Abeloff's Clinical Oncology, 4th Edition

Part I – Science of Clinical Oncology

Section A – Biology and Cancer

Chapter 4 – Control of the Cell Cycle

Jacqueline Lees

SUMMARY OF KEY POINTS

  

   

Cells in most postnatal tissues are quiescent. Exceptions include cells of the hematopoietic system, skin, and gastrointestinal mucosa.

  

   

The key challenges for proliferating cells are to make an accurate copy of the 3 billion bases of DNA (S phase) and to segregate the duplicated chromosomes equally into daughter cells (mitosis).

  

   

Progression through the cell cycle is dependent on both extrinsic and intrinsic factors.

  

   

Extrinsic factors include cell-to-cell contact, basement membrane attachments, and growth factor or cytokine exposure.

  

   

The internal cell cycle machinery is controlled largely by oscillating levels of cyclin proteins and by modulation of cyclin-dependent kinase activity.

  

   

One way in which growth factors regulate cell cycle progression is by affecting the levels of the D-type cyclins in the G1 phase of the cell cycle.

  

   

The restriction point of the cell cycle occurs in late G1 and is the point beyond which the cell is committed to progress through the rest of the cell cycle. It is governed by a known tumor suppressor, the retinoblastoma protein.

  

   

Cell cycle checkpoints are surveillance mechanisms that link the rate of cell cycle transitions to the timely and accurate completion of prior dependent events.

  

   

Cells can arrest at cell cycle checkpoints temporarily to allow for (1) the repair of cellular damage; (2) the dissipation of an exogenous cellular stress signal; or (3) availability of essential growth factors, hormones, or nutrients.

  

   

The major function of the p53 tumor suppressor protein is to induce cell cycle arrest, senescence, or death in response to cellular stress.

  

   

Activation of the G1, S, and G2 phase checkpoints after DNA damage minimizes replication of damaged DNA templates or their segregation to daughter cells.

  

   

Activation of the mitotic spindle checkpoint prevents defects in chromosome segregation and protects against aneuploidy.

  

   

Disruption of cell cycle controls is a hallmark of all malignant cells. Disruption can manifest as alterations of growth factor signaling pathways, dysregulation of the core cell cycle machinery, and/or disruption of cell cycle checkpoint controls.

  

   

Because cell cycle control is disrupted in virtually all tumor types, the cell cycle-related gene products that are mutated in tumors provide therapeutic targets that might preferentially affect tumor cells more than normal tissues.

INTRODUCTION

The majority of the cells in the adult body are arrested in a quiescent state, called the G0 state. Most of these cells are terminally differentiated and never divide. However, specific populations retain the ability to proliferate throughout the adult life span, and this is essential for viability. For example, cells of the hematopoietic compartment and the gut have a high rate of turnover, and high rates of proliferation are therefore essential for the maintenance of these tissues. On average, about 2 trillion cell divisions occur in an adult human every 24 hours (about 25 million per second). The decision to proliferate or not is very tightly regulated. It is influenced by a variety of exogenous signals, including nutrients, mitogenic (e.g., epidermal growth factor and platelet-derived growth factor) and inhibitory (e.g., transforming growth factor-β) growth factors, and the interaction of the cell with its neighbors and with the underlying extracellular matrix. Each of these factors stimulates intracellular signaling pathways that can either promote or suppress proliferation. The cell integrates all of these signals, and if the balance is favorable, the cell will initiate the proliferation process. Anything that disrupts this balance can lead to either the reduction or expansion of a particular cell population. It is now clear that such changes are a hallmark of tumor cells. They carry mutations that impair signaling pathways that suppress proliferation and/or activate pathways that promote proliferation.

It is essential that proliferating cells copy their genomes and segregate them to the daughter cells with high fidelity. Over the past three decades, extensive effort has been placed on unraveling the basic molecular events that control this process. Studies in a variety of organisms have identified evolutionarily conserved machinery that controls eukaryotic cell cycle transitions through the action of key enzymes called cyclin-dependent kinases (CDKs). Eukaryotic cells have also evolved a series of surveillance pathways, termed cell cycle checkpoints, that monitor for potential problems during the cell cycle process. Human cells are continuously exposed to external agents (e.g., reactive chemicals and ultraviolet light) and to internal agents (e.g., by-products of normal intracellular metabolism, such as reactive oxygen intermediates) that can induce DNA damage. The cell cycle checkpoints detect DNA damage and activate cell cycle arrest and DNA repair mechanisms, thereby maintaining genomic integrity. Most, if not all, human tumor cells have mutations within key components of both the cell cycle machinery and checkpoint pathways. This has important clinical implications, as the presence of these defects can modulate cellular sensitivity to chemotherapeutic regimens that induce DNA damage. This chapter focuses on the mechanics of the cell cycle and checkpoint-signaling pathways and discusses how this knowledge can lead to the efficient use of current anticancer therapies and to the development of novel agents.

CELL CYCLE MACHINERY

Overview of Cell Cycle Phases

Cell proliferation proceeds through a well-defined series of stages ( Fig. 4-1 ). First, the cell moves from the quiescent G0 state into the first gap phase, or G1, in which the cell is essentially readying itself for the cell division process. This involves a dramatic upregulation of both transcriptional and translational programs not only to yield the proteins that are required to regulate cell division but also to essentially double the complement of macromolecules so that one cell can give rise to two cells without a loss of cell size. This protein synthesis phase is frequently referred to as cell growth. Not surprisingly, this takes a significant amount of time (anywhere from 8 to 30 hours) and energy. Studies with cultured cells show that mitogenic growth factors are essential for continued passage through G1. Specifically, if growth factors are withdrawn at any point during this phase, the cell will not divide. However, as it nears the end of G1, the cell passes through a key transition point, called the restriction point, at which it becomes growth factor independent and is fully committed to undergoing cell division. Within an hour or two, the cell enters the synthesis phase, or S phase, in which each of the chromosomes is replicated once and only once. The cell then enters a second gap phase, called G2, which lasts 3 to 5 hours, and then initiates mitosis, or M phase, in which the chromosomes are segregated. On completion of mitosis, the daughter cells can enter quiescence or initiate a second round of cell division, depending on the milieu.

 
 

Figure 4-1  The cell cycle. One round of cell division requires high-fidelity duplication of DNA during the S phase of the cell cycle and proper segregation of duplicated chromosomes during mitosis, or M phase. Before and after the S phase and M phase, the cell transits through “gap” phases, termed G1 and G2. The appropriate transition through these stages is controlled by the action of specific cyclin/CDK complexes.

 

 

Cyclin and Cyclin-Dependent Kinase Complexes

The CDKs constitute a large subfamily of highly conserved ser/thr kinases that are defined by their dependence on a regulatory subunit, called a cyclin. [1] [2] Only a subset of these CDKs are specifically involved in cell cycle regulation. Yeast use a single CDK for cell cycle control that is called cdc2 in Schizosaccharomyces pombe and cdc28 in Saccharomyces cerevisiae. In contrast, mammals employ multiple CDKs.[3] The first identified human CDK, called CDK1 (originally cdc2), was cloned by virtue of its ability to complement a mutant cdc2 yeast strain.[4] Subsequent studies identified additional human CDKs and determined that they regulate distinct cell cycle stages; CDK4 and CDK6 regulate passage through G1, CDK2 regulates the G1-to-S transition and S phase, and CDK1 controls G2 and mitosis. The activity of these kinases is controlled by multiple regulatory mechanisms. [5] [6] Most important, the CDKs act in association with a cyclin subunit that binds to the conserved PSTAIRE helix within the kinase. [3] [5] Cyclin binding causes a reorientation of residues within the active sites that is essential for kinase activity.[3] The associated cyclin also determines the substrate specificity of the resulting cyclin-CDK complex. The cyclins are quite divergent, especially in their N-terminal sequences, but they all share a highly conserved 100-amino-acid sequence, called the cyclin box, that mediates CDK binding and activation. As their name implies, cyclins were originally identified as proteins whose expression was restricted to a particular stage of the cell cycle.[7] This is due to cell cycle-dependent regulation of both cyclin gene transcription and protein degradation. Notably, there is frequently a delay between the formation of a particular cyclin/CDK complex and the appearance of kinase activity (Fig. 4-2 ). This reflects considerable post-translational regulation of the cyclin/CDK complex. [8] [9] [10] [11] First, kinase activation is absolutely dependent on phosphorylation of a threonine residue that is adjacent to the active site (thr 160 in CDK2). This is catalyzed by a kinase, called CDK-activating kinase (CAK). [11] [12] [13] In mammalian cells, phosphorylation occurs after cyclin binding. Although there appear to be at least two mammalian CAKs, the major CAK is a trimolecular complex composed of CDK7, cyclin H, and Mat1. [12] [13] [14] This kinase is constitutively active, and to date, there is no evidence that it is cell cycle regulated. Indeed, the CDK7-cyclinH-Mat1 complex is also required for the control of basal transcription via regulation of RNA polymerase II function. [12] [15] Second, when it is first formed, the cyclin/CDK complex is frequently subject to inhibitory phosphorylation of Thr-14 and Tyr-15 residues within the CDK's active site by the Wee1 (Tyr-15) and Myt1 (Thr-14 and Tyr-15) kinases.[9] Activation of the cyclin/CDK complex is now dependent on the action of a dual-specificity phosphatase called Cdc25.[10] Mammalian cells have three different Cdc25 proteins, called Cdc25A, Cdc25B, and Cdc25C, which show some specificity for different cyclin/CDK complexes.[10]

 
 

Figure 4-2  The expression of the cyclin subunits in tightly linked to cell cycle phases. Extracellular stimuli, such as mitogenic growth factors and hormones, induce the expression of the D-type cyclins early in G1. Cyclin E expression occurs late in G1, synonymous with the restriction point, and its levels peak at the G1-to-S transition and then decline during S phase. Cyclin A is induced a little later than cyclin E and is degraded at the metaphase of mitosis. The β-type cyclins are expressed primarily in G2, and the levels remain high until the anaphase of mitosis. There is typically a delay between the expression of the cyclin protein and the appearance of cyclin-associated kinase activity that results from post-translational regulation. This is most pronounced in the case of cyclin B, for which activation of the cyclin B/CDK1-associated kinase activity does not occur until the G2-to-M transition.

 

 

The mammalian cyclins are divided into four distinct classes—D-type cyclins, E-type cyclins, α-type cyclins, and β-type cyclins—on the basis of both their sequence homology and the stage of the cell cycle at which they act (see Figs. 4-1 and 4-2 [1] [2]).[16] Each of these classes has two or three paralogs (cyclin D1, D2, and D3; cyclin E1 and E2; cyclin A1 and A2; and cyclin B1 and B2). The relative roles of these paralogs are still unclear. In some cases, differences exist (e.g., in subcellular localization) that suggest that particular paralogs will have distinct activities or regulation. However, other studies (particularly the analysis of mouse models) show that there is considerable functional redundancy, or at least an ability to substitute for one another, between paralogs of a particular cyclin type. [17] [18] For simplicity, we will focus largely on the core properties of the cyclin types.

The D-type cyclins represent an unusual class of cyclins. [16] [19] First, they do not participate the cell division process itself. Instead, they play a critical role in determining whether a cell will divide under the direction of external cues. Second, their expression is not really cell cycle regulated; D-type cyclins are present at very low levels in quiescent cells, in large part because they are phosphorylated by an abundant G0 kinase called GSK3b and then exported to the cytoplasm for degradation, but their expression is induced during G1, and it persists through all subsequent cell cycle stages.[20] This G1induction is a direct consequence of mitogenic signaling. This inhibits expression of GSK3b and activates transcription of the D-type cyclins. Notably, individual mitogenic signaling pathways induce different D-type cyclins. For example, signaling by EGF/ras (via the AP-1 transcription factor) and Wnt/β-catenin (via the Tcf/Lef transcription factor) specifically induces cyclin D1, while c-myc specifically induces cyclin D2. [21] [22] [23] [24] [25] [26] This specificity helps to ensure that the presence of multiple proproliferative signals gives rise to more D-type cyclins than a single mitogen does. Importantly, the analysis of mouse models shows that cyclins D1, D2, and D3 are functionally redundant; the key factor appears to be the total level of D-type cyclin that is present in the cell.[27] This promotes cell cycle entry through two distinct mechanisms. First, the D-type cells titrate inhibitory molecules, called CDK inhibitors, away from other CDK-kinase complexes and thereby promote their activation. [5] [28] [29] Second, cyclins D1, D2, and D3 associate with CDK4 and CDK6, and the resulting complexes phosphorylate the retinoblastoma protein (pRB), a key gatekeeper for cell cycle reentry.[30] [31] [32] [33] [34] We will discuss both the CDK inhibitors and pRB in more detail later because of their central importance in controlling cell cycle reentry and their frequent disruption in cancer.

E-type cyclins are expressed during late in G1 under the control of the E2F transcription factors. [35] [36] Cyclin E binds specifically to CDK2, and the resulting complex is required for cells to move through the G1-to-S transition. [37] [38] [39] To date, several cyclin E/CDK2 substrates have been identified. Some cyclin E/CDK2 substrates play a positive role in cell cycle progression. For example, cyclin E/CDK2 phosphorylates NPAT, a transcription factor that mediates transcriptional activation of histone gene clusters. [40] [41] [42] The resulting increase in histone pools is essential for the appropriate packaging of newly replicated DNA in S phase cells. Cyclin E/CDK2 also induces the duplication of centrosomes that is required for formation of the mitotic spindle. [43] [44] [45] [46] Other cyclin E/CDK2 substrates are key cell cycle inhibitors. First, cyclin E/CDK2 phosphorylates pRB at completely different sites from those that have already been modified by the cyclin D/CDK4/6 kinases, and this is sufficient to inactivate pRB's growth suppressive function. [31] [34] [47] Second, cyclin E/CDK2 phosphorylates the CDK inhibitor p27 on Thr-187. [48] [49] This creates a high-affinity binding site for a ubiquitin ligase, called SCF, which plays a very important role in G1/S control. [50] [51] [52] [53] SCF has three core components: a RING finger protein, called Rbx1, which recruits the E2-ubiquitin conjugate; a cullin (Cul1); and Skp1 ( Fig. 4-3 ). [52] [53] [54] Skp1 acts to recruit a family of proteins, called F-box proteins, that determine the target specificity of the SCF complex (see Fig. 4-3 ). In the case of p27, the F-box protein is Skp2, and the SCF complex is therefore designated as SCFSkp2. [55] [56] Once SCF binds its substrate, it transfers a ubiquitin molecule to lysine residues within the target protein and subsequently to lysine residues in the ubiquitin molecule to create a polyubiquitin chain.[54] This polyubiquitin chain targets the substrate to the proteasome for degradation.[54] Notably, SCF is also responsible for the decline in cyclin E/CDK2 levels that is activated during S phase (see Fig. 4-2 ). First, cyclin E/CDK2 actually phosphorylates itself on multiple sites, creating a recognition site for SCFFbw7/Cdc4 and thereby ensuring its own destruction. [57] [58] Second, cyclin A/CDK2 (the S phase kinase) phosphorylates E2F-1, the transcription factor that activates cyclin E transcription, and this is targeted for degradation by SCFSkp2. [59] [60] [61] Together, these mechanisms restrict the action of cyclin E/CDK2 to a small window in the cell cycle.

 
 

Figure 4-3  Ubiquitin ligases. The SCF and APC ubiquitin ligases play a key role in enabling forward passage through key cell cycle transitions. These are both large complexes that include three core components: a scaffolding protein called a cullin, a protein that recruits the E2 and its associated ubiquitin molecule, and a specificity factor (called the F-box protein in SCF and the activator in APC) that recruits the substrate. SCF and APC catalyze polyubiquitination of their substrates, and this acts as a signal for substrate degradation by the 26S proteasome. SCF has numerous substrates whose degradation promotes passage through the early stages of the cell cycle, including p27Kip1 (the restriction point) and cyclin E, E2F-1, and Cdt1 (S-phase). APC is essential for completion of mitosis (by promoting degradation of securin and the mitotic cyclins) and to allow origin licensing (by promoting degradation of geminin and thereby allowing accumulation of cdt1 during G1).

 

 

The α-type cyclins are first transcribed during late G1 under the control of the E2F transcription factors in a similar manner to that of cyclin E. However, in contrast to cyclin E, cyclin A associates with both CDK2 and CDK1 and it acts at two distinct cell cycle stages. [62] [63] First, cyclin A/CDK2 is absolutely required for S phase progression. This was established by showing that injection of either cyclin A antisense constructs or antibodies was sufficient to block S phase progression. [62] [64] Notably, cyclin A/CDK2 enters the nucleus at the start of S phase, and it is specifically localized at nuclear replication foci and therefore is thought to be actively involved in the firing of replication origins.[65] As was described previously, cyclin A/CDK2 is also required to phosphorylate E2F-1 and mediate its degradation, and this is required to prevent E2F1 from triggering apoptosis. [59] [60] [61] Second, cyclin A/CDK1 complexes act during G2 and at the beginning of mitosis.[62] Here, they are thought to play a key role in initiating the condensation of chromatin and might also participate in the activation of the cyclin B/CDK1 complexes. Cyclin A/CDK2 is destroyed in the prometaphase of mitosis through the action of another ubiquitin ligase, called anaphase-promoting complex (APC).[66] APC is a much larger complex that SCF, but it also contains a RING finger protein, called Apc11, to recruit the E2-ubiquitin conjugate, a core cullin subunit (Apc2), and it binds a variety of activators that are required for APC activity and, in a manner comparable to that of the F-box proteins of SCF, establish substrate specificity (see Fig. 4-3 ). [54] [66] In the case of cyclin A/CDK2, the activator is CDC20, and the APC complex is therefore designated as APCCdc20.

There are two β-type cyclins, B1 and B2, which show significant differences in their subcellular localization. The analysis of mutant mouse models shows that cyclin B2 loss has no detectable effect on development, while cyclin B1 is absolutely required for embryogenesis.[67] This indicates that cyclin B1 is the major β-type cyclin in vivo; therefore, we will restrict our discussion to this isoform. Cyclin B1 protein first appears at the beginning of G2. It accumulates steadily through G2 and associates specifically with CDK1.[63] However, the resulting cyclin B1/CDK1 complex is mostly sequestered in the cytoplasm, and it is retained in an inactive form throughout G2 via the inhibitory phosphorylation of Thr-14 and Tyr-15 in CDK1′s active site by the Myt1 and, to a lesser extent, Wee1 kinases. [68] [69] [70]Activation of cyclin B1/CDK1 occurs in a highly synchronous manner during the first stage (called prophase) of mitosis (see Fig. 4-2 ).[71] This activation is mediated by two changes. First, the activities of myt1 and wee1 are dramatically downregulated at the transition between G2 and M. Second, there is a dramatic increase in the activity of the Cdc25A and C phosphatases that relieves the inhibitory phosphorylation of Thr-14 and Tyr-15.[10] These activity changes are triggered by the phosphorylation of Myt1, Wee1, Cdc25A, and Cdc25C. Three different kinases are thought to contribute to this phosphorylation: polo-like kinase, cyclin A/CDK1, and cyclin B1/CDK1 itself. The involvement of cyclin B1/CDK1 creates a powerful feedforward loop; once a small amount of cyclin B1/CDK1 is activated, it simultaneously inactivates its own inhibitors and activates its activators, enabling a rapid transformation of the entire cyclin B1/CDK1 pool from the inactive state to the active state. Once active, cyclin B1/CDK1 phosphorylates components of the centrosomes and initiates a process called centrosome separation, in which the centrosomes move to opposing poles of the nascent spindle, an event that is essential for formation of the mitotic spindle.[72] Cyclin B1/CDK1 then translocates across the nuclear membrane (which is still intact at this point in the mitosis) to orchestrate mitotic events.[73] Notably, cyclin B1/CDK1 is degraded at the end of metaphase (see Fig. 4-2 ).[74] This is triggered by the ubiquitination of cyclin B1 by the ubiquitin ligase APC and its subsequent recognition and degradation by the proteosome. [66] [75] [76] This downregulation of mitotic CDKs is required for cytokinesis (the separation of the daughter cells) and reentry into G0/G1.[74]

Cyclin-Dependent Kinase Inhibitors

The CDK inhibitors (CDKIs) play a key role in establishing the activity of the cyclin/CDK complexes in response to either external signals or internal stresses.[5] The CDKIs can be divided into two distinct families based on their biological properties. The first CDKI family is named INK4, based on their roles as inhibitors of CDK4. The INK4 family has four members called p16INK4a, p15INK4b, p18INK4c, and p19INK4d. These INK4 proteins specifically target CDK4 and CDK6 and not other CDKs. They preferentially target the monomeric CDK and prevent cyclin binding. Consistent with their inhibitory role, the alterations in the INK4 genes are observed in human tumors.[77] Ink4a appears to be most the most frequently affected; it was identified as a tumor suppressor that is associated with familial melanoma, and it is inactivated by point mutation, deletion, and/or promoter methylation in approximately 30% of all human tumors. [78] [79] In contrast, point mutations in p15INK4b, p18INK4c, and p19INK4d are rare, but promoter methylation of Ink4c has been detected in Hodgkin lymphomas and medulloblastomas, and reduced p18INK4c protein expression has been seen in a variety of tumor types.[80] [81] [82] [83] [84]

The second CDKI family is named CIP/KIP and includes three members: p21Cip1 (also called p21Waf1), p27Kip1, and p57Kip2.[5] These CIP/KIP proteins have two major activities. First, they associate with, and inhibit the activity of, the G1/S and S phase kinases cyclin E/CDK2 and cyclin A/CDK2. Second, p21Cip1 and p27Kip1 bind to the D-type cyclins outside of the CDK binding site and actually promote assembly of cyclin D/CDK4/6 complexes.[85] These two activities are clearly paradoxical. However, they are critical in establishing how the cell decides whether or not to divide in response to external signals. In general, the expression and/or activity of CDKIs is promoted by growth suppressive signals and inhibited by proproliferative signals. For example, the inhibitory growth factor transforming growth factor-β induces transcription of p15INK4b, while several mitogenic signaling pathways cause Akt to phosphorylate p21Cip1 and p27Kip1 and induce their cytoplasmic sequestration. [86] [87] [88] [89] [90] Importantly, signaling pathways have the opposite effect on the D-type cyclins: Growth suppressive signals inhibit their expression and activity while mitogens are activating. The opposing regulation of CDKIs and D-type cyclins controls cell cycle entry by creating a tipping point ( Fig. 4-4 ). G0/G1 cells have low levels of D-type cyclins and high levels of CDKIs; thus, cell cycle entry is blocked. However, an increase in mitogenic signals boosts the levels of cyclin D, and this eventually exceeds the level of INK4 proteins, which are simultaneously declining. At this point, the D-type cyclins begin to bind to the CIP/KIP proteins. This helps the D-type cyclins to assemble into active cyclin D/CDK4/6 complexes and draws the CIP/KIP proteins away from the cyclin E/CDK2 complexes as they begin to accumulate. The cyclin D/CDK4/6 and cyclin E/CDK2 complexes cooperate in the phosphorylation and inactivation of the pRB protein. This appears to be the tipping point in commitment to cell cycle entry.

 
 

Figure 4-4  The CDK inhibitors (CDKIs) play a key role in regulating the G1-to-S transition. The levels of nuclear INK4 and Cip/Kip CDKIs are typically elevated by growth inhibitory stimuli and reduced by mitogenic stimuli. The INK4 family members bind specifically to CDK4 and CDK6 and inhibit their association with cyclin D. The Cip/Kip CDKIs interact with both the cyclin and CDK components and have highest affinity with the intact cyclin–CDK complex. Cip/Kip binds in a different way to cyclin D versus cyclin E complexes, and this has opposing effects on their activity. Cip/Kip binding enables formation of active cyclin D–CDK4/6 complexes. In contrast, Cip/Kip associates with cyclin E–CDK complexes and blocks their activity. Cyclin D complexes have a higher affinity for Cip/Kip than do cyclin E complexes. The mitogen-induced accumulation of D-type cyclins during G1 titrates Cip/Kip from cyclin E–CDK2 and facilitates activation of both cyclin D–CDK4/6 and cyclin E–CDK2 kinases. Cyclin E–CDK2 can also phosphorylate p27 to promote its ubiquitin-mediated degradation.

 

 

pRB Tumor Suppressor

The retinoblastoma protein (pRB) was originally identified by virtue of its association with hereditary retinoblastoma protein.[91] It behaves as a classic tumor suppressor: Affected individuals inherit a germline mutation within one Rb-1 allele, and loss of heterozygosity is seen in all of the tumors. Subsequent studies showed that the transforming ability of small DNA tumor viruses, including human papilloma virus, adenovirus, and simian virus, was dependent on the ability of virally encoded oncoproteins (E7, E1A, and SV40, respectively) to bind and inhibit pRB.[92] Moreover, the RB-1 gene was found to be inactivated in approximately one third of all sporadic human tumors.[91] Thus, pRB is a major human tumor suppressor.

To date, numerous pRB-associated proteins have been identified.[93] However, studies in mouse models indicate that pRB's tumor suppressive activity is largely dependent on its ability to prevent cell cycle entry through inhibition of the E2F transcription factors. [94] [95] [96] The E2F proteins regulate the cell cycle-dependent transcription of numerous targets, including core components of the cell cycle control (e.g., cyclin E and cyclin A) and DNA replication (e.g., cdc6, CDT1, and the MCM proteins) machineries. [47] [97] [98] [99] pRB regulates E2F through two distinct mechanisms. First, its association with E2F is sufficient to block its transcriptional activity.[100] Second, the pRB-E2F complex can recruit histone deacetylases (HDACs) to the promoters of E2F-responsive genes and thereby actively repress their transcription. [101] [102] [103] Cell cycle entry requires the sequential phosphorylation of pRB by cyclin D/CDK4/6 and cyclin E/CDK2 complexes and the consequent dissociation of pRB from E2F. [34] [47] [104] Importantly, tumors that retain wild-type pRB almost always carry activating mutations in cyclin D1 or CDK4 or inactivating mutations in the CDK4-inhibitor, p16.[104] This suggests that the functional inactivation of pRB, and the resulting deregulation of E2F, is an essential step in tumorigenesis.

Studies to date have identified eight E2f genes that encode nine different E2F proteins.[99] pRB and its relatives p107 and p130 (collectively called the pocket proteins) regulate a subset of the E2Fs: E2F1, E2F2, E2F3a, E2F3b, E2F4, and E2F5. These E2F proteins associate with a dimerization partner, called DP, and the resulting complexes function primarily as either activators (E2F1, E2F2, and E2F3a) or repressors (E2F4 and E2F5) of transcription under the direction of the pocket proteins.[47] Most classic E2F target genes are regulated by the coordinated action of these repressor and activator E2Fs ( Fig. 4-5 ). In G0/G1 cells, the DP-E2F4 and DP-E2F5 complexes associate with the promoters of E2F-responsive genes and recruit p107 and p130, along with their associated HDACs, to actively repress their transcription. [105] [106] At the same time, the activating E2Fs are bound by pRB, inhibiting their potential to activate transcription. Whether complexes containing pRB and activating E2Fs contribute to the repression of E2F target genes is still unclear.[107] In response to mitogenic signaling, CDK activity increases, and the phosphorylation of the pocket proteins causes them to release their associated DP-E2Fs. E2F4 and E2F5 dissociate from the DNA and translocate to the cytoplasm because they have potent nuclear export signals. [108] [109] The free E2F complexes—DP-E2F1, DP-E2F2, and DP-E2F3—now occupy the promoters and activate their transcription. Thus, in every cell cycle, there is a coordinated switch from the repressive to the activating E2Fs that enables the simultaneous activation of genes promoting cell cycle progression. Since cyclin E is itself an E2F-responsive gene, this regulation creates a strong feedforward loop: The appearance of a small amount of the cyclin E/CDK2 kinase promotes pRB inactivation and further cyclin E expression. This signal is further amplified by cyclin E/CDK2′s ability to phosphorylate p27 and signal its destruction. Importantly, pRB-inactivation is largely synonymous with the restriction point, defined as the point at which cells become committed to divide even in the absence of mitogenic stimuli. [110] [111] Consistent with this model, the exogenous expression of any individual activating E2F in cell culture is sufficient to stimulate DNA synthesis in the absence of growth signals. [112] [113] [114] [115] [116]

 
 

Figure 4-5  The retinoblastoma protein (pRB) and the restriction point. The pocket proteins—pRB, p107, and p130—regulate a subset of the E2F family of transcription factors. The pocket proteins bind to these E2Fs during G1 and suppress their activity through two mechanisms. First, pRB binds to E2F1, E2F2, and E2F3a (collectively called the activating E2Fs) and blocks their transcriptional activity. Second, p107 and p130 associate with E2F4 and E2F5 (together called the repressive E2Fs), and the resulting complexes recruit histone deacetylases (HDACs) to the promoters of E2F-responsive genes and actively repress their transcription. E2F-responsive genes encode core components of the cycle control and DNA replication machinery, and cell cycle entry is impossible without these products. Mitogenic signaling leads to the sequential activation of cyclin D–CDK4/6 and cyclin E/CDK2, and these phosphorylate the pocket proteins and release their associated E2Fs. This causes the repressive E2Fs to dissociate from E2F-responsive promoters and allows the activating E2Fs to bind and activate their transcription.

 

 

DNA Replication

The DNA replication machinery is optimized to ensure that the genome is copied once—and only once—in each cell cycle. [117] [118] [119] [120] This is achieved through a two-step process that first establishes a prereplication complex (pre-RC) at each origin of replication, a process that is frequently referred to as origin licensing, and subsequently transforms pre-RCs into the preinitiation (pre-IC) complex that activates DNA replication ( Fig. 4-6 ). These two steps occur at distinct stages of the cell cycle to ensure that origins are only licensed once per cell cycle, and rereplication cannot occur. Pre-RC formation takes place during G1. The first step in this process is the recruitment of the multiprotein complex called the origin recognition complex (ORC) to the origin DNA. [121] [122] Although ORC binds a subset of genomic sites, there is no evidence that ORC exhibits sequence-specific DNA binding, and it is still unclear how ORC is recruited to specific sequences. Once bound, ORC recruits additional proteins including Cdc6, Cdt1, and finally the MCM complex, a helicase that is required to unwind the DNA strands to form the pre-RC. Once cells enter S phase, the transformation of the pre-RC to the pre-IC requires the activity of two kinases: a CDK (likely, but not yet proven, to be cyclin A/CDK2) and the Ddf4-dependent kinase, which is composed of the Dbf4 regulatory subunit and the Cdc7 kinase. [123] [124] In mammals, the precise target(s) of these kinases is still unclear. However, the action of these kinases allows numerous additional proteins to associate with the pre-RC and form the pre-IC.[125] Assembly of the pre-IC is thought to trigger DNA unwinding by the MCM complex, recruitment of the DNA polymerases, and initiation of the replication process, frequently called origin firing.

 
 

Figure 4-6  Origin licensing and firing. The origin replication complex (ORC) associates with replication origins. During G1, Cdc6 and Cdt1 are loaded on chromatin, and they in turn load the MCM complex on chromatin, at which point licensing is considered complete, and the multiprotein complex is called the pre-RC. Once cells pass the G1-to-S transition, this complex is activated to form the pre-IC, and DNA replication is initiated. Activation requires both CDK and Ddf4-dependent kinase activity. It results in recruitment of numerous proteins and activation of the MCM complex, which unwinds the DNA. Subsequently, core components of the replication machinery, including DNA polymerase a and DNA polymerase e, are recruited to initiation sites. The transition from pre-RC to pre-IC results in inhibition of cdt1 by ubiquitin-mediated degradation and geminin binding. Origin licensing cannot occur again until activation of APC at the end of mitosis allows accumulation of cdt1.

 

 

The transformation of the pre-RC to the pre-IC can occur at different time points in S phase, depending on whether the origin fires early or late. [118] [119] [120] The system can tolerate this heterogeneity because the pre-RC is disassembled after firing and cannot reform until the subsequent cell cycle. This occurs through several mechanisms. The MCM complex travels with the replication fork in its role as the DNA helicase. There is also some evidence that phosphorylation of ORC1 reduces its ability to bind to origins. Finally, and most important, Cdt1 is prevented from participating in pre-RC formation outside of G1 phase in two distinct ways. First, Cdt1 is marked for destruction by ubiquitination.[126] This is mediated by SCFSkp2 and particularly by an E4 ubiquitin ligase that includes Rbx1 (to recruit the E2-ubiquitin), a cullin (Cul4), Ddb1, and Dtl/Cdt2 (the substrate specificity factor). [127] [128] [129] Importantly, this Cul4-Ddb1Dtl/Cdt2 complex functions independently of Cdt1 phosphorylation. Instead, Cdt1 is targeted only when proliferative cell nuclear antigen is present on the DNA, which occurs primarily as a consequence of the initiation of DNA replication.[130] Second, cells possess a protein called geminin that sequesters Cdt1 and prevents it from participating in pre-RC formation. Geminin is present specifically in S, G2, and early M phase cells. However, the two major mitotic APC complexes, APCCdc20 and APCCdh1, ubiquitinate geminin and thereby trigger its destruction. This creates a window between anaphase of mitosis and late G1 (when APCCdh1 is inactivated) in which geminin is absent and therefore Cdt1 is free to participate in pre-RC formation. The importance of both the Cul4-Ddb1Dtl/Cdt2 complex and geminin is underscored by the finding that the loss of either one of these regulators is sufficient to trigger inappropriate Cdt1 accumulation and rereplication of the genome. [128] [129] [131] [132]

Mitosis

The mitotic machinery is optimized to ensure that the replicated chromosomes are faithfully segregated to the daughter cells. This is achieved through the use of a specialized microtubule-based structure, the mitotic spindle, on which the original chromosomes and their newly replicated copies, called sister chromatids, align and are then partitioned to opposite poles of the cell. The appropriate side-by-side alignment of the sister chromatids, termed biorientation, is facilitated by the physical tethering of the sister chromatids to one another. This process, called cohesion, actually occurs in S phase in a manner that is coordinated with the replication process. [133] [134] Cohesin is mediated by four proteins that together make up the cohesin complex. Two of these proteins, Smc1 and Smc3, have a long coiled-coil structure with a dimerization domain at one end that allows them to heterodimerize to form a V-like structure. Importantly, the remaining ends of Smc1 and Smc3 can associate with each another to form a functional ATP domain. This acts in an ATP-dependent manner to recruit two additional proteins, Scc1 and Scc3, that form a closed ring structure.[135] It is still unclear precisely how this ring links the sister chromatids; some investigators hypothesize that the cohesin complex encircles the chromosomes; others argue that the ring (which is known to be approximately 50 nM) is too small to surround complex chromatin structures. Regardless of the mechanism, the cohesin complex links the sister chromatids at the centromeres and at periodic intervals along the arms. The sister chromatids are essentially strung out and become entangled. Consequently, the chromosome structure must be modified before segregation. This occurs toward the end of G2 and the beginning of mitosis. Largely on the basis of morphologic features, mitosis is divided into five different stages—prophase, prometaphase, metaphase, anaphase, and telophase ( Fig. 4-7 )—prior to separation of the daughter cells or cytokinesis.

 
 

Figure 4-7  Key stages of mitosis. As the parent cell enters prophase, the chromosomes begin to condense, and proteins associate to form the kinetochores. The centrosomes segregate to the poles to begin formation of the mitotic spindle. Nuclear envelope (NE) breakdown denotes the start of prometaphase. In this phase, the sister chromatids continue to condense, and they attach to spindle microtubules via their kinetochores. During metaphase, the sister chromatids align at the metaphase plate and eventually achieve appropriate biorientation. At the onset of anaphase, the sister chromatids separate and move toward the poles of the spindle. During telophase, the parent cell is divided into two daughter cells by cytokinesis.

 

 

Prophase is essentially a preparative stage. One of the major events is the modification of the DNA. In a process called resolution, the sister chromatids are untangled via the action of topoisomerase II. [133] [134] Resolution requires removal of the chromosome arm cohesin through phosphorylation of Scc3 by polo-like kinase and histone H3 by the aurora B kinase. Importantly, the cohesin complex at the centromere is somehow protected from this modification by a protein called shugosin (Sgo).[136] This is the glue that keeps the sister chromatids together until the appropriate point in mitosis. In addition to resolution, the sisters undergo condensation, essentially packaging into a more compact chromatin structure. This process involves two multimeric complexes, condensin I and II, which also contribute to sister chromatid resolution, and it requires phosphorylation by mitotic CDKs. [133] [134] During prophase, the nuclear envelope is still intact; consequently, differences in subcellular localization of the condensin and CDK complexes allow only condensin II and cyclin A/CDK1 (nuclear), and not condensin I and cyclin B/CDK1 (cytoplasmic), to initiate condensation. The second major event in prophase is the activation of the cytoplasmic cyclin B1/CDK1. This initiates formation of the mitotic spindle by triggering the centrosomes, which are located in the cytoplasm and are already nucleating microtubules, to segregate to the opposite poles of the nascent spindle. The active cyclin B1/CDK1 complex then translocates into the nucleus. Once there, it phosphorylates components of the nuclear envelope and triggers its breakdown. [137] [138] This defines the transition from prophase to prometaphase.

During prometaphase, the condensation process is accelerated because condensin I and cyclin B/CDK1 now have access to the DNA. The sister chromatids become attached to spindle microtubules through a structure called the kinetochore, which is assembled onto centromeric DNA.[139] Microtubules nucleated from the centrosomes attach to the kinetochore through a process called search and capture, in which individual microtubules grow and shrink until they contact and bind the kinteochore.[140] Typically, one sister chromatid of the pair attaches first, and this attachment is further stabilized through the recruitment of additional microtubules from the same pole of the mitotic spindle to create a kinetochore fiber: highly bundled microtubules bound to the kinetochore. The sister chromatids oscillate in the cell until the second sister chromatid is captured by microtubules emanating from the other pole. These oscillations continue until all of the chromosomes are properly aligned on the metaphase plate during metaphase.

Metaphase is defined as the point at which all of the chromosome pairs are fully condensed, attached to the mitotic spindle, and aligned at the center—termed the metaphase plate. The pulling of the kinetochore fibers toward the poles creates tension through the cohesin complex at the kinetochores that indicates that the sister chromatids have achieved appropriate biorientation. The cell constantly monitors the attachments of microtubules to the chromosomes, and the tension that is generated by microtubules on the kinetochores ensures that the sister chromatids are properly aligned at the metaphase plate.[141] This is one of several cell cycle checkpoints, called the mitotic spindle checkpoint, that we will describe in more detail in the following sections.

Anaphase is characterized by the segregation of the chromosomes. This event is controlled by the mitotic ligase APCCdc20. [75] [76] [141] [142] [143] [144] APCCdc20 ubiquitinates, and thereby triggers the degradation of, a protein called securin that exists to bind and inhibit a protease called separase. Once released, separase cleaves the Scc1 component of the cohesin complex. This opens the cohesin ring, unlinking the sister chromatids and allowing them to be pulled to opposite poles. The spindle poles then move farther apart to ensure that the chromosomes are fully segregated. APCCdc20 also activates the ubiquitination and degradation of geminin, allowing accumulation of Cdt1 for origin relicensing in the subsequent G1 phase, and the mitotic cyclins, allowing loss of CDK kinase activity. This latter event is critical to the completion of mitosis and cytokinesis.

During telophase, the mitotic spindle disassembles, leaving a single centrosome and a single set of chromosomes with each nascent daughter cell. As the DNA begins to decondense the nuclear envelope reforms around the segregated chromosomes to create two nuclei. These events are dependent on the loss of CDK kinase activity and the dephosphorylation of CDK substrates.

Finally, the cell undergoes cytokinesis, or cytoplasmic division. This involves formation of an actin- and myosin-containing structure, called the contractile ring, on the inner face of the cell membrane. The position of the contractile ring is carefully controlled. For most mammalian cells (ones that are not undergoing asynchronous division), the ring begins to form in anaphase and its position is established by the position of the metaphase plate. As the membrane grows, the contractile ring contracts steadily to form a constriction, termed the cleavage furrow, which ultimately separates the two nuclei and forms the two daughter cells. These cells can adopt either a G0 or a G1 state, depending on the extrinsic signals that exist.

CELL CYCLE CHECKPOINTS

At key transitions during eukaryotic cell cycle progression, signaling pathways monitor the successful completion of events in one phase of the cell cycle before proceeding to the next phase. These regulatory pathways are commonly referred to as cell cycle checkpoints. [145] [146] [147] In a broader context, cell cycle checkpoints are signal transduction pathways that link the rate of cell cycle phase transitions to the timely and accurate completion of prior dependent events. Checkpoint surveillance functions are not confined to monitoring normal cell cycle progression; they are also activated by both external and internal stress signals. To minimize the possibility of errors, checkpoints exist at four different points in the cell cycle: G1/S, intra-S, G2/M, and at the metaphase to anaphase transition (called the spindle checkpoint).

The best-studied of the cell cycle checkpoints are those that monitor the status and structure of chromosomal DNA during cell cycle progression. [147] [148] [149] In particular, cells scan the chromatin for partially replicated DNA as well as DNA strand breaks and other DNA lesions that can result from both extrinsic (e.g., chemicals, ionizing or ultraviolet radiation) and intrinsic (e.g., by-products of intracellular metabolism) DNA-damaging agents. The checkpoint pathways include sensor proteins that detect these DNA lesions and simultaneously trigger two processes: They recruit additional complexes to repair the DNA and activate signaling pathways that induce a temporary cell cycle arrest. In certain situations, which are determined by the cell type and the degree of damage, the checkpoint pathways can induce permanent cell cycle arrest (a process called senescence) or apoptosis.

The central components of the DNA damage response (DDR) are two members of the phosphoinositide 3-kinase-related kinase family: ATM and ATR. [147] [148] ATM was original identified by virtue of its mutation in a hereditary syndrome, ataxia-telangiectasia, which is associated with radiation hypersensitivity and cancer predisposition.[150] ATR is also associated with a hereditary syndrome called Seckel syndrome. Early studies suggested that ATM and ATR played distinct roles in the response to double-stranded DNA breaks (ATM) versus replicative defects and single-stranded breaks (ATR). However, we now know that the regulation is more complex; there is considerable cross-talk between ATM and ATR, and they share many mediators and effectors, but the precise composition and role of the DDR complexes vary depending on both the type of the damage and the stage of the cell cycle.[151] In this chapter, we focus primarily on how the DDR activates cell cycle checkpoints ( Fig. 4-8 ). However, it is important to note that many of the components of the core DDR machinery are affected in hereditary disease syndromes and/or abrogated in human tumors (e.g., NBS1, BRCA1, BRCA2, and the Franconi's anemia proteins).

 
 

Figure 4-8  ATM/ATR signaling is activated by DNA damage and replication stress. The cell constantly monitors the chromatin for lesions, using complex signal transduction pathways that center on the ATM and ATR kinases. The precise mechanism of response varies according to the type of DNA damage and the cell cycle stage. Double-stranded breaks (DSBs) are the most deleterious form of DNA damage. DSBs are recognized by the MRN complex that consists of Mre11, Rad50, and Nbs1. This complex recruits ATM to the site of damage. ATM phosphorylates histone H2AX, to form gH2AX, and this creates a binding platform for additional proteins that propagate the DNA damage response and activate repair. For S and G2 phase cells, but not G1 cells, ATR is also recruited to the damage site. ATR and/or ATM signal to their effector kinases—CHK1 and CHK2—respectively, to influence cell cycle progression as described in Figure 4-9 . Errors in DNA replication can also activate the DNA damage response machinery through the presence of single-stranded DNA (ssDNA) that is a hallmark of the replication fork. The ssDNA is coated with RPA and bound by ATR. Active ATR then recruits the DNA damage and repair machinery, including ATM, leading to the sequential activation of CHK1 and then CHK2.

 

 

G1/S Checkpoint

In G1 cells, double-stranded DNA breaks (DSBs) are the most common and most deleterious type of DNA damage. These DSB breaks are recognized by the multifunctional Mre11-Rad50-Nbs1 (MRN) complex.[147] This complex recruits ATM to the site of damage. It is still unclear whether ATM activation occurs before or in response to MRN binding. The active ATM then recruits proteins to modify the chromatin at the region of the break and activate repair and signaling. As a first step in this process, ATM phosphorylates histone H2AX, to form γH2AX. This helps to hold the damaged ends together and acts as a binding platform for additional factors, including Mdc1, 53BP1, and BRCA1, as well as more MRN and ATM. In contrast to the S and G2 response, there is no recruitment of ATR to DSB in G1cells; therefore, ATM is solely responsible for checkpoint activation. The recruitment of additional ATM amplifies the signal, and ATM acts via phosphorylation and activation of the effector kinase CHK2.[152] [153]

CHK2 influences G1 cell cycle arrest via two mechanisms ( Fig. 4-9 ). First, it phosphorylates all three members of the Cdc25 family. Phospho-Cdc25A is ubiquitinated by the SCFTrcpβ and degraded, while phospho-Cdc25B and phospho-Cdc25C are bound and sequestered by a cytoplasmic protein called 14–3-3. [10] [154] [155] This is a rapid response that can take effect within minutes after DNA damage, and it has a widespread effect on cell cycle progression by preventing activation of all CDK2 and CDK1 complexes. In the case of the G1/S checkpoint, cyclin E/CDK2 is the relevant target. Second, CHK2 phosphorylates p53, a critical regulator of cell cycle checkpoints.[156] In normal, nonstressed cells, p53 protein is maintained at low steady-state levels because it has a very short half-life. This half-life is a result of rapid ubiquitination of p53 by HDM2 (the human ortholog of murine MDM2 protein) and its consequent degradation. The importance of MDM2 for maintenance of appropriate p53 levels in vivo is highlighted by the fact that absence of MDM2 in knockout mice results in early embryonic lethality that is rescued by a dual knockout of MDM2 and p53.[157] Phosphorylation of p53 by CHK2 is sufficient to prevent its association with HDM2/MDM2.[158] This leads to an accumulation of p53, which functions as a transcriptional activator. p53 induces expression of many genes. One of the key targets for the G1/S (and also G2/M) checkpoint is the CDK inhibitor p21Cip1. [159] [160] This p53-mediated arrest takes longer to develop than Cdc25 response (because it requires transcription and protein synthesis) but appears to be much more robust. Moreover, in addition to inducing cell cycle arrest, p53 has the capacity to induce apoptosis through the transcriptional activation of proapoptotic regulators (e.g., the BH3-only proteins PUMA and NOXA).[161] How p53 chooses to activate arrest versus apoptosis targets is not fully understood, but it is clearly influenced by both the cell type and the level of damage.[161]

 
 

Figure 4-9  DNA damage, replicative stress, and oncogenic stress induce cell cycle arrest. DNA damage and replication stress lead to the rapid phosphorylation and activation of the CHK1 and/or CHK2 kinases. These enforce cell cycle arrest through two mechanisms. CHK1 and CHK2 both phosphorylate the cdc25 phosphatases, and this triggers their ubiquitination and degradation (cdc25A) or binding and inhibition by 14-3-3 (cdc25B and cdc25C), thereby preventing activation of either cyclin/CDK2 or cyclin/CDK1 kinases. CHK1 and CHK2 also phosphorylate p53 and prevent it from being targeted by HDM2 for ubiquitin-mediated degradation. As a result, p53 accumulates and activates transcription of p21Cip1, inhibiting CDK2 and CDK1 kinase complexes, or proapoptotic genes. Oncogenic stress also leads to cell cycle arrest by activating replicative stress and/or inducing transcription or the p14Arf tumor suppressor and suppressing HDM2-mediated inhibition of p53.

 

 

Importantly, p53 is also activated by other stress signals (see Fig. 4-9 ). In particular, it is now well established that numerous oncogenes trigger a stress response (called oncogene-induced stress) that leads to the activation of p53. [162] [163] [164] [165] The emerging view is that this occurs through two distinct mechanisms. First, oncogene activation is thought to yield replicative stress that activates p53 via activation of CHK kinases and phosphorylation of HDM2/MDM2 as just described. [166] [167] Second, many oncogenes activate transcription of Arf.[168] This gene is encoded by the INK4a/Arf locus, and it actually shares two coding exons, which are read in alternate reading frames (hence the name ARF), with the p16Ink4A tumor suppressor.[169] The Arf protein product, called p14Arf in humans and p19Arfin mouse, binds to HDM2/MDM2 and prevents it from regulating p53. [170] [171] [172] [173] [174] As with the DDR, this frees p53 to activate the transcription or proarrest or proapoptotic targets. The central importance of this p53 pathway is underscored by the finding that the majority of human tumors carry mutations in p53, have upregulated HDM2 (typically by gene amplification), or have inactivated p19Arf.[169] This is very analogous to the selective pressure to deregulate pRB pathway components (pRB, cyclin D/CDK4, and p16Ink4A) that was discussed previously.[104] Together, the pRB and p53 pathways are critical gatekeepers of G1-to-S progression in normal cell cycle and stress response.

As an additional DNA damage response in G1 cells, genotoxic agents also inhibit origin licensing by way of an ATM/ATR-independent process. This is achieved through regulation of Cdt1. [127] [128] [129] [175] As was described previously, Cdt1 is required for pre-RC formation. In an undamaged cell, Cdt1 is available during G1 but is inhibited after origin firing by degradation (mediated by the SCFskp2 and Cul4-Ddb1-Dtl/Cdt2 ubiquitin ligases) and geminin binding. As a key feature of this regulatory system, Cdt1 is completely resistant to Cul4-Ddb1-Dtl/Cdt2 in the G1 phase. However, DNA damage allows Cul4-Ddb1-Dtl/Cdt2 complex to ubiquitinate Cdt1 and induce its degradation. This process requires binding of Cdt1 to proliferative cell nuclear antigen, but the mechanism by which this induces Cdt1 ubiquitinylation is not understood. Importantly, the degradation of Cdt1 is extremely rapid, occurring within minutes of the DNA damage. As a result, origin licensing is completely blocked until the damage is repaired and Cdt1 is resynthesized.

Intra-S Phase Checkpoint

One of the major goals of cell cycle checkpoints is to prevent the deleterious consequences of replicating damaged DNA. Therefore, S phase cells must respond virtually instantaneously to DNA damage to halt initiation of new replication forks throughout the S phase.[149] The most deleterious damage is DSBs. These can occur through the action of DNA damaging agents (from either extrinsic or intrinsic sources) or as a consequence of the replication process itself, for example, if the replication fork passes through nicked DNA or replication stalls at sites of DNA damage. The cell senses the damage in different ways depending on whether or not the lesion is associated with replication. Ultimately, both ATM and ATR are recruited to the site of damage, but the order of binding is different. [149] [151]Replication-linked DSBs are distinguished by the presence of single-stranded DNA, a hallmark of the replication process. The single-stranded DNA is coated by RPA and bound by ATR and its regulator subunit ATRIP, even during the normal replication process. In response to DNA damage, the ATR kinase is activated, and it then recruits a variety of complexes that mediate both repair and checkpoint activation, including ATM. In contrast, nonreplication-associated DSBs initially recruit and activate ATM through the MRN-dependent process described previously for G1/S checkpoint. However, in S phase cells, DSB resection causes the formation of single-stranded DNA (through the action of the MRN endonuclease), and this is then bound by RPA and ATR/ATRIP. [149] [151] Thus, in S phase cells, ATR and ATM jointly orchestrate the DDR. ATR contributes to the checkpoint response in a similar manner to ATM: It activates an effector kinase, called CHK1, which can also phosphorylate the cdc25 proteins and p53. [176] [177] [178] [179]

G2 Checkpoint

The G2 checkpoint is required to prevent the passage of DNA lesions to two daughter cells during mitosis. [147] [180] DSBs are detected exactly as we described previously for the S phase nonreplication-associated DSBs. Similarly, the ATR/CHK1 and ATM/CHK2 pathways enforce arrest through inhibition of G2 and mitotic CDK complexes via the rapid removal of the cdc25 phosphates and the p53-dependent induction of the p21Cip1 CDKI.

Spindle Checkpoint

The preceding sections focused on the steps the cell takes to prevent the propagation of DNA errors to the daughter cells. In contrast, the spindle checkpoint acts to ensure that there is appropriate partitioning of the chromosomes.[181] We have already introduced the concept that chromosome segregation is prevented until all of the condensed sister chromatid pairs are aligned at the metaphase plate with the appropriate biorientation. This is actually controlled by a signaling network that constitutes the spindle checkpoint ( Fig. 4-10 ). The core components of the spindle checkpoint—called MAD1, MAD2, BUBR1, and BUB1 in humans—were originally identified through screens in yeast for “mitotic arrest deficient” (MAD) and “budding uninhibited by benzimidazole” (BUB) mutants.[181] These proteins become active in the prometaphase of mitosis (see Fig. 4-10 ). They associate with the kinetochore and, in the absence of biorientation, prevent the CDC20 activator from binding to the APC. As a result, separase is sequestered by securin and unable to cleave the centromeric cohesin (see Fig. 4-10 ). It is still unclear precisely how the spindle checkpoint is inactivated by appropriate biorientation. However, it involves monitoring the tension through the cohesin complex at the kinetochores (created by the pulling of the spindle fibers toward the poles) and the dissociation of MAD2 from the attached kinetochore (see Fig. 4-10 ). Because aneuploidy is a shared feature of many cancer cells, there has been considerable speculation that disruption of the spindle checkpoint could occur during tumor progression. [182] [183] [184] Notably, inactivating mutations in Bub1 have been identified in human colon carcinoma cell lines, which are known to have a high degree of aneuploidy.[185] Moreover, haploinsufficiency of Mad2 has been shown to cause elevated rates of lung tumor development in Mad2+/- mice compared with age-matched wild-type mice.[186] However, it is still an open question whether spindle checkpoint defects make a significant contribution to tumor development.

 
 

Figure 4-10  The spindle checkpoint. Improper chromosome alignment on the mitotic spindle, disruption of microtubule dynamics, or unattached kinetochores can activate the spindle checkpoint. Spindle checkpoint signaling is mediated by the Bub1, Bub3, BubR1, and Mad2 proteins, which all localize to kinetochores. These core spindle checkpoint regulators prevent the activator protein Cdc20 from binding to APC and therefore protects securin, a major APCcdc20 target, from ubiquitin-mediated degradation. As a result, securin remains bound to separase, and this prevents cleavage of Scc1 and loss of centromeric cohesin. The spindle checkpoint is relieved at the end of the metaphase by the appropriate biorientation of the sister chromatids at the metaphase plate. The sensing mechanism involves detecting tension through the cohesin complex at the kinetochores that is created by the pulling of the spindle fibers toward the poles. Mad2 then dissociates from the attached kinetochore, and this allows cdc20 to activate APC and trigger sister chromatid segregation.

 

 

CELL CYCLE DEREGULATION IN HUMAN CANCERS

Molecular analysis of human tumors demonstrates that alterations in components of the cell cycle machinery and checkpoint-signaling pathways occur in the majority of human tumors ( Table 4-1 ). This finding underscores how important maintenance of cell cycle control is in the prevention of human cancer. The alterations in the cell cycle machinery that occur most frequently include loss or mutation of the pRB tumor suppressor; overexpression of cyclins, CDKs, and Cdc25 phosphatases; and loss of expression of CDKIs. The most frequently altered cell cycle checkpoint-signaling molecule is the p53 tumor suppressor. Proteins that reside upstream of p53 (including ATM and CHK2) are also targeted for mutation in human tumors, and their discovery and analysis have greatly deepened our insight into DNA damage response-signaling pathways. Mutations that affect the pRB pathway have been identified in the majority of human cancers. [91] [187] The RB-1 gene was originally identified by virtue of its mutation in both familial and sporadic retinoblastoma, but it is defective in many other tumor types, especially osteosarcoma and lung cancer. Indeed, more than 90% of small-cell lung cancers have mutantRB-1, suggesting that disruption of the pRB pathway (through the genetic or epigenetic targeting of RB-1 or upstream signaling components) is a requirement for the genesis of lung cancer.[188] It is important to note that inactivation of the parallel and interconnecting p14Arf-p53 axis is also essential in functionally pRB-deficient lung cells to bypass efficient apoptosis.[169] In breast cancer, loss of normal pRB function due to RB-1 mutation is observed in 20% of tumors.[189] In the 80% of breast carcinomas that lack RB-1 mutations, alterations in components of the signaling pathways that regulate pRB are frequently found, including cyclin D1 and cyclin E overexpression and cdk4 and cdk6 gene amplification. [190] [191] [192] Nearly 50% of invasive breast cancers have elevated cyclin D expression compared with surrounding normal breast epithelium, while transgenic mice with overexpression of human cyclin D1 or cyclin E in mammary gland cells develop mammary adenocarcinomas. [193] [194] [195] Similarly, cdk4 and cdk6 gene amplification occurs in breast cancers, sarcomas, gliomas, and melanomas.[196]


Table 4-1   -- Mutations of Cell Cycle Checkpoint Regulators in Human Tumors[*]

Gene/Protein

Tumors Associated with Mutations or Altered Expression

Hereditary Syndromes Associated with Germline Mutations

ATM

Breast carcinomas, lymphomas, leukemias

Ataxia-telangiectasia

Bub1

Colorectal carcinomas

NR

BRCA1

Breast and ovarian carcinoma

Familial breast and ovarian cancer

Cdc25A

Carcinomas of breast, lung, head and neck, and lymphoma

NR

Cdc25B

Carcinomas of breast, lung, head and neck, and lymphoma

NR

Cdk4

Wide array of cancers

NR

Cdk6

Wide array of cancers

NR

Chk1

Colorectal and endometrial carcinomas

NR

Chk2

Carcinomas of breast, lung, colon, urogenital tract, and testis

Li-Fraumeni syndrome

Cyclin D1

Wide array of cancers

NR

Cyclin D2

Lymphoma and carcinomas of the colon, testis and ovary

NR

Cyclin D3

Lymphoma, pancreatic carcinoma

NR

Cyclin E

Wide array of cancers

NR

MDM2

Soft tissue tumors, osteosarcomas, esophageal carcinomas

NR

MRE11

Lymphoma

Ataxia-telangiectasia-like disorder

NBS

Lymphomas, leukemias

Nijmegen breakage syndrome

p15INK4b

Wide array of cancers

NR

p16INK4a

Wide array of cancers

Familial melanoma

p27KIP1

Wide array of cancers

NR

p53

Wide array of cancers

Li-Fraumeni syndrome

p57KIP2

Bladder carcinomas

NR

p130

Wide array of cancers

NR

pRB

Wide array of cancers

Familial retinoblastoma

NR, not reported.

 

*

Only alterations that are present in more than 10% of primary tumors are represented.

 

Modifications of CDKIs that act upstream of pRB activity are also commonly found in human tumors. The CDK inhibitor p27Kip1 is often aberrantly expressed in human breast cancer, and reduced p27Kip1protein levels are correlated with more aggressive breast tumors. [197] [198] Likewise, decreased expression of the CDK inhibitor p57Kip2 is found in human bladder cancers.[199] Germline mutations inp16INK4a predispose individuals to melanoma, while deletion of p15INK4b and p16INK4a is linked to the pathogenesis of lymphomas, mesotheliomas, and pancreatic cancers. [78] [79] [196] [200] In tumor types in which p15INK4b and p16INK4a are not deleted, methylation of the gene locus leads to transcriptional repression and loss of gene expression. In some tumors, hypermethylation prevents expression of both p16INK4a and p14Arf, which are encoded by alternative reading frames of the Ink4a/Arf locus.[199] Both Cdc25A and Cdc25B phosphatases are overexpressed in more than 30% of primary breast tumors, 40% to 60% of non-small-cell lung cancers, 50% of head and neck tumors, and a significant fraction of non-Hodgkin's lymphomas. [10] [201] [202] Elevation of these oncogenic phosphatases can result in increased activation of CDK and override of checkpoint arrest.

p53 mutation is the most frequently observed mutation in the majority of human tumors. The importance of p53-dependent signaling in tumor suppression is underscored by the frequency of mutation in sporadic tumors and the finding that germline mutations of p53 result in Li-Fraumeni syndrome, a highly penetrant familial cancer syndrome that is associated with significantly increased rates of brain tumors, breast cancers, and sarcomas. [203] [204] In human tumors that lack p53 gene mutation, p53 function may be disrupted by alterations in cellular proteins that modulate the levels, localization, and biochemical activity of p53. For example, in some tumors with wild-type p53 alleles, MDM2 gene amplification occurs, resulting in MDM2 protein overexpression and subsequent p53 inactivation.[205] In human papillomavirus-induced cervical carcinoma, p53 is typically not mutated; however, the human papillomavirus E6 protein binds p53 and targets it for degradation, abrogating p53-dependent signaling.[206]

Mutation in components of the DNA damage response pathway also leads to enhanced tumorigenesis, as was discussed previously. For example, ATM mutations occur in ataxia-telangiectasia, a disorder in which patients have increased sensitivity to radiation and an elevated incidence of leukemias, lymphomas, and breast cancer. [150] [207] ATM-null mice exhibit growth retardation, neurologic dysfunction, infertility, defective T lymphocyte maturation, and sensitivity to ionizing radiation. [208] [209] The majority of ATM-deficient animals develop malignant lymphomas by 4 months of age, while ATM-/-fibroblasts have abnormal radiation checkpoint function after exposure to ionizing radiation. [208] [209] The DNA double-strand break repair gene MRE11 is mutated in individuals with an ataxia-telangiectasia-like disorder.[210] Mutations of Chk2 and Chk1 also arise in human cancers. Chk2 mutations have been reported in several cancers, including lung, while Chk1 mutations have been observed in human colon and endometrial cancers. [211] [212] In addition, heterozygous alteration of Chk2 occurs in a subset of individuals with Li-Fraumeni syndrome who lack p53 gene mutations.[213] These findings support the theory that in human tumors in which p53 is intact, the function of this tumor suppressor might be disrupted by alterations in cellular proteins that modulate the levels or activity of p53. In addition, the breast cancer susceptibility tumor suppressors BRCA1 and BRCA2 are known to participate in the DNA damage response and repair.[214] Similarly, the Fanconi's anemia proteins, which were originally identified by virtue of their association with a recessive development disorder called Fanconi's anemia, which is associated with increased cancer predisposition (particularly acute myeloid leukemia), also function in the DNA damage response.[214]

The spindle checkpoint disruption has also been linked to the pathogenesis of several human tumors. BUB1 mutations have been identified in human colon carcinoma cells, and Bub1 mutation facilitates the transformation of cells that lack the breast cancer susceptibility gene, BRCA2. [185] [215] Moreover, Michel and colleagues demonstrate that Mad2+/- mice have significantly higher rates of lung tumor development than do age-matched wild-type mice.[186]

THERAPEUTIC MANIPULATION OF CELL CYCLE CONTROLS

Research over the past two decades has shown that alterations in cell cycle machinery and checkpoint signaling lead to tumorigenesis. These findings have important implications for the optimization of current therapeutic regimens and for the selection of novel cell cycle targets for the future development of anticancer agents. A leading goal of cancer-based research is to identify compounds that will target key cell cycle controls in a tumor-specific manner.

Targeting Cyclin-Dependent Kinase Activity

There has been considerable debate about whether inhibition of CDK activity is a rational strategy for anticancer therapies. CDK activity is frequently elevated in human tumors, but it is also required to maintain specific cells populations in the adult (e.g., the hematopoietic compartment and gut) that are essential for viability. Thus, the key issue is whether there is sufficient difference in the CDK activity in tumor versus normal cells to create a therapeutic window. Over the last few years, the analysis of CDK and cyclin mouse models has yielded considerable insight into this question but has also raised additional questions.[18] On the positive side, studies in mouse models clearly show that tumors can be more dependent on CDK activity, or at least a specific CDK activity, than normal tissues can. For example, loss of D-type cyclins has been shown to have little or no effect on the development and maintenance of many tissues, but loss of cyclin D-associated kinase activity can greatly suppress the development of certain tumor types, depending on the tissue and the identity of the initiating oncogenic lesions. [22] [29] [216] On the negative side, the mouse models also show that the cell cycle machinery is extremely robust; it adapts easily to the loss of CDKs or cyclins by using other CDKs or cylins to substitute for the missing activity. For example, CDK2 knockout mice are fully viable because CDK4/6 and CDK1 now form novel cyclin/CDK complexes and assume roles that are normally specific to CDK2. [217] [218] [219] This raises the possibility that tumor cells will rapidly develop resistance to CDK-inhibitory drugs by simply adapting their cell cycle machinery. In light of these complexities, efforts have been placed on generating pharmacologic inhibitors of CDKs that either are CDK-specific or have pan-CDK activities. Numerous small molecule inhibitors have been developed, and many are in clinical trials. [220] [221]

One of the first compounds to be tested, flavopiridol, is a pan-CDK inhibitor that inhibits CDK4/6, CDK2, and CDK1 kinase activity. Consistent with this broad action, flavopiridol arrests cells at G1/S (in a pRB-dependent manner) and G2/M. This antiproliferative activity against a variety of human cancer cell lines produced favorable clinical responses in phase I and phase II studies of patients with renal, colorectal, gastric, lung, and esophageal carcinomas. [222] [223] [224] Notably, it was also determined that if target cells are first induced to induced to enter S phase, then treatment with flavopiridol had significant cytotoxic effects.[221] This arises through two mechanisms. First, flavopiridol inhibits the action of cyclin A/CDK2 and thereby prevents the phosphorylation of E2F1 and its subsequent degradation. [59] [61] The persistence of E2F1 in late stages of the cell cycle is known to trigger apoptosis, and this effect shows strong specificity for tumor cells versus normal cells, presumably because of higher E2F1 levels. Second, flavopiridol suppresses the activity of CDK7 (which functions both as a component of both CAK and as a RNA polymerase II CTD kinase that promotes transcriptional elongation) and CDK9 (which acts in association with cyclin T to form another CTD kinase called P-TEFb). [225] [226] Inhibition of CDK7 and CDK9 suppresses mRNA synthesis, and this leads to a rapid loss of transcripts that have short half-lives, including many cell cycle regulators (e.g., cyclins) and antiapoptosis regulators. As a result of these observations, clinical trails have been conducted using sequential treatment of an S phase chemotherapeutic agent, gemcitabine, and then flavopiridol.[221] On a similar theme, sequential treatment with paclitaxel (which inhibits mitotic spindle function) and then flavopiridol also yields cytotoxic synergy.[221] In this case, flavopiridol is acting by inhibiting cyclin B/CDK1 and thus prevents phosphorylation and stabilization of a protein, called survivin, that is required to maintain the spindle checkpoint. Thus, the cells proceed through cytokinesis and enter G1 without segregating their chromosomes, and this triggers apoptosis. Phase I and phase II trials have been conducted with paclitaxel and flavopiridol, and currents efforts are focused on optimizing the dosage and the time interval of administration.[221] More selective CDK inhibitors are also being analyzed. [220] [221] These include small molecules that show a strong selectively for CDK2 and CDK1 or are highly specific for CDK4/6. Cell- and animal-based studies show that these drugs yield the anticipated affects. For example, the CDK4/6 inhibitor PD0332991 yields a G1/S arrest in an pRB-dependent manner, and it can yield regression of xenografts generated from pRB-positive cell lines.[227]Many of these drugs have yet to be tested in clinical trials.

Targeting DNA Damage Response Proteins

In the last decade, there has been a growing appreciation that many tumors cells carry mutations that disrupt their DNA damage response (DDR). This is a major factor in establishing the resistance of tumors to chemotherapeutic agents, many of which work by causing DNA damage and triggering apoptosis through induction of DNA damage pathways. Therefore, considerable attention has focused on designing cancer treatments that would be effective in cells with an impaired DDR. Since it is hard to restore the function of mutant or missing proteins, the prevailing strategy is to identify drugs that would synergize with the defective DDR to selectively kill the tumor cells and not the normal cells. For example, inhibitors of poly(ADP-ribose) polymerase selectively kill cells that lack either BRCA1 or BRCA2. [228] [229] [230] The rationale for this is that these proteins provide two alternative repair mechanisms in response to DNA damage: homologous recombination (BRCA1 and BRCA2) and base excision repair (poly(ADP-ribose) polymerase). Therefore, loss of one but not both of these pathways can be tolerated. As a second example, inhibition of CHK1 sensitizes p53 mutant cells to DNA damage.[220] Since p53 is mutated in approximately half of all human tumors and the absence of p53 is a major predictor of poor response to classic chemotherapeutic agents, considerable efforts are being made to develop small molecular inhibitors of CHK1.

SUMMARY

Over the past several decades, investigators have uncovered a wealth of information about the proteins that control cell growth and division in human cells. A key finding is that deregulation of the cell cycle machinery and/or checkpoints is a universal alteration that has been identified in human cancer. [104] [231] Although numerous genetic alterations can result in loss of normal checkpoints, the hope is that common strategies will be developed against a wide variety of cancers. Even though several of the currently used anticancer therapies target nonselective and non-mechanism-based targets, their effectiveness, albeit limited in many cases, is likely due to the fact that they ultimately target cell cycle regulatory or DDR-signaling pathways, the status of which is different in normal cells versus tumor cells. Identifying all the components of the cellular machinery that control the cell cycle both positively and negatively is vital to the continued development of anticancer agents that can preferentially eliminate cancer cells and minimize the toxicity to normal tissues. The information that is generated by the genomic and proteomic approaches using eukaryotic model systems will continue to reveal new cell cycle regulatory molecules. As our understanding of cell cycle regulation and checkpoint signaling improves, the goal is to use this knowledge in the design of mechanism-based therapeutics that will bring anticancer therapy to a new level. There can be little doubt of the value of targeting the cell cycle in drug discovery.

REFERENCES

  1. Malumbres M, Barbacid M: Mammalian cyclin-dependent kinases.  Trends Biochem Sci2005; 30:630-641.
  2. Malumbres M, Barbacid M: Cell cycle kinases in cancer.  Curr Opin Genet Dev2007; 17:60-65.
  3. Morgan DO: Cyclin-dependent kinases: engines, clocks, and microprocessors.  Annu Rev Cell Dev Biol1997; 13:261-291.
  4. Lee MG, Nurse P: Complementation used to clone a human homologue of the fission yeast cell cycle control gene cdc2.  Nature1987; 327:31-35.
  5. Sherr CJ, Roberts JM: CDK inhibitors: positive and negative regulators of G1-phase progression.  Genes Dev1999; 13:1501-1512.
  6. Pavletich NP: Mechanisms of cyclin-dependent kinase regulation: structures of Cdks, their cyclin activators, and Cip and INK4 inhibitors.  J Mol Biol1999; 287:821-828.
  7. Evans T, Rosenthal ET, Youngblom J, et al: Cyclin: a protein specified by maternal mRNA in sea urchin eggs that is destroyed at each cleavage division.  Cell1983; 33:389-396.
  8. Russo A, Jeffrey PD, Pavletich NP: Structural basis of cyclin-dependent kinase activation by phosphorylation.  Nat Struct Biol1996; 3:696-700.
  9. Kellogg DR: Wee1-dependent mechanisms required for coordination of cell growth and cell division.  J Cell Sci2003; 116(pt 24):4883-4890.
  10. Boutros R, Lobjois V, Ducommun B: CDC25 phosphatases in cancer cells: key players?.  Good targets? Nat Rev Cancer2007; 7:495-507.
  11. Lolli G, Johnson LN: CAK-Cyclin-dependent activating kinase: a key kinase in cell cycle control and a target for drugs?.  Cell Cycle2005; 4:572-577.
  12. Harper JW, Elledge SJ: The role of Cdk7 in CAK function: a retro-retrospective.  Genes Dev1998; 12:285-289.
  13. Kaldis P: The cdk-activating kinase (CAK): from yeast to mammals.  Cell Mol Life Sci1999; 55:284-296.
  14. Kaldis P, Solomon MJ: Analysis of CAK activities from human cells.  Eur J Biochem2000; 267:4213-4221.
  15. Fisher RP: Secrets of a double agent: CDK7 in cell-cycle control and transcription.  J Cell Sci2005; 118(pt 22):5171-5180.
  16. Sherr CJ: Mammalian G1 cyclins.  Cell1993; 73:1059-1065.
  17. Deshpande A, Sicinski P, Hinds PW: Cyclins and cdks in development and cancer: a perspective.  Oncogene2005; 24:2909-2915.
  18. Lee YM, Sicinski P: Targeting cyclins and cyclin-dependent kinases in cancer: lessons from mice, hopes for therapeutic applications in human.  Cell Cycle2006; 5:2110-2114.
  19. Sherr CJ: D-type cyclins.  Trends Biochem Sci1995; 20:187-190.
  20. Diehl JA, Cheng M, Roussel MF, Sherr CJ: Glycogen synthase kinase-3beta regulates cyclin D1 proteolysis and subcellular localization.  Genes Dev1998; 12:3499-3511.
  21. Aktas H, Cai H, Cooper GM: Ras links growth factor signaling to the cell cycle machinery via regulation of cyclin D1 and the Cdk inhibitor p27KIP1.  Mol Cell Biol1997; 17:3850-3857.
  22. Robles AI, Rodriguez-Puebla ML, Glick AB, et al: Reduced skin tumor development in cyclin D1-deficient mice highlights the oncogenic ras pathway in vivo.  Genes Dev1998; 12:2469-2474.
  23. Tetsu O, McCormick F: Beta-catenin regulates expression of cyclin D1 in colon carcinoma cells.  Nature1999; 398:422-426.
  24. Shtutman M, Zhurminsky J, Simcha I, et al: The cyclin D1 gene is a target of the beta-catenin/LEF-1 pathway.  Proc Natl Acad Sci USA1999; 96:5522-5527.
  25. Bouchard C, Dittrich O, Kiermaier A, et al: Regulation of cyclin D2 gene expression by the Myc/Max/Mad network: Myc-dependent TRRAP recruitment and histone acetylation at the cyclin D2 promoter.  Genes Dev2001; 15:2042-2047.
  26. Yu Q, Ciemerych MA, Sicinski P: Ras and Myc can drive oncogenic cell proliferation through individual D-cyclins.  Oncogene2005; 24:7114-7119.
  27. Kozar K, Ciemerych MA, Rebel VI, et al: Mouse development and cell proliferation in the absence of D-cyclins.  Cell2004; 118:477.471
  28. Cheng M, Sexl V, Sherr CJ, Roussell MF: Assembly of cyclin D-dependent kinase and titration of p27Kip1 regulated by mitogen-activated protein kinase kinase (MEK1).  Proc Natl Acad Sci USA1998; 95:1091-1096.
  29. Landis MW, Pawlyk BS, Li T, et al: Cyclin D1-dependent kinase activity in murine development and mammary tumorigenesis.  Cancer Cell2006; 9:13-22.
  30. Ewen ME, Sluss HK, Sherr CJ, et al: Functional interactions of the retinoblastoma protein with mammalian D-type cyclins.  Cell1993; 73:487-497.
  31. Connell-Crowley L, Harper JW, Goodrich DW: Cyclin D1/Cdk4 regulates retinoblastoma protein-mediated cell cycle arrest by site-specific phosphorylation.  Mol Biol Cell1997; 8:287-301.
  32. Kato J, Matsushime H, Herbert SW, et al: Direct binding of cyclin D to the retinoblastoma gene product (pRb) and pRb phosphorylation by the cyclin D-dependent kinase CDK4.  Genes Dev1993; 7:331-342.
  33. Connell-Crowley L, Elledge SJ, Harper JW: G1 cyclin-dependent kinases are sufficient to initiate DNA synthesis in quiescent human fibroblasts.  Curr Biol1998; 8:65-68.
  34. Adams PD: Regulation of the retinoblastoma tumor suppressor protein by cyclin/cdks.  Biochim Biophys Acta2001; 1471:M123-M133.
  35. Ohtani K, DeGregori J, Nevins JR: Regulation of the cyclin E gene by transcription factor E2F1.  Proc Natl Acad Sci USA1995; 92:12146-12150.
  36. Geng Y, Eaton EN, Picón M, et al: Regulation of cyclin E transcription by E2Fs and retinoblastoma protein.  Oncogene1996; 12:1173-1180.
  37. Tsai LH, Lees E, Faha B, et al: The cdk2 kinase is required for the G1-to-S transition in mammalian cells.  Oncogene1993; 8:1593-1602.
  38. Koff A, Giordano A, Desai D, et al: Formation and activation of a cyclin E-cdk2 complex during the G1 phase of the human cell cycle.  Science1992; 257:1689-1694.
  39. Koff A, Cross F, Fisher A, et al: Human cyclin E, a new cyclin that interacts with two members of the CDC2 gene family.  Cell1991; 66:1217-1228.
  40. Ye X, Wei Y, Nalepa G, Harper JW: The cyclin E/Cdk2 substrate p220(NPAT) is required for S-phase entry, histone gene expression, and Cajal body maintenance in human somatic cells.  Mol Cell Biol2003; 23:8586-8600.
  41. Zhao J, Dynlacht B, Imai T, et al: Expression of NPAT, a novel substrate of cyclin E-CDK2, promotes S-phase entry.  Genes Dev1998; 12:456-461.
  42. Zhao J, Kennedy BK, Lawrence BD, et al: NPAT links cyclin E-Cdk2 to the regulation of replication-dependent histone gene transcription.  Genes Dev2000; 14:2283-2297.
  43. Hinchcliffe EH, Li C, Thompson EA, et al: Requirement of Cdk2-cyclin E activity for repeated centrosome reproduction in Xenopus egg extracts.  Science1999; 283:851-854.
  44. Hinchcliffe EH, Sluder G: Centrosome duplication: three kinases come up a winner!.  Curr Biol2001; 11:R698-R701.
  45. Lacey KR, Jackson PK, Stearns T: Cyclin-dependent kinase control of centrosome duplication.  Proc Natl Acad Sci USA1999; 96:2817-2822.
  46. Okuda M, Horn HF, Tarapore P, et al: Nucleophosmin/B23 is a target of CDK2/cyclin E in centrosome duplication.  Cell2000; 103:127-140.
  47. Trimarchi JM, Lees JA: Sibling rivalry in the E2F family.  Nat Rev Mol Cell Biol2002; 3:11-20.
  48. Montagnoli A, Fiore F, Eytan E, et al: Ubiquitination of p27 is regulated by Cdk-dependent phosphorylation and trimeric complex formation.  Genes Dev1999; 13:1181-1189.
  49. Sheaff RJ, Groudine M, Gordon M, et al: Cyclin E-CDK2 is a regulator of p27Kip1.  Genes Dev1997; 11:1464-1478.
  50. Carrano AC, Eytan E, Hershko A, Pagano M: SKP2 is required for ubiquitin-mediated degradation of the CDK inhibitor p27.  Nat Cell Biol1999; 1:193-199.
  51. Nakayama KI, Hatakeyama S, Nakayama K: Regulation of the cell cycle at the G1-S transition by proteolysis of cyclin E and p27Kip1.  Biochem Biophys Res Commun2001; 282:853-860.
  52. Nakayama KI, Nakayama K: Regulation of the cell cycle by SCF-type ubiquitin ligases.  Semin Cell Dev Biol2005; 16:323-333.
  53. Cardozo T, Pagano M: The SCF ubiquitin ligase: Insights into a molecular machine.  Nat Rev Mol cell Biol2004; 5:739-751.
  54. Nakayama KI, Nakayama K: Ubiquitin ligases: Cell-cycle control and cancer.  Nat Rev Cancer2006; 6:369-381.
  55. Sutterluty H, Chatelain E, Marti A, et al: p45SKP2 promotes p27Kip1 degradation and induces S phase in quiescent cells.  Nat Cell Biol1999; 1:207-214.
  56. Malek NP, Sundberg H, McGrew S, et al: A mouse knock-in model exposes sequential proteolytic pathways that regulate p27Kip1 in G1 and S phase.  Nature2001; 413:323-327.
  57. Koepp DM, Schaefer LK, Ye X, et al: Phosphorylation-dependent ubiquitination of cyclin E by the SCFFbw7 ubiquitin ligase.  Science2001; 294:173-177.
  58. Clurman BE, Sheaff RJ, Thress K, et al: Turnover of cyclin E by the ubiquitin-proteasome pathway is regulated by cdk2 binding and cyclin phosphorylation.  Genes Dev1996; 10:1979-1990.
  59. Dynlacht BD, Flores O, Lees JA, Harlow E: Differential regulation of E2F transactivation by cyclin/cdk2 complexes.  Genes Dev1994; 8:1772-1786.
  60. Kitagawa M, Higashi H, Suzuki-Takahashi I, et al: Phosphorylation of E2F-1 by cyclin α-cdk2.  Oncogene1995; 10:229-236.
  61. Krek W, Xu G, Livingston DM: Cyclin α-kinase regulation of E2F-1 DNA binding function underlies suppression of an S phase checkpoint.  Cell1995; 3:1149-1158.
  62. Pagano M, Pepperkok R, Verde F, et al: Cyclin A is required at two points in the human cell cycle.  Embo J1992; 11:961-971.
  63. Draetta G, Luca F, Westendorf J, et al: Cdc2 protein kinase is complexed with both cyclin A and B: evidence for proteolytic inactivation of MPF.  Cell1989; 56:829-838.
  64. Girard F, Strausfeld U, Fernandez A, Lamb NJ: Cyclin A is required for the onset of DNA replication in mammalian fibroblasts.  Cell1991; 67:1169-1179.
  65. Cardoso MC, Leonhardt H, Nadal-Ginard B: Reversal of terminal differentiation and control of DNA replication: cyclin A and Cdk2 specifically localize at subnuclear sites of DNA replication.  Cell1993; 74:979-992.
  66. Peters JM: The anaphase-promoting complex: proteolysis in mitosis and beyond.  Mol Cell2002; 9:931-943.
  67. Brandeis M, Rosewall I, Carrington M, et al: Cyclin B2-null mice develop normally and are fertile whereas cyclin B1-null mice die in utero.  Proc Natl Acad Sci USA1998; 95:4344-4349.
  68. Heald R, McLoughlin M, McKeon F: Human wee1 maintains mitotic timing by protecting the nucleus from cytoplasmically activated Cdc2 kinase.  Cell1993; 74:463-474.
  69. Parker LL, Piwnica-Worms H: Inactivation of the p34cdc2-cyclin B complex by the human WEE1 tyrosine kinase.  Science1992; 257:1955-1957.
  70. Lundgren K, Walworth N, Booher R, et al: mik1 and wee1 cooperate in the inhibitory tyrosine phosphorylation of cdc2.  Cell1991; 64:1111-1122.
  71. Jin P, Hardy S, Morgan DO: Nuclear localization of cyclin B1 controls mitotic entry after DNA damage.  J Cell Biol1998; 141:875-885.
  72. Nigg EA, Blangy A, Lane HA: Dynamic changes in nuclear architecture during mitosis: on the role of protein phosphorylation in spindle assembly and chromosome segregation.  Exp Cell Res1996; 229:174-180.
  73. Porter LA, Donoghue DJ: Cyclin B1 and CDK1: Nuclear localization and upstream regulators.  Prog Cell Cycle Res2003; 5:335-347.
  74. Pines J: Mitosis: A matter of getting rid of the right protein at the right time.  Trends Cell Biol2006; 16:55-63.
  75. Morgan DO: Regulation of the APC and the exit from mitosis.  Nat Cell Biol1999; 1:E47-E53.
  76. Sullivan M, Morgan DO: Finishing mitosis, one step at a time.  Nat Rev Mol Cell Biol2007; 8:894-903.
  77. Roussel MF: The INK4 family of cell cycle inhibitors in cancer.  Oncogene1999; 18:5311-5317.
  78. Ruas M, Peters G: The p16INK4a/CDKN2A tumor suppressor and its relatives.  Biochim Biophys Acta1998; 1378:F115-F177.
  79. Ortega S, Malumbres M, Barbacid M: Cyclin D-dependent kinases, INK4 inhibitors and cancer.  Biochim Biophys Acta2002; 1602:73-87.
  80. Bartkova J, Thullberg M, Rajpert-De Meyts E, et al: Cell cycle regulators in testicular cancer: loss of p18INK4C marks progression from carcinoma in situ to invasive germ cell tumours.  Int J Cancer2000; 85:370-375.
  81. Buchwald PC, Akerstrom G, Westin G: Reduced p18INK4c, p21CIP1/WAF1 and p27KIP1 mRNA levels in tumours of primary and secondary hyperparathyroidism.  Clin Endocrinol (Oxf)2004; 60:389-393.
  82. Sanchez-Aguilera A, Delgado J, Camacho FI, et al: Silencing of the p18INK4c gene by promoter hypermethylation in Reed-Sternberg cells in Hodgkin lymphomas.  Blood2004; 103:2351-2357.
  83. Morishita A, Masaki T, Yoshiji H, et al: Reduced expression of cell cycle regulator p18(INK4C) in human hepatocellular carcinoma.  Hepatology2004; 40:677-686.
  84. Uziel T, Zindy F, Sherr CJ, Roussel MF: The CDK inhibitor p18Ink4c is a tumor suppressor in medulloblastoma.  Cell Cycle2006; 5:363-365.
  85. LaBaer J, Garrett MD, Stevenson LF, et al: New functional activities for the p21 family of CDK inhibitors.  Genes Dev1997; 11:847-862.
  86. Hannon GJ, Beach D: p15INK4B is a potential effector of TGF-βeta-induced cell cycle arrest.  Nature1994; 371:257-261.
  87. Zhou BP, Liao Y, Xia W, et al: Cytoplasmic localization of p21Cip1/WAF1 by Akt-induced phosphorylation in HER-2/neu-overexpressing cells.  Nat Cell Biol2001; 3:245-252.
  88. Liang J, Zubovitz J, Petrocelli T, et al: PKB/Akt phosphorylates p27 impairs nuclear import of p27 and opposes p27-mediated G1 arrest.  Nat Med2002; 8:1153-1160.
  89. Shin I, Yakes FM, Rojo F, et al: PKB/Akt mediates cell-cycle progression by phosphorylation of p27(Kip1) at threonine 157 and modulation of its cellular localization.  Nat Med2002; 8:1145-1152.
  90. Viglietto G, Motti ML, Bruni P, et al: Cytoplasmic relocalization and inhibition of the cyclin-dependent kinase inhibitor p27(Kip1) by PKB/Akt-mediated phosphorylation in breast cancer.  Nat Med2002; 8:1136-1144.
  91. Weinberg RA: The retinoblastoma gene and gene product.  Cancer Surv1992; 12:43-57.
  92. Helt AM, Galloway DA: Mechanisms by which DNA tumor virus oncoproteins target the Rb family of pocket proteins.  Carcinogenesis2003; 24:159-169.
  93. Morris EJ, Dyson NJ: Retinoblastoma protein partners.  Adv Cancer Res2001; 82:1-54.
  94. Yamasaki L, Bronson R, Williams BO, et al: Loss of E2F-1 reduces tumorigenesis and extends the lifespan of Rb1+/- mice.  Nat Genet1998; 18:360-364.
  95. Ziebold U, Lee EY, Bronson RT, Lees JA: E2F3 loss has opposing effects on different pRB-deficient tumors, resulting in suppression of pituitary tumors but metastasis of medullary thyroid carcinomas.  Mol Cell Biol2003; 23:6542-6552.
  96. Lee EY, Cam H, Ziebold U, et al: E2F4 loss suppresses tumorigenesis in Rb mutant mice.  Cancer Cell2002; 2:463-472.
  97. Ren B, et al: E2F integrates cell cycle progression with DNA repair, replication, and G(2)/M checkpoints.  Genes Dev2002; 16:245-256.
  98. Cam H, Cam H, Takahasi Y, et al: A common set of gene regulatory networks links metabolism and growth inhibition.  Mol Cell2004; 16:399-411.
  99. DeGregori J, Johnson DG: Distinct and overlapping roles for E2F family members in transcription, proliferation and apoptosis.  Curr Mol Med2006; 6:739-748.
  100. Helin K, Harlow E, Fattaey A: Inhibition of E2F-1 transactivation by direct binding of the retinoblastoma protein.  Mol Cell Biol1993; 13:6501-6508.
  101. Brehm A, Miska EA, McCance DJ, et al: Retinoblastoma protein recruits histone deacetylase to repress transcription.  Nature1998; 391:597-601.
  102. Magnaghi-Jaulin L, Groisman R, Naguibneva I, et al: Retinoblastoma protein represses transcription by recruiting a histone deacetylase.  Nature1998; 391:601-605.
  103. Brehm A, Kouzarides T: Retinoblastoma protein meets chromatin.  Trends Biochem Sci1999; 24:142-145.
  104. Sherr CJ: Cancer cell cycles.  Science1996; 274:1672-1677.
  105. Rayman JB, Takahasi Y, Indjeian VB, et al: E2F mediates cell cycle-dependent transcriptional repression in vivo by recruitment of an HDAC1/mSin3B corepressor complex.  Genes Dev2002; 16:933-947.
  106. Takahashi Y, Rayman JB, Dynlacht BD: Analysis of promoter binding by the E2F and pRB families in vivo: distinct E2F proteins mediate activation and repression.  Genes Dev2000; 14:804-816.
  107. Stevaux O, Dyson NJ: A revised picture of the E2F transcriptional network and RB function.  Curr Opin Cell Biol2002; 14:684-691.
  108. Verona R, Moberg K, Estes S, et al: E2F activity is regulated by cell cycle-dependent changes in subcellular localization.  Mol Cell Biol1997; 17:7268-7282.
  109. Gaubatz S, Lees JA, Lindeman GJ, Livingston DM: E2F4 is exported from the nucleus in a CRM1-dependent manner.  Mol Cell Biol2001; 21:1384-1392.
  110. Bartek J, Bartkova J, Lukas J: The retinoblastoma protein pathway and the restriction point.  Curr Opin Cell Biol1996; 8:805-814.
  111. Planas-Silva MD, Weinberg RA: The restriction point and control of cell proliferation.  Curr Opin Cell Biol1997; 9:768-772.
  112. Johnson DG, Schwarz JK, Cress WD, Nevins JR: Expression of transcription factor E2F1 induces quiescent cells to enter S phase.  Nature1993; 365:349-352.
  113. Qin XQ, Livingston DM, Kaelin Jr WG, Adams PD: Deregulated transcription factor E2F-1 expression leads to S-phase entry and p53-mediated apoptosis.  Proc Natl Acad Sci USA1994; 91:10918-10922.
  114. Shan B, Lee WH: Deregulated expression of E2F-1 induces S-phase entry and leads to apoptosis.  Mol Cell Biol1994; 14:8166-8173.
  115. Kowalik TF, DeGregori J, Schwarz JK, Nevins JR: E2F1 overexpression in quiescent fibroblasts leads to induction of cellular DNA synthesis and apoptosis.  J Virol1995; 69:2491-2500.
  116. Lukas J, Petersen BO, Holm K, et al: Deregulated expression of E2F family members induces S-phase entry and overcomes p16INK4A-mediated growth suppression.  Mol Cell Biol1996; 16:1047-1057.
  117. Diffley JF: Once and only once upon a time: Specifying and regulating origins of DNA replication in eukaryotic cells.  Genes Dev1996; 10:2819-2830.
  118. Bell SP, Dutta A: DNA replication in eukaryotic cells.  Annu Rev Biochem2002; 71:333-374.
  119. Takeda DY, Dutta A: DNA replication and progression through S phase.  Oncogene2005; 24:2827-2843.
  120. Arias EE, Walter JC: Strength in numbers: Preventing rereplication via multiple mechanisms in eukaryotic cells.  Genes Dev2007; 21:497-518.
  121. Bell SP, Stillman B: ATP-dependent recognition of eukaryotic origins of DNA replication by a multiprotein complex.  Nature1992; 357:128-134.
  122. Bell SP, Mitchell J, Leber J, et al: The multido-main structure of Orc1p reveals similarity to regulators of DNA replication and transcriptional silencing.  Cell1995; 83:563-568.
  123. Nougarède R, Della Setta F, Zarzov P, Schwob E: Hierarchy of S-phase-promoting factors: yeast Dbf4-Cdc7 kinase requires prior S-phase cyclin-dependent kinase activation.  Mol Cell Biol2000; 20:3795-3806.
  124. 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 Biol2000; 20:3086-3096.
  125. Walter J, Newport J: Initiation of eukaryotic DNA replication: origin unwinding and sequential chromatin association of Cdc45, RPA, and DNA polymerase alpha.  Mol Cell2000; 5:617-627.
  126. Arias EE, Walter JC: Replication-dependent destruction of Cdt1 limits DNA replication to a single round per cell cycle in Xenopus egg extracts.  Genes Dev2005; 19:114-126.
  127. Higa LA, Banks D, Wu M, et al: L2DTL/CDT2 Interacts with the CUL4/DDB1 complex and PCNA and regulates CDT1 proteolysis in response to DNA damage.  Cell Cycle2006; 5:1675-1680.
  128. Jin J, Arias EE, Chen J, et al: A family of diverse Cul4-Ddb1-interacting proteins includes Cdt2, which is required for S phase destruction of the replication factor Cdt1.  Mol Cell2006; 23:709-721.
  129. Sansam CL, Shepard JL, Lai K, et al: DTL/CDT2 is essential for both CDT1 regulation and the early G2/M checkpoint.  Genes Dev2006; 20:3117-3129.
  130. Arias E, Walter J: PCNA functions as a molecular platform to trigger Cdt1 destruction and prevent rereplication.  Nat Cell Biol2005; 8:90.
  131. Tachibana KE, Gonzalez MA, Guarguaglini G, et al: Depletion of licensing inhibitor geminin causes centrosome overduplication and mitotic defects.  EMBO Rep2005; 6:1052-1057.
  132. Saxena S, Dutta A: Geminin and p53: Deterrents to rereplication in human cancer cells.  Cell Cycle2003; 2:283-286.
  133. Haering CH, Nasmyth K: Building and breaking bridges between sister chromatids.  Bioessays2003; 25:1178-1191.
  134. Nasmyth K, Haering CH: The structure and function of SMC and kleisin complexes.  Annu Rev Biochem2005; 74:595-648.
  135. Gruber S, Haering CH, Nasmyth K: Chromosomal cohesin forms a ring.  Cell2003; 112:765-777.
  136. Salic A, Waters JC, Mitchison TJ: Vertebrate shugoshin links sister centromere cohesion and kinetochore microtubule stability in mitosis.  Cell2004; 118:567-578.
  137. Peter M, Nakagawa J, Doree M, et al: In vitro disassembly of the nuclear lamina and M phase-specific phosphorylation of lamins by cdc2 kinase.  Cell1990; 61:591-602.
  138. Margalit A, Vlcek S, Gruenbaum Y, Foisner R: Breaking and making of the nuclear envelope.  J Cell Biochem2005; 95:454-465.
  139. Fukagawa T: Assembly of kinetochores in vertebrate cells.  Exp Cell Res2004; 296:21-27.
  140. O'Connell CB, Khodjakov AL: Cooperative mechanisms of mitotic spindle formation.  J Cell Sci2007; 120(pt 10):1717-1722.
  141. Hoyt MA: Cell biology: extinguishing a cell cycle checkpoint.  Science2006; 313:624-625.
  142. de Gramont A, Cohen-Fix O: The many phases of anaphase.  Trends Biochem Sci2005; 30:559-568.
  143. Fry AM, Yamano H: APC/C-mediated degradation in early mitosis: how to avoid spindle assembly checkpoint inhibition.  Cell Cycle2006; 5:1487-1491.
  144. Yu H: Cdc20: a WD40 activator for a cell cycle degradation machine.  Mol Cell2007; 27:3-16.
  145. Kastan MB, Bartek J: Cell-cycle checkpoints and cancer.  Nature2004; 432:316-323.
  146. Lukas J, Lukas C, Bartek J: Mammalian cell cycle checkpoints: signalling pathways and their organization in space and time.  DNA Repair (Amst)2004; 3:997-1007.
  147. Bartek J, Lukas J: DNA damage checkpoints: from initiation to recovery or adaptation.  Curr Opin Cell Biol2007; 19:238-245.
  148. Motoyama N, Naka K: DNA damage tumor suppressor genes and genomic instability.  Curr Opin Genet Dev2004; 14:11-16.
  149. Gottifredi V, Prives C: The S phase checkpoint: when the crowd meets at the fork.  Semin Cell Dev Biol2005; 16:355-368.
  150. Taylor AM, Harnden DG, Arlett CF, et al: Ataxia telangiectasia: a human mutation with abnormal radiation sensitivity.  Nature1975; 258:427-429.
  151. Cuadrado M, Martinez-Pastor B, Murga M, et al: ATM regulates ATR chromatin loading in response to DNA double-strand breaks.  J Exp Med2006; 203:297-303.
  152. Matsuoka S, Huang M, Elledge SJ: Linkage of ATM to cell cycle regulation by the Chk2 protein kinase.  Science1998; 282:1893-1897.
  153. Matsuoka S, Rotman G, Ogawa A, et al: Ataxia telangiectasia-mutated phosphorylates Chk2 in vivo and in vitro.  Proc Natl Acad Sci USA2000; 97:10389-10394.
  154. Busino L, Donzelli M, Chiesa M, et al: Degradation of Cdc25A by beta-TrCP during S phase and in response to DNA damage.  Nature2003; 426:87-91.
  155. Hermeking H, Benzinger A: 14–3-3 proteins in cell cycle regulation.  Semin Cancer Biol2006; 16:83-192.
  156. Hirao A, Kong YY, Matsuoka S, et al: DNA damage-induced activation of p53 by the checkpoint kinase Chk2.  Science2000; 287:1824-1827.
  157. Jones SN, Roe AE, Donehower LA, Bradley A: Rescue of embryonic lethality in Mdm2-deficient mice by absence of p53.  Nature1995; 378:206-208.
  158. Montes de Oca Luna R, Wagner DS, Lozano G: Rescue of early embryonic lethality in mdm2-deficient mice by deletion of p53.  Nature1995; 378:203-206.
  159. Waldman T, Kinzler KW, Vogelstein B: p21 is necessary for the p53-mediated G1 arrest in human cancer cells.  Cancer Res1995; 55:5187-5190.
  160. el-Deiry WS, Tokino T, Velculescu VE, et al: WAF1, a potential mediator of p53 tumor suppression.  Cell1993; 75:817-825.
  161. Vousden KH, Lu X: Live or let die: the cell's response to p53.  Nat Rev Cancer2002; 2:594-604.
  162. Serrano M, Lin AW, McCurrach ME, et al: Oncogenic ras provokes premature cell senescence associated with accumulation of p53 and p16INK4a.  Cell1997; 88:593-602.
  163. de Stanchina E, McCurrach ME, Zindy F, et al: E1A signaling to p53 involves the p19(ARF) tumor suppressor.  Genes Dev1998; 2:2434-2442.
  164. Dimri GP, Itahana K, Acosta M, Campisi J: Regulation of a senescence checkpoint response by the E2F1 transcription factor and p14(ARF) tumor suppressor.  Mol Cell Biol2000; 20:273-285.
  165. Zindy F, Eischen CM, Randle DH, et al: Myc signaling via the ARF tumor suppressor regulates p53-dependent apoptosis and immortalization.  Genes Dev1998; 12:2424-2433.
  166. Bartkova J, Razaei N, Liontos M, et al: Oncogene-induced senescence is part of the tumorigenesis barrier imposed by DNA damage checkpoints.  Nature2006; 444:633-637.
  167. Di Micco R, Fumagalli M, Cicalese A, et al: Oncogene-induced senescence is a DNA damage response triggered by DNA hyper-replication.  Nature2006; 444:638-642.
  168. Zindy F, Williams RT, Baudino TA, et al: Arf tumor suppressor promoter monitors latent oncogenic signals in vivo.  Proc Natl Acad Sci USA2003; 100:15930-15935.
  169. Sherr CJ: The INK4a/ARF network in tumour suppression.  Nat Rev Mol Cell Biol2001; 2:731-737.
  170. Kamijo T, Weber JD, Zambetti G, et al: Functional and physical interactions of the ARF tumor suppressor with p53 and Mdm2.  Proc Natl Acad Sci USA1998; 95:8292-8297.
  171. Honda R, Yasuda H: Association of p19(ARF) with Mdm2 inhibits ubiquitin ligase activity of Mdm2 for tumor suppressor p53.  EMBO J1999; 18:22-27.
  172. Llanos S, Clark A, Rowe J, Peters G: Stabilization of p53 by p14ARF without relocation of MDM2 to the nucleolus.  Nat Cell Biol2001; 3:445-452.
  173. Pomerantz J, Schreiber-Agus N, Liégeois NJ, et al: The Ink4a tumor suppressor gene product, p19Arf, interacts with MDM2 and neutralizes MDM2′s inhibition of p53.  Cell1998; 92:713-723.
  174. Weber JD, Taylor LJ, Roussel MF, et al: Nucleolar Arf sequesters Mdm2 and activates p53.  Nat Cell Biol1999; 1:20-26.
  175. Higa LA, Mihaylov IS, Banks DP, et al: Radiation-mediated proteolysis of CDT1 by CUL4-ROC1 and CSN complexes constitutes a new checkpoint.  Nat Cell Biol2003; 5:1008-1015.
  176. Sanchez Y, Wong C, Toma RS, et al: Conservation of the Chk1 checkpoint pathway in mammals: linkage of DNA damage to Cdk regulation through Cdc25.  Science1997; 277:1497-1501.
  177. Furnari B, Blasina A, Boddy MN, et al: Cdc25 inhibited in vivo and in vitro by checkpoint kinases Cds1 and Chk1.  Mol Biol Cell1999; 10:833-845.
  178. Liu Q, Guntuku S, Cui XS, et al: Chk1 is an essential kinase that is regulated by Atr and required for the G2M DNA damage checkpoint.  Genes Dev2000; 14:1448-1459.
  179. Jin J, Shirogane T, Xu L, et al: SCFbeta-TRCP links Chk1 signaling to degradation of the Cdc25A protein phosphatase.  Genes Dev2003; 17:3062-3074.
  180. O'Connell MJ, Walworth NC, Carr AM: The G2-phase DNA-damage checkpoint.  Trends Cell Biol2000; 10:296-303.
  181. Musacchio A, Salmon ED: The spindle-assembly checkpoint in space and time.  Nat Rev Mol Cell Biol2007; 8:379-393.
  182. Cimini D, Degrassi F: Aneuploidy: a matter of bad connections.  Trends Cell Biol2005; 15:442-451.
  183. Kops GJ, Weaver BA, Cleveland DW: On the road to cancer: aneuploidy and the mitotic checkpoint.  Nat Rev Cancer2005; 5:773-785.
  184. Baker DJ, Chen J, van Deursen JM: The mitotic checkpoint in cancer and aging: what have mice taught us?.  Curr Opin Cell Biol2005; 17:583-589.
  185. Cahill DP, Lengaur C, Yu J, et al: Mutations of mitotic checkpoint genes in human cancers.  Nature1998; 392:300-303.
  186. Michel LS, Liberal V, Chatterjee A, et al: MAD2 haploinsufficiency causes premature anaphase and chromosome instability in mammalian cells.  Nature2001; 409:355-359.
  187. Sellers WR, Kaelin Jr WG: Role of the retinoblastoma protein in the pathogenesis of human cancer.  J Clin Oncol1997; 15:3301-3312.
  188. Kaye FJ: RB and cyclin dependent kinase pathways: defining a distinction between RB and p16 loss in lung cancer.  Oncogene2002; 21:6908-6914.
  189. Varley JM, Armour J, Swallow JE, et al: The retinoblastoma gene is frequently altered leading to loss of expression in primary breast tumours.  Oncogene1989; 4:725-729.
  190. Zheng L, Lee WH: The retinoblastoma gene: aprototypic and multifunctional tumor suppressor.  Exp Cell Res2001; 264:2-18.
  191. Nobori T, Miura K, Wu DJ, et al: Deletions of the cyclin-dependent kinase-4 inhibitor gene in multiple human cancers.  Nature1994; 368:753-756.
  192. Ravaioli A, Bagli L, Zucchini A, Monti F: Prognosis and prediction of response in breast cancer: the current role of the main biological markers.  Cell Prolif1998; 31:113-126.
  193. Weinstat-Saslow D, Merino MJ, Manrow RE, et al: Overexpression of cyclin D mRNA distinguishes invasive and in situ breast carcinomas from non-malignant lesions.  Nat Med1995; 1:1257-1260.
  194. Wang TC, Cardiff RD, Zukerberg L, et al: Mammary hyperplasia and carcinoma in MMTV-cyclin D1 transgenic mice.  Nature1994; 369:669-671.
  195. Bortner DM, Rosenberg MP: Induction of mammary gland hyperplasia and carcinomas in transgenic mice expressing human cyclin E.  Mol Cell Biol1997; 17:453-459.
  196. Elsayed YA, Sausville EA: Selected novel anticancer treatments targeting cell signaling proteins.  Oncologist2001; 6:517-537.
  197. Porter PL, Malone KE, Heagerty PJ, et al: Expression of cell-cycle regulators p27Kip1 and cyclin E, alone and in combination, correlate with survival in young breast cancer patients.  Nat Med1997; 3:222-225.
  198. Catzavelos C, Bhattacharya N, Ung YC, et al: Decreased levels of the cell-cycle inhibitor p27Kip1 protein: prognostic implications in primary breast cancer.  Nat Med1997; 3:227-230.
  199. Esteller M, Herman JG: Cancer as an epigenetic disease: DNA methylation and chromatin alterations in human tumours.  J Pathol2002; 196:1-7.
  200. Cannon-Albright LA, Goldgar DE, Meyer LJ, et al: Assignment of a locus for familial melanoma, MLM, to chromosome 9p13-p22.  Science1992; 258:1148-1152.
  201. Gasparotto D, Maestro R, Piccinin S, et al: Overexpression of CDC25A and CDC25B in head and neck cancers.  Cancer Res1997; 57:2366-2368.
  202. Wu W, Fan YH, Kemp BL, et al: Overexpression of cdc25A and cdc25B is frequent in primary non-small cell lung cancer but is not associated with overexpression of c-myc.  Cancer Res1998; 58:4082-4085.
  203. Nigro JM, Baker SJ, Preisinger AC, et al: Mutations in the p53 gene occur in diverse human tumour types.  Nature1989; 342:705-708.
  204. Ozbun MA, Butel JS: Tumor suppressor p53 mutations and breast cancer: a critical analysis.  Adv Cancer Res1995; 66:71-141.
  205. Momand J, Jung D, Wilczynski S, Niland J: The MDM2 gene amplification database.  Nucleic Acids Res1998; 26:3453-3459.
  206. Scheffner M, Werness BA, Huibregtse JM, et al: The E6 oncoprotein encoded by human papillomavirus types 16 and 18 promotes the degradation of p53.  Cell1990; 63:1129-1136.
  207. Khanna KK: Cancer risk and the ATM gene: a continuing debate.  J Natl Cancer Inst2000; 92:795-802.
  208. Barlow C, et al: Atm-deficient mice: a paradigm of ataxia telangiectasia.  Cell1996; 86:159-171.
  209. Xu Y, Ashley T, Brainerd EE, et al: Targeted disruption of ATM leads to growth retardation, chromosomal fragmentation during meiosis, immune defects, and thymic lymphoma.  Genes Dev1996; 10:2411-2422.
  210. Petrini JH: The Mre11 complex and ATM: collaborating to navigate S phase.  Curr Opin Cell Biol2000; 12:293-296.
  211. Matsuoka S, Nakagawa T, Masuda A, et al: Reduced expression and impaired kinase activity of a Chk2 mutant identified in human lung cancer.  Cancer Res2001; 61:5362-5365.
  212. Bertoni F, Codegoni AM, Furlan D, et al: CHK1 frameshift mutations in genetically unstable colorectal and endometrial cancers.  Genes Chromosomes Cancer1999; 26:176-180.
  213. Bell DW, Varley JM, Szydlo TE, et al: Heterozygous germ line hCHK2 mutations in Li-Fraumeni syndrome.  Science1999; 286:2528-2531.
  214. Wang W: Emergence of a DNA-damage response network consisting of Fanconi anaemia and BRCA proteins.  Nat Rev Genet2007; 8:735-748.
  215. Lee H, Trainer AH, Friedman LS, et al: Mitotic checkpoint inactivation fosters transformation in cells lacking the breast cancer susceptibility gene, Brca2.  Mol Cell1999; 4:1-10.
  216. Yu Q, Geng Y, Sicinski P: Specific protection against breast cancers by cyclin D1 ablation.  Nature2001; 411:1017-1021.
  217. Satyanarayana A, Hilton MB, Kaldis P: p21 inhibits Cdk1 in the absence of Cdk2 to maintain the G1/S phase DNA damage checkpoint.  Mol Biol Cell2007;[Epub ahead of print].
  218. Berthet C, Aleem E, Coppola V, et al: Cdk2 knockout mice are viable.  Curr Biol2003; 13:1775-1785.
  219. Ortega S, Prieto I, Odajima J, et al: Cyclin-dependent kinase 2 is essential for meiosis but not for mitotic cell division in mice.  Nat Genet2003; 5:25-31.
  220. Collins I, Garrett MD: Targeting the cell division cycle in cancer: CDK and cell cycle checkpoint kinase inhibitors.  Curr Opin Pharmacol2005; 5:366-373.
  221. Shapiro GI: Cyclin-dependent kinase pathways as targets for cancer treatment.  J Clin Oncol2006; 24:1770-1783.
  222. Schwartz GK, Ilson D, Saltz L, et al: Phase II study of the cyclin-dependent kinase inhibitor flavopiridol administered to patients with advanced gastric carcinoma.  J Clin Oncol2001; 19:1985-1992.
  223. Senderowicz AM: Flavopiridol: the first cyclin-dependent kinase inhibitor in human clinical trials.  Invest New Drugs1999; 17:313-320.
  224. Stadler WM, Vogelzang NJ, Amato R, et al: Flavopiridol, a novel cyclin-dependent kinase inhibitor, in metastatic renal cancer: a University of Chicago Phase II Consortium study.  J Clin Oncol2000; 18:371-375.
  225. Lu X, Burgan WE, Cerra MA, et al: Transcriptional signature of flavopiridol-induced tumor cell death.  Mol Cancer Ther2004; 3:861-872.
  226. Meinhart A, Kamenski T, Hoeppner S, et al: A structural perspective of CTD function.  Genes Dev2005; 19:1401-1415.
  227. Fry DW, Harvey PJ, Keller PR, et al: Specific inhibition of cyclin-dependent kinase 4/6 by PD 0332991 and associated antitumor activity in human tumor xenografts.  Mol Cancer Ther2004; 3:1427-1438.
  228. Tutt AN, Lord CJ, McCabe N, et al: Exploiting the DNA repair defect in BRCA mutant cells in the design of new therapeutic strategies for cancer.  Cold Spring Harb Symp Quant Biol2005; 70:139-148.
  229. Farmer H, McCabe N, Lord CJ, et al: Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy.  Nature2005; 434:917-921.
  230. Bryant HE, Schultz N, Thomas HD, et al: Specific killing of BRCA2-deficient tumours with inhibitors of poly(ADP-ribose) polymerase.  Nature2005; 434:913-917.
  231. Hartwell LH, Kastan MB: Cell cycle control and cancer.  Science1994; 266:1821-1828.