Cell Cycle, Regulation of


Christopher W. McAndreW, Department of Chemistry and Biochemistry, University of California, San Diego, California

Randy F. Gastwirt, Department of Biomedical Sciences, University of California, San Diego, California

Daniel J. Donoghue, Department of Chemistry and Biochemistry, Moores UCSD Cancer Center, University of California, San Diego, California

doi: 10.1002/9780470048672.wecb057


Normal regulation of the cell cycle ensures the passage of genetic material without mutations and aberrations. Proper completion of each phase is critical to the initiation of the following phase, and the pathways that cell division occur in an ordered, sequential, and irreversible procession. The two major cell-cycle events that are regulated tightly are DNA replication and cell division. Progression through each phase transition is regulated by extracellular signaling, transcription factors, cyclin-dependent kinases (CDKs), and checkpoints, which prevent uncontrolled cell division. Cyclin/CDK complexes are the primary factors responsible for the timely order of cell-cycle progression, which include entry into S phase, initiation of DNA replication, and mitotic entry. Each phase of the cell cycle and the different cyclin/CDK complexes, as well as other important factors that regulate cell-cycle progression and checkpoints, will be discussed.

The cell cycle is the sequence of events by which growing cells duplicate and divide into two daughter cells. In mammalian cells and other eukaryotes, cell division represents a process of highly ordered and tightly regulated molecular events. The cell cycle is composed of five phases in mammals, including G0, G1, S, G2, and M phases. Replication of DNA occurs during S phase and division occurs during M phase. During the two gap phases, G1 and G2, cells produce RNA and proteins required for the subsequent S and M phases, respectively. Cells in a resting, quiescent state are in G0 phase. Stimulation by external growth factors or mitogens triggers quiescent cells to reenter the cell cycle in G1 by activating numerous signaling cascades, and it leads to the sequential activation of cyclin dependent kinases (CDKs). Activation of CDKs requires interaction with a cyclin partner, T-loop phosphorylation at T160 (CDK2) or T161 (CDK1) catalyzed by CDK activating kinase (CAK), and dephosphorylation at T14 and Y15 by CDC25 dual phosphatases. The inhibitory phosphorylations at T14 and Y15 are catalyzed by the serine/threonine kinase Wee1 and threonine/tyrosine kinase Myt1, and these cause misalignment of the glycine-rich loop and the ATP phosphate moiety. CDKs phosphorylate multiple substrates. The proper regulation of CDKs is necessary for orderly cell-cycle phase transitions. A general representation of the key players and events during the cell cycle can be observed in Fig. 1.

Numerous checkpoints also exist to ensure normal cell-cycle progression and transmission of an unaltered genome. These checkpoints are conserved signaling pathways that monitor cell growth conditions, cell-cycle progression, and structural and functional DNA defects; they are critical for cell survival or death. Checkpoint responses induce and sustain a delay in cell-cycle progression, and activate machinery to respond to changes in cell growth conditions, repair DNA, and stall replication. When cellular damage cannot be repaired, these checkpoints can induce apoptosis. The mammalian checkpoints include the quiescent checkpoint, G1/S checkpoint, replicative checkpoint, G2 checkpoint, mitotic checkpoint, and the DNA damage checkpoints. Improper checkpoint control promotes tumorigenesis through increased mutation rates, aneuploidy, and chromosome instability. The following sections will give an overview of the regulation of the various phases of the mammalian cell cycle, activation of specific checkpoints, and the molecules involved in the mechanisms that regulate these processes.



Figure 1. Regulation of the mammalian cell cycle by cyclin/CDKs. Activation of growth factor receptors in GQ leads to activation of many signaling cascades that lead to the expression of cyclin D. Progression into S phase is mediated by Rb and E2Fs that lead to the initiation and progression of DNA synthesis through cyclin E/A/CDK2 activity. On completion of DNA replication, cyclin B/CDK1 activity promotes phosphorylation of substrates required for entry into mitosis and eventual cytokinesis, which produces two identical daughter cells.


From Quiescence to the Point of No Return

G0-G1 transition

After cell division, the daughter cell enters into Go phase where it becomes ready to divide again before entering into G1. In most cases, the newly formed cell increases in size and mass for division to occur again, by enhancing ribosome biosynthesis (1). This task is accomplished by phosphorylation of the S6 ribosomal subunit by S6 kinase (2). This kinase is regulated by members of the PI3K family, including TOR, PDK1, and PI3K, which are activated by insulin receptor signaling (3, 4). These family members phosphorylate the translational inhibitor 4E-BP1, leading to dissociation of the initiation factor eIEF4E, which promotes cyclin D and Myc translation (5). In the absence of growth factors, these kinases are inactive and cannot signal progression from quiescence to G1. Acetylation and phosphorylation of the tumor suppressor p53 also seems to be involved in maintaining cellular quiescence (6, 7).


G1 phase

In the presence of growth factors during the Go and G1 phases, ras and mitogen-activated protein kinase (MAPK) cascades are activated and subsequently regulate cell cycle progression (8). MAPK regulates cyclin D expression directly by controlling the activation protein-1 and ETS transcription factors, which transactivate the cyclin D promoter (9, 10). Consequently, the MAPK cascade activates cyclin D-dependent kinases (CDK4 and CDK6) and regulates cell proliferation. Additionally, the MAPK cascade regulates directly the synthesis of the CIP/KIP family of CDK inhibitors (CKIs), specifically p21CIP and p27KIP, which regulate CDK activity negatively and influence cyclin D/CDK4/6 complex formation in G1 (11, 12). The growth factor-dependent synthesis of D-type cyclins occurs during the G0/G1 transition and peak in concentration in late G1 phase (13). These proteins have a very short half-life and are degraded rapidly after removal of mitogenic stimulation. The INK family of CKIs primarily inhibits cyclin D/CDK4/6 complexes. Only when the concentration of cyclin D exceeds that of the INK proteins can these cyclin D/CDK4/6 complexes overcome their inhibition (14, 15).

In early to mid G1 phase, active cyclin D/CDK4/6 complexes phosphorylate the three Rb pocket proteins (Rb, p130, and p107), which results in their partial repression (13). The phosphorylation status of these proteins controls E2F transcriptional activity and S-phase entry by mediating passage through the restriction point in late Gi (16-18). E2F proteins (E2F1-6) form heterodimers with a related family of DP proteins (DP1-3), and can act as both activators and repressors of transcriptional activity. In G0 and early G1, Rb is in an active, hypophosphorylated form. Active Rb represses the activity of the E2F transcription factor family by binding directly to the transactivation domain of E2F proteins and recruiting histone deacetylases, methyltransferases, and chromatin remodeling complexes to E2F-regulated promoters (19, 20). This activity results in the modification of histones, compaction of chromatin structure, and prevention of promoter access by transcription machinery (20). Phosphorylation of Rb by cyclin D/CDK4/6 complexes during G1 releases histone deacetylase, which alleviates transcriptional repression partially (13, 19, 20). As a result, the E2F/DP transcription factors activate the transcription of cyclin E and many genes responsible for the G1/S transition and DNA synthesis including CDK2, cyclin A, cyclin E, RPA1, MAT1, PCNA, DHFR, c-Myc, DNA polymerase-α, p220NPAT, and CDC25A (21).


G1/S transition

Cyclin E expression in mid to late G1 results in the formation of cyclin E/CDK2 complexes, which are required for S-phase entry and the initiation of DNA replication. Cyclin E/CDK2 also phosphorylates Rb, except on different residues than those catalyzed by cyclin D/CDK4/6 complexes (22). Cyclin E/CDK2 phosphorylation of Rb promotes the dissociation of E2F transcription factors from Rb, which results in complete relief of transcriptional repression (23). Thus, Rb inactivation occurs through the sequential phosphorylation by CDK4/6 and CDK2. Additional E2F and cyclin E/CDK2 activity increases through a positive feedback mechanism because cyclin E is one of the many genes activated by E2F (24). Cyclin E/CDK2 activity enhances this positive feedback even more by promoting the degradation of its own inhibitor, p27KIP. These complexes have been shown to phosphorylate p27KIP at T187, which promotes its association with the Skp-Cullin-F-boxSKP2 (SCFSKP2) complex to target p27KIP for ubiquitination and proteasomal degradation (25). Cyclin D/CDK4/6 complexes have been hypothesized to sequester the bound CKI inhibitor p27KIP away from cyclin E/CDK2 complexes to facilitate their activation (26). However, recently p27KIP was shown to be phosphorylated by Src-family tyrosine kinases at Y88, which reduces its steady-state binding to cyclin E/CDK2. This action facilitates p27KIP phosphorylation at T187 by cyclin E/CDK2 to promote its degradation (27, 28). Thus, rather than cyclin D/CDK4/6 sequestration of p27KIP, these tyrosine kinases may be responsible for activation of p27KIP-bound cyclin E/CDK2 complexes at the G1 /S transition.

The c-myc proto-oncogene encodes another transcription factor involved in many processes, which include E2F regulation (29). Its expression is induced by mitogenic stimulation, promotes S-phase entry in quiescent cells, and increases total cell mass. Myc activates the transcription of cyclin E, CDC25A, and several other genes (30). The Myc-induced proliferation mechanism activates cyclin E/CDK2 activity directly through increased cyclin E levels and CDC25A activity, which removes T14 and Y15 inhibitory CDK2 phosphorylation catalyzed by Wee1/Myt1 (31). Additionally, this activity is enhanced indirectly through Myc by mediating the sequestration of p27KIP from cyclin E/CDK2 into cyclin D/CDK4/6 complexes, which in turn promotes the cyclin E/CDK2 catalyzed phosphorylation and degradation of p27KIP (32). Cul-1, a component of the SCFSKP2 complex, was shown to be a transcriptional target of Myc, which may explain the link between p27KIP degradation and Myc activation (33).

Cyclin E/CDK2 also phosphorylates p220NPAT, which is a protein involved in the regulation of histone gene expression. This phosphorylation is a major event that occurs as cells begin to enter S phase (34). The phosphorylation of p220NPAT by cyclin E/CDK2 is required for histone gene expression activation at the onset of S phase (35). Once cells have passed through the restriction point, they are committed to initiate DNA synthesis and complete mitosis. Cell-cycle progression continues independently of the presence of growth factor stimulation after passage through the restriction point.


Regulation of DNA Synthesis and Mitotic Entry

S phase

At the G1/S transition, the cell enters S phase in which DNA synthesis occurs and each chromosome duplicates into two sister chromatids. During S-phase entry, the initiation of replication occurs at sites on chromosomes termed origins of replication. Replication origins are found in two states within cells: a pre-replicative complex (pre-RC) that is present in G1 before DNA replication initiation, and the the post-replicative complex (post-RC) that exists from the onset of S phase until the end of M phase (36). At the onset of S phase, an increase in cyclin A expression and cyclin A/CDK2 activity occurs (37, 38), and the protein kinase GSK-3P phosphorylates cyclin D and signals its relocalization to the cytoplasm where it is degraded by the proteasome (39, 40). Cyclin A/E/CDK2 activity controls each round of DNA replication, and this dictates the state of the replicative complexes. Low CDK activity permits the assembly of the pre-RC to form a licensed origin at the end of M phase, whereas the increase in CDK activity during the G1 /S transition triggers initiation of DNA replication and converts origins to the post-RC form (41). Reformation of the pre-RC is prevented by high CDK activity, which acts to inhibit re-replication events that would result in numerous copies of chromosomes.

The initiation of DNA replication requires both the assembly of the pre-RC complex at origins of replication and the activation of these complexes by CDKs and other kinases to initiate DNA synthesis (42-44). Numerous proteins are required for pre-RC formation and DNA replication initiation, which include the Origin Recognition Complex (ORC), cdc6/18, cdc45, cdtl, the GINS complex, and mini chromosome maintenance (MCM) proteins (43). ORC proteins (ORC1-6) bind directly to replication origins as a hexamer and facilitate the loading of other components of the pre-RC (45, 46). The cdc6/18 and cdt1 proteins play a central role in coordinating chromatin licensing. They bind directly to the ORC complex independently of each other (47). Here, they facilitate cooperatively the loading of the MCM proteins (MCM2-7), which form a hexameric ring-complex that possesses ATP-dependent helicase activity (48, 49). Cyclin E/CDK2 is recruited to replication origins through its interaction with cdc6, and this event regulates cdtl, cdc45, and MCM loading, which makes chromatin replication competent. After binding of the MCM proteins, the affinity of both cdc6/18 and cdtl for the ORC is reduced, and they dissociate (48, 49). Then, cyclin A/CDK2 phosphorylates cdc6 to promote its export from the nucleus and cdt1 to target its ubiq- uitination by the SCFSKP2 complex (50, 51). In this way, after initiation and release of these factors from the ORC, cyclin A/CDK2 activity acts to prevent re-replication by inhibiting reformation of the pre-RC. However, cyclin E/CDK2 activity acts primarily to promote the initiation of DNA synthesis (52).

Dbf4-dependent kinase (DDK) contains the kinase subunit cdc7, and it is required for DNA replication initiation (53). DDK targets MCMs for phosphorylation, thereby increasing the affinity of these proteins for cdc45, which is a factor required for the initiation and completion of DNA replication (49, 54, 55). The GINS complex, which consists of the four subunits Sld5, Psf1, Psf2, and Psf3, is required for the initiation and progression of eukaryotic DNA replication (56). This complex associates with Cdc45 and the MCM proteins to activate their helicase activity. As a result of GINS and cdc45 binding to the MCM complex, the DNA is unwound, which results in single stranded DNA (ssDNA) (49, 57). Replication protein A (RPA) is recruited to single stranded DNA, and it is required for the subsequent binding and activation of DNA polymerase-a (58-60). The GINS complex also interacts with, and stimulates the polymerase activity of the DNA polymerase-α-primase complex (61).


G2 phase

After completion of DNA duplication, the cell enters the second restriction point of the cell cycle; referred to as the G2 phase. Similar to what happens during G1, in this second gap phase the cell halts to synthesize factors required for initiation and completion of mitosis and to check for any aberrations that result from DNA synthesis (62, 63).

Cyclin B/CDK1 is the primary regulator of the G2/M transition, and its activity is required for entry into mitosis. It was termed the maturation-promoting factor (MPF) because it was originally shown to be essential for Xenopus oocytes maturation after hormonal stimulation, and it was found subsequently to be equivalent to a mitosis-promoting activity (64). CDK1 activity is regulated primarily by localization of cyclin B, CDC25C activity, and p21CIP levels, which are controlled by checkpoint machinery (65). Cyclin B/CDK1 complexes remain inactive until their activity is required for mitosis entry in late G2. Toward the end of S phase, cyclin B expression is increased. However, during the onset of G2, cyclin B is retained in the cytoplasm by its cytoplasmic retention signal (CRS), and the CKI p21CIPinhibits CAK-mediated activation of cyclin/CDKs (66). Additionally, Wee1 and Myt1 phosphorylate T14 and Y15 on cyclin B/CDK1 in the cytoplasm to keep these complexes inactive even when CDK1 is phosphorylated by CAK (67). The transcription factor p53 also mediates the inhibition of cyclin B/CDK1 activity by promoting p21 expression, and it down-regulates expression of CDK1 (63, 68). Furthermore, cyclin A/CDK2 phosphorylates and inactivates members of the E2F transcription family in G2 to suppress cell growth during this gap (69-71).


G2/M transition

During the G2/M transition, the localization of cyclin B changes dramatically and regulates CDK1 activity (72). The CRS is phosphorylated by MAPK and polo-like kinase 1 (Plk1), which promotes its nuclear translocation (73, 74). Contomitant with nuclear import, cyclin B is phosphorylated to a greater extent to prevent association with CRM1, which promotes its nuclear retention (75, 76). This relocalization occurs at the onset of mitosis toward the end of the G2/M transition when the cell is ready to begin the mitotic process (77). Activation of cyclin B/CDK1 in late G2 is achieved by preventing the access of cytoplasmic Wee1/Myt1 kinases to the complex and by promoting shuttling of the CDC25 phosphatases to the nucleus, where they dephosphorylate and activate CDK1 (78-80). Cyclin B/CDK1 complexes phosphorylate CDC25A to promote its stability and CDC25C to promote its activity (81). Both CDC25A and CDC25C activate CDK1 even more, which results in a positive feedback loop that sustains cyclin B/CDK1 activity in the nucleus to signal mitotic entry (82, 83). ERK-MAP kinases also regulate cyclin B/CDK1 activity by phosphorylating CDC25C at T48 (84). ERK1/2 activation of CDC25C leads to removal of inhibitory phosphorylations of cyclin B/CDK1 complexes and is required for efficient mitotic induction. Thus, MAPKs are involved in the positive feedback loop that leads to cyclin B/CDK1 activation.

The increase in nuclear cyclin B/CDK1 activity promotes phosphorylation of nuclear substrates that are necessary for mitosis, such as nuclear envelope breakdown, spindle formation, chromatin condensation, and restructuring of the Golgi and endoplasmic reticulum (85, 86). Numerous cyclin B/CDK1 substrates have been defined, which include nuclear lamins, nucleolar proteins, centrosomal proteins, components of the nuclear pore complex, and microtubule-associated proteins (87-89). Cyclin B/CDK1 complexes also phosphorylate MCM4 to block replication of DNA, the TFIIH subunit of RNA polymerase II to inhibit transcription, and the ribosomal S6 protein kinase to prevent translation during mitosis (90-92).


Regulation of Cell Division

The centrosome

Normally, the centrosome is composed of two centrioles and the pericentriolar material. It functions not only as a microtubule nucleation center, but also as an integrated regulator of cell-cycle checkpoints. Recent data indicates it is required for cell-cycle progression (93). The centrosome duplication process begins in late G1 and is regulated primarily by CDK2 activity (94). Cyclin A/E/CDK2 phosphorylates the Mps1p kinase and nucleophosmin, which are two centrosome associated proteins. CDK2 activity is required for Mps1p stability and Mps1p-dependent centrosome duplication (95). Cyclin E/CDK2 phosphorylates nucleophosmin at T199, releasing it from unduplicated centrosomes, which are a requirement for centrosome duplication (96). Completion of centrosome duplication and initiation of their separation occur in G2 and are dependent on cyclinA/E/CDK2 activity. These processes are necessary for proper spindle formation and for balanced chromosome separation during mitosis.

The Aurora kinase family members play a role in centrosome function, spindle assembly, and chromosome alignment, and they are essential for mitosis. Specifically, Aurora-A activity is maximal during G2/M; it regulates mitotic spindle assembly, centrosome separation, and it facilitates the G2/M transition by phosphorylating CDC25B at the centrosome, which is an important event for cyclin B localization to the nucleus (97). Aurora-B activity is maximal from metaphase to the end of mitosis and regulates chromatin protein modification, chromatid separation, and cytokinesis (98). During mitosis, a complex process of degradation and phosphorylation regulates Aurora kinase activity to ensure proper mitotic advancement. Aurora-A is activated mainly by autophosphorylation (99), Ajuba (100), TPX2 (101), and HEF1 (102), whereas INCENP is thought to activate Aurora-B (103). Both Aurora-A and B are degraded rapidly at the end of mitosis.


M phase

The mitotic phase is divided into five phases, which include prophase, prometaphase, metaphase, anaphase, and telophase. During prophase, nucleoli disappear, chromatin condensation takes place, and the mitotic spindle is formed at centrosomes that contain centrioles. In prometaphase, fragmentation of the nuclear envelope occurs and mitotic spindles extend from the poles toward the center of the cell. At metaphase, centrioles pair at opposite poles, and the chromosomes align in the cell center along the metaphase plate. Then, microtubules bind to the kinetochores located at the centromeres of each chromatid of the chromosomes. The transition from metaphase to anaphase is triggered by MPF inactivation through the degradation of cyclin B by the E3 ubiquitin ligase anaphase-promoting complex (APC/C) (104). Cdc20 is required for activation of the ubiquitin ligase activity of APC/C, which promotes degradation of securin. Subsequently, a release mechanism activates the protease known as separase, which cleaves cohesion and promotes sister chromatid separation and anaphase entry (105, 106). This mechanism induces the separation of chromatids in anaphase as microtubules from each pole pull them apart through their kinetochore. Because of cyclin-B/Cdk1 inactivation in late anaphase, the major ubiquitin ligase activity is switched from APC/C-Cdc20 to APC/C-Cdh1. The latter continues to regulate many proteins whose degradation is required for cell-cycle progression, including Cdc20, which becomes one of its targets and a substrate of the Aurora kinases. (107-111).

In telophase, nuclei for each daughter cell form at the two poles, and the mitotic spindle apparatus disappears. Furthermore, nuclear membranes, nuclear lamina, nuclear pores, and nucleoli are reformed. The cell is now ready for cytokinesis, which is physical division of the cytoplasm. The cytoplasm divides as actin/myosin filaments contract and pinch off the plasma membrane, which results in two daughter cells that enter into G0 or G1 for another round of division. The main checkpoint that exists during M phase in mammalian cells is the spindle checkpoint; it is in place to ensure proper microtubule assembly, proper cell division, and that each daughter cell receives one copy of DNA.


Spindle checkpoint

The spindle checkpoint is activated when microtubules fail to attach to the kinetochores of each sister chromatid and/or when misalignment of chromosomes occurs along the metaphase plate (112-114). This mechanism blocks entry into anaphase and ensures proper segregation of the chromatids to opposite spindle poles. Misregulation of this checkpoint results in aneuploid daughter cells after division (115, 116). Checkpoint proteins associated with kinetochores monitor microtubule-kinetochore attachment and tension; these proteins regulate this checkpoint by preventing cdc20 binding to the APC/C (117-119).

The main spindle checkpoint proteins include Mad1, Mad2, BubR1, Bub1, Bub3, Mps1p, and CENP-E. These proteins act both independently and dependently of their interaction with kinetochores. Association of Mad2 with kinetochores and cdc20 requires the presence of Mad1 (120). At the kine- tochore, Mad2 is converted to a form that can bind and sequester cdc20 away from the APC/C, which results in its inhibition (121). Additionally, formation of the mitotic checkpoint complex BubR1/Bub3/Mad2/cdc20 occurs independently of interaction with unattached kinetochores, and it signals anaphase to wait by binding and inhibiting the APC/C (122, 123). An unattached kinetochore activates a kinase cascade that involves the dual-specificity kinase Mps1p and the serine/threonine kinases BubR1 and Bub1 that amplifies this wait signal (124, 125). Furthermore, BubR1 interacts directly with the kinesin-like protein CENP-E to regulate microtubule tension at kinetochores, which is also involved in regulation of the spindle checkpoint (126, 127). Thus, this checkpoint serves to inhibit the APC/C indirectly through cdc20 sequestration and directly through association with the mitotic checkpoint complex, and to regulate the tension at kinetochores required for anaphase entry (128).


DNA Damage Checkpoints

In addition to checkpoints that ensure normal cell-cycle progression, numerous DNA damage checkpoints exist in mammalian cells. These checkpoints exist to regulate the highly conserved mechanisms that control DNA replication and mitosis to ensure mutations within the genome are not passed on to the daughter cells. Misregulation of these pathways is associated with genomic instability and cancer development. The key players involved in the DNA damage checkpoint cascade (Fig. 2) include the DNA damage sensors ATM (Ataxia Telangiectasia Mutated), ATR (ATM and Rad3 Related), Rad1, Rad9, Hus1, and ATRIP, and the effectors Chk1/2 (Checkpoint Kinase 1/2), and CDC25.



Figure 2. Brief model of DNA damage checkpoint signaling. DNA damage elicits a conserved response headed by the ATM and ATR kinases. Phosphorylation cascades and localization of mediators to sites of damage allows for signaling to the effector kinases Chk1 and Chk2. Chk1/2 elicit cell-cycle arrest through phosphorylation-dependent degradation of the Cdc25 family of phosphatases. Parallel activation of p53 by both ATM/ATR and Chk1/Chk2 leads to upregulation of the CDK inhibitor p21, which enforces cell-cycle arrest to a greater extent. See text for in-depth discussion of the checkpoint pathways.


G1/S-phase checkpoint

The primary DNA damage checkpoint is the G1/S checkpoint, which acts to prevent the replication initiation of damaged DNA. During G1 and even after passage through the restriction point (but prior to initiation of DNA synthesis), DNA damage activates two checkpoint-signaling pathways sequentially, and both pathways function to inhibit CDK2 activity. The first pathway initiated is p53-independent and is rapid and short-lived (129). This pathway results in phosphorylation and degradation of CDC25A (130, 131). DNA damage leads to the activation of ATM and ATR, which phosphorylate and activate Chk1 and Chk2 (132, 133). CDC25A is phosphorylated by these kinases, which target its ubiquitination and proteasomal degradation (134). As a result, the inhibitory phosphorylations of CDK2 are increased, which diminishes CDK2 activity. Ultimately, this lack of CDK2 activity inhibits the cdc45 loading to pre-RCs and the subsequent initiation of DNA replication to halt the cell cycle and to allow time to repair damaged DNA (133).

The second pathway activated in the presence of DNA damage prior to initiation of DNA synthesis acts in a p53-dependent manner. As stated above, the tumor suppressor p53 is a transcription factor, which acts primarily to increase expression of the CKI p21CIP during DNA damage. Like CDC25A, the activation of ATM/ATR promotes the phosphorylation of p53, which enhances the stability of p53 by preventing efficient interaction with the E3 ubiquitin ligase MDM2, which is a protein responsible for targeting p53 degradation (135). This mechanism leads to the transcription and accumulation of p21, which silences CDK2 activity to prevent cell-cycle progression and to allow for DNA repair (136). MDM2 is also a target of p53 transcription, which creates a negative feedback loop with p53 (137). After repair of damaged DNA has been completed, the checkpoint is turned off and progression into S phase resumes.


S-phase checkpoints

Cells that have passed the G1/S checkpoint are ready to begin S phase and DNA replication. The S-phase checkpoints are a group of three mechanistically distinguishable checkpoints (138) of which two respond directly to DNA damage. One is independent of ongoing replication and is activated in response to DNA double-stranded breaks (DSBs), which is known as the intra-S-phase checkpoint. The second checkpoint, the replication checkpoint, responds to replication fork stalling caused by the collision of replication machinery with DNA damage, direct inhibition of polymerases, or depletion of dNTPs. Although these two checkpoints respond to different forms of stress, both checkpoints prevent cell-cycle advance, inhibit ongoing replication, prevent origin firing, and stabilize the replication fork so that repair and replication resumption can occur. The third type of S-phase checkpoint is the S/M checkpoint. Currently, this checkpoint is not understood as well as the previous two, but it is known to prevent entry into mitosis when replication is stalled or incomplete. It acts to preserve genomic stability by preventing premature chromatin condensation and breaks at common fragile sites.


The replication checkpoint

The replication checkpoint is activated when the replication machinery encounters DNA damage or when the replicative polymerase is inhibited and stalls (139, 140). This checkpoint stabilizes stalled replication forks and signals for DNA damage repair while preventing exit from S phase. Stalling causes uncoupling of the helicase from the polymerase, which leads to DNA unwinding without subsequent new strand polymerization. This action leads to accumulation of ssDNA, which is a trigger for checkpoint activation (141-143). ssDNA is also believed to activate other checkpoints, which include those initiated by DNA repair mechanisms such as nucleotide excision repair (144, 145) or recession of DSBs generated during homologous recombination (146, 147). The ssDNA is coated by RPA proteins (148, 149), which set up a scaffold for the recruitment and localization of DNA damage sensors in S phase. ATR is central to the replication checkpoint and is recruited to RPA coated ssDNA through its interaction with its binding partner, ATRIP (150-152). In addition, other sensors of DNA damage, including Rad17, which is an RFC-like clamp loader, and the 9-1-1 complex, which is a heterotrimeric clamp composed of Rad9, Rad1, and Hus1, are recruited to RPA coated ssDNA and serve to activate ATR and to help recruit and activate downstream mediators of the checkpoint (153-155).

After ATR activation and recruitment/activation of other sensors, numerous proteins are recruited to the site of damage and act as mediators of the DNA damage-signaling cascade. Most mediators are involved in the activation of the effecter kinase Chk1 (156). One mediator, Claspin, is recruited to sites of damage, is phosphorylated by ATR, and recruits Chk1 subsequently. Direct interaction between Claspin and Chk1 is required for phosphorylation and activation by ATR (157-160). Other mediators include BRCA1 and BRCA1C-terminal motif (BRCT)-containing proteins. These mediators form large multimeric complexes and are often visualized as nuclear foci by immunofluorescence microscopy (156, 161). MDC1 (Mediator of DNA damage-checkpoint protein 1) recruits mediators of the checkpoint, such as 53BP1 and NBS1 (162-164). These proteins function to maintain foci oligomerization and to promote ATR mediated phosphorylation of its substrates, which include all of these mediators and SMC1 (Structural maintenance of chromosomes 1). SMC1 is part of the cohesin complex and is required for sister chromatid cohesion in S phase (165, 166).

Finally, Chk1 is recruited to the nuclear foci that contain the large scaffold of BRCT-containing proteins and is activated in an ATR/Rad17/9-1-1/BRCA1/Claspin dependent fashion (157-159). Then, Chk1 facilitates the checkpoint by phosphorylating CDC25 family members (167) and p53 (see above for more detail on these events); this results in cell-cycle arrest, DNA repair, and survival choices.


The intra-S-phase checkpoint

Unlike the replication checkpoint, the intra-S-phase checkpoint does not require replication to be activated (138, 168). At the head of this checkpoint is the ATM protein kinase, which is a member of the PI3K family of protein kinases (including ATR and DNA-PK). ATM and the intra-S-phase checkpoint are activated by the detection of DSBs, which can be achieved without direct interaction of the replication machinery with sites of damage. Another interesting difference between the replication checkpoint and the intra-S-phase checkpoint is that activation of the latter does not alter the progression of active replication units, only the inhibition of late origin firing (169). Thus, the intra-S-phase checkpoint causes delays in, but not complete arrest of, S-phase progression (138).

Although the sensors of DSBs are not definitively known, two protein complexes serve as excellent candidates because of their ability to enhance ATM activity. These complexes are the MRN (Mre11-Nbs1-Rad50) complex and the Rad17/9-1-1 complex (discussed above). The MRN complex has nuclease activity and localizes to DSBs independently of ATM. At sites of damage, it plays a role in activation of ATM, efficient phosphorylation of ATM substrates, and recession of DSBs (170-172). Although much of the checkpoint from here out involves the same mediators including 53 BP1, BRCA1, MDC1, and SMC1, it has two more distinct features compared with the replication checkpoint.

The first feature involves the recession of DSBs, which activates a parallel ATR/ssDNA signaling cascade similar to that discussed above (146, 173, 174). The second feature involves the activation of Chk2. Unlike Chk1, which is only present in S and G2 phases, Chk2 is present throughout the entire cell cycle (175, 176). Chk2 also differs from Chk1 in that it must dimerize to be fully active (177-179), and in response to DNA damage, it becomes soluble in the nucleus and dissipates from damage sites as a mechanism to enhance signaling (180, 181). When phosphorylated by ATM, Chk2 plays similar roles as Chk1 in the degradation of CDC25 family members and phosphorylation of p53.

Although the replication and intra-S-phase checkpoints have distinct mechanisms of activation and signaling, the final goal is the same: to delay or to inhibit S-phase progression providing time and signaling events that lead to DNA repair, so that mutations are not transmitted to daughter cells in the ensuing mitotic division.


S/M checkpoint

The S/M checkpoint can be activated by replication inhibition or when DNA replication is not completed (182-186). This checkpoint signals through the ATR/Chk1 pathways and prevents premature chromatin condensation (PCC) and entry into mitosis (183, 185, 187). Depletion of ATR in Xenopus egg extracts or Chk1 in embryonic stem cells results in premature entry into mitosis prior to completion of replication (183, 185). In addition, different regions of the genome replicate at different rates, and common fragile sites are known to be late replicating regions. These common fragile sites are often left unreplicated during mitotic entry (188-191). PCC causes breaks when fragile sites are not fully replicated (189). Therefore, mitotic delay is required to ensure the proper replication of the entire genome to prevent breaks that might occur because of PCC. Both ATR (187) and Chk1 (188) are involved in the stability of common fragile sites, which indicates that the S/M checkpoint is required to maintain genomic stability by ensuring proper replication prior to mitotic entry.


G2/M-phase checkpoint

The G2/M checkpoint acts to ensure that cells that experience DNA damage in G2 or that contain unresolved damage from the previous G1 or S phase do not initiate mitosis. Much like the G1checkpoint and in contrast to the S checkpoints, cell-cycle arrest or delay that results from the G2 checkpoint involves a combination of acute/transient and delayed/sustained mechanisms. The acute/transient mechanisms involve the rapid posttranslational modification of effector proteins, whereas the delayed/sustained mechanism involves the alteration of transcriptional programs (192).

Of all molecules targeted in the G2/M checkpoint, cyclin B/CDK1 seems to be the most important as its activity stimulates mitotic entry directly. DNA damage in the G2 phase activates ATM/ATR pathways (as described above), which results in Chk1/Chk2-mediated inhibition of the Cdc25C phosphatase that would normally activate CDK1 and trigger transition through the G2/M boundary. In G2, Cdc25B is also targeted for degradation by Chk1 and Chk2 via the mechanisms described above, and it is the only known mechanism of cell-cycle arrest that is shared across all checkpoints. Cdc25 degradation is one of the key mechanisms of the acute/transient branch of the checkpoint.

The more delayed and prolonged mechanisms by which the checkpoint silences CDK1 activity is through the activation of the p53 pathway. Activation of p53 is achieved by phosphorylation by ATM/ATR or Chk1/Chk2 and results in nuclear localization, tetramerization, and stimulation of p53 transcriptional activity toward p21CIP. In G2, BRCA1 can stimulate p21 expression in a p53 independent fashion (193); along with two other p53 targets, GADD45 and 14-3-3e, BRCA1 may cooperate to achieve maximal inhibition of CDK1 and to prevent mitotic entry to allow for repair of DNA lesions (68).

The centrosome also regulates the G2/M DNA damage response, and numerous checkpoint proteins are associated with the centrosome (194). Centrosome separation is regulated by the kinases Nek2 and Plk1, and this process is inhibited by DNA damage in an ATM-dependent manner. ATM activation leads to Plk1 and Nek2 inhibition, which results in deregulation of the centrosome (195). By this mechanism, centrosome separation is inhibited, and it contributes to maintaining the G2/M checkpoint (196). Plk1 is also known to phosphorylate and activate CDC25C (197). Thus, Plk1 inhibition also results in CDC25C inhibition, inactivation of cyclin B/CDK1, and a halt in cell-cycle progression.

Normally, cell-cycle progression resumes when DNA damage repair is completed; otherwise, apoptosis prevents genomic instability if the damage is excessive and beyond repair. However, data from Saccharomyces cerevisiae, Xenopus, and human cells, suggests that pathways to re-enter cell-cycle progression exist even when unrepaired DNA damage is present. This process of “checkpoint adaptation” has been shown to allow mitotic entry in response to ionizing radiation in human cells in a Plk1 dependent manner, and it may promote carcinogenesis and genomic instability (198, 199). It has been speculated that activation of centrosomal cyclin B/CDK1 plays a central role in this process, and it may occur through Plk1 mediated degradation of Wee1 and/or inhibition of Chk1 activity that leads to stabilization of CDC25 (200). Although its function is not well understood, checkpoint adaptation has been proposed to move cells into a phase where they can die, allow progression into other phases where difficult DNA damage can be repaired, and even exist to allow natural evolution (201).



The mammalian cell cycle is controlled by numerous factors involved in regulation of CDKs and checkpoint responses. Although many proteins involved in the pathways that lead to activation or inactivation of these have been elucidated over the years, much remains to be explored. Although most CDKs control the cell division cycle, regulation of the cell cycle is clearly more than progression from growth to DNA synthesis to division and transmission of genetic material. Growing evidence exists for the role of CDKs in controlling the balance between senescence, cell growth, checkpoint activation, and apoptotic signaling. Clearly, the inability to respond properly to DNA damage and cellular stress through checkpoint activation and apoptosis has a role in oncogenic potential and therapeutic considerations. The identification of novel factors and signal cascades that mediate the regulation of the cell cycle will lead to new drug targets in the fight against cancer and numerous other diseases.



1. Geyer PK, Meyuhas O, Perry RP, Johnson LF. Regulation of ribosomal protein mRNA content and translation in growth- stimulated mouse fibroblasts. Mol. Cell Biol. 1982; 2:685-693.

2. Dufner A, Thomas G. Ribosomal S6 kinase signaling and the control of translation. Exp. Cell. Res. 1999; 253:100-109.

3. Long X, Muller F, Avruch J. TOR action in mammalian cells and in Caenorhabditis elegans. Curr. Top. Microbiol. Immunol. 2004; 279:115-138.

4. Um SH, D’Alessio D, Thomas G. Nutrient overload, insulin resistance, and ribosomal protein S6 kinase 1, S6K1. Cell. Metab. 2006; 3:393-402.

5. Costa LJ. Aspects of mTOR biology and the use of mTOR inhibitors in non-Hodgkin’s lymphoma. Cancer Treat. Rev. 2007; 33:78-84.

6. Brooks CL, Gu W. Ubiquitination, phosphorylation and acetylation: the molecular basis for p53 regulation. Curr. Opin. Cell Biol. 2003; 15:164-171.

7. Knights CD, Catania J, Di Giovanni S, Muratoglu S, Perez R, Swartzbeck A, Quong AA, Zhang X, Beerman T, Pestell RG, Avantaggiati ML. Distinct p53 acetylation cassettes differentially influence gene-expression patterns and cell fat e. J. Cell Biol. 2006; 173:533-544.

8. Torii S, Yamamoto T, Tsuchiya Y, Nishida E. ERK MAP kinase in G cell cycle progression and cancer. Cancer Sci. 2006; 97:697-702.

9. Shaulian E, Karin M. AP-1 in cell proliferation and survival. Oncogene 2001; 20:2390-2400.

10. Shen Q, Uray IP, Li Y, Krisko TI, Strecker TE, Kim HT, Brown PH. The AP-1 transcription factor regulates breast cancer cell growth via cyclins and E2F factors. Oncogene. In Press.

11. Milde-Langosch K, Bamberger AM, Methner C, Rieck G, Loning T. Expression of cell cycle-regulatory proteins rb, p16/MTS1, p27/KIP1, p21/WAF1, cyclin D1 and cyclin E in breast cancer: correlations with expression of activating protein-1 family members. In t. J. Cancer 2000; 87:468-472.

12. Sherr CJ, Roberts JM. CDK inhibitors: positive and negative regulators of G1-phase progression. Genes Dev. 1999; 13:1501-1512.

13. Coqueret O. Linking cyclins to transcriptional control. Gene 2002; 299:35-55.

14. Ekholm SV, Reed SI. Regulation of G(1) cyclin-dependent kinases in the mammalian cell cycle. Curr. Opin. Cell Biol. 2000; 12:676-684.

15. Ohtani N, Yamakoshi K, Takahashi A, Hara E. The p16INK4a- RB pathway: molecular link between cellular senescence and tumor suppression. J. Med. Invest. 2004; 51:146-153.

16. Weinberg RA. The retinoblastoma protein and cell cycle control. Cell 1995; 81:323-330.

17. Harbour JW, Dean DC. The Rb/E2F pathway: expanding roles and emerging paradigms. Genes Dev. 2000; 14:2393-2409.

18. Sherr CJ. The Pezcoller lecture: cancer cell cycles revisited. Cancer Res. 2000; 60:3689-3695.

19. Harbour JW, Dean DC. Chromatin remodeling and Rb activity. Curr. Opin. Cell Biol. 2000; 12:685-689.

20. Wang C, Fu M, Mani S, Wadler S, Senderowicz AM, Pestell RG. Histone acetylation and the cell-cycle in cancer. Front. Biosci. 2001; 6:D610-629.

21. Muller H, Bracken AP, Vernell R, Moroni MC, Christians F, Grassilli E, Prosperini E, Vigo E, Oliner JD, Helin K. E2Fs regulate the expression of genes involved in differentiation, development, proliferation, and apoptosis. Genes Dev. 2001; 15:267-285.

22. Lundberg AS, Weinberg RA. Functional inactivation of the retinoblastoma protein requires sequential modification by at least two distinct cyclin-cdk complexes. Mol. Cell Biol. 1991; 8:753-761.

23. Dynlacht BD, Flores O, Lees JA, Harlow E. Differential regulation of E2F transactivation by cyclin/cdk2 complexes. Genes Dev. 1994; 8:1772-1786.

24. Geng Y, Eaton EN, Picon M, Roberts JM, Lundberg AS, Gifford A, Sardet C, Weinberg RA. Regulation of cyclin E transcription by E2Fs and retinoblastoma protein. Oncogene 1996; 12:1173-1180.

25. Nakayama, K.I., S. Hatakeyama, and K. Nakayama. 2001. Regulation of the cell cycle at the G1-S transition by proteolysis of cyclin E and p27Kip1. Biochem Biophys Res Commun 282:853-860.

26. Soos TJ, Kiyokawa H, Yan JS, Rubin MS, Giordano A, DeBlasio A, Bottega S, Wong B, Mendelsohn J, Koff A. Formation of p27-CDK complexes during the human mitotic cell cycle. Cell Growth Differ. 1996; 7:135-146.

27. Chu I, Sun J, Arnaout A, Kahn H, Hanna W, Narod S, Sun P, Tan CK, Hengst L, Slingerland J. p27 phosphorylation by Src regulates inhibition of cyclin E-Cdk2. Cell 2007; 128:281-294.

28. Grimmler M, Wang Y, Mund T, Cilensek Z, Keidel EM, Waddell MB, Jakel H, Kullmann M, Kriwacki RW, Hengst L. Cdk-inhibitory activity and stability of p27Kip1 are directly regulated by oncogenic tyrosine kinases. Cell 2007; 128:269-280.

29. Obaya AJ, Mateyak MK, Sedivy JM. Mysterious liaisons: the relationship between c-Myc and the cell cycle. Oncogene 1999; 18:2934-2941.

30. Leone G, DeGregori J, Sears R, Jakoi L, Nevins Jr. Myc and Ras collaborate in inducing accumulation of active cyclin E/Cdk2 and E2F. Nature 1997; 387:422-426.

31. Berns K, Hijmans EM, Bernards R. Repression of c-Myc responsive genes in cycling cells causes G1 arrest through reduction of cyclin E/CDK2 kinase activity. Oncogene 1997; 15:1347-1356.

32. Muller D, Bouchard C, Rudolph B, Steiner P, Stuckmann I, Saffrich R, Ansorge W, Huttner W, Eilers M. Cdk2-dependent phosphorylation of p27 facilitates its Myc-induced release from cyclin E/cdk2 complexes. Oncogene 1997; 15:2561-2576.

33. O’Hagan RC, Ohh M, David G, de Alboran IM, Alt FW, Kaelin Jr WG, DePinho RA. 2000. Myc-enhanced expression of Cul1 promotes ubiquitin-dependent proteolysis and cell cycle progression. Genes Dev 14:2185-2191.

34. Zhao J, Kennedy BK, Lawrence BD, Barbie DA, Matera AG, Fletcher JA, Harlow E. NPAT links cyclin E-Cdk2 to the regulation of replication-dependent histone gene transcription. Genes Dev. 2000; 14:2283-2297.

35. Stein GS, van Wijnen AJ, Stein JL, Lian JB, Montecino M, Zaidi SK, Braastad C. An architectural perspective of cell-cycle control at the G1/S phase cell-cycle transition. J. Cell Physiol. 2006; 209:706-710.

36. Zannis-Hadjopoulos M, Sibani S, Price GB. Eucaryotic replication origin binding proteins. Front. Biosci. 2004; 9:2133-2143.

37. Desdouets C, Sobczak-Thepot J, Murphy M, Brechot C. Cyclin A: function and expression during cell proliferation. Prog. Cell Cycle Res. 1995; 1:115-123.

38. Yam CH, Fung TK, Poon RY. Cyclin A in cell cycle control and cancer. Cell Mol. Life Sci. 2002; 59:1317-1326.

39. Benzeno S, Lu F, Guo M, Barbash O, Zhang F, Herman JG, Klein PS, Rustgi A, Diehl JA. Identification of mutations that disrupt phosphorylation-dependent nuclear export of cyclin D1. Oncogene 2006; 25:6291-6303.

40. Diehl JA, Cheng M, Roussel MF, Sherr CJ. Glycogen synthase kinase-3beta regulates cyclin D1 proteolysis and subcellular localization. Genes Dev. 1998; 12:3499-3511.

41. Diffley JF. Regulation of early events in chromosome replication. Curr. Biol. 2004. 14:R778-786.

42. Stillman B. Cell cycle control of DNA replication. Science 1996; 274:1659-1664.

43. Teer JK, Dutta A. Regulation of S phase. Results Probl. Cell Differ. 2006; 42:31-63.

44. Ritzi M, Knippers R. Initiation of genome replication: assembly and disassembly of replication-competent chromatin. Gene 2000; 245:13-20.

45. Da-Silva LF, Duncker BP. ORC function in late G1: maintaining the license for DNA replication. Cell Cycle 2007; 6:128-130.

46. DePamphilis ML. The ‘ORC cycle’: a novel pathway for regulating eukaryotic DNA replication. Gene 2003; 310:1-15.

47. Hua XH, Newport J. Identification of a preinitiation step in DNA replication that is independent of origin recognition complex and cdc6, but dependent on cdk2. J Cell Biol 1998; 140:271-281.

48. Maiorano D, Lutzmann M, Mechali M. MCM proteins and DNA replication. Curr. Opin. Cell Biol. 2006; 18:130-136.

49. Lei M, Tye BK. Initiating DNA synthesis: from recruiting to activating the MCM complex. J. Cell. Sci. 2001; 114:1447-1454.

50. Liu, E., X. Li, F. Yan, Q. Zhao, and X. Wu. Cyclin-dependent kinases phosphorylate human Cdt1 and induce its degradation. J. Biol. Chem. 2004; 279:17283-17288.

51. Petersen BO, Lukas J, Sorensen CS, Bartek J, Helin K. Phosphorylation of mammalian CDC6 by cyclin A/CDK2 regulates its subcellular localization. EMBO J. 1999; 18:396-410.

52. Woo, R.A., and R.Y. Poon. 2003. Cyclin-dependent kinases and S phase control in mammalian cells. Cell Cycle 2:316-324.

53. Jiang, W., D. McDonald, T.J. Hope, and T. Hunter. 1999. Mammalian Cdc7-Dbf4 protein kinase complex is essential for initiation of DNA replication. Embo J 18:5703-5713.

54. Masai H, Taniyama C, Ogino K, Matsui E, Kakusho N, Matsumoto S, Kim JM, Ishii A, Tanaka T, Kobayashi T, Tamai K, Ohtani K, Arai K. Phosphorylation of MCM4 by Cdc7 kinase facilitates its interaction with Cdc45 on the chromatin. J. Biol. Chem. 2006; 281:39249-39261.

55. Sheu YJ, Stillman B. Cdc7-Dbf4 phosphorylates MCM proteins via a docking site-mediated mechanism to promote S phase progression. Mol. Cell 2006; 24:101-113.

56. Chang YP, Wang G, Bermudez V, Hurwitz J, Chen XS. Crystal structure of the GINS complex and functional insights into its role in DNA replication. Proc. Natl. Acad. Sci. U.S.A. 2007; 104:12685-12690.

57. Zou L, Stillman B. Formation of a preinitiation complex by S-phase cyclin CDK-dependent loading of Cdc45p onto chromatin. Science 1998; 280:593-596.

58. Sharova, N.P., and E.B. Abramova. Initiation of DNA replication in eukaryotes is an intriguing cascade of protein interactions. Biochemistry 2002; 67:1217-1223.

59. Walter J, Newport J. Initiation of eukaryotic DNA replication: origin unwinding and sequential chromatin association of Cdc45, RPA, and DNA polymerase alpha. Mol. Cell 2000; 5:617-627.

60. Zou L, Stillman B. Assembly of a complex containing Cdc45p, replication protein A, and Mcm2p at replication origins controlled by S-phase cyclin-dependent kinases and Cdc7p-Dbf4p kinase. Mol. Cell Biol. 2000; 20:3086-3096.

61. De Falco M, Ferrari E, De Felice M, Rossi M, Hubscher U, Pisani FM. The human GINS complex binds to and specifically stimulates human DNA polymerase alpha-primase. EMBO Rep. 2007; 8:99-103.

62. Stark GR, Taylor WR. Analyzing the G2/M checkpoint. Methods Mol. Biol. 2004; 280:51-82.

63. Stark GR, Taylor WR. Control of the G2/M transition. Mol. Biotechnol. 2006; 32:227-248.

64. Pines J. Regulation of the G2 to M transition. Results Probl. Cell. Differ. 1998; 22:57-78.

65. Jackman MR, Pines JN. Cyclins and the G2/M transition. Cancer Surv. 1997; 29:47-73.

66. Aprelikova O, Xiong Y, Liu ET. Both p16 and p21 families of cyclin-dependent kinase (CDK) inhibitors block the phosphorylation of cyclin-dependent kinases by the CDK-activating kinase. J. Biol. Chem. 1995; 270:18195-18197.

67. Fattaey A, Booher RN. Myt1: a Wee1-type kinase that phosphorylates Cdc2 on residue Thr14. Prog. Cell Cycle Res. 1997; 3:233-240.

68. Taylor WR, Stark GR. Regulation of the G2/M transition by p53. Oncogene 2001; 20:1803-1815.

69. Dynlacht BD, Moberg K, Lees JA, Harlow E, Zhu L. Specific regulation of E2F family members by cyclin-dependent kinases. Mol. Cell. Biol. 1997; 17:3867-3875.

70. Kitagawa M, Higashi H, Suzuki-Takahashi I, Segawa K, Hanks SK, Taya Y, Nishimura S, Okuyama A. Phosphorylation of E2F-1 by cyclin A-cdk2. Oncogene 1995; 10:229-236.

71. Xu M, Sheppard KA, Peng CY, Yee AS, Piwnica-Worms H. Cyclin A/CDK2 binds directly to E2F-1 and inhibits the DNA-binding activity of E2F-1/DP-1 by phosphorylation. Mol. Cell. Biol. 1994; 14:8420-8431.

72. Porter LA, Donoghue DJ. Cyclin B1 and CDK1: nuclear localization and upstream regulators. Prog. Cell Cycle Res. 2003; 5:335-347.

73. Li J, Meyer AN, Donoghue DJ. Nuclear localization of cyclin B1 mediates its biological activity and is regulated by phosphorylation. Proc. Natl. Acad. Sci. U.S.A. 1997; 94:502-507.

74. Toyoshima-Morimoto F, Taniguchi E, Shinya N, Iwamatsu A, Nishida E. Polo-like kinase 1 phosphorylates cyclin B1 and targets it to the nucleus during prophase. Nature 2001; 410:215-220.

75. Hagting A, Karlsson C, Clute P, Jackman M, Pines J. MPF localization is controlled by nuclear export. EMBO J. 1998; 17:4127-4138.

76. Yang J, Bardes ES, Moore JD, Brennan J, Powers MA, Kornbluth S. Control of cyclin B1 localization through regulated binding of the nuclear export factor CRM1. Genes Dev. 1998; 12:2131-2143.

77. Jackman M, Lindon C, Nigg EA, Pines J. Active cyclin B1-Cdk1 first appears on centrosomes in prophase. Nat. Cell. Biol. 2003; 5:143-148.

78. Kallstrom H, Lindqvist A, Pospisil V, Lundgren A, Rosenthal CK. Cdc25A localisation and shuttling: characterisation of sequences mediating nuclear export and import. Exp. Cell Res. 2005; 303:89-100.

79. Takizawa CG, Morgan DO. Control of mitosis by changes in the subcellular location of cyclin-B1-Cdk1 and Cdc25C. Curr. Opin. Cell. Biol. 2000; 12:658-665.

80. Uchida S, Ohtsubo M, Shimura M, Hirata M, Nakagama H, Matsunaga T, Yoshida M, Ishizaka Y, Yamashita K. Nuclear export signal in CDC25B. Biochem. Biophys. Res. Commun. 2004; 316:226-232.

81. Perdiguero E, Nebreda AR. Regulation of Cdc25C activity during the meiotic G2/M transition. Cell Cycle 2004; 3:733-737.

82. Hoffmann I, Clarke PR, Marcote MJ, Karsenti E, Draetta G. Phosphorylation and activation of human cdc25-C by cdc2-cyclin B and its involvement in the self-amplification of MPF at mitosis. EMBO J. 1993; 12:53-63.

83. Pomerening JR, Sontag ED, Ferrell Jr JE. Building a cell cycle oscillator: hysteresis and bistability in the activation of Cdc2. Nat. Cell Biol. 2003; 5:346-351.

84. Wang R, He G, Nelman-Gonzalez M, Ashorn CL, Gallick GE, Stukenberg PT, Kirschner MW, Kuang J. Regulation of Cdc25C by ERK-MAP kinases during the G2/M transition. Cell 2007; 128:1119-1132.

85. Lowe M, Rabouille C, Nakamura N, Watson R, Jackman M, Jamsa E, Rahman D, Pappin DJ, Warren G. Cdc2 kinase directly phosphorylates the cis-Golgi matrix protein GM130 and is required for Golgi fragmentation in mitosis. Cell 1998; 94:783-793.

86. Moore JD, Kirk JA, Hunt T. Unmasking the S-phase-promoting potential of cyclin B1. Science 2003; 300:987-990.

87. Nigg EA. The substrates of the cdc2 kinase. Semin. Cell. Biol. 1991; 2:261-270.

88. Peter M, Nakagawa J, Doree M, Labbe JC, Nigg EA. In vitro disassembly of the nuclear lamina and M phase-specific phosphorylation of lamins by cdc2 kinase. Cell 1990; 61:591-602.

89. Sirri V, Hernandez-Verdun D, Roussel P. Cyclin-dependent kinases govern formation and maintenance of the nucleolus. J. Cell. Biol. 2002; 56:969-981.

90. Hendrickson M, Madine M, Dalton S, Gautier J. Phosphorylation of MCM4 by cdc2 protein kinase inhibits the activity of the minichromosome maintenance complex. Proc. Natl. Acad. Sci. U.S.A. 1996; 93:12223-12228.

91. Long JJ, Leresche A, Kriwacki RW, Gottesfeld JM. Repression of TFIIH transcriptional activity and TFIIH-associated cdk7 kinase activity at mitosis. Mol. Cell. Biol. 1998; 18:1467-1476.

92. Papst PJ, Sugiyama H, Nagasawa M, Lucas JJ, Maller JL, Terada N. Cdc2-cyclin B phosphorylates p70 S6 kinase on Ser411 at mitosis. J. Biol. Chem. 1998; 273:15077-15084.

93. Fu J, Bian M, Jiang Q, Zhang C. Roles of Aurora kinases in mitosis and tumorigenesis. Mol. Cancer Res. 2007; 5:1-10.

94. Winey M. Cell cycle: driving the centrosome cycle. Curr. Biol. 1999; 9:R449-452.

95. Fisk HA, Mattison CP, Winey M. Human Mps1 protein kinase is required for centrosome duplication and normal mitotic progression. Proc. Natl. Acad. Sci. U.S.A. 2003; 100:14875-14880.

96. Tokuyama Y, Horn HF, Kawamura K, Tarapore P, Fukasawa K. Specific phosphorylation of nucleophosmin on Thr(199) by cyclin-dependent kinase 2-cyclin E and its role in centrosome duplication. J. Biol. Chem. 2001; 276:21529-21537.

97. Cazales M, Schmitt E, Montembault E, Dozier C, Prigent C, Ducommun B. CDC25B phosphorylation by Aurora-A occurs at the G2/M transition and is inhibited by DNA damage. Cell Cycle 2005; 4:1233-1238.

98. Katayama H, Brinkley WR, Sen S. The Aurora kinases: role in cell transformation and tumorigenesis. Cancer Metastasis Rev. 2003; 22:451-464.

99. Cheetham GM, Knegtel RM, Coll JT, Renwick SB, Swenson L, Weber P, Lippke JA, Austen DA. Crystal structure of aurora-2, an oncogenic serine/threonine kinase. J. Biol. Chem. 2002; 277:42419-42422.

100. Hirota T, Kunitoku N, Sasayama T, Marumoto T, Zhang D, Nitta M, Hatakeyama K, Saya H. Aurora-A and an interacting activator, the LIM protein Ajuba, are required for mitotic commitment in human cells. Cell 2003; 114:585-598.

101. Tsai MY, Wiese C, Cao K, Martin O, Donovan P, Ruderman J, Prigent C, Zheng Y. A Ran signalling pathway mediated by the mitotic kinase Aurora A in spindle assembly. Nat. Cell. Biol. 2003; 5:242-248.

102. Pugacheva EN, Golemis EA. The focal adhesion scaffolding protein HEF1 regulates activation of the Aurora-A and Nek2 kinases at the centrosome. Nat. Cell. Biol. 2005; 7:937-946.

103. Bolton MA, Lan W, Powers SE, McCleland ML, Kuang J, Stukenberg PT. Aurora B kinase exists in a complex with sur- vivin and INCENP and its kinase activity is stimulated by survivin binding and phosphorylation. Mol. Biol. Cell. 2002; 13:3064-3077.

104. Harper JW, Burton JL, Solomon MJ. The anaphase-promoting complex: it’s not just for mitosis any more. Genes Dev. 2002; 16:2179-2206.

105. Yu H. Regulation of APC-Cdc20 by the spindle checkpoint. Curr. Opin. Cell. Biol. 2002; 14:706-714.

106. Waizenegger I, Gimenez-Abian JF, Wernic D, Peters JM. Regulation of human separase by securin binding and autocleavage. Curr. Biol. 2002; 12:1368-1378.

107. Baker DJ, Dawlaty MM, Galardy P, van Deursen JM. Mitotic regulation of the anaphase-promoting complex. Cell. Mol. Life Sci. 2007; 64:589-600.

108. Kramer ER, Scheuringer N, Podtelejnikov AV, Mann M, Peters JM. Mitotic regulation of the APC activator proteins CDC20 and CDH1. Mol. Biol. Cell. 2000; 11:1555-1569.

109. Bembenek J, Yu H. Regulation of the anaphase-promoting complex by the dual specificity phosphatase human Cdc14a. J. Biol. Chem. 2001; 276:48237-48242.

110. Prinz S, Hwang ES, Visintin R, Amon A. The regulation of Cdc20 proteolysis reveals a role for APC components Cdc23 and Cdc27 during S phase and early mitosis. Curr. Biol. 1998; 8:750-760.

111. Reis A, Levasseur M, Chang HY, Elliott DJ, Jones KT. The CRY box: a second APCcdh1-dependent degron in mammalian cdc20. EMBO Rep. 2006; 7:1040-1045.

112. Malmanche N, Maia A, Sunkel CE. The spindle assembly checkpoint: preventing chromosome mis-segregation during mitosis and meiosis. FEBS Lett. 2006; 580:2888-2895.

113. May KM, Hardwick KG. The spindle checkpoint. J. Cell. Sci. 2006; 119:4139-4142.

114. Musacchio A, Salmon ED. The spindle-assembly checkpoint in space and time. Nat. Rev. Mol. Cell Biol. 2007; 8:379-393.

115. Bharadwaj R, Yu H. The spindle checkpoint, aneuploidy, and cancer. Oncogene 2004; 23:2016-2027.

116. Kops GJ, Weaver BA, Cleveland DW. On the road to cancer: aneuploidy and the mitotic checkpoint. Nat. Rev. Cancer 2005; 5:773-785.

117. Chan GK, Liu ST, Yen TJ. Kinetochore structure and function. Trends Cell. Biol. 2005; 15:589-598.

118. Pinsky BA, Biggins S. The spindle checkpoint: tension versus attachment. Trends Cell. Biol. 2005; 15:486-493.

119. Vanoosthuyse V, KG Hardwick. Bub1 and the multilayered inhibition of Cdc20-APC/C in mitosis. Trends Cell. Biol. 2005; 15:231-233.

120. Yu H. Structural activation of Mad2 in the mitotic spindle checkpoint: the two-state Mad2 model versus the Mad2 template model. J. Cell. Biol. 2006; 173:153-157.

121. Fang G, Yu H, Kirschner MW. The checkpoint protein MAD2 and the mitotic regulator CDC20 form a ternary complex with the anaphase-promoting complex to control anaphase initiation. Genes Dev. 1998; 12:1871-1883.

122. Fang G. Checkpoint protein BubR1 acts synergistically with Mad2 to inhibit anaphase-promoting complex. Mol. Biol. Cell. 2002; 13:755-766.

123. Sudakin V, Chan GK, Yen TJ. Checkpoint inhibition of the APC/C in HeLa cells is mediated by a complex of BUBR1, BUB3, CDC20, and MAD2. J. Cell. Biol. 2001; 154:925-936.

124. Morrow CJ, Tighe A, Johnson VL, Scott MI, Ditchfield C, Taylor SS. Bub1 and aurora B cooperate to maintain BubR1-mediated inhibition of APC/CCdc20. J. Cell. Sci. 2005; 118:3639-3652.

125. Yu H, Tang Z. Bub1 multitasking in mitosis. Cell Cycle 2005; 4:262-265.

126. Chan GK, Jablonski SA, Sudakin V, Hittle JC, Yen TJ. Human BUBR1 is a mitotic checkpoint kinase that monitors CENP-E functions at kinetochores and binds the cyclosome/APC. J. Cell. Biol. 1999; 146:941-954.

127. Jablonski SA, Chan GK, Cooke CA, Earnshaw WC, Yen TJ. The hBUB1 and hBUBR1 kinases sequentially assemble onto kinetochores during prophase with hBUBR1 concentrating at the kinetochore plates in mitosis. Chromosoma 1998; 107:386-396.

128. Zhou J, Yao J, Joshi HC. Attachment and tension in the spindle assembly checkpoint. J. Cell. Sci. 2002; 115:3547-3555.

129. Rotman G, Shiloh Y. ATM: a mediator of multiple responses to genotoxic stress. Oncogene 1999; 18:6135-6144.

130. Falck J, Mailand N, Syljuasen RG, Bartek J, Lukas J. The ATM-Chk2-Cdc25A checkpoint pathway guards against radioresistant DNA synthesis. Nature 2001; 410:842-847.

131. Mailand N, Falck J, Lukas C, Syljuasen RG, Welcker M, Bartek J, Lukas J. Rapid destruction of human Cdc25A in response to DNA damage. Science 2000; 288:1425-1429.

132. Niida H, Nakanishi M. DNA damage checkpoints in mammals. Mutagenesis 2006; 21:3-9.

133. Sancar A, Lindsey-Boltz LA, Unsal-Kacmaz K, Linn S. Molecular mechanisms of mammalian DNA repair and the DNA damage checkpoints. Annu. Rev. Biochem. 2004; 73:39-85.

134. Busino L, Chiesa M, Draetta GF, Donzelli M. Cdc25A phosphatase: combinatorial phosphorylation, ubiquitylation and proteolysis. Oncogene 2004; 23:2050-2056.

135. Tibbetts RS, Brumbaugh KM, Williams JM, Sarkaria JN, Cliby WA, Shieh SY, Taya Y, Prives C, Abraham RT. A role for ATR in the DNA damage-induced phosphorylation of p53. Genes Dev. 1999; 13:152-157.

136. Nayak BK, Das GM. Stabilization of p53 and transactivation of its target genes in response to replication blockade. Oncogene 2002; 21:7226-7229.

137. Wu X, Bayle JH, Olson D, Levine AJ. The p53-mdm-2 autoregulatory feedback loop. Genes Dev. 1993; 7:1126-1132.

138. Bartek J, Lukas C, Lukas J. Checking on DNA damage in S phase. Nat. Rev. Mol. Cell. Biol. 2004; 5:792-804.

139. Feehan HF, Mancusi Ungaro A. The use of cocaine as a topical anesthetic in nasal surgery. A survey report. Plast. Reconstr. Surg. 1976; 57:62-65.

140. Andreassen PR, Ho GP, D’Andrea AD. DNA damage responses and their many interactions with the replication fork. Carcinogenesis 2006; 27:883-892.

141. Byun TS, Pacek M, Yee MC, Walter JC, Cimprich KA. Functional uncoupling of MCM helicase and DNA polymerase activities activates the ATR-dependent checkpoint. Genes Dev. 2005; 19:1040-1052.

142. Cortez D. Unwind and slow down: checkpoint activation by helicase and polymerase uncoupling. Genes Dev. 2005; 19:1007-1012.

143. Costanzo V, Gautier J. Single-strand DNA gaps trigger an ATR- and Cdc7-dependent checkpoint. Cell Cycle 2003; 2:17.

144. Bomgarden RD, Lupardus PJ, Soni DV, Yee MC, Ford JM, Cimprich KA. Opposing effects of the UV lesion repair protein XPA and UV bypass polymerase eta on ATR checkpoint signaling. EMBO J. 2006; 25:2605-2614.

145. Costa RM, Chigancas V, Galhardo Rda S, Carvalho H, Menck CF. The eukaryotic nucleotide excision repair pathway. Biochimie 2003; 85:1083-1099.

146. Garcia-Muse T, Boulton SJ. Distinct modes of ATR activation after replication stress and DNA double-strand breaks in Caenorhabditis elegans. EMBO J. 2005; 24:4345-4355.

147. Valerie K, Povirk LF. Regulation and mechanisms of mammalian double-strand break repair. Oncogene 2003; 22:5792-5812.

148. Zou L, Liu D, Elledge SJ. Replication protein A-mediated recruitment and activation of Rad17 complexes. Proc. Natl. Acad. Sci. U.S.A. 2003; 100:13827-13832.

149. Lao Y, Gomes XV, Ren Y, Taylor JS, Wold MS. Replication protein A interactions with DNA. III. Molecular basis of recognition of damaged DNA. Biochemistry 2000; 39:850-859.

150. Ball HL, Myers JS, Cortez D. ATRIP binding to replication protein A-single-stranded DNA promotes ATR-ATRIP localization but is dispensable for Chk1 phosphorylation. Mol. Biol. Cell. 2005; 16:2372-2381.

151. Namiki Y, Zou L. ATRIP associates with replication protein A-coated ssDNA through multiple interactions. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:580-585.

152. Zou L, Elledge SJ. Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 2003; 300:1542-1548.

153. Parrilla-Castellar ER, Arlander SJ, Karnitz L. Dial 9-1-1 for DNA damage: the Rad9-Hus1-Rad1 (9-1-1) clamp complex. DNA Repair 2004; 3:1009-1014.

154. Zou L, Cortez D, Elledge SJ. Regulation of ATR substrate selection by Rad17-dependent loading of Rad9 complexes onto chromatin. Genes Dev. 2002; 16:198-208.

155. Thelen MP, Venclovas C, Fidelis K. A sliding clamp model for the Rad1 family of cell cycle checkpoint proteins. Cell 1999; 96:769-770.

156. Gottifredi V, Prives C. The S phase checkpoint: when the crowd meets at the fork. Semin. Cell. Dev. Biol. 2005; 16:355-368.

157. Wang X, Zou L, Lu T, Bao S, Hurov KE, Hittelman WN, Elledge SJ, Li L. Rad17 phosphorylation is required for claspin recruitment and Chk1 activation in response to replication stress. Mol. Cell. 2006; 23:331-341.

158. Lin SY, Li K, Stewart GS, Elledge SJ. Human Claspin works with BRCA1 to both positively and negatively regulate cell proliferation. Proc. Natl. Acad. Sci. U.S.A. 2004; 101:6484-6489.

159. Chini CC, Chen J. Human claspin is required for replication checkpoint control. J. Biol. Chem. 2003; 278:30057-30062.

160. Lee J, Kumagai A, Dunphy WG. Claspin, a Chk1-regulatory protein, monitors DNA replication on chromatin independently of RPA, ATR, and Rad17. Mol. Cell. 2003; 11:329-340.

161. Wang Y, Cortez D, Yazdi P, Neff N, Elledge SJ, Qin J. BASC, a super complex of BRCA1-associated proteins involved in the recognition and repair of aberrant DNA structures. Genes Dev. 2000; 14:927-939.

162. Goldberg M, Stucki M, Falck J, D’Amours D, Rahman D, Pappin D, Bartek J, Jackson SP. MDC1 is required for the intra-S-phase DNA damage checkpoint. Nature 2003; 421:952-956.

163. Lou Z, Chini CC, Minter-Dykhouse K, Chen J. Mediator of DNA damage checkpoint protein 1 regulates BRCA1 localization and phosphorylation in DNA damage checkpoint control. J. Biol. Chem. 2003; 278:13599-13602.

164. Stewart GS, Wang B, Bignell CR, Taylor AM, Elledge SJ. MDC1 is a mediator of the mammalian DNA damage checkpoint. Nature 2003; 421:961-966.

165. Kim ST, Xu B, Kastan MB. Involvement of the cohesin protein, Smc1, in Atm-dependent and independent responses to DNA damage. Genes Dev. 2002; 16:560-570.

166. Kitagawa R, Bakkenist CJ, McKinnon PJ, Kastan MB. Phosphorylation of SMC1 is a critical downstream event in the ATM-NBS1-BRCA1 pathway. Genes Dev. 2004; 18:1423-1438.

167. Xiao Z, Chen Z, Gunasekera AH, Sowin TJ, Rosenberg SH, Fesik S, Zhang H. Chk1 mediates S and G2 arrests through Cdc25A degradation in response to DNA-damaging agents. J. Biol. Chem. 2003; 278:21767-21773.

168. Falck J, Petrini JH, Williams BR, Lukas J, Bartek J. The DNA damage-dependent intra-S phase checkpoint is regulated by parallel pathways. Nat. Genet. 2002; 30:290-294.

169. Merrick CJ, Jackson D, Diffley JF. Visualization of altered replication dynamics after DNA damage in human cells. J. Biol. Chem. 2004; 279:20067-20075.

170. D’Amours D, Jackson SP. The Mre11 complex: at the crossroads of dna repair and checkpoint signalling. Nat. Rev. Mol. Cell. Biol. 2002; 3:317-327.

171. Lee JH, Paull TT. Direct activation of the ATM protein kinase by the Mre11/Rad50/Nbs1 complex. Science 2004; 304:93-96.

172. Uziel T, Lerenthal Y, Moyal L, Andegeko Y, Mittelman L, Shiloh Y. Requirement of the MRN complex for ATM activation by DNA damage. EMBO J. 2003; 22:5612-5621.

173. Cuadrado M, Martinez-Pastor B, Murga M, Toledo LI, Gutierrez - Martinez P, Lopez E, Fernandez-Capetillo O. ATM regulates ATR chromatin loading in response to DNA double-strand breaks. J. Exp. Med. 2006; 203:297-303.

174. Jazayeri A, Falck J, Lukas C, Bartek J, Smith GC, Lukas J, Jackson SP. ATM- and cell cycle-dependent regulation of ATR in response to DNA double-strand breaks. Nat. Cell. Biol. 2006; 8:37-45.

175. Kaneko YS, Watanabe N, Morisaki H, Akita H, Fujimoto A, Tominaga K, Terasawa M, Tachibana A, Ikeda K, Nakanishi M. Cell-cycle-dependent and ATM-independent expression of human Chk1 kinase. Oncogene 1999; 18:3673-3681.

176. Lukas C, Bartkova J, Latella L, Falck J, Mailand N, Schroeder T, Sehested M, Lukas J, Bartek J. DNA damage-activated kinase Chk2 is independent of proliferation or differentiation yet correlates with tissue biology. Cancer Res. 2001; 61:4990-4993.

177. Ahn JY, Li X, Davis HL, Canman CE. Phosphorylation of threonine 68 promotes oligomerization and autophosphorylation of the Chk2 protein kinase via the forkhead-associated domain. J. Biol. Chem. 2002; 277:19389-19395.

178. Oliver AW, Paul A, Boxall KJ, Barrie SE, Aherne GW, Garrett MD, Mittnacht S, Pearl LH. Trans-activation of the DNA-damage signalling protein kinase Chk2 by T-loop exchange. EMBO J. 2006; 25:3179-3190.

179. Xu X, Tsvetkov LM, Stern DF. Chk2 activation and phosphorylation-dependent oligomerization. Mol. Cell. Biol. 2002; 22:4419-4432.

180. Li J, Stern DF. DNA damage regulates Chk2 association with chromatin. J. Biol. Chem. 2005; 280:37948-37956.

181. Lukas C, Falck J, Bartkova J, Bartek J, Lukas J. Distinct spatiotemporal dynamics of mammalian checkpoint regulators induced by DNA damage. Nat. Cell. Biol. 2003; 5:255-260.

182. Guo Z, Kumagai A, Wang SX, Dunphy WG. 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. 2000; 14:2745-2756.

183. Hekmat-Nejad M, You Z, Yee MC, Newport JW, Cimprich KA. Xenopus ATR is a replication-dependent chromatin-binding protein required for the DNA replication checkpoint. Curr. Biol. 2000; 10:1565-1573.

184. Nghiem P, Park PK, Kim Y, Vaziri C, Schreiber SL. ATR inhibition selectively sensitizes G1 checkpoint-deficient cells to lethal premature chromatin condensation. Proc. Natl. Acad. Sci. U.S.A. 2001; 98:9092-9097.

185. Niida H, Tsuge S, Katsuno Y, Konishi A, Takeda N, Nakanishi M. Depletion of Chk1 leads to premature activation of Cdc2-cyclin B and mitotic catastrophe. J. Biol. Chem. 2005; 280:39246-39252.

186. Petermann E, Caldecott KW. Evidence that the ATR/Chk1 pathway maintains normal replication fork progression during unperturbed S phase. Cell Cycle 2006; 5:2203-2209.

187. Casper AM, Nghiem P, Arlt MF, Glover TW. ATR regulates fragile site stability. Cell 2002; 111:779-789.

188. Durkin SG, Arlt MF, Howlett NG, Glover TW. Depletion of CHK1, but not CHK2, induces chromosomal instability and breaks at common fragile sites. Oncogene 2006; 25:4381-4388.

189. El Achkar E, Gerbault-Seureau M, Muleris M, Dutrillaux B, Debatisse M. Premature condensation induces breaks at the interface of early and late replicating chromosome bands bearing common fragile sites. Proc. Natl. Acad. Sci. U.S.A. 2005; 102:18069-18074.

190. Le Beau MM, Rassool FV, Neilly ME, Espinosa R 3rd, Glover TW, Smith DI, McKeithan TW. Replication of a common fragile site, FRA3B, occurs late in S phase and is delayed further upon induction: implications for the mechanism of fragile site induction. Hum. Mol. Genet. 1998; 7:755-761.

191. Wang L, Darling J, Zhang JS, Huang H, Liu W, Smith DI. Allele-specific late replication and fragility of the most active common fragile site, FRA3B. Hum. Mol. Genet. 1999; 8:431-437.

192. Lukas J, Lukas C, Bartek J. Mammalian cell cycle checkpoints: signalling pathways and their organization in space and time. DNA Repair 2004; 3:997-1007.

193. Nyberg KA, Michelson RJ, Putnam CW, Weinert TA. Toward maintaining the genome: DNA damage and replication checkpoints. Annu. Rev. Genet. 2002; 36:617-656.

194. Fletcher L, Muschel RJ. The centrosome and the DNA damage induced checkpoint. Cancer Lett 2006; 243:1-8.

195. Smits VA, Klompmaker R, Arnaud L, Rijksen G, Nigg EA, Medema RH. Polo-like kinase-1 is a target of the DNA damage checkpoint. Nat. Cell. Biol. 2000; 2:672-676.

196. Lange BM. Integration of the centrosome in cell cycle control, stress response and signal transduction pathways. Curr. Opin. Cell. Biol. 2002; 14:35-43.

197. Toyoshima-Morimoto F, Taniguchi E, Nishida E. Plk1 promotes nuclear translocation of human Cdc25C during prophase. EMBO Rep. 2002; 3:341-348.

198. Bartek J, Lukas J. DNA damage checkpoints: from initiation to recovery or adaptation. Curr. Opin. Cell. Biol. 2007; 19:238-245.

199. Syljuasen RG, Jensen S, Bartek J, Lukas J. Adaptation to the ionizing radiation-induced G2 checkpoint occurs in human cells and depends on checkpoint kinase 1 and Polo-like kinase 1 kinases. Cancer Res. 2006; 66:10253-10257.

200. van Vugt MA, Bras A, Medema RH. Polo-like kinase-1 controls recovery from a G2 DNA damage-induced arrest in mammalian cells. Mol. Cell. 2004; 15:799-811.

201. Syljuasen RG. Checkpoint adaptation in human cells. Oncogene 2007; 26:5833-5839.


Further Reading

Humphrey T, Brooks G, eds. Cell Cycle Control: Mechanisms and Protocols. Humana Press, Inc., Totowa, NJ. pp. 113-153.

Schonthal AH, ed. Checkpoint Controls and Cancer, Volume 1: Reviews and Model Systems. 2004. Humana Press, Inc., Totowa, NJ. pp. 1-49, 99-161.


See Also

Cell Cycle

Cell Division, Small Molecules to Study

DNA Damage: An Overview

DNA Damage, Sensing of

DNA Replication, an Overview