Damage induced in the DNA after exposure of cells to ionizing radiation activates checkpoint pathways that inhibit progression of cells through the G1 and G2 phases and induce a transient delay in the progression through S phase. Checkpoints together with repair and apoptosis are integrated in a circuitry that determines the ultimate response of a cell to DNA damage. Checkpoint activation typically requires sensors and mediators of DNA damage, signal transducers and effectors. Here, we review the current state of knowledge regarding mechanisms of checkpoint activation and proteins involved in the different steps of the process. Emphasis is placed on the role of ATM and ATR, as well on CHK1 and CHK2 kinases in checkpoint response. The roles of downstream effectors, such as P53 and the CDC25 family of proteins, are also described, and connections between repair and checkpoint activation are attempted. The role of checkpoints in genomic stability and the potential of improving the treatment of cancer by DNA damage inducing agents through checkpoint abrogation are also briefly outlined.
It has been known for over 50 years that exposure of cells to ionizing radiation (IR) delays the normal progression through the cell cycle (Maity et al., 1994; Bernhard et al., 1995; Iliakis, 1997). While a delay in G2 is the most evident, significant delays also occur in G1, as well as throughout S phase. These cell cycle delays were initially interpreted as passive cellular responses resulting from the induction by IR of damage in the DNA. However, these early studies also provided circumstantial evidence that the delays actually reflect induction of cellular processes assisting the irradiated cell to cope with the induced damage by somehow facilitating repair (Walters et al., 1974; Tobey, 1975; Lücke-Huhle, 1982). Strong support for this alternative came with the observation that cells from ataxia telagiectasia (AT) patients have significantly reduced delays in their progression through the cell cycle and are, at the same time, sensitive to killing after exposure to IR (Houldsworth and Lavin, 1980; Painter and Young, 1980; Painter, 1981, 1986; Lavin and Schroeder, 1988).
Decisive in the further development of the field was the isolation, first in the budding and later in the fission yeast, of repair-proficient, radiation-sensitive mutants showing defects in radiation-induced cell cycle delays (Lee and Nurse, 1988; Hartwell and Weinert, 1989; Nurse, 1990; Norbury and Nurse, 1992; Hartwell and Kastan, 1994; Paulovich et al., 1997). Extensive genetic studies with these organisms uncovered a complex network of genes that cooperate to delay the normal progression through the cycle as soon as damage is registered in the genome. We now know that in these genome-surveillance mechanisms, delay in cell cycle progression is but one manifestation, captured by the term DNA damage checkpoint. The present state of knowledge allows the integration of delays induced after irradiation in the different phases of the cell cycle under the rubric of a single checkpoint activated by damage in the DNA (Elledge, 1996; Zhou and Elledge, 2000).
A parallel development that proved essential for the molecular understanding of the DNA damage checkpoint was the delineation of events underlying progression of cells through the cycle. It is now established that the cell cycle is controlled by interdependent regulatory transitions that bring the DNA to a state competent for duplication and division (Nurse, 1994; Elledge, 1996; Sherr, 1996; Stillman, 1996). These transitions are achieved by the activation of cyclin-dependent kinases (CDKs), the operation of proteolytic pathways and alterations in the state of chromatin. CDKs are essential components of the cell cycle machinery and are subject to strict regulation by independent mechanisms including association with cognate cyclins, phosphorylation at specific serine/threonine or tyrosine residues and the association with specific inhibitory proteins. Not surprisingly, CDKs turn out to also function as key targets of the checkpoint response. Progression through the cycle mediated by the above cell cycle machinery is ‘checked’ by surveillance mechanisms to ensure that cells will not progress to the next phase of the cell cycle before events of the preceding phase have been completed (Hartwell and Weinert, 1989; Hartwell and Kastan, 1994; Nurse, 1997; Paulovich et al., 1997). These surveillance mechanisms, known as cell cycle checkpoints, are conceptually distinct from the DNA damage checkpoint. However, as our understanding of the molecular mechanisms underlying the different checkpoints increases important similarities become apparent.
Here, we briefly outline the current state of knowledge of the DNA damage checkpoint. The progress made since the initial radiobiological observations is spectacular and represents the combined achievements of genetics and biochemistry in yeast, Xenopus and mammalian cell systems. The review mainly focuses on the DNA damage checkpoint as presently delineated in higher eukaryotes exposed to IR. Studies with other systems are only mentioned when the results have direct relevance to higher eukaryotes. Despite the recent dramatic accumulation of knowledge, key issues of checkpoint response remain unresolved and are discussed briefly.
Like many fields in modern biology, the field of checkpoints is nurtured by the conviction that detailed knowledge of the molecular mechanisms will enable a better understanding of the processes underlying genomic instability and cancer. Virtually every gene implicated in the DNA damage checkpoint contributes to genomic stability or is associated with genetic disorders or predisposition to cancer development. Moreover, because checkpoint abrogation seems to be an important feature of the cancer cell, considerable effort is presently placed towards the development of compounds that specifically abrogate the remaining checkpoint responses in cancer cells. It is hoped that these reagents will significantly improve our ability to combat cancer, alone or in combination with other treatment modalities such as IR.
General concepts and key players
The DNA damage checkpoint can be defined as a network of interacting pathways operating in concert to recognize damage in the DNA and elicit the response (Elledge, 1996; Zhou and Elledge, 2000; Nyberg et al., 2002). It shares characteristics of a signal transduction pathway, and the participating proteins can be formally divided into sensors, transducers and effectors (Figure 1). Sensor proteins recognize DNA damages, directly or indirectly, and function to signal the presence of these abnormalities and initiate the biochemical cascade. Transducers are typically protein kinases that relay and amplify the damage signal from the sensors by phosphorylating other kinases or downstream target proteins. Effector proteins include the ultimate downstream targets of the transducer protein kinases. Modification of effector proteins by upstream kinases, directly or indirectly, mediates the inhibition in cell cycle progression. The effector stage is where the DNA damage checkpoint, interphases with the cell cycle machinery. Proteins presently implicated as sensors and signal transducers are summarized in Table 1, and key players are described next. Effector proteins and detailed mechanisms of signal transmission and arrest in the different phases of the cell cycle are described in the subsequent sections.
It comes as a surprise that from the basic steps of the DNA damage checkpoint, the one involved in the initial sensing of abnormal DNA structures is the less well understood. Two proteins were initially considered good candidates as sensors owing to their ability to be directly activated by single- and double-strand breaks (DSBs), respectively, poly(ADP-ribose) polymerase (PARP) and DNA-dependent protein kinase (DNA-PK) (Lücke-Huhle, 1982; Anderson and Carter, 1996; Lees-Miller, 1996; Jeggo, 1997; D'Amours et al., 1999; Smith and Jackson, 1999; Smith, 2001; Ziegler and Oei, 2001). However, they do not seem to be required for the initiation of the global DNA damage response, although a possible role at the local level remains unexplored, and DNA-PK seems to somehow assist the recovery from the S phase delay (DiBiase et al., 2000; Guan et al., 2000).
There are several fundamental difficulties associated with the identification and characterization of sensor proteins. First, it is not known whether sensors recognize the initial DNA damage in all its diversity using a number of proteins or whether they recognize relatively common repair intermediates with the help of only a few proteins. Early work in bacteria and more recent work in eukaryotes points to single-stranded DNA (ssDNA), a common intermediate of several otherwise distinct DNA repair pathways, as the initiating signal (Friedberg et al., 1995). The prevailing view at present is that checkpoint signaling is initiated by ssDNA and DNA DSBs, but recent results also implicate changes in chromatin conformation (Zhou and Elledge, 2000; Nyberg et al., 2002; Bakkenist and Kastan, 2003). If ssDNA is a signal initiating the checkpoint response, repair pathways that generate large quantities of ssDNA should induce a stronger response than pathways generating low quantities of ssDNA, making the choice of the repair pathway important for the checkpoint response. For DNA DSBs, a lesion initiating a strong checkpoint response, this means that repair by nonhomologous end-joining (NHEJ) (Jeggo, 1998; Thompson and Schild, 2001, 2002) will only generate a weak checkpoint response owing to the fast removal of the break and the limited generation ssDNA, whereas homology-directed repair (Thompson and Schild, 2001, 2002) will generate a strong response as long stretches of ssDNA are known intermediates of this pathway (see below).
Despite difficulties, recent work in yeast has provided candidate-sensor protein complexes, and considerable effort is presently placed on characterizing the functions of homologs in higher eukaryotes. Two potentially cooperating complexes, RAD17–RFC and the 9-1-1, are presently in the center of interest. These protein complexes have functional homologs in DNA replication, and it is thought that they function similarly as part of the checkpoint response. The RAD17–RFC complex comprises RAD17 and the four small subunits of replication factor C (RFC; RFC2, RFC3, RFC4, RFC5) (Griffiths et al., 1995; Green et al., 2000; Lindsey-Boltz et al., 2001). RFC, which includes subunits RFC1–5, recognizes and binds single-strand/double-strand DNA junctions and functions to load on the DNA a homotrimeric protein with a clamp-like structure needed for increased processivity for DNA polymerases δ and ɛ (Stillman, 1994; Brush and Kelly, 1996). There is evidence that the RAD17–RFC complex functions as a loader of the 9-1-1 complex on damaged DNA (Lindsey-Boltz et al., 2001; Zou et al., 2002; Bermudez et al., 2003). The 9-1-1 complex is composed of the RAD9, HUS1 and RAD1 subunits that interact to form a PCNA-like structure that loads on the DNA like a sliding clamp similar to PCNA. It has also been speculated that the 9-1-1 complex functions to generate more ssDNA to enhance checkpoint-associated signaling (Lydall and Weinert, 1995; Parker et al., 1998; Volkmer and Karnitz, 1999; Xu et al., 2001). It will be interesting to see whether and how the RAD16–RFC and 9-1-1 complexes are involved in checkpoint signaling originating from sites of DNA DSBs. It will also be important to establish how, if at all, these protein complexes function within the known pathways of DNA DSB repair.
Other proteins such as the Mre11–RAD50–NBS1 (Petrini, 2000) complex and the BRCA1 protein (Scully and Livingston, 2000; Wang et al., 2000; Venkitaraman, 2001; Xu et al., 2001) have been implicated in the sensing of DNA DSBs. BRCA1 has been proposed to function as an adaptor of checkpoint initiation by localizing additional substrates for transducer kinase phosphorylation and by perhaps linking checkpoint arrest to DNA damage repair. Indeed, BRCA1 is known to interact with several proteins involved in DNA repair and to form a complex that may function in signaling (Wang et al., 2000). However, the existing evidence is not sufficient for determining whether what is experimentally seen reflects sensing or effector functions (Zhou and Elledge, 2000; Venkitaraman, 2001). It is likely that in the near future, more sensing proteins will surface or that known proteins will be recognized as sensors of the DNA damage checkpoint. This process will be greatly assisted by a better characterization of the aberrant DNA structure that is sensed by the checkpoint pathway.
A class of proteins termed mediators has also been implicated in the transduction of the DNA damage signal. The prototype of this class of proteins is the RAD9 gene of Saccharomyces cerevisiae. It contains two repeat domains originally found in the carboxyl terminus of BRCA1 and therefore termed BRCT domains. BRCT-repeat containing proteins have been identified in mammals, and some of them are thought to have functions similar to those of RAD9. Of these proteins, BRCA1 (Venkitaraman, 2001), TopBP1 (topoisomerase II binding protein 1) (Yamane et al., 2002), 53BP1 (P53 binding protein 1) (Schultz et al., 2000; Anderson et al., 2001) and the recently described MDC1 (mediator of DNA damage checkpoint protein 1) (Goldberg et al., 2003; Lou et al., 2003; Stewart et al., 2003) are the most likely candidates. All of these proteins have been implicated in checkpoint response after DNA damage and are thought to be involved in the recognition of the damage and the recruitment of additional proteins that facilitate downstream signaling and repair.
Two extensively studied kinases of the phosphoinositide 3-kinase (PI3K)-related family of proteins, ATM and ATR, are located immediately downstream of the damage sensors. ATM is the gene mutated in patients suffering from AT, a disease associated with immune deficiency, cerebellar degeneration and an increased predisposition to cancer (Rotman and Shiloh, 1998, 1999; Shiloh, 2001). At the cellular level, ATM mutation is associated with gross chromosomal rearrangements, radioresistant DNA synthesis, as well as a reduction in G1 and G2 arrest. ATM plays an important role in the DNA damage checkpoint by controlling the initial phosphorylation of several key proteins of the overall response such as P53, MDM2, BRCA1, CHK2, MDC1 and NBS1 (Giaccia and Kastan, 1998; Kastan and Lim, 2000). Phosphorylation of these proteins follows in AT cell-delayed kinetics (but is not absent) and is due to the activation of alternative signaling pathways. ATR (ATM and RAD3-related) shares a number of substrates with ATM, but its function is less well studied (Abraham, 2001). This is because ATR mutations have not been associated with human disease, and because ATR−/− mice die early in embryogenesis and ATR−/− cells are not viable (Cliby et al., 1998; Brown and Baltimore, 2000; Shiloh, 2001). This essential role of the protein in the life cycle of the cell and the organism compromises genetic studies.
It is not entirely clear how ATM and ATR are activated in response to the various stimuli and, in fact, a direct-sensor role is occasionally attributed to these proteins (Tibbetts et al., 1999; Abraham, 2001; Bao et al., 2001; Cortez et al., 2001; Durocher and Jackson, 2001; Zou et al., 2002; Bakkenist and Kastan, 2003). In line with this, there are reports for direct stimulation of ATM kinase activity by DNA damage in vivo (Banin et al., 1998; Canman et al., 1998; Khanna et al., 1998; Durocher and Jackson, 2001). On the other hand, direct stimulation by DNA in vitro remains controversial (Banin et al., 1998; Smith et al., 1999; Chan et al., 2000a), and recent results point to changes in chromatin conformation rather than DNA breakage per se as the signal activating ATM (Bakkenist and Kastan, 2003). A key question for this type of studies is whether ATM is aided by other proteins in its DNA binding, much the same way as Ku aids DNA binding and activation of DNA-PKcs. Interestingly, ATR has recently been shown to form a heterodimer with ATR interacting protein (ATRIP), which is required for checkpoint signaling (Edwards et al., 1999; Paciotti et al., 2000; Rouse and Jackson, 2000; Cortez et al., 2001; Wakayama et al., 2001; Wolkow and Enoch, 2002), although the mechanism remains elusive. It will be particularly interesting to see whether similarly functioning ATM partners also exist.
Despite similarities, important differences also distinguish ATM and ATR, the most prevalent probably being the kinetics of activation and the types of damage to which they respond best. Existing data support the hypothesis that ATM is the main determinant of the early checkpoint response induced by IR damage, whereas ATR responds later to processed IR-induced lesions, as well as to lesions induced by UV and by blocked replication forks (Zhou and Elledge, 2000; Nyberg et al., 2002).
The further propagation of signal generated by the DNA damage checkpoint relies to a considerable degree on the activation of the CHK1 and CHK2 kinases (Rhind and Russell, 2000; Bartek et al., 2001; Walworth, 2001). These structurally unrelated serine/threonine kinases share some overlapping substrate specificity. CHK2 is phosphorylated by ATM in response to ionizing radiation, and this phosphorylation is required for its activation (Matsuoka et al., 1998b, 2000; Melchionna et al., 2000). CHK1 is phosphorylated by ATR (Sanchez et al., 1997; Liu et al., 2000). Phosphospecific antibodies recognizing phosphoserine 345 of CHK1 reveal that this site is strongly phosphorylated in response to hydroxyurea and UV light and moderately in response to IR (Kim et al., 1999; Liu et al., 2000). Of note, mice lacking CHK1 die early in embryogenesis similarly to ATR−/− mice (Liu et al., 2000; Takai et al., 2000). As result of these observations, ATM is usually linked to CHK2 and ATR to CHK1.
In the following sections, we dissect the molecular characteristics of checkpoint responses in G1, S and G2 phases of the cell cycle. The outline specifies the roles of the above kinases in checkpoint signaling and indicates alternative strategies utilized by the cells as part of the overall response to DNA damage.
Molecular mechanisms of the G1-phase checkpoint
Accurate replication of the genome is paramount to genomic stability. Therefore, organisms have developed a complex network of regulatory processes to impose strict regulation upon DNA replication under both normal circumstances, as well as following DNA damage. Key regulatory steps occur in G1 phase and determine whether a cell will enter the cell cycle. Once the division cycle is initiated, surveillance mechanisms monitor the order and quality of events and halt progression if DNA damage is encountered (Hartwell and Weinert, 1989; Hartwell and Kastan, 1994; Nurse, 1994, 1997; Elledge, 1996; Paulovich et al., 1997; Zhou and Elledge, 2000).
Initiation of cell cycle progression is affected by the activation of two key regulatory kinases, CDK4 and CDK2, in association with D-type cyclins and cyclin E, respectively. Phosphorylation of target proteins by these kinases promotes entry into S phase, which requires inactivation of inhibitors such as the retinoblastoma protein (pRB) and the activation of S phase-promoting kinases such as CDC45 (Sherr, 1995, 1996; Stillman, 1996). A further contribution for this transition is provided by a pathway involving the Myc proto-oncogene, a transcription factor (Amati et al., 1992; Marcu et al., 1992; Atchley and Fitch, 1995; Eisenman and Cooper, 1995) acting by a variety of mechanisms including the transcriptional activation of cyclins D1, D2 and E of the CDC25A phosphatase, as well as of E2Fs (Galaktionov et al., 1996; Leone et al., 1997; Santoni-Rugiu et al., 2000; Bartek and Lukas, 2001a, 2001b; Seoane et al., 2002; Sheen and Dickson, 2002). The RB and the Myc pathways cooperate to increase abundance and activity of the CDK2-Cyclin E kinase, which is thought to initiate DNA replication by ultimately activating the CDC45 kinase. CDC45 is important for the firing of replication origins licensed by the Orc complex, CDC6 and the MCM family of proteins (Stillman, 1996; Dutta and Bell, 1997; DePamphilis, 1999; Tye, 1999; Donaldson and Blow, 2001; Bell and Dutta, 2002). Thus, cyclin E-CDK2 is positioned at the convergence point of two regulatory pathways contributing to the initiation of the cell cycle in nonirradiated cells. As we describe next, the same kinase is the target of two branches of the DNA damage-induced process that delays cell cycle progression in G1 (Bartek and Lukas, 2001a, 2001b).
Our current understanding of the molecular mechanism underlying DNA damage-induced delay in G1 is summarized in Figure 2. The checkpoint response in this phase of the cell cycle has two kinetically distinct components. The initial, acute phase operates by ‘locking’ CDK2-Cyclin E kinase in an inactive state through inhibition/destruction of the activating phosphatase CDC25A and is followed by a more delayed and sustained G1 arrest, also mediated by CDK2-Cyclin E inactivation through stabilization of the P53 tumor suppressor protein (Bao et al., 2001; Lukas et al., 2001). Of note, both components of the checkpoint response utilize the ATM/ATR and CHK1/CHK2 kinases (see above).
To ensure fast response, the initial acute component of the G1 delay utilizes ubiquitin/proteasome-mediated protein degradation rather than the relatively slower process of transcriptional activation and new protein synthesis employed by the P53 branch. Recent reports indicate a rapid decrease, independent of P53 status, of the abundance and activity of CDC25A in mammalian cells exposed to IR or UV light. The reduction in CDC25A abundance results from the ubiquitination and proteasome-mediated degradation of the protein, caused, in the case of IR-induced DNA damage, by phosphorylation at Ser123 by CHK2 (Mailand et al., 2000; Falck et al., 2001). A similar process involving phosphorylation by CHK1 is thought to be active after exposure to UV light (Mailand et al., 2000). There is evidence implicating ATM in the activation of CHK2, presumably through phosphorylation of the Thr68 residue of the kinase, and of CHK1 by ATR through phosphorylation of Ser317/345 (Costanzo et al., 2000; Zhou and Elledge, 2000; Abraham, 2001; Falck et al., 2001; Shiloh, 2001; Nyberg et al., 2002). Downstream, inactivation of CDC25A deprives the cell from a central activator of the CDK2-Cyclin E kinase, operating by removing inhibitory phosphates from Thr14 and Tyr15 of CDK2. This ‘locks’ the kinase in an inactive state and inhibits further downstream events required for the progression of cells into S phase. At least for UV, inactivation of CDC25A may also result in the maintenance of the inhibitory TYR17 phosphorylation on CDK4 that would additionally inhibit entry into S phase (Terada et al., 1995).
An analogous mechanism based on protein degradation in response to IR has also been reported for cyclin D1 (Agami and Bernards, 2000). Inhibition of CDK2 by this pathway is thought to reflect redistribution of the P21 CDK inhibitor (see below) from the CDK4-Cyclin D1 complex, where it serves as an assembly factor, to cyclin E-CDK2 complexes, which are inhibited by P21 (Sherr and Roberts, 1999; Agami and Bernards, 2000; Yu et al., 2001). This putative component of the checkpoint response is not shown in Figure 2 and would operate independently of ATM.
The central element of the second branch of the IR-induced G1 component of the DNA damage checkpoint is the stabilization of the P53 protein and the activation of its transcriptional activity. These effects lead to the transcription of a large number of genes, among them the P21Waf1/Cip1, an inhibitor of CDK4-Cyclin E kinase that mediates the arrest in G1 (Figure 2) (Sherr and Roberts, 1995; Dotto, 2000; Vogelstein et al., 2000; Zhou and Elledge, 2000; Lukas et al., 2001). It remains to be seen whether P21 is the only critical target downstream of P53 in this G1 response. The present state of knowledge assumes activation within minutes by DNA damage of ATM or ATR and phosphorylation by these kinases of P53 on Ser15. A concomitant phosphorylation of Ser20 by a similarly activated CHK2, and possibly CHK1, disrupts the normal interaction between P53 and MDM2 that targets in nonirradiated cells P53 for ubiquitination and proteasome-mediated degradation. This leads to the observed P53 increase in abundance after IR. Other modifications of P53 itself, as well as phosphorylation of MDM2 by ATM/ATR and CHK1/CHK2, reinforce the induced delay (Larner et al., 1999; Matsuoka et al., 2000; Bao et al., 2001). Furthermore, MDM2 is itself transcriptionally activated by P53 generating a negative feedback loop that keeps P53 in check, and dynamic changes in the subcellular localization of P53 and MDM2 provide additional levels of regulation for this complex response (Melchionna et al., 2000; Lukas et al., 2001).
The above dual response is carefully designed for inducing a delay in G1, starting from a single set of damage-responsive kinases and targeting predominantly a single kinase of the cell cycle engine. One of the functions of the G1 checkpoint may be to delay DNA replication (Larner et al., 1997, 1999) (see below), but significant contributions to DNA repair and genomic stability are also considered. Despite significant progress in understanding the checkpoint itself, the molecular interphase with genetic instability and DNA repair remain unknown. The fact that P53 is frequently mutated or deleted in human tumors suggests an important role of the G1 checkpoint in maintaining genomic integrity. Although it is widely accepted that cell cycle delays induced by checkpoint activation facilitate repair, there is no direct evidence that the G1 checkpoint promotes DNA DSB repair. Of the two pathways implicated in the repair of DNA DSBs, NHEJ is fast compared with the G1 delay and has not been linked to the checkpoint response, while homology-directed repair is thought not to occur efficiently in G1 cells. Alternatively, the correlation between the G1 checkpoint and genomic integrity might be via the elimination of DNA damage containing cells by apoptosis, and it is well known that P53 is involved in the regulation of this process (Figure 1) (Hickman et al., 2002). It will be important to put into perspective the different aspects of the G1 checkpoint with the repair of DNA DSBs, genomic stability and apoptosis.
Molecular mechanisms of the S phase checkpoint
The sophisticated mechanisms described above prevent a cell irradiated in G1 from entering S phase with a full load of DNA damage, but are of no benefit to a cell irradiated during S phase. Moreover, although the vast majority of cells in an adult organism are in G1 at any given time, damage registered during S can interfere with the functioning DNA replication machinery and lead to serious genomic abnormalities. Not surprisingly, therefore, cells have developed mechanisms that detect DNA damage during S phase and transiently halt the firing of replicons still waiting to be replicated (Painter, 1986; Lavin and Schroeder, 1988; Larner et al., 1997). First indications for a checkpoint in S phase (also termed intra-S phase checkpoint) came from the observation that exposure of cells to ionizing radiation causes a transient inhibition in the incorporation of radioactive precursors into nascent DNA (Painter, 1986; Lavin and Schroeder, 1988; Larner et al., 1997). Although the recognition that this effect reflects an active cellular response initiated by DNA damage is relatively recent (Lamb et al., 1989; Cleaver et al., 1990; Wang et al., 1995, 1999), early work allowed the detailed characterization of its salient cellular manifestations.
The dose–response curve describing the inhibition of DNA replication after exposure to IR is biphasic with a radiosensitive component reflecting inhibition in the firing of replicon clusters and a radioresistant component deriving from inhibition in chain elongation (Painter, 1986; Lavin and Schroeder, 1988; Larner et al., 1997). By its nature, and contrary to the delays in G1 or G2, which inhibit globally and for the most part persistently transition to a subsequent cell cycle phase, the S phase delay is short and can target only parts of the genome. It is active to a comparable degree throughout S phase (Larner et al., 1997) (G1 and G2 checkpoints arrest at a defined stage), and its length may be determined by the repair requirements imposed in this phase of the cell cycle, the distributed nature of its activity, the lability of stalled replicons and the availability of an additional checkpoint in the immediately following G2 phase. The radioresistance of wild-type S phase cells, particularly when compared with the exquisite radiosensitivity of AT cells, attests, on the other hand, to a particularly efficient and relevant checkpoint. The term radioresistant DNA synthesis, RDS, is frequently used to signify the absence or simply a reduction in this checkpoint.
The present state of knowledge regarding the S phase checkpoint is summarized in Figure 3. In contrast to the G1-phase checkpoint (Figure 2), a role for P53 or P21 could not be established for the S phase checkpoint (Lee et al., 1997; Guo et al., 1999). In fact, it is not known whether P53 is fully activated during S phase after exposure to IR. Accumulation of P53 after drug-induced inhibition of DNA replication was reported during S phase, but did not cause full transcriptional activation. It was reasoned that full-scale P53 transcriptional activation during S phase might cause unwanted apoptotic cell death in cooperation with the E2F-1 transcription factor and may therefore be suppressed (Gottifredi et al., 2001). Other reports indicate, however, a protective function for P53 during S phase that prevents aberrant entrance into G2/M (Taylor et al., 1999) and suggest that further investigations on the topic are warranted.
Upon exposure to IR, ATM is activated and phosphorylates several substrates that are candidate components of the S phase checkpoint including CHK2 (Matsuoka et al., 2000; Melchionna et al., 2000). A downstream target of CHK2 is CDC25A, whose phosphorylation leads to proteolytic degradation by the proteasome in a process similar to that described above for the G1 phase checkpoint (Falck et al., 2001; Kastan, 2001). Degradation of CDC25A will deprive the cells from an essential activator of CDK2, either in association with cyclin A or cyclin E, and will block replicon firing by inhibiting CDC45 and other cellular targets. Indeed, inhibition of CDK2 activity through CDC25A degradation leads to delay in S phase progression that correlates timely with the S phase checkpoint response. These observations when combined with the above outline of the G1 checkpoint suggest that the ATM–CHK2–CDC25A–CDC45 axis forms a rapid response system for the inhibition of various cell cycle-related processes in the cell, including DNA replication. Predictably, interference with the CHK2–CDC25A–CDK2 cascade at any of these steps downstream of ATM results in radioresistant DNA synthesis.
Recent work points to additional contributions for the IR-induced S phase checkpoint. Delayed inhibition of DNA replication in AT cells could be attributed to the activation of the ATR/CHK1 pathway supporting the notion that this branch of the checkpoint response is also activated in cells exposed to IR (Zhou et al., 2002). Furthermore, results with HeLa cells indicate radioresistant DNA synthesis (RDS) after disruption of CHK1 that correlates with the accumulation of nondegradable, hypophosphorylated CDC25A (Zhao et al., 2002). These new results in aggregate emphasize the importance of an ATR–CHK1–CDC25A axis for the S phase checkpoint and suggest Ser123 of CDC25A as one candidate target site for CHK1. In contrast to the ATM response that reaches a maximum about 30 min after irradiation, ATR-mediated inhibition is slower and reaches a maximum 2–4 h later. How the coordination of the two pathways is designed will be an interesting topic of examination in the future.
In addition to CHK1 and CHK2, other targets of ATM and possibly also ATR include BRCA1 (Cortez et al., 1999; Gatei et al., 2000a; Li et al., 2000) and NBS1 (Gatei et al., 2000b; Lim et al., 2000; Wu et al., 2000; Zhao et al., 2000), a component of the Mre11–NBS1–RAD50 complex (Petrini, 2000; Desai-Mehta et al., 2001). There is evidence that the ATM-mediated phosphorylation of NBS1 is required for the proper execution of the intra-S phase checkpoint, since mutating the targeted serine residues (Ser278, 343, 397) to alanine results in RDS. Recent reports implicate MDC1, a newly discovered BRCT-repeat-containing protein, in the S phase checkpoint (Goldberg et al., 2003; Lou et al., 2003; Stewart et al., 2003), although the mechanism of action remains unclear. Results support the function of MDC1 as a mediator sharing an intimate relation with H2AX and helping to recruit other repair or signaling proteins to the damage sites (Stewart et al., 2003). Other results are compatible with a function on a checkpoint pathway operating in parallel to the CDC25A pathway (Figure 3) downstream of RAD50–Mre11–NBS1 (Goldberg et al., 2003), and finally other results demonstrate a direct interaction between MDC1 and phosphorylated CHK2 and implicate the protein in the main pathway shown in Figure 3 (Lou et al., 2003).
In addition to the above pathways and interactions, several reports document interesting interplays between CHK2, BRCA1 and NBS1 (Wang, 2000; Wang et al., 2000; Lee and Chung, 2001), between the Mre11 complex and E2F-1 (Maser et al., 2001), and point to S phase checkpoint defects in BRCA1-deficient cells (Xu et al., 2001), and an important role for SMC1 protein (Kim et al., 2002; Yazdi et al., 2002). These observations are potentially very interesting, but it is not clear at present how they fit in the overall picture of the S phase checkpoint indicated in Figure 3. Evidence accumulates for parallel pathways that cooperate to inhibit DNA replication after exposure to IR (Falck et al., 2002), and it is relevant to recall recent studies suggesting a requirement for active DNA replication in S phase checkpoint activation (Lupardus et al., 2002; Stokes et al., 2002).
Molecular mechanisms of the G2 checkpoint
A temporary arrest of mammalian cell division is one of the first effects of radiation to be documented and investigated (Bernhard et al., 1995, 1999; McKenna, 1995; Iliakis, 1997). Early on, it could be established that IR-induced division arrest derives from a block in the progression of cells from the G2 into the M phase of the cell cycle. As with the delay during S phase, G2 arrest was initially considered a passive consequence of the presence of damaged DNA, but extensive investigations led to speculations for an active response with a role in DNA repair (Walters et al., 1974; Tobey, 1975; Lücke-Huhle, 1982; Hartwell and Weinert, 1989). Indeed, a long G2 delay on a DNA repair proficient background has been associated with radioresistance to killing (McKenna et al., 1991; Su and Little, 1993; Smeets et al., 1994).
The molecular characterization of the events causing the transition of a cell from G2 into M phase provided a solid foundation for the molecular delineation of the G2 checkpoint. Figure 4 summarizes important molecular components of the G2 checkpoint.
A key effector of the G2 checkpoint is the CDC2 (CDK1) kinase. Activation of this kinase, by association with cyclin B, subcellular translocation events and a series of phosphorylation and dephosphorylation events, is essential in initiating mitosis (Nurse, 1990; Norbury and Nurse, 1992; Morgan, 1995). Given the pivotal role of this kinase in the transition from G2 to M, its equally central role in the G2 checkpoint comes as no surprise. Indeed, activation and maintenance of the G2 arrest require targeting of multiple regulatory processes involved in the normal progression of the cell from G2 into M phase, including phosphatases promoting mitosis, kinases blocking CDC2 function, the abundance and translocation of cyclin B, and other proteins regulating these regulators (Zhou and Elledge, 2000; Smits and Medema, 2001; Nyberg et al., 2002).
An essential and precisely controlled event in the initiation of mitosis is the removal of inhibitory phosphorylations from CDC2 on TYR15 and Thr14, added earlier in the cycle by Wee1 and Myt1, respectively (Parker and Piwnica-Worms, 1992; Booher et al., 1997; Liu et al., 1997). Indeed, a CDC2 mutant that cannot be phosphorylated at these sites reduces G2 arrest (Jin et al., 1996). Several processes contributing to IR-induced G2 arrest inhibit the processes mediating the removal of these inhibitory phosphorylations. Not surprisingly therefore, central in the regulation of the G2 checkpoint is the inhibition of the CDC25C phosphatase (Draetta and Eckstein, 1997; Peng et al., 1997). Due to the central role of this phosphatase in the regulation of CDC2 activity, several pathways of regulation appear to converge at this site. Thus, activation of ATM leads to CHK2 activation through phosphorylation at Thr68, which in turn phosphorylates CDC25C at Ser216 to block its function (Matsuoka et al., 1998a; Zhou et al., 2000; Lopez-Girona et al., 2001). The inhibition is mediated by binding of the phosphorylated form of CDC25C to the 14-3-3 protein that is thought to render CDC25C catalytically less active and to cause its sequestration in the cytoplasm (Peng et al., 1997; Blasina et al., 1999; Furnari et al., 1999; Graves et al., 2000; Lopez-Girona et al., 2001). The relevance of cytoplasmic sequestration of CDC25C remains uncertain, however, as mammalian cells, in contrast to the fission yeast, sequester Cyclin B-CDC2 in the cytoplasm by active export until just before mitosis (see below), and even in the fission yeast forced nuclear localization of CDC25 does not override the G2 arrest (Lopez-Girona et al., 2001; O'Connell et al., 2002).
A parallel pathway is thought to operate through the activation of ATR, which causes the phosphorylation of CHK1 at Ser317/345 (Guo et al., 2000), which then negatively regulates CDC25C by phosphorylation at Ser216 (Peng et al., 1997; Sanchez et al., 1997). More recent results with HeLa cells point to CDC25A as an additional target of this pathway and, therefore, a determinant of the G2 checkpoint response (Zhao et al., 2002). Furthermore, experiments in Xenopus indicate that CHK1, when activated, phosphorylates Wee1 on Ser549. This phosphorylation promotes binding of the 14-3-3 protein that enhances the inhibitory activity of the kinase towards CDC2 (Lee et al., 2001) and adds in this way another layer of control and consolidation of the G2 arrest.
DNA damage also regulates CDC2 activity by regulating cyclin B levels and subcellular localization (Bulavin et al., 2002). In some cell lines, cyclin B mRNA levels decline after IR, possibly due to decreased stability (Muschel et al., 1992; Hwang et al., 1995; Maity et al., 1996; Crawford and Piwnica-Worms, 2001), and recent results indicate that this effect contributes to the maintenance of the G2 arrest (DeSimone et al., 2003). Subcellular compartmentalization is an important mechanism of cyclin regulation in higher eukaryotes even in the absence of DNA damage. Early studies demonstrate that in contrast to cyclin A, which remains nuclear after synthesis, Cyclin B (with CDC2) is initially localized in the cytoplasm during S and G2 phases and translocates to the nucleus at the beginning of mitosis (Pines and Hunter, 1991). A cytoplasmic retention signal at the N-terminal part of the protein, containing a hydrophobic nuclear export signal that binds the nuclear export factor CRM1 (exportin 1), appears responsible for the nuclear exclusion during S and G2 (Pines and Hunter, 1994; Li et al., 1995; Hagting et al., 1998; Toyoshima et al., 1998; Yang et al., 1998). After induction of DNA damage, Cyclin B remains sequestered in the cytoplasm, and in certain types of cells nuclear targeting causes premature mitotic events (Jin et al., 1998; Toyoshima et al., 1998; Kao et al., 1999). There is evidence that 14-3-3σ is required for sequestration of cyclin B in the cytoplasm in response to DNA damage (Chan et al., 1999, 2000b; Hermeking et al., 2000).
Other studies suggest additional regulation for the G2 arrest by two newly discovered regulators of CDC25C, PLK1 and PLK3. Both proteins are members of the Polo-like kinase (PLK) family that play a crucial role in several mitotic events including initiation and exit from mitosis, as well as centrosome function (Glover et al., 1998; Nigg, 1998). A hallmark feature of this family of kinases is the presence of a highly conserved carboxyl terminal region that includes two blocks of strong similarity termed polo boxes. A recent report identifies a novel phosphopeptide-binding domain in the carboxyl region of the protein that encompasses the two polo domains (polo box domain, PBD). It is thought that PBD is essential for recognition of the appropriate substrates by the PLK family of kinases. PLK1 is a positive regulator of CDC25C activity in nonirradiated cells and acts to promote mitotic entry by phosphorylating CDC25C. These interactions are facilitated by the PBD that recognizes CDK phosphorylation consensus sites such as those present in CDC25C (Elia et al., 2003; Sillje and Nigg, 2003). It has been proposed that ATR and ATM phosphorylate and inactivate PLK1, thus depriving the cell from a CDC25C activator and consolidating the delay in the entry of cells into M (Smits et al., 2000; van Vugt et al., 2001). Along these lines, it has been demonstrated that PLK1 phosphorylation correlates with a decrease in CDC2-cyclin B kinase activity, and that this inhibition is alleviated by caffeine, which inhibits ATM and ATR. On the other hand, expression of an unphosphorylatable PLK1 cannot contribute to the G2/M arrest induced by DNA damage (Smits et al., 2000). An additional exciting observation is that PLK1 protein stability is regulated by the checkpoint protein Chfr, which delays entry into mitosis when cells are under mitotic stress (Scolnick and Halazonetis, 2000; Kang et al., 2002). Chfr in Xenopus extracts ubiquitinates the human PLK1 to target it for degradation, thereby inhibiting CDC25C activation. The mechanism of Chfr activation after IR is not known. However, these results indicate an intricate web of interactions between pathways ultimately leading to the silencing of CDC2 activity. The second member of the family, PLK3, is activated by DNA damage in an ATM-dependent manner and is known to interact and to phosphorylate CDC25C at Ser216, which leads to inhibition of its activity (Smits et al., 2000; Xie et al., 2001).
Additional inhibition in the progression into M phase in the presence of DNA damage may be accomplished by the CDK inhibitor P21 through PCNA (Ando et al., 2001). The latter protein binds P21, CDC25C and CDC2-cyclin B, but not simultaneously, and may act as a platform for protein interactions (Xiong et al., 1992; Harper et al., 1993; Kawabe et al., 2002). It appears that binding of P21 and CDC25C to the PCNA–CDC2–cyclin B complex is mutually exclusive (Ando et al., 2001); as a result, P21 interaction with PCNA–CDC2–cyclin B will exclude CDC25C from interacting with CDC2 to dephosphorylate it for mitotic progression. In addition, P21 may act by blocking CAK (Pines, 1995), which activates CDC2 through phosphorylation on Thr161 (Smits et al., 2000). The involvement of P21 in the G2 checkpoint also implicates P53 in the response. Indeed, there is evidence for a role of P53 in the maintenance of the G2 delay through activation of three of its transcriptional target proteins, Gadd45, P21 and 14-3-3σ, and repression in the transcription of CDC2 and Cyclin B (Powell et al., 1995; Hermeking et al., 1997; Chan et al., 1999; Piwnica-Worms, 1999; Taylor and Stark, 2001; DeSimone et al., 2003). It may also be relevant in this regard that CHK2 null ES cells are defective in maintaining, but not in initiating, G2 arrest in response to IR (Hirao et al., 2000), and that CHK1-deficient DT40 cells are viable but fail to arrest in G2 in response to IR (Zachos et al., 2003). Finally, there is evidence for an involvement of BRCA1 in the G2 checkpoint, either through ATM/ATR or possibly also by directly activating CHK1 (Larson et al., 1997; Somasundaram et al., 1997; Tibbetts et al., 2000; Xu et al., 2001; Yarden et al., 2002), but the mechanistic significance of these observations remains unknown.
The above-outlined network of interactions regulating progression through G2 after DNA damage is further enriched by the recognition that the actual mechanism of the checkpoint may also be determined by the phase in the cell cycle where radiation is given. Recent results suggest the activation after DNA damage of two molecularly distinct checkpoints in G2 (Xu et al., 2002). One occurs early after IR, pertains to cells in G2 at the time of irradiation, is ATM dependent, transient and dose independent, and is reflected by an abrupt reduction in mitotic index. The second, which occurs later, pertains to cells irradiated in earlier phases of the cell cycle, is ATM independent, dose dependent and reflected by an accumulation of cells in G2. Earlier studies provided evidence for strong dependence of G2 arrest on the phase of the cell cycle at the time of radiation and demonstrated an order-of-magnitude longer delay for cells irradiated in G2. How these results correlate with the above observations and the nature of the underlying molecular mechanisms are likely to be active areas of investigation in the coming years.
DNA damage checkpoints and the repair of DNA DSBs
The widely held view that checkpoints aid DNA repair is logical, supported by circumstantial evidence but not rigorously proven. Although an arrest in cell cycle progression provides time before the initiation of the next cell cycle transition, it is clear that the checkpoint response is more than a waiting period in the cycle. Indeed, simply providing time for repair by manipulating growth conditions in protocols devised to evaluate repair of potentially lethal damage (PLD) (Iliakis, 1988) cannot modulate radiosensitivity in AT cells (Weichselbaum et al., 1978). Furthermore, genetic manipulations that restore checkpoint response in defective cell lines without correcting for radiosensitivity to killing have been described (Xu et al., 2002), and there is a report for an AT derivative cell line with wild-type radiosensitivity to killing that still displays RDS (Lehmann et al., 1986).
To put the DNA damage checkpoint and DNA repair in perspective, it is perhaps useful to attempt integration with the essential events of the normal cell cycle. The discussion centers on DNA DSBs and their repair, but it could be expanded to include other types of DNA damage as well. One major task of the regulatory transitions that drive the cell cycle is to bring the DNA to a state competent for duplication and division (Stillman, 1996). Along these lines, the regulatory transitions associated with checkpoint activation should bring the DNA to a state competent for DSB repair. DNA DSBs can be induced in all phases of the cell cycle, but their repair is likely to proceed optimally only in certain phases of the cell cycle, and alterations in chromatin conformation associated with cell cycle transitions may interfere with it. A checkpoint response may therefore be useful on several fronts: (1) allow time for repair in the phase of the cell cycle where DSBs are induced; (2) allow partial processing and safe transition of the cell and the lesion into a phase of the cycle where repair proceeds optimally; (3) prevent an imminent transition in chromatin conformation that will undermine correct processing.
Before further discussion of these aspects of checkpoint response, it will be instructive to review salient features of DNA DSB repair. Cells of higher eukaryotes remove IR-induced DNA DSBs mainly by NHEJ (Jeggo, 1998; Jackson, 2002). Homology-directed repair, although an important determinant of cell radiosensitivity to killing, does not appear to measurably contribute to the removal from the genome of IR-induced DNA DSBs (Wang et al., 2001a, 2001b; Xia et al., 2001). NHEJ is an extremely fast process removing over 75% of the induced breaks in less than 20 min in some cell lines and is active in every phase of the cell cycle including S phase (Metzger and Iliakis, 1991). It is unlikely, therefore, that this process will benefit significantly by checkpoint responses whose maximal manifestation requires at least 1–2 h. Indeed, evidence has been presented that NHEJ is a checkpoint-independent process (Wang et al., 2003b).
On the other hand, evidence accumulates for linkages between checkpoint response and homology-directed repair (Wang et al., 2003a). Several of the proteins participating in the checkpoint pathways outlined above have been implicated, directly or indirectly, in this form of DNA DSB repair (Hoeijmakers, 2001; Jackson, 2002), and the slower kinetics of this pathway are in line with those of checkpoint response. We are thus faced with the apparent contradiction that a major pathway of DNA DSB repair does not seem to take advantage of the checkpoint response, while an apparently minor one does.
We attempted to resolve this discrepancy, as well as other discrepancies mentioned elsewhere, by proposing that NHEJ is the first step in the process of DSB repair aiming at restoring continuity in the genome, albeit with low fidelity. It is followed by homology-directed processes aiming at restoring, whenever critical, the sequence around the break (Asaad et al., 2000; Wang et al., 2001b). This second phase would also restore fidelity in the overall repair process. In this highly speculative view, repair of DNA DSBs is a multistep process that starts with the synapsis of the DNA ends and completes with the restoration of the DNA sequence. The initial rapid step of end-joining is likely to be sensitive to abrupt alterations in chromatin conformation such as those occurring at the beginning of the S phase as well as at mitosis, as they may cause end separation that will undermine correct rejoining. Checkpoints may serve here by delaying these transitions. Subsequent steps of the checkpoint response are likely to be more integrated with the subsequent repair steps and may serve to bring the DNA to a state where homology-directed repair proceeds efficiently. At the same time, they may block the biochemical processes associated with DNA duplication and division. Only more research will show whether such a model is tenable, how the repair process is implemented in its details, where in the cell cycle the different steps take place, and what cell cycle transitions may be required for its completion. This information may also clarify the important question of the activating signal for the checkpoint response.
Checkpoints and cancer
The essential nature of checkpoint responses in human health and genomic stability is most clearly indicated by the observation that defects in nearly all identified component proteins are associated with severe hereditary human genetic disorders or a predisposition to cancer (Hoeijmakers, 2001; Khanna and Jackson, 2001; van Gent et al., 2001). Therefore, detailed understanding of the pathways underlying checkpoint response, as well as identification and characterization of the participating proteins, will significantly advance our ability to unravel the complex processes leading to the development of cancer.
Furthermore, understanding of the mechanisms underlying checkpoint response after DNA damage will benefit existing therapeutic modalities and likely contribute to the development of novel cancer-treatment approaches (Harris et al., 1998; Sampath and Plunkett, 2001). This is because IR and the vast majority of drugs used in cancer therapy act on the DNA, either by directly inflicting DNA damage, or by affecting important aspects of DNA metabolism and causing damage indirectly. As a result, checkpoint activation becomes an important determinant of the ultimate response to the treatment, and targeted interference, specifically in the tumor cell, a promising strategy for an improved outcome. The frequent abrogation of important checkpoint responses in tumor cells offers an initial advantage towards this goal.
Therapeutic strategies have been conceived that manipulate CDK activity indirectly by altering the checkpoint pathways that regulate CDK function or directly by affecting the catalytic subunit of CDK. Thus, expression of P21 in a variety of tumor cell lines into P53 or P21 negative cells causes increased apoptosis, reduced cell proliferation and reduced tumor growth. Small-molecule inhibitors of CDKs further hold great promise as antineoplastic agents because they induce cell cycle arrest by directly inhibiting CDKs, generally by insertion into the ATP binding site. Some of these compounds, such as UCN-01 and flavopiridol, are currently in clinical trials (Senderowicz and Sausville, 2000; Sampath and Plunkett, 2001). Also, peptide inhibitors of CHK1 and CHK2 have been developed and are being evaluated (Suganuma et al., 1999). It is thus evident that the recent advances made in understanding the cell cycle machinery and the DNA damage checkpoint response have identified a wealth of new targets for the development of new antineoplastic agents, or of response modifiers for existing ones.
Abraham RT . (2001). Genes Dev., 15, 2177–2196.
Agami R and Bernards R . (2000). Cell, 102, 55–66.
Amati B, Dalton S, Brooks MW, Littlewood TD, Evans GI and Land H . (1992). Nature, 359, 423–426.
Anderson CW and Carter TH . (1996). Curr. Top. Microbiol. Immunol., 217, 91–111.
Anderson L, Henderson C and Adachi Y . (2001). Mol. Cell. Biol., 21, 1719–1508.
Ando T, Kawabe T, Ohara H, Ducommun B and Itoh M . (2001). J. Biol. Chem., 276, 42971–42977.
Asaad NA, Zeng Z-C, Guan J, Thacker J and Iliakis G . (2000). Oncogene, 19, 5788–5800.
Atchley WR and Fitch WM . (1995). Proc. Natl. Acad. Sci. USA, 92, 10217–10221.
Bakkenist CJ and Kastan MB . (2003). Nature, 421, 499–506.
Banin S, Moyal L, Shieh S-Y, Taya Y, Anderson CW, Chessa L, Smorodinsky NI, Prives C, Reiss Y, Shiloh Y and Ziv Y . (1998). Science, 281, 1674–1677.
Bao S, Tibbetts RS, Brumbaugh KM, Fang Y, Richardson DA, All A, Chen SM, Abraham RT and Wang X-F . (2001). Nature, 411, 969–974.
Bartek J, Falck J and Lukas J . (2001). Nat. Rev. Mol. Cell Biol., 2, 877–886.
Bartek J and Lukas C . (2001a). FEBS Lett., 490, 117–122.
Bartek J and Lukas J . (2001b). Curr. Opin. Cell Biol., 13, 738–747.
Bell SP and Dutta A . (2002). Annu. Rev. Biochem., 71, 333–374.
Bermudez VP, Lindsey-Boltz LA, Cesare AJ, Maniwa Y, Griffith JD, Hurwitz J and Sancar A . (2003). Proc. Natl. Acad. Sci. USA, 100, 1633–1638.
Bernhard EJ, Maity A, Muschel RJ and McKenna WG . (1995). Radiat. Environ. Biophys., 34, 79–83.
Bernhard EJ, McKenna WG and Muschel RJ . (1999). Cancer J., 5, 194–204.
Blasina A, Price BD, Turenne GA and McGowan CH . (1999). Curr. Biol., 9, 1135–1138.
Booher RN, Holman PS and Fattaey A . (1997). J. Biol. Chem., 272, 22300–22306.
Brown EJ and Baltimore D . (2000). Genes Dev., 14, 397–402.
Brush GS and Kelly TJ . (1996). Mechanisms for Replication DNA. Cold Spring Harbor Laboratory Press: New York.
Bulavin DV, Amundson SA and Fornace Jr AJ . (2002). Curr. Opin. Genet. Dev., 12, 92–97.
Canman CE, Lim D-S, Cimprich KA, Taya Y, Tamai K, Sakaguchi K, Appella E, Kastan MB and Siliciano JD . (1998). Science, 281, 1677–1679.
Chan DW, Son S-C, Block W, Ye R, Khanna KK, Wold MS, Douglas P, Goodarzi AA, Pelley J, Taya Y, Lavin MF and Lees-Miller SP . (2000a). J. Biol. Chem., 275, 7803–7810.
Chan TA, Hermeking H, Lengauer C, Kinzler KW and Vogelstein B . (1999). Nature, 401, 616–620.
Chan TA, Hwang PM, Hermeking H, Kinzler KW and Vogelstein B . (2000b). Genes Dev., 14, 1584–1588.
Cleaver JE, Rose R and Mitchell DL . (1990). Radiat. Res., 124, 294–299.
Cliby WA, Roberts CJ, Cimprich KA, Stringer CM, Lamb JR, Schreiber SL and Friend SH . (1998). EMBO J., 17, 159–169.
Cortez D, Guntuku S, Qin J and Elledge SJ . (2001). Science, 294, 1713–1716.
Cortez D, Wang Y, Qin J and Elledge SJ . (1999). Science, 286, 1162–1166.
Costanzo V, Robertson K, Ying CY, Kim E, Avvedimento E, Gottesman M, Grieco D and Gautier J . (2000). Mol. Cell, 6, 649–659.
Crawford DF and Piwnica-Worms H . (2001). J. Biol. Chem., 276, 37166–37177.
D'Amours D, Desnoyers S, D'Silva I and Poirier GG . (1999). Biochem. J., 342, 249–268.
DePamphilis ML . (1999). BioEssays, 21, 5–16.
Desai-Mehta A, Cerosaletti KM and Concannon P . (2001). Mol. Cell. Biol., 21, 2184–2025.
DeSimone JN, Bengtsson U, Wang Q, Lao XY, Redpath JL and Stanbridge EJ . (2003). Radiat. Res., 159, 72–85.
DiBiase SJ, Zeng Z-C, Chen R, Hyslop T, Curran Jr WJ and Iliakis G . (2000). Cancer Res., 60, 1245–1253.
Donaldson AD and Blow JJ . (2001). Curr. Biol., 11, R979–R982.
Dotto GP . (2000). Biochim. Biophys. Acta, 1471, M43–M56.
Draetta G and Eckstein J . (1997). Biochim. Biophys. Acta, 1332, M53–M63.
Durocher D and Jackson SP . (2001). Curr. Opin. Cell Biol., 13, 225–231.
Dutta A and Bell SP . (1997). Ann. Rev. Cell Biol., 13, 293–332.
Edwards RJ, Bentley NJ and Carr AM . (1999). Nat. Cell Biol., 1, 393–398.
Eisenman RN and Cooper JA . (1995). Nature, 378, 438–439.
Elia AEH, Cantley LC and Yaffe MB . (2003). Science, 299, 1228–1231.
Elledge SJ . (1996). Science, 274, 1664–1672.
Falck J, Mailand N, Syljuasen RG, Bartek J and Lukas J . (2001). Nature, 410, 842–847.
Falck J, Petrini JHJ, Williams BR, Lukas J and Bartek J . (2002). Nat. Genet., 30, 290–294.
Friedberg EC, Walker GC and Siede W . (1995). DNA Repair Mutagenesis. ASM Press: Washington, DC.
Furnari B, Blasina A, Boddy MN, McGowan CH and Russell P . (1999). Mol. Biol. Cell, 10, 833–845.
Galaktionov K, Chen X and Beach D . (1996). Nature, 382, 511–517.
Gatei M, Scott SP, Filippovitch I, Soronika N, Lavin MF, Weber B and Khanna KK . (2000a). Cancer Res., 60, 3299–3304.
Gatei M, Young D, Cerosaletti KM, Desai-Mehta A, Spring K, Kozlov S, Lavin MF, Gatti RA, Concannon P and Khanna KK . (2000b). Nat. Genet., 25, 115–119.
Giaccia AJ and Kastan MB . (1998). Genes Dev., 12, 2973–2983.
Glover DM, Hagan IM and Tavares AAM . (1998). Genes Dev., 12, 3777–3787.
Goldberg M, Stucki M, Falck J, D'Amours D, Rahman D, Pappin D, Bartek J and Jackson SP . (2003). Nature, 421, 952–956.
Gottifredi V, Shieh SY, Taya Y and Prives C . (2001). Proc. Natl. Acad. Sci. USA, 98, 1036–1041.
Graves PR, Yu L, Schwarz JK, Gales J, Sausville EA, O'Connor PM and Piwnica-Worms H . (2000). J. Biol. Chem., 275, 5600–5605.
Green CM, Erdjument-Bromage H, Tempst P and Lowndes NF . (2000). Curr. Biol., 10, 39–42.
Griffiths DJ, Barbet NC, McCready S, Lehmann AR and Carr AM . (1995). EMBO J., 14, 5812–5823.
Guan J, DiBiase S and Iliakis G . (2000). Nucleic Acids Res., 28, 1183–1192.
Guo CY, D'Anna JA and Larner JM . (1999). Radiat. Res., 151, 125–132.
Guo Z, Kumagai A, Wang SX and Dunphy WG . (2000). Genes Dev., 14, 2745–2756.
Hagting A, Karlsson C, Clute P, Jackman M and Pines J . (1998). EMBO J., 17, 4127–4138.
Harper JW, Adami GR, Wei N, Keyomarsi K and Elledge SJ . (1993). Cell, 75, 805–816.
Harris EE, Kao GD, Muschel RJ and McKenna WG . (1998). Cancer Treat. Res., 93, 169–190.
Hartwell LH and Kastan MB . (1994). Science, 266, 1821–1828.
Hartwell LH and Weinert TA . (1989). Science, 246, 629–634.
Hermeking H, Lengauer C, Polyak K, He T-C, Zhang L, Thiagalingam S, Kinzler KW and Vogelstein B . (1997). Mol. Cell, 1, 3–11.
Hermeking H, Rago C, Schuhmacher M, Li Q, Barrett JF, Obaya AJ, O'Connell BC, Mateyak MK, Tam W, Kohlhuber F, Dang CV, Sedivy JM, Eick D, Vogelstein B and Kinzler KW . (2000). Proc. Natl. Acad. Sci. USA, 97, 2229–2234.
Hickman ES, Moroni MC and Helin K . (2002). Curr. Opin. Genet. Dev., 12, 60–66.
Hirao A, Kong YY, Matsuoka S, Wakeham A, Ruland J, Yoshida H, Liu D, Elledge SJ and Mak TW . (2000). Science, 287, 1824–1827.
Hoeijmakers JHJ . (2001). Nature, 411, 366–374.
Houldsworth J and Lavin MF . (1980). Nucleic Acids Res., 8, 3709–3720.
Hwang A, Maity A, McKenna WG and Muschel RJ . (1995). J. Biol. Chem., 270, 28419–28424.
Iliakis G . (1988). Int. J. Radiat. Biol., 53, 541–584.
Iliakis G . (1997). Semin. Oncol., 24, 602–615.
Jackson SP . (2002). Carcinogenesis, 23, 687–696.
Jeggo PA . (1997). Mutat. Res., 384, 1–14.
Jeggo PA . (1998). Adv. Genet., 38, 186–218.
Jin P, Gu Y and Morgan DO . (1996). J. Cell. Biol., 134, 963–970.
Jin P, Hardy S and Morgan DO . (1998). J. Cell Biol., 141, 875–885.
Kang D, Chen J, Wong J and Fang G . (2002). J. Cell Biol., 156, 249–259.
Kao GD, McKenna WG and Muschel RJ . (1999). J. Biol. Chem., 274, 34779–34784.
Kastan MB . (2001). Nature, 410, 766–767.
Kastan MB and Lim D-S . (2000). Nat. Rev. Mol. Cell Biol., 1, 179–186.
Kawabe T, Suganuma M, Ando T, Kimura M, Hori H and Okamoto T . (2002). Oncogene, 21, 1717–1726.
Khanna KK and Jackson SP . (2001). Nat. Genet., 27, 247–254.
Khanna KK, Keating KE, Kozlov S, Scott S, Gatei M, Hobson K, Taya Y, Gabrielli B, Chan D, Lees-Miller SP and Lavin MF . (1998). Nat. Genet., 20, 398–400.
Kim S-T, Lim D-S, Canman CE and Kastan MB . (1999). J. Biol. Chem., 274, 37538–37543.
Kim S-T, Xu B and Kastan MB . (2002). Genes Dev., 16, 560–570.
Lamb JR, Petit-Frere C, Broughton BC, Lehmann AR and Green MHL . (1989). Int. J. Radiat. Biol., 56, 125–130.
Larner JM, Lee H and Hamlin JL . (1997). Cancer Surv., 29, 25–45.
Larner JM, Lee H, Little RD, Dijkwel PA, Schildkraut CL and Hamlin JL . (1999). Nucleic Acids Res., 27, 803–809.
Larson JS, Tonkinson JL and Lai MT . (1997). Cancer Res., 57, 3351–3355.
Lavin MF and Schroeder AL . (1988). Mutat. Res., 193, 193–206.
Lee C-H and Chung JH . (2001). J. Biol. Chem., 276, 30537–30541.
Lee H, Larner JM and Hamlin JL . (1997). Proc. Natl. Acad. Sci. USA, 94, 526–531.
Lee J, Kumagai A and Dunphy WG . (2001). Mol. Biol. Cell, 12, 551–563.
Lee M and Nurse P . (1988). Trends Genet., 4, 287–290.
Lees-Miller SP . (1996). Biochem. Cell Biol., 74, 503–512.
Lehmann AR, Arlett CF, Burke JF, Green MHL, James MR and Lowe JE . (1986). Int. J. Radiat. Biol., 49, 639–643.
Leone G, DeGregori J, Sears R, Jakoi L and Nevins JR . (1997). Nature, 387, 422–425.
Li J, Meyer AN and Donoghue DJ . (1995). Mol. Biol. Cell, 6, 1111–1124.
Li S, Ting NSY, Zheng L, Chen P-L, Ziv Y, Shiloh Y, Lee EY-HP and Lee W-H . (2000). Nature, 406, 210–215.
Lim D-S, Kim S-T, Xu B, Maser RS, Lin J, Petrini JHJ and Kastan MB . (2000). Nature, 404, 613–617.
Lindsey-Boltz LA, Bernudez VP, Hurwitz J and Sancar A . (2001). Proc. Natl. Acad. Sci. USA, 98, 11236–11241.
Liu F-F, Stanton JJ, Wu Z and Piwnica-Worms H . (1997). Mol. Cell. Biol., 17, 571–583.
Liu Q, Guntuku S, Cui XS, Matsuoka S, Cortez D, Tamai K, Luo G, Carattini-Rivera S, DeMayo F, Bradley A, Donehower LA and Elledge SJ . (2000). Genes Dev., 14, 1448–1459.
Lopez-Girona A, Kanoh J and Russell P . (2001). Curr. Biol., 11, 50–54.
Lou Z, Minter-Dykhouse K, Wu X and Chen J . (2003). Nature, 421, 957–961.
Lücke-Huhle C . (1982). Radiat. Res., 89, 298–308.
Lukas C, Bartkova J, Latella L, Falck J, Mailand N, Schroeder T, Sehested M, Lukas J and Bartek J . (2001). Cancer Res., 61, 4990–4993.
Lupardus PJ, Byun T, Yee M-c, Hekmat-Nejad M and Cimprich KA . (2002). Genes Dev., 16, 2327–2332.
Lydall D and Weinert T . (1995). Science, 270, 1488–1491.
Mailand N, Falck J, Lukas C, Syljuasen RG, Welcker M, Bartek J and Lukas J . (2000). Science, 288, 1425–1429.
Maity A, Hwang A, Janss A, Phillips P, McKenna WG and Muschel RJ . (1996). Oncogene, 13, 1647–1657.
Maity A, McKenna WG and Muschel RJ . (1994). Radiother. Oncol., 31, 1–13.
Marcu KB, Bossone SA and Patel AJ . (1992). Annu. Rev. Biochem., 61, 809–860.
Maser RS, Mirzoeva OK, Wells J, Olivares H, Williams BR, Zinkel RA, Farnham PJ and Petrini JHJ . (2001). Mol. Cell. Biol., 21, 6006–6016.
Matsuoka S, Huang M and Elledge SJ . (1998a). Science, 282, 1893–1897.
Matsuoka S, Huang M and Elledge SJ . (1998b). Science, 282, 1893–1897.
Matsuoka S, Rotman G, Ogawa A, Shiloh Y, Tamai K and Elledge SJ . (2000). Proc. Nat. Acad. Sci. USA, 97, 10389–10394.
McKenna WG . (1995). 37th Annual Meeting, American Society for Therapeutic Radiology and Oncology. Miami Beach, FL.
McKenna WG, Iliakis G, Weiss MC, Bernhard EJ and Muschel RJ . (1991). Radiat. Res., 125, 283–287.
Melchionna R, Chen X-B, Blasina A and McGowan CH . (2000). Nat. Cell Biol., 2, 762–765.
Metzger L and Iliakis G . (1991). Int. J. Radiat. Biol., 59, 1325–1339.
Morgan DO . (1995). Nature, 374, 131–134.
Muschel RJ, Zhang HB, Iliakis G and McKenna WG . (1992). Radiat. Res., 132, 153–157.
Nigg EA . (1998). Curr. Opin. Cell Biol., 10, 776–783.
Norbury C and Nurse P . (1992). Annu. Rev. Biochem., 61, 441–470.
Nurse P . (1990). Nature, 344, 503–508.
Nurse P . (1994). Cell, 79, 547–550.
Nurse P . (1997). Cell, 91, 865–867.
Nyberg KA, Michelson RJ, Putnam CW and Weinert TA . (2002). Annu. Rev. Genet., 36, 617–656.
O'Connell MJ, Walworth NC and Carr AM . (2002). Trends Cell Biol., 10, 296–303.
Paciotti V, Clerici M, Lucchini G and Longhese MP . (2000). Genes Dev., 14, 2046–2059.
Painter RB . (1981). Mutat. Res., 84, 183–190.
Painter RB . (1986). Int. J. Radiat. Biol., 49, 771–781.
Painter RB and Young BR . (1980). Proc. Nat. Acad. Sci. USA, 77, 7315–7317.
Parker AE, Van de Weyer I, Laus MC, Oostveen I, Yon J, Verhasselt P and Luyten WHML . (1998). J. Biol. Chem., 273, 18332–18339.
Parker LL and Piwnica-Worms H . (1992). Science, 257, 1955–1957.
Paulovich AG, Toczyski DP and Hartwell LH . (1997). Cell, 88, 315–321.
Peng C-Y, Graves PR, Thoma RS, Wu Z, Shaw AS and Piwnica-Worms H . (1997). Science, 277, 1501–1505.
Petrini JH . (2000). Curr. Opin. Cell Biol., 12, 293–296.
Pines J . (1995). Semin. Cancer Biol., 6, 63–72.
Pines J and Hunter T . (1991). J. Cell Biol., 115, 1–17.
Pines J and Hunter T . (1994). EMBO J., 13, 3772–3781.
Piwnica-Worms H . (1999). Nature, 401, 535–537.
Powell SN, DeFrank JS, Connell P, Eogan M, Preffer F, Dombkowski D, Tang W and Friend S . (1995). Cancer Res., 55, 1643–1648.
Rhind N and Russell P . (2000). J. Cell Sci., 113, 3889–3896.
Rotman G and Shiloh Y . (1998). Human Mol. Genet., 1998, 1555–1563.
Rotman G and Shiloh Y . (1999). Oncogene, 18, 6135–6144.
Rouse J and Jackson SP . (2000). EMBO J., 19, 5801–5812.
Sampath D and Plunkett W . (2001). Curr. Opin. Oncol., 13, 484–490.
Sanchez Y, Wong C, Thoma RS, Richman R, Wu Z, Piwnica-Worms H and Elledge SJ . (1997). Science, 277, 1497–1501.
Santoni-Rugiu E, Falck J, Mailand N, Bartek J and Lukas J . (2000). Mol. Cell. Biol., 20, 3497–3509.
Schultz LB, Chehab NH, Malikzay A and Halazonetis TD . (2000). J. Cell Biol., 151, 1381–1390.
Scolnick DM and Halazonetis TD . (2000). Nature, 406, 430–435.
Scully R and Livingston DM . (2000). Nature, 408, 429–442.
Senderowicz AM and Sausville EA . (2000). J. Nat. Cancer Inst., 92, 376–387.
Seoane J, Le H-V and Massague J . (2002). Nature, 419, 729–734.
Sheen J-H and Dickson RB . (2002). Mol. Cell. Biol., 22, 1819–1833.
Sherr CJ . (1995). Trends Biol. Sci., 20, 187–191.
Sherr CJ . (1996). Science, 274, 1672–1677.
Sherr CJ and Roberts JM . (1995). Genes Dev., 9, 1149–1163.
Sherr CJ and Roberts JM . (1999). Genes Dev., 13, 1501–1512.
Shiloh Y . (2001). Curr. Opin. Genet. Dev., 11, 71–77.
Sillje HHW and Nigg EA . (2003). Science, 299, 1190–1192.
Smeets MFMA, Mooren EHM, Abdel-Wahab AHA, Bartelink H and Begg AC . (1994). Radiat. Res., 140, 153–160.
Smith GCM, Cary RB, Lakin ND, Hann BC, Teo S-H, Chen DJ and Jackson SP . (1999). Proc. Natl. Acad. Sci. USA, 96, 11134–11139.
Smith GCM and Jackson SP . (1999). Genes Dev., 13, 916–934.
Smith S . (2001). Trends Biochem. Sci., 26, 175–180.
Smits VAJ, Klompmaker R, Arnaud L, Rijksen G, Nigg EA and Medema RH . (2000). Nat. Cell Biol., 2, 672–676.
Smits VAJ and Medema RH . (2001). Biochim. Biophys. Acta, 1519, 1–12.
Somasundaram K, Zhang H, Zeng YX, Houvras Y, Peng Y, Wu GS, Licht JD, Weber BL and El-Deiry WS . (1997). Nature, 389, 187–190.
Stewart GS, Wang B, Rignell CR, Taylor AMR and Elledge SJ . (2003). Nature, 421, 961–966.
Stillman B . (1994). Cell, 78, 725–728.
Stillman B . (1996). Science, 274, 1659–1664.
Stokes MP, Van Hatten R, Lindsay HD and Michael WM . (2002). J. Cell Biol., 158, 863–872.
Su L and Little JB . (1993). Radiat. Res., 133, 73–79.
Suganuma M, Kawabe T, Hori H, Funabiki T and Okamoto T . (1999). Cancer Res., 59, 5887–5891.
Takai H, Tominaga K, Motoyama N, Minamishima YA, Nagahama H, Tsukiyama T, Ikeda K, Nakayama K and Nakanishi M . (2000). Genes Dev., 14, 1439–1447.
Taylor WR, Agarwall ML, Agarwal A, Stacey D and Stark GR . (1999). Oncogene, 18, 283–295.
Taylor WR and Stark GR . (2001). Oncogene, 20, 1803–1815.
Terada Y, Tatsuka M, Jinno S and Okayama H . (1995). Nature, 376, 358–362.
Thompson LH and Schild D . (2001). Mutat. Res., 477, 131–153.
Thompson LH and Schild D . (2002). Mutat. Res., 509, 49–78.
Tibbetts RS, Brumbaugh KM, Williams JM, Sarkaria JN, Cliby WA, Shieh S-Y, Taya Y, Prives C and Abraham RT . (1999). Genes Dev., 13, 152–157.
Tibbetts RS, Cortez D, Brumbaugh KM, Scully R, Livingston D, Elledge SJ and Abraham RT . (2000). Genes Dev., 14, 2989–3002.
Tobey RA . (1975). Nature, 254, 245–247.
Toyoshima F, Moriguchi T, Wada A, Fukuda M and Nishida E . (1998). EMBO J., 17, 2728–2735.
Tye BK . (1999). Annu. Rev. Biochem., 68, 649–686.
van Gent DC, Hoeijmakers JHJ and Kanaar R . (2001). Nat. Rev. Genet., 2, 196–206.
van Vugt MATM, Smits VAJ, Klompmaker R and Medema RH . (2001). J. Biol. Chem., 276, 41656–41660.
Venkitaraman AR . (2001). J. Cell Sci., 114, 3591–3598.
Vogelstein B, Lane DP and Levine AJ . (2000). Nature, 408, 307–310.
Volkmer E and Karnitz LM . (1999). J. Biol. Chem., 274, 567–570.
Wakayama T, Kondo T, Ando S, Matsumoto K and Sugimoto K . (2001). Mol. Cell. Biol., 21, 755–764.
Walters RA, Gurley LR and Tobey RA . (1974). Biophys. J., 14, 99–118.
Walworth NC . (2001). Curr. Opin. Genet. Dev., 11, 78–82.
Wang H, Wang X, Iliakis G and Wang Y . (2003a). Radiat. Res., 159, 420–425.
Wang H, Zeng Z-C, Bui T-A, DiBiase SJ, Qin W, Xia F, Powell SN and Iliakis G . (2001a). Cancer Res., 61, 270–277.
Wang H, Zeng Z-C, Bui T-A, Sonoda E, Takata M, Takeda S and Iliakis G . (2001b). Oncogene, 20, 2212–2224.
Wang JYY . (2000). Nature, 405, 404–405.
Wang X, Wang H, Iliakis G and Wang Y . (2003b). Radiat. Res.. (in press).
Wang Y, Cortez D, Yazdi P, Neff N, Elledge SJ and Qin J . (2000). Genes Dev., 14, 927–939.
Wang Y, Huq MS, Cheng X and Iliakis G . (1995). Radiat. Res., 142, 169–175.
Wang Y, Zhou XY, Wang H-Y and Iliakis G . (1999). J. Biol. Chem., 274, 22060–22064.
Weichselbaum RR, Nove J and Little JB . (1978). Nature, 271, 261–262.
Wolkow TD and Enoch T . (2002). Mol. Biol. Cell, 13, 480–492.
Wu X, Ranganathan V, Weisman DS, Heine WF, Ciccone DN, O'Neill TB, Crick KE, Pierce KA, Lane WS, Rathbun G, Livingston DM and Weaver DT . (2000). Nature, 405, 477–482.
Xia F, Taghian DG, DeFrank JS, Zeng Z-C, Willers H, Iliakis G and Powell SN . (2001). Proc. Natl. Acad. Sci. USA, 98, 8644–8649.
Xie S, Wu H, Wang Q, Cogswell P, Husain I, Conn C, Stambrook P, Jhanwar-Uniyal M and Dai W . (2001). J. Biol. Chem., 276, 43305–43312.
Xiong Y, Zhang H and Beach D . (1992). Cell, 71, 505–514.
Xu B, Kim S-t and Kastan MB . (2001). Mol. Cell. Biol., 21, 3445–3450.
Xu B, Kim S-T, Lim D-S and Kastan MB . (2002). Mol. Cell. Biol., 22, 1049–1059.
Yamane K, Wu X and Chen J . (2002). Mol. Cell. Biol., 22, 555–566.
Yang J, Bardes ESG, Moore JD, Brennan J, Powers MA and Kornbluth S . (1998). Genes Dev., 12, 2131–2143.
Yarden RI, Pardo-Reoyo S, Sgagias M, Cowan KH and Brody LC . (2002). Nat. Genet., 30, 285–289.
Yazdi PT, Wang Y, Zhao S, Patel N, Lee EY-HP and Qin J . (2002). Genes Dev., 16, 571–582.
Yu Q, Geng Y and Sicinski P . (2001). Nature, 411, 1017–1021.
Zachos G, Rainey MD and Gillespie DAF . (2003). EMBO J., 22, 713–723.
Zhao H, Watkins JL and Piwnica-Worms H . (2002). Proc. Natl. Acad. Sci. USA, 99, 14795–14800.
Zhao S, Weng Y-C, Yuan S-SF, Lin Y-T, Hsu HC, Lin S-CJ, Gerbino E, Song M-h, Zdzienicka MZ, Gatti RA, Shay JW, Ziv Y, Shiloh Y and Lee EY-HP . (2000). Nature, 405, 473–477.
Zhou B-BS, Chaturvedi P, Spring K, Scott SP, Johanson RA, Mishra R, Mattern MR, Winkler JD and Khanna KK . (2000). J. Biol. Chem., 275, 10342–10348.
Zhou B-BS and Elledge SJ . (2000). Nature, 408, 433–439.
Zhou X-Y, Wang X, Hu B, Guan J, Iliakis G and Wang Y . (2002). Cancer Res., 62, 1598–1603.
Ziegler M and Oei SL . (2001). BioEssays, 23, 543–548.
Zou L, Cortez D and Elledge SJ . (2002). Genes Dev., 16, 198–208.
We are indebted to Jutta Mueller and Nancy Mott for secretarial assistance. The work in our is supported by NIH Grants CA42026, CA56706, CA76203 and 2P01 CA56690, EU Contracts FIS5-2002-00078 and FIGH-CT-2002-00218, as well as a Grant from the Volkswagenstiftung.
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