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Carcinogenesis Advance Access originally published online on January 3, 2008
Carcinogenesis 2008 29(2):321-332; doi:10.1093/carcin/bgm276
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© The Author 2008. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

Take a break—resveratrol in action on DNA

Susanne A. Gatz and Lisa Wiesmüller1,*

University Children's Hospital, Eythstrasse 24, D-89075 Ulm, Germany
1 Department of Obstetrics and Gynecology, University of Ulm, Prittwitzstrasse 43, D-89075 Ulm, Germany

* To whom correspondence should be addressed. Tel: +49 731 50058800; Fax: +49 731 50058810; Email: lisa.wiesmueller{at}uni-ulm.de


   Abstract
Top
 Abstract
Introduction
 Effects of RV on...
 Modulation of DNA damage...
 Conclusions
 Funding
 References
 
The phytochemical resveratrol (RV) has become a focus of intense research owing to its roles in promoting longevity and in cancer prevention. As an anticancer agent, RV has primarily been linked to growth and death regulatory pathways. There is now growing evidence that, under physiological conditions, RV additionally contributes to the maintenance of genome stability. Thus, at the stage of DNA damage formation, RV protects the genome as an antioxidant via inhibition of inflammation, suppression of metabolic carcinogen activation, de novo expression of genes that encode detoxifying proteins and possibly even via radical scavenging properties. However, results demonstrating RV-dependent DNA breakage in the presence of Cu(II) ions and inhibition of DNA polymerases {alpha} and {delta} produced some controversy regarding RV's role as a caretaker compound. Significantly, recent studies have revealed that activation of ataxia telangiectasia mutated and ataxia telangiectasia Rad3 related could be a central effect of RV that underlies cell-cycle regulation and the newly described activation of fidelity control mechanisms in DNA double-strand break repair involving Nbs1 and p53. In this review, we discuss the existing data on RV's direct and indirect effects on genome integrity, in the light of future chemopreventive and chemotherapeutic protocols involving RV or related compounds.

Abbreviations: ATM, ataxia telangiectasia mutated; ATP, adenosine triphosphate; ATR, ataxia telangiectasia Rad3 related; APE/Ref-1, apurinic/apyrimidinic endonuclease-1/redox factor-1; BER, base excision repair; COX, cyclooxygenase; CYP450, cytochrome P450; DSBs, double-strand breaks; GE, genistein; HR, homologous recombination; hTert, human telomerase reverse transcriptase; MN, micronuclei; MGMT, O6-methylguanine-DNA methyltransferase; NADPH, nicotinamide adenine dinucleotide phosphate; NF-{kappa}B, nuclear factor-{kappa}B; NHEJ, non-homologous end-joining; NO, nitric oxide; NOS, nitric oxide synthases; PI3K, phosphatidylinositol 3-kinase; PKC, protein kinase C; pol, polymerase; PTEN, phosphatase and tensin homolog; QR, quinone reductase; QU, quercetin; RNS, reactive nitrogen species; ROS, reactive oxygen species; RR, ribonucleotide reductase; RV, resveratrol; SCE, sister chromatid exchange; Sir2, silent information regulator 2; topo, topoisomerase


   Introduction
Top
 Abstract
 Introduction
Effects of RV on...
 Modulation of DNA damage...
 Conclusions
 Funding
 References
 
The genome of mammalian cells is constantly exposed to various harmful influences stemming from environmental sources (e.g. sunlight, cigarette smoke and pollution) or from normal processes in the organism (metabolic intermediates and products, errors in replication, recombination and mitosis). Spontaneous occurrence of DNA damage with e.g. 10 000 depurinations/day x cell (1) or 50 endogenous DNA double-strand breaks (DSBs) repairs /replicating cell x cell cycle (2) is, indeed, a common event in the life of a cell. The immediate consequences, if repair mechanisms fail, are gene mutations, genome rearrangements and apoptosis. However, mammalian cells exhibit a multitude of mechanisms to guarantee the maintenance of genome stability.

These mechanisms include caretaker pathways, which are directly involved in DNA damage removal. More indirectly acting gatekeeper pathways give time for DNA repair through cell-cycle arrest or inhibit the manifestation of DNA damage through apoptosis induction.

Carcinogenesis is a multistep process that can be roughly divided into tumor initiation, when the first genome mutations are set, tumor promotion and tumor progression (3,4). Over the last years, data emerged indicating that natural constituents of the regular diet influence the process of carcinogenesis. By means of their action such phytochemicals can be divided into primarily tumor-blocking and tumor-suppressing agents. Tumor-blocking agents prevent DNA damage formation or promote DNA damage removal, and tumor suppressing agents slow down the process of initiated cells to become invasive cancer cells. This separation is not strict and several chemopreventive agents are shown to act in both categories (3–5). This is particularly true for the most thoroughly investigated chemopreventive food constituent resveratrol (RV) (4–8). Since the early work of Pezzuto et al. (6) 10 years ago, it has become an accepted fact that RV decelerates carcinogenesis via these steps.

More specifically, as an anticancer agent RV turned out to be involved in the maintenance of genomic stability, through direct and indirect mechanisms. A large body of evidence has accumulated to suggest influence on the indirect gatekeeper pathways controlling cell cycle and apoptosis, and most reviews published on RV so far focused on these activities. Here, we cover recent findings on the more direct genome stabilizing roles of RV in DNA damage formation, removal and signaling. However, DNA repair and checkpoint activation are intricately linked. Consequently, a complete disentanglement of the pathways is hard to achieve and often might not convey the physiologic relevance. Nevertheless, we made an effort to discuss indirect and direct effects of RV on genomic integrity separately. In the following chapter, we briefly recapitulate effects on the cell cycle and apoptosis. The chapters thereafter will provide in-depth analyses of the different actions of the stilbene RV (3,4',5-trihydroxy-trans-stilbene) on DNA. In addition, activities of the flavonoids genistein (4',5,7-trihydroxyisoflavone, GE)—an isoflavonoid—and quercetin (3,3',4',5,7-pentahydroxyflavone, QU)—a flavonol—will be discussed, as far as they are relevant with respect to the main focus of this review, i.e. physical and functional interactions with relevance for DNA repair (Figure 1, Table I).


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  		Table I. Physical interactions and regulation of enzymatic activities with relevance for DNA repair

 


Figure 1
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  		Fig. 1. Chemical structure of RV, GE and QU.

 
It is important to note beforehand that RV dosage varied dramatically from study to study and often lacked physiological relevance: In in vitro studies, RV concentrations of 15–100 µM were typically applied and, in fact, were required to inhibit proliferation and to induce cell death in most studies (33,34). In human beings, due to the low bioavailability, concentrations of unmodified RV in the lower nanomolar and micromolar range were estimated to be reached after moderate consumption of red wine and pharmacological treatment, respectively (34,35). However, in vitro and in vivo responses to the same dose did not necessarily correlate (35). Moreover, up to 10–20 times higher concentrations (i.e. 20–40 µM) were detected in animal studies after intravenous administration (35). Unmodified RV has a short half-life and is extensively metabolized in the body, and some flavonoids like QU may increase the bioavailability of RV by inhibition of its glucuronidation. This is why available data are insufficient to predict peak levels of unmodified RV in most tissues. Moreover, although in vivo concentrations of RV metabolites can be >10 times higher, it is not clear in how far these metabolites can take over the effects attributed to RV (8,34,35). Nevertheless, several studies in animals have been performed and indeed document chemopreventive and even chemotherapeutic effects of RV to, however, highly variable extent (34–36). Since the end points of most of these studies were tumor initiation, cell proliferation and apoptosis, in-depth discussion of these in vivo studies is beyond the scope of this review.


   Effects of RV on growth and death regulatory pathways
Top
 Abstract
 Introduction
 Effects of RV on...
Modulation of DNA damage...
 Conclusions
 Funding
 References
 
RV has been demonstrated to mediate cell-cycle arrest and apoptosis induction via the main signal transduction pathways for cell survival. Thus, RV mostly inhibits signaling through mitogen-activated protein kinase and phosphatidylinositol 3-kinase (PI3K)/AKT pathways (4,5,7,8,33). Consistently, RV suppresses the activity of the downstream transcription factors AP-1 and nuclear factor-{kappa}B (NF-{kappa}B) (RelA/p65) (4,5,7,8). The list of genes well described to be transcriptionally influenced by RV with impact on the cell cycle and apoptosis encompasses cyclins, cyclin-dependent kinases, caspases, p53, p21(Cip1/WAF1), p300, NF-{kappa}B, bcl2, bax and IAPs (5,7,8). Changes in posttranslational modifications have also been documented for RV-treated cells, particularly regarding protein phosphorylation and acetylation (5,7,8 and discussed in The ataxia telangiectasis mutated (ATM)/ataxia telangiectasia Rad3 related (ATR)-signaling network and The sirtuin connection). Regarding the effect of RV on the cell cycle, highly divergent observations have been made because they heavily depend on the cell type investigated, dosage and treatment protocol. In most cases, RV caused a reversible cell-cycle arrest in S- or G1 phase. Concerning apoptosis induction by RV, receptor- and mitochondria-mediated mechanisms seem to be involved: RV induces apoptosis through the CD95/CD95L interaction, through translocation of Bax (7,37) and inhibitor of apoptosis protein depletion (38,39). More recently, RV and other phytochemicals have been documented to also suppress the STAT3 pathway, which leads to cell-cycle arrest and apoptosis (40–42; for review see ref. 43). Despite these observations, the apoptosis inducing effect of RV alone is weak, but potentiates the apoptotic effects of cytokines (e.g. tumor necrosis factor-related apoptosis-inducing ligand) and chemotherapeutic agents (39,44,45).


   Modulation of DNA damage formation, removal and signaling by RV
Top
 Abstract
 Introduction
 Effects of RV on...
 Modulation of DNA damage...
Conclusions
 Funding
 References
 
The impact of RV on genome stability is highlighted by findings indicating that RV affects all aspects of DNA metabolism, i.e. DNA replication, recombination, repair, relaxation and telomere maintenance (Figure 2). New clues to the underlying mechanisms have come from a number of reports suggesting physical and biochemical interactions between RV and DNA. Aside from direct interactions with DNA, RV also influences the redox state of cells and thereby indirectly modulates the integrity of genomic DNA.


Figure 2
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  		Fig. 2. Nuclear activities of RV with relevance for DNA repair. Gray color marks the immediate targets of RV.

 
Metabolic sources of DNA damage.
Reactive oxygen species (ROS), such as hydrogen peroxide (H2O2), superoxide anion (Formula) and hydroxyl radical (.OH), are ubiquitous within the cell. Free radicals interact with proteins, lipids and DNA and lead to oxidative damage. The resulting DNA lesions include base modifications, base loss, single-strand DNA breaks and through single-strand DNA breaks clustering even DSBs (46).

ROS are exogenously generated by {gamma}-ray and ultraviolet light irradiation and endogenously by the normal cellular metabolism and by inducible membrane-bound enzyme systems such as the nicotinamide adenine dinucleotide phosphate (NADPH) oxidase complex. The principal source of cellular ROS is mitochondrial respiration since adenosine triphosphate (ATP) synthesis is coupled with continuous electron leakage (47). Aside from the electron transport chain in mitochondria, low levels of ROS are formed in peroxisomes and by cytochrome P450 (CYP450). CYP450 represents the main phase I enzyme that together with phase II enzymes such as UDP-glucuronyltransferase and NADPH:quinone oxidoreductase catalyzes the metabolic conversion of a multitude of endogenous substrates and detoxification of xenobiotic reagents. Biotransformation bears certain risks: procarcinogens are converted into carcinogens; ROS and reactive nitrogen species (RNS) arise as by-products. Moreover, ROS and RNS levels are also elevated during inflammation by neutrophils, eosinophils and macrophages. During this pathophysiological process, nitric oxide (NO.) is produced and reacts with ROS to form the RNS peroxynitrite (ONOO–), a highly oxidizing and nitrating substance. NO itself reacts with proteins, prosthetic groups and DNA (pyrimidine bases). To maintain the physiological redox potential, ROS and reaction products are eliminated by sophisticated intracellular defense mechanisms involving the antioxidatve enzymes superoxide dismutase, catalase, glutathione peroxidase, the protein-specific oxidoreductases, the copper and iron transport and storage proteins as well as low-molecular-weight antioxidants like glutathione, vitamin C and uric acid (3,48,49).

RV as an antioxidant.
One of the most direct effects of cancer-blocking agents is to quench ROS by an intrinsic antioxidant capacity. In fact, it was speculated that RV would do big parts of its job as an inhibitor of tumor initiation through this effect (6). In support of this idea, in vitro studies revealed that RV can act as radical scavenger in hydroxyl, superoxide and transition metal-based radical generating systems, although the physiological relevance of these investigations is questionable due to the very high doses of RV (≥325 µM) used (50). RV itself can be oxidized by peroxidases, i.e. competes with the redox probe in several assays used to prove scavenging activities. Excluding this bias in a cell-based system, Poolman et al. (51) detected antioxidative activity of RV in monocytes even at moderate RV concentrations of 0.1–50 µM. Concomitantly, RV interfered with stress signaling via the PI3K/AKT pathway and thereby with cellular NADPH oxidase activity. Moreover, antioxidants including RV are known to induce de novo expression of genes that encode detoxifying proteins like superoxide dismutase, catalase and glutathione peroxidase (4,34).

All in all, these results show that RV elicits antioxidative effects, but it remains a matter of debate whether, in vivo, RV possesses radical scavenging properties, modulates downstream protective mechanisms or both (34). In this context it should be noted that some antioxidants, among them several plant phenols, turn into pro-oxidants under certain in vitro conditions, particularly via interaction with transition metal ions (48). At low concentrations (4–8 µM), a pro-oxidant effect was also demonstrated for RV in HL-60 leukemia cells and was characterized by NADPH oxidase-dependent elevation of intracellular superoxide (52). Depending on treatment conditions and the intracellular milieu, RV might, therefore, be able to act as antioxidant or pro-oxidant and to differentially influence transcription factors and cellular signaling cascades, which are extremely sensitive to even subtle changes in the redox setting (53).

Modulation of oxygen metabolism.
Several pieces of evidence indicate that RV affects the mitochondrial respiration chain. In rat brain, RV inhibited complexes I–III and complex V comprising the F0F1-ATPase/ATP synthase with an EC50 of 23 µM (14). Enzymatic inhibition of the ATPase/ATP synthase by RV was confirmed by other groups (IC50 value 6–28 µM), extended to rat liver, rat heart, the bovine enzyme and to other phytochemicals like GE and QU and demonstrated for both RV and GE to be non-competitive (15–17; Table I). Using high-resolution crystallography, Walker et al. (17) provided a structural basis for the inhibition of F0F1-ATPase/ATP synthase by RV. Although the authors speculated that inhibition of the enzyme would cause enhanced ROS production and apoptosis induction in tumor cells (17), Watabe et al. (54) showed that this is not the case in neuroblastoma cells treated with the well-known F0F1-ATP synthase inhibitor oligomycin. Moreover, Zini et al. (55) even saw a protective effect on ROS production by RV at a concentration as low as 1 nM.

On the other hand, when mice were fed on a RV-containing diet, they showed an increase in mitochondria in the liver (56), brown adipose tissue and muscle fibers (57). This increase was accompanied by transcription and deacetylation of peroxisome proliferator-activated receptor-{gamma} coactivator 1{alpha}, which is known to upregulate genes for oxidative phosphorylation and mitochondrial biogenesis. Furthermore, RV (10 µM) has been demonstrated to activate adenosine monophosphate-activated protein kinase-mediated mitochondrial biogenesis in neurone cells in a manner depending on the upstream kinase LKB1 (58). In support of the idea that RV modulates signaling upstream of adenosine monophosphate-activated protein kinase, activation of adenosine monophosphate-activated protein kinase through phosphorylation was also reported by another group, here in the human hepatocyte cell line HepG2 after RV treatment (10 µM) (59). In accord with these findings, chronic treatment (≥10 days) with subapoptotic concentrations of RV was recently shown to induce senescence in various cancer cell lines due to a rise in endogenous ROS largely originating from mitochondria (60). Taken together, both scenarios, prevention and stimulation of ROS production via mitochondria are possible. Future studies will have to answer which effect is relevant for normal versus cancer cells.

Xenometabolism.
Procarcinogenic compounds like the aryl hydrocarbon 7,12-dimethylbenz[a]anthracene and the polycyclic aromatic hydrocarbon benzo[a]pyrene require activation mainly by phase I enzymes to form covalent DNA adducts and thereby to develop their carcinogenic potential. Induction of phase II enzymes, on the other hand, is critical for removal of electrophilic and oxidative toxicants from the cell before they are able to damage the DNA. Suppression of metabolic activation of carcinogens (phase I) and enhancement of their detoxification (phase II) are two important features of a cancer-blocking chemopreventive agent (3,4). RV has been demonstrated to fulfill both criteria in several ways: Utilizing intact cells, microsomes or recombinant CYP450 proteins, RV was demonstrated to abrogate the enzymatic activities of different CYP450 proteins (Ki = 1 µM for CYP450 1A1 and CYP450 1B1 and 16 µM for CYP1A2) (61,62; reviewed in refs 8,33,34). RV also inhibits CYP1A1 expression, both at the level of basal and aryl hydrocarbon receptor-mediated transcription (61,63).

RV was further shown to enhance expression of phase II enzymes like hem oxygenase 1 and quinone reductase (QR) 1 in vitro and in vivo (reviewed in ref. 34). The cytosolic flavoenzymes QR1 and QR2 catalyze a two-electron reduction of quinones to hydroquinones, which effectively prevents the formation of ROS by other QRs (64). At first sight somewhat contradictory, RV was identified as a strong inhibitor of QR2. With a Kd of 35 nM, QR2 even represents the target molecule of RV with the highest affinity reported so far (18; Table I). Three-dimensional structure analysis on QR2–RV cocrystals confirmed the results from binding studies, as it showed specific binding to the unique active-site cleft in QR2. Other natural polyphenols also fit into the binding pocket and, indeed, high-affinity binding was demonstrated for QU (18,65; Table I) and also for the hormone melatonine and the antimalaria drug chloroquine (65). Interestingly, Buryanovskyy et al. (18) noticed that RV-treated K562 erythroleukemia cells displayed the same increase in antioxidant and detoxification enzyme expression as was observed in K562 cells upon RNA interference-mediated QR2 knockdown. The authors, therefore, speculated that inhibition of QR2 increases the response of other antioxidant and phase II enzymes after oxidative stress, which would be in line with RV's role as an anticarcinogenic agent.

Inflammation.
Generation of ROS and other activated species is a major mechanism during inflammation. The enzymes cyclooxygenase (COX) and lipoxygenase, which generate eicosanoids from arachidonic acid, are central to the inflammatory process and of particular clinical relevance. RV is known to influence both enzyme classes. The major COX enzymes are the constitutive form COX1, which produces prostaglandins involved in tissue homeostasis, and COX2, which is inducibly expressed during inflammation and neoplasia. COX1 and COX2 suppress apoptosis and promote DNA synthesis, cell proliferation, invasiveness, angiogenesis and thereby carcinogenesis (66). In vitro studies demonstrated that RV inhibits both COX (ED50 15 µM) and hydroperoxidase activity (ED50 3.7 µM) of COX1. While RV caused enzymatic inhibition of COX2 only at much higher doses (hydroperoxidase, ED50 85 µM) or not at all (COX) (6), it downregulated COX2 at the messenger RNA level. NF-{kappa}B and protein kinase C (PKC) pathways are good candidates as mediators for this effect of RV on COX2 expression (reviewed in ref. 7,8,33). RV suppresses the activity of the upstream transcription factor NF-{kappa}B (see Effects of RV on growth and death regulatory pathways) and intervenes with phorbol myristate acetate treatment-triggered and PKC-mediated COX2 promoter activation (67–69). The two-enzyme activities of lipoxygenase, dioxygenase and hydroperoxidase, are differentially regulated by RV. Whereas the dioxygenase activity is enzymatically inhibited by RV with an IC50 value of 5–40 µM, the hydroperoxidase function is not altered (70,71).

Besides the enzymes COX and lipoxygenase, the NO metabolism, which also plays a critical role in inflammatory reactions, is influenced by RV. Three nitric oxide synthases (NOS) exist. The endothelial NOS and neuronal NOS are constitutively expressed and are important for generation and maintenance of low basal levels of NO. Inducible NOS is found in various cell types including epithelial cells, macrophages and a variety of cancer cells; its expression is induced by activation of several transcription factors, among them the RV targets NF-{kappa}B and AP-1 (49). RV inhibits NO production and inducible NOS expression in cancer cells and lipopolysaccharide-stimulated macrophages (8). In conclusion, RV counteracts key inflammatory reactions, which are directly or indirectly associated with ROS or RNS formation, suggesting that RV decreases the load of these DNA reactive substances both at the level of the cell and the organism.

Direct DNA interactions and breakage
In vitro studies applying ultraviolet absorption spectral analysis and Fourier transform infrared spectroscopy revealed that RV directly binds DNA and RNA, which probably involves H-bonding (9,10,72; Figure 2). After RV binding, the B-form of DNA and the A-family conformation of RNA were unaltered (10). Comparable results were obtained with the chemopreventive agents GE and QU (10,11; Table I). Along this line, it could further be demonstrated that DNA intercalation by the dyes ethidium bromide and acridine orange was significantly reversed by RV (50 µM) and GE (100 µM) (73). In this context, it should be noted that in contrast with several other polyphenols (including QU), RV and GE themselves are not DNA intercalators (12; Table I). From these observations, it is conceivable that the interactions of RV (and GE) that stabilize the double-helical structure of the DNA represent the molecular basis that protects against genotoxic effects by these and other mutagenic drugs, i.e. form the basis of RV's function as antimutagen.

On the other hand, in vitro experiments with circular plasmid and with calf thymus DNA showed that RV mediates DNA strand scission (9,74,75). In addition to DNA nicking, Subramanian et al. (76) even saw DSB induction by RV. However, these activities turned out to be absolutely dependent on the presence of Cu(II) and oxidative conditions. Further data indicated that RV promotes hydroxyl radical formation by DNA-bound Cu(II) ions was able to reduce Cu(II) to Cu(I) and that Cu(I) is a requisite intermediate in this process. Other metal ions could not substitute for Cu(II) (9,74–77). In the absence of Cu(II), RV did not induce DNA cleavage at concentrations of up to 100 µM or even 900 µM (9,50). Given that RV binds DNA, it was, then, proposed that a ternary complex between RV, Cu(II) and DNA is formed, which facilitates target delivery of reactive oxygen to DNA (9,75,77–79). For comparison, GE in combination with Cu(II) did not cause DNA nicking, rather it even reduced DNA cleavage induced by hydrogen peroxide or hydroquinone, indicating that GE effectively scavenges relevant reaction intermediates (77).

Consistent with DNA cleavage, transformation efficiencies with RV/Cu(II)-treated plasmid DNAs decreased upon transfer into competent bacteria. Sequencing of the bacterially amplified DNA revealed point mutations, in specific, mostly deletions of guanine nucleotides (78). Copper ions are known to specifically associate with DNA bases, particularly with guanine bases (80). Mobilization of such endogenous copper by RV may, thus, result in pro-oxidant DNA cleavage and mutagenesis at these sites. In vitro, copper concentrations of 25–50 µM were necessary to cause effective DNA cleavage by RV (9). For comparison, the copper level in blood is ~16 µM. Since in cancer cells the level is known to be even higher, it was hypothesized that RV's anticancer activity could be a consequence of tumor cell-specific DNA degradation (78). However, the physiological significance of the in vitro findings was questioned because in the presence of ascorbic acid or glutathione, RV did not induce DNA cleavage and rather switched to its antioxidant performance (74). The relevance of these copper-dependent effects seems even more questionable since in healthy humans, metal ions appear to be largely sequestered and cannot catalyze free radical reactions (48). Raising further doubts about the physiological significance, RV (<200 µM) treatment of vital human blood lymphocytes did not lead to significant DNA breakage as judged from tail length changes in the comet assay (81). Only upon use of RV at a concentration of 200 µM or, as in the in vitro assays, of 50 µM RV together with Cu(II), RV-dependent DNA damage was noticeable (82). Nevertheless, since the DNA cleaving activity of 200 µM RV was reversed by exogenously added radical scavengers or the Cu(I)-specific sequestering agent neocuproine, the authors conclude that RV in high doses leads to DNA breakage by oxidative damage through endogenous copper mobilization (81). Therefore, indeed, endogenous copper mobilization could be involved in RV's influence on DNA damage induction in experiments utilizing high RV concentrations.

DNA metabolism
Inhibition of enzymes required for DNA synthesis: polymerases {alpha} and {delta}, ribonucleotide reductase.
DNA polymerases (pols) are required for de novo synthesis of undamaged DNA and additionally appear to be involved in repair replication of damaged DNA. The B-family members {alpha}, {delta} and {varepsilon} represent the eukaryotic replicative pols (83). Pols generally accepted to be directly implicated in DNA repair in mammalian cells are the X-family pols β [base excision repair (BER)], µ and {lambda} [non-homologous end-joining (NHEJ)] (84). From studies in yeast, it is known that the B-family pols {alpha}, {delta} and {varepsilon} are required for DNA synthesis during DSB repair, pols {delta} and {varepsilon} during nucleotide excision repair and BER and pol {delta} during mismatch repair (83,85).

A first hint for RV's inhibitory potential on replicative pols came from experiments using the SV40 DNA replication bioassay in CV-1 cells (86). Subsequently, this finding was verified for pols {alpha} and {delta} by in vitro polymerase assays with isolated enzymes (20,21). Biochemical characterization by Locatelli et al. (21) showed that RV is a non-competitive inhibitor of pol {alpha} with respect to the reaction substrates. Indicating a high specificity of inhibition, activities of pols β and {lambda} were not compromised (21). For comparison, QU is a very potent inhibitor of pol β (22; Table I, Figure 2).

Mammalian ribonucleotide reductases (RRs) are tetrameric enzymes ({alpha}2β2), which convert the four standard ribonucleotides into their 2'-deoxyribonucleotide counterparts, thereby providing the precursors for DNA synthesis and repair (87). The large homodimer {alpha}2 (R1) harbors the active, substrate-binding site, and the small β2 homodimer (R2) complexes two iron ions. The radical needed for reduction of the ribonucleotides is generated on a tyrosine residue in R2 close to the di-iron center deeply inside the subunit protected from the solvent (87). Inhibition of R2 by tyrosyl radical scavenging, as known for hydroxyurea, disturbs normal DNA replication and causes replication fork arrest.

The tyrosyl radical was totally destroyed after co-incubation of pure recombinant murine R2 protein with as little as 10 µM RV (23). However, enzyme inhibition studies with cellular extracts of L1210-R2 murine leukemia cells revealed an IC50 value of 100 µM (Table I, Figure 2). The authors speculated that cell lines with limiting amounts of R2 rather than with 15- to 20-fold R2 overexpression as in L1210-R2 should reveal a lower IC50 value. However, for the well-established RR inhibitor hydroxyurea, an IC50 value of 1 mM was determined in the L1210-R2 extract, i.e. in the concentration range previously reported for hepatoma cells (88). This suggests that the data obtained with L1210-R2 cells are generally valid. For comparison, [3H]thymidine incorporation studies revealed IC50 values of 8–10 µM for RV in K562 cells (23). This concentration range argues for polymerase rather than RR inhibition as the cause underlying replication inhibition (Table I). However, since [3H]thymidine incorporation as the end point does neither discriminate between the inhibitory influence of RV on polymerases and RRs nor between these enzymes and other effects of RV such as on the cell cycle (see Effects of RV on growth and death regulatory pathways), it is not yet clear which precise mechanism causes the replication inhibition observed.

Topoisomerases.
Topoisomerases (topos) function to maintain normal DNA topology, a critical requirement for each cellular activity that involves DNA strand separation, including DNA replication, recombination and RNA transcription. Furthermore, they take part in condensation and decondensation of chromosomes (89). Three different classes of topos have been described in mammals, type IA (comprising topos III{alpha} and β), IB (comprising topo I) and II (comprising topo II). Type I topos are monomeric enzymes, which ATP independently relax the DNA double helix by introducing a single-strand DNA break, subsequently passing a single DNA strand through the break and rejoining the DNA. Type II topos are homodimeric proteins. They catalyze an ATP-dependent process, which introduces DSBs into DNA and enables another intact DNA double helix to pass through the break (89).

Two different classes of topo inhibitors have been described. Class I drugs prevent completion of the topoisomerization reaction by stabilizing the enzyme–DNA complex and are referred to as topo poisons, whereas class II drugs are catalytic inhibitors that interfere with the enzymatic function without DNA strand break formation. In two studies, RV was characterized as a class II (catalytic) inhibitor of topo II with an IC50 of 66–79 µM (24,27; Table I, Figure 2). Activities of RV toward topo I or topo II as a poison were not detected up to doses of >400 µM (24). This is in disagreement with one report postulating topo I poisoning activity for RV (12). However, only a limited number of experiments were performed in this study. For comparison, QU was demonstrated to be a catalytic inhibitor of both, topo I and topo II, and GE acts as an effective topo II poison and catalytic inhibitor (24; Table I). Interestingly, the clastogenicity of GE, which is provoked by its topo II poison activity, was antagonized by mere topo II catalytic inhibitors like biochanin, galangin and daidzein in V79 Chinese hamster cells (90). A similar antagonistic activity is, therefore, expected for the catalytic inhibitor RV.

Telomerase.
Telomeres are DNA–protein complexes that cap the end of linear eukaryotic chromosomes preventing them from degradation, recombination, fusion with other chromosomes and suppressing DSB signaling (91). The maintenance of functional telomeres is crucial for continued proliferation and—with the exception of certain tumor cells—dependent on the ribonucleoprotein complex telomerase. Telomerase is an RNA-dependent DNA polymerase with reverse transcriptase activity that adds hexameric repetitive sequences (TTAGGG) to chromosome ends (92).

Telomerase activity was recently shown to be downregulated after treatment with relatively high doses of RV (64% at 88 µM for 48 h or 67% at 175 µM for 24 h) in human breast cancer MCF-7 cells; some inhibition (up to 19%) was also seen after treatment with lower doses (22–44 µM) (29; Table I, Figure 2). Downregulation was confirmed in human colon carcinoma HT-29 and WiDr cells (93). Since the rate-limiting telomerase subunit, human telomerase reverse transcriptase (hTert), has to shuttle from the cytoplasm to the nucleus for holoenzyme assembly (91), it was of interest that elevated hTert levels were found in the cytoplasm of RV-treated MCF-7 cells, while nuclear accumulation failed (29). RV is known to inhibit PKC, Protein kinase B/AKT and NF-{kappa}B pathways, which play a role in regulating hTert through phosphorylation and nuclear shuttling, respectively (91). Therefore, RV's effects on telomerase are probably indirect via these pathways. For comparison, GE was demonstrated to inhibit telomerase activity through inhibition of AKT-dependent hTert phosphorylation (30) and to suppress hTert transcription via repression of the transcription factor c-Myc (30,31; Table I). If RV exhibits a specific effect on hTert transcription has not yet been examined. However, PKC and NF-{kappa}B were shown to promote hTert transcription via increased c-Myc promoter binding (94). Moreover, RV has been documented to suppress c-Myc activity (41,95) and to inhibit STAT3, an important activator of c-Myc (43). Therefore, it is conceivable that RV also antagonizes hTert transcription via these molecular targets. Given that hTert is frequently overexpressed in tumor cells, hTert is an interesting target for therapeutic approaches involving polyphenolic compounds.

DNA damage and repair
The cellular mechanisms, which are responsible for the removal of DNA damage, encompass BER, nucleotide excision repair, mismatch repair and, following DSB formation or stalling of replication forks, the DSB repair pathways NHEJ and homologous recombination (HR) (96). From the results of genotoxicity studies and molecular target screens, RV has been implicated in the modulation of DNA damage formation and removal.

Indirect evidence for RV-mediated effects on DNA repair.
Several studies have been carried out to study RV's influence on the integrity of the genome, but gave ambiguous results. Explanations for these discrepancies are probably to stem from the diverse assay principles applied. Classically, assessment of genomic damage and its removal has made use of a series of genotoxicity assays that rely on end points such as appearance of DNA breaks, micronuclei (MN), sister chromatid exchanges (SCEs) or chromosome aberrations. Unfortunately, these end points do not tell much about the causative role of specific DNA repair mechanisms. Thus, MN formation and SCEs can be induced by a large spectrum of genotoxic treatments that enroll multiple DNA repair pathways. DNA breaks scored in the comet assay (single-cell gel electrophoresis) may reflect incomplete BER, nucleotide excision repair or DSB repair and are not informative about the quality of the DNA repair process (97). Moreover, indirect effects on DNA repair by changes in growth characteristics or cell lethality cannot easily be ruled out, which is even true for the majority of HR assay systems, as they rely on clonal outgrowth in selective medium. This aspect together with the estrogenic activity of RV in promoting growth in estrogen receptor-positive cells (98) emphasizes the need of appropriate model systems and the danger of overinterpreting data obtained with individual cell lines. Despite the assay limitations listed, the studies summarized in the following provided first hints for a potential regulatory role of RV in the maintenance of genomic stability.

Stopper et al. (98,99) observed induction of MN in L5178Y mouse lymphoma and Chinese hamster V79 cells by RV. Consistently, Matsuoka et al. (100,101) scored a dose-dependent increase of MN in Chinese hamster lung cells for up to 44 µM RV and additionally detected SCEs and chromosomal aberrations. Notably, however, RV treatment in Chinese hamster lung cells was paralleled by S-phase cell-cycle arrest and apoptosis induction, which is why these data have to be interpreted with caution. In the Ames test, which measures the reversion of mutations in bacteria, RV was negative (100). This reduces the likelihood that RV-induced MN formation in mammalian cells was caused by mutagenic base damage. When Matsuoka et al. (102) used SPD8/V79 Chinese hamster cells to assay HR between partial hprt gene duplications, they found recombination stimulation with a concentration optimum ~20 µM RV, as was similarly observed for SCEs. This observation led the authors to hypothesize that in analogy to hydroxyurea, RV blocks replication forks through inhibition of RR (see Inhibition of enzymes required for DNA synthesis: polymerases {alpha} and {delta}, ribonucleotide reductase), thereby initiating recombination and SCEs.

Genotoxic effects were also seen with GE, whereas contradictory results were obtained with QU (99). GE treatment consistently caused DNA breakage in the comet assay (99), which was convincingly demonstrated also for QU in a recent study applying the neutral comet assay on CD34+ cells (103). In most of the studies, RV prevented break formation probably due to its antioxidant activities (see Metabolic sources of DNA damage). Additionally, RV may accelerate DNA repair as was indicated by the observation of unscheduled DNA synthesis together with protection against DNA breakage after treatment with the alkylating agent N-methyl-N'-nitro-N-nitrosoguanidine (104). Taken together, straightforward conclusions on the mechanistic basis for the observed effects on genome stability cannot be drawn, as they very much depend on the chemopreventive agent chosen and the treatment conditions applied.

DNA repair factors as molecular targets of RV.
Indications that RV treatment induces the expression of DNA repair genes were provided by studies in the human breast carcinoma cell lines HBL100, MCF-7 and MDA-MB-231 and the non-malignant breast epithelial cell line MCF10A (105,106). Treatment with 30–50 µM RV raised messenger RNA levels in all four cell lines for the DSB repair surveillance genes BRCA1, BRCA2 and TP53 and in all lines except MCF-7 for the genes encoding the HR recombinase Rad51 and the chromatin factor p300. Given that MCF10A cells are estrogen receptor-negative cells, RV appears to alter transcription of these genes independently of its estrogen-like properties. Pathways responsible could involve AP-1 or NF-{kappa}B (4,5,7,8). Contrary to the transcriptional profiles, protein expression was unchanged for BRCA1 and BRCA2 in the same breast carcinoma cell lines (105). However, concerning p53, several investigators observed RV-induced protein accumulation. This increase often correlated with phosphorylation of p53 on serine 15, although enhancement of p53 phosphorylation without a rise in protein expression was also observed (7,8). Accumulation of p300 was documented for the prostate carcinoma cell line LNCaP after treatment with 100 µM RV for 24–48 h (107). The Rad51 protein was found to be upregulated in RV-treated lymphoblastoid cells (S.A. Gatz, M. Keimling, C. Baumann, T. Dörk, S. Fulda, and L. Wiesmüller, Carcinogenesis, in press). In this context, it is interesting to note that exposure of MCF-7 cells to even low concentrations (0.1–1000 nM) of RV, GE and QU increased phosphatase and tensin homolog (PTEN) protein levels up to 3-fold (108), although a strict dose dependency was not observed. PTEN is commonly known as tumor suppressor, which inhibits PI3K and thereby antagonizes the proliferation-stimulatory AKT-dependent pathways (109). Very recently, PTEN has additionally been demonstrated to play a fundamental role in the maintenance of chromosomal integrity through physical interactions with centromeres and via upregulation of Rad51 protein (109). In conclusion, RV may contribute to genomic stability through a rise in Rad51, the central enzyme involved in the safest DSB repair pathway HR (110), and in p53, which controls the fidelity of HR (111–113; Figure 2).

Apart from the above-mentioned DSB repair factors, potential links between RV and O6-methylguanine-DNA methyltransferase (MGMT) have been investigated (114). This enzyme directly removes alkylating lesions at O6 of guanine. MGMT represents the first line of defense against the permanent threat of alkylation damage of DNA from metabolic (endogenous) and common environmental (exogenous) sources. On the other hand, in tumors, MGMT confers resistance to chemotherapy involving alkylating agents (115). While searching for novel MGMT-targeted chemoprevention strategies, Niture et al. (114) identified several cysteine/glutathione-enhancing drugs and several plant antioxidants, which led to an up to 3-fold increase in the enzymatic activity of MGMT in different cell lines. Induction of MGMT expression was noticeable both at the messenger RNA and the protein level. RV (10 µM) caused only a marginal, 1.2-fold increment in enzyme activity in HT29 colon carcinoma and UW228 medulloblastoma cells. Clear changes in protein expression, as determined by western blotting, were not detectable (114). These data argue against a major impact of RV on the direct repair pathway of DNA base damage executed by MGMT.

Another attractive target for chemoprevention and tumor therapy studies is apurinic/apyrimidinic endonuclease-1/redox factor-1 (APE/Ref-1), which coordinates DNA repair and cell-cycle regulatory pathways (116). Mammalian APE/Ref-1 is indispensable for life because it has essential functions in BER. APE/Ref-1 additionally engages in redox regulation of many transcription factors like NF-{kappa}B, AP-1 and p53, which is also involved in DSB repair and BER (112,113,116). BER is an important mechanism for the removal of oxidized/reduced, alkylated, deaminated bases and of base mismatches. Following the removal of the damaged base, APE/Ref-1 catalyzes the incision step 3' to the AP site and, through physical interactions with downstream enzymes, promotes the gap tailoring, DNA synthesis and sealing steps in both short-patch and long-patch BER (96).

Very recently, APE/Ref-1 was identified as a new potential molecular target of RV in a compound screen based on three-dimensional structure modeling (19). The proposed interaction site for RV was located in the redox regulation domain within the human APE/Ref-1 molecule. To investigate the functional consequences of the putative RV–APE/Ref-1 interaction, the authors prepared nuclear extracts from, unfortunately, only one particular melanoma cell line (c83-2c) with increased APE/Ref-1 expression and performed DNA cleavage and DNA-binding assays. Conversion of the supercoiled form of depurinated plasmid DNA into the nicked form, which is indicative of APE/Ref-1 incision activity, was found to be reduced after incubation with RV at doses ≥20 µM. It is of interest that no effect on APE/Ref-1 expression or enzyme activity was noticeable after RV treatment of living cells, which the authors interpreted as further support for a direct interaction of RV with the protein in the extract (19; Table I, Figure 2). Testing the influence of RV on the APE/Ref-1 target molecule AP-1 in electrophoretic mobility shift assays revealed that RV antagonizes DNA binding by AP-1 in a manner strictly depending on the presence of APE/Ref-1. However, whether RV, indeed, physically interacts with APE/Ref-1 and counteracts its enzymatic functions will have to await molecular interaction studies and cleavage assays with defined protein composition to exclude the involvement of other DNA-metabolizing enzymes (see DNA metabolism). So far, others failed to show similar effects regarding both inhibition of endonuclease and DNA-binding activities of APE/Ref-1 by RV (116). In the light of the important role of APE/Ref-1 in the maintenance of genomic stability, an inhibitory effect on APE/Ref-1 endonuclease by RV would definitely provoke a conflict regarding its propagated role in chemoprevention.

The ATM/ATR-signaling network.
Lesion detection is the first essential step in the cellular response to a DSB. Within seconds after DSB formation, the complex consisting of the nuclease Mre11, the ATPase Rad50 and the regulatory component Nbs1 (MRN complex) associates with and recruits the PI3K-related kinase ATM to the break. Apart from an intact MRN complex, activation of ATM requires phosphorylation at three sites within the ATM molecule, particularly intermolecular trans-phosphorylation at serine 1981 (117–120; reviewed in ref. 121). The histone acetyltransferases hMOF and TIP60 were suggested to also be involved in ATM activation, thereby linking changes in the chromatin structure to sensing and repairing DSBs (reviewed in ref. 122). Once activated, ATM phosphorylates several key DSB repair and checkpoint control factors like H2AX, p53, Nbs1, BRCA1, SMC1, Chk2 and Chk1. H2AX phosphorylation occurs over megabase-pair regions and is required for the retention of mediator proteins like p53BP1, MDC1, BRCA1 and the MRN complex at the break. The MRN complex, thus, represents both an upstream activator and a downstream target of ATM (122–124). The PI3K-related kinase ATR phosphorylates almost the same substrates as ATM, however, in response to replication fork blockage. ATR activation requires complex formation with the ATR-interacting protein and association with double-stranded DNA–single-stranded DNA transition sites containing replication protein A-coated single-stranded DNA, Claspin, TopBP1 and the 9-1-1 (Rad9-Rad1-Hus1) complex that stabilizes the TopBP1–ATR interaction (125,126).

Very recently, RV was reported to induce S-phase arrest through activation of an ATM/ATR–Chk1/Chk2–Cdc25C pathway (127). Compatible with these data RV has been demonstrated to activate ATM and ATR pathways in human lymphoblastoid cell lines leading to phosphorylation of several downstream targets like p53, Nbs1, Chk1 and Chk2 (S.A. Gatz, M. Keimling, C. Baumann, T. Dörk, S. Fulda, and L. Wiesmüller, Carcinogenesis, in press). In this study use of a fluorescence-based assay further revealed that RV controls the fidelity of HR and NHEJ through activation of ATM/ATR–p53 and ATM/ATR–Nbs1 pathways, respectively (Figure 2). Importantly, the fast readout of the repair assay combined with comparatively low RV dosage (5–30 µM) enabled to separate this effect from induction of cell-cycle arrest or apoptosis. The finding that p53 is necessary for RV-triggered HR control reconciles these results and earlier reports on the induction of SCEs and MN formation after RV treatment because the latter were based on experiments with cells carrying dysfunctional p53 (98–102).

Reminiscent of the model describing RV as an activator of the PI3K-related kinases ATM and ATR, several other putative chemopreventive substances, namely apigenin, kaempferol, luteolin, QU and GE have been demonstrated to lead to phosphorylation of ATM and ATM/ATR downstream targets (128–130). Reasons for the ATM/ATR-activating role of RV and the other substances have been sought in potential DNA-damaging effects (127–130). However, in contrast to GE, which is an established topo II poison (Table I), thereby indirectly introducing DSBs, the causal relationship with RV is less clear. So far, only one group reported on modest topo I poisoning by RV at a concentration of 100 µM (12). The results from other groups rather indicated catalytic inhibition of topo II (Table I). The latter findings are compatible with published comet assay data, which demonstrate that RV concentrations of up to 100 µM do not lead to an increased tail moment and even show a protective effect (see Direct DNA interactions and breakage). Since RV was reported to inhibit pol {alpha} and {delta} in vitro (Table I) and to induce oxidative stress under certain conditions in cancer cells (60), activation of ATM/ATR pathways could be mediated by replication fork stalling. Independently of the mechanism, activation of ATR and/or ATM is further corroborated by phosphorylation of H2AX in the nucleus, although distinct focal accumulations as typically seen at sites of DSBs are not observed (60,127). For the elucidation of the activation mechanism, it is of interest that RV is known to bind DNA and to modulate DNA intercalation of substances such as ethidium bromide and acridine orange (see Direct DNA interactions and breakage). Bakkenist et al. (117) successfully applied chromatin-active compounds like the intercalating drug chloroquine, when establishing the central role of ATM in the DNA damage response. By analogy, RV could as well induce structural changes that lead to ATM and ATR activation in the absence of immediate DNA damage.

The sirtuin connection.
The sirtuins form a large and ancient family of proteins with two enzymatic functions: Mono-ADP-ribosyl transferase and NAD+-dependent protein deacetylase activity. So far, four sirtuins (Sir1–4) have been described in yeast and seven sirtuins in mammals (SIRT1–7). The prototype is silent information regulator 2(Sir2) from Saccharomyces cerevisiae. The histone deacetylase Sir2 functions as a chromatin silencer to regulate transcription, recombination, genomic stability and aging. It is of particular interest for this review that Sir2 appears to be involved in DSB repair by NHEJ and suppresses rDNA recombination, which generates extrachromosomal rDNA circles (reviewed in ref. 131).

RV treatment (10 and 100 µM) in yeast was reported to lead to a Sir2-dependent inhibition of ribosomal recombination of up to 60%, as determined by reduction of rDNA::ADE2 marker gene loss (28). In apparent contradiction, Kaeberlein et al. (132) could not confirm Sir2 activation by RV with three different yeast strains. Nevertheless, since murine Sir2{alpha} and human SIRT1 are the closest Sir2 homologues, it was attractive to search for links between Sir2{alpha}/SIRT1 and DNA repair. However, the data accumulated so far are scarce and hardly convincing: Unlike yeast sir2 mutants, murine embryonic stem cells from Sir2{alpha} knockout mice were not sensitive to ionizing radiation (133). Furthermore, SIRT1–/– DT40 chicken cells showed plasmid-based NHEJ and HR activities without statistically significant differences from wild-type cells (134). However, two other groups independently described sirtuin-dependent effects on NHEJ, though diametrically opposed (135,136). By use of the neutral comet assay, Wojewodzka et al. (135) measured slightly faster repair of ionizing radiation-induced DSBs in chinese hamster ovary cells after treatment with sirtuin inhibitor GPI 19015. However, these differences were noticeable only in cells dysfunctional for the classical, DNA-dependent protein kinase-dependent NHEJ pathway. From their results obtained after overexpression or knockdown of SIRT1, Jeong et al. (136) concluded that SIRT1 stimulates repair of DNA strand breaks.

RV was identified as the most potent SIRT1 activator in a small molecule screen based on deacetylation assays, which applied synthetic SIRT1 peptide substrates, labeled with a fluorophore (Fluor de Lys kit) (28). This effect was mainly attributed to RV's ability to lower the Km of SIRT1 for both the acetylated substrate and NAD+ (28; Table I). However, these biochemical data were subsequently challenged by two groups independently showing that RV had no effect on SIRT1-dependent binding and deacetylation of target peptides lacking the fluorophore. This fluorophore dependency can be explained by the fact that the fluorophore decreased the binding affinity of the peptide, and that, in the presence of RV, fluorophore-containing substrates bound more tightly to SIRT1 (132,137). These data cast doubts on the proposed allosteric mechanism. On the other hand, RV at low micromolar concentrations reduced accumulation of acetylated p53 after ultraviolet irradiation in HEK293 and U2OS cells with wild-type SIRT1, but not in their counterparts with dominant-negative SIRT1 (28). Finally, studies with mice showed that RV-containing diet promotes longevity and improves glucose homeostasis most probably by stimulating SIRT1-mediated deacetylation of the transcriptional coactivator peroxisome proliferator-activated receptor-{gamma} coactivator 1{alpha} (56,57). Therefore, accumulating in vitro and in vivo data on an activating interaction between RV and mammalian SIRT1 exist; however, the molecular characterization of the underlying mechanisms awaits further investigation.

The growing list of non-histone SIRT1 targets includes several proteins that might directly or indirectly be involved in DNA repair and regulation of genomic stability: p53, Nbs1, KU70, Werner helicase, NF-{kappa}B and FOXO proteins (FOXO1, FOXO3a and FOXO4) (131,138,139). As a key surveillance factor of the DNA damage response, p53 is a particularly interesting molecule regarding SIRT1-dependent modulation of not only transcriptional and proapoptotic responses but also of DSB repair (111–113). So far, contradictory data have been presented on the influence of p53 acetylation on its transcriptional activities (140–145). RV was demonstrated to enhance SIRT1-mediated deacetylation of p53, which correlated with enhanced cell survival after ionizing radiation (28). Given that Nbs1 was identified as another SIRT1 substrate, it is interesting that deacetylation of Nbs1 is a prerequisite for Nbs1 phosphorylation by ATM (139). From these data, it is conceivable that SIRT1 is involved in RV-mediated DSB repair modulation by p53 and Nbs1. RV was further established to enhance SIRT1-mediated deacetylation of the DNA-dependent protein kinase component KU70 (146). KU70 deacetylation appears to stabilize the cytoplasmic KU70–Bax complex, thereby preventing apoptosis through Bax sequestration (146). In this context it is of interest that Jeong et al. (136) observed that overexpression of SIRT1 led to increased repair of DNA strand breaks. This repair increase correlated with KU70 deacetylation, but formal proof for a causal relationship is still missing.

Through the discovery of an upstream link to Chk2, SIRT1 has itself been brought in line with DSB signaling by the kinase cascade involving ATM and Chk2 (147). Clearly, further research will be necessary to establish the precise role of SIRT1 in regulating p53, Nbs1 and KU70 functions and to correctly position RV within these interactions (Figure 2). Other targets like NF-{kappa}B and FOXO4, for which RV-triggered and SIRT1-mediated modulation was also documented, appear to influence genomic stability more indirectly such as via stress gene or apoptosis regulation (148,149). Regarding the sirtuin SIRT6, convincing evidence for a role in DNA repair has been provided (150). Thus, from studies with SIRT6-deficient mice, SIRT6 is a nuclear, chromatin-associated protein that suppresses genomic instability and has been implicated in BER. Investigating a possible role of RV in SIRT6-dependent repair regulation, therefore, remains a challenge for the future.


   Conclusions
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 Effects of RV on...
 Modulation of DNA damage...
 Conclusions
Funding
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The most straightforward mechanism possibly underlying RV's effects is direct enzyme interaction, as was postulated for QR2 and APE/Ref-1 (18,19; Table I and Figure 2). One alternative way of RV action, which could well explain a number of cell-based findings, is its interaction with DNA. DNA intercalation is an attractive model for multiple small molecules (151). RV, though itself not intercalating with DNA, has been shown to revert DNA intercalation through stabilization of double-stranded DNA structure, thereby exerting a protective effect (10,72). On the other hand, RV may give access to reactive molecules such as the transition metal Cu(II), as was shown previously for some flavonoids (13). RV may also counteract DNA-metabolizing processes through blockage of enzymatic interaction sites on the DNA, resulting in the catalytic inhibition of topo II and polymerases (Table I). Consequently, mere DNA binding by RV could be the basis for RV-induced replication stress and downstream ATM and ATR signaling (60,127). Adding another layer of complexity to RV's influence on the genome structure, RV may also contribute to chromatin remodeling through upregulation of p300 and activation of SIRT1 (8; Figure 2). Intriguingly, changes in the chromatin structure are known to promote ATM activation (120). From this it is conceivable that RV-induced changes in the primary DNA and chromatin structure synergistically trigger ATM signaling, which is reminiscent of the functions attributed to the MRN complex (118).

It is tempting to speculate that activation of ATM/ATR by RV is not only essential for its effects on Chk1, Chk2, p53 and Nbs1 in DSB repair and checkpoint control (Figure 2), but also the clue to many other properties of this substance. In support of this idea, RV has been linked to multiple ATM targets such as NF-{kappa}B, STAT3, AP-1, Bid/Bax, p38 mitogen-activated protein kinase and AKT (5,7,8). This concept of RV action involving DNA interaction and ATM/ATR activation is not restricted to RV. Rather, other polyphenols, QU and GE in particular, seem to share the same principle, although they are distinct with respect to the pattern of DNA interaction and the biological consequences such as regarding the introduction of DNA damage (Table I, Figure 3). These differences may have an impact on cancer preventive and therapeutic strategies. Given that RV triggers defensive mechanisms without causing serious DNA damage, RV is expected to prepare the cell against devastating stimuli and argues for RV as a chemopreventive rather than a therapeutic substance. This is in line with the fact that RV showed convincing effectiveness mostly in in vivo models for cancer prevention, such as in adult rats where it inhibits 7,12-dimethylbenz[a]anthracene-induced formation of ductal lesions in the mammary gland (36). For comparison, the topo II poison GE, which causes DSB formation, ought to be applied with caution in chemoprevention. This is exemplified best by a recent study of Schooten et al. (103), which indicates that biologically relevant concentrations of GE can induce leukemogenic abnormalities in the MLL breakpoint cluster region in primary hematopoietic progenitor cells. On the other hand, GE might be a promising compound for therapeutic approaches, as treatment with GE inhibited development of tumors derived from adult T-cell leukemia cells and inhibited tissue invasion of these cells in a recent study in mice (152). However, effectiveness of polyphenols could be limited because mutations in key genes of the ATM/ATR cascade are particularly frequent during the process of carcinogenesis. The data summed up here add another layer to the already highly complex world of RV and related compounds and may serve as the basis for the design of future chemoprevention and therapy studies.


Figure 3
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  		Fig. 3. Model for the functional interactions of RV with ATM and ATR, which may also be true for QU and GE.

 

   Funding
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 Abstract
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 Effects of RV on...
 Modulation of DNA damage...
 Conclusions
 Funding
References
 
Deutsche Forschungsgemeinschaft, Clinical Research Group: ‘Regulation of apoptosis and its dysfunction in diseases’, project: ‘Role of apoptosis in the induction of carcinogenic chromosome instabilities’ to L.W.; The University of Ulm (‘Baustein 3.4’, I.Z06) to S.A.G.


   Acknowledgments
 
Conflict of Interest Statement: None declared.


   References
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 Abstract
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 Modulation of DNA damage...
 Conclusions
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 References
 

  1. Lindahl T, et al. Rate of depurination of native deoxyribonucleic acid. Biochemistry (1972) 11:3610–3618.[CrossRef][Web of Science][Medline]
  2. Vilenchik MM, et al. Endogenous DNA double-strand breaks: production, fidelity of repair, and induction of cancer. Proc. Natl Acad. Sci. USA (2003) 100:12871–12876.[Abstract/Free Full Text]
  3. Manson NM. Cancer prevention—the potential for diet to modulate molecular signalling. Trends Mol. Med. (2003) 9:11–18.[CrossRef][Web of Science][Medline]
  4. Surh YJ. Cancer chemoprevention with dietary phytochemicals. Nat. Rev. Cancer (2003) 3:768–780.[CrossRef][Web of Science][Medline]
  5. Aggarwal BB, et al. Molecular targets of dietary agents for prevention and therapy of cancer. Biochem. Pharmacol. (2006) 71:1397–1421.[CrossRef][Web of Science][Medline]
  6. Jang M, et al. Cancer chemopreventive activity of resveratrol, a natural product derived from grapes. Science (1997) 275:218–220.[Abstract/Free Full Text]
  7. Pervaiz S. Chemotherapeutic potential of the chemopreventive phytoalexin resveratrol. Drug Resist. Updat. (2004) 7:333–344.[CrossRef][Medline]
  8. Signorelli P, et al. Resveratrol as an anticancer nutrient: molecular basis, open questions and promises. J. Nutr. Biochem. (2005) 16:449–466.[CrossRef][Web of Science][Medline]
  9. Fukuhara K, et al. Resveratrol as a new type of DNA-cleaving agent. Bioorg. Med. Chem. Lett. (1998) 8:3187–3192.[CrossRef][Medline]
  10. Usha S, et al. Interaction of resveratrol and genistein with nucleic acids. J. Biochem. Mol. Biol. (2005) 38:198–205.[Medline]
  11. Alvi NK, et al. Interaction of quercetin with DNA. Biosci. Rep. (1986) 6:861–868.[CrossRef][Web of Science][Medline]
  12. Webb MR, et al. Comparative analysis of topoisomerase IB inhibition and DNA intercalation by flavonoids and similar compounds: structural determinates of activity. Biochem. J. (2004) 384:527–541.[CrossRef][Web of Science][Medline]
  13. Snyder RD, et al. Evaluation of the clastogenic, DNA intercalative, and topoisomerase II-interactive properties of bioflavonoids in Chinese hamster V79 cells. Environ. Mol. Mutagen. (2002) 40:266–276.[CrossRef][Web of Science][Medline]
  14. Zini R, et al. Effects of resveratrol on the rat brain respiratory chain. Drugs Exp. Clin. Res. (1999) 25:87–97.[Web of Science][Medline]
  15. Zheng J, et al. Inhibition of mitochondrial proton F0F1-ATPase/ ATP synthase by polyphenolic phytochemicals. Br. J. Pharmacol. (2000) 130:1115–1123.[CrossRef][Web of Science][Medline]
  16. Kipp JL, et al. Effect of estradiol, diethylstilbestrol, and resveratrol on F0F1-ATPase activity from mitochondrial preparations of rat heart, liver, and brain. Endocrine (2001) 15:165–175.[CrossRef][Web of Science][Medline]
  17. Gledhill JR, et al. Mechanism of inhibition of bovine F1-ATPase by resveratrol and related polyphenols. Proc. Natl Acad. Sci. USA (2007) 104:13632–13637.[Abstract/Free Full Text]
  18. Buryanovskyy L, et al. Crystal structure of quinone reductase 2 in complex with resveratrol. Biochemistry (2004) 43:11417–11426.[CrossRef][Web of Science][Medline]
  19. Yang S, et al. Alterations in the expression of the apurinic/apyrimidinic endonuclease-1/redox factor-1 (APE/Ref-1) in human melanoma and identification of the therapeutic potential of resveratrol as an APE/Ref-1 inhibitor. Mol. Cancer Ther. (2005) 4:1923–1935.[Abstract/Free Full Text]
  20. Stivala LA, et al. Specific structural determinants are responsible for the antioxidant activity and cell cycle effects of resveratrol. J. Biol. Chem. (2001) 276:22586–22594.[Abstract/Free Full Text]
  21. Locatelli GA, et al. Inhibition of mammalian DNA polymerases by resveratrol: mechanism and structural determinants. Biochem. J. (2005) 389:259–268.[CrossRef][Web of Science][Medline]
  22. Ono K, et al. Mechanisms of inhibition of various cellular DNA and RNA polymerases by several flavonoids. J. Biochem. (Tokyo) (1990) 108:609–613.[Abstract/Free Full Text]
  23. Fontecave M, et al. Resveratrol, a remarkable inhibitor of ribonucleotide reductase. FEBS Lett. (1998) 421:277–279.[CrossRef][Web of Science][Medline]
  24. Cho KH, et al. Selection of cancer chemopreventive agents based on inhibition of topoisomerase II activity. Eur. J. Cancer (2000) 36:2146–2156.[CrossRef][Medline]
  25. Lopez-Lazaro M, et al. Cells lacking DNA topoisomerase II beta are resistant to genistein. J. Nat. Prod. (2007) 70:763–767.[CrossRef][Medline]
  26. Filipski J. Competitive inhibition of nicking-closing enzymes may explain some biological effects of DNA intercalators. FEBS Lett. (1983) 159:6–12.[CrossRef][Medline]
  27. Jo JY, et al. Effects of grape cell culture extracts on human topoisomerase II catalytic activity and characterization of active fractions. J. Agric. Food Chem. (2005) 53:2489–2498.[CrossRef][Medline]
  28. Howitz KT, et al. Small molecule activators of sirtuins extend Saccharomyces cerevisiae lifespan. Nature (2003) 425:191–196.[CrossRef][Medline]
  29. Lanzilli G, et al. Resveratrol down-regulates the growth and telomerase activity of breast cancer cells in vitro. Int. J. Oncol. (2006) 28:641–648.[Web of Science][Medline]
  30. Jagadeesh S, et al. Genistein represses telomerase activity via both transcriptional and posttranslational mechanisms in human prostate cancer cells. Cancer Res. (2006) 66:2107–2115.[Abstract/Free Full Text]
  31. Ouchi H, et al. Genistein induces cell growth inhibition in prostate cancer through the suppression of telomerase activity. Int. J. Urol. (2005) 12:73–80.[CrossRef][Web of Science][Medline]
  32. Ku WC, et al. Inhibition of telomerase activity by PKC inhibitors in human nasopharyngeal cancer cells in culture. Biochem. Biophys. Res. Commun. (1997) 241:730–736.[CrossRef][Web of Science][Medline]
  33. Le Corre L, et al. Resveratrol and breast cancer chemoprevention: molecular mechanisms. Mol. Nutr. Food Res. (2005) 49:462–471.[CrossRef][Web of Science][Medline]
  34. Baur JA, et al. Therapeutic potential of resveratrol: the in vivo evidence. Nat. Rev. Drug Discov. (2006) 5:493–506.[CrossRef][Web of Science][Medline]
  35. Howells LM, et al. Predicting the physiological relevance of in vitro cancer preventive activities of phytochemicals. Acta Pharmacol. Sin. (2007) 28:1274–1304.[CrossRef][Medline]
  36. Athar M, et al. Resveratrol: a review of preclinical studies for human cancer prevention. Toxicol. Appl. Pharmacol. (2007) 224:274–283.[CrossRef][Medline]
  37. Mahyar-Roemer M, et al. P21 waf1/cip1 can protect human colon carcinoma cells against p53-dependent and p53-independent apoptosis induced by natural chemopreventive and therapeutic agents. Oncogene (2001) 20:3387–3398.[CrossRef][Medline]
  38. Hayashibara T, et al. Resveratrol induces downregulation in survivin expression and apoptosis in HTLV-1-infected cell lines: a prospective agent for adult T cell leukemia chemotherapy. Nutr. Cancer (2002) 44:193–201.[CrossRef][Medline]
  39. Fulda S, et al. Sensitization for anticancer drug-induced apoptosis by the chemopreventive agent resveratrol. Oncogene (2004) 23:6702–6711.[CrossRef][Medline]
  40. Kotha A, et al. Resveratrol inhibits Src and Stat3 signaling and induces the apoptosis of malignant cells containing activated Stat3 protein. Mol. Cancer Ther. (2006) 5:621–629.[Abstract/Free Full Text]
  41. Faber AC, et al. Resveratrol induces apoptosis in transformed follicular lymphoma OCI-LY8 cells: evidence for a novel mechanism involving inhibition of BCL6 signaling. Int. J. Oncol. (2006) 29:1561–1566.[Medline]
  42. Bhardwaj A, et al. Resveratrol inhibits proliferation, induces apoptosis, and overcomes chemoresistance through down-regulation of STAT3 and nuclear factor-kappaB-regulated antiapoptotic and cell survival gene products in human multiple myeloma cells. Blood (2007) 109:2293–2302.[Abstract/Free Full Text]
  43. Aggarwal BB, et al. Targeting signal-transducer-and-activator-of-transcription-3 for prevention and therapy of cancer: modern target but ancient solution. Ann. N. Y. Acad. Sci. (2006) 1091:151–169.[CrossRef][Web of Science][Medline]
  44. Fulda S, et al. Sensitization for tumor necrosis factor-related apoptosis-inducing ligand-induced apoptosis by the chemopreventive agent resveratrol. Cancer Res. (2004) 64:337–346.[Abstract/Free Full Text]
  45. Jazirehi AR, et al. Resveratrol modifies the expression of apoptotic regulatory proteins and sensitises non-Hodgkin's lymphoma and multiple myeloma cell lines to paclitaxel-induced apoptosis. Mol. Cancer Ther. (2004) 3:71–84.[Abstract/Free Full Text]
  46. Dianov GL, et al. Co-ordination of DNA single strand break repair. DNA Repair (2007) 6:454–460.[Medline]
  47. Finkel T, et al. Oxidants, oxidative stress and the biology of ageing. Nature (2000) 408:239–247.[CrossRef][Medline]
  48. Guetens G, et al. Oxidative DNA damage: biological significance and methods of analysis. Crit. Rev. Clin. Lab. Sci. (2002) 39:331–457.[Web of Science][Medline]
  49. Bonavida B, et al. Therapeutic potential of nitric oxide in cancer. Drug Resist. Updat. (2006) 9:157–173.[CrossRef][Medline]
  50. Leonard SS, et al. Resveratrol scavenges reactive oxygen species and effects radical-induced cellular responses. Biochem. Biophys. Res. Commun. (2003) 309:1017–1026.[CrossRef][Web of Science][Medline]
  51. Poolman TM, et al. Inhibition of the respiratory burst by resveratrol in human monocytes: correlation with inhibition of PI3K signalling. Free Radic. Biol. Med. (2005) 39:118–132.[CrossRef][Medline]
  52. Ahmad KA, et al. Resveratrol inhibits drug-induced apoptosis in human leukemia cells by creating an intracellular milieu nonpermissive for death execution. Cancer Res. (2004) 64:1452–1459.[Abstract/Free Full Text]
  53. Kamata H, et al. Redox regulation of cellular signalling. Cell. Signal. (1999) 11:1–14.[CrossRef][Web of Science][Medline]
  54. Watabe M, et al. ATP depletion does not account for apoptosis induced by inhibition of mitochondrial electron transport chain in human dopaminergic cells. Neuropharmacology (2007) 52:536–541.[CrossRef][Medline]
  55. Zini R, et al. Resveratrol-induced limitation of dysfunction of mitochondria isolated from rat brain in an anoxia-reoxygenation model. Life Sci. (2002) 71:3091–3108.[CrossRef][Web of Science][Medline]
  56. Baur JA, et al. Resveratrol improves health and survival of mice on a high-calorie diet. Nature (2006) 444:337–342.[CrossRef][Medline]
  57. Lagouge M, et al. Resveratrol improves mitochondrial function and protects against metabolic disease by activating SIRT1 and PGC-1alpha. Cell (2006) 127:1109–1122.[CrossRef][Web of Science][Medline]
  58. Dasgupta B, et al. Resveratrol stimulates AMP kinase activity in neurons. Proc. Natl Acad. Sci. USA (2007) 104:7217–7222.[Abstract/Free Full Text]
  59. Zang M, et al. Polyphenols stimulate AMP-activated protein kinase, lower lipids, and inhibit accelerated atherosclerosis in diabetic LDL receptor-deficient mice. Diabetes (2006) 55:2180–2191.[Abstract/Free Full Text]
  60. Heiss EH, et al. Chronic treatment with resveratrol induces redox stress- and ATM-dependent senescence in p53-positive cancer cells. J Biol. Chem. (2007) 282:26759–26766.[Abstract/Free Full Text]
  61. Ciolino HP, et al. Inhibition of aryl hydrocarbon-induced cytochrome P-450 1A1 enzyme activity and CYP1A1 expression by resveratrol. Mol. Pharmacol. (1999) 56:760–777.[Abstract/Free Full Text]
  62. Chang TK, et al. Differential inhibition and inactivation of human CYP1 enzymes by trans-resveratrol: evidence for mechanism-based inactivation of CYP1A2. J. Pharmacol. Exp. Ther. (2001) 299:874–882.[Abstract/Free Full Text]
  63. Ciolino HP, et al. Resveratrol inhibits transcription of CYP1A1 in vitro by preventing activation of the aryl hydrocarbon receptor. Cancer Res. (1998) 58:5707–5712.[Abstract/Free Full Text]
  64. Vasiliou V, et al. Update of the NAD(P)H:quinone oxidoreductase (NQO) gene family. Hum. Genomics (2006) 2:329–335.[Medline]
  65. Vella F, et al. NRH:quinone reductase 2: an enzyme of surprises and mysteries. Biochem. Pharmacol. (2005) 71:1–12.[CrossRef][Medline]
  66. Cervello M, et al. Cyclooxygenases in hepatocellular carcinoma. World J. Gastroenterol. (2006) 12:5223–15121.[Medline]
  67. Subbaramaiah K, et al. Resveratrol inhibits cyclooxygenase-2 transcription and activity in phorbol ester-treated human mammary epithelial cells. J. Biol. Chem. (1998) 273:21875–21882.[Abstract/Free Full Text]
  68. Stewart JR, et al. Resveratrol preferentially inhibits protein kinase C-catalyzed phosphorylation of a cofactor-independent, arginine-rich protein substrate by a novel mechanism. Biochemistry (1999) 38:13244–13251.[CrossRef][Web of Science][Medline]
  69. Slater SJ, et al. Inhibition of protein kinase C by resveratrol. Biochim. Biophys. Acta (2003) 1637:59–69.[Medline]
  70. Pinto MC, et al. Resveratrol is a potent inhibitor of the dioxygenase activity of lipoxygenase. J. Agric. Food Chem. (1999) 47:4842–4846.[CrossRef][Medline]
  71. MacCarrone M, et al. Resveratrol prevents apoptosis in K562 cells by inhibiting lipoxygenase and cyclooxygenase activity. Eur. J. Biochem. (1999) 265:27–34.[Web of Science][Medline]
  72. Usha S, et al. Possible inhibition of group I intron RNA by resveratrol and genistein. J. Biomol. Struct. Dyn. (2006) 24:25–32.[Medline]
  73. Usha S, et al. Modulation of DNA intercalation by resveratrol and genistein. Mol. Cell. Biochem. (2006) 284:57–64.[CrossRef][Medline]
  74. Burkitt MJ, et al. Effects of trans-resveratrol on copper-dependent hydroxyl-radical formation and DNA-damage: evidence for hydroxyl-radical scavenging and a novel, glutathione-sparing mechanism of action. Arch. Biochem. Biophys. (2000) 381:253–263.[CrossRef][Medline]
  75. Ahmad A, et al. DNA breakage by resveratrol and Cu(II): reaction mechanism and bacteriophage inactivation. Cancer Lett. (2000) 154:29–37.[CrossRef][Web of Science][Medline]
  76. Subramanian M, et al. A mechanistic study on the nuclease activities of some hydroxystilbenes. Bioorg. Med. Chem. (2004) 12:1231–1237.[CrossRef][Medline]
  77. Win W, et al. Different effects of genistein and resveratrol on oxidative DNA damage in vitro. Mutat. Res. (2002) 513:113–120.[Web of Science][Medline]
  78. Ahmad A, et al. Prooxidant activity of resveratrol in the presence of copper ions: mutagenicity in plasmid DNA. Toxicol. Lett. (2005) 159:1–12.[CrossRef][Medline]
  79. Fukuhara K, et al. Structural basis for DNA-cleaving activity of resveratrol in the presence of Cu(II). Bioorg. Med. Chem. (2006) 14:1437–1443.[CrossRef][Medline]
  80. Kagawa TF, et al. Covalent modification of guanine bases in double-stranded DNA. The 1.2-A-Z-DNA structure of d(CGCGCG) in the presence of CuCl2. J. Biol. Chem. (1991) 266:20175–20184.[Abstract/Free Full Text]
  81. Azmi AS, et al. Plant polyphenols mobilize endogenous copper in human peripheral lymphocytes leading to oxidative DNA-breakage: a putatative mechanism for anticancer properties. FEBS Lett. (2006) 580:533–538.[CrossRef][Web of Science][Medline]
  82. Azmi AS, et al. Resveratrol-Cu(II) induced DNA breakage in human peripheral lymphocytes: implications for anticancer properties. FEBS Lett. (2005) 579:3131–3135.[CrossRef][Web of Science][Medline]
  83. Hubscher U, et al. Eukaryotic DNA polymerases. Annu. Rev. Biochem. (2002) 71:133–163.[CrossRef][Web of Science][Medline]
  84. Ramadan K, et al. The DNA-polymerase-X family: controllers of DNA quality? Nat. Rev. Mol. Cell Biol. (2004) 5:1038–1043.[CrossRef][Web of Science][Medline]
  85. Lydeard JR, et al. Break-induced replication and telomerase-independent telomere maintenance require Pol32. Nature (2007) 448:820–823.[CrossRef][Web of Science][Medline]
  86. Sun NJ, et al. DNA polymerase and topoisomerase II inhibitors from Psoralea corylifolia. J. Nat. Prod. (1998) 61:362–366.[CrossRef][Medline]
  87. Kolberg M, et al. Structure, function, and mechanism of ribonucleotide reductases. Biochim. Biophys. Acta (2004) 1699:1–34.[Medline]
  88. Elford HL. Effect of hyroxyurea on ribonucleotide reductase. Biochem. Biophys. Res. Commun. (1968) 33:129–135.[CrossRef][Medline]
  89. Woo LL, et al. The broken genome: genetic and pharmacologic approaches to breaking DNA. Ann. Med. (2007) 39:208–218.[CrossRef][Web of Science][Medline]
  90. Snyder RD, et al. Reduction of genistein clastogenicity in Chinese hamster V79 cells by daidzein and other flavonoids. Food Chem. Toxicol. (2003) 41:1291–1298.[CrossRef][Web of Science][Medline]
  91. Zimmermann S, et al. Telomeres and telomerase as targets for cancer therapy. Cell. Mol. Life Sci. (2007) 64:906–921.[CrossRef][Web of Science][Medline]
  92. Bailey SM, et al. Telomeres, chromosome instability and cancer. Nucleic Acids Res. (2006) 34:2408–2417.[Abstract/Free Full Text]
  93. Fuggetta MP, et al. Effect of resveratrol on proliferation and telomerase activity of human colon cancer cells in vitro. J. Exp. Clin. Cancer Res. (2006) 25:189–193.[Medline]
  94. Sheng WY, et al. A major role of PKC theta and NFkappaB in the regulation of hTERT in human T lymphocytes. FEBS Lett. (2006) 580:6819–6824.[CrossRef][Medline]
  95. Zhang P, et al. c-Myc downregulation: a critical molecular event in resveratrol-induced cell cycle arrest and apoptosis of human medulloblastoma cells. J. Neurooncol. (2006) 80:123–131.[CrossRef][Medline]
  96. Sancar A, et al. Molecular mechanisms of mammalian DNA repair and the DNA damage checkpoints. Annu. Rev. Biochem. (2004) 73:39–85.[CrossRef][Web of Science][Medline]
  97. Speit G, et al. Chromosomal mutagen sensitivity associated with mutations in BRCA genes. Cytogenet. Genome Res. (2004) 104:325–332.[CrossRef][Medline]
  98. Schmitt E, et al. Hormonal and genotoxic activity of resveratrol. Toxicol. Lett. (2002) 136:133–142.[CrossRef][Medline]
  99. Stopper H, et al. Genotoxicity of phytoestrogens. Mutat. Res. (2005) 574:139–155.[Medline]
  100. Matsuoka A, et al. Resveratrol, a naturally occurring polyphenol, induces sister chromatid exchanges in a Chinese hamster lung (CHL) cell line. Mutat. Res. (2001) 494:107–113.[Medline]
  101. Matsuoka A, et al. The 4'-hydroxy group is responsible for the in vitro cytogenetic activity of resveratrol. Mutat. Res. (2002) 521:29–35.[Medline]
  102. Matsuoka A, et al. Correlation of sister chromatid exchange formation through homologous recombination with ribonucleotide reductase inhibition. Mutat. Res. (2004) 547:101–107.[Web of Science][Medline]
  103. van Waalwijk van Doorn-Khosrovani SB, et al. Dietary flavonoids induce MLL translocations in primary human CD34+ cells. Carcinogenesis (2007) 28:1703–1709.[Abstract/Free Full Text]
  104. Chakraborty S, et al. Prevention and repair of DNA damage by selected phytochemicals as measured by single cell gel electrophoresis. J. Environ. Pathol. Toxicol. Oncol. (2004) 23:215–226.[CrossRef][Medline]
  105. Fustier P, et al. Resveratrol increases BRCA1 and BRCA2 mRNA expression in breast tumour cell lines. Br. J. Cancer (2003) 89:168–172.[CrossRef][Medline]
  106. Le Corre L, et al. Effects of resveratrol on the expression of a panel of genes interacting with the BRCA1 oncosuppressor in human breast cell lines. Clin. Chim. Acta (2004) 344:115–121.[CrossRef][Web of Science][Medline]
  107. Narayanan BA, et al. Differential expression of genes induced by resveratrol in LNCaP cells: P53-mediated molecular targets. Int. J. Cancer (2003) 104:204–212.[CrossRef][Medline]
  108. Waite KA, et al. Phytoestrogen exposure elevates PTEN levels. Hum. Mol. Genet. (2005) 14:1457–1463.[Abstract/Free Full Text]
  109. Shen WH, et al. Essential role for nuclear PTEN in maintaining chromosomal integrity. Cell (2007) 128:157–170.[CrossRef][Web of Science][Medline]
  110. Valerie K, et al. Regulation and mechanisms of mammalian double-strand break repair. Oncogene (2003) 22:5792–5812.[CrossRef][Web of Science][Medline]
  111. Bertrand P, et al. P53's double life: transactivation independent repression of homologous recombination. Trends Genet. (2004) 20:235–324.[CrossRef][Web of Science][Medline]
  112. Sengupta S, et al. p53: traffic cop at the crossroads of DNA repair and recombination. Nat. Rev. Mol. Cell Biol. (2005) 6:44–55.[CrossRef][Web of Science][Medline]
  113. Gatz SA, et al. p53 in recombination and repair. Cell Death Differ. (2006) 13:1003–1016.[CrossRef][Web of Science][Medline]
  114. Niture SK, et al. Increased expression of the MGMT repair protein mediated by cysteine prodrugs and chemopreventative natural products in human lymphocytes and tumor cell lines. Carcinogenesis (2007) 28:378–389.[Abstract/Free Full Text]
  115. Kaina B, et al. MGMT: key node in the battle against genotoxicity, carcinogenicity and apoptosis induced by alkylating agents. DNA Repair (Amst.) (2007) 6:1079–1099.[CrossRef][Medline]
  116. Fishel ML, et al. The DNA base excision repair protein Ape1/Ref-1 as a therapeutic and chemopreventive target. Mol. Aspects Med. (2007) 28:375–395.[CrossRef][Medline]
  117. Bakkenist CJ, et al. DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature (2003) 421:499–506.[CrossRef][Medline]
  118. Lee JH, et al. ATM activation by DNA double-strand breaks through the Mre11-Rad50-Nbs1 complex. Science (2005) 308:551–554.[Abstract/Free Full Text]
  119. Pellegrini M, et al. Autophosphorylation at serine 1987 is dispensable for murine Atm activation in vivo. Nature (2006) 443:222–225.[CrossRef][Medline]
  120. Berkovich E, et al. Roles of ATM and NBS1 in chromatin structure modulation and DNA double-strand break repair. Nat. Cell Biol. (2007) 9:683–690.[CrossRef][Web of Science][Medline]
  121. Lavin MF, et al. ATM activation and DNA damage response. Cell Cycle (2007) 6:931–942.[Medline]
  122. Scott SP, et al. The cellular control of DNA double-strand breaks. J. Cell. Biochem. (2006) 99:1463–1475.[CrossRef][Web of Science][Medline]
  123. Bakkenist CJ, et al. Initiating cellular stress responses. Cell (2004) 118:9–17.[CrossRef][Web of Science][Medline]
  124. O'Driscoll M, et al. The role of double-strand break repair–insights from human genetics. Nat. Rev. Genet. (2006) 7:45–54.[CrossRef][Web of Science][Medline]
  125. Kumagai A, et al. How cells activate ATR. Cell Cycle (2006) 5:1265–1268.[Medline]
  126. Delacroix S, et al. The Rad9-Hus1-Rad1 (9-1-1) clamp activates checkpoint signaling via TopBP1. Genes Dev. (2007) 21:1472–1477.[Abstract/Free Full Text]
  127. Tyagi A, et al. Resveratrol causes Cdc2-tyr15 phosphorylation via ATM/ATR-chk1/2-Cdc25C pathway as a central mechanism for S-phase arrest in human ovarian carcinoma Ovcar-3 cells. Carcinogenesis (2005) 26:1978–1987.[Abstract/Free Full Text]
  128. O'Prey J, et al. Effects of dietary flavonoids on major signal transduction pathways in human epithelial cells. Biochem. Pharmacol. (2003) 66:2075–2088.[CrossRef][Web of Science][Medline]
  129. Ye R, et al. The plant isoflavonoid genistein activates p53 and chk2 in an ATM-dependent manner. J. Biol. Chem. (2001) 276:4828–4833.[Abstract/Free Full Text]
  130. Ye R, et al. The isoflavonoids genistein and quercetin activate different stress signalling pathways as shown by analysis of site-specific phosphorylation of ATM, p53 and histone H2AX. DNA Repair (2004) 3:235–244.[CrossRef][Medline]
  131. Michan S, et al. Sirtuins in mammals: insights into their biological function. Biochem. J. (2007) 404:1–13.[CrossRef][Web of Science][Medline]
  132. Kaeberlein M, et al. Substrate-specific activation of sirtuins by resveratrol. J. Biol. Chem. (2005) 280:17038–17045.[Abstract/Free Full Text]
  133. McBurney MW, et al. The absence of SIR2alpha protein has no effect on global gene silencing in mouse embryonic stem cells. Mol. Cancer Res. (2003) 1:402–409.[Abstract/Free Full Text]
  134. Matsushita N, et al. Role of NAD-dependent deacetylases SIRT1 and SIRT2 in radiation and cisplatin-induced cell death in vertebrate cells. Genes Cells (2005) 10:321–332.[Abstract/Free Full Text]
  135. Wojewodzka M, et al. Sirtuin inhibition increases the rate of non-homologous end-joining of DNA double strand breaks. Acta Biochim. Pol. (2007) 54:63–69.[Medline]
  136. Jeong J, et al. SIRT1 promotes DNA repair activity and deacetylation of Ku70. Exp. Mol. Med. (2007) 39:8–13.[Medline]
  137. Borra MT, et al. Mechanism of human SIRT1 activation by resveratrol. J. Biol. Chem. (2005) 280:17187–17195.[Abstract/Free Full Text]
  138. Garske AL, et al. SIRT1 top 40 hits: use of one-bead, one-compound acetyl-peptide libraries and quantum dots to probe deacetylase specificity. Biochemistry (2006) 45:94–101.[CrossRef][Web of Science][Medline]
  139. Yuan Z, et al. SIRT1 regulates the function of the Nijmegen breakage syndrome protein. Mol. Cell (2007) 27:149–162.[CrossRef][Medline]
  140. Luo J, et al. Negative control of p53 by Sir2alpha promotes cell survival under stress. Cell (2001) 107:137–148.[CrossRef][Web of Science][Medline]
  141. Vaziri H, et al. hSIR2(SIRT1) functions as an NAD-dependent p53 deacetylase. Cell (2001) 107:149–159.[CrossRef][Web of Science][Medline]
  142. Langley E, et al. Human SIR2 deacetylates p53 and antagonizes PML/p53-induced cellular senescence. EMBO J. (2002) 21:2383–2396.[CrossRef][Web of Science][Medline]
  143. Cheng HL, et al. Developmental defects and p53 hyperacetylation in Sir2 homolog (SIRT1)-deficient mice. Proc. Natl Acad. Sci. USA (2003) 100:10794–10799.[Abstract/Free Full Text]
  144. Kamel C, et al. SirT1 fails to affect p53-mediated biological functions. Aging Cell (2006) 5:81–88.[CrossRef][Medline]
  145. Solomon JM, et al. Inhibition of SIRT1 catalytic activity increases p53 acetylation but does not alter cell survival following DNA damage. Mol. Cell. Biol. (2006) 26:28–38.[Abstract/Free Full Text]
  146. Cohen HY, et al. Calorie restriction promotes mammalian cell survival by inducing the SIRT1 deacetylase. Science (2004) 305:390–392.[Abstract/Free Full Text]
  147. Abdelmohsen K, et al. Phosphorylation of HuR by Chk2 regulates SIRT1 expression. Mol. Cell (2007) 25:543–557.[CrossRef][Web of Science][Medline]
  148. Yeung F, et al. Modulation of NF-kappaB-dependent transcription and cell survival by the SIRT1 deacetylase. EMBO J. (2004) 23:2369–2380.[CrossRef][Web of Science][Medline]
  149. Kobayashi Y, et al. SIRT1 is critical regulator of FOXO-mediated transcription in response to oxidative stress. Int. J. Mol. Med. (2005) 16:237–243.[Web of Science][Medline]
  150. Mostoslavsky R, et al. Genomic instability and aging-like phenotype in the absence of mammalian SIRT6. Cell (2006) 124:315–329.[CrossRef][Web of Science][Medline]
  151. Snyder RD. Assessment of atypical DNA intercalating agents in biological and in silico systems. Mutat. Res. (2007) 623:72–82.[Medline]
  152. Yamasaki M, et al. Soy-derived isoflavones inhibit the growth of adult T-cell leukemia cells in vitro and in vivo. Cancer Sci. (2007) 98:1740–1746.[CrossRef][Medline]
Received September 6, 2007; revised November 26, 2007; accepted November 27, 2007.


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