Checkpoint Controls and Targets in Cancer Therapy (Cancer Drug Discovery and Development)

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Author: Zahid H. Siddik

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Checkpoint Controls and Targets in Cancer Therapy

Cancer Drug Discovery and Development Series Beverly A. Teicher, PhD, Series Editor

For other titles published in the series, go to www.springer.com/humana select the subdiscipline search for your title

Checkpoint Controls and Targets in Cancer Therapy Edited by

Zahid H. Siddik University of Texas, M.D. Anderson Cancer Center, Houston, Texas, USA

Editor Zahid H. Siddik Department of Experimental Therapeutics The University of Texas M.D. Anderson Cancer Center 1515 Holcombe Boulevard Houston, TX 77030 USA [email protected]

Series Editor Beverly A. Teicher, PhD Department of Oncology Research Genzyme Corporation Framingham, MA, USA

ISBN 978-1-60761-177-6 e-ISBN 978-1-60761-178-3 DOI 10.1007/978-1-60761-178-3 Springer Dordrecht Heidelberg London New York Library of Congress Control Number: 2009933775 © Humana Press, a part of 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 (Humana Press, 999 Riverview Drive, Suite 208, Totowa, NJ 07512 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)

For Jaan, Bubbloo, Bubbly and Pupsee, The Essentials of Life

Preface

Much work over the last two decades has firmly established that loss of cell cycle checkpoint regulation, and resultant unabated cellular proliferation, is an inherent characteristic of cancer. This loss can occur through aberration in any one single component of the many signal transduction pathways that orchestrate checkpoint regulation, and results in either a failure to activate the checkpoint or a failure to respond to the activated checkpoint. In normal cells, checkpoint pathways are activated when genetic or cellular homeostasis is compromised, and signals are then transduced to re-establish basal conditions, and, failing this, to activate the apoptotic machinery to induce a cellular suicidal response. This implies that both survival and cell death pathways are induced following checkpoint activation, and that the final decision by the cell to live or die is dependent on the net result from integrating two opposing sets of signals. It is intriguing that checkpoint pathways are also critical in cancer therapy to provide an apoptotic stimulus in tumor cells when cellular damage induced by the therapeutic agent is detected by the sensor system. Therefore, it is not surprising that failure in pro-apoptotic signals following DNA damage induces therapeutic resistance. Understanding the intricacies of checkpoint response is, therefore, central not only for identifying key checkpoint targets in tumor cells, but also for the design of therapeutic regimen that will enhance antitumor effects. Although early versions of this design entail empirical combinations of cytotoxic agents with cell cycle or checkpoint inhibitors, a greater understanding of the concepts could make such combinations more rational and, thereby, clinically

more effective. Toward that goal, the contributions in this book will consolidate the current state of knowledge on checkpoint responses and provide the status of targets under investigation for potential therapeutic exploitation. The immediate attraction of the book to the scientific community is that it represents a timely opportunity to build upon existing views of checkpoints and expand our understanding of the inner workings of the critical checkpoint machinery. The accumulating knowledge over the past several decades has provided ample appreciation that response to checkpoint activation is manifested through coordinated inhibition of cyclin-dependent kinase (CDK) complexes in G1, S, and/or the G2 phase in order to arrest the cell cycle. Kinase inhibition can occur through several mechanisms, including inhibitory phosphorylation of CDK, destruction of the cognate cyclins, and recruitment of CDK inhibitors from the INK and WAF1/CIP1 families. However, the wealth of information from recent discoveries needs to be examined critically to expand our horizons. At the same time, there is acute awareness that checkpoint response varies depending on the cytotoxic agent, and this serves as a reminder of the magnitude of complexity that is inherent in checkpoint regulation. This volume is intended to bring the cancer research community closer toward a better understanding of this regulation, and how checkpoint abnormalities can promote cancer progression and impact negatively on chemotherapeutic outcome. This book brings together renowned experts, who are defining the field of checkpoints, and as vii

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such it represents a unique collection of insightful contributions that will serve as an important resource for both the research community and the medical oncologists. Since checkpoint regulation

Preface

is inextricably linked to cancer development and cancer therapy, the contents of this book will be of significant interest and appeal to both basic and translational investigators alike. Zahid H. Siddik, Ph.D.

Contents

Preface...................................................................................................................................................

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Part I Circuitry of Checkpoint Response 1. Evasion of G1 Checkpoints in Cancer........................................................................................... Krijn K. Dijkstra, Cristophe Blanchetot, and Johannes Boonstra

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2. Distinct Pathways Involved in S-Phase Checkpoint Control......................................................... Paula J. Hurley and Fred Bunz

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3. Mechanisms of G2 Phase Arrest in DNA Damage-Induced Checkpoint Response...................... Jian Kuang and Ruoning Wang

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4. Centrosomes in Checkpoint Responses......................................................................................... Alwin Krämer

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5. Interplay of 14-3-3 Family of Proteins with DNA Damage-Regulated Molecules in Checkpoint Control.................................................................................................................... Mong-Hong Lee, Sai-Ching Jim Yeung, and Heng-Yin Yang

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Part II Checkpoint Response and the Aetiology of Cancer 6. Chromatin Modifications and Orchestration of Checkpoint Response in Cancer......................... Makoto Nakanishi

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7. DNA Damage Response and the Balance Between Cell Survival and Cell Death........................ Bernd Kaina, Wynand P. Roos, and Markus Christmann

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8. Dysfunction of the RB Retinoblastoma Gene in Cancer............................................................... 109 Francesca Pentimalli, Letizia Cito, and Antonio Giordano 9. G1 Phase Cyclins in Cancer Development and Progression.......................................................... 123 John Patrick Alao

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Contents

10. The BRCA1/2 Pathway Prevents Some Leukemias and Lymphomas in Addition to Breast/Ovarian Cancers: Malignancies that Overcome Checkpoint Controls......................... 155 Bernard Friedenson Part III Targeting Checkpoint Response in Cancer Therapy 11. Regulation of p53 Activity and Associated Checkpoint Controls............................................... 171 Sean M. Post, Alfonso Quintás-Cardama, and Guillermina Lozano 12. The Importance of p53 Signaling in the Response of Cells to Checkpoint Inhibitors................ 189 Alan Eastman 13. Targeting p21-Dependent Pathways for Cell Death in Cancer Therapy...................................... 199 Zahid H. Siddik 14. p27Kip1 as a Biomarker and Target for Treatment of Cancer..................................................... 215 Xiao-Feng Le and Robert C. Bast Jr. 15. Targeting Cyclin-Dependent Kinases with Small Molecule Inhibitors....................................... 235 Paolo Pevarello, James R. Bischoff, and Ciro Mercurio 16. Chk1 and Chk2 as Checkpoint Targets........................................................................................ 245 Haiying Zhang, Zhan Xiao, and Tom Sowin 17. Targeting Cdc25 Phosphatases in Cancer Therapy...................................................................... 261 Johannes Rudolph Index..................................................................................................................................................... 271

Contributors

John Patrick Alao, Ph.D. Department of Cell and Molecular Biology, Lundberg Laboratory, University of Gothenburg, S-405 30 Göteborg, Sweden, [email protected] Robert C. Bast, Jr., M.D. Department of Experimental Therapeutics, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA, [email protected] James R. Bischoff, Ph.D. Experimental Therapeutics Programme, Centro Nacional de Investigaciones Oncologicas (CNIO), 28029 Madrid, Spain Cristophe Blanchetot, Ph.D. Cellular Architecture and Dynamics, Institute of Biomembranes, 3584 CH Utrecht, The Netherlands Johannes Boonstra, Ph.D. University College Utrecht, 3508 BE Utrecht, The Netherlands Cellular Architecture and Dynamics, Institute of Biomembranes, 3584 CH Utrecht, The Netherlands, [email protected] Fred Bunz, M.D., Ph.D. Department of Radiation Oncology and Molecular Radiation Sciences and the Sidney Kimmel Comprehensive Cancer Center, Johns Hopkins University School of Medicine, Baltimore, MD 21231, USA, [email protected] Markus Christmann, Ph.D. Department of Toxicology, University of Mainz, D-55131 Mainz, Germany Letizia Cito, Ph.D. CROM – Center of Oncology Research Mercogliano, Avellino, Italy Krijn K. Dijkstra, B.Sc. University College Utrecht, 3508 BE Utrecht, The Netherlands

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Alan Eastman, Ph.D. Department of Pharmacology and Norris Cotton Cancer Center, Dartmouth Medical School, Lebanon, NH 03756, USA, [email protected] Bernard Friedenson, Ph.D. Department of Biochemistry and Molecular Genetics, College of Medicine, University of Illinois Chicago, Chicago, IL 60607, USA, [email protected] Antonio Giordano, M.D., Ph.D. CROM – Center of Oncology Research Mercogliano, Avellino, Italy Sbarro Institute for Cancer Research and Molecular Medicine, Center for Biotechnology, College of Science and Technology, Temple University, Philadelphia, PA 19122, USA, [email protected] Department of Human Pathology and Oncology, University of Siena, Siena, Italy Paula J. Hurley, Ph.D. Department of Radiation Oncology and Molecular Radiation Sciences and the Sidney Kimmel Comprehensive Cancer Center, Johns Hopkins University School of Medicine, Baltimore, MD 21231, USA Bernd Kaina, Ph.D. Department of Toxicology, University of Mainz, D-55131 Mainz, Germany [email protected] Alwin Krämer, M.D. Clinical Cooperation Unit Molecular Hematology/Oncology, German Cancer Research Center and Department of Internal Medicine V, University of Heidelberg, 69120 Heidelberg, Germany, a.kraemer@dkfz-heidelberg Jian Kuang, Ph.D. Department of Experimental Therapeutics, Division of Cancer Medicine, M. D. Anderson Cancer Center, Houston, TX 77030, USA, [email protected] Xiao-Feng Le, M.D., Ph.D. Department of Experimental Therapeutics, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA Mong-Hong Lee, Ph.D. Department of Molecular and Cellular Oncology, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA, [email protected] The Program in Genes & Development, The University of Texas Graduate School of Biomedical Sciences at Houston, Houston, TX, USA Guillermina Lozano, Ph.D. Department of Genetics, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA, [email protected] Ciro Mercurio, Ph.D. DAC Srl c/o Campus IFOM-IEO, Via Adamello 16, 20139 Milano, Italy

Contributors

Contributors

Makoto Nakanishi, M.D., Ph.D. Department of Cell Biology, Graduate School of Medical Sciences, Nagoya City University, Nagoya 467-8601, Japan, [email protected] Francesca Pentimalli, Ph.D. CROM – Center of Oncology Research Mercogliano, Avellino, Italy Sbarro Institute for Cancer Research and Molecular Medicine, Center for Biotechnology, College of Science and Technology, Temple University, Philadelphia, PA 19122, USA Paolo Pevarello, Ph.D. Experimental Therapeutics Programme, Centro Nacional de Investigaciones Oncologicas (CNIO), 28029 Madrid, Spain, [email protected] Sean M. Post Department of Genetics, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA Alfonso Quintás-Cardama Department of Genetics and Department of Leukemia, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA Wynand P. Roos, Ph.D. Department of Toxicology, University of Mainz, D-55131 Mainz, Germany Johannes Rudolph, Ph.D Department of Chemistry and Biochemistry, University of Colorado at Boulder, Boulder, CO 80309, USA, [email protected] Zahid H. Siddik, Ph.D. Department of Experimental Therapeutics, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA, [email protected] Tom Sowin, Ph.D. Advanced Technology, Global Pharmaceutical Research and Development, Abbott Laboratories, Abbott Park, IL 60064, USA Ruoning Wang, Ph.D. Department of Experimental Therapeutics, Division of Cancer Medicine, M. D. Anderson Cancer Center, Houston, TX 77030, USA Zhan Xiao, Ph.D. MedImmune, LLC, One MedImmune Way, Gaithersburg, MD 20878, USA Heng-Yin Yang, Ph.D. Department of Molecular and Cellular Oncology, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA Sai-Ching Jim Yeung, M.D., Ph.D. Department of Endocrine Neoplasia and Hormonal Disorders,

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Department of General Internal Medicine, Ambulatory Treatment & Emergency Care, The University of Texas M. D. Anderson Cancer Center, Houston, TX 77030, USA Haiying Zhang, Ph.D. Cancer Research, Global Pharmaceutical Research and Development, Abbott Laboratories, Abbott Park, IL 60064, USA, [email protected]

Contributors

Part I

Circuitry of Checkpoint Response

Chapter 1

Evasion of G1 Checkpoints in Cancer Krijn K. Dijkstra, Cristophe Blanchetot, and Johannes Boonstra

Abstract A cell progressing through the cell c ycle encounters various checkpoints in G1 that can lead to an exit from the cell cycle. This ensures that cells only proliferate when destined to do so. Cancer, however, is characterized by uncontrolled proliferation and cancer cells thus bypass these checkpoints. In this chapter, the various G1 checkpoints will be scrutinized with an emphasis on how alterations in cancer can lead to their evasion. The molecular core driving cells through G1 and central in all checkpoints consists of CDKs, cyclins, activating enzymes, and CKIs. Shortly after mitosis, cells meet a first checkpoint that may direct them to apoptosis in the presence of cellular stress. After passing this checkpoint, cells meet the restriction point (R), which monitors the presence of growth factors and in which pRb plays a central role. Two checkpoints can halt the cell cycle beyond R and induce apoptosis or senescence: the classic p53dependent stress checkpoint and a recently identified energy checkpoint. In contrast to an exit of the cell cycle in the absence of growth factor signaling, hypermitogenic signaling may also result in cell cycle arrest and senescence. This seems to be a central mechanism to prevent tumorigenesis. Finally, the role of reactive oxygen species in control of the G1 phase will be considered, with emphasis on their possible involvement in the hypermitogenic arrest. Keywords Cell cycle • Early G1 checkpoint • Restriction point • Stress checkpoint • Hypermitogenic stress • Energy checkpoint • ROS

1.1 Introduction To proliferate, a cell must undergo a cell cycle consisting of four phases. The first gap phase (G1) is characterized by cellular growth and prepares the cell for replication of its DNA, which occurs in the synthesis (S) phase. After duplication of its chromosomes, the cell enters a second gap phase (G2) and finally divides during mitosis (M). The cell cycle is under tight control to ensure that all phases are completed correctly. The regulation of G1 phase progression is particularly interesting, because this phase contains several decision points, called checkpoints, which determine whether a cell continues to proliferate or exits the cell cycle definitively or transiently. If all G1 checkpoints are passed, the cell enters the S phase. However, various events can arrest the cell at one of the checkpoints and prevent further progression through the cell cycle. This is an adaptive mechanism because it ensures that cells only proliferate when they are destined to do so. Cancer is marked by uncontrolled cell proliferation, indicating that the checkpoints that would normally have arrested the cells do not function properly in tumor cells. A characterization of the alterations in tumors that render them irresponsive to the ‘stop’ signals from checkpoints is important because it allows for specific anti-cancer therapies. This chapter will discuss the various checkpoints in the G1 phase of the cell cycle and how they are evaded by cancer cells. First, an overview of the central players

From: Checkpoint Controls and Targets in Cancer Therapy, Cancer Drug Discovery and Development, Edited by: Zahid H. Siddik, DOI 10.1007/978-1-60761-178-3_1, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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that drive cells through G1 is provided. This is followed by a discussion of the various checkpoints that a cell would normally meet when going through the G1 phase, starting shortly after mitosis and ending just before onset of the S phase.

K.K. Dijkstra et al.

This molecular core controlling G1 progression is central in all checkpoints that will be discussed in the following sections. Many alterations in cancer will affect components of this molecular core [2], either altering their expression by abnormal upstream signaling or by directly mutating the genes involved.

1.2 The Molecular Core Controlling Progression Through G1

1.2.1 CDKs are the Engine Behind the Cell Cycle

In the G1 phase of the cell cycle, cells have to make a decision between continuing proliferation or exiting the cell cycle to become quiescent, differentiated, senescent or apoptotic (rev. in 1). While various proteins control this process, a central role is played by the cyclin-dependent kinases (CDKs). In early G1, progression through the cell cycle is promoted by CDK4 or CDK6, depending on the cell type. In late G1, the actions of CDK2 prepare the cell for entry into S phase and DNA replication. CDKs are regulated at various levels. Their activation state is subject to a tug-of-war between activating binding partners (cyclins) and cyclin-dependent kinase inhibitors (CKIs). Upon cyclin binding, the cyclin–CDK complexes undergo activating phosphorylations and dephosphorylations that enable them to phosphorylate their target proteins ultimately promoting entry into S phase (Fig. 1.1).

In yeast, the organism where CDKs were initially discovered, only one CDK exists. In eukaryotes, however, up to eleven CDKs have been found, with different CDKs being active in different phases of the cell cycle (rev. in 2, 3). In G1, the most important CDKs are CDK2 and CDK4/6 (rev. in 2, 4). The most important target of these three kinases is the retinoblastoma protein, pRb. In short, in its hypophosphorylated form, pRb binds to E2F transcription factors, preventing transcription of genes essential for progression through late G1 and S. When CDK4/6 and CDK2 phosphorylate pRb, these transcription factors are released and the cell cycle progresses from G1 to S. The targets of CDK6 are limited to members of the pRb family, and CDK4 only has three other targets: Smad3, the replication licensing factor Cdt1, and Marcks, the myristoylated alanine-rich C-kinase substrate [2, 4].

Figure 1.1. The central molecular core controlling progression through G1. CDKs require binding to cyclins to become activated. Cyclin binding induces a conformational change, exposing a site that can be phosphorylated by CAK. Normally, CDKs are inactive due to inhibitory phosphorylations on two other residues, which are removed by CDC25A, leading to CDK activation. When the CDK is fully activated, it can phosphorylate several target proteins. CKIs compete with cyclins for CDKs, thereby inhibiting CDK activity. In addition, CKI binding induces conformational changes in CDKs that prevent activating phosphorylation by CAK. CDK cyclin-dependent kinase; CAK CDK-activating kinase; CDC25 A cell division cycle 25 A; CKI cyclin-dependent kinase inhibitor; P phosphorylated residue.

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1. Evasion of G1 Checkpoints in Cancer

In contrast, in addition to pRb family members, CDK2 has a wide range of other targets, which are mainly involved in the initiation of DNA replication, such as the mini chromosome maintenance (MCM) proteins ([5]; rev. in 4, 6). The different CDK isoforms show considerable functional redundancy (rev. in 4). Mice with single null-mutations in CDK2, CDK4, or CDK6 develop relatively normally, with only some effects in specific cell types, such as defects in the erythroid lineage or in pancreatic b cells ([7–9]; rev. in 4). Even mice with triple knock-outs of the interphase CDKs (CDK2, CDK4 and CDK6) can develop until midgestation; it was shown that CDK1 compensated for their loss by binding to cyclins D and E [10]. This indicates that CDK1 is sufficient to drive the cell cycle in most cell types. The requirement of interphase CDKs in certain specific cell types may be due to specific substrates unique to those cells. Alternatively, it is conceivable that when interphase cyclins bind to their normal CDK partners, the complex has higher activity than when they bind to CDK1. Possibly, in certain specific cell types, CDK1 in a complex with interphase cyclins cannot generate the required kinase activity for progression through the cell cycle [10]. This functional redundancy has implications for the design of anti-cancer drugs, as targeting only one CDK will probably not be effective enough in halting the cell cycle. In some cases, tumor cells have mutations in genes encoding for CDKs [2, 4]. For example, amplification of the CDK4 gene can occur in breast cancer [11]. Furthermore, in some cases of family melanomas, a mutation in CDK4 renders it insensitive to inhibition by INK4 [12]. In addition, in a few cases of splenic marginal zone lymphoma and B-cell lymphoma, translocation of the CDK6 gene resulted in its overexpression [13, 14]. However, mutations in the genes encoding for CDKs are relatively rare [4], which is perhaps surprising considering their crucial role in controlling G1 phase progression. This may be partly explained by the notion that only few mutations would result in a gain of function and most mutations would render CDKs nonfunctional, prohibiting cell cycle progression. This implies that abnormal functioning of the cell cycle machinery in tumor cells is often not due to alterations in CDKs themselves, but rather in their regulators.

1.2.2 Cyclins are Essential Binding Partners for CDKs While CDKs are constitutively expressed throughout the cell cycle, different CDKs are active in different cell cycle phases (rev. in 3, 4). The temporal control of their activation is regulated by cyclins, with different types of cyclins expressed in different phases of the cell cycle. Up to 16 different eukaryotic cyclins have been found, with the D-type and E-type cyclins being the most important ones in the G1 phase (rev. in 3, 15). Cyclin D binds to CDK4/6 and cyclin E to CDK2. CDK2 can also bind to cyclin A, but that is considered to be the beginning of the S phase and therefore lies beyond the scope of this chapter. Cyclins are mainly regulated at the transcriptional level. As will be discussed in detail in the next section, CDKs can both bind to cyclins and to certain inhibitory proteins. When the amounts of cyclins increase, they can overrule these inhibitors in the fight for CDKs and thereby promote cell cycle progression. One way in which abnormal cell cycle progression can occur in cancer is therefore by overexpression of cyclins. Indeed, the cyclin D gene is mutated in many cancers [15, 16]. While the three D-type cyclins (D1, D2 and D3) are highly homologous, cyclin D1 is most frequently associated with cancer (rev. in 15, 16). Overexpression of cyclin D1 can have various causes, from chromosomal translocations or gene amplification to specific mutations in the coding region that alter its activity. For example, in approximately 70% of all mantle cell lymphomas, chromosomal translocation of the cyclin D1 gene results in its overexpression [17]. Amplification of 11q13, where the cyclin D1 gene is located, occurs in many tumors, for example in 15–32% of nonsmall-cell lung cancers ([18, 19]; rev. in 20) and 25% of pituitary adenomas [21]. In addition to control on the level of transcription, cyclins are also regulated by their stability and subcellular localization. Phosphorylation of cyclin D1 by glycogen synthase kinase-3b (GSK-3b) on a specific threonine residue is required for its polyubiquination and its nuclear export [22–25]. Cyclin D1 can be mutated such that it is insensitive to cytoplasmic degradation. A single nucleotide-mutation (G870A) leads to alternative splicing of the cyclin D1 gene resulting in a protein lacking this threonine residue (rev. in 16).

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Cyclin E is also frequently overexpressed in several human cancer cell lines and primary tumors [6]. Most mutations affect upstream regulators of cyclin E expression, but amplification of the cyclin E gene itself can also contribute to tumor formation [6]. This has been found with frequencies ranging from 2 to 20% in endometrial, ovarian, gastric, colorectal and breast cancers, in some cases leading to increased mRNA expression [23–29]. In conclusion, alterations in cyclins can promote cell cycle progression in various ways, from overexpression to increased stability and nuclear localization. Since cyclins are such crucial elements of the cell cycle machinery, tumors frequently target these essential proteins for mutations.

1.2.3 CKIs Prevent Activation of CDKs While cyclin binding to CDKs is required for their activation, they can be inhibited by another class of proteins, the cyclin-dependent kinase inhibitors (CKIs). Two different families of CKIs can be distinguished according to their affinity for specific cyclin–CDK complexes (rev. in 30–32). Members of the INK4 family (Inhibitors of CDK4) bind to CDK4 and CDK6, preventing their association with cyclin D (Fig. 1.1). In contrast, members of the Cip/Kip family (CDK inhibitory protein/kinase inhibitory protein) bind both to cyclins and to CDKs and show a broader specificity, affecting cyclin D-, E- and A-dependent kinases. The INK4 family consists of four members: p16(INK4A), p15(INK4B), p18(INK4C) and p19(INK4D). The INK4 inhibitors bind to CDK4 and CDK6 and induce conformational changes that prevent cyclin D binding, thus leading to cell cycle arrest (rev. in 30–33). p19INK4D is not associated with tumorigenesis, but the p18INK4C gene is located on a chromosomal region linked to abnormalities in human tumors [34] and p18-knock out mice develop pituitary and testicular tumors. However, the locus containing the p15INK4B and p16INK4A genes is the most frequently mutated in cancer. Alternative splicing of this locus gives rise to the protein ARF, whose tumor-suppressor function is mainly through upregulation of p53 by binding to and inhibiting its negative regulator, murine double minute 2 (MDM2) (rev. in 35, 36). Much research has been aimed at identifying which of the genes on the locus is the most potent tumor-suppressor,


and p16 seems the most likely candidate. In many cancers, p16 is one of the most frequent mutations found, and specific loss of p16 (without p15 or ARF) has been found in many tumors ([37]; rev. in 32, 35, 38). Inactivation of p16 has been found in as much as 85% of studied tumor cell lines and in many primary tumors [32]. Two important mechanisms of p16 inactivation are gene deletions and inappropriate methylation of the promoter leading to gene silencing [38]. It remains unclear to what extent inactivation of p15 or ARF contributes to tumorigenesis, as they are often codeleted with each other, so specific genetic lesions inactivating only one of these proteins and not the other two on this locus (i.e., single knock-outs of p15 or ARF) are not well described [35, 38]. However, there are some reports suggesting that in some cases mutation of p15 or ARF specifically is associated with cancer, for example through methylation silencing [39–43]. The Cip/Kip family consists of three proteins, p21 (Cip1), p27(Kip1) and p57(Kip2) (rev. in 31, 32, 44). These proteins share a conserved domain that can bind to both cyclins and CDKs, but the remaining sequence is different, indicating functional differences. Indeed, p21Cip1 seems to play an important role in p53-mediated cell cycle arrest upon DNA damage, whereas p27Kip1 usually stops the cell cycle upon mitogen starvation or other inducers of quiescence. p57 Kip2 likely plays an important role during embryonic development. While the INK4 family of CKIs clearly inhibits CDK4/6 by preventing binding of cyclin D, the role of members of the Cip/Kip family in inhibiting CDKs is a little more complicated [31, 44]. They can effectively inhibit CDK2 in late G1, preventing entry into the S phase. However, their inhibition of CDK4/6 is controversial. Some studies report inhibition of CDK4/6 by Cip/Kip family members [45–47], while others found no inhibition in vivo or in vitro [48–50]. It has even been suggested that Cip/Kip CKIs may act as activators of CDK4/6 as they promote their assembly with cyclin D [31]. Moreover, it is important to realize that CDK4/6 can sequester Cip/Kip CKIs away from CDK2, thereby promoting cell cycle progression. A recent study sheds some light on this controversy by showing that p27Kip1 can be both inhibitory and noninhibitory [50]. In arrested cells, p27 has an inhibitory effect on CDK4/6. However, in proliferating cells,

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p27 becomes phosphorylated on two critical tyrosine residues, rendering it noninhibitory. Furthermore, p27 may also have oncogenic functions in some situations [51, 52]. Low p27 levels do not always correlate with proliferation rate and p27+/ − mice are more susceptible than p27-null mice to the development of certain cancers. Possibly, p27 has cyclin-CDK-independent functions that are oncogenic. Mice with a mutation in p27 that prevents its interaction with cyclins or CDKs but retains other functions develop more tumors than complete p27 knock-outs. In addition, cytoplasmic localization of p27 – where it cannot inhibit CDKs but may retain potentially oncogenic functions – is a negative prognostic factor in cancer in some situations ([53, 54]; rev. in 55). This suggests that cancer cells may select against the tumorsuppressive roles of p27, while retaining their oncogenic functions. Expelling Cip/Kips from the nucleus could retain their possibly cytoplasmic roles, while preventing inhibition of CDKs. It has been proposed that cytoplasmic Cip/Kip activity is essential for regulation of the cytoskeleton via inhibition of RhoA [44, 51, 55, 56]. Regarding the different functions of Cip/Kip CKIs depending on the situation, it is not surprising that the p21 and p27 genes are rarely mutated in cancer [44]. Deleting the genes would circumvent not only their inhibitory effects, but also any other functions that may actually promote cell cycle progression. For p57, no oncogenic role has been reported, which may explain why its gene is inactivated in different ways in various tumors (rev. in 44). Mutations in the gene are also found in 5–10% of Beckwith–Wiedemann syndrome cases, an overgrowth syndrome with increased risk of embryonic tumor development [57]. In conclusion, while it is clear that both the INK4 and Cip/Kip families of CKIs are important players in the control of the cell cycle and in cancer, some recent findings only underscore that much can still be learned about these intriguing proteins. Nevertheless, the fact that CKIs play an important role in many tumors remains. In some situations, as with p16, mutations in the gene itself can inactivate the inhibitors. In other cases, such as with p21 and p27, mutations in the gene itself are not always favorable, but alterations in upstream signaling pathways may still affect the function of these CKIs.

1.2.4 Phosphorylation and Dephosphorylation of Cyclin–CDK Complexes Upon binding to their cyclin partners, CDKs need to be both phosphorylated and dephosphorylated to be activated. The enzyme CDK-activating kinase (CAK) can provide an activating phosphorylation of the CDK on a specific threonine residue (rev. in 58). This leads to a conformational change of the cyclin–CDK complex, resulting in improved binding and transfer of phosphoryl groups to its substrates [59–61]. CAK is a complex consisting of CDK7, cyclin H and the assembly factor ménage a trois1 (MAT1). Binding of cyclins changes the conformation of the CDK and exposes the target phosphorylation site. Therefore, CDKs not bound to their cyclin partners are only poorly phosphorylated by CAK. On the other hand, binding of CKIs changes the conformation of CDKs such that the phosphorylation site is inaccessible (Fig. 1.1). In this way, CAK activity is mainly regulated by its substrate conformation. During the cell cycle, levels of all CAK subunits remain fairly constant [62]. Moreover, levels of CDK7 show little or no decline after serum starvation [62]. This suggests that CAK activity is not a limiting factor in controlling cell cycle progression [63]. Consistent with this notion, CDK7 is only marginally increased in tumor cells [62]. However, CDK7 may still be a relevant target for anti-cancer therapy, as its inhibition would affect the activity of all other CDKs [58]. As has been discussed above, a general CDK inhibitor is likely to be more effective than inhibitors of specific CDKs due to redundancy among the CDKs. Of course, whether this redundancy also applies to CDK7 and whether other CDKs could overcome its inhibition is still doubtful. CDKs are normally held in an inactive state through phosphorylation on two other residues. CDK2 is phosphorylated by the WEE1 kinase, but the kinase responsible for inhibitory phosphorylations on CDK4/6 has not yet been identified [64– 67]. These inhibitory phosphorylations are removed by the cell division cycle 25 (CDC25) phosphatases (rev. in 68). This class of phosphatases consists of three isoforms: CDC25A, CDC25B, and CDC25C. While the latter two play important roles in the S phase and G2/M transition, CDC25A is primarily

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involved in the G1/S transition [68, 69]. However, some redundancy has been reported between different CDC25 isoforms, since CDC25B−/− and CDC25C−/− double knock-outs develop normally [70]. Unlike CAK, levels of CDC25A fluctuate throughout the cell cycle. It is predominantly expressed in G1 and it forms an auto-amplification (positive feedback) loop with cyclin E-CDK2 through their mutual activation [71]. CDC25A participates also in the DNA damage checkpoint, as CHK1 can lead to cell cycle arrest via the degradation of CDC25A upon DNA damage [68]. Overexpression of CDC25 family members is a recurrent theme in a wide range of tumors [68]. For example, CDC25A was found overexpressed in 52% of breast carcinoma samples and in 56% of hepatocellular carcinoma samples [72, 73]. There are various ways in which CDC25 can become overexpressed in cancer. No evidence exists for gene amplification or rearrangement or other specific genetic mutations of the CDC25 gene. However, the transcription factors Myc and E2F play a role in the transcriptional regulation of CDC25A [74–76]. Levels of these transcription factors are frequently elevated in cancer, as will be discussed in the next section. An increase in CDC25A levels might also occur at the posttranslational level. For example, breast cancer cell lines that overexpress CDC25A showed increased stability of the CDC25A protein [77]. Normally, CDC25A is degraded in S and G2 by bTRCP (beta-transducin repeat containing protein), a component of the SCF ubiquitin ligase complex. Mutations leading to decreased levels of bTRCP have been found in various tumors, and may lead to increased levels of CDC25A [68]. In addition, suppression of WEE1 is also associated with several cancers, including nonsmall-cell lung cancer and colon cancer [78, 79]. In short, both phosphorylation by CAK and dephosphorylation by CDC25A are required for the activation of G1 CDKs. While CAK does not seem to play a big role in tumorigenesis, mutations in CDC25A can contribute to increased proliferation in some cancers. As discussed in this and previous sections, the molecular core functioning as the engine of the cell cycle consists of CDKs, cyclins, CKIs, and specific phosphatases and kinases. Mutations in many of these components make cells more susceptible to tumorigenesis. The following sections will take a closer look at specific checkpoints in the G1 phase


of the cell cycle. It will become evident that these checkpoints are activated in response to different stimuli, use different molecular mechanisms and steer the cell in different directions, but all of them will modulate the molecular core defined in this section in some way or another.

1.3 Early G1 Checkpoint The study of cell cycle progression is usually performed with cells synchronized by serum starvation. Cells become quiescent in the absence of growth factors, but upon serum addition, quiescent cells re-enter the cell cycle in the G1 phase. Unfortunately, there are several limitations of this method. The transition from G0 to G1 is difficult to establish precisely and the cells have to recover from serum starvation, which is a rather nonphysiological event that could be reflected in the state of the cell [80]. Another disadvantage is that events in the G1 phase that occur before the point of re-entry are seldom studied. By synchronizing cells with mitotic shake-off – a technique that isolates mitotic cells by making use of the fact that mitotic cells loosen their attachment to each other and to a substratum – many of these problems can be circumvented [80]. When cells synchronized in mitosis were deprived of serum immediately after mitosis, they were unable to enter S phase. This in itself is not so surprising, as cells only become independent of growth factors after passage of the restriction point (see following section). However, in contrast to quiescent cells and cells serum-starved 2 h after mitosis, these cells did not re-enter the cell cycle upon readdition of serum. This indicates that a previously unknown checkpoint early after mitosis can lead to permanent withdrawal from the cell cycle [81]. Cells arrested at this state, called G0−, were characterized by the absence of cyclin D, phosphorylated mitogen-activated protein kinase (MAPK) and phosphorylated focal adhesion kinase (FAK) and the presence of p27KIP1. MAPK normally gets phosphorylated as early as 10 min after mitosis, indicating that these cells were arrested immediately after mitosis [81, 82]. Of note, cells became apoptotic after prolonged serum starvation: after 3 days 40% of the cells seemed to be induced for apoptosis. This suggests that early after mitosis the


cell decides whether it further progresses through the cell cycle or enters apoptosis [80]. An important protein involved in survival pathways is the phosphoinositide 3-kinase (PI3K). Its activators include, but are not limited to growth factor receptors, attachment to the extracellular matrix, and cellular stress, including heat shock and hypoxia [83, 84]. PI 3-kinase not only controls cellular survival, but also cell proliferation. This important role for PI3K in both anti-apoptotic and proliferation pathways make it a likely candidate to be involved in the early G1 checkpoint. Indeed, cells treated with inhibitors of PI3K 0 to 2 h after mitosis showed significantly less incorporation of radioactive thymidine 14 h after the start of the experiment, in contrast to cells where PI3K was inhibited later in G1 [85]. Cells with inhibited PI3K arrested shortly after mitosis, indicated by the absence of cyclin D and the absence of phosphorylated MAPK. These cells also showed increased expression of the apoptotic protein caspase-3 and underwent more apoptosis [85]. Protein kinase B (PKB)/Akt is an important downstream target of PI3K known to block apoptosis and promote survival. Inactivation of PKB/Akt was probably involved in this cell cycle arrest, as inhibition of PKB/Akt produced largely similar phenotypes [80]. There are several ways in which PI3K promotes cell cycle progression and survival via PKB/Akt (rev. in 86, 87). Firstly, Akt promotes the stability and synthesis of cyclin D1 by inhibiting glycogen synthase kinase-3b (GSK-3b). GSK-3b promotes cyclin D1 degradation by the proteasome and it can decrease transcription of cyclin D1 via b-catenin and LEF transcription factors. PKB/Akt can inhibit GSK-3b, thereby leading to increased levels of cyclin D1. PKB can also phosphorylate p21CIP1 and p27KIP1, expelling them from the nucleus and preventing inhibition of CDKs. A particularly interesting target of PKB/Akt is the family of forkhead transcription factors (FOXO1, 3, 4 and 6) [86, 88]. They induce the transcription of genes involved in both apoptosis and cell cycle arrest making it attractive to consider a role for FOXO in the early G1 checkpoint. Activation of FOXO can lead to cell cycle arrest by inducing the transcription of the CKIs p21CIP1 and p27KIP1 and of the Rb family member p130, which leads to cell cycle arrest by binding to inhibitory E2F transcription factors (see next section) [89–91]. FOXO can also halt the cell cycle by

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repressing expression of cyclin D1 and D2 [92, 93]. In addition, FOXO induces apoptosis by increasing expression of proapoptotic genes such as BIM [94] and down-regulating expression of prosurvival genes, such as Bcl-xL ([95]; rev. in 88). PKB phosphorylates FOXO, which leads to translocation of FOXO to the cytoplasm, thereby preventing transcription of its target genes and promoting cellular survival and proliferation ([96, 97]; rev. in 86, 88). The signaling pathway proposed to play a central role in this checkpoint is depicted in Fig. 1.2. At least three situations can be imagined in which this checkpoint would be activated. First, PI3K, PKB, and FOXO are all involved in responses to cellular stress [83, 86, 88]. Therefore, this checkpoint could prevent proliferation of damaged cells by inducing apoptosis. Second, this checkpoint could play a role in apoptosis during normal development. For example, many cells become apoptotic during development of the rat cerebral cortex [98]

Figure 1.2. The PI3K-PKB-FOXO signaling pathway proposed to be an important player in the early G1 checkpoint. PI3K activates PKB/Akt, which inactivates p21Cip1 and p27Kip1 by expelling them from the nucleus. PKB can also promote proliferation by removing the repression of transcription of cyclin D1 by inhibiting GSK-3b. PKB inactivates FOXO, which has anti-proliferative and proapoptotic roles. FOXO leads to cell cycle arrest via promoting transcription of p21Cip1 and p27Kip1 and repressing transcription of cyclin D, and promotes apoptosis by inducing transcription of proapoptotic and repressing transcription of prosurvival genes. PTEN acts upstream of this signaling cascade by inhibiting PI3K. PTEN phosphatase and tensin homolog; PI3K phosphoinositide 3-kinase; PKB protein kinase B; GSK-3b glycogen synthase kinase 3beta.

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and 50% of oligodendrocytes in the developing rat optical nerve undergo apoptosis after mitosis [99, 100]. Third, absence of growth factors could induce a withdrawal from the cell cycle. In vivo, unavailability of growth factors to cells may be most relevant in abnormal situations, as growth factors normally diffuse freely and reach most, if not all cells. It is possible, however, that cells desensitize growth factor receptors or regulate downstream pathway components that would give rise to the same phenotype as growth factor deprivation [1]. This could be in response to various physiological stimuli, such as detachment from the extracellular matrix or increased cell density. It is conceivable that cancer cells overrule this checkpoint and continue to grow even though stress or other events that would normally activate this checkpoint are present. Many of the regulators of cell cycle progression thought to be involved in this checkpoint are frequently mutated in tumors. Mutations of cyclin D and p21CIP1/p27KIP1 have been described in the previous section. However, alterations in any upstream pathway regulating the molecular core described at the beginning of the chapter may in potential be tumorigenic. PI3K and PKB have been implicated in this checkpoint and mutations in the PI3K-PKB signaling pathway are found in most tumors (rev. in 86, 88, 99, 100). Moreover, it seems conceivable that FOXO is also involved in this checkpoint. Therefore, this section will round off with a discussion of the mutations in these proteins in correlation with tumor growth. First, PI3K itself is frequently mutated in cancer, most often by amplification or activating mutations of the p110 catalytic subunit [101, 102]. This occurs for example in up to 50% of ovary, cervical, lung, and bowel cancers [101]. A negative regulator of the PI3K signaling pathway is the tumor-suppressor phosphatase with tensin homology (PTEN) (rev. in 99, 100). Mutations in the PTEN gene leading to its decreased expression are found in over 50% of several primary tumors, including glioma, melanoma, prostate cancer, endometrial cancer, breast cancer and leukemia [101]. Congenital defects in PTEN are also associated with several syndromes, most importantly Cowden syndrome, which is a hamartoma syndrome with increased disposition to thyroid, breast, and endometrial cancer [103]. In line with its central role in controlling proliferation and survival, PKB/Akt is also frequently


found overexpressed or overactivated in many human cancers. For example, the Akt gene is often amplified in ovarian cancer [104]. Furthermore, PKB was found overexpressed in over 60% of prostate cancers [105]. Nevertheless, mutations in the PKB gene itself are not very common [87, 101], indicating that its abnormal expression and activation is mainly the result of abnormal upstream signaling. Expression of constitutively active forms of FOXO1, FOXO3 or FOXO4 can suppress tumor growth in mice, and cytoplasmic localization of FOXO is associated with poor prognosis in breast cancer patients [106, 107]. Knock-out of only one member of the FOXO family does not induce cancer, probably due to compensation by other family members [88]. However, the deletion of FOXO1/3/4 in adult cells gives rise to thymic lymphomas and hemangiomas in mice [108]. In humans, FOXO is often present at chromosomal breakpoints in cancer, suggesting that the absence of a proper FOXO gene may promote tumorigenesis [88, 109–113]. In conclusion, recent experiments with cells synchronized in mitosis led to the discovery of a novel G1 checkpoint termed G0− [81], located immediately after mitosis. As inhibition of PI3K and PKB resulted in a similar arrest, these proteins may very well be central players in this checkpoint. Because of its important roles in apoptosis and cell cycle arrest, further studies should investigate the role of the PI3K/PKB/FOXO pathway in this checkpoint. As PI3K, PKB and FOXO are involved in sensing cellular stress, it will be of interest to investigate whether this checkpoint is truly another G1 stress checkpoint. Cancer cells can avoid this checkpoint in several ways, by mutations of cyclins or CKIs, and by mutations in upstream pathways affecting these central players.

1.4 The Restriction Point The most well-known checkpoint in the G1 phase is the restriction point (R). Already in the 1970s it was observed that cells were dependent on the presence of growth factors in the first part of G1, but lost this dependency a few hours before the onset of the S phase [114]. The term ‘restriction point’ (R) was coined to designate the point after which cells become committed to completion of the cell cycle regardless of the presence of growth factors [115].


Time-lapse analysis of mammalian cells in culture showed that R was somewhere between 3 and 4 h after mitosis [116, 117]. It was suggested that passage of R was determined by the accumulation of a labile protein following growth factor stimulation [115, 116, 118, 119]. Cyclin D fits these criteria. Its accumulation is highly regulated and it acts in conjunction with CDK4/6 to phosphorylate its main target, the retinoblastoma protein pRb (rev. in 120). pRb, together with p130 and p107, is part of the pocket-protein family. These proteins act as guardians of the cell cycle by controlling the E2F transcription factors ([121–124]; rev. in 125–127). During quiescence and in early G1, pRb is hypophosphorylated and sequesters the activating E2F family members (E2F1-3). At the same time, E2F4 and E2F5 are bound to p130 and inhibit transcription of E2Fresponsive genes, which is necessary for entry into S phase. When cyclin D binds to CDK4/6, the complex phosphorylates pRb, which starts releasing the activating E2Fs. This starts a positive feedback loop, in which cyclin E is transactivated by E2F, and – once bound to CDK2 – accelerates pRb phosphorylation and thus release of more E2F. p130 is also phosphorylated and subsequently ubiquinated, releasing the repressor E2Fs, which then probably are translocated from the nucleus to the cytoplasm. Growth factors are necessary for the induction of cyclin D and once pRb has begun to release E2F and cyclin E is being transcribed, cells lose their dependence on growth factors and are thus past the restriction point. However, if growth factors are absent earlier, cells get arrested at R and become quiescent or differentiate ([115, 116]; rev. in 1). As discussed earlier, depletion of growth factors is likely not commonly encountered in vivo. Nevertheless, other more physiological stimuli can also lead to an arrest at R, such as loss of adhesion, contact inhibition and possibly induction by differentiating factors [1]. Immediately after the identification of R, it was proposed that malignant cells have lost control of the restriction point [115]. Indeed, many cancer cells are less dependent on growth factors for proliferation (rev. in 128). Correspondingly, mutations in molecular components of the restriction point are the most common alterations in human cancer. As described at the beginning of the chapter, the expression of cyclins D and E is frequently elevated

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in cancer and mutations in p16 prevent inhibition of CDK4/6. Consistent with its central role in R, pRb is frequently mutated in a variety of cancers, including retinoblastomas, osteosarcomas and lung, renal and bladder carcinomas [127, 129]. Interestingly, pRbpositive lung cancer samples often lack p16INK4a, but pRb-negative samples may retain p16INK4A [130]. This illustrates that in tumorigenesis, mutations in components of the same signaling axis often are mutually exclusive or redundant. Altered levels of the E2F transcription factors are found in several types of cancer and often associated with poor prognosis [126]. Even though this often results from abnormalities in upstream signaling pathways, such as mutation of pRb, some mutations in the E2F gene itself have also been reported. For example, the E2F3 gene is often amplified and overexpressed in bladder carcinoma [131] and gene amplification of the E2F1 gene has been found in several cases of colorectal and gastric carcinoma [132, 133]. Moreover, the E2F4 gene contains a serine trinucleotide repeat, which is mutated in several cancers [134–136]. However, except for the latter, mutations in the E2F family are relatively rare [126]. This might be partly explained by the notion that E2Fs have various functions. For example, E2F1 also has a role in apoptosis (rev. in 126), so overexpression would not necessarily promote tumor growth. The central molecular core defining R is regulated by many different growth factor-dependent signaling cascades (Fig. 1.3). The most widely studied is the mitogen-activated protein kinase (MAPK) pathway. The MAPK family consists of different isoforms, each being activated by different extracellular signals and having different intracellular substrates. One of the most important pathways is the MAPK/ERK pathway [137]. An important player in this pathway is the small GTPase Ras, which is activated by growth factor receptors and then activates Raf. Via activation of MEK (MAPK- or ERK kinase) this leads to activation of extracellular signal-related kinases 1 and 2 (ERK1 and ERK2). ERK can phosphorylate targets in the cytoplasm or activate transcription factors in the nucleus. Activation of the MAPK pathway leads to the expression of cyclin D and plays a role in the degradation of p27. In this way, growth factor signaling leads to the activation of CDK4/6 and promotes passage through R. Ras can also play

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Figure 1.3. The MAPK signaling cascade promotes cell cycle progression via induction of cyclin D and phosphorylation of pRb. MAPK mitogen-activated protein kinase; GF-R growth factor-receptor; MEK MAPK- or ERK-kinase; ERK extracellular signal-related kinases; CDK cyclin-dependent kinase; pRb retinoblastoma protein; S genes and proteins downstream of the restriction point essential for entry into S phase.

a role in cell cycle progression via activation of the PI3K pathway [137]. Ras is a very potent oncogene and can transform cells without the aid of another oncogene [138]. It is mutated in a great variety of human cancers – albeit with varying incidence – where it accumulates in the GTP-bound, active form [139, 140]. Ras can also be activated by mutation of upstream growth factor receptors. Overexpression of members of the epidermal growth factor receptor family (EGFR and ERBB2) are most common [141]. Mutations in downstream targets of Ras also exist, such as activating mutations in BRAF, one of the members of the Raf family, especially in melanomas [142]. Common mutations in the PI3K pathway have been described in an earlier section and can contribute to evasion of the restriction point too. In conclusion, the restriction point is a crucial point in G1 that monitors the presence of growth factors. After phosphorylation of pRb and activation of cyclin E-CDK2, cells are no longer growth factordependent. In tumors, several mechanisms are used to avoid this checkpoint, varying from overactivation of the MAPK pathway to mutation of pRb and E2F itself. This explains why some cancer cell lines show less stringent growth factor requirements and why in vivo, tumors are not restricted by increased cell density which would normally arrest cells at R.

1.5 The Stress Checkpoint As discussed in the previous section, the cell has evolved mechanisms to transiently stop cell cycle progression in the absence of growth factors. However, there are also situations in which a permanent withdrawal from the cell cycle is desirable. Generally, the most important event that makes the cell unfit for further replication is DNA damage, which can occur in various situations, such as in the case of telomere attrition or oxidative damage (rev. in 143). After leaving the cell cycle upon stress, cells become apoptotic or senescent (rev. in 1, 143, 144). Interestingly, cancer cells seem to evade cell cycle arrest induced by all these forms of stress. In contrast to normal cells, tumor cells avoid cell cycle arrest upon DNA damage, allowing for genetic instability and further accumulation of tumorigenic mutations (rev. in 145). Furthermore, while normal cells can only divide a finite number of times due to shortening of telomeres, cancer cells are immortal [146–149]. Finally, as will be elaborated on in a later section, tumors feature elevated levels of reactive oxygen species. Apparently, mechanisms designed to arrest the cell cycle upon oxidative stress are evaded in cancer. These observations suggest that critical players in this checkpoint must be altered in cancer. This section will explore the

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molecular details of the stress checkpoint and how mutations therein can lead to cancer. A crucial factor in the stress checkpoint is p53. It plays a central role by integrating various stress stimuli and has a broad range of downstream targets, some of which lead to cell cycle arrest [150]. In the absence of stress, the protein MDM2 inactivates p53 by blocking its transcriptional activities and promoting its degradation (rev. in 36, 151). Normally, Mdm2 and p53 are kept in balance through a regu-latory feedback loop [151, 152]. Various forms of stress, however, can distort this balance. For example, several components of the DNA damage response, such as ATM (a critical sensor of DNA damage), CHK1/CHK2 (transducers of the DNA damage signal), and BRCA1 (involved in DNA repair) activate p53. ATM and CHK1/2 also target MDM2 (rev. in 145). As functional telomeres maintain the integrity and stability of DNA, telomere attrition can lead to DNA damage and can activate p53 as well (rev. in 153). The most important mechanism through which p53 can arrest the cell cycle is via induction of p21CIP1 [154–156]. Increased levels of this CKI prevent binding of cyclin E to CDK2. This arrests the cell cycle by preventing complete phosphorylation of pRb and by interfering with pRbindependent functions of cyclin E-CDK2 that are necessary for entry into S phase, such as aiding in the initiation of DNA replication by loading the MCM chromosome maintenance proteins to their origins of replication ([5]; rev. in 4). As p21Cip1 is the most important CKI involved in the checkpoint and cyclin E/CDK2 only plays a role after the initial phosphorylation of pRb has already occurred, the stress checkpoint has been situated temporally beyond the restriction point. Indeed, senescent cells showed chromosome condensation more similar to cells in late G1/S than in early G1/G0 [157]. Perhaps not surprisingly, together with the pRb, the p53 pathway is the most common deregulated pathway in cancer (rev. in 158). The p53 gene is inactivated in about 50% of all human tumors. The mutation signature is highly heterogeneous, varying from total gene deletions to mutants that have lost apoptotic or antiproliferative functions [158]. In line with its role as a negative regulator of p53, the MDM2 gene was found to be amplified in 19 different tumor types [159]. In addition, deregulation of upstream pathway components also affects levels

of activated p53 [145]. For example, mutation of ATM occurs in the genomic instability syndrome ataxia– telangiectasia that predisposes to cancer and is also found in other cancers, such as leukemia (rev. in 160, 161). The role of CHK in cancer seems to be more complex and evidence for the role of mutations in abnormal cell proliferation is mostly obtained in mouse models [145]. Yet, inherited mutations in one allele of the CHEK2 gene are found in some families with the Li-Fraumeni syndrome, which predisposes very strongly to cancer development [162]. Finally, inherited mutations in BRCA1/2 predispose women to ovarian and breast cancer [163]. In conclusion, this late G1 checkpoint plays an important role in halting cell cycle progression upon stress and may direct cells towards apoptosis or senescence. Mutations in various components of this checkpoint enable cancer cells to evade this checkpoint.

1.6 Hypermitogenic Arrest The checkpoint described in the previous section is located temporally beyond the restriction point and depletion of growth factors does not cause cell cycle arrest at the stress checkpoint. Nevertheless, an interesting idea proposed by Blagosklonny suggested that instead of the absence of growth factors, an arrest beyond the restriction point may result from an overstimulation of growth factors, i.e., a hypermitogenic arrest [164, 165]. This paradoxical statement may well be explained by the finding that many growth factor signaling pathways not only induce cyclins, but also CKIs, such as p21Cip1 and p16INK4A [164, 166–169]. For example, activation of Raf can lead to the expression of both cyclin D1 and p21Cip1 [167, 168]. The duration and intensity of signaling is essential for the effects of Raf: moderately strong signaling leads to induction of cyclin D1 and entry into S phase, while high-intensity signaling induced p21Cip1 and caused G1 arrest. This finding is interesting in light of the complex interaction of CKIs with the cell cycle machinery. As discussed in an earlier section, Cip/Kip CKIs can stabilize the cyclin D–CDK4/6 complex (rev. in 31). One could imagine that under moderate-intensity mitogenic signaling, Cip/Kip CKIs stimulate cell cycle progression by promoting cyclin D–CDK4/6 interactions. They would not

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inhibit CDK2 to a large extent, because they are sequestered by the cyclin D–CDK4/6 complex. However, when growth factor signaling increases, Cip/Kip levels also increase to such an extent that the amounts of CDK4/6 are no longer sufficient to sequester them. They are then free to inhibit cyclin E-CDK2 and thereby lead to cell cycle arrest. This would result in a hypermitogenic arrest, in which growth factor pathways are active and cyclins abundant, but where CDKs are inhibited by CKIs. A hypermitogenic arrest, in which the same signaling pathway can both lead to cell cycle progression and cell cycle arrest, seems to have evolved to prevent tumorigenesis [164, 165]. Cells facing persistent oncogenic stress are automatically shut down and enter senescence (Fig. 1.4). Cells cannot circumvent the hypermitogenic arrest by increasing growth factor signaling upstream of the inhibited kinases – this would in theory only strengthen the arrest [164, 165]. Hypermitogenic senescence can be avoided or reversed either by removing the inhibition, or by activating downstream components of the cell cycle machinery. Cancer cells are therefore likely to have abnormal activation of both upstream and downstream kinases and transcription factors [164]. For example, combinations of abnormally active Ras and


myc are particularly carcinogenic, as Ras provides the necessary growth factor signaling, while myc can act downstream of the hypermitogenic block [170]. Hypermitogenic arrest has been associated with senescence [164, 165], so it seems logical to temporally localize the hypermitogenic arrest after the restriction point, as senescence induced by the stress checkpoint has also been localized beyond R. However, at least three observations cast some doubt on this suggestion. Firstly, senescent cells expressing unphosphorylated pRb have been reported [171], suggesting that R – as defined in the classical sense – has not been passed yet. Secondly, there is also a role for p16INK4a in the induction of senescence (rev. in 1). For example, p16 is induced upon overactivation of the Ras pathway [166] and is involved in the induction of senescence [172]. Yet, the target of p16 (CDK4/6) is active before the passage of R. Thirdly, inactivation of pRb can prevent Ras-induced senescence [165]. These three observations suggest that early events that prevent phosphorylation of pRb by cyclin D-CDK4/6 may play a role in hypermitogenic senescence as well. Hypermitogenic arrest may well be an anti-cancer mechanism acting throughout the entire G1 phase. Before passage of R, hyperproliferative signals halt the cell cycle via the induction of CKIs, preventing phosphorylation of pRb. Yet, after passage of the restriction point, inhibition of CDK2 upon induction of CKIs by hypermitogenic signaling can lead to cell cycle arrest through inhibition of its pRb-independent functions required for entry into S phase.

1.7 AMPK and an Energy Checkpoint? Figure 1.4. Hypermitogenic arrest results from highintensity mitogenic signaling. Low-intensity signaling through the MAPK pathway both induces cyclin D and p21 expression. However, p21 does not inhibit cyclin E–CDK2 because it is sequestered by cyclin D–CDK4/6 complexes. Yet when the intensity of MAPK signaling increases, the amounts of p21 become so high that it can no longer be sequestered by cyclin D–CDK4/6. p21 then inhibits CDK2 and prevents entry into S phase.

As outlined in the previous two sections, the cell can exit the cell cycle in response to various forms of stress, including DNA damage, telomere attrition, and hypermitogenic stress. Another form of stress is energy depletion. Sufficient amounts of ATP are needed to complete the cell cycle and to sustain long-term viability of the cell and its daughter. A critical sensor of the cell’s energy state, AMP-activated protein kinase (AMPK), has been


implicated in regulation of the cell cycle [173–178]. Many cancer cells have an adapted, distinct metabolism and can grow under harsh conditions in terms of substrate availability (rev. in 179, 180). This section will discuss how cells can normally be arrested by this ‘energy checkpoint’ and how it may be evaded in cancer. When in energy-consuming cells the ratio of ADP to ATP increases, two molecules of ADP are converted to one molecule of ATP, thereby generating AMP as a sideproduct. The AMP:ATP ratio is therefore an indicator of the cellular energy state. AMPK is activated by AMP and is therefore a sensor of low-energy states (rev. in 181). In addition, AMPK can be phosphorylated by at least two upstream kinases, the tumor-suppressor LKB1 and the calcium-sensor CaMKKb [182–187]. Upon activation, it attempts to restore the cell’s energy state by inhibiting anabolic pathways and stimulating catabolic pathways ([176]; rev. in 181). It would be adaptive to transiently arrest the cell cycle when ATP levels are low to allow the cell to recover. Indeed, cells grown under limiting glucose conditions could not enter S phase, but when glucose was re-added the cells recovered from the arrest [176]. Apparently, AMPK was involved in this arrest, as expression of a dominant negative form of AMPK resulted in significantly more cells entering S phase despite low-energy conditions. Interestingly, p53 was shown to be necessary for the cell cycle arrest, as loss of p53 disrupted the arrest induced by glucose restriction. AMPK could activate p53 by phosphorylation at Ser-15 [176]. The involvement of p53 in this ‘energy checkpoint’ suggests that it is related to the p53-dependent stress checkpoint discussed in an earlier section. This is not so surprising when considering energy depletion simply as yet another form of cellular stress. Moreover, if the energy checkpoint would be situated earlier in G1, many events could still deplete energy after having passed the checkpoint. By placing the checkpoint just before entry into the S phase, a ‘last check’ can be performed before initiating DNA synthesis. As suggested by Blomen and Boonstra [1], p53 can induce an initial and reversible G1 arrest, whereupon further signals direct the cell towards apoptosis or senescence. Indeed, it was shown that the amount of MEF cells entering senescence after

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passage 4 was markedly increased by transfection of constitutively active AMPK, as shown by enhanced levels of senescence-associated b-galactosidase [176]. However, besides entering senescence, cells induced by p53 to exit the cell cycle can also become apoptotic. It has been suggested that the presence or absence of survival factors dictates the decision between senescence and apoptosis [1]. Interestingly, it has been found that upon energy depletion, AMPK can inhibit IRS-1 (insulin receptor substrate-1), which is a positive regulator of the PI3K/Akt pathway [177]. In line with the important role of this pathway in cellular survival, cells grown under prolonged glucoselimiting conditions became apoptotic [177]. It is conceivable that the decision of the cell to enter senescence or apoptosis depends on the severity of the insult: when energy levels are sufficient to maintain the viability of the cell, the cell expresses survival signals (such as the PI3K pathway) and becomes senescent. However, if energy levels become so low that the cell cannot be functionally sustained, survival signals will be inhibited and the cell will enter apoptosis. How AMPK exactly mediates cell cycle arrest remains unclear. While involvement of p53 and the PI3K pathway have been suggested [176, 177], a recent study showed that AMPK can also inhibit ERK, a key kinase in the MAPK pathway, as well as p70S6K, which promotes protein synthesis at the ribosome [178]. Involvement of AMPK in the Ras/ERK pathway has also been reported by others [174]. Future studies should elucidate the mechanisms in which AMPK can control the cell cycle to provide a better understanding of the relationship between energy supply and proliferation. Then how is this energy checkpoint relevant to cancer? Tumor cells can grow under harsh conditions with limited nutrient and oxygen availability and display a distinct metabolism (rev. in 179, 180). It was already shown decades ago that tumor cells favor glycolysis over oxidative phosphorylation, even in the presence of oxygen [188]. Initially, the option that this altered metabolism could be a cause of cancer was seriously considered. When evidence accumulated that cancer was caused by mutations, this idea was slowly discredited [179]. However, the dual role of AMPK as a cell cycle regulator and an energy sensor provoked renewed

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interest in the relation between metabolism, the cell cycle, and cancer. The role of AMPK in tumorigenesis is paradoxical, however. On the one hand, its activation can speed up metabolism thereby favoring growth, but on the other, it causes cell cycle arrest. Ashrafian provided a hypothesis in which this paradox could be resolved [189]. It is important to realize that adaptations in premalignant tumors might be different from those in advanced tumors. Premalignant cells normally have sufficient supplies of glucose and oxygen and therefore do not need AMPK activation. AMPK is therefore kept at basal levels or suppressed by excess glycogen. When substrate availability becomes a limiting factor for growth of advanced tumors, AMPK activation could provide a selective advantage by upregulating glucose uptake and glycolysis. The reason AMPK activation does not lead to cell cycle arrest may be explained by the fact that advanced tumors often show mutations in proteins that would actually mediate that arrest, such as p53. Mutations in genes coding for AMPK subunits have been found [190, 191], but no associations with cancer have been reported. Nevertheless, it was reported that pancreatic cancer cells better tolerated nutrient deprivation than hepatic cancer cells [192]. Interestingly, pancreatic cells expressed higher levels of AMPK and transfection of AMPK antisense decreased their tolerance to glucose deprivation and inhibited tumor growth in nude mice. In addition, incubation of some cancer cells with the AMPK-activator AICAR decreased cellular proliferation or survival [193]. The tumor-suppressor LKB-1 is an activator of AMPK [182–185]. Inactivating mutations have been found in patients with the Peutz–Jehgers syndrome, a genetic syndrome associated with multiple hamartomatous polyps and an increased risk of cancer development, especially in the digestive system (rev. in 194). Furthermore, in about 30% of patients with sporadic lung adenocarcinomas inactivating mutations in the LKB-1 gene have been found [195, 196]. The role AMPK plays in tumorigenesis is interesting because it could answer long-standing questions about the relation between metabolism and cancer. More research should be directed at the role of AMPK in the etiology of cancer, as it might open new therapeutic windows.


1.8 The Role of Reactive Oxygen Species in Cancer The previous sections scrutinized specific checkpoints. This last section, however, discusses an important player in the entire G1 phase (and beyond). An increasing body of evidence points towards a role for reactive oxygen species (ROS) in the control of the cell cycle. While ROS are most commonly regarded as toxic and unwanted side products of aerobic life, they deserve some attention in a discussion on cancer for at least three reasons. First, the concentration of ROS determines whether cells progress through the cell cycle or not: while low levels are required for cell cycle progression, higher levels elicit checkpoint responses and lead to cell cycle arrest. Second, ROS are tightly integrated in the cell cycle machinery. Third, cancer cells have higher ROS levels compared to normal cells. Reactive oxygen species consist of several kinds of reactive oxygen radicals and hydrogen peroxide, which can form the very reactive hydroxyl radical in a reaction with transition metals (rev. in 197). They normally arise as a side-product of various cellular processes, most notably during oxidative phosphorylation in mitochondria. In addition, ROS production can be a goal in itself, such as its production by the NADPH oxidase (NOX) system upon activation of phagocytic cells, resulting in a respiratory burst that kills invading pathogens (rev. in 69). Interestingly, NOX homologs have also been found in nonphagocytic cells (rev. in 69, 198), which will be discussed below. Since ROS are so reactive, they can easily oxidize various cellular components, such as lipids, proteins, and nucleic acids, and thereby lead to cellular damage [197]. To prevent this, the cell is endowed with a battery of defense molecules against oxidative damage, called antioxidants. Antioxidants can be enzymes, such as superoxide dismutase – which catalyzes the conversion of O2− to H2O2 – and catalase – which converts H2O2 into water. Antioxidants can also be small molecule buffers, such as the tripeptide glutathione, which reacts with H2O2 to change from its reduced (GSH) to its oxidized form (GSSG) [197]. For a long time it has been thought that ROS were mainly unwanted and potentially dangerous side-products, as illustrated by its implication in


various diseases [197]. Recently, however, a function for ROS in cellular signaling is beginning to be appreciated. One way in which ROS can affect cellular function is by changing the conformation of proteins through redox chemistry, for example, by changing the redox state of critical cysteine residues [199]. These residues might be more or less susceptible to redox modifications – much like differences in susceptibility to acid–base chemistry – giving rise to the notion that ROS can have specific effects depending on its concentration, exposure time, and the type of ROS involved. Indeed, at low concentrations ROS leads to enhanced proliferation, but when concentration and exposure time increase, ROS induce cell cycle arrest, apoptosis, or necrosis [197]. For example, when NIH3T3 fibroblasts were treated with a low concentration of H2O2, their proliferation increased; in contrast, cells treated with a higher concentration of H2O2 showed growth arrest and cellular death [200]. ROS thus act as a double-edged sword: low concentrations are required, but higher concentrations are detrimental for cell cycle progression. How can this be explained molecularly? As many proteins have sites sensitive to redox chemistry, ROS can modulate the activity of a wide variety of proteins. For example, ROS can induce phosphorylation of tyrosine growth factor receptors and inactivation of phosphatases, activate components of the MAPK pathway, and modulate various transcription factors [197]. Many of its effects in G1 seem to converge at the transcription of the cyclin D1 gene [201]. In line with its role in integrating various signals, the cyclin D1 promoter contains binding sites for many different transcription factors, including AP-1, NF-kB and Sp-1, which have all been shown to be regulated by ROS [201]. A good example is the regulation by ROS of c-Fos and Fra-1, members of the AP-1 transcription factor family that are activated by ERK (rev. in 69, 201). For expression of cyclin D1, degradation of c-Fos and nuclear translocation of Fra-1 is required. Transient or low levels of ROS cause replacement of c-Fos by Fra-1 and lead to cyclin D transcription. Constitutive or higher levels of ROS, however, lead to prolonged ERK activation which results in retention of c-Fos in the nucleus, preventing cyclin D expression. Low levels of ROS can thus promote cell cycle progression, but since ROS are toxic at higher

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concentrations, oxidative stress can induce cell cycle arrest, for example via the p53-dependent stress checkpoint [202]. Such a system, in which low-intensity signaling promotes cell cycle progression, but high-intensity signaling represses it, reminds one of the hypermitogenic arrest discussed in an earlier section. Could ROS be involved in this arrest? One requirement would be that growth factor signaling and ROS production are intimately linked. Interestingly, this seems to be the case of ROS production by NADPH oxidase homologs in nonphagocytic cells (rev. in 69, 198). In phagocytic cells, the membrane-localized NADPH oxidase complex (NOX) generates large amounts of superoxide, which is converted to H2O2 and in further reactions to other ROS, which can effectively kill invading pathogens (rev. in 203, 204). The discovery of NOX homologs in nonphagocytic cells has led to the idea that ROS produced by NOX may also serve other purposes (rev. in 69, 198). Consistent with the observation that low levels of ROS are required for cell cycle progression, the regulation of NOX can be linked to growth factor stimulation via the small GTP-binding protein Rac (rev. in 69, 198, 201). In phagocytic cells, NOX activation requires Rac [205, 206]. In nonphagocytic cells, EGF, PDGF, TNF-a, and IL-10 induce ROS production and this also depends on Rac, since a dominant negative Rac copy inhibited the increase in ROS production upon stimulation by these growth factors or cytokines [207]. In addition, it is known that PI3K, which can activate Rac, is required for PDGF-induced ROS production [208]. Finally, activation of NOX by EGF requires both PI3K and Rac1 [209]. The tight link between growth factor signaling and ROS production by NOX would lead to a situation in which normal signaling produces low amounts of ROS that can promote cell cycle progression. When the signaling becomes more intense and the concentration of ROS becomes toxic, checkpoints are activated and the cell cycle is arrested to prevent cellular damage (Fig. 1.5). Because its dual nature – toxic at high concentrations and a signaling molecule at low concentrations – ROS are ideally suited for a place in hypermitogenic arrest to prevent tumorigenesis. The role of ROS in oncogene-induced senescence and the hypermitogenic arrest will be an exciting area of future research [210].

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Figure 1.5. ROS are linked to the cell cycle machinery. Signals initiated by growth factor-receptors activate Ras and Rac, which directly contribute to mitogenic signaling. Simultaneously, Rac induces ROS production by NOX. When mitogenic signaling remains within certain limits, the concentration of ROS is such that it contributes to mitogenic signaling via redox modifications. However, when mitogenic signaling intensifies, this is reflected in the ROS concentration which reaches a threshold and arrests the cell cycle via induction of the stress checkpoint and by inactivation of cell cycle proteins by redox modifications. GF-R growth factor-receptor; NOX NADPH oxidase; ROS reactive oxygen species.

ROS are tightly interwoven in the control of the cell cycle. One could see ROS as a way to prevent the engine of the cell cycle from overheating by shutting it down prematurely once it starts running too fast. In cancer, however, this shutdown does not occur. Cancer cells have elevated ROS levels compared to normal cells [211–213], which can drive them through the cell cycle. One way in which they can elevate ROS levels is by overexpression of NOX1, as found in prostate cancer cells with increased tumor potential [214] and in adenomas and adenocarcinomas in the colon [215]. Since ROS have many targets, they may promote tumor growth in various ways, such as mitogenic signaling or resistance against apoptosis [69]. ROS are both a cause and a result of mitogenic signaling. In some situations, cancer cells are therefore favored by increased ROS production, as this would promote cell cycle progression. However, in other situations, the oxidative environment created by increased ROS production due to mitogenic signaling may be maladaptive. For example, in the nucleus, a reducing environment is favored for binding of transcription factors and gene expression [213]. Therefore, another way in which cancer cells may respond to elevated ROS levels is by upregulation


of antioxidants. For example, thioredoxin reductase 1 (TR1) is upregulated in many malignant cells and knocking out TR1 is antitumorigenic [216]. Apparently, sometimes upregulation of antioxidants is favored, while at other times an increase in ROS is tumorigenic. This may depend on the type of ROS or antioxidant involved, as well as the moment and cellular location of its production, concentration, and exposure time. It remains to be investigated how the balance between ROS and antioxidants is regulated precisely in tumors.

1.9 Conclusions In conclusion, the cell has to successfully pass several checkpoints in the G1 phase of the cell cycle to enter the S phase (Fig. 1.6). The various checkpoints encountered in this chapter each have different inducing signals as well as different outcomes. Firstly, various forms of stress can make cells unfit for further replication, making it adaptive to direct these cells towards senescence or apoptosis. The novel checkpoint early after mitosis may serve this role, in addition to the more traditional p53-dependent stress checkpoint. Moreover, a checkpoint in which AMPK plays a central role ensures the cell only enters S phase if enough energy is available. Secondly, the cell depends on the presence of growth factors until it has passed the restriction point. Desensitization of growth factor signaling pathways in response to, for example, detachment from the extracellular matrix may provide a physiological stimulus to arrest cells at R. Thirdly, besides insufficient mitogenic signaling, cells may also be arrested in response to hypermitogenic stress, as a mechanism to prevent tumorigenesis. The production of ROS is essential for many events in G1 phase progression, but may be particularly relevant to this hyperproliferative arrest. Cancer is characterized by uncontrolled cell proliferation. Various checkpoints can impede this proliferation and therefore cells with mutations that render cells insensitive to these checkpoints gain a selective advantage with regard to proliferation. Direct mutation of components of the essential molecular core described in an earlier section is an effective way to evade these checkpoints, which is for example the case with the many mutations in p16INK4A and cyclin D1. As the case with the

19


Figure 1.6. A cell progressing through the cell cycle encounters various checkpoints in G1. Shortly after mitosis, the cell meets an early G1 checkpoint, in which PI3K signaling plays a dominant role. In mid-G1, the restriction point monitors the presence of growth factors; pRb–E2F complexes are important elements of this checkpoint. Beyond R, a p53-dependent stress checkpoint controls the integrity of the cell. Finally, an energy checkpoint in which AMPK plays a key role controls whether the cell has enough energy to complete the rest of the cell cycle. It is proposed that all throughout G1, a hypermitogenic arrest in which ROS signaling could be an important factor may occur.

Cip/Kip CKIs, however, it is apparent that mutation of the genes itself is not always favorable because some functions of the protein need to be retained for cell cycle progression, while other functions can be dismissed. Some questions about the nature of these checkpoints or their role in tumorigenesis remain open. Particularly of interest are the relatively ‘new’ checkpoints. Much remains to be learned about how the early G1 checkpoint operates molecularly and the signals of which activate it in vivo. The discovery of this checkpoint also illustrates the importance of studying the ongoing cell cycle in addition to cells entering G1 from quiescence. Another novel checkpoint is the energy checkpoint, which needs further molecular characterization. Its role in cancer is especially interesting considering the altered metabolism of cancer cells. A better understanding of how AMPK activation or inactivation may play a role in various stages of cancer may help explain this interesting observation [189]. The concept of a hypermitogenic arrest as a mechanism to prevent tumorigenesis is relatively new and especially the role ROS play in this arrest can be further explored. ROS are tightly linked to the cell cycle via NADPH oxidase homologs, but mitochondrial ROS production may also be relevant for G1 phase progression. Investigating the interaction between mitochondrial activity and cell cycle progression may be a

fruitful undertaking in the understanding of cancer. Finally, the central molecular core underlying G1 phase progression, consisting of CDKs, cyclins, activating enzymes, and CKIs, remains surprising, even after decades of thorough investigation. New roles for Cip/Kip CKIs may alter our view on regulation of the cell cycle. All in all, the complexity of G1 phase regulation and the many ways in which it is altered in cancer will keep inspiring researchers for many years.

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Chapter 2

Distinct Pathways Involved in S-Phase Checkpoint Control Paula J. Hurley and Fred Bunz

Abstract The S-phase checkpoint is activated when DNA damage occurs during DNA synthesis or when DNA replication intermediates accumulate. Depending on the type and magnitude of damage, cells activate one of the three distinct S-phase checkpoint pathways: (1) an intra-S-phase checkpoint induced by double strand break, (2) a replication checkpoint by the stalled replication fork, and (3) a S–M checkpoint to block premature mitosis. These checkpoint pathways coordinate a network of signaling molecules and are thought to ensure the fidelity of the replicating genome. Keywords Intra-S-phase checkpoint • Replication checkpoint • S–M checkpoint • ATM • ATR • Chk1 • Chk2 • Double strand break • DNA replication

2.1 Introduction Cellular proliferation involves numerous processes that are tightly regulated to maintain the integrity of the genome. Complete and faithful DNA replication before cellular division is essential. In human cells, this tightly monitored and highly controlled process involves the duplication of the entire diploid genome, consisting of nearly six billion base pairs, during the relatively brief period of S-phase. Damage to replicating DNA can arise from both endogenous sources such as reactive metabolites or replication intermediates and exogenous sources such as ionizing radiation (IR), ultraviolet (UV) irradiation, and chemicals

that intercalate between the DNA strands, form adducts or interstrand crosslinks. Eukaryotes have developed genetically defined surveillance mechanisms called “checkpoints” that monitor many aspects of cell growth and division. This chapter focuses on the S-phase checkpoint that monitors the progress of DNA replication and prevents damaged DNA from being replicated [1]. Checkpoints protect the replicating cell from genotoxic stress by coordinating DNA replication with DNA damage sensing processes, DNA repair, and cell cycle progression. In response to DNA damage, cells activate pathways leading to apoptosis, senescence, or survival and cell cycle resumption. Germline inheritance of mutated checkpoint genes confers cancer predisposition, and sporadic tumors commonly accumulate checkpoint defects. These observations provide evidence that altered cell cycle regulation contributes to tumorigenesis [2].

2.2 Unperturbed S-Phase Fork Initiation and Progression A large number of highly specialized molecules function coordinately to replicate genomic DNA efficiently. Eukaryotic genomes contain numerous origins of replication that fire at different intervals during S-phase. Replication origins have differential firing times that can be categorized as early, intermediate, and late. At any given time during S-phase, some origins may be progressing while


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others have finished or have yet to fire. When origins are fired, protein–DNA complexes form structures known as replication forks, which proceed from the origin in a bidirectional manner [3]. Before the initiation of DNA synthesis, protein complexes that facilitate DNA unwinding and the subsequent loading of DNA polymerase complexes assemble at replication origins. This critical process, known as licensing, occurs before S- phase, during G1 [4, 5]. At the initial stage, a protein complex known as the origin of recognition complex (ORC) assembles each origin of replication. The ORC is a complex of six subunits that serves as an essential scaffold for the assembly of other replication proteins on origin-containing chromatin. Following ORC assembly, cell division cycle 6 (Cdc6), an ATPase, and then Cdt1 bind ORC. Together, these two proteins load the putative DNA helicase, the enzyme responsible for unwinding the pare-ntal duplex DNA, onto the origin [6, 7]. Through the hydrolysis of ATP, Cdt1 and Cdc6 mediate the binding of the mini-chromosome maintenance (MCM) proteins to the origin of replication [8, 9]. MCM proteins, designated MCM2–7, form a highly conserved heterohexameric complex essential for the final phase of origin licensing [10]. The MCM complex is thought to be the catalytic core of the helicase necessary for DNA unwinding ahead of the advancing replication fork [11, 12]. MCM proteins 2–7 loaded onto the chromatin form the pre-replication complex (pre-RC) that is the defining characteristic of a licensed origin. Although the origin is licensed once the pre-RC binds the chromatin, the MCM complex remains inactive until the initiation of DNA replication. Activation of MCM2–7 is not fully understood, but it involves the recruitment of several proteins to the pre-RC during S-phase. Cyclin-dependent kinase (Cdk) and Cdc7/DBf4-dependent kinase bind the pre-RC and are necessary for the loading of Cdc45. Cdc45 functions as both an initiation and an elongation factor. Cdc45 stably associates with MCM proteins at the fork and is required for duplex unwinding at the fork and for elongation of the nascent strands [13]. Firing of DNA replication origins occurs once, and only once, during each cell cycle. To prevent refiring of replication origins once S-phase has begun, MCM dissociates from chromatin and a protein known as Geminin binds to Cdt1 and

P.J. Hurley and F. Bunz

prevents its interaction with the MCM complex, thereby inhibiting further MCM2–7 recruitment [14, 15]. Once initiated, DNA replication essentially consists of two tightly coupled processes: unwinding of the parental duplex DNA, and simultaneous synthesis of the nascent DNA on the leading and lagging strands. Chromatin unwinding allows the replicative DNA polymerases to access the DNA template. Eukaryotic DNA replication depends on three DNA polymerases: e, a, and d. DNA Polymerase e is required for the continuous synthesis of the leading strand. DNA Polymerase a/primase generates the short RNA/DNA primer needed for DNA synthesis on both the leading and the lagging strands, while DNA Polymerase d is required for the complete synthesis of the Okazaki fragments on the lagging strand. Proliferating cell nuclear antigen assembles into a sliding clamp, required for the processivity of DNA Polymerase d, that is loaded onto the chromatin by the clamp loader ATPase RF-C. The unwinding of DNA by the replicative helicase exposes short sections of single stranded DNA (ssDNA), which if exposed to nucleases would be susceptible to degradation. Replication protein A (RPA) tightly binds and thus protects ssDNA. These highly conserved proteins, along with a number of accessory factors, compose the molecular machines that replicate genomic DNA.

2.3 S-Phase Checkpoint Overview The progress of DNA replication is very closely monitored. Stalled replication forks or DNA lesions that occur during S-phase, and which could impair fork progression and thereby impede DNA synthesis, generate signals that are transduced throughout the cell [16]. Among the signaling pathways that respond to such stimuli are the S-phase checkpoints. Three distinct S-phase checkpoint responses have been identified: the intra-S-phase checkpoint, the replication checkpoint and the S–M checkpoint [1]. The intra-S-phase checkpoint is activated when double strand breaks occur in the genome outside actively synthesizing replicons, thus activation of this checkpoint is independent of the replication fork. In contrast, the replication checkpoint occurs at replication forks that have stalled due to inhibition

2. Distinct Pathways Involved in S-Phase Checkpoint Control

of DNA polymerases, depletion of deoxyribonucleotides (dNTPs), or collision with damaged or aberrantly structured DNA. As with the replication checkpoint, the S–M checkpoint is dependent on the replication machinery and ensures that cell division does not occur until the genome is fully duplicated [1]. The different S-phase checkpoints share some of the same signaling molecules, but are elicited under distinct experimental conditions (Fig. 2.1). Unlike other checkpoints that can invoke either a permanent or a transient cell cycle arrest, the S-phase checkpoints are generally transient in nature. The length of delay depends of the type and magnitude of the genomic insult. DSBs caused by stimuli such as IR, trigger the intra-S-phase checkpoint and cause DNA synthesis inhibition in a dose dependent manner that can last a few hours.

Figure 2.1. S-Phase checkpoints. Depending on the type and the degree of DNA damage, three distinct S-phase checkpoints can be activated: (1) the S–M checkpoint, (2) the replication checkpoint (RC), and (3) the intra-Sphase checkpoint (ISC). The S–M checkpoint activates a delayed cell cycle arrest of cells damaged during S-phase and thus ensures that the genome is fully replicated before the onset of mitosis. The replication checkpoint is activated when replication forks stall due to damaged DNA or to inhibition of replication mediators, and is consequently fork dependent. The intra-S-phase checkpoint is independent of the replication fork, as it is activated by DSB occurring in the genome outside of active replicons.

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The S–M checkpoint is experimentally observable only after several hours [17]. Similarly, the replication checkpoint can cause longer delays in DNA replication.

2.3.1 The Intra-S-Phase Checkpoint Even before the development of the checkpoint concept by Hartwell and colleagues [18], it was observed that exposure of human cells to IR causes a rapid and marked reduction in DNA synthesis [19, 20]. By incubating asynchronous cell populations with radiolabeled thymidine, investigators noted that IR treatment caused a dramatic reduction in thymidine incorporation and therefore a transient halt in DNA replication [20]. The DNA replication pause in response to IR is not caused by the encounter of a moving replication fork with a DSB. Rather, DSBs caused by IR trigger an active process that halts DNA replication throughout the nucleus. The IR-dependent pause in DNA replication was, in retrospect, the first observation of a checkpoint in human cells. The intra-S-phase checkpoint is activated by DSBs, which in the laboratory or clinical setting are typically generated by exposure to IR or to a class of drugs known as radiomimetics. The agents that cause these relatively severe lesions are sometimes referred to as clastogens, defined as agents that cause cytogenetically observable chromosome breaks. However, DSBs are also generated endogenously by physiological processes such as DNA replication and recombination. If left unrepaired, DSBs can lead to chromosome translocations. DSBs can occur anywhere in the genome, and are most often distant from active replication forks. That such breaks are sufficient to transiently suppress DNA replication indicates that the origin of the DSB signal is DNA replication fork independent. IR induces a transient and a partial inhibition of DNA synthesis. Following IR, essential initiation proteins are not recruited to the origin of replication. Active replication forks are mostly unaltered by the DSB response; IR does not inhibit the rate of fork progression nor increase fork stalling [21]. Thus, the delay in S-phase progression can be attributed primarily to an inhibition of unfired replication origins [1]. The continued progression of actively synthesizing replication forks accounts

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for the significant amount of thymidine that is still incorporated into IR treated cells. Genetic defects in what is now understood to be the intra-S-phase checkpoint were first seen in patients with the autosomal recessive disease ataxia-telangiectasia (AT) [19, 20, 22]. Individuals with AT are highly sensitive to IR and are predisposed to cancers, particularly to leukemias and lymphomas. The collective symptoms in AT patients, which also include immune deficiency and cerebellar ataxia highlight the important role that the DNA damage responses play in diverse cellular processes and diseases. Cells from AT patients exhibit a diminished capacity to inhibit DNA synthesis in response to IR. The phenomenon of undiminished DNA replication after IR was termed “radioresistant DNA synthesis” (RDS) [19]. RDS is a direct indication of a defective intra-S-phase checkpoint and is diagnostic for AT. The cell-based assay first used to detect RDS has since served as the gold standard to detect S-phase checkpoint deficiencies. Unperturbed cells are initially incubated with 14C-thymidine. This label is incorporated into cells that are actively replicating DNA and therefore reflects both the total cell number and the overall rate of cell proliferation. After removal of the unincorporated 14C-thymidine label, cells are treated with IR and then incubated briefly with 3H-thymidine. The ratio at which these two isotopes are incorporated allows for the comparison of the level of DNA synthesis before and after IR treatment. This is a quantitative assay, but it has an inherent level of background due to the incorporation of thymidine during the ongoing elongation of previously fired origins. This background can limit the detection of partial RDS. Recently a more descriptive assay alternatively called DNA fiber labeling or DNA combing has been used to examine the intra-S-phase checkpoint. Unlike the RDS assay, this technique can assess dynamic features of individual DNA replication forks. DNA combing utilizes two thymidine analogs that can be differentially detected by florescent in situ hybridization. Cells are initially labeled with one thymidine analog such as 5-bromo-2deoxyuridine (BrdU). After removing unincorporated BrdU, cells are exposed to IR and then labeled with a different thymidine analog such as 5-iodo-2-deoxyuridine (IdU) [23]. Following treatment, DNA spreads are made by spotting cells on


a glass slide and then lysing those cells in situ. The slides are then tilted to allow DNA fibers to spread over the slide surface [24]. Incorporated thymidine analogs are detected by the use of fluorescent antibodies (green for BrdU and red for IdU, for example). Active replication forks can be visualized as labeled tracks under a fluorescence microscope. The length of each green and red track is a rough indicator of fork velocity before and after IR, respectively. Ongoing forks are labeled in two colors while a stalled fork will only be labeled with one color, indicating that the second analog was not incorporated at that DNA replication fork after IR treatment (see Fig. 2.2). A typical analysis involves the quantification of hundreds of individual DNA replication forks. Detailed analysis of the number and length of the fluorescent labels allows for the detection of fork slowing, inhibited origin firing, and fork stalling.

Figure 2.2. DNA fiber labeling. DNA fiber labeling is a tool used to examine replication fork dynamics. S-phase cells are initially labeled with a thymidine homolog such as BrdU. After removal of unincorporated label, cells are treated with IR and then incubated with a second thymidine analog such as IdU. After cell membranes are disrupted, DNA strands are “combed” to elongate labeled tracks. Differentially colored fluorescent antibodies are used to detect thymidine analog incorporation. The length of second label incorporation correlates to the rate of the replication fork after IR treatment. Absence of the second label is an indication of fork stalling. Incorporation of second label alone identifies new replication origins that fire after IR treatment.


2.3.1.1 Molecules Involved in the Intra-S-Phase Checkpoint Advanced assays such as DNA fiber labeling have aided in the identification and characterization of intra-S-phase checkpoint mediators. This section will give a brief overview of some of the molecules involved in this process. The molecular defect that causes AT, biallelic mutations in ataxia-telangiectasia mutated (ATM), points to the central importance of the ATM protein in the intra-S-phase checkpoint. Similar to other checkpoints, the intra-S-phase checkpoint pathway is the composed of sensor, transducer, and effector components. ATM serves as a proximal sensor of DSBs. As a member of the phosphoinositide 3-kinase-like kinase family (PIKK), ATM is a high molecular mass serine/threonine kinase [25]. In response to DSBs, ATM triggers the phosphorylation of over 700 proteins at more than 900 known sites, and can thus affect cellular processes as diverse as cell motility and replication origin licensing [26]. In unperturbed cells ATM exists as an inactive dimer. DSBs lead to ATM autophosphorylation at several residues including serine 1981 and concurrent monomerization and partial activation of kinase activity [27]. Phosphorylation at 1981 marks active ATM, but its necessity for ATM activation has been called into question [28]. Once activated, ATM becomes tightly associated with damaged chromatin. Full ATM activation is facilitated by a protein complex composed of MRE11, Rad50, and NBS1, known as the MRN complex. Notably, mutations in MRE11 and NBS1 cause ataxia-telangiectasia-like disorder and Nijmegen breakage syndrome, respectively. These disorders clinically overlap with AT and similarly feature immunodeficiency, radiosensitivity, and increased cancer risk [29]. The MRN complex localizes to the DSB independent of ATM and other known checkpoint proteins, and it may also serve as a sensor of the DSB. Nbs1 binds to ATM and is involved in the recruitment of ATM to the DSB [30–32]. ATM, in turn, phosphorylates proteins in the MRN complex [33]. Cells deficient for Nbs1 or for the ATM-dependent phosphory-lation of Nbs1 are defective for the intra-S-phase checkpoint and exhibit an RDS phenotype [33, 34]. The reciprocal interactions between ATM and the MRN

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complex in a common DNA damage response pathway explains the overlapping nature of the rare disorders caused by inactivation of ATM, Nbs1, Mre11, or Rad50. ATM signaling leads to the inhibition of unfired competent replication origins throughout the S-phase nucleus. ATM affects origins of DNA replication that may be distant from the DSB site through a network of signaling molecules. The primary transducer of ATM signaling is checkpoint kinase 2 (Chk2) [35]. Studies of cells derived from knockout mice have shown that like ATM deficiency, Chk2 deficiency causes RDS, demonstrating conclusively the importance of Chk2 in the intra-S-phase checkpoint [36]. Chk2 mediates ATM signaling by phosphorylating the Cdc25A phosphatase. Cdc25A is targeted for proteasomemediated degradation by Chk2, thus causing Cdc25A inhibition [37]. During unperturbed proliferation, Cdc25A removes inhibitory phosphate moieties on Cyclin Dependent Kinase 2 (Cdk2), thus maintaining Cdk2 in an active state. Cdk2dependent loading of CDC45 onto the chromatin is necessary for DNA polymerase a binding. When Cdk2 is inhibited by the Chk2-mediated degradation of Cdc25A, DNA polymerase a is not loaded and late firing origins are effectively inhibited. Thus, through a well defined signaling pathway, ATM activation by DSBs inhibits the firing of late origins and leads to a transient pause in DNA synthesis during S-phase. In response to IR, ATM indirectly activates another PIKK family member, ATM- and Rad3-related (ATR) [38–42]. ATR is recruited to IR-induced DSBs subsequent to ATM activation [38]. Enzymatic cleavage of DSBs during the early stages of DNA repair generates RPA coated ssDNA, which then stimulates ATR recruitment to the chromatin through its binding partner ATR interacting protein (ATRIP). The contribution of ATR to the intra-S-phase checkpoint is debated. ATM-dependent ATR activation in response to DSBs occurs in cells that are in S phase, suggesting that ATR could contribute to the intra-S-phase checkpoint [41]. ATR is the predominant activator of the checkpoint kinase Chk1 [43]. Like Chk2, Chk1 can mediate Cdc25A degradation [44]. Cells with biallelic hypomorphic ATR mutations do not display RDS, which provides strong evidence that ATR does not play a significant role in the intra-S-phase checkpoint [40].

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Little is known about recovery from the intra-Sphase checkpoint and resumption of DNA synthesis than about its activation. During the period in which origin firing is suppressed, Claspin, which is necessary for Chk1 activation, is phosphorylated by polo like kinase 1 (Plk1) and is consequently marked for proteosome-mediated degradation by the SCF (b-TrCP) ubiquitin ligase complex [45–47]. How Plk1 is activated is not currently known. Following Claspin degradation, Cdc25A protein levels increase and presumably the recovery of Cdc25A levels promotes the resumption of DNA synthesis. The mechanism of recovery from the S-phase checkpoint remains an active area of study.

2.3.2 The Replication Checkpoint In addition to DSBs, many other insults to the chromatin or to the replication machinery can occur during S-phase. Replication stress during S-phase in the form of damaged DNA, aberrant DNA structures, DNA polymerase inhibition, or dNTP inhibition can cause stalling of the replication fork [16]. Replication fork stalling can also arise in unperturbed cells when the fork encounters replication slow zones, repetitive sequences, or fragile sites [48–50]. Upon stalling, the replication fork must remain associated with the replisome in order to allow subsequent resumption of replication progression following impediment removal or bypass. Collapse of the replication fork will cause disassembly of the replisome and consequent replication termination. Collapse of one fork may be rescued by proximate fork replication through the collapsed fork, but extensive fork collapse that cannot be overcome by neighboring forks can lead to breaks or gaps in the chromatin. Fork dynamics can be visualized by DNA fiber labeling as described above or by two-dimensional (2D) gel electrophoresis. 2D electrophoresis resolves replication intermediates based upon their mass and topology [51, 52]. As DNA is replicated, it doubles in mass and forms bubbled or branched structures, depending on the proximity of the fork to the replication origin. Varying gel electric field and agarose content in two separate dimensions allows for initial separation by mass and subsequent separation by topology. Replication intermediates are then visualized by Southern blot.


Eukaryotes respond to replication stress by activating the replication checkpoint. Like the other S-phase checkpoints, the replication checkpoint functions to monitor DNA integrity and replication fidelity and to coordinate replication with repair and cell cycle progression. The replication checkpoint is activated upon stalling or collapse of the replication fork and leads to inhibition of late origin firing, maintenance of fork stability, promotion of DNA repair, and re-initiation of stalled forks once repairs are completed [53]. Replication checkpoint mutant yeast exhibit increased genomic rearrangements when exposed to replication inhibitors, suggesting that the replication checkpoint may be similarly essential for maintaining the integrity of human chromosomes [54, 55]. In contrast to the intra-S-phase checkpoint, replication checkpoint-induced signaling depends predominantly on ATR and its downstream effector kinase Chk1. Both ATR and Chk1 function during unperturbed cell growth as well as in the DNA damage response. Much research is focused on uncoupling the dual roles of ATR and Chk1. The genes that encode ATR and Chk1 are essential for viability in mouse [56–59] as well as human cells [60]. The loss of function of either gene leads to cell cycle arrest and death even in the absence of exogenous DNA damage. Deficiency of ATR or Chk1 leads to the accumulation of replicationassociated chromosome breaks [61]. Insights into the role ATR plays in the unperturbed cell cycle have been gained by studies of Mec1 (ATR homolog in S. cerevisiae) mutants. Mec1 has been shown to regulate dNTP synthesis. In the absence of exogenous stress, lethal Mec1 mutants can be rescued by augmenting dNTP levels [62]. In human cells, insufficient dNTP pools cause fork stalling [21]. ATR may be necessary for dNTP synthesis and/or fork stabilization in the absence of sufficient dNTPs. Chk1 also regulates dNTP synthesis. However, modulation of dNTP levels in Chk1 depleted mouse embryonic fibroblasts (MEFs) does not rescue them from S-phase arrest, suggesting that additional ATR/Chk1 regulated pathways in mammalian cells contribute to cellular viability in the absence of exogenous stress [63]. In addition to its role in unperturbed cell proliferation, ATR maintains fork integrity in response to replication inhibition. DNA damage-induced fork stalling destabilizes the replisome and thus


inhibits replicon recovery. ATR appears to function to stabilize the replisome and prevent fork collapse [64, 65]. In the absence of ATR, replicons are unable to recover from many DNA damaging agents. Stalled forks generate large areas of RPA coated ssDNA, which recruits ATR, through its binding partner ATRIP, to the chromatin [66]. Mutations in the yeast ATR homolog Mec1 cause the loss of DNA polymerases from stalled forks [67]. MCM2, a component of the replicative helicase, normally advances with the replication fork, but is irreversibly lost from the chromatin during replication stress in the absence of ATR [68]. ATR phosphorylates MCM2 in response to multiple DNA damaging agents, but the cellular outcome of this regulation is not known [26, 69, 70]. By protecting MCM and DNA polymerases at the replisome, ATR may protect the fork by preventing uncoupling of DNA unwinding and DNA synthesis. Chromatin bound ATR activates the transducer kinase Chk1 through phosphorylation at serines 317 and 345 [43]. Like ATR, Chk1 is necessary for maintaining the replisome at stalled forks, but Chk1 does not appear to facilitate all of the functions of ATR [53, 71]. This suggests that ATR activates additional transducers of the replication checkpoint. ATR may also function in the replication checkpoint by stimulating DNA repair. In response to replication stress, ATR directly interacts with members of several DNA repair pathways. ATR phosphorylates FANCD2, a member of the Fanconi Anemia pathway, as well as BRCA1 and BRCA2 [72–75]. FANCD2, and BRCA2 are important for the repair of interstand DNA crosslinks, and BRCA1 and BRCA2 are regulatory components of protein complexes that facilitate homologous recombination-mediated repair. Following damage repair, checkpoint recovery involves restarting of impeded replication forks and firing later replication origins. Checkpoint recovery may be partially mediated by Chk1 downregulation. Immediately following ATR induction, Chk1 is activated, but is subsequently degraded several hours later [76]. Investigation of this phenomenon has revealed that phosphorylated Chk1 is targeted by the SCF ubiquitin ligase complex for proteasome-mediated degradation. Interestingly, both activation and destabilization of Chk1 are regulated through phosphorylation of serine 345.

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2.3.3 The S–M Checkpoint The S–M checkpoint ensures that cellular division does not occur until the genome is fully replicated, and thereby blocks premature entry into mitosis. This checkpoint differs from the well defined G2/M checkpoint in that it is ATM independent, is measurable only several hours after DNA damage, and is initiated in cells that were in S-phase at the time of insult [17]. Similar to the replication checkpoint, the S–M checkpoint is triggered at the replication fork and involves many of the same mediators. The role of Chk1 in this checkpoint varies between organisms and experimental systems. Fission yeast doubly deficient, but not singly deficient, for cds1 (Chk2 homolog) and Chk1 lack an effective S–M checkpoint, suggesting that Chk2 and Chk1 may have overlapping roles in the S–M checkpoint [77]. DT40 lymphoma somatic cells deficient for Chk1 enter mitosis with incompletely replicated DNA when DNA synthesis is blocked [78] and Chk1 deficient mouse blastocysts also enter mitosis in the presence of replication inhibitors, suggesting that Chk1 is necessary for the S–M checkpoint in vertebrates [59]. However, conditional deletion of ATR in MEFs and failure of Chk1 activation does not cause premature mitosis [79]. As with the other S-phase checkpoints, the S–M checkpoint targets CDC25A for degradation, and consequently inhibits Cyclin B-Cdk1, which is needed for mitotic entry. The role of this pathway in cell survival is not fully understood and remains an area of active interest. Acknowledgment: This work was supported by the Flight Attendant Medical Research Institute (P.J. Hurley and F. Bunz).

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52. Ferguson BM, Brewer BJ, Reynolds AE, Fangman WL (1991) A yeast origin of replication is activated late in S phase. Cell 65:507–515 53. Paulsen RD, Cimprich KA (2007) The ATR pathway: fine-tuning the fork. DNA Repair (Amst) 6:953–966 54. Myung K, Datta A, Kolodner RD (2001) Suppression of spontaneous chromosomal rearrangements by S phase checkpoint functions in Saccharomyces cerevisiae. Cell 104:397–408 55. Myung K, Kolodner RD (2002) Suppression of genome instability by redundant S-phase checkpoint pathways in Saccharomyces cerevisiae. Proc Natl Acad Sci USA 99:4500–4507 56. Brown EJ, Baltimore D (2000) ATR disruption leads to chromosomal fragmentation and early embryonic lethality. Genes Dev 14:397–402 57. de Klein A, Muijtjens M, van Os R et al (2000) Targeted disruption of the cell-cycle checkpoint gene ATR leads to early embryonic lethality in mice. Curr Biol 10:479–482 58. Liu Q, Guntuku S, Cui XS et al (2000) Chk1 is an essential kinase that is regulated by Atr and required for the G(2)/M DNA damage checkpoint. Genes Dev 14:1448–1459 59. Takai H, Tominaga K, Motoyama N et al (2000) Aberrant cell cycle checkpoint function and early embryonic death in Chk1(−/−) mice. Genes Dev 14:1439–1447 60. Cortez D, Guntuku S, Qin J, Elledge SJ (2001) ATR and ATRIP: partners in checkpoint signaling. Science 294:1713–1716 61. Syljuasen RG, Sorensen CS, Hansen LT et al (2005) Inhibition of human Chk1 causes increased initiation of DNA replication, phosphorylation of ATR targets, and DNA breakage. Mol Cell Biol 25:3553–3562 62. Desany BA, Alcasabas AA, Bachant JB, Elledge SJ (1998) Recovery from DNA replicational stress is the essential function of the S-phase checkpoint pathway. Genes Dev 12:2956–2970 63. Naruyama H, Shimada M, Niida H et al (2008) Essential role of Chk1 in S phase progression through regulation of RNR2 expression. Biochem Biophys Res Commun 374:79–83 64. Lopes M, Cotta-Ramusino C, Pellicioli A et al (2001) The DNA replication checkpoint response stabilizes stalled replication forks. Nature 412:557–561 65. Tercero JA, Diffley JF (2001) Regulation of DNA replication fork progression through damaged DNA by the Mec1/Rad53 checkpoint. Nature 412:553–557 66. Sogo JM, Lopes M, Foiani M (2002) Fork reversal and ssDNA accumulation at stalled replication forks owing to checkpoint defects. Science 297:599–602

36 67. Cobb JA, Bjergbaek L, Shimada K, Frei C, Gasser SM (2003) DNA polymerase stabilization at stalled replication forks requires Mec1 and the RecQ helicase Sgs1. EMBO J 22:4325–4336 68. Dimitrova DS, Gilbert DM (2000) Temporally coordinated assembly and disassembly of replication factories in the absence of DNA synthesis. Nat Cell Biol 2:686–694 69. Cortez D, Glick G, Elledge SJ (2004) Minichromosome maintenance proteins are direct targets of the ATM and ATR checkpoint kinases. Proc Natl Acad Sci USA 101:10078–10083 70. Yoo HY, Shevchenko A, Dunphy WG (2004) Mcm2 is a direct substrate of ATM and ATR during DNA damage and DNA replication checkpoint responses. J Biol Chem 279:53353–53364 71. Cobb JA, Schleker T, Rojas V, Bjergbaek L, Tercero JA, Gasser SM (2005) Replisome instability, fork collapse, and gross chromosomal rearrangements arise synergistically from Mec1 kinase and RecQ helicase mutations. Genes Dev 19:3055–3069 72. Pichierri P, Rosselli F (2004) The DNA crosslinkinduced S-phase checkpoint depends on ATR-CHK1 and ATR-NBS1-FANCD2 pathways. EMBO J 23: 1178–1187

P.J. Hurley and F. Bunz 73. Andreassen PR, D’Andrea AD, Taniguchi T (2004) ATR couples FANCD2 monoubiquitination to the DNA-damage response. Genes Dev 18:1958–1963 74. Chen J (2000) Ataxia telangiectasia-related protein is involved in the phosphorylation of BRCA1 following deoxyribonucleic acid damage. Cancer Res 60:5037–5039 75. Tibbetts RS, Cortez D, Brumbaugh KM et al (2000) Functional interactions between BRCA1 and the checkpoint kinase ATR during genotoxic stress. Genes Dev 14:2989–3002 76. Zhang YW, Otterness DM, Chiang GG et al (2005) Genotoxic stress targets human Chk1 for degradation by the ubiquitin-proteasome pathway. Mol Cell 19:607–618 77. Boddy MN, Furnari B, Mondesert O, Russell P (1998) Replication checkpoint enforced by kinases Cds1 and Chk1. Science 280:909–912 78. Zachos G, Rainey MD, Gillespie DA (2005) Chk1dependent S–M checkpoint delay in vertebrate cells is linked to maintenance of viable replication structures. Mol Cell Biol 25:563–574 79. Brown EJ, Baltimore D (2003) Essential and dispensable roles of ATR in cell cycle arrest and genome maintenance. Genes Dev 17:615–628 Paula J. Hurley and Fred Bunz

Chapter 3

Mechanisms of G2 Phase Arrest in DNA Damage-Induced Checkpoint Response Jian Kuang and Ruoning Wang

Abstract DNA damage activates cell cycle checkpoints and inhibits cell cycle progression. In G2 phase cells, the checkpoint response results in the inhibition of Cdc2/cyclin B activity, which, thereby, prevents G2/M transition, and, as a result, cells accumulate at the G2/M boundary. Both p53-depen-dent and -independent mechanisms are involved in the inhibition of Cdc2 kinase activity during the DNA damage-induced G2 phase checkpoint response. The p53-independent mechanism causes an acute but transient inhibition of the Cdc2/cyclin B activity through posttranslational modifications of the Cdc2activating phosphatase Cdc25, whereas the p53dependent mechanism causes a delayed but sustained inhibition of the Cdc2/cyclin B activity through both transactivation of p21, GADD45 and 14-3-3 and tran-srepression of Cdc2 and cyclin B. Because the p53-dependent mechanism is often defective in tumor cells, abrogation of the p53-independent mechanism to preferentially negate the G2 checkpoint response and induce programmed cell death in tumor cells has become an attractive adjuvant strategy in cancer therapy. Keywords G2 checkpoint • Cdc2 • Cdc25 • PP2A • 14-3-3 • 14-3-3s • p53 • p21 • GADD45

3.1 Introduction DNA-damaging agents in the form of radiation and chemotherapeutic drugs are commonly used in the treatment of human cancers [1, 2]. The DNA damage induced by these anti-tumor agents results

in the unwinding and bending of the DNA helical structure, and specialized recognition proteins detect these specific distortions and propagate signals to initiate the cell cycle checkpoint response [3, 4]. The checkpoint response stops the cell cycle progression at G1, S and/or G2 phases. This results from activation of one or multiple cell cycle checkpoints, which are signal transduction pathways that generate inhibitory signals for key cell cycle regulators, most notably the cyclindependent kinase (Cdk) complexes (Fig. 3.1). In particular, the early G1-phase checkpoint inhibits Cdk4/cyclin D activity and prevents G1 phase progression; the G1/S checkpoint attenuates Cdk2/ cyclin E activity and inhibits G1/S transition; the intra S phase checkpoint inhibits Cdk2/cyclin A activity, which is normally required to support S phase progression; and the G2 phase checkpoint inhibits the activity of Cdk1/cyclin B (also known as Cdc2/cyclin B) and arrests cells at the G2/M boundary [5–7]. The inhibition of Cdk activity by different cell cycle checkpoints can be achieved through a variety of mechanisms, inclu-ding downregulation of cyclins [8–13], prevention of the complex formation between a Cdk and its cognate cyclin partner [14], induction of inhibitory phosphorylations in Cdks [13–19], upregulation of the Cdk inhibitor p21 [20–24] and/or retention of the Cdk complex in the cytoplasm [25, 26]. The checkpoint response permits cells to repair damaged DNA, and prevents DNA synthesis or cell division in the presence of damaged chromosomes, and, thus, is one of the important factors that affect the outcome of chemotherapy and radiotherapy


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J. Kuang and R. Wang

DNA damage

Cdk4/ Cyclin D

G1

Cdk2/ Cyclin E

Cdk2/ Cyclin A

DNA synthesis

Cdk1/ Cyclin B

G2

Mitosis

Figure 3.1. Activation of multiple cell cycle checkpoints by DNA damage. Each checkpoint is associated with the inhibition of a specific Cdk complex so that cell cycle progression can be halted in any phase.

[5, 27–33]. This chapter discusses the current knowledge on the major mechanisms that inhibit Cdc2/cyclin B activity in the DNA damage-induced G2 phase checkpoint response, and provides the molecular basis for the weakened G2 phase checkpoint response in tumor cells.

3.2 Checkpoint Inhibition of Cdc2/Cyclin B Activity by Multiple Mechanisms To understand the major mechanisms that inhibit Cdc2/cyclin B activity in the DNA damage-induced G2 phase checkpoint response, it is first important to understand the basic regulation of Cdc2/cyclin B activity in a normal cell cycle. Intensive studies from 1989 to 1995 using both genetic and biochemical approaches have established a generally accepted model on the critical events that regulate Cdc2 kinase activity in a normal cell cycle, which are illustrated in Fig. 3.2. In this model, Cdc2 kinase activity is regulated by interplays between the degradation status of the activating subunit cyclin B and the phosphorylation status of the catalytic subunit Cdc2. Cyclin B is absent from G1 to early S phase due to active degradation of the protein by APC (Anaphase promoting complex), but begins to accumulate from mid S phase through G2 phase as a result of the shutting down of cyclin B degradation machinery by G1/S

phase Cdk activity [34–36]. On the contrary, the level of Cdc2 protein is quite stable throughout the cell cycle and always in great excess of cyclin B. Thus, most, if not all, of cyclin B is in a complex with Cdc2, whereas the majority of Cdc2 is in a free form, which has little catalytic activity. Importantly, cyclin B binding to Cdc2 in S and G2 phase cells not only provides an essential activating subunit to Cdc2, but also promotes Cdc2 phosphorylation both at the activating site (Thr161 in Xenopus Cdc2 and T160 in human Cdc2) by CAK (Cdk activating kinase) and at the inhibitory sites Thr14 and Tyr15 by Wee1 family protein kinases [37–40]. Because the inhibitory phosphorylation of Cdc2 plays a dominant role, the Cdc2/cyclin B complex is kept inactive from S phase through G2 phase. Only at the G2/M transition point, the preexisting dual-specific protein phosphatase Cdc25 becomes activated, and Wee1 family kinases become inactivated, leading to the robust dephosphorylation of Cdc2 at the inhibitory sites Thr14 and Tyr15. Since now only the activating phosphorylation at Thr161 remains, the suppressed state of the Cdc2/cyclin B complex is relieved, and the activated Cdc2/cyclin B complex is ready to drive the G2/M transition provided that the complex translocates from the cytoplasm into the nucleus as a result of cyclin B phosphorylation [25, 26]. Toward the end of mitosis, cyclin B is degraded by the activated APC, and the activating phosphorylation is removed due to the loss of cyclin B protection from a constitutive phosphatase activity [41–44]. Then, a complete loss of Cdc2/cyclin B activity triggers the exit of mitosis, and Cdc2 goes back to its original G1 state that is not significantly phosphorylated at any of the three sites. Based on these well established principles, the inhibition of Cdc2/cyclin B activity during the DNA damage-induced G2 phase checkpoint response can in principle be achieved through inhibition of cyclin B synthesis, inhibition of cyclin B binding to Cdc2, inhibition of the CAK-catalyzed phosphorylation of the activating site in Cdc2, inhibition of Cdc25 activation and inhibition of the nuclear translocation of the Cdc2/cyclin B complex. In fact, all of these mechanisms have been reported to be involved in the inhibition of -Cdc2/cyclin B activity during the DNA damage-induced G2 phase checkpoint response at least in some cell types.

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3. Mechanisms of G2 Phase Arrest in DNA Damage-Induced Checkpoint Response Stabilization Cyclin B

T14

Cd

c2 Cyclin B

Cdc2

(G1)

/Y1

Wee1 Myt1

Cdc

2

Cyclin B

T16

CAK

1

(S & G2)

(G2) G2 / M transition

Cdc25 activation

KAP Cdc2

T161 Dephosphorylation

APC Cdc2

Cyclin B Degradation

P

Cyclin B

Cdc2 P T161

T161

(G1)

5

(anaphase)

(M)

Figure 3.2. Regulation of Cdc2 kinase activity during the cell cycle. The level of Cdc2/cyclin B complex is determined by cyclin B degradation, and the activity of Cdc2/cyclin B complex is determined by the phosphorylation status of Cdc2.

3.3 Two Major Mechanisms are Critically Involved in Cdc25 Activation During G2/M Transition Inhibition of Cdc25 activation is the most prominent and universal mechanism in the DNA damage-induced G2 checkpoint response. Cdc25 exists as a single protein in yeasts, two very similar proteins in frogs but quite different A, B, C isoforms in mammalian cells. As shown by sequence alignment of the single Cdc25 in Xenopus oocyte (xCdc25) and the A, B, C isoforms of human Cdc25 (hCdc25A, B, C). Cdc25 proteins contain a highly conserved C-terminal catalytic domain of ~ 170 residues and an N-terminal regulatory domain of 300–400 residues, that are poorly conserved (Fig. 3.3). Previous studies on the regulation of Cdc25 in Xenopus oocytes (xCdc25) and its closest relative of xCdc25 in human (hCdc25C) in cultured cells have identified two major mechanisms that are

critically involved in their activations during the G2/M transition [25, 30, 45–47]. One of the mechanisms is the extensive phosphorylation of the N-terminal regulatory domain of Cdc25, and the other is the removal of the 14-3-3 protein from a conserved region within the N-terminal regulatory domain of Cdc25 (Fig. 3.4). The extensive phosphorylation of the N-terminal regulatory domain of Cdc25 directly increases the phosphatase activity of Cdc25 [48–50]. On the other hand, why the 14-3-3 removal is important for Cdc25 activation is not fully understood. In interphase cells, 14-3-3 stoichiometrically binds to preexisting Cdc25 that is phosphorylated at a conserved serine residue in the regulatory domain of Cdc25, i.e., S287 in xCdc25 and S216 in hCdc25C [51–54]. The phosphorylation can be catalyzed by multiple protein kinases, including TAK-1, protein kinase A, calmodulindependent kinase II and potentially other functionally redundant protein kinases [51, 52, 55, 56]. While compromising the 14-3-3 interaction with

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J. Kuang and R. Wang Regulatory-domain

Catalytic-domain Human Cdc25A Human Cdc25B

15% identity

50% identity

Human Cdc25C 70% identity

30% identity Xenopus Cdc25C 375

1

550

Figure 3.3. Regulatory and catalytic domains of Cdc25. The catalytic domain of Cdc25 is highly conserved (50–70% identity), whereas the homology of the regulatory domain between the three human and one Xenopus forms is less conserved (15–30% identity).

14-3-3

Inactive Cdc25C (in interphase)

p-S287

Regulatory domain

Catalytic domain

14-3-3

14-3-3

Inactivating dephosphorylation S287 phosphorylation 14-3-3 binding

Active Cdc25C (in mitosis)

14-3-3 removal Activating phosphorylation S287 dephosphorylation

p-T48/T67/T138/S205/S285/T308 and more

Regulatory domain

Catalytic domain

Figure 3.4. Mechanisms involved in Cdc25C activation. Hyperphosphorylation of the regulatory domain and removal of 14-3-3 are the two mechanisms responsible for activating Cdc25 during the G2/M transition.

Cdc25 accelerates Cdc25 activation and the G2/M transition, enhancing this interaction has opposite effects [51, 53]. These findings provide the basis for the current understanding that 14-3-3 binding to Cdc25 is critical for keeping Cdc25 inactive in interphase cells and that 14-3-3 removal from Cdc25 is critical for Cdc25 activation during the G2/M transition. However, 14-3-3 has weak or no inhibitory effects on the catalytic activity of Cdc25

[51, 53], and thus its removal from Cdc25 should have mild or no stimulatory effects on Cdc25 activity. Although the removal of 14-3-3 has been demonstrated to unmask the nuclear localization signal in Cdc25 and thus promote translocation of Cdc25 from the cytoplasm into the nucleus [25, 57], this mechanism does not explain why dissociation of 14-3-3 from Cdc25 is important for Cdc25 activation in the cytoplasm [56, 58]. Thus,

3. Mechanisms of G2 Phase Arrest in DNA Damage-Induced Checkpoint Response

it is likely that 14-3-3 removal both promotes the nuclear translocation of Cdc25 and plays yet-to-be identified roles in Cdc25 activation. One potential role that 14-3-3 removal may play in Cdc25 activation is to unmask yet-to-be identified activating phosphorylation sites in the nearby region of the regulatory domain of Cdc25 through of relieving a physical hindrance. If such is the case, the activating phosphorylation of Cdc25 and the removal of 14-3-3 from Cdc25 may form a positive feedback loop in the process of robust mitotic activation of Cdc25, and the inhibition of Cdc25 activation during the DNA damage-induced G2 phase checkpoint response may involve the inhibition of either or both of these processes.

3.3.1 Inhibition of 14-3-3 Removal During DNA Damage-Induced G2 Phase Checkpoint Response Studies in the last decade using multiple experimental systems have provided abundant and solid evidence that inhibition of 14-3-3 removal is critically involved in the inhibition of Cdc25 activation during the DNA damage-induced G2 phase checkpoint response [51, 54, 59]. Also established

OLD MODEL DNA damage Activate ATM / ATR

41

is the critical role of the DNA damage-induced activation of the ATM/ATR-Chk1/Chk2 pathways and/or the p38-MAPKAPK pathway in the inhibition of the 14-3-3 removal from Cdc25 [60, 61]. Then, because all of these checkpoint kinases directly phosphorylate Cdc25 at the 14-3-3 docking site, these discoveries led to a commonly accepted thinking that these checkpoint kinases inhibit the 14-3-3 removal and prevent Cdc25 activation during the DNA damage-induced G2 phase checkpoint response via promoting Cdc25 phosphorylation at the 14-3-3 docking site [60, 61] (Fig. 3.5, “Old Model”). However, this simple and straightforward model has not considered the facts that the preexisting Cdc25 in interphase cells is already phosphorylated at the 14-3-3 docking site [55, 62] and that the phosphorylation site is protected by the bound 14-3-3 from the opposing phosphatase activity [63]. With these starting points, it seems unlikely that the DNA damageactivated checkpoint kinases play key roles in the phosphorylation status of the preexisting Cdc25 at the 14-3-3 docking site. In fact, there have been reports that the DNA damage-induced G2 phase checkpoint response does not significantly change the phosphorylation status of Cdc25 at the 14-3-3 docking site.

NEW MODEL DNA damage Activate ATM / ATR Activate Chk1/2

Activate Chk1 / 2 Phosphorylate Cdc25 at S287 Bind 14-3-3 to Cdc25 Inhibit Cdc25 activation or function

Maintain the activity of Cdc25 bound PP2A Prevent Cdc25-T138 phosphorylation

Inhibit kinase activation? Inhibit phosphatase inactivation? Prevent phosphorylation of intermediate filament proteins

Inhibit 14-3-3 removal Inhibit Cdc25 activation or function

Inhibit Cdc2 activation Inhibit Cdc2 activation G2 / M arrest

G2 / M arrest

Figure 3.5. Mechanisms inhibiting 14-3-3 dissociation from Cdc25C in DNA damage-induced G2 checkpoint response. The figure provides a comparison between the “Old Model” and the “New Model” for inhibiting Cdc25C activity and inducing G2/M arrest.

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Then, how may activation of the checkpoint kinases during the DNA damage-induced G2 phase checkpoint response inhibit the 14-3-3 removal and Cdc25 activation? To understand this issue, it is first important to understand the regulation of 14-3-3 removal in a normal cell cycle. Following a series of pilot studies [62–64], Kornbluth and colleagues have generated a conceptual framework for the regulation of the removal of 14-3-3 from xCdc25 in Xenopus egg extracts [47, 65]. In their recently proposed model [62] (Fig. 3.5, “New Model”), two events are rate-limiting for the 14-3-3 removal during the G2/M transition. One of them is phosphorylation of Cdc25 at T138, which somehow loosens the interaction between 14-3-3 and xCdc25. The other event is the phosphorylation of intermediate filament proteins such as keratin by a yet-to-be identified mitotic kinase. The phosphorylated intermediate filament protein(s) sequesters already loosened 14-3-3 away from xCdc25. Although the regulation of the phosphorylation of intermediate proteins is yet to be understood, the T138 phosphorylation is regulated by a Cdc25-associated phosphotase activity. In interphase extracts, the constitutively active Cdk2 is able to phosphorylate Cdc25 at T138. However, a specific PP2A holoenzyme (PP2A/B56d) is stoichiometrically associated with xCdc25, taking off the T138 phosphorylation in Cdc25. During the G2/M transition, this Cdc25 specific phosphatase gets inactivated as a result of the loss of an activating phosphorylation on B56d, which can be catalyzed by Chk1. The inhibition of the PP2A/ B56d activity then results in a net increase in the phosphorylation of T138 in Cdc25 [62]. While it remains to be determined whether the model described above is of general significance and accuracy, the model does provide several novel insights on the potential mechanisms that regulate the 14-3-3 removal during the DNAdamage-induced checkpoint response. First, since it is the mitotic phosphorylation of Cdc25 and intermediate filament proteins rather than dephosphorylation of Cdc25 at the 14-3-3 docking site that are rate-limiting for the 14-3-3 removal, inhibition of either or both of these events should be the key to the inhibition of the 14-3-3 removal during the DNA damage-induced G2 phase checkpoint response. Second, because a non-mitotic kinase may catalyze a mitosis-specific phosphory-


lation of Cdc25 due to the cell cycle regulation of a Cdc25-bound protein phosphatase, and a similar principle may apply to the mitotic-specific phosphorylation of intermediate proteins, protein phosphatases can be crucial targets of checkpoint kinases during the DNA damage-induced G2 checkpoint response.

3.3.2 Inhibition of Activating Phosphorylation of Cdc25 in DNA Damage-Induced G2 Phase Checkpoint Response Both xCdc25 and hCdc25C undergo an activationassociated dramatic gel mobility shift during the G2/M transition as a result of the phosphorylation of numerous sites in the N-terminal regulatory domain [50, 66–68] (Fig. 3.4). The activating phosphorylation sites include T138, implying that there is a cross talk between the two major mechanisms of Cdc25 regulation. Since the activationassociated dramatic gel mobility shift of Cdc25 is invariantly inhibited during the DNA damageinduced G2 phase checkpoint response [51, 54, 59], the checkpoint response must include the activation of signaling pathways that potently inhibit the activating phosphorylation of Cdc25. However, the molecular mechanisms that inhibit the activating phosphorylation of Cdc25 during the DNA damage-induced G2-phase checkpoint response are not well understood. It is generally accepted that activating phosphorylation of Cdc25 during the G2/M transition consists of the initiation and amplification steps [46, 69]. The initiation step involves phosphorylation of Cdc25 by kinases that can be activated prior to Cdc2 activation and serves to generate a threshold level of Cdc25 activity. In the amplification step, the kinases that can be directly or indirectly activated by Cdc2 further phosphorylate Cdc25 and generate a peak level of Cdc25 activity that is sufficient to induce the G2/M transition. Based on these principles, DNA damage-induced checkpoint inhibition of the activating phosphoryaltion of Cdc25 may target either or both of the initiation and amplification steps of the process (Fig. 3.6). To understand how DNA damage-induced checkpoint response may inhibit the initiation of Cdc25 activation in any experimental system, the


DNA damage

Initiation

Amplification ERK,Plk1

ERK Plk1

Cdc25C

Cdc2/cyclin B

Figure 3.6. Inhibition of activating phosphorylation of Cdc25 in DNA damage-induced G2 checkpoint response. Activating phosphorylation of Cdc25 consists of the initiation and amplification steps. Therefore, inhibition of these steps will be necessary to prevent activation of Cdc25.

first important task is to identify the kinase(s) that initiates Cdc25 activation during the G2/M transition. Polo-like kinase 1 has been suggested to initiate Cdc25 activation in certain biological systems [70], and thus its activation may be targeted by DNA damage-induced G2 phase checkpoint response [71, 72]. Since the nonmitotic Cdk2 complex is able to phosphorylate Cdc25 in an M-phase specific manner due to the regulation of Cdc25associated PP2A activity as described above, it is possible that the Chk-catalyzed phosphorylation of the Cdc25-associated PP2A is one of the mechanisms that inhibit the initiating activation of Cdc25 by Cdk2 during DNA damage-induced G2 phase checkpoint response. Moreover, the recent finding that ERK-MAP kinases can be both activated prior to and independent of Cdc2 activation and catalyze some of the activating phosphorylations in Cdc25 during the G2/M transition [50] has generated the insight that growth promoting signaling kinases may also play important roles in the initiating activation of Cdc25. Because these signaling kinases are already activated in interphase cells in somatic cell cycles, it is conceivable that the Cdc25-bound PP2A that inhibits the Cdk2-catalyzed phosphorylation of T138 in Cdc25 also plays a role in the

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inhibition of activating phosphorylation of Cdc25 by the signaling kinases in interphase cells. If such is the case, the DNA damage-induced checkpoint response may inhibit the initiating activation of Cdc25 and the 14-3-3 removal through a common mechanism. The activated Cdc25 not only activates Cdc2/ cyclin B, but also is phosphorylated and activated by the activated Cdc2/cyclin B [66, 67]. These findings led to the current thinking that Cdc2 kinase forms a direct positive feedback loop with Cdc25, responsible for their switch-like and coordinated activations during the G2/M transition. In addition to this direct positive feedback loop, the activated Cdc2/cyclin B both activates polo-like kinases and catalyzes the priming phosphorylation in Cdc25 that promotes the binding and phosphorylaiton by polo-like kinases [73]. In Xenopus oocytes, the activated Cdc2/cyclin B also enhances the ERK2 activation through regulating the upstream regulators [74–76]. These indirect positive feedback loops should also contribute to the robust activation of Cdc25 activation during the G2/M transition and be targets of the DNA damage-induced G2 checkpoint response. There have not been reports on the presence of specific mechanisms that inhibit the amplification step of the Cdc25 activation. However, it is conceivable that the Cdc25-bound PP2A, whose activity is maintained during the DNA damage-induced checkpoint response, inhibits the amplification step of the Cdc25 activation.

3.4 p53-Dependent Mechanisms in the DNA Damage-Induced G2 Checkpoint Response The DNA damage activated ATM/Chk1 and ATR/ Chk2 pathways not only inhibit the Cdc25 activation in the absence of de novo RNA and protein syntheses, but also post-translationally stabilize and activate the transcription factor p53. The induced p53 then transactivates or transrepresses genes that determine the level, activity or the localization of the Cdc2/cyclin B complex [77]. Among the identified mechanisms illustrated in Fig. 3.7, high levels of p53 inhibit the transcription of both cyclin B and Cdc2, partially through directly repressing

44


window of opportunity to preferentially negate the G2 checkpoint and induce programmed cell death in tumor cells.

DNA damage ATM Chk2

ATR Chk1

3.4.1 Role of p21 Transcriptional activation

p21

14-3-3

Reduced functions of Cdc2/cyclin B complex

p53

GADD45

Transcriptional repression

Cdc2

Cyclin B

Reduced levels of Cdc2 / cyclin B complex

Complete G2 / M arrest

Figure 3.7. p53-dependent mechanisms in DNA damage-induced G2 checkpoint response. Following activation, p53 transactivates and transrepresses target genes to induce the G2/M arrest.

their promoters, and this results in the reduction or elimination of the Cdc2/cyclin B complex [78–82]. The induced p53 also activates the transcription of p21, 14-3-3 and GADD45, which inhibit the Cdc2/ cyclin B activity through varied mechanisms [77, 83–85]. All of these p53 dependent mechanisms require de novo RNA and protein synthesis, and thus they cause a delayed inhibition of Cdc2/cyclin B activity as compared to the p53-independent mechanisms that inhibit Cdc25 activation discussed above. On the other hand, because p53-dependent mechanisms are more stable than the p53-independent mechanisms that inhibit Cdc25 activation, p53-dependent mechanisms often determine the length and stability of the DNA damage-induced G2 checkpoint response [77]. Importantly, p53 is often deleted or mutated in tumor cells [83–85]. This makes the DNA damage-induced G2 phase checkpoint arrest less sustained in tumor cells than in their normal counterparts and provides a

p21 is a Cdk interacting protein that is able to bind various Cdk/cyclin complexes and inhibit their activities [24, 86]. The p21 promoter has two p53 response elements, which have avid affinity for activated p53 [87–90]. Thus, induction of p53 in the DNA damage-induced checkpoint response leads to a robust induction of p21 [91]. The involvement of p21 in the G2 phase checkpoint arrest has been demonstrated in a number of reports, but the most definitive evidence has come from studies in p21-knockout HCT-116 colorectal carcinoma cells [21]. These cells, like parental p21-proficient HCT-116 cells, arrested at G2/M after exposure to ionizing radiation. However, unlike parental cells, p21-deficient cells failed to maintain the G2 arrest, and this coincided with restoration of Cdc2/cyclin B activity. These findings established that p21 is required to sustain the inhibition of the Cdc2/cyclin B complex and prevent cells from prematurely entering into mitosis. While the involvement of p21 in the G2 checkpoint arrest is established, the mechanism by which the induced p21 maintains the inhibition of the Cdc2/ cyclin B activity is not defined. Since induced p21 is able to bind the Cdc2/cyclin B complex, it may directly inhibit the Cdc2/cyclin B activity if expressed at stoichiometrically high levels. The problem with this idea is that p21 is much less efficient in binding and inhibiting the Cdc2/cyclin B complex than binding and inhibiting various G1 and S phase Cdk complexes, including Cdk4/cyclin D, Cdk2/cyclin E, and Cdk2/cyclin A, in vitro [86]. Alternatively, since the Cdk2 activity may be critically involved in the initial activation of Cdc25 [63] and has been shown to be required for the activation of the Cdc2/cyclin B activity [41, 42], the induced p21 may maintain the inhibition of the Cdc2/cyclin B activity indirectly through the inhibition of the Cdk2 activity. In addition to these potential mechanisms, the induced p21 may inhibit CAK-catalyzed activating phosphorylation of Cdc2 [77]. Another alternative mechanism is through reduction or elimination of the Cdc2/cyclin


B complex. Although this has been ascribed to high induction levels of p53 inhibiting transcription of both Cdc2 and cyclin B, there is solid evidence that p53-dependent repression of these kinase components requires an interplay between p53 and the induced p21 [92]. This is particularly reinforced by the demonstration that ionizing radiation represses Cdc2 and cyclin B1 expression in wild-type MEFs, but not in p21- or p53-null MEFs [93, 94]. One plausible explanation for these observations is that the inhibition of the G1 phase Cdk activity by p21 leads to the inhibition of the inactivating RB phosphorylation and consequent E2F derepression, which then prevents the transcription of both Cdc2 and cyclin B [95–104]. Although the role of p21 in maintaining the G2 checkpoint arrest is irrefutable, the p21 induction alone may not be sufficient to initiate the inhibition of the Cdc2/cyclin B activity and G2 phase checkpoint arrest. A clear example of this point is the cell cycle perturbations induced by the cisplatin analog DAP [105]. In contrast to the parental compound cisplatin, which activates both the ATM-Chk1 pathway [106] and the p53-p21 pathway [107] and inhibits both the G1/S and G2/M transitions [108–110], DAP treatment led to a robust p21 induction in a p53-dependent manner in ovarian A2780 tumor cells without activating the ATM/ Chk1 and ATR/Chk2 pathways that affect Cdc25 activity [91]. Although the induced p21 potently inhibited both Cdk4 and Cdk2 activities and cell cycle progression at the G1/S boundary, it inhibited neither the Cdc2/cyclin B activation nor the G2/M transition. These findings suggest that high levels of p21 do not cause the G2 phase arrest without the contribution of other mechanisms.

3.4.2 Role of 14-3-3s 14-3-3s is an interacting protein that binds the Cdc2/Cyclin B complex without inhibiting its catalytic activity [111]. The 14-3-3s promoter has a p53 response element, and thus its expression is induced in the DNA damage-induced checkpoint response [112]. Because 14-3-3s deficient cells have defects in maintaining the DNA damageinduced G2 phase arrest and undergo premature mitotic entry in colon cancer HCT116 cells that had been exposed to either adriamycin or ioniz-

45

ing radiation [113], 14-3-3s is implicated in the maintenance of the G2 phase checkpoint arrest. In understanding the mechanism of 14-3-3s functions, the induced 14-3-3s was found to sequester the Cdc2/cyclin B complex in the cytoplasm. Since Cdc2/cyclin B must be localized in the nucleus for its mitosis inducing function, the effect of 14-3-3s prevents the Cdc2/cyclin B complex from inducing mitosis [111]. Interestingly, 14-3-3s expression is not only reduced in p53-deficient cells, but also compromised by other mechanisms in the presence of p53 functions. For example, silencing of 14-3-3s due to promoter hypermethylation occurs in several tumor types, including breast and prostate. This promotes early entry into mitosis, increase genomic instability, and induce carcinogenesis [114]. A second example is with BRCA1, which has been reported to cooperate with p53 to induce the expression of 14-3-3s in mouse embryonic stem cells after treatment with ionizing radiation [115]. However, BRCA1 is mutated in breast and ovarian tumors [115]. The presence of multiple mechanisms that inactivate the 14-3-3s-dependent checkpoint function in tumor cells indicates the important roles of 14-3-3s in the maintenance of genetic stability.

3.4.3 Role of GADD45 GADD45 is another p53 target that gets upregulated in the DNA damage-induced checkpoint response [116]. GADD45 deficient lung carcinoma cells do not stably arrest at the G2/M boundary following DNA damage, whereas microinjection of GADD45 into primary human fibroblasts or ectopic expression of GADD45 in cycling retinoblastoma proteinnegative cells causes cell cycle arrest at the G2/M boundary [117, 118]. Moreover, GADD45 nullmice are more susceptible to developing tumors following exposure to ionizing radiation as a result of genomic instability [119]. These findings implicate GADD45 in the maintenance of the G2 phase checkpoint arrest. In understanding the mechanism by which GADD45 specifically targets the G2 phase checkpoint, the induced GADD45 was found to bind the monomeric Cdc2 and prevent the assembly of the Cdc2/cyclin B complex [77]. In contrast, GADD45 did not bind Cdk2 or inhibit Cdk2/cyclin E activity [14], explaining its preferential effects

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on the G2 phase checkpoint. In hematopoietic cells, GADD45 requires the presence of p53 to induce G2 arrest [120], suggesting that the induced GADD45 alone may not be sufficient to quantitatively prevent the formation of the Cdc2/cyclin B complex. It is possible that p53-induced GADD45 cooperates with p53-induced 14-3-3s and p21 to quantitatively inhibit the Cdc2/cyclin B activity and induce a stable G2 phase arrest.

3.5 Weakened G2 Phase Checkpoint in Tumor Cells and Its Exploitation in Cancer Therapy Many of the commonly used chemotherapeutic agents achieve their effects by damaging DNA damage. In most cases, these DNA damaging agents do not kill cancer cells directly. Rather, they initially activate the cell cycle checkpoints and stops cell cycle progression. This not only permits cells to repair the DNA, but also prevents DNA synthesis or cell division in the presence of damaged chromosomes. These effects promote cell survival and are against the ultimate goal of chemotherapy, killing cancer cells. Only when the level of DNA damage induced is high enough to directly trigger programmed cell death (apoptosis) or impossible to be eliminated by DNA repair within a reasonable period of time do these DNA damaging agents induce cell death. Based on these principles, agents that are able to preferentially abrogate cell cycle checkpoint response in cancer cells would increase the therapeutic effect of these DNA damaging agents. Although DNA damage activates multiple cell cycle checkpoints and stops cell progression at G1, S and/or G2 phases, the G2 phase checkpoint response is often the most prominent checkpoint response in cancer cells for the following three reasons. First, cancer cells often have p53 deletion or mutation and thus are defective in p53-dependent G1 phase checkpoint response [83–85]. Second, in p53 proficient cells, the DNA damage-induced G1 phase checkpoint arrest, which is primarily induced by the delayed p53-dependent induction of p21, is much slower than the G2 checkpoint arrest [91]. As a result, DNA damaged G1 phase cells


progress into S and G2 phases until the delayed checkpoint mechanism kicks in. Third, although the S phase checkpoint arrest in S phase cells is prompt, it often does not last whether the damaged DNA is all repaired or not. After a transient arrest, these cells pile up at the next G2 phase checkpoint for completion of repair. The prominence of G2 phase checkpoint response in tumor cells provides the basis for developing a strategy that preferentially abrogate the G2 phase checkpoint response in cancer cells for enhancing the therapeutic effect of these DNA damaging agents. How can this be achieved based on the current understanding of the mechanisms of the G2 phase arrest in DNA damageinduced checkpoint response? Since p53 is often deleted or mutated in tumor cells, abrogation of the p53-independent mechanisms that inhibit the Cdc25 activation would preferentially abrogate the G2 phase checkpoint response in cancer cells (Fig. 3.8) but not in normal cells. In fact, this is one of the adjuvant strategies that are currently being explored in cancer therapy, and multiple G2 checkpoint abrogators have already been developed for the use of this strategy [121–124]. Apparently, an eventual successful use of this strategy will need a more complete understanding of the p53-independent mechanisms that inhibit the Cdc25 activation during the DNA damageinduced checkpoint response and the availability of drugs that have satisfying phamacodynamics and pharmacokinetics properties.

3.6 Conclusion The DNA damage-induced G2 checkpoint response involves multiple p53-dependent and -independent mechanisms, which cooperate to generate a rapid and durable arrest of the cell cycle at the G2/M until DNA damage is fully repaired. A variety of molecules are involved, and available data have provided important insights on their functions. However, many key questions remain to be answered. A more complete understanding of the mechanisms that support the DNA damage induced G2 phase arrest provides a more solid basis for rational design and effective use of the G2 checkpoint abrogators in cancer therapy.

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p53 deficient cancer cells

ATM/ATK CHK1/2 PP2A

Abrogation of p53-independent checkpoint response ATM/ATR CHK1/2 PP2A ??

Inhibitor

Intact p53-independent Checkpoint response

14-3-3

Inactive Cdc25C

Active Cdc25C

14-3-3

G2

Mitosis G2 arrest & DNA repair

G2

Mitosis

Premature M-phase entry & Mitotic catastrophe

Figure 3.8. Strategies that may specifically abrogate the G2 checkpoint response in tumor cells. Approaches that inhibit known or undetermined targets and activate Cdc25 in the presence of DNA damage will cause premature M phase entry and cell death due to mitotic catastrophe.

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116. Kastan MB, Zhan Q, el-Deiry WS et al (1992) A mammalian cell cycle checkpoint pathway utilizing p53 and GADD45 is defective in ataxia-telangiectasia. Cell 71:587–597 117. Vairapandi M, Balliet AG, Hoffman B, Liebermann DA (2002) GADD45b and GADD45g are cdc2/ cyclinB1 kinase inhibitors with a role in S and G2/M cell cycle checkpoints induced by genotoxic stress. J Cell Physiol 192:327–338 118. Fan W, Richter G, Cereseto A, Beadling C, Smith KA (1999) Cytokine response gene 6 induces p21 and regulates both cell growth and arrest. Oncogene 18:6573–6582 119. Hollander MC, Sheikh MS, Bulavin DV et al (1999) Genomic instability in Gadd45a-deficient mice. Nat Genet 23:176–184 120. Liebermann DA, Hoffman B (2007) Gadd45 in the response of hematopoietic cells to genotoxic stress. Blood Cells Mol Dis 39:329–335 121. Calonge TM, O’Connell MJ (2008) Turning off the G2 DNA damage checkpoint. DNA Repair (Amst) 7:136–140 122. Bucher N, Britten CD (2008) G2 checkpoint abrogation and checkpoint kinase-1 targeting in the treatment of cancer. Br J Cancer 98:523–528 123. Tse AN, Carvajal R, Schwartz GK (2007) Targeting checkpoint kinase 1 in cancer therapeutics. Clin Cancer Res 13:1955–1960 124. Levesque AA, Eastman A (2007) p53-Based cancer therapies: Is defective p53 the Achilles heel of the tumor? Carcinogenesis 28:13–20

Chapter 4

Centrosomes in Checkpoint Responses Alwin Krämer

Abstract Centrosomes consist of a pair of barrelshaped centrioles, surrounded by a pericentriolar matrix. Their best characterized function is to organize both interphase microtubule arrays and the mitotic spindle, which mediates the strictly bipolar separation of chromosomes during cell division. In addition, centrosomes have come into focus as part of a network that integrates cell cycle arrest and repair signals in response to genotoxic stress. Recent evidence suggests that centrosomes are involved in both, regulation of the G2/M transition in response to DNA damage and induction of cell death via centrosome amplification and mitotic catastrophe as a backup mechanism for the elimination of cells that evade DNA damage checkpoints operating earlier during the cell cycle. While other aspects of the G2/M checkpoint are described elsewhere, this chapter will focus on the emerging role of centrosomes as regulators and effectors of DNA damage at mitotic entry. Keywords Centrosome • DNA damage • Checkpoint • Chk1 • Pericentrin • Microcephalin • cdc25 • Seckel syndrome • Microcephaly • Centrosome amplification • Mitotic catastrophe

4.1 Centrosomes A single mammalian centrosome comprises two centrioles, embedded in a protein meshwork known as the pericentriolar material. Vertebrate centrioles are cylindrical organelles made up of triplet microtubules [1]. Of the two centrioles present in a single

centrosome, only the older or mature one is equipped with appendages at its distal end. Appendages have been implicated in microtubule anchoring, but their precise roles are still elusive. To date, more than hundred centrosomal proteins have been identified [2]. Many of them contain predicted coiled-coil structures, suggesting that they play scaffolding roles. In particular, the pericentriolar material serves as a scaffold for g-tubulin ring complexes that function in microtubule nucleation [3]. Many proteins associate with the pericentriolar matrix in a cell cycle dependent manner, demonstrating cell cycle regulation of centrosome function. Conversely, centrosomes transiently recruit components of signaling cascades and thus function as platforms for enhancing the efficiency of cell cycle regulatory steps, specifically the G2/M transition [4, 5]. As illustrated in Fig. 4.1, cells in G1 phase of the cell cycle contain a single centrosome consisting of one older or mature and one younger or daughter centriole. Before cell division, this whole structure needs to be duplicated once, so that a G2 phase cell harbors two centrosomes, each comprising two closely linked centrioles. Centrosome division can be subdivided into several distinct steps, with centrosome duplication occurring during S phase and centrosome segregation during M phase [6]. Later in M phase the two centrioles of every centrosome are separated from each other in a process termed disengagement. This disengagement, which seems to constitute the licensing event for subsequent centriole duplication, might be mediated by separase activity and ensures that centrioles are duplicated


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Figure 4.1. Centrosome reproduction cycle. Centrioles are represented as barrels. Gray shading represents the mature mother centriole. The immature daughter centriole is displayed in white.

only once per cell cycle [7, 8]. During S phase, centriole duplication is characterized by formation of exactly one procentriole next to each parental centriole. Procentrioles then continue to elongate until in G2 phase the centrosome consists of two pairs of centrioles. At the G2/M transition, the putative linker connecting the two parental centrioles is severed and the two centrosomes are separated by microtubule-dependent motor proteins [9–14]. Upon formation of the bipolar mitotic spindle, the two centrosomes then associate with the two spindle poles and segregate into the incipient daughter cells, thereby completing the centrosome cycle. Both centrosome and DNA replication are simultaneously initiated by the activation of cyclin-dependent kinase 2 (Cdk2), in association with cyclins E and/or A in late G1 phase of the cell cycle [15–17]. However, since a direct action of Cdk2 at the centrosome has not been demonstrated, it remains possible that Cdk activity is required primarily to advance cells to a cell cycle stage that is permissive for centrosome duplication. In contrast, polo-like kinase 4 (Plk4) has unequivocally been identified as a positive regulator of centriole duplication [18, 19]. Of note, overexpression of Plk4 in human cells results in the production of multiple centriole precursors surrounding a single parental centriole and Plk4+/− mice are prone to develop tumors [18, 20].

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It has been long known that ciliated epithelial cells and male gametes of lower plants are able to generate large numbers of centrioles de novo in the absence of a template [21]. This acentriolar pathway for centriole biogenesis was thought to be restricted to a few specialized cell types. Recently, however, the de novo formation of centrioles was shown to be inducible in normal and transformed human cells after resident centrosomes were removed by laser ablation or microsurgery [22–24]. This finding suggests that two distinct centriole assembly pathways coexist with the de novo acentriolar pathway being normally suppressed by templated centriole duplication [25, 26]. Although centrosomes are the major organizers of animal cell division, they are not essential for mitotic spindle assembly. Some animal cells normally organize their spindles without canonical centrosomes, and cultured cells that have had their centrosomes removed can still form bipolar spindles [27, 28]. Also, it has recently been shown that DSas-4 mutant flies, which lack centrosomes altogether because DSas-4 is absolutely required for centrosome biogenesis, develop with near normal timing into morphologically normal adults [29]. In these cases, the mitotic chromosomes seem to initiate the assembly of a bipolar spindle and thereby compensate for the lack of centrosomes. However, when present, centrosomes exert a strong influence on the number of spindle poles formed [30].

4.2 Centrosomal Regulation of the G2/M Transition 4.2.1 Centrosomal Regulation of Unperturbed G2/M Progression 4.2.1.1 Cdc25 During unperturbed cell cycles, entry into mitosis requires Cdk1/cyclin B activity [31]. It has long been known that a fraction of the Cdk1/cyclin B complex localizes to the centrosome [32, 33]. More recently, it became evident that the initial activation of Cdk1/cyclin B in early prophase, including Cdk1 dephosphorylation at Thr14/Tyr15 and cyclin B phosphorylation at Ser126 and Ser133, indeed takes place at the centrosomes and from there spreads to induce both cytoplasmic and nuclear

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mitotic events like spindle formation, nuclear envelope breakdown and chromosome condensation [4]. Dephosphorylation of Cdk1 at Thr14 and Tyr15 depends on the activity of the dual-specificity phosphatase Cdc25 [34]. In human cells, the Cdc25 family includes three members, Cdc25A, Cdc25B and Cdc25C [35, 36]. Although all three Cdc25 forms have been shown to function during mitosis, Cdc25B is thought to act as an initiator of early mitotic events, because its overexpression in interphase cells induces premature mitotic entry [37–39]. Moreover, Cdc25B has been shown to localize to centrosomes from the beginning of S phase until mitosis [5, 40, 41]. Cdc25A and Cdc25C, on the other hand, are thought to further induce Cdk1 activity in the nucleus and sustain a positive feedback loop that rapidly amplifies both Cdc25 and Cdk1 activity to trigger mitotic entry [41, 42]. Most recent data suggest that, in addition to Cdc25B, Cdc25C might reside at centrosomes as well and cooperate with Cdc25B in activating Cdk1/cyclin B at the G2/M transition [43–45].

4.2.1.2 Aurora-A The centrosomal recruitment of Cdc25B seems to be brought about by phosphorylation at Ser353 by Aurora-A [40]. Aurora-A, one of three family members of the mammalian aurora kinase family, is upregulated at the onset of mitosis and localizes to centrosomes during interphase and to both spindle poles and spindle microtubules during early mitosis [46, 47]. Of those substrates of Aurora-A identified to date, many play important roles in spindle assembly, chromosome segregation, and cytokinesis. Consistently, disruption of the Aurora-A gene leads to defects in centrosome and chromosome separation, spindle assembly, and mitotic progression. Remarkably, increasing evidence links Aurora-A to oncogenesis and Aurora-A overexpression due to gene amplification can be found in a wide variety of human tumors. Analogous to Cdk1, Aurora-A kinase activity is essential for mitotic entry [48]. Importantly, centrosomal Aurora-A becomes activated prior to Cdk1/cyclin B and is a prerequisite for initial activation of Cdk1/cyclin B at the centrosome. Since centrosomes of Aurora-A-depleted cells lack significant cyclin B loading, Aurora-A might even be required for the recruitment of Cdk1/ cyclin B to the centrosomes. Centrosomal activation of

Aurora-A itself seems to be mediated by an Aurora-Ainteracting protein called Ajuba [48]. Ajuba, a LIM domain-containing protein, concentrates on centrosomes during the G2/M phase of the cell cycle and interacts with Aurora-A during mitosis. As both proteins form a complex at the centrosome, Ajuba facilitates autophosphorylation and thereby activation of Aurora-A on Thr288.

4.2.1.3 Chk1 While Aurora-A acts as a positive upstream regulator of centrosomal Cdc25B, Chk1 has been shown to serve as its opponent within the centrosomal Cdc25B-Cdk1 pathway during unperturbed cell cycle progression [5, 49]. Chk1 is a serine/threonine kinase that is activated in response to diverse genotoxic insults by the proximal checkpoint kinases ATR and ATM and localizes, in addition to the nuclear compartment, to interphase but not mitotic centrosomes [5, 49–51]. Chk1 plays central roles in the intra-S and G2/M DNA damage checkpoints which slow DNA replication and delay mitotic entry, respectively, in response to DNA damage [50, 52, 53]. Chk1 expression is largely restricted to S and G2 phases, and it is active even during unperturbed cell cycles, although it is further activated in response to DNA damage or stalled replication [54–57]. Accordingly, mammalian cells haploinsufficient for Chk1 exhibit a variety of abnormal cell cycle phenotypes including inappropriate S phase progression, spontaneous DNA damage, and premature entry into mitosis [58]. Consistent with this latter observation, several studies have clearly implicated Chk1 as a key regulator of the initiation, progression, and fidelity of unperturbed mitosis. Remarkably, during interphase Chk1 seems to restrain centrosome-associated Cdk1 activity through local inhibition of Cdc25B, thereby preventing premature activation of Cdk1/cyclin B kinase activity and counterbalancing the effect of Aurora-A [5]. Although the mechanism of this inhibition is not fully understood, Chk1 phosphorylates Cdc25B at multiple sites, including Ser230 and Ser563, in vitro and in vivo during unperturbed cell cycle progression [5, 59, 60]. Ser230 phosphorylated Cdc25B localizes to centrosomes and displays reduced phosphatase activity [60], supporting a model in which, in the absence of DNA damage, Chk1 constitutively phosphorylates

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unstable cells by inducing cell death [64–66]. The biological significance of the DNA damage response is apparent from documented links between DNA damage response malfunction and numerous pathological states including cancer, immune deficiency, and neurological disorders [64–66]. The term “cell cycle checkpoint” refers to the mechanisms by which the cell actively halts progression through the cell cycle until it can ensure that an earlier process, such as DNA replication or mitosis, is complete [67]. Multiple checkpoints exist and are conserved in the cell cycle in higher eukaryotes to ensure that if one fails, others will take care of genomic integrity and cell survival, since maintenance of genomic Figure 4.2. Schematic representation of a simplified integrity is essential to avoid cellular transformamodel for centrosome-localized processes that control tion, neoplasia, or cell death. mitotic entry of mammalian cells through regulation of The DNA damage checkpoint network is comCdk1/cyclin B. posed of DNA damage sensors, signal transducers, and various effector pathways, and its central comCdc25B during interphase and thereby prevents ponents are the phosphoinositide 3-kinase related the premature initiation of mitosis by negatively kinases ATM and ATR, which relay their signals regulating the activity of Cdc25B at the centrosome to the downstream checkpoint kinases Chk1 and (Fig. 4.2). Dissociation of Chk1 from centrosomes Chk2 [50, 65]. ATM is activated primarily by at the end of G2 phase, together with positive DNA double strand breaks induced by ionizing regulatory phosphorylation of Cdc25B at Ser353 radiation, whereas ATR also responds to ultramediated by Aurora-A, then enables Cdc25B to violet (UV) radiation or stalled replication forks. activate the centrosomal pool of Cdk1/cyclin B While ATM predominantly phosphorylates Chk2, and to initiate mitosis [5, 40, 49, 61]. Inhibition of ATR is required for Chk1 phosphorylation after Cdc25B activity after Ser230 phosphorylation by exposure to UV or replicative stress. A plethora Chk1 might be brought about by the creation of a of proteins is known to play a role as DNA dam14-3-3 protein binding site at Ser230, together with age sensors and, as such, are promptly recruited Ser151 and Ser323 [62, 63]. 14-3-3 protein binding to sites of DNA damage. The Mre11/Rad50/Nbs1 to these phosphorylation sites has been reported (MRN) complex, MDC1, 53BP1, RPA, Rad17, and to inactivate Cdc25B by blocking the access of its microcephalin (MCPH1) are all components of the early DNA damage response complex. Defects in substrates to the catalytic site. these molecules disrupt the DNA damage response and lead not only to genomic instability and cancer 4.2.2 Centrosomal Regulation of the development but also immunological and neuroDNA-Damage-Induced G2/M Checkpoint logical defects including reduced brain size and mental retardation. Prominent examples 4.2.2.1 DNA Damage Checkpoint Network are ataxia telangiectasia, Nijmegen breakage synThe cellular machinery that responds to dam- drome, AT-like disorder, and possibly Li-Fraumeni aged DNA encompasses a dynamic network of syndrome due to aberrations in ATM, NBS1, hierarchically ordered proteins and multiprotein Mre11, and Chk2 [68], respectively and the microcomplexes capable of detecting DNA lesions cephalic disorders, Seckel syndrome and primary and signaling their presence to activate path- microcephaly, caused by hypomorphic mutations ways that delay cell cycle progression, repair in ATR or defective ATR signaling [69, 70] and the DNA lesions, or eliminate the genetically mutations in MCPH1 [71], respectively.


4.2.2.2 Role of Centrosomes While it has been unambiguously shown that centrosomes are involved in the regulation of progression through unperturbed cell cycles, the question remains whether regulatory pathways localized to the centrosome are also implicated in the checkpoint response to DNA damage. However, growing data indeed argue in favor of such a possibility. First, both Chk1 and Chk2 have been shown to accumulate at centrosomes of several human cell lines in response to DNA damage induced by UV or ionizing irradiation and bleomycin or hydroxyurea treatment [5, 72, 73]. In all cases, the DNA damage-induced centrosomal pools of the checkpoint kinases have been found to be phosphorylated at ATR/ATM target sites. These findings are in agreement with earlier data showing that, in syncytial Drosophila embryos, DNA damage leads to an increased loading of DmChk2 onto centrosomes and results in mitotic catastrophe with disruption of spindle assembly and chromosome segregation [74, 75]. Genetic ablation of ATR or chemical inhibition of ATM/ATR by caffeine led to nuclear export and pronounced centrosomal accumulation of Chk1 even in the absence of DNA damage [72]. These findings might be explained by data suggesting that basal activity of ATR is required to keep the chromatinassociated pool of Chk1 within the nucleus [76]. However, others have reported opposite findings with phosphorylation of Chk1 at serines 317 and 345 being required for its cytoplasmic and centrosomal localization [73]. Therefore, another explanation is that an impaired response to deregulated DNA replication leads to pronounced centrosomal accumulation of Chk1 after caffeine treatment and in ATR-deficient Seckel fibroblasts without exogenous DNA damage. As a consequence, centrosomal accumulation of Chk1 might prevent CDK1/cyclin B activation as a backup mechanism in the absence of ATR/ATM activity, a finding that is consistent with the low proliferative potential of ATR-Seckel fibroblasts. Second, specific interference with centrosomal Chk1 activity is sufficient to disrupt DNA damageinduced G2/M phase arrest, thereby allocating a specific role for the centrosomal pool of Chk1 during this checkpoint response [72]. While expression

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of wild-type Chk1 forced to the centrosome via a centrosomal targeting domain rapidly downregulates the proportion of cells in mitosis after DNA damage has occurred, reflecting an intact G2/M checkpoint response, its kinase-dead counterpart led to checkpoint deficiency with impaired reduction of the mitotic fraction [72]. Also, nonphosphorylatable S317A/S345A Chk1 versions were only capable of inhibiting cell death in Chk1-deficient cells after DNA damage when they were tagged to the centrosomes [73]. In contrast, neither centrosome-tagged wild-type nor S317A/S345A Chk1 versions rescue defective intra-S phase checkpoint function, reflecting the fact that the target of the intra-S phase checkpoint is nuclear Cdk2 and not centrosomal Cdk1, as is the case for the G2/M phase checkpoint. From these findings it can be concluded that (1) phosphorylation of Chk1 at ATR/ATM target sites seems to play a role in centrosomal targeting of the checkpoint kinase and that (2) the centrosomal pool of Chk1 appears to be, in addition to its role during unperturbed cell cycle progression, involved in the response to DNA damage as well. Although the data for a centrosomal role of Chk2 in mammalian cells are sparse, the recruitment of activated Chk2 to the centrosome might additionally contribute to prevent Cdk1 activation and thereby mitotic entry after DNA damage, consistent with a model for the cooperation between nuclear Chk1 – a “workhorse” that is active even during normal, unperturbed cell cycles – and nuclear Chk2 – an “amplifier” which is additionally active only after DNA damage [50]. Fitting to this overall concept, it has recently been reported that, upon activation of the G2/M checkpoint by DNA damage, activating phosphorylation of centrosomal Cdc25B at Ser353 does not occur [77]. The lack of centrosomal Cdc25B phosphorylation in cells arrested at the G2/M DNA damage checkpoint could be attributed to inactivation of Aurora-A following DNA damage. Chemical inhibition of Chk1 does not only abrogate the G2/M checkpoint but has been shown to induce centrosomal Aurora-A activation as evidenced by phosphorylation at Thr288 and subsequent Cdc25 activation via phosphorylation at Ser 353, implying that centrosomal Aurora-A kinase activity is directly or indirectly controlled by Chk1 following DNA damage.

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4.2.2.3 Factors Impacting Centrosomal Regulation Recently, it has been found that in addition to hypomorphic ATR mutations, mutations in pericentrin cause Seckel syndrome with defective ATRdependent DNA damage signaling [78]. Almost simultaneously, pericentrin mutations have been identified as the cause of microcephalic osteodysplastic primordial dwarfism type II (MOPD II) [79]. Pericentrin is a large coiled-coil protein with a C-terminal PACT domain that targets it to the centrosome [80]. Pericentrin localizes to the pericentriolar material, where it serves as a scaffolding protein and, as such, interacts with several structural and regulatory centrosomal proteins, including g-tubulin, protein kinase A as well as protein kinase CbII [81–84] and plays an important role in microtubule nucleation and spindle organization [81, 85]. Accordingly, antibodies to pericentrin disrupt mitosis, suggesting that it is essential for mitotic progression [82]. In cells from patients with pericentrin-Seckel syndrome, pericentrin is completely lost from centrosomes, a finding that is explained by the fact that truncating mutations within the protein lead to loss of the C-terminal PACT domain [78]. Pericentrin-Seckel like ATR-Seckel cells show defects in ATR-dependent G2/M checkpoint control, for the first time providing evidence for a role of a structural centrosomal protein in cell cycle and checkpoint regulation. This idea is further supported by the description of pericentrin mutations in MOPD II [79]. In this disorder, loss of pericentrin from the centrosome is associated with mitotic failure, cell death, and growth restriction [86]. Mechanistically, it has been speculated that loss of pericentrin might preclude centrosomal recruitment of Chk1, thereby preventing the regulation of mitotic entry via control of the initial activation of cyclin B/Cdk1 at the centrosome both during unperturbed cell cycles and after DNA damage [5, 78]. As already pointed out above, primary microcephaly is a neurodevelopmental disorder that is, analogous to Seckel syndrome and MOPD II, characterized by markedly reduced brain size [87]. The first causative gene identified encodes microcephalin (MCPH1), a protein implicated in the ATR DNA damage response pathway [71, 88]. How exactly MCPH1 is involved in ATR checkpoint signaling remains controversial [89]. Evidence for both functions up- and

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down-stream of Chk1 have been reported [88–90]. In addition to its nuclear localization, microcephalin has been shown to localize to centrosomes as well [91, 92]. Microcephaly of patients with MCPH1 mutations reflects defects in neurogenesis due to defective mitosis in neural precursor cells. Initial studies suggested that microcephalin could play at least two roles in cell physiology, in the regulation of unperturbed mitotic cell cycles, and in response to genotoxic stress. Importantly, in addition to its role in primary microcephaly, decreased levels and diverse mutations of microcephalin have been found in several types of human cancer [90].

4.3 DNA-Damage-Induced Centrosome Amplification 4.3.1 Mechanisms of DNA-Damageinduced Centrosome Amplification Abnormal centrosome numbers are common in tumors and occur both after DNA damage and in cells defective in DNA damage response and repair. Centrosome amplification has been frequently observed in both solid tumors and hematological malignancies and is linked to aneuploidy and tumorigenesis [93–98]. Centrosome abnormalities in several types of human neoplasias do correlate with the occurrence of karyotype aberrations and with the clinical aggressiveness of the respective disorder [96, 98–103]. Mechanistically, centrosome amplification may lead to chromosomal instability and/or mitotic catastrophe through the formation of multipolar mitotic spindles resulting in chromosome missegregation. There are many observations linking DNA damage to changes in centrosome number. Amplified centrosomes and the formation of multipolar spindles after exposure to DNA damaging agents have been known since the early 1960s [104–106]. Centrosome amplification and multipolar spindle formation have been reported to occur in several tumor as well as nontransformed cell types, e.g. U2OS osteosarcoma cells, KK47 bladder cancer cells, HeLa cervical cancer cells, Jurkat leukemia cells, Hct116 colon carcinoma cells, BJ fibroblasts, CHO hamster cells, and DT40 chicken


cells after exposure to various ionizing radiation dose levels ranging from 2 to 10 Gy [107–112]. UV irradition and DNA damaging agents including hydroxyurea, mitomycin C, arsenite, chromate, benzo[a]pyrene diol epoxide, and 6-thioguanine also induce centrosome amplification and spindle multipolarity [72, 109, 113–115].

4.3.1.1 Genetic Factors Contributing to Centrosome Amplification Homologous recombination is a key pathway for the repair of severe DNA damage such as double strand breaks. Impaired homologous recombination causes deleterious genetic rearrangements, which could have a causative role in malignant transformation [116]. A number of studies have recently shown that loss or inactivation of homologous recombination repair proteins results in centrosome amplification. Prominent among those are Rad51, the Rad51-like proteins Rad51B, Rad51C, Rad51D, XRCC2, and XRCC3, as well as BRCA1 and BRCA2 [117–123]. Also, primary mouse embryonic fibroblasts deficient in the DNA mismatch repair gene Msh2 show centrosome amplification, defective mitotic spindle organization, and unequal chromosome segregation [124]. The poly(ADP-ribose)polymerase (PARP) family of proteins which catalyze poly(ADP-ribosyl)ation and function in DNA damage response and repair, have been linked to centrosome amplification as well. In particular, cells from PARP1-deficient mice exhibit a high frequency of centrosome amplification [125]. Mutational inactivation of p53, reduction of p53 activity by overexpression of the p53 inactivating protein Mdm2 and reduced activity of the downstream targets of p53, p21, and Gadd45 are all known to induce supernumerary centrosomes [126–130]. In addition, overexpression of the DNA damage sensor ATR and its downstream transducer Chk1 lead to centrosome amplification [72, 131].

4.3.1.2 The Process Involved in Centrosome Amplification From a mechanistic point of view centrosome amplification can occur through either overreplication of centrosomes within a single cell cycle or by a cell division/cytokinesis defect associated

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with amplification of both centrosomes and cellular DNA content. Alternatively, centrosomes may fragment into multiple microtubule organizing centers. Hut and coworkers have reported multipolar spindle formation in CHO cells that enter mitosis with damaged or incompletely replicated DNA [109]. In these cases, during mitosis centrosomes split into fragments containing only one centriole, leading to multipolar spindles with extra centrosome-like structures without prior centrosome amplification [109]. Despite spindle multipolarity, cells did exit from mitosis, resulting in severe division errors including cytokinesis defects with subsequent polyploidization. Earlier observations by Sato and colleagues pointed in the same direction by showing that after g-irradiation the fraction of cells containing intact, functionally competent centrosomes decreased over time [106]. Instead, disintegrated centrosomal components with reduced microtubule nucleating capacity appeared, leading the authors to conclude that g-irradiation causes disintegration and loss of functionality of centrosomes during mitosis. In the absence of DNA damage, tetraploidization has been proposed as a major route to centrosome amplification. Meraldi and coworkers reported that overexpression of the mitotic kinases Aurora-A, Aurora-B, and Plk1 – independent from their kinase activity – gives rise to extra centrosomes through defects in cell division and consequent tetraploidization, most pronounced in p53-deficient cells [132]. Whether or not this mechanism applies to DNA damage-dependent centrosome amplification remains to be elucidated. Most data available so far suggest that centrosome amplification induced by DNA damage occurs during a prolonged G2 phase arrest. Initially, it was shown that chicken DT40 cells which conditionally lack Rad51 recombinase and thereby incur high levels of spontaneous DNA damage, arrest in G2 phase and acquire supernumerary centrosomes [133]. This centrosome amplification occurred without additional rounds of DNA replication and was consequently not the result of cytokinesis failure. In addition, under these conditions supernumerary centrosomes were shown to contain paired centrioles by serial section electron microscopy and therefore resulted from bona fide centrosome duplication, rather than centrosome splitting or

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fragmentation. Similar results were obtained when DNA damage was introduced by g-irradiation. In both situations the induction of supernumerary centrosomes was sensitive to G2/M checkpoint abrogation by caffeine, wortmannin, or ATM gene targeting in DT40 cells. Later work demonstrated by electron microscopy that DNA damage induces bona fide centrosome amplification – not fragmentation – in several human cell types including lymphoblastoid cells, U2OS osteosarcoma cells, Hct116 colon carcinoma cells, and Jurkat T cell lymphoma cells as well [111]. Inhibition of ATM/ ATR by caffeine and suppression of Chk1 activity by either siRNA or drug treatment suppressed DNA-damage-induced centrosome amplification [72, 111]. Also, a kinase-dead Chk1 version or Chk1 mutants, which cannot be phosphorylated by ATM/ATR were unable to restore centrosome amplification, showing that signaling to Chk1 and Chk1 kinase activity are necessary to induce supernumerary centrosomes after DNA damage. Corroborating these data, it has been reported recently that exogenous DNA damage leads to centrosomal accumulation of Chk1 in normal human fibroblasts and also in ATM- or ATR-deficient cells [72]. Importantly, the degree of centrosome amplification correlated with the centrosomal Chk1 load in response to DNA damage in the respective cell lines, indicating that DNA damage-induced centrosome amplification might be regulated by centrosomal Chk1. Since expression of a centrosome-tagged version of Chk1 is sufficient to induce centrosome amplification even in the absence of exogenous DNA damage – a phenotype that can be suppressed by addition of caffeine – it can be concluded that both centrosomal localization of Chk1 and its phosphorylation by ATR/ATM are required for centrosome amplification after DNA damage. Taken together, the centrosomal fraction of Chk1 plays an important role for both induction of a G2/M arrest and centrosome amplification in response to DNA damage. The principal role of Chk1 during these processes appears to be to directly relay the DNA damage signal to the centrosome, where it imposes a cell cycle delay by inhibiting Cdk1 activation. In a fraction of cells, centrosomal Chk1 subsequently induces centrosome amplification. The mechanistic basis of this decision in terms of which of the cells with centrosome-accumulated

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Chk1 would undergo centrosome amplification, remains unknown.

4.3.2 Consequences of DNA-Damageinduced Centrosome Amplification Centrosomal aberrations including numerical amplification are common in a variety of cancers, including premalignant and early lesions [94, 134, 135], in which, on the other hand, activation of the DNA damage response pathway does occur frequently [136, 137]. Hence, one might speculate that Chk1dependent centrosome amplification in premalignant lesions and early cancer serves as a checkpoint mechanism helping to eliminate malignant cells via induction of multipolar mitoses with subsequent mitotic catastrophe. It has long been known that the most frequent mode of cell death after irradiation is mitosis-linked death in which cells manifesting chromosomal aberrations during division form nonclonogenic cells containing micronuclei [138, 139]. Importantly, recent data suggest that a significant proportion of cell death induced by ionizing radiation seems to occur as a consequence of centrosome amplification through the induction of multipolar mitoses with subsequent mitotic catastrophe [110, 112]. Notably, in addition to DNA damageinduced centrosome amplification, recent evidence suggests that subsequent formation of multipolar mitoses followed by mitotic catastrophe does also depend on Chk1 kinase activity [115]. Aboli-tion of Chk1, on the other hand, prevents DNA damage-induced centrosome amplification and multipolar mitosis-associated cell death. These data lend support to the above hypothesis that DNA damage-induced centrosome amplification constitutes a mechanism which ensures death of cells that evade the DNA damage or spindle assembly checkpoints.

4.3.3 Centrosome Clustering Evolution from early premalignant lesions to advanced cancer has been shown to be associated with mutations compromising an effective response to DNA damage [136, 137]. Similarly, cancer progression might be paralleled by abrogation of a

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centrosome amplification checkpoint, since it has been shown that many cancer cell lines are capable of coalescing supernumerary centrosomes into two functional spindle poles, a phenomenon that has been termed centrosomal clustering and prevents the occurrence of mitotic catastrophe [140–145]. Since supernumerary centrosomes almost exclusively occur in a wide variety of neoplastic disorders but not in nontransformed cells, it has been argued that the inhibition of centrosomal clustering with consequential induction of multipolar mitotic spindles and subsequent cell death might specifically target tumor cells with no effect on normal cells with a regular centrosome content [144]. Indeed, a recent phenotype-based small molecule screen led to the identification of a drug, griseofulvin, that inhibits centrosomal clustering and subsequently leads to mitotic catastrophe, specifically in tumor cells with multiple centrosomes [144]. In addition to tumor cells, healthy cells, forced to contain supernumerary centrosomes by either ablation of cytokinesis or overexpression of SAK, the ortholog of Plk4 in Drosophila melanogaster, are capable of centrosomal clustering as well [142, 145]. In Drosophila lines with amplified centrosomes, many cells with extra centrosomes initially form multipolar spindles, but these spindles ultimately become bipolar by centrosome coalescence and inactivation. As a result, chromosomal instability does not dramatically increase in these flies which are viable and maintain a stable diploid genome over many generations. Nevertheless, in this model system larval brain cells with extra centrosomes can generate metastatic tumors when transplanted into the abdomen of wild-type hosts, thereby demonstrating that centrosome amplification can initiate tumorigenesis in Drosophila. It is not clear how centrosome amplification initiates tumor formation. In the brains of the Drosophila lines with extra centrosomes the rate of aneuploidy is low due to efficient centrosomal clustering, although it is higher than that observed in wildtype brain cells. Therefore, it is possible that a modest increase in aneuploidy ensures survival by preventing mitotic catastrophe and/or continuous loss of genetic information and at the same time allows some cells with extra centrosomes to initiate tumorigenesis in flies. Untill now, no data on a causal relationship between supernumerary

centrosomes and tumor formation in higher organisms are available.

4.4 Centrosome Damage Checkpoint Two fundamental roles of centrosomes within the DNA damage response have been described above: Firstly, there is plenty of evidence, which demonstrate that centrosomes serve as spatiotemporal organizers where checkpoint components come into proximity to each other allowing for an ordered regulation of cell cycle progression through the G2/M checkpoint. Second, centrosomes seem to be directly subjected to regulation by DNA damage in a way that centrosome amplification might serve as an effector of the DNA damage response. In addition to these DNA damage-induced effects on centrosomes, the existence of a centrosome damage checkpoint has been described. Cell cycle progression through G1 and S phases is not affected in cells with extra centrosomes [146, 147]. In contrast, recent studies indicate that disruption of centrosome integrity is associated with both cytokinesis defects and G1 phase arrest. For example, when centrosomes are removed or disrupted by microsurgery, laser ablation or microinjection of centrosome antibodies, cytokinesis is impaired and cells arrest in G1 [27, 148, 149]. Moreover, recent data demonstrate that siRNA-mediated depletion of several different, single centrosomal proteins induces a p53-dependent cell cycle arrest in G1 phase [150, 151]. The arrest was shown to require p38, p53, and p21, and is preceded by p38dependent activation and centrosomal recruitment of p53. These results suggest the presence of a cell cycle checkpoint that prevents cells from entering S phase when they acquire defects in centrosome structure and/or function. Abrogation of this proposed centrosome damage checkpoint might have deleterious downstream consequences, in that in p53-deficient cells depletion of centrosomal components has been shown to induce spindle defects, cytokinesis failure, and aneuploidy. Similarly, p53deficiency may abrogate this checkpoint in many human tumors, thereby contributing to centrosome defects, spindle dysfunction, and chromosomal instability [94, 126].

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4. Centrosomes in Checkpoint Responses trosomal clustering in a phenotype-based screen. Cancer Res 67(13):6342–6350 145. Basto R, Brunk K, Vinadogrova T et al (2008) Centrosome amplification can initiate tumorigenesis in flies. Cell 133(6):1032–1042 146. Wong C, Stearns T (2005) Mammalian cells lack checkpoints for tetraploidy, aberrant centrosome number, and cytokinesis failure. BMC Cell Biol 6(1):6 147. Uetake Y, Sluder G (2004) Cell cycle progression after cleavage failure: mammalian somatic cells do not possess a "tetraploidy checkpoint". J Cell Biol 165(5):609–615 148. Khodjakov A, Rieder CL (2001) Centrosomes enhance the fidelity of cytokinesis in vertebrates

67 and are required for cell cycle progression. J Cell Biol 153(1):237–242 149. Balczon R, Simerly C, Takahashi D, Schatten G (2002) Arrest of cell cycle progression during first interphase in murine zygotes microinjected with anti-PCM-1 antibodies. Cell Motil Cytoskeleton 52(3):183–192 150. Mikule K, Delaval B, Kaldis P, Jurcyzk A, Hergert P, Doxsey S (2007) Loss of centrosome integrity induces p38–p53-p21-dependent G1-S arrest. Nat Cell Biol 9(2):160–170 151. Srsen V, Gnadt N, Dammermann A, Merdes A (2006) Inhibition of centrosome protein assembly leads to p53-dependent exit from the cell cycle. J Cell Biol 174(5):625–630

Chapter 5

Interplay of 14-3-3 Family of Proteins with DNA Damage-Regulated Molecules in Checkpoint Control Mong-Hong Lee, Sai-Ching Jim Yeung, and Heng-Yin Yang

Abstract DNA damage causes cell cycle delay, and this process involves a number of highly-regulated proteins that sense DNA damage and regulate the cell cycle machinery. For example, tumor suppressor p53 is activated to maintain genome stability, while CDC25 phosphatase is inhibited to prevent further cell cycle progression in response to DNA damage. The 14-3-3 family of phosphoserine/phosphothreonine binding proteins are involved in the regulation of DNA damage check point response, and they coordinate with DNA damage response regulators such as p53 and CDC25 to regulate the cell cycle to promote DNA repair or survival. Here, we discuss recent data that define the roles of 14-3-3 in DNA-damage checkpoint pathway. Keywords Checkpoint response • 14-3-3 • Akt • Mdm2 • p53 • DNA damage

5.1 Introduction The 14-3-3 is a family of about 30 kDa homodimeric proteins found in eukaryotic cells. The 14-3-3 family is highly conserved over a wide range of mammalian species, and its members include seven isotypes b,e, h g, t (also called q), z, and s[1, 2]. 14-3-3 proteins bind to phosphoserine/phosphothreoninecontaining peptide motifs: Mode-1 (RSXpSXP) or Mode-2 (RXXXpSXP) sequences. Notably, 14-3-3 proteins interact with more than 200 target proteins in a phosphoserine-dependent (mode-1 and -2) as well as phosphoserine-independent manner [3–6].

By interacting with these various proteins, 14-3-3 family members have many diverse functions, including critical roles in signal transduction pathways and cell cycle regulation in response to DNA damage checkpoint control [7–9]. The DNA damage checkpoint is a signal transduction pathway, induced by DNA damage, that inhibits cell cycle progression in G1, G2, or M-phases or slows the progression rate of S phase. Following DNA damage, DNA repair and cellcycle checkpoints will activate several important regulators (Fig. 5.1) that are regulated by or collaborate with 14-3-3 family proteins to regulate cell cycle arrest, cell survival, or preserve the integrity of chromosomes (Table 1). Recent studies have provided new mechanisms and revealed new functions of 14-3-3 in DNA damage control. Here, we review the interplay of 14-3-3 family proteins with key DNA damage-regulated molecules to offer insights on DNA damage checkpoint control.

5.2 p53 Loss of control of genomic stability is crucial in the development of cancer, and p53 is important in regulating responses to DNA damage to maintain this stability [10]. p53 is an important tumor suppressor involved in DNA damage check point control to regulate DNA repair and thus maintain genome integrity. DNA damage induces the activation of ATM, ataxia telengiectasia mutated, which in turn


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Figure 5.1. The cellular response to DNA damage. Proteins regulating various signaling events including cell cycle and DNA damage checkpoint responses are shown. 14-3-3 family members regulate or collaborate with these key proteins. See text for details.

controls genome stability and cell cycle progression [11]. When chromatin is altered structurally after DNA damage, it triggers a signal that causes a cross-phosphorylation of Ser1981 in ATM dimers and subsequent disassociation of dimers [11]. The ATM monomer then phosphorylates downstream targets for DNA damage response, including p53 and MDM2 [12, 13]. ATM can phosphorylate p53 at Ser15, which leads to the transactivating function of p53 by facilitating the interaction between p53 and p300 [14]. ATM signaling can also lead to phosphorylation at p53 Ser 20 [15, 16], which together with phosphorylation at Ser 15 interferes with the binding of p53 to MDM2 [17]. In addition, ATM can phosphorylate MDM2 at Ser 395 and block the p53 nuclear exporting activity of MDM2 [13], thereby stabilizing p53. The significance of ATM-mediated phosphorylation is demonstrated by the observation that substitution of Ser 395 to aspartic acid, which mimics phosphorylation, can compromise MDM2-mediated p53 degradation and nuclear export [12].

We will first discuss how 14-3-3 regulates p53 during the DNA damage checkpoint control. ATM also has impacts on MDM2 or Chk, which are regulated by 14-3-3, and this aspect will be discussed later. p53 is a transcription factor and the 14-3-3s gene is a p53-inducible gene involved in cell-cycle checkpoint control after DNA damage [18]. 14-3-3s functions as a negative regulator of cyclin-dependent kinases (CDKs) [19]. Thus, DNA damage-induced upregulation of p53 leads to the expression of 14-3-3 s to control cell cycle progression in addition to p21, another major CDK inhibitor induced by p53. Unlike p21, 14-3-3s has a positive feed back regulation on the activity of p53 [20]. First, 14-3-3s promotes cooperative p53 dimer–dimer interaction [20] to stabilize p53 as a homotetramer for DNA binding and potentiates p53s transcriptional activity [20]. Members of the 14-3-3 protein family have been shown to act as adapter proteins binding to many signal proteins to exert biological function [21]. Thus, 14-3-3s seems to use this characteristic to positively regulate p53.

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Second, 14-3-3s binds MDM2, an ubiquitin ligase and negative regulator of p53, to compromise MDM2mediated p53 ubiquitination and thus stabilize p53 [20]. The 14-3-3s-mediated p53 stabilization is primarily due to an increase in p53 halflife [20]. At the same time, elevated levels of 14-3-3s lead to a decrease in the halflife of MDM2 [22]. These observations indicate that 14-3-3s impacts the p53 negative regulator, MDM2, to stabilize p53. 14-3-3 members can affect target protein enzyme activity [1]. In this case, 14-3-3s is able to affect the enzyme activity of MDM2 by enhancing MDM2 self-ubiquitination [22]. Third, 14-3-3s blocks MDM2-mediated p53 nuclear export very efficiently, thereby stabilizing and retaining p53 in the nucleus [20]. This is due to the fact that 14-3-3s translocated MDM2 from the nucleus to the cytoplasm [22]. Thus,14-3-3 members can influence the subcellular localization of target protein, and it appears that 14-3-3 uses its activity to affect the subcellular localization of both MDM2 and p53 [22]. Other 14-3-3 isoforms (g, e, t) also bind to p53, supposedly through Ser378 phosphorylation by ATM, to induce putative conformation change of p53 [23]. However, the details and the impact of this binding remain unclear. Although 14-3-3 family members are structurally related, their impacts on p53 activity are different. Recent data have shown that p53 expression is reduced in 14-3-3z transgenic mice, and 14-3-3z appears to activate MDM2-mediated p53 degradation [24].

of MDM2 [22], and antagonizes MDM2-mediated p53 nuclear export. Fourth, 14-3-3s blocks MDM2mediated p53 NEDDylation [22]. In addition to ubiquitination activity, MDM2 uses its C-terminal RING domain to promote p53 NEDDylation at three lysine residues of p53 [26]. NEDDylation appears to have a negative impact on the transactivation activity of p53 [26], and 14-3-3s can antagonize MDM2-mediated p53 NEDDylation [22], which in turn will take away the inhibitory effect of NEDDylation on p53 transcriptional activity. This mechanism, in which 14-3-3s causes MDM2 downregulation to stabilize p53 and inhibit tumor growth, has been confirmed in animal tumor studies [22]. These results provide evidence that 14-3-3s is a pivotal MDM2 regulator. Recent data indicate that 14-3-3z can enhance MDM2 nuclear localization to facilitate p53 degradation [24]. Also, MDMX, a close homologue of MDM2, is implicated in suppressing p53 transcriptional activity through binding to its transactivation domain [27]. Studies indicated that MDMX can bind to and stabilize MDM2. Importantly, 14-3-3e can bind to MDMX at the C terminus (Ser367) in a phosphorylation-dependent manner [28]. Protein kinase B (PKB, also called Akt) phosphorylates MDMX at Ser367 to recruit 14-3-3 to bind the MDM2MDM4 complex, leading to stabilization of both MDMX and MDM2. Again, these observations indicate that 14-3-3 isoforms have different impact on MDM2.

5.3 MDM2

5.4 Akt

MDM2 is an E3 ubiquitin ligase for p53 that decreases the stability and halflife of p53 through the ubiquitin-proteasome pathway [25]. As a negative regulator of p53, MDM2 is recognized as an oncogene that is dysregulated in cancer. Detailed biochemical studies also indicate that 14-3-3s has important impact on this oncogene. First, 14-3-3s interacts with MDM2 at the RING domain [22], which is important for the ubiquitin ligase activity of MDM2. This association is phosphorylationdependent as the interaction is compromised when phosphatases in the cells are not inhibited [22]. Second, 14-3-3s decreases the stability of MDM2 by facilitating the self-ubiquitination of MDM2 [22]. Third, 14-3-3s induces the cytoplasmic location

Akt is the cellular homologue of the oncogene of the AKT8 oncovirus (v-Akt). Akt is activated when particular extracellular signals activate receptor tyrosine kinases to enhance phosphatidylinositide 3-OH kinase (PI3K) activity on phospholipids. Akt is an oncogene crucial for a variety of cellular processes, including cell survival and proliferation. Several line of evidence has demonstrated its role in the DNA damage checkpoint. First, Akt activity is down regulated in response to DNA damage [29]. Second, Akt also affects p53 activity by phosphorylating MDM2 [30, 31] at Ser 166 and Ser 188 [30], inhibiting MDM2 self-ubiquitination [32]. Akt-mediated phosphorylation of MDM2 also promotes nuclear localization of MDM2 and

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inhibits interaction between MDM2 and p19ARF, thereby decreasing p53 stability [30]. Third, Akt can phosphoryate Miz1 to prevent its DNA binding activity [33]. Miz1 is required for DNA damage response. Akt-mediated phosphorylation of Miz1 compromises the role of Miz1 in regulating DNA damage response [33]. Lastly, Akt can phosphorylate and reduce the nuclear localization of Chk1 [34], thereby interfering with Chk1-mediated p53 phosphorylation and subsequent p53 stabilization. Given that activated Akt can have these aforementioned effects on the DNA damage response, Akt activity must be properly restrained in response to DNA damage. 14-3-3s can regulate the subcellular localization of MDM2 despite the presence of activated Akt [29]. It is conceivable that 14-3-3s can antagonize Akt-mediated MDM2 activation to stabilize p53. 14-3-3s can antagonize the survival benefits of Akt in the presence of DNA damaging drugs [29]. Increased physical association between 14-3-3s and Akt correlates with inactivation of Akt in response to DNA damage [29] Thus, it is clear that 14-3-3 s plays an important role in restraining Akt in response to DNA damage. Recent data shows that 14-3-3z can induce hyperactivation of the phosphoinositide 3-kinase/ Akt pathway, which in turn activates phosphorylation and translocation of the MDM2 E3 ligase to antagonize p53 activity [24]. Also, 14-3-3e collaborates with Akt to stabilize MDM2-MDM4 complex, which will enhance MDM2-mediated p53 ubiquitination [28]. 14-3-3h serves as a regulator of Akt to abolish the DNA binding activity of Miz1 in inducing p21 in response to DNA damage [33]. It is worthwhile to mention that several isotypes of 14-3-3 are substrates of Akt and are involved in suppressing apoptosis by binding to the phosphorylated death agonist BAD [35], inhibiting apoptosis signal-regulating kinase 1 [36], or binding to the forkhead transcription factor to block the gene expression involved in apoptosis [37]. Therefore, some 14-3-3 family members, unlike 14-3-3s, are indeed mediators of antiapoptotic signals and antagonize the proapoptotic function of p53. The mammalian target of rapamycin complex 1 (mTORC1) is positively regulated by amino acids and insulin, and play important roles in cellular regulation and human disease such as cancer [38] [39]. mTORC1 regulates the phosphorylation proteins

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involved in the control of mRNA translation, such as the eukaryotic initiation factor 4E-binding proteins (4E-BPs) [40]. It is known that Akt-mediated phosphorylation and inhibition of TSC2 (tuberous sclerosis complex 2), which inhibits its ability to promote hydrolysis of GTP bound to the G-protein Rheb (Ras homologue enriched in the brain), a positive regulator of mTORC1, thereby promoting mTORC1 activity [40, 41]. Recently, Akt has another layer of regulation to activate mTORC1 through its activity on PRAS40. PRAS40, (prolinerich Akt substrate of 40 kDa) is associated with mTORC1 complex and binds to the c-terminal kinase domain of mTOR. PRAS40 can negatively regulate the mTOR pathway [42, 43]. Importantly, PRAS40 is phosphorylated by Akt at thr 246, resulting in 14-3-3 binding and loose association of PRAS 40 with mTOR, thus PRAS40 no longer inhibits mTOR activity [42]. Together, Akt and other 14-3-3 isoforms enhance the mTOR activity, which in turn will have a positive impact on the cap-dependent translation of mRNA as mTOR is phosphorylating 4EBP1, thus allowing the eIF4E factor for participating in the cap-dependent translation of mRNA. As described below, this capdependent mRNA translation is antagonized by 14-3-3s through its interaction with eIF4B [44]. Given that 14-3-3s is an inhibitor of Akt, 14-33s may also inhibit the Akt-PRAS40 axis, thus antagonizing mTORC1 activity. These two kinds of activity contribute to 14-3-3s’s activity in suppressing cap-dependent translation of mRNA, which will affect cytokinesis (see below) [44] and have roles in DNA-damage responses.

5.5 Myc and miz1 Miz1, a Myc-interacting zinc finger protein, is a transcription factor required for DNA-damageinduced cell-cycle arrest. Miz1 is involved in the transcriptional control of the p15Ink4b and p21Cip1 genes. Recently, loss of Miz1 can compromise the induction of DNA damage (UV)-induced expression of p21 [33]. 14-3-3h interacts with the DNA binding domain of Miz1, which is phosphorylated by Akt at ser 428, thereby inhibiting Miz1 binding activity to DNA [33]. Also, Myc represses the transcriptional activation by Miz1. Thus, both Akt and


Myc modulate Miz1 activity in response to DNA damage. Given that 14-3-3s is an inhibitor of Akt [29], it will be worthwhile to investigate whether 14-3-3s will inhibit Akt-mediated Miz1 phosphorylation at ser428 and preserve the DNA binding activity of Miz1 to regulate genes in response to DNA damage. It is possible that 14-3-3s and 14-3-3 h are antagonizing each other to regulate Miz1 activity in response to DNA damage. c-Myc, a transcription factor that can induce tumorigenesis by promoting cell proliferation and blocking cell differentiation [45]. Importantly, c-Myc also contributes to tumorigenesis as an important antagonist of the DNA damage-sensing mechanism [46]. Modest overexpression of c-Myc is sufficient to disrupt the checkpoint control and impede the repair of DNA double-strand breaks aggravating genomic instability thereby accelerating tumor progression. As a transcription repressor, c-Myc can impair the activation of p53-target gene promoters such as p21WAF1, 14-3-3s, MDM2, GADD45, PERP, and NOXA [47]. Given that 14-3-3s is activated to potentiate the transcriptional activity of p53, the 14-3-3-mediated positive feedback loop regulation is, therefore, compromised. Thus, c-Myc can inactivate the p53–14-3-3s or p53–p21 axes to regulate the DNA damage response. It is important to point out that Fbxw7, a p53 target gene product, is an ubiquitin ligase responsible for c-Myc degradation [48] [49]. c-Myc degradation involves Pin1/Fbxw7-mediated ubiquitination. Basically, Pin1 binds to the phosphorylated Thr58Pro motif of c-Myc, leading to isomerization of c-Myc, PP2A-mediated dephosphorylation of Ser62, and subsequent Fbxw7-mediated ubiquitination [50]. The ubiquitinated form of c-Myc is degraded through 26S proteasomes [49]. Therefore, p53 can reciprocally regulate c-Myc stability through accelerating the ubiquitin-mediated degradation process. It is important to note that Fbxw7 is a tumor suppressor downregulated or mutated in many types of cancer [51, 52]. Mutation of FBXW7 occurs in human malignancies of diverse tissue origins. Some of these mutants can have dominant-negative impact or allow oncogenic substrate to escape Fbxw7-mediated degradation. Loss of Fbxw7 leads to genomic instability, which is at least in part due to the loss of the activity to degrade c-Myc [52].

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5.6 Chk1 and Chk2 Cells respond to DNA damage by activating a complex DNA damage response pathway that induces cellcycle arrest and transcriptional and posttranscriptional activation of a subset of genes. This process provides time for repair, prevents mutations from being propagated and is very important for maintaining the integrity of the genome. Double strand breaks, a common form of DNA damage, activate the important ataxia telangiectasia mutated (ATM) and ataxia telangiectasia Rad3related (ATR) kinases [53]. ATM and ATR directly and indirectly mediate a series of posttranslational modifications to activate and stabilize p53 [54, 55] [56], which is the guardian of genome integrity. Activated ATM or ATR also phosphorylates checkpoint kinase CHK2 or CHK1 [57], which in turn inactivates the phosphatase Cdc25C and leads to inhibition of Cdc2 and G2 arrest. This process requires CHK1 and CHK2 to phosphorylate a conserved site (Ser216) on protein phosphatase Cdc25C [58], which results in its sequestration in the cytoplasm by the 14-3-3 isoforms [59-62]. Importantly, CHK2 is mutated in patients with Li-Fraumeni syndrome type 2 (Li-Fraumeni-like syndrome) [63] in the absence of p53 mutations. Also, the CHK2 gene mutations are detected in sporadic colon, lung, and breast cancers. These observations suggest that CHK2 functions as a tumor suppressor [64, 65]. It was shown that CHK2-/embryonic stem cells failed to maintain G2 arrest in response to g-irradiation [66] [16]. Also, CHK2/- cells are defective in the stabilization of p53 [15] and fail to induce p21 in response to DNA damage [66]. Methods for inhibition of CHK1 or CHK2 kinase activity may selectively sensitize cells to DNA-damaging agents including chemotherapeutic drugs or g-irradiation and may be useful for the treatment of cancer [67] [68]. It has been shown that 14-3-3 proteins can directly interact with Chk1 [69] although detailed biological significance of this interaction is not clear. One possibility is that it can direct Chk1 to certain substrate. For example, Chk1 can phosphorylate MDMX at Ser 367, facilitating the 14-3-3g-MDMX binding and the cytoplasmic retaining of MDMX [70]. Akt can phosphorylate CHK1 at Ser 280, which in turn leads to ubiquitination [34] and cytoplasmic

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CHK1 localization [34]. Elevated Akt activity can increase cytoplasmic CHK1 and promote aneuploidy [34]. Thus Akt activity can compromise activity of Chk1 through phosphorylation, ubiquitination, and reduced nuclear localization to promote genomic instability. Chk1 is important in regulating CDC25B or C to cause their subcellular localization change by phosphorylation, which will facilitate 14-3-3 isoforms and CDC25 interaction, thereby compromising the positive effect of CDC25 toward Cyclin B/ CDC2 activity that is required for G2/M progression. Given that 14-3-3s can inhibit Akt activity, it is possible that 14-3-3s will have impact on Chk1-CDC25 axis to regulate DNA damage responses.

5.7 BRCA1 BRCA1 is a tumor suppressor mutated in 5% of breast and ovarian cancers with important roles in DNA repair and transcription [71]. After DNA damage BRCA1 protein is phosphorylated and activated by ATM, ATR, and Chk2 protein kinases. BRCA1 is a transcriptional co-activator of p53 protein [72], and loss of BRCA1 will result in decreased expression of several groups of genes including stress response genes, cytoskeleton genes, and genes involved in protein synthesis and degradation. Surprisingly, 14-3-3s is the only p53 target downregulated in BRCA1 knockout cells, while the majority of p53 target genes remained without any change [73]. Indeed, BRCA1 interacts with p53 to transcriptionally activate the gene expression of 14-3-3s [73]. Transcription of 14-3-3s is important for BRCA1’s role as a regulator for cell cycle arrest and checkpoint control as BRCA1deficient cells were unable to sustain G2/M arrest after DNA damage caused by ionizing radiation. This link is biologically significant since BRCA1 has important roles in DNA repair and transcription [71, 74]. Mutations identified in the C terminus of BRCA1 from patients with breast cancer cannot activate transcription of 14-3-3s [73], suggesting that the tumor-suppressive function of BRCA1 involves 14-3-3s. It is important to point out that BRCA1 has no impact on 14-3-3s expression in the absence of p53, which is consistent with the observation that BRCA1 requires wt p53 to activate p53 response elements [72]. The interplay between

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14-3-3s, p53, and BRCA1 provide important concept of the checkpoint control among these tumor suppressor genes. Synergistic interaction between BRCA1 and p53 in regulating 14-3-3s can enhance 14-3-3s’s role as a CDK and Akt inhibitor to potentiate p53 activity. On the other hand, haploinsufficiency in either BRCA1 or p53 genes might lead to an inadequate 14-3-3s response to DNA damage. Given that both BRCA1 and 14-33s associate with p53, it remains to be determined whether 14-3-3s affects the biological activity of BRCA1 in DNA double strand break repair or cell cycle checkpoint function. For example, it will be interesting to learn whether 14-3-3s affects BRCA1’s activity on the promoter of GADD45 [75, 76], a protein involved in DNA checkpoint. Additionally, it is important to know whether 14-3-3s affects the heterodimeric complex formation between BRCA1/BARD1 and the complex’s ubiquitination activity after DNA damage [77, 78]. In addition to its positive impact on 14-3-3s, it is important to point out that BRCA1 also plays roles in activating Chk1 kinase and regulating CDC25C subcellular localization [79]. BRCA1 expression potentiates Chk1 kinase activity toward CDC25C. Also BRCA1 expression leads to CDC25C exclusion from the nucleus [79]. BRCA1 also increased the amount of Wee1 kinase, another inhibitor of Cdc2 activity, resulting in increased phosphorylation of Tyr on CDC2 [79].

5.8 Cell Cycle Regulators 14-3-3 family is known to regulate cell cycle progression, and several molecular mechanisms behind cell cycle regulation have been studied. For example, first, 14-3-3e and 14-3-3b have been isolated in a yeast two hybrid screen designed to identify proteins that interact with the human CDC25A and CDC25B phosphatases. They bind to CDC25 but do not affect the phosphatase activities of CDC25 [80-82]. Second, the 14-3-3z has been found to interact with the Wee1 kinase [83], which plays a key role in cell cycle progression by inactivating cyclin-dependent kinases. Recent data show that the binding of 14-3-3 to Wee1 can positively regulate the activity of Wee1 [84]. Third, 14-3-3 isoforms have been shown to interact with polyoma middle T antigen [85] involved in cell proliferation,


but it remains to be elucidated how dysregulation of 14-3-3 proteins contributes to the development of neoplasia. Fourth, in yeast, two checkpoint genes, rad24 and rad25, encode 14-3-3 protein homologues that together provide a function that is essential for cell proliferation [60, 86]. We will discuss the interaction between 14-3-3 family members with some of these cell cycle regulators (CDC25, CDK, and CDK11) and their roles in the DNA-damage checkpoint regulation.

5.8.1 CDC25 For cells to enter the mitosis, Cyclin B/cdc2 needs to be activated. CDC25 family of dual specificity phosphatases activates cdc2/cyclin B in mitosis by removing inhibitory phosphorylations from Thr-14 and Tyr-15 residues in cdc2 [87]. CDC25B and CDC25C are two isoforms that are responsible for activating cdc2/cyclin B in mitosis. CDC25B is active during G2 phase while CDC25C is activated at the G2/M transition. Overexpression of dominant-negative mutants of CDC25B and CDC25C leads to G2 phase arrest, confirming their roles in mitosis. Thus the role of CDC25 proteins is to ensure activation of cdc2/cyclin B and facilitate mitosis entry. 14-3-3 isoforms binds to CDC25B and regulates the subcellular redistribution of CDC25B from the nucleus to the cytoplasm during G2 checkpoint after CDC25B is phosphorylated by Chk. 14-3-3 isoforms can bind to CDC25B phosphorylated at Ser323 and facilitates CDC25B to stay in the cytoplasm [81, 88]. As expected, mutation of Ser323 to alanine can completely block the cytoplasmic localization of CDC25B [81]. It is important to point out that not every isoform bind to CDC25B tightly; Cdc25B binds tightest to 14-3-3b, h, and z. During interphase, Cdc25C is phosphorylated on Ser216, which can be recognized and bound by 14-3-3 proteins [58, 82, 89]. The binding of 14-3-3 leads to the masking of the NLS on Cdc25C. This complex is then excluded from the nucleus through the NES on CDC25C. At the entry of mitosis, Ser216 phosphorylation is dephosphorylated, and 14-3-3 is released. The resultant exposure of NLS of CDC25C then causes relocation into the nucleus in prophase to activate cyclinB/ CDC2. It is important to point out that Chk1 is activated by DNA damage and will be phosphorylating CDC25C on Ser216, causing the

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14-3-3 (14-3-3g, e, and z) binding again and thus interfering with CDC25C’s activity in activating cyclinB/CDC2. Recently, MAPKAP kinase -2, a new DNA damage checkpoint kinase, is able to phosphorylate CDC25B and C to cause 14-3-3 binding in response to UV-induced DNA damage in mammalian cells [90]. Again, 14-3-3 isoforms play an important role in regulating CDC25 activity in the MAPKAP kinase 2 pathway.

5.8.2 CDK Cyclin-dependent kinases (CDKs) and their cyclin partners are positive regulators that induce cell cycle progression, whereas important negative regulators, such as cyclin-dependent kinase inhibitors (CKIs), act as brakes to stop cell cycle progression in response to regulatory signals [87]. For cells to grow normally, the positive and negative regulators of the cell cycle machinery must be very carefully controlled. Any deregulation of this machinery will result in abnormal cell growth. CDKs may be the most highly regulated enzymes characterized, and multiple mechanisms exist to regulate their activity. The 14-3-3s gene is induced by p53 for involvement in G2 cell cycle checkpoint control after DNA damage [18] because it has activity to inhibit cyclin B1/Cdc2 activity [19, 91]. Interestingly, purified 14-3-3s cannot inhibit cyclin-CDK activity in vitro although overexpression of 14-3-3s in the cells can inhibit Cdc2-associated kinase activity and cell cycle progression [19], suggesting that a biological environment is required to assay its CDK inhibitory activity. Molecular mechanistic studies showed that 14-3-3s sequesters cyclin B1/CDC2 complexes in the cytoplasm to cause G2 arrest [19, 91], and its absence allows cyclin B1/Cdc2 complexes to enter the nucleus, causing mitotic catastrophe. During this process, the NES of 14-33s is required to sequester Cdc2 in the cytoplasm [19]. In addition to Cdc2, 14-3-3s has also been shown to specifically interact with CDK2 and CDK4 and can inhibit CDK activities to block cell cycle progression, thus defining it as a new class of CKI [19, 87]. Thus, 14-3-3s can directly bind and sequester CDK into the cytoplasm to inhibit its activity required for cell cycle progression, which may contribute to radiation-induced p53-mediated cell cycle arrest. CDK activity controls cell proliferation and is involved in the development of

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human cancer. The expression of 14-3-3s, like the function of CDK inhibitors such as p21, p27, and p57, is one of the mechanisms that cells employ to ensure the maintenance of a nonproliferative state [87]. Recent data indicate that 14-3-3s has a positive impact on retinoblastoma protein RB [92], a tumor suppressor and G1-S restriction point gatekeeper, thus controlling cell cycle progression. First, it is shown that disabling MDM2 leads to inhibition of DNA synthesis in an RB-dependent pathway [93]. That MDM2 can degrade RB in a proteasome-dependent pathway, providing an explanation for such a phenomenon [93, 94]. Importantly, 14-3-3s is required to block MDM2-meditated RB degradation [92], and expression of 14-3-3s efficiently blocks MDM2-mediated RB degradation. Second, 14-3-3s can inhibit activity of G1 cyclin CDK [19], such as CDK4, which usually catalyzes the phosphorylation of Rb, allowing cell cycle to pass the G1-S restriction point gate. It then provides reason why 14-3-3s expression level is significantly reduced in many types of cancer.

5.8.3 CDK11 14-3-3s are known to regulate the cell cycle. Cdk11/ PITSLRE, a member of the Cdc2-like protein kinase family, is also regulated by 14-3-3s [44]. CDK11 undergoes a cap-independent translation from an internal ribosome entry site during mitosis to produce a 58-kDa isoform that is important for proper mitotic progression. 14-3-3s plays a pivotal role in normal mitosis to suppress the cap-dependent translation through binding eIF4B, allowing the cap-independent translation of important mitotic regulators [44]. Cells lacking 14-3-3s cannot suppress cap-dependent translation and fail to stimulate cap-independent translation during and immediately after mitosis. One of the important genes regulated in this manner is CDK11. Depletion of 14-3-3s leads to blocking of IRES-dependent mitotic translation of p58 PITSLRE kinase [44]. Failure of expression of CDK11 results in failed cytokinesis, loss of Polo-like kinase-1 at the midbody, and accumulation of binucleated cells [44]. It is conceivable that loss of 14-3-3s can cause genomic instability and 14-3-3s deficiency leads to abnormal chromosomal number and DNA breakage.

M.-H. Lee et al.

5.9 Conclusion We have discussed several aspects of the DNAdamage checkpoint pathways regulated by members of the 14-3-3 family. 14-3-3 proteins are a class of regulatory proteins found in all eukaryotic cells that have phosphoserine/phosphothreonine binding activities, and they bind a large number of proteins to regulate various signaling events including cell cycle and DNA damage checkpoint responses, apoptosis, and transcriptional control. We particularly focused on several checkpoint response regulators in Fig. 5.1, and on mechanisms that govern how 14-3-3 family members regulate the function of these regulators. The specific involvement of 14-3-3 family in regulating some of these mediators in DNA damage response is interesting, especially the Akt signaling pathway, adding a new landscape to the activities of 14-3-3s. For example, Akt can impact the subcellular localization of MDM2 and Chk1. Akt regulates myc-Miz1 axis to affect Miz1 DNA binding activity, thus regulating p21 gene expression. In addition, Akt regulates MTORC1 complex activity via PRAS40, which in turn impacts eIF4B and the cap-dependent translation of mRNA. We are now beginning to appreciate these modifications which may be regulated by 14-3-3s, and is the family member known to inhibit the kinase activity of Akt. Other 14-3-3 isoforms also actively participate in this Akt signaling pathway to regulate DNA damage pathway, albeit in a positive way. It is important to point out that 14-3-3s has unique amino acids (Met202, Asp204, and His206) that may be responsible for binding to its particular set of binding targets that are not recognized by other 14-3-3 family members [95, 96], thus making 14-3-3s function differently from other isoforms in response to DNA damage. We expect that more 14-3-3-mediated regulatory mechanisms in DNA checkpoint response will emerge because 14-3-3 has several binding targets involved in DNA damage response [5] [6]. Given that DNA damage response plays an important role in tumorigenesis, understanding the role of 14-3-3 in DNA damage checkpoint will facilitate the discovery of therapeutic strategies for cancer treatment.


Acknowledgments: We would like to thank the Susan Komen Breast Cancer Foundation, NIH grant (RO1CA 089266 to Lee MH), Directed Medical Research Programs (DOD SIDA BC062166 to Yeung S J & Lee M H) and Cancer Center Core Grant (CA16672) for their research support. We apologize to our many colleagues whose work we were unable to cite due to space constraints.

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Part II

Checkpoint Response and the Aetiology of Cancer

Chapter 6

Chromatin Modifications and Orchestration of Checkpoint Response in Cancer Makoto Nakanishi

Abstract Cell cycle checkpoints are important surveillance systems to maintain genomic integrity. Once checkpoint systems sense the abnormal chromosomal DNA structures such as DNA damage and stalled DNA replication, which pose a great threat to genome stability, they execute cell cycle arrest through inhibiting the activity of cell cycle regulators and coordinating it with the DNA repair process. Thus, aberrant checkpoint responses probably result in loss or amplification of chromosome activity, which may result in transformation of normal cells into malignant ones. Checkpoint responses also execute apoptosis and cellular senescence when cells sense unrepairble and extensive chromosomal abnormalities. Numerous key players have been identified in terms of checkpoint sensor proteins, transducer kinases and effectors, but the coordination, interconnectedness, and mechanisms by which they execute important anti-tumor protective responses such as apoptosis or senescence have only recently become evident. In this chapter, changes in chromatin modification and the molecular mechanisms of checkpoint activation in response to abnormal chromatin structure as well as the regulation of checkpoint kinases and effectors are discussed. These checkpoint responses are important for determining the potential effects of current cancer therapies in terms of toxicity and efficacy. Keywords Chk1 • Chk2 • DNA damage • DNA replication • p53

6.1 Changes in Chromatin Modification at Abnormal Chromosomal Sites Checkpoint response is mediated through a series of events involving sensors, mediators, transducers, and effectors that activate cell cycle arrest, DNA repair, apoptosis, and senescence (Fig. 6.1). Chromatin modification is an important early event. Normally, eukaryotic genomic DNA is packaged with histone and nonhistone proteins into highly condensed chromatin structures. Hence, efficient repair of DNA damage should require modification and remodeling to render the chromatin structures more accessible to DNA repair enzyme and checkpoint signaling proteins. Nucleosomes constitute the fundamental unit of chromatin. This supramolecular assembly includes a nucleosomal core and a variable DNA linker. The nucleosomal core particle comprises about 146 bp of DNA wrapped around an octameric complex containing two copies each of histones H2A, H2B, H3, and H4. Alteration of the chromatin structure can be achieved by covalent modification of histone tails or by altering histone composition, as listed in Table 6.1. Most wellcharacterized histone modification at DNA damage sites is phosphorylation of H2AX at Ser139 by ATM, ATR, and DNA-PK [1–3]. This phosphorylation (g-H2AX) covers a region that extends up to megabases away from the break site [4] and can be


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M. Nakanishi DNA damage Sensors recognize DNA damage Chromatin modifications at damage sites Recruitment of mediators at damage sites Activation of transducer kinases Regulation of effector activity

Activation of DNA repair

Cell cycle arrest Apoptosis

Senescence

Figure 6.1. Conceptual organization of the signal transduction of checkpoint responses. DNA damage is recognized by sensor proteins and induce chromatin modifications at damage sites that can function as a platform for mediator proteins and DNA repair proteins. These signals are transmitted to transducers (mainly kinases) and the transducer molecules regulate effectors, thereby arresting the cell cycle, inducing apoptosis or senescence, and activating DNA repair mechanisms.

a platform for protein complexes required for transmitting the signals to downstream mediator kinases [5]. These complexes involve MDC1 [6–8], 53BP1, BRCA1, and the MRN complex (Mre11, Rad50, Nbs1) [9]. In yeast, a platform of g-H2AX surrounding the damage sites is also required for the recruitment of chromosomal modification and remodeling factors, NuA4 histone acetyltransferase and INO80SWR1 remodeling complexes that probably unravel the packed chromatin, allowing repair enzymes to access the DNA [10–13]. The Tip60-HAT complex is also required for DNA damage repair [14]. Tip60 phosphorylates histone H4 at multiple sites and these phosphorylation are required for the recruitment of DNA repair proteins [15]. Tip60-dependent histone acetylation is also essential for exchange of g-H2AX with unphosphorylated H2AX in mammals for the termination of damage signaling [16]. Histone methylation has played an important role in recruitment of protein complexes required for damage signaling. For example, methylation of histone H3 at K79 by Dot1L is indispensable for the recruitment of 53BP1 at damage sites [17]. Collectively, histone modification and remodeling result in changes in the higher order chromatin structure and recruitment of protein complexes required for DNA repair and transmission of signaling to transducer kinases.

Table 6.1. Histone Modifications at DNA Damage Sites. DNA damage induces several types of histone modifications. These modifications are mediated by distinct enzymes and have unique functions as shown. Modifications

Enzyme

Functions

H2AX Phosphorylation

ATM, ATR, DNA-PK

H2AX Ubiquitination

RNF8, UBC13

H4 acetylation

Tip60

H3-K79 methylation H3-K20 methylation

Dot1

Recruitment of 53BP1, BRCA1, Nbs1, MDC1 Recruitment of BRCA1-BARD1CCDC98-RAP80 complex Recruitment of DNA repair enzyme; gH2AX exchange Recruitment of 53BP1

Set9

Chk1 activation

6.2 Recognition of Abnormal Chromatin by Sensor Proteins Abnormal chromosomal structures, including DNA damage and stalled replication forks are effectively recognized by the so called sensor

6. Chromatin Modifications and Orchestration of Checkpoint Response in Cancer

proteins. For example, studies in yeasts and mammals have demonstrated that Rad9, Rad1, Hus, and Rad17 are essential factors that activate checkpoint signaling in response to various types of DNA damage [18]. Rad9, Rad1, and Hus1 form a heterotrimeric complex (the 9-1-1 complex) whose structure resembles a PCNA-like sliding clamp [19]. Rad17 forms an RFC-related complex with four small RFC subunits, Rfc2, Rfc3, Rfc4, and Rfc5 that acts as a clamp-loading complex [20]. The Rad17/Rfc2-5 complex is recruited to ssDNA where it is loaded in an RPA-dependent manner [21]. The presence of a dsDNA–ssDNA junction, as might be found at a stalled replication fork, activates this complex to load a second complex, the PCNA-related 9-1-1 clamp (Rad9–Rad1–Hus1). The chromatin-bound 9-1-1 complex then facilitates phosphorylation mediated by ATR and ATM. Chromosomal double-strand breaks (DSBs) are highly toxic DNA lesions that reduce cell survival. DSBs are first recognized by the multifunctional sensor complex, MRN, that consists of the structure specific nuclease Mre11, the ATPase and adenylate kinase subunit Rad50, and the adapotor protein NBS1 (Nijmegen breakage syndrome 1) [22,23]. MRN complexes can bind to the exposed DNA ends and unwind them in an ATP-dependent manner [24,25]. MRN complexes recruit and activate ATM [26]. An activated ATM then catalyzes the phosphorylation of H2AX (gH2AX) [27,28], which subsequently binds MDC1 through its BRCT domain. ATM also phosphorylates a BRCA1-BARD1-CCDC98RAP8 complex [29,30]. gH2AX bound MDC1 is phosphorylated by ATM and subsequently functions as a scaffold for the recruitment and assembly of DNA damage response mediators and effectors [31]. For example, phosphorylated MDC1 is recognized by the FHA daomain of RNF8 that subsequently forms a complex to UBC13 [31–34]. This ubiquitin ligase complex catalyzes K63-linked polyubiquitination of H2A and gH2AX and the resultant polyubiquitin chains on H2A or H2AX bind to RAP80, thereby facilitating the recruitment of the BRCA1-BARD1CCDC98-RAP8 complex to DSB sites. The recruitment of these factors and complexes is prerequisite for proper DNA repair and checkpoint signaling [35].

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6.3 Activation of Transducer Kinases In mammals, once sensor complexes recognize DNA damage, ATM and ATR kinases, which are both extremely large proteins that phosphorylate a great number of substrates, are rapidly activated. Patients bearing an ATM mutation suffer from a devastating syndrome called ataxia telangiectasia that causes immunodeficiency, genome instability, clinical radio-sensitivity and a predisposition to cancer [36]. Although cells lacking ATM are viable, suggesting that ATM is a nonessential gene for normal cell cycle progression and development [37], its kinase activity is strongly stimulated by DSBs. The identification of a damage-induced phosphorylation site (Ser1981) revealed a new mechanism for ATM regulation by which a rapid and sensitive switch for checkpoint signals is permitted [38]. ATM under unperturbed condition exists as a homodimer complex in which its kinase active site is physically blocked by tight intermolecular binding to a protein domain around Ser1981. In response to DSBs, a conformational change in the ATM protein stimulates it to autophosphorylate Ser1981 in an intermolecular manner. In contrast, the auto-phosphorylation of ATM at Ser1981 is suppressed under unperturbed condition by the constitutive interaction with PP2A that dephosphorylates Ser1981 [39]. Acetylation of ATM by Tip60 was also reported to be important for its full activation [40]. ATR was discovered from its sequence similarity to ATM and Rad3 [41], and was shown to play an essential role in DNA damage and DNA replication checkpoint activation [42,43]. Mutations in ATR gene have been reported in a subset of patients with Seckel syndrome [44], which is a human autosomal recessive disorder causing severe intrauterine growth retardation, proportionate dwarfism, microcephaly, with skeletal and brain abnormalities, and cancer predisposition. ATR constitutively forms a heterodimer with ATRIP that binds to UV-damaged DNA or to RPAcoated single-stranded DNA [45]. In addition to ATRIP, in response to DNA damage, TopBP1, a mediator protein containing eight BRCT phospho-recognition motifs, binds and activates ATR/ ATRIP complexes in a manner distinct from the

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role of TopBP1 in the initiation of DNA replication [46,47]. TopBP1 binds the constitutively phosphorylated C-terminal tail of Rad9 on the 9-1-1 complex, at damage sites and this binding occurs via its first pair of BRCT repeats on TopBP1. Therefore, although the precise physical mechanisms by which the interactions between TopBP1 and ATR–ATRIP elicit increased ATR activity remain to be determined, TopBP1 appears to be implicated in early events of signaling following recruitment of ATR– ATRIP to sites of DNA damage and replication stress. For the downstream targets, ATR is capable of specifically phosphorylating serine or threonine residues in SQ/TQ sequences as with ATM, sharing common downstream substrates such as p53 [48–51] and BRCA1 [52,53], although they primarily respond to different stimuli [22].

6.4 Regulation of Checkpoint Kinases The checkpoint kinases Chk1 and Chk2 were first identified in fission yeast as essential for cell cycle arrest before mitosis, in response to DNA damage or DNA replication blockage, respectively. These kinases were also identified in vertebrate cells based on their homology with fission yeast Chk1 and Cds1. Chk1 is phosphorylated at Ser317 and Ser345 by ATR in response to DNA damage or DNA replication stress. This phosphorylation is blocked in cells lacking the kinase ATR [54] and markedly inhibited in cells with a reduced amount of Rad17 [55] or lacking Hus1 [56]. Chk1 is a constitutively active enzyme and the ATR/ATMdependent phosphorylation appears not to regulate its kinase activity but rather its subcellular localization [57,58]. For example, following ATR/ATMdependent phosphorylation, Chk1 is targeted to centrosomes [58,59], where cyclin B1-cdk1 is first activated at the onset of mitosis [60]. In undamaged cells, a significant proportion of Chk1 is chromatin associated and ATR/ATM-dependent Chk1 phosphorylation following DNA damage results in rapid Chk1 dissociation from chromatin. One of the major downstream targets of Chk1 is a family of Cdc25 phosphatases [61,62]. Cdc25 phosphatases catalyze dephosphorylation of the inhibitory phosphorylation of cdk1 and cdk2 at

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T14 and Y15 [63], and thus activate their kinase activity. Studies in yeasts, Xenopus, and mammals have demonstrated that phosphorylation of these Cdc25 proteins by Chk1 creates binding sites for 14-3-3 proteins and downregulates their phosphatase activities [61]. In addition to 14-3-3 binding, in the presence of DNA damage during S phase progression, activated ATR-Chk1 phosphorylates Cdc25A triggering its ubiquitination and degradation [64]. The downregulated Cdc25A suppresses cdk2 and cdk1 activation that blocks the loading of Cdc45 [65,66], a protein required for the initiation of DNA replication through the recruitment of DNA polymerase alpha into prereplication complexes, onto chromatin. Analyses using mice deficient in Chk1 revealed its essential role in early embryonic development [54,67]. Chk1-deficient ES cells demonstrated that Chk1 is prerequisite for cell cycle arrest before mitosis in response to DNA damage or DNA replication block as described above. In addition to its involvement in checkpoints, Chk1, like ATR [42], plays a role at every point in the cell cycle and loss of Chk1 results in the premature onset of mitosis through the dephosphorylation of cdk1 at Tyr15. Premature mitosis leads to the activation of caspases 3 and 9 triggered by cytoplasmic release of cytochrome c and the subsequent mitotic catastrophes [68]. Furthermore, Chk1 plays an important role in transcriptional regulation. Chromatin-bound Chk1 phosphorylates histone H3-threonine 11 (H3-T11) [69]. Phosphorylation of H3-T11 significantly enhances the binding affinity between GCN5 histone acetyltransferase and histone tails. Thus, changes in the phosphorylation status of H3-T11 in response to DNA damage probably influence GCN5 recruitment at promoters and thus transcription of GCN5-dependent genes. GCN5 is utilized as an accessory acetyltransferase for E2Fs [70], which regulate the expression of the many genes involved in DNA replication [71]. In addition, GCN5 is recruited to the promoters of many cell cycle genes [72] and GCN5-deficient cells have been shown to exhibit a significant decrease in their growth capability which is associated with a reduction in the expression of many cell cycle regulatory genes [73]. Therefore, Chk1-dependent repression of GCN5-dependent gene expression serves as an alternative checkpoint mechanism to


promote cell cycle delay or arrest, besides regulation of the inhibitory Y15 phosphorylation of cdk1 and cdk2. In contrast to Chk1, Chk2 is dispensable for normal cell survival, cell proliferation, and prenatal development [74,75]. Interestingly, heterozygous germline mutations in the human Chk2 gene are found in a subset of patients with Li-Fraumeni syndrome, a familial cancer syndrome, without any mutation in their p53 gene, suggesting its role as a bona fide tumor suppressor [76]. Chk2 is mainly activated by phosphorylation of its Thr68 in an ATM-dependent manner in response to DSBs [77]. Although biochemical analyses revealed that activated Chk2 is capable of phosphorylating Cdc25A at Ser123, Cdc25C at Ser216, BRCA1 at Ser988 and p53 at several sites, including Ser20 [78], examination of Chk2-deficient mice and cells showed that this enzyme functions mainly in p53dependent apoptosis and DNA repair [75], but not cell cycle arrest in response to DNA damage. Chk2-deficient mice are resistant to IR as a result of the preservation of splenic lymphocytes, thymocytes, neurons of the developing brain apoptosis of which is known to be p53 dependent [75,79]. Although ATM-Chk2 signal at the intra-S phase checkpoint in response to IR is considered to be important because phosphorylation of Cdc25A by Chk2 triggers its ubiquitination and degradation in vitro [80], Chk2-deficient cells showed that it is dispensable for the intra-S phase checkpoint response. Thus, the function of Chk2 in cell cycle arrest upon DNA damage remains questionable. In addition to Chk1 and Chk2, Plk1 also functions in cell cycle checkpoints as a checkpoint kinase. Plk(s) are serine/threonine kinases possessing one or two polo-box domains. Plk1 is highly expressed at G2-M phase and is degraded at the end of mitosis [81–83]. Although Plk1 is expressed in G2, its kinase activity is first detected at the G2/M transition and reaches its maximum during mitosis [84]. Several lines of evidence demonstrate that Plk1 is implicated in the regulation of different cell cycle processes, including mitotic entry, spindle formation, centrosomal duplication, and cytokinesis [85]. Plk1 appears to regulate cyclin B-cdk1 at several levels. In Xenopus and human system, Plk1 directly phosphorylates and activate Cdc25C and consists a part of the MPF amplification loop [86–89]. However, Plk1-depleted cells enter mitosis

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with active cyclin B-cdk1 but incompletely phosphorylated cdc25C [90] and cdc25C-depleted cells grow normally [91]. Therefore, physiological significance of cdc25C phosphorylation by Plk1 still remains to be determined. Plk1 also regulates cdk1 activation through direct phosphorylation of Wee1 [90]. In response to a variety of DNA damage, Plk1 appears to be catalytically inactivated and this inhibition is dependent on functional ATM or ATR [88]. Hence, ectopic expression of constitutively active Plk1 mutants can result in impaired G2 arrest in response to DNA damage. Intriguingly, Plk1 is reported to be overexpressed in a variety of human tumors, including prostate cancer, ovarian cancer, hepatoblastomas, and melanomas, and its overexpression coincides with bad prognosis [92–95]. Therefore, Plk1 dysfunction is likely to induce malignant transformation into normal cells.

6.5 Effector Molecules 6.5.1 p53 The p53 tumor suppressor protein is one of the ultimate targets of cell cycle checkpoints. The p53 protein plays a central role in the decision of a cell to undergo either cell cycle arrest, apoptosis, or cellular senescence after diverse stresses, including DNA damage, hypoxia, and the activation of oncogenes [96–98]. The p53 is probably the most frequently mutated and extensively studied tumor suppressor protein that acts as a highly regulated sequence-specific DNA binding transcription factor for many target genes. The amount and transcriptional activity of p53 is regulated by posttranscriptional modification, such as phosphorylation, sumorylation, neddation, and acetylation [99]. In normal cells, p53 protein level is under precise control by the protein Mdm2 which acts as an E3 ligase and targets p53 for ubiquitin-dependent degradation, acting as a critical negative regulator [100]. This is clinically important as Mdm2 level is amplified in at least 10–20% of human cancers. In addition, the importance of Mdm2 as negative regulators of p53 has been established both biochemically and genetically. In response to DNA damage, p53 is phosphorylated at several sites in its transactivation domain, including Ser15 and Ser20 [101]. ATM and ATR phosphorylate p53 at

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Ser15 [48,49], which inhibits the interaction of p53 with Mdm2, resulting in p53 stabilization [102]. Mdm2 phosphorylation by ATM also reduces its capability to promote nucleo-cytoplasmic shuttling and the subsequent degradation of p53 [103]. Phosphorylation of p53 at Ser20 (murine Ser23) by Chk1 and Chk2 also appears to play an important role in its activity. Mice with p53 point mutation at Ser23 die between 1 and 2 years with tumors that were most commonly of B cell lineage [104]. Although MEFs with p53 mutation at Ser23 do not show dramatic reduction in IR-induced p53 stabilization or p53-dependent cell cycle arrest, thymocytes and developing cerebellum with the same mutation demonstrate a significant reduction in IR-induced p53 stabilization and a resistance to apoptosis. Acetylation of p53 is critical for the function of p53 as a transcriptional regulator [105]. The unacetylated p53 with mutations in all major acetylation sites retains its ability to induce p53Mdm2 feedback loop, but fails to induce cell cycle arrest and apoptosis in response to DNA damage. Acetylation of p53 abrogates Mdm2-mediated repression by docking the recruitment of Mdm2 to p53-responsive promoters, which results in p53 activation in a phosphorylation independent manner. Furthermore, acetylation of p53 at K120 by Tip60 is a prerequisite for p53-dependent apoptosis but is dispensable for its mediated growth arrest upon DNA damage [106,107], suggesting its role in the decision between cell cycle arrest and apoptosis. Particularly, given that K120 is a recurrent site for p53 mutation in human cancer, Tip60dependent acetylation of p53 upon DNA damage may protect tumorigenesis in vivo. p53 is thought to be essential for G1 arrest in response to DNA damage. The key transcriptional target of p53 is the p21 Cdk inhibitor (p21 CKI) [108], which inhibits cyclin E-cdk2 activity, thereby inhibiting G1/S transition [109]. p21 CKI also binds to the cyclin D-cdk4 or cyclin D-cdk6 complexes and prevents them from phosphorylating Rb, thereby suppressing the Rb/E2F pathway. Thus, the G1 checkpoint signal targets two independent and critical tumor suppressor pathways that are most commonly deregulated in human cancers. However, p21-deficient cells are apparently normal at the beginning of G1 arrest although the maintenance of G1 arrest appears to be impaired

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[110], suggesting that an alternative target should exist downstream of p53. In this regard, a rapid decrease, independent of p53 status, in the abundance and activity of Cdc25A in response to DNA damage was reported to play an important role in the initiation of G1 arrest [64,80]. This reduction in Cdc25A is triggered by phosphorylation of its Ser123 residue by Chk1 or Chk2. Furthermore, cyclin D1 degradation has also been reported to be essential for initiation of DNA damage induced G1 arrest [111]. The reduced cyclin D1 protein then decreases the amount of cyclin D1-cdk4 or cyclin D1-cdk6 complex, resulting in the redistribution of p21 CKI to cyclin E-cdk2 complex and the inhibition of its activity. However, this checkpoint response is independent of ATM-p53 and triple knockout mice lacking cyclin D1, D2, and D3 show apparently normal cell cycle control without significant defects in their checkpoint response [112]. Therefore, the main pathway by which p53 regulates the initiation of G1 arrest in response to DNA damage remains elusive. p53 also regulates other tumor protective pathways such as apoptosis and senescence. Activation of the p53 pathway lead to apoptosis in certain cell types, notably cells of hematopoietic origin [113]. Stimuli such as DNA damage, withdrawal of growth factors, and ectopic expression of the nuclear oncogenes stimulate p53-dependent apoptosis. Several mechanisms have been implicated in p53 mediated apoptosis. For example, p53 activation leads to up-regulation of proapoptotic Bax, PUMA, and Noxa, and down-regulation of prosurvival Bcl-2. Functional p53 is also indispensable for premature senescence induced by nuclear oncogenes and DNA damage [114] although the precise mechanisms how p53 regulates their pathways are largely unknown.

6.5.2 Cdc25 Family of Phosphatases In mammals, three dual specific phosphatases, Cdc25A, Cdc25B, and Cdc25C, dephosphorylate the Cdks that act on kinases directly to regulate cell cycle transitions [63]. Cdk activation depends not only on the association with a cyclin subunit but also on proper phosphorylation of the catalytic subunit [115]. Cdk-activating kinase mediates phosphorylation of a threonine residue in the T-loop that is required for full activatin of Cdks. In contrast, phosphorylation of the tyrosine


and threonine residues located in the N-terminus ATP binding domain inhibits the Cdk activity. This inhibitory phosphorylation is catalyzed by the Wee1 and Myt1 kinases, and subsequently dephosphorylated by the Cdc25 phosphatases. Cdc25 phosphatases are critical for timely Cdk activation in cell cycle progression. Although the expression patterns of the three Cdc25 proteins are mostly overlapping, Cdc25C was initially thought to be the major effector of the G2/M DNA damage checkpoint response. However, recent reports have revealed that both Cdc25C-deficeint [91] and Cdc25B-deficient [116] cells have a normal G2/M checkpoint, suggesting that Cdc25A is also the main effector at G2/M checkpoints. In response to DNA damage, Chk1 phosphorylates Cdc25A on multiple sites including Ser76, Thr124, and Ser178. While Ser178 phosphorylation creates a binding site for 14-3-3 protein, Ser76 phosphorylation promotes subsequent phosphorylation of Ser82 and Ser88 by undefined kinases [117,118]. Phosphorylation of Ser82 and Ser88 facilitates the association with the SCF ubiquitin ligase complex. Hence, the Chk1-dependent priming phosphorylation of Cdc25A initiates the multistep process of ubiquitin-mediated proteasomal degradation. Cdc25B and Cdc25C are also substrates of Chk1 and Chk2 although the significance of Cdc25B phosphorylation remains unclear. The cell cycle promoting action of Cdc25s and their role as p53-independent checkpoint targets suggest that the expression level of these phosphatases must be tightly regulated in normal cells and dysregulation of these activities easily result in impaired checkpoints and genomic instability, a hallmark of cancer. Indeed, Cdc25A overexpression has been reported in a variety of human cancers, including breast, liver, esophageal, endometrial, and colorectal cancers [119].

6.6 Conclusion The coordinated activation of cell cycle checkpoints, DNA repair, and apoptosis are essential for the maintenance of genome integrity and tumor suppression. Given that mutations or decreased expression of the genes implicated in checkpoint control are detected in the most of cancers, proper checkpoint signaling is essential for preventing

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cancer. Since abrogation of the G2 checkpoint might be more detrimental in cancer cells lacking p53 than in normal cells, new anticancer drugs targeting the G2 checkpoint inhibitor appear to be important for the development of therapeutic strategies with fewer side effects. An approach that combines conventional anticancer treatments such as radiation and chemotherapy with the use of new small molecule inhibitors for checkpoint regulators should prove to be effective in eliminating cancer cells.

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Chapter 7

DNA Damage Response and the Balance Between Cell Survival and Cell Death Bernd Kaina, Wynand P. Roos, and Markus Christmann

Abstract DNA damage induces the activation of a cascade of kinases that trigger the DNA damage response (DDR). Downstream are targets that either help cells to survive or undergo cell death. DNA damage-induced cell death is executed by apoptosis, necrosis, mitotic catastrophe, and autophagy. Of these different forms of cell inactivation, apoptosis is often the main route of cell death following DNA damage. Cells undergo apoptosis upon genotoxic stress via the death receptor and/or the intrinsic mitochondrial damage pathway, with p53 and AP-1 involved decisively. Not every type of DNA damage induces apoptosis. Many DNA lesions are tolerated by the cell, some are mutagenic without being toxic and some are more toxic than mutagenic. Severe DNA damages are O6-alkylguanines, bulky lesions, and DNA double-strand breaks, that activate DDR and downstream survival and death signals. The survival and death pathways triggered by upstream DDR functions will be discussed in this chapter. Keywords Checkpoints • Cell cycle • DNA damage response • DNA repair • p53 • Apoptosis • Death receptors • Fas • ATM • ATR • Anticancer drugs

7.1 DNA Damage Triggers Signaling that Stimulates DNA Repair and Apoptosis DNA damaging agents, such as ultraviolet light, oxidants and alkylating agents, are able to activate immediate-early signaling cascades by stimulation

of membrane bound receptors. A paradigmatic example is the EGF receptor, which is phosphorylated upon genotoxic treatment. This is due to the inhibition of receptor tyrosine dephosphorylation [1] caused by blocking the activity of receptor-directed tyrosine phosphatases [2]. For death receptors (Fas/CD95/Apo1) it has also been shown that activation can occur in a direct way, i.e. without the mediation of DNA damage [3–5]. Therefore, for quite a long time, it was a matter of debate whether cell death induced by genotoxic agents requires DNA damage or is solely dependent on direct receptor activation. There are three lines of evidence that cell death executed by apoptosis, which is induced by genotoxic agents, is caused by DNA damage: a) Inability of cells to repair DNA lesions results in hypersensitivity to the killing effect (an exception to the rule is DNA mismatch repair, which is required for O6methylguanine triggered cell death), which has been shown for mutants defective in O6-methylguanineDNA methyltransferase (MGMT), base excision repair (BER), nucleotide excision repair (NER), DNA double-strand break (DSB) repair and DNA crosslink repair; b) Modified nucleotides such as 6-thioguanine or gancyclovir incorporated in the DNA induce apoptosis; and c) DSBs induced in the cellular genome by restriction enzymes induce a strong apoptotic response. Apoptosis in these situations is rarely associated with necrosis [6]. The apoptotic pathways activated by specific DNA lesions such as O6-methylguanine (O6MeG) and cyclobutane pyrimidine dimers (CPDs), as well as its interplay with DNA repair pathways has


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been studied in great detail (for recent reviews see [7,8]). Thus it has been shown that O6MeG does not block DNA replication. It mispairs with thymine, giving rise to mismatch repair substrates that are recognized by MutSa (MSH2 and MSH6) and MutLa (MLH1 and PMS2), which provoke a futile mismatch repair cycle leading to the formation of DSBs and apoptosis signaling [7]. CPDs provide a representative example of bulky DNA lesions that block DNA replication and transcription. They were shown to trigger apoptosis either by forming DSBs or blocking the transcription of antiapoptotic genes [9]. A critical down-regulated gene is MAP kinase phosphatase 1 (MKP1) that has been shown to be blocked in expression upon UV-C light, which via dephosphorylation of Jun kinase (JNK) leads to downregulation of the MAP kinase pathway, resulting in low AP-1 levels that has an impact on the regulation of anti-apoptotic (e.g. repair genes)

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7.2 The DNA Damage Response: The Mrn/Atm/Atr Connection DSBs and other replication blocking lesions, notably DNA interstrand crosslinks, are lethal DNA lesions and, therefore, cells have to be equipped with sensors that recognize these lesions immediately upon formation. DNA damage recognition by these sensors starts a protein cascade, which finally results in cell cycle arrest, thereby giving the cell time for repair. The most important sensors are the phosphatidyl inositol 3-kinase-like kinases, ataxia-telangiectasia mutated (ATM) and ATM- and Rad3-related (ATR). The activation of ATM and ATR is highly complex

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or proapoptotic (e.g. Fas ligand) genes [10]. The network of receptor and DNA damage-triggered early responses are outlined in Fig. 7.1.

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Figure 7.1. Signaling pathways induced by receptor activation and DNA damage. Molecular signals in the cytoplasm and the nucleus cooperate to orchestrate DNA repair and the final fate of the cell by impacting the balance between pro and anti-apoptotic signals.

7. DNA Damage Response and the Balance Between Cell Survival and Cell Death

and dependent on additional proteins. While ATM is mainly activated by ionizing radiation-induced DSBs, ATR is activated in response to UV light and presumably all chemical agents that give rise to stalled DNA replication forks. ATM/ATR is implicated in three crucial functions: regulation and stimulation of DSB repair (homologous recombination), signaling cell cycle checkpoints, and signaling apoptosis via p53 (Fig. 7.2). ATM activation is dependent on autophosphorylation and on a functional MRN complex consisting of Mre11, NBN (alias Nbs1), and Rad50. Autophosphorylation mainly occurs at Ser1981 [11], which leads to dissociation of inactive ATM dimers into catalytic active monomers. Additional autophosphorylation on Ser367 and pSer1893 has also been detected in vitro and in vivo upon ionizing radiation. Mutations in each autophosphorylation sites detected so far (S367A, S1893A and S1981A) abrogate ATM signaling upon exposure to ionizing radiation [12]. However, in mouse, mutation of the autophosphorylation sites does not influence ATM activity [13] and in vitro activation of ATM protein kinase activity is not strictly dependent on autophosphorylation on S1981 [14]. ATM monomerization was also shown to occur by the interaction of ATM with the MRN complex and single stranded DNA [15]. As noted above, ATM activation requires the MRN complex, which recognizes and migrates to DSBs upon ionizing radiation [16] and retains at the site of DNA damage in small granular foci. In the absence of the MRN complex, the catalytic activity of ATM and Ser 1981 phosphorylation is reduced [17] and ATM is not recruited to the DSB [15]. The exact mechanism of how the MRN complex regulates ATM activation is still unclear. Whereas ATM can interact with the MRN complex independent of Nbs1 [18], Nbs1 is required for nuclear translocation of MRN [19], for ATM autophosphorylation and the recruitment of ATM to DSBs [20] resulting in increased ATM activation [21]. In vitro, unwinding of DNA ends by the MRN complex, but not ATM autophosphorylation, was required for monomerization of ATM and ATM activation [15]. The presence and absence of MRN differentially affects the activation of several ATM targets [22] and can be used to explain the mechanism of ATM activation. In this model, ATM is autophosphorylated via signals generated by structural changes of the chromatin, resulting in the release of preactived ATM

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monomers. Parallel to ATM autophosphorylation, H2AX is phosphorylated and the MRN complex as well as BRCA1 and SMC1 are recruited to DNA double-strand breaks independently. Preactivated ATM is now recruited onto the DSB via NBS1 and BRCA1. ATM can then interact and phosphorylate NBS1, BRCA1, and SMC1. If ATM is activated in the absence of DSBs or not recruited to the DSB, only the nucleoplasmic substrates (p53, CHK2) can be phosphorylated. Besides the MRN complex, additional co-factors seem to be involved in determining the substrate-specific effects of ATM. Thus, loss of p53 binding protein 1 (p53BP1) decreases ATM autophosphorylation and ATM mediated phosphorylation of Chk2 and SMC1 [23,24]. Activation of ATR is mediated via blockage of the DNA polymerases and large stretches of singlestranded DNA (Fig. 7.2), which are created upon genotoxic stress and covered by RPA (replication protein A) [25]. RPA labeled ssDNA induces the recruitment of ATR in the complex with ATRIP (ATR-interacting protein) and the 9–1–1 complex consisting of Rad9, Hus1, and Rad1 (see review [26]), which cooperate in the activation of downstream targets. The ATRIP–ATR complex is bound to DNA via direct interaction between ATRIP and RPA [27–30] and ATRIP oligomerization [31], whereas the loading of the 9–1–1 complex is a multi step process. Rad9 interacts with TopBP1 [32] and recruits TopBP1 to the stalled replication fork [33,34]. Upon replication stress TopBP1 directly activates the ATR–ATRIP complex, most likely via a conformational change of the ATR– ATRIP complex [35]. ATM and ATR do not work totally independently. Thus, upon the formation of DSBs, ATR chromatin loading depends on ATM [36] since phosphorylation of TopBP1 by ATM is necessary for ATR activation [37]. If the replication fork has stalled. e.g. following UV-C treatment, ATM is phosphorylated and activated at the autophosphorylation site Ser1981 by ATR [38].

7.3 The DNA Damage Response: Cell Cycle Arrest The most important target proteins for ATM and ATR are CHK1, CHK2, and p53. ATM phosphorylates the checkpoint kinase-2 (CHK2) after the formation of DSBs at Thr68 [39,40], whereas ATR

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Figure 7.2. DNA damage responses upstream and downstream of ATM and ATR. A major outcome of DNA damage signaling is checkpoint response through inhibition of CDK/cyclin activity, which can occur in a p53-dependent andindependent manner, and this induces cell cycle arrest to allows DNA repair to proceed.


phosphorylates CHK1 at Ser345 after DNA replication blockade [41,42]. In turn, CHK2 and CHK1 phosphorylate p53 at Ser20, whereby it becomes activated [43]. Phosphorylation of p53 by these checkpoint kinases masks the MDM2 binding site thus leading to a release of p53 from MDM2. Since MDM2 normally targets p53 for ubiquitin-dependent degradation, the dissociation of the MDM2-p53 complex results in stabilization of the p53 protein [44–47]. ATM and ATR can also directly phosphorylate p53 at Ser15, thereby increasing its transactivation activity [48,49], and the transactivated MDM2 provoking its down-regulation by negative feedback [50]. As a consequence, the nuclear translocation of p53 and its DNA binding activity to target genes for transcriptional activation are tightly regulated. One of the p53 target genes is p21, whose upregulation inhibits the Cdk2–cyclin E–PCNA complex, resulting in G1/S blockage. Additionally, p53 independent cell cycle arrest can be induced by CHK1 via the phosphorylation of Cdc25a and Cdc25c. Phosphorylation of Cdc25a at Ser123 leads to ubiquitination and degradation of Cdc25a [51], which can no longer activate the Cdk2–cyclin E complex by dephosphorylation of CDK2 at Thr14 and Tyr15, thereby inducing a G1/S arrest. Phosphorylation of Cdc25c at Ser216 induces binding of Cdc25c to the 14-3-3s protein [52,53]. Within this complex, Cdc25c is transported out of the nucleus and is therefore unable to dephosphorylate/activate Cdk1-CyclinB, which finally results in G2/M arrest [54] (lower part of Fig. 7.2). Besides arresting at the G1/S and the G2/M checkpoint, cells can also be blocked during replication. Intra-S damage leads to stabilization of stalled replication forks and prevents firing of late and dormant origins (for review see [55]). The regulation of this intra-S-arrest is only marginally characterized, however, also in this process, ATM and ATR appear to play the most important role. In yeast, the ATR homolog Mec1 and the protein Mrc1 (Mediator of the Replication Checkpoint) are involved in the intra-S phase arrest induction. Mrc1 is required for the activation of the CHK2 homolog Rad53 [56,57] and in Xenopus, the Mrc1 related protein Claspin mediates Chk1 activation in response to replication blockage [58]. Besides this, Mrc1 is a replication fork component, which is hyperphosphorylated upon DNA damage by Mec1 [56,59].

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A recent study showed that Mrc1 interacts with the Pol2 amino and the carboxy terminus throughout the cell cycle. Upon hyperphosphorylation, the interaction between Mrc1 and the amino terminus of Pol2 is abolished [60]. In addition, Mrc1 interacts with MCM2, which might enhance interaction between Pol2 and the replication helicase. This leads to the speculation that upon phosphorylation of Mrc1 the coupling between polymerases and helicases is abrogated, thus arresting cells within the S-phase. This is substantiated by the fact that DNA synthesis and unwinding is uncoupled in Mrc1-null strains in the presence of replication stress [61]. An unexpected role in the intra-S phase arrest is exerted by the proapoptotic protein Bid. Upon the induction of DSBs by the topoisomerase II inhibitor etoposide, Bid is phosphorylated by ATM and translocates into the nucleus. Here it plays a yet unknown function in the induction of the intra-Sphase arrest, which is supported by the fact that BID cells failed to accumulate in the S-phase upon etoposide treatment [62,63].

7.4 DNA Damage Response and Apoptosis Pathways The main apoptosis players activated by DNA damage are outlined in Fig. 7.3. Three pathways are evident. One is involved in the upregulation of p53/p73, which in turn upregulates the fas receptor/caspase-8 dependent apoptotic pathway. In p53-inactive cells, DNA damage activates the mitochondrial (also called endogenous) apoptotic pathway. A hallmark of mitochondrial damagemediated apoptosis is decline of Bcl-2 protein level that leads to leakiness of mitochondria, cytochrome c release and activation of the apoptosome – a complex consisting of Apaf-1, ATP, procaspase-9, and cytochrome c. Caspase-9 in turn cleaves caspase-3 (and other downstream caspases such as caspase-7), which degrades the inhibitor of caspase-activated DNase (ICAD), and this enables CAD to cleave DNA into the typical nucleosomal fragments. There is crosstalk between both pathways: p53 upregulates the genes encoding Noxa and Puma, and caspase-8 cleaves Bid, which releases Bax and Bak. These proapoptotic players target the mitochondrial membrane, thus enhancing

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Apoptosis Figure 7.3. Main pathways of DNA damage triggered apoptosis. The pathways involve p53/p73-dependent and -independent Fas/Caspase-8 signaling, and p53/p73-dependent and -independent apoptotsome (cytochrome c/Apaf-1/ ATP/procaspase 9) formation. In all cases, signals converge on Caspase-3 to induce apoptosis.

the effect of down-regulation of Bcl family members in response to DNA damage (Fig. 7.3). It is important to note that in some cell systems, such as glioma cells, the p53 driven apoptotic pathway can be stimulated by low level DNA damage (notably O6MeG), whereas in p53 mutated cells higher DNA damage levels are required in order to elicit the same apoptotic response [64]. Important factors that modulate the effect of DNA damage are p53, Jun kinase/p38 kinase, caspase-2, NF-kB and Akt, which will be briefly discussed below.

7.4.1 p53 As outlined above, p53 becomes activated in response to DNA damage due to phosphorylation by ATM/ATR. Once phosphorylated, p53 translocates into the nucleus and acts as the transcription factor of a large number of genes. In this context, pro and anti-apoptotic genes are of interest. Proapoptotic genes encode for Fas-R, Bax, PUMA,

NOXA, Apaf-1, and Pidd (Fig. 7.4). Anti-apoptotic genes regulated by p53 are DDB2, XPC, Fen1, MGMT, and MSH2 [65] – all of these are involved in DNA repair (Fig. 7.4). Therefore, it appears that p53 regulates some DNA repair genes and reduces the level of DNA lesions, whereas under certain circumstances it exacerbates the deleterious effects of genotoxins leading to apoptosis. This is exemplified in glioblastoma (GBM) cells, in which wild-type p53 imparts greater sensitivity to the methylating drug temozolomide than mutant p53, and this is probably due to p53-dependent upregulation of Fas [64]. Contrary to this are p53 wild-type GBM cells more resistant to the chloroethylating and crosslink inducing agent nimustine (ACNU), which was related to an upregulation of the DNA repair genes ddb2 and xpc, the products of which are supposed to participate in the repair of DNA chloroethylation lesions [66]. In this case, the decision of p53 to stimulate apoptosis or DNA repair is obviously dependent on the genotoxic agent,


following DNA damage. JNK/p38 kinase becomes activated by EGF receptor phosphorylation. As outlined above, this is counteracted by MKP1 that dephosphorylates JNK and, therefore, downregulates its activity [68,69]. Genotoxic agents may provoke a reduction in the level of MKP1, which is probably due to transcription blockage that is a result of bulky lesions in the DNA, and therefore ameliorates apoptosis by sustained upregulation of JNK/p38 kinase [10,67] (Fig. 7.3).

DSBs ATM p53 pro-apoptotic Fas-Receptor Bax Puma Noxa Apaf1 Pidd

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anti-apoptotic DDB2 XPC Fen1 MGMT MSH2

Figure 7.4. p53 triggers survival and death pathways. DNA damage activates p53, which upregulates both pro and anti-apoptotic genes. The balance between the two sets of signals dictate the final fate of the cell, and this depends on the extent of DNA damage and cell context.

but it may also depend on the dose level of a given genotoxin. Thus, it is reasonable to speculate that with low dose levels, DNA repair is stimulated whereas with high dose levels apoptosis is activated. Careful dose–response studies are needed in order to substantiate this hypothesis.

7.4.2 Sustained JNK and p38 Kinase Activation Some DNA damaging agents induce the activation of stress-activated protein kinase/c-Jun N-terminal kinase (SAPK/JNK) and p38 kinase, which results in an increase in c-Jun level and AP-1 activity. This sustained activation of AP-1 is accompanied by transcriptional activation of the Fas-L gene. Nucleotide excision repair (NER) defective mutants display a higher level of sustained JNK/p38 kinase activation, indicating that DNA damage is responsible for the response [67]. Together with DNA damage-induced p53 upregulation, which triggers transcription of the Fas receptor, the sustained upregulation of JNK/p38 kinase and concomitant AP-1 drives Fas ligand-dependent apoptosis

7.4.3 Caspase-2 Activation A very promising line of research has focused on finding a possible link between the nucleus (DNA damage) and the apoptotic machinery in the cytoplasm (mitochondria). It was argued that caspases, proteases crucial in the initiation and execution of apoptosis, might play a role here. The only procaspase constitutively present in the nucleus is caspase-2 [70,71], which may, therefore, have a role in DNA damage triggered apoptosis. Indeed, caspase-2 is required for etoposide, cisplatin, and UV-light induced apoptosis [72]. Furthermore, germ cells and oocytes from caspase-2 knockout mice are more resistant to doxorubicin [73]. Caspase-2 induced apoptosis also requires caspase-9 activation [74] indicating that caspase-2 acts through the mitochondria damage pathway. The action of caspase-2 may facilitate the mitochondrial apoptotic pathway in two ways: 1) Caspase-2 may activate the proapoptotic Bcl-2 family proteins and thereby causes release of cytochrome c from mitochondria, or 2) caspase-2 may promote the release of cytochrome c independent of the Bcl-2 family proteins. In the first mode of action, caspase-2 acts upstream of the mitochondria by inducing Bid cleavage, Bax translocation, and subsequent cytochrome c release [72,74,75]. In the second proposed mode of action, caspase-2 directly activates the mitochondrial pathway [76], which is independent of its enzymatic activity. Caspase-2 can also disrupt the interaction of cytochrome c with anionic phospholipids, notably cardiolipin, and thereby enhance the release of cytochrome c [77]. Despite the evidence supporting a role of caspase-2 in DNA damage induced apoptosis, there is no model linking caspase-2 to a given type of DNA damage such as pyrimidine dimers,

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O6-methylguanine, or DSBs. Therefore, caspase-2 may be considered as a candidate for providing a link between the primary DNA damage and downstream apoptosis players. Recently, it has been shown that caspase-2 physically interacts with DNA-PKCS and, thereby, stimulates the activity of this DNA damage activated kinase [111]. Therefore, caspase-2 may be considered to play a crucial role in regulating the balance between DNA damage, DNA repair, and apoptosis.

B. Kaina et al.

TRAF1 and 2 [87]. Bcl-xL and A1/Bfl-1 are both members of the Bcl-2 family of proteins that inhibit apoptosis by heterodimerization with the proapoptotic Bcl-2 family members [88–90]. Since c-IAP2, TRAF1, and TRAF2 are involved in the inhibition of the death receptor pathway, the induction of these proteins by NF-kB suppresses the activation of caspase-8 dependent apoptosis [87]. Interestingly, NFkB also up-regulates MKP1 [91], which counteracts JNK phosphorylation and JNK driven apoptosis observed after sustained DNA damage induction (see above).

7.4.4 Survival Signaling Mediated by NF-kB

7.4.5 Survival Signaling Mediated by Akt

The NF-kB family of transcription factors [78] is grouped into two classes. Class 1 comprises the proteins NF-kB1 and NF-kB2, which are respectively expressed as large proteins, p105 and p100, and then processed by the ubiquitin/proteosome pathway to the mature proteins p58 and p52 [79]. This processing occurs by the degradation of the C-terminal regions of p105 and p100. Class 2 comprises the proteins RelA, RelB and cRel. Both classes of proteins have a Rel homology domain in their N-terminus and are, therefore, referred to as the NF-kB/Rel proteins. NF-kB is found in an inactive state in the cytoplasm of the cells [80] and becomes rapidly activated and translocated into the nucleus upon signaling from the receptor activator of nuclear factor k B (RANK) [81], tumor necrosis factor receptor (TNFR) [82] or toll-like receptors (TLR) [83] on the cell membrane. NF-kB is kept inactive in the cytoplasm due to its binding to its inhibitor IkB (inhibitor of kB) that shields the nuclear localization signal (NLS) of NF-kB. For the activation of NF-kB, IkB is phosphorylated by IkB kinase (IKK) at serines 32 and 36 (human IkB), which leads to the ubiquitination and proteasomal degradation of IkB [84]. NF-kB then becomes nuclear localized and can mediate the transcription of its target genes. Transcriptional activation by NF-kB leads to changes in the inflammatory and immune response as well as cell cycle control. The focus of this discussion will rest on the fact that NF-kB regulates the expression of anti-apoptotic proteins and, therefore, has a profound prosurvival function. NF-kB causes the expression of the anti-apoptotic proteins Bcl-xL, A1/Bfl-1 [85], c-IAP2 [86] and

Akt (also known as protein kinase B) is a serinethreonine kinase that plays a central role in cellular signaling. Dependent on the downstream target proteins, Akt can have both an influence on protein translation, apoptosis, cell cycle control, longevity, and metabolism. The overall function and control that Akt has in the cell is complex, incompletely characterized and far beyond the scope of this discussion. This section will focus on the influence that Akt has on apoptosis. Akt activation by phosphatidylinositol 3-kinase (PI3-kinase) is required for the suppression of apoptosis [92]. The inhibition of apoptosis by Akt has been characterized in many cancer cell systems [93–96]. Apoptosis can be suppressed by Akt through the use of three basic strategies. Firstly, by directly causing the inhibition of apoptotic factors involved in the execution of apoptosis. Secondly, Akt can influence the transcriptional control needed during apoptosis and, thirdly, Akt can indirectly influence apoptosis by the impact it has on cellular signaling related to p53 regulation (Fig. 7.5). Direct inhibition of apoptosis by Akt occurs via the phosphorylation of BAD on Ser136 [97,98]. BAD is a member of the Bcl-2 family proteins and acts by binding to Bcl-2 or Bcl-XL and thereby inhibiting their anti-apoptotic function [90]. When Akt phosphorylates BAD, phosphoBAD becomes sequestrated by 14-3-3 proteins and is inactivated, as the phosphorylated BAD has a lower binding affinity for Bcl-XL [99]. The release of Bcl-2 and Bcl-XL free these anti-apoptotic proteins and, thus, they can fulfill their apoptosis suppression function.

7. DNA Damage Response and the Balance Between Cell Survival and Cell Death Akt

Direct inhibition of apoptosis by phosphorylation of: Bad

Caspase-9

inactivation

inactivation

Inhibition of mitochondrialmediated apoptosis

Transcriptional Inhibition of apoptosis by phosphorylation of: -B

MDM2

NF -B activation

p53 inhibition

Upregulation of survival genes

No induction of apoptotic genes

Figure 7.5. Anti-apoptotic pathways triggered by Akt. The anti-apoptotic events are regulated by posttranslational modification by phosphorylation, which either directly inactivates proapoptoic Bad and caspase-9 or indirectly upregulates anti-apoptotic genes (through activation of NF-kB) and transrepresses pro-apoptotic genes (by inhibition of p53).

Another direct inhibition target of Akt phosphorylation is caspase-9. This caspase together with Apaf-1 and cytochrome c constitute the apoptosome (Fig. 7.3), which is one of the centrally required components for mitochondrial mediated apoptosis [100]. It has been shown that Akt can phosphorylate human caspase-9 on Ser196, and thus attenuates the apoptotic activity of caspase-9 [101]. Interestingly, suppression of caspase-9 activity only seems to apply to humans as Akt does not seem to be able to phosphorylate mouse caspase-9 [102]. The direct inhibition of apoptosis by Akt occurs via phosphorylation at Ser83 of apoptosis signal-regulating kinase 1 (ASK1) [103]. ASK1 is upstream of JNK and p38 kinase, which is further upstream of Bid cleavage, Bax translocation, and cytochrome c release [104]. Collectively, it is clear that Akt, through its direct phosphorylations of BAD, caspase-9 and ASK1, suppresses the mitochondrial apoptotic pathway. The major transcriptional control that Akt has on the suppression of apoptosis is due to the activation of the transcription factor nuclear factor-kB (NF-kB) and the prevention of the nuclear localization of p53. As outlined above, NF-kB transcribes the anti-apoptotic Bcl-2 family proteins Bfl1/A1, Bcl-XL and Nrl3 as well as the direct apoptosis inhibitors IAPs [105]. The mechanism whereby Akt promotes the activity of NF-kB is by advancing

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the degradation of the NF-kB inhibitor IkB [106] that then releases NF-kB for transcribing its prosurvival target genes. As already stated, the second mode for transcriptional suppression of apoptosis by Akt is by preventing the nuclear localization of p53. Akt binds to and phosphorylates at Ser166 and Ser186 of Mdm2, the E3 ubiquitin ligase murine double minute 2, and thereby induces its nuclear import or increases its ubiquitin ligase activity [107,108]. Mdm2 is greatly involved in the inactivation of p53, and this increase in nuclear Mdm2 level and activity will inactivate the proapoptotic transcriptional function of p53 (Fig. 7.5). As Akt is activated via growth factors binding to membrane receptors and PI-3 kinase signaling [109], the question now arises as to what Akt has to do with DNA damage triggered apoptosis. The answer can be found in the p53 protein, which not only has a profound influence on apoptosis, but it also plays a central role in DNA damage signaling and repair [110]. The suppression of p53 function by Akt following the induction of DNA damage may lead to a high nontolerable level of unrepaired DNA lesions. The cell will then have to overcome the prosurvival signaling of Akt in order to undergo apoptosis. The consequences for the cell in failing to do this might lead to necrotic cell death, which in the case of chemo- or radiation therapy will lead to inflammation, or fixation of DNA damage, with resultant mutation and/or tumorigenesis. This shows that upon the induction of DNA damage, various pathways are both converging and competing, regulating survival, upregulating DNA repair genes, inducing cell cycle arrest through checkpoint response to allow more time for repair, presumably also promoting damage tolerance functions and, in the worst scenario, all of these confounding signals pushing the cell into controlled cell death – apoptosis.

7.5 Conclusion Following DNA damage, a number of pathways are activated; some transduce cell death signals, with others transducing survival signals. The numerous signals of varying intensities converge and compete, and are carefully regulated by cell context and the level of DNA damage, with the net effect being either cell survival or death by apoptosis.

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Understanding signaling pathways that negatively perturb the balance between cell survival and cell death is essential not only in defining the process of tumorigenesis, but also for devising therapeutic concepts that are more effective in the management of cancer.

Acknowledgements: This work of the authors is supported by Deutsche Forschungsgemeinschaft, Mildred-Scheel Stiftung für Krebsforschung und Stiftung Rheinland-Pfalz.

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Chapter 8

Dysfunction of the RB Retinoblastoma Gene in Cancer Francesca Pentimalli, Letizia Cito, and Antonio Giordano

Abstract The retinoblastoma gene RB, which was the first tumor suppressor gene to be identified, is a key regulator of the cell cycle and its inactivation, either direct or indirect, underlies multiple types of human tumors. Consistent with its role as tumor suppressor, it is well established that RB inhibits cell proliferation by binding to the E2F family of transcription factors thereby repressing genes that are required for the G1–S transition of the cell cycle. However, in the past decade, a myriad of studies focusing on the role of RB in cancer development implicated RB in many cellular processes that could all contribute to its tumor suppressor function, suggesting that the role of RB in cancer is much more complex than previously thought. To further complicate matters, the other members of the RB family, retinoblastoma-like 1 (RBL1 or p107) and retinoblastoma-like 2 (RBL2 or p130), have both overlapping and distinct functions compared with RB and many cellular functions of RB are mediated by over a hundred interacting proteins and numerous transcriptional targets. Now, emerging evidence shows that RB status can influence the response to different anti-cancer therapeutics according to the context. Therefore, a thorough understanding of all RB functions in cancer is more crucial than ever. Keywords Retinoblastoma • RB family • Tumor suppressors • Cell cycle • G1–S transition • Checkpoints • E2F

8.1 Introduction The discovery of the retinoblastoma gene RB has been a milestone in cancer research. At the beginning of the twentieth century Boveri suggested that individuals inheriting chromosomal alterations are predisposed to cancer development [1]. Much later, in 1971, Knudson, by studying the epidemiology of the paediatric tumor retinoblastoma, postulated a model according to which two mutational events or ‘two hits’ are required for tumor development; therefore, an individual who inherits a specific mutation in a cancer gene will be predisposed to cancer because only another somatic mutation in the same gene will be enough for the tumor to develop [2]. Only a few years later the work from Cavenee and colleagues provided the molecular tools, through loss of heterozygosity (LOH) studies, which led to the mapping and subsequent cloning of the RB gene [3–6]. The discovery that biallelic RB mutations were indeed causative of retinoblastoma tumors [7] confirmed Knudson hypothesis and identified RB as the first tumor suppressor gene, the inactivation of which confers increased susceptibility to cancer. Then, RB was found to be mutated in several other human tumors, such as osteosarcomas 4, small-cell lung cancer (SCLC) [8], breast [9] and bladder cancer [10]. Now it is well established that alterations of the RB pathway occur in the majority of human tumors.


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Subsequent work identified the RB gene product as a phosphoprotein that was phosphorylated during specific phases of the cell cycle [11, 12] and that was bound by the E1A [13], T antigen [14] and E7 [15] oncoproteins expressed by DNA tumor viruses, such as adenovirus, SV40 and papillomavirus, respectively. These studies were the first to implicate RB function in the regulation of cell proliferation. In 1991, the transcription factor E2F was identified as the first cellular target of RB [16] and a few years later Weinberg proposed the still valid model according to which RB regulates cell cycle progression by binding E2F and blocking the transcription of genes necessary for the G1–S transition [17]. Despite decades of intensive research, however, many gaps remain in our understanding of RB function, probably because of several confounding issues. First of all, beyond its role as regulator of the G1 checkpoint, RB has key functions in a wide range of biological processes including differentiation, apoptosis, senescence, DNA repair, and replication, all of which are likely to contribute to its tumor suppressor function. Second, RB belongs to a family of proteins and it shares with the other members, the retinoblastoma-like 1 (RBL1 or p107) and retinoblastoma-like 2 (RBL2 or p130) proteins, both overlapping and distinct functions [18–20], which makes more difficult to dissect the specific role of each protein. Third, RB has been shown to interact with over a hundred cellular proteins [21] and for most of these interactions the biological significance still has to be assessed. Moreover, RB acts as a transcriptional co-factor with the ability to either inhibit or stimulate the function of various transcription factors thereby affecting the transcription of a wide range of genes [22, 23] and it also participates in the epigenetic regulation of chromatin, with effects not only on gene expression but also on proper chromosome segregation and telomere maintenance [24]. Dissecting and understanding all the functions of RB family members that contribute to tumor suppression in different cell types and at different steps of tumorigenesis is crucial to identify new possible therapeutic strategies. Moreover, recent evidence is showing that RB status can be predictive of the therapeutic response to different anti-cancer agents according to the context, therefore RB expression should be taken into account when considering therapeutic strategies.

F. Pentimalli et al.

8.2 The RB Family The RB family consists of three structurally and functionally related proteins: RB (also known as p105), p107 and p130, with their genes being located on chromosomes 13q14.2, 20q11.2 and 16q12.2, respectively. All family members are also called ‘pocket proteins’ because they share sequence homology in a bipartite region that folds into a globular pocket-like domain. The pocket region is structurally characterized by two subdomains, A and B, which are highly conserved among the three family members and are separated by a spacer region, which is longer and more conserved between p107 and p130 rather than RB [18, 25]. The pocket domain mediates the interaction with members of the E2F family of transcription factors and with proteins containing an LXCXE motif, such as viral oncoproteins, D-type cyclins and histone deacetylases (HDACs), whereas the spacer domain allows the assembly of the A and B subdomains into the pocket-like structure. A nuclear localization signal (NLS) located in the carboxy terminal region of the RB family members allows protein transport to the nucleus and can act as a carrier for proteins, such as the E2F4 and E2F5 transcription factors, that lack an NLS [25]. The three pocket proteins are differentially expressed throughout the cell cycle: RB expression is steady whereas p107 expression peaks during the S phase and p130 expression is higher in arrested cells [18, 26]. All pocket proteins are implicated in the regulation of cell cycle: their overexpression cause cell cycle arrest in G1 in most cell types; they all interact with E2F family members (although in different combinations) and repress E2F-mediated transcription; they are all substrates of cyclindependent kinases (CDKs) and undergo phosphorylation and dephosphorylation during cell cycle. However, only p107 and p130 possess cyclindependent kinase (CDK)-inhibitory activity [19, 26].

8.3 RB and the Cell Cycle Cell cycle control is the key cellular process governing cell proliferation and its malfunctioning is the leading cause of cancer. From the mid 1990s

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it is known that RB has a crucial role in cell cycle control, mainly acting as the gatekeeper of the G1–S transition through its interaction with E2F proteins. Following mitogenic clues, progression through the cell cycle is driven by the sequential activation of cyclin–CDK complexes, which are specific for each cell-cycle phase and phosphorylate different sets of target proteins, whereas CDK inhibitors (CKIs), which are activated by anti-proliferative stimuli, counteract their activity by acting as cell-cycle brakes. In quiescent cells (G0) and in early G1, RB is hypophosphorylated and is able to repress E2F by direct binding with the highest affinity (Fig. 8.1). Mitogenic signals converging on the cell cycle machinery activate the G1 phase cyclin D–CDK4 and cyclin D–CDK6 complexes that initiate RB phosphorylation with the consequent release of E2F inhibition and the transcriptional activation of a panel of genes required for the S phase and cell cycle progression, including genes that activate CDK2 and further promote RB phosphorylation [25, 27, 28]. RB in fact is maintained hyperphosphorylated and thereby inactive until mitosis by the sequential activation of different cyclin–CDK complexes, including cyclin E–CDK2, cyclin A–CDK2, and cyclin B–CDK1. During mitosis, RB inhibitory activity is restored through dephosphorylation by phosphatase activity [29]. Anti-proliferative stimuli, however, can restore RB activity during the cell cycle by induction of CKIs, modulation of cyclin expression or of phosphatase activity [28], suggesting that RB is the hub of cell cycle regulation. This model clearly explains how RB loss or mutation leads to E2F deregulation and consequent aberrant proliferation and how other oncogenic events, directly or indirectly affecting RB, can impair cell cycle regulation (see the following sections for further discussion). However, this early model of cell cycle regulation has been much more complicated by the fact that both RB and E2F belong to families of proteins and they all form a network with many more functions than previously thought. It was shown that pocket proteins can inhibit E2F-mediated transcription not only by blocking the E2F transactivation domain, but also actively repressing transcription [30] through the recruitment of chromatin remodelling enzymes

such as HDACs, the SWI/SNF complex and the histone methyltransferase SUV39H1 [19]. Moreover, the RB family–E2Fs network has been shown to regulate not only genes required for S phase entry and progression but also genes required for mitosis, checkpoint control, DNA damage response, differentiation, apoptosis, and development suggesting that deregulation of this network does not only lead to increased proliferation, but also accounts for other mechanisms inducing tumorigenesis [19].

8.3.1 The E2F Family and Its Pocket Protein-Mediated Regulation At least eight members belong to the E2F transcription factor family in addition to several isoforms that are generated by alternative splicing. Most of them require heterodimerization with members of the DP family of transcription factors for high affinity DNA binding. E2F1, E2F2, and E2F3a are strong transcriptional activators that are inhibited by RB binding and stimulate gene expression in late G1, whereas they are generally absent or expressed at low levels in quiescent cells. E2F4 and E2F5 are transcriptional repressors that are expressed throughout the cell cycle but only in G0 and early G1 they are recruited to the nucleus through the binding with p107 and p130. The E2F3b isoform is constitutively expressed, like E2F4 and E2F5, suggesting that it could function as a transcriptional repressor as well, however, recent evidence suggests that it has overlapping roles and significant functional redundancy with the other activator E2F factors [31, 32]. E2F6 is also a transcriptional repressor that acts through recruitment of the Polycomb group of proteins to establish a repressive chromatin structure on E2Fresponsive promoters. The recently identified E2F7 and E2F8 bind to DNA in a DP-independent manner and also repress E2F site-dependent transcription in a RB-independent manner [19, 25]. In G0 and early G1, most E2F-responsive genes are repressed by complexes that include repressor E2Fs (E2F4 or E2F5), which are recruited to the nucleus by p130 or p107, and HDACs, which promote nucleosome formation by removing acetyl groups from histones, whereas RB holds E2F1–3 inactive and away from their responsive promoters.

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Figure 8.1. Schematic representation of RB throughout the cell cycle. In quiescent cells (G0) and in early G1, RB is hypophosphorylated and inhibits cell-cycle progression by binding and repressing E2F. Then, mitogenic signals activate the G1 phase cyclin D–CDK4 and cylclin D–CDK6 complexes that initiate RB phosphorylation with the consequent release of E2F inhibition. The E2F-mediated gene expression program drives progression through S phase and G2/M. RB is maintained hyperphosphorylated until mitosis by the sequential activation of different cylclin–CDK complexes. During mitosis, RB inhibitory activity is restored through dehpsphorylation by phosphatase activity. However, anti-proliferative stimuli can restore RB activity during the cell cycle by induction of CKIs and modulation of cyclin expression or of phosphatase activity.

In late G1, pocket proteins are targeted by phosphorylation and release their E2F partners: E2F4 and E2F5 relocate to the cytoplasm whereas E2F1–3 can bind their responsive promoters. Gene transcription is often facilitated through the recruitment of histone acetyltransferases (HATs) [19, 25, 33]. Pocket proteins can also repress E2F-responsive genes through SUV39H1-mediated methylation of the histone H3 lysine 9 (H3K9), which creates a high-affinity binding site for members of the heterochromatin protein 1 (HP1) family that induce transcriptional silencing. Higher levels of methylated H3K9 have been found at E2F-responsive promoters in senescent cells compared with quiescent cells [34], suggesting that pocket proteins have an important role in determining and/or maintaining

a senescent or differentiated state by mediating a stable gene repression. So, E2F-responsive genes are regulated through both the inactivation of the activator E2Fs by RB and the formation of repressor complexes by p107 and p130. However, the control of E2F-responsive genes by pocket proteins is not redundant because the genes that are deregulated owing to the loss of p107 and p130 differ from those associated with RB loss [19]; it seems that RB, p107 and p130 regulate overlapping but also distinct sets of genes and their function at a given promoter depends on additional cofactors and on the specific context such as cell type or stage (for example during quiescence, senescence or differentiation) [19, 25, 35]. This is currently a matter of intensive investigation.

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8.3.2 E2F-Independent Mechanisms of Cell Cycle Regulation Pocket proteins can affect cell cycle progression also through E2F-independent mechanisms. For example, RB has been found to inhibit the S-phase kinase-associated protein 2 (SKP2)-mediated ubiquitylation and degradation of the CDK inhibitor p27KIP1 with consequent inhibition of CDK activity and resulting cell cycle arrest [36]. More recently, Dyson and colleagues have further dissected this pathway showing that RB, but not p107 and p130, interacts with the anaphase-promoting complex/cyclosome (APC)/C through its activator CDH1 and this interaction promotes the ubiquitylation and degradation of SKP2 and the subsequent stabilization of p27KIP1. These authors propose that RB might use ubiquitylation and the turnover of more APC/C targets to induce cell cycle exit and maintain a differentiated state [37]. Moreover, given that SKP2 has also been shown to be directly regulated by E2F1 at the transcriptional level, Assoian has recently provided a model according to which there is a positive feedback loop including RB–E2F, SKP2, p27KIP1 and cyclin E–CDK2 whose regulation is crucial for passage through the restriction point (the point during G1 at which cell cycle progression becomes independent of mitogenic stimuli) [38]. Interestingly, p107 and p130 contain, in the N-terminal and in the spacer region respectively, a kinase inhibitory domain that has been shown to specifically inhibit CDK2 kinase activity in vivo for p107 [39, 40] and in vitro [41] and in vivo [42] for p130. In late G1, if the conditions favor cell division, CDK2 activity overpowers repression by p27KIP1 and p130 and, in concert with cyclin E, inactivates the RB family members enabling progression through the cycle [33].

8.4 RB and Cancer Cancer is a disease in which the mechanisms that govern normal cell proliferation and homeostasis are impaired. Given that more than 100 distinct types of cancer exist, often with many different variants in specific organs, it is a very complex disease. In 2000, Hanahan and Weinberg defined the hallmarks of cancer cells identifying six dif-

ferent essential alterations in cell physiology that collectively dictate malignant growth [43]: in some way RB is involved in most of these processes. Soon after the seminal discovery of the RB gene as the first tumor suppressor, owing to its role in retinoblastoma, it was clear that RB is frequently mutated in a variety of human cancers. Then, mutations affecting the RB pathway were found to occur with such a high frequency in cancer that it was proposed that disabling the RB pathway is essential for tumorigenesis [44]. Indeed, RB is altered in tumors not only directly, through different mechanisms, but also indirectly, through deregulation of other pathway members.

8.4.1 Different Mechanisms of RB Inactivation in Cancer RB is known to be directly inactivated by different mechanisms. First of all, as we discussed above, the genetic inactivation of RB underlies its role in the susceptibility to and occurrence of retinoblastoma, both in familial and sporadic cases, and in osteosarcomas [4]. Genetic or epigenetic inactivation of RB is found in many human tumors such as lung, prostate, breast, bladder, oesophageal, and liver cancer as well as in advanced gliomas and in some cases of chronic myeloid leukaemia 45. In lung cancer, in particular, germline mutations of RB predispose to SCLC and RB is mutated in more than 90% of sporadic SCLCs, whereas in only 15–30% of non-SCLC cases [45]. RB mutations are rarely found in other cancer types such as colorectal, pancreatic, renal cell, endometrial, ovarian, thyroid and gastric carcinomas as well as in myeloma, nonHodgkin lymphoma and HPVnegative head and neck carcinoma, even though in many of these cases mutations affecting the RB pathway in other ways are present [45]. Soon after the discovery of the RB protein, as mentioned above, it was found that viral proteins, such as adenovirus E1A [13, 46, 47], SV40 T antigen [14] and papillomavirus E7 [15], could bind to and inactivate RB and its family members. These small DNA viruses take over the DNA replication machinery of host cells to induce viral genome replication by prematurely stimulating cell cycle S-phase entry, through the dissociation of E2Fs from RB. Infection with oncoviruses has been

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shown to contribute to tumorigenesis in cervical carcinoma [48], squamous cell carcinoma of the head and neck [49] and mesothelioma [50], although it is difficult to dissect the precise mechanism of action given that oncoviral proteins can often inhibit all RB family members and other crucial tumor suppressors such as p53. Recently, another mechanism has been found that could underlie viral-induced tumorigenesis through inactivation of RB. The authors found that the RNAdependent RNA polymerase, nonstructural protein 5B (NS5B) of the hepatitis C virus (HCV) mediates RB ubiquitylation and its subsequent degradation via the proteasome; this mechanism is likely to induce the proliferation of HCV-infected liver cells and might contribute to liver tumorigenesis [51]. Human liver cancers also express high levels of gankyrin, a protein that inactivates RB at multiple levels. Gankyrin directly binds RB, but not p107 or p130, and promotes its phosphorylation and degradation. But gankyrin also binds CDK4, precluding its inhibition by p16INK4A, which results in increased RB phosphorylation and consequent stimulation of E2F transcriptional activity, thereby causing cell cycle progression [52, 53]. These findings suggest that multiple mechanisms, acting both directly and indirectly, can act together to achieve RB inactivation in particular contexts. Similarly, the inverted CCAAT box binding protein of 90 kDa (ICBP90) has the ability to regulate RB both at the transcriptional level, through binding of the RB promoter, and at the protein level. Overexpression of ICBP90, which has been found in cancer cells, downregulates RB expression and favors entry of cells into the S phase [54]. As RB has been shown to interact with many other proteins, it is likely that many of these interactions can affect its tumor suppressive role. Furthermore, recent evidence showed that members of the RB family can be posttranscriptionally regulated by microRNAs suggesting yet another possible way of RB deregulation that could occur in cancer and might have been overlooked in the past decades [55, 56].

8.4.2 RB Pathway Deregulation in Cancer Consistent with the importance of RB in regulating cell proliferation by controlling the G1–S transition, other key players of the RB path-


way acting in this context are often altered in cancer [44]. Mitogenic signals converge to stimulate the expression of D-type cyclins and to increase CDK4/6 activity whereas the CDK inhibitor p16INK4A counteracts the activity of cyclin D–CDK4/6 complexes; RB-mediated activation of E2F will depend on the final effect between these balancing forces (Fig. 8.1). Cyclin D is often found to be overexpressed in a variety of human cancers by different mechanisms, such as chromosomal translocation, gene amplification, provirus integration or reduced degradation [57]. For example, the genetic feature of mantle cell lymphoma is the chromosomal translocation t(11;14)(q13;q32) that juxtaposes CCND1, which encodes cyclin D1, to the immunoglobulin heavy chain gene and results in the constitutive overexpression of cyclin D1. This leads to increased cell proliferation and is thought to be the primary event in the pathogenesis of this tumor [58]. Also in multiple myeloma translocations involving the CCND1 locus occur; whereas in 5% of multiple myeloma cases it is the cyclin D3 to be overexpressed as a result of a chromosomal translocation [59]. Gene amplification is another common mechanism leading to aberrant overexpression of cyclin D1 and has been found to occur in nonSCLC, head and neck squamous cell carcinomas, pancreatic carcinomas, bladder cancer, pituitary adenomas, and breast cancer [60]. Other mechanisms may also account for the increased overexpression of cyclin D in many human cancers. For example, Diehl and colleagues found mutations impairing the nuclear export of cyclin D that favor its nuclear accumulation and seem to contribute to carcinogenesis in oesophageal tumors [61]. Moreover, these authors recently identified the FBX4 protein of the ubiquitin protein ligase complex SCF as the E3 ligase for cyclin D, which is responsible for its degradation. As FBX4 is often altered in human cancer, this seems to be another mechanism that induces tumorigenesis by increasing cyclin D expression [62]. Similar to cyclin D, CDK4 is also found to be amplified in sarcomas and gliomas [44], whereas mutations impairing CDK4 or CDK6 interaction with p16INK4A have been described in melanoma [63] and neuroblastoma [64], respectively. Beyond RB, the most frequent mutation in human cancers affecting the RB–E2F pathway involves the CDK inhibitor p16INK4A, the loss of which results

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in high cyclin D–CDK4 activity and consequent RB inactivation by phosphorylation. Inherited mutations of the gene encoding p16INK4A, CDKN2A, and subsequent loss of the wild-type allele in tumors were first observed in familial melanoma [65]; however, similar to RB, loss of p16INK4A function is much more prevalent in sporadic cancers of many cancer types [66] and occurs through different mechanisms including epigenetic silencing. Usually mutational events resulting in RB or p16INK4A loss, or in the overexpression of cyclin D or CDK4/6, are mutually exclusive, suggesting that these molecules normally function in a linear pathway that suppresses tumorigenesis. Although RB mutations are so frequent in human cancers, mutations in other members of the RB family are not common events, with few exceptions. For example, loss of function mutations within the p130 gene have been found in SCLC lung carcinoma cell lines [67] and lung cancer samples [68]. According to Nevins,what is most striking is the general lack of mutational events affecting members of the E2F family. Even though E2F1 loss in the mouse contributes to tumorigenesis, E2F3 has a key role in the control of cellular proliferation and E2F4 in determining cellular differentiation; direct alteration of these genes does not seem to contribute significantly to cancer development. A potential explanation for this phenomenon could rely on the fact that E2Fs widely overlap in function or it might imply that E2F deregulation could be so disruptive that would not allow a productive event [66].

8.4.3 RB, Angiogenesis, and Metastasis The growth of solid tumors depends on the generation of new vascular supply, which involves the recruitment of endothelial cells to form new blood vessels and results from the balance between proangiogenic and anti-angiogenic factors that regulate the socalled “angiogenic switch.” Recently, several findings showed that the RB family is involved in different facets of the angiogenic process [69]. Both RB and p130 have been shown to modulate the expression of important angiogenesis modulators. For example, a functional RB is required for the cyclin A-induced autocrine expression of the vascular endothelial growth factor (VEGF) in prostate cancer [70], and in mice, RB null tumors showed reduced levels of VEGF [71]. Instead

p130 overexpression has been shown to cause regression of established tumor grafts in mice, by inhibiting angiogenesis through the downregulation of VEGF [72]. Even though RB relation with VEGF expression still needs to be clarified, it seems that VEGF itself can regulate RB proteins in certain contexts. It has been shown that VEGF stimulation leads to the dissociation of RB family members, E2F4, and E2F5 from the human metallothionein 1G promoter – the expression of which is required in endothelial cells during angiogenesis – and increases the binding of E2F1–3 [73]. This suggests that VEGF could regulate the expression of angiogenic factors through modulation of the RB–E2F pathway. Interestingly, many genes involved in angiogenesis contain E2F binding sequences in their promoters [69]. Another possible mechanism for RB involvement in angiogenesis is through its interaction with other protein partners. For example, in a mouse model of pituitary carcinogenesis RB loss releases the ID2 protein, which inhibits the DNA binding of transcription factors containing the basic helixloop-helix motif, and ID2 consequently promotes growth and angiogenesis by functioning as a master regulator of VEGF [74]. Recent work also showed that blocking RB binding to the RAF1 kinase with a specific small molecule is able to inhibit angiogenesis both in vitro and in vivo [75]. RB alterations seem also to be involved in other processes affecting cancer progression, such as invasion and metastasis. These effects could be mediated by E2Fs given that several genes targeted by these transcription factors have a role in these processes [45]. However, a thorough understanding of these mechanisms requires further studies.

8.4.4 Mouse Models of RB Loss and Cancer Mouse models of cancer have largely contributed to uncover RB functions in the past decades. The first studies modelling RB loss in mice were quite surprising. Although human patients carrying an RB germline mutation develop predominant retinoblastomas, the targeted disruption of one RB allele in mice does not predispose to this type of cancer but in contrast leads to pituitary and thyroid tumors. This difference was tentatively attributed to species-specific differences in the number of susceptible cells, in the timing of susceptibility

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during development, or in the number of mutational requirements. Subsequent studies in fact have shown that retinoblastoma development in mice requires other genetic mutations in addition to RB loss, such as alterations resulting in the loss of p107 or of the p53 tumor suppressor. These studies also showed that RB loss is not necessarily a rate-limiting event for tumor initiation but can also provide a growth advantage during later stages of disease. This is consistent with the fact that, whereas RB germline mutations predispose to a limited range of tumors, in sporadic cancer RB loss underlies the occurrence of many cancer types. The development of mice null for the other members of the RB family, conditional and tissue specific knockout mice and the crossing with mice carrying targeted mutations in other genes have further contributed to understanding RB function in tumorigenesis and have been extensively reviewed elsewhere [26, 76, 77].

8.5 RB Functions Affecting Its Tumor Suppressor Role In the past decade, many new functions of RB were uncovered, implicating it in various cellular processes. Each of these functions is likely to contribute to RB tumor suppressor function, suggesting that its role in cancer is much more complex than previously thought. For example, the RB family has a key role in promoting differentiation in various contexts (see previous discussion and below) and loss of this function can clearly affect tumor onset and/or progression. In fact, RB mutations, which impair RB ability to bind and regulate E2Fs but retain RB ability of inducing differentiation, can prevent tumorigenesis as shown in some families with low penetrant retinoblastomas [45]. Several examples of how RB’s role in different cellular processes can affect tumorigenesis are described below.

8.5.1 RB and Transcriptional Activation As discussed above, RB’s role as a repressor of E2F-mediated transcription is not only well established but it is also consistent with its tumor suppressor function. However, there is also evidence that RB can act as an activator of gene transcrip-


tion. For example, in epithelial cells RB has been shown to interact with the AP2 transcription factor and activate the transcription of the E-cadherin promoter, the master gene of the epithelial phenotype [78]. Now, Taya and colleagues show that RB depletion in breast cancer cells results in the specific down-regulation of E-cadherin expression and induction of an epithelial-to-mesenchymal transition [79]. This is consistent with the finding that, in cases of mesenchymal-like invasive breast cancer, there is a concomitant downregulation of RB and E-cadherin expression, suggesting that RB inactivation contributes to tumor progression not only by causing loss of cell proliferation control but also promoting an invasive phenotype. RB’s ability to inhibit epithelial-to-mesenchymal transition through the activation of genes that are crucial for maintaining the epithelial differentiation would therefore seem another tumor suppressor function of RB [79]. In epithelial cells, RB has also been shown to upregulate the transcription of the CDK inhibitor p21CIP1 through the SP1 family of transcription factors that, beyond regulating the expression of specific epithelial markers, are involved in the RB-mediated activation of several other promoters, such as FOS, MYC, TGFb1, and IGFII. Interestingly, the high levels of p21CIP1 achieved in this context maintain RB in a hypophosphorylated state allowing its interaction with transcription factors such as SP1 and AP2 in order to activate differentiation-specific genes [80]. A growing body of evidence supports the hypothesis of a key role for RB as an activator of transcription. To cite another example of how this function of RB can impact on tumorigenesis, Thomas and colleagues showed that RB physically interacts with the osteoblast transcription factor, CBFA1 and associates with osteoblast-specific promoters in vivo, whereas RB loss blocks osteoblast differentiation [81]. This impairment of differentiation due to RB loss seems to be another mechanism that, together with inappropriate proliferation, can account for the occurrence of osteosarcomas in both familial and sporadic cases.

8.5.2 RB and Apoptosis Many studies in the past have shown that RB has a role in apoptosis [82]. In particular, RB inhibits apoptosis and this, at least in part, is due to RB

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ability of blocking E2F1, which regulates the transcription of crucial apoptotic genes, such as those for the apoptotic peptidase activating factor 1 (APAF1) and for several caspases including the effector caspases 3 and 7 [82]. In response to certain apoptotic stimuli, RB needs to be inactivated by caspase-mediated degradation. Several results suggest that the RB tumor suppressor function of inhibiting cell proliferation, which is inactivated by phosphorylation, and the RB tumor promoting function of inhibiting apoptosis, which is inactivated by caspase cleavage, can be dissociated [83, 84]. Beyond caspase-mediated degradation, however, other mechanisms of RB inactivation, such as gene mutation or binding to viral oncoproteins, have been associated with sensitization to apoptosis [82]. So, RB inactivation can both promote cell proliferation and apoptosis raising the issue of how cells decide whether to proliferate or die. Different models have been proposed to provide an explanation (for a review see ref. [82]), however, the dual role of RB in the regulation of cell proliferation and cancer has deep implications in tumor development. It seems intuitive that RB inactivation needs to be coupled with defects in the apoptotic pathways for cancer to occur. Indeed, p53 inactivation contributes to tumorigenesis and suppression of apoptosis in RB-null mice and it is intriguing that DNA tumor viruses have developed oncoproteins that can bind and inactivate both RB and p53 [82].

8.5.3 RB and Senescence Senescence consists in the irreversible growth arrest of cells, which can be triggered by various stresses, including oxidative damage, telomere dysfunction, DNA damage, and several chemotherapeutic drugs. Cellular senescence not only contributes to ageing but it is also an important tumor suppressive mechanism that protects organisms against cancer. The RB–p16INK4A and the p53–ARF pathways are responsible for the execution of the proliferative arrest that characterizes senescence [85]. In particular, the acute loss of RB in senescent cells leads to reversal of the cellular senescence programme [86] and RB pathway integrity is necessary to maintain the stable form of cell cycle arrest typical of senescence [34]. Interestingly, there is also

evidence that other members of the RB family, in particular p130, have a role in sustaining cell cycle arrest during cellular senescence [87]. Therefore, this function of the RB family members needs to be carefully examined for its potential impact on cancer development.

8.5.4 RB and Genomic Instability Many studies focusing on RB function showed that RB is involved not only in controlling the G1–S transition but also in processes such as DNA replication and chromosomal segregation in mitosis, thereby functioning in checkpoint controls also during the S phase and the G2–M transition and following different types of DNA damage. In some contexts RB deficiency can induce, through these mechanisms, genomic instability, which is one of the main features of cancer cells. RB can inhibit ongoing DNA replication in S phase following DNA damage or other stimuli (such as cAMP signalling or viral factors inhibiting cellular DNA replication) through different mechanisms. It has been proposed that RB can inhibit DNA replication factors by both direct binding and transcriptional repression. Also, RB downregulates cyclin A reducing CDK2 activity and consequently blocking the function of important replication factors, such as proliferating cell nuclear antigen (PCNA) and the DNA polymerase processivity factor [27]. Several data also hint at a role for RB in the G2–M transition. In fact, many crucial factors involved in the G2–M transition are regulated by the RB–E2F pathway and the RB-mediated repression of some of these (cyclin B and CDK1 in particular) is required following DNA damage to maintain the G2–M checkpoint [27]. Also p107 and p130 seem to be crucial for the regulation of genes required during the G2 and M phases [88]. Unrepaired DNA damage is detrimental to an organism because it can cause genomic instability and subsequent susceptibility to various diseases including cancer. Therefore, following DNA damage, cells need to activate cell cycle checkpoints to slow down the cell cycle and allow the DNA repair systems to repair the lesions before proceeding to DNA replication, chromosomal segregation, and mitosis. Normally, if the type or amount of damage overwhelms the survival response machinery,

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apoptosis is triggered. Obviously any defect in the cellular response to DNA damage is potentially dangerous because it is likely to induce chromosomal aberrations and more or less severe genomic instability depending on the damage. As RB is crucial for most of these processes and in all the phases of the cell cycle it is not unexpected that RB loss has been associated with increased aneuploidy [45]. Recently, another mechanism has been described that accounts for the genomic instability induced by RB loss. It has been shown that RB family proteins have a direct role in the assembly of pericentric and telomeric heterochromatin domains and that lack of the RB family results in chromosome segregation defects and abnormal telomere elongation, two processes that are frequently altered in human cancer [24].

8.6 Exploiting RB Pathway Therapeutically Given the fact that the RB pathway is altered in most human cancers, it represents an appealing target for cancer therapies. However, in comparison with other tumor suppressors such as p53, there have been fewer studies aiming at restoring RB function in cancer cells [89]. Early studies suggested that RB’s ability to inhibit tumor cell growth is variable whereas the inhibition of p53-induced apoptosis can be deleterious. So, RB phosphorylation mutants and truncated variants have been considered to improve RB tumor suppressor function [89]. Data on the possible use in cancer therapy of the other RB family members are very limited, although a preclinical study using retrovirusmediated transfer of p130 proved to be promising for suppressing the growth of lung carcinoma cells in vitro and in vivo [68]. Other approaches that directly exploit RB loss have been attempted quite successfully with oncolytic viruses. For example, Fueyo and colleagues used an oncolytic virus carrying a mutated E1A adenoviral protein that renders viral replication dependent on RB inactivation and generates a tumor-selective virus, which has an anticancer effect in gliomas [90]. Although targeting tumor suppressor genes is an attractive strategy for cancer gene therapy, so far


results from clinical trials have not mirrored the preclinical studies. A greater understanding of the biology of RB and its family members is required, owing to their involvement in so many different processes, before these therapeutic approaches can be successful. Other strategies, however, have been considered to target the RB pathway. For example, the use of inhibitors of the cyclin D–CDK4 assembly, inhibitors of the mitogenic pathway activating cyclin D synthesis, and CDK inhibitors has been considered [91]. However, although the use of CDK inhibitors has been the focus of intense research in the past decades, the compounds developed so far still need to be improved much [92]. We recently identified a small molecule, Spa310, based on the p130 spacer domain, which is able to inhibit CDK2 activity and suppress tumor formation in nude mice, suggesting that Spa310 could be a good candidate for further research in this field [42]. Recently, it has emerged that the consequences of RB inactivation in tumors can be very different depending on the context and the type of cancer. Moreover, loss of RB function has been associated with a different therapeutic response according to the tumor type and the therapeutic agent. Therefore, it has been suggested that RB status should be analysed and carefully considered when designing antitumor strategies in patients [28]. Long after RB discovery it seems that a thorough understanding of its role in cancer is more crucial than ever.

Acknowledgments: The authors would like to thank the members of Giordano’s lab for their helpful comments. The authors apologize for having cited reviews in some instances, rather than many excellent primary papers, owing to the broadness of the topic and the space constraints.

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Chapter 9

G1 Phase Cyclins in Cancer Development and Progression John Patrick Alao

Abstract Cyclin D1 and cyclin E are rate limiting for progression through G1 and entry into S phase during the cell cycle. Cyclin D1 plays a crucial role in linking favorable environmental conditions with cell cycle progression. Cyclin D1 indirectly induces cyclin E expression and activation, which is required for entry into S phase. The deregulated expression of either cyclin would thus greatly impact on a cell’s ability to effectively regulate cell cycle progression. It is perhaps not surprising therefore, that both cyclin D1 and cyclin E play important roles in the development and progression of cancer. This chapter explores the roles of these cyclins in cancer. For clarity, the major focus will be on the well-characterized roles of cyclin D1 and cyclin E and thus to a lesser extent on other G1 cyclins such as cyclins D2 and D3. This chapter summarizes the evidence linking cyclin D1 and cyclin E to cancer development and progression. The cellular pathways that underlie the deregulated expression of the G1 cyclins in cancer as well as their oncogenic roles are also discussed. Lastly, the potential of these cyclins to serve as therapeutic targets in cancer therapy is explored. Keywords Cell cycle • G1 phase • Cyclin D1 • Cyclin E • Cancer • Oncogene • Ubiquitin • 26 S proteasome • Cancer therapy

9.1 Introduction Cell cycle re-entry and progression through G1 is tightly controlled by the retinoblastoma tumor suppressor protein (pRB) [1, 2]. D-type cyclins form active kinase complexes with their cyclin dependent kinase partners 4 and 6 (CDK4/6) that phosphorylate and inactivate pRb. As a consequence, pRbsequestered E2F transcription factors are released and genes required for S phase entry are expressed [1, 3]. Cyclin D1 expression is absolutely dependent on mitogenic signalling and thus links the extracellular environment to the regulation of cell division [4]. Overcoming this dependence on mitogenic signalling was the first characteristic or hallmark of cancer cell proliferation to be clearly characterized [5]. It is therefore not surprising that the deregulated expression of G1 cyclins often plays a central role in oncogenesis. Several mechanisms underlie the deregulated expression of G1 cyclins in cancer cells. These include alterations such as gain of function mutations in upstream transducers of mitogenic signals, gene amplification, chromosomal translocations, deregulated splice variant expression, and impaired protein degradation. In the current chapter, the role of G1 cyclins in cancer cell proliferation will be discussed. Emphasis will be laid on the mechanisms that underlie deregulated G1 cyclin expression and the roles of these cyclins in cancer cell proliferation.


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9.2 Identification of Cyclin D1 as a Putative Proto-oncogene 9.2.1 Chromosomal Translocations Associated with Cyclin D1 Overexpression The cyclin D1 gene (CCND1) was originally identified as PRAD1 (parathyroid adenomatosis), a gene mapping adjacent to the chromosomal breakpoint in a parathyroid adenoma bearing a rearrangement of the parathyroid hormone (PTH) gene [6]. CCND1 was found to be highly overexpressed in this subset of adenomas, probably as a result of coming under the control of regulatory elements within the promoter of the PTH gene. Further studies identified CCND1 as a gene mapping close to the 11q13 locus, rearranged in the B-cell lymphoma associated with the t(11;14) translocation [7, 8]. In this study, CCND1 was found to be juxtaposed with the immunoglobulin locus on chromosome 14 in 70% of centrocytic or mantlezone B-cell lymphomas. As in subsets of parathyroid adenomas, cyclin D1 was also observed to be highly expressed in these lymphomas [8]. Further studies using transgenic mice expressing the CCND1 gene under the control of an immunoglobulin heavy chain enhancer (Eµ), provided further evidence that the cyclin is an oncogene [9, 10]. Eµ-CCND1 mice developed few spontaneous tumors in contrast to mice expressing both EµCCND1 and Eµ-myc transgenes. Interestingly, the levels of endogenous cyclin D1 were frequently found to be elevated in lymphomas from Eµ-myc mice. Similarly, cyclin D1 mRNA induction was frequently observed in cell lines derived from Eµ-myc mice and transformed with the raf or ras oncogenes [9]. Transgenic mice expressing Eµ-L-myc in both B- and T-cells develop mainly T-cell lymphomas after a long latency period. Co-expression of Eµ-CCND1 increased the incidence of tumor development and reduced the latency periods. The oncogenic effects of cyclin D1 were, however, restricted to B-lymphocytes. This observation suggested that cyclin D1 tumorigenic activity is dependent on both the presence of cooperating oncogenes and cell lineage [10]. In fact, earlier studies in rat fibroblasts had demonstrated

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a transforming potential for cyclin D1 only in the presence of Ha-ras [11]. Similar studies demonstrated that the ability of cyclin D1 to transform mouse or rat fibroblasts was dependent on the parental strain [12, 13]. Together, these studies suggested that cyclin D1 is a weak oncogene that can nevertheless, cooperate with other oncogenes to drive tumorigenesis.

9.2.2 Cyclin D1 Overexpression Promotes Tumor Development Transgenic mice have also been used to study the role of cyclin D1 in breast cancer. The expression of cyclin D1 under control of the mouse mammary tumor virus (MMTV) promoter induced abnormal mammary cell proliferation and the development of adenocarcinomas in this tissue [14]. Later studies demonstrated that cyclin D1 deficient mice expressing activated Her2/neu or ras were resistant to mammary tumors induced by these oncogenes [15]. Cyclin D1 ablation did not, however, protect against tumors induced by the myc or Wnt1 oncogenes. Cyclin D1 is overexpressed in a large percentage of breast cancers (see Sect. 3) [16–18]. These studies thus provided evidence to support a causal role for cyclin D1 in breast cancer development. It should be noted that cyclin D1 can cooperate with Myc in B-cell lymphoma but not breast cancer development [10, 15]. This further highlights the fact that the oncogenic potential of cyclin D1 depends on additional genomic changes and is cell type specific. Hence, cyclin D1 ablation does not suppress tumor development in other cell types [15, 19].

9.3 Impact of Deregulated Mitogenic Signaling on Cyclin D1 Expression 9.3.1 Mitogenic Signaling Pathways Converge on Cyclin D1 Several mitogenic signaling pathways have been shown to control cell division in part, by regulating cyclin D1 expression. In fact, the CCND1 promoter contains binding sites for several transcription

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factors (reviewed in [20–22]). Several regulators of cyclin D1 expression have themselves been identified as oncogenes. These include protein kinase B (AKT), Ras, Raf-1, Wnt/b-Catenin, the oestrogen alpha receptor (ERa), ERK1/ERK2, and the fibroblast growth factor receptor (FGFR) [23–27]. A thorough review of these pathways is beyond the scope of this chapter. For illustrative purposes, however, the impact of deregulated Wnt/b-catenin and ERa on cyclin D1 expression in cancer are discussed.

9.3.2 Deregulated Wnt Signaling and Cyclin D1 in Cancer The canonical Wnt/b-catenin pathway regulates the expression of target genes through b-catenin/Tcell factor (TCF) [28, 29]. Deregulated Wnt signaling has been implicated in the development of several cancers including those of the breast and colon [30–32]. Wnt-1 has been shown to transform murine epithelial cells and reduce the growth factor requirements of rodent fibroblasts, [32, 33] and furthermore, mice overexpressing Wnt-1 develop mammary carcinomas [15, 33, 34]. The binding of the Wnt ligand to its receptor, Frizzled, results in the inhibition of GSK3b and b-catenin stabilization [30, 35]. As a consequence, b-catenin accumulates within the nucleus, forms complexes with TCF and regulates the expression of target genes including CCND1 [36]. GSK3b also regulates cyclin D1 stability and subcellular localization at the posttranslational level (see Sect. 5.2) [23]. Wnt mediated GSK3b inhibition thus impacts on cyclin D1 expression at both the mRNA and protein levels. Overexpression of Wnt-1 in mouse fibroblasts resulted in decreased GSK3b activity and induced the accumulation of cyclin D1 [37]. Wnt-1 overexpression, however, only moderately induced cyclin D1 expression in this system. Consequently, coexpression of Wnt-1 with activated MEK1 (an upstream activator of ERK1/2) was required to facilitate mitogen independent S-phase entry [37]. Wnt-1 is thus most effective in promoting tumor formation in cells where it enhances CCND1 expression and cyclin D1 stability. It should also be noted, that cyclin D1 ablation did not prevent mammary tumor formation in mice overexpressing Wnt-1 [15]. The ability of Wnt-1 to effectively

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promote tumor development through cyclin D1 thus appears to be tissue specific [38].

9.3.3 Oestrogen Receptor Signaling and Cyclin D1 in Cancer Oestradiol (E2) is an important mediator of CCND1 expression in cells that express the oestrogen receptor alpha (ERa) [39]. Close to 70% of human breast cancers express ERa [40–42] and the antioestrogen tamoxifen exerts its antiproliferative effects in part by inhibiting ERa mediated CCND1 expression [43, 44]. The cyclin D1 promoter region contains several putative oestrogen responsive sites [36, 39, 45–50]. Studies by Sabbah et al. [50] demonstrated the role of the cyclic adenosine monophosphate (cAMP) response - like element within the CCND1 promoter in mediating the ERa induced expression of this gene. Further studies by Cicatiello et al. [51] using stably expressed reporter constructs identified an additional role for the AP1 site within the CCND1 promoter. More recent studies by Eeckhoute et al. [52] have indentified a novel E2 responsive and cell type specific enhancer downstream of the CCND1 coding region. This enhancer (enh2) is required for ERa recruitment to the CCND1 gene and facilitates E2 mediated cyclin D1 expression in breast cancer cells. Eeckhoute et al. also identified a role for the FoxA1 transcription factor in mediating ERa induced cyclin D1 expression. Together, these studies show that E2 mediates cyclin D1 expression through complex interactions with regulatory sequences proximal and distal to the CCND1 open reading frame. ERa is thus an important regulator of cyclin D1 expression and hence cell proliferation. Indeed, small interfering RNA (siRNA) mediated knockdown of ERa resulted in the downregulation of cyclin D1 expression in MCF-7 breast cancer cells. Furthermore, the siRNA mediated knockdown of ERa inhibited MCF-7 cell proliferation to the same degree as the siRNA mediated knockdown of cyclin D1 expression [53]. Many of the signaling pathways activated in cancer regulate cell proliferation at least partly, by regulating the cellular levels of cyclin D1. The deregulated activity of mitogenic signaling pathways is thus likely to underlie cyclin D1 accumulation in some cancers.

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9.4 CCND1 amplification in cancer 9.4.1 CCND1 Amplification is Common in Cancer CCND1 maps to chromosome 11q13, a region that is frequently amplified in human cancers [54–57]. Faust and Meeker [58] observed CCND1 amplification in approximately 48% of human cancer cell lines investigated. Although cyclin D1 expression was detected in all samples, amplification of CCND1 showed only a partial correlation with cyclin D1 overexpression in these studies. Further studies by Zhang et al. [59] detected CCND1 amplification in 13% of samples from patients with hepatocellular carcinoma (HCC). Immunohistochemical analyses demonstrated a close link between CCND1 amplification and cyclin D1 overexpression. Studies by Jares et al. [60] also detected CCND1 amplification in 37% of human laryngeal carcinoma cases. CCND1 amplification was associated with advanced local invasion, lymph node metastases and advanced disease stage. Furthermore, cyclin D1 mRNA overexpression was detected in 35% of the cases examined. CCND1 amplification was also strongly associated with increased mRNA expression. Indeed, the degree of gene amplification was closely correlated with the levels of mRNA expression. Cyclin D1 overexpression has also been linked to CCND1 amplification in lung cancer [61].

9.4.2 Importance of CCND1 Amplification in Breast Cancer CCND1 amplification has also been reported in breast cancer by numerous investigators ([62] and references therein). These studies show a variation in the frequency of CCND1 amplification ranging from 9 to 24%. In a recent study using chromogenic in situ hybridization (CISH) and a well characterized rabbit monoclonal antibody, Reis-Filho et al. [62] detected cyclin D1 overexpression in 67% of cases and CCND1 amplification in 15% of cases. Furthermore, there was a strong association between CCND1 amplification and cyclin D1 overexpression. Cyclin D1 expression was also associated with oestrogen and progesterone receptor expression. Cyclin D1 expression

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and CCND1 amplification was not detected (or showed an inverse correlation) in breast cancers with basal-like phenotypes or in tumor samples from BRCA1 carriers [62, 63]. Since tumors from BRCA1 carriers frequently display a basal-like phenotype, Reis-Filho et al. have suggested that cyclin D1 does not play a significant role in the biology of these cancers [62]. A more recent study by Elsheikh et al. [64] detected CCND1 amplification in 10% and cyclin D1 overexpression in 44% of breast cancer samples examined. Cyclin D1 overexpression was positively correlated to CCND1 amplification. Both CCND1 amplification and cyclin D1 overexpression were associated with oestrogen receptor expression. The study by Elsheikh et al. also strengthened previous findings, which have suggested a limited role for cyclin D1 in the development of basal-like cancers [62–64]. The strong correlation between CCND1 amplification and cyclin D1 overexpression demonstrates the importance of gene amplification in mediating the deregulated expression of this cyclin. These studies also indicate, however, that CCND1 amplification occurs less frequently than cyclin D1 overexpression. Additional mechanisms must thus facilitate cyclin D1 overexpression in the absence of CCND1 amplification. As noted in the previous section, E2 can drive cell cycle progression in part by inducing cyclin D1 expression in oestrogen receptor (ER) positive breast cancer cells. The strong association between ER expression and cyclin D1 overexpression thus provides further evidence for the importance of this pathway in some breast cancers. The study by Reis-Filho et al. [62] did not reveal a significant correlation between CCND1 amplification and ER expression. In their study, CCND1 amplification was detected in only 19% of the ER positive cancer samples. In contrast, Elsheikh et al. [64] found that roughly 83% of the samples with concomitant CCND1 amplification and cyclin D1 overexpression were ER positive. The reasons for this discrepancy remain to be determined. The study by Reis-Filho et al. [62] suggests that ER overexpression may be sufficient to drive cyclin D1 overexpression in the absence of CCND1 amplification. Although the ER regulates cyclin D1 expression at the transcriptional level [50–52], cyclin D1 also regulates ER transcriptional activity [65, 66] (see Sect. 7). These findings may explain the high percentage of ER


positive tumors with both CCND1 amplification and cyclin D1 overexpression reported by Elsheikh et al. [64]. Based on their findings, Elsheikh et al. [64] have suggested that CCND1 amplification may confer a proliferative advantage to ER positive breast cancer cells. The nuclear accumulation of cyclin D1 has recently been shown to enhance genomic instability [67] (see Sect. 7). It remains to be determined, however, if CCND1 amplification precedes ER overexpression. It is also unclear as to how concomitant ER expression and CCND1 amplification as opposed to ER expression in the absence of CCND1 amplification impacts on the biology of breast cancer cells.

9.4.3 CCND1 Amplification Plays a Limited Role in Mediating Cyclin D1 Overexpression The studies discussed above demonstrate that CCND1 amplification often underlies the elevated levels of cyclin D1 protein detected in cancer cells. It is clear, however, that cyclin D1 overexpression can be frequently detected in the absence of CCND1 amplification. As noted in Sect. 3, the deregulated activity of mitogenic signaling pathways might be sufficient to induce cyclin D1 overexpression in the absence of gene amplification. The posttranslational regulation of cyclin D1 has also been shown to be important for regulating the cellular levels of cyclin D1 [68]. In fact, deregulated cyclin D1 degradation via the ubiquitindependent degradation pathway is now known to be responsible for the elevated levels of cyclin D1 frequently observed in cancer. The importance of this pathway in regulating cyclin D1 in normal and cancer cells is discussed below.

9.5 Deregulated cyclin D1 degradation is common in cancer 9.5.1 Cyclin D1 Degradation During the Normal Cell Cycle Cyclin D1 is an intrinsically unstable protein with a half-life of less than 30 min [68]. Serum withdrawal or the inhibition of protein synthesis thus leads to a rapid decline in cyclin D1 protein lev-

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els [4, 69, 70]. The rapid degradation of cyclin D1 occurs at the G1/S phase boundary during the normal cell cycle [4]. The loss of cyclin D1 expression following the inhibition of mitogenic signaling thus serves to link cell proliferation with the extracellular environment. Cyclin D1 degradation also coincides with its nuclear export during its entry into S phase [4]. The overexpression of cyclin D1 prevented S phase entry and proliferating cell nuclear antigen (PCNA) function following DNA damage in human fibroblasts [71]. Further studies demonstrated that overexpressed cyclin D1 inhibits cell proliferation by binding to and inhibiting the activity of both PCNA and cyclin dependent kinase 2 (CDK2). The inhibitory effect of cyclin D1 on cell proliferation was observed in both normal human fibroblast and transformed cell lines [72]. Cyclin D1 degradation at the S phase boundary thus appears to be necessary for proper DNA replication [72, 73]. In fact, the generation of stable clones expressing a cyclin D1 mutant that cannot be efficiently degraded has been reported to be difficult [73].

9.5.2 Phosphorylation-Dependent Cyclin D1 Degradation Early studies by Diehl et al. [68] demonstrated that cyclin D1 is targeted for degradation via the ubiquitin dependent degradation pathway. Phosphorylation of threonine residue 286 (T286) was shown to greatly enhance the degradation of cyclin D1 via this pathway. Mutation of T286 to alanine (T268A) extended the half life of the cyclin from approximately 30 min to 3.5 h. Binding of cyclin D1 to CDK4 enhanced T286 phosphorylation but was not dependent on the kinase activity of CDK4, suggesting the involvement of an additional kinase [68]. Further studies identified glycogen synthase kinase 3b (GSK3b) as the kinase responsible for mediating T286 phosphorylation [23]. Purified GSK3b phosphorylated cyclin D1 specifically on T286 in vitro and depletion of the kinase from mammalian cell extracts prevented T286 phosphorylation. Similarly, overexpression of a kinase dead GSK3b mutant significantly inhibited the phosphorylation of cyclin D1 by endogenous kinases in insect cells or by ectopically expressed wild type GSK3b. In addition, GSK3b was also

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shown to play a role in regulating the subcellular localization of cyclin D1, as overexpression of this kinase induced the nuclear export of cyclin D1. GSK3b was also observed to transiently accumulate within the nucleus at the S phase boundary, coinciding with cyclin D1 nuclear export and degradation [23]. As noted in Sect. 3, the mitogen activated Ras signaling pathway has been shown to induce cyclin D1 expression [74–76]. Ras can also induce Akt activation via PI3K resulting in the inhibition of GSK3b by Akt [77]. Deregulated Ras signaling thus induces cyclin D1 overexpression by enhancing CCND1 activation and preventing the phosphorylation dependent degradation of the cyclin [23]. Overexpression of constitutively active Akt was observed to prolong the halflife of cyclin D1 while inhibition of PI3K induced the increased turnover of the cyclin [23]. Given its role in regulating GSK3b, deregulated Wnt signalling can be expected to facilitate cyclin D1 accumulation through similar mechanisms [23, 30]. Further studies demonstrated that the GSK3b dependent phosphorylation of T286 facilitates the nuclear export of cyclin D1 by enhancing its interaction with CRM1 [78]. Overexpression of CRM1 induced the cytoplasmic redistribution of wild type but not T286A mutant cyclin D1. Inhibition of CRM1 with leptomycin B (LMB) or GSK3b with lithium chloride (LiCl) prevented the nuclear export of cyclin D1 at S phase. Similarly, treatment with LMB prevented the cytoplasmic redistribution of cyclin D1 in cell overexpressing GSK3b. CRM1 thus facilitates the nuclear export of T286 phosphorylated cyclin D1 in a GSK3b dependent manner. Mutation of T286 to alanine thus greatly enhances the stability of cyclin D1 and results in constitutively nuclear localization throughout the cell cycle [23, 78]. Although T286 clearly plays a role in regulating the stability and subcellular localization of cyclin D1, recent studies have questioned the importance of GSK3b in mediating the phosphorylation of this residue. Guo et al. [73] confirmed the importance of T286 phosphorylation in mediating cyclin D1 degradation and observed a clear increase in the levels of phosphorylated cyclin D1 at S phase. These authors did not, however, detect changes in the activity of PI3K, Akt or GSK3b during S phase. Furthermore, inhibition of GSK3b did not influence cyclin D1 phosphorylation or stability in

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these studies. Overexpression of constitutively active GSK3b similarly failed to affect cyclin D1 phosphorylation or stability [73, 79]. Other studies suggest that GSK3b mediates the regulation of cyclin D1 stability in cancer cells. Inhibition of GSK3b induced the nuclear accumulation of cyclin D1 in the MCF-7 human breast cancer cell line. The subcellular loclization of GSK3b was not, however, observed to vary with cell cycle progression in these cells [69]. GSK3b has also been shown to facilitate rapamycin mediated cyclin D1 degradation in human breast cancer cell lines [80]. Furthermore, GSK3b has been shown to enhance differentiation-inducing factor-1 (DIF-1) and differentiation-inducing factor-3 (DIF-3) induced cyclin D1 degradation in cell lines derived from human tongue squamous cell carcinomas (NA and SAS) and a human cervical cancer cell line (HeLa) [81, 82]. More recently, the GSK3b dependent phosphorylation was shown to facilitate the hydroxyurea (HU) induced degradation of cyclin D1 in mammalian cell lines. Accordingly, mutation of cyclin D1 T286 to alanine or the siRNA mediated knockdown of GSK3b in HuH7 human hepatoma cells prevented HU induced cyclin D1 degradation [83]. The dimerization of F-box proteins has been shown to be necessary for the formation of functional SCF complexes ([84] and references therein). Interestingly, GSK3b has recently been shown to mediate Fbx4 dimerization by phosphorylating serine residue 12 (S12). GSK3b can thus regulate the stability of cyclin D1 by directly phosphorylating T286 and by facilitating the phosphorylation dependent dimerization of its E3 ligase Fbx4 [85] (see Sect. 5.5.3). Together these studies suggest that GSK3b plays an important role in mediating cyclin D1 T286 phosphorylation and its subsequent nuclear export and degradation by 26 S proteasomes. The studies by Guo et al. and Yang et al. [73, 79] suggest that GSK3b mediated cyclin D1 phosphorylation may be cell lineage specific. Alternatively, GSK3b may regulate cyclin D1 stability in a general manner throughout the cell cycle and not specifically during S phase. Further studies will be needed to determine with certainty, the exact role(s) that GSK3 plays in the regulation of cyclin D1 stability. In recent years, additional kinases that mediate cyclin D1 degradation have been identified. Furthermore, the phosphorylation and indeed ubiquitin independent


degradation of cyclin D1 has also been shown to occur (Fig. 9.1). These pathways are discussed in the following two sections.

9.5.3 GSK3b-Independent Cyclin D1 Phosphorylation 9.5.3.1 Mirk/Dyrk1B Mirk/Dyrk1B is an arginine directed kinase that has been shown to mediate cyclin D1 degradation by phosphorylating threonine residue 288 (T288) [86]. Overexpression of Mirk in colon cancer cell lines induced T288 phosphorylation and the rapid turnover of cyclin D1. In contrast, knockdown of Mirk expression by siRNA induced the accumulation of cyclin D1 [87, 88]. Mirk activity is restricted to the G0 and early G1 phases of the cell cycle, suggesting that it is not required for cyclin D1 degradation at the S phase boundary [88, 89]. Mirk1 affects numerous cellular processes but its expression is elevated in skeletal muscle. Mirk may thus exert its effects on cellular physiology in a cell type specific manner [86, 90]. Overexpression of Mirk delayed cell cycle entry in mink lung epithelial cells (Mv1Lu), suggesting that it enforces G0 arrest by destabilizing cyclin D1 [86, 88]. Mirk has also been shown to cooperate with GSK3b to enhance DIF-3 induced cyclin D1 degradation in a human cervical cancer cell line HeLa [82]. These studies suggest that in some cell lines, GSK3b and Mirk1 function together to mediate the rapid turnover of cyclin D1.

9.5.3.2 p38SAPK2a The stress activated protein kinase 2a (SAPK2a, p38, p38a, p38SAPK2a) has also been shown to phosphorylate cyclin D1 on T286, resulting in the increased turnover of the cyclin. p38SAPK2a is activated in response to diverse environmental and cellular stresses [91–94]. Early studies demonstrated that p38SAPK2a inhibits cyclin D1 expression in the chinese hamster lung fibroblast cell line CCL39 [26]. Studies by Casanovas et al. [91] show, however, that p38SAPK2a phosphorylates cyclin D1 in response to various environmental stresses and thus enhances its degradation via the ubiquitin pathway. Using the Granta 519 cell line (derived from a high grade nonHodgkin’s lymphoma patient [95]), these inves-

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tigators demonstrated that osmotic and oxidative stress as well as sodium arsenite induced the rapid degradation of cyclin D1 via the ubiquitin dependent degradation pathway. Inhibition of p38SAPK2a activity abolished the osmotic stress but not oxidative stress or arsenite induced degradation of cyclin D1. The p38SAPK2a kinase thus specifically targets cyclin D1 for ubiquitin dependent degradation in response to osmotic stress (50 mM sodium chloride (NaCl)). In vitro studies further demonstrated that p38SAPK2a specifically phosphorylates cyclin D1. p38SAPK2a dependent cyclin D1 phosphorylation was inhibited by the p38SAPK2a specific inhibitor SB203580 [91]. Mutation of T286 to alanine abolished p38SAPK2a mediated cyclin D1 phosphorylation and prevented its ubiquitylation in vitro [91]. Both aspirin and its active component sodium salicylate have been shown to activate p38SAPK2a in mammalian cells [96, 97]. Aspirin also induced the rapid degradation of cyclin D1 via the ubiquitin dependent degradation pathway [97]. Inhibition of p38SAPK2a with a chemical inhibitor (PD169316) or knockdown of its expression by siRNA partially abolished aspirin induced cyclin D1 degradation. These observations suggest that aspirin induces cyclin D1 degradation in a p38SAPK2a dependent manner [97]. p38SAPK2a also mediates cellular responses to ultraviolet (UV) radiation [98, 99]. Interestingly, UV radiation has been shown to induce the rapid degradation of cyclin D1 mRNA and protein levels in the Bac1.2F5 mouse macrophage cell line [100]. Moreover, the activation of p38SAPK2a has been shown to repress CCND1 expression and target cyclin D1 for ubiquitin dependent proteasomal degradation [26, 91, 101]. UV radiation may thus induce cyclin D1 degradation in a p38SAPK2a dependent manner.

9.5.3.3 ATM and ATR The ataxia telangiectasia mutated (ATM) and ATM and Rad3 related (ATR) kinases play essential roles in mediating cellular responses in response to DNA damage. Furthermore, recent findings indicate that ATM is also involved in the regulation of cellular responses to osmotic and oxidative stresses [102–104]. Studies by Hitomi et al. [105] have demonstrated that both ATM and ATR can induce the phosphorylation of cyclin D1 on amino

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Figure 9.1. Regulation of cyclin D1 degradation. (a). Cyclin D1 does not contain a nuclear localization signal (NLS) and its sequestration may result in accumulation within the cytoplasm. (b). Cytoplasmic cyclin D1 is transported into the nucleus in association with its binding partners e.g. CDK4 and possibly various transcription


residue T286. ATR is activated in S phase during normal cell division and is specifically activated by UV during this period ([105] and references therein). Accordingly, ATR induced cyclin D1 T286 phosphorylation in response to UV exposure predominantly during S phase. The siRNA directed knockdown of ATR attenuated the UV induced phosphorylation of cyclin D1 T286. Interestingly, knockdown of ATR also inhibited T286 phosphorylation in normally dividing cells. ATR thus induces cyclin D1 phosphorylation on T286 during both the normal cell cycle and following UV induced DNA damage. Since ATR is activated in response to normal DNA replication ([105] and references therein), Hitomi et al. suggest that ATM may be the kinase that directs cyclin D1 phosphorylation and subsequent degradation during this period. The study by Hitomi et al. [105] also demonstrated that ATM phosphorylates cyclin D1 on T286 in response to DNA double strand breaks. It remains unclear, however, if the ATR and ATM kinases directly phosphorylate cyclin D1 on T286. ATM and ATR thus mediate G1 cell cycle arrest in part, by inducing cyclin D1 phosphorylation and increasing its rate of degradation via the ubiquitin pathway. ATR mediated phosphorylation and hence cyclin D1 degradation may also be necessary to allow efficient DNA repair and progression through S phase [71, 73, 79, 105].

9.5.3.4 IkB Kinase a The IkB kinase a (IKKa) is a component of the IKK complex and activates NF-kB by phospho-

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rylating (and thus targeting for degradation) the NF-kB inhibitory protein IkB [106–108]. IKKa also regulates cellular processes independently of its role in facilitating NF-kB [109, 110]. Studies by Kwak et al. [111] have also identified IKKa as a regulator of cyclin D1 subcellular localization and stability. Cyclin D1 expression is elevated and exhibits an exclusively nuclear localization pattern in proliferating IKKa−/− primary mouse embryonic fibroblast (MEF) cells. In contrast, cyclin D1 localization was predominantly cytoplasmic in wild type (WT), IKKb−/− and IKKa+/+ reconstituted MEF cells, indicating a role for this kinase in regulating cyclin D1 localization. IKKa interacts with cyclin D1 in vivo and in vitro and phosphorylates T286. Furthermore, the levels of phosphorylated cyclin D1 were substantially reduced in IKKa−/− cells and this was associated with a fourfold increase in the halflife of the cyclin [111]. IKKa thus regulates cyclin D1 localization and stability by phosphorylating T286 within the cyclin [111]. The siRNA mediated knockdown of IKKa in HeLa cells similarly resulted in cyclin D1 stabilization and a predominantly nuclear localization pattern. In addition, pRb phosphorylation was substantially reduced in these cells. IKKa mediated cyclin D1 degradation thus impacts directly on the cyclin’s ability to regulate G1 progression in terms of pRb phosphorylation [2, 111]. A more recent study indicates that IKKa also plays a role in the regulation of cyclin D1 activity in LNCaP prostate cancer cells [111] (see Sect. 5.5.5). IKKa mediated regulation of cyclin D1 is therefore not restricted to MEFs

Figure 9.1. (continued) factors (TF). (c). p38SAPK2a has been shown to phosphorylate cyclin D1 on threonine residue 286 (T286) and induce its proteasomal degradation. It is unclear if the F-box proteins FBX4 and FBXW8 are involved in mediating p38SAPK2 induced cyclin D1 degradation in the cytoplasm. (D, E). Within the nucleus, active cyclin dependent kinase 4 (CDK4)- or CDK6-cyclin D1 complexes phosphorylate the retinoblastoma protein (RB). Cyclin D1 can also influence the activity of various transcription factors independently of CDK4/ 6. (F). Free cyclin D1 is degraded through the ubiquitin dependent 26 S proteasomal degradation pathway independently of phosphorylation on T286 and/ or T288. Antizyme mediates cyclin D1 degradation via the 26 S proteasome independently of ubiquitin. Some studies suggest that cyclin D1 may also be targeted for proteasome independent degradation. (g, h). GSK3b phosphorylates cyclin D1 on T286 which facilitates its nuclear export by the CRM1. GSK3b influences cyclin D1 stability since the phosphorylated form of the cyclin is subsequently degraded within the cytoplasm. Additional kinases known to phosphorylate cyclin D1 on T286 are ATR and ERK1/ 2. ATM phosphorylates T286 following exposure to genotoxic agents. (i). Phosphorylation of T288 is mediated by the mirk/Dyrk 1b kinase and can induce cyclin D1 degradation. (j). IKKa regulates cyclin D1 localization and stability by phosphorylating T286. (k). FBX4, FBXW8, and b-TrCP ubiquitylate phosphorylated cyclin D1 within the cytoplasm, targeting it for 26 S proteasomal degradation.

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and can contribute to the regulation of cyclin D1 activity in cancer cells.

9.5.4 Phosphorylation- and Ubiquitylation-Independent Degradation of Cyclin D1 9.5.4.1 Phosphorylation-Independent Cyclin D1 Degradation Although phosphorylation of cyclin D1 T826 (and to a lesser extent T288) greatly enhances the rate of its turnover, cyclin D1 ubiquitylation and degradation can nonetheless occur independently of phosphorylation [113]. Mutation of human cyclin D1 T286 and T288 to alanine resulted in a threefold increase in the half life of the cyclin in U2OS human osteosarcoma cells. These mutations did not abolish the degradation completely as low levels of cyclin D1 proteolysis were still observable. Coexpression of CDK4 however, abolished the low level of cyclin D1 degradation. These observations suggested that the degradation of “free” i.e. nonCDK4 bound cyclin D1 can occur independently of T286 and/ or T288 phosphorylation [113]. Furthermore, low levels of cyclin D1 ubiquitylation could be detected in the T286A/T288A mutant suggesting that “free” cyclin D1 can be degraded independently of phosphorylation via the ubiquitin dependent degradation pathway. Cyclin D1 was also effectively ubiquitylated in vitro, independently of T286 and T288 phosphorylation. Phosphorylation independent degradation may provide a second degradation pathway that regulates cellular levels of nonCDK4 bound cyclin D1 [113]. The rapid phosphorylation independent degradation of cyclin D1 in several mammalian cell lines, following cellular exposure to genotoxic agents has also been reported [114]. The Cyclin D1 T286A mutant was effectively degraded following exposure to ionizing radiation (IR). Furthermore, the culture of MCF-7 breast cancer cells with LiCl (a GSK3b inhibitor) did not prevent IR induced cyclin D1 degradation. T286 phosphorylation is therefore not required for the rapid induction of cyclin D1 degradation following exposure to IR [114]. These studies also demonstrated a requirement for a conserved RxxL destruction box within the N-terminal region of cyclin D1. Cyclin D1 ubiquitylation in response to IR was also shown to

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be dependent on the anaphase promoting complex (APC). Interestingly, mutation of the cyclin D1 destruction box (Fig. 9.2) prevented IR induced cyclin D1 degradation but did not result in the accumulation of the cyclin in nonirradiated cells. APC mediated cyclin D1 destruction following exposure to IR thus represents a distinct degradation pathway. Taken together with the findings of Hitomi et al. [105] (Sect. 5.3.3), the findings by Agami and Bernards [114] suggest that DNA damage triggers the ubiquitin dependent degradation of cyclin D1 both dependent and independent of T286 phosphorylation. It is currently unclear if these pathways operate simultaneously following exposure to genotoxic agents or in a cell and context dependent manner. The rapid degradation of cyclin D1 in response to DNA damage is necessary, however, to facilitate rapid cell cycle arrest in G1 following DNA damage and occurs independently of T286 phosphorylation in MCF-7 cells [114].

9.5.4.2 Ubiquitin-Independent Cyclin D1 Degradation Antizyme targets ornithine decarboxylase (ODC) for ubiquitin independent degradation via the 26 S proteasome [115, 116]. Initial studies demonstrated that antizyme induction in hamster malignant oral keratinocytes (HCPC-1) and Dunning rat prostate carcinoma cell lines (AT2.1, AT3.1, MatLyLu, Nbe, FB2) results in G1 arrest [117]. Subsequent studies by Newman et al. [118] demonstrated that antizyme targets cyclin D1 for proteasomal degradation. Antizyme interacts with cyclin D1 both in vitro and in vivo. Mutation of T286 to alanine did not prevent antizyme mediated cyclin D1 degradation which interestingly, also occurs independently of ubiquitin [118]. Together these studies suggest that antizyme induction induces G1 arrest by targeting cyclin D1 for ubiquitin independent degradation via the 26 S proteasome. The precise role of antizyme in regulating normal cell cycle progression is currently unclear. One possibility is that antizyme functions as a tumor suppressor by contributing towards the regulation of cellular cyclin D1 levels [118]. Additional studies in mantle cell lymphoma (MCL) cells have also provided evidence suggesting that cyclin D1 degradation can occur independently of 26 S proteasomes. Studies using the Jeko1 mantle cell lymphoma (MCL)

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9. G1 Phase Cyclins in Cancer Development and Progression Mirk/ Dyrk1b ATM, ATR, GSK3b, ERK1/ 2,

?

IKKa, p38SAPK2

APC

**

NH2

NH2

COOH

COOH

5-9 = RB interacting site

29-32 = RxxL motif

56-145 = Cyclin box

179-193 = D-domain

241-290 = PEST motif

251-255 = LxxLL motif

241-275 = Intron 4 sequence

279-286 = TrCP binding motif

**

= T286, T288

Figure 9.2. Schematic representation of cyclin D1 (top) and cyclin D1b (bottom) regulatory sequences. See text for details. Adapted from Knudsen et al. [184] and Alao [209].

line demonstrated that the specific inhibition of 26 S proteasomal activity by PS431 (bortezomib/ Velcade) [119] resulted in a decline in cyclin D1 protein levels [120]. Similarly, lactacystin (a specific proteasome inhibitor) [121] failed to prevent the actinomycin D and lovastatin induced loss of cyclin D1 expression in PC-3-M prostate cancer cells [70]. Lactacystin also failed to block the loss of cyclin D1 expression in MCF-7 cells treated with cycloheximide [122]. The studies by Choi et al. suggested a possible role for calpains in regulating cyclin D1 stability [70]. In the absence of further studies, however, the role of these proteases in regulating cyclin D1 stability remains unclear. More recent studies by Feng et al. [123] have demonstrated that cyclin D1 variants, which cannot be ubiquitylated, are nevertheless still subject to proteolysis and do not accumulate significantly in vivo. Together, these studies indicate that cyclin D1 degradation can occur independently of ubiquitin and indeed independently of the 26 S proteasome. Antizyme clearly facilitates cyclin D1 proteasomal degradation independently of ubiquitin. In fact, other proteins that are targeted for ubiquitin independent proteasomal degradation have been identified [124]. The pathway(s) that regulate cyclin D1 degradation independently of proteasomes have yet to be characterized. The studies by Choi et al. [70] and others [120, 122], however, provide compelling evidence that one or more such pathways exist.

9.5.5 Role of SCF Complexes in Regulating Cyclin D1 Stability 9.5.5.1 Cyclin D1 Interacts with SCF Complexes Early studies by Yu et al. [125] indicated that cyclin D1 and p21 interacted with cullin 1 (Cul1) in a human keratinocyte cell line. Cullins interact with Skp1, Rbx1 and one of several F-box proteins to form E3 ligase complexes known as Skp1Cullin-F-box (SCF) complexes. Particular SCF complexes specifically catalyse the ubiquitylation of one or more substrate proteins, targeting them for ubiquitin dependent 26 S proteasomal degradation. F-box proteins recognize specific subsets of target proteins and thus function to confer substrate specificity to SCF complexes [126–128]. Antisense mediated suppression of Skp1, Skp2/p45 (see Sect. 5.5.2) and Cul1 induced the accumulation of cyclin D1 and p21 in the HaCat human keratinocyte and DLD1 human colon adenocarcinoma cell lines. Inhibition of 26 S proteasomal activity similarly induced the accumulation of cyclin D1 and p21 in HaCat and RKO human colorectal carcinoma cells [125]. Subsequent studies by Russell et al. [129] detected the elevated expression of cyclin D1 and cyclin D3 in primary breast cancer samples. Elevated cyclin D1 expression was not strictly correlated with increased mRNA expression, confirming previous findings that cyclin D1 protein levels can accumulate despite the lack of increased

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mRNA expression. Similar experiments detected increased cyclin D1 protein expression without an accompanying increase in mRNA levels in several cancer cell lines [129]. The study by Russell et al. [129] also confirmed previous findings demonstrating an interaction between cyclin D1 and Cul1, and the involvement of the ubiquitin dependent degradation pathway [125]. Similar observations were also made regarding the regulation of cyclin D3 stability [129]. Together, these findings suggested that SCF complexes target cyclin D1 for ubiquitin dependent degradation.

9.5.5.2 Skp2/p45 The initial observation that the suppression of Skp2 expression induced the accumulation of cyclin D1, suggested that the F-box protein targets this cyclin for 26 S proteasomal degradation. In their study, the suppression of Skp2 expression also induced the accumulation of p21 but not p27 [125]. Subsequent studies in the SK-UT-1B uterine sarcoma cell line appeared to provide further evidence that Skp2 mediates cyclin D1 degradation [130]. SK-UT-1B cells are defective in targeting cyclin D1 and p27 for ubiquitin dependent degradation [113, 130, 131]. Ganiatsas et al. [130] identified a novel Skp2 splice variant (Skp2-CTV) that fails to efficiently bind to Skp1 and is constitutively cytoplasmic in contrast to Skp2, which is predominantly nuclear. The expression of wild type Skp2 in SK-UT-1B cells also leads to a reduction in the level and halflife of both the cyclin D1 and p27 proteins. It should be noted that neither the study by Yu et al. [125] nor that of Russell et al. [129] demonstrated a direct interaction between Skp2 and cyclin D1, or the ability of this F-box protein to directly catalyse cyclin D1 ubiquitylation in vitro. More recent studies also suggest that Skp2 is not a bona fide cyclin D1 E3 ligase (see Sects. 5.5.3–5.5.5). Skp2 is now known to directly target p27 [132]. The observation by Yu et al. [125] that the suppression of Skp2 expression did not induce the accumulation of p27 is therefore surprising. Furthermore, the siRNA mediated knockdown of Skp2 suppressed cyclin D1 expression in HeLa cells [133]. Since cyclin D1 exists in complexes with p21 and p27 [134] and can be ubiquitylated on multiple lysine residues [123], the possibility remains that Skp2 indirectly catalyses the ubiquitylation of cyclin D1.

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9.5.5.3 Fbx4 The SCFFbx4-aB crystallin complex has been identified as a bona fide E3 ligase for cyclin D1 [135]. aB crystallin was identified as a cyclin D1 interacting protein by affinity chromatography and co-immunoprecipitation experiments. The interaction of aB crystallin with cyclin D1 was dependent on T286 phosphorylation since the mutation of this residue to alanine abolished the interaction between the two proteins. aB crystallin interacts with the Fbx4 F-box protein [136] and further experiments demonstrated that both aB crystallin and Fbx4 interact with cyclin D1 in vivo. In vitro studies demonstrated that Fbx4 interacts specifically with T286 phosphorylated cyclin D1 in a aB crystallin dependent manner [135]. In this study, phosphorylated cyclin D1 peptides interacted specifically with Fbx4 but not with the F-box proteins Fbw2 or Skp2. Expression of dominant negative Fbx4 constructs doubled the halflife and induced the accumulation of cyclin D1 (but not the T286A mutant) in a aB crystallin dependent manner. In addition, the expression of dominant negative Fbx4 induced a decline in the cellular levels of polyubiquitylated cyclin D1. Similarly, short-hairpin RNA (shRNA) directed knockdown of Fbx4 or aB crystallin extended the halflife of cyclin D1 but did not affect the stability of cyclin A or cyclin E. The shRNA directed knockdown of Fbx4 or aB crystallin also induced a decline in the cellular levels of polyubiquitylated cyclin D1. Conversely, the overexpression of FBX4 or aB crystallin induced a phosphorylation and proteasome dependent decrease in cyclin D1 levels [135]. The study by Lin et al. [135] also demonstrated that purified SCFFbx4-aB crystallin complexes can catalyse the phosphorylation dependent ubiquitylation of cyclin D1 in vitro. Additional experiments demonstrated that both Fbx4 and aB crystallin are predominantly cytoplasmic proteins that show increased binding to phosphorylated cyclin D1 during S phase. As discussed in Sects. 5.1 and 5.2, cyclin D1 has been shown to undergo increased phosphorylation, nuclear export, and degradation during S phase. Furthermore, the inhibition of Fbx4 activity accelerated progression through G1 in wild type but not cyclin D1−/− NIH 3 T3 cells. Interestingly, the expression of both Fbx4 and aB crystallin is often downregulated in a wide range of tumors and cancer cell lines and


is frequently accompanied by increased cyclin D1 levels [135]. Re-expression of aB crystallin in MCF-7 cells substantially reduced the halflife of cyclin D1 in this cell line. The findings of Lin et al. [135] thus identified Fbx4 as the first bona fide cyclin D1 E3 ligase.

9.5.5.4 Fbxw8 Additional studies have identified the Fbxw8 F-box protein as a bona fide cyclin D1 E3 ligase [137]. Co-immunoprecipitation experiments identified Fbxw8 and Fbxl12 (but not b-TrCP, Fbxw5, Fbxw7, Fbxw11, Fbxl5 or Skp2) as F-box proteins that interact with cyclin D1. Maximal binding of Fbxw8 but not Fbxl12 was dependent on T286 phosphorylation. However, only Fbxw8 was able to catalyse the polyubiquitylation of cyclin D1 in vitro in a phosphorylation dependent manner [137]. Fbxw8 catalyzed cyclin D1 ubiquitylation as part of SCF complexes containing either Cul1 or Cul7. Interestingly, Okabe et al. also identified a role for the Ras/ Raf/MEK/ERK MAPK pathway in regulating cyclin D1 T286 phosphorylation in cancer cell lines. Purified ERK2 phosphorylated “free” and CDK4/6 bound cyclin D1 on T286 in vitro. The ability of ERK2 to phosphorylate T286 was dependent on a conserved kinase docking site (D-domain) spanning residues 179–193 within cyclin D1 (Fig. 9.2) [137, 138]. Furthermore, ERK2 enhanced cyclin D1 polyubiquitylation in vitro, in a T286 and D-domain dependent manner. In vivo, chemical inhibition of ERK2 in HCT116 cells with U0126 resulted in a rapid decline of T286 phosphorylated cyclin D1 and increased the halflife of the cyclin by approximately twofold [137]. Overexpression of Fbxw8 in HCT116 and other cancer cell lines induced a decline in the levels of cyclin D1 but not cyclin E. In contrast, expression of a dominant negative Fbxw8 construct induced the accumulation of cyclin D1. Similarly, the siRNA mediated knockdown of Cul1, Cul7 or Fbxw8 induced the accumulation of T286 phosphorylated cyclin D1 in various cancer cell lines. The knockdown of these SCF components also extended the halflife of cyclin D1 to varying degrees [137]. Together, these findings demonstrate that Fbxw8 targets cyclin D1 for ubiquitin dependent degradation.

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9.5.5.5 b-TrCP The b-transducin repeat-containing protein (b-TrCP) is a well characterized F-box protein that controls various cellular processes by targeting multiple substrates for ubiquitin dependent degradation [139]. Activators of the peroxisome proliferator activated receptor g (PPARg) have been shown to target cyclin D1 for ubiquitin-dependent proteasomal degradation in human cancer cell lines [140–142]. Interestingly, the effect of these agents has been shown to be independent of their PPARg agonist activity and has allowed for the development of derivatives that do not target this receptor [140]. Recent findings indicate that PPARg agonists and their derivatives target cyclin D1 for degradation in a b-TrCP dependent manner [111]. The PPARg derivative, STG28, has been shown to induce cyclin D1 T286 phosphorylation, nuclear export and ubiquitin dependent degradation independently of GSK3 b [140]. IkB kinase a (IKKa) has previously been shown to phosphorylate cyclin D1 on T286 [111] (see Sect. 5.3.4). Inhibition of IKKa with Bay11-708 [143] attenuated STG28 induced cyclin D1 degradation in LNCaP prostate cancer cells. Knockdown of IKKa by siRNA or the expression of the pIKK2M, IKKa dominant negative mutant similarly attenuated STG28 induced cyclin D1 degradation. Immunoprecipitation experiments demonstrated an interaction between cyclin D1 and both b-TrCP and Fbxw8 (see Sect. 5.5.4). Exposure of LNCaP cells to STG28, however, induced an increased association of cyclin D1 with Cul1 and b-TrCP but not Fbxw8. No interaction between cyclin D1 and the F-box proteins Fbx4, Fbw7, or Skp2 was detected in this study [111]. Accordingly, the siRNA mediated knockdown of b-TrCP and not Fbxw8 attenuated STG28 induced cyclin D1 degradation. Furthermore, overexpression of b-TrCP and not Skp2 induced a decline in cellular cyclin D1 levels and enhanced STG28 induced cyclin D1 degradation [111]. b-TrCP interacted with cyclin D1 even in untreated cells, suggesting that this F-box protein plays a physiological role in regulating cyclin D1 stability in unstressed cells. Cyclin D1 lacks a canonical b-TrCP binding motif [111, 127]. Instead, a sequence between glutamine residue 279 (E279) and T286 provides a variant b-TrCP recognition site [111]. Mutation of T286

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to alanine abolished b-TrCP binding. Mutation of E279 and E280 to alanine alone or in combination with the T286A mutation abolished STG28 induced cyclin D1 degradation. The E279A, E280A mutant was still phosphorylated following exposure to STG28, and in contrast to the T286A mutant localized predominantly within the cytoplasm. In vitro studies have also demonstrated that the co-mutation of E279, E280, and T286 to alanine substantially reduces the ability of b-TrCP to bind cyclin D1 [111]. IKKa thus facilitates cyclin D1 nuclear export by phosphorylating T286 which in turn facilitates the b-TrCP mediated ubiquitylation and degradation of cyclin D1 within the cytoplasm [111]. Wei et al. have also demonstrated that b-TrCP mediates cyclin D1 degradation in LNCaP cells in response to glucose starvation [111]. Glucose starvation induced b-TrCP expression and enhanced its interaction with cyclin D1. In contrast, the expression of Fbw7, Fbx4, and Skp2 was repressed under these conditions and none of the F-box proteins co-immunoprecipitated with cyclin D1 in glucose starved cells. Fbxw8 interacted with cyclin D1 but this association did not increase following glucose withdrawal [111].

9.5.6 Deciphering the Interactions Between Different Cyclin D1 Degradation Pathways The foregoing discussions indicate that several kinases and F-box proteins regulate cyclin D1 stability. In addition, several lines of evidence point to the existence of phosphorylation, ubiquitylation and/ or 26 S proteasome independent degradation (Fig. 9.1). The kinases ATR, Dyrk1b, ERK1/ 2, GSK3b, and IKKa appear to regulate cyclin D1 stability during the normal cell cycle by mediating T286 or T288 (Dyrk1b) phosphorylation. In contrast, p38SAPK2a appears to mediate cyclin D1 phosphorylation in response to environmental stresses and DNA damaging agents. Similarly, both ATM and ATR phosphorylate cyclin D1 in response to DNA damaging agents. Initial studies in NIH-3 T3 mouse fibroblasts indicated a role for GSK3b but not ERK2 or p38SAPK2a in mediating cyclin D1 phosphorylation on T286 [23]. More recent studies suggest, however, that ATR and not GSK3b is the major regulator of cyclin D1 phosphorylation in

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normally dividing NIH-3 T3 cells [79, 105, 144]. Since IKKa has also been identified as a mediator of cyclin D1 phosphorylation in MEF cells [111], it is unclear if these kinases co-operate to regulate cyclin D1 stability. IKKa inhibition has been shown to induce the accumulation of cyclin D1 in cancer cell lines [111, 113]. Furthermore, GSK3b activity has been linked to the regulation of cyclin D1 stability in HeLa, HuH7, MCF-7, and NA human cancer cell lines [69, 81, 82]. Fbx4 S12 phosphorylation, required for its dimerization, has also been shown to correlate inversely with Akt activity in human oesophageal and breast cancer cell lines [84], providing further evidence for GSK3b mediated regulation of cyclin D1 stability in these cells. The activity of these kinases is therefore, not restricted to normal cells and can clearly play an important role in regulating cyclin D1 stability in cancer cells. Consequently, the deregulated activity of these kinases may result in the accumulation of cyclin D1. The relative importance of these kinases in regulating cyclin D1 stability may vary however, in cell and context specific manner. At least five F-box proteins have been linked to the regulation of cyclin D1 stability to date. Recent studies suggest that Skp2 is not a bona fide cyclin D1 E3 ligase but might nevertheless, indirectly target the cyclin for degradation [125, 130, 145]. In contrast, Fbx4, Fbxw8, and bTrCP directly interact with cyclin D1 in vitro and in vivo [111, 135, 137]. Fbx4 and aB crystallin expression has been shown to be suppressed in a large percentage of tumor samples and breast cancer cell lines [135]. The suppression of Fbx4 has also been shown to be sufficient to induce the acquisition of a transformed phenotype in MEF cells [84]. Mutations that affect the ability of Fbx4 to dimerize have also been identified in oesophageal tumor samples [84]. The deregulated activity of Fbx4 thus appears to underlie the elevated levels of cyclin D1 expression in at least some cancers. These studies also suggest that while Fbx4 is important for regulating cyclin D1 stability in normal cells, it may play a lesser role in the regulation of the cyclin’s stability in cancer cells. Fbxw8 activity appears to play an essential role in facilitating the proliferation of some cancer cell lines. The siRNA mediated knockdown of Fbxw8 in HCT116 cells induced cyclin D1 accumu lation and significantly reduced the rate of cell


proliferation [137]. Unlike Fbx4, which requires the GSK3b mediated phosphorylation of cyclin D1 T286, Fbxw8 appears to be dependent on ERK1/ 2 mediated phosphorylation of this residue. ERK1/ 2 activity is often up-regulated in cancer cells [38]. In contrast, elevated Akt activity often results in the inhibition of GSK3b activity in cancer cells ([23] and references therein). Since cyclin D1 overexpression can inhibit cell cycle progression by interfering with DNA replication [72], some regulation of cyclin D1 stability may be required even in cancer cells. In fact, cyclin D1 can still be effectively degraded in MCF-7 cells that do not express aB crystallin [69, 135]. ERK1/ 2 dependent degradation of cyclin D1 by Fbxw8 may thus serve to maintain cyclin D1 levels below a certain threshold in cancer cells [137, 146]. Interactions between cyclin D1 and Fbxw8 have been reported in the LnCaP, HCT116, SW480 (human colorectal adenocarcinoma), U2OS and T98G (human glioblastoma) cell lines [111, 137]. Barbash et al. [84], however, failed to detect Fbxw8 expression in normal or tumor derived oesophageal epithelial cells. The role of Fbxw8 in regulating cyclin D1 stability and cell cycle progression in MCF-7 cells has not been reported. Interestingly, Fbxw8 does not appear to regulate cyclin D1 stability in LnCaP cells despite its interaction with this cyclin and the expression of activated ERK1/ 2 in these cells [111]. Instead, b-TrCP appears to be the major regulator of cyclin D1 stability in LnCaP cells. The shRNA directed knockdown of b-TrCP did not, however, affect cyclin D1 stability in U2OS cells [135]. Furthermore, b-TrCP was not observed to co-immunoprecipitate with cyclin D1 in the study by Okabe et al. [137]. b-TrCP expression has also been shown to be induced by different environmental stresses in 293 T human embryonic kidney cells [147] and glucose starved LnCaP cells [111]. b-TrCP may thus target cyclin D1 for degradation when cells are exposed to environmental stresses. Together, these observations indicate a differential role for Fbx4, Fbxw8, and b-TrCP in regulating the stability of cyclin D1 in different cell types. Further studies will be required to establish more precisely, the relative roles of these F-box proteins in regulating cyclin D1 stability in normal and cancer cells. Fbx4, Fbxw8, and b-TrCP require the prior phosphorylation of T286 in order to bind to and target cyclin D1 for ubiquitin dependent degradation.

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Cyclin D1 degradation can, however, occur independently of T286/ T288 phosphorylation ([113] see Sect. 5.4). Furthermore, the cyclin D1 variant, cyclin D1b (see Sect. 6), which lacks residue T286 does not always display an extended halflife [148]. It is currently uncertain if additional E3 ligases that target cyclin D1 independently of phosphorylation exist or if Fbx4, Fbxw8, and b-TrCP can mediate the low level degradation of nonphosphorylated cyclin D1. Alternatively, degradation pathways that do not involve prior substrate ubiquitylation or proteasomal activity may play a more central role in this context (see Sect. 5.4.2). Further studies are needed to create a comprehensive and integrated model of the various cyclin D1 degradation pathways in normal and cancer cells (Fig. 9.1).

9.6 The cyclin D1b splice variant in cancer Cyclin D1b is a splice variant of cyclin D1 that lacks the C-terminal regulatory sequences including T286 and T288 [149]. As a consequence, cyclin D1b is localized predominantly within the nucleus [148, 150]. Other studies suggest, however, that a significant amount of cyclin D1b can be detected in the cytoplasm of mantle cell lymphoma (MCL) cells [151]. The G/ A870 polymorphism has been linked to increased expression of the cyclin D1b variant. Carriers of the A allele have an increased propensity to express cyclin D1b but additional factors are expected to play a role in determining relative expression levels [149, 152–155]. Notably, a recent study failed to detect cyclin D1b in samples from MCL patients despite its expression at the mRNA level [156]. Cyclin D1b expression has been detected in several cancer subtypes [150–152, 157, 158]. Based on its similarity to cyclin D1, cyclin D1b may be expected to mimic cyclin D1 activity. Surprisingly, cyclin D1b is not effective at mediating Rb phosphorylation in vivo [148, 159]. Furthermore, cyclin D1b does not always show enhanced stability despite the absence of C-terminal regulatory sequences [148]. These observations suggest that CDK independent effects and, in particular, its constitutive nuclear localization may be important in cancer development and progression [148] (see below).

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9.7 Roles of D-Type Cyclins in Cancer 9.7.1 Oncogenic Roles of Cyclin D1 9.7.1.1 Cyclin D1 as an Oncoprotein Given its role in cell cycle regulation and its frequent overexpression in cancer cells, numerous studies have attempted to define the role of cyclin D1 in cancer development. As outlined in Sect. 2, studies in transgenic mice suggest that cyclin D1 is weakly oncogenic but can cooperate with other oncogenes to promote tumor development. Early studies indicated that cyclin D1 overexpression accelerates cell cycle progression and reduces serum dependence in NIH-3 T3 MEF cells as well as in Rat2 and Rat6 embryonic fibroblasts [12, 13]. The ability of cyclin D1 overexpression to induce oncogenic transformation or promote tumor development in mice in these studies appeared to be cell strain specific, possibly reflecting a role for additional oncogenes [12, 13]. Similar studies by Musgrove et al. [160] demonstrated that cyclin D1 overexpression accelerated cell cycle progression and reduced the growth factor dependence of breast cancer cells. Given that a reduced dependence on mitotic stimuli is a hallmark of cancer cells, deregulated cyclin D1 expression may facilitate cancer development by reducing the requirement for growth factors. The study by Quelle et al. [13] suggests, however, that this activity alone may be insufficient to drive the acquisition of a transformed phenotype. For instance, cyclin D1 overexpression in breast cancer infrequently correlates with markers of increased proliferation [161–167]. These studies suggest that the oncogenic activity of cyclin D1 may be independent of its interaction with CDK4.

9.7.1.2 Importance of Cyclin D1 Localization in Cancer Cells control the activity of cyclin D1 in part, by tightly regulating its subcellular localization (reviewed in [168]). The observation that overexpression of the constitutively nuclear T286 mutant greatly enhanced the transforming potential of cyclin D1 highlighted the importance of cyclin D1 nuclear accumulation in cancer [78]. The nuclear

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accumulation of cyclin D1 is highly regulated in normal cells [23, 78, 169, 170] and continues to be regulated in some cancer cells [69, 171–177]. Alt et al. hypothesized that the constitutive nuclear localization of cyclin D1 would enhance its transforming potential [78]. Long term overexpression of cyclin D1 in NIH-3 T3 cells did not result in focus formation and did not facilitate anchorage independent growth. In contrast, NIH-3 T3 cells expressing the T286A mutant formed large numbers of foci and exhibited anchorage independent growth. Furthermore, NIH-3 T3 cells overexpressing the T286A mutant but not wild type cyclin D1 induced tumor formation in SCID mice [78]. In these studies, the expression levels of wild type and T286A cyclin D1 were observed to be identical. The enhanced transforming potential of the T286A mutant was thus attributed to its constitutive nuclear localization [78]. Unlike wild type cyclin D1, the T286A mutant has also been shown to induce B-cell lymphomas in mice [178]. Further evidence for the enhanced oncogenic potential of the T286A mutant has come from studies in transgenic mice. Mice expressing the T286A mutant under the control of the MMTV-LTR, developed mammary adenocarcinomas at an increased frequency compared to mice expressing wild type cyclin D1 [67]. Furthermore, tumors from mice expressing the T286A mutant display characteristics that differ from those of mice expressing wild type cyclin D1. These include an increased mitotic index, increased genetic instability, and differences in tumor histology [67]. The clinical importance of these observations has recently come from oesophageal cancer cell lines and tumor samples. Benzeno et al. [179] have detected CCND1 mutations at low frequency in tumor samples from oesophageal cancer cell lines and tumor samples. These mutations prevent GSK3b mediated T286 phosphorylation, thus stabilizing the protein and rendering it constitutively nuclear. Mutations that prevent Fbx4 dimerization (and hence its ability to mediate cyclin D1 degradation), have also been detected in oesophageal tumor samples [84]. Similar to the T286A mutation, functional inactivation of Fbx4 results in the nuclear accumulation of cyclin D1 [84]. Furthermore, the suppression of Fbx4 expression is sufficient to induce the cellular transformation of MEF cells [84]. Several studies have, however, detected cyclin D1 expression with


a predominantly cytoplasmic localization pattern in tumor samples ([177] and references therein). Similarly, cyclin D1 localization within the cytoplasm was associated with cell cycle exit and quiescence in cardiomyocytes, postmitotic neurons and hepatocellular carcinoma cells [169, 170, 176, 180]. A predominantly cytoplasmic cyclin D1 localization pattern was also associated with lower grade tumors in prostate cancer [177]. Together, these studies indicate, that the activity and oncogenic potential of cyclin D1 is greatly enhanced by its accumulation within the nucleus. Given its role in cell cycle regulation, cyclin D1 overexpression likely confers a proliferate advantage to cancer cells. The tight regulation of cyclin D1 localization, however, provides a barrier against the full activation of its oncogenic potential [168]. Constitutively nuclear cyclin D1 mutants retain their capacity to catalyse the phosphorylation of Rb [179]. Furthermore, the attenuation of cyclin D1 catalytic activity significantly diminishes the transforming capacity of the T286 mutant in vitro [179]. The constitutive nuclear localization of cyclin D1 may thus increase its capacity to drive cell cycle progression [179]. The experimental evidence suggests that constitutive cyclin D1 nuclear localization can also drive cellular transformation independently of its catalytic activity [181]. Cells expressing the T286 mutant differentially express a number of genes associated with DNA replication and repair [67]. Cyclin D1 overexpression has also been shown to enhance gene amplification [182, 183]. Furthermore, tumors from mice expressing the T286 mutant display enhanced the stabilization of Cdt1 and increased aneuploidy relative to tumors from mice expressing the wild type cyclin [67]. Cyclin D1 accumulation within the nucleus may thus induce additional changes that enhance its capacity to transform cells. Studies on the Cyclin D1b protein suggest that this is indeed the case. Cyclin D1b is often localized predominantly within the nucleus but is not necessarily resistant to degradation [148, 150]. Expression of cyclin D1b has also been reported in primary breast, oesophageal, prostate, endometrial, and other cancers [150, 152, 184, 185]. In contrast to cyclin D1, cyclin D1b induces the transformation of NIH-3 T3 cells [148]. Given its weak catalytic activity, cyclin D1b has been proposed to induce cellular transformation

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as a result of either its constitutive nuclear localization, or its capacity to differentially interact with other proteins [135, 148]. Cyclin D1 modulates the activity of several transcription factors independently of CDK4 [186–188] (see Sect. 7.1.3). Studies in prostate cancer cells have demonstrated differential effects of cyclin D1 and cyclin D1b on androgen receptor (AR) activity. Cyclin D1 represses the ligand dependent activity of the AR and inhibits the cell cycle progression of androgen responsive prostate cancer cells. In contrast, cyclin D1b retained the capacity to bind AR but stimulated LnCaP cell proliferation [152]. The loss of cyclin D1 T286 phosphorylation or the regulatory sequences within the C-terminal region, thus have the capacity to modulate its activity and enhance its oncogenic potential [67, 152].

9.7.1.3 CDK4/6-Independent Activity of Cyclin D1 in Cancer Cyclin D1 regulates the activity of transcription factors independently of CDK4 [186, 187]. Cyclin D1 has been shown to induce the ligand independent activation of the oestrogen receptor a (ERa) independently of CDK4 [65, 66]. Ligand induced activation of ERa results in conformational changes within the receptor that facilitate its interaction with members of the steroid receptor coactivator (SRC) family [189]. Cyclin D1 facilitates the interaction between ERa and SRC members by virtue of its capacity to bind both proteins in the absence of oestrogen [190, 191]. Furthermore, cyclin D1 can interact synergistically with oestrogen to facilitate ERa activation [66]. It has thus been proposed that ERa and cyclin D1 interact in an autostimulatory loop where ERa induces Cyclin D1 expression, which in turn binds to and further enhances the activity of the receptor [192]. The interaction between cyclin D1 and ERa has important therapeutic implications. Antioestrogen therapy is limited by the development of resistance to these agents [189]. Overexpression of cyclin D1 has been shown to confer resistance to tamoxifen and reverse the growth inhibitory effects of antioestrogens [193–195]. Cyclin D1 has been shown to be important for the proliferation of tamoxifen resistant and tamoxifen induced MCF-7 breast cancer cells [43, 44]. Resistance to antioestrogens has also been associated with a shift towards ERa

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independent cyclin D1 expression [189]. Cyclin D1 overexpression can thus facilitate cell proliferation and resistance to antioestrogens by facilitating oestrogen independent ERa activation. Cyclin D1 has also been shown to regulate a transcriptional program independently of CDK4 [188]. Cyclin D1 interacts with the C/EBPb transcription factor to regulate the expression of a subset of genes including those that encode molecular chaperones and their regulators. C/EBPb regulates several cellular processes including differentiation and deregulated C/EBPb expression has been associated with tumorigenesis. It has been proposed, that cyclin D1 exerts its oncogenic actions partly by deregulating the expression of chaperones [187]. Evidence for this assertion has recently come from studies in B-cell lymphoma cell lines. In these studies, low level constitutive cyclin D1 expression induced the accumulation of molecular chaperones including Hsp70 and mediated resistance to apoptosis [196]. Cyclin D1 may also exert its oncogenic effects by preventing the activation of cellular differentiation pathways [86, 197, 198]. Cyclin D1 overexpression attenuates the capacity of C/EBPb to regulate cellular programs including differentiation [187]. Cyclin D1 overexpression may also interfere with differentiation programs by repressing peroxisome proliferator activated receptor g (PPARg) activity [199]. Additional studies similarly suggest that cyclin D1 overexpression prevents the initiation of differentiation programs in rhabdomyosarcoma cell lines. Accordingly, the pharmacological inhibition of CDK4/ 6 complexes blocked proliferation and promoted the differentiation of cultured myoblasts and rhabdomyosarcoma cell lines [200]. Together, these studies indicate that CDK4/6 independent cyclin D1 activity plays a major role in oncogenesis by perturbing transcription factor activity.

9.7.2 Roles of Cyclin D2 and D3 in Cancer Although less extensively characterized than cyclin D1, there is evidence to suggest that the other D-type cyclins may also contribute to tumor formation. The structural and biochemical characteristics of the D-type cyclins suggest that they perform similar functions [201, 202]. Although many cell types express one or more D-type cyclins, their role

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in regulating G1 progression appears to be cell type specific [203, 204]. Cyclin D2 and D3 expression has been shown to be elevated in multiple myeloma (MM) as a consequence of chromosomal translocations. Cyclin D1 has also been shown to be overexpressed together with cyclin D3 in primary breast cancer samples and cell lines [129]. Overexpression of cyclin D2 in rat fibroblasts shortened G0 to S phase progression and reduced serum dependence [13]. Recent studies have also demonstrated that the overexpressed cyclin D2 in human myeloma cell lines (HMCLs) is functional and modulates cell cycle progression [205]. The overexpression of cyclin D2 and D3 may thus induce increased proliferation in a cell type specific manner [203, 205]. Additional studies indicate that where coexpressed, individual cyclins may mediate distinct functions. Cyclin D3 appears to play a lesser role in mediating cell cycle progression in response to mitogenic signaling in cultured hepatocytes and regenerating mouse liver tissue [206–208]. Furthermore, both cyclin D1 and D3 are expressed in MCF-7 cells [129], but the siRNA mediated knockdown of cyclin D1 alone was sufficient to inhibit the proliferation of this cell line [53]. As discussed above, cyclin D1 appears to partly exert its oncogenic effects independently of CDK4. Cyclin D1 interacts with several transcription factors and modulates their activity [186]. It is currently unclear, if cyclin D2 and D3 similarly modulate transcription factor activity. A recent study suggests, that the 3 D-type cyclins exert distinct proliferative and transcriptional effects in cultured hepatocytes [206]. Future studies will need to determine the role of cyclin D2 and D3 in modulating transcription factor activity in cancer cells.

9.8 Pharmacological Targeting of Cyclin D1 in Cancer Numerous clinical and experimental cancer therapeutic agents inhibit cyclin D1 expression in cancer cell lines [209]. The identification of cyclin D1 as a target common to these agents probably reflects its role in coupling the extracellular environment and intracellular status to cell cycle progression. The labile nature of cyclin D1 results in its rapid disappearance in the absence of continued synthesis [23]. Alternatively, the increased activity of the


degradation pathways that target cyclin D1 can also lead to a rapid decline in protein levels [85] [209]. Entry into a subsequent round of cell division is dependent on the elevation of cyclin D1 levels in G2 [210]. This system thus enables cells to rapidly exit the cell cycle under unfavorable conditions. In fact, DNA damage and environmental stresses induce the rapid degradation of cyclin D1 [91, 105, 114, 210]. Given its importance in oncogenesis, cyclin D1 has been proposed as an attractive target for anticancer therapy [211–213]. Furthermore, proof of principle trials in the clinic indicate that targeting cyclin D1 degradation may prove useful in treating cancer [214]. Therapeutic agents that target cyclin D1 expression and/ or stability can be grouped into three main groups. Firstly, agents that inhibit the mitogenic signaling pathways regulating cyclin D1 expression induce a decline in the cellular levels of the cyclin [21]. Hence, ERa inhibition by antioestrogens suppresses cyclin D1 expression in cells that express the receptor. Such a therapy is likely, however, to select for cell populations that are less dependent on oestrogen and can result in drug resistance [43, 44, 50, 215, 216]. The second group of agents directly targets the catalytic activity of cyclin D1–CDK4/ 6 complexes [217, 218]. PD0332991 is a potent and specific inhibitor of CDK4 and CDK6 [219]. Treatment of various Rb positive cell lines with PD0332991 induces Rb dephosphorylation and G1 arrest. PD0332991 also showed antitumor activity in mouse xenograft models. Analyses of tumor samples from PD0332991 treated mice demonstrated a reduction in the levels of phosphorylated Rb, reduced proliferation indicated by the elimination of Ki-67 staining and the downregulation of E2F regulated gene expression [219]. These studies demonstrated that PD0332991 inhibits tumor growth by preventing cyclin D1–CDK4/ 6 mediated cell cycle progression. They also suggest, that Cyclin D1 regulates the proliferation of several cancer cell lines in a CDK4/ 6 dependent manner. Cyclin D1 expression also underlies the development of mantle cell lymphoma (MCL) and treatment of primary MCL cells with PD0332991 induced G0/ G1 arrest accompanied by reduced pRb phosphorylation [156]. Additional studies suggest that PD0332991 may be useful in combination with other anticancer agents 220 565. 5T33MM multiple myeloma (MM) tumor cells depend on

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cyclin D2–CDK4 complexes for cell cycle progression. Treatment of 5T33MM cells with PD0332991 induced cell cycle arrest at G1 in vitro. In ex vivo experiments, this agent induced tumor suppression and increased survival. PD0332991 induced G1 arrest has also been reported to sensitize MM cells to bortezomib induced cytotoxicity. The effect of PD0332991 is thus not limited to cyclin D1–CDK4/ 6 complexes and may thus be useful in situations where cyclin D1 ablation is not applicable (see below). Furthermore, agents like PD0332991 may be useful as chemosensitizers [220]. Related studies suggest that PD0332991 may also be useful in combination with antioestrogens such as tamoxifen in the treatment of breast cancer [67]. A number of therapeutic agents have also been shown to induce the increased degradation of cyclin D1 in cancer cells [209]. These studies suggest that the specific targeting of cyclin D1 is feasible and small molecule cyclin D1 ablative agents are currently in development [140]. Clinical therapeutics known to induce cyclin D1 degradation includes the retinoids, PPARg agonists, and histone deacetylase inhibitors [209]. These agents induce cyclin D1 degradation via the ubiquitin dependent degradation pathway but show differential requirements for the kinase mediated phosphorylation of T286. In general, these agents also suppress the expression of cyclin D1 at the mRNA level [53, 213, 221, 222]. Retinoids such as All-trans retinoic acid (RA) induce cyclin D1 degradation in a GSK3b and T286 dependent manner. The specific E3 ligases that mediate retinoid induced cyclin degradation have not been identified. RA also induced cyclin D3 degradation via 26 S proteasomes [212, 223]. Bexarotene (a retinoid X receptor agonist) has also been shown to target cyclin D1 for degradation in vitro and in vivo [224, 225]. Combination of bexarotene with erlotinib (an EGFR inhibitor) was more effective at inhibiting proliferation and repressing cyclin D1 expression in vitro than either agent alone [225]. Clinical studies using combination therapy with these agents for the treatment of nonsmall-cell lung cancer (NSCLC) have also shown promising results with limited toxicity [213, 225]. It should be noted, however, that the efficacy of retinoid induced cyclin D1 degradation appears to be cell line specific [226–228].. Further studies will thus be necessary to optimize the use of these agents in cancer therapy.

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PPARg agonists have also been shown to induce the ubiquitin dependent degradation of cyclin D1 in cancer cell lines [140–142]. Initial studies demonstrated that prostaglandin J2 (PGJ2) and synthetic PPARg agonists induce the cytoplasmic redistribution and degradation of cyclin D1 in a human breast cancer cell line. This study suggested that PPARg agonists suppress the proliferation of this cell line in part by targeting cyclin D1 [142]. Later studies determined that the PPARg agonist troglitazone induces cyclin D1 degradation independently of its PPARg agonist activity 141 474. This has lead to the development of troglitazone derivatives that lack PPARg agonist activity but nevertheless potently induce cyclin D1 degradation [140]. In contrast to the T286A mutation, inhibition of GSK3b did not suppress cyclin D1 degradation induced by PPARg agonists and troglitazone derivatives [140, 141]. More recent studies have demonstrated a role for IKKa in mediating T286 phosphorylation in a human prostate cancer cell line treated with the troglitazone derivative STG28 [111] (see Sects. 5.3.4 and 5.5.5). This study also determined that STG28 induces the b-TrCP mediated degradation of cyclin D1 in a IKKa manner [111]. PPARg agonists thus suppress cancer cell proliferation partly by targeting cyclin D1 for degradation. The development of troglitazone derivatives that lack PPARg agonist activity is important, since these agents are likely to induce fewer side effects [140]. Histone deacetylase (HDAC) inhibitors have been shown to be effective in suppressing cancer cell proliferation both in vitro and in vivo [229–232]. The HDAC inhibitor suberoylanilide hydroxamic acid (SAHA/ Vorinostat/ Zolinza) has recently been approved for the treatment of cutaneous T-cell lymphoma (CTCL) [230]. SAHA and other HDAC inhibitors are also in clinical trials for the treatment of other solid and haematological cancers [230, 233, 234]. The prototype HDAC inhibitor, trichostatin A (TSA), has been shown to induce cyclin D1 degradation via 26 S proteasomes in human breast cancer cell lines [53, 122]. TSA induced cyclin D1 degradation preceded its suppression of cyclin D1 mRNA expression and was only partially dependent on GSK3b or nuclear export [53, 122]. Although Skp2 upregulation has been shown to be associated with TSA induced cyclin D1 degradation, it is unlikely that this F-box protein is directly involved [53, 139, 235].

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The kinases and E3 ligases that directly facilitate HDAC inhibitor induced cyclin D1 degradation thus remain to be indentified. The effects of these agents on cyclin D1b have not been reported. It is unlikely, however, that those agents which require prior T286 phosphorylation will target cyclin D1b for degradation. Similar problems can be expected in the context of naturally occurring mutations within cyclin D1 that prevent T286 phosphorylation 179 233. It will thus be important to identify additional small molecule cyclin D1 ablative agents that simultaneously target cyclin D1 and cyclin D1b for degradation. Numerous other clinical and experimental therapeutic agents have also been shown to target cyclin D1 for ubiquitin dependent degradation ([209] and references therein). The recent identification of novel kinases and E3 ligases that regulate cyclin D1 degradation will provide further insight into the molecular pharmacology of these agents. Interestingly, many agents that induce cyclin D1 degradation are naturally occurring compounds found in traditional remedies and dietary substances. Cyclin D1 is thus also an important target for the development of chemopreventive strategies [209, 224].

9.9 Cyclin E in Cancer Cyclin E is essential for entry into S phase and thus regulates G1 cell cycle progression together with cyclin D1 [236] [2] [1]. Like cyclin D1, the aberrant expression of cyclin E1 has also been associated with cancer cell development and proliferation [237, 238]. Elevated cyclin E levels have been detected in numerous cancer subtypes [239–246]. Studies in transgenic mice suggest that cyclin E like cyclin D is a weak oncogene [247–249]. Like cyclin D1 cyclin E can collaborate with other oncogenes to induce tumor development [249–251]. Like cyclin D1, the overexpression of cyclin E is not always associated with amplification of the cyclin E gene (CCNE) [239, 244, 245]. Cyclin E is highly unstable and degraded via the ubiquitin dependent degradation pathway [252–255]. The deregulated degradation of cyclin E has been determined to be responsible for its accumulation in some cancers [238]. The Fbw7 F-box protein mediates the phosphorylation dependent degradation


of cyclin E. Mutations that affect the ability of Fbw7 to target its substrates have been identified in cancer cells [256–258]. The precise role of deregulated Fbw7 activity in cancer development is unclear, since the F-box protein targets several key substrates [258]. The degradation of cyclin E that is not bound to CDK2 (free cyclin E) occurs independently of Fbw7 [252, 254]. The precise E3 ligase(s) that mediate the degradation of free cyclin E have not been identified. It thus remains unclear if the deregulated activity of these pathways also underlies the accumulation of cyclin E in some cancer cells [238, 257]. Low molecular weight isoforms of cyclin E have also been detected in cancer cells [259]. It has been proposed that these isoforms are unique to cancer cells and enhance cyclin E oncogenic activity [259]. Interestingly, the mechanisms by which cyclin E promotes oncogenesis resemble those of cyclin D1. This might be expected, given their role in regulating cell cycle progression [29] [2, 236]. The overexpression of cyclin E lessened mitogenic dependence and accelerated progression through G1 in mammalian fibroblasts [260] [261]. Overexpression of cyclin E, however, also induces a delay in cell cycle progression [262, 263]. Cyclin E is now known to regulate several other cellular processes in addition to regulating pRb phosphorylation [238]. The effect of deregulated cyclin E activity on these pathways is now believed to underlie the oncogenic effects of this cyclin [238]. As reported for cyclin D1, cyclin E overexpression also induces genomic instability in mammalian cells [262–264]. Importantly, the suppression of Fbw7 activity similarly induces genomic instability in a cyclin E dependent manner [265]. The perturbation of proper S phase progression as a result of cyclin E accumulation may underlie the genetic instability observed in these studies [238]. CDK2 inhibitors have been under development for several years [217, 218]. Cyclin E can, however, function independently of CDKs and this is probably reflected by its proposed role in oncogenesis [238]. For these reasons, targeting Cyclin E-CDK activity in cancer may not be useful [238]. Agents that induce cyclin E degradation have not been identified. Targeting cyclin E expression itself may prove to be a more effective approach in cancer therapy. A better understanding of the pathways that target cyclin E for degradation

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independently of Fbw7 may facilitate this therapeutic approach.

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Chapter 10

The BRCA1/2 Pathway Prevents Some Leukemias and Lymphomas in Addition to Breast/Ovarian Cancers: Malignancies that Overcome Checkpoint Controls Bernard Friedenson Abstract In individuals who do not inherit BRCA1 or BRCA2 gene mutations, the encoded proteins prevent breast/ovarian cancer. However BRCA1 and BRCA2 proteins have multiple functions including participating in a DNA damage response subroutine that mediates repair of DNA double strand breaks by error-free methods. Inactivation of BRCA1, BRCA2, or any other critical protein within this “BRCA pathway” due to a gene mutation inactivates this error-repair process. DNA fragments produced by double strand breaks are then left to non-specific processes that rejoin them without any regard for preserving normal gene regulation or function, so rearrangements of DNA segments are more likely. These kinds of rearrangements are typically associated with some lymphomas and leukemias, so risk of these cancers is also increased. Therefore, along with preventing breast/ovarian cancers, preventing a subgroup of human lymphomas and leukemias is a physiologically important function of the pathway mediated by BRCA1 and BRCA2 gene products. Substantial percentages of these cancers, however, include non-random, characteristic gene rearrangements that bear witness to misrepair of DNA double strand breaks. In at least some of these diseases, there are specific mechanisms that overcome checkpoint controls. Keywords BRCA1 • BRCA2 • DNA repair • Leukemias • Lymphomas • Mantle cell lymphoma • Acute myeloid leukemia

10.1 Introduction BRCA1 and BRCA2 proteins are thought to be essential to prevent breast/ovarian cancer largely because carriers of mutations in the corresponding genes have high lifetime risks. More modest increases in risk for other cancers have also been noted [1–5]. Basic science studies find multiple biologic functions for BRCA1 and BRCA2 proteins [6–15], including participating within a pathway that mediates error-free repair of DNA double strand breaks by homologous recombination [15]. Much evidence indicates that BRCA and Fanconi pathways influence homologous recombination-mediated DNA repair. For example, human BRC repeats within BRCA2 mediate homologous recombination through control of RAD51 recombinase [16]. Fanconi anemia cells have a several fold increase in the frequency of aberrant rearrangements [17]. Figure 10.1 incorporates a model [18] for this error-free double strand break repair pathway which also has multiple connections to checkpoint controls of necessity [8, 19–24]. These controls help ensure optimal repair of double strand breaks by halting DNA replication and mitosis during the repair [21]. If repair is not possible, the cell checkpoints cause apoptosis or permanent arrest. BRCA1 and BRCA2 gene products are placed within a sequence encompassing the MRE11, Rad50 and NBS1 complex (MRN complex), ATM, CHEK2, BRCA1, BRCA2, and Fanconi anemia proteins. For the purposes here, this model will be referred to as the “BRCA pathway.”


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Cellular responses to DNA damage, such as DNA repair, cell cycle arrest, chromatin remodeling, and apoptosis are well coordinated [25]. There is enormous complexity in these controls. For example, yeast has seven different pathways operating to prevent gross chromosomal rearrangements [26]. Additional natural controls exist because cells may not multiply if genes encoding essential processes are damaged so that they are no longer compatible with either cell division or life. The pathway outlined in Fig. 10.1 is simplified but it has a major advantage because epidemiologic data exist for most individual components shown. The model can, therefore, be used to test theory against cancer susceptibility, helping to sort out overall physiologically relevant activities. A critical protein function lost from anywhere within this error-free repair BRCA pathway may force repair of DNA double strand breaks by lower fidelity, error prone methods. If cells containing illegitimate, pathogenic DNA repairs are able to re-enter the cell cycle and to bypass checkpoints, then risks of some cancers should increase. Lymphomas and leukemias can be associated with large gene rearrangements, which can be pictured as arbitrary rejoining of broken DNA fragments. For example, almost all mantle cell lymphomas have a characteristic interchange between pieces of chromosomes 11 and 14 [t(11;14)(q13;q32)]. In some leukemias, the make-up of a fusion protein may bear witness to other abnormal repairs (e.g., [17, 27]). Error-tolerant repair may also leave other signs such as in the acute myeloid leukemias, where there may be evidence of abnormal gene fusions, duplications, inversions, deletions, or reciprocal translocations [28]. In some cases, a translocation may attenuate the spindle checkpoint, making aneuploidy more likely [29]. The present study was designed to test the hypothesis that inactivation of a critical component of the BRCA pathway would increase risks of some lymphomas and leukemias by favoring pathogenic gene rearrangements. The results show that risks of a subset of leukemias and lymphomas increase up to nearly 2,000-fold. These findings may have clinical implications for surveillance and chemotherapy.

10.2 Methods used to Study the BRCA Pathway in Hematologic Cancers This analysis evaluated the risks of leukemias and lymphomas associated with a deleterious mutation

B. Friedenson

within a prototype BRCA pathway (Fig. 10.1). The purpose of this article is not to examine functionality of specific gene variants, but rather to examine the effect of the loss of gene function anywhere within a testable pathway on risk of specific hematologic cancers. For many of the studies examined, especially case series, the exact genetic variant is unknown, but loss of gene function (regardless of the reason) was confirmed by other means (e.g., RNA, protein or other tests). Genes examined were ATM, NBS1, MRE11, BRCA1, BRCA2, Fanconi anemia genes usually studied as a group including 13 known genes, and CHEK2. PubMed, PubMed Central, Google, and Google scholar searches collected about 2,500 relevant research articles related to the model BRCA pathway. Case-control, cohort, prevalence studies and basic science studies were reviewed. Data from studies that measured cancer incidences associated with epigenetic modification of pathway genes and/or alterations in protein or mRNA levels were also included. Epidemiologic studies were excluded in whole or in part if they did not provide required data or permit calculation of required information or if they were superseded or subsequently invalidated. The rarity of mutations in some molecules limited the analysis of some BRCA pathway components. As far as possible, statistical analyses were limited to gene variants either known to eradicate normal protein function or to severely lower normal levels. All the mutations were spontaneously occurring and/ or inherited except for therapy related (somatic) inactivation of BRCA1 in acute myeloid leukemia (AML). To verify that therapy related disease did not bias the results, it was compared to data for primary AML. Epidemiologic data were tabulated as odds ratios or relative risks: for ATM associations with NHL as MCL, with ALL, CLL, and PLL; for Fanconi anemia gene associations (13 known genes) with primary AML, with leukemia before age 15 and with ALL; for BRCA1 associations with primary and therapy related AML and with CML; for BRCA2 associations with AML, ALL, and CLL; for NBS1 associations with lymphomas, ALL and NHL; and for CHEK2 associations with CLL. Data used for these calculations are available at http:// www.biomedcentral.com/1471-2407/7/152. For meta-analyses, the DerSimonian-Laird random effects model [30] was used throughout since

10. The BRCA1/2 Pathway Prevents Some Leukemias and Lymphomas

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Figure 10.1. Brief overview of components within the BRCA pathway used here as a working model. Pathway components are also integral to checkpoint responses and some known relationships to checkpoints are indicated in the figure. Numbers in brackets refer to literature references. “Error-free” double strand break repair by homologous recombination involves many other proteins but the discussion is limited by available data to the proteins shown (with the exception of RAD50). Not shown are additional components including EMSY, PALB2, a whole family of RAD51 related proteins, DCC, cohesins, and accessory proteins. Large epidemiologic studies are uncommon for these additional proteins and deficiency states may be unknown. Other protein kinases related to ATM carry out similar functions in response to other genotoxic stresses, and some of them collaborate with ATM. Associations with further additional proteins are also likely.

it relaxes the assumption of a common effect due to mutation. This may be more appropriate here than fixed effects models since inactivating mutations can in theory have different targets with different effect sizes. However, the uncertainty bounds for random effects are more conservative and often larger. When at least three studies were available, meta-analysis was performed. For the nine studies available for ATM mutations in MCL, potential methodological confounders were ruled out by

generating subgroups without the potential confounder. Statistical associations were compared to independent basic science experiments and to basic science theory. Data from the NCI Surveillance Epidemiology and End Result (SEER) program were used to compare incidences in the general population versus Fanconi anemia groups. The NCI “DevCan” program was used to calculate cumulative control incidences for cancers affecting Fanconi anemia patients.

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The prevalence of ATM mutation heterozygotes in the general population is widely cited as 0.3–1%. The incidence of biallelic mutations which are required to cause the hereditary disease A-T is much smaller (3/million to 11/million) [31, 32]. Use of any value within this range as a control would give much larger risks. However, a population prevalence of 1% for ATM mutations was used to prevent overstating differences from the general population. None of the meta-analyses presented showed evidence for heterogeneity by the Breslow–Day method, from the inconsistency statistic [33], by a moment based method and graphically from L’Abbe plots. Bias was assessed using the method of Egger and by inspecting funnel plots for asymmetry [34]. There was no statistical evidence of publication bias for summary estimates (results not shown).

10.3 Defects Impacting the BRCA Pathway in Hematologic Cancers To determine if lymphoid/myeloid tissues were at increased risk due to mutations within the BRCA pathway, risks due to mutations or aberrations throughout the model pathway are summarized in Table 10.1. As each of these genes within the BRCA pathway is considered below, the conclusion emerges that a deficit in almost any of these genes greatly increases risks of a subgroup of hematologic cancers. This establishes what must be an important physiologic function of the overall arrangement of interconnected pathways.

10.3.1 Inactivation of the BRCA Pathway Gene ATM Allows a Translocation Associated with Mantle Cell Lymphoma Nine studies of the incidence of ATM mutations in MCL from a total of 363 patient samples were summarized. Meta-analysis seemed appropriate initially because all the studies found very strong odds ratios for an ATM–MCL association, so all nine studies have the same general pattern (criteria used by the Cochrane Review group).

B. Friedenson

Combining the nine studies then as described in methods, gave 70.26 [95% CI = 34.59–142.72] as the minimum odds ratio that a mantle cell lymphoma contains an ATM mutation (Table 10.1). The c2 test value that the pooled odds ratio differs from 1 was 138.30, P 700-fold increase in combined odds ratios for AML (Table 10.1). In Fanconi anemia, the BRCA pathway deficiency leads to visibly increased numbers of chromosome breaks, gaps, rearrangements, and quadriradii in the presence of DNA damaging agents. It is easy to see pieces of one chromosome inappropriately attached to another. This may result from a documented increase in repair by the less specific process of non-homologous end joining. Checkpoint controls fail in Fanconi anemia, and this failure is probably essential to disease. An example of a translocation capable of creating a differentiation block is the recurring t(3;12) (q26;p13) translocation. In a Fanconi anemia patient, this rearrangement was present in the bone marrow at the time of initial diagnosis of myelodysplastic syndrome (often a precursor of AML). The patient had a normal constitutional karyotype but AML then developed. When acute transformation to AML occurred, cytogenetic analysis found multiple chromosome deletions and rearrangements typical of Fanconi anemia [82]. Fanconi anemia is a rarely inherited disease, but the same t(3;12) translocation is sometimes the first and the only cytogenetic abnormality found in AML patients who do not have hereditary Fanconi anemia. This particular rearrangement is thought

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to predispose to AML as follows [83]. EVI-1 becomes driven by the TEL promoter [82, 84] so EVI-1 is overexpressed. Normally EVI-1 is expressed in early myeloid progenitor cells where it helps determine whether progenitors differentiate or proliferate. Abnormal EVI-1 expression probably contributes to AML by interfering with other genes controlling the commitment to differentiate. These progenitors are designed to proliferate rapidly and then to differentiate. Failure to induce timely differentiation might result in a prolonged proliferation phase. This mechanism of prolonging cell multiplication bypasses cell cycle related checkpoint controls, favoring the accumulation of additional cooperating events. This places the progenitors at much higher risk for leukemia. A background of hereditary Fanconi anemia would greatly increase chances for gene rearrangements and deletions in progenitors both as initial and as cooperating events. In myeloid leukemias, certain sites may associate with up to 40 different gene partners and chromatin structural elements closely associate with such breakpoints [85]. Some of these translocations have prognostic significance. Defects in various DNA damage and S-phase checkpoints permit differing degrees of increased spontaneous gross chromosomal rearrangements, increased chromosome loss, and increased recombination. Functioning checkpoints should censor which rearrangements that persist in AML cells. Perhaps certain chromosome regions are selected for rearrangements because they are more actively transcribed and exposed in a transcription complex [85, 86]. The proximity between neighboring chromosomes may also be an influence. A variety of gene rearrangements due to misrepaired double strand breaks also occur frequently in other diseases associated here with BRCA pathway deficiencies. In some cases of T-PLL, one gene rearrangement deregulates the expression of the T-cell receptor. Similarly, any of several recurring chromosomal translocations can be detected in substantial numbers of cases of childhood ALL. Gene fusions or other rearrangements often found in some of these tumors bear witness to a double strand break repaired by error-prone methods. Inactivation of a single gene can increase risks for multiple cancers, and inactivation of a different gene in the same pathway may have similar effects.

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Error-free repair is predominant in S/G2 whereas non-homologous end joining is more frequently used in G0/G1 because of the lack of homology sequences in these phases. Nbs1 promotes the processing of DNA double strand breaks, thereby facilitating recruitments of pathway molecules, such as Brca1 and Rad51, to DNA breaks to conduct homologous recombination. Therefore, mammalian cells have evolved a delicate repair mechanism modulated through the MRN complex. By favoring homologous recombination, cells can maximize their capacity to correctly repair lethal double strand breaks and, by repressing non-homologous end joining, cells minimize the generation of “dangerous liaisons” between broken chromosomes [87, 88]. The deficiencies that increase risk for leukemias and lymphomas may well be helpful in understanding other cancers in BRCA1/2 mutation carriers. Reciprocal translocations and other chromosome rearrangements also occur in breast and in ovarian tumors [89, 90]. The results here add the information that the same BRCA pathway can be disabled in both breast and hematological cancers. These deficits suggest the need for improved surveillance. They also present a vulnerability that may be exploited during therapy.

10.5 Conclusions This work shows that the pathway containing BRCA1/2 gene products is essential to prevent a group of leukemias and lymphomas. The results may have clinical implications for surveillance and chemotherapy in these and perhaps in other cancers. The genetic defect accompanying BRCA pathway deficiencies studied here is probably a chromosomal misrepair syndrome enabled by nonfunctioning or ineffective checkpoint controls. Acknowledgment: I am very grateful and wish to extend special thanks to Dr. Jack Kaplan and the UIC College of Medicine for their general support.

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B. Friedenson 52. Luciani J, Depetris D, Usson Y, Metzler-Guillemain C, Mignon-Ravix C, Mitchell M, Megarbane A, Sarda P, Sirma H, Moncla A, Feunteun J, Mattei M-G (2006) PML nuclear bodies are highly organised DNA-protein structures with a function in heterochromatin remodeling at the G2 phase. J Cell Sci 119:2518–2531 53. Xu Z-X, Timanova-Atanasova A, Zhao R-X, Chang K-S (2003) PML colocalizes with and stabilizes the DNA damage response protein TopBP1. Mol Cell Biol 23:4247–5256 54. Sugita K, Taki T, Hayashi Y, Shimaoka H, Kumazaki H, Inoue H, Konno Y, Taniwaki M, Kurosawa H, Eguchi M (2000) MLL–CBP fusion transcript in a therapy-related acute myeloid leukemia with the t(11;16)(q23;p13) which developed in an acute lymphoblastic leukemia patient with Fanconi anemia. Genes Chromosomes Cancer 27(3):264–269 55. La Starza R, Aventin A, Matteucci C, Crescenzi B, Romoli S, Testoni N, Pierini V, Ciolli S, Sambani C, Locasciulli A, Di Bona E, Lafage-Pochitaloff M, Martelli MF, Marynen P, Mecucci C (2006) Genomic gain at 6p21: a new cryptic molecular rearrangement in secondary myelodysplastic syndrome and acute myeloid leukemia. Leukemia 20(6):958–964 56. Friedman L, Thistlethwaite F, Patel K, Yu V, Lee H, Venkitaraman A, Abel K, Carlton M, Hunter S, Colledge W, Evans M, Ponder B (1998) Thymic lymphomas in mice with a truncating mutation in Brca2. Cancer Res 58:1338–1343 57. Mak T, Hakem A, McPherson J, Shehabeldin A, Zablocki E, Migon E, Duncan G, Bouchard D, Wakeham A, Cheung A, Karaskova UJ, Sarosi I, Squire J, Marth J, Hakem R (2000) Brca1 required for T cell lineage development but not TCR loci rearrangement. Nat Immunol 1:77–82 58. Scardocci A, Guidi F, D’Alou F, Gumiero D, Fabiani E, DiRuscio A, Martini M, Larocca L, Zollino M, Hohaus S, Leone G, Voso M (2006) Reduced BRCA1 expression due to promoter hyermethylation in therapy-related acute myeloid leukaemia. Br J Cancer 95:1108–1113 59. Risch H, McLaughlin J, Cole D, Rosen B, Bradley L, Fan I, Tang J, Li S, Zhang S, Shaw P, Narod S (2006) Population BRCA1 and BRCA2 mutation frequencies and cancer penetrances: a kin-cohort study in Ontario, Canada. J Natl Cancer Inst 98(23):1694–706 60. Shih H, Nathanson K, Seal S, Collins N, Stratton M, Rebbeck T, Weber B (2000) BRCA1 and BRCA2 mutations in breast cancer families with multiple primary cancers. Clin Cancer Res 6:4259–4264 61. Evans H, Lewis C, Robinson D, Bell C, Moller H, Hodgson S (2001) Cancer risks in women with 2 breast or ovarian cancers: clues to genetic cancer susceptibility. Int J Cancer 94:758–759

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Part III Targeting Checkpoint Response in Cancer Therapy

Chapter 11 Regulation of p53 Activity and Associated Checkpoint Controls Sean M. Post, Alfonso Quintás-Cardama, and Guillermina Lozano

Abstract Cancer develops when numerous genetic changes accumulate in pathways that regulate cell proliferation, differentiation, and apoptosis. In eukaryotes, failure to control cell division and loss of genomic stability are the hallmarks of neoplastic cells. Eukaryotic cells have evolved intricate signaling pathways, called checkpoints that can both halt uncontrolled proliferation and delay cell cycle progression in response to genotoxic stress, oncogenic activation, and aberrant proliferative signals. The p53 tumor suppressor encodes one such checkpoint protein. Although various mechanisms activate numerous positive and negative regulators required for cell cycle signaling cascades, these proteins relay their signals through many of the same essential components. Given the importance of these proteins in regulating cell cycle progression, many are altered during tumorigenesis. Herein, we provide an overview of the signaling mechanisms that control cell cycle progression and a primer of some of the most clinically promising agents targeting p53 and associated pathways that impact the cell cycle. Keywords p53 • Mdm2 • p19Arf • Tumorigenesis • Cancer • Ras • Cell cycle • Rb • Cyclin • CDK • ATM

11.1 Overview of the Cell Cycle The eukaryotic cell cycle is a process in which cells grow and then divide into two genetically identical cells. The cell cycle is divided into four discrete

phases allowing for the orderly transition through DNA replication, chromosomal condensation, spindle formation, and cytokinesis. The phases of the cell cycle are G1, in which a cell prepares for DNA replication; S, in which DNA synthesis takes place; G2, the period in which cells ensure their genetic information has been faithfully replicated; and M, in which homologous chromosomes are aligned equatorially and cell division takes place. To ensure normal unidirectional progression through the cell cycle, multiple and overlapping systems have evolved in eukaryotes. The first to be identified was the cyclin dependent kinase (CDK)cyclin system [1, 2]. CDKs are normally inactive protein kinases, which become activated by posttranslational modification and upon interaction with their regulatory cyclin partners. Throughout the cell cycle, CDK protein levels are relatively constant, while cyclin (hence the name) levels fluctuate. For example, CDKs 2, 4, 6, 7, and Cdc2 (also known as CDK1) are constitutively expressed throughout the cell cycle, but cyclin D is expressed in early G1 and is present through S and G2/M phases [3–5]. Cyclin E is expressed at the G1/S transition [6, 7], cyclin A during S-phase [8–11], and cyclin B in late S and at the G2/M transition [12, 13]. Cyclin H is expressed throughout all cell cycle phases [14–17]. Most cyclins are rapidly degraded at different times during the cell cycle. The cell cycle dependent flux in levels of various cyclins is one mechanism that regulates the kinase activity of the individual CDKs. These active CDK-cyclin kinase complexes exert their proliferative role on the cell cycle by pushing the cell

From: Checkpoint Controls and Targets in Cancer Therapy, Cancer Drug Discovery and Development Edited by: Zahid H. Siddik, DOI 10.1007/978-1-60761-178-3_11, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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from one phase to the next via phosphorylation of specific target proteins. The kinase activity of individual CDKs is not only controlled by changes in levels of the cyclins but also by its association with cyclin dependent kinase inhibitors (CKIs) and by post-translational modifications to the CDK-cyclin complexes. CKIs bind to and inhibit the CDK-cyclin complex and suppress their potential to influence the cell cycle. These CKIs include p15, which inhibits early G1 phase CDK-cyclins, such as CDK4-cyclin D and CDK6-cyclin D [18]. Loss of p15 expression is associated with several cancer types, such as acute lymphoblastic leukemia (ALL), chronic lympho cytic leukemia (CLL), and glioblastomas [18, 19]. Another CKI, p16 inhibits G1 phase kinases, such as CDK6-cyclin D [18]. Additionally, p16 expression, like that of p15, is lost in many human cancers [20]. The CKI p21, which is a target of the p53 tumor suppressor, inhibits CDK2-cyclin E in G1 and arrest cells at the G1/S boundary [21]. Inhibition of this late G1 phase CDK is consolidated by the closely related CKI p27 [22–24]. Post-translational modification is another mechanism controlling the activity of CDK-cyclin complexes. Phosphorylation of threonine and tyrosine residues in the amino terminus of CDKs by the kinases Wee1 and Myt1 [25–29] suppresses CDKcyclin activity, while it is activated following phosphorylation on threonine residues at the carboxy terminus by CDK-activating kinases (CAK) [30] such as CDK7 and CDK9. The inhibitory phosphorylation on T14 and Y15 is relieved by dephosphorylation of these residues by the phosphatase Cdc25 [31]. Thus, Cdc25 phosphatase activity allows the CDK-cyclin complexes to become active and drive the cell cycle. These multiple positive and negative regulatory signals allow for the tight control of entry into and exit from each transition point of the cell cycle. The first is the G1/S phase transition, which prevents replication of damaged DNA. The second occurs during S-phase, named the intra-S phase checkpoint, which slows replication if the genome is damaged. Third, the G2/M boundary checkpoint prevents aberrant mitosis if the chromosomes are damaged. A vast array of proteins influences cell cycle progression. Some are cell type specific or present only during a specific developmental stage. For example, loss of cyclin D1 results in viable mice

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that have retinal defects, mammary proliferation defects during pregnancy, and die early in life [32]. Mice expressing only cyclin D1 are viable but die prematurely due to megaloblastic anemia, while the cyclin D2-only and cyclin D3-only mice develop neurological defects, and lack normal cerebella, respectively [33]. Deletion of all three cyclin D genes results in embryo lethality due to anemia and heart defects [34]. Thus, the D type cyclins control cell cycle progression in a context specific manner. Likewise, CDK1-null mice are embryo lethal [35, 36], while CDK2 knockout mice are viable but sterile, suggesting a critical role for CDK2 only in the germline [37]. The myriad of tissue-specific defects resulting from deletion of these seemingly overlapping cell cycle pathways provides overwhelming evidence that tissue specificity or developmental differences have a critical role in determining cell cycle progression. Alterations to one or another of these cell cycle proteins are common in many cancers.

11.2 DNA Damage-Dependent Cell Cycle Arrest Cells can also arrest the cell cycle following exogenous insults to the genome, arising from exposure to ultraviolet light (UV), ionizing radiation (IR), and chemotherapeutic agents that cause DNA damage or interfere with DNA replication. Damage to the genome poses a grave threat, as mutations left unrepaired can ultimately lead to loss of cell cycle control and development of cancer. A large body of work has shown that damage-dependent checkpoint proteins negatively regulate cell cycle progression by inhibiting many of the aforementioned cell cycle kinases [38]. In response to DNA damage, protein complexes sense aberrant DNA structures (unwinding and/or bending) and initiate signal transduction cascades resulting in activation of cell cycle arrest or apoptotic programs. DNA damage-dependent signal transduction cascades directly phosphorylate the cell cycle machinery and CDK-cyclin complexes, resulting in delayed cell cycle progression. This delay provides cells the opportunity to repair the damage before moving to the next cell cycle phase. Loss of DNA damage dependent checkpoint

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activation can be disastrous, resulting in genomic instability (a hallmark of cancer). In cancer cells, however, many of these cell cycle arrest proteins or pathways are rendered nonfunctional. As such, cancer cells treated with chemotherapeutic agents that cause DNA damage will inefficiently repair DNA leading to cell death, while normal cells will be able to repair the damage.

11.3 Inactivation of p53-Dependent Cell Cycle Arrest Pathway The p53 tumor suppressor pathway is one of the most commonly inactivated pathways in human cancers. Mutations in or deletions of the p53 gene occur in over 50% of cancers [39]. The p53 protein is a sequencespecific transcription factor required for the activation of numerous DNA damage-dependent checkpoint response and apoptotic genes, such as p21, Gadd45, cyclin G, Bad, and Bax [40]. The N-terminus of p53 constitutes the transactivation domain that also binds its negative regulators, Mdm2 and Mdm4 [39]. The vast majority of the mutations found in tumors are in the central DNA binding domain. At the C-terminus lies the tetramerization domain. p53 also contains nuclear import and export motifs. p53 is induced and activated in response to DNA damage caused by a multitude of drugs, UV, and IR. Activation of p53 results in pleiotropic effects aimed at regulating physiological processes such as apoptosis, cell cycle arrest, and senescence. Cell type, the extent of damage, and the levels of the p53 protein (which is regulated by its negative inhibitors) contribute to the cellular response. While p53 regulates a multitude of pathways, its activity is also regulated by numerous mechanisms. Tumors that do not harbor deletions or mutations in p53 often have changes in other components of the p53 pathway resulting in p53 inactivation [39]. These include upregulation of the p53 inhibitors Mdm2 and Mdm4. The Mdm2 gene encodes an E3 ubiquitin ligase that binds to and ubiquitinates p53, thereby targeting it for degradation, and effectively removing p53 activity [41–43]. Interestingly, while Mdm2 inhibits p53 levels and activity, the transcription of Mdm2 is positively regulated by p53, creating a negative feedback loop whereby Mdm2 down regulates p53 while p53 upregulates Mdm2

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[44–46]. Mdm2 is amplified in over one third of human sarcomas, which retain a wild-type p53 gene, [39] and overexpressed by unknown mechanisms in many other tumors, suggesting that Mdm2 inactivates p53 function. Mdm4, a homologue of Mdm2, also binds to and negatively regulates p53 but lacks the E3 ubiquitin ligase activity of Mdm2 [47]. Like Mdm2, Mdm4 is overexpressed in multiple human cancers that retain wild type p53 [48]. The best evidence for the importance of p53 inhibition by Mdm2 and Mdm4 was demonstrated in mouse models. Loss of Mdm2 leads to a cell-death phenotype, which results in embryonic lethality at 3.5 days post coitum (dpc) [49, 50], while Mdm4 loss leads to cell cycle arrest and embryo lethality at 7.5 dpc [51]. These early embryonic lethal phenotypes are completely rescued by the concomitant deletion of p53, indicating that Mdm2 and Mdm4 are critical inhibitors of p53 activity in vivo. Changes in the levels of Mdm2 and Mdm4 can have profound effects on tumorigenesis, as both are either amplified or overexpressed in multiple tumor types that harbor wild-type p53. This fact has been recapitulated in transgenic mouse models in which overexpression of Mdm2 or Mdm4 results in increased tumorigenesis [52] (and G. Lozano, unpublished observations). The recent identification of a single nucleotide polymorphism (SNP) in the promoter of Mdm2 has shown that subtle and physiological changes to Mdm2 can likewise have severe consequences in tumor progression [53]. This polymorphic change creates a strong and stable Sp1 binding site that constitutively increases MDM2 transcription and therefore decreases p53 levels. Li-Fraumeni syndrome (LFS) patients harboring a p53 mutation who are also homozygous for the G nucleotide at SNP309 have an earlier onset of tumors as well as multiple primary tumors when compared to LFS patients lacking this polymorphism [53]. The generation of a mouse with this SNP will help elucidate the potential causative role of this SNP in cancer progression. The importance of the Mdm2-p53 pathway in cancer development is further highlighted by the fact that murine p19Arf (p14Arf in human) interacts with Mdm2 and sequesters it in the nucleolus [54–56]. This interaction blocks the E3-ubiquitin ligase function of Mdm2 resulting in the stabilization and activation of p53 [55, 57]. p19Arf is the protein product encoded by the alternate reading

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frame (ARF) of the INK4a locus and is an important tumor suppressor often deleted in cancers [58, 59]. Many tumors that express high levels of p19Arf harbor functionally inactivate p53, and those that maintain wild-type p53 often have silenced or deleted p19Arf 60. Hence, loss of p19Arf allows the released Mdm2 to bind to and inhibit p53 function. Thus, disruption or changes to p53 or other components of the p53 pathway, such as increasing Mdm2 or Mdm4 levels, or losing p19Arf, are essential steps in the process of cell transformation. These data indicate that while inactivation of the p53 pathway is observed in the vast majority of human cancers, the mechanisms leading to disruption of the pathway can vary greatly. As such, drugs affecting the stoichiometry of p19Arf, Mdm2, or p53 may be useful targets for reactivating the p53 pathway in human tumors with wild type p53.

11.4 p53 Mouse Models Another complication of the p53 pathway is the fact that most alterations in p53 result in a mutant protein that has gain-of-function activities [61]. However, the first generated and most widely studied p53 mouse model is the p53 knockout mouse [62]. While the p53-null mouse succumbs to tumors, unfortunately it does not represent a model for the most common type of p53 alterations, that of missense mutations [63]. To this end, researchers have begun to generate p53 knock-in mouse models that specifically match the germline mutations observed in the LFS families. Two mouse models have recently been generated: the p53R172H mouse (corresponding to the human p53 R175H mutation) [64, 65] and the p53R270H mouse model (corre sponding to the human p53 R273H mutation) [65]. In human cancers, these represent “hot-spot” mutations that lie in the DNA binding domain of p53. These mice, unlike those lacking p53, develop metastatic disease. This illustrates the subtle but critical differences in generating mouse models that more faithfully replicate human diseases. The establishment of in vivo mouse models with p53 missense mutations has also led to other novel discoveries. p53 mutant proteins are inherently unstable and become stable in tumors because of tumor specific alterations [64–66]. Additionally, many of the same signals that stabilize and activate

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wild type p53, such as disruption of interactions with MDM2 and DNA damage, also stabilize mutant p53 [66]. The stabilization of mutant p53 correlates with an increased potential for metastasis. Thus, differences in alterations of the p53 pathway provide different targets for therapy. A current list of mutant p53 mouse models that have been generated can be obtained from the website: (http://p53.free.fr/Database/p53_database.html). Such mouse models provide potential tools for developing and evaluating novel therapies against tumors with dysfunctions in the p53 pathways.

11.5 Targeting the p53 Pathway Directly Given the complexity of regulating either the wild type or mutant p53 protein, an understanding of the specific molecular defects disrupting the p53 pathway must be identified in each tumor to develop specific treatment strategies for each individual cancer patient. If Mdm2 is overexpressed and p53 is wild-type, then treatment with drugs that inhibit the Mdm2-p53 interaction would be crucial for tumor response. In contrast, if p53 mutations are present, this treatment option would not be viable, and in fact could stabilize mutant p53 with worse prognosis. In this case, methods to reintroduce wild-type p53 function into tumors would more likely yield a beneficial therapeutic response, if wild type p53 can overcome the activity of mutant p53.

11.5.1 Small-Molecule Inhibitors of the MDM2-p53 Interaction As previously mentioned, the activity of p53 is tightly regulated by MDM2. This regulation takes place through an autoregulatory feedback loop that involves direct protein-protein interaction between MDM2 and p53. MDM2 inhibits p53 through several mechanisms that include the exporting of p53 out of the nucleus and the targeting of the p53 protein for proteasome-mediated degradation. The interaction of MDM2 and p53 involves four key hydrophobic residues (Phe19, Leu22, Trp23, and Leu26) mapping to a short amphipathic helix formed by p53 and a deep hydrophobic groove in MDM2.


X-ray crystallography-based studies have delineated the topography of the MDM2-p53 interface and provided the rationale for the design of small-molecule inhibitors that interfere with this interaction. Several approaches have been taken to identify inhibitors of the MDM2-p53 interaction. One of the most fruitful approaches has been the use of high throughput screening of large chemical libraries. In 2004, Vassilev et al described a series of potent MDM2 inhibitors called Nutlins, containing a cis-imidazoline core structure [67]. Nutlin-1, Nutlin-2, and Nutlin-3 disrupted the interaction between MDM2 and p53 with IC50 values of 260 nM, 140 nM, and 90 nM, respectively. Nutlin-1 and Nutlin-2 are racemic mixtures whereas Nutlin-3a is an active enantiomer derived from racemic Nutlin-3. Crystallographic studies showed that Nutlin-2 binds MDM2 within the p53 binding domain [67]. Nutlin-3a has been tested against a series of cell lines representing different tumor types such as colon, breast, lung, prostate, melanoma, osteosarcoma, and renal cancer, all of them expressing wild-type p53 [68]. Nutlin3a-induced arrest at G1/S and/or G2/M phases in all these cell lines [67, 68], primarily due to induction of p53 activity that resulted in transcriptional activation of the CDK inhibitor p21 [68]. Treatment with Nutlin-3 (racemic) of primary B-cell chronic lymphocytic leukemia, [69–72] acute myeloid leukemia, [73] and multiple myeloma [74] cells resulted in marked apoptosis. Nutlin-3 treatment rendered complete inhibition of tumor growth in several xenograft models of human cancer with wild-type p53 [67, 68, 75]. RITA (reactivation of p53 and induction of tumor cell apoptosis) is another low-molecular weight compound with the potential to activate wild-type p53 in human cancers and was identified by screening the National Cancer Institute (NCI) chemical library [76]. Further characterization of this compound showed that RITA binds to the p53 N-terminus and causes a conformational change that disrupts the MDM2-p53 interaction in vitro, which results in p53 accumulation, upregulation of p53 target genes, and massive apoptosis of tumor cells, with little effect on normal cells [76]. Indeed, RITA suppressed the growth of human fibroblasts and lymphoblasts only upon the expression of an activated oncogene such as c-MYC and demonstrated high antitumor activity with minimal toxicity in mice carrying human tumor xenografts [76].

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MI-219, a novel spiro-oxindole analog, has also shown to be a potent inhibitor of the MDM2-p53 interaction with potent cellular activity in cancer cells with wild-type p53 and an excellent cell permeability and pharmacokinetic profile [77, 78]. MI-219 was rationally designed to mimic all four key residues in the MDM2-p53 interface [78]. MI-219 binds MDM2 with a Ki value of 5 nM, whereas Nutlin-3 has a Ki value of 36 nM [78]. Treatment with MI-219 resulted in cellular accumulation of p53 and growth inhibition of cancer cells with wild-type p53 [78]. These studies indicate that drugs that interfere with the respective binding domains on Mdm2 and p53 can reactivate p53. Caution must be taken to ensure that these tumor cells have wild type p53 as mutant p53 is also stabilized by Mdm2 loss. In summary, small-molecule compounds that activate wild-type p53 in cancer cells are active and appear not to induce significant toxicity in normal cells. This therapeutic window reinforces the notion that nutlin-3, RITA, and MI-219 represent promising candidates for drug development in human cancers with wild-type p53.

11.5.2 p53-Reactivating Small Molecules The above examples of drugs that activate p53 are dependent on the presence of wild-type p53 in cells. However, inactivation of the p53 gene via missense mutations is the most common mechanism of inactivating p53 in human tumors. The fact that cancer cells carrying mutant p53 proteins were susceptible to the reintroduction of wildtype p53 spurred the development of activators of mutant p53. Low-molecular-weight compounds such as PRIMA-1 and MIRA-1 activate mutant p53 in cell-based assays [79]. Although the mechanism of action of these drugs is largely unknown, structural studies have shown how small peptides bind the DNA-binding domain of mutant p53 and refold the protein into an active form [80]. In vitro, PRIMA-1 restores the wild-type conformation and reactivates the transcriptional function of mutant p53. Treatment with PRIMA-1 causes inhibition of xenograft tumor growth in SCID mice, and in the NCI diversity set, targets preferentially mutant p53-expressing tumor cells [81]. PRIMA-1 induces apoptosis selectively in mutant

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p53-expressing colorectal carcinoma cells through a mechanism that involves the JNK pathway in a caspase-independent manner [82]. However, a recent report has shown that PRIMA-1 can induce mutant p53-dependent mitochondria-mediated apoptosis through activation of caspase-2 leading to cytochrome c release and subsequent activation of caspase-9 and caspase-3 [83]. PRIMA-1MET, a methylated and more potent form of PRIMA-1, was found to be synergistic with chemotherapeutic drugs such as adriamycin and cisplatin as assessed both in vitro and in an in vivo SCID mouse model [84]. In addition to PRIMA-1, other molecules that target mutant p53 such as MIRA-1, the amifostine derivative WR1065, [85] ellipticine, and derivatives thereof, [86] can also restore the conformation of p53 to wild-type and induce p53-dependent checkpoint and pro-apoptotic targets, thus widening the repertoire of agents with clinical potential for patients with tumors expressing mutant forms of p53.

11.6 Oncogenic Stimulation of the Cell Cycle and the p53 Pathway In vivo, cells exist in either a cycling (G1/S/G2/M) or quiescent (G0) state. G0 cells can be thrust into the cell cycle by environmental cues, such as mitogenic signaling or aberrant oncogenic signaling. Receptors on the cell surface bind to various growth factors, cytokines, or mitogens which in turn activate downstream signaling cascades [87]. This results in the increased expression of genes that regulate proliferation. Conversely, many of these same signals activate apoptotic programs as well, creating dual but competing signals for life and death. Given the propensity to drive two major growth regulating pathways, mutations or aberrant regulation of the genes involved in these pathways are commonly observed in human cancers, as cancer cells either over proliferate (by increasing the regulation of proliferative genes), and mutate or delete apoptotic programs in order to survive. The literature detailing the slew of signaling cascades emanating from various receptor molecules is overwhelming. However, as an example, one common pathway altered in cancer is the Ras-Raf-Mek-Erk pathway. Receptors on the cell surface bind to various growth factors or mitogens which in turn

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stimulate the Ras-Raf-Mek-Erk signaling cascade [88, 89]. Upon binding to growth factors or ligands on the cell surface, the receptors become activated and bind macromolecule complexes in the cytosol of the cell. This converts Ras from its inactive (GDP bound) to active (GTP bound) state, which can then bind to and activate Raf. Raf is a protein kinase that regulates downstream kinases, such as Mek [90], which in turn phosphorylates protein kinase Erk that results in activation of gene expression via phosphorylation of downstream targets [91, 92]. The importance of this pathway can not be understated as point mutations that mimic the constitutively active form of Ras, which releases Ras from its growth factor dependence, have been commonly reported in many human cancers [93]. Additionally, activating mutations of BRaf (one of the three Raf proteins) are common in melanoma and other cancers [87, 94]. Deregulation of the Ras-Raf-Mek-Erk pathway has the ability to stimulate cellular proliferation, which in and of itself can also activate the p53 pathway. As a consequence of Ras activation, levels of cyclins D1, D2, and D3 increase [95]. Furthermore, overexpression of Raf leads to increased levels of cyclins and CDKs, as well as decreased levels of CKIs [96]. Aberrant expression of cyclins and CDKs, stemming from the deregulation of Ras-Raf, has the potential to inactivate the cell cycle inhibitory complex Retinoblastoma protein (Rb)-E2F, ultimately resulting in the activation of the p53 pathway in some cell types (Fig. 11.1). Expression of activated Ras has been shown to result in phosphorylation of Rb [97]. Additionally, expression of Ras results in increased levels of the transcription factor E2F [98], as well as the levels of E2F targets [99, 100]. Moreover, when Ras function is inhibited by the compound Farnesyl thiosalicylic acid, E2F is inactivated [101]. The potential impact of Rb-E2F on the p53 pathway will be further elaborated below. Another example of how the Ras-Raf-MekErk pathway impacts the p53 pathway is through its regulation of the transcription factor Dmp1. Dmp1 in conjunction with c-Jun and JunB directly binds to and activates the p19Arf promoter [102]. Expression of oncogenic Ras in Dmp1-null cells failed to result in transactivation of both p19Arf and p21. The activation of Dmp1 by the Ras/Raf/Mek/Erk

11. Regulation of p53 Activity and Associated Checkpoint Controls $

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Figure 11.1. Schematic representation of the convergence of the cell cycle and the DNA damage-dependent signaling pathways through the p19Arf/Mdm2/p53 axis. Symbols: # loss of gene expression; F inactivating gene mutations; + activating gene mutations; $ protein overexpression.

pathway demonstrates the ability of Ras to directly impact p53 activity. Ultimately, aberrant activation of the Ras/Raf/ Mek/Erk pathway stimulates p53 function resulting in cell cycle arrest or apoptosis. In order to counteract the antiproliferative effects of p53, cells must bypass the p53 pathway through inactivation or mutations. As such, Ras activating and p53 inactivating mutations are common in human cancers, and thus a series of chemotherapeutic agents are currently being employed in the clinic to address these defects.

11.6.1 Inhibitors of the Ras-Raf-MekErk Pathway The Ras-Raf-Mek-Erk pathway is an evolutionary conserved signaling pathway. Ras gene mutations that constitutively activate Ras are commonly encountered in a variety of hematologic disorders and solid tumors [103]. Ras is a small G-protein that is initially synthesized in the cytoplasm as an inactive protein that eventually migrates to the inner surface of the plasma membrane [104]. This is a critical step in Ras function and is accomplished through a post-translational reaction termed prenylation, whereby a 15-carbon isoprenyl (farnesyl) group is attached to the Ras C-terminal cysteine by an enzyme called farnesyltransferase

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and to a lesser extent by geranylgeranyl-protein transferases [103]. Inhibition of these enzymes has been investigated as a means of interfering with Ras processing and signaling. Several farnesyltransferase inhibitors (FTIs) have been developed. However, suppression of tumor cells by FTIs also occurs in the absence of Ras mutations, suggesting that other proteins equally dependent on prenylation might be as relevant for this effect. In this regard, another mechanism by which FTIs may decrease the proliferation of cancer cells is by interfering with bipolar spindle formation during transition from prophase to metaphase in mitosis. Centromeric proteins such as CENP-E and CENP-F are substrates for farnesyltransfeRase [105]. Tipifarnib (R115777) is an oral nonpeptidomimetic FTI with significant in vitro antiproliferative activity against a variety of cell types. Treatment with tipifarnib of patients with hematologic malignancies indicates that this agent has activity in this setting [106]. Unfortunately, several phase II studies conducted in patients with different tumor types have shown that this activity, when present, is modest [107–112]. Results with lonafarnib, (SCH66336), another oral nonpeptidomimetic with similar mechanism of action as tipifarnib, have been largely disappointing [113–115]. Despite the modest activity of FTIs in vivo, alternative strategies to target the Ras-Raf-Mek-Erk pathway have been developed. One such strategy is that of using small-molecule kinase inhibitors against BRaf, Mek, and/or Erk. Sorafenib is a bisaryl urea analog with potent activity against a wide range of kinases [116] that has shown promising results in clinical trials. Although sorafenib was originally designed as a BRaf inhibitor, subsequent studies demonstrated that in addition to being a potent BRaf kinase inhibitor (IC50 of 6 nM), sorafenib was also a potent inhibitor of vascular endothelial growth factor receptors 1, 2, and 3, platelet-derived growth factor receptor-b, and other receptor tyrosine kinases involved in tumorigenesis such as KIT (IC50 6 nM) [116]. Treatment with sorafenib reduced the levels of phosphorylated (p)Erk in tumor cell lines in which the Ras-Raf–Mek–Erk pathway was upregulated by oncogenic Ras and/or Raf mutations, including colon and pancreatic tumor cells (both K-Ras), melanoma tumor cells (b-Raf V600E), and breast tumor cells (K-Ras and b-Raf G463V) [117].

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Sorafenib also inhibited tumor growth and angiogenesis and induced apoptosis via Mcl-1 downregulation, through Ras-Raf–Mek–Erk-dependent or -independent pathways, depending on the type of tumor being investigated [117]. Activating somatic mutations in the BRaf proto-oncogene have been described in a wide variety of malignancies, including melanoma, papillary thyroid cancer, and other cancers [94]. For instance, in melanoma, BRaf mutations are found in 59% of cases arising on the skin without signs of chronic sun-induced damage [118]. The potent activity of sorafenib against the BRaf kinase makes it an attractive candidate for clinical development, and several studies have demonstrated the activity of this agent in patients with acute myeloid leukemia, [119] melanoma, [120] renal cell carcinoma, [121, 122] and advanced hepatocellular carcinoma [123].

11.7 Crosstalk Between RB and p53 Pathways in Early G1 In response to aberrant proliferative signals, pathways have evolved that inhibit cell cycle progression. One of the classic negative regulators of cell cycle progression is the Rb pathway. Rb was the first tumor suppressor cloned, and three Rb family members have subsequently been identified; p105, p107, and p130 [124]. Each of these proteins share similar structure and function. The effect that Rb exerts on the cell cycle is similar to that of CKIs, although through a different mechanism. Rb negatively regulates progression through early G1, where CDK4/6 have important roles [125]. Rb influences cell cycle regulation by binding to and inhibiting the function of the transcription factor E2F [126, 127]. To date, at least six E2F proteins have been identified [128]. E2F binds to the promoter of many genes required for transition from G1 to S by regulating the expression of cyclins A and E as well as components of the DNA replication machinery [11, 129]. Conversely, the cell cycle arrest function of Rb is negatively regulated by various CDK-cyclin complexes. CDK4-cyclin D and CDK6-cyclin D can hyperphosphorylate Rb, resulting in the release of E2F and thus progression through the cell cycle [14]. Therefore, interplay between Rb-E2F, CDK-cyclin complexes, and CKIs exists in order to ensure proper regulation of

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cell cycle progression in response to proliferative signals (Fig. 11.1). As alluded to above, the Rb-E2F pathway also directly influences the activation of the p53 pathway. Both human p14Arf and murine 19Arf are directly induced by E2F1 over-expression in cell culture [130, 131]. In addition, Rb-E2F3 binds to the p19Arf promoter and dampens its transactivation [132]. As a function of cell cycle dependent phosphorylation of Rb, this complex is released allowing for p19Arf activation. E2F transactivation leads to a senescent phenotype in a p19Arf-p53-p21 dependent manner [132, 133]. However, in tumors isolated from mouse models, a less robust effect of Rb loss or increased E2F levels has been observed, suggesting the potential for a possible E2F-independent activation of the p19Arf-p53 pathway [134, 135]. A further upstream regulator of the p53 pathway is the CKI p16. Deletion of p16 in a homozygous mutant p53R172H background results in the stabilization of p53R172H [66]. In further support of p16dependent regulation of p53, p16 loss in human mammary epithelial cells leads to stabilization of p53 and conversely, overexpression of p16 leads to p53 instability via an Rb-dependent manner [136]. It is easy to envision that loss of p16, resulting in the activation of the CDK4- or CDK6-cyclinD complexes could then increase phospho-Rb and thus release its inhibition on E2F, which would then be free to activate the p19Arf promoter, resulting in increased p53 levels. Taken together, normal regulators of the cell cycle (CDKs, cyclins, and CKIs) influence Rb-E2F interactions and sit at a crossroad between cell cycle progression and DNA damage checkpoint activation via the p53 tumor suppressor pathway. Since Rb loss leads to p53 activation through these regulators in some cell types, tumors simultaneously need to mutate p53 to proceed through the cell cycle. Targeting the Rb pathway in the clinic has been challenging. Rb-mutant retinoblastomas have been shown to harbor wild type p53. Recently, however, 80% of retinoblastomas have been found to overexpress Mdm4, a potent inhibitor of p53 [137]. Using the preclinical drug nutlin-3, which disrupts the p53-Mdm2 interaction and, at higher concentrations, the p53-Mdm4 interaction, Rb mutant tumors have been targeted for p53-dependent destruction [137]. This study


proposes a novel approach to targeting tumors possessing Rb mutations but retaining wild type p53.

11.7.1 Small Molecule Agent PD0332991 Impacts the CDK4/CDK6-Rb Pathway From the preceding section, it follows that disruption of the CDK4/6-Rb pathway has the potential for therapy. One possibility is by inhibition of CDK activity. The first demonstration that single-agent therapy with a highly selective inhibitor of CDK4 and CDK6 as a potential candidate for clinical use has been recently reported with the development of the pyridopyrimidine PD0332991 (CDK4/D1: IC50, 0.011 mmol/L; CDK6: IC50, 0.016 mmol/L) [138, 139]. This compound is endowed with >100fold higher selectivity for CDK4 and CDK6 than for CDKs 1, 2 and 5, and has >1000-fold selectivity against a panel of 35 other kinases [139]. In vitro, rapid and reversible inhibition of Rb phosphorylation at serines 780 and 795 (CDK4 and CDK6 specific sites) was demonstrated in MDA-MB-435 and Colo-205 tumor cells upon treatment with PD0332991 [138]. Treatment of tumor cells expressing wild-type retinoblastoma (Rb) results in induction of G1 arrest accompanied by reduction of phosphorylation at serines 780 and 795 on Rb. Treatment with PD0332991 of eight different cell lines carrying wild-type Rb indicated potent antiproliferative activity. Not surprisingly, in tumor cell lines lacking Rb (e.g., MDA-MG-468 breast carcinoma cell line), in which p16INK4A is present at high levels and already associated with CDK4/6, PD0332991 is inactive. In xenografts of cell lines harboring functional Rb, remarkable activity was observed upon oral administration of PD0332991, with tumor regression observed in Colo-205 colon and SF-295 glioblastoma xenografts, accompanied by the abrogation of Rb phosphorylation at serine 780 and reduction of the Ki-67 proliferation marker [138]. Subsequent oral administration to mice bearing Colo-205 xenografts resulted in potent tumor regression although tumor growth was observed after treatment discontinuation [138]. A phase I study (NCT00141297) of oral PD0332991 in patients with advanced malignancies and wild-type Rb has completed accrual. Several other phase I studies using this agent are currently underway for patients with different malignancies such as

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relapsed mantle cell lymphoma (NCT00420056), multiple myeloma (in combination with the proteasome inhibitor bortezomib; NCT00555906), and hormone-sensitive advanced breast cancer (in combination with the oral non-steroidal aromatase inhibitor letrozole; NCT00721409).

11.8 Activation of Cell Cycle Checkpoints Through DNA Damage-Dependent Kinases ATM (Ataxia Telengiectasia-Mutated) is a serine/ threonine dependent protein kinase required for regulating checkpoints following DNA damage. The ATM gene was first identified as the gene mutated in patients with the rare autosomal recessive disorder ataxia-telangiectasia (A-T) [140]. A-T patients suffer from cerebellar ataxia, telangiectasia, immunodeficiencies, predisposition to cancer, and sterility. Cells isolated from A-T patients and cells derived from Atm-deficient mice have defects in activating the G1/S, S-phase, and G2/M cell cycle checkpoints are sensitive to ionizing irradiation (IR) and exhibit radioresistant DNA synthesis, a condition where DNA replication is not slowed after treatment with IR [141, 142]. These results indicate that ATM is involved in cellular responses to lesions caused by IR. Activation of the cell cycle checkpoint following DNA damage requires ATM to phosphorylate cell cycle arrest proteins, such as p53 [143–145], Chk1 [146, 147], and Chk2 [148]. ATM, as well as Chk1 and Chk2, directly and indirectly regulates checkpoint response by two main mechanisms. The first and most studied mechanism is the p53dependent checkpoint pathway, which requires ATM to directly phosphorylate p53 on Ser15 [149]. ATM also indirectly regulates p53 by activating Chk1 and Chk2, which in turn phosphorylate p53 on Ser20 [150, 151]. Phosphorylation of Ser20 disrupts the interaction between p53 and MDM2, the E3 ubiquitin ligase that targets p53 for degradation [152]. This disruption results in the accumulation of p53 protein and activation of p53-dependent targets such as p21 and GADD45. p21 prevents G1/S transition by inhibiting G1 phase CDKs, and by interacting with and thereby inhibiting the function of the DNA replication protein, proliferating cell nuclear antigen (PCNA) [153]. Additionally,

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p21 is a potent activator of cell senescence [154157]. GADD45 inhibits S-phase progression by also binding to and inactivating PCNA [158] and inhibits the G2/M phase by inhibiting the function of Cdc2 [159]. These data indicate that checkpoint kinases maintain a functional cell cycle checkpoint by controlling the levels and transcriptional activity of p53 in response to DNA damage. Regulating the activity of p53 is a complex process. Not only is p53 function regulated by Mdm2 and Mdm4, but the p53 transcriptional activities are also controlled by a myriad of posttranslational modifications. Positive effectors of p53 activation include phosphorylation by kinases, such as ATM, Chk1, and Chk2, as well as ubiqitination, acetylation, neddylation, and sumoylation [160]. Phosphorylation at the N-terminus of p53 by these and others kinases has the ability to activate p53. Deletions of any of these cell cycle checkpoint kinases impair but do not eliminate p53 activity in vivo, suggesting that phosphorylation by itself is insufficient to completely and properly regulate p53 activity in vivo. Moreover, the observation that mutations in p53 phosphorylated amino acids have not been identified in human cancers suggests that modifications to p53 are not likely the most critical factor regulating p53 function in vivo, and suggests these post-translational modifications play a subtle role in either regulating activation of specific p53-dependent targets or have some effect on specific cell types. In fact, many mouse models harboring point mutations to specific sites of posttranslational modification have been generated and show only mild phenotypes at best [161–164]. Furthermore, knock-in mice with a p53 that is unable to be acetylated due to lysine to arginine substitutions in all seven residues in the C-terminal tetramerization domain also failed to demonstrate an in vivo phenotype [165]. Together, these results indicate that p53 post-translation modifications do not have a profound impact on tumorigenesis in vivo, but rather, suggest that they may have more subtle effects on p53 that may become apparent under experimental conditions. Another G1/S checkpoint regulator is the ATMChk1/Chk2-Cdc25A pathway. Upon activation, ATM and Chk1 impinge on the cell cycle by disrupting the activation of the CDK-cyclin complex. Chk1 phosphorylates Cdc25A leading to its ubiquitin-

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mediated degradation [166, 167]. This negative regulation of the cell cycle activating phosphatase Cdc25A results in a delay of the cell cycle [29, 166]. Cdc25A normally removes the inhibitory cons traints of the Thr-14 and Tyr-15 phosphorylation on CDK2, stimulating its kinase activity and thus driving the transition from G1 to S phase [31]. As a consequence of CDC25A inhibition, the CDK2-cyclin E kinase remains inactive, and cell cycle progression is halted. Failure to activate this DNA damage-dependent checkpoint pathway results in aberrant cell cycle proliferation. By regulating two different G1/S checkpoint mechanisms, ATM and Chk1/Chk2 provide cells with tightly controlled checkpoint responses.

11.8.1 Small-Molecule CDK Inhibitors An alternative to inhibiting cell cycle progression by endogenous p53-dependent and –independent checkpoint mechanisms is to inhibit CDK directly with small molecule drugs. More than 50 smallmolecule compounds that target the ATP-binding pocket of the catalytic site of CDKs have been identified and many of them are currently undergoing preclinical and clinical evaluation [168]. An important question regarding these agents is whether promiscuous CDK inhibition is preferable to selective CDK inhibition. Selective inhibition of CDK4/6 by PD0332991 was discussed earlier. Similarly, several compounds with nanomolar or low micromolar potency for CDK2/CDK1 inhibition but markedly lower activity against CDK4/6 have been identified, including seliciclib, SNS032, SU9516 and AZ703. As expected, these compounds have been reported to induce S and G2 arrest followed by apoptosis. In addition, given that CDK2/CDK1 inhibition might enhance apoptosis induced by DNA-damaging agents that affect S-phase progression (e.g., anthracyclines) or by microtubule stabilizing agents (e.g., taxanes), the use of CDK inhibitors in combination with more traditional chemotherapeutic agents may render superior results compared with the use of each group of agents individually. Confirmation of this hypothesis will require the conduction of randomized trials, a task that will undoubtedly be facilitated by the fact that many of these agents are orally bioavailable.


11.8.1.1 Flavopiridol (alvocidib, L868275; HMR1275) The semisynthetic flavonoid flavopiridol was the first CDK inhibitor to enter the clinical arena, having been tested in more than 50 clinical trials so far. Flavopiridol is the most potent known inhibitor of CDK9 (IC50 3 nM versus 30 nM and 100 nM for CDK1 and CDK2, respectively) [169, 170]. In proliferating cell populations, flavopiridol blocks cell-cycle progression at the G1/S and G2/M boundaries. A majority of tumor cells exposed in vitro to flavopiridol arrest at G1/S but significant fractions of cells can be detected in sub-G1 and at the G2/M transition indicating significant levels of apoptosis and mitotic arrest. Flavopiridol downregulates cyclin D1 and D3, c-Myc, and the apoptosis regulators Mcl-1 and XIAP [171]. Flavopiridol also activates p53 through down regulation of MDM2, thereby promoting apoptosis [171]. Flavopiridol has proven efficacious in vitro against chronic lymphocytic leukemia (CLL), a disease where tumor cells survive on account of persistent expression of CDK9-dependent antiapoptotic proteins. In a phase I trial, 42 patients with fludarabinerefractory CLL received a 30-min loading dose of flavopiridol followed by a 4-h intravenous infusion, which was pharmacokinetically modeled to achieve and sustain micromolar concentrations for several hours [172]. Therapy was given weekly for 4–6 weeks. The observed dose-limiting toxicity (DLT) was hyperacute tumor lysis syndrome (massive lysis of malignant cells leading to severe electrolyte imbalance). Safe administration of flavopiridol was achieved by carefully monitoring and correcting potassium levels. Of the 42 patients treated, 19 (45%) achieved a partial response that lasted for a median of more than 12 months [172]. Responses were observed in patients with genetically high-risk disease, including 5 (42%) of 12 patients with del(17p13.1), associated with loss of the p53 gene, and 13 (72%) of 18 patients with del(11q22.3), associated with loss of the ATM gene [172]. Combinations of flavopiridol with the mainstay of CLL therapy, the purine analog fludarabine and rituximab, have demonstrated substantial but manageable toxicity, with high clinical response rates. Remarkable response rates to flavopiridol have also been reported among patients with mantle

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cell lymphoma [173]. The highest activity of flavopiridol as a single-agent has been observed in patients with B-cell malignancies (e.g,. CLL, mantle cell lymphoma, multiple myeloma). However, clinical trials in solid tumors are underway, both as single-agent as well as in combination with standard chemotherapeutic agents, where the sequential addition of the former appears to potentiate the antimitotic activity of the taxane docetaxel, as evidenced in clinical trials for patients with refractory metastatic pancreatic cancer (NCT00331682) or locally advanced breast cancer (NCT00020332).

11.8.1.2 SNS-032 (formerly BMS-387032) SNS-032 is an aminothiazole CDK inhibitor with potent activity against CDK2/cyclinE (IC50=48 nM) and CDK9 (IC50=4 nM), which has been shown to inhibit cell cycle and transcription [174]. The excellent pharmacokinetic profile of SNS-032 in both rodents and dogs contributed to its selection as a clinical candidate [174]. In a dose-finding study in 21 patients with metastatic solid tumors or refractory lymphoma, SNS-032 was given as a 1 h intravenous infusion in 3 weekly doses over 21 days at a starting dose of 4 mg/m2 [174]. In this study, the half-life of SNS-032 in humans was shown to be 5–10 h. SNS-032 was well tolerated and 15% of the patients exhibited stable disease [174]. Two separate phase I multicenter trials are currently underway to address the tolerability of SNS-032 given as short infusions to patients with B-cell malignancies (NCT0446342) or advanced solid tumors (NCT00292864).

11.8.1.3 Seliciclib (CYC-202, R-roscovotine) Seliciclib is a trisubstituted purine analog with selectivity of CDK1, CDK2, CDK7, and CDK9 [175, 176]. Seliciclib has also been shown to downregulate Mcl-1, prior to the induction of apoptosis, thus suggesting a transcription inhibition mechanism of action [177, 178]. Roscovitine also inhibits the repair of double-strand breaks by either homologous recombination or nonhomologous end joining, which may also contribute to its chemosensitizing properties [179]. In a recently reported phase I study, seliciclib was administered orally for 7 consecutive days at either 100 mg, 200 mg, or 800 mg on a daily twice schedule to 21 patients

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with a variety of cancers [180]. Dose-limiting toxicities were seen at 800 mg twice daily in the form of grade 3-4 fatigue, Rash, hyponatremia, and hypokalemia. Inhibition of retinoblastoma protein phosphorylation was not demonstrated in peripheral blood mononuclear cells. No complete responses were observed but disease stabilization was reported in 8 patients [180]. The APPRAISE trial is a double-blinded, randomized study of single agent seliciclib versus best supportive care in patients with non-small cell lung cancer (NSCLC) who have received at least two prior systemic therapies. Of the first 173 patients, 45 achieved stable disease for six weeks and were entered into the blinded portion of the study in which they were randomized either to continue seliciclib or to receive placebo with best supportive care. The primary efficacy endpoint is progression free survival. Data will remain blinded until the last patient has completed follow-up.

11.9 Conclusions Cells utilize highly intricated and tightly regulated cell cycle signaling systems to sense DNA damage, aberrant growth signals, and oncogenic activation to initiate a checkpoint-signaling cascade to halt cycle progression. With our ever expanding knowledge of the cell cycle, DNA damage-dependent checkpoints, and the obvious crosstalk between these pathways, classic non-targeted chemotherapeutic agents are now beginning to give way to biologybased therapeutics aimed at treating specific tumor types by targeting specific molecular drivers. This “more personalized” approach to cancer therapy has already paid remarkable dividends for patients with some cancers (e.g., such as chronic myeloid leukemia and gastrointestinal stromal tumors) for which key pathogenetic underpinnings have been identified and matched with appropriate targeted agents (e.g., imatinib, and sunitinib, respectively). The continued investigation of critical components of the cell cycle and the p53 pathway will undoubtedly contribute fundamental insights into aspects inextricably linked to cancer cells such as cell division, apoptosis, and senescence, which potentially will facilitate the development of more rational therapeutics with the potential to improve clinical outcomes.

S.M. Post et al.

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11. Regulation of p53 Activity and Associated Checkpoint Controls 128. Trimarchi JM, Lees JA (2002) Sibling rivalry in the E2F family. Nat Rev Mol Cell Biol 3(1):11–20 129. Lees E, Faha B, Dulic V, Reed SI, Harlow E (1992) Cyclin E/cdk2 and cyclin A/cdk2 kinases associate with p107 and E2F in a temporally distinct manner. Genes Dev 6(10):1874–1885 130. Bates S, Phillips AC, Clark PA et al (1998) p14ARF links the tumour suppressors RB and p53. Nature 395(6698):124–125 131. DeGregori J, Leone G, Miron A, Jakoi L, Nevins JR (1997) Distinct roles for E2F proteins in cell growth control and apoptosis. Proc Natl Acad Sci U S A 94(14):7245–7250 132. Aslanian A, Iaquinta PJ, Verona R, Lees JA (2004) Repression of the Arf tumor suppressor by E2F3 is required for normal cell cycle kinetics. Genes Dev 18(12):1413–1422 133. Rowland BD, Denissov SG, Douma S, Stunnenberg HG, Bernards R, Peeper DS (2002) E2F transcriptional repressor complexes are critical downstream targets of p19ARF/p53-induced proliferative arrest. Cancer Cell 2(1):55–65 134. Russell JL, Powers JT, Rounbehler RJ, Rogers PM, Conti CJ, Johnson DG (2002) ARF differentially modulates apoptosis induced by E2F1 and Myc. Mol Cell Biol 22(5):1360–1368 135. Tolbert D, Lu X, Yin C, Tantama M, Van Dyke T (2002) p19(ARF) is dispensable for oncogenic stress-induced p53-mediated apoptosis and tumor suppression in vivo. Mol Cell Biol 22(1):370–377 136. Zhang J, Pickering CR, Holst CR, Gauthier ML, Tlsty TD (2006) p16INK4a modulates p53 in primary human mammary epithelial cells. Cancer Res 66(21):10325–10331 137. Laurie NA, Donovan SL, Shih CS et al (2006) Inactivation of the p53 pathway in retinoblastoma. Nature 444(7115):61–66 138. Fry DW, Harvey PJ, Keller PR et al (2004) Specific inhibition of cyclin-dependent kinase 4/6 by PD 0332991 and associated antitumor activity in human tumor xenografts. Mol Cancer Ther 3(11):1427–1438 139. Toogood PL, Harvey PJ, Repine JT et al (2005) Discovery of a potent and selective inhibitor of cyclin-dependent kinase 4/6. J Med Chem 48(7): 2388–2406 140. Savitsky K, Bar-Shira A, Gilad S et al (1995) A single ataxia telangiectasia gene with a product similar to PI-3 kinase. Science 268(5218):1749–1753 141. Barlow C, Hirotsune S, Paylor R et al (1996) Atmdeficient mice: a paradigm of ataxia telangiectasia. Cell 86(1):159–171 142. Xu Y, Baltimore D (1996) Dual roles of ATM in the cellular response to radiation and in cell growth control. Genes Dev 10(19):2401–2410

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188 division age in both normal and SV40-transformed human fibroblasts. Oncogene 10(5):835–840 158. Azam N, Vairapandi M, Zhang W, Hoffman B, Liebermann DA (2001) Interaction of CR6 (GADD45gamma ) with Proliferating Cell Nuclear Antigen Impedes Negative Growth Control. J Biol Chem 276(4):2766–2774 159. Zhan Q, Antinore MJ, Wang XW et al (1999) Association with Cdc2 and inhibition of Cdc2/ Cyclin B1 kinase activity by the p53-regulated protein Gadd45. Oncogene 18(18):2892–2900 160. Olsson A, Manzl C, Strasser A, Villunger A (2007) How important are post-translational modifications in p53 for selectivity in target-gene transcription and tumour suppression? Cell Death Differ 14(9):1561–1575 161. Chao C, Hergenhahn M, Kaeser MD et al (2003) Cell type- and promoter-specific roles of Ser18 phosphorylation in regulating p53 responses. J Biol Chem 278(42):41028–41033 162. Sluss HK, Armata H, Gallant J, Jones SN (2004) Phosphorylation of serine 18 regulates distinct p53 functions in mice. Mol Cell Biol 24(3):976–984 163. Iwakuma T, Parant JM, Fasulo M et al (2004) Mutation at p53 serine 389 does not rescue the embryonic lethality in mdm2 or mdm4 null mice. Oncogene 23(46):7644–7650 164. MacPherson D, Kim J, Kim T et al (2004) Defective apoptosis and B-cell lymphomas in mice with p53 point mutation at Ser 23. Embo J 23(18):3689–3699 165. Krummel KA, Lee CJ, Toledo F, Wahl GM (2005) The C-terminal lysines fine-tune P53 stress responses in a mouse model but are not required for stability control or transactivation. Proc Natl Acad Sci U S A 102(29):10188–10193 166. Furnari B, Rhind N, Russell P (1997) Cdc25 mitotic inducer targeted by chk1 DNA damage checkpoint kinase. Science 277(5331):1495–1497 167. Sanchez Y, Wong C, Thoma RS et al (1997) Conservation of the Chk1 checkpoint pathway in mammals: linkage of DNA damage to Cdk regulation through Cdc25. Science 277(5331):1497–1501 168. Shapiro GI (2006) Cyclin-dependent kinase pathways as targets for cancer treatment. J Clin Oncol 24(11):1770–1783 169. Chao SH, Fujinaga K, Marion JE et al (2000) Flavopiridol inhibits P-TEFb and blocks HIV-1 replication. J Biol Chem 275(37):28345–28348 170. Chao SH, Price DH (2001) Flavopiridol inactivates P-TEFb and blocks most RNA polymerase II transcription in vivo. J Biol Chem 276(34):31793–31799

S.M. Post et al. 171. Lam LT, Pickeral OK, Peng AC, et al. (2001) Genomic-scale measurement of mRNA turnover and the mechanisms of action of the anticancer drug flavopiridol. Genome Biol; 2(10):RESEARCH0041 172. Byrd JC, Lin TS, Dalton JT et al (2007) Flavo piridol administered using a pharmacologically derived schedule is associated with marked clinical efficacy in refractory, genetically highrisk chronic lymphocytic leukemia. Blood 109(2): 399–404 173. Kouroukis CT, Belch A, Crump M et al (2003) Flavopiridol in untreated or relapsed mantle-cell lymphoma: results of a phase II study of the National Cancer Institute of Canada Clinical Trials Group. J Clin Oncol 21(9):1740–1745 174. Heath EI, Bible K, Martell RE, Adelman DC, Lorusso PM (2008) A phase 1 study of SNS032 (formerly BMS-387032), a potent inhibitor of cyclin-dependent kinases 2, 7 and 9 administered as a single oral dose and weekly infusion in patients with metastatic refractory solid tumors. Invest New Drugs 26(1):59–65 175. McClue SJ, Blake D, Clarke R et al (2002) In vitro and in vivo antitumor properties of the cyclin dependent kinase inhibitor CYC202 (R-roscovitine). Int J Cancer 102(5):463–468 176. Meijer L, Borgne A, Mulner O et al (1997) Biochemical and cellular effects of roscovitine, a potent and selective inhibitor of the cyclin-dependent kinases cdc2, cdk2 and cdk5. Eur J Biochem 243(1–2):527–536 177. Alvi AJ, Austen B, Weston VJ et al (2005) A novel CDK inhibitor, CYC202 (R-roscovitine), overcomes the defect in p53-dependent apoptosis in B-CLL by down-regulation of genes involved in transcription regulation and survival. Blood 105(11):4484–4491 178. Hahntow IN, Schneller F, Oelsner M et al (2004) Cyclin-dependent kinase inhibitor Roscovitine induces apoptosis in chronic lymphocytic leukemia cells. Leukemia 18(4):747–755 179. Crescenzi E, Palumbo G, Brady HJ (2005) Roscovitine modulates DNA repair and senescence: implications for combination chemotherapy. Clin Cancer Res 11(22):8158–8171 180. Benson C, White J, De Bono J et al (2007) A phase I trial of the selective oral cyclin-dependent kinase inhibitor seliciclib (CYC202; R-Roscovitine), administered twice daily for 7 days every 21 days. Br J Cancer 96(1):29–37

Chapter 12

The Importance of p53 Signaling in the Response of Cells to Checkpoint Inhibitors Alan Eastman

Abstract Many anticancer agents induce DNA damage resulting in arrest of cell cycle progression and thereby permit time for repair and recovery. Accordingly, these cell cycle checkpoints limit the efficacy of DNA-damaging agents. Chk1 inhibitors have been developed to prevent arrest and enhance cell killing. This approach would also kill normal cells if it were not for the protective role played by the p53 tumor suppressor. This chapter discusses how p53 can protect cells from DNA damage rather than induce apoptosis, and how activation of p53 in nontumor tissues could enhance the therapeutic index when a patient with a p53 defective tumor is administered a combination of a DNA-damaging agent plus Chk1 inhibitor. Keywords DNA damage • Cell cycle arrest • p53 • p21waf1 • Cyclin B • Chk1 inhibitors • UCN-01 • Topoisomerase I inhibitors • Nucleoside analogs

12.1 The Discovery of UCN-01 as a Checkpoint Inhibitor The observation that caffeine sensitized cells to ultraviolet light was reported more than 40 years ago [1]. Fifteen years later, this sensitization was attributed to the ability of caffeine to overcome DNA damage-induced G2 cell cycle arrest, driving the cells through a lethal mitosis [2], but identification of its target as ataxia telangiactasia mutated (ATM) and ATM and RAD3-related (ATR) did not occur until 1999 [3, 4]. Unfortunately, neither caffeine nor its analogs were selective or potent

enough to overcome cell cycle arrest in patients. Our studies on the mechanism of action of caffeine led to the discovery in 1996 that 7-hydroxystaurosporine (UCN-01) was an alternate and highly potent checkpoint inhibitor [5]. The serendipitous events that led to this discovery have not been told before, but it provides a prelude to a discussion of the role of p53 in checkpoint regulation. Qizhi Wang was a graduate student in the Ph.D. program in pharmacology and toxicology at Dartmouth Medical School. Two years into the program, her husband completed his Ph.D. and obtained a new position in Washington, DC. So that she could follow her husband, Qizhi completed her M.S. degree, but with the understanding that she would seek out a new mentor in Bethesda and complete her Ph.D. at a distance while remaining a student in our graduate program. Qizhi found a new mentor in Dr. Edward Sausville at the National Cancer Institute (NCI), and began to study the biological effects of the protein kinase C inhibitor UCN-01. Qizhi and Ed visited Dartmouth in 1995 so that she could report on her research findings to her thesis committee, which I chaired. At that meeting, Qizhi showed that UCN-01 had an unexpected impact on cell cycle progression; it appeared to activate Cdk1 and Cdk2, and reduce the length of time cells spent in G2 [6]. This observation should be put in context with the research that was ongoing in my laboratory at that time. For many years, I had been studying the mechanism of action of cisplatin, and by 1995 my group had published several papers on its impact on cell cycle arrest [7–10]. Todd Bunch, a postdoctoral fellow in my laboratory, was continuing studies on the impact of


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caffeine on cisplatin-induced cell cycle arrest. Upon seeing Qizhi’s data, I questioned whether UCN-01 might have a similar effect as caffeine on arrested cells. Within a week, Ed had sent us the compound, and in another week Todd had performed the experiments and sent back the answer that UCN-01 was a potent checkpoint inhibitor. This original work was performed in our Chinese hamster ovary cell model [5]. Meanwhile, three laboratories had just reported that the checkpoint action of caffeine and its analog pentoxyfylline was much more effective at enhancing cytotoxicity in cell lines defective for p53 [11–13]. One of these groups was led by Patrick O’Connor at NCI. Qizhi, Ed, and I then collaborated with Patrick to extend our observation to human cell lines, and demonstrated that UCN-01 was also much more effective in cell lines that were defective in p53 [14]. In summary, these results showed that UCN-01 was 100,000 times more potent than caffeine at abrogating cell cycle arrest, and it selectively killed p53 defective cells. Importantly, the levels of UCN-01 that were required to inhibit Chk1 and enhance cell killing could be achieved in animals. These combined observations resulted in tremendous interest because of the potential to selectively kill tumor cells, and the impact of this work can be judged by the numerous pharmaceutical companies who have subsequently developed their own Chk1 inhibitors, of which at least five are currently in clinical trials [15].

12.2 The Target of UCN-01 At the time UCN-01 was discovered as a checkpoint inhibitor, Chk1 had not yet been discovered. Human Chk1 was discovered in 1997 [16], which led rapidly to the realization that Chk1 was the target for UCN-01 [17, 18]. UCN-01 was originally shown to inhibit Chk1 but not Chk2, although a later paper provided evidence that Chk2 was also a target [19]. Both Chk1 and Chk2 possess autophosphorylation sites (ser296-Chk1 and ser516-Chk2) and the availability of antibodies to these phosphorylation sites allowed us to demonstrate that 10–30 nM UCN-01, a concentration effective at abrogating arrest, inhibited only Chk1 autophosphorylation [20]. We have seen no inhibition of the Chk2 autophosphorylation at concentrations up to 1 µM. Hence, UCN-01 does not inhibit Chk2 in cells. UCN-01 inhibits many other kinases. Using an in vitro screen of 25 different kinases, UCN-01

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was shown to inhibit 12 of them by at least 50% at 1 µM [21]. Interestingly, MAPKAPK2 (MK2) was one of the kinases not inhibited in this screen, although a recent paper has suggested it is inhibited, and that this partly explains the efficacy of UCN-01 [22]. We have been unable to replicate this observation, and have concluded that UCN-01 does not inhibit MK2 [20]. Our conclusion is supported by other work demonstrating that UCN-01 can only abrogate arrest in several normal cell lines when p38, the upstream activator of MK2, is suppressed [23]. The fact that UCN-01 could not abrogate arrest alone suggests it is unable to inhibit MK2 in these models. Another interesting target in vitro is CDK2, which is inhibited at only slightly higher concentrations than Chk1 [24]. However, this inhibitory action cannot be occurring in cells because it would prevent the abrogating action of UCN-01. In MDA-MB-231 breast cancer cells, we find that 300 nM UCN-01 can induce G1 arrest, while 5 µM induces G2 arrest (unpublished observations). The G1 arrest may relate to UCN-01-mediated inhibition of PDK1 [25], an upstream activator of AKT. However, the impact of UCN-01 on the PDK1–AKT pathway is very complex, as AKT has many other potential roles in checkpoint regulation such as via inhibitory phosphorylation on ser280 of Chk1 [26, 27], or by acting in concert with DNA−protein kinase to enhance transcriptional activation of p21waf1 [28]. One of the confounding problems in studies with UCN-01 relates to the use of unnecessarily high concentrations that inhibit many of these other targets. It has been common practice to use concentrations in the range of 100–1,000 nM UCN-01 (it is this range that inhibits PDK1), whereas Chk1 is inhibited in cells at 15 nM, which is the concentration we have used to abrogate arrest in a variety of cell lines. The availability of a phosphospecific antibody to ser296-Chk1 now makes it feasible to directly measure the concentration that inhibits Chk1 activity in every cell system. With the development of new inhibitors of Chk1, some of these off-target effects can be avoided, yet many of these inhibitors also have other inhibitory activities. In particular, many also inhibit Chk2 [15]. The impact of this remains to be established. As Chk2 may contribute to the induction of p53, it is possible that a dual inhibitor of both kinases may be detrimental, as it may overcome the protection p53 provides to normal tissues.

12. The Importance of p53 Signaling in the Response of Cells to Checkpoint Inhibitors

12.3 The Importance of p53 in Response of Cells to Checkpoint Inhibition The observation that p53 wild-type cells are resistant to Chk1 inhibitors has frequently been rationalized as follows. Cells with functional p53 can arrest in G1 when their DNA is damaged, while in the absence of p53 this G1 checkpoint is defective and damaged tumor cells will arrest in S or G2. As Chk1 inhibitors are effective only on S and G2 cells, this will selectively enhance killing in the p53 defective tumor. Unfortunately, this rationale is not well supported by data. Most of the DNAdamaging agents used cause damage selectively in S phase. Nucleoside analogs do not impact passage through G1 but only damage DNA during S phase. Topoisomerase I inhibitors (such as SN38 used in many of our studies) cause DNA double-strand breaks only when the replication complex collides with a stalled topoisomerase I. Hence, there is little if any G1 arrest in either p53 wild-type or mutant cells incubated with these agents. We have compared the impact of SN38 on p53 mutant MDA-MB-231 cells and p53 wild-type MCF10A cells [29]. In both cell lines, SN38 induces G2 arrest at low concentrations and S phase arrest at higher concentrations. Upon addition of UCN-01 to S-arrested cells, only the p53 mutant cells progressed through S and G2 phase, and then through a lethal mitosis. When p53 was suppressed in MCF10A using shRNA (MCF10A/Dp53), the cells still arrested in S phase when incubated with SN38, but were now sensitive to UCN-01, progressed through the cell cycle, and also underwent a lethal mitosis [30]. Hence, p53 primarily protects cells by preventing the abrogation of S and G2 arrest, not by causing a G1 arrest.

12.4 How Does p53 Protect Cells from Chk1 Inhibitors? p53 is a transcriptional activator and many genes are induced: most notably for cell cycle checkpoint regulation is the CDK inhibitor p21waf1 (hereafter called p21). Less often considered is the fact that activation of p53 represses far more genes than it induces. We have investigated the impact of p21 on checkpoint abrogation in MCF10A cells using shRNA to prevent its induction

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upon DNA damage [31]. As expected, upon incubation with SN38, these MCF10A/Dp21 cells arrested in S phase. Upon addition of UCN-01, these cells were no longer resistant but progressed to G2. Surprisingly, the cells failed to undergo a lethal mitosis. With regard to S phase progression, it is logical to expect that p21 will inhibit Cdk activity. However, our results were inconsistent with this simple explanation. The activation of Chk1 by DNA damage causes phosphorylation and degradation of Cdc25A, thereby preventing activation of cyclin/Cdk complexes (primarily cyclin E/Cdk2 in S phase). Upon inhibition of Chk1 in p53 defective cells, Cdc25A reaccumulates and cells progress through the cycle. Therefore, we expected Cdc25A to accumulate in p53 wild-type cells incubated with UCN-01, but this was not observed. We found that DNA damage suppressed Cdc25A at the transcriptional level, and this occurred in a p21-dependent manner. The MCF10A/Dp21 cells were unable to repress Cdc25A mRNA, and therefore progressed to G2 when incubated with UCN-01. These results are summarized in Fig. 12.1. As noted above, MCF10A cells lacking p21 still arrested in G2 when Chk1 was inhibited. This is due to the p53 repressor function, and the critical genes repressed are cyclins A and B. MCF10A and MCF10A/Dp21 cells both exhibited strong repression of cyclin B, whereas the MCF10A/Dp53 cells were unable to repress cyclin B and therefore progressed through a lethal mitosis. As will be discussed below, these observations are critical to understanding which cells will respond to Chk1 inhibitors in the clinical setting. The ability of p53 to protect cells from Chk1 inhibitors has been questioned. Several reports have shown that UCN-01 can abrogate DNA damageinduced arrest independent of the p53 status [32, 33]. These differences have been resolved by considering the different drug schedules used [34]. When a Chk1 inhibitor is administered simultaneously with the DNA-damaging agent, there is no time for induction of the p53 response before the checkpoint is inhibited, and hence UCN-01 will increase cell killing independent of p53 status. However, if the DNA damage is allowed to first activate p53, the subsequent addition of a Chk1 inhibitor will have no impact on those cells. In addition to the immortalized MCF10A cells, we have also seen this phenotype in three p53 wildtype tumors: CAKI-1, U87MG, and SUM102 [31].

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Figure 12.1. Impact of p53 signaling on the pathways regulating DNA damage-induced S phase arrest (top box) and G2 arrest (bottom box). p53 induces expression of p21 while repressing transcription of cyclin B. The induced p21 then represses both transcription of Cdc25A and cyclin/Cdk activity.

This schedule with delayed addition of UCN-01 has been an important criterion in the design of many clinical trials. For example, in our initial clinical trial, cisplatin was administered to patients 24 h prior to commencing the UCN-01 infusion [35].

12.5 Some p53 Wild-Type Cells Are Still Sensitive to Chk1 Inhibitors While activation of p53 can protect many cells from subsequent incubation with a Chk1 inhibitor, it has become apparent that many cell lines commonly considered to be p53 wild-type elicit an ineffective p53 response when damaged. These cells remain

sensitive to UCN-01. Accordingly, there may be many p53 wild-type tumors that respond to the DNA damage plus Chk1 inhibitor combination. Tumor cell lines that we have found to be sensitive to this combination include HCT116, MCF7, and U2OS [31]. The HCT116 colorectal cancer cell line has been used extensively to study p53 signaling, and invaluable derivatives are available that are deleted for p53, p21 or other genes. These cells are well known to be defective for mismatch DNA repair, but it is less frequently realized that they also have a defect in another DNA damage response pathway consisting of the Mre11/Rad50/Nbs1 (MRN) complex [36]. When incubated with SN38, these cells arrest primarily in G2 rather than S phase, although the S phase arrest was reinstated by transfection with Mre11 [31, 37].


The addition of UCN-01 to G2-arrested HCT116 cells induces a mitotic catastrophe (further discussion of this point appears below) [31]. While SN38 induced p21 in the HCT116 cells, comparison to MCF10A cells showed that this induction was very limited. In addition, HCT116 cells were unable to repress cyclin B. Hence, they were readily able to undergo mitosis when incubated with UCN-01. In search of an explanation for the attenuated induction of p21, we first assessed the impact of DNA damage on the mRNA levels and found it to be significantly induced; yet the protein did not accumulate. In ongoing studies, we have discovered that the half-life of p21 protein is much shorter in HCT116 cells than MCF10A cells. The half-life of the protein in MCF7 and U2OS cells is also short and, as these cells are not deficient in MRN signaling, this does not appear to contribute to the mechanism. The reason for this rapid rate of degradation of p21 is currently under investigation. There are other interesting reports on a p53 wildtype cell line that is sensitive to Chk1 inhibition. The myeloid leukemia cell line ML1 has been used in a variety of studies with nucleoside analogs such as gemcitabine and fluodarabine nucleoside which stall the progression of replication forks [38–40]. These cells were first arrested in S phase with the nucleoside analog, and then UCN-01 was added after 24 h. The cells did not progress through the cell cycle but rather the replication forks collapsed and the cells underwent rapid apoptosis. This difference in fate may relate to the fact that leukemia cells, like the bone marrow, are more prone to apoptosis than epithelial cells. Importantly, these investigators showed that incubation with UCN-01 caused recovery of Cdc25A protein [39]. Considering the observation discussed above that p21 suppresses transcription of Cdc25A, this suggests that the p53 response had not been effectively induced. Unfortunately, the level of induction of p21 was not reported (only its binding to cyclin/Cdk complex was shown), but its level must have been insufficient to suppress the transcription of Cdc25A. However, as the cells did not need to progress through the cell cycle prior to dying, it remains to be determined whether p53 and/ or p21 can protect the cells from collapse of the stalled replication forks. Similar to the observations with nucleoside analogs, we have also reported a situation in which S phase-arrested cells can die without first

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progressing through S phase [29]. This occurred when MDA-MB-231 cells were incubated with SN38 to arrest them in S phase followed by a high concentration of UCN-01 (>100 nM). An important part of this experimental protocol was the use of the DNA polymerase inhibitor aphidicolin which prevented S phase progression under all conditions but only prevented cell death induced by low concentrations of UCN-01. This suggested a secondary target that was inhibited at high concentrations of UCN-01. We proposed that this second target could be C-TAK1, a kinase that constitutively phosphorylates Cdc25C to prevent premature mitosis and is another known target of UCN-01 [18]. Accordingly, we concluded that when both Chk1 and C-TAK1 are inhibited in S phase, cells can directly undergo a lethal mitosis without having to complete DNA synthesis. The p53 wild-type MCF10A cells remained resistant to UCN-01 at all concentrations, suggesting that p53 could prevent death even when cell cycle progression was not required, but whether p21 or cyclin B is more critical remains to be determined.

12.6 Two Experimental Caveats Aphidicolin has commonly been used as a model of S phase arrest, but caution should be taken in considering this a checkpoint response. While aphidicolin does stall replication forks and activate Chk1, it is impossible for cell cycle progression to occur even if Chk1 is inhibited because the DNA polymerase will remain inhibited. Similarly, hydroxyurea arrests cells in S phase by inhibiting ribonucleotide reductase, thereby depleting the deoxyribonucleotides that are essential for DNA synthesis. Finally, the nucleoside analog 5-fluoruracil pseudo-irreversibly inhibits thymidine synthase, thereby preventing synthesis of essential thymidine. Unless these drugs are removed, and in the latter case synthesis of new thymidine synthase occurs, the cells will remain arrested independent of Chk1 activity. These points have frequently not been taken into consideration in some of the published literature. For example, it has been concluded that cells lacking ATM and ATR have an additional kinase that is responsible for aphidicolin- or hydroxyurea-mediated cell cycle arrest [23, 41]. Similarly, a novel checkpoint response

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was reported in 5-fluoruracil-treated cells [42]. Regrettably, these conclusions are incorrect because of the failure to realize that these drugs will arrest DNA synthesis without the need for a checkpoint response. Clearly, it is critical to keep in mind the mechanism of action of the DNA-damaging agents under investigation. A second experimental caveat arises from the reliance on flow cytometry to assess the phase of the cell cycle and in particular whether cells have undergone mitosis. Flow cytometry does not discriminate between cells in G2, M, or post-mitotic cells that have failed to undergo cytokinesis (i.e., cells in a tetraploid G1 state). In our initial experiments with HCT116 cells incubated with SN38, we concluded that the addition of UCN-01 did not induce mitosis. However, we noted microscopically that the cells incubated with SN38 plus UCN-01 underwent mitotic catastrophe as judged by cells with multiple small nuclei. Analysis of geminin, a marker of S and G2, also showed that these cells were post-mitotic, yet they still had a G2/M DNA content because they had failed to undergo cytokinesis [31]. An alternative method to discriminate G2 from M, the mitotic phosphorylation of histone H3, would also fail to detect these post-mitotic tetraploid cells. Accordingly, the reliance on flow cytometry to assess sensitivity to Chk1 inhibition may impact the interpretation of experimental results and could lead to erroneous conclusions.

12.7 Why Doesn’t Activation of p53 Kill the Patient? The focus of the majority of research on p53 as a therapeutic target has been on the ability to activate or manipulate its expression in the tumor, but of more relevance to the current discussion is the impact of p53 on normal cells. The proposal that p53 protects normal cells may appear contradictory to the well-established role of p53 as an inducer of apoptosis. We have discussed this point in a recent review where we suggested that induction of apoptosis as a consequence of p53 activation has been greatly exaggerated [34]. Briefly, we note that much of the literature is influenced by the seminal report that thymocytes from p53-knockout mice are resistant to g-radiation [43]. A subsequent

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report showed that a fibrosarcoma derived from the same p53-knockout mice was also resistant to g-radiation [44]. In contrast, a spontaneous T-cell lymphoma derived from the same mice showed no differential response to g-radiation [45], yet this report is frequently ignored. Furthermore, there are numerous reports of p53 defective cells undergoing apoptosis following DNA damage, and many cases where the loss of p53 appears to sensitize cells to DNA damage. The explanation of these discrepancies relates to the cellular context which impacts the differential expression of pro-apoptotic vs. growth arrest genes. I refer the reader to our previous review for a detailed discussion [34]. Of particular relevance to the current review is the impact of p53 on the response of normal cells in vivo. Whole-body irradiation of mice induces extensive apoptosis in thymus, spleen, intestinal epithelium, and oral mucosa, with mice dying as a consequence of the bone marrow suppression [46]. Activation of p53 can be seen in multiple tissues, but the induction of p21 rather than pro-apoptotic genes is observed in tissues that do not undergo apoptosis [47]. Interestingly, while p53-knockout mice are more resistant to lethal bone marrow toxicity, they are more sensitive to gastrointestinal toxicity, and this sensitivity is also observed in p21 knockout mice [46]. It is evident that p21 can protect many tissues by preventing cell cycle progression, particularly when the cells are stressed by DNA damage. Much of the research on the role of p53 has been performed with g-radiation. The profile of toxicity resulting from other DNA-damaging agents can vary significantly from that of radiation. While DNA-damaging agents induce myelosuppression, the limiting toxicities are frequently mediated at other organs. Even p53-mediated apoptosis induced by g-radiation is limited in the patient by fractionated low doses and by targeting the radiation to the tumor rather than sensitive tissues. This discussion raises the important question as to whether the observations with many of these experimental models are relevant to the therapeutic situation. It is critical to resolve whether activation of p53 contributes significantly to patient toxicity or whether it protects patients. Perhaps the most striking example showing resistance of normal cells to p53 activation comes from the recent development of small molecules such as nutlin, RITA, and MI-219, which inhibit the binding


of p53 to MDM2 (sometimes known as HDM2 in human cells) [48–50]. The rationale for their development has been that such inhibitors will activate p53 in p53 wild-type tumors and thereby kill them. Yet should they not have the same impact on normal cells? This is clearly not the case. In a very exciting advance, it has been shown that MI-219 is selectively toxic to tumors grown in mice, and can induce complete tumor regression [50]. Importantly, it was nontoxic to the host. In several tissues studied, there was a robust upregulation of p21 in a p53-dependent manner. It must be concluded that activation of p53 in normal cells is not pro-apoptotic but it can be used to protect cells by arresting their progression through the cell cycle.

12.8 A Strategy to Enhance the Activity of Checkpoint Inhibitors In Vivo I have so far described the evidence that activation of p53 can protect cells from checkpoint inhibitors and have suggested this strategy may be selectively cytotoxic to tumors. I have explained the need for DNA damage to fully activate p53 in the normal tissues prior to adding the Chk1 inhibitor as a means to avoid toxicity to the normal cells. The addition of the Chk1 inhibitor after DNA damage is also based on the premise that cells require time to repair DNA damage, and that inhibition of Chk1 will limit the time available, thereby enhancing cell death. Because of the delayed addition of the Chk1 inhibitor, this raises the question as to how much DNA repair may have occurred already, and whether earlier inhibition of Chk1 would provide greater cell killing. These possibilities and the following discussion are graphically represented in Fig. 12.2. There is a second advantage to earlier inhibition of Chk1, or even concurrent administration of a DNA-damaging agent and Chk1 inhibitor. The amount of DNA damage a cell experiences is self-limiting because of the activation of Chk1. For example, inhibition of topoisomerase I induces relatively few double-strand breaks before the checkpoint is activated to prevent ongoing replication and thereby limiting further DNA breaks. However, if replication is forced to continue by inhibition of Chk1, the number of DNA breaks

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will increase and, presumably, cytotoxicity will be enhanced. Instead of just limiting the time for DNA repair, concurrent incubation can also enhance the initial amount of DNA damage. While the concurrent addition of UCN-01 and DNA damage may be more effective at killing tumor cells, this will be of no value if it also enhances killing in p53 wild-type cells, and is thereby toxic to the patient. In the previous section, I discussed the observation that an inhibitor of p53:MDM2 activated p53 in normal tissues but was not toxic to the host. We therefore propose that these inhibitors could be used to activate p53 in the patient’s normal cells and thereby protect the patient from the damage plus checkpoint inhibitor combination. As originally proposed, one would only have considered prescribing a p53:MDM2 inhibitor to a patient with a p53 wild-type tumor, as it would have no impact if p53 were defective. However, we believe that these p53:MDM2 inhibitors should be clinically tested for their ability to protect patients from DNA-damaging agents either when administered alone or when used in combination with Chk1 inhibitors. We believe this will enhance the therapeutic window for killing of p53 defective tumors.

12.9 It Still Isn’t That Simple: Inhibition of Chk1 Does Not Abrogate Arrest in All p53 Defective Tumors I have so far implied that arrested p53 defective tumors will be sensitive to UCN-01 and other Chk1 inhibitors, while some but not all p53 wild-type tumors will be resistant. I have not yet addressed the question of whether all p53 defective tumors will be sensitive, but the answer unfortunately appears to be no. We have found several cell lines that arrest in S phase when incubated with SN38 but do not progress through S when incubated with UCN-01. The mechanism of resistance is not related to drug uptake or metabolism as the cells are also resistant to incubation with caffeine. For example, the breast cancer cell lines T47D and the human embryo kidney cell line HEK293 both remain arrested in S phase and Cdc25A does not reaccumulate when UCN-01 is added. Preliminary results have shown that DNA

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Figure 12.2. Impact of the schedule of DNA damage and Chk1 inhibition on cell death in normal p53 wildtype normal cells vs. p53 defective tumor cells. (a) Impact of sequential administration of DNA damage and Chk1 inhibitor. (b) Impact of concurrent administration of the two agents. (c) Impact of prior activation of the protective p53 response using approaches such as a p53:MDM2 inhibitor.

damage induces loss of Chk1 in the HEK293 cells through a post-transcriptional mechanism; other proteins are also decreased which likely explains their persistent S phase arrest despite the depletion of Chk1. In contrast, Chk1 is not decreased in T47D cells suggesting there are at least two different mechanisms of resistance to UCN-01. Finally, we have recently reported an unexpected phenotype in MDA-MB-231 cells in which shRNA-mediated

suppression of Chk1 prevented SN38-induced S phase arrest, but the cells still arrested in G2 [20]. There obviously remain additional checkpoint regulators to be discovered. Perhaps the most critical consequence of these resistant phenotypes is that the same can be expected in some patients’ tumors and this could occur on either a frequent or rare occasion. It is possible that the optimism for Chk1 inhibitors in the clinical setting will be unrealized if


the drugs are tested against resistant tumors. Hence, there is a critical need to better understand which tumors will respond to these inhibitors. Acknowledgments: The research from my laboratory that is reviewed here represents the work of many dedicated post-doctoral fellows, students, and technicians. The main contributors to this research program over the past 15 years were Catherine Demarcq, Todd Bunch, Ethan Kohn, Aime Levesque, Edward Bresnick, Wen-Hui Zhang, Andrew Fanous, and Alissa Poh. The research has been supported by the Norris Cotton Cancer Center and the American Cancer Society, and continues to be supported by the National Cancer Institute (CA117874).

References 1. Rauth AM (1967) Evidence for dark reactivation of ultraviolet light damaged DNA in mouse L cells. Radiat Res 31:121–138 2. Lau CC, Pardee AB (1982) Mechanism by which caffeine potentiates lethality of nitrogen mustard. Proc Natl Acad Sci USA 79:2942–2946 3. Blasina A, Price BD, Turenne GA, McGowan CH (1999) Caffeine inhibits the checkpoint kinase ATM. Curr Biol 9:1135–1138 4. Sarkaria JN, Busby EC, Tibbetts RS et al (1999) Inhibition of ATM and ATR kinase by the radiosensitizing agent, caffeine. Cancer Res 59:4375–4382 5. Bunch RT, Eastman A (1996) Enhancement of cisplatin-induced cytotoxicity by 7-hydroxystaurosporine (UCN-01), a new G2-checkpoint inhibitor. Clin Cancer Res 2:791–797 6. Wang Q, Worland PJ, Clark JL, Carlson BA, Sausville EA (1995) Apoptosis in 7-hydroxystaurosporinetreated T lymphoblasts correlates with activation of cyclin-dependent kinases 1 and 2. Cell Growth Different 6:927–936 7. Eastman A, Jennerwein MM, Nagel D (1988) Characterization of bifunctional adducts produced in DNA by trans-diamminedichloroplatinum(II). Chem Biol Interact 67:71–80 8. Sorenson CM, Eastman A (1988) Mechanism of cisdiamminedichloroplatinum(II)-induced cytotoxicity: role of G2 arrest and DNA double-strand breaks. Cancer Res 48:4484–4488 9. Sorenson CM, Barry MA, Eastman A (1990) Analysis of events associated with cell cycle arrest at G2 and cell death induced by cisplatin. J Nat Cancer Inst 82:749–754

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10. Demarcq C, Bunch RT, Creswell D, Eastman A (1994) The role of cell cycle progression in cisplatininduced apoptosis in Chinese hamster ovary cells. Cell Growth Different 5:983–993 11. Russell KJ, Wiens LW, Demers GW, Galloway DA, Plon SE, Groudine M (1995) Abrogation of the G2 checkpoint results in differential radiosensitization of G1 checkpoint-deficient and G1 checkpointcompetent cells. Cancer Res 55:1639–1642 12. Powell SN, DeFrank JS, Connell P et al (1995) Differential sensitivity of p53(−) and p53(+) cells to caffeine-induced radiosensitization and override of G2 delay. Cancer Res 55:1643–1648 13. Fan S, Smith ML, Rivet DJ et al (1995) Disruption of p53 function sensitizes breast cancer MCF-7 cells to cisplatin and pentoxifylline. Cancer Res 55:1649–1654 14. Wang Q, Fan S, Eastman A, Worland PJ, Sausville EA, O’Connor PM (1996) UCN-01: a potent abrogator of G2 checkpoint function in cancer cells with disrupted p53. J Nat Cancer Inst 88:956–965 15. Bucher N, Britten CD (2008) G2 checkpoint abrogation and checkpoint kinase-1 targeting in the treatment of cancer. Br J Cancer 98:523–528 16. Sanchez Y, Wong C, Thoma RS et al (1997) Conservation of the Chk1 checkpoint pathway in mammals: linkage of DNA damage to Cdk regulation through Cdc25. Science 277:1497–1501 17. Graves PR, Yu L, Schwarz JK et al (2000) The Chk1 protein kinase and the Cdc25C regulatory pathways are targets of the anticancer agent UCN-01. J Biol Chem 275:5600–5605 18. Busby EC, Leistritz DF, Abraham RT, Karnitz LM, Sarkaria JN (2000) The radiosensitizing agent 7-hydroxystaurosporine (UCN-01) inhibits the DNA damage checkpoint kinase hChk1. Cancer Res 60:2108–2112 19. Yu Q, Rose JL, Zhang H, Takemura H, Kohn KW, Pommier Y (2002) UCN-01 inhibits p53 up-regulation and abrogates g-radiation-induced G2-M checkpoint independently of p53 by targeting both of the checkpoint kinases, Chk2 and Chk1. Cancer Res 62:5743–5748 20. Zhang W-H, Poh A, Fanous AA, Eastman A (2008) DNA damage-induced S phase arrest in human breast cancer depends on CHK1, but G2 arrest can occur independently of Chk1, Chk2 or MAPKAPK2. Cell Cycle 7:1668–1677 21. Davies SP, Reddy H, Caivano M, Cohen P (2000) Specificity and mechanism of action of some commonly used protein kinase inhibitors. Biochem J 351:95–105 22. Reinhardt HC, Aslanian AS, Lees JA, Yaffe MB (2007) p53-Deficient cells rely on ATM- and ATR-mediated checkpoint signaling through the p38MAPK/MK2 pathway for survival after DNA damage. Cancer Cell 11:175–189

198 23. Rodriguez-Bravo V, Guaita-Esteruelas S, Salvador N, Bachs O, Agell N (2007) Different S/M checkpoint responses of tumor and non-tumor cell lines to DNA replication inhibition. Cancer Res 67:11648–11656 24. Zhao B, Bower MJ, McDevitt PJ et al (2002) Structural basis for Chk1 inhibition by UCN-01. J Biol Chem 277:46609–46615 25. Sato S, Fujita N, Tsuruo T (2002) Interference with PDK1-Akt survival signaling pathway by UCN-01 (7-hydroxystaurosporine). Oncogene 21:1727–1738 26. King FW, Skeen J, Hay N, Shtivelman E (2004) Inhibition of Chk1 by activated PKB/Akt. Cell Cycle 3:634–637 27. Puc J, Keniry M, Li HS et al (2005) Lack of PTEN sequesters CHK1 and initiates genetic instability. Cancer Cell 7:193–204 28. Bozulic L, Surucu B, Hynx D, Hemmings BA (2008) PKBa/Akt1 acts downstream of DNA-PK in the DNA double-strand break response and promotes survival. Mol Cell 30:203–213 29. Kohn EA, Ruth ND, Brown MK, Livingstone M, Eastman A (2002) Abrogation of the S phase DNA damage checkpoint results in S phase progression or premature mitosis depending on the concentration of UCN-01 and the kinetics of Cdc25C activation. J Biol Chem 277:26553–26564 30. Levesque AA, Kohn EA, Bresnick E, Eastman A (2005) Distinct roles for p53 transactivation and repression in preventing UCN-01-mediated abrogation of DNA damage-induced S and G2 cell cycle checkpoints. Oncogene 24:3786–3796 31. Levesque AA, Fanous AA, Poh A, Eastman A (2008) Defective p53 signaling in p53 wildtype tumors attenuates p21waf1 induction and cyclin B repression rendering them sensitive to Chk1 inhibitors that abrogate S and G2 arrest. Mol Cancer Therap 7:252–262 32. Husain A, Yan X-J, Rosales N, Aghajanian C, Schwartz GK, Spriggs DR (1997) UCN-01 in ovary cancer cells: effective as a single agent and in combination with cis-diamminedichloroplatinum(II) independent of p53 status. Clin Cancer Res 3:2089–2097 33. Hirose Y, Berger MS, Pieper RO (2001) Abrogation of the Chk1-mediated G2 checkpoint pathway potentiates temozolomide-induced toxicity in a p53-independent manner in human glioblastoma cells. Cancer Res 61:5843–5849 34. Levesque AA, Eastman A (2007) p53-based cancer therapies: is defective p53 the Achilles heel of the tumor? Carcinogenesis 28:13–20 35. Perez RP, Lewis LD, Beelen AP et al (2006) Modulation of cell cycle progression in human tumors: a pharmacokinetic and tumor molecular pharmacodynamic study of cisplatin plus the Chk1 inhibitor UCN-01 (NSC 638850). Clin Cancer Res 12:7079–7085

A. Eastman 36. Giannini G, Ristori E, Cerignoli F et al (2002) Human MRE11 is inactivated in mismatch repairdeficient cancers. EMBO Rep 3:248–254 37. Takemura H, Rao VA, Sordet O et al (2006) Defective Mre11-dependent activation of Chk2 by Ataxia telangiectasia mutated in colorectal carcinoma cells in response to replication-dependent DNA double strand breaks. J Biol Chem 281:30814–30823 38. Shi Z, Azuma A, Sampath D, Li Y-X, Huang P, Plunkett W (2001) S-phase arrest by nucleoside analogues and abrogation of survival without cell cycle progression by 7-hydroxystaurosporine. Cancer Res 61:1065–1072 39. Sampath D, Shi Z, Plunkett B (2002) Inhibition of cyclin-dependent kinase 2 by the Chk1-Cdc25A pathway during the S phase checkpoint activated by fludarabine: dysregulation by 7-hydroxystaurosporine. Mol Pharmacol 62:680–688 40. Ewald B, Sampath D, Plunkett W (2007) H2AX phosphorylation marks gemcitabine-induced stalled replication forks and their collapse upon S-phase checkpoint abrogation. Mol Cancer Therap 6:1239–1248 41. Brown EJ, Baltimore D (2003) Essential and dispensable roles of ATR in cell cycle arrest and genome maintenance. Genes Dev 17:615–628 42. Xiao Z, Xue J, Sowin TJ, Rosenberg SH, Zhang H (2005) A novel mechanism of checkpoint abrogation conferred by Chk1 downregulation. Oncogene 24:1403–1411 43. Lowe SW, Schmitt EM, Smith SW, Osborne BA, Jacks T (1993) p53 is required for radiation-induced apoptosis in mouse thymocytes. Nature 362:847–849 44. Lowe SW, Bodis S, McClatchey A et al (1994) p53 status and the efficacy of cancer therapy. Science 266:807–810 45. Strasser A, Harris AW, Jacks T, Cory S (1994) DNA damage can induce apoptosis in proliferating lymphoid cells via p53-independent mechanisms inhibitable by Bcl-2. Cell 79:329–339 46. Komarova EA, Kondratov RV, Wang K et al (2004) Dual effect of p53 on radiation sensitivity in vivo: p53 promotes hematopoietic injury, but protects from gastrointestinal syndrome in mice. Oncogene 23:3265–3271 47. Fei P, Bernhard EJ, El-Deiry WS (2002) Tissuespecific induction of p53 targets in vivo. Cancer Res 62:7316–7327 48. Vassilev LT, Vu BT, Graves B et al (2004) In vivo activation of the p53 pathway by small-molecule antagonists of MDM2. Science 303:844–848 49. Issaeva N, Bozko P, Enge M et al (2004) Small molecular RITA binds to p53, blocks p53-HDM-2 interaction and activates p53 function in tumors. Nat Med 10:1321–1328 50. Shangary S, Qin D, McEachern D et al (2008) Temporal activation of p53 by a specific MDM2 inhibitor is selectively toxic to tumors and leads to complete tumor growth inhibition. Proc Natl Acad Sci USA 105:3933–3938

Chapter 13

Targeting p21-Dependent Pathways for Cell Death in Cancer Therapy Zahid H. Siddik

Abstract Checkpoint activation is a natural response to DNA damage that enables cells to take appropriate action, which results in cells either living or dying. A major target in checkpoint response is the tumor suppressor p53, which becomes stabilized and activated following the post-translation modifications. The activated p53 then transactivates the downstream p21 gene to inhibit cyclin-dependent kinase (CDK), induce cell cycle arrest to allow DNA repair, and enable cells to survive. If DNA damage is excessive, then p53-dependent apoptosis is affected. Since cell cycle arrest precedes cell death, it is not surprising that checkpoint response has been correlated with apoptosis, and the possibility that these two processes cross-communicate is supported by evidence that p21, like p53, is a tumor suppressor, and its expression is downregulated in apoptosis-defective cancers. In addition to affecting apoptosis through checkpoint response, p21 can also induce this mode of cell death through a different mechanism that involves repression of anti-apoptotic genes. It becomes apparent that p21 may be an important component of not only checkpoint response, but also apoptosis. Keywords DNA damage • p21 • Cyclin-dependent kinases • G1 checkpoint response • Cell cycle arrest • Transactivation • Gene repression • ApoptosisZ.H. Siddik

13.1 Introduction Research over the past two decades has established that DNA damage does not kill cancer cells imme-

diately. Instead, pre-determined steps are initiated that allow checkpoint control pathways to sense the type of DNA damage and then transduce appropriate signals to stop the cell cycle at the G1/S and/or G2/M transition points [1, 2]. This permits DNA repair and prevents DNA replication or mitosis in the presence of damaged chromosomes. As may be anticipated, these effects promote cell survival. However, if DNA damage is high and exceeds the capacity of the DNA repair machinery to complete the repair in a timely manner, the cells go on to die, possibly through signaling pathways activated by checkpoint response that trigger apoptosis. This is consistent with the literature reports that tumor cells retaining an intact G1 checkpoint response and are susceptible to G1-phase arrest are also drug-sensitive [3–5]. Such a relationship has been reinforced with the class of platinum antitumor agents in the NCI 60-cell line tumor panel [6]. In this regard, there is a contradiction to consider: tumor cells generally have deregulated G1-S transitions to promote proliferation but often respond to antitumor agents by activating G1 checkpoint response and arrest in G1/S [7]. It is likely that pathways inducing G1 cell cycle arrest following DNA damage over-ride deregulated pathways, which promote cell proliferation. The DNA damage checkpoint is an integrated network of signal transduction pathways that allow cells to transfer information from the DNA lesion to the cell cycle machinery via sensor, transducer, and effector signaling proteins [8]. This results in the inhibition of CDK activities, which arrests or slows the cell cycle. Inhibition of CDK4/cyclin D1


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and CDK2/cyclin E activities is associated with G1-phase checkpoint response to prevent G1-S transition, CDK2/cyclin A with S-phase checkpoint response, and Cdc2/cyclin A and Cdc2/cyclin B with the G2-phase checkpoint response to prevent G2-M transition [1, 8]. CDK inhibition may be caused by downregulation of cyclins, inhibition of binding between CDK and its cyclin partner, induction of inhibitory tyrosine phosphorylation of CDKs, inhibition of activating threonine phosphorylation of CDKs, or binding of INK4 or Cip/ Kip inhibitors to CDK [2]. In general, cyclin degradation and CDK tyrosine phosphorylation cause a rapid but transient inhibition of CDK activities. In contrast, inhibition of CDK complex formation and upregulation of CDK inhibitors (CDKI) cause delayed but sustained inhibition of CDK activities. CDKI from the INK4 family (p16INK4A, p15INK4B and p18INK4C) specifically inhibits the CDK4 complex during the G1 phase of the cell cycle. In contrast, the Cip/Kip family members (p21, p27 and p57) inhibit most CDK/cyclin complexes in all phases of the cell cycle. Among G1-phase CDKIs, the role of p21 may be the most significant in DNA damage-induced checkpoint response since p21 deletion alone prevents G1-phase arrest [1, 2]. Although p21 can also inhibit CDK activities in S- and G2-phase, its requirement is not as critical since other mechanisms exist to inhibit these CDKs [2]. The positive relationship between the presence of an intact G1 checkpoint response and drug sensitivity implicates p21 as a central player in antitumor response, and indeed its recognition as a tumor suppressor attests to this claim [9]. Consequently, this specific CDK inhibitor, with its involvement in G1 arrest and cell death, is the focus of attention in this chapter.

13.2 Molecular Aspects of CDK Inhibition by p21 CDK complexed with its cognate cyclin partner is active, and its inhibition by p21 is necessary to arrest cells at G1/S when DNA is damaged. In such arrested cells, p21 in fact is found in the nucleus where CDKs are also located [10]. CDK inhibition by p21 is a bimodal process: for cell cycle arrest with ionizing radiation, the most commonly studied DNA damaging agent, a p53-independent inhibition first

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occurs within 10 h resulting from redistribution of p21 from the CDK4/cyclin D1 complex to CDK2/ cyclin E, and then maintenance of this inhibition by an increase in p21 levels in a p53-dependent manner [11]. This implies that p21 is already present in the active CDK4/cyclin D1 complex, where reports suggest that it has an important role as an assembly factor [12, 13]. This role conveys the appearance that p21 doesn’t have an inhibitory effect against the CDK4 complex. However, increases in p21 levels following DNA damage are known to inhibit both CDK4/cyclin D1 and CDK2/cyclin E [14]. Therefore, it follows that separate regulatory mechanisms exist that permit p21 to be not only an effective assembly factor of the CDK complex, but also a potent inhibitor of CDK activity. To explain this paradox, several reports have offered the explanation that a single molecule of p21 is necessary to promote assembly of the CDK complex, whereas further recruitment of p21, when stoichiometry relative to CDK is ³2, leads to CDK inhibition [15–17]. This model, however, is not universally accepted as independent studies have demonstrated that a single molecule of p21 is sufficient to inhibit the CDK complex [18, 19]. Although it is clear that p21 can inhibit CDK directly, an indirect inhibition is also reported through induction of p27, which has a close structural homology with p21 and is functionally similar to p21 in slowing cell growth by inhibiting G1-phase CDK [20–22]. Although induction of p27 is observed with several DNA-damaging agents, including UV and chemotherapeutic drugs such as platinum-based drugs [20, 23, 24], coinduction of p21 and p27, and concomitant growth inhibition has also been demonstrated [25–27]. This is consistent with the established role of 27 as a tumor suppressor also, and, moreover, decreased levels of p27 have been associated with a number of cancers [28, 29] and drug resistance [30]. Evidence suggests that induction of p27 is through a stabilization process, and in a scenario that is well accepted, this involves p27 bound specifically to the CDK2/cyclin E complex, but not CDK4/cyclin D1. This binding is normally transient, primarily since bound p27 is immediately phosphorylated at Thr-187 by active CDK2/cyclin E, then recognized by the E3 ubiquitin ligase SCF/Skp2 complex and rapidly degraded by the proteosomal system [21, 28, 29, 31, 32]. In the stabilization process, the p21 induced by DNA damage inhibits CDK2/cyclin E,

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prevents p27 phosphorylation and, thereby, p27 degradation [33, 34]. In this manner, p27 cooperates with p21 to robustly and independently inhibit CDK2/ cyclin E, and a recent estimate suggests that about 75% of the kinase complex molecules are inhibited by p21, and 25% by p27 [24]. In addition to CDK, cyclin, and p21, PCNA can also be found in the G1-phase CDK complex to form a quartenary structure [35, 36]. PCNA is a multi-function protein that binds to a variety of partner proteins through different domains, including a number of cell cycle proteins such as CDK4, CDK2, p21, and cyclin D [37–39]. This binding appears to be transformation sensitive, in that PCNA is absent or much less abundant in CDK complexes from transformed cells [40]. However, it is not clear how PCNA may impact CDK activities. In some reports, addition of PCNA to the CDK2 complex in a cell free system does not appear to increase or inhibit CDK activities [18, 41]. This lack of effect is consistent with studies indicating that CDK complexes containing PCNA are active in exponentially growing cells [35, 36], and is further supported by the finding that PCNA is dispensable for p21-dependent G1-arrest induced by sodium butyrate [42]. On the other hand, contradictory evidence is also available to demonstrate that preventing p21 from binding PCNA attenuates its CDK-inhibitory functions [43–46]. In this regard, the most potent p21 peptide fragment for CDK inhibition contains the PCNA consensus sequence of amino acids 141-160 at the C-terminus of p21 [47, 48]. It appears, therefore, that the interaction between p21 and PCNA within the CDK complex may be critical for CDK inhibition, but the exact mechanism is not certain. It is possible that p21 may be stabilized within the CDK complex by PCNA that allows CDK inhibition to persist [49], but generalization of this concept requires caution since other reports suggest that proteosomal degradation of p21 is in fact increased by PCNA [50]. Thus, the significance of PCNA in the CDK complex remains unclear and requires further studies.

13.3 Regulation of p21

processes are usually involved in upregulating p21, but post-translational stabilization also has a major role in inducing this CDKI. Moreover, the functional activity of p21 in checkpoint response is dependent on nuclear levels of this inhibitor, and regulation of its distribution across the nuclear membrane is an important consideration.

13.3.1 Transcriptional Mechanisms Upregulating p21 Although p21 expression can be upregulated by posttranscriptional modifications that lead to enhanced stability of p21 mRNA [17, 51], transcriptional activation is usually the major mechanism for the induction of p21. The promoter region of the p21 gene is well endowed with consensus sites (response elements) for a number of transcriptional activators (Fig. 13.1), with p21 promoter activation involving both p53-dependent and -independent mechanisms [17]. Differentiation agents, including butyrate, SAHA, retinoic acid, and TGF-b, activate p21 transcription by p53-independent mechanisms, which may involve cooperative binding of other transcription factors (such as Sp1, Sp3, C/ EBPa, RAR and Smad3/4) to corresponding cisacting elements located within the p21 promoter. Similar p53-independent activation of p21 promoter by DNA-damaging agents has also been reported as with UV radiation [52, 53] and cisplatin [54]. In the case of cisplatin, the induction of p21 is thought to occur through the upregulation of E1AF, which then binds to the “ets” site to activate the promoter. p53 Sp1/3

p73 Vitamin D receptor (VDR)

Retinoic acid receptor (RAR)

C-EBP�/�

STAT1 / 3 / 5 p21 p21 Smad2 / 3 / 4

ets / E1AF Activator protein 2 (AP2)

CREB-binding protein (CBP) p300

E2F1 / 3 BRCA1

Insofar as p21 is critical for checkpoint response following DNA damage, it is of vital interest to understand how its induction is regulated. Transcriptional

Figure 13.1. Transcriptional activating factors of the p21 promoter. A number of activators upregulate p21 either alone or in cooperation with other factors.

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Of the various transcriptional activators that lead to p21 increases, p53-dependent transactivation of p21 is the most significant when DNA is damaged by cytotoxic agents [17]. This is consistent with reports that absence of p53 generally fails to inhibit G1-phase CDKs and, consequently, G1-arrest is not observed [8, 55]. Other transcriptional activators also cooperate with p53 to transactivate p21 and include Sp1, Sp3, CBP/p300, and BRCA1. Since p53 and HDAC1 both bind to the C-terminus of Sp1, the deacetylase can antagonize p53-dependent transactivation of p21 [56], and this raises the rationale for combining an HDAC inhibitor with a p53-dependent antitumor agent to increase p21 upregulation and suppress tumor growth. Indeed, the reported synergy between HDAC inhibitors with VP-16, ellipticine, doxorubicin, or cisplatin may be due to this effect [57]. Transactivation of p21 by p53 is one of the strongest responses to DNA damage by the cell, and this is consistent with the finding that promoter occupancy of p53 for the p21 gene is one of the highest among the various downstream gene targets [58, 59]. The strong response could also be due to the presence of at least two consensus sites for p53 on the p21 promoter located about 1.4 and 2.3 kb upstream of the transcription start site [17]. Although there is potential for cooperation between the two sites, it appears that this is not the case since binding of p53 to one site was shown not to influence the binding to the other [60]. However, there are quantitative differences in binding that are of interest. In tumor models exposed to 5-fluorouracil, the induced p53 binds to both the proximal and distal sites within a few hours of treatment, but the distal site has greater affinity for the tumor suppressor, with up to tenfold greater binding of p53 at this site [60, 61]. As may be expected from such a high affinity, minor conformational changes in p53 through mutations are tolerated by the promoter and permit p21 transactivation, albeit at reduced levels, although the majority of p53 mutants fail to bind to the p21 promoter and to transactivate [58, 62].

13.3.2 Post-translational Mechanisms of p21 Induction Increase in protein stability is an important mechanism for p21 induction. This stabilization can occur by protein–protein interactions that prevent proteosomal degradation, including the binding

Z.H. Siddik

with PCNA [49] or WISp39 [63]. An alternative mechanism for p21 stabilization is via post-translational modification, which in addition can regulate p21 function. This modification usually involves phosphorylation following the activation of specific kinases [64, 65]. Ser130 of p21 is one phosphorylation site, which is targeted by MAPKs [66, 67], and this likely involves activation of MAPK family members p38a, JNK1, and ERK following DNA damage by antitumor agents, as has been observed with cisplatin [68–70]. Similarly, the PI3K/Akt kinase pathway is essential for p53-dependent induction of p21 following DNA damage by cisplatin, taxol, or ionizing radiation and involves phosphorylation of p21 at the Thr145 and Ser146 sites [71, 72]. These two neighboring sites are within the PCNA binding domain of p21, and interestingly their phosphorylation is reported to inhibit p21–PCNA interaction and impede CDK inhibition and apoptosis [43–45, 73]. The possibility is raised, therefore, that PCNA within the tetrameric CDK/cyclin/p21/ PCNA complex may regulate functions of p21 in cell cycle arrest and drug sensitivity and could explain why p21 is capable of both promoting and inhibiting apoptosis [64].

13.4 Role of p21 in Apoptotic Response The significance of p21 in cellular homeostasis is so critical that tumor cells do not select for mutant p21, which, therefore, is a rare occurrence in human cancers [17, 74, 75]. Moreover, p21 is now considered to be a tumor suppressor, primarily since p21 knockout mice develop several types of cancers, including histiocytic sarcomas, hemangiomas and B-cell lymphomas, and all p21-deficient mice die substantially earlier than p21-proficient mice [9]. This tumor suppressive role has been confirmed independently in other systems [76, 77] and implies that p21 should facilitate cell death if DNA damage is detected and goes unrepaired. Indeed, a number of reports have provided definitive data to support the pro-apoptotic activity of p21, particularly when combined with antitumor agents, including cisplatin and 5-FU [64, 78]. This includes the demonstration that ectopic p21 can enhance drug-dependent cytotoxicity in ovarian OVCAR-3, SKOV-3, and 2774 tumor cell lines


that have defective p53 [79, 80]. These preclinical data are consistent with the clinical finding that cancers expressing p21 are associated with both a superior therapeutic outcome and an improved patient survival rate [81, 82]. Despite such apoptotic significance of p21, there is paucity of data, outside of its role in CDK inhibition and cell cycle regulation, to provide a mechanism for its proapoptotic effects. One of the mechanisms gaining acceptance involves p21-dependent suppression of anti-apoptotic genes, including survivin, stathmin, B-mYB, and aB-crystallin that are implicated in inhibiting apoptosis [80, 83]. Ironically, Chk1 as an activator of p53 is also known to be anti-apoptotic, and its eventual downregulation by p21 following antitumor drug treatment is necessary to induce cell death [84, 85]. Therefore, at least some of the well-documented gene repressive and apoptotic functions of p53 may actually be carried out indirectly by transactivated p21 (Fig. 13.2). It remains to be demonstrated convincingly whether p21 facilitates apoptosis through inhibition of CDK, more specifically G1-phase CDKs, which are more dependent on inhibition by p21 for checkpoint response [86]. However, there is ample evidence that tumor cells retaining G1 checkpoint response are more sensitive to DNA-damaging

DNA damage

ATM

Checkpoint signaling

Chk2

p53

Drug

ATR

Chk1

p21

CDK/cyclin inhibition Apoptosis

Anti-apoptotic proteins (e.g., Survivin, B-mYB)

Figure 13.2. Pathways of p21-dependent apoptosis. DNA damage upregulates ATR/Chk1 and/or ATM/Chk2 pathways, which induce and activate p53. Resultant transactivation of p21 gene by p53 is the major mechanism of p21 induction, which induces cell death by either suppressing anti-apoptotic genes or inhibiting G1-phase CDKs.

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agents [3–6], including the observation that some potent p53-dependent and -independent apoptotic drugs induce only G1 checkpoint response in an exclusively p21-dependent manner, with prominent absence of S- or G2/M-checkpoint responses [14, 87, 88]. Conversely, dominant negative CDK mutants prevent drug-induced apoptosis [89]. Furthermore, small molecule inhibitors of CDK activity not only arrest cells, but also induce apoptosis [74, 89, 90]. However, many of the chemical inhibitors are not specific, particularly at concentrations being employed in the literature reports, and so it is unclear whether they induce apoptosis specifically by targeting CDKs in the G1-phase or elsewhere. Nevertheless, studies demonstrate that the antitumor activity of roscovitine (CYC-202) and BMS-387032 is likely mediated through preferential inhibition of CDK2/Cyclin E [74, 89, 90]. Similarly, the triaminopyrimidine derivative CINK4 [91] and pyrido-[2,3-d]pyrimidin7-one derivative PD0332991 [92] demonstrate preferential CDK4 inhibition and potent cytotoxicity but interestingly were less effective against Rb-negative tumor cells [90]. This suggests that E2F sequestration by hypophosphorylated Rb, as seen in G1 checkpoint response with p21, may be a requirement for antitumor activity with these chemical inhibitors. Although there are ample demonstrations of apoptotic effects associated with p21, it is worth noting that p21 has also been implicated in inhibiting apoptosis and inducing drug resistance [64, 78]. A strong support for the anti-apoptotic role has come from studies in HCT-116 colorectal cells, which harbor wild-type p53, but following p21-knockout are sensitized to DNA-damaging agents, including ionizing radiation, adriamycin, etoposide, cisplatin, and nitrogen mustard [93, 94]. Sensitization is not universal, as it was not observed with taxol or vincristine, nor reproducible in an independent study, as in the case of ionizing radiation and adriamycin [94]. Moreover, similar cytotoxic sensitization with these agents are also observed when p53 is disrupted in parental HCT116 cells [94], and this suggests that HCT-116 cells are among a minority of model systems that have an unusual cell context where wild-type p53 is actually promoting the drug resistance phenotype. On the other hand, these HCT-116 cells were sensitive to the combination of either 5-FU and the

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antioxidant PDTC [95] or taxol and manumycin [96] in a p53-independent, but p21-dependent manner. In these combinations, PDTC and manumycin induced p21 and its induction then supported the cell killing activity of 5-FU and taxol, respectively. It appears, therefore, that in tumors cells harboring wild-type p53, p21 transactivation either in a p53-dependent or -independent manner facilitates apoptosis induced by the antitumor agent.

13.5 Loss of p21 Regulation in Drug Resistance Drug resistance is a major limiting factor in the treatment of cancer, and each tumor normally has several mechanisms of resistance that co-exist, and the combination of mechanisms represents a unique signature for that tumor. However, some mechanisms, such as loss of p53 function, are more critical and induce a greater degree of resistance. Since p21 is a tumor suppressor and has a role in apoptosis, it is not surprising that loss of p21 regulation impacts apoptosis and also contributes to the drug resistance phenotype. Several mechanisms of resistance specific to p21 have been identified, and available evidence for each will be provided in this section. For p21 to play a role in apoptosis, its presence in the nucleus is essential to either repress transcription of anti-apoptotic genes or facilitate apoptotic signaling by inhibiting G1-phase CDKs (see Fig. 13.2). Nuclear exclusion of p21 appears to be a frequent occurrence that impedes the apoptotic process and, thereby, allows cells to survive toxic insults by DNA damaging agents. Sequestration of p21 in the cytoplasm is largely mediated by post-translational modification and, as discussed earlier, this process is also required for p21 induction by protein stabilization [65]. Indeed, many of the same kinases are involved in both effects, and this raises conflicts in our understanding. A likely explanation for contradictory observations is that specific kinases are activated transiently for p21 stabilization, but their constitutive activation following deregulation induces resistance by maintaining p21 in a phosphorylated state in the cytoplasm. The PI3K/Akt pathway best exemplifies this, and it is localized in the cytoplasm where it phosphorylates p21 at Thr145 and Ser146 sites [71, 72]. This pathway

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is deregulated and constitutively active in a number of cancers, which sequesters p21 in the cytoplasm, disrupts the ability of p21 to attenuate proliferation, and induces drug resistance [97–99]. Since the PI3K/Akt pathway is regulated by a number of pathways, any deregulation upstream will also attenuate p21 function by nuclear exclusion and induce resistance. This is best seen in tumors demonstrating amplification or overexpression of HER-2/neu, which encodes the transmembrane receptor tyrosine kinase p185 and upregulates the PI3K/Akt pathway [99]. Thus, the poor 5-year survival rate in breast cancer patients overexpressing HER-2/neu has been ascribed to Akt-dependent phosphorylation of p21 at Thr145 and its cytoplasmic sequestration [100]. An alternative mode of increasing PI3K/Akt activity and sequestering p21 in the cytoplasm is through downregulation of the negative phosphatase regulator PTEN that is frequently encountered in refractory human cancers [101]. Defective p21 regulation also involves repression of its promoter activity as a result of deregulation of any one of the many transcriptional factors, including mutation, deletion or MDM2-dependent inhibition of p53 [102]. Interference of transcription factors by negative regulators is an alternative mechanism of p21 repression. For instance, c-Myc interacts with Sp1/Sp3 and prevents their ability to play a role in inducing p21 [103]. Similarly, the Rb binding protein HBP1 counters the function of the E2F1/E2F3 transcriptional factors to block p21 expression [104]. According to Kastan, activation of p53 alone is sufficient to kill tumor cells [105], presumably, in part, through p53-dependent upregulation of p21. However, this scenario is not always observed in drug resistant model systems, where the antitumor agent may activate the wild-type p53, but p21 fails to become transactivated. Such a defect is well demonstrated in tumor cells resistant to platinum drugs (cisplatin, oxaliplatin, satraplatin and BBR3464), with resistance correlated to loss of p21 expression and/or G1 checkpoint response [6, 106–110]. The mechanism involved in the inability of the activated wild-type p53 to transactivate p53 is unclear, but epigenetic silencing through hypermethylation of the p21 promoter is a possibility as it has been observed in tumor cells [102, 111]. Silencing usually involves hypermethylation at 5¢-CpG islands, which are located in the proximal


region, around position 50-119 upstream of the start site that harbors six conserved “GC box” binding sites for Sp1 and/or Sp3. Hypermethylation in this region, therefore, disrupts the ability of Sp1/Sp3 to bind to the promoter and activates the p21 promoter. However, CpG islands are not prerequisites to affect silencing, and methylation at a single CpG can be sufficient to disrupt p21 expression in some cases, as has been demonstrated with position 692, which is within the STAT response element of the promoter [102]. Indeed, p21 silencing through this STAT site has been found in 50% of rhabdomyosarcoma tumors [112], and hypermethylation of the proximal CpG islands has been observed in bone marrow cells from over 40% of acute lymphoblastic leukemia (ALL) patients [113]. This hypermethylation in ALL patients is associated with a poor survival rate of only 6–8% compared to about 60% when the promoter is hypomethylated. Hypermethylation of p21 promoter has also been observed in 34% of NK cell cancers [114].

13.6 Targeting p21 for Reversing Drug Resistance As indicated earlier, p21 is an important component of checkpoint response that is also strongly associated with tumor sensitivity. Therefore, in tumor cells with deregulated p21, restoring p21-dependent functions would seem to be a rational approach for treating such cancers. Where cytoplasmic sequestration of phospho-p21 is suspected as a mechanism of resistance, identification of the appropriate kinase (e.g., Akt) will determine the logical selection of one of several kinase inhibitors currently available [97]. It may be questioned, however, why such inhibitors should be effective, primarily since inhibition could prevent p21 induction, as may be possible in the case of Akt inhibition [71]. This is an important consideration, mainly because induced p21 is the essential component involved in apoptosis, whereas basal p21, which is expressed in resistant cells, is not [115]. Surprisingly, this has not been the case, and downregulation of Akt actually did enhance the sensitivity of tumor cells to antitumor agents, including cisplatin [116], although it is uncertain whether such an effect was mediated by inhibition of p21 phosphorylation. In general, p21 induction following DNA damage involves transcriptional processes and, therefore, a

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failure to increase levels of p21 is predominantly due to defects in transactivation. It follows then that designing small molecules to restore endogenous transcription of p21 would be the most logical and effective approach toward sensitizing tumor cells to antitumor drugs. This is consistent with reports that transfection of p21 vectors in tumor cells harboring defective p53 sensitizes these cells to DNAdamaging agents, including cisplatin, doxorubicin, and methotrexate [79, 117, 118]. The small molecule strategy to transcriptionally upregulate p21 can be either p53-dependent or p53-independent.

13.6.1 p53-Dependent Approach to Restoring p21 Expression Loss of p21 expression through gene silencing represents an important mechanism. Silencing involves promoter hypermethylation that impacts transcription because of several reasons, including interference with the binding of transcription factors to the promoter [119]. Hypermethylation occurs at cytosine residues within CpG dinucleotides by DNA methyltransferase 1 (DNMT1), which as a therapeutic target can be inhibited by several classes of chemical agents [119]. Of those available, the most widely used is decitabine, which is a DNA damaging agent that initially becomes incorporated into DNA and traps the DNMT1 by an irreversible covalent interaction. The result is that methylation is prevented following DNA replication, and gene expression is eventually restored. Because of the kinetics of decitabine-mediated inhibition of DNMT1, it normally takes a few cell cycles in the presence of low concentrations of the inhibitor to restore gene expression. Such concentrations, however, are not sufficient to reach a threshold of DNA damage that induces p53 and upregulates p21 transcription [120], and so decitabine is normally used in combination with another agent. That this strategy is feasible has been demonstrated in rhabdomyosarcoma cells, where p21 promoter hypermethylation at the STAT-binding site was reversed by a 48-h exposure to decitabine and sensitivity restored to STAT-1 when activated by interferon-g [112]. Silencing of the p21 promoter is rare in many tumor types, and mechanisms incapacitating p53 also have to be considered for loss of p21 expression. In this regard, it is notable that about 10–60% of advanced cancers retaining wild-type p53, including

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non-small cell lung, ovarian and breast cancers, are chemoresistant [121–126]. Indeed, the level of resistance in wild-type p53 cancers can be greater than in mutant p53 counterparts [106, 127]. There is evidence that in such cancers, p53 is induced but p21 expression remains at basal levels [107, 109]. This defect in the p53/p21 pathway, however, is drug specific in some cancers, in which case another agent with a different mechanism of action, even from the same class of antitumor drugs, could restore the pathway. This is amply demonstrated in two examples with platinum-based agents. In one, the p53/p21 pathway in cisplatin resistant ovarian 2780CP tumor cells is defective when cisplatin is used as a DNA-damaging agent, but responds effectively, as indicated by p21 expression, to the non-cross-resistant analog 1R,2R-diaminocyclohexane-diacetato-dicholoroPtIV (DAP) [107] or ionizing radiation [128]. In the second example, ovarian tumor cells resistant to a novel trinuclear platinum drug BBR-3464 did not express p21 when exposed to BBR-3464, but the p21 was inducible in a p53-dependent manner by cisplatin [109]. Induction of p21 in a tumor model with one agent and not another is further highlighted in a recent report with Rat-1 cells [111]: UV upregulated p21 in a p53-dependent manner and induced cell cycle arrest and apoptosis, whereas IR failed to induce p21 mRNA or protein, failed to arrest cells in G1, and failed to induce apoptosis. Moreover, transfection of Rat-1 cells with an artificial chromosome containing a genomic copy of the p21 gene restored p21 expression, G1 arrest and sensitivity to IR. Indeed, there are several examples in the literature of resistance to one agent and sensitivity to another, but these differences have not been correlated to relative p21 expression. This is best illustrated in MCF-7 and ZR-75-1 breast tumor models that have wildtype p53 and represent drug sensitive and resistant phenotype, respectively: in comparison to MCF-7 cells, ZR-75-1 cells demonstrated tenfold resistance to 5-fluorouracil (5-FU), but conversely were twofold more sensitive to doxorubicin [129]. Thus, resistance of ZR-75-1 cells to 5-FU can be circumvented by doxorubicin, and using such an understanding to develop novel agents will likely have clinical benefits against wild-type p53 cancers, presumably by circumventing resistance due to agent-specific loss of p21.

Z.H. Siddik

The understanding that loss of p21 upregulation is associated with resistance in wild-type p53 tumor models and restoration of p21 induction by another agent circumvents resistance, lends to the conclusion that the machinery for upregulating p21 is essentially intact. Since transcriptional mechanisms are usually involved, the p21 promoter becomes a focus of speculation to explain differential p21 expression between two antitumor agents. In this regard, differences in post-translational phosphorylation of p53 by the two agents could be a likely mechanism. For instance, in the case of differential cytotoxicity between UV and ionizing radiation against Rat-1 cells [111] or cisplatin and DAP against ovarian 2780CP cells [107], one agent (UV or cisplatin) induces p53 phosphorylation at Ser-392 while the other agent (ionizing radiation or DAP) does not [107, 130]. Similarly, DNA damage by UV [131], cisplatin [131], and 5-FU [132] activates ATR and/or Chk1, whereas ionizing radiation [133] and doxorubicin [134] activate ATM and/or Chk2, and these kinases will cause differences in post-translational modification of p53 at other sites [135, 136]. However, all these agents are known to be effective inducers of p21 in a p53-dependent manner, and one can conclude that transactivation of p21 in sensitive cells is independent of

Anti-tumor Agent 1 (e.g., cisPt)

Anti-tumor Agent 2 (e.g., DAP)

p53

p53

IL-6, EGF, IFN-γ

STAT

SAHA, TSA, Na Butyrate HDAC

Sp1/3 p21

−2301 −1394

−692

−119 to −50

Figure 13.3. Induction of p21 by p53-dependent and -independent mechanisms. Activated p53 can be qualitatively distinct depending on post-translational modifications that are specific to the DNA-damaging agent, such as cisplatin (cisPt) or the non-cross-resistant analog 1R,2R-diaminocyclohexane-diacetato-dicholoro-PtIV (DAP), and it can then bind to the two p53 response elements on the p21 promoter to induce p21. Other agents, such as cytokines or HDAC inhibitors, can upregulate different activating factors, which can similarly induce p21 in a p53-independent manner.


the qualitatively different form of p53 (Fig. 13.3). Observations in resistant cells, on the other hand, would strongly suggest that one form of p53 with its unique modifications might lose its capacity to affect p21 transactivation, whereas another form with a different combination of post-translational modifications is fully functional. Although it is not certain whether the two forms of p53 actually bind differentially to the consensus site on the p21 promoter in resistant cells, the concept that p21 expression can be re-activated in resistant cells by a qualitatively different form of p53 may allow screening for small molecule DNA-damaging agents that are more effective in restoring p21 and, therefore, in treating refractory cancers.

13.6.2 p53-Independent Approach to Restoring p21 Expression The p53-dependent approach for restoring p21 expression has an obvious limitation, in that it is restricted to tumor cells having wild-type p53. Therefore, a p53-independent tactic that may be applicable to both mutant and wild-type p53 cancers would have definite advantages. As noted earlier, tumor cells can be effectively sensitized to PDTC/5-FU or manumycin/taxol combination, where the role of antioxidant PDTC or the farnesyltransferase inhibitor manumycin is to induce p21 in order to sensitize cells to the cytotoxic agent [95, 96]. In these studies, the mechanism of p53-independent induction of p21 is uncertain. A rational approach would be to activate the p21 promoter through a known mechanism. In this respect, upregulating activating factors that bind to the p21 promoter downstream of p53 consensus sites may be the most logical approach, and this would allow induced p21 to inhibit tumor growth by itself or in synergy with DNA-damaging agents. There are a number of activating factors that have potential in this regard, including STAT1/3/5 and Sp1/3 [17, 102]. Each specific factor can be upregulated by several agents, and this is exemplified with STATdependent agents, such as IL-6, EGF, and IFN-g, and Sp-dependent agents, such as HDAC inhibitors SAHA, trichostatin A (TSA), and sodium butyrate (Fig. 13.3). Several HDAC inhibitors have shown synergy in combination with DNA damaging agents at the preclinical level, and such combinations are now entering clinical trials [137].

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However, the clearest evidence that antitumor synergy of such approaches is through expression of p21 is largely from preclinical investigations. For instance, the combination of SAHA and lexatumumab (an agonist monoclonal antibody to death receptor TRAIL-R2) was synergistic against parental HCT-116p21+/+ colorectal cells but not against p21-knockdown HCT-116p21−/− cells [138, 139]. More importantly, this synergy was also observed in TRAIL-resistant HCT8 and HT29 colorectal models and was independent of p53.

13.7 Conclusion The CDK inhibitor p21 has a critical role in G1 checkpoint response, and this response is intimately linked to cell death. Cytotoxic effects mediated by p21 may also occur through repression of antiapoptotic genes, such as survivin. It is not surprising, therefore, that loss of p21 function in the nuclear compartment is a feature of drug resistant tumor cells. This loss can occur through several mechanisms, including cytoplasmic sequestration and loss of transcription through promoter hypermethylation and downregulation of transactivating function of p53. Since p21 is not mutated or deleted in tumor cells, its induction through one of several approaches will likely re-sensitize tumor cells to cell death stimulus. Drug combinations that target p21 for upregulation, therefore, have the potential of reversing the drug-resistance phenotype, and need to be explored thoroughly at both the preclinical and clinical levels. Acknowledgements: Supported by NIH Grant RO1 CA127263.

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Chapter 14 p27Kip1 as a Biomarker and Target for Treatment of Cancer Xiao-Feng Le and Robert C. Bast Jr.

Abstract p27Kip1 (p27) is a critical cyclin dependent kinase inhibitor that negatively regulates cell cycle progression from G1 to S phase. Levels of nuclear p27 are reduced in many human cancers. Both the levels and function of p27 are regulated by transcriptional, translational, and posttranslational mechanisms. In regulating p27, posttranslational phosphorylation of the protein is particularly important for determining susceptibility to ubiquitination and proteolytic degradation, as well as the levels, subcellular localization, and functional activity of p27. In addition to the control of cell cycle, cytoplasmic expression of p27 can enhance cell motility, increase renewal of stem cells, inhibit apoptosis, induce autophagy, and stimulate tumor metastasis. Persistence of nuclear p27 provides an independent favorable prognostic factor in many human cancers. Expression or induction of p27 may provide a predictive biomarker for response to chemotherapy, hormonal therapy, and at least some targeted therapies including trastuzumab and Src inhibitors. In addition, restoration of nuclear p27 levels might be achieved pharmacologically, inhibiting proliferation of cancer cells. Inhibition of SKP2-mediated p27 ubiquitination provides one additional promising approach. Keywords p27Kip1 • Cell cycle • G1 arrest • Autophagy • Tumor suppressor • Targeted therapy • Trastuzumab

14.1 Introduction p27Kip1 (hereafter p27) belongs to a family of Cip/Kip cyclin-dependent kinase (CDK) inhibitors. Although p27 was first reported as an inhibitor of the cyclin E/A-CDK2 complex in growth-arrested cells [1, 2], it is now known to perform multiple cell cycle-dependent and cell cycle-independent roles that affect proliferation, motility, differentiation, survival, and autophagy [3–9]. The function and level of p27 are regulated by an array of signaling pathways that have an impact on transcriptional, translational, and posttranslational events [4–8]. Downregulation of p27 occurs during malignant transformation, affecting proliferation, prognosis, and response to treatment. In normal cells, p27 blocks cell cycle progression by inhibiting cyclin E-CDK2 and cyclin A-CDK2 activity [3]. Intact autocrine and paracrine growth inhibitory pathways can upregulate p27, producing cell cycle arrest [1–3]. Reduced levels of p27 protein are found in a variety of human cancers and are associated with a poor prognosis [3–5]. Loss of p27 in murine models leads to enhanced growth of normal organs and of malignant tumors [3]. With human and murine cancer cells, changes in levels of p27 can also predict response to treatment with chemotherapy, radiotherapy, endocrine manipulation, and molecularly targeted agents. Many of these interventions may depend upon functional p27 to be effective [4, 6, 9–13]. A germline mutation in


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the p27 gene or CDKN1B occurs rarely [3], but phosphorylation-mediated, posttranslational mechanisms largely control the expression and localization of p27 in normal and in cancer cells [3–5]. A significant number of anti-cancer drugs that are now available can modulate the posttranslational regulation of p27. In this chapter, the function of p27 as a tumor suppressor, prognostic factor, and a predictive biomarker for cancer therapeutics will be discussed in depth, as will the opposing functions of p27 as a protein that can enhance tumor mobility, survival, and metastasis.

14.2 Regulation of the p27 Gene and Protein To understand the role of p27 in normal cells and particularly in cancer cells, it is critical to know how levels and activity of p27 are regulated. The expression and function of p27 protein can be regulated at multiple levels through different mechanisms.

14.2.1 Transcriptional Regulation of the p27 Gene (CDKN1B) A number of transcription factors and microRNAs (miR) have been shown to regulate p27 gene expression, which are summarized in Table 14.1. Inhibitors of differentiation 3(Id3), a bHLH transcriptional repressor, directly associate with and regulate the activity of several families of transcriptional regulators including E2F [16]. Downregulation of CDKN1B expression by Id3 is a novel mechanism [17, 18]. miR221/222 are

X.-F. Le and R.C. Bast Jr.

also novel mechanisms that regulates both mRNA and protein levels of p27 [8]. It is worthy to point out that the factors involved in transcriptional regulation of CDKN1B may participate in other mechanisms to regulate p27 protein. In fact, FoxO transcriptional factors, under the control of PI3K signaling pathway, primarily modulate activity and localization of the p27 protein through posttranslational mechanism (see below) [19].

14.2.2 Translational Regulation of the p27 Protein Eukaryotes initiate translation through two major mechanisms: cap-dependent scanning and internal ribosome entry [23, 24]. For the majority of cellular mRNAs, translation is thought to be initiated by cap-dependent scanning [23]. Internal ribosome entry requires specific structures known as internal ribosome entry site (IRES) elements in the 5¢ untranslated region (5¢UTR) of an mRNA and is used by some viruses and a specific set of cellular mRNAs that are induced in response to stress signals [24–27]. p27 is one of the proteins that can depend upon IRES for initiation of transcription [25–27]. Embryonic lethal abnormal vision-like 1 in Drosophila (ELAVL1, Hu antigen R in humans) and ELAVL4 (Hu antigen D in humans) encode proteins that bind to the IRES of the p27 gene and inhibit translation of p27 protein [26, 27]. Conversely, two other proteins, the heterogeneous nuclear ribonucleoprotein C1/C2 [26] and polypyrimidine tract-binding protein (PTB) [28] have been shown to bind to the IRES of p27 mRNA and to promote translation. Most recently, miR221 and miR222 have been shown to bind independently to 3¢UTR of the p27 gene and to inhibit translation of p27 [8, 29, 30].

Table 14.1. Transcription factors and miR that regulate the p27 gene (CDKN1B) expression. Repressors c-Myc (14) HES1 (15) Inhibitor of DNA binding/differentiation 3 (Id3) (16-18) miRNA 221/222 (miR221/222) (8) Activators Forkhead box class O family (FoxO) (19) Menin (20) E2F1 (21)

14.2.3 Posttranslational Regulation of the p27 Protein After synthesis, p27 protein can be further modified by phosphorylation on serine, threonine, and tyrosine residues, mediating the modulation of p27 levels, location, and activity [4, 31–33]. To date, at least 7 important phosphorylation sites have been identified on the p27 protein. These include serine-10, tyrosine-74, tyrosine-88, tyrosine-89,

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14. p27Kip1 as a Biomarker and Target for Treatment of Cancer

Src /Abl / Lyn/ Bcr-Abl

hKIS/ AKT / MIRK/ MAPK S10

Y74/88/ 89

N

NES / CRM1 CDK /cyclin

Cytosolic localization; Nuclear export; p27 stabilization

p27/ K4 or K6/ D; Destabilization of p27

AKT

CDK2

AMPK AKT RSK

T157

T187

T198

C

NLS

Jab1

Nuclear export

Cytosolic localization; Nuclear export; Protein stabilization

p27/ 14-3-3; Cytosolic localization; p27 stabilization; Autophagy; p27/ K4 or K6/D

Figure 14.1. Schematic diagram of p27 unique structure, and its phosphorylation sites, the respective kinases involved and possible consequences. NES, nuclear export signal; CRM1, chromosomal region maintenance 1 (also called exportin 1); Jab1, c-Jun co-activator; NLS, nuclear localization signal; hKIS, human kinase-interacting stathmin; MIRK, minibrain-related kinase; AMPK, AMP-activated protein kinase; RSK, p90 ribosomal protein S6 kinases; K4 or K6/D, CDK4 or CDK6/cyclin D.

threonine-157, threonine-187, and threonine-198 (Fig. 14.1). Phosphorylation at serine-10 accounts for 75% of the phosphorylation observed on this protein [34]. At least four kinases have been shown to phosphorylate serine-10, including MAPK [35], human kinase interacting stathmin (hKIS) [36], AKT [37] and minibrain-related kinase (MIRK) [38]. The physiological significance of phosphorylation at serine-10 is contextual and controversial. Nevertheless, three functions have been proposed. Serine-10 phosphorylation activates the ubiquitylation– proteasome pathway in the cytosol during early G1 phase [39], promotes p27 nuclear export [35, 37, 40], and stabilizes p27 protein [38, 41]. Our own data, obtained after trastuzumab-induction of p27 protein, support the role of serine-10 phosphorylation in stabilizing p27 [13]. Tyrosine phosphorylation of p27 protein at sites 74, 88, and 89 can be carried out by nonreceptor tyrosine kinases such as Src, Lyn, Abl, and the oncoprotein Bcr-Abl (4, 43-45; Fig. 14.1). Tyrosine phosphorylation of p27 is reported to modulate its inhibitory activity by converting p27 from a bound inhibitor to a bound noninhibitor of p27-CDK4/6cyclin D complexes and by transforming p27 from an inhibitor to a substrate of CDK2-cyclin E complexes [4, 42–44]. Tyrosine phosphorylation of p27 can destabilize p27 protein by promoting subsequent threonine-187 phosphorylation [4, 42, 43]. Phosphorylation of threonine 157 can cause p27

to exit the nucleus, which relieves CDK2 from p27induced inhibition and also makes p27 vulnerable to proteasome degradation in the cytosol [45–47]. Phosphorylation of threonine-187 is the most intensively studied event for p27 protein, which is well known to be catalyzed by CDK2 [31–33, 48]. This phosphorylation prepares p27 protein for binding to an ubiquitin E3 ligase, S-phase kinase-associated protein 2 (SKP2), which leads to 26S proteasome degradation [31–33, 49]. Phosphorylation of threonine-198, like phosphorylation of serine-10, can be achieved by multiple kinases. AKT [50], AMPactivated protein kinase (AMPK) [9] and p90 ribosomal protein S6 kinases (RSK) [51] have been shown to phosphorylate this site (Fig. 14.1). Threonine-198 phosphorylation is shown to promote p27 binding to 14-3-3, retaining p27 in the cytosol [50, 51]. Threonine-198 phosphorylation can stabilize p27 protein [9, 52] and induce autophagy [9]. Furthermore, this phosphorylation is required to form p27/cyclin D/CDK4 complexes [52]. Figure 14.1 summarizes important posttranslational regulation of the p27 protein based on current research data.

14.2.4 p27 Proteolysis Three mechanisms are involved in the proteolysis of p27. The best described is mediated by SKP2. This mechanism depends on phosphorylation of

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threonine-187 and occurs in the late G1 phase through G2 phases in the nucleus [31, 32, 49]. The SKP2 ubiquitin ligase complex consists of SKP1, CUL1, CKS1B, RBX1, or a tripartite motifcontaining protein 21 (TRIM21) and SKP2 itself [31, 32, 49]. Changes in any of these components can affect p27 degradation and expression. A second proteolytic pathway occurs in cytosol in early G1 phase of the cell cycle and is totally independent of phosphorylation of threonine-187 [32]. Proteolysis is mediated by a Kip1 ubiquitylation– promoting complex (KPC). While it is not certain that this mechanism depends on phosphorylation of p27, it may be facilitated by phosphorylation of serine-10 and threonine-157 [32]. Pirh2 (p53-induced ring-H2-type ubiquitin ligase)-mediated p27 proteolysis is the third mechanism. It occurs in late G1 phase and continues through G2 phase in both the nucleus and cytoplasm [53]. This mechanism is independent of phosphorylation on threonine-187, levels of SKP2, and p53 function [53].


tion, and protein stability [56]. In addition to Jab1/CSN5, overexpression of CSN3, -6, -7, and -8 have also been reported to induce nuclear export and downregulation of p27 protein [55]. AKT phosphorylates p27 within its NLS on Threonine-157, impairs the p27-importin binding, promoting p27 binding to 14-3-3 and preventing its nuclear localization [45–47, 50, 57, 58]. By similar mechanisms, phosphorylation of p27 on serine-10 and threonine-198 promotes the binding of the protein to members of the 14-3-3 protein family that retain the protein in the cytoplasm [35, 37, 40, 50, 51, 59]. A novel mechanism that potentially regulates p27 translocation has recently been reported. Cyclin D2, but not cyclin D1 or D3, is able to promote p27 nuclear export and degradation in early G1 phase of the cell cycle in response to growth stimuli [60]. This mechanism depends upon its phosphorylation on threonine-280 by glycogen synthase kinase-3 beta (GSK3B) [33, 60]. Thus, the PI3K/PKB pathway can regulate p27 localization by several mechanisms [45–47].

14.2.5 p27 Translocation p27 activity is critically regulated by subcellular localization [3, 4]. In response to stimuli from mitogens or growth factors, p27 is rapidly translocated from the nucleus to cytosol in early G1 phase of the cell cycle, releasing inhibition of CDK2 [3, 33]. To act as a cell cycle inhibitor, p27 must be located in the nucleus, whereas its cytoplasmic sequestration allows cell cycle progression and other cell cycle-independent functions [5–7] to occur. Localization of p27 is not only influenced by its unique protein structures such as nuclear export signal (NES), nuclear localization signal (NLS), and c-Jun co-activator (Jab1) binding site (Fig. 14.1), but p27 localization is also tightly controlled by phosphorylation on p27 serine-10, threonine-157, and threonine-198 [4–7, 33]. Jab1, also known as CSN5, is involved in the nuclear export of p27 protein and induces its degradation in the cytoplasm [54]. Jab1/CSN5 functions as an adaptor between p27 and the nuclear exporting protein CRM1 to induce nuclear export and subsequent degradation [55]. Endogenous Jab1/CSN5 is incorporated into the COP9 signalosome, a multiprotein complex involved in modulating signal transduction, gene transcrip-

14.2.6 p27 Sequestration p27 positively regulates CDK by assembly and import of D-type cyclin-CDK4/6 [3]. Cyclin D is one of the early responding genes whose expression increases after stimulation with growth factors [3]. CDK inhibitors such as p27 and p21Cip1 would then shift p27 from CDK2-cyclin E complexes to cyclin D-CDK4 or CDK6 complexes and thus activate CDK2-cyclin E [3, 61]. Such p27 sequestration into cyclinD-CDK4/6 complexes is important for regulating p27 function during cell cycle progression. Both the MEK/ERK pathway and expression of the proto-oncogene c-Myc can induce cyclin D and cyclin E transcription and/or stabilize the proteins, thus stimulating the assembly of the cyclin D-CDK4/6 complexes, and the sequestration of p27 in these complexes [61–64].

14.3 Nuclear p27 as a CDK Inhibitor and Tumor Suppressor p27 suppresses tumor growth through its ability to inhibit cyclin-dependent kinases and to block cell proliferation. Knock-out of p27 in murine models


supports the assumption that p27 functions as a CDK inhibitor and a tumor suppressor. Homozygous p27 null mice are much bigger than wild-type mice and develop pituitary tumors and organ hyperplasia more frequently [65–67]. Heterozygous mice are hypersensitive to chemical carcinogens and gamma-irradiation [68]. The impact of “haploinsufficiency” points to the possible role of p27 as a rheostat for tumor suppression [69]. Other mouse models demonstrate that loss of p27 can cooperate with mutations in several oncogenes and tumor suppressor genes to facilitate tumor growth. Concomitant inactivation of one Phosphatase and Tensin homolog deleted on chromosome 10 (PTEN) allele and one or both p27 alleles accelerate spontaneous neoplastic transformation and increase the incidence of different tumors [70]. Concomitant inactivation of one retinoblastoma (Rb) tumor suppressor gene and both p27 alleles also accelerates tumorigenesis in both the pituitary and thyroid [71]. Combined deficiency of the p14/16 and p27 genes is reported to significantly increase in the number of metastatic tumors when compared to either gene deletion alone [72]. Deficiency of p27 can work collaboratively with deficiencies of a number of tumor suppressors (e.g. p53, APC, inhibin) to promote tumorigenesis in a broad range of tissues [73–75]. In transgenic lymphoma models, p27 deficiency has been shown to cooperate with Myc oncogene expression to promote lymphomagenesis [76].

14.4 Cytoplasmic p27 Possesses Multiple Cell Cycle-Independent Functions 14.4.1 Regulation of the Actin Cytoskeleton and Cell Migration Discovery of the cell cycle-independent function of p27 was prompted by the observation that p27-null fibroblasts exhibit a dramatic decrease in motility when compared with wild-type cells [77]. Indeed, p27 was found to co-localize with F-actin [77]. p27 was further found to bind to and inhibit RhoA activity by interfering with the interaction between RhoA and its activators, the guanine-nucleotide exchange factors [78]. p27-null fibroblasts exhibit

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decreased cell motility [78]. Cytoplasmic p27 expression in human breast cancer cells can increase cell motility and tumorigenicity [79]. This was the first oncogenic function attributed to p27, independent of CDK function [7]. Conversely, p27 can inhibit cell migration in other cell types. For example, p27 inhibits migration by binding to stathmin, a microtubule-destabilizing protein, in fibrosarcoma cells [80].

14.4.2 Regulation of Stem and Progenitor Cells Observations with a knock-in mouse model, where p27 without inhibitory activity for CDK (p27CK-) was re-expressed in mice, showed that p27CKexpression in cytoplasm could expand the numbers of stem and progenitor cells from several lineages in vivo, especially in the lung, retina, blood, and glial systems [81]. Increased numbers of stem and progenitor cells were associated with increased hyperplastic lesions and tumors in the lung, retina, pituitary, ovary, adrenals, spleen, and lymphomas in the p27CK- mice [81]. This also suggests that p27 may exert oncogenic activity, depending upon the context in which it is expressed.

14.4.3 Regulation of Apoptosis p27 can protect cells from apoptosis by constraining CDK2 activity [82]. The p27 protein can itself be cleaved by caspases to yield smaller fragments, which protect leukemic cells from apoptosis, whereas noncleavable forms of p27 do not [83]. The anti-apoptotic function of cytosolic p27 is mediated at least in part through the activation of AKT kinase [79]. While the ability of intact p27 to prevent apoptosis in cancer cells is still not fully documented and this property of the molecule could contribute to drug resistance. Upregulation of p27 could affect the impact of some chemotherapeutic drugs in certain cell types, both by limiting the number of cycling cells and by reducing their sensitivity to apoptosis [84].

14.4.4 Regulation of Malignancy and Tumor Metastasis In human cancer cells, tumors with high levels of exclusively nuclear p27 were almost well differentiated

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or of low grade, whereas cancers with cytoplasmic p27 localization were more poorly differentiated [45–47, 85–87]. Targeted cytoplasmic expression of p27 induces cancer cell motility, tumorigenicity, and metastases [79, 88]. Thus, data from clinical studies, mouse models, and cell culture seem to support an oncogenic function for p27, especially cytoplasmic p27.

14.5 p27 as a Prognostic Factor in Human Cancers 14.5.1 p27 as an Independent Prognostic Indicator in Breast Cancer More than 10 years ago, decreased levels of p27 levels were first observed in human breast cancers and found to correlate with poor patient outcome [89–92]. Over the last decade, p27 has been extensively studied in breast cancer. Details of these studies can be found in several excellent review articles [4, 86, 87, 93]. Although not all the studies agree, most support the original conclusion that low levels of p27 are associated with poor overall prognosis [4]. Reduced p27 in patients with breast cancer has correlated with increased tumor proliferation index, high tumor grade, decreased expression of estrogen receptor (ER) or progesterone receptor (PR), and increased risk of relapse [4, 86, 87].

14.5.2 p27 as an Independent Prognostic Indicator in Human Cancers Reduction or loss of p27 protein occurs in a wide variety of human tumors, including carcinomas of the colon, prostate, lung, gastrointestinal tract, liver, ovary, and cervix as well as brain tumors, endocrine tumors, lymphomas, and soft tissue sarcomas [4, 86, 87, 93]. Multivariate analysis indicates that reduced levels of p27 constitute an independent prognostic factor in multiple tumor types [4, 86, 87, 93]. There are some cancers with “normal” or even increased p27 expression. However, increased p27 protein is located largely in the cytoplasm rather than in the nucleus in most cases [93]. As discussed above, cytoplasmic p27 may mediate motility, resistance to apoptosis, and metastatic potential [78–88].


14.5.3 Reduction of p27 in Early Oncogenesis The strong correlation between p27 levels and patient survival in a variety of human malignancies underscores the potential contribution of p27 loss or mislocalization to human carcinogenesis. Reduction in p27 may happen early in oncogenesis. Normal mammary epithelial cells express high levels of nuclear p27, whereas p27 levels gradually decrease during the progression from hyperplasia to ductal carcinoma in situ (DCIS) and from DCIS to invasive breast cancer [90, 94–96]. In other cancer types, similar results were obtained. Reduced expression of p27 is a rather early event in gastric oncogenesis before dysplastic changes occur. [97]. A loss or absence of p27 expression is an early pathogenic event in mast cell and histiocyte tumorigenesis [98]. Loss of both p27 and E-cadherin expression occurred early during the preneoplastic stages of head and neck carcinogenesis [99].

14.6 Oncogenic Mechanisms of p27 Inactivation in Human Cancer Understanding the mechanisms by which p27 is inactivated in carcinogenesis is important to understand the role of p27 in cancer and also might suggest new therapeutic strategies. Two major mechanisms have been identified for p27 inactivation in cancer: downregulation of p27 expression by ubiquitylation and proteasomal degradation; and exclusion from the nuclear compartment mediated by phosphorylation of p27.

14.6.1 Accelerated p27 Degradation During normal cell cycle progression, levels of p27 protein decline following degradation by ubiquitin–proteasome dependent proteolysis [31, 32, 49]. Similar regulation occurs in cancer cells. Protein lysates obtained from patients with colon cancers with low p27 levels exhibited proteolytic activity toward recombinant p27 in vitro [91]. Several other mechanisms may cause accelerated p27 degradation in human cancers as summarized in Table 14.2.

14. p27Kip1 as a Biomarker and Target for Treatment of Cancer Table 14.2. Mechanisms of accelerated p27 degradation in cancer. 1. Activation of SKP2 2. Activation of PI3K signaling 3. Activation of Src family kinases 4. Activation of Ras-MAPK 5. Activation of TRIM21

14.6.1.1 Activation of SKP2 As discussed above, the best described proteolytic mechanism for p27 protein is SKP2-dependent proteolysis. Once p27 protein is phosphorylated on threonine-187, the SKP2 ubiquitin ligase complex is able to conjugate with the protein and activate S26 proteasomes [31, 32, 49]. Overexpression of SKP2 has been observed in many human cancers including lymphoma, colon, breast, head and neck, prostate, lung, gastric, and oral cancers [100–103]. High SKP2-mediated protein degradation is significantly correlated with reduced p27 expression [100, 101].

14.6.1.2 Activation of PI3K Signaling An increase in PI3K activity can result from (1) activating mutations in PI3K components, (2) inactivating mutations in PTEN, (3) amplification and overexpression of PI3K or AKT, (4) cross talk from activation of the Ras/MAP pathway, or (5) autocrine or paracrine growth factor stimulation of growth factor/cytokine receptors that activate PI3Ks [104]. Mutations of the epidermal growth factor receptor (EGFR) and Ras oncogene, and amplification/ overexpression of human epidermal growth factor receptor 2 (HER2) have been shown to increase the levels of the PI3K lipid products [104, 105]. Loss of the tumor suppressor PTEN, either by deletion of the gene or by deactivating mutations, causes constitutive PI3K signaling through the build up of PI3K lipid products [104]. Overexpression of HER2 is correlated tightly with decreased p27 levels in clinical samples [11, 106, 107]. Our own data clearly indicate that overexpression of HER2 dramatically lowers p27 protein in cultured cancer cells [108]. The underlying mechanism by which increased HER2 downregulates p27 is considered at least in part due to activation of PI3K signaling [108–111]. The tumor suppressor function of PTEN is carried out through p27 induction [112–114]. Loss of PTEN produces activation of PI3K and AKT activity and reduces p27 levels [114]. As discussed earlier, activation of AKT

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could promote threonine phosphorylation and proteasome degradation of the p27 protein [4, 45–47]. PTEN may also inhibit SKP2 activity and partially upregulate p27 protein [115].

14.6.1.3 Activation of Src Family Kinases Signaling from a number of receptor tyrosine kinases (c-Met, EGFR, HER2, PDGFR), integrins, and G-protein coupled receptors can activate Src [43, 116, 117]. As a vital nonreceptor tyrosine kinase in the cytosol, Src mediates multiple signaling activities that control cell proliferation, migration, adhesion, survival, and angiogenesis [116, 117]. Src activation is frequently detected in human cancers [43, 116, 117]. Activation of Src promotes tyrosine phosphorylation of p27 protein and activates cyclin E/D-CDK2/4/6 complexes and subsequent proteasome degradation of p27 protein [4, 42–44]. Activation of other Src family kinases (such as Lyn, Lck and Fyn) and Abl kinase that also phosphorylate p27 can promote p27 proteolysis in hematopoietic malignancies and lymphoma [118, 119].

14.6.1.4 Activation of Ras-MAPK Both in vivo and in vitro data suggest that Ras activity regulates p27 activity [35, 120, 121]. Ras activity modulates p27 levels through the RhoA and MAPK pathways [120–123]. MAPK may directly phosphorylate p27 and facilitate the proteolysis of p27 degradation [120, 121]. HER2 overexpression is also reported to activate MAPK and to degrade p27 protein [122]. Rho A has been shown to activate CDK2 activity [123] or ROCK kinase [124] and to reduce p27 protein. Interestingly, p27 is able to bind to and inhibit RhoA activity as well [78]. The feedback loop between RhoA and p27 may contribute to Ras-dependent regulation of p27 protein. Additionally, RhoA is able to decrease the translational efficiency of p27 mRNA [125].

14.6.1.5 Activation of TRIM21 A RING finger B-box protein TRIM21 (also known as Ro52) is a component of the SKP2 E3 ubiquitin ligase [126]. Activated TRIM21 activity promotes the ubiquitination of Threonine187-phosphorylated p27 in a RING-dependent manner in vitro. Dysregulation of TRIM21 has not yet been reported in human cancers.

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14.6.2 Mislocalization of p27 Mislocalization of p27 protein, resulted from a variety of mechanisms (Table 14.3), abrogates its inhibitory function of cell growth. Oncogenic signaling from activation of the PI3K-AKT, Ras-MAPK, and Src pathways can result in phosphorylation of p27 protein on serine 10, threonine 157, and threonine 198 and lead to cytoplasmic localization [4–7, 33]. Cytoplasmic localization of p27 not only inactivates the inhibitory function of p27, but also allows p27 to perform a number of cell cycle-independent functions as described previously [78–88]. Jab1 is involved in the nuclear export of p27 protein [54]. Jab1 overexpression has been observed in a number of human cancers and inversely correlates with p27 levels [127–130]. Overexpression of Jab1 and other family members such as CSN-3, -6, -7, and -8 may also induce nuclear export of p27 protein [55]. Overexpression of 14-3-3 protein can retain p27 protein in the cytoplasm [35, 37, 40, 50, 51, 59] and lead to poor prognosis in cancer [177]. Cyclin D2 is reported to regulate the p27 translocation to cytosol [33, 60]. Overexpression of cyclin D2 has been demonstrated to promote tumor formation and poor prognosis in animal models and cancer patients [178–180]. The serine/threonine kinases Pim1/2/3 have been recently shown to bind to and directly phosphorylate p27 at threonine-157 and threonine-198 residues, which also result in nuclear export of p27 [131]. Pim kinases are overexpressed in leukemias, lymphomas, prostate cancer and head/ neck carcinomas [132–135].

14.6.3 Sequestration of p27 Sequestration by cyclinD-CDK4/6 is another mechanism that leads to functional inactivation of p27. Table 14.3. Mechanisms of p27 mislocalization in cancer. 1. Threonine 157 phosphorylation by activated PI3K signaling 2. Tyrosine 77/88/89 phosphorylation by activated Src and Abl kinase signaling 3. Serine 10 and threonine 198 phosphorylation by activated Ras-MAPK signaling 4. Overexpression of Jab1/CSN5 and related CSN family members 5. Overexpression of 14-3-3 protein 6. Overexpression of cyclin D2 7. Overexpression of Pim kinases


Both Myc-dependent and independent pathways cause the shift p27 from CDK2-cyclin E complexes to cyclin D-CDK4 or CDK6 complexes and thus activate CDK2-cyclin E [61]. Various oncogenic and mitogenic signals transmitted through the Ras-MAPK and PI3K pathways induce cyclin D transcription and/or stabilize the cyclin D protein, increase the assembly of the cyclin D-CDK4/6 complexes and thus sequester p27 in inactive cytoplasmic complexes [3, 61]. Myc can induce cyclin D and cyclin E transcriptions, stimulate the assembly of the cyclin D-CDK4/6 complexes, and sequester p27 [61–64]. Due to chromosomal translocation, gene amplification and stabilization of mRNA and protein, cyclin D overexpression is already implicated in several types of human tumors, including parathyroid adenomas and B-cell neoplasms, as well as gastric, esophageal, and breast cancers [136, 137]. Myc expression is deregulated in a wide range of human cancers and is often associated with aggressive, poorly differentiated tumors [138]. Sequestration of p27 is only one mechanism by which Myc functions, since Myc as a transcription factor regulates a variety of cellular processes, including not only cell growth, proliferation, and cell-cycle progression, but also transcription, differentiation, apoptosis, and cell motility [138].

14.6.4 Decreased Translation of p27 IRES-mediated p27 translation is an important pathway for p27 synthesis [25–27]. Patients with X-linked dyskeratosis congenita (DKC) usually harbor DKC1 gene mutation [139]. The DKC1 gene encodes a pseudouridine synthase that modifies ribosomal RNA [139]. DKC1 mutation produces IRES defects, impairs translation of p27 as well as the antiapoptotic factors Bcl-xL and the X-linked Inhibitor of Apoptosis Protein (XIAP) and increases the susceptibility of mutation carriers to various cancers [139]. Overexpression (especially cytoplasmic expression) of ELAVL1 that negatively regulates p27 translation [26, 27] has been detected in ovarian [140], colon [141], breast [142], prostate [143], laryngeal [144], and lung [145]. Most recently, miR-221 and miR-222 have been shown to bind to p27 3¢ UTR and to inhibit its translation [8, 29, 30]. Overexpression of miR-221


and miR-222 has been detected in thyroid carcinomas with decreased p27 levels [146]. The expression of miR-221 and miR- 222 is also significantly elevated in HER2-positive primary human breast cancer tissues [147], associated with downregulation of p27 levels [108–111]. A significant inverse correlation between miR-221 and p27 is reported in over 70% human hepatocellular carcinomas [148].

14.6.5 Impaired p27 Synthesis in Cancers Overexpression of p27 transcriptional repressors such as c-Myc, HES1, Id3, and miRNA 221/222 can impair p27 synthesis [14–18]. c-Myc overexpression is found in a significant fraction of human cancers and may inhibit p27 expression by several mechanisms including repression of transcription [138]. The Notch signaling marker HES1 is overexpressed in glioblastomas [149] and osteosarcomas [150]. The transcriptional inhibitor of differentiation Id3 is overexpressed in lung cancers associated with aggressive metastatic potential and a poor prognosis [151, 152]. Overexpression of miR-221 and miR-222 has been detected in multiple human cancers [146–148], potentially contributing to transcriptional suppression of the p27 gene in these neoplasms [8]. Similarly, downregulation or functional inhibition of p27 transcriptional activators such as FoxO, menin, E2F1, and SP1 are thought to impair p27 synthesis as well [19–22]. Under control of AKT and SKP2, the FoxO factors function as tumor suppressors in a variety of cancers by upregulating p27 and proapoptotic factors such as Fas ligand, tumor necrosis factor-related apoptosisinducing ligand, Bim, bNIP3, and Bcl-xL [153]. Phosphorylated–FoxO1 staining was significantly lower in metastatic colon cancer than in primary tumors in human specimens [154]. Cytoplasmic FoxO3 (functional inactivation) correlated with the expression of IKKb or phospho-Akt and was associated with poor survival in many tumors [155]. Menin, encoded by multiple endocrine neoplasia type 1 (MEN1) gene, is mutated in heritable and sporadic endocrine tumors [156]. Inactivation of menin by mutation leads to leukemogenesis [156] and accelerated cell proliferation by downregulating

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p27 and upregulating cyclin D1 and CDK4 [157]. The transcription factor E2F1 is a critical regulator of the cell cycle due to its ability to promote S phase entry [158]. On the other hand, E2F1 not only induces proapoptotic proteins such as caspase 3, 7, 9, and Apaf1 [158], but also directly induces expression of CDK inhibitors such as p19ARF [158] and p27 [21, 159]. E2F1 primarily binds to the promoter region of p27 and enhances its transcription [21]. Based on similar mechanisms, the transcription factor SP1 in combination with other co-factors binds to the promoter region of p27 and enhances its transcription [22, 160]. Dysregulation of both E2F1 and SP1 in human cancer may contribute to the dysregulation of p27 protein. Finally, oncogenic kinases Pim1/2/3 have been recently shown to inactivate forkhead transcription factors (FoxO1a and FoxO3a) and decrease the synthesis of p27 mRNA [131].

14.7 p27 as a Biomarker for Treatment Response in Human Cancer 14.7.1 Efficacy of Chemotherapy Among eight reports that assessed the value of p27 levels for predicting efficacy of chemotherapy in human cancer, four indicated that low p27 levels prior to treatment predicted benefits from chemotherapy in colorectal and ovarian cancers [4, 11, 161, 162]. Reduced p27 levels were correlated with a high proliferative rate in cancer cells, and cycling cancer cells may be more sensitive to chemotherapy [4, 11, 162]. Four additional reports showed that low p27 levels predicted poor benefit from chemotherapy in breast, lung, and ovarian cancers [163–166]. All these studies were not prospectively designed, however, to test the predictive value of p27. To date, p27 has repeatedly observed to be an independent prognostic factor [4, 86, 87, 93], but observations regarding p27 as a predictor of treatment response have been less numerous. Prospective, carefully designed and randomized trials to test the predictive value of p27 for chemotherapy in human cancer are needed.

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14.7.2 Efficacy of Hormonal Therapy in Breast Cancer Since p27 is an important mediator of the G1 arrest induced by anti-estrogens [10], p27 levels could predict the potential activity of these drugs in particular patients. Indeed, two prospective randomized clinical trials have shown that high levels of p27 predicted a favorable response to adjuvant tamoxifen therapy in breast cancer [167, 168]. A third retrospective clinical study also confirmed that high levels of p27 predicted a favorable response to adjuvant hormonal therapy [4].

14.7.3 Efficacy of Radiation Therapy Among three reports that evaluated the predictive value of p27 for response to radiotherapy in cancers, two studies showed that high p27 levels in pretreatment tumor were associated with longer survival following radiotherapy in cervical and laryngeal cancers [12, 169]. Another report, in contrast, showed that reduced p27 correlated with improved outcome after radiotherapy in aggressive esophageal cancers [170]. Further studies are warranted to assess the true value of p27 in predicting the outcomes of radiotherapy.

14.7.4 Efficacy of Trastuzumab We and others have shown that p27 is required for the anti-HER2 antibody trastuzumab to induce G1 arrest of the cell cycle and to suppress growth of human breast cancers that overexpress HER2 [13, 108–111]. Downregulation of p27 with siRNA or anti-sense oligonucleotides against human p27 abrogates trastuzumab-mediated cell cycle arrest [13]. Therefore, p27 levels and subcellular localization may dictate the efficacy of this targeted antibody therapy. In vitro, the earliest p27 upregulation in trastuzumab-sensitive breast cancer cell lines can be detected at 14 h after antibody treatment [13]. Using this criteria, we were able to predict correctly the response to trastuzumab treatment in 11 breast cancer cell lines that express high levels of HER2 (Le XF and Bast Jr RC, unpublished data). In support of the value of p27 to predict response to trastuzumab treatment, trastuzumab resistance has been reported to associate with decreased p27 levels [171].


14.8 p27 as a Target for Therapy 14.8.1 Rationale for Therapy Restoration of functional p27 to the nuclei of cancer cells could provide a useful therapeutic strategy based on the ability of p27 to arrest the cell cycle to induce differentiation, and to induce autophagy or type 2 programmed cell death. Cell cycle dependent functions of p27 have been studied in depth, but less is known regarding its effect on differentiation. Loss of p27 leads to hyperplasia and tumors in mice [87, 172], whereas levels of p27 increase in terminally differentiated cells [173, 174]. Human cancers with high levels of nuclear p27 are almost all well differentiated or of low grade, whereas those with low nuclear p27 and localization of p27 in the cytoplasm are poorly differentiated [45–47, 85–87]. Although the precise molecular mechanism(s) of p27-induced differentiation are not clear, p27 has been shown to promote neuronal differentiation through mechanisms that differ from those required for neuronal migration [175]. Two recent studies document the role of p27 in the induction of autophagy. AMP-activated protein kinase (AMPK)-mediated pathway phosphorylates p27 at threonine-198, stabilizes p27, and permits cells to undergo autophagy [9]. Specific targeting of SKP2 protein preventing degradation of p27 protein leads to effective and selective killing of leukemic cells through a caspase-independent programmed cell death or autophagy [176]. Depending upon the context, upregulating p27 might not always exert an anti-cancer effect. Decreasing the fraction of cycling cancer cells could decrease sensitivity to some cycle-specific drugs. The ability of p27 to prevent apoptosis and to induce drug resistance is also a concern [81–83]. Increased cytoplasmic p27 expression is reported to increase cancer cell motility and tumor metastasis [78, 79, 88]. The ability of p27 to increase stem and progenitor cell numbers in several organs/systems might also mediate paradoxical effects [81]. Proteosome inhibitors such as bortezomib (Velcade®) can upregulate p27 [181], but also affect levels of many other proteins. Consequently, the precise role of p27 in the response of cancer cells to proteosome inhibitors is difficult to assess. As SKP2 mediates unbiquitination of p27, cancers that overexpress SKP2 could be targeted by small molecular


weight SKP2 inhibitors or siRNA [176]. For cancers with cytoplasmic localization of p27, inhibitors of PI3K/AKT, Ras/MAP, and Src could restore nuclear p27. Tumors with overexpression of miR221/222 should be targeted with antagomiR gene therapy.

14.8.2 Linking Molecular Therapeutics and Diagnostics Although there are a number of potential problems associated with therapeutic strategies based on p27-upregulation, the value of p27 as an independent prognostic factor and a predictive biomarker of response to targeted therapy is better documented. As detailed above, upregulation of p27 has been observed after treatment with trastuzumab in those HER2-overexpressing breast cancer cell lines that exhibit growth inhibition [13, 108–111]. Cell lines that fail to respond to trastuzumab do not upregulate p27. In this setting p27 may not only be a biomarker for response, but also an effector of inhibiting cell proliferation. Thus, therapies that upregulate p27 might be considered in breast cancer patients who have developed trastuzumab resistance. A similar example is observed with inhibitors of Src such as dasatinib where cancer cell lines that respond and upregulate p27. Our unpublished data strongly suggest that p27 upregulation is critical mediator of Src inhibition therapy.

14.9 Conclusion Significant progress has been achieved in understanding the role of p27 in normal and malignant cells. A number of fundamental questions are still unanswered. How is p27 phosphorylation coordinated at multiple sites and by multiple serine, threonine, and tyrosine kinases? What are the physiological roles of protein phosphorylation at threonine 198 and serine 10? Which phosphorylation events are observed most frequently in human cancers? How can convenient and rapid assays for these events be developed for clinical use? Although the value of p27 as an independent biomarker has been suggested in a number of studies, prospective validation has not been achieved and the clinical application of p27 in making decisions related to patient care is not clear. As molecular therapeutics

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develop, it may well become possible to manipulate p27 levels and localization. SKP2 inhibition appears to be one of the most accessible targets. Monitoring of p27 before and during therapy with trastuzumab, PI3K/AKT inhibitors, Ras/MAP inhibitors, and Src inhibitors needs to occur not only in the laboratory, but also in the clinic. Acknowledgments: This work was supported in part by a grant CA39930 from the National Cancer Institute, a grant from the Commonwealth Foundation (Goodwin Family), and the generous support of the Zarrow Foundation.

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Chapter 15

Targeting Cyclin-Dependent Kinases with Small Molecule Inhibitors Paolo Pevarello, James R. Bischoff, and Ciro Mercurio

Abstract In spite of a long running effort take out by most of the principal pharmaceutical companies, CDK inhibitors are not yet available for use by clinical oncologists. This is due to several reasons; among them is a changing perspective on how many and which CDK subtypes have to be inhibited for optimal antitumor activity coupled with minimal toxicity, and on the opportunity of targeting additional relevant kinases in oncology. CDK inhibitors can be classified into three classes based on the pattern of their kinase inhibition properties: the first group includes compounds like AT-7519 or R-547 with broad CDK profile; the second group features, for example, selective CDK4/6 inhibitors like PD-0332991 and P276-00; in the third group, we find compounds with additional kinase activity besides CDK inhibition, e.g., ZK304709 and PHA848125. In this chapter, we review the current status of the CDK inhibitor pipeline across the industry, taking into consideration those compounds that have progressed into clinical testing. Keywords Cyclin-dependent kinase • Small molecule inhibitor • Selectivity • Multi-target • Clinical trials

15.1 Introduction In recent years, several targeted molecular therapeutics have received market approval from the FDA (Food & Drug Administration) and/or the EMEA (European Medicinal Agency) as anticancer agents. Either small molecules (SM) such as Gleevec®,

Velcade®, Iressa®, Tarceva®, Nexavar®, Zolinza®, Sutent®, Sprycel®, Lapatinib®, Nilotinib®, and Torisel® or biologics like Alemtuzumab®, Cetuximab®, Bevacizumab®, Trastuzumab®, and Panitumumab® have generated considerable expectations that the recent advances in molecular oncology and the ever-increasing understanding of relevant pathways and biological targets in tumorigenesis and cancer progression can effectively be translated into new treatment options. Most of the aforementioned drugs target growth factors or growth factor receptors or tyrosine kinases in the signal transduction pathways that govern tumor cell proliferation and/or angiogenesis. Many of the programs that led to these marketed compounds began in the mid 1990s or even later. Concurrently, programs targeting cell cycle regulatory proteins, and in particular the CDKs, were underway, sometimes within the same companies. It is, therefore, surprising that as of yet no new drug targeting cell cycle progression, and specifically CDKs, has made it to the market in the oncology arena. The delayed development of CDK inhibitors is probably due to a convergence of different factors such as: (a) the reappraisal of the role CDKs plays during the cell cycle in both normal and genetically compromised tumor cells; (b) off-target effects of the compounds that entered clinical trials; (c) a less-than-optimal preclinical optimization of first generation compounds with regard to their pharmacokinetic or pharmacodynamic profile [1]. In this chapter, we would like to briefly comment on the changing view about the use of a CDK inhibitor as an anti-tumor agent and review the


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current status of CDK inhibitors (to the best of the currently accessible knowledge) that are currently investigated in clinical trials. Compounds whose key preclinical and clinical data are publicly available or for which confirmed clinical trials have been initiated are briefly reviewed and data driven considerations relating to specific features or opportunities that these compounds present are discussed.

15.2 CDK Inhibitors in Clinical Trials One explanation currently gaining acceptance for the generally disappointing results of a number of CDK inhibitors that underwent clinical trials in the past was that a less than optimal schedule of administration was used, often mandated by safety and tolerability concerns which in turn are also linked to potential off-targets effects [1]. The administration schedules commonly employed, e.g., acute intravenous (iv.) olus with long intervals between administrations, are probably not the best way of controlling proliferation with a CDK inhibitor. The estimated doubling time of carcinoma cells is in the scale of the days [2], which implies that the tumor should be exposed to a CDK inhibitor for days in order to target all the cells that are cycling in the tumor. Another emerging concept derives from what genetics is telling us about the possible therapeutic uses of a CDK inhibitor [1, 3, 4, 5]. Murine genetic models suggest that specific inhibition of one CDK, e.g., CDK2 or CDK4, should have a limited toxic effect, but that increasing the number of kinases targeted will increase toxicity. A third important consideration is that inhibition of single CDK family members might only be efficacious in specific cell types in a limited genetic background [1]. This consideration calls for a proper choice of patients to be treated based on the genetic context of their tumors, which is a concept that nowadays is widely accepted in Clinical Oncology. The three points noted here are beginning to be addressed with the new generation of CDK inhibitors that are entering or will soon be entering into clinical development, and there is optimism that the next few years will witness the success of at least some of them. In general, CDK inhibitors that have entered clinical trials can be categorized into three main classes: Group 1 – those with a broad CDK inhibition profile (also called pan-CDK inhibitors);

P. Pevarello et al.

Group 2 – those exclusively or preferentially inhibiting either CDK4/CDK6 or CDK2 activity; and Group 3 – those inhibiting CDKs and additional kinase targets of interest in oncology.

15.2.1 Group 1 Inhibitors Targeting a Broad Range of CDKs Inhibitors in this group usually display sub-micromolar to low nanomolar potency against a range of CDK/ cyclin complexes, such as CDK1/cyclin B, CDK2/ cyclin A and CDK2/cyclin E, Cdk4/cyclin D, CDK6/cyclin D, CDK7/cyclin C, and CDK9/cyclin T. The rationale for their development is based on their potent anti-proliferative activity in a variety of tumor cell types and tissues and on the anticipation that their unavoidable toxicological impact on rapidly dividing normal tissues (e.g., gastrointestinal and bone marrow) will be offset by strong efficacy in a cancer setting. Compounds of this type were the most common outcome of past CDK programs and belong to many different chemical classes. First generation compounds such as Alvocidib® (flavopiridol) and Seliciclib® ((R)-roscovitine) are representative of this class. Examples of second generation pan-CDK inhibitors are SNS-032, AT-7519, AT-9311, PHA-793887, R-547, and SCH-727965 (Table 15.1).

15.2.1.1 Alvocidib® (flavopiridol) After a rather long clinical development history, Alvocidib®, jointly developed by NCI and SanofiAventis, may be the first CDK inhibitor to reach the market as early as 2010. The compound was granted Orphan Drug status by the EMEA in 2007 for the chronic lymphocytic leukemia (CLL) indication. Clinical studies in a number of other malignancies have not revealed significant efficacy at the doses and administration schedules examined. It seems, therefore, important (and the same conclusion may be drawn for other CDK inhibitors) to re-examine using both the therapeutic context (e.g., CLL) and the administration regimen (e.g., long infusion for an intravenous agent) that have greater potential.

15.2.1.2 Selicilib® ((R)-roscovitine) Selicilib® is another broad spectrum CDK inhibitor being developed by Cyclacel. It is orally active,

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15. Targeting Cyclin-Dependent Kinases with Small Molecule Inhibitors Table 15.1. Compound with a broad CDK activity profile (pan-CDK inhibitors).

Company

Code number

NCI

Alvocidib (flavopiridol)

Disclosed or bona fide structure or chemotype H N

Admin route Conditions

1,2,4,5,6,7,9,

III (2008)

i.v.

1,2,4,5,7,9

II (2007)

p.o.

1,2,4,7,9

I/II (2007)

i.v.

1,2,4,5

I/IIa (2007) i.v.

1,2,4,5

I (2007)

i.v.

1,2,4,7

I (2006)

i.v.

Advanced solid tumors

1,2,9

I (2008)

i.v.

Non-Hodgkin’s lymphoma, multiple myeloma

Chronic lymphocytic leukemia (CLL)

Cl

HO

H

HO

O

O

OH

Cyclacel

Clinical phase (last Reported CDK reported activity date)

Seliciclib (R-roscovitine) HN N

N HO

Sunesis

SNS-032 (BMS-387032)

N H

N

N

S

N H

S

O

Astex

AT-7519

O

O

H N

Cl

AT-9311

PHA-793887

ScheringPlough

SCH-727965

R-547 (Ro-4584820)

H N

N H

N H

Advanced breast cancer, melanoma, or nonsmall cell lung cancer (NSCLC); B-Cell malignancies Advanced or metastatic solid tumors; Refractory non-Hodgkin’s lymphoma

O N H

NH

H N

HN N

Nerviano medical sciences (NMS) HoffmannLaRoche

NH

N

Cl

Astex

O

N N

O N

N

na H N O S

N

N

O

N

O

F NH2 O

F

OH N

N N N HN

+

N

−

O

with good in vitro biochemical activity against CDK1, 2, 4, 5, 7, and 9, but with a weaker antiproliferative activity (as compared to other CDK inhibitors) against most tumor cell lines. In spite of this drawback and by virtue of its high bioavailability and safety/tolerability, it underwent several clinical trials, not limited to the treatment of cancer. Evidence of clinical efficacy, at least as a single agent, is still inconclusive, and the activity of the compound is being actively investigated in combination with various agents.

15.2.1.3 SNS-032 SNS-032 is a thiazole derivative, in-licensed by Sunesis from BristolMyers Squibb (former code BMS-387032). It is administered by the iv route and preferentially inhibits CDK2/cyclin A/E (IC50: 38 nM), CDK7/cyclin H (IC50: 62 nM), and CDK9 (IC50: 4 nM), which indicates that it acts on both proliferation and transcription processes. These kinases are selectivity inhibited in vitro when compared with CDK1/cyclin B (IC50: 480 nM), CDK5/p35

238

(IC50: 340 nM), CDK4/cyclin D1 (IC50: 925 nM), and CDK4/cyclin D2 (IC50: > 1000 nM), and its pattern of CDK inhibition is different from other Group 1 compounds such as Alvocidib (CDK2/cyclin A: IC50: 100 nM; CDK7/cyclin H: IC50, ca. 300 nM; CDK9: IC50: 6 nM; CDK1/cyclin B: IC50, 41 nM; CDK4/cyclin D1 and CDK6/cyclin D2: IC50: 65 and ca. 100 nM, respectively) [6]. The inhibitor is currently in Phase I clinical trials in patients with multiple myeloma and CLL.

P. Pevarello et al.

15.2.1.6 PHA-793887 PHA-793887 is a multi-targeting CDK inhibitor, which was identified at Pharmacia and now developed by Nerviano Medical Sciences (NMS). Although preclinical data on this compound have not yet been made available, it entered Phase I clinical trials as an 1-h i.v. infusion for 5 consecutive days in a 2-week cycle at the Cancer Therapy & Research Center of the University of Texas Health Science Center [7].

15.2.1.7 R-547 15.2.1.4 AT-7519 AT-7519, which preferentially inhibits CDK1/cyclin B, CDK2/cyclin A, CDK4/cyclin D, and CDK9/ cyclin T, is being developed by Astex Technology as a potential intravenous (i.v.) treatment for cancer. The compound is undergoing PhaseI/IIa studies in patients with solid tumors or hematological malignancies (e.g., non-Hodgkin’s Lymphoma). A typical administration schedule for this compound is a 1–3 h i.v. infusion on days 1, 4, 8, and 11. Preclinical data show that AT-7519 preferentially inhibits CDK2/ cyclin A (IC50: 44 nM), CDK4/cyclin D1 (IC50: 67 nM); CDK5/p35 (IC50: 18 nM), CDK9/cyclin T1 (IC50: 34.4 mg/m2). Maximum dose administered (MDA) was 259 mg/m2/day and RDPII was 185 mg/m2/day. As for PD-0332991, multiple myeloma is currently pursued, primarily since preclinical tests of the compound demonstrated activity against a range of multiple myeloma cells resistant to standard therapy and synergy with bortezomib (Velcade®). In NOD-SCID mice bearing GFP/MM xenografts, significant tumor growth inhibition was observed at 25 mg/kg, the compound being given 3 times a week for 3 weeks.

15.2.2.2 PD-0332991 PD-0332991, originated by Onyx and WarnerLambert and developed by Pfizer, is a preferential CDK4/Cyclin D1-CDK6/Cyclin D2 inhibitor currently in Phase I/II clinical trials for indications like

Table 15.2. Compounds with exclusive or preferential CDK4/CDK6 vs. CDK2 or CDK1 activity or viceversa. Company

Code number

Disclosed or bona fide structure or chemotype

Reported Stage (last CDK Activity reported date)

Admin. route Conditions

Nicholas piromal

P276-00

Rohitukine derivative

1,4,6

I/II (2007)

p.o.

Pfizer

PD-0332991

HN

4,6

I (2006)

p.o.

2

I (2008)

i.v.

O N

N

N N H

Nerviano medical sciences (NMS)

PHA-690509

S

N

N

N N

N

O

Advanced refractory neoplasms; multiple myeloma Neoplasms; non-Hodgkin’s lymphoma

240

P. Pevarello et al.

refractory multiple myeloma (in combination with bortezomib and dexamethasone) and mantle cell lymphoma. In a previous Phase I trial, 57 patients with Rb positive breast and colorectal tumors, liposarcoma, and melanoma were treated with two schedules, either 25–150 mg once-daily for 21 days (of a 28-day cycle) or 100–225 mg over 14 days of a 21-day cycle. An MTD of 125 mg was established. Six patients achieved stable disease using the first schedule. As seen with other CDK inhibitors, main adverse effects were neutropenia, anemia, fatigue, nausea, constipation, vomiting, and diarrhea though the DLT was due to myelosuppression [13]. This compound may have the potential in a combination therapy treatment schedule as hinted by several preclinical studies, which showed additive or synergistic effect with other investigational agents (like the MEK inhibitor PD-0325901) or marketed drugs (e.g., bortezomib (Velcade®)). In vitro, PD-0332991 is characterized by preferential CDK4/Cyclin D1-3 inhibition (IC50: 9–15 nM), with substantially less potency against other CDKs (e.g., CDK2/CyclinA and CDK1/Cyclin B, IC50s > 5 mM) and other general kinases [14]. Activity in inhibiting tumor cell growth was in the low micromolar range (e.g., IC50: 0.12 mM

for H1299 lung carcinoma and 0.13 mM for Colo205 colon carcinoma cells). In line with its expected mechanism of action, PD-0332991 was able to block Rb-positive tumor cell lines in G1 phase, preventing S-phase entry. The compound was also active in several animal tumor models, in some cases displaying clear tumor regression [15].

15.2.2.3 PHA-690509 PHA-690509, discovered at Pharmacia and currently pursued by NMS, shares with SNS-032 a 5-substituted-2-aminocarbonyl core and is reported to inhibit CDK2/CyclinA [16]. Multicenter Phase I dose escalation trials had been initiated in 2005, and its clinical development program was ongoing in April 2008 [17].

15.2.3 Group 3 Inhibitors Targeting CDKs and Additional Kinases of Oncologic Interests The agents listed in (Table 15.3) not only inhibit CDKs, but also additional kinases of interest in cancer chemotherapy. These additional targets may be

Table 15.3. CDK Inhibitors with dual or multiple mechanism of action.

Company Bayer schering pharma AG

Code number ZK304709

R1 SO2 R4

R5

PHA848125

RGB286638

N

NH

Relapsed/ refractory solid tumors

1,2,4,5,

Trk-A

p.o.

Relapsed/ refractory solid tumors

1,2,3,5,7,9 GSK-3b, Enters Mek-1, Lyn, Phase I Fyn, c-Src (2008)

i.v.

Multiple myeloma (MM)

1

i.v.

Myelodysplasia (MDS)

NH

N

N N

ON-01910. Na

O

N

OMe

MeO

S O

O NH +

COO- Na

I (2008)

O N

O

N H

ONO Pharmaceuticals

p.o.

N

O N H

VEGFR1I (2006) 3PDGFR-b Flt-3

NH

N

N

Admin. route Conditions

1,2,4,7,9

N R2

N

O

Stage (last reported date)

O

N H

GPC Biotech

Other kinase activity

R3

N H

Nerviano medical science (NMS)

Reported CDK activity

Disclosed or bona fide structure or chemotype

OMe O

Plk-1, PDGFR

I (2008)

15. Targeting Cyclin-Dependent Kinases with Small Molecule Inhibitors

within the cell cycle (e.g., Plk-1 for ON-01910Na) or in signal transduction pathways (e.g., Trk-A for PHA-848125, VEGFRs for ZK304709, GSK3b, Mek1, Jnk and others for RGB-286638). Development of these dual activity compounds follows a change of perception within the kinase community about selective vs. multi-target inhibition after the establishment of the clinical proof of concept of multi-targeted compounds such as Sorafenib® and Sunitinib®, but also, retrospectively, with Gleevec®. Therefore, this class of molecules, which co-target CDKs and non cyclin-dependent kinases, may prove to be more effective than CDKselective compounds.

15.2.3.1 ZK304709 ZK304709 is a small molecule belonging to the pyrimidine class, and it was the first novel multi-target growth inhibitor, combining inhibition of cyclin-dependent kinase (IC50 values of 4 nM, 50 nM, and 60 nM against, respectively, CDK2/ Cyclin E, CDK1/Cyclin B and CDK4/Cyclin D) with tyrosine kinase activities (IC50 values between 1 and 30 nM on VEGFR1,2,3 and PDGFR-b). Inhibition of multiple kinases by ZK304709 correlated with antiproliferative activity in different cancer cells at low micromolar concentrations, and with evident effects on tumor vasculature and angiogenesis [18]. ZK304709 has shown better in vivo activity than Doxorubicin,® Paclitaxel,® and Gemcitabine® in a large number of xenograft models, and also in an orthotopic pancreatic carcinoma model with effects both on primary tumors and on metastases. The interesting preclinical profile of the molecule warranted the start of two clinical studies in 2004. In one study, ZK304709 was administered on days 1–7 of a 21-day cycle to fasting patients at a starting dose of 15 mg/qd. Eight dose levels were explored ranging from 15 to 360 mg/day. The drug was well tolerated and the most common adverse event were nausea (58%), vomiting (55%), diarrhoea, and lymphopenia (both 35%), fatigue (23%), anaemia (20%), lethargy (18%), anorexia (13%), and dizziness (10%). However, the lack of increment in blood concentration above the 90 mg dose, which based on preclinical studies may not be sufficient to result in meaningful pharmacological

241

inhibition of kinase activity, caused an anticipated shutdown of the study and, thus, an inability to determine an MTD. The poor solubility/physicochemical properties of the compound may be a possible reason for the absence of dose-dependent pharmacokinetics [19]. In the second dose escalation study [20], in which ZK304709 was administered for 14 days of a 28-day cycle, eight dose levels were explored, ranging from 15 to 285 mg/day. The main adverse effects were similar to the first study, including vomiting (77.1%), nausea (74.3%), and fatigue (51.4%), with dose limiting toxicities being identified as grade 3 dizziness, hypertension, and fatigue. Disease stabilization was reported in a few patients.

15.2.3.2 PHA-848125 This compound, which also originated from Pharmacia and is currently pursued by NMS, is an oral multi-CDK inhibitor (mainly acting on CDK1/Cyclin B (IC50 400.nM), CDK2/Cyclin A (IC50 32.nM), CDK4/Cyclin D (IC50 160.nM), with additional inhibition of tyrosine kinase Trk-A (IC50 50.nM). Inhibition of tumor cell proliferation against a panel of > 100 tumor cell lines was observed in the range 0.1–3.0 mM. In preclinical animal models, this compound showed consistent tumor growth inhibition (TGI: 73–91% in four human xenograft mice models) [21]. PHA-848125 recently completed a Phase I dose escalation study in patients with advanced/metastatic tumors. The compound was given orally, once daily for 7 days in a 2-week cycle in a dose range of 50–300 mg/ day. Recommended dose for phase II (RDPII) was set at 150 mg/day, while the maximum tolerated dose (MTD, based on tremors and reversible ataxia) was 200 mg/day. At RDPII, Cmax and AUC were respectively 1.47 mM and 25 mM.h, with a dose-dependent increase. Drug elimination of the drug was slow, with a half-life of 30–40 h. Six patients with stable disease were documented (four at RPDII) [22, 23]. Different administration schedule (e.g., 4 days on, 3 days off for 3 weeks of a 4/week cycle) are currently being tested to maximize its potential effect. Another Phase I/II clinical study of PHA-848125 given daily for 14 consecutive days every 3 weeks is ongoing in recurrent glioblastoma multiforme patients [24].

242

15.2.3.3 RGB-286638 RGB-286638 is a novel indenopyrazole with a broad spectrum inhibition of CDK activity (IC50 values of 1, 2, 3, 4, 44, and 55 nM against, respectively, CDK9, CDK1/Cyclin B, CDK2/Cylin A, CDK4/Cyclin D, and CDK6/Cyclin D). In addition, it potently inhibits several other kinases, including, GSK3b, Mek-1, Jnk, c-Src, Lyn, and Fyn, with IC50s 50,000 (Chk2) >50,000 (Aur2)

Undisclosed Undisclosed

Undisclosed Undisclosed

Sentinel Oncology Alethia Biotherapeutics

UD Undisclosed

the goal of a trial with AZD-7762 administered as a single intravenous agent and in combination with weekly standard dose irinotecan in patients with advanced solid malignancies (clinicaltrials.gov; ASCO 2008 Analyst Briefing).

16.7.1.3 PF-477736 PF-477736 inhibits Chk1 with a Ki value of 0.49 nM. It is about 100-fold selective for Chk1 over Chk2 (Ki, 47nM). PF-477736 potentiates the effect of a range of DNA-damaging agents (SN-38, doxorubicin, carboplatin and mytomycin-C), especially with gemcitabine.

PF-477736 potentiates the antiproliferative activity of gemcitabine in Colo-205, PC-3, a colon and prostate cancer cell lines, respectively. A 12-fold potentiation of gemcitabine activity was observed. Consistent with other reports, the potentiation was only observed in p53 deficient cell lines. The in vivo efficacy was demonstrated in a Colo205 xenograft model, where PF-477736 significantly potentiated the antitumour activity of docetaxel, leading to enhanced tumor regression and prolonged survival compared to docetaxel alone, with no concomitant systemic toxicity. PF-477736 also potentiates gemcitabine and irinotecan in HT29 mouse xenograft

253

16. Chk1 and Chk2 as Checkpoint Targets

models. The radiation and PF-477736 combination showed a 2.7 enhancement ratio of antitumor efficacy in mice bearing A431 tumors. The treatment with 5 Gy radiation + PF-477736 resulted in a tumor growth delay of 6.7 days, compared to a delay of 2.9 days with radiation alone [74, 84, 85]. Currently, a non-randomized, open-label, uncontrolled Australian and US Phase I trial is ongoing in 30 patients with advanced solid tumors. The key objectives are to determine the primary endpoint of MTD and overall safety of PF-477736 + gemcitabine. Secondary endpoints include safety of PF-477736 administered alone in a single cycle, pharmacokinetics, pharmacodynamics, and to preliminarily assess antitumor activity of the combination.

16.7.1.4 IC-83 (LY-2603618) IC-83 is a highly selective Chk1 inhibitor, under development by ICOS (Lilly) as a chemo/radiosensitizer. Only limited preclinical data are available in the public domain although ICOS disclosed in a company Web Page that lead compounds have been evaluated in safety, pharmacology, and proof-ofefficacy studies in animal models. Recently, IC-83 has entered a Phase I study in combination with Pemetrexed. It is a US non-randomized, openlabel, uncontrolled, dose escalation Phase I trial which will enroll 80 cancer patients to assess the primary endpoint of safety and tolerability of IC-83 administered after pemetrexed treatment at 500 mg/m2 q 21 days. Secondary endpoints include pharmacokinetics and antitumour activity. Completion of the trial was expected in December 2008 (clinicaltrials.gov).

16.7.1.5 CBP-501 CBP-501 is a synthetic peptide mimetic of CDC25C, where an Ala replaces the Ser216 residue. It is developed by CanBas and licensed by Takeda as a substrate inhibitor of Chk1, Chk2, MK2, and cTak with the EC50 values of 3.4, 6.5, 0.9 and 1.4 mM, respectively. CBP-501 abrogates the G2 checkpoint and enhances cytotoxicity of cisplatin and Bleomycin in Jurkat, Miapaca, and other cancer cell lines. In xenograft models, CBP-501 augments the antitumor activity of cisplatin and Bleomycin without increasing adverse effects. The t1/2 and MTD of CBP-501 in mice are 3 h and 99 mg/kg ip, respectively. The effective doses in SCID/human

xenograft models are in the range 5-40 mg/kg. It is in Phase I trials against malignant mesothelioma and other type cancers [75]. In a US dose-escalation Phase I trial, 176 refractory solid tumour patients were given CBP-501 iv as a 1-hr infusion on days 1, 8, and 15, q 28 day, it decreased phosphorylation of Cdc25C. The MTD was not reached. No DLT or grade 3–4 toxicity was reported at up to 9.5 mg/m2, and the main toxicity associated with treatment was grade 2 allergic reaction without haemodynamic or respiratory changes. Stable disease of > 5 months was reported in one heavily pretreated ovarian cancer patient. It is also in US phase I and II trials in combination with cisplatin. The phase I part is a dose-finding study of escalating doses of CBP-501 combined with full-dose cisplatin and pemetrexed in patients with histologically confirmed solid malignancy where MTD will be determined based on DLTs occurring during the first treatment cycle and pharmacokinetics of the triplet combination will be assessed. The phase II part will evaluate full-dose cisplatin and pemetrexed combined with CBP-501 at the MTD. Previously untreated, unresectable malignant pleural mesothelioma patients will be randomized in a 2:1 ratio to pemetrexed, cisplatin, and CBP-501 or to pemetrexed and cisplatin (clinicaltrials.gov).

16.7.1.6 UCN-01 UCN-01 is a non-selective kinase inhibitor and inhibits Chk1 (IC50 = 10 nM), Chk2 (IC50 = 10 nM), PKC-alpha (IC50 = 530 nM) [86–88], and many other kinases including several of the cyclindependent kinases. It is conceivable that its clinical effect may not be solely due to the result of inhibition of Chk1 activity [89]. UCN-01 is the first checkpoint kinase inhibitor as a sensitizing agent to enter the clinical trial. However, the clinical development of UCN-01 has been hampered by its unfavorable pharmacokinetic properties and toxicities, for instance, hypotension, hyperglycemia, and pulmonary dysfunction, most of which are likely attributable to kinase targets other than Chk1 [90, 91]. Nevertheless, it has been studied in clinical phase I trials in combination with perifosine, cisplatin, and irinotecan [92–94] (clinicaltrials.gov) and in a phase II trial as a single agent. In one phase I trial in 12 advance solid tumor patients, UCN

254

01 + topoteacan showed antitumor activity with one partial response (ovarian cancer) and three stable diseases. The recommended dose for phase II study in advanced solid tumor patients is 70 mg/M2 of UCN-01 in combination with topoteacan 1 mg/M2 [95](clinicaltrials.gov).

16.7.2 Chk1 Inhibitors in Preclinical Studies Several Chk1 inhibitors are in preclinical development, and the available information, albeit limited, is discussed below. Others identified as leads by Merck, Sareum, BioFocus, Amphora, and Hutchison China MediTech collaborating with Merck KGaA are in very early stages in their program to optimize cellular and in vivo antitumor activity and will not be discussed here.

H. Zhang et al.

16.7.2.3 S-024 and S-144 S-024 and S-144 are two different leads in a series of Chk1 inhibitors, under development by Sentinel Oncology for the treatment of cancer. S-024 is in preclinical development and testing against in vivo models of cancer are underway to evaluate efficacy (Pharmaprojects, Informa UK Ltd).

16.7.2.4 AB-IsoG AB-IsoG is a checkpoint kinase 1 inhibitor, under development by Alethia Biotherapeutics for the treatment of ovarian cancer. AB-IsoG sensitizes ovarian cancer cells lines to topotecan or doxorubicin. In xenograft models, AB-IsoG displays significant tumor shrinkage without toxicity to normal cells. It has also shown efficacy in breast and colon cancer (Company Web Page, Alethia 23 Apr 2007; Pharmaprojects, Informa UK Ltd).

16.7.2.1 A-776574 There are a number of reports published by researchers at Abbott covering three series of Chk1 inhibitors (benzodiazepinone, tricyclic pyrazole, and macrocyclic urea) that are under development for the treatment of cancer. Preclinical results indicate that all three series of compounds are potent and selective Chk1 inhibitors. In particular, the benzodiazepinone series of compounds are very potent, with IC50 values in the low nM range against Chk1 and achieve more than 500-fold selectivity over a range of kinases. It broadly potentiates chemotherapeutics and broadly sensitizes cancer cell lines, with colon cancer and melanoma being indicated as most likely to respond to Chk1 inhibitors. A-776574 (a benzodiazepinone compound) potentiates the efficacy of irinotecan in Colo205 and DLD-1 colon xenografts models [41, 96].

16.7.2.2 CHIR-124 CHIR-124 is a quinolone-based small molecule in development by Novartis. It is a very potent and selective Chk1 inhibitor. The IC50 values against Chk1 and Chk2 are 0.3 and 697 nM, respectively [42, 97]. CHIR-124 abrogates S and G2/M checkpoints and potentiates both camptothecin and SN-38 in p53-deficient tumor cells in vitro and in vivo (especially in the MDA-MD-435 breast cancer orthotopic xenograft model) [43].

16.8 Biomarkers for a Chk1 Inhibitor Biomarkers are important tools for stratifying patients, establishing proper drug dose and schedule, and for quantifying drug benefits in the clinical phase of drug development. The resulting information is critical for making informed decisions on drug safety and efficacy, which help reduce the attrition rate in clinical studies, and thereby, decrease the overall cost of drug development. More importantly, biomarkers that are capable of predicting drug efficacy can be used to stratify patient populations and bring the new medicine to the right patients more rapidly than the conventional hit and miss approach [98]. There are three main types of biomarkers, each serving different purposes; (1) cancer or disease progression related markers that can be used to evaluate drug efficacy; (2) efficacy predictive markers that can predict the activity of the drug in different patient categories and hence help stratify patients; (3) pharmacodynamic (PD) markers that can provide guidance in determining the most effective drug dose and schedule. Having a reliable PD marker constitutes a viable alternative to the conventional MTD, which may reduce undesirable adverse events. This marker is especially important

255

16. Chk1 and Chk2 as Checkpoint Targets

for drugs that are devoid of any apparent toxicity, which makes it impossible or impractical to obtain an MTD in the case of selective Chk1 inhibitors.

16.8.1 Biomarkers Predictive of Chk1 Inhibitor Efficacy It has been demonstrated that Chk1 is the major mediator of cell-cycle checkpoints in response to various chemotherapeutics [22, 51, 69, 71]. Elimination of Chk1 will potentiate chemotherapeutics by abrogating these checkpoints leading to increased apoptosis and cell death [22, 63, 86]. It is now clear that Chk1 phosphorylates the family of Cdc25 phosphatases, which in turn inhibit CDK2 and Cdc2 activities and prevent their premature activation, thus maintaining the highly regulated temporal order of cell cycle progression. In the event of cell cycle alteration due to DNA damage, Chk1 is activated to phosphorylate Cdc25 leading to its degradation via the proteasomal pathway or sequestration to cytoplasm by binding to 14-3-3 protein [57]. As a consequence, Cdc2 is inactivated and cells are arrested at checkpoints until the damaged DNA is repaired. Under this circumstance, inhibition of Chk1 leads to the improper activation of Cdc2 resulting in checkpoint abrogation and presumably mitotic catastrophe. Therefore, in the event of G2/M checkpoint abrogation, Cdc2 activity is a key factor inducing mitotic catastrophe and cell death. It is also well known that cyclin B1 is a cofactor required for Cdc2 activity. An increase in cyclin B1 expression in the late G2-phase is essential for Cdc2 activation leading to M phase entry. Consequently, cyclin B1 may also be responsible as a “rate limiting factor” for the improper activation of Cdc2 activity resulting in premature mitosis and mitotic catastrophe incurred through the checkpoint abrogation. Therefore, it is conceivable that the cyclin B1 level can be a major factor in determining the cellular efficacy of a Chk1 inhibitor in potentiation of topoisomerase inhibitors. Indeed, the published results are consistent with this notion, and it has been demonstrated that the cellular levels of cyclin B1 positively correlate with the degree of potentiation achieved in various cancer cell lines. Down regulation of cyclin B1 leads to impairment of the induction of mitotic catastrophe as well as decreased potentiation of topoisomerase inhibitors by either Chk1 siRNA

or a small molecule Chk1 inhibitor. Of additional interest, colon cancer cell lines in general appear to express higher levels of cyclin B1 and also display higher sensitivity to Chk1 inhibitors, implying that Chk1 inhibitor may be more efficacious in treating colon cancers. Thus, cyclin B1 has been proposed and tested as a biomarker predictive of the efficacy of Chk1 inhibitors across different types of cancer cells [96]. Further validation of this biomarker in a clinical study is warranted. This finding will potentially be very useful for the stratification of patients for Chk1 inhibitor clinical trials and hence, maximize its chance of success.

16.8.2 Pharmacodynamic Markers for Establishing Dose and Schedule The ability to effectively monitor the effects of Chk1 inhibition in vivo is important for establishing the most effective dose and schedule of a Chk1 inhibitor. It has been shown that upon activation of Chk1, Cdc25A is rapidly degraded and the phosphorylation at Y15 of Cdc2 is increased. As a result, cyclin B1 accumulates due to cells arrested in S or G2. Conversely, in the presence of a Chk1 inhibitor, Cdc25A is protected from degradation, and Y15 site of Cdc2 is dephosphorylated. Concomitantly, the level of cyclin B1 decreases, because of the effect of checkpoint abrogation [22, 32, 71, 96, 99]. Together, they serve as pharmacodynamic (PD) markers that may help determine the effective dose and schedule for Chk1 inhibitors in animal models. So far, tumor samples from in vivo studies showed the expected profile changes with respect to these candidate markers and demonstrated that Cdc25A, cyclin B1 and Cdc2-Y15P can be used as PD markers in vivo to determine the most effective dose of a Chk1 inhibitor [96]. More detailed quantitative studies are required to develop a guideline to determine the most effective dose for a Chk1 inhibitor. Notably, tissue sample accessibility is always a difficult issue for biomarker studies in clinical settings. Using circulating tumor cells may be a potential means to resolve this. It has been reported that tumor cells disseminate into blood stream even at early stage when the tumor burden is small [100]. Thus, isolation of circulating tumor cells (CTCs) in a blood sample may be a convenient and powerful tool to collect cancer cells for the purposes of biomarker study [101]. The Cellsearch™ system is

256

a simple blood test that captures and assesses CTC to determine the prognosis of patients, and which has been cleared by FDA for clinical trials [102]. Molecular profiling of CTCs using cytogenetics and cytochemistry analysis allows researchers to measure the copy number of a given gene and the expression level of a specific protein in CTCs [103, 104]. This technology is considered as a real time “biopsy” in cancer patients and certainly can be a valuable tool to apply in biomarker studies.

16.9 Conclusion Over the last two decades, significant advances in the management of cancer and development of novel targeted therapies have emerged. However, cancer still remains a leading cause of death, and chemotherapy and radiation therapy remain the mainstay of treatment for most patients with cancer. Success in developing a Chk1 inhibitor as a chemo or radio-sensitizer can greatly improve the efficacy and increase the therapeutic ratio of the traditional cancer therapy and address unmet medical needs. Currently, several Chk1 inhibitors have entered phase I clinical studies. The key objectives are to determine the primary endpoint of MTD, to assess the overall safety and tolerability for the combination treatment, and to obtain information on pharmacokinetics. Since a selective Chk1 inhibitor should not display any single agent activity, the conventional MTD may not be applicable for some of the inhibitors. Thus, PD markers will be a useful alternative to establish effective drug dose and schedule. Even though the proof of an improved schedule ultimately needs to come from well-run Phase III trials, the search among schedules could be shortened by the use of surrogate endpoints. The Chk1 inhibitor certainly represents a promising new area of anticancer drugs. The outcome of early clinical trials involving these agents is still awaited. The results of current and upcoming clinical trials should demonstrate (1) if the inhibitors exacerbate the toxicity of standard therapies and affect the therapeutic ratio, (2) if simultaneous inhibition of Chk2 activity is beneficial for the treatment, and (3) if a Chk1 inhibitor can overcome MDR resistance in cancer.

H. Zhang et al.

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16. Chk1 and Chk2 as Checkpoint Targets 19. Martinho RG et al (1998) Analysis of Rad3 and Chk1 protein kinases defines different checkpoint responses. EMBO J 17(24):7239–7249 20. Shieh SY et al (2000) The human homologs of checkpoint kinases Chk1 and Cds1 (Chk2) phosphorylate p53 at multiple DNA damage-inducible sites. Genes Dev 14(3):289–300 21. Furnari B et al (1999) Cdc25 inhibited in vivo and in vitro by checkpoint kinases Cds1 and Chk1. Mol Biol Cell 10(4):833–845 22. Xiao Z et al (2003) Chk1 mediates S and G2 arrests through Cdc25A degradation in response to DNA damaging agents. J Biol Chem 278:21767–21773 23. Chehab NH et al (2000) Chk2/hCds1 functions as a DNA damage checkpoint in G(1) by stabilizing p53. Genes Dev 14(3):278–288 24. Zhao H, Piwnica-Worms H (2001) ATR-mediated checkpoint pathways regulate phosphorylation and activation of human Chk1. Mol Cell Biol 21(13):4129–4139 2 5. de Klein A et al (2000) Targeted disruption of the cell-cycle checkpoint gene ATR leads to early embryonic lethality in mice. Curr Biol 10(8):479–482 26. Matsuoka S, Huang M, Elledge SJ (1998) Linkage of ATM to cell cycle regulation by the Chk2 protein kinase. Science 282(5395):1893–1897 27. Cliby WA et al (2002) S phase and G2 arrests induced by topoisomerase I poisons are dependent on ATR kinase function. J Biol Chem 277(2):1599–1606 28. Flaggs G et al (1997) Atm-dependent interactions of a mammalian chk1 homolog with meiotic chromosomes. Curr Biol 7(12):977–986 29. Kaneko YS et al (1999) Cell-cycle-dependent and ATM-independent expression of human Chk1 kinase. Oncogene 18(25):3673–3681 30. Liu Q et al (2000) Chk1 is an essential kinase that is regulated by Atr and required for the G(2)/M DNA damage checkpoint. Genes Dev 14(12):1448–1459 31. Furuta T et al (2003) Phosphorylation of histone H2AX and activation of Mre11, Rad50, and Nbs1 in response to replication-dependent DNA double-strand breaks induced by mammalian DNA topoisomerase I cleavage complexes. J Biol Chem 278(22): 20303–20312 32. Busino L et al (2003) Degradation of Cdc25A by beta-TrCP during S phase and in response to DNA damage. Nature 426(6962):87–91 33. Mailand N et al (2002) Regulation of G(2)/M events by Cdc25A through phosphorylation- dependent modulation of its stability. EMBO J 21(21):5911– 5920 34. Donzelli M et al (2002) Dual mode of degradation of Cdc25 A phosphatase. EMBO J 21(18):4875

257 35. Peng CY et al (1997) Mitotic and G2 checkpoint control: Regulation of 14–3-3 protein binding by phosphorylation of Cdc25C on serine-216. Science 277(5331):1501–1505 36. Graves PR et al (2001) Localization of human Cdc25C is regulated both by nuclear export and 14-3-3 protein binding. Oncogene 20(15):1839–1851 37. Hirao A et al (2002) Chk2 Is a Tumor Suppressor That Regulates Apoptosis in both an Ataxia Telangiectasia Mutated (ATM)-Dependent and an ATM-Independent Manner. Mol Cell Biol 22(18):6521–6532 38. Takai H et al (2000) Aberrant cell cycle checkpoint function and early embryonic death in Chk1(–/–) mice. Genes Dev 14(12):1439–1447 39. Chen Z et al (2002) Human Chk1 expression is dispensable for somatic cell death and critical for sustaining G2 DNA damage checkpoint. Mol Cancer Ther 2:543–548 40. Matthews DJ et al (2007) Pharmacological abrogation of S-phase checkpoint enhances the anti-tumor activity of Gemcitabine in vivo. Cell Cycle 6(6):104–110 41. Xiao Z et al (2006) The discovery of a Chk1 inhibitor and the biomarkers. The 2nd Protein Kinases in Drug Discovery conference at Boston 42. Tse AN et al (2007) CHIR-124, a novel potent inhibitor of Chk1, potentiates the cytotoxicity of topoisomerase I poisons in vitro and in vivo. CCR06-1424. Clin Cancer Res 13(2):591–602 43. Hibner B et al (2004) Small molecule inhibitors of CHK1 cause abrogation of the G2/M cell cycle checkpoint in vitro and tumor regression in a murine xenograft model. AACR Meet Abstr 2004(1):537-a 44. Abraham RT (2001) Cell cycle checkpoint signaling through the ATM and ATR kinases. Genes Dev 15(17):2177–2196 45. Lukas C et al (2001) DNA damage-activated kinase Chk2 is independent of proliferation or differentiation yet correlates with tissue biology. Cancer Res 61(13):4990–4993 46. Sorensen CS et al (2003) Chk1 regulates the S phase checkpoint by coupling the physiological turnover and ionizing radiation-induced accelerated proteolysis of Cdc25A. Cancer cell 3:247–258 47. Gatei M et al (2003) Ataxia-telangiectasia-mutated (ATM) and NBS1-dependent phosphorylation of Chk1 on Ser-317 in response to ionizing radiation. J Biol Chem 278(17):14806–14811 48. Chen M-S et al (2001) Absence of apparent phenotype in mice lacking Cdc25C protein phosphatase. Mol Cell Biol 21(12):3853–3861 49. Mailand N et al (2000) Rapid destruction of human Cdc25A in response to DNA damage. Science 288(5470):1425–1429

258 50. Zhou BB, Elledge SJ (2000) The DNA damage response: Putting checkpoints in perspective. Nature 408(6811):433–439 51. Zhao H, Watkins JL, Piwnica-Worms H (2002) Disruption of the checkpoint kinase 1/cell division cycle 25A pathway abrogates ionizing radiationinduced S and G2 checkpoints. Proc Natl Acad Sci USA 99(23):14795–14800 52. Uto K et al (2004) Chk1, but not Chk2, inhibits Cdc25 phosphatases by a novel common mechanism. EMBO J 23:3386–3396 53. Sorensen CS et al (2004) ATR, claspin and the Rad9-Rad1-Hus1 complex regulate Chk1 and Cdc25A in the absence of DNA damage. Cell Cycle 3(7):941–945 54. Ahn J, Prives C (2002) Checkpoint kinase 2 (Chk2) monomers or dimers phosphorylate Cdc25C after DNA damage regardless of threonine 68 phosphorylation. J Biol Chem 277(50):48418–48426 55. Zhu Y et al (2004) Intra-S-phase checkpoint activation by direct CDK2 inhibition. MCB.24.14.62686277.2004. Mol Cell Biol 24(14):6268–6277 56. Rainey MD et al (2007) Chk2 is required for optimal mitotic delay in response to irradiation-induced DNA damage incurred in G2 phase. Oncogene 27(7):896–906 57. Bartek J, Lukas J (2003) Chk1 and Chk2 kinases in checkpoint control and cancer. Cancer Cell 3(5):421–429 58. Lukas J et al (1997) Cyclin E-induced S phase without activation of the pRb/E2F pathway. Genes Dev 11(11):1479–1492 59. Suganuma M et al (1999) Sensitization of cancer cells to DNA damage-induced cell death by specific cell cycle G2 checkpoint abrogation. Cancer Res 59(23):5887–5891 60. Koniaras K et al (2001) Inhibition of Chk1-dependent G2 DNA damage checkpoint radiosensitizes p53 mutant human cells. Oncogene 20(51):7453–7463 61. Luo Y et al (2001) Blocking Chk1 expression induces apoptosis and abrogates the G2 checkpoint mechanism. Neoplasia (New York) 3(5):411–419 62. Zehan Chen ZX, Wen-zhen Gu, John Xue, Mai H. Bui, Peter Kovar, Gaoquan Li, Gary Wang, Zhi-Fu Tao, Yunsong Tong, Nan-Horng Lin, Hing L. Sham, Jean Y.J. Wang, Thomas J. Sowin, Saul H. Rosenberg, Haiying Zhang (2006) Selective Chk1 inhibitors differentially sensitize p53-deficient cancer cells to cancer therapeutics. Int J Cancer 119(12):2784–2794 63. Eastman A et al (2002) A novel indolocarbazole, ICP-1, abrogates DNA damage-induced cell cycle arrest and enhances cytotoxicity: Similarities and differences to the cell cycle checkpoint abrogator UCN-01. Mol Cancer Ther 1(12):1067–1078

H. Zhang et al. 64. Jackson JR et al (2000) An indolocarbazole inhibitor of human checkpoint kinase (Chk1) abrogates cell cycle arrest caused by DNA damage. Cancer Res 60(3):566–572 65. Donzelli M, Draetta GF (2003) Regulating mammalian checkpoints through Cdc25 inactivation. EMBO Rep 4(7):671–677 66. Zaugg K et al (2007) Cross-talk between Chk1 and Chk2 in double-mutant thymocytes. Proc Natl Acad Sci 104(10):3805–3810 67. Hirao A et al (2000) DNA damage-induced activation of p53 by the checkpoint kinase Chk2. Science 287(5459):1824–1827 68. Xiao Z et al (2005) Novel indication for cancer therapy: Chk1 inhibition sensitizes tumor cells to antimitotics. Int J Cancer 115:528–538 69. Shao RG, Cao CX, Pommier Y (2004) Abrogation of Chk1-mediated S/G2 checkpoint by UCN-01 enhances ara-C-induced cytotoxicity in human colon cancer cells. Acta Pharmacol Sin 25(6):756–762 70. Xiao Z et al (2006) Differential roles of checkpoint kinase 1, checkpoint kinase 2, and mitogen-activated protein kinase-activated protein kinase 2 in mediating DNA damage-induced cell cycle arrest: Implications for cancer therapy. Mol Cancer Ther 5(8):1935–1943 71. Xiao Z et al (2005) A novel mechanism of checkpoint abrogation conferred by Chk1 downregulation. Oncogene 24(8):1403–1411 72. Janetka J et al (2007) Inhibitors of checkpoint kinases: From discovery to the clinic. Curr Opin Drug Discov Devel 10(4):473–486 73. Ashwell S, Zabludoff S (2008) DNA damage detection and repair pathways – recent advances with inhibitors of checkpoint kinases in cancer therapy. Clin Cancer Res 14(13):4032–4037 74. Teng M et al (2007) Structure-Based Design and Synthesis of (5-Arylamino-2H-pyrazol-3-yl)biphenyl-2¢, 4¢-diols as Novel and Potent Human CHK1 Inhibitors. J Med Chem 50(22):5253–5256 75. Sha S-K et al (2007) Cell cycle phenotype-based optimization of G2-abrogating peptides yields CBP501 with a unique mechanism of action at the G2 checkpoint. Mol Cancer Ther 6(1):147–153 76. Duhem C, Ries F, Dicato M (1996) What does multidrug resistance (MDR) expression mean in the clinic? Oncologist 1(3):151–158 77. Ou Y-H et al (2005) p53 C-terminal phosphorylation by CHK1 and CHK2 participates in the regulation of DNA-damage-induced C-terminal acetylation. Mol Biol Cell 16(4):1684–1695 78. Craig A et al (2003) Allosteric effects mediate CHK2 phosphorylation of the p53 transactivation domain. EMBO Rep 4(8):787–792

16. Chk1 and Chk2 as Checkpoint Targets 79. Pereg Y et al (2006) Differential roles of ATM- and Chk2-mediated phosphorylations of Hdmx in response to DNA damage. Mol Cell Biol 26(18):6819–6831 80. Chen L et al (2005) ATM and Chk2-dependent phosphorylation of MDMX contribute to p53 activation after DNA damage. EMBO J 24:3411–3422 81. Komarova EA et al (2004) Dual effect of p53 on radiation sensitivity in vivo: p53 promotes hematopoietic injury, but protects from gastro-intestinal syndrome in mice. Oncogene 23(19):3265–3271 82. Takai H et al (2002) Chk2-deficient mice exhibit radioresistance and defective p53-mediated transcription. EMBO J 21:5195–5205 83. Jack MT et al (2002) Chk2 is dispensable for p53mediated G1 arrest but is required for a latent p53mediated apoptotic response. Proc Natl Acad Sci 99(15):9825–9829 84. Blasina A et al (1999) A human homologue of the checkpoint kinase Cds1 directly inhibits Cdc25 phosphatase. Curr Biol 9(1):1–10 85. Blasina A et al (2008) Breaching the DNA damage checkpoint via PF-00477736, a novel small-molecule inhibitor of checkpoint kinase 1. Mol Cancer Ther 7(8):2394–2404 86. Zhao B et al (2002) Structural Basis for Chk1 Inhibition by UCN-01. J Biol Chem 277(48):46609–46615 87. Seynaeve C et al (1994) Differential inhibition of protein kinase C isozymes by UCN-01, a staurosporine analogue. Mol Pharmacol 45(6):1207–1214 88. Jiang X et al (2004) Inhibition of Chk1 by the G2 DNA damage checkpoint inhibitor isogranulatimide. Mol Cancer Ther 3(10):1221–1227 89. Dasmahapatra GP et al (2004) In vitro combination treatment with perifosine and UCN-01 demonstrates synergism against prostate (PC-3) and lung (A549) epithelial adenocarcinoma cell lines. Clin Cancer Res 10(15):5242–5252 90. Dees EC et al (2005) A phase I and pharmacokinetic study of short infusions of UCN-01 in patients with refractory solid tumors. Clin Cancer Res 11(2):664–671 91. Hotte SJ et al (2006) Phase I trial of UCN-01 in combination with topotecan in patients with advanced solid cancers: A Princess Margaret Hospital Phase II Consortium study. Ann Oncol 17(2):334–340 92. Senderowicz AM (2003) Small-molecule cyclindependent kinase modulators. Oncogene 22(42): 6609–6620 93. Perez RP et al (2006) Modulation of cell cycle progression in human tumors: A pharmacokinetic and tumor molecular pharmacodynamic study of cisplatin

259 plus the Chk1 inhibitor UCN-01. Clin Cancer Res 12(23):7079–7085 94. Fracasso PM et al (2004) Phase I study combining UCN-01 with irinotecan in resistant solid tumor malignancies. J Clin Oncol (Meet Abstr) 22(14_Suppl):3139 95. Hirte HW (2005) A phase II study of UCN-01 in combination with topotecan in patients with advanced recurrent ovarian cancer: A princess margaret phase II consortium trial. J Clin Oncol (Meet Abstr) 23(16_Suppl):3127 96. Xiao Z et al (2008) Cyclin B1 is an efficacy-predicting biomarker for Chk1 inhibitors. Biomarkers 6:1–18 97. Seiler JA et al (2007) The intra-S-phase checkpoint affects both DNA replication initiation and elongation: Single-cell and -DNA fiber analyses. Mol Cell Biol 27(16):5806–5818 98. Frank R, Hargreaves R (2003) Clinical biomarkers in drug discovery and development. Nat Rev Drug Discov 2(7):566–580 99. Molinari M et al (2000) Human Cdc25 A inactivation in response to S phase inhibition and its role in preventing premature mitosis. EMBO Rep 1(1):71–79 100. Nakagawa T et al (2007) Detection of circulating tumor cells in early-stage breast cancer metastasis to axillary lymph nodes. Clin Cancer Res 13(14): 4105–4110 101. Fiegl M et al (2004) Circulating epithelial cells in breast cancer. N Engl J Med 351(23):2452–2454 102. Riethdorf S et al (2007) Detection of circulating tumor cells in peripheral blood of patients with metastatic breast cancer: A validation study of the cell search system. Clin Cancer Res 13(3):920–928 103. Ntouroupi TG et al (2008) Detection of circulating tumour cells in peripheral blood with an automated scanning fluorescence microscope. Br J Cancer 99:789–795 104. Tewes M et al (2008) Molecular profiling and predictive value of circulating tumor cells in patients with metastatic breast cancer: An option for monitoring response to breast cancer related therapies. Breast Cancer Res Treat 111(3):391–400 105. Galaktionov K, Beach D (1991) Specific activation of cdc25 tyrosine phosphatases by B-type cyclins: Evidence for multiple roles of mitotic cyclins. Cell 67:1181–1194 106. Chen L, Liu TH, Walworth NC (1999) Association of Chk1 with 14-3-3 proteins is stimulated by DNA damage. Genes Dev 13(6):675–685

Chapter 17

Targeting Cdc25 Phosphatases in Cancer Therapy Johannes Rudolph

Abstract The Cdc25 phosphatases serve as important regulators of the eukaryotic cell cycle, both during normal cell division and in the checkpoint response to DNA damage. Additionally, overexpression of Cdc25 phosphatases has been linked to numerous forms of cancer with frequent correlation to a poor clinical outcome. Thus, an attractive approach to cancer therapy would be to inhibit the Cdc25 phosphatases with small molecules. Toward this end, numerous screens of actual and virtual chemical libraries have been undertaken, with some synthetic efforts directed at lead optimization. These efforts have yielded many different classes of molecules, the most important of which are highlighted herein. Keywords Cdc25 phosphatase • Cell cycle • Cancer • Anti-proliferative agents

17.1 Introduction Regulated duplication of cells in a process known as the cell division cycle is essential for the growth and maintenance of all living organisms. The dysfunction of the normal cell division cycle yielding uncontrolled cell growth is a hallmark of cancer, a leading cause of death worldwide [1]. Misregulation of the cell division cycle can occur either through gain-of-function or loss-of-function mutations acquired by viral infection (e.g., human papilloma virus), inheritance (e.g., breast cancer associated with BRCA1 or BRCA2), or most commonly by sporadic mutations that occur in response

to exposure to radiation, chemicals, or spontaneously without apparent cause. In gain-of function mutations (e.g., oncogenic ras) the hyperactivity of oncogenes leads to an accelerated cell cycle, whereas in loss-of-function mutations (e.g., deletion of p53) a normal tumor-suppressive activity is lost. Although these genetic alterations can impinge on any of the multitude of signal transduction processes that exist inside the cell, they all ultimately lead to an increase in the activity of the cyclin-dependent kinases (Cdk/cyclins). The Cdk/cyclins serve as the central regulators of the eukaryotic cell cycle (Fig. 17.1). When Cdk/ cyclin activity is high, cells duplicate their DNA and undergo mitosis. When Cdk/cyclin activity is low, cell division halts. Thus, attempting to inhibit the activity of the Cdk/cyclins seems a valuable approach to control the rampant cell growth that is characteristic of cancer, regardless of the genetic origin of the cancer. It might seem most logical to develop small molecules directly toward the Cdk/cyclins, thus targeting dividing cells much like other chemotherapeutics (e.g., cis-platinum, 6-mercaptopurine) [2]. In fact, the Cdk/cyclins, as protein kinases, belong to a class of well-studied enzymes that contain a large binding pocket for their co-substrate ATP that is also suitable for binding small drug-like molecules. Despite the high homology among the greater than 500 known protein kinases, subtle differences in the active sites have allowed for the development of specific kinase inhibitors [3]. Clinically, protein kinase inhibitors have also proven to be of great therapeutic value. For example, inhibition of the


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Figure 17.1. Overview of the role of Cdc25 phosphatases in cell cycle regulation. Cdks associated with various cyclins are responsible for driving cell cycle progression through G1, S, G2, and M phases. Cdk/cyc activity is regulated in part by phosphorylation (by Wee1 and Myt1 kinases) and dephosphorylation (by Cdc25 phosphatases). A primary signaling route for halting cell cycle progression in response to DNA damage is mediated through the Cdc25 phosphatases.

protein kinase Bcr-Abl by imatinib is used in the treatment of chronic myelogenous leukemia (CML) [4] and gefitinib inhibition of epidermal growth factor repressor (EGFR) is used in the treatment of nonsmall cell lung cancer (NSCLC) [5]. However, despite much effort in the pharmaceutical industry, inhibiting the Cdk/cyclins directly has yet to yield an FDA-approved drug. The difficulties arise in part because of the existence of a large family of Cdk/cyclins (Cdk1 – Cdk9, variable association with multiple cyclins) with partially overlapping functions. Thus, developing a molecule that targets the selected subset of the Cdk/cyclins most associated with cell growth (Cdk1 and Cdk2) while avoiding those associated with neural function (Cdk5), as well as the other related kinases, is not a trivial task.

17.2 Targeting Cdc25 An alternative approach to inhibiting the Cdk/ cyclins is to inhibit one of the regulatory proteins that directly activates these protein kinases. Newly formed Cdk/cyclin complexes initially accumulate in an inactivate state wherein the Cdk subunit is phosphorylated on two adjacent residues (Thr14 and Tyr15 in human Cdk1 and Cdk2) by the Wee1 and Myt1 kinases (Fig. 17.1) [6]. Dephosphorylation of these two sites, and thereby activation of kinase activity, is catalyzed by the Cdc25 phosphatases. In humans, there are three Cdc25s (A, B, and C) that have overlapping substrate preferences for the different Cdk/cyclin complexes. The three Cdc25s thus

participate in both the G1-S and G2-M transitions of cell cycle regulation during normal cell division. All three Cdc25s also contribute to the checkpoint response. Insults to the cell such as DNA damage, oxidative stress, or radiation lead to re-localization or degradation of the Cdc25s [7]. Although there exists functional redundancy among the Cdc25s toward the Cdk/cyclins most associated with cell cycle progression (i.e., Cdk1/cyclin B and Cdk2/ Cyclin A), the Cdc25s are not known to activate the neuronal Cdk5 or other noncyclin-dependent kinases. Therefore, targeting the Cdc25s as a means to block Cdk/cyclin activity and cell growth may actually afford higher selectivity and efficacy than inhibiting all Cdk/cyclins or a single member of the Cdk/cyclin family, respectively. Making the Cdc25 phosphatases even more attractive candidates for the development of cancer therapeutics are the proof-of-principle studies performed in cell lines as well as their observed overexpression in numerous cancers (reviewed in [8]). Cdc25A and Cdc25B, and not Cdc25C, are proto-oncogenes and selected overexpression of these two phosphatases leads to an accelerated cell cycle. Importantly, inhibiting one or more of the Cdc25s by microinjection of antibodies or treatment with antisense or siRNA leads to a block in S-phase and/or G2 arrest. Most recently, it has been shown that siRNA targeting Cdc25B significantly inhibited the growth of hepatocellular carcinoma cells in culture as well their xenografts in nude mice [9]. Naturally occurring overexpression of Cdc25A and Cdc25B, and not Cdc25C, has been detected in tumors ranging from breast to

17. Targeting Cdc25 Phosphatases in Cancer Theraphy

NSCLC, from ovarian to colorectal carcinoma [10]. In many of these cancers the overexpression of the Cdc25s is frequently associated with a high histological grade and/or a higher rate of relapse.

17.3 Cdc25 Isoforms as Targets Until recently it has appeared wisest to target all three Cdc25 (A, B, and C) phosphatases nonspecifically. Mice lacking both Cdc25B and Cdc25C are viable and completely normal except for female infertility, as also seen for cdc25B−/− mice [11]. Along with the apparent overlapping substrate specificities and expression patterns seen in cellular studies, this has further reinforced the ideas of a functional redundancy among the Cdc25s. Recently, however, in vivo studies of Cdc25A expression have confirmed that Cdc25A alone can serve as a primary driver of tumorigenesis. Transgenic mice that overexpress both H-ras and Cdc25A develop mammary tumors with a significantly shorter latency than the H-ras overexpressing mice alone and cells derived from these tumors are characterized by aneuploidy and deletion of fragile chromosome regions [12]. Additionally, although Cdc25A(−/−) mice are embryonic lethal, the heterozygous Cdc25A(+/−) mice are resistant to tumorigenesis via Ras activation, further indicating a critical and rate-limiting role for Cdc25A in vivo [13]. Although these results suggest that a small molecule inhibitor targeting Cdc25A alone may be sufficient to block cancerous growth, such a solution may not be applicable to all cancer types (i.e., cancers driven by overexpression of Cdc25B). Additionally, achieving selective inhibition among the Cdc25 isoforms may not be practical from a drug discovery perspective because of the high homology among the three Cdc25s, particulary within the active site pocket (see below).

17.4 Characteristics of the Cdc25 Target Cdc25s are enzymes that dephosphorylate both phospho-threonine and phospho-tyrosine on their Cdk/cyclin substrates; they thus belong to the

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subclass of dual specificity phosphatases in the larger family of protein tyrosine phosphatases (PTPs). The Cdc25 proteins can be divided into two primary regions (Fig. 17.2). The N-terminii are 150–300 residues in length, highly divergent in sequence, and contain numerous sites for phosphorylation and ubiquitination. The modifications of the N-terminii are involved in regulating phosphatase activity [14,15], protein levels [16–18], association with other proteins [19–21], and intracellular localization (nuclear vs. cytoplasmic) [22–24]. The C-terminal regions display ~ 60% pairwise identity over ~ 200 amino acids and contain the signature motif (HCX5R) that defines the active sites of all PTPs. The structures of the catalytic domains of the Cdc25s are unlike other PTPs, sharing only the superimposable active site loop that cradles the phosphate of the protein substrate (Fig. 17.2) [25,26]. Sitting beneath the phosphate cradle is the catalytic cysteine, responsible for initiating the dephosphorylation of Cdk/cyclin substrates. As in other PTPs, this catalytic cysteine exists as an unprotonated thiolate and is therefore highly susceptible to oxidation. The highly reactive cysteine serves as a potential means for redox-regulation of phosphatase activity [27,28] as well as a target for irreversible inhibition of the Cdc25s using small reactive molecules (see below). Like many other phosphatases, and in contrast to many other enzymes that utilize protein substrates (e.g., proteases), the Cdc25s lack an apparent substrate recognition site into which the flanking sequence of the site to be dephosphorylated could bind (Fig. 17.2). The surfaces nearby the active sites of the Cdc25s are relatively flat and the active site pockets are extremely shallow and are thus able to accommodate both phospho-threonine and phospho-tyrosine containing substrates. This makes the Cdc25 phosphatases not as readily amenable for binding small molecule competitive inhibitors as compared to other enzymatic targets (e.g., protein kinases). On the other hand, characterization of the interaction between Cdc25s and their Cdk/cyclin substrates has identified a binding pocket on the Cdc25s that is remote from the active site (>20 Å) and is critical for substrate docking both in vitro and in vivo (Fig. 17.2) [29,30]. This remote pocket may be amenable to small molecule inhibition of Cdc25 activity (see below).

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S477 E478 S476 F475

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Figure 17.2. Structural features of the three human Cdc25 phosphatases. Top: Schematized sequence alignment demon strates the diversity of the N-terminal regions, the high homology in the C-terminal catalytic domains, and the completely conserved sequence of the catalytic loop. Bottom left: The catalytic domain of Cdc25B is shown in gray ribbon with the catalytic loop shown in black. A sulfate is shown in space filling diagram above the catalytic loop, representing the phosphate of a bound substrate. The critical arginine of the remote binding pocket for protein substrate is shown in licorice near the bottom of the structure. Bottom right: Blowup of the active site loop with a bound sulfate demonstrating the importance of backbone (not sidechain) interactions in coordinating the sulfate (in space-filling) in the active site.

17.5 Screening for Small Molecule Inhibitors Over the past 10+ years, extensive efforts in both academia and industry, have been devoted to inhibiting Cdc25 phosphatases. Thus far all library screens have been based on inhibiting the phosphatase activity of Cdc25s toward artificial small molecule substrates such as para-nitrophenyl phosphate, thereby targeting the shallow active site pocket. These screens have mostly lead to the identification of quinoid-based compounds, with some representation by more diverse chemical classes including dysidiolides, sulfiricins, steroid derivatives, and phosphate surrogates [31–33]. Because of the recent comprehensive compilation of Cdc25 inhibitors by the Ducommun group [33], we review

herein only selected examples, in part to demonstrate the diversity of compounds and the difficulty in generating potent and selective inhibitors against the Cdc25 phosphatases. The most potent and best-characterized inhibitors of the Cdc25s are the para-quinonoids, whose study has been complicated in that they can serve as electrophiles (i.e., form covalent adducts) and undergo redox cycling (i.e., oxidize susceptible residues). For example, JUN-1111 (1, Fig. 17.3; ~ 0.2 µM) inhibits the Cdc25s in vitro in a time-dependent and redox-dependent manner [34]. Treatment of cells with JUN-1111 yields intracellular reactive oxygen species, an oxidized (and inactivated) Cdc25B, and cell cycle arrest. Modifications of the heterocyclic core appear to be tolerated and provide the opportunity to build greater specificity toward the Cdc25s [35]. Another para-quinone-based inhibitor is

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6: ZL-I-196

Figure 17.3. Representative inhibitors of Cdc25 phosphatases.

BN82685 (2, Fig. 17.3, ~ 0.2 µM), shown to inhibit Cdc25 activity in vitro, in cultured cells, and slow the growth of xenografted tumors in mice following oral administration [36]. In addition, pilot studies in colon cancer cells indicate the potential for combination therapies of BN82685 and paclitaxel, a microtubule inhibitor [37]. Although Cdc25s are not known to dimerize, the most potent compound described thus far is the bis-quinonoid IRC-083864 (3, Fig. 17.3; ~ 20 nM) [38]. Although the reactivity and redox cycling of the quinones suggests a lack of specificity for Cdc25s, not all quinones are able to inactivate the Cdc25s. For example, some modifications of the heterocycle that is fused to the quinone ring in derivatives of JUN-1111 reduce the potency of these compounds toward the Cdc25s [39]. Besides the para-quinoids, there have been a fair number of ortho-quinoids reported as inhibitors of the Cdc25 phosphatases. The most advanced of these is the naphthopyranedione ARQ-501 (4, Fig. 17.3), which entered phase 1 clinical trials as an intravenously administered drug in 2004. According to ArQule, ARQ-501 has now completed a phase 2 proof-of-principle trial as a standalone therapy in leiomysarcoma and head and

neck cancer, having undergone an additional trial in combination with the nucleoside analog gemcitabine. It is probable, however, that ARQ-501 is not directly a Cdc25 inhibitor, but rather functions by some other mechanism. Other reported orthoquinoid inhibitors of the Cdc25s include 5169131 (5, Fig. 17.3; ~ 10 µM), [40] and the indolyldihydroquinone ZL-1-196 (6, Fig. 17.3; ~ 1 µM) [41], of which only 5169131 shows cellular activity. Interestingly, the ortho-quinoids, in contrast to the para-quinoids, appear to be reversible noncovalent inhibitors. Given the highly reactive cysteine in the active site pocket of the Cdc25s, some efforts at inhibiting Cdc25s have focused on covalently trapping the cysteine with an electrophile. For example, the maleimide PM-20 (7, Fig. 17.4; ~ 1–10 µM) shows surprising selectivity for Cdc25s versus other cysteine-containing phosphatases [42]. Perhaps this specificity can be understood in light of the high reactivity of the cysteine thiolate in the active sites of the Cdc25s, as reflected in their greater susceptibility to oxidation as compared to other PTPs [28]. PM-20 selectively inhibits the growth of human Hep3B tumor cells compared to normal rat hepatocytes and

266

J. Rudolph

slows the growth of rat hepatoma cells in vivo. In a different approach, phosphate mimetics have been studied as inhibitors of the Cdc25s. Dysidiolide (8, Fig. 17.4; ~ 10 µM), with a g-hydroxybutenolide serving as an analog of phosphate [43], has been used as a basis for designing other inhibitors, such as the neodysidiolides (9, Fig. 17.4; ~ 1 µM) [44]. Some compounds have even managed to incorporate both an electrophilic moiety as well as a phosphate mimetic, such as the diterpenoid isolated from a marine anemone (10, Fig.17.4; ~ 4 µM)

[45] and the cinnamic acid derivative TPY-835 (11, Fig. 17.4; ~ 5 µM) [46]. Despite their dual function, neither of these inhibitors shows exceptional potency nor selectivity, and their large size does not bode well for further improvements in potency.

17.6 Virtual Screening Although some progress toward discovering and optimizing small molecule inhibitors of Cdc25s

COOH COOH H

O S

HO

OH

N HO

H

O

S O O

10: diterpenoid

7: PM-20

HO HO HO

8: dysidiolide

HO

9: neodysidiolide

O O

O

O

N Cl

S

N COOH

O

O

11: TPY-835

O

O

O O

N

H S

H OH O 12: macrocycle

HO

Figure 17.4. More representative inhibitors of Cdc25 phosphatases.

O N

13: virtual lead

267

17. Targeting Cdc25 Phosphatases in Cancer Theraphy

has been made, the nanomolar binding constants, cellular potency, and specificity expected of clinical candidates have in general not been achieved. Therefore, virtual screening and docking methods have more recently been applied to discovering novel inhibitors of Cdc25 phosphatases. These approaches take advantage of the multiple known structures of the catalytic domains of Cdc25A [47] and especially Cdc25B [30,48,49], along with advances in computational techniques. For example, a macrocycle (12, Fig. 17.4) was designed with a carbonyl group that is suggested to mimic phosphate binding in the active site. This macrocycle displays only modest selectivity toward the Cdc25s compared to other phosphatases and does not have cellular activity, presumably because of its lipophilicity [50]. In the absence of crystal structures of Cdc25s with bound inhibitors, modeling of many of the known active site inhibitors using molecular docking programs has provided important guidelines towards the design of more potent inhibitors [51]. Most recently, careful selection and computational preparation of the active site of Cdc25B was followed by a multistep virtual ligand screen. After further re-docking, ~ 100 compounds were identified that experimentally inhibited Cdc25B, including a novel compound with an IC-50 of 13 µM in vitro (13, Fig. 17.4) [52]. A similar approach has been suggested but not yet reported for the docking pocket remote from the active site that is critical for binding protein substrate [8].

17.7 Conclusion Targeting the Cdc25 phosphatases as a strategy for an anti-cancer drug continues to be an attractive and intriguing possibility. Proof-of-principle experiments using siRNA and knockout cell lines provide significant, albeit not “home-run” support for the idea that a Cdc25 inhibitor would block rapid cell division in a cancer patient. Although many groups have striven toward discovering inhibitors against the Cdc25s using a multitude of approaches and chemical libraries, the potency and selectivity have not been sufficient enough to progress further and use any compound in the clinic to date. One can hope that continued interest and research in the field of Cdc25 phosphatases will someday overcome this hurdle.

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J. Rudolph 31. Pestell KE, Ducruet AP, Wipf P, Lazo JS (2000) Small molecule inhibitors of dual specificity protein phosphatases. Oncogene 19(56):6607–12 32. Prevost GP, Brezak MC, Goubin F et al (2003) Inhibitors of the Cdc25 phosphatases. Prog Cell Cycle Res 5:225–34 33. Contour-Galcera M-O, Sidhu A, Prevost G, Bigg D, Ducommun B (2007) What’s new on Cdc25 phosphatase inhibitors. Parmacol Therapeut 115:1–12 34. Brisson M, Nguyen T, Wipf P et al (2005) Redox regulation of Cdc25B by cell-active quinolinediones. Mol Pharmacol 68(6):1810–20 35. Cossy J, Belotti D, Brisson M, Skoko JJ, Wipf P, Lazo JS (2006) Biological evaluation of newly synthesized quinoline-5, 8-quinones as Cdc25B inhibitors. Bioorg Med Chem 14(18):6283–7 36. Brezak M-C, Quaranta M, Contour-Galcera M-O et al (2005) Inhibition of human tumor cell growth in vivo by an orally bioavailable inhibitor of CDC25 phosphatases. Mol Cancer Ther 4(9):1378–487 37. Cazales M, Boutros R, Brezak M-C, Chaumeron S, Prevost G, Ducommun B (2007) Pharmacologic inhibition of CDC25 phosphatases impairs interphase microtubule dynamics and mitotic spindle assembly. Mol Cancer Ther 6(1):318–25 38. Galcera MO, Bigg D, Prevost G, Sidhu A (2006) inventors; Cdc25 phosphatase inhibitors patent WO 2006/067311 39. Lazo JS, Aslan DC, Southwick EC et al (2001) Discovery and biological evaluation of a new family of potent inhibitors of the dual specificity protein phosphatase Cdc25. J Med Chem 44:4042–9 40. Brisson M, Nguyen T, Vogt A et al (2004) Discovery and characterization of novel small molecule inhibitors of human Cdc25B dual specificity phosphatase. Mol Pharmacol 66:824–33 41. Sohn J, Kiburz B, Li Z et al (2003) Inhibition of Cdc25 phosphatases by indolyldihydroxyquinones. J Med Chem 46(13):2580–8 42. Kar S, Wang MZ, Yao W, Michejada CJ, Carr BI (2006) PM-20, a novel inhibitor of Cdc25A, induces extracellular signal-regulated kinase 1/2 phosphorylation and inhibits hepatocellular carcinoma growth in vitro and in vivo. Mol Cancer Ther 5:1511–9 43. Gunasekera SP, McCarthy PJ, Kelly-Broger M, Lobkovsky E, Clardy J (1996) Dysidiolide: A novel protein phosphatase inhibitor from the caribbean Sponge Diysidea etherria de Laubenfels. J Am Chem Soc 118:8759–60 44. Shimazawa R, Suzuki T, Dodo K, Shirai R (2004) Design and synthesis of dysidiolide analogs from vitamin D3: novel class of Cdc25A inhibitors. Bioorg Med Chem Lett 14:3291–4

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Index

A Abrogation of checkpoint response, 37, 46, 60–61, 89, 248, 251 Acute myeloid leukemia (AML), 156, 161–164, 178 Akt, 9, 10, 15, 71–74, 76, 100, 102–103, 125, 128, 136, 137, 190, 202, 204, 205, 217–219, 221–223, 225 Anticancer drugs, 89, 245, 256 Apoptosis, 8–11, 13, 15, 17, 18, 27, 46, 72, 76, 83, 84, 87–89, 95–97, 99–103, 110, 111, 116–118, 140, 156, 162, 173, 175–178, 180–182, 193, 194, 199, 202–206, 219, 220, 222–224, 242, 246, 247, 249, 250, 255 Ataxia telangiectasia mutated (ATM), 13, 31, 33, 41, 43, 45, 55–57, 60, 69–71, 73, 74, 83, 85–88, 96–100, 129–131, 136, 155–159, 179–181, 189, 193, 203, 206, 246–248 Ataxia telangiectasia Rad3-related (ATR), 31–33, 41, 43, 45, 55–60, 73, 74, 83, 85–87, 96–100, 129–131, 136, 189, 193, 203, 206, 246–248 Autophagy, 95, 215, 217, 224 B BRCA1, 13, 33, 45, 59, 74, 84–87, 97, 126, 155–164, 202, 261 BRCA2, 33, 59, 155–164, 261 C Cdc2, 37–39, 42–46, 73–76, 171, 180, 200, 247, 248, 251, 255 Cdc25 phosphatases, 7, 31, 38, 55, 73, 86, 89, 172, 180, 247, 255, 261–267 Cell cycle, 3–19, 27, 37, 53, 69, 83, 96–99, 102, 103, 110–114, 117, 118, 123, 126–129, 131, 132, 136–143, 156, 171–172, 176–182, 189, 199, 215, 218–220, 222–224, 235, 245, 261, 262, 264

Cell cycle arrest, 6, 8–10, 12–17, 29, 32, 45, 61, 69, 72–75, 83, 86–88, 96–99, 103, 110, 113, 117, 131, 132, 142, 156, 162, 172–174, 177–179, 189, 190, 193, 199, 200, 202, 206, 215, 224, 245, 247, 249, 264 Cell division cycle 25 (Cdc25), 7, 8, 38–47, 54–55, 57, 74, 75, 86, 88–89, 172, 247, 255, 261–267 Centrosome amplification, 58–61 Centrosomes, 53–61, 86 Checkpoint response(s), 16, 28, 37–47, 53–61, 70, 76, 83–89, 98, 103, 157, 173, 179, 180, 193, 194, 199–201, 203–205, 207, 246–249, 262 Chemoprotection, 250–251 Chk1, 8, 13, 31–33, 41–43, 45, 55–60, 72–76, 86–89, 97, 99, 179, 180, 190–197, 203, 206, 245–256 Chk2, 13, 31, 33, 41, 43, 45, 56, 57, 73–74, 86–89, 97, 99, 179, 180, 190, 203, 206, 245–256 Chk1 inhibitors, 190–193, 195, 196, 247, 249–256 Clinical trials, 118, 142, 177, 181, 190, 192, 207, 224, 235–242, 251–256 Cyclin B, 37–39, 43–46, 54–58, 74, 75, 87, 117, 171, 191–193, 200, 236–242, 247, 248, 262 Cyclin D1, 5, 88, 114, 123–143, 162, 172, 199, 218, 238 Cyclin-dependent kinase (CDK), 4–8, 12, 28, 37, 38, 44, 45, 54, 70, 74–76, 88, 89, 98, 110–114, 116, 118, 138, 171, 172, 175, 178–182, 191–193, 199–203, 207, 215, 218–219, 223, 235–243, 246, 248, 261–263 Cyclin E, 5, 6, 8, 11–14, 37, 44, 45, 88, 99, 111, 113, 134, 135, 142–143, 171, 172, 180, 191, 200, 201, 203, 215, 217, 218, 221, 222, 236, 238, 241, 248 Cyclins, 4–8, 10, 14, 19, 37, 75, 88, 98, 114, 123, 124, 126, 128, 129, 131, 132, 135–141, 143, 171, 172, 176, 178, 191, 192, 200, 201, 219, 236, 249, 261–263

271

272 D Death receptors, 95, 102, 207 DNA damage, 6, 27, 37, 55–61, 69–77, 83–89, 95, 111, 156, 172, 189, 191–196, 199–203. 205, 206, 245, 262 DNA damage response, 13, 30–32, 56, 58–61, 70, 72–74, 76, 85, 95–104, 111, 192 DNA repair, 13, 27, 31–33, 46, 69, 74, 83–85, 87, 89, 95–96, 98, 100–103, 110, 117, 131, 155, 156, 162, 173, 192, 195, 199, 245, 249 DNA replication, 4, 13, 27–31, 54–57, 59, 85, 86, 96, 97, 113, 117, 127, 131, 137, 155, 171, 172, 179, 199, 205, 245, 247 Double strand breaks (DSB), 28, 29, 31, 32, 56, 59, 73, 74, 85, 87, 95–97, 99, 102, 131, 155–158, 162–164, 181, 191, 195, 249 E Early G1 checkpoint, 8–10, 19 E2F, 4, 8, 9, 11, 12, 19, 45, 88, 110–117, 123, 141, 176, 178, 203, 216 Energy checkpoint, 14–16, 19 F Fas, 95, 96, 99–101, 223 14-3-3, 39–44, 56, 69–77, 86, 89, 102, 217, 218, 222, 247, 248, 255 14-3-3s, 45, 46, 70–76, 99 G GADD45, 44–46, 59, 73, 74, 173, 179, 180 G1 arrest, 13, 15, 88, 132, 141, 179, 200–202, 206, 224, 250, 251 G1 checkpoint response, 199, 200, 203, 204, 207 G2 checkpoints, 39, 41, 43–47, 75, 89, 246–248 Gene repression, 112 G1 phase, 3–5, 8, 10, 14, 16, 18, 19, 37, 45, 46, 111, 112, 123–143, 172, 179, 199–204, 217, 218, 240, 245 G1-S transition, 8, 37, 88, 110, 111, 114, 117, 162, 171, 172, 179, 180, 200 H Hypermitogenic stress, 14, 18 I Inhibitors, 4, 31, 70, 86, 99, 110, 127, 172, 189–190, 200, 215, 235, 245–256, 261 Intra-S-phase checkpoint, 28–32 L Leukemias, 10, 13, 30, 58, 155–164, 172, 175, 178, 181, 182, 193, 205, 222, 236, 251, 262

Index Lymphomas, 5, 10, 30, 60, 113, 114, 124, 129, 132, 137, 138, 140–142, 155–164, 179, 181, 194, 202, 219–222, 238, 240, 247, 249, 251 M Mantle cell lymphoma (MCL), 5, 114, 132, 137, 140, 141, 156–158, 162, 178, 179, 181, 240 Mdm2, 6, 13, 59, 70–73, 76, 87, 88, 99, 103, 173–175, 177–181, 195, 196, 204 Microcephalin (MCPH1), 56, 58 Microcephaly, 56, 58, 85 Mitotic catastrophe, 47, 57, 58, 60, 61, 75, 86, 193, 194, 249, 255 Multi-target, 238, 241, 242 N Nucleoside analogs, 191, 193, 265 O Oncogene, 12, 55, 71, 87, 123–125, 138, 159, 175, 219, 249, 261 P p21, 6, 37, 59, 70, 88, 99, 133, 172, 191, 199–207, 218, 245 p53, 6, 13, 15–19, 43–46, 59, 61, 69–75, 86–89, 97–103, 114, 116–118, 171–182, 189–197, 200–207, 218, 219, 245, 246, 249–252, 254, 261 p19Arf, 72, 173, 174, 176–178, 223 Pericentrin, 58 p27Kip1, 6, 8–10, 113, 215–225 PP2A, 42, 43, 73, 85 Predictive biomarker, 216, 225 p21waf1, 73, 190, 191 R Ras, 11, 12, 14, 15, 18, 72, 124, 125, 128, 135, 176–178, 221, 222, 225, 239, 261, 263 Rb, 9, 45, 76, 88, 109–118, 131, 137, 139, 178–179, 203, 204, 219, 238, 240, 249, 141176 RB family, 4, 5, 9, 110, 111, 113–118, 178 Reactive oxygen species (ROS), 12, 16–19, 264 Replication checkpoint, 28, 29, 32–33, 85, 99, 246 Restriction point, 8, 10–14, 18, 19, 76, 113 Retinoblastoma, 4, 11, 12, 45, 76, 109–118, 123, 131, 162, 176, 178, 179, 182, 219 ROS. See Reactive oxygen species S Seckel syndrome, 56, 58, 85 Selectivity, 179, 181, 237, 239, 251, 254, 262, 265–267 Siddik, Z.H., 199–207

Index Small molecule inhibitors, 89, 174–175, 203, 235–243, 248, 263, 264, 266 S–M checkpoint, 28, 29, 33 26 S proteasome, 128, 131–133, 136, 141, 142 Stress checkpoint, 10, 12–15, 17–19 T Targeted therapy, 225, 250 Topoisomerase I inhibitors, 191, 247 Transactivation, 71, 87, 99, 111, 173, 176, 178, 202–207 Trastuzumab, 217, 224, 225, 235

273 Tumorigenesis, 6, 8, 10, 11, 14, 16–19, 27, 58, 61, 73, 76, 88, 103, 104, 110, 111, 113–117, 124, 140, 173, 177, 180, 219, 220, 235, 239, 249, 263 Tumor suppressor, 6, 10, 15, 16, 69, 73, 74, 76, 87, 88, 109, 110, 113, 114, 116–118, 123, 132, 162, 172–174, 178, 200, 202, 204, 216, 218–219, 221, 223, 249 U Ubiquitin, 8, 32, 33, 71–74, 85–87, 89, 99, 102, 103, 114, 127–129, 131–135, 137, 141, 142, 173, 179, 180, 200, 217, 218, 220, 221, 247, 248, 263 UCN-01, 189–196, 253–254

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