DNA damage-induced cell death: From specific DNA lesions to the DNA damage response and apoptosis

Cancer Letters xxx (2012) xxx–xxx

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Mini-review

DNA damage-induced apoptosis: From specific DNA lesions to the DNA damage response and apoptosis Wynand P. Roos, Bernd Kaina ⇑ Department of Toxicology, University of Mainz, Obere Zahlbacher Str. 67, D-55131 Mainz, Germany

a r t i c l e

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Article history: Available online xxxx Keywords: DNA damage DNA damage response Signaling ATM ATR p53 DNA repair Apoptosis

a b s t r a c t DNA damaging agents are potent inducers of cell death triggered by apoptosis. Since these agents induce a plethora of different DNA lesions, it is firstly important to identify the specific lesions responsible for initiating apoptosis before the apoptotic executing pathways can be elucidated. Here, we describe specific DNA lesions that have been identified as apoptosis triggers, their repair and the signaling provoked by them. We discuss methylating agents such as temozolomide, ionizing radiation and cisplatin, all of them are important in cancer therapy. We show that the potentially lethal events for the cell are O6-methylguanine adducts that are converted by mismatch repair into DNA double-strand breaks (DSBs), nonrepaired N-methylpurines and abasic sites as well as bulky adducts that block DNA replication leading to DSBs that are also directly induced following ionizing radiation. Transcriptional inhibition may also contribute to apoptosis. Cells are equipped with sensors that detect DNA damage and relay the signal via kinases to executors, who on their turn evoke a process that inhibits cell cycle progression and provokes DNA repair or, if this fails, activate the receptor and/or mitochondrial apoptotic cascade. The main DNA damage recognition factors MRN and the PI3 kinases ATM, ATR and DNA-PK, which phosphorylate a multitude of proteins and thus induce the DNA damage response (DDR), will be discussed as well as the downstream players p53, NF-jB, Akt and survivin. We review data and models describing the signaling from DNA damage to the apoptosis executing machinery and discuss the complex interplay between cell survival and death. Ó 2012 Elsevier Ireland Ltd. All rights reserved.

1. Introduction Chemical carcinogens, ionizing radiation (IR) and the ‘classic’ genotoxic anticancer drugs all attack the DNA, which is at the very heart of their genotoxic and toxic attributes. It is therefore important to shed light on the processes taking place on the level of damaged DNA, in order to facilitate our understanding of carcinogens and the effect of anticancer drugs on cells and tissues. Potentially lethal events for the cell are DNA double-strand breaks (DSBs) and DNA lesions that prevent the replication and transcription of DNA. Cells have developed sensors that detect these lesions. Sensor systems recognize the damage and relay the signal via kinases to ‘executors’, which start a process that either inhibits cell cycle progression and strengthens DNA repair or induces apoptosis that destroys the cell. The main ‘players’ of DNA damage recognition are ATM, ATR and DNA-PK, which phosphorylate a multitude of proteins and thus induce the DNA damage response (DDR), in which p53 and BRCA1/2 play important roles. Downstream are targets that either help cells to survive or destine them to undergo cell ⇑ Corresponding author. Address: Institut für Toxikologie, Obere Zahlbacher Str. 67, D-55131 Mainz, Germany. Tel.: +49 6131 393 3246; fax: +49 6131 230506. E-mail address: [email protected] (B. Kaina).

death. Here, we discuss the main pathways of survival and death, focusing on specific DNA lesions that were identified as triggers of apoptosis. We address methylating genotoxic agents, ionizing radiation and cisplatin as paradigmatic examples of DNA damaging agents that play an extremely important role in cancer therapy, and discuss the findings describing the repair of critical lesions and its interplay with downstream signaling finally leading to cell death by apoptosis. 2. Critical DNA lesions triggering apoptosis Genotoxic anticancer drugs such as methylating agents, IR and cisplatin modify DNA thereby triggering a cascade of events leading to apoptosis. These DNA lesions impede the normal function of DNA such as transcription and DNA replication. The evidence that DNA lesions are able to trigger apoptosis comes from studies on DNA repair defective cell lines, either due to mutation in or knockdown of essential repair proteins, and studies where cells were fed with cytotoxic modified nucleotide precursors that are incorporated into the DNA. Consequently, nearly all DNA repair defective cell lines are hypersensitive to killing by genotoxins, with the exception of mismatch repair (MMR) mutants. This has been shown for cells exhibiting defects in O6-methylguanine-DNA

0304-3835/$ - see front matter Ó 2012 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.canlet.2012.01.007

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methyltransferase (MGMT), base excision repair (BER), nucleotide excision repair (NER), repair of DSBs and repair of DNA crosslinks [1]. Also, feeding HSV-1 thymidine kinase expressing cells with ganciclovir, these cells are able to incorporate the modified nucleotide into its DNA, triggers a strong apoptotic response [2]. We should note, however, that over-expression of DNA repair genes does not always protect against genotoxin-induced apoptosis. For example, the amount of MGMT molecules per cell clearly correlates with cell survival up to a saturation level [3] while over-expression of N-methylpurine-DNA glycosylase (MPG) [4] has a detrimental effect. As a rule, over-expression of repair proteins involved in complex repair pathways seem to have a harmful effect on cells because of imbalances in the execution of the repair steps that are normally highly regulated. DNA damage is a generic term for many different DNA modifications that have the ability for activating apoptosis. The mechanisms whereby these different modifications initiate apoptosis differ based on the chemistry of the lesion and the manner in which the cell process or detect the DNA modification. In this review, we have therefore chosen three examples that are representative of different DNA damaging noxes that form different DNA adducts and lesions and consequently trigger apoptosis by distinct mechanisms. The following sections will focus on methylating agents, IR and cisplatin. 2.1. Methylating agents Methylating genotoxic agents are found in the environment [5,6], in processed meat [7], in cured tobacco [8] and are produced during endogenous metabolic processes [9]. These methylating species cause mutations during DNA replication and are therefore carcinogens. It is important to note that the amount of DNA lesions achieved by chronic low level exposure to these methylating agents are usually insufficient to trigger apoptosis and therefore, premalignant cells will not always be eliminated. Methylating agents are often used as anticancer drugs; these are procarbazine, dacarbazine (DTIC), streptozotocin and temozolomide. Because the levels of DNA lesions achieved during therapy with these chemotherapeutics are high enough, they trigger apoptosis in cancer cells [10,11]. All methylating anticancer drugs are SN1 agents. These agents methylate DNA at 13 positions, forming the main products N7-methylguanine (N7MeG), N3-methyladenine (N3MeA) and N3methylguanine (N3MeG). One of the minor products is O6-methylguanine (O6MeG) [12]. Depending on the cellular system, O6MeG, N3MeG and N3MeA are cytotoxic lesions that are able to trigger apoptosis (Fig. 1A). 2.1.1. Processing of O6-methylguanine into an apoptotic lesion The minor DNA lesion O6MeG, induced at less than 8% of total methylations [12,13], is of special importance because of its mispairing properties during DNA replication. O6MeG does not block DNA synthesis significantly, it causes the formation of GC to AT point mutations after two rounds of DNA replication and, consequently, is a DNA adduct [14] important for cancer induction [15] and cancer progression [16]. As O6MeG does not block DNA synthesis, it is interesting that O6MeG is also a cytotoxic DNA lesion [3,17]. Cytotoxicity is inversely related to mutagenesis [18], meaning that mutation does not cause cytotoxicity. This cytotoxic lesion is repaired in a stoichiometric reaction by MGMT. One methyl group is transferred from the O6-position of guanine to a cystein (Cys145) in MGMT, thereby inactivating MGMT and targeting it for proteosomal degradation [19]. The resistance of cells to O6MeG is therefore directly proportional to the amount of MGMT in the cell and the rate at which the cell can re-synthesize the enzyme, with protection increasing linearly with increasing MGMT up to the point when N-methylations such as N3MeG and

N3MeA become cytotoxic [3]. O6MeG was shown to be a DNA lesion capable of triggering apoptosis in cell systems were MGMT activity or expression was modulated, cells expressing MGMT are almost completely resistant to methylating agents for the dose range where N-methylations are not cytotoxic and conversely cells defective in repair of O6MeG by MGMT were sensitive to methylating agent-induced apoptosis in the same dose range [20]. As MGMT plays such a important role in protecting cells against methylating chemotherapeutics, it is not surprising that these chemotherapeutics have found application in the tumor types that express the lowest amounts of MGMT [21], namely malignant glioma [22] and metastatic melanoma [23]. O6MeG does not trigger apoptosis directly; it requires MMR [24,25] and DNA replication [26]. As a result, non-replicating cells and cells defective in MMR are resistant to methylating agents at doses where O6MeG triggers apoptosis. At higher doses, replication and MMR are no longer required [26,27]. During DNA synthesis in S-phase, O6MeG mispairs with thymine [28], this mismatch is a substrate for MMR and is bound by the heterodimer MutSa [29], comprised of MSH2 and MSH6. After recruitment of the heterodimer MutLa [30], comprised of PMS2 and MLH1, and exonuclease I [31] thymine is excised, but because of the mispairing properties of O6MeG, still present in the DNA, thymine is simply reinserted during the re-synthesis of the double-stranded DNA. As this remains a substrate for MMR, the futile removal and re-insertion of thymine will occur causing the formation of long persistent stretches of single-stranded DNA [32] until the cell again enters S-phase. At this stage, in a DNA synthesis [26] and futile MMR dependent manner [33], DSBs are formed [34]. MMR deficient cells became tolerant to O6MeG DNA lesions because they are unable to undergo apoptosis in response to this lesion. In fact, the level of MMR expression contributes to the sensitivity of cells to O6MeG; cells that express high levels of MMR are more sensitive to O6MeG-triggered apoptosis than cells expressing lower levels [35,36]. This tolerance of O6MeG lesions in the absence of MMR occurs at the expense of high levels of GC to AT point mutations [18]. If the DSBs that are formed due to futile MMR are not repaired by homologous recombination (HR) [37] in the post-treatment cell cycle [38] the cells die by apoptosis (Fig. 1A, left panel). 2.1.2. Processing of N-methylations can trigger apoptosis Unlike O6MeG, the N-methylations N7MeG, N3MeA and N3MeG are repaired by BER. Repair by BER depends on the following steps: Recognition, base removal, incision and re-synthesis. The recognition step is facilitated by a specific DNA glycosylase which recognizes and removes modified bases by hydrolyzing the Nglycosidic bond. In human cells, the glycosylase responsible for the removal of N-methylated bases is MPG (also known as 3-alkyladenine-DNA glycosylase (AAG) or alkylpurine-DNA N-glycosylase (APNG)). MPG is a type I glycosylase [39]. Type I glycosylases remove modified bases leaving an apurinic/apyrimidinic site (AP site) in DNA. MPG releases N3MeA, N7MeG, and N3MeG from DNA [40] and imbalanced expression renders cells more sensitive to methylating agents [4]. MPG-knockout mice are viable and show an increased level of mutations following methylating agent treatment [41]. Following base release by MPG, incision into the phosphodiester bond of the AP site occurs by AP endonuclease (APE1) (also known as APEX or Ref-1) resulting in 50 -deoxyribose-5-phosphate (50 dRP) and 30 OH [42]. The insertion of the first nucleotide is performed by DNA polymerase b (Polb) [43]. Polb has lyase activity and is able to catalyze the release of the hemiacetal form of 50 -dRP residues from incised AP sites by b-elimination during short patch BER [44]. During long patch BER further DNA synthesis is accomplished by Pole or Pold together with PCNA [45], resulting in repair patches of up to 20 nucleotides. The deoxyribosephosphate flap structure (50 -dRPflap)

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Fig. 1. Critical DNA lesions induced by methylating agents, ionizing radiation and cisplatin. (A) Methylating agents induce methyl DNA adducts that require processing in order to form DSBs. They also induce DNA replication blocking lesions ether directly or due to repair intermediates formed during BER. (B) Ionizing radiation induces DSBs directly. Similar to methylating agents, IR also induces DNA replication blocking lesions that may be converted into DSBs. (C) Cisplatin induces different DNA adducts that may have different consequences for transcription and replication.

is removed by PCNA-stimulated flap endonuclease (FEN1) [46]. The ligation step is performed by DNA ligase III in complex with XRCC1 during short patch BER or DNA ligase I during long patch BER. Deficiency in any one of the BER steps causes a severe repair defect. Down-regulation of APE1 sensitizes cells to methylating agents

[47], and complete loss of APE1 is lethal even on cellular level. Conditional APE1 knockouts undergo apoptosis as soon as APE1 is gone [48], which may reflect the presence of spontaneous AP sites that are formed at the frequency of 10,000 per cell per day. Alternatively, apoptosis observed in conditional APE1 knockout cells

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may be due to a lack of the APE1’s redox function, originally described as Ref-1. APE1 (Ref-1) regulates the activity of various transcription factors of which activator protein-1 (AP-1) and p53 are two examples [49]. Consequently, apoptosis observed in APE1 knockout cells might be due to transcription imbalance in addition to the presence of spontaneous AP sites. Cells knockout for Polb are viable, although Polb is indispensable during embryogenesis [50]. Polb deficient cells are hypersensitive to methylating agents [50] due to the induction of apoptosis [51]. Similar to O6MeG, the N-methylations N7MeG, N3MeA and N3MeG do likely not signal apoptosis directly, they require processing into an apoptosis triggering DNA lesion (Fig. 1A). This is accomplished by mechanisms that are DNA replication dependent and independent. Both mechanisms rely partly on BER. The evidence for DNA replication independent activation of apoptosis comes from studies on non-replicating blood cells exposed to methylating agents [26,52]. Peripheral non-stimulated blood lymphoid cells (PBLC) undergo apoptosis when treated with a sufficiently high dose of methylating agent [26]. This onset of apoptosis is preceded by the accumulation of DSBs. In non-replicating monocytes, methylating agents trigger apoptosis at much lower dose levels. This is because monocytes have a defect in BER, they do not express XRCC1 and ligase III [52]. In both PBLCs and monocytes the model for apoptosis initiation requires DSBs. During repair of N-methylations DNA single-strand breaks (SSBs) are formed as BER intermediates. If sufficient N-methylation lesions are present, repair in overlapping patches may lead to DSBs. The chance for these DSBs forming increases with the loss of XRCC1 and ligase III, as the BER intermediates, i.e. SSBs, persist. It is conceivable that during cancer therapy with alkylating agents, DSBs and apoptosis are induced as a result of mis-repair of Nalkylpurines. However, to induce a sufficient amount of N-methylations to cause overlapping BER patches that give rise to DSBs is very unlikely during cancer therapy, as the attainable clinical doses are likely not high enough. N-methylations also induce apoptosis in a DNA replication dependent manner. The DNA replication blocking lesions are N3MeA and N3MeG as well as the BER repair intermediates, i.e. AP sites and SSBs. When the replication fork encounters one of these lesions it is blocked, which may lead to replication fork collapse and the formation of DSBs. Consistent with this model, cancer cells transiently over-expressing MPG that cause a higher frequency of AP sites are sensitized toward methylating agent-triggered apoptosis [53]. Additionally, in Polß knockout cells that cannot perform BER to completion, non-sealed SSBs are transformed into DSBs during S-phase, which in turn trigger apoptosis [54]. Interestingly, HR also protect cells against these replication dependent DSBs arising from N-methylation lesions [55]. It should be clear from the above discussion that the response of cells to methylating agents is a complex matter that depends on multiple repair factors such as MGMT, BER and DSB repair by HR. Therefore, depending on the DNA repair status of any given cell and the dose level of the methylating agent the response of the cell may be a mixed function of both O6MeG and N-methylpurines (Fig. 1A). This notwithstanding, both O6MeG and N-methylpurines are processed into DSBs that seem to be the ultimate apoptosis trigger. It should be noted that it is not clear whether tumors are generally BER competent. For BER deficient or imbalanced tumors, both N-methylpurines and O6MeG might provoke cancer cell kill. Therefore, it is reasonable to focus current research on strategies targeting the BER pathway together with inhibiting MGMT. 2.2. Ionizing radiation Any discussion on DNA damage caused by oxidative stress is incomplete without mentioning IR. Radiation is unique amongst

DNA-damaging agents in the broadness of the different DNA lesions it induces and in the manner that it does this. Due to ionization tracts, reactive oxygen species (ROS) are generated at localized regions in the nucleus. Of the total DNA lesions induced by IR, 80% are base modifications and 20% are damage to the sugar phosphate backbone [56]. One Gy of IR induces approximately 2000 base modifications per cell [57]. Of the 20 different base lesions induced by IR, 8-oxo-7,8-dihydroguanine (8-oxoG) and 5,6-dihydroxy-5,6-dihydrothymine (thymine glycol, ThG) are probably the best known [56]. In addition to the base lesions, one Gy also induces the formation of 1000 DNA single-strand breaks (SSBs) [58] and 35 DSBs [59] per cell. Today it is mostly accepted that DSBs are the most important cytotoxic lesion induced by IR. Cells defective in repairing DSBs by non-homologous end joining (NHEJ) or HR are sensitive to IR-induced cell kill, with NHEJ playing the dominant protective role [60]. NHEJ is based on the interplay of the proteins Ku70, Ku80 and DNA-PKcs (DNA-dependent protein kinase, catalytic subunit). Ku70/Ku80 recognizes the DNA ends and is bound there, and DNA-PKcs is eventually bound to these proteins, forming a complex that is described as DNA-PK (DNA-dependent protein kinase) [61]. This complex has kinase activity. Not only does the complex phosphorylate H2AX, but also XRCC4 (X-ray repair complementing defective repair in Chinese hamster cells 4), ligase IV, artemis and XLF (XRCC4-like factor), which then remain at the lesion and bring about the joining of the broken DNA ends. Although DSBs are clearly responsible for cell kill upon IR, their contribution to IR-induced apoptosis is still unclear as there is no clear correlation between cell killing and apoptosis in some cell types [62]. The identification of specific DNA lesions responsible for the activation of apoptosis following IR is made extremely difficult due to the broad spectrum of damages induced by IR (Fig. 1B). ThG is a DNA replication blocking lesion [63] and, therefore, may account for replication-dependent activation of apoptosis if not repaired by BER following recognition by Nth1 (thymine-glycol-DNA N-glycosylase) [64] or the backup glycosylase Neil1 (Nei like 1) [65]. Experimental evidence that ThG is able to initiate apoptosis is still lacking although loss of Neil1 does sensitize cells to IR induced cell kill [66]. A critical mutagenic lesion induced by IR is 8-oxoG (the isomeric form is 8-hydroxyguanine). 8-oxoG mispairs with adenine during DNA replication giving rise to GC to TA transversion mutations. It is repaired by Ogg1 (8oxoguanine-DNA glycosylase) utilizing the BER pathway [67]. Ogg1 knockout cells are not hypersensitive to IR and OGG1/CSB double knockouts, in which 8-oxoG is neither removed in the transcribed nor un-transcribed strand, are only slightly more sensitive. MMR recognizes 8-oxoG/A mispairs [68], correcting them post-replicatively. Evidence for a contribution of MMR in 8oxoG-triggered apoptosis is lacking. Similar to our discussion on methylating agents where BER repair intermediates like SSBs and AP-sites block DNA replication and thereby trigger apoptosis, it should be kept in mind that 80% of DNA damage induced by IR are base modifications - the majority of which will be subject to BER. Therefore, the contribution of BER repair intermediates to apoptosis induction in IR treated replicating cells should not be ignored. The most convincing evidence that DSBs are effective triggers of apoptosis comes from studies where ‘‘clean’’ DSB were induced by electroporating restriction enzymes into cells. By introducing the blunt-end cutter Pvu II into cells, a dose-dependent increase in DSBs was observed and these DSBs induced a strong apoptotic response [69]. Although ‘‘clean’’ DSBs are able to trigger apoptosis effectively, the apoptotic response following IR may reflect a mixed function of the DSBs that activate apoptosis directly as well as the base modifications and BER repair intermediates that block DNA replication and induce DSBs indirectly (Fig. 1B).

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2.3. Cisplatin Cisplatin (cis-diammine-dichloroplatinum (II), cDDP, cisPtCl2(NH3)2) enters the cells ether through passive diffusion or by active transport by Ctr1 (copper transporter 1) [70] or Oct2 (organic cation transporter 2) [71]. The intracellular concentration of cisplatin is counteracted by export accomplished by ATP7B (ATPase, Cu++ transporting, beta polypeptide) and ATP7A (coppertransporting ATPase 1) [72] as well as by inactivation by glutathione [73] or metallothionein [74]. Resistance/sensitivity of cancer cells have been attributed to different expression levels of these mechanisms that control the intracellular concentration of cisplatin. This review will not focus on this; instead it will focus on the DNA lesions induced by cisplatin along with how these DNA lesions activate the apoptotic machinery. The reader should however keep in mind that the intracellular cisplatin concentration will determine how many DNA lesions are formed and therefore how effectively the apoptosis machinery is activated. Unlike methylating agents, cisplatin induces crosslinked DNA adducts. Upon entering the cell, cisplatin is no longer protected by the high chloride anionic concentration in the blood and loses a chloride ligand through hydrolyses forming aquated cisplatin (cis-PtCl(NH3)2(H2O)). Aquated cisplatin binds to N7 of guanine and water is displaced forming the cisplatin monofunctional adduct. The second chloride ligand is lost through hydrolysis and the bifunctional adduct is formed through cisplatin binding to N7 of adjacent guanine or adenine or nonadjacent guanine [75]. The resulting crosslinks (Fig. 1C) are distributed in their frequency as follows: guanine-guanine intrastrand crosslinks comprise 47–50% of lesions, guanine-adenine intrastrand crosslinks comprise 23– 28% of lesions, guanine-base-guanine intrastrand and guanineguanine interstrand crosslinks (ICLs) of two nonadjacent guanines comprise 8–10% of lesions while monofunctional binding to guanine comprise 2–3% of lesions [76,77]. The intrastrand crosslinks are repaired by nucleotide excision repair (NER) while the ICLs are repaired by a complex mechanism, which makes use of components of NER, DSB repair and TLS. Cells defective in NER due to mutations in ERCC1 (excision repair cross-complementing rodent repair deficiency complementation group 1), ERCC4 (XPF), ERCC2 (XPD) or ERCC3 (XPB) are hypersensitive to cisplatin [78], which provide evidence that cisplatin-induced DNA lesions are the primary cause of cell death. ERCC1 and ERCC3 mutants also show higher levels of apoptosis following cisplatin treatment, supporting the hypothesis that non-repaired cisplatin-induced DNA lesions trigger apoptosis [79]. ERCC1 was shown to be involved in the repair of ICLs rather than the repair of DNA-intrastrand crosslinks [80], and ERCC1 mutants are more sensitive to cisplatin than ERCC3 mutants [79], indicating that cisplatin-induced ICLs play a bigger role in inducing apoptosis than cisplatin intrastrand crosslinks. Furthermore, human testis tumor cell lines express low levels of ERCC1 and XPF, and consequently these cell lines are defective in ICL repair and extremely sensitive to cisplatin-induced apoptosis; over-expression of ERCC1/XPF in these cells enhanced ICL repair and reduced the level of apoptosis [81]. If not repaired, these adducts can theoretically trigger apoptosis via two mechanisms: they may block transcription and/or DNA replication (Fig. 1C).

Fig. 2. Activation of ATM and ATR during the DNA damage response. (A) ATM is activated by DSBs. (B) ATR becomes activated by stalled DNA replication forks. See text for detailed discussion on the activation of ATM and ATR.

apoptosis. Important sensors of these DNA lesions are the phosphatidyl inositol 3-kinase-like kinases ATM (ataxia-telangiectasia mutated) and ATR (ATM- and Rad3-related). ATM is mainly activated by DSBs formed directly by IR, or indirectly by methylating agents and cisplatin due to sustained replication blockage causing collapse of replication forks [82,83]. ATR is activated in response to stalled DNA replication forks. Since IR, methylating agents and cisplatin all induce DNA base damage that can block DNA replication (Fig. 1) they are able to activate ATR [82,84,85]. ATM/ATR is implicated in three crucial functions: regulation and stimulation of DSB repair, activation of cell cycle checkpoints, and signaling to apoptosis. Therefore, they are key nodes in making the decision between survival and death following genotoxin exposure. This review will focus on how ATM and ATR initiate the apoptosis machinery. Although the signaling from ATM and ATR overlap, they are not identical and will therefore be discussed separately.

3. DNA damage response

3.1. Detection of DSBs

DSBs and DNA replication blocking lesions are apoptotic DNA lesions and, therefore, cells are equipped in detecting these lesions upon formation (Fig. 2). Recognition of these DNA lesions starts a protein cascade, which finally results in cell cycle arrest (checkpoint activation) and DNA repair. If DNA repair fails, or is overwhelmed by to many DNA lesions, these sensors initiate

DSBs are detected very rapidly. The detection and subsequent downstream signaling from DSBs requires the interplay between the MRN complex and ATM (Fig. 2A). It is possible that PARP-1 (poly(ADP-ribose) polymerase) is also involved as indicated by some recent data [86]. PARP-1 catalyzes the formation of poly(ADP-ribose) chains that were proposed to facilitate the docking

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of the MRN complex to the DSB [87]. The MRN complex consists of NBN (NBS1 or Nibrin), MRE11 and RAD50. MRE11, which exhibits exo- and endonuclease activity and RAD50 have DNA binding capabilities while NBS1 is responsible for shuttling the MRN complex to the nucleus [88]. RAD50 holds the DSB ends together while MRE11 makes use of its nuclease activity to process the DSB ends. Once the MRN complex is localized to the DSB, ATM is recruited to this site. The serine/threonine protein kinase ATM is found in its inactive state as a homodimer that upon recruitment to the DSB-MRN site activates itself by phosphorylation [89]. This autophosphorylation occurs at Ser1981 [89], which leads to the disassociation of inactive ATM dimers into catalytic active monomers. Additional autophosphorylation sites on Ser367 and Ser1893 were also detected in vitro and in vivo. In humans, mutations in these phosphorylation sites abrogate ATM signaling upon ionizing radiation [90]. However, in mouse, mutation in these phosphorylation sites do not influence ATM activity [91]. Interestingly, the in vitro activation of ATM kinase activity is not strictly dependent on autophosphorylation on Ser1981 [92]. In the absence of the MRN complex, the catalytic activity of ATM and Ser1981 phosphorylation is reduced [93] and ATM is not recruited to the DSB [94]. The absence of MRN also affects the activation of several ATM targets [95]. ATM monomerization was also shown to occur by interaction of ATM with the MRN complex and single-stranded DNA [94]. In vitro, unwinding of DNA ends by the MRN complex, but not ATM autophosphorylation, is required for monomerization of ATM and ATM activation [94]. In vivo, active ATM monomers are responsible for DSB dependent signaling, as it phosphorylates a large number of proteins. Among them is NBN within the MRN complex, the activity of the MRN complex is strengthened by this phosphorylation. The presence and absence of MRN differentially affects the activation of several ATM targets [95] and can be used to explain the mechanism of ATM activation. In this model, ATM is autophosphorylated via signals generated by structural changes of the chromatin, resulting in the release of activated ATM monomers. Following ATM autophosphorylation, H2AX is phosphorylated and the MRN complex, BRCA1 (breast cancer type 1 susceptibility protein) and SMC1 (structural maintenance of chromosomes 1) are recruited to the DSB independently. Activated ATM is now recruited to the DSB via NBN and BRCA1. ATM can now interact and phosphorylate NBN, BRCA1 and SMC1. If ATM is activated in the absence of DSBs or not recruited to the DSB, then only nucleoplasmic substrates (p53, Chk2) are phosphorylated. Beside the MRN complex, additional co-factors seem to be involved in determining the substrate-specificity of ATM. Thus, loss of 53BP1 (p53 binding protein 1) decreases ATM autophosphorylation and ATM mediated phosphorylation of Chk2 and SMC1 [96,97]. 3.2. Detection of blocked DNA replication forks One of the essential processes of life is DNA synthesis. Normal DNA polymerases cannot synthesize new DNA if the template DNA contains modifications, for example intrastrand crosslinks, or replication blocking DNA lesions such as N3MeA and AP sites. These DNA lesions are normally bypassed by translesion synthesis (TLS) polymerases. However, if too many of these lesions are present in replicating DNA, the replicating fork is stalled and may lead to DSBs within the fragile DNA single-stranded regions. To prevent the formation of DSBs following replication arrest, a detection system has evolved that stabilizes DNA in these areas and initiates repair. This detection system uses RPA (replication protein A), the ATR-ATRIP complex, TopBP1 (DNA topoisomerase 2-binding protein 1) and the 9-1-1 complex (Fig. 2B). During normal DNA replication, RPA binds to single-stranded DNA and prevents it from winding back on itself or from forming secondary structures. Un-

der replication blocking conditions these long stretches of RPAbound single-stranded DNA [98] persist and the serine/threonine protein kinase ATR is recruited to these sites by ATRIP (ATR-interacting protein) [98]. The ATR-ATRIP complex is bound to RPAbound single-stranded DNA by direct interaction of ATRIP with RPA [99] and oligomerization [100]. The recruitment of the 9-1-1 complex is a multi-step process. The 9-1-1 complex consists of Rad9, Hus1 and Rad1 [101]. Rad9 interacts with TopBP1 [102] and recruits TopBP1 to the stalled replication fork [103]. TopBP1 directly activates the ATR-ATRIP complex, most likely via a conformational change of the ATR-ATRIP complex [104]. ATM and ATR do not work independently. Thus, upon the formation of DSBs, ATR chromatin loading depends on ATM [105] since phosphorylation of TopBP1 by ATM is necessary for ATR activation [106]. Upon stalling of the replication fork, ATM is phosphorylated by ATR at Ser1981 and activated [107]. Similar to ATM, activated ATR phosphorylates many proteins (BRCA1 and BRCA2, Rad51 and FANCJ, among others), whose activity is modulated as a result. One of the main pathways activated by ATR is homologous recombination (HR), which is required for the abolition of replication blockage. 3.3. Phosphorylation of histone 2AX DSBs and stalled replication forks cause the phosphorylation of neighboring histone 2AX at Ser139 (H2AX for the unphosphorylated form and cH2AX for the phosphorylated form) via ATM and ATR respectively. For DSBs [108], the distribution of cH2AX within the nucleus is localized around the DSB. These cH2AX foci already appear 10 min after DSB induction. Over time the amount and intensity of these cH2AX foci reaches a maximum (at about 1 h) and later on decreases due to the activity of the phosphatase PP4 [109], showing that DSB repair has occurred. As stated above, stalled replication forks activate ATR and ATR phosphorylates H2AX. The functional significance of cH2AX is assumed to be a signal that facilitates the repair of free DSBs or DSBs that are formed at stalled replication forks, presumably by causing the chromatin to be more accessible for DNA repair. 3.4. DSB repair by homologous recombination The repair of DSB has been studied intensively over the past years; it has to be seen as part of the cell’s DNA damage response and will therefore be dealt with in this context. HR plays a significant role as resistance factor for genotoxic anticancer drugs, including methylating agents [37] and cisplatin [110], because it facilitates repair of DSBs that are formed at blocked replication forks. HR is complex. It requires additional systems beside damage recognition, which finds the intact homologous DNA strand, transports it to the lesion, partially denatures the DNA, initiates and executes DNA synthesis at the intact matrix, separates the DNA strands and restores the undamaged and intact DNA [111]. Describing this complex process is beyond the scope of this review, especially since the three-dimensional DNA and chromatin structure have to be considered. What is important is that ATM/ATR signaling activates proteins by phosphorylation that are essential for HR. These are, among others, BRCA1, BRCA2, FANCD2, WRN and BLM. 4. Apoptosis The current paradigm states that if DNA repair fails, cells undergo death by activating one of the programmed death pathways, i.e. apoptosis. This implies that cells containing sufficient DNA damage to overwhelm its capability of repairing its DNA will be eliminated from the population in a given tissue. There are

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various experimental systems, which show that defects in the execution of apoptosis increase cancer incidence (e.g. [112]). Thus apoptosis following DNA damage is a protective mechanism that prevents carcinogenesis. DNA damage most often activates the extrinsic death receptor apoptosis pathway (Fas, CD95, Apo-1) and/or the intrinsic mitochondrial apoptosis pathway [113]. The Fas pathway depends on receptor activation, assembling of the DIS-complex consisting of FADD (Fas-associated protein with death domain) and pro-caspase-8 and -10 which leads to the activation of the caspase cascade culminating in the digestion of DNA by CAD (caspase-activated DNAse) and, at the same time, proteolytic inactivation of cellular proteins by the executing caspases-3 and -7. The mitochondrial apoptosis pathway depends on the regulation of cytochrome c release from the mitochondria into the cytoplasm, which is governed by pro-apoptotic proteins like Bax (Bcl-2-associated X protein) and Bak (Bcl-2 homologous antagonist/killer) and anti-apoptotic proteins like Bcl-2 (B-cell lymphoma 2) and Bcl-XL (B-cell lymphoma-extra large). Upon cytochrome c release the apoptosome forms, consisting of pro-caspase-9, Apaf-1 (Apoptotic protease activating factor 1) along with cytochrome c. Similar to the DIScomplex, the apoptosome initiates the caspase cascade leading to protein and DNA inactivation. How does the DNA damage response in the nucleus activate the apoptosis machinery in the cytoplasm?

increasing its transactivation activity [118]. Consequently, the nuclear localization of p53 and its DNA binding activity increases and its target genes are transcriptionally induced (Fig. 3A). p53 transcriptionally regulates both pro- and anti-apoptotic genes. Proapoptotic genes are Fas-R, Bax, Puma (p53 upregulated modulator of apoptosis), Noxa, Apaf-1 and Pidd (p53-induced protein with death domain), all encode downstream DNA damage response proteins. Anti-apoptotic genes are DDB2, XPC, Fen1, MGMT and MSH2 [119], all encode upstream DNA repair proteins. The transcription factor p53 therefore plays a dual role: a protective role by increasing DNA repair and a sensitization role by increasing apoptosis execution. This dual role is illustrated by way of examples: For methylating agents p53 stimulates apoptosis due to its up-regulation of Fas [120]. For IR, p53 also stimulates apoptosis due to its upregulation of Noxa [121]. For cisplatin, in testicular germ cancer cell lines that are defective in the repair of ICLs, p53 further sensitizes by inducing Noxa and Puma [122]. Conversely, p53 caused resistance due to increased p53 mediated XPA nuclear localization [123] after cisplatin and XPC and DDB2 expression [124] after alkylating agent treatment and therefore stimulates repair of crosslinks and ICLs. In this case, the decision of p53 to stimulate apoptosis or DNA repair is obviously dependent on the genotoxic agent. It may also be dependent on the dose level of a given genotoxin.

4.1. Role of p53

4.2. Transcriptional inhibition

Important phosphorylation targets of ATM and ATR are Chk1, Chk2 and p53. ATM phosphorylates Chk2 (checkpoint kinase-2) after the formation of DSBs at Thr68 [114], while ATR phosphorylates Chk1 (checkpoint kinase-1) at Ser345 upon stalled DNA replication forks [115]. In turn, Chk2 and Chk1 phosphorylate the transcription factor p53 at Ser20 [116]. This prevents the ubiquitination of p53 by the ubiquitin E3 ligase MDM2 and prevents its proteosomal degradation and the p53 protein is stabilized [117]. ATM and ATR can also directly phosphorylate p53 at Ser15, thereby

The inhibition of transcription is able to trigger apoptosis [125]. Since bulky DNA lesions, like those induced by cisplatin, in the transcribed DNA strand block transcription [126], it is reasonable to propose that this is a mechanism whereby DNA damage triggers apoptosis [127]. Apoptosis may occur via this mechanism because vital proteins, required for life, are no longer produced. In support of this, it was shown that Cockayne’s syndrome cells that exhibit a defect in transcription coupled repair (TCR, a NER sub-pathway) are sensitive to bulky DNA damage triggered apoptosis and that

Fig. 3. Downstream signaling from the DNA damage response to the apoptosis machinery. (A) ATM and ATR cause stabilization of p53. p53 plays a critical role in regulating DNA repair and DNA damage-triggered apoptosis. (B) Inhibition of transcription causes sustained SAPK/JNK signaling due to lack of MKP-1 expression, thereby activating apoptosis. (C) ATM activates the NF-jB pathway. Activated NF-jB has pro- and anti-apoptotic function. (D) PTEN abolishes the suppression of apoptosis by Akt. See text for description on how p53, SAPK/JNK, NF-jB and Akt functions following the induction of DNA damage by methylating agents, IR and cisplatin.

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apoptosis correlated with the amount of transcriptional inhibition [128]. Indeed, NER mutants display a stronger block in transcription than wild-type cells [125]. Although it is accepted that inhibition of transcription does contribute to cisplatin-induced apoptosis [79,129] it is still a matter of debate how much it contributes. As non-repaired bulky lesions have the ability to inhibit both transcription and DNA replication, the possibility remains that apoptosis also results from DNA breaks produced as a consequence of replication blockage. Experiments with the cyclophosphamide derivative maforfamide showed that cells seem to be more sensitive to apoptosis triggered by blockage of DNA replication than transcription [130]. Thus, more research is required to solve this important question. 4.3. Sustained JNK and p38 kinase activation The mechanism whereby sustained JNK and p38 kinase activation triggers apoptosis provides an example of how transcriptional inhibition may trigger apoptosis. DNA damaging agents have been shown to activate SAPK/JNK (stress-activated protein kinase/c-Jun N-terminal kinase) and p38 kinases, which results in an increase in c-Jun level and the activity of the transcription factor AP-1 (Fig. 3B). This sustained activation of AP-1 is accompanied by increased Fas-L expression. NER repair defective mutants display a higher level of sustained JNK/p38 kinase activation, indicating DNA damage is responsible for the response [131]. Together with DNA damage-induced p53 up-regulation and subsequent Fas expression, the AP-1 mediated Fas-L expression drives the extrinsic apoptosis pathway upon DNA damage. How is sustained activation of JNK/p38 kinase achieved? JNK/p38 kinases become activated by EGF receptor phosphorylation. This is counteracted by the phosphatase MKP1 that dephosphorylates JNK and, therefore, downregulates its activity [132,133]. Genotoxic agents provoke a reduction in the level of MKP1, which is likely due to transcription blockage due to bulky lesions in the DNA, and therefore ameliorates apoptosis by sustained up-regulation of JNK/p38 kinase [131,134]. We should note that sustained JNK activation also activates the mitochondrial apoptosis pathway via Bax phosphorylation and mitochondrial localization [135]. 4.4. Caspase-2 activation The only pro-caspase constitutively present in the nucleus is caspase-2 [136,137], which is therefore a possible player in DNA damage-triggered apoptosis. Caspase-2 has been shown to play a role in cisplatin and IR induced apoptosis [138,139]. Caspase-2 induced apoptosis requires caspase-9 activation [140] indicating that caspase-2 acts most often through the mitochondria damage pathway. Caspase-2 may activate apoptosis in the following manners. (1) It acts upstream of the mitochondria by inducing Bid cleavage, Bax translocation and subsequent cytochrome c release [138]. (2) It directly activates the mitochondrial pathway [141], which is independent of its enzymatic activity. Caspase-2 can also disrupt the interaction of cytochrome c with anionic phospholipids, notably cardiolipin, and thereby enhances the release of cytochrome c [142]. (3) This caspase is able to activate apoptosis following IR and Chk1 inhibition in a p53, Bcl-2 and caspase-3 independent manner [143]. 4.5. Role of NF-jB First, a short primer in NF-jB signaling. Nuclear factor jB (NFjB) is found in an inactive state in the cytoplasm [144] and becomes rapidly activated and translocated into the nucleus upon signaling from the receptor activator of NF-jB (RANK) [145], tumor necrosis factor receptor (TNFR) [146] or toll-like receptors (TLR)

[147] on the cell membrane. NF-jB is kept inactive in the cytoplasm due to its binding to its inhibitor IjB (inhibitor of jB) that shields the nuclear localization signal (NLS) of NF-jB. For the activation of NF-jB, IjB is phosphorylated by IjB kinase (IKK) at Ser32 and Ser36 (human IjB), which leads to the ubiquitination and proteasomal degradation of IjB [148]. NF-jB then becomes nuclear localized and can mediate the transcription of its target genes. Apart from its activation by cell receptors it can also be activated by DNA damage (Fig. 3C). DSB causes the nuclear localization of NEMO (NF-jB essential modulator) [149] followed by NEMOs phosphorylation by ATM [149]. NEMO and ATM forms a complex that is shuttled to the cytoplasm [150]. In the cytoplasm, IjB kinase (IKK) binds to NEMO/ATM. IKK phosphorylates the NF-jB inhibitor IjBa that traps NF-jB in the cytoplasm. Phosphorylated IjBa is subsequently ubiquitinated and degraded via the 26S proteasome pathway, liberating NF-jB and NF-jB becomes nuclear localized where it can perform its transcriptional function. Transcriptional activation by NF-jB leads to changes in pro- and antiapoptotic factors. Thus, it causes the expression of Bcl-xL, A1/Bfl1 [151], c-IAP2 [152] and TRAF1 and 2 [153]. Bcl-xL and A1/Bfl-1 are both members of the Bcl-2 family of proteins that inhibit apoptosis by heterodimerization with the pro-apoptotic Bcl-2 family members [154–156]. Since c-IAP2, TRAF1 and TRAF2 are involved in the inhibition of the death receptor pathway; the induction of these proteins by NF-jB suppresses the activation of caspase-8 dependent apoptosis [153]. Interestingly, NF-jB also up-regulates MKP-1 [157], which counteracts JNK phosphorylation and JNK driven apoptosis observed after sustained JNK activation following DNA damage [158]. Thus, NF-jB plays an important role as an anti-apoptotic player. However, recently NF-jB has also been implicated in the DDR where NF-jB activation leads to increased DNA damage and therefore sensitization of cells to genotoxins [159].

4.6. Role of Akt Akt is a serine-threonine kinase that plays a central role in multiple cellular processes. Akt activation by PI3-kinases (these are receptor activated kinases; to our knowledge ATM, ATR and DNA-PK do not phosphorylate Akt) causes suppression of apoptosis [160]. This inhibition of apoptosis by Akt has been characterized in many cancer cell systems [161–164]. Akt suppresses apoptosis by the following mechanisms (Fig. 3D): (1) It phosphorylates BAD on Ser136 [165,166]. BAD is a member of the Bcl-2 family proteins and acts by binding to Bcl-2 or Bcl-XL and thereby inhibiting their anti-apoptotic function [156]. When Akt phosphorylates BAD, BAD become sequestrated by 14-3-3 proteins and is inactivated, as the phosphorylated form of BAD has a lower binding affinity for Bcl-XL [167]. The release of Bcl-2 and Bcl-XL liberates these anti-apoptotic proteins and thus they can fulfill their apoptosis suppression function. (2) Akt phosphorylates human caspase-9 on Ser196 [168] thereby inhibiting it. Caspase-9, along with Apaf-1 and cytochrome c, form the apoptosome, which is required for mitochondrial mediated apoptosis [169]. (3) Akt phosphorylates apoptosis signal-regulating kinase 1 (ASK1) on Ser83 and this results in the inhibition of apoptosis induced by ASK1 [170]. ASK1 is upstream to JNK and p38 kinase, which is upstream to Bid cleavage, Bax translocation and cytochrome c release [171]. (4) Akt stimulates the activity of NF-jB by promoting the degradation of the NF-jB inhibitor IjB [172]. (5) Akt prevents the nuclear localization of p53. It binds to and phosphorylates Mdm2, the ubiquitin E3 ligase, on Ser166 and Ser186 thereby inducing its nuclear import or increasing its ubiquitin ligase activity [173,174]. Mdm2 is greatly involved in the inactivation of p53. Thus, increase in nuclear Mdm2 can inactivate the pro-apoptotic transcriptional function of p53.

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4.7. Role of PTEN Genotoxins have to overcome the suppression of apoptosis by Akt in order to activate apoptosis effectively. A player involved is PTEN (Fig. 3D). IR and cisplatin are both able to activate PTEN [175,176]. PTEN is a phosphatase that counteracts the suppression of apoptosis by preventing the phosphorylation of Akt, thereby initiating apoptosis. Another link between Akt and DNA damage is p53 [177]. The suppression of p53 by Akt following DNA damage may lead to a high non-tolerable level of unrepaired DNA lesions since p53 is inefficient in up-regulating DNA repair genes as outlined above. In order to undergo apoptosis efficiently, the cell needs to overcome the pro-survival signaling of Akt. The consequences for the cell in failing to do this might be (a) necrotic cell death, which in the case of chemo- or radiation therapy will lead to inflammation, or (b) fixation of DNA damage, which might lead to mutation and/or tumorigenesis. This is an example that shows the complex interplay between multiple pathways in the DNA damage response. 4.8. Role of survivin An interesting example where the same player is able to provoke both suppression of apoptosis and stimulation of DNA repair is survivin. Survivin is a member of the inhibitor of apoptosis (IAP) family; it directly inhibits the activity of caspases, notably caspase3 [178]. Recently it was shown that survivin also stimulates DSB repair by its interaction with DNA-PK [179]. Therefore, cancer cells that have increased levels of survivin may be protected against IR on two fronts: by blocking apoptosis and stimulating DSB repair by NHEJ. It is unlikely that survivin also stimulates repair of alkylation and cisplatin-induced lesions since DSBs formed in response to these agents are mostly repaired by HR [37,110]. More experimentation, however, is required to clarify this open question. 5. Conclusions It should be clear from the above discussion that the cell’s response to DNA damaging insults is complex. We make the case that most DNA lesions are converted into at least one of these DNA structures: DSBs and DNA replicating blocking lesions. IR induces DSBs directly while methylating agents and cisplatin induce replicating blocking lesions that require processing in order to become DSBs. The cascade of events initiated by ATM and ATR leads to cell cycle checkpoint activation and DNA repair. In order for this cascade to initiate apoptosis DNA repair has to be overwhelmed. If DNA repair is overwhelmed, DSBs and DNA replicating blocking lesions will persist and the signaling cascade will not be turned off. A second point we would like to stress pertains to the current paradigm stating that low levels of DNA damage trigger their repair whereas high levels of DNA damage (or residual DNA damage) activates apoptosis. This paradigm receives support from the dual function of p53 and NF-jB. They both transcribe pro-survival and pro-apoptosis genes. Therefore, p53 and NF-jB initially protect cells against DNA damage while, if the damage is not repaired, they becomes pro-apoptotic. Further support is found in the interaction of the DDR and the Akt pathway. When DNA damage is not repaired, the suppression of apoptosis by Akt is abolished. A similar argument can be made for the long-lasting activation of JNK. In order to switch off JNK signaling, cell’s have to up-regulate MKP-1 that is prevented however by DNA adducts blocking transcription. The apoptosis pathways finally activated following this interplay of pro-survival and pro-apoptosis signaling are the death receptor and/or the mitochondrial pathway, depending on the cellular background. This is further complicated in cancer cell systems, as can-

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cer cells suppress the apoptosis machinery by for example increasing survivin levels, thereby effecting both DNA repair and the execution of apoptosis. Under these conditions cancer cells are successful in attaining resistance against DNA damaging therapeutics. Acknowledgements Work of the authors is supported by the German Research Council (DFG), Deutsche Krebshilfe, Stifung Rheinland-Pfalz and University Medical Center Mainz. References [1] B. Kaina, DNA damage-triggered apoptosis: critical role of DNA repair, doublestrand breaks, cell proliferation and signaling, Biochem. Pharmacol. 66 (2003) 1547–1554. [2] M.T. Tomicic, R. Thust, B. Kaina, Ganciclovir-induced apoptosis in HSV-1 thymidine kinase expressing cells: critical role of DNA breaks, Bcl-2 decline and caspase-9 activation, Oncogene 21 (2002) 2141–2153. [3] B. Kaina, G. Fritz, S. Mitra, T. Coquerelle, Transfection and expression of human O6-methylguanine-DNA methyltransferase (MGMT) cDNA in Chinese hamster cells: the role of MGMT in protection against the genotoxic effects of alkylating agents, Carcinogenesis 12 (1991) 1857–1867. [4] T. Coquerelle, J. Dosch, B. Kaina, Overexpression of N-methylpurine-DNA glycosylase in Chinese hamster ovary cells renders them more sensitive to the production of chromosomal aberrations by methylating agents – a case of imbalanced DNA repair, Mutat. Res. 336 (1995) 9–17. [5] R. Peto, R. Gray, P. Brantom, P. Grasso, Nitrosamine carcinogenesis in 5120 rodents: chronic administration of sixteen different concentrations of NDEA, NDMA, NPYR and NPIP in the water of 4440 inbred rats, with parallel studies on NDEA alone of the effect of age of starting (3, 6 or 20 weeks) and of species (rats, mice or hamsters), IARC Sci. Publ. (1984) 627–665. [6] H. Bartsch, R. Montesano, Relevance of nitrosamines to human cancer, Carcinogenesis 5 (1984) 1381–1393. [7] W. Fiddler, The occurrence and determination of N-nitroso compounds, Toxicol. Appl. Pharmacol. 31 (1975) 352–360. [8] M. Christmann, B. Kaina, O(6)-methylguanine-DNA methyltransferase (MGMT): impact on cancer risk in response to tobacco smoke, Mutat. Res. (2011). [9] L.J. Marnett, P.C. Burcham, Endogenous DNA adducts: potential and paradox, Chem. Res. Toxicol. 6 (1993) 771–785. [10] S.C. Naumann, W.P. Roos, E. Jost, C. Belohlavek, V. Lennerz, C.W. Schmidt, M. Christmann, B. Kaina, Temozolomide- and fotemustine-induced apoptosis in human malignant melanoma cells: response related to MGMT, MMR, DSBs, and p53, Br. J. Cancer 100 (2009) 322–333. [11] W.P. Roos, L.F. Batista, S.C. Naumann, W. Wick, M. Weller, C.F. Menck, B. Kaina, Apoptosis in malignant glioma cells triggered by the temozolomideinduced DNA lesion O6-methylguanine, Oncogene 26 (2007) 186–197. [12] D.T. Beranek, Distribution of methyl and ethyl adducts following alkylation with monofunctional alkylating agents, Mutat. Res. 231 (1990) 11–30. [13] D.T. Beranek, C.C. Weis, D.H. Swenson, A comprehensive quantitative analysis of methylated and ethylated DNA using high pressure liquid chromatography, Carcinogenesis 1 (1980) 595–606. [14] K. Becker, J. Dosch, C.M. Gregel, B.A. Martin, B. Kaina, Targeted expression of human O(6)-methylguanine-DNA methyltransferase (MGMT) in transgenic mice protects against tumor initiation in two-stage skin carcinogenesis, Cancer Res. 56 (1996) 3244–3249. [15] S.L. Gerson, MGMT: its role in cancer aetiology and cancer therapeutics, Nat. Rev. Cancer 4 (2004) 296–307. [16] K. Becker, C.M. Gregel, B. Kaina, The DNA repair protein O6-methylguanineDNA methyltransferase protects against skin tumor formation induced by antineoplastic chloroethylnitrosourea, Cancer Res. 57 (1997) 3335–3338. [17] R.S. Day 3rd, C.H. Ziolkowski, D.A. Scudiero, S.A. Meyer, A.S. Lubiniecki, A.J. Girardi, S.M. Galloway, G.D. Bynum, Defective repair of alkylated DNA by human tumour and SV40-transformed human cell strains, Nature 288 (1980) 724–727. [18] P. Karran, M. Bignami, DNA damage tolerance, mismatch repair and genome instability, Bioessays 16 (1994) 833–839. [19] A.E. Pegg, M.E. Dolan, R.C. Moschel, Structure, function, and inhibition of O6alkylguanine-DNA alkyltransferase, Prog. Nucl. Acid Res. Mol. Biol. 51 (1995) 167–223. [20] W. Meikrantz, M.A. Bergom, A. Memisoglu, L. Samson, O6-alkylguanine DNA lesions trigger apoptosis, Carcinogenesis 19 (1998) 369–372. [21] M. Christmann, B. Verbeek, W.P. Roos, B. Kaina, O(6)-Methylguanine-DNA methyltransferase (MGMT) in normal tissues and tumors: enzyme activity, promoter methylation and immunohistochemistry, Biochim. Biophys. Acta (2011). [22] J.L. Villano, T.E. Seery, L.R. Bressler, Temozolomide in malignant gliomas: current use and future targets, Cancer Chemother. Pharmacol. 64 (2009) 647– 655.

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[23] A.M. Eggermont, J.M. Kirkwood, Re-evaluating the role of dacarbazine in metastatic melanoma: what have we learned in 30 years?, Eur J. Cancer 40 (2004) 1825–1836. [24] R. Pepponi, G. Marra, M.P. Fuggetta, S. Falcinelli, E. Pagani, E. Bonmassar, J. Jiricny, S. D’Atri, The effect of O6-alkylguanine-DNA alkyltransferase and mismatch repair activities on the sensitivity of human melanoma cells to temozolomide, 1,3-bis(2-chloroethyl)1-nitrosourea, and cisplatin, J. Pharmacol. Exp. Ther. 304 (2003) 661–668. [25] M.J. Hickman, L.D. Samson, Role of DNA mismatch repair and p53 in signaling induction of apoptosis by alkylating agents, Proc. Natl. Acad. Sci. USA 96 (1999) 10764–10769. [26] W. Roos, M. Baumgartner, B. Kaina, Apoptosis triggered by DNA damage O6methylguanine in human lymphocytes requires DNA replication and is mediated by p53 and Fas/CD95/Apo-1, Oncogene 23 (2004) 359–367. [27] L. Stojic, P. Cejka, J. Jiricny, High doses of SN1 type methylating agents activate DNA damage signaling cascades that are largely independent of mismatch repair, Cell Cycle 4 (2005) 473–477. [28] J.S. Eadie, M. Conrad, D. Toorchen, M.D. Topal, Mechanism of mutagenesis by O6-methylguanine, Nature 308 (1984) 201–203. [29] D.R. Duckett, J.T. Drummond, A.I. Murchie, J.T. Reardon, A. Sancar, D.M. Lilley, P. Modrich, Human MutSalpha recognizes damaged DNA base pairs containing O6-methylguanine, O4-methylthymine, or the cisplatin-d(GpG) adduct, Proc. Natl. Acad. Sci. USA 93 (1996) 6443–6447. [30] G.M. Li, P. Modrich, Restoration of mismatch repair to nuclear extracts of H6 colorectal tumor cells by a heterodimer of human MutL homologs, Proc. Natl. Acad. Sci. USA 92 (1995) 1950–1954. [31] J. Genschel, L.R. Bazemore, P. Modrich, Human exonuclease I is required for 50 and 30 mismatch repair, J. Biol. Chem. 277 (2002) 13302–13311. [32] N. Mojas, M. Lopes, J. Jiricny, Mismatch repair-dependent processing of methylation damage gives rise to persistent single-stranded gaps in newly replicated DNA, Genes Dev. 21 (2007) 3342–3355. [33] P. Karran, C. Stephenson, Mismatch binding proteins and tolerance to alkylating agents in human cells, Mutat. Res. 236 (1990) 269–275. [34] K. Ochs, B. Kaina, Apoptosis induced by DNA damage O6-methylguanine is Bcl-2 and caspase-9/3 regulated and Fas/caspase-8 independent, Cancer Res. 60 (2000) 5815–5824. [35] J. Dosch, M. Christmann, B. Kaina, Mismatch G-T binding activity and MSH2 expression is quantitatively related to sensitivity of cells to methylating agents, Carcinogenesis 19 (1998) 567–573. [36] W.P. Roos, M. Christmann, S.T. Fraser, B. Kaina, Mouse embryonic stem cells are hypersensitive to apoptosis triggered by the DNA damage O(6)methylguanine due to high E2F1 regulated mismatch repair, Cell Death Differ. 14 (2007) 1422–1432. [37] W.P. Roos, T. Nikolova, S. Quiros, S.C. Naumann, O. Kiedron, M.Z. Zdzienicka, B. Kaina, Brca2/Xrcc2 dependent HR, but not NHEJ, is required for protection against O(6)-methylguanine triggered apoptosis, DSBs and chromosomal aberrations by a process leading to SCEs, DNA Repair (Amst) 8 (2009) 72–86. [38] S. Quiros, W.P. Roos, B. Kaina, Processing of O6-methylguanine into DNA double-strand breaks requires two rounds of replication whereas apoptosis is also induced in subsequent cell cycles, Cell Cycle 9 (2010) 168–178. [39] T.R. O’Connor, Purification and characterization of human 3-methyladenineDNA glycosylase, Nucl. Acids Res. 21 (1993) 5561–5569. [40] D. Chakravarti, G.C. Ibeanu, K. Tano, S. Mitra, Cloning and expression in Escherichia coli of a human cDNA encoding the DNA repair protein Nmethylpurine-DNA glycosylase, J. Biol. Chem. 266 (1991) 15710–15715. [41] R.H. Elder, J.G. Jansen, R.J. Weeks, M.A. Willington, B. Deans, A.J. Watson, K.J. Mynett, J.A. Bailey, D.P. Cooper, J.A. Rafferty, M.C. Heeran, S.W. Wijnhoven, A.A. van Zeeland, G.P. Margison, Alkylpurine-DNA-N-glycosylase knockout mice show increased susceptibility to induction of mutations by methyl methanesulfonate, Mol. Cell Biol. 18 (1998) 5828–5837. [42] D.M. Wilson, D. Barsky, The major human abasic endonuclease: formation, consequences and repair of abasic lesions in DNA, Mutat. Res. 485 (2001) 283–307. [43] R.W. Sobol, S.H. Wilson, Mammalian DNA beta-polymerase in base excision repair of alkylation damage, Prog. Nucl. Acid Res. Mol. Biol. 68 (2001) 57–74. [44] R. Prasad, W.A. Beard, P.R. Strauss, S.H. Wilson, Human DNA polymerase beta deoxyribose phosphate lyase. Substrate specificity and catalytic mechanism, J. Biol. Chem. 273 (1998) 15263–15270. [45] M. Stucki, B. Pascucci, E. Parlanti, P. Fortini, S.H. Wilson, U. Hubscher, E. Dogliotti, Mammalian base excision repair by DNA polymerases delta and epsilon, Oncogene 17 (1998) 835–843. [46] A. Klungland, T. Lindahl, Second pathway for completion of human DNA base excision-repair: reconstitution with purified proteins and requirement for DNaseIV (FEN1), EMBO J. 16 (1997) 3341–3348. [47] G. Fritz, S. Grösch, M. Tomicic, B. Kaina, APE/Ref-1 and the mammalian response to genotoxic stress, Toxicology 193 (2003) 67–78. [48] T. Izumi, D.B. Brown, C.V. Naidu, K.K. Bhakat, M.A. Macinnes, H. Saito, D.J. Chen, S. Mitra, Two essential but distinct functions of the mammalian abasic endonuclease, Proc. Natl. Acad. Sci USA 102 (2005) 5739–5743. [49] S. Xanthoudakis, G. Miao, F. Wang, Y.-C.E. Pan, T. Curran, Redox activation of Fos-Jun DNA binding activity is mediated by a DNA repair enzyme, EMBO J. 11 (1992) 3323–3335. [50] R.W. Sobol, J.K. Horton, R. Kühn, H. Gu, R.K. Singhai, R. Prasad, K. Rajewski, S.H. Wilson, Requirement of mammalian DNA polymerase-ß in base-excision repair, Nature 379 (1996) 183–186.

[51] K. Ochs, R.W. Sobol, S.H. Wilson, B. Kaina, Cells deficient in DNA polymerase beta are hypersensitive to alkylating agent-induced apoptosis and chromosomal breakage, Cancer Res. 59 (1999) 1544–1551. [52] M. Briegert, B. Kaina, Human monocytes, but not dendritic cells derived from them, are defective in base excision repair and hypersensitive to methylating agents, Cancer Res. 67 (2007) 26–31. [53] M. Rinne, D. Caldwell, M.R. Kelley, Transient adenoviral N-methylpurine DNA glycosylase overexpression imparts chemotherapeutic sensitivity to human breast cancer cells, Mol. Cancer Ther. 3 (2004) 955–967. [54] K. Ochs, J. Lips, S. Profittlich, B. Kaina, Deficiency in DNA polymerase ß provokes replication-dependent apoptosis via DNA breakage, Bcl-2 decline and caspase-3/9 activation, Cancer Res. 62 (2002) 1524–1530. [55] T. Nikolova, M. Ensminger, M. Lobrich, B. Kaina, Homologous recombination protects mammalian cells from replication-associated DNA double-strand breaks arising in response to methyl methanesulfonate, DNA Repair (Amst) 9 (2010) 1050–1063. [56] J. Cadet, T. Delatour, T. Douki, D. Gasparutto, J.P. Pouget, J.L. Ravanat, S. Sauvaigo, Hydroxyl radicals and DNA base damage, Mutat. Res. 424 (1999) 9– 21. [57] D.T. Goodhead, Initial events in the cellular effects of ionizing radiations: clustered damage in DNA, Int. J. Radiat. Biol. 65 (1994) 7–17. [58] H. Nikjoo, S. Uehara, W.E. Wilson, M. Hoshi, D.T. Goodhead, Track structure in radiation biology: theory and applications, Int. J. Radiat. Biol. 73 (1998) 355– 364. [59] K. Rothkamm, M. Lobrich, Evidence for a lack of DNA double-strand break repair in human cells exposed to very low x-ray doses, Proc. Natl. Acad. Sci. USA 100 (2003) 5057–5062. [60] K. Rothkamm, I. Kruger, L.H. Thompson, M. Lobrich, Pathways of DNA doublestrand break repair during the mammalian cell cycle, Mol. Cell Biol. 23 (2003) 5706–5715. [61] G.C. Smith, S.P. Jackson, The DNA-dependent protein kinase, Genes Dev. 13 (1999) 916–934. [62] G.G. Steel, The case against apoptosis, Acta Oncol. 40 (2001) 968–975. [63] J. Evans, M. Maccabee, Z. Hatahet, J. Courcelle, R. Bockrath, H. Ide, S. Wallace, Thymine ring saturation and fragmentation products: lesion bypass, misinsertion and implications for mutagenesis, Mutat. Res. 299 (1993) 147–156. [64] T.P. Hilbert, W. Chaung, R.J. Boorstein, R.P. Cunningham, G.W. Teebor, Cloning and expression of the cDNA encoding the human homologue of the DNA repair enzyme, Escherichia coli endonuclease III, J. Biol. Chem. 272 (1997) 6733–6740. [65] M. Takao, S. Kanno, K. Kobayashi, Q.M. Zhang, S. Yonei, G.T. van der Horst, A. Yasui, A back-up glycosylase in Nth1 knock-out mice is a functional Nei (endonuclease VIII) homologue, J. Biol. Chem. 277 (2002) 42205–42213. [66] T.A. Rosenquist, E. Zaika, A.S. Fernandes, D.O. Zharkov, H. Miller, A.P. Grollman, The novel DNA glycosylase, NEIL1, protects mammalian cells from radiation-mediated cell death, DNA Repair (Amst) 2 (2003) 581–591. [67] T.A. Rosenquist, D.O. Zharkov, A.P. Grollman, Cloning and characterization of a mammalian 8-oxoguanine DNA glycosylase, Proc. Natl. Acad. Sci. USA 94 (1997) 7429–7434. [68] A. Mazurek, M. Berardini, R. Fishel, Activation of human MutS homologs by 8oxo-guanine DNA damage, J. Biol. Chem. 277 (2002) 8260–8266. [69] J. Lips, B. Kaina, DNA double-strand breaks trigger apoptosis in p53-deficient fibroblasts, Carcinogenesis 22 (2001) 579–585. [70] S. Ishida, J. Lee, D.J. Thiele, I. Herskowitz, Uptake of the anticancer drug cisplatin mediated by the copper transporter Ctr1 in yeast and mammals, Proc. Natl. Acad. Sci. USA 99 (2002) 14298–14302. [71] G. Ciarimboli, T. Ludwig, D. Lang, H. Pavenstadt, H. Koepsell, H.J. Piechota, J. Haier, U. Jaehde, J. Zisowsky, E. Schlatter, Cisplatin nephrotoxicity is critically mediated via the human organic cation transporter 2, Am. J. Pathol. 167 (2005) 1477–1484. [72] R. Safaei, S.B. Howell, Copper transporters regulate the cellular pharmacology and sensitivity to Pt drugs, Crit. Rev. Oncol. Hematol. 53 (2005) 13–23. [73] C. Meijer, N.H. Mulder, H. Timmer-Bosscha, W.J. Sluiter, G.J. Meersma, E.G. de Vries, Relationship of cellular glutathione to the cytotoxicity and resistance of seven platinum compounds, Cancer Res. 52 (1992) 6885–6889. [74] P. Surowiak, V. Materna, A. Maciejczyk, M. Pudelko, E. Markwitz, M. Spaczynski, M. Dietel, M. Zabel, H. Lage, Nuclear metallothionein expression correlates with cisplatin resistance of ovarian cancer cells and poor clinical outcome, Virchows Arch. 450 (2007) 279–285. [75] E.R. Jamieson, S.J. Lippard, Structure, recognition, and processing of CisplatinDNA adducts, Chem. Rev. 99 (1999) 2467–2498. [76] A.M. Fichtinger-Schepman, J.L. van der Veer, J.H. den Hartog, P.H. Lohman, J. Reedijk, Adducts of the antitumor drug cis-diamminedichloroplatinum(II) with DNA: formation, identification, and quantitation, Biochemistry 24 (1985) 707–713. [77] A.M. Fichtinger-Schepman, A.T. van Oosterom, P.H. Lohman, F. Berends, cisDiamminedichloroplatinum(II)-induced DNA adducts in peripheral leukocytes from seven cancer patients: quantitative immunochemical detection of the adduct induction and removal after a single dose of cisdiamminedichloroplatinum(II), Cancer Res. 47 (1987) 3000–3004. [78] G. Damia, L. Imperatori, M. Stefanini, M. D’Incalci, Sensitivity of CHO mutant cell lines with specific defects in nucleotide excision repair to different anticancer agents, Int. J. Cancer 66 (1996) 779–783. [79] T.R. Dunkern, G. Fritz, B. Kaina, Cisplatin-induced apoptosis in 43-3B and 27-1 cells defective in nucleotide excision repair, Mutat. Res. 486 (2001) 249–258.

Please cite this article in press as: W.P. Roos, B. Kaina, DNA damage-induced apoptosis: From specific DNA lesions to the DNA damage response and apoptosis, Cancer Lett. (2012), doi:10.1016/j.canlet.2012.01.007

W.P. Roos, B. Kaina / Cancer Letters xxx (2012) xxx–xxx [80] F. Larminat, V.A. Bohr, Role of the human ERCC-1 gene in gene-specific repair of cisplatin-induced DNA damage, Nucl. Acids Res. 22 (1994) 3005–3010. [81] S. Usanova, A. Piee-Staffa, U. Sied, J. Thomale, A. Schneider, B. Kaina, B. Koberle, Cisplatin sensitivity of testis tumour cells is due to deficiency in interstrand-crosslink repair and low ERCC1-XPF expression, Mol. Cancer 9 (2010) 248. [82] S. Caporali, S. Falcinelli, G. Starace, M.T. Russo, E. Bonmassar, J. Jiricny, S. D’Atri, DNA damage induced by temozolomide signals to both ATM and ATR: role of the mismatch repair system, Mol. Pharmacol. 66 (2004) 478–491. [83] K. Yoshida, T. Ozaki, K. Furuya, M. Nakanishi, H. Kikuchi, H. Yamamoto, S. Ono, T. Koda, K. Omura, A. Nakagawara, ATM-dependent nuclear accumulation of IKK-alpha plays an important role in the regulation of p73-mediated apoptosis in response to cisplatin, Oncogene 27 (2008) 1183–1188. [84] J.S. Myers, D. Cortez, Rapid activation of ATR by ionizing radiation requires ATM and Mre11, J. Biol. Chem. 281 (2006) 9346–9350. [85] N. Pabla, S. Huang, Q.S. Mi, R. Daniel, Z. Dong, ATR-Chk2 signaling in p53 activation and DNA damage response during cisplatin-induced apoptosis, J. Biol. Chem. 283 (2008) 6572–6583. [86] J.F. Haince, S. Kozlov, V.L. Dawson, T.M. Dawson, M.J. Hendzel, M.F. Lavin, G.G. Poirier, Ataxia telangiectasia mutated (ATM) signaling network is modulated by a novel poly(ADP-ribose)-dependent pathway in the early response to DNA-damaging agents, J. Biol. Chem. 282 (2007) 16441–16453. [87] J.F. Haince, D. McDonald, A. Rodrigue, U. Dery, J.Y. Masson, M.J. Hendzel, G.G. Poirier, PARP1-dependent kinetics of recruitment of MRE11 and NBS1 proteins to multiple DNA damage sites, J. Biol. Chem. 283 (2008) 1197–1208. [88] J.P. Carney, R.S. Maser, H. Olivares, E.M. Davis, M. Le Beau, J.R. Yates 3rd, L. Hays, W.F. Morgan, J.H. Petrini, The hMre11/hRad50 protein complex and Nijmegen breakage syndrome: linkage of double-strand break repair to the cellular DNA damage response, Cell 93 (1998) 477–486. [89] C.J. Bakkenist, M.B. Kastan, DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation, Nature 421 (2003) 499–506. [90] S.V. Kozlov, M.E. Graham, C. Peng, P. Chen, P.J. Robinson, M.F. Lavin, Involvement of novel autophosphorylation sites in ATM activation, Embo J. 25 (2006) 3504–3514. [91] M. Pellegrini, A. Celeste, S. Difilippantonio, R. Guo, W. Wang, L. Feigenbaum, A. Nussenzweig, Autophosphorylation at serine 1987 is dispensable for murine Atm activation in vivo, Nature 443 (2006) 222–225. [92] J.T. Powers, S. Hong, C.N. Mayhew, P.M. Rogers, E.S. Knudsen, D.G. Johnson, E2F1 uses the ATM signaling pathway to induce p53 and Chk2 phosphorylation and apoptosis, Mol. Cancer Res.. 2 (2004) 203–214. [93] T. Uziel, Y. Lerenthal, L. Moyal, Y. Andegeko, L. Mittelman, Y. Shiloh, Requirement of the MRN complex for ATM activation by DNA damage, Embo J. 22 (2003) 5612–5621. [94] J.H. Lee, T.T. Paull, ATM activation by DNA double-strand breaks through the Mre11-Rad50-Nbs1 complex, Science 308 (2005) 551–554. [95] R. Kitagawa, C.J. Bakkenist, P.J. McKinnon, M.B. Kastan, Phosphorylation of SMC1 is a critical downstream event in the ATM-NBS1-BRCA1 pathway, Genes Dev. 18 (2004) 1423–1438. [96] I.M. Ward, J. Chen, Histone H2AX is phosphorylated in an ATR-dependent manner in response to replicational stress, J. Biol. Chem. 276 (2001) 47759– 47762. [97] B. Wang, S. Matsuoka, P.B. Carpenter, S.J. Elledge, 53BP1, a mediator of the DNA damage checkpoint, Science 298 (2002) 1435–1438. [98] T.S. Byun, M. Pacek, M.C. Yee, J.C. Walter, K.A. Cimprich, Functional uncoupling of MCM helicase and DNA polymerase activities activates the ATR-dependent checkpoint, Genes Dev. 19 (2005) 1040–1052. [99] L. Zou, S.J. Elledge, Sensing DNA damage through ATRIP recognition of RPAssDNA complexes, Science 300 (2003) 1542–1548. [100] H.L. Ball, D. Cortez, ATRIP oligomerization is required for ATR-dependent checkpoint signaling, J. Biol. Chem. 280 (2005) 31390–31396. [101] E.R. Parrilla-Castellar, S.J. Arlander, L. Karnitz, Dial 9-1-1 for DNA damage: the Rad9-Hus1-Rad1 (9-1-1) clamp complex, DNA Repair (Amst) 3 (2004) 1009– 1014. [102] M. Makiniemi, T. Hillukkala, J. Tuusa, K. Reini, M. Vaara, D. Huang, H. Pospiech, I. Majuri, T. Westerling, T.P. Makela, J.E. Syvaoja, BRCT domaincontaining protein TopBP1 functions in DNA replication and damage response, J. Biol. Chem. 276 (2001) 30399–30406. [103] J. Lee, A. Kumagai, W.G. Dunphy, The Rad9-Hus1-Rad1 checkpoint clamp regulates interaction of TopBP1 with ATR, J. Biol. Chem. 282 (2007) 28036– 28044. [104] A. Kumagai, J. Lee, H.Y. Yoo, W.G. Dunphy, TopBP1 activates the ATR–ATRIP complex, Cell 124 (2006) 943–955. [105] M. Cuadrado, B. Martinez-Pastor, M. Murga, L.I. Toledo, P. Gutierrez-Martinez, E. Lopez, O. Fernandez-Capetillo, ATM regulates ATR chromatin loading in response to DNA double-strand breaks, J. Exp. Med. 203 (2006) 297–303. [106] H.Y. Yoo, A. Kumagai, A. Shevchenko, A. Shevchenko, W.G. Dunphy, Ataxiatelangiectasia mutated (ATM)-dependent activation of ATR occurs through phosphorylation of TopBP1 by ATM, J. Biol. Chem. 282 (2007) 17501–17506. [107] T. Stiff, S.A. Walker, K. Cerosaletti, A.A. Goodarzi, E. Petermann, P. Concannon, M. O’Driscoll, P.A. Jeggo, ATR-dependent phosphorylation and activation of ATM in response to UV treatment or replication fork stalling, Embo J. 25 (2006) 5775–5782. [108] E.P. Rogakou, D.R. Pilch, A.H. Orr, V.S. Ivanova, W.M. Bonner, DNA doublestranded breaks induce histone H2AX phosphorylation on serine 139, J. Biol. Chem. 273 (1998) 5858–5868.

11

[109] S. Nakada, G.I. Chen, A.C. Gingras, D. Durocher, PP4 is a gamma H2AX phosphatase required for recovery from the DNA damage checkpoint, EMBO Rep. 9 (2008) 1019–1026. [110] Q.E. Wang, K. Milum, C. Han, Y.W. Huang, G. Wani, J. Thomale, A.A. Wani, Differential contributory roles of nucleotide excision and homologous recombination repair for enhancing cisplatin sensitivity in human ovarian cancer cells, Mol. Cancer 10 (2011) 24. [111] J. San Filippo, P. Sung, H. Klein, Mechanism of eukaryotic homologous recombination, Annu. Rev. Biochem. 77 (2008) 229–257. [112] S. Wirtz, G. Nagel, L. Eshkind, M.F. Neurath, L.D. Samson, B. Kaina, Both base excision repair and O6-methylguanine-DNA methyltransferase protect against methylation-induced colon carcinogenesis, Carcinogenesis 31 (2010) 2111–2117. [113] W.P. Roos, B. Kaina, DNA damage-induced cell death by apoptosis, Trends Mol. Med. 12 (2006) 440–450. [114] S. Matsuoka, G. Rotman, A. Ogawa, Y. Shiloh, K. Tamai, S.J. Elledge, Ataxia telangiectasia-mutated phosphorylates Chk2 in vivo and in vitro, Proc. Natl. Acad. Sci. USA 97 (2000) 10389–10394. [115] Z. Guo, A. Kumagai, S.X. Wang, W.G. Dunphy, Requirement for Atr in phosphorylation of Chk1 and cell cycle regulation in response to DNA replication blocks and UV-damaged DNA in Xenopus egg extracts, Genes Dev. 14 (2000) 2745–2756. [116] S.Y. Shieh, J. Ahn, K. Tamai, Y. Taya, C. Prives, The human homologs of checkpoint kinases Chk1 and Cds1 (Chk2) phosphorylate p53 at multiple DNA damage-inducible sites, Genes Dev. 14 (2000) 289–300. [117] N.H. Chehab, A. Malikzay, M. Appel, T.D. Halazonetis, Chk2/hCds1 functions as a DNA damage checkpoint in G(1) by stabilizing p53, Genes Dev. 14 (2000) 278–288. [118] C.E. Canman, D.S. Lim, K.A. Cimprich, Y. Taya, K. Tamai, K. Sakaguchi, E. Appella, M.B. Kastan, J.D. Siliciano, Activation of the ATM kinase by ionizing radiation and phosphorylation of p53, Science 281 (1998) 1677–1679. [119] M. Christmann, G. Fritz, B. Kaina, Induction of DNA repair genes in mammalian cells in response to genotoxic stress, in: D. Lankenau (Ed.), Genome Dynamics and Stability, vol. 1, Springer Verlag, Berlin-Heidelberg, 2007, pp. 383–398. [120] W.P. Roos, L.F. Batista, S.C. Naumann, W. Wick, M. Weller, C.F. Menck, B. Kaina, Apoptosis in malignant glioma cells triggered by the temozolomideinduced DNA lesion O(6)-methylguanine, Oncogene 26 (2007) 186–197. [121] E. Oda, R. Ohki, H. Murasawa, J. Nemoto, T. Shibue, T. Yamashita, T. Tokino, T. Taniguchi, N. Tanaka, Noxa, a BH3-only member of the Bcl-2 family and candidate mediator of p53-induced apoptosis, Science 288 (2000) 1053– 1058. [122] M. Gutekunst, M. Oren, A. Weilbacher, M.A. Dengler, C. Markwardt, J. Thomale, W.E. Aulitzky, H. van der Kuip, p53 hypersensitivity is the predominant mechanism of the unique responsiveness of testicular germ cell tumor (TGCT) cells to cisplatin, PLoS One 6 (2011) e19198. [123] Z. Li, P.R. Musich, Y. Zou, Differential DNA damage responses in p53 proficient and deficient cells: cisplatin-induced nuclear import of XPA is independent of ATR checkpoint in p53-deficient lung cancer cells, Int. J. Biochem. Mol. Biol. 2 (2011) 138–145. [124] L.F. Batista, W.P. Roos, M. Christmann, C.F. Menck, B. Kaina, Differential sensitivity of malignant glioma cells to methylating and chloroethylating anticancer drugs: p53 determines the switch by regulating xpc, ddb2, and DNA double-strand breaks, Cancer Res. 67 (2007) 11886–11895. [125] T.R. Dunkern, G. Fritz, B. Kaina, Ultraviolet light-induced DNA damage triggers apoptosis in nucleotide excision repair-deficient cells via Bcl-2 decline and caspase-3/-8 activation, Oncogene 20 (2001) 6026–6038. [126] E.C. Friedberg, DNA damage and repair, Nature 421 (2003) 436–440. [127] M. Ljungman, D.P. Lane, Transcription – guarding the genome by sensing DNA damage, Nat. Rev. Cancer 4 (2004) 727–737. [128] M. Ljungman, F. Zhang, Blockage of RNA polymerase as a possible trigger for u.v. light-induced apoptosis, Oncogene 13 (1996) 823–831. [129] B.C. McKay, C. Becerril, M. Ljungman, P53 plays a protective role against UVand cisplatin-induced apoptosis in transcription-coupled repair proficient fibroblasts, Oncogene 20 (2001) 6805–6808. [130] M. Goldstein, W.P. Roos, B. Kaina, Apoptotic death induced by the cyclophosphamide analogue mafosfamide in human lymphoblastoid cells: contribution of DNA replication, transcription inhibition and Chk/p53 signaling, Toxicol. Appl. Pharmacol. 229 (2008) 20–32. [131] M. Hamdi, J. Kool, P. Cornelissen-Steijger, F. Carlotti, H.E. Popeijus, C. van der Burgt, J.M. Janssen, A. Yasui, R.C. Hoeben, C. Terleth, L.H. Mullenders, H. van Dam, DNA damage in transcribed genes induces apoptosis via the JNK pathway and the JNK-phosphatase MKP-1, Oncogene 24 (2005) 7135–7144. [132] C.C. Franklin, A.S. Kraft, Conditional expression of the mitogen-activated protein kinase (MAPK) phosphatase MKP-1 preferentially inhibits p38 MAPK and stress-activated protein kinase in U937 cells, J. Biol. Chem. 272 (1997) 16917–16923. [133] D.D. Hirsch, P.J. Stork, Mitogen-activated protein kinase phosphatases inactivate stress-activated protein kinase pathways in vivo, J. Biol. Chem. 272 (1997) 4568–4575. [134] M. Christmann, M.T. Tomicic, D. Aasland, B. Kaina, A role for UV-light-induced c-Fos: Stimulation of nucleotide excision repair and protection against sustained JNK activation and apoptosis, Carcinogenesis 28 (2007) 183–190. [135] C. Tournier, P. Hess, D.D. Yang, J. Xu, T.K. Turner, A. Nimnual, D. Bar-Sagi, S.N. Jones, R.A. Flavell, R.J. Davis, Requirement of JNK for stress-induced activation of the cytochrome c-mediated death pathway, Science 288 (2000) 870–874.

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[136] M. Mancini, C.E. Machamer, S. Roy, D.W. Nicholson, N.A. Thornberry, L.A. Casciola-Rosen, A. Rosen, Caspase-2 is localized at the Golgi complex and cleaves golgin-160 during apoptosis, J. Cell. Biol. 149 (2000) 603–612. [137] B. Zhivotovsky, A. Samali, A. Gahm, S. Orrenius, Caspases: their intracellular localization and translocation during apoptosis, Cell Death Differ. 6 (1999) 644–651. [138] P. Lassus, X. Opitz-Araya, Y. Lazebnik, Requirement for caspase-2 in stressinduced apoptosis before mitochondrial permeabilization, Science 297 (2002) 1352–1354. [139] V. Hanoux, C. Pairault, M. Bakalska, R. Habert, G. Livera, Caspase-2 involvement during ionizing radiation-induced oocyte death in the mouse ovary, Cell Death Differ. 14 (2007) 671–681. [140] Y. Guo, S.M. Srinivasula, A. Druilhe, T. Fernandes-Alnemri, E.S. Alnemri, Caspase-2 induces apoptosis by releasing proapoptotic proteins from mitochondria, J. Biol. Chem. 277 (2002) 13430–13437. [141] J.D. Robertson, V. Gogvadze, A. Kropotov, H. Vakifahmetoglu, B. Zhivotovsky, S. Orrenius, Processed caspase-2 can induce mitochondria-mediated apoptosis independently of its enzymatic activity, EMBO Rep. 5 (2004) 643–648. [142] M. Enoksson, J.D. Robertson, V. Gogvadze, P. Bu, A. Kropotov, B. Zhivotovsky, S. Orrenius, Caspase-2 permeabilizes the outer mitochondrial membrane and disrupts the binding of cytochrome c to anionic phospholipids, J. Biol. Chem. 279 (2004) 49575–49578. [143] S. Sidi, T. Sanda, R.D. Kennedy, A.T. Hagen, C.A. Jette, R. Hoffmans, J. Pascual, S. Imamura, S. Kishi, J.F. Amatruda, J.P. Kanki, D.R. Green, A.A. D’Andrea, A.T. Look, Chk1 suppresses a caspase-2 apoptotic response to DNA damage that bypasses p53, Bcl-2, and caspase-3, Cell 133 (2008) 864–877. [144] J.A. Molitor, W.H. Walker, S. Doerre, D.W. Ballard, W.C. Greene, NF-kappa B: a family of inducible and differentially expressed enhancer-binding proteins in human T cells, Proc. Natl. Acad. Sci. USA 87 (1990) 10028–10032. [145] B.G. Darnay, V. Haridas, J. Ni, P.A. Moore, B.B. Aggarwal, Characterization of the intracellular domain of receptor activator of NF-kappaB (RANK). Interaction with tumor necrosis factor receptor-associated factors and activation of NF-kappab and c-Jun N-terminal kinase, J. Biol. Chem. 273 (1998) 20551–20555. [146] S. Schutze, T. Machleidt, M. Kronke, The role of diacylglycerol and ceramide in tumor necrosis factor and interleukin-1 signal transduction, J. Leukoc. Biol. 56 (1994) 533–541. [147] G. Zhang, S. Ghosh, Molecular mechanisms of NF-kappaB activation induced by bacterial lipopolysaccharide through Toll-like receptors, J. Endotoxin Res. 6 (2000) 453–457. [148] M. Karin, Y. Ben-Neriah, Phosphorylation meets ubiquitination: the control of NF-[kappa]B activity, Annu. Rev. Immunol. 18 (2000) 621–663. [149] T.T. Huang, S.M. Wuerzberger-Davis, Z.H. Wu, S. Miyamoto, Sequential modification of NEMO/IKKgamma by SUMO-1 and ubiquitin mediates NFkappaB activation by genotoxic stress, Cell 115 (2003) 565–576. [150] K. Brzoska, I. Szumiel, Signalling loops and linear pathways: NF-kappaB activation in response to genotoxic stress, Mutagenesis 24 (2009) 1–8. [151] H.H. Lee, H. Dadgostar, Q. Cheng, J. Shu, G. Cheng, NF-kappaB-mediated upregulation of Bcl-x and Bfl-1/A1 is required for CD40 survival signaling in B lymphocytes, Proc. Natl. Acad. Sci. USA 96 (1999) 9136–9141. [152] Z.L. Chu, T.A. McKinsey, L. Liu, J.J. Gentry, M.H. Malim, D.W. Ballard, Suppression of tumor necrosis factor-induced cell death by inhibitor of apoptosis c-IAP2 is under NF-kappaB control, Proc. Natl. Acad. Sci. USA 94 (1997) 10057–10062. [153] C.Y. Wang, M.W. Mayo, R.G. Korneluk, D.V. Goeddel, A.S. Baldwin Jr., NFkappaB antiapoptosis: induction of TRAF1 and TRAF2 and c-IAP1 and c-IAP2 to suppress caspase-8 activation, Science 281 (1998) 1680–1683. [154] H. Zhang, S.W. Cowan-Jacob, M. Simonen, W. Greenhalf, J. Heim, B. Meyhack, Structural basis of BFL-1 for its interaction with BAX and its anti-apoptotic action in mammalian and yeast cells, J. Biol. Chem. 275 (2000) 11092–11099. [155] A.B. Werner, E. de Vries, S.W. Tait, I. Bontjer, J. Borst, Bcl-2 family member Bfl1/A1 sequesters truncated bid to inhibit is collaboration with pro-apoptotic Bak or Bax, J. Biol. Chem. 277 (2002) 22781–22788. [156] E. Yang, J. Zha, J. Jockel, L.H. Boise, C.B. Thompson, S.J. Korsmeyer, Bad, a heterodimeric partner for Bcl-XL and Bcl-2, displaces Bax and promotes cell death, Cell 80 (1995) 285–291. [157] A. Zhou, S. Scoggin, R.B. Gaynor, N.S. Williams, Identification of NF-kappa Bregulated genes induced by TNFalpha utilizing expression profiling and RNA interference, Oncogene 22 (2003) 2054–2064.

[158] A. Brozovic, G. Fritz, M. Christmann, J. Zisowsky, U. Jaehde, M. Osmak, B. Kaina, Long-term activation of SAPK/JNK, p38 kinase and fas-L expression by cisplatin is attenuated in human carcinoma cells that acquired drug resistance, Int. J. Cancer 112 (2004) 974–985. [159] S. Karl, Y. Pritschow, M. Volcic, S. Hacker, B. Baumann, L. Wiesmuller, K.M. Debatin, S. Fulda, Identification of a novel pro-apopotic function of NF-kappaB in the DNA damage response, J. Cell Mol. Med. 13 (2009) 4239– 4256. [160] R. Yao, G.M. Cooper, Requirement for phosphatidylinositol-3 kinase in the prevention of apoptosis by nerve growth factor, Science 267 (1995) 2003– 2006. [161] H.G. Wendel, E. De Stanchina, J.S. Fridman, A. Malina, S. Ray, S. Kogan, C. Cordon-Cardo, J. Pelletier, S.W. Lowe, Survival signalling by Akt and eIF4E in oncogenesis and cancer therapy, Nature 428 (2004) 332–337. [162] S. Bao, G. Ouyang, X. Bai, Z. Huang, C. Ma, M. Liu, R. Shao, R.M. Anderson, J.N. Rich, X.F. Wang, Periostin potently promotes metastatic growth of colon cancer by augmenting cell survival via the Akt/PKB pathway, Cancer Cell 5 (2004) 329–339. [163] Q. Shi, S. Bao, J.A. Maxwell, E.D. Reese, H.S. Friedman, D.D. Bigner, X.F. Wang, J.N. Rich, Secreted protein acidic, rich in cysteine (SPARC), mediates cellular survival of gliomas through AKT activation, J. Biol. Chem. 279 (2004) 52200– 52209. [164] D. Gupta, N.A. Syed, W.J. Roesler, R.L. Khandelwal, Effect of overexpression and nuclear translocation of constitutively active PKB-alpha on cellular survival and proliferation in HepG2 cells, J. Cell Biochem. 93 (2004) 513–525. [165] L. del Peso, M. Gonzalez-Garcia, C. Page, R. Herrera, G. Nunez, Interleukin-3induced phosphorylation of BAD through the protein kinase Akt, Science 278 (1997) 687–689. [166] S.R. Datta, H. Dudek, X. Tao, S. Masters, H. Fu, Y. Gotoh, M.E. Greenberg, Akt phosphorylation of BAD couples survival signals to the cell-intrinsic death machinery, Cell 91 (1997) 231–241. [167] J. Zha, H. Harada, E. Yang, J. Jockel, S.J. Korsmeyer, Serine phosphorylation of death agonist BAD in response to survival factor results in binding to 14-3-3 not BCL-X(L), Cell 87 (1996) 619–628. [168] M.H. Cardone, N. Roy, H.R. Stennicke, G.S. Salvesen, T.F. Franke, E. Stanbridge, S. Frisch, J.C. Reed, Regulation of cell death protease caspase-9 by phosphorylation, Science 282 (1998) 1318–1321. [169] I. Budihardjo, H. Oliver, M. Lutter, X. Luo, X. Wang, Biochemical pathways of caspase activation during apoptosis, Annu. Rev. Cell Dev. Biol. 15 (1999) 269– 290. [170] A.H. Kim, G. Khursigara, X. Sun, T.F. Franke, M.V. Chao, Akt phosphorylates and negatively regulates apoptosis signal-regulating kinase 1, Mol. Cell Biol. 21 (2001) 893–901. [171] V.V. Sumbayev, I.M. Yasinska, Regulation of MAP kinase-dependent apoptotic pathway: implication of reactive oxygen and nitrogen species, Arch. Biochem. Biophys. 436 (2005) 406–412. [172] L.P. Kane, V.S. Shapiro, D. Stokoe, A. Weiss, Induction of NF-kappaB by the Akt/PKB kinase, Curr. Biol. 9 (1999) 601–604. [173] T.M. Gottlieb, J.F. Leal, R. Seger, Y. Taya, M. Oren, Cross-talk between Akt, p53 and Mdm2: possible implications for the regulation of apoptosis, Oncogene 21 (2002) 1299–1303. [174] L.D. Mayo, D.B. Donner, A phosphatidylinositol 3-kinase/Akt pathway promotes translocation of Mdm2 from the cytoplasm to the nucleus, Proc. Natl. Acad. Sci. USA 98 (2001) 11598–11603. [175] J.S. Kim, X. Xu, H. Li, D. Solomon, W.S. Lane, T. Jin, T. Waldman, Mechanistic analysis of a DNA damage-induced, PTEN-dependent size checkpoint in human cells, Mol. Cell Biol. 31 (2011) 2756–2771. [176] T. Schondorf, M. Becker, U.J. Gohring, B. Wappenschmidt, H. Kolhagen, C.M. Kurbacher, Interaction of cisplatin, paclitaxel and adriamycin with the tumor suppressor PTEN, Anticancer Drugs 12 (2001) 797–800. [177] M. Christmann, M.T. Tomicic, W.P. Roos, B. Kaina, Mechanisms of human DNA repair: an update, Toxicology 193 (2003) 3–34. [178] D.C. Altieri, Survivin and IAP proteins in cell-death mechanisms, Biochem. J. 430 (2010) 199–205. [179] S. Reichert, C. Rodel, J. Mirsch, P.N. Harter, M.T. Tomicic, M. Mittelbronn, B. Kaina, F. Rodel, Survivin inhibition and DNA double-strand break repair: a molecular mechanism to overcome radioresistance in glioblastoma, Radiother. Oncol. 101 (2011) 51–58.

Please cite this article in press as: W.P. Roos, B. Kaina, DNA damage-induced apoptosis: From specific DNA lesions to the DNA damage response and apoptosis, Cancer Lett. (2012), doi:10.1016/j.canlet.2012.01.007