Cell Cycle Deregulation in Cancer (Contemporary Cancer Research)

Contemporary Cancer Research Jac A. Nickoloff, S ERIES E DITOR For other titles published in this series, go to http:/...

Author: Greg H. Enders

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Contemporary Cancer Research Jac A. Nickoloff, S ERIES E DITOR

For other titles published in this series, go to http://www.springer.com/series/7675

Greg H. Enders Editor

Cell Cycle Deregulation in Cancer

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Editor Greg H. Enders Fox Chase Cancer Center Department of Medicine 333 Cottman Ave. Philadelphia PA 19111-2497 USA [email protected]

ISBN 978-1-4419-1769-0 e-ISBN 978-1-4419-1770-6 DOI 10.1007/978-1-4419-1770-6 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010921202 © Springer Science+Business Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Contents

Part I

Starting the Cell Division Cycle

1 Escape from Cellular Quiescence . . . . . . . . . . . . . . . . . . Elena Sotillo and Xavier Graña

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2 Interplay Between Cyclin-Dependent Kinases and E2F-Dependent Transcription . . . . . . . . . . . . . . . . . . Jun-Yuan Ji and Nicholas J. Dyson

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3 Regulation of Pre-RC Assembly: A Complex Symphony Orchestrated by CDKs . . . . . . . . . . . . . . . . . . . . . . . . A. Kathleen McClendon, Jeffry L. Dean, and Erik S. Knudsen

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Part II

Proliferation Under Duress

4 Mitotic Checkpoint and Chromosome Instability in Cancer . . . . Haomin Huang and Timothy J. Yen

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5 Mitotic Catastrophe . . . . . . . . . . . . . . . . . . . . . . . . . . Jeremy P.H. Chow and Randy Y.C. Poon

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6 p53, ARF, and the Control of Autophagy . . . . . . . . . . . . . . Robert D. Hontz and Maureen E. Murphy

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Part III Long-Term Proliferation 7 Regulation of Self-Renewing Divisions in Normal and Leukaemia Stem Cells . . . . . . . . . . . . . . . . . . . . . . Andrea Viale and Pier Giuseppe Pelicci 8 Maintenance of Telomeres in Cancer . . . . . . . . . . . . . . . . Eros Lazzerini Denchi 9 The Senescence Secretome and Its Impact on Tumor Suppression and Cancer . . . . . . . . . . . . . . . . . . . . . . . Alyssa Kennedy and Peter D. Adams

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Part IV

Contents

Applications in Preventing and Treating Cancer

10 Cell Cycle Deregulation in Pre-neoplasia: Case Study of Barrett’s Oesophagus . . . . . . . . . . . . . . . . . . . . . . . Pierre Lao-Sirieix and Rebecca C. Fitzgerald

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11 Targeting Cyclin-Dependent Kinases for Cancer Therapy . . . . . Neil Johnson and Geoffrey I. Shapiro

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors

Peter D. Adams Cancer Research UK Beatson Labs, University of Glasgow, Glasgow G61 1BD, UK, [email protected] Jeremy P.H. Chow Department of Biochemistry, Hong Kong University of Science and Technology, Clear Water Bay, Hong Kong, China Jeffry L. Dean Department of Cancer Biology, Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, PA 19107, USA Eros Lazzerini Denchi Laboratory of Chromosome Biology and Genomic Stability, Department of Genetics, The Scripps Research Institute, La Jolla, CA 92037, USA, [email protected] Nicholas J. Dyson Department of Pathology, Harvard Medical School, Massachusetts General Hospital Cancer Center, Charlestown, MA 02129, USA, [email protected] Rebecca C. Fitzgerald Cancer Cell Unit, Hutchison-MRC Research Centre, Cambridge, CB2 0XZ, UK, [email protected] Xavier Graña Fels Institute for Cancer Research and Molecular Biology, Temple University School of Medicine, Philadelphia, PA 19140, USA; Department of Biochemistry, Temple University School of Medicine, Philadelphia, PA 19140, USA, [email protected] Robert D. Hontz Program in Molecular and Translational Medicine, Fox Chase Cancer Center, Philadelphia, PA 19111, USA Haomin Huang Fox Chase Cancer Center, Philadelphia, PA 19111, USA, [email protected] Jun-Yuan Ji Department of Pathology, Harvard Medical School, Massachusetts General Hospital Cancer Center, Charlestown, MA 02129, USA; Department of Molecular and Cellular Medicine, Texas A&M Health Science Center, College Station, TX 77843, USA, [email protected] Neil Johnson Department of Medical Oncology, Dana-Farber Cancer Institute, Boston, MA 02115, USA; Department of Medicine, Brigham and Women’s vii

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Contributors

Hospital, Boston, MA 02115, USA; Harvard Medical School, Boston, MA 02115, USA, [email protected] Alyssa Kennedy Fox Chase Cancer Center, Philadelphia, PA 19111, USA Erik S. Knudsen Department of Cancer Biology, Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, PA 19107, USA, [email protected] Pierre Lao-Sirieix Cancer Cell Unit, Hutchison-MRC Research Centre, Cambridge, CB2 0XZ, UK, [email protected] A. Kathleen McClendon Department of Cancer Biology, Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, PA 19107, USA, [email protected] Maureen E. Murphy Program in Molecular and Translational Medicine, Fox Chase Cancer Center, Philadelphia, PA 19111, USA, [email protected] Pier Giuseppe Pelicci Department of Experimental Oncology, European Institute of Oncology at IFOM-IEO-Campus, Milan 20139, Italy, [email protected] Randy Y.C. Poon Department of Biochemistry, Hong Kong University of Science and Technology, Clear Water Bay, Hong Kong, China, [email protected]; http://ihome.ust.hk/∼rycpoon Geoffrey I. Shapiro Department of Medical Oncology, Dana-Farber Cancer Institute, Boston, MA 02115, USA; Department of Medicine, Brigham and Women’s Hospital, Boston, MA 02115, USA; Harvard Medical School, Boston, MA 02115, USA, [email protected] Elena Sotillo Department of Pathology, Children’s Hospital of Philadelphia, Philadelphia, PA 19104-4399, USA Andrea Viale Department of Experimental Oncology, European Institute of Oncology at IFOM-IEO-Campus, Milan 20139, Italy, [email protected] Timothy J. Yen Fox Chase Cancer Center, Philadelphia, PA 19111, USA, [email protected]

Part I

Starting the Cell Division Cycle

Chapter 1

Escape from Cellular Quiescence Elena Sotillo and Xavier Graña

Abstract Quiescent: From Latin quies, referring to a state of being at rest, dormant, inactive, quiet, still (Merriam-Webster, 2009, Online Dictionary: http://www.merriam-webster.com/dictionary/quiescent). This term refers to a state of dormancy as opposed to a proliferative state. However, quiescent cells are in any other regard metabolically active. In many tissues with relative fast cell renewal rates the primary function of a small group of undifferentiated cells is limited to selfrenewal (stem cells). These cells remain quiescent most of the time dividing only occasionally. In other tissues, key cell types perform fundamental tissue functions while remaining quiescent. Both stem cells and cells from tissues that renew via simple duplication can remain quiescent for long periods of time while retaining the capacity to re-enter the cell cycle. This chapter will discuss the mechanisms emerging as responsible for the maintenance of quiescence as well as those pathways that mediate quiescence entry and exit. We will also review signaling pathways deregulated during infection by Simian Virus 40 (SV40) and oncogenic transformation, which result in unscheduled exit from quiescence into the cell cycle, with focus on SV40 small t antigen.

1.1 Quiescence: The Reversible State Eukaryotic cells can be in a dividing proliferative state or they can enter nondividing states. There are four possible non-dividing states: quiescence (G0), senescence, differentiation, and apoptosis. Importantly, only quiescent cells can reversibly re-enter the cell cycle upon appropriate stimuli, whereas terminally differentiated (for the most part) and senescent cells, which can also survive for long periods of time, have permanently withdrawn the cell cycle (Fig. 1.1). In multicellular organisms, commitment to a round of DNA replication and cell division X. Graña (B) Fels Institute for Cancer Research and Molecular Biology, Temple University School of Medicine, Philadelphia, PA 19140, USA e-mail: [email protected]

G.H. Enders (ed.), Cell Cycle Deregulation in Cancer, Contemporary Cancer Research, C Springer Science+Business Media, LLC 2010 DOI 10.1007/978-1-4419-1770-6_1,

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requires adequate concentration of mitogens in the environment, space, and for adherent cells, a substrate to attach to. Thus, deprivation of mitogens, lack of adhesion, or growth to high density drive normal cells into quiescence (Fig. 1.1). Recent studies have uncovered that each of these cell cycle exit-initiating signals elicits a distinct gene expression signature (Coller et al., 2006). However, to preserve the reversibility of the quiescent state, a shared “quiescent gene expression program” that includes genes that suppress differentiation and apoptosis is implemented in all instances. It is well established that the quiescent state is associated with an increase in the expression of the CDK inhibitor p27 (Sherr and Roberts, 1999). Unexpectedly, the study of the gene expression fingerprints that characterize quiescence has also revealed that quiescence is not equivalent to growth arrest induced via inhibition of CDKs. Cells ectopically expressing the p21/p27 CDK inhibitors exhibit a distinctive program of gene expression that includes a portion of the genes found

Fig. 1.1 Fate of proliferating normal cells upon cell cycle exit. Upon cell cycle exit, cells can enter three non-dividing stable states: terminal differentiation, senescence, and quiescence. Of these, only cellular quiescence is reversible. Cellular quiescence can be triggered by mitogenic starvation, growth to high density, and lack of attachment to substratum. The restriction point (R) is the point in G1 phase where cells commit to a round of DNA replication and cell division. Cells require mitogens in the post-mitotic G1 prior to the R. Mitogens activate G1 CDKs, which cooperatively inactivate pocket proteins and activate the E2F program of gene expression required for cell cycle progression (see text)

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downregulated by all quiescent signals mentioned above, but it does not induce upregulation of genes that suppress differentiation or inhibit apoptosis (Coller et al., 2006). In agreement with the observation that CKI inhibitors are upregulated during differentiation along particular lineages, overexpression of p21 in dermal fibroblasts induced growth arrest but did not prevent MyoD-induced differentiation. In contrast, fibroblasts forced into quiescence by contact inhibition or mitogenic withdrawal are resistant to differentiation signals (Coller et al., 2006). These results show that cellular quiescence is not a mere consequence of cell cycle exit but rather a unique resting state that preserves cells in environments that are not suitable for proliferation. More recently, the mechanisms that control the reversibility of cellular quiescence have started to be unveiled. Because the transcriptional repressor Hairy and Enhancer of Split1 (HES1) is induced by signals that force fibroblasts into quiescence but is not regulated when cell cycle exit is induced by overexpression of CKIs (Coller et al., 2006), Sang et al. tested whether HES1 modulates the reversibility of cellular quiescence (Sang et al., 2008). Remarkably, it was found that ectopic expression of HES1 in dermal fibroblasts prevents p21-induced irreversible senescence, although it cannot reverse this phenotype if senescence is attained prior to HES1 expression. More importantly, their work also demonstrated that MyoD-induced differentiation of proliferating fibroblasts is prevented by ectopic expression of HES1 and that inactivation of HES1 in quiescent fibroblasts is sufficient to induce spontaneous senescence or trigger myogenic differentiation in response to MyoD activation. Thus, HES1 emerges as a pivotal candidate to control the reversibility of the quiescent state.

1.2 Overcoming the Restriction Point 1.2.1 The Restriction Point In unicellularorganisms such as yeast, the availability of nutrients in the environment primarily determines their proliferation rate. In contrast, nutrients in the environment of cells in multicellular organisms are not typically limiting, and thus proliferation rates are determined by mitogens produced by other cells or by genetic developmental programs. The cell cycle can be subdivided in two functionally distinct parts based on their dependency on mitogens for cell cycle progression (Fig. 1.1). The mitogen-dependent phase spans the period of the cell cycle beginning with initiation of post-mitotic G1 to the Restriction point (R), which was first defined by Arthur Pardee (Pardee, 1974). Once cells surpass R, they are committed to a round of DNA replication and cell division, and the progression and continuity from one phase to the next depend solely on the cell’s efficiency to faithfully complete DNA replication, chromosomal segregation, cytokinesis, and other required intermediate steps. On the other hand, normal post-mitotic early G1 cells that encounter an environment with limiting mitogens, extracellular substrate attachment, or space, enter a reversible quiescence state.

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The main challenge faced by a cell exiting quiescence is to synthesize de novo all the gene products required for successful cell cycle entry and passage through R. E2F transcription factors control the expression of many genes whose products are essential, or at least important, for cell cycle progression. In quiescent cells, repressor E2Fs (E2Fs 4 and 5) form complexes with pocket proteins (typically p130 and the retinoblastoma protein, pRB) which silence E2F-dependent gene expression (reviewed in Graña et al., 1998; Mulligan and Jacks, 1998; Blais and Dynlacht, 2004; Rowland and Bernards, 2006). Mitogens activate intracellular signaling pathways that trigger activation of G1 cyclin/CDK complexes, which in turn disrupt E2F/pocket protein complexes via phosphorylation of the pocket protein. It is thought that once the E2F-gene expression program is set in motion to warrant the expression of sufficient levels of DNA replication enzymes and other cell cycle proteins and regulators, cell cycle progression becomes insensitive to both positive and negative external mitogenic stimuli (Fig. 1.1).

1.2.2 G1-Cyclins/CDK, pRB, and E2F Transcription Factors Since this book contains a chapter devoted to the interplay between CDKs and E2Fdependent transcription, the focus of this section will be restricted to the events important for quiescence exit/entry. G1-cyclins, together with their catalytic partners, the CDKs, are the key effectors of mitogenic signaling that drive cells out of quiescence in propitious environmental conditions. There are three mammalian isoforms of cyclin D (D1, D2, and D3) that exhibit tissue-specific expression. D-type cyclins bind to CDK4 or CDK6 (CDK46) and are activated in mid-G1. E-type cyclins, E1 and E2, bind to CDK2 leading to its activation later in G1. Mitogenic stimulation activates RAS, which induces cyclin D1 transcription (Albanese et al., 1995) and stabilization through RAF/MAPK and PI3K/AKT mitogenic pathways (Diehl et al., 1998; Henry et al., 2000). Cyclin D/CDK4-6 complexes promote activation of cyclin E/CDK2 complexes through sequestration of CDK inhibitors (CKIs) from the CIP/KIP family (p21, p27, and p57). The trimeric complex cyclin D/CDK4-6/CKI shuttles into the nucleus, where it phosphorylates multiple sites on p130/pRB, relieving repressor E2Fs from pocket protein inhibition to initiate expression of early E2F-dependent genes, which in turn will generate more cyclin E (Fig. 1.2). The increase in cyclin E expression and the sequestration of CKIs by cyclin D/CDK4-6 complexes ensure accumulation of CKI-free cyclin E/CDK2 complexes that can be phosphorylated on the activating T-loop of CDK2 by the CDK-activating kinase (CAK) (Kato et al., 1994; Kaldis et al., 1998; Sherr and Roberts, 2004). As cyclin E/CDK2 active complexes emerge, a positive feedback loop ensures rapid activation of CDK2 through direct phosphorylation of CKIs, triggering their degradation, and hyperphosphorylation of pocket proteins facilitating additional accumulation of cyclin E and CDK2. CDK2 completes the inactivation of pocket proteins initiated by CDK4-6, which results in forceful elimination of repressor E2F complexes at the promoters and the expression of activator

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Fig. 1.2 Mitogens stimulate cell cycle re-entry via activation of the E2F-program of gene expression. Transition into the G1 phase of the cell cycle from quiescence requires activation of E2F-dependent gene expression. Expression of E2F-dependent genes is silent in quiescent cells. Promoters of E2F-dependent genes are occupied by E2F complexes containing repressors E2Fs and p130, as well as homologs of C. elegans synthetic multivulva class B gene products (MuvB pep). Mitogenic stimulation results in activation of CDKs by inducing G1 cyclin accumulation and inactivation of CKIs through various mechanisms. G1 CDKs phosphorylate pocket proteins disrupting their interaction with repressor E2Fs coinciding with the expression of gene products. Among the upregulated proteins are activator E2Fs (E2F1-3) that are recruited to promoters coinciding with recruitment of HATs and promoter activity. Cyclin E is an E2F-regulated gene product that helps inactivate pocket proteins, but also targets other substrates for phosphorylation that are important for DNA replication and centriole duplication. Antimitogenic signaling negatively regulates CDKs through upregulation of CKIs

E2Fs (E2F1-3), which are subsequently recruited to multiple E2F-dependent promoters coinciding with expression of E2F-dependent genes. Obviously, there are other players that participate in the activation of these CDKs and E2F-dependent gene expression, so readers are directed to more comprehensive reviews (Blais and Dynlacht, 2004; Rowland and Bernards, 2006; Blais and Dynlacht, 2007). It is important to note at this time that whereas cyclin D/CDK4-6 primary substrates are pocket proteins and Smad3 (Liu and Matsuura, 2005), both involved in repression of cell cycle-dependent gene expression, cyclin E/CDK2 functions are not limited to pocket protein inactivation in G1. Cyclin E/CDK2 phosphorylates multiple factors involved in centrosomal duplication, replication origin licensing and firing, and control of histone synthesis (Moroy and Geisen, 2004) (Fig. 1.2). Overexpression of G1 cyclins is common in primary tumors and derived tumor cell lines (Malumbres and Barbacid, 2001). Considering that mitogenic signaling converges in the activation of G1 cyclin/CDK complexes, deregulation of G1 cyclins in tumor cells may reduce the threshold of mitogenic stimulation required for passage through the R or for escaping quiescence. In this regard, early studies showed

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that overexpression of either D1 or E shortens G1 phase upon mitogenic stimulation. However, quiescent primary non-transformed fibroblasts that ectopically express cyclin D1 and/or E do not exit quiescence if the environment is deprived of mitogens or if the cells are arrested by growth to high density (Ohtsubo and Roberts, 1993; Quelle et al., 1993; Resnitzky et al., 1994; Sotillo et al., 2008, 2009). In contrast, similar expression of cyclin D1 and E in certain tumor cell lines is sufficient to trigger exit from quiescence in the absence of any mitogenic stimulation (Calbó et al., 2002). Experiments performed in our laboratory have shown that in quiescent tumor-derived T98G cells forced expression of cyclin E leads to formation of active CDK2 complexes, pocket protein phosphorylation, and activation of the E2F program concomitantly with cell cycle entry. Under the same conditions expression of cyclin E in quiescent normal human fibroblasts (NHF) leads to formation of inactive complexes failing to trigger cell cycle entry. Concentrations of serum as low as 0.1% make quiescent NHF responsive to deregulated cyclin E expression, suggesting that other mitogen-dependent events, besides cyclin E accumulation, are required to fully activate CDK2 and exit G0. This is consistent with previous work showing that microinjection of active G1 cyclin/CDK complexes into the nucleus of primary human WI38 fibroblasts is sufficient to induce DNA synthesis (Connell-Crowley et al., 1998). Despite the clear important role of G1 CDKs in mediating passage through R and triggering E2F-dependent gene expression, ablation of G1 CDKs and cyclins in mice has evidenced a high level of functional redundancy and compensation among these G1 cyclin/CDK complexes in triggering inactivation of pocket proteins and other essential events during the cell cycle (Malumbres and Barbacid, 2009). Targeted disruption of D-type cyclins, E-type cyclins, CDK4-6, or CDK2 reduces inactivation of pocket proteins but not below a threshold that could prevent E2Fdependent gene expression in both proliferating and serum starved and re-stimulated MEFs (Lee and Sicinski, 2006; Berthet and Kaldis, 2007; Malumbres and Barbacid, 2009). Indeed, even fibroblasts obtained from mouse embryos that simultaneously lack expression of CDK2, CDK4, and CDK6 proliferate and exit quiescence in response to serum stimulation (Santamaria et al., 2007). Thus, CDK1 via its binding with cyclin E appears sufficient to inactivate pocket proteins and induce passage through R. Of note, serum-starved cyclin E1–/–; E2–/– double knock-out MEFs are unable to re-enter cell cycle when stimulated with mitogens. However, this is due to a defect in loading of MCM2 onto chromatin, as pocket proteins are inactivated and E2F-dependent genes expressed (Geng et al., 2003). These results indicate great plasticity and compensation among cyclins and CDKs in many cell types, with function of some of them only essential in particular cell types.

1.2.3 Is Inactivation of Pocket Proteins Beyond a Certain Threshold Sufficient for Passage Through R? Ablation of the three pocket proteins in MEFs makes these cells bypass cell cycle exit signals induced by mitogen withdrawal, contact inhibition, and loss of attachment, but cells become apoptotic (Dannenberg et al., 2000; Sage et al., 2000). Pocket

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protein mutant MEFs also fail to arrest in response to a variety of signals that cause G1 arrest. Thus, pocket proteins are important for establishment of a G1 growth arrest and exit into quiescence, as triple mutant MEFs fail to become quiescent in response to three independent signals that mediate reversible growth arrest. Ectopic expression of E2F1 drives quiescent rodent fibroblasts into S phase (Johnson et al., 1993). Activator E2Fs (E2F1-3) are likely required for cell cycle re-entry, while E2F-repressor activities are critical for contact inhibition induced cell cycle exit and other signals that induce G1 arrest (Rowland and Bernards, 2006). Therefore, there is firm evidence that pocket protein/E2F pathways play critical roles in cell cycle re-entry/exit, as loss of these pathways makes cells insensitive to extracellular control in both cycling and quiescent cells. It is important to stress that despite the high degree of compensation among pocket proteins, it has become clear that distinct complexes play specialized functions. An evolutionarily conserved complex designated DREAM has been identified in mammalian cells that contains p130, E2F4, and mammalian homologs of Caenorhabditis elegans synthetic multivulva class B (synMuvB) gene products, including LIN-9, LIN-37, LIN-52, LIN-53, and LIN-54 (Litovchick et al., 2007). The DREAM complex binds the promoters of cell cycleregulated genes in serum starved quiescent human T98G cells and is required for their repression. It is yet to be known if other G0 and G1 arrest-inducing signals result in formation of the same DREAM complex, and whether assembly of this complex is associated with formation of the common gene expression signature that defines quiescence in NHF (Coller et al., 2006). Conceivably, E2F-repressor complexes formed in response to G1 arrest signals that do not result in cell cycle exit into quiescence may be different from the DREAM complex in one or several subunits. It is also possible that expression of HES1 in cells exiting into G0 is independent of the formation of DREAM, but it might play a role in defining the type of E2F/pocket protein complexes that assemble at cell cycle promoters.

1.2.4 What Are Cells Doing as They Exit Quiescence Back into G1? The classical video-microscopy experiments of Zetterberg and Larsson defined the period of time between post-mitotic G1 and R in NIH 3T3 cells (Zetterberg and Larsson, 1985). In these experiments, removal of mitogens for 1 h affected cell cycle length of only very young post-mitotic cells progressing through early G1. Mitogen removal for 1 h in early G1 resulted in elongation of the cell cycle by as much as 8 h. Others have shown that the time that cells require to exit quiescence is proportional to the time that they have spent in this state (Owen et al., 1989). One logical explanation that was drawn from these early studies is that cells need time to de novo transcribe and translate the gene products required for passage to R. More recently, changes in the assembly of DNA pre-replication factors onto replication origins have also been linked to quiescence. Blow and Hodgson proposed to define quiescence as a reversible withdrawal from the cell cycle characterized by unlicensed origins and lack of CDK activity (Blow and Hodgson, 2002). Origins of replication in metazoans are bound by ORC complexes in quiescent and

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cycling cells, but other components of the pre-replication complex are not loaded onto chromatin in quiescent cells (see Chapter 3 by McClendon et al., this volume). These include CDC6 and CDT1, both required for assembly of the multisubunit helicase composed of MCM2-7. Expression of MCM-2 in cells of the colonic crypt corroborates findings of cultured cells, as MCM2 is expressed at high levels in amplifying cells that are actively proliferating and is not expressed in terminally differentiated cells. MCM2 expression levels in the stem cells at the base of the crypts are lower than in the amplifying cells, which are consistent with their infrequent division (reviewed in Blow and Hodgson, 2002). A number of the subunits of the pre-replication complex are not expressed in quiescent cells because their genes are repressed by E2F-dependent mechanisms and because their protein products are targets of the APCCdh1 ubiquitin ligase, which is active when CDK activity is low (Diffley, 2004). In this regard, it is important to highlight that CDK-mediated phosphorylation of CDC6 has been linked to its stabilization and accumulation, as it prevents APC-mediated ubiquitination (Mailand and Diffley, 2005). CDC6 stabilization, accumulation, and loading onto replication origins may occur concomitantly with or downstream of its E2F-dependent transcription. Alternatively, CDC6 accumulation may be a mitogenically regulated CDK event partially independent of the E2F program. It is also notable that CDC6 expression has been linked to an “attachment checkpoint” that apparently operates at least partially independently of E2F-dependent transcription in NRK fibroblasts (Jinno et al., 2002). Thus, assembly of pre-RC emerges as another process linked to G1 checkpoints that mediate quiescence entry and exit.

1.3 Oncogenes That Cooperate to Bypass Quiescence Cellular oncogenic transformation is associated with unresponsiveness to antiproliferative and differentiation signals, bypassing of mitogenic extracellular requirements and an increase in proliferative lifespan. As multicellular organisms consist mostly of quiescent cells, critical oncogenic alterations may primarily deal with ensuring that cells initiating the transformation process remain in a stable proliferative state most of the time. In this regard, as the term “Restriction Point” was coined it was already proposed to be bypassed during malignancy (Pardee, 1974). A number of studies have focused on identifying activated oncogenes, viral transforming proteins, and/or tumor suppressor genes that, when deregulated alone or in combination, confer mitogen independency, the ability to bypass contact inhibition, and insensitivity to lack of substratum attachment. In other words, defining genetic alterations that bypass signals that ensure entry into quiescence in normal cells. The most informative studies have been performed testing the effect of combinations of oncogenes/inactivation of tumor suppressor genes in primary cells of different species in culture. That is, testing the ability of these cells to grow in suboptimal concentrations of serum, to form foci in cell monolayers and/or to grow in an anchorage-independent manner. Often, these studies have been followed up by testing if cells that appear transformed in culture form tumors when injected into

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nude mice. If so, these cells are typically designated “malignantly transformed.” A conclusion of these studies is the fact that the ability of certain oncogenes to bypass cellular quiescence varies among species, and human cells are more resistant to oncogenic transformation than murine cells (Hahn and Weinberg, 2002). It is also important to mention here that cells need to become immortal for malignant transformation. This is accomplished in many human cells via stabilization of chromosomal ends (telomeres), which shorten with each DNA replication cycle, as somatic cells do not express telomerase. In contrast, the majority of human tumor cells express telomerase or exhibit an alternative mechanism (ALT) to maintain chromosomal length (Stewart and Weinberg, 2006; Johnson and Broccoli, 2007) (see Chapter 8 by Denchi, this volume). In contrast, most studies performed using murine cells show that these cells become immortal by overcoming stress checkpoints, as most cell types used in these studies exhibit telomerase activity. For an in-depth analysis on immortalization, senescence, cancer, and aging readers are directed to specific reviews (Sherr and DePinho, 2000; Hahn, 2002; Blasco and Hahn, 2003; Serrano and Blasco, 2007). Thus, what oncogenes or inactivated tumor suppressor genes help bypass quiescence? Ectopic expression of c-MYC and an activated RAS oncogene in normal immortal rat fibroblasts (REF-52 cells) induce cyclin E/CDK2 activity and exit from quiescence in low concentrations of serum (Leone et al., 1997). In quiescent REF-52 cells, cellular RAS is required for activation of CDK activity and E2F-dependent gene expression in response to mitogenic stimulation, but activated RAS is insufficient to induce CDK activation in quiescent REF-52 cells placed in low serum. Coexpression of c-MYC allows cyclin E/CDK2 activation likely via downregulation of CKIs (Leone et al., 1997). Others have shown that ectopic expression of an oncogenic version of RAS induces premature senescence in human and mouse cells through activation of the RAF/MAPK pathway leading to activation of p53/ARF. This arrest is bypassed by depletion of p53 function in mouse cells, but disruption of both the pRB and p53 pathways is necessary to bypass RAS-induced senescence in human cells (Serrano, 1997, 1998) (see Chapter 9 by Adams, this volume). Thus, the c-MYC/RAS pair fails to transform mouse fibroblasts unless the p53/ARF pathway is mutated, and human cells require alteration of both the p53 and pRB pathways, as well as activation of telomerase in order to become transformed (Hahn et al., 2002; Hahn and Weinberg, 2002). This likely explains why normal human fibroblasts as opposed to immortal REF-52 cells fail to exit quiescence upon coexpression of c-MYC and an activated RAS oncogene (Sotillo et al., 2008). Thus, because the same oncogenic signals that induce escape from quiescence also induce senescence, alteration of the p53/ARF pathway is critical for oncogenesis in mouse cells, while alteration of both the p53 and pRB pathways is required in human cells. Thorough transformation assays from the Hahn and Weinberg laboratories have demonstrated that expression of oncogenic RAS, SV40 large T (LT) and small t (st) antigens, and the catalytic subunit of telomerase (hTERT) is sufficient to transform NHF (Hahn et al., 1999), human mammary epithelial cells (Elenbaas et al., 2001), and kidney epithelial cells (Hahn et al., 2002). Subsequent studies have shown that in this setting LT and st can be substituted by inactivation of p53, pRB, and PTEN plus constitutive

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expression of c-MYC (Boehm et al., 2005). The role of the SV40 tumor antigens will be discussed in more detail in the next section. Examination of NHF with various combinations of genetic alterations showed that NHF-expressing hTERT, a dominant negative version of p53 and deregulated c-MYC, exit quiescence in low concentrations of serum, but subsequent disruption of pRB via shRNA made these cells insensitive to serum starvation (Boehm et al., 2005). Interestingly, as mentioned above disruption of the three pocket proteins in MEFs appears to be required to make these cells largely insensitive to mitogen withdrawal, as disruption of either pRB or p130/p107 delays but does not prevent cell cycle exit (Sage et al., 2000). Interestingly, in addition to its role in irreversible senescence and apoptosis, p53 has been suggested to participate in cell cycle exit into quiescence, as p53 expression and activity increase in quiescent NHF, and p53 inactivation by different means delays reversible cell cycle exit in response to mitogenic withdrawal (Itahana et al., 2002). c-MYC has been shown to induce expression of the so-called E2F-activators (E2F1-3) and to directly interact with their promoters upon mitogenic stimulation (Leone et al., 2001; Fernandez et al., 2003; Leung et al., 2008). Of note, c-MYC binding to E2F-promoters seems critical for the loading of E2F1 to these promoters. Besides those genes required for cell cycle entry, E2F1 also regulates a group of genes involved in apoptosis. It has been shown that activation of the PI3K/AKT pathway during mitogenic stimulation inhibits E2F1 pro-apoptotic targets, favoring the role of E2F1 as an inducer of proliferation rather than apoptosis (Hallstrom et al., 2008; Hallstrom and Nevins, 2009). The importance that activation of PI3K/AKT pathway has on oncogenic transformation is underscored by the fact that mutations in PTEN, a key negative regulator of this pathway, and amplification and abnormal activation of PI3K and AKT are associated with many types of human cancers (Keniry and Parsons, 2008; Yuan and Cantley, 2008). It has been shown that ectopic expression of the active subunit of PI3K, p110a, can substitute for st when coexpressed with LT and hTERT in human epithelial cells, promoting growth in low concentrations of serum as well as proliferation in soft agar (Zhao et al., 2003). In this scenario, st can also be substituted by coexpression of activated alleles of AKT1 and RAC, a downstream effector of the PI3K/AKT pathway. Also, forced coactivation of the RAS/RAF/MEK pathway with AKT elicited a robust proliferative response leading to activation of G1 cyclin/CDKs resulting from cyclin D1 accumulation and p27 repression, as well as removal of p21 from cyclin E/CDK2 complexes (Mirza et al., 2004). In this regard, the PI3K pathway negatively regulates FOXO transcription factors via AKT-mediated phosphorylation and exclusion from the nucleus. Activation of FOXO transcription factors is associated with cell cycle exit into quiescence in nonhematopoietic cells and has been implicated in the transcriptional activation of p27 as well as downregulation of D-type cyclins (Medema et al., 2000; Schmidt et al., 2002). Moreover, the PI3K pathway has been shown to cooperate with c-MYC in the expression of c-MYC-dependent genes by inactivating FOXO, which is implicated in the negative regulation of multiple c-MYC genes (Bouchard et al., 2004). FOXO transcription factors have been implicated in long-term survival of quiescent cells (Burgering and Medema, 2003).

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The adenoviral oncoprotein E1A has also been long known to have the ability to stimulate exit from cellular quiescence. Two recent studies suggest how E1A triggers cell cycling and inhibits the cellular antiviral response and differentiation (Ferrari et al., 2008; Horwitz et al., 2008). E1A expression in quiescent fibroblasts results in the global relocation of pocket proteins and the p300/CBP acetyltransferase on cellular promoters. This process occurs in a sequential manner, leading to acetylation of lysine 18 on histone H3 and promoter transactivation of a restricted number of genes involved in proliferation and growth. However, both E1A and SV40 LT cause a global decrease in the acetylation of histone H3 at this site, which is apparently due to the restriction of HATs to the subset of proliferation/growth genes concomitant to the exclusion of these proteins on other gene promoters. Thus, hypoacetylation at histone H3 lysine 18 may be a general consequence of DNA tumor oncogenesis, which is linked to quiescence exit (reviewed in Ferrari et al., 2009). It is also important to highlight recent findings that suggest that HES1, whose increased expression in quiescent cells has been associated with conferring the reversible nature of this state, is found expressed at high levels in rhabdomyosarcoma tumors and derived cell lines (Sang et al., 2008). Rhabdomyosarcomas are aggressive tumors that express the muscle differentiation factor MyoD, but exhibit a block in myogenic differentiation. Forced inactivation of HES1 in a rhabdomyosarcoma cell line via expression of a dominant negative HES1 mutant or pharmacological inhibition of Notch, which positively regulates HES1 expression, promoted MHC expression and differentiation of these cells. In summary, to endow normal cells with the capability of exiting quiescence in unfavorable environments, oncogenes/inactivated tumor suppressor genes must mimic mitogenic signals. Cells with these alterations may produce their own mitogens, force surrounding cells to do so, or exhibit constitutively activated downstream signaling pathways independently of mitogenic stimulation. Some of these alterations as well as others will also help bypass antiproliferative signals from the environment or will make the cell independent of substrate feeding. Phosphorylation of pocket proteins and activation of the E2F transcription program are critical, but exit from quiescence does not always lead to effective proliferation. Thus, other pathways, such as those that inhibit apoptosis and cell senescence and/or are involved in monitoring faithful DNA replication, must also be altered.

1.4 SV40 and Exit from Quiescence 1.4.1 SV40 Tumor Antigens and Their Cellular Targets As described in Section 1.3, transformation assays designed to identify the precise combination of altered pathways that are required for transformation of a variety of normal human cells have revealed that expression of SV40 LT and st antigens, oncogenic RAS, and hTERT suffices to ensure the required alterations (Hahn et al., 1999; Elenbaas et al., 2001; Hahn et al., 2002). Transformation is also attained with

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expression of oncogenic RAS, c-MYC, hTERT, st and inactivation of both p53 and pRB (Boehm et al., 2005). However, expression of st, hTERT, and pRB inactivation is not required when comparable transformation assays are performed using rodent cells. The effects of st on transformation of human cells are thought to be, at least in part, due to its ability to facilitate proliferation in conditions that promote quiescence. This indicates that the ability of st to facilitate a bypass of the quiescent state may be uniquely critical for transformation of human cells. Thus, in this section we will discuss the effects of expression of SV40 antigens in human cells with a major focus on st. In the early 1960s, Polyomavirus Simian Virus 40 (SV40) was discovered as a viral contaminant during the production of poliovirus vaccines from cultures from rhesus monkey kidney cells (Eddy et al., 1962). Soon after its discovery, SV40 was shown to induce tumors in newborn hamsters (Girardi et al., 1962; Gerber, 1963). However, it took several years and work from multiple laboratories to demonstrate that the expression of proteins encoded in the Early Region (ER) of SV40 was responsible for oncogenic transformation (reviewed in Chen and Hahn, 2003). SV40 ER encodes three proteins that share 82 amino acids in their aminoterminal end, which includes a DnaJ chaperone domain. However, their unique carboxy-terminal extensions are generated through alternative splicing resulting in the synthesis of three separate protein products designated large T (LT), small t (st), and 17KT antigens (reviewed in Ali and DeCaprio, 2001; Chen and Hahn, 2003; Pipas, 2009). The role of both LT and st in cellular transformation and tumorigenesis has been extensively studied and both proteins have been major tools for identification and characterization of key signaling pathways commonly altered during cancer development (Pipas, 2009). The C-terminus of LT targets the tumor suppressor gene product p53 while an LXCXE motif present in the amino-terminal end of the unique domain targets the three pocket proteins pRB, p130, and p107 (reviewed in Ali and DeCaprio, 2001) (Fig. 1.3). Thus, LT inactivates two major suppressor pathways that are found inactivated in most tumor cells. On the other hand, the unique carboxy-terminal domain of st associates with and inhibits PP2A activities that are not yet completely defined (Pallas et al., 1990; Yang et al., 1991; Pallas et al., 1992). PP2A is an heterotrimeric serine/threonine phosphatase that consists of a catalytic subunit (PP2A/C), a structural subunit (PP2A/A), and a variable B subunit that dictates subcellular localization and substrate specificity (reviewed in Virshup and Shenolikar, 2009). There are two isoforms each for PP2A/C and PP2A/A subunits, and four distinct families of conserved PP2A/B subunits encoded from at least 15 different genes, many with multiple splice variants. Thus, the combination of PP2A/A/B/C subunits yields multiple possible heterotrimers to specifically target a variety of substrates that play key roles in cellular proliferation, DNA damage, and viability among many other cellular processes. st binds to the PP2A/A/C dimer through a cystein-rich region interfering with the binding of the PP2A/B subunit, thus likely precluding specific substrate recognition and/or proper subcellular localization. While in human cells the transforming pathways targeted by LT have been clearly linked to disruption of pRB/pocket proteins and p53-dependent pathways

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Fig. 1.3 SV40 st antigen disrupts PP2A heterotrimeric complexes and upregulates a number of cell cycle proteins. The early region of SV40 encodes large T antigen (LT) that inactivates p53 and pocket proteins and small t antigen (st) that targets a still not well-defined set of PP2A heterotrimeric complexes. st binds to the PP2A dimer composed of a catalytic subunit (PP2A/C) and a scaffold subunit (PP2A/A), displacing B regulatory subunits. Displacement of B subunits of the B56 family has been linked to transformation and oncogenic upregulation (c-MYC upregulation). st expression in quiescent fibroblasts has been shown to upregulate a variety of pathways resulting in unscheduled expression of key cell cycle regulators and replication factors

by making cells insensitive to checkpoint and antiproliferative signaling, the unique pathways altered by st are still poorly defined due to the vast number of potentially critical cellular pathways where distinct trimeric PP2A holoenzymes play critical but not well understood roles (reviewed in Chen and Hahn, 2003; Skoczylas et al., 2004). In the next section, we will primarily focus on the analysis of those pathways targeted by st that promote exit from quiescence.

1.4.2 SV40 Small t Antigen Promotes Exit from Quiescence The effects of st on transformation of human cells are thought to be, at least in part, due to its ability to facilitate proliferation in reduced concentrations of growth factors. This function is dependent on its ability to bind and inhibit PP2A, as st mutants unable to bind PP2A fail to cooperate with LT driving cell proliferation in

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a limiting mitogenic environment (Skoczylas et al., 2004). Expression of st stimulates growth of monkey kidney cells (CV-1) maintained in 0.1% serum to an extent comparable to serum. In this scenario quiescence exit is accompanied by activation of the RAF/MAPK pathway (Sontag et al., 1993). It was later shown that activation of PKCζ through st-mediated inhibition of PP2A would also contribute to further activate the MAPK pathway, as well as activate NFkB-dependent gene expression (Sontag et al., 1997) in both CV-1 and NIH3T3 cells in 0.1% serum. In the same study it was determined that pharmacological inhibition of the PI3K pathway or a dominant negative mutant of p85 blocks st-mediated activation of PKCζ and NFκB, as well as st-induced cell proliferation (Fig. 1.3). The importance of the PI3K pathway for st-mediated transformation has been discussed earlier in this chapter when referring to combinations of oncogenes able to induce exit from quiescence in normal cells (Fig. 1.3). Progress has more recently been made identifying PP2A heterotrimers that could mediate st transforming activities. In human embryonic kidney (HEK) cells, PP2A/B56γ has been identified as a potential target of st transforming activity. In these cells, knockdown of B56γ substitutes for st in transformation assays with defined combinations of altered genes (Chen et al., 2004). However, it was subsequently shown that B56γ cannot substitute for st in cells maintained in suboptimal mitogen concentrations (Moreno et al., 2004), suggesting that st has other targets important to bypass serum requirements. A second member of the same family of B subunits, B56α, has been identified as a negative regulator of c-MYC protein stability, providing a mechanism to explain how st increases c-MYC expression. Interestingly, a stable c-MYC mutant that cannot be dephosphorylated substitutes for st in transformation assays (Yeh et al., 2004; Arnold and Sears, 2006). Downstream targets of st have also been studied. A number of reports have described transcriptional activation of cyclin D1 and cyclin A promoters in reporter assays (Porras et al., 1996; Watanabe et al., 1996; Skoczylas et al., 2005). Cyclin A protein levels were also shown to be upregulated by st in density-arrested human fibroblasts stimulated with serum, concomitantly with downregulation of p27. In this scenario, cyclin A is inactive and cells do not enter the cell cycle unless LT is coexpressed (Porras et al., 1999). In search for oncogenes that cooperate to bypass quiescence induced by complete depletion of mitogens in NHF, our lab found that coexpression of st and cyclin E, but not their individual expression, was sufficient to bypass quiescence and induce DNA synthesis (Sotillo et al., 2008). This same combination of oncogenes bypassed quiescence induced by growth to high density and lead to continued proliferation and foci formation in hTERT-NHF. These events are at least partially dependent on PP2A inhibition. Expression of st alone in quiescent NHF did not lead to accumulation of cyclins A or D1, but strikingly led to accumulation and loading of the essential replication factor CDC6 (Sotillo et al., 2009). As MCM2 loading onto chromatin was also observed in density-arrested cells expressing st, these results suggest that st induces steps toward licensing of replication origins, a characteristic of cells exiting G0 into G1 (see Section 1.2.4 above). When cyclin E and st were coexpressed in quiescent NHF, CDC6 further accumulated coinciding with CDK2 activation and

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DNA synthesis. In addition, we and others have observed that CDC6 expression, as well as the expression of other pre-RC components, is linked to phosphorylation of CDK2 on its activating T-loop (Nevis et al., 2009; Sotillo et al., 2009). Therefore, deregulation of cyclin E expression in the context of normal cells apparently driven out of quiescence by st leads to the cooperative and coordinated activation of an essential pre-replication complex factor (CDC6) and an activity required for origin firing (CDK2) (Sotillo et al., 2009). Importantly, it was also found that in the context of this oncogenic-driven exit from G0 and proliferation, CDK2 activity appeared to be essential (Sotillo et al., 2008). While the direct target of st in this case is unknown the current data suggest that the selective accumulation of the CDC6 transcript is dependent on E2F promoter elements but independent of CDK activation. This suggests that st controls factors that can selectively regulate the expression of a gene(s) whose expression is associated with exit from G0. Finally, it is important to point out that despite the observation that CDC6 expression is required for passage through an “attachment checkpoint” in NRK fibroblasts (Jinno et al., 2002), expression of cyclin E and st failed to induce anchorage independent growth of hTERT NHF, indicating that this oncogene pair does not reverse quiescence induced by all signals. Altogether these studies show that st changes a fundamental property of quiescent cells that differentiates them from post-mitotic G1 cells, which is the status of the cell replication origins. By facilitating passage through the G0/G1 transition, st may trick certain cells to create an environment proper for viral replication despite extracellular signaling that would otherwise keep cells in a quiescent state. Understanding how st disrupts the mechanisms that ensure maintenance of the quiescent state will provide insight into the molecular nature of this state, increase our understanding of how DNA tumor viruses promote their own replication, and may unveil novel mechanisms for cellular transformation that are exclusive to human cells.

1.5 Future Directions The defining characteristic of cells in the quiescent state is maintenance of their ability to re-enter the cell cycle in propitious conditions. Recent work has shown that while cells exiting the cell cycle into quiescence initiate programs of gene expression that are coupled to the quiescent inducing signal, there is a common gene expression signature that emerges in a time-dependent manner. This signature appears associated with acquisition of resistance to differentiation, senescence, and cell death. A transcription factor designated HES1 has been shown to be upregulated during cell cycle exit into quiescence and appears to be required for maintenance of the quiescent state, as it blocks both differentiation and senescence in human fibroblasts. This factor is found deregulated in rhabdomyosarcomas and mediates a block in myogenic differentiation exhibited by these tumor cells. Work over the past several decades has led to the identification of oncogenes and tumor suppressor genes that, when deregulated, facilitate mitogen-independent cell cycle progression and insensitivity to other signals that induce exit into quiescence. SV40 st antigen

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has emerged as a key player in the specific transformation of human cells, as its unique transforming activity is not required in rodent cells. st appears to facilitate transformation in certain environments that favor cell cycle exit into quiescence, such as mitogen starvation and contact inhibition. Recent progress in identification of the PP2A heterotrimers that are targeted by st, as well as the downstream cell cycle players that mediate st activities that drive cells out of quiescence are likely to provide important insights in the near future. Acknowledgments We thank Manuel Serrano, David G. Johnson, Alison Kurimchak, and Judit Garriga for critically reading this manuscript and helpful suggestions. Work in this lab has been supported by a grant project under CA095569 and a Career Development Award (K02 AI01823) to XG of the National Institutes of Health.

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Pallas DC, Weller W, Jaspers S, Miller TB, Lane WS, Roberts TM (1992) The third subunit of protein phosphatase 2A (PP2A), a 55-kilodalton protein which is apparently substituted for by T antigens in complexes with the 36- and 63-kilodalton PP2A subunits, bears little resemblance to T antigens. J Virol 66: 886–893. Pardee AB (1974) A restriction point for control of normal animal cell proliferation. Proc Natl Acad Sci U S A 71: 1286–1290. Pipas JM (2009) SV40: cell transformation and tumorigenesis. Virology 384: 294–303. Porras A, Bennett J, Howe A, Tokos K, Bouck N, Henglein B, Sathyamangalam S, Thimmapaya B, Rundell K (1996) A novel simian virus 40 early-region domain mediates transactivation of the cyclin A promoter by small-t antigen and is required for transformation in small-t antigendependent assays. J Virol 70: 6902–6908. Porras A, Gaillard S, Rundell K (1999) The simian virus 40 small-t and large-T antigens jointly regulate cell cycle reentry in human fibroblasts. J Virol 73: 3102–3107. Quelle DE, Ashmun RA, Shurtleff SA, Kato JY, Bar SD, Roussel MF, Sherr CJ (1993) Overexpression of mouse D-type cyclins accelerates G1 phase in rodent fibroblasts. Genes Dev 7: 1559–1571. Resnitzky D, Gossen M, Bujard H, Reed SI (1994) Acceleration of the G1/S phase transition by expression of cyclins D1 and E with an inducible system. Mol Cell Biol 14: 1669–1679. Rowland BD, Bernards R (2006) Re-evaluating cell-cycle regulation by E2Fs. Cell 127: 871–874. Sage J, Mulligan GJ, Attardi LD, Miller A, Chen S, Williams B, Theodorou E, Jacks T (2000) Targeted disruption of the three Rb-related genes leads to loss of G(1) control and immortalization. Genes Dev 14: 3037–3050. Sang L, Coller HA, Roberts JM (2008) Control of the reversibility of cellular quiescence by the transcriptional repressor HES1. Science 321: 1095–1100. Santamaria D, Barriere C, Cerqueira A, Hunt S, Tardy C, Newton K, Caceres JF, Dubus P, Malumbres M, Barbacid M (2007) Cdk1 is sufficient to drive the mammalian cell cycle. Nature 448: 811–815. Schmidt M, Fernandez de Mattos S, van der Horst A, Klompmaker R, Kops GJ, Lam EW, Burgering BM, Medema RH (2002) Cell cycle inhibition by FoxO forkhead transcription factors involves downregulation of cyclin D. Mol Cell Biol 22: 7842–7852. Serrano M, Blasco MA (2007) Cancer and ageing: convergent and divergent mechanisms. Nat Rev Mol Cell Biol 8: 715–722. Serrano M, Lin AW, McCurrach ME, Beach D, Lowe SW (1997) Oncogenic ras provokes premature cell senescence associated with accumulation of p53 and p16INK4a. Cell 88: 593–602. Sherr CJ, DePinho RA (2000) Cellular senescence: mitotic clock or culture shock? Cell 102: 407–410. Sherr CJ, Roberts JM (1999) CDK inhibitors: positive and negative regulators of G1-phase progression. Genes Dev 13: 1501–1512. Sherr CJ, Roberts JM (2004) Living with or without cyclins and cyclin-dependent kinases. Genes Dev 18: 2699–2711. Skoczylas C, Fahrbach KM, Rundell K (2004) Cellular targets of the SV40 small-t antigen in human cell transformation. Cell Cycle 3: 606–610. Skoczylas C, Henglein B, Rundell K (2005) PP2A-dependent transactivation of the cyclin A promoter by SV40 ST is mediated by a cell cycle-regulated E2F site. Virology 332: 596–601. Sontag E, Fedorov S, Kamibayashi C, Robbins D, Cobb M, Mumby M (1993) The interaction of SV40 small tumor antigen with protein phosphatase 2A stimulates the map kinase pathway and induces cell proliferation. Cell 75: 887–897. Sontag E, Sontag JM, Garcia A (1997) Protein phosphatase 2A is a critical regulator of protein kinase C zeta signaling targeted by SV40 small t to promote cell growth and NF-kappaB activation. EMBO J 16: 5662–5671.

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Sotillo E, Garriga J, Kurimchak A, Cook J, Grana X (2008) Cyclin E and SV40 small T antigen cooperate to bypass quiescence and contribute to transformation by activating CDK2 in human fibroblasts. J Biol Chem 283: 11280–11292. Sotillo E, Garriga J, Padgaonkar A, Kurimchak A, Cook J, Grana X (2009) Coordinated activation of the origin licensing factor CDC6 and CDK2 in resting human fibroblasts expressing SV40 small T antigen and cyclin E. J Biol Chem 284: 14126–14135. Stewart SA, Weinberg RA (2006) Telomeres: cancer to human aging. Annu Rev Cell Dev Biol 22: 531–557. Virshup DM, Shenolikar S (2009) From promiscuity to precision: protein phosphatases get a makeover. Mol Cell 33: 537–545. Watanabe G, Howe A, Lee RJ, Albanese C, Shu IW, Karnezis AN, Zon L, Kyriakis J, Rundell K, Pestell RG (1996) Induction of cyclin D1 by simian virus 40 small tumor antigen. Proc Natl Acad Sci U S A 93: 12861–12866. Yang SI, Lickteig RL, Estes R, Rundell K, Walter G, Mumby MC (1991) Control of protein phosphatase 2A by simian virus 40 small-t antigen. Mol Cell Biol 11: 1988–1995. Yeh E, Cunningham M, Arnold H, Chasse D, Monteith T, Ivaldi G, Hahn WC, Stukenberg PT, Shenolikar S, Uchida T, Counter CM, Nevins JR, Means AR, Sears R (2004) A signalling pathway controlling c-Myc degradation that impacts oncogenic transformation of human cells. Nat Cell Biol 6: 308–318. Yuan TL, Cantley LC (2008) PI3K pathway alterations in cancer: variations on a theme. Oncogene 27: 5497–5510. Zetterberg A, Larsson O (1985) Kinetic analysis of regulatory events in G1 leading to proliferation or quiescence of Swiss 3T3 cells. Proc Natl Acad Sci U S A 82: 5365–5369. Zhao JJ, Gjoerup OV, Subramanian RR, Cheng Y, Chen W, Roberts TM, Hahn WC (2003) Human mammary epithelial cell transformation through the activation of phosphatidylinositol 3-kinase. Cancer Cell 3: 483–495.

Chapter 2

Interplay Between Cyclin-Dependent Kinases and E2F-Dependent Transcription Jun-Yuan Ji and Nicholas J. Dyson

Abstract Precise control of cell proliferation is essential for normal development and survival of all multi-cellular organisms. The deregulation of cell proliferation is a fundamental feature of all types of cancer. One of the key regulators of cell proliferation is the E2F transcription factor. E2F controls the expression of many genes that are required for cells to divide and elevated E2F activity is found in most tumor cells. The activation and inactivation of E2F are tightly linked to the activation of cyclin-dependent kinases (CDKs). In normal cells, these connections allow the periodic oscillations in CDK cycle to be coupled with temporal programs of gene expression. Multiple CDK–cyclin complexes (including CDK1/2–CycA, CDK1/2– CycB, and CDK7–CycH) have been shown to directly phosphorylate E2F or its dimerization partner DP. However, in recent genetic studies, one of the strongest modifiers of E2F-dependent phenotypes was cdk8, a kinase that had not previously been linked to E2F. In this review, we summarize the effects of CDKs on E2F1 activity and describe a model that may explain the role of CDK8–CycC in E2F regulation. Since CDKs can both increase and decrease E2F activity, understanding the interplay between E2F and CDK–cyclin complexes may suggest therapeutic approaches to efficiently block cancer cell proliferation.

2.1 Cell Cycle Progression Is Driven by the Integrated Action of Cyclin-Dependent Kinases and a Transcriptional Network The mitotic cell cycle is composed of an S phase (for DNA synthesis) and an M phase (for mitosis), separated by two gap phases, G1 and G2. Progression through

J.-Y. Ji (B) Department of Pathology, Harvard Medical School, Massachusetts General Hospital Cancer Center, Charlestown, MA 02129, USA e-mail: [email protected]


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these phases is driven by the periodic activation of CDK (cyclin-dependent kinase)– cyclin complexes. One of the most important discoveries in the field of cell biology was that these central components of the cell cycle machinery have been conserved during eukaryote evolution (Nasmyth, 1995; Nurse, 2000). The conservation of CDK function is so extensive that the human CDK1 (CDC2) gene is able to substitute for its functional orthologs in yeast (Lee and Nurse, 1987). Genome sequencing has revealed that mammalian cells contain at least 13 CDKs and 29 cyclins (Liu and Kipreos, 2000; Malumbres and Barbacid, 2009). CDK1– CDK6, CDK10, CDK11, and the CDK-activating kinase, CDK7, are all involved in cell cycle control (Fisher, 2005; Loyer et al., 2005; Malumbres and Barbacid, 2009). Of these, CDK1 is the most significant: genetic and molecular experiments show that CDK1 is both necessary and sufficient to drive cell cycle progression in species as diverse as yeast and humans (Nurse, 1990; Malumbres and Barbacid, 2009). To some degree, the large number of human CDKs may reflect tissue-specific and finetuned regulation of cell cycle regulation in metazoans. In addition, it has become clear that some CDKs have activities that are distinct from cell cycle control. For example, CDK7, CDK8, and CDK9 are all able to phosphorylate the carboxylterminal domain (CTD) of RNA polymerase II and have functions in transcriptional regulation (Fisher, 2005; Loyer et al., 2005; Phatnani and Greenleaf, 2006). CDK12 and CDK13 bind to L-type cyclins (CycL) and regulate alternative RNA splicing (Chen et al., 2006; Chen et al., 2007). Analysis of CDK function in early embryos and embryo extracts showed that fluctuations in CDK activity are sufficient to regulate cell cycle transitions in the absence of transcription. However, in most eukaryotic cells, cell cycle progression is accompanied by dynamic changes in gene expression patterns. The recent analysis of transcriptional profiles in yeast has shown that the transcription of more than 70% of periodically expressed genes continues to oscillate after B-type cyclins have been removed (Orlando et al., 2008). Although these cells cannot complete DNA replication or mitosis, some cell cycle events continue to occur in a periodic manner (Haase and Reed, 1999). Computational analysis of the transcriptional network shows that it has the properties of an oscillator (Orlando et al., 2008). These findings suggest that cell cycle regulation results from the tightly integrated activities of two oscillators (a CDK oscillator and a transcriptional oscillator) that fluctuate in tandem. Transcriptional events that respond to changes in CDK activity, and the periodic expression of CDK regulators, couple these oscillators together and ensure that changes in CDK activity are coordinated with the appropriate programs of gene expression. The interwoven nature of the CDK and transcriptional oscillators is beautifully illustrated by studies of the RB/E2F network. In mammalian cells, RB/E2F proteins play a critical role in the control of the G1 to S transition. CDKs contribute to both activation and inactivation of E2F-dependent transcription. Moreover, E2F complexes control the expression of genes encoding CDKs, cyclins, and their regulators. As described below, genetic studies have revealed a new player in this network of interactions. Unexpectedly, the levels of CDK8 have a significant impact on E2F transcriptional activity. Evidence that CDK8 is frequently amplified

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and overexpressed in colorectal cancer cells suggests that CDK8 levels may be particularly important in this type of cancer.

2.2 Rb and E2F Proteins Regulate the G1 to S-Phase Transition in Higher Eukaryotes The G1 to S-phase transition is a critical event for the control of cell proliferation. With few exceptions, once cells complete this transition they are committed to progress through the remainder of the cycle and will divide into two daughter cells. In higher eukaryotes, the retinoblastoma (RB) tumor suppressor protein and E2F transcription factors play pivotal roles in the regulation of the G1 to S-phase transition (reviewed by Dyson, 1998; Lipinski and Jacks, 1999; Müller and Helin, 2000; Zhu, 2005). The E2F transcription factor provides a temporal control over the expression of hundreds of target genes whose products are necessary for accurate DNA replication and mitosis (Müller et al., 2001; Ren et al., 2002; Bracken et al., 2004). In non-dividing cells E2F proteins act together with RB-family proteins to repress the transcription of these targets, helping to maintain the quiescent state (G0) (reviewed by Classon and Dyson, 2001; Frolov and Dyson, 2004; Burkhart and Sage, 2008). This repression must be relieved for cells to proliferate. In response to the activation of G1 cyclins, E2F switches from a repressor to an activator, and drives the expression of E2F target genes as cells leave G0/G1 and progress through the cell cycle (Trimarchi and Lees, 2002; Bracken et al., 2004; Dimova and Dyson, 2005). E2F was initially identified as a factor required for the activation of the E2 promoter of adenovirus (see Nevins, 1992 for a review) and it is the composite transcriptional activity of a group of proteins that share similar DNA-binding domains. The basic unit of E2F is a heterodimer composed of an E2F and a DP subunit (Dyson, 1998). Eight E2F genes and three DP genes have been identified in mammals (reviewed in Dimova and Dyson, 2005; DeGregori and Johnson, 2006). E2F1–E2F6 bind to DNA in association with a DP subunit. Structural studies show that both DP and E2F subunits contain DNA-binding domains (Zheng et al., 1999) and E2F/DP dimerization is essential for high-affinity DNA binding and transcriptional activity (reviewed in Dyson, 1998). E2F proteins are often subdivided into groups that primarily activate (E2F1, E2F2, and E2F3a) or repress transcription (E2F3b, E2F4, E2F5, E2F6, E2F7, and E2F8) (reviewed in Dimova and Dyson, 2005; DeGregori and Johnson, 2006). Repressor E2Fs are relatively abundant and broadly expressed, while activator E2Fs are potent and their activities are under tight control (Trimarchi and Lees, 2002; Attwooll et al., 2004). E2F7 and E2F8 act without a DP subunit and, like E2F6, are thought to repress transcription without directly binding to RB family proteins (Maiti et al., 2005; Milton et al., 2006). In contrast, the transcriptional properties of E2F1–E2F5 are controlled by a direct physical interaction with the RB family members. Activator E2Fs have the ability to potently reverse the effects of repressor E2Fs at target promoters.

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Figure 2.1 summarizes the general properties of the RB-E2F regulatory network. In early G1 phase of the cell cycle, RB family proteins (pRB, p107, and p130) bind to E2F family members and recruit co-repressor complexes, thereby repressing E2Fdependent transcription (Fig. 2.1a). Upon growth factor stimulation, CDK–cyclin (Cyc) complexes, such as CDK4/6–CycD, phosphorylate pRB and partially relieve its repressive activity. This allows the expression of CycE, which binds to and activates CDK2. CDK2–CycE further phosphorylates pRB, resulting in its complete inactivation (Fig. 2.1b). Hyperphosphorylated pRB can no longer bind to E2F, effectively allowing E2F transcription factors to activate transcription (Dyson, 1998; Nevins, 1998; Müller and Helin, 2000; Fig. 2.1c). Some of the best-known transcriptional targets for E2F are components or regulators of CDKs, such as Cyclin A2 (CCNA2; Schulze et al., 1995; DeGregori et al., 1995; Shan et al., 1996; Ren et al., 2002), Cyclin B1 (CCNB1; Zhu et al., 2004), Cyclin D1 (CCND1; Lee et al., 2000),

G1 phase

A Co-repressor complexes

target gene expression

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OFF DP CDK2-CycE CDK2-CycA

B growth factor Stimulations (eg., EGF)

CDK4-CycD

pRB

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C p

p

p

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target gene expression E2F

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cyclin A cyclin E cdc25 cdk1 ……

E2F-DP

ON

expression of target genes

G1 to S-phase transition and mitosis

PCNA TK ORC MCM ……

DNA replication

Apaf1 p73 ……

Apoptosis

Fig. 2.1 A summary of the interplay between RB/E2F and CDKs during the G1 to S-phase transition. (a) During G1 phase, pRB blocks the transcriptional activity of E2Fs by directly binding to the transactivation domain of activator E2Fs and/or by recruiting transcriptional corepressor complexes to target promoters. (b) Growth factor stimulation leads to activation of G1 CDKs, the phosphorylation of pRB (and p107 and p130), the disruption of repressor complexes, and increased transcription. E2F targets include genes with key functions in cell cycle progression, DNA replication, and genes that can potentially sensitize cells toward apoptosis. Note that increased transcription of CycE triggers a feedback loop that promotes the phosphorylation of pRB. E2F-induction of CycA also increases pRB phosphorylation but CycA-associated kinases can also phosphorylate E2F1 and down-regulate its activity. (c) In early S-phase, the phosphorylation of pRB-family members by CDK4–CycD, CDK2–CycE, and CDK2–CycA results in the complete release of repressor complexes allowing activator E2Fs to drive gene expression. For simplicity, we have used pRB to refer the three pRB family proteins; the specific and overlapping functions of pRB family proteins are reviewed by Classon and Dyson (2001) and Burkhart and Sage (2008)

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Cyclin D3 (CCND3; Müller et al., 2001), Cyclin E1/2 (CCNE1/2; DeGregori et al., 1995; Ohtani et al., 1995; Botz et al., 1996; Geng et al., 1996; Shan et al., 1996), Cyclin G2 (CCNG2; Müller et al., 2001), CDK1 (CDC2; DeGregori et al., 1995; Shimizu et al., 1995; Tommasi and Pfeifer, 1995; Ren et al., 2002; Zhu et al., 2004), CDK2 (Shan et al., 1996; Ren et al., 2002), and CDC25A (Vigo et al., 1999; Ren et al., 2002). The importance of the RB/E2F network for normal control of cell proliferation is highlighted by evidence that this regulation is inactivated in most types of tumor cells (Weinberg, 1995; Dyer and Bremner, 2005; Tsantoulis and Gorgoulis, 2005; Burkhart and Sage, 2008). Mutation of both copies of the RB gene is rate-limiting for the development of both familial and sporadic retinoblastoma. The RB tumor suppressor is also mutated at a high frequency in small cell lung carcinomas and osteosarcoma (Weinberg, 1995; Dyer and Bremner, 2005). In other cancers, a variety of alternative events have been shown to functionally inactivate pRB. These include (a) the loss of the CDK inhibitor p16INK4a by deletion, point mutation, or promoter methylation, (b) the over-expression of CycD1 by gene rearrangement or amplification, (c) gain-of-function mutations in genes encoding either CDK4 or CycD1, (d) the expression of viral oncoproteins, (e) amplification of the E2F3 gene, and (f) additional changes resulting in increased CDK2–CycE activity (Weinberg, 1995; Sherr, 1996; Nevins, 2001; Johnson and Degregori, 2006). All of these events have a net effect of deregulating E2F-dependent transcription, generating a cellular environment that is permissive for cell proliferation.

2.3 CDK Phosphorylation Is One of Several Mechanisms That Regulate E2F Activity The levels and activity of E2Fs are regulated at multiple levels. In general, activator E2Fs are not highly expressed in quiescent cells but are transcribed in response to growth factor stimulation. The E2F1 promoter contains E2F-binding sites (Johnson et al., 1993; Hsiao et al., 1994) and this positive-feedback loop helps to amplify the levels of activator E2Fs. E2F1–E2F3 are constitutively localized in the nucleus because they contain a nuclear localization signal (NLS). This signal is absent in E2F4 (Verona et al., 1997) and E2F4-containing complexes have been observed to translocate from cytoplasm to nucleus during the transition from G0–G1 and S phases (Lindeman et al., 1997; Verona et al., 1997). E2F proteins are subject to a variety of post-translational modifications (summarized in Fig. 2.2). E2F1 is activated by acetylation in response to DNA damage, a change that appears to stabilize the protein (Martínez-Balbás et al., 2000; Ianari et al., 2004) and this modification may also help to direct E2F1 to specific subset of target genes (Pediconi et al., 2003). E2F1 is also phosphorylated by kinases that regulate DNA damage responses, such as Chk2 (Checkpoint kinase 2) and ATM (ataxia-telangiectasia mutated)/ATR (ATM and Rad3 related) (Fig. 2.2). Chk2 modifies E2F1 at Ser364, which stabilizes E2F1 (Stevens et al., 2003). Similarly,

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J.-Y. Ji and N.J. Dyson CDK1/2-CycA

CBP/ P/CAF ATM/ATR

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Ser375 Lys120 Lys117 Lys125

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380

283

dimerization domain with Dp

Ser403

transactivation domain (TAD)

409

Thr433

426 437

pRB binding CDK8-CycC binding p14ARF binding; CBP/ P/CAF binding

Fig. 2.2 The structure and regulation of the human E2F1 protein. Binding sites and functional domains that have been mapped within the 437 amino acid residues of E2F1 are shown. Mapped sites of lysine acetylation (Lys117, Lys120, and Lys125) and serine/threonine phosphorylation are indicated. See text for details and references

ATM/ATR was shown to phosphorylate E2F1 at Ser31, a site that is not conserved in E2F2 or E2F3, during the DNA damage response (Lin et al., 2001). Phosphorylation of E2F1 at Ser31 also increases its stability (Lin et al., 2001). These DNA damageinduced modifications are thought to enhance the transcriptional activity of E2F1 and to promote apoptosis through the selective activation of key target genes, such as Apaf1 (Moroni, et al., 2001) and p73 (Urist et al., 2004). Other post-translational modifications are thought to suppress E2F activity. CDK7–CycH, which is the kinase component of the general transcription factor TFIIH complex, can phosphorylate E2F1 at Ser403 and Thr433. Both of these sites are located within the TAD (transactivating domain) of E2F1 (Vandel and Kouzarides, 1999). Phosphorylation of E2F1 on these two sites promotes its degradation, as mutation of these residues to Ala significantly increased the stability of E2F1-TAD (Vandel and Kouzarides, 1999). Several mammalian CDK–cyclin complexes have been reported to directly phosphorylate E2F1 or its dimerization partner DP1, including CDK1/2–CycA (Dynlacht et al., 1994; Krek et al., 1994; Xu et al., 1994; Krek et al., 1995; Dynlacht et al., 1997), CDK1/2–CycB (Dynlacht et al., 1997), CDK7–CycH (Vandel and Kouzarides, 1999), and CDK8–CycC (Morris et al., 2008). These studies paint a complicated picture because CDK–cyclin complexes target different sites on E2F1 (Fig. 2.2) and these modifications can have different effects on E2F activity. For example, phosphorylation of Ser375 of E2F1 by CDK1–CycA has been reported to promote the binding of E2F1 to pRB (Peeper et al., 1995), whereas the more general phosphorylation of E2F1 and/or DP1 by CDK2–CycA disrupts the formation of

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E2F1–DP1 heterodimers and reduces their DNA-binding properties (Dynlacht et al., 1994; Krek et al., 1994; Kitagawa et al., 1995). CycA uses a hydrophobic patch to bind directly to an RXL motif at the N-terminus of the mammalian activator E2Fs in S-phase (Schulman et al., 1998) and E2F1 has been reported to be phosphorylated on at least 6 sites in vivo and 7–9 sites in vitro by CDK2–CycA (Xu et al., 1994). Because different CDKs have similar substrate specificities, it is unclear which kinases are responsible for phosphorylating E2F1 or DP1 in vivo. Biochemical studies show that E2F1 and DP1 each contain at least one site that is phosphorylated by CDK2–CycA but not by CDK2–CycE (Dynlacht et al., 1994) and these results have suggested a model in which the different substrate specificities of CDK2–CycE and CDK2–CycA kinases generate a specific period of E2F1 activity: with E2F1 being activated late in G1 by the phosphorylation of RB-family members by CDK2–CycE and inactivated in S-phase by CDK2–CycA (Fig. 2.1b). The levels of activator E2Fs are limited by proteasome-dependent degradation (Hateboer et al., 1996; Hofmann et al., 1996; Campanero and Flemington, 1997; Marti et al., 1999). E2Fs have been proposed to be targeted for degradation by SCFSkp2 complex in mammals (Marti et al., 1999) or by SCFslmb in Drosophila (Hériché et al., 2003). Degradation of human E2F1 requires the Cul1-E3 ubiquitin ligase (Marti et al., 1999; Ohta and Xiong, 2001). More recent studies in Drosophila have shown that the abrupt degradation of dE2F1 that occurs when cells enter Sphase is mediated by the Cul4Ctd2 E3 ubiquitin ligase and requires PCNA and a PIP (PCNA-interacting protein) motif located near the N terminus of dE2F1 (Shibutani et al., 2008). No PIP boxes have been identified in mammalian activator E2Fs, suggesting that different species may have evolved different mechanisms for the ubiquitylation of activator E2Fs.

2.4 How Do E2Fs Activate Transcription? When activator E2Fs accumulate in late G1 and are released from any inhibitory effects of pRB-family members, they are potent activators of transcription, but precisely how they exert this effect is unclear. E2F proteins have been reported to interact with the general transcription machinery, with various transcription factors, and with proteins that change chromatin structure (Fig. 2.3a). However, at present it is unclear which of these interactions are the most important for the biological functions of E2F in vivo. E2F1 has been shown to physically interact with TATA-box binding protein (TBP) (Hagemeier et al., 1993; Emili and Ingles, 1995; Pearson and Greenblatt, 1997), a subunit of TFIID complex (Hahn, 2004). TBP can recruit general transcription factors, such as TFIIA and TFIIB, to DNA. pRB has been shown to repress E2F-dependent transcription in chromatin-free in vitro transcription assays by blocking recruitment of TFIIA and TFIID, thereby preventing formation of a preinitiation complex (PIC) (Ross et al., 1999). Whether E2F1 directly binds to TBP in vivo is uncertain and many E2F-regulated promoters are TATA-less promoters

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A Co-activators TBP (TFIID) P300 or P/CAF Tip60 HCF Mediator

Functions recruiting and PIC assembly histone and E2F1 acetylation histone acetylation recruiting HCF-containing HMTases recruiting general transcription factors

B

modification of histone tails Co-activator complexes

chromatin in open configuration

DP

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GTFs

DP

E2F

D

target gene expression RNA Pol II

ON

small Mediator complex CDK8 sub-module

target gene expression

p

E2F

DP GTFs

RNA Pol II

OFF

Fig. 2.3 A model for E2F1-mediated transcriptional activation. (a) Transcriptional co-activators that have been linked to E2F1 are listed. Please see text for references. (b–d) Shows a model for E2F1 regulation that is suggested by the discovery of an interaction between E2F1 and the CDK8 module. After phosphorylation and release of pRB, E2F–DP complexes recruit transcriptional co-activators that modify histone tails and render the chromatin in open configuration (b). P/CAF-mediated acetylation of E2F1 may stabilize E2F1 and enhance DNA binding of E2F1–DP1 complexes. E2F1–DP recruitment of small mediator complex, general transcriptional factors, and RNA polymerase II activates the expression of E2F target genes (c). Recruitment of the CDK8 sub-module blocks the re-initiation of E2F-dependent transcription, possibly by CDK8-mediated phosphorylation of E2F1 and disruption of E2F1–DP heterodimers (d)

that do not require TBP (Majello et al., 1998). E2F1 has also been shown to directly interact with p62 subunit of the TFIIH both in vitro (Pearson and Greenblatt, 1997) and in vivo (Vandel and Kouzarides, 1999). TFIIH consists of 10 subunits that form a core complex and a CAK (Cdk-activating kinase) complex, which is composed

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of CDK7, CycH, and an assembly subunit MAT1 (reviewed in Hahn, 2004; Fisher, 2005). A different set of studies have described physical interactions between E2F1 and histone acetyl transferases (HATs). Examples include p300/CBP (CREB (cAMPresponse element-binding) protein-binding protein) (Fry et al., 1999), PCAF (p300/CBP-associated factor) (Lang et al., 2001), and the Tip60 complex (Taubert et al., 2004). These transcriptional cofactor complexes are best known to acetylate histone tails, presumably allowing E2Fs to promote chromatin with an open conformation (Fig. 2.3a, b). E2F1, E2F3a, and E2F4 have also been shown to directly bind to the transcription cofactor HCF1 (Host Cell Factor-1) (Luciano and Wilson, 2003; Knez et al., 2006; Tyagi et al., 2007). HCF1 enhances transcription by recruiting co-activator complexes, such as MLL (mixed-lineage leukemia) and hSet1 histone H3 lysine 4 (K4) methyltransferases (HMTs) (Tyagi et al., 2007) and H4K16 HAT MOF (Dou et al., 2005; Smith et al., 2005). Methylation of histone H3K4 at the promoter regions often positively correlates with active transcription (Barski et al., 2007). Interestingly, HCF1 can also associate with a transcriptional co-repressor complex that contains mSin3A/B and HDAC1/2 (Tyagi et al., 2007; Wysocka et al., 2003), and this property may account for the repressive effect of E2F4 and hypoacetylation of E2F responsive promoters during early G1 phase (Tyagi et al., 2007; Wilson, 2007). A transition from E2F1-bound pRB-mediated or E2F4-mediated repressive complexes during early G1 to E2F1-bound HCF1-mediated MLL family HMTs has been proposed to facilitate the activation of E2F target genes required for proliferation (Tyagi et al., 2007). Despite these extensive studies, there are still many unanswered questions. It is unclear, for example, which of these potential mechanisms of E2F regulation are rate-limiting at key E2F targets. It is possible that the availability of a particular co-activator may be regulated in vivo to limit E2F-dependent cell proliferation. Alternatively, given that E2F proteins can interact with several different cofactors, it is possible that no single co-activator is essential for E2F to drive cell proliferation.

2.5 Drosophila as a Model System to Study E2F Activity In Vivo The studies described above illustrate a complicated pattern of interactions between E2F proteins and CDK activity. In different contexts E2F complexes can either repress or activate the expression of genes encoding cyclins and CDKs, which can either activate or inactivate the transcriptional activity of E2Fs at various points in the cell cycle. Understanding the relationship between these groups of proteins is a difficult problem and one of the limitations of our current models is that they are largely based on biochemical studies and on experiments in which mutant proteins have been over-expressed. Genetic studies in animal models provide a valuable complement to these molecular analyses, helping to identify components of the regulatory network that have

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the greatest impact in vivo. Drosophila has proven to be a powerful model system for studies of E2F. The Drosophila E2F network is less complex than the mammalian E2F family, containing just two E2F homologs, dE2F1 (activator), and dE2F2 (repressor). These two E2Fs provide functions that are analogous to those provided by the mammalian E2Fs (Stevaux and Dyson, 2002; Trimarchi and Lees, 2002). The streamlined nature of the Drosophila E2F/RBF families has enabled clear insights into the organization and action of this complex regulatory network (for a recent review, see van den Heuvel and Dyson, 2008). One drawback of the Drosophila E2F/RBF network is that dE2f1 homozygous mutant animals die early in development (Duronio et al., 1995) and this early lethality represents an obstacle to the analysis of E2F function in the context of animal development. Although one can produce mutant clones using mitotic recombination, such clones are small and are generated randomly thus difficult to follow. This makes the large-scale genetic analysis of mutant clones labor-intensive and unfeasible. This hurdle was circumvented in recent studies by the use of transgenic lines that use RNA interference (RNAi) to selectively target dE2F1 in tissues that are not needed for animal viability (Morris et al., 2008). Lowering the levels of the endogenous dE2F1 protein reduces transcription from dE2F1-dependent promoters and impairs cell proliferation. These changes give visible and reproducible phenotypes in adult eyes and wings that can be modified by additional genetic changes (Morris et al., 2008). dE2f1-RNAi-induced phenotypes have been used to screen mutant collections and the results provide the first glimpse of the spectrum of genes that are ratelimiting for E2F1-dependent cell proliferation in vivo. Unexpectedly, one of the strongest interactions identified in more than 200 dominant modifiers was provided by an allele of cdk8 (Morris et al., 2008). Mutation of cdk8 strongly suppressed dE2f1-RNAi phenotypes in both the eye and the wing. Experiments using cultured Drosophila S2 cells and cdk8 mutant embryos indicate that CDK8 negatively regulates dE2F1-dependent transcription (Morris et al., 2008). In similar genetic tests, mutant alleles of cycA, cycB, cycE, and cdk2 all weakly enhanced dE2f1-RNAi phenotypes, suggesting that they functionally co-operate with dE2F1 (unpublished observations). Such genetic experiments suggest that, of the CDKs and cyclins tested, CDK8 is the clearest negative regulator of E2F1 in vivo.

2.6 CDK8–Cyclin C Negatively Regulates E2F1-Dependent Transcription CDK8 and its regulatory cyclin partner (cyclin C) are subunits of the CDK8 module of the Mediator complex, a transcriptional cofactor that acts as an interface between specific transcription factors and the basal RNA polymerase II (Pol II) transcription apparatus in yeast and human cells (reviewed by Boube et al., 2002; Conaway et al., 2005; Kornberg, 2005; Malik and Roeder, 2005; Myers and Kornberg, 2000; Taatjes et al., 2004; Woychik and Hampsey, 2002).

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Biochemical fractionation experiments indicate that the 30 or more mediator subunits form two distinct types of complexes: a small mediator complex (also known as CRSP, or Positive Cofactor 2, PC2) that activates transcription of Pol II-dependent genes (Malik and Roeder, 2005; Myers and Kornberg, 2000; Näär et al., 2001), and a large mediator complex that generally represses transcription (Mittler et al., 2001). The large mediator complex contains all but one (CRSP70/MED26) of the subunits of the small mediator complex, but also has the CDK8 module, which contains CDK8, CycC, MED12, and MED13 (Näär et al., 2002; Taatjes et al., 2002; Malik et al., 2004). Current models of mediator function suggest that transcription factors bind to promoter sequences and recruit the small mediator complex. This complex recruits Pol II and promotes the initiation of transcription (Malik and Roeder, 2005). With the release of Pol II, many of the general transcription factors are left behind. This “scaffold complex” contains transcription activator, TFIIA, TFIID, TFIIE, TFIIH, and the mediator complex (Yudkovsky et al., 2000) and can potentially recruit free RNA Pol II, TFIIB, and TFIIF, and thereby allowing multiple rounds of transcription reinitiation to occur. Such reinitiation occurs more efficiently than the first round of transcription (Yudkovsky et al., 2000). In general, association of the CDK8 module with the small mediator complex negatively regulates transcription (Malik and Roeder, 2005). Precisely how the suppression occurs is uncertain, and at least four mechanisms have been proposed: (a) CDK8 can phosphorylate the RNA Pol II CTD (on Ser5) and Med2 and disrupt the interaction between RNA Pol II and the mediator complex (Hengartner et al., 1998; Hallberg et al., 2004; Liu et al., 2004; van de Peppel et al., 2005); (b) CDK8 can phosphorylate the CycH subunit of TFIIH, suppressing the CTD kinase activity of CDK7–CycH, effectively preventing the initiation of transcription (Akoulitchev et al., 2000); (c) only the small mediator complex (lacking the CDK8 module) interacts with Pol II and forms a holoenzyme complex, suggesting that the CDK8 submodule may serve as a physical barrier between the small mediator complex and RNA Pol II general transcriptional machinery (Näär et al., 2002; Samuelsen et al., 2003; Bjorklund and Gustafsson, 2005; van de Peppel et al., 2005; Elmlund et al., 2006; Knuesel et al., 2009b); and (d) CDK8 has been shown to directly phosphorylate and inactivate several transcription activators. For example, studies in mammalian cells have shown that CDK8 complex, when recruited by Mastermind, promotes the phosphorylation and ubiquitin-mediated destruction of Notch intracellular domain (Fryer et al., 2004). Phosphorylation by SRB10 (a budding yeast homolog of CDK8) promotes the nuclear export of MSN2 (Chi et al., 2001) or triggers the degradation of GCN4 (Chi et al., 2001). SRB10 has been shown to directly interact with the activating domains of several different transcriptional activators, potentially explaining why activator phosphorylation is often concurrent with transcriptional activation (Ansari et al., 2002; reviewed in Tansey, 2001). It is worth noting that, in some cases, CDK8/SRB10-mediated phosphorylation activates, rather than represses, transcription, such as the cases for Gal4 (Hirst et al., 1999) and Sip4 (Vincent et al., 2001) in yeast. In addition, CDK8 positively regulates p53-mediated transcription of p21 in mammalian cells treated with Nutlin3

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(Donner et al., 2007), a selective small molecule antagonist of MDM2 (Vassilev, 2007). Moreover, CDK8-mediated phosphorylation of Ser10 on histone H3 may also potentiate transcription (Knuesel et al., 2009a). These mediator studies provide potential explanations for the genetic interaction between CDK8 and E2F1 (Fig. 2.3c, d). The fact that CDK8 is a negative regulator of E2F1 suggests that the small mediator complex functions as a co-activator at some dE2F1-dependent genes (Fig. 2.3c). As has been observed in other systems, the CDK8 module may suppress re-initiation and thus limit the burst of transcriptional activation generated by promoter-bound dE2F1. In keeping with this, the effects of depleting dE2F1 were partially suppressed by reduction of CDK8, CycC, MED12, or MED13 (Morris et al., 2008). There are several lines of evidence to suggest that E2F proteins may also be substrates for the CDK8 kinase. Experiments using cultured fly and human cells show that CDK8 associates with dE2F1/E2F1. Moreover, the E2F1 associated with CDK8 becomes phosphorylated when CDK8 immunoprecipitates are incubated in a kinase buffer (Morris et al., 2008). Although CDK2–CycA has been shown to phosphorylate and inhibit the DNA-binding activity of E2F1–DP1 heterodimers, it is conceivable that this mechanism of regulation is normally carried out by CDK8– CycC (Fig. 2.3d). Such a model could explain why dE2F1/E2F1 is stable when bound to RBF1/pRB but is rapidly degraded when it is able to activate transcription. The fact that mutant alleles of cdk8 suppress dE2F1-RNAi phenotypes, while cycA and cdk2 alleles do not, is consistent with the idea that CDK8 down-regulates dE2F1 in vivo. However, we note that genetic interactions between cycA and dE2f1 may be harder to detect if CycA-associated kinases have both positive and negative effects on dE2F1 activity. Future experiments that map the specific residues of E2F1/DP1 that are phosphorylated by CDK8 may help to clarify this issue. The recruitment of a CDK to promoters is the salient feature of current models for CDK8 action, and it is also possible that CDK8–mediated repression of E2F1 may involve other proteins.

2.7 Deregulation of CDK8–CycC in Human Cancers Consistent with the idea that the levels of CDK8–CycC can have a significant impact on cell proliferation, the CDK8 and CCNC (encoding CycC) genes are amplified, mutated, or deleted in various types of human cancers. For example, a point mutation of CDK8 (D189N) has been found in diverse tumor samples (Greenman et al., 2007), but the functional consequence of this mutation is still unknown. The CCNC gene is frequently deleted in acute lymphoblastic leukemia (Li et al., 1996), osteosarcoma (Ohata et al., 2006), and gastric cancer patients (Yang et al., 2007), but is highly expressed in colorectal adenocarcinoma, leukemia, and lymphoma cells (Su et al., 2004). The variety of changes seen in tumor cells may reflect the fact that the mediator complex regulates a vast number of genes, and the role or importance of CDK8–CycC will vary depending on the identity of the transcription factors that

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drive tumor cell proliferation. A striking example of such context-specific effects is provided by the recent finding that CDK8 is an oncoprotein in colorectal cancers (Firestein et al., 2008). CDK8 was identified in a shRNA screen for genes that promote β-catenin-driven transcription and are needed for proliferation of colorectal tumor cell lines. Subsequent experiments demonstrated that the chromosomal region harboring CDK8 gene shows copy number gains in about 47% of colon cancers. Over-expression of wild-type CDK8, but not the kinase dead mutant, was found to promote anchorage-independent growth in untransformed 3T3 cells (Firestein et al., 2008). In an unexpected convergence of events, E2F1 was found to suppress the transcriptional activity of β-catenin (Morris et al., 2008). CDK8 and RB1 both reside on chromosome 13q and copy gains of both genes are found in many colon cancer patients (Gope et al., 1990; Firestein et al., 2008). The amplification of CDK8 seems to be particularly advantageous for colorectal cancers. For reasons that are not yet clear, the components of the CDK8 module enhance β-catenin-driven transcription, and increased levels of CDK8 helps to promote β-catenin activity in colon cancer cells. In addition, elevated CDK8 prevents E2F1 from inhibiting β-catenin. In this cell type, CDK8 appears to co-operate with pRB to sustain β-catenin activity, perhaps explaining why the RB1 gene is never lost, and is occasionally overexpressed, in colorectal cancers (Morris et al., 2008).

2.8 Conclusions and Future Directions The identification of CDK8–CycC as a negative regulator of dE2F1 illustrates the value of unbiased genetic screens. Because of the relative ease and low cost of genetic screens and phenotypic analyses, studies in model systems like Drosophila and Caenorhabditis elegans provide a powerful complement to the molecular and biochemical analyses that traditionally have been carried out in mammalian cells. Genetic screens identify components that have a significant functional impact on a phenotype regardless of the underlying mechanism. Such studies have the advantage that they can discover important and unsuspected connections between genes and processes. Genetic interactions are, however, only a starting point and need to be complemented by biochemical studies that characterize the underlying molecular events. The genetic interaction between CDK8–CycC and E2F1 raises a new series of questions for future studies. This interaction suggests that the small mediator complex plays an important role in E2F-mediated activation. It is uncertain, however, whether the mediator complex acts at all E2F targets or just a subset. Do activator E2Fs interact with a particular component of the mediator complex, and does E2F1 recruit mediator, or does mediator recruit E2F1? A second issue is the significance of CDK8–CycC phosphorylation of E2F1. The genetic data and the evidence that CDK8–CycC can phosphorylate E2F1 raise the possibility that CDK8–CycC may be the key CDK activity that is responsible for the attenuation of activator E2Fs. Further work is clearly necessary to determine which of the many CDKs that

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can phosphorylate E2F1/DP1 dimers are actually responsible for this regulation in vivo. Answering this question will provide a better understanding of how the E2F transcriptional program is switched on and off, and how it can be coupled to, or uncoupled from, the CDK cycle. In different contexts, deregulated E2F1 can promote cell proliferation or induce cell death. Knowing how to tip the balance between these outcomes toward cell death may have many applications in the treatment of cancer cells. The fact that CDK8 is an oncoprotein in colorectal cancers has raised the possibility that CDK8 inhibitors might have utility in cancer cells that harbor CDK8 amplification (Firestein et al., 2008). Since CDK8 is also an inhibitor of E2F1, CDK8 antagonists may generally enhance E2F1 activity. In combination with appropriate pro-apoptotic stimuli, CDK8 inhibitors may be useful for stimulating E2F1-induced apoptosis in many different types of cancer cells. Acknowledgments We thank Drs. Erick Morris, Gerold Schubiger, and Fajun Yang for critical comments on this review. J.Y.J. is supported by a post-doctoral fellowship from the MGH Fund for Medical Discovery. This work was supported by a grant from the NIH (RO1 GM53203). N.J.D. is the MGH Saltonstall Foundation Scholar.

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Chapter 3

Regulation of Pre-RC Assembly: A Complex Symphony Orchestrated by CDKs A. Kathleen McClendon, Jeffry L. Dean, and Erik S. Knudsen

Abstract DNA replication is a tightly regulated process that has critical implications for human cancers. Pre-replication (pre-RC) assembly is required for initiation of DNA replication, and virtually all components of the pre-RC are regulated by complex mechanisms that center around the activity of cyclin-dependent kinases (CDKs). CDK/cyclin complexes both positively and negatively regulate pre-RC components, including their expression, activity, stabilization, and degradation. Together, these complex mechanisms orchestrate DNA replication and cell cycle progression in a manner by which genetic material is duplicated only once. Deregulated pre-RC activity has been shown to result in DNA re-replication and genomic instability, a hallmark of cancer. Furthermore, deregulation of pre-RC components and CDK/cyclins is observed in a multitude of human cancers. Thus, regulation of pre-RC assembly is a critical facet of normal cell biology that has profound implications related to cancer etiology and diagnosis.

3.1 The Pre-replication Complex DNA replication is a tightly regulated process that requires coordinated assembly of protein complexes at distinct periods of the cell cycle. Sites of DNA replication are determined by binding of origin replication complexes (ORCs) (Bell and Stillman, 1992; Bell and Dutta, 2002; Chesnokov, 2007). The ORC is conserved from yeast to humans and is believed to remain bound to DNA throughout the cell cycle (Bell and Dutta, 2002; Chesnokov, 2007). At the end of mitosis, replication control mechanisms are “reset” when Cdc6 and Cdt1 are recruited to ORCs (Bell and Dutta, 2002; Liang et al., 1995; Coleman et al., 1996; Cocker et al., 1996; Nishitani et al., 2000; Maiorano et al., 2000). Binding of Cdc6 and Cdt1 to ORCs results in the subsequent E.S. Knudsen (B) Department of Cancer Biology, Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, PA 19107, USA e-mail: [email protected]


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recruitment of the mini-chromosome maintenance (Mcm) complex (Mcm2-7) during late M-phase and G1 (Bell and Dutta, 2002; Tye, 1999; Maiorano et al., 2006). Recruitment of Mcm2-7 completes the “pre-replication complex” (pre-RC), the critical substrate upon which replication initiates. Formation of the pre-RC is essential for replication initiation and ultimately progression into S phase (Bell and Dutta, 2002; Lau et al., 2007). At the beginning of S phase, the pre-RC is converted to the initiation complex (IC) through the recruitment of factors such as cdc45 and Mcm10 (Bell and Dutta, 2002; Maiorano et al., 2006; Merchant et al., 1997; Mimura and Takisawa, 1998; Sawyer et al., 2004). IC formation results in the subsequent recruitment of components of the “replisome” (RFC, RPA, and PCNA), the active complex responsible for DNA replication (Waga and Stillman, 1998; Diffley and Labib, 2002; Mendez and Stillman, 2003). While the formation of all protein complexes leading to DNA replication is highly controlled, regulation of pre-RC assembly for replication licensing is particularly crucial for maintaining genome integrity. Multiple ORCs form along the genome to ensure efficient replication of DNA at each cell division (Bell and Dutta, 2002; Chesnokov, 2007; DePamphilis, 2005). Additionally, pre-RC formation is tightly regulated to ensure that each of these regions is replicated only once (Bell and Dutta, 2002; Lau et al., 2007). Alterations in the control of replication licensing can lead to loss or duplication of genetic material, predisposing cells to mutations and chromosomal instability, factors that have been implicated in promoting tumorigenesis (Lau et al., 2007; Storchova and Pellman, 2004; Pellman, 2007). A crucial mechanism by which cells regulate pre-RC formation and insure faithful replication of the genome is manifested via the complex actions of cyclin-dependent kinases (CDKs) (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009).

3.2 Cyclin-Dependent Kinases (CDKs) and General Cell Cycle Control Cyclin-dependent kinases (CDKs) are serine/threonine kinases that play critical roles in cell cycle progression (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009). As eukaryotic cells progress through the cell cycle, CDK activity is stimulated via association with cyclins. Mammalian cells utilize CDK1, CDK2, CDK4, and CDK6 for cell cycle progression (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009). CDK2, CDK4, and CDK6 are considered to be the interphase CDKs, and while these proteins are required for proliferation of specific cell types, they are not essential genes in mice (Malumbres et al., 2004; Kozar et al., 2004; Ortega et al., 2003; Berthet et al., 2003; Barriere et al., 2007). Furthermore, while CDK2/4/6 have been shown to act cooperatively to modulate cell cycle progression, some functional redundancy has been indicated in both cells and mice lacking one or more of the interphase CDKs (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009; Malumbres et al., 2004; Kozar et al., 2004; Barriere et al., 2007; Sherr and Roberts, 2004). In contrast, CDK1 is involved in driving mitosis and is essential for cell

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division (Santamaria et al., 2007). Each of these CDKs is tightly regulated, and modulation of CDK activation is critical in determining progression versus arrest of the cell cycle (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009). The general model of cell cycle regulation by CDKs begins with mitogenic signaling to stimulate accumulation of D-type cyclins, which form complexes with and activate CDK4 and CDK6 at the onset of G1 (Malumbres and Barbacid, 2001). Cyclin D–CDK4/6 complexes function to phosphorylate and inactivate the retinoblastoma tumor suppressor (RB) or related pocket proteins (p107 and p130) (Mittnacht, 1998; Harbour et al., 1999). Inactivation of RB relieves repression of the E2F family of transcription factors, allowing for the expression of genes essential for DNA replication and cell cycle progression (Cobrinik, 2005; Markey et al., 2002). This initial step in cell cycle progression is antagonized by a family of CDK inhibitors (CKIs) including the INK4 proteins (p16INK4a , p15INK4b , p18INK4c , and p19INK4d ). These CKIs specifically inhibit CDK4 and CDK6 by directly disrupting association to D-type cyclins, ultimately promoting RB-dependent cell cycle arrest (Sherr and Roberts, 1999; Roussel, 1999; Sherr and McCormick, 2002). Alleviation of RB-mediated transcriptional repression allows for the expression of genes such as E-type and A-type cyclins (Cobrinik, 2005; Markey et al., 2002). E-type cyclins complex with and activate CDK2 to promote further phosphorylation and complete inactivation of RB, thus allowing for expression of genes required for driving DNA replication and the G1 to S phase transition (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009; Harbour et al., 1999; Hochegger et al., 2008). Much like the negative regulation of cyclin D–CDK4/6 complexes by INK4 proteins, cyclin–CDK2 complexes are inhibited by a second class of CKIs made up of the Cip/Kip family of proteins (including p21Cip1 and p27Kip1 ). These CKIs bind both CDKs and cyclins and inhibit the functions of both cyclin A- and cyclin E-CDK2 complexes (Sherr and Roberts, 1999; Besson et al., 2008). Interestingly, Cip/Kip proteins also function as assembly factors for cyclin D–CDK complex formation. Inhibition of cyclin D/CDK4 by p16INK4a results in the displacement of p27Kip1 , which allows for the inhibition of cyclin E/CDK by p27Kip1 and subsequent G1 arrest (Sherr and Roberts, 1999; Besson et al., 2008). Upon successful initiation and progression of DNA replication, cyclin B1–CDK1 complexes form to drive mitosis (Malumbres and Barbacid, 2009; Hochegger et al., 2008; Malumbres and Barbacid, 2005). Final inactivation and degradation of cyclin B1, as well as various other factors, via the anaphase-promoting complex (APC/C) results in mitotic exit (Acquaviva and Pines, 2006; Peters, 2006). Furthermore, degradation of cyclin B1 effectively eliminates CDK activity, ultimately allowing for new origin licensing and progression of cells into the next cell cycle (Acquaviva and Pines, 2006; Peters, 2006). CDKs enforce cell cycle order via a complex array of protein interactions and mechanisms, all of which dramatically impact pre-RC assembly and replication initiation. These mechanisms involve both positive regulation of pre-RC formation in G1 phase of the cell cycle and negative regulation of pre-RCs to prevent rereplication through S phase and mitosis (Malumbres and Barbacid, 2009; Doonan

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Origin Fig. 3.1 Complex regulation of pre-RC assembly by CDK/cyclins. (a) Mitogenic signaling stimulates CDK4/6–cyclin D activity at the G0–G1 transition of the cell cycle. CDK4/6–cyclin D complexes inactivate the retinoblastoma tumor suppressor (RB) to relieve repression of the E2F family of transcription factors, allowing for the expression of genes essential for pre-RC assembly. CDK4/6–cyclin D function is antagonized by the INK4 family of CDK inhibitors. (b) CDK2– cyclin E complexes play dual roles in pre-RC regulation in G1. First, CDK2–cyclin E complexes function to promote further inactivation of RB, thus allowing for expression of genes required for driving DNA replication and the G1 to S phase transition. Second, CDK2–cyclin E complexes phosphorylate and stabilize Cdc6. Interestingly, cyclin E also associates with Cdt1 and the Mcm

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and Kitsios, 2009). Both aspects of CDK regulation are critical for maintaining genomic stability and promoting faithful replication of the genome.

3.3 Positive Impact of CDKs on Pre-RC Assembly (G0–G1 Phase) CDKs are involved in the regulation of pre-RC assembly from the onset of cell cycle initiation (Fig. 3.1). Activation of the cyclin D–CDK4/6 complex is the first step in exiting G0 in response to mitogenic signaling (Malumbres and Barbacid, 2001). This kinase complex functions to phosphorylate and partially inactivate the RB tumor suppressor, allowing for the release of E2F transcription factors and subsequent transcription of E2F target genes, including Cdc6, Cdt1, and Mcm2-7 (Mittnacht, 1998; Harbour et al., 1999; Cobrinik, 2005; Markey et al., 2002). Transcription of Cdc6, Cdt1, and Mcm2-7, and the accumulation of these proteins at ORCs, is required for formation of a functional pre-replication complex (Bell and Dutta, 2002; Liang et al., 1995; Coleman et al., 1996; Cocker et al., 1996; Nishitani et al., 2000; Maiorano et al., 2000; Lau et al., 2007). In contrast, activation of the RB pathway and subsequent G1 arrest is characterized by transcriptional repression of pre-RC components (Markey et al., 2002; Braden et al., 2006, 2008). Furthermore, targeted inhibition of CDK4/6 results in the down-regulation of pre-RC components (Braden et al., 2008). Thus, pre-RC assembly is initially regulated by the ability of cyclin D–CDK4/6 to release transcriptional repression of pre-RC components through inactivation of RB, an event that is considered critically important for cells exiting quiescence. Once pre-RC components have been transcribed and begin assembling at ORCs, a second interphase cyclin–CDK complex, cyclin E–CDK2, is responsible for facilitating and maintaining pre-RC formation for subsequent replication complex assembly (Malumbres and Barbacid, 2009; Harbour et al., 1999; Lundberg and Weinberg, 1998). Cyclin E–CDK2 complexes display some functional redundancy with cyclin D–CDK4/6 complexes in their ability to phosphorylate and inactivate RB, ultimately promoting transcription of S phase proteins that bind the pre-RC and initiate DNA replication (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009; Harbour et al., 1999; Hochegger et al., 2008). Additionally, cyclin E–CDK2

Fig. 3.1 (continued) to promote pre-RC assembly independent of CDKs. Additionally, geminin is degraded at the onset of G1 to allow Cdt1 association at chromatin. (c) Recruitment of Cdc45 and Mcm10 in S phase promotes loading of the replisome and initiation of replication. At the onset of DNA replication, CDK2–cyclin E/A complexes take on a negative regulatory role by phosphorylating pre-RC components, resulting in their dissociation from chromatin and subsequent degradation. (d) As cells transition through the G2–M phases of the cell cycle, the chromatin has been stripped of all pre-RC components. The cell is now “reset” in G0 for another round of cell cycle

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has been shown to directly phosphorylate Cdc6, thereby preventing Cdc6 degradation via the anaphase-promoting complex/cyclosome (APC/C), which remains active until late G1 phase (Mailand and Diffley, 2005). This cyclin E–CDK2mediated stabilization is specific to Cdc6, as Cdt1 is not a target of the APC/C (Mailand and Diffley, 2005; Fujita, 2006). Thus, stabilization of Cdc6 by cyclin E–CDK2 ensures that the critical components of the pre-RC are not only assembled for replication licensing, but are also maintained during G1 progression. In addition to the kinase-dependent function of cyclin E, this interphase cyclin functions to promote replication complex assembly independent of CDKs. Recent studies have shown that cyclin E can interact with replication complex components, Cdt1 and Mcm, to promote loading of Mcm proteins to ORCs (Geng et al., 2007). Additionally, while CDK2 null fibroblasts were shown to undergo normal cell cycles (Sherr and Roberts, 2004), cyclin E null cells fail to progress from quiescence to S phase due to a defect in Mcm protein loading (Geng et al., 2003). A cyclin E mutant that fails to activate CDKs displayed the same interactions with pre-RCs as the wild-type cyclin E protein and was able to rescue the G0 to S phase transition defect in cyclin E null fibroblasts (Geng et al., 2007). Thus, cyclin E plays dual roles in promoting replication complex assembly through kinase-dependent and kinaseindependent mechanisms, both of which are critical for progression of cells from a quiescent state. The positive regulation of pre-RC assembly and subsequent replication initiation is modulated by interphase cyclins and CDKs via a complex array of mechanisms. While all of these mechanisms are essential for promoting DNA replication and cell cycle progression, there must exist counterbalances to ensure that DNA replication is occurring faithfully and that cell cycle progression is not occurring under inappropriate conditions. Interestingly, CDKs are critical modulators of the negative regulation of pre-RC assembly as well.

3.4 Negative Impact of CDKs on Pre-RC Assembly (S–M Phase) While elevated levels of G1 CDKs function to activate pre-RC assembly and replication initiation in G1 phase, this same cellular pool of CDKs is also required to prevent re-initiation during S, G2, and M phases (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009). One method of negatively regulating pre-RCs is through inhibition of Cdc6. As discussed above, phosphorylation on specific residues of Cdc6 by CDK2 results in stabilization of Cdc6 in G1 phase (Mailand and Diffley, 2005). However, phosphorylation at distinct sites of the protein has been observed to result in Cdc6 dissociation from ORCs, nuclear export, and proteosomal degradation (Petersen et al., 1999; Mendez and Stillman, 2000; Petersen et al., 2000; Borlado and Mendez, 2008). The importance of Cdc6 nuclear export is somewhat controversial, as significant amounts of Cdc6 have also been observed to remain bound to chromatin throughout S and G2 phases (Mendez and Stillman, 2000; Borlado and Mendez, 2008; Coverley et al., 2000; Fujita et al., 1999). However, CDK-mediated

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phosphorylation of ORC, Cdt1, and Mcm proteins has also been implicated in the dissociation of pre-RCs, indicating that the phosphorylation of various components of the pre-RC by CDKs at least partially contributes to the inhibition of pre-RC assembly and replication initiation (Lau et al., 2007; Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009). In addition to the CDK-dependent dissociation and degradation of Cdc6, there are two main modes of negative regulation of Cdt1 in mammalian cells. First, Cdt1 is targeted for ubiquitin-mediated proteolysis via interactions with two distinct E3 ubiquitin ligase complexes, CUL4-DDB1CDT2 and SCFSkp2 . CUL4-DDB1CDT2 targets Cdt1 for degradation once Cdt1 is bound to PCNA on chromatin (Fujita, 2006; Zhong et al., 2003; Hu et al., 2004; Senga et al., 2006). This degradation event occurs in S phase and is stimulated by export/degradation of cyclin D1 (Aggarwal et al., 2007). Unlike CUL4-DDB1CDT2 , SCFSkp2 -mediated degradation of Cdt1 is dependent on the phosphorylation of Cdt1 by CDKs and occurs throughout the cell cycle (Liu et al., 2004; Li et al., 2003). Additionally, both cyclin A–CDK2 and cyclin A–CDK1 have been implicated in the phosphorylation and subsequent degradation of Cdt1 (Fujita, 2006; Liu et al., 2004; Sugimoto et al., 2004). The second mechanism of negative regulation of Cdt1 consists of direct binding and inhibition of Cdt1 by geminin. Geminin was originally identified as an APC/C substrate that inhibits pre-RC formation via prevention of Mcm complex loading (McGarry and Kirschner, 1998). Geminin is degraded by the APC/C complex at the metaphase/anaphase transition of mitosis, allowing for Cdt1 accumulation and ORC binding in the next round of the cell cycle (McGarry and Kirschner, 1998; Wohlschlegel et al., 2000; Tada et al., 2001). More recently, geminin has been identified as a target of E2F transcription that accumulates as cells enter S phase (Yoshida and Inoue, 2004). Thus, regulation of Cdt1 by geminin is ultimately controlled by upstream CDKs and the release of RB-mediated transcriptional repression that is required for geminin expression. Negative regulation of pre-RC components is essential for preventing DNA rereplication. As DNA re-replication has obvious implications in genome instability and human cancers, it is not surprising that cells have evolved to utilize multiple mechanisms to control pre-RC assembly and replication initiation.

3.5 Perturbations of Pre-RC Assembly and Cancer 3.5.1 Functional Effects of Deregulated Pre-RC Assembly Over-expression of pre-RC components (i.e., Cdc6 and Cdt1) and loss of preRC regulatory proteins (i.e., geminin) result in deregulated DNA replication and cell cycle progression, which can have significant impact on genome stability. For example, over-expression of pre-RC components has been implicated in DNA re-replication and chromosomal instability in eukaryotic cells (Lau et al., 2007; Tatsumi et al., 2006; Vaziri et al., 2003; Karakaidos et al., 2004). Additionally, overexpression of Cdc6 and Cdt1 has been shown to promote cellular transformation and

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tumorigenesis in mouse models either alone or in cooperation with known oncogenic lesions (Lau et al., 2007; Seo et al., 2005; Gonzalez et al., 2006; Arentson et al., 2002; Blow and Gillespie, 2008). Furthermore, depletion of geminin has also been shown to result in DNA re-replication and genomic instability similar to overexpression of Cdt1 (Zhu et al., 2004; Melixetian et al., 2004; Saxena and Dutta, 2005). Thus, it is not surprising that deregulation of pre-RC components and pre-RC regulatory factors is frequently observed in various types of human cancer.

3.5.2 Deregulation of Pre-RC Components in Cancer While mutations of pre-RC components have not been identified in human cancers, deregulation of these proteins has been observed in various model systems of human cancers and tumor specimens. For example, over-expression of Cdt1 and/or Cdc6 has been observed in a multitude of tumors and tumor-derived cell lines (Lau et al., 2007; Karakaidos et al., 2004; Williams et al., 1998; Murphy et al., 2005; Ohta et al., 2001; Bravou et al., 2005). Additionally, deregulation of Mcm2-7 has also been observed in various types of human malignancies, including breast, cervical, and esophageal cancers (Lau et al., 2007; Williams et al., 1998; Going et al., 2002; Shetty et al., 2005). Furthermore, since deregulation of pre-RC components induces characteristics that are considered to be critical for tumorigenesis, expression signatures of pre-RC proteins have been proposed as potential biomarkers for cancer diagnosis (Lau et al., 2007; Williams and Stoeber, 2007; Gonzalez et al., 2005). Ultimately, deregulation of pre-RC assembly is driven by aberrations in CDK/cyclin signaling (Malumbres and Barbacid, 2009; Doonan and Kitsios, 2009). Thus, mutations and deregulated activities of pre-RC regulatory CDK/cyclin proteins are a frequent occurrence in human cancers. For example, deregulation of CDK4/6, via direct mutation within the genes, loss of p16 or over-expression of D-type cyclins, has been implicated in a variety of tumors (Malumbres and Barbacid, 2009; Ortega et al., 2002; Knudsen and Knudsen, 2008). Additionally, while mutations of CDK2 have not yet been observed in human cancers, deregulation of cyclin A and cyclin E via various mechanisms has been observed in a multitude of cancers and is associated with poor prognosis (Malumbres and Barbacid, 2009; Yasmeen et al., 2003). (Kitahara et al., 1995) Each of these alterations in cyclin/CDK activity ultimately results in enhanced pre-RC assembly and subsequent DNA replication, processes that have proven to be critical for tumorigenesis.

3.6 Conclusions Regulation of pre-RC assembly and replication initiation are critical facets of normal cell biology that have implications for human cancers. Cyclin/CDK complexes maintain tight control of over these processes via a complex array of mechanisms.

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Perturbations of this regulation can lead to abnormal DNA re-replication, which can ultimately promote genome instability and cancer. Thus, it is not surprising that deregulation of either pre-RC components themselves or the cyclin/CDK signaling that controls pre-RC assembly is a common occurrence in many tumors and has implications as both biomarkers for cancer diagnosis and targets for cancer therapies.

3.7 Future Directions Since CDKs play such a prominent role in controlling the cell cycle progression, these proteins have become key therapeutic targets for not only cancer, but numerous other diseases as well (Lee and Sicinski, 2006). However, like many targeted therapies, not all CDK inhibitors have proven to be effective in clinical trials (Malumbres and Barbacid, 2009). It is likely that in order to efficiently target CDKs and/or cyclins for treatment of diseases such as cancer, it will be critical to elucidate the specific requirements for distinct CDKs and cyclins in different cell types and tissues. Furthermore, tumors are frequently characterized by a spectrum of mutations that could interfere with the desired outcome of CDK/cyclin inhibition. Thus, understanding the dependencies of specific CDKs and cyclins in distinct tissues, as well as potential cooperating or impeding lesions, will be important for designing treatment regimens involving specific CDK inhibition alone or in combination with other established therapies.

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Part II

Proliferation Under Duress

Chapter 4

Mitotic Checkpoint and Chromosome Instability in Cancer Haomin Huang and Timothy J. Yen

Abstract Chromosome instability (CIN) results in aneuploidy that may promote tumorigenesis and chemoresistance. Chromosome missegregation allows cells to rapidly change gene expression patterns on a global scale and provides a mechanism that promotes genetic and biochemical diversity that is used by cells to achieve a transformed state or to survive suboptimal growth conditions. The mitotic checkpoint is a fail-safe mechanism that monitors the fidelity of the microtubule attachments to the kinetochore, a macromolecular structure that is situated at centromeres and essential for chromosome segregation. Over the past two decades many kinetochore proteins have been identified and characterized in detail. Among these are the mitotic checkpoint proteins that monitor the integrity of kinetochore:microtubule attachments and initiate and propagate the “wait anaphase” signal that inactivates the anaphase-promoting complex/cyclosome (APC/C). The CIN phenotype of many cancer cell lines may be attributed to the lack of a robust mitotic checkpoint as these cells fail to arrest in response to microtubule inhibitors such as taxol and nocodazole. Interestingly, mutations in the essential mitotic checkpoint genes are rare. The checkpoint defects exhibited by CIN cells may be due to a combination of factors that affect the mechanical efficiency of the spindle, as well as the threshold level of “wait anaphase” signal that is required to inhibit the global pool of the APC/C. In this chapter, we will review our current understanding of CIN and the mitotic checkpoint and discuss the recent views of the relationships among the mitotic checkpoint, CIN, tumorigenesis, and chemoresistance.

H. Huang (B) Fox Chase Cancer Center, Philadelphia, PA 19111, USA e-mail: [email protected]


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4.1 Chromosome Instability (CIN) 4.1.1 Chromosome Missegregation, Aneuploidy, and CIN Genetic instability is often considered to be the driving force of tumorigenesis. At the nucleotide level, germline mutations in several different mismatch repair (MMR) genes were found to result in microsatellite instability (MIN) in hereditary nonpolyposis colon cancer (HNPCC) (Fishel et al., 1993; Leach et al., 1993; Parsons et al., 1993; Shibata et al., 1994; Thibodeau et al., 1993; Umar et al., 1994). Even though MIN is strongly linked to the development of HNPCC, it only represents 15% of solid tumors (Tomonaga, 2008; Weaver and Cleveland, 2006). In contrast to MIN cancer cells that are generally neardiploid (Eshleman et al., 1998; Jallepalli and Lengauer, 2001), CIN (chromosome instability) cancers exhibit increased rates of chromosome loss and gains at each division. CIN cells therefore exhibit dramatic differences in karyotype within a population (Lengauer et al., 1997, 1998). Interestingly, CIN tumors generally do not exhibit microsatellite instability (Jallepalli and Lengauer, 2001). Aneuploidy is clearly an outcome of CIN but the relationship is complex. Aneuploid tumor cells are not necessarily classified as CIN if they do not gain and lose chromosomes at each division (Kops et al., 2005b). Aneuploidy may be the result of a one-time event that provided the cell with a growth advantage. Once the desired traits are achieved, the ploidy is stably maintained in subsequent generations (unlike CIN). The CIN phenotype is also not simply caused by aneuploidy. For example, addition of an extra copy of chromosome 3 into the near-diploid HCT116 MIN cell line, or its fusion with another MIN cell line, did not result in a CIN progeny cells (Lengauer et al., 1997). By contrast, a fusion between a MIN (DLD1) and a CIN (HT29) tumor cells produced cells that retained the CIN phenotype (Lengauer et al., 1997). These findings suggest that the determinants for CIN may be considered as dominant gain-of-function mutations. A recent report demonstrated that when diploid non-CIN cell lines, HCT116 and RPE1, were exposed to a single treatment of anti-mitotic drugs that increased the rate of chromosome missegregation, the progeny cells exhibited a transient CIN phenotype for 20–30 generations (Thompson and Compton, 2008). The number of cells with deviant chromosome numbers eventually declined, and the ploidy returned to near basal levels and was stabilized. Not surprisingly, when a CIN cell line (HT29) was subjected to the identical protocol, it maintained its CIN phenotype. Collectively, these findings indicate that missegregation of one or multiple chromosomes is not sufficient to convert cells to the CIN phenotype and, most importantly, cannot convert near-diploid cells to aneuploid cells. The latter point is consistent with the general notion that cells do not tolerate chromosome imbalance and that cells must first overcome this barrier before becoming CIN (Yuen and Desai, 2008).

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4.1.2 What Are the Defects That Result in Chromosome Missegregation in CIN Cells? Research into mechanisms of chromosome segregation over the past 20 years has identified many genes that if lost or mutated will result in chromosome missegregation. Defects in the mitotic checkpoint can result in aneuploidy as cells divide regardless of whether their chromosomes are properly attached to the spindle or not (Chan et al., 2000, 1999; Gorbsky et al., 1998; Huang et al., 2008a; Jelluma et al., 2008a, b; Jones et al., 2005; Kops et al., 2005a; Logarinho et al., 2008; Tang et al., 2004; Taylor and McKeon, 1997). Mutations in proteins that specify proper attachment of the spindle microtubules to the kinetochore, and those important for sister chromatid cohesion, will also cause chromosome missegregation (Bakhoum et al., 2009; Cimini et al., 2001; DeLuca et al., 2006; Feng et al., 2006; Huang et al., 2007, 2008b; Jallepalli et al., 2001; Kaplan et al., 2001; Pfleghaar et al., 2005; Vong et al., 2005). Finally, defects during cytokinesis will produce polyploid cells (Ganem et al., 2007; Storchova and Pellman, 2004). There are literally hundreds of proteins essential for accurate chromosome segregation, and the loss of any single protein would result in aneuploidy. Indeed, mutations in genes that are essential for mitotic checkpoint signaling, as well as spindle functions have been identified in some tumor cells. However, no single gene mutation has been found to dominate and account for the high frequency of aneuploid cancer cells. Efforts to sequence the genomes of cancer cells should eventually lead to an exhaustive screen for mutations in genes that are known to be critical for mitosis.

4.2 The Mitotic Checkpoint The mitotic, or spindle assembly, checkpoint is a fail-safe mechanism that monitors the fidelity of chromosome segregation in space and time (Chan et al., 2005; Chan and Yen, 2003; Liu et al., 2003; May and Hardwick, 2006; Musacchio and Salmon, 2007; Yen and Kao, 2005). The molecular details of this checkpoint pathway have been extensively reviewed so we will cite the features that are most relevant to this chapter. Until all chromosomes are properly aligned at the spindle equator, the mitotic checkpoint inhibits the anaphase-promoting complex/cyclosome (APC/C), an E3 ubiquitin ligase that promotes the degradation of substrates that prevent cells from entering anaphase (King et al., 1995; Peters et al., 1994; Sudakin et al., 2001, 1995). The core components of the mitotic checkpoint are the Bub1, BubR1, Mps1 kinases, along with Bub3, Mad1, and Mad2 (Cahill et al., 1998; Chan et al., 1999; Chen et al., 1996; Davenport et al., 1999; Hardwick and Murray, 1995; Hoyt et al., 1991; Li and Murray, 1991; Li and Benezra, 1996; Roberts et al., 1994; Sudakin et al., 2001; Taylor et al., 1998; Taylor and McKeon, 1997; Weiss and Winey, 1996; Winey et al., 1991). All of these proteins seem to act at multiple steps of the signaling pathway that includes monitoring the integrity of the kinetochore–microtubule

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attachments, generating, and amplifying the “wait anaphase signal” that then propagates throughout the cell to inhibit the APC/C (Campbell and Gorbsky, 1995; Li and Nicklas, 1995; McIntosh, 1991; Nicklas et al., 1995; Rieder et al., 1995, 1994). Loss of any single component will cause cells to exit mitosis regardless of whether their chromosomes establish proper bipolar attachments and achieve alignment at the spindle equator. The mitotic checkpoint appears to monitor two aspects of kinetochore– microtubule attachments. The Mad1 and Mad2 checkpoint proteins are thought to monitor the microtubule occupancy at the kinetochore as they are recruited to kinetochores that lack microtubule attachments (Chen et al., 1996; Skoufias et al., 2001; Waters et al., 1998). When the kinetochore is saturated with microtubules (mammalian kinetochores can bind approximately 20–25 microtubules), the amounts of Mad1 and Mad2 on kinetochores are reduced by nearly 100-fold (Hoffman et al., 2001). The dissociation of Mad1 and Mad2 from kinetochores is not sufficient to silence the checkpoint until the functionality of the microtubule attachment is satisfied. Productive microtubule attachments occur as a result of their end-on attachments to kinetochores. This geometry appears to be mediated by a molecular sleeve that is composed of the Nuf2/Ndc80 complex (Cheeseman et al., 2006; DeLuca et al., 2005; McCleland et al., 2004). Upon end-on attachment, the intrinsic dynamic properties of the microtubule in conjunction with various microtubulebinding proteins at the kinetochore generate opposing poleward-directed forces that are critical for generating tension between the sister kinetochores. Owing to the random nature by which microtubules encounter kinetochores, non-end-on attachments do occur that are unable to generate proper levels of kinetochore tension. If these defective attachments go undetected, their persistence can be a source of lagging chromosomes when cells exit mitosis (Cimini et al., 2001). In addition to monitoring microtubule occupancy, kinetochores possess an elaborate error correction system that will sever non-productive attachments to allow for new rounds of attachment (Pinsky et al., 2006; Tanaka et al., 2002). Aurora B kinase is a core component of the error system that is thought to regulate the severing activity of the microtubule depolymerase, MCAK (mitotic centromere-associated kinesin) (Andrews et al., 2004; Lampson et al., 2004; Lan et al., 2004). Aurora B kinase has been reported to specify the differential sensitivity of the mitotic checkpoint to the microtubule inhibitors, taxol and nocodazole (Ditchfield et al., 2003). Microtubules in taxol-treated cells are able to establish kinetochore attachments but fail to generate tension due to the loss of dynamicity of the microtubule (Waters et al., 1998). Taxol induces a mitotic delay that is dependent on Aurora B. By contrast, the mitotic arrest induced with nocodazole, a drug that blocks microtubule polymerization that leads to unoccupied kinetochores, was less dependent on Aurora B (Ditchfield et al., 2003; Famulski and Chan, 2007). Given that Aurora B promotes the detachment of microtubules from kinetochores, it remains possible that it indirectly activates the checkpoint by transiently creating unoccupied kinetochores. This may explain why in taxol-arrested cells, Mad2 can still be detected at a few kinetochores even though the majority of the kinetochores are attached to microtubules and lack detectable Mad2 (Waters et al., 1998).

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Although all of the checkpoint proteins are localized at kinetochores, their mechanistic roles remain unresolved. It is possible that each protein or subsets of proteins monitor the activities of discrete sets of microtubule-binding proteins and motors within the kinetochore. Alternatively, the combined activities of all the different microtubule-binding proteins that act on a single kinetochore are monitored by a single complex of checkpoint proteins. Regardless of the mechanism, some of the checkpoint proteins must act as mechanosensors that convert microtubule attachment status to a diffusible signal that is propagated from a defective kinetochore. The amplification process that allows a localized defect to alter the global biochemical state of the cell may be mediated in part by the rapid turnover of checkpoint proteins at kinetochores. The proteins that are released from a defective kinetochore may assume an activated state that is capable of inhibiting the APC/C (May and Hardwick, 2006; Musacchio and Salmon, 2007). On the other hand, the amplification might also be explained by a conventional signal transduction cascade that is mediated by the various mitotic checkpoint kinases that reside at kinetochores (Chen, 2004; Ditchfield et al., 2003; Elowe et al., 2007; Huang et al., 2008a; Huang and Yen, 2009; King et al., 2007; Matsumura et al., 2007; Qi et al., 2006; Rancati et al., 2005; Wong and Fang, 2007). Early studies in PtK1 cells showed that a single unattached kinetochore is sufficient to delay mitotic exit (Rieder et al., 1994). More recent studies of a variety of human cell lines have shown that the length of the delay can vary among different cell lines (Gascoigne and Taylor, 2008; Rieder and Maiato, 2004) and perhaps due to the number of defective kinetochores in a cell (Feng et al., 2006; Huang et al., 2007, 2008a). These observations suggest that the checkpoint signaling must achieve a threshold whose magnitude may vary depending on the cumulative biochemical activities of the checkpoint and spindle proteins. Depending on the number of unattached kinetochores, a cell may vary the length of the delay. As discussed below, different cell lines or cell types may express different amounts of checkpoint and spindle proteins and thus influence the magnitude of the threshold that is necessary to maintain cells in a checkpoint-inhibited state. As a complex network, a functional mitotic checkpoint consists of many upstream regulators and downstream effectors. The hZW10–ROD complex and CENP-E are examples of proteins that facilitate the actions of the conserved mitotic checkpoint proteins (Abrieu et al., 2000; Chan et al., 2000; Famulski and Chan, 2007; Famulski et al., 2008; Yao et al., 2000). For instance, hZW10 complex is essential for recruitment of Mad1 to the unattached kinetochore (Kops et al., 2005a; Liu et al., 2006), while CENP-E is a binding partner of BubR1 (Chan et al., 1998) and is a regulator of BubR1 kinase activity (Mao et al., 2003, 2005). More recently, Chk1 kinase, a component of the DNA damage checkpoint (Zachos et al., 2007; Zachos and Gillespie, 2007), was found to be required for cells to arrest in mitosis in the presence of taxol, but not nocodazole. The underlying mechanism of differential sensitivity of the mitotic checkpoint to these different drugs suggests that the checkpoint may distinguish between microtubule binding and the functionality of the attachments. Chk1 may be part of the checkpoint pathway that with Aurora B and BubR1 monitors the quality of microtubule attachments (tension) as opposed to

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whether the kinetochore is occupied by a microtubule. This suggests that there may be distinct signaling pathways that are activated or more prevalent in response to the different types of kinetochore attachment defects. This possibility is also supported by the finding that the phosphorylation patterns of various subunits of the APC/C differed between taxol and nocodazole treatment (Steen et al., 2008). The identities of the kinases that are responsible for the different phosphorylation patterns are currently being pursued.

4.3 Aneuploidy/CIN, Mitotic Checkpoint, and Cancer While detailed mechanisms of checkpoint protein function remain under intensive investigation, the identification of these proteins has fueled efforts to study their contribution to the mechanism of aneuploidy and tumorigenesis. Not surprisingly, mitotic checkpoint defects are commonly seen in various types of tumors as well as cancer cell lines (Kops et al., 2005b; Weaver and Cleveland, 2006). Checkpoint defective cells are classified based on their inability to arrest in mitosis in response to spindle poisons such as taxol and the vinca alkaloids. Even in the absence of spindle poisons, these cells often exhibit lagging chromosomes in anaphase or non-disjunction events that result from premature mitotic exit in the presence of improperly aligned chromosomes. Unlike some tumor suppressors, oncogenes, and other cell cycle checkpoint genes that are not essential (Liu and Yen, 2008), all the mitotic checkpoint genes examined to date in mouse models are essential (Baker et al., 2004; Dobles et al., 2000; Iwanaga et al., 2007; Kalitsis et al., 2000; Wang et al., 2004). It appears that the efficiency of the spindle machinery of embryonic mammalian cells is not high enough to operate without a safeguard. In the absence of the mitotic checkpoint, the rate of chromosome missegregation may be so high that it is incompatible with life. Cells that exhibit a defective mitotic checkpoint are therefore more likely to harbor mutations that compromise rather than eliminate this activity. Mutations in the core mitotic checkpoint proteins are uncommon in human cancers. For example, SNP (single nucleotide polymorphisms) analysis of the promoter and coding regions of the six essential spindle checkpoint genes (BUBR1/BUB1B, BUB3, CENP-E, MAD2/MAD2L1, MAD2L2, and MPS1/TTK) in 441 German familial breast cancer cases did not reveal a strong correlation with increased risk of breast cancer (Vaclavicek et al., 2007). By contrast, changes in expression levels of mitotic checkpoint genes and their proteins are frequently seen (Table 4.1) (Coe et al., 2006; Grabsch et al., 2003; Lewis et al., 2005; Miura et al., 2002; Ouyang et al., 2002; Qian et al., 2002; Seike et al., 2002; Shichiri et al., 2002; Shigeishi et al., 2001a, b; Sze et al., 2004; Wang et al., 2002, 2000; Yuan et al., 2006). While mutations were not identified for Bub1, BubR1, Bub3, Mad1, Mad2, and Mps1/TTK in a panel of aneuploid breast cancer cell lines, the levels of RNA and protein encoded by these genes were all elevated as compared to the control breast epithelial cell line MCF10A (Yuan et al., 2006). Furthermore, the same study showed that 77%

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Table 4.1 Status of mitotic checkpoint genes in human cancers that have been examined Gene

Mutation

Misregulation

BUB1 BUBR1 BUB3 MAD1 MAD2

5.4% (13/238) 11.1% (3/27) NA NA 32% (25/77)

47% (122/261) 35.7% (70/196) 80% (66/82) 90% (38/42) 55% (61/110)

Total

11.9% (41/342)

51.7% (357/691)

Compiled from summary by Weaver and Cleveland (2006).

of 270 primary breast tumor samples exhibited a two-fold to three-fold increase in BubR1 levels when compared to 18 normal breast ductal tissues. A similar analysis of a panel of small-cell lung cancer (SCLC) cell lines revealed the presence of extra copies of 7p22.3, a locus that contains the Mad1 gene (Coe et al., 2006). This raises a possibility that overexpression of Mad1 may contribute to the development of this type of cancer. The significance of these observations has been strengthened by the finding that overexpression of Mad2 in mice promoted aneuploidy and tumorigenesis (Sotillo et al., 2007). Precisely how overexpression of checkpoint proteins promotes aneuploidy or tumorigenesis is not known. Increased levels of checkpoint proteins may delay mitosis and thus increase the chances that kinetochores establish defective microtubule attachments. Another possibility is that overexpression may be viewed as a “gain-of-function” mutation where higher levels of checkpoint proteins promote interactions with other growth pathways that they normally do not interact with. Regardless of the defects that promote chromosome missegregation, aneuploidy might promote tumorigenesis by two ways. One possibility is that chromosome instability results in large-scale changes in gene expression patterns that provide the diversity that is necessary for promoting tumorigenesis (Duesberg, 2005; Duesberg et al., 2005). A second possibility is that chromosome instability might facilitate loss of heterozygosity (LOH) of genes that are frequently associated with tumorigenesis. This is illustrated in a study of colorectal cancer where over 50% of early stage colorectal tumors that were examined exhibited an allelic imbalance (AI) of chromosome 5q, which is where the adenomatous polyposis coli (APC) tumor suppressor gene is located. Thus, chromosome instability might act at an early step in the development of colorectal cancer by promoting LOH (Shih et al., 2001). Whether or not chromosome instability directly drives tumorigenesis remains a topic for debate (Hahn et al., 1999; Marx, 2002; Zimonjic et al., 2001). Mice with heterozygous mutations in genes that play critical roles in mitosis, such as BUB1, BUBR1, BUB3, MAD1, MAD2, CENP-E, RAE1, NUP98, SECURIN, RANBP2, and CHFR, are haploinsufficient because their cells exhibit increased rates of aneuploidy (Ricke et al., 2008). In spite of this, the level of spontaneous tumor formation is low. Of the 13 different CIN mice (including Rae1–Bub3 and Rae1–Nup98 compound heterozygotes) that were characterized, 10 were found to be prone to cancer only

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after exposure to carcinogens (Ricke et al., 2008). The frequency of tumorigenesis was also not increased after crossing Bub3+/– mice with p53 or Rb heterozygous mutant mice. The expectation was that chromosome instability resulting from a reduction in Bub3 would promote LOH of either p53 or Rb and thus increase the rate of tumorigenesis. The discrepancy between the expected and the observed may be due to the possibility that the rate of chromosome loss for Bub3+/– mice may be too low to promote LOH of the tumor suppressor genes (Kalitsis et al., 2005). Thus, frequency of LOH may increase in mice that are haploinsufficient for other checkpoint genes. The ability of aneuploidy to increase cell transformation and tumorigenesis may depend on other genetic factors. For example, loss of p16ink4a tumor suppressor gene from hypomorphic BubR1H/H mice (these mutants express < 20% normal BubR1 levels) increased rates of lung adenocarcinoma by nearly 18-fold over the BubR1H/H mutant. Interestingly, loss of the p19Arf tumor suppressor did not increase the incidence of tumor formation relative to parental strain (Baker et al., 2008). The different outcomes seen with these two tumor suppressors may reflect differences in their functions. In these studies, loss of p16ink4a decreased the frequency of senescent cells and premature aging in tissues, while p19Arf acted as an anti-aging factor and prevented cellular senescence in hypomorphic BubR1H/H mice. It is possible that the loss of p16ink4a may increase the chance that aneuploid hypomorphic BubR1 cells survive and become transformed. The complex relationship between tumor suppressor and mitotic checkpoint genes is also seen between APC (adenomatous polyposis coli) and BubR1. BubR1+/– mutant mice do not exhibit spontaneous tumors. When BubR1+/– mice were crossed with heterozygous APCmin/+ mice, the compound mutant progenies had 10 times more colonic tumors (adenocarcinomas) than did APCmin/+ mice (Rao et al., 2005). Despite the increase in the frequency of high-grade colonic tumors, there was a significant decrease in the number of adenomas (polyps) in the small intestine of APCmin/+ /BubR1+/– mice relative to the APCmin/+ mice (Rao et al., 2005). The reduced number of adenomas might be due to the fact that chromosome instability resulting from BubR1 deficiency promoted apoptosis. However, sufficient number of cells with the appropriate chromosome content survived and became tumorigenic. Striking a delicate balance between aneuploidy that can lead to cell death and pathways that allow aneuploid cells to survive is likely a key factor in the early steps of tumorigenesis. The complex relationship between chromosome instability and tumorigenesis is also exemplified in studies of cells and mice that were depleted of the kinetochore motor protein, CENP-E (Ricke et al., 2008; Weaver et al., 2007). Aneuploid mouse embryo fibroblasts (MEFs) from CENP-E heterozygous mutant mice increased their efficiency of transformation with the loss of the p19Arf , or when SV40 large T antigen is overexpressed. Xenograft experiments showed that these transformed MEFs also produced tumors at higher rates than normal MEFs. Analysis of tumor formation in CENP-E+/– mice, however, revealed a more complex picture. Ten percent of CENP-E+/– mice spontaneously developed lymphomas of the spleen and lung tumors after 19–21 months. This frequency is significantly higher than wild-type

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mice. However, the number of spontaneous liver tumors in 19–21-month-old CENPE+/– mice was reduced by 50% when compared to wild-type mice. Even after exposure to the carcinogen, 7,12-dimethylbenz[a]anthracene (DMBA), CENP-E+/– mice developed 20% less total tumors than wild-type controls. The overall rate of tumorigenesis was also delayed in CENP-E+/– /p19Arf null mice relative to mice just lacking p19Arf (Weaver et al., 2007). Based on these complex outcomes, it was suggested that aneuploidy promoted by loss of CENP-E can either promote or suppress tumor formation. Aneuploidy is generally not well tolerated by cells and thus may induce apoptosis and indirectly reduce the frequency of tumor formation. This is supported by studies of yeast and mouse cells where it was shown that chromosome gains can impair proliferation and alter metabolic properties (Torres et al., 2007; Williams et al., 2008). Interestingly, depending on what chromosome was gained, some cells became more sensitive to immortalization, while other cells were less sensitive to immortalization (Hernando, 2008; Torres et al., 2007; Williams et al., 2008). The combined studies suggest that aneuploidy generates the genetic and biochemical diversity that is required to achieve the transformed state. However, the intrinsic and extrinsic factors that influence the cell’s ability to tolerate aneuploidy are also critical determinants at the early steps of tumorigenesis.

4.4 Mitosis as a Target for Chemotherapy Research into the mechanisms that specify chromosome segregation has revealed a plethora of candidate anti-cancer drug targets. As many of the proteins appear to function specifically in mitosis, such inhibitors should exhibit increased specificity and reduce undesirable side effects, such as neuropathies, that are associated with current anti-microtubule agents (Jablonski et al., 2003; Liu and Yen, 2008). Inhibitors that are being developed for the clinics target the Aurora A, Aurora B, Plk1 kinases, and the microtubule motors, CENP-E and kinesin-5 (Jackson et al., 2007; Weaver and Cleveland, 2005). In spite of these new discoveries, the mechanism by which inhibitors of mitosis kill tumor cells remains poorly understood. An important concern with this class of inhibitors is that they can promote aneuploidy that drives the selection of drug-resistant cells (Ganem et al., 2007; Rieder and Maiato, 2004; Storchova and Pellman, 2004). A key toward improving tumor cell response to anti-mitotic agents is to obtain insights into how the mitotic checkpoint is linked to cell survival and death pathways. A prolonged mitotic arrest induced by drugs does not always culminate in death. Time-lapse studies have revealed that there is considerable variability in how cells within a population respond to anti-mitotic drugs (Gascoigne and Taylor, 2008; Orth et al., 2008; Shi et al., 2008). Inhibitors of spindle function will cause the cells to block in mitosis for variable lengths of time. Some cells will die during mitosis but the conditions that lead to this outcome are not known. However, there does not appear to be a strong correlation between death in mitosis and the length of

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the arrest. This is consistent with histological studies that suggested the lack of correlation between the mitotic index and apoptosis in tumors from patients and mouse xenografts that were treated with paclitaxel and docetaxel (Milross et al., 1996; Schimming et al., 1999; Symmans et al., 2000). From a clinical standpoint, an enhanced ability of anti-mitotic drugs to kill cells before they exit mitosis should significantly reduce tumor recurrance. Although studies have reported that an active mitotic checkpoint is an important determinant for activating apoptosis in response to anti-mitotic agents (Masuda et al., 2003; Sudo et al., 2004; Tao et al., 2005; Taylor and McKeon, 1997). Recent studies that monitored the fates of individual cells that were arrested in mitosis indicated that the length of the checkpoint arrest did not correlated with death in mitosis (Gascoigne and Taylor, 2008; Orth et al., 2008; Shi et al., 2008). New data show that death in mitosis does not depend on the ability of the mitotic checkpoint to prolong mitosis. Instead, direct inhibition of factors that promote mitotic exit is more effective at killing cells in mitosis. This suggests that activation of apoptosis may result from the cumulative biochemical affects of an extended mitosis (i.e., super-physiological phosphorylation of cdk1 substrates) (Huang et al., 2009). How cells chose to die during mitosis as opposed to after they have exited mitosis is not understood. One model proposes that death in mitosis occurs if apoptosis is activated before cyclin B1 is degraded and cells exit mitosis (Gascoigne and Taylor, 2008). Consistent with this idea, cyclin B1/cdk1 has been shown to phosphorylate caspase-9, inhibiting its ability to induce apoptosis in cells arrested in mitosis by nocodazole (Allan and Clarke, 2007). It is likely that competing factors dictate whether apoptosis is initiated before the cells exit mitosis. Whether these factors regulate different pathways that dictate death in mitosis or death after mitotic exit is not known. Cells with defective spindles that do not die in mitosis will exit as a result of a crippled checkpoint or “slippage” (Rieder and Maiato, 2004). Slippage is not due to the silencing of the mitotic checkpoint but is thought to be due to a background level of APC/C activity that slowly degrades cyclin B1, and thus cdk1 activity, to levels below that required to maintain the mitotic state (Chibazakura, 2004; Foley et al., 1999; Foley and Sprenger, 2001; Maiato et al., 2002; Tsuiki et al., 2001). After cells exit mitosis, they may senesce, die, or proliferate (Andreassen et al., 2001; Rieder and Maiato, 2004; Weaver and Cleveland, 2005), and these responses may be influenced by p53 status (Andreassen et al., 2001). The link between mitotic stress and apoptosis is complex and this connection has been further complicated by the existence of a caspase-independent mitotic death (CIMD) pathway (Niikura et al., 2007). CIMD appears to respond to defective microtubule–kinetochore attachments in the presence of nocodazale, taxol, or 17-AAG, a Hsp90 inhibitor that disrupts localization of several kinetochore proteins. More importantly, CIMD appears to be dependent on p73 but not p53 and is preferentially activated in cells with an impaired spindle checkpoint. CIN cell lines with reduced Bub1 levels were found to undergo CIMD in response to spindle defects. Thus, Bub1 expression may be an indicator of sensitivity of some tumors to mitotic death mediated by anti-mitotic agents.

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4.5 Conclusions and Future Directions Chromosome instability allows cancer cells to rapidly change their gene expression patterns in a single cell division. These large-scale changes can provide cells with the genetic and biochemical diversity to undergo transformation and overcome poor growth conditions. The mechanism by which cells adopt the CIN phenotype appears to be complex as mutations in genes that encode key components of the mitotic spindle or its regulatory machinery are rare. The underlying defects for CIN have not been defined and may ultimately be due to epigenetic changes that alter the expression levels of multiple proteins so that the spindle functions or activities of the checkpoint proteins operate suboptimally. In addition, mutations in some tumor suppressor genes that allow cells to tolerate the aneuploid state are likely to facilitate CIN. Thus, tolerance of aneuploidy has dual functionality for the neoplastic cell, in that it can promote tumorigenesis and cellular drug resistance. On the other hand, aneuploid cells tend to be less fit than wild-type cells, and this weakness may be exploited to improve response of cancer cells to chemotherapy (Rancati et al., 2008; Torres et al., 2007). Of the many genes whose loss has been shown to result in aneuploidy, no single gene mutation has been found to be responsible for chromosome missegregation in CIN cancer cell lines. The exception may be in colorectal cancer cells where mutations in the APC gene were found to disrupt spindle functions and increase rates of chromosome missegregation (Kaplan et al., 2001). In spite of this, the mechanism that renders the CIN phenotype remains unclear. The defects may be a combination of reduced spindle checkpoint activities and deficient capacity to establish stable kinetochore–microtubule attachments. Comprehensive analysis of mRNA and protein expression along with sequencing of the genomes of CIN cells would undoubtedly be revealing. Identification of deficiencies in specific networks and pathways that provide spindle functions may allow us to exploit them to enhance the killing of CIN cells. As has been suggested, aneuploid cells with defects in kinetochore–microtubule capture might be more sensitive to inhibitors of the mitotic checkpoint given that they may not survive high rates of chromosome missegregation. Establishment of the CIN condition requires not only crippling the mitotic apparatus, but additional mechanisms that allow cells to tolerate the aneuploid state. A comprehensive survey of the activities of various cell death and survival pathways in CIN cells would likely yield important insights about this complex relationship. It is possible that mutations that block cell death pathways may play a critical role. However, it is also possible that aneuploidy itself may select for survivors. Inhibiting the key survival mechanisms in CIN cells should resensitize them to aneuploidy. Mitotic checkpoint inhibitors may be used in conjunction with anti-mitotic drugs to reduce the viability of cells that overcome a checkpoint arrest and thus to improve treatment outcome. The mitotic checkpoint guards against aneuploidy and is likely to play a critical role in determining the sensitivity of tumors to new and existing anti-mitotic agents.

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A weakened checkpoint would promote cells treated with anti-mitotic drugs to exit mitosis and provide an opportunity for the aneuploid cells to survive and proliferate. While it is obvious that targeting cells to die in mitosis would significantly enhance treatment outcome, the link between mitotic arrest and cell death remains to be revealed. Regardless, the fact that cells can die in mitosis suggests a connection between mitotic events and apoptosis. A priori, it seems that cells have evolved a mechanism to kill themselves in response to an inability to properly segregate chromosomes. Improving the efficiency of anti-mitotic drugs would benefit from efforts to identify the molecular links between mitosis and apoptosis.

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Chapter 5

Mitotic Catastrophe Jeremy P.H. Chow and Randy Y.C. Poon

Abstract Mitotic catastrophe is generally defined as a mode of cell death associated with aberrant mitotic activity. It is characterized by unscheduled activation of cyclin B1–CDK1, premature chromosome condensation, and cell death. Progress in the past several years has unraveled a myriad of pathways that can trigger mitotic catastrophe. In this review, we will discuss several common forms of mitotic catastrophe that are highly relevant to cancer. These include mitotic catastrophe triggered by prolonged mitotic arrest induced by microtubule poisons and the abrogation of DNA damage and replication checkpoints. Failure of cells to undergo mitotic catastrophe is believed to contribute to tumorigenesis. Furthermore, mitotic catastrophe is exploited as a strategy to induce cell death in cancer treatments.

5.1 Introduction Proper control of cell number is dictated by a delicate balance between cell cycle and cell death. Deregulation of these controls is the root of genome instability and disorders such as cancer. Progress in the past several years has unraveled some of the underlying principles of a specific form of cell death termed mitotic catastrophe. Although the biological significance and mechanism of mitotic catastrophe remain to be fully defined, the prevailing view is that mitotic catastrophe plays a critical role in maintaining genome stability. Mitotic catastrophe is of particular interest to cancer research because it integrally links cell death to checkpoints and the cell cycle. Understanding mitotic catastrophe may reveal principles of tumorigenesis as well as leads for novel therapeutic designs. Historically, the term “mitotic catastrophe” was first used (Russell and Nurse, 1987) to describe the lethal phenotype associated with the fission yeast strain that R.Y.C. Poon (B) Department of Biochemistry, Hong Kong University of Science and Technology, Clear Water Bay, Hong Kong, China e-mail: [email protected]; http://ihome.ust.hk/~rycpoon


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contain an activated allele of cdc2 (cdc2–3w) and a recessive temperature-sensitive wee1 mutation (wee1-50) (Russell and Nurse, 1986). These mutations overactivate the mitotic engine, resulting in premature and aberrant mitosis, particularly with respect to chromosome segregation and septum formation (Molz et al., 1989). Mitotic catastrophe is now generally defined as a mode of cell death associated with premature or inappropriate entry into mitosis. It should be noted that there is still no consensus definition of mitotic catastrophe. Because of that, there are disagreements and uncertainties about the precise causes, mechanisms, and outcomes of mitotic catastrophe. Here, we adopt a relatively simple definition of mitotic catastrophe to mean a mode of cell death associated with aberrant mitotic activity (Castedo et al., 2004b). Some scholars have defined mitotic catastrophe with mainly morphological terms including the formation of large cells with multiple micronuclei and decondensed chromatin that is clearly distinctive from apoptosis (Roninson et al., 2001). Other groups have considered mitotic catastrophe as a state generally, but not necessary, associated with cell death as an outcome (Erenpreisa and Cragg, 2001; Roninson et al., 2001; Vakifahmetoglu et al., 2008). We are restricting our definition to a process that ultimately results in cell death, thus excluding situations in which cells survive aberrant mitosis, such as after mitotic slippage and the formation of multinucleated giant cells. A myriad of pathways can trigger mitotic catastrophe, including premature mitotic entry, prolonged mitotic block, and defective mitosis. Likewise, several underlying mechanisms appear to be involved in the execution of mitotic catastrophe. In this review, we will discuss several common forms of mitotic catastrophe that are highly relevant to cancer. These include mitotic catastrophe triggered by prolonged mitotic arrest induced by microtubule poisons and the abrogation of DNA damage and replication checkpoints. These conditions are characterized by unscheduled activation of cyclin B1–CDK1, premature chromosome condensation, and cell death.

5.2 Normal Control of Mitosis and the Spindle-Assembly Checkpoint To appreciate how abnormal mitotic events are brought about during mitotic catastrophe, we first briefly review the current paradigm of normal mitotic control in mammalian cells (Fig. 5.1). We refer the reader in this section to a number of relevant review articles. It is well established that CDK1 (also called CDC2) is the key driving force for mitosis. CDK1 is activated by binding to cyclin B, which then phosphorylates substrates that are critical for entry into mitosis. Destruction of cyclin B provides a mechanism to rapidly inactivate CDK1 and allow the cell to exit mitosis. In mammalian cells, cyclin B1 is believed to be the major mitotic B-type cyclin (Fung and Poon, 2005). Cyclin B3 is restricted only to developing germ cells and adult testis, and cyclin B2 does not appear to have an essential function in mice. Different B-type

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ATR/ATM

Spindle Disruption SCF

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PLK1 EMI1 CHK1/CHK2 APC/C WEE1 MYT1 Bora

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Cyclin A2-CDK1/2 CDK1 Cyclin B

Aurora A

PLK1

CDC25

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Fig. 5.1 Control of cyclin B1–CDK1. Cyclin B1–CDK1 is kept inactive during G2 phase by Thr14/Tyr15 phosphorylation. Activation of cyclin B1–CDK1 is orchestrated by feedback loops involving CDC25A/B/C and WEE1/MYT1. How the system is kick-started is uncertain, but may involve in the Aurora A-PLK1–CDC25 axis. After DNA replication block or DNA damage, the cyclin B1–CDK1 activation system is suppressed by the ATM/ATR–CHK1/CHK2–CDC25/WEE1 axis. Once the cell is in mitosis, unattached kinetochores activate the spindle-assembly checkpoint. This checkpoint is mediated by MAD2’s inhibition of APC/CCDC20 . When the checkpoint is satisfied, APC/CCDC20 is turned on to destroy cyclin B1. This inactivates CDK1, leading to the activation of APC/CCDH1 , which in part is responsible in keeping a low level of cyclin B1 expression during G1 phase. During S phase, APC/CCDH1 is turned off by cyclin A–CDK1/2 and EMI1, allowing the re-accumulation of cyclin B1 for the next mitosis. See text for details

cyclins are also differentially localized: While cyclin B1 is cytoplasmic during interphase and translocates into the nucleus during mitotic entry, cyclin B2 colocalizes with the Golgi apparatus and contributes to its fragmentation during mitosis. CDK1 is present throughout the cell cycle. In contrast, cyclin B1 accumulates from S phase and forms a complex with CDK1. The complex is kept inactive by phosphorylation of CDK1Thr14/Tyr15 by MYT1 and WEE1. At the end of G2 phase, the stockpile of inactive cyclin B1–CDK1 complexes is activated abruptly by members of the CDC25 family. Cyclin B1–CDK1 catalyzes its own activation by an intricate network of feedback loops that simultaneously stimulate CDC25 activation and WEE1 inactivation (Lindqvist et al., 2009). Thus, cyclin B1–CDK1 is essentially a bistable system that becomes autocatalytic once a critical portion is activated (Ferrell, 2002). Despite many years of research, however, how the initial batch of cyclin B1–CDK1 complexes is activated remains one of the key outstanding questions in the field. Activation of CDC25 by PLK1 may kick-start the system (van Vugt and Medema, 2005). PLK1 is also involved in the inactivation of WEE1

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and MYT1. The activation of PLK1 in G2 phase requires phosphorylation of Thr210 by Aurora A, an event that is assisted by Bora. Binding of Bora to PLK1 is stimulated by cyclin B1–CDK1-dependent phosphorylation, creating yet another positive feedback loop in the activation of cyclin B1–CDK1 (Lindqvist et al., 2009). Cyclin A–CDK1/2, another cyclin–CDK pair that is activated slightly earlier, may also participate in the activation of cyclin B1–CDK1 (Fung et al., 2007). Phosphorylation of another residue in CDK1, Thr161, is absolutely required for the activation of the kinase activity. However, the activity of the enzyme responsible for Thr161 phosphorylation (CAK) does not appear to be regulated during the cell cycle (Kaldis, 1999). At the end of mitosis, cyclin B1 is removed by the ubiquitin–proteasome system. Specifically, a ubiquitin ligase, designated the anaphase-promoting complex/cyclosome (APC/C) loaded with a targeting subunit termed CDC20, is responsible (Wolf et al., 2007). The timely destruction of cyclin B1 is conferred by a short sequence at the NH2 -terminal region known as the destruction box (D box). APC/C is also responsible for degradation of several other substrates including geminin and securin (Yu, 2007). Degradation of securin releases separase, which in turn cleaves cohesin to allow sister chromatid separation (Yanagida, 2000). Proteolysis of geminin releases CDT1 from the complex to form the pre-replication complex required for the next round of DNA replication (Seo and Kroll, 2006). Activated cyclin B1–CDK1 also negatively regulates itself by stimulating the activity of APC/CCDC20 through phosphorylation of several subunits of APC/C including CDC16, CDC23, and CDC27. APC/C is also activated by PLK1 and inactivated by PKA through phosphorylation. Moreover, phosphorylation of CDC20 by cyclin B1–CDK1 is also necessary for APC/CCDC20 activation. Hence, cyclin B1–CDK1 paves the path for its own destruction by stimulating APC/CCDC20 in a negative feedback loop (Yu, 2007). Activation of APC/CCDC20 is initiated only when all the chromosomes have achieved bipolar attachment to the mitotic spindles. Only one single unattached kinetochore is sufficient to delay the onset of anaphase (Musacchio and Salmon, 2007). Unattached kinetochores or the absence of tension between the paired kinetochores activates a surveillance mechanism termed the spindle-assembly checkpoint. This checkpoint inhibits APC/CCDC20 , thereby maintaining a high level of active cyclin B1–CDK1. A growing body of evidence indicates that unattached kinetochores attract the components of the checkpoint machinery (including BUB1, BUB3, MAD1, MAD2, MAD3/BUBR1, MPS1, and CENP-E), catalyzing the formation of diffusible mitotic checkpoint complexes (MCC; components include MAD2, BUBR1, and BUB3), which in turn inhibit APC/CCDC20 (see Chapter 4). It is generally accepted that the sequestration of CDC20 by MAD2 is a key step in the spindle-assembly checkpoint. Binding to CDC20 requires a conformational change in MAD2 from a less stable open conformation (known as O-MAD2) to the more stable closed conformation (C-MAD2). Upon conformational change from O-MAD2 to C-MAD2, the C-terminal CDC20-binding site (which resembles a “seatbelt” structure) is exposed, thus allowing interaction with CDC20. According to a currently favored MAD2-template model, the C-MAD2 that binds to MAD1

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at the kinetochores serves as a template for the conversion of the cytosolic pool of O-MAD2 into C-MAD2. The newly activated MAD2 then leaves the kinetochore to bind and inhibit CDC20. As MAD2 has the same conformation when complexed with both MAD1 and CDC20, the C-MAD2–CDC20 complexes may autoamplify the checkpoint signal by recruiting more O-MAD2 to the complex. This model provides an elegant mechanism for cytosolic propagation of the checkpoint signal away from kinetochores (Musacchio and Salmon, 2007). Within minutes after all kinetochores are properly attached, the spindle-assembly checkpoint is terminated to allow APC/CCDC20 activation and anaphase onset. How the checkpoint is silenced is not entirely clear, but evidence suggests that several mechanisms may be involved. These include the stripping of checkpoint components from the kinetochore by a dynein motility-dependent mechanism, an energy-dependent MAD2–CDC20 complex dissociation mechanism, an APC/Cdependent mechanism, and the neutralization of MAD2 by binding to p31comet (Musacchio and Salmon, 2007). In marked contrast to CDC20, phosphorylation of CDH1 by cyclin B1–CDK1 alters the conformation of CDH1 and prevents its binding to APC/C, thus keeping APC/CCDH1 inactive during mitosis. Destruction of cyclin B1 at the end of mitosis downregulates CDK1 activity and therefore relieves the inhibition of APC/CCDH1 (Baker et al., 2007; Pesin and Orr-Weaver, 2008). The phosphatase CDC14, which is believed to remove most of the phosphorylation carried out by MPF at the end of mitosis (Bembenek and Yu, 2003; Stegmeier and Amon, 2004), antagonizes the inhibitory phosphorylation on CDH1. The activated APC/CCDH1 then degrades CDC20 and takes over the task of degrading any remaining cyclin B1. APC/CCDH1 remains active during G1 phase to curb the unscheduled accumulation of mitotic cyclins. Cell cycle-dependent transcription of cyclin B1 provides an additional level of regulation of cyclin B1 (Fung and Poon, 2005). At the G1 –S transition, APC/CCDH1 itself is turned off by phosphorylation, allowing the re-accumulation of cyclin B1. Cyclin A–CDK1/2 phosphorylates CDH1, resulting in the dissociation of CDH1 from the APC/C core. APC/CCDH1 is also turned off by EMI1, which begins to accumulate at the G1 /S transition (van Leuken et al., 2008). During normal prophase, the SCFβ-TrCP targets EMI1 for ubiquitin-mediated degradation. This degradation of EMI1 is promoted by PLK1-dependent phosphorylation (Eckerdt and Strebhardt, 2006).

5.3 Mitotic Catastrophe Caused by Mitotic Block and Mitotic Slippage Agents that suppress microtubule dynamics can artificially activate the spindleassembly checkpoint by leaving the kinetochores unoccupied. These include chemicals that inhibit microtubules depolymerization (e.g., Taxol) or polymerization (e.g., vinca alkaloid and nocodazole). Spindle-disrupting drugs are among the most

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important chemotherapeutic agents available for a variety of cancers (Weaver and Cleveland, 2005). Activation of the spindle-assembly checkpoint leads to persistent activation of cyclin B1–CDK1 and stabilization of APC/C targets. While numerous targets have been proposed, the precise mechanism underlying spindle toxins-mediated mitotic catastrophe remains controversial (see below). A direct link between the spindle-assembly checkpoint and mitotic catastrophe is yet to be fully established. Paradoxical evidence implies that the spindle-assembly checkpoint is either required or inhibitory to apoptosis. Checkpoint disruption with dominant-negative BUB1 (Taylor and McKeon, 1997) or BUBR1 (Shin et al., 2003), siRNA against MAD2 or BUBR1 (Sudo et al., 2004), or overexpression of Aurora A (Anand et al., 2003) inhibits the cell death induced by nocodazole or Taxol. Similarly, several human cancer cell lines that contain an impaired checkpoint are resistant to nocodazole treatment (Masuda et al., 2003). In disagreement with these findings, disruption of the spindle-assembly checkpoint by trichostatin A (Dowling et al., 2005), a mutant CDC20 (Sihn et al., 2003), or siRNA targeting BUB1 (Niikura et al., 2007) or BUBR1 (Lee et al., 2004) sensitizes cells to nocodazole- or Taxol-induced killing. The role of cyclin B1–CDK1 in spindle poison-induced cell death is also controversial. Inhibition of CDK1 by chemical inhibitors, a dominant-negative CDK1, or cyclin B1 antisense oligonucleotides prevents Taxol-induced apoptosis (Yu et al., 1998; Shen et al., 1998). Likewise, inhibition of CDK1 with the specific inhibitor RO3306 reduced nocodazole-induced apoptosis (Chan et al., 2008). However, no reduction of nocodazole-induced apoptosis was found in the presence of the CDK inhibitor olomoucine (Masuda et al., 2003). Moreover, addition of CDK inhibitors to Taxol-treated cells enhances apoptosis (O’Connor et al., 2002; Wall et al., 2003; Pennati et al., 2005). Results from kinase-dead CDK1 expressing or CDK1 conditional knockout cells similarly point to an apoptosisprotecting role of cyclin B1–CDK1 (O’Connor et al., 2002). Furthermore, continuous spindle-assembly checkpoint function, but not cyclin B1-CDK1 activity, is required for caffeine-induced apoptosis in nocodazole-arrested cells (Gabrielli et al., 2007). Spindle toxin-stimulated cell death is further complicated by mitotic slippage (also called adaptation) (Weaver and Cleveland, 2005; Rieder and Maiato, 2004). After a prolonged block in mitosis, some cells can inactivate cyclin B1–CDK1 precociously and exit mitosis without chromosome segregation and cytokinesis. The nuclear envelope then randomly reforms around groups of chromosomes. Although the exact mechanism of mitotic slippage is not known, the central event seems to be a slow but continuous degradation of cyclin B1. It was found that slippage is required for the apoptosis induced in response to mitotic kinesin Eg5 inhibition, which leads to monopolar spindle formation and spindle-assembly checkpoint activation (Mayer et al., 1999). Slippage-refractory cells are resistant to Eg5 inhibitor-induced apoptosis, but promotion of mitotic slippage with CDK inhibitors enhances cell death (Tao et al., 2005). However, contradictory evidence suggests that inhibiting Eg5 activity leads to apoptosis regardless of spindle-assembly checkpoint function (Chin and Herbst, 2006).

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A solution to the problem may be provided by the findings that although cell death is reduced by mitotic slippage, it is induced in the subsequent multipolar mitosis (Chan et al., 2008). Mitotic slippage generates cells that contain tetraploid DNA contents and two centrosomes, both of which can be duplicated during the subsequent S phase. Because centrosomes are microtubule organization centers, cells with supernumerary centrosomes form multipolar mitotic spindles. The uneven segregation of genetic material into the daughter cells may result in different fates, including mitotic catastrophe (Chan et al., 2008), aneuploidy, and transformation. Several studies have provided evidence that tetraploidization increases chromosome instability in yeast (Mayer and Aguilera, 1990; Storchova et al., 2006) and mammalian cells (Cowell, 1980; Fujiwara et al., 2005). A p53-dependent tetraploidy checkpoint has been proposed to prevent S phase entry in cells that have undergone adaptation or aborted cytokinesis (Andreassen et al., 2001). However, the existence of the tetraploidy checkpoint has been disputed (Fujiwara et al., 2005; Uetake and Sluder, 2004; Wong and Stearns, 2005). One of the possibilities is that the p53-dependent arrest after tetraploidization is mainly due to DNA damage or centrosomal stress during the aberrant mitosis (Storchova and Kuffer, 2008). Mitotic catastrophe per se does not seem to depend on p53 as it occurs in both p53-positive and p53-negative cells (Lanni and Jacks, 1998; Minn et al., 1996). Nevertheless, p53 may affect mitotic catastrophe by indirectly regulating cyclin B1–CDK1 activity. The abundance of cyclin B1 mRNA is negatively regulated by p53 through a transcriptional repression mechanism (Dan and Yamori, 2001).

5.4 Normal Control of the DNA Damage and Replication Checkpoints Several checkpoints that monitor DNA integrity prevent precocious entry into mitosis (Fig. 5.1). Progress in the past several years has unraveled very similar underlying principles in the DNA replication checkpoint, the intra-S DNA damage checkpoint, and the G2 DNA damage checkpoint in preventing the activation of CDK1. In essence, DNA damage or replication stress activates sensors that facilitate the activation of the PI-3 (phosphoinositide 3-kinase)-related protein kinases ATM and ATR. ATM/ATR then activates CHK1 or CHK2, which in turn inactivates CDC25s and activates WEE1, culminating in the inhibitory phosphorylation of CDK1 (Kastan and Bartek, 2004). We refer the reader below to relevant reviews. Following exposure to ionizing radiation or other genotoxic insults that elicit DNA double-strand breaks, ATM is autophosphorylated at Ser1981, leading to dimer dissociation and activation of the kinase. ATR is activated by a broader spectrum of stress including ultraviolet irradiation, hypoxia, and replication stress. ATM and ATR phosphorylate residues in the SQ/TQ domain of CHK1/CHK2, thereby stimulating the kinase activity of these effector kinases. The upstream sensors that initiate the activation of ATM/ATR consist of an intricate network of large protein complexes, of which many components

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contain the BRCT domain. These include the RAD9–HUS1–RAD1 (9-1-1) clamp and the RAD17–RFC clamp loader that facilitate ATR-mediated activation of CHK1 (Parrilla-Castellar et al., 2004). Another large complex that participates in ATM/ATR activation is the so-called BRCA1-associated genome surveillance complex composed of BRCA1, BLM, and MRN (MRE11–RAD50–NBS1) (Wang et al., 2000; Jhanwar-Uniyal, 2003). Stalled replication forks mainly activate the ATR–CHK1 pathway. Replication fork progression can be impaired by insufficient nucleotide supply or lesions and obstacles on the DNA. Several proteins including ATRIP (ATR-interacting protein), TopBP1, and Claspin appear to be required for recruiting ATR to single-stranded DNA present at stalled replication forks to phosphorylate CHK1 (Cimprich and Cortez, 2008). The ATR–CHK1 pathway is essential even in the absence of exogenous stresses during unperturbed S phase, probably for maintaining high rates of replication fork progression (Petermann and Caldecott, 2006). Claspin is usually degraded by SCFβ-TrCP -mediated ubiquitination following the phosphorylation of Claspin by PLK1. This pathway is inhibited after DNA damage (Freire et al., 2006). In response to genotoxic stress in G2 phase, the phosphatase CDC14B translocates from the nucleolus to the nucleoplasm and activates APC/CCDH1 . This degrades PLK1 and consequently stabilizes Claspin, allowing the G2 DNA damage checkpoint to be maintained (Bassermann et al., 2008). CHK1 and CHK2 are believed to be involved in the inactivation of all three isoforms of the CDC25 family (CDC25A, CDC25B, and CDC25C) (Boutros et al., 2006). Phosphorylation of CDC25CSer216 by CHK1/CHK2 inactivates its phosphatase activity either directly or indirectly through the creation of a 14-3-3 binding site. Binding of 14-3-3 masks a proximal nuclear localization sequence and anchors CDC25C in the cytoplasm, preventing efficient access of CDC25C to cyclin B1–CDK1. Interestingly, phosphorylation of a proximal site (Ser214) by cyclin B1–CDK1 inhibits further phosphorylation of CDC25CSer216 . This provides an elegant mechanistic explanation for the suppression of DNA damage-mediated CDC25C inactivation during mitosis (Chen and Poon, 2008). CDC25B is believed to possess a unique role in activating cyclin B1–CDK1 at the centrosome. A growing body of evidence indicates that CHK1 may shield centrosomal cyclin B1–CDK1 from unscheduled activation by CDC25B during normal G2 phase and presumably also during the G2 DNA damage checkpoint. The molecular basis of this activity may be due to CHK1-dependent phosphorylation of CDC25BSer323 , creating a docking site for 14-3-3 that prevents access of substrates to the catalytic site. Dissociation of CHK1 from the centrosomes at the end of G2 phase, together with positive regulatory phosphorylation of CDC25BSer353 by Aurora A, enables CDC25B to activate the centrosomal cyclin B1–CDK1 and initiate mitosis (Boutros et al., 2007). CDC25A is arguably the most important member of the CDC25 family due to its nonredundant role in mouse cells. CDC25A is targeted for rapid degradation by CHK1/CHK2 through a ubiquitin-mediated mechanism. CDC25A stability is controlled by the APC/CCDH1 complex during mitotic exit and early G1 , but by the SCFβ-TrCP complexes during interphase. Importantly, the SCFβ-TrCP -dependent

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turnover of CDC25A is enhanced in response to DNA damage. Phosphorylation of CDC25ASer76 by CHK1 is required for the phosphorylation of a phosphodegron centered at Ser82 (by an as-yet-unidentified kinase), creating a binding site for β-TrCP. Interestingly, β-TrCP also binds to a separate nonphosphorylated sequence in CDC25A (the DDG motif) and plays a role in CHK1-induced ubiquitination and degradation of CDC25A (Boutros et al., 2007). There is also evidence that CHK1 can phosphorylate and activate WEE1 by promoting 14-3-3 binding (Rothblum-Oviatt et al., 2001; Lee et al., 2001). Suppression of CDC25s or activation of WEE1 promotes CDK1Thr14/Tyr15 phosphorylation, thus preventing damaged cells from entering mitosis. Other mechanisms are also known to play critical roles in the G2 DNA damage checkpoint. For example, 14-3-3σ is involved in sequestering cyclin B1–CDK1 in the cytoplasm after DNA damage (Chan et al., 1999). In addition to its well-known role in the G1 DNA damage checkpoint, the p53–p21CIP1/WAF1 axis is also important for the G2 arrest after DNA damage (Bunz et al., 1998). In p53-deficient cells, a pathway involving the MAP kinase p38 and MK2 (MAPKAP kinase-2) is important for the G2 DNA damage checkpoint (Reinhardt et al., 2007). Activation of this pathway by genotoxic stress is also dependent on ATM/ATR.

5.5 Mitotic Catastrophe Caused by Abrogation of DNA Integrity Checkpoints Conceptually, it is vital to prevent the precocious activation of cyclin B1–CDK1 and mitotic entry as long as replication or DNA repair remains incomplete. It is well known that bypass of the classic checkpoint pathways described above promotes mitotic catastrophe. For example, cells lacking p53, p21CIP1/WAF1 , or 14-3-3σ fail to arrest in G2 after DNA damage and undergo mitotic catastrophe (Chan et al., 1999; Bunz et al., 1998). Depletion of p38 MAP kinase in p53-deficient cells leads to checkpoint deregulations and mitotic catastrophe during DNA damage (Reinhardt et al., 2007). Furthermore, the G2 DNA damage checkpoint is partially impaired in many cancer cells (Fingert et al., 1986). They are unable to maintain G2 arrest and eventually undergo aberrant mitosis and mitotic catastrophe (Chang et al., 2000). Uncoupling of the ATM/ATR–CHK1/CHK2 axis can ablate the DNA damage and replication checkpoints, inducing unscheduled activation of cyclin B1–CDK1 and mitotic catastrophe (Chan et al., 1999; Castedo et al., 2004b; Vogel et al., 2007). Cells that contain defective ATM, such as those derived from ataxia telangiectasia, often exhibit radio-resistant DNA synthesis and mitotic catastrophe (Lavin and Khanna, 1999). In support of a role of CHK2 in the G1 DNA damage checkpoint, the IR-induced G1 arrest is impaired in CHK2–/– mouse embryonic fibroblasts (Takai et al., 2002). CHK2–/– mice are relatively normal and fertile. In contrast, ATR- and CHK1-deficient mice die at an early embryonic stage with morphological abnormalities similar to mitotic catastrophe (Brown and Baltimore, 2000; Takai et al., 2000; Liu et al., 2000). Studies using conditional CHK1 knockout mice also revealed that

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CHK1 deficiency causes inappropriate S phase entry, accumulation of DNA damage during replication, and mitotic catastrophe (Lam et al., 2004; Niida et al., 2005). Since the effect of the ATM/ATR–CHK1/CHK2 pathway is on the inhibitory phosphorylation of CDK1, it is not surprising that expression of a nonphosphorylatable mutant of CDK1 can trigger mitotic catastrophe (Blasina et al., 1997; Chow et al., 2003; Niida et al., 2005). Agents that can inhibit the ATM/ATR–CHK1/CHK2 pathway can induce mitotic catastrophe and many are potential chemotherapeutic agents. Caffeine is a classic agent that inhibits ATM/ATR (Blasina et al., 1999; Hall-Jackson et al., 1999; Sarkaria et al., 1999) The checkpoints can also be uncoupled with CHK1 inhibitors such as UCN-01 (Wang et al., 1996; Graves et al., 2000; Tse and Schwartz, 2004). However, UCN-01 also inhibits MK2 and may induce mitotic catastrophe through this pathway (Reinhardt et al., 2007). Likewise, inhibition of CHK2 promotes mitotic catastrophe after DNA damage (Castedo et al., 2004a). The spindle-assembly checkpoint is required for mitotic catastrophe induced by abrogation of the DNA damage checkpoint (Nitta et al., 2004; Vogel et al., 2004), suggesting a trap in mitosis is required for these types of cell death.

5.6 Mitotic Catastrophe as a Specialized Form of Cell Death Involving CDK1 Mitotic catastrophe is generally defined as a type of cell death linked to abnormal activation of mitotic kinases. It has been argued whether mitotic catastrophe is a special case of apoptosis (Castedo et al., 2004b). Mitotic catastrophe frequently displays phenotypic characteristic of apoptosis and shares several molecular hallmarks with apoptosis such as caspase activation. Other studies suggest that mitotic catastrophe represents a form of cell death distinct from apoptosis (Roninson et al., 2001). The issue whether mitotic catastrophe is a mode of cell death or a process leading to apoptosis is still controversial (Chu et al., 2004; Skwarska et al., 2007; Vakifahmetoglu et al., 2008). The many overlapping effects of mitotic catastrophe and apoptosis add to the uncertainty. For example, CDK1-dependent phosphorylations are important in triggering nuclear envelop breakdown and chromatin condensation during normal mitosis and presumably also in mitotic catastrophe (Porter and Donoghue, 2003). During apoptosis, these events also occur but are carried out at least in part by caspases. Caspase-6 is essential for cleavage of nuclear lamins (Orth et al., 1996; Takahashi et al., 1996) and for chromatin condensation (Ruchaud et al., 2002), especially in cells that express lamin A/C (Slee et al., 2001). In this connection, caspase-6 also cleaves cyclin B1 itself during mitotic catastrophe, generating a non-degradable version of cyclin B1 (Chan et al., 2009). What is the molecular basis of cell death during mitotic catastrophe? Cyclin B1–CDK1 kinase activity is increased in apoptotic cells induced by different death signals, suggesting that premature or prolonged activation of CDK1 may cause apoptosis. Indeed, the fusion of mitotic cells with cells in S or G2 phase results

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in mitotic catastrophe, probably due to the unscheduled active cyclin B1–CDK1 complexes (Mackey et al., 1996). Overexpression of cyclin B1 and CDK1 induces premature chromosome condensation and mitotic catastrophe (Heald et al., 1993; Jin et al., 1998). Prolonged accumulation of cyclin B1 is also the key requirement for cell death after spindle disruption (Chan et al., 2008). Knockdown of cyclin B1 inhibits mitotic catastrophe induced by the bypass of DNA damage checkpoints (Chan et al., 2009). While the underlying principles of CDK1-mediated toxicity in general remain largely unresolved, a few targets that affect survival have been identified. Several members of the BCL-2 family can be phosphorylated by cyclin B1–CDK1. BAD is phosphorylated by cyclin B1–CDK1 at Ser128 (Berndtsson et al., 2005; Konishi et al., 2002). This reduces the interaction between phosphorylated Ser136 and 14-3-3, allowing BAD to translocate to mitochondria and induce cell death. BCL2 is also phosphorylated by cyclin B1–CDK1, but the role in mitotic catastrophe remains to be clarified (Brichese et al., 2002; Furukawa et al., 2000; Ling et al., 1998; Scatena et al., 1998). Kinases other than CDK1 have also been implicated in controlling the BCL-2 family during mitotic catastrophe. For example, disruption of microtubules activates p38 MAP kinase, which induces the translocation of BAX to mitochondria and enhances mitotic catastrophe (Deacon et al., 2003). Survivin is another potential target of cyclin B1–CDK1 during mitotic catastrophe. Survivin is a member of the inhibitors of apoptosis (IAPs) family and is believed to inhibit apoptosis. Survivin is also a component of the chromosome passenger complex, requiring for a sustained spindle-assembly checkpoint arrest in response to the lack of tension at the kinetochore (Carvalho et al., 2003; Lens et al., 2003). Survivin accumulates during G2 /M phase and its degradation is controlled by ubiquitination. Phosphorylation of survivinThr34 by CDK1 stabilizes the protein (O’Connor et al., 2000). Since depletion of survivin by siRNA abolishes Taxol-induced mitotic arrest and cell death (Carvalho et al., 2003; Lens et al., 2003), the regulation of survivin by CDK1 probably contributes to the mitotic arrest rather than apoptosis. In fact, inhibition of CDK1 with chemical inhibitors downregulates survivin expression and induces MYC-dependent apoptosis (Goga et al., 2007). Given that the role of survivin in mitotic catastrophe is ambiguous, it is not clear whether the cytotoxicity of chemicals that downregulates survivin (such as the COX-2 inhibitor Celecoxib) involves mitotic catastrophe at all.

5.7 Mitotic Catastrophe and Cancer: Future Directions Mitotic catastrophe has been studied from two perspectives in cancer research. On the one hand, failure of some cells to undergo mitotic catastrophe is believed to contribute to tumorigenesis. On the other hand, mitotic catastrophe is exploited as a strategy to induce cell death in cancer treatments. In cells that contain defective checkpoints, mitotic catastrophe represents a final mechanism to prevent genome instability. Although most cells that bypass the DNA

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integrity checkpoints eventually die through mitotic catastrophe, surviving cells may undergo aberrant mitosis, resulting with mis-segregation of chromosomes and cytokinesis failure. Likewise, cells that evade mitotic catastrophe after prolonged activation of the spindle-assembly checkpoint enter a tetraploidy G1 state. If additional checkpoints are lacking (such as being p53 defective), aneuploid cells can further undergo DNA replication and centrosome duplication, paving the way to multipolar mitosis and further genome instability (Fig. 5.2). It has long been realized that many tumors contain a population of polyploid cells (Goga et al., 2007). It is likely that the ability to execute cytokinesis reduces as the ploidy increases, giving rise to multinucleated giant cells (King, 2008). Stresses

Checkpoints

Spindle disruption

Spindle-assembly checkpoint

Replication block

Replication checkpoint

Outcomes

Checkpoint recovery

Checkpoint bypass Mitotic catastrophe DNA damage

DNA damage checkpoint

Cell cycle genome instability

Fig. 5.2 Mitotic catastrophe is an important mechanism to prevent genome instability. Checkpoints are turned on in response to various stresses to halt the cell cycle. After the stresses are resolved, the checkpoints are turned off to allow cell cycle progression. Prolonged activation of the spindle-assembly checkpoints frequently results in mitotic catastrophe. Bypass of the DNA integrity checkpoints may also induce precocious activation of cyclin B1–CDK1 and mitotic catastrophe. Cells that failed to undergo mitotic catastrophe with defective checkpoint execution may complete mitosis inappropriately, resulting in genome instability

The presence of polyploid giant cells that fail to be killed by mitotic catastrophe may also account for resistant to cancer therapy. Following DNA damage (in particular with relatively low dose of DNA damaging agents), many polyploid cells appear after an initial phase of mitotic catastrophe and survive for weeks as monoor multinucleated giant cells (Blagosklonny, 1999; Puig et al., 2008). Whether these cells still retain proliferative potential is controversial. Some groups claim that giant cells have reduced or no proliferative potential (Therman and Kuhn, 1989). Others showed that giant cells can undergo multipolar mitosis or depolyploidization to return to near diploid state (Erenpreisa and Cragg, 2001). These studies suggest that the multistep process of escaping mitotic catastrophe through polyploidization and then depolyploidization may account for tumor relapse after initial efficient cancer therapy. Conversely, strategies have also been developed to sensitize cancer cells to undergo mitotic catastrophe. Small molecules that inhibit the G2 DNA damage checkpoint should in principle promote mitotic catastrophe when used in

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combination with DNA damaging agents. Among these, the CHK1 inhibitor UCN-01 is in relatively more advanced development (Senderowicz, 2003). Inhibition of CHK1 with UCN-01 after DNA damage overcomes the DNA damage checkpoints, inducing premature activation of cyclin B1–CDK1 and mitotic catastrophe. However, UCN-01 is also a potent inhibitor of protein kinase C, CDKs, MK2, AKT (through inhibition of phosphoinositide-dependent kinase 1), and other kinases. This promiscuous nature of UCN-01 (and protein kinase inhibitors in general) makes defining its precise role difficult. For example, the two kinases that can phosphorylate CDC25CSer216 – cTAK1 and CHK1 – can both be inhibited by UCN01 (Busby et al., 2000). Preclinical results using UCN-01 are promising and, thus, several clinical trials that combine various DNA damaging drugs with UCN-01 are under way (Blagden and de Bono, 2005; Kortmansky et al., 2005). CDK1 inhibitors, such as roscovitine, are expected to inhibit mitotic catastrophe. Indeed, some studies indicate that treatment with roscovitine blocks Adriamycininduced mitotic catastrophe (Park et al., 2005). However, since roscovitine itself is a potent chemotherapeutic agent, there are many more reports showing that roscovitine enhances DNA damage-induced cell death (the underlying mechanism is not obvious, but MYC-overexpressing tumors are particularly sensitive to roscovitine (Goga et al., 2007)). As with other potential therapies against cell cycle regulators, it is not immediately obvious why agents that induce genotoxic stress and mitotic catastrophe should selectively target cancer cells and spare normal cells (Hunt, 2008). The usual explanation is that the balance of various cell cycle regulators is severely altered in cancer cells. This sensitizes cancer cells to agents that further disrupt cell cycle and checkpoint control. A greater understanding of precisely how checkpoints are controlled in normal and cancer cells is essential for designing better therapeutic agents. Acknowledgments We apologize for those whose work that could not be cited due to space constraints. Related works in our laboratory are supported by Research Grants Council grant HKUST6439/06 M to R.Y.C.P.

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Chapter 6

p53, ARF, and the Control of Autophagy Robert D. Hontz and Maureen E. Murphy

Abstract The p53 and ARF genes encode well-known tumor suppressor proteins that respond to oncogenic and genotoxic signals in order to induce growth arrest or apoptosis. Recently, both of these proteins were found to be intimately tied to metabolic pathways and to play surprising roles in autophagy. Autophagy (“selfeating”) is a critical response of eukaryotic cells to stress. During this process, portions of the cytosol, including cytoplasmic organelles, are sequestered into characteristic double-membrane vesicles called autophagosomes that are delivered to the lysosome for degradation. This rather non-specific degradation process allows the cell to adapt to its bioenergetic needs and to prolong survival. The following sections will outline the evidence for a role of p53 and ARF in autophagy, the role of this pathway in cancer, and what questions remain to be answered.

6.1 The ARF Tumor Suppressor and Autophagy The p14ARF tumor suppressor (p19ARF in mouse, and hereafter referred to as ARF) is transcribed from within the INK4a/ARF locus using an alternate reading frame from the p16INK4a tumor suppressor. The two overlapping tumor suppressor genes comprise a locus that is commonly mutated and/or deleted in human cancer (Quelle et al., 1995; Sherr et al., 2005). While a number of transcription factors coordinately control the expression of the INK4a/ARF locus, the ARF and INK4a genes have separate promoters and are chiefly regulated in an independent manner (Gil and Peters, 2006). The ARF gene is tightly suppressed at the transcriptional level in normal, non-transformed cells, but is upregulated in response to mutational activation of various oncogenes, including Ha-ras, c-MYC, or β-catenin, or in response

M.E. Murphy (B) Program in Molecular and Translational Medicine, Fox Chase Cancer Center, Philadelphia, PA 19111, USA e-mail: [email protected]


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to chronic mitogenic stimuli, such as would occur in cultured cells (for review see Sherr et al., 2005). ARF is a predominantly nucleolar protein whose best understood function is as a positive regulator of the p53 tumor suppressor protein. ARF stabilizes p53 by sequestering and inhibiting the E3 ubiquitin ligase MDM2, which normally ubiquitylates p53 and targets it for proteasome-mediated degradation. As a result, cells with transcriptionally upregulated ARF undergo cell cycle arrest in a p53-dependent manner and are likewise sensitized to p53-dependent apoptosis (for review see Sherr et al., 2005). Interestingly, ARF can also induce cell growth arrest and apoptosis in a p53-independent manner (Kelly-Spratt et al., 2004; Weber et al., 2000); this is believed to occur via physical interaction with, and inhibition of, a number of cell cycle regulators, including nucleophosmin (B23/NPM), DP1, E2F, c-myc (for review see Sherr et al., 2005). When overexpressed in cells, ARF inhibits the function(s) of these proteins by sequestering them in the nucleolus. Additionally, ARF can inhibit nucleophosmin and MDM2 function by enhancing the sumoylation of these proteins (Tago et al., 2005). ARF has also been shown to suppress cell growth by inhibiting the processing of precursor rRNA transcripts (Sugimoto et al., 2003). Each of these diverse activities of ARF is believed to underlie at least part of the tumor suppressor function of this protein.

6.2 ARF Induces Autophagy It was first reported in 2006 that both human and mouse ARF can be translated from an internal methionine (Met48 in human, Met45 in mouse) to generate a short version of the protein denoted smARF (short mitochondrial ARF) (Reef et al., 2006). smARF lacks the N terminus of ARF, which encodes much of ARF’s tumor suppressor functions; specifically, smARF lacks the MDM2-binding domain (which mediates p53-dependent growth arrest), the nucleolar localization sequence, the nucleophosmin-binding domain, and the region required to inhibit rRNA processing. Immunolocalization and biochemical fractionation studies determined that, in contrast to full-length ARF, smARF predominantly localized to mitochondria (Reef et al., 2006). The mitochondrial localization of smARF is supported by reports that both human and mouse smARF interact with the protein p32, which is mitochondrial (Reef et al., 2007; Itahana and Zhang, 2008). It should be noted that one group reported that, in addition to the mitochondria, smARF also localizes to the nucleus and cytoplasm (Ueda et al., 2008). Full-length ARF and smARF are generated in equal proportions using in vitro translation reactions; however, smARF has a very short half-life in vivo, as it is rapidly eliminated by proteasome-mediated degradation. Consequently, smARF is present at very low steady-state levels (less than 5% of total ARF protein) (Reef et al., 2006). Kimchi and colleagues found that enforced expression of smARF resulted in abnormalities in mitochondrial membrane potential, which they found were caused by increased autophagy. While autophagy is primarily regarded as a cell survival pathway, chronic induction of autophagy is believed to lead to cell death.

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Kimchi’s group showed that acute overexpression of smARF induces autophagic cell death, in a p53- and caspase-independent manner. Conflicting reports exist regarding the potential role of full-length ARF in autophagy. Abida and Gu engineered a version of ARF in which the internal methionine was mutated and showed that this form could induce autophagy in transfected cells (Abida and Gu, 2008). In contrast, Reef and Kimchi published findings that full-length (nucleolar) ARF is incapable of inducing autophagy except when ARF was expressed at non-physiological levels, at which time ARF was observed in extra-nuclear compartments (Reef and Kimchi, 2008). This group concluded that, at physiologically relevant concentrations, only smARF was able to induce autophagy, and that this occurred predominantly at mitochondria. Because both groups relied heavily on transient transfection and overexpression of ARF and smARF in cells, the contribution of full-length ARF to autophagy remains unresolved.

6.3 ARF-Mediated Autophagy Can Enhance Cell Survival and Promote Tumor Progression Recently our group reported that ARF has a pro-survival role in autophagy. Rather than relying on overexpression of this protein in transfected cells, we took advantage of the fact that p53 is a potent repressor of the ARF gene, and that p53-null cells express high levels of endogenous ARF (for review see Gil and Peters, 2006). Building upon reports that ARF is frequently overexpressed in B-cell tumors (Eischen et al., 1999), we silenced this gene in B-cell lymphomas using two different short hairpins for ARF and found that silencing ARF impaired autophagy, as well as the ability of these cells to establish tumors in vivo (Humbey et al., 2008). These data support a cytoprotective (rather than cytotoxic) role for ARF in autophagy. Consistent with this premise, we found that silencing ARF reduced the survival of lymphoma cells and mouse embryo fibroblasts exposed to nutrient-deprived conditions. These data may explain the large percent of human tumors with mutant p53 that retain high levels of ARF expression and support the premise that ARF and autophagy promote the survival of some tumors. The cytoprotective role of ARF and autophagy may be restricted to certain tumor types. Specifically, we found that silencing ARF in three different primary lymphoma cell lines reduced survival and progression in vivo. In contrast, silencing ARF in a p53-null sarcoma cell line enhanced the progression of this tumor in a xenograft assay, despite clearly inhibiting autophagy (Pimkina and Murphy, 2009). These data indicate that autophagy may be cytoprotective only for certain tumors. Alternatively, ARF’s cytoprotective role may be balanced by its p53-independent growth suppressive functions, and in the sarcoma cell line tested the latter effect predominated. Recently our group used the technique of two-dimensional in-gel electrophoresis to identify ARF-interacting proteins at the mitochondria. This study revealed that at the mitochondria, ARF physically interacts with the Bcl2 family member Bcl-xl (Pimkina et al., 2009). Bcl-xl plays a known role in autophagy; it interacts with

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Beclin-1 and negatively regulates the kinase activity of the Vps34/Beclin-1 complex, which is a key mediator of autophagy (Maiuri et al., 2007; Pattingre et al., 2005). Our group showed that ARF inhibits the ability of Bcl-xl to associate with Beclin-1 complexes. Interestingly, the Beclin-1/Bcl-xl complex is notoriously difficult to detect by co-immunoprecipitation; our studies showed that in cells with high levels of ARF, the Beclin-1/Bcl-xl complex is undetectable. However, in these same cells, when ARF is silenced, the complex is readily detectable by immunoprecipitation followed by immunoblotting (Pimkina et al., 2009). While regulation of Beclin-1/Bcl-xl complex formation is one mechanism whereby ARF induces autophagy, the fraction of ARF that interacts with Bcl-xl in cells is low, and there are undoubtedly other mechanisms remaining to be identified.

6.4 The p53 Tumor Suppressor and Autophagy: p53 Induces Autophagy The groups of Levine and Jin were the first to demonstrate that the p53 tumor suppressor regulates autophagy. Activation of p53 by DNA damaging agents led to inhibition of mammalian target of rapamycin (mTOR) and induction of autophagy, as assessed by autophagosome formation and LC3 lipidation and cleavage (Feng et al., 2005). There were two mechanisms proposed for the inhibition of mTOR activity by p53: first, that in a p53-dependent manner, genotoxic stress induced the activation of the enzyme AMPK, which is a kinase that normally responds to metabolic stress. This increase in AMPK activity was not as impressive as that induced by glucose starvation, but it correlated well with the decrease in mTOR induced by p53. Notably, the authors showed that compound C, which inhibits AMPK, completely alleviated the inhibition of mTOR by p53, supporting the premise that p53’s negative regulation of mTOR was via AMPK (Feng et al., 2005). These authors later showed that in some cell types p53 induction led to the transcriptional activation of PTEN, TSC2, and AMPK β1, which negatively regulate mTOR; this effect of p53 on mTOR was estimated to occur in a slower time frame than activation of AMPK by p53, which takes minutes to hours (Feng et al., 2005, 2007). More recently Budanov and Karin showed that the products of two well-defined p53 target genes, Sestrin1 and Sestrin2, can bind and activate AMPK, causing it to phosphorylate TSC2 and thereby inhibit mTOR (Budanov and Karin, 2008). The combined data indicate that p53 uses multiple avenues in order to inhibit mTOR and subsequently induce autophagy.

6.5 p53 Transactivates the Autophagy Gene DRAM Another mechanism whereby p53 induces autophagy emerged with the discovery by the group of Ryan that p53 directly transactivates the gene encoding DRAM (damage-regulated autophagy modulator). DRAM encodes a highly conserved lysosomal protein with six putative transmembrane domains (Crighton et al., 2006).

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p53 binds directly to a consensus p53-binding site in the DRAM promoter following genotoxic stress. Notably, silencing DRAM was shown to markedly inhibit p53-mediated apoptosis, while overexpression of DRAM alone had limited cytotoxic effect on cells. Therefore, DRAM was found to be necessary but not sufficient for programmed cell death mediated by p53. Interestingly, in support of a role for autophagy in p53-dependent apoptosis, the authors found that silencing ATG5 likewise inhibited apoptosis induced by p53. A role for autophagy in p53-mediated apoptosis is likewise supported by the findings of Vousden and colleagues, who showed that the p53 target genes PUMA and Bax induce mitophagy (autophagy of mitochondria) in response to mitochondrial perturbations, and that inhibition of PUMA- or Bax-induced mitophagy dampens the apoptotic response (Yee et al., 2009). These data firmly support a role for autophagy in the successful execution of apoptosis. Why autophagy might be required for the completion of apoptosis is not presently clear.

6.6 Nutrient Stress Signals to p53 The groups of Jin and Thompson first showed that nutrient deprivation signals to p53. As early as 20 min after glucose deprivation, p53 becomes phosphorylated on serine 15, a hallmark signal of p53 activation (Feng et al., 2005; Jones et al., 2005). This phosphorylation is mediated by the kinase AMPK, which is thought to serve as a fuel sensor in the cell by assessing the ratio of AMP to ATP. Phosphorylation of p53 induces a p53-dependent cell cycle arrest, indicating that like genotoxic stress, nutrient stress may also use the p53 pathway in order to achieve cessation of cell proliferation. Whereas Thompson and colleagues showed that p53, along with phosphorylation on serine 15, was essential for AMPK-mediated growth arrest in low glucose, it remains unclear whether p53 is a direct or indirect target of AMPK, and it seems likely that there is an intermediate kinase involved. Interestingly, Thompson’s group pursued the connection between nutrient stress and p53 further and showed recently that the anti-diabetes drug metformin and the AMPK-activating drug AICAR both effectively diminish the growth of p53–/– tumors, but not matched p53+/+ counterparts (Buzzai et al., 2007).

6.7 p53 Negatively Regulates Autophagy in Unstressed Cells Seemingly in contrast to the abundant data that p53 induces autophagy in stressed cells, Kroemer and colleagues reported that p53 inhibits autophagy in unstressed cells. This group showed that silencing of p53 or pharmacological inhibition of this protein led to an increase in the steady-state level of basal autophagy, in both mammalian cells and in Caenorhabditis elegans (Tasdemir et al., 2008a, b). Consistent with this premise, certain tissues of the p53 knockout mouse, such as the liver, pancreas, and kidney, have increased basal levels of autophagy compared

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to wild-type mice. Similarly, p53–/– MEFs survive more efficiently in response to metabolic stress compared to wt MEFs (Tasdemir et al., 2008a, b). This group found that autophagy induced by p53 silencing occurs only in cells in the G1 and S phases of the cell cycle (Tasdemir et al., 2008c). Further, they showed that silencing p53 in C. elegans increases the life span of this organism by causing increased autophagy (Tavernarakis et al., 2008). Somewhat surprisingly, the Kroemer group obtained data supporting a role for cytoplasmic p53 in the inhibition of autophagy. Using overexpressed mutant forms of p53 in transfected cells, this group found that the nuclear export sequence (NES) of p53 is required for its ability to inhibit autophagy, although whether the NES mutant was otherwise functional or capable of oligomerizing appropriately was not tested (Tasdemir et al., 2008a). More surprisingly, this group reported that some tumor-derived point mutants of p53 could likewise inhibit autophagy; again in transfected cells, this group found that tumorderived mutant forms of p53 that typically localize to the nucleus (such as R282W) fail to suppress autophagy, while mutants that consistently localize to the cytosol (such as R273H) were highly efficient autophagy inhibitors (Morselli et al., 2008). Unclear from these studies is whether tumor cell lines engineered to stably express these mutants differ in the basal level of autophagy or whether this was an effect of acute p53 overexpression. A notable difference between many of the mutant forms of p53 that varied in autophagy suppression is that several of the mutants incapable of inhibiting autophagy are typically globally denatured (such as R282W), while those that can inhibit autophagy are largely native in conformation (such as R273H). These findings may have interesting implications for our understanding of why different tumor types tend to select for different “hotspot” mutations in p53, particularly in lieu of the findings that different tumor types may have different sensitivity to autophagy inhibition.

6.8 Conclusions and Future Directions The findings presented here on the role of p53 and ARF in autophagy are at first blush somewhat contradictory. For example, stress-activated p53 induces autophagy, while “unstressed” p53 represses basal levels of autophagy (Fig. 6.1). How these findings are reconciled is not presently known. One clue may be that p53, particularly the p53-induced genes PUMA and Bax, may selectively induce mitochondrial autophagy (mitophagy), and this may allow for the release of mitochondrial inducers of apoptosis such as cytochrome c more effectively (Yee et al., 2009). In contrast Kroemer’s group found that silencing IREα, which controls one arm of the endoplasmic reticulum (ER) stress pathway, effectively inhibits the ability of p53 to suppress autophagy; in other words, p53 may suppress the ER stress pathway, and silencing of p53 may predominantly signal ER stress and concomitant autophagy. As such, more careful delineation of the types of autophagy being induced (that is, mitophagy versus reticulophagy versus non-specific autophagy) along with the upstream signaling pathway (such as nutrient deprivation, ER stress, or genotoxic

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Fig. 6.1 The mechanisms whereby p53 and ARF, in stressed and unstressed cells, regulate autophagy

stress) may clarify some of these apparent discrepancies. The contribution of ARF to the autophagy pathway(s) regulated by p53 is unclear: Because the majority of tumor cell lines inactivate either p53 or ARF, this question can only be addressed in primary mouse embryo fibroblasts or in tissues from the p53 knockout mouse, compared to the p53/ARF double knockout. Additionally, how ARF induces autophagy and the rules that govern whether ARF-mediated autophagy is cytoprotective or cytotoxic need to be delineated. The combined data suggest that the p53 and ARF tumor suppressors are very intimately tuned into the metabolic state of the cell, perhaps even more so than to the DNA damage pathway. It is tempting to speculate that the functions of p53 and ARF originally evolved to monitor the metabolic state of the cell and to induce growth arrest, autophagy, or apoptosis in response to nutrient signals; these pathways may then have been adapted to monitor genotoxic stress.

References Abida WM, Gu W (2008) p53-dependent and independent activation of autophagy by ARF. Cancer Res 68: 352–357. Budanov AV, Karin M (2008) p53 target genes sestrin1 and sestrin2 connect genotoxic stress and mTOR signaling. Cell 134: 451–460. Buzzai M, Jones RG, Amaravadi RK, Lum JJ, DeBerardinis RJ, Zhao F, Viollet B, Thompson CB (2007) Systemic treatment with the antidiabetic drug metformin selectively impairs p53-deficient tumor cell growth. Cancer Res 67: 6745–6752. Crighton D, Wilkinson S, O’Prey J, Syed N, Smith P, Harrison PR, Gasco M, Garrone O, Crook T, Ryan KM (2006) DRAM, a p53-induced modulator of autophagy, is critical for apoptosis. Cell 126: 121–134. Eischen CM, Weber JD, Roussel MF, Sherr CJ, Cleveland JL (1999) Disruption of the ARFMdm2-p53 tumor suppressor pathway in Myc-induced lymphomagenesis. Genes Dev 13: 2658–2669.

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Feng Z, Hu W, de Stanchina E, Teresky AK, Jin S, Lowe S, Levine AJ (2007) The regulation of AMPK beta1, TSC2, and PTEN expression by p53: stress, cell and tissue specificity, and the role of these gene products in modulating the IGF-1-AKT-mTOR pathways. Cancer Res 67: 3043–3053. Feng Z, Zhang H, Levine AJ, Jin S (2005) The coordinate regulation of the p53 and mTOR pathways in cells. Proc Natl Acad Sci USA 102: 8204–8209. Gil J, Peters G (2006) Regulation of the INK4b-ARF-INK4a tumour suppressor locus: all for one or one for all. Nat Rev Mol Cell Biol 7: 667–677. Humbey O, Pimkina J, Zilfou JT, Jarnik M, Dominguez-Brauer C, Burgess DJ, Eischen CM, Murphy ME (2008) The ARF tumor suppressor can promote the progression of some tumors. Cancer Res 68: 9608–9613. Itahana K, Zhang Y (2008) Mitochondrial p32 is a critical mediator of ARF-induced apoptosis. Cancer Cell 13: 542–553. Jones RG, Plas DR, Kubek S, Buzzai M, Mu J, Xu Y, Birnbaum MJ, Thompson CB (2005) AMPactivated protein kinase induces a p53-dependent metabolic checkpoint. Mol Cell 18: 283–293. Kelly-Spratt KS, Gurley KE, Yasui Y, Kemp CJ (2004) p19Arf suppresses growth, progression, and metastasis of Hras-driven carcinomas through p53-dependent and -independent pathways. PLoS Biol 2: E242. Maiuri MC, Le Toumelin G, Criollo A, Rain JC, Gautier F, Juin P, Tasdemir E, Pierron G, Troulinaki K, Tavernarakis N et al. (2007) Functional and physical interaction between Bcl-X(L) and a BH3-like domain in Beclin-1. EMBO J 26: 2527–2539. Morselli E, Tasdemir E, Maiuri MC, Galluzzi L, Kepp O, Criollo A, Vicencio JM, Soussi T, Kroemer G (2008) Mutant p53 protein localized in the cytoplasm inhibits autophagy. Cell Cycle 7: 3056–3061. Pattingre S, Tassa A, Qu X, Garuti R, Liang XH, Mizushima N, Packer M, Schneider MD, Levine B (2005) Bcl-2 antiapoptotic proteins inhibit Beclin 1-dependent autophagy. Cell 122: 927–939. Pimkina J, Humbey O, Zilfou JT, Jarnik M, Murphy ME (2009) ARF induces autophagy by virtue of interaction with and inhibition of Bcl-xl. J Biol Chem 284: 2803–2810. Pimkina J, Murphy ME (2009) Arf, autophagy and tumor suppression. Autophagy 5: 1–3. Quelle DE, Zindy F, Ashmun RA, Sherr CJ (1995) Alternative reading frames of the INK4a tumor suppressor gene encode two unrelated proteins capable of inducing cell cycle arrest. Cell 83: 993–1000. Reef S, Kimchi A (2008) Nucleolar p19ARF, unlike mitochondrial smARF, is incapable of inducing p53-independent autophagy. Autophagy 4: 866–869. Reef S, Shifman O, Oren M, Kimchi A (2007) The autophagic inducer smARF interacts with and is stabilized by the mitochondrial p32 protein. Oncogene 26: 6677–6683. Reef S, Zalckvar E, Shifman O, Bialik S, Sabanay H, Oren M, Kimchi A (2006) A short mitochondrial form of p19ARF induces autophagy and caspase-independent cell death. Mol Cell 22: 463–475. Sherr CJ, Bertwistle D, Den Besten W, Kuo ML, Sugimoto M, Tago K, Williams RT, Zindy F, Roussel MF (2005) p53-dependent and -independent functions of the Arf tumor suppressor. Cold Spring Harb Symp Quant Biol 70: 129–137. Sugimoto M, Kuo ML, Roussel MF, Sherr CJ (2003) Nucleolar Arf tumor suppressor inhibits ribosomal RNA processing. Mol Cell 11: 415–424. Tago K, Chiocca S, Sherr CJ (2005) Sumoylation induced by the Arf tumor suppressor: a p53independent function. Proc Natl Acad Sci USA 102: 7689–7694. Tasdemir E, Chiara Maiuri M, Morselli E, Criollo A, D’Amelio M, Djavaheri-Mergny M, Cecconi F, Tavernarakis N, Kroemer G (2008b) A dual role of p53 in the control of autophagy. Autophagy 4: 810–814. Tasdemir E, Maiuri MC, Galluzzi L, Vitale I, Djavaheri-Mergny M, D’Amelio M, Criollo A, Morselli E, Zhu C, Harper F, Nannmark U, Samara C, Pinton P, Vicencio JM, Carnuccio R, Moll UM, Madeo F, Paterlini-Brechot P, Rizzuto R, Szabadkai G, Pierron G, Blomgren K,

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Part III

Long-Term Proliferation

Chapter 7

Regulation of Self-Renewing Divisions in Normal and Leukaemia Stem Cells Andrea Viale and Pier Giuseppe Pelicci

Abstract The identification of a rare population of tumour cells capable of transplanting the disease in animal models has radically modified our perception of the biological organization of tumours. These cells, named tumour-initiating cells (TICs), or cancer stem cells (CSCs), share phenotypic and functional characteristics with normal stem cells, most notably their ability to self-renew. Here we discuss regulation of self-renewal in normal haematopoietic and leukaemic stem cells, focusing on the role of cell quiescence and inhibitors of the cell cycle. In the last decade the identification of a rare population of tumour cells capable of transplanting the disease in animal models has radically modified our perception of the biological organization of tumours. These cells, named tumour-initiating cells (TICs), or cancer stem cells (CSCs) because they share phenotypic and functional characteristics with normal stem cells, have been initially identified in the myeloid acute leukaemias (Lapidot et al., 1994), and only more recently in solid tumours (Al-Hajj et al., 2003; Collins et al., 2005; Kim et al., 2005; Li et al., 2007; O’Brien et al., 2007; Ricci-Vitiani et al., 2007; Schatton et al., 2008; Singh et al., 2004). Their discovery in tumours of various histotypes has reinforced the hypothesis that cancer is in reality an anomalous tissue recapitulating the hierarchical organization found in the normal tissue from which it originates. In this model, a rare population of stem cells (SCs) would then be a valuable source of progenitor cells in continuous expansion, able to generate more differentiated cells and initiate tumours upon transplantation into a new host (Bonnet and Dick, 1997). In the same way as normal haemopoiesis has been the paradigm for studies on adult stem cells, acute myeloid leukaemias are the paradigm for investigations on cancer stem cells (Huntly and Gilliland, 2005). Indeed, a huge amount of data is now available with regard to both the normal and the leukaemic haemopoietic system, which will be the focus of our review. A. Viale (B) Department of Experimental Oncology, European Institute of Oncology at IFOM-IEO-Campus, Milan, Italy e-mail: [email protected]; [email protected]


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7.1 Self-Renewal Potential of Normal Haematopoietic Stem Cells Is Limited Haemopoiesis is a tightly regulated process which controls the continuous replacement of differentiated blood cells. It has been estimated that throughout one’s life the haematopoietic system produces more than 1016 cells. This process relies on the presence of rare haematopoietic stem cells (HSCs) that divide giving rise to highly proliferating progenitor cells, sometimes known as transit-amplifying cells, which after several cycles of division become terminally differentiated. Thus, long-lived multicellular organisms have developed a strategy which leads to the amplification of a relatively small number of HSCs into a huge numbers of progeny. The unique ability of a stem cell to originate two daughter cells with different replicative and differentiative potentials (one transit-amplifying progenitor and a new stem cell) is known as self-renewal. Because stem cells maintain tissue homeostasis throughout the lifespan, one could assume that they have, in contrast to other somatic cells, an infinite replication potential. However, since the 1960s–1970s, it has been clear that during transplantation of bone marrow from a mouse to another, an assay known as serial transplantation, the ability of HSCs to sustain long-term multilineage reconstitution of the host is limited to ∼5–6 rounds (Harrison, 1979; Harrison and Astle, 1982; Harrison et al., 1978), suggesting that HSCs have a limited replication potential. Recovering the entire haemopoiesis in irradiated mice is in fact a big challenge for transplanted HSCs. Not only are they immediately induced to proliferate after transplantation to quickly regenerate the damaged tissue, but a conspicuous fraction continues to proliferate for longer period, at least 4 months (Allsopp et al., 2001). Perpetuation of this hyperproliferative state leads ultimately to functional exhaustion of the HSCs. The number of divisions that one HSC can undergo during the lifespan of a mammal is relatively constant (∼80–200) and evolutionarily conserved (Shepherd et al., 2007); as a result of more frequent divisions, a stem cell reaches its replicative ceiling much sooner, loosing the ability to self-renew.

7.2 Haematopoietic Stem Cells Are Deeply “Dormant” Another supposed hallmark of SCs, as compared to their more differentiated progenitors, is that of infrequent division, often referred to as “quiescence” or “G0”. A variety of evidence supports this idea. For example, several groups described HSCs as resistant to diverse cell cycle specific cytotoxic agents such as 5-fluorouracil, that eliminates rapidly dividing cells while sparing non-cycling (long-term reconstituting) stem cells (Hodgson and Bradley, 1979; Lerner and Harrison, 1990; Randall and Weissman, 1997). Due to the growing interest in stem cells, different assays have been developed to address the issue of quiescent or “resting” cells in vivo. Amongst these, the

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method of direct evaluation of total RNA, hypothesizes that quiescent cells have less RNA than their proliferating counterparts (Darzynkiewicz et al., 1979). Indeed, detection in a stem cell population of both RNA and DNA content (using RNAspecific flourochrome Pironin Y and DNA-specific dye Hoechst) demonstrated that 75% of HSCs have low levels of RNA and DNA, compatible with a quiescent state (Cheshier et al., 1999). Another method most frequently used to analyse in vivo cell cycle status uses the 5 -bromo-2 -deoxyuridine (BrdU). BrdU, a synthetic nucleoside analogous to thymidine and incorporated in newly synthesized DNA during replication, has been extensively used to study both proliferation (after acute incorporation) and quiescence. The latter requires exposure of mice to BrdU for long intervals of time (pulse), followed by several months of no BrdU intake (chase); the cells that retain BrdU, known as label retaining cell (LRC), are supposed to be “quiescent” or slowly proliferating cells. Using these approaches authors have demonstrated that HSCs proliferate slowly: ∼8% enters the cell cycle per day but in due course all HSCs are recruited into cycle with a frequency of division of around 2 months (Cheshier et al., 1999). These experimental evidences are coherent with a model of haemopoiesis in which the haemopoietic SC compartment consists of a homogeneous population of cells that divide asynchronously and infrequently, and where quiescence is not a functional defined state in which stem cells are “frozen,” but rather a simple period of time between divisions, however long it may be. One of the biggest obstacles to studying the cell cycle of HSCs is their extreme rarity (3 μM (Fry et al., 2004). Similarly, in mantle cell lymphoma and multiple myeloma primary cells and cell lines, characterized

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Targeting Cyclin-Dependent Kinases for Cancer Therapy

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by cyclin D1 overexpression resulting from genomic rearrangement, PD0332991 also induces cytostatic G1 arrest without apoptosis (Baughn et al., 2006; Marzec et al., 2006). Additionally, PD0332991 has demonstrated efficacy in a broad spectrum of human tumor xenografts. In vivo activity is also Rb-dependent, with little effect against Rb-deficient xenografts (Fry and Garrett, 2000). Among breast, glioblastoma, non-small cell lung cancer, and prostate carcinoma xenografts, nearcomplete stasis was observed. Interestingly, in mice bearing Colo-205 colon cancer xenografts, PD0332991 produced rapid tumor regression. The mechanism for this effect remains unclear. It has been postulated that in some cases, the xenograft represents a balance of proliferating and apoptotic cells, and inhibition of the proliferative compartment allows naturally dying cells to predominate. Alternatively, inhibition of cdk4/6 activity via the p16INK4A tumor suppressor protein has been shown to reduce VEGF expression and compromise angiogenic signaling (Harada et al., 1999), observations that may account for effects of a cdk4/6 inhibitor compound observed in vivo that did not occur in vitro. Additionally, it has also been demonstrated in SW480 colon cancer cells that selective cdk4 inhibition causes degradation of the NFκB suppressor protein, IκB, and induces translocation of RelA from cytoplasm to nucleoplasm and eventually to the nucleolus. The nucleolar translocation of RelA is accompanied by repression of NFκB-driven transcription as well as apoptosis, providing a mechanism by which a cdk4 inhibitor may induce apoptosis in certain cell types (Thoms et al., 2007). It is possible that the association of cdk4 inhibition with repression of NFκB activity also contributes to the synergism of PD0332991 with bortezomib (Menu et al., 2008) and dexamethasone (Baughn et al., 2006) recently reported in multiple myeloma cells. PD0332991 has completed Phase 1 testing in advanced solid tumors with documented expression of Rb. Dose-limiting toxicities included neutropenia and thrombocytopenia, so that treatment was administered 21 of every 28 days or 14 of every 21 days. Nineteen of 74 patients had stable disease ≥16 weeks, including patients with breast, colon, and ovarian carcinoma who remained on trial >20 months (O’Dwyer et al., 2007). In the context of this trial, three patients with growing teratoma syndrome were treated. Each of the mature teratomas expressing high levels of the Rb protein was inoperable and chemotherapy- and radiation-resistant. These patients achieved either partial response or stable disease lasting at least 18 months (Vaughn et al., 2009). Additionally, in a preliminary report of a pharmacodynamic study of PD0332991 performed in mantle cell lymphoma, 8 of 17 heavily pre-treated patients achieved partial response or stable disease lasting up to 60 weeks. PD0332991 treatment caused reduced 18 F-fluoro-3 -deoxy-3 -L-fluorothymidine (FLT) uptake on positron emission tomography (PET) scans, consistent with G1 arrest, as well as reduced phosphorylation of Rb at cdk4 sites, documented immunohistochemically in lymph node biopsies obtained before and during treatment (Leonard et al., 2008). Mechanisms of resistance have also been investigated, illustrated by the effects of PD0332991 in acute myeloid leukemia (AML) cell lines. In Flt3 internal tandem duplication (ITD)-expressing cell lines, PD0332991 induces sustained cell cycle

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arrest, followed in some cases by apoptosis. In contrast, in Flt3 wild-type AML cells, PD0332991 induced an initial G1 arrest that was partially overcome after 120 h through the downregulation of p27Kip1 and the reactivation of cdk2 (Wang et al., 2007). This is similar to the cell cycle progression observed in cdk4null/null /cdk6–/– mouse embryonic fibroblasts, in which cdk2 compensates for the loss of the other G1 cdks in order to facilitate cell cycle progression (Malumbres et al., 2004). Therefore, in some cases, agents that inhibit both cyclin D-dependent kinases and cyclin E–cdk2 complexes may produce stronger G1 arrest phenotypes with delayed emergence of resistance. Alternative mechanisms of cdk2 activation or G1 progression may well occur in other cellular backgrounds, so that the development of cell line derivatives resistant to PD0332991 is likely to be instructive.

11.3 Targeting Cdk2 and Cdk1 Cdk2 was initially considered to be a therapeutically relevant candidate for anticancer drug design emanating from experiments that demonstrated growth arrest or apoptosis when cells were treated with a dominant-negative mutant (Hu et al., 2001), cdk2 inhibitory peptides (Chen et al., 1999), targeted cyclin A degradation (Chen et al., 2004), or ectopic expression of p27Kip1 (Wang et al., 1997). Consequently, many compounds have since been developed with cdk2 inhibitory activity. However, initial experiments did not take into account the effects of these treatments on cdk1 activity. Ectopic p27Kip1 expression or loss of cyclin Adependent kinase activity was likely to inhibit both cdk2 and cdk1; expression of the dominant-negative mutant also reduced cdk1 activity, and reported cdk2 inhibitory peptides were capable of cdk1 inhibition at high concentration. Therefore, it is not altogether surprising that experiments with embryonic fibroblasts from cdk2–/– knockout mice (Berthet et al., 2003; Ortega et al., 2003), as well as tumor cells subjected to antisense- or siRNA-mediated cdk2 depletion (Cai et al., 2006; Tetsu and McCormick, 2003), have demonstrated either absent or minor defects in cell cycle proliferation, including only slight slowing of the G1/S transition in synchronized cells. In cdk2-depleted cells, cyclin E–cdk1 complexes (Aleem et al., 2005), as well as cyclin D-dependent kinase activity, compensate during the G1/S transition, while cyclin A–cdk1 complexes govern S and G2 progression (Cai et al., 2006; L’Italien et al., 2006). One exception has been found in melanoma, where there is correlation in expression between the microphthalmia-associated transcription factor (MITF) and cdk2. Melanomas with low MITF expression have low levels of cdk2 and are particularly susceptible to G1 arrest induced by siRNA-mediated cdk2 depletion (Du et al., 2004). In contrast to results for other cell cycle cdks, genetic ablation of cdk1 in the mouse germline results in very early embryonic lethality, indicating cdk1 is required for cell division during the early stages of embryonic development (Santamaria et al., 2007). However, no conditional cdk1 knockout strain is available, and the effects of cdk1 ablation on different tissue types or mature adult cells have not been assessed. Nonetheless, consistent with data in knockout mice, shRNA-mediated depletion of cdk1 from non-transformed retinal pigment epithelial cells leads to

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potent G2 arrest (Johnson et al., 2009). In contrast, depletion of cdk1 from cancer cells results in the formation of novel cyclin B–cdk2 complexes, so that cells are able to proliferate with only slight slowing of the G2–M transition (Cai et al., 2006; L’Italien et al., 2006). Therefore, effects of cdk ablation in embryonic mouse systems may not always predict outcomes in human cancer cells. Although in most instances, individual depletion of cdk2 or cdk1 cannot completely block cancer cell proliferation, dual cdk2 and cdk1 depletion or inhibition produces S phase retardation and G2 cell cycle arrest, and in some instances cell death (Cai et al., 2006; L’Italien et al., 2006; Payton et al., 2006). The ability of cdk2 and cdk1 to easily compensate for one another in transformed cells should guide the prioritization of compounds with equipotency against cdk2 and cdk1 over those identified in cdk2-selective screens.

11.4 Combined Targeting of Cdks and Anti-apoptotic Proteins 11.4.1 Transcriptional Cdk Inhibition Several cdk family members play a role in mRNA transcription and processing. In addition to its contribution to cdk-activating kinase (CAK) activity, cdk7 phosphorylates Ser5 of the heptapeptide repeat of the C-terminal domain (CTD) of RNA polymerase II. Cdk9, in complex with T cyclins (P-TEFb), phosphorylates the CTD preferentially at Ser2. These events are required for transcriptional initiation and the elongation of nascent mRNAs, respectively (Meinhart et al., 2005). Inhibition of cdk9 has largely been studied with potent inhibitory compounds, including flavopiridol and seliciclib (Shapiro, 2006). The interaction of flavopiridol with the cdk9 ATP-binding site occurs at a Ki of 3 nM, such that it is difficult to demonstrate competition by ATP (Chao et al., 2000; Chao and Price, 2001; de Azevedo et al., 2002). Flavopiridol has profound effects on cellular transcription, causing depletion of labile mRNAs with short half-life, including those encoding cyclin D1, c-Myc, mitotic regulatory kinases, and Mdm2 (Lam et al., 2001; Lu et al., 2004). Additionally, transcripts encoding Mcl-1 and XIAP are rapidly depleted following cdk9 inhibition (Lam et al., 2001). Depletion of Mcl-1 may contribute, in part, to the activity of flavopiridol recently demonstrated in chronic lymphocytic leukemia (Byrd et al., 2007; Chen et al., 2005) and has prompted the evaluation of cdk9 inhibitors in other Mcl-1-dependent hematologic malignancies, including multiple myeloma and mantle cell lymphoma (Gojo et al., 2002; Kouroukis et al., 2003; Venkataraman et al., 2006). The depletion of anti-apoptotic proteins such as Mcl-1 and XIAP via cdk9 depletion or inhibition has also induced apoptosis in solid tumor cell lines in which cell cycle progression has been disrupted by combined inhibition of cdk2 and cdk1 (Cai et al., 2006). In this scenario, cdk9 inhibition lowers the apoptotic threshold and facilitates cell death from the S and G2 phases. Therefore, cdk2, cdk1, and cdk9 may together represent a promising subset of the cdk family for drug targeting. Several such compounds with cell cycle and transcriptional cdk inhibitory activity have been described, including AZD5438 (Byth et al., 2009), SNS-032 (Conroy

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et al., 2009; Heath et al., 2008), and SCH727965, the latter compound a novel pyrazolo[1,5-α]pyrimidine that has demonstrated promise in early-phase clinical trials (Nemunaitis et al., 2009; Parry et al., 2007; Shapiro et al., 2008). Seliciclib, which also targets cell cycle and transcriptional cdks, has recently demonstrated activity in nasopharyngeal carcinoma, with evidence of reduced cdk2-mediated phosphorylation of Rb, as well as reduced transcription of proliferation and survival genes, with decreased expression of cyclin D1 and Mcl-1, consistent with cdk9 inhibition. Seven of 14 evaluable patients had clinical evidence of tumor reduction; some responses were associated with increased tumor apoptosis, necrosis, and decreases in plasma EBV DNA post-treatment (Hsieh et al., 2009). Finally, a selective cdk7 inhibitor has recently been described with preclinical activity in vitro and in vivo related to both cell cycle and transcriptional cdk inhibitory effects (Ali et al., 2009). In some cell types, it is possible that cdk2 and cdk1 may contribute to phosphorylation of the CTD of RNA polymerase II. For example, in U2OS osteosarcoma cells, combined cdk2 and cdk1 depletion alone resulted in reduced RNA polymerase II phosphorylation, XIAP depletion, and apoptosis (Cai et al., 2006). Cdk2 depletion has also been shown to induce apoptosis in human diffuse large cell lymphoma cells, with concomitant reduction in Mcl-1 expression (Faber and Chiles, 2007).

11.4.2 Survivin as a Target of Cdk1 Survivin is an inhibitor of apoptosis protein (IAP) phosphorylated by cyclin B-cdk1 and is rapidly degraded in the absence of phosphorylation by cdk1. MYCtransformed cells have been shown to be survivin dependent, and the combination of MYC overexpression and cdk1 inhibition causes synthetic lethality in both in vitro and in vivo models of MYC-induced lymphomas and hepatoblastomas (Goga et al., 2007). Similarly, survivin expression is critical for cancer cell survival after exposure to microtubule-disrupting agents (O’Connor et al., 2002; Wall et al., 2003). Following mitotic arrest induced by taxanes or vinca alkaloids, cdk1 inhibition results in compromised survivin expression and substantial apoptosis. This has led to the development of several sequential taxane–cdk inhibitor combinations (Shapiro, 2004).

11.5 Reduced Cyclin D1 Expression Mediated by Transcriptional Cdk Inhibition The cyclin D1 transcript also has short half-life and is depleted by cdk9 inhibition. Recent experiments have demonstrated in vitro and in vivo synergism of the anti-HER2 antibody trastuzumab, as well as small molecule EGFR inhibitors with inhibitors of cdk9, including flavopiridol and seliciclib (Fleming et al., 2008; Nahta et al., 2002; Wu et al., 2002). The combination of an ErbB targeting agent with a cdk9 inhibitor results in enhanced loss of expression of cyclin D1 compared to

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either drug alone, which in part accounts for synergistic anti-proliferative effects. Cyclin D1 is essential for HER2-mediated transformation (Yu et al., 2001) and is also a critical downstream effector of mutant EGFR (Kobayashi et al., 2006). In the case of non-small cell lung cancer cell lines, cooperativity was also observed in EGFR wild-type cells, suggesting the combination may expand the range of tumors that may benefit from EGFR inhibition (Fleming et al., 2008). These results also raise the possibility that direct cdk4/6 inhibition may synergize with ErbB pathway inhibitors (Yang et al., 2004; Yu et al., 2006).

11.6 Modulation of the p53 and p21Waf1/Cip1 by Transcriptional Cdk Inhibition Mdm2 represents a transcript targeted by cdk9 inhibition, resulting in elevated levels of p53 (Alonso et al., 2003; Demidenko and Blagosklonny, 2004). In cells particularly prone to p53-dependent apoptosis, including acute lymphocytic leukemia (Jackman et al., 2008) and germ cell tumor cells (Mayer et al., 2005), p53 accumulation may directly contribute to cell death. In addition, transcriptional repression can also prevent the induction of p21Waf1/Cip1 , so that p53 induction can occur without a concomitant rise in p21Waf1/Cip1 . The same is true after DNA damage in p53 wildtype cells. For example, in p53 wild-type colon carcinoma cells that arrest after CPT-11-mediated stimulation of p53 and p21Waf1/Cip1 , flavopiridol-mediated inhibition of the transcriptional induction of p21Waf1/Cip1 can contribute to cell death (Motwani et al., 2001). Additionally, cdk inhibitor-mediated suppression of p53independent induction of p21Waf1/Cip1 following histone deacetylase inhibition can convert the response from cytostatic to cytotoxic (Almenara et al., 2002). Interestingly, cdk2 is synthetically lethal to neuroblastoma cells harboring MYCN amplification. In these cells, selective cdk2 depletion results in apoptosis that is dependent on the continued expression of MYCN, as well as accumulation of wild-type p53 (Molenaar et al., 2009). While not yet formally tested, it is tempting to speculate that increased p53 levels result from reduced RNA polymerase II CTD phosphorylation that occurs following cdk2 depletion. Therefore, in these cells, as well as in lymphoma cells, in which cdk2 depletion is accompanied by Mcl-1 depletion (Faber and Chiles, 2007), it is possible that cdk2 contributes substantially to CTD phosphorylation.

11.7 Cdks and E2F-1 Activity 11.7.1 Cdk Inhibition and the Prevention of Neutralization of E2F-1 Activity During S Phase The cell cycle cdks regulate E2F-1 activity in a complex manner (Chapter 2 by Ji and Dyson). Following cdk-mediated phosphorylation of Rb during G1, E2F-1 activity is derepressed and E2F-1 is released, so that it can direct transcription

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of genes required for S phase. However, this transcription is activated only transiently. Orderly S phase progression requires the downregulation of E2F-1 activity, accomplished in part by phosphorylation mediated by both cdk2 and cdk1 (Dynlacht et al., 1994; Kitagawa et al., 1995; Krek et al., 1994; Peeper et al., 1995; Xu et al., 1994). E2F-1 also interacts with the cyclin H-cdk7 as a component of the TFIIH muti-subunit protein complex. Phosphorylation of E2F-1 by cyclin H-cdk7 is a prerequisite for ubiquitination and degradation (Vandel and Kouzarides, 1999). The targeting of E2F-1 phosphorylation via cdk inhibition may lead to the selective cell death of malignant cells. The disrupted cyclin D–cdk4/6–INK4–Rb pathway in tumor cells produces high levels of E2F-1 activity. A reduction in cdk activity during S phase may lead to persistence of E2F-1 activity that is inconsequential for normal cells, but may leave transformed cells with inappropriately persistent high-level E2F-1 activity that is sufficient to induce apoptosis (Chen et al., 1999; Jiang et al., 2003). E2F-1-dependent apoptosis following cdk inhibition may occur in a p73-dependent fashion (Chen et al., 2004). Alternatively, inappropriately persistent E2F-1 activity may suppress transcription of Mcl-1, which may also lead to apoptosis (Croxton et al., 2002). Therefore, in addition to suppression of Mcl-1 transcription by cdk9 inhibition, Mcl-1 expression may also be reduced by modulation of E2F-1 activity afforded by inhibition of cdks 2, 1, and 7 (Ma et al., 2003). Interestingly, baseline E2F-1 activity has recently distinguished multiple myeloma cells that undergo apoptosis in response to cdk inhibitor compounds from cells that are resistant (Eguchi et al., 2009). Cells lacking p18INK4C have higher baseline activity of E2F-1 than cells expressing p18INK4C , related to high cdk6 activity in p18INK4C -deficient cells. These cells, therefore, have lower baseline levels of Mcl-1 and more readily reach the threshold for mitochondrially-induced apoptosis after transcriptional cdk inhibition than cells with higher baseline levels of Mcl-1. Therefore, in this setting, the pretreatment level of p18INK4C serves as a surrogate to define E2F-1 activity and the predicted response to cdk inhibition.

11.7.2 Role of Cdk8 in Modulation of E2F-1 Activity E2F-1 has recently been shown to be a potent and specific inhibitor of β-catenin/Tcell factor-dependent transcription (Morris et al., 2008). Colorectal tumors that depend on β-catenin for abnormal proliferation thus select conditions that suppress E2F-1 and enhance the activity of β-catenin. These conditions include retention of wild-type Rb, as well as amplification of CDK8. Cdk8 is a member of the mediator complex, which serves to convey information from gene-specific regulatory proteins to the basal RNA polymerase II transcription machinery. Cdk8 represses E2F-1 activity, and elevated levels of cdk8 protect β-catenin-mediated transcription from inhibition by E2F-1 (Morris et al., 2008). Cdk8 has been shown to be a colorectal oncoprotein and its kinase activity is necessary for β-catenin-driven transformation. Suppression of cdk8 expression inhibits proliferation of colon cancer cells characterized by high levels of cdk8 and β-catenin hyperactivity (Firestein et al.,

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2008). Presumably, small molecule-mediated inhibition of cdk8 or inhibition of cdks 2, 1, or 7 may also serve to activate E2F-1 activity and suppress β-catenin-mediated gene expression in colorectal cancer cells.

11.8 Cdk Inhibition and DNA Damage 11.8.1 Induction of DNA Damage by Cdk Inhibition Cdk inhibition during S phase, mediated by pharmacologic inhibitors, dn-cdk2, and siRNA targeting cdk2, elicits an intra-S phase checkpoint that shares components of the pathway activated by double-strand DNA breaks (Zhu et al., 2004). In A2780 ovarian cancer cells, cdk inhibition has been shown to induce the accumulation of activated forms of the phosphatidylinositol 3-kinase family member ataxiatelangiectasia mutated (ATM) and the checkpoint kinase Chk2, as well as nuclear foci containing phosphorylated substrates of ATM, including p53 and histone H2AX (γ-H2AX), suggesting that cdk inhibition can induce or predispose to DNA damage. Cdk2 phosphorylates a number of proteins during S phase, including minichromosome maintenance proteins (MCM), which are required for origin replication firing. Cdk2 inhibition during DNA replication leads to increased MCM complex association with DNA and triggers the rereplication of DNA. Overreplication likely results in the formation of DSB and ssDNA intermediates, which activate ATM and ATR, and subsequently p53 (Zhu et al., 2004). An alternative mechanism by which cdk inhibition could mediate DNA damage appears related to the reduced expression of Chk1 (Enders, 2008; Maude and Enders, 2005). Chk1, along with Chk2, is typically activated by DNA damage in order to constrain cdk activity. However, following prolonged (i.e., 24 h) cdk inhibition during S phase, the response to cell cycle slowing involves downregulation of Chk1, perhaps part of a negative feedback loop promoting cell cycle recovery. Reduced cdk activity may slow or stall DNA replication forks. This block in replication is detected by ATR (ATM and Rad3-related), which primarily activates Chk1. Stalled replication forks are dependent on the ATR–Chk1 pathway for stabilization; when Chk1 activity is compromised double-strand breaks may occur. This mechanism may, in part, explain the increased cytotoxicity and frequency of double-strand DNA breaks observed when cdk inhibitors are combined with DNA-damaging treatments such as topoisomerase inhibitors (Crescenzi et al., 2005; Maude and Enders, 2005).

11.8.2 Cdk Inhibition in DNA Damage-Induced Checkpoint Control In addition to the replicative stress and DNA damage response elicited by cdk inhibition during S phase, cdks have been implicated in checkpoint control following

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exposure to standard DNA-damaging agents. Cdk1 and cdk2 are inhibited downstream in the DNA damage pathway, affording cell cycle arrest and time for DNA repair processes to occur. However, emerging evidence indicates that cdks play critical roles upstream in the initiation of checkpoint control, prior to the terminal events in the checkpoint cascade when their activities are inhibited. Following double-stranded DNA breaks, ATM is recruited by the MRN (Mre11–Rad50–Nbs1) complex (Lee and Paull, 2005). MRN, together with the endonuclease CtIP (Sartori et al., 2007), carries out the process of end resection in order to produce regions of single-stranded DNA. Single-stranded DNA is coated by RPA proteins, which serve to recruit ATR (Jazayeri et al., 2006; Zou and Elledge, 2003). In budding yeast, cdk1 (cdc28) is required for activation of DNA end resection and ultimately the Mec1 (ATR homolog)-dependent DNA damage checkpoint following a doublestrand break (Ira et al., 2004). In mammalian cells, inhibition or depletion of cdk1 alone did not affect end resection, likely because of compensation by cdk2. However, brief exposure to small molecule-mediated combined inhibition of cdk2 and cdk1 in concert with DNA damage did compromise end resection (Johnson et al., 2009). Activation of endonuclease activity requires cdk-mediated phosphorylation of CtIP at Thr 847, analogous to events that occur in yeast (Huertas and Jackson, 2009). Cdk inhibition therefore compromises ATR recruitment and the phosphorylation of Chk1, necessary to initiate checkpoint signaling cascades in response to DNA damage, and ultimately may sensitize cells to DNA-damaging treatments. DNA end resection is just one level at which cdk activity regulates the DNA damage response. When cells are treated with hydroxyurea (HU), which results in DNA end resection-independent ssDNA break formation and direct ATR activation, Chk1 phosphorylation is also abrogated with cdk inhibition. Recent studies have demonstrated that Chk1 and other ATR and ATM targets are not phosphorylated as efficiently when cdk1 is depleted or inhibited in non-small cell lung cancer (NSCLC) cell lines (Johnson et al., 2009). Loss of DNA damage-induced checkpoint control was caused by a reduction in formation of BRCA1-containing foci. Furthermore, expression in BRCA1-deficient cells of BRCA1 mutated at cdk phosphorylation sites S1497 and S1189/S1191 resulted in compromised formation of BRCA1-containing foci compared to those formed in cells engineered to express wild-type BRCA1. ATR- and ATM-mediated phosphorylation of nonchromatin-bound proteins is dependent on BRCA1 focus formation (Foray et al., 2003; Yarden et al., 2002); therefore, when cdk1 is inhibited and BRCA1 foci formation is reduced, Chk1 and other BRCA1-dependent ATM/ATR substrates are not phosphorylated (Fig. 11.1). Selective cdk1 depletion or inhibition also sensitized NSCLC cells to DNA-damaging agents. Importantly, this sensitization only occurred in transformed cells. In contrast to transformed cells, which continued to proliferate in the absence of cdk1 because of compensation by cdk2, non-transformed cells underwent potent G2 arrest in response to cdk1 depletion, which antagonized the response to subsequent DNA damage (Johnson et al., 2009).

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Fig. 11.1 Cdk activity is required to initiate the DNA damage response. Although cdks have traditionally been considered to be the downstream targets of DNA damage-induced checkpoint cascades, cdk activity remains elevated initially after exposure to DNA damage. Cdks participate in several upstream processes. Cdk5 phosphorylates ATM at S794. Cdks also phosphorylate CtIP, necessary for DNA end resection following a double-strand break (not shown). Cdk2 phosphorylates ATRIP at S224. Additionally, cdk1 phosphorylates BRCA1 at S1497 and S1189/S1191, events required for efficient BRCA1 focus formation. Therefore, cdk activity facilitates the ATMand ATR-dependent phosphorylation of non-chromatin bound substrates, including Chk1 and Chk2. Eventually, activation of checkpoint kinases leads to inhibition of cdk1 and cdk2 activities, permitting cell cycle arrest. The inhibition of cdk2 permits interaction of BRCA2 with Rad51 (see text), so that homologous recombination repair can occur

It is of interest that while cdk2 can compensate for reduced cdk1 activity during DNA end resection in NSCLC cells, the same was not true for BRCA1 phosphorylation. With individual cdk depletion, only cdk1 depletion sensitized to DNA-damaging treatments in these cells. However, cell type-specific differences may exist. For example, in MCF-7 breast cancer cells, selective cdk2 depletion resulted in significant abrogation of Chk1 and p53 phosphorylation after treatment with ionizing radiation (Deans et al., 2006). Cdk2 has also been shown to phosphorylate ATRIP and is required to maintain G2 arrest after DNA damage. Cells reconstituted with mutated ATRIP at cdk phosphorylation sites were not as strongly G2 arrested after ionizing radiation as cells containing wild-type ATRIP (Myers et al., 2007). These results suggest that in some cell types, selective cdk2 depletion or inhibition may sensitize to DNA-damaging treatments. Of note, cdk5 has recently been shown to directly phosphorylate ATM at Ser794, an essential event for the activation of ATM kinase activity in response to DNA damage in post-mitotic neurons (Tian et al., 2009). While cdk5 depletion also affects checkpoint control in non-neuronal cells, the precise mechanism has not yet been defined (Turner et al., 2008).

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11.8.3 Cdk Inhibition and DNA Repair In addition to phosphorylation of CtIP at Thr847 to stimulate endonuclease activity, there is also cdk-mediated phosphorylation at Ser327, which is critical for the interaction of CtIP with BRCA1 and recruitment of CtIP to sites of DNA damage (Yun and Hiom, 2009). Following DNA damage, during S and G2, the cdk-dependent CtIP–BRCA1 interaction directs the cell toward homologous recombination repair, as opposed to the non-homologous end-joining repair that occurs during G1, when cdk2 and cdk1 activities are low. Additionally, the role of cdk1 in BRCA1 phosphorylation and recruitment to sites of DNA damage is likely also important for homologous recombination repair. Cdk inhibition, therefore, may sensitize to DNAdamaging treatments not only by compromising checkpoint signaling but also by preventing homologous recombination repair. By extension, cdk inhibition may sensitize cancer cells to PARP inhibition (Ashworth, 2008). Cdk2 has also been linked to repair of double-strand breaks by non-homologous end joining (Crescenzi et al., 2005; Deans et al., 2006). In MCF7 breast cancer cells, combined cdk2 and BRCA1 depletion resulted in a marked reduction in colony formation compared to individual knockdowns, suggesting that the targeting of several repair pathways may be synthetically lethal (Deans et al., 2006), as is the case with PARP inhibition in a background of BRCA deficiency. Coupled with high cyclin E and low p27Kip1 expression found in BRCA-deficient cells (Aaltonen et al., 2008; Chappuis et al., 2005; Deans et al., 2004), it is possible that these cells are particularly cdk2 dependent. Whether cdk2 inhibition has therapeutic value in BRCA1-deficient cancers is yet to be clinically tested. Of note, cdk2 has also been shown to phosphorylate BRCA2, which impairs its interaction with Rad51, thereby inhibiting homologous recombination (Esashi et al., 2005). Although this activity of cdk2 appears paradoxical, it is representative of the interaction of cdks with BRCA proteins designed to insure that checkpoint control and DNA repair are properly coordinated. Immediately after DNA damage, cdk activity remains high. Cdk1 and cdk2 activities regulate DNA end resection and BRCA1 function and ultimately ATR–Chk1 signaling, while cdk2 phosphorylates BRCA2 and prevents homologous recombination. Later, only after cdk activity is reduced downstream in the checkpoint cascade to promote cell cycle arrest is the interaction of BRCA2 and Rad51 facilitated, permitting homologous recombination repair (Jazayeri et al., 2006). Finally, cdk9 inhibition may also serve to suppress homologous recombination DNA repair. This has been shown to occur in a p53-dependent fashion; although homologous recombination is considered an error-free repair mechanism, its upregulation can lead to genomic instability, such that p53 interestingly plays a role in its suppression, in an activity separable from its classic tumor suppressor functions in transcriptionally transactivating target genes implicated in growth control and apoptosis (Bertrand et al., 2004; Gatz and Wiesmuller, 2006). In p53 wildtype colon cancer cells, cdk9 inhibition enhances the degree of p53 accumulation following exposure to camptothecin-mediated DNA damage. Initially, phosphorylated p53 binds and inhibits Rad51 and then ultimately mediates transcriptional

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repression of Rad51. This inhibition of DNA repair caused by cdk9 inhibition augments camptothecin-induced apoptosis in p53 wild-type cells (Ambrosini et al., 2008). Coupled with effects of cdk inhibition on p21Waf1/Cip1 expression, it is of interest that irinotecan/flavopiridol combinations have produced the greatest benefit in p53 wild-type gastrointestinal malignancies (Shah et al., 2005).

11.9 Future Perspectives New generation cdk inhibitor compounds remain under evaluation. The selective cdk4/6 inhibitor PD0332991 has demonstrated both expected pharmacodynamic and clinical activity in tumors retaining wild-type Rb, albeit with a primary outcome of cytostasis when tumor control is achieved (Leonard et al., 2008). Because of compensatory activity among cdks in transformed cells, equipotent cdk2/cdk1 inhibitors are more likely to arrest cancer cells than drugs more selective for cdk2, although melanoma, lymphoma, and neuroblastoma represent the first examples for which selective cdk2 depletion or inhibition may induce cell cycle arrest or apoptosis (Du et al., 2004; Faber and Chiles, 2007; Molenaar et al., 2009). The G1 or S/G2 cycle arrest afforded by cdk inhibitors has reduced tumor burden in carcinogen-induced models of esophageal and colon adenocarcinoma (Boquoi et al., 2009; Lechpammer et al., 2005), suggesting these compounds could ultimately play a role in the prevention of premalignant cell progression. Several cdk2/cdk1 inhibitors also inhibit transcriptional cdk activity, so that the single agent activity of cdk2/cdk1/cdk9 inhibition will ultimately be tested in both hematologic malignancies and solid tumors. Mcl-1-dependent cancer cells may be particularly susceptible, and cells with high baseline E2F-1 activity may be most prone to apoptosis (Eguchi et al., 2009). Elevation of E2F-1 activity by cdk2/1 inhibition (or cdk8 inhibition) in colon cancer cells may serve to suppress β-catenin activity, also expected to translate to clinical benefit (Morris et al., 2008). In addition to modulation of anti-apoptotic protein expression and E2F-1 activity, the role of cdks in DNA damage-induced checkpoint control and repair suggests multiple mechanisms by which cdks may be effectively combined with DNAdamaging agents. While combined cdk2/cdk1 inhibition may be preferable for inducing cell cycle arrest by a cdk inhibitor alone, selective inhibition of these cdks may be easier to combine with DNA damage to prevent cell cycle arrest in malignant cells that could inhibit a DNA damage response (Deans et al., 2006; Johnson et al., 2009). The clinical development of these combinations remains difficult, since sequence dependence may affect their optimization and a commitment to randomized trials is required. Nonetheless, results to date with several chemotherapy/cdk inhibitor combinations have demonstrated promise (Bible et al., 2005; Goffin et al., 2003; Shah et al., 2005). Whether used alone or in combination, pharmacodynamic assessment of cdk compounds in early-phase trials will be essential for establishing that the expected targets are modulated so that clinical outcomes can be better interpreted (Haddad

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et al., 2004; Hsieh et al., 2009; Tan et al., 2004). Despite the challenges, further work is justified to determine whether the strategy of cdk inhibition will play a part in the anti-cancer armamentarium. Acknowledgments Supported by NIH grants R01 CA090687, P50 CA089393 [DanaFarber/Harvard Center (DF/HCC) Specialized Program of Research Excellence (SPORE) in Breast Cancer], P50 CA090578 (DF/HCC SPORE in Lung Cancer), Susan G. Komen Post-doctoral Fellowship Award KG080773, and Lymphoma Research Foundation Mantle Cell Lymphoma Research Initiative and Correlative Grants MCLI-03-006 and MCLC-07-015.

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Index

Note: The letters ‘t’ and ‘f’ following the locators refer to tables and figures respectively

A Aaltonen, K., 178 Abrieu, A., 63 Acosta, M., 143–145, 147 Acquaviva, C., 45 AC, see Adenocarcinoma (AC) Acute myeloid leukemia (AML) cell lines, 117–118, 169–170 Adams P. D., 11, 139–149 Adenocarcinoma (AC), 159–162, 179 colorectal, 34 lung, 66 oesophageal, 158–159, 161 Adenomatous polyposis coli (APC), 10, 45, 48–49, 59, 61–66, 68–69, 82–84, 86, 158 Aggarwal, P., 49 Aguilera, A., 85 Ailles L. E., 116 Aisner D. L., 128–129 Albanese, C., 6 Aleem, E., 170 Al-Hajj, M., 109 Ali, S., 180 Ali S. H., 14 Allan L. A., 68 Allsopp R. C., 110 Almenara, J., 173 Alonso, M., 173 Alternative lengthening of telomeres (ALT), 11, 127–128, 131–132, 134 Ambrosini, G., 179 AML, see Acute myeloid leukemia (AML) cell lines Amon, A., 83 Anand, S., 84 Anaphase-promoting complex/cyclosome (APC/C), 45, 48–49, 61–64, 68, 82–84

Andreassen P. R., 68, 85 Andrews P. D., 62 Ansari A. Z., 33 Antonchuk, J., 114 APC/C, see Anaphase-promoting complex/cyclosome (APC/C) APC, see Adenomatous polyposis coli (APC) Apoptosis, 3–5, 12–13, 28, 36, 66–68, 80, 84, 88–89, 98, 101–103, 119, 129, 144, 162, 169–174, 178–179 Arai, F., 114 Arentson, E., 50 Arnold H. K., 16 Artandi S. E., 133 Ashworth, A., 178 Astle C. M., 110 Ataxia-telangiectasia mutated (ATM), 27–28, 85–88, 115, 129–130, 175–177 ATM and Rad3 (ATR), 27–28, 85–88, 129–130, 133, 143, 175–178 ATM, see Ataxia-telangiectasia mutated (ATM) ATR, see ATM and Rad3 (ATR) Attwooll, C., 25 Avilion A. A., 130 B Bacchetti, S., 131 Bahram, F., 131 Baird D. M., 128 Baker D. J., 64, 66, 83 Bakhoum S. F., 61 Baldwin A. S Jr, 144–145 Baltimore, D., 87 Bani-Hani, K., 161 Banumathy, G., 142 Barbacid, M., 7–8, 24, 44–45, 47–51, 167 Barrett M. T., 157–162

G.H. Enders (ed.), Cell Cycle Deregulation in Cancer, Contemporary Cancer Research, C Springer Science+Business Media, LLC 2010 DOI 10.1007/978-1-4419-1770-6,

187

188 Barrett’s oesophagus, case study, 157–162 See also Pre-neoplasia, cell cycle deregulation in Barriere, C., 44 Barski, A., 31 Bartek, J., 85 Bartkova, J., 141, 148 Basak, C., 145 Bassermann, F., 86 Batlle, E., 160 Baughn L. B., 169 Bavik, C., 142 Bell S. P., 43–44, 47 Bembenek, J., 83 Benezra, R., 61 Bennett D. C., 146 Bernardi, R., 142 Bernards, R., 6–7, 9 Berndtsson, M., 89 Berthet, C., 8, 44, 170 Bertrand, P., 178 Besson, A., 45 Betts J. C., 144 Bible K. C., 179 Bilaud, T., 129 Bjorklund, S., 33 Blackburn E. H., 127 Blagden, S., 91 Blagosklonny M. V., 90, 173 Blais, A., 6–7 Blasco M. A., 11, 133 Blasina, A., 88 Blow J. J., 9–10, 50 Bodnar A. G., 130 Boehm J. S., 12, 14 Bonnet, D., 109, 116 Boquoi, A., 179 Borlado L. R., 48 Botz, J., 27 Boube, M., 32 Bouchard, C., 12 Boutros, R., 86–87 Boynton R. F., 161 Bracken A. P., 25 Braden W. A., 47 Bradley T. R., 110 Braig, M., 145 Bravou, V., 50 Breakage–Fusion–Bridge (BFB cycles), 129, 133 Bremner, R., 27 Brichese, L., 89 Brito M. J., 161

Index Broccoli, D., 11, 129 Brown E. J., 87 Brown J. P., 129, 141 Bruce W. R., 116 Bryan T. M., 131 Budanov A. V., 100 Bulavin D. V., 140, 145 Bunz, F., 87 Burgering B. M., 12 Burkhart D. L., 25–27 Busby E. C., 91 Buscemi, G., 130 Buzzai, M., 101 Byrd J. C., 171 Byth K. F., 171 C Cahill D. P., 61 Cai, D., 170–172 CAK, see Cdk-activating kinase (CAK) Calbó, J., 8 Caldecott K. W., 86 Calegari, F., 115 Campanero M. R., 29 Campbell M. S., 62 Cancer stem cells (CSCs), 109, 120–121 Cantley L. C., 12 Cao, Y., 131 Carboxy-terminal domain (CTD), 14, 24, 29, 33, 168, 171–173 Carvalho, A., 89 Caspase-independent mitotic death (CIMD) pathway, 68 Castedo, M., 80, 87 Cathepsin, 143 Cdk-activating kinase (CAK), 6, 24, 30, 82, 168, 171 Cdk inhibition and DNA repair, 178–179 ATR–Chk1 signaling, 178 BRCA-deficient cells, 178 cdk9 inhibition, 178 CtIP–BRCA1 interaction, 178 DNA damaging treatments, 178 p21Waf1/Cip1 expression, 179 p53 wildtype colon cancer cells, 178 transcriptional, 171–172 anti-apoptotic proteins (Mcl-1/XIAP) depletion, 171 AZD5438/SNS-032/SCH727965, 172 flavopiridol, 171

Index mRNA transcription and process, 171 seliciclib, 172 U2OS osteosarcoma cells, 172 CDK inhibitors (CKIs) family, 4–7, 11, 45, 51, 84, 168, 175, 179 CDKI, see Cyclindependent kinase inhibitor (CDKI) CDKs for cancer therapy, 167–180 Cdk2 and Cdk1, targeting, 170–171 antisense- or siRNA-mediated cdk2 depletion, 170 cyclin A degradation or ectopic expression of p27Kip1 , 170 depletion of cdk2 or cdk1, 171 genetic ablation of cdk1, 170 MITF and cdk2, correlation, 170 Cdk4 and Cdk6, targeting, 168–170 cyclin D–cdk4/6–INK4 pathway, 168 PD0332991, inhibitor, see PD0332991 treatment Cdk inhibition in DNA damage-induced checkpoint control, 175–177 and DNA repair, 178–179 induction of DNA damage, 175 Cdks and E2F-1 activity Cdk8 in modulation, role of, 174–175 prevention of neutralization during S phase, 173–174 combined targeting and anti-apoptotic proteins survivin as target of Cdk1, 172 transcriptional Cdk inhibition, 171–172 cyclin D1 expression, reduced, 172–173 anti-HER2 antibody trastuzumab, 172 EGFR wild-type cells, 172 future perspectives, 179–180 p53 and p21Waf1/Cip1 , modulation of, 173 acute lymphocytic leukemia/germ cell tumor cells, 173 p21Waf1/Cip1 , 173 p53 wild-type colon carcinoma cells, 173 CDKs/E2F-dependent transcription, 23–36 CDK8–CycC in human cancers, deregulation, 34–35 colorectal tumor cell lines, 35 point mutation of CDK8 (D189N), 34 CDK8–CycC negative regulation, 32–34 biochemical fractionation experiments, 33 CDK8/SRB10-mediated phosphorylation, 32–33

189 E2F1 with CDK8, association, 34 RNA polymerase II (Pol II) transcription, 32 “scaffold complex”, 32 suppression mechanisms, 32 CDK phosphorylation, 27–29 CDK7–CycH, 28 CDK–cyclin complexes, 28 E2F1 or DP1 in vivo, phosphorylation of, 29 E2F1, phosphorylation by kinases, 27 G0–G1 transition and S phases, 27 human E2F1 protein, structure/regulation, 28f NLS, 27 proteasome-dependent degradation, 29 target genes (Apaf1/p73), 28 cell cycle progression, 23–25 B-type cyclins, 24 CDK oscillator/transcriptional oscillator, 24 cell cycle machinery, 24 genetic and molecular experiments, 24 genome sequencing, 24 L-type cyclins (CycL), 24 mitotic cell cycle, 23 RB/E2F proteins, role of, 24 Drosophila model system to study E2F activity in vivo, 31–32 dE2f1-RNAi-induced phenotypes, 32 drawbacks, 32 E2F homologs (dE2F1/dE2F2), 32 E2Fs activate transcription, 29–31 cell proliferation, 31 E2F1 and HATs, physical interactions, 31 E2F1 interaction with p62 subunit (TFIIH in vitro and in vivo), 30 E2F1-mediated transcriptional activation model, 30f TBP, transcriptional factors, 29 future directions, 35–36 Rb and E2F proteins, 25–27 CDK–cyclin (Cyc) complexes, 26 CDKs, components or regulators of, 26–27 DNA replication and mitosis, 25 E2F proteins, classification, 25 eight E2F genes/three DP genes, mammals, 25 G1 to S-phase transition, 25 higher eukaryotes, 25 pRB inactivation, events of, 27

190 CDKs/E2F-dependent transcription (cont.) RB-E2F regulatory network, general properties, 26f small cell lung carcinomas and osteosarcoma, 27 Cell cycle machinery, 24, 167 Celli G. B., 132 Cellular quiescence, 3–18 future directions, 17–18 oncogenes to bypass quiescence, 10–13 adenoviral oncoprotein E1A, 13 ALT, 11 c-MYC-dependent genes, 12 combinations of oncogenes, effects, 10 DNA replication cycle, 11 E2F-activators (E2F1–3), 12 FoxO transcription factors, 12 inactivation of tumor suppressor genes, effects, 11 “malignantly transformed”, 11 p53/ARF pathway, 11 PI3K/AKT pathway, 12 pRB and p53 pathways, 11 RAF/MAPK pathway, 11 RAS/RAF/MEK pathway, 12 “Restriction Point”, 10 Rhabdomyosarcomas, 13 shRNA, serum starvation, 12 SV40 large T (LT)/small t (st) antigens/hTERT, 11 restriction point, 5–10 exit quiescence back into G1, role of cells to, 9–10 G1-cyclins/CDK, pRB, and E2F transcription factors, 6–8 reversible state, 3–5 cell cycle exit, 4f inhibition of CDKs, 4 MyoD-induced differentiation, 5 non-dividing states, four, 3 “quiescent gene expression program”, 4 spontaneous senescence, 5 transcriptional repressor HES1, 5 SV40 and exit from quiescence, 17–18 small t antigen promotes exit, 15–17 tumor antigens and cellular targets, 13–15 three pocket proteins, ablation of, 8–9 Cesare A. J., 132 Chang B. D., 87 Chan G. K., 61–63 Chan T. A., 87 Chan Y. W., 84–85, 88–89

Index Chao S. H., 160–162, 171 Chappuis P. O., 178 Checkpoint kinase 2 (Chk2), 27–28, 85–88, 129–130, 175 Cheeseman I. M., 62 Chen, C., 113, 116 Cheng, T., 113, 115 Chen H. H., 24 Chen L. Q., 162 Chen, R., 171 Chen R. H., 61–63 Chen, W., 14–16, 170, 174 Chen, Y., 86 Chen Y. N., 170, 174 Chen, Z., 139, 145 Cheshier S. H., 111 Chesnokov I. N., 43–44 Chibazakura, T., 68 Chiles T. C., 172–173, 179 Chin G. M., 84 Chin, L., 133 Chi, Y., 33 Chk2, see Checkpoint kinase 2 (Chk2) Chong, L., 129 Chow J. P., 88 Chow J. P. H., 79–91 Chromosome instability (CIN), 60–61 chromosome missegregation/ aneuploidy/CIN, 60 aneuploidy, 60 anti-mitotic drugs treatment, 60 CIN/MIN cancer cells, advantage, 60 dominant gain-of-function mutations, 60 germline mutations, 60 missegregation of one or multiple chromosomes, 60 chromosome missegregation, defects, 61 aneuploidy, 61 genes, lost or mutated, 61 mutations in genes/proteins, 61 sister chromatid cohesion, 61 Chu, K., 88 CIMD, see Caspase-independent mitotic death (CIMD) pathway Cimini, D., 61–62 Cimprich K. A., 86 CIN, see Chromosome instability (CIN) CKIs, see CDK inhibitors (CKIs) family Clarke P. R., 68 Clarkson B. D., 117 Classon, M., 25–26 Clement, G., 160

Index Cleveland D. W., 60, 64–65, 67–68, 84 Cobrinik, D., 45, 47 Cocker J. H., 43, 47 Coe B. P., 64–65 Cohen S. B., 130 Coleman T. R., 43, 47 Collado, M., 145 Coller H. A., 4–5, 9 Collins A. T., 109 Compton D. A., 60 Conaway R. C., 32 Connell-Crowley, L., 8 Conroy, A., 171–172 Coppe J. P., 141–142 Coppola, D., 161 Cortez, D., 86 Cosme-Blanco, W., 146 Courtois-Cox, S., 145 Coverley, D., 48 Cowell J. K., 85 Cragg M. S., 80, 90 Crescenzi, E., 175, 178 Crighton, D., 100 Croxton, R., 174 CSCs, see Cancer stem cells (CSCs) CTD, see Carboxy-terminal domain (CTD) Cyclindependent kinase inhibitor (CDKI), 112–114 D D’Adda di Fagagna, F., 140–141, 148 Dai CY., 145 Damage-regulated autophagy modulator (DRAM), 100–101 Dankort, D., 146 Dannenberg J. H., 8 Dan, S., 85 Darzynkiewicz, Z., 111 Davenport J. W., 61 Deacon, K., 89 Dean J. L., 43–51 Deans A. J., 177–179 De Azevedo W. F., 171 De Bono, J., 91 DeCaprio J. A., 14 Degregori, J., 27 DeLancey J. O., 159 De Lange, T., 129–130 DeLuca J. G., 61–62 Demidenko Z. N., 173 Denchi E. L., 11, 127–134 Deng, Q., 140, 145

191 DePamphilis M. L., 44 DePinho R. A., 11 Desai, A., 60 Dick J. E., 109, 116–117 Diehl J. A., 6, 144 Di Fagagna F. D., 129 Diffley J. F., 10, 44, 48 7, 12-dimethylbenz[a]anthracene (DMBA), 67 Di Micco, R., 148 Dimova D. K., 25 Dimri G. P., 145 Ditchfield, C., 62–63 DMBA, see 7, 12-dimethylbenz[a]anthracene (DMBA) DNA damage by Cdk inhibition induced checkpoint control, 175–177, 177f cdk depletion, 177 checkpoint control initiation, 176 HU, cells treated with, 176 MRN (Mre11–Rad50–Nbs1), 176 NSCLC cell lines, 176 induction of, 175 A2780 ovarian cancer cells, 175 ATR–Chk1 pathway, 175 MCM, 175 topoisomerase inhibitors, 175 DNA damage signaling, 140 aberrant or shortened telomeres, 140–141 SDFs and TIFs, 141 Dobles, M., 64 Donner A. J., 34 Donoghue D. J., 88 Doonan J. H., 44–45, 48–50 Double-stranded breaks (DSBs), 130 Dou, Y., 31 Dowling, M., 84 DRAM, see Damage-regulated autophagy modulator (DRAM) DREAM complex, 9 DSBs, see Double-stranded breaks (DSBs) Duesberg, P., 65 Duhrsen, U., 112 Du, J., 170, 179 Dunham M. A., 131 Duronio R. J., 32 Dutta, A., 43–44, 47, 50 Dyer M. A., 27 Dynein motility-dependent mechanism, 83 Dynlacht B. D., 6–7, 28–29, 174 Dyson, N., 25–26 Dyson N. J., 23–36, 114, 141, 173

192 E E2F-1 phosphorylation, 174 E2F transcription program, 13 Eckerdt, F., 83 ECM, see Extracellular stroma or matrix (ECM) Eddy B. E., 14 EGFR, see Epidermal growth factor receptor (EGFR) EGF, see Epidermal growth factor (EGF) Eguchi, T., 174, 179 Eischen C. M., 99 El-Deiry W. S., 146 Elenbaas, B., 11, 13 Elledge S. J., 131, 176 Elmlund, H., 33 Elowe, S., 63 Emili, A., 29 Enders G. H., 148, 175 Endoplasmic reticulum (ER) stress pathway, 102 “End replication problem”, 128 Engelhardt, M., 132 Epidermal growth factor (EGF), 161 Epidermal growth factor receptor (EGFR), 161, 172–173 Erenpreisa, J., 80, 90 Esashi, F., 178 Eshleman J. R., 60 Essers M. A., 112, 121 Extracellular stroma or matrix (ECM), 143 F Faber A. C., 172–173, 179 Famulski J. K., 62–63 Fang, G., 63 Feldser D. M., 146 Feng, J., 61, 63, 130 Feng, Z., 100–101 Ferber, A., 142 Fernandez P. C., 12 Ferrari, R., 13 Ferrell J. E Jr, 81 Filipe M. I., 161 Finco T. S., 144–145 Fingert H. J., 87 Firestein, R., 35–36, 174–175 Fishel, R., 60 Fisher R. P., 24, 31 Fitzgerald R. C., 157–162 Fkh6, see Forkhead homologue transcription factor (Fkh6) Flejou J. F., 161

Index Fleming I. N., 172–173 Flemington E. K., 29 Fogal, V., 142 Foley, E., 68 Foray, N., 176 Forkhead homologue transcription factor (Fkh6), 160 Forkhead O (FoxO) subfamily, 113 cell cycle checkpoints, 113 FoxO1–4 (triple knockout), 113 FoxO transcription factors, 12 G0–G1 transition, G1–S and G2–M, 113 Forrester, K., 161 Forsyth N. R., 129–130 FoxO, see Forkhead O (FoxO) subfamily Freeman, A., 158 Freire, R., 86 Frolov M. V., 25 Fry C. J., 31 Fry D. W., 168–169 Fryer C. J., 33 Fujita, M., 48–49 Fujiwara, T., 85, 134 Funayama, R., 142 Fung T. K., 80, 82–83 Furukawa, Y., 89 G Gabrielli, B., 84 Galiana, C., 161 Galipeau P. C., 162 Gan, B., 113 Ganem N. J., 61, 67 Garrett M. D., 169 Gascoigne K. E., 63, 67–68 Gatz S. A., 178 Gazin, C., 142 Geddert, H., 161 Geisen, C., 7 Geng, Y., 8, 27, 48 Gerber, P., 14 Gfi–1, see Growth factor independent 1 (Gfi-1) Gil, J., 97, 99, 144 Gillen, P., 160 Gillespie D. A., 63 Gillespie P. J., 50 Gilliland D. G., 109 Girardi A. J., 14 Goffin, J., 179 Goga, A., 89–91, 172 Going J. J., 50, 145, 160 Gojo, I., 171

Index Gonzalez M. A., 50 Gonzalez, S., 50, 143 Gonzalez-Suarez, E., 133 Goodfellow, H., 142 Gope, R., 35 Gorbsky G. J., 61–62 Gordon K. E., 132 Gorgoulis V. G., 27 Grabsch, H., 64 Graña, X., 3–18, 114 Graves P. R., 88 Greenberg R. A., 133 Greenblatt, J., 29–30 Greenleaf A. L., 24 Greenman, C., 34 Greider C. W., 127, 146 Griffin J. D., 116 Griffith J. D., 130 Growth factor independent 1 (Gfi-1), 113 Guan, Y., 117 Guerra, N., 147 Guo, A., 142 Guo, X., 130 Gustafsson C. M., 33 Gu, W., 99 H H2B-GFP, see Histone H2B–green fluorescent protein (H2B–GFP) Haase S. B., 24 Haddad R. I., 179–180 Haematopoietic stem cells (HSCs), 110–119 Hagemeier, C., 29 Hahn, S., 29, 31 Hahn W. C., 11, 13–15, 65 Haigh C. R., 162 Hairy and Enhancer of Split1 (HES1), 5, 9, 13, 17 Ha, L., 146 Hallberg, M., 33 Hall-Jackson C. A., 88 Hall, M., 168 Hallstrom T. C., 12 Hampsey, M., 32 Hannink, M., 144 Harada, H., 169 Harbour J. W., 45, 47 Hardwick K. G., 61, 63 Hardwick R. H., 161 Harper J. W., 168 Harrison D. E., 110 Harris S. L., 141 Harvey, M., 141

193 Hastie N. D., 132 Hateboer, G., 29 HATs, see Histone acetyl transferases (HATs) HCF1, see Host Cell Factor-1 (HCF1) Heald, R., 89 Heath E. I., 172 Heinrich P. C., 147 HEK, see Human embryonic kidney (HEK) cells Helin, K., 25–26 Hemann M. T., 133 Hengartner C. J., 33 Henry D. O., 6 Henson J. D., 132 Herbig, U., 141, 145, 148 Herbst J. J., 160 Herbst, R., 84 Hereditary nonpolyposis colon cancer (HNPCC), 60 Hériché J. K., 29 Hernando, E., 67 Herrera, E., 133 HES1, see Hairy and Enhancer of Split1 (HES1) Hiom, K., 178 Hirst, M., 33 Histone acetyl transferases (HATs), 13, 31 Histone H2B–green fluorescent protein (H2B–GFP), 111 HNPCC, see Hereditary nonpolyposis colon cancer (HNPCC) Hochegger, H., 45, 47 Hockemeyer, D., 129, 134 Hock, H., 113 Hodgson, B., 9–10 Hodgson G. S., 110 Hoffman D. B., 62 Hofmann, F., 29 Homologous recombination (HR)-based mechanism, 131, 178 Hong M. K., 160 Hontz R. D., 97–103 Hope K. J., 117 Horng, T., 147 Horwitz G. A., 13 Host Cell Factor-1 (HCF1), 31 Houghtaling B. R., 129 Hoyt M. A., 61 HR, see Homologous recombination (HR)-based mechanism HSCs, see Haematopoietic stem cells (HSCs) Hsiao K. M., 27 Hsieh W. S., 172, 180

194 Huang, H., 59–70 Huang H. C., 63, 68 Huang P. S., 61, 63 Huang, X., 61 Hu, B., 170 Huertas, P., 176 Hu, J., 49 Human embryonic kidney (HEK) cells, 16 Humbey, O., 99 Huntly B. J., 109 Hunt, T., 91 HU, see Hydroxyurea (HU) Huttner W. B., 115 Hydroxyurea (HU), 176 I Ianari, A., 27 IAP, see Inhibitor of apoptosis protein (IAP) IC, see Initiation complex (IC) IEN, see Intraepithelial neoplasia (IEN) Iftikhar S. Y., 160 Iihara, K., 161 Inadomi J. M., 159 Ingles C. J., 29 Inhibitor of apoptosis protein (IAP), 89, 172 Initiation complex (IC), 44 Inoue, I., 49 Internal tandem duplication (ITD)-expressing cell lines, 169–170 Intraepithelial neoplasia (IEN), 157–159 Ira, G., 176 Ishikawa, F., 117 Itahana, K., 12, 98 ITD, see Internal tandem duplication (ITD)-expressing cell lines Ito, K., 116, 140, 145 Iwanaga, Y., 64 Iwasa, H., 140, 145 J Jablonski S. A., 67 Jackman K. M., 173 Jackson J. R., 67 Jackson S. P., 176 Jacks, T., 6, 25, 85 Jallepalli P. V., 60–61 Jang Y. Y., 115 Jankowski, J., 161 Jazayeri, A., 176, 178 Jelluma, N., 61 Jemal, A., 120 Jhanwar-Uniyal, M., 86 Jiang, J., 174 Jiang W. Q., 132

Index Ji J.-Y., 23–36, 141, 173 Jinno, S., 10, 17 Jin, P., 89 Jin, S., 100, 101 Jin, Z., 162 Johnson D. G., 9, 25, 27 Johnson J. E., 11 Johnson, N., 167–180 Johnson P. F., 145 Jones M. H., 61 Jones R. G., 101 K Kaestner K. H., 160 Kaldis, P., 6, 8, 82 Kalitsis, P., 64, 66 Kamijo, T., 141 Kao G. D., 61 Kaplan K. B., 61, 69 Karakaidos, P., 49–50 Karin, M., 100 Karlseder, J., 130 Karnoub A. E., 140 Kastan M. B., 85 Kato, J., 6 Kaufman P. D., 142 Kaur, B., 162 Kelly-Spratt K. S., 98 Keniry, M., 12 Khanna K. K., 87 Kiel M. J., 115 Kim C. F., 109 Kimchi, A., 98–99 Kim S. H., 129 King E. M., 63 King R. W., 61, 90 Kipreos E. T., 24 Kirschner M. W., 49 Kitagawa, M., 29, 174 Kitahara, K., 50 Kitsios, G., 44–45, 47–50 Kiyono, T., 139, 141 Knez, J., 31 Knudsen E. S., 43–51 Knudsen K. E., 50 Knuesel M. T., 33–34 Kobayashi, S., 173 Koda, M., 112 Konishi, Y., 89 Kops G. J., 60–61, 63–64 Kornberg R. D., 32–33 Kortlever R. M., 143 Kortmansky, J., 91

Index Kouroukis C. T., 171 Kouzarides, T., 28, 30, 174 Kozar, K., 44 Krek, W., 28–29, 174 Krizhanovsky, V., 143, 147 Kroemer, G., 101–102 Kroll K. L., 82 Kuffer, C., 85 Kuhn E. M., 90 Kuilman, T., 142–145, 147 Kumble, S., 161 Kunsch, C., 144–145 L Label retaining cell (LRC), 111 Labib, K., 44 Lam L. T., 171 Lam M. H., 88 Lampson M. A., 62 Lang S. E., 31 Lanni J. S., 85 Lan, W., 62 Lao-Sirieix, P., 157–162 Lapidot, T., 109, 116 Larsson, O., 9 Lau, E., 44, 47, 49–50 Lavin M. F., 87 Leach F. S., 60 Lechpammer, M., 179 Lee B. Y., 145 Lee E. A., 84 Lee, J., 87 Lee J. H., 176 Lee M. G., 24 Lee R. J., 26 Lees J. A., 25, 32 Lee Y. M., 8, 51 Lengauer, C., 60 Lens S. M., 89 Leonard, J., 169, 179 Leone, G., 11–12 Lerner, C., 110 Lessard, J., 116 Leukaemia stem cells (LSCs), 109–121 Leung J. Y., 12 Levine A. J., 100, 141 Lewis T. B., 64 Liang, C., 43, 47 Li, B., 129 Li, C., 109 Li, H., 34 Lindeman G. J., 27 Lindqvist, A., 81–82

195 Ling Y. H., 89 Linskens M. H., 142 Lin S. Y., 131 Lin W. C., 28 Lipinski M. M., 25 Li, R., 61 L’Italien, L., 170–171 Litovchick, L., 9 Liu, F., 7 Liu, D., 129 Liu, E., 49 Liu, J., 24, 116 Liu, Q., 87 Liu S. T., 61, 63–64, 67 Liu, Y., 33 Li, X., 49, 62 Li, Y., 61 Loayza, D., 129 LOH, see Loss of heterozygosity (LOH) of genes Lord R. V., 161 Loss of heterozygosity (LOH) of genes, 65–66, 162 Lowenberg, B., 116 Loyer, P., 24 LSCs, see Leukaemia stem cells (LSCs) Luciano R. L., 31 Lundberg A. S., 47 Lu, X., 171 M Mackey M. A., 89 MAD2 by binding to p31comet, neutralization of, 83 MAD2–CDC20 complex dissociation mechanism, 83 Maiato, H., 63, 67–68, 84 Maier J. A., 143 Mailand, N., 10, 48 Maiorano, D., 43–44, 47 Maiti, B., 25 Maiuri M. C., 100 Majello, B., 30 Maley C. C., 162 Malik, S., 32–33 Malumbres, M., 7–8, 24, 44–45, 47–51, 167, 170 Mammalian target of rapamycin (mTOR), 100 Mao, Y., 63 Markey M. P., 45, 47 Marti, A., 29 Martínez-Balbás M. A., 27 Marx, J., 65

196 Marzec, M., 169 Masuda, A., 68, 84 Matrix metalloproteases (MMP1 and MMP3), 143 Matsumura, S., 63 Matsuoka, S., 113 Matsuura, I., 7 Maude S. L., 175 Ma, Y., 174 Mayer, F., 173 Mayer T. U., 84 Mayer V. W., 85 May K. M., 61, 63 Maynard, S., 116 MCAK, see Mitotic centromere-associated kinesin (MCAK) McCleland M. L., 62 McClendon A. K, 10, 43–51 McClintock, B., 129 McCormick, F., 45, 170 MCC, see Mitotic checkpoint complexes (MCC) components McDonald E. R., 146 McGarry T. J., 49 McIntosh J. R., 62 McKeon, F., 61, 68, 84 Mcm, see Mini-chromosome maintenance (Mcm) complex Medema R. H., 12, 81 Meeker A. K., 132 MEFs, see Mouse embryo fibroblasts (MEFs) Meinhart, A., 168, 171 Melixetian, M., 50 Meltzer S. J., 161 Mendez, J., 44, 48 Menu, E., 169 Merchant A. M., 44 Meyerson, M., 167 Michaloglou, C., 145–146, 148 Microphthalmia-associated transcription factor (MITF), 170 Microsatellite instability (MIN), 60 Millis A. J., 143 Milross C. G., 68 Milton, A., 25 Mimura, S., 44 Mini-chromosome maintenance (Mcm) complex, 44, 46–50 Minn A. J., 85 MIN, see Microsatellite instability (MIN) Mirza A. M., 12 Mismatch repair (MMR) genes, 60

Index MITF, see Microphthalmia-associated transcription factor (MITF) Mitotic catastrophe, 79–91 and cancer: future directions, 89–91 depolyploidization, 90 mis-segregation of chromosomes, 90 multipolar mitosis and further genome instability, 90f roscovitine, 91 UCN-01, 91 caused by abrogation of DNA integrity checkpoints, 87–88 ATM/ATR–CHK1/CHK2 axis, uncoupling method, 87–88 replication or DNA repair, 87 UCN-01 (CHK1 inhibitors), 88 caused by mitotic block/slippage, 83–85 checkpoint disruption, 84 cyclin B1–CDK1, role of, 84 microtubules depolymerization/polymerization, 83 mitotic slippage or adaptation, 84–85 p53-dependent tetraploidy checkpoint, 85 slippage-refractory cells, 84 spindle-assembly checkpoint disruption, 84 spindle-disrupting drugs, 83–84 description/definition, 79–80 as form of cell death involving CDK1, 88–89 apoptosis, caspase activation, 88 BCL-2 family, 89 caspase-6, 88 CDK1-dependent phosphorylations, 88 cyclin B1–CDK1 kinase activity, 88–89 survivin, 89 mitotic control/spindle-assembly checkpoint, 80–83 APC/CCDC20, activation of, 82 APC/CCDH1 inactive during mitosis, 83 CDC25 activation/WEE1 inactivation, 81 CDK1 or CDC2, 80 CDK1Thr14/Tyr15 , phosphorylation, 81 components of checkpoint machinery, 82 control of cyclin B1–CDK1, 81f cyclin B1/B3, 80–81 MCC components, 82 O-MAD2/C-MAD2, “seatbelt” structure, 82–83 PLK1, activation of, 81–82

Index proteolysis of geminin, 82 spindle-assembly checkpoint, termination mechanism, 83 subunits of APC/C, 82 ubiquitin–proteasome system, 82 normal control of DNA damage/replication checkpoints, 85–87 ATM/ATR, activation of, 85–86 ATR–CHK1 pathway, 86 BRCA1-associated genome surveillance complex, 86 CDC25 family, isoforms of, 86–87 claspin by PLK1, phosphorylation of, 86 DNA damage or replication stress, 85 G1/G2 DNA damage checkpoint, 87 Mitotic cell cycle, 23 gap phases (G1 and G2), 23 M phase (for mitosis), 23 S phase (for DNA synthesis), 23 Mitotic centromere-associated kinesin (MCAK), 62 Mitotic checkpoint and CIN in cancer, 59–70 aneuploidy/CIN, mitotic checkpoint, and cancer, 64–67 13 different CIN mice, 65 aneuploidy, genetic factors, 66 APC and BubR, relationship between, 66 core mitotic checkpoint proteins, mutations of, 64 DMBA, 67 “gain-of-function” mutation, 65 heterozygous mutations in genes, 65 high-grade colonic tumors, 66 immortalization, cell sensitivity, 67 kinetochore motor protein, CENP-E, 66 liver tumors, 67 LOH of genes, 65 mitotic checkpoint defects, occurrences, 64, 65t primary breast tumor samples, 65 spindle checkpoint genes, 64 taxol and vinca alkaloids, spindle poisons, 64 tumorigenesis, 65 tumor suppressors/oncogenes/cell cycle checkpoint genes, 64 CIN, see Chromosome instability (CIN) future directions, 69–70 mitosis for chemotherapy, 67–68 anti-microtubule agents, side effects, 67 apoptosis, activation of, 68

197 CIMD pathway, 68 clinical inhibitors, 67 crippled checkpoint or “slippage”, 68 mitotic drug arrest, 67 mitotic index/apoptosis tumors, correlation between, 68 mitotic checkpoint, 61–64 Aurora B kinase, 62 checkpoint proteins (mechanosensors), 63 core components, 61 hZW10–ROD complex and CENP-E, 63 inhibits APC/C, 61 kinetochores elaborate error correction system, 62 Mad1 and Mad2 checkpoint proteins, 62 MCAK, 62 Nuf2/Ndc80 complex, 62 taxol and nocodazole treatment, differentiation, 63–64 upstream regulators/downstream effectors, 63 “wait anaphase signal”, 62 Mitotic checkpoint complexes (MCC) components, 82 Mittler, G., 33 Mittnacht, S., 45, 47 Miura, K., 64 Mixed-lineage leukemia (MLL), 31 Miyamoto, K., 113, 116 MLL, see Mixed-lineage leukemia (MLL) MMP1 and MMP3, see Matrix metalloproteases (MMP1 and MMP3) MMR, see Mismatch repair (MMR) genes Molenaar J. J., 173, 179 Molz, L., 80 Moore M. A., 116 Moreno C. S., 16 Morgan, C., 162 Morgan D. O., 168 Moroni M. C., 28 Moroy, T., 7 Morris E. J., 28, 32, 34–35, 174, 179 Morselli, E., 102 Moshkin Y. M., 142 Motwani, M., 173 Mouse embryo fibroblasts (MEFs), 8–9, 12, 66, 99, 102–104, 144 MRN (Mre11–Rad50–Nbs1), 86, 132, 176

198 MTOR, see Mammalian target of rapamycin (mTOR) Mukaida, N., 144–145 Müller, H., 25–27 Mulligan, G., 6 Multani Muntoni, A., 132 Murphy M. E., 97–103 Murphy, N., 50 Murray A. W., 61 Musacchio, A., 61, 63, 82–83 Myers J. S., 177 Myers L. C., 32–33 N Näär A. M., 33 Naef A. P., 159 Nahta, R., 172 Nakajima, T., 144–145 Nakamura, T., 161 Nakamura T. M., 130 Nakanishi, M., 129 Nasmyth, K., 24 Nemunaitis, J., 172 Nerlov, C., 144 Neshat, K., 162 NES, see Nuclear export sequence (NES) Nevins J. R., 12, 25–27 Nevis K. R., 17 NHEJ, see Non-homologous end joining (NHEJ) NHF, see Normal human fibroblasts (NHF) Nicklas R. B., 62 Niida, H., 88, 129 Niikura, Y., 68, 84 Nishitani, H., 43, 47 Nitta, M., 88 NK2 homeobox transcription factor (Nkx2–3), 160 Nkx2–3, see NK2 homeobox transcription factor (Nkx2–3) NLS, see Nuclear localization signal (NLS) Non-homologous end joining (NHEJ), 129, 132, 178 Non-small cell lung cancer (NSCLC) cell lines, 169, 173, 176–177 Normal human fibroblasts (NHF), 8–9, 11–12, 16–17 NSCLC, see Non-small cell lung cancer (NSCLC) cell lines Nuclear export sequence (NES), 102 Nuclear localization signal (NLS), 27 Nurse, P., 24, 79–80

Index O O’Brien C. A., 109 O’Connor D. S., 84, 89, 172 O’Dwyer, P., 169 Ohata, N., 34 Ohtani, K., 27 Ohta, S., 50 Ohta, T., 29 Ohtsubo, M., 8 Olovnikov A. M., 128 Oncogene signaling pathways, 140 chronic signaling, 140 mutated oncogenic Ras (K-Ras/N-Ras/ H-Ras), 140 Raf–MEK–ERK signaling, 140 Onwuegbusi BA., 161 ORCs, see Origin replication complexes (ORCs) Orford K. W., 115 Origin replication complexes (ORCs), 43–44, 47–48 Orlando D. A., 24 Orr-Weaver T. L., 83 Ortega, S., 44, 50, 170 Orth J. D., 67–68 Orth, K., 88 O’Shaughnessy J. A., 157 Ouatu-Lascar, R., 162 Ouyang, B., 64 Owen T. A., 9 P P18INK4C -deficient cells, 174 P300/CBP-associated factor (PCAF), 31 P53/ARF/control of autophagy, 97–103 ARF induces autophagy, 98–99 full-length ARF in autophagy, role of, 99 full-length ARF and smARF, 98 smARF, 98 ARF tumor suppressor and autophagy, 97–98 ARF, functional aspects, 98 oncogenes (Ha-ras/c-MYC/β–catenin), 97, 98 p14ARF tumor suppressor, 97 p16INK4a tumor suppressor, 97 cell survival and promote tumor progression, enhancement, 99–100 Bcl-xl, 99–100 Beclin-1/Bcl-xl complex, 100 cytoprotective role, 99 overexpressed in B-cell tumors, 99

Index p53-null sarcoma cell line, 99 pro-survival role in autophagy, 99 two-dimensional in-gel electrophoresis technique, 99 future directions autophagy pathway(s), 103 ER stress pathway, 102 stress-activated/“unstressed” p53, 102, 103f nutrient stress signals to p53, 101 p53 negative autophagy regulation in unstressed cells, 101–102 “hotspot” mutations in p53, 102 Kroemer group, 102 mammalian cells/Caenorhabditis elegans, 101 tumor-derived point mutants (R282W/R273H), 102 p53 transactivates autophagy gene DRAM, 100–101 p53 target genes, PUMA and Bax induce mitophagy, 101 p53 tumor suppressor and autophagy, 100 mTOR, inhibition mechanisms, 100 p53 by DNA damaging agents, activation of, 100 p53 target genes, sestrin1 and sestrin2, 100 Pabst, O., 160 Page-McCaw, A., 143–144 PAI, see Plasminogen activator inhibitor (PAI) Pallas D. C., 14 Palmero, I., 141 Palm, W., 129, 132 Pandolfi P. P., 142 Pantic, M., 134 Pardee A. B., 5, 10 Park I. K., 116 Park S. S., 91 Parmar, K., 115 Parrilla-Castellar E. R., 86 Parrilla, P., 162 Parry, D., 172 Parsons, R., 12, 60 Passegue, E., 114–115 Pattingre, S., 100 Paull T. T., 176 Paulson T. G., 161–162 Payton, M., 171 PC2, see Positive Cofactor 2 (PC2) PCAF, see P300/CBP-associated factor (PCAF) PCNA-interacting protein (PIP), 29

199 PD0332991 treatment, 168–170 bortezomib and dexamethasone, 169 dose-limiting toxicities, 169 effects on AML cell lines, 169 IC50 values, 168 induces cytostatic G1 arrest, 169 inhibits recombinant cdk4 and cdk6, 168 in mice bearing Colo-205 colon cancer xenografts, 169 p16INK4A tumor suppressor protein, 169 pharmacodynamic study, 169 Rb-dependent/Rb-deficient xenografts, 169 SW480 colon cancer cells, 169 Pearson, A., 29–30 Pearson, M., 142 Pediconi, N., 27 Peeper D. S., 28, 142, 145, 174 Pelicci P. G., 109–121 Pellish L. J., 160 Pellman, D., 44, 61, 67 Pennati, M., 84 Perkins N. D., 144 Perry, S., 117 Persons D. L., 161 Petermann, E., 86 Petersen B. O., 48 Peters F. T. M., 162 Peters, G., 97, 99, 168 Peters J. M., 45, 61 Petit, I., 112 Pfeifer G. P., 27 Pfleghaar, K., 61 Phatnani H. P., 24 Phelps-Durr T. L., 142 Phosphatase and tensin homologue (PTEN), 11–12, 100, 113 PI3K–Akt pathway, 113, 140 PI3K–PTEN–Akt pathway, 113 PIC, see Pre initiation complex (PIC) Pimkina, J., 99–100 Pines, J., 45, 168 Pinsky B. A., 62 Pipas J. M., 14 PIP, see PCNA-interacting protein (PIP) Plasminogen activator inhibitor (PAI), 143 Plentz R. R., 134 PML, see Promyelocytic leukemia (PML) Pocket proteins p130 and pRB, 6 Polyomavirus Simian Virus 40 (SV40), 11, 13–17, 66 Poon R. Y., 83, 86 Poon R. Y. C., 79–91

200 Porras, A., 16 Porter L. A., 88 Positive Cofactor 2 (PC2), 33 Potts P. R., 132 PP2A catalytic subunit (PP2A/C), 14 PP2A/B subunits, four families, 14 PP2A/C and PP2A/A, isoforms, 14 structural subunit (PP2A/A), 14 variable B subunit, 14 PRB and p53 tumor suppressor pathways, 140 PRB, see Retinoblastoma protein (pRB) Pre initiation complex (PIC), 29–30 Pre-neoplasia, cell cycle deregulation in, 157–162 Barrett’s carcinogenesis, proliferation, 160 Fkh6 and Nkx2–3, 160 luminal surface, proliferation at, 160 squamous oesophagus, 160 Wnt pathway, 160 BE and oesophageal cancer, 158–159 BE with dysplasia, 159 definition of BE, 158 duodeno-gastro-oesophageal reflux, 158–159 haematoxylin/eosin representative of BE with dysplasia, 158f identification of biomarkers, 159 oesophageal AC with dysplasia, 159 surveillance-detected cancers, 159 cell cycle progression, factors influencing, 160–161 growth factors and oncogenes, role of, 161–162 c-myc oncogene, amplification, 161 methylation of p16, 161–162 ras family (h, k and n), oncogenes, 161 luminal factors, role of, 162 complete/incomplete normalisation acid exposure, 162 sodium–hydrogen exchanger (Na+ /H+ ), 162 pre-neoplasia IEN, risk factors, 157–158 lowgrade dysplasia to IEN, 158 molecular abnormalities, 158 Pre-RC assembly, regulation of, 43–51 CDKs and general cell cycle control, 44–47 APC/C, 45 cell cycle order, 45 cell cycle progression, 44 Cip/Kip family of proteins, 45 CKIs family, 45

Index complex regulation by CDK/cyclins, 46f cyclin D–CDK4/6 complexes, 45 E-type and A-type cyclins, 45 mitogenic signaling, 45 serine/threonine kinases, 44 functional effects of deregulated Pre-RC assembly, 49–50 depletion of geminin, 50 over-expression of Cdc6 and Cdt1, 49 over-expression of pre-RC components, 49 negative impact of CDKs, 48–49 direct binding/inhibition of Cdt1 by geminin, 49 DNA re-replication, 49 inhibition of Cdc6, 48 S, G2, and M phases, re-initiation of, 48 ubiquitin-mediated proteolysis, 49 positive impact of CDKs, 47–48 CDK2 null fibroblasts, 48 cyclin–CDK complex/cyclin E–CDK2, 47–48 cyclin D–CDK4/6 complex, activation, 47 cyclin E null fibroblasts, 48 RB pathway, activation, 47 Pre-RC components in cancer, 50 deregulation of CDK4/6, 50 deregulation of cyclin A and cyclin E, 50 deregulation of Mcm2–7, 50 deregulation of pre-RC components, 50 over-expression of Cdt1 and/or Cdc6, 50 pre-replication complex, 43–44 DNA replication, 43 initiation complex (IC), 44 “Pre-replication complex” (pre-RC), 10, 17, 43–51, 47, 82 Price D. H., 171 Prieur, A., 145 Promyelocytic leukemia (PML), 118, 132, 142 PTEN, see Phosphatase and tensin homologue (PTEN) Puig P. E., 90 Pulse–chase system, 111 PUMA and Bax induce mitophagy, 101 Q Qian, Z., 64 Qi, W., 63 Quelle D. E., 8, 97

Index R Rabinovitch P. S., 161 Rae1–Bub3/Rae1– Nup98 (compound heterozygotes), 65 Rancati, G., 63, 69 Randall T. D., 110 Rao C. V., 66 Reactive oxygen species (ROS), 115–116, 139–140 Reddel R. R., 132 Reed S. I., 24 Reef, S., 98–99 Reid B. J., 160–161 Reinhardt H. C., 87–88 Ren, B., 25–27 Resnitzky, D., 8 Restriction point, 5–10 cell cycle, functional parts, 5 E2F transcription factors, 6 exit quiescence back into G1, role of cells to, 9–10 APC-mediated ubiquitination, 10 “attachment checkpoint”, 10 cell cycle, elongation of, 9 DNA pre-replication factors, 9 MCM2 expression levels, 10 quiescence, definition (Blow and Hodgson), 9 video-microscopy experiments (Zetterberg and Larsson), 9 G1-cyclins/CDK, pRB, and E2F transcription factors, 6–8 activation of E2F-program of gene expression, 7f CAK and CKIs, 6 D-type cyclins/E-type cyclins, 6 histone synthesis, 7 inactivation of pocket proteins, 8 mitogenic stimulation, 6 primary tumors and derived tumor cell lines, 7 quiescent NHF, 8 quiescent tumor-derived T98G cells, 8 role of G1 CDKs, 8 phosphorylation of pocket protein, 5 pocket proteins inactivation, 8–9 apoptotic cells, 8 DREAM complex, 9 E2F1–3, for cell cycle re-entry, 9 pocket protein/E2F pathways, roles, 9 proliferation rate, 5 Retinoblastoma protein (pRB), 6–8, 11–12, 14–15, 27–31, 35, 140–141, 167

201 Ricci-Vitiani, L., 109 Ricke R. M., 65–66 Rieder C. L., 62–63, 67–68, 84 RNA interference (RNAi), 32, 34 RNAi, see RNA interference (RNAi) Roberts A. I., 143 Roberts B. T., 61 Roberts J. M., 4, 6, 8, 44–45, 48, 141, 168 Rocha, W., 142 Roeder R. G., 32–33 Roninson I. B., 80, 88 ROS, see Reactive oxygen species (ROS) Rossi D. J., 112, 116 Ross J. F., 29 Rothblum-Oviatt C. J., 87 Roussel M. F., 45 Rowland B. D., 6–7, 9 Ruchaud, S., 88 Rudolph K. L., 132–133, 146 Russell, P., 79–80 Ryan K. M., 100 S Sage, J., 8, 12, 25–27 Salmon E. D., 61, 63, 82–83 Sampliner R. E., 159 Samuelsen C. O., 33 Sancho, E., 160 Sang, L., 5, 13 Santamaria, D., 8, 45, 170 Sarbia, M., 161 Sarkaria J. N., 88 Sarkisian C. J., 146 Sartori A. A., 176 SASP, see Senescence-associated secretory phenotype (SASP) Sausville E. A., 167 Sauvageau, G., 116 Sawyer S. L., 44 Saxena, S., 50 Scadden D. T., 115 “Scaffold complex”, 33 Scatena C. D., 89 SCFSkp2 complex in mammals, 29 Schatton, T., 109 Schimming, R., 68 Schmidt, M., 12 Schmitt C. A., 146 Schmittwolf, C., 114 Schulman B. A., 29 Schulmann, K., 161

202 Schulze, A., 26 Schwartz G. K., 88 Sciallero, S., 161 SCID leukaemia-initiating cell (SL-IC), 116 SCs, see Stem cells (SCs) SDFs, see Senescence associated DNA damage foci (SDFs) Sears R. C., 16 Sebastian, T., 144–145 Sedelnikova O. A., 145 Seike, M., 64 Self-renewing divisions in normal/LSCs, regulation of, 109–121 existence of LSCs, 116–117 evidence, 116 flow cytometry methods, 116 “long-” and “short-repopulating” cells, 117 SL-IC, 116 xenotransplantation models, 116 future directions, 120–121 anti-tumour drugs, 120 cancer stem cell hypothesis, 120 genetic models of SC exhaustion, 112–114 abnormal signalling, 113 c-Myc, 113 deletion of Fbw7, 113 HoxB4 over-expression, 114 niche, 113 p21cip1/kip1, 112–113 SNAG domain, 113 HSCs are deeply “dormant”, 110–112 5-fluorouracil, 110 “dormant” and “activated” HSCs, comparison, 112 “dormant” HSCs, 111–112 G-CSF and IFNα, 112 GFP-positive LRC/GFP-negative non-LRC SCs, 111 low levels of RNA and DNA, 111 LRC, 111 multiparametric flow cytometry, 111 pulse–chase system, 111 “quiescence” or “G0”, 110 quiescent or “resting” cells in vivo, 110 “stem cell mobilization”, 112 LSCs are quiescent, 117 molecular mechanisms of SC exhaustion, 115–116 Bmi1, Polycomb family, 116 DNA damage/genomic damage, 116 “dormant” niche/vascular niche, 115 G1 phase of cell cycle, 115

Index replicative stress, 116 ROS/SNO, 115 molecular mechanisms of SC quiescence, 114–115 cdk actions opposed by INK4/Cip/Kip family, 114 cycD-cdk4/6 complexes, 114 myelosuppressive chemotherapy or mobilization, 114 paradigm in SC exhaustion, 118–120 damage repair mechanism, 119 DNA repair mechanism, effects, 120 role of p21 and DNA damage, 119f regulation of quiescence and self-renewal, 117–118 AML1–ETO oncogene, 118 bone marrow transplantation, 117 cell cycle inhibitor p21, 118 cell cycle “pause”, 118 PML-RAR oncogene, 118 self-renewal potential of HSCs, 110 daughter cells, SCs originate, 110 haemopoiesis, 110 self-renewal, 110 serial transplantation assay, 110 Senderowicz AM., 91 Senescence associated DNA damage foci (SDFs), 141 Senescence-associated heterochromatin foci (SAHF), 142–143 Senescence-associated secretory phenotype (SASP), 141–142 Senescence-associated-β-galactosidase activity (SA-βgal), 145 Senescence-messaging secretome (SMS), 141–142 Senescence secretome, impact on tumor suppression/cancer, 139–149 altered secretory phenotype of senescent cells growth regulators, 142–143 inflammatory regulators, 143 regulation of secretome, 144–145 stromal regulators, 143–144 See also Senescent cells, altered secretory phenotype of senescence secreteome in tumor suppression, 145–148, 146f benign clonal neoplasms, 145 cancerous neoplasms, 146 chemokines, 147 clonal benign human nevi, 148 components, role of other, 147–148

Index human NK-like cell line, 147 immune regulators, 147 immune surveillance, 147 liver cancer, 147 mutagenic processes, 146 pre-cancerous neoplasms, 145 proliferation arrest, 146 SA-βgal, 145 senescence program, aspects, 145 tumor suppression process, 148 senescence signaling pathways, 140–141 cell senescence program, 140 DNA damage signals, 140 oncogene signaling pathways, 140 p53 and pRB tumor suppressor pathways, 141 triggers of cell senescence, 139–140 oxidative stress or ROS, 139 replicative senescence, 139 telomeres, 139 triggers/signals/effectors of cell senescence, 140f Senescent cells, altered secretory phenotype of growth regulators, 142–143 cyclin A2, 142 formation of SAHF, 142 gene silencing or nuclear heterochromatinization, 142 histone chaperones, 142 SAHF/HIRA/WNT2, connections and significance, 142 inflammatory regulators, 143 cytokines/chemokines receptors, 143 NKG2D signaling, 143 regulation of secretome, 144–145 C/EBPβ and NFκB, 145 CXRC2 ligands, 144 MEFs, 144 oncogenic stress, 144 regulators of senescence, 144 stromal regulators, 143–144 ECM, 143 MMPs, effects and function, 143–144 Senga, T., 49 Seo, J., 50 Seo, S., 82 Serrano, M., 11, 139 Shah M. A., 179 Shamma, A., 145 Shan, B., 26–27 Shapiro G. I., 167–180 Sharkis S. J., 115 Sharp J. A., 142

203 Shay J. W., 131, 139, 141 Shelton D. N., 142–144 Shenolikar, S., 14 Shen S. C., 84 Shepherd B. E., 110 Sherr C. J., 4, 6, 11, 27, 44–45, 48, 97–98, 114, 141, 167–168 Shetty, A., 50 Shibata, D., 60 Shibutani S. T., 29 Shichiri, M., 64 Shigeishi, H., 64 Shih I. M., 65 Shi, J., 67–68 Shimizu, M., 27 Shin H. J., 84 Short mitochondrial ARF (smARF), 98–99 Sicinski, P., 8, 51 Sihn C. R., 84 Singer M. S., 142 Singh S. K., 109 Single nucleotide polymorphisms (SNP) analysis, 64 Sirieix P. S., 160 Skipper H. E., 117 Skoczylas, C., 15–16 Skoufias D. A., 62 Skwarska, A., 88 Slee E. A., 88 Sluder, G., 85 SmARF, see Short mitochondrial ARF (smARF) Smith E. R., 31 Smogorzewska, A., 129 SMS, see Senescence-messaging secretome (SMS) Smyth M. J., 147 SNO, see Spindle-shaped N-cadherin-positive osteoblasts (SNO) SNP, see Single nucleotide polymorphisms (SNP) analysis Sontag, E., 16 Sotillo, E., 3–18, 114 Sotillo, R., 65 Sottile, J., 141 Souza R. F., 160, 162 Spindle-shaped N-cadherin-positive osteoblasts (SNO), 115 Sprenger, F., 68 Srinivas, G., 112 Stearns, T., 85 Steen J. A., 64 Stegmeier, F., 83

204 Stein, B., 144–145 Stem cells (SCs), 109–110, 114 Stevaux, O., 32 Stevens, C., 27 Stewart S. A., 11 Stillman, B., 43–44, 48 Stoeber, K., 50 Storchova, Z., 44, 61, 67, 85 Strebhardt, K., 83 Su A. I., 34 Sudakin, V., 61 Sudo, T., 68, 84 Sugimoto, M., 98 Sugimoto, N., 49 Sun, P., 146 SV40 and exit from quiescence, 17–18 small t antigen promotes exit, 15–17 “attachment checkpoint”, 17 cyclin A protein levels, 16 effects on transformation of human cells, 15 foci formation in hTERT-NHF, 16 HEK cells, 16 monkey kidney cells (CV-1), 16 pre-replication complex factor (CDC6), 17 RAF/MAPK pathway, 16 tumor antigens and cellular targets, 13–15 carboxy-terminal domain, 14 cell cycle proteins, upregulation, 15f DnaJ chaperone domain, 14 PP2A, see PP2A SV40 antigens in human cells, effects/expression, 14 SV40, discovery of, 14 SV40, see Polyomavirus Simian Virus 40 (SV40) Swann J. B., 147 Symmans W. F., 68 SynMuvB, see Synthetic multivulva class B (synMuvB) gene products Synthetic multivulva class B (synMuvB) gene products, 9 Sze K. M., 64 T Taatjes D. J., 32–33 Tada, S., 49 TAD, see Transactivating domain (TAD) Tagami, H., 142 Tago, K., 98 Takahashi, A., 88 Takai, H., 87, 129, 141

Index Takisawa, H., 44 Tanaka T. U., 62 Tan A. R., 180 Tang, Z., 61 Tansey W. P., 33 Tao, W., 68, 84 Tasdemir, E., 101–102 TATA-box binding protein (TBP), 29–30 Tatsumi, Y., 49 Taubert, S., 31 Tavernarakis, N., 102 Taylor S. S., 61, 63, 67–68, 84 TBP, see TATA-box binding protein (TBP) Telomerase or alternative mechanism (ALT), 11, 127, 131–132, 134 Telomere dysfunction-induced foci (TIFs), 141, 145 Telomeres in cancer, maintenance, 127–134 chromosome end protection, shelterin complex, 129–130 DNA damage machinery, 129 DSBs, 130 localization of POT1, 130 localization of shelterin, 129–130 molecular mechanisms, 130 six proteins in mammalian cells, 129 telomeric TTAGGG, 129 future perspectives, 134 telomere dysfunction, 128–129 ATM/ATR, mammalian DNA damage response pathways, 129 BFB cycles, 129 end-to-end chromosomal fusions, 129 functions, 128 NHEJ, 129 telomere elongation in cancer, 130–132 expression of TERT, 130–131 PML nuclear bodies (APBs), 132 telomerase complex, composed of, 130 telomere elongation mechanisms, 131f T-SCE, 132 telomere instability and cancer progression, 132–133 colorectal carcinomas/breast carcinomas, 132 mus musculus (Terc–/– mice), 133 oral squamous cell carcinomas, 132 skin carcinogenesis/liver carcinogenesis, 133 telomere deregulation in cancer, 133f telomere shortening, 132

Index telomere length in mammals, 127–128 “end replication problem”, 128 mammalian telomeres/shelterin complex/T-loop, 128f telomerase, 127–128 telomerase-mediated telomere elongation/erosion, 127 Telomere sister chromatid recombination (T-SCE), 132 Te Poele R. H., 146 Tetsu, O., 170 Therman, E., 90 Thibodeau S. N., 60 Thompson C. B., 101 Thompson S. L., 60 Thoms H. C., 169 Tian, B., 177 TICs, see Tumour-initiating cells (TICs) TIFs, see Telomere dysfunction-induced foci (TIFs) Tissue-type plasminogen activator (tPA), 143 Tommasi, S., 27 Tomonaga, T., 60 Toogood P. L., 168 Torres E. M., 67, 69 Tothova, Z., 113, 116 TPA, see Tissue-type plasminogen activator (tPA) Transactivating domain (TAD), 28 Trautmann, B., 161 Tremain, R., 142 Triadafilopoulos, G., 160 Trimarchi J. M., 25, 32 Tsantoulis P. K., 27 TSC1, see Tuberous sclerosis complex 1 (TSC1) T-SCE, see Telomere sister chromatid recombination (T-SCE) Tse A. N., 88 Tselepis, C., 161 Tsuiki, H., 68 Tuberous sclerosis complex 1 (TSC1), 113 Tumbar, T., 111 Tumour-initiating cells (TICs), 109 Turner N. C., 177 Tyagi, S., 31 Tye B. K., 44 U Ueda, Y., 98 Uetake, Y., 85 Umar, A., 60 Urist, M., 28

205 V Vaclavicek, A., 64 Vakifahmetoglu, H., 80, 88 Vandel, L., 28, 30, 174 Van den Heuvel, S., 32 Van de Peppel, J., 33 Van Der Gaag, H., 116 Van Leuken, R., 83 Van Steensel, B., 129 Van Vugt M. A., 81 Vassilev L. T., 34 Vaughn D. J., 169 Vaziri, C., 49 Venezia T. A., 115 Venkataraman, G., 171 Ventura, A., 146 Verona, R., 27 Verreault, A., 142 Viale, A., 109–121 Viale, A., 109–121 Vigo, E., 27 Vincent, O., 33 Virshup D. M., 14 Vogel, C., 87–88 Vong Q. P., 61 Vousden K. H., 101 W Waga, S., 44 Wajapeyee, N., 142, 147 Walch, A., 161 Wall N. R., 84, 172 Wang K. K., 159 Wang, L., 170 Wang, M., 144 Wang, Q., 64, 88 Wang, X., 64, 170 Wang, Y., 86 Warner J. K., 117 Watanabe, G., 16 Waters J. C., 62 Waugh D. J., 147 Weaver B. A., 60, 64–68, 84 Weber J. D., 98 Weinberg R. A., 11, 27, 47, 140 Weiss, E., 61 Weissman I. L., 110 West M. D., 141, 143 Wiesmuller, L., 178 Williams B. R., 67 Williams G. H., 50 Wilson, A., 111–113, 115 Wilson A. C., 31

206 Wilson, C., 147 Winey, M., 61 Wingless and Int pathway (Wnt), 160 Wnt, see Wingless and Int pathway (Wnt) Wohlschlegel J. A., 49 Wolf, F., 82 Wong, C., 85 Wong O. K., 63 Woychik N. A., 32 Wright W. E., 139 Wu, K., 172 Wysocka, J., 31 X Xiong, Y., 29 Xue, W., 143, 146–147 Xu, M., 28–29, 174 Y Yamori, T., 85 Yanagida, M., 82 Yang, C., 173 Yang, L., 114 Yang, S., 34 Yang S. I., 14 Yao, X., 63 Yarden R. I., 176 Yasmeen, A., 50 Yee K. S., 101–102 Yeh, E., 16 Ye J. Z., 129

Index Yen T. J., 59–70 Ye, X., 142, 147 Yoshida, K., 49 Young A. R., 145 Yuan, B., 64 Yuan T. L., 12 Yuan, Y., 114 Yu, D., 84 Yudkovsky, N., 33 Yuen K. W., 60 Yu, H., 82–83 Yu H. T., 132 Yun M. H., 178 Yu, Q., 173 Z Zachos, G., 63 Zetterberg, A., 9 Zhang, J., 113, 115 Zhang, R., 142 Zhang, Y., 98 Zhao J. J., 12 Zheng, N., 25 Zhong, W., 49 Zhong Z. H., 132 Zhu, L., 25 Zhu, W., 26–27, 50 Zhu, Y., 175 Zimonjic, D., 65 Zou, L., 176

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