Checkpoint Responses in Cancer Therapy (Cancer Drug Discovery and Development)

Checkpoint Responses in Cancer Therapy Cancer Drug Discovery and Development™ Beverly A. Teicher, Series Editor Check...

Author: Wei Dai

135 downloads 2205 Views 7MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!
Report copyright / DMCA form

DOWNLOAD PDF

Checkpoint Responses in Cancer Therapy

Cancer Drug Discovery and Development™ Beverly A. Teicher, Series Editor Checkpoint Responses in Cancer Therapy, edited by Wei Dai, 2008 Cancer Proteomics: From Bench to Bedside, edited by Sayed S. Daoud, 2008 Antiangiogenic Agents in Cancer Therapy, Second Edition, edited by Beverly A. Teicher and Lee M. Ellis, 2007 Apoptosis and Senescence in Cancer Chemotherapy and Radiotherapy, Second Edition, edited by David A. Gerwitz, Shawn Edan Holtz, and Steven Grant, 2007 Molecular Targeting in Oncology, edited by Howard L. Kaufman, Scott Wadler, and Karen Antman, 2007 In Vivo Imaging of Cancer Therapy, edited by Anthony F. Shields and Patricia Price, 2007 Transforming Growth Factor- in Cancer Therapy, Volume II: Cancer Treatment and Therapy, edited by Sonia Jakowlew, 2008 Transforming Growth Factor- in Cancer Therapy, Volume 1: Basic and Clinical Biology, edited by Sonia Jakowlew, 2008 Microtubule Targets in Cancer Therapy, edited by Antonio T. Fojo, 2007 Cytokines in the Genesis and Treatment of Cancer, edited by Michael A. Caligiuri, Michael T. Lotze, and Frances R. Balkwill, 2007

Regional Cancer Therapy, edited by Peter M. Schlag and Ulrike Stein, 2007 Gene Therapy for Cancer, edited by Kelly K. Hunt, Stephan A. Vorburger, and Stephen G. Swisher, 2007 Deoxynucleoside Analogs in Cancer Therapy, edited by Godefridus J. Peters, 2006 Cancer Drug Resistance, edited by Beverly A. Teicher, 2006 Histone Deacetylases: Transcriptional Regulation and Other Cellular Functions, edited by Eric Verdin, 2006 Immunotherapy of Cancer, edited by Mary L. Disis, 2006 Biomarkers in Breast Cancer: Molecular Diagnostics for Predicting and Monitoring Therapeutic Effect, edited by Giampietro Gasparini and Daniel F. Hayes, 2006 Protein Tyrosine Kinases: From Inhibitors to Useful Drugs, edited by Doriana Fabbro and Frank McCormick, 2005 Bone Metastasis: Experimental and Clinical Therapeutics, edited by Gurmit Singh and Shafaat A. Rabbani, 2005 The Oncogenomics Handbook, edited by William J. LaRochelle and Richard A. Shimkets, 2005 Camptothecins in Cancer Therapy, edited by Thomas G. Burke and Val R. Adams, 2005

Checkpoint Responses in Cancer Therapy

Edited by

Wei Dai, phd Department of Environmental Medicine, New York University School of Medicine, Tuxedo, NY

Editor Wei Dai, PhD Department of Environmental Medicine New York University School of Medicine Tuxedo, NY

Series Editor Beverly A. Teicher, PhD Genzyme Corporation Framingham, MA

ISBN: 978-1-58829-930-7

e-ISBN: 978-1-59745-274-8

Library of Congress Control Number: 2007940763 ©2008 Humana Press, a part of Springer Science+Business Media, LLC 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. Cover illustration: Fig. 2A-C, Chapter 11, “Antiproliferation Inhibitors Targeting Aurora Kinases,” by Kishore Shakalya and Daruka Mahadevan. Printed on acid-free paper 987654321 springer.com

Preface The cell cycle is tightly regulated by a number of molecular entities that maintain the genetic integrity of the cell and ensure that genetic information is passed correctly to the daughter cells. Starting in the 1980s, extensive research efforts have revealed the existence of evolutionarily conserved surveillance mechanisms—commonly referred to as checkpoints—that regulate the cell cycle by monitoring specific cellcycle processes and block progression through the cell cycle until these processes have been completed accurately. If any of the molecular events underlying these processes are impaired, the cell cycle stops to allow the cell time for repair. If the cell is damaged beyond repair, it will commit suicide by activating apoptotic processes. The loss of checkpoint function often results in infidelity of DNA replication, chromosome mis-segregation, or both, thereby predisposing the cell to genetic instability and neoplastic transformation. Cancer frequently results from damage to multiple genes controlling cell division. Existing evidence indicates that cancer cells are frequently defective in regulating one cell-cycle checkpoint, which makes them very susceptible to insults to a second checkpoint, resulting in apoptosis. Based on their mechanism of action, current antitumor drugs that target the cell cycle can generally be divided into three categories: inhibition of DNA synthesis, induction of DNA damage, and disruption of mitotic processes. Most cancer drugs used in clinical settings kill cancer cells by altering the DNA structure and inhibiting DNA replication. Such events inevitably lead to activation of the DNA replication or the DNA damage checkpoint. Since the 1980s, much effort has also been directed to the discovery of mitotic targets or processes, which, if altered, can lead to a mitotic catastrophe (a specialized case of apoptosis). It is known that mitotic processes are closely monitored by several surveillance mechanisms, including the G2/M transition checkpoint, the prophase stress checkpoint, and the spindle checkpoint. A defect in the regulation of any mitotic checkpoint often results in genomic instability, which predisposes the cell to malignant transformation. By characterizing the molecular components of cell-cycle checkpoint mechanisms and exploring differences in the checkpoint status between normal cells and malignant cells, we may be v

vi

Preface

able to facilitate the discovery and development of chemotherapeutic compounds that are more effective and more specific for tumor cells. Checkpoint Responses in Cancer Therapy presents summaries of the advances made since the 1980s in identifying various components of cell-cycle checkpoints, elucidating their molecular regulation during checkpoint activation, and validating the use of checkpoint proteins as targets for the development of anticancer drugs. Proteins important for G1 progression are known to mediate cell-cycle checkpoint responses, especially after treatment with chemotherapeutic agents. Considerably less is known about the G1 or G1/S checkpoint than other cell-cycle checkpoints, in terms of their usefulness as targets for anticancer drug discovery. We do know that the Rb protein (pRb), a major tumor suppressor in the cells, plays a pivotal role in the negative control of the cell cycle. The pRb is responsible for regulating the G1 checkpoint, blocking the transition from G1 to S, thereby preventing cell proliferation. Our current knowledge of pRb is summarized in chapter 1. Many chemotherapeutic drugs used in clinical settings target key molecules regulating DNA synthesis or modulating the response to DNA damage. Great progress has been made recently in characterizing the structure and functions of proteins responsible for activating checkpoints that monitor DNA integrity, and many pharmaceutical companies have explored molecular targets that control the S phase checkpoint for therapeutic leads. Detailed discussions of the major molecular players affecting DNA synthesis and the response to DNA damage, from the viewpoint of biology and cancer drug development, are provided in chapters 2 through 6. Many checkpoints monitor the successful execution of the various stages of mitosis to ensure that the cell produces two daughter cells with identical genetic contents. Several protein kinase families are critical for the regulation of these checkpoints. Since the 1990s, intensive drug screening, coupled with studies of kinase structure and function, have led to the identification of a variety of chemical compounds that are capable of inhibiting individual mitotic kinases. Chapters 7 through 12 are devoted to advances in these areas. Posttranslational modifications of the histone tails are closely associated with regulation of the cell cycle as well as chromatin structure. Multiple histone tail acetylations result in the destabilization and decondensation of chromatin fibers. Given that histone acetylation and deacetylation play an important role in the regulation of gene expression and eventually the fate of the cell, much effort has been made to develop histone deacetylase inhibitors to serve as anticancer

Preface

vii

agents. Some of these inhibitors may affect the regulation of cell-cycle checkpoints. Our current knowledge of histone deacetylase inhibitors is summarized in chapter 13. The overall goal of Checkpoint Responses in Cancer Therapy is to enhance our understanding of the many cell-cycle checkpoint molecules that have already been identified as effective targets for anticancer drug development, and to explore the possibility of developing a new generation of anticancer drugs with improved therapeutic indices based on their ability to target a number of checkpoint components that are yet to be fully studied. It is anticipated that this book will serve as a valuable source of information, not only for researchers in the pharmaceutical and biotechnology industries, but also for academic scientists studying cell-cycle regulation, signal transduction, and apoptosis, as well as those involved in cancer research. Wei Dai Tuxedo, NY

Contents Preface .......................................................................................

v

Contributors ...............................................................................

xi

1 RB-Pathway: Cell Cycle Control and Cancer Therapy..... Erik S. Knudsen and Wesley A. Braden

1

2 Targeting the p53/MDM2 Pathway for Cancer Therapy ... 19 Christian Klein and Lyubomir T. Vassilev 3 DNA Topoisomerases as Targets for the Chemotherapeutic Treatment of Cancer .......................... 57 Ryan P. Bender and Neil Osheroff 4 Targeting ATM/ATR in the DNA Damage Checkpoint.... 93 Joseph M. Ackermann and Wafik S. El-Deiry 5 Compounds that Abrogate the G2 Checkpoint................... 117 Takumi Kawabe 6 CDK Inhibitors as Anticancer Agents ................................ 135 Timothy A. Yap, L. Rhoda Molife, and Johann S. de Bono 7 CHFR as a Potential Anticancer Target ............................. 163 Minoru Toyota, Lisa Kashima, and Takashi Tokino 8 Antimicrotubule Agents ...................................................... 177 Miguel A. Villalona-Calero, Larry Schaaf, and Robert Turowski 9 Kinesin Motor Inhibitors as Effective Anticancer Drugs.............................................................. 207 Vasiliki Sarli and Athanassios Giannis 10 Targeting the Spindle Checkpoint in Cancer Chemotherapy................................................................... 227 Jungseog Kang and Hongtao Yu

ix

x

Contents

11 Antiproliferation Inhibitors Targeting Aurora Kinases ...... 243 Kishore Shakalya and Daruka Mahadevan 12 Plks as Novel Targets for Cancer Drug Design ................. 271 Wei Dai, Yali Yang, and Ning Jiang 13 Do Histone Deacetylase Inhibitors Target Cell Cycle Checkpoints that Monitor Heterochromatin Structure?... 291 Brian Gabrielli, Frankie Stevens, and Heather Beamish Index .......................................................................................... 311

Contributors Joseph M. Ackermann, PhD • Laboratory of Molecular Oncology and Cell Cycle Regulation, University of Pennsylvania School of Medicine, Philadelphia, PA Heather Beamish, PhD • Lions Research Fellow, Cancer Biology Program, Centre for Immunology and Cancer Research, University of Queensland, Brisbane, Queensland, Australia Ryan P. Bender, PhD • Department of Biochemistry, Vanderbilt University School of Medicine, Nashville, TN Wesley A. Braden, MD • Department of Cell and Cancer Biology, University of Cincinnati, Cincinnati, OH Wei Dai, PhD • Department of Environmental Medicine, New York University School of Medicine, Tuxedo, NY Johann S. de Bono, MD, FRCP, MSc, PhD • Senior Lecturer and Consultant, Section of Medicine, Institute of Cancer Research, Royal Marsden Hospital, Drug Development Unit, Sutton, Surrey, UK Wafik S. El-Deiry, MD, PhD • Radiation Biology Program, Abramson Comprehensive Cancer Center, Associate Director for Physician-Scientist Training, Hematology-Oncology Division, University of Pennsylvania School of Medicine, Philadelphia, PA Brian Gabrielli, PhD • NHMRC Senior Research Fellow, Head, Cell Cycle Group Diamantina Institute for Cancer Immunology and Metabolic Medicine, University of Queensland, R Wing, Princess Alexandra Hospital, Brisbane, Queensland, Australia Athanassios Giannis, PhD, MD • Institute of Organic Chemistry, Leipzig University, Leipzig, Germany Ning Jiang, PhD • Department of Pathology, New York Medical College, Valhalla, NY Jungseog Kang, PhD • Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, TX Lisa Kashima, MS • Department of Molecular Biology, Cancer Research Institute, Sapporo Medical University, Sapporo, Japan Takumi Kawabe, MD, PhD • President and CEO, CanBas Co. Ltd., Makiya, Numazu, Japan Christian Klein, PhD • Pharma Research, Roche Diagnostics, Penzberg, Germany xi

xii

Contributors

Erik S. Knudsen, PhD • Department of Cell and Cancer Biology, University of Cincinnati, Cincinnati, OH Daruka Mahadevan, MD, PhD • Department of Medicine, Hematology/Oncology, Arizona Cancer Center, The University of Arizona College of Medicine, Tucson, AZ L. Rhoda Molife, MRCP, MSc, MD • Section of Medicine, Institute of Cancer Research, Royal Marsden Hospital, Drug Development Unit, Sutton, Surrey, UK Neil Osheroff, PhD • Professor of Biochemistry and Medicine, John G. Coniglio Chair in Biochemistry, Department of Biochemistry, Vanderbilt University School of Medicine, Nashville, TN Vasiliki Sarli, PhD • Institute for Organic Chemistry, University of Leipzig, Leipzig, Germany Larry Schaaf, PhD • Clinical Treatment Unit, The Ohio State University, Comprehensive Cancer Center, Columbus, OH Kishore Shakalya, BSc, MS • University of Arizona Cancer Center, Tucson, AZ Frankie Stevens, PhD • Cancer Biology Program, Centre for Immunology and Cancer Research, University of Queensland, Brisbane, Queensland, Australia Takashi Tokino, PhD • Department of Molecular Biology, Cancer Research Institute, Sapporo Medical University, Sapporo, Japan Minoru Toyota, MD, PhD • Department of Molecular Biology, Cancer Research Institute, Sapporo Medical University, Sapporo, Japan Robert Turowski, MBA, RPh • Departments of Internal Medicine and Pharmacology, The Ohio State University, Columbus, OH Lyubomir T. Vassilev, PhD • Discovery Oncology, Roche Research Center, Hoffmann-La Roche Inc., Nutley, NJ Miguel A. Villalona-Calero, MD, FACP • Associate Professor of Internal Medicine and Pharmacology, Division of Hematology and Oncology, The Ohio State University Medical Center, Columbus, OH Yali Yang, PhD • The Ohio State University Medical Center, Department of Environmental Medicine, New York University School of Medicine, Tuxedo, NY

Contributors

Timothy A. Yap, BSc, MBBS, MRCP • Section of Medicine, Institute of Cancer Research, Royal Marsden Hospital, Drug Development Unit, Sutton, Surrey, UK Hongtao Yu, PhD • Associate Professor, Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, TX

xiii

1

RB-Pathway Cell Cycle Control and Cancer Therapy

Erik S. Knudsen and Wesley A. Braden CONTENTS Dysregulation of the RB Pathway in cancer Cell Cycle Control Through the RB-Pathway Influence of the RB-Pathway Genotoxic Therapies Impact of RB Pathway on Antimetabolites Influence of RB on Antimicrotubule Agents Targeted Therapeutics and the Impact of RB Pathway Synopsis

Abstract The retinoblastoma tumor suppressor (RB) is a negative regulator of cellular proliferation that impacts multiple facets of cell cycle control. To this end, RB function is commonly abrogated in tumorigenesis, thereby contributing to the uncontrolled proliferation of From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

1

2

Knudsen and Braden

cancer cells. However, it is becoming increasingly clear that RB is not only involved in the etiology and progression of cancer, but also modifies the response to specific therapeutic modalities. Here we discuss the role of RB in cell cycle control as it relates to the response to specific classes of therapeutic agents. Key Words: Tumor suppressor; retinoblastoma; cyclins; chemotherapy; molecularly targeted therapeutics; cell cycle; E2F; cyclins

1. DYSREGULATION OF THE RB PATHWAY IN CANCER The retinoblastoma tumor suppressor protein, RB, is subject to functional inactivation in a multitude of different tumor types. The Rb gene was identified based on bialleleic inactivation in the pediatric eye tumor, retinoblastoma (1–3). Subsequent analyses demonstrated that loss of heterozygosity of the Rb gene or histochemical deficiency of RB protein occurs in a wide fraction of human cancers and is heterogeneous in nature, depending on the specific tumor type (4–10). In addition to direct effects on Rb gene expression, the RB protein is subject to post-translational modifications that have a significant impact on its function. For example, viral oncoproteins (E1A, E7, T-Ag) can bind to and directly inactivate RB. Such proteins function by a combination of sequestration and targeting RB for degradation as key facets of their transforming activity (11–13). Additionally, RB activity is compromised as a consequence of CDK-mediated phosphorylation (14–16). These phosphorylation events modify RB conformation and disrupt virtually all of the biochemical activities associated with the tumor suppressor (17). Therefore, it is perhaps not surprising that deregulated phosphorylation of RB is a key factor in human cancer (15,18). Specifically, deregulation of CDK-activity occurs through the overexpression of cyclin D1 or loss of the CDK4/6-inhibitor, p16ink4a, and is believed to trigger the aberrant phosphorylation and hence inactivation of RB (14–16,19–21). Importantly, loss of RB function is found in many tumor types but typically in a nonredundant manner, such that deregulation of RB phosphorylation (e.g., p16ink4a loss), or RB loss, are mutually exclusive events (19–22). Thus, these studies have hastened the need to understand how RB functions, and correspondingly if defining RB status in a particular tumor can be used as a determinant for efficacious therapeutic intervention.

Chapter 1 / RB-Pathway

3

2. CELL CYCLE CONTROL THROUGH THE RB-PATHWAY RB is a critical regulator of the G1/S transition of the cell cycle, which is responsive to CDK/cyclin activity. Conventionally, RB inhibits S-phase entry by binding to the E2F family of transcription factors and preventing transcription of genes required for DNA replication and productive passage through mitosis (23–26). When hypophosphorylated, RB is bound to the transactivation domain of multiple E2F proteins, eliciting transcriptional repression of E2F-regulated genes (27–29). To achieve promoter repression, RB recruits corepressors such as histone deacetylases (HDACs), histone methyltransferases, members of the SWI/SNF complex, and polycomb group proteins to E2F-regulated promoters (30–37). Upon mitogenic signaling, RB becomes hyperphosphorylated as a consequence of CDK-mediated phosphorylation, leading to release of E2F and transcription of downstream target genes (18). Through this action, it is generally believed that phosphorylation of RB by CDK/Cyclins is essential for entry into S-phase. Coordination between CDK/Cyclin kinase activity and E2F-transcription define the RB-pathway as a module that is functionally inactivated in most cancer (14–17,19–22,38). Antimitogenic signals, which can be broadly defined as those that impede cellular proliferation, often are dependent on RB to inhibit cell cycle progression (15,38). Such signals, are often targeted in existing cancer therapy or are under investigation in the context of new therapies (39,40). For example, DNA damage elicits cell cycle checkpoints that are dominant to the function of mitogenic signals (41), and DNA damaging agents represent a mainstay of current cancer therapy (42,43). Under conditions of such anti-mitogenic signals, RB phosphorylation is prevented, leading to subsequent reactivation of the protein (15,18,38). Strikingly, the mechanisms through which RB can become activated are quite varied, involving effects on both CDK/cyclin activity and potential dephosphorylation of RB by phosphatase activities (18,44,45). Such signaling pathways have led to a concerted analysis of how RB activation functions in discrete phases of the cell cycle. Additionally, RB status as a determinant of the response to each signaling pathway and impact on therapeutic signaling is currently being resolved.

4

Knudsen and Braden

2.1. The RB Pathway in S-Phase Control DNA replication is a very tightly regulated process. As cells exit mitosis and enter G1, origin recognition complexes (ORCs) are bound to sites of potential replication firing and recruit 2 proteins, cdc6 and cdt1, independently (46–50). Mini-chromosome maintenance proteins (MCM2-7), which are thought to function as the replicative helicases, are recruited by cdc6 and cdt1 to form the prereplicative complex (preRC) (48,51–53). As the cell progresses into S-phase, a multitude of other proteins necessary for DNA replication are subsequently recruited to the preRC, allowing for maturation of the replisome complex that facilitates duplication of the genome (48,49,54). The RB pathway is a critical molecular node that impinges upon the DNA replication machinery. More clearly, many of the genes necessary for DNA replication have been shown to be regulated by the E2F transcription factor (55,56). As such, loss or activation of RB activity significantly impacts RNA levels, protein levels and activity associated with discrete DNA replication factors. Several studies have dissected the impact of RB on replication control, and these studies have uniformly demonstrated that RB has the capacity to actively inhibit DNA replication (57–59). Functional analyses of the mechanism underlying this action have illustrated that RB has the capacity to disrupt PCNA chromatin association through a pathway associated with the down-regulation of CDK2 activity. Consistent with this finding, several laboratories have subsequently demonstrated that inhibition of CDK2 activity via distinct mechanisms, mediates similar influences on PCNA function (57,59,60). More recently, it was determined that p16ink4a, which signals upstream of RB to inhibit CDK4 activity, can inhibit MCM chromatin association in G1 (58). Perhaps even more surprising was that p16ink4a expression was able to inhibit total protein levels of cdc6 and cdt1, whereas RB was not (58). Although RB has no effect on replication control in G1, p16ink4a has no effect on replication in S-phase. This is most likely because of low CDK4 activity in S-phase, thus compromising p16ink4a signaling. These data suggest a fundamental difference by which RB and p16ink4a influence DNA replication and underscore that although these proteins exist within the same pathway, they have differential effects on cell cycle and replication control (61). In addition to these specific models of RB-mediated arrest, RB controls the expression of a wide spectrum of replication factors (e.g., MCM2, MCM5, cdc6, and PCNA) and thus, deficiency of RB has the capacity to dramatically influence the coordination of S-phase (57,62,63).


5

2.2. Involvement of RB Pathway in G2/M Control Akin to the processes involved in DNA replication, progression through mitosis is achieved via a number of discrete steps associated with mitotic entry and subsequent exit from mitosis into G1. Mitotic entry is dependent on the activation of CDK1/cyclin B complexes that initiate nuclear envelope breakdown and chromosome condensation (64,65). However, mitotic progression from metaphase is dependent on passage through the spindle checkpoint. This checkpoint stalls progression to anaphase, and ensures that all chromosomes are associated with the mitotic spindle (66–68). Once all chromosomes are bound by the spindle, specific signals coalesce to promote the activation of the anaphase promoting complex (APC) to mediate the destruction of multiple substrates and allow progression through anaphase, and ultimately exit from mitosis (69). The extent to which the RB-pathway contributes to G2/M progression is less clearly defined than is the role of RB in other phases of the cell cycle. Activation of RB by multiple means has not demonstrated a pronounced effect on the progression through mitosis, as cells fail to accumulate in G2 or any mitotic stages (70). However, as is the case with DNA replication, multiple critical factors that are required both for mitotic entry (e.g., Cdk1 and Cyclin B1) and exit (e.g., Cdc20, Plk1, and Mad2) are regulated by RB (56,71–73). Thus, one would predict that the RB pathway impinges on the regulation of mitosis. Consistent with this supposition, loss of RB has recently been associated with delayed mitotic progression occurring as a consequence of MAD2 upregulation (71).

3. INFLUENCE OF THE RB-PATHWAY GENOTOXIC THERAPIES The majority of cytotoxic regimens utilized in the treatment of cancer cause DNA damage (74). For example, cisplatin, cyclophosphamide and ionizing radiation directly damage DNA, whereas agents such as the topoisomerase poisons, irinotecan, and doxorubicin induce DNA damage as a result of indirect effects (75–79). Following such treatment, S-phase is slowed or halted to allow for adequate repair of the lesions before progressing into mitosis (80–83). Combined, these controls limit the segregation of damaged DNA to daughter cells. It has been shown by multiple groups that RB is required for the solicitation of cell cycle arrest in the presence of DNA damage (17,45, 84,85). Initially, it was observed that RB-deficient embryonic fibroblasts failed to undergo the cessation of DNA replication following

6

Knudsen and Braden

DNA damage (85,86). However, these cells still mount a G2/M checkpoint response, and as such fail to progress through mitosis and harbor a propensity to over-replicate their genomes (45,87). Using RNAi technology, RB was subsequently shown to play similar roles in response to cytotoxic therapy in tumor cells, whereas other components of the RB pathway (e.g., p16ink4a loss) did not influence the checkpoint response (84). The consequence of RB loss in both fibroblasts and tumor cell models is associated with increased sensitivity to DNA damaging agents (84,85,88). Thus, although RB loss enables checkpoint bypass, invariably this has been associated with enhanced cell killing. The mechanisms through which RB influences survival are not intrinsically clear, but could be dependent on additional secondary damage as a consequence of ongoing DNA replication in the presence of damage (89). Alternatively, it has been reported that RB loss deregulates the expression of many proapoptotic factors, thus shifting the balance toward cell death following DNA damage (90–92). Irrespective of the mechanism, these findings suggest that RB-deficient tumors may be particularly amenable to treatment with cytotoxic agents. However, an ongoing concern would be that tumors lacking checkpoint function would be genomically unstable, and thus although initial therapy may be quite effective, mutations afforded at the time of treatment could contribute to rendering the tumor more aggressive and ultimately therapy resistant.

4. IMPACT OF RB PATHWAY ON ANTIMETABOLITES Some of the most commonly utilized agents in the treatment of cancer are anti-metabolites that largely function by perturbing dNTP pools (93,94). For example, the thymidylate synthase inhibitors (5-fluorouracil, tomudex) are first line agents in the treatment of many cancers and have a pronounced deleterious effect on DNA replication (93). Interestingly, RB controls many facets of DNA replication, including the levels of many metabolic enzymes, such as thymidylate synthase, dihydrofolate reductase and ribonucleotide reductase subunits (60,95–99). As such, RB-deficient cells express highly elevated levels of these enzymes and RB has the capacity to dramatically alter dNTP pools (97). In cell culture models, RB loss enables ongoing DNA replication in the presence of such agents, suggesting that RB-deficiency negatively affects therapeutic response (97,98).


7

5. INFLUENCE OF RB ON ANTIMICROTUBULE AGENTS A number of chemotherapeutic agents function by perturbing microtubule dynamics and correspondingly the function of the mitotic spindle. Specifically, taxanes and vinca-alkaloid agent are utilized clinically and prevent appropriate progression through mitosis (100–104). A consequence of this protracted mitotic-deficit is cell death, making these compounds very effective cytotoxic agents. As discussed above, there is evidence that RB-status impacts the regulation of mitotic progression. Taxanes promote microtubule stability and have been shown to influence RB phosphorylation (105–107). However, vinca-alkaloids, which destabilize microtubules, are one class of compounds for which the absence of RB has little effect on cell cycle response (86). In contrast, RB deficiency facilitates ongoing DNA replication and changes in cell ploidy following treatment with nocodazole that also destabilizes microtubule arrays (108,109). Strikingly, there is little indication of how RB loss ultimately modifies sensitivity to these clinically relevant agents.

6. TARGETED THERAPEUTICS AND THE IMPACT OF RB PATHWAY Recently, there has been a dramatic initiative to define anti-cancer agents that function upon specific molecular targets. Such therapeutics currently encompass a wide-range of targets (e.g., steroid hormone receptors, growth factor signaling pathways, cell cycle machinery, and chromatin). Many of these agents function by antagonizing mitogenic signaling cascades and thus inhibiting proliferation of tumor cells. As such, it may be suspected that loss of RB would have a significant impact on the response elicited downstream of such agents. Here we will briefly discuss several targeted therapeutics for which the role of RB has been investigated:

6.1. Staurosporine and 7-Hydoxystaurosporine Staurosporine and related analogs where classically believed to function as general PKC inhibitors. However, such agents impact a diverse range of targets including cyclin dependent kinases and Chk1 (110). Studies have demonstrated that cells containing functional RB respond to staurosporine treatment, resulting in diminished CDK4 activity and G1 arrest (110). Additionally, RB is required for

8

Knudsen and Braden

staurosporine to elicit this G1 arrest (111). Similar data exists for 7-hydroxystaurosporine (UCN-01) implying RB-status is a key determinant of cell cycle response (112). In the case of UCN-01, RBdeficiency and corresponding cell cycle progression is associated with enhanced cell death (113). 7-Hydroxystaurosporine (UCN-01), which was originally identified as a protein kinase C selective inhibitor, is currently in clinical trials as an anticancer drug (114). It has been previously shown that UCN-01 induced preferential G1-phase accumulation in tumor cells and this effect was associated with the RB and its regulatory factors, such as CDK2 and CDK inhibitors p21Cip1 and p27Kip1 (115). However, recent evidence suggests UCN-01 arrests cells in G1 regardless of RB-status (113). Perhaps more striking was that UCN-01 induced apoptosis in RB-deficient cell lines, but not in RB-proficient cell lines (113). These observations suggest that G1-checkpoint function might be important for cell survival during UCN-01 treatment (116). Combined, the data implicate a functional RB pathway as a significant determinant of the sensitivity of tumor cells to UCN-01.

6.2. Geldanamycins Geldanamycin (GM) is an ansamycin antibiotic that functions to inhibit heat shock protein 90 (Hsp90) chaperone function and results in the subsequent degradation of important signaling molecules (e.g., cyclin D1). Although GM has been shown to diminish CDK2 and CDK4 activity resulting in G1 arrest in RB proficient backgrounds, GM had no effect on cell cycle kinetics in an RB negative background (117). Furthermore, release of RB negative cells from G2/M in the presence of GM continued through G1 and the subsequent cell cycle unperturbed (117). Thus, RB is necessary for effective GM treatment of human tumors.

6.3. CDK Inhibitors Flavopiridol, roscovitine, PD0332991 and other compounds in development inhibit CDK activity, preventing phosphorylation of various CDK target proteins and ultimately leading to cell cycle arrest (118). Although flavopiridol functions as a generic CDK inhibitor molecule, roscovitine can inhibit CDK1 and CDK2 function, and PD0332991 specifically inhibits CDK4 kinase activity. In xenograft models, these agents halt tumor growth and can reduce tumor volume (114,118,119). Although these results are promising, RB status has


9

been shown to play a critical role in response to these agents. Particularly, the growth suppressive functions of PD0332991 are entirely dependent on RB, such that RB-deficient tumor models fail to respond to treatment. Therefore, understanding the functional status of RB will be an absolute requirement for the effective usage of such agents (58).

7. SYNOPSIS Combined, these studies indicate that RB status is a determinant of a variety of therapeutic approaches utilized in the treatment of cancer. In certain instances, RB-deficiency is associated with increased sensitivity to specific agents. This is most clearly established in the case of specific genotoxic agents. However, in other instances, RB-deficiency has an apparent deleterious impact on response, as exemplified with therapies directed at CDK activity. These studies provide an important basis for further investigation of RB action on therapeutic response and for devising methodologies to most efficaciously treat cancer rationally. However, to build upon these basic findings will require a number of further advances that will facilitate the utilization of RB-status as a determinant of therapeutic response.

ACKNOWLEDGMENTS The authors regret any omissions in the preparation of this article and thank our colleagues for though-provoking discussion. Members of Karen Knudsen’s and Erik Knudsen’s laboratories assisted in the preparation and editing of the manuscript.

REFERENCES 1. Knudson AG, Jr. Mutation and cancer: statistical study of retinoblastoma. Proc Natl Acad Sci U S A 1971;68:820–823. 2. Cavenee WK, Dryja TP, Phillips RA, et al. Expression of recessive alleles by chromosomal mechanisms in retinoblastoma. Nature 1983;305:779–784. 3. Khidr L, Chen PL. RB, the conductor that orchestrates life, death and differentiation. Oncogene 2006;25:5210–5219. 4. Abraham R, Pagano F, Gomella LG, Baffa R. Chromosomal deletions in bladder cancer: shutting down pathways. Front Biosci 2007;12:826–838. 5. Backus HH, Van Groeningen CJ, Vos W, et al. Differential expression of cell cycle and apoptosis related proteins in colorectal mucosa, primary colon tumours, and liver metastases. J Clin Pathol 2002;55:206–211.

10

Knudsen and Braden

6. Friend SH, Bernards R, Rogelj S, et al. A human DNA segment with properties of the gene that predisposes to retinoblastoma and osteosarcoma. Nature 1986;323:643–646. 7. Lee WH, Bookstein R, Hong F, Young LJ, Shew JY, Lee EY. Human retinoblastoma susceptibility gene: cloning, identification, and sequence. Science 1987;235(4794):1394–1399. 8. Malumbres M, Barbacid M. To cycle or not to cycle: a critical decision in cancer. Nat Rev Cancer 2001;1:222–231. 9. Scambia G, Lovergine S, Masciullo V. RB family members as predictive and prognostic factors in human cancer. Oncogene 2006;25: 5302–5308. 10. Wikenheiser-Brokamp KA. Retinoblastoma regulatory pathway in lung cancer. Curr Mol Med 2006;6:783–793. 11. DeCaprio JA, Ludlow JW, Figge J, et al. SV40 large tumor antigen forms a specific complex with the product of the retinoblastoma susceptibility gene. Cell 1988;54:275–283. 12. Munger K, Werness BA, Dyson N, Phelps WC, Harlow E, Howley PM. Complex formation of human papillomavirus E7 proteins with the retinoblastoma tumor suppressor gene product. Embo J 1989;8: 4099–4105. 13. Whyte P, Buchkovich KJ, Horowitz JM, et al. Association between an oncogene and an anti-oncogene: the adenovirus E1A proteins bind to the retinoblastoma gene product. Nature 1988;334:124–129. 14. Sherr CJ. Cancer cell cycles. Science 1996;274(5293):1672–1677. 15. Wang JY, Knudsen ES, Welch PJ. The retinoblastoma tumor suppressor protein. Adv Cancer Res 1994;64:25–85. 16. Weinberg RA. The retinoblastoma protein and cell cycle control. Cell 1995;81:323–330. 17. Bartek J, Bartkova J, Lukas J. The retinoblastoma protein pathway in cell cycle control and cancer. Exp Cell Res 1997;237:1–6. 18. Mittnacht S. Control of pRB phosphorylation. Curr Opin Genet Dev 1998;8:21–27. 19. Bartkova J, Lukas J, Bartek J. Aberrations of the G1- and G1/Sregulating genes in human cancer. Prog Cell Cycle Res 1997;3: 211–220. 20. Kaelin WG, Jr. Alterations in G1/S cell-cycle control contributing to carcinogenesis. Ann N Y Acad Sci 1997;833:29–33. 21. Sherr CJ, McCormick F. The RB and p53 pathways in cancer. Cancer Cell 2002;2:103–112. 22. Nevins JR. The Rb/E2F pathway and cancer. Hum Mol Genet 2001; 10:699–703. 23. Dyson N. The regulation of E2F by pRB-family proteins. Genes Dev 1998;12:2245–2262.


11

24. Helin K, Lees JA, Vidal M, Dyson N, Harlow E, Fattaey A. A cDNA encoding a pRB-binding protein with properties of the transcription factor E2F. Cell 1992;70:337–350. 25. Kaelin WG, Jr., Krek W, Sellers WR, et al. Expression cloning of a cDNA encoding a retinoblastoma-binding protein with E2F-like properties. Cell 1992;70:351–364. 26. Nevins JR, Chellappan SP, Mudryj M, et al. E2F transcription factor is a target for the RB protein and the cyclin A protein. Cold Spring Harb Symp Quant Biol 1991;56:157–162. 27. Flemington EK, Speck SH, Kaelin WG, Jr. E2F-1-mediated transactivation is inhibited by complex formation with the retinoblastoma susceptibility gene product. Proc Natl Acad Sci U S A 1993;90:6914–6918. 28. Helin K, Harlow E, Fattaey A. Inhibition of E2F-1 transactivation by direct binding of the retinoblastoma protein. Mol Cell Biol 1993;13:6501–6508. 29. Weintraub SJ, Prater CA, Dean DC. Retinoblastoma protein switches the E2F site from positive to negative element. Nature 1992;358:259–261. 30. Brehm A, Miska EA, McCance DJ, Reid JL, Bannister AJ, Kouzarides T. Retinoblastoma protein recruits histone deacetylase to repress transcription. Nature 1998;391:597–601. 31. Dahiya A, Wong S, Gonzalo S, Gavin M, Dean DC. Linking the Rb and polycomb pathways. Mol Cell 2001;8:557–569. 32. Dunaief JL, Strober BE, Guha S, et al. The retinoblastoma protein and BRG1 form a complex and cooperate to induce cell cycle arrest. Cell 1994;79:119–30. 33. Nielsen SJ, Schneider R, Bauer UM, et al. Rb targets histone H3 methylation and HP1 to promoters. Nature 2001;412:561–565. 34. Strobeck MW, Knudsen KE, Fribourg AF, et al. BRG-1 is required for RB-mediated cell cycle arrest. Proc Natl Acad Sci U S A 2000;97: 7748–7753. 35. Strober BE, Dunaief JL, Guha, Goff SP. Functional interactions between the hBRM/hBRG1 transcriptional activators and the pRB family of proteins. Mol Cell Biol 1996;16:1576–1583. 36. Weintraub SJ, Chow KN, Luo RX, Zhang SH, He S, Dean DC. Mechanism of active transcriptional repression by the retinoblastoma protein. Nature 1995;375:812–815. 37. Zhang HS, Gavin M, Dahiya A, et al. Exit from G1 and S phase of the cell cycle is regulated by repressor complexes containing HDAC-RbhSWI/SNF and Rb-hSWI/SNF. Cell 2000;101:79–89. 38. Cobrinik D. Pocket proteins and cell cycle control. Oncogene 2005;24:2796–2809. 39. Collins I, Garrett MD. Targeting the cell division cycle in cancer: CDK and cell cycle checkpoint kinase inhibitors. Curr Opin Pharmacol 2005;5:366–373.

12

Knudsen and Braden

40. Malumbres M. Preclinical models for cell cycle-targeted therapies. Adv Exp Med Biol 2006;587:139–147. 41. Kao J, Rosenstein BS, Peters S, Milano MT, Kron SJ. Cellular response to DNA damage. Ann N Y Acad Sci 2005;1066:243–258. 42. Sanchez-Perez I. DNA repair inhibitors in cancer treatment. Clin Transl Oncol 2006;8:642–646. 43. Madhusudan S, Hickson ID. DNA repair inhibition: a selective tumour targeting strategy. Trends Mol Med 2005;11:503–511. 44. Lundberg AS, Weinberg RA. Functional inactivation of the retinoblastoma protein requires sequential modification by at least two distinct cyclin-cdk complexes. Mol Cell Biol 1998;18:753761. 45. Avni D, Yang H, Martelli F, et al. Active localization of the retinoblastoma protein in chromatin and its response to S phase DNA damage. Mol Cell 2003;12(3):735–746. 46. Bell SP, Dutta A. DNA replication in eukaryotic cells. Annu Rev Biochem 2002;71:333–374. 47. Blow JJ, Hodgson B. Replication licensing–defining the proliferative state? Trends Cell Biol 2002;12:72–78. 48. Diffley JF, Labib K. The chromosome replication cycle. J Cell Sci 2002;115:869–872. 49. Dutta A, Bell SP. Initiation of DNA replication in eukaryotic cells. Annu Rev Cell Dev Biol 1997;13:293–332. 50. Stillman B. Origin recognition and the chromosome cycle. FEBS Lett 2005;579:877–884. 51. Dimitrova DS, Prokhorova TA, Blow JJ, Todorov IT, Gilbert DM. Mammalian nuclei become licensed for DNA replication during late telophase. J Cell Sci 2002;115:51–59. 52. Mendez J, Stillman B. Chromatin association of human origin recognition complex, cdc6, and minichromosome maintenance proteins during the cell cycle: assembly of prereplication complexes in late mitosis. Mol Cell Biol 2000;20:8602–8612. 53. Okuno Y, McNairn AJ, den Elzen N, Pines J, Gilbert DM. Stability, chromatin association and functional activity of mammalian pre-replication complex proteins during the cell cycle. Embo J 2001;20:4263–4277. 54. Diffley JF. Regulation of early events in chromosome replication. Curr Biol 2004;14:R778–786. 55. Bosco G, Du W, Orr-Weaver TL. DNA replication control through interaction of E2F-RB and the origin recognition complex. Nat Cell Biol 2001;3:289–295. 56. Markey MP, Angus SP, Strobeck MW, et al. Unbiased analysis of RBmediated transcriptional repression identifies novel targets and distinctions from E2F action. Cancer Res 2002;62:6587–6597.


13

57. Angus SP, Mayhew CN, Solomon DA, et al. RB reversibly inhibits DNA replication via two temporally distinct mechanisms. Mol Cell Biol 2004;24:5404–5420. 58. Braden WA, Lenihan JM, Lan Z, et al. Distinct action of the retinoblastoma pathway on the DNA replication machinery defines specific roles for cyclin-dependent kinase complexes in prereplication complex assembly and S-phase progression. Mol Cell Biol 2006;26:7667–7681. 59. Sever-Chroneos Z, Angus SP, Fribourg AF, et al. Retinoblastoma tumor suppressor protein signals through inhibition of cyclin-dependent kinase 2 activity to disrupt PCNA function in S phase. Mol Cell Biol 2001;21:4032–4045. 60. Naderi S, Wang JY, Chen TT, Gutzkow KB, Blomhoff HK. cAMPmediated inhibition of DNA replication and S phase progression: involvement of Rb, p21Cip1, and PCNA. Mol Biol Cell 2005;16: 1527–1542. 61. Tort F, Bartkova J, Sehested M, Orntoft T, Lukas J, Bartek J. Retinoblastoma pathway defects show differential ability to activate the constitutive DNA damage response in human tumorigenesis. Cancer Res 2006;66:10258–10263. 62. Markey M, H. Siddiqui, and E.S. Knudsen. Geminin is targeted for repression by the retinoblastoma tumor suppressor pathway through intragenic E2F sites. J Biol Chem 2002;279:29255–29262. 63. Vernell R, Helin K, Muller H. Identification of target genes of the p16INK4A-pRB-E2F pathway. J Biol Chem 2003;278:46124–46137. 64. Arellano M, Moreno S. Regulation of CDK/cyclin complexes during the cell cycle. Int J Biochem Cell Biol 1997;29:559–573. 65. King RW, Jackson PK, Kirschner MW. Mitosis in transition. Cell 1994;79:563–571. 66. Doncic A, Ben-Jacob E, Barkai N. Evaluating putative mechanisms of the mitotic spindle checkpoint. Proc Natl Acad Sci U S A 2005;102: 6332–6337. 67. Nigg EA. Origins and consequences of centrosome aberrations in human cancers. Int J Cancer 2006;119:2717–27123. 68. Georgi AB, Stukenberg PT, Kirschner MW. Timing of events in mitosis. Curr Biol 2002;12:105–114. 69. Morgan DO. Regulation of the APC and the exit from mitosis. Nat Cell Biol 1999;1:E47–53. 70. Angus SP, Fribourg AF, Markey MP, et al. Active RB elicits late G1/S inhibition. Exp Cell Res 2002;276:201–213. 71. Hernando E, Nahle Z, Juan G, et al. Rb inactivation promotes genomic instability by uncoupling cell cycle progression from mitotic control. Nature 2004;430:797–802.

14

Knudsen and Braden

72. Gunawardena RW, Siddiqui H, Solomon DA, et al. Hierarchical requirement of SWI/SNF in retinoblastoma tumor suppressor-mediated repression of Plk1. J Biol Chem 2004;279:29278–29285. 73. Jackson MW, Agarwal MK, Yang J, et al. p130/p107/p105Rb-dependent transcriptional repression during DNA-damage-induced cell-cycle exit at G2. J Cell Sci 2005;118:1821–1832. 74. Gewirtz DA. Growth arrest and cell death in the breast tumor cell in response to ionizing radiation and chemotherapeutic agents which induce DNA damage. Breast Cancer Res Treat 2000;62:223–235. 75. Pommier Y. Topoisomerase I inhibitors: camptothecins and beyond. Nat Rev Cancer 2006;6:789–802. 76. Bowen JM, Gibson RJ, Stringer AM, et al. Role of p53 in irinotecaninduced intestinal cell death and mucosal damage. Anticancer Drugs 2007;18:197–210. 77. Kawato Y, Aonuma M, Hirota Y, Kuga H, Sato K. Intracellular roles of SN-38, a metabolite of the camptothecin derivative CPT-11, in the antitumor effect of CPT-11. Cancer Res 1991;51:4187–4191. 78. Cummings J, Anderson L, Willmott N, Smyth JF. The molecular pharmacology of doxorubicin in vivo. Eur J Cancer 1991;27: 532–535. 79. Cutts SM, Nudelman A, Rephaeli A, Phillips DR. The power and potential of doxorubicin-DNA adducts. IUBMB Life 2005;57:73–81. 80. Jeggo PA, Lobrich M. Contribution of DNA repair and cell cycle checkpoint arrest to the maintenance of genomic stability. DNA Repair (Amst) 2006;5:1192–1198. 81. O’Connor PM, Fan S. DNA damage checkpoints: implications for cancer therapy. Prog Cell Cycle Res 1996;2:165–173. 82. Searle JS, Sanchez Y. Stopped for repairs: a new role for nutrient sensing pathways? Cell Cycle 2004;3:865–868. 83. O’Connell MJ, Cimprich KA. G2 damage checkpoints: what is the turnon? J Cell Sci 2005;118:1–6. 84. Bosco EE, Wang Y, Xu H, et al. The retinoblastoma tumor suppressor modifies the therapeutic response of breast cancer. J Clin Invest 2007;117:218–228. 85. Knudsen KE, Booth D, Naderi S, et al. RB-dependent S-phase response to DNA damage. Mol Cell Biol 2000;20:7751–7763. 86. Harrington EA, Bruce JL, Harlow E, Dyson N. pRB plays an essential role in cell cycle arrest induced by DNA damage. Proc Natl Acad Sci U S A 1998;95:11945–11950. 87. Lan Z, Sever-Chroneos Z, Strobeck MW, et al. DNA damage invokes mismatch repair-dependent cyclin D1 attenuation and retinoblastoma signaling pathways to inhibit CDK2. J Biol Chem 2002;277:8372–8381.


15

88. Wang JY, Naderi S, Chen TT. Role of retinoblastoma tumor suppressor protein in DNA damage response. Acta Oncol 2001;40:689–695. 89. Bosco EE, Mayhew CN, Hennigan RF, Sage J, Jacks T, Knudsen ES. RB signaling prevents replication-dependent DNA double-strand breaks following genotoxic insult. Nucleic Acids Res 2004;32:25–34. 90. Guo Z, Yikang S, Yoshida H, Mak TW, Zacksenhaus E. Inactivation of the retinoblastoma tumor suppressor induces apoptosis protease-activating factor-1 dependent and independent apoptotic pathways during embryogenesis. Cancer Res 2001;61:8395–8400. 91. Chau BN, Wang JY. Coordinated regulation of life and death by RB. Nat Rev Cancer 2003;3:130–138. 92. Nahle Z, Polakoff J, Davuluri RV, et al. Direct coupling of the cell cycle and cell death machinery by E2F. Nat Cell Biol 2002;4:859–864. 93. Longley DB, Harkin DP, Johnston PG. 5-fluorouracil: mechanisms of action and clinical strategies. Nat Rev Cancer 2003;3:330–338. 94. Schweitzer BI, Dicker AP, Bertino JR. Dihydrofolate reductase as a therapeutic target. Faseb J 1990;4:2441–2452. 95. DeGregori J, Kowalik T, Nevins JR. Cellular targets for activation by the E2F1 transcription factor include DNA synthesis- and G1/S-regulatory genes. Mol Cell Biol 1995;15:4215–4224. 96. Noe V, Alemany C, Chasin LA, Ciudad CJ. Retinoblastoma protein associates with SP1 and activates the hamster dihydrofolate reductase promoter. Oncogene 1998;16:1931–1938. 97. Angus SP, Wheeler LJ, Ranmal SA, et al. Retinoblastoma tumor suppressor targets dNTP metabolism to regulate DNA replication. J Biol Chem 2002;277:44376–44384. 98. Almasan A, Yin Y, Kelly RE, et al. Deficiency of retinoblastoma protein leads to inappropriate S-phase entry, activation of E2F-responsive genes, and apoptosis. Proc Natl Acad Sci U S A 1995;92:5436–5440. 99. Siddiqui H, Solomon DA, Gunawardena RW, Wang Y, Knudsen ES. Histone deacetylation of RB-responsive promoters: requisite for specific gene repression but dispensable for cell cycle inhibition. Mol Cell Biol 2003;23:7719–7731. 100. Himes RH, Kersey RN, Heller-Bettinger I, Samson FE. Action of the vinca alkaloids vincristine, vinblastine, and desacetyl vinblastine amide on microtubules in vitro. Cancer Res 1976;36:3798–3802. 101. Adjei AA, Rowinsky EK. Novel anticancer agents in clinical development. Cancer Biol Ther 2003;2:S5–15. 102. Rowinsky EK. The development and clinical utility of the taxane class of antimicrotubule chemotherapy agents. Annu Rev Med 1997;48: 353–374. 103. Geney R, Chen J, Ojima I. Recent advances in the new generation taxane anticancer agents. Med Chem 2005;1:125–139.

16

Knudsen and Braden

104. Bensch KG, Malawista SE. Microtubule crystals: a new biophysical phenomenon induced by Vinca alkaloids. Nature 1968;218: 1176–1177. 105. Lanzi C, Cassinelli G, Cuccuru G, et al. Cell cycle checkpoint efficiency and cellular response to paclitaxel in prostate cancer cells. Prostate 2001;48:254–264. 106. Morita M, Suyama H, Igishi T, et al. Dexamethasone inhibits paclitaxel-induced cytotoxic activity through retinoblastoma protein dephosphorylation in non-small cell lung cancer cells. Int J Oncol 2007;30:187–192. 107. Motwani M, Rizzo C, Sirotnak F, She Y, Schwartz GK. Flavopiridol enhances the effect of docetaxel in vitro and in vivo in human gastric cancer cells. Mol Cancer Ther 2003;2:549–555. 108. Iovino F, Lentini L, Amato A, Di Leonardo A. RB acute loss induces centrosome amplification and aneuploidy in murine primary fibroblasts. Mol Cancer 2006;5:38. 109. Khan SH, Wahl GM. p53 and pRb prevent rereplication in response to microtubule inhibitors by mediating a reversible G1 arrest. Cancer Res 1998;58:396–401. 110. McGahren-Murray M, Terry NH, Keyomarsi K. The Differential Staurosporine-Mediated G1 Arrest in Normal versus Tumor Cells Is Dependent on the Retinoblastoma Protein. Cancer Res 2006;66: 9744–9753. 111. Zhou W, Lin Y, Wersto R, Chrest J, Gabrielson E. Staurosporineinduced G(1) arrest in cancer cells depends on an intact pRB but is independent of p16 status. Cancer Lett 2002;183:103–107. 112. Orr MS, Reinhold W, Yu L, Schreiber-Agus N, O’Connor PM. An important role for the retinoblastoma protein in staurosporine-induced G1 arrest in murine embryonic fibroblasts. J Biol Chem 1998;273: 3803–3807. 113. Akiyama T, Sugiyama K, Shimizu M, Tamaoki T, Akinaga S. G1checkpoint function including a cyclin-dependent kinase 2 regulatory pathway as potential determinant of 7-hydroxystaurosporine (UCN01)-induced apoptosis and G1-phase accumulation. Jpn J Cancer Res 1999;90:1364–1372. 114. Senderowicz AM. Novel small molecule cyclin-dependent kinases modulators in human clinical trials. Cancer Biol Ther 2003;2:S84–95. 115. Akiyama T, Yoshida T, Tsujita T, et al. G1 phase accumulation induced by UCN-01 is associated with dephosphorylation of Rb and CDK2 proteins as well as induction of CDK inhibitor p21/Cip1/WAF1/Sdi1 in p53-mutated human epidermoid carcinoma A431 cells. Cancer Res 1997;57:1495–1501.


17

116. Mack PC, Gandara DR, Bowen C, et al. RB status as a determinant of response to UCN-01 in non-small cell lung carcinoma. Clin Cancer Res 1999;5:2596–604. 117. Srethapakdi M, Liu F, Tavorath R, Rosen N. Inhibition of Hsp90 function by ansamycins causes retinoblastoma gene product-dependent G1 arrest. Cancer Res 2000;60:3940–3946. 118. Lee YM, Sicinski P. Targeting cyclins and cyclin-dependent kinases in cancer: lessons from mice, hopes for therapeutic applications in human. Cell Cycle 2006;5:2110–2114. 119. Fry DW, Harvey PJ, Keller PR, et al. Specific inhibition of cyclindependent kinase 4/6 by PD 0332991 and associated antitumor activity in human tumor xenografts. Mol Cancer Ther 2004;3:1427–1438.

2

Targeting the p53/MDM2 Pathway for Cancer Therapy Christian Klein and Lyubomir T. Vassilev CONTENTS Introduction The p53 Tumor Suppressor Protein Reactivation of Mutant p53 in Tumors Inhibition of the p53-MDM2 Interaction Inhibition of MDM2 E3 Ligase Activity Modulation of p53 Activity for Protection of Normal Tissues During Chemotherapy

Key Words: 53; MDM2; tumor suppressor; genome guadian; protein-protein interaction

1. INTRODUCTION Nearly 40,000 research and review articles published since 1980 have elevated the tumor suppressor p53 to the rank of the most studied protein in cancer research (1–5). However, the translation of this From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

19

20

Klein and Vassilev

knowledge into therapies has been lagging. This is mainly because of the fact that p53 is a tumor suppressor and its function is frequently lost in human tumors. Therefore, p53 has to be activated, rather than inhibited, a task considerably more challenging in the pharmacological setting. Recently, several major drug discovery advances in the exploration of the p53 pathway raised the hopes that the powerful growth suppressive and proapoptotic activity of p53 could be harnessed for the benefit of the cancer patient. In this chapter, we summarize the current state of knowledge about p53 function and examine novel therapeutic approaches for targeting the p53 pathway focusing primarily on peptides and small molecules. Gene therapy and antisense approaches for targeting p53 have been reviewed earlier (6,7).

2. THE p53 TUMOR SUPPRESSOR PROTEIN The p53 protein has been described for the first time in 1979 as an oncoprotein. However, in the following years, it has been recognized that only mutated p53 acts as oncoprotein whereas the normal wild-type protein possess the properties of a tumor suppressor (8,9). Lane first proposed that p53 functions as a “guardian of the genome” protecting mammalian cells from the consequences of DNA damage (10). Later this protective role has been expanded to include a wide variety of oncogenic stresses thus establishing p53 as the “cellular gatekeeper” (11). The p53 tumor suppressor serves as a central node in a complex signal transduction network known as the p53 pathway, which has evolved as a major defense against cancer. This pathway recognizes diverse forms of oncogenic stress within the cellular environment and translates them into appropriate cellular responses to minimize tumorigenic consequences. In response to stress, p53 halts cell proliferation to prevent the propagation of DNA damage and/or directly helps in its repair. If the damage is too severe and beyond repair, p53 can induce programmed cell death (apoptosis) or senescence as a last resort to avoid possible malignant transformation. Because of its critical antitumor role, p53 is the most frequent target of genetic alterations in cancer. It is inactivated by mutations and/or deletions in half of all human tumors (12–14). Somatic mutations of p53 are the genetic basis of the Li-Fraumeni Syndrome, that is characterized by a high familiar tumor incidence (15,16). In tumors without p53 mutation, the function of the p53 pathway is frequently attenuated or disabled by other mechanisms, e.g., nuclear exclusion, interaction with viral proteins such as E1B and E6, overexpression of its negative regulator MDM2, or inactivation of the tumor suppressor

Chapter 2 / Targeting the p53/MDM2 Pathway for Cancer Therapy

21

protein p14ARF (17). If one takes into account these additional defects in p53 regulation, the p53 pathway is likely altered in more than 70% of all human tumors. Under physiological conditions, p53 has a short half-life and is present in cells at low concentrations. This is essential because high levels of p53 are detrimental to their ability to proliferate. In response to cellular stress (e.g., DNA damage, hypoxia, oncogene activation, etc.), p53 is activated by a post-translational mechanism that increases its stability. p53 Rapidly accumulates in cell nuclei and, as a potent transcription factor, activates or inhibits multiple genes (18) involved in cell cycle control, apoptosis, senescence, and angiogenesis (11,19–21). Functions independent of its transcriptional activity have also been described (22,23). For a detailed review of post-translational mechanisms regulating p53 see (24). p53 plays a central role in tumor suppression but it is not essential for mouse development and normal physiology. p53 deficient mice are normal but develop multiple tumors very early in life (25,26). Transgenic mice with elevated p53 activity have increased resistance to spontaneous tumors but display early signs of aging (27). It has been speculated that prolonged activation of p53 by therapeutic means might lead to premature ageing (28). However, other mouse experiments in which p53 levels are increased by genetic means have not shown any signs of premature aging (29). The discovery of p53 in Drosophila melanogaster (30–32) and Clostridium elegans (CEP-1) (33,34) proved that tumor suppression mechanisms have evolved early in the evolution of multicellular organisms. In addition, 2 close p53 homologues, p63 and p73, have been identified with functions during embryogenesis, differentiation, and possibly tumorigenesis (35–38). The p53 tumor suppressor is made up of different functional domains: i) an N-terminal flexible transactivation domain (1-50) that interacts with proteins of the transcriptional machinery and its functional antagonist MDM2 ii) a proline-rich domain (63-97), iii) an evolutionary highly conserved central DNA binding domain (98), iv) a tetramerization domain (320–356), and v) a C-terminal regulatory domain (363–393) (39,40). The majority of p53 mutations in human tumors are missense mutations within the DNA binding domain, most frequently at 6 mutation hot-spots: R175, G245, R248, R249, R273, and R282 (12,13) which underlines the importance of DNA binding for the physiological function of p53. As a transcription factor, p53 binds at the promoter region of target genes to a double-stranded DNA sequence composed of 2 consecutive palindromic consensus sites

22

Klein and Vassilev

5’-PuPuPuC(A/T)(T/A)GPyPyPy-3’) where Pu represent purine and Py represents pyrimidine bases, respectively (41). It has been well established that p53 binds DNA as a tetramer but despite the extensive efforts the crystal structure of tetrameric p53 has not been determined yet. The difficulties are likely because of the fact that p53 is intrinsically unstructured in large portions of the molecule (42,43). However, several crystal structure and NMR spectroscopy studies of isolated p53 domains have been published. X-ray, NMR, and modeling data have helped to gain insight into p53 structure and function. Just recently, the structure of full-length p53 has been reconstructed using electron microscopy combined with crystallography (44). The crystal structure of the p53 DNA binding domain in complex with its consensus DNA has helped to clarify the consequences of hot-spot p53 mutations (45). The DNA binding domain of p53 is made of a 4-stranded and a 5-stranded antiparallel sheet, a loop-sheet-helix (LSH) motif, comprising loop L1 (residues 113–123), a 3-stranded sheet and helix H2 (residues 278–286). The loop L2 (residues 164–194), which is interrupted by the short helix H1, and the loop L3 (residues 237–250) are stabilized by a single coordinated zinc atom (45,46). The hot-spot residues R273 within the H2 helix of the LSH motif and R248 within the L3 loop contact directly the major and minor groove of bound DNA. The other hot-spot mutations are involved in the stabilization of the surrounding structure via a hydrogen bond network. Thus, there are 2 major classes of mutations preventing DNA binding: (i) Mutations involved in direct DNA contact and (ii) mutations destabilizing the structural integrity of the DNA binding region (R249S, G245S) and/or causing a globally denatured p53 structure (R175H, C242S) (47–49). The major classes of mutations can be distinguished by 2 antibodies, PAb1620 and PAb240. PAb1620, binds preferentially to the wild-type conformation of p53 DBD including solvent-exposed “DNA contact mutants,” which do not alter its native conformation. PAb240 binds specifically to non-natively folded “structural mutants” of the p53 DBD (50,51). Furthermore, allosteric binding sites on p53 have been identified by phage display (52) and epitopes specific for different conformational states of p53 can be distinguished (53,54). Wild-type p53 exhibits only moderate thermodynamic stability and can easily lose its native conformation and DNA binding activity upon relatively small changes such as point mutations or increase of temperature (48,55). NMR spectroscopic studies and molecular dynamics simulations have revealed that local conformational changes result in the thermodynamic destabilization of these mutants (47,56,57).


23

Based on a detailed thermodynamic and biophysical analysis of p53 mutants, Fersht and co-workers have distinguished 3 folding phenotypes: i) DNA contact mutations that have little or no effect on folding; ii) mutations that disrupt local structure, and iii) mutations that cause denaturation. Further division of these molecular phenotypes into DNA-binding competent and incompetent states defines 1 wild-type and 5 distinct classes of p53 mutation (47–49). Recent biophysical studies using full length p53 instead of the p53 DNA-binding domain have confirmed that common cancer mutations affect p53 stability and DNA binding properties and that thermodynamic stability of tetrameric p53 is dictated by its DNA binding domain (58).

3. REACTIVATION OF MUTANT p53 IN TUMORS Restoration of wild-type p53 conformation and function to tumors that have lost p53 activity because of mutation has been considered one of the most attractive strategies for cancer therapy. It is believed that the p53 pathway of cancer cells with mutant p53 is otherwise intact and by reactivation of p53 one could take full advantage of its antitumor potential. Several experimental studies using macromolecular tools to modulate mutant p53 have validated this approach in vitro (59). One straightforward p53 activation approach is the introduction of wild-type p53 in tumors with mutated p53 by gene therapy. The proofof-concept for this therapeutic strategy has been well established in tissue culture and animal models of human cancer. However, technical difficulties and severe side effects associated with the methods of gene introduction have delayed the clinical validation of p53 gene therapy. In October of 2003, the State Food and Drug Administration of China (SFDA) approved a recombinant adenoviral p53 gene therapy with the generic name Gendicine (Shenzen SiBiono GenTech, Shenzen, China) for the treatment of head and neck squamous cell carcinoma. Other p53-based gene therapies are currently in clinical phase II/III trials (e.g., Introgen, Austin, TX) (60,61). The coming years will show whether restoration of p53 activity by gene therapy is viable in the clinical setting. Given the current concerns regarding the application of gene therapy in humans there remains a strong demand for smallmolecule drugs that can activate p53 in human tumors. There are 2 principle mechanisms to achieve activation of mutant p53 by small molecules: The largest proportion of p53 structural mutants can adopt both mutant and wild-type conformation. Wild-type p53 conformation could be rescued by small molecules that bind to

24

Klein and Vassilev

wild-type conformation and thus shift p53 conformational equilibrium in favor of the wild-type. For a small proportion of DNA contact mutants or more heavily destabilized p53 mutants, however, small molecules would need to recognize a specific mutant conformation and induce wild-type-like conformation and function e.g., by forming additional stabilizing interactions (48). The small molecule should prevent aggregation and maintain mutant p53 in a folded state so that p53 can regulate its target genes and/or bind to its interaction partners (62). These requirements make the identification of small molecules that can restore p53 wild-type conformation highly challenging. Nevertheless, early proof-of-concept studies have shown that wild-type p53 conformation can be rescued by several means. Based on structural information, Wieczorek at al. have introduced basic residues into the DNA binding domain to establish new contacts between p53 and the DNA phosphate backbone. Replacement of Thr 284 with Arg substantially increased DNA binding affinity and activated transactivation and tumor suppressor functions in 3 out of 7 most common p53 mutants (63). A second-site suppressor mutant in yeast have been identified that can overcome the deleterious effects of common p53 cancer mutations in cells (64). Analyzing the mechanism of second-site suppressor mutants by double-mutant cycles. Nikolova et al. have been able to identify specific second-site mutants and have concluded that the function of p53 mutants could be restored by small molecules that are capable of stabilizing the native p53 structure (65). Finally, semirational protein design (66) and directed evolution (67,68) have helped to generate p53 mutants with wild-type p53 function and increased thermodynamic stability. Nikolova et al. have designed a superstable quadruple mutant M133L/V203A/N239Y/N268D of the p53 DNAbinding domain with second-site suppressor mutations N239Y and N268D that have specifically restored the activity and stability of several oncogenic mutants (66). The crystal structure of this superstable quadruple mutant illustrated how an increase in rigidity results in elevated thermostability and explained how N268D and N239Y rescue some of the common cancer mutants (69). It has been shown that addition of consensus DNA and heparin resulted in a thermodynamic stabilization of the p53 DNA-binding domain (66,70). The chemical chaperones glycerol and TMAO and also aminothiol WR1065 have been shown to stabilize the active wild-type conformation of temperature-sensitive p53 mutants in vivo (71–76). In summary, these studies support the idea that wild-type p53 conformation of mutant p53 can be stabilized by a second-site suppressor


25

mutation and suggested that it might be amenable to stabilization by peptides and small molecules (77). The first published small molecule claimed to restore wild-type conformation to mutant p53 is CP-31398 (78) (Fig. 2A). Foster et al. (1999) have screened a large compound library searching for small molecules that could stabilize the PAb 1620-epitope of wild-type p53 against thermal stress/denaturation. They have used a recombinant p53 DNA-binding domain and an ELISA-based assay (78). CP-31398, the most potent representative of the identified compound classes, has been found to be active in cellular assays. CP-31398 stabilized wildtype p53 conformation, caused the accumulation of transcriptionally active p53 in cells with mutant p53, and inhibited the growth of tumor xenograft with mutant p53 in nude mice (78). Medicinal chemistry efforts on lead compounds and its derivatives have allowed to derive a structure-activity-relationship, but has not improved substantially the potency (78). In addition, independent laboratories have been unable to confirm the mode-of-action of CP-31398. Several groups showed that CP-31398 induces a specific p53 response and can induce apoptosis in tumor cells, (79–84) but others have not found evidence that this effect is dependent on the presence of mutant p53 (85,86). Based on the structure activity relationship of the described compounds that is characterized by a hydrophobic (R1) and a hydrophilic (R2) group connected via a defined linker (L), it has been hypothesized that these analogs might interact with 2 different binding sites on the surface of p53 and thus could stabilize the wild-type p53 conformation. However, since its discovery no evidence for direct binding of CP-31398 with p53 or the p53 DNA-binding domain has been reported. In contrast, it has been found that CP-31398 does not stabilize p53 thermodynamically but intercalates into DNA. In addition, it has been reported that CP31398 exhibits unspecific toxicity and is unable to reactivate mutant p53 protein (86). Taken together, the mode-of-action of CP-31398 remains controversial and requires further investigation. PRIMA-1 (for p53 reactivation and induction of massive apoptosis) was identified from the small NCI diversity set chemical library of ca. 2,000 cpds in a cellular proliferation assay screen for novel p53 reactivators (87) (Fig. 2A). Compounds have been tested for their ability to inhibit the growth of Saos-2 tumor cell lines transfected with the p53 mutant His-273. PRIMA-1 has been found to restore p53-dependent apoptosis in several tumor cell lines with different p53 status and to restore wild-type p53 conformation and DNAbinding activity in cell extracts as well as in living cells. In contrast to CP31398, PRIMA-1 appears to be able to rescue wild-type p53

26

Klein and Vassilev

conformation of both DNA contact and structural p53 mutants and inhibits tumor growth in xenograft models (87). A statistical analysis of the effect of PRIMA-1 on cells of the NCI tumor cell panel have shown a preferential inhibition of cells with mutant rather than wild-type p53 whereas 44 known antitumor agents have not shown such preference (88). Recently, it has been shown that a methylated form of PRIMA-1MET is more active than PRIMA-1 and synergizes with chemotherapeutic agents (e.g., cisplatin) (89) in cellular assays. In the meantime, a maleimide analog with similar activity originating from the same screening campaign, called MIRA-1 (for mutant p53-dependent induction of rapid apoptosis) (Fig. 2A), has also been published. MIRA-1 has been able to reactivate DNA binding properties of mutant p53 and to preserve its active conformation in vitro and in living cells. Another chemical derivative, MIRA-3, have shown antitumor activity in vivo (90). Work conducted by independent laboratories have supported part of these findings and have reported a selective induction of apoptosis by PRIMA-1 in mutant p53 cells via the c-JunNH2-kinase pathway (91). Recently, a new family of p53 reactivators derived from the backbone of PRIMA-1 were described (92). Myers et al. (2005) have found that PRIMA-1 and their derivatives selectively eliminate mutant p53 cells, however, they have been unable to find any evidence for restoration of wild-type p53 properties including p53-induced reporter gene activation in vitro (92). Despite multiple attempts to show interaction of CP-31398 and PRIMA1 with p53 by biophysical methods there have been no data reported supporting a direct interaction of CP-31398 (86) or PRIMA-1 with p53. One hypothetical explanation for the lack of direct interaction is that p53 preparations used for biophysical studies may not represent the correct native wild-type conformation whereas in the cellular context CP-31398 and PRIMA1 have access to and can stabilize p53 DNA binding conformation during protein translation. In agreement with this hypothesis are results from (86) showing that CP-31398 can induce the PAb1620 epitope in cells expressing mutant p53. This has been prevented by cycloheximide, suggesting that CP31398 might exclusively act on newly synthesized p53. However, the simple structure and low molecular weight of PRIMA-1 have raised concerns about its ability to restore the DNA-binding activity of structurally diverse DNA contact as well as structural mutants. In addition, the high concentrations needed to mediate CP-31398 and PRIMA-1 action on p53 (up to 150 μM in in vitro and cellular assays) compared to a lower IC50 in cell proliferation assays (ca. 20 μM for PRIMA-1) suggests


27

that PRIMA-1 might exhibit off-target activities. Taken together the mode of action of both CP-31398 and PRIMA-1 remains elusive and somewhat controversial. The first proof-of-concept for rescuing of mutant p53 by a peptide stabilizing wild-type p53 conformation has come from studies with a short SH3 domain peptide CDB3. This peptide has been derived from the crystal structure of the p53 binding protein, 53BP2, in complex with the p53 DNA-binding domain (93). Biophysical and NMR spectroscopy studies have shown that CDB3 peptide binds the p53 DNA binding domain specifically and stabilizes it against thermal unfolding in a chaperone-like manner (94). NMR data have indicated that the CDB3 peptide is capable of binding to the native structure of p53 and to revert the chemical shifts of the R249S hot-spot mutant back to a wild-type (95). It has been demonstrated that CDB3 peptide penetrates cells, induces active wild-type p53 conformation in the hot-spot mutants, His-175 and His-273, resulting in activation of p53 target genes and partial restoration of apoptosis (96). Based on these experiments, the authors have concluded that in order to rescue the conformation of unstable p53 mutants, drugs with a mode of action similar to CDB3 peptide have to bind p53 during or immediately after biosynthesis (62). Recently, a cellular screening approach has been described for identification of small molecules targeting p53 family members e.g., p73 that can exert antitumor effects in p53 mutant cell systems (97). Several small molecules have been discovered that can activate p53 reporter activity and p53 target genes (e.g., p21 WAF1 , KILLER/DR5), induce apoptosis in p53-deficient cells, and demonstrate potent antitumor activity in vivo (97). Although substantial progress has been made in proving the principle feasibility of restoring wild-type conformation and function to mutant p53, several issues need to be resolved before this approach becomes a clinical reality. None of the described p53-activating small molecules have been proven to act directly on p53 in a clear mechanistic fashion. In addition, their potency and druglike properties need substantial improvement. Even more importantly, there are multiple forms of mutant p53 in human cancer. It still remains to be demonstrated that a small-molecule compound could effectively restore the function of a substantial number of these mutants.

28

Klein and Vassilev

4. INHIBITION OF THE p53-MDM2 INTERACTION 4.1. Function and Structure of MDM2 The cellular oncoprotein MDM2 (murine double minute 2, also termed HDM2 in humans) acts as a master cellular regulator of the p53 protein (11,20,98,99). MDM2 has been originally identified as oncoprotein that binds to p53 and inhibits p53-mediated transactivation (100). The mdm2 gene was found to be up-regulated in human tumors and tumor cell lines by gene amplification, increased transcription and enhanced translation. The overall frequency of mdm2 gene amplification in human tumor tissue samples is ca. 7% with the highest frequency observed in soft tissue tumors (20–30%), osteosarcomas (16%), and esophageal carcinomas (13%) (101–103). MDM2 encodes a multi domain protein with i) an N-terminal domain that contains the binding site for p53 and p73; ii) an acidic domain that contains a binding site for the tumor suppressor p14ARF ; iii) a putative Zn finger and binding site for the retinoblastoma protein Rb, and iv) a RING finger and E3 ligase domain that is responsible for the ubiquitination of p53. In addition, MDM2 contains nuclear import and export signals (100). MDM2 blocks transcriptional activation of p53 by physical binding to the N-terminal transactivation domain (residues 15–29) and by targeting p53 for ubiquitination and degradation through its E3 ligase activity and export from the nucleus to the cytosol (104). As a consequence of MDM2 regulation, p53 exhibits a short half-life and the concentration of active p53 in the nucleus is kept at low levels. Moreover, the MDM2 gene itself contains 2 p53 responsive elements in its promoter region and MDM2 transcription is induced following p53 activation and accumulation. Thus p53 and MDM2 mutually regulate their protein levels through a negative feedback loop (105,106) (Fig. 1). In tumors overexpressing MDM2, this feedback loop is dysregulated and even upon p53 activation its levels are no longer sufficient to exert effective tumor suppressor function. Inhibition of MDM2 expression or its binding to p53 is expected to overcome the oncogenic consequences of MDM2 overexpression and stabilize nuclear p53, resulting in the concurrent activation of p53 downstream genes and induction of cell cycle arrest and apoptosis. Genetic and biochemical studies have mapped the p53-MDM2 binding site to the N-terminal part of the transactivation domain of p53 (also References 107–109 termed BOX1 domain) and the N-terminal domain of MDM2. The structural basis of this interaction has been


29

Degradation

MDM2

p53 Inhibition

Activation MDM2 gene

Fig. 1. Scheme of the p53-MDM2 autoregulatory feedback loop. Modified from Trends Mol Med., 2007.

revealed by the crystal structure of the N-terminal MDM2 domain in complex with a short peptide derived from the N-terminal p53 domain (residues 15–29) (Fig. 3A) (110): The unstructured transactivation domain of p53 (42,111) forms an amphiphilic -helix that projects their hydrophobic residues Phe (19), Trp (23), and Leu (26) into a

A

B Cl

O

N

N

Cl

O

N

N

O

N

N

O

O

Cl

S

N Cl N H

O

CP-31398

R

I

N

N

C

NH

O

Benzodiazepines

Nutlin-3

N O

Pifithrin-α

Cl O

N

OH NH

CH2 CH2

N

O

O

N H

Cl

O

Spiro-oxindoles

O

MIRA-1

O + N – O

O

O

HN

S N

Pifithrin-µ

Quilinols

OH

O

S NH2

N

OH

PRIMA-1

O

NH OH

O

O

N

N

O

RITA

S

HLI98C OH

Cl

Fig. 2. Small-molecule modulators of the p53 pathway: A Activators of mutant p53, B Inhibitors of the p53-MDM2 interaction and E3 ligase activity of MDM2, and C p53 inhibitors. See text for details.

30

Klein and Vassilev

A

B

L26 W23

C

F19

D

Fig. 3. Co-crystal structures of MDM2 with A The natural p53 peptide (110), B The high affinity AP peptide (125,149), C Benzodiazepinedione (184) and D Nutlin-2 (126).

relatively deep hydrophobic cleft on the surface of MDM2 (110). The MDM2 binding site on p53 coincides with the -helical F FFmotif (F: phenylalanine, : hydrophobic residue; in p53: Trp (23)) that is responsible for the interaction with factors of the transcriptional machinery such as hTAFII31 (112). This mode of binding explains why MDM2 binding inhibits the transcriptional activity of p53. The structure of the p53-binding pocket on MDM2 evoked major interest in designing small-molecule antagonists that could block p53 binding by occupying the hydrophobic binding pocket on MDM2 surface (113). A survey of protein–protein interactions with potential as drug targets revealed that the overall dimension of the MDM2 binding pocket is slightly above the arbitrary defined non-amenable range (114). This analysis suggested that finding a druglike inhibitor of the p53-MDM2 interaction would be possible but likely very difficult (114). NMR spectroscopic studies with synthetic peptides showed that residues Phe (16) and Trp (23) are indispensable for binding and provided detailed insight into the structural requirements for the interaction of p53 peptides with MDM2 (115). These studies have revealed global conformational changes in the MDM2 molecule upon ligand


31

binding, stretching far beyond the binding cleft (115,116). Indeed, the p53 binding domain of MDM2 exhibits significant changes in conformation upon binding suggesting that the p53 binding domain of MDM2 is flexible (115,117). NMR studies with the free p53-binding domain of human MDM2 (apo-MDM2) have found that MDM2 residues 16–24, which are strictly conserved in mammals, form a lid that closes over the well ordered p53-binding domain and might help to stabilize apoMDM2 (118). Post-translational modification e.g., phosphorylation within the MDM2 lid may disrupt the p53–MDM2 binding (118). In addition, the existence of MDM2 lid may explain how MDM2 may differ from its close homologue MDM4 (MDMX). MDM4 binds p53 and inhibits its transcriptional activity thus acting as a critical p53 regulator in vivo. However, unlike MDM2, MDM4 does not ubiquitinylate and degrade p53 (119–122). The highest degree of sequence similarity between MDM2 and MDM4 is in their p53 binding domains, (123) whereas the MDM2 lid is not conserved. Therefore, one can speculate that lid modifications may be responsible for differential regulation of the binding affinity of MDM2 and MDM4 for p53 in the cellular context (118). Recently, the structure of human apo-MDM2 has been solved by NMR spectroscopy (124). Data analysis has confirmed that p53 binding is accompanied by displacement of the flexible lid and adjustments of secondary structure. These results suggest that human MDM2 becomes more rigid and stable upon binding to p53. They also indicate that the binding pocket of human apo-MDM2 is relatively shallow whereas MDM2 in complex with peptides (110,125) or small molecule antagonists (126,127) exhibits a more open and accessible conformation (124). Based on these observations the authors have suggested that the structure of ligand-complexed MDM2 but not apo-MDM2 should serve as a template for rational drug design.

4.2. Validation of MDM2 as a Target The discovery of a relatively deep p53-binding pocket on MDM2 molecule further strengthened the interest to MDM2 as a target for therapeutic intervention (110). In the following years, several laboratories applied a variety of experimental approaches to examine the biological and chemical feasibility of targeting the p53-MDM2 interaction. Microinjection of a monoclonal antibody, 3G5, that blocks the p53 binding site on MDM2 have shown an induction of p53 transcriptional activity (128,129). Inhibition of MDM2 expression by antisense oligonucleotides has resulted in p53 activation in several cancer cell

32

Klein and Vassilev

types, p53-mediated cell cycle arrest and apoptosis. Also, MDM2 inhibition have potentiated the effects of chemotherapeutic agents both in vitro and in vivo (130–134). Knockdown of MDM2 by RNA interference delivered via a protamine-antibody fusion protein has activated p53 and caused growth inhibition of subcutaneous B16 tumors in nude mice (135). Microinjection or intracellular expression of fusion proteins of the phage-display optimized IP3 peptide with Thioredoxin or GST, respectively, have resulted in induction of p53 transcriptional activity and p53-mediated apoptosis (129,136). A p53 dependent cell cycle arrest and apoptosis have been induced also by the transduction of cancer cells with IP3 and AP peptides (see the following for details) (137–139) coupled with peptide transduction domains penetration and Tat. In another study, MDM2 downregulation has caused regression of retinoblastoma tumors grown in rabbit eyes (140). Further validation of the p53-MDM2 interaction for tumor therapy comes from genetic modulation of MDM2 expression in mice. Targeted disruption of the mdm2 gene results in embryonic lethality but the mice can be saved by simultaneous deletion of TP53. These studies support the critical function of MDM2 in p53 regulation in vivo (141,142). Studies with conditional MDM2 knockouts provide further support to the rationale for inhibiting the p53MDM2 interaction for cancer therapy (143). Mice with a hypomorphic MDM2 allele that express approx 30% of the normal level of MDM2 are viable and have a normal lifespan. p53 levels have been found elevated in mouse tissues but the only adverse effects were moderately decreased white blood cell count and elevated apoptosis in the thymus and small intestine. These results imply that inhibition of MDM2 in mice does not cause major target-related toxicity with the possible exception of certain hematopoietic defects. Moreover, they suggest that even partial inhibition of MDM2 may be sufficient to activate the p53 pathway in vivo. In summary, several biochemical and genetic studies have validated the p53-MDM2 interaction as a target for pharmacological intervention. However, the development of pharmacologically relevant antagonists has met significant difficulties.

4.3. Peptidic Inhibitors of the p53-MDM2 Interaction Since the discovery of the role of MDM2 as a negative regulator of p53 and the existence of a well-defined hydrophobic p53-binding pocket on MDM2 (110) the p53-MDM2 interaction has attracted major


33

interest as a therapeutic target. (3,104,144,145) The first peptidic antagonists of the p53-MDM2 binding have been identified by modeling the interactions of the MDM2 binding peptide derived from the N-terminal domain of p53. Bottger et al. have used phage-display to find novel MDM2 binding peptides with enhanced affinity (146). Using this approach, the phage IP3 peptide has been generated with a 30-fold higher affinity than the native p53 peptide. The alignment of the phage peptide sequences has allowed to define PxFxDYWxxL as a p53 consensus motif for binding to MDM2 (146,147). Using a semirational design in combination with NMR spectroscopy Garcia-Echeverria et al. (2000) generated a highly potent version of the IP3 peptide, the AP peptide (19-26) (148). To optimize the affinity, artificial amino acids have been introduced into the IP3 peptide. It has been proposed that these residues stabilize entropically the helical conformation of the peptide (Aib-21 for Asp-21 and Ac3c-25 for Leu-25) and form additional polar (Pmp-22 for Tyr-22) and hydrophobic van-der-Waals interactions (Ser-20 by Met-20 and Trp-23 by 6-Cl-Trp-23) with MDM2. As a result, the affinity of the AP peptide has increased 60-fold in comparison to the phage IP3 and nearly 2,000-fold in comparison with the native p53 peptide (148). The crystal structures of human MDM2 and of a humanized version of Xenopus MDM2 (hXDM2) in complex with the AP peptide confirmed the modeling predictions of Garcia-Echeverria et al. that steric shape complementarity, covered lipophilic surface and entropic stabilization could substantially enhance the affinity of a MDM2 ligand (Fig. 3B) (125,149). The experimental data obtained with peptidic MDM2 antagonists have validated the predictions of molecular dynamics approaches (150) and suggested a quantitative structure activity relationship (QSAR) for the development of small molecule antagonists (151).

4.4. Small-Molecule Inhibitors of the p53-MDM2 Interaction Targeting protein-protein interactions with small-molecules has long been considered as high risk and has met skepticism throughout the drug discovery community (152–154). This view has been challenged during the recent years by several proof-of-concept studies that targeted protein interactions with peptidomimetic and small-molecule antagonists (114,154–157). However, most of the compounds described in the literature have exhibited major liabilities such as relatively low potency and selectivity as well as inadequate pharmacological properties. Therefore, the development of drug-like small molecule MDM2 inhibitors is still in its infancy. In the quest for small-molecule

34

Klein and Vassilev

antagonists of the p53-MDM2 interaction several ELISA, fluorescence and NMR-based binding assays amenable for high throughput screening of small molecules have been developed (158–162). During the last few years, putative antagonists of the p53-MDM2 interaction have been identified representing several different compound classes. The first reported small-molecule MDM2 antagonists have been identified from a class of phenoxy acetic acid and phenoxymethyl tetrazole, also known as chalcones (116). Chalcones inhibit p53MDM2 interaction with IC50 values in the high micromolar range by binding to the p53-binding pocket on MDM2 as demonstrated by NMR spectroscopy but their potency and selectivity has been inadequate. They have also shown inhibition of glutathione-S-transferase activity (116). Recently, boronic-chalcone derivatives have been described as putative MDM2 antagonists. However, the authors have not provided evidence that the action of these compounds is related to the direct inhibition of the p53-MDM2 interaction (163). Using a computer-aided design Zhao et al. have synthesized putative nonpeptidic polycyclic MDM2 antagonists (164). Their initial evaluation have shown that some of these inhibitors have a moderate affinity for MDM2 and can induce the p53 pathway in tumor cell lines but their cellular potency is low (164). The fungal cyclic nonapeptide Chlorofusin has been identified by screening a library of microbial extracts (165). Chlorofusin binds to MDM2 and inhibits the p53-MDM2 interaction. Because of its low KD of 4.6 μM, complex chemical structure, and a molecular mass of 1364 Da, chlorofusin does not represent a candidate drug but has been proposed as lead for design of more potent analogues (165,166). The synthesis of the peptide portion of the molecule and the biosynthesis of chlorofusin have been described recently but no cellular or structural data have been reported. (167–169). 4.4.1. The Nutlins Recently, the first potent and selective small-molecule antagonists of the p53-MDM2-interaction both in vitro and in vivo were identified from a class of cis-imidazoline derivatives, also termed nutlins (Fig. 2B) (126). The lead compounds were found by high-throughput screening and improved by multidimensional optimization and rational structure-based design. NMR spectroscopy using a humanized version of Xenopus MDM2 (127) was applied to select compounds that reversibly bind to the primary binding pocket of MDM2 for further optimization (170,171). Nutlins bind to MDM2 with high affinity and


35

displace p53 with IC50 in the 100–300 nM range (126). The first high resolution crystal structure of MDM2 in complex with a small molecule, nutlin-2, confirmed that these compounds bind MDM2 in the p53-binding pocket (Fig. 3D) (126). The structure revealed that the high binding affinity of nutlins is due to the geometry of a rigid imidazoline scaffold projecting its 3 aromatic substituents deeply into the hydrophobic sub-pockets. An overlay of the structure coordinates of the MDM2-bound nutlin-2 and the MDM2-bound p53 peptide (110) revealed that the interactions of the 3 p53 peptide side chains, known to be essential for MDM2 binding, are successfully mimicked by the inhibitor in the 3 subpockets. Leu 26 and Trp 23 are mimicked by the 2 chlorophenyl groups, and Phe 19 is mimicked by the isopropoxy group (114,126). The structure of MDM2-nutlin complex provided clear evidence that it is possible to generate potent small-molecule inhibitors of protein–protein interactions if these compounds can effectively mimic essential binding interactions of the natural protein ligand (114). Treatment of a panel of tumor cell lines expressing wild-type p53 with MDM2 antagonists stabilized p53 and activated the p53 pathway as revealed by the induction of multiple p53-regulated genes. This led to a cell cycle arrest in G1 and G2 phases and apoptosis with IC50s in the low micromolar range. This mode of action occurred only in cells with wild-type but not mutant p53 confirming that the cellular activity of nutlins is derived from intervention in MDM2-mediated p53 regulation. Human and mouse fibroblasts showed comparable sensitivity to nutlin treatment suggesting that mouse xenograft models can assess adequately the potential therapeutic window of this new strategy (172). Oral administration of nutlin-3 to nude mice bearing established osteosarcoma xenografts tumors overexpressing MDM2 protein (SJSA-1) caused dose-dependent tumor inhibition and tumor regression without major target-related toxicities (126). Nutlins are not genotoxic agents and stabilize p53 without the induction of p53 phosphorylation and likely any other modifications that p53 acquires in response to oncogenic stress (126). Therefore, they represent valuable molecular tools to study the role of posttranslational modifications in p53 function (173). Comparison of the activity of nutlin-induced unphosphorylated p53 and phosphorylated p53 induced by genotoxic drugs in cancer cells revealed no difference in their sequence-specific DNA binding, ability to transactivate p53 target genes and to induce p53-dependent apoptosis (174). The observation that p53 can be effectively activated without the need for

36

Klein and Vassilev

phosphorylation supports the use of MDM2 antagonists as single agents in the treatment of tumors with wild-type p53 (173). Nutlins act directly on MDM2 to prevent its interaction with p53 and therefore they can compensate for any signaling defects in cancer cells upstream of p53 including MDM2 overexpression. However, they require intact downstream p53 signaling for effective execution of p53-dependent cell cycle arrest and apoptosis (173). Studies using nutlin-3 to probe p53 downstream signaling in multiple tumor-derived cancer cell lines expressing wild-type p53 revealed that cell cycle arrest function of p53 is well preserved in cancer. However, apoptotic response to MDM2 antagonists is highly variable among cancer lines, suggesting that many tumors might have suffered aberrations in p53-dependent apoptotic signaling (175,176). Not surprisingly, cells with mdm2 gene amplification were found most sensitive to inhibition of the p53-MDM2 interaction. Usually it is rare to find more than one genetic defect on the same pathway. Cancer cells in which mdm2 gene is amplified express wild-type p53 protein because MDM2 overexpression can effectively disable p53 function (100). In these cells, p53 function should be effectively restored by MDM2 antagonists. Indeed, nutlin-3 proved most efficacious against osteosarcoma cells and xenografts in which mdm2 gene is amplified. However, it was also efficacious against tumors with normal MDM2 expression in which at least the major components of p53–dependent apoptotic signaling are preserved, suggesting that patients with wild-type p53 tumors and normal MDM2 expression may benefit from MDM2 antagonists (175). Recent studies with nutlin-3 demonstrated that nongenotoxic p53 activation may be an effective new strategy for the therapy of hematological cancers such as acute myeloid leukemia, multiple myeloma, and B-CLL. Treatment of multiple myeloma tumor samples with nutlin-3 resulted in an increase of p53 levels, and induced p53 target genes (e.g., p21 and MDM2) and apoptotic cell death (177). Tumor cells underwent effective apoptosis even in the presence of stromal cells, which themselves appeared to tolerate exposure to nutlin-3 (177). Kojima et al. (2005) demonstrated a similar mode-of-action in acute myeloid leukemia (AML) cells that frequently overexpress MDM2 (178). Treatment with nutlins triggered several molecular events consistent with induction of apoptosis whereas no induction of apoptosis was observed in AML samples harboring mutant p53. Colony formation of AML progenitors was inhibited in a dosedependent fashion, whereas normal CD34(+) progenitor cells were less affected. Mechanistic studies suggested that nutlin-induced apoptosis


37

was mediated by both transcriptional activation of proapoptotic Bcl-2 family proteins, and transcription-independent mitochondrial permeabilization resulting from mitochondrial p53 translocation. Although MDM2 antagonists are efficacious as single agents they may also have additive or synergistic effects with other cytotoxic agents that activate p53 signaling by virtue of being genotoxic. MDM2 inhibition synergistically enhanced cytotoxicity of cytosine arabinoside and doxorubicin in AML blasts but not in normal hematopoietic progenitor cells (178). Two recent studies showed that nutlin-3 can induce the p53 pathway and apoptosis in nearly 100% of B-CLL and synergizes with fludarabine and chlorambucil. Importantly, nutlins exhibit low cytotoxicity on T cells, bone marrow and CD34(+) cells despite induction of p53 (179,180). The p53-MDM2 interaction might thus represent an attractive therapeutic option for combination therapy in B-CLL, especially as p53 inactivation by mutation occurs rarely in B-CLL at the time of diagnosis and usually occurs late during the disease (179,180). In the last 2 years, over 30 studies have been published using nutlins-3 as a tool to study p53 biology or explore potential therapeutic applications. A recent study using a large-scale RNA interference screen to identify genes important for nutlins-3 activity revealed that p53 together with the p53 binding protein 53BP1 that plays a role in DNA damage response are critical for response to nutlins (181). Taken together, the studies conducted so far with nutlins have provided the proof-of-principle that the p53-MDM2 interaction can be effectively inhibited by small molecules and that MDM2 antagonists might offer therapeutic use in the treatment of tumors that have retained wild-type p53. Nutlins’ discovery also strengthened the notion that pharmacological inhibitor of protein-protein binding can be developed as a therapeutic modality in cancer and other diseases. 4.4.2. Other Recent Small Molecule MDM2 Antagonists Following the discovery of the nutlins, several reports have been published that describe novel small-molecule MDM2 antagonists with in vitro and cellular activity (reviewed in (173,182,183)). Recently, a novel series of benzodiazepinedione antagonists of the HDM2-p53 interaction have been described (Fig. 2B). The comparison of the co-crystal structures of nutlin-2 and a 1,4-benzodiazepine2,5-dione in complex with MDM2 (Fig. 3C) showed that indeed the p53-MDM2 interaction can be mimicked by chemically diverse

38

Klein and Vassilev

small molecules (126,184). The initial 1,4-benzodiazepine-2,5-diones exhibited low cellular penetrability. However, novel representatives of the 1,4-benzodiazepine-2,5-dione scaffold have improved cellular activity and pharmacokinetic properties, induce stabilization of p53 and subsequent transcription of p53 target genes and inhibit the proliferation of tumor cells in the low μM range (184–190). Their relatively low cellular potency and lack of published in vivo xenograft data makes it difficult to judge the potential of this class of compounds for tumor therapy. Based on the available structural information Spiro(oxindole3,3 pyrrolidine) derivatives with a spiro-oxindole core structure (Fig. 2B) have been designed (191,192). The most potent inhibitor has shown an in vitro Ki value of 3 nM and greater than 10,000-fold selectivity to MDM2 over Bcl-2/BclxL proteins. It activates p53 function and inhibits the growth of wild-type p53 tumor cells in the low micromolar range with nearly 30-fold selectivity between cells with wildtype and mutant p53. Their biological properties and in vivo activity need to be further characterized (192). Using an integrated, virtual database screening strategy (193) the same group reported recently another class of quinolinol MDM2 antagonists (Fig. 2B) with good cellular activity and selectivity for wild-type p53 (194). Additional inhibitors of the p53-MDM2 interaction for which primarily biochemical and only a limited set of cell biological data are reported include nonpeptidic sulfonamides, (151) N-(2phenoxybenzoyl)tryptophan-derivatives, (161) Isoindolinone derivatives, (195,196) Terphenyl-based -helical mimetics, (157,197,198) -hairpin peptidomimetics, (199–201) and retroinverso p53 peptides (202). In contrast to classical MDM2 inhibitors that block the p53MDM2 interaction by binding to the p53 binding site, a 2,5-bis (5-hydroxymethyl-2thienyl)furan derivative termed RITA (for reactivation of p53 and induction of tumor cell apoptosis) (Fig. 2B) has been identified recently by a cell-based screen. RITA has been reported to activate the p53 pathway, induce p53 dependent growth arrest and apoptosis and inhibit the growth of HCT116 human tumor xenograft tumors in a p53-dependent manner. Surprisingly, RITA was found to block the p53-MDM2 interaction by binding to the N-terminal domain of p53 (203). However, independent NMR studies have indicated that RITA does not interfere with the p53-MDM2 binding, suggesting that the compound has to work through a different still unknown mechanism (204).


39

In summary, the recent development of potent and selective MDM2 antagonists proved that the p53-MDM2 interaction represents a tractable target for pharmacological intervention and suggested that nongenotoxic p53 activation might offer a new therapeutic option to patients with tumors expressing wild-type p53 (173,183,205).

5. INHIBITION OF MDM2 E3 LIGASE ACTIVITY p53 has a short half-life and its cellular level is controlled primarily by ubiquitin-dependent degradation in the proteasome. Although several E3 ligases that can ubiquitinate p53 have been reported recently e.g., Pirh2, (206) COP1, (207) and ARF-BP1/Mule, (208) their role in maintaining p53 stability is not clear yet. On the other hand, it has been well established that MDM2 plays a critical role in regulating p53 stability. Therefore, inhibition of the E3 ubiquitin ligase activity of MDM2 should lead to p53 stabilization and represents an (209,210) alternative strategy for activation of the tumor suppressor p53. However, inhibitors of MDM2 E3 ligase activity are not expected to inhibit p53-MDM2 binding and may not restore the transcriptional activity of MDM2-bound p53. Whether or not E3 ligase inhibitors can exhibit comparable efficacy compared to p53-MDM2 interaction inhibitors has not been shown. The feasibility to block MDM2-mediated ubiquitinylation of p53 by small-molecule inhibitors has been established in vitro (209). Lai et al. (2001) have used an enzyme assay, monitoring MDM2-catalyzed ubiquitin transfer from preconjugated ubiquitin-Ubc4 to p53, and have identified the first class of small-molecule inhibitors of this enzyme. Three distinct types of reversible inhibitors have been described with benzsulfonamide, urea, and imidazolone scaffolds and potency in the range of 3-15 μM. All 3 types of compounds have displayed selective inhibition of MDM2 E3 ligase activity, with little or no effect on other non-p53 related molecules involved in ubiquitin-dependent TRCP degradation ˜ (e.g., E1, Nedd4, SCF or even on MDM2 autoubiquitinylation. In addition, it has been confirmed that none of the compounds interfered with the physical interaction of p53 and MDM2 (209). However, no data on the cellular potency of these compounds has been reported (209). Recently, the first class of potent small-molecule inhibitors of the MDM2 E3 ubiquitin ligase with cellular activity has been described (210,211). Davydov et al. have developed an assay for the autoubiquitination activity of the E3 ligase of human MDM2

40

Klein and Vassilev

applicable to high-throughput screening (211) and have screened a small library of 10,000 compounds. They have identified a class of phenyl-7-nitro-10H-pyrimido(4,5-b)quinoline-2,4-dione derivatives termed HLI98 (Fig. 2B). These compounds have shown specificity for MDM2 in vitro, although their IC50s are high (20–50 μM) and effects on unrelated E3 ligases have been detected at higher concentrations. In cellular assays, HLI98 compounds have induced stabilization of p53 and MDM2 and activation of p53-dependent transcription and apoptosis at concentrations greater than 50 μM, but have also shown significant p53-independent toxicity (210). The potential of HLI98 derivatives as therapeutics is limited by their low solubility, potency and off-target activity (210).

6. MODULATION OF p53 ACTIVITY FOR PROTECTION OF NORMAL TISSUES DURING CHEMOTHERAPY One of the main functions of activated p53 is to arrest cell cycle progression. Nutlins effectively arrest cells with wild–type p53 in G1 and G2 phases of the cell cycle. If proliferating cells are prevented from entering mitosis they will be largely protected from the cytotoxicity of mitotic inhibitors (e.g., taxanes, vinca alkaloids) that require cells to transition through the division phase for their activity. Thus cell cycle modulation in normal (212–214) proliferating tissues may offer a chemoprotective strategy. MDM2 antagonists depend on the presence of wild-type p53 for their antitumor activity in cancer cells. They do not affect cancer cells in which p53 is mutated and inactive as a transcription factor. At the same time, all normal proliferating cells express wild-type p53 and should be arrested effectively by MDM2 antagonists. These principle features, allow for designing of a strategy to selectively protect normal proliferating cells in patients undergoing antimitotic therapy of their mutant p53 tumors. This approach was tested in vitro using cultured human skin fibroblasts subjected to pretreatment with nutlins-3 for 24 h followed by high doses of paclitaxel for 48 h. Pretreatment of proliferating fibroblasts with MDM2 antagonist reduces significantly the toxicity of paclitaxel. The same treatment scheme, however, did not reduce paclitaxel cytotoxicity to breast cancer cells with mutant p53, (215) suggesting a possible clinical applicability. However, cultured fibroblasts are not a good model for normal tissues that are most sensitive to p53 activation (e.g., hematopoietic system) and further validation of this approach in animal models is needed to assess its potential clinical relevance.


41

Majority of currently used chemotherapeutics are genotoxic agents that can activate the p53 pathway not only in tumor but also in normal tissues (216–220). Although some p53-deficient/mutant tumors have been found to be resistant to chemotherapy (221). in general, the presence or absence of wild-type p53 is not a good clinical predictor of response to chemotherapy (222,223). It is believed that induction of apoptosis because of activation of p53 in normal tissue during tumor therapy is one of the reasons for severe side effects observed with conventional broad-spectrum tumor therapy (218). Therefore, a temporary inhibition of p53 transcriptional activity is proposed as a strategy to protect normal tissue from the harmful consequences of radiotherapy or chemotherapy of p53-mutant tumors (224). A cellular screening for compounds that inhibited the transcriptional activity of p53 has identified Pifithrin-, for “p– fifty three inhibitor”, as a putative p53 inhibitor (Fig. 2C) (225). Pifithrin- has decreased the side effects of chemo- and radiotherapy in mice without affecting the efficacy against p53-mutant tumors and without promoting tumor formation in the short term (224–231). Despite the specificity of Pifithrin- to the p53 pathway, its direct effect on p53 has not been demonstrated so far and the mode-of-action remains largely unknown (232). In an attempt to separate pro-apoptotic from transcriptional activation function of p53, Strom et al., (2006) designed a cellular screen for compounds that affect solely the apoptotic activity of p53 via the mitochondrial pathway (e.g., interaction with Bcl-2 and Bcl-xl) (233). Pifithrin-μ (Fig. 2C) has been identified from this cellular screen as inhibitor of the p53 interaction with the anti-apoptotic proteins Bcl-xL and Bcl-2, whereas it does not affect p53-mediated transactivation (233). These approaches are still awaiting the development of pharmacologically relevant compounds that can be tested in animal models and ultimately in the. clinic.

REFERENCES 1. Gomez-Lazaro M, Fernandez-Gomez FJ, Jordan J. p53: twenty five years understanding the mechanism of genome protection. J Physiol Biochem 2004; 60:287–307. 2. Bourdon JC, Laurenzi VD, Melino G, Lane D. p53: 25 years of research and more questions to answer. Cell Death Differ 2003; 10:397–399. 3. Lane D. Anthony Dipple Carcinogenesis Award. p53 from pathway to therapy. Carcinogenesis 2004; 25:1077–1081. 4. Hofseth LJ, Hussain SP, Harris CC. p53: 25 years after its discovery. Trends Pharmacol Sci 2004; 25:177–181.

42

Klein and Vassilev

5. Braithwaite AW, Royds JA, Jackson P. The p53 story: layers of complexity. Carcinogenesis 2005; 26:1161–1169. 6. Lane DP. Exploiting the p53 pathway for the diagnosis and therapy of human cancer. Cold Spring Harb Symp Quant Biol 2005; 70:489–497. 7. Roth JA. Adenovirus p53 gene therapy. Expert Opin Biol Ther 2006; 6:55–61. 8. May P, May E. Twenty years of p53 research: structural and functional aspects of the p53 protein. Oncogene 1999; 18:7621–7636. 9. Lane DP, Benchimol S. p53: oncogene or anti-oncogene? Genes Dev 1990; 4:1–8. 10. Lane DP. Cancer. p53, guardian of the genome. Nature 1992; 358:15–6. 11. Levine AJ. p53, the cellular gatekeeper for growth and division. Cell 1997; 88:323–331. 12. Hainaut P, Hernandez T, Robinson A, et al. IARC Database of p53 gene mutations in human tumors and cell lines: updated compilation, revised formats and new visualisation tools. Nucleic Acids Res 1998; 26:205–213. 13. Hollstein M, Hergenhahn M, Yang Q, Bartsch H, Wang ZQ, Hainaut P. New approaches to understanding p53 gene tumor mutation spectra. Mutat Res 1999; 431:199–209. 14. Soussi T, Beroud C. Assessing TP53 status in human tumours to evaluate clinical outcome. Nature Rev Cancer 2001; 1:233–240. 15. Chompret A. The Li-Fraumeni syndrome. Biochimie 2002; 84:75–82. 16. Birch JM, Alston RD, McNally RJ, et al. Relative frequency and morphology of cancers in carriers of germline TP53 mutations. Oncogene 2001; 20:4621–4628. 17. Agarwal ML, Taylor WR, Chernov MV, Chernova OB, Stark GR. The p53 network. J Biol Chem 1998; 273:1–4. 18. El-Deiry WS. Regulation of p53 downstream genes. Semin Cancer Biol 1998; 8:345–357. 19. Vousden KH. p53. Death star. Cell 2000; 103:691–694. 20. Vogelstein B, Lane D, Levine AJ. Surfing the p53 network. Nature 2000; 408:307–310. 21. Itahana K, Dimri G, Campisi J. Regulation of cellular senescence by p53. Eur J Biochem 2001; 268:2784–91. 22. Moll UM, Wolff S, Speidel D, Deppert W. Transcription-independent pro-apoptotic functions of p53. Curr Opin Cell Biol 2005; 17:631–636. 23. Erster S, Moll UM. Stress-induced p53 runs a transcription-independent death program. Biochem Biophys Res Commun 2005; 331:843–850. 24. Bode AM, Dong Z. Post-translational modification of p53 in tumorigenesis. Nat Rev Cancer 2004; 4:793–805. 25. Lozano G, Zambetti GP. What have animal models taught us about the p53 pathway? J Pathol 2005; 205:206–220.


43

26. Attardi LD, Donehower LA. Probing p53 biological functions through the use of genetically engineered mouse models. Mutat Res 2005; 576:4–21. 27. Tyner SD, Venkatachalam S, Choi J, et al. p53 mutant mice that display early ageing-associated phenotypes. Nature 2002; 415:45–53. 28. Donehower LA. p53: guardian AND suppressor of longevity? Exp Gerontol 2005; 40:7–9. 29. Garcia-Cao I, Garcia-Cao M, Martin-Caballero J, et al. “Super p53” mice exhibit enhanced DNA damage response, are tumor resistant and age normally. Embo J 2002; 21:6225–6235. 30. Jin S, Martinek S, Joo WS, et al. Identification and characterization of a p53 homologue in Drosophila melanogaster. Proc Natl Acad Sci U S A 2000; 97:7301–7306. 31. Brodsky MH, Nordstrom W, Tsang G, Kwan E, Rubin GM, Abrams JM. Drosophila p53 binds a damage response element at the reaper locus. Cell 2000; 101:103–113. 32. Ollmann M, Young LM, Di Como CJ, et al. Drosophila p53 is a structural and functional homolog of the tumor suppressor p53. Cell 2000; 101:91–101. 33. Derry WB, Putzke AP, Rothman JH. Caenorhabditis elegans p53: role in apoptosis, meiosis, and stress resistance. Science 2001; 294:591–595. 34. Schumacher B, Hofmann K, Boulton S, Gartner A. The C. elegans homolog of the p53 tumor suppressor is required for DNA damageinduced apoptosis. Curr Biol 2001; 11:1722–1727. 35. Yang A, McKeon F. P63 and P73: P53 mimics, menaces and more. Nat Rev Mol Cell Biol 2000; 1:199–207. 36. Melino G, Lu X, Gasco M, Crook T, Knight RA. Functional regulation of p73 and p63: development and cancer. Trends Biochem Sci 2003; 28:663–670. 37. Melino G, De Laurenzi V, Vousden KH. p73: Friend or foe in tumorigenesis. Nat Rev Cancer 2002; 2:605–615. 38. Moll UM, Slade N. p63 and p73: roles in development and tumor formation. Mol Cancer Res 2004; 2:371–386. 39. Arrowsmith CH, Morin P. New insights into p53 function from structural studies. Oncogene 1996; 12:1379–1385. 40. Arrowsmith CH. Structure and function in the p53 family. Cell Death Differ 1999; 6:1169–1173. 41. El-Deiry WS, Kern SE, Pietenpol JA, Kinzler KW, Vogelstein B. Definition of a consensus binding site for p53. Nat Genet 1992; 1:45–49. 42. Dawson R, Muller L, Dehner A, Klein C, Kessler H, Buchner J. The N-terminal domain of p53 is natively unfolded. J Mol Biol 2003; 332: 1131–41.

44

Klein and Vassilev

43. Bell S, Klein C, Müller L, Hansen S, Buchner J. p53 contains large unstructured regions in its native state. J Mol Biol 2002; 322:917–927. 44. Okorokov AL, Sherman MB, Plisson C, et al. The structure of p53 tumour suppressor protein reveals the basis for its functional plasticity. Embo J 2006; 25:5191–5200. 45. Cho Y, Gorina S, Jeffrey PD, Pavletich NP. Crystal structure of a p53 tumor suppressor-DNA complex: understanding tumorigenic mutations. Science 1994; 265:346–355. 46. Zhao K, Chai X, Johnston K, Clements A, Marmorstein R. Crystal Structure of the Mouse p53 Core DNA-binding Domain at 2.7 A Resolution. J Biol Chem 2001; 276:12120–12127. 47. Bullock AN, Henckel J, Fersht AR. Quantitative analysis of residual folding and DNA binding in mutant p53 core domain: definition of mutant states for rescue in cancer therapy. Oncogene 2000; 19:1245–56. 48. Bullock A, Fersht A. Rescuing the function of mutant p53. Nat Rev Cancer 2001; 1:68–76. 49. Joerger AC, Ang HC, Veprintsev DB, Blair CM, Fersht AR. Structures of p53 cancer mutants and mechanism of rescue by second-site suppressor mutations. J Biol Chem 2005; 280:16030–16037. 50. Cohen PA, Hupp TR, Lane DP, Daniels DA. Biochemical characterization of different conformational states of the Sf9 cell-purified p53His175 mutant protein. FEBS Lett 1999; 463:179–184. 51. Milner J. Flexibility: the key to p53 function? Trends Biochem Sci 1995; 20:49–51. 52. Ravera MW, Carcamo J, Brissette R, et al. Identification of an allosteric binding site on the transcription factor p53 using a phage-displayed peptide library. Oncogene 1998; 16:1993–1999. 53. Stephen CW, Helminen P, Lane DP. Characterisation of epitopes on human p53 using phage-displayed peptide libraries: insights into antibody-peptide interactions. J Mol Biol 1995; 248:58–78. 54. Stephen CW, Lane DP. Mutant conformation of p53. Precise epitope mapping using a filamentous phage epitope library. J Mol Biol 1992; 225:577–583. 55. Hansen S, Hupp TR, Lane DP. Allosteric regulation of the thermostability and DNA binding activity of human p53 by specific interacting proteins. J Biol Chem 1996; 271:3917–3924. 56. Wright JD, Noskov SY, Lim C. Factors governing loss and rescue of DNA binding upon single and double mutations in the p53 core domain. Nucleic Acids Res 2002; 30:1563–1574. 57. Wong KB, DeDecker BS, Freund SM, Proctor MR, Bycroft M, Fersht AR. Hot-spot mutants of p53 core domain evince characteristic local structural changes. Proc Natl Acad Sci U S A 1999; 96: 8438–8442.


45

58. Ang HC, Joerger AC, Mayer S, Fersht AR. Effects of common cancer mutations on stability and DNA binding of full-length p53 compared with isolated core domains. J Biol Chem 2006; 281:21934–41. 59. Lane DP, Hupp TR. Drug discovery and p53. Drug Discov Today 2003; 8:347–355. 60. Edelman J, Nemunaitis J. Adenoviral p53 gene therapy in squamous cell cancer of the head and neck region. Curr Opin Mol Ther 2003; 5:611–617. 61. Zeimet AG, Riha K, Berger J, et al. New insights into p53 regulation and gene therapy for cancer. Biochem Pharmacol 2000; 60:1153–1163. 62. Friedler A, Veprintsev DB, Hansson LO, Fersht AR. Kinetic instability of p53 core domain mutants: implications for rescue by small molecules. J Biol Chem 2003; 278:24108–24112. 63. Wieczorek AM, Waterman JL, Waterman MJ, Halazonetis TD. Structure-based rescue of common tumor-derived p53 mutants. Nat Med 1996; 2:1143–1146. 64. Brachmann RK, Yu K, Eby Y, Pavletich NP, Boeke JD. Genetic selection of intragenic suppressor mutations that reverse the effect of common p53 cancer mutations. Embo J 1998; 17:1847–1859. 65. Nikolova PV, Wong KB, DeDecker B, Henckel J, Fersht AR. Mechanism of rescue of common p53 cancer mutations by second-site suppressor mutations. Embo J 2000; 19:370–378. 66. Nikolova PV, Henckel J, Lane DP, Fersht AR. Semirational design of active tumor suppressor p53 DNA binding domain with enhanced stability. Proc Natl Acad Sci U S A 1998; 95:14675–14680. 67. Matsumura I, Ellington AD. In vitro evolution of thermostable p53 variants. Protein Sci 1999; 8:731–740. 68. Xirodimas DP, Lane DP. Molecular evolution of the thermosensitive PAb1620 epitope of human p53 by DNA shuffling. J Biol Chem 1999; 274:28042–28049. 69. Joerger AC, Allen MD, Fersht AR. Crystal structure of a superstable mutant of human p53 core domain. Insights into the mechanism of rescuing oncogenic mutations. J Biol Chem 2004; 279:1291–1296. 70. Bullock AN, Henckel J, DeDecker BS, et al. Thermodynamic stability of wild-type and mutant p53 core domain. Proc Natl Acad Sci U S A 1997; 94:14338–14342. 71. Brown CR, Hong-Brown LQ, Welch WJ. Correcting temperaturesensitive protein folding defects. J Clin Invest 1997; 99:1432–1444. 72. Maurici D, Monti P, Campomenosi P, et al. Amifostine (WR2721) restores transcriptional activity of specific p53 mutant proteins in a yeast functional assay. Oncogene 2001; 20:3533–3540. 73. North S, Pluquet O, Maurici D, El Ghissassi F, Hainaut P. Restoration of wild-type conformation and activity of a temperaturesensitive mutant

46

74.

75.

76.

77. 78.

79. 80.

81.

82.

83.

84.

85.

86.

87.

Klein and Vassilev of p53 (p53(V272M)) by the cytoprotective aminothiol WR1065 in the esophageal cancer cell line TE-1. Mol Carcinog 2002; 33:181–188. Ohnishi K, Ota I, Takahashi A, Ohnishi T. Glycerol restores p53dependent radiosensitivity of human head and neck cancer cells bearing mutant p53. Br J Cancer 2000; 83:1735–1739. Ohnishi T, Matsumoto H, Wang X, Takahashi A, Tamamoto T, Ohnishi K. Restoration by glycerol of p53-dependent apoptosis in cells bearing the mutant p53 gene. Int J Radiat Biol 1999; 75:1095–1098. Ohnishi T, Ohnishi K, Wang X, Takahashi A, Okaichi K. Restoration of mutant TP53 to normal TP53 function by glycerol as a chemical chaperone. Radiat Res 1999; 151:498–500. Bykov VJ, Selivanova G, Wiman KG. Small molecules that reactivate mutant p53. Eur J Cancer 2003; 39:1828–1834. Foster BA, Coffey HA, Morin MJ, Rastinejad F. Pharmacological rescue of mutant p53 conformation and function. Science 1999; 286: 2507–2510. Smith ML, Fornace AJ. Chemotherapeutic targeting of p53. Cancer Biology and Therapy 2002:47–55. Takimoto R, Wang W, Dicker DT, Rastinejad F, Lyssikatos J, S. E-DW. The mutant p53-conformation modifying drug, CP-31398, can induce apoptosis of human cancer cells and can stabilize wild-type p53 protein. Cancer Biology and Therapy 2002:47–55. Demma MJ, Wong S, Maxwell E, Dasmahapatra B. CP-31398 restores DNA-binding activity to mutant p53 in vitro but does not affect p53 homologs p63 and p73. J Biol Chem 2004; 279:45887–45896. Wischhusen J, Naumann U, Ohgaki H, Rastinejad F, Weller M. CP31398, a novel p53-stabilizing agent, induces p53-dependent and p53independent glioma cell death. Oncogene 2003; 22:8233–8245. Luu Y, Bush J, Cheung KJ, Jr., Li G. The p53 stabilizing compound CP31398 induces apoptosis by activating the intrinsic Bax/mitochondrial/caspase-9 pathway. Exp Cell Res 2002; 276:214–222. Luu Y, Li G. The p53-stabilizing compound CP-31398 enhances ultraviolet-B-induced apoptosis in a human melanoma cell line MMRU. J Invest Dermatol 2002; 119:1207–1209. Tanner S, Barberis A. CP-31398, a putative p53-stabilizing molecule tested in mammalian cells and in yeast for its effects on p53 transcriptional activity. J Negat Results Biomed 2004; 3:5. Rippin TM, Bykov VJ, Freund SM, Selivanova G, Wiman KG, Fersht AR. Characterization of the p53-rescue drug CP-31398 in vitro and in living cells. Oncogene 2002; 21:2119–2129. Bykov VJ, Issaeva N, Shilov A, et al. Restoration of the tumor suppressor function to mutant p53 by a low-molecular-weight compound. Nat Med 2002; 8:282–288.


47

88. Bykov VJ, Issaeva N, Selivanova G, Wiman KG. Mutant p53dependent growth suppression distinguishes PRIMA-1 from known anticancer drugs: a statistical analysis of information in the National Cancer Institute database. Carcinogenesis 2002; 23:2011–2018. 89. Bykov VJ, Zache N, Stridh H, et al. PRIMA-1(MET) synergizes with cisplatin to induce tumor cell apoptosis. Oncogene 2005; 24:3484–3491. 90. Bykov VJ, Issaeva N, Zache N, et al. Reactivation of mutant p53 and induction of apoptosis in human tumor cells by maleimide analogs. J Biol Chem 2005; 280:30384–30391. 91. Li Y, Mao Y, Brandt-Rauf PW, Williams AC, Fine RL. Selective induction of apoptosis in mutant p53 premalignant and malignant cancer cells by PRIMA-1 through the c-Jun-NH2-kinase pathway. Mol Cancer Ther 2005; 4:901–909. 92. Myers MC, Wang J, Iera JA, et al. A new family of small molecules to probe the reactivation of mutant p53. J Am Chem Soc 2005; 127: 6152–6153. 93. Gorina S, Pavletich NP. Structure of the p53 tumor suppressor bound to the ankyrin and SH3 domains of 53BP2. Science 1996; 274: 1001–1005. 94. Friedler A, Hansson LO, Veprintsev DB, et al. A peptide that binds and stabilizes p53 core domain: chaperone strategy for rescue of oncogenic mutants. Proc Natl Acad Sci U S A 2002; 99:937–942. 95. Friedler A, DeDecker BS, Freund SM, Blair C, Rudiger S, Fersht AR. Structural distortion of p53 by the mutation R249S and its rescue by a designed peptide: implications for “mutant conformation”. J Mol Biol 2004; 336:187–196. 96. Issaeva N, Friedler A, Bozko P, Wiman KG, Fersht AR, Selivanova G. Rescue of mutants of the tumor suppressor p53 in cancer cells by a designed peptide. Proc Natl Acad Sci U S A 2003; 100:13303–13307. 97. Wang W, Kim SH, El-Deiry WS. Small-molecule modulators of p53 family signaling and antitumor effects in p53-deficient human colon tumor xenografts. Proc Natl Acad Sci U S A 2006; 103:11003–11008. 98. Iwakuma T, Lozano G. MDM2, an introduction. Mol Cancer Res 2003; 1:993–1000. 99. Bond GL, Hu W, Levine AJ. MDM2 is a central node in the p53 pathway: 12 years and counting. Curr Cancer Drug Targets 2005; 5:3–8. 100. Momand J, Wu HH, Dasgupta G. MDM2-master regulator of the p53 tumor suppressor protein. Gene 2000; 242:15–29. 101. Momand J, Jung D, Wilczynski S, Niland J. The MDM2 gene amplification database. Nucleic Acids Res 1998; 26:3453–3459. 102. Watanabe T, Ichikawa A, Saito H, Hotta T. Overexpression of the MDM2 oncogene in leukemia and lymphoma. Leuk Lymphoma 1996; 21:391–397, color plates XVI following 5.

48

Klein and Vassilev

103. Rayburn E, Zhang R, He J, Wang H. MDM2 and human malignancies: expression, clinical pathology, prognostic markers, and implications for chemotherapy. Curr Cancer Drug Targets 2005; 5:27–41. 104. Moll UM, Petrenko O. The MDM2-p53 interaction. Mol Cancer Res 2003; 1:1001–1008. 105. Juven-Gershon T, Oren M. Mdm2: the ups and downs. Mol Med 1999; 5:71–83. 106. Freedman DA, Wu L, Levine AJ. Functions of the MDM2 oncoprotein. Cell Mol Life Sci 1999; 55:96–107. 107. Picksley SM, Vojtesek B, Sparks A, Lane DP. Immunochemical analysis of the interaction of p53 with MDM2;–fine mapping of the MDM2 binding site on p53 using synthetic peptides. Oncogene 1994; 9: 2523–2529. 108. Freedman DA, Epstein CB, Roth JC, Levine AJ. A genetic -2 interaction domains. Mol Cell Biol 1993; 13:4107–4114. 109. Chen J, Marechal V, Levine AJ. Mapping of the p53 and mdm-2 interaction domains. Mol Cell Biol 1993; 13:4107–4114. 110. Kussie PH, Gorina S, Marechal V, et al. Structure of the MDM2 oncoprotein bound to the p53 tumor suppressor transactivation domain. Science 1996; 274:948–953. 111. Lee H, Mok KH, Muhandiram R, et al. Local structural elements in the mostly unstructured transcriptional activation domain of human p53. J Biol Chem 2000; 275:29426–29432. 112. Uesugi M, Verdine GL. The alpha-helical FXXPhiPhi motif in p53: TAF interaction and discrimination by MDM2. Proc Natl Acad Sci U S A 1999; 96:14801–14806. 113. Picksley SM, Darr DA, Mansoor MS, Loadman PM. Current advances in the inhibition of the auto-regulatory interaction between the p53 tumour suppressor protein and MDM2 protein. Expert Opin. Ther. Patents 2001; 11:1825–1835. 114. Fry DC, Vassilev LT. Targeting protein-protein interactions for cancer therapy. J Mol Med in press. 115. Schon O, Friedler A, Bycroft M, Freund SM, Fersht AR. Molecular mechanism of the interaction between MDM2 and p53. J Mol Biol 2002; 323:491–501. 116. Stoll R, Renner C, Hansen S, et al. Chalcone derivatives antagonize interactions between the human oncoprotein MDM2 and p53. Biochemistry 2001; 40:336–144. 117. Schon O, Friedler A, Freund S, Fersht AR. Binding of p53-derived ligands to MDM2 induces a variety of long range conformational changes. J Mol Biol 2004; 336:197–202. 118. McCoy MA, Gesell JJ, Senior MM, Wyss DF. Flexible lid to the p53–binding domain of human Mdm2: implications for p53 regulation. Proc Natl Acad Sci U S A 2003; 100:1645–1648.


49

119. Jackson MW, Berberich SJ. MdmX protects p53 from Mdm2-mediated degradation. Mol Cell Biol 2000; 20:1001–1007. 120. Stad R, Little NA, Xirodimas DP, et al. Mdmx stabilizes p53 and Mdm2 via two distinct mechanisms. EMBO Rep 2001; 2:1029–1034. 121. Parant J, Chavez-Reyes A, Little NA, et al. Rescue of embryonic lethality in Mdm4-null mice by loss of Trp53 suggests a nonoverlapping pathway with MDM2 to regulate p53. Nat Genet 2001; 29:92–95. 122. Marine JC, Jochemsen AG. Mdmx as an essential regulator of p53 activity. Biochem Biophys Res Commun 2005; 331:750–760. 123. Bottger V, Bottger A, Garcia-Echeverria C, et al. Comparative study of the p53-mdm2 and p53-MDMX interfaces. Oncogene 1999; 18:189–199. 124. Uhrinova S, Uhrin D, Powers H, et al. Structure of free MDM2 Nterminal domain reveals conformational adjustments that accompany p53-binding. J Mol Biol 2005; 350:587–598. 125. Klein C, Breitenlechner C, Hesse F, et al. Crystal structure of humanized Xenopus MDM2 in complex with a high-affinity peptide antagonist, Proceedings of the 96th Annual Meeting of the American Association for Cancer Research, Anaheim, California, Abstract 4826, April 16–20, 2005, 2005. 126. Vassilev LT, Vu BT, Graves B, et al. In vivo activation of the p53 pathway by small-molecule antagonists of MDM2. Science 2004; 303:844–848. 127. Fry DC, Emerson SD, Palme S, Vu BT, Liu CM, Podlaski F. NMR structure of a complex between MDM2 and a small molecule inhibitor. J Biomol NMR 2004; 30:163–173. 128. Blaydes JP, Gire V, Rowson JM, Wynford-Thomas D. Tolerance of high levels of wild-type p53 in transformed epithelial cells dependent on auto-regulation by mdm-2. Oncogene 1997; 14:1859–1868. 129. Bottger A, Bottger V, Sparks A, Liu WL, Howard SF, Lane DP. Design of a synthetic Mdm2-binding mini protein that activates the p53 response in vivo. Curr Biol 1997; 7:860–869. 130. Geiger T, Husken D, Weiler J, et al. Consequences of the inhibition of Hdm2 expression in human osteosarcoma cells using antisense oligonucleotides. Anticancer Drug Des 2000; 15:423–430. 131. Chen L, Lu W, Agrawal S, Zhou W, Zhang R, Chen J. Ubiquitous induction of p53 in tumor cells by antisense inhibition of MDM2 expression. Mol Med 1999; 5:21–34. 132. Wang H, Nan L, Yu D, Agrawal S, Zhang R. Antisense anti-MDM2 oligonucleotides as a novel therapeutic approach to human breast cancer: in vitro and in vivo activities and mechanisms. Clin Cancer Res 2001; 7:3613–3624. 133. Wang H, Zeng X, Oliver P, et al. MDM2 oncogene as a target for cancer therapy: An antisense approach. Int J Oncol 1999; 15:653–660.

50

Klein and Vassilev

134. Zhang Z, Li M, Wang H, Agrawal S, Zhang R. Antisense therapy targeting MDM2 oncogene in prostate cancer: Effects on proliferation, apoptosis, multiple gene expression, and chemotherapy. Proc Natl Acad Sci U S A 2003; 100:11636–11641. 135. Song E, Zhu P, Lee SK, et al. Antibody mediated in vivo delivery of small interfering RNAs via cell-surface receptors. Nat Biotechnol 2005; 23:709–717. 136. Wasylyk C, Salvi R, Argentini M, et al. p53 mediated death of cells overexpressing MDM2 by an inhibitor of MDM2 interaction with p53. Oncogene 1999; 18:1921–1934. 137. Chene P, Fuchs J, Bohn J, Garcia-Echeverria C, Furet P, Fabbro D. A small synthetic peptide, which inhibits the p53-hdm2 interaction, stimulates the p53 pathway in tumour cell lines. J Mol Biol 2000; 299: 245–253. 138. Garcia-Echeverria C, Furet P, Chene P. Coupling of the antennapedia third helix to a potent antagonist of the p53/hdm2 protein-protein interaction. Bioorg Med Chem Lett 2001; 11:2161–2164. 139. Chene P, Fuchs J, Carena I, Furet P, Garcia-Echeverria C. Study of the cytotoxic effect of a peptidic inhibitor of the p53-hdm2 interaction in tumor cells. FEBS Lett 2002; 529:293–2937. 140. Harbour JW, Worley L, Ma D, Cohen M. Transducible peptide therapy for uveal melanoma and retinoblastoma. Arch Ophthalmol 2002; 120:1341–1346. 141. Montes de Oca Luna R, Wagner DS, Lozano G. Rescue of early embryonic lethality in mdm2-deficient mice by deletion of p53. Nature 1995; 378:203–206. 142. Jones SN, Roe AE, Donehower LA, Bradley A. Rescue of embryonic lethality in Mdm2-deficient mice by absence of p53. Nature 1995; 378:206–208. 143. Mendrysa SM, McElwee MK, Michalowski J, O’Leary KA, Young KM, Perry ME. mdm2 Is critical for inhibition of p53 during lymphopoiesis and the response to ionizing irradiation. Mol Cell Biol 2003; 23: 462–472. 144. Lane DP, Hupp TR. Drug discovery and p53. Drug Discov Today 2003; 8:347–355. 145. Chene P. Inhibiting the p53-MDM2 interaction: an important target for cancer therapy. Nat Rev Cancer 2003; 3:102–109. 146. Bottger V, Bottger A, Howard SF, et al. Identification of novel mdm2 binding peptides by phage display. Oncogene 1996; 13:2141–2147. 147. Bottger A, Bottger V, Garcia-Echeverria C, et al. Molecular characterization of the hdm2-p53 interaction. J Mol Biol 1997; 269:744–756. 148. Garcia-Echeverria C, Chene P, Blommers MJ, Furet P. Discovery of potent antagonists of the interaction between human double minute 2 and tumor suppressor p53. J Med Chem 2000; 43:3205–3208.


51

149. Sakurai K, Schubert C, Kahne D. Crystallographic analysis of an 8-mer p53 peptide analogue complexed with MDM2. J Am Chem Soc 2006; 128:11000–11001. 150. Massova I, Kollmann PA. Computational Alanine Scanning To Probe Protein-Protein Interactions: A Novel Approach To Evaluate Binding Free Energies. J. Am. Chem. Soc. 1999; 121:8133–8143. 151. Galatin PS, Abraham DJ. QSAR: hydropathic analysis of inhibitors of the p53-mdm2 interaction. Proteins 2001; 45:169–175. 152. Jones S, Thornton JM. Principles of protein-protein interactions. Proc Natl Acad Sci U S A 1996; 93:13–20. 153. Stites WE. Protein-Protein Interactions: Interface Structure, Binding Thermodynamics, and Mutational Analysis. Chem Rev 1997; 97: 1233–1250. 154. Arkin MR, Wells JA. Small-molecule inhibitors of protein-protein interactions: progressing towards the dream. Nat Rev Drug Discov 2004; 3:301–317. 155. Toogood PL. Inhibition of protein-protein association by small molecules: approaches and progress. J Med Chem 2002; 45:1543–1558. 156. Cochran AG. Antagonists of protein-protein interactions. Chemistry & Biology 2000; 7:R85–94. 157. Yin H, Hamilton AD. Strategies for targeting protein-protein interactions with synthetic agents. Angew Chem Int Ed Engl 2005; 44:4130–4163. 158. Lai Z, Auger KR, Manubay CM, Copeland RA. Thermodynamics of p53 binding to hdm2(1–126): effects of phosphorylation and p53 peptide length. Arch Biochem Biophys 2000; 381:278–284. 159. Kane SA, Fleener CA, Zhang YS, Davis LJ, Musselman AL, Huang PS. Development of a binding assay for p53/HDM2 by using homogeneous time-resolved fluorescence. Anal Biochem 2000; 278:29–38. 160. Knight SM, Umezawa N, Lee HS, Gellman SH, Kay BK. A Fluorescence Polarization Assay for the Identification of Inhibitors of the p53-DM2 Protein-Protein Interaction. Anal Biochem 2002; 300:230–6. 161. Zhang R, Mayhood T, Lipari P, et al. Fluorescence polarization assay and inhibitor design for MDM2/p53 interaction. Anal Biochem 2004; 331:138–146. 162. D’Silva L, Ozdowy P, Krajewski M, Rothweiler U, Singh M, Holak TA. Monitoring the effects of antagonists on protein-protein interactions with NMR spectroscopy. J Am Chem Soc 2005; 127:13220–13226. 163. Kumar SK, Hager E, Pettit C, Gurulingappa H, Davidson NE, Khan SR. Design, synthesis, and evaluation of novel boronic-chalcone derivatives as antitumor agents. J Med Chem 2003; 46:2813–28135. 164. Zhao J, Wang M, Chen J, et al. The initial evaluation of non-peptidic small-moleculre HDM2 inhibitors based on p53-HDM2 complex structure. Cancer Letters 2002; 183:69–77.

52

Klein and Vassilev

165. Duncan SJ, Gruschow S, Williams DH, et al. Isolation and structure elucidation of Chlorofusin, a novel p53-MDM2 antagonist from a Fusarium sp. J Am Chem Soc 2001; 123:554–560. 166. Duncan SJ, Cooper MA, Williams DH. Binding of an inhibitor of the p53/MDM2 interaction to MDM2. Chem Commun (Camb) 2003: 316–317. 167. Desai P, Pfeiffer SS, Boger DL. Synthesis of the chlorofusin cyclic peptide: assignment of the asparagine stereochemistry. Org Lett 2003; 5:5047–5050. 168. Malkinson JP, Zloh M, Kadom M, Errington R, Smith PJ, Searcey M. Solid-phase synthesis of the cyclic peptide portion of chlorofusin, an inhibitor of p53-MDM2 interactions. Org Lett 2003; 5:5051–5054. 169. Duncan SJ, Williams DH, Ainsworth M, Martin S, Ford R, Wrigley SK. On the biosynthesis of an inhibitor of the p53/MDM2 interaction. Tetrahedron Letters 2002; 43:1075–1078. 170. Fry DC, Graves BJ, Vassilev LT. Exploiting protein-protein interactions to design an activator of p53. Protein-Protein Interactions: A Molecular Cloning Manual. New York in press: Cold Spring Harbor Laboratory Press. 171. Fry DC, Graves BJ, Vassilev LT. Development of E3-substrate (MDM2p53) binding inhibitors: structural aspects. Meth Enzymol in press. 172. Vassilev LT. Small-Molecule Antagonists of p53-MDM2 Binding: Research Tools and Potential Therapeutics. Cell Cycle 2004; 3:419–421. 173. Vassilev LT. p53 Activation by small molecules: application in oncology. J Med Chem 2005; 48:4491–4499. 174. Thompson T, Tovar C, Yang H, et al. Phosphorylation of p53 on key serines is dispensable for transcriptional activation and apoptosis. J Biol Chem 2004; 279:53015–530122. 175. Tovar C, Rosinski J, Filipovic Z, et al. Small-molecule MDM2 antagonists reveal aberrant p53 signaling in cancer: implications for therapy. Proc Natl Acad Sci U S A 2006; 103:1888–1893. 176. Harris CC. Protein-protein interactions for cancer therapy. Proc Natl Acad Sci U S A 2006; 103:1659–1660. 177. Stuhmer T, Chatterjee M, Hildebrandt M, et al. Non-genotoxic activation of the p53 pathway as a therapeutic strategy for multiple myeloma. Blood 2005; 106:3609–3617. 178. Kojima K, Konopleva M, Samudio IJ, et al. MDM2 antagonists induce p53-dependent apoptosis in AML: implications for leukemia therapy. Blood 2005; 106:3150–3159. 179. Coll-Mulet L, Iglesias-Serret D, Santidrian AF, et al. MDM2 antagonists activate p53 and synergize with genotoxic drugs in B-cell chronic lymphocytic leukemia cells. Blood 2006; 107:4109–4114.


53

180. Secchiero P, Barbarotto E, Tiribelli M, et al. Functional integrity of the p53-mediated apoptotic pathway induced by the nongenotoxic agent nutlin-3 in B-cell chronic lymphocytic leukemia (B-CLL). Blood 2006; 107:4122–4129. 181. Brummelkamp TR, Fabius AW, Mullenders J, et al. An shRNA barcode screen provides insight into cancer cell vulnerability to MDM2 inhibitors. Nat Chem Biol 2006; 2:202–206. 182. Fotouhi N, Graves B. Small molecule inhibitors of p53/MDM2 interaction. Curr Top Med Chem 2005; 5:159–165. 183. Vassilev LT. MDM2 inhibitors for cancer therapy. Trends Mol Med:, in press. 184. Grasberger BL, Lu T, Schubert C, et al. Discovery and cocrystal structure of benzodiazepinedione HDM2 antagonists that activate p53 in cells. J Med Chem 2005; 48:909–912. 185. Parks DJ, Lafrance LV, Calvo RR, et al. 1,4-Benzodiazepine-2,5diones as small molecule antagonists of the HDM2-p53 interaction: discovery and SAR. Bioorg Med Chem Lett 2005; 15:765–770. 186. Raboisson P, Marugan JJ, Schubert C, et al. Structure-based design, synthesis, and biological evaluation of novel 1,4-diazepines as HDM2 antagonists. Bioorg Med Chem Lett 2005; 15:1857–1861. 187. Koblish HK, Zhao S, Franks CF, et al. Benzodiazepinedione inhibitors of the Hdm2:p53 complex suppress human tumor cell proliferation in vitro and sensitize tumors to doxorubicin in vivo. Mol Cancer Ther 2006; 5:160–169. 188. Leonard K, Marugan JJ, Raboisson P, et al. Novel 1,4benzodiazepine2,5-diones as Hdm2 antagonists with improved cellular activity. Bioorg Med Chem Lett 2006; 16:3463–3468. 189. Marugan JJ, Leonard K, Raboisson P, et al. Enantiomerically pure 1, 4-benzodiazepine-2,5-diones as Hdm2 antagonists. Bioorg Med Chem Lett 2006; 16:3115–3120. 190. Parks DJ, LaFrance LV, Calvo RR, et al. Enhanced pharmacokinetic properties of 1,4-benzodiazepine-2,5-dione antagonists of the HDM2p53 protein-protein interaction through structure-based drug design. Bioorg Med Chem Lett 2006; 16:3310–3314. 191. Ding K, Lu Y, Nikolovska-Coleska Z, et al. Structure-based design of potent non-peptide MDM2 inhibitors. J Am Chem Soc 2005; 127:10130–10131. 192. Ding K, Lu Y, Nikolovska-Coleska Z, et al. Structure-based design of spiro-oxindoles as potent, specific small-molecule inhibitors of the MDM2-p53 interaction. J Med Chem 2006; 49:3432–3435. 193. Lu F, Chi SW, Kim DH, Han KH, Kuntz ID, Guy RK. Proteomimetic libraries: design, synthesis, and evaluation of p53-MDM2 interaction inhibitors. J Comb Chem 2006; 8:315–325.

54

Klein and Vassilev

194. Lu Y, Nikolovska-Coleska Z, Fang X, et al. Discovery of a nanomolar inhibitor of the human murine double minute 2 (MDM2)-p53 interaction through an integrated, virtual database screening strategy. J Med Chem 2006; 49:3759–3762. 195. Hardcastle IR, Ahmed SU, Atkins H, et al. Isoindolinone-based inhibitors of the MDM2-p53 protein-protein interaction. Bioorg Med Chem Lett 2005; 15:1515–1520. 196. Hardcastle IR, Ahmed SU, Atkins H, et al. Small-molecule inhibitors of the MDM2-p53 protein-protein interaction based on an isoindolinone scaffold. J Med Chem 2006; 49:6209–6721. 197. Yin H, Lee GI, Park HS, et al. Terphenyl-based helical mimetics that disrupt the p53/HDM2 interaction. Angew Chem Int Ed Engl 2005; 44:2704–2707. 198. Chen L, Yin H, Farooqi B, Sebti S, Hamilton AD, Chen J. p53 alphaHelix mimetics antagonize p53/MDM2 interaction and activate p53. Mol Cancer Ther 2005; 4:1019–1025. 199. Fasan R, Dias RL, Moehle K, et al. Using a beta-hairpin to mimic an alpha-helix: cyclic peptidomimetic inhibitors of the p53-HDM2 proteinprotein interaction. Angew Chem Int Ed Engl 2004; 43:2109–2112. 200. Kritzer JA, Hodsdon ME, Schepartz A. Solution structure of a betapeptide ligand for hDM2. J Am Chem Soc 2005; 127:4118–4119. 201. Kritzer JA, Lear JD, Hodsdon ME, Schepartz A. Helical beta-peptide inhibitors of the p53-hDM2 interaction. J Am Chem Soc 2004; 126:9468–9469. 202. Sakurai K, Chung HS, Kahne D. Use of a retroinverso p53 peptide as an inhibitor of MDM2. J Am Chem Soc 2004; 126:16288–16289. 203. Issaeva N, Bozko P, Enge M, et al. Small molecule RITA binds to p53, blocks p53-HDM-2 interaction and activates p53 function in tumors. Nat Med 2004; 10:1321–1328. 204. Krajewski M, Ozdowy P, D’Silva L, Rothweiler U, Holak TA. NMR indicates that the small molecule RITA does not block p53-MDM2 binding in vitro. Nat Med 2005; 11:1135–1136; author reply 1136–1137. 205. Klein C, Vassilev LT. Targeting the p53-MDM2 interaction to treat cancer. Br J Cancer 2004; 91:1415–1419. 206. Leng RP, Lin Y, Ma W, et al. Pirh2, a p53-induced ubiquitin-protein ligase, promotes p53 degradation. Cell 2003; 112:779–791. 207. Dornan D, Wertz I, Shimizu H, et al. The ubiquitin ligase COP1 is a critical negative regulator of p53. Nature 2004; 429:86–92. 208. Chen D, Kon N, Li M, Zhang W, Qin J, Gu W. ARF-BP1/Mule is a critical mediator of the ARF tumor suppressor. Cell 2005; 121: 1071–1083. 209. Lai Z, Ferry KV, Diamond MA, et al. Human mdm2 mediates multiple mono-ubiquitination of p53 by a mechanism requiring enzyme isomerization. J Biol Chem 2001; 276:31357–31367.


55

210. Yang Y, Ludwig RL, Jensen JP, et al. Small molecule inhibitors of HDM2 ubiquitin ligase activity stabilize and activate p53 in cells. Cancer Cell 2005; 7:547–559. 211. Davydov IV, Woods D, Safiran YJ, et al. Assay for ubiquitin ligase activity: high-throughput screen for inhibitors of HDM2. J Biomol Screen 2004; 9:695–703. 212. Blagosklonny MV, Pardee AB. Exploiting cancer cell cycling for selective protection of normal cells. Cancer Res 2001; 61:4301–4305. 213. Blagosklonny MV. Sequential activation and inactivation of G2 checkpoints for selective killing of p53-deficient cells by microtubule-active drugs. Oncogene 2002; 21:6249–6254. 214. Blagosklonny MV, Robey R, Bates S, Fojo T. Pretreatment with DNAdamaging agents permits selective killing of checkpoint-deficient cells by microtubule-active drugs. J Clin Invest 2000; 105:533–539. 215. Carvajal D, Tovar C, Yang H, Vu BT, Heimbrook DC, Vassilev LT. Activation of p53 by MDM2 antagonists can protect proliferating cells from mitotic inhibitors. Cancer Res 2005; 65:1918–1924. 216. Lakkaraju A, Dubinsky JM, Low WC, Rahman YE. Neurons are protected from excitotoxic death by p53 antisense oligonucleotides delivered in anionic liposomes. J Biol Chem 2001; 276:32000–32007. 217. Sellers WR, Fisher DE. Apoptosis and cancer drug targeting. J Clin Invest 1999; 104:1655–1661. 218. Blagosklonny MV. P53: an ubiquitous target of anticancer drugs. Int J Cancer 2002; 98:161–166. 219. Reed JC. Apoptosis-based therapies. Nat Rev Drug Design 2002; 1: 111–121. 220. Pruschy M, Rocha S, Zaugg K, et al. Key targets for the execution of radiation-induced tumor cell apoptosis: the role of p53 and caspases. Int J Radiat Oncol Biol Phys 2001; 49:561–567. 221. Brown JM, Wouters BG. Apoptosis, p53, and tumor cell sensitivity to anticancer agents. Cancer Res 1999; 59:1391–1399. 222. Pirollo KF, Bouker KB, Chang EH. Does p53 status influence tumor response to anticancer therapies? Anticancer Drugs 2000; 11:419–432. 223. Johnstone RW, Ruefli AA, Lowe SW. Apoptosis: a link between cancer genetics and chemotherapy. Cell 2002; 108:153–164. 224. Komarova EA, Gudkov AV. Could p53 be a target for therapeutic suppression? Semin Cancer Biol 1998; 8:389–400. 225. Komarov PG, Komarova EA, Kondratov RV, et al. A chemical inhibitor of p53 that protects mice from the side effects of cancer therapy. Science 1999; 285:1733–1737. 226. Komarova EA, Gudkov AV. Chemoprotection from p53-dependent apoptosis: potential clinical applications of the p53 inhibitors. Biochem Pharmacol 2001; 62:657–667.

56

Klein and Vassilev

227. Komarova EA, Gudkov AV. Suppression of p53: a new approach to overcome side effects of antitumor therapy. Biochemistry (Mosc) 2000; 65:41–48. 228. Wiesmuller L. The tumor suppressor p53 in the center of a strategy aiming at the alleviation of side effects in cancer therapies. Angewandte Chemie-International Edition 2000; 39:1768. 229. Bassi L, Carloni M, Fonti E, Palma de la Pena N, Meschini R, Palitti F. Pifithrin-alpha, an inhibitor of p53, enhances the genetic instability induced by etoposide (VP16) in human lymphoblastoid cells treated in vitro. Mutat Res 2002; 499:163–176. 230. Culmsee C, Zhu X, Yu QS, et al. A synthetic inhibitor of p53 protects neurons against death induced by ischemic and excitotoxic insults, and amyloid beta-peptide. J Neurochem 2001; 77:220–228. 231. Gudkov A, Komarov P, Komarova E. p53 inhibitors and therapeutic use of the same. Patentschrift 2000:PCT/US00/02104 beruhend auf WO00/44364. 232. Gudkov AV, Komarova EA. Prospective therapeutic applications of p53 inhibitors. Biochem Biophys Res Commun 2005; 331:726–736. 233. Strom E, Sathe S, Komarov PG, et al. Small-molecule inhibitor of p53 binding to mitochondria protects mice from gamma radiation. Nat Chem Biol 2006; 2:474–479.

3

DNA Topoisomerases as Targets for the Chemotherapeutic Treatment of Cancer Ryan P. Bender and Neil Osheroff CONTENTS Introduction DNA Topology and Topoisomerases Topoisomerase-Targeted Anticancer Drugs Other Topoisomerase Poisons Topoisomerase II and Leukemia Checkpoint Responses and Repair of Topoisomerase-Mediated DNA Damage Summary

Abstract DNA topoisomerases are ubiquitous enzymes that play important roles in a variety of critical nuclear processes. These enzymes control the topological structure of DNA in the cell, regulate under- and overwinding of the double helix, and remove knots and tangles from the genetic material. In order to perform their critical cellular From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

57

58

Bender and Osheroff

functions, topoisomerases generate DNA strand breaks as requisite intermediates in their catalytic cycles. Consequently, while these enzymes are essential to cell survival, they also have the potential to fragment the genome. The DNA cleavage activity of topoisomerases has been exploited to generate some of the most important drugs currently used to treat human malignancies. This article will familiarize the reader with many aspects of topoisomerase enzymology and cell biology, their interactions with anticancer drugs, and the cellular consequences of topoisomerase-mediated DNA strand breaks. It also will discuss the checkpoint functions and repair pathways that are triggered in response to topoisomerase-generated DNA damage. Key Words: Topoisomerase II; topoisomerase II; topoisomerase II; topoisomerase II poisons; anticancer drugs; cancer chemotherapy; genotoxicity; cancer; leukemia; DNA; DNA supercoiling; DNA lesion

1. INTRODUCTION Malignant cells are frequently distinguished by rapid growth coupled with an impaired ability to activate cell cycle checkpoints and DNA repair pathways (1,2). Consequently, DNA in cancerous tissues often sustains elevated rates of replication and transcription, despite a decreased competence to restore genomic integrity following damage. This dual property of high DNA metabolism and low genetic stability makes the double helix an attractive target for cancer chemotherapy. Indeed, several classes of widely used anticancer drugs act by damaging DNA, either directly or indirectly (3). Multiple therapeutic strategies are used to damage DNA. Ultimately, these strategies block DNA replication or other essential nucleic acid processes, generate mutations, create DNA strand breaks, or induce gross chromosomal abnormalities (3). For example, methotrexate, which inhibits dihydrofolate reductase, decreases cellular thymine pools and promotes incorporation of deoxyuridine into chromosomes (4). The unnatural nucleotide AraC is incorporated into the genetic material and impairs DNA replication (5). Mechlorethamine (nitrogen mustard) (6) and cisplatin (7) alkylate bases or crosslink the 2 strands of the double helix. Finally, bleomycin (8) generates DNA strand breaks by a chemical mechanism and etoposide (9) does so by altering the activity of topoisomerase II. Even when damage induced by chemotherapeutic drugs is moderate, catastrophic events can occur if DNA or RNA polymerases traverse

Chapter 3 / DNA Topoisomerases

59

the damage or if the cell enters mitosis with conjoined or broken chromosomes (2,10). To prevent these events from occurring, cells activate pathways known as checkpoints (1,10,11). When activated, checkpoints halt cell cycle progression and allow DNA damage to be repaired in the absence of replication or cellular division. Thus, all things being equal, malignant cells with impaired checkpoint responses are more sensitive to chemotherapeutic agents that damage the genetic material (1,2). Some of the most effective anticancer drugs currently in clinical use damage DNA by targeting topoisomerases (3). These agents act in an insidious manner and convert topoisomerases from essential enzymes to potent cellular toxins that fragment the genome. Therefore, this article will focus on mammalian topoisomerases and the mechanism by which anticancer drugs exploit the natural activity of these enzymes to kill malignant cells.

2. DNA TOPOLOGY AND TOPOISOMERASES DNA is essentially an extremely long double-stranded rope in which the 2 strands are interwound about one another (12). As a result, the topological properties of the genetic material profoundly influence virtually every major DNA process. DNA is globally unwound (i.e., negatively supercoiled) in all vertebrate species (13–15). This unwinding makes it easier to separate complementary DNA strands from one another and therefore greatly facilitates replication and transcription. Once the replication or transcription machinery begins to travel along the DNA template, however, deleterious effects of topology are manifested. Because helicases separate, but do not unwind the 2 strands of the double helix, fork progression results in acute overwinding (i.e., positive supercoiling) of the DNA ahead of the tracking systems (13,15–17). In contrast to underwinding, overwinding dramatically increases the difficulty of opening the double helix. Therefore, the accumulation of positive supercoils represents a formidable block to all DNA processes that require strand separation (17–20). The effects of DNA topology are further compounded by the extreme length of the double helix. The genetic material from a single human cell, which approaches 2 meters in length, exists in a nucleus that is only 5–10 microns in diameter. Consequently, the double helix is subjected to the same forces and constraints as a room tightly packed from floor to ceiling with rope. Nuclear processes such as

60

Bender and Osheroff

recombination and replication naturally generate knots and tangles in DNA, respectively (21,22). If knots accumulate in the genome, DNA tracking systems are unable to separate the 2 strands of the double helix (13,15,17,21). Moreover, if tangled (i.e., catenated) daughter chromosomes are not separated before cell division, cells will die of mitotic failure (16,17,23–25). The topological state of the double helix in eukaryotic cells is regulated by ubiquitous enzymes known as topoisomerases, which act by creating transient breaks in the double helix (16,17,23–27). To maintain genomic integrity during this process, topoisomerases form covalent bonds between active site tyrosyl residues and the phosphate moieties of newly generated DNA termini (16,17,23–30). These covalent protein-cleaved DNA complexes are referred to as cleavage complexes. There are 2 classes of topoisomerases that are distinguished by the number of DNA strands that they cleave during their catalytic cycles.

2.1. Type I Topoisomerases Type I topoisomerases act by generating a transient single-stranded break in the double helix followed by a single-stranded DNA passage event or controlled rotation about the break (24,27,31,32). As a result, these enzymes are able to alleviate torsional stress (i.e., remove superhelical twists) in duplex DNA and resolve single-stranded hemicatenated recombination intermediates. Type I topoisomerases are involved in all DNA processes that involve tracking systems and play important roles in maintaining genomic integrity (16,27,29,32). There are 2 subclasses of type I enzymes. Type IA enzymes (topoisomerase III and topoisomerase III) are homologous to bacterial topoisomerase I ( protein) (33). They link to the 5´terminal phosphate of the DNA during scission and use a singlestrand passage mechanism (17,24). The physiological functions of these enzymes are not well understood. However, in concert with recQ helicases, they appear to play important roles in resolving double Holliday junctions that arise during a variety of recombination pathways (34–36). Although topoisomerase III and topoisomerase III are fascinating enzymes in their own right, they are not targets for any known anticancer drugs. Therefore, they will not be discussed further in this article. Additional information on these enzymes can be found in several reviews (17,24,34,36). The type IB enzyme, topoisomerase I, is distinct from the type IA enzymes. It links to the 3 -terminal phosphate during scission


61

and modulates DNA under- and overwinding by a controlled rotation mechanism (24,27,31,37). As a result, topoisomerase I is important for maintaining appropriate global levels of DNA supercoiling in the cell and alleviating torsional stress that accumulates ahead of nucleic acid tracking systems, especially transcription and replication complexes (24,27). Topoisomerase I is a nuclear enzyme that is enriched in the nucleolus, reflecting a heavy involvement in the synthesis of ribosomal RNAs (24,27). Topoisomerase I was the first eukaryotic topoisomerase to be isolated (37) and is well characterized, both structurally and enzymologically (24,27,31,32). The enzyme carries out its complete DNA nicking, rotation, and ligation cycle without the need for a high-energy cofactor, such as ATP (38). In addition, whereas the activity of topoisomerase I is stimulated by the presence of divalent cations, such as Mg(II) or Co(II), it does not require them (39,40). Topoisomerase I appears to be expressed constitutively in all cell types, irrespective of proliferative status, and protein levels remain constant over the cell cycle (41). The enzyme is not essential at the cellular level, but is required for proper development (24,27,32,42,43). There is evidence that in some cases, it is expressed at higher levels in malignant cells (41). Mammalian cells encode a second type IB topoisomerase, mttopoisomerase I, that is targeted to the mitochondria (24,27,44). This recently discovered enzyme is involved in the synthesis of the mitochondrial genome.

2.2. Type II Topoisomerases Mammalian type II topoisomerases are homodimeric enzymes that interconvert different topological forms of DNA by a double-stranded DNA passage mechanism (16,17,24,26,28,29). Briefly, these enzymes: 1) bind 2 separate segments of DNA, 2) create a double-stranded break in one of the segments, 3) translocate the second DNA segment through the cleaved nucleic acid “gate,” 4) rejoin (i.e., ligate) the cleaved DNA, and 5) release the translocated segment through a gate generated at an interface between the 2 protein subunits (16,17,24,26,28,29,45). The 2 scissile bonds that are cut by type II topoisomerases are staggered and located across the major groove from one another (16,17,24,26,29). Thus, these enzymes generate cleaved DNA molecules that contain 4-base single-stranded cohesive ends at their 5 -termini (46,47). During the cleavage event, all known type II topoisomerases covalently attach to the newly generated 5 -DNA termini.

62

Bender and Osheroff

In contrast to topoisomerase I, type II enzymes require 2 cofactors to carry out the catalytic double-stranded DNA passage reaction. First, they need a divalent cation for all steps beyond enzyme-DNA binding. Mg(II) appears to be used in vivo (46–50). Second, they use the energy of ATP to drive the overall DNA strand passage reaction (48,51–53). As a result of their double-stranded DNA passage reaction, type II enzymes are able to remove superhelical twists from DNA and resolve knotted or tangled duplex molecules (16,17,24,26,29). They function in numerous DNA processes and are required for recombination, the separation of daughter chromosomes, and proper chromosome structure, condensation, and decondensation (16,17,24,26,29). Mammals express 2 distinct isoforms of the type II enzyme, topoisomerase II and topoisomerase II (24,26,54,55). These isoforms display a high degree (∼70%) of amino acid sequence identity and similar enzymological characteristics, but differ in their protomer molecular masses (170 vs. 180 kDa, respectively) and are encoded by separate genes (17,24–26,54–59). Topoisomerase II and topoisomerase II are both nuclear enzymes (mammals have no known mitochondrial type II enzymes), but have distinct patterns of expression and cellular functions. Topoisomerase II is essential for the survival of actively growing cells and its concentration is upregulated dramatically during periods of proliferation (60–63). Furthermore, enzyme levels increase over the cell cycle and peak in G2/M (62,64,65). Topoisomerase II is found at replication forks and remains tightly associated with chromosomes during mitosis (16,63,66,67). Thus, it is believed to be the isoform that functions in growth-dependent processes, such as DNA replication and chromosome segregation (16,17). In contrast to topoisomerase II, topoisomerase II is dispensable at the cellular level (68). However, the isoform is required for proper neural development (69). Expression of topoisomerase II is independent of proliferative status and cell cycle, and the enzyme dissociates from chromosomes during mitosis (16,59,62,70). Topoisomerase II cannot compensate for the loss of topoisomerase II in mammalian cells, suggesting that these 2 isoforms do not play redundant roles in replicative processes (59,63,71,72). Although the physiological functions of topoisomerase II have yet to be defined, recent evidence indicates roles in the transcription of hormonally- or developmentally-regulated genes (73,74).


63

3. TOPOISOMERASE-TARGETED ANTICANCER DRUGS As discussed above, all topoisomerases cleave DNA during their critical cellular functions. Although the strand breaks generated by these enzymes are transient in nature, they are potentially deleterious to the cell (Fig. 1). When a nucleic acid tracking system, such as a replication or transcription complex, attempts to traverse a topoisomeraseDNA cleavage complex, it converts the transient enzyme-DNA interaction to a permanent double-stranded break (9,16,26,29). The resulting strand breaks, as well as the inhibition of essential DNA processes, initiate replication restart (which generates additional DNA breaks) and recombination/repair pathways. If the accumulation of strand breaks overwhelms the cell, they trigger apoptotic pathways (25,75–79). However, if the DNA strand breaks are not able to overwhelm the cell, surviving populations may contain chromosomal translocations or other aberrations (25,75–79). Agents that increase levels of topoisomerase-DNA cleavage complexes are known as “topoisomerase poisons” because they convert these enzymes to potent cellular toxins (9,25,27,80–84). Topoisomerase poisons work by 2 nonmutually exclusive mechanisms. Some poisons act by inhibiting the ability of their topoisomerase target to ligate cleaved DNA intermediates (9,25,27,32,81,82,85–87). Other poisons have little effect on the rate of enzyme-mediated ligation and are believed to act primarily by enhancing the forward rate of cleavage complex formation (9,25,88–92). The exact mechanism by which this second group of drugs increases levels of DNA cleavage is unknown. They may specifically affect the forward rate of DNA scission. Alternatively, they may affect the enzyme-DNA binding equilibrium, as the level of topoisomerase-mediated DNA cleavage is proportional to the amount of enzyme bound. Clinically relevant topoisomerasetargeted agents appear to act primarily by inhibiting DNA ligation (9,25,27,81,82,86). A diverse group of natural and synthetic compounds have been found to increase levels of topoisomerase-DNA cleavage complexes in vitro and in human cells. Several of these agents are in wide clinical use as anticancer therapeutics and represent some of the most successful drugs currently used for the treatment of human malignancies (3).

3.1. Topoisomerase I-Targeted Drugs Topoisomerase I is the target for an emerging class of drugs based on camptothecin, a natural product derived from the bark of the Chinese yew tree, Camptotheca acuminata (Fig. 2) (93). These drugs represent

64

Bender and Osheroff

Topoisomerase I

Topoisomerase II

Transient Cleavage Complexes

Transient Cleavage Complexes Topoisomerase Poison

Increased Cleavage Complexes

Increased Cleavage Complexes Replication or Transcription Machinery

Permanent DSB

Permanent DSB

Arrest and Repair

Cellular Recovery ?? Stable 11q23 Translocations (Leukemia) ?? Apoptosis

Apoptosis

Cell Death

Fig. 1. Topoisomerases as cellular toxins. Levels of transient DNA cleavage complexes generated by topoisomerase I or topoisomerase II are increased following exposure to a variety of topoisomerase poisons. These cleavage complexes are converted to permanent DNA strand breaks (DSB) as a result of collisions with replication or transcription complexes. If checkpoints and DNA repair pathways function properly and if levels of DSB do not overwhelm the cell, DNA damage is repaired, resulting in cellular recovery. If checkpoints or DNA repair pathways are impaired or if levels of DSB overwhelm the cell, apoptotic pathways are initiated, resulting in cell death. Finally, in some cases,


65

Camptothecins

A

O

O

B CN N D

N

OH

N

E

O

O

O

OH

OH

Camptothecin (closed lactone ring form)

+

Na

O

Camptothecin (open ring form)

N N N

O

F

O

HO

N O

N

N

O

O

OH

O

OH

O

Topotecan

Diflomotecan

Non-camptothecins

O

N

N

O

Irinotecan/CPT-11

O

N F

O OH

O

O N

F

F

O N N O OH

HO HO

H3CO

O

H3CO F

Indolocarbazoles

O

O

N

N

H3CO

R

O

O

Phenanthradines

O

H3CO

5,11-Diketoindenoisoquinolines

Fig. 2. Selected topoisomerase I-targeted anticancer drugs.

some of the most active new agents in the clinic and show promise against malignancies that respond poorly to existing therapies, such as non-small cell lung cancer, metastatic ovarian cancer, and colorectal cancer (3,27,32,94). The parental compound, camptothecin, is clinically problematic for 2 reasons. First, the active “ring closed lactone form” of the drug Fig. 1. (Continued) treatment of cells with topoisomerase II poisons induces the formation of stable leukemic translocations involving the MLL gene at chromosomal band 11q23. The source of the DNA breaks that ultimately lead to the translocations is controversial and is postulated to result from DNA cleavage mediated by topoisomerase II, apoptotic nucleases (coupled with abortive apoptosis), or both.

66

Bender and Osheroff

(see the E ring in Fig. 2) is rapidly and reversibly converted to the inactive “ring open form” under physiological conditions (27). Second, camptothecin readily binds human serum albumin, making it inaccessible for cellular uptake (27,95,96). Subsequent generations of drugs, such as topotecan and irinotecan (Fig. 2), have a more stable lactone ring and display decreased albumin binding (27). A new generation of camptothecin-like drugs, called homocamptothecins (Fig. 2), currently is doing well in clinical trials. These drugs are as active as the camptothecins, but contain a 7-member E ring (as opposed to the 6-member ring of camptothecin) (97,98). The ring-closed lactone form of the homocamptothecins has a longer half-life in blood than the camptothecins, and ring opening is essentially irreversible (97,98). Consequently, the pharmacokinetics of homocamptothecins are more predictable than those of camptothecins (97,98). These attributes make homocamptothecins much easier to schedule and dramatically reduce inter-patient variability. The lead compound in this drug class is diflomotecan (97,98). Other drugs that target topoisomerase I, including the indolocarbazoles (99), the 5,11-diketoindenoisoquinolines (100), and the phenanthridines (101) (Fig. 2), currently are under clinical development. These drugs are not encumbered by the presence of the unstable lactone ring and appear to induce very stable cleavage complexes (27). Thus, they have clinical advantages over the camptothecins.

3.2. Topoisomerase II-Targeted Drugs Topoisomerase II is the target for some of the most successful anticancer drugs currently used to treat human malignancies (102). It is estimated that one half of all chemotherapy regimens include these agents (102). Six topoisomerase II-targeted drugs are approved for use in the United States, with others being used worldwide (3,102,103). Several of these agents are shown in Fig. 3. One of the first topoisomerase II-targeted agents to be discovered was etoposide, which is derived from podophyllotoxin (9,103). This natural product is found in Podophyllum peltatum, more commonly known as the May apple or mandrake plant (9,103). Podophyllotoxin has been used as a folk remedy for over a thousand years (9,103). It has well-established antimitotic properties, which are related to its potent inhibition of tubulin polymerization (9,103). Although clinical use of podophyllotoxin as an antineoplastic agent was prevented by high toxicity, 2 synthetic analogs, etoposide and teniposide, were developed

Chapter 3 / DNA Topoisomerases H

OH

O

R

67

HN(CH2)2NH(CH2)2OH O

O

O

OH

C

O O OH

R2

OH

HO O

H

OH

O

HN(CH2)2NH(CH2)2OH R1

O O O

O

OH

Mitoxantrone

H

O

H3C

O

O

+

NH3 NHSO2 CH3

OH

OCH3

H3CO OH

R= H3 C

CH3 O

= Etoposide

NH

= Teniposide S

N

R1 =

R2 =

CH2OH

OCH3

= Doxorubicin

CH3

OCH3

= Daunorubicin

CH2OH

H

= Idarubicin

Amsacrine

Fig. 3. Selected topoisomerase II-targeted anticancer drugs.

(9,103). These analogs display increased antineoplastic activity and decreased toxicity as compared to the parent compound. Etoposide was approved for clinical use against cancer in the mid-1980s (9,103). Surprisingly (and in marked contrast to podophyllotoxin), etoposide displays no ability to inhibit tubulin polymerization. Rather, it kills cells by acting as a topoisomerase II poison (9,103–107). Etoposide and other drugs such as doxorubicin are front-line therapy for a variety of systemic cancers and solid tumors, including leukemias, lymphomas, sarcomas, and breast, lung, and germline cancers (3). Every form of cancer that is considered to be curable by systemic chemotherapy uses drugs that target topoisomerase II in treatment regimens (108). Although topoisomerase II is believed to be the cytotoxic target of the drugs shown in Fig. 3, the relative contributions of topoisomerase II and topoisomerase II to the chemotherapeutic effects of these agents has yet to be resolved. Some drugs appear to favor one isoform or the other; however, no truly “isoform-specific” agents have been identified. The issue of isoform specificity has potential clinical ramifications. For example, because topoisomerase II is present in all cell types, it may be responsible for mediating some of the toxic side effects of topoisomerase II poisons in nonmalignant tissues (109). Alternatively, because topoisomerase II and topoisomerase II are involved in different cellular processes, it may be that cleavage complexes formed with one or the other isoform are more likely to be converted to a permanent DNA strand break.

68

Bender and Osheroff

4. OTHER TOPOISOMERASE POISONS In addition to the synthetically derived topoisomerase poisons that are used to treat cancer [and antibacterial quinolones that target the prokaryotic type II enzymes, DNA gyrase and topoisomerase IV (110–112)], 3 other categories of topoisomerase poisons have been characterized. These include natural products that are normal dietary components (bioflavonoids), toxic metabolites of drugs or industrial chemicals (quinones), and DNA-damaging agents (Fig. 4). Thus far, compounds in the first 2 categories have been found to affect primarily the type II enzyme. However, DNA damaging agents increase cleavage mediated by both type I and II topoisomerases.

4.1. Bioflavonoids The most prominent natural products with activity against mammalian topoisomerases are the bioflavonoids (i.e., phytoestrogens) (113–116) (Fig. 4). Bioflavonoids represent a diverse group of polyphenolic compounds that are components of many fruits, vegetables, and plant leaves (117–120). These compounds affect human cells through a variety of pathways; they are strong antioxidants and efficient Quinone Metabolites

Bioflavanoids OH HO

OH

O HO

O

O

O Cl

OH

O

OH

OH

Genistein

Quercetin

O N

HO

CH2 H

O N

N

N

NH2 HO H

OH

DNA Base Oxidation (8-oxo-dG)

OH

O

H H

OH

PCB Quinones (4’Cl-2,5pQ)

DNA Damage

O

H

Benzoquinone

NH

O N

O

O

OHO

H OH

Abasic Site

N H

O

H

H OH

HO

N N

H

H

OH

OH

H

H

DNA Base Alkylation (εdG)

Fig. 4. Other topoisomerase poisons. Structures are shown for selected bioflavonoids, quinones, and DNA lesions.


69

inhibitors of growth factor receptor tyrosine kinases (117–120). In addition, they are potent topoisomerase II poisons (113–116). It has been suggested that genistein (an isoflavone that is abundant in soy) has chemopreventative properties, and that ingestion of this compound contributes to the low incidence of breast and colorectal cancers observed in the Pacific rim (117–120). However, as discussed below, there also is evidence associating genistein consumption during pregnancy to the development of infant leukemias (114,121–124).

4.2. Quinones Quinones are highly reactive compounds that are often produced in the body as a result of detoxification or metabolism pathways. Quinones damage cells by generating oxidative radicals and by covalently modifying proteins and (to a lesser extent) nucleic acids (125–128). Several quinone-based compounds also are strong topoisomerase II poisons. Among these compounds are N-acetyl p-benzoquinone imine, the toxic metabolite of acetaminophen [the most widely used analgesic in the world (129)] (130), 1,4-benzoquinone [a reactive metabolite of benzene (131)] (132,133), and a variety of PCB (polychlorinated biphenyl) quinone metabolites (134) (Fig. 4). Menadione (the quinone known as vitamin K3) also displays activity against topoisomerase II (135). Quinones are unique among characterized topoisomerase II poisons, in that their activity requires covalent attachment to the enzyme (130,132–135). The detailed mechanism of quinone action has yet to be determined. However, studies suggest that quinones increase levels of topoisomerase II-DNA cleavage complexes by multiple effects on the enzyme, including crosslinking the N-terminal protein gate of topoisomerase II and inhibiting rates of enzyme-mediated DNA ligation (130,134).

4.3. DNA Damage Unlike the 2 categories of topoisomerase II poisons discussed above, DNA lesions are potent enhancers of DNA scission mediated by both the type I and II enzymes (Fig. 4). Topoisomerase I is most sensitive to abasic sites, oxidative lesions, and alkylated bases (136–139). Topoisomerase II prefers lesions that distort the double helix, and is particularly sensitive to abasic sites and alkylated bases that contain exocyclic rings (89,90,92,140–142).

70

Bender and Osheroff

DNA damage increases cleavage at naturally occurring sites of topoisomerase I or topoisomerase II action (89–92,136,138,140–143). In all cases, lesions must be located proximal to the sites of cleavage to act as enzyme poisons (89–92,136,138,140–143). Whereas topoisomerase I is generally sensitive to lesions immediately upstream or downstream from the scissile bond (138,143), topoisomerase II requires that damage be localized within the 4-base stagger that separates the 2 scissile bonds on the opposite strands of the double helix (89–92,140–142). Although DNA damage increases levels of topoisomerase I and topoisomerase II cleavage complexes, the mechanism by which lesions alter the activity of these enzymes differs. DNA damage increases the concentration of topoisomerase I-DNA cleavage complexes primarily by inhibiting rates of enzyme-mediated DNA ligation (136–140). In contrast, damage has no obvious effects on rates of topoisomerase IImediated DNA ligation and appears to act primarily by enhancing the forward rate of scission (89–92,140,141). The physiological benefits of DNA lesions as topoisomerase poisons, if any, are unclear. However, it is notable that topoisomerase I and topoisomerase II both appear to play roles in fragmenting genomic DNA during apoptosis (143–146). It has been suggested that the apoptotic activities of topoisomerases are enhanced (or perhaps triggered) by DNA lesions that are generated following the release of oxidative radicals from permeable mitochondria in apoptotic cells (143–146).

5. TOPOISOMERASE II AND LEUKEMIA In addition to its role as an essential cellular protein and target for anticancer drugs, clinical studies suggest that topoisomerase II initiates chromosomal translocations that lead to specific types of leukemia (see Fig. 1). For example, ∼2–3% of patients treated with regimens that include etoposide ultimately develop acute myelocytic leukemia (78,147–149). Recently, correlations between the rising use of mitoxantrone to treat breast cancer and the development of secondary leukemias also have been noted (150). The common feature in ∼50% of these leukemias is the presence of translocations within an 8.3 kb breakpoint cluster region in the MLL (mixed lineage leukemia) gene at chromosome band 11q23 (77,78,147–149). The basis for the development of topoisomerase II-initiated leukemias has not been elucidated, but it appears to be related to


71

the function of the protein product of the MLL gene. MLL is the human homolog of the Drosophila trithorax and yeast Set1 proteins, and is a histone methyltransferase that is involved in transcriptional regulation in hematopoietic cells (149,151–154). Accumulating evidence suggests that the fusion of the MLL protein with other cellular partner proteins alters MLL function and affects the differentiation of pluripotent hematopoietic stem cells or committed myeloid or lymphoid stem cells by deregulating the expression of the HOX gene (149,151,152,154,155). Infant acute lymphoblastic leukemias and some benzene-induced leukemias also display translocations involving chromosomal band 11q23 (156,157). Epidemiological and biochemical studies have established potential links between these malignancies and topoisomerase II. For example, the maternal consumption (during pregnancy) of foods that are high in genistein or other naturally occurring topoisomerase II poisons increases the risk of developing infant acute lymphoblastic leukemias ∼3-fold (114,121–124). In addition, individuals with chronic exposure to benzene display an increasingly higher risk for leukemias with 11q23 chromosomal translocations if they are heterozygous or homozygous for the C609T polymorphism of the NAD(P)H:quinone oxidoreductase 1 (NQO1) (158–161). NQO1 is the enzyme that reduces 1,4-benzoquinone (a highly active topoisomerase II poison) to the less reactive 1,4-hydroquinone (133,158,159,161–163). Although the involvement of topoisomerase II-mediated DNA cleavage in the development of leukemias with MLL translocations is widely accepted, the role of the enzyme-associated DNA strand breaks in triggering the chromosomal aberrations is controversial. Two hypotheses have been proposed (see Fig. 1). The first postulates that the breaks induced by topoisomerase II play a direct role in the translocation process (77,149,156,164). In this case, the enzyme cleaves within the MLL gene and following processing and recombination/repair, the breaks are reattached to other sites in the genome (presumably that also were cleaved by the type II enzyme). Supporting this hypothesis, all MLL (and partner) chromosomal breakpoints identified in patient samples, including those with secondary and infant leukemias, are located in close proximity to in vitro sites of topoisomerase II-mediated DNA cleavage (165–167). Furthermore, leukemias with 11q23 chromosomal translocations are only observed in patients treated with topoisomerase II poisons, and are not seen following other anticancer therapies (78,147,168,169). This is despite

72

Bender and Osheroff

the fact that radiation, DNA alkylation and crosslinking agents, and drugs such as bleomycin that chemically cleave DNA all generate chromosomal breaks and induce apoptotic pathways. The second hypothesis states that the breaks induced by topoisomerase II play an indirect role in the translocation process. In this case, enzyme-mediated DNA cleavage induces apoptosis, which initiates chromosomal fragmentation (164,170,171). Occasionally (by processes that have yet to be described), apoptosis aborts and nuclease-generated breaks within the MLL gene are processed and reattached to other sites in the genome (172–174). Supporting this theory, a major site of apoptotic cleavage is located in the breakpoint cluster region of the MLL gene (170,172–174). Moreover, translocations involving 11q23 can be induced in cultured human cells by agents that trigger apoptosis, but do not target topoisomerase II (170,172–174). The discrepancy between patients and cellular studies regarding the requirement for topoisomerase II poisons to induce leukemic translocations is notable. The apoptotic model reconciles this discrepancy by further postulating that the inhibition of topoisomerase II function following treatment with poisons alters chromatin structure and that these alterations, coupled with nuclease action, are required for the translocation event in patients (170). It is likely that the process that translates topoisomerase II-mediated DNA cleavage into 11q23 chromosomal translocations is highly complex and multifaceted. Both (or neither) of the above hypotheses may contribute to the process. Clearly, this is an area of considerable clinical relevance that deserves greater attention and future studies.

6. CHECKPOINT RESPONSES AND REPAIR OF TOPOISOMERASE-MEDIATED DNA DAMAGE 6.1. Checkpoint Responses If DNA damage is not repaired, attempts to continue ongoing nuclear processes can have catastrophic consequences. For example, continued replication of chromosomes that contain double-stranded DNA breaks can trigger cell death pathways or lead to the loss of acentromeric fragments of the genome during mitosis. Therefore, in response to DNA damage, cycling cells induce kinase cascades that halt the cell cycle and provide a window in which the damage can be repaired. Because these cascades prevent the cell from incurring further injury, they are said to function as checkpoints.


73

There are 2 major kinases that are activated in response to DNA damage that occurs during S phase, ATM and ATR. ATM is activated primarily in response to double-stranded DNA breaks, such as those created by exposure to ionizing radiation (175–178). When switched on, ATM phosphorylates Chk2 (among other proteins), which in turn phosphorylates a host of target proteins involved in cell cycle progression and DNA repair (175,176,179). ATR is activated primarily in response to long patches of RPA-coated single-stranded DNA (176,180–183). Typically, these single-stranded regions are generated when replication forks are blocked by damaged DNA and helicases are uncoupled from the replicative machinery (182–185). When switched on, ATR phosphorylates Chk1 (among other proteins), which also targets a number of regulatory and repair proteins (175,185–187). It should be noted that there is considerable cross-talk between the downstream targets of ATM and ATR (185,187). Finally, ATM is not an essential enzyme. However, mutations in this protein cause ataxia telangiectasia, a pleiotropic autosomal recessive disorder that results in a profound cancer predisposition and radiosensitivity (188–190). ATR, in contrast, is essential for cell survival (191,192). Upon treatment with either topoisomerase I- or topoisomerase IItargeted drugs, the cell cycle arrests in an ATR-dependent fashion (193–197). This is a surprising finding, in light of the fact that topoisomerase poisons generate double-stranded DNA breaks, which would presumably elicit an ATM-dependent response. Three implications can be drawn from this discovery. First, the major pathway by which topoisomerase poisons kill cells may be the inhibition of major nuclear process, such as replication, rather than the generation of DNA strand breaks per se. Second, the double-stranded DNA breaks that result from drug treatment may be created indirectly by replication restart pathways, rather than directly by disruption of cleavage complexes. Third, because the termini of topoisomerase-cleaved DNA are protein-associated, double-stranded breaks that are generated by disruption of cleavage complexes may not be recognized as “genuine” strand breaks by ATM or other repair proteins. In this latter regard, it is notable that Ku and the catalytic subunit of DNA-dependent protein kinase (DNA-PKcs), proteins that interact with DNA termini during nonhomologous end joining, do not bind to disrupted topoisomerase II-DNA cleavage complexes unless the bulk of the terminally bound protein is first removed (198).

74

Bender and Osheroff

6.2. Processing Topoisomerases from DNA Termini Before DNA termini generated by topoisomerase-mediated DNA cleavage can be rejoined, or potentially even recognized by repair proteins, topoisomerases must be proteolyzed and/or removed from the DNA. Three distinct processes appear to be involved. First, considerable circumstantial evidence suggests that both topoisomerase I and topoisomerase II can be removed from DNA termini by the actions of endonucleases. The most direct studies point to a role for the Mre11 nuclease, which functions together with Rad50 and Nbs1 in the first stages of DNA recombination (199,200). Although other nucleases have been implicated, it is not clear whether they actually remove topoisomerases from cleaved nucleic acid chains or are involved in downstream recombination events (201). Second, before cleavage complexes can be recognized by repair proteins, the bulk of topoisomerase I or topoisomerase II needs to be removed by proteolysis (202). In this regard, all evidence points to an important role for ubiquitination followed by the actions of the 26S proteosome (202,203). Proteolysis leaves a short polypeptide attached to the DNA termini that is small enough to allow ring proteins such as Ku or DNA-PKcs to slip onto the nucleic acid end (198,204). As discussed below, it also allows the terminal protein-DNA complex to be processed by tyrosyl-DNA phosphodiesterase 1 (Tdp1). Third, proteolyzed topoisomerases can be removed from DNA termini by the actions of Tdp1 (205). This enzyme contains an active site tyrosyl residue that displaces the topoisomerase tyrosyl residue that is covalently linked to the terminal DNA phosphate moiety (206–208). Tdp1 is then removed from the DNA by hydrolysis and recycled (209,210). The enzyme was originally proposed to act only on the 3 protein-DNA linkage generated by topoisomerase I-mediated cleavage (205). However, recent studies potentially extend the range of Tdp1 substrates to the 5 protein-DNA linkage generated by topoisomerase II (206,207). Depletion of Tdp1 from human or yeast cells causes mild hypersensitivity to either camptothecin or etoposide and overexpression confers ∼2–fold resistance to these agents (206,207,211,212). At least in the case of topoisomerase I-generated DNA strand breaks, it appears that Tdp1 is used primarily to remove the enzyme from cleavage complexes stabilized by collisions with transcription complexes rather than DNA replication forks (212). Finally, it is notable that a genetic defect in Tdp1 results in a rare recessive disease known as spinocerebellar ataxia with axonal neuropathy (SCAN1) (210,213). Not only does the SCAN1 Tdp1


75

mutant have impaired activity, but it maintains a covalent bond with the terminal DNA phosphate moiety that is considerably more stable than that of the wild-type enzyme (212). SCAN1 cells display increased sensitivity to camptothecin (211,212).

6.3. Repair of Topoisomerase-Generated DNA Strand Breaks Following processing, double-stranded DNA breaks generated by topoisomerase I or topoisomerase II are repaired by classic recombination pathways (9,138,139,214,215). Both homologous recombination and nonhomologous end joining (NHEJ) have been implicated. It appears that NHEJ is the favored pathway in human cells (215). Collisions between the transcription machinery and topoisomerase I-DNA cleavage complexes often result in the generation of singlestranded DNA breaks (27,139). In these cases, topoisomerase I is removed from the DNA terminus by Tdp1, and the processed nick is repaired by single strand DNA break repair pathways (27,139).

7. SUMMARY Topoisomerase I and topoisomerase II are widely used targets for a variety of successful anticancer drugs. These drugs take advantage of normal catalytic events mediated by topoisomerases and convert them to potent cellular toxins that generate breaks in the genome. The ability of cells to trigger checkpoints and DNA repair pathways in response to topoisomerase-targeted drugs has a major influence on the clinical efficacy of these agents. Therefore, it is important to further our understanding of the relationships between topoisomerasemediated DNA damage and the mechanisms by which cells arrest nuclear processes and restore the integrity of the genetic material after this damage has occurred.

ACKNOWLEDGMENTS Work in the senior author’s laboratory was supported by National Institutes of Health research grants GM33944 and GM53960. RPB was a trainee under National Institutes of Health grant 5 T32 CA09582. We are grateful to O. J. Bandele and J. E. Deweese for critical reading of the manuscript.

76

Bender and Osheroff

REFERENCES 1. Ishikawa K, Ishii H and Saito T. DNA damage-dependent cell cycle checkpoints and genomic stability. DNA Cell Biol 2006;25:406–411. 2. Nakanishi M, Shimada M and Niida H. Genetic instability in cancer cells by impaired cell cycle checkpoints. Cancer Sci 2006;97:984–989. 3. Chabner B and Longo D, eds. Cancer Chemotherapy and Biotherapy: Principles and Practice. 4th ed. Philadelphia: Lippincott; 2006. 4. Schilsky RL. Methotrexate: An Effective Agent for Treating Cancer and Building Careers. The Polyglutamate Era. Oncologist 1996;1:244–247. 5. Miura S, Yoshimura Y, Satoh H and Izuta S. The antitumor mechanism of 1-(2-deoxy-2-fluoro-4-thio-beta-D-arabinofuranosyl)-cytosine: effects of its triphosphate on mammalian DNA polymerases. Jpn J Cancer Res 2001;92:562–567. 6. Povirk LF and Shuker DE. DNA damage and mutagenesis induced by nitrogen mustards. Mutat Res 1994;318:205–226. 7. Zorbas H and Keppler BK. Cisplatin damage: are DNA repair proteins saviors or traitors to the cell? Chembiochem 2005;6:1157–1166. 8. Povirk LF. DNA damage and mutagenesis by radiomimetic DNAcleaving agents: bleomycin, neocarzinostatin and other enediynes. Mutat Res 1996;355:71–89. 9. Baldwin EL and Osheroff N. Etoposide, topoisomerase II and cancer. Curr Med Chem Anti-Canc Agents 2005;5:363–372. 10. Yang XH and Zou L. Checkpoint and coordinated cellular responses to DNA damage. Results Probl Cell Differ 2006;42:65–92. 11. Houtgraaf JH, Versmissen J and van der Giessen WJ. A concise review of DNA damage checkpoints and repair in mammalian cells. Cardiovasc Revasc Med 2006;7:165–172. 12. Watson JD and Crick FHC. Molecular structure of nucleic acids. A structure for deoxyribose nucleic acid. Nature 1953;171:737–738. 13. Cozzarelli NR and Wang JC. DNA Topology and its Biological Effects. Cold Spring Harbor: Cold Spring Harbor Laboratory Press; 1990. 14. Kanaar R and Cozzarelli NR. Roles of supercoiled DNA structure in DNA transactions. Curr Opin Struct Biol 1992;2:369–379. 15. Schvartzman JB and Stasiak A. A topological view of the replicon. EMBO Rep 2004;5:256–261. 16. Nitiss JL. Investigating the biological functions of DNA topoisomerases in eukaryotic cells. Biochim Biophys Acta 1998;1400:63–81. 17. Wang JC. Cellular roles of DNA topoisomerases: a molecular perspective. Nat Rev Mol Cell Biol 2002;3:430–440. 18. Brill SJ, DiNardo S, Voelkel-Meiman K and Sternglanz R. DNA topoisomerase activity is required as a swivel for DNA replication and for ribosomal RNA transcription. Nci Monographs 1987;4:11–15.


77

19. Kim RA and Wang JC. Function of DNA topoisomerases as replication swivels in Saccharomyces cerevisiae. J Mol Biol 1989;208: 257–267. 20. Peter BJ, Ullsperger C, Hiasa H, Marians KJ and Cozzarelli NR. The structure of supercoiled intermediates in DNA replication. Cell 1998;94:819–827. 21. Zechiedrich EL, Khodursky AB and Cozzarelli NR. Topoisomerase IV, not gyrase, decatenates products of site-specific recombination in Escherichia coli. Genes Dev 1997;11:2580–2592. 22. Sherratt DJ, Soballe B, Barre FX, Filipe S, Lau I, Massey T and Yates J. Recombination and chromosome segregation. Philos Trans R Soc Lond B Biol Sci 2004;359:61–69. 23. Osheroff N. DNA Topoisomerases. Biochim Biophys Acta 1998;1400. 24. Champoux JJ. DNA topoisomerases: structure, function, and mechanism. Annu Rev Biochem 2001;70:369–413. 25. Wilstermann AM and Osheroff N. Stabilization of eukaryotic topoisomerase II-DNA cleavage complexes. Curr Top Med Chem 2003;3:321–338. 26. Velez-Cruz R and Osheroff N. DNA Topoisomerases: Type II. In: Lennarz W and Lane MD, eds. Encyclopedia of Molecular Biology. San Diego: Elsevier Science; 2004:806–811. 27. Pommier Y. Topoisomerase I inhibitors: camptothecins and beyond. Nat Rev Cancer 2006;6:789–802. 28. Berger JM, Gamblin SJ, Harrison SC and Wang JC. Structure and mechanism of DNA topoisomerase II. Nature 1996;379:225–232. 29. Wang JC. Moving one DNA double helix through another by a type II DNA topoismerase: the story of a simple molecular machine. Quart Rev Biophys 1998;31:107–144. 30. Sabourin M and Osheroff N. Topoisomerases. In: Creighton TE, ed. Encyclopedia of Molecular Medicine. New York: John Wiley and Sons, Inc.; 2002:3192–3197. 31. Stewart L, Redinbo MR, Qiu X, Hol WG and Champoux JJ. A model for the mechanism of human topoisomerase I. Science 1998;279:1534–1541. 32. Leppard JB and Champoux JJ. Human DNA topoisomerase I: relaxation, roles, and damage control. Chromosoma 2005;114:75–85. 33. Srivenugopal KS, Lockshon D and Morris DR. Escherichia coli DNA topoisomerase III: purification and characterization of a new type I enzyme. Biochemistry 1984;23:1899–1906. 34. Oakley TJ and Hickson ID. Defending genome integrity during S-phase: putative roles for RecQ helicases and topoisomerase III. DNA Repair (Amst) 2002;1:175–207.

78

Bender and Osheroff

35. Heyer WD, Ehmsen KT and Solinger JA. Holliday junctions in the eukaryotic nucleus: resolution in sight? Trends Biochem Sci 2003;28:548–557. 36. Laursen LV, Bjergbaek L, Murray JM and Andersen AH. RecQ helicases and topoisomerase III in cancer and aging. Biogerontology 2003;4:275–287. 37. Champoux JJ and Dulbecco R. An activity from mammalian cells that untwists superhelical DNA–a possible swivel for DNA replication (polyoma-ethidium bromide-mouse-embryo cells-dye binding assay). Proc Natl Acad Sci U S A 1972;69:143–146. 38. Rowe TC, Rusche JR, Brougham MJ and Holloman WK. Purification and properties of a topoisomerase from Ustilago maydis. J Biol Chem 1981;256:10354–10361. 39. Wang JC. Type I DNA Topoisomerases. Enzymes 1981: 331–344. 40. Attardi DG, De Paolis A and Tocchini-Valentini GP. Purification and characterization of Xenopus laevis type I topoisomerase. J Biol Chem 1981;256:3654–3661. 41. Larsen AK and Gobert C. DNA topoisomerase I in oncology: Dr Jekyll or Mr Hyde? Pathol Oncol Res 1999;5:171–178. 42. Morham SG, Kluckman KD, Voulomanos N and Smithies O. Targeted disruption of the mouse topoisomerase I gene by camptothecin selection. Mol Cell Biol 1996;16:6804–6809. 43. Zhang CX, Chen AD, Gettel NJ and Hsieh TS. Essential functions of DNA topoisomerase I in Drosophila melanogaster. Dev Biol 2000;222:27–40. 44. Zhang H, Barcelo JM, Lee B, Kohlhagen G, Zimonjic DB, Popescu NC and Pommier Y. Human mitochondrial topoisomerase I. Proc Natl Acad Sci USA 2001;98:10608–10613. 45. Roca J. The path of the DNA along the dimer interface of topoisomerase II. J Biol Chem 2004;279:25783–25788. 46. Liu LF, Rowe TC, Yang L, Tewey KM and Chen GL. Cleavage of DNA by mammalian DNA topoisomerase II. J Biol Chem 1983;258: 15365–15370. 47. Sander M and Hsieh T. Double strand DNA cleavage by type II DNA topoisomerase from Drosophila melanogaster. J Biol Chem 1983;258:8421–8428. 48. Osheroff N, Shelton ER and Brutlag DL. DNA topoisomerase II from Drosophila melanogaster. Relaxation of supercoiled DNA. J Biol Chem 1983;258:9536–9543. 49. Osheroff N. Role of the divalent cation in topoisomerase II-mediated reactions. Biochemistry 1987;26:6402–6406.


79

50. Osheroff N and Zechiedrich EL. Calcium-promoted DNA cleavage by eukaryotic topoisomerase II: trapping the covalent enzyme-DNA complex in an active form. Biochemistry 1987;26:4303–4309. 51. Osheroff N. Eukaryotic topoisomerase II. Characterization of enzyme turnover. J Biol Chem 1986;261:9944–9950. 52. Lindsley JE and Wang JC. Proteolysis patterns of epitopically labeled yeast DNA topoisomerase II suggest an allosteric transition in the enzyme induced by ATP binding. Proc Natl Acad Sci USA 1991;88:10485–10489. 53. Lindsley JE and Wang JC. On the coupling between ATP usage and DNA transport by yeast DNA topoisomerase II. J Biol Chem 1993;268:8096–8104. 54. Drake FH, Zimmerman JP, McCabe FL, Bartus HF, Per SR, Sullivan DM, Ross WE, Mattern MR, Johnson RK, Crooke ST and Mirabelli CK. Purification of topoisomerase II from amsacrine-resistant P388 leukemia cells. Evidence for two forms of the enzyme. J Biol Chem 1987;262:16739–16747. 55. Drake FH, Hofmann GA, Bartus HF, Mattern MR, Crooke ST and Mirabelli CK. Biochemical and pharmacological properties of p170 and p180 forms of topoisomerase II. Biochemistry 1989;28: 8154–8160. 56. Tsai-Pflugfelder M, Liu LF, Liu AA, Tewey KM, Whang-Peng J, Knutsen T, Huebner K, Croce CM and Wang JC. Cloning and sequencing of cDNA encoding human DNA topoisomerase II and localization of the gene to chromosome region 17q21–22. Proc Natl Acad Sci USA 1988;85:7177–7181. 57. Jenkins JR, Ayton P, Jones T, Davies SL, Simmons DL, Harris AL, Sheer D and Hickson ID. Isolation of cDNA clones encoding the beta isozyme of human DNA topoisomerase II and localisation of the gene to chromosome 3p24. Nucleic Acids Res 1992;20:5587–5592. 58. Tan KB, Dorman TE, Falls KM, Chung TD, Mirabelli CK, Crooke ST and Mao J. Topoisomerase II alpha and topoisomerase II beta genes: characterization and mapping to human chromosomes 17 and 3, respectively. Cancer Res 1992;52:231–234. 59. Austin CA and Marsh KL. Eukaryotic DNA topoisomerase IIbeta. BioEssays 1998;20:215–226. 60. Heck MM and Earnshaw WC. Topoisomerase II: A specific marker for cell proliferation. J Cell Biol 1986;103:2569–2581. 61. Hsiang YH, Wu HY and Liu LF. Proliferation-dependent regulation of DNA topoisomerase II in cultured human cells. Cancer Res 1988;48:3230–3235. 62. Woessner RD, Mattern MR, Mirabelli CK, Johnson RK and Drake FH. Proliferation- and cell cycle-dependent differences in expression of the

80

63.

64.

65.

66.

67.

68.

69. 70.

71.

72.

73.

74.

Bender and Osheroff 170 kilodalton and 180 kilodalton forms of topoisomerase II in NIH-3T3 cells. Cell Growth Diff 1991;2:209–214. Grue P, Grasser A, Sehested M, Jensen PB, Uhse A, Straub T, Ness W and Boege F. Essential mitotic functions of DNA topoisomerase II are not adopted by topoisomerase II in human H69 cells. J Biol Chem 1998;273:33660–33666. Heck MM, Hittelman WN and Earnshaw WC. Differential expression of DNA topoisomerases I and II during the eukaryotic cell cycle. Proc Natl Acad Sci USA 1988;85:1086–1090. Kimura K, Saijo M, Ui M and Enomoto T. Growth state- and cell cycledependent fluctuation in the expression of two forms of DNA topoisomerase II and possible specific modification of the higher molecular weight form in the M phase. J Biol Chem 1994;269:1173–1176. Bauman ME, Holden JA, Brown KA, Harker WG and Perkins SL. Differential immunohistochemical staining for DNA topoisomerase II alpha and beta in human tissues and for DNA topoisomerase II beta in non-Hodgkin’s lymphomas. Mod Pathol 1997;10:168–175. Christensen MO, Larsen MK, Barthelmes HU, Hock R, Andersen CL, Kjeldsen E, Knudsen BR, Westergaard O, Boege F and Mielke C. Dynamics of human DNA topoisomerases IIalpha and IIbeta in living cells. J Cell Biol 2002;157:31–44. Chen M and Beck WT. DNA topoisomerase II expression, stability, and phosphorylation in two VM-26-resistant human leukemic CEM sublines. Oncol Res 1995;7:103–111. Yang X, Li W, Prescott ED, Burden SJ and Wang JC. DNA topoisomerase IIbeta and neural development. Science 2000;287:131–134. Isaacs RJ, Davies SL, Sandri MI, Redwood C, Wells NJ and Hickson ID. Physiological regulation of eukaryotic topoisomerase II. Biochim Biophys Acta 1998;1400:121–137. Dereuddre S, Delaporte C and Jacquemin-Sablon A. Role of topoisomerase II beta in the resistance of 9-OH-ellipticine-resistant Chinese hamster fibroblasts to topoisomerase II inhibitors. Cancer Res 1997;57:4301–4308. Sakaguchi A and Kikuchi A. Functional compatibility between isoform alpha and beta of type II DNA topoisomerase. J Cell Sci 2004;117: 1047–1054. Haince JF, Rouleau M and Poirier GG. Transcription. Gene expression needs a break to unwind before carrying on. Science 2006;312:1752–1753. Ju BG, Lunyak VV, Perissi V, Garcia-Bassets I, Rose DW, Glass CK and Rosenfeld MG. A topoisomerase IIbeta-mediated dsDNA break required for regulated transcription. Science 2006;312:1798–1802.


81

75. Kaufmann SH. Cell death induced by topoisomerase-targeted drugs: more questions than answers. Biochim Biophys Acta 1998;1400:195–211. 76. Kaufmann SH, Gore SD, Miller CB, Jones RJ, Zwelling LA, Schneider E, Burke PJ and Karp JE. Topoisomerase II and the response to antileukemic therapy. Leukemia Lymph 1998;29:217–237. 77. Rowley JD. The critical role of chromosome translocations in human leukemias. Ann Rev Genet 1998;32:495–519. 78. Felix CA. Secondary leukemias induced by topoisomerase-targeted drugs. Biochim Biophys Acta 1998;1400:233–255. 79. Sordet O, Khan QA, Kohn KW and Pommier Y. Apoptosis induced by topoisomerase inhibitors. Curr Med Chem Anti-Canc Agents 2003;3:271–290. 80. Kreuzer KN, McEntee K, Geballe AP and Cozzarelli NR. Lambda transducing phages for the nalA gene of Escherichia coli and conditional lethal nalA mutations. Mol Gen Genet 1978;167:129–137. 81. Bjornsti M-A, Benedetti P, Viglianti GA and Wang JC. Expression of human DNA topoisomerase I in yeast cells lacking DNA topoisomerase I: restoration of sensitivity of the cells to the antitumor drug camptothecin. Cancer Res 1989;49:6318–6323. 82. Pommier Y, Pourquier P, Fan Y and Strumberg D. Mechanism of action of eukaryotic DNA topoisomerase I and drugs targeted to the enzyme. Biochim Biophys Acta 1998;1400:83–106. 83. Li TK and Liu LF. Tumor cell death induced by topoisomerase-targeting drugs. Annu Rev Pharmacol Toxicol 2001;41:53–77. 84. Meng LH, Liao ZY and Pommier Y. Non-camptothecin DNA topoisomerase I inhibitors in cancer therapy. Curr Top Med Chem 2003;3: 305–320. 85. Hsiang YH and Liu LF. Identification of mammalian DNA topoisomerase I as an intracellular target of the anticancer drug camptothecin. Cancer Res 1988;48:1722–1726. 86. Osheroff N. Effect of antineoplastic agents on the DNA cleavage/religation reaction of eukaryotic topoisomerase II: inhibition of DNA religation by etoposide. Biochemistry 1989;28:6157–6160. 87. Staker BL, Hjerrild K, Feese MD, Behnke CA, Burgin AB, Jr. and Stewart L. The mechanism of topoisomerase I poisoning by a camptothecin analog. Proc Natl Acad Sci U S A 2002;99:15387–15392. 88. Robinson MJ, Martin BA, Gootz TD, McGuirk PR, Moynihan M, Sutcliffe JA and Osheroff N. Effects of quinolone derivatives on eukaryotic topoisomerase II. A novel mechanism for enhancement of enzyme-mediated DNA cleavage. J Biol Chem 1991;266:14585–14592. 89. Kingma PS and Osheroff N. Spontaneous DNA damage stimulates topoisomerase II-mediated DNA cleavage. J Biol Chem 1997;272: 7488–7493.

82

Bender and Osheroff

90. Cline SD, Jones WR, Stone MP and Osheroff N. DNA abasic lesions in a different light: solution structure of an endogenous topoisomerase II poison. Biochemistry 1999;38:15500–15507. 91. Cline SD and Osheroff N. Cytosine arabinoside (araC) lesions are positionspecific topoisomerase II poisons and stimulate DNA cleavage mediated by the human type II enzymes. J Biol Chem 1999;274:29740–29743. 92. Velez-Cruz R, Riggins JN, Daniels JS, Cai H, Guengerich FP, Marnett LJ and Osheroff N. Exocyclic DNA lesions stimulate DNA cleavage mediated by human topoisomerase II alpha in vitro and in cultured cells. Biochemistry 2005;44:3972–3981. 93. Wall ME and Wani MC. Camptothecin and taxol: discovery to clinic–thirteenth Bruce F. Cain Memorial Award Lecture. Cancer Res 1995;55:753–760. 94. Garcia-Carbonero R and Supko JG. Current perspectives on the clinical experience, pharmacology, and continued development of the camptothecins. Clin Cancer Res 2002;8:641–661. 95. Burke TG and Mi Z. Preferential binding of the carboxylate form of camptothecin by human serum albumin. Anal Biochem 1993;212:285–287. 96. Burke TG and Mi Z. The structural basis of camptothecin interactions with human serum albumin: impact on drug stability. J Med Chem 1994;37:40–46. 97. Bailly C. Homocamptothecins: potent topoisomerase I inhibitors and promising anticancer drugs. Crit Rev Oncol Hematol 2003;45:91–108. 98. Osheroff N. Diflomotecan. Ipsen. IDrugs 2004;7:257–263. 99. Yamashita Y, Fujii N, Murakata C, Ashizawa T, Okabe M and Nakano H. Induction of mammalian DNA topoisomerase I mediated DNA cleavage by antitumor indolocarbazole derivatives. Biochemistry 1992;31:12069–12075. 100. Antony S, Jayaraman M, Laco G, Kohlhagen G, Kohn KW, Cushman M and Pommier Y. Differential induction of topoisomerase I-DNA cleavage complexes by the indenoisoquinoline MJ-III-65 (NSC 706744) and camptothecin: base sequence analysis and activity against camptothecinresistant topoisomerases I. Cancer Res 2003;63):7428–7435. 101. Li TK, Houghton PJ, Desai SD, Daroui P, Liu AA, Hars ES, Ruchelman AL, LaVoie EJ and Liu LF. Characterization of ARC-111 as a novel topoisomerase I-targeting anticancer drug. Cancer Res 2003;63:8400–8407. 102. Hande KR. Clinical applications of anticancer drugs targeted to topoisomerase II. Biochim Biophys Acta 1998;1400:173–184. 103. Hande KR. Etoposide: four decades of development of a topoisomerase II inhibitor. Eur J Cancer 1998;34:1514–1521.


83

104. Ross W, Rowe T, Glisson B, Yalowich J and Liu L. Role of topoisomerase II in mediating epipodophyllotoxin-induced DNA cleavage. Cancer Res 1984;44:5857–5860. 105. Chen GL, Yang L, Rowe TC, Halligan BD, Tewey KM and Liu LF. Nonintercalative antitumor drugs interfere with the breakagereunion reaction of mammalian DNA topoisomerase II. J Biol Chem 1984;259:13560–13566. 106. Yang L, Rowe TC, Nelson EM and Liu LF. In vivo mapping of DNA topoisomerase II-specific cleavage sites on SV40 chromatin. Cell 1985;41:127–132. 107. Nitiss JL. Using yeast to study resistance to topoisomerase II-targeting drugs. Cancer Chemother Pharmacol 1994;34:S6–13. 108. Burden DA and Osheroff N. Mechanism of action of eukaryotic topoisomerase II and drugs targeted to the enzyme. Biochim Biophys Acta 1998;1400:139–154. 109. Jensen PB, Sorensen BS, Sehested M, Grue P, Demant EJ and Hansen HH. Targeting the cytotoxicity of topoisomerase II-directed epipodophyllotoxins to tumor cells in acidic environments. Cancer Res 1994;54:2959–2963. 110. Sissi C and Palumbo M. The quinolone family: from antibacterial to anticancer agents. Curr Med Chem Anticancer Agents 2003;3:439–450. 111. Mitscher LA. Bacterial topoisomerase inhibitors: quinolone and pyridone antibacterial agents. Chem Rev 2005;105:559–592. 112. Robicsek A, Jacoby GA and Hooper DC. The worldwide emergence of plasmid-mediated quinolone resistance. Lancet Infect Dis 2006;6:629–640. 113. Constantinou A, Mehta R, Runyan C, Rao K, Vaughan A and Moon R. Flavonoids as DNA topoisomerase antagonists and poisons: structureactivity relationships. J Nat Prod 1995;58:217–225. 114. Strick R, Strissel PL, Borgers S, Smith SL and Rowley JD. Dietary bioflavonoids induce cleavage in the MLL gene and may contribute to infant leukemia. Proc Natl Acad Sci U S A 2000;97:4790–4795. 115. Martin-Cordero C, Lopez-Lazaro M, Pinero J, Ortiz T, Cortes F and Ayuso MJ. Glucosylated isoflavones as DNA topoisomerase II poisons. J Enzyme Inhib 2000;15:455–460. 116. Galati G and O’Brien PJ. Potential toxicity of flavonoids and other dietary phenolics: significance for their chemopreventive and anticancer properties. Free Radic Biol Med 2004;37:287–303. 117. Adlercreutz H, Markkanen H, and Watanabe S. Plasma concentrations of phyto-oestrogens in Japanese men. Lancet 1993;342:1209–1210. 118. Lamartiniere CA. Protection against breast cancer with genistein: a component of soy. Am J Clin Nutr 2000;71:1705S–1709S.

84

Bender and Osheroff

119. Sarkar FH, Adsule S, Padhye S, Kulkarni S, and Li Y. The role of genistein and synthetic derivatives of isoflavone in cancer prevention and therapy. Mini Rev Med Chem 2006;6:401–407. 120. Siddiqui IA, Adhami VM, Saleem M, and Mukhtar H. Beneficial effects of tea and its polyphenols against prostate cancer. Mol Nutr Food Res 2006;50:130–143. 121. Ross JA, Potter JD, and Robison LL. Infant leukemia, topoisomerase II inhibitors, and the MLL gene. Journal of the National Cancer Institute 1994;86(22):1678–1680. 122. Ross JA, Potter JD, Reaman GH, Pendergrass TW, and Robison LL. Maternal exposure to potential inhibitors of DNA topoisomerase II and infant leukemia (United States): a report from the Children’s Cancer Group. Cancer causes & control : CCC 1996;7:581–590. 123. Ross JA. Dietary flavonoids and the MLL gene: A pathway to infant leukemia? Proc Natl Acad Sci U S A 2000;97:4411–4413. 124. Spector LG, Xie Y, Robison LL, Heerema NA, Hilden JM, Lange B, Felix CA, Davies SM, Slavin J, Potter JD, Blair CK, Reaman GH, and Ross JA. Maternal diet and infant leukemia: the DNA topoisomerase II inhibitor hypothesis: a report from the children’s oncology group. Cancer Epidemiol Biomarkers Prev 2005;14:651–655. 125. Shen Y, Shen HM, Shi CY, and Ong CN. Benzene metabolites enhance reactive oxygen species generation in HL60 human leukemia cells. Hum Exp Toxicol 1996;15:422–427. 126. Kuo ML, Shiah SG, Wang CJ, and Chuang SE. Suppression of apoptosis by Bcl-2 to enhance benzene metabolites-induced oxidative DNA damage and mutagenesis: A possible mechanism of carcinogenesis. Mol Pharmacol 1999;55:894–901. 127. Lovern MR, Cole CE, and Schlosser PM. A review of quantitative studies of benzene metabolism. Crit Rev Toxicol 2001;31:285–311. 128. Rappaport SM, Waidyanatha S, Qu Q, Shore R, Jin X, Cohen B, Chen LC, Melikian AA, Li G, Yin S, Yan H, Xu B, Mu R, Li Y, Zhang X, and Li K. Albumin adducts of benzene oxide and 1,4benzoquinone as measures of human benzene metabolism. Cancer Res 2002;62:1330–1337. 129. Prescott LF. Paracetamol: past, present, and future. Am J Ther 2000;7:143–147. 130. Bender RP, Lindsey RH, Jr., Burden DA, and Osheroff N. N-acetyl-pbenzoquinone imine, the toxic metabolite of acetaminophen, is a topoisomerase II poison. Biochemistry 2004;43:3731–3739. 131. Ross D. The role of metabolism and specific metabolites in benzeneinduced toxicity: evidence and issues. J Toxicol Environ Health A 2000;61:357–372.


85

132. Lindsey RH, Jr., Bromberg KD, Felix CA, and Osheroff N. 1,4-Benzoquinone is a topoisomerase II poison. Biochemistry 2004;43:7563–7574. 133. Lindsey RH, Jr., Bender RP, and Osheroff N. Effects of benzene metabolites on DNA cleavage mediated by human topoisomerase II alpha: 1,4-hydroquinone is a topoisomerase II poison. Chem Res Toxicol 2005;18:761–770. 134. Bender RP, Lehmler HJ, Robertson LW, Ludewig G, and Osheroff N. Polychlorinated Biphenyl Quinone Metabolites Poison Human Topoisomerase IIalpha: Altering Enzyme Function by Blocking the N-Terminal Protein Gate. Biochemistry 2006;45:10140–10152. 135. Wang H, Mao Y, Chen AY, Zhou N, LaVoie EJ, and Liu LF. Stimulation of topoisomerase II-mediated DNA damage via a mechanism involving protein thiolation. Biochemistry 2001;40:3316–3323. 136. Pourquier P, Waltman JL, Urasaki Y, Loktionova NA, Pegg AE, Nitiss JL, and Pommier Y. Topoisomerase I-mediated cytotoxicity of N-methyl-N’-nitro-N-nitrosoguanidine: trapping of topoisomerase I by the O6-methylguanine. Cancer Res 2001;61:53–58. 137. Pourquier P and Pommier Y. Topoisomerase I-mediated DNA damage. Adv Cancer Res 2001;80:189–216. 138. Pommier Y, Redon C, Rao VA, Seiler JA, Sordet O, Takemura H, Antony S, Meng L, Liao Z, Kohlhagen G, Zhang H, and Kohn KW. Repair of and checkpoint response to topoisomerase I-mediated DNA damage. Mutat Res 2003;532:173–203. 139. Pommier Y, Barcelo JM, Rao VA, Sordet O, Jobson AG, Thibaut L, Miao ZH, Seiler JA, Zhang H, Marchand C, Agama K, Nitiss JL, and Redon C. Repair of topoisomerase I-mediated DNA damage. Prog Nucleic Acid Res Mol Biol 2006;81:179–229. 140. Kingma PS and Osheroff N. The response of eukaryotic topoisomerases to DNA damage. Biochim Biophys Acta 1998;1400:223–232. 141. Sabourin M and Osheroff N. Sensitivity of human type II topoisomerases to DNA damage: stimulation of enzyme-mediated DNA cleavage by abasic, oxidized and alkylated lesions. Nucleic Acids Res 2000;28:1947–1954. 142. Khan QA, Kohlhagen G, Marshall R, Austin CA, Kalena GP, Kroth H, Sayer JM, Jerina DM, and Pommier Y. Position-specific trapping of topoisomerase II by benzo[a]pyrene diol epoxide adducts: implications for interactions with intercalating anticancer agents. Proc Natl Acad Sci U S A 2003;100:12498–12503. 143. Sordet O, Khan QA, and Pommier Y. Apoptotic topoisomerase I-DNA complexes induced by oxygen radicals and mitochondrial dysfunction. Cell Cycle 2004;3:1095–1097.

86

Bender and Osheroff

144. Pourquier P, Kohlhagen G, Urasaki Y, and Pommier Y. Induction of topoisomerase I cleavage complexes by staurosporin in apoptotic leukemia cells: Could topoisomerase I act as an apoptotic endonuclease? Proc Am Assoc Cancer Res 2001:303. 145. Solovyan VT, Bezvenyuk ZA, Salminen A, Austin CA, and Courtney MJ. The role of topoisomerase II in the excision of DNA loop domains during apoptosis. J Biol Chem 2002;277:21458–21467. 146. Belyaev IY. DNA loop organization and DNA fragmentation during radiation-induced apoptosis in human lymphocytes. Radiats Biol Radioecol 2005;45:541–548. 147. Felix CA. Leukemias related to treatment with DNA topoisomerase II inhibitors. Medical & Pediatric Oncology 2001;36:525–535. 148. Smith MA, Rubinstein L, Anderson JR, Arthur D, Catalano PJ, Freidlin B, Heyn R, Khayat A, Krailo M, Land VJ, Miser J, Shuster J, and Vena D. Secondary leukemia or myelodysplastic syndrome after treatment with epipodophyllotoxins. J Clin Oncol 1999;17:569–577. 149. Felix CA, Kolaris CP, and Osheroff N. Topoisomerase II and the etiology of chromosomal translocations. DNA Repair (Amst) 2006;5:109–1108. 150. Mistry AR, Felix CA, Whitmarsh RJ, Mason A, Reiter A, Cassinat B, Parry A, Walz C, Wiemels JL, Segal MR, Ades L, Blair IA, Osheroff N, Peniket AJ, Lafage-Pochitaloff M, Cross NC, Chomienne C, Solomon E, Fenaux P, and Grimwade D. DNA topoisomerase II in therapy-related acute promyelocytic leukemia. N Engl J Med 2005;352:1529–1538. 151. Hess JL. MLL: a histone methyltransferase disrupted in leukemia. Trends Mol Med 2004;10:500–507. 152. Hess JL. Mechanisms of transformation by MLL. Crit Rev Eukaryot Gene Expr 2004;14:235–254. 153. Tenney K and Shilatifard A. A COMPASS in the voyage of defining the role of trithorax/MLL-containing complexes: linking leukemogensis to covalent modifications of chromatin. J Cell Biochem 2005;95:429–436. 154. Lappin TR, Grier DG, Thompson A, and Halliday HL. HOX genes: seductive science, mysterious mechanisms. Ulster Med J 2006;75:23–31. 155. Li ZY, Liu DP, and Liang CC. New insight into the molecular mechanisms of MLL-associated leukemia. Leukemia 2005;19:183–190. 156. Felix CA and Lange BJ. Leukemia in infants. Oncologist 1999;4:225–240. 157. Smith MT. Chromosome damage from biological reactive intermediates of benzene and 1,3-butadiene in leukemia. Adv Exp Med Biol 2001;500:279–287. 158. Larson RA, Wang Y, Banerjee M, Wiemels J, Hartford C, Le Beau MM, and Smith MT. Prevalence of the inactivating 609C–>T polymorphism in the NAD(P)H:quinone oxidoreductase (NQO1) gene in patients with primary and therapy-related myeloid leukemia. Blood 1999;94:803–807.


87

159. Smith MT. Benzene, NQO1, and genetic susceptibility to cancer. Proc Natl Acad Sci U S A 1999;96:7624–7626. 160. Smith MT, Wang Y, Kane E, Rollinson S, Wiemels JL, Roman E, Roddam P, Cartwright R, and Morgan G. Low NAD(P)H:quinone oxidoreductase 1 activity is associated with increased risk of acute leukemia in adults. Blood 2001;97:1422–1426. 161. Smith MT, Wang Y, Skibola CF, Slater DJ, Lo Nigro L, Nowell PC, Lange BJ, and Felix CA. Low NAD(P)H:quinone oxidoreductase activity is associated with increased risk of leukemia with MLL translocations in infants and children. Blood 2002;100: 4590–4593. 162. Wiemels JL, Pagnamenta A, Taylor GM, Eden OB, Alexander FE, and Greaves MF. A lack of a functional NAD(P)H:quinone oxidoreductase allele is selectively associated with pediatric leukemias that have MLL fusions. United Kingdom Childhood Cancer Study Investigators. Cancer Res 1999;59:4095–4099. 163. Lindsey RH, Bender RP, and Osheroff N. Stimulation of topoisomerase II-mediated DNA cleavage by benzene metabolites. Chem Biol Interact 2005;153–154:197–205. 164. Aplan PD. Chromosomal translocations involving the MLL gene: molecular mechanisms. DNA Repair (Amst) 2006;5:1265–1272. 165. Felix CA, Lange BJ, Hosler MR, Fertala J, and Bjornsti M-A. Chromosome band 11q23 translocation breakpoints are DNA topoisomerase II cleavage sites. Cancer Res 1995;55:4287–4292. 166. Lovett BD, Lo Nigro L, Rappaport EF, Blair IA, Osheroff N, Zheng N, Megonigal MD, Williams WR, Nowell PC, and Felix CA. Near-precise interchromosomal recombination and functional DNA topoisomerase II cleavage sites at MLL and AF-4 genomic breakpoints in treatmentrelated acute lymphoblastic leukemia with t(4;11) translocation. Proc Natl Acad Sci U S A 2001;98:9802–9807. 167. Lovett BD, Strumberg D, Blair IA, Pang S, Burden DA, Megonigal MD, Rappaport EF, Rebbeck TR, Osheroff N, Pommier YG and Felix CA. Etoposide metabolites enhance DNA topoisomerase II cleavage near leukemia-associated MLL translocation breakpoints. Biochemistry 2001;40:1159–1170. 168. Super HJ, McCabe NR, Thirman MJ, Larson RA, Le Beau MM, Pedersen-Bjergaard J, Philip P, Diaz MO, and Rowley JD. Rearrangements of the MLL gene in therapy-related acute myeloid leukemia in patients previously treated with agents targeting DNA-topoisomerase II. Blood 1993;82:3705–3711. 169. Andersen MK, Christiansen DH, Jensen BA, Ernst P, Hauge G, and Pedersen-Bjergaard J. Therapy-related acute lymphoblastic leukaemia with MLL rearrangements following DNA topoisomerase II inhibitors,

88

170.

171.

172.

173.

174.

175. 176. 177.

178.

179. 180. 181.

182. 183. 184.

185.

Bender and Osheroff an increasing problem: report on two new cases and review of the literature since 1992. Br J Haematol 2001;114:539–543. Betti CJ, Villalobos MJ, Jiang Q, Cline E, Diaz MO, Loredo G, and Vaughan AT. Cleavage of the MLL gene by activators of apoptosis is independent of topoisomerase II activity. Leukemia 2005;19:2289–2295. Hars ES, Lyu YL, Lin CP, and Liu LF. Role of apoptotic nuclease caspase-activated DNase in etoposide-induced treatment-related acute myelogenous leukemia. Cancer Res 2006;66:8975–8979. Betti CJ, Villalobos MJ, Diaz MO, and Vaughan AT. Apoptotic triggers initiate translocations within the MLL gene involving the nonhomologous end joining repair system. Cancer Res 2001;61:4550–4555. Betti CJ, Villalobos MJ, Diaz MO, and Vaughan AT. Apoptotic stimuli initiate MLL-AF9 translocations that are transcribed in cells capable of division. Cancer Res 2003;63:1377–1381. Vaughan AT, Betti CJ, Villalobos MJ, Premkumar K, Cline E, Jiang Q, and Diaz MO. Surviving apoptosis: a possible mechanism of benzeneinduced leukemia. Chem Biol Interact 2005;153–154:179–185. Shiloh Y. ATM and ATR: networking cellular responses to DNA damage. Curr Opin Genet Dev 2001;11:71–77. Abraham RT. Cell cycle checkpoint signaling through the ATM and ATR kinases. Genes Dev 2001;15:2177–2196. Abraham RT. Checkpoint signaling: epigenetic events sound the DNA strand-breaks alarm to the ATM protein kinase. Bioessays 2003;25:627–630. Bakkenist CJ and Kastan MB. DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature 2003;421:499–506. Blasina A, Price BD, Turenne GA, and McGowan CH. Caffeine inhibits the checkpoint kinase ATM. Curr Biol 1999;9(19):1135–1138. Kornbluth S, Smythe C, and Newport JW. In vitro cell cycle arrest induced by using artificial DNA templates. Mol Cell Biol 1992;12:3216–3223. Zou L, Cortez D, and Elledge SJ. Regulation of ATR substrate selection by Rad17-dependent loading of Rad9 complexes onto chromatin. Genes Dev 2002;16:198–208. Carr AM. DNA structure dependent checkpoints as regulators of DNA repair. DNA Repair (Amst) 2002;1:983–994. Cortez D. Unwind and slow down: checkpoint activation by helicase and polymerase uncoupling. Genes Dev 2005;19:1007–1012. Sogo JM, Lopes M, and Foiani M. Fork reversal and ssDNA accumulation at stalled replication forks owing to checkpoint defects. Science 2002;297:599–602. McGowan CH and Russell P. The DNA damage response: sensing and signaling. Curr Opin Cell Biol 2004;16:629–633.


89

186. Liu Q, Guntuku S, Cui XS, Matsuoka S, Cortez D, Tamai K, Luo G, Carattini-Rivera S, DeMayo F, Bradley A, Donehower LA, and Elledge SJ. Chk1 is an essential kinase that is regulated by Atr and required for the G(2)/M DNA damage checkpoint. Genes Dev 2000;14:1448–1459. 187. Harrison JC and Haber JE. Surviving the breakup: the DNA damage checkpoint. Annu Rev Genet 2006;40:209–235. 188. Lavin MF and Shiloh Y. The genetic defect in ataxia-telangiectasia. Annu Rev Immunol 1997;15:177–202. 189. Rotman G and Shiloh Y. ATM: a mediator of multiple responses to genotoxic stress. Oncogene 1999;18:6135–6144. 190. Bott L, Thumerelle C, Cuvellier JC, Deschildre A, Vallee L, and Sardet A. Ataxia-telangiectasia: a review. Arch Pediatr 2006;13:293–298. 191. Brown EJ and Baltimore D. ATR disruption leads to chromosomal fragmentation and early embryonic lethality. Genes Dev 2000;14:397–402. 192. de Klein A, Muijtjens M, van Os R, Verhoeven Y, Smit B, Carr AM, Lehmann AR, and Hoeijmakers JH. Targeted disruption of the cell-cycle checkpoint gene ATR leads to early embryonic lethality in mice. Curr Biol 2000;10:479–482. 193. Cliby WA, Lewis KA, Lilly KK, and Kaufmann SH. S phase and G2 arrests induced by topoisomerase I poisons are dependent on ATR kinase function. J Biol Chem 2002;277:1599–1606. 194. Costanzo V, Shechter D, Lupardus PJ, Cimprich KA, Gottesman M, and Gautier J. An ATR- and Cdc7-dependent DNA damage checkpoint that inhibits initiation of DNA replication. Mol Cell 2003;11: 203–213. 195. Siu WY, Lau A, Arooz T, Chow JP, Ho HT, and Poon RY. Topoisomerase poisons differentially activate DNA damage checkpoints through ataxia-telangiectasia mutated-dependent and -independent mechanisms. Mol Cancer Ther 2004;3:621–632. 196. Ho CC, Siu WY, Chow JP, Lau A, Arooz T, Tong HY, Ng IO, and Poon RY. The relative contribution of CHK1 and CHK2 to Adriamycininduced checkpoint. Exp Cell Res 2005;304:1–15. 197. Flatten K, Dai NT, Vroman BT, Loegering D, Erlichman C, Karnitz LM, and Kaufmann SH. The role of checkpoint kinase 1 in sensitivity to topoisomerase I poisons. J Biol Chem 2005;280:14349–14355. 198. Martensson S, Nygren J, Osheroff N, and Hammarsten O. Activation of DNA-dependent protein kinase by drug and radiation-induced DNA strand breaks. Radiat Res 2003:in press. 199. Vance JR and Wilson TE. Yeast Tdp1 and Rad1-Rad10 function as redundant pathways for repairing Top1 replicative damage. Proc Natl Acad Sci U S A 2002;99:13669–13674.

90

Bender and Osheroff

200. Neale MJ, Pan J, and Keeney S. Endonucleolytic processing of covalent protein-linked DNA double-strand breaks. Nature 2005;436:1053–1057. 201. Deng C, Brown JA, You D, and Brown JM. Multiple endonucleases function to repair covalent topoisomerase I complexes in Saccharomyces cerevisiae. Genetics 2005;170:591–600. 202. Zhang A, Lyu YL, Lin CP, Zhou N, Azarova AM, Wood LM and Liu LF. A protease pathway for the repair of topoisomerase II-DNA covalent complexes. J Biol Chem 2006. 203. Desai SD, Zhang H, Rodriguez-Bauman A, Yang JM, Wu X, Gounder MK, Rubin EH, and Liu LF. Transcription-dependent degradation of topoisomerase I-DNA covalent complexes. Mol Cell Biol 2003;23:2341–2350. 204. Hammarsten O, DeFazio LG, and Chu G. Activation of DNAdependent protein kinase by single-stranded DNA ends. J Biol Chem 2000;275:1541–1550. 205. Yang SW, Burgin AB, Jr., Huizenga BN, Robertson CA, Yao KC, and Nash HA. A eukaryotic enzyme that can disjoin dead-end covalent complexes between DNA and type I topoisomerases. Proc Natl Acad Sci USA 1996;93:11534–11539. 206. Barthelmes HU, Habermeyer M, Christensen MO, Mielke C, Interthal H, Pouliot JJ, Boege F, and Marko D. TDP1 overexpression in human cells counteracts DNA damage mediated by topoisomerases I and II. J Biol Chem 2004;279:55618–55625. 207. Nitiss KC, Malik M, He X, White SW, and Nitiss JL. Tyrosyl-DNA phosphodiesterase (Tdp1) participates in the repair of Top2-mediated DNA damage. Proc Natl Acad Sci U S A 2006;103:8953–8958. 208. Raymond AC and Burgin AB, Jr. Tyrosyl-DNA phosphodiesterase (Tdp1) (3’-phosphotyrosyl DNA phosphodiesterase). Methods Enzymol 2006;409:511–524. 209. Connelly JC and Leach DR. Repair of DNA covalently linked to protein. Mol Cell 2004;13:307–316. 210. El-Khamisy SF and Caldecott KW. DNA single-strand break repair and spinocerebellar ataxia with axonal neuropathy-1. Neuroscience 2006;145:1260–1266. 211. Interthal H, Chen HJ, Kehl-Fie TE, Zotzmann J, Leppard JB, and Champoux JJ. SCAN1 mutant Tdp1 accumulates the enzyme– DNA intermediate and causes camptothecin hypersensitivity. Embo J 2005;24:2224–2233. 212. Miao ZH, Agama K, Sordet O, Povirk L, Kohn KW, and Pommier Y. Hereditary ataxia SCAN1 cells are defective for the repair of transcription-dependent topoisomerase I cleavage complexes. DNA Repair (Amst) 2006;5:1489–1494.


91

213. Takashima H. Molecular genetics of inherited neuropathies. Rinsho Shinkeigaku 2006;46:1–18. 214. Sabourin M, Nitiss JL, Nitiss KC, Tatebayash H, Ikeda H, and Osheroff N. Yeast recombination pathways triggered by topoisomerase II-mediated DNA breaks. Nucl Acids Res 2003;313:4373–4384. 215. Valerie K and Povirk LF. Regulation and mechanisms of mammalian double-strand break repair. Oncogene 2003;22:5792–5812.

4

Targeting ATM/ATR in the DNA Damage Checkpoint Joseph M. Ackermann and Wafik S. El-Deiry CONTENTS Introduction ATM/ATR Gene Organization ATM and ATR are Key Mediators of the DNA Damage Response Pathway Prospects of Regulating ATM/ATR Pathways

Key Words: ATM; ATR; Ataxia telangiectasia; DNA damage checkpoint; ionizing radiation; phosphorylation

1. INTRODUCTION DNA damage elicits an intricate, coordinated response in the cell leading to cell cycle arrest, DNA repair, or cell death. The DNA damage response is mobilized in the very early stages of tumorigenesis—a process that selects for cells deficient in a properly functioning DNA damage response (1,2). Interest in the disease ataxia telangiectasia (A-T) spurred significant advances in our current understanding of the DNA damage response. Recognized in mid-1960, From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

93

94

Ackermann and El-Deiry

A-T was named after its indications—cerebellar ataxia (uncoordinated movement) and ocular telangiectasia (dilated blood vessels of the eye) (3). It is an autosomal recessive neurological disorder that afflicts victims in early childhood and is also characterized by immunodeficiency, hypogonadism, and susceptibility to ionizing radiation (IR), radiomimetic compounds, and cancer (4,5). A breakthrough occurred in 1995 when Yosef Shiloh and colleagues used positional cloning to identify the gene responsible for A-T, ATM (A-T mutated) (6). In the decade plus since this pivotal discovery, our understanding of the DNA damage response has improved immensely. Key players have been identified, but our limited knowledge of their precise roles and dynamic interactions suggest that we have only scratched the surface. ATM and A-T and Rad3 related protein (ATR) have emerged as central proteins in the DNA damage response. ATM and ATR belong to the phosphatidylinositol-3 kinase related kinase (PIKK) family of proteins that also includes the catalytic subunit of DNA-dependent protein kinase (DNA-PKcs)—all of which are important in the DNA damage response. DNA is susceptible to damage induced by exogenous agents (ultraviolet light [UV], IR, radiomimetic chemicals, DNA replication inhibitors, viral infection) as well as endogenous processes (replication fork stalling, oxygen radicals generated from normal metabolism, meiosis in germinal cells, V(D)J recombination and class-switch recombination). If left unchecked, the DNA damage may lead to cancer. Therefore, mechanisms are in place to detect DNA damage that trigger an appropriate response—cell cycle inhibition, activation of DNA repair systems, and if the damage is irreparable, apoptosis. The type of DNA damage dictates the response. ATM responds primarily to DNA double strand breaks (DSB), such as those induced by IR and radiomimetic compounds. ATR is activated in response to replication interference, UV, and bulky lesions—such as those caused by DNA alkylating agents. Unlike ATM and ATR, DNAPKcs does not appear to be directly involved in coordinating the DNA damage response. Instead, DNA-PKcs serves as a critical mediator of DNA repair. ATM and ATR are also activated by hypoxia—which does not induce DNA damage—and reoxygenation—which does induce DNA damage (7–13). ATM and ATR have also been implicated in the maintenance of naturally occurring DNA DSBs—teleomeres (14–16). This chapter will focus on understanding the activation and function of ATM and ATR within the scope of the DNA damage response.

Chapter 4 / Targeting ATM/ATR in the DNA Damage Checkpoint

95

2. ATM/ATR GENE ORGANIZATION ATM and ATR share a similar overall structural organization. The ATM gene encodes a 3,056 amino acid polypeptide of approximately 369 kDa; the ATR gene encodes a slightly smaller polypeptide of 2,644 amino acids (17). ATM and ATR proteins both contain a PI 3-kinase related catalytic domain near the carboxyl terminus flanked by 2 loosely conserved domains: a FAT (FRAP/ATM/TTRAP) and FATC (“C” indicates proximity to the carboxy-terminus) (18). The function of the FAT domains is unknown, although it has been proposed to interact with the kinase domain to stabilize the carboxy-terminal end of the protein (19). The catalytic domain functions as a S/T-Qdirected kinase (serine or threonine followed by glutamine) directed solely to proteins—it has no detectable lipid kinase activity (20). In the canonical model, ATM and ATR carry out unique and mutually exclusive roles in the DNA damage response (21). ATR is activated by replication stress—DNA damage that arises at the replication fork that could be induced by stalled replication forks, the DNA replication inhibitor hydroxyurea (HU), and bulky lesions caused by ultraviolet light (UV) or DNA-alkylating agents, such as methyl methanesulfonate (MMS). ATM is activated in response to DSBs that are not associated with replication stress, such as DSBs caused by IR and radiomimetic chemicals. The emerging picture points to a more complex, intertwined relationship between ATM and ATR.

3. ATM AND ATR ARE KEY MEDIATORS OF THE DNA DAMAGE RESPONSE PATHWAY 3.1. Regulation of ATM by Phosphorylation ATM is predominantly a nuclear protein that exists in an inactive dimeric or higher-order oligomeric form. Intermolecular autophosphorylation of serine 1981, located in the FAT domain, disrupts the ATM oligomer freeing the monomer to interact with substrate proteins (19). This phosphorylation event is very rapid—it occurs within 5 min after exposure to low levels of IR. Two additional autophosphorylation sites on ATM, serine 367 and serine 1893, are involved in ATM signaling (22). Introduction of ATM into A-T cells restores the radiosensitivity of these cells to levels similar to the parental cell line with functional ATM; transfection of S367A or S1893A mutants are unable to do so. These three phosphorylation sites appear to be important in regulating the activity of ATM, but are they essential?

96


It may be premature to regard phosphorylation of serines 367, 1893, and 1981 as essential for ATM activity. In in vitro assays, the ATM S1981A mutant does not block dimer formation or ATM kinase activity (19,23). ATM phosphorylation at S1981 is not required for ATM monomerization because ATM monomers not phosphorylated at S1981 have been detected (24). In vivo expression of a S1987A (mouse amino acid corresponding to human S1981) mutant as the sole ATM species does not abrogate the ATM-dependent response (25). These authors went on to show that the S1987A mutant ATM does not have a dominant-negative effect on ATM heterozygous mice when overexpressed or expressed at physiological levels. Additionally, both S367 and S1893 are autophosphorylated in vitro, but only S1893 has been demonstrated to be phosphorylated in vivo (22). Together, these data indicate that phosphorylation of these residues likely plays a role in the regulation of the ATM signaling pathway, but raises questions as to whether these are essential events for ATM activation or consequences of ATM activation. A number of possibilities could explain the discrepancies on whether a single phosphorylated residue is sufficient to activate ATM. It is possible that a combination of post-translational modifications is necessary for ATM activation or that there is redundancy between sites. Phosphatases can also regulate ATM phosphorylation. Evidence is emerging that protein phosphatase 2A (PP2A) interacts with ATM to maintain S1981 in an unphosphorylated state (26). Upon activation by IR, PP2A dissociates from ATM allowing ATM to be activated. Importantly, treatment with the protein phosphatase inhibitor okadaic acid (OA) induces ATM S1981 autophosphorylation comparable to IR-induced levels, but without causing detectable DNA DSBs. However, even though phosphorylated at S1981, OA treatment is not sufficient to activate ATM kinase activity. Another phosphatase, Wip1 (PPM1D), has also been demonstrated to regulate ATM S1981 phosphorylation in vitro and in vivo and to be a critical regulator of the ATM pathway (27,28). Overexpression of Wip1 prevents activation of the ATM signaling cascade, whereas deletion of Wip1 leads to the activation of ATM. In addition to regulation by phosphorylation, ATM is also directly regulated by acetylation (29,30). Tip60, a histone acetyltransferase, interacts with the FATC domain of ATM and mediates rapid acetylation of ATM in response to IR and bleomycin (30). In this study, depletion of Tip60 was able to attenuate the ATM DNA damage response as measured by phosphorylation of ATM targets Chk2 and p53.


97

In summary, ATM activation is likely controlled by multiple posttranslational modifications, but even when activated, a functional DNA damage response relies on a concerted cooperation from other mediators of the DNA damage response pathway.

3.2. MRE11/RAD50/NBS1 Complex is a Key Regulator of ATM The MRE11/RAD50/NBS1 (MRN) complex clearly plays a role in regulating ATM in response to IR by linking ATM to the site of DNA DSBs (Fig. 1). All 3 MRN components are encoded by essential genes—loss of function of each gene is embryonic lethal in mice (31–33). Hypomorphic mutations of MRE11 and NBS1 cause ataxia-telangiectasia-like disease (ATLD) and Nijmegan breakage syndrome (NBS) respectively—diseases related to A-T. ATLD patients develop many of the symptoms of A-T, but at a later onset and with slower progression. Patients afflicted with NBS are prone to radiation sensitivity, lymphoid malignancies, and genomic instability as well as immunological and mental deficiencies. NBS1 contains two domains commonly associated with proteins involved in cell cycle regulation: a fork-head associated (FHA) domain—a phospho-residuebinding domain—and a BRCT (BRCA1 carboxy-terminal) domain that mediates protein–protein interactions. Through these domains, NBS1 is able to interact with other proteins at the site of DNA damage, such as phosphorylated H2AX (H2AX)—a histone H2A variant (34,35). Localization of NBS1 to the site of DNA damage then triggers recruitment of the MRE11-Rad50 (MR) complex to NBS1. Rad50 belongs to the structural maintenance of chromosomes (SMC) family of proteins. Rad50 forms antiparallel homodimers that form a flexible hinge region. This configuration allows Rad50 to interact with broken DNA ends via its Walker binding motifs to keep the ends in close proximity thereby facilitating DNA repair (36). MRE11 contains a phosphoesterase domain that exhibits single- and double-stranded DNA endonuclease activity, and can act as a 3 –5 double-stranded DNA exonuclease. The MRN complex therefore has the ability to be recruited to the site of DNA damage, interact with the damaged DNA ends, and process the ends. NBS1 regulates two important functions of the MRN complex—it targets MRN to the site of DNA damage and recruits and activates ATM. Elegant in vitro studies conducted by Lee and Paull demonstrated that dimeric ATM requires both the MRN complex and DNA for significant ATM activity, as measured by phosphorylation of the

98


DNA DSB

DNA replication stress

ATR ATM

ATRIP

ATR

TopBP1

ATRIP

Rad50 S1 NB

MRE11

53BP1

Rad17 BRCA1

H2AX

RPA

DNA

9-1-1

Cell cycle arrest DNA repair Apoptosis

Fig. 1. Activation of the ATM and ATR Pathways. ATM is activated by ionizing radiation and DNA damaging agents that cause DNA double-strand breaks (DSB). Upon activation, ATM is recruited to the DSB by the MRN complex (MRE11Rad50-NBS1) where it associates with other regulators including 53BP1, H2AX, and BRCA1. Evidence suggests that changes in chromatin structure can also activate ATM, but through a different mechanism (see text for details). In response to DNA replication stress, ATR is activated. ATR exists as a heterodimer with ATRIP. After stimulation, ATR-ATRIP relocalizes to the site of DNA damage. RPA coats the DNA ends to protect them from degradation and from forming inhibitory secondary structures. ATR interacts with several proteins at the site of damage including Rad17 and 9-1-1 (Rad9-Hus1-Rad1). ATR-ATRIP also transiently interacts with TopBP1, which regulates the activity of ATR. An increasing number of reports demonstrate crosstalk between the ATM and ATR pathways. It is unlikely that these pathways function in a mutually exclusive fashion (see text for details). Activation of ATM and ATR results in cell cycle arrest, DNA repair, or apoptosis.

ATM substrates p53 and Chk2 (23). As a monomer, ATM efficiently phosphorylates its substrates independent of the presence of MRN or DNA, but the MRN complex is necessary for ATM binding to DNA. The FHA domain of NBS1 targets the MRN complex to H2AX at the site of DNA DSBs (35). However, the FHA domain of NBS1 is reportedly not essential for ATM activation (37–40). This suggests that either NBS1 can be recruited to the site of DNA damage independent of its FHA domain or that ATM can be partially activated without translocating to the site of DNA damage. Because the FHA domain of NBS has been demonstrated to be essential for colocalization with H2AX


99

at the site of IR-induced DNA DSBs (40), the latter is more likely to be true. Indeed, exposure to hypotonic buffer, which alters chromatin structure but does not cause DNA damage, activates ATM (41). ATM remains diffuse—it does not localize to foci. Hypotonic buffer also has been shown to activate ATM independently of MRN (40). ATM recruitment to the site of DNA damage is also dependent on the MRN complex. Whereas the NBS1 N-terminus FHA domain regulates the interaction between MRN and DNA, the C-terminus of NBS1 has been implicated in regulating the interaction of MRN with ATM (42,43). However, in vitro data suggests a role for Rad50 in recruiting ATM to the site of DSBs (23).

3.3. ATM is Differentially Activated Most reports regarding ATM activation have specifically focused on IR-induced ATM activation, but it is becoming apparent that the mechanism of ATM activation is dependent upon the inducing agent. IR causes DNA DSBs, but ATM is activated by agents that modify chromatin structure even if DNA damage does not occur. In vitro IRinduced phosphorylation of ATM Ser-1981 is the direct result of DNA double strand breaks that can be recapitulated in cells by introducing the restriction enzyme I-Sce 1 (19). However, chloroquine or hypotonic buffer—agents known to alter higher-order chromatin structure but not cause DNA breaks—induce phosphorylation at S1981. Additionally, treating G0 fibroblasts with camptothecin, a topoisomerase I poison, activates ATM in the absence of DNA strand breaks (41). After IR exposure, phosphorylated S1981 ATM relocates to discrete foci at the site of DNA damage (19,22,41). However, after treatment with chloroquine or hypotonic buffer, ATM undergoes phosphorylation at S1981 but it remains diffuse (41). This data provides evidence that the formation of DNA DSBs is not necessary for ATM activation—it can also be activated through alteration in chromatin structure (which can be the result of DNA damage). These findings raised interesting questions. It was believed that once recruited to the site of DNA damage ATM functions to activate downstream effectors. How is ATM activated in the absence of DNA damage and what is its function? It is evident that ATM regulation and function is even more complex than initially appreciated. Some insight into the regulation of ATM by different agents has recently come to light. DNA damaging and chromatin modifying agents differentially regulate ATM, but surprisingly, DNA damaging agents also differentially regulate ATM. Difilippantonio et al. generated transgenic mice that lack the endogenous mouse Nbs1 gene, but express

100


either wild-type or mutant forms of the human Nbs1 gene (40). NBS1 is required for ATM S1981 autophosphorylation in response to IR, but NBS1 is not required for ATM S1981 autophosphorylation induced by replication stress, UV, hydroxyurea, or aphidicolin even though DNA DSBs are detected (monitored by -H2AX) after treatment. Hypotonic conditions, which alter chromatin structure but do not induce DNA DSBs, do not require NBS1 to activate ATM. A recent report describes the colocalization of ATMIN (ATM Interacting protein) and ATM in response to chloroquine and hypotonic stress, but not IR (44). Exposure to IR interferes with the ATMINATM association, however IR has no effect on the ATMIN-ATM association in cells with impaired NBS1. This suggests that NBS1 and ATMIN compete for ATM. Phosphorylation of the ATM targets SMC1, Chk2, and p53 is impaired in ATMIN-deficient resting cells and chloroquine and hypotonic stress-induced cells; IR-induced phosphorylation of these targets is unaffected in ATMIN-deficient cells. Together this suggests a model in which ATM is activated by different regulatory proteins in response to different stimuli—NBS1 regulates ATM in response to IR, ATMIN in response to chloroquine and hypotonic stress.

3.4. ATM Activates Targets at the site of DNA Damage Once recruited to the site of DNA damage and activated, ATM initiates activation of downstream pathways through phosphorylation of a number of key mediators including p53, Mdm2, H2AX, NBS1, SMC1, BRCA1, MDC1, and Chk2. Activation of these pathways ultimately leads to a functional response to DNA damage—cell cycle arrest, apoptosis, or DNA repair. The MRN complex is necessary to recruit ATM to the site of DNA damage and then becomes an ATM substrate itself. The NBS1 component of MRN is phosphorylated at serines 278, 343, 397, and 615 in response to IR in an ATM-dependent manner, which is necessary for the activation of the S-phase checkpoint (45–49). MDC1 (Mediator of DNA damage Checkpoint protein 1) may function to retain ATM at the site of DNA damage after recruitment by MRN. MDC1 regulates IR-induced apoptosis and the intra-S and G2 /M checkpoints (50,51). MDC1 contains a BRCT and FHA domain that enables it to interact with H2AX and ATM, respectively (52). One can envision a model in which MDC1 assists in anchoring ATM to the site of DNA damage thereby enabling ATM to phosphorylate targets and to amplify the DNA damage response. Recently, it has been demonstrated that the reduction of RNA polymerase I (Pol I) activity


101

induced by DNA damage is not the direct result of the DNA damage, but Pol I transcription is inhibited by an ATM/NBS1/MDC1-dependent pathway. At the site of DNA damage, ATM is known to regulate a number of proteins. ATM phosphorylates H2AX on serine 139—a (S/T)-Q motif (53). 53BP1 (p53 Binding Protein 1) is also phosphorylated by ATM on several residues in response to IR (54,55). Both 53BP1 and H2AX are detected at the sites of DNA DSBs. Mice lacking either H2AX (56) or 53BP1 (57) have increased susceptibility to cancer and defective cell cycle checkpoints. H2AX and 53BP1 are clearly involved in the DNA damage response (1) and there is great interest to identify the specific roles of these proteins in response to DNA damage. BRCA1 is also found co-localized with MRN, H2AX, and 53BP1 at IR-induced foci and is a target of ATM phosphorylation. Germline mutations in BRCA1 render the carrier susceptible to breast cancer—a 50–80% risk of developing breast cancer by the age of 70 (58). BRCA1 functions in both cell cycle checkpoints and DNA repair; mutations to BRCA1 thereby lead to genetic instability. Serine residues 988, 1387, 1423, 1497, and 1524 of BRCA1 are phosphorylated in an ATM-dependent manner in response to IR (59). Interestingly, differential phosphorylation of BRCA1 selectively regulates its involvement in different checkpoints. Phosphorylation of S1387 is required for the S-phase checkpoint (59), whereas S1423 mediates the G2 /M checkpoint (60). Additionally, after IR ATM phosphorylates the BRCA1 interacting protein CtIP that causes it to dissociate from BRCA1 (61). This results in transcriptional derepression of genes required for activation of the G2 /M checkpoint. To date, twelve mutations in ATM that cause A-T have also been identified as breast cancer susceptibility alleles, further implicating the relationship between BRCA1 and ATM (62).

3.5. p53 is a Major ATM Target The regulation of p53 by ATM is the best known pathway by which ATM regulates the cell cycle (63). p53 is a transcription factor that regulates the ATM-induced G1 /S checkpoint. One target of p53 transcriptional activity is the cyclin-dependent kinase inhibitor p21waf1/cip1 , which prevents the cyclin E/Cdk2 complex from promoting the G1 /S transition. The activity of p53 is intimately controlled by the E3 ubiquitin ligase Mdm2. Binding of Mdm2 to p53 conceals the transactivating domain of p53 and exports p53 from the nucleus to the cytoplasm, thus preventing the transcriptional activity of p53. Mdm2 also targets p53 for degradation via the ubiquitin-proteasome pathway.

102


Through a negative feedback loop, p53 is able to regulate itself. Mdm2 is a transcriptional target of p53 and when p53 levels are elevated, p53 is responsible for increasing the transcription of Mdm2, which in turn reduces p53 levels. ATM directly phosphorylates serine 15 on p53 in vitro, which enhances its transcriptional activity (20,64) and IR-induced phosphorylation of this residue is ATM-dependent in vivo (65,66). ATM is also responsible for indirect phosphorylation of p53 through Chk2. IR induces ATM-mediated phopshorylation of Chk2 on threonine 68 (67); activated Chk2 subsequently phosphorylates p53 on serine 20, which disrupts the interaction between Mdm2 and p53 (68,69). ATM also regulates p53 through Mdm2 (Mdm2 in mice; Hdm2 in humans) and the related protein Mdmx (Mdmx or Mdm4 in mice; Hdmx in humans) (Fig. 2). ATM phosphorylates Hdm2 on serine 395, which prevents Hdm2-mediated degradation on p53 (70,71). Mdmx is structurally related to Mdm2. It lacks the ubiquitin ligase activity of Mdm2, however, it binds to p53 and inhibits its transcriptional activity (72). Mdmx also interacts with Mdm2 through their respective RING finger domains which stabilizes Mdm2; this interaction is postulated to enhance Mdm2-mediated inhibition of p53 (73). IR-induced DNA damage leads to ATM-dependent phosphorylation of Mdmx at serine 342, serine 367, and serine 403 resulting in enhancement of p53 activity (74,75). ATM directly phosphorylates S403 (75); Chk2 is responsible for S342 and S367 phosphorylation (74). Chen et al. also demonstrated that Mdm2 targets Mdmx for degradation after IR-induced DNA damage, which enhances the p53 response. Interestingly, another E3 ubiquitin ligase known to regulate p53, COP1, is also phosphorylated by ATM (76). DNA damaging agent-induced ATM-dependent phosphorylation of COP1 at serine 387 disrupts the COP1-p53 complex thereby stabilizing p53 (Fig. 2). In response to DNA damage, p53 regulates ATM through a very intricate network including direct and indirect regulation—this highlights the importance of p53 and the G1 /S checkpoint in the DNA damage response.

3.6. ATM Substrates Regulate Cell Cycle Checkpoints In addition to the ATM substrates previously discussed, the mechanisms of regulation for several other ATM substrates with roles in cell cycle control are also established. Previously, it was believed that ATM and ATR signal through the downstream checkpoint kinases Chk2 and Chk1, respectively. However, ATM has been demonstrated to signal through both Chk2 and Chk1 to regulate Cdc25A (77–79).

Chapter 4 / Targeting ATM/ATR in the DNA Damage Checkpoint A.

B.

C. ATM

ATM P S15 p53

P T68 Chk2

p53 enhanced transcriptional activity

Mdm2

p53

increased activity

D.

S367 P

Chk2

S342 P p53

Mdm2

P S403 Mdmx

destabilize Mdm2 increased p53 activity

inhibition, degradation

E. ATM

p53

ATM

Chk2 P

103

ATM P S395

Mdm2

inhibition, degradation

Mdm2

p53 increased activity

p53 COP1

degradation

P S387 COP1

p53 increased activity

Fig. 2. ATM Regulation of p53. The multiple mechanisms by which ATM regulates p53 reflect the importance of p53 in the DNA damage response. ATM directly phosphorylates p53 at S15 to enhance p53 transcriptional activity (A). ATM indirectly and directly regulates the E3 ubiquitin ligase Mdm2 to control p53 activity. Chk2 is phosphorylated by ATM at T68, which stimulates Chk2 kinase activity. In turn, Chk2 phosphorylates p53 at S20, disrupting the interaction of Mdm2 with p53, thereby increasing p53 activity (B). Mdmx interacts with Mdm2 to enhance Mdm2 stability. Phosphorylation of Mdmx at S403 by ATM and S342, S367 by Chk2 leads to the destabilization of Mdm2 and subsequent increase in p53 activity (C). ATM can also enhance p53 activity by direct phosphorylation of Mdm2 at S395 (D). Like Mdm2, COP1 is an E3 ubiquitin ligase with activity against p53. Phosphorylation of COP1 at S387 disrupts the interaction between COP1 and p53 resulting in elevated p53 (E).

Cdc25A, along with Cdc25B and Cdc25C, comprise the Cdc25 family of phosphatases. These phosphatases remove the inhibitory phosphates from the appropriate cyclin dependent kinase (Cdk), thus promoting cell cycle progression. Cdc25A was initially believed to be involved exclusively in G1 and S phases, but is now known to be essential in G2 /M as well (80). In response to IR, ATM phosphorylates Chk2 (77) and Chk1 (79), which in turn, phosphorylate Cdc25A. The phosphorylation of Cdc25A accelerates its proteolysis, which can lead to intra-S-phase or G2 /M arrest. DNA damage induced Chk1 and Chk2 mediated phosphorylation of Cdc25 family members also creates a 14-3-3 binding site (80). 14-3-3 sequesters the Cdc25 member, thus

104


preventing it from removing the inhibitory phosphates from the Cdk. Other agents, including UV (81) and the topoisomerase inhibitors camptothecin and doxorubicin (82), also enhance Chk1-mediated Cdc25A degradation, although the involvement of ATM or ATR was not investigated. Furthermore, in response to DNA damage, evidence suggests that Chk2 phosphorylates Cdc25C on S216, in an ATMdependent manner (83). This inactivating phosphorylation of Cdc25C prevents G2 /M progression. A number of other studies describe DNA damage-induced Chk1 and Chk2 regulation of Cdc25 phosphatases (reviewed in (83)), however the involvement of ATM and ATR was not investigated. SMC1 (Structural Maintenance of Chromosomes 1) is a component of the cohesin complex that is required for the maintenance of sister chromatid cohesion during DNA replication and mitosis. In response to almost every type of DNA damage, SMC is phosphorylated at two serines, S957 and S966 (41). ATM phosphorylates these sites after IR; ATR is responsible for phosphorylating SMC1 in response to UV and hydroxyurea. In mouse fibroblasts that express SMC1 in which S957 and S966 have been mutated to alanines (SMC1-S957A,S966A cells), phosphorylated ATM, H2AX, NBS1, 53BP1, BRCA1, and T68 phosphorylated Chk2 are found in foci at the site of DNA damage (84). This suggests that SMC1 is downstream of these proteins, and in fact, SMC1 phosphorylation is dependent on ATM, NBS1, and BRCA1. SMC1-S957A,S966A cells have a defective intra-S-phase checkpoint in response to IR, are hypersensitive to DNA damaging agents, and exhibit phenotypic traits similar to ATM-deficient cells (41). It is clear that ATM-dependent phosphorylation of SMC1 in response to DNA damage leads to DNA repair but the details remain to be elucidated. The ATM target NEMO has emerged as key regulator in the DNA damage response. NEMO is the regulatory subunit of the IKK complex that activates NF-B by negatively regulating IB, an inhibitor of NF-B. The NF-B family of transcription factors regulate immune and inflammatory responses, cell migration, and apoptosis (85). In response to DNA damaging agents, ATM phosphorylates NEMO, which then shuttles NF-B to the cytoplasm where it interacts with IKK and its regulatory subunit ELKS (86). Activated IKK phosphorylates IB which frees NF-B to regulate its numerous target genes. This represents a new, distinct pathway regulated by ATM in response to DNA damage. Overall, activated ATM is known to directly or indirectly phosphorylate over 30 substrates (5,45). New targets are emerging, including


105

the polo-like kinases, which regulate entry and exit from the cell cycle (for review see (87)). Proteomic schemes have been implemented to identify novel ATM and ATR targets (88,89). Matsuoka et al. identified over 700 proteins apparently phosphorylated by ATM, ATR, or DNAPK. Analysis of a subset of these revealed that more than 90% of the proteins were indeed involved in the DNA damage response. Approximately half of the assigned functions for the proteins with known functions fell into 3 categories: (1) nucleoside, nucleotide, and nucleic acid metabolism (including mRNA transcription and DNA replication, recombination, and repair), (2) protein metabolism and modification, and (3) cell cycle. Proteomic schemes, such as this, carry the potential to accelerate the identification of currently unknown ATM and ATR substrates and pathways that will greatly assist in forming a more complete picture of the DNA damage response.

3.7. ATR Unlike ATM, ATR is an essential gene in mice—loss of ATR is embryonic lethal (90). ATR mutations have been associated with the human disorder, Seckels syndrome. A mutation within ATR results in hypomorphic expression of ATR (91). This disease is characterized by growth retardation, microcephaly, and mental retardation; clinical sensitivity to DNA damaging agents and cancer predisposition have not been described in these patients (91). Human cells and mouse embryonic fibroblasts, in which ATR has been disrupted, undergo cell cycle arrest or apoptosis even in the absence of exogenous DNA damage (92,93). Likewise disruption of Chk1, a downstream target of ATR, in mice is also embryonic lethal, which further suggests the importance of this pathway (94,95). Whereas DNA damaging agentand IR-induced DSBs have been identified as the primary activator of ATM, ATR responds to a broader spectrum of DNA insults. DNA replication interference, including stalled replication forks, is the primary activator of ATR (see reviews (93,96)).

3.7.1. Activation of ATR ATR exists as a stable heterodimeric complex with ATR-interacting protein (ATRIP) (92) (Fig. 1). The heterodimer does not undergo apparent phosphorylation or dissociation in response to DNA replication stress, and constitutively maintains the ability to phosphorylate its substrates (42,97,98). This led to a hypothesis that ATR is

106


regulated spatially. Upon DNA insult, ATR relocalizes to the stalled replication fork or site of DNA damage where it interacts with its substrates. DNA repair and stressed DNA replication forks lead to the formation of single-stranded DNA and double-stranded/singlestranded junctions (96). Replication protein A (RPA) coats the singlestranded DNA ends protecting them from nucleases, further DNA damage, and prevents the formation of inhibitory secondary structures (99). ATR-ATRIP and other proteins, including Rad17 and the PCNA-related Rad9-Hus1-Rad1 (9-1-1) complex, are recruited to the site of RPA-coated damaged DNA (97,100). Several phosphorylation events take place at these foci. In vitro, Rad17 and RPA are phosphorylated by ATR (97,101,102). It is not known whether these proteins are phosphorylated in vivo or what importance it may carry— it may assist in recruiting other proteins. ATR also phosphorylates ATRIP (103). TopBP1, which contains a BRCA1 carboxy-terminal (BRCT) domain, directly interacts with ATR in an ATRIP-dependent manner (104). Phosphorylation of ATRIP by ATR conceivably facilitates the interaction of the complex with TopBP1. ATR phosphorylates TopBP1, which strongly stimulates the kinase activity of ATR (104,105). The interaction between TopBP1 with ATR-ATRIP appears to be transient and therefore difficult to detect experimentally. Depletion of TopBP1 does not affect the relocalization of ATR-ATRIP to sites of DNA damage, but it significantly impairs phosphorylation of ATR targets, including Chk1, Nbs1, Smc1, and H2AX (106). Claspin is also phosphorylated by ATR and participates downstream of TopBP1 to activate Chk1, but is not required for activation of other ATR targets (106,107). Together these findings suggest that TopBP1 is an important mediator of ATR activity and, downstream of TopBP1, distinct pathways exist to activate specific ATR targets. As previously noted, ATR is activated by replication stress, which can be induced by UV, HU, and MMS, to control cell cycle checkpoints. ATR plays a central role in the intra-S phase replication checkpoint (93). ATR activates Chk1 resulting in downstream activation of checkpoint kinases that block cell cycle progression. ATR has also been implicated in the G1 /S and G2 /M checkpoints. In response to UV, ATR-activated Chk1phosphorylates Cdc25A prevents the activation of the cyclinE/Cdk2 complex necessary for G1 to S progression—similar to ATM-Chk2 control of Cdc25A discussed previously (81). Analogous to the ATR-Chk2-Cdc25A pathway, ATRChk2 can regulate the G2 /M checkpoint through phosphorylation of Cdc25C, resulting in the inhibition of Cdk1 activity (108).


107

3.7.2. ATM and ATR Signaling Pathways: Mutually Exclusive or Inclusive? Undoubtedly, ATR is essential for the DNA damage response associated with replication stress. Likewise, ATM is activated in response to DSBs that are not associated with replication stress, such as DSB induced by IR and radiomimetic chemicals. Chk1 predominantly mediates the ATR pathway, whereas Chk2 is the foremost mediator of ATM signaling. However, as we have gained a better understanding of the DNA damage response, it has become apparent that these pathways are not mutually exclusive. ATM and ATR phosphorylate many of the same substrates including p53, H2AX, BRCA1, 53BP1, Chk1 and Chk2 (17,55,67,78,109). Indeed, ATR can phosphorylate Chk2 in response to IR and UV even in the absence of ATM (109). Reciprocally, IR-induced ATM can activate Chk1 (78). A growing amount of evidence demonstrates that ATR is activated by DNA DSBs caused by IR and DNA damaging agents (109–113). In this case, activation of ATR is ATM dependent and the MRN complex is also necessary (110–112). Furthermore, ATM has been found to phosphorylate TopBP1 (113). Large stretches of RPA coated single-stranded DNA are detected at the site of damage (110,111). ATR relocalizes to these sites and phosphorylates Chk1 (111). Taken together, these findings suggest a model in which ATM activates the ATR pathway in a similar fashion to UV and replication stress. Possibly, the DNA ends at the IRinduced DSB are processed to resemble the structure generated by UV and replication stress. ATM has been shown to be activated in the canonical ATR pathway. In response to UV and replication fork stalling, ATM is phosphorylated at S1981 and activated by an ATR-dependent mechanism (114). In conclusion, ATM and ATR share many of the same substrates and both are found activated in response to DNA damage thought to stimulate one pathway or the other.

4. PROSPECTS OF REGULATING ATM/ATR PATHWAYS Unquestionably, ATM and ATR are central to the DNA damage response, which is activated in the early stages of tumorigenesis (1,2). The cell senses damaged DNA, arrests until the DNA is repaired, or is eliminated by apoptosis thus preventing it from replicating. However, when defects in the DNA damage response arise cells with DNA lesions can elude detection and promote tumorigenesis. Defects in ATM and associated proteins result in diseases including A-T, ATLD,

108


and NBS. Likewise, ATR is an essential gene in mice and hypomorphic expression is associated with Seckels syndrome. ATM/ATR activate known tumor suppressors, p53 and BRCA1. Therefore, restoring functional ATM/ATR pathways would be beneficial, albeit therapeutically difficult to accomplish. Targeted inhibition of ATM/ATR pathways is likely to yield more fruitful results. As well as being a cause of cancer, DNA damage is used to treat cancer—radiation and cytotoxic DNA damaging agents are widely used cancer treatments. Cells lacking functional ATM/ATR pathways are radiosensitive. Therefore, one strategy is to inhibit ATM/ATR pathways to enhance the sensitivity to DNA damaging cancer therapies. A number of key proteins involved in ATM/ATR pathways are protein kinases, which make good drug targets. Over 500 protein kinases are encoded by the human genome (115) and share a conserved catalytic domain (116). This “druggable” ATP binding pocket has been the focus of programs aimed at developing protein kinase inhibitors. Because the ATP binding pocket is conserved among protein kinases, many existing protein kinase inhibitors have broad action against the family of protein kinases. These ATP binding site inhibitors include the competitive inhibitor quercetin and its derivative LY294002 and the natural products wortmannin and demethoxyviridin, which are irreversible inhibitors (see reviews (87,117)). An additional ATP-competitive inhibitor, KU-55933, is reportedly specific to ATM (118). Caffeine, a well known disruptor of the cell cycle, has been demonstrated to inhibit ATM, ATR, and Chk2 (119,120). A number of other Cdk inhibitors also exist, including flavopridol, indirubin, and roscovitine. A small peptide that disrupts the DNA damage response has recently been described (121). This peptide, which contains the C-terminal sequence of NBS1, abrogates the DNA damage response in an ATM-dependent manner and enhances radiosensitivity. The most promising results have come from UCN01—a small molecule known to inhibit Chk1, Cdk2/cyclinA, and PDK1 (116). UCN-01 is currently undergoing clinical trials for the treatment of leukemia and other cancers. In targeting a pathway for therapeutic purposes, it is a definite advantage to know as much possible about the pathway—how it is activated and regulated, what are the downstream targets, and what are the functions of these targets. The discovery of ATM, and subsequently ATR, has been instrumental in understanding the DNA damage response pathway. The challenge remains to understand additional details of the pathway, particularly elucidating the overlap between


109

the ATM and ATR pathways, defining how DNA damage is sensed, and identifying the scope of proteins involved in the DNA damage response.

REFERENCES 1. Gorgoulis VG, Vassiliou LV, Karakaidos P, et al. Activation of the DNA damage checkpoint and genomic instability in human precancerous lesions. Nature 2005; 434:907–913. 2. Bartkova J, Horejsi Z, Koed K, et al. DNA damage response as a candidate anti-cancer barrier in early human tumorigenesis. Nature 2005; 434:864–870. 3. Hecht F, Koler RD, Rigas DA, et al. Leukaemia and lymphocytes in ataxia-telangiectasia. Lancet 2 1966; 1193. 4. McKinnon PJ. ATM and ataxia telangiectasia. EMBO Rep 2004; 5:772–5776. 5. Lavin MF, Delia D, Chessa L. ATM and the DNA damage response. Workshop on ataxia-telangiectasia and related syndromes. EMBO Rep 2006; 7:154–160. 6. Savitsky K, Bar-Shira A, Gilad S, et al. A single ataxia telangiectasia gene with a product similar to PI-3 kinase. Science 1995; 268:1749–1753. 7. Hammond EM, Denko NC, Dorie MJ, Abraham RT, Giaccia AJ. Hypoxia links ATR and p53 through replication arrest. Mol Cell Biol 2002; 22:1834–1843. 8. Hammond EM, Dorie MJ, Giaccia AJ. ATR/ATM targets are phosphorylated by ATR in response to hypoxia and ATM in response to reoxygenation. J Biol Chem 2003; 278:12207–12213. 9. Hammond EM, Giaccia AJ. The role of ATM and ATR in the cellular response to hypoxia and re-oxygenation. DNA Repair (Amst) 2004; 3:1117–1122. 10. Hammond EM, Dorie MJ, Giaccia AJ. Inhibition of ATR leads to increased sensitivity to hypoxia/reoxygenation. Cancer Res 2004; 64:6556–6562. 11. Hammond EM, Freiberg RA, Giaccia AJ. The roles of Chk 1 and Chk 2 in hypoxia and reoxygenation. Cancer Lett 2006; 238:161–167. 12. Gibson SL, Bindra RS, Glazer PM. Hypoxia-induced phosphorylation of Chk2 in an ataxia telangiectasia mutated-dependent manner. Cancer Res 2005; 65:10734–10741. 13. Gibson SL, Bindra RS, Glazer PM. CHK2-dependent phosphorylation of BRCA1 in hypoxia. Radiat Res 2006; 166:646–651. 14. Purdy A, Su TT. Telomeres: not all breaks are equal. Curr Biol 2004; 14:R613–614.

110


15. Slijepcevic P. The role of DNA damage response proteins at telomeres– an “integrative” model. DNA Repair (Amst) 2006; 5:1299–1306. 16. Denchi EL, de Lange T. Protection of telomeres through independent control of ATM and ATR by TRF2 and POT1. Nature 2007; 448: 1068–1071. 17. Goodarzi AA, Block WD, Lees-Miller SP. The role of ATM and ATR in DNA damage-induced cell cycle control. Prog Cell Cycle Res 2003; 5:393–411. 18. Bosotti R, Isacchi A, Sonnhammer EL. FAT: a novel domain in PIKrelated kinases. Trends Biochem Sci 2000; 25:225–227. 19. Bakkenist CJ, Kastan MB. DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature 2003; 421:499–506. 20. Canman CE, Lim DS, Cimprich KA, et al. Activation of the ATM kinase by ionizing radiation and phosphorylation of p53. Science 1998; 281:1677–1679. 21. Cuadrado M, Martinez-Pastor B, Fernandez-Capetillo O. “ATR activation in response to ionizing radiation: still ATM territory”. Cell Div 2006; 1:7. 22. Kozlov SV, Graham ME, Peng C, Chen P, Robinson PJ, Lavin MF. Involvement of novel autophosphorylation sites in ATM activation. Embo J 2006; 25:3504–3514. 23. Lee JH, Paull TT. ATM activation by DNA double-strand breaks through the Mre11-Rad50-Nbs1 complex. Science 2005; 308:551–554. 24. Dupre A, Boyer-Chatenet L, Gautier J. Two-step activation of ATM by DNA and the Mre11-Rad50-Nbs1 complex. Nat Struct Mol Biol 2006; 13:451–457. 25. Pellegrini M, Celeste A, Difilippantonio S, et al. Autophosphorylation at serine 1987 is dispensable for murine Atm activation in vivo. Nature 2006; 443:222–225. 26. Goodarzi AA, Jonnalagadda JC, Douglas P, et al. Autophosphorylation of ataxia-telangiectasia mutated is regulated by protein phosphatase 2A. Embo J 2004; 23:4451–4461. 27. Shreeram S, Hee WK, Demidov ON, et al. Regulation of ATM/p53-dependent suppression of myc-induced lymphomas by Wip1 phosphatase. J Exp Med 2006; 203:2793–2799. 28. Shreeram S, Demidov ON, Hee WK, et al. Wip1 phosphatase modulates ATM-dependent signaling pathways. Mol Cell 2006; 23:757–764. 29. Jiang X, Sun Y, Chen S, Roy K, Price BD. The FATC domains of PIKK proteins are functionally equivalent and participate in the Tip60dependent activation of DNA-PKcs and ATM. J Biol Chem 2006; 281:15741–15746.


111

30. Sun Y, Jiang X, Chen S, Fernandes N, Price BD. A role for the Tip60 histone acetyltransferase in the acetylation and activation of ATM. Proc Natl Acad Sci U S A 2005; 102:13182–13187. 31. Xiao Y, Weaver DT. Conditional gene targeted deletion by Cre recombinase demonstrates the requirement for the double-strand break repair Mre11 protein in murine embryonic stem cells. Nucleic Acids Res 1997; 25:2985–2991. 32. Luo G, Yao MS, Bender CF, et al. Disruption of mRad50 causes embryonic stem cell lethality, abnormal embryonic development, and sensitivity to ionizing radiation. Proc Natl Acad Sci U S A 1999; 96:7376–7381. 33. Zhu J, Petersen S, Tessarollo L, Nussenzweig A. Targeted disruption of the Nijmegen breakage syndrome gene NBS1 leads to early embryonic lethality in mice. Curr Biol 2001; 11:105–109. 34. Tauchi H, Kobayashi J, Morishima K, et al. The forkhead-associated domain of NBS1 is essential for nuclear foci formation after irradiation but not essential for hRAD50[middle dot]hMRE11[middle dot]NBS1 complex DNA repair activity. J Biol Chem 2001; 276:12–15. 35. Kobayashi J, Tauchi H, Sakamoto S, et al. NBS1 localizes to gammaH2AX foci through interaction with the FHA/BRCT domain. Curr Biol 2002; 12:1846–1851. 36. D’Amours D, Jackson SP. The Mre11 complex: at the crossroads of dna repair and checkpoint signalling. Nat Rev Mol Cell Biol 2002; 3:317–327. 37. Horejsi Z, Falck J, Bakkenist CJ, Kastan MB, Lukas J, Bartek J. Distinct functional domains of Nbs1 modulate the timing and magnitude of ATM activation after low doses of ionizing radiation. Oncogene 2004; 23:3122–3127. 38. Lee JH, Xu B, Lee CH, et al. Distinct functions of Nijmegen breakage syndrome in ataxia telangiectasia mutated-dependent responses to DNA damage. Mol Cancer Res 2003; 1:674–681. 39. Cerosaletti KM, Concannon P. Nibrin forkhead-associated domain and breast cancer C-terminal domain are both required for nuclear focus formation and phosphorylation. J Biol Chem 2003; 278:21944–21951. 40. Difilippantonio S, Celeste A, Fernandez-Capetillo O, et al. Role of Nbs1 in the activation of the Atm kinase revealed in humanized mouse models. Nat Cell Biol 2005; 7:675–685. 41. Kitagawa R, Kastan MB. The ATM-dependent DNA damage signaling pathway. Cold Spring Harb Symp Quant Biol 2005; 70:99–109. 42. Falck J, Coates J, Jackson SP. Conserved modes of recruitment of ATM, ATR and DNA-PKcs to sites of DNA damage. Nature 2005; 434: 605–611.

112


43. You Z, Chahwan C, Bailis J, Hunter T, Russell P. ATM activation and its recruitment to damaged DNA require binding to the C terminus of Nbs1. Mol Cell Biol 2005; 25:5363–5379. 44. Kanu N, Behrens A. ATMIN defines an NBS1-independent pathway of ATM signalling. Embo J 2007; 26:2933–2941. 45. Kastan MB, Lim DS. The many substrates and functions of ATM. Nat Rev Mol Cell Biol 2000; 1:179–186. 46. Zhao S, Weng YC, Yuan SS, et al. Functional link between ataxiatelangiectasia and Nijmegen breakage syndrome gene products. Nature 2000; 405:473–477. 47. Wu X, Ranganathan V, Weisman DS, et al. ATM phosphorylation of Nijmegen breakage syndrome protein is required in a DNA damage response. Nature 2000; 405:477–482. 48. Lim DS, Kim ST, Xu B, et al. ATM phosphorylates p95/nbs1 in an S-phase checkpoint pathway. Nature 2000; 404:613–617. 49. Gatei M, Young D, Cerosaletti KM, et al. ATM-dependent phosphorylation of nibrin in response to radiation exposure. Nat Genet 2000; 25:115–119. 50. Goldberg M, Stucki M, Falck J, et al. MDC1 is required for the intraS-phase DNA damage checkpoint. Nature 2003; 421:952–956. 51. Stewart GS, Wang B, Bignell CR, Taylor AM, Elledge SJ. MDC1 is a mediator of the mammalian DNA damage checkpoint. Nature 2003; 421:961–966. 52. Lou Z, Minter-Dykhouse K, Franco S, et al. MDC1 maintains genomic stability by participating in the amplification of ATM-dependent DNA damage signals. Mol Cell 2006; 21:187–200. 53. Rogakou EP, Pilch DR, Orr AH, Ivanova VS, Bonner WM. DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 1998; 273:5858–5868. 54. Ward IM, Minn K, Jorda KG, Chen J. Accumulation of checkpoint protein 53BP1 at DNA breaks involves its binding to phosphorylated histone H2AX. J Biol Chem 2003; 278:19579–19582. 55. Jowsey P, Morrice NA, Hastie CJ, McLauchlan H, Toth R, Rouse J. Characterisation of the sites of DNA damage-induced 53BP1 phosphorylation catalysed by ATM and ATR. DNA Repair (Amst) 2007. 56. Celeste A, Difilippantonio S, Difilippantonio MJ, et al. H2AX haploinsufficiency modifies genomic stability and tumor susceptibility. Cell 2003; 114:371–383. 57. Ward IM, Minn K, van Deursen J, Chen J. p53 Binding protein 53BP1 is required for DNA damage responses and tumor suppression in mice. Mol Cell Biol 2003; 23:2556–2563. 58. Deng CX. BRCA1: cell cycle checkpoint, genetic instability, DNA damage response and cancer evolution. Nucleic Acids Res 2006; 34:1416–1426.


113

59. Xu B, O’Donnell AH, Kim ST, Kastan MB. Phosphorylation of serine 1387 in Brca1 is specifically required for the Atm-mediated S-phase checkpoint after ionizing irradiation. Cancer Res 2002; 62:4588–4591. 60. Xu B, Kim S, Kastan MB. Involvement of Brca1 in S-phase and G(2)-phase checkpoints after ionizing irradiation. Mol Cell Biol 2001; 21:3445–3450. 61. Li S, Ting NS, Zheng L, et al. Functional link of BRCA1 and ataxia telangiectasia gene product in DNA damage response. Nature 2000; 406:210–215. 62. Renwick A, Thompson D, Seal S, et al. ATM mutations that cause ataxia-telangiectasia are breast cancer susceptibility alleles. Nat Genet 2006; 38:873–875. 63. Lvine AJ. p53, the cellular gatekeeper for growth and division. Cell 1997; 88:323–331. 64. Banin S, Moyal L, Shieh S, et al. Enhanced phosphorylation of p53 by ATM in response to DNA damage. Science 1998; 281:1674–1677. 65. Khanna KK, Keating KE, Kozlov S, et al. ATM associates with and phosphorylates p53: mapping the region of interaction. Nat Genet 1998; 20:398–400. 66. Siliciano JD, Canman CE, Taya Y, Sakaguchi K, Appella E, Kastan MB. DNA damage induces phosphorylation of the amino terminus of p53. Genes Dev 1997; 11:3471–3481. 67. Matsuoka S, Rotman G, Ogawa A, Shiloh Y, Tamai K, Elledge SJ. Ataxia telangiectasia-mutated phosphorylates Chk2 in vivo and in vitro. Proc Natl Acad Sci U S A 2000; 97:10389–10394. 68. Chehab NH, Malikzay A, Stavridi ES, Halazonetis TD. Phosphorylation of Ser-20 mediates stabilization of human p53 in response to DNA damage. Proc Natl Acad Sci U S A 1999; 96:13777–13782. 69. Hirao A, Kong YY, Matsuoka S, et al. DNA damage-induced activation of p53 by the checkpoint kinase Chk2. Science 2000; 287:1824–1827. 70. Khosravi R, Maya R, Gottlieb T, Oren M, Shiloh Y, Shkedy D. Rapid ATM-dependent phosphorylation of MDM2 precedes p53 accumulation in response to DNA damage. Proc Natl Acad Sci U S A 1999; 96:14973–14977. 71. Maya R, Balass M, Kim ST, et al. ATM-dependent phosphorylation of Mdm2 on serine 395: role in p53 activation by DNA damage. Genes Dev 2001; 15:1067–1077. 72. Marine JC, Jochemsen AG. Mdmx as an essential regulator of p53 activity. Biochem Biophys Res Commun 2005; 331:750–760. 73. Stad R, Little NA, Xirodimas DP, et al. Mdmx stabilizes p53 and Mdm2 via two distinct mechanisms. EMBO Rep 2001; 2:1029–1034. 74. Chen L, Gilkes DM, Pan Y, Lane WS, Chen J. ATM and Chk2dependent phosphorylation of MDMX contribute to p53 activation after DNA damage. Embo J 2005; 24:3411–3422.

114


75. Pereg Y, Shkedy D, de Graaf P, et al. Phosphorylation of Hdmx mediates its Hdm2- and ATM-dependent degradation in response to DNA damage. Proc Natl Acad Sci U S A 2005; 102:5056–5061. 76. Dornan D, Shimizu H, Mah A, et al. ATM engages autodegradation of the E3 ubiquitin ligase COP1 after DNA damage. Science 2006; 313:1122–1126. 77. Falck J, Mailand N, Syljuasen RG, Bartek J, Lukas J. The ATMChk2-Cdc25A checkpoint pathway guards against radioresistant DNA synthesis. Nature 2001; 410:842–847. 78. Gatei M, Sloper K, Sorensen C, et al. Ataxia-telangiectasia-mutated (ATM) and NBS1-dependent phosphorylation of Chk1 on Ser-317 in response to ionizing radiation. J Biol Chem 2003; 278:14806–14811. 79. Sorensen CS, Syljuasen RG, Falck J, et al. Chk1 regulates the S phase checkpoint by coupling the physiological turnover and ionizing radiationinduced accelerated proteolysis of Cdc25A. Cancer Cell 2003; 3:247–258. 80. Boutros R, Dozier C, Ducommun B. The when and wheres of CDC25 phosphatases. Curr Opin Cell Biol 2006; 18:185–191. 81. Mailand N, Falck J, Lukas C, et al. Rapid destruction of human Cdc25A in response to DNA damage. Science 2000; 288:1425–1429. 82. Xiao Z, Chen Z, Gunasekera AH, et al. Chk1 mediates S and G2 arrests through Cdc25A degradation in response to DNA-damaging agents. J Biol Chem 2003; 278:21767–21773. 83. Matsuoka S, Huang M, Elledge SJ. Linkage of ATM to cell cycle regulation by the Chk2 protein kinase. Science 1998; 282:1893–1897. 84. Kitagawa R, Bakkenist CJ, McKinnon PJ, Kastan MB. Phosphorylation of SMC1 is a critical downstream event in the ATM-NBS1-BRCA1 pathway. Genes Dev 2004; 18:1423–1438. 85. Habraken Y, Piette J. NF-kappaB activation by double-strand breaks. Biochem Pharmacol 2006; 72:1132–1141. 86. Wu ZH, Shi Y, Tibbetts RS, Miyamoto S. Molecular linkage between the kinase ATM and NF-kappaB signaling in response to genotoxic stimuli. Science 2006; 311:1141–1146. 87. Schmit TL, Ahmad N. Regulation of mitosis via mitotic kinases: new opportunities for cancer management. Mol Cancer Ther 2007; 6:1920–1931. 88. Matsuoka S, Ballif BA, Smogorzewska A, et al. ATM and ATR substrate analysis reveals extensive protein networks responsive to DNA damage. Science 2007; 316:1160–1166. 89. Mu JJ, Wang Y, Luo H, et al. A proteomic analysis of ataxia telangiectasia-mutated (ATM)/ATM-Rad3-related (ATR) substrates identifies the ubiquitin-proteasome system as a regulator for DNA damage checkpoints. J Biol Chem 2007; 282:17330–17334.


115

90. de Klein A, Muijtjens M, van Os R, et al. Targeted disruption of the cell-cycle checkpoint gene ATR leads to early embryonic lethality in mice. Curr Biol 2000; 10:479–482. 91. O’Driscoll M, Gennery AR, Seidel J, Concannon P, Jeggo PA. An overview of three new disorders associated with genetic instability: LIG4 syndrome, RS-SCID and ATR-Seckel syndrome. DNA Repair (Amst) 2004; 3:1227–1235. 92. Cortez D, Guntuku S, Qin J, Elledge SJ. ATR and ATRIP: partners in checkpoint signaling. Science 2001; 294:1713–1716. 93. Paulsen RD, Cimprich KA. The ATR pathway: Fine-tuning the fork. DNA Repair (Amst) 2007; 6:953–966. 94. Liu Q, Guntuku S, Cui XS, et al. Chk1 is an essential kinase that is regulated by Atr and required for the G(2)/M DNA damage checkpoint. Genes Dev 2000; 14:1448–1459. 95. Takai H, Tominaga K, Motoyama N, et al. Aberrant cell cycle checkpoint function and early embryonic death in Chk1(-/-) mice. Genes Dev 2000; 14:1439–1447. 96. Zou L. Single- and double-stranded DNA: building a trigger of ATRmediated DNA damage response. Genes Dev 2007; 21:879–885. 97. Zou L, Elledge SJ. Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 2003; 300:1542–1548. 98. Bakkenist CJ, Kastan MB. Initiating cellular stress responses. Cell 2004; 118:9–17. 99. Ishino Y, Nishino T, Morikawa K. Mechanisms of maintaining genetic stability by homologous recombination. Chem Rev 2006; 106: 324–339. 100. Zou L, Liu D, Elledge SJ. Replication protein A-mediated recruitment and activation of Rad17 complexes. Proc Natl Acad Sci U S A 2003; 100:13827–13832. 101. Barr SM, Leung CG, Chang EE, Cimprich KA. ATR kinase activity regulates the intranuclear translocation of ATR and RPA following ionizing radiation. Curr Biol 2003; 13:1047–1051. 102. Binz SK, Sheehan AM, Wold MS. Replication protein A phosphorylation and the cellular response to DNA damage. DNA Repair (Amst) 2004; 3:1015–1024. 103. Itakura E, Umeda K, Sekoguchi E, Takata H, Ohsumi M, Matsuura A. ATR-dependent phosphorylation of ATRIP in response to genotoxic stress. Biochem Biophys Res Commun 2004; 323:1197–1202. 104. Kumagai A, Lee J, Yoo HY, Dunphy WG. TopBP1 activates the ATRATRIP complex. Cell 2006; 124:943–955. 105. Hashimoto Y, Tsujimura T, Sugino A, Takisawa H. The phosphorylated C-terminal domain of Xenopus Cut5 directly mediates ATR-dependent activation of Chk1. Genes Cells 2006; 11:993–1007.

116


106. Liu S, Bekker-Jensen S, Mailand N, Lukas C, Bartek J, Lukas J. Claspin operates downstream of TopBP1 to direct ATR signaling towards Chk1 activation. Mol Cell Biol 2006; 26:6056–6064. 107. Kumagai A, Dunphy WG. Repeated phosphopeptide motifs in Claspin mediate the regulated binding of Chk1. Nat Cell Biol 2003; 5:161–165. 108. Donzelli M, Draetta GF. Regulating mammalian checkpoints through Cdc25 inactivation. EMBO Rep 2003; 4:671–677. 109. Wang XQ, Redpath JL, Fan ST, Stanbridge EJ. ATR dependent activation of Chk2. J Cell Physiol 2006; 208:613–619. 110. Adams KE, Medhurst AL, Dart DA, Lakin ND. Recruitment of ATR to sites of ionising radiation-induced DNA damage requires ATM and components of the MRN protein complex. Oncogene 2006; 25:3894–3904. 111. Jazayeri A, Falck J, Lukas C, et al. ATM- and cell cycle-dependent regulation of ATR in response to DNA double-strand breaks. Nat Cell Biol 2006; 8:37–45. 112. Myers JS, Cortez D. Rapid activation of ATR by ionizing radiation requires ATM and Mre11. J Biol Chem 2006; 281:9346–9350. 113. Yoo HY, Kumagai A, Shevchenko A, Dunphy WG. Ataxiatelangiectasia mutated (ATM)-dependent activation of ATR occurs through phosphorylation of TopBP1 by ATM. J Biol Chem 2007; 282:17501–17506. 114. Stiff T, Walker SA, Cerosaletti K, et al. ATR-dependent phosphorylation and activation of ATM in response to UV treatment or replication fork stalling. Embo J 2006; 25:5775–5782. 115. Manning G, Whyte DB, Martinez R, Hunter T, Sudarsanam S. The protein kinase complement of the human genome. Science 2002; 298:1912–1934. 116. Noble ME, Endicott JA, Johnson LN. Protein kinase inhibitors: insights into drug design from structure. Science 2004; 303:1800–1805. 117. Stein RC. Prospects for phosphoinositide 3-kinase inhibition as a cancer treatment. Endocr Relat Cancer 2001; 8:237–248. 118. Hickson I, Zhao Y, Richardson CJ, et al. Identification and characterization of a novel and specific inhibitor of the ataxia-telangiectasia mutated kinase ATM. Cancer Res 2004; 64:9152–9159. 119. Blasina A, Price BD, Turenne GA, McGowan CH. Caffeine inhibits the checkpoint kinase ATM. Curr Biol 1999; 9:1135–1138. 120. Sarkaria JN, Busby EC, Tibbetts RS, et al. Inhibition of ATM and ATR kinase activities by the radiosensitizing agent, caffeine. Cancer Res 1999; 59:4375–4382. 121. Cariveau MJ, Tang X, Cui XL, Xu B. Characterization of an NBS1 C-Terminal Peptide That Can Inhibit Ataxia Telangiectasia Mutated (ATM)-Mediated DNA Damage Responses and Enhance Radiosensitivity. Mol Pharmacol 2007; 72:320–326.

5

Compounds that Abrogate the G2 Checkpoint Takumi Kawabe CONTENTS Why Focus on the G2 Checkpoint? Molecular Mechanism of the G2 Checkpoint Screening Protocol to Identify G2 Checkpoint Abrogators Currently Available Compounds with G2 Checkpoint-Abrogating Potential Compounds in the Clinic with G2 Checkpoint-Inhibiting Activity Future Directions

Abstract This chapter summarizes the theoretical background of cancer therapeutics that abrogate the cell cycle G2 checkpoint and reviews the molecular mechanism of the G2 checkpoint s cascade. In addition, screening procedures to identify candidate compounds, currently available compounds, and compounds in clinical trials with G2 checkpoint-abrogating potential are discussed.

From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

117

118

Kawabe

Key Words: Cell cycle; G1 checkpoint; G2 checkpoint; S checkpoint; oncogenesis; G2 abrogation; cancer therapy; DNA damage; DNA repair; mutation

1. WHY FOCUS ON THE G2 CHECKPOINT? Since the 1970s, environmental mutagens, oncogenic viruses, and inherited mutations have been suspected causative agents of oncogenesis. In recent years, these factors have been shown to affect the cell cycle G1 checkpoint mainly by impairing p53 and Rb pathways, thus leading to genomic instability (1,2). The term “genomic instability” implies the accumulation of chromosomal breaks/rearrangements, gene amplifications/deletions, and aneuploidy. Cancer cells exhibit increased mutation rates (3,4). Theoretically, an impaired G1 checkpoint increases the likelihood that damaged DNA will be replicated. DNA damage is repaired by one or more of the following mechanisms, depending on the type of damage and the phase of the cell cycle: mismatch repair, nuclear excision repair, base excision repair, methylguanine-DNA methyltransferase, homologous recombination repair, nonhomologous end joining, and translesion synthesis (TLS) (5). Among these, TLS employs specialized DNA polymerases such as Pol, Pol, Pol, and Pol and the accessory protein Rev1. TLS may facilitate mutation because these DNA polymerases exhibit extremely low fidelity during the copying of undamaged DNA substrates (6). The molecular mechanism of TLS implies that it preferentially transpires during S phase; however, there is a report of TLS in G1 phase (7). Like TLS, other repair mechanisms are prone to errors (8,9). Thus, further study is warranted to elucidate the relationship between increased mutation rate and a defective G1 checkpoint. Mutations of p53 are found in more than half of all human tumors (1), and the G1 checkpoint can be defective in the presence of wild-type p53 (10). Thus, more than half of all human tumor cells have an impaired G1 checkpoint. The normal cell cycle progresses sequentially through the phases of G0/G1, S, G2, and M, at which point the cell divides into 2 daughter cells that return to G0/G1. When genomic DNA is damaged normal cells arrest at G1 and G2 checkpoints with some delay in S phase, whereas G1 checkpoint-defective cancer cells arrest at the G2 checkpoint. Because most normal cells in the body are in G0 or G1 phase many arrest at the G1 checkpoint. Thus, G2 checkpoint abrogation in the presence of DNA damage would affect cancer cells more than

Chapter 5 / Compounds that Abrogate the G2 Checkpoint

119

normal cells because cells cannot survive in the absence of a checkpoint to repair DNA damage (2). Furthermore, because there are multiple G2 checkpoint signal cascades and some of them are dependent on p53 (11), disruption of p53-independent G2 checkpoints may affect only p53-defective cancer cells (12). In the absence of other treatments G2 checkpoint inhibitors alone may effectively target cancer cells that tend to have more DNA damage than normal cells because of increased superoxide production and defective repair systems. However, G2 checkpoint inhibitors function better in combination with DNA-damaging treatments. Interestingly, most of the conventional cytotoxic treatments and anticancer medicines including ionizing radiation, hyperthermia, pyrimidine/purine antimetabolites, alkylating agents, DNA topoisomerase inhibitors, and platinum compounds cause DNA damage. Patients tend to suffer severe adverse effects from these treatments because they are not selective for cancer cells, but only for rapidly cycling and repair-defective cells. The use of G2 checkpoint abrogators in combination with M phaseacting compounds such as paclitaxel has been proposed, in which low doses of DNA-damaging agents are used to arrest normal cells at G1 and G2 phases, presumably in a p53-dependent manner. Cancer cells are selectively killed by paclitaxel in M phase because p53deficient cells cannot arrest at G1 (because of p53 deficiency) nor at G2 (because of p53 deficiency and p53-independent G2 checkpoint abrogation) (12).

2. MOLECULAR MECHANISM OF THE G2 CHECKPOINT DNA damage is detected by a variety of sensor protein complexes (13). DNA damage sensors include Rad 17, Rfc2-5, 9-1-1 (Rad9, Rad1, and Hus1), and MRN (Mre11, Rad50, and Nbs1), but the precise nature of each sensor complex is yet to be fully determined. These sensor complexes activate ATM and/or ATR in the presence of DNA damage or replication block with the help of ATRIP (14), Mbs1 (15), MCM (16), and RPA (17). ATM and ATR then activate CHK1 and CHK2, aided by Claspin, BRCA1, TopBP1, 53BP1, and/or Mdc1 (13). Activated CHK1 (18,19) and CHK2 (20) phosphorylate serine 216 of CDC25C and inhibit its ability to trigger mitosis. CHK1 also phosphorylates and activates WEE1, an inhibitor of mitosis, and promotes binding of 14-3-3 to WEE1 (21). CHK1 stability is regulated

120

Kawabe

by Hsp90 (22). The activating phosphate group on CHK2 threonine 68 can be added by TTK/hMps1 (23) and removed by Wip1/PPMD1, a PP2C phosphatase (24). Wip1/PPMD1 also removes the activating phosphates on CHK1 and p53 and inhibits G2 checkpoint signals (25). In addition, AKT-mediated phosphorylation of CHK1 (26) and CHK2 (27) suppresses G2 checkpoint signals. Inhibitory phosphorylation of serine 216 on CDC25C can also be accomplished by MAPKAPK2 (28) and ERK1/2 (29). CDC25C inhibitory phosphorylation is maintained by the binding of 14-3-3 and reversed by PP2A (30). The involvement of MAPKAP-K2 (28) and p38 MAPK (31,32) in the UV damage-induced phosphorylation of CDC25B was also documented. WEE1 and Myt1 (33) add and CDC25C removes a phosphate group

ERK1/2

MAPKAP-K2

CHK1

CHK2

C-Tak1

p38

P CDC25C

CDC25C PP2A

P WEE1

14-3-3

P

M-phase

CDC25B

CDC25B P CDC2

CDC2

CDC2 Cyclin B Cyclin B

Cyclin B

14-3-3σ

p21

GADD45

p53

Fig. 1. Molecular mechanism of the G2 checkpoint. The G2 checkpoint signal cascade begins at the site of DNA damage. A variety of proteins recognize DNA damage and transduce signals in the form of phosphorylation relays. Shown is a schema of the downstream portion of the G2 checkpoint, which blocks G2 to M phase transition by inhibiting CDK1 (CDC2/Cyclin B) activation. Bold shapes indicate proteins that receive damage signals from upstream proteins such as ATM and ATR. Bold lines and arrows indicate inhibitory signals for G2 to M phase transition. Arrows are promoting reactions and short lines are inhibitory reactions.


121

from tyrosine 15 of CDC2 in the CDC2/Cyclin B complex (CDK1), which is the executor of G2 to M phase transition. CHK1- and CHK2mediated phosphorylation of CDC25A and its subsequent degradation by the ubiquitin-proteosome pathway has been described as another S/G2 checkpoint mechanism in G1-defective cells (34,35), whereas CDC25A functions at the G1 to S phase transition in normal cells. Another arm of the G2 checkpoint signal cascade is the p53dependent pathway. ATM activated by DNA damage phosphorylates p53 in addition to CHK1 and CHK2 (36). Activated p53 increases transcription of p21 (37), GADD45 (38), and 14-3-3 (39,40) to inhibit G2 to M phase transition via CDK1 inhibition. In addition, p53 suppresses the expression of cyclin B and CDC2 in a p21- and Rb-dependent manner, which further ensures G2 arrest (41,42). The stabilization of p21 by Hsp90-binding protein WISp39 has also been shown to play a role in p21-mediated G2 arrest (43). The downstream signaling cascades of the G2 checkpoint are depicted in Fig. 1. G2 and S phase checkpoints share many components, such as sensor proteins, CHK1, Claspin, and CDC25A/B. Thus, currently available G2 checkpoint abrogators tend to also inhibit the S checkpoint. One report showed that S checkpoint but not G2 checkpoint abrogation was the reason for clonogenic suppression by CHK1 inhibition (44). For the therapeutic use of checkpoint abrogators, S and G2 checkpoint inhibition may be equally specific in sensitizing G1 checkpointdefective cells (45). However, it may be preferable to disrupt the G2 checkpoint but not the S checkpoint because there is a greater risk of increasing the mutation rate by S checkpoint abrogation. More study is required to clarify this issue.

3. SCREENING PROTOCOL TO IDENTIFY G2 CHECKPOINT ABROGATORS There are 2 types of screening protocols to identify G2 checkpoint abrogators. One strategy is dependent on the identification of molecular target(s). This approach typically uses in vitro enzyme inhibition analyses with or without virtual screening. Inhibitors of CHK1, CHK2, and WEE1 have been sought by this method. The other type of screening protocol employs live cells, and 2 different strategies have been reported. In the method established by Roberge et al. (46), G1-defective cells are treated with a DNA-damaging agent, and then with nocodazole plus candidate compounds. The cells normally arrest at G2 phase, but in the presence

122

Kawabe

of a G2-abrogating compound they enter M phase where they are trapped by nocodazole. G2-abrogating compounds identified by this method include isogranulatimides (46), debromohymenialdisine (47), 13-hydroxy-15-ozoapathin (48), dinophysistoxin 1, 27-O-acetylokadaic acid, and 27-O-acetyldinophysistoxin 1 (49). Another method uses flow cytometry (50). Cells are treated with candidate compounds under 2 different conditions, one with a DNA-damaging agent and the other with an M phase-arresting compound such as colchicine. The M phasearresting compound is used to analyze the specificity of G2 abrogation. G2-abrogating compounds boromycin (51), simaomicin (52), 4-chlorobenzoic acid 4-methoxyphenyl ester (Patent No. WO 2003104181), and CBP501 (50) have been identified by this method. Each of the 2 screening protocols, isolated molecule-based or live cell-based screening, has merits and disadvantages. In the former protocol, the mechanism of action of the identified drug candidate will be clear. However, it requires a molecular target, and it is always uncertain whether the isolated molecule from the in vitro screening will have a pharmacological effect in live organisms. The latter method avoids the problem of in vitro/in vivo discrepancies, and it does not require knowledge of a target molecule. On the other hand, the molecular mechanism of action must be determined after a candidate molecule is identified.

4. CURRENTLY AVAILABLE COMPOUNDS WITH G2 CHECKPOINT-ABROGATING POTENTIAL Caffeine, pentoxifylline, and other methylxanthine derivatives are the oldest G2 checkpoint-abrogating compounds. Caffeine inhibits ATM and ATR (53). However, a caffeine concentration of at least 2 mM is required for G2 abrogation in vitro. Thus, the clinical use of caffeine as a G2 checkpoint abrogator is limited because it exhibits many other biological activities at much lower doses (54,55). Interestingly, although caffeine abrogates the G2 checkpoint, it does not abrogate the G1 checkpoint (56). Originally identified as PKC inhibitors, Staurosporin (57), UCN01 (58), and Go6976 (59) are indolocarbazole-type inhibitors with CHK1 inhibitory activity. SB-218078 (60) and ICP-1 (61) are relatively specific CHK1 inhibitors with indolocarbazole structures. Another novel CHK1 inhibitor, CEP-3891, has been reported (62), and an increasing number of CHK1 inhibitors are being identified (63,64, Patent No. WO 2001053268/US 2005148643, US 2003162785/WO


123

2004076424/WO 2004080973/CAN 144:468120, WO 2003093297, WO 2004081008/WO 2005016909/WO 2005066163, WO 2005009435, WO 2005047294, WO 2005072733/WO 2005118583, WO 2005103036, US 2005256157, WO 2006012308/WO 2006014359/WO 2006021002, WO 2006074281/WO 2006086255). In addition to affecting the G2 checkpoint, CHK1 inhibition has also been shown to abrogate the S phase and mitotic spindle checkpoints (65–67). Three compounds have been identified that inhibit both CHK1 and CHK2: synthetic peptide TAT-S216A (68), a marine sponge-derived debromohymenialdisine (47), and XL844 (http://www.exelixis.com/ pipeline.shtml#XL844). Although the IC50 s of TAT-S216A and debromohymenialdisine for inhibiting purified kinases in vitro are high compared to many small molecule inhibitors, the differences between IC50 and ED50 in live cells tend to be less than for other inhibitors (47,68). The difference in IC50 s of UCN-01 between recombinant CHK-2 versus immunoprecipitated CHK-2 may indicate a disparity between in vitro and in vivo efficacy (69). A CHK2-specific inhibitor, CEP-6367, has been reported without biological data (70). A novel pyridopyrimidine class WEE1 inhibitor, PD0166285, has been reported (71). The effects of WEE1 inhibitors on normal cells need to be further investigated. Contrary to the prediction that WEE1 inhibition would profoundly affect the normal cell cycle, PD0166285 seems to affect p53-defective cancer cell lines more than p53 wildtype lines (72). There is a continuing effort to find inhibitors of WEE1, some of which also inhibit CHK1 (Pat No. WO2003091255, US 2005250836). PP2A inhibitors such as okadaic acid (73) and fostriecin (74) have been shown to abrogate the G2 checkpoint. Okadaic acid is considered a tumor promoter (75) and a food poison (76). Additional PP2A inhibitors have been reported; however, they are not suitable for therapeutic use because of high toxicities (49). An inhibitor of Hsp90, 17-allylamino-17-demethoxygeldanamycin (17-AAG), was shown to abrogate the G2 checkpoint, presumably by degrading CHK1 (22). As yet, there have been no reports of compounds targeting 14-3-3. Given the redundancy of the checkpoint pathways, it may be useful to target multiple cascades at once, although it will likely be difficult to obtain a molecule capable of this task unless it is a substrate mimic such as TAT-S216A (68) or CBP501 (50). However, inhibition of CHK2 or MAPKAP2 in addition to CHK1 inhibition was no more effective in checkpoint abrogation than CHK1 inhibition alone (77); similarly, CHK1 and PLK1 inhibitors did not exhibit synergistic effects (78).

124

Kawabe

5. COMPOUNDS IN THE CLINIC WITH G2 CHECKPOINT-INHIBITING ACTIVITY A brief history of G2 checkpoint abrogators is shown in Fig. 2. In the 1960s and 1970s, caffeine and other methylxanthines were studied as sensitizers of DNA-damaging anticancer medicines in vitro (2). G2 checkpoint abrogation by caffeine was suggested in 1980 (79,80) and in 1982 (81). A clinical study of cancer patients with caffeine and cytotoxics (MeCCNU) was published in 1980 with a negative result (82). In 1989, a phase II study on the combination of caffeine, cisplatin, and cytarabine was reported (83); however, the subsequent phase III study had a negative result (84). In 1992 and 1998, potentially positive results were published for osteosarcoma patients with arterial infusions of caffeine and cisplatin, with or without doxorubicin (85,86). However, to date there has been no established clinical use of caffeine as a G2 checkpoint abrogator.

1960 1970

1980

1990

2000 2005

DNA repair defect by caffeine reported Caffeine and G2 phase relation published Three major theories of oncogenesis (heredity, mutagen, virus) G2 passing (in human cell) by caffeine reported (80) First Clinical study with Caffeine and MeCCNU reported (82) Idea of G2 checkpoint (in mammalian cells) published (81,99) Explosion of molecular biological techniques Concept of G2 checkpoint (in yeast) published (100) Clinical study with Caffeine, CDDP, ARA-C reported (83) Revealed function of major tumor suppressors in G1 checkpoint Enhancing activity of DNA damaging anti-cancer medicine in vitro by G2 checkpoint inhibition reported (Fostriecin (74), Caffeine (101), UCN-01 (58)) First Molecular cascade of G2 checkpoint revealed (18,19) New generation of G2 checkpoint inhibitors start clinical trials (94) (CBP501, XL844)

Fig. 2. History of G2 checkpoint inhibitors as anti-cancer medicine. There is an almost 40-year history of G2 checkpoint inhibitors as anti-cancer candidates, although elucidation of the molecular mechanism of the G2 checkpoint did not begin until the 1980s. The new generation of G2 checkpoint inhibitors entered clinical trials in 2005, a quarter century from the first recognizable clinical trial of caffeine, which is viewed retrospectively as a G2 checkpoint inhibitor.


125

Table 1 The New Generation of Checkpoint Inhibitors that Began Clinical Trials in 2005 Compound Screening (Company) method

Screening target

Develop mental stage

Trial target Route in clinic

CBP501 (CanBas)

Live cell- G2 Phase I based abrogation (US)

Solid tumors

XL844 (Exelixis)

Enzyme inhibition

CLL/ Oral Lymphoma

CHK1/ CHK2

Phase I (US)

Intravenous

Clinical studies of UCN-01 have been ongoing since the late 1990s. The expected mechanisms of action of UCN-01 are inhibition of PKC (87), promotion of apoptosis (AKT inhibition) (88), arrest of the cell cycle at G1/S (CDK inhibition) (89), and abrogation of the DNA damage checkpoint. The strong binding of UCN-01 to a human serum protein, alpha-1-acid glycoprotein (90), hampers treatment. Fostriecin has been tested in a phase I study (91,92), but no information is currently available on the phase II study. The Hsp90 inhibitor 17-AAG has been in clinical study since 1999, although it is not viewed as a G2 checkpoint abrogator (93). In 2005, 2 compounds originally intended to disrupt the G2 checkpoint entered phase I studies (94). CBP501 is an optimized version of TAT-S216, which was originally designed as a competitive inhibitor of CDC25C serine 216-targeting molecules (68). CBP501 inhibits MAPKAP-K2, CHK1, C-Tak1, and less efficiently, CHK2 (50). A phase I clinical trial of CBP501 was initiated in April 2005. Another new-generation G2 checkpoint-inhibiting compound, XL844, was identified by enzyme inhibition screening. XL844 is an orally available compound that inhibits CHK1 and CHK2 (http://www.exelixis.com/pipeline.shtml#XL844), and its clinical study was initiated in September 2005 (Table 1).

6. FUTURE DIRECTIONS The molecular mechanism of the G2 checkpoint has been largely revealed in the past 10 years, with continuing efforts to further characterize key components. Molecular targeting approaches have focused on CHK1, CHK2, and WEE1, and recent studies revealed additional

126

Kawabe

molecular targets such as MAPKAP-K2 (28), 14-3-3 (95), and Hsp90 (22). Phenotype-based screening for novel G2 checkpoint abrogators will continue to be important until the entire picture of the molecular mechanism of the G2 checkpoint is revealed. The potential for secondary cancer development by therapeutic G2 checkpoint abrogation should be considered because the oncogenic potential of permanent G2 checkpoint abrogation has been proposed (96–98). While ongoing clinical trial results are eagerly awaited, more clinical candidates should be identified to further augment this therapeutic approach. The theory behind G2 checkpoint abrogation for cancer treatment is well supported by both in vitro and in vivo results that suggest G2 checkpoint abrogation could affect the vast majority of cancer cells.

REFERENCES 1. Levine AJ. p53, the cellular gatekeeper for growth and division. Cell 1997;88:323–331. 2. Kawabe T. G2 checkpoint abrogators as anticancer drugs. Mol Cancer Ther 2004;3:513–519. 3. Bentle MS, Bey EA, Dong Y, Reinicke KE, Boothman DA. New tricks for old drugs: the anticarcinogenic potential of DNA repair inhibitors. J Mol Histol 2006;37: 1567–2379. 4. Sjöblom T, Jones S, Wood LD, et al. The consensus coding sequences of human breast and colorectal cancers. Science Published Online September 7, 2006/ Science Express 5. Fleck O, Nielsen O. DNA repair. J Cell Sci 2004;117:515–517. 6. Shcherbakova PV, Fijalkowska IJ. Translesion synthesis DNA polymerases and control of genome stability. Front Biosci 2006;11:2496–2517. 7. Sarkar S, Davies AA, Ulrich HD, McHugh PJ. DNA interstrand crosslink repair during G1 involves nucleotide excision repair and DNA polymerase zeta. EMBO J 2006;25:1285–1294. 8. Chan KK, Zhang QM, Dianov GL. Base excision repair fidelity in normal and cancer cells. Mutagenesis 2006;21:173–178. 9. Hsu GW, Ober M, Carell T, Beese LS. Error-prone replication of oxidatively damaged DNA by a high-fidelity DNA polymerase. Nature 2004;431:217–221. 10. Wyllie FS, Haughton MF, Blaydes JP, Schlumberger M, WynfordThomas D. Evasion of p53-mediated growth control occurs by 3 alternative mechanisms in transformed thyroid epithelial cells. Oncogene 1995;10:49–59.


127

11. Taylor WR, Stark GR. Regulation of the G2/M transition by p53. Oncogene 2001;20:1803–1815. 12. Blagosklonny MV. Sequential activation and inactivation of G2 checkpoints for selective killing of p53-deficient cells by microtubule-active drugs. Oncogene 2002;21:6249–6254. 13. O’Connell MJ, Cimprich KA. G2 damage checkpoints: what is the turnon? J Cell Sci 2005;118:1–6. 14. Yang XH, Zou L. Recruitment of ATR-ATRIP, Rad17, and 9–1-1 complexes to DNA damage. Methods Enzymol 2006;409:118–131. 15. Stiff T, Reis C, Alderton GK, Woodbine L, O’Driscoll M, Jeggo PA. Nbs1 is required for ATR-dependent phosphorylation events. EMBO J 2005;24(1):199–208. 16. Bailis JM, Forsburg SL. MCM proteins: DNA damage, mutagenesis and repair. Curr Opin Genet Dev 2004;14:17–21. 17. Yang XH, Zou L. Recruitment of ATR-ATRIP, Rad17, and 9–1-1 complexes to DNA damage. Methods Enzymol 2006;409:118–131. 18. Sanchez Y, Wong C, Thoma RS, Richman R, Wu Z, Piwnica-Worms H, Elledge SJ. Conservation of the Chk1 checkpoint pathway in mammals: linkage of DNA damage to Cdk regulation through Cdc25. Science 1997;277:1497–1501. 19. Peng CY, Graves PR, Thoma RS, Wu Z, Shaw AS, Pownica-Worms H. Mitotic and G2 checkpoint control: regulation of 14–3-3 protein binding by phosphorylation of Cdc25C on serine-216. Science 1997;277: 1501–1505. 20. Matsuoka S, Huang M, Elledge SJ. Linkage of ATM to cell cycle regulation by the Chk2 protein kinase. Science 1998;282:1893–1897. 21. Lee J, Kumagai A, Dunphy WG. Positive regulation of Wee1 by Chk1 and 14–3-3 proteins. Mol Biol Cell 2001;12:551–563. 22. Arlander SJ, Eapen AK, Vroman BT, McDonald RJ, Toft DO, Karnitz LM. Hsp90 inhibition depletes Chk1 and sensitizes tumor cells to replication stress. J Biol Chem 2003;278:52572–52577. 23. Wei JH, Chou YF, Ou YH, et al. TTK/hMps1 participates in the regulation of DNA damage checkpoint response by phosphorylating CHK2 on threonine 68. J Biol Chem 2005;280:7748–7757. 24. Oliva-Trastoy M, Berthonaud V, Chevalier A, et al. The Wip1 phosphatase (PPM1D) antagonizes activation of the Chk2 tumour suppressor kinase. Oncogene 2006; Advance online publication. 25. Lu X, Nannenga B, Donehower LA. PPM1D dephosphorylates Chk1 and p53 and abrogates cell cycle checkpoints. Genes Dev 2005;19: 1162–1174. 26. Puc J, Keniry M, Li HS, et al. Lack of PTEN sequesters CHK1 and initiates genetic instability. Cancer Cell 2005;7:193–204. 27. Hirose Y, Katayama M, Mirzoeva OK, Berger MS, Pieper RO. Akt

128

28.

29.

30.

31.

32. 33.

34.

35. 36.

37.

38. 39. 40.

41.

Kawabe activation suppresses Chk2-mediated, methylating agent-induced G2 arrest and protects from temozolomide-induced mitotic catastrophe and cellular senescence. Cancer Res 2005;65:4861–4869. Manke IA, Nguyen A, Lim D, Stewart MQ, Elia AE, Yaffe MB. MAPKAP kinase-2 is a cell cycle checkpoint kinase that regulates the G2/M transition and S phase progression in response to UV irradiation. Mol Cell 2005;17:37–48. Eymin B, Claverie P, Salon C, Brambilla C, Brambilla E, Gazzeri S. p14ARF triggers G2 arrest through ERK-mediated Cdc25C phosphorylation, ubiquitination and proteasomal degradation. Cell Cycle 2006;5:759–765. Hutchins JR, Clarke PR. Many fingers on the mitotic trigger: post-translational regulation of the Cdc25C phosphatase. Cell Cycle 2004;3:41–45. Bulavin DV, Higashimoto Y, Popoff IJ, et al. Initiation of a G2/M checkpoint after ultraviolet radiation requires p38 kinase. Nature 2001;411:102–107. Lemaire M, Froment C, Boutros R, et al. CDC25B phosphorylation by p38 and MK-2. Cell Cycle 2006;5:1649–1653. Wang Y, Decker SJ, Sebolt-Leopold J. Knockdown of Chk1, Wee1 and Myt1 by RNA interference abrogates G2 checkpoint and induces apoptosis. Cancer Biol Ther 2004;3:305–313. Falck J, Mailand N, Syljuasen RG, Bartek J, Lukas J. The ATMChk2-Cdc25A checkpoint pathway guards against radioresistant DNA synthesis. Nature 2001;410:842–847. Mailand N, Falck J, Lukas C, et al. Lukas J. Rapid destruction of human Cdc25A in response to DNA damage. Science 2000;288:1425–1429. Kurz EU, Lees-Miller SP. DNA damage-induced activation of ATM and ATM-dependent signaling pathways. DNA Repair 2004;3: 889–900. Agarwal ML, Agarwal A, Taylor WR, Stark GR. p53 controls both the G2/M and the G1 cell cycle checkpoints and mediates reversible growth arrest in human fibroblasts. Proc Natl Acad Sci U S A 1995;92:8493–8497. Wang XW, Zhan Q, Coursen JD, et al. GADD45 induction of a G2/M cell cycle checkpoint. Proc Natl Acad Sci U S A 1999;96:3706–3711. Hermeking H, Lengauer C, Polyak K, et al. 14–3-3 sigma is a p53regulated inhibitor of G2/M progression. Mol Cell 1997;1:3–11. Chan TA, Hermeking H, Lengauer C, Kinzler KW, Vogelstein B. 14-3-3Sigma is required to prevent mitotic catastrophe after DNA damage. Nature 1999;401:616–620. Manni I, Mazzaro G, Gurtner A, et al. NF-Y mediates the transcriptional inhibition of the cyclin B1, cyclin B2, and cdc25C promoters upon induced G2 arrest. J Biol Chem 2001;276:5570–5576.


129

42. Flatt PM, Tang LJ, Scatena CD, Szak ST, Pietenpol JA. p53 regulation of G(2) checkpoint is retinoblastoma protein dependent. Mol Cell Biol 2000;20:4210–4223. 43. Jascur T, Brickner H, Salles-Passador I, et al. Regulation of p21(WAF1/CIP1) stability by WISp39, a Hsp90 binding TPR protein. Mol Cell 2005;17:237–249. 44. Tse AN, Schwartz GK. Potentiation of cytotoxicity of topoisomerase i poison by concurrent and sequential treatment with the checkpoint inhibitor UCN-01 involves disparate mechanisms resulting in either p53independent clonogenic suppression or p53-dependent mitotic catastrophe. Cancer Res 2004;64:6635–6644. 45. Cho SH, Toouli CD, Fujii GH, Crain C, Parry D. Chk1 is essential for tumor cell viability following activation of the replication checkpoint. Cell Cycle 2005;4:131–139. 46. Roberge M, Berlinck RG, Xu L, et al. High-throughput assay for G2 checkpoint inhibitors and identification of the structurally novel compound isogranulatimide. Cancer Res 1998;58:5701–5706. 47. Curman D, Cinel B, Williams DE, et al. Inhibition of the G2 DNA damage checkpoint and of protein kinases Chk1 and Chk2 by the marine sponge alkaloid debromohymenialdisine. J Biol Chem 2001;276: 17914–17919. 48. Rundle NT, Xu L, Andersen RJ, Roberge M. G2 DNA damage checkpoint inhibition and antimitotic activity of 13-hydroxy-15-oxozoapatlin. J Biol Chem 2001;276:48231–48236. 49. Britton R, Roberge M, Brown C, van Soest R, Andersen RJ. New okadaic acid analogues from the marine sponge Merriamum oxeato and their effect on mitosis. J Nat Prod 2003;66:838–843. 50. Sha S, Sato T, Kobayashi H, et al. Cell cycle phenotype-based optimization of G2 abrogating peptides yields CBP501 with a unique mechanism of action at the G2 checkpoint. Mol Cancer Ther 2006 in press. 51. Arai M, Koizumi Y, Sato H, Kawabe T, et al. Boromycin abrogates bleomycin-induced G2 checkpoint. J Antibiot 2004;57:662–668. 52. Arai M, Sato H, Kobayashi H, et al. Selective inhibition of bleomycininduced G2 cell cycle checkpoint by simaomicin alpha. Biochem Biophys Res Commun 2004;317:817–822. 53. Sarkaria JN, Busby EC, Tibbetts RS, et al. Inhibition of ATM and ATR kinase activities by the radiosensitizing agent, caffeine. Cancer Res 1999;59:4375–4382. 54. Williams M, Jarvis MF. Adenosine antagonists as potential therapeutic agents. Pharmacol Biochem Behav 1988;29:433–441. 55. Mante S, Minneman KP. Caffeine inhibits forskolin-stimulated cyclic AMP accumulation in rat brain. Eur J Pharmacol 1990;175:203–205.

130

Kawabe

56. Kaufmann WK, Heffernan TP, Beaulieu LM, et al. Caffeine and human DNA metabolism: the magic and the mystery. Mutat Res 2003;532: 85–102. 57. Tam SW, Schlehel R. Staurosporine overrides checkpoints for mitotic onset in BHK cells. Cell Growth Differ 1992;3:811–817. 58. Wang Q, Fan S, Eastman A, Worland PJ, Sausville EA, O’Connor PM. UCN-01: a potent abrogator of G2 checkpoint function in cancer cells with disrupted p53. J Natl Cancer Inst 1996;88:956–965. 59. Kohn EA, Yoo CJ, Eastman A. The protein kinase C inhibitor Go6976 is a potent inhibitor of DNA damage-induced S and G2 cell cycle checkpoints. Cancer Res 2003;63:31–35. 60. Jackson JR, Gilmartin A, Imburgia C, Winkler JD, Marshall LA, Roshak A. An indolocarbazole inhibitor of human checkpoint kinase (Chk1) abrogates cell cycle arrest caused by DNA damage. Cancer Res 2000;60:566–572. 61. Eastman A, Kohn EA, Brown MK, et al. 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 2002;1:1067–1078. 62. Syljuasen RG, Sorensen CS, Nylandsted J, Lukas C, Lukas J, Bartek J. Inhibition of Chk1 by CEP-3891 accelerates mitotic nuclear fragmentation in response to ionizing Radiation. Cancer Res 2004;64: 9035–9040. 63. Foloppe N, Fisher LM, Howes R, Potter A, Robertson AG, Surgenor AE. Identification of chemically diverse Chk1 inhibitors by receptor-based virtual screening. Bioorg Med Chem 2006;14:4792–4802. 64. Tao ZF, Lin NH. Chk1 inhibitors for novel cancer treatment. Anticancer Agents Med Chem 2006;6:377–388. 65. Xiao Z, Xue J, Sowin TJ, Rosenberg SH, Zhang H. A novel mechanism of checkpoint abrogation conferred by Chk1 downregulation. Oncogene 2005;24:1403–1411. 66. Xiao Z, Xue J, Semizarov D, Sowin TJ, Rosenberg SH, Zhang H. Novel indication for cancer therapy: Chk1 inhibition sensitizes tumor cells to antimitotics. Int J Cancer 2005;115:528–538. 67. Vogel C, Kienitz A, Muller R, Bastians H. The mitotic spindle checkpoint is a critical determinant for topoisomerase-based chemotherapy. J Biol Chem 2005;280:4025–4028. 68. Suganuma M, Kawabe T, Hori H, Funabiki T, Okamoto T. Sensitization of Cancer Cells to DNA Damage-induced Cell Death by Specific Cell Cycle G2 Checkpoint Abrogation. Cancer Res 1999;59:5887–5891. 69. Yu Q, La Rose J, Zhang H, Takemura H, Kohn KW, Pommier Y. UCN01 inhibits p53 up-regulation and abrogates -radiation-induced G(2)-M checkpoint independently of p53 by targeting both of the checkpoint kinases, Chk2 and Chk1. Cancer Res 2002;62:5743–5748.


131

70. Sorensen CS, Syljuasen RG, Falck J, et al. Chk1 regulates the S phase checkpoint by coupling the physiological turnover and ionizing radiation-induced accelerated proteolysis of Cdc25A. Cancer Cell 2003;3:247–258. 71. Wang Y, Li J, Booher RN, et al. Radiosensitization of p53 mutant cells by PD0166285, a novel G(2) checkpoint abrogator. Cancer Res 2001;61:8211–8217. 72. Li J, Wang Y, Sun Y, Lawrence TS. Wild-type TP53 inhibits G(2)-phase checkpoint abrogation and radiosensitization induced by PD0166285, a WEE1 kinase inhibitor. Radiat Res 2002;157:322–330. 73. Ghosh S, Schroeter D, Paweletz N. Okadaic acid overrides the S-phase check point and accelerates progression of G2-phase to induce premature mitosis in HeLa cells. Exp Cell Res 1996;227:165–169. 74. Roberge M, Tudan C, Hung SM, Harder KW, Jirik FR, Anderson H. Antitumor drug fostriecin inhibits the mitotic entry checkpoint and protein phosphatases 1 and 2A. Cancer Res 1994;54:6115–6121. 75. Suganuma M, Fujiki H, Suguri H, et al. Okadaic acid: an additional nonphorbol-12-tetradecanoate-13-acetate-type tumor promoter. Proc Natl Acad Sci U S A. 1988 85:1768–1771. 76. Dickey RW, Fryxell GA, Granade HR, Roelke D. Detection of the marine toxins okadaic acid and domoic acid in shellfish and phytoplankton in the Gulf of Mexico. Toxicon 1992;30:355–359. 77. Xiao Z, Xue J, Sowin TJ, Zhang H. Differential roles of checkpoint kinase 1, checkpoint kinase 2, and mitogen-activated protein kinaseactivated protein kinase 2 in mediating DNA damage-induced cell cycle arrest: implications for cancer therapy. Mol Cancer Ther 2006;5: 1935–1943. 78. Gao Q, Huang X, Tang D, et al. Influence of chk1 and plk1 silencing on radiation- or cisplatin-induced cytotoxicity in human malignant cells. Apoptosis 2006;11:1789–1800. 79. Iseki S, Ebina T, Ishida N. Caffeine-induced recovery from G2 block caused by neocarzinostatin. Gann 1980;71:567–571. 80. Bases R, Mendez F, Liebeskind D, Elequin F, Neubort S. DNA of HeLa cells during caffeine-promoted recovery from X-ray induced G2 arrest. Int J Radiat Biol Relat Stud Phys Chem Med 1980;37:437–445. 81. Lucke-Huhle C. Alpha-irradiation-induced G2 delay: a period of cell recovery. Radiat Res 1982;89:298–308. 82. Cohen MH, Schoenfeld D, Wolter J. Randomized trial of chlorpromazine, caffeine, and methyl-CCNU in disseminated melanoma. Cancer Treat Rep 1980;64:151–153. 83. Dougherty JB, Kelsen D, Kemeny N, Magill G, Botet J, Niedzwiecki D. Advanced pancreatic cancer: a phase I-II trial of cisplatin, highdose cytarabine, and caffeine. J Natl Cancer Inst 1989;81: 1735–1738.

132

Kawabe

84. Kelsen D, Hudis C, Niedzwiecki D, et al. A phase III comparison trial of streptozotocin, mitomycin, and 5-fluorouracil with cisplatin, cytosine arabinoside, and caffeine in patients with advanced pancreatic carcinoma. Cancer 1991;68:965–969. 85. Tsuchiya H, Yasutake H, Yokogawa A, Baba H, Ueda Y, Tomita K. Effect of chemotherapy combined with caffeine for osteosarcoma. J Cancer Res Clin Oncol 1992;118:567–569. 86. Tsuchiya H, Tomita K, Mori Y, et al. Caffeine-assisted chemotherapy and minimized tumor excision for nonmetastatic osteosarcoma. Anticancer Res 1998;18:657–666. 87. Hofmann J. Protein kinase C isozymes as potential targets for anticancer therapy. Curr Cancer Drug Targets 2004;4:125–146. 88. Tsuruo T, Naito M, Tomida A, et al. Molecular targeting therapy of cancer: drug resistance, apoptosis and survival signal. Cancer Sci 2003;94:15–21. 89. Nahta R, Hortobagyi GN, Esteva FJ. Signal transduction inhibitors in the treatment of breast cancer. Curr Med Chem Anticancer Agents 2003;3:201–216. 90. Fuse E, Tanii H, Kurata N, et al. Unpredicted clinical pharmacology of UCN-01 caused by specific binding to human alpha1-acid glycoprotein. Cancer Res 1998, 58:3248–3253. 91. Le LH, Erlichman C, Pillon L, et al. Phase I and pharmacokinetic study of fostriecin given as an intravenous bolus daily for 5 consecutive days. Invest New Drugs 2004;22:159–167. 92. de Jong RS, Mulder NH, Uges DR, et al. Phase I and pharmacokinetic study of the topoisomerase II catalytic inhibitor fostriecin. Br J Cancer 1999;79:882–887. 93. Goetz MP, Toft D, Reid J, et al. Phase I trial of 17-allylamino-17demethoxygeldanamycin in patients with advanced cancer. J Clin Oncol 2005;23:1078–1087. 94. Garber K. New checkpoint blockers begin human trials. J Natl Cancer Inst 2005;97:1026–1028. 95. Qi W, Martinez JD. Reduction of 14–3-3 proteins correlates with increased sensitivity to killing of human lung cancer cells by ionizing radiation. Radiat Res 2003;160:217–223. 96. Paules RS, Levedakou EN, Wilson SJ, et al. Defective G2 checkpoint function in cells from individuals with familial cancer syndromes. Cancer Res 1995;55:1763–1773.


133

97. Kawabe T, Muslin AJ, Korsmeyer SJ. HOX11 interacts with protein phosphatases PP2A and PP1 and disrupts a G2/M cell-cycle checkpoint. Nature 1997;385:454–458. 98. Nakada S, Katsuki Y, Imoto I, et al. Early G2/M checkpoint failure as a molecular mechanism underlying etoposide-induced chromosomal aberrations. J Clin Invest 2006;116:80–89. 99. Lau CC, Pardee AB. Mechanism by which caffeine potentiates lethality of nitrogen mustard. Proc Natl Acad Sci U S A 1982;79:2942–2946. 100. Weinert T, Hartwell L. Control of G2 delay by the rad9 gene of Saccharomyces cerevisiae. J Cell Sci Suppl 1989;12:145–148. 101. Russell KJ, Wiens LW, Demers GW, Galloway DA, Plon SE, Groudine M. Abrogation of the G2 checkpoint results in differential radiosensitization of G1 checkpoint-deficient and G1 checkpointcompetent cells. Cancer Res 1995;55:1639–1642.

6

CDK Inhibitors as Anticancer Agents Timothy A. Yap, L. Rhoda Molife, and Johann S. de Bono CONTENTS Introduction Cyclin-Dependent Kinases Therapeutic CDK Inhibitors Future Strategies

Abstract Eukaryotic organisms depend on the cell cycle for their survival though a cyclical biochemical process consisting of tightly controlled, enzymatically driven reactions that result in cell division and the generation of new cells. The cell cycle is regulated by kinases such as cyclin-dependent kinases (CDKs), and non-CDKs, which include the Aurora and Polo-like kinases, as well as checkpoint proteins and mitotic kinesins. The CDK family of serine-theronine kinases is a common target for genetic or epigenetic events, resulting in the amplification or overexpression of these kinases in a myriad of tumor types. Such findings make CDKs rational and attractive targets for cancer therapeutics as their inhibition could potentially result in preferential targeting of malignant cells. However, several cyclin-CDK complexes have been found to be dispensable for cell proliferation owing to functional redundancy, promiscuity, and From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

135

136

Yap, Molife, and de Bono

compensatory mechanisms. Although these issues have hampered their progress into the clinic, several novel compounds are currently in various phases of clinical trial development. This chapter will introduce the role of CDKs in facilitating the cell cycle, their aberrations in malignant progression and pharmacological strategies targeting them. Key Words: Cell cycle; cyclin-dependent kinase; cyclin, inhibitor; flavopiridol; UCN-01; E7070; seliciclib; BMS 387032; ZD 304709

1. INTRODUCTION The cell cycle ensures the survival of multicellular organisms by enabling the generation of new cells through a cyclical biochemical process consisting of highly regulated, enzymatically driven reactions that result in cell division. There are 4 distinct phases in each cell cycle: Gap 1 (G1 ), S (DNA synthesis), Gap 2 (G2 ) and M (mitosis) (Fig. 1). Cell cycle progression is controlled by a family of nonmembrane bound serine-threonine kinases, which interact with other protein

Fig. 1. The 4 phases of the cell cycle. Gap 1 (G1) phase, cell grows; S phase, replication of DNA; Gap 2 (G1) phase, cell prepares to divide; M phase, cell division. Cell cycle progression is driven by the association of cyclins with their respective CDKs. DNA damage leads to the activation of checkpoint kinase proteins (Chk1 and Chk2), which cause cell cycle arrest by phosphorylating CDC25.

Chapter 6 / CDK Inhibitors as Anticancer Agents

137

families, to permit systematic and coordinated cell cycle activities. Each individual phase of the cell cycle is driven by these kinases, which are in turn tightly controlled by undergoing inactivation or destruction on completion of their respective tasks. The largest and most important family of kinases is the cyclin-dependent kinases (CDKs). Other groups of non-CDK kinases are also required for normal cell cycle progression. Under normal circumstances, CDKs permit cell cycle progression. However, in the event of an error occurring, the cell cycle is halted by these kinases until the aberration is corrected. The development of faults or mutations in this finely tuned control system may thus have great potential to lead to uncontrolled cell cycle progression and cellular transcription, as well as aberrant apoptosis, all hallmarks of malignancy. Genetic or epigenetic events have been found to lead to amplification or overexpression of CDKs and cyclins, or to the loss of function of their corresponding inhibitors. These may thus result in the dysregulation of cell division and the development of cancer. There is considerable evidence implicating CDK anomalies in carcinogenesis, and thus, strategies directed at CDK-related targets may form an attractive and rational area for novel anticancer drug development.

2. CYCLIN-DEPENDENT KINASES The CDKs were first described 30 years ago when Nurse et al (1) identified a gene in fission yeast responsible for entry into mitosis, whilst Matsui (2) and Smith (3) identified a maturation-promoting factor (MPF) complex in amphibian oocytes. This heterodimeric complex was later purified (4) and found to consist of CDK1 and cyclin B. The amount of cyclin B, which drives the activity of the MPF complex, rises through the cell cycle until mitosis, when it drops abruptly because of degradation. Cell cycle kinases phosphorylate proteins on both serine and threonine amino acid residues and are thus known as serine/threonine kinases. By interacting with other protein families, they systematically regulate cell cycle progression. The CDKs form the largest group in this family of kinases. There are at least nine structurally related CDKs, including the cell cycle CDKs 1, 2, 4, and 6, and the noncell cycle CDKs 3, 5, 7, 8 and 9 (Fig. 1). The latter CDKs are considered “housekeeping” kinases because of their involvement in noncell cycle related activities. For example, CDKs 7, 8, and 9 phosphorylate RNA polymerase II and regulate RNA transcription, whilst CDK5 is

138


involved in synaptic vesicle recycling, insulin secretion, and neuronal survival (5). CDKs are closely associated with proteins called cyclins, so named because their concentration fluctuates in a cyclical manner during the cell cycle (Table 1) (6,7). This specific cyclin expression pattern defines the relative position of the cell within the cell cycle. There are a substantial number of cyclins identified thus far (cyclins A to cyclin T) (Fig. 1). When levels of cyclins rise, they form stable active heterodimeric complexes with CDKs, activating the CDK protein kinase function. These cyclin-CDK complexes are activated by phosphorylation at specific sites on the CDK by cyclin H-CDK7, also known as CDK-activating kinase (CAK) (8). When their concentration falls, cyclins detach from and inhibit CDKs, resulting in the loss of CDK phosphorylation and their catalytic abilities. These cyclin (regulatory) - CDK (catalytic) complexes may be classified in 2 separate groups based on their respective roles, either in cell cycle progression or in the regulation of transcription (Table 1) (5,9). Progression through the G1 phase is driven by the association of cyclin D isoforms (cyclins D1, D2, and D3) with CDKs 2, 4, and 6, whilst cyclin E interacts with CDK2 at the G1 /S transition and drives its entry into S phase (Fig. 1). They facilitate cell cycle progression by mediating the sequential phosphorylation of the retinoblastoma (Rb) protein (10,11). This inactivates Rb and facilitates the G1 to S transition

Table 1 Cyclins and Their Respective CDK Partners and Role Cyclin

CDK

Role

A B

2 1

C D (D1 , D2 , D3 isoforms) E F (orphan) H

8 1, 4, 6 2 − 7

T

9

S phase progression Transcription regulation G2 to M phase transition Transcription regulation G1 phase progression G1 /S transition Cell cycle progression Transcription regulation CDK-activating kinase(CAK) Transcription regulation


139

through its interaction with E2F transcription factors (Fig. 1) (12). S phase progression is facilitated by cyclin A-CDK2 complex, with cyclin A-CDK1 (also referred to as CDC2) necessary in the G2 phase (13). Cyclin B-CDK1 is important during the mitosis (M) phase for cell division to take place (14). There are also “orphan” cyclins, such as cyclin F, which do not possess a CDK partner, but are still essential for cell cycle progression (6,7). Those complexes involved in regulating transcription include cyclin B-CDK1, cyclin H-CDK7, cyclin C-CDK8 and cyclin T-CDK9 (pTEFb) (5,15). These complexes all phosphorylate the carboxyterminal domain (CTD) of RNA polymerase II (15,16). Cyclin T-CDK9 drives the initiation and elongation of nascent RNA transcripts. Cyclin B-CDK1 and cyclin C-CDK8 suppress mRNA production during mitosis and inhibit protein factors required to initiate transcription. Tumor cells seem to be especially sensitive to RNA polymerase II inhibition, suggesting that targeting the CTD of RNA polymerase II may be a rational choice in the development of novel anti-CDK inhibitors (14). Although these cyclin-CDK complexes are regulatory proteins, they are themselves regulated by corresponding CDK inhibitors (CDKIs), which serve as negative regulators of the cell cycle by stopping cells from progressing through the cell cycle (Table 2). These cell cycle inhibitory proteins include the CDK inhibitor protein/kinase inhibitor protein (Cip/Kip) class of CDKIs, inhibitor of CDK4 (INK4) class of CDKIs, as well as various kinases and phosphatases (17,18). CAK also binds activating phosphate groups to the cyclin-CDK complexes, whilst another kinase Wee1 adds inhibitory phosphates, both resulting in the inactivation of the complexes. These phosphate groups are removed Table 2 CDKs and Their Respective Inhibitors Cell Cycle Kinases

Inhibitory Proteins

Class of Inhibitor

Cyclin D-associated kinases (CDK2, 4 and 6)

Cyclin E-CDK2 and cyclin A-CDK2 complexes

Inhibitor of cdk4 (INK4) class of CDKIs

Cyclin E-CDK2 and cyclin A-CDK2 complexes

p21waf1 p27kip1 p57kip2

Kinase inhibitor protein (KIP) class of CDK inhibitors

140


by the phosphatase CDC25, leading to the subsequent activation of the complexes (19). Genetic or epigenetic events may lead to the overexpression of CDKs and cyclins, or the inactivation of CDKIs. This may in turn lead to the loss of cell cycle checkpoint integrity, and to uncontrolled cell division and potentially carcinogenesis. This observation, as well as the knowledge that cell cycle arrest by CDK inhibition could induce apoptosis, has led to the rationale that CDK inhibitors present attractive and rational targets for the development of novel anticancer therapeutics (14).

3. THERAPEUTIC CDK INHIBITORS There is considerable evidence implicating CDK aberrations in carcinogenesis. CDK4 has been demonstrated to be amplified in gliomas (20,21), breast cancer (22), lung cancer (23), and different sarcoma types (24,25). Since this initial finding that CDK4 gene amplification and overexpression in sporadic breast carcinomas is associated with increased tumor cell proliferation (22), it has been reported that mice lacking cyclin D1 are resistant to breast cancer triggered by the Erb-2 (HER-2) oncogene (26). Also, studies have shown that the deregulation of cyclin D1-CDK4/6 interactions is an essential target of erbB2 function in both murine and human breast tumors, and that the overexpression of erbB2 may be sufficient to deregulate cyclin D1-CDK4/6 activity in breast cancer (27). Two recent studies further demonstrated that this CDK complex was a rational target for novel drug development (28,29). In the first study, “knock-in” mice expressing mutant cyclin D1 deficient in activating CDK4/6 were resistant to breast cancers initiated by ErbB-2, but still had normal cyclin D1-dependent murine development (28). Also, primary human breast cancers were analyzed and found to have high cyclin D1 levels in about a quarter of ErbB-2-overexpressing tumors (29). Sustained CDK4-associated kinase activity was also found to be essential to maintain breast tumorigenesis. It is reported that the ability of cyclin D1 to activate CDK4 underlies the key role for cyclin D1 in breast cancer formation (29). Thus, cyclin D1-CDK4/6 kinase inhibition may be of therapeutic benefit to patients with this subtype of breast cancer. CDK2 has been shown to be over-expressed in colorectal (CRC) (30) and lung cancers (31). Mouse knockout studies have however suggested that CDK2 is not essential for mitotic cell division and may thus not be an optimal therapeutic target (32,33).


141

Epigenetic silencing by hypermethylation of the promoter region of the inhibitory INK4 gene p16 results in transcription inhibition and loss of its gene expression, potentially resulting in uncontrolled cellular proliferation (14). Accordingly, the functional loss of p16 has been demonstrated in a range of cancers, including CRC, melanoma, lung and breast tumors (14), providing evidence implicating CDKI mutations in carcinogenesis. Targeting these cell cycle aberrations may potentially restore cell cycle checkpoints and control the cycling ability of tumors, and even facilitate apoptotic induction. Significant research has been carried out in the development of novel pharmacological compounds targeting CDKs in cancer therapeutics, especially small-molecule inhibitors (34). These agents target CDKs either directly, by inhibiting the catalytic CDK subunit, or indirectly by targeting regulatory pathways that control CDK activity (14).

3.1. First Generation CDK Inhibitors The first generation of CDK inhibitors developed include flavopiridol, staurosporine and its analogue UCN-01, butyrolactone, the paullones and indirubin. These agents are nonselective and inhibit both CDKs and other kinases. 3.1.1. Flavopiridol Preclinical Activity. Flavopiridol (Alvocidib; Aventis-NCI), a derivative of the plant alkaloid rohitukine was the first pharmacological CDK inhibitor to enter clinical trials. It is a pan-CDK inhibitor with multiple mechanisms of antitumoral activity (35). Flavopiridol targets CDKs 1, 2, 4, 6, and 7 through its interaction at the adenosine triphosphate (ATP) binding site (35). It also has activity against the cyclin T-CDK9 complex, which inhibits transcription and reduces cyclin D1 mRNA expression (36). Other kinases affected include PKA and PKC. In vitro studies have shown that Flavopiridol induces G1 and G2 cell cycle arrest, is cytotoxic to cells undergoing DNA synthesis and induces apoptosis (37). Clinical Experience. Based on preclinical data, clinical trials of flavopiridol primarily evaluated 2 schedules—a 72-h infusion administered every 14 d and a daily 1-h bolus schedule administered over 1, 3, or 5 d every 21 d (38–41). In phase-I trials of the infusional schedule, the dose limiting toxicity (DLT) was secretory diarrhoea (38,39). At higher doses, a proinflammatory syndrome comprising fever, fatigue,

142


and reversible hypotension, as well as tumor pain was seen (38). With the bolus schedule, the DLT was neutropenia. Other toxicities consisted of gastrointestinal symptoms, and again a proinflammatory syndrome (Table 3) (40). Another phase-I trial of a 24-h infusion repeated weekly demonstrated colonic ulceration and abdominal pain as the DLTs (Table 3) (42). With the 72-hour infusion schedule administered every 14 d, a complete response was observed in 1 patient with gastric adenocarcinoma, who remained disease-free for more than 2 yr after the completion of treatment (39). In addition, there was 1 partial response (PR) in a patient with renal cell carcinoma (RCC) and minor responses in 3 patients with RCC, CRC, and non-Hodgkins lymphoma (NHL) (38). However, despite these initial promising results, phase-II studies of these tumor types, as well as non small cell lung cancer (NSCLC), mantle cell lymphoma, hormone refractory prostate cancer and melanoma all demonstrated negative results (Table 3) (43–50). A high incidence of arterial and venous thrombo-embolic events were observed in these studies. Re-evaluation of data from the phase-I trials revealed that these thrombotic events had actually occurred previously, but were not previously fully appreciated (39). Although there was no clear dose- or concentration-dependent relationship established between flavopiridol and the occurrence of thrombosis, these studies suggested that the toxicities seen when using a 72-h schedule were unacceptable. Mixed results were seen with phase-II trials of the daily bolus schedule in mantle cell lymphoma, melanoma, chronic lymphocytic leukemia, endometrial cancer, RCC and multiple myeloma (Table 3) (50–55). The response rate in the study of mantle cell lymphoma was modest at 11%, with stable disease (SD) in 70% of patients for a median of 3.4 mo (51). These results contrasted with the findings of the infusional study in the same tumor type, where no responses were seen (46). In the chronic lymphocytic leukemia study by Byrd and colleagues (50), the bolus schedule was compared with the infusional schedule in a sequential trial, and demonstrated an 11% response rate with the former schedule, and no activity with the latter schedule. Modest activity was seen in RCC, with a response rate similar to that obtained in the infusional phase-II study done by Stadler and colleagues (43,54). All other studies showed negative results. Preclinical studies demonstrated that flavopiridol can enhance cytotoxic chemotherapy-induced apoptosis, and induce sequence dependent cytotoxic synergy (56,57). Flavopiridol was shown to signif-

143

I I

I

Seliciclib BMS 387032

ZK304709

−

− −

SCCHN, Mel, NSCLC, CRC

Gastrointestinal, metabolic, skin Gastrointestinal, metabolic, skin, myelosuppresion Hypertension, dizziness, fatigue, gastrointestinal

(102), (103)

(79), (80), (81), (82), (84), (85), (87), (86) (92), (94), (93), (95) (98), (99), (100)

(38), (39), (40), (42), (43), (44), (45), (46), (47), (48), (49), (50), (51), (52), (53), (54), (55) (60), (61), (62), (63), (64) (70), (71), (72), (77) (74), (76)

References

RCC, renal cell cancer; NSCLC, nonsmall cell lung cancer; MCL, mantle cell lymphoma; CRC, colorectal carcinoma; HRPC, hormone refractory prostate cancer; CLL, chronic lymphocytic lymphoma; Mel, metastatic melanoma; EC, endometrial carcinoma; MM, multiple myeloma; SCCHN, squamous cell carcinoma head and neck.

a

I, II

RCC −

I, II I

E7070

Myelosuppression, gastrointestinal, fatigue Gastrointestinal, hyperglycaemia Arrhythmia, gastrointestinal, hyperglycaemia,sepsis, renal impairment, Myelosuppression, skin, constitutional, gastrointestinal

−

I

Flavopiridol and chemotherapy UCN-01 UCN-01 and chemotherapy

Toxicities Diarrhoea, pro-inflammatory syndrome, gastrointestinal

Phase I, II

Flavopiridol (single agent)

Tumor Types(phase II)a RCC, gastric, NSCLC, MCL,CRC HRPC, CLL, Mel, EC, MM

Trial status

Compound

Table 3 Clinical Trials of CDK Inhibitors

144


icantly enhance paclitaxel-induced apoptosis in the human gastric and breast cancer cell lines MKN-74 and MCF-7 if administered following paclitaxel (57). This synergistic effect of flavopiridol on paclitaxel-treated cells is caused by an increase in caspase 3 activation (57). When flavopiridol was administered after docetaxel to tumorbearing xenografts, there was an increase in chemotherapy-induced apoptosis and antitumor activity, as shown by tumor growth delay and regression (58). These synergistic effects were not observed if the sequence of treatments were reversed. It is hypothesized that pretreatment with flavopiridol inhibits cyclin B1-CDC2 kinase activity preventing cells from entering mitosis, and thus antagonizing the effect of docetaxel (58). On this basis, several combination studies have been reported (Table 3) (57,59). When paclitaxel was followed by flavopiridol (24-h infusion), the DLTs were neutropenia and pulmonary toxicity. Clinical activity was seen in patients with esophageal, lung and prostate cancer, including patients who had previously progressed on paclitaxel (60). In other phase-I studies, flavopiridol has been combined with docetaxel, irinotecan, cis– and carboplatin, as well as mitozantrone, indicating that these combinations are feasible. It is also clear from these and other studies that the sequence of administration of these agents is important (61–64). 3.1.2. UCN-01 Preclinical Activity. The second CDK inhibitor to enter clinical trials was UCN-01 (7-hydroxystaurosporine), a chemical derived from staurosporine, an alkaloid from Streptomyces bacteria (35). UCN-01, like flavopiridol, has several mechanisms of action and is not just a pan-CDK inhibitor, but also a phospholipid-dependent protein kinase C (PKC) inhibitor. It potently abrogates the G2 checkpoint control in cancer cells with disrupted p53 function by targeting the CDC25 kinases, Chk1, and Chk2. At concentrations above 100 nM, this leads to cell-cycle arrest and apoptosis (65–67). UCN-01 has also demonstrated activity against the 3-phosphoinositide-dependent protein kinase-1 (PDK1)-Akt survival signaling pathway by the inhibition of upstream Akt/PKB kinase PDK1 (68). In addition, UCN-01 may also potentate the cytotoxicity of S-phase-active chemotherapeutics (69). Clinical Experience. The first phase-I study of UCN-01 involved a 72-h infusion administered every 14 days (Table 1) (70). This was subsequently modified to a 72-h infusion given every 28 d because of the observation of an unexpectedly long half-life and high plasma


145

concentration because of avid binding of drug to plasma -1-acid glycoprotein. The final recommended phase-II schedule and dose was an initial 72-h infusion of 95 mg/m2 , followed by monthly 36-h infusions at 42.5 mg/m2 /d. DLTs were hyperglycaemia caused by peripheral insulin resistance, pulmonary toxicity, nausea, vomiting, and hypotension. Activity was seen in a patient with large cell lymphoma (prolonged SD) and a second patient with melanoma (PR lasting 6 months). In other studies, a monthly 3-h infusion was tested (71,72). Toxicities were mild and included nausea, hyperglycemia, and hypotension (72). With this schedule, the recommended phase-II dose was of an initial 95 mg/m2 infusion, followed by 47.5 mg/m2 for subsequent courses (72). No objective responses were seen. Like flavopiridol, UCN-01 was shown to potentiate the activity of cytotoxics in a sequence-dependent manner. UCN-01 has been shown to suppress thymidine synthase (TS) protein expression in a human gastric cancer cell line SK-GT5 in a dose-dependent manner, because of the suppression of E2F-1 protein expression, a crucial transcription factor of TS (73). In addition, this same model showed a significant increase in apoptosis when cells were treated with a combination of 5-fluorouracil (5-FU) and UCN-01 as opposed to either agent alone; however this was seen only when UCN-01 was administered after 5-FU. These preclinical findings prompted a phase-I combination study of 5-FU administered pre-UCN-01. UCN-01 was administered every 28 d at a fixed dose of 135 mg/m2 over 72 h at cycle 1, then 67.5 mg/m2 over 36 h in subsequent cycles. 5-FU was administered as a weekly 24-h infusion and dose-escalated in successive cohorts. The DLTs were syncope, arrhythmia, and hyperglycemia (74). Other toxicities were headache and emesis. No objective responses were seen but 1 patient demonstrated prolonged SD for more than 6 mo (74). Other combination studies have been performed (Table 1) (73–76). When cisplatin was combined with UCN-01, the DLTs were grade-5 sepsis with respiratory failure and grade-3 creatinine in 1 patient with subsequent death, and grade-3 atrial fibrillation in a second patient (76). The study was thus terminated. Despite this, tumor biopsies obtained from 3 patients pre- and posttreatment showed decreased levels of Chk1 and CDC25. One patient with adenocarcinoma of unknown primary had a PR associated with an increased expression of p27KIP1 . A phase-II study of the 3-h infusion administered every 21 d in 21 patients with RCC failed to meet its efficacy goal of time to disease progression (TTP) of 4.5 mo (Table 3) (77).

146


In a third study, 33 patients were treated with UCN-01 and topotecan once every 3 wk (69). The recommended dose was 70 mg/m2 of UCN01 with 1.0 mg/m2 of topotecan. The observed DLTs were grade-3 emesis, and grade-4 neutropenia, whereas other toxicities included transient hyperglycaemia, fatigue, and hypotension. Twelve patients had SD for at least 6 cycles, whereas 1 patient with ovarian cancer demonstrated a PR – this has led to a phase-II study of this drug combination in ovarian cancer (69).

3.2. Second Generation CDK Inhibitors The next generation of compounds developed, when compared with 1st generation CDK inhibitors, displayed more selective activity against their targeted CDKs, and included the agents E7070, seliciclib, and BMS-387032. 3.2.1. E7070 Preclinical Activity. E7070 is a chloroindolyl sulphonamide that has been demonstrated to induce G1/S cell cycle arrest at low nanomolar concentrations (78). Evidence from in vitro cell lines showed that this compound not only inhibits cyclin E and CDK2, but also down-regulates cyclin H. At higher concentrations, E7070 may result in apoptosis through the upregulation of p53 and p21 (78). This agent was also found to have antitumor activity in 42 different cell lines (78). In addition, E7070 demonstrated schedule-dependent antitumor activity in vivo (78). Clinical Experience. Five phase-I studies investigating different schedules of this chloroindoyl sulfonamide have been published (Table 3) (79–83). The recommended doses of the various schedules were as follows: 1. Daily for 5 consecutive days every 3 wk; 130 mg/m2 (79). 2. 1-h infusion every 21 d; 700 mg/m2 and 800 mg/m2 in minimally and heavily pretreated groups respectively (80). 3. 5-d continuous infusion every 21 d; 96 mg/m2 /day (81). 4. weekly infusions for 4 wk every 6 wk; 400 mg/m2 /week (82).

The DLTs were neutropenia and thrombocytopenia; other toxicities were primarily gastrointestinal (stomatitis, nausea, vomiting), dermatological (folliculitis), and constitutional (fatigue). With the first schedule, 1 patient (out of 33) with heavily pretreated breast cancer demonstrated a PR (79). In the study of a 1-h infusion administered every 21 d, 14 out of 27 assessable patients—with a variety of tumor types including metastatic RCC and CRC—had SD for up to 23 mo,


147

including patients with previously documented disease progression before study entry, who went on to demonstrate SD for more than 6 mo (80). With the schedule involving a weekly infusion for 4 wk every 6 wk, 1 PR (out of 55 patients) was seen in a patient with endometrial adenocarcinoma (82). Another patient with metastatic melanoma received 21 cycles of treatment. On completion of treatment, this patient’s disease remained stable for a further year. Twelve patients achieved SD for a median of 5 mo (82). These findings suggest that more frequent dosing of E7070 may result in greater clinical benefit. The 3-week schedule of E7070 has been tested in the phaseII setting, but revealed negative results in metastatic squamous cell carcinoma of the head and neck (SCCHN) and melanoma (84,85). In the study of SCCHN, 3 patients showed evidence of reduced phosphorylation of RB after treatment, which was thought to be related to the modulation of CDK activity by E7070 (84). Minimal activity was seen in a phase-II trial in patients with CRC, where 9% of patients achieved a PR (86). Similarly, in a phase-II study in NSCLC, minimal activity with a 2% response rate was seen (87). 3.2.2. Seliciclib Preclinical Activity. Seliciclib (R-roscovitine; CYC202; Cyclacel Ltd, Dundee, United Kingdom) is a purine-based CDK inhibitor, particularly CDKs 1, 2, 7, and 9 (88,89). A recent study demonstrated that seliciclib regulates invasive breast cancer cell (MDA-MB231) proliferation and survival through CDK5 (90). Seliciclib has also been shown to inhibit RB phosphorylation in tumor cells and murine tumor xenografts (91). Clinical Experience. Seliciclib was tested as a single agent in 2 dose schedules—as a twice daily dose for 5 d every 21 d, and as a twice daily dose for 7 d every 21 d (92–94). The recommended dose for the former study was 1,250 mg (Table 3) (92), whereas the maximum dose reached for the latter study was 1,600 mg (93,94). The DLTs in both studies were similar and included vomiting, hypokalaemia and skin rash. Other toxicities included nausea and renal impairment. No responses were reported in either study, although in the 5-d study, 3 patients showed SD for 6 mo. Seliciclib, in combination with gemcitabine and cisplatin, was tested in chemotherapy-naïve patients with metastatic NSCLC (Table 3). The recommended phase-II dose was 800 mg twice daily of seliciclib on days 1–4, 8–11, and 15–18, with intravenous gemcitabine at a dose

148


of 1,000 mg/m2 administered on days 5 and 12, and cisplatin at a dose of 75 mg/m2 given on day 5 (95). DLTs comprised of nausea, vomiting, elevated glutamyl transferase and transient hypokalaemia. Six out of 14 patients demonstrated a PR, and 7 had SD. However, this level of response may have been achievable with chemotherapy alone, and thus a randomized controlled trial is necessary to distinguish any added benefits from seliciclib.

3.2.3. BMS-387032 Preclinical Activity. BMS-387032 (N-[5-[[[5-(1,1-dimethylethyl)2-oxazolyl]methyl]thio]-2-thiazolyl]-4-pipe ridinecarboxamide) is a potent, selective, and competitive small molecule inhibitor of the cyclin E-CDK2 complex, with an IC50 of 48 nM (96). It also inhibits CDKs 1 and 4, but has less potency against all other tested non-CDK kinases (96). In vitro, this compound inhibits CDK2 phosphorylation in the A2780 ovarian carcinoma cell line, inhibiting the phosphorylation of downstream targets of CDK2, such as RB, histone H1, and DNA polymerase-. It was shown to display in vitro cytotoxicity against this cell line, with an IC50 of 50 nM. In vivo studies involving BMS-387032 demonstrated oral bioavailability and activity against a wide range of cell lines, including A2780 ovarian, A431 human squamous cell carcinoma, and P388 murine leukemia (96). There was also antitumor activity against a murine mammary carcinoma generated in transgenic mice overexpressing cyclin E. Combination studies suggest that BMS-387032 is synergistic with cisplatin in SV-1 colon carcinoma cells, with the synergy dependent on drug sequence, dose and cyclin D1 expression (97). Optimal cytotoxic activity was demonstrated when the compound was administered 24 hours before the cisplatin, whereas antagonism was observed when this administration sequence was reversed. Clinical Experience. Preliminary results of 3 BMS-387032 phaseI trials have been reported (Table 1) (98–100). With a schedule involving a 1-h infusion administered every 3 wk, the most common toxicities observed were fatigue, nausea, vomiting, anorexia, diarrhoea, constipation and transient liver transaminitis (98). SD lasting more than 6 mo was reported in patients with RCC, NSCLC, sarcoma and oropharyngeal carcinoma. Similar toxicities were seen with a 1-h weekly infusion, and a schedule involving a 24-h infusion administered every 3 wk (99,100). Further development of this agent has


149

ceased in view of the disappointing results in these initial studies. It has been suggested that the compound may be too selective for the cyclin E-CDK2 complex. 3.2.4. ZK304709 Preclinical Activity. ZK304709 (Schering AG) is a selective inhibitor of CDKs 1, 2, 4, 7, and 9 (Table 3) (101). It also possesses antiangiogenic effects through its inhibition of the tyrosine kinase activity of vascular endothelial growth factor-receptor (VEGF-R) 1, 2, 3, and platelet derived growth factor-beta receptor (PDGF-R) (101). It has been shown to have greater efficacy than standard chemotherapeutic compounds, both in human tumor xenografts and orthotopic human pancreatic carcinoma models (101). Clinical Experience. Two phase-I studies exploring different oral schedules of ZK304709 have been reported (102,103). In the first study, a 14-d 4-weekly schedule was tested, whereas the second study evaluated a 7-d 3-weekly schedule. Doses ranging from 15 mg to 180 mg were tested. The DLTs observed included nausea, vomiting, dizziness, fatigue, and hypertension. Limited activity was demonstrated, with SD observed for more than 4 mo in patients with a variety of tumor types. 3.2.5. PD 0332991 PD 0332991 (Pfizer Global Research, New York), a pyrido[2,3day]pyrimidine-7-ones, is a new generation CDK inhibitor, with relative selectivity for CDK4/6 (104,105). This selectivity for CDK4 was demonstrated in RB-positive cell lines in vitro. These studies also reported exclusive G1 arrest and dephosphorylation of RB by PD 0332991 at known CDK4-specific phosphorylation sites. The IC50 values of this compound against RB-positive cell lines were between 40 and 400 nmol/L, in contrast to the IC50 values of more than 3 μmol/L against 2 RB-negative cell lines. The compound was cytostatic in breast and colon carcinoma cells, and also reported to cause significant tumor regression in mice bearing Colo-205 colon carcinoma xenografts (106). In MDA-MB-453 human breast cancer cells, PD 0332991 inhibited 50% of Rb phosphorylation at the CDK4-specific phosphorylation site Ser780 in vivo, and a decreased expression of E2F-1-dependent genes and Ki67 staining. It showed no activity against Rb-negative tumor xenografts (106). A phase-I clinical trial with PD 0332991 in patients with RB-positive advanced solid tumors has recently commenced (14).

150


3.3. Key Issues with CDK Inhibitors CDKs are essential for cell cycle progression and checkpoint function—both of which lead to oncogenic transformation if disrupted. Studies have shown that certain cell cycle kinases are amplified or overexpressed in a range of tumor types, suggesting that their inhibition could result in preferential targeting of malignant cells. Essential proteins such as pRB, which are downregulated in tumor cells, may also be restored to their normal function. In addition, ATP-binding sites essential for cell cycle kinase activation offer good drug target sites. All these factors undoubtedly make the development of cell cycle kinase inhibitors an attractive area for cancer therapeutics. However, to date, CDK inhibitors such as flavopiridol have demonstrated limited clinical activity as single agents in advanced solid tumors, the reasons for which remain unclear. It is hypothesized that the antitumor activity of flavopiridol has been limited by the finding that it is highly protein bound. This was supported by the observation that administrating the agent to patients with refractory chronic lymphocytic leukemia for 30 min, followed by a 4-hour continuous infusion, resulted in tumor lysis and increased biological activity (107). This is thought to be because of the inhibition of CDK complexes that phosphorylate CTD of RNA polymerase II, resulting in the suppression of genes that inhibit apoptosis. It however remains to be seen if this approach applies to solid tumors too. Apart from the initiation of apoptosis, it also appears that chronic dosing of CDK inhibitors is key for sustained antitumor effects. Oral dosing rather that IV administration of these agents may thus be essential in their development as single agents in advanced solid tumors (14). There also remain unanswered questions with regards to the selectivity and promiscuity of these CDK inhibitors. Affinity chromatography with panels of CDKs have revealed the unselective nature of CDK inhibitors developed thus far. Although one potentially potent and selective CDK4/6 inhibitor has been identified (106), most other “specific” CDK inhibitors have now been shown to inhibit other kinases, and even possibly other unknown proteins, demonstrating that selectivity remains an issue (35). The nonselectivity of CDK inhibitors may be demonstrated with seliciclib and BMS-387032. Although thought to be relatively selective for CDK2, these agents have also been found to inhibit CDK1. However, there is increasing evidence that CDK2 is not crucial for cell proliferation as shown by:


151

• Evidence from in vivo studies, demonstrating that proliferation continues even after specific deletion of CDK2 by siRNA, or antisense nucleotides (33). • Induction of G1 arrest in CDK2−/− cells by Cip/Kip proteins (108). • Viability of CDK2 knockout mice (8,32).

Thus, agents that selectively target CDK2 may not be of therapeutic benefit in all tumor types. It thus remains unclear if selective CDK inhibitors will show superior efficacy in the clinic over pan-CDK inhibitors such as flavopiridol. Functional redundancy of a number of CDKs may limit the effects of highly selective CDK inhibitors. In vitro studies have shown that cell proliferation proceeds despite the absence of individual G1 CDKs, probably because of the presence of other compensatory kinases, or because these kinases are nonfunctioning. This implies that highly selective inhibitors of individual G1 CDKs may potentially be ineffective as single agent cancer therapeutics. Regardless, certain tumors have highly amplified CDKs, with their survival dependent on its expression. Thus, for specific CDK inhibitors to succeed as single agents, patient selection will be essential, and may be aided by novel microarray technology. For example, microarray studies have demonstrated that CDK4 is highly amplified in certain liposarcomas (109). By identifying such “at risk” group of susceptible patients, one might increase the chances of an antitumor response with these specific CDK inhibitors. Nonselective CDK inhibitors might not always be disadvantageous. It is now thought that much of the clinical activity observed with the first and second generation CDK inhibitors may in part be because of their effects on noncell cycling “housekeeping” CDKs 3, 5, 7, 8, and 9, rather than the ones they were originally intended to target. CDKs 7, 8, and 9 are involved in RNA transcription through the phosphorylation of the CTD of RNA polymerase II, whereas CDK 5 is associated with insulin secretion, thus explaining the hyperglycaemia observed with flavopiridol therapy (5). Although the inhibition of multiple CDKs may enhance antitumor activity, this must be balanced against the risk of compromising selective cytotoxicity in tumor cells. The promiscuous activity of nonspecific CDK inhibitors may involve normal cycling cells, and result in toxicity to normal dividing cells, including significant gastrointestinal and bone marrow toxicity. This has implications for the clinical development of these inhibitors, not just with regards to safety and tolerance, but also the MTD that may be reached with these

152


inhibitors. These drug-related toxicities may also limit the potential for combination with standard cytotoxic chemotherapies, where the future of these agents may lie.

4. FUTURE STRATEGIES Our improved understanding of the molecular biology that belies the cell cycle, together with novel data from techniques such as combinatorial chemistry, high throughput screening and x-ray crystallography have allowed the continued identification of novel cell cycle kinase targets, as well as therapeutic small molecule and antibody inhibitors. Computational approaches will continue to improve our understanding of molecular recognition and reveal novel target sites for drug design. Alternative targeting strategies are already being evaluated, such as the inhibition of protein–protein interactions, kinase-substrate protein interactions, or the phosphorylation sites on the protein kinases. A wide range of other kinases may offer potential alternatives in “drugging” the cell cycle, including Bub1, BubR1, and Mps1, which are involved in the spindle assembly checkpoint, as well as the human Nek family of kinases. Other agents indirectly target the cell cycle, such as 17-(Allylamino)-17-demethoxygeldanamycin (17-AAG), which suppresses Chk1 expression (110), and histone deacetylase (HDAC) inhibitors, such as suberoylanilide hydroxamic acid (SAHA), which upregulate the transcription of p21 and induce cell cycle arrest (111). Potent and selective small molecule antagonists of MDM2 have also recently been developed. MDM2 has a high affinity for the p53 tumor suppressor protein and affects its transcriptional activity and stability. Thus, the inhibition of the interaction between MDM2 and p53 may result in p53 stabilization; this has been demonstrated to lead to cell cycle arrest, apoptosis, and growth inhibition of human tumor xenografts in nude mice (112). This novel class of compounds may thus represent another potential cell cycle inhibitor. In view of the growing list of potential antitumor agents available, clear prioritization of the clinical development of these novel compounds based on detailed target evaluation is required. CDK inhibitors allow cell cycle aberrations of malignancy to be manipulated directly, and this may be further exploited by combining such inhibitors with standard chemotherapeutic agents. This strategy appears to be where the future application of these and other similar targeted therapies lies. However, in view of cell cycle mediated drug


153

resistance, regimens combining these CDK inhibitors with standard cytotoxics need to be rationally planned before their advancement into the clinical arena. Drug sequencing and scheduling must be considered when combining a CDK inhibitor with an agent that works on cycling cells (e.g., cytotoxic chemotherapy), as erroneous combinations may result in negating effects. Apart from sequencing and scheduling, appropriate dosing and time intervals between different agents is also likely to be crucial to clinical outcome (58). Cytotoxic agents are not the only potential therapeutic partners for these agents; combinations involving other molecular targeted agents are also of interest, as evidenced by the preclinical synergy observed with bortezimib (113,114). Combining CDK inhibitors with other targeted agents may allow simultaneous inhibition of multiple pathways “horizontally” or “vertically,” and inhibit both the target, as well as any “cross-talking” between pathways (115). The increased risk of drug-induced toxicity will again require attention with combination studies. In order to address this issue, sequential rather than concomitant administration of such agents may have to be evaluated in clinical trials. General issues relating to the development of any novel therapy also apply here. This includes determining the optimal dose schedule to be administered, the best tumor types to be targeted, and demonstration of target modulation from tumor specimens by using appropriate and reliable PD biomarkers. These PD endpoints should be consistent with the phenotype known to be associated with target inhibition. Although PD endpoints have been defined in some studies (88,91), the optimum targets and their clinical relevance are yet to be defined. Patient selection will also remain critically important, and may be aided by the application of novel expression or proteomic arrays to the study of individual tumor characteristics. Modern functional imaging modalities, such as FLT-PET may also be useful in monitoring responses to inhibitors (116). In the future, staging systems for different tumor types will need to be modified to integrate molecular biomarkers, which could potentially be utilized not just as tools for appropriate patient selection for treatment, but also as diagnostic and prognostic predictors. All these novel developments could eventually lead to our ultimate goal of personalized medicine.

REFERENCES 1. Nurse P. Genetic control of cell size at cell division in yeast. Nature 1975; 256:547–551.

154


2. Masui Y, Markert CL. Cytoplasmic control of nuclear behavior during meiotic maturation of frog oocytes. J Exp Zool 1971; 177:129–145. 3. Smith LD Ecker RE. The interaction of steroids with Rana pipiens Oocytes in the induction of maturation. Dev Biol 1971;25:232–247. 4. Lohka MJ, Masui Y. (1983) Formation in vitro of sperm pronuclei and mitotic chromosomes induced by amphibian ooplasmic components. Science 1983;220:719–721. 5. Sausville EA. Complexities in the development of cyclin-dependent kinase inhibitor drugs. Trends Mol Med 2002;8:S32–37. 6. Bai C, Richman R, Elledge SJ. Human cyclin F. Embo J 1994; 13: 6087–6098. 7. Kong M, Barnes EA, Ollendorff V, Donoghue DJ. Cyclin F regulates the nuclear localization of cyclin B1 through a cyclin-cyclin interaction. Embo J 2000;19:1378–1388. 8. Berthet C, Aleem E, Coppola V, Tessarollo L, Kaldis P. Cdk2 knockout mice are viable. Curr Biol 2003;13:1775–1785. 9. Meyerson M, Enders GH, Wu CL et al. A family of human cdc2-related protein kinases. Embo J, 1992;11:2909–2917. 10. Lundberg AS, Weinberg RA. Functional inactivation of the retinoblastoma protein requires sequential modification by at least two distinct cyclin-cdk complexes. Mol Cell Biol 1998;18:753–761. 11. Harbour JW, Luo RX, Dei Santi A, Postigo AA, Dean DC. (1999) Cdk phosphorylation triggers sequential intramolecular interactions that progressively block Rb functions as cells move through G1. Cell 1999;98:859–869. 12. Sherr CJ. G1 phase progression: cycling on cue. Cell, 1994;79, 551–555. 13. Pines J. Cyclins: wheels within wheels. Cell Growth Differ, 1991;2, 305–310. 14. Schwartz GK, Shah MA. Targeting the cell cycle: a new approach to cancer therapy. J Clin Oncol, 2005;23,9408–9421. 15. Meinhart A, Kamenski T, Hoeppner S, Baumli S, Cramer P. A structural perspective of CTD function. Genes Dev, 2005;19,1401–1415. 16. Prelich G. RNA polymerase II carboxy-terminal domain kinases: emerging clues to their function. Eukaryot Cell, 2002;1:153–162. 17. Harper JW, Elledge SJ. The role of Cdk7 in CAK function, a retroretrospective. Genes Dev, 1998;12:285–289. 18. Sherr CJ, Roberts JM. Inhibitors of mammalian G1 cyclin-dependent kinases. Genes Dev, 1995;9:1149–1163. 19. Coleman TR, Dunphy WG. Cdc2 regulatory factors. Curr Opin Cell Biol, 1994;6:877–882. 20. Collins VP. Gene amplification in human gliomas. Glia, 1995;5: 289–296. 21. Reifenberger G, Reifenberger J, Ichimura K, Meltzer PS, Collins VP.


22.

23.

24.

25.

26. 27.

28.

29. 30.

31.

32.

33. 34.

155

Amplification of multiple genes from chromosomal region 12q13–14 in human malignant gliomas: preliminary mapping of the amplicons shows preferential involvement of CDK4, SAS, and MDM2. Cancer Res, 1994;54:4299–4303. An HX, Beckmann MW, Reifenberger G, Bender HG, Niederacher D. Gene amplification and overexpression of CDK4 in sporadic breast carcinomas is associated with high tumor cell proliferation. Am J Pathol, 1999;154:113–118. Wikman H, Nymark P, Vayrynen A et al. CDK4 is a probable target gene in a novel amplicon at 12q13.3-q14.1 in lung cancer. Genes Chromosomes Cancer, 2005;42:193–199. Forus A, Larramendy ML, Meza-Zepeda LA et al. Dedifferentiation of a well-differentiated liposarcoma to a highly malignant metastatic osteosarcoma: amplification of 12q14 at all stages and gain of 1q22q24 associated with metastases. Cancer Genet Cytogenet, 2001; 125: 100–111. Khatib ZA, Matsushime H, Valentine M, Shapiro DN, Sherr CJ, Look AT. Coamplification of the CDK4 gene with MDM2 and GLI in human sarcomas. Cancer Res, 1993;53:5535–5541. Yu Q, Geng Y, Sicinski P. Specific protection against breast cancers by cyclin D1 ablation. Nature, 2001;411:1017–1021. Yang C, Ionescu-Tiba V, Burns K et al. The role of the cyclin D1dependent kinases in ErbB2-mediated breast cancer. Am J Pathol, 2004;164:1031–1038. Landis MW, Pawlyk BS, Li T, Sicinski P, Hinds PW. Cyclin D1dependent kinase activity in murine development and mammary tumorigenesis. Cancer Cell, 2006;9:13–22. Yu Q, Sicinska E, Geng Y et al. Requirement for CDK4 kinase function in breast cancer. Cancer Cell, 2006;9:23–32. Yamamoto H, Monden T, Miyoshi H et al. Cdk2/cdc2 expression in colon carcinogenesis and effects of cdk2/cdc2 inhibitor in colon cancer cells. Int J Oncol, 1998;13:233–239. Dobashi Y, Shoji M, Jiang SX, Kobayashi M, Kawakubo Y, Kameya T. Active cyclin A-CDK2 complex, a possible critical factor for cell proliferation in human primary lung carcinomas. Am J Pathol, 1998; 153: 963–972. Ortega S, Prieto I, Odajima J et al. Cyclin-dependent kinase 2 is essential for meiosis but not for mitotic cell division in mice. Nat Genet, 2003;35: 25–31. Tetsu O, McCormick F. Proliferation of cancer cells despite CDK2 inhibition. Cancer Cell, 2003,3:233–245. Senderowicz AM. Small-molecule cyclin-dependent kinase modulators. Oncogene, 2003;22:6609–6620.

156


35. Senderowicz AM, Sausville EA. Preclinical and clinical development of cyclin-dependent kinase modulators. J Natl Cancer Inst, 2000; 92: 376–387. 36. Carlson B, Lahusen T, Singh S et al. Down-regulation of cyclin D1 by transcriptional repression in MCF-7 human breast carcinoma cells induced by flavopiridol. Cancer Res, 1999;59:4634–4641. 37. Patel V, Senderowicz AM, Pinto D, Jr. et al. Flavopiridol, a novel cyclin-dependent kinase inhibitor, suppresses the growth of head and neck squamous cell carcinomas by inducing apoptosis. J Clin Invest, 1998;102:1674–1681. 38. Senderowicz AM, Headlee D, Stinson SF et al. Phase I trial of continuous infusion flavopiridol, a novel cyclin-dependent kinase inhibitor, in patients with refractory neoplasms. J Clin Oncol, 1998;16:2986–2999. 39. Thomas JP, Tutsch KD, Cleary JF et al. Phase I clinical and pharmacokinetic trial of the cyclin-dependent kinase inhibitor flavopiridol. Cancer Chemother Pharmacol, 2002;50:465–472. 40. Tan AR, Headlee D, Messmann R et al. Phase I clinical and pharmacokinetic study of flavopiridol administered as a daily 1-hour infusion in patients with advanced neoplasms. J Clin Oncol, 2002; 20: 4074–4082. 41. Arguello F, Alexander M, Sterry JA et al. Flavopiridol induces apoptosis of normal lymphoid cells, causes immunosuppression, and has potent antitumor activity In vivo against human leukemia and lymphoma xenografts. Blood, 1998;91:2482–2490. 42. Sasaki Y, Sasaki T, Minami H et al. A phase I pharmacokinetic (PK)pharmacodynamic (PD) study of flavopiridol by 24 hours continuous infusion (CI) repeating every week. Proc Am Soc Clin Oncol 21:2002 (abstr 371). 43. Stadler WM, Vogelzang NJ, Amato R et al. Flavopiridol, a novel cyclindependent kinase inhibitor, in metastatic renal cancer: a University of Chicago Phase II Consortium study. J Clin Oncol, 2000; 18: 371–375. 44. Schwartz GK, Ilson D, Saltz L et al. Phase II study of the cyclindependent kinase inhibitor flavopiridol administered to patients with advanced gastric carcinoma. J Clin Oncol, 2001;19:1985–1992. 45. Shapiro GI, Supko JG, Patterson A et al. A phase II trial of the cyclin-dependent kinase inhibitor flavopiridol in patients with previously untreated stage IV non-small cell lung cancer. Clin Cancer Res, 2001;7:1590–1599. 46. Lin TS, Howard OM, Neuberg DS, Kim HH, Shipp MA. Seventy-two hour continuous infusion flavopiridol in relapsed and refractory mantle cell lymphoma. Leuk Lymphoma, 2002;43:793–797.


157

47. Aklilu M, Kindler HL, Donehower RC, Mani S, Vokes EE. Phase II study of flavopiridol in patients with advanced colorectal cancer. Ann Oncol, 2003;14:1270–1273. 48. Liu G, Gandara DR, Lara PN, Jr. et al. A Phase II trial of flavopiridol (NSC #649890) in patients with previously untreated metastatic androgen-independent prostate cancer. Clin Cancer Res, 2004;10: 924–928. 49. Flinn IW, Byrd JC, Bartlett N et al. Flavopiridol administered as a 24hour continuous infusion in chronic lymphocytic leukemia lacks clinical activity. Leuk Res, 2005;29:1253–1257. 50. Byrd JC, Peterson BL, Gabrilove J et al. Treatment of relapsed chronic lymphocytic leukemia by 72-hour continuous infusion or 1-hour bolus infusion of flavopiridol: results from Cancer and Leukemia Group B study 19805. Clin Cancer Res, 2005;11:4176–4181. 51. Kouroukis CT, Belch A, Crump M et al. 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, 2003;21:1740–1745. 52. Burdette-Radoux S, Tozer RG, Lohmann RC et al. Phase II trial of flavopiridol, a cyclin dependent kinase inhibitor, in untreated metastatic malignant melanoma. Invest New Drugs, 2004;22:315–322. 53. Grendys EC, Jr., Blessing JA, Burger R, Hoffman J. A phase II evaluation of flavopiridol as second-line chemotherapy of endometrial carcinoma: a Gynecologic Oncology Group study. Gynecol Oncol, 2005;98,249–253. 54. Van Veldhuizen PJ, Faulkner JR, Lara PN, Jr. et al. A phase II study of flavopiridol in patients with advanced renal cell carcinoma: results of Southwest Oncology Group Trial 0109. Cancer Chemother Pharmacol, 2005;56:39–45. 55. Dispenzieri A, Gertz MA, Lacy MQ et al. Flavopiridol in patients with relapsed or refractory multiple myeloma: a phase 2 trial with clinical and pharmacodynamic end-points. Haematologica, 2006;91: 390–393. 56. Matranga CB, Shapiro GI. Selective sensitization of transformed cells to flavopiridol-induced apoptosis following recruitment to S-phase. Cancer Res, 2002;62:1707–1717. 57. Motwani M, Delohery TM, Schwartz GK. Sequential dependent enhancement of caspase activation and apoptosis by flavopiridol on paclitaxel-treated human gastric and breast cancer cells. Clin Cancer Res, 1999;5:1876–1883. 58. Motwani M, Rizzo C, Sirotnak F, She Y, Schwartz GK. Flavopiridol enhances the effect of docetaxel in vitro and in vivo in human gastric cancer cells. Mol Cancer Ther, 2003;2:549–555.

158


59. Jung CP, Motwani MV, Schwartz GK. Flavopiridol increases sensitization to gemcitabine in human gastrointestinal cancer cell lines and correlates with down-regulation of ribonucleotide reductase M2 subunit. Clin Cancer Res, 2001;7:2527–2536. 60. Schwartz GK, O’Reilly E, Ilson D et al. Phase I study of the cyclindependent kinase inhibitor flavopiridol in combination with paclitaxel in patients with advanced solid tumors. J Clin Oncol, 2002;20: 2157–2170. 61. Tan AR, Yang X, Berman A et al. Phase I trial of the cyclin-dependent kinase inhibitor flavopiridol in combination with docetaxel in patients with metastatic breast cancer. Clin Cancer Res, 2004;10:5038–5047. 62. Shah MA, Kortmansky J, Motwani M et al. A phase I clinical trial of the sequential combination of irinotecan followed by flavopiridol. Clin Cancer Res, 2005;11:3836–3845. 63. Bible KC, Lensing JL, Nelson SA et al. Phase 1 trial of flavopiridol combined with cisplatin or carboplatin in patients with advanced malignancies with the assessment of pharmacokinetic and pharmacodynamic end points. Clin Cancer Res, 2005;11:5935–5941. 64. Karp JE, Passaniti A, Gojo I et al. Phase I and pharmacokinetic study of flavopiridol followed by 1-beta-D-arabinofuranosylcytosine and mitoxantrone in relapsed and refractory adult acute leukemias. Clin Cancer Res, 2005;11:8403–8412. 65. Wang Q, Fan S, Eastman A, Worland PJ, Sausville EA, O’Connor PM. UCN-01: a potent abrogator of G2 checkpoint function in cancer cells with disrupted p53. J Natl Cancer Inst, 1996;88: 956–965. 66. Busby EC, Leistritz DF, Abraham RT, Karnitz LM, Sarkaria JN. The radiosensitizing agent 7-hydroxystaurosporine (UCN-01) inhibits the DNA damage checkpoint kinase hChk1. Cancer Res, 2000;60: 2108–2112. 67. Yu Q, La Rose J, Zhang H, Takemura H, Kohn KW, Pommier Y. UCN01 inhibits p53 up-regulation and abrogates gamma-radiation-induced G(2)-M checkpoint independently of p53 by targeting both of the checkpoint kinases, Chk2 and Chk1. Cancer Res, 2002;62:5743–5748. 68. Sato S, Fujita N, Tsuruo T. Interference with PDK1-Akt survival signaling pathway by UCN-01 (7-hydroxystaurosporine). Oncogene, 2002;21:1727–1738. 69. Hotte SJ, Oza A, Winquist EW et al. 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, 2006;17:334–340. 70. Sausville EA, Arbuck SG, Messmann R et al. Phase I trial of 72-hour continuous infusion UCN-01 in patients with refractory neoplasms. J Clin Oncol, 2001;19:2319–2333.


159

71. Tamura T, Sasaki Y, Minami H et al. Phase I study of UCN-01 by 3-hour infusion. 1999, ASCO Annual Meeting (Abstract 611). 72. Dees EC, Baker SD, O’Reilly S et al. A phase I and pharmacokinetic study of short infusions of UCN-01 in patients with refractory solid tumors. Clin Cancer Res, 2005;11:664–671. 73. Hsueh CT, Kelsen D, Schwartz GK. UCN-01 suppresses thymidylate synthase gene expression and enhances 5-fluorouracil-induced apoptosis in a sequence-dependent manner. Clin Cancer Res, 1998;4:2201–2206. 74. Kortmansky J, Shah MA, Kaubisch A et al. Phase I trial of the cyclindependent kinase inhibitor and protein kinase C inhibitor 7-hydroxystaurosporine in combination with Fluorouracil in patients with advanced solid tumors. J Clin Oncol, 2005;23:1875–1884. 75. Pollack IF, Kawecki S, Lazo JS. Blocking of glioma proliferation in vitro and in vivo and potentiating the effects of BCNU and cisplatin: UCN-01, a selective protein kinase C inhibitor. J Neurosurg, 1996;84:1024–1032. 76. Lara PN, Jr., Mack PC, Synold T et al. The cyclin-dependent kinase inhibitor UCN-01 plus cisplatin in advanced solid tumors: a California cancer consortium phase I pharmacokinetic and molecular correlative trial. Clin Cancer Res, 2005;11:4444–4450. 77. Rini BI, Weinberg V, Shaw V et al. Time to disease progression to evaluate a novel protein kinase C inhibitor, UCN-01, in renal cell carcinoma. Cancer, 2004;101:90–95. 78. Ozawa Y, Sugi NH, Nagasu T et al. E7070, a novel sulphonamide agent with potent antitumour activity in vitro and in vivo. Eur J Cancer, 2001;37:2275–2282. 79. Punt CJ, Fumoleau P, van de Walle B, Faber MN, Ravic M, Campone M. Phase I and pharmacokinetic study of E7070, a novel sulfonamide, given at a daily times five schedule in patients with solid tumors. A study by the EORTC-early clinical studies group (ECSG). Ann Oncol, 2001;12:1289–1293. 80. Raymond E, ten Bokkel Huinink WW, Taieb J et al. Phase I and pharmacokinetic study of E7070, a novel chloroindolyl sulfonamide cell-cycle inhibitor, administered as a one-hour infusion every three weeks in patients with advanced cancer. J Clin Oncol, 2002;20: 3508–3521. 81. Terret C, Zanetta S, Roche H et al. Phase I clinical and pharmacokinetic study of E7070, a novel sulfonamide given as a 5-day continuous infusion repeated every 3 weeks in patients with solid tumours. A study by the EORTC Early Clinical Study Group (ECSG). Eur J Cancer, 2003;39:1097–1104. 82. Dittrich C, Dumez H, Calvert H et al. Phase I and pharmacokinetic study of E7070, a chloroindolyl-sulfonamide anticancer agent, administered on a weekly schedule to patients with solid tumors. Clin Cancer Res, 2003;9:5195–5204.

160


83. Droz JP. Phase I trial of five-days continuous infusion E7070 [N(3-chloro-7-indolyl)-1,4-benzenedisulfonamide] in patients with solid tumors. Proc Am Associ Cancer Res, 2000;41:609. 84. Haddad RI, Weinstein LJ, Wieczorek TJ et al. A phase II clinical and pharmacodynamic study of E7070 in patients with metastatic, recurrent, or refractory squamous cell carcinoma of the head and neck: modulation of retinoblastoma protein phosphorylation by a novel chloroindolyl sulfonamide cell cycle inhibitor. Clin Cancer Res, 2004;10:4680–4687. 85. Smyth JF, Aamdal S, Awada A et al. Phase II study of E7070 in patients with metastatic melanoma. Ann Oncol, 2005;16:158–161. 86. Mainwaring PN, Van Cutsem E, Van Laethem J et al. 2002, ASCO Annual Meeting (Abstract 611). 87. Talbot D, Norbury C, Slade M et al. 2002, ASCO Annual Meeting (Abstract 1306). 88. Whittaker SR, Walton MI, Garrett MD and Workman P. The Cyclin-dependent kinase inhibitor CYC202 (R-roscovitine) inhibits retinoblastoma protein phosphorylation, causes loss of Cyclin D1, and activates the mitogen-activated protein kinase pathway. Cancer Res, 2004;64:262–272. 89. McClue SJ, Blake D, Clarke R et al. In vitro and in vivo antitumor properties of the cyclin dependent kinase inhibitor CYC202 (Rroscovitine). Int J Cancer, 2002;102:463–468. 90. Goodyear S, Sharma MC. Roscovitine regulates invasive breast cancer cell (MDA-MB231) proliferation and survival through cell cycle regulatory protein cdk5. Exp Mol Pathol 2007 Feb;82(1):25–32. 91. Raynaud FI, Whittaker SR, Fischer PM et al. In vitro and in vivo pharmacokinetic-pharmacodynamic relationships for the trisubstituted aminopurine cyclin-dependent kinase inhibitors olomoucine, bohemine and CYC202. Clin Cancer Res, 2005;11:4875–4887. 92. Pierga J, Faivre S, Vera K et al. A phase I and pharmacokinetic (PK) trial of CYC202, a novel oral cyclin-dependent kinase (CDK) inhibitor, in patients (pts) with advanced solid tumors. Proc Am Soc Clin Oncol 22:2003 (abstr 840). 93. Benson C, White J, Twelves C et al. A phase I trial of the oral cyclin dependent kinase inhibitor CYC202 in patients with advanced malignancy. Proc Am Soc Clin Oncol 2003;22:2003 (abstr 838). 94. White JD, Cassidy J, Twelves C et al. A phase I trial of the oral cyclin dependent kinase inhibitor CYC202 in patients with advanced malignancy. J Clin Oncol, 2004 ASCO Annual Meeting Proceedings (PostMeeting Edition). Vol 22, No 14S (July 15 Supplement), 2004: 3042. 95. Siegel-Lakhai WS, Rodenstein DO, Beijnen JH, Gianella-Borradori A, Schellens JH, Talbot DC. Phase I study of seliciclib (CYC202 or R-roscovitine) in combination with gemcitabine (gem)/cisplatin (cis) in


96.

97. 98.

99.

100.

101.

102.

103.

104.

105. 106.

161

patients with advanced Non-Small Cell Lung Cancer (NSCLC). J Clin Oncol, 2005 ASCO Annual Meeting Proceedings. Vol 23, No. 16S, Part I of II (June 1 Supplement), 2005:2060. Misra RN, Xiao HY, Kim KS et al. N-(cycloalkylamino)acyl-2aminothiazole inhibitors of cyclin-dependent kinase 2. N-[5-[[[5-(1,1 -dimethylethyl)-2-oxazolyl]methyl]thio]-2-thiazolyl]-4-piperidinecarboxamide (BMS-387032), a highly efficacious and selective antitumor agent. J Med Chem, 2004;47:1719–1728. Blagden S, de Bono J. Drugging cell cycle kinases in cancer therapy. Curr Drug Targets, 2005;6:325–335. Jones SF, Burris HA, Kies M et al. A phase I study to determine the safety and pharmacokinetics (PK) of BMS-387032 given intravenously every three weeks in patients with metastatic refractory solid tumors. Proc Am Soc Clin Oncol 22:2003 (abstr 798). McCormick J, Gadgeel SM, Helmke W et al. Phase I study of BMS387032, a cyclin dependent kinase (CDK) 2 inhibitor Proc Am Soc Clin Oncol 22:2003 (abstr 835) Shapiro G, Lewis N, Bai S et al. A phase I study to determine the safety and pharmacokinetics (PK) of BMS-387032 with a 24-hr infusion given every three weeks in patients with metastatic refractory solid tumors Proc Am Soc Clin Oncol 22:2003 (abstr 799). Siemeister G, Luecking U, Wagner C, Detjen K, Mc Coy C and Bosslet K. Molecular and pharmacodynamic characteristics of the novel multi-target tumor growth inhibitor ZK 304709. Biomed Pharmacother, 2006;60:269–272. Graham J, Wagner K, Plummer R et al. Phase I dose-escalation study of novel oral multi-target tumor growth inhibitor (MTGI) ZK 304709 administered daily for 7 days of a 21-day cycle to patients with advanced solid tumors. J Clin Oncol, 2006 ASCO Annual Meeting Proceedings Part I. Vol 24, No. 18S (June 20 Supplement), 2006: 2073. Ahmed S, Molife R, Shaw H et al. Phase I dose-escalation study of ZK 304709, an oral multi-target tumor growth inhibitor (MTGI), administered for 14 days of a 28-day cycle. J Clin Oncol, 2006 ASCO Annual Meeting Proceedings Part I. Vol 24, No. 18S (June 20 Supplement), 2006:076. Toogood PL, Harvey PJ, Repine JT et al. Discovery of a potent and selective inhibitor of cyclin-dependent kinase 4/6. J Med Chem, 2005;48:2388–2406. Shapiro GI. Cyclin-dependent kinase pathways as targets for cancer treatment. J Clin Oncol, 2006;24: 1770–1783. Fry DW, Harvey PJ, Keller PR et al. Specific inhibition of cyclindependent kinase 4/6 by PD 0332991 and associated antitumor activity in human tumor xenografts. Mol Cancer Ther, 2004;3: 1427–1438.

162


107. Lin TS, Dalton JT, Wu D et al. Flavopiridol given as a 30-min intravenous (IV) bolus followed by 4-hr continuous IV infusion (CIVI) results in clinical activity and tumor lysis in refractory chronic lymphocytic leukemia (CLL). J Clin Oncol, 2004 ASCO Annual Meeting Proceedings (Post-Meeting Edition). Vol 22, No 14S (July 15 Supplement), 2004:6564. 108. Martin A, Odajima J, Hunt SL et al. Cdk2 is dispensable for cell cycle inhibition and tumor suppression mediated by p27(Kip1) and p21(Cip1). Cancer Cell, 2005;7:591–598. 109. Fritz B, Schubert F, Wrobel G et al. Microarray-based copy number and expression profiling in dedifferentiated and pleomorphic liposarcoma. Cancer Res, 2002;62:2993–2998. 110. Arlander SJ, Eapen AK, Vroman BT, McDonald RJ, Toft DO, Karnitz LM. Hsp90 inhibition depletes Chk1 and sensitizes tumor cells to replication stress. J Biol Chem, 2003;278:52572–52577. 111. Richon VM, Sandhoff TW, Rifkind RA, Marks PA. Histone deacetylase inhibitor selectively induces p21WAF1 expression and gene-associated histone acetylation. Proc Natl Acad Sci U S A, 2000;97: 10014–10019. 112. Vassilev LT, Vu BT, Graves B et al. In vivo activation of the p53 pathway by small-molecule antagonists of MDM2. Science, 2004;303: 844–848. 113. Kim DM, Koo SY, Jeon K et al. Rapid induction of apoptosis by combination of flavopiridol and tumor necrosis factor (TNF)-alpha or TNFrelated apoptosis-inducing ligand in human cancer cell lines. Cancer Res,2003;63:621–626. 114. Dai Y, Rahmani M, Grant S. Proteasome inhibitors potentiate leukemic cell apoptosis induced by the cyclin-dependent kinase inhibitor flavopiridol through a SAPK/JNK- and NF-kappaB-dependent process. Oncogene, 2003;22:7108–7122. 115. Senderowicz AM. Targeting cell cycle and apoptosis for the treatment of human malignancies. Curr Opin Cell Biol, 2004;16:670–678. 116. McArthur GA, Raleigh J, Blasina A et al. Imaging with FLT-PET demonstrates that PF-477736, an inhibitor of CHK1 kinase, overcomes a cell cycle checkpoint induced by gemcitabine in PC-3 xenografts. Journal of Clinical Oncology, 2006 ASCO Annual Meeting Proceedings Part I. Vol 24, No. 18S (June 20 Supplement), 2006:3045.

7

CHFR as a Potential Anticancer Target Minoru Toyota, Lisa Kashima, and Takashi Tokino CONTENTS Introduction Identification of CHFR The Role of CHFR in the Early Prophase Checkpoint Molecular Mechanisms Modifying CHFR Function Genetic And Epigenetic Alteration of CHFR In Human Tumors Methylation of CHFR as a Molecular Marker to Predict Sensitivity to Microtubule Inhibitors CHFR as a Molecular Target for Cancer Therapy Concluding Remarks

Key Words: CHFR; mitotic checkpoint; prophase checkpoint; tubulin; E3 ubiquitin ligase; mitosis


163

164

Toyota, Kashima, and Tokino

1. INTRODUCTION Cancer arises through the accumulation of multiple genetic changes that include both gene deletion and amplification (1). Of the affected genes, those encoding proteins that comprise the mitotic checkpoints have been of particular interest to us. During the mitotic cell division cycle, cells make an exact copy of their chromosomes and physically tether the original and the replica together to form pairs of sister chromatids. Throughout this process, mitotic checkpoints guard against chromosome mis-segregation by delaying cell cycle progression. These checkpoints are each comprised of several molecules involved in what are termed the early mitotic checkpoint, the spindle assembly checkpoint and the late mitotic checkpoint. The best characterized of the three is the spindle assembly checkpoint, which mediates inhibition of the anaphase-promoting complex (APC) through the recruitment of several checkpoint proteins, including Bub1, Bub1R, Bub3, Mad1, and Mad2 (2–4). Notably, defects in the spindle assembly checkpoint have been implicated in tumorigenesis. For instance, Mad2+/− mice develop lung tumors at high rates after long latencies (5). However, extensive analysis of mitotic checkpoint genes in various types of cancers has shown that mutation of spindle assembly checkpoint genes is rare (6–8). Recently, another checkpoint functioning during early prophase was identified (Fig. 1) (9,10). In the absence of mitotic stress, deformation of the nuclear envelop associated with penetration by microtubules from maturing centrosomes occurs during prophase and is followed by breakdown of the nuclear envelope (11,12). In the presence of mitotic Interphase

Prophase

Centrosome

Chromosome condensation

Activation of cdc2/cyclin B1

Checkpoint off

Phosphorylation of histone H3 ser10

Checkpoint on

α-tubulin Cyclin B1

Centrosome maturation

Nuclear membrane breakdown

Fig. 1. Prophase checkpoint. Cellular phenotypes sen during interphase and prophase are shown. During the prophase, cdc2/Cyclin B1 is activated, the duplicated interphase chromatin condenses into chromosomes, and phosphorylation of ser10 of histone H3 takes place. In the presence of mitotic stress, the prophase checkpoint is switched on, and entry into mitosis is blocked.

Chapter 7 / CHFR as a Potential Anticancer Target

165

stress, however, breakdown of the nuclear envelope is blocked the early prophase checkpoint. In this review, we will focus on the activities of CHFR, a novel component of the early prophase checkpoint. Because the decision whether or not to break down the nuclear envelope is a critical determinant of a cancer cell’s susceptibility to microtubule inhibitors, CHFR could be a useful target for cancer therapy.

2. IDENTIFICATION OF CHFR CHFR was first identified as a yeast homologue containing an FHA domain (10). Normal primary cells and tumor cell lines that express CHFR exhibit delayed entry into metaphase when centrosome separation is inhibited by mitotic stress. By contrast, tumor cell lines in which CHFR expression is silenced enter metaphase without delay. Ectopic expression of wild-type CHFR restores the cell cycle delay and increases the likelihood that the cells will survive the mitotic stress. Comprised of 664 amino acids, CHFR contains an FHA domain, which mediates interaction with phosphorylated proteins, and a ring finger domain, which contributes to the protein’s ubiquitin ligase activity. Blast analysis of CHFR cDNA revealed the gene to contain 18 exons spanning about 60 kb on chromosome 12q24. The sequence of human CHFR is similar to that of fission yeast DMA1 and the budding yeast homologues DMA1 and DMA2, which are involved in a later mitotic checkpoint that delays a cell’s exit from mitosis in the event of spindle damage (13,14).

3. THE ROLE OF CHFR IN THE EARLY PROPHASE CHECKPOINT In normal cells, chromosome condensation is delayed by drugs such as paclitaxel and docetaxel, which induce mitotic stress by disrupting the structure of microtubules (10). That delay reflects in part suppression of cdc2/cyclin B1 activity, which is controlled by the molecule’s localization. Normally, cyclin B1 accumulates in the cytoplasm during S and G2 phases and is then translocated to the nucleus during prophase. Toyoshima et al. showed that the cytoplasmic localization of cyclin B1 during interphase is directed by a nuclear export signal (NES)-dependent transport system and that introduction of NES-disrupted cyclin B1 into cells activates a DNA damage-induced G2 checkpoint (15). When mitotic stress occurs in cancer cells, cyclin B1 accumulates in cytoplasm of cells that express wild-type CHFR, but accumulates in nucleus of cells that lack CHFR expression (16–18). Thus, CHFR appears to suppress entry into mitosis by sequestering cyclin B1 in cytoplasm.

166


Activation of cyclin B1 by cyclin B1/cdc2 kinase and its subsequent translocation to the nucleus precede the apoptosis induced by microtubule inhibitors (19). Conversely, inhibition of cyclin B1/cdc2 kinase activity by the inhibitor p21WAF1 reduces the incidence of apoptosis. In addition, we have shown that nuclear localization of cyclin B1 is strongly associated with the sensitivity of cancer cells to microtubule inhibitors (16,17), which is consistent with an earlier report showing that nuclear localization of cyclin B1 correlates with sensitivity to chemotherapeutic drugs and that leptomycin B, an inhibitor of nuclear export of cyclin B1, sensitizes cancer cells to chemotherapeutic drugs (20). The mechanism by which nuclear localization of cyclin B1 triggers apoptotic signaling remains unknown, however. CHFR contains a ring finger domain characteristic to E3 ubiquitin ligases, and its ubiquitin ligase activity has been demonstrated both in vitro and in vivo (21–23). For instance, CHFR is known to ubiquitinate itself and accumulate during G2/M phase. Another potential target for ubiquitination by CHFR is the oncogenic serine/threonine kinase PLK1, which has several substrates, including cyclin B1, and is essential for mitosis (Fig. 2) and for maintenance of genomic mitotic stress G2

mid prophase

early prophase

PKB/Akt

late prophase

CHFR Aurora B

PML

K63 Ub Ub

Ub

P Histone H3

Aurora A

chromosome condensation

PIk1 P Aurora A

p38

repress activate interact

Cdc25

Cdc2 Cyclin B1

nuclear envelope breakdown

Wee1

nuclear translocation

Fig. 2. Schematic representation of a prophase checkpoint involving CHFR. CHFR delays entry into mitosis by inhibiting PLK1 and Cyclin B1.


167

integrity (24,25). Notably, PLK1 is overexpressed in various human cancers (26–29), leading to phosphorylation of cyclin B1 and its accumulation in nucleus, where it mediates entry into mitosis (30). Kang et al. reported that CHFR can mediate degradation of PLK1, leading to inhibition of Cdc25C and Cdc2/cyclin B1 in Xenopus cell free extracts (23), though other evidence suggests that CHFR is a noncanonical ubiquitin ligase that does not target proteins for degradation, but rather utilizes its ubiquitin ligase activity for signal transduction (21). Consistent with that idea is the finding that no significant changes in the levels of endogenous PLK1 are seen in CHFR-delayed cells (18). Another target for CHFR-mediated ubiquitination is Aurora-A kinase. Aurora-A is an oncogene located on human chromosome 20q.13, and is overexpressed in various human tumors as a result of gene amplification (31–34). Indeed, mouse NIH-3T3 cells transfected with Aurora A develop into tumors when implanted in nude mice. Recently, Yu et al. found that CHFR physically interacts with and ubiquitinates Aurora-A both in vivo and in vitro (35). Moreover, levels of Aurora A were lower in mouse embryonic fibroblasts expressing wild-type CHFR than in CHFR-deficient cells, and levels of Aurora A and its association with CHFR were increased after the cells were treated with the proteosome inhibitor MG132. Thus, CHFR appears to control the expression levels of both PLK1 and Aurora A, 2 oncogenic proteins involved in mitosis. The molecular pathway via which CHFR activates the prophase checkpoint is poorly understood, but the fact that CHFR has noncanonical ubiquitin ligase activity suggests it may be associated with a known signaling pathway. For instance, Matsuoka and Pines proposed that activated TAK1 activates CHFR, which in turn activates the p38 stress kinase pathway (36). They also found that p38 kinases are required for the early prophase checkpoint, and that introduction of p38 or p38 into cells activates the checkpoint. Conversely, inhibition of p38 abrogated the early prophase checkpoint. Activation of the early prophase checkpoint also was lost in cancer cells in which CHFR expression was silenced, suggesting p38 is a downstream effector of CHFR. TAK1 is a MAP kinase kinase kinase in the p38 kinase pathway and can be activated by TRAF6 (37). In this pathway, activated TRAF6 functions as a ubiquitin ligase and polyubiquitinates itself. The polyubiquitinated mediator then activates TAK1, which leads to the activation of NF-kB. Ubc13-Mms heterodimer is

168


the ubiquitin-conjugating enzyme and Lys63-polyubiquitin chains are formed. In that regard, Bothos et al. found that CHFR catalyzes the formation of noncanonical Lys63-polyubiquitin chains, with Ubc13Mms2 acting as the polyubiquitin-conjugating enzyme (21).

4. MOLECULAR MECHANISMS MODIFYING CHFR FUNCTION CHFR activity is known to be modulated through both phosphorylation and ubiquitination, though the precise molecular mechanisms remain to be determined. For instance, CHFR is phosphorylated by protein kinase B (PKB)/AKT, the central effector of the phosphatidylinositol 3-kinase (PI3K) signal transduction pathway, which is now known to be frequently activated in human tumors. Expression of a CHFR mutant that could not be phosphorylated by PKB led to a reduction in PLK1 levels and inhibition of entry into mitosis (38). It may be that the deregulated PKB activity frequently seen in tumors inhibits CHFR activity and thereby promotes the accumulation of PLK1, leading to disruption of the checkpoint. CHFR is known in localize in the nucleus, but the precise mechanism governing the distribution of the protein remains unknown. Daniels et al. reported that the intranuclear distribution of CHFR is controlled by PML (39), a target of the t(15;17) chromosomal translocation typical of promyelocytic acute myeloid leukemia, which reciprocally fuses PML to retinoic acid receptor (40). Overexpression of PML in cancer cell lines suppresses cell growth and induces apoptosis, suggesting PML is a tumor suppressor (41). Moreover, following microtubule disruption, PML-/- cells do not exhibit the delayed entry into mitosis that characterizes the CHFR-dependent checkpoint, suggesting PML also plays a role in the checkpoint (39).

5. GENETIC AND EPIGENETIC ALTERATION OF CHFR IN HUMAN TUMORS Studies indicate that CHFR is not expressed in a subset of human tumor cell lines (10). Mariatos et al. identified CHFR gene mutations in 3 of 53 (5.7%) lung cancers (42). All of the tumor-associated mutations were missense mutations, and the CHFR proteins encoded by the mutants were defective and unable to serve as checkpoints against mitotic stress. No CHFR mutations were detected in any other types of cancer; in fact, genetic inactivation of CHFR appears to be a rare event in human tumors (17,43).


169

DNA methylation is another important molecular event that leads to gene silencing in cancer (44,45). The targets of methylation are CpG islands, which are genetic regions with guanine plus cytosine contents of >55% (46) situated within the 5 untranslated regulatory sequences and first exons of genes containing >500 bp. About 40% of mammalian genes contain CpG islands within their promoters, and the consequence of their methylation is transcriptional silencing. As is the case with other cancer related genes, CHFR gene is primarily silenced by methylation of its 5 CpG island, not by gene deletion or mutation. Aberrantly methylated CHFR has been detected in a variety of tumor types including colon (43,47), stomach (17), lung (47,48), breast (49,50), nasopharyngeal (51), oral (16) and hematopoietic tumors (52). Methylated CHFR also has been detected in premalignant tissues, such as colorectal adenomas, indicating that epigenetic silencing of CHFR is an early event during tumorigenesis (43). That expression of CHFR is restored when silenced cells are treated with a methyltransferase inhibitor (5-aza-dC) confirms that the absence of CHFR expression is the result of DNA methylation and not, for example, altered expression of a transcription factor. The molecular mechanism by which DNA methylation occurs in cancer remains unknown. In mammals, methylation of cytosine is mediated by 3 DNA methyltransferases, DNMT1, DNMT3A and DNMT3B, and DNMT1 appears to be highly expressed in human tumors (53), though several reports suggest DNMT expression does not correlate with gene methylation (54). On the other hand, there does appear to be a correlation between DNA methylation and viral infection (55,56), and Kusano et al. frequently detected methylation of CHFR in Epstein-Barr virus-associated gastric cancers (57). Subsets of cancers show simultaneous methylation of multiple genes, which is characteristic of the CpG island methylator phenotype (CIMP) (58–60). In colorectal and gastric cancer, CHFR methylation is frequently detected in tumors expressing CIMP, and Jass et al. proposed that serrated adenoma is a precursor of CIMP-positive colorectal cancers (61,62). Those investigators also observed extensive DNA methylation in normal colon mucosa from patients with multiple hyperplastic polyps, suggesting there may be a genetic basis for CIMP. Because CHFR is involved in a mitotic checkpoint, it has been speculated that the process of chromosomal segregation is defective in tumors with CHFR methylation, which leads to chromosomal instability. Consistent with that idea, CHFR-/- mouse embryonic fibroblasts show increased numbers of aneuploid cells, and inhibition of Aurora A using short hairpin (sh)RNA reduced the frequency of such cells. Thus,

170


another function of CHFR may be to maintain chromosomal integrity. In contrast to mouse cells, however, human cancer cells with CHFR methylation tend to be diploid, suggesting CHFR methylation is not a cause of chromosomal instability in those cases (63). Instead, colorectal cancers with CHFR methylation show microsatellite instability (63,64), which suggests that inactivation of the prophase checkpoint may be a factor contributing to the accumulation of mutations.

6. METHYLATION OF CHFR AS A MOLECULAR MARKER TO PREDICT SENSITIVITY TO MICROTUBULE INHIBITORS Cell cycle checkpoint dysfunction is often associated with sensitivity to chemotherapeutic agents. For example, overexpression of MAD2 sensitizes cancer cells to both cisplatin and vincristine (65,66), whereas overexpression of Aurora A induces chemoresistance (34). In addition, Satoh et al. found that the microtubule inhibitors paclitaxel and docetaxel induced apoptosis among gastric cancer cells showing CHFR methylation and that adenoviral introduction of CHFR into methylated cancer cell lines restores the checkpoint and reduces the incidence of apoptosis (17). This correlation between CHFR methylation and sensitivity to microtubule inhibitors appears to be specific, as there was no correlation between CHFR methylation and sensitivity to other chemotherapeutic agents (e.g., VP16) or to UV. These findings suggest that CHFR methylation could serve as a molecular marker predictive of the sensitivity of tumors to microtubule inhibitors. Consistent with that idea, Koga et al. (67) found that 6 (86%) of 7 patients with methylated CHFR tumors showed some regression or no progression of their disease when treated with a microtubule inhibitor, whereas 4 (80%) of 5 patients with unmethylated CHFR tumor showed progressive deterioration. A correlation between CHFR methylation and sensitivity to microtubule inhibitors also has been noted in oral squamous cell carcinoma (16). Thus, CHFR methylation may be a clinically useful indicator of the responsiveness of cancers to treatment with microtubule inhibitors.

7. CHFR AS A MOLECULAR TARGET FOR CANCER THERAPY The fact that CHFR is frequently inactivated by genetic or epigenetic alteration in human cancers suggests that this cancer-specific checkpoint defect also could be a useful therapeutic target. Bearing


Microtubule inhibitors e.g. Docetaxel

Cancer with CHFR checkpoint

171

CHFR

Mitotic stress

Arrest at G2/M

Repair damage Proliferation

DNA methylation of CHFR Microtubule inhibitors e.g. Docetaxel

Cancer without CHFR checkpoint

Mitotic stress

Knock down by shRNA

Continue mitosis Apoptosis

Fig. 3. Sensitivity of cancer cells to microtubule inhibitors is dependent on whether the CHFR checkpoint is functional: top, cancer with normal checkpoint; bottom, cancer with an impaired checkpoint. Both DNA methylation and shRNAmediated silencing of CHFR can increase the sensitivity of otherwise resistant cells to microtubule inhibitors.

that in mind, we recently established a system to knock down CHFR expression using shRNA (Fig. 3) (16). We found that CHFR expression was significantly suppressed in cancer cells transfected with shRNA. The resultant impairment of the prophase checkpoint led to an increased mitotic index in cells treated with microtubule inhibitors, which in turn led to an increased incidence of apoptosis among the cells. This effect was specific to microtubule inhibitors, as no effect was seen when a DNA damaging agent (cisplatin or VP16) was used. In addition, the earlier finding that E3 ubiquitin ligases can be targeted using small molecules (68) suggests drugs that inhibit CHFR’s ubiquitin ligase activity also could be useful for enhancing the sensitivity of cancer cells to microtubule inhibitors.

8. CONCLUDING REMARKS About half of human neoplasias show an intact checkpoint against mitotic stress, which diminishes the efficacy of paclitaxel and docetaxel. We propose that disruption of the prophase checkpoint by suppressing CHFR expression using shRNA or inhibiting CHFR’s

172


ubiquitin ligase activity using small molecules could represent novel strategies for enhancing the sensitivity of cancer cells to microtubule inhibitors.

ACKNOWLEDGMENTS The authors thank Dr. William F. Goldman for editing the manuscript. This study was supported in part by Grants-in-Aid for Scientific Research on Priority Areas from the Ministry of Education, Culture, Sports, Science, and Technology (T.T. and M.T.).

REFERENCES 1. Kinzler, K. W., Vogelstein, B. Lessons from hereditary colorectal cancer. Cell 1996;87: 159–70. 2. Fang, G. Checkpoint protein BubR1 acts synergistically with Mad2 to inhibit anaphase-promoting complex. Mol Biol Cell 2002;13: 755–66. 3. Sudakin, V., Chan, G. K., Yen, T. J. Checkpoint inhibition of the APC/C in HeLa cells is mediated by a complex of BUBR1, BUB3, CDC20, and MAD2. J Cell Biol 2001;154: 925–936. 4. Tang, Z., Bharadwaj, R., Li, B., Yu, H. Mad2-Independent inhibition of APCCdc20 by the mitotic checkpoint protein BubR1. Dev Cell 2001;1: 227–237. 5. Michel, L. S., Liberal, V., Chatterjee, A., et al. MAD2 haplo-insufficiency causes premature anaphase and chromosome instability in mammalian cells. Nature 2001;409: 355–359. 6. Cahill, D. P., da Costa, L. T., Carson-Walter, E. B., et al. Characterization of MAD2B and other mitotic spindle checkpoint genes. Genomics 1999;58: 181–187. 7. Cahill DP„ Lengauer C, Yu J, et al. Mutations of mitotic checkpoint genes in human cancers. Nature 1998;392: 300–303. 8. Wang Z, Cummins JM, Shen D, et al. Three classes of genes mutated in colorectal cancers with chromosomal instability. Cancer Res 2004;64: 2998–3001. 9. Cortez D, Elledge SJ. Conducting the mitotic symphony. Nature 2000;406: 354–356. 10. Scolnick DM, Halazonetis TD. Chfr defines a mitotic stress checkpoint that delays entry into metaphase. Nature 2000;406: 430–435. 11. Georgatos SD, Pyrpasopoulou A, Theodoropoulos PA. Nuclear envelope breakdown in mammalian cells involves stepwise lamina disassembly and microtubule-drive deformation of the nuclear membrane. J Cell Sci 1997;110 (Pt 17): 2129–2140.


173

12. Salina D, Bodoor K, Eckley DM, et al. Cytoplasmic dynein as a facilitator of nuclear envelope breakdown. Cell 2002;108: 97–107. 13. Fraschini R, Bilotta D, Lucchini G, Piatti S. Functional characterization of Dma1 and Dma2, the budding yeast homologues of Schizosaccharomyces pombe Dma1 and human Chfr. Mol Biol Cell 2004;15: 3796–3810. 14. Murone M, Simanis V. The fission yeast dma1 gene is a component of the spindle assembly checkpoint, required to prevent septum formation and premature exit from mitosis if spindle function is compromised. Embo J 1996;15: 6605–6616. 15. Toyoshima F, Moriguchi T, Wada A, Fukuda M, Nishida E. Nuclear export of cyclin B1 and its possible role in the DNA damage-induced G2 checkpoint. Embo J 1998;17: 2728–2735. 16. Ogi K, Toyota M, Mita H, et al. Small interfering RNA-induced CHFR silencing sensitizes oral squamous cell cancer cells to microtubule inhibitors. Cancer Biol Ther 2005;4: 773–778. 17. Satoh A, Toyota M, Itoh F, et al. Epigenetic inactivation of CHFR and sensitivity to microtubule inhibitors in gastric cancer. Cancer Res 2003;63: 8606–8613. 18. Summers MK, Bothos J, Halazonetis TD. The CHFR mitotic checkpoint protein delays cell cycle progression by excluding Cyclin B1 from the nucleus. Oncogene 2005;24: 2589–2598. 19. Heliez C, Baricault L, Barboule N, Valette A. Paclitaxel increases p21 synthesis and accumulation of its AKT-phosphorylated form in the cytoplasm of cancer cells. Oncogene 2003;22: 3260–3268. 20. Porter LA, Cukier IH, Lee JM. Nuclear localization of cyclin B1 regulates DNA damage-induced apoptosis. Blood 2003;101: 1928–1933. 21. Bothos J, Summers MK, Venere M, Scolnick DM, Halazonetis TD. The Chfr mitotic checkpoint protein functions with Ubc13-Mms2 to form Lys63-linked polyubiquitin chains. Oncogene 2003;22: 7101–7107. 22. Chaturvedi P, Sudakin V, Bobiak ML, et al. Chfr regulates a mitotic stress pathway through its RING-finger domain with ubiquitin ligase activity. Cancer Res 2002;62: 1797–1801. 23. Kang D, Chen J, Wong J, Fang G.. The checkpoint protein Chfr is a ligase that ubiquitinates Plk1 and inhibits Cdc2 at the G2 to M transition. J Cell Biol 2002;156: 249–259. 24. Strebhardt K, and Ullrich A. Targeting polo-like kinase 1 for cancer therapy. Nat Rev Cancer 2006;6: 321–330. 25. Xie S, Xie B, Lee MY, Dai W. Regulation of cell cycle checkpoints by polo-like kinases. Oncogene 2005;24: 277–286. 26. Takahashi T, Sano B, Nagata T, et al. Polo-like kinase 1 (PLK1) is overexpressed in primary colorectal cancers. Cancer Sci 2003;94: 148–152.

174


27. Weichert W, Denkert C, Schmidt M, et al. Polo-like kinase isoform expression is a prognostic factor in ovarian carcinoma. Br J Cancer 2004;90: 815–821. 28. Weichert W, Schmidt M, Gekeler V, et al. Polo-like kinase 1 is overexpressed in prostate cancer and linked to higher tumor grades. Prostate 2004;60: 240–245. 29. Weichert W, Schmidt M, Jacob J, et al. Overexpression of Polo-like kinase 1 is a common and early event in pancreatic cancer. Pancreatology 2005;5: 259–265. 30. Toyoshima-Morimoto F, Taniguchi E, Shinya N, Iwamatsu A, Nishida E. Polo-like kinase 1 phosphorylates cyclin B1 and targets it to the nucleus during prophase. Nature 2001;410: 215–220. 31. Jeng YM, Peng SY, Lin CY, Hsu HC. Overexpression and amplification of Aurora-A in hepatocellular carcinoma. Clin Cancer Res 2004;10: 2065–2071. 32. Gritsko TM, Coppola D, Paciga JE, et al. Activation and overexpression of centrosome kinase BTAK/Aurora-A in human ovarian cancer. Clin Cancer Res 2003;9: 1420–1426. 33. Li D, Zhu J, Firozi PF, et al. Overexpression of oncogenic STK15/BTAK/Aurora A kinase in human pancreatic cancer. Clin Cancer Res 2003;9: 991–997. 34. Anand S, Penrhyn-Lowe S, Venkitaraman AR. AURORA-A amplification overrides the mitotic spindle assembly checkpoint, inducing resistance to Taxol. Cancer Cell 2003;3: 51–62. 35. Yu X, Minter-Dykhouse K, Malureanu L, et al. Chfr is required for tumor suppression and Aurora A regulation. Nat Genet 2005;37: 401–406. 36. Matsusaka T, and Pines J. Chfr acts with the p38 stress kinases to block entry to mitosis in mammalian cells. J Cell Biol 2004;166: 507–516. 37. Wang C, Deng L, Hong M, et al. TAK1 is a ubiquitin-dependent kinase of MKK and IKK. Nature 2001;412: 346–351. 38. Shtivelman E. Promotion of mitosis by activated protein kinase B after DNA damage involves polo-like kinase 1 and checkpoint protein CHFR. Mol Cancer Res 2003;1: 959–969. 39. Daniels MJ, Marson A, Venkitaraman AR. PML bodies control the nuclear dynamics and function of the CHFR mitotic checkpoint protein. Nat Struct Mol Biol 2004;11: 1114–1121. 40. Melnick A, and Licht JD. Deconstructing a disease: RARalpha, its fusion partners, and their roles in the pathogenesis of acute promyelocytic leukemia. Blood 1999;93: 3167–3215. 41. Takahashi Y, Lallemand-Breitenbach V, Zhu J, de The H. PML nuclear bodies and apoptosis. Oncogene 2004;23: 2819–2824. 42. Mariatos G, Bothos J, Zacharatos P, et al. Inactivating mutations targeting the chfr mitotic checkpoint gene in human lung cancer. Cancer Res 2003;63: 7185–7189.


175

43. Toyota M, Sasaki Y, Satoh A, et al. Epigenetic inactivation of CHFR in human tumors. Proc Natl Acad Sci U S A 2003;100: 7818–7823. 44. Jones PA, Baylin S B. The fundamental role of epigenetic events in cancer. Nat Rev Genet 2002;3: 415–428. 45. Toyota M, Issa JP. Epigenetic changes in solid and hematopoietic tumors. Semin Oncol 2005;32: 521–530. 46. Takai D, Jones PA. Comprehensive analysis of CpG islands in human chromosomes 21 and 22. Proc Natl Acad Sci U S A 2002;99: 3740–3745. 47. Corn PG., Summers MK, Fogt F, et al. Frequent hypermethylation of the 5 CpG island of the mitotic stress checkpoint gene Chfr in colorectal and non-small cell lung cancer. Carcinogenesis 2003;24: 47–51. 48. Mizuno K, Osada H, Konish, H, et al. Aberrant hypermethylation of the CHFR prophase checkpoint gene in human lung cancers. Oncogene 2002;21: 2328–2333. 49. Erson AE, Petty EM. CHFR-associated early G2/M checkpoint defects in breast cancer cells. Mol Carcinog 2004;39: 26–33. 50. Tokunaga,E, Oki E, Nishida K, et al. Aberrant hypermethylation of the promoter region of the CHFR gene is rare in primary breast cancer. Breast Cancer Res Treat 2006;97: 199–203. 51. Cheung HW, Ching YP, Nicholls JM, et al. Epigenetic inactivation of CHFR in nasopharyngeal carcinoma through promoter methylation. Mol Carcinog 2005;43: 237–245. 52. van Doorn R, Zoutman WH, Dijkman R, et al. Epigenetic profiling of cutaneous T-cell lymphoma: promoter hypermethylation of multiple tumor suppressor genes including BCL7a, PTPRG, and p73. J Clin Oncol 2005;23: 3886–3896. 53. Etoh T, Kanai Y, Ushijima S, et al. Increased DNA methyltransferase 1 (DNMT1) protein expression correlates significantly with poorer tumor differentiation and frequent DNA hypermethylation of multiple CpG islands in gastric cancers. Am J Pathol 2004;164: 689–699. 54. Eads CA, Danenberg KD, Kawakami K, et al. CpG island hypermethylation in human colorectal tumors is not associated with DNA methyltransferase overexpression. Cancer Res 1999;59: 2302–2306. 55. Kaneto H, Sasaki S, Yamamoto H, et al. Detection of hypermethylation of the p16(INK4A) gene promoter in chronic hepatitis and cirrhosis associated with hepatitis B or C virus. Gut 2001;48: 372–377. 56. Suzuki M, Toyooka S, Shivapurkar N, et al. Aberrant methylation profile of human malignant mesotheliomas and its relationship to SV40 infection. Oncogene 2005;24: 1302–1308. 57. Kusano M, Toyota M, Suzuki H, et al. Genetic, epigenetic, and clinicopathologic features of gastric carcinomas with the CpG island methylator phenotype and an association with Epstein-Barr virus. Cancer 2006;106: 1467–1479.

176


58. Toyota M, Ahuja N, Ohe-Toyota M, et al. CpG island methylator phenotype in colorectal cancer. Proc Natl Acad Sci U S A 1999;96: 8681–8686. 59. Toyota M, Ahuja N, Suzuki H, et al. Aberrant methylation in gastric cancer associated with the CpG island methylator phenotype. Cancer Res 1999;59: 5438–5442. 60. Toyota M, Ohe-Toyota M, Ahuja N, Issa JP. Distinct genetic profiles in colorectal tumors with or without the CpG island methylator phenotype. Proc Natl Acad Sci U S A 2000;97: 710–715. 61. Wynter CV, Walsh MD, Higuchi T, et al. Methylation patterns define two types of hyperplastic polyp associated with colorectal cancer. Gut 2004;53: 573–580. 62. Minoo P, Baker K, Goswami R, et al. Extensive DNA methylation in normal colorectal mucosa in hyperplastic polyposis. Gut 2006;55: 1467–1474. 63. Bertholon J, Wang Q, Falette N, et al. Chfr inactivation is not associated to chromosomal instability in colon cancers. Oncogene 2003;22: 8956–8960. 64. Brandes JC, van Engeland M, Wouters KA, Weijenberg,MP, Herman JG. CHFR promoter hypermethylation in colon cancer correlates with the microsatellite instability phenotype. Carcinogenesis 2005;26: 1152–1156. 65. Cheung HW, Jin DY, Ling MT et al. Mitotic arrest deficient 2 expression induces chemosensitization to a DNA-damaging agent, cisplatin, in nasopharyngeal carcinoma cells. Cancer Res 2005;65: 1450–1458. 66. Wang X, Jin DY, Wong HL, et al. MAD2-induced sensitization to vincristine is associated with mitotic arrest and Raf/Bcl-2 phosphorylation in nasopharyngeal carcinoma cells. Oncogene 2003;22: 109–116. 67. Koga Y, Kitajima Y, Miyoshi A, et al. The significance of aberrant CHFR methylation for clinical response to microtubule inhibitors in gastric cancer. J Gastroenterol 2006;41: 133–139. 68. Sun Y. Targeting E3 ubiquitin ligases for cancer therapy. Cancer Biol Ther 2003;2: 623–629.

8

Antimicrotubule Agents Miguel A. Villalona-Calero, Larry Schaaf, and Robert Turowski CONTENTS Tubulin, Cellular Organization and Properties Tubulin Interactive Antitumor Agents: Discovery and Mechanism of Action Clinical Development and Approved Indications Tumor Resistance to Tubulin Interactive Agents Novel Tubulin Interactive Agents in Clinical Development

Abstract The structural protein tubulin has been validated as an antineoplastic target by the successful clinical development of several anticancer agents. Natural product discovery led to the initial compounds, while refinement of their chemical structure and bioavailability has led to a few others. In this chapter we will discuss the cellular organization and properties of tubulin, as well as the discovery, mechanism of action, toxicity, and the clinical trials that support the clinical indications of currently approved tubulinFrom: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

177

178

Villalona-Calero, Schaaf, and Turowski

interactive agents. We will also discuss novel antimicrotubular chemical entities in development, which feature enhanced biodelivery or circumvention of tumoral drug resistance. Key Words: Tubulin; vinca alkaloids; taxanes; epothilones; tumor resistance; albumin-bound

1. TUBULIN, CELLULAR ORGANIZATION AND PROPERTIES Mammalian cells regulate intracellular transport and the assembly of the mitotic spindle by specialized filaments named microtubules. They are polarized structures composed of heterodimers of and tubulin, distinct proteins that have been highly conserved through evolution. Both monomers share about 40% similarity in sequence, and resolution through crystallography reveals a core of 2 beta sheets surrounded by helices (1). The dimers bind guanosine triphosphate (GTP) within the subunit and polymerize onto a growing microtubule end coinciding with GTP hydrolysis to GDP. The subunit in 1 dimer aligns with the subunit on the next, longitudinally around a hollow core. Two dynamic processes control microtubule function. The first is mediated by GTP hydrolysis, which weakens the affinity of tubulin for other tubulin molecules. Because it is irreversible, net growing occurs at the plus end of the microtubule, but net shortening at the minus end, a process that has been termed treadmilling (2). The second, dynamic instability, features the switching at microtubule ends between slow sustained growth and rapid shortening (3,4). Both processes can coexist and the degree in which they are present is highly dependent on cellular conditions. For example, during mitosis highly dynamic microtubules help in the attachment and movement of the chromosomes, while rapid treadmilling help in the flow of signals from kinetochores to the poles during metaphase and anaphase (5,6). Other than GTP and concentrations of free tubulin, various chemical mediators, such as magnesium or calcium, have been shown to influence the promotion of assembly or disassembly of microtubules (7). In addition, the binding of microtubule-associated proteins (MAPs) to modified regions of polymerized tubulin stabilize microtubules against disassembly and mediate interactions with other cellular components (8). Microtubules assemble in 13 protofilaments around the microtubule organizing centers (MTOC) or centrosomes. A third member of the tubulin superfamily of proteins, tubulin has been shown to be in

Chapter 8 / Antimicrotubule Agents

179

direct contact with the minus end of the microtubules and is thought to be responsible for the nucleation of microtubule assembly by MTOC (9). Before cell division, the MTOC duplicates and separates forming the two poles of the mitotic spindle.

2. TUBULIN INTERACTIVE ANTITUMOR AGENTS: DISCOVERY AND MECHANISM OF ACTION 2.1. Discovery The first tubulin interactive antitumor agents available to oncologists were the naturally occurring members of the Vinca alkaloids, vinblastine and vincristine (10). Both of these compounds were isolated from the leaves of the Madagascar periwinkle plant (Catharanthus roseus, formerly Vinca rosea). In the late 1950s, the antimitotic activity and chemotherapeutic potential was discovered by groups both at the University of Western Ontario and Eli Lilly Research Laboratory (11,12). In contrast to taxol, the discovery of the vinca alkaloids was a “chance” discovery. In 1958, researchers led by Dr. Noble at Western Ontario were looking for a source for oral hypoglycemic agents to treat diabetes. Rats administered oral periwinkle extracts had no change in blood sugar or glucagon levels. However, injection of periwinkle extracts into rats resulted in a decline in white blood cell count. This unexpected decrease in white blood cell count was the first hint that compounds in the periwinkle plant possessed cytotoxic activity (13,14). A colleague, Dr. Charles Beer, extracted, isolated, and purified a chemical from periwinkle, which he named “vincaleukoblastine” (i.e., vinblastine). Researchers at Eli Lilly subsequently isolated and characterized another compound, vincristine. Vincristine was approved by the FDA in 1963 and vinblastine was approved 2 years later. These 2 agents gained widespread use for the single-agent treatment of childhood hematological and solid malignancies, and shortly after, for adult hematological malignancies. During the ensuing 40 years, they have become the cornerstone of several combination therapies and this success has led to the development of various novel semi-synthetic analogues, including vinorelbine, vindesine, and vinflunine. The development of paclitaxel (taxol) from basic science discovery to viable pharmaceutical product took an exciting but circuitous route that lasted approximately 30 years (15–18). Taxol was discovered through a collaborative agreement in 1960 between the National Cancer Institute (NCI) and the US Department of Agriculture (USDA) to screen plant tissues in North America for antitumor activity. In August

180


1962, USDA botanists collected 650 plant samples including bark, twigs, leaves and fruit of the Pacific yew tree (Taxus brevifolia) in Washington State. The initial screening of yew extracts demonstrated 9KB cytotoxicity in 1964 and a recollection was assigned to the research team headed by Dr. Monroe E. Wall and Dr. Mansukh C. Wani at the Research Triangle Institute for fractionation in 1965. “Taxol” was isolated as the active constituent in 1967, and the report of its isolation and structural elucidation was published in 1971 (18,19). Taxol showed strong activity in vivo against P1534 leukemia, but the NCI did not consider this predictive of clinical activity, especially because taxol was not very active against either L1210 or P388 leukemia models (17). Thus, the NCI considered it no more promising a candidate for further development than a number of other available compounds, especially considering the compound was present in extremely low amounts, the extraction and isolation was difficult, and the supply of the yew tree was limited (15). Fortunately, 2 major developments occurred that revitalized interest in taxol. The first was the discovery that it demonstrated very strong activity against the B16 melanoma assay that had been introduced by the NCI in 1975 (15). Subsequent observation of its significant activity against several human tumor xenograft systems, including the MX-1 mammary tumor, provided further evidence of its superior spectrum of activity. This led the NCI to accept taxol as a drug candidate for preclinical development in 1977. The second development that stimulated interest in taxol was a series of studies by Dr. Susan Horwitz and coworkers at Albert Einstein Medical Center in New York demonstrating a unique mechanism of action in promoting tubulin polymerization and stabilization of microtubules against depolymerization (20,21). Despite these exciting discoveries, the development of taxol followed a tortuous route (21,22). Approval was granted for initiation of Phase I clinical trials in 1983. The initial trials were fraught with serious problems of toxicity, particularly allergic reactions including anaphylaxis, and the drug was very close to being dropped from clinical study for reasons of safety. This can be traced back to the extremely poor solubility of taxol in aqueous systems and its relatively high dose requirements compared to other antitumor natural products, such as the Vinca alkaloids, vinblastine and vincristine. Development of a suitable formulation for parenteral administration was extremely difficult, and ultimately required the use of a surfactant formulation in which Cremophor EL, a polyethoxylated castor oil derivative, is a major component (18). Cremophor is known to cause histamine release


181

in dogs and is believed to be implicated in hypersensitivity reactions to other drugs where it is a component of the formulation. The concentration of Cremophor EL in the taxol formulation developed for clinical trials results in a dose of Cremophor EL that is at least twice that used with any other experimental drug. Throughout the 1980s, taxol progressed through Phases I, II, and III of clinical trials. However, the effort to supply drug was difficult and expensive. In an effort to obtain adequate supplies of taxol, the NCI advertised in 1989 that it had a Cooperative Research and Development (CRADA) to grant to the pharmaceutical company submitting the best proposal (15). The award was issued to Bristol-Myers Squibb. The company moved rapidly and in 1992, 21 years after the original publication on taxol, the FDA approved its use for treatment of refractory ovarian cancer (21). At present, semi-synthesis remains the only commercially viable route for producing this drug in the West. The method relies on the use of a precursor obtained from the less-endangered European yew; and from needles, rather than bark. The compound baccatin III, otherwise known as DAB III, is isolated, and through a still-complex series of synthetic steps, it is transformed into taxol. In 1986, during research on a semisynthetic route to taxol from 10-deacetylbaccatin III obtained from needles of Taxus baccata (European yew tree), another effective chemotherapeutic agent, docetaxel (marketed as Taxotere by SanofiAventis) was discovered. Human studies with docetaxel were initiated in 1989 and the drug was approved by the FDA to treat certain patients with locally advanced or metastatic breast cancer in 1996. Although docetaxel is more water soluble than taxol, it still requires polysorbate80 and ethanol in its traditional formulation and premedications are needed to minimize hypersensitivity reactions. Other chemical changes to the taxane molecule have led to a number of paclitaxel and docetaxel analogues. Alterations in the structure of the taxane side chains have led to the development of water-soluble analogues and agents with substantial oral bioavailability. Conjugates of paclitaxel to other moieties, including polyethylene glycol and polyglutamate, and liposomal encapsulation of the molecule are also strategies being investigated. Only a few of these have reached the point of adequate testing in humans and their clinical utility is not known.

182


2.2. Mechanism of Action As noted above, microtubules are key components of the cytoskeleton. They are essential in all eukaryotic cells because they are critical in the development and maintenance of cell shape, in the transport of vesicles, mitochondria and other cell components throughout cells. They also play a major role in cell signaling, and in cell division and mitosis (10). It is during cell division that microtubules form the mitotic spindle that promotes alignment of chromosomes to the equatorial plane and mediates the subsequent division of chromosomes to the 2 daughter cells (23,24). It is this key role in cell division that makes microtubules an important target for the development of chemotherapeutic drugs that effect rapidly dividing cancer cells. There is considerable evidence in the literature demonstrating that subtle alterations in microtubule dynamics by microtubule-targeting agents can cause abnormal attachment of chromosomes, impair kinetochore tension, and prevent anaphase onset and chromosome segregation (24,25). The net result is that the cell undergoes apoptosis (24,26). Despite more than 25 years of focused research, the exact mechanism of action of tubulin-binding agents on microtubules has not been identified. These agents bind to soluble tubulin and/or directly to tubulin in the microtubules. The abundant amount of tubulin in neurons and the role of microtubules in axonal transport are thought to explain the neurologic toxicity observed with most tubulin-binding agents in the clinic (27,28). Although most of the currently marketed antitubulin agents are antimitotic agents and inhibit cell proliferation by acting on the polymerization dynamics of spindle microtubules and altering proper spindle function, the specific effects of individual microtubuletargeted drug on microtubule polymer mass and on the stability and dynamics of the microtubules are much more complex than initially thought (10,24,26,29–31). Microtubule-targeted drugs have traditionally been classified into 2 main groups. The first group, denoted as “microtubuledestabilizing agents,” inhibits microtubule polymerization at high concentrations. Examples of this group include the Vinca alkaloids (vinblastine, vincristine, vinorelbine, vindesine and vinflunine), cryptophycins, halichondrins, estramustine, colchicines, and combretastatins (10,24,29). The second group is known as “microtubule-stabilizing agents” because at high concentrations these agents stimulate microtubule polymerization and stabilize the microtubule (i.e., result in microtubule “bundling”) (10,24,29,30,36). Examples include paclitaxel, docetaxel, eleutherobins, epothilones, laulimalide, sarcodictyins,


183

and discodermolide (24,29). However, this simplistic classification of drugs as microtubule “stabilizers” or “destabilizers” can lead to confusion because it implies that the influence of these drugs on microtubule polymer mass represents their most important action responsible for the anticancer activity. However, recent work has demonstrated that drugs that increase or decrease microtubule polymerization at high concentrations can dramatically suppress microtubule dynamics at 10–100-fold lower concentrations, and thus kinetically stabilize the microtubules without changing the microtubule-polymer mass (10,24,29,32–35). In fact, it now seems that the most important mechanism of action of these drugs is the suppression of spindlemicrotubule dynamics, which results in the slowing and/or blocking of mitosis at the metaphase-anaphase transition and induction of apoptotic cell death. Although these drugs appear to have a common mechanism of action, it should be pointed out that each one appears to bind to diverse sites on tubulin and they have different effects on microtubule dynamics. This variability in the modes of action of these agents might be responsible not only for the differences in efficacy and therapeutic windows, but also for the differing specificities of these drugs for different tumor types and for variations in their potential to cause neurotoxicity and other adverse events (29). The recent development of new techniques such as hydrogen/deuterium exchange coupled to liquid chromatography-electrospray ionization mass spectroscopy will permit investigators to more fully understand the molecular mode of action of these important drugs in cancer chemotherapy (31).

3. CLINICAL DEVELOPMENT AND APPROVED INDICATIONS 3.1. Naturally Derived Vinca Alkaloids The impetus for the clinical development of vincristine and vinblastine as antitumor agents was in great measure originated by the observation of single agent clinical antitumor activity in lymphoproliferative disorders, Wilms tumor’s and even in lung cancer (37–40). However, it took incorporation into chemotherapy regimens with other cytotoxic agents to best exploit their clinical benefit. It is of interest that despite the common origin and similarities in the chemical structure of vincristine and vinblastine, their spectrum of antitumor activity and toxicities are quite different.

184


Neurological toxicity is the cardinal undesirable effect of vincristine and empirically has resulted in a dose cap of 2 mg per week. The most usual effects are peripheral bilateral paresthesias (sensory impairment), but cumulative doses can result in neuritic pain, motor dysfunction, lost of deep tendon reflexes, autonomic effects and even paralysis. Vinblastine most commonly produces myelosuppression, especially neutropenia, and much less neurotoxicity than vincristine. Inflammation of the oro-pharyngeal mucosa is more common with vinblastine than vincristine. Although the reasons for the differences in the toxicity profile between these agents are not completely clear, the more extensive tissue binding and lower clearance of vincristine at the doses and schedule commonly used with these agents may explain some of these differences. In fact, prolonged infusions of vinblastine are associated with more neurologic effects. It is also plausible that variation in post-translational modifications of microtubule polymers in neurological tissues (41) may produce functional differences rendering these tissues more susceptible to the increased retention of vincristine. Both drugs are heavily metabolized in the liver with subsequent biliary excretion, thus dose adjustments are mandated in patients with liver dysfunction. Table 1 depicts the United States Food and Drug Administration (FDA) approved indications (42) for vincristine and vinblastine, as well as for other tubulin-interactive agents. Compendia listing for additional indications reimbursed by Medicare and major insurers (43,44) are also listed. It cannot be over-emphasized that both agents are still to this date, important ingredients of curative regimens in lymphoproliferative malignancies, germ cell tumors and pediatric malignancies (45–48).

3.2. Vindesine and Vinorelbine Vindesine is a synthetic derivative of vinblastine. It differs structurally from the parent compound mainly by a carboxamide group in place of a methyl ester (Fig. 1). Toxicity and activity appear very similar to vinblastine (49), and although used in European countries, it does not have an approved indication in the United States. Vinorelbine, another synthetic derivative of vinblastine, includes 2 tartaric acid residues in its structural formula (Fig. 1). The same as the other vinca alkaloids, the main toxicities are hematologic and neurologic. Neurotoxicity is less prominent than for vincristine, as it was predicted by preclinical studies, for which it demonstrated lower affinity for axonal microtubules than for mitotic spindle microtubules (50). The development of this agent focused on nonsmall cell lung and breast cancer,

185

Vinblastine Lymphoma: Hodgkin’s, lymphocytic, histiocytic, Mycosis fungoides Advanced carcinoma of the testes Kaposi’s sarcoma, Letterer-Siwe disease Choriocarcinoma Breast carcinoma

Vincristine Acute leukemia. In combination in: Hodgkin’s disease Non-Hodgkin’s malignant lymphomas Rhabdomyosarcoma Neuroblastoma Wilms’ tumor

FDA approved indications

Bladder Kidney Lung Melanoma Ovary (germ cell) Prostate

Brain Breast Cervical Chron. Lymph Leukemia Chron. Myel Leukemia Colorectal Cutaneous Lymphoma Ewing’s Sarcoma Kaposi’s Sarcoma ITP*

(Continued)

ITP Head and Neck Neuroblastoma Trophoblastic Neoplasm

Kidney Liver Lung Melanoma Multiple Myeloma Osteosarcoma Ovary (germ cell) Retinoblastoma Soft Tissue Sarcomas Trophoblastic disease Waldenstrom’s

Drug compendia uses

Table 1 Clinical Indications for Tubulin Interactive Agents

186

Vinorelbine Single agent or with cisplatin for the first-line treatment in advanced nonsmall cell lung cancer Docetaxel Breast cancer: metastatic, after failure of prior chemotherapy; adjuvant, in combination with doxorubicin and cyclophosphamide Nonsmall cell lung cancer: Locally advanced or metastatic after failing platinum-based chemo In combination with cisplatin for the treatment of of unresectable, chemo-naïve patients Prostate: with prednisone for androgen independent disease Gastric: with cisplatin and fluorouracil for chemo-naïve advanced disease. Head and Neck: with cisplatin and 5FU

FDA approved indications

Table 1 (Continued)

Bladder Esophageal Lung (small cell) Ovarian

Breast Ovary

Drug compendia uses Cervix

187

* Idiopathic Immune Thrombocytopenia

Breast cancer: -Adjuvant treatment of nodepositive disease, sequential to doxorubicin containing chemo; after failure of combination chemo for metastatic disease or relapse within 6 months of adjuvant. Nonsmall cell lung cancer: with cisplatin, for first-line in patients not candidates for surgery and/or radiation therapy. AIDS related Kaposi’s sarcoma, as second line

Paclitaxel Ovary: first-line, in combination with cisplatin Bladder Carcinoma of Unknown Primary Cervix Endometrial Esophageal Fallopian Tube Head & Neck Peritoneal Prostate Stomach Testes

188


Fig. 1. Chemical structure of the vinca alkaloids in clinical use.

although it also has demonstrated activity in ovarian and cervical cancer (42–44). In nonsmall cell lung cancer, in combination with cisplatin, vinorelbine have been shown to be superior to cisplatin-vindesine and to cisplatin alone, both in antitumor response rate and overall survival (51,52). Furthermore, the highest degree of evidence that adjuvant chemotherapy is able to increase the number of long term nonsmall cell lung cancer survivors after curative surgery has been accomplished with vinorelbine-cisplatin regimens (53,54). In breast cancer, single agent antitumor response rates for vinorelbine for chemo-naïve and previously treated patients range from 35 to 40% (55,56).

3.3. Paclitaxel The success of paclitaxel needs to be measured not only by its broad clinical antitumor activity, but also because (through its large profitability) it revolutionized corporate mentality and industry investment in the development of anticancer drugs. The biggest challenge in its early development was water solubility and the hypersensitivity reactions induced by its diluent, chremophor EL (polyoxyethylated castor oil). Incorporation of premedication permitted the shortening of the infusion to make it feasible to administer in the outpatient clinic. The shorter infusions revealed that at higher doses paclitaxel has nonlinear pharmacokinetics because of saturable distribution and saturable elimination. Hepatic cytochrome


189

P450 oxidases are largely responsible for the metabolism and clearance of paclitaxel (57). CYP2C is responsible for the formation of its principal metabolite 6 hydroxypaclitaxel, and CYP3A for the formation of minor metabolites. Paclitaxel demonstrated substantial antitumor activity in patients with relapsed ovarian cancer, both in 24 h (135–175 mg/m2 ) and 3 h (175 mg/m2 ) infusion schedules (58). This eventually led to its FDA approval for ovarian cancer utilizing either of the above 2 schedules. Prominent activity with repercussions in survival was also noted in combination regimens in first line ovarian cancer and nonsmall cell lung cancer (59–61). In breast cancer, its most common use has stemmed from the realization that consolidation with this drug adds to survival in the adjuvant setting, following administration of older established regimens (62,63). Most common toxicities of paclitaxel include: 1) hypersensitivity reactions, which are for the most part preventable with the preadministration of steroids and H1 and H2 blockers; 2) non cumulative neutropenia; 3) alopecia; 4) asymptomatic bradyarrythmias; 5) myalgias/arthralgias, and 6) neurotoxicity. The latter, is the most troublesome and difficult to manage toxicity of paclitaxel. Symmetrical distal loss of sensation carried by large (propioception, vibration) and small (temperature, pin prick) fibers occurs, as well as loss of deep tendon reflexes (64). Motor and autonomic dysfunctions may occur in patients with previous neuropathy because of diabetes and alcoholism. At usual doses, these symptoms appear after multiple treatments, but attempts to escalate doses with colony stimulating factor support have resulted in early severe peripheral and even central neuropathy (65,66). Nevertheless, despite its toxicity profile, the broad antitumor activity of paclitaxel, as a single or combination therapy, has made this agent, one of the most commonly prescribed antitumor agents ever; effectively setting the ground for modern chemotherapy and the development of agents to come.

3.4. Docetaxel The clinical development of docetaxel faced the same challenges that any analogs of a successful medication would. Superior activity, better safety profile, or both were necessary for this semisynthetic taxane-derivative of 10-deacetyl-baccatin III to find its rightful place in the clinic. Preclinical studies were encouraging in that despite sharing a similar binding site on tubulin as paclitaxel, its affinity was 1.9-fold higher

190


and assembly of microtubules needed a lower critical protein concentration (67). Docetaxel also was not completely cross resistant with paclitaxel (68). Docetaxel is slightly more water soluble and formulated with polysorbate 80 instead of cremophor EL. However, hypersensitivity reactions including flushing, back and chest pain can occur. Premedication with steroids with or without H1 or H2 blockers is able to manage this hypersensitivity, which in our experience appears milder than paclitaxel. Extensive tissue distribution, protein binding, and high clearance, are features shared with paclitaxel. Biliary excretion after P450 mediated metabolism is largely the most prominent route of excretion. CYP3A4 is the most predominant isoform. As it is true for paclitaxel, toxicities of docetaxel are scheduledependent, with higher rate of severe neutropenia than paclitaxel when both are administered at recommended doses, every 3 wk. Weekly administration for both drugs has lesser impact in myelosuppression, and neurophatic effects, which are dose-limiting for paclitaxel in this schedule, appear lower for docetaxel. However, fatigue/asthenia are more prominent and become the dose-limiting for docetaxel on the weekly schedule (42). Toxic effects idiosyncratic to docetaxel are fluid retention at cumulative doses, an erythematous rash in hands and forearms, palmo-plantar erythrodysesthesias, sclerosis of tear ducts and atrophy, soreness and brittleness of fingernails (69). Initial efficacy trials established prominent activity in breast cancer (70,71), edging paclitaxel in subsequent postapproval randomized clinical trials (72). As with paclitaxel, additional clinical value for this drug has been reported in the adjuvant breast cancer setting following doxorubicin and cyclophosphamide (73). Prospective randomized trials in nonsmall cell lung cancer established the clinical value of docetaxel in this disease showing superior survival compared to supportive care in previously treated patients and equivalence in combination to cisplatin to other commonly used doublets in chemo-naïve patients (74–76). Other approved indications include hormone refractory prostate cancer (77), gastric (78) and head and neck carcinomas (79).

4. TUMOR RESISTANCE TO TUBULIN INTERACTIVE AGENTS In vitro studies of cells exposed to tubulin interactive agents have suggested 2 major mechanisms of acquired resistance. The first implicate an amplified gene that codes for a transmembrane


191

P-glycoprotein, PGP (80–82). This protein functions as an efflux pump, binding substrate to a high affinity binding site. Hydrolysis of ATP at the binding site produces a conformational change that shifts the substrate to a lower affinity site, releasing the substrate into the outer leaflet of the membrane (83). Hydrolysis of ATP at a second binding site returns the protein to the conformation able to bind new drug. Despite some diversity in this protein, the mechanism of resistance that it mediates affects several types of drugs including: vinca alkaloids, taxanes, anthracyclines, epipodophyllotoxins, actinomycin D, and colchicine. Thus, this phenomenon has been termed multidrug resistance (MDR) and the amplified gene responsible for it is mdr1 (81–84). This gene is a member of a family of transporter genes named ATP-binding cassette (ABC) transporters, thus it is also referred as ABCB1. Amplification has been found constitutively in some human tissues, e.g., large bowel, kidney, brain, liver. ABCB1 amplification in these tissues plays an important role in preventing the absorption and penetration of toxins through the blood brain barrier, as well as increasing their hepatic and renal elimination. Its presence in tumors derived from these tissues is thought to mediate many instances of primary chemotherapy resistance. Although other ABC transporters like the ABCC family (MRP) and ABCG2 (MXR, BRCP) have been associated with drug resistance, the vinca alkaloids and the taxanes are poor substrates, so ABCB1 is still considered the major mechanism of efflux pump for these agents (82). Another protein, the major vault protein LRP (not an ABC transporter), has also been studied, but has less convincing evidence of its association to taxanes and vinca alkaloids resistance. PGP phenotype in vitro has been shown to be reversible with a number of drugs utilized in the clinic for other indications. The list includes antimalarials, calcium channel blockers and other antiarrhytmic agents, as well as immunosuppressive drugs like cyclosporine. Unfortunately, the high toxicity associated with these agents at levels that result in inhibition of PGP has made their use in association with chemotherapy impractical in the clinic. Nevertheless, the appeal of the concept has fueled interest in the development of more tolerable agents. To this end several second generation drugs have advanced to clinical trials: PSC 833 (valspodar), a derivative of cyclosporine; VX-710 (biricodar), which restores drug sensitivity to both MDR1 and MRP1- expressing cells; the PGP specific inhibitors with lesser affinity to CYP3A, XR9576 (tariquidar) and LY335979 (zosuquidar); and the oral non-CYP3A4-dependent R101933 and OC144-093.

192


Significant challenges faced clinical trials, even with the less toxic second generation agents. These included: 1) difficulties in the pretreatment identification of PGP overexpressing tumors and PGP quantification, thus inability to study the subgroup of patients most likely to benefit from this approach; 2) interactions with the clearance of the anticancer agents, thus the need for chemotherapy dose reductions; 3) lack of adequate in vivo correlates of the amount of PGP tumor inhibition achieved by these agents; and 4) complexity of widely used chemotherapy regimens in which non-PGP and PGP substrates are used concurrently. However, a few randomized trials of first generation agents have demonstrated significant differences in overall survival using a PGP inhibitor in AML, breast and lung cancer (85–87), but no differences in ovarian, myeloma and small cell lung cancer (88–90). The development of non-CYP-dependent agents, and incorporation of Sestamibi (a PGP substrate) imaging, as well as more sophisticated methods to quantify PGP in tumors, is anticipated to advance this field. Creativity in trial design, e.g., induction with PGP inhibitors before chemotherapy or oral therapy with poorly absorbed PGP substrate chemotherapy has also the potential of a positive impact. A second potential mechanism of resistance to tubulin interactive agents is alterations in and tubulin. Theoretically, either mutations at the binding sites or posttranslational modifications could result in drug resistance. Monzo and collaborators described tubulin mutations in about a third of 49 nonsmall cell lung cancer patients in exons 1 or 4 of the TUBB gene that encodes the most highly expressed isoform of tubulin (91). Exon 4 encodes more than half of the protein including the ATP binding site. The mutations were thought to affect the ability of the patients to respond to paclitaxel treatment and resulted in decreased median survival. However, subsequent investigation showed that TUBB mutations are very uncommon and that the reported mutations likely were an artifact of coamplification of pseudogenes. The observed improved outcome associations were attributed to tumor aneuploidy (92,93). More recently, attention has been focused on microtubule associated proteins, which theoretically can also affect the binding of tubulin interactive agents. These ongoing investigations include: 1) microtubuleassociated protein 4 (MAP-4), the major microtubule-associated protein in nonneuronal tissues, which promotes microtubule polymerization through binding to the COOH termini of - and - tubulin (94); 2) Stahmin, a soluble cytoplasmic protein that can bind to tubulin


193

dimers and stimulate microtubule catastrophes (95); and 3) the microtubule associated protein Tau, which binds to both the inner and outer surface of tubulin and may bind to the same inner-surface pocket as paclitaxel (96). Given the differences in the mechanism of action and tubulin binding sites of the vinca alkaloids compared to the taxanes, the presence or absence of these proteins would be expected to affect sensitivity to these agents differently. Because no inhibitors/activators of these proteins have been developed, the focus of current investigation is to evaluate them as predictors of drug sensitivity and potential tools to individualize treatment choice.

5. NOVEL TUBULIN INTERACTIVE AGENTS IN CLINICAL DEVELOPMENT The success of the vinca alkaloids and taxanes has encouraged the search of new chemical entities with similar mechanisms of action, yet with improved therapeutic index. Three major areas comprise these efforts: 1) compounds that overcome the problems of formulation and delivery of the taxanes; 2) compounds that can overcome multidrug resistance; and 3) compounds that bind microtubules at different sites than the taxanes. The most advanced compound is Abraxane (ABI-007), a nanometersized albumin-bound paclitaxel particle. This drug takes advantage of the ability of albumin to carry lipophilic molecules in humans and to preferentially deliver paclitaxel to tumors by biologically interacting with albumin receptors mediating drug transport (97). The synthetic solvents needed for paclitaxel (cremophor EL /ethanol) and docetaxel (polysorbate 80/ethanol) are not necessary for albuminbound compounds, thus the hypersensitivity reactions observed with paclitaxel and docetaxel do not occur following administration of Abraxane. In addition, the absence of the cremophor binding micelles results in a higher volume of distribution and higher clearance for nanoparticle albumin bound paclitaxel compared to paclitaxel formulated with cremophor (98). A phase III randomized trial was performed to compare the antitumor activity and impact of survival of every 3 wk Abraxane (260 mg/m2 ) to that of paclitaxel (175 mg/m2 ) in patients with metastatic breast cancer (99). A significantly higher response rate (33% vs. 19%) was observed for the Abraxane group as well as a significant longer time to tumor progression. At a median follow up of 103 wk, the

194


differences in overall survival had not reached statistical significance. No hypersensitivity reactions were observed in the abraxane arm, and grade 4 neutropenia was lower, suggesting a role for the paclitaxel solvent in this toxicity. However, the incidence of neuropathy was higher in the Abraxane arm. Based on this trial, Abraxane had received FDA approval in patients with metastatic breast cancer with disease progression after first line therapy. Despite this initial success in breast cancer, it is not clear however, if in tumors that require a higher dose of paclitaxel for efficacy, the suggested improved therapeutic index of Abraxane will be sustained. In fact, initial trials in lung cancer have been somewhat disappointing (100). Nevertheless, the approach is very intriguing and a series of other nanoparticle albumin bound paclitaxel formulations, which potentially have even better therapeutic index are undergoing early human investigation. Other agents in development that promise enhanced delivery of paclitaxel include CT-2103, a large macromolecule conjugate of paclitaxel and poly-L-glutamate (101); docosahexanoic acid (DHA)- paclitaxel, a conjugate of the fatty acid from human breast milk DHA to paclitaxel (102); and the vitamin E-based paclitaxel emulsion S-8184 (103). CT-2103 enters tumor cells by endocytosis and concentrate in tumor tissues after metabolic cleavage by lysosomal enzymes, whereas DHA-paclitaxel utilizes the increased fatty acid uptake characteristic of tumor tissue to enhance tumor targeting (101,102). Alpha tocopherol PEG succinate acts as a surfactant in S-8184 to enhance paclitaxel solubility and to increase the circulation times and AUC of paclitaxel (103). The second category of microtubule-interactive drugs in development includes agents that are less susceptible to multidrug resistance. The epothilones, macrolides initially isolated as cytotoxic metabolites from the myxobacterium Sorangium cellulosum, have led the charge in the clinic. Similar to paclitaxel and docetaxel, epothilones induce microtubule bundling, formation of multipolar spindles and mitotic arrest (104). The epothilones compete with paclitaxel for binding to microtubules, and cell lines selected for resistance to epothilones contain mutations in tubulin, very near to the taxane-binding site (105). Structure-activity studies showed that modifications at or near the C12-13 epoxide can affect microtubule-stabilizing activity. For example, a methyl group at position 12, the difference between the natural products epothilone A and B results in a doubling in tubulin polymerization induction (106).


195

S

S O

N

O OH

N

O

O OH

O

O OH Epothilone B (patupilone) S

Epothilone D (KOS-862) S

H N

N

O OH

O OH

N

O

O OH

O

O OH BMS-247550 (ixabepilone)

O OH

KOS-1584

Fig. 2. Epothilone Derivatives in Development.

Figure 2 depicts the chemical structure of epothilone B (patupilone) and derivatives undergoing clinical development. Note the oxygen in the 16-member ring; the lactone is the primary site of metabolic attack and once opened, the compound loses cytocidal activity. The synthetic ixabepilone (BMS-247550, aza-epothilone B) has oxygen in the ring replaced by a nitrogen; the resultant lactam is more resistant to ringopening. Because the epoxide group is highly reactive and thought to contribute to epothilone toxicity, desoxyepothilone B (epothilone D, KOS-862) has the epoxide at Carbon 12–13 reduced to a double bond in an effort to give this compound less toxicity. The second generation KOS-1584 makes only one alteration compared to KOS862: an additional reduced double bond at Carbon 9-10, flattening the 16-member ring. Although all studied epothilones have broad antitumor activity in vitro and in vivo, the small chemical changes discussed above result in very different compounds in the clinic. It is of interest that drug resistance mechanisms differ between the epothilones and the taxanes, with P-glycoprotein minimally affecting the cytotoxicity of

196


the epothilones, suggesting that epothilones may be more active than the taxanes in patients with tumors with high PGP expression (107). KOS-862 appears less affected by PGP among the first generation epothilones. In fact, the concentrations required for 50% growth inhibition (IC) measured in CCRF-CEM/VBL1000 cells (2,048-fold resistance to vinblastine) were 0.029, 0.092, 2.99, and 5.17 μM for KOS-862, desoxyepothilone F, ixabepilone, and paclitaxel, respectively. These values represent 4-, 33.5-, 1,423-, and 3,133-fold resistance, respectively, when compared with the corresponding IC in the parent [nonmultiple drug-resistant (MDR)] CCRF-CEM cells (108). Studies in mouse xenografts by the same investigators also show a higher efficacy for the desoxyepothilone compounds. In addition, a comparison of KOS-862 and the second generation KOS-1584 using the MV522 lung cancer line showed greater efficacy at a 4-fold lower dose and a greater duration of activity for KOS-1584 (personal communication, A. Hannah, June, 2006). In the clinic, most of the data has been generated with epothilone B and ixabepilone. Polyethylene glycol 300 is used as a solvent for epothilone B and ixabepilone is prepared on a cremophor-based formulation. As expected, hypersensitivity has been observed with the latter, thus premedication is required. Broad antitumor activity with responses in multiple tumor types has been observed in Phase I/II trials for both compounds, even in patients with previous exposure to taxanes or in p-glycoprotein producing tumors (107). However, the therapeutic index is narrow with frequent neutropenia, mucositis, neuropathy and diarrhea at active doses. Interestingly, toxicities of these 2 compounds differ, with more diarrhea, and less neuropathy and myelosuppression observed with epothilone B. Preferred schedules of administration are weekly for 3 wk followed by 1 wk of rest for epothilone B and daily for 5 d every 3 wk for ixabepilone. Pharmacokinetics appear linear for both compounds and microtubule bundle formation in peripheral mononuclear cells have been shown to correlate with ixabepilone AUC (109). Furthermore, microtubule bundle formation was observed even at 24 h after infusion in biopsies of treatment responsive chest wall metastases of a breast cancer patient (109). Other epothilones in phase I/II clinical trials include: 1) the water soluble epothilone B analog, BMS-310705; 2) KOS-862; and 3) KOS-1584. Antitumor responses have been observed with all these compounds (110–112). Diarrhea is dose-limiting for BMS-310705


197

(110), central neuropathy for KOS-862 (111) and yet to be defined for KOS-1584 (112). The third class of compounds in development includes agents that do not compete with the taxanes or the vinca alkaloids in the binding of tubulin. An alternative binding site to the vinca alkaloids and taxanes is the site used by colchicine. Although the toxicity of this agent at meaningful concentrations for microtubule assembly inhibition has prevented its development as an anticancer agent, several natural products have been found to bind to tubulin near or at the colchicine site. Examples are cornigerine, podophyllotoxin, steganicin and the combrestatins (113). A series of synthetic compounds targeting this site have followed and are undergoing in vivo or early clinical evaluation, along with compounds that interfere with tubulin polymerization by alkylating critical tubulin sulfhydryl groups near that site (114–116). Although it is early to judge how many of the agents in development will produce a significant impact in the clinic, the wide scope of these investigations and the scientific knowledge they have generated are quite admirable. Undoubtedly, the pioneers that participated in the discovery and development of the early compounds should be recognized for opening the interest in this field and for how their foresight have led to amelioration of symptoms, life prolongation and even cures in myriads of cancer patients.

REFERENCES 1. Downing K, Nogales E. Crystallographic structure of tubulin: Implications for dynamics and drug binding. Cell Structure and function. 1999;24: 269–275. 2. Margolis RL, Wilson L. Opposite end assembly and disassembly of microtubules at steady state in vitro. Cell 1978;13:1–8. 3. Mitchinson T, Kirschner MW. Dynamic instability of microtubule growth. Nature 312:237–242, 1984. 4. Horio T, Hotani H. visualization of the dynamic instability of individual microtubules by dark-field microscopy. Nature, 1986;321:605–607. 5. Belmont L, Hyman A, Sawin K, Mitchinson T. Real-time visualization of cell cycle dependent changes in microtubule dynamics in cytoplasmic extracts. J Cell Biol 1990;40:95–107. 6. Mitchinson T, Salmon E. Pleward kinetochore fiber movement occurs during both metaphase and anaphase A in newt lung cell mitosis. J Cell Biol 1992;119:569–582. 7. Carlier MF. Role of nucleotide hydrolysis in the polymerization of actin and tubulin. Cell Byophys 1988;12:105.

198


8. Valle RB, Bloom GS, Theurkauf WE. Microtubule associated Proteins: subunits of the cytomatrix. J Cell Biol 1984;99: 38s. 9. Oakley B, Akkari Y. -tubulin at ten: progress and prospects. Cell Structure and Function 1999;24: 365–372. 10. Jordan MA, Wilson L. Microtubules as a target for anticancer drugs. Nature Reviews Cancer 2004; 4:253–265. 11. Johnson IS, Wright HF, Svoboda GH. Experimental basis for clinical evaluation of anti-tumor principles derived from Vinca rosea Linn. J. Lab. Clin. Med. 1958; 54:830–837. 12. Noble RL, Beer CT, Cutts JH. Further biological activities of vincaleukoblastine: an alkaloid isolated from Vinca rosea (L.). Biochem. Pharmacol. 1958; 1:347–348. 13. Duffin J. Poisoning the spindle: serendipity and discovery of the antitumor properties of the vinca alkaloids (Part 1). Pharmacy in History 2002; 44(2):64–76. 14. Duffin J. Poisoning the spindle: serendipity and discovery of the antitumor properties of the vinca alkaloids (Part 2). Pharmacy in History 2002; 44(3):105–119. 15. Wall ME. Camptothecin and taxol: discovery to clinic. Med. Res. Rev. 1998 Sep; 18(5):299–314. 16. Wall, M.E., & Wani, M.C. Camptothecin and taxol: discovery to clinic– thirteenth Bruce F. Cain Memorial Award Lecture. Cancer Research 1995; 55(4):753–760. 17. Oberlies NH, Kroll DJ. Camptothecin and taxol: historic achievements in natural products research. J. Nat. Prod. 2004; 67(2), 129–135. 18. Cragg GM. Paclitaxel (Taxol® ): A success story with valuable lessons for natural product drug discovery and development. Med. Res. Rev. 1998 Sep; 18(5):315–331. 19. Wani MC, Taylor HL, Wall ME, Coggon P and McPhail AT. Plant antitumor agents. VI. The isolation and structure of taxol, a novel antileukemic and antitumor agent from Taxus brevifolia. J. Am. Chem. Soc.,1971; 93(9):2325–2327. 20. Schiff PB, Fant J, Horwitz SB. Promotion of microtubule assembly in vitro by taxol. Nature 1979; 277:665–667. 21. Horwitz SB. Personal recollections on the early development of taxol. J. Nat. Prod. 2004: 67(2):136–138. 22. Rowinsky EK, Cazenave LA, Donehower RC. Taxol: a novel investigational antimicrotubule agent. Journal of the National Cancer Institute 1990; 82(15):1247–1259. 23. McIntosh JR, Grishchuk EL, West RR. Chromosome-microtubule interactions during mitosis. Annual Review of Cell and Developmental Biology 2002; 18:193–219.


199

24. Zhou J, Giannakakou P. Targeting Microtubules for Cancer Chemotherapy. Curr. Med. Chem. – Anti-Cancer Agents 2005; 5:65–71. 25. Zhou J, Panda D, Landen JW, Wilson L, Joshi HC. Minor alteration of microtubule dynamics causes loss of tension across kinetochore pairs and activates the spindle checkpoint. J. Biol. Chem. 2002; 277(19):17200–17208. 26. Chen J-G, Yang CP, Cammer M, Horwitz SB. Gene Expression and Mitotic Exit Induced by Microtubule-Stabilizing Drugs. Cancer Res. 2003; 63: 7891–7899. 27. Dumontet C, Sikic BI. Mechanisms of action and resistance to antitubulin agents: microtubule dynamics, drug transport, and cell death. J. Clin. Oncol. 1999; 17(3):1061–1070. 28. Lee JJ, Swain SM. Peripheral neuropathy induced by microtubulestabilizing agents. J. Clin. Oncol. 2006; 24(10):1633–1644 (erratum 2401–2403). 29. Jordan MA. Mechanism of action of antitumor drugs that interact with microtubules and tubulin. Curr. Med. Chem. – Anti-cancer Agents 2002; 2:1–17. 30. Checchi PM, Nettles JH, Zhou J, Snyder JP, Joshi HC. Microtubuleinteracting drugs for cancer treatment. Trends in Pharmacol. Sci. 2003; 24(7):361–365. 31. Xiao H, Verdier-Pinard P, Fernandez-Fuentes N, Burd B, Angeletti R, Fiser A, Horowitz SB, Orr GA. Insights into the mechanism of microtubule stabilization by Taxol. Proc. National Aced. Sci. 2006; 103(27):10166–10173. 32. Derry WB, Wilson L, Jordan, M A. Low potency of taxol at microtubule minus ends: implications for its antimitotic and therapeutic mechanism. Cancer Res. 1998; 58(6): 1177–1184. 33. Yvon A-M, Wadsworth P, Jordan MA. Taxol suppresses dynamics of individual microtubules in living human tumor cells. Mol. Biol. Cell 1999; 10:947–949. 34. Jordan MA, Thrower D, Wilson L. Mechanism of inhibition of cell proliferation by Vinca alkaloids. Cancer Res. 1991; 51:2212–2222. 35. Derry WB, Wilson L, Jordan MA. Substoichiometric binding of taxol suppresses microtubule dynamics. Biochemistry 1995; 34:2203–2211. 36. Schiff PB, Horwitz SB. Taxol stabilizes microtubules in mouse fibroblast cells. Proceedings of the National Academy of Sciences 1980; 77(3): 1561–1565. 37. Evans AE, Farber S, Brunet S, Mariano PJ. Vincristine in the treatment of acute leukemia in children. Cancer. 16:1302–1306, 1963. 38. Macmahon HE, Bedizel M, Ellis CA. Vincristine (leurocristine) sulfate in the treatment of children with metastatic Wilms’ tumor.

200

39. 40.

41. 42. 43. 44. 45.

46.

47. 48.

49.

50.

51.

52.

53.

Villalona-Calero, Schaaf, and Turowski Pediatric Division, Southwest Cancer Chemotherapy Group. Pediatrics 32:880–887, 1963. Hill JM, Loeb E. Treatment of leukemia, lymphoma, and other malignant neoplasms with Vinblastine. Cancer Chemother Rep 15:41–61, 1961. Frei E 3rd, Franzino A, Shnider et al. The treatment of inoperable carcinoma of the lung by vinblastine sulphate: a preliminary report. Br J Dis Chest 56:64–69, 1962. Schultze E, Asai D, Bulinski J, Kirschner M. post-translational modifications and microtubule stability. J Cell Biol 105:2167, 1987. www.fda.gov/cder/cancer/druglistframe.htm. USP DI. Thomson Micromedex. 6200 S. Syracuse Way, Greenwood Village, CO 80111–4740. AHFS Drug Information. American Society of HealthSystem Pharmacists, Inc. 7272 Wisconsin Avenue, Bethesda, MD 20814. Rowe, JM, Buck, G, Burnett, AK, et al. Induction therapy for adults with acute lymphoblastic leukemia: results of more than 1500 patients from the international ALL trial: MRC UKALL XII/ECOG E2993. Blood 2005; 106:3760. Canellos GP, Anderson JR, Propert KJ et al. Chemotherapy of advanced Hodgkin’s disease with MOPP, ABVD, or MOPP alternating with ABVD. N Engl J Med 1992; 327:1478–1484. D’Angio GJ, Breslow N, Beckwith JB et al. Results of The Third National Wilms’ Tumor Study. Cancer 1989;64:349–360. Williams SD, Birch R, Einhorn LH et al. Treatment of disseminated germ-cell tumors with cisplatin, bleomycin, and either vinblastine or etoposide. N Engl J Med. 1987; 316:1435–1440. Kris MG, Gralla RJ, Kalman LA, et al. Randomized trial comparing vindesine plus cisplatin with vinblastine plus cisplatin in patients with non-small cell lung cancer, with an analysis of methods of response assessment. Cancer Treatment Rep 1985a 69(4):387–395. Binet S, Fellous A, Lataste H, et al. In situ analysis of the action of Navelbine or various types of microtubules using immunofluorescence. Semin Oncol 1989, 16 (sppl 4): 5. Wozniak AJ, Crowley JJ, Balcerzak SP et al. Randomized trial comparing cisplatin with cisplatin plus vinorelbine in the treatment of advanced non-small-cell lung cancer: a Southwest Oncology Group study. J Clin Oncol. 1998; 16:2459–2465. Le Chevalier T, Brisgand D, Soria JC et al. Long term analysis of survival in the European randomized trial comparing vinorelbine/cisplatin to vindesine/cisplatin and vinorelbine alone in advanced non-small cell lung cancer. Oncologist. 2001; 6 Suppl 1:8–11. Douillard JY, Rosell R, De Lena M, et al. Adjuvant vinorelbine plus cisplatin versus observation in patients with completely resected stage


54.

55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

201

IB-IIIA non-small-cell lung cancer (Adjuvant Navelbine International Trialist Association [ANITA]): a randomised controlled trial. Lancet Oncol. 2006 7(9):719–727. Winton T, Livingston R, Johnson D, et al. Vinorelbine plus cisplatin vs. observation in resected non-small-cell lung cancer. N Engl J Med. 2005 23;352(25):2589–2597. Weber B, Vogel C, Jones S et al. Intravenous vinorelbine (NAVELBINE) as first- and second-line therapy in advanced breast cancer. J Clin Oncol. 1995; 13:2722–2730. Fumoleau P, Delgado FM, Delozier T et al. Phase II trial of weekly intravenous vinorelbine in first-line advanced breast cancer chemotherapy. J Clin Oncol. 1993; 11:1245–1252. Cresteil T, Monsarrat B, Alvinerie P, et al. Taxol metabolism by human liver microsomes: identification of cytochrome p450 isoenzymes involved in its biotransformation. Cancer Res 1994, 54:386. Eisenhauer E, ten Bokkel Huinink W, Swenerton K, et al. EuropeanCanadian randomized trial of taxol in relapsed ovarian cancer: high vs low dose and long vs short infusion. J Clin Oncol 1994, 12:2654. Piccart, MJ, Bertelsen, K, James, K, et al. Randomized intergroup trial of cisplatin-paclitaxel versus cisplatin-cyclophosphamide in women with advancd epithelial ovarian cancer: three-year results. J Natl Cancer Inst 2000; 92:699. Muggia, FM, Braly, PS, Brady, MF, et al. Phase III randomized study of cisplatin versus paclitaxel versus cisplatin and paclitaxel in patients with suboptimal stage III or IV ovarian cancer: A Gynecologic Oncology Group study. J Clin Oncol 2000; 18:106. Bonomi P, Kim K, Fairclough D, et al. Comparison of survival and quality of life in advanced non-small-cell lung cancer patients treated with two dose levels of paclitaxel combined with cisplatin versus etoposide with cisplatin: results of an Eastern Cooperative Oncology Group trial. J Clin Oncol. 2000 Feb;18(3):623–631. Henderson IC, Berry DA, Demetri GD et al. Improved outcomes from adding sequential paclitaxel but not from escalating doxorubicin dose in an adjuvant chemotherapy regimen for patients with node-positive primary breast cancer. J Clin Oncol 2003; 21:976–983. Mamounas EP, Bryant J, Lembersky BC et al. Paclitaxel after doxorubicin plus cyclophosphamide as adjuvant chemotherapy for nodepositive breast cancer: results from NSABP-28. J Clin Oncol 2005; 23:2694–2702. Rowinsky EK, Donehower R. Antimicrotubule agents. In Cancer Chemotherapy and Biotherapy, second edition. Chabner B and Longo D editors. Lippincot-Raven, Philadelphia, 1996.

202


65. Nieto Y, Cagnoni PJ, Bearman SI, et al. Acute encephalopathy: a new toxicity associated with high-dose paclitaxel. Clin Cancer Res. 1999, 5(3):501–506. 66. Rowinsky EK. On pushing the outer edge of the outer edge of paclitaxel’s dosing envelope. Clin Cancer Res. 1999, 5(3):501–506. 67. Diaz JF, Andreu JM. Assembly of purified GDP-tubulin intro microtubules induced by taxol and taxotere; reversibility, ligand stoichiometry and competition. Biochemistry 1993, 32:2747. 68. Garcia P, Braguer D, Carles G, et al. Comparative effects of taxol and taxotere on different human carcinomas cell lines. Cancer Chemother Pharmacol 1994, 34:335. 69. Hainsworth J, Burris H, Erland J, et al. Phase I trial of docetaxel administered by weekly infusion in patients with advanced refractory cancer. J Clin Oncol 1998, 16(6): 2164–2168. 70. Ravdin PM, et al. Phase II trial of docetaxel in advanced anthracyclineresistant or anthracenedione-resistant breast cancer. J. Clin Oncol. 1995;13(12):2879–2885. 71. Valero V, et al. Phase II trial of docetaxel: a new, highly effective antineoplastic agent in the management of patients with anthracyclineresistant metastatic breast cancer. J Clin Oncol. 1995;13(12):2886–2894. 72. Jones SE, Erban J, Overmoyer B, et al Randomized phase III study of docetaxel compared with paclitaxel in metastatic breast cancer. J Clin Oncol. 2005 Aug 20;23(24):5542–5551. 73. Martin M, Pienkowsky T, Mackey J, et al. Phase III trial of adjuvant docetaxel for node positive breast cancer. N Engl J Med 2005;352(22):2302–2313. 74. Shepherd FA, Dancey J, Ramlau R, et al. Prospective randomized trial of docetaxel versus best supportive care in patients with non-small cell lung cancer previously treated with platinum-based chemotherapy. J Clin Oncol. 2000;18(10):2095–2103. 75. Fossella F, Pereira JR, von Pawell J, et al. Randomized, multinational, phase III study of docetaxel plus platinum combinations versus vinorelbine plus cisplatin for advanced non-small cell lung cancer. J Clin Oncol. 2003;21(16):3016–3024. 76. Schiller JH, Harrington D, Belani CP, et al. Comparison of four chemotherapy regimens for advanced non-small cell lung cancer. N Engl J Med 2002;346:92–98. 77. Tannock IF, de Witt R, Berry WR et al. Docetaxel plus prednisone or mitoxantrone plus prednisone for advanced prostate cancer. N Engl J Med 2004;351(15):1502–1512. 78. Cutsem VE, Moiseyenko VM, Tjulandin S et al. Phase III study of docetaxel and cisplatin plus fluorouracil compared with cisplatin and


79.

80. 81.

82. 83.

84.

85.

86.

87.

88.

89.

90.

203

fluorouracil as first-line therapy for advanced gastric cancer: a report of the V325 Study Group. J Clin Oncol 2006; 24:4991–4997. Remenar, E, et al. Tax 323. A randomized phase III multicenter trial of neoadjuvant docetaxel plus cisplatin and 5-flourouracil (TPF) versusneoadjuvant PF in patients with locally advanced unresectable squamous cell carcinoma of the head and neck (SCCHN): final analysis of EORTC 24971. Proc Am Soc Clin Oncol. 2006. Abstract 5516. Dano K. Active outward transport of daunomycin in resistant Ehrlich ascites tumor cells. Biochim Biophys Acta 1973; 323; 466–483. Juliano RL, Ling V. A surface glycoprotein modulating drug permeability in Chinese hamster ovary cell mutants. Biochim Biophys Acta 1976, 455: 152–162. Leonard GD, Fojo T, Bates SE. The role of ABC transporters in clinical practice. The Oncologist 2003; 8(5):411–424. Sauna ZE, Smith MM, Muller M, et al. The mechanism of action of multidrug-resistance-linked P glycoprotein. J Bioenerg Biomembr 2001, 33:481–491. Ueda K, Clark DP, Chen CJ, et al. The human multidrug resistance (mdr1) gene. cDNA cloning and transcription initiation. J Biol Chem 1987, 262:505–508. List AF, Kopecky KJ, Willman CL, et al. Benefit of cyclosporine modulation of drug resistance in patients with poor-risk acute myeloid leukemia: a Southwest Oncology Group study. Blood 2001, 98: 3212–3220. Belpomme D, Gauthier S, Pujade-Lauraine E, et al. Verapamil increases the survival of patients with anthracycline-resistance metastatic breast carcinoma. Ann Oncol 2000, 11: 1471–1476. Millward MJ, Cantwell BM, Munro NC, et al. Oral verapamil with chemotherapy for advanced non-small cell lung cancer: a randomized study. Br J Cancer 1993, 67: 1031–1035. Joly F, Mangioni C, Nicoletto M, et al. A phase 3 study of PSC 833 in combination with paclitaxel and carboplatin (PC-PSC) versus paclitaxel and carboplatin (PC) alone in patients with stage III epithelial ovarian cancer or primary cancer of the peritoneum. Proc Am Soc Clin Oncol 2002, 21: 202a. Sonneveld P, Suciu S, Wijermans P, et al. Cyclosporin A combined with vincristine, doxorubicin and dexamethasone (VAD) compared with VAD alone in patients with advanced refractory multiple myeloma; an EORTC-HOVON randomized phase III study (06914). Br J Haematolo 2001, 115; 895–902. Milroy R. A randomized clinical study of verapamil in addition to combination chemotherapy in small cell lung cancer. West of Scotland Lung Cancer Research Group and the Aberdeen Oncology Group. Br J Cancer 1993, 68: 813–818.

204


91. Monzo M, Rosell R, Sanchez J, et al. Paclitaxel resistance in non-small cell lung cancer associated with beta tubulin gene mutations. J Clin Oncol 1999, 17: 1786–1793. 92. Kelley M, Li S, Harpole D. Genetic analysis of the B-tubulin gene, TUBB, in non-small cell lung cancer. J Natl Cancer Inst 2001, 93: 1886–1888. 93. Sale S, Song R, Shen P, et al. conservation of the class 1 -tubulin gene in human populations and lack of mutations in lung cancers and paclitaxel-resistant ovarian cancers. Mol Cancer Therapeutics 2002, 1:215–225. 94. Zhang CC, Yang JM, Bash-Babula J, et al. DNA damage increases sensitivity to vinca alkaloids and decreases sensitivity to taxanes through p53-dependent repression of microtubule-associated protein 4. Cancer Res. 1999 59(15):3663–3670. 95. Alli E, Bash-Babula J, Yang J and Hait W. Effect of Stathmin on the Sensitivity to Antimicrotubule Drugs in Human Breast Cancer. Cancer Research 2002, 62, 6864–6869, 2002. 96. Wagner P, Wang B, Clark E, et al. Microtubule associated protein (MAP)-Tau, a novel mediator of paclitaxel sensitivity in vitro and in vivo. Cell Cycle 2005, 4: 1149–1152. 97. Desai N, Trieu V, Yao R, et al: Increased antitumor activity, intratumor paclitaxel concentrations, and endothelial cell transport of cremophorfree, albumin-bound paclitaxel, ABI-007, compared with cremophorbased paclitaxel. Clin Cancer Res. 2006 12(4):1317–1324. 98. Sparreboom A, Scripture CD, Tieu V, et al. Comparative preclinical and clinical pharmacokinetics of a cremophor-free, nanoparticle albuminbound paclitaxel (ABI-007) and paclitaxel formulated in Cremophor (Taxol). Clin Cancer Res. 2005 11(11):4136–4143. 99. Gradishar W, Tjulandin S, Davidson N, et al. Phase III trial of nanoparticle albumin-bound paclitaxel compared with polyethylated castor oilbased paclitaxel in women with breast cancer. J Clin Oncol. 2005, 23(31):7794–7803. 100. Green M, Manikhas G, Orlov S, et al. Abraxane, a novel Cremophor-free, albumin-bound particle form of paclitaxel for the treatment of advanced non-small-cell lung cancer. Ann Oncol. 2006 Aug;17(8):1263–1268. 101. Singer J, Baker B, De vries P, et al. Poly-(L)-glutamic acid-paclitaxel (CT-2103) [XYOTAX], a biodegradable polymeric drug conjugate: Characterization, preclinical pharmacology, and preliminary clinical data. Adv Exp Med Biol 2003 519: 81–99. 102. Bradley M, Webb N, Anthony F, et al. Tumor targetting by covalent conjugation of a natural fatty acid to paclitaxel. Clin Cancer Res 2001, 7:3229–3238.


205

103. Lundberg B, Risovic V, Ramaswamy M, et al. A lipophilic paclitaxel derivative incorporated in a lipid emulsion for parenteral administration. J Control Release 2003, 86:93–100. 104. Bollag DM, McQueney PA, Zhu J, et al. Epothilones, a new class of microtubule-stabilizing agents with a taxol-like mechanism of action. Cancer Res 1995, 55: 2325–2333. 105. Giannakakou P, Gussio R, Nogales E, et al. A common pharmacophore for epothilone and taxanes: molecular basis for drug resistance conferred by tubulin mutations in human cancer cells. Proc Natl Acad Sci USA 2000, 97:2904–2909. 106. Nicolaou K, Winssinger N, Pastor J, et al. Synthesis of epothilones A and B in solid and solution phase. Nature 1997, 387: 268–272. 107. Goodin S, Kane M, Rubin E. Epothilones; mechanism of action and biologic activity. J Clin Oncol 2004, 22: 2015–2025. 108. Chou T, O’Connor O, Tong W, et al. The synthesis, discovery, and development of a highly promising class of microtubule stabilization agents: curative effects of desoxyepothilones B and F against human tumor xenografts in nude mice. Proc Natl Acad Sci U S A. 2001 Jul 3; 98(14):8113–8118. 109. McDaid H, Mani S, Shen H, et al. Validation of the pharmacodynamics of BMS-247550, an analogue of epothilone B, during a phase I clinical study. Clin cancer Res 2002, 8:2035–2043. 110. Mekhail T, Chung C, Holden S, et al. Phase I trial of novel epothilone B analog BMS-310705 IV q 21 days. Proc Am Soc Clin Oncol 2003, 22, abstr 515 P–129. 111. Rizvi N, Villalona-Calero M, Lynch T, et al. A Phase II study of KOS862 (Epothilone D) as second line therapy in non-small cell lung cancer. Proc 11th World Conference on Lung Cancer, 2005: abstr. P-565. 112. Villalona-Calero M, Goel S Schaaf L, et al. First-in-Human Phase 1 Trial of a Novel Epothilone, KOS-1584. Proc. Am Soc Clinical Oncology 2006. abstr 2003 P-79. 113. Sackett D. Podophyllotoxin, steganacin and combretastatin: natural products that bind at the colchicine site of tubulin. Pharmacol Ther 1993, 59:163–228. 114. Yee K, Hagey A, Verstovsek S, et al. Phase 1 Study of ABT-751, a Novel Microtubule Inhibitor, in Patients with Refractory Hematologic Malignancies. Clin Cancer Res 2005, 11: 6615–6624. 115. Nguyen T, McGrath C, Hermone A, et al. A common pharmacophore for a diverse set of colchicine site inhibitors using a structure-based approach. J Med Chem 2005, 48:6107–6116. 116. Combeau C, Provost J, Lancelin F, et al. RPR112378 and RPR115781: two representatives of a new family of microtubule assembly inhibitors. Mol Pharmacol 2000, 57:553–563.

9

Kinesin Motor Inhibitors as Effective Anticancer Drugs Vasiliki Sarli and Athanassios Giannis CONTENTS Introduction Kinesins KSP Inhibitors Chemically Modified Antisense Oligonucleotides Conclusion

Abstract Mitotic kinesins are essential for the formation and function of the mitotic spindle during mitosis. The following pages describe some of the different roles of this class of enzymes on cell division and our current understanding of kinesin inhibitors in anticancer drug development. Key Words: Kinesins; antimitotics; human Eg5; inhibitors; cancer


207

208

Sarli and Giannis

1. INTRODUCTION Currently, there is an increasing interest in exploring drug candidates for cancer chemotherapy by blocking proteins exclusively involved in the spindle assembly and function. The spindle itself is a highly dynamic structure, mainly composed of microtubules, which organises and directs the chromosomes of the mother cell to divide equally between the 2 daughter cells. For the proper function of the spindle in a normal cell division, numerous proteins are involved in the spatial organisation of both microtubules and chromosomes, including motor proteins of the kinesin family. Deregulation of some mitotic kinesins can perturb the spindle orchestration and inhibit the cell cycle progression at mitosis. Several members of the kinesin family are potential targets for the discovery of novel ways to inhibit proliferation of tumour cells.

2. KINESINS Kinesins are microtubule-based proteins that participate in different transport systems, as well as in chromosomal and spindle movements during mitosis (1–4). They have a highly conserved, ∼ 340 amino acid motor domain, which contains the ATP-binding and microtubules binding sites. The attachment area with specific cargoes occurs outside the motor domain and shows few similarities between the different members. Kinesin superfamily (KIFs) consists of 45 proteins, which are able to convert the energy produced by ATP hydrolysis to mechanical force. They move step by step along the microtubules by repeating cycles of attachment, sliding and dissociation (5). Three major types of kinesins have been described depending on the location of the motor domain. The N-kinesins have the motor domain at the N-terminus, the M-kinesins at the middle of the molecule and C-kinesins at the COOH-terminal domain. The position of the motor determines the direction of their movement to minus or plus ends of microtubules, whereas the central motor domain in M-kinesins is responsible for microtubule depolymerisation. Mitosis itself can be divided into 5 key stages, named prophase, prometaphase, metaphase, anaphase and telophase (Fig. 1). During prophase chromatin condenses into chromosomes and big changes in microtubular network occur. The nuclear envelope breaks down and the mitotic spindle is formed with chromosomes attached to microtubules through their kinetochores, as the cell enters prometaphase. During metaphase the chromosomes align properly at the metaphase

Chapter 9 / Kinesin Motor Inhibitors as Effective Anticancer Drugs

209

Nucleus

G1

Microtubules

Prometaphase Prophase

Kinetochore

Centrosome

Midbody

Anaphase

Metaphase

Cytokinesis

Fig. 1. Different phases of cell division.

plate and when the spindle checkpoint is satisfied the cell enters anaphase by sister chromatids separation and migration towards the poles of the spindle. At the end of mitosis, the spindle elongates as the distance between poles increases and the cell is divided by cytokinesis. Gene disruption experiments have showed that kinesins play important roles in all the phases of mitosis and their depletion causes several mitotic defects such as monopolar spindle formation, chromosome misalignment, anaphase delay, and failure of cytokinesis. Recently the analysis of human motor proteins by RNA interference demonstrated the existence of at least 13 kinesins involved in mitosis and cytokinesis (Table 1) (6). Eg5 (Kinesin-5 family), Kif2A (Kinesin13 family) and KifC1 (Kinesin-13 family) have essential roles in the earliest stages of mitosis, responsible for bipolar spindle formation. Homo sapiens Eg5 (KSP) is a slow plus-end-directed motor of the Kinesin-5 family (BimC family) (7), localises along the interpolar spindle microtubules and spindle poles (8). This motor has attracted significant attention as drug target because of its central role in cell division. Eg5 inhibition results in mitotic arrest giving a characteristic monoastral phenotype (9) BimC kinesins have a catalytic domain to the N-terminus and form bipolar homotetramers (10,11), with 2 motor domains at each end of a rod (12). Members of this kinesin family can crosslink and slide antiparallel interpolar microtubules apart (13–15). On the other hand, HSET (KifC1) is a minus end-directed motor that

210

Sarli and Giannis Table 1 Mitotic Kinesins and Their Function

Kinesin

Family

Function

Kif14 Kif4A Kif4B KSP MKLP1 MKLP2 MPP1 CENP-E Kif18 Kid Kif2A KifC1

Kinesin-3 Kinesin-4 Kinesin-4 Kinesin-5 Kinesin-6 Kinesin-6 Kinesin-6 Kinesin-7 Kinesin-8 Kinesin-10 Kinesin-13 Kinesin-13

MCAK

Kinesin-13

chromosome congression and alignment midbody formation cytokinesis midbody formation cytokinesis centrosome separation spindle bipolarity midbody formation cytokinesis midbody formation cytokinesis cytokinesis chromosome congression and alignment chromosome congression and alignment chromosome congression and alignment spindle bipolarity spindle bipolarity chromosome congression and alignment chromosome congression and alignment

provide inward-directed forces oppositely oriented to the forces of the plus end directed motor Eg5 (16,17). Kif2A is a depolymerising enzyme that localises to centrosomes and spindle poles. Depletion of Kif2A inhibits cell cycle progression by the assembly of monopolar or asymmetrical bipolar spindles (18). Other kinesins are involved in attachment of microtubules to kinetochores, chromosome movement and alignment. For example MCAK (Kinesin-13 family) (19) and CENP-E (Kinesin-7 family) (20) are associated with kinetochores. Kinesin MCAK can interact with curved protofilaments at microtubule ends accelerating their depolymerisation process (21,22). MCAK is regulated and phosphorylated by Aurora B kinase and is required for chromosome congression and correction of improper kinetochore-microtubule attachments (23). The plus-enddirected motor CENP-E appears to be essential for kinetochore capture of spindle microtubules and congression of chromosomes to the metaphase plate and seems to be an attractive anticancer target (24,25). Inhibition of CENP-E function results in appearance of some unaligned chromosomes at metaphase and mitotic arrest with activation of the mitotic checkpoint proteins at the kinetochores (26). Moreover, CENPE has a CAAX box (where C is a cysteine, A is an aliphatic amino acid, and X is a methionine, glutamine, serine or threonine) and is


211

farnesylated. The presence of farnesyl transferase inhibitors alters the association between CENP-E and the microtubules and cause accumulation of cells before metaphase and the activation of G2/M checkpoint (27). Theoretically, this inhibition could result in the known antiproliferative effects of the FTIs in human tumors. In addition to the spindle- and kinetochore-associated kinesins there is a group of molecular motors, named chromokinesins, which associate with chromosome arms and are involved in chromosome movement during mitosis (28). All the chromokinesins belong to Kinesin-4 or Kinesin-10 subfamilies and contain a DNA-binding region. For example the chromokinesin Kid (Kinesin-10 family) that is a plus-end-directed motor is necessary for spindle morphogenesis (29), orientation of chromosome arms (30), chromosome oscillation and movement (31). Kif4A and Kif4B, 2 closely related motors (Kinesin 4 family), associate with chromosomes and are necessary for faithful chromosome segregation and regulation of the midzone formation and maintenance during cytokinesis (32). RNA interference depletion of Kif4A resulted in defective prometaphase organisation, chromosome misalignment, spindle defects, chromosome missegregation and cytokinesis failures (33). Although human kinesin Kif14 (Kinesin-3 family) was at first reported to be involved in chromosome congression, recent studies show that is important in cell abscission (34). Further insight on the role of Kif14 has come from the finding that Kif14 interacts with PCR1 (protein-regulating cytokinesis 1) and citron kinase and is required for the process of cytokinesis (35). Kif14 is overexpressed in some human tumors and is identified as a candidate oncogene (36). Further studies on the function of this motor and the connection between its elevated expression and oncogenesis are necessary for the evaluation of Kif14 as an anticancer target. Cytokinesis failures are also caused by the depletion of the 2 closely related kinesins MKLP1 (37) and MKLP2 (38). These motors belong to the Kinesin-6 family and are spindle midzone- or midbody-associated proteins. MKLP1 is crucial for the formation of midboby matrix and MKLP2 is essential for the late stages of cytokinesis (39). A third member of this family MPP1 is a slow plus-end-directed motor that localises to the spindle midzone (40). It is involved in midzone microtubule bundling and microtubule sliding. MPP1 is required for proper progression of cytokinesis and successful cell cleavage in human cells.

212

Sarli and Giannis

3. KSP INHIBITORS Several reports have described the modulation of KSP activity by small molecule inhibitors. High throughput screening approaches yielded promising chemical diverse compounds, many of which show high antiproliferative activity. Interestingly, the first MCAK inhibitors were recently identified (41). Inhibitors for other mitotic motors have not been reported so far and their discovery using phenotypic screenings seems to be a difficult task, probably owning to functional redundancy between different kinesins (42). Furthermore, monoester formation is not an exclusively property of Eg5 inhibitors (43). In this chapter we emphasise to the potential use of KSP inhibitors as novel anticancer agents. The reader is referred recent reviews for a more detailed discussion on this subject (44,45).

3.1. Monastrol Monastrol 1 is the first characterised small molecule inhibitor of Eg5 (Scheme 1) (46). This compound was identified in a phenotype-based screen and arrests cells at G2/M phase leading to monoastral spindles. After its discovery, monastrol became a prototype compound in understanding the spindle assembly mechanisms in mitotic progression (47). S-monastrol is more potent inhibitor than the R-enantiomer in vitro and in vivo with an IC50 value of 14 μM. Monastrol inhibits reversibly Eg5 and ATP hydrolysis both alone and in the presence of microtubules (48,49). Remarkably, it does not target to the nucleotide pocket of Eg5, but is an allosteric inhibitor that binds to an induced-fit pocket 12 Å away from the catalytic centre of the enzyme. Crystallographic models of KSP with monastrol and Mg2+ . ADP, show that it induces a variety of conformational changes in the motor (50). These include a folding of loop L5 to form an induced-fit cavity and reorientation of the neck-linker/switch-2 cluster. In monastrol treated cells the mitotic arrest-deficient protein 2 (MAD2) localises to kinetochores, indicating the activation of the spindle assembly checkpoint. The spindle assembly checkpoint is the mechanism that monitors the mitotic spindle and the proper attachment of chromosomes to spindle microtubules. Incomplete kinetochoremicrotubule attachments induce a signal for anaphase delay to prevent errors in chromosome segregation. The activation of spindle checkpoint is a common chemotherapeutic strategy also employed by the known antimicrotubule drugs like taxanes and vinca alkaloids.

Chapter 9 / Kinesin Motor Inhibitors as Effective Anticancer Drugs OH

OH

O

R R

S

N H

F

OH NH

NH

1:Monastrol

OH

OH

O

O

N H

213

NH

NH

S

N H

N H

S

S

5: VS-83

2: R = H, Enastron 4: Enastrol 3: R = CH3, Dimethylenastron

H O

N O

OH

O

N N

N

N O

H

N H

N

Cl

O H 8: Terpendole E

Cl

N

NH2

OH

6: CK0106023 7: Ispinesib

Br

F

F

O N

N R

N

N H

NH2

S

O

HO OH

F F N

OH

O

N

O

H N 15

O N

16

14: KSP-IA

13: S-trityl-L-cysteine

OH

N

NH2

O

9: R = (CH2)3CH3, HR22C16 10: R = (CH2)4CH2NH2 11: R = (CH2)2CH3, HR22C16-A1 12: R = CH2Ph

N

O

CH2C6H5

N

Cl

N

O

S N N OH H

H2N 17

18

Scheme 1. Eg5 inhibitors discussed in the text.

Like antimicrotubule drugs, long-term monastrol-induced mitotic arrest can activate the apoptotic machinery, which eventually can lead to cell death. Such effects of monastrol compared to taxol were studied on AGS and HT29 cell lines from gastric and colon carcinomas (51). The 2 cell division inhibitors induce mitotic arrest by attacking the mitotic spindle with different mechanism and seem to trigger different apoptotic pathways, which can be cell specific. In both cell lines

214

Sarli and Giannis

monastrol inhibits cell proliferation and induces apoptosis through mitochondrial membrane depolarisation. However, the AGS cells were found to be more sensitive to monastrol effect. Neuropathy is the most serious side effect of the currently available antimitotic agents, attributed to non-specific specific interaction with microtubules. These drugs alter the microtubule dynamics and interfere with other cellular processes, including intracellular transport and organelle positioning. Eg5 and other mitotic kinesins are also expressed in terminally postmitotic neurons and developing brains, but the roles of these motors are still elusive. There is hope that anti-kinesin drugs will cause less toxic effects than the microtubule-directed agents, and one important question that needs to be answered is the effect of kinesin inhibitors in neurons. Baas and coworkers shed some light on this issue and investigated the effect of monastrol in cultured post mitotic neurons (52). These studies showed that exposure of sensory or sympathetic neurons to monastrol for a few hours had a positive short-term effect and increased the rate of axonal growth. Over longer exposure times there is no indication of toxicity of monastrol on the neurons. The effect of monastrol on the growth of dendrites and axons was also examined in developing neurons (53). Inhibition of Eg5 induced a distinctive growth profile of dendrites and axons. The modest toxic effects on the neurons over time compared with the deleterious effect of the known antimicrotubule drugs are promising for the application of anti-kinesin drugs in cancer treatment. Another important limitation of the efficacy of cytostatic agents in cancer chemotherapy is the development of multidrug resistance. One major cause of multidrug resistance is the over-expression of ATP-binding cassette transporters termed MDR proteins, such as the P-glycoprotein. P-gp, localised on the plasma membrane of resistant cancer cells, can bind and transport the antitumor drugs and, hence can influence the absorption and bioavailability of these drugs (54). The interaction of monastrol with P-glycoprotein was recently investigated (55). These studies demonstrated that monastrol is a very weak inhibitor of Pgp and it weakly induces MDR1 mRNA in LS180 cells. However, this effect did not result in increased Pgp function and monastrol appears not to be transported by Pgp. This is further supported by the similar growth-inhibiting potency of the drug between native and Pgp over-expressing cell lines.

3.2. Monastrol Analogues In an attempt to develop more potent and specific inhibitors of KSP based on monastrol scaffold, a small library of 40 compounds,


215

was synthesised and tested for their inhibitory activity against Eg5 using an in vitro steady-state ATPase assay (56). Three of these compounds i.e., enastron 2 (IC50 = 2 μM), enastrol 4 (IC50 = 2 μM) and dimethylenastron 3 (IC50 = 200 nM) proved to be potent inhibitors and arrested cultured human cervical carcinoma cells in G2/M phase with the formation of monoastral spindles (Scheme 1). Furthermore, 3,4-dihydrophenylquinazoline-2(1H)-thiones such as VS83 5 (IC50 =1.2 μM) have been also reported to display inhibitory effects against Eg5 (57). Enastron, dimethylenastron, and VS-83 display high antiproliferative activity on human glioblastoma cells (58). In contrast with paclitaxel, they induced no cytocidal effect on quiescent cells. Furthermore, they are neither modulators nor substrates of p-glycoprotein (Pgp). Additional studies on the effect of dimethylenastron in human multiple myeloma cells showed that suppression of Eg5 induces apoptosis and upregulates Hsp70 through the phosphatidylinositol 3-kinase (PI3K/Akt) pathway (59). Upregulation of Hsp70 is cytoprotective and Hsp70 can bind or antagonise the function or several proapoptotic proteins, such as apoptosis-inducing factor (AIF), Apaf-1 and apoptosis signal-regulating kinase 1 (ASK1). In this study it was found that the FTI277, a farnesyltransferase inhibitor interacts synergistically with dimethylenastron in inducing apoptosis through disrupting the Akt/Hsp70 signaling axis. These findings provide the first evidence for Eg5 inhibitor activity in hematologic malignancy and identify Hsp70 up-regulation as a critical mechanism responsible for modulating myeloma cell sensitivity to Eg5 inhibitors. In addition, these findings suggest that a combination of Eg5 inhibitors with agents abrogating Hsp70 induction would be more useful for myeloma therapy.

3.3. Quinazolinones Another class of KSP inhibitors is based on quinazolinone core. CK0106023 6 (Scheme 1) is a potent and allosteric inhibitor of KSP motor domain function. Among 5 kinesins tested including MKLP1, Kif1A, KHC and CENP-E, CK0106023 was found to be specific for KSP (60). The R-enantiomer of CK0106023 is >1,000-fold more potent to inhibit KSP ATPase activity and >100-fold more effective at cell growth inhibition than the S-enantiomer. The mean growth inhibitory activity of CK0106023 toward a variety of human tumor cell lines was 364 nM, including the multidrug resistant lines NCI/ADR-RES, HCT-15, and A2780ADR. In tumor-bearing mice CK0106023 showed

216

Sarli and Giannis

antitumor activity comparable to paclitaxel, with the formation of monopolar spindles, similar to monoasters in CK0106023-treated cells. A member of this chemical class of inhibitors SB-715992 7 (Ispinesib) demonstrated broad antitumor activity in murine tumor models is currently in phase II of clinical trials (61). Recently the effect of SB-715992 in PC-3 human prostate cancer cells was investigated (62). These studies revealed that SB-715992 inhibited cell proliferation and induced apoptotic cell death. From gene expression profiles altered by SB-715992, it was found that SB-715992 increased the expression of cyclin-dependent kinase inhibitors including p27KIP1 , p15 , and p57Kip2 that arrest the cell cycle and decreased the expression of genes such as FGF (fibroblast growth factor) and EGFR (epidermal growth factor) that are responsible for cell survival and cell proliferation. Additionally, it was shown that genistein, a broad-spectrum protein tyrosine kinase inhibitor enhanced the induction of apoptosis by SB-715992 in PC-3 cells, suggesting that genistein could be used in combination with SB-715992 for better therapeutic effects for the treatment of prostate cancer.

3.4. Indole Derivatives Terpendole E 8 is a natural product isolated from the fermentation broth of a soil-isolated fungus and arrests cells at M phase through inhibition of Eg5 (Scheme 1) (63). TerE inhibits the microtubulestimulated ATPase activity of Eg5 with an IC50 value of 23 μM, but it does not inhibit conventional kinesins. Another indole-based inhibitor of Eg5 is HR22C16 9 (IC50 = 800 ±10nM), which was identified from a forward-chemical-genetic screen of 16000 compounds (Scheme 1) (64). A solid phase solid synthesis of HR22C16 analogues provided a small library of 50 compounds, which were tested for their ability to inhibit Eg5. One derivative, HR22C16 10 (Scheme 1) inhibited Eg5 motility with an IC value of 90 nM. In these studies photocaging of HR22C16 was also reported for the examination of Eg5 function with increased temporal precision. Furthermore, the efficacy of HR22C16 and its analogues in Taxolresistant and –sensitive human ovarian carcinoma cells, which have PgP overexpression or acquired -tubulin mutations, was evaluated (65). It was shown that HR22C16-A1 11 (Scheme 1) inhibits cell survival and retains its efficacy in PgP-overexpressing cells, suggesting that it is not a PgP substrate. Additionally, this compound induces cell death through the intrinsic apoptotic pathway with a dose-dependent increase in both caspace-9 and PARP cleavage. Further studies on the importance of stereochemistry in HR22C16 derivatives showed that all


217

4 diastereomers of HR22C16 are active against Eg5 (66). From a small library of 60 tetrahydro- -carbolines compound 12 with a N-Benzyl side chain was identified also as a specific and potent inhibitor Eg5 with IC50 = 0.65 μM.

3.5. S-Trityl-l-Cysteine S-trityl-l-cysteine 13 (Scheme 1) is a potent tumor growth inhibitor in the NCI 60 tumor cell line screen that targets human Eg5 (67). It was identified from an in vitro assay screening of small-molecule libraries of a total 2,869 compounds and inhibits Eg5 with IC50 of 1.0 μM for the inhibition of basal ATPase activity and 140 nM for the microtubule-activated ATPase activity. It causes mitotic arrest in HeLa cells with an IC50 = 1.0 μM and shows an average of inhibition of growth in 60 different tumor cell lines with GI = 1.3 μM. Among 9 different human kinesins tested, S-trityl-l-cysteine specifically inhibits Eg5 (68). Both S- and R-enantiomers of this unnatural amino acid are inhibiting Eg5 with nearly equal potencies. Hydrogen-deuterium (H/D) exchange mass spectroscopy and directed mutagenesis showed that S-trityl-l-cysteine and monastrol bind to the same region of Eg5 and cause similar conformational changes within the interaction site (69). Recently, the ´´key residues´´ crucial for inhibition by monastrol and S-trityl-l-cysteine in the binding pocket of Eg5 were investigated (70). Eleven residues in the binding cavity were mutated and the effects were monitored by kinetic analysis and mass spectroscopy. Of the mutations tested, Leu214 (helix 3) was identified as critical for inhibition by both monastrol and S-trityl-l-cysteine. This study demonstrates that single point mutations in the common binding pocket on Eg5 can prevent the binding of inhibitors and abolish their inhibitory effects. Such resistance mechanism has just been shown for ispinesib in HCT116 colorectal tumor cell lines and implicates 2 distinct point mutations in amino acid substitutions, D130V and A133D in loop L5. It is now well established that genomic instability is an important factor in the development of drug resistance and further investigations on the identification of resistant KSP mutants in different cancer lines are necessary. Possibly, KSP inhibitors from different chemical classes that will interact with different residues on the KSP binding cavity might be used to overcome this type of resistance.

3.6. Dihydropyrrole Derivatives Recently Tao and colleagues performed a high-throughput screen and identified KSP-IA 14 (Scheme 1) as a potent (IC50 = 11 nM)

218

Sarli and Giannis

and specific Eg5 inhibitor (71). This study offered the first insights to the mechanism of inducing cancer cell apoptosis by a kinesin inhibitor. In cultured A2780 ovarian carcinoma cells KSP-IA induced monoastral spindles and activated the spindle assembly checkpoint as determined by detection of the phosphorylation of the spindle checkpoint component BubR1. Similar results were obtained in spindle checkpoint-competent HCT116 cells. Prolonged exposure to KSPIA activates the spindle checkpoint and induces caspace-dependent apoptosis. This apoptosis induction is coupled with mitotic slippage, which is the process that cells exit mitosis to form tetraploid cells in a pseudo G1 phase. The apoptotic mechanism is initiated by activation of the pro-apoptotic protein Bax of the Bcl-2 family. On the other hand, the slippage-refractory colon cancer cell line HT29 was resistant to KSP-IA-induced of apoptosis. Sequential suppression of Cdk1 with purvananol A (Purv) facilitates the exit from mitosis and synergies with the KSP inhibitor in inducing cell death. These results indicate that activation of the spindle checkpoint and subsequent mitotic slippage increases the lethality to KSP inhibitors. A better understanding of the cell signaling mechanisms that lead to apoptosis after long-term mitotic arrest is clearly necessary for the clinical development of these inhibitors and the improvement of their clinical efficacy. Recently, a detailed study on modifications of dihydropyrrole lead structure and optimization efforts was published. Potency and aqueous solubility were improved through introduction of basic amides and ureas moieties to the dihydropyrrole scaffold, and provided inhibitors that cause mitotic arrest in A2780 human ovarian carcinoma cells with EC50 < 10nM (72).

3.7. Other Inhibitors Based on high-throughput screen, 3,5-diaryl-4,5-dihydropyrazole derivatives were identified as potent and selective Eg5 inhibitors that bind to the same allosteric pocket of KSP as monastrol (73). It was shown that dihydropyrazole 15 (Scheme 1) is the most potent inhibitor in this study with an IC50 value of 26 nM and causes apoptotic cell death through caspase-3 activation in A2780 human ovarian carcinoma cells with an IC50 = 15 nM. In another high-throughput effort the 4-phenyl-tetrahydroisoquinoline scaffold was identified to give access to potent inhibitors of human Eg5 (74). Compound 16 (Scheme 1) was identified as the most potent inhibitor with an IC50 value of 104 nM. The X-Ray crystal structure of 16 in complex with Mg2+ and ADP showed that it binds to the known


219

allosteric pocket of Eg5. The inhibitor was also tested for its cytotoxic activity in a panel of tumor cell lines and revealed antiproliferative activity against all the cell lines tested, including both A2780S taxolsensitive (IC50 = 0.267 μM) and A2780-TAXR taxol-resistant (IC50 = 0.215 μM) cell lines. Synthesis and SAR of substituted pyrrolotriazine-4-one analogues as Eg5 inhibitors has also been reported (75). These compounds bind to the same allosteric site on the Eg5 protein as monastrol and other inhibitors. Inhibitor 17 shows high cytotoxicity in A2780 human ovarian carcinoma cells and in vivo antitumor activity in the iv P388 murine leukaemia model. More recently phenothiazine 18 was reported to inhibit the microtubule-stimulated ATPase activity of Eg5 with an IC50 value of 1.52 μM and showed strong toxicity in the transformed WI38VA13, HL-60 and HeLa cell lines (76).

4. CHEMICALLY MODIFIED ANTISENSE OLIGONUCLEOTIDES A library of 2 -methoxy-modified antisense oligonucleotides (2 MOE ASO) targeting 1,510 different genes has been used to identify cell cycle regulatory genes through G2/M phase (77). One of the most effective ASOs targeted Eg5 and was effective against a variety of tumor cells (lung, bladder, pancreas, prostate, cervix and liver) with monoastral phenotypes. The ASO-mediated Eg5 inhibition induced mitotic arrest, increased apoptosis and inhibited cell proliferation in U87-MG and HeLa cells. In vivo experiments showed that treatment with Eg5 ASO significantly reduced growth of U87-MG and MDAMB-231 tumor xenografts and Eg5 expression with no ASO-mediated toxicities. Although antisense oligonucleotides have been successfully used to screen for gene function in high-throughput cell based assays and validate gene targets in vivo, their therapeutic utility remains to be shown.

5. CONCLUSION Mitotic kinesins are key enzymes in mitotic progression and their inhibition represents a new approach to attack the mitotic spindle. Especially, kinesin KSP has been attracted particular attention and has been regarded as a validated mitotic cancer target. Small molecule inhibitors of KSP are currently evaluated in clinical trials and early results show a promising anti-cancer activity and acceptable toxicity profiles. These drug candidates may represent the next generation

220

Sarli and Giannis

of anti-mitotic cancer drugs offering an important alternative for the treatment of cancer and other proliferative diseases such as diabetic retinopathy, restenosis, pulmonary and liver fibrosis, Sjogren´s, lupus erythematosis, lymphoproliferative disorders, etc.

REFERENCES 1. Wittmann T, Hyman A, Desai A. The spindle: a dynamic assembly of microtubules and motors. Nat Cell Biol 2001;3:E28–34. 2. Vale RD. The molecular motor toolbox for intracellular transport. Cell 2003;112:467–480. 3. Hirokawa N, Takemura R. Kinesin superfamily proteins and their various functions and dynamics. Exp Cell Res 2004;301:50–59. 4. Sharp DJ, Rogers GC, Scholey JM. Microtubule motors in mitosis. Nature 2000;407:41–47. 5. Yildiz A, Selvin PR. Kinesin: walking, crawling or sliding along? Trends Cell Biol 2005;15:112–120. 6. Zhu C, Zhao J, Bibikova M, Leverson JD, Bossy-Wetzel E, Fan JB, Abraham RT, Jiang W. Functional analysis of human microtubule-based motor proteins, the kinesins and dyneins, in mitosis/cytokinesis using RNA interference. Mol Biol Cell 2005;16:3187–3199. 7. Lawrence CJ, Dawe RK, Christie KR, Cleveland DW, Dawson SC, Endow SA, Goldstein LS, Goodson HV, Hirokawa N, Howard J et al. A standardized kinesin nomenclature. J Cell Biol 2004;167: 19–22. 8. Cochran JC, Sontag CA, Maliga Z, Kapoor TM, Correia JJ, Gilbert SP. Mechanistic analysis of the mitotic kinesin Eg5. J Biol Chem 2004;279:38861–38870. 9. Koller E, Propp S, Zhang H, Zhao C, Xiao X, Chang M, Hirsch SA, Shepard PJ, Koo S, Murphy C, Glazer RI, Dean NM. Use of a chemically modified antisense oligonucleotide library to identify and validate Eg5 (kinesin-like 1) as a target for antineoplastic drug development. Cancer Res 2006;66:2059–2066. 10. Kashina AS, Baskin RJ, Cole DG, Wedman KP, Saxton WM, Scholey JM. A bipolar kinesin. Nature 1996;379:270–272. 11. Turner J, Anderson R, Guo J, Beraud C, Fletterick R, Sakowicz R. Crystal structure of the mitotic spindle kinesin Eg5 reveals a novel conformation of the neck-linker. J Biol Chem 2001;276:25496–25502. 12. Kapoor TM, Mitchison TJ. Eg5 is static in bipolar spindles relative to tubulin: evidence for a static spindle matrix. J Cell Biol 2001;154: 1125–1133.


221

13. Kapitein LC, Peterman EJ, Kwok BH, Kim JH, Kapoor TM, Schmidt CF. The bipolar mitotic kinesin Eg5 moves on both microtubules that it crosslinks. Nature 2005;435:114–118. 14. Cameron LA, Yang G, Cimini D, Cancan JC, Kisurina-Evgenieva O, Khodjakov A, Danuser G, Salmon ED. Kinesin 5-independent poleward flux of kinetochore microtubules in PtK1 cells. J Cell Biol 2006;173: 173–179. 15. Valentine MT, Fordyce PM, Krzysiak TC, Gilbert SP, Block SM. Individual dimers of the mitotic kinesin motor Eg5 step processively and support substantial loads in vitro. Nat Cell Biol 2006;8:470–476. 16. Mountain V, Simerly C, Howard L, Ando A, Schatten G, Compton DA. The kinesin-related protein, HSET, opposes the activity of Eg5 and cross-links microtubules in the mammalian mitotic spindle. J Cell Biol 1999;147:351–365. 17. DeLuca JG, Newton CN, Himes RH, Jordan MA, Wilson L. Purification and characterization of native conventional kinesin, HSET, and CENP-E from mitotic hela cells. J Biol Chem 2001;276:28014–28021. 18. Ganem NJ, Compton DA. The KinI kinesin Kif2a is required for bipolar spindle assembly through a functional relationship with MCAK. J Cell Biol 2004;166:473–478. 19. Wordeman L, Mitchison TJ. Identification and partial characterization of mitotic centromere-associated kinesin, a kinesin-related protein that associates with centromeres during mitosis. J Cell Biol 1995;128:95–104. 20. Yucel JK, Marszalek JD, McIntosh JR, Goldstein LS, Cleveland DW, Philp AV. CENP-meta, an essential kinetochore kinesin required for the maintenance of metaphase chromosome alignment in Drosophila. J Cell Biol 2000;150:1–11. 21. Moore A, Wordeman L. The mechanism, function and regulation of depolymerizing kinesins during mitosis. Trends Cell Biol 2004;14, 537–546. 22. Helenius J, Brouhard G, Kalaidzidis Y, Diez S, Howard J. The depolymerizing kinesin MCAK uses lattice diffusion to rapidly target microtubule ends. Nature 2006;441:115–119. 23. Lan W, Zhang X, Kline-Smith SL, Rosasco SE, Barrett-Wilt GA, Shabanowitz J, Hunt DF, Walczak CE, Stukenberg PT. Aurora B phosphorylates centromeric MCAK and regulates its localization and microtubule depolymerization activity. Curr Biol 2004;14:273–286. 24. Schaar BT, Chan GK, Maddox P, Salmon ED, Yen TJ. CENP-E function at kinetochores is essential for chromosome alignment. J Cell Biol 1997;139:1373–1382. 25. Kapoor TM, Lampson MA, Hergert P, Cameron L, Cimini D, Salmon ED, McEwen BF, Khodjakov A. Chromosomes can congress to the metaphase plate before biorientation. Science 2006;311:388–391.

222

Sarli and Giannis

26. Tanudji M, Shoemaker J, L’Italien L, Russell L, Chin G, Schebye XM. Gene silencing of CENP-E by small interfering RNA in HeLa cells leads to missegregation of chromosomes after a mitotic delay. Mol Biol Cell 2004;15:3771–3781. 27. Ashar HR, James L, Gray K, Carr D, Black S, Armstrong L, Bishop WR, Kirschmeier P. Farnesyl transferase inhibitors block the farnesylation of CENP-E and CENP-F and alter the association of CENP-E with the microtubules. J Biol Chem 2000;275:30451–30457. 28. Mazumdar M, Misteli T. Chromokinesins: multitalented players in mitosis. Trends Cell Biol 2005;15:349–355. 29. Tokai-Nishizumi N, Ohsugi M, Suzuki E, Yamamoto T. The chromokinesin Kid is required for maintenance of proper metaphase spindle size. Mol Biol Cell 2005;16:5455–5463. 30. Levesque AA, Compton DA. The chromokinesin Kid is necessary for chromosome arm orientation and oscillation, but not congression, on mitotic spindles. J Cell Biol 2001;154:1135–1146. 31. Brouhard GJ, Hunt AJ. Microtubule movements on the arms of mitotic chromosomes: polar ejection forces quantified in vitro. Proc Natl Acad Sci USA 2005;102:13903–13908. 32. Zhu C, Jiang W. Cell cycle-dependent translocation of PRC1 on the spindle by Kif4 is essential for midzone formation and cytokinesis. Proc Natl Acad Sci USA 2005;102:343–348. 33. Mazumdar M, Sundareshan S, Misteli T. Human chromokinesin KIF4A functions in chromosome condensation and segregation. J Cell Biol 2004;166:613–620. 34. Carleton M, Mao M, Biery M, Warrener P, Kim S, Buser C, Marshall CG, Fernandes C, Annis J, Linsley PS. RNA interference-mediated silencing of mitotic kinesin KIF14 disrupts cell cycle progression and induces cytokinesis failure. Mol Cell Biol 2006;26:3853–3863. 35. Gruneberg U, Neef R, Li X, Chan EHY, Chalamalasetty RB, Nigg EA, Barr FA. KIF14 and citron kinase act together to promote efficient cytokinesis. J Cell Biol 2006;172:363–372. 36. Corson TW, Huang A, Tsao MS, Gallie BL. KIF14 is a candidate oncogene in the 1q minimal region of genomic gain in multiple cancers. Oncogene 2005;24:4741–4753. 37. Zhu C, Bossy-Wetzel E, Jiang W. Recruitment of MKLP1 to the spindle midzone/midbody by INCENP is essential for midbody formation and completion of cytokinesis in human cells. Biochem J 2005;389: 373–381. 38. Gruneberg U, Neef R, Honda R, Nigg EA, Barr FA. Relocation of Aurora B from centromeres to the central spindle at the metaphase to anaphase transition requires MKlp2. J Cell Biol 2004;166:167–172.


223

39. Neef R, Klein UR, Kopajtich R, Barr FA. Cooperation between mitotic kinesins controls the late stages of cytokinesis. Curr Biol 2006;16: 301–307. 40. Abaza A, Soleilhac JM, Westendorf J, Piel M, Crevel I, Roux A, Pirollet F. M phase phosphoprotein 1 is a human plus-end-directed kinesin-related protein required for cytokinesis. J Biol Chem 2003;278: 27844–27852. 41. Aoki S, Ohta K, Yamazaki T, Sugawara F, Sakaguchi K. Mammalian mitotic centromere-associated kinesin (MCAK): a new molecular target of sulfoquinovosylacylglycerols novel antitumor and immunosuppressive agents. FEBS 2005;272:2132–2140. 42. Goshima G, Vale RD. The roles of microtubule-based motor proteins in mitosis: comprehensive RNAi analysis in the Drosophila S2 cell line. J Cell Biol 2003;162:1003–1016. 43. Crespo NC, Delarue F, Ohkanda J, Carrico D, Hamilton AD, Sedti SM. The farnesyltransferase inhibitor, FTI-2153, inhibits bipolar spindle formation during mitosis independently of transformation and Ras and p53 mutation status. Cell Death Differ 2002;9:702–709. 44. Bergnes G, Brejc K, Belmont L. Mitotic kinesins: prospects for antimitotic drug discovery. Curr Top Med Chem 2005, 5, 127–145. 45. Sarli V, Giannis A. Inhibitors of mitotic kinesins: next-generation antimitotics. Chem Med Chem 2006;13,293–298. 46. Mayer TU, Kapoor TM, Haggarty SJ, King RW, Schreiber SL, Mitchison TJ. Small molecule inhibitor of mitotic spindle bipolarity identified in a phenotype-based screen. Science 1999;286:971–974. 47. Kapoor TM, Mayer TU, Coughlin ML, Mitchison TJ. Probing spindle assembly mechanisms with monastrol, a small molecule inhibitor of the mitotic kinesin, Eg5. J Cell Biol 2000;150:975–988. 48. Maliga Z, Kapoor TM, Mitchison TJ. Evidence that monastrol is an allosteric inhibitor of the mitotic kinesin Eg5. Chem Biol 2002;9: 989–996. 49. DeBonis S, Simorre JP, Crevel I, Lebeau L, Skoufias DA, Blangy A, Ebel C, Gans P, Cross R, Hackney DD, Wade RH, Kozielski F. Interaction of the mitotic inhibitor monastrol with human kinesin Eg5. Biochemistry 2003;42:338–349. 50. Yan Y, Sardana V, Xu B, Homnick C, Halczenko W, Buser CA, Schaber M, Hartman GD, Huber HE, Kuo LC. Inhibition of a mitotic motor protein: where, how, and conformational consequences. J Mol Biol 2004;335:547–554. 51. Leizerman I, Avunie-Masala R, Elkabets M, Fich A, Gheber L. Differential effects of monastrol in two human cell lines. Cell Mol Life Sci 2004;61,2060–2070.

224

Sarli and Giannis

52. Haque SA, Hasaka TP, Brooks AD, Lobanov PV, Baas PW. Monastrol, a prototype anti-cancer drug that inhibits a mitotic kinesin, induces rapid bursts of axonal outgrowth from cultured postmitotic neurons. Cell Motil Cytoskeleton 2004;58:10–16. 53. Yoon SY, Choi JE, Huh JW, Hwang O, Lee HS, Hong HN, Kim D. Monastrol, a selective inhibitor of the mitotic kinesin Eg5, induces a distinctive growth profile of dendrites and axons in primary cortical neuron cultures. Cell Motil Cytoskeleton 2005;60:181–190. 54. Naito M, Hamada H, Tsuruo T. ATP/Mg2+-dependent binding of vincristine to the plasma membrane of multidrug-resistant K562 cells. J Biol Chem 1988;263:11887–11891. 55. Peters T, Lindenmaier H, Haefeli WE, Weiss J. Interaction of the mitotic kinesin Eg5 inhibitor monastrol with P-glycoprotein. NaunynSchmiedebergs Arch Pharmacol 2006;372:291–299. 56. Gartner M, Sunder-Plassmann N, Seiler J, Utz M, Vernos I, Surrey T, Giannis A. Development and biological evaluation of potent and specific inhibitors of mitotic Kinesin Eg5. Chem Bio Chem 2005;5:1173–1177. 57. Sarli V, Huemmer S, Sunder-Plassmann N, Mayer TU, Giannis A. Synthesis and biological evaluation of novel EG5 inhibitors. Chem Bio Chem 2005;6:2005–2013. 58. Muller C, Gross D, Sarli V, Gartner M, Giannis A, Bernhardt G, Buschauer A. Inhibitors of kinesin Eg5: antiproliferative activity of monastrol analogues against human glioblastoma cells. Cancer Chemother Pharmacol 2006; Epub ahead of print. 59. Liu M, Aneja R, Liu C, Sun L, Gao J, Wang H, Dong JT, Sarli V, Giannis A, Joshi HC, Zhou J. Inhibition of the mitotic kinesin Eg5 upregulates Hsp70 through the phosphatidylinositol 3-kinase/Akt pathway in multiple myeloma cells. J Biol Chem 2006;281:18090–18097. 60. Sakowicz R, Finer JT, Beraud C, Crompton A, Lewis E, Fritsch A, Lee Y, Mak J, Moody R, Turincio R, Chabala JC, Gonzales P, Roth S, Weitman S, Wood KW. Antitumor activity of a kinesin inhibitor. Cancer Res 2004;64:3276–3280. 61. Johnson RK, McCabe FL, Cauder E, Innlow L, Whitacre M, Winkler JD, Bergnes G, Feng B, Smith WW, Morgans D, Wood K, Jackson JR. SB-715992, a potent and selective inhibitor of KSP mitotic kinesin, demonstrates broad-spectrum activity in advanced murine tumors and human tumor xenografts. Proc Am Assoc Cancer Res 2002; 43:269. 62. Davis DA, Sarkar SH, Hussain M, Li Y, Sarkar FH. Increased therapeutic potential of an experimental anti-mitotic inhibitor SB715992 by genistein in PC-3 human prostate cancer cell line. BMC Cancer 2006;6:22. 63. Nakazawa J, Yajima J, Usui T, Ueki M, Takatsuki A, Imoto M, Toyoshima YY, Osada H. A novel action of terpendole E on the motor activity of mitotic Kinesin Eg5. Chem Biol 2003;10:131–137.


225

64. Hotha S, Yarrow JC, Yang JG, Garrett S, Renduchintala KV, Mayer TU, Kapoor TM. HR22C16: a potent small-molecule probe for the dynamics of cell division. Angew Chem 2003;115:2481–2484. 65. Marcus AI, Peters U, Thomas SL, Garrett S, Zelnak A, Kapoor TM, Giannakakou P. Mitotic kinesin inhibitors induce mitotic arrest and cell death in Taxol-resistant and -sensitive cancer cells. J Biol Chem 2005;280:11569–11577. 66. Sunder-Plassmann N, Sarli V, Gartner M, Utz M, Seiler Z, Huemmer S, Mayer TU, Surrey T, Giannis A. Synthesis and biological evaluation of new tetrahydro-beta-carbolines as inhibitors of the mitotic kinesin Eg5. Bioorg Med Chem 2005;13:6094–6111. 67. DeBonis S, Skoufias DA, Lebeau L, Lopez R, Robin G, Margolis RL, Wade RH, Kozielski F. In vitro screening for inhibitors of the human mitotic kinesin Eg5 with antimitotic and antitumor activities. Mol Cancer Ther 2004;3:1079–1090. 68. Skoufias DA, DeBonis S, Saoudi Y, Lebeau L, Crevel I, Cross R, Wade RH, Hackney D, Kozielski F. S-trityl-L-cysteine is a reversible, tight binding inhibitor of the human kinesin Eg5 that specifically blocks mitotic progression. J Biol Chem 2006;281:17559–17569. 69. Brier S, Lemaire D, DeBonis S, Forest E, Kozielski F. Identification of the protein binding region of S-trityl-L-cysteine, a new potent inhibitor of the mitotic kinesin Eg5. Biochemistry 2004;43:13072–13082. 70. Brier S, Lemaire D, DeBonis S, Forest E, Kozielksi F. Molecular dissection of the inhibitor binding pocket of mitotic kinesin Eg5 reveals mutants that confer resistance to antimitotic agents. J Mol Biol 2006;360:360–376. 71. Tao W, South VJ, Zhang Y, Davide JP, Farell L, Kohl NE, Sepp-Lorenzino L, Lobell RB. Induction of apoptosis by an inhibitor of the mitotic kinesin KSP requires both activation of the spindle assembly checkpoint and mitotic slippage. Cancer Cell 2005;8:49–59. 72. Fraley ME, Garbaccio RM, Arrington KL, Hoffman WF, Tasber ES, Coleman PJ, Buser CA, Walsh ES, Hamilton K, Fernandes C, Schaber MD, Lobell RB, Tao W, South VJ, Yan Y, Kuo LC, Prueksaritanont T, Shu C, Torrent M, Heimbrook DC, Kohl NE, Huber HE, Hartman GD. Kinesin spindle protein (KSP) inhibitors. Part 2: the design, synthesis, and characterization of 2,4-diaryl-2,5-dihydropyrrole inhibitors of the mitotic kinesin KSP. Biorg Med Chem Let 2006;16:1775–1779. 73. Cox CD, Breslin MJ, Mariano BJ, Coleman PJ, Buser CA, Walsh ES, Hamilton K, Huber HE, Kohl NE, Torrent M, Yan Y, Kuo LC, Hartman GD. Kinesin spindle protein (KSP) inhibitors. Part 1: The discovery of 3,5-diaryl-4,5-dihydropyrazoles as potent and selective inhibitors of the mitotic kinesin KSP. Biorg Med Chem Let 2005;15:2041–2045.

226

Sarli and Giannis

74. Tarby CM, Kaltenbach RF, Huynh T, Pudzianowski A, Shen H, Ortega-Nanos M, Sheriff S, Newitt JA, McDonnell PA, Burford N, Fairchild CR, Vaccaro W, Chen Y, Borzilleri RM, Naglich J, Lombardo LJ, Gottardis M, Trainor GL, Roussell DL. Inhibitors of human mitotic kinesin Eg5: characterization of the 4-phenyltetrahydroisoquinoline lead series. Bioorg Med Chem 2006;16: 2095–2100. 75. Kim KS, Lu S, Cornelius LA, Lombardo LJ, Borzilleri RM, Schroeder GM, Sheng C, Rovnyak G, Crews D, Schmidt RJ, Williams DK, Bhide RS, Traeger SC, McDonnell PA, Mueller L, Sheriff S, Newitt JA, Pudzianowski AT, Yang Z, Wild R, Lee FY, Batorsky R, Ryder JS, Ortega-Nanos M, Shen H, Gottardis M, Roussell DL. Synthesis and SAR of pyrrolotriazine-4-one based Eg5 inhibitors. Bioorg Med Chem Lett 2006;16:3937–3942. 76. Okumura H, Nakazawa J, Tsuganezawa K, Usui T, Osada H, Matsumoto T, Tanaka A, Yokoyama S. Phenothiazine and carbazolerelated compounds inhibit mitotic kinesin Eg5 and trigger apoptosis in transformed culture cells. Toxicol Let 2006;166:44–52. 77. Koller E, Propp S, Zhang H, Zhao C, Xiao X, Chang M, Hirsch SA, Shepard PJ, Koo S, Murphy C, Glazer RI, Dean NM. Use of a chemically modified antisense oligonucleotide library to identify and validate Eg5 (kinesin-like 1) as a target for antineoplastic drug development. Cancer Res 2006;66:2059–2066.

10

Targeting the Spindle Checkpoint in Cancer Chemotherapy Jungseog Kang and Hongtao Yu CONTENTS Introduction Molecular Mechanisms of Chromosome Segregation and the Spindle Checkpoint A Defective Spindle Checkpoint Contributes to Tumorigenesis Role of the Spindle Checkpoint in Apoptosis Caused by Antimitotic Drugs Targeting the Spindle Checkpoint Proteins for Cancer Chemotherapy

Abstract Proper chromosome segregation during mitosis is critical for cells to inherit the correct number of chromosomes and maintain genetic stability. The spindle checkpoint is a cell-cycle surveillance mechanism that prevents premature sister-chromatid separation and ensures the fidelity of chromosome segregation. A defective spindle checkpoint results in aneuploidy and contributes to tumorigenesis. On the other hand, many tumor cells still exhibit a partially From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

227

228

Kang and Yu

functional spindle checkpoint and undergo prolonged mitotic arrest followed by apoptosis when treated with the antimitotic class of anticancer drugs, such as paclitaxel (Taxol). Recent studies have shown that a more complete inactivation of the spindle checkpoint reduces the efficacy of these drugs in eliciting apoptosis in cultured cancer cells. Therefore, quantitative differences in the strength of the spindle checkpoint may influence the efficacy of antimitotic drugs in cancer chemotherapy. Key Words: Mitosis; chromosome segregation; the spindle checkpoint; aneuploidy; the anaphase-promoting complex; ubiquitination; antimitotics; Mad2; Cdc20; Bub1

1. INTRODUCTION Equal partitioning of genetic material during mitosis is a challenging task for eukaryotic cells. After chromosome duplication, microtubule fibers emanating from the two opposite spindle poles attach to the opposing kinetochores of each pair of sister chromatids (1). Dissolution of sister-chromatid cohesion then allows the segregation of separated chromatids to each daughter cell (2). Chromosome missegregation results in aneuploidy, which contributes to tumorigenesis and causes birth defects and Down syndrome (3,4). To avoid these disastrous consequences, eukaryotic cells have developed a sophisticated surveillance mechanism called the spindle checkpoint to ensure the fidelity of chromosome segregation during mitosis and meiosis (3,5). In response to chromatids that have not yet achieved proper attachment to the mitotic spindle, this checkpoint delays the onset of anaphase (3,5). Most cancer cells are aneuploid. Aneuploidy, in combination with other genetic alterations, is believed to confer proliferative advantage to cancer cells by altering the gene dosage of tumor suppressors and protooncogenes (4). Recent genetic studies in mice have demonstrated that a defective spindle checkpoint can lead to aneuploidy and tumorigenesis. On the other hand, mutations of the known spindle checkpoint genes are rare in human cancers. Indeed, most cancer cells undergo prolonged mitotic arrest when exposed to the antimitotic class of anticancer drugs, such as vinblastine and Taxol, indicating that they still possess a somewhat functional spindle checkpoint. In this article, we review how chromosome segregation is regulated by the spindle checkpoint, how defects of the spindle checkpoint contribute to tumorigenesis, and finally how the spindle checkpoint influences apoptosis caused by antimitotic drugs.

Chapter 10 / The Spindle Checkpoint in Cancer Chemotherapy

229

2. MOLECULAR MECHANISMS OF CHROMOSOME SEGREGATION AND THE SPINDLE CHECKPOINT When eukaryotic cells enter mitosis, the replicated sister chromatids are held together by the cohesin protein complex (2). At early prophase, the cohesin complex on chromosome arms is released by Plk1/Aurora B-dependent phosphorylation, but the centromeric cohesin is protected by shugoshin (6). At the metaphase to anaphase transition, a large ubiquitin ligase called the anaphase-promoting complex or cyclosome in complex with its mitotic activator Cdc20 (APC/CCdc20 ) mediates the destruction of several important cell-cycle regulators, such as securin and cyclin B1 (7) (Fig. 1). Degradation of securin causes activation of separase, which cleaves the centromeric pool of cohesin and triggers sister-chromatid separation. Destruction of cyclin B1 inactivates Cdk1 and allows mitotic exit and cytokinesis. The unattached kinetochores

A

Unattached kinetochore or lack of tension

Plk1 and Aurora B

Prophase

Metaphase

B

Anaphase Ub Ub Ub Cyclin B

Cdc20

Spindle Checkpoint

Spindle Checkpoint

APC/C

Mitotic Exit

Cdk1

Ub Ub Ub Securin Separase

Sister chromatid separation

Fig. 1. Chromosome segregation and the spindle checkpoint. (A) Sister-chromatid cohesion is resolved by a Plk1- and Aurora B-dependant pathway at early prophase while the centromeric cohesion is removed by separase. The spindle checkpoint prevents the activation of separase and the removal of centromeric cohesion until bipolar attachment of all kinetochores to the mitotic spindle. (B) The spindle checkpoint inhibits the activation of APC/C, prevents degradation of securin and cyclin B1, and blocks separation of sister chromatids and mitotic exit.

230

Kang and Yu

activate the spindle checkpoint, which inhibits APC/CCdc20 , stabilizes securin and cyclin B1, prevents the activation of separase, and delays the onset of anaphase (8). Several spindle checkpoint components are conserved from yeast to man, including Bub1, Bub3, Mad1, Mad2, BubR1, Mps1, and the Aurora B-INCENP-survivin-borealin chromosome passenger complex (3,5). Higher eukaryotes have additional players that are not conserved in yeast, such as the Rod-Zw10-Zwilch complex, presumably because of increased complexity in their mitotic processes (3,5). It is generally believed that the spindle checkpoint senses both the lack of occupancy of spindle microtubules on kinetochores and the lack of tension across the two opposing kinetochores of a given pair of sister chromatids (9,10). Though it is not exactly clear how the spindle checkpoint detects these defects, most if not all spindle checkpoint proteins localize to unattached kinetochores and their kinetochore localization is critical for checkpoint signaling. Numerous studies in Xenopus egg extracts and mammalian cells have established a hierarchy of kinetochore recruitment for various checkpoint proteins (11–13). The chromosome passenger complex consisting of Aurora B, INCENP, survivin, and borealin lies at the top of this cascade. It is required for the kinetochore localization of Bub1 and Mps1, two protein kinases that are interdependent for their own kinetochore localization. Bub1 and Mps1 are in turn required for the kinetochore localization of BubR1 (a protein kinase related to Bub1), Mad1, and Mad2. Interestingly, the kinetochore localization of many checkpoint proteins is dynamic, i.e., the kinetochore-bound and cytosolic pools of these proteins exchange rapidly (14–16). Furthermore, several spindle checkpoint proteins are direct inhibitors of APC/CCdc20 . In vitro, Mad2 or BubR1 directly binds to Cdc20 and acts as stoichiometric inhibitors of APC/CCdc20 (17–19) whereas Bub1 phosphorylates Cdc20 and inhibits APC/CCdc20 catalytically (20). In vivo, BubR1, Bub3, Mad2, and Cdc20 can be incorporated in to a single complex, called the mitotic checkpoint complex (MCC) (19). Collectively, these studies support a model in which the unattached/untense kinetochores emit multiple diffusible inhibitory signals that block the activity of APC/CCdc20 upon checkpoint activation, thus stabilizing securin and delaying the onset of anaphase (3). After each pair of sister chromatid achieves bipolar attachment to the mitotic spindle, several mechanisms contribute to the silencing of the spindle checkpoint. First, the kinetochore concentrations of many spindle checkpoint proteins, including Mad1 and Mad2, drop


231

sharply upon microtubule attachment, partly because of their dyneindependent transport to the spindle poles (21). The Rod-Zw10-Zwilch complex also plays important roles in this process, because its inactivation prevents the recruitment of the dynein-dynactin complex to the kinetochores (22,23). Second, p31comet selectively binds to Mad1or Cdc20-bound forms of Mad2 (24,25). In doing so, it prevents the generation of Mad2–Cdc20-containing APC/C-inhibitory checkpoint complexes and neutralizes the APC/C-inhibitory activity of Mad2 in these complexes (25–27). Together, these mechanisms help to silence the spindle checkpoint, although it remains unclear whether and how the Mad2–p31comet interaction is regulated by upstream signals of the spindle checkpoint.

3. A DEFECTIVE SPINDLE CHECKPOINT CONTRIBUTES TO TUMORIGENESIS To study the role of the spindle checkpoint in tumorigenesis, several mouse models have been established, such as Mad2-, Bub3-, and BubR1-null mice. Homozygous Mad2-null mice die around day 7.5 of embryogenesis (E7.5), although a small set of embryos remain viable until E10.5 (28). The lethality of mouse embryonic fibroblast (MEF) cells with Mad2 depletion was rescued by inactivation of p53, suggesting that cells with high rate of chromosome missegregation resulted in p53-dependent apoptosis (29). Heterozygous deletion of the MAD2 allele results in a defective spindle checkpoint and increased rate of lung tumors after a long latency (30). MEFs from these mice exhibit premature sister-chromatid separation and elevated chromosome missegregation. Mice with homozygous deletion of Bub3 also fail to survive beyond E7.5 (31). MEFs from heterozygous Bub3deleted mice show an increase of chromosome missegregation and aneuploidy because of a spindle checkpoint defect (32). Furthermore, when treated with carcinogen, the Bub3+/− heterozygous mice have a higher incidence of lung cancer. Complete ablation of BubR1 also causes embryonic lethality in mice (33). Heterozygous BubR1+/− mice develop lung and intestinal cancers when challenged with carcinogens (34). Interestingly, mice with their BubR1 level reduced to about 10% of that of the wild-type have a short life space and develop multiple aging-associated phenotypes at an earlier age (35), suggesting a role for BubR1 in preventing premature aging. The effect of a defective spindle checkpoint in tumorigenesis becomes more apparent when combined with mutations of other tumor suppressor genes. For example, the

232

Kang and Yu

incidence of colonic tumors in mice harboring one mutant allele of the Apc tumor suppressor is significantly increased when one of their BubR1 alleles is deleted (36). Homozygous Brca2 mutant mice develop thymic lymphomas, in which Bub1 or BubR1 mutation were often observed, suggesting that inactivation of spindle checkpoint facilitates tumorigenesis in Brca2-null background (37). Collectively, these studies in mice suggest that a defective spindle checkpoint contributes to tumorigenesis. The first case of genetic cancer predisposition syndrome with germline mutations in a spindle checkpoint gene has been reported recently (38,39). Patients with mosaic variegated aneuploidy (MVA) syndrome have a high propensity to develop cancers, such as Wilms tumor, rhabdomyosarcoma, and leukemia (40). In one study, 5 of 8 MVA families have been shown to carry biallelic BubR1 mutations with one allele deleted and the other containing missense mutations (38). In another study, all 7 families have monoallelic BubR1 mutations that decreases the total expression levels of BubR1 by more than 50% (39). These findings strongly suggest that aneuploidy resulting from a defective spindle checkpoint is a causal factor for cancer formation. Consistently, various human tumor cell lines contain mutations in the spindle checkpoint genes, including Bub1, BubR1, Mad1, Mad2, Zw10, Zwilch, or ROD (4). Some of the mutations in Bub1 or Mad1 have been shown to interfere with normal spindle checkpoint signaling (41,42). However, mutations of spindle checkpoint genes in cancer cell lines are relatively rare. Instead, misregulation of protein levels of these checkpoint components by expression of oncogenes or mutations in tumor suppressors is more frequently observed in human cancers (4). For instance, BRCA1 positively regulates the transcription of Mad2 through binding to the promoter of Mad2 (43). BRCA1 is also involved in the transcriptional regulation of BubR1 (44). p53 directly binds to the Mad1 promoter and regulates its expression (45,46). Thus, mutations of these tumor suppressors are expected to decrease the levels of these checkpoint proteins. Furthermore, the Tax oncoprotein from HTLV-1 binds to Mad1 and may interfere with its function (47). Breast cancer specific gene 1, BCSG1, was reported to bind to BubR1 directly, causing its degradation (48). In Rb-negative tumor cells, E2F binds to the Mad2 promoter and upregulates Mad2 expression (49,50). In this case, however, it is unclear how an increase in Mad2 levels contributes chromosome instability. Taken together, multiple mechanisms exist to weaken the spindle checkpoint, possibly contributing to aneuploidy and tumorigenesis.


233

4. ROLE OF THE SPINDLE CHECKPOINT IN APOPTOSIS CAUSED BY ANTIMITOTIC DRUGS The antimitotic class of chemotherapeutic drugs, including the taxanes and the vinca alkaloids, target spindle microtubules and disrupt normal progression through mitosis of cancer cells. Prolonged exposure (>24 h) of tumor cells to these drugs at concentrations around 100 nM in vitro activates the spindle checkpoint, resulting in mitotic arrest followed by apoptosis. How tumor cells respond to treatment of antimitotic drugs in vivo is not well understood. The concentration of paclitaxel in the plasma after intravenous infusion to cancer patients decreases quickly because of detoxification by the liver (51–54). However, the concentration of paclitaxel in the plasma is above 100 nM (the concentration sufficient to trigger the spindle checkpoint in vitro) for longer than 20 h following each administration. Furthermore, the concentration of paclitaxel in cells is higher than in the medium because cells accumulate paclitaxel because of its high affinity toward microtubules (55). Therefore, it is formally possible that tumor cells can be exposed to high concentrations of paclitaxel in vivo to maintain a prolonged spindle checkpoint-dependent mitotic arrest followed by apoptosis. Tumor cells that are exposed to the antimitotic drugs can have several possible fates. In the first scenario, cells are refractory to the treatment and continue to proliferate in the presence of these drugs. In the second scenario, cells undergo a prolonged mitotic arrest in the presence of high drug concentrations, but resume normal cell cycle after drug clearance. In a third scenario, cells experience a brief mitotic delay and then undergo mitotic exit without cytokinesis, resulting in polyploid interphase cells. A major mechanism for human cells to escape from mitosis in the presence of nocodazole is chronic cyclin B degradation without silencing of the spindle checkpoint (56). It has also been suggested that p53-positive tumor cells can remain arrested in G1 following this type of mitotic exit because of a tetraploidy checkpoint, although the existence of such a checkpoint has recently been challenged (57). Indeed, most human tumor cell lines continue to replicate their DNA once they successfully escape from mitosis in the presence of paclitaxel. After drug clearance, these polyploid cells resume their cell cycle, but may have difficulty to complete subsequent mitosis because of their larger numbers of chromosomes and centrosomes. Some progenies of these cells may commit apoptosis whereas others may become grossly aneuploid. All of the above scenarios will not lead to efficient killing of tumor cells. In the final scenario,

234

Kang and Yu

in the presence of high concentrations of paclitaxol, cells undergo a mitotic arrest followed by apoptosis without an intervening interphase of significant duration. Several lines of evidence suggest that this mechanism requires at least a partially functional spindle checkpoint and may explain the cytotoxic effects of antimitotic drugs. Earlier studies by Taylor and McKeon (58) showed that overexpression of a dominant-negative mutant of Bub1 (DN-Bub1) significantly decreases the apoptotic sub-G1 population in nocodazole-treated cells. Correspondingly, 73% of DN-Bub1-overexpressing cells escape from mitosis in the presence of nocodazole and have DNA content greater than 4N whereas only 30% of control cells contain greater than 4N DNA. This suggests that inactivation of the spindle checkpoint facilitates mitotic slippage and survival upon nocodazole treatment. A more recent study also showed that inactivation of BubR1 or Bub1 decreases apoptotic cell death and increases population of polyploidy cells in the presence of spindle poisons, indicating a requirement of the spindle checkpoint in apoptosis triggered by antimitotic drugs (59). Another study also demonstrated that inactivation of Mad2 or BubR1 in breast cancer cells result in paclitaxel-resistance (60). When paclitaxel was added to these cells, the mitotic index of Mad2- or BubR1-depleted cells by RNAi does not increase because of a spindle checkpoint defect. Prolonged paclitaxel treatment for 48 h causes about 70% of cell death in control cells but only 30% of cell death in Mad2 or BubR1-depleted cells. Furthermore, overexpression of Mad2 in two breast cancer cell lines exhibiting low expression of Mad2 restores the spindle checkpoint and increases cell death about 2-fold when treated with paclitaxel. Consistent with a requirement of the spindle checkpoint in antimitotic drug-induced cell killing, there is a significant positive correlation between nocodazole-induced cytotoxicity and the status of the spindle checkpoint in several human lung cancer cell lines (61). In 12 lung cancer cell lines examined, 4 out of 6 cell lines exhibiting less cytotoxicity to nocodazole had an impaired spindle checkpoint whereas 6 out of 6 cell lines showing high toxicity to nocodazole displayed a normal spindle checkpoint. Finally, the importance of the spindle checkpoint in antimitotics-induced cytotoxicity was again demonstrated in a recent study using a chemical inhibitor of the mitotic kinesin KSP (62). The mitotic kinesin KSP is required for the formation of the bipolar spindle in mitosis. Inhibition of KSP generates monopolar spindles and activates the spindle checkpoint. Activation of the spindle checkpoint by the KSP inhibitor is reversible, but longer treatment of the inhibitor over 24 h triggers irreversible apoptotic cell death. Activation


235

of caspase-3 during apoptosis correlates with the disappearance of mitotic markers such as phosphorylation of nucleolin, suggesting that mitotic slippage is coupled with apoptosis. Indeed, inhibition of Cdk1 in KSP inhibitor-resistant cell lines promotes mitotic slippage and increases the population of apoptotic cells. Overexpression of the dominant-negative Bub1 mutant in KSA inhibitor- or paclitaxel-treated cells reduced apoptosis. Reduction of the Mad2 level also increases survival of cancer cells treated with the KSA inhibitor. These and other lines of evidence suggest that activation of the spindle checkpoint is required for efficient cancer cell killing by antimitotic drugs and other drugs that target proteins with essential mitotic functions (Fig. 2). Considering the long duration of high plasma paclitaxel concentration during cancer chemotherapy, cancer cells may experience a prolonged mitotic arrest that is dependent on the spindle checkpoint and proceed to commit apoptosis. In the presence of these antimitotic drugs, cells

Functional/partially functional spindle checkpoint

Interphase cells

Defective spindle checkpoint

Interphase cells

Paclitaxel

Paclitaxel

No or transient mitotic delay

Prolonged mitotic arrest Mitotic cells

Apoptotic cells

Mitotic cells

Polyploid interphase cells Apoptotic cells

Polyploid interphase cells

Fig. 2. Role of the spindle checkpoint in paclitaxel-induced apoptosis in tumor cells. Upon treatment of paclitaxel, cancer cells with a competent spindle checkpoint undergo prolonged mitotic arrest followed by apoptosis. On the other hand, tumor cells that are deficient in the spindle checkpoint progress through mitosis without cytokinesis and become polyploid.

236

Kang and Yu

without a sufficiently strong spindle checkpoint will experience no or transient mitotic delay and undergo mitotic exit without cytokinesis, resulting in polyploid interphase cells that will continue cell division following the decline of drug concentrations. Activation of the spindle checkpoint is required for antimitotic cancer drug-induced apoptosis, but the molecular connection between these two processes is not clear. Because several components of the spindle checkpoint, including Mps1 (63) and BubR1 (59), were shown to promote apoptosis, the spindle checkpoint might be capable of generating proapoptotic signals. It is also possible that the apoptotic cell death is the secondary consequence of the prolonged mitotic arrest caused by the spindle checkpoint. Future studies are needed to uncover the mechanisms that connect spindle checkpoint activation to apoptosis.

5. TARGETING THE SPINDLE CHECKPOINT PROTEINS FOR CANCER CHEMOTHERAPY A complete inactivation of the spindle checkpoint causes gross chromosome missegregation and results in lethality in multicellular organisms. On the other hand, partial inactivation of the spindle checkpoint increases the frequency of chromosome missegregation and facilitates tumorigenesis. Nevertheless, the weakened spindle checkpoint in many tumor cells still arrest them in mitosis when challenged with high concentrations of antimitotic drugs, which is required for efficient cell killing by these drugs. One possible explanation is that the compromised spindle checkpoint in tumor cells is insufficient to detect a few unattached kinetochores during normal mitosis, but can respond to gross perturbations at all kinetochores in the presence of antimitotic drugs. Therefore, for tumors with a completely functional spindle checkpoint, inhibitors that partially inactivate the spindle checkpoint will synergize with the antimitotic drugs. For tumors with an inactive spindle checkpoint, compounds that enhance the spindle checkpoint may be beneficial. Along this line, inactivation of p31comet (a Mad2 inhibitor) sensitizes HeLa cells toward Taxol-induced mitotic arrest (25). In conclusion, as different tumor cells vary in the strength of their spindle checkpoint, tailored strategies that manipulate the spindle checkpoint in a refined manner are needed to improve the efficacy of antimitotic drugs.


237

REFERENCES 1. Cleveland DW, Mao Y, Sullivan KF. Centromeres and kinetochores: from epigenetics to mitotic checkpoint signaling. Cell 2003;112:407–421. 2. Nasmyth K. Segregating sister genomes: the molecular biology of chromosome separation. Science 2002;297(5581):559–565. 3. Bharadwaj R, Yu H. The spindle checkpoint, aneuploidy, and cancer. Oncogene 2004;23(11):2016–2027. 4. Kops GJ, Weaver BA, Cleveland DW. On the road to cancer: aneuploidy and the mitotic checkpoint. Nat Rev Cancer 2005;5(10):773–785. 5. Musacchio A, Hardwick KG. The spindle checkpoint: structural insights into dynamic signalling. Nat Rev Mol Cell Biol 2002;3(10):731–741. 6. Yu H, Tang Z. Bub1 multitasking in mitosis. Cell Cycle 2005;4(2): 262–265. 7. Peters JM. The anaphase promoting complex/cyclosome: a machine designed to destroy. Nat Rev Mol Cell Biol 2006;7(9):644–656. 8. Yu H. Regulation of APC-Cdc20 by the spindle checkpoint. Curr Opin Cell Biol 2002;14(6):706–714. 9. Pinsky BA, Biggins S. The spindle checkpoint: tension versus attachment. Trends Cell Biol 2005;15(9):486–493. 10. Andrews PD. Aurora kinases: shining lights on the therapeutic horizon? Oncogene 2005;24(32):5005–5015. 11. Vigneron S, Prieto S, Bernis C, Labbe JC, Castro A, Lorca T. Kinetochore localization of spindle checkpoint proteins: who controls whom? Mol Biol Cell 2004;15(10):4584–4596. 12. Chen RH. BubR1 is essential for kinetochore localization of other spindle checkpoint proteins and its phosphorylation requires Mad1. J Cell Biol 2002;158(3):487–496. 13. Sharp-Baker H, Chen RH. Spindle checkpoint protein Bub1 is required for kinetochore localization of Mad1, Mad2, Bub3, and CENP-E, independently of its kinase activity. J Cell Biol 2001;153(6):1239–1250. 14. Howell BJ, Hoffman DB, Fang G, Murray AW, Salmon ED. Visualization of Mad2 dynamics at kinetochores, along spindle fibers, and at spindle poles in living cells. J Cell Biol 2000;150(6):1233–1250. 15. Shah JV, Botvinick E, Bonday Z, Furnari F, Berns M, Cleveland DW. Dynamics of centromere and kinetochore proteins; implications for checkpoint signaling and silencing. Curr Biol 2004;14(11):942–952. 16. Howell BJ, Moree B, Farrar EM, Stewart S, Fang G, Salmon ED. Spindle checkpoint protein dynamics at kinetochores in living cells. Curr Biol 2004;14(11):953–964. 17. Tang Z, Bharadwaj R, Li B, Yu H. Mad2-Independent inhibition of APCCdc20 by the mitotic checkpoint protein BubR1. Dev Cell 2001;1(2):227–237.

238

Kang and Yu

18. Fang G. Checkpoint protein BubR1 acts synergistically with Mad2 to inhibit anaphase-promoting complex. Mol Biol Cell 2002;13(3):755–766. 19. Sudakin V, Chan GK, Yen TJ. Checkpoint inhibition of the APC/C in HeLa cells is mediated by a complex of BUBR1, BUB3, CDC20, and MAD2. J Cell Biol 2001;154(5):925–936. 20. Tang Z, Shu H, Oncel D, Chen S, Yu H. Phosphorylation of Cdc20 by Bub1 provides a catalytic mechanism for APC/C inhibition by the spindle checkpoint. Mol Cell 2004;16(3):387–397. 21. Howell BJ, McEwen BF, Canman JC, et al. Cytoplasmic dynein/dynactin drives kinetochore protein transport to the spindle poles and has a role in mitotic spindle checkpoint inactivation. J Cell Biol 2001;155(7): 1159–1172. 22. Starr DA, Williams BC, Hays TS, Goldberg ML. ZW10 helps recruit dynactin and dynein to the kinetochore. J Cell Biol 1998;142(3):763–774. 23. Buffin E, Lefebvre C, Huang J, Gagou ME, Karess RE. Recruitment of Mad2 to the kinetochore requires the Rod/Zw10 complex. Curr Biol 2005;15(9):856–861. 24. Habu T, Kim SH, Weinstein J, Matsumoto T. Identification of a MAD2binding protein, CMT2, and its role in mitosis. Embo J 2002;21(23): 6419–6428. 25. Xia G, Luo X, Habu T, Rizo J, Matsumoto T, Yu H. Conformationspecific binding of p31(comet) antagonizes the function of Mad2 in the spindle checkpoint. Embo J 2004;23(15):3133–3143. 26. Mapelli M, Filipp FV, Rancati G, et al. Determinants of conformational dimerization of Mad2 and its inhibition by p31comet. Embo J 2006;25(6):1273–1284. 27. Yu H. Structural activation of Mad2 in the mitotic spindle checkpoint: the two-state Mad2 model versus the Mad2 template model. J Cell Biol 2006;173(2):153–157. 28. Dobles M, Liberal V, Scott ML, Benezra R, Sorger PK. Chromosome missegregation and apoptosis in mice lacking the mitotic checkpoint protein Mad2. Cell 2000;101(6):635–645. 29. Burds AA, Lutum AS, Sorger PK. Generating chromosome instability through the simultaneous deletion of Mad2 and p53. Proc Natl Acad Sci U S A 2005;102(32):11296–11301. 30. Michel LS, Liberal V, Chatterjee A, et al. MAD2 haplo-insufficiency causes premature anaphase and chromosome instability in mammalian cells. Nature 2001;409(6818):355–359. 31. Kalitsis P, Earle E, Fowler KJ, Choo KH. Bub3 gene disruption in mice reveals essential mitotic spindle checkpoint function during early embryogenesis. Genes Dev 2000;14(18):2277–2282. 32. Babu JR, Jeganathan KB, Baker DJ, Wu X, Kang-Decker N, van Deursen JM. Rae1 is an essential mitotic checkpoint regulator that


33. 34.

35.

36.

37.

38.

39.

40.

41. 42.

43.

44.

45.

239

cooperates with Bub3 to prevent chromosome missegregation. J Cell Biol 2003;160(3):341–353. Wang Q, Liu T, Fang Y, et al. BUBR1 deficiency results in abnormal megakaryopoiesis. Blood 2004;103(4):1278–1285. Dai W, Wang Q, Liu T, et al. Slippage of mitotic arrest and enhanced tumor development in mice with BubR1 haploinsufficiency. Cancer Res 2004;64(2):440–445. Baker DJ, Jeganathan KB, Cameron JD, et al. BubR1 insufficiency causes early onset of aging-associated phenotypes and infertility in mice. Nat Genet 2004;36(7):744–749. Rao CV, Yang YM, Swamy MV, et al. Colonic tumorigenesis in BubR1+/−ApcMin/+ compound mutant mice is linked to premature separation of sister chromatids and enhanced genomic instability. Proc Natl Acad Sci U S A 2005;102(12):4365–4370. Lee H, Trainer AH, Friedman LS, et al. Mitotic checkpoint inactivation fosters transformation in cells lacking the breast cancer susceptibility gene, Brca2. Mol Cell 1999;4(1):1–10. Hanks S, Coleman K, Reid S, et al. Constitutional aneuploidy and cancer predisposition caused by biallelic mutations in BUB1B. Nat Genet 2004;36(11):1159–1161. Matsuura S, Matsumoto Y, Morishima K, et al. Monoallelic BUB1B mutations and defective mitotic-spindle checkpoint in seven families with premature chromatid separation (PCS) syndrome. Am J Med Genet A 2006;140(4):358–367. Matsuura S, Ito E, Tauchi H, Komatsu K, Ikeuchi T, Kajii T. Chromosomal instability syndrome of total premature chromatid separation with mosaic variegated aneuploidy is defective in mitotic-spindle checkpoint. Am J Hum Genet 2000;67(2):483–486. Cahill DP, Lengauer C, Yu J, et al. Mutations of mitotic checkpoint genes in human cancers. Nature 1998;392(6673):300–303. Tsukasaki K, Miller CW, Greenspun E, et al. Mutations in the mitotic check point gene, MAD1L1, in human cancers. Oncogene 2001;20(25):3301–3305. Wang RH, Yu H, Deng CX. A requirement for breast-cancer-associated gene 1 (BRCA1) in the spindle checkpoint. Proc Natl Acad Sci U S A 2004;101(49):17108–17113. Chabalier C, Lamare C, Racca C, Privat M, Valette A, Larminat F. BRCA1 downregulation leads to premature inactivation of spindle checkpoint and confers paclitaxel resistance. Cell Cycle 2006;5(9): 1001–1007. Chun AC, Jin DY. Transcriptional regulation of mitotic checkpoint gene MAD1 by p53. J Biol Chem 2003;278(39):37439–37450.

240

Kang and Yu

46. Iwanaga Y, Jeang KT. Expression of mitotic spindle checkpoint protein hsMAD1 correlates with cellular proliferation and is activated by a gainof-function p53 mutant. Cancer Res 2002;62(9):2618–2624. 47. Jin DY, Spencer F, Jeang KT. Human T cell leukemia virus type 1 oncoprotein Tax targets the human mitotic checkpoint protein MAD1. Cell 1998;93(1):81–91. 48. Gupta A, Inaba S, Wong OK, Fang G, Liu J. Breast cancer-specific gene 1 interacts with the mitotic checkpoint kinase BubR1. Oncogene 2003;22(48):7593–7599. 49. Hernando E, Nahle Z, Juan G, et al. Rb inactivation promotes genomic instability by uncoupling cell cycle progression from mitotic control. Nature 2004;430(7001):797–802. 50. Ren B, Cam H, Takahashi Y, et al. E2F integrates cell cycle progression with DNA repair, replication, and G(2)/M checkpoints. Genes Dev 2002;16(2):245–256. 51. Hurwitz CA, Relling MV, Weitman SD, et al. Phase I trial of paclitaxel in children with refractory solid tumors: a Pediatric Oncology Group Study. J Clin Oncol 1993;11(12):2324–2329. 52. Gianni L, Kearns CM, Giani A, et al. Nonlinear pharmacokinetics and metabolism of paclitaxel and its pharmacokinetic/pharmacodynamic relationships in humans. J Clin Oncol 1995;13(1):180–190. 53. Sonnichsen DS, Hurwitz CA, Pratt CB, Shuster JJ, Relling MV. Saturable pharmacokinetics and paclitaxel pharmacodynamics in children with solid tumors. J Clin Oncol 1994;12(3):532–538. 54. Huizing MT, Keung AC, Rosing H, et al. Pharmacokinetics of paclitaxel and metabolites in a randomized comparative study in platinum-pretreated ovarian cancer patients. J Clin Oncol 1993;11(11):2127–2135. 55. Jordan MA, Toso RJ, Thrower D, Wilson L. Mechanism of mitotic block and inhibition of cell proliferation by taxol at low concentrations. Proc Natl Acad Sci U S A 1993;90(20):9552–9556. 56. Brito DA, Rieder CL. Mitotic checkpoint slippage in humans occurs via cyclin B destruction in the presence of an active checkpoint. Curr Biol 2006;16(12):1194–1200. 57. Uetake Y, Sluder G. Cell cycle progression after cleavage failure: mammalian somatic cells do not possess a “tetraploidy checkpoint”. J Cell Biol 2004;165(5):609–615. 58. Taylor SS, McKeon F. Kinetochore localization of murine Bub1 is required for normal mitotic timing and checkpoint response to spindle damage. Cell 1997;89(5):727–735. 59. Shin HJ, Baek KH, Jeon AH, et al. Dual roles of human BubR1, a mitotic checkpoint kinase, in the monitoring of chromosomal instability. Cancer Cell 2003;4(6):483–497.


241

60. Sudo T, Nitta M, Saya H, Ueno NT. Dependence of paclitaxel sensitivity on a functional spindle assembly checkpoint. Cancer Res 2004;64(7):2502–2508. 61. Masuda A, Maeno K, Nakagawa T, Saito H, Takahashi T. Association between mitotic spindle checkpoint impairment and susceptibility to the induction of apoptosis by anti-microtubule agents in human lung cancers. Am J Pathol 2003;163(3):1109–1116. 62. Tao W, South VJ, Zhang Y, et al. Induction of apoptosis by an inhibitor of the mitotic kinesin KSP requires both activation of the spindle assembly checkpoint and mitotic slippage. Cancer Cell 2005;8(1):49–59. 63. Bhonde MR, Hanski ML, Budczies J, et al. DNA damage-induced expression of p53 suppresses mitotic checkpoint kinase hMps1: the lack of this suppression in p53MUT cells contributes to apoptosis. J Biol Chem 2006;281(13):8675–8685.

11

Antiproliferation Inhibitors Targeting Aurora Kinases Kishore Shakalya and Daruka Mahadevan CONTENTS Introduction Biology of Aurora Kinases Linking Aurora Kinases to Oncogenesis Inhibitors in Pre-Clinical and Clinical Development Structural Biology: Insight into Aurora Inhibition Conclusions and Future Directions

Abstract The family of serine-threonine (S/T) protein kinases Aurora A, B, and C regulate distinct functions and phases of mitosis within the cell cycle. Aurora A regulates centrosome maturation and spindle assembly while Aurora B (and C) a chromosome passenger protein (CPP) regulates chromosome orientation on the spindle and cytokinesis. All 3 Auroras are overexpressed in a multitude of human cancers with an associated polyploid phenotype containing multiple centrosomes. Aurora A and B when aberrantly overexpressed and dysregulated in an appropriate genetic background function as oncogenes by overriding cell-cycle checkpoints leading to errors From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

243

244

Shakalya and Mahadevan

in mitosis (aneuploidy) with subsequent chromosomal instability, a hallmark of human cancer. Over the past 5 years the biology of Aurora kinases in mitotic regulation has been further elucidated by RNA interference (RNAi), dominant-negative kinase mutants and several small molecular ATP-site competitive inhibitors. The availability of several crystal structures with bound ADP, several drugs and peptide fragments from interacting proteins have provided a detailed insight of the active site, biological functions, rationale for the mode of binding of a broad range of small molecular inhibitors and clues to the likelihood of developing resistance through point mutations. Homology models and crystal structures have also aided in fragment-based rational drug discovery of Aurora inhibitors including compounds specific for Aurora A and B respectively. Moreover, several Aurora kinase inhibitors have shown promising antitumor activity in rodent models of cancer. However, overexpression of Aurora A results in paclitaxel resistance owing to overriding of a spindle checkpoint and combination therapy studies are warranted to evaluate mechanisms of resistance. Despite these concerns, excitingly, over the past 2 years Aurora inhibitors have entered the phase I arena and are being evaluated in solid and hematological malignancies. Key Words: Aurora Kinase (A, B, C); mitosis; cell cycle; protooncogenes; protein kinase inhibitors; aneuploidy; cytokinesis

1. INTRODUCTION Aurora kinases (Aurora A and B) were first discovered as mutant alleles in Drosophila melanogaster that lead to defective spindle-pole assembly (1,2). Of the fungi, Saccharomyces cerevisiae and Saccharomyces pombe, have a single Aurora kinase known as increase-inploidy 1 (Ipl1) (3) and Aurora-related kinase 1 (ARK1) respectively (4). Evolutionary relationships ascertained from phylogenetic trees suggest that the 3 vertebrate Auroras (Aurora A, B and C) evolved from a single urochordate ancestor all of which is related to the prototype molecule Ipl1. Aurora A is an orthologous lineage in cold-blooded vertebrates whereas Aurora B and C evolved more recently from a gene duplication of an ancestral Aurora B/C gene in mammals. Comparison of the crystal structures of Aurora A and B to their respective catalytic domain sequences supports the notion that Aurora B and C are closely related paralogs (5). The human Aurora kinases range in length from 309 to 403 amino acids with a carboxy terminal catalytic domain of ∼300 residues, a

Chapter 11 / Antiproliferation Inhibitors Targeting Aurora Kinases

245

variable length amino terminal extension and a short carboxy terminal extension. The Aurora kinase families from fly to human (orthologs and homologs) share a highly conserved catalytic domain but their amino terminal extensions are of variable length with no sequence similarity (6). A structure based multiple sequence alignment of the 3 human Aurora kinases using Clustal W demonstrated that the length of the amino terminal extension of human Aurora A is 97 residues, Aurora B is 34 residues and Aurora C has no extension (7). Accumulated evidence suggests that the amino terminal regions of Aurora A and B provide specificity, are species specific; localize the kinases to their respective sub-cellular sites and substrate recognition. Within the carboxy terminal catalytic domain of Aurora A 2 sequences mediate degradation at the end of the mitotic phase (M-phase), which includes the destruction box (D-box) and the D-box activating sequence (Abox). In contrast, Aurora B and C have a putative D-box but lack an A-box and are not targeted to proteolysis during M-phase exit (1). Figure 1 shows a schematic representation of the 3 human Auroras including their lengths, % identity, domain organization and location of A and D-boxes. Several protein kinases (CDK1, Plk1, Nek2 and Auroras) tightly regulate the M-phase of the cell cycle, a complex biological process whereby a complete copy of the duplicated genome is precisely segregated by the microtubule spindle apparatus into 2 daughter cells. To preserve the integrity of this complex system multiple high fidelity checkpoint systems have evolved in parallel to ensure correct spatial-

N-terminal Extension 97

1 Aurora-A

402 1

57%

35

60%

344

Aurora-B 1

75%

Aurora-C

309 Catalytic Domain B A-Box

Kinase

D-Box

Fig. 1. shows a schematic representation of the 3 human Aurora kinases (A, B, C) including their lengths, % identity, domain organization and location of A and D-boxes.

246


temporal coordination (8). Aurora kinases are mitotic kinases that associate with chromosomes, chromosome-associated proteins and cytoskeletal components that drive cell division and are thus critical regulators of genomic stability. They appear at specific locations during the M-phase: Aurora A known as the “polar kinase” associates with the duplicating centrosomes; Aurora B known as the ‘equatorial kinase’ is a chromosomal passenger protein; Aurora C localizes to the centrosome from anaphase to telophase (1,9). All three human Aurora kinases are strongly associated with human cancer because of frequent overexpression and map to distinct regions of chromosomal loci known to be tumor associated amplicons. This implies Aurora kinases play important roles in tumor initiation and/or progression in an appropriate genetic context. In humans Aurora A maps to chromosome 20q13.2-q13.3, Aurora B to 17p13.1 (close to p53) and Aurora C to 19q13.43 (10). Given the oncogenic amplification of Aurora kinases in tumors, inhibiting its enzyme activity with a specific small molecular inhibitor (SMI) to the catalytic domain ATP-binding site is regarded as feasible for targeted cancer therapy. Several Aurora kinase SMIs have been developed, which has not only allowed further elucidation of the M-phase and its regulation with the attendant phenotypic changes but more importantly as antiproliferative inhibitors targeting both solid and hematological malignancies where their efficacy, potency and safety are currently being evaluated in clinical trials (11).

2. BIOLOGY OF AURORA KINASES Despite the highly conserved catalytic domains of Aurora kinases, they have different sub-cellular locations and functions. Model organisms (Drosophila Saccharomyces, Caenorhabditis and Xenopus) have provided detailed descriptions of the pivotal roles of Aurora function during cell division. During the M-phase, Aurora kinases are necessary for chromosome segregation, condensation and orientation in the metaphase plate, spindle assembly and to complete cytokines (Table 1). The details of these functions have been extensively reviewed (1,2,12).

2.1. Aurora A: Regulator of Centrosome & Spindle Assembly Normal proliferating cells express Aurora A, which peaks during the G2 /M phase of the cell cycle. At the beginning of M-phase, Aurora A is


247

Table 1 Biological Roles of Aurora Kinases Aurora kinase

Functions

Aurora A

• Localizes to centrosomes; centrosome duplication and maturartion; spindle assembly; • allosterically activated by TPX2, ajuba and phosphatase inhibitor-2 • phosphorylates TPX2, Eg5 kinesin-like protein, CDC25B phosphatase, p53, BRCA-1, CPEB (cytoplasmic polyadenylation element binding protein) • Associates with centromeres; chromosome bi-orientation; cytokinesis; • allosterically activated by INCENP and survivin; forms a complex with INCENP-survivin-borealin • phosphorylates histone H3, borealin, MgcRacGAP, vimentin, desmin, myosin II regulatory light chain, glial fibrillatory acidic protein, central-spindlin, microtubule depolymerase MCAK • Functions similar to Aurora B • expression restricted to reproductive organs

Aurora B

Aurora C

extensively confined to the spindle poles where it localizes to the duplicated centrosome and regulates their separation and maturation (1,13). TPX2 (target protein for xenopus kinesin-like protein 2) a microtubule associated protein (14) and Ajuba (LIM domain containing protein) (15) binds to Aurora A and regulates its kinase activity by auto-catalysis with consequent phosphorylation. Activated Aurora A is then localized to the spindle pole through the activated form of Ran (Ran-GTP) - TPX2 dependent mechanism by inhibiting importin- and . Further, TPX2 and microtubules promote Aurora A phosphorylation by preventing protein phosphatase I (PPI) induced dephosphorylation (14) that negatively regulates this pathway. Other potential substrates that are phosphorylated by activated Aurora A are transforming acidic coiled-coil protein (TACC) (16), centrosomin (CNN) (17), p53 (18), MDB3 (activator of histone deacetylase 1 [HDAC-1]) (19) and BRCA1 (20) which all localize to the centrosome. Several methods to abrogate Aurora A activity have confirmed the assigned roles for this kinase isoform, which include RNA interference

248


(RNAi), which delays mitotic entry (15), anti-Aurora A antibody microinjection into late G2 cells prevents centriole pair separation at prophase (19) and chromosome mal-alignment in the metaphase plate when injected after centriole separation (21). These studies imply additional roles for Aurora A in the later stages of the M-phase. At the end of the G2 /M phase Aurora A is degraded by the anaphasepromoting complex (APC) via the ubiquitin-proteosome pathway (22). In contrast, overexpression of Aurora A compromises the spindle check point (23) and inhibits cytokinesis (24).

2.2. Aurora B: Chromosomal Passenger and Cytokinesis Regulator Normal proliferating cells also express Aurora B with levels peaking at the G2 /M phase of the cell cycle. Aurora B is a ‘chromosomal passenger protein’ (CPP) that forms a quaternary complex with three other CPPs the inner centromere protein (INCENP) (25), survivin (26), and borealin (27), which are required to localize the kinase to centromeres and the spindle with functional interdependence. At the beginning of prophase, Aurora B is decorated along the length of the chromosomes, at metaphase it becomes concentrated at the inner centromere, at anaphase migrates to the central region of the mitotic spindle and finally in telophase to the site of the contractile ring (1,24). Hence, Aurora B is an important regulator of kinetochore function required for correct chromosomal alignment, and segregation as well as spindle checkpoint and cytokinetic functions. Chromosome passenger proteins are substrates of Aurora B, however additional well characterized substrates include histone H3 a structural component of chromatin (28), topoisomerase II a chromosomal scaffolding protein (29) and vimentin, which localizes to the cleavage furrow (30). During M-phase Aurora B phosphorylates histone H3 on serine 10 and this is thought to be important for chromosomal condensation and sister chromatid cohesion (31). Abrogation of Aurora B by gene knockdown or RNAi most likely disrupts the multi-protein chromosomal passenger complex that has structural and catalytic functions, hence affect stoichiometry that disrupts the normal localization and function of binding partners. However, normal cells treated with a SMI to Aurora targeting only its catalytic function demonstrates maloriented chromosomes at anaphase, a defective spindle checkpoint and inhibition of cytokinesis leading to a predictable phenotype of enlarged polyploid cells (32). However, Xenopus egg extracts treated with ZM447439 showed that chromosome


249

condensation was initiated but failed to progress appropriately. Instead the chromosomes underwent premature decondensation during mitotic progression and mitotic spindle assembly was inhibited (33).

2.3. Aurora C: Complements Aurora B Functions In normal tissue Aurora C expression is highest in the reproductive system (sperm and oocytes) (34) but appears to be universally expressed among normal human tissues (35). Aurora C is a CPP similar to Aurora B with expression peaking at G2 /M and later-on in M-phase (36). Evidence that Aurora C has similar functions to Aurora B is provided by overexpression of a kinase-dead form that leads to a multinucleated phenotype whereas kinase-active form corrects for the multinucleated phenotype induced by silencing Aurora B (35,36). Hence, Aurora C functionally complements Aurora B.

3. LINKING AURORA KINASES TO ONCOGENESIS All 3 Aurora kinase genes are localized to known poor prognosis chromosomal amplicons that are frequently amplified in human malignancies (10). Aurora A was established as a proto-oncogene because of its overexpression in human breast cancer cell lines (BTAK, “breast tumor activated kinase”) (37), primary colorectal carcinoma biopsies and its ability to transform rodent cells (NIH3T3 and rat1) by ectopic overexpression and transformed cell induced tumor formation in nude mice (38,39). Further support for its oncogenic role(s) was provided when Xenopus Aurora A transformed NIH 3T3 fibroblasts and led to development of tumors in mice (40). Moreover, Aurora A is overexpressed in a variety of human tumors including pancreatic, ovarian, gastric, breast, colorectal and aggressive B-cell non-Hodgkin’s lymphoma. Overexpression appears to correlate with severity of the grade and may also be associated with stage of the disease (10,11). More recently Aurora A was identified as a candidate low-penetrance tumor-susceptibility gene because of a T-to-A polymorphism leading to an amino acid change from F31I and associated aneuploidy in colon (41) and esophageal cancer (42). However, ectopic overexpression of Aurora A in primary MEF cells alone or with a trans-dominant p53 (eliminates DNA damage checkpoint) or with BCL-2 (eliminates apoptosis) does not induce colony formation (23). Because NIH3T3 or rat1 cells are immortalized with inherent genetic abnormalities it is likely that other defects cooperate with Aurora A to initiate the malignant process. This cooperativity is

250


supported by the observation that Aurora A overexpression potentiates the oncogenic activity of H-ras by influencing cell growth (43) and p53 phosphorylation by Aurora A leads to inhibition of its transactivation activity and degradation (44) with enhanced cell survival. Accumulating data implicate Aurora B to be a proto-oncogene in an appropriate genetic background similar to that observed for Aurora A. One report provides tantalizing data where overexpression of Aurora B in CHE diploid fibroblasts carrying the G245S mutation in both p53 alleles, overrides the p53-dependent G1-checkpoint and when these cells are injected into nude mice form metastases. These murine fibroblasts also had an aneuploid phenotype with an increased histone H3 serine 10 phosphorylation. Further, p53 (G245S) nontransfected cells also form tumors but these did not metastasize (45). In cells that overexpress Aurora A and B defects in p53 appear to play a critical role in stabilizing polyploidy. An extensive systematic analysis of Aurora A, B and C expression levels utilizing mRNA in multiple primary tumor samples of different stages and types (breast, bladder, colon, endometrium, esophagus, liver, lung, ovary, pancreas, prostate, rectum, skin, stomach, and thyroid) demonstrated that Aurora A and B expression levels increase or decrease in parallel. Aurora B overexpression is associated with aneuploidy, higher tumor grade and anaplastic features indicative of enhanced tumor proliferation. However, Aurora C was not overexpressed compared to corresponding normal tissue and did not correlate with either Aurora A or B expression (11). More recent data show that Aurora C is indeed expressed in tumor cell lines (36) and patient biopsy material (46). The functional inter-relationships of the 3 human Auroras in initiating and maintaining malignancy needs further elucidation as feedback mechanisms may play a role in regulating their respective expression.

4. INHIBITORS IN PRE-CLINICAL AND CLINICAL DEVELOPMENT Evidence to date provide a strong rationale for targeting Aurora kinases in human malignancies because overexpression of Aurora A and B in the context of underlying genetic defects transforms cells and induces tumor metastasis respectively. In both cases the catalytic activity of the S/T kinase is essential to tumor development. They are thus effective targets for anticancer drug discovery, design and development for clinical utility in treating malignant disease. Although there are many ways to target the Aurora kinases, selectively targeting


251

the ATP binding site with a SMI and disrupting its enzymatic activity is the most effective and efficient method to proceed toward the clinic. Several chemical classes of SMIs of Aurora kinases have been developed and evaluated for specificity, efficacy and potency utilizing in vitro kinase assays, cell based assays and mouse xenograft models of cancer (47). First and second generation Aurora kinase inhibitors available in the public domain are shown with their pertinent medicinal chemistry profiles (Table 2).

4.1. Dual-Action Aurora Kinase Inhibitors Structure based multiple sequence alignment of the catalytic domains of Aurora (A, B and C) demonstrate that of the 26 residues lining the active site pocket only 3 residues vary (Leu215, Thr217, Arg220) and are all specific to Aurora A (5,7). Hence early drug discovery of Aurora kinase inhibitors were nonselective dual action antitumor compounds that inhibited Aurora A and B/C. However, second generation compounds are being developed that target predominantly Aurora A versus Aurora B/C respectively (48). The first SMIs of Aurora to be described were hesperadin (49,49), ZM447439 (32,32) and VX-680 (50,50). All 3 compounds induced a similar phenotype when evaluated in cell based assays in that there were inhibitions of histone H3 Ser10 phosphorylation and cell division but no effect on cell cycle progression. The end result is a second round of endoreduplication that leads to tetraploidy and subsequent apoptosis or arrest in pseudo-G1 dependent on the status of the p53 post mitotic checkpoint (51). The phenotype observed with Aurora kinase SMIs appear to be consistent with Aurora B inhibition and correlates with S. cerevisiae Aurora Ipl1 that functions to resolve syntelic chromosomal orientations in yeast (52). Aurora B inhibition has no effect on nondividing cells but in dividing cells exposure to hesperadin or ZM447439 leads to several checkpoint proteins (BUB1 and BUBR1) not localizing correctly to kinetochores with associated inhibition of BUBR1 phosphorylation. Despite these effects Aurora kinase SMIs do not totally inhibit checkpoint signaling because treated cells exposed to microtubule toxins that prevent microtubule assembly (nocodazole) undergo mitotic arrest for several hours while in the presence of microtubule stabilizing agents (paclitaxel) there is no mitotic arrest. This is however a paradox because dual-action Aurora SMIs induce a phenotype characteristic of inhibition of Aurora B. The explanation for this observation may be that Aurora A inhibition leads to abnormal

252

Compound Class

Aurora A and aurora B

Aurora A and aurora B (AZ have aurora A specific inhibitors)

MW: 576.67 log Po/w: 4.476 tPSA:121.21

MW: 516.66 log Po/w: 3.752 tPSA: 94.3

Selectivity

Chemical propertiesa

Boehringer Ingleheim

Astra Zeneca

Company

Unknown

Phase 1 in Aug 2005

Clinical status

Table 2 First and second generation Aurora kinase inhibitors available in the public domain with their pertinent medicinal chemistry profiles

253


Aurora A and aurora B and Flt-3

MW: 513.6 log Po/w: 4.27 tPSA:97.85

MW: 464.59 log Po/w: 3.33 tPSA:102.1

Vertex/Merck

Astra Zeneca

(Continued)

Phase 1, Aug 2005

Unknown

254

Compound Class

Aurora A and aurora B and CDK

Aurora A > aurora B

MW: 513.72 log Po/w: 2.99 tPSA: 96.6

MW: 394.35 log Po/w: 0.47 tPSA: 146.0

Selectivity

Chemical propertiesa

Table 2 (Continued) Clinical status

Johnson & Johnson

Phase I 2006

Pfizer/Nerviano PHA-739358 Medical (related to Sciences PHA-680632) Phase II Sep 2006

Company

255

Aurora A and aurora B and Flt-3


ImClone Systems Inc.

Arizona Cancer Center and Supergen

Unknown

Unknown

a The MW and log Po/w were calculated using QikProp (ADME package from Schrodinger - www.schrodinger.com) and the tPSA property was calculated using Molinspiration (www.molinspiration.com)

MW: 468.22 log Po/w: 3.91 tPSA: 92.94

MW: 605.69 log Po/w: 3.45 tPSA: 150.5

256


spindle assembly that can only be observed in cells having a functional spindle checkpoint. In the presence of dual-action SMIs the abnormal spindles arising from Aurora A inhibition may not be visible because in the absence of Aurora B activity they do not trigger mitotic arrest. Mitosis will duly progress with formation of an abnormal spindle until cytokinetic arrest in the absence of Aurora B activity. This leads to a phenotype that appears like the one observed after RNAi knockdown of Aurora B (44). An alternative explanation is that Aurora B inhibition bypasses the requirement for Aurora A activation (53) because Aurora A inhibition activates the BUBR1 checkpoint, which is regulated by Aurora B kinase. However, most tumors overexpress all 3 Aurora kinases and therefore the best anticancer agents are likely to be dualaction inhibitors.

4.2. AstraZeneca (ZM447439 and AZD1152) AstraZeneca pharmaceuticals have pursued the development of Aurora A and B specific inhibitors to determine kinase specific phenotypes and development of novel antiproliferative agents. Molecular and chemical genetic approaches demonstrate that inhibition of Aurora B kinase activity phenocopies ZM447439 (48). Further, AZD1152 (54) and one of its analogs ZM2 (48), which is >100 times selective for Aurora B over Aurora A, induced an identical phenotype that was antiproliferative. AZD1152 also potently suppressed the expression of phosphor-histone H3 in tumor cells and in tumor xenografts in mice (54). Using similar approaches mentioned above simultaneous repression of Aurora A plus induction of a catalytic mutant induces a monopolar phenotype (48). Consistently, ZM3 (analog of AZD1152 and ZM2) (48,54), which is >20 times potent versus Aurora A compared to ZM447439, also induces a monopolar phenotype (48). The expression of a drug-resistant Aurora A mutant reverses the monopolar phenotype, indicating that Aurora A activity drives spindle bipolarity in human cells. These studies with Aurora A and B specific SMIs provides credence to the idea of developing dual-specificity Aurora kinase inhibitors for anticancer drug therapy in human malignancies.

4.3. Vertex-Merck (VX-680) Several Aurora compounds are close to entering the clinic or are in the clinic in phase I dose escalation studies. VX-680 (Vertex


257

Pharmaceuticals) is currently in clinical development. It is a 4,6di-amino pyrimidine that potently inhibits all 3 Aurora kinases (Ki 0.6–4.6 nM). VX-680 is relatively specific but does inhibit Flt-3 (30 nM), Fyn (52 nM), ITK (22 nM), Lck (80nM) and Src (35 nM) (50). It inhibits tumor cell proliferation in a panel of tumor cell lines (breast, cervical, colorectal, leukemia, melanoma, pancreatic and prostate) with EC50 15-113 nM after 4 d of treatment with no effect on viability of noncycling peripheral blood mononuclear cells (10 μM). The compound was also effective in abolishing colony formation in primary leukemic cells refractory to standard chemotherapy and cells possessing Flt-3 internal tandem duplications (35–100 nM). The compound inhibits cell cycle progression and induces apoptosis as determined by annexin V staining. VX680 inhibits tumor growth in xenograft models of leukemia (HL60), colon (HCT-116) and pancreatic (MiaPaCa-2) cancers leading to tumor regression at well tolerated doses when administered by intraperitoneal or intravenous routes. The main toxicity (in rats and not mice) was neutropenia, which was reversible when the drug was discontinued. Histopathological studies showed marked reduction in histone H3 phosphorylation and higher incidence of apoptosis by TUNEL immunohistochemical staining compared to untreated controls (50). VX-680 is currently being evaluated in phase-I clinical studies in solid tumors and acute myeloid leukemias.

4.4. Pfizer-Nerviano Medical Sciences (PHA-680632) PHA-680632 is a 1,4,5,6-tetrahydropyrrolo[3,4-c] pyrazole scaffold based SMI (55) with IC50 ranging from 27, 135, and 120 nM for Aurora A, B and C (56) respectively with minimal cross reactivity to other kinases evaluated. When compared to other Aurora kinase SMIs PHA-680632 phenocopies Aurora B kinase inhibition. The compound showed activity against a wide range of cancer cell lines and had antitumor activity by suppression of tumor growth in mouse xenograft models of leukemia (HL60), ovarian cancer (A2780), colon cancer (HCT116) and mammary carcinoma (MMTV v-Ha-ras transformed cells). Further, immunohistochemical analysis of the tumor biopsy samples for histone H3 phosphorylation as a specific biomarker for PHA-680632 inhibition of Aurora B correlated with the antitumor response (56).

258


4.5. Johnson & Johnson (JNJ-7706621) JNJ-7706621 is a dual cell cycle inhibitor with activity against cyclin-dependent kinases (CDK1, 2, 3, 6) with an IC50 of 3-175 nM and Aurora kinases (A and B) with an IC50 of 11–15 nM respectively (57). The compound was selective in inhibiting proliferation of cancer cell lines of various origins (cervical, colon, ovarian, prostate, melanoma, breast, uterine) irrespective of p53, retinoblastoma (Rb) and MDR-1/Pglycoprotein status and was ˜10-fold less effective in inhibiting normal cell growth (lung fibroblasts, smooth muscle cells, endothelial cells). JNJ-7706621 delayed progression through G1 with growth arrest at the G2 -M phase, induced endoreduplication and inhibited histone H3 serine phosphorylation consistent with Aurora inhibition. Treatment of human A375 melanoma mouse tumor xenograft model with JNJ-7706621 at different doses and schedules indicated a direct correlation between total cumulative dose and antitumor efficacy irrespective of dosing schedules. Daily dosing at 125 mg/kg gave the optimal antitumor response but with increased toxicity (death) without obvious weight loss. Dosing at 7 On / 7 Off or 7 On / 14 Off at 125 mg/kg were less effective as daily dosing with a tumor growth inhibition (TGI) of 93% and 88% respectively. Dosing at 100mg/kg daily was equivalent to 125 mg/kg 7 On /7 Off for TGI (57). The identified dosing schedules provide a rationale for phase-I clinical trials.

4.6. Other Bio-Pharmaceuticals Developing Aurora Kinase Inhibitors Several other novel and potent Aurora kinase ATP-site competitive SMIs have been developed by several bio-pharmaceutical groups demonstrating an intense interest in targeting the Aurora kinases in M-phase of the cell cycle. ImClone Systems have identified a pyrimido-oxazepine as a template that selectively inhibits both Aurora and Flt-3 kinases but their biological activity has not been reported (58). An academic group identified a series of Aurora kinase inhibitors with quinazoline and pyrimidine-based tricyclic scaffolds (59) utilizing homology modeling, in silico fragment search and interactive docking (7,60). The lead compound 4-(6,7-dimethoxy-9H1,3,9-triaza-fluoren-4-yl)-piperazine-1-carbothioic acid [4-9pyrimidin2-ylsulfamoyl)–phenyl]-amide (MP235) demonstrated selectivity for Aurora kinases with a IC50 94 nM, inhibited histone H3 serine phosphorylation in a dose-dependent manner and led to cell cycle arrest in the G2 /M phase (59) consistent with Aurora B inhibition. MP235


259

and derivatives are being developed by Supergen Pharmaceuticals. Several companies (ASTEX Therapeutics, SGX) are now utilizing fragment-based high throughput X-ray crystallography to design novel, potent and effective small molecular inhibitors to protein kinases (61) including the Aurora kinases.

4.7. Aurora Inhibitors in Phase I Clinical Trials Several Aurora kinase inhibitors are now in phase I clinical trials being evaluated for safety, toxicity, dosing, schedule and efficacy in both solid and hematological malignancies (Table 2). The effects of inhibiting the M-phase by Aurora kinase SMIs are more dramatic in transformed cells than normal proliferating cells in cell culture studies. Hence, there is appears to be a therapeutic window for safety and selectivity in animals and humans. Despite these observations it is likely that therapy with Aurora kinase SMIs would lead to myelotoxicity, gastrointestinal toxicity (mucositis & diarrhea) and possibly neuropathy. The most advanced are VX-680, AZD1152, PHA-739358 (related structure to PHA-680632), AT9283, JNJ-7706621, and SNS-314. Interestingly, VX-680 (62) and AT9283 also inhibits BCR-Abl tyrosine kinase (Thr315Ile) mutant enzyme providing beneficial off-target effects in chronic myeloid leukemia. Many novel Aurora kinase SMIs are ready to enter phase-I clinical trials to be evaluated in human malignancies that will provide proofof-principal of the efficacy of inhibiting this target, phase-II dosing, mechanisms of resistance and rational combinations to overcome resistance. Dual CDK/Aurora inhibitors such as JNJ-7706621 are likely to be more effective than those SMIs specifically targeting Aurora kinases.

5. STRUCTURAL BIOLOGY: INSIGHT INTO AURORA INHIBITION Three dimensional structures of Aurora kinases representing several activation states with bound adenine nucleotides with and without interacting proteins provide insights of how this family of enzymes is regulated. More importantly this information is critical to the development of novel SMIs (dual-action and/or specific to Aurora A or B) with good resident times and biological half-lives for cancer therapy. Further, if mutations were to arise in the catalytic residues (e.g., gatekeeper residues) as observed in the tyrosine kinases such as BCRAbl in chronic myeloid leukemia or c-Kit in gastrointestinal stromal

260

Shakalya and Mahadevan (A) N-Terminal Domain

TPX2

Hinge ADP

T-Loop with Ser & Thr Phosphate

C-Terminal Domain

(B)

PHA-680632 T-Loop Disordered

(C) In-Box INCENP

Hesperadin C-Terminal Extension

Fig. 2.

T-Loop with Ser-Phosphate


261

tumors with chronic imatinib mesylate therapy, then structural biology coupled with medicinal chemistry approaches may provide a powerful tool to overcome drug resistance.

5.1. Aurora A The crystal structures of the carboxy terminal catalytic domain of Aurora A to bound adenosine, ADP, drug (PHA-680632) or short peptide fragment (TPX2) from binding proteins have been determined (10). Both Aurora A and B exhibit the well established bilobar protein kinase “fold.” The amino terminal -sheet domain containing a single -helix (C-helix) is implicated in Mg-ATP binding and interacts with kinase regulators. The carboxy terminal -helical domain is the docking site for substrates and is implicated in phosphate transfer. The hinge region connects the 2 domains and the adenosine moiety of ATP binds to the main chain hinge region via hydrogen bond(s) common to all protein kinases. The Mg-ATP and substrate binding sites reside between the 2 domains in a well defined large essentially hydrophobic active site pocket (Fig. 2). The activation loop (T-loop) originating from the carboxy terminal domain has differing conformations depending on the active or inactive states of the enzyme. For Aurora A bound to adenosine (63) the T-loop folds into the active site producing a hydrophobic pocket. However, Aurora A bound to ADP (64) the T-loop is positioned in an active conformation by a bound phosphate ion (crystallization buffer) despite the absence of phosphorylation. Moreover, the crystal structure of Aurora A and Fig. 2. (Continued) Crystal structure of Aurora kinase (A and B) show the classic bilobar protein kinase fold composed of an N-terminal -sheet domain, C-terminal -helical domain and a hinge region that connects the two domains. The Mg2+ ATP and substrate binding sites reside in the pocket between the two domains. (A). Aurora A-TPX2-ADP complex (pdb: 1OL5) shows that the N-terminal segment of TPX2 binds near the N-terminal domain of Aurora A in close proximity to the C-helix and ADP binds in the catalytic pocket; (B). Aurora A bound to the small molecular inhibitor PHA-680632 (pdb: 2BMC) shows that the T-loop is disordered and the drug occupies the large ‘fluorophenyl’ pocket; (C). Aurora B in complex with INCENP and Hesperadin (pdb:2BFY) shows the In-box peptide interacting with the C-terminal extension and the N-terminal domain forming a crown-like motif, the T-loop in an active conformation with a phospho-serine and bound drug that makes no conformational changes compared to unbound Aurora B-INCENP complex.

262


Aurora A-TPX2 complex (65) (pdb: 1OL5) shows that the N-terminal segment of TPX2 binds near the amino terminal domain at a similar location to where cyclins bind CDKs (Fig. 2A). However, TPX-2 binding molds an extended active T-loop conformation, which requires phosphorylation on Thr 287 and Thr 288 in the Aurora consensus target sequence [(R/K)X(T/S)(I/L/V)]. In the absence of the TPX2 peptide, the phospho-Thr 288 is exposed to solvent whereas when bound leads to a buried position locking the active conformation. Aurora A bound to small molecular inhibitors (ZM447439 analog compound 13) (66) and PHA-680632 (67) (pdb: 2BMC) (Fig. 2B) show that the T-loop is disordered but the drug occupies the “fluorophenyl” pocket originally identified in mitogen-activated protein kinases (MAPK) (68).

5.2. Aurora B Crystal structure of Aurora B bound to INCENP (69) sheds insight into the activation of Aurora B by INCENP, which is a 2-step process in which the latter partially activates the former, but full activation requires phosphorylation of a conserved Thr-Ser-Ser (TSS) motif near the carboxy terminus of INCENP (26). In the Aurora B-IN-box (INCENP) complex the latter forms a molecular “crown” extending across (˜70A) perimeter of the amino terminal domain with predominant hydrophobic interactions and extends to make further interactions with the carboxy terminal extension of Aurora B (pdb:2BFY) (Fig. 2C). This complex represents an intermediate state of activation in comparison to the fully active state shown by the Aurora A-TPX2 complex (65) for two reasons. The first is the loss of an ion-pair interaction between Lys 122 and Glu 141, which in the active state forms a buried ion-pair that orients the and phosphates of ATP for phosphotransfer (70). The second is the opening of the catalytic cleft of Aurora B between the 2 domains that is 15° more open than Aurora A. This positions the C-helix and more specifically Glu 141 in the Aurora B-INCENP complex to prevent the interaction with Lys 122 that is necessary for closure of the catalytic cleft and full activity (69). The structure of the Aurora B-INCENP-hesperadin ternary complex compared to the binary complex shows very little conformational change except for the glycine-rich loop that becomes perfectly ordered when drug is bound. Hesperadin slots into the open catalytic pocket of the enzyme where the indolinone moiety binds to the main chain carbonyl and amide of Glu 171 and Ala 173 respectively. Hence, hesperadin binds to an Aurora B conformation where


263

the T-loop is fully extended (69) and this conformation is likely to be clinically relevant for optimal inhibition of the enzyme.

6. CONCLUSIONS AND FUTURE DIRECTIONS Absence or overexpression of Aurora kinases A and B lead to a tetraploid phenotype. However, the mechanism and lethality of tetraploidy are different. Absence of Aurora A leads to spindle defects (monopolar and monoastral bipolar spindles) with formation of tetraploid cells arrested in G2 /M by the postmitotic checkpoint and consequent apoptosis. Therefore, a lack of Aurora A is not well tolerated by cells. Absence of Aurora B leads to overriding of the mitotic checkpoint with inhibition of cytokinesis. This leads to a tetraploid state with 2 centrosomes and avoidance of apoptosis. Hence, a lack of Aurora B is better tolerated by cells. Overexpression of Aurora A (active and inactive kinase) leads to polyploidy and provides a means to escape the p53 dependent postmitotic check point. Overexpression of Aurora B (active and inactive kinase) also induces polyploidy, the inactive enzyme being more efficient than the active enzyme indicating disruption of the stoichiometry of protein-protein interactions. Because cancer cells overexpress both active Aurora A and B simultaneously in an abnormal genetic background (lack or mutated p53 or activated H-ras) it is more than likely that these kinases collaborate to produce a complex polyploid state that leads to transformation (Aurora A) and induction of metastasis (Aurora B). Therefore, inhibiting both Aurora A and B kinases (including Aurora C) simultaneously would be a feasible and rational approach to cancer therapy. Many mechanistic and biological questions remain unanswered at the basic, translational and clinical levels that hopefully will be unraveled in the next few years. At the basic and translational levels understanding exactly how Aurora kinases function and identification of genetic defects that contribute to making Aurora A and B oncogenes in the carcinogenesis process should throw light on how SMIs kill tumors and potential mechanisms of resistance. At the clinical level establishing target inhibition at a safe dose in humans is being evaluated for several SMIs. However, to determine that Aurora activity is being inhibited in human studies should include biomarkers for kinase inhibition rather than the traditional dose finding studies for maximum tolerated dose and dose limiting toxicity. For Aurora B, lack of phosphorylation of histone H3 (Ser 10) could be a useful biomarker whereas for Aurora A additional studies are needed to

264


identify specific biomarkers. Other factors that need clarification are types of tumor most likely to respond to these SMIs, p53 status, kinase selectivity profile and proliferative rate of the tumor. Finally it would be important to understand the pharmacokinetics of Aurora inhibitors when combined with other anticancer agents such as taxanes or vinca alkaloids that interfere with microtubule function and other targeted agents such as CDK inhibitors.

REFERENCES 1. Carmena M, Earnshaw WC. The cellular geography of aurora kinases. Nat Rev Mol Cell Biol 2003;4:842–854. 2. Nigg EA. Mitotic kinases as regulators of cell division and its checkpoints. Nat Rev Mol Cell Biol 2001;2:21–32. 3. Francisco L, Wang W, Chan CS. Type 1 protein phosphotase acts in opposition to Ipl1 protein kinase in regulating yeast chromosome segregation. Mol Cell Biol 1994;14;4731–4740. 4. Petersen J, Paris J, Willer M, Phillipe M, Hagan IM. The S.pombe aurora-related kinase Ark1 associates with mitotic structures in a stage dependent manner and is required for chromosome segregation. J Cell Sci 2001;114:4371–4384. 5. Brown JR, Koretke KK, Birkeland ML, Sanseau P, Patrick DR. Evolutionary relationships of Aurora kinases: implications for model organism studies and the development of anti-cancer drugs. BMC Evol Biol 2004;4(1):39. 6. Giet R, Prigent C. Aurora/Ipl1p-related kinases, a new oncogenic family of mitotic serine-threonine kinases. J Cell Sci 1999;112 (Pt 21): 3591–601. 7. Mahadevan D, Bearss D, Vankayalapati H. Structure Based Design of Novel Anti-Cancer Agents Targeting Aurora Kinase. Curr Med Chem – Anti-Cancer Agents 2003;3:25–34. 8. Largauer C, Kinzler KW, Vogelstein B. Genetic instabilities in human cancers. Nature 1998;396:643–649. 9. Kimura M, Matsuda Y, Yoshioka T, Okano Y. Cell cycle-dependent expression and centrosome localization of a third human aurora/Ipl1related protein kinase, AIK3. J Biol Chem 1999;274:7334–7340. 10. Matthews N, Visintin C, Hartzoulakis B, Jarvis A, Selwood D. Aurora A and B kinases as targets for cancer: will they be selective for tumors? Expert Rev Anticancer Ther 2006;61(1):109–120. 11. Keen N, Taylor S. Aurora-kinase inhibitors as anticancer agents. Nature Reviews Cancer 2004;4:927–936. 12. Adams RR, Carmena M, Earnshaw WC. Chromosomal passengers and the (aurora) ABC’s of mitosis. Trends Cell Biol 2001;11:49–54.


265

13. Kufer TA, Nigg EA, Sillje HH. Regulation of Aurora-A kinase on the mitotic spindle. Chromosoma 2003;112(4):159–163. 14. Tsai MY, Wiese C, Cao K, et al. A Ran signaling pathway mediated by the mitotic kinase Aurora A in spindle assembly. Nat Cell Biol 2003;5(3):242–248. 15. Hirota T, Kunitoku N, Sasayama T, et al.. Aurora A and an interacting activator, the LIM protein Ajuba, are required for mitotic commitment in human cells. Cell 2003;114(5):585–598. 16. Giet R, McLean D, Descamps S, et al. Drosophila Aurora A kinase is required to localize D-TACC to centrosomes and to regulate astral microtubules. J Cell Biol 2002;156(3):437–451. 17. Terada Y, Uetake Y, Kuriyama R. Interaction of Aurora A and centrosomin at the microtubule-nucleating site in Drosophila and mammalian cells. J Cell Biol 2003;162(5):757–763. 18. Liu Q, Kaneko S, Yang L, et al.. Aurora A abrogation of p53 DNA binding and transactivation activity by phosphorylation of serine 215. J Biol Chem 2004;279(50):52175–52182. 19. Sakai H, Urano T, Ookata K, et al.. MBD3 and HDAC1, two components of the NuRD complex, are localized to Aurora A-positive centrosomes in M phase. J Biol Chem 2002;277(50):48714–48723. 20. Ouchi M, Fujiuchi N, Sasai K, et al.. BRCA1 phosphorylation by Aurora A in the regulation of G2 to M transition. J Biol Chem 2004;279(19):19643–19648. 21. Marumoto T, Honda S, Hara T, et al. Aurora-A kinase maintains the fidelity of early and late mitotic events in HeLa Cells. J Biol Chem 2003;278(51):51786–51795. 22. Walter AO, Seghezzi W, Korver W, Sheung J, Lees E. The mitotic serine/threonine kinase Aurora2/AIK is regulated by phosphorylation and degradation. Oncogene 2000;19(42):4906–4916. 23. Anand S, Penrhyn-Lowe S, Venkitaraman AR. AURORA-A amplification overrides the mitotic spindle assembly checkpoint including resistance to Taxol. Cancer Cell 2003;3(1):51–62. 24. Meraldi P, Honda R, Nigg EA. Aurora-A overexpression reveals tetraploidation as a major route to centrosome amplification in p53-/cells. EMBO J 2002;21:483–492. 25. Bolton MA, Lan W, Powers SE, McCleland ML, Kuang J, Stukenberg PT. Aurora B kinase exists in a complex with survivin and INCENP and its kinase is stimulated by surviving binding and phosphorylation. Mol Cell Biol 2002;13(9):3064–3077. 26. Honda R, Korner R, Nigg EA. Exploring the functional interactions between Aurora B, INCENP and survivin in mitosis. Mol Biol Cell 2003;14(8):3325–3341.

266


27. Gassmann R, Carvalho A, Henzing AJ, et al. Borealin: a novel chromosomal passenger required for stability of the bipolar mitotic spindle. J Cell Biol 2004;166(2):179–191. 28. Hsu JY, Sun ZW, Li X, et al. Mitotic phosphorylation of histone H3 is governed by Ipl1/aurora kinase and Glc7/PPI phosphatase in budding yeast and nematode. Cell 2000;102(3):279–291. 29. Morrison C, Henzing AJ, Jensen ON, et al.. Proteomic analysis of human metaphase chromosomes reveals topoisomerase II alpha as an Aurora B substrate. Nucleic Acid Res 2002;30(23):5318–5327. 30. Goto H, Yasui Y, Kawajiri A, et al.. Aurora-B regulates the cleavage furrow-specific vimentin phosphorylation in the cytokinetic process. J Biol Chem 2003;278(10):8526–8530. 31. Giet R, Glover DM. Drosophila aurora B kinase is required for histone H3 phosphorylation and condensin recruitment during chromosome condensation and to organize the central spindle during cytokinesis. J Cell Biol. 2001;152(4):669–682. 32. Ditchfield C, Johnson VL, Tighe A, et al. Aurora B couples chromosome alignment with anaphase by targeting BubR1, Mad2, and Cenp-E to kinetochores. J Cell Biol. 2003;161(2):267–280. 33. Gadea BB, Ruderman JV. Aurora kinase inhibitor ZM447439 blocks chromosome-induced spindle assembly, the completion of chromosome condensation, and the establishment of the spindle integrity checkpoint in Xenopus egg extracts. Mol Biol Cell. 2005;16(3):1305–1318. 34. Tseng TC, Chen SH, Hsu YP, Tang TK. Protein kinase profile of sperm and eggs: cloning and characterization of two novel testis specific protein kinases (AIE1, AIE2) related to yeast and fly chromosome segregation regulators. DNA Cell Biol 1998;17(10):823–833. 35. Yan X, CaoL, LiQ, et al. Aurora C is directly associated with Survivin and required for cytokinesis. Genes Cells. 2005;10(6):617–626. 36. Sasai K, Katayama H, Stenoien D, et al.. Aurora-C kinase is a novel chromosomal passenger protein that can complement Aurora-B kinase function in mitotic cells. Cell Motil Cytoskeleton. 2004;59(4):249–263. 37. Sen S, Zhou H, White RA. A putative serine/threonine kinase encoding gene BTAK on chromosome 20q13 is amplified and overexpressed in human breast cancer cell lines, Oncogene 1997;14(18):2195–2200. 38. Bischoff JR, Anderson L, Zhu Y, et al. A homologue of Drosophila Aurora kinase is oncogenic and amplified in human colorectal cancers. EMBO J 1998;17(11):3052–3065. 39. Zhou H, Kuang J, Zhong L, et al. Tumor amplified kinase STK15/BTAK induces centrosome amplification, aneuploidy and transformation. Nat Genetics 1998;20:189–193. 40. Littlepage LE, Wu H, Andresson T, Deanehan JK, Amundadottir LT, Ruderman JV. Identification of phosphorylated residues that affect


41.

42.

43.

44.

45.

46.

47.

48. 49.

50.

51. 52.

53.

267

the activity of the mitotic kinase Aurora-A. Proc Natl Acad Sci. 2002;99(24):15440–15445. Ewart-Toland A, Briassouli P, de Koning JP, et al. Identification of stk6/STK15 as a candidate low-penetrance tumor-susceptibility gene in mouse and human. Nat Genetics 2003;34:403–412. Miao X, Sun T, Wang Y, Zhang X, Tan W, Lin D. Functional STK15 Phe31Ile polymorphism is associated with the occurrence and advanced disease status of esophageal squamous cell carcinoma. Cancer Res. 2004;64:2680–2683. Tatsuka M, Sato S, Kitajima S, et al. Over-expression of Aurora A potentiates HRAS-mediated oncogenic transformation and is implicated in oral carcinogenesis. Oncogene 2005;24:1122–1127. Giet R, Petretti C, Prigent C. Aurora kinases, aneuploidy and cancer, a coincidence or a real link? TRENDS in Cell Biology 2005;15(5): 241–250. Ota T, Suto S, Katayama H, et al. Increased mitotic phosphorylation of histone H3 attributable to AIM-1/Aurora B over-expression contributes to chromosome number instability. Cancer Res. 2002;62(8): 5168–5177. Takahashi T, Futamura M, Yoshimi N, et al. Centrosomal kinases, HsAIRK1 and HsAIRK3, are overexpressed in primary colorectal cancers. Jpn J Cancer Res. 2000;91(10):1007–1014. Mortlock AA, Keen NJ, Jung FH, et al. Progress in the Development of Selective Inhibitors of Aurora Kinases. Current Topics in Medicinal Chemistry. 2005;5:807–821. Girdler F, Gascoigne KE, Eyers PA, et al. Validating Aurora B as an anti-cancer drug target. J Cell Sci. 2006;119:3664–3675. Hauf S, Cole RW, LaTerra S, et al. The small molecule Hesperadin reveals a role for Aurora B in correcting kinetochore microtubule attachment and in maintaining the spindle assembly checkpoint. J Cell Biol. 2003;161:281–294. Harrington EA, Bebbington D, Moore J, et al. VX-680, a potent and selective small-molecular inhibitor of the Aurora kinases, suppresses tumor growth in vivo. Nat Medicine 2004;10(3):262–267. Margolis RL, Lohez OD, Andreassen PR. G1 tetraploidy checkpoint and the suppression of tumorigenesis. J Cell Biochem. 2003;88:673–683. Tanaka TU, Rachidi N, Janke C, et al. Evidence that the Ipl1-Sli15 (Aurora kinase-INCENP) complex promotes bi-orientation by altering kinetochore-spindle pole correctors. Cell 2002;108:317–329. Yang H, Burke T, Dempsey J, et al. Mitotic requirement for Aurora A kinase is bypassed in the absence of Aurora B kinase. FEBS Lett. 2005;579(16):3385–3391.

268


54. Jung FH, Pasquet G, Lambert-van der Brempt C, et al. Discovery of Novel and Potent Thiazoloquinazolines as Selective Aurora A and B Kinase Inhibitors. J Med Chem. 2006;49:955–970. 55. Fancelli D, Berta D, Bindi S, et al. Potent and selective Aurora inhibitors identified by the expansion of a novel scaffold for protein kinase inhibition. J Med Chem. 2005;48(8):3080–3084. 56. Soncini C, Carpinelli P, Gianellini L, et al. PHA-680632, a novel Aurora kinase inhibitor with potent anti-tumoral activity. Clin Cancer Res. 2006;12(13):4080–4089. 57. Emanuel S, Rugg CA, Gruninger RH, et al. The In vitro and In vivo Effects of JNJ-7706621: A Dual Inhibitor of Cyclin-Dependent Kinases and Aurora Kinases. Cancer Res. 2005;65(19):9038–9046. 58. Pan W, Liu H, Xu Y-J, et al. Pyrimido-oxazepine as a versatile template for the development of inhibitors of specific kinases. Bioorganic & Medicinal Chemistry Letters. 2005;15:5474–5477. 59. Warner SL, Bashyam S, Vankayalapati H, et al. Identification of a lead small-molecule inhibitor of the Aurora kinases using a structure-assisted, fragment-based approach. Mol Cancer Ther. 2006;5(7):1764–1773. 60. Vankayalapati H, Rojanala S, Saldanha J, et al. Targeting Aurora Kinase 2 in Oncogenesis: A Structural Bioinformatics Approach to Target Validation and Rational Drug Design. Mol Cancer Ther. 2003;3:283–294. 61. Gill A, Cleasby A, Jhoti H. The Discovery of Novel Protein Kinase Inhibitors by Using Fragment-Based High-Throughput X-ray Crystallography. ChemBioChem 2005;6:506–512. 62. Young MA, Shah NP, Chao LH, et al. Structure of the kinase domain of an imatinib-resistant Abl mutant in complex with the Aurora kinase inhibitor VX-680. Cancer Res. 2006;66:1007–1014. 63. Cheetham GM, Knegtel RM, Coll JT, et al. Crystal structure of Aurora2, an oncogenic serine/threonine kinase. J Biol Chem. 2002;277(45): 42419–42422. 64. Nowakowski J, Cronin CN, McRee DE, et al. Structures of the cancerrelated Aurora-A, FAK and EphA2 protein kinases from nanovolume crystallography. Structure. 2002;10(12):1659–1667. 65. Bayliss R, Sardon T, Vernos I, Conti E. Structural basis of AuroraA activation by TPX2 at the mitotic spindle. Mol Cell 2003;12(4): 851–862. 66. Heron NM, Anderson M, Bowers DP, et al. SAR and inhibitor complex structure determination of a novel class of potent and specific Aurora kinase inhibitors. Bioorg. Med. Chem. Lett. 2005;16: 1320–1323. 67. Fancelli D, Berta D, Bindi S, et al. Potent and selective Aurora kinase inhibitors identified by the expansion of a novel scaffold for protein kinase inhibition. J. Med. Chem. 2005;48(8):3080–3084.


269

68. Wang Z, CanagarajahBJ, Boehm JC, et al. Structural basis of inhibitor selectivity in MAPKs. Structure. 1998;6(9):1117–1128. 69. Sessa E, Mapelli M, Ciferri C, et al. Mechanism of Aurora B activation by INCENP and inhibition by hesperadin. Mol Cell. 2005;18(3):379–391. 70. Huse M, Kuriyan J. The conformational plasticity of protein kinases. Cell 2002;109:275–282.

12

Plks as Novel Targets for Cancer Drug Design Wei Dai, Yali Yang, and Ning Jiang CONTENTS Introduction Structural Features of Plks Function of Plks Deregulation of Plks in Cancer Compounds Inhibiting Plks Alternative Approaches to Inhibit Plk1 Summary

Abstract Protein tyrosine kinases have served as targets for successful cancer drug development. Protein serine/threonine (Ser/Thr) kinases are more prevalent than protein tyrosine kinases among the molecular entities that regulate cell proliferation. They have been extensively explored to determine their effectiveness as targets for discovery of novel anticancer agents. Polo-like kinases (Plks) comprise a family of conserved proteins that mediate numerous molecular processes critical for cell division. In addition to the highly conserved protein kinase domain in their amino terminal, the Plks share a common structural motif, called the Polo box domain (PBD), in their noncatalytic carboxyl terminal. The PBD functions as a docking site for From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

271

272

Dai, Yang, and Jiang

certain Ser/Thr-phosphorylated proteins, although the structural basis for distinguishing one group of these proteins from another remains unclear. Plks are important regulators of various cell-cycle checkpoints. Deregulation of Plks occurs in a variety of human malignancies. During the past decade, much effort has been directed toward the discovery of novel chemical agents that target Plks specifically. We attempt to summarize recent developments in this field and discuss the structural characteristics of the Plk family that appear to make them uniquely suitable for cancer drug discovery and development. Key Words: Polo-like kinases; plk1; polo-box domain; mitosis; checkpoint; inhibitors; anti-cancer drugs; phosphorylation

1. INTRODUCTION Protein kinases are pivotal in the control of the cell-cycle. In addition to the master regulators, the cyclin-dependent kinases (Cdks), several other families of protein kinases, including the Polo-like kinases (Plks) and the Aurora family of kinases are also essential for cell cycle progression. The Plks regulate numerous cellular and molecular events during cell division (1,2). Polo, the founding member of this kinase family, was identified and characterized in Drosophila melanogaster (3). Its essential role in cell division was established through genetic studies in which Polo mutations were associated with aberrant patterns of mitotic and meiotic division (4). Cdc5 and Plo1, which are structural and functional orthologs of Polo found in budding yeast and fission yeast, respectively, play a critical role in regulating cell division in these low eukaryotes (5,6). The Plks are conserved throughout evolution. Polo homologs have been identified and characterized in every major eukaryotic model system studied thus far (7). Mammalian cells contain 4 proteins (Plk1, Snk/Plk2, Prk/Fnk/Plk3, and Sak/Plk4) (Fig. 1) that are highly homologous to Polo (8–12). Extensive research conducted over the past decade demonstrated that the Plks are major regulators of cell division and that their importance in the cell cycle, especially during mitosis, may rival that of cyclin-dependent kinases. Moreover, deregulation of the expression of Plk1 and other members of the kinase family has been detected in various types of cancer. Given their importance in cell division and their deregulation in malignant cells, much effort has been made to discover and develop small-chemical compounds that can inhibit or modulate Plk activity. In this review, we present a

Chapter 12 / Plks as Novel Targets for Cancer Drug Design Kinase Domain

273

Polo-box Doamin

Plk1

603 a.a. S137 T210

D-Box PB1

Plk2

PB2

685 a.a. S166 T239

Plk3

646 a.a. S107 T219

E4

970 a.a.

Plk4 T169

Fig. 1. Human Plks. Schematic representations of 4 known Plks identified in human cells. Major structural features in Plk1 are indicated, including the kinase domain, PBD, phosphorylation sites, and D-box. Similar structural features in other members of the Polo family are also highlighted. The amino acid sequences encoded by exon 4 (E4) in Plk3 are missing from the major structure in the mouse counterpart. Plk, Polo-like kinase; PBD, Polo-box domain; D-box, destruction box.

summary of recent advances in the discovery of anticancer drugs that use Plks as their molecular targets.

2. STRUCTURAL FEATURES OF PLKS 2.1. The Kinase Domain The protein kinase domain in the N terminal of the Plk is its most highly conserved structure. The sequence G-X-G-XX-A, which is presumed to be involved in ATP-binding in sub-domain I, differs from the typical sequence (G-X-G-XX-G) seen in other Ser/Thr protein kinase families (13). Posttranslational modification of the Plk plays an important role in the regulation of Plk activity. For example, Plk1 is phosphorylated during the cell cycle and the phosphorylation occurs at least at Ser 137 and Thr 210 (T210) within the kinase domain (14). Phosphorylation of Plk1 at T210 in the so-called T-loop appears to activate the kinase activity as T210 mutations (e.g., T210N) that mimic T210 phosphorylation sufficiently induce Plk1 activity (14). The fact that Thr residues equivalent to T210 in Plk1 are also conserved in Plk2, Plk3, and Plk4 (Fig. 1), as well as in the Plks of low eukaryotes (11) suggests the importance of T210 in the regulation of Plk activity. It has been postulated that phosphorylation within the T-loop of the kinase

274


domain of Plk1 allows it to maintain a conformation that is favorable to its interaction with substrates or prevents it from interacting with inhibitors, thereby increasing in its kinase activity.

2.2. The Polo-Box Domain The Polo-box domain (PBD) consists of either 1 or 2 identifiable Polo boxes (15). Although little amino acid homology is observed between Polo boxes, their 3-dimensional conformations are very similar. Elegant studies by the Yaffe group have shown that each Polo box consists of a 6 motif and that the sequence (Polo-cap) at the N-terminal end of Polo-box 1 wraps around Polo-box 2, thus tethering it to the N terminal (15). The same research group also demonstrated that the PBD in all human Plks functions as a docking site for various Ser/Thr-phosphorylated proteins (16). Several studies indicate that the PBD of Plk1 and Plk3 may contain a signal for the subcellular localization of the Plk molecule to centrosomes or spindle poles, given that mutations in conserved residues of the PBD abolish such subcellular localization. (17,18). Alternatively, a phospho-protein(s) at the centrosome recruits Plk1 or Plk3 specifically through the PBD.

2.3. Other Structural Features The region that joins the kinase domain and PBD of Plks is rather diverse and shares little sequence homology. For example, a destruction box (D-box) has been identified in Plk1, but not in Plk2 or Plk3 (19). This is not surprising, because the Plk1 protein level peaks during G2 and M; it is degraded at the end of mitosis and its degradation is dependent on the anaphase-promoting complex (APC) (19). This region may also exhibit functions that have yet to be studied. For example, the primary structure of human Plk3 includes a sequence encoded by exon 4 that is not present in the major form of murine Plk3; otherwise, the human Plk3 sequence is highly homologous with its murine counterpart (11).

3. FUNCTION OF PLKS Plks are important regulators of many cellular events that are essential for cell division, including centrosome duplication and maturation, Golgi apparatus dynamics, responses to DNA damage and stress, DNA repair, the onset of mitosis, bipolar spindle formation, sister chromatid cohesion, chromosome segregation, and cytokinesis

Chapter 12 / Plks as Novel Targets for Cancer Drug Design

275

DNA Damage Checkpoint

DNA Damage Response Centriole Duplication

DNA Repair S

Centrosome Maturation & Seperation

G1 Plks G2

ito tic

Ex

it

y

tr En

M

tic ito

Bipolar Spindle Formation

Golgi Dynamics

M

M

Spindle Checkpoint Activation M Ch itot i Sister Chromatid Separation c ec kp Ex oi it Cytokinesis nt

Ch G ec 2/M kp oi nt

Post-mitotic Functions

Spindle Checkpoint

Fig. 2. Schematic representation of the major functions of Plks during the cell cycle. Major checkpoints that are influenced or regulated by Plks are also indicated. The pie chart is not proportional in terms of the time the cell spends in each phase of the cell cycle.

(Fig. 2). Plk2 and Plk3 also have postmitotic functions that contribute to the regulation of synapse remodeling and synaptic plasticity (20,21). Current antitumor drugs used in clinical settings act primarily by or disrupting the structural integrity of DNA or mitotic processes, and Plks play a major role in checkpoint responses to these events. Therefore, Plks may be ideal targets for the design of anticancer drugs.

3.1. DNA Damage Responses Plk1 activity is inhibited by DNA damage (22–24). The DNA damage-induced inhibition of Plk1 appears to be mediated by blocking its activation because expression of active mutants of Plk1 can override the G2 /M arrest induced by DNA damage (22). This mode of Plk1 inhibition is at least in part dependent on ataxia telangiectasia-mutated (ATM) or A-T and rad3-related (ATR) kinase activity (23). Plk2

276


mRNA expression is induced rapidly in human thyroid cells that have been exposed to X-irradiation; this effect is partly mediated by p53 (25). Plk4 interacts physically with p53, and this interaction is dependent on the PBD (26). Plk3, another important mediator of checkpoint activity in response to DNA damage, is activated by oxidative stress as well as DNA damage. Its activity in response to DNA damage increases rapidly in an ATM-dependent manner (27,28). Plk3 interacts with and phosphorylates p53, targeting its Ser-20 in vitro (27). In separate studies, Plk3 was found to interacts functionally and physically with Chk2 during DNA damage checkpoint activation (28,29). Together, these findings suggest that Plk3 links DNA damage with cell-cycle arrest or apoptosis.

3.2. Mitotic Regulation Plk1 has numerous targets during mitosis. Plk1, as well as Plk3, is required for the completion of mitosis because point mutation, PBD disruption, or N-terminal truncation causes abnormal mitosis and cytokinesis failure (17,18). RNAi-induced inhibition of Plk1 results in the accumulation of cells in a pre-anaphase state or cells with incompletely separated chromatids (30,31). It is rather intriguing to observe that the silencing of Plk1 is correlated with a high level of Cdc2/cyclin B activity (30). In a separate study, RNAi-induced depletion of Plk1 leads to prometaphase arrest (32). A detailed analysis of the timing of mitotic entry indicates that the majority of cells depleted of Plk1 initiate mitotic entry with a short delay (31); thus, Plk1 is apparently not required for mitotic entry. Together, these observations indicate that Plk1 is essential for the progression of mitosis, but it is probably not critical for the onset of mitosis in mammalian cells. The observation that Plk1 targets Cdc25 (33) may reflect the need to maintain the activated state of this phosphatase during early mitosis, which is essential for keeping the activity of Cdk1 high before anaphase entry.

4. DEREGULATION OF PLKS IN CANCER Plk1 undergoes a cyclic change during the cell cycle, whose levels peak at mitosis. Numerous studies show that Plk1 is an excellent tumor marker (34). Overexpression of Plk1 is detected in a majority of human tumor cells, including nonsmall-cell lung cancer, head and neck cancer, breast cancer, colorectal cancer, esophageal and gastric cancer, endometrial cancer, thyroid cancer, and melanomas (34). Increased


277

expression of Plk1 is closely associated with a poor prognosis for cancer patients and increased metastatic potential for certain types of tumors (35,36). It is believed that Plk1 plays a causative role in neoplastic transformation. Interestingly, Plk1 mutations are also detected in several types of cancer and cell lines derived from primary tumors (37). Deregulated expression and structural abnormalities have also been detected in other members of the kinase family. For example, transcriptional silencing of Plk2 is a frequent event in B-cell lymphomas (38); and Plk3 expression is down-regulated in a significant proportion of lung carcinomas (11), head and neck squamous cell carcinomas (39), and colon cancer (40). No mutations were detected in a survey of 40 lung tumor cell lines, but several polymorphisms were identified (41). The loss of heterozygosity of Plk4 is a frequent event in hepatoma at polymorphic markers around the Plk4 locus; reduced activity of Plk4 contributes to enhanced development of cancer as a result of mitotic errors (42). These observations indicate that Plks are aberrantly expressed in malignant cells, which may play a causative role in the genesis of cancer. During the past decade, many categories of novel molecules that affect the activity of Plks have been described. Given that of Plks suppression often leads to mitotic arrest and mitotic catastrophe, these kinases are being considered as excellent targets for anticancer drug discovery.

5. COMPOUNDS INHIBITING PLKS 5.1. Scytonemin Scytonemin is a yellow pigment originally isolated from cyanobacteria that functions in nature as an effective shield against solar radiation in bacteria. It is one of the earliest known compounds capable of providing protection from the effects of ultraviolet (UV) light (43,44). The UV-protective property of scytonemin is provided through its unique structure (45,46)—now referred to as the scytonemin skeleton—which consists of phenolic and indolic subunits (Table 1, Fig. 3) and is believed to be the result of a combination of tryptophan and phenylpropanoid derivatives (44). Several other structural features of scytonemin—including its lack of chirality, multiple dissection points, and phenolic groups—are attractive targets for chemical modifications. Because of its close relationship with antiproliferative

278

In vitro IC50

2.0 μM

N/A

24 nM

24 nM

9–10 nM

0.8 nM

20 nM

Compound

Scytonemin

HMN-214

Wortmannin

LY294002

ON01910

BI2536

Cyclapolin 1

No

Yes

No

Yes

Yes

No

Yes

ATP-Competitive inhibitor

Inhibiting Plks

Inhibiting Plk1, PI3K, PI4K, ATM/ATR, DNA-PK, mTor Inhibiting Plk1, PI3K, PI4K, ATM/ATR, mTor, DNA-PK Inhibiting Plk1, Plk2, PDGFR, Abl, Flt-1, Cdk1, Src, Fyn Inhibiting Plk1

Inhibiting Plk1, Chk1, Myt1, Cdk1/cyclin B, PKC 2 Inhibiting Plks and Cdks

In vitro function

Table 1 Inhibitors of Polo-like kinases

Phase I in patients with advanced solid tumors

Phase I

Phase I in patients with advanced solid tumors

Status of clinical development

36

20,38,53

17

35,64

35,50,66

14,57,59

13,42,51,52

Reference


279

compounds such as nostodione A, scytonemin has been regarded as an attractive candidate for development into an antiproliferative and anticancer agent. Scytonemin is the first chemical compound known to be capable of inhibiting Plk1. It does so through a competitive and noncompetitive mechanism with regard to ATP (45). It also exhibits in vitro inhibition of many other Ser/Thr kinases—including Myt1, Chk1, and Cdk1/cyclin B—with an IC50 value of approximately 2 μM, an effective concentration for inhibiting Plk1 (45). Scytonemin suppresses the growth of a variety of actively proliferating or transformed cells and induces apoptosis (45). The effect of scytonemin on nonproliferating human monocytes has also been studied; at the same concentrations effective for inhibiting proliferating cells, scytonemin does not affect the viability of the monocytes (45). This study strongly suggests that scytonemin specifically targets proliferating cells, making it very attractive for exploration for its therapeutic efficacy in tumor cells.

5.2. HMN-214 HMN-214 ([E]-4-{2-[2-(N-acetyl-N-[4-methoxybenzenesulfonyl] amino) stilbazole}]1-oxide) is a stilbene derivative that blocks the progression of mitosis without significantly affecting the dynamics of microtubules (47). HMN-214 was synthesized as an oral prodrug because of the poor oral absorption of HMN-176, an active metabolite.

Fig. 3. Chemical structures of Polo-like kinase inhibitors (57,60,71).

280


HMN-214 binds with Plk1, thereby interfering with its proper subcellular localization and mitotic function. It remains to be determined whether HMN-214 affects the spatial positioning of other members of the kinase family, given that they share a similar localization pattern (e.g., to centrosomes) during the cell cycle (47). HMN-176 exhibits a potent cytotoxic effect on various human tumor cell lines (47–49). In mouse xenograft models, it has exhibited potent antitumor activity that is at least as effective as that of antitumor drugs currently used in the clinic (48). In preclinical trials, HMN-214 has demonstrated favorable antitumor activity without exhibiting neurotoxicity (50), a side effect frequently observed with microtubule-disrupting agents such as paclitaxel and vincristine. In in vitro studies, HMN214 had demonstrated its ability to down-regulate the expression of MDR1 (47), which suggests that it may not induce common multidrugresistant phenotypes. Because of the promising results seen in preclinical studies, HMN214 was selected for a phase I clinical trial (N = 33) (50). In the 29 patients who completed the study, the best tumor response was characterized by stable disease (n = 7) and a transient decline of carcinoembryonic antigen (n = 1). The recommended dosage is 8 mg/m2 daily, based on the phase I administration schedule (50). Its doselimiting toxicities are bone pain syndrome and hyperglycemia. It has been recommended that patients with prostate and pancreatic cancers be selected for phase II clinical trials, because high levels of Plk1 are usually detected in tumors from these patients.

5.3. Wortmannin and LY294002 Wortmannin is a metabolite originally isolated from the fungus, Pencillium wortmannii, that potently inhibits phosphatidylinositol 3-kinase (PI3K) activity with a low (nanomolar) IC50 (51). LY294002 is a synthetic compound with a structure based on that of the flavanoid quercetin that was specifically designed as a PI3K inhibitor (52). Although LY294002 exhibits a much higher IC50 than wortmannin, it is widely used in in vitro studies because of its stability. Wortmannin targets the ATP-binding site of PI3K and is a competitive inhibitor of ATP binding (53). Although it has been widely considered a specific inhibitor of PI3K and related kinases (e.g., ATM and ATR), a recent study indicates that wortmannin interacts directly with and potently inhibits Plk1 with a IC50 of 24 nM (54). LY294002 also inhibits Plk1 in a manner similar to that of wortmannin (54). Most importantly, the potency of wortmannin against Plk1 is approx 3 orders of magnitude greater than that of scytonemin, and its efficacy in


281

blocking Plk1 activity has been demonstrated in intact cells (54). We have recently observed that wortmannin is equally effective against Plk3, both in vivo and in vitro (55); this suggests that its spectrum of substrate specificity is broader than was originally thought. Given that a significant amount of information is already available regarding the structure and function of wortmannin and LY294002, these compounds can provide valuable information for the further development and optimization of Plk inhibitors for use as targets of anticancer interventions.

5.4. ON01910 ON01910 belongs to a group of novel small-molecule kinase inhibitors that are unrelated to ATP. Known to induce mitotic arrest and exhibits antitumor activity, it has recently been shown to be capable of inhibiting Plk1 kinase activity, as well, at an IC50 value as low as 10 nM (56). At higher concentrations, it also inhibits other protein kinases (56). In an in vitro screening assay, ON01910 induced apoptosis in 94 different tumor cell lines, including multidrug-resistant cell lines, with half-growth inhibition (GI50 ) values of 50 to 200 nM. ON01910-induced cell death appears to result from spindle abnormalities and prolonged G2/M arrest. In murine xenograft models, it inhibits the growth of a wide variety of tumors from humans, including those derived from the liver, breast, and pancreas, and acts synergistically with several chemotherapeutic agents, such as oxaliplatin and doxorubicin, to induce the complete regression of tumor growth. This compound is very well tolerated in animals, who exhibit no apparent side effects such as myelotoxicity, neuropathy, or cardiotoxicity (56). Among the known Plk inhibitors, ON01910 appears to have a unique mode of action. Instead of targeting the ATP-binding pocket of Plk1, in which frequent mutations in the ATP-binding pocket have been observed, it targets the substrate-binding region (56). This property makes it attractive for use in the development of antineoplastic compounds, because compounds that target substrate-binding regions exert less selection pressure for mutations and, thus, may be more stable. Moreover, this category of inhibitors will not compete directly with a very high concentration of ATP. Therefore, it is predicted that this compound will induce drug resistance slowly, if at all.

5.5. BI 2536 BI 2536 is an ATP-dihydropteridinone derivative (57) that potently and specifically inhibits Plk1. In an in vitro study, BI

282


2536 demonstrated high Plk1 specificity, inhibiting this kinase but not the tyrosine kinases or major categories of Ser/Thr kinases, and doing so with an IC50 of 0.8 nM (57–59). BI 2536 causes mitotic arrest leading to apoptosis in a wide range of tumor cells. In murine xenograft models, BI 2536 has exhibited potent inhibition of tumor growth that results in tumor regression. Because of its excellent efficacy profile in in vitro and animal studies, this compound has been used in phase I clinical trials. Preliminary results indicate that it is well tolerated, causing no cumulative or irreversible toxicities. At high doses, however, it can cause neutropenia and thrombocytopenia (58,59). BI 2536 is currently being explored for the possibility of further clinical development.

5.6. Cyclapolin 1 Cyclapolin 1 and its derivatives were discovered through the virtual screening of chemical compound libraries using high-throughput docking software based on a hypothetical structure of the Plk1 kinase domain (60). These compounds share a benzthiazole N-oxide core structure. Cyclapolin 1 is a noncompetitive inhibitor of ATP, even though it was modeled on the ATP-binding pocket in Plk1. It has been hypothesized that a potential covalent interaction between cyclapolin 1 and a residue in the ATP-binding pocket is responsible for its lack of dependence on ATP concentrations. Cyclapolin 1 and its derivatives are highly specific for Plk1. In an extensive study of a panel of 20 protein kinases, cyclapolin 1 did not demonstrate a significant level of inhibitory activity against other major Ser/Thr kinases, including major mitotic kinases (e.g., Aurora A and Cdk1). In fact, the inhibitory concentrations for all of the other kinases in that study exceeded 100 μM, compared with approximately 20 nM for Plk1 (60). Exposure of HeLa cells to cyclapolin 1 resulted in a mitotic arrest that correlated with spindle abnormalities (specifically, collapsed spindles). Its effects can be explained by the observation that it adversely affects the recruitment, activation, and maintenance of the microtubule-nucleating activity of centrosomes. It also exhibits favorable in vitro antitumor activity with a phenotype that is consistent with mitotic arrest and kinase inhibition (60). Interestingly, the phenotype of cells exposed to cyclapolin 1 in vivo is different from that observed in cells depleted of Plk1. Unlike the effect of cyclapolin 1 on intact cells, RNAi-induced Plk1 silencing induces the accumulation of metaphase cells with apparently functional bipolar spindles.


283

6. ALTERNATIVE APPROACHES TO INHIBIT PLK1 During the past decade, alternative molecular approaches to targeting Plk1 have been developed (Table 2). During the mid 1990s, researchers introduced an anti-Plk1 antibody into HeLa cells by means of microinjection, which inhibited Plk1 function and led to mitotic catastrophe (61). The expression of dominant-negative Plk1 through the transduction of recombinant adenovirus also induces apoptosis in most tumor cell lines. Interestingly, normal human mammary epithelial cells exhibit only a minimal apoptotic response when Plk1 is

Table 2 Alternative Approaches to Inhibit Plk1 Approaches to targeting PLKs

Phenotype

Reference

Microinjection of Plk1 antibody

Mitotic catastrophe in HeLa cells; transient G2 arrest in normal human diploid fibroblasts Programmed cell death in most tumor cells but not in normal HMECsa Decreased tumor cell viability in vitro; reduced growth of tumor xenografts in nude mice; no inhibitory effect on primary cells Inhibiting proliferation and inducing significant mitotic arrest and apoptosis in tumor cells but in normal cells Inhibiting proliferation of tumor cells but not of HMECs, normal hTERT-RPE1 or MCF10A cells Inhibiting proliferation of HeLa and A549 tumor cells in vitro and reduced growth of tumor xenografts in vivo

28

DN-Plk1 via adenoviral vector Antisense oligodeoxynucleotides to PLK1

Polo box 1 sequence of Plk1 fused to an antennapedia peptide siRNAs directed against PLK1

PLK1 shRNAs driven by S6 promoter

a

HMEC, human mammary epithelial cells.

14,43

9,48

71

33,48,61

49

284


inhibited (18,62). Specific phosphorothioate antisense oligonucleotides (ASOs) have been designed to inhibit Plk1 expression (63). In fact, selected ASOs effectively inhibit Plk1 expression in tumor cell lines, resulting in a greatly reduced cell viability in vitro and tumorigenesis in vivo (63,64). Attempts have been made to block Plk1 activity through the delivery of a competitive peptide directly into the cell (65). When it is fused to an antennapedia peptide capable of penetrating the plasma membrane, Plk1-PB1 sequences can cause the formation of multiple centrosomes in tumor cells. This is also associated with the inhibition of cell viability and increased cell death (65). Recently, RNAi has been widely used to silence Plk1. Small interfering RNAs (siRNAs) are capable of an effective knock-down or silencing of Plk1; this, in turn, leads to metaphase or pre-anaphase arrest followed by mitotic catastrophe in various cancer cell lines (63,66,67). Consistent with the notion that specificity can be achieved through the perturbation of this kinase, the silencing of Plk1 has little effect on normal cell growth (63). Because of its excellent efficacy in vitro, RNAi is currently being considered as a new strategy for cancer intervention in vivo (68,69). In a recent study, Plk1 shRNAs driven by a U6 promoter efficiently silenced kinase expression for an extended period of time and that silencing is correlated with the suppression of tumor growth in nude mice (70). At present, it remains uncertain whether the RNAi approach results in side effects in vivo because of the possible off-target activity of siRNAs. Nevertheless, it has opened up a new avenue of research for the design and development of antitumor drugs targeting Plk1 and possibly other Plks, as well.

7. SUMMARY Over the past several decades, several categories of new compounds that target Plks have been developed, most of them through the study of Plk1. Some of these compounds are currently being investigated in preclinical and clinical trials. Many of these compounds appear to be superior to existing compounds that are currently widely used as chemotherapeutic agents in clinical settings. Collectively, they exhibit promising features, such as specificity and potency against the target molecules, low cyto- and neurotoxicity, and a reduced potential for developing drug resistance. Given that various mitotic kinases are currently being considered as potential targets of anticancer drug development, structure-function analyses of Plks, especially their PBDs, are expected to lead to the development of a new generation of compounds


285

with better therapeutic indices. Our recent studies show that the Plk3 PBD, when ectopically expressed, exhibits more potent activity than the Plk1 PBD in inducing mitotic catastrophe (17). This suggests that Polo family members other than Plk1 need to be studied, in addition to Plk1, in the drug discovery process. Mitotic catastrophe is a special type of programmed cell death caused in part by the abnormal activation or inactivation of mitotic kinases. Two subtypes of mitotic catastrophe have been identified in recent studies: those that occur during or close to metaphase and those seen after failed attempts to undergo successful mitosis. In both situations, the cell dies of apoptosis as a result of caspase activation and the induction of mitochondrial membrane permeability. Many types of tumor cells have defective mitotic checkpoints, making them more susceptible or sensitive to the inhibition of cell-cycle kinases by initiating apoptosis or mitotic catastrophe. Further study of Plk regulation during mitotic progression will allow us to identify additional targets for the rationale design of drugs for cancer intervention.

ACKNOWLEDGMENTS We thank the members of Dr. Dai’s laboratory for their helpful discussions. We also thank Ms. Heather Carnisi for her editorial and administrative assistance. This work is supported in part by grants from the National Institutes of Health to WD (CA90658 and CA74229).

REFERENCES 1. Barr FA, Sillje HH, Nigg EA. Polo-like kinases and the orchestration of cell division. Nat Rev Mol Cell Biol. 2004;5:429–440. 2. Xie S, Xie B, Lee MY, Dai W. Regulation of cell cycle checkpoints by polo-like kinases. Oncogene. 2005;24:277–286. 3. Fenton B, Glover DM. A conserved mitotic kinase active at late anaphasetelophase in syncytial Drosophila embryos. Nature. 1993;363:637–640. 4. Glover DM, Hagan IM, Tavares AA. Polo-like kinases: a team that plays throughout mitosis. Genes Dev. 1998;12:3777–3787. 5. Kitada K, Johnson AL, Johnston LH, Sugino A. A multicopy suppressor gene of the Saccharomyces cerevisiae G1 cell cycle mutant gene dbf4 encodes a protein kinase and is identified as CDC5. Mol Cell Biol. 1993;13:4445–4457. 6. Ohkura H, Hagan IM, Glover DM. The conserved Schizosaccharomyces pombe kinase plo1, required to form a bipolar spindle, the actin ring, and septum, can drive septum formation in G1 and G2 cells. Genes Dev. 1995;9:1059–1073.

286


7. Dai W. Polo-like kinases, an introduction. Oncogene. 2005;24:214–216. 8. Donohue PJ, Alberts GF, Guo Y, Winkles JA. Identification by targeted differential display of an immediate early gene encoding a putative serine/threonine kinase. J Biol Chem. 1995;270:10351–10357. 9. Hamanaka R, Maloid S, Smith MR et al. Cloning and characterization of human and murine homologues of the Drosophila polo serine-threonine kinase. Cell Growth Differ. 1994;5:249–257. 10. Simmons DL, Neel BG, Stevens R, Evett G, Erikson RL. Identification of an early-growth-response gene encoding a novel putative protein kinase. Mol Cell Biol. 1992;12:4164–4169. 11. Li B, Ouyang B, Pan H et al. Prk, a cytokine-inducible human protein serine/threonine kinase whose expression appears to be down-regulated in lung carcinomas. J Biol Chem. 1996;271:19402–19408. 12. Leung GC, Hudson JW, Kozarova A et al. The Sak polo-box comprises a structural domain sufficient for mitotic subcellular localization. Nat Struct Biol. 2002;9:719–724. 13. Hanks SK, Quinn AM, Hunter T. The protein kinase family: conserved features and deduced phylogeny of the catalytic domains. Science. 1988;241:42–52. 14. Tsvetkov L, Stern DF. Phosphorylation of Plk1 at S137 and T210 is inhibited in response to DNA damage. Cell Cycle. 2005;4:166–171. 15. Elia AE, Rellos P, Haire LF et al. The molecular basis for phosphodependent substrate targeting and regulation of Plks by the Polo-box domain. Cell. 2003;115:83–95. 16. Elia AE, Cantley LC, Yaffe MB. Proteomic screen finds pSer/pThrbinding domain localizing Plk1 to mitotic substrates. Science. 2003;299:1228–1231. 17. Jiang N, Wang X, Jhanwar-Uniyal M, Darzynkiewicz Z, Dai W. Polo box domain of Plk3 functions as a centrosome localization signal, overexpression of which causes mitotic arrest, cytokinesis defects, and apoptosis. J Biol Chem. 2006;281:10577–10582. 18. Seong YS, Kamijo K, Lee JS et al. A spindle checkpoint arrest and a cytokinesis failure by the dominant-negative polo-box domain of Plk1 in U-2 OS cells. J Biol Chem. 2002;277:32282–32293. 19. Lindon C, Pines J. Ordered proteolysis in anaphase inactivates Plk1 to contribute to proper mitotic exit in human cells. J Cell Biol. 2004;164:233–241. 20. Pak DT, Sheng M. Targeted protein degradation and synapse remodeling by an inducible protein kinase. Science. 2003;302:1368–1373. 21. Kauselmann G, Weiler M, Wulff P et al. The polo-like protein kinases Fnk and Snk associate with a Ca(2+)- and integrin-binding protein and are regulated dynamically with synaptic plasticity. EMBO J. 1999;18: 5528–5539.


287

22. Smits VA, Klompmaker R, Arnaud L et al. Polo-like kinase-1 is a target of the DNA damage checkpoint. Nat Cell Biol. 2000;2:672–676. 23. van Vugt MA, Smits VA, Klompmaker R, Medema RH. Inhibition of Polo-like kinase-1 by DNA damage occurs in an ATM- or ATRdependent fashion. J Biol Chem. 2001;276:41656–41660. 24. Ando K, Ozaki T, Yamamoto H et al. Polo-like kinase 1 (Plk1) inhibits p53 function by physical interaction and phosphorylation. J Biol Chem. 2004;279:25549–25561. 25. Shimizu-Yoshida Y, Sugiyama K, Rogounovitch T et al. Radiationinducible hSNK gene is transcriptionally regulated by p53 binding homology element in human thyroid cells. Biochem Biophys Res Commun. 2001;289:491–498. 26. Swallow CJ, Ko MA, Siddiqui NU, Hudson JW, Dennis JW. Sak/Plk4 and mitotic fidelity. Oncogene. 2005;24:306–312. 27. Xie S, Wu H, Wang Q et al. Plk3 functionally links DNA damage to cell cycle arrest and apoptosis at least in part via the p53 pathway. J Biol Chem. 2001;276:43305–43312. 28. Xie S, Wu H, Wang Q et al. Genotoxic stress-induced activation of Plk3 is partly mediated by Chk2. Cell Cycle. 2002;1:424–429. 29. Bahassi eM, Conn CW, Myer DL et al. Mammalian Polo-like kinase 3 (Plk3) is a multifunctional protein involved in stress response pathways. Oncogene. 2002;21:6633–6640. 30. Liu X, Erikson RL. Activation of Cdc2/cyclin B and inhibition of centrosome amplification in cells depleted of Plk1 by siRNA. Proc Natl Acad Sci U S A. 2002;99:8672–8676. 31. Sumara I, Gimenez-Abian JF, Gerlich D et al. Roles of polo-like kinase 1 in the assembly of functional mitotic spindles. Curr Biol. 2004;14: 1712–1722. 32. van Vugt MA, van de Weerdt BC, Vader G et al. Polo-like kinase-1 is required for bipolar spindle formation but is dispensable for anaphase promoting complex/Cdc20 activation and initiation of cytokinesis. J Biol Chem. 2004;279:36841–36854. 33. Kumagai A, Dunphy WG. Purification and molecular cloning of Plx1, a Cdc25-regulatory kinase from Xenopus egg extracts. Science. 1996;273:1377–1380. 34. Takai N, Hamanaka R, Yoshimatsu J, Miyakawa I. Polo-like kinases (Plks) and cancer. Oncogene. 2005;24:287–291. 35. Kanaji S, Saito H, Tsujitani S et al. Expression of polo-like kinase 1 (PLK1) protein predicts the survival of patients with gastric carcinoma. Oncology. 2006;70:126–133. 36. Kneisel L, Strebhardt K, Bernd A et al. Expression of polo-like kinase (PLK1) in thin melanomas: a novel marker of metastatic disease. J Cutan Pathol. 2002;29:354–358.

288


37. Simizu S, Osada H. Mutations in the Plk gene lead to instability of Plk protein in human tumour cell lines. Nat Cell Biol. 2000;2:852–854. 38. Syed N, Smith P, Sullivan A et al. Transcriptional silencing of Polo-like kinase 2 (SNK/PLK2) is a frequent event in B-cell malignancies. Blood. 2006;107:250–256. 39. Dai W, Li Y, Ouyang B et al. PRK, a cell cycle gene localized to 8p21, is downregulated in head and neck cancer. Genes Chromosomes Cancer. 2000;27:332–336. 40. Dai W, Liu T, Wang Q, Rao CV, Reddy BS. Down-regulation of PLK3 gene expression by types and amount of dietary fat in rat colon tumors. Int J Oncol. 2002;20:121–126. 41. Wiest J, Clark AM, Dai W. Intron/exon organization and polymorphisms of the PLK3/PRK gene in human lung carcinoma cell lines. Genes Chromosomes Cancer. 2001;32:384–389. 42. Ko MA, Rosario CO, Hudson JW et al. Plk4 haploinsufficiency causes mitotic infidelity and carcinogenesis. Nat Genet. 2005;37:883–888. 43. Garcia-Pichel F, Sherry ND, Castenholz RW. Evidence for an ultraviolet sunscreen role of the extracellular pigment scytonemin in the terrestrial cyanobacterium Chlorogloeopsis sp. Photochem Photobiol. 1992;56: 17–23. 44. Proteau PJ, Gerwick WH, Garcia-Pichel F, Castenholz R. The structure of scytonemin, an ultraviolet sunscreen pigment from the sheaths of cyanobacteria. Experientia. 1993;49:825–829. 45. Stevenson CS, Capper EA, Roshak AK et al. The identification and characterization of the marine natural product scytonemin as a novel antiproliferative pharmacophore. J Pharmacol Exp Ther. 2002;303: 858–866. 46. Stevenson CS, Capper EA, Roshak AK et al. Scytonemin–a marine natural product inhibitor of kinases key in hyperproliferative inflammatory diseases. Inflamm Res. 2002;51:112–114. 47. Tanaka H, Ohshima N, Ikenoya M et al. HMN-176, an active metabolite of the synthetic antitumor agent HMN-214, restores chemosensitivity to multidrug-resistant cells by targeting the transcription factor NF-Y. Cancer Res. 2003;63:6942–6947. 48. Takagi M, Honmura T, Watanabe S et al. In vivo antitumor activity of a novel sulfonamide, HMN-214, against human tumor xenografts in mice and the spectrum of cytotoxicity of its active metabolite, HMN-176. Invest New Drugs. 2003;21:387–399. 49. Medina-Gundrum L, Cerna C, Gomez L, Izbicka E. Investigation of HMN-176 anticancer activity in human tumor specimens in vitro and the effects of HMN-176 on differential gene expression. Invest New Drugs. 2005;23:3–9. 50. Garland LL, Taylor C, Pilkington DL, Cohen JL, Von Hoff DD. A phase I pharmacokinetic study of HMN-214, a novel oral stilbene derivative with


51. 52.

53. 54.

55.

56.

57. 58.

59.

60. 61.

62.

63.

64.

65.

289

polo-like kinase-1-interacting properties, in patients with advanced solid tumors. Clin Cancer Res. 2006;12:5182–5189. Wipf P, Halter RJ. Chemistry and biology of wortmannin. Org Biomol Chem. 2005;3:2053–2061. Vlahos CJ, Matter WF, Hui KY, Brown RF. A specific inhibitor of phosphatidylinositol 3-kinase, 2-(4-morpholinyl)-8-phenyl-4H-1benzopyran-4-one (LY294002). J Biol Chem. 1994;269:5241–5248. Stein RC. Prospects for phosphoinositide 3-kinase inhibition as a cancer treatment. Endocr Relat Cancer. 2001;8:237–248. Liu Y, Shreder KR, Gai W et al. Wortmannin, a widely used phosphoinositide 3-kinase inhibitor, also potently inhibits mammalian polo-like kinase. Chem Biol. 2005;12–99-107. Liu Y, Jiang N, Wu J, Dai W, Rosenblum JS. Polo-like kinases inhibited by wortmannin. Labeling site and downstream effects. J Biol Chem. 2007;282:2505–2511. Gumireddy K, Reddy MV, Cosenza SC et al. ON01910, a non-ATPcompetitive small molecule inhibitor of Plk1, is a potent anticancer agent. Cancer Cell. 2005;7:275–286. Strebhardt K, Ullrich A. Targeting polo-like kinase 1 for cancer therapy. Nat Rev Cancer. 2006;6:321–330. Munzert G, Steinbild S, Frost A, Hedbrn S, Rentschler J, Kraiser D et al. A phase I study of two administration schedules of the Polo-like kinase 1 inhibitor BI 2536 in patients with advanced solid tumors. J Clin Oncol 24 (18S), 3069. 2006. Hofheinz R, Hochhaus A, Al-Batran A, Nanci V, Renchardt V, Trommeshauser D et al. A phase I repeated dose escalation study of the Polo-like kinase inhibitor BI 2536 in patients with advanced solid tumours. J Clin Oncol 24 (18S), 2038. 2006. McInnes C, Mazumdar A, Mezna M et al. Inhibitors of Polo-like kinase reveal roles in spindle-pole maintenance. Nat Chem Biol. 2006;2: 608–617. Lane HA, Nigg EA. Antibody microinjection reveals an essential role for human polo-like kinase 1 (Plk1) in the functional maturation of mitotic centrosomes. J Cell Biol. 1996;135:1701–1713. Cogswell JP, Brown CE, Bisi JE, Neill SD. Dominant-negative polo-like kinase 1 induces mitotic catastrophe independent of cdc25C function. Cell Growth Differ. 2000;11:615–623. Spankuch B, Heim S, Kurunci-Csacsko E et al. Down-regulation of Pololike kinase 1 elevates drug sensitivity of breast cancer cells in vitro and in vivo. Cancer Res. 2006;66:5836–5846. Elez R, Piiper A, Giannini CD, Brendel M, Zeuzem S. Polo-like kinase1, a new target for antisense tumor therapy. Biochem Biophys Res Commun. 2000;269:352–356. Yuan J, Kramer A, Eckerdt F, Kaufmann M, Strebhardt K. Efficient internalization of the polo-box of polo-like kinase 1 fused to an Antennapedia

290

66. 67. 68.

69.

70.

71.

Dai, Yang, and Jiang peptide results in inhibition of cancer cell proliferation. Cancer Res. 2002;62:4186–4190. Liu X, Erikson RL. Polo-like kinase (Plk)1 depletion induces apoptosis in cancer cells. Proc Natl Acad Sci U S A. 2003;100:5789–5794. van Vugt MA, Medema RH. Polo-like kinase-1: activity measurement and RNAi-mediated knockdown. Methods Mol Biol. 2005;296:355–369. Guan R, Tapang P, Leverson JD et al. Small interfering RNA-mediated Polo-like kinase 1 depletion preferentially reduces the survival of p53defective, oncogenic transformed cells and inhibits tumor growth in animals. Cancer Res. 2005;65:2698–2704. Nogawa M, Yuasa T, Kimura S et al. Intravesical administration of small interfering RNA targeting PLK-1 successfully prevents the growth of bladder cancer. J Clin Invest. 2005;115:978–985. Spankuch B, Matthess Y, Knecht R et al. Cancer inhibition in nude mice after systemic application of U6 promoter-driven short hairpin RNAs against PLK1. J Natl Cancer Inst. 2004;96:862–872. Jiang N, Wang X, Yang Y, Dai W. Advances in mitotic inhibitors for cancer treatment. Mini Rev Med Chem. 2006;6:885–895.

13

Do Histone Deacetylase Inhibitors Target Cell Cycle Checkpoints that Monitor Heterochromatin Structure? Brian Gabrielli, Frankie Stevens, and Heather Beamish CONTENTS Introduction Cell Cycle Responses to HDACi Treatment HDACi Kill Tumor Cells by Synthetic Lethality Molecular Mechanism of HDACi-Induced Aberrant Mitosis Why is There an HDACi Sensitive G2 Checkpoint?

Abstract HDACi are showing promise as anticancer drugs owing to their ability to block proliferation and promote differentiation or apoptosis in a wide range of cancer cell lines, and inhibit tumor growth in vivo. The antiproliferative effects of HDACis are caused by drug induced G1 or G2 phase cell cycle arrests. They also promote aberrant mitosis, which in cycling cells can result in the rapid From: Cancer Drug Discovery and Development Checkpoint Responses in Cancer Therapy Edited by: W. Dai © Humana Press, Totowa, NJ, a part of Springer Science+Business Media

291

292

Gabrielli, Stevens, and Beamish

onset of apoptosis. Hyperacetylation of the constitutive centromeric heterochromatin upon HDACi treatment is believed to disrupt the normal functioning of the kinetochore associated spindle assembly checkpoint machinery. The G2 phase checkpoint arrest appears to be in response to perturbation of heterochromatin structure, thereby ensuring genomic stability. The ability of cells to detect and respond to perturbations in heterochromatin structure is likely to be critical in determining cell fate, and its disruption in cancers would permit the silencing and unsilencing of genes often observed in modified transcriptomes of cancer cells as compared to their normal tissue counterparts. Key Words: Histone deacetylase inhibitor; heterochromatin; checkpoint; synthetic lethality; chromosomal passenger proteins; spindle assembly checkpoint

1. INTRODUCTION Histone deacetylase inhibitors (HDACis) are a structurally diverse family of drugs that inhibit the class-I and class-II histone deacetylases (HDACs) with relatively little selectivity for specific isoforms (1). HDACi treatment of cells inhibits the deacetylation of a wide range of proteins including transcription factors, and the resulting hyperacetylated proteins affect a broad spectrum of cellular events. Transcriptional changes underlie many of the altered cellular responses to HDACis, although HDACi effects that are independent of altered transcription are being identified. These include affecting the stability of proteins by stabilizing the acetylation of HSP90 thereby inhibiting its chaperone activity (2). A number of HDACis are currently undergoing clinical trials as single agents, and are showing promise for specific clinical indications (3–6). Specific cancer types are likely to be highly susceptible to HDACis, particularly where dysregulation of transcription factors are a significant contributor to the transformed phenotype (7). Other nontranscriptional targets of HDACi may also provide tumor selective action, for example in cancers where HSP90 stabilization of transforming oncogenes is a major contributing factor (2,8). HDACis also show promise in combination with other traditional chemotherapeutic drugs, although in many cases the basis for this synergy is not well understood (9). This chapter will be restricted to a review of our current understanding of the effects of HDACi treatment upon the cell

Chapter 13 / Do HDACis Target Cell Cycle Checkpoints?

293

cycle, the molecular mechanisms by which these cell cycle effects are imposed, and how this may be utilized to provide tumor selective cytotoxicity by targeting cell cycle checkpoints that are defective in tumor cells.

2. CELL CYCLE RESPONSES TO HDACI TREATMENT The most common consequence of HDACi treatment of cells is reduced proliferation, usually related to G1 or G2/M phase cell cycle arrests, but may also be associated with differentiation or increased apoptosis. HDACi also promote aberrant mitosis, which results in the rapid onset of apoptosis. These cell cycle effects are dependent on the cell lines being examined, and the dose of HDACi used. Transcriptional changes appear to be responsible for the G1 phase arrest and these effects are observed at low doses of HDACi that cause only modest increases in histone acetylation (10,11). This G1 arrest is the most common cell cycle response to HDACi treatment and is observed in the majority of cell lines tested, including nonimmortalized primary cells. The G2 arrest is less commonly observed, and generally requires a relatively high dose of HDACi which causes extensive histone hyperacetylation (10,12) (Fig. 1). The mitotic defects are only observed in cells that fail to arrest in G2 phase with high doses of HDACi, although the mechanisms by which HDACis affect G2 and mitosis are not yet fully understood.

2.1. HDACi-Induced G1 Phase Arrest 2.1.1. G1/S Phase Progression Progression through the cell cycle is controlled by the ordered activation of a family of protein kinases known as the cyclin dependent kinases (cdks). Individual cdks associate with a limited repertoire

Fig. 1. Low doses of HDACi initiate a G1 phase arrest in the majority of cell lines, whereas high doses of HDACi are required to affect G2 phase and mitosis, and these effects correspond to the selective cytotoxicity of HDACi.

294


of cyclins at specific stages of the cell cycle, and the activity of discrete cyclin/cdks complexes control progression through each cell cycle stage. Cell cycle progression from G1 into S phase requires the sequential activation of cyclin D/cdk4 or cdk6, and cyclin E/cdk2, although cyclin E/cdk2 can perform both functions in cells that lack cyclin D/cdk4 activity (13). These two cdk complexes phosphorylate Rb to allow full activation of the Rb-bound E2F, a transcription factor required for the expression of an array of genes required for progression into and though S phase (reviewed in (14)). Cyclin E/cdk2 and cyclin A/cdk2 also have critical roles in the initiation and maintenance of DNA replication (15–17). 2.1.2. HDACi Induces Expression of p21WAF1/CIP1 and Other cdk Inhibitors HDACi treatment of the majority of cell lines examined results in a G1 phase cell cycle arrest. The G1 arrest is associated with the inhibition or loss of the G1 phase cdk/cyclin complexes responsible for Rb phosphorylation resulting in hypophosphorylation of Rb, loss of the S phase population and DNA synthesis (12,18,19). The arrest is a consequence of HDACi-induced regulation of a number of genes that control G1/S phase progression. The most prominent of the upregulated genes is the cdk inhibitor p21WAF1/CIP1 . HDACi treatment increases acetylation of chromatin in the p21WAF1/CIP1 promoter region thereby allowing increased access to an Sp1/Sp3 site that is correlated with increased gene transcription (10,18,20,21), however additional modifications to the transcriptional machinery also appear to be required for full transcriptional activation to occur (22,23). P21WAF1/CIP1 up-regulation is independent of the tumor suppressor p53. In the DNA damage response, p53 is downstream of the cell cycle checkpoint signaling complex containing ATM (24,25). Although HDACi treatment does not appear to produce DNA strand breaks, there does appear to be a role for ATM, independent of p53, in the HDACiinduced up-regulation of p21WAF1/CIP1 expression, as this response is reduced both in ATM defective cell lines and in the presence of caffeine (26). However, where ATM activation has been assessed by examination of ATM autophosphorylation on S1981, there was no evidence of ATM activation following HDACi treatment (27). Thus the role of ATM in p21WAF1/CIP1 up-regulation remains controversial, as does the idea that this is a bone fide checkpoint response. Increased p21WAF1/CIP1 levels bind and inhibit cyclin E and cyclin A/cdk2 complexes that are responsible for phosphorylation and inacti-


295

vation of Rb and essential for progression into and through S phase. Deletion of the p21WAF1/CIP1 gene or failure of HDACi treatment to up-regulate p21WAF1/CIP1 expression, possibly because of caspase dependent destruction of p21WAF1/CIP1 , correlates with a lack of G1 phase arrest (19,28,29). There are also reports of other cdk inhibitors such as the cdk4 and cdk6 specific p15INK4B and p19INK4D inhibitors and the p21WAF1/CIP1 -related p57KIP2 inhibitor being up-regulated in various cell lines with HDACi treatment (30–33). These cdk inhibitors may contribute to a G1 phase arrest in some cell lines by inhibiting either the cdk4/cyclin D complexes by the INK4 family proteins, or cdk2/cyclin E and cdk2/cyclin A complexes by the p21WAF1/CIP1 – related proteins (34). The INK4 proteins can also cause redistribution of p21WAF1/CIP1 family proteins off cdk4 complexes where they have little inhibitory activity in vivo, onto cdk2 complexes of which they are potent inhibitors, thereby reinforcing the G1 arrest (35). The upregulation of p21WAF1/CIP1 is widely observed in both normal and tumor cell lines (12,29), and is one of the most commonly up-regulated genes in response to diverse HDACis in multiple cell lines (36). It is not clear how common the up-regulation of the other cdk inhibitor genes is following HDACi treatment. 2.1.3. Other G1 Phase Targets The up-regulation of p21WAF1/CIP1 and other cdk inhibitors may not be the sole cause of the G1 arrest observed. Loss of expression of cyclin D and cyclin A are also commonly observed following HDACi treatment and are likely to contribute to the loss of cdk2 and cdk4 kinase activities leading to the hypophosphorylated Rb (12,18,19,37,38). Surprisingly, HDACi treatment also increases cyclin E expression although this is not associated with increased cdk2 activity, presumably through inhibition of the kinase because of increased p21WAF1/CIP1 levels (18,19,37) (Fig. 2A). Genes involved in DNA synthesis such as CTP synthase, thymidylate synthetase and thymidine kinase have been reported as targets for HDACi-induced down-regulation (32,36). Loss of these enzymes would have a similar effect to treatment with antimetabolites such as hydroxyurea, blocking S phase progression and thereby contributing to the G1/S arrest. The resulting reduced nucleotide pools also effect p21WAF1/CIP1 levels reinforcing the G1 arrest, possibly as part of a checkpoint response (39) (Fig. 2B). It is clear that many mechanisms combine to impose the observed HDACi-induced G1 phase arrest, all of which involve transcriptional changes.

296

Gabrielli, Stevens, and Beamish (A)

(B)

Fig. 2. A. G1/S transition is regulated by cdk4/cyclin D and cdk2/cyclin E kinase activities, whereas cdk2/cyclin A is required for S phase progression. HDACi treatment up-regulates the expression of the cdk2 inhibitor p21WAF1/CIP1 and cdk4 inhibitor p19INK4D while down-regulating cyclin D and cyclin A levels, thereby blocking Rb phosphorylation and promoting the G1 phase arrest. Cyclin E levels are up-regulated but the increased p21WAF1/CIP1 expression can block the activity of this complex. B. HDACi treatment down regulates the expression of genes required for expansion of the nucleotide pools required for S phase, thus imposing a G1/S arrest that is dependent on p21WAF1/CIP1 expression.

Up to 20% of genes are affected by HDACi treatment, the percentage being dependent on how the expression changes are defined (36,40), but it is clear that only a very small proportion of these changes have any significance in the cell cycle changes observed. A common feature is up-regulation of p21WAF1/CIP1 , as is the regulation of genes that could contribute to a G1 arrest (36). Thus the G1 arrest


297

is a consequence of transcriptional changes in response to HDACi treatment, although the mechanisms used to impose the arrest may be HDACi and cell line dependent. Interestingly, HDACi treatment during S phase did not appear to affect the time taken to transit through S phase, suggesting that drug treatment had little direct effect on DNA replication (12).

2.2. Up-Regulated Expression of p21WAF1/CIP1 Reduces HDACi Induced Cytotoxicity The antiproliferative activity of HDACis is a result of cell cycle arrest but also of increased apoptosis. The up-regulated expression of p21WAF1/CIP1 and associated G1 arrest are observed at relatively low doses of drug, whereas higher doses, which do not significantly further increase p21WAF1/CIP1 levels, are associated with cytotoxicity (10,11,29). However, in a small subset of cell lines where HDACi treatment fails to up-regulate p21WAF1/CIP1 protein expression, low doses of drug can achieve high levels of apoptosis (29). Similarly, knockout of p21WAF1/CIP1 expression increases the sensitivity of cells to killing by HDACis (29,41). Other studies have shown that other drugs that block p21WAF1/CIP1 expression increased the cytotoxicity of HDACis (42–46). The hypersensitivity to HDACis was reversed by treating with inhibitors of apoptosis (29), suggesting that increased p21WAF1/CIP1 expression reduces HDACi cytotoxicity by blocking apoptosis. p21WAF1/CIP1 is also a substrate of caspase 3, and cleavage of p21WAF1/CIP1 appeared to destroy its antiapoptotic effect, thus the kinetics of p21WAF1/CIP1 expression and caspase 3 activation may determine the relative sensitivity of cells to HDACi treatment (29). The INK4 family cdk4 inhibitor p16 has also been reported to block HDACi-induced apoptosis in a leukemic cell line (47,48). However, this effect was not reproduced in other cell lines with either stable or inducible p16 expression as these cells remained sensitive to HDACiinduced cell death (49). The decreased sensitivity was not because of a block in proliferation, as HDACi treatment induced cell death even in nonproliferating cells.

2.3. HDACi-Induced G2 Phase Checkpoint G1 phase arrest is detected in majority of cells following HDACi treatment, whereas G2 phase arrest is observed only at high doses of HDACi (10,19,29), and in comparatively few cell lines (12). Cell lines that arrest in G2 phase are resistant to the cytotoxic effects of

298


these drugs (10,12,50). In these studies it is important to demonstrate where in the cell cycle cells are arrested, G2 phase or mitosis, by other methods together with single dimension FACS analysis demonstrating increased cells with 4n DNA content, as this could represent cells that have undergone aberrant mitosis and failed cytokinesis to produce G1 phase cells with 4n DNA (12). This is especially relevant for HDACi as treatment of cells is capable of inducing aberrant mitosis (see the following). 2.3.1. Mechanism of the HDACi-sensitive G2 phase checkpoint The mechanism of the G2 checkpoint imposed arrest appears to be dependent on the timing of HDACi addition with addition of the drug in S phase sufficient to produce the G2 arrest (10,12,51). This HDACi-sensitive G2 checkpoint appears to operate independently of the DNA damage G2 checkpoint, as DNA damaging agents can initiate a G2 checkpoint arrest in cells that failed to arrest in response to HDACi treatment (12). However, the HDACi-sensitive checkpoint does appear to utilize the caffeine-sensitive ATM/ATR checkpoint signaling pathway common to other agents (50), although the lack of detectable ATM activation suggests that ATR may be responsible for signaling the arrest (27). In common with other G2 checkpoint mechanisms, HDACi treatment blocks the activation of cyclin B/cdk1 and down-regulates cyclin B1 protein levels (12,51–53). The expression of Gadd45, a growth arrest and DNA damage inducible gene that can cause a G2/M arrest by inhibiting cdc2/cyclin B activity (54), has been found to be up-regulated following HDACi treatment and may also participate the G2 arrest observed (55). A second G2 checkpoint mechanism is used upon HDACi treatment when cells are in antephase, immediately prior to nuclear membrane breakdown. This arrest is independent of ATM, instead utilizing the p38MAPK-MAPKAPK2-cdc25B pathway to inhibit entry into mitosis (27,56). This mechanism, the antephase checkpoint, appears to be separate from the G2 phase checkpoint initiated in response to S phase HDACi addition. 2.3.2. The HDACi-Sensitive G2 Checkpoint Distinguishes HDACi Resistant Cells from HDACi Sensitive Cells In the majority of immortalized, virally transformed or tumor cell lines, the HDACi sensitive G2 checkpoint is defective. Correlative evidence from a large panel of cell lines treated with high dose


299

HDACi demonstrated that the presence of a G2 phase checkpoint arrest 24 h after drug treatment was always associated with resistance to cell killing (12). Furthermore, reintroducing a G2 checkpoint arrest in HDACi-sensitive cells rescues the cells from HDACi-induced apoptosis, whereas knockout of the ATM/ATR-dependent G2 checkpoint mechanism by expression of the Epstein-Barr virus EBNA 3 family proteins converts an HDACi-resistant cell line to an HDACi sensitive cell line (12,50). Thus, the functional status of the HDACisensitive G2 checkpoint determines the tumor-selective cytotoxicity of these drugs. Nonimmortalized primary cells with a functional G2 checkpoint are resistant to killing by even high doses of HDACi, whereas this checkpoint is defective in the majority of immortalized, virally transformed and tumor cell lines tested, which are as a consequence sensitive to killing by these drugs.

2.4. HDACi Treatment Disrupts Mitosis and the Spindle Assembly Checkpoint A number of studies have demonstrated that HDACi treatment results in aberrant mitosis, with the condensed chromosomes failing to congress to the metaphase plate thereby segregating unequally into the two daughter cells (12,57–62). These cell lines have a defective HDACi-sensitive G2 checkpoint, as cells with an intact checkpoint do not progress into mitosis in the presence of drug. The molecular basis of the mitotic defect is becoming clearer although the hyperacetylated proteins that are responsible for the defect are yet to be elucidated. The aberrant mitosis initiates the spindle assembly checkpoint that blocks anaphase chromosome segregation until all of the chromosomes have achieved bipolar attachment to mitotic spindle (63). There is some controversy about whether the spindle assembly checkpoint is activated under these conditions, however, all research groups reported delays in mitosis suggesting that some checkpoint function is maintained. My own laboratory has observed Mad2 loading on the unaligned chromosomes (Stevens et al., in press), but others have reported a loss of hBub1 and BubR1 staining to kinetochores (62,64). Nevertheless, in all studies the checkpoint fails as cells exit mitosis without correctly separating their sister chromatids (57,62) (Stevens et al., in press). Levels of mitotic histone H3 Ser10 phosphorylation are unaffected by HDACi treatment (60,62), although binding of the heterochromatin protein HP1 to the centromeric regions is reduced (58,60,62). There are effects on the expression of Plk1 (51), and the chromosomal passenger proteins Aurora B (62), Borealin, INCENP and Survivin,

300


which, although expressed at normal levels, fail to accumulate at the centromere in mitosis (Stevens et al., in press). The outcome of the mitotic defects is dependent on the dose and timing of HDACi treatment. Treatment with relatively low doses of HDACis that cause little cytotoxicity, cause cells to delay in mitosis (57,59,60), presumably under the influence of the spindle assembly checkpoint, resulting in limited cytokinesis disruption. The timing of drug addition is also critical. The HDACi is required to present at least through S phase to produce the aberrant mitotic phenotype as addition of high doses of drug in early G2 phase failed to produce aberrant mitosis, whereas exposure to high doses of HDACi during S phase followed by wash off produced high levels of aberrant mitosis although the cells were unable to overcome the spindle checkpoint arrest (57).

3. HDACI KILL TUMOR CELLS BY SYNTHETIC LETHALITY The consequence of HDACi induced aberrant mitosis is rapid apoptosis upon exit from mitosis. The premature mitotic exit is a requirement for apoptosis as blocking mitotic exit inhibited apoptosis (57). The cell death is consequence of synthetic lethality. This concept, originally used to describe yeast mutations, defines mutations of different genes that individually may reduce viability to a minor extent, but when combined with a second mutation results in complete loss of viability (65). The two mutant genes often operate in compensatory pathways, with loss of function in one pathway being compensated for by the activity of the second gene pathway. In the case of the HDACi, the loss of the G2 phase checkpoint in HDACi treated cells forces cells to undergo aberrant mitosis, which activates the spindle assembly checkpoint to delay mitotic exit until the defects in mitosis are rectified. With high doses of HDACi the spindle assembly checkpoint is eventually overcome and cells aberrantly exit mitosis and rapidly initiate apoptosis. Extending the mitotic delay decreases the apoptosis observed, demonstrating that the premature mitotic exit is responsible for the apoptosis observed (57). Thus the cell death observed upon HDACi treatment is the consequence of the combination of the defective HDACi-sensitive G2 checkpoint, an intrinsic feature of the cancer cell, and HDACi-induced bypass of the compensating spindle assembly checkpoint. Normal cells are protected from cell death by their intact G2 checkpoint, and do not proceed into mitosis until the drug has been removed and the hyperacetylation reversed (Fig. 3).


301

Fig. 3. HDACi resistant cells have an intact HDACi-sensitive G2 checkpoint which blocks progression into mitosis. In cells sensitive to killing by HDACi, the G2 checkpoint is defective, and the cells undergo an aberrant mitosis. This would normally be detected by the spindle assembly checkpoint, but the drugs eventually bypass this arrest and prematurely exit mitosis resulting in either multinuclear cells that are likely to have limited proliferative potential, or rapid onset of apoptosis.

4. MOLECULAR MECHANISM OF HDACI-INDUCED ABERRANT MITOSIS This leaves unanswered questions: how does HDACi treatment trigger the G2 checkpoint response, and how does HDACi treatment cause the disruption or bypass of the subsequent spindle assembly checkpoint in cell lines with a defective G2 checkpoint? It is clear that transcriptional changes underlie the G1 phase arrest, and this is likely to account for the cytostatic activity of HDACis, but this requires only relatively low drug doses which do not promote significant cell death. Up-regulation of p21WAF1/CIP1 expression is detected at these low doses (10,29), suggesting that many of the HDACi dependent transcriptional changes may be elicited by low drug concentrations. The contribution of HDACi induced transcriptional changes to the G2 and mitotic defects is less clear. Although changes in the expression of key G2/M regulators have been reported (12,18,66), the effects of HDACi on G2/M requires higher doses of drug, suggesting that the effects may be because of hyperacetylation of nonhistone proteins (12,57,61). One potential target is -tubulin which is acetylated at Lys40 (67), but the mitotic defects observed upon HDACi treatment are independent of tubulin acetylation (57,61). A clue to the target(s) maybe the very distinctive aberrant mitosis resulting from HDACi treatment. This is a phenocopy of mutations of kinetochore and centromeric proteins (68–71), suggesting that HDACi treatment disrupts normal centromere/kinetochore function. The most obvious effect of the drugs is hyperacetylation of the normally hypoacetylated centromeric heterochromatin, possibly disrupting its

302


higher order structure. HDACi treatment does not effect either the expression or localization of centromere-specific proteins such as the histone H3-like CENP-A (12,58), nor affect mitotic H3 Ser10 phosphorylation which is required for entry into mitosis and, in some organisms, proper chromosome segregation (60,72). HDACi treatment does, however, result in loss of HP1 binding to the heterochromatin (58,73), which is required for the higher order structure of the centromeric heterochromatin. Deletion of SUV39H1, the methyltransferase responsible for histone H3 Lys 9 methylation which acts as the binding site for HP1, or mutation of the H3 Lys 9 residue both result in aberrant mitosis (74–76). Thus HDACi treatment may instigate aberrant mitosis and disrupt kinetochore function by hyperacetylating the centromeric heterochromatin thereby blocking HP1 binding. A centromere specific HDAC containing Sin3 co-repressor complex has have been described in both yeast and mammals, and depletion of Sin3 affects chromosome segregation in mitosis (77,78). This centromere specific HDAC containing complex may explain the requirement for higher HDACi doses to affect G2 and mitosis if it is less sensitive to inhibition by HDACi. The cell cycle dependent timing of HDACi addition on G2 and mitosis may be explained by examining the acetylation state of the heterochromatin. Heterochromatin is normally hypoacetylated, with transient acetylation during deposition of histones onto newly replicated DNA in S phase. This contrasts with the continuously changing acetylation state of the actively transcribed euchromatin which is readily affected by low doses of HDACi (79–81). Together, these observations provide compelling evidence that the centromeric heterochromatin is the likely target of the HDACi stabilized acetylation that causes the mitotic effects of these drugs.

5. WHY IS THERE AN HDACI SENSITIVE G2 CHECKPOINT? The existence of numerous checkpoint pathways that ensure the integrity of genomic DNA emphasizes the paramount importance the cell places on ensuring genomic integrity. The genome contains all possible genes and other noncoding regulatory sequences that the whole organism requires for normal growth and development. The large number of modifications that chromatin can undergo, and the diverse biological outcomes of these modifications demonstrate that changes to the heterochromatin can radically affect the biology of the cell (82,83). Thus, whereas the DNA codes for all possible genes and


303

thereby all possible cell fates, the heterochromatin defines the potential transcriptome in each cell by silencing unwanted genes, thereby determining cell fate and function. It is certain that such a critical function would be protected by checkpoint responses to ensure the fidelity of this process. Equally, loss of this checkpoint function would be expected to be a common feature of cancer, permitting changes in heterochromatinization of genes with the consequent silencing or unsilencing which may underlie many of the gene expression changes detected in cancer versus normal tissue of origin. This checkpoint would also be critical in G2/M progression where the specialized centromeric heterochromatin has an essential function ensuring the fidelity of partitioning the replicated genome by influencing the activity of the kinetochore associated spindle assembly checkpoint complexes. This would underlie the G2 checkpoint. However, the mechanism that detects changes in heterochromatinization also appears to operate in noncycling cells. In this case it also signals apoptosis but only in cells with a defective HDACi-sensitive checkpoint, the only difference from cycling cells is the time to onset of apoptosis is delayed by 24 h (49). The more rapid onset of apoptosis in actively cycling cells is a consequence of the disrupted centromeric heterochromatin and failed mitosis. The trigger for apoptosis in the nonproliferating cells is at present unknown. Thus there is good evidence that HDACi treatment blocks heterochromatin histone deacetylation to produce its clinically important tumor-selective cytotoxicity, but rather than this necessarily being via changes in transcription, may be through changes in heterochromatin structure and function.

ACKNOWLEDGMENTS This work was supported by funding from the National Health and Medical Research Council of Australia.

REFERENCES 1. Marks PA, Richon VM, Breslow R, Rifkind RA. Histone deacetylase inhibitors as new cancer drugs. Curr Opin Oncol 2001;13(6):477–483. 2. Drysdale MJ, Brough PA, Massey A, Jensen MR, Schoepfer J. Targeting Hsp90 for the treatment of cancer. Curr Opin Drug Discov Devel 2006;9(4):483–495. 3. Kelly WK, Marks PA. Drug insight: Histone deacetylase inhibitors– development of the new targeted anticancer agent suberoylanilide hydroxamic acid. Nat Clin Pract Oncol 2005;2(3):150–157.

304


4. Fouladi M, Furman WL, Chin T, et al. Phase I study of depsipeptide in pediatric patients with refractory solid tumors: a Children’s Oncology Group report. J Clin Oncol 2006;24(22):3678–3685. 5. Ryan QC, Headlee D, Acharya M, et al. Phase I and pharmacokinetic study of MS-275, a histone deacetylase inhibitor, in patients with advanced and refractory solid tumors or lymphoma. J Clin Oncol 2005;23(17):3912–3922. 6. Reid T, Valone F, Lipera W, et al. Phase II trial of the histone deacetylase inhibitor pivaloyloxymethyl butyrate (Pivanex, AN-9) in advanced nonsmall cell lung cancer. Lung Cancer 2004;45(3):381–386. 7. Minucci S, Pelicci PG. Histone deacetylase inhibitors and the promise of epigenetic (and more) treatments for cancer. Nat Rev Cancer 2006;6(1):38–51. 8. Aoyagi S, Archer TK. Modulating molecular chaperone Hsp90 functions through reversible acetylation. Trends Cell Biol 2005;15(11):565–567. 9. Karagiannis TC, El-Osta A. The paradox of histone deacetylase inhibitormediated modulation of cellular responses to radiation. Cell Cycle 2006;5(3):288–295. 10. Richon VM, Sandhoff TW, Rifkind RA, Marks PA. Histone deacetylase inhibitor selectively induces p21WAF1 expression and gene-associated histone acetylation. Proc Natl Acad Sci U S A 2000;97(18):10014–10019. 11. Gabrielli B, Warrener R, Burgess A, Beamish H. Defining the chemotherapeutic targets of histone deacetylase inhibitors. Ann N Y Acad Sci 2004;1030:627–635. 12. Qiu L, Burgess A, Fairlie DP, Leonard H, Parsons PG, Gabrielli BG. Histone deacetylase inhibitors trigger a G2 checkpoint in normal cells that is defective in tumor cells. Mol Biol Cell 2000;11(6):2069–2083. 13. Gray-Bablin J, Zalvide J, Fox MP, Knickerbocker CJ, DeCaprio JA, Keyomarsi K. Cyclin E, a redundant cyclin in breast cancer. Proc Natl Acad Sci U S A 1996;93(26):15215–15220. 14. Sherr CJ. The Pezcoller lecture: cancer cell cycles revisited. Cancer Res 2000;60(14):3689–3695. 15. Ohtsubo M, Theodoras AM, Schumacher J, Roberts JM, Pagano M. Human cyclin E, a nuclear protein essential for the G1-to-S phase transition. Mol Cell Biol 1995;15(5):2612–2624. 16. Pagano M, Pepperkok R, Verde F, Ansorge W, Draetta G. Cyclin A is required at two points in the human cell cycle. Embo J 1992;11(3): 961–971. 17. Strausfeld UP, Howell M, Descombes P, et al. Both cyclin A and cyclin E have S-phase promoting (SPF) activity in Xenopus egg extracts. J Cell Sci 1996;109(Pt 6):1555–1563. 18. Sambucetti LC, Fischer DD, Zabludoff S, et al. Histone deacetylase inhibition selectively alters the activity and expression of cell cycle


19.

20.

21.

22.

23.

24.

25.

26. 27.

28.

29.

30. 31.

305

proteins leading to specific chromatin acetylation and antiproliferative effects. J Biol Chem 1999;274(49):34940–3494. Sandor V, Senderowicz A, Mertins S, et al. P21-dependent g(1)arrest with downregulation of cyclin D1 and upregulation of cyclin E by the histone deacetylase inhibitor FR901228. Br J Cancer 2000;83(6):817–825. Huang L, Sowa Y, Sakai T, Pardee AB. Activation of the p21WAF1/CIP1 promoter independent of p53 by the histone deacetylase inhibitor suberoylanilide hydroxamic acid (SAHA) through the Sp1 sites. Oncogene 2000;19(50):5712–5719. Xiao H, Hasegawa T, Isobe K. p300 collaborates with Sp1 and Sp3 in p21(waf1/cip1) promoter activation induced by histone deacetylase inhibitor. J Biol Chem 2000;275(2):1371–1376. Kim YK, Han JW, Woo YN, et al. Expression of p21(WAF1/Cip1) through Sp1 sites by histone deacetylase inhibitor apicidin requires PI 3-kinase-PKC epsilon signaling pathway. Oncogene 2003;22(38): 6023– 6031. Gui CY, Ngo L, Xu WS, Richon VM, Marks PA. Histone deacetylase (HDAC) inhibitor activation of p21WAF1 involves changes in promoterassociated proteins, including HDAC1. Proc Natl Acad Sci U S A 2004;101(5):1241–1246. Canman CE, Lim DS, Cimprich KA, et al. Activation of the ATM kinase by ionizing radiation and phosphorylation of p53. Science 1998;281(5383):1677–1679. Khanna KK, Keating KE, Kozlov S, et al. ATM associates with and phosphorylates p53: mapping the region of interaction. Nat Genet 1998;20(4):398–400. Ju R, Muller MT. Histone deacetylase inhibitors activate p21(WAF1) expression via ATM. Cancer Res 2003;63(11):2891–2897. Mikhailov A, Shinohara M, Rieder CL. Topoisomerase II and histone deacetylase inhibitors delay the G2/M transition by triggering the p38 MAPK checkpoint pathway. J Cell Biol 2004;166(4):517–526. Archer SY, Meng S, Shei A, Hodin RA. p21(WAF1) is required for butyrate-mediated growth inhibition of human colon cancer cells. Proc Natl Acad Sci U S A 1998;95(12):6791–6796. Burgess AJ, Pavey S, Warrener R, et al. Up-regulation of p21(WAF1/CIP1) by histone deacetylase inhibitors reduces their cytotoxicity. Mol Pharmacol 2001;60(4):828–837. Hitomi T, Matsuzaki Y, Yokota T, Takaoka Y, Sakai T. p15(INK4b) in HDAC inhibitor-induced growth arrest. FEBS Lett 2003;554(3):347–350. Sato N, Fukushima N, Maitra A, et al. Discovery of novel targets for aberrant methylation in pancreatic carcinoma using high-throughput microarrays. Histone acetylation-mediated regulation of genes in leukaemic cells. Gene expression profiling of multiple histone deacetylase

306

32.

33.

34. 35.

36.

37.

38.

39.

40.

41.

42.

Gabrielli, Stevens, and Beamish (HDAC) inhibitors: defining a common gene set produced by HDAC inhibition in T24 and MDA carcinoma cell lines. Cancer Res 2003;63(13):3735–3742. Mitsiades CS, Mitsiades NS, McMullan CJ, et al. Transcriptional signature of histone deacetylase inhibition in multiple myeloma: biological and clinical implications. Proc Natl Acad Sci U S A 2004;101(2):540–545. Yokota T, Matsuzaki Y, Miyazawa K, Zindy F, Roussel MF, Sakai T. Histone deacetylase inhibitors activate INK4d gene through Sp1 site in its promoter. Oncogene 2004;26:26. Sherr CJ, Roberts JM. Inhibitors of mammalian G1 cyclin-dependent kinases. Genes Dev 1995;9(10):1149–1163. McConnell BB, Gregory FJ, Stott FJ, Hara E, Peters G. Induced expression of p16(INK4a) inhibits both CDK4- and CDK2-associated kinase activity by reassortment of cyclin-CDK-inhibitor complexes. Mol Cell Biol 1999;19(3):1981–1989. Glaser KB, Staver MJ, Waring JF, Stender J, Ulrich RG, Davidsen SK. Gene expression profiling of multiple histone deacetylase (HDAC) inhibitors: defining a common gene set produced by HDAC inhibition in T24 and MDA carcinoma cell lines. Mol Cancer Ther 2003;2(2):151–163. Kim YB, Lee KH, Sugita K, Yoshida M, Horinouchi S. Oxamflatin is a novel antitumor compound that inhibits mammalian histone deacetylase. Oncogene 1999;18(15):2461–2470. Wharton W, Savell J, Cress WD, Seto E, Pledger WJ. Inhibition of mitogenesis in Balb/c-3T3 cells by Trichostatin A. Multiple alterations in the induction and activation of cyclin-cyclin-dependent kinase complexes. J Biol Chem 2000;275(43):33981–33987. Deng C, Zhang P, Harper JW, Elledge SJ, Leder P. Mice lacking p21CIP1/WAF1 undergo normal development, but are defective in G1 checkpoint control. Cell 1995;82(4):675–684. Peart MJ, Smyth GK, van Laar RK, et al. Identification and functional significance of genes regulated by structurally different histone deacetylase inhibitors. Proc Natl Acad Sci U S A 2005;102(10): 3697–3702. Vrana JA, Decker RH, Johnson CR, et al. Induction of apoptosis in U937 human leukemia cells by suberoylanilide hydroxamic acid (SAHA) proceeds through pathways that are regulated by Bcl-2/Bcl-XL, c-Jun, and p21CIP1, but independent of p53. Oncogene 1999;18(50):7016–7025. Rosato RR, Wang Z, Gopalkrishnan RV, Fisher PB, Grant S. Evidence of a functional role for the cyclin-dependent kinase-inhibitor p21WAF1/ CIP1/MDA6 in promoting differentiation and preventing mitochondrial dysfunction and apoptosis induced by sodium butyrate in human myelomonocytic leukemia cells (U937). Int J Oncol 2001;19(1):181–191.


307

43. Rahmani M, Yu C, Dai Y, et al. Coadministration of the heat shock protein 90 antagonist 17-allylamino- 17-demethoxygeldanamycin with suberoylanilide hydroxamic acid or sodium butyrate synergistically induces apoptosis in human leukemia cells. Cancer Res 2003;63(23): 8420–8427. 44. Rosato RR, Almenara JA, Yu C, et al. The cyclin-dependent kinase inhibitor flavopiridol disrupts sodium butyrate-induced p21WAF1/CIP1 expression and maturation while reciprocally potentiating apoptosis in human leukemia cells. Mol Pharmacol 2004;65(3):571–581. 45. Rahmani M, Yu C, Reese E, et al. Inhibition of PI-3 kinase sensitizes human leukemic cells to histone deacetylase inhibitor-mediated apoptosis through p44/42 MAP kinase inactivation and abrogation of p21(CIP1/WAF1) induction rather than AKT inhibition. Oncogene 2003;22(40):6231–6242. 46. Rosato RR, Almenara JA, Grant S. The histone deacetylase inhibitor MS-275 promotes differentiation or apoptosis in human leukemia cells through a process regulated by generation of reactive oxygen species and induction of p21CIP1/WAF1 1. Cancer Res 2003;63(13):3637–3645. 47. Bernhard D, Ausserlechner MJ, Tonko M, et al. Apoptosis induced by the histone deacetylase inhibitor sodium butyrate in human leukemic lymphoblasts. Faseb J 1999;13(14):1991–2001. 48. Peart MJ, Tainton KM, Ruefli AA, et al. Novel mechanisms of apoptosis induced by histone deacetylase inhibitors. Cancer Res 2003;63(15): 4460–4471. 49. Burgess A, Ruefli A, Beamish H, et al. Histone deacetylase inhibitors specifically kill nonproliferating tumour cells. Oncogene 2004;23(40):6693–6701. 50. Krauer KG, Burgess A, Buck M, Flanagan J, Sculley TB, Gabrielli B. The EBNA-3 gene family proteins disrupt the G2/M checkpoint. Oncogene 2004;23(7):1342–1353. 51. Lallemand F, Courilleau D, Buquet-Fagot C, Atfi A, Montagne MN, Mester J. Sodium butyrate induces G2 arrest in the human breast cancer cells MDA-MB-231 and renders them competent for DNA rereplication. Exp Cell Res 1999;247(2):432–440. 52. Goldstone S, Pavey S, Forrest A, Sinnamon J, Gabrielli B. Cdc25dependent activation of cyclin A/cdk2 is blocked in G2 phase arrested cells independently of ATM/ATR. Oncogene 2001;20(8):921–932. 53. Maity A, Hwang A, Janss A, Phillips P, McKenna WG, Muschel RJ. Delayed cyclin B1 expression during the G2 arrest following DNA damage. Oncogene 1996;13(8):1647–1657. 54. Jin S, Antinore MJ, Lung FD, et al. The GADD45 inhibition of Cdc2 kinase correlates with GADD45-mediated growth suppression. J Biol Chem 2000;275(22):16602–16608.

308


55. Hirose T, Sowa Y, Takahashi S, et al. p53-independent induction of Gadd45 by histone deacetylase inhibitor: coordinate regulation by transcription factors Oct-1 and NF-Y. Oncogene 2003;22(49):7762–7773. 56. Manke IA, Nguyen A, Lim D, Stewart MQ, Elia AE, Yaffe MB. MAPKAP kinase-2 is a cell cycle checkpoint kinase that regulates the G2/M transition and S phase progression in response to UV irradiation. Mol Cell 2005;17(1):37–48. 57. Warrener R, Beamish H, Burgess A, et al. Tumor cell-selective cytotoxicity by targeting cell cycle checkpoints. Faseb J 2003;17(11):1550–1552. 58. Taddei A, Maison C, Roche D, Almouzni G. Reversible disruption of pericentric heterochromatin and centromere function by inhibiting deacetylases. Nat Cell Biol 2001;3(2):114–120. 59. Shin HJ, Baek KH, Jeon AH, et al. Inhibition of histone deacetylase activity increases chromosomal instability by the aberrant regulation of mitotic checkpoint activation. Oncogene 2003;22(25):3853–3858. 60. Cimini D, Mattiuzzo M, Torosantucci L, Degrassi F. Histone hyperacetylation in mitosis prevents sister chromatid separation and produces chromosome segregation defects. Mol Biol Cell 2003;14(9):3821–3833. 61. Blagosklonny MV, Robey R, Sackett DL, et al. Histone deacetylase inhibitors all induce p21 but differentially cause tubulin acetylation, mitotic arrest, and cytotoxicity. Mol Cancer Ther 2002;1(11):937–941. 62. Robbins AR, Jablonski SA, Yen TJ, et al. Inhibitors of histone deacetylases alter kinetochore assembly by disrupting pericentromeric heterochromatin. Cell Cycle 2005;4(5):717–726. 63. Musacchio A, Hardwick KG. The spindle checkpoint: structural insights into dynamic signalling. Nat Rev Mol Cell Biol 2002;3(10):731–741. 64. Dowling M, Voong KR, Kim M, Keutmann MK, Harris E, Kao GD. Mitotic spindle checkpoint inactivation by trichostatin a defines a mechanism for increasing cancer cell killing by microtubule-disrupting agents. Cancer Biol Ther 2005;4(2):197–206. 65. Kaelin WG, Jr. The concept of synthetic lethality in the context of anticancer therapy. Nat Rev Cancer 2005;5(9):689–698. 66. Noh EJ, Lee JS. Functional interplay between modulation of histone deacetylase activity and its regulatory role in G2-M transition. Biochem Biophys Res Commun 2003;310(2):267–273. 67. Piperno G, LeDizet M, Chang XJ. Microtubules containing acetylated alpha-tubulin in mammalian cells in culture. J Cell Biol 1987;104(2): 289–302. 68. Mackay AM, Ainsztein AM, Eckley DM, Earnshaw WC. A dominant mutant of inner centromere protein (INCENP), a chromosomal protein, disrupts prometaphase congression and cytokinesis. J Cell Biol 1998;140(5):991–1002.


309

69. Nishihashi A, Haraguchi T, Hiraoka Y, et al. CENP-I is essential for centromere function in vertebrate cells. Dev Cell 2002;2(4):463–476. 70. Schaar BT, Chan GK, Maddox P, Salmon ED, Yen TJ. CENP-E function at kinetochores is essential for chromosome alignment. J Cell Biol 1997;139(6):1373–1382. 71. Taylor SS, McKeon F. Kinetochore localization of murine Bub1 is required for normal mitotic timing and checkpoint response to spindle damage. Cell 1997;89(5):727–735. 72. Prigent C, Dimitrov S. Phosphorylation of serine 10 in histone H3, what for? J Cell Sci 2003;116(Pt 18):3677–3685. 73. Ekwall K, Olsson T, Turner BM, Cranston G, Allshire RC. Transient inhibition of histone deacetylation alters the structural and functional imprint at fission yeast centromeres. Cell 1997;91(7):1021–1032. 74. Rea S, Eisenhaber F, O’Carroll D, et al. Regulation of chromatin structure by site-specific histone H3 methyltransferases. Nature 2000;406(6796):593–599. 75. Bannister AJ, Zegerman P, Partridge JF, et al. Selective recognition of methylated lysine 9 on histone H3 by the HP1 chromo domain. Nature 2001;410(6824):120–124. 76. Mellone BG, Ball L, Suka N, Grunstein MR, Partridge JF, Allshire RC. Centromere silencing and function in fission yeast is governed by the amino terminus of histone H3. Curr Biol 2003;13(20):1748–1757. 77. David G, Turner GM, Yao Y, Protopopov A, DePinho RA. mSin3associated protein, mSds3, is essential for pericentric heterochromatin formation and chromosome segregation in mammalian cells. Genes Dev 2003;17(19):2396–2405. 78. Silverstein RA, Richardson W, Levin H, Allshire R, Ekwall K. A new role for the transcriptional corepressor SIN3; regulation of centromeres. Curr Biol 2003;13(1):68–72. 79. Sobel RE, Cook RG, Perry CA, Annunziato AT, Allis CD. Conservation of deposition-related acetylation sites in newly synthesized histones H3 and H4. Proc Natl Acad Sci U S A 1995;92(4):1237–1241. 80. Taddei A, Roche D, Sibarita JB, Turner BM, Almouzni G. Duplication and maintenance of heterochromatin domains. J Cell Biol 1999;147(6):1153–1166. 81. Mello JA, Almouzni G. The ins and outs of nucleosome assembly. Curr Opin Genet Dev 2001;11(2):136–141. 82. Dillon N, Festenstein R, Cheung WL, Briggs SD, Allis CD. Unravelling heterochromatin: competition between positive and negative factors regulates accessibility. Acetylation and chromosomal functions. Trends Genet 2002;18(5):252–258. 83. Jenuwein T, Allis CD. Translating the histone code. Science 2001;293(5532):1074–1080.

Index 1,27-O-acetylokadaic acid, 122 1,4-benzodiazepine-2,5-dione, 39 13-hydroxy-15-ozoapathin, 122 17-allylamino-17demethoxygeldanamycin, 123 27-O-acetyldinophysistoxin, 122 4-chlorobenzoic acid 4-methoxyphenyl ester, 122 5-aza-dC, 169 5-fluorouracil, 6 5,11-diketoindenoisoquinolines, 67 7-hydroxystaurosporine (UCN-01), 7

Bortezimib, 153 Bradyarrythmias, 189 Butyrolactone, 141

Actinomycin D, 191 Alkylating agents, 94, 119 Alopecia, 189 Amphiphilic, 29 Amplicon, 242 AMT and A-T and Rad3 related (ATR), 94 Anaphase promoting complex (APC), 5, 224, 269 Anaphase-promoting complex/cyclosome (APC/C), 164 Aneuploidy, 192, 224, 239 Angiogenesis, 21 Ansamycin, 8 Antagonists, 33 Anthracyclines, 191 Antimetabolites, 119 Antimitogenic, 3 Aphidicolin, 99 Arrhythmia, 145 Arthralgias, 189 Ataxia telangiectasia mutated (ATM), 93, 275 ATM-interaction protein (ATMIN), 100 Base excision repair, 118 Benzodiazepinedione, 39 Benzoquinone, 70 Benzsulfonamide, 41 Bioavailability, 148 Bioflavonoids, 69 Bleomycin, 58, 72, 96

Caffeine, 108, 122 Camptotheca acuminata, 66 Camptothecin, 66, 99 Cellular gatekeeper, 20 Centriole separation, 248 Centrosome, 165, 178 Centrosome duplication, 270 Chalcones, 34 Chaperone, 27 Checkpoint, 74 Checkpoint abrogators, 119 Checkpoint dysfunction, 170 Chlorambucil, 39 Chlorofusin, 34 Chloroindolyl sulphonamide, 146 Chromokinesins, 211 Chromosome missegregation, 211, 224 Chromosome passenger, 226 Chromosome passenger protein (CPP), 239 Chromosome segregation, 270 Cis-imidazoline, 34 Cisplatin, 5, 58, 170 Cleavage furrow, 248 Clostridium elegans, 21 Cohesin, 225 Colchicines, 191 Crystallography, 22 Cyclapolin, 277 Cycloheximide, 26 Cyclophosphamide, 5 Cyclosome, 225 Cytokinesis, 209, 225, 244, 259, 270 Cytosine arabinoside, 39 Cytotoxic, 119 Cytotoxicity, 293 Debromohymenialdisine, 122 Destruction box, 269 Diarrhea, 196

311

312 Diflomotecan, 67 Dihydropyrazole, 218 Dihydropyrrole, 217 Dimethylenastron, 215 Dinophysistoxin, 122 Docetaxel, 144, 165, 277 Drosophila melanogaster, 21, 240 Druggable, 108 E3 ubiquitin ligase, 44, 102, 166 Eg5, 207 Embryogenesis, 21 Enastron, 215 Endocytosis, 194 Environmental mutagens, 118 Epigenetic, 135 Epipodophyllotoxins, 191 Epothilone, 194 Equilibrium, 24 Erythrodysesthesias, 190 Etoposide, 68 Euchromatin, 298 Farnesyl transferase, 210 FAT and FATC domains, 94 FHA domain, 165 Flavopiridol, 9, 108, 141 Fludarabine, 39 Fork-head associated (FHA), 98 Fostriecin, 123 G1/S transition, 3 G2 checkpoint, 117 H2AX, 101 Geldanamycin, 8 Genistein, 70 Genomic instability, 118 Glutathione-S-transferase, 34 Golgi apparatus, 270 Guardian of the genome, 20 Helicases, 3 Heterochromatin, 288 Heterodimer, 97 Heterozygosity, 2 Histone deacetylases (HDACs), 3, 152, 288 Histone methyltransferases, 3 Holliday junctions, 60 Homologous recombination, 118 Hydrophilic, 25

Index Hydrophobic, 29 Hydroxyurea, 95, 295 Hyperacetylation, 287 Hyperglycemia, 145 Hypermethylation, 141 Hypersensitivity, 76, 188, 297 Hyperthermia, 119 Hypogonadism, 93 Hypomorphic, 97, 105 Hypoxia, 94 Idiosyncratic, 190 Imidazolone, 41 Immunodeficiency, 93 Indirubin, 108, 141 Indolocarbazole, 67, 122 Ionizing radiation (IR), 5, 93 Irinotecan, 5, 67 Isogranulatimides, 122 Isoindolinone, 40 Kinesin, 207 Kinesin superfamily (KIFs), 208 Kinetochore, 178, 210, 225 Leptomycin B, 166 Li-Fraumeni Syndrome, 20 Lipophilic, 33 Macromolecular, 23 Maleimide, 26 Maturation-promoting factor (MPF), 137 Mechlorethamine, 60 Meiotic, 267 Methotrexate, 60 Methyl methanesulfonate (MMS), 95 Methyltransferase inhibitor, 169 Methylxanthine, 122 Microtubule-associated proteins (MAPs), 178 Microtubule-destabilizing agents, 182 Microtubule organizing center (MTOC), 178 Microtubules, 178 Midbody, 211 Mini-chromosome maintenance (MCM), 3 Mismatch repair, 118 Mitotic catastrophe, 276 Mitotic checkpoint, 163, 259 Monastrol, 212 Monoastral, 209

Index Mosaic variegated aneuploidy (MVA), 232 MRE11/RAD50/NBS1 (MRN) complex, 96 Mucositis, 196 Myalgias, 189 Myelosuppression, 184 Neuropathy, 76, 196, 213 Neurotoxicity, 184, 189, 275 Neutropenia, 146, 184, 196, 277 Nijmegan breakage syndrome (NBS), 97 Nocodazole, 121–122 Non-Hodgkins lymphoma, 142 Nonhomologous end joining, 118 Nonpeptidic sulfonamides, 40 Nuclear excision repair, 118 Nutlins, 34–37 Off-target effect, 259 Okadaic acid, 96, 123 Oncoprotein, 20 Origin recognition complexes (ORCs), 3 Oxaliplatin, 277 Paclitaxel, 144, 165, 224 Palindromic, 21 Paullones, 141 PCB quinones, 70 Pentoxifylline, 122 Peptidomimetics, 40 P-glycoprotein, 214 Pharmacokinetics, 67, 188 Phenanthradines, 67 Phenoxymethyl tetrazole, 34 Phosphatidylinositol-3 kinase, 94 Phospho-residue-binding domain (BRCT), 98 Phytoestrogens, 69 Pifithrin-, 43 Platinum compounds, 119 Podophyllotoxin, 68 Polo-box domain (PBD), 269 Polo-like kinases (Plks), 267 Polyploidy, 248 Polysorbate, 80, 190 Preclinical models, 10 Prophase checkpoint, 164 Propioception, 189 Pyridopyrimidine, 123

313 Quercetin, 70, 108 Quinazolinone, 215 Quinones, 70

Rad9-Hus1-Rad1 (9–1-1) complex, 105 Radiomimetic, 94 Reoxygenation, 94 Replication protein A (RPA), 105 Replisome, 4 Retinoblastoma, 1, 32 RING finger, 28 Ring finger domain, 103, 166 RNA polymerase, 101, 150 Roscovitine, 9, 108

Saccharomyces cerevisiae, 240 Saccharomyces pombe, 240 Sclerosis, 190 Scytonemin, 277 Seckels syndrome, 105 Senescence, 20 Separase, 226 Simaomicin, 122 Sister chromatid cohesion, 270 Sister chromatids, 164 Spectroscopy, 31 Spindle checkpoint, 5, 224 Spindle midzone, 211 Spinocerebellar ataxia, 76 Staurosporin, 7, 122, 141 S-trityl-L-cysteine, 217 Structural maintenance of chromosomes (SMC), 98 Suberoylanilide hydroxamic acid (SAHA), 152 Survivin, 226 SWI/SNF complex, 3 Syncope, 145

Taxanes, 7, 42, 191 Teniposide, 68 Tetraploid, 259 Therapeutic index, 194 Therapeutic modalities, 2 Thermodynamic, 22 Thrombocytopenia, 146, 277 Thymidine kinase, 295 Thymidine synthase, 295

314 Tomudex, 6 Topoisomerase, 5, 59, 119 Topotecan, 67 Transactivation domain, 3 Translesion synthesis, 118 Treadmilling, 178 Tubulin, 177 Tumor suppressor, 20 Tumor xenografts, 149 Tumorigenesis, 21, 107 Tyrosyl-DNA phosphodiesterase 1 (Tdp1), 75

Index Ubiquitination, 224 Ubiquitinylation, 41 Ultraviolet (UV), 94, 277 Vinblastine, 179 Vinca alkaloid, 7, 42, 179, 191 Vincristine, 170, 179 VS-83, 215 Wortmannin, 108, 276 Xenograft, 40

Checkpoint Responses in Cancer Therapy (Cancer Drug Discovery and Development)

Antiangiogenic Agents in Cancer Therapy (Cancer Drug Discovery and Development) (Cancer Drug Discovery and Development)

DNA Repair in Cancer Therapy (Cancer Drug Discovery and Development)

Fluoropyrimidines in Cancer Therapy (Cancer Drug Discovery and Development)

Camptothecins in Cancer Therapy (Cancer Drug Discovery and Development)

Deoxynucleoside Analogs in Cancer Therapy (Cancer Drug Discovery and Development)

Farnesyltransferase Inhibitors in Cancer Therapy (Cancer Drug Discovery and Development)

Farnesyltransferase Inhibitors in Cancer Therapy (Cancer Drug Discovery and Development)

Antiangiogenic Agents in Cancer Therapy (Cancer Drug Discovery and Development)

Chemoradiation in Cancer Therapy (Cancer Drug Discovery and Development)

Death Receptors in Cancer Therapy (Cancer Drug Discovery and Development)

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

Regional Cancer Therapy (Cancer Drug Discovery and Development)

Gene Therapy for Cancer (Cancer Drug Discovery and Development)

Cancer Drug Resistance (Cancer Drug Discovery and Development)

Genomics in Cancer Drug Discovery and Development

Chemokine Receptors in Cancer (Cancer Drug Discovery and Development)

Tumor Models in Cancer Research (Cancer Drug Discovery and Development)

DNA Topoisomerases and Cancer (Cancer Drug Discovery and Development)

Apoptosis, Senescence and Cancer (Cancer Drug Discovery and Development)

Stem Cells and Cancer (Cancer Drug Discovery and Development)

Targets for Cancer Chemotherapy (Cancer Drug Discovery and Development)

Immunotherapy of Cancer (Cancer Drug Discovery and Development)

Anticancer Drug Development Guide (Cancer Drug Discovery and Development)

Principles of Anticancer Drug Development (Cancer Drug Discovery and Development)

Cancer Drug Design and Discovery

Checkpoint Controls and Cancer

Transforming Growth Factor-Beta in Cancer Therapy, Volume II: Cancer Treatment and Therapy (Cancer Drug Discovery and Development)

Checkpoint Responses in Cancer Therapy (Cancer Drug Discovery and Development)

Antiangiogenic Agents in Cancer Therapy (Cancer Drug Discovery and Development) (Cancer Drug Discovery and Development)

DNA Repair in Cancer Therapy (Cancer Drug Discovery and Development)

Fluoropyrimidines in Cancer Therapy (Cancer Drug Discovery and Development)

Camptothecins in Cancer Therapy (Cancer Drug Discovery and Development)

Deoxynucleoside Analogs in Cancer Therapy (Cancer Drug Discovery and Development)

Farnesyltransferase Inhibitors in Cancer Therapy (Cancer Drug Discovery and Development)

Farnesyltransferase Inhibitors in Cancer Therapy (Cancer Drug Discovery and Development)

Antiangiogenic Agents in Cancer Therapy (Cancer Drug Discovery and Development)

Chemoradiation in Cancer Therapy (Cancer Drug Discovery and Development)

Death Receptors in Cancer Therapy (Cancer Drug Discovery and Development)

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

Regional Cancer Therapy (Cancer Drug Discovery and Development)

Gene Therapy for Cancer (Cancer Drug Discovery and Development)

Cancer Drug Resistance (Cancer Drug Discovery and Development)

Genomics in Cancer Drug Discovery and Development

Chemokine Receptors in Cancer (Cancer Drug Discovery and Development)

Tumor Models in Cancer Research (Cancer Drug Discovery and Development)

DNA Topoisomerases and Cancer (Cancer Drug Discovery and Development)

Apoptosis, Senescence and Cancer (Cancer Drug Discovery and Development)

Stem Cells and Cancer (Cancer Drug Discovery and Development)

Targets for Cancer Chemotherapy (Cancer Drug Discovery and Development)

Immunotherapy of Cancer (Cancer Drug Discovery and Development)

Anticancer Drug Development Guide (Cancer Drug Discovery and Development)

Principles of Anticancer Drug Development (Cancer Drug Discovery and Development)

Cancer Drug Design and Discovery

Checkpoint Controls and Cancer

Transforming Growth Factor-Beta in Cancer Therapy, Volume II: Cancer Treatment and Therapy (Cancer Drug Discovery and Development)

Recommend Documents