YEAST AS TOOL IN CANCER RESEARCH
Yeast as Tool in Cancer Research
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JOHN L.. NITISS St. Jude Children’s Res...
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YEAST AS TOOL IN CANCER RESEARCH
Yeast as Tool in Cancer Research
Edited by
JOHN L.. NITISS St. Jude Children’s Research Hospital, Memphis, TN, U.S.A.
and
JOSEPH HEITMAN Duke University Medical Center, Durham, NC, U.S.A.
A C.I.P. Catalogue record for this book is available from the Library of Congress.
ISBN 978-1-4020-5962-9 (HB) ISBN 978-1-4020-5963-6 (e-book)
Published by Springer, P.O. Box 17, 3300 AA Dordrecht, The Netherlands. www.springer.com
Printed on acid-free paper
All Rights Reserved © 2007 Springer No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work.
TABLE OF CONTENTS Foreword…………………………………………………………………….ix List of Contributors………………………………………………………….xi Introduction………………………………………………………………....xv Chapter 1 FROM DNA REPLICATION TO GENOME INSTABILITY IN SCHIZOSACCHAROMYCES POMBE: PATHWAYS TO CANCER………………………………………………………………...1 Julie M. Bailis and Susan L. Forsburg Chapter 2 DISSECTING LAYERS OF MITOTIC REGULATION ESSENTIAL FOR MAINTAINING GENOMIC STABILITY…………...37 Jennifer S. Searle and Yolanda Sanchez Chapter 3 YEAST AS A TOOL IN CANCER RESEARCH: NUCLEAR TRAFFICKING……………………………………………….75 Anita H. Corbett and Adam C. Berger Chapter 4 STUDIES OF PROTEIN FARNESYLATION IN YEAST …………...….101 Nitika Thapar and Fuyuhiko Tamanoi Chapter 5 FROM BREAD TO BEDSIDE: WHAT BUDDING YEAST HAS TAUGHT US ABOUT THE IMMORTALIZATION OF CANCER CELLS…………………………………………………..…123 Soma S. R. Banik and Christopher M. Counter
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Table of Contents
Chapter 6 HSP90 CO-CHAPERONES IN SACCHAROMYCES CEREVISIAE……………………………………………………………...141 Marija Tesic and Richard F. Gaber Chapter 7 YEAST AS A MODEL SYSTEM FOR STUDYING CELL CYCLE CHECKPOINTS………………………………………….179 Carmela Palermo and Nancy C. Walworth Chapter 8 METABOLISM AND FUNCTION OF SPHINGOLIPIDS IN SACCHAROMYCES CEREVISIAE: RELEVANCE TO CANCER RESEARCH…………………………………………….…191 L. Ashely Cowart, Yusuf A. Hannun and Lina M. Obeid Chapter 9 EXPLORING AND RESTORING THE p53 PATHWAY USING THE p53 DISSOCIATOR ASSAY IN YEAST………………….211 Rainer K. Brachmann Chapter 10 FUNCTIONAL ANALYSIS OF THE HUMAN p53 TUMOR SUPPRESSOR AND ITS MUTANTS USING YEAST………………....233 Alberto Inga, Francesca Storici and Michael A. Resnick Chapter 11 ABC TRANSPORTERS IN YEAST – DRUG RESISTANCE AND STRESS RESPONSE IN A NUTSHELL………………………….289 Karl Kuchler and Christoph Schüller Chapter 12 THE FHCRC/NCI YEAST ANTICANCER DRUG SCREEN………..…315 Susan L. Holbeck and Julian Simon Chapter 13 YEAST AS A MODEL TO STUDY THE IMMUNOSUPPRESSIVE AND CHEMOTHERAPEUTIC DRUG RAPAMYCIN………………....347 John R. Rohde, Sara A. Zurita-Martinez and Maria E. Cardenas Chapter 14 USE OF YEAST AS A MODEL SYSTEM FOR IDENTIFYING AND STUDYING ANTICANCER DRUGS…………………………..…375 Jun O. Liu and Julian A. Simon
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Chapter 15 GENETIC ANALYSIS OF CISPLATIN RESISTANCE IN YEAST AND MAMMALS…………………………………………...393 Seiko Ishida and Ira Herskowitz Chapter 16 USING YEAST TOOLS TO DISSECT THE ACTION OF ANTICANCER DRUGS: MECHANISMS OF ENZYME INHIBITION AND CELL KILLING BY AGENTS TARGETING DNA TOPOISOMERASES………………………………409 Anna T. Rogojina, Zhengsheng Li, Karin C. Nitiss and John L. Nitiss Index…......….…………………………………………………………….429
FOREWORD Leland H. Hartwell Director, Fred Hutchinson Cancer Research Center, Nobel Laureate for Medicine, 2001
Yeast has proved to be the most useful single-celled organism for studying the fundamental aspects of cell biology. Resources are now available for yeast that greatly simplify and empower new investigations, like the presence of strains with each gene deleted, each protein tagged and databases on protein–protein interactions, gene regulation, and subcellular protein location. A powerful combination of genetics, cell biology, and biochemistry employed by thousands of yeast researchers has unraveled the complexities of numerous cellular processes from mitosis to secretion and even uncovered new insights into prion diseases and the role of prions in normal biology. These insights have proven, time and again, to foretell the roles of proteins and pathways in human cells. The collection of articles in this volume explores the use of yeast in pathway analysis and drug discovery. Yeast has, of course, supplied mankind’s most ubiquitous drug for thousands of years. In one aspect, the role of yeast in drug discovery is much like the role of yeast in other areas of biology. Yeast offers the power of genetics and a repetoire of resources available in no other organism. Using yeast in the study of drug targets and metabolism can help to make a science of what has been largely an empirical activity. A science of drug discovery would permit rigorous answers to important questions. What is the target of the drug? Is there more than one target and what are the relative affinities? What is the physiological consequence of inactivating a particular protein? Which drug in a panel is the most specific? How many ways can a cell mutate to resistance? What is the consequence of inhibiting two proteins? Which proteins in the cell if inhibited would produce a desired physiological outcome? Are all the proteins in a pathway equivalent targets? Can one identify drugs that alter the location or interactions of proteins without affecting their activity? Each of these questions can be rigorously answered in yeast but not in most other
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systems. Questions like these have rarely been answered in the field of drug discovery. A more challenging question is: Can yeast be made more applicable for the discovery of drugs against human targets? While many human drugs are active in yeast many are not. The lack of effect in yeast can be due to the fact that yeast does not have the targets at all – e.g., cell surface hormone receptors, or because the orthologous protein in yeast is sufficiently different, or that yeast cells have redundant proteins and inhibition of one is masked by the second. However, even if drugs against human targets are active in yeast, the yeast orthologue is likely to be different enough to preclude optimization of drug identification in yeast. One way to solve this problem is by substitution of human orthologues for yeast ones. One could even substitute human transport proteins and drug-metabolizing proteins to further optimize the yeast system. Moreover, our ultimate interest in drugs is to alter physiology, which is the product not of single proteins but of pathways and networks of proteins acting in concert. The most effective use of yeast would probably result from substituting entire pathways of yeast proteins with their human counterparts. With the aid of reporters that quantitatively reveal the activity of the pathway at different points one could enter a new era of drug discovery that interrogates modulation of the pathway at different sites. Finally, we should think about using the same approach for pathways that do not normally exist in yeast – for example pathways that synthesize hormones. By constructing the pathway in yeast de novo with appropriate reporters one could screen for drugs that modulate hormone synthesis and easily localize the target in the pathway. This sounds like fun. I suspect the dough has only begun to rise on what yeast has to offer in the arena of drug discovery.
LIST OF CONTRIBUTORS JULIE M. BAILIS The Salk Institute for Biological Studies 10010 North Torrey Pines Road La Jolla, CA 92037
ANITA H. CORBETT Department of Biochemistry and Graduate Program in Biochemistry Cell and Developmental Biology Emory University School of Medicine 1510 Clifton Rd., NE Atlanta, GA 30322
SOMA S. R. BANIK Departments of Pharmacology and Cancer Biology and Radiation Oncology Duke University Medical Center, Box 3813 Durham, NC, USA 27710
CHRISTOPHER M. COUNTER Departments of Pharmacology and Cancer Biology and Radiation Oncology Duke University Medical Center, Box 3813 Durham, NC, USA 27710
ADAM C. BERGER Department of Biochemistry and Graduate Program in Biochemistry Cell and Developmental Biology Emory University School of Medicine 1510 Clifton Rd., NE Atlanta, GA 30322
L. ASHELY COWART Ralph H. Johnson Veterans Administration and the Departments of Medicine and Biochemistry and Molecular Biology Medical University of South Carolina Charleston, South Carolina
RAINER K. BRACHMANN Department of Medicine University of California at Irvine Irvine, CA, USA 92697
SUSAN L. FORSBURG Molecular and Computational Biology Section University of Southern California 835 W. 37th St., SHS 172 Los Angeles, CA 90089-1340
MARIA E. CARDENAS Department of Molecular Genetics and Microbiology Duke University Medical Center Durham, NC, USA 27710
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xii RICHARD F. GABER Department of Biochemistry, Molecular Biology and Cell Biology Northwestern University Evanston, Illinois 60208 YUSUF A. HANNUN Ralph H. Johnson Veterans Administration and the Departments of Medicine and Biochemistry and Molecular Biology Medical University of South Carolina Charleston, South Carolina IRA HERSKOWITZ Department of Biochemistry and Biophysics University of California San Francisco, CA SUSAN L. HOLBECK National Cancer Institute Developmental Therapeutics Program Information Technology Branch, Rockville, MD ALBERTO INGA Laboratory of Molecular Genetics National Institute of Environmental Health Sciences NIH, P.O. Box 12233 Research Triangle Park NC 27709 SEIKO ISHIDA Department of Biochemistry and Biophysics University of California San Francisco, CA
List of Contributors KARL KUCHLER Mqx F. Perutz Laboratories Department of Medical Biochemistry Division of Molecular Genetics Medical University Vienna Campus Vienna Biocenter Vienna, Austria ZHENGSHENG LI Department of Molecular Pharmacology St. Jude Children’s Research Hospital Memphis, TN 38105 JUN O. LIU Departments of Pharmacology and Neuroscience Johns Hopkins School of Medicine Baltimore, MD 21205 JOHN L. NITISS Department of Molecular Pharmacology St. Jude Children’s Research Hospital Memphis, TN 38105 KARIN C. NITISS Department of Molecular Pharmacology St. Jude Children’s Research Hospital Memphis, TN 38105 LINA M. OBEID Ralph H. Johnson Veterans Administration and the Departments of Medicine and Biochemistry and Molecular Biology Medical University of South Carolina Charleston, South Carolina
List of Contributors CARMELA PALERMO Department of Pharmacology UMDNJ-Robert Wood Johnson Medical School and Joint Graduate Program in Cellular and Molecular Pharmacology UMDNJ-Graduate School of Biomedical Sciences and Rutgers The State University of New Jersey 675 Hoes Lane Piscataway, NJ 08854-5635 MICHAEL A. RESNICK Laboratory of Molecular Genetics National Institute of Environmental Health Sciences NIH, P.O. Box 12233 Research Triangle Park, NC 27709 ANNA T. ROGOJINA Department of Molecular Pharmacology St. Jude Children’s Research Hospital Memphis, TN 38105 JOHN R. ROHDE Department of Molecular Genetics and Microbiology Duke University Medical Center Durham, NC, USA 27710 YOLANDA SANCHEZ Department of Pharmacology and Toxicology Dartmouth Medical School 7650 Remsen Hanover, NH 03755 USA
xiii CHRISTOPH SCHÜLLER Mqx F. Perutz Laboratories Department of Medical Biochemistry Division of Molecular Genetics Medical University Vienna Campus Vienna Biocenter Vienna, Austria JENNIFER S. SEARLE Department of Molecular Genetics, Biochemistry and Microbiology The University of Cincinnati 231 Ablert Sabin Way Cincinnati, OH 45267-0524 JULIAN A. SIMON Divisions of Clinical Research, Basic Sciences and Human Biology Fred Hutchinson Cancer Research Center Seattle, WA 98109 FRANCESCA STORICI Laboratory of Molecular Genetics National Institute of Environmental Health Sciences NIH, P.O. Box 12233 Research Triangle Park, NC 27709 FUYUHIKO TAMANOI Department of Microbiology, Immunology & Molecular Genetics Genetics Jonsson Comprehensive Cancer Center Molecular Biology Institute University of California Los Angeles, 405 Hilgard Ave. Los Angeles, CA 90095-1489
xiv MARIJA TESIC Department of Biochemistry, Molecular Biology and Cell Biology Northwestern University Evanston, Illinois 60208 NITIKA THAPAR Department of Microbiology, Immunology & Molecular Genetics Jonsson Comprehensive Cancer Center Molecular Biology Institute University of California Los Angeles, 405 Hilgard Ave. Los Angeles, CA 90095-1489
List of Contributors NANCY C. WALWORTH Department of Pharmacology UMDNJ-Robert Wood Johnson Medical School and Joint Graduate Program in Cellular and Molecular Pharmacology UMDNJ-Graduate School of Biomedical Sciences and Rutgers The State University of New Jersey 675 Hoes Lane Piscataway, NJ 08854-5635 SARA A. ZURITA-MARTINEZ Department of Molecular Genetics and Microbiology Duke University Medical Center Durham, NC, USA 27710
INTRODUCTION John L. Nitiss Department of Molecular Pharmacology, St. Jude Children’s Research Hospital, Memphis, TN 38105
In 1970, Lee Hartwell reported a series of genetic experiments showing that progression through the cell cycle in yeast was amenable to genetic analysis. At about the same time, several investigators, including Michael Resnick and Brian Cox identified the first yeast mutants that were shown to be defective in DNA repair processes. Walt Fangman and his co-workers were characterizing the basics of yeast chromosomes and yeast DNA replication (even though cytogenetics was not practical, and because yeast lack thymidine kinase, pulse labeling of DNA was not possible). Gerry Fink was identifying the many ways a eukaryote regulates gene expression, while Fred Sherman carried out studies on cytochrome c that illuminated translation (and much else). Other investigators were becoming convinced that yeast could shed light on many fundamental processes that were not accessible in multicellular eukaryotes. Since many investigators committed to using yeast as an experimental system, there was also considerable efforts to increase the scope of yeast genetics by developing new genetic tools, which became an effort to develop molecular, biochemical, and cell biological tools. The important tools developed in yeast are too numerous to mention (although the discovery of gene replacement by homologous recombination surely requires note). Contemporary biologists, even those studying “large” eukaryotes, continue to learn from yeast systems. Despite the impressive roster of accomplishments in basic biology obtained using yeast as a model, there are areas of importance of cancer research where yeast has not been extensively utilized. In addition, many investigators and clinicians working in many areas of cancer research tend not to think of yeast as being relevant to their areas of interest. In some, cases, yeast researchers have not made the appropriate effort to communicate their results to the cancer research community. The goal of this book is to highlight the contributions that yeast systems have made to in a variety of
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areas of cancer research. Accordingly, this volume is intentionally directed more to workers outside the “yeast world” and toward investigators interested in cancer. We have requested the authors to highlight areas where yeast-based systems have made contributions not readily accessible with other experimental systems, and to try to communicate clearly to workers who may not be familiar with yeast. This book is broadly organized into three sections. The first section, including Chapters 1 through 8 highlight areas of biology that are particularly relevant to cancer research. These include studies of DNA metabolism (Chapters 1, 2, and 7), protein localization and trafficking (Chapters 3, 4, and 6), and cell immortalization (Chapter 5). Chapter 8, a discussion of sphingolipids, is relevant both to the biology, and potentially, the development of novel cancer treatments. The second section, Chapters 9 and 10 describe how yeast can be used to study human p53. These chapters highlight the ability to learn about the function of human oncoproteins using yeast. The third section is broadly concerned with studying anticancer drugs in yeast. Some of the chapters discuss concerns broadly relevant to drug action (Chapters 11 and 14), while the actions of specific anticancer drugs, such as rapamycins, platinum compounds, and topoisomerase inhibitors are explored in Chapters 13, 15, and 16. Finally, Chapter 12 describes one broad effort to use yeast as a tool for drug discovery. There are many other areas of interest not included in this volume where yeast systems have made important contributions to cancer research. These areas include important methodologies such as yeast two hybrid, areas of basic biology such as the study of yeast Ras proteins and yeast kinases, and areas of great relevance to anticancer drugs, such as yeast systems of DNA repair. While we hope to include such topics in future volumes, we also felt that there were other superb sources already available for topics such as a general introduction to yeast. This book would not have been possible without the efforts of Peggy Vandiveer in the Word Processing Center at St. Jude Children’s Hospital. Peggy carefully formatted all of the chapters and cheerfully and quickly handled a huge amount of work. Thanks are also due to Jeffrey Berk and Aman Seth in the Nitiss laboratory, who carefully checked all of the chapters and caught many things that might have slipped through. Support for the generation of this book was provided to JLN by the American Lebanese Syrian Associated Charities (ALSAC).
Chapter 1 FROM DNA REPLICATION TO GENOME INSTABILITY IN SCHIZOSACCHAROMYCES POMBE: PATHWAYS TO CANCER
Julie M. Bailis1 and Susan L. Forsburg2 1
The Salk Institute for Biological Studies, 10010 North Torrey Pines Road, La Jolla, CA 92037; 2Molecular and Computational Biology Section, University of Southern California, 835 W. 37th St., SHS 172, Los Angeles, CA 90089-1340
1
INTRODUCTION
The genetic integrity of cells depends on the complete, accurate replication of each genome cell cycle. Cells are particularly susceptible to genetic changes during the DNA synthesis (S) phase of the cell cycle, because only one complete copy of the DNA template exists, and potentially damaging breaks and unwinding occur as part of the replication process. Genome instability may result in deletion or amplification of genetic information within a chromosome, translocation of part of a chromosome to another chromosome, or gain or loss of whole chromosomes. These changes, in turn, can have important consequences for chromatin structure and gene expression. In wild-type cells, DNA replication is tightly regulated and involves multiple mechanisms that prevent amplification or loss of genetic information. In addition, internal controls such as checkpoints delay or arrest replication if active replication forks are blocked. Cancer cells, in contrast, are characterized by uncontrolled proliferation and chromosome instability. The fission yeast Schizosaccharomyces pombe provides an outstanding model for studies of replication and chromosome dynamics, with replication origins and centromeres that are similar to those of metazoans [46, 205]. This review will focus on DNA replication in S. pombe and its role in maintenance of genome integrity. We will consider the choice and 1 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 1–35. © 2007 Springer.
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organization of origins of DNA replication, the proteins that assemble at origins and those that are responsible for actual DNA synthesis. We will also describe regulatory mechanisms that promote origin firing, control the timing of the replication program, and limit replication to once per cell cycle. Then, we will discuss the cellular response to blocks to DNA replication. Finally, we will evaluate how studies of DNA replication in fission yeast and other model organisms have demonstrated relevance to cancer biology, concentrating on those pathways that, when disrupted, lead to genome instability and progression toward cancer.
1.1
DNA replication origins and the origin recognition complex
The fission yeast genome, like that of other eukaryotes, is divided among multiple linear chromosomes each requiring multiple origins of replication. DNA synthesis must therefore be coordinated both between the different chromosomes and within each individual chromosome. The identification of specific fragments of the genome that could replicate autonomously on plasmids provided the first step toward identifying S. pombe origins of replication. The autonomously replicating sequence (ARS) elements in S. pombe that have been described thus far are at least 500 bp in size and correspond to intergenic regions of the genome [24, 53, 114, 203]. Interestingly, centromeric regions of S. pombe chromosomes appear to be enriched for DNA fragments with ars activity [170]. In some chromosomal regions, two or more origins are clustered close together [40, 147]. Replication initiates from discrete, defined sites within each origin region [24, 39, 40, 53, 147, 203]. S. pombe replication origins preferentially lie in adenine–thymine (A–T) rich regions of the genome where the local sequence of As and Ts is asymmetric [210]. Although short consensus sequences similar to that of the S. cerevisiae ARS element have been identified within some of the S. pombe origins, these are not essential for origin activity [114]. Fine structure examination of individual origins has revealed multiple short stretches of DNA where replication initiates as well as adjacent accessory sequences that promote origin function [29, 39, 85]. Thus, the organization of S. pombe replication origins appears to be modular, with redundant, dispersed, and degenerate elements contributing to efficient activity. Different replication origins display distinct firing efficiencies [147]. It is not known whether the choice of origins is regulated or stochastic. Within those regions of the chromosome where origins are clustered, there may be a hierarchy of preferential origin usage determined at least in part by local enhancer sequences [84]. In the S. pombe genome, the number of potential
Chapter 1: Replication in Fission Yeast
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replication origins has been estimated at 1 per 20 kb [114] to 1 per 55 kb [203]. The recent publication of the S. pombe genome sequence [205] should facilitate genome-wide identification of replication origins as has recently been described for budding yeast [153, 206], as well as provide information about origin usage under different growth conditions. Replication initiation at origins requires the association of multiple conserved proteins with the origin region [41, 80]. As in other eukaryotes, the S. pombe origin recognition complex (ORC) binds to the origin DNA and marks it as a potential site of replication initiation [27, 28, 179]. ORC is composed of six related proteins (ORC1–6), each one of which is essential and conserved in eukaryotic organisms [13]. In some organisms, several of the ORC subunits may contact the origin DNA [13]. In S. pombe, ORC binding to the origin DNA is mediated by the Orp4 subunit, which contains multiple A–T hooks [27, 28, 90]. This motif, which is not present in the Orc4 subunit of other eukaryotes, is thought to promote protein binding within the DNA minor groove in regions that are A–T rich [27]. Unexpectedly, S. pombe ORC can bind to multiple specific sites within a single replication origin [28, 179]. These sites correspond to the regions of the origin where replication initiates as determined by two-dimensional gel analysis [179]. Thus, the position of the ORC complex within the origin may determine the location and direction of the replication machinery. It is not known whether S. pombe ORC also associates with nonorigin DNA, as has been described for other eukaryotes and which may reflect the additional role of ORC in heterochromatic silencing [13, 206]. The majority of ORC protein in S. pombe associates with chromatin, including replication origins, throughout the cell cycle [107, 146]. The Orp2 (Orc2) subunit, originally identified as a cyclin-dependent kinase (CDK) binding protein, contains consensus sites for CDK phosphorylation [95]. Orp2 becomes phosphorylated in a CDK-dependent manner during mitosis, and is dephosphorylated during G1/S of the cell cycle [107, 197]. It remains to be determined whether other ORC subunits also become modified during the cell cycle.
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THE PREREPLICATIVE COMPLEX AND LICENSING OF ORIGINS
The preparation for S phase begins as early as mitosis (M phase) or G1, when additional proteins required for DNA replication assemble sequentially at replication origins to form the prereplicative complex (preRC). A series of additional steps is required to convert the preRC into an initiation complex that actively synthesizes DNA. While the proteins and general pathways
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involved in this process are conserved in eukaryotes, many of the details and order of their functions are still incomplete. Figure 1 provides a model of our current view of the assembly and activation of proteins at replication origins.
Figure 1. Model of assembly and activation of the preRC based on information from multiple systems. Cdc18 and Cdt1 bind to ORC at the origin and facilitate the loading of the MCM protein complex. The preRC is activated by the actions of the CDK and Hsk1/Dfp1 kinases (“P” indicates phosphorylation). This promotes the association of other replication factors with the complex localized to the origin, and leads to initiation of DNA synthesis. Many of the relevant substrates of Hsk1 and CDK have yet to be identified.
The S. pombe Cdc18 (Cdc6 in other eukaryotes) and Cdt1 proteins bind to origin sites marked by ORC [55, 95, 141]. ORC and Cdc18 physically interact [28], as do Cdc18 and Cdt1 [141]. ORC, Cdc18, and Cdt1 together recruit the minichromosome maintenance (MCM) proteins to the complex assembling at origins [79, 146]. Genetic interactions between the MCM proteins and Cdc18, and the MCM proteins and ORC, have been demonstrated in S. pombe [49, 55, 101, 102]. Although each of the preRC components is essential for DNA replication, the different proteins carry out distinct functions in assembly and activation of replication origins. The Cdc18/Cdc6 and Cdt1 proteins are thought to act as a “licensing factor” that is a critical determinant of the onset of DNA replication [81, 109, 142]. Expression of S. pombe Cdc18 and Cdt1, which is controlled by the Cdc10 transcription factor [65, 81], is restricted to the G1/S window of the cell cycle [81, 141]. In addition, the Cdc18 protein is regulated by phosphorylation, which targets Cdc18 for ubiquitin-mediated proteolysis [68, 69]. It is not known whether S. pombe Cdt1 protein levels are also regulated. In Xenopus, Cdt1 activity is inhibited by association with another protein, geminin [16]; however, a geminin homolog has not been described in yeast. The cell cycle-regulated activity of Cdc18 and Cdt1 both promote
Chapter 1: Replication in Fission Yeast
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preRC assembly and prevents its reassembly until the next cell cycle [17, 80]. The MCM complex is not only a preRC component, but also a compelling candidate for a replicative helicase [41, 80, 98, 148]. The complex is composed of six homologous subunits (MCM2–7) with similarity to a large family of ATPases [91]. Each subunit is conserved in other eukaryotes, and each is essential for replication and cell viability [148, 193]. Mutation of individual S. pombe MCM genes results in defects in replication, irreversible S-phase arrest and delocalization of the entire MCM complex from the nucleus [34, 49, 101, 149]. The mcm mutant arrest occurs with the bulk of replication completed and requires the DNA damage checkpoint, suggesting that the chromosomes are damaged in mcm mutant cells [101, 102, 108, 123, 178]. Interestingly, although the MCM proteins are estimated to be at least tenfold more abundant than the predicted number of origins, reducing the amount of a single MCM protein results in chromosome instability and defects in the completion of S phase [49, 102]. Replication can still initiate with low levels of MCM protein, suggesting that the amount of MCM protein required to initiate DNA replication is much less than that needed to complete S phase [102]. Genetic interactions between S. pombe MCM genes and factors involved in the elongation step of DNA replication suggest that MCM complex function is required throughout S phase [49, 102]. This is consistent with more direct experiments in S. cerevisiae, where specific degradation of one of the six MCM subunits during S phase blocks further DNA replication [94, 103]. The current model suggests that a heterohexameric MCM complex with all six MCM subunits is present at replication origins, and then travels with the replication fork [78, 148, 193]. Although all six S. pombe MCM subunits interact [1], distinct MCM subcomplexes have been identified in vitro and in vivo, suggesting that individual MCM proteins have different relative affinities for each other [96, 162, 163]. The Mcm4, 6, and 7 proteins are thought to form a “core” complex that is tightly associated [96, 163]. Mcm2 associates with this core through interactions with Mcm4 [163]. The Mcm3 and 5 proteins form a dimer that is also loosely associated with the MCM core proteins [162], probably through interaction with Mcm7 [101]. Similar subcomplexes of MCM proteins have also been described in other eukaryotes [67, 160]. The core complex of Mcm4, 6, and 7, but not the heterohexameric complex of Mcm2, 3, 4, 5, 6, and 7, demonstrates weak helicase activity in vitro in S. pombe [96] and in human cells [67]. Curiously, point mutations of conserved residues in each Mcm protein that are predicted to inhibit ATPase or helicase activity display different effects in vivo in both S. pombe [49, 52] and S. cerevisiae [160] depending on the subunit mutated. This suggests division of labor
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amongst the MCM subunits, and may explain why six related proteins are required for MCM activity. Although MCM nuclear localization is regulated in budding yeast, MCM proteins localize to the nucleus constitutively throughout the cell cycle in fission yeast and in metazoans [149, 193]. In these organisms, the association of the MCM proteins with chromatin varies: MCM proteins localize to chromatin, including replication origins, in late M phase and dissociate during S phase [79, 146]. Thus, regulation of MCM chromatin binding is one mechanism of control of MCM complex function. Part of this regulation is provided by the Cdc18 and Cdt1 loading factors, which limit MCM binding to M/G1 of the cell cycle [80]. In some organisms, Mcm4 is also regulated by CDK-dependent phosphorylation. In Xenopus, this promotes Mcm4 dissociation from chromatin [62, 152]; in S. cerevisiae, Mcm4 phosphorylation leads to its exclusion from the nucleus [140]. However, CDK consensus site mutants of S. pombe Mcm4 do not display obvious phenotypes in vivo [52].
3
REPLICATION INITIATION REQUIRES TWO DISTINCT PROTEIN KINASE ACTIVITIES
Although several of the proteins required for DNA replication are assembled at replication origins prior to the start of S phase, origin firing and initiation of DNA synthesis requires the activities of two protein kinase complexes, the CDK [17] and DDK (Dbf4-dependent) kinases [73, 110, 161]. While some substrates of these protein kinases have been identified, it is clear that our understanding of their role is still incomplete. In fission yeast, there is a single CDK, cdc2+, that functions as the major regulator of cell cycle transitions including S phase onset [127]. Importantly, the ability of S. pombe Cdc2 to functionally complement cross-species [11, 97] suggests that the principles of function and regulation of the CDKs are conserved in eukaryotes. S. pombe Cdc2 associates with different cyclins in different stages of the cell cycle. The G1/S phase transition is promoted by the assembly of Cdc2 with the B-type cyclin Cig2 [45, 125]. In mitosis, Cdc2 associates with the Cdc13 cyclin [127]. Fission yeast CDK activity varies through the cell cycle, and this global regulation controls the dependency of S phase on completion of mitosis and prevents multiple rounds of replication within a single cell cycle [21, 31, 126].
Chapter 1: Replication in Fission Yeast
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Figure 2. Control mechanisms that prevent rereplication. CDK activity is regulated so that the preRC can only assembly once each cell cycle. CDK phosphoyrlates Drc1, which promotes association with Rad4/Cut5 and replication initiation. CDK phosphorylation of Cdc18 and Orc2 prevents origins from refiring.
Remarkably, the CDK kinase complex plays both positive and negative roles in S phase progression (Figure 2). Replication initiation is positively activated by CDK-mediated phosphorylation of Drc1, which results in the association of Drc1 with Rad4/Cut5 [143]. Rad4/Cut5 is essential for the initiation of DNA synthesis and also acts in the DNA damage checkpoint [44, 156, 157]. Although the molecular role of Rad4/Cut5 is unclear, the interaction between Rad4 and Drc1 may promote the association of the DNA polymerases α and ε with origins [143]. The S. pombe homologs of Rad4/Cut5 in S. cerevisiae (Dpb11) and in human cells (TopBP1) interact with DNA polymerase as part of their replication function [112, 182]. Each of these proteins contains BRCT motifs, which are also found in the human DNA repair gene XRCC1 and the BRCA1 tumor suppressor [19]. CDK activity negatively regulates DNA replication by preventing origins from refiring in a single cell cycle [17, 80]. CDK-mediated phosphorylation of Cdc18 and its subsequent degradation prevent reassembly of the preRC until the next cell cycle, because recruitment of preRC factors such as the MCM proteins depends on Cdc18 [68]. Overproduction of Cdc18 or expression of a nonphosphorylated version of Cdc18 induces re-replication, presumably by resetting origins to the G1 state [80]. Mutation of the CDK phosphorylation sites in Orp2 also allows re-replication [197], although the mechanism by which CDK-mediated phosphorylation of Orp2 prevents re-replication is not clear [95, 107, 197]. Unlike Cdc18, Orp2 appears to remain associated with the ORC complex at the origin [107]; it is not known whether phosphorylation of Orp2 changes its affinity for other origin-associated proteins. Manipulation of the Cdc2 protein kinase itself can result in rereplication. Cells that lack mitotically active Cdc2 [21] or the mitotic cyclin Cdc13 [61] re-replicate DNA, as do cells overproducing the CDK inhibitor
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Rum1 [126]. These observations demonstrated that increased levels of CDK kinase activity after the G1/S transition are required to prevent reinitiation of DNA replication within a single cell cycle. Re-replication appears to mimic a normal S phase because it requires all of the proteins that function in DNA synthesis in a normal S phase [172]. This suggests that re-replication results solely from a disruption of the order of the cell cycle. CDK kinase activity is not the only control exerted over the onset of S phase. Initiation of DNA replication also requires the activity of the Cdc7 protein kinase [161]. Hsk1, the S. pombe homolog of Cdc7 [23, 111] associates with a subunit, Dfp1 (Dbf4 in other eukaryotes), which is required for Hsk1 kinase activity toward its substrates [23, 180]. Although the levels of Hsk1 protein appear to be constant in different phases of the cell cycle, Dfp1 levels and the associated Hsk1-dependent protein kinase activity are specifically upregulated during G1/S [22]. The Mcm2 protein is a substrate of Hsk1/Dfp1 in vitro [22] and in vivo [181]. However, the biological effect of Mcm2 phosphorylation remains to be determined. It is also possible that Hsk1 has additional, as yet unidentified, substrates in vivo; in S. cerevisiae, DNA polymerase α and another replication factor, Cdc45 (the homolog of S. pombe Sna41), are targets of Cdc7 [82, 145, 201]. S. pombe Hsk1 has three apparent roles in the cell. First, Hsk1/Dfp1 may activate individual origins of replication by phosphorylating components of the preRC, as has been described for Cdc7 in S. cerevisiae [20, 37]. This role for Hsk1 may also involve control of the temporal order of origin firing, since the kinase is a potential target of the replication checkpoint [22, 171, 181]. Second, Hsk1 is involved in the recovery from replication blocks such as hydroxyurea [171]. Third, Hsk1 may influence the establishment of sisterchromatid cohesion during S phase [8, 171, 181]. The potential mechanisms of these activities are discussed below.
4
ADDITIONAL REPLICATION FACTORS
Once the preRC is formed and activated by the CDK and Cdc7 protein kinases, additional components of the replication machinery become associated with the complex, which becomes the active replication fork. Many of these proteins are essential for viability and conserved in all eukaryotes, including Cdc23/Mcm10, Sna41/Cdc45, the single-strand DNA binding protein replication protein A (RPA), DNA polymerase α DNA polymerase δ DNA polymerase ε DNA, replication factor C (RFC), and proliferating cell nuclear antigen (PCNA) [198]. S. pombe cdc23 mutants, like the corresponding mutants in other eukaryotes, display defects in replication and demonstrate genetic
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interactions with other replication mutants [7, 77, 100]. A mutant of the S. cerevisiae homolog of cdc23+, MCM10, can be complemented by S. pombe cdc23+, suggesting conservation of gene function [7]. S. cerevisiae MCM10 appears necessary to recruit the MCM complex to chromatin [66], and also functions in replication elongation [116]. In both S. cerevisiae and S. pombe, Cdc23 associates with chromatin throughout the cell cycle [66, 101]. In Xenopus, MCM10 is dispensable for MCM complex binding to chromatin but is required to localize Cdc45/Sna41 [204]. Sna41/Cdc45 appears to be required both for replication initiation and elongation. The association of Sna41 with chromatin requires the Sld3 protein, which has been described in both the budding and fission yeasts [74, 136]. Sna41/Cdc45 is required to load other replication factors such as RPA, DNA polymerase α, DNA polymerase ε, and PCNA [182]; in S. pombe, Sna41 [124, 194] has been shown to interact with DNA polymerase α [195] and recruit it to the MCM protein complex at replication origins [195]. A role for Sna41/Cdc45 in origin DNA unwinding has been suggested from experiments in Xenopus [199] and in S. cerevisiae [74]. In S. cerevisiae, Cdc45 function has been shown to be required throughout S phase [189]. Many of the genes involved in DNA replication in S. pombe are homologous to those described in other eukaryotes, and the enzymology of the gene products is conserved [198]. RPA is needed to recruit the DNA polymerases α, δ, and ε to replication origins [198]. Interestingly, DNA polymerase α may have an additional function beyond replication initiation and DNA synthesis (see section VIII). The eukaryotic DNA polymerase processivity factor PCNA becomes associated with the replication fork through the actions of the clamp loader, RFC [198]. S. pombe PCNA also interacts with DNA polymerase δ [155] and promotes its processivity [6]. Components of S. pombe RFC and primase are involved in the replication checkpoint as well as in DNA synthesis [56, 164]. Several proteins involved in lagging strand metabolism, such as DNA ligase [72], the Dna2 helicase [54, 75], and the homolog of the FEN1 endonuclease, Rad2 [132] have also been characterized in S. pombe. The dna2 mutant can be suppressed by overproduction of DNA polymerase δ, DNA ligase or Rad2, suggesting that Dna2 has a central function in Okazaki fragment maturation [75]. Mutants of a novel S. pombe replication factor, cdc24+, are suppressed by overproduction of Dna2, as well as by overproduction of PCNA or RFC [54, 184]. S. pombe cdc24+ has no known sequence homolog in other systems [54]. Mutants of cdc24+ are defective in the completion of S phase and display chromosome breakage, which is not typical of replication mutants [54]. Cdc24 thus may play a role in
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maintenance of chromosome integrity during S phase, perhaps through a function in Okazaki fragment maturation.
5
TIMING OF REPLICATION ORIGIN FIRING
Coordination of multiple replication origins on multiple chromosomes is essential to maintain stability of the genome. In wild-type cells, there is a temporal program of origin firing: some origins fire early in S phase, others are active in the middle of S phase, and still other replication origins do not initiate synthesis until late in S phase. In general, heterochromatic regions are late-replicating in most organisms [208]. However, a recent analysis of replication origins in S. pombe [86] suggests that a centromeric origin in S. pombe replicates early in S phase. In contrast, origins located within other regions of heterochromatin, in particular the telomere and the rDNA, replicate late in S phase. Although most [14, 16] of the origins analyzed in this study [86] replicate early in S phase, it remains to be determined whether these are representative of other S. pombe origins. Control of the timing of origin firing has been investigated using the drug hydroxyurea (HU) to block cells in S phase. HU treatment causes an array of lesions in the cell, including inhibition of the enzyme ribonucleotide reductase, which depletes the nucleotide pools and prevents further DNA replication [80, 106, 158, 188]. For the few origins that have been analyzed in S. pombe, HU treatment prevents the activity of normally late-firing origins, but does not restrain early origin firing [86]. This mechanism is checkpoint-dependent [86], as both early and late origins are active in mutants lacking the checkpoint kinases Rad3 (a homolog of human ATM) or Cds1 (the equivalent of human CDS1/CHK2 and budding yeast Rad53). Checkpoint-dependent inhibition of late origin firing was previously described in budding yeast [158, 166], and the conservation of replication timing control between these divergent yeasts suggests that a similar mechanism should also operate in human cells.
6
RESPONSE TO REPLICATION BLOCKS
During S phase progression, active replication forks may encounter blocks to further synthesis. In S. pombe, HU-induced arrest has been most often used to study the effects of blocks to replication. Cells treated with HU undergo checkpoint-mediated arrest in S phase that depends on Cdc2 and six “checkpoint rad” proteins [25]. Wild-type cells can recover from HU treatment, resulting in restart of DNA replication, the completion of S phase
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and subsequent progress through the cell cycle. This suggests that replication fork structure is preserved during S phase arrest caused by HU treatment. The checkpoint kinase Cds1 may play a role in stabilization of stalled replication forks. S. pombe cds1 mutants arrest in HU, but are defective in recovery from the arrest [104]. The HU-induced arrest of cds1 cells results from the activation of the damage checkpoint kinase Chk1, suggesting that Cds1 is needed to prevent chromosomal damage in the presence of HU [104]. Consistent with this hypothesis, aberrant replication structures are observed in S. pombe cds1 and rad3 mutants treated with HU [86]. Loss of replication fork integrity has also been observed in rad53 mutants of S. cerevisiae, suggesting a conserved mechanism of damage tolerance [106, 188]. S. cerevisiae Rad53 is phosphorylated in response to S phase damage [150, 188] and phosphorylates components of the replication machinery to restrain its progression [150]. Similarly, S. pombe Cds1 becomes phosphorrylated in response to HU treatment [104]. This in turn leads to Cds1dependent phosphorylation of both Hsk1 and its activator Dfp1, suggesting that Hsk1 and Dfp1 are targets of the cellular response to replication blocks [22, 171]. These phosphorylation events may serve to regulate the interaction of Hsk1/Dfp1 with proteins localized to replication origins and to prevent the activation of origins under conditions unfavorable to the cell [70, 110]. Although S. pombe Hsk1 and Dfp1 have an essential role in activation of replication origins, Cds1 is dispensable during a normal S phase [104, 129]. Recently, an S-phase specific upstream activator of Cds1, Mrc1, was identified [3, 185]; both Mrc1 and Cds1 are nonessential for cell viability [3, 129]. Future work should determine how the cell senses blocks to replication and how this information is directed to Mrc1 and Cds1. Recovery from HU-induced S phase arrest also requires Rqh1, the fission yeast homolog of the Escherichia coli RecQ helicase [175]. The wild-type function of Rqh1 during S phase is not known but may involve preventing inappropriate recombination events when replication forks are stalled [175]. Mitotic recombination is elevated in rqh1 mutants that are treated with HU [131, 175]. Mutants of rqh1+ also display synthetic genetic interactions with components of the replication machinery [131, 171]. In S. cerevisiae, the Rqh1 homolog SGS1 localizes with the Rad53 kinase and promotes its phosphorylation in response to HU [51]. However, SGS1 mutants are hyperrecombinant even in the absence of HU, suggesting there may be some differences in the function between SGS1 and Rqh1 [51].
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RECOMBINATION PROTEINS IN S PHASE
Several replication mutants display increased levels of recombination, suggesting a link between these processes. Certain alleles of S. pombe DNA polymerase α, DNA ligase, and rad2+ have mutator phenotypes [105]. The increase in mutation frequency in these mutants suggests that the corresponding wild-type proteins prevent genome changes and rearrangements, which may result from recombination during S phase. Recombination is elevated in mcm mutant cells that have been arrested in S phase [102]. In addition, S. pombe rad2 mutants are synthetically lethal in combination with mutants of rad50, rhp51, or rhp54 (the S. pombe homologs of RAD50, RAD51, and RAD54), suggesting that recombination functions become essential when Okazaki fragment metabolism is compromised [58, 130, 132]. The association of impaired replication function with increased recombination has also been described in S. cerevisiae [36, 117] and prokaryotes [119], suggesting this is a general feature of S phase. Conversely, certain recombination mutants display S phase defects. In the S. pombe rad50 mutant, S phase is delayed relative to wild type and the cells are sensitive to HU [58]. In vertebrate cells, inactivation of the recombination proteins Rad51 [173] or Mre11 [32] leads to DNA strand breaks and cell lethality. These and other observations have led to the suggestion that recombination proteins are normal components of S-phase progression in eukaryotes that protect genome integrity [133, 135]. Thus, replication fork stalls and starts may occur as part of normal S phase in eukaryotes, as has been described in prokaryotes [33, 119]. There are several possible consequences of a stalled replication fork, which may depend on its cause. Ideally, fork structure is protected and its components remain assembled during the arrest (Figure 3). However, the fork may lose structural integrity if this protection fails, resulting in its collapse and the generation of DNA breaks; these breaks are likely to be lethal to the cell if they are not repaired [33, 119]. Recombination is one mechanism that can reestablish a replication fork from a DNA break [33, 120]. Although recombination-dependent replication has been best characterized in prokaryotes, there is evidence that a similar process operates in eukaryotes. In S. cerevisiae, break-induced replication (BIR) can replicate hundreds of kilobases of DNA starting from a chromosomal break [92]. In S. pombe, cells lacking telomerase can replicate telomere sequences, presumably by a recombinational mechanism [137]. Importantly, replication mediated by recombination is predicted to be independent of replication origins and origin proteins. Thus, there may be mechanistic links between recombination and replication throughout S phase
Chapter 1: Replication in Fission Yeast
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in S. pombe and other eukaryotes, which are likely to be significant for the maintenance of overall genome stability.
Figure 3. Model for replication restart (based on 33,119). When cells are treated with HU, replication forks stall. If the structure of the fork can be maintained through the arrest, then the fork may resume synthesis once HU is removed from the media. If the fork structure cannot be maintained, the fork may collapse, generating DNA double-strand breaks. Recombination is one mechanism that may repair DNA breaks and reestablish stalled replication forks.
8
COORDINATION OF S PHASE EVENTS WITH MITOSIS
The events of S phase are closely linked with the later events of the cell cycle. As described above, maintenance of the order of the cell cycle and alternation of S phase with mitosis preserves the genome integrity. In addition, chromosomal processes such as sister-chromatid cohesion and silencing are regulated coordinately with the replication of the DNA. Intriguingly, the replication fork is the one structure that contacts all of the DNA, once, in a single cell cycle. This positions the replication machinery in a unique state to monitor and modify any region of the genome during S phase. Cohesion between newly replicated sister chromatids holds them together from S phase until their separation during mitosis. This arrangement is essential for the proper attachment of kinetochores to the microtubule spindles and correct segregation of the chromatids during mitosis [139]. The
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close apposition of sister chromatids throughout G2 and M phase likely also facilitates repair of damaged DNA off the homologous template [169]. Cohesion is mediated by a conserved complex of proteins first identified in S. cerevisiae, the cohesins [57, 118]. Homologs of the cohesins also exist in fission yeast [15, 187, 192] and have provided important insights as to how cohesion assembly is regulated during S phase. The fission yeast Eso1 protein is critical for the activation of cohesion function, although it is not required for the assembly of cohesin proteins with chromatin [186]. Interestingly, Eso1 appears to be a fusion of two budding yeast proteins: the cohesin activator Eco1 and the DNA damage bypass polymerase Rad30. This homology suggests a direct link between cohesion and DNA damage repair. In fission yeast, cohesion sites along the chromosome have been identified and are particularly concentrated at the repeat regions of centromeres [14, 192]. Like human centromeres, S. pombe centromeres are relatively large (compared to S. cerevisiae centromeres) and contain heterochromatin. Two recent studies indicate that the heterochromatin protein Swi6 (the equivalent of mammalian HP1) is specifically required for the efficient assembly of cohesion proteins to the centromere [14, 144]. Cohesion association with chromatin arms occurs independently of Swi6. Unexpectedly, the Hsk1 kinase also has a role in sister-chromatid cohesion [171, 181]. Like Swi6, Hsk1 specifically influences the association of the cohesin complex with centromeres [8]. Swi6 physically interacts with both Hsk1 and its subunit Dfp1, suggesting direct recruitment of cohesin to centromeres by these replication proteins [8]. These findings suggest a role for Hsk1 and Dfp1 in sister-chromatid cohesion that may be separable from their role in replication initiation. Replication proteins also affect heterochromatin function. Within regions of heterochromatin, including centromeres, most genes are normally silenced. At least in budding yeast, silencing involves progression through S phase, although perhaps not fork passage per se [50, 88, 99, 121]. As noted above, eukaryotic ORC proteins are required for gene silencing and localize to heterochromatin regions in addition to replication origins [13]. In S. cerevisiae, the PCNA [209] [43], RFC, and Cdc45 proteins also contribute to silencing [43]. In S. pombe, mutants of DNA polymerase α display defects in silencing [138]. DNA polymerase α interacts with Swi6 and is required for its proper localization to heterochromatin [2, 138] suggesting that disruption in silencing can be attributed to defective heterochromatin structure. hsk1 mutants also display a defect in silencing, even though the Swi6 protein is localized to heterochromatin [8]. This suggests that heterochromatin function depends on the assembly of other proteins with Swi6; whether this requires a
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passing replication fork or other aspects of S-phase progression remains to be determined.
9
DNA REPLICATION IN FISSION YEAST MEIOSIS
Meiosis is a specialized cell cycle that generates recombinant, haploid progeny cells from a diploid cell. The meiotic cell cycle differs from the vegetative cell cycle in two outstanding respects. First, the S phase that occurs prior to meiosis (premeiotic S) is followed by two successive rounds of chromosome segregation, rather than alternating between S phase and mitosis. Second, the first meiotic division is reductional, resulting in the maintenance of cohesion between sister chromatids but the separation and segregation of homologous chromosomes. The preparation for this modified cell division cycle involves lengthy interaction between homologous chromosomes during the prophase stage of meiosis I but is likely to initiate as early as premeiotic S phase. Premeiotic S phase is longer than S phase in vegetative cells in most organisms [26]. The cause of this difference is unclear, since experiments in budding yeast suggest that the same replication origins are active in vegetative and meiotic cells [30]. However, other experiments in S. cerevisiae suggest that meiosis-specific chromosomal factors required during prophase might assemble during premitotic STET [26]. Recent studies in S. pombe have addressed the question of whether the replication machinery that functions during premeiotic S phase is the same as that utilized in the vegetative cell cycle [48, 103, 128]. S. pombe proteins required for the actual synthesis of the DNA in vegetative cells, such as DNA polymerase α and ribonucleotide reductase, also are essential for premeiotic S phase [48]. In contrast, mutants defective in initiation of DNA replication, such as the mcm mutants and cdc18, display different phenotypes in meiosis and mitosis. In mitosis, conditional alleles of these mutants allow bulk DNA replication but cause cells to arrest in late S phase [49, 81, 102]. In contrast, in similar conditions these mutants can proceed through the meiotic divisions and sporulate [48]. With more extreme conditions, these mutants delay replication and the subsequent meiotic divisions [103, 128]. This may reflect a quantitative difference: meiotic cells may tolerate a lower amount of certain replication proteins than that needed during the vegetative cell cycle. It is also possible that other meiotic factors, perhaps recombination proteins, can contribute to premeiotic replication. The S. pombe MCM protein complex is associated with chromatin during premeiotic S phase [103], consistent with the hypothesis that these proteins
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function in premeiotic DNA replication. However, MCM proteins are not localized to chromatin in between the meiotic divisions, when an additional round of DNA replication is suppressed [103]. Similar to the vegetative cell cycle, the meiotic cell cycle is subject to checkpoint controls. Fission yeast cells that have been induced to enter meiosis block the cell cycle when treated with HU [48, 128]. As in the vegetative cell cycle, HU-induced arrest during meiosis is likely to be checkpoint-dependent. Future work should resolve the components of this response, which are likely to be essential for the viability of gametes. Importantly, premeiotic S phase is closely coupled to the downstream events of meiosis such as recombination [47]. This has been best demonstrated in budding yeast, where blocks to premeiotic DNA synthesis prevent meiotic recombination and changes in the timing of premeiotic replication result in corresponding changes in the timing of initiation of meiotic recombination [18]. The molecular mechanism by which this occurs is still unclear [47].
10
REPLICATION: A CONSERVED PROCESS
Most, if not all, of the proteins involved or implicated in DNA replication in S. pombe have homologs in other eukaryotes including human cells. Analysis of these proteins in S. pombe and other model organisms has greatly facilitated the identification and characterization of the human counterparts. However, many of the details of regulation of the replication process differ between yeast and human cells, which may reflect internal differences in cell cycle control between the different organisms as well as the additional complexity required in multicellular organisms. S. pombe cells maintain genome stability and order of the cell cycle through multiple and overlapping pathways. In particular, the process of DNA replication itself is tightly regulated so that the genome is duplicated once, entirely and accurately, in each cell cycle. Mechanisms are also in place to restrain passage into mitosis until replication is complete. It is possible that additional mechanisms, as yet uncovered, also act to maintain the integrity of the genetic material. S. pombe cells that have lost this control capability may re-replicate DNA, lose chromosomes, and die. In comparison, deregulation of S-phase control in human cells may result in genome instability and contribute to cancer progression.
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17
PATHWAYS TO CANCER FROM REPLICATION DEFECTS
The recent completion of the S. pombe genome [205] revealed an impressive number of fission yeast genes with human homologs implicated in cancers. Interestingly, many of these genes have known or implied functions in DNA replication in S. pombe (Table 1). Current evidence suggests that multiple pathways of control of eukaryotic DNA replication can be disrupted to result in genome instability and predisposition to cancer [133]. Thus, deregulation of CDK activity, impaired origin firing, changes in the timing of firing, loss of control in the order of S phase and M phase, and inability to limit replication to once per cell cycle are all mechanisms that may lead to changes in chromosome structure and gene function [17, 80]. In addition, defects in the checkpoint response to replication blocks, and the inability to respond appropriately to stalled replication forks, also contribute to genome instability. Ultimately, the gain or loss of genetic information may lead to inappropriate expression of proto-oncogenes or loss of tumor-suppressor function [59]. When the normal timing of origin firing is disrupted, cells are susceptible to deregulated cell cycle progression. This could result either through refiring of origins in a single cell cycle, or through firing late origins of replication under conditions where they are normally prevented from firing. Treatment of S. cerevisiae cells with the antitumor drug adozelesin changes the normal pattern of replication such that active replication forks are blocked, but silent origins are activated [200]. Translocations or amplifications of mammalian chromosomes also may alter replication timing of a particular sequence [165, 177]. Conversely, uncontrolled cell proliferation may result in deregulation of replication timing. This is observed both in checkpoint mutants in S. pombe [86] and in human cancers [38]. Thus, disruption of the timing and coordination of replication is one pathway toward genome instability. Cells extend multiple, overlapping control mechanisms to restrict DNA replication to once per cell cycle. In S. pombe, this is accomplished by regulation of CDK kinase activity, phosphorylation, and destruction of Cdc18, and phosphorylation of STET [80]. In human cells, the Cdc18 equivalent Cdc6 is also negatively regulated by CDK phosphorylation [115], suggesting that regulation of human Cdc6 likewise contributes to prevention of re-replication. The MCM proteins are another CDK target, at least in some organisms [17]. There are several examples of deregulated CDK activity associated with cancers [42, 174].
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Table 1. Fission yeast replication-related proteins and their human homologs S. pombe protein PreRC Orp1–6
ORC1–6
MCM2–7
MCM2–7
Human equivalent
Cdc18 Cdc6 Cdt1 Cdt1 Initiation Rad4/Cut5 TopBP1 Sld3 ? Sna41 Cdc45 Drc1 ? Hsk1 Cdc7 Dfp1 Dbf4, ASK Synthesis Pol1/Swi7 Pol α Cdc6 Pol δ Cdc20 Pol ε Cdc17 DNA ligase PCNA PCNA RFC1–5 RFC1–5 RPA RPA Rad2 FEN1 Cdc24 ? Checkpoint/Cell Cycle Rad3 ATR, ATM Cds1 Rad53/CHK2 Chk1 CHK1 Rad17 RAD17 Cdc2 CDC2 Mrc1 Claspin Swi1 ? Recombination/Repair Rad50 RAD50 Mre11 MRE11 Nbs1 NBS1 Rhp51 RAD51 Rhp54 RAD54 Rqh1
BLM, WRN, RecQL4
Cohesion/Chromatin Swi6 HP1 Rad21 Rad21 Eso1 Rad30/Pol µ
Role in cancer (References)
Upregulated in cancer cells [4, 64, 154, 167, 183, 202]; Mcm2 (Bm28/CDCL1) is associated with acute myeloid leukemia [122, 190] Upregulated in proliferating cells [202] Upregulated in cancer cells [5] BRCT domains found in BRCT and XRCC1 [19]
Upregulated in cancer cells [63]
Upregulated in cancer cells [176] Upregulated in proliferating cells [183, 202] Overproduced in cancer cells [191]
Ataxia telangiectasia [159] Li-Fraumeni [12] Overexpressed in cancer cells [9] [42]
Ataxia-telangiectasia-like disorder (Mre11), Nijmegen syndrome (Nbs1) [35]
Bloom’s syndrome (BLM), Werner’s syndrome (WRN), Rothmund-Thomson Syndrome (RecQL4) [51, 196] Downregulated in cancer cells [89] Downregulated in cancer cells [83] Xeroderma pigmentosum variant [71, 113]
Overexpression of certain replication proteins, such as Cdt1, can promote tumor formation in mammals [5]. In addition, many replication proteins are specifically upregulated in cancer cells. Human Cdc7 (the homolog of the S. pombe Hsk1 kinase) is overexpressed in certain tumor cells [63]. Furthermore, human MCM proteins are specifically expressed (or overexpressed) in cycling cells and are not detectable in quiescent cells. An
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important consequence of these findings is that the presence of the MCM proteins in cells provides a sensitive diagnostic marker for proliferating cells. MCM proteins are detected in cells that have exited quiescence and reentered the cell cycle; thus, MCM proteins are detected in precancerous cells as well as in tumor cells [4, 64, 154, 183, 190, 202]. MCM transcription is further upregulated by activated oncogenes [167]. Interestingly, human BM28/CDCL1 (the homolog of Mcm2), maps to a chromosomal locus associated with acute myeloid leukemia, suggesting BM28/CDCL1 as a candidate oncogene [122]. Damage tolerance and repair mechanisms are also essential to prevent genome instability. In S. pombe, Rqh1 is needed for recovery from replication blocks [131, 175]. Human cells have at least five Rqh1 homologs, three of which are linked with cancer susceptibility syndromes. Mutations in BLM are associated with Bloom’s syndrome, mutations in WRN lead to Werner’s syndrome, and mutation of RecQL4 results in Rothmund-Thomson syndrome [51, 196]. Hyperrecombination and cancer susceptibility are characteristic of both Bloom’s and Werner’s syndromes. Inappropriate recombination due to the loss of other S-phase functions may generate deletions or expansions in the genetic information, as has been demonstrated in S. cerevisiae [134, 135]. Polymerase slippage may contribute to the formation of triplet repeat sequences, which are associated with several disorders including Huntington’s disease [93]. Checkpoint genes are important gatekeepers of genome stability [59]. Mutations in the ATM checkpoint kinase are linked to ataxia telangiectasia [159], and mutations in the checkpoint kinase Cds1 (also called CHK2) are found in a subset of patients with Li-Fraumeni syndrome [10, 12]. In addition, Rad17 (one of the checkpoint rad proteins) is overexpressed in certain types of human cancers [9]. The corresponding S. pombe proteins (Rad3, Cds1, and Rad17) are all involved in the cellular response to replication blocks [25]. The S. pombe Rad4/Cut5 protein, which also has a role in cellular checkpoints [156], contains a BRCT motif that is also present in the human BRCA1 tumor suppressor and the XRCC1 DNA repair protein [44, 157]. Thus, mutations that disrupt function of the replication checkpoint are also implicated in predisposition to cancer. Genomic instability leading to cancers may also result from chromosome structure defects caused by errors in S-phase processes linked to DNA replication. In S. pombe, the Eso1 protein is needed to activate cohesion so that sisterchromatids are held together until mitosis [186]. Part of the Eso1 protein is homologous to DNA polymerase η (Rad30), which is defective in the xeroderma pigmentosum variant syndrome characterized by predisposition to skin cancers [71, 113]. In addition, human securin, which
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normally prevents premature sister-chromatid separation, can induce cell transformation and tumorigenesis when overexpressed [211]. Expression of the Rad21 cohesin is downregulated in certain tumors [83]. In addition, HP1 is downregulated in breast cancer cells that are metastatic or invasive [89]. The S. pombe homolog of HP1, Swi6, recruits Rad21 to centromeres and other regions of heterochromatin [14, 144]. Recently, phosphorylation of another cohesin subunit, Smc1, has been shown to be required for the S-phase checkpoint in human cells [87, 207]. Taken together, these connections suggest a direct role for chromatin structure in maintenance of genome stability. Work from multiple experimental systems in recent years has revealed striking similarities in basic cellular processes among eukaryotes. In addition, synergy between studies of DNA replication, recombination, and chromosome structure has provided important insights into how DNA replication is integrated with other cellular processes. Studies of DNA replication in S. pombe and in other model systems has provided the proteins and pathways that serve as a framework for identification and characterization of the human counterparts. Collections of yeast mutants, as well as information about which genes together are essential for a process (synthetic lethality), are an important resource for understanding how mutations in distinct genes can result in similar phenotypes such as predisposition to cancers [60]. Several recent studies have supported the validity of yeast mutants with chromosome instability defects as models for the cellular response to cancer treatment drugs [76, 151, 168]. These simple eukaryotes continue to lead the way in fundamental understanding of normal and abnormal cell division.
ACKNOWLEDGMENTS Work in our laboratory was supported by grants from the NIH, NSF, and the American Cancer Society. S.L.F. is a Stohlman Scholar of the Leukemia & Lymphoma Society. J.M.B. is a fellow of the Damon Runyon Cancer Research Fund (DRG-1634). We thank Michael Catlett and Han-Kuei Huang for comments on the manuscript.
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ADDENDUM Since this chapter was written, several important discoveries have advanced our understanding of how DNA replication contributes to genome stability. First, progress has been made toward uncovering the functions of proteins associated with replication origins. Unexpectedly, an additional, conserved protein complex essential for DNA replication has been identified. The GINS complex consists of four subunits (Sld5/Cdc105, Psf1/Cdc101, Psf2/Cdc102, and Psf3) that associate with replication origins and are required for replication initiation and elongation [10, 12, 21]. In S. pombe, the Psf2 subunit was identified in a screen for mutants that prevent rereplication; similar to its homologs in other organisms, S. pombe Psf2 is required for normal S-phase progression [4]. Other recent reports have uncovered details about the requirements for replication protein association with origins and activity on their targets. The S. pombe Cdc23 protein has been shown to be required for Hsk1-dependent phosphorylation of the MCM complex [13], as well as for the recruitment of Cdc45 to chromatin [6]. Cdc45 appears to be one of the last proteins to associate with the prereplicative complex, as its localization requires formation of the prereplicative complex, as well as Hsk1 and Rad4 [3, 6]. Cdc45 has been suggested to travel with the replication fork in S. cerevisiae [1, 22] and is required for the in vitro helicase activity of the heterohexameric MCM complex [14]. A second area of research progress has been in understanding how cells respond to S-phase damage and replication blocks. S. pombe Cds1, like its homolog Rad53 in S. cerevisiae, is required to prevent replication fork collapse in the presence of HU [15]. Importantly, two proteins that mediate Cds1 function, Mrc1 and Swi1/Tof1, have been shown to associate with the replication fork even in the absence of DNA damage, suggesting a mechanism to couple detection of damage by the moving fork [11, 15, 17]. In mammalian cells, HU-induced checkpoint activation leads to the association of MCM7 with the Rad17 and ATRIP/ATR checkpoint proteins [23] and phosphorylation of MCM2 and MCM3 [2, 9, 24]. Interestingly, the mammalian checkpoint kinases appear to affect not only fork stabilization during damage, but also the timing of origin firing during normal S-phase progression [19]. Together, these findings strongly suggest a central, conserved link between S-phase checkpoints, MCM proteins, and replication fork stability. Interestingly, fission yeast checkpoint mutants predicted to cause replication fork collapse do not cause cell inviability in meiosis [18]. Meiotic cells may tolerate a higher level of DNA damage because programmed DNA damage is generated during meiotic recombination. Lastly, our understanding of the links between replication and human disease has been strengthened. Seckel syndrome, a human disease that predisposes to chromosome instability and shares phenotypes with DNA
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repair disorders [16] is associated with mutation of the checkpoint protein kinase ATR (Rad3 in fission yeast). Genes involved in the Fanconi anemia pathway (such as BRCA and Rad51) are required for the response to replication blocks [8]. Interestingly, Rad51, which is implicated in leukemia [20] has been shown to influence the rate of replication fork progression during S-phase DNA damage [7]. Additionally, there is an ever-broadening literature on the use of MCMs and other conserved replication proteins as markers of hyperproliferating cells, and as measures of cancer prognosis (rev. in [5]). Many of these recent discoveries have been carried out in human cells, but clearly build on basic research in yeast that continues to identify new genes and regulatory interactions. Thus, studying DNA replication in simple model systems continues to provide a crucial foundation for understanding pathways of cancer initiation and progression.
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10. Kanemaki, M., A. Sanchez-Diaz, A. Gambus, and K. Labib. 2003. Functional proteomic identification of DNA replication proteins by induced proteolysis in vivo. Nature 423:720–724. 11. Katou, Y., Y. Kanoh, M. Bando, H. Noguchi, H. Tanaka, T. Ashikari, K. Sugimoto, and K. Shirahige. 2003. S-phase checkpoint proteins Tof1 and Mrc1 form a stable replication-pausing complex. Nature 424:1078–1083. 12. Kubota, Y., Y. Takase, Y. Komori, Y. Hashimoto, T. Arata, Y. Kamimura, H. Araki, and H. Takisawa. 2003. A novel ring-like complex of Xenopus proteins essential for the initiation of DNA replication. Genes Dev. 17:1141–1152. 13. Lee, J. K., Y. S. Seo, and J. Hurwitz. 2003. The Cdc23 (Mcm10) protein is required for the phosphorylation of minichromosome maintenance complex by the Dfp1-Hsk1 kinase. Proc. Nat. Acad. Sci. USA 100:2334–2339. Epub 2003 Feb 25. 14. Masuda, T., S. Mimura, and H. Takisawa. 2003. CDK- and Cdc45-dependent priming of the MCM complex on chromatin during S-phase in Xenopus egg extracts: possible activation of MCM helicase by association with Cdc45. Genes Cells 8:145– 161. 15. Noguchi, E., C. Noguchi, W. H. McDonald, J. R. R. Yates, and P. Russell. 2004. Swi1 and Swi3 are components of a replication fork protection complex in fission yeast. Mol. Cell Biol. 24:8342–8355. 16. O’Driscoll, M., V. L. Ruiz–Perez, C. G. Woods, P. A. Jeggo, and J. A. Goodship. 2003. A splicing mutation affecting expression of ataxia-telangiectasia and Rad3-related protein (ATR) results in Seckel syndrome. Nat. Genet. 33:497–501. Epub 2003 Mar 17. 17. Osborn, A. J., and S. J. Elledge. 2003. Mrc1 is a replication fork component whose phosphorylation in response to DNA replication stress activates Rad53. Genes Dev. 17:1755–1767. 18. Pankratz, D. G., and S. L. Forsburg. 2005. Meiotic S-phase damage activates recombination without checkpoint arrest. Mol. Biol. Cell 2005 Feb 2; [Epub ahead of print]. 19. Shechter, D., V. Costanzo, and J. Gautier. 2004. ATR and ATM regulate the timing of DNA replication origin firing. Nat. Cell Biol. 6:648–655. Epub 2004 Jun27. 20. Slupianek, A., C. Schmutte, G. Tombline, M. Nieborowska-Skorska, G. Hoser, M. O. Nowicki, A. J. Pierce, R. Fishel, and T. Skorski. 2001. BCR/ABL regulates mammalian RecA homologs, resulting in drug resistance. Mol. Cell 8:795–806. 21. Takayama, Y., Y. Kamimura, M. Okawa, S. Muramatsu, A. Sugino, and H. Araki. 2003. GINS, a novel multiprotein complex required for chromosomal DNA replication in budding yeast. Genes Dev. 17:1153–1165. 22. Tercero, J. A., K. Labib, and J. F. Diffley. 2000. DNA synthesis at individual replication forks requires the essential initiation factor Cdc45p. EMBO J. 19:2082–2093. 23. Tsao, C. C., C. Geisen, and R. T. Abraham. 2004. Interaction between human MCM7 and Rad17 proteins is required for replication checkpoint signaling. EMBO J. 23:4660– 4669. Epub 2004 Nov 11. 24. Yoo, H. Y., A. Shevchenko, A. Shevchenko, and W. G. Dunphy. 2004. Mcm2 is a direct substrate of ATM and ATR during DNA damage and DNA replication checkpoint responses. J. Biol. Chem. 279: 53353–53364.
Chapter 2 DISSECTING LAYERS OF MITOTIC REGULATION ESSENTIAL FOR MAINTAINING GENOMIC STABILITY Jennifer S. Searle 1 and Yolanda Sanchez 2 1
Department of Molecular Genetics, Biochemistry and Microbiology, The University of Cincinnati, 231 Ablert Sabin Way, Cincinnati, OH 45267-0524; 2Department of Pharmacology and Toxicology, Dartmouth Medical School, 7650 Remsen, Hanover, NH 03755, USA
Cell division is the process by which a cell copies its DNA and cellular constituents and gives rise to two cells. One complete cell division is termed a cell cycle. (Although you can have a cell cycle without cell division.) This process is divided into four distinct phases for most eukaryotic cells: G1, S, G2, and M (Figure 1A). S phase (for synthesis) is the stage in which the DNA is replicated and M phase, (for mitosis), is the stage in which the replicated DNA condenses and the chromosomes are divided equally into two cells. G1 and G2 are “gap phases” that separate DNA replication and mitosis. This chapter is focused on how yeast have served as a useful model system that was instrumental in the identification of proteins that promote cell cycle progression in all eukaryotes. We will discuss the pioneering work of individuals that used budding yeast to reveal the layers of regulation that coordinate progression through the cell cycle. In addition, we will discuss how this model organism that is amenable for genetics has been used to identify proteins that regulate cell cycle, and in particular mitosis, in order to safeguard the integrity of the genome. We will end by linking this work to the specific problem of cancer, and mention the utility of this system in research designed to establish and validate targets for anticancer drug discovery.
37 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 37–73. © 2007 Springer.
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1
THE MANY LAYERS OF CELL CYCLE REGULATION
1.1
The cell cycle engine is composed of a cyclin and a cyclin-dependent kinase (Cdk)
For many years, scientists sought to identify the molecular components that regulate the duplication and segregation of chromosomes. Several groups of biochemists working on clam, sea urchin, and frog eggs had been studying the activity of maturation-promoting factor (MPF), which causes immature eggs to undergo meiosis and prepares them for fertilization [78, 110]. MPF from many species could drive mitosis when injected into cells that were at stages other than mitosis. MPF could also induce replication and condensation cycles of sperm DNA, mimicking the chromosome division cycle in a cell-free system [76]. MPF activity rose in cells that were entering meiosis or mitosis and would then drop precipitously after cell division. A new burst of activity was followed by another chromosomal division [34]. At the time, MPF was coined as the “cell cycle clock” and many groups attempted to purify its components but failed. Tim Hunt reasoned that a protein functioning to promote mitosis should have a pattern of expression that coincided with the induction of mitosis in a cell-free extract system. Hunt used sea urchin and Xenopus extracts to identify a protein whose accumulation levels cycled (cyclin) in a similar pattern as did the activity of the MPF, and the translation of which was required for Xenopus extracts to enter mitosis [28, 83]. Meanwhile, the geneticists interested in this problem took a different approach. They postulated that if proteins had a role in the progression through the cell cycle by either promoting a transition (G1 to S or G2 to M), or by acting as a substrate for the next transition to occur, then the genes encoding these proteins could be identified in a genetic screen. The screen consisted of randomly mutagenizing yeast and selecting for conditional mutants (functional at one temperature, and inactive at a higher or restrictive temperature) that would cause the cells to arrest at particular stages of the cell cycle when the cells were shifted to the restrictive temperature. Budding yeast was the optimal organism for geneticists doing cell cycle research because the cells divide by budding and the size of the bud indicates their position in the cell cycle. Yeast is also easy to grow and is amenable to genetic manipulation. Leland Hartwell carried out a genetic screen to identify conditional mutants in the budding yeast Saccharomyces cerevisiae. Since many of the components that drive cell division are essential (the cell
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cannot live without them), some of the mutants that Hartwell identified had defects in cell cycle progression (cdc = cell division cycle mutants). Hartwell identified dozens of mutants that arrested at different stages of the cell cycle when the cells were raised to the restrictive temperature. The arrest point was easy to identify as the cells arrested with no bud (G1) or with a uniform bud size (Figure 1B). One such mutant, cdc28, was defective in the initiation of DNA replication [43]. The cdc28 mutation caused cells to arrest without a bud and with unreplicated DNA at the restrictive temperature. Paul Nurse and Pierre Thuriaux carried out a screen similar to Hartwell’s using the fission yeast Schizosaccharomyces pombe. They identified a mutant, cdc2, which arrested prior to mitosis under restrictive conditions [92]. When the CDC2 gene was cloned, it was found to encode a protein with homology to kinases. Protein kinases are enzymes that modify other proteins by adding phosphate groups, a common mechanism used by the cell to transmit information or regulate protein activity. David Beach, while working with Nurse, also found that the cdc2 mutation was in a gene homologous to the budding yeast CDC28 gene identified by Hartwell [5] and cloned earlier by Nasmyth and Reed [87]. In the late 1980s, investigators in the fields of biochemistry and genetics came together and initiated an explosion of cell cycle research. At that point, Manfred Lohka and James Maller succeeded in the purification of MPF by monitoring an activity that could promote chromosome condensation. Jean Gautier and Maller’s group joined with Nurse, and William Dunphy and John Newport joined with David Beach and both groups discovered that MPF is composed of two subunits; a kinase that is the product of the CDC28 and cdc2+ genes in the yeasts and the mitotic cyclin [27, 33]. Thus, it was found that the cell cycle engine is universal or conserved, and that in its bare bones is a complex, composed of a cyclindependent kinase (Cdk) and an activating partner, the cyclin [27, 33, 65, 72]. Interaction of the Cdk and cyclin activates the complex to phosphorylate substrates, allowing the cell to move into the next cell cycle stage. The cyclin partner not only activates the Cdk but also provides specificity for substrates that will drive a particular cell cycle transition. For example, a complex containing a Cdk and a member of the G1 cyclin family promotes entry into a new cell division cycle; whereas a Cdk/Mitotic cyclin complex promotes entry into mitosis. Thus, the availability of the cyclins serves to regulate the cell cycle stage-specific activity of the Cdks. The 2001 Nobel Prize for medicine was awarded to three individuals who used biochemistry and genetics to discover the Cdk complex. Tim Hunt discovered the mitotic cyclin and, importantly, he found that degradation of the cyclin coincided with the end of mitosis and, identified a layer of regulation of the cell cycle engine. Leland Hartwell carried out the famous
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genetic screen that led to the identification of the Cdc28/Cdk. Paul Nurse identified cdc2+ in a similar genetic screen and found that mammals had a gene that could function in the place of cdc2+. In doing so, he showed that the proteins that comprise the cell cycle engine are conserved between yeast and mammals [68]. These individuals, combined forces with other biochemists studying the activity of the cell cycle engine, which sparked the intensity of the field that studies how cell division, chromosome duplication, and segregation are coordinated. [86, 98]. Many years after the discovery of the Cdk complex, Pavletich and coworkers analyzed the crystal structure of the cyclin/Cdk complex, which revealed the mechanism by which the cyclin activates the Cdk. It turns out that the helix containing a conserved amino acid involved in ATP binding is positioned away from the active site of the Cdk. The cyclin, by virtue of its binding, realigns the residues involved in ATP binding and keeps domains from blocking the catalytic cleft of the enzyme. Thus, binding of the cyclin removes a stearic block that allows the cdk to phosphorylate its substrates [55]. This is the reason why MPF activity coincides with a peak of cyclin levels. Phosphorylation of the activation loop is required to stabilize the active form of the Cdk complexed with the cyclin, and thus adds another level of regulation to this complex [95].
Figure 1A. The eukaryotic cell cycle and the universal cell cycle engine. The eukaryotic cell cycle (with the exception of germ cells and early embryonic cells) is divided into four distinct phases G1, S, G2, and M. S phase (for synthesis) is the stage in which the DNA is replicated and M phase, mitosis, is the stage in which the replicated DNA condenses and the chromosomes are divided equally into two daughter cells. G1 and G2 are “gap phases” that separate DNA replication and mitosis. The cyclin-dependent kinase/cyclin (Cdk/cyclin) complex which drives cell cycle transitions is regulated by phosphorylation of the Cdk.
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Figure 1B. Progression of Saccharomyces cerevisiae cells through the cell cycle can be monitored by bud and nuclear morphology. Bud emergence coincides with the initiation of DNA synthesis (DNA replication), G2 cells have large buds and an undivided nucleus at the neck. Cells in anaphase have an elongated bipolar nucleus (dumbbell shape) and elongated spindles. Telophase cells have two distinctly divided nuclei in the mother and daugher cells. Cytokinesis gives rise to two unbudded cells that are in G1. Indicated is the morphology with which the cdc mutants arrest at each point in the cell cycle.
2
LAYERS OF REGULATION: PART I. HOW YEAST GENETICS AND BIOCHEMISTRY REVEALED THE FIRST LAYER OF REGULATION
2.1
Phosphorylation and proteolysis regulate the activity of the cell cycle engine
Using genetic and biochemical analyses, researchers in the last 15 years have deciphered a complex regulatory web, involving phosphorylation, ubiquitination-mediated proteolysis and control of subcellular localization that functions to achieve the exquisite order and timing of progression through the stages of the cell cycle. For the remainder of this chapter, we will focus on the layers of regulation that ensure that mitosis results in the transmission of genetic information with high fidelity. Therefore, it is important to review here the stages of mitosis that result in the segregation of chromosomes and the generation of two daughter cells.
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Following activation of MPF and chromosome condensation, and prior to separation of the replicated chromosomes, the cells are in the metaphase stage of mitosis (see Figure 2). Metaphase. The condensed chromosomes are attached to the spindle and the sister chromatids remain associated to each other via their centromeres (kinetochores), and by the presence of cohesin and condensing complexes (see Figure 2) [39, 46, 62]. In metazoans the chromosomes align themselves in the mid-zone in a structure called the metaphase plate. Anaphase. The cohesion between the sister chromatids is lost and the sisters separate due to the force exerted by the spindle, which is the physical structure that separates the chromosomes so that the two daughter cells receive a complete copy of the genome [62]. This is achieved by the cleavage of the cohesin complexes by the enzyme separase (Esp1 in yeast) [16]. Telophase. The separated chromatids move to opposite poles of the cell. Mitotic exit. The mitotic Cdk/B-type cyclin complexes are inactivated via many mechanisms (discussed below). This allows the pre-replication complexes to assemble on the chromatin so that the cell is ready to begin DNA replication at the next S phase. Cytokinesis. The physical separation of the cytoplasm that results in the two daughter cells.
Figure 2. Progression through mitosis. Following activation of MPF and chromosome condensation and prior to separation of the replicated chromosomes, the cells are in the metaphase stage of mitosis. In metaphase, the condensed chromosomes are attached to the spindle, and the sister chromatids remain associated to each other via their centromeres (kinetochores), and by the presence of cohesin complexes (Scc1, Smc1, and Smc3). Ubiquitinmediated destruction of securin (Pds1) leads to release of separase (Esp1), which cleaves cohesin complexes. The cohesion between the sister chromatids is lost which triggers
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anaphase where the sisters separate due to the force exerted by the spindle. The separated chromatids move to opposite poles of the cell. The mitotic Cdk/B-type cyclin complexes are inactivated via many mechanisms, which allows the pre-replication complexes to assemble on the chromatin so that the cell is ready to begin DNA replication at the next S phase.
2.1.1
Regulation of the Cdk/cyclin complex by phosphorylation
Biochemical and genetic experiments identified kinases and phosphatases (enzymes that remove the phosphates added by the kinases) that regulate Cdk complexes (see Figure 1). The Cdk/cyclin complex can be regulated by (1) positive (see above) and negative phosphorylation on the Cdk subunit; (2) the expression and destruction of the cyclin component; (3) binding of inhibitors (Cdk inhibitors=CKI’s); and (4) regulating the subcellular localization of the complex [95]. Experiments carried out in the fission yeast S. pombe, and the Xenopus extract system provided the first clues as to how this complex is regulated by phosphorylation. The studies of fission yeast mutants carried out by Nurse identified the Wee1 family of kinases, that function to maintain the Cdk/cyclin complex in an inactive state until the cell is ready to enter the next phase of cell division [91]. Genetic and biochemical studies also led to the identification of the Cdc25 family of phosphatases, which catalyze the removal of the inhibitory phosphates added by the Wee1 kinases to the Cdk subunit [32, 103]. For example, during the late stages of DNA replication, the mitotic cyclins are expressed and assemble with their cdk partner (Cdk1); however, this complex is maintained in an inactive state by inhibitory phosphorylation carried out by the Wee1 kinases. At the end of DNA replication, and before the onset of mitotic events, the inhibitory phosphates are removed by the Cdc25 family of phosphatases, thus setting off an amplification of mitotic Cdk/Cyclin activity. This regulatory network is utilized in the fission yeast and metazoans to prevent premature onset of mitosis [9, 96, 105, 111, 130] and in the budding yeast to regulate mitosis until the cell has reached a critical bud size [45, 79]. In response to perturbations in bud construction, Swe1 (budding yeast Wee1) and Mih1 (budding yeast Cdc25 phosphatase) are regulated to delay mitotic activation of Cdc28 [118]. Failure to block mitosis when bud construction or critical size have not been completed results in binucleate cells [42, 45, 112]. 2.1.1.1 Regulation of the cell cycle by ubiquitin-mediated proteolysis The biochemists Hunt, Kirchner, and Ruderman, among others [4, 28, 60, 85] had identified that the cyclins were regulated by proteolysis, and more importantly, that degradation of the B-type cyclin was required for cycling
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Xenopus extracts to exit mitosis. Expression of a nondegradable cyclin mutant would lock the extracts in mitosis [85]. Using purification and fractionation approaches, these investigators identified an activity that would ligate ubiquitin to the cyclin component and trigger its destruction. A structure called the proteosome would then recognize the proteins with polyubiquitin chains and process them down to their amino acid components (degradation). This activity of ligating ubiquitin to the B-type cyclin was given the name of the anaphase-promoting complex or cyclosome (APC/C) [116, 4]. Many of the mutants identified by Hartwell exhibited an arrest either prior to DNA replication (but were able to bud) or exhibited a large-budded arrest, which indicated that the cells were able to replicate their DNA, but were unable to complete mitosis. These mutants turned out to be deficient in components of the multi-protein complexes involved in the ordered destruction of cell cycle regulators. There are three critical cell cycle transitions that are regulated by proteolysis: the G1 to S, metaphase to anaphase, and anaphase to G1 transitions. Several of the cdc mutants that arrested at the G1 to S transition had high protein levels of the G1 cyclins (Clns) and of the Cdk kinase inhibitor (CKI) Sic1. Similarly, mutants that arrested in mitosis before anaphase did so with high levels of the B-type (mitotic) cyclin Clb2 and, presumably, with high levels of an anaphase inhibitor [54]. Mutants that had completed anaphase but arrested at telophase did so with high levels of the B-type cyclins, which mimicked the block to mitotic exit observed with the nondegradable B-type cyclin in metazoan extracts (Figure 3). Since cyclin degradation was a requirement for mitotic exit in the cell-free extract system, scientists predicted that the strains that arrested with high levels of the mitotic cyclin carried deficiencies in components of multi-protein complexes involved in ligating ubiquitin to cell cycle regulators in order to trigger their ordered destruction. There are two complexes that regulate the cell cycle via ubiquitin-mediated proteolysis: the Skip1, Cdc53, F box complex (SCF) and the APC/C. We will focus our attention on the APC, or cyclosome, which is a multi-protein complex that acts as a ubiquitin ligase and is active in mitosis and G1. Once again, the geneticists and biochemists joined efforts in order to identify the enzymatic activity of a large multi-protein complex that is required for progression through mitosis in all eukaryotes. Stefan Irniger, when in Nasmyth’s laboratory, showed that two of the cdc mutants that showed a pre-anaphase arrest phenotype, cdc16 and cdc23, carried mutations in genes encoding proteins that were required for the proteolysis of the B-type cyclins [54]. Around the same time, Hieter’s group found that the Cdc16 and Cdc23 proteins formed a complex with Cdc27, another protein also required for
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proteolysis of the B-type cyclin [66]. Then Kirschner’s and Hieter’s groups joined forces to determine that Cdc27 and Cdc16 are indeed components of a large complex that catalyzes the conjugation of ubiquitin to the B-type cyclins and are thus, components of the evolutionarily conserved APC/C [60]. The biochemical experiments with the nondegradable B-type cyclin mentioned above always led to a block in mitotic exit but not to chromosome segregation at anaphase indicating that proteolysis of the B-type cyclins was not required for anaphase to occur. The observation that APC mutants that arrested in pre-anaphase did so with high levels of the B-type cyclin Clb2, and that B-type cyclin degradation is not required for anaphase to take place (biochemical experiments with nondegradable B-type cyclin and [118]), suggested that the APC/C had different substrates in order to promote anaphase and mitotic exit. The anaphase substrate of the APC/C, Pds1, was identified in a genetic screen carried out by Doug Koshland’s group that identified genes encoding proteins involved in chromosome segregation [135, 136]. In this screen, the investigators were looking for mutations that caused the cell to separate sister chromatids prematurely (Pds = precocious dissociation of the sisters). Orna Cohen-Fix, while in Koshland’s laboratory, showed that Pds1 was destroyed prior to anaphase and that it was a substrate of the APC/C [20]. The APC/C components and its substrates are conserved, thus Pds1 orthologues are now called securin, due to their function in keeping sister chromatids together until anaphase. The mechanism by which Pds1 keeps sister chromatids together is by acting as an inhibitor of a caspase-like protease Esp1 (also known as separase). Esp1 cleaves Scc1, a protein that is part of the cohesin complex (Scc1, Smc1, Smc3, and Scc3) that forms during DNA replication to keep the sister chromatids together. Cleavage of Scc1 by Esp1 leads to loss of cohesion along the arms of the chromosomes, and the tension provided by the spindle apparatus is then able to pull the sister chromatids apart at anaphase (see Figure 2). In summary, the APC promotes progression through mitosis by the ordered ubiquitinmediated proteolysis of the anaphase inhibitor, Pds1 (securin), and the B-type (mitotic) cyclins, Clb2 and Clb5 [66, 97, 123, 126, 127]. The APC relies on proteins that act as specificity factors in order to ubiquitinate the correct substrates in an ordered and timely manner. These proteins are the WD repeat (tryptophan and aspartic acid repeats) containing proteins Cdc20 and Hct1/Cdh1. Thus, cdc20, a mutant identified in the Hartwell screen that arrests prior to anaphase, is defective in the ubiquitination and destruction of both Pds1 (securin) and Clb2 (B-type cyclin). Since destruction of the B-type cyclins is not required for anaphase [118], this finding suggested that there existed a regulatory step that ensured that the
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destruction of securin preceded the destruction of the cyclins in order to ensure that the sister chromatids separate before the cell exits mitosis. This finding also explains why cells with defects in the destruction of securins would arrest with high levels of the B-type cyclins [19, 121]. In summary, entrance into mitosis is regulated at the level of activation of the Cdk/mitotic cyclin complex (Figures 1 and 3). High activity of the mitotic Cdk complex triggers mitosis but is inhibitory to the pathway that promotes exit from mitosis (Figure 3). Exit from mitosis requires the inactivation of this complex and reversal of Cdk phosphorylation, achieved by the APC-mediated proteolysis of the mitotic cyclins and release of the phosphatase Cdc14 from the nucleolus [107, 127, 127]. The mechanism that regulates the interdependence of anaphase and mitotic exit has recently come to light from work carried out in the laboratories of Kim Nasmyth, Angelika Amon, Doug Koshland, David Morgan, Uttam Surana, and Ray Deshaies among others. We will simplify the intricate regulatory network that regulates mitotic exit (MEN), however, the details can be found in the following papers and references therein [56, 97, 107, 114, 137]. Following a successful anaphase, the initial degradation of the B-type cyclins, Clb5 and Clb2 [109], leads to the activation of a signaling network that results in the release of the phosphatase Cdc14 from the nucleolus. This involves the activation of Tem1, a GTP-binding protein of the Ras superfamily, which is a positive regulator of Clb destruction (Figure 3). Tem1 is negatively regulated by a heteromeric GTPase-activating protein (GAP) composed of Bfa1 and Bub2. Inactivation of Bfa1 and Bub2 allows activation of Tem1 and triggers the release of Cdc14 from the nucleolus. Activation of the polo-like kinase Cdc5, also promotes Cdc14 release from the nucleolus. Cdc14 then reaches its targets to remove phosphates that have been added by the Cdk complex [127]. The activity of Cdc14 allows full activation of Hct1/Cdh1, the APC specificity factor for Clb2 ubiquitination, and dephosphorylation of Sic1, which causes this Cdk inhibitor to accumulate. These two events serve to lower the activity of the mitotic Cyclin/Cdk complexes even further, allowing mitotic exit and the establishment of pre-replication complexes [123, 126, 127]. cdc5 and cdc14 were identified in the Hartwell screen as mutations that caused cells to arrest after anaphase but before cytokinesis (telophase arrest, see Figure 1) and the proteins encoded by the genes mutated in these strains are positive regulators of Clb2 inactivation and mitotic exit.
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2.2
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Why does the cell have so many proteins to regulate cell division?
Cells must replicate chromosomes with high fidelity and segregate identical copies to two cells [17]. They must also duplicate their cellular constituents and maintain a critical size. In order to carry out this feat every time, cells must overcome two problems: The alternation problem: DNA must be replicated once per cell cycle. One example of a control that addresses the alternation problem is the exquisite regulation of Cdk kinase activity by the APC during mitotic progression, which ensures that cells complete mitosis prior to the initiation of another round of DNA replication. When these mechanisms fail, cells go through more than one replication cycle without an intervening mitosis. This results in hyperploidy, or too many sets of chromosomes. In some specialized cell types, such as trophoblasts, DNA replication without mitosis leads to the formation of tetraploid cells. The completion problem: Initiating a cell cycle transition is dependent upon the completion of the previous stage. For example, cells must finish DNA replication before beginning mitosis. When this dependence fails, cells will attempt to segregate incompletely replicated chromosomes, which can lead, in a worst case scenario, to a catastrophic or lethal mitosis. The first evidence that the cell had evolved mechanisms to address these two problems came from elegant experiments carried out in the 1970s by Potu Rao and Robert Johnson. These scientists used cell fusion experiments to show that there was an activity in S-phase cells that could promote initiation of DNA replication in a G1 cell in a fashion similar to MPF promotion of mitosis. That is, when a cell in the replication stage of the cell cycle (S phase) was fused to a G1 cell the G1 nucleus in the heterokaryon (the cell resulting from the fusion that had two nuclei) would begin DNA replication. Today we know that this S-phase-promoting factor is none other than Cdk complexes with S-phase cyclins. When Rao and Johnson fused a cell in S phase with a cell that had completed DNA replication but had not initiated its mitotic program (G2 cell) they made a very important discovery. The first observation was that the G2 nucleus in the heterokaryon would delay entry into mitosis until DNA replication was completed in the S-phase nucleus. The second observation was that the G2 nucleus did not reinitiate DNA replication. These observations indicated that there was a signal from the S-phase nucleus that would delay mitosis until replication was completed (completion problem); and that there was a mechanism in place in the G2 nucleus that prevented the reinitiation of DNA replication until the cell had gone through mitosis (the alternation problem). Therefore, with these now historical studies, Rao and Johnson demonstrated that cells had mechanisms
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in place that regulated the order and timing of cell cycle transitions. These mechanisms are referred to as checkpoint controls [98]. The controls identified by Rao and Johnson become essential when the relative timing of cell cycle transitions is challenged and cannot be merely regulated by an oscillator, or the “cell cycle clock”; such is the case when DNA replication is prolonged due to DNA damage or other stress conditions (Figures 4 and 5). Elegant experiments carried out by Smythe and Newport with cell free extracts from Xenopus eggs provided biochemical evidence of this response [111] by showing that incompletely replicated DNA blocks the activation of the mitotic Cdk/cyclin complex by maintaining the Cdk subunit in a phosphorylation form. The mechanisms that are activated when the relative timing of cell cycle transitions is altered are also important when the chromosomes are not correctly attached or oriented on the spindle or when a cell has not reached a critical size prior to division. These checkpoint controls are mediated by signal transduction pathways that ensure the interdependence of cell cycle events. For example, during DNA replication a signal is produced (probably at the replication fork, or when forks pause or stall] that will cause the cell to delay mitosis until all forks have completed replication. Because of their function, these biochemical pathways are called checkpoints, a term coined by Hartwell, and they represent the next layer of regulation of the cell cycle that is covered in this chapter.
Figure 3. Phosphorylation and ubiquitin-mediated proteolysis regulate ordered progression through mitosis. Entrance into mitosis is regulated by the abundance of the cyclin and by positive and negative phosphorylation of the Cdk subunit. The activation of the mitotic Cdk complex triggers mitosis but is inhibitory to the pathway that promotes exit from mitosis. The APC promotes progression through mitosis by the ordered ubiquitin-mediated proteolysis of
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the anaphase inhibitor, Pds1 (securin), and the B-type (mitotic) cyclins, Clb2, and Clb5. The APC/Cdc20 targets Pds1 for ubiquitination, this leads to activation of the separase and loss of cohesion (see Figure 2). The APC/Cdc20 then targets the Clbs for ubiquitination leading to the activation of the mitotic exit network (MEN) and release of the phosphatase Cdc14 from the nucleolus. Dephosphorylation of Hct1 and Sic1 serve to further inactivate the mitotic Cdk/Cyclin complexes and allows the assembly of pre-replication complexes and exit from mitosis.
3
LAYERS OF REGULATION: PART II. THE YEAST MODEL SYSTEM WAS INSTRUMENTAL IN THE DISCOVERY OF CHECKPOINT PATHWAYS
The genetic material of all organisms is constantly challenged by DNAdamaging agents from endogenous sources, such as the byproducts of respiration and from exogenous sources, such as UV radiation and environmental toxicants. In addition, during every S phase a number of the replication forks stall or collapse invoking mechanisms under study today to reactivate the fork or fix the damage resulting from the collapsed fork [23, 139]. Accurate transmission of chromosomes to each daughter cell requires that cells do not begin anaphase until all chromosomes have been completely replicated and correctly aligned on the spindle. Similarly, cells that have incurred DNA damage in G1 delay DNA replication until the damage has been repaired. Cells that have incurred damage in S phase slow down DNA replication and repair damage before they progress through mitosis [17, 44, 74] (Figure 4). Genetic analyses in yeast have shown that checkpoint pathways regulate cell cycle transitions in response to perturbations and solve the completion and alternation problem [17, 74].
3.1
The S phase and DNA damage checkpoints. The genetic screens that uncovered the signal transduction pathways
The Rad screen. By the late 1960s, many groups had carried out screens to identify yeast mutants that were sensitive to ionizing radiation, UV radiation, or both; these mutants were named rad mutants. Brian Cox, John Game, and Robert Mortimer among others, identified a collection of radiation sensitive mutants [22]. In 1970 scientists came to an agreement that mutations that conferred sensitivity to ionizing radiation would continue to be named rad mutants, numbered from RAD50 upward and they would be loci separate from those identified in screens for UV-sensitive mutants.
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Game and Mortimer took a collection of mutants sensitive to ionizing radiation and decided to assign x-ray sensitive mutants to genetic loci and to name them according to the new nomenclature. In doing so, they assigned the RAD50 to RAD57 loci. There are at least three explanations as to why mutations would lead to a radiation-sensitive phenotype. The mutated gene could encode a protein that is required in order for the cell to: (1) repair DNA damage; (2) stop the cell cycle before a critical transition (i.e., mitosis) in order to allow DNA repair; or (3) regulate the increased expression of proteins involved in DNA repair or cell cycle arrest. Mutation in the second class of genes usually confers sensitivity to both types of radiation, although this cannot be used as the only criterion for assigning them to the cell cycle arrest category. The Mec screen. Although many groups embarked on studies of the rad mutants that affected repair of the UV and IR-induced lesions, Lee Hartwell and Ted Weinert analyzed the rad mutants for their ability to stop the cell cycle in G2 in response to DNA damage and called the surveillance mechanism responsible for monitoring the successful completion of cell cycle events checkpoints. Investigators had long noted that yeast cells delayed cell cycle progression in response to DNA damage (refs in [133], but the genes required for the G2/M arrest had not been uncovered. Ted Weinert, while working with Lee Hartwell, characterized the rad mutations that were required for cell cycle arrest following a DNA damage signal. He also carried out a screen that would identify mutations in additional genes involved in this checkpoint response and named them mec mutants (mitotic entry checkpoint). Using cdc mutants that accumulate damage lesions in G2 at the restrictive temperature, Weinert showed that a rad9 mutation (originally identified as a UV-sensitive mutant [22] and subsequently mutations in RAD17 and RAD24 allowed cells to go through mitosis in the presence of DNA damage. Weinert then used a genetic screen with the damage-inducing cdc mutants and identified the mec1, mec2, and mec3 mutants as defective for the G2/M checkpoint [132]. The Sad and Dun Screens. Meanwhile Stephen Elledge’s group carried out a screen to identify mutants that were defective for the S phase checkpoint-induced arrest (Sad = S phase arrest defective) that would lead to catastrophic mitosis of unreplicated chromosomes. That screen identified SAD1 and SAD3 as essential components for this response [2]. The Elledge lab also carried out a screen for mutants that failed to turn on the transcriptional response as measured by the upregulation of ribonucleotide reductase 3 mRNA (RNR3). These mutants were called dun (DNA damage uninducible) and led to the identification of DUN1 and DUN2 as genes required for the transcriptional response following replication blocks and DNA damage [89, 142].
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At around the same time, Kato and Ogawa characterized a mutant named esr1, which showed sensitivity to the alkylating agent methyl–methane– sulfonate (MMS) and to UV radiation [57]. In addition, the esr1 mutants displayed meiotic defects. David Stern’s group took a biochemical approach and isolated Spk1 in a screen that used a bacterial system to identify dual specificity kinases (kinases that can phosphorylate proteins on serines/ threonine and tyrosine residues) from yeast [141]. When all of the genes mutated in these strains were cloned it turned out that many alleles of the same genes had been identified in the different screens and this pointed not only to their important role in various responses but also provided hints as to their biochemical functions. From the genes that were cataloged by Game and Mortimer as x-ray sensitive [30], RAD51, RAD52, and RAD54–RAD57 encoded proteins involved in recombinationmediated DNA repair. However, RAD53 encoded an essential kinase that is required for the cell to arrest not only in response to DNA damage but also when DNA replication is slowed down or blocked [2, 133, 141]. RAD53 alleles were identified not only in the rad screens but also in the sad (sad1), and mec (mec2) screens and Rad53 was the kinase identified in David Stern’s screen for dual specificity kinases (Spk1) (Table II). Another essential kinase that regulates this response is Mec1, and it was identified in the mec screen, sad screen (sad3) and in Ogawa’s screen as the gene mutated in esr1. Dun1 turned out to be another kinase [143] that shared phosphor–peptide recognition domains (FHA forked-head associated domains [47]) with Rad53 outside of the kinase domain. Dun2 is the catalytic subunit of DNA polymerase epsilon [89]. The fact that a component of DNA polymerase showed defects in a checkpoint response sparked enthusiasm and speculation that the DNA polymerase complexes, by the nature of their function, made attractive candidates for sensors and scanning machines [88]. Three kinases had been identified that are required in order to signal DNA damage and replication blocks. Many groups began to organize the rad, mec, and sad mutants (and other mutants that displayed sensitivity to agents that damage DNA or cause replication blocks) into the DNA repair category or as components of signal transduction pathways that signal the presence of these lesions. There were several criteria and several readouts that were used by many laboratories in order to piece together the checkpoint signal transduction pathways. The readouts included the effect of mutations on: (1) the cell’s ability to delay cell cycle progression when encountering damage at different stages of the cell cycle; (2) the activation of the kinases by phosphorylation; and (3) the upregulation of RNR3 mRNA and other damage-inducible gene transcripts. Using these readouts, proteins required for the checkpoint
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response with similarity to proteins involved in DNA replication (DNA polymerase or polymerase-associated proteins) or lesion processing (proteins with homology to nucleases or other DNA associated complexes) were considered to act as sensors that would trigger the response. Proteins that acted by regulating phosphorylation status of targets, such as protein kinases, were assigned the role of transducers due to their biochemical function. The proteins phosphorylated by the kinases (and presumably also regulated by phosphatases) were considered as effectors. These turned out to be crude assignments since most signal transduction pathways involve feedback loops that cause the kinases to phosphorylate upstream components. Nevertheless, at the time, it was a good plan of attack.
3.2
The S phase and DNA damage checkpoints. The biochemical properties of the components of the signal transduction pathways
Although the response to different forms of DNA damage employ common transducers, like Mec1 and Rad53, the sensors and downstream effectors differ depending on the signal type and the cell cycle stage during which the lesion is encountered (see below and [88]). An important aspect of checkpoint function concerns the mechanisms by which the transducers integrate different signals and regulate different cell cycle transitions. Here we will discuss the pathways that regulate progression through S phase and mitosis in order to prevent the segregation of incompletely replicated or damaged chromosomes. In 1995, Yosef Shilo’s group cloned the gene mutated in patients with a cancer-prone syndrome, ataxia telangectasia. These patients also display sensitivity to radiation and their cells have checkpoint defects [94, 108]. The product of the ATM gene is a large protein with homology to phosphatidylinositol 3-kinases (PI(3)-kinase-related protein kinases) and belongs to a family of proteins that include the checkpoint kinases Mec1 and Tel1 [84] from S. cerevisiae and Rad3 from S. pombe [7]. This discovery sparked renewed interest in the yeast checkpoint field since once again it was shown, as was the case for CDC28 and cdc2+, that the checkpoint pathways are also conserved between yeast and humans (Table II). The sensors: The first sensors identified for the S-phase checkpoint are proteins associated with the replication machinery [58, 88]. Proteins other than those associated with the replication machinery have also been ascribed a role as sensors for the S-phase checkpoint. Three of these proteins are part of a trimeric complex: Mre11, Rad50, and Xrs2 (Nibrin in mammals), and have been implicated in lesion processing for DNA repair [10, 15].
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Mutations in the genes encoding these proteins leads to checkpoint signaling defects in yeast [24, 36] and in humans mutations in NIBRIN and MRE11 lead to a cancer-prone syndrome similar to that of ataxia telangiectasia [13, 115]. The proteins in this complex are also targets of the Mec1 kinase and its orthologues. It remains to be determined whether other proteins that are involved in lesion processing and repair are also involved in checkpoint signaling, and if this role is different from their role in generating the singlestranded DNA that could act as the signal [124] to recruit other sensor complexes (see below). The picture of how the DNA damage sensor complexes activate the checkpoint kinases following DNA damage has begun to emerge. So goes the model: due to their association with DNA and predicted protein structure, there are at least three complexes that have been assigned the role of DNA damage sensors in yeast and mammals (Figure 4). Two complexes include the Rad24-RFC “clamp loading” complex (Table 2) [37], which is associated with replication Factor C-like proteins, and the Ddc1-Mec3Rad17 (“clamp like” complex [14, 63, 64, 113, 128], which has been proposed to act like a PCNA clamp. The clamp complex is referred to as the 9-1-1 complex due to the nomenclature of the S. pombe (and mammalian) orthologous proteins (S. pombe and human Rad9, Hus1 and Rad1, see Table II), where the similarity to the PCNA clamp was originally observed [126]. A third complex that contains Mec1 and Ddc2/Lcd1/Pie1 (Atr/Atrip in mammals, see Table II) requires both the Rad24 and Mec3 complexes for activation of the DNA damage response [61, 81, 101, 129]. Following recognition or processing of the lesion, the Rad24 complex recruits the Ddc1-Mec3-Rad17 clamp complex to DNA in an analogous mechanism to replication factor C-mediated loading of PCNA onto DNA at the primer-template junction during DNA replication, where it serves to act as a platform for the processive DNA polymerase complexes [122]. The Ddc1 protein is phosphorylated by Mec1 [93], and the human orthologue of Rad24 becomes phosphorylated by the Mec1 orthologue in a Mec3-dependent manner [144]. The Jackson laboratory used yeast genetics to define the role of Ddc2/Lcd1/Pie1 (Atrip homologue) in localizing Mec1 to chromatin [101]. Although Mec1 and the human homologue Atr localize to chromatin independently of the Rad24 complex, the phosphorylation of the yeast Mec1 and mammalian Atr substrates, and thus checkpoint signaling, is regulated by the sensor complexes containing Rad24 and Mec3 [21, 81, 144]. The fact that the kinases phosphorylated the sensor complexes generated confusion as to which component was upstream; however, work published in 2001 and 2002 (reviewed in [80]) gave rise to the model that there are at least three chromatin associated complexes: Mec1/Ddc2/Lcd1, Rad24-RFC and the Mec3 complex. The Rad24 and Mec3 complexes are thought to provide
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substrate specificity to the central checkpoint kinases of the Atm family, which include Mec1 and mammalian Atm and Atr [144] (Figure 4), although phosphorylation of a subset of Mec1 substrates (like Ddc2) does not require Rad24 and Mec3. All of these stepwise interactions argue that the activation of the DNA damage response could be lesion, cell cycle, and protein complex specific. Future work will determine all the factors and interactions that provide such specificity. The checkpoint responses triggered during DNA replication have been classified as the intra-S phase and S/M checkpoints. The former, monitors replication fork integrity, serves to stabilize replication forks, and prevents the firing of late replication origins. The S/M checkpoint blocks mitosis until DNA replication has been completed. What is the nature of the signal for activating the intra-S phase and the S/M checkpoints? Recent analyses of the structures arising from stalled replication forks point to single-stranded DNA that accumulates at a stalled fork as the possible signal. Single-stranded DNA has previously been proposed as the signal that activates the DNA damage response [31, 75]. The trimeric Rpa complex and Rad51 protein, which can bind single-stranded DNA, could be the molecules that trigger a response when they interact with the sensor complexes in the context of a stalled or collapsed replication fork, which have been estimated to occur every cell cycle [101]. rpa mutations in yeast confer S phase and G2/M checkpoint defects [58], and Ddc1 (the S. cerevisiae homologue of hRad9) has been found to co-localize with Rad51, during meiosis in yeast [48]. Work carried out with both yeast and the Xenopus systems put forth the idea that the primase enzyme, which is part of DNA Polα complex and synthesizes the RNA primer during DNA replication, was required for the S phase checkpoint signal to be generated [77, 82]. This suggested that the synthesis of the RNA primer itself could act as the signal that elicits the checkpoint response. This is in agreement with the fact that the Rfc and PCNA-like molecules, which require the RNA primer to load on to DNA are also required for checkpoint signaling [138]. Newport’s and Dunphy’s groups have used a Xenopus in vitro system to study the association of the Atm-related protein Atr and other checkpoint proteins with chromatin. An interesting find from these and other laboratories was that DNA replication is required for Atr and Hus1, a component of the clamp, to associate with chromatin [138]. Depletion experiments showed that Rpa was necessary for Atr and Hus1 to associate with chromatin and drug experiments suggested that DNA Polα was required for Hus1 association with chromatin [138]. However, these studies do not rule out the possibility that the effect of Rpa depletion is due to the inhibition of DNA replication, which is required for these proteins to associate with chromatin. These studies also do not
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differentiate between the scenarios where the RNA primer is the signal or where the synthesis of the primer is a step required for the sensor complexes to assemble on DNA [138] (Figure 5). Nevertheless, with some of the known players in hand, the answer to these questions may be around the corner. The transducers. In S. cerevisiae, replication blocks and DNA damage which occur in S-phase activate a checkpoint response regulated by the kinase Mec1 (Atr orthologue), which acts upstream of the kinase Rad53 to stabilize replication forks, prevent firing of late replicating origins and prevent anaphase entry [2, 106, 120, 133]. A role for Rad53 in the regulation of replication origin stability and firing would suggest that Rad53 complexes may also be localized in close proximity to proteins involved in DNA replication. Mec1 has also been shown to mediate the DNA-damage checkpoint that regulates mitotic progression by signaling to both Chk1 and Rad53. While in the Elledge laboratory we showed that Chk1 and Rad53 via Dun1 form parallel branches in the DNA-damage checkpoint that function downstream of Mec1 to regulate anaphase entry and mitotic exit, respectively (Figure 4, [103]). This is accomplished by blocking the destruction of two substrates of the anaphase-promoting complex or cyclosome. The effectors. The effectors of the S-phase checkpoint that mediate a block to anaphase in the budding yeast remain elusive, although in the fission yeast and vertebrates Chk1 and Cds1/Chk2 (Rad53 orthologue) target the Cdc25 phosphatase to elicit a G2/M arrest. However the effectors of the DNA damage checkpoint in the budding yeast have been identified. The DNA damage-induced checkpoint arrest that prevents progression through mitosis in S. cerevisiae occurs after the formation of a spindle, a stage that requires high activity of the mitotic cyclin/Cdk kinase (Clb2/Cdc28). At the checkpoint-induced arrest, which occurs at the metaphase to anaphase transition (M–A), the sister chromosomes are paired, held together by cohesins (such as Scc1, Mcd1) and can presumably interact with the bipolar pre-anaphase (short) spindle that has assembled at this stage. Progression through mitosis in all eukaryotic cells is regulated by the activity of the APC or cyclosome, which triggers chromosome segregation and mitotic exit through the ubiquitin-mediated degradation of anaphase inhibitors, such as Pds1 and mitotic cyclins, respectively. Orna Cohen-Fix, when in Koshland’s laboratory, showed that Pds1 was phosphorylated in response to DNA damage in a Mec1-dependent manner [18]. Phosphorylation of Pds1 correlates with accumulation of Pds1 following DNA damage [104]. While in the Elledge laboratory, we also showed that Chk1 inhibits the M–A transition by phosphorylation and stabilization of Pds1 and Rad53 blocks Clb2 degradation and possibly exit from mitosis by blocking the activation of the MEN. The Elledge laboratory continued to look for the targets of the
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Rad53 pathway and found that Bfa1, a component of the MEN pathway is regulated by Rad53 and Dun1. Phosphorylation of Bfa1 in a Rad53dependent manner inhibits the activity of the MEN pathway to help maintain the levels of Clb2 and thus allow cells to maintain their arrest in mitosis by maintaining active Cdk/B-type complexes [50]. However, the first wave of Clb2 destruction occurs at or right after anaphase, and the Cross laboratory showed that the first wave of Clb2 destruction could be sufficient for mitotic exit [131] indicating that Rad53 must have other targets in order to maintain high levels of Clb2/Cdk activity. Other roles of the checkpoint pathways involve the transcriptional induction of genes encoding products involved in DNA metabolism and DNA repair. The ribonucleotide reductase (RNR) genes are transcriptional targets of the S phase and DNA damage checkpoints that have been studied extensively in yeast and are conserved in mammals. The Rnr proteins catalyze the rate-limiting step of DNA synthesis, which is the generation of the deoxyribonucleotide pools. The Sphase and DNA damage checkpoint mediated by Mec1, Rad53, and Dun1 controls the transcriptional upregulation of the RNR genes in yeast [51, 139, 142]. Two of the effectors for the checkpoint-induced upregulation of Rnr activity are the transcriptional repressor Crt1 and the Rnr regulatory subunit Sml1 [139, 143]. In response to DNA damage or stalled replication forks, Crt1 is hyperphosphorylated in a Dun1-dependent manner and released from the DNA allowing transcription of the RNR genes [51]. In addition, phosphorylation of Sml1 in a Dun1-dependent fashion allows the activation of ribonucleotide reductase, presumably by mediating the degradation of Sml1 [141]. The Chk1 and Rad53 (Cds1/Chk2) proteins from both S. pombe and S. cerevisiae function downstream of the Atm homologues Rad3 and Mec1 and regulate mitotic progression following DNA damage, albeit through different mechanisms. Although S. cerevisiae displays a unique cell cycle organization, orthologues of both Pds1 and Esp1 have been identified in S. pombe (cut2 and cut1, respectively) and mammals (separin and securin, see Table II).
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Figure 4. The DNA damage checkpoint regulates mitotic progression by blocking the ubiquitin-mediated destruction of Pds1 and the inactivation of the mitotic Cdk/Cyclin complexes. Checkpoint activation is achieved by the interaction of two complexes that are independently recruited to sites of DNA damage Mec1-Ddc2/Lcd1/Pie1 and Mec3-Ddc1Rad17 (9-1-1 complex in mammals). The Rad24-RFC “clamp loading” complex recruits the Ddc1-Mec3-Rad17 “clamp like” complex following recognition or processing of the lesion. Mec1 and Ddc2/Lcd1/Pie1 requires both the Rad24 and Mec3 complexes for activation of the DNA damage response. The Ddc1 and Ddc2 proteins are phosphorylated by Mec1. The Rad24 and Mec3 complexes provide substrate specificity to the central checkpoint kinase Mec1 to direct phosphorylation of substrates such as Rad9, and the effector kinases Rad53 and Chk1, which then associate with Rad9. Chk1 and Rad53 form parallel branches in the DNA damage checkpoint to regulate anaphase entry and mitotic exit. Chk1 phosphorylates and blocks the ubiqutin-mediated destruction of the securin Pds1, and Rad53 activates Dun1 to block the inactivation of Bfa1 and activation of MEN. Dun1 also phosphorylates the Crt1
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transcriptional repressor and the RNR inhibitor Sml1 (not shown) in order to elicit a transcriptional response.
3.3
The spindle checkpoint, another checkpoint mechanism that monitors the integrity of the spindle
3.3.1
The genetic screens
Andrew Murray, who has studied the regulation of mitosis using both biochemistry (MPF) and yeast genetics, identified mitotic arrest deficient (mad) mutants with increased sensitivity to anti-microtubule drugs. These mutants were defective in the mitotic checkpoint arrest in response to a drug that interferes with mitotic spindle assembly by depolymerizing microtubules [70]. Murray identified mad1, mad2, and mad3 in this screen. Andrew Hoyt also identified mutants that were sensitive to microtubule destabilizing agents and failed to restrain subsequent cell cycles as evidenced by budding and DNA replication when in the presence of an antimicrotubule drug; these mutants were named bub (budding uninhibited by benzimidazole) [49]. Bub1 was later shown by Hoyt to be a kinase, again pointing to a signal transduction pathway in checkpoint control that is activated in response to microtubule perturbation. Mark Winey identified mutants defective in the duplication of the spindle poles. Duplication of spindle pole bodies is a requisite step in order to form a bipolar spindle that will separate the chromosomes at mitosis. He called his mutants mps for (monopolar spindle) [134]. One of the proteins identified by Whiney, Mps1, is a kinase [67], and the function of Mps1 is required for both spindle pole body duplication and the spindle checkpoint. The spindle checkpoint pathways, like the DNA damage checkpoint, bifurcate to control anaphase and mitotic exit [1, 8, 69, 119]. Epistasis analyses determined that the Mad2 and Bub2 acted in two separate pathways: Mad/Bub pathway functions to block M–A and the Bub2 pathway functions to block mitotic exit. The kinase Mps1 is thought to act upstream of the identified spindle checkpoint proteins as its overexpression activates both the Mad/Bub pathway and the Bub2 pathway, and causes cells to arrest both in metaphase and before mitotic exit [41].
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Figure 5. What is the signal that activates the S/M checkpoint during DNA replication and in response to replication problems? Some of the same proteins involved in the DNA damage response, such as the clamp loader proteins and the clamp-like complex, are involved in activation of the S/M phase checkpoint during DNA replication. During DNA replication the primase and DNA polymerase α complex synthesizes an RNA primer followed by short stretch of DNA synthesis. This acts to recruit the PCNA clamp via the RFC complex to the primer/template junction. PCNA and other proteins have a role in recruiting the processive DNA polymerase enzymes δ and ε. Genetic and biochemical experiments implicate proteins that are located at the replication fork (Rpa, DNA polymerase α/Primase complex) in the recruitment of both the clamp loader, clamp and Atr (Mec1) kinases in order to signal ongoing replication and stalled forks and to activate a checkpoint response that will block mitotic progression until DNA replication has been completed.
3.3.2
The biochemical pathways that are emerging from the studies of the genes identified in the genetic screens
As was the case for the DNA damage and S-phase-checkpoint proteins, the biochemical properties of the proteins that regulate the spindle checkpoint comprise a signal transduction pathway. Because the spindle and DNA damage checkpoints both regulate mitotic progression, they share some of the same targets, however, the transducers and sensors are different due to the nature and location of the signal. In the spindle checkpoint the
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signal seems to be generated at least in part by an unattached or monooriented kinetochore (the structure on the chromosome that allows attachment to the spindle, see Figure 6) (reviewed in [11]). Both Mad2, which recognizes unattached kinetochores, and the Aurora Kinase Ipl1, which is activated by mono-oriented kinetochores, activate the downstream signaling pathway to arrest cells in mitosis. The precise composition of the signaling complexes that elicit a metaphase arrest has yet to be elucidated, however, many of the proteins involved have been identified via genetic screens. Mad1, 2, and 3, as well as Bub1 and Bub3, comprise the Mad/Bub pathway, and these proteins are involved in the metaphase arrest in response to a spindle-activated checkpoint. Bub1, a kinase in the Mad/Bub pathway, acts interdependently with Mps1 to inhibit anaphase. Mad1 is hyperphosphorylated by Mps1 and this phosphorylation is dependent on Bub1 as well as the proteins Mad2 and Bub3 (reviewed in [3]). Mad2 binds to the APC specificity factor Cdc20 and this interaction is thought to inhibit the APC/C from targeting Pds1 for degradation, thereby inhibiting anaphase [53, 59]. cdc20 mutants that cannot bind Mad2 are deficient in the spindle checkpoint [40, 53]. Other components of the Mad/Bub pathway, including Mad3 and Bub3, are also found in a complex with Cdc20. Whether the interaction of Mad3 and Bub3 with Cdc20 results in inhibition of the APC/C has yet to be determined. Therefore, Cdc20 acts as the effector of the Mad/Bub pathway in response to spindle checkpoint activation. Mitotic exit is regulated by a second pathway, the Bub2 pathway, which also acts downstream of Mps1. Bfa1 is the effector of the DNA damage checkpoint and it interacts with Bub2 to form the GTPase activating protein involved in the MEN pathway. Recently, the Bfa1-interacting protein Ibd2 was identified and was shown to act upstream of Bub2/Bfa1 in the spindle checkpoint [52]. Activation of Bub2 by unattached or mono-oriented kinetochores inhibits the MEN pathway and the cells arrest before exiting mitosis (Figure 6). Thus, the Bub2/Bfa1 complex integrates different signals in order to block activation of the MEN. It remains to be determined whether other pathways that coordinate cell cycle transitions also regulate this complex.
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Figure 6. The spindle checkpoint blocks mitotic progression when the chromosomes are not attached or bi-oriented to the spindle. Unattached kinetochores as well as mono-oriented kinetochores (not bi-oriented) activate the spindle checkpoint. The spindle checkpoint is a signal transduction pathway that bifurcates to control anaphase and mitotic exit. Mps1 is thought to act upstream and it signals to both the Mad/Bub pathway and the Bub2 pathway, to bock anaphase and mitotic exit. Mps1 phosphorylates Mad1, and this phosphorylation is dependent on Bub1, Mad2, and Mad3. Phosphorylated Mad1 interacts with Mad2 and Bub3, Mad3 and Mad2, can bind to Cdc20. Interaction of these proteins with Cdc20 prevents the activation of the APC/Cdc20 blocking the ubiquitination of Pds1 and the Clbs. Mps1 activation of Bub2 blocks activation of the MEN. Ipl1, an Aurora kinase that localizes to kinetochres, activates the spindle checkpoint when the spindles bound to the kinetochores are not connected to opposite spindle pole bodies. The signal for this event is the lack of tension on the spindles bound to the sister chromatids. There are two possible mechanisms for Ipl1 mediated activation of the spindle checkpoint, (a) Ipl1 causes the dissociation of the spindle from a kinetochore which is detected by Mad2 and activates the Mps1-dependent checkpoint,
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and (b) Ipl1 could activate the Mad2 checkpoint directly. The detailed mechanisms of this surveillance pathway remain to be determined.
3.4
Roles of the S phase, DNA damage checkpoints and spindle (mitotic) checkpoints
From the studies carried out in S. cerevisiae, the following roles have been elucidated for the checkpoint that monitors DNA replication status and DNA integrity during S phase: The sensors are part of or associated with the replication machinery, or their structure mimics the complexes involved in DNA replication. The kinases Mec1 and Rad53 regulate effectors that (1) stabilize replication forks and allow reinitiation of DNA replication once the problem has been overcome [26, 73, 120]; (2) prevent firing of late replication origins [106]; (3) block anaphase until DNA replication is complete [2, 133]; and (4) through regulation of Dun1, activate transcription of genes encoding proteins involved in DNA metabolism and DNA repair [51, 139, 142, 143]. The DNA damage checkpoint pathway that operates at G1/S functions to block entry into S phase until the damage is repaired. The targets of this response have been better characterized in metazoans than in yeast. Due to differences in the organization of the cell cycle, the effectors for the DNA damage-induced arrest at G2/M differ between the budding yeast and mammals. In the budding yeast, the arrest occurs at the metaphase to anaphase transition (Figure 4); whereas in metazoans and fission yeast, the Chk1 and Cds1/Chk2 (sp and hRad53)-mediated arrest occurs in G2 by inactivation of the Cdc25 phosphatase and activation of the Wee1 kinases [29, 38, 96, 100, 105]. Budding yeast cells do activate a G2/M checkpoint in response to perturbations in bud construction and/or bud size [45, 79]; therefore, it remains to be determined whether mammalian cells also regulate the metaphase to anaphase transition in response to DNA damage. The mitotic or spindle checkpoint monitors the proper attachment and orientation of chromosomes on the spindle. This ensures the accurate transmission of genetic material to daughter cells. The signals that activate the S phase and spindle checkpoints originate at the replication fork or the kinetochore, and serve to indicate lack of completion of that particular cell cycle stage, much like the checkpoint mechanisms identified by Rao Johnson. Similarly the checkpoints that monitor replication forks and DNA integrity use similar mechanisms, and sometimes some of the same proteins, to delay cell cycle transitions, and in addition play an important role in the stability of replication forks and in DNA repair.
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One aspect of checkpoint signaling that eludes investigators is the mechanism by which these proteins provide specificity to activate the kinases in a cell cycle and lesion-specific context. For example, in S. cerevisiae Rad9 is involved in the amplification of the signal to both Rad53 and Chk1, however Chk1 is not activated in G1/S, whereas Rad53 is. In addition, Mec1 and the paralogue Tel1 can both signal to Rad53 and possibly to Chk1 but the pattern of Chk1 phosphorylation differs depending on the type of lesion that activates the response. Cells use a relatively limited number of molecules to integrate a variety of signals to coordinate cell cycle progression. As in all signal transduction studies, the burning question in checkpoint signaling is how specificity is achieved in these responses.
4
THE CANCER CONNECTION
Mutations in checkpoint genes, compromise the response to DNA damage at the cellular level and at the level of the organism lead to a predisposition to cancer. The low complexity of architecture of the yeast genome along with its amenability to genetic manipulation has allowed rapid alternation between the yeast and vertebrate model systems to elucidate the circuitry of the DNA damage checkpoints in eukaryotic cells. An important question in the study of checkpoints that has become the challenge for the next few years concerns the organization of the signal transduction pathways and the possible crosstalk or relationship between the checkpoint response and the proteins that carry out DNA repair. One of the hallmarks of tumor cells is that they have accumulated mutations that not only relieve the cells from growth regulatory signals but also allow them to invade the circulatory system, avoid the immune response, and grow at distant sites usually in a different tissue environment. The current hypothesis for the genetic changes in tumor cells is that they have gone through a period of genomic instability and loss of checkpoint control [71]. It is not surprising then that human orthologues of the checkpoint genes identified in yeast Atm (Mec1/Tel1), Chk2 (Rad53), Brca1 (Rad9) are tumor suppressor genes and that mutations in these genes predispose individuals to cancer [12]. Another characteristic of tumor cells is that they are aneuploid (too many chromosomes or missing chromosomal regions). This is not surprising since some cancer cells have mutations in spindle checkpoint genes that cause defects in the surveillance of the correct orientation and attachment of chromosomes to the mitotic spindle [12].
64 4.1
J. S. Searle and Y. Sanchez Therapeutic targets for cancer therapy
Most cancer cells are partially compromised in their ability to respond to DNA damage or replication blocks, which limits the effectiveness of current cancer treatments that involve the use of DNA-damaging agents such as chemotherapy and radiation. Therefore, manipulation of these pathways may allow us to design more effective therapeutic regimes for the treatment of cancer. For example, cells could be rendered more sensitive to genotoxic agents by inactivation of an additional branch or signaling point of the checkpoint pathway. Our studies with the yeast pathways regulated by Rad53/Dun1 and Chk1 have shown that indeed this is the case. Such treatments would render cancer cells more sensitive to chemotherapy or radiation treatment. Several investigators are using both S. cerevisiae and S. pombe to test this hypothesis by using the yeasts as genetic tools to inactivate interactions among different players in the response to DNA damage. By the year 2001, not only had the yeast genome been sequenced but there was a collection of deletion mutants in every nonessential yeast gene which has been used in screens to identify genes involved in several physiological processes, including DNA repair. Michael Resnick’s group, who had identified some of the original RAD genes in standard genetic screens, used this collection of deletion mutants to identify additional genes involved in the response to ionizing radiation [6]. This screen uncovered old favorites and new mutants that had varying degrees of sensitivity to ionizing radiation and to other agents that damage DNA or block DNA replication. The sensitive strains had mutations in genes that encode proteins involved in DNA repair, cell cycle arrest, chromatin remodeling, nuclear architecture, and endocytosis. In these studies the budding yeast system, proved once again, to be a powerful genetic tool for the identification of proteins that function in pathways that regulate cell cycle progression and genomic stability. And the lessons that we have learned from yeast include the finding that the efficacy of treatment depends on the integrity of checkpoint response in the cell. Due to the conservation of the pathways that guard the integrity of the genome, we have the tools today that allow investigators to use the budding yeast in genome-wide approaches to screen peptide and chemical agents as potential therapeutic agents. The utility of this system for the development of peptide inhibitors that target signaling and checkpoint pathways has already been demonstrated in a screen that identified peptides that specifically inhibited components of MAP kinase signaling and mitotic checkpoint pathways [90]. Checkpoint pathways, by virtue of their role in protecting the integrity of the genome, are essential to mammals for the avoidance of diseases such as
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cancer. When the checkpoint pathways go awry, they become targets for cancer drug discovery. Thus, studies that shed light on the mechanisms of the DNA damage response in mammalian cells will allow us to further understand both the therapeutic effects and the individual risk of exposure to genotoxic agents, chemotherapeutic agents, and to radiation.
ACKNOWLEDGMENTS In this chapter we have covered the work of many laboratories spanning nearly 40 years. Broad strokes were necessary to convey the big picture of how the players of the many layers of regulation of mitosis were identified. We apologize to our colleagues whose work we did not cite due to limited space. We are grateful to Ted Weinert, Craig Tomlinson, Todd Stukenberg, and members of the Sanchez laboratory for helpful comments and discussions and to Doug Kellogg for communicating data prior to publication. We thank Eunice Ablordeppey for doing the footwork necessary to gather many of the published papers included in the references.
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139. Zhao, X., E. G. Muller, and R. Rothstein. 1998. A suppressor of two essential checkpoint genes identifies a novel protein that negatively affects dNTP pools [In Process Citation]. Mol.Cell 2:329–340. 140. Zhao, X., and R. Rothstein. 2002. The Dun1 checkpoint kinase phosphorylates and regulates the ribonucleotide reductase inhibitor Sml1. Proc. Natl. Acad. Sci. USA 99:3746–3751. 141. Zheng, P., D. S. Fay, J. Burton, H. Xiao, J. L. Pinkham, and D. F. Stern. 1993. SPK1 is an essential S-phase-specific gene of Saccharomyces cerevisiae that encodes a nuclear serine/threonine/tyrosine kinase. Mol. Cell. Biol. 13:5829–5842. 142. Zhou, Z., and S. J. Elledge. 1993. DUN1 encodes a protein kinase that controls the DNA damage response in yeast. Cell 75:1119–1127. 143. Zhou, Z., and S. J. Elledge. 1992. Isolation of crt mutants constitutive for transcription of the DNA damage inducible gene RNR3 in Saccharomyces cerevisiae. Genetics 131:851–866. 144. Zou, L., D. Cortez, and S. J. Elledge. 2002. Regulation of ATR substrate selection by Rad17-dependent loading of Rad9 complexes onto chromatin. Genes Dev. 16:198–208.
Chapter 3 YEAST AS A TOOL IN CANCER RESEARCH: NUCLEAR TRAFFICKING Anita H. Corbett and Adam C. Berger Department of Biochemistry and Graduate Program in Biochemistry, Cell and Developmental Biology, Emory University School of Medicine, 1510 CLifton Rd., NE, Atlanta, GA 30322
Recently, scientists have begun to appreciate the important role that dynamic intracellular trafficking plays in regulating the function of growth regulatory factors that have been implicated in cancer. Importantly, numerous examples of proteins whose function is regulated by entry into or exit from the nucleus have been identified [107]. The classic example of this regulation is the translocation of a transcription factor into the nucleus in response to a signal [63]. One such example is the transcription factor p53, which is transported into the nucleus, where it can act in various nucleic acid transactions in response to DNA damage [27, 89]. Since p53 function is lost in a large number of human tumors [110], understanding what mechanisms regulate p53 function is obviously of critical importance to understanding cell transformation. It has become clear that one of the mechanisms that contributes to regulation of p53, as well as other proteins that have been implicated in cancer, is the orchestrated movement of the protein between the cytoplasm and the nucleus [27, 89]. Our understanding of the various mechanisms that regulate such nuclear protein import and export has been spurred by recent advances in the field of nucleocytoplasmic trafficking. An eukaryotic cell is defined by the advantageous presence of a membrane-bound nucleus, which separates the nuclear genetic material from the cytoplasmic protein translation machinery to allow exquisite regulation of gene expression. This separation dictates the need for a transport system that can connect the cytoplasm to the nucleus. The basic machinery that mediates this transport has been conserved from yeast to humans [82]. In fact, studies of nuclear transport in yeast have identified critical regulators of nuclear transport and provided important insight into the mechanism by which the process is orchestrated. 75 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 75–100. © 2007 Springer.
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The goal of this chapter is to provide a brief overview of the recent advances in the study of nuclear transport and describe how studies in yeast model systems have contributed to these advances. The chapter primarily focuses on the budding yeast, Saccharomyces cerevisiae, but also incorporates findings from the fission yeast, Schizosaccharomyces pombe. The chapter first presents a general overview of the mechanism and mediators of the nuclear transport process. It then describes three specific examples of critical growth regulatory yeast proteins that are modulated by nuclear transport and are models for cancer-implicated processes. The first example is the cell cycle-regulated localization of cyclin B, primarily focusing on studies with S. cerevisiae cyclin B, Clb2p. The second example is the S. pombe phosphatase, cdc25. Finally, the third example is the S. cerevisiae Pho4p transcription factor, which serves as a paradigm for signal-mediated transcription factor import and export. These examples highlight the regulation that can be imparted by dynamic localization between the cytoplasm and the nucleus.
1
THE NUCLEAR PORE COMPLEX
All macromolecular traffic between the cytoplasm and the nucleus flows through a large channel called the nuclear pore complex (NPC). The NPC is a large macromolecular machine that spans the double membrane of the nuclear envelope (see [28, 98, 100] for recent reviews). Studies over the years have revealed that nuclear pores have eightfold rotational symmetry within the membrane but have distinct nuclear and cytoplasmic faces. The cytoplasmic face has filaments that emanate from the pore and the nuclear face has a basket-like structure. These distinct faces are thought to contribute to the mechanism of targeting and transport through the pores. Nuclear pores are composed of proteins termed nucleoporins or Nups. A large number of these nucleoporins contain characteristic phenylalanine–glycine (FG) repeat motifs that are likely to play a critical role in the translocation of substrates through the pore. Several recent studies have identified all the proteins that make up both the yeast and the vertebrate NPC. This work was led by a pioneering study in S. cerevisiae where Rout and colleagues used a proteomic approach to identify the proteins that comprise the yeast NPC [86]. This work paved the way for a similar study in higher eukaryotes [19], which revealed that although the vertebrate pore complexes are somewhat larger than those in yeast, the number of distinct proteins is approximately the same. Spatial analysis of the yeast nuclear pore revealed that many of the nuclear pore proteins exist in multiple copies that are symmetrically distributed between the nuclear and cytoplasmic faces of the pore [86].
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Many of the FG-repeat-containing nucleoporins are predicted to line the central channel of the NPC and make direct contact with transport receptors (section 2.2) moving through the pore. In contrast, there are a few nucleoporins that are asymmetrically localized to either the nuclear or cytoplasmic face of the pore. These asymmetric proteins probably comprise the distinct structures seen on either face of the pore, the nuclear basket and the cytoplasmic filaments. It seems likely that these structures may serve as docking or assembly/disassembly sites for the cargoes that are transported through the pore. A more detailed understanding of the overall structure of the pore will be required before the precise mechanism of movement through can be fully defined. There are several examples of gene translocation events involving genes encoding nucleoporins that have been implicated in human cancers. The most common translocations of this type, which occur in acute myelogenous leukemia, involve the Nup98 nuclear pore protein [68]. There have been very few studies to understand the functional importance of these translocations and the resulting oncogenic fusion proteins. Another example of a translocation that has been linked to leukemia is the translocation that places the Met kinase under the control of the Tpr (Translocated promoter region) nuclear pore protein [23].
2
PROTEIN TRAFFICKING
2.1
Nuclear transport targeting signals
As with most intracellular targeting mechanisms, trafficking between the nucleus and the cytoplasm depends on amino acid sequences within the protein cargoes to be transported. However, unlike other targeting mechanisms, such as mitochondrial and ER targeting, nucleocytoplasmic trafficking signals are not cleaved and remain an integral part of the protein. This mechanism allows the cargo proteins to undergo multiple rounds of import and/or export and allows the cell to exploit this cycling as a regulatory mechanism. Although there are still numerous nucleocytoplasmic-targeting signals that are ill-defined or completely unknown, canonical signals for both nuclear protein import and nuclear protein export have been identified and studied in some detail. Nuclear import signals are generally termed nuclear localization signals (NLS), while signals that target proteins for export from the nucleus are termed nuclear export signals (NES). The classical NLS consists of either a single cluster of basic amino acid residues (monopartite) or two clusters of basic amino acid residues separated by a 10–13 amino acid linker sequence (bipartite) [56, 85]. The monopartite
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NLS sequence is typified by the NLS found in the SV40 large T antigen (PKKKRKV), whereas the bipartite NLS is typified by the nucleoplasmin NLS (KRPAATKKAGQAKKKK). These sequences mediate binding to a classical nuclear import receptor, importin/karyopherin α (discussed in more detail in section 2.2 below). Although numerous NLS sequences have been identified, the best consensus sequence that has emerged thus far is: K(K/R)x(K/R) where x is any amino acid [44]. The classical NES is comprised of a series of hydrophobic amino acids, generally leucine, isoleucine, or valine [32]. The founding member of the NES family is the protein kinase A inhibitor protein, which contains the NES sequence LALKLAGLDI, where the underlined residues are critical for NES function [108]. Numerous variations on this theme have been identified and thus far the best consensus sequence for an NES is LxxxLxxLxL, where the spacing between the leucines can vary and, in fact, the leucines can be substituted with virtually any hydrophobic amino acid. Obviously, it is possible to scan the sequence of any given protein and identify putative classical NLS or NES sequences, however, substantial experimental evidence is required to prove that an amino acid sequence is both necessary and sufficient to serve as a functional targeting signal [21]. This is particularly critical given the rather weak consensus sequences for the known nuclear targeting signals. Current efforts are being made to better define the important characteristics of both NLS and NES sequences. Recent studies have examined the interaction between NLS sequences and the classical NLS receptor, importin α, by determining the binding energy contributed by each residue within NLS sequences [44]. This study has been extended by using budding yeast to determine how the binding affinity of the NLS for the NLS receptor relates to the actual accumulation of the cargo within the nucleus [43]. A complementary study used an in vivo randomization-selection assay to determine which residues within an NES are absolutely required for export function [11]. Mutations or modifications that alter the nuclear-targeting signals of growth regulatory proteins are associated with some instances of cellular transformation. There is evidence that phosphorylation that occurs proximal to an NLS sequence can alter the nuclear targeting of the cargo protein [40, 51]. One notable example where an alteration in nuclear targeting is associated with cellular transformation is the v-jun oncogene. Normal cellular jun (c-jun) contains a classical NLS with a flanking cysteine [13]. In the v-jun oncogene the sequence is altered such that the cysteine is replaced by a serine [13, 102]. It seems that phosphorylation of this serine regulates the nuclear import of v-jun and thus imparts an inappropriate function on the oncoprotein [13, 102]. There are a number of other mechanisms that can be used to regulate the function of a nuclear targeting signal. For example, there are cases where an NLS is masked by a binding partner as in the case
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of IκB binding to the transcription factor NFκB. IκB binds to NFκB and masks the NLS and only when IκB is degraded in a signal-dependent manner does NFκB enter the nucleus, trigger gene expression, and cause changes in cell growth [71, 80]. As we learn more about the signals that target cargo proteins to and from the nucleus, it is likely that we will uncover other regulatory mechanisms that are critical for proper control of cell growth. In fact, recent work with the Bcr–Abl oncoprotein indicates that trapping this protein within the nucleus in its active state can induce transformed cells to undergo apoptosis [105]. Thus, there is potential therapeutic value in uncovering ways to alter the nuclear transport properties of certain proteins. Experiments carried out in the yeast model system are making important contributions to our understanding of nuclear targeting signals.
2.2
Nuclear transport receptors
Targeting signals within cargo proteins are recognized by soluble receptors that direct those cargoes to the nuclear pore for transport. These receptors are a family of related molecules that are generally referred to as either importins (for import receptors), exportins (for export receptors), or generally as transport receptors or karyopherins [14, 35, 111]. The budding yeast proteins are generally referred to as karyopherins, or Kaps, and are coupled with their molecular weight to generate the standard yeast name (e.g., Kap95 for the 95 kDa karyopherin). There are 14 of these functionally and structurally related Kap proteins in S. cerevisiae (see Table 1) and 23 that have been identified thus far in humans [99]. In many cases, the cargoes bind directly to these Kap proteins for import into the nucleus. However, an additional adaptor is required to mediate recognition and binding of the classical basic NLS [34]. This protein, often called importin α in higher eukaryotes and Srp1p or Kap60p in S. cerevisiae, binds directly to the NLS sequence within the cargo and then also interacts with a member of the karyopherin family, generally referred to as importin β in higher eukaryotes and Kap95p in S. cerevisiae [35, 82]. As described in more detail below, this heterotrimeric complex is then imported into the nucleus. Thus, the Kap60p/NLS cargo complex actually serves as a specialized cargo for import by the karyopherin, Kap95p. Although the S. cerevisiae genome encodes only a single Kap60 protein, there are at least six corresponding importin α genes that have been identified in humans [62].
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Table 1. Soluble nuclear transport factors Common protein names Other names (S. cerevisiae) Ran GTPase Cycle Ran (Gsp1p/Gsp2p)
RanGAP (Rna1p)
RanGEF (Prp20p)
Cnr1p/Cnr2p (S. cerevisiae) Cst17p (S. cerevisiae) Spi1p (S. pombe) Ran (metazoans) Rna1p (S. pombe) Segregation distorter (D. melanogaster) RanGAP1 (vertebrates) Srm1p (S. cerevisiae) Mtr1p (S. cerevisiae) Pim1p (S. pombe) BJ1 (D. melanogaster) RCC1 (vertebrates)
Function
Small GTPase that regulates cargo-receptor interactions
GTPase activating protein for Ran
Guanine nucleotide exchange factor for Ran
Nuclear transport receptors* Importin/karyopherin α (Kap60p) Importin/karyopherin 95 (Kap95p)
Srp1p (S. cerevisiae)
NLS receptor
Rsl1p (S. cerevisiae)
CRM1 (Crm1p/Xpo1p) CAS (Cse1p)
Kap124p (S. cerevisiae) Exportin 1 (vertebrates) Kap109p (S. cervisiae)
Complexes with Kap60p to import NLS cargo/import receptor for ribosomal proteins and other cargoes NES receptor
Transportin (Kap104p) Kap121p
Pse1p (S. cerevisiae)
Kap123p Kap108p
Yrb4p (S. cerevisiae) Sxm1p (S. cerevisiae)
Kap120p Kap122p Kap119p
Pdr6p (S. cerevisiae) Nmd5p (S. cerevisiae)
Kap111p Los1p Kap114p Kap142p
Mtr10p (S. cerevisiae) Exportin-t (vertebrates) HRC1004 (human) Msn5p (S. cerevisiae)
Kap60p recycling to the cytoplasm hnRNP import Ribosomal protein/Pho4p/Spo12p import Ribosomal protein import Ribosomal protein import/La protein import 60S ribosomal subunit export TFIIA import TFIIS transcription factor and Hog1p import Npl3p import tRNA export TATA-binding protein import RP-A import and Pho4p/Far1p/Mig1p/Ste5p export
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*There are a large number of importin/exportin/karyopherin receptors in yeast and vertebrates. In some cases (as indicated) vertebrate homologues with the same apparent function as the corresponding S. cerevisiae protein have been identified. However, in many cases no obvious single functional vertebrate homologue corresponding to the S. cerevisiae receptor has been assigned. For this reason the nuclear transport receptors listed focus on the S. cerevisiae proteins where the family of receptors is most extensively characterized.
With the exception of the NLS adaptor protein, Kap60p, Kaps share a common domain structure [34, 35]. Each has a conserved N-terminal domain that mediates binding to the small GTPase Ran, which regulates cargo binding to the Kap proteins as described in more detail in the following section (2.3) on the Ran GTPase cycle. The central domain of the Kaps is an FG binding domain that mediates interactions with the FG-repeat-containing nucleoporins located in the nuclear pore. Finally, the least well understood domain is the C-terminal domain, which mediates binding to the cargoes. Some of the receptor/cargo interactions are starting to be defined at the molecular level by structural studies of both yeast and vertebrate proteins. The most structural information has been gathered on the heterodimeric import receptor for classical NLS-containing cargoes. Structures of both mouse and yeast importin α/Kap60p have provided important information about how an NLS is recognized and how binding to the NLS cargo is regulated [17, 61]. These studies also revealed that despite relatively low sequence conservation between the yeast and vertebrate NLS receptor proteins (~30% identity), the overall structure of the proteins is identical. The structural basis for the interaction between vertebrate importin α and importin β has also been defined [16]. In addition, crystallization of a complex of vertebrate importin β with an FG-repeat peptide has revealed the structural basis for the interaction of the transport receptor with the FGcontaining nucleoporins [5]. Although many of these structural studies have examined vertebrate proteins, subsequent studies in yeast, which exploited this structural information to generate rationally designed site-directed mutants, strongly argue for functional conservation of these proteins between yeast and higher eukaryotes [4, 82]. This lends confidence to the assumption that studies of these nuclear transport proteins in the yeast model system can provide important insight into their function in higher eukaryotes. Alteration in several nuclear transport receptors have been implicated in human cancer. The export receptor required for the recycling of the NLS receptor, importin α, is a protein named CAS, which stands for cellular apoptosis susceptibility [7]. The human CAS gene is amplified in several transformed cell lines including breast, colon, and bladder cancer [12]. In addition, its expression pattern and intracellular localization is altered in
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70–90% of tumor cells in a population of invasive ductal and lobular breast carcinomas [6]. The functional yeast homologue of CAS is a protein called Cse1p, which was originally isolated in a screen for yeast mutants with defects in chromosome segregation [113]. It is possible that CAS/Cse1p serves as a critical regulator of growth control that when altered can cause uncontrolled cell growth or cancer. It seems likely that other transport receptors may also be implicated in tumorigenesis in the future. Consistent with this notion, a recent study identified a translocation of a novel human karyopherin, RanBP17, in acute lymphoblastic leukemia [39]. In addition, a truncated form of the NLS receptor, importin α, has been identified in the human breast cancer cell line, ZR-75-1 [59].
2.3
The Ran GTPase cycle
One of the major requirements for efficient transport of cargoes into and out of the nucleus is that the system must have inherent directionality. For example, in the case of nuclear protein import, the cargo protein must be bound by the receptor in the cytoplasm, translocated into the nucleus, and then dissociated from the receptor. There must be some mechanism to switch the receptor from cargo pick up to cargo delivery. For most nuclear transport processes, this switch is the Ran GTPase [22, 97, 107]. Work from a number of laboratories over the years has provided an insight into how Ran imparts directionality on nuclear transport (For review see [35]). The current model (Figure 1) relies on the compartmentalization of RanGDP and RanGTP, where RanGDP is primarily cytoplasmic and RanGTP is within the nucleus associated with chromatin [78]. The underlying mechanism for this compartmentalization is the differential localization of the proteins that regulate the GTPase activity of Ran. Like many other small GTPases [70], isolated Ran hydrolyzes GTP very slowly. The rate of hydrolysis is increased ~10,000-fold in the presence of the Ran GTPase Activating Protein (RanGAP) [8], which is called Rna1p in S. cerevisiae [9]. The Ran Guanine nucleotide Exchange Factor (RanGEF) enhances the rate of conversion of RanGDP to RanGTP [10]. The S. cerevisiae RanGEF is Prp20p [30]. RanGAP is localized to the cytoplasm [46], which increases the cytoplasmic pool of RanGDP. In contrast, RanGEF is enriched within the nucleus where it is bound to chromatin at least in part through interactions with histones [78, 79]. This enriches the nuclear pool of RanGTP. Many of the studies that helped to elucidate the Ran cycle were carried out in yeast. All proteins of the Ran cycle are highly conserved. For example, the amino acid sequence of human Ran is 85% identical to that of the S. cerevisiae Ran protein, Gsp1p. Human RanGEF, RanGAP, and NTF2 can all functionally replace their yeast counterparts [18, 30, 72]. This
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conservation of function means that researchers can take advantage of the numerous conditional alleles of these essential genes that have been generated in yeast over the years [21]. These experimental tools have been critical for the studies that have elucidated the mechanism of Ran action. The mechanism by which the different nucleotide-bound states of Ran regulate nuclear transport is illustrated in Figure 1 (Import and Export). Import cargoes bind their karyopherin receptors in the cytoplasm. These receptor/cargo complexes are imported into the nucleus where the complex encounters RanGTP. RanGTP binds with high affinity to the N-terminus of the karyopherin. This induces a conformational change that releases the cargo into the nucleus. In contrast, export karyopherins bind their cargoes as an obligate heterotrimeric complex that contains the receptor, the cargo, and RanGTP. This heterotrimeric complex is exported to the cytoplasm where it encounters RanGAP. Conversion of RanGTP to RanGDP in the presence of RanGAP dissociates the export complex and delivers the cargo to the cytoplasm. In both cases the receptors are recycled so that they can be used for another round of transport. In addition it is necessary to replenish the nuclear pools of RanGTP, which would be depleted by ongoing nuclear export. This is accomplished by the small RanGDP-binding protein, NTF2, which binds RanGDP in the cytoplasm and imports it into the nucleus where it can be converted to RanGTP [84, 95]. How NTF2 is then recycled to the cytoplasm has not yet been determined. Much recent work on the Ran GTPase has revealed a more general role for RanGTP in marking the position of the chromatin within the cell and hence the position of the nucleus [107]. This more general function underlies the well-characterized role of RanGTP in regulating import and export cargo binding to the transport receptors. In order for import complexes to dissociate in the proper cellular compartment, there must be a signal that they have reached the nucleus. This signal is the contact with RanGTP, which is concentrated at the chromatin due to the chromatin association of the exchange factor [78]. For nuclear protein export, the converse is true. Export complexes are formed only in the presence of RanGTP, in the vicinity of chromatin, and then disassembled in the cytoplasm where the RanGTP is hydrolyzed by the action of the cytoplasmic RanGAP. The realization that RanGTP serves as a general marker for the nuclear compartment and specifically the chromatin explains several recent observations about additional roles for Ran. Work from a number of laboratories suggests that RanGTP is required for mitotic spindle function during cell division [55]. While this finding could suggest that RanGTP binds directly to a component of the spindle apparatus, more recent experiments suggest that RanGTP is required to release proteins that are required for spindle function from the importin β family of transport receptors [38, 75, 109]. A similar role for RanGTP in mediating nuclear envelope assembly has also been identified both in vertebrates and in yeast
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[88, 106, 122]. In this case, the RanGTP likely releases nuclear pore proteins, which are required for the association and assembly of early NPCs with the chromatin. In all three cases, nuclear transport, mitotic spindle assembly, and nuclear envelope assembly, RanGTP serves to dissociate proteins from the transport receptor in the vicinity of chromatin. This concentrates proteins at the chromatin or within the nucleus where they function. Thus, although RanGTP was originally identified as a critical regulator of nuclear transport, this role is probably only one aspect of Ran function. Ran GTPase Cycle cytoplasm Ran GDP
nucleus
Ran GDP Ran GDP
Ran GDP
Ran GEF
Ran GAP
Ran GTP
Import
Cargo
Cargo Ran GDP
Ran GDP Ran GDP
Export
Ran GTP
Ran GTP
Ran GTP
β
β
Ran GTP
Cargo
Ran GTP Ran GTP Ran GTP
Cargo
β
β
Ran GDP
Ran GTP
Figure 1. The Ran GTPase cycle regulates the directionality of nuclear protein transport. The RanGAP protein is localized to the cytoplasm, which leads to a high level of cytoplasmic RanGDP. The RanGEF is localized to the nucleus, which leads to a high level of nuclear RanGTP. This asymmetric distribution of RanGTP and RanGDP regulates the assembly of receptor cargo complexes. For Import, the transport receptor (β) binds to the import cargo in the cytoplasm. This complex is translocated through the nuclear pore into the nucleus. In the nucleus RanGTP binds to the transport receptor, which causes a conformational change that releases the cargo. For Export, the export receptor (β) binds to the export cargo in the nucleus in an obligate trimeric complex with RanGTP. The trimeric complex is translocated through the nuclear pore into the cytoplasm. In the cytoplasm, the complex encounters the RanGAP, which hydrolyzes the RanGTP to RanGDP and dissociates the complex to release the cargo into the cytoplasm.
Given the critical role of Ran in nuclear transport as well as other important cellular processes, it may not be surprising that no mutations in the Ran cycle components have yet been linked directly to human cancer. It seems likely that the functions of these proteins are so critical to cellular viability that mutations are incompatible with life. However, as we learn
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more about the regulatory mechanisms, we may uncover changes in regulators of the Ran cycle that do contribute to human disease.
2.4
Classical nuclear protein import/export
cytoplasm Cargo
nucleus
Cargo
NLS
(4)
NLS
(1)
β
Cargo (2) NLS
α β
α β
(3)
β
Cargo
Cargo
NLS
α
NLS
Ran GTP
α
α α Cse1
Ran GAP
α Cse1 Ran GTP
α (5)
Cse1 Ran GTP
α
Cse1
Cse1 Ran GTP
Ran GDP
Figure 2. The nuclear import pathway for cargoes that contain a classical NLS is the best characterized nuclear transport pathway. This pathway can be dissected into at least five distinct steps: (1) recognition and binding of the NLS cargo to the Kap60p(α)/Kap95p(β) heterodimeric import receptor in the cytoplasm; (2) targeting to the nuclear pore through interactions between Kap95p and the nuclear pore; (3) translocation through the pore through transient interactions between Kap95p and the FG-repeat containing nucleoporins; (4) delivery into the nucleus when RanGTP binds to Kap95p to cause a conformational change that releases the Kap60p/NLS cargo; and (5) recycling of Kap60p to the cytoplasm in a heterotrimeric complex with the export receptor, Cse1p, and RanGTP.
The best understood nuclear transport process is the import of protein cargoes that contain a classical basic NLS (Figure 2). Thus, this process can be used most readily to illustrate the steps that occur when a transport cargo is moved into or out of the nucleus. Historically nuclear protein import was divided into two steps, docking at the nuclear pore, an energy independent step, and translocation into the nucleus, an energy-dependent step. Advances in our understanding of the process and the players now lead us to define at least five distinct steps for import of an S. cerevisiae cargo that contains a classical NLS: (1) recognition and binding of the NLS cargo to the Kap60p(α)/Kap95p(β) heterodimeric import receptor in the cytoplasm; (2) targeting to the nuclear pore through interactions between Kap95p and the nuclear pore; (3) translocation through the pore via transient interactions between Kap95p and the FG-repeat containing nucleoporins; (4) delivery into the nucleus where RanGTP binds to Kap95p to cause a conformational change that releases the Kap60p/NLS cargo; and (5) recycling of Kap60p to the cytoplasm in a heterotrimeric complex with Cse1p and RanGTP, and
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Kap95 presumably in complex with RanGTP. Note that the only energy expenditure in this process occurs when the karyopherin proteins are recycled to the cytoplasm and the accompanying RanGTP is hydrolyzed. Export of cargoes from the nucleus is very similar to the import process except that the export complex forms in the presence of RanGTP (in the nucleus) and is dissociated upon GTP hydrolysis (in the cytoplasm). Thus far all export mechanisms seem to involve the direct recognition of the cargo protein by the export Kap protein, but the molecular details of the recognition have not been elucidated for the NES export receptor/cargo complex. The export of cargoes that contain a leucine rich NES, which is recognized by the export Kap, Crm1p/Xpo1p (CRM1 in vertebrates), serves as a paradigm for transport receptor mediated nuclear export. The steps involved in NES export are: (1) recognition and binding of the NES cargo in a trimeric complex with Crm1p/Xpo1p; (2) targeting to the nuclear pore through interactions between Crm1p/Xpo1p and the FG nucleoporins; (3) translocation through the pore; (4) delivery of the cargo upon RanGTP hydrolysis in the cytoplasm; and (5) recycling of the export receptor and RanGDP to the nucleus. This cycle of export is virtually identical to the recycling of Kap60p(α) by Cse1p that is illustrated in Figure 2. This demonstrates the conservation of mechanism in the nuclear transport process.
3
RNA EXPORT
Multiple classes of RNAs, including mRNAs, tRNAs, and U snRNAs, are transcribed and processed within the nucleus and then transported to their sites of action in the cytoplasm. Competition experiments have shown that the export mechanisms of different classes of RNA are distinct from one another [67, 76]. A great deal of information has emerged regarding factors that are essential for RNA export [76], but like protein trafficking, the exact mechanism of translocation through the NPC remains elusive. As compared with proteins, RNAs require extensive processing before they reach their mature form and are ready to exit the nucleus, and so it has sometimes been difficult to separate activities required for RNA processing (such as splicing) from bona fide mediators of export.
3.1
Export of RNA via classical nuclear transport receptors
The export of tRNA, U snRNA, and rRNA follows pathways analogous to nuclear protein export. For example, tRNA is recognized directly by the
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importin-β family carrier exportin-t/Los1p [1, 42, 67, 90] and is exported as a complex with RanGTP, which is disassembled in the cytoplasm when RanGTP is hydrolyzed to RanGDP. Preferential export of mature tRNAs seems to be achieved at least in part by the specificity of exportin-t for the mature processed, modified, and appropriately aminoacylated tRNAs [94]. U snRNAs are synthesized in the nucleus, transported to the cytoplasm where they associate with protein components of mature snRNPs, and are then reimported to the nucleus where they function in mRNA splicing [50]. The monomethylated G cap of the initial export substrate is recognized by the cap-binding complex (CBC) [50] and then exported from the nucleus in a CRM1-dependent manner [31]. As there is no evidence that the CBC binds directly to CRM1, it seems likely that an adaptor protein mediates this interaction. rRNAs are exported in the context of large assembled RNP complexes. Although export depends on Ran [48, 74], it is controversial whether Ran plays a direct role in export or whether instead its activity may be essential for the import of components required for RNP assembly [87].
3.2
mRNA export
Export of poly (A)+ RNA remains the least well understood of the RNA export mechanisms. mRNAs are not exported to the cytoplasm as naked nucleic acids, but rather as ribonucleoprotein complexes and it is generally agreed that the export machinery recognizes signals within the proteins of these complexes rather than the RNA itself [49, 77]. For example, export of unspliced HIV transcripts from the nucleus is mediated by Rev through its export by the NES receptor [20]. While this mechanism is exploited by HIV, it seems that none of the karyopherin receptors, or even Ran itself, play a central role in mRNA export. Although the mechanistic details of mRNA export have not yet been fully elucidated, it appears that there are two classes of proteins that are required to achieve export of mature messages. First, there is a family of evolutionarily conserved heterogeneous nuclear ribonucleoproteins (hnRNPs) that interact with poly (A)+ RNA in vivo [25]. A number of these hnRNP proteins shuttle between the cytoplasm and the nucleus and escort the poly (A)+ RNAs as they are exported through the NPC [76, 93]. Current models suggest that at least some of the hnRNP proteins may be involved in RNA processing steps that occur co-transcriptionally. The hnRNPs that remain bound to the maturing messages may serve as markers that the different processing steps have been successfully accomplished. The second class of proteins consists of those proteins that have been implicated more directly in the export process, including the helicase, Sub2p (UAP56 in humans), and the heterodimeric export receptor, Mex67p/Mtr2p (TAP/p15 in humans) [20, 58, 92]. TAP was originally identified as a factor necessary for
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the export of simian type D viral RNAs that contain a CTE (constitutive transport element). Subsequent experiments showed that TAP/Mex67p is also required for the export of endogenous mRNAs [57, 58] and that mutations in the yeast MEX67 gene cause a rapid onset defect in the export of poly (A)+ RNA [92]. TAP/Mex67p shuttles between the cytoplasm and the nucleus [58], and in complex with p15/Mtr2p, binds both to mRNA and to nucleoporins [2]. Thus, it could potentially target bound RNAs directly to the NPC for export. Recent work has led to the identification of the TREX (TRanscription/EXport) complex, which links mRNA transcription and export from the nucleus [83]. This evolutionarily conserved complex contains both factors required for transcription of mRNA and factors that will be required for export of the mature message. A great deal of effort is currently focused on understanding how the hnRNP proteins, the export factors, and other accessory proteins cooperate to export mature mRNA from the nucleus. There are a number of studies that demonstrate increased expression of several different hnRNP proteins in tumor cells as compared to normal control cells. For example, Snead et al. [96] found that hnRNP B1 was overexpressed in 84% of malignant tumors in lung disease as compared to only 18% of benign tumors. This and other related studies have led to the suggestion that hnRNP expression could be a useful marker for the early detection of specific tumors [112].
4
REGULATION THROUGH NUCLEAR TRANSPORT
We are only just beginning to reveal the role that nucleocytoplasmic transport plays in cancer cell transformation and studies in S. cerevisiae and S. pombe have been critical in advancing our knowledge. Studies of the cell cycle regulatory proteins cyclin B1 (Clb2p in S. cerevisiae) and Cdc25C (cdc25 in S. pombe), as well as the yeast transcription factor, Pho4p, have helped to delineate the mechanisms whereby these growth regulatory proteins traffic between the cytoplasm and the nucleus [37, 116]. Here we discuss advances made using yeast, as well as other model systems to understand the nuclear translocation of cyclin B1/Clb2p, Cdc25p, and Pho4p. Specifically we will review how the cell has utilized this transport to regulate the functions of these proteins and thereby the timing and accuracy of cellular processes.
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Cyclin B/Clb2p
Cyclin was originally identified as a protein that gradually increases in concentration during the cell cycle and then is degraded at the onset of each cellular division [26]. A family of proteins known as the cyclins has since been identified, with homologues in all eukaryotes, that all contain a conserved protein kinase-binding site called the cyclin box. These proteins function as regulatory subunits of these kinases, known as cyclin-dependent kinases [24]. Studies in yeast showed that these cyclin/cyclin-dependent kinase complexes regulate the timing of cell cycle progression. Furthermore, their regulated activity is necessary for spindle disassembly, cytokinesis, and G1 transition [101]. Mitosis is triggered by the activity of the cyclin/cyclin-dependent kinase complex in eukaryotes. To ensure that mitosis is not entered prematurely, the activity of this complex is regulated through the steady state cytoplasmic compartmentalization of the kinase and cyclin subunits [104], inactivating phosphorylation of the complex subunits, as well as degradation of the cyclin subunit, which occurs through the ubiquitin-mediated pathway [33]. These control mechanisms ensure that the cell does not prematurely enter the next phase of the cell cycle. At the beginning of S-phase, cyclin B/Clb2p begins to accumulate within the cytoplasm of cells, but as the cell cycle progresses into prophase, cyclin B/Clb2p is relocalized to the nucleus [81]. As shown in Figure 3 (Cyclin B and Cdc25) this nuclear localization is due to import through the classical import pathway of Kap60p(α) and Kap95p(β) in yeast [45] or through a direct interaction with importin β in humans [73]. The protein is retained within the nucleus because of the inability of the export receptor, CRM1, to bind to the NES of a phosphorylated form of cyclin B/Clb2p [114]. Once cyclin B/Clb2p is localized within the nucleus, cyclin-dependent kinase 1 (Cdc2 in mammalian cells/Cdc28p in S. cerevisiae) is activated to promote mitosis. Studies using S. cerevisiae have shown that as the APC/cyclosome, a ubiquitin ligase activity containing complex [60], regains activity during mitosis, cyclin B/Clb2p is targeted for destruction [3, 117] and this destruction is dependent on CRM1-mediated export [120] to the cytoplasmic 26S proteasome, allowing for transition into G1 of the next cell cycle. Thus, the cell utilizes the intrinsic nuclear localization and export sequences contained within cyclin B/Clb2p, in addition to the transport machinery, to modulate the activity of the Cdc2–cyclin B complex.
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Figure 3. Cyclin B. Mitotic progression through the cell cycle depends on translocation of the Cdc2–Cyclin B complex into the nucleus. The dimeric complex is dephosphorylated on Cdc2p by Cdc25p, allowing access to the import–receptor complex consisting of importin α/β. The quaternary complex translocates through the nuclear pore and dissociates upon RanGTP binding to β in the nucleus. The Cdc2–Cyclin B complex is retained in the nucleus by inhibitory phosphorylation of Cyclin B. At the end of mitosis, the complex is dephosphorylated, allowing CRM1 to bind to the NES of Cyclin B. Upon binding, the complex is exported from the nucleus and targeted for destruction by the 26S proteasome. Cdc25. The 14-3-3 protein binds to and holds Cdc25p in an inactive state in the cytoplasm during interphase by blocking access to its intrinsic NLS. Upon activating phosphorylation of Cdc25p at prophase, importin α/β binds to the Cdc25p NLS and the trimeric complex is translocated into the nucleus. The complex is dissociated by binding of RanGTP to β. Cdc25p loses activity upon dephosphorylation, exposing its intrinsic NES, which is then bound by CRM1. The dimeric complex is then translocated through the nuclear pore into the cytoplasm where the complex is dissociated. Pho4. Under low phosphate conditions, the activity of the Pho80p/Pho85p kinase complex is downregulated, leading to an underphosphorylated form of Pho4p. Dephosphorylation reveals the NLS of Pho4p and allows binding by its import receptor Pse1p. The dimeric complex is translocated into the nucleus where it is dissociated by RanGTP binding to Pse1p. After Pho4p activates transcription of the phosphate responsive genes, Pho4p becomes hyperphosphorylated, creating a binding site for its export receptor Msn5p to form a trimeric complex with RanGTP. Following binding by Msn5p, Pho4p is rapidly recycled to the cytoplasm where the trimeric complex is dissociated by hydrolysis of RanGTP to RanGDP.
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Cyclin B1 misexpression has been found in numerous cancers ranging from prostate carcinomas [29] to esophageal squamous cell carcinomas [103]. In normal esophageal squamous cells, cyclin B1 is localized in the nucleus and expressed in only a small fraction of cells. However, in esophageal squamous cell carcinomas, cyclin B1 is localized to the cytoplasm and expressed in a large number of cells [103]. This mislocalization of cyclin B1 has also been seen in other solid tumors as well as lymphomas and leukemias [119]. These observations suggest that the proper localization of cyclin B1 may be important in the maintenance of normal cell growth and the prevention of tumor formation.
4.2
Cdc25
Evidence from S. pombe has provided much of the foundation for the regulation of mitotic entry in eukaryotic cells (Figure 3). This entry is triggered by the cyclin-dependent kinase, Cdc2/Cdc28p. As discussed above, the activity of Cdc2/Cdc28p is dependent upon interaction with its regulatory subunit, cyclin B/Clb2p. Prior to mitosis, Cdc2–cyclin B complexes are held in an inactive state by phosphorylation of Cdc2/Cdc28p. The cell division cycle 25C protein phosphatase, Cdc25C/Cdc25p, acts on the Cdc2– cyclin B complex to dephosphorylate Cdc2/Cdc28p, thus activating the Cdc2–cyclin B complex to promote mitosis. In order to prevent early activation, the cell must regulate the activity of Cdc25C/Cdc25p. This is accomplished by regulating the nuclear transport of Cdc25C/Cdc25p. Studies in both mammalian cells and fission yeast have shown that Cdc25C/Cdc25p shuttles between the nucleus and cytoplasm throughout the cell cycle, but has a steady state cytoplasmic localization during interphase and a nuclear localization during prophase [41, 69]. Cdc25C/Cdc25p contains both a classical NLS and an NES and studies using Xenopus and S. pombe have shown that Cdc25C/Cdc25p is imported through interactions with importin-α/Kap60p [65, 115] and importin-β/Kap95p [15]. Posttranslational modifications near these targeting sequences control the steady state localization of the protein by interfering with the recognition of these signals by the receptors. During interphase in both Xenopus [66] and S. pombe, [121] phosphorylation of Cdc25C/Cdc25p near its intrinsic NLS creates a binding site for the phosphoserine-binding protein, 14-3-3. The 14-3-3 protein binds to Cdc25C/Cdc25p and inhibits nuclear import probably by sterically hindering the access of importin α to the NLS. This causes a redistribution of Cdc25C/Cdc25p from the nucleus to the cytoplasm. This model is supported by experiments from fission yeast, which show that in the absence of the 14-3-3 protein, Rad24p, Cdc25C/Cdc25p is predominantly nuclear
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[69]. Thus, 14-3-3 proteins regulate Cdc25C/Cdc25p function by inhibiting its nuclear import. In addition to 14-3-3 inhibition of nuclear import, Cdc25C/Cdc25p contains a leucine-rich NES that is recognized by the export factor CRM1 [31, 69]. Experiments in both S. pombe and human cells show that mutation or inhibition of CRM1 leads to nuclear accumulation of Cdc25C/Cdc25p [36, 69]. This suggests that Cdc25C/Cdc25p is actively exported from the nucleus throughout interphase and that this export is CRM1-dependent. Cdc25C is involved in the formation of acute myeloid leukemia [47] and cervical adenocarcinoma [37], and is the target of a number of anticancer agents [37, 118]. Treatment of HeLa cells with the anticancer drug, UCN-01, directly perturbs the normal cytoplasmic localization of Cdc25C causing a nuclear accumulation [37]. In combination with DNA damage, this nuclear accumulation of Cdc25C could potentially drive the cells to enter mitosis prematurely by activating the Cdc2-cyclin B complex and bypassing the normal G2/M checkpoint, leading to apoptosis. Thus, understanding the regulation of Cdc25C nucleocytoplasmic transport may provide new potential therapeutic targets for the treatments of leukemia and cervical cancer.
4.3
Pho4p
Cellular response to environmental signaling allows for adaptation to changes in growth conditions. This generally occurs through alterations in gene expression by a signal-cascade event originating in the plasma membrane, progressing through the cytoplasm, and ending with upregulation of gene expression in the nucleus. Cells can be primed for rapid response to extracellular signals by constitutively synthesizing the protein effectors while at the same time inhibiting their function. These effectors are generally transcription factors and one method for inhibiting their function is the retention of these molecules in the cytoplasm until the signal needs to be transduced into the nucleus. Cellular signaling can be accomplished through phosphorylation of the transcription factor to either activate or inactivate the protein. This phosphorylation can occur through complexes such as cyclin–CDK. A paradigm for signal transduction in S. cerevisiae is the cyclin–CDK complex, Pho80p–Pho85p (Figure 3). This complex recognizes phosphate depletion from the environment and transduces its signal through the phosphorylation of the transcriptional activator Pho4p [52]. Under conditions where phosphate levels are low, the Pho80p–Pho85p complex activity is downregulated [91]. This results in the accumulation of an unphosphorylated form of Pho4p [52] that is translocated into the nucleus by its import receptor, Pse1p [54], a Kap protein that preferentially associates
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with the unphosphorylated form of Pho4p [54]. Pho4p then activates the transcription of the phosphate-responsive genes. When phosphate levels rise, the Pho80p–Pho85p complex regains activity and hyper-phosphorylates Pho4p. This phosphorylation causes Pho4p to be rapidly recycled to the cytoplasm through its export receptor, Msn5p [53], an importin β family member that specifically recognizes the phosphorylated form of Pho4p [53], thus repressing the transcription of the phosphate-responsive genes. The Pho80p–Pho85p complex phosphorylates Pho4p on multiple serine resides [64]. Phosphorylation of Pho4p on two sites is necessary and sufficient to cause nuclear export by Msn5p [64]. Additional phosphorylation of Pho4p on a third residue within the nuclear localization signal inhibits its interaction with the import receptor, Pse1p [64]. Both of these phosphorylation events serve to ensure that Pho4p remains cytoplasmic under conditions of high phosphate levels. Thus, phosphorylation of these residues serves to regulate the nucleocytoplasmic transport of Pho4p. By examining nuclear transport of Pho4p, we can better understand how the activity of transcription factors is regulated in response to environmental signals.
5
CONCLUSIONS/IMPLICATIONS FOR THE FUTURE
Obviously the use of yeast as a model system to study the nuclear transport process has been extremely valuable in the past. The prediction is that future studies of this highly conserved process in model organisms including yeast will provide further insights. The goal is to define the mechanisms that regulate nuclear transport of critical growth regulatory molecules. These transport mechanisms could then be specifically targeted for novel therapeutic regimes.
ACKNOWLEDGMENTS We would like to acknowledge the members of the Corbett laboratory for helpful discussions and comments. We thank Dr. Deanna Green for careful reading of the manuscript. This work was supported by grants from the NIH to AHC and ACB.
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Chapter 4
STUDIES OF PROTEIN FARNESYLATION IN YEAST
Nitika Thapar and Fuyuhiko Tamanoi Department of Microbiology, Immunology & Molecular Genetics, Jonsson Comprehensive Cancer Center, Molecular Biology Institute, University of California, Los Angeles, 405 Hilgard Ave., Los Angeles, CA 90095-1489
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PROTEIN FARNESYLATION
Protein farnesylation has emerged as one of the important classes of posttranslational modification of proteins [84, 109]. Farnesylation is the first step in a three-step series of protein processing and involves the addition of a farnesyl group to a cysteine located in a C-terminal motif called the CaaX motif (C is cysteine, a is aliphatic amino acid, and X is the C-terminal amino acid that is usually methionine, cysteine, glutamine, alanine, or serine). These proteins subsequently undergo proteolytic cleavage and carboxylmethylation (Figure 1). Significance of protein farnesylation in human cancer has been underscored by the findings that many proteins undergoing farnesylation play roles in signal transduction. Of these, Ras proteins are particularly noteworthy.
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CaaX Farnesyl Pyrophosphate (FPP)
Protein Farnesyltransferase
Pyrophosphate (PP) CaaX aaX
CaaX Protease
C
Carboxyl Methyltransferase
C
Me
Figure 1. Posttranslational modification of proteins containing the CaaX motif at their Cterminus. Protein farnesyltransferase transfers a farnesyl group on the cysteine residue which is followed by cleavage of the last three amino acids and subsequent methylation of the Cterminal carboxyl group. (In the CaaX motif C is cysteine, a is an aliphatic amino acid and X is the C-terminal amino acid.)
Protein farnesylation is catalyzed by protein farnesyltransferase (FTase) that is conserved from yeast to human [84, 109]. This enzyme recognizes the CaaX motif and transfers a farnesyl group from farnesyl pyrophosphate (FPP), an intermediate in cholesterol biosynthesis, resulting in the formation of a thioether bond. The catalytic mechanism of action of the FTase has been elucidated [91]. The enzyme contains one molecule of a tightly bound zinc ion (Zn2+) that participates in the catalytic reaction [51, 91, 97]. With yeast FTase, the reaction proceeds by an ordered mechanism with FPP binding first. In the case of mammalian enzyme, it has been suggested from kinetic and structural studies that the bound FPP forms part of the binding surface of the protein substrate [36]. Three-dimensional structures of the rat and human enzymes have been determined with and without bound substrates [65, 82, 95]. The structure mainly consists of α-helices with the β-subunit forming a barrel like structure and the α-subunit wrapping around the β-subunit from one side of the barrel. FTase belongs to a family of protein prenyltransferases which includes protein geranylgeranyltransferase type I (GGTase I) and type II (GGTase II). GGTase I catalyzes geranylgeranylation of proteins such as RhoA, Cdc42, or
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Rac proteins [24, 39, 86]. The modification involves a 20-carbon geranylgeranyl group added to a cysteine in the CaaL motif (similar to the CaaX motif except that the C-terminal amino acid is leucine or phenylalanine) [74, 108]. GGTase I shares a common α-subunit with FTase, while its β-subunit shares about 30% homology with the β-subunit of FTase. GGTase II catalyzes the addition of a geranylgeranyl group to both cysteines within the CC or the CXC motif that are found in a number of Rab protein [88]. In addition to α- and β-subunits that share homology with their counterparts in FTase, GGTase II contains an additional component-Rep (Rab escort protein) [96]. Recently, anticancer drugs based on the inhibition of protein farnesylation have been developed. These small molecular weight compounds called farnesyltransferase inhibitors (FTIs) selectively inhibit farnesyltransferase [7, 38, 84, 94]. A variety of compounds including peptidomimetic inhibitors, farnesyl pyrophosphate analogues, bisubstrate inhibitors, and natural compounds have been identified. FTIs have been shown to reverse rasmediated phenotypes in ras-transformed cells. FTIs inhibit the growth of tumors or even regress tumors in animal model systems [62]. Currently, FTIs are being evaluated in a variety of clinical trials [57]. In this review, we discuss how yeast studies contributed to the overall study of protein farnesylation. We will first describe yeast studies concerning FTase as well as yeast-based assays that led to the identification of one of the first generation FTI compounds. We will then focus on identification and characterization of farnesylated proteins. Yeast has provided a comprehensive analysis of farnesylated proteins that was instrumental in the identification of a number of novel farnesylated proteins. Of particular interest is Rheb, a novel family of the Ras-superfamily G-proteins. This protein is highly conserved and plays a role in the regulation of cell cycle at the G1/S phase. We will summarize the current understanding of Rheb in S. cerevisiae and S. pombe.
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CONTRIBUTION OF YEAST TO THE DISCOVERY AND CHARACTERIZATION OF PROTEIN FARNESYLTRANSFERASE
2.1
Identification of genes encoding subunits of protein farnesyltransferase
DPR1/RAM1 and RAM2 encode β and α subunits of FTase in S. cerevisiae, respectively [40, 41, 48]. These genes were identified from the study to isolate mutants defective in membrane association of Ras proteins. The isolation of the mutants involved reversion of ras activated phenotypes in S. cerevisiae. Briefly, mutants were sought that reverse phenotypes due to the activation of Ras2 such as heat shock sensitivity, decrease in glycogen accumulation, and temperature sensitivity [34, 80]. DPR1/RAM1 and RAM2 encode proteins with 431aa and 316 aa, respectively. They share approximately 30% identity with corresponding subunits of mammalian FTase. Sequence characterization of Ram2 and the α subunit led to the finding of five tandem sequence repeats in these subunits [11]. Identification of these genes was important in the study of FTase in mammalian cells. The presence of two distinct genes affecting farnesylation provided hints for the idea that the mammalian enzyme consists of two subunits. Sequence of DPR1/RAM1 determined in 1987 by our group provided the first information on one of the FTase subunits [40]. Subsequent determination of mammalian FTase βsubunit sequence in 1991 revealed sequence homology with Dpr1/Ram1 [21]. Sequence homology between Ram2 and the α-subunit was also demonstrated [61]. Recently, we have identified genes encoding subunits of FTase in S. pombe. The cwp1+ and cpp1+ genes encode α and β subunits of FTase, respectively [4, 105]. Co-expression of Cpp1 and Cwp1 in Escherichia coli produces a recombinant enzyme that is active in farnesylation [105]. The cwp1+ was identified from the yeast two hybrid screen using cwg2+ [4]. cwg2+ encodes the β-subunit of GGTase I, bearing similarity to its homologue in S. cerevisiae-CAL1/CDC43 [73], and is involved in cell wall synthesis [30].
2.2
Mutational analysis of protein farnesyltransferase
One of the unique properties of FTase is its ability to recognize the CaaX motif. This observation was critical in the development of peptidomimetic FTIs, as they were derived from a tetrapeptide such as CVFM that inhibited the activity of FTase. Thus, it was of interest to understand which amino
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acid residues were involved in the CaaX recognition. We have employed PCR-mediated random mutagenesis of S. cerevisiae FTase to gain insight into this issue. The DPR1/RAM1 gene was randomly mutagenized and the mutant library was screened for its ability to suppress temperature-sensitive growth of cal1 mutant which is defective in GGTase I [28, 71]. The intention here was to identify amino acid changes that converted the recognition of the CaaX motif to the CaaL motif. This screen led to the identification of amino acid changes at three residues-S159, Y362, and Y366 [28]. A change in either of these residues was sufficient to convert FTase to mutants that had increased recognition of the CaaL motif. Y362 and Y366 are perfectly conserved in all farnesyltransferase subunits examined. The Y362 mutant is well characterized but Y366 awaits further investigation. The residues Y362 and Y366 correspond to residues Y361 and Y365 in the mammalian enzyme. Alteration of Y361 to leucine in the human enzyme led to an increase in the utilization of GST–CIIL substrate [29], suggesting that this residue is important in the recognition of the CAAL motif. However, a recent kinetic study suggests that the change in the CaaX motif recognition by FTase by the alteration of Y361 reflects a kinetic effect due to the complex mechanism of FTase [90]. In the crystal structure of mammalian FTase [92, 95], Y361 is located in a pocket that would be occupied by the second aliphatic amino acid in the CaaX motif. In addition, P152, which corresponds to S159, is located close to the C-terminal amino acid of the CaaX motif.
2.3
Identification of inhibitors of protein farnesyltransferase by yeast-based assays
By using yeast-based assays, we have obtained one of the first generation compounds of FTIs [46, 72]. This microbial screen was based on our observation that Ste18 protein is farnesylated [33]. This γ-subunit of the heterotrimeric G-protein is involved in yeast-mating functions to inhibit cell growth by activating a MAP kinase cascade that consists of Ste11, 7, and Kss/Fus [44, 49]. This growth inhibitory effect is normally blocked, as Ste18 is complexed with the α-subunit. Thus, the disruption of the α-subunit is lethal. However, this lethality is suppressed when the farnesylation of Ste18 is inhibited. Culture media from a variety of microbial sources were screened that led to the identification of manumycin as an inhibitor of FTase [46]. Manumycin inhibited FTase by competing with FPP and inhibited rasactivated phenotypes in yeast as well as in C. elegans [47, 72]. Manumycin also inhibited proliferation of pancreatic cancer cells and the growth of tumors in mice [54, 56].
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YEAST STUDIES GREATLY EXPANDED OUR UNDERSTANDING OF FARNESYLATED PROTEINS
One of the current emphases of the farnesylation study is to determine substrates of farnesylation. The first group of farnesylated proteins discovered included fungal mating factors, nuclear lamins, and proteins involved in visual signal transduction such as transducin [15, 24]. The significance of farnesylation in cancer came into prominence when Ras was found to be farnesylated [19, 45, 85]. Ras is the product of an oncogene and mutations of Ras have been found in approximately 30% of human cancers [12]. Farnesylation is found to be critical for membrane localization of Ras and it has also been demonstrated that mutations of the CaaX motif inhibit the function of Ras [66]. A number of other proteins are also farnesylated. Thus, identification and characterization of novel farnesylated proteins becomes crucial for understanding how farnesylation regulates various cellular mechanisms.
3.1
Farnesylated proteins in S. cerevisiae
The sequencing of the S. cerevisiae genome has greatly facilitated the search for proteins which are possible substrates of FTase, based on the presence of the CaaX motif at their C-termini. Table 1a lists the proteins in S. cerevisiae which are farnesylated or predicted to be farnesylated. Ras1 and 2 proteins are GTP-binding proteins and are the homologues of the mammalian proto-oncogene product Ras [27, 79]. They are the major players in the signal transduction pathway involving adenylyl cyclase (Cyr1) [98]. Farnesylated Ras2–GTP complex was found to activate Cyr1 in vitro more efficiently than either the unprenylated protein or a C-terminal deletion mutant [63]. Invasive growth is regulated via the Cdc42–Ste20–MAPK cascade as well as the cAMP-dependent protein kinase pathway [75]. The critical role of farnesylation was studied by making CaaX box mutations as well as deletions. The C319S mutant of Ras2 is defective in farnesylation and all subsequent modifications i.e., proteolytic cleavage and methylation. Spores containing this mutant fail to yield colonies in the absence of Ras1 [10].
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Table 1a. S. cerevisiae CaaX motif-containing proteins which have been shown to be farnesylated or are candidates for farnesylation based on sequence similarity Protein CAAX motif Function/cellular role Ras1 -CIIC G-protein, cAMP pathway Ras2 -CIIS G-protein, cAMP pathway Mfa1 -CVIA Mating pheromone a-factor Mfa2 -CVIA Mating pheromone a-factor Stel8 -CTLM Gγ involved in mating Ydj1 -CASQ DnaJ homolog, protein transport Xdj1j -CCIQ DnaJ homolog Rho3 -CTIM Cell polarity Rho4 -CIIM Cell polarity/Actin cytoskeleton organization Rheb -CSIM GTP binding/Amino acid metabolism Skt5 -CVIM Cell wall maintenance Gis4 -CAIM Signal transduction/Cell stress Rcy1 -CCIM Vesicular transport Pex19 -CKQQ Peroxisome biogenesis Atr1 -CTVA Small molecule transport N1142 -CSIM Hemoprotein Ykt6 -CIIM v-SNARE, vesicular transport YCP4 -CTVM Unknown YDL009C -CAVS Unknown YPL191C -CVVM Unknown YGL082W -CVIM Unknown YMR265C -CSNA Unknown YML133C -CCPS Unknown YPR203W -CCPS Unknown YFL065C -CCPS Unknown YHL049C -CCPS Unknown YDL151C -CYPA Unknown YJL118W -CCCS Unknown YIR007W -CVIS Unknown
Other Ras-superfamily G-proteins that are farnesylated are Rho3, 4, and Rheb. Rho3 is involved in cell polarity [1], and we have shown that it directly binds to Exo70, a subunit of the exocyst, a multiprotein complex affecting fusion of secretory vesicles with the plasma membrane [83]. Rheb is a recently discovered GTP, binding protein belonging to the Ras superfamily of G proteins. It bears similarity to the Ras, Ral, and Rap proteins. In S. cerevisiae, it has been shown to be involved in the uptake of arginine and lysine. This appears to be due to the modulation of the activity of the permeases Can1 and probably Lyp1 [101]. Mutation of the CaaX motif (C206S) eliminates its function. The null mutant is viable but displays hypersensitivity to canavanine and thialysine, the toxic analogues of arginine and lysine respectively. Later in the chapter, we provide a more detailed description of Rheb.
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The Mfa1 and 2 proteins are mating pheromones (mating factor a) exported from the cells by Ste6, which interact with α cells to produce cell cycle arrest and mating responses [70]. Mutant forms of the proteins with different substitutions in the CaaX motif are poor substrates for in vitro farnesylation [99]. C-terminal farnesylation was shown to be required for the production and export of a-factor-related peptide (AFRP) [22]. Ste18 is the γ subunit of the guanine nucleotide-binding protein that mediates signal transduction by pheromones during mating [9]. It is anchored to the plasma membrane through hydrophobic interactions with its farnesylated Cterminus. It activates the pheromone response pathway with Ste4 [81]. Mutation in the CaaX motif (C107S) prevents farnesylation and results in a decrease of its steady state levels [33, 52, 103]. This mutant has decreased response to pheromones [33, 103]. Ydj1 is a protein involved in protein import into mitochondria and endoplasmic reticulum and is the homologue of E. coli DnaJ [17]. It belongs to the Hsp40 family of co-chaperones and is required for the ubiquitinationdependent degradation of short-lived and abnormal proteins [64, 67]. It associates with the membrane through its farnesyl group, and C406S mutant exhibits decreased membrane association [18, 55]. C-terminal farnesylation is also required for growth at elevated temperatures [18]. Xdj1, a nonessential homologue of DnaJ was identified as an ORF in the genomic DNA, however no mRNA or protein for it has been detected [87]. Hdj2, a human homologue of Ydj1, has recently been shown to provide a valuable tool to follow inhibition of farnesylation in peripheral blood of patients treated with FTI (2). Pex19 is a protein involved in peroxisome membrane formation and maintenance, which interacts with Pex3 only when it is farnesylated [42, 50]. Farnesylation at the cysteine in its CKQQ motif is essential for its function in vivo [42]. Several other proteins predicted to be farnesylated include known proteins such as Gis4 (protein implicated to be involved in the cAMP pathway) [8], Skt5 (a killer toxin-resistant protein) [13, 14], Rcy1 (protein involved in endocytic membrane traffic) [37], Ykt6 (synaptobrevin homologue) [69, 100] as well as proteins with unknown function. There are other proteins which have been predicted to be farnesylated but not yet characterized.
3.2
Farnesylated proteins in S. pombe
The fission yeast S. pombe has a few proteins functionally related to the S. cerevisiae proteins which have been shown to be farnesylated. As summarized in Table 1b, these include Mfm1, 2 and 3, the functionally redundant precursor polypeptides for the mating pheromone M factor
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produced by h– cells which are structurally similar to the a-factor of S. cerevisiae [60]. They are posttranslationally cleaved to yield the sequence YTPKVPYMCVIA which is isoprenylated on the cysteine [26]. A synthetic M-factor peptide lacking any modification is inactive. Ras1 is the homologue of mammalian Ras and regulates cell morphogenesis and is required for the nutrient starvation signal transduction pathway that leads to mating [3, 35]. Rheb is a member of the Rheb GTPases but divergent from the S. cerevisiae ortholog. It is known to perform functions distinct from Ras1 [68]. The null mutant is lethal and the inhibition of Rheb farnesylation leads to an arrest in the G0/G1 phase [68, 106]. Farnesylation-deficient mutant of Rheb (SVIA) fails to rescue the rhb1– mutant suggesting that farnesylation is important for its activity [106]. Rho2 and Rho 3 are members of the Rho family of proteins involved in the control of cell morphology and cell polarity [5]. Rho2 is involved in the regulation of cell wall synthesis through the interaction with PKC homologues [16]. Lack of Rho3 causes abnormal cell morphology and cytokinesis defects [105]. Table 1b. S. pombe CaaX motif-containing proteins which have been shown to be farnesylated or are candidates for farnesylation based on sequence similarity Protein CAAX motif Function/cellular role SpRas -CVIC GTP binding/mating response Mfm1 -CVIA Signal transduction/mating response Mfm2 -CVIA Signal transduction/mating response Mfm3 -CVIA Signal transduction/mating response Rhb1 -CVIA Cell cycle regulator Spj1 -CAQQ Protein folding/cell stress Rho2 -CIIS Cell polarity/cell wall maintenance Rho3 -CIIA Cell polarity/cell wall maintenance Git11 -CTIS Signal transduction/mating response SPBC405.06 -CQAQ Unknown (has DnaJ domain) SPBC13G1.11 -CIIA Unknown (predicted SNARE) SPCC417.05C -CIIS Unknown (predicted chitin biosynthesis) SPAC24B11.10C -CVVM Unknown (predicted chitin biosynthesis) SPAC607.09C -CALT Unknown (predicted cellular pH homeostasis) SPAC17C9.14 -CPTQ Unknown (predicted peroxisome biogenesis)
4
RHEB, A NOVEL CLASS OF FARNESYLATED PROTEINS INVOLVED IN THE REGULATION OF CELL CYCLE PROGRESSION
Among many farnesylated proteins, Rheb has recently emerged as an important protein. This protein belongs to the Ras superfamily of G- proteins.
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There are features unique to the Rheb subfamily. Recent studies indicate the need of Rheb to be farnesylated for its activity. In the following sections, we describe characterization of this novel member of the Ras family.
4.1
Rheb binds GTP and is farnesylated
Rheb is a small GTP-binding protein belonging to the Ras superfamily. It was first identified as the product of an inducible gene in the rat hippocampal granule cells, which was induced in response to seizures and by NMDAdependent synaptic activity in the long-term potentiation paradigm [104]. It is well conserved and homologues have been identified in human [43, 73], Xenopus, zebrafish, Drosophila, sea squirt, Dictyostelium, Aspergillus [77], S. cerevisiae, and S. pombe [101, 102] (Figure 2). In mammals, two genes for Rheb exist – Rheb1 and Rheb2, unlike lower species which have a single gene only [6, 78]. Its amino acid sequence shows the presence of highly conserved GTP-binding regions as well as the CaaX motif. Interestingly, Rheb proteins have an arginine residue at the third position in the G1-box (corresponding to the 12th amino acid in Ras, which is involved in the intrinsic GTPase activity of the protein) unlike most other Ras superfamily proteins which have a glycine residue (Figure 3). The effector domain (F-V-E/ D-S-Y-Y/D-P-T-I-E-N-E/Q/T-F-T/S/N-R/K-x-x) is well conserved among Rheb members. Rheb shares the highest amino acid homology with human H-Ras (36%), human Rap2 (38%), and yeast Ras1 (43%). It is localized to the plasma membrane like the Ras protein and found to be farnesylated in vivo; treatment with farnesyltransferase inhibitors (FTIs) leads to its mislocalization to the cytoplasm [23]. In humans it is ubiquitously expressed, though high levels of expression were observed in the skeletal
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ScRheb SpRheb HsRheb DmRheb
G1 MEYATMSSSNSTHNFQRKIALIGARNVGKTTLTVRFVESR M----------APIKSRRIAVLGSRSVGKSSLTVQYVENH M----------PQSKSRKIAILGYRSVGKSSLTIQFVEGQ M----------P-TKERHIAMMGYRSVGKSSLCIQFVEGQ * . *.**..* *.***..* ...**..
40 30 30 29
ScRheb SpRheb HsRheb DmRheb
Effector Domain G3 FVESYYPTIENEFTRIIPYKSHDCTLEILDTAGQDEVSLL FVESYYPTIENTFSKNIKYKGQEFATEIIDTAGQDEYSIL FVDSYDPTIENTFTKLITVNGQEYHLQLVDTAGQDEYSIF FVDSYDPTIENTFTKIERVKSQDYIVKLIDTAGQDEYSIF **.** *****.*.. .... ...******* *..
80 70 70 69
ScRheb SpRheb HsRheb DmRheb
NIKSLTGVRGIILCYSIINRASFDLIPILWDKLVDQLGKD NSKHSIGIHGYVLVYSITSKSSFEMVKIVRDKILNHTGTE PQTYSIDINGYILVYSVTSIKSFEVIKVIHGKLLDMVGKV PVQYSMDYHGYVLVYSITSQKSFEVVKIIYEKLLDVMGKK . .* .* **... **... .. .*... *.
120 110 110 109
ScRheb SpRheb HsRheb DmRheb
G4 NLPVILVGTKADLGRSTKGVKRCVTKAEGEKLASTIGSQD WVPIVVVGNKSDLHM-----QRAVTAEEGKALANE----QIPIMLVWNKKDLHM-----ERVISYEEGKALAES----YVPVVLVGNKIDLHQ-----ERTVSTEEGKKLAES----.*...* .* ** .* .. .**. **..
160 140 140 139
ScRheb SpRheb HsRheb DmRheb
G5 KRNQAAFIECSAELDYNVEETFMLLLKQMERVEGTLGLDA --WKCAWTEASARHNENVARAFELIISEIEKQAN--PSPP --WNAAFLESSAKENQTAVDVFRRIILEAEKM-D--GAAS --WRAAFLETSAKQNESVGDIFHQLLILIENE-N--GNP. *. * ** . .. * .. *. .
200 176 175 173
ScRheb SpRheb HsRheb DmRheb
CAAX ENNNKCSIM GDGKGCVIA QGKSSCSVM QEKSGCLVS . . * .
209 185 184 182
Figure 2. Amino acid sequence alignment of Rheb protein from various species (Sc – S. cerevisiae; Sp – S. pombe; Hs – Human; Dm – D. melanogaster). The G-boxes (G1–G5) are indicated by overlining. The invariant arginine in the G1 box, the effector domain and the CaaX motif are shown in bold. Asterisks represent identical residues and dots represent similar residues.
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G1
1
G2
G3
G1
GSRSVGKS
G5
S. cerevisiae Rheb
GARNVGKT
1
G4
G2
G3
G4
S. pombe Rheb
209
CSIM
G5
185
CVIA
Figure 3. Structure of S. cerevisiae and S. pombe Rheb proteins. The G-boxes are represented as dark segments. The invariant arginine in the G1 box as well as the C-terminal CaaX motif are indicated.
and cardiac muscle [43]. Rheb mRNA has been found to be induced in cerebral-ischemia elicited events through NMDA receptor action in rat brain [58] as well as in human fibroblasts exposed to UV radiation implying a possible role in UV sensitization of cells [59]. Human Rheb was reported to be upregulated in several transformed cells [43]. The growth regulatory activities of Rheb were explored by Clark et al. [23] who found that neither the wild type nor the presumed constitutively active (Q64L) Rheb protein could induce transformation in normal NIH3T3 cells. At the same time, a mutant Rheb, S20N that carries an amino acid change analogous to the dominant negative form of Ras (S17N) did not inhibit growth of the cells. Rheb was found to behave more like Rap1A [23], which is a negative regulator of Ras function, as it was also found to antagonize the oncogenic potential of Ras. It was also speculated that since Rheb and Ras show strong identity in the effector domain region which is crucial for Raf binding, Rheb may be binding to Raf in a nonproductive manner and hence titrating it away from Ras and preventing the downstream cascade [23]. On the other hand, Yee and Worley [107] found that Rheb stimulates transformation of NIH3T3 fibroblasts when expressed in conjunction with Raf-1. Thus, a synergistic interaction of Rheb with Raf-1
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was suggested, as the transforming potential was much lower when Rheb or Raf-1 alone was expressed singly. A report showing that Rheb inhibits B-Raf has been published [53]. The Rheb protein became a prime focus of attention recently after Drosophila and mammalian studies showed that it is a component of the insulin/TOR/S6K pathway. Rheb activates S6K-mediated by TOR. Rheb was found to be the direct target of the Tsc1/Tsc2 complex which serves as a GAP for Rheb [6, 78]. Mutations in the Tsc1/Tsc2 genes have been shown to be responsible for the development of tuberous sclerosis (TSC), a genetic disease characterized by the presence of benign tumors known as hamartomas in various organs and occurrence of seizures and mental retardation. Studies involving Drosophilia and human Rheb indicate a potentially critical role of Rheb in the pathogenesis of this disease. Thus, a further investigation of the function of Rheb may provide insights into possible therapies for TSC.
4.2
Genetic study of Rheb in S. cerevisiae revealed the involvement of Rheb in arginine uptake
Rheb was first identified and characterized in S. cerevisiae (ScRheb) by Urano et al. [101]. It shares an overall 26% sequence identity and 57% similarity with human and fly Rheb proteins. The genetic study was undertaken by creating a deficient strain and analyzing its phenotypes in the total absence of endogenous Rheb or in the presence of mutant forms of the protein. The disruption strain showed no loss in viability, and no growth, temperature, and morphological defects were detected [101]. Mating, sporulation, and secretion were found to be normal. Phenotypes observed by the disruption of RAS1 or RAS2 genes were not observed in the ScRheb deficient strain. A unique phenotype exhibited by this strain was an increased sensitivity to the arginine and lysine analogues – canavanine and thialysine, respectively [101]. This phenotype was confirmed to be due to the deficiency in RHEB by complementation studies involving wild-type ScRheb as well as the Rheb from S. pombe (SpRheb), which were able to rescue the defect. The increased sensitivity to the amino acid analogues appears to be due to an enhanced uptake of arginine and lysine [3- to 3.5-fold and 1.5- to 2-fold respectively). The strain specifically showed an increased uptake of the two basic amino acids amongst all others tested [101]. The hypersensitivity to canavanine appears to be due to a lack of regulation of Can1 – the permease involved in arginine uptake. Chattopadhyay and Pearce [20] recently reported that Rheb interacts with Btn2p, a yeast protein involved in the
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regulation of vacuolar H+-ATPase. The interaction with Btn2p appears to alter cellular localization of Rheb [20]. The roles of various conserved regions in the sequence were confirmed by creating mutations in the effector domain, the G1 box and the CaaX motif [101]. Effector domain mutants (Y46D and N51D) could not complement the sensitivity to canavanine and thialysine, while the wild-type Rheb could. Similarly, when the highly conserved arginine in the G1 box was mutated to glycine, the mutant protein showed reduced ability to rescue the phenotype. Farnesylation was found to be critical for ScRheb function since a change of cysteine in the CaaX box to serine rendered the protein incapable of complementing the hypersensitivity to canavanine as well as the increased arginine uptake. The connection between arginine uptake and a signal transduction pathway mediated by Rheb needs to be explored which could highlight novel functions of this protein.
4.3
Genetic study of Rheb in S. pombe uncovered the second function of Rheb-regulation of cell cycle progression.
The S. pombe homologue of Rheb (SpRheb) was simultaneously identified by Mach et al. [68] and Urano et al. [101]. Like the ScRheb, the SpRheb shares a 26% sequence identity with the human and fly Rheb proteins. The function of SpRheb in S. pombe was analyzed by creating a disruption mutant as was done previously for ScRheb [106]. Unlike the ScRheb disruption mutant which did not exhibit any growth defects, the sprheb– strain did not grow. Use of a conditional mutant showed that the inhibition of Rheb leads to the arrest of the cell cycle at the Go/G1 phase. This phenotype could be rescued by the expression of the wild-type SpRheb and human Rheb but not by the ScRheb. In another study Mach et al. [68] made a similar conditional mutant of Rheb (rhb1–) and showed that the cells arrested at the G0/G1 phase as small round cells. The rhb1– allele could be complemented by wild-type rhb1+ and rhb1Q64L (analogous to constitutively active H-Ras Q61L mutant). Interestingly rhb1– cells arrested growth with a terminal phenotype similar to that of nitrogen starved cells [25, 32, 93]. Cells depleted of nitrogen enter stationary phase, are smaller than actively growing cells and remain viable for several weeks. Nitrogen-starved cells are known to induce fnx1 mRNA, a probable proton-driven plasma membrane transporter of the multidrug resistance group [31]. Overexpression of fnx1 has also been shown to cause growth arrest like nitrogen-starved cells [31]. A mutant of Rheb – rhb1–
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D121A (a hypomorphic mutant analogous to S. cerevisiae tem1–3, a temperature-sensitive allele [8]) was used to characterize the Rheb function [68]. The involvement of SpRheb in cell cycle was implicated by studies of a strain deficient in the β subunit of farnesyltransferase encoded by cpp1+ [106]. This farnesylation-deficient strain (cpp1–) was found to exhibit enrichment of G0/G1 cells cycle as well as canavanine hypersensitivity. Expression of SpRheb–CVIL, capable of being geranylgeranylated in the cpp1– background could restore normal cell cycle profile. On the other hand, the expression of SpRheb–SVIA which is incapable of being farnesylated could not rescue this defect in both the cpp1– and rhb1– strains implying the critical role of farnesylation of this protein in maintaining a normal cell cycle. Since previously it had been shown that ScRheb regulates arginine uptake as well as canavanine sensitivity [101], a similar role was predicted for SpRheb. In agreement with this idea, the cpp1– strain exhibited hypersensitivity to canavanine. Furthermore, the increased ability to uptake arginine could be rescued by the expression of SpRheb–CVIL but not by wild-type Rheb since it is not capable of being farnesylated in that strain. These observations suggest that SpRheb has dual functions, one to regulate cell cycle and the other to regulate arginine uptake. The results again imply a crucial role of farnesylation of the Rheb protein [105]. Figure 4 shows a scheme for the function of S. cerevisiae and S. pombe Rheb.
5
SUMMARY
Yeast studies have significantly contributed to the understanding of protein farnesylation. Identification and characterization of yeast mutants defective in protein farnesyltransferase were critical for understanding the structure and function of this enzyme in higher eukaryotes. Random mutagenesis of yeast FTase β-subunit revealed the significance of three residues, S159, Y362, and Y366 in the recognition of the CaaX motif. The two tyrosine residues, perfectly conserved in all FTases so far examined, appear to be of particular interest. Precise roles of the corresponding residues in mammalian enzymes, Y361 and Y365, need to be investigated. Finally, yeast provided an assay that was used to screen natural compounds to identify inhibitors of FTase.
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Rheb
Cys
Arginine uptake
S. cerevisiae
Rheb
Arginine uptake
Cys
Cell cycle progression
S. pombe
Figure 4. Schematic drawing showing the function of Rheb in S. cerevisiae and S. pombe. Rheb is farnesylated at the C-terminal cysteine residue and localized to the plasma membrane. In S. cerevisiae, Rheb regulates arginine uptake; in S. pombe, Rheb regulates cell cycle progression as well as arginine uptake.
In addition, yeast studies have provided information on the physiological function of a variety of farnesylated proteins. In this review, we focused on a recently characterized novel member of the Ras superfamily G-proteins – Rheb. Both S. cerevisiae and S. pombe have proven to be excellent systems to study the function of Rheb. The availability of FTase-deficient mutants has further been instrumental in defining the critical role of farnesylation of Rheb. Although experiments with S. cerevisiae indicated a link between Rheb and arginine uptake, the significance of this effect on cell physiology is yet to be determined. On the other hand, experiments with S. pombe do indicate a role of Rheb in regulating cell cycle. Simultaneously, the study also revealed the critical role of farnesylation in maintaining cell cycle progression. Thus, in both these systems, farnesylation was deemed necessary for proper function of Rheb.
ACKNOWLEDGMENTS We thank Angel Tabancay and Dr. Jun Urano for discussion. This work is supported by NIH grant CA41996.
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Chapter 5 FROM BREAD TO BEDSIDE: WHAT BUDDING YEAST HAS TAUGHT US ABOUT THE IMMORTALIZATION OF CANCER CELLS Soma S.R. Banik and Christopher M. Counter Departments of Pharmacology and Cancer Biology and Radiation Oncology, Duke University Medical Center, Box 3813, Durham, NC, USA 27710
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INTRODUCTION
The budding yeast Saccharomyces cerevisiae is a formidable model system indeed. With the entire genome sequenced, unparalleled genetic malleability, and an eukaryotic background, this system is virtually beyond compare for studying the multitude of biological pathways that are conserved amongst eukaryotes. Importantly with regards to human cancer, many of the cellular processes of the mammalian cell can be found, admittedly in a stripped-down version, in yeast. In this regard, yeasts are fertile ground for elucidating mechanisms and identifying the key players in cellular processes. This invaluable information can then act as a guide for the cancer cell biologist attempting to navigate the analogous pathway in the far more complex and elaborate system of the mammalian cell. One example of where yeast has been used in this regard is in understanding how cancer cells acquire the ability to divide indefinitely. Here we highlight the enormous contributions made by studies performed in the model system of S. cerevisiae to our understanding of this tumourigenic process.
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CELLULAR IMMORTALIZATION
It has long been appreciated that normal human somatic cells adapted to grow outside the body, or in culture, divide a limited number of times [29, 123 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 123–139. © 2007 Springer.
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40]. In contrast, cancer cells have the potential to divide indefinitely, displaying immortality [6]. HeLa cells are perhaps the best example of this. First isolated in 1951 from a cervical cancer specimen, these cells have been cultured to this day in one form or another in laboratories around the world [56]. The aberrantly long lifespan unique to cancer cells in culture suggests that the immortalization process may represent a key event in tumourigenesis. Indeed, the clonal nature of tumour progression supports the notion that cancer cells undergo numerous cycles of cell division in the process of giving rise to a tumour [12, 25, 26, 79]. This basic and fundamental difference between normal cells and cancer cells suggests the tantalizing possibility that understanding the cellular mechanisms of cell immortalization might reveal novel molecular targets that might be exploited in cancer therapy. Importantly, as is the case for other therapeutic targets, much of the groundwork for understanding these processes was first established by studying the analogous system in yeast.
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TELOMERES
Remarkably, one of the key determinants of cellular immortalization is found in non-coding regions at the extreme ends of chromosomes. These terminal telomere structures are complex with a host of specialized cellular proteins to cap and protect the ends of eukaryotic chromosomes from illegitimate recombination and degradation [9]. The telomere itself ends in a 3′ single-stranded overhang that invades the double-stranded telomeric tract, forming a t-loop configuration that is thought to constitute an essential structure for chromosome protection [33]. The proteins that complex with telomeric DNA do so in an organized and structured fashion. For example, in humans, hTRF1 is known to bind to the double-stranded region [11, 85], whereas the protein hTRF2 is found to foster the formation of a triplex DNA structure where the terminal 3′ overhang is tucked into the double-stranded telomere tract [33]. Lastly, it has been suggested that the telomeric overhang may be bound by the protein hPot1 [8]. To add an additional layer of complexity to this structure, many other proteins then associate with these core telomere-binding proteins [9]. Together, the DNA and associated proteins form a functional telomere. In the absence of functional telomeres, mammalian chromosomes become highly unstable and recombinogenic, resulting in the formation of dicentric and ring chromosomes [16, 41]. Indeed, the protective nature of telomeric DNA was characterized in yeast where cloned telomeric repeats were shown to have the property of stabilizing linear DNA [74]. In fact, telomeric sequences provided a final essential structure for the formation of artificial yeast chromosomes (YACs), now an integral tool in mammalian genetics [60].
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Complete replication of telomeric DNA must contend with two hurdles. First, the most terminal primer for lagging strand replication leaves a gap following removal of the RNA primer but that cannot be filled in by any conventional DNA polymerase, resulting in telomere shortening [64, 80]. Second, leading strand synthesis forms a flush end that must be processed to produce the G-strand overhang necessary to produce the telomeric loop structure, presumably by the partial degradation of the C-strand, again resulting in a net loss of telomeric DNA [82]. Thus, another feature of telomeres must be a unique mode of replication, as continual erosion of chromosomes clearly cannot be tolerated over time, from one generation to the next. In fact, this process of telomere shortening also limits the maximum lifespan of normal human somatic cells [10, 37, 76].
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TELOMERE LENGTH DICATES THE LIFESPAN OF HUMAN CELLS
The inability to completely replicate the ends of chromosomes was first postulated to account for the finite lifespan of human somatic cells in 1971 [64]. Twenty years after this hypothesis was put forth, it was demonstrated that telomeres in normal somatic cells were far shorter than those found in germ cells, which are considered to be “biologically immortal” [15]. The actual process of telomere shortening during DNA replication was subsequently documented in cell culture and in vivo. Specifically, it was shown that telomere length decreased with increasing cell division in cultured human fibroblasts [37], or with increasing donor age in hematopoetic cells [39]. Indeed, telomere shortening with cell division or donor age has now been documented in a variety of other human somatic cells [7, 13, 22, 45, 46, 58, 73, 77, 78, 83]. In the case of cells growing in culture, it is now apparent that the proliferative capacity of the culture is directly proportional to the telomere length of the founding cells and the number of chromosome replications, as opposed to time, further supporting the model that telomere shortening dictates cell lifespan [1–3]. Although these observations made in mammalian cells were telling, it was in yeast that the most direct and compelling evidence was found to suggest that continual telomeric erosion would ultimately limit cellular lifespan. In an elegant yeast genetic screen undertaken by Lundblad and Szostak, a yeast mutant was identified that lost the ability to maintain telomere length [55]. Since this mutation caused ever shortening telomeres, the gene harbouring this mutation was named EST1. The specific importance of the EST1 protein was ascertained by the direct genetic disruption of the EST1 gene, which lead to telomere shortening and eventually a severe compromise in cellular viability. Thus, the ability to easily disrupt genes in yeast provided the first
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direct evidence for a relationship between telomere length and cellular lifespan in eukaryotic cells [55].
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TELOMERE SHORTENING IS ARRESTED IN IMMORTAL CANCER CELLS
As per the model predicting that telomere shortening limits cellular lifespan, it follows that immortal cancer cells must employ a specialized mechanism to overcome the problem associated with telomere shortening. This phenomenon of cellular immortalization and the pathways involved can be recapitulated in transformed cells growing in culture. Here, somatic cells are driven to divide beyond their normally allotted lifespan by perturbing the function of cell cycle regulatory molecules such as p53, p21, and Rb [67, 71]. As telomeres in these cells nonetheless do continue to shorten, a period of severe genomic instability ensues eventually leading to massive cell death. This cycle of instability and cell death selects for rare cells that have acquired genetic alterations that allow for indefinite proliferation. Telomere length analysis of these cells revealed that the mortal cells in the population perished with extremely short and unstable telomeres, whereas telomere shortening and gross genetic instability were suspended in the immortal cell clones [16]. This phenomenon of telomere maintenance appeared to suggest a general mechanism, as it was independent of both cell type, and the mechanism used to derail normal cellular growth regulatory mechanisms [16, 17, 47, 69, 75]. Like the immortalized cells, cancer cells have also been shown to maintain telomere length in culture, presumably reflecting a similar selection of mutations that stabilizes telomere length in vivo [6, 18]. Taken together, it was proposed that if unchecked, loss of telomeric DNA would be lethal, as already noted in yeast; however cancer cells can surmount this proliferative blockade through the illegitimate activation of a telomere maintenance mechanism [16].
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TELOMERASE
In 1985 Greider and Blackburn published a seminal paper in which they identified a biochemical activity in extracts isolated from the single-celled ciliate Tetrahymena. This activity could elongate a single-stranded telomeric oligonucleotide by the addition of telomeric repeats [31]. The discovery of this “telomere terminal transferase” or “telomerase” activity in Tetrahymena established a foundation for studying telomere biology in simple-model organisms such as ciliates and yeast. The telomerase RNA
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subunit was subsequently cloned and shown to contain a telomeric template [32], which was copied by a protease-sensitive subunit onto telomeres as DNA, suggesting that the catalytic subunit functions as a reverse transcriptase. Later, it was shown that telomerase was present in a human cell line [59], and that the enzyme was activated when cells became immortal [16]. Importantly, telomerase activity was not detectable in extracts prepared from normal somatic cells but was found in extracts prepared from actual cancer cells [18, 44, 68]. We now know that telomerase activity counteracts telomere shortening in aberrantly proliferating cancer cells and allows them to escape the “telomere barrier” to cell division. In fact, telomerase activity has been detected in thousands of tumours from every cancer type, making activation of this enzyme one of the most common changes in human cancers [68]. Understanding how telomerase is activated during cancer, and what components comprise the enzyme is critical in elucidating its role in cancer, and in developing ways to target the immortalization process in cancer cells. As mentioned, the enzyme was known to be composed of at least two subunits, an RNA and a protease-sensitive subunit. To account for the activation of telomerase activity in cancer cells, it was hypothesized that one or both of the genes encoding these subunits were transcriptionally silent in normal cells, but upregulated during tumourigenesis. The first subunit identified in humans was the RNA, whose expression was found to be ubiquitous [27]. This left the possibility that the protease-sensitive subunit(s) were normally repressed in somatic cells, but upregulated in cancers. To test this hypothesis, it was of utmost importance to clone the catalytic subunit of telomerase.
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YEAST TO THE RESCUE
The cloning of the catalytic subunit of telomerase proved to be a far more formidable task than the cloning of the RNA subunit. A number of putative protein catalytic subunits proved to be red herrings as telomerase activity could not be attenuated upon protein ablation. The difficulty proved to be related to the exceedingly low level of protein expression in most sys tems. Comprehensive gene expression studies estimate that only 1 to 2 transcripts encoding this subunit are found in cancer cells [49]. Such low levels made biochemical purification of human telomerase or transcript-based comparisons to identify the catalytic subunit transcript challenging. Such low expression levels, however, do not pose a problem for genetic-based searches in yeast. Furthermore, the identification of the yeast protein could then be used to seek out the human homologue from human genome or
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expressed sequence tag sequence databases, or as a probe to screen libraries for homologues from progressively more complex eukaryotes. Four features of the budding yeast cerevisiae made it an ideal system to search for the catalytic subunit. First, sequencing of the yeast genome was under full steam at the time that the catalytic subunit was cloned, providing a seamless union between genetic screens and gene identification. Second, an assay had been developed to detect telomerase activity in cerevisiae, which would be essential to test if a candidate gene actually encoded the catalytic subunit of telomerase [14]. Third, the unparalleled genetic malleability of cerevisiae, allowed genetic screens to be done that would be impossible in other organisms. Fourth, the cloning of the cerevisiae telomerase RNA subunit allowed for the characterization a phenotype associated with a loss of telomerase activity. Loss of this RNA-encoding gene, termed TLC1, gave rise to telomere shortening and a decrease in cell viability, a phenotype easily scored [72]. All of these features proved to be invaluable in the cloning of the catalytic subunit of telomerase.
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GENETIC AND BIOCHEMICAL APPROACHES IDENTIFY THE YEAST CATALYTIC SUBUNIT OF TELOMERASE
To definitively clone the catalytic subunit of telomerase, two parallel approaches were employed. In both experimental schemes, the use of yeast as a model system was paramount to the identification of this elusive protein. One of these experimental paradigms involved the initial biochemical purification of telomerase from the ciliate Euplotes crassus – yet another simple model organism specifically used for its high level of telomerase activity. Here, the telomerase complex was purified using an oligonucleotide complementary to the telomerase RNA template [51]. Intriguingly, upon microsequence analysis, one of the proteins purified through this approach proved to show sequence homology with the Est2 protein [53] previously shown to lie in the telomere maintenance pathway in S. cerevisiae [50]. This suggested that the Euplotes protein p123 as well as Est2 may in fact function as the catalytic protein of the telomerase enzyme [53]. The second approach used to clone the catalytic subunit of telomerase was based completely on a genetic screen performed in yeast. This screen was designed in consideration of the two mechanisms used by yeast to maintain telomeres, namely telomerase-mediated addition of telomeric repeats via copying of a telomeric template, and the RAD52-mediated recombination of telomeric elements. Importantly, the genetic ablation of both of these
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telomere-maintaining pathways was known to induce yeast to undergo senescence and cell death [54, 72]. This known phenomenon was exploited in order to reveal the catalytic subunit of telomerase. As such, yeast as were randomly mutated in hopes of disrupting the gene encoding the telomerase catalytic subunit. The mutant yeasts were then induced to spontaneously extrude the RAD52 gene involved in telomere recombination. Those yeast as that did not survive in the absence of this second telomere-maintaining gene, were presumed to harbour a telomere defect, and thus were screened for the presence of short telomeres and an absence of the in vitro primer-elongating activity characteristic of the telomerase enzyme. Two populations of mutant yeasts were found to satisfy the telomerase-defining criteria outlined above, thus identifying genes-encoding components of the core telomerase enzyme. The first of these genes was TLC1, encoding the previously identified telomerase RNA. The second was the gene encoding the Est2 protein [20]. Thus, through an entirely independent, experimental approach, EST2 was predicted to encode the protein catalytic subunit of telomerase. To prove that the Est2 protein was the catalytic subunit of telomerase, both studies relied heavily upon genetic manipulations that were easily accomplished in yeast, but terribly difficult in other eukaryotes. As EST2 was shown to contain weak sequence homology with reverse transcriptases, invariant amino acids in the putative catalytic core of Est2p were specifically mutated to decipher possible telomeric effects [20, 53]. Indeed, mutations of these conserved amino acids resulted in senescence, telomere shortening, and importantly, loss of telomerase catalytic activity. These experiments conclusively demonstrated that Est2p was not only essential for telomerase activity, but actually encoded the catalytic subunit of telomerase.
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TELOMERASE IS ESSENTIAL FOR IMMORTALIZATION AND TUMOURIGENESIS
Having first identified the telomerase catalytic subunit from model eukaryotic organisms, the cloning of human TERT followed almost immediately. As is the case for numerous proteins that show sequence homology between lower eukaryotes and vertebrates, telomerase subunits from Euplotes and yeast were successfully used to identify the human hTERT homologue through databases searches of human gene sequences [38, 43, 57, 61]. Clearly, in the absence of such information about the yeast and ciliate proteins, the cloning of hTERT would have been an incredibly tedious endeavour, slowed by the inability to make rapid genetic manipulations in mammalian cells. Instead, once again, the characterization of yeast and ciliate proteins was able to catapult cancer researchers into the realm of elucidating uncharted cancer processes in humans. Accordingly,
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upon the cloning of hTERT, the pace of research in cancer and cellular immortalization hastened dramatically. First, it was shown that the hTERT mRNA was indeed upregulated during the immortalization process in cultured cells, and present in cancer cells but generally not in normal somatic cells [43, 48, 57, 61]. More importantly, ectopic expression of the hTERT cDNA in telomerase-negative mortal cells was shown to restore telomerase activity, arrest telomere shortening and immortalize cells, resembling all of the characteristics of telomerase function that had already been delineated in yeast [10, 19, 38, 62, 76, 81]. The ability to immortalize normal human cells has now given rise to a plethora of opportunities with respect to both research and therapeutics. For example, isogenic normal cells from patients can now be immortalized to provide isogenic controls for cancer cell lines isolated from tumours from the same patient, thus representing an invaluable resource for assessing a normal baseline in studies of drug action and gene activation [28]. In addition, the ability to immortalize normal cells has widespread applications beyond the realm of cancer research. In the case of often rare genetic disorders, fibroblasts can be isolated and stably immortalized to provide an unlimited supply of cells that can be used for disease characterization and to develop possible therapeutic interventions [65]. With respect to tissue engineering, activation of telomerase in normal cells may prove to be important for increasing the proliferative lifespan necessary for making genetic manipulations in vitro [70]. While the role of telomerase had been documented in the process of cell immortalization, its function in tumourigenesis was conclusively established in the first study that converted normal cells into cancer cells through the introduction of defined genetic elements. Human kidney or mammary epithelial, fibroblast, or astrocyte cells were made to be tumourigenic upon infection with the SV40 early region to derail cell cycle checkpoints, oncogenic ras to provide proliferative signals, and hTERT to allow for cellular immortalization [23, 35, 66]. These cells clearly showed hallmarks of cancer, namely, having the capability to form colonies in a semi-solid medium and giving rise to tumours upon injection into nude mice; however, these phenotypes were markedly absent in cells not infected with hTERT. Thus, the link between the processes of hTERT expression, cellular immortalization, and tumourigenesis was firmly established.
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CHARACTERIZATION OF YEAST TELOMERASE PROVIDES THE FIRST METHOD TO INHIBIT THE HUMAN ENZYME IN CANCER CELLS
Telomerase is activated in approximately 85% of human cancers [68] in a wide variety of tumour types where its function is critical for cellular immortalization. In contrast, most normal cells lack telomerase activity. Thus, it stands to reason that the inhibition of telomerase could represent a novel and selective strategy of killing tumour cells via induction of telomere shortening and cellular crisis, whilst having minimal detrimental effects on normal tissue. To test this possibility, researchers were again guided by experiments done first in yeast. Specifically, expression of a catalytically inactive Est2p protein in otherwise normal telomerase-positive yeast, led to the shortening of telomeres. This phenotype suggested that the catalytically dead protein could act as a dominant-negative by sequestering factors away from the active enzyme [53]. To test this, the same mutation that inactivated telomerase catalytic activity in yeast was made in hTERT, and was found to abrogate the catalytic activity of the enzyme [62, 81]. Ectopic expression of this mutant did in fact have a dominant-negative effect in telomerasepositive transformed or tumour cell lines, where it reduced or even abolished measurable telomerase activity. Successful expression of the dominantnegative in human cells also led to telomere shortening and even cell death [30, 36, 84]. Notably, when injected into nude mice, cancer cells expressing this dominant-negative hTERT, did not produce tumours, or formed only small tumours that regressed [34, 36]. Importantly, telomerase-negative cells were unaffected by the expression of this dominant-negative protein [36]. These experiments, which find their origins in yeast, have now spawned efforts to generate small molecule inhibitors of telomerase. In an effort to recapitulate the phenotype of the dominant-negative version of hTERT, several groups have attempted to inhibit telomerase enzyme activity through a variety of experimental approaches. Perhaps the most promising of these has been the use of a small-molecule telomerase inhibitor [21]. Administration of this inhibitor to cancer cells in culture led to telomere shortening and cellular senescence. Furthermore, drug-treated cells also had a reduced tumourigenic potential when injected into nude mice. The pursuit of telomerase inhibitors is an excellent example of how a phenotype first documented in yeast has laid the foundation for the development of compounds that might be clinically useful in the treatment of human disease.
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BACK TO BASICS
Having identified the core components of human telomerase and the basic mechanism through which telomerase functions, we are now poised to dissect telomerase function and the mechanisms governing cancer cell immortalization in more precise detail. In particular, to further understand telomerase function, it will be necessary to elucidate the mechanism by which telomerase-dependent elongation of telomeres occurs in immortal human cells. Once again, studies done in yeast are leading the way. Much work has been accomplished in yeast towards understanding one mode of telomerase-dependent elongation of telomeres; namely to regulation of telomerase access to telomeres. Specifically, several of the proteins now known to be involved in telomere-targetting were in fact discovered in initial genetic screens that identified the proteins Est1, Est2, Est3, and Cdc13 which function in the telomerase pathway [50, 55]. While Est1p has been shown to associate with telomerase, it does not appear to directly target telomerase to the telomere end. Cdc13p on the other hand, binds the extreme end of the telomere, having high affinity for single-stranded telomeric DNA [63]. This implies that telomerase may be recruited to telomere ends through these two intermediate proteins. This model was further substantiated in experiments where the fusion of Est1 and Cdc13 led to greatly elongated telomeres [24], and fusion of the telomerase enzyme to Cdc13 was able to bypass the requirement for a functional Est1 protein within the cell [24]. As the mechanism of telomerase recruitment may also be conserved between yeast and humans, considerable effort is now being aimed at understanding this process in human cells. Support for the existence of this mode of telomerase regulation in humans has come from telomerase fusion experiments done in human cells. Specifically, we identified a mutant of hTERT that retained telomerase activity yet was unable to elongate telomeres [4] – a phenotype reminiscent of yeast mutants defective in telomerase recruitment [24, 42, 52]. To test if this mutant of hTERT was indeed defective in telomere targetting, it was fused to the telomere-binding protein TRF2. While the mutant hTERT protein alone was unable to elongate telomeres or immortalize cells, the function of the mutant protein was rescued upon artificially targetting it to telomeres via the TRF2 protein [5], indicating that human telomerase is regulated by its access to telomeres. Presumably such regulation is mediated by protein–protein interactions.
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In this regard we note that the human protein hPot1 [8] may be a functional homologue of the yeast protein Cdc13, which is involved in the recruitment of telomerase to telomeres. While a possible telomerase recruitment function for Pot1 analogous to that of Cdc13 remains speculative at this time, future studies guided by the yeast literature will certainly help in defining this function.
12
CONCLUSION
Cellular immortalization is one of the fundamental and defining characteristics of cancer. In the majority of human cancers, immortalization is also conferred by the telomere-maintaining activities of the enzyme telomerase. As the telomerase pathway is highly conserved amongst almost all eukaryotic organisms, many studies that have elucidated the function of human telomerase have relied heavily on studies previously done in yeast. As yeast constitute an excellent model system highly amenable to elucidating genes responsible for a variety of cellular functions, many proteins in the telomerase pathway including the catalytic subunit of telomerase were first identified in yeast. Indeed, many proteins in the telomerase pathway, including the catalytic subunit of telomerase, were first identified in yeast. As new functions of telomerase are characterized and new members of the telomerase pathway are identified, the yeast telomerase pathway will certainly continue to be an invaluable system to aid in elucidating the contribution of telomerase to cellular immortalization and tumourigenesis.
ACKNOWLEDGMENTS We would like to thank Dr. Raymund Wellinger for reviewing this manuscript, and providing helpful comments.
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Chapter 6 HSP90 CO-CHAPERONES IN SACCHAROMYCES CEREVISIAE
Marija Tesic and Richard F. Gaber Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, Evanston, Illinois 60208
Heat-shock response is a general mechanism through which cells cope with external, as well as internal, stresses. Although the heat-shock response is elicited under conditions of stress, proteins involved in this set of pathways are also necessary for basic maintenance of cellular constituents, which involves regulation of proper levels, folded states, and conformations of proteins. Heat shock proteins play key roles in ensuring the fine-tuning and integrity of the signal transduction pathways, including those involved in cellular proliferation and cell cycle regulation. Understanding the function of the heat shock proteins is, therefore, central to our knowledge of proper cell regulation and how it might go awry; it is also essential for developing productive interventions to prevent the loss of this regulation. Hsp90 is an essential, abundant, and highly conserved heat shock protein. It interacts with a large number and variety of wild type and mutant proteins that are partially folded, or adopt unstable or inactive conformations. Because of its ability to buffer the morphologic variability within the cell and allow mutant or partially folded proteins to perform wild-type functions Hsp90 is considered a molecular “evolutionary capacitor” [142]. Hsp90 is also a global signal transduction regulator that can modulate responses of numerous signaling molecules to intra- and extracellular stimuli. A subset of Hsp90 substrate proteins are involved in the regulation of cellular proliferation and transformation. Deeper understanding of the regulation of these substrates by Hsp90 will further our knowledge of tumorigenesis in general, and thus lead to approaches through which it can be prevented. Hsp90 is a molecular chaperone that performs its roles in concert with other proteins, collectively referred to as Hsp90 co-chaperones. Most mammalian Hsp90 co-chaperones are conserved in Saccharomyces 141 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 141–177. © 2007 Springer.
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cerevisiae (see Table 1). On the one hand, this facilitates (the) analysis of Hsp90 co-chaperones in vivo with a facile and genetically tractable system. On the other hand, the absence of certain mammalian homologs of the Hsp90 complex in yeast affords an opportunity to use yeast as a “bag of enzymes” to study in vivo precisely those components that are missing. Table 1. Conservation of Hsp90 co-chaperones in mammalian and S. cerevisiae cells. The third column lists relevant structural domains of the listed co-chaperones. The fourth column lists selected Hsp90 substrates known to interact with either mammalian or S. cerevisiae cochaperones. Applicable references are provided in the text. Mammalian Hsp90α, Hsp90β
S. cerevisiae Hsp82, Hsc82
Domains ATPase
Hsp70
Ssal-4
ATPase
Hsp40/Hdj1, Hdj2
Ydj1, Sis1
J domain, Zn
Substrates N/A N/A 2+
GR, v-Src, Hap1
finger Hop/p60
Sti1
TPR
GR, PR, v-Src, HSF,
Hip/p48
___
TPR, GGMP repeat
GR, PR
p23
Sbal1
___
GR, PR, AR, MR,
Gen2, p53
ER, TR, PKR, telomerase, p53, HSF, Gen2, AhR CyP40
Cpr6, Cpr7
TPR, PPIase
GR, v-Src, Hfl1, p53
FKBP51, FKBP52
___
TPR- PPIase
GR, PR
PP5
Ppt1
TPR, phosphatase
GR, p53
p50/Cdc37
Cdc37
Kinase binding
v-Src, Ste11, Raf1, Cdk4, Cdk6, AR, Gen2
ARA9/XAP2/AIP
___
TPR, FKBP domain
AhR
TTC4
Cns1
TPR
GR, Hsf1
AKT1,2,3, kinases?
Sch9
S/T kinase, C2
GR, Hap1, Ste11
domain Hsp110
Sse1
Calmodulin binding
GR, HSF
Bag1
?
Ub-like, Hsp 70-
GR, p53
binding CHIP
___
TPR, U-Box
GR, CFTR
?
Hch1
___
___
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The high degree of conservation of most Hsp90 complex components suggests that the mechanisms of function of Hsp90 are conserved. Since Hsp90 is capable of playing multiple roles in the regulation of its substrates, the co-chaperones might aid Hsp90 at a number of points of regulation, which may differ considerably among different substrates. Additionally, some of the co-chaperones may have regulatory functions in their own right, independent of their roles in the Hsp90 complex. This discussion is focused on the role that Hsp90 co-chaperones play in helping Hsp90 to control proper expression levels, activity, and localization of client proteins. Another essential and highly conserved heat shock protein, Hsp70, plays a prominent role in the regulation of some Hsp90 substrates, but the discussion of this chaperone and its co-chaperones is beyond the scope of this article (for review, see [104]).
1
HSP90 AS A FINELY TUNED CHAPERONE MACHINE
Hsp90 is a highly conserved molecular chaperone. Eukaryotic Hsp90 is essential (for recent reviews, see [123, 181]) and abundant, constituting up to 1–2% total cellular protein [75]. Although the number of proteins with which it is known to interact is large and still growing, Hsp90 does not interact with substrates and co-chaperones indiscriminately, but apparently in a highly specific and regulated manner. It is of interest to consider the question of how one molecule can provide at the same time such a wide scope of productive interactions, and the specificity required for control. While part of the answer is provided by the versatile nature of the Hsp90 molecule itself, the full answer to this question requires understanding of the interacting partners of Hsp90, its co-chaperones. Hsp90 consists of three domains: the amino terminal domain, the middle region, and the carboxy terminal domain. The amino terminus contains an ATPase domain of about 23 kDa; in yeast, it is approximately 210 amino acids in length [145]. Hsp90 homologs in S. cerevisiae have a considerably higher ATPase activity [0.1–0.2 1/min) [122, 135] than human Hsp90 (0.02 1/min [119]), but the biological significance of this difference is still unclear. Although the ATPase activity of Hsp90 as compared with other ATPases is low, it is essential for its in vivo functions [117, 122]. Geldanamycin (GA), a potential antitumor agent that mimicks the structure of ATP, inhibits Hsp90 ATPase activity by blocking the ATP-binding pocket [134]. In S. cerevisiae, the middle region of Hsp90 extends approximately from residues 210 to 272. It enhances the affinity of the amino terminus for unfolded substrates [145], but is dispensable for viability [94].
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The carboxy terminus of S. cerevisiae Hsp90 binds some of the Hsp90 substrates, for example steroid receptors [145]. The carboxy terminus also binds those co-chaperones that contain the tetratricopeptide (TPR) domain, including Hop, CyP40, FKBP51 and FKBP52, and PP5 (see Table 1). TPRs are structural motifs involved in protein–protein interactions of a variety of proteins (for review, see [9]). The principal interaction between cochaperones containing TPR domains and Hsp90 is mediated by the carboxyterminal sequence of Hsp90, MEEVD. The carboxy terminus may also be the location of a second, cryptic ATP-binding site [96], that is exposed upon binding of the nucleotide at the N-terminus [159]. Hsp90 exists in solution as a dimer, and it is the dimeric form that binds substrates and co-chaperones. Although the primary, stable dimerization is mediated by the carboxy terminus [107], both amino and carboxy termini of Hsp90 are involved in dimerization. The amino terminus mediates transient dimerization of Hsp90 in response to ATP binding [20, 133, 172]. Dimerization may not be required for interaction with substrates, or viability. A short deletion in the carboxy terminus of chicken Hsp90 leads to loss of dimerization, but has no effect on the interaction of the chaperone with estrogen receptor in vivo; additionally, the mutant chicken protein confers viability to S. cerevisiae cells lacking endogenous Hsp90 [101]. Although both amino and carboxy termini of Hsp90 can bind substrates and ATP in vitro [145, 182], suggesting that the domains might function independently, work from several laboratories reveals that there is a finely tuned communication between the two termini of the molecule that enable it to undergo activation cycles with its clients and co-chaperones [20, 133, 140, 146, 159, 172]. Through conformational changes involving the entire Hsp90 molecule, the ATPase activity of the amino terminus is tightly coupled to the chaperoning of substrates, which is most likely performed by the carboxy terminus.
1.1
Hsp90 in S. cerevisiae
In budding yeast, two genes, HSP82 and HSC82, encode Hsp90 proteins that are 97% identical. HSP82 was initially identified using differential plaque filter hybridization performed to find genes expressed specifically in heat shocked cells [54]. HSC82 was subsequently identified by hybridization of HSP82 to yeast genomic library [12]. Cells harboring deletions of either HSP82 or HSC82 are viable, but the double deletion is lethal [12]. Under normal conditions, Hsc82 is present at approximately 20-fold greater levels than Hsp82, and is induced only about twofold upon heat shock. In contrast, when cells are heat shocked, HSP82 is induced some 20fold, to a level approximately the same as Hsc82 levels [12]. Deletions of
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HSC82 or HSP82 do not exhibit dosage compensation and have no discernible effects on the expression of other genes [12]. Cells in which either gene is deleted exhibit weak temperature sensitivity (ts) at 37°C. Underscoring the abundance of Hsp90, cells remain viable even when its levels are decreased to 10% of wild type, although this decrease leads to partial loss of some Hsp90 functions. Overexpression of Hsp90 does not offer increased protection from heat shock, nor does it enhance thermotolerance [30]. A number of mutant alleles in HSP82 and HSC82 have been generated [11, 84, 113] and have proven useful in studying the function of this protein, especially in investigating the effects of loss of Hsp90 function on the client proteins in vivo.
2
HSP90 SUBSTRATES IN S. CEREVISIAE
Most Hsp90 substrates identified to date are signaling molecules and fall largely into two groups: transcription factors and protein kinases (for review, see [130]), although there is a growing list of substrates not involved in signal transduction. Common to several Hsp90 substrates is their propensity to adopt multiple conformations. The requirement for the Hsp90 complex arises from the need to chaperone each client into its proper conformation, in which the molecule is poised to receive a signal and perform its cellular role. Table 2 outlines heterologous Hsp90 substrates that have been expressed in budding yeast cells and known endogenous substrates. In this section, selected substrates are treated in detail according to the following criteria: historical importance and widespread use in the studies of Hsp90 (steroid receptors), exemplification of regulation by a co-chaperone (aryl hydrocarbon receptor), and involvement in cellular proliferation and tumorigenesis (p53, Ste11, and telomerase).
2.1
Heterologous substrates
Some of mammalian Hsp90 client proteins that are not conserved in S. cerevisiae, but were known from biochemical studies to bind the Hsp90 machinery, have been expressed in S. cerevisiae cells in order to study their in vivo regulation by the Hsp90 complex. 2.1.1
Steroid receptors
When rat glucocorticoid receptor (GR) is expressed in S. cerevisiae, it is capable of enhancing transcription driven by a glucocorticoid response element (GRE) in a ligand-dependent fashion [147]. Genetically decreasing
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yeast Hsp90 levels to about 5% of wild type leads to reduction in GR signaling [126]. This was the first evidence that Hsp90 plays a functional role in steroid receptor complexes with which it was known to physically interact. Maturation of steroid receptors in yeast has since become a commonly used assay for general Hsp90 function. For example, a number of Hsp90 mutants have been isolated in a screen for decreased GR signaling [11]. To date, several steroid receptors have been expressed in yeast (see Table 6.2) [11, 53]. Although a functional dependence of androgen receptor (AR) on Hsp90 was not demonstrable using in vitro approaches [115, 118], expression of AR in S. cerevisiae cells carrying mutant Hsp90 allowed demonstration of the requirement for a functional Hsp90 complex [53]. Table 2. Hsp90 substrates that have been expressed or identified in S. cerevisiae. The right column lists known S. cerevisiae co-chaperones that have been experimentally demonstrated to interact with the listed substrates either physically, genetically, or in functional assays. The applicable references are given in the text.
Endogenous substrates
2.1.2
Progesterone receptor (PR) Estrogen receptor (ER) Mineralocorticoid receptor (MR) Androgen receptor (AR) Aryl hydrocarbon (dioxin) receptor (AhR) Retinoic acid receptor (RXR) v-Src c-Src p53 PKR Heat Shock Factor (Hsf1) Hap1 Ste11 Gcn2 Cna2
S. cerevisiae co-chaperones interacting with substrates Ydj1, Sti1, Sba1, Cpr7, Cns1, Sch9, Sse1 Sti1, Sba1 Sba1 Sba1 Sba1, Cdc37 ⎯ ⎯ Ydj1, Sti1, Cpr7, Cdc37 ⎯ ⎯ Sba1, Cdc37 Sti1, Cpr7, Cns1, Sch9, Sse1 Ydj1, Sch9 Cdc37, Sch9 Sti1, Sba1, Cdc37 ⎯
Telomerase
⎯
Steroid receptors
Heterologous substrates
Hsp90 substrates in S. cerevisiae Glucocorticoid receptor (GR)
Aryl hydrocarbon receptor (AhR)
AhR is a nuclear receptor that, like steroid receptors, binds its ligands (mostly planar aromatic molecules such as dioxin) inside the cell. Upon ligand binding, AhR heterodimerizes with ARNT (AhR nuclear translocator). Both AhR and ARNT contain bHLH domain for DNA binding. AhR was initially expressed in yeast cells as a LexA fusion in order
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to eliminate the need for ARNT coexpression, demonstrating for the first time the need for Hsp90 in the signaling of this nuclear receptor [18]. Another fusion, with the DNA-binding domain of human GR to eliminate the need for the dioxin-responsive elements, has also been expressed in yeast [175]. In order to more faithfully replicate the mammalian system, AhR and ARNT were coexpressed in yeast cells [106] where they exhibited dependence on components of the Hsp90 complex [105]. An AhR co-chaperone, ARA9/XAP2/AIP, was identified in a yeast twohybrid screen for proteins that interact with AhR [17]. Its expression in S. cerevisiae, which does not harbor a homolog, enhanced the ligand responsiveness of AhR [17]. It is evident from the work of several laboratories that ARA9/XAP2/AIP determines the specificity of the Hsp90 complex for AhR [6, 19, 82, 83, 90, 102, 103, 125]. Several important lessons can be learned from the studies of ARA9/ XAP2/AIP that might also apply to other Hsp90 co-chaperones. First, ARA9/XAP2/AIP interacts directly both with Hsp90 and AhR [102], and the interaction with AhR is mediated by the TPR domain of ARA9/XAP2/AIP, a domain that is primarily implicated in binding to Hsp90. This suggests a possibility that the TPR domains of other TPR-containing Hsp90 cochaperones direct the interaction between the co-chaperones and various substrates, in addition to mediating the interaction between co-chaperones and Hsp90. Second, ARA9/XAP2/AIP regulates AhR at least in part by increasing the proportion of the receptor in the cytoplasm [82, 90, 125]. Other co-chaperones may also play roles in the proper localization of Hsp90 substrates, possibly even bringing together the substrates and the Hsp90 complex. Third, it has been shown that ARA9/XAP2/AIP protects AhR against proteasome-dependent degradation [82]. ARA9/XAP2/AIP may have a chaperone activity on its own, possibly mediated by the FKBP-like immunophilin domain situated in the amino terminus of the protein. Several co-chaperones (p23, CyP40, FKBPs) have in vitro chaperoning activities [13, 58], which may be important in the maintenance of specific Hsp90 substrates in proper folded states. 2.1.3
v-Src and c-Src
It has been known for some time that the oncogenic protein kinase v-Src can be co-immunoprecipitated with Hsp90 [14]. Expression of v-Src in S. cerevisiae cells is toxic, presumably due to nonspecific phosphorylation of yeast proteins [15, 88]. Hsp90 maintains normal levels and specificity of the kinase [177]. It was initially thought that only v-Src, but not c-Src depends on the presence of the functional Hsp90 complex. However, use of a particularly stringent Hsp90 mutant in S. cerevisiae revealed that c-Src also
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requires Hsp90 for full activity [178]. The regulation of v-Src by Hsp90 complex reveals aspects of both positive and negative control: the first step, in which Hsp90 helps the kinase adopt the proper conformation, is followed by a second step in which Hsp90 prevents the kinase from being activated. The intrinsic structural differences between the c-Src and v-Src presumably contribute to the varying extent of dependence on Hsp90 [178]. The comparison between the extent of dependence of the two kinases on Hsp90 could provide useful insights into the nature of the Hsp90 substrate recognition. 2.1.4
p53
p53 is a transcription factor with a well-studied role as a tumor suppressor. In rabbit reticulocyte lysates, a mutant form of the p53 protein (p53V143A) interacts with Hsp90 whereas the wild type does not [8]. Addition of Hsp90 inhibitors, geldanamycin (GA) and macbecin I, impaired the ability of Hsp90 to interact with several p53 mutant forms [8, 176]. Expression of wild type and mutant p53 in S. cerevisiae showed that the mutant Hsp90G313N is unable to bind mutant p53V143A, demonstrating authenticity of the interaction [8]. In contrast, purified Hsp90 interacts with the purified wild type p53, but not a mutant, p53R175H [87]. A co-chaperone, Bag-1, is capable of dissociating wild-type p53–Hsp90 complexes in vitro [87]. Overall, the fact that p53 (both in wild type and mutant forms) interacts with the components of the Hsp90 complex provides a direct connection between tumorigenic signaling in cells and chaperone complexes that might regulate this signaling cascade.
2.2
Endogenous substrates
Relatively few endogenous yeast substrates of Hsp90 are known. Most of them were initially discovered through their similarity to the binding partners of Hsp90 in mammalian cells. 2.2.1
Ste11
Ste11 occupies a role in MAPK pathways analogous to that of the mammalian Raf serine/threonine kinase. Several groups have shown that Raf forms GA-sensitive complexes with Hsp90 [60, 149, 150, 160, 161, 171]. One of the signaling pathways involving Ste11 is the yeast pheromone response, a MAPK cascade activated by the binding of a mating pheromone to a cell surface receptor, leading ultimately to cell cycle arrest. S. cerevisiae cells in which Hsp90 is mutated are unable to undergo arrest in
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the presence of a mating pheromone [93]. In Hsp90 mutant cells, both the accumulation of Ste11 and its activity are decreased. A screen for multicopy suppressors of this phenotype led to the isolation of Ste5, a component of the MAPK-signaling module that functions as a scaffold to maintain association of the other components of the cascade (Ste11, Ste7, Fus3/Kss1) [93]. It is possible that the specificity of Hsp90 interaction with Ste11 is determined by the Hsp90 co-chaperone Cdc37. Cdc37 has been dubbed a “kinase-targeting subunit” of the Hsp90 complex [165], and cdc37 mutant cells have reduced Ste11 function [2]. Cdc37, Ste11, and Hsp90 form a heterocomplex in yeast cells [2]. Since such complexes have also been identified in mammalian cells [60, 160], there appears to be strong conservation in the way components of the MAPK cascade are regulated by the Hsp90 complex, thus allowing researchers to extend studies of yeast cell cycle to mammalian systems. 2.2.2
Telomerase
Hsp90 was identified as a multicopy suppressor of mutant Stn1 [61], a protein that interacts with a telomerase regulator Cdc13. Overexpression of Hsp90 leads to telomere shortening in both stn1 mutant and wild-type cells, but genetically decreasing Hsp90 levels to 5% wild-type has no effect on telomere length [61]. Interestingly, hTERT, a human catalytic subunit of telomerase, was identified in a yeast two-hybrid screen for proteins that interact with a Hsp90 co-chaperone, p23 [70]. Both Hsp90 and p23 enhance telomerase activity in vitro and in cell culture. Future studies should reveal whether the regulation of telomerase by the Hsp90 complex resembles that of other client proteins. It has been proposed that telomerase expression is necessary for the immortalization of cancer cells [35, 65]. Therefore, inhibition of Hsp90 function might lead to the reversal of cellular transformation, at least partly, through the telomerase-based mechanism and contribute to the antitumorigenic effect of Hsp90 inhibitor drugs, such as GA.
3
HSP90 CO-CHAPERONES IN BUDDING YEAST
Mechanism of Hsp90 function has primarily been studied using in vitro approaches, especially reconstitution of the assembly of Hsp90 heterocomplexes, where progesterone receptor (PR) or glucocorticoid receptor (GR) served as model substrates (for review, see [31] and [131]). Other studies have, in one way or another, corroborated the model of overall Hsp90 function that has emerged from the in vitro reconstitution
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experiments. The picture of Hsp90 that arises from these studies is that of a highly dynamic molecule that joins and departs from complexes containing its substrates and co-chaperones (Figure 1). During the course of activation by the Hsp90 complex, the substrate is transformed from an inactive and possibly partially unfolded state, to a conformation in which it is poised to respond to appropriate cellular signals.
Figure 1. Sequence of events leading to the activation of a model substrate by Hsp90 and Hsp70 and their co-chaperones. Although steps in this sequence have been elucidated using mammalian cells, protein names indicated here are those of S. cerevisiae (co-chaperones). Details of dynamic interactions between the proteins are described in the text.
Steroid receptors have been used as model substrates that are chaperoned by Hsp90 from a conformation incapable of binding the hormone ligand to a hormone-binding form [31, 131]. A highly conserved and essential chaperone, Hsp70, is thought to be the first to bind the substrate, thus defining the early phase of the steroid receptor maturation cycle [74, 132, 155]. A recent report suggests, however, that Hsp40/Hdj1, a co-chaperone of Hsp70, may, in fact, be the first to bind, at least to PR [66]. Another Hsp70 regulator, Hip, is also associated with progesterone receptors [129], and might aid the formation of this early complex by stabilizing the interaction between Hsp70 and the substrate [59]. Regardless of their relative order, it is by now well established that Hsp70 and its co-chaperones Hip and Hsp40/Hdj1 participate in the early stages of substrate activation. Hsp90 is the next component to enter the complex. Its initial binding may be mediated by the co-chaperone, Hop, which is capable of binding both Hsp90 and Hsp70 concurrently [27, 28, 43, 76, 156]. The presence of Hop defines the intermediate complex. Hop is replaced by another set of co-chaperones, which bind to the same acceptor site on Hsp90 as Hop. These are most often
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large immunophilins of the cyclophilin (CyP40) or FKBP class (FKBP51 and FKBP52) [68, 120, 138, 143, 144, 168]. The late or mature complex is defined by the presence of immunophilins and is thought to be stabilized by another co-chaperone, p23 [42, 79, 157]. In the mature complex, the steroid receptor has been activated and is, therefore, capable of binding ligand. Most likely, the Hsp90 machinery disassembles upon ligand binding [132]. At this point, the continued interaction of client proteins with the Hsp90 machinery prevents them from being activated without proper signaling. Therefore, Hsp90, and the proteins that assist it, can regulate their targets both positively and negatively. This dual role of the Hsp90 complex in substrate control is an important aspect of its cellular function. To understand the involvement of Hsp90 co-chaperones in the regulation of Hsp90 substrates, it will be necessary to delineate different points of the Hsp90 cycle at which co-chaperones are thought to play a role. The exact make up of the Hsp90 machinery that is necessary and sufficient for the transformation of steroid receptors is still unclear. For example, while purified Hsp70, Hsp90, and Hop can transform GR, p23 is still necessary for the stabilization of complexes [42]. Additionally, it has been reported that, under some conditions, Hsp90 and Hsp70 alone can mediate steroid receptor maturation [136]. However, the in vivo mechanism is likely to be far more complex and the yeast system has been of great use in addressing the question of the requirement for the various co-chaperones in Hsp90 functions in the cell.
3.1
TRP-containing co-chaperones
3.1.1
Hop/Sti1
The homolog of mammalian Hop, STI1 was first identified in S. cerevisiae as a stress-inducible mRNA [116]. sti1∆ cells are viable at 30°C, but exhibit a slow-growth phenotype at both higher and lower temperatures. In mammalian cells, Hop has been established as a common co-chaperone for Hsp90 and Hsp70 that brings the two chaperones together and thus facilitates formation of the intermediate complexes in the steroid receptor maturation cycle [27, 28, 43, 76, 156]. The structures of human Hop TPR domains corroborate this view [148]. Hop/Sti1 has nine TPRs. The three amino terminal TPRs bind to Hsp70. Of the other six TPRs, the three central repeats are sufficient to bind Hsp90. The crystal structures of two TPR domains of Hop bound to their appropriate Hsp90 and Hsp70 peptides explain how Hop might serve as a bridge between Hsp90 and Hsp70. It is of interest that the carboxy-terminal motif of Hsp90, MEEVD, binds to the TPR
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domain of Hop with a very similar dissociation constant (Kd = 11 µM) to the entire C-terminal domain of Hsp90 (Kd = 6 µM). This indicates that the main (and possibly sole) structural determinants for binding to TPR domains lie within this small region of Hsp90. Reconstitution of steroid receptor complexes in reticulocyte lysates and formation of complexes from purified components defined the minimal chaperone machinery required to transform steroid receptors into an active state: Hsp90, Hsp70, Hop, Hsp40, and p23 [130, 131]. Because of the enhancement in the rate of steroid receptor activation in the presence of Hop [109], it can be concluded that this co-chaperone at least enhances the assembly of the complex and its activity on substrates, although there may not be an absolute requirement for Hop in Hsp90 complexes. Combined genetic and biochemical data from a number of laboratories suggest that Hop/Sti1 helps Hsp90 in the formation of productive complexes i.e., those capable of chaperoning substrates. In sti1∆ cells, signaling through eIF2α kinase Gcn2 is decreased [46]. Deletion of STI1 in S. cerevisiae leads to at least a threefold decrease in the activation of a GRE-LacZ reporter gene, and significant loss of v-Src activity [21]. While sti1∆ affects only the activity and not the level of v-Src protein, mutations in hsp90 decrease both. It is therefore possible that Sti1 is involved specifically with one aspect of vSrc chaperoning by Hsp90, aimed at maintaining its activity. However, a closer look at the Hsp90 and Sti1 complexes in S. cerevisiae reveals a more puzzling picture. When Hsp90 and GR complexes are precipitated from wild type and sti1∆ cells, there is no observable difference in the association between Hsp90 and Hsp70 [21]. Therefore, either there is a considerable difference between the yeast and mammalian complexes with respect to the requirement for Hop/Sti1 to link Hsp90 and Hsp70, or Hop/Sti1 is not required to bring together the two chaperones in vivo. Indeed, because Hsp70 and Hsp90 can interact independently with substrate molecules, Hop/Sti1 may be required only under some circumstances. How Hop/Sti1 exerts the observed stimulatory influence on the maturation of Hsp90 substrates remains an important question. In addition to its bridging role, which may or may not be important in vivo, Hop/Sti1 protein is apparently involved in the regulation of Hsp90 ATPase activity. In the presence of purified Sti1, the ATPase activity of purified yeast Hsp90 is reduced [135]. Sti1 also elicits a significant conformational change in Hsp90 upon binding to the chaperone in vitro [135]. Since the ATPase and chaperoning activities of Hsp90 are tightly coupled, Sti1 may regulate the chaperoning of the Hsp90 substrates through its effect on the ATPase activity.
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CyP40/Cpr6, Cpr7
CyP40 is a large immunophilin of the cyclophilin family, named thus because of binding to the immunosuppressive drug cyclosporin A [63]. CyP40 was discovered as a component of estrogen-receptor complexes [138]. The two yeast homologs of CyP40, Cpr6 and Cpr7, were first identified in a yeast two-hybrid screen by their interaction with Rpd3, a transcriptional regulator [49]. Cpr6 and Cpr7 are 41% identical, and share the organization of CyP40: amino-terminal peptidyl-prolyl cis-trans isomerase (PPIase) domain and a carboxy-terminal domain consisting of three TPR units, ending with a putative calmodulin-binding domain. Whereas CPR6 is heat-shock inducible, CPR7 is not [44]. cpr6∆ has no discernable phenotype, while cpr7∆ cells grow slowly at all temperatures. Overexpression of bovine CyP40, or overexpression of Cpr6, does not alleviate the slow growth of cpr7∆ cells [44]. Both Cpr6 and Cpr7 bind directly to Hsp90 through their TPR domains [47]. Although Cpr6 shares a higher degree of homology with human CyP40 (47%) than does Cpr7 [35%), cpr6∆ has no effect on the signaling of steroid receptors in S. cerevisiae [170]. A requirement for a cyclophilin in two Hsp90 functions was demonstrated by deleting CPR7, which leads to a decrease in both GR maturation and v-Src activity [47]. Cpr6 and Cpr7 are peptidyl-prolyl isomerases, enzymes that catalyze the conversion of peptide bonds adjacent to a proline from cis- to trans-. Since Hsp90 is involved in the folding of substrates into proper conformations, isomerase activities of immunophilins (both CyP40 and FKBPs) may help Hsp90 in this aspect of its cellular role. The isomerase activities of Cpr6 and Cpr7 are substantially different [99]. The PPIase activity of Cpr6 is 100-fold higher than that of Cpr7 when RCM-RNAse T1 is used as a substrate, and sixfold higher when two short synthetic peptides were assayed [99]. It has been proposed that Cpr7 might have evolved to isomerize a specific, still unidentified, substrate, and thus provide specificity for a subset of Hsp90 complexes [99]. However, the Cpr7 TPR domain alone is sufficient both for the normal growth of yeast cells, and apparently for full Hsp90 chaperone action [50, 44]. It is possible that, by binding to Hsp90, the TPR domains of co-chaperones modify the conformation Hsp90 itself is adopting, and thereby affect the physical and functional state of Hsp90. The two models of cyclophilin function in the Hsp90 complex are not mutually exclusive, and the regulation involving the isomerization of substrates could exist in addition to the TPR-mediated regulation of Hsp90 itself. CyP40 and its homologs may be partly regulated by localization. In mammalian cells, CyP40 is mainly nucleolar at normal conditions [97, 120], but is redistributed throughout the nucleus upon heat shock [97]. One
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possibility regarding the function of Hsp90 co-chaperones is that they serve to properly localize Hsp90 in the cell. For example, FKBP52 interacts with dynein through the isomerase domain [152]. The binding of hormone to the steroid receptor leads to the exchange of FKBP51 for FKBP52 in concert with the latter’s recruitment to dynein; transport of the steroid receptorHsp90–FKBP52–dynein complex from the cytosol to the nucleus is most likely mediated by the interaction of dynein with microtubules [39]. Therefore, the cyclophilin class of immunophilins might also be involved in the regulation of Hsp90 by cellular localization, in a manner similar to FKBPs. In vivo functions of CyP40 homologs are also known in other organisms. An Arabidopsis thaliana CyP40 homolog, SQN, is required for the vegetative growth of the plant shoot [7]. A Schizosaccharomyces pombe CyP40 homolog, Wis2, was found as a multicopy suppressor of mutations in cell cycle regulation [173], suggesting another possible connection between Hsp90 and the regulation of cell proliferation. 3.1.3
TTC4/Cns1
Cns1 was identified as a multicopy suppressor of the slow growth phenotype of cpr7∆ cells [44, 98] and in a multicopy screen for suppressors of the temperature sensitive phenotype of some hsp90ts– mutants [114]. Cns1 is essential in S. cerevisiae. It has three TPRs in the amino terminus, while the carboxy terminus contains a domain of unknown function. Cns1 binds to Hsp90 through the TPR domain, and appears to share complexes with Cpr7, but not with Cpr6 [44, 98]. In cpr7∆ cells, overexpression of Cns1 can restore both normal growth and Hsp90-related functions, including maturation of GR, and negative regulation of HSF [98]. Like CPR7, CNS1 is not heat-inducible. In contrast with Sti1/Hop, there is no evidence that Cns1 can bind directly to Hsp70. CNS1 also shows strong genetic interactions with CPR7 [188 – added in proof ]. This suggests that Cns1 and Cpr7 either share a common function, or are involved in the same complex, which may or may not contain Hsp90. The possibility that Hsp90 complexes exist which contain multiple TPRdomain co-chaperones has interesting implications for our understanding of the functional role of these co-chaperones. Immunophilins such as CyP40 and FKBP52 that bind to Hsp90 through their TPR domain were initially shown to exist in distinct Hsp90 complexes [79, 121]. This led to a hypothesis that Hsp90 contains only one TPR-accepting pocket, for which different TPR domain-containing co-chaperones compete. However, Sti1, like Hsp90 is a dimer in solution and binds to Hsp90 in a 1:1 molar ratio [135]. It has been shown that purified Cpr6 also binds to a Hsp90 dimer in a
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1:1 molar ratio, although Cpr6 is a monomer in solution [99, 135]. These findings suggest that two TPR proteins can bind to the Hsp90 dimer simultaneously and that TPRs may not necessarily compete for binding to Hsp90; instead, their binding may be independent of each other, or even cooperative. Consistent with this, others and we have shown that Cns1 and Cpr7 can be found in the same Hsp90 complexes in vivo [44, 98]. This model of binding corresponds well with our current understanding of the interactions between TPR proteins and Hsp90, mediated by the MEEVD motif. Since each of the two Hsp90 monomers contains this motif, it is easy to envision how two TPR domains can be accommodated by the Hsp90 dimer. Interestingly, the observation of at least one mixed TPR heterocomplex, containing Cns1 and Cpr7, suggests that other combinations may exist and that they may play a role in Hsp90 substrate specificity. By combining even a small set of different TPR co-chaperones, a large number of distinct Hsp90 complexes can be achieved, perhaps providing the specificity of the Hsp90 complexes for their substrates. Cns1 belongs to a protein family with homologs in S. pombe, Caenorhabditis elegans, Drosophila melanogaster, and humans. The Drosophila Dpit47 interacts with the DNA polymerase alpha subunit [36]. The human Cns1 homolog, TTC4, is of particular interest as a potential tumor suppressor. The TTC4 gene maps to the region of chromosome 1p31 which undergoes loss of heterozygosity (LOH) in up to 50% of breast cancers [69, 167]. In addition, in samples from patients with malignant melanoma, six different point mutations in TTC4 were found [128]. We have found that TTC4 can interact with yeast Hsp82 in vivo (Tesic, M. and R. Gaber, unpublished data), providing a possible connection between the Hsp90 chaperone machinery and human tumorigenesis. 3.1.4
PP5/Ppt1
Both human protein phosphatase 5 (PP5), and its S. cerevisiae homolog, Ppt1, were identified as TPR domain containing phosphatases [26]. The two comprise a unique phosphatase subfamily [162]. They share 46% identity in the phosphatase domain, and 36% identity in the TPR domain. The interaction of PP5 with Hsp90 in COS cells is mediated by its TPR domain [24]. Interestingly, when the TPR domain alone of PP5 is overexpressed, it lowers the transcriptional activity of GR in CV-1 cells, suggesting a stimulatory role for PP5 in steroid receptor maturation [24]. However, the use of antisense oligonucleotides leads to an increase in GR-dependent signaling [187]. Therefore, it is not clear whether PP5 antagonizes or enhances steroid receptor-mediated transcriptional activation [32]. So far, possible effects on cellular Hsp90 functions of deleting PPT1 in S. cerevisiae have not
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been investigated and may provide interesting insights into the intricacies of the mechanism of action of this phosphatase in Hsp90 complexes. The analysis of PP5-containing Hsp90 complexes demonstrates that this phosphatase competes with other TPR domain co-chaperones for binding to the Hsp90 [151]. Interestingly, PP5 is also able to compete with the binding of Hop, which could not be out-competed by the immunophilins [151]. This lends credence to the hypothesis that there is a hierarchy of binding to the TPR-accepting pocket of Hsp90, which is determined by the relative-binding constants, concentration, and availability of particular chaperones. The crystal structure of the amino terminus of PP5 was the first structure of a Hsp90-binding TPR domain to be solved [38], providing insight into the structure-function relationships of this highly versatile protein–protein interaction motif [89]. Each individual TPR consists of two antiparallel α helices at a 24° angle, which are tandemly repeated. Eight conserved residues, mostly small and large hydrophobic amino acids, determine the highly degenerate signature of any TPR motif, and are buried within the structure. Certain exposed charged residues likely to be involved in protein– protein interactions were mutated in PP5 and shown to be important for the binding of PP5 to Hsp90 [141]. These lysines and arginines are highly conserved among all TPR-domain containing Hsp90-binding co-chaperones. A structure of the TPR domain of Hop co-crystallized with the MEEVD peptide from Hsp90 corroborated these findings by showing that the very same charged residues made crucial interactions with Hsp90 [148]. Interestingly, mutations in the MEEVD sequence of Hsp90 significantly reduce, but do not abrogate, the binding of PP5 TPRs to Hsp90 [141], suggesting that there may be additional binding determinants between Hsp90 and its TPR co-chaperones. It is still unclear whether the phosphatase domain of PP5 is involved in the regulation of Hsp90-dependent events. The simplest model would suggest a role for PP5 in targeting the Hsp90 multichaperone complex to a substrate to be dephosphorylated. However, it is possible, as was shown for Cpr7 in S. cerevisiae [50], that the phosphatase domain of PP5 is dispensable for its Hsp90-related role. In support of the idea that TPR domains regulate Hsp90 function, the TPR domain of PP5 is emerging as a highly versatile functional unit. It can negatively regulate the phosphatase activity of PP5 by steric hindrance, which is released by the addition of polyunsaturated fatty acids [25, 154]. A study of a series of mutations in PP5 made in order to investigate how the TPR domain accommodates the interactions both with Hsp90 and the phosphatase domain of PP5 itself reveals that the residues involved in binding to Hsp90 are distinct from those involved in the
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autoregulatory role [81]. In general, multiple functions may exist for TPR domains of other Hsp90 binding proteins. Although ppt1∆ cells grows normally, treatment with antisense PP5 oligonucleotides leads to G1 cell cycle arrest in S. cerevisiae cells [186]. Investigation of possible downstream effects of PP5 loss of function reveals a strong increase in the level of p53 phosphorylation, and increased affinity for its genomic response elements; PP5 is also able to dephosphorylate p53 in vitro. Given these results, and the observed negative regulation of the GR, it is possible that PP5 is involved in cancers through effects on both GR and p53 regulation [187]. Since both GR and p53 interact with Hsp90, PP5 may be one of the co-chaperones dedicated to these Hsp90 substrates.
3.2
Other co-chaperones
3.2.1
p23/Sba1
Although it has been known for some time that steroid receptor complexes contain a protein of 23 kDa in size termed p23 [158], the S. cerevisiae homolog, Sba1, was identified only recently [10, 52]. SBA1 is constitutively expressed, and sba1∆ cells are viable at 30°C and 37°C, but grow slowly at 16°C. In S. pombe, a p23 homolog Wos2 interacts with the Hsp90 substrate Wee1, which is involved in cell cycle control [111]. In L cell cytosol and rabbit reticulocyte lysate reconstitution assays, p23 stabilizes Hsp90-steroid receptor complexes [42]. There are at least two requirements for the high-affinity binding of p23 to the amino terminus of Hsp90 [29]: dimerization of Hsp90 [62] and ATP binding (but not ATP hydrolysis) [20, 133]. p23 seems to play a role in the “coupling” of the ATPase activity and the substrate release by Hsp90 [180]. p23 may accomplish this, at least in part, by inhibiting the ATPase activity of Hsp90, thus extending the duration of the interaction between the client protein and the chaperone, allowing for the productive folding to occur [100]. This observation is in contrast with a previous report in which no effect on the ATPase activity of purified yeast Hsp90 was observed in the presence of purified Sba1 [117]. It has been somewhat difficult to establish whether Sba1 in S. cerevisiae is truly a functional homolog of p23. sba1∆ cells are sensitive to benzoquinoid ansamycins, a class of molecules such as GA, a potent inhibitor of Hsp90 ATPase activity. In general, sensitivity to GA of yeast cells carrying mutations in certain genes points to involvement of those genes in Hsp90-related pathways. Sba1 was found in Hsp90 complexes, along with Cpr6 and a small amount of Sti1; this interaction was inhibited by
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GA and another benzoquinoid ansamycin, macbecin I (Mb I) [52]. However, deletion of SBA1 does not suppress the lethality of cells in which v-Src is overexpressed. Similarly, AR and GR signaling is only marginally affected by sba1∆ [52]. On the other hand, the hypersensitivity of sba1∆ cells to benzoquinone ansamycins can be rescued by expression of human p23 [10]. A recent report, however, finds strong differential effects of both p23 and Sba1 on a number of steroid receptors [57]. It is notable that, in these assays, the effects of p23 and Sba1 are virtually indistinguishable. While the role of Hsp90 may be to control hormone-binding affinity of the steroid receptors, p23 could be affecting their transactivational activity, which may take place significantly later [57]. In agreement with this is the observation that the interaction between Sba1 and two different steroid receptors (GR and MR) persists even in the absence of Hsp90 [57]. Two other groups also observed Hsp90-independent binding of p23 to endogenous yeast Hsp90 substrates, Gcn2 and Hap1 [45, 72]. p23 is likely involved in the regulation of Hsp90 substrates in two ways: by binding the substrate proteins directly, and by controlling the coordination between the ATPase cycle and client refolding performed by Hsp90. 3.2.2
Hsp40 (Hdj)/Ydj1
Ydj1 is a S. cerevisiae homolog of bacterial DnaK co-chaperone, DnaJ, a founding member of a highly conserved family of Hsp40 chaperones [16]. Hsp40 enhances the ATPase activity of Hsp70, as well as the release of ATP from Hsp70 [23, 183]. Therefore, it is primarily an Hsp70 co-chaperone. However, it is present in Hsp90 complexes and affects Hsp90 functions. Hsp40 homologs in all organisms have a similar domain structure: an amino terminal J-domain stimulates ATP hydrolysis by Hsp70 and is flanked by a centrally located glycine and phenylalanine rich region; the carboxy terminus contains a domain capable of binding unfolded substrates. ydj1∆ S. cerevisiae cells grow slowly at 23°C, and are inviable at 37°C. This phenotype can be suppressed by the expression of human Hdj2, but not Hdj1 [56]. S. cerevisiae contains another Hsp40 homolog, Sis1, which is essential and involved in some Hsp70 functions [95]. The substrate-binding domain of either Sis1 or Ydj1 performs an essential function in yeast cells, and acts in cis with the J domain [77]. The involvement of Ydj1 in Hsp90 complexes was revealed by synthetic lethality of Ydj1G135D in combination with Hsp82G170D temperature-sensitive mutant [86]. By itself, Ydj1G315D increases both basal and hormone-induced activities of two steroid receptors (GR and ER) expressed in yeast. Ydj1G135D can also rescue from lethality S. cerevisiae cells in which v-Src is overexpressed. Although v-Src activity, but protein level is not reduced in
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ydj1∆ cells, mutations in the J-domain can reduce both mRNA and protein levels of v-Src [40]. Interestingly, mutations in Ydj1 have differential effects on various Hsp90 substrates. In ydj1∆ cells, androgen receptor-mediated signaling is compromised [56]. In contrast, in most ydj1 mutant cells, signaling mediated by GR and ER was enhanced, yet the levels and the activity of v-Src were mostly decreased [78]. Together with Hsp70, Ydj1 represses the activation of an endogenous yeast substrate, Hap1, in the absence of heme [71]. The negative effects of Ydj1 on the activation of Hsp90 substrates appear to be mediated by its J domain [71, 78]. Collectively, these findings suggest that Ydj1 is involved in multiple steps of client protein regulation by the Hsp90 machinery, including direct interactions with both the chaperones as well as the substrates themselves. 3.2.3
Sch9
Sch9 was discovered as a multicopy suppressor of a cdc25ts mutant [169], and subsequently found in a transposon mutagenesis screen as a suppressor of the G2/M cell cycle arrest phenotype exhibited by a HSF truncation mutant [1–583] [108]. It has homology with the serine/threonine class of protein kinases. S. cerevisiae cells carrying the HSF[1–583] mutant also show loss of some Hsp90-dependent activities: GR maturation, Hap1dependent transcription, and Ste11-mediated pheromone responsiveness. Loss of Sch9 activity restores normal function to these cells, suggesting that Sch9 can act as a negative regulator of Hsp90. Interestingly, deletion of SCH9 has no effect on pheromone response upon addition of the pheromone ligand; however, the basal level of activity is strongly enhanced [108]. Therefore, Sch9 probably acts to modulate Hsp90 function at the point of control of client proteins in the absence of their activating signals. Sch9 was also found to regulate stress responses and chronological life span in S. cerevisiae cells [51]. Although it is known that a decrease in Hsp90 function can slow down chronological aging of yeast cells [64], it is not clear whether Hsp90 plays a role in aging processes through Sch9. 3.2.4
Hsp110/Sse1
SSE1 and its homolog, SSE2, were identified in a calmodulin-binding fraction of a yeast expression library [110]. The two are 76% identical, and have homology to Hsp70 family members. sse1∆, but not sse2∆, causes slow growth at all temperatures; the combination of two deletions does not show synthetic effects. Overexpression of human Hsp110 does not rescue sse1∆ cells from slow growth. sse1∆ exhibits genetic interaction with sti1∆, and is hypersensitive to benzoquinone ansamycins; Sse1 physically
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associates with Hsp82 [92]. Loss of Sse1 function compromises GR maturation, and decreased basal HSF repression [92]. It is clear that Sse1 plays a role in the Hsp90 complexes, but the details of its involvement remain to be determined. 3.2.5
Cdc37
It has been known for a long time that Cdc37 is involved in the regulation of cell cycle at start [139]. Both Cdc37 and Hsp90 are required for the signaling by the sevenless receptor tyrosine kinase in D. melanogaster [37]. Like Hsp90, Cdc37 in S. cerevisiae is an essential protein and a dimer, but unlike Hsp90, its cellular concentration is low (about 0.01% total protein) and is not increased by heat shock [85]. Cdc37 was first seen in steroid receptor complexes as a protein of about 50 kDa size [124], and virtually all cytosolic Cdc37 is associated with Hsp90 [174]. Cdc37 has been dubbed the “protein kinase targeting subunit” of Hsp90 [164, 165]. It interacts with a number of kinases that also bind to Hsp90, including cell cycle regulator Cdk4 [165], Raf-1 [160], and v-Src [41]. However, it is now clear that its cellular role in general, and its role as an Hsp90 co-chaperone in particular, is more versatile than just as a determinator of specificity for Hsp90. Cdc37 is also a molecular chaperone in its own right. Like Hsp90, it is capable of maintaining unfolded β-galactosidase in a folding-competent state [85]. Genetic interactions between HSP90 and CDC37 in both budding yeast and fruit flies indicated that their roles overlap, but are not identical. Consistent with the idea that Cdc37 interacts solely with Hsp90 kinase substrates, overexpression of Cdc37 in the Hsp82G170D mutant restored normal v-Src, but not GR, activity [85]. However, signaling by a different steroid receptor, AR, was strongly affected by deletion of CDC37 [55]. Extending these findings, AR, but not GR, interacted with recombinant Cdc37 in a GA-sensitive manner [137]. Therefore, Cdc37 is found predominantly, but not exclusively, in Hsp90-protein kinase complexes. Apparently, even if the substrates of Hsp90 belong to the same functional class, like AR and GR, and share some structural similarities, they can have distinct requirements from the Hsp90 complex. It is possible that the structural differences between these substrates are significant enough to require the use of a different set of chaperones and co-chaperones by the cell. Binding to the Hsp90 complex may be determined by the folded state of the substrate, rather than simply by the structure of folded domains. It is conceivable that, in a partially folded state, AR resembles protein kinases more closely than does GR, and is therefore recruited by Cdc37–Hsp90 complexes.
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Cdc37 binds to Hsp90 at a site adjacent to the PP5 binding site. Full length PP5, but not the TPR domain alone, competes with Cdc37 for the binding to Hsp90, suggesting steric obstruction between Cdc37 and the phosphatase domain of PP5 [153]. A direct role for Cdc37 in neoplastic processes is beginning to emerge. Mouse mammary tumor virus promoter driven MMTV-Cdc37 transgenic mice develop tumors at a similar rate as MMTV-cyclin D1 mice [163]. In addition, human prostate cancer cells show increased expression levels of CDC37 [166]. How CDC37 realizes its oncogenic property is not clear. A possible mechanism involves its known interaction with oncogenes such as Cdk4 and Raf1. In addition, both Cdc37 and Hsp90 have been found to associate with oncogenic protein-tyrosine kinases, v-Yes, v-Fps, and v-Fgr [73]. Cdc37 may regulate on its own, and help Hsp90 in regulating, a number of proteins involved in cellular proliferation. Understanding the mechanism and selectivity of this co-chaperone will remain particularly important in the attempts to prevent cellular transformation by targeting Hsp90 and its substrates.
4
S. CEREVISIAE AS A SYSTEM TO STUDY HSP90 CO-CHAPERONES
Key to the use of S. cerevisiae as a model system to study conserved cellular processes are the similarities between yeast and higher eukaryotes with respect to that particular set of processes. As mentioned earlier, most components of the Hsp90 machinery, and the Hsp90 complexes themselves, are conserved in S. cerevisiae (see Table 1 and [22]). In addition, the mechanism of function of this machinery is similar to that in mammalian cells sufficiently to enable proper regulation of nonconserved substrates, such as steroid receptors and Src kinases. Some important co-chaperones, however, are not found in S. cerevisiae. A class of TPR co-chaperones that bind to Hsp90, consisting of large immunophilins FKBP51 and FKBP52, have no S. cerevisiae homologs. Hip, a co-chaperone for Hsp70, is also not conserved in S. cerevisiae. Two other co-chaperones missing in budding yeast are the regulators of Hsp90 and Hsp70, Bag-1 [80] and CHIP [5, 33]. It is possible that the functions these proteins play are conserved, but are performed by evolutionarily divergent homologs. Conversely, some co-chaperones of Hsp90 that have been identified in S. cerevisiae apparently do not have functional homologs in higher organisms. Hch1, a protein with no apparent homologs to any proteins in higher eukaryotes [34, 67], has been identified as a multicopy suppressor of the Hsp82E381K ts mutant [114]. Genetic analysis predicts that
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this protein is a general regulator of Hsp90, and it is surprising that it is not conserved. Therefore, although most components of Hsp90 complexes are conserved, some may be specific to S. cerevisiae, probably because they supply functions unique to the needs of a yeast cell. The absence of some Hsp90 co-chaperones in S. cerevisiae may prove to be advantageous in the study of nonconserved Hsp90 complex components. In the same way that expression of heterologous substrates provides a tool for investigating the role of Hsp90 complex components in vivo, expression in S. cerevisiae of mammalian-specific Hsp90 co-chaperones, for example FKBPs, may shed light on their in vivo roles. There are considerable differences in the intrinsic ATPase activity of Hsp90 and the way it is regulated between S. cerevisiae and metazoa. The ATPase activity of purified yeast Hsp90 is at least several fold higher than the ATPase activity of human Hsp90 [100, 112, 119, 135]. Furthermore, effects of yeast co-chaperones on the ATPase activity of yeast Hsp90 and effects of mammalian co-chaperones on the ATPase activity of mammalian Hsp90 are significantly different. While Sti1 decreases the ATPase activity of Hsp90 [135], Hop has no effect on the basal rate of ATP hydrolysis, but inhibits the substrate protein-stimulated rate [100]. In addition, while Sba1 apparently has no effect on the ATP hydrolysis by yeast Hsp90 [117], p23 stimulates basal and substrate-stimulated ATPase activity [100]. If these are bona fide differences between S. cerevisiae and human Hsp90, and not just discrepancies in experimental approaches, then findings from the yeast system may not be quite so readily extrapolated to mammalian cells, at least regarding the regulation of the ATPase activity. To compare the degree of conservation between fungal and mammalian Hsp90 complexes, it is instructive to consider those substrates that are conserved between yeast and higher eukaryotes. For example, HSF complexes in S. cerevisiae, Xenopus laevis, and an in vitro reconstituted system have all been studied [3, 48, 185]. While in S. cerevisiae Cpr7 helps Hsp90 in the attenuation of the heat shock response under both basal and induced conditions [48], immunodepletion of its mammalian homolog CyP40 in the in vitro reconstitution assay fails to elicit HSF1 activation [185]. Apparently, metazoan and yeast regulation of HSF differs with regards to the involvement of cyclophilins. On the other hand, regulation of some substrates exhibits striking similarities. For example, both Ste11 and its mammalian homolog Raf require Cdc37 for full function and association with the Hsp90 complex [2, 60]. Along the same lines, the models of Hsp90 regulation of eIF-2α kinases, fungal Gcn2, and mammalian PKR, demonstrate similar mechanisms with multiple steps of Hsp90 association and involvement of at least one common co-chaperone, Sba1/p23 [45, 46]. Identical requirements for the components of the Hsp90 complex by a
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particular substrate in S. cerevisiae and mammalian cells cannot be assumed, and will have to be established on an individual basis.
5
THE ROLE OF CO-CHAPERONES IN HSP90 COMPLEXES
It is clear from a large number of studies in model systems, as well as from in vitro approaches, that co-chaperones are versatile modulators of Hsp90 action that are involved with virtually every aspect of Hsp90 functioning. Underlying this adaptability is the fact that Hsp90 shares at least some of its co-chaperones with other molecular chaperones. Hop/Sti1, for example, binds both Hsp70 and Hsp90. In S. cerevisiae, Sti1, Cpr7, and Cns1 interact with a molecular chaperone Hsp104, which is responsible for both thermotolerance and maintenance of prion phenotypes in yeast cells [1]. Therefore, co-chaperones are likely to have very versatile roles both in the regulation of Hsp90 and other cellular functions in general. As we have seen, some co-chaperones can regulate the ATPase activity of Hsp90. Presence of purified Sti1 decreases the ATPase activity of purified yeast Hsp82 [135]. The ATPase activity of human Hsp90 is inhibited by the addition of p23 [100]. Given the coupling of the ATPase cycle to the chaperoning activities of Hsp90, the regulation of ATPase activity by the cochaperones is likely to be one of the most important aspects of the cochaperone involvement in the Hsp90 function. Chaperoning of substrates by Hsp70 and Hsp90 has been well established, but this activity may also be performed by the co-chaperones themselves. p23, CyP40, and FKBP52 all have chaperone activities in vitro [13, 58]. Some co-chaperones, including Ydj1, p23, and ARA9/XAP2/AIP are capable of binding client proteins directly [57, 71, 78, 102]. The formation of such ternary complexes may be important for the adoption of proper conformations by the substrates, or for the progress of the substrate to the next step in the refolding cycle. Hsp90 co-chaperones may also play a role in the proper localization of the substrates. For example, FKBPs apparently use their interaction with dynein to facilitate nuclear translocation of the steroid receptors [39]. Since CyP40 localizes to the nucleolus [97, 120], it may serve to localize Hsp90 to this cellular compartment. In further support for this role of co-chaperones, ARA9/XAP2/AIP regulates cellular distribution of AhR [82, 90, 125]. In spite of numerous experiments affirming the presence of co-chaperones in Hsp90 complexes, and mounting evidence that co-chaperones play important functional roles in these complexes, the question of whether any of the co-chaperones are absolutely required for a specific Hsp90 function
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remains unanswered. In vitro, two chaperones alone, Hsp70 and Hsp90, have been reported to mediate maturation of steroid receptors [136], arguing against an unconditional requirement for the co-chaperones. Furthermore, although the Hsp90 MEEVD sequence is considered to be sufficient for the binding of TPR domain co-chaperones, its deletion has no effect on the viability of S. cerevisiae cells [94]. This suggests that interactions of Hsp90 with all proteins that bind to it through their TPR motifs are dispensable. If this is the case, then TPR domain co-chaperones may simply be modulators of Hsp90 functions. Alternatively, it may not be the physical association with Hsp90 per se that is necessary for substrate regulation, but the presence of co-chaperones in the Hsp90 complex, which can also be achieved through binding to substrates directly. While some co-chaperones, like p23, are most likely generally required by Hsp90, others may be specific for certain classes of substrates, or only in some cell types, and under certain cellular conditions. Although it chaperones a large number of substrates, Hsp90 does not interact indiscriminately with partially unfolded or inactive proteins, suggesting that triage decisions are made in the cell that determine Hsp90 substrate specificity. One of the models of co-chaperone action, which posits that co-chaperones determine specificity of the Hsp90 complex for individual substrates, has been difficult to prove. Some Hsp90 co- chaperones show clear preference for one class of substrates. ARA9/XAP2/AIP is the only co-chaperone identified so far that apparently interacts with only one Hsp90 substrate, AhR [6, 19, 82, 83, 90, 102, 103, 125]. All other cochaperones show a less stringent predilection for Hsp90 substrates. Cdc37, for example, is present almost exclusively in protein kinase–Hsp90 complexes. However, Cdc37 also interacts with at least one steroid receptor, AR. Proteins containing TPR domains are the most obvious candidates for co-chaperones that determine specificity of the Hsp90 complex because they bind Hsp90 directly through the TPR domain, while the other domain in each co-chaperone (isomerase in the case of immunophilins, or phosphatase in the case of PP5/Ppt1) can mediate binding to the appropriate substrate. Especially intriguing is the idea that Hsp90 can accommodate more than one TPR domain co-chaperone (Cns1 and Cpr7, for example), thus increasing the number of possible co-chaperone combinations and, consequently, the number of substrates Hsp90 acts upon. However, it is still equally likely that co-chaperones determine specificity of Hsp90 function not by bridging the chaperone to the substrate, but by keeping Hsp90 itself in a conformation that favors some client proteins over others. Consistent with this idea, defining the substrate recognition site for Hsp90 has proven to be difficult, suggesting that this is a highly versatile chaperone capable of discriminating between similar proteins. Future work should be
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able to distinguish the two hypotheses of Hsp90 substrate determination, and, given the variety among co-chaperones, it is likely that the cochaperones will have widely differing roles in this process.
6
HSP90 CO-CHAPERONES IN CANCER
Hsp90 is of major interest to the area of cancer research as a direct target of an antitumor agent which is currently in clinical trials, a GA derivative 17-allylaminogeldanamycin (17-AAG) [127]. Although it is known that GA inhibits the ATPase activity of Hsp90, the mechanistic basis for the antineoplastic properties of this class of benzoquinone ansamycins is not clearly understood. Therefore, it is possible that a small number of key tumorigenic substrates become unregulated when Hsp90 function is partially lost. It is also possible that, given the involvement of Hsp90 in a number of processes regulating cellular growth at multiple steps, there is a cumulative requirement for Hsp90 by a large number of regulatory proteins, which leads to the observed effect of GA-related drugs. Several components of the Hsp90 complex are involved in the regulation of cell cycle proteins. For example, phosphorylation and subsequent degradation of Cln3, a yeast cyclin, depends on Ydj1 [179]. Wis2, a CyP40 homolog in S. pombe, can suppress a cell cycle defect [173]. As mentioned, Cdc37 interacts with Cdk4 and Cdk6 (but not Cdk2 and Cdc2) [165]. Hsp90 itself interacts with Wee1 kinase, which phosphorylates cyclin B [4]. Therefore, various components of the Hsp90 complex may be involved at multiple points of control of cell cycle, and mutations in these proteins could lead to the loss of regulation and cellular transformation. Certain substrates of Hsp90 have been implicated in tumorigenesis. Some are oncogenic kinases, such as v-Src, v-Yes, v-Fps, and v-Fgr [14, 91, 184]. Telomerase, which is involved in cellular immortalization, also requires Hsp90 complex for function [70]. A particularly important Hsp90 substrate is p53, a well-known tumor suppressor; differences in the interaction of Hsp90 with p53 wild type and mutant forms may have significant impact on our understanding of cancer development [8]. As the list of known Hsp90 client proteins is growing, the number of those involved in tumorigenic processes is likely to increase as well. Some co-chaperones, such as Cdc37 and TTC4, are themselves potential oncogenes or tumor suppressors [128, 163, 166, 167]. It is not clear, however, whether the role of Cdc37 as an oncogene is directly related to its involvement in Cdk4–Hsp90 complexes, and maybe also complexes with other oncogenic kinases, or the oncogenic role is an unrelated property of this co-chaperone.
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Because a large number of proteins depend on Hsp90 for normal levels and activity, it is a concern that indiscriminate inhibition of Hsp90 function, as is achieved by inhibiting its function through GA, may have widely pleotropic effects. If co-chaperones indeed determine the specificity for Hsp90, instead of inhibiting Hsp90 function directly, it might be more advantageous to inhibit the function of a specific co-chaperone involved in a complex with a tumorigenic substrate. Additionally, instead of completely inhibiting the function of a certain co-chaperone, specific inhibition of the physical interaction between Hsp90 and the co-chaperone might provide even greater specificity for the desired antitumor agent. In order to accomplish this, it will be necessary to understand the details of the physical interaction between the co-chaperones and Hsp90, as well as the various aspects of regulation by the co-chaperones of both Hsp90 and the substrates, in the context of an in vivo cell; for this approach, the yeast system is particularly well suited.
ACKNOWLEDGMENTS The authors would like to thank Ellen Nollen, Joshua Schnell, and Sricharan Bandhakavi for their critical reading of the manuscript and helpful suggestions.
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Chapter 7 YEAST AS A MODEL SYSTEM FOR STUDYING CELL CYCLE CHECKPOINTS
Carmela Palermo and Nancy C. Walworth Department of Pharmacology, UMDNJ-Robert Wood Johnson Medical School and Joint Graduate Program in Cellular and Molecular Pharmacology, UMDNJ-Graduate School of Biomedical Sciences and Rutgers, The State University of New Jersey, 675 Hoes Lane, Piscataway, NJ 08854-5635
The survival of any organism depends on a cell’s ability to accurately duplicate the genome and equally distribute one copy to each daughter cell. Successful cell reproduction entails passage through the discrete G1 (preDNA synthesis), S (DNA synthesis), G2 (post-DNA synthesis), and M (mitotic) phases of the cell cycle. Remarkably, the events of the cell cycle have been well conserved among eukaryotic organisms ranging from the unicellular yeasts to humans [23]. As a result, there is a diverse range of experimental organisms with evolutionary distinctions that contribute to our understanding of the mechanics that drive the cell cycle, including the yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe, the filamentous fungus Aspergillus nidulans, genetically tractable multicellular eukaryotes including the fruit fly Drosophila melanogaster and the nematode Caenorhabditis elegans, systems easily amenable to cell-free analysis including eggs and embryos of Xenopus laevis and cell culture systems derived from a variety of mammalian tissues. The timing and accuracy of cell division are critical for maintaining the integrity of the genome during normal growth and development. For example, the commencement of chromosome segregation is dependent on the completion of sequential events including DNA replication, spindle pole body or centrosome duplication, chromosome condensation, and spindle formation. Studies in the fission yeast S. pombe revealed that at the core of the cell cycle machinery is the cdc2+ gene product, p34cdc2 kinase, which is
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required at both the onset of S-phase and M-phase [4, 24]. When activated, Cdc2 controls mitotic entry via phosphorylation of a number of proteins. The activation of Cdc2 is itself tightly regulated requiring association with a cyclin regulatory subunit (cyclin B), appropriate subcellular localization, as well as critical phosphorylation and dephosphorylation events on an activating threonine residue and inhibitory tyrosine (Tyr 15) residue [20].
Figure 1. A schematic model of the DNA damage checkpoint pathway. DNA damage triggers several proteins to localize to presumptive DNA damage sites. Chk1 becomes phosphorylated by Rad3/Rad26, concomitant with binding to the 14-3-3 protein, Rad24 or Rad25. Chk1 phosphorylates Cdc25 and Wee1 to control their activity and/or subcellular localization resulting in a delay to activation of Cdc2 and therefore, a delay to mitotic entry.
To ensure timely entry into mitosis, activation of the cyclin B/cdc2 complex is tightly regulated by cell cycle checkpoints. Checkpoints monitor the status of a cell’s DNA and prevent inappropriate transitions into subsequent phases of the cell cycle when DNA damage or other perturbations to the cell cycle are sensed. Specifically, in the event of potentially catastrophic damage to the genome, checkpoints communicate with the basic cell cycle machinery to transiently arrest cell cycle progression. Such delayed mitotic entry prevents cells from initiating M-phase before DNA damage is repaired or DNA replication is complete. Chemicals, radiation, and DNA metabolism itself, incessantly threaten genomic stability by generating DNA damage. Theoretically, the absence of a checkpointinduced delay would allow propagation of unrepaired bases, broken chromosomes, or cells with an abnormal complement of chromosomes [25].
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The ramifications of such events could be detrimental to the cell resulting in cell death or to the organism, if unrepaired damage leads to uncontrolled proliferation. Mutations that affect the fidelity of cell cycle progression could understandably lead to disease states, such as cancer, characterized by uncontrolled cell divisions. In order to utilize checkpoint pathways as pharmacological targets it is critical to fully understand how the surveillance mechanism normally ensures proper cell division. The fission yeast, S. pombe has been an extraordinarily useful organism for studying the cell cycle, identifying components of the DNA damage checkpoint pathways for the development of a checkpoint-signaling model [26]. S. pombe is a rodshaped unicellular ascomycete fungus, which grows by apical extension, and executes many cellular processes similar to multicellular eukaryotes. In fact, this characteristic apical growth allowed for the initial identification of cell division cycle (cdc) genes whose function is required for the completion of cell division. Specifically, cdc mutants become blocked at specific stages of the cell cycle where they continue to elongate but fail to divide. Thus, S. pombe cell division cycle mutants could be easily identified microscopically as elongated cells. The DNA damage checkpoint pathway has largely been defined through the identification of loss-of-function mutants that fail to delay mitotic entry following DNA damage in S. pombe. Remarkably, components of the checkpoint pathway have been well conserved spanning the evolutionary distance from unicellular yeast to flies, frogs, mice, and humans. Indeed, many of the human checkpoint proteins that have been identified to date have been found exclusively based on their homology to the fission or budding yeast counterparts (Table 1). A model of checkpoint signaling has emerged from studies in fission and budding yeast, complemented by studies in Drosophila, mice, Xenopus, and cultured cells [19]. At its most rudimentary level, the checkpoint pathway consists of DNA damage sensors, signal transducers to relay the signal, and effectors that specifically interact with the cell cycle machinery. In yeast, checkpoints are generally not required for survival, only becoming essential when DNA damage or perturbations to DNA replication emerge. Thus, checkpoint components can be identified through genetic screens that select for mutants that are sensitive to DNA-damaging agents. For example, when normal cells are damaged by ultraviolet (UV) irradiation, most will transiently arrest cell cycle progression, repair DNA damage, and upon completing repair proceed through the cell cycle [35]. Mutations that render cells sensitive to DNA damage can be classified as either repair defective or checkpoint defective [13]. Specifically, mutations that render cells repair defective will result in a permanent cell cycle arrest. On the other hand,
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cells harboring mutant checkpoint components bypass the arrest and proceed through subsequent phases of the cycle with unrepaired DNA damage, ultimately resulting in cell death. Table 1. Checkpoint pathway components Functional class ATM family Complex with Rad3 PCNA-like
Protein kinases BRCT containing RFC Homology
S. pombe Red3 Rad26 Rad1 Rad9 Hus1 CHK1 Cds1 Cut5 Crb2 Rad17
H. sapiens ATR ATRIP hRad1 hRad9 hHus1 Chk1 Cds1/Chk2 53BP1 BRCA1 hRad17
S. cerevisiae MEC1 LCD1/DDC2 RAD17 DDC1 MED3 CHK1 RAD53 RAD9 RAD24
Early genetic studies of radiation-sensitive (rad) mutants of fission yeast, originally isolated due to their sensitivity to UV and ionizing radiation, led to the identification of some of the first checkpoint pathway components [22]. Further studies revealed that four of the original rad mutants (rad1, rad3, rad9, rad17) exhibited a complete loss of viability and aberrant attempts at mitosis following exposure to hydroxyurea (HU), an inhibitor of DNA synthesis, [2]. This demonstrates that rad1, rad3, rad9, and rad17 are normally responsible for recognizing incomplete DNA replication and ensuring that mitosis is dependent on the completion of S-phase. Additionally, epistatic analysis of the rad mutants with mutant components of the cell cycle machinery provided evidence that the DNA damage checkpoint pathway directly impinges on the cell cycle regulatory machinery [11]. The major cell cycle regulatory kinase controlling mitotic entry is Cdc2, which is activated by the Cdc25 phosphatase [27] and inhibited by the kinase activity of Wee1 and Mik1 proteins [17]. A temperature-sensitive loss-of-function allele of wee1 (wee1-50) renders cells viable at both permissive and restrictive temperature, though at restrictive temperature causes cells to divide at a smaller size than wild type. Double mutants of rad1, rad3, rad9, or rad17 with wee1-50 are inviable at restrictive temperature and display mitotic abnormalities as a consequence of inappropriately advancing into mitosis when wee1 activity is decreased [2]. This synthetic lethality between the checkpoint genes and cell cycle regulatory genes was the first evidence in S. pombe demonstrating that the checkpoint pathway that responds to DNA damage must also communicate with the cell cycle machinery. While the first checkpoint components emerged by testing the ability of radiation-sensitive mutants to initiate mitosis amidst a block to DNA
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replication, additional genetic screens followed to expand the search for specific components of the pathway. A genetic screen in which nitrosoguanidine-mutagenized cells were analyzed for sensitivity to HU led to the identification of the hus mutants (hydroxyurea sensitive) [10]. Interestingly, in addition to the “new” hus genes, alleles of the aforementioned rad1, rad3, and rad17 genes emerged in this genetic screen as well. The hus1 mutant, like the rad mutants, can be characterized as checkpoint deficient, evidenced by its increased radiation sensitivity and synthetic lethality with a wee mutant [10]. However, viability assays of synchronous cultures of the hus1, rad1, rad3, and rad17 mutants revealed that the cells became irreversibly arrested in S-phase after HU treatment, whereas a dominant mutant allele of cdc2, cdc-3w which is independent of cdc25 activation, loses viability as they undergo abortive mitosis [10]. This finding suggests that while the hus and rad genes play a role in the checkpoint pathway that couples mitosis to completion of DNA replication, additionally they assume a secondary function that is required for recovery from DNA damage. An alternative method of searching for new checkpoint mutants employed ethyl methanesulfonate (EMS) mutagenesis of cells harboring a temperature-sensitive allele of DNA ligase (cdc17-K42) and subsequent screening for UV- or HU-sensitive mutants. At restrictive temperature DNA ligase activity is decreased in these cells, leading to unligated DNA fragments that may generate a DNA damage response. The rationale for employing the DNA ligase mutant background was to distinguish components of the checkpoint that respond to DNA damage rather than to a block in S phase, such as that imposed by the HU treatment [3]. Two additional checkpoint components, rad26 and rad27, were isolated through this unique genetic screen. Functional characterization of the rad26 gene by deletion of an internal region of the ORF and subsequent replacement with a functional gene for the ura4 auxotrophic marker, revealed that it was inessential for viability. Consistent with the phenotype of a checkpoint defect, deletion of rad26 conferred increased sensitivity to HU treatment and ionizing radiation. Furthermore, similar to the hus1 mutant, the rad26-deleted strain was unable to reverse the S-phase arrest imposed by HU treatment. Strikingly, a point mutant of this gene, rad26-T12, was found to be radiation sensitive, yet displayed a normal mitotic arrest in response to ionizing radiation [3]. The phenotypic discrepancy between the rad26 point mutant and null allele can be resolved by proposing that the rad/hus gene products form a complex [3]. Mutations in any particular component of the complex could lead to loss of the specific function of that gene product, for example ability to repair DNA damage, but would not affect the integrity of the complex as a whole. On the other hand, complete loss of the gene product
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might structurally disrupt the complex and hence lead to complete failure of the signal transduction system. This finding began to illustrate the complex interactions and multiple feedback controls that exist, yet converge on a single signal transduction pathway, whose ultimate task is to delay mitotic progression. Interestingly, the predictions made based on genetic observations are now supported by biochemical evidence that the Rad1, Rad9, and Hus1 gene products indeed form a heterotrimeric complex that is related in structure to the PCNA processivity factor for DNA polymerase δ [7]. As sophisticated genetic screens began to establish the checkpoint pathway in S. pombe, serendipitous findings also contributed candidates for the central cast of players that couple the checkpoint pathway to the mitotic apparatus. As one example, a genetic screen aimed to identify elements that directly interact with Cdc2 utilized a cold-sensitive cdc2-r4 mutant that grows as wild type at 36°C, yet undergoes cell cycle arrest at the restrictive temperature of 25°C [17]. The cdc2-r4 mutation itself suppressed the lethality resulting from simultaneous loss-of-function of the inhibitory kinases wee1 and mik1, allowing cells to survive in the absence of tyrosine phosphorylation of Cdc2 and indicating that other mechanisms of Cdc2 regulation are in operation [17]. Introduction of a multicopy gene library into the cdc2-r4 strain led to isolation of plasmids that suppressed the effects of the cold-sensitive mutation at 18°C [33]. Consequently, this screen revealed one of the core components of the checkpoint pathway known as chk1 [33]. Deletion of the chk1 gene does not affect cell viability; however, a characteristic checkpoint defect is evident when cells are exposed to UV irradiation. Notably the UV sensitivity of a chk1 disrupted strain can be largely rescued by imposing an “artificial” G2 delay, discriminating this checkpoint component from gene products that are involved in DNA repair processes [33]. Chk1 is a member of the serine/threonine family of protein kinases and itself was found to undergo phosphorylation in response to DNA damage [34]. The phosphorylated form, observable by SDS-PAGE as a decreased mobility form of the protein, has proven to be a useful tool to dissect how previously isolated and novel checkpoint components interact with Chk1 to arrest cell cycle progression in response to DNA damage. In fact, epistatic analysis with the rad checkpoint genes indicated that damage induced phosphorylation of Chk1 depends on the function of rad1, rad3, rad9, rad17, and rad26, [34], rad24 [8], and crb2 [28]. To further define the role of Chk1 in the checkpoint pathway, the full-length gene was utilized as bait in a yeast two-hybrid screen of an S. pombe cDNA library for proteins that interact with Chk1. Two fission yeast 14-3-3 proteins, Rad 24 and Rad25, emerged from this screen [8]. Structural data
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supports a role for the 14-3-3 family of proteins in signal-transduction pathways, perhaps as scaffolding or adaptor proteins that bring together signaling molecules [1]. Intriguingly, rad24 and rad 25 also emerged in the genetic screen that identified rad26 as a pathway element [3, 12]. Accordingly, the Chk1-Rad24/Rad25 interaction was further pursued and molecular analysis demonstrated that the interaction indeed occurs in vivo. Furthermore, the association is stimulated by DNA damage, with the phosphorylated form of Chk1 preferentially interacting with the 14-3-3 proteins [8]. This data suggests that association of 14-3-3 proteins with Chk1 following DNA damage plays a significant, yet to date undetermined, role in the checkpoint pathway. In an independent genetic screen, the cds1 gene (Checking DNA Synthesis) was isolated as a multicopy suppressor of a temperature-sensitive DNA polymerase-α mutant [21]. Phenotypic characterization of cds1 revealed that the gene was dispensable for normal growth or survival in response to UV light, yet essential for survival when DNA replication was inhibited by treatment with HU. Further assessment of genetic interactions demonstrated that the wild-type gene almost completely rescued the HU sensitivity, but not the UV sensitivity, of the rad1, rad3, and rad9 mutants [21]. The ability of Cds1 to suppress HU sensitivity of the checkpoint mutants implied that its primary role was to instigate an S-phase specific checkpoint arrest. However, cells in which the cds1 gene has been eliminated by a knockout mutation, do arrest in the presence of HU, but fail to recover from that arrest. Thus, as cells resume cell cycle progression upon resuming DNA synthesis, they enter a catastrophic mitosis [21]. It has been proposed, therefore, that Cds1 plays an important role in stabilizing replication forks that have stalled as a result of depleted pools of ribonucleotides [15]. Consistent with this suggestion, cells lacking Cds1 that have been exposed to HU exhibit activation of the DNA damage checkpoint as manifested by phosphorylation of Chk1 [15]. Thus, while Cds1 and Chk1 are both critical effectors of the checkpoint response to inhibition of DNA replication, they seem to perform distinct roles in the process. In addition, the results of assays that measure the response of Cds1 and Chk1 to DNA damage suggest that while both proteins require the function of Rad3 to be activated, there is a temporal distinction as to when one or the other kinase can respond to DNA damage [18]. Loss of mitotic checkpoint control can be plausibly associated with the development and progression of cancer. Interestingly two additional fission yeast checkpoint proteins, Cut 5 and Crb2, contain BRCT domains (BRCA1 C Terminus) that are also found in the human breast and ovarian cancer susceptibility gene, BRCA1 [28, 32]. BRCT domains are found in a growing number of proteins involved in DNA metabolism and can promote
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protein–protein interactions [5]. The cut5 gene was initially identified as a temperature-sensitive mutation that disrupts the normal coordination between nuclear division and cytokinesis leading to what is known as the “cut” phenotype (cell untimely torn) [29, 30]. Subsequent genetic analysis of this gene revealed that cut5 is required for both the onset of DNA replication and to establish a DNA damage-dependent mitotic arrest [29]. Interestingly, cut5 is the only gene that has been assigned two mutually exclusive roles in DNA replication and checkpoint control. The second BRCT-containing protein was identified in a yeast two-hybrid screen for Cut5 interacting proteins [28]. Deletion of the crb2 gene compromised the DNA damage checkpoint, as in the rad mutants, and therefore warranted investigation into Crb2’s interaction with the other checkpoint proteins. Consequently it was found that wild-type copies of the rad1, rad3, rad9, rad17, or rad26 genes could not suppress the UV sensitivity of a crb2 null strain, while overexpression of chk1 did improve the survival of the UV-damaged strain [28]. Thus, the interactions hint that Crb2 may act downstream or in parallel to the Rad checkpoint proteins, while functioning upstream of Chk1 in the signaling pathway. Furthermore, molecular characterization of Crb2 indicated that a Rad-dependent modified form of the protein appears in response to UV irradiation at the same time that phosphorylated Chk1 appears. One interpretation of these results assigns Crb2 a role in the checkpoint pathway in which it is modified in response to DNA damage and subsequently transmits a signal through Chk1 to mediate cell cycle arrest. It seems that from the moment that the DNA damage checkpoint was discovered in yeast it was anticipated that a correlation would be made between loss of checkpoint control and onset of cancer. Indeed, one might assume that loss of checkpoint control is a favorable condition for promoting unregulated cell division and, therefore, that oncogenic processes might target key checkpoint regulators to achieve this hallmark trait. Hence the discovery of checkpoint components, cut5 and crb2, that contain structural features in common with domains characterized in two human cancer susceptibility genes increase suspicion that a cancer connection is likely. Evidence is mounting that other checkpoint genes may play critical roles in the development of cancer as well. For example, the human ATM protein, which is a member of a family of proteins homologous to the S. pombe Rad3 protein, also plays a role in signaling the presence of DNA double-strand breaks to delay cell cycle progression [14]. Mutations in ATM lead to the recessive human disorder ataxia telangiectasia (AT), in which patients suffer from progressive neurodegeneration and the tendency to develop cancers, particularly lymphomas [14]. Experimental analysis of cells isolated from AT patients demonstrated that they were extremely sensitive to ionizing
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radiation as a result of failure to arrest the cell cycle in response to DNAdouble-strand breaks. On the other hand, analysis of Chk1 null mice embryos revealed that some checkpoint proteins are essential during embryogenesis, as cells succumb early on in development and exhibit mitotic abnormalities [16, 31]. Similar phenotypes are observed with mice null for the Rad3 family member ATR [6, 9]. Undoubtedly in the near future, further genetic analysis will help to demonstrate how disruption of checkpoint surveillance may lead to changes in genomic stability that are key to tumor evolution.
ACKNOWLEDGMENT Work in the Walworth laboratory is supported by the NIH (GM53194).
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Aitken, A. 1996. 14-3-3 and its possible role in co-ordinating multiple signalling pathways. Trends Cell Biol. 6:341–347. 2. al-Khodairy, F., and A. M. Carr. 1992. DNA repair mutants defining G2 checkpoint pathways in Schizosaccharomyces pombe. EMBO J. 11:1343–1350. 3. al-Khodairy, F., E. Fotou, K. S. Sheldrick, D. J. F. Griffiths, A. R. Lehmann, and A. M. Carr. 1994. Identification and characterization of new elements involved in checkpoints and feedback controls in fission yeast. Mol. Biol. Cell 5:147–160. 4. Beach, D., B. Durkacz, and P. Nurse. 1982. Functionally homologous cell cycle control genes in budding and fission yeast. Nature 300:706–709. 5. Bork, P., K. Hofmann, P. Bucher, A. F. Neuwald, S. F. Altschul, and E. V. Koonin. 1997. A superfamily of conserved domains in DNA damage-responsive cell cycle checkpoint proteins. FASEB J. 11:68–76. 6. Brown, E. J., and D. Baltimore. 2000. ATR disruption leads to chromosomal fragmentation and early embryonic lethality. Genes Dev. 14:397–402. 7. Caspari, T., M. Dahlen, G. Kanter-Smoler, H. D. Lindsay, K. Hofmann, K. Papadimitriou, P. Sunnerhagen, and A. M. Carr. 2000. Characterization of Schizosaccharomyces pombe Hus1: a PCNA-related protein that associates with Rad1 and Rad9. Mol. Cell. Biol. 20:1254–1262. 8. Chen, L., T.-H. Liu, and N. C. Walworth. 1999. Association of Chk1 with 14-3-3 proteins is stimulated by DNA damage. Genes Dev. 13:675–685. 9. de Klein, A., M. Muijtjens, R. van Os, Y. Verhoeven, B. Smit, A. M. Carr, A. R. Lehmann, and J. H. J. Hoeijmakers. 2000. Targeted disruption of the cell-cycle checkpoint gene ATR leads to early embryonic lethality in mice. Curr. Biol. 10:479–482. 10. Enoch, T., A. M. Carr, and P. Nurse. 1992. Fission yeast genes involved in coupling mitosis to the completion of DNA replication. Genes Dev. 6:2035–2046. 11. Enoch, T., and P. Nurse. 1990. Mutation of fission yeast cell cycle control genes abolishes dependence of mitosis on DNA replication. Cell 60:665–673.
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12. Ford, J. C., F. al-Khodairy, E. Fotou, K. S. Sheldrick, D. J. Griffiths, and A. M. Carr. 1994. 14-3-3 protein homologs required for the DNA damage checkpoint in fission yeast. Science 265:533–535. 13. Hartwell, L. H., and T. A. Weinert. 1989. Checkpoints: controls that ensure the order of cell cycle events. Science 246:629–634. 14. Lavin, M. F., and Y. Shiloh. 1997. The genetic defect in Ataxia-Telangiectasia. Annu. Rev. Immunol. 15:177–202. 15. Lindsay, H. D., D. J. F. Griiffithes, R. J. Edwards, P. U. Christensen, J. M. Murray, F. Osman, N. Walworth, and A. M. Carr. 1998. S-phase-specific activation of Cds1 kinase defines a subpathway of the checkpoint response in Schizosaccharomyces pombe. Genes Dev. 12:382–395. 16. Liu, Q., S. Guntuku, X.-S. Cui, S. Matsuoka, D. Cortez, K. Tamai, G. Luo, S. Carattini-Rivera, F. DeMayo, A. Bradley, L. A. Donehower, and S. J. Elledge. 2000. Chk1 is an essential kinase that is regulated by Atr and required for the G2/M DNA damage checkpoint. Genes Dev. 14:1448–1459. 17. Lundgren, K., N. Walworth, R. Booher, M. Dembski, M. Kirschner, and D. Beach. 1991. mik1 and wee1 cooperate in the inhibitory tyrosine phosphorylation of cdc2. Cell 64:1111–1122. 18. Martinho, R. G., H. D. Lindsay, G. Flaggs, A. J. DeMaggio, M. F. Hoekstra, A. M. Carr, and N. J. Bentley. 1998. Analysis of Rad3 and Chk1 protein kinases defines different checkpoint responses. EMBO J. 17:7239–7249. 19. Melo, J., and D. Toczyski. 2002. A unified view of the DNA-damage checkpoint. Curr. Opin. Cell Biol. 14:237–245. 20. Morgan, D. O. 1997. Cyclin-dependent kinases: engines, clocks, and microprocessors. Annu. Rev. Cell Dev. Biol. 13:261–291. 21. Murakami, H., and H. Okayama. 1995. A kinase from fission yeast responsible for blocking mitosis in S phase. Nature 374:817–819. 22. Nasim, A., and B. P. Smith. 1975. Genetic control of radiation sensitivity in Schizosaccharomyces pombe. Genetics 79:573–582. 23. Nurse, P. 1990. Universal control mechanism regulating the onset of M-phase. Nature 344:503–508. 24. Nurse, P., and Y. Bissett. 1981. Gene required in G1 for commitment to cell cycle and in G2 for control of mitosis in fission yeast. Nature 292:558–560. 25. Nyberg, K. A., R. J. Michelson, C. W. Putnam, and T. A. Weinert. 2002. Toward maintaining the genome: DNA damage and replication checkpoints. Annu. Rev. Genet. 36:617–656. 26. O’Connell, M. J., N. C. Walworth, and A. M. Carr. 2000. The G2-phase DNAdamage checkpoint. Trends Cell Biol. 10:296–303. 27. Russell, P., and P. Nurse. 1986. cdc25+ functions as an inducer in the mitotic control of fission yeast. Cell 45:145. 28. Saka, Y., F. Esashi, T. Matsusaka, S. Mochida, and M. Yanagida. 1997. Damage and replication checkpoint control in fission yeast is ensured by interactions of Crb2, a protein with BRCT motif, with Cut5 and Chk1. Genes Dev. 11:3387–3400. 29. Saka, Y., and M. Yanagida. 1993. Fission yeast cut5+, required for S phase onset and M phase restraint, is identical to the radiation-damage repair gene rad4+. Cell 74:383–393.
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30. Samejima, I., T. Matsumoto, Y. Nakaseko, D. Beach, and M. Yanagida. 1993. Identification of seven new cut genes involved in Schizosaccharomyces pombe mitosis. J. Cell Sci. 105 ( Pt 1):135–143. 31. Takai, H., K. Tominaga, N. Motoyama, Y. A. Minamishima, H. Nagahama, T. Tsukiyama, K. Ikeda, K. Nakayama, and M. Nakanishi. 2000. Aberrant cell cycle checkpoint function and early embryonic death in Chk1-/- mice. Genes Dev. 14:1439–1447. 32. Verkade, H. M., and M. J. O’Connell. 1998. Cut5 is a component of the UVresponsive DNA damage checkpoint in fission yeast. Mol. Gen. Genet. 260:426–433. 33. Walworth, N., S. Davey, and D. Beach. 1993. Fission yeast chk1 protein kinase links the rad checkpoint pathway to cdc2. Nature 363:368–371. 34. Walworth, N. C., and R. Bernards. 1996. rad-dependent response of the chk1-encoded protein kinase at the DNA damage checkpoint. Science 271:353–356. 35. Weinert, T. A., and L. H. Hartwell. 1988. The RAD9 gene controls the cell cycle response to DNA damage in Saccharomyces cerevisiae. Science 241:317–322.
Chapter 8 METABOLISM AND FUNCTION OF SPHINGOLIPIDS IN SACCHAROMYCES CEREVISIAE: RELEVANCE TO CANCER RESEARCH
L. Ashely Cowart*, Yusuf A. Hannun* and Lina M. Obeid# #Ralph H. Johnson Veterans Administration and the Departments of Medicine and Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, South Carolina
Over the last two decades, sphingolipids have emerged as important cell regulators. Included in this lipid class are the lipid mediator sphingosine, its phosphorylated derivative sphingosine-1-phosphate, and ceramide. In humans, the major known functions of sphingolipid-mediated cell regulation is the modulation of signaling pathways which control fundamental cellular processes including cell division, senescence, apoptosis, angiogenesis, and differentiation [40, 41, 74], all of direct relevance to cancer pathogenesis and progression. Because of these myriad activities, the enzymes that generate sphingolipid mediators are potential targets for cancer treatment. Thus, characterization of these enzymes with respect to activity, regulation, localization, and cellular function is fundamental to developing strategies for modulating sphingolipid levels in vivo. The yeast system has emerged as an invaluable tool for the identification, cloning, and characterization of enzymes of sphingolipid metabolism. Furthermore, insights gained from yeast studies of sphingolipid metabolism and function continue to push the forefront of sphingolipid research, especially since the roles of sphingolipids in stress and other cell responses appear to be conserved from yeast to human.
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OVERVIEW OF SPHINGOLIPID METABOLISM
As with many fundamental cellular pathways, sphingolipid metabolic pathways are highly conserved between yeast and mammalian cells. This high degree of evolutionary conservation underscores the fundamental importance of this lipid class and, importantly, allows the use of yeast as a model system for sphingolipid metabolism.
1.1
Long chain base synthesis
In both yeast and mammalian cells, sphingolipid biosynthesis begins with the production of 3-keto-dihydrosphingosine by condensation of palmitoylCoA with serine (Figure 1). This enzymatic activity is referred to as serine palmitoyltranferase (SPT), and the reaction is thought to be the rate-limiting step in sphingolipid biosynthesis and in yeast is catalyzed by the proteins lcb1p and lcb2p [86], which form a heterodimer [32]. Molecular modeling has identified a putative catalytic site in the cleft of the interface between the two peptides [32]. This structural model is supported by experiments demonstrating that both components are required for activity [75], and, in yeast, deletion of either of these subunits is lethal, rendering the cells auxotrophic for sphingoid bases (the products of the reaction) [85]. Thus, this pathway is vital to Saccharomyces cerevisiae. SPT has been identified as the target of the immunosupressant myriocin, which has structural similarity to sphingosine [14]. Inhibition of SPT by myriocin is conserved from yeast to mammalian SPT isoforms. Complementation screens in S. cerevisiae were used to clone both subunits in this organism [11, 75], and the subsequent characterization and cloning of mammalian isoforms was aided by information gained from this initial work in yeast [37, 76]. Interestingly, a novel subunit of the lcb1p/lcb2p complex was recently identified in yeast, tsc3p [33]. This gene is required for maximal SPT activity in yeast; however, deletion of this gene is not lethal under normal laboratory conditions as it is not required for basal SPT activity. TSC3 belongs to the group of genes identified as temperature suppressors of calcium sensitivity, many of which are involved in sphingolipid biosynthesis. Experimental data demonstrate that accumulation of the downstream sphingolipid product inositolphosphoceramide (IPC) leads to increased cell sensitivity to extracellular calcium; thus, deletion of any gene upstream which would decrease production of this metabolite, suppresses the calcium-sensitive phenotype. Importantly, the calcium-sensitive phenotype in yeast has led to the cloning of several enzymes in sphingolipid biosynthetic pathways (see Figure 1). Though a mammalian homologue of TSC3 remains to be identified, yeast model systems that enabled isolation of
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the TSC genes were and continue to be vital to cloning mammalian homologues of other enzymes of sphingolipid biosynthesis. Figure 1
Figure 1. Pathways of sphingolipid metabolism. Steps represented in bold text are shared by yeast and mammalian cells. Yeast-specific enzymes and compounds in italics and mammalian-specific enzymes and compounds in plain font.
The product of SPT is reduced to dihydrosphingosine by another gene in the TSC group, 3-keto-dihydrosphingosine reductase (TSC10). Long-chain bases have cell regulatory activities of their own; however, a variety of other lipid mediators are produced by their further metabolism. At this point in sphingolipid biosynthesis, yeast and mammalian pathways slightly diverge. In yeast, dihydrosphingosine is hydroxylated to form phytosphingosine by the SUR2/SYR2 gene product. This gene was identified originally as suppressing a defect in starvation survival [20] and by conferring sensitivity to the toxin syringomycin E [17]. Its biochemical activity was identified based on homology to another yeast sphingolipid hydroxylase [35]. Thus, in yeast, there are two major sphingoid base species formed by de novo
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pathways. In mammalian systems, however, dihydrosphingosine is the only major sphingoid base formed from de novo biosynthesis.
1.2
Phosphorylation and dephosphorylation of sphingoid bases
Sphingoid bases are subject to two known downstream metabolic pathways, one of which is their phosphorylation. Dihydrosphingosine and, in yeast, phytosphingosine can each be phosphorylated at C1 yielding dihydrosphingosine-1-phosphate (DHS-1-P) and phytosphingosine-1-phosphate, respectively. Additionally, sphingosine, which results from ceramide catabolism in mammalian cells (see below), can also be phosphorylated at C1 yielding sphingosine-1-phosphate (S-1-P). Cloning of genes responsible for sphingoid base kinase activity in yeast, LCB4 and LCB5 [78], laid important groundwork for the later cloning of mammalian isoforms [54, 62]. DHS-1-P can be cleaved by the DHS-1-P lyase into hexadecanal (palmitaldehyde) and ethanolamine phosphate in yeast, and in mammals, S-1-P is cleaved into hexadecenal and ethanolamine phosphate. The yeast gene responsible for this activity, DPL1, was cloned, and its deletion confers an increased sensitivity to growth inhibition by sphingoid bases [88]. This phenotype was integral to the cloning of the sphingoid base kinases LCB4 and LCB5 (above), as it was speculated that the reason for DPL1-dependent sphingoid base sensitivity was actually due to accumulation of sphingoid base phosphates [92]. Thus, deletion of the sphingoid base kinases reversed the phenotype of the DPL1 deletion [52]. This is but one example of the simple and elegant experimental strategies possible in yeast which have allowed the cloning of a multitude of genes. In addition to DPL1-mediated cleavage, sphingoid base phosphates can be dephosphorylated by sphingoid base phosphate phosphatases. The yeast enzymes YSR2/LCB3/LBP1 and YSR3/LBP2 were cloned and characterized in yeast [68, 70], and this facilitated the subsequent cloning of a murine sphingoid base phosphate phosphatase by sequence homology, especially in the highly conserved active site regions [67]. Both yeast sequences and the murine sequence facilitated the cloning of two human sphingosine phosphate phosphatase isoforms which may eventually become targets for cancer therapeutics [55, 80].
1.3
Ceramide biosynthesis
In addition to their metabolism by phosphorylation, both dihydrosphingosine and phytosphingosine can be N-acylated to form dihydro- or
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phytoceramide, respectively, by yeast ceramide synthases. There is little production of phytoceramide in mammalian cells, rather, dihydrosphingosine is converted to dihydroceramide by (dihydro)ceramide synthase before reduction to ceramide by dihydroceramide desaturase. This divergence is potentially very important, as dihydroceramide has few, if any, known cellular effects [4]. Thus, dihydroceramide desaturase may be a key regulatory point in mammalian sphingolipid metabolism, regulating the relative levels of the active lipid ceramide and the inactive lipid dihydroceramide. Sphingolipid biosynthesis in yeast continues to diverge slightly from mammalian pathways at the point of ceramide biosynthesis. Acyl groups detected in yeast ceramides typically have very long carbon chains (C26); however, in mammalian ceramides, N-acyl groups range in length from 16 to 24 carbons with varied degrees of saturation [21]. Despite these differences, overall the biosynthesis is highly conserved at a biochemical as well as a functional level (see below, Figure 1, Table 1). Table 1. Comparison of sphingolipid metabolism and function in Saccharomyces cerevisiae and mammals. (Fungal toxin inhibitors are listed in parentheses.) Activity Enzyme
Yeast Function
Enzyme
Mammalian Function
Production
LCB1 LCB2
Heat stress [49],
LBC1 LBC2
Heat stress [48],
of 3-keto-
heterodimer
cell cycle arrest [49],
heterodimer
hereditary sensory
dihydro-
(myriocin,
endocytosis [104],
neuropathy type I
sphingosine
syringomycin)
intracellular protein
[32]
transport [101] N-acylation
LAG1, LAC1
of sphingoid
(fumonisin)
Influence life span [18]
UOG1
and differentiation
bases Sphingoid
Regulator of growth [98]
LCB4, LCB5
base kinase
Clear sphingoid base
hSPP1, hSPP2, mSPP1
Clear sphingoid
uptake [70], clear
base phosphates;
sphingoid base
change balance
phosphates, cell
between ceramide
proliferation [69],
and S-1-P
heat stress [69] Ceramidase
YPC1, YDC1
Clear ceramide/
Neutral ceramidase,
Clear ceramide/
produce sphingoid
Alkaline ceramidase
produce sphingoid
bases
bases
Addition of
AUR1
Calcium sensitivity
Sphingomyelin
Correlate with
headgroups
(aureobasidin)
[23]
synthase, Glucosyl-
transformed pheno-
ceramide synthase
type [65]; drug
to ceramide
resistance [63, 64] Breakdown
ISC1
Cell wall in
Neutral sphingomye-
Central link between
of complex
Schizosaccharomyes
linase 1, neutral
apoptosis-inducing
sphingolipids
pombe [27]
sphingomyelinase 2,
factors and ceramide
acid sphingomyelinase
formation [38]; lipidstorage disease [29]
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Yeast enzymes required for sphingoid base N-acylation are LAG1 and LAC1 [91]. It is not yet established whether the enzymes encoded by these genes produce ceramide directly or act as necessary activators of an as yet unidentified ceramide synthases; however, the dependence of ceramide formation on these proteins as well as their ability to control intracellular ceramide levels have been well established biochemically [91]. (Longevity assurance gene LAG1) was initially identified as a gene whose deletion conferred increased lifespan in yeast [91]. This is particularly interesting as studies in mammalian cells indicate that increased ceramide is correlated with cell senescence (see below). The functions of lag1p and lac1p are partially redundant, thus, deletion strains of LAG1 or LAC1 are viable; however, deletion of both genes causes a severe slow growth phenotype, indicating the importance of ceramide biosynthetic pathways in yeast [91]. Very interestingly, LAG1 homologues have been identified throughout eukaryotic organisms, including tomato [9], Caenorhabditis elegans [51], Drosophila [97], and human [51] based on sequence similarity to the yeast gene. In particular, a mouse homologue (UOG) was recently identified as necessary for N-stearoyl-sphinganine biosynthesis, and of particular interest is that this gene had a suspected role in the regulation of cell differentiation prior to discovery of its role in ceramide synthesis, suggesting not only biochemical conservation but possibly some degree of functional conservation as well [98]. Ceramide synthase was found to be a target of the group of fungal toxins known as fumonisins. Fumonisin inhibition of ceramide synthase was first reported in rat liver microsomes, but provided insight into mechanisms of the observed correlation between fumonisin consumption and development of cancer [99]. It appears that not all LAG1 homologues are fumonisin targets; however, the bulk of ceramide synthase activity in mammalian cells is inhibited by fumonisins. Similarly to sphingoid bases, ceramide can be phosphorylated at C1. A mammalian ceramide kinase was cloned based on sequence similarity to mouse and human isoforms of sphingosine kinase [93]. Interestingly, no yeast homologue for ceramide kinase has been identified. Presently, however, at least 50% of the yeast genome (i.e., protein-encoding open reading frames) are of unknown function. Thus, further characterization of ceramide kinase may well lead to identification of a yeast isoform, and, if so, yeast systems may be useful in functional characterization of ceramide1-phosphate.
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Formation of complex sphingolipids
A variety of complex sphingolipids are formed in both yeast and mammalian cells by the addition of various headgroups to the ceramide molecule. Again, though there is significant divergence between yeast and mammalian pathways, several aspects of these pathways are conserved between the two. In yeast, a glycerophospholipid-derived phosphoinositol headgroup is added to ceramide forming IPC. IPC can be mannosylated to form mannosyl-IPC (MIPC), and yet another phosphoinositol headgroup is added to MIPC, forming M(IP)2C. Together, these comprise the set of complex sphingolipids in yeast. Biosynthesis of these lipids is carried out by the IPC synthase AUR1, the MIPC synthases CSG1 and CSG2, and the M(IP)2C synthase IPT1 [23]. AUR1 was originally identified as the target of the antifungal agent aureobasidin A, as mutations in this gene conferred resistance to aureobasidin A [43]. Deletion of this gene is lethal, indicating a strict requirement for complex sphingolipids in yeast (and/or toxicity of the accumulated ceramide). Later biochemical studies identified the function of AUR1 as the IPC synthase and confirmed the inhibition of this enzyme by aureobasidin A [77]. Of particular interest is that the homologue of this gene in Cryptococcus neoformans, IPC1, has been shown to be involved in the pathogenesis of this virulent fungus [66]. As with the TSC group of genes, CSG1 and CSG2 also play roles in cell sensitivity to extracellular calcium, as a previous study demonstrated that mutants in either gene were hypersensitive to extracellular calcium [2, 3]. Thus, it appears that mannosylated IPC derivatives are important for resistance to high extracellular calcium concentrations. The mammalian set of complex sphingolipids is much larger and more diverse. Whereas in yeast the inositol headgroup derives from phosphatidylinositol, the analogous mammalian headgroup, phosphocholine, derives from phosphatidylcholine through an analogous transferase reaction which also generates diacylglycerol from the glycerophospholipid (reviewed in [39]). Unlike ceramide, which is associated with attenuation of cell growth through cell cycle arrest, apoptosis, or senescence (reviewed in [38, 42]), diacylglycerol is recognized as the physiological agonist of protein kinase C and is often associated with mitogenesis [7]. Importantly, in producing sphingomyelin, sphingomyelin synthase catalyzes both the consumption of ceramide and the formation of diacylglycerol, perhaps shifting the cellular balance of growth-attenuating mediators to growth-promoting mediators [39]. Therefore, this activity may be a key point of cell growth regulation, making it an excellent target for chemotherapeutic agents. Unfortunately, mammalian sphingomyelin synthase has evaded purificationand cloning.
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Hopefully further knowledge of the yeast orthologue will assist in this goal as well. Though mannose is the only sugar in S. cerevisiae sphingolipids, mammalian complex sphingolipids incorporate a variety of carbohydrate groups including lactose, glucose, galactose, and sialic acid. These activities in mammalian cells give rise to glycosphingolipids, gangliosides, and cerebrosides. It should be noted that in some other fungal species including Candida albicans, glucosylceramide synthase has been detected [60], though the significance of this activity is not known [59].
1.5
Sphingolipid catabolism
Though complex sphingolipids in yeast are structurally different from complex sphingolipids in mammalian cells, there is surprising functional conservation of sphingolipid catabolic pathways. These pathways are important not only because of their ability to regulate sphingolipid levels, but also with respect to their participation in formation of ceramide and sphingoid bases as they break down more complex sphingolipids. Thus, two major routes contribute to formation of bioactive sphingolipids: de novo biosynthesis and catabolic pathways. In yeast, IPCs and their mannosylated derivatives are hydrolyzed to produce phytoceramide by the enzyme isc1p. The analogous enzyme in mammalian cells, neutral sphingomyelinase, hydrolyzes sphingomyelin to produce ceramide. The extended family of sphingomyelinases consists of several isoforms based on expression, intracellular localization, and pH optima [61]. Just as hydrolysis of inositolphosphoceramide and sphingomyelin produces phytoceramide and ceramide, respectively, hydrolysis of ceramide by ceramidases produces sphingoid bases. In yeast, the phytoceramidase YPC1 was first identified by yeast genetic screens, and the dihydroceramidase YDC1 was cloned based on sequence homology [71, 72]. The sequences of these two genes greatly assisted the cloning of several mammalian ceramidase isoforms [73]. Importantly, the same pathways that metabolize de novo sphingolipids also may metabolize ceramide and sphingoid bases derived from catabolic reactions.
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SPHINGOLIPID FUNCTIONS
2.1
Modulation of signaling pathways
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Studies in yeast have demonstrated the modulation of signaling pathways by sphingolipids, including the modulation of the protein kinases pkh1p and pkh2p by sphingoid bases [31]. These kinases act upstream of pkc1p and the YPK kinases 1 and 2, which control endocytosis and other cell processes [90]. Ongoing research is focused on discovering new pathways of sphingolipid action in yeast since these pathways may be conserved between yeast and mammalian systems. One of the original signaling activities attributed to sphingolipids was the modulation of protein kinases [5]. Indeed, modulation of protein kinase C activity by sphingosine has been implicated in a variety of downstream effects of sphingolipid production. Other studies led to the identification of the ceramide-activated protein phosphatase (CAPP) [24]. This phosphatase activity was demonstrated to be directly activated by exogenous ceramide, and to have characteristics of protein phosphatase 2A (PP2A) including cation independence and inhibition by okadaic acid [24]. Other studies in mammalian systems also identified protein phosphatase 1 (PP1)-dependent CAPP activity, suggesting that other phosphatases may also act as CAPPs [12]. Work in yeast demonstrated the conservation of this activity in S. cerevisiae, where phosphatase activity was shown to mediate ceramide induced growth inhibition [30]. Importantly, genetic studies in cerevisiae demonstrated that the activity is comprised of the regulatory subunits TPD3 and CDC55 and the catalytic subunit SIT4 [79]. Ceramide interactions with the mammalian CAPP have been linked to numerous cellular activities including downregulation of c-myc in response to TNF [103], dephosphorylation of the oncogene c-jun [87], induction of cell cycle arrest through dephosphorylation of the retinoblastoma gene product Rb [56], regulation of alternative mRNA splicing via dephosphorylation of SR proteins [13], and the inactivation of PKCα and Bcl2 by dephosphorylation [57]. Other studies have demonstrated direct effects of ceramide on the kinase KSR (kinase suppressor of ras) which may be involved in mediating effects of ceramide on MAP kinases [105]. The protease cathepsin D was also identified as a direct target for ceramide, and, given its lysosomal localization, cathepsin D may define a compartment-specific pathway for ceramide action [45]. The discovery of these pathways regulated by ceramide underscores the significant influence of sphingolipid metabolism in the regulation of cell division, apoptosis, and differentiation. Thus, elucidation of the pathways of
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ceramide action in yeast and mammals is important, as these pathways may become excellent targets for development of chemotherapeutics.
2.2
Stress responses
Perhaps the most studied activities of sphingolipids in yeast involve yeast stress responses. In yeast, heat stress induces an acute upregulation of de novo sphingolipid biosynthesis. As early as 5 min, increases of several fold are seen in dihydrosphingosine and phytosphingosine, immediately followed by increases in phosphorylated sphingoid bases [92]. Subsequently the ceramides are observed to rise, around 20–30 min, when sphingoid bases decrease [22, 50]. Several components of the yeast heat stress response, including cell cycle arrest and proteolysis are dependent on de novo sphingolipid biosynthesis (Figure 2), as revealed in studies using a temperature-sensitive mutant in LCB1, lcb1-100, which cannot generate sphingolipids upon heat stress. This mutant does not undergo transient cell cycle arrest [49], and addition of exogenous phytosphingosine induces a transient cell cycle arrest [15, 16, 50]; thus, demonstrating an essential role for the acute formation of sphingolipids in the heat stress response.
Figure 2. Heat stress induces many activities in yeast, some of which are mediated by stressinduced de novo sphingolipid synthesis. Direct targets for sphingolipids in yeast include ceramide-activated protein phosphatase, comprised of sit4p, cdc55p, and tpd3p. Additionally, heat stress induces sphingolipid synthesis in some mammalian cell types. Interestingly,
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several chemotherapeutic agents induce de novo sphingolipid production and thus, their effects are likely mediated in part by sphingolipid biosynthesis. (See text for further discussion.)
Another key feature of the yeast heat stress response is the internalization and ubiquitin-mediated proteolysis of nutrient permeases. Previous studies in yeast have demonstrated that de novo sphingolipid biosynthesis is required for these activities, and that exogenously added phytosphingosine initiates this degradation in the absence of heat stress [15, 16]. This sphingolipid activity is at least somewhat conserved in some mammalian cells, as ceramide has been demonstrated to modulate the ubiquitin-mediated proteolysis of c-myc [82]. More investigation is needed in both yeast and mammalian systems to determine the specific mechanisms of sphingolipid modulation of this pathway. Interestingly, a recent study implicates lipid rafts in the transport and degradation of the nutrient permease Fur4p [44], and thus, the propensity of sphingolipids to form lipid microdomains [10, 34, 36, 47] may facilitate these activities. These findings in yeast led to the hypothesis that mammalian heat stress responses may depend at least partially on de novo generated sphingolipids. Indeed, subsequent studies in MOLT-4 human leukemia cells [48] confirmed this hypothesis. Heat stress in these cells induced a flux through sphingolipid biosynthetic pathways similar to those observed in yeast. Heat-induced de novo sphingolipid biosynthesis in MOLT-4 cells was shown to influence mRNA splicing by the modulation of SR protein phosphorylation state [48]. One difference between heat-induced de novo sphingolipid biosynthesis in yeast and in the MOLT-4 cells is that yeast accumulate sphingoid bases prior to ceramide formation, whereas the MOLT-4 cells showed no accumulation in sphingoid bases after heat stress, but accumulated ceramide, which is thought to be the active intermediate in modulation of many mammalian signaling pathways. Whether this situation is typical of mammalian systems and whether the sphingoid bases may play specific roles remains to be determined. Downstream regulatory roles of sphingolipids generated during stress responses is of interest to cancer research, as several chemotherapeutic agents including etoposide, daunorubicin, and gemciytabine have been demonstrated to cause increased de novo sphingolipid synthesis, and furthermore, many effects of these drugs are blocked by inhibitors of ceramide synthesis ([8], reviewed in [81, 89]). Thus, these agents act at least partially through the de novo pathway of sphingolipid generation. Therefore, understanding the mechanisms of formation and action of sphingoid bases and ceramides may lead to direct insight into the mechanisms of action of several chemotherapeutic agents and, especially, the mechanisms by which they activate stress response pathways.
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Ceramide and cell senescence
Many lines of evidence in a spectrum of systems indicate that ceramide levels are correlated with regulation of life span and cell senescence. These studies stemmed from early work indicating elevated ceramide levels in senescent human diploid fibroblasts (HDF) [96]. Furthermore, addition of ceramide to proliferating HDF induced several markers of senescence including cessation of DNA replication and dephosphorylation of Rb [19], suggesting a role for ceramide in mediating the conversion of cell phenotype from proliferating to senescent. Several clues exist as to the specific mechanisms by which ceramide may mediate the regulation of lifespan. A key phenotype of senescent cells is the lack of response to mitogens. Investigations into this particular facet of cell senescence led to the discovery that lack of mitogenic response is mediated through inhibition of phospholipase D (PLD). Senescent HDF showed ceramide-dependent inability to activate PLD, resulting in lack of PKC activation which in turn mediates the transcriptional response to mitogens. Treatment of quiescent young HDF with exogenous ceramide mimicked the inactivation of PLD and its sequelae observed in senescent HDF [82, 83, 96]. Other investigation has demonstrated that ceramide inhibits telomerase activity in a lung cancer cell line via inactivation of c-Myc [82]. This inactivation partially involves increased ubiquitylation and subsequent proteolytic degradation, as do some stress responsive events, as discussed above [82]. Current investigation focuses on the roles of ceramide in coordinating the development of these senescent phenotypes. Importantly, links between ceramide generation and cell senescence appear to be conserved in yeast. In S. cerevisiae, deletion of the longevity assurance genes LAG1 and LAC1 confers an increased life span in yeast by 30% [91]. As is the case with mammalian cells, each yeast cell has a defined number of cell division cycles it can undergo before reaching a nondividing state, or senescence. It may follow that LAG1 and LAC1 regulate yeast life span via the regulation of ceramide levels. Intriguingly, some of the Lac1/Lag1 homologues in other eukaryotes appear to contain not only LAG-1 homologous domains, but transcriptional regulatory HOX-motifs, suggesting the possibility of coupling between ceramide binding and transcriptional regulation [97]. The structural and functional conservation of these proteins, as well as their potential roles in the regulation of ceramide formation, is of intense interest. Indeed, ongoing studies in yeast and mammalian system will likely shed light on the relationship of LAG1 and its homologues to ceramide levels and cell growth control. In sum, studies in both yeast and mammalian systems have enabled the compilation of a general picture of ceramide influences on cell senescence and aging; this apparent functional conservation attests to the evolutionary
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importance of these pathways. Eventually these studies may provide novel targets for cancer therapeutics.
2.4
Sphingosine kinase-mitogenesis and vascularization
Tumor progression depends on successful vascularization to provide factors which enable tumor growth. These ideas have led to the hypothesis that reducing tumor vascularization is an effective way to block tumor progression. Novel insights came from work demonstrating that S-1-P is a high affinity ligand for the endothelium development and growth receptor-1 (EDG-1) [58] and other EDG receptors, recently renamed S-1-P receptors [46]. These are G-protein coupled receptors (GPCRs) which are involved in many cell processes including vascularization and mitogenesis [25]. Subsequent studies have further supported roles for S-1-P and sphingosine kinase in activation of these receptors [1, 53, 84, 95, 100, 102]. Of particular interest is that a member of this receptor family, the mammalian EDG-2 lysophosphatidic acid receptor, when heterologously expressed in yeast, was shown to couple to the pheromone signaling pathway [26], indicating evolutionary conservation between these two receptors. It will be interesting to see if sphingoid base phosphate activation of GPCRs is conserved in all eukaryotic systems.
3
CONCLUSIONS
Yeast studies have contributed to our understanding of sphingolipid formation, metabolism, and function in mammalian cells and yeast sphingolipid studies have become intertwined with mammalian studies, complementting, supporting, and indeed, often spurring research into mammalian sphingolipid biosynthesis and function. Key areas for further investigation include mechanisms of ceramide biosynthesis induction by chemotherapeutic agents and factors such as TNF, and determining functions of individual sphingolipids such as sphingoid bases and sphingoid base phosphates, and their mechanisms of action. In general, more research is needed into how sphingolipid metabolic pathways are coordinated to regulate relative cellular levels of sphingolipid and other metabolites including diacylglycerols. Undoubtedly, yeast systems will continue to be exploited to maximum advantage for increasing our understanding of sphingolipid formation and function in all eukaryotic cells.
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ACKNOWLEDGMENTS This work was supported by a VA merit award (LMO), and NIH grants AG16583 (LMO), GM62887 (LMO), GM63465 (YAH), and GM4825 (YAH).
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Chapter 9 EXPLORING AND RESTORING THE p53 PATHWAY USING THE p53 DISSOCIATOR ASSAY IN YEAST
Rainer K. Brachmann Department of Medicine, University of California at Irvine, Irvine, California 92697, USA
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BRIEF INTRODUCTION TO p53 BIOLOGY
For more than a decade, the tumor suppressor protein p53 has been recognized as a, and maybe the, central protein that protects humans from cancer. Its role as “guardian of the genome” [51] and its frequent inactivation in human cancers has drawn numerous researchers into the p53 field, guaranteeing a continuous flow of new research data that is regularly summarized in excellent reviews (see for example, 20, 49, 56, 79, 97, 98; see also the chapter by Inga and colleagues for an in-depth review of p53 biology). Therefore, this introduction will be short, but nevertheless sufficient to appreciate the p53-related questions that are being pursued in the p53 dissociator assay. The p53 protein was initially isolated in 1979 as a 53 kD protein that was associated with SV40 large T antigen [53, 57]. It was a decade before p53 was recognized as an important tumor suppressor because of its frequent mutation in human cancers [38, 52, 73]. This key finding raised two central questions: Why was p53 so important and what was its mode of action? It was quickly established that wild-type p53 exerts its effects, at least in part, as a transcription factor that recognizes its direct target genes through binding of its core domain (amino acids 96–292) to specific DNA sequences. In unstressed cells, p53 is relatively inactive and has a short halflife of 15 min because of negative feedback loops, such as with MDM2. 211 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 217–232. © 2007 Springer.
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p53 drives expression of MDM2, and the Mdm2 protein then binds to the p53 amino-terminus and induces ubiquitin-mediated degradation [55]. Activation of the p53 protein occurs through upstream signals, such as DNA damage, hypoxia, overexpression of oncogenes and others. These signals typically lead to posttranslational modifications of p53, in particular aminoterminal phosphorylation and carboxy-terminal acetylation. These modifycations stabilize and activate p53. p53 is then able to induce its numerous target genes and causes cell cycle arrest, DNA repair, and/or apoptosis [20, 49, 56, 79, 97, 98]. The central role of p53 in protecting humans from cancer is reflected in the enormous number of human cancers with p53 mutations. It is estimated that 50% of all human cancers have p53 mutations, and such mutations have been catalogued in large international databases [5, 18, 32, 36, 37, 62, 77]. The latest update of the IARC TP53 Mutation Database contains 19,809 somatic p53 mutations. 14,123 or 71% of these are missense mutations affecting the p53 core domain, and close to 1,000 different amino acid changes have been described for this p53 region (R9, http://www. iarc.fr/p53/). It has been, and continues to be, a significant challenge to understand the complexity of wild-type p53, p53 cancer mutants and downstream target genes. The chapter by Inga and colleagues summarizes how powerful genetic approaches in yeast assays have made important contributions to characterizing the p53 network. This chapter will focus on two aspects of p53 research that are addressed in the p53 dissociator assay: restoring function to p53 cancer mutants and identifying proteins that play important roles in p53 biology, be it as a p53 inhibitor, a p53 coactivator, or a protein that is regulated by p53.
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THE p53 DISSOCIATOR ASSAY IN YEAST
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The URA3 gene and counterselection in the yeast Saccharomyces cerevisiae
The p53 dissociator assay is based on the principles of yeast experimental systems developed by Fields and others that allow for the study of macromolecular interactions by simple phenotypic readouts [22, 23, 30, 100]. Like these systems, the p53 dissociator assay uses a reporter gene, URA3, activation of which enables yeast cells to grow on media lacking uracil, thus allowing for the selection of a macromolecular interaction. But
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the dissociator assay has a significant difference to the other systems in that it also allows for selection against a particular interaction. This is possible because the URA3 gene product is involved in converting 5-fluoro-orotic acid (Foa) into a toxic substance, thereby arresting cell growth [7]. This twist has been exploited as the “reverse one- or two-hybrid system”, a powerful tool that allows for the selection of novel factors, “dissociators”, able to interfere with a macromolecular interaction [10, 95, 103]. The p53 dissociator assay consists of a tightly repressed reporter construct, 1cUAS53::URA3, typically integrated into the yeast genome, in which a p53 consensus DNA-binding site in the context of the artificially introduced SPO13 promoter is placed upstream of the URA3 gene [11, 95]. p53 binds to its DNA-binding site and activates transcription of URA3. The activity of wild-type p53 can be phenotypically scored in two ways: yeast growth on plates without Uracil (Ura+) and lack of growth on Foa-containing plates, i.e., Foa-sensitive (FoaS).
Figure 1. Principles of the p53 dissociator assay in yeast. The p53-dependent reporter gene URA3 allows for selection against active p53, because URA3 expression results in conversion of 5-fluoro-orotic acid (Foa) into a toxic substance, thus preventing yeast growth. The concept of counterselection can be exploited for the identification of human proteins that reduce p53 activity in the yeast assay. Not only p53 inhibitors score as “dissociators”, but also p53 coactivators and proteins that are regulated by p53. This is likely the case because the human proteins are overexpressed, often amino-terminally truncated (due to the process of the library construction) and evaluated in an artificial yeast assay. Since the p53 dissociator assay relies on native p53 and a consensus p53 DNA-binding site (contrary to, for example, two-hybrid assays for p53), a variety of p53-inhibitory mechanisms can be envisioned: for example, a “dissociator” (“?” in the figure) may conceal the p53 transactivation domain (A), but could also achieve its effect by binding to the p53 DNA-binding site (B).
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The FoaS phenotype of the p53 dissociator assay provides the essential tool to perform cDNA expression library screens (“dissociator screens”). Proteins that interfere with wild-type p53 activity will reduce URA3 expression and allow growth on Foa plates, i.e., Foa-resistant (FoaR). p53 dissociators might utilize a variety of mechanisms, such as binding to and masking the transactivation domain of p53 (Figure 1A) or the actual p53 DNA-binding site (Figure 1B), p53 degradation, or posttranslational modifications of p53. The p53 dissociator assay should clearly identify proteins that negatively regulate p53. But proteins with quite different functions in human cells might be isolated as well, since the yeast assay is different from human cells in many ways. The library proteins are expressed at higher levels than p53 and outside their usual context, thus potentially lacking important regulators of their activity. For example, a p53 enhancer may require a protein complex for its activity in human cells. Missing these important cofactors in yeast, the p53 enhancer may now have the opposite effect by binding to and sequestering p53. Additionally, due to the mode of library construction, most of the cDNA expression plasmids encode for amino-terminally truncated proteins that may behave in a dominant-negative fashion because they lack, for example, a regulatory domain.
2.2
Established p53 inhibitors in the p53 dissociator assay
A useful p53 dissociator assay requires that already known regulators of p53 are able to change the yeast phenotype from FoaS to FoaR. The first validation of the assay came from the protein that led to the identification of p53, SV40 large T antigen. As predicted, this viral protein was able to interfere with p53 activity in the p53 dissociator assay and resulted in Foaresistance [101]. The p53 dissociator assay can also recapitulate more complex situations, such as the interplay of p53, E6-associated protein (E6-AP) and E6 of highrisk human papilloma viruses (HPV). Exposure to high-risk HPV is linked to cervical cancers, and the E6 protein of high-risk HPV has been shown to induce p53 degradation. To this purpose, E6 recruits the human host factor E6-AP, an E3 in the ubiquitin cascade that forms a direct thioester bond with ubiquitin. The trimeric complex of p53, high-risk HPV E6, and E6-AP results in the degradation of p53 via the ubiquitin cascade (Figure 2A) [20, 28, 70, 84, 96]. Expression of E6-AP and high-risk HPV E6 in the p53 dissociator assay conferred the FoaR phenotype. Controls demonstrated that both high-risk HPV E6 and E6-AP were required for this phenotype. The E6 protein from a
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low-risk HPV, unable to cause cervical cancers or to induce p53 degradation, was inactive in the p53 dissociator assay (Figure 2B). Western blot analysis for p53 in the presence of high-risk HPV E6 and E6-AP showed decreased p53 protein levels as compared to the controls suggesting that the two proteins induced degradation of p53 also in yeast (Figure 2C). Experiments with two E6-AP mutants with mutations in cysteine 833 (provided by J. Huibregtse, Rutgers University, Piscataway, NJ) confirmed that the thioester bond between E6-AP and ubiquitin was essential for the FoaR phenotype in yeast (Figure 2D; Brachmann and Boeke, unpublished data). Similar results were obtained with an assay in Schizosaccharomyces pombe that utilized the principle of p53 counterselection through the p53-dependent ura4+ reporter gene [99].
2.3
p53 dissociator screens to identify proteins important for p53 biology
In order to perform large-scale screens, further optimization of the assay to reduce the number of false-positive clones was required. False-positives were mainly due to recessive mutations in the URA3 reporter gene and to a lesser extent in the p53 cDNA. The use of a diploid reporter strain with two copies each of the URA3 reporter gene and the p53 cDNA eliminated most of this undesired background [11, 101]. Thus far, three dissociator screens have been performed. The discussed modifications of the assay resulted in a powerful and very stringent selection system: approximately 1 × 107 yeast transformants resulted in 600 FoaR clones, and one third of those passed all confirmatory tests. Thus, only one in 50,000 plasmids was able to confer an FoaR phenotype (Figure 3) [101]. Besides a large group of proteins without connection to p53 biology in the literature, several established p53-interacting proteins were isolated. This included the known inhibitor of p53, MDM2 [101]. High-risk HPV E6 was also isolated when a screen was performed in the presence of E6-AP with a cDNA expression library from HeLa cells, a cervical cancer cell line known to express E6 and other HPV proteins (see Figure 2; Brachmann, and Boeke unpublished data). Besides p53 inhibitors, the screens also isolated 53BP1, a protein that physically interacts with the p53 core domain and that, in mammalian assays, can enhance p53 activity [43, 44].
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A
B
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D
Figure 2. Recapitulating the biology of p53, E6-associated protein and high-risk human papilloma virus (HPV) E6 in the p53 dissociator assay. Infection with high-risk HPV is associated with more than 80% of human cervical cancers. (A) The E6 protein induces ubiquitin-mediated degradation of p53 through recruitment of the host factor E6-associated protein (E6-AP), an E3 in the ubiquitin cascade whose cysteine 833 forms a direct thioester with ubiquitin (“~S-” in the figure). (B) The complex interplay between two human and one viral protein(s) can be recreated in the p53 dissociator assay, since the combination of E6-AP and high-risk HPV E6 results in an Foa resistant (FoaR) phenotype on plates containing Foa (“+Foa0.1%”). The interference with p53 is highly specific, since low-risk E6 from an HPV unable to cause cervical cancer does not confer Foa resistance. (C) The FoaR phenotype corresponds to reduced p53 protein levels that are seen not only in galactose
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media (high expression of E6-AP), but also in glucose media (minimal expression of E6-AP). (D) E6-AP mutants in which cysteine 833 is changed to alanine or serine suggest that thioester formation between E6-AP and ubiquitin is essential for inhibition of p53. In all experiments (B–D), E6-AP proteins were expressed under the control of the GAL1 promoter that is induced on galactose, “Gal”, but not induced on glucose-containing media. E6 was expressed under the control of the constitutive ADH1 promoter. The two plasmids containing the marker genes LEU2 or TRP1 were selected on media lacking leucine and tryptophane (Leu–Trp). The ADH1-p53 expression cassette was either integrated into the genome at the LYS2 locus (B and D) or maintained on a plasmid with the HIS3 marker gene (C).
Figure 3. Summary of three p53 dissociator screens. The use of a diploid yeast reporter strain with two copies each of the assay components (p53 expression cassette and p53-dependent URA3 reporter gene) results in very stringent screens that allow only 1 in 50,000 plasmids to score with an FoaR phenotype in the p53 dissociator assay.
2.4
The example of hADA3 as a p53 dissociator
hADA3 was one of the first proteins to be isolated in the HeLa p53 dissociator screen. At the time, this human homologue of the yeast protein Ada3p had just been described as a part of histone acetyltransferase (HAT) complexes containing the human HATs hGCN5 or PCAF [76]. The yeast gene ADA3 had previously been identified because its inactivation protected Saccharomyces cerevisiae from the toxic effects of an overexpressed artificial transcription factor, the GAL4–VP16 fusion protein [78]. Subsequent work established that yeast Ada3p physically interacts with Ada2p and the yeast HAT Gcn5p, forming HAT complexes termed Ada or SAGA (if additional proteins are present) [29]. These HAT complexes acetylate histones thereby relaxing tightly packed chromatin and giving transcription factors access to promoters. They also appear to regulate other proteins by acetylating them. Both Ada2p and Ada3p interact with the transactivation domains of transcription factors, and the whole Ada complex is required for transcriptional activity of many transcription factors [29, 60, 89, 90].
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Interestingly, prior to any studies of hADA3, work by Candau and colleagues had already suggested that Ada proteins are important for transcriptional activity of p53 in yeast [17]. They took advantage of the yeast Ada complex to further dissect the transactivation domain of p53. In this context, they showed that gene knockouts of ada2, ada3, and gcn5 resulted in loss of transcriptional activity of p53 in yeast. However, it was not further explored why the Ada complex was required for transcriptional activity of p53. The characterization of full-length hADA3 in human cells by us indicated that hADA3, very much like its yeast counterpart, serves as an adaptor or docking protein that allows transcription factors to communicate with HAT complexes. In the case of p53, the interaction with hADA3 appears to be very tightly regulated. Although a basal interaction between p53 and hADA3 can be detected, p53 that is activated by upstream stress signals shows a dramatically enhanced interaction with p53. This enhanced interaction depends on amino-terminal phosphorylation of p53. Carboxyterminally truncated hADA3 acts as a dominant-negative protein and abrogates p53 transcriptional activity and p53-mediated apoptosis. Not only does hADA3 interact with activated p53, but it is also required for p53 activity following stress stimuli, since absence of hADA3 (through antisense approaches) results in significantly reduced p53 transcriptional activity following DNA damage [101]. Thus, the p53 dissociator assay identified a new coactivator of p53 whose further characterization suggested novel mechanisms of blunting the p53 pathway. For instance, mutations that truncate hADA3 may result in dominant-negative proteins that prevent access of p53 to HAT complexes and, thus, full activation of p53. This hypothesis is readily testable by screening human cancer specimens for hADA3 mutations and aberrant hADA3 protein sizes. A knockout mouse model for ADA3 will be another important strategy to assess the significance of ADA3 for the p53 pathway. It was recently shown that high-risk HPV E6 not only targets p53 for degradation, but also hADA3 [50], suggesting that removal of hADA3 is an important step in HPV-mediated carcinogenesis.
2.5
p53 dissociator screens to characterize specific p53 inactivations in cancers
Considering the central role of p53 and its frequent inactivation through p53 gene mutations, it is very likely that the remaining human cancers with wild-type p53 must significantly reduce the activity of the p53 pathway. But compared to the detailed knowledge regarding p53 mutations, much less is
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known about the p53 status of the remaining 50% of human cancers with no p53 mutations. One obvious way of eliminating the p53 pathway would be to abrogate p53 gene expression. This is indeed the case for a subset of breast cancers. Raman and colleagues determined that the transcription factor HOXA5 is important for expression of the p53 gene [81]. In breast cancers with loss of p53 expression, the promoter of the HOXA5 gene was frequently methylated resulting in lack of HoxA5 expression and HOXA5 protein. It is currently unknown whether this mechanism is found in other cancers as well. A second efficient mode to eliminate the p53 pathway is direct inactivation of the p53 protein. There are two well-documented examples: a large fraction of sarcomas, as well as a small percentage of other human cancers show overexpression of MDM2, thus turning the physiological negative feedback loop for p53 into a tool to rid the cell of p53 activity [20, 28, 56, 59, 68, 96]. Also, high-risk HPV cause more than 80% of cervical cancers, and the HPV E6 protein is responsible for inducing the degradation of p53 [20, 28, 70, 84, 96]. Firm numbers are not available, but it is unlikely that the majority of human cancers with wild-type p53 utilize inactivation of p53 by high-risk HPV E6 or MDM2 overexpression. There are two additional ways of inactivating wild-type p53 protein whose underlying molecular mechanisms have not yet been fully elucidated. A variety of human cancers show complete or partial mislocalization of p53 to the cytoplasm. This has been described for primary tissues and/or cell lines of breast cancers, colon cancers, neuroblastomas, malignant, melanomas and retinoblastomas [8, 9, 65, 67, 85, 91, 102], and impairs p53’s ability to induce G1 arrest and cause growth inhibition [48, 66]. Several underlying genetic alterations can be envisioned, including loss of factors required for active import of p53 into the nucleus (Figure 4A, I) and overexpression or constitutive activation of factors that actively hold p53 in the cytoplasm (Figure 4A, II). The latter model is supported by indirect evidence from several studies that suggest that an unidentified short-lived anchor protein might tether p53 to the vimentin scaffold [26, 47, 48] and the identification of a cytoplasmic anchor protein, Parc, for p53 [45, 74]. Stommel and colleagues provided strong support for a third possibility, inappropriate nuclear export of p53 [88]. They showed that disruption of p53 homo-tetramers results in exposure of a nuclear export signal in the carboxy-terminus. Inappropriate activity of the nuclear factor(s) involved in this mechanism could thus result in excessive nuclear export of p53 (Figure 4A, III).
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Figure 4. p53 dissociator screens to elucidate specific modes of wild-type p53 inactivation in human cancers. (A) A significant percentage of human cancers carry wild-type p53 that is mislocalized to the cytoplasm, thus rendering it inactive. Several mechanisms may underlie this finding. I. p53 may be imported into the nucleus inefficiently. II. p53 may be actively held in the cytoplasm. III. p53 may be excessively exported from the nucleus because of inappropriate exposure of a normally hidden nuclear export signal (NES) in the region of the tetramerization domain. (B) Testicular cancers carry wild-type p53 that is stabilized and transcriptionally inactive, suggesting the presence of a p53-inhibitory protein in the nuclei of these cancers. Targeted p53 dissociator screens with cDNA expression libraries representative of these p53 inactivations should be able to decipher the underlying molecular mechanisms.
Testicular cancers have a distinctly different phenotype of p53 inactivation. They typically have elevated p53 protein levels, yet the p53 genes do not show mutations [69]. Based on work with murine teratocarcinoma cell lines [61], wild-type p53 in this type of cancer is not transcriptionally active. But the inactivation can be overcome through DNA damage or induction of differentiation with retinoic acid. These findings are highly suggestive of the presence of a protein that sequesters and functionally inactivates p53 in the nucleus (Figure 4B).
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The p53 dissociator assay provides an excellent tool to perform targeted dissociator screens with cDNA expression libraries of cell lines that reflect the two specific inactivations of p53.
3
INTRAGENIC SUPPRESSOR MUTATIONS AND THE QUEST TO ENDOW FUNCTION TO p53 CANCER MUTANTS
3.1
Strategies to restore p53 function to human cancers
Because of its central role in tumor suppression, p53 and its pathways have been recognized as a prime target for developing new cancer therapies [13, 24, 27, 35, 40, 46, 58, 64], and various strategies have been pursued. Wild-type p53 may be delivered through gene therapy approaches regardless of the p53 status of the cancer [3, 19, 71, 83, 92, 105]. Cancer cells lacking p53 function can be specifically targeted with mutant adenovirus [6, 63] or adeno-associated virus [80]. Inactive wild-type p53 and some p53 cancer mutants may be reactivated through targeting of the very carboxy-terminal putative autoregulatory domain of p53 with antibodies [1, 33, 41, 72] or peptides spanning part of this region [2, 42, 86]. If MDM2 overexpression is responsible for p53 inactivation, the inappropriate interaction between MDM2 and p53 may be disrupted by peptides and compounds [21, 40, 87]. The pursuit of this strategy has resulted in very potent small-molecule inhibitors of MDM2 [54, 94]. Yet, all these strategies have their limitations, such as lack of efficient local and/or systemic delivery, undesirable responses of the host immune system to the therapeutic agent, lack of specificity for the cancer cell and/or targeting of a small subset of human cancers. Besides these strategies, one approach appears particularly appealing: small chemical compounds that endow p53 cancer mutants with function by restoring the integrity of the p53 core domain. Such compounds are highly desirable, because of the high frequency of p53 mutations and the large number of patients who could potentially benefit. It has been estimated that every year approximately 360,000 patients in the USA and 2.6 million patients worldwide are diagnosed with cancers that contain p53 mutations [34, 35]. This subset of human cancers (including lung, prostate, colorectal, breast, head and neck, pancreatic, and gastric cancers) is often resistant to conventional therapies and difficult to treat at advanced stages [5, 18, 24, 31, 35, 38, 58].
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The strategy is, at least in theory, possible because of the unique pattern of p53 missense mutations in human cancers. Typically, p53 cancer mutants are full-length proteins with an intact transactivation domain and carboxyterminal tetramerization domain. The problem is the altered core domain (amino acids 96–292) whose structural integrity needs to be restored in order to have functional p53. Compounds able to achieve this goal are predicted to be specific for the cancer cells, since the structurally intact wild-type p53 core domain of surrounding normal cells should not be affected. Furthermore, p53 cancer mutants are typically present at very high levels (thus providing a large drug target), since their lack of transcriptional activity abrogates negative feedback loops, such as with MDM2 [55]. The approach also has the advantage of combining systemic delivery with lack of a host immune response. The attractive features of this strategy are easily rivaled by the enormous challenges it poses: such compounds may simply not exist, or, if they do, they may rescue only a very small fraction of the close to 1,000 different known p53 cancer mutants that have one amino acid change in the core domain. On a more optimistic note, looking at missense mutations of the p53 core domain (71% of the database entries; R9, http://www.iarc.fr/p53/), the 50 most frequent amino acid changes account for 55% of the total number, and of these 50, the 8 most common amino acid changes comprise the majority of them (30% of the total). Thus, chemical compounds that rescue a subset of these p53 mutants are likely to correspond to a large number of cancer patients. Also, as a proof of principle, two classes of compounds have already been identified in large drug screens that appear to have the desired characteristics. The first screen utilized an antibody-based screening strategy that relied on partially unfolded wild-type p53 due to elevated temperatures. The isolated compounds appeared to partially restore function to a few p53 cancer mutants [25]. However, further characterization of the lead compound CP-31398 has thus far not established the exact mechanism of rescue. In addition, CP-31398 appears to have strong p53-independent activities [13, 16, 82, 93]. The second class of compound was isolated using a mammalian growth suppression assay [16]. Very impressively, this compound rescued the most common p53 cancer mutant R175H whose core domain is very likely entirely denatured [13–16, 106]. It still remains to be determined how many others of the most common p53 cancer mutants will be rescued by this compound and what the exact mechanism of rescue is.
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Intragenic suppressor mutations for p53 cancer mutants
The p53 yeast dissociator assay has allowed a broader look at the challenge of rescuing p53 cancer mutants; it has been used for identifying intragenic suppressor mutations that overcome the deleterious effects of cancer mutations. Such suppressor mutations are highly informative, as they identify key regions within the p53 core domain that, upon alteration, provide increased stability to the core domain. Such suppressor mutations have also been pursued by computer modeling, but this strategy generated only one successful suppressor mutation and several false predictions [104]. Additionally, “suppressor mutation design” was not as wide in scope as genetic screens in yeast.
Figure 5. Search for intragenic suppressor mutations that endow function to common p53 cancer mutants. (A) Starting with a common p53 cancer mutant (with mutation “C”) that is transcriptionally inactive (Ura– phenotype), (B) the up- or downstream regions of the cDNA are mutagenized using PCR mutagenesis. (C) In a separate reaction, the cDNA expression plasmid is gapped in the same region. (D) Both products are transformed into yeast, and the overlap between PCR product and gapped plasmid allows the yeast to repair the plasmid by homologous recombination. After confirmatory experiments, the plasmid is eventually sequenced to identify the suppressor mutation(s) (“S”).
Screens for suppressor mutations in the p53 dissociator assay take advantage of PCR mutagenesis and gap repair in yeast through homologous recombination. Starting with a p53 cancer mutant that is Ura– (Figure 5A), the regions up- and downstream of the cDNA are PCR-amplified under mutagenic conditions (Figure 5B). The expression plasmid for the p53
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cancer mutant is gapped, so as to create an overlap with the PCR products (Figure 5C). PCR products and gapped plasmids are then transformed into a yeast reporter strain that repairs the plasmid by homologous recombination utilizing a PCR product (Figure 5D). A pilot study identified several suppressor mutations for three cancer mutations: N268D for V143A, T123P, N239Y and S240N for G245S and T123A in combination with H168R for R249S [12]. The suppressor mechanism for H168R appeared to be very specific to R249S [12, 75], but two other isolated suppressor mutations, N268D and N239Y, increased the overall stability of two cancer mutants, G245S and V143A. This suggested that, in principle, small molecules should be able to bind to and stabilize the core domain of such p53 cancer mutants in a similar manner [75].
3.3
Identification of a global suppressor motif for p53 cancer mutants
The initial findings with the p53 dissociator assay were very promising, yet several technical limitations needed to be overcome to allow for a systematic analysis of the most common p53 cancer mutants. To this end, an engineered open reading frame for p53 was constructed [39] that contained numerous silent restriction sites [4]. Analysis for the eight most common p53 cancer mutants (30% of core domain missense mutants) identified suppressor mutations for four of them (G245S, R249S, R273C, and R273H; 12% of core domain missense mutants). Several combinations of suppressor mutations were identified, and, intriguingly, N239Y or S240R were involved in the rescue of all four p53 mutants [4]. This suggested that amino acids 239 and 240 may be part of a global suppressor motif, a hypothesis that was further pursued by oligonucleotide-mediated mutagenesis of a small region of the core domain, codons 225–241. The mutagenesis included directed changes of codons 239 and 240 to all other 19 possible amino acids plus a background mutagenesis resulting in approximately 1 in 100 nucleotide misincorporations. This strategy resulted in the rescue of 16 out of 30 of the most common p53 mutants tested, representing 18% of all reported p53 core domain missense mutants. As expected, the suppressor mutations included changes of codons 239 and/or 240, but, in addition, changes of amino acid 235 to lysine were also frequently found. The resulting p53 mutants were not only transcriptionally active in yeast, but had also activity in mammalian reporter gene and apoptosis assays [4]. These findings strongly argue that manipulation of a small region of the p53 core domain is sufficient to endow p53 cancer mutants with function. This region therefore comprises a global suppressor motif. Structural studies
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of this motif will determine the exact rescue mechanism and provide the basis to exploit this motif further by designing small compounds able to recreate the suppressor mechanism (Figure 6). By restoring function to p53 cancer mutants, these small molecules would, in effect, reactivate a major apoptotic pathway and “relicense” the cancer cell to kill itself.
Figure 6. Strategy for the development of a new anticancer therapy that aims to endow p53 cancer mutants with wild-type function. (A) A pilot study established that the concept of and search for intragenic suppressor mutations was a viable strategy to achieve a better understanding of how mutant p53 core domains can be stabilized. (B) The subsequent comprehensive screen using a new engineered p53 open reading frame identified amino acids 235, 239, and 240 a part of an important global suppressor motif that rescued 16 of the 30 most common p53 cancer mutants. (C) Crystal structures for p53 cancer mutants and p53 suppressor/cancer mutants will establish the underlying mechanism that allows the 235–239– 240 suppressor motif to rescue p53 cancer mutants. (D) This information will lay the foundation to pursue the most challenging aspect of the project: design, identify, and characterize small compounds able to recreate the rescue mechanism of the suppressor motif. This step requires a multidisciplinary approach that combines the expertise of computer scientists, chemists, structural biologists, and molecular biologists.
4
CONCLUSIONS
Baker’s yeast or S. cerevisiae is a highly valuable research tool for studies of the human tumor suppressor protein and transcription factor p53. Reporter gene assays for p53 in yeast have made important contributions to our understanding of this central tumor suppressor protein, and the chapter by Inga and colleagues provides an excellent overview on these efforts. This chapter highlighted the specific contributions of the p53 dissociator assay. The hunt for intragenic suppressor mutations illustrates that, similar to other p53 yeast assays, it is able to efficiently analyze and select for p53 activity. But the p53 dissociator assay also provides additional genetic strategies in
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yeast, as it exploits the concept of selection against p53 activity, a twist that allows for powerful screens for proteins important to p53 biology. As the biology of p53 is bound to become more and more complex, p53 yeast assays will continue to contribute to our understanding of this crucial tumor suppressor protein. The focus of this review was p53, but many of the discussed concepts can be readily applied to the studies of other fundamental cellular mechanisms. Undoubtedly, many human proteins, particularly transcription factors, are just waiting to be plugged into a yeast assay of their own.
ACKNOWLEDGMENTS R.K.B. thanks Carrie Baker Brachmann for critically reading the manuscript and Jef D. Boeke whose vision and support were instrumental for the development of the p53 dissociator assay.
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103. White, M. A. 1996. The yeast two-hybrid system: forward and reverse. Proc. Natl. Acad. Sci. USA 93:10001–10003. 104. Wieczorek, A. M., J. L. Waterman, M. J. Waterman, and T. D. Halazonetis. 1996. Structure-based rescue of common tumor-derived p53 mutants. Nat. Med. 2:1143–1146. 105. Wolf, J. K., G. B. Mills, L. Bazzet, R. C. Bast, Jr., J. A. Roth, and D. M. Gershenson. 1999. Adenovirus-mediated p53 growth inhibition of ovarian cancer cells is independent of endogenous p53 status. Gynecol. Oncol. 75:261–266. 106. Wong, K. B., B. S. DeDecker, S. M. Freund, M. R. Proctor, M. Bycroft, and A. R. Fersht. 1999. Hot-spot mutants of p53 core domain evince characteristic local structural changes. Proc. Natl. Acad. Sci. USA 96:8438–8442.
Chapter 10 FUNCTIONAL ANALYSIS OF THE HUMAN p53 TUMOR SUPPRESSOR AND ITS MUTANTS USING YEAST
Alberto Inga, Francesca Storici and Michael A. Resnick Laboratory of Molecular Genetics, National Institute of Environmental Health Sciences, NIH, P.O. Box 12233, Research Triangle Park, NC 27709
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FUNCTIONAL ALTERATION OF p53: A HALLMARK OF CANCER
Multiple perturbations of the complex network of signaling pathways that define the precise role of a cell in its tissue microenvironment and that regulate stress responses are accumulated through genetic and epigenetic changes during processes of transformation that lead to cancer [129, 201]. In particular, the acquisition of self-sufficiency in growth signal and the resistance to growth inhibitory and apoptotic signals are likely to be required for initiation and early progression [77, 174]. Limitless replicative potential, associated with the selection of canonical or alternative ways of maintaining the stability of chromosome ends, the ability to induce angiogenesis and to acquire invasive potential are additional steps in the transformation process that could lead to a tumor [77]. Stepwise and orderly accumulation of functional changes at protooncogenes and tumor suppressors has been associated with different stages of cancer in specific types of tissue [70, 129]. However, accumulating evidence of the multiple levels of complex interactions between survival, proliferation, and growth restraint pathways suggest that the underlying genetic alterations may be codependent and specific combinations of changes may be selected according to the microenvironment and the genetic/ epigenetic makeup of the transforming cell [212]. 233 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 233–288. © 2007 Springer.
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The tumor suppressor gene p53 plays a major role in cellular response to various environmental stresses [107, 114]. The p53 protein is a sequencespecific transcription factor that can lead to transactivation of over 50 genes, many of which are involved in apoptotic or cell cycle arrest responses (Figure 1). Its role in biological responses may also relate to the ability to interact with a vast number of proteins [164]. Functional alterations in the p53 pathway likely occur in nearly all human cancers. In almost half the human malignancies, there is a mutation in the p53 gene itself [71, 79]. Interestingly, ~80% of p53 mutations are missense changes that lead to single amino acid substitutions, a feature that distinguishes p53 from other tumor suppressor genes (e.g., APC, NF1, BRCA1) [22, 67, 155]. The incidence of p53 mutations and the types of mutations can vary among tumors in different tissues or populations [71]. Yeast provides unique advantages for finding and characterizing human p53 mutations.
Figure 1. The tumor suppressor p53 is a sequence-specific transcription factor. Many types of stress induce signal transduction pathways that lead to p53 stabilization and activation, mainly by posttranslational modifications. The p53 protein is shown as a tetrameric (dimer of dimers) sequence-specific transcription factor that recognizes promoter response elements (RE). The solid arrows indicate the organization of the RE sequence as a closely spaced pair of
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dimer-binding sites each consisting of two monomer-binding sites in inverted orientation (see text for details). A list of the principal p53 target genes and their functions is shown.
In this chapter we first describe the role of p53 in mammalian cells and the approaches that are commonly used to identify and evaluate the consequences of p53 mutations. This is followed by a broad discussion of the many yeast-based assays that have been developed for investigating p53.
1.1
Anatomy of the p53 protein and p53 tumor mutations
The human p53 protein consists of 393 amino acids. Starting from the amino terminus (Nter), the following functional domains have been identified: 2 transactivation domains, a proline-rich sequence likely involved in protein–protein interactions, a DNA-binding domain, a tetramerization domain and a basic carboxy-terminal (Cter) region with regulatory functions [189] (see Figure 2). The protein contains three nuclear localization signals toward the Cter and one or possibly two nuclear export signals [107, 219].
Figure 2. p53 structural organization, topological domains, functional regulation by posttranslational modifications and location of tumor mutations.
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a
Functional domains. TA I and II = acidic transactivation domains; amino acids 22 and 23 are required for TA I while 53 and 54 are crucial for TA II. One nuclear export signal NES whose activity is influenced by the phosphorylation status of p53 overlaps with TA I (220). PxxP = proline-rich domain with possible role in protein–protein interactions [175]. TD = tetramerization domain; NLS = nuclear localization signal; RR = basic regulatory region. b Conserved domains. The positions of the 5 evolutionary-conserved regions along the 393 p53 amino acid sequence are indicated. Four of them correspond to sequences within the sequence-specific DNA-binding domain. With the exception of domains I and II, the major p53 hotspots are located in the conserved domains. Presented is the distribution of the tumor p53 mutations along the protein (hotspots are indicated by the number in the graph). Data from the R5 release of the IARC p53 mutation database (containing more than 15,000 mutations; http://www.iarc.fr/P53/Somatic.html) were used to generate the graph. Posttranslational modifications. The numbers indicate the amino acids at which posttranslational modifications have been described [5]. The p53 sequence-specific DNAbinding domain does not appear to be modified. P = phosphorylation; Ac = acetylation; Ubi = ubiquitination (sumolation of lysine 386 is also shown [173]). Phosphorylation of p53 residues at the Nter might affect interaction with acetylases and acetylation of p53 (dotted arrows). Acetylation of lysines in the p53 Cter inhibits Mdm2-mediated ubiquitination. The serine/threonine protein kinases that can phosphorylate p53 are also indicated above the posttranslational modification sites.
A database of p53 mutations reported in tumor samples and cell lines has been developed and is updated yearly at the International Agency for Cancer Research (IARC), Lyon, France [75, 79]. Most (~80%) of the p53 mutations associated with cancer occur as missense changes in the sequence-specific DNA-binding domain that comprises about half of the entire protein sequence (amino acid 100–300). This suggests that most tumor-associated changes are likely to affect the ability to transcribe p53 responsive genes. The unusually high frequency of missense changes in the p53 mutation spectrum has provided opportunities in the field of molecular epidemiology to investigate the role of carcinogens as etiologic agents of cancer [83]. The similarity between p53 mutational fingerprints in tumors among exposed populations and mutational fingerprints obtained following in vitro treatment with an agent has been used to indicate its role in the etiology of some cancers [15, 78] (discussed in section 2). Germline p53 mutations lead to the dominant tumor-susceptibility Li– Fraumeni syndrome (LFS) characterized by a high incidence of various cancers with clear tissue-specific effects that in some cases relate to the nature of the p53 mutations in terms of impact on protein function [74]. The germline p53 mutation spectrum is similar to the somatic tumor spectrum. An LFS-like phenotype has also been identified in families that do not harbor germline p53 defects. Mutations in the checkpoint kinase Chk2 gene, an upstream regulator of p53 activity, have been observed in these families suggesting that the p53 pathway can be altered in multiple steps [112]. Alterations in the upstream regulators or downstream effectors of p53 may lead to a hypomorphic functional state of the protein that could impact
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on the selective pressure for additional p53 gene mutations during cancer development. An issue that has received relatively little attention is that partial inactivation or changes in the spectrum of p53 functions may be sufficient for, or even favor, tumor progression depending on the cell type and physiological state (see sections 1.4, 2.1.3, and 2.5). This is relevant to the prognostic use of tumor p53 status to estimate tumor aggressiveness and therapeutic responses. Although reports differ possibly also because of lack of standardization, p53 status can be a prognostic factor based on multivariate analyses, with loss-of-function p53 mutations correlating with poor prognosis [34, 51, 120, 124, 153, 181, 188]. The analysis of p53 mutation spectra from tumors suggests that much of the p53 gene is a target for missense mutations that can inactivate its function. In fact, more than 1,300 different amino acid changes have been identified in the protein domain that provides for sequence-specific DNA binding [126]. Although there are several strong hotspots, the majority of tumor-associated p53 alleles do not occur at hot spot codons [79] (Figure 2). Information on the specific functional defects associated with p53 mutations, particularly at non-hot spot codons, is limited. These functional defects could range from total loss of transactivation potential to partial loss that affects only a subset of p53 target genes, to subtle differences in activity compared with the wild-type (WT) protein, and also to enhanced and altered DNA binding specificity leading to gain of functions (see sections 2.1.3, 2.5, and 2.8). The crystal structure of the human p53 DNA-binding domain and the comparison of p53 sequences among many species show that the most frequent mutations affect amino acids directly involved in establishing DNA contacts and that hot spots tend to occur at evolutionary conserved residues [36, 209]. The p53 DNA-binding domain has an immunoglobulin-like fold. Three loops are involved in DNA contacts, and hydrophobic interactions of two beta-sheets plus the coordination of a zinc atom provide for correct positioning of the loops. This conformation may explain why many single amino acid changes would affect the activity of the protein, even if distant from the DNA contact sites. However, structural information for the complete protein is not available, which limits structure/function analysis. Several tumor-associated mutations affect amino acids that belong to the DNA-binding domain but do not face the DNA [72]. Since the active form of p53 is a tetramer (a dimer of dimers) (Figure 1), these mutations might affect protein–protein interactions that stabilize the tetramer and indirectly lead to inefficient DNA contacts [130]. The observation that p53 is active as a tetramer has led to the hypothesis that the selection for missense changes in the DNA-binding domain can be explained by the capacity of the mutant proteins to form hetero-tetramers
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with WT p53, leading to dominant-negative effects [24]. The extent, the mechanisms and the physiological relevance of this dominant-negative potential are still under debate, especially since the WT allele is generally lost [8, 19, 59]. It is possible that the requirement for WT p53 activity, a specific partial function, or complete loss-of-function may change during the evolution of a tumor. Notably, many tumor p53 mutant proteins are present at high levels in tumor cells and some p53 mutants acquire oncogenic attributes [73, 162].
1.2
Biochemical functions of the p53 protein
Different activities and many protein–protein interactions have been ascribed to the WT p53 protein [164]. Some of these suggest a direct role for p53 in preserving genome stability, particularly in the regulation of homologous recombination and modulation of DNA repair capacity [3, 47, 111, 176]. However, mounting evidence indicates that the main p53 function in tumor suppression results from its activity as a rapidly inducible, sequence-specific transcription factor [206, 208] (see Figure 1). 1.2.1
Regulation of the p53 transcription factor
In normal unstressed cells, p53 is expressed constitutively but the protein has a short half-life. Various types of stress signals, including proliferative stimuli, viral infection, nucleotide pool unbalance, hypoxia, with DNA damage being the best characterized, lead to p53 stabilization and activation. These are achieved mainly through posttranslational modifications, comprising phosphorylation, acetylation, and sumolation in the Nter and the Cter regions [7, 69, 154]. Interestingly, the large sequence-specific DNA-binding domain appears to be largely excluded from posttranslational modifications (see Figure 2), with the exception of proline isomerization [227]. The precise effect of these changes is not completely understood. Multiple Nter serine/threonine phosphorylations (~10 sites) can lead to a change in the relative binding affinity of p53 toward negative (e.g., MDM2, an E3ubiquitin ligase that targets p53 for proteasome-mediated degradation) and positive (e.g., p300/CBP, a histone acetylase) regulatory factors, or basal elements of the transcription machinery (e.g., TAFII40, 60, 31) [154]. This results in increased availability of nuclear p53 protein and stimulation of transcription [5, 165]. p53 activity is tightly regulated [7, 68] (see Figure 2). Different types of stress including DNA damage may lead to specific phosphorylations of p53 via different upstream protein kinases. This provides flexibility in p53 abundance, localization, and DNA-binding activity. Notably, the regulation
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of p53 activity also involves one major, and possibly more, negative feedback loop(s). The MDM2 gene is directly activated by p53 [163]. The importance of this negative regulation is exemplified by the oncogenic potential of Mdm2 gene amplification and by the observation that Mdm2 knockout (KO) mice are embryonic lethal and exhibit rapid and massive apoptosis, while the double knockout with p53 is viable [135]. 1.2.2
Transactivation of p53 responsive genes
The induction of p53 responses can lead to different biological effects according to the cell type or the activating stimuli. For example, temporary G1 or G2 cell cycle arrest, premature senescence, and programmed cell death can all be induced (or maintained) by p53 [208]. The mechanisms of how the choice between these pathways is brought about are still elusive, although several possibilities have been proposed. Differential regulation of downstream genes, both in terms of the extent and the kinetics of transactivation and repression, is likely to be an important factor for dictating the specificity of p53 biological responses. Locus-specific features of chromatin assembly and remodeling are proposed to affect the transcriptional status of specific genes [110]. Possibly, the combinatorial action of different transcription factors, coregulators and corepressors provides for specific gene regulation and could be the basis of cell-type specific effects of p53 responses. Over 100 genes can be modulated (both activation and repression) by p53, as revealed by genome-wide expression profiles [220]. In vivo binding of the p53 protein to specific promoter sequences has been demonstrated for a limited number of p53-regulated genes using chromatin immunoprecipitation (ChIP) assays [12, 101]. For other p53 targets, sequence response elements that are bound in vitro by p53 in gel shift assays have been identified in promoter or intronic regions [49]. These sequences are sufficient for transcripional activation by p53 of reporter genes during transient transfection of mammalian cells. A degenerate p53 consensus response element (RE) has been identified that contains two DNA repeats each of which contains a head-to-head inverted sequence (5′RRRCWWGYYYN(0-13)RRRCWWGYYY-3′ where R = purine, Y = Pyrimidine; W = A or T) [50]. The single inverted repeat can bind a p53 protein dimer, so that the two inverted repeats provide for the binding of a p53 tetramer (see diagram of a p53 tetramer bound to DNA in Figure 1). There are many variations which are expected to affect intrinsic RE binding affinity and differential transactivation. The p53 dimer-binding sites can be separated by up to 13 nucleotides with neutral sequence, and p53 REs identified in regulatory regions of p53-target genes can deviate considerably
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from the consensus sequence. Also, there are variations in the number or repeats, position relative to the ATG start site, and the arrangement of REs. In vitro DNA-binding experiments with naked DNA indicate that the ability of p53 to act as a transcription factor depends on posttranslational modifications that activate the protein, enabling it to bind DNA [82]. In particular, Cter modifications have been proposed to relieve its inhibitory effect by triggering a conformational switch. Models whereby covalent modifications activate p53 DNA-binding and transactivation function are consistent with the observation that p53 is induced by stress responses. Moreover, different posttranslational modifications might have differential effects on p53 activity, thus providing a direct link between a particular activating signal and a specific response [48]. Conclusions from gel shift assays, however, should be taken with caution since it is questionable how reliably this in vitro assay reproduces the nuclear environment. Recently the role of chromosome structure in p53 transactivation potential has been examined in vitro using chromatinized DNA preparations and in vivo using ChIP assays [12, 52, 101, 196]. Interestingly, acetylation of p53 does not appear to affect its DNA-binding affinity, yet the histone acetylase activity of p300/CBP is important in p53-mediated transactivation [12]. It is possible that by virtue of its specific DNA-binding capacity and protein–protein interaction with p300/CBP, p53 can stimulate chromatin modifications at promoter sites that stimulate transcription. The acetylation of p53 might be a by-product of these molecular interactions [165]. Thus, p53 intrinsic DNA-binding affinity might be an important component in the regulation of p53-dependent transcriptional modulation. Since p53 is active as a tetramer, its intranuclear concentration may strongly affect binding kinetics and transactivation. The importance of p53 levels is often overlooked (discussed below), especially for mutants with altered function, since assays in mammalian cells usually involve high expression. In fact, both the chromatin structure of the reporter plasmid and the high levels of p53 protein expressed from constitutive viral promoters result in assessments of transactivation activity under conditions that are not physiological.
1.3
Identification of p53 mutations in tumor samples
Several techniques have been used to determine p53 status in fresh, frozen, or archived tumor specimens [132]. These include immunohistochemistry (IHC), DNA sequencing, single-stranded conformational polymorhphism (SSCP), denaturing gradient gel electrophoresis (DGGE), loss of heterozygosity (LOH), and functional assessment in yeast. The levels
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of p53 expression in individual cells from tumor sections can be determined by IHC. Since tumors are generally highly heterogeneous, single cell microscopic analysis is useful. High expression of p53, which is detectable by antibody staining in individual cells, can be indicative of p53 mutations. p53 missense mutants often lack the control of downstream pathways which regulate p53 levels, as is the case for p53 mutants defective in MDM2 transactivation. However, IHC positivity has only limited correlation (~60%) with the appearance of p53 mutations [10, 66, 80, 131]. Methods are also available for the direct detection of mutations in the p53 gene based on the conformational changes they produce in isolated DNA fragments. However, these are limited by the heterogeneity of cells within tumors with respect to p53 status. These electrophoresis-based approaches (SSCP, DGGE) are time-consuming, since short PCR-generated DNA fragments corresponding to at best a single exon are examined separately in the gels. Fragments with altered mobility can be detected only when a mutant allele is present in at least 10–20% of the cells in the specimen [29, 81]. Given the high frequency of p53 mutations in tumors and the modest size of the p53 gene, direct DNA sequencing has been used as an alternative method. This approach has, however, the same sensitivity issues as SSCP and DGGE and is relatively time-consuming. Recently, an oligonucleotidebased array (p53 GeneChip) was developed that contains almost all possible p53 point mutations, but not deletions/insertions. DNA fragments obtained by multiplex PCR of p53 exons from tumor samples are fragmented, fluorescein-labeled and then hybridized to the chip. p53 mutations are detected and identified by the presence and position of hybridization signals that differ from WT sequences. This method is very sensitive (detecting as few as 1–2% of mutant alleles in the cell population) and can also be applied to microdissected tumor sections [2, 121, 213]. However, the high cost of this approach is a limiting factor. An arrayed primer extension (APEX) assay for detection and identification of p53 mutations has also been developed recently [203]. Similar to the GeneChip, a DNA sample is amplified, fragmented enzymatically, but it is then annealed to arrayed primers on a chip. There are two primers to probe p53 coding nucleotides (both sense and antisense strands). Four fluorescently labeled dideoxynucleotides are used for template-dependent DNA polymerase extension reactions, and mutants are detected as novel extension reactions. One limitation which is common with all the methods described above is that they do not provide information on the functional consequences of a given amino acid change in the p53 protein.
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Characterization of p53 alleles that retain function in mammalian cells
Expression of p53 alleles under constitutive viral promoters from expression vectors in mammalian cell lines have provided means to evaluate the tumor suppressor potential of WT and mutant forms of the protein in different cell types. Generally, ectopic expression in tumor cell lines of WT p53 leads to growth suppression through cell cycle arrest and apoptosis. Loss-of-function p53 mutants usually lose this activity. However, tumorassociated amino acid changes in the p53 DNA-binding domain do not always result in complete loss of growth suppressing activity and biological function. For example, some p53 mutants retain the ability to induce cell cycle arrest, but cannot induce apoptosis when expressed in tumor cell lines. This biological effect correlates with the ability to induce the cyclindependent kinase inhibitor p21 but not the apoptotic gene bax [19, 122]. Transactivation activity of p53 mutants in human cells can also be assessed using luciferase reporter assays in transient transfection. Other p53 mutants that are temperature-sensitive, show loss-of-function at 39°C but normal or partial activity at 32°C [116, 161]. Some p53 mutants acquire oncogenic gain-of-function features such as the ability to deregulate chromosome segregation, upregulate genes conferring resistance to chemotherapeutic agents and promote angiogenesis [20, 58, 64, 142]. A group of tumor-associated mutants was identified that appeared to retain WT levels of activity toward growth suppression, apoptosis, and transactivation [187]. Interestingly, they were observed in familial cases of breast cancer associated with BRCA1 germline mutations where the incidence of p53 mutations is much higher than those found for sporadic breast cancer [72]. Furthermore the spectrum of p53 mutations associated with familial breast cancer is different from that of sporadic breast cancer. The actual functional status of p53 alleles, not simply whether or not there is a mutation, may correlate with clinical features of a tumor and represent a relevant prognostic marker for patient management in conventional chemo- and radiotherapy protocols. There are, however, severe limitations on the functional analysis of p53 mutations in mammalian cells including, but not limited to, the nonphysiological high level of ectopic expression of p53 obtained with viral promoters.
1.5
Cancer-therapy approaches based on p53 functional status
Although the development of neoplasia involves a global change in the complex architecture of signaling pathways that define and regulate the
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growth characteristics of a cell, the extensive interconnections between these pathways suggest that targeting a specific branch may lead to global effects [129, 206]. Given the high frequency of p53 alterations in almost every tumor type and the tumor suppressive role of the protein, p53 is an appealing target for cancer therapy. Various approaches have been used to restore p53 function in tumor cells or exploit the loss of p53 activity to selectively destroy cancer cells [128]. The observation that some p53 alleles can retain partial function has stimulated investigations into the biophysical/biochemical features of p53 mutant alleles and the possibility of reactivating mutant proteins using small molecules (chemicals or peptides) [27, 61, 147, 171, 183]. The same approach, but with the opposite aim to partially inactivate WT p53, has been explored with the idea that normal cells surrounding a tumor may be protected from the inactivating effects of conventional therapy [108]. The viral delivery to tumor cells of WT copies of p53 capable of inducing apoptosis has also been evaluated [38, 195, 214]. Another strategy for treatment of p53 defective tumor cells is replication of viruses that only propagate in p53-defective cells. Promising results have been reported both with a modified adenovirus, that has a mutation in the protein responsible for WT p53 inactivation [18, 128] and with a WT adenoassociated virus that induces cell cycle arrest in normal WT p53 cells but apoptosis in p53-deficient tumor cells [167]. Given the many differences in p53 mutations, a detailed knowledge of the functional status of p53 mutants could have clinical value, especially for therapies tailored to specific tumors. Several methods for functionally classifying p53 mutants have been used that are based on physical/chemical, or immunological/structural parameters [1, 26, 35, 146]. However, many of these approaches to functional analysis have had limited utility.
2
YEAST-BASED p53 FUNCTIONAL ASSAYS
There are now many examples where the yeast Saccharomyces cerevisiae has proven an invaluable in vivo test tube for examining diverse human gene functions and disease including, for example, cell signaling, DNA metabolism, and mitochondrial function [102, 169]. No p53 homologue has been reported in yeast. In fact, the evolution of the p53 gene seems to coincide with the development of multicellular organisms, possibly because of the need for tissue remodeling via programmed cell death and also because the elimination of individual cells in an organism in response to DNA damage can be beneficial. However, damage recognition and cell cycle responses induced by signaling pathways
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in yeast are very sensitive and elaborate and they involve protein kinases and complexes that are well conserved during evolution [208]. The high evolutionary conservation between yeast and mammals is also evident in basic cellular processes such as DNA replication and repair, chromosome segregation, transcription, and translation [14]. Because of these features and its genetic tractability, we have referred to yeast as honorary mammal [169]. As described in the following sections, yeast-based assays have provided the means for structure/function characterization of p53, functional classification of p53 mutant alleles, identification and evaluation of p53 response elements, and screening of chemicals capable of reactivating p53 mutants. These assays currently provide the only practical means to rapidly identify p53 mutations from individual patients and to assess functional status. In a broader sense, the development of a p53 mutant functionality database along with linkage to the vast IARC human p53 tumor mutation database will be valuable in understanding the correlation between p53 functional status, tumor aggressiveness, and responsiveness to therapy.
2.1
Human p53 can function as a transcription factor in yeast
The evolutionary conservation of transcriptional machinery prompted early investigations of human p53 as a sequence-specific transcription factor in yeast. Schärer and Iggo [180] constructed a hybrid yeast promoter where the yeast upstream activating sequences were replaced by a p53 RE. They demonstrated that expression of WT, full-length p53 but not tumorassociated mutants (four were tested) could induce a reporter gene downstream of the promoter. Interestingly, a murine p53 mutant that appeared temperature-sensitive in mammalian cells, was also temperature-sensitive for transactivation in yeast [175]. This work showed that the key function of the p53 tumor suppressor gene could be rapidly assessed in yeast. 2.1.1
p53 functional assays for screening cancer cell lines and tumor samples
The observations in yeast prompted the development of assays using cDNAs that could more rapidly evaluate p53 functional status in tumors or cell lines. The general features are summarized in Figure 3A. p53 cDNAs can be expressed in yeast from constitutive or inducible promoters (Figure 3A). Most investigations have employed the strong, constitutive promoter of ADH1. WT and mutant p53s do not inhibit the growth of yeast at the level of expression obtained with this promoter. The inducible GAL1 promoter has
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also been used. Its expression is repressed on glucose media but is induced to very high levels on media containing galactose as carbon source.
Figure 3A. General features of the p53 functional assay in yeast. Expression of p53 in yeast and transactivation by p53. The complete cDNA of a given p53 allele can be expressed under either a constitutive or an inducible promoter from a selectable plasmid. The centromeric plasmids are stably transmitted at low copy number. WT p53 can act as a transcription factor in yeast stimulating the activity of a promoter whose upstream activating sequences have been replaced with a p53 response element. Mutant p53 would change the level of transcription of the reporter gene thus leading to phenotypic changes. If the p53 is not active, no transactivation is observed. If the p53 allele is less active, higher levels of GAL1 induction are needed; if more active, less induction is required for a phenotypic change.
Overexpression of WT p53 under the GAL1 promoter leads to severe growth inhibition [17, 144]. The mechanism awaits further investigation, but the phenotype has been exploited in different types of selection screens [89] (see sections 2.1.4 and 2.3). Recently, we developed a protocol using the GAL1 promoter that allows for both inducible and rheostatable expression of p53 alleles. This method further expands the sensitivity of the transactivation assay as a means to classify p53 tumor alleles [87] (see sections II.3 and II.6) since it allows quantitative assessment of mutant activity compared to WT. The p53-dependent reporter gene can be transcribed from a plasmid or an integrated copy in the yeast genome. p53 transactivation is assessed using different phenotypic assays according to the reporter gene [24, 54, 95] (see Figure 3B). For example, HIS3 gene expression allows growth on plates lacking histidine (i.e., only WT p53 expressing cells will grow). The URA3 gene can be similarly used on plates lacking uracil, but it also allows for
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counterselection of colonies where p53 function is lost since Ura cells are sensitive to 5-fluoro-orotic acid (5-FOA) [21]. Phenotypic characterization of p53 null alleles based on transactivation of a reporter gene Reporter gene: HIS3 Assay:
URA3 -
growth on his
ADE2 -
growth on ura or 5-FOA
growth on plates with low adenine
Response: Wild type p53: p53 mutant:
growth no growth
growth on ura- and 5-FOA sensitive -
no growth on ura and growth on 5-FOA
growth, white color slow growth, red color
Figure 3B. Summary of reporter genes and the assays used for phenotypic analysis of p53 mutant alleles under conditions of high expression.
Transcription of the ADE2 gene provides for a simple color assay on plates containing low amounts of adenine. Cells expressing WT p53 and nonfunctional p53 mutants will grow but the colonies have different sizes and colors. As shown in Figure 3C, colonies expressing nonfunctional alleles are small and red, while WT p53 colonies are normal size and white. Blockage of adenine biosynthesis upstream of ADE2 gene leads to the accumulation of a red pigment. On plates containing a low amount of adenine, the ade2 mutants appear as small red colonies. Expression of WT + p53 leads to an Ade2 phenotype resulting in white colonies. Three other reporters of p53 activity have been used: the betagalactosidase gene [200, 202], Green fluorescent protein (GFP) [184], and the firefly luciferase gene [222]. While their levels of expression can be quantified, they are not as useful in genetic screens of p53 mutations, since they do not provide for rapid identification of colonies expressing p53 mutants.
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Figure 3C. Example of phenotypic assays using the ADE2 red/white reporter gene. Under conditions of high constitutive expression of WT p53, transactivation of ADE2 results in white colonies. Loss-of-function p53 mutants will result in red colonies since they cannot turn on ADE2. When the GAL1-based inducible expression system is used, transactivation is proportional to the levels of WT (or mutant) p53 protein expressed. Presented are yeast transformants with WT p53 grown on plates containing different amounts of galactose inducer (plus raffinose carbon source). Also shown are western blots of protein extracts from cells grown under the same conditions. The appearance of colonies extends from red to pink to white as the amount of p53 is increased.
2.1.2
Characterization of tumors with single and multiple p53 mutations
Tumors containing p53 mutations may have cells that are heterozygous or homozygous for WT p53 or have more than one p53 mutation. As indicated above, the yeast-based approaches provide for the isolation and functional analysis of separated alleles (FASAY) [28, 41, 185]. The procedure utilizes the exquisite recombination abilities of yeast as described in Figure 3D. First, total cDNAs are prepared by reverse transcription of total RNA extracted from fresh or frozen tumor biopsies. The p53 cDNA is then amplified using a pair of specific primers, typically for amino acids 43 through 374 [54]. This region corresponds to part of exon 4 through the beginning of exon 11 and includes the entire sequence-specific DNA-binding domain where most tumor-associated mutations are found (see Figure 2).
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Unpurified PCR products are directly transformed into yeast together with an expression plasmid that contains a selectable marker, an origin of replication, a centromere for stable propagation at low copy number, and consecutive sequences corresponding to the Nter and Cter regions of p53, coding for amino acids 1–64 and 351–393, respectively. The vector is linearized between these regions. Thus, the PCR product and plasmid have 63 and 69 nucleotides of common sequence at each end.
Figure 3D. Diagram of the gap repair assay for isolation of individual p53 alleles directly from tumor samples. Total RNA is extracted from fresh or frozen tumor biopsies and total cDNA is prepared by RT-PCR. The p53 cDNA is PCR amplified using specific primers. Unpurified PCR products are transformed into yeast together with a linearized p53 expression vector whose ends are identical to the ends of the PCR fragment. The highly efficient homologous recombination system of yeast generates a complete p53 expression vector. The functionalily of the p53 allele is then assessed using a transactivation assay.
Cotransformation of PCR product and linearized vector leads to efficient gap repair of the plasmid by the PCR fragments yielding a circular vector capable of expressing a complete p53 cDNA. Using 50 ng of PCR product and 10 ng of vector, more than 1,000 colonies can be obtained in a typical transformation experiment. Each colony is selected for the presence of the reconstituted vector and corresponds to the isolation of a single p53 allele. Thus, the intrinsic heterogeneity of tumor materials is not a major limitation with this approach and mixed samples containing various mutant or wild type alleles can be analyzed effectively. In the case of the ADE2 p53 reporter gene, WT p53 alleles expressed in normal cells or in heterozygous tumor cells would result in variable percentages of white colonies and red colonies corresponding to the frequency of the loss-of-function p53 alleles. The nature of the mutations can be easily determined by recovering the plasmid
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from the red colonies or using a colony PCR approach followed by DNA sequencing. There are two limitations with the yeast functional assay. It requires extraction from fresh or frozen samples and errors may result during transcription and amplification. These limitations hold true for many analyses of clinical samples by PCR. Typically there are 2–5% red colonies, possibly due to RT-PCR-generated errors. Amplification fidelity can be assessed by parallel RT-PCR of each sample [34]. The critical requirement for RNA can be avoided by modifying the functional assay into an exon-byexon DNA analysis that uses formalin-fixed, paraffin-embedded archival samples [41]. When compared to other systems for p53 mutant identification, the yeast functional assay has greater sensitivity than electrophoresis-based methods (SSCP, DGGE) [113, 132, 197, 198] and is comparable to the GeneChip approach [213]. In addition, the yeast assay is rapid and relatively inexpensive and, furthermore, it enables functional defects in p53 to be assessed. Functional analysis based on transactivation capacity in yeast is robust and appears more sensitive than transient transfection gene reporter assays in mammalian cells. Over 100 different mutant p53 alleles isolated from clinical samples were found to lack transactivation function in the red/white ADE2-based assays [31, 55, 172]. The tissues have included fresh and frozen specimens from brain (glioblastoma and astrocytoma), upper aero-digestive tract, breast, colon, bladder, prostate, lymphocytes, and skin [29, 30, 34, 46, 65, 93, 103, 123, 125, 134, 145, 151, 159, 160, 168, 172, 186, 204, 211, 215]. Included in these analyses are dysplasias, primary carcinomas, and advanced tumors. The physical and functional separation of p53 alleles also provides an assay to monitor intra-tumor WT p53 delivery by viral vectors for cancer gene therapy approaches to treat brain, head and neck, ovarian, and lung cancer [38, 106, 195, 199, 214]. Specific primers can be used to amplify the mRNA obtained from endogenous p53 alleles or the virally encoded WT cDNA to estimate the contribution of viral WT p53 mRNA in tumor samples. 2.1.3
Identification of p53 mutants with differential effects on the transactivation of specific promoters
The yeast-based systems provide opportunities to evaluate p53 mutants for ability to transactivate at various REs. As described above, the consensus sequence of the p53 response element is degenerate and REs from mammalian p53-regulated genes exhibit many deviations from the consensus [42, 49, 50]. This observation prompted the evaluation of the transactivation
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activity of p53 mutants with different p53 REs. Most screenings of tumor samples have utilized high p53 expression and the RGC RE to determine if p53 is capable of transactivation. The RGC sequence was originally found within the human ribosome gene cluster and was shown to be a bona fide p53 RE based on both in vitro binding and transactivation assays [11]. The WAF1 REs of the key p53-regulated genes such as p21 and bax have also been used [55]. Highly expressed WT p53 is capable of transactivation from these REs and results in white colonies in the ADE2 based assay. Flaman et al. [54] examined 51 different tumor-associated mutations observed in human tumors. Nine of them scored as WT with P21, while none were able to transactive the BAX or the RGC RE. Another 77 mutants were examined by Campomenosi et al. [30] using REs of RGC, BAX, and P21. Consistent with the previous report, about 15% of the mutants were WT toward the P21, but none of them was able to transactivate BAX. This finding suggests that inactivation of the apoptotic function of p53 is critical for tumor development. DiComo and Prives [42] further expanded the investigation of p53 transactivation ability using nine different REs upstream of the HIS3 reporter. Nineteen tumor-derived mutants were analyzed at different temperatures. While p53 hot spot mutants appeared transcriptionally inactive, four mutants in the DNA-binding domain scored as WT with P21 and a strong consensus RE at 37°C, and one of them [R283H) appeared fully active with several REs. An additional group of four mutants was active with the same REs at lower temperatures. These mutations affected amino acids belonging to the beta-sheet scaffold of the p53 DNA-binding domain. All the mutations were inactive with RGC. Consistent with this, WT p53 was less active with the RGC and BAX REs compared to P21 and the strong consensus sequence. A sensitive screen was recently developed to specifically isolate p53 mutants retaining partial function (in preparation). The assay was based on the observation that in the ADE2 reporter system white and red colonies represent the extreme phenotypes in a range of p53 transactivation capabilities from WT to complete loss-of-function. With the view that it might be possible to identify mutants with reduced function, p53 mutations were generated by random mutagenesis and directly cloned into yeast using gap repair (see Figure 3). Examination of colony color and morphology allowed the identification of intermediate phenotypic classes (pink, sectored, and speckled colonies) that were due to the expression of p53 mutants with reduced transactivation activity. Approximately 60 different single amino acid changes were found that could result in proteins with reduced activity. The changes in activity were quantified with a luciferase reporter assay that was developed in yeast. A group of mutants that were tested for transactivation and growth suppression in human cells validated the yeast
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assay (in preparation). Interestingly, the mutations were spread uniformly across the entire mutagenized region (amino acids 50–350) which extends beyond the DNA-binding domain. Since there were no repeats, the screen was far from saturation. Based on the number of these mutants that correspond to p53 tumor mutations in the IARC p53 database, we estimate that up to 20% of tumor-associated p53 mutants retain partial function. The functional evaluation in yeast of many p53 alleles associated with cancer has revealed that a large fraction of mutants retain residual activity. Interestingly, transactivation assays in mammalian cells along with in vitro measurements of conformation stability and DNA-binding potential have confirmed the yeast results [26, 28], although the number of p53 mutants examined is small. We have concluded that it is not possible a priori to predict the functional effect of most mutations based on sequence conservation, topology, or structural models, although loss-of-function alleles appear to affect amino acids with a higher degree of evolutionary conservation [31]. These results further support the need for a detailed rational classification of p53 mutants associated with cancer based on in vivo transactivation at many promoters. 2.1.4
p53 functional assays to evaluate dominance
Many of the p53 mutants appear to be dominant, which helps explain their frequent occurrence in tumors. The levels of expression of p53 alleles and the ratio between WT and mutant proteins within a cell can significantly influence the dominant-negative potential of p53 mutants. In fact, since p53 is active as a tetramer, dominance could result from the formation of mixed tetramers that would be conformationally unstable and incapable of DNA binding and transactivation. Yeast-based functional assays have provided for rapid assessment of the dominance potential of p53 mutants when equally expressed with WT p53. Brachmann et al. [24] developed a mutation selection screen for dominant-negative p53 mutants that was based on the counterselectable URA3 reporter gene. Briefly, WT p53 can induce URA3 transcription leading both to uracil prototrophy and sensitivity to 5-FOA. If a p53 mutant is co-expressed with WT p53 and acts in a dominant fashion, cells will grow on 5-FOA plates. The screen identified 31 dominant mutants of which 13 conferred at least partial growth even in the presence of 2 copies of WT p53. The mutations clustered in the p53 mutational hot spots, supporting the view that dominant-negative interactions are important in the selection of p53 missense mutations in human cancers. The ADE2 color assay has also been used to evaluate the dominance of 71 different p53 alleles [85, 136]. In this assay, a recessive p53 mutant in the presence of WT p53 would lead to white colonies, while a dominant mutant
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would lead to pink/red colonies. Approximately 30% of the mutations exhibited dominance. These mutants affected residues that are highly conserved and that are more frequently associated with cancer; they appeared to cluster in the L3 loop and H1 and H2 helices of the DNAbinding domain (for an extensive description of p53 structure, see [36, 189]). The possibility that the mutants exhibited different degrees of dominance according to the p53 RE used in the transactivation assay was also explored. A significantly higher fraction of mutants was dominant over WT p53 with the BAX compared to the P21 RE. This result is likely due to the intrinsic difference in activity of these two REs and suggests that dominance in vivo could be p53 gene-target specific. Interestingly, a recent study in mammalian cells using bi-cistronic vectors that allows for equal expression of WT and mutant p53 from a strong viral promoter revealed that dominant-negative effects were visible only with the bax gene [8]. A recent investigation employed the ADE2-based yeast functional assay to evaluate transactivation and dominance of p53 alleles expressed in endometrial carcinomas. Both the frequency of p53 mutations and the ratio of dominant-negative mutants was higher among endometrioid-type compared to serous-type tumors [177]. 2.1.5
Reactivation of p53 mutants using yeast-based screens
The restoration of cellular pathways in tumor cells that lead to cell death by apoptosis is an appealing cancer therapy approach. In this respect p53 represents a potentially powerful target. The identification of a large group of p53 mutants that appear to retain some transactivation function and the observation that the majority of loss-of-function alleles are recessive when heterozygous with WT p53 [136] suggest that many tumor-associated mutants retain native conformation or do not dramatically perturb protein conformation. Different types of yeast assays have been used to address the possibility that small molecules could partially reactivate p53 function either by stabilizing the structure of the mutants and/or by enhancing DNA-binding capability. Brachmann et al. [25] used an elegant screen to identify p53 mutations that act as intragenic suppressors of common hot spot p53 tumor mutations. The assay utilizes random PCR mutagenesis or degenerate oligonucleotides [13] to induce mutations in defined regions of the p53 cDNA. Gap repair was used to clone the mutagenized fragments within a p53 cDNA that contains a specific mutation corresponding to a p53 hot spot. Compensating mutations were identified that restore p53 function based on transcription of the p53 URA3 reporter. Single amino acid changes were identified that were capable either of restoring function to specific p53
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mutations or had a broad reactivating effect on several hots pots. The intragenic suppressors affected amino acids that belonged either to the L1 loop (T123A, T123P) or the L3 loop (N239Y, S240N). Another suppressor (N268D) was found in the S10 beta-strand. This mutation might have a stabilizing effect on the DNA-binding domain. It is interesting that intragenic suppressors were identified in both L1 and L3 loops. According to the crystal structure, these loops are exposed regions when the DNA-binding domain is bound to a consensus RE [36]. Thus, the loops might be good targets for small molecules that could provide in trans functional reactivation of p53 hot spot mutants. Expression of the reactivated double mutants in mammalian cells resulted in restoration of transactivation and apoptotic function in transient assays. No suppressor for the common tumor p53 mutation R175H has been identified with the yeast screen [13]. According to studies with conformational sensitive antibodies and also in vitro analysis with purified core domains, R175H has a denatured conformation that might prevent this type of rescue [36]. However, a recent report based on a mammalian cell-based screen of molecules capable of reactivating p53 function reported the identification of a compound that can activate R175H [27]. Another approach can also reveal suppressor mutations. We developed a screen for p53 mutants exhibiting enhanced transactivation function compared to WT p53 [87]. Unlike other assays described above, our approach was based on rheostatable expression of p53 under the GAL1 promoter (see Figure 3A & C). At low-expression levels WT p53 cannot activate transcription of the ADE2 reporter gene and yeast colonies appear as p53 mutants (i.e., red colonies). Thus, p53 alleles with enhanced function can be easily selected (i.e., white colonies) at low levels of p53 expression. Interestingly, several single amino acid changes produced by PCR mutagenesis were identified both in the L1 and L3 loops, in the H2 helix and in exposed regions of the beta-strands located on the opposite side of the protein:DNA surface. This supertrans group of mutants included the intragenic suppressors reported by Brachmann et al. [25]. Consistent with the enhanced function, the supertrans alleles were not affected by dominantnegative p53 mutants when heterozygous. A recent study showed that the radio/chemoprotective agent amifostine could activate WT p53 in mammalian cells [148]. Maurici et al. [127] used the yeast functional assay to evaluate whether amifostine had a direct effect on p53 activity. A temperature-sensitive p53 mutant (V272M) was activated by amifostine at the nonpermissive temperature and a small number of p53 mutants that retain function with the P21 but not the RGC or BAX REs were partially reactivated by the compound. These results suggest that the yeast functional assay could be applied to screening of compounds to identify
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leads for p53 reactivating drugs. However, the results were based on the color assay (red/pink/white) that might also be influenced by the effects of the drug on the growth of yeast. A quantitative assessment of the effect of amifostine both on WT and mutant p53 activity is needed.
2.2
p53 mutational fingerprints identified with the yeast functional assay: a tool for molecular epidemiology
As indicated above, p53 mutation spectra associated with various cancers in specific populations may be helpful in identifying carcinogens and understanding underlying events [205]. Exposure to some environmental agents has been associated with the features of p53 mutation spectra in tumors [15, 83]. The yeast-based p53 functional assay has been adapted to identify p53 mutations induced in vitro or in vivo by mutagens and carcinogens [86, 140]. p53 DNA has been treated directly with several mutagens and carcinogens and DNA changes were determined using in vivo yeast functional assays after transformation with the treated DNA. p53 functional mutants identified by the lack of reporter gene expression (color or growth assay) were sequenced and p53 mutational fingerprints were developed. The following agents have been examined: UV, 8-methoxy-psoralen, CCNU, Me-lex, nitric oxide, and reactive oxygen species [23, 62, 84, 86, 90, 105, 137, 138, 141]. The UV-induced p53 mutation spectra identified by the yeast system included tandem mutations at dipyrimidine sites, a hallmark of UV mutagenesis [90, 137, 140]. Further analysis revealed that the yeast p53 spectra and the reported p53 mutations associated with nonmelanoma skin cancer are indistinguishable. This result exemplifies the utility of determining mutation spectra in defined experimental conditions using the same in vivo target of mutagenesis that is used in molecular epidemiological studies [62]. Similarly mutation spectra induced by bifunctional alkylating agents were consistent with earlier mutagenesis studies [86]. The correlation between lesion hot spots and mutation hot spots has also been evaluated using a minor-groove specific methylating agent as well as oxidative damage [23, 105]. Based on results with UV the functional assay can also be used to examine the effects on p53 of DNA damage induced in vivo [139]. So far, only direct-acting mutagens have been used. The study of chemicals that require metabolic activation may also be feasible if yeast are provided with a repertoire of human or mammalian cytochrome genes [156].
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Yeast assays to characterize p53 amino- and carboxy-terminal functional domains
The Nter and Cter domains play important roles in p53 structural organization, interaction with other proteins and transactivation. The p53 functional assays can also be used to address changes in these regions. Ishioka et al. [94] reported that the p53 Cter is generally refractory to mutations that inactivate transactivation functions and p53 deleted for the last 40 amino acids (Q354Stop) retains WT activity. However, single amino acid changes in the tetramerization domain (L344P, K351E) can nearly inactivate p53 transactivation. A germline p53 mutation in the tetramerization domain (R337C) that is found in cancer prone families [119] results in yeast colonies that are pink in the ADE2 assay, suggesting partial retention of transactivation activity. It is possible that this amino acid change affects the stability of the p53 tetramer; however, the mutant protein is unaffected by temperature changes (23°–34°C), unlike many pink colony mutants that we have examined (unpublished). A second germline mutation has been identified at codon 337 (R337H) in Brazilian families that were characterized by increased incidence of childhood adrenal carcinoma (ACC) [170]. Surprisingly, there was no effect on the incidence of other types of cancer. A recent study showed that R337H retained almost normal function in in vitro DNA-binding assays, but exhibited temperature and pH-sensitivity [43]. The reason why this specific functional defect is highly tissue-specific remains to be determined, although it has been proposed that the adrenocortical cellular environment would lead to functional inactivation of this p53 mutant thus increasing the risk of cancer development. It will be interesting to compare the R337C and R337H phenotypes in the yeast functional assay. Candau et al. [32] used a yeast-based assay to analyze Nter functions of p53. Instead of expressing the complete p53 cDNA, chimeras were created between p53 and the DNA-binding domain of the GAL4 transcription factor. The use of p53 missense and deletion mutants revealed that the Nter region of p53 contains a second transactivation domain mapping between amino acids 40–83 with amino acids W53 and F54 playing a critical role. The activity of this transactivation domain has been confirmed in mammalian cell experiments [221]. It has also been proposed that this element might display target gene specificity, thereby contributing to p53-induced differential transactivation. However, the underlying mechanisms for this effect remains to be established. Coincident changes at amino acids 22 and 23 (L22E and W23S) in the first p53 trnsactivation domain [42] can inhibit binding to MDM2, p300/CBP, and TAF proteins [53, 118, 152, 163]. Consistent with this, the
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double mutant was not functional in mammalian cells nor in transgenic mice even though it had no effect on p53 DNA-binding affinity [97]. Unexpectedly, the mutant retained transactivation similar to WT when tested with a strong consensus RE in yeast [42]. By way of comparison, this suggested that the transactivation domain at amino acids 53 and 54 is relatively more important to p53 activity in yeast than in mammalian cells. However, the functional effect of mutating amino acids 22 and 23 as well as deleting the last 40 amino acids [94] might have been masked since p53 was expressed at high levels from a constitutive ADH1 promoter. Experiments at low expression levels indicate that both transactivation domains have similar contributions to p53 activity (unpublished). In addition, we have found that deletion of the last 40 amino acid at the carboxy terminus also reduces p53 transactivation in yeast (unpublished). This is contrary to the results of gel shift experiments with naked DNA, but consistent with DNA-binding experiments with chromatinized DNA that may mimic more closely the nuclear environment [52].
2.4
Yeast assays to characterize and identify p53 interacting proteins
The contribution of yeast transcriptional adaptor proteins in p53 transactivation can also be explored with yeast functional assays. The yeast ADA2, ADA3, and GCN5 genes, whose product are involved in multiprotein complexes that have histone acetylating activities, were shown to contribute to transactivation mediated by the p53 Nter [32]. A recent study showed that the histone H4 and H2A acetyltransferase activity of the NuA4 complex can contribute to p53 activity in yeast [149]. Transcriptional activation by p53 was associated with targeted NuA4-dependent histone H4 hyperacetylation. A genetic screen for human proteins that interfere with p53 transactivation function identified the human homologue of ADA3 as an important cofactor in p53 activity [210]. This “yeast dissociator screen” is based on the URA3 reporter gene and selection against p53 activity on 5-FOA plates (described in section 2.1.4; see also Figure 3). A human protein that is capable of binding to p53 might reduce transactivation activity thereby enhancing growth on 5-FOA plates. This protein–protein interaction could identify proteins that are truly negative regulators of p53 activity or that instead cooperate with p53 in human cells but appear as inhibitors in the heterologous system in yeast since additional protein components are missing. Experiments in mammalian cells showed that hADA3, which is part of histone acetyltransferase complexes, was capable of stimulating p53 activity and that its physical interaction with p53 is possibly modulated by stress responses leading to posttranslational modifications of p53.
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A similar system was developed in the fission yeast Schizosaccharomyces pombe [207]. As in the budding yeast, p53 can stimulate transcription in this organism and its overexpression leads to severe growth inhibition [17]. An S. pombe strain was constructed containing two p53-dependent reporter genes, one of which (ura4+) is counterselected on 5-FOA plates and positively selected on uracil-deficient plates. The assay was validated using SV40 large-T antigen and MDM2, both known interactors and inhibitors of p53 activity. It was also shown that tumor-derived p53 mutants are functionally distinguishable from WT p53 based on transactivation. Finally, the assay revealed that the p53 homologue from Drosophila could also be functionally tested in S. pombe [207]. p53 protein–protein interactions have also been identified in yeast 2hybrid screens [96]. However, since p53 also can activate transcription with its transactivation domains, the entire p53 protein cannot be used. The screening relies on the construction of two chimeric proteins, one containing a DNA-binding domain and the other a transactivation domain [109]. The GAL4 DNA-binding domain is fused to the desired p53 protein fragment for which interacting proteins are sought. This chimeric protein is presented as a “bait” to a library of proteins, each of which is fused to the transactivation domain of the GAL4 protein. Physical interaction between the bait and an unknown protein would bridge the DNA-binding and the transactivation domain of GAL4 leading to selectable transactivation of a reporter gene. The 2-hybrid screen was performed using a murine p53 sequence lacking the Nter domain leaving amino acids 73–390 [96]. Two p53 interacting proteins were discovered, 53BP1 and 53BP2. Binding of p53 with either protein inhibited its binding to REs in vitro. 53BP1 contains two Brca1 C-terminal (BRCT) domains, a common protein–protein interaction motif found in proteins involved in the DNA damage and repair responses [98]. Structural analysis confirmed that 53BP1 binds p53 at a region that overlaps with the DNA-binding surface of p53 and involves p53 residues that are mutated in cancer. Analysis of intracellular localization of 53BP1 in human cells revealed discrete nuclear accumulation within 5–15 min after exposure to ionizing radiation (IR) [4]. These results suggest 53BP1 functions early in the cellular response to DNA DSBs and may activate p53 responses [182]. The 53BP2 contains an SH3 domain and also ankyrin repeats that may mediate protein–protein interactions. 53BP2 is a 528 amino acid fragment of the carboxy terminal region of the ASPP2 protein [179]. Interestingly, ASPP2 and a related protein ASPP1 can specifically enhance p53 transactivation function but only of specific genes leading to p53-induced apoptosis but not cell cycle arrest. The expression of ASPP2 is subject to stress signals and frequently appears downregulated in breast carcinomas expressing WT but not mutant p53 [179]. This result provides additional
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evidence that functional alteration of the p53 pathway may occur through upstream regulation of p53 protein activity. Differential transactivation is important in the biological responses to stress, particularly apoptosis vs cell cycle arrest. The finding that ASPP2 acts as cofactor for p53 but possibly only for the activation of specific target genes provides support to a proposed mechanism explaining how differential transactivation by p53 and the specificity of the p53-induced cellular responses are achieved [110, 208]. Namely, cell- type-dependent availability of cofactors may decide which pathway, cell cycle arrest or apoptosis, is induced by p53 in response to a stress signal. Consistent with this, a recent 2-hybrid screen identified the homeodomain-interacting protein kinase-2 (HIPK2) as a p53 cofactor. HIPK2 phosphorylates p53 at serine 46, stimulates p53 transactivation and mediates the induction of apoptosis [40]. In addition to p53-binding factors and modifying proteins, the intrinsic DNA-binding affinity of p53 toward the many related REs in target genes can play an important role in differential transactivation. As discussed below (see section 2.6), yeast-based functional assays have provided tools to assess this possibility.
2.5
Identification of p53 mutants exhibiting altered transactivation specificity
Several mammalian transcription factors including p53 recognize a broad variety of REs that belong to a rather loose consensus [133]. Little is known about the mechanisms determining recognition and levels of binding. The proteins interact with only some of the nucleotides in the response elements and also have contacts with the sugar/phosphate backbone. The DNAbinding affinity is affected by conformational changes of the protein:DNA complex and this is particularly important when the transcription factor is a multimer (as hetero- and homodimers or tetramers) [130, 143]. The effects of DNA sequence context in determining structural features of the DNA elements, such as bending of the double helix, are not well understood. In the case of the p53 transcription factor which coordinates multiple biological responses, the loose consensus sequence suggests that p53 variants with altered DNA sequence binding specificity might lead to selective downstream responses among the multiple p53 pathways. For example, changes in the amino acids 120, 248, 277 (including C277R), and 283 that contact the DNA have been examined with in vitro DNA-binding assays and transactivation in yeast [200]. The impact of base changes in the p53 consensus was also examined. Interestingly, while all mutations showed a defect in binding to the canonical DNA element, some mutants exhibited WT or even greater activity toward specific sequences.
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Yeast-based selection screens have been developed to isolate p53 mutants with altered DNA sequence binding specificity. Freeman et al. [60] selected p53 mutants with enhanced activity using only one of the two inverted dimers within a RE (i.e., a half-site consisting of 5′ RRR-CWWGYYY) to mediate transactivation. Two mutations in the conserved domain II of the DNA-binding domain (L1 loop, S121F, and T123A) were identified. They also varied the content of the Gs and As in the purine (R) tract and found that both 121F and 123A were more active than WT with a G-rich RE (5′ GGGCW-3′). With other sequences, the mutants were less than or equal to WT p53 in acitivity. An alternative approach to identifying p53 mutants has involved reducing the amount of p53 protein to a low level using the GAL1 rheostatable promoter and using complete REs [87]. Using this approach we have identified p53 toxic mutants, that prevent growth at moderate to high levels of expression and can lead to inviability [89]. This screen was based on previous results that showed a direct correlation between the growth inhibition phenotype and the ability of p53 to act as transcription factor in yeast. We hypothesized that mutants with a greater impact than WT p53 on growth of yeast may possess enhanced DNA binding/transactivation. Among the many mutants some (including T123A) appeared more active with all the p53 REs examined at low levels of p53 and were termed supertrans (see section 2.1.5). Other mutants exhibited altered DNA-binding selectivity including enhanced, reduced, and loss-of-function depending on the RE used. These mutants had amino acid changes in the L1 loop (positions 122 and 125) and in the helix II (the DNA contact sites 277 and 279). Most mutants were at amino acids that do not directly contact DNA. Saller et al. [178] investigated the impact of S121F and C277R when expressed in mammalian primary and tumor cell lines. Interestingly, the altered DNA-binding specificity in yeast was also observed in mammalian cells both with gene reporter assays and endogenous gene expression. This suggests that the intrinsic DNA-binding affinities of p53 mutants toward specific REs assessed by the yeast functional assay can predict the behavior of a p53 mutant in mammalian cells. In a subsequent study this group compared the in vivo DNA binding of WT p53, S121F, and C277R at six promoters using chromatin immunoprecipitation assays in mammalian cells [101]. Also they correlated DNA binding with transactivation determined by real-time PCR of four genes. S121F appeared defective for binding and transactivation of p21, MDM2, and PIG3, but was slightly more active than WT p53 at the bax promoter. 277R was defective or partially defective at p21 and bax, but active at MDM2 and PIG3. These results are again consistent with predictions of the yeast functional assays. Furthermore, the
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ChIP data showed differences in promoter occupancy by WT p53 as well as the different p53 mutants. Overall, these results suggest that differences in intrinsic DNA binding of p53 toward the many REs are important factors in p53 differential transactivation and can play a significant role in the biological consequences of p53 mutants. This is exemplified by our recent analysis of the murine T122L p53 mutation hot spot [88]. This mutation was found in 40% of UVinduced skin tumors in Xpc–/– p53+/– mice but was never detected in UVinduced skin cancers in WT mice and was rarely observed in Xpc+/– p53+/– Xpc–/– p53+/+, or Xpa–/– mice. The yeast functional assay at low levels of T122L expression revealed altered transactivation specificity that included enhanced, reduced, and loss-of-function for specific REs. Interestingly, transactivation was impaired with some REs from apoptotic genes but it was normal or even slightly enhanced with REs from cell cycle control and DNA repair genes. The altered DNA-binding affinity and specificity of the T122L mutant protein suggest that it would lead to an altered pattern of expression of downstream p53-regulated genes in vivo in skin cells and that this could contribute to its selection. The balance between the defect in apoptosis induction and the retained activity in DNA repair and cell cycle control may become advantageous specifically in XPC–/– cells that are proficient for transcription coupled repair and exhibit a WT like p53 response to DNA damage. In WT and Xpa–/– cells dominant-negative loss-of-function p53 mutants would instead be more advantageous to reduce p53-dependent and independent apoptosis.
2.6
Functional classification of p53 response elements in yeast reveals a wide range of transactivation capacities
The yeast functional assay provides the means to specifically address the role of p53 protein levels and p53 intrinsic DNA-binding affinity in differential transactivation of target genes. Recent work using in vitro preparation of chromatinized DNA and in vivo ChIP assays suggested that p53 protein in human cells could be active for DNA binding even in the absence of posttranslational modifications [9, 12, 52, 196]. Furthermore, changes in promoter occupancy resulting from stress-induced signals reflected more directly the changes in p53 levels rather than modifications in relative binding affinity [101]. However, chromatin modification induced by p53 binding to promoter sequences and recruitment of histone acetylases is likely very important in transactivation although effects are locus dependent [165].
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We have developed a set of isogenic yeast strains that differ only for a single p53 response element cloned upstream of a minimal CYC1promoterADE2 open reading frame at the ADE2 locus [91]. Thus, all the possible factors affecting p53 transactivation are kept equal and the role of binding affinity and p53 protein amount can be precisely evaluated. To construct these strains we took advantage of a recently developed in vivo mutagenesis approach (discussed below) that allows for rapid modifications of yeast sequences using oligonucleotides that are transformed directly into yeast [192]. Thirty strains, each with a different p53 RE derived from mammalian p53-responsive genes, were constructed [91]. Genes involved in cell cycle control, apoptosis, DNA repair/replication, and p53 regulation were included in this broad panel of REs to examine various features of WT and mutant p53s. To understand the importance of p53 levels in differential transactivation, we defined conditions for the rheostatable induction of p53 under the GAL1 promoter. Nearly linear increase of p53 over a 100-fold range could be accomplished using a range of galactose concentrations from 0.001% to 0.12% added to medium containing 2% raffinose [91]. Transactivation of the ADE2 reporter gene could be assessed using the phenotypic (red/pink/white) assay and also by quantification of ADE2 mRNA with real-time PCR measurements. The p53 REs were ranked according to their relative activity. Surprisingly, there was a 1000-fold range in transcriptional activation between the various RE sequences. Statistical models that estimate binding energy based on nucleotide usage and the degree of heterology of a RE from the consensus [16] predicted only small variations and these did not correlate with the functional results. We found that the DNA sequence CATG at the junction of p53 monomer binding sites greatly affects p53 transactivation capacity, possibly due to structural properties such as bending [143]. The absence of spacing between dimmer-binding sites is also crucial for high transactivation capacity by WT p53. This result contrasts with in vitro DNAbinding assays [50] but confirms previous observation with yeast-based transactivation [202]. Our results suggest that intrinsic DNA-binding affinity as well as p53 protein levels are important contributors to p53-induced differential transactivation. WT p53 had weak activity toward half the apoptotic REs, particularly those active on the mitochondrial pathway of programmed cell death. Activation of these targets requires high levels of p53. Possibly specific coactivators that are missing in yeast or the combinatorial action of multiple transcription factors play an important role in transactivation of these REs. Qian et al. [166] recently reported the construction of an even larger panel of REs and surrounding promoter regions into a yeast expression
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plasmid as well as a luciferase vector for transactivation experiments in mammalian cells. While they only used high, constitutive expression of p53 and a different p53-dependent reporter, their results are consistent with ours. Interestingly, luciferase assays in mammalian cells were completely consistent with the yeast results, suggesting that the yeast-based observation can predict p53 transactivation capacity in transient assays in cell lines [166].
2.7
Functional assays for p53 family members and their interactions with p53 mutants
Given the key role of p53 protein in the stress-induced cellular responses it was surprising that lack of this gene does not result in severe defects during embryonic development in mice. However, inappropriate expression of p53 can lead to embryonic lethality or increased risk of malformations [37, 44, 76]. Two p53 homologues, p63 and p73, have been identified [99, 115]. The p63 and p73 genes might compensate for the p53 defects in germline and somatic cells. p73 was initially proposed to be a tumor suppressor gene in neuroblastoma due to its chromosomal location at 1p36.2 [191]. This region is frequently associated with loss of heterozygosity (LOH) in other types of cancer. Although there is limited evidence of an alternative spliced form of p53 [56], multiple spliced variants at the Cter and an alternative promoter at the Nter have been described for p63 and p73. Six variants of p73 with differing Cter structures (alpha, beta, gamma, delta, epsilon, and xi) and three of p63 have been reported [191]. The relative abundance of these protein variants varies according to the cell type and in tumor cells. For example, the Νter variants of both p63 and p73 are highly expressed (as much as 10 times more than p53) in epithelial and brain tumors where they appear to have oncogenic properties [39, 45]. The structural organization of the p63 and p73 proteins resembles p53 and comprises a transactivation domain (~20–30% homology to p53), a DNA-binding domain (~60–70% homology), a tetramerization domain (~30–40% homology), and a long carboxy terminus of uncertain function [6]. KO mice have been obtained both for p63 and p73. Unlike the p53 KO, growth defects were observed. The p73 KO showed chronic inflammation, infections, and hippocampal dysgenesis [218]. It has been proposed that the Nter variants act as neuroprotective factors possibly by inhibiting p53-dependent apoptosis. Interestingly, no increase in tumor incidence was reported in these mice. The p63 KO had a lack of mammary glands and defects in limb formation [217]. Mutations in the DNA-binding mutations of the p63 gene have been observed in families affected by the autosomal dominant disorder EEC
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characterized by ectrodactyly, ectodermal dysplasia, and facial clefts [33]. These and other observations suggest that defects in p63 are associated with alteration of the asymmetric cell division of epithelial stem cells [158]. Recent evidence suggests that p63 and p73 play a role in stress-responses; both genes exhibit changes in expression levels and posttranslational modifications [57, 117]. For example, UV-radiation can lead to a reduction in the expression of the Nter p63 variants and this correlates with p53 stabilization. This suggested that p63- as well as p73-variants that lack the transactivation domain may function to override and restrain p53 function. However, there appears to be cooperation of p63 and p73 with p53 in the induction of apoptosis. p63 and p73 variants that contain the transactivation domain can transactivate p53 target genes in experimental systems [157, 216]. Different transactivation capacities were reported for the various Cter variants. It is possible that while the Nter variants inhibit apoptosis physiologically during development, in defined stages or cell types and upon aberrant expression in somatic tumor cells, the other variants could exhibit tumor suppressive functions and be particularly active in stress-induced apoptotic responses [92, 100, 190]. In this respect, it is also possible that abundantly expressed mutant forms of p53 could inhibit p63 and p73 (as well as p53 when heterozygous) through physical interactions that are not mediated by the tetramerization domains [63, 193, 194]. Yeast-based functional assays have been developed to address the transactivation function of p63 and p73 isoforms, to identify functional mutants in tumor samples and to evaluate the effects on functional interaction between p73 isoforms and hotspots p53 mutants. A functional assay for p63 transactivation function has been developed based on the HIS3 reporter gene and a quantifiable assay based on expression of GFP [104, 184]. p63 can use p53 REs to stimulate transcription but the degree of transactivation differs from that of p53. Although creation of p53 equivalent hot spot mutants led to loss of p63 function, the rare p63 missense changes identified in tumors do not appear to affect protein activity. This confirms that p63 is not a target for mutational inactivation in tumors. Nozaki et al. [150] developed an ADE2 - based functional assay for p73 splice variants. They reported that different p73 variants were active to different degrees with p53 response elements. Functional analysis showed that p73 is not mutated in meningiomas, although LOH at 1p36 is common in these tumors.
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Future directions: determining functional fingerprints of tumor-associated p53 alleles using quantitative yeast-based assays
There is considerable evidence suggesting that knowledge of the p53 status of tumors and of the functional defect of a specific p53 mutation may be relevant to prognosis and to tailoring cancer therapy to individual patient needs. The precise functional status of p53 mutations may influence tumor responsiveness to chemo- and radiotherapy, the effect of viral particles targeting p53-defective cells or reintroducing WT p53 and the efficacy of small molecules in reactivating p53 alleles. Furthermore, the residual capacity of each mutant to induce a set of responses, to influence the activity of the remaining WT p53 allele or that of the p73 and p63 protein variants may drive tumor development through different selective pressures. In fact, p53-dependent modulation of transcription is broad and flexible and can lead to different biological outcomes, and mutants may preferentially affect the expression of downstream genes involved in specific pathways (see section 2.1.3 and 2.1.4). It is, therefore, important to evaluate transactivation capacity of p53 mutants using many REs under conditions where protein expression can be varied. The same holds true for investigations of dominance, the ability of a mutant to interfere with p53 family members or the reactivation of p53 mutants. As transient transfection experiments in mammalian cells have shown, transactivation assays with overexpressed p53 alleles under viral promoters may poorly represent physiological conditions. The yeast assay we described has several features that make it appealing for p53 studies. It is rapid, relatively inexpensive, versatile, and highly reproducible; it can assess p53 missense as well as nonsense mutant proteins, and loss-of-function as well as subtle mutations. Here we present a strategy that improves the throughput of the screen. To take into account the levels of p53 expression in the transactivation assay, we have developed a protocol that provides for functional classifycation of any p53 mutant using quantitative yeast assays with tight control of p53 expression [91]. The scheme is presented in Figure 4A and 4B. p53 can be expressed from a selectable centromeric plasmid. For rapid modification of the p53 sequence we have also constructed a yeast host strain by integrating a single copy of the GAL1:p53 expression cassette into the genome in order to utilize our recently developed delitto perfetto in vivo mutagenesis system [192]. A CORE cassette (CO = counterselectable KLURA3; RE = reporter kanMX4 gene) is inserted at a specific location within the p53 cDNA sequence. Four isogenic derivatives of the host strain, each with a single CORE cassette positioned at 200 nucleotides intervals of
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Figure 4A. Development of a p53 mutant functionality database. Construction of any missense p53 mutant can be accomplished rapidly with the delito perfetto in vivo mutagenesis system [192]. The GAL1:p53 expression cassette is cloned into a yeast chromosomal locus to obtain a p53-host strain. A number of isogenic derivatives of this strain are created that have a CORE cassette (containing a counterselectable gene and a reporter gene that identifies cassette integration) [192] inserted at different sites (amino acid 250 in the example, Jordan and Resnick, unpublished). By placing a CORE cassette at 200 nucleotide intervals, the entire p53 can be subjected to mutagenesis. Introduction of oligonucleotides that surround the CORE allows for creation of specific mutants (R273H in the example) or the generation of many different mutants in a small region if degenerate oligonucleotides are used. Only DNA sequencing of the region surrounding the replacement site is needed to confirm the nature of the induced mutation(s) [192].
the p53 cDNA, enables easy production of all possible single missense mutants between amino acids 50 and 350. The mutant construction is achieved by simple replacement of the CORE cassette (selected on 5-FOA and confirmed by kanamycin sensitivity) with oligonucleotides. The oligonucleotide can be specific for a single desired mutation or degenerate in order to randomly mutate the region of interest. Limited DNA sequencing of the replaced fragment can confirm and/or identify the mutants [192]. The host strain with a given p53 allele is then mated (Figure 4B) to a set of 20 isogenic strains representing a p53 functional array. Each of these strains can test p53 transactivation using a reporter gene construct that only differs for the sequence of a p53 RE. Cell cycle checkpoint, DNA repair/replication,
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apoptosis (both receptor and mitochondrial pathway), and p53-regulatory genes are included. The functional array also covers a wide range of intrinsic DNA-binding affinities (see section 2.6). Diploids are selected and the transactivation capacity of the p53 mutants is assessed at different levels of protein expression using replica-plating of purified diploids on different concentrations of the galactose inducer. The same procedure can be used to evaluate p53 dominance toward WT p53, p63, or p73 when expressed by an identical cassette from a centromeric plasmid. Pharmacological reactivation of p53 mutants can be similarly evaluated and also more precisely quantified using the luciferase reporter gene. Strategy for the determination of p53 allele functional fingerprints using the p53 Functional Array
x
Mate or MATa Yeast “p53-host” strain contains an integrated p53 allele or a p53 expression vector
Select diploids
MATα p53 Functional Array: > 20 isogenic strains that differ only in the sequence of a p53 response element (e.g., P21, BAX, MDM2, PIG3, AIP1, NOXA, GADD45, etc.) cloned upstream of the ADE2 reporter gene
Evaluate transactivation capacity using the color (red/pink/white) assay by replica plating onto plates containing different amounts of galactose to induce variable p53 expression.
Figure 4B. Transactivation assays using the functional array of p53 response elements. Over 20 isogenic yeast strains that differ only in the sequence of a p53 response element upstream of the p53 reported gene ADE2 (the luciferase reporter gene can also be used to quantify transactivation) are mated with the p53-host strain expressing a given p53 mutant. Diploids are selected on suitable media and then tested with the phenotypic transactivation assay to determine the functional fingerprint of the p53 allele being tested. The analysis of the dominance potential of p53 mutants can be performed when the p53-host strain is transformed with a second GAL1:WT p53 (or p63, or p73) expression cassette.
We have analyzed 20 tumor-associated p53 alleles for transactivation [222] and dominance (unpublished) using this protocol. The alleles represent different functional classes including loss-of-function, WT, and promoterselective. In particular, we tested a group of p53 mutations associated with familial breast cancer from patients affected by BRCA1 germline mutations. Under conditions of high expression these p53 mutants appeared as active as WT p53, both in mammalian and yeast functional assays [31, 187]. Interestingly, analysis with many p53 response elements at low and variable
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p53 expression clearly distinguished all mutants from WT p53. Several mutations showed residual function with at least half of the REs. The group of mutants associated with familial breast cancer exhibited subtle defects comprising both enhanced and reduced function with different promoters. Promoter-selective mutants that were classified as WT with a P21 RE (but mutant with BAX) instead showed reduced activity compared to WT p53 at low levels of expression. Similar reduction, but not loss of activity, was observed with other REs that are highly responsive to p53. The dominance assay at various low p53 expression levels was sensitive to p53 gene dosage in that one and two copies of the GAL1:WT p53 expression cassette were phenotypically distinguishable. Different degrees of dominance were revealed with a group of loss-of-function p53 mutants when heterozygous. Among the p53 alleles retaining partial function some were completely recessive (i.e., it appeared as if two WT copies of p53 were expressed) while others led to a phenotype corresponding to a reduction in gene dosage (i.e., hemizygous for WT p53).
2.9
Concluding comments: development of a p53-mutant functionality database and relevance to other transcription factors
While many p53 mutations associated with cancer may retain partial transactivation function and although several classification methods for p53 mutants have been attempted, it is not presently possible to predict a priori the behavior of a mutant protein. The yeast functional assays provide an important contribution to the functional classification of p53 alleles. It appears that the combination of low and variable expression of p53 alleles and the assessment of the transactivation capacity using many p53 response elements provides a greater sensitivity for the functional classification of p53 mutants than the standard yeast assays using high-constitutive p53 expression. When combined with the efficient method for constructing p53 alleles using in vivo mutagenesis and with the high-throughput selection and screening system using diploids formed between “p53-host” and “the p53 functional array” strains (see Figure 4) these tools allow for rapid development of p53 functional fingerprints. Hence, the yeast functional assay offers a practical means to develop a p53 mutant functionality database that enables the functional fingerprints for all the p53 mutants relevant to cancer to be linked with the IARC p53 mutation database. This information will become valuable in understanding the correlation between p53 functional status and tumor aggressiveness and responsiveness to therapy, thus providing directions for effective patient management in the clinical setting.
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An example of functional classification of 24 p53 mutants with 15 p53 REs is presented in Figure 5 (222; see also Recent Developments). As a general trend, transactivation capacity toward REs from apoptotic genes is more severely affected compared to cell cycle arrest and DNA repair genes. Five functional classes of p53 mutants can be identified [222]. For example, supertrans mutants, such as 121F and 123A, showed enhanced capacity with all or most of the REs. The V122A mutation showed instead a preferential increase in activity with a subset of Res. The mutant S215C that has been reported in familial breast cancer associated with BRCA1-germline mutations, showed subtle differences compared to WT p53. Interestingly, it showed a slight increase in transactivation capacity toward a few REs. The mutant R282Q is weak and shows a reduction in transactivation for all REs that range from ~50% to ~5%. Two mutants at codon 277 have split functions with enhanced activity toward some REs of the cell cycle arrest group, but loss-of-function with all apoptotic REs. Finally, three mutants appear to have almost completely lost transactivation potential. The system developed for assessment of p53 function can be applied to many sequence-specific transactivation factors. The combination of tight regulation of protein expression, rapid construction of responsive promoters, and phenotypic/quantitative assessment of transcription levels offers opportunities to investigate transactivation specificity for several families of transcription factors that recognize diverged promoter elements. The approach provides the means to study altered selectivity, intrinsic DNA binding, and differential transactivation in vivo, to isolate and characterize enhancer elements as well as cofactors, corepressors, and external modifiers of the transcription complex assembly.
ACKNOWLEDGMENTS Our thanks to Drs. Robbert Slebos, Daniel Menendez, Dmitry Gordenin, Tom Darden, Gilberto Fronza for advice, helpful discussions and comments on the manuscript. We also want to acknowledge that there are many additional valuable contributions and directions that have been made in the vast p53 field which we were unable to include in this review because of space limitations.
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ADDENDUM
RECENT DEVELOPMENTS Since the original submission, there have been several additional publications related to this topic. Rather than greatly extending this paper, we have chosen to incorporate a small number of recent, yeast-based studies relevant to our review. Using variable expression of p53 and measuring transactivation at many REs, we have shown that single amino acid changes in the p53 DNA-binding domain can have diverse effects on the ability to induce transcription from many target promoters [222] – see also section 2.9 and Figure 5. Based on these results, we have proposed to include p53 in a newly defined group of genes which we have called master genes of diversity. We defined a master gene for genetically derived phenotypic diversity in terms of “a single transcriptional activator (or repressor) that regulates many genes through different REs. Mutations of the master gene can lead to a variety of simultaneous changes in both the selection of targets and the extent of transcriptional modulation at the individual targets, resulting in a vast number of potential phenotypes that can be created with minimal mutational changes without altering existing protein–protein interactions” [222]. We have expanded our functional analysis of p53 REs [91] to establish response element rules that can be used for in silico searches of genomes. In particular, we have focused on the impact of the spacer between dimmerbinding sites and on sequence features in the core element of the RE (i.e., CWWG – see section 1.2.2) as well as local sequence-context effects in the homopurine (RRR) and homopyrimidine (YYY) repeats flanking the core. Contrary to predictions based on in vitro binding assay, our results indicated that the spacer is critical for RE transactivation potential, with a 2 nt spacer reducing activity of a strong RE down to ~10%. Spacers of 5 and 10 nt led to even stronger reduction in activity to ~1% of a full site (Inga, Menendez, and Resnick, in preparation). Our analyses also revealed that half-site REs (i.e., dimmer - binding sites) with consensus sequence and the CATG core signature can mediate low level of p53 transactivation and that a dimmer binding site with an adjacent monomer site (i.e., 3/4 sites) can mediate moderate transactivation. These findings have implications for addressing the functional impact of sequence variation in p53 REs and for identifying target genes in the p53 transcriptional network.
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Figure 5. Relative transactivation capacity of p53 alleles with mutations in the DNA binding domain [222]. Twenty-four human p53 mutations were examined in 15 isogenic yeast strains each containing a different p53 RE that regulates expression of the ADE2 reporter gene. The REs are ordered based on decreasing transactivation capacity with wild type p53 [91]. The p53 mutants are ordered according to their position in the primary sequence. The transactivation capacity of each allele toward each RE was determined using variable expression of p53 under the GAL1 promoter and compared to the activity of wild type p53. The relative transactivation capacities of mutants with respect to wt is presented in a form similar to that for expression arrays, with red greatly increased, green greatly decreased, blue loss-offunction, and black equal to wild type. The quantification is based on the amounts of p53 protein required for transactivation with wt or with a mutant allele. This was derived from the minimal amount of galactose required for transactivation to occur – see section 2.6.
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Besides mutations in p53 or in proteins involved in the p53 response pathway, genetic variation in promoter REs of individual p53 target genes can be expected to alter biological responses to stress. Based on our set of response elements rules, we have developed custom bioinformatics tools to address the evolution of p53 networks between human and mouse due to variation in RE sequences. We had previously observed that mouse and human p53 proteins exhibited identical transactivation capacities toward a large panel of REs [222]. We focused on 26 p53 target genes that belong to different functional groupings (including cell cycle control, apoptosis, and DNA repair). First, we examined the conservation of established p53 target REs in humans in the homologous promoter regions in mouse. Then we searched in both species for additional p53 functional REs in a 10 kb region encompassing the transcriptional start site. We found that ~60% of the REs are functionally conserved. Surprisingly, most of the genes involved in DNA metabolism and repair showed no conservation of RE sequence nor compensatory REs in the 10 kb promoter regions examined (Inga, Menendez, Jegga, Aronow, and Resnick, in preparation). A similar strategy was developed to identify single nucleotide polymorphisms (SNPs) in p53 REs that can lead to functional variation in the p53 regulatory network between individuals [223]. An approach was developed that combines a custom bioinformatics search to identify candidate SNPs and functional assays to assess their effect on p53 transactivation. Among ~2 million human SNPs examined, we identified over 200 located within putative p53 REs that are located in the proximity of transcriptional start site of an established gene. We then ranked the putative p53 REs based on predicted transactivation potential and functional impact of the SNP. Forty REs were predicted to mediate p53 transactivation in one allele and to be functionally altered by the SNP. Eight of these were evaluated in functional assays to determine both the activity of the putative REs and the impact of the SNPs on transactivation. All eight candidate p53 REs were functional, and in every case the SNP pair exhibited differential transactivation capacities. Additionally, six of the eight genes adjacent to these SNPs were induced by genotoxic stress or activated directly by transfection with p53 cDNA in human cells. Thus, the overall strategy efficiently identifies SNPs that may affect gene expression responses in the p53 regulatory pathway [223]. We have proposed that these SNPs in p53 response elements could contribute to human differences in responses to stress. The group led by Ishioka has reported the completion of a comprehensive analysis of the functional impact of single amino acid changes in the p53 protein [224]. Using site-directed mutagenesis they constructed 2314 p53 mutants (an average of nearly six amino acid changes at each codon) and measured transactivation in yeast using quantitative
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reporter systems. To improve the throughput of the functional assay they chose fluorescent reporter genes (EGFP and Ds-Red) and utilized eight different p53-binding sequences. To increase the readout from the fluorescent reporters, multicopy plasmids (based on a 2 µm replication origin) were used, although this choice may have led to high experimental variability (see Figure 4 [224]). The general conclusion from this impressive study was that nearly two-third of the p53 mutants in the DNA-binding domain were inactive; activity was instead mostly unaffected by amino acid changes both at the Nter and Cter of the protein, with the exception of the tetramerization domain [224]. Unfortunately, this publication contained mostly summary data for the whole library; hence a comparison of the results for individual mutants with other yeast-based assays was not possible. In a subsequent study [225], the same group reported that 142 of the 2314 p53 mutants exhibited temperature sensitivity (ts) toward at least one of the eight p53-binding sequences examined. Fifty-four mutants were tested in mammalian cell-based assays and nearly 90% of them were ts [225]. Standard quantitative criteria were adopted to classify ts mutants, based on the difference in activity between 30°C and 37°C. However, the screening identified as ts only 11 of 59 p53 mutations that had been described as ts by other groups and were available in the collection [225]. This discrepancy is indicative of significant differences in sensitivity or classification parameters between the various functional assays and emphasizes the need to compare transactivation data for individual mutants obtained with the different methodologies available. An effort to standardize and compare results of the various functional assays has been taken by the curators of the IARC p53 mutation database (http://www-p53.iarc.fr/index.html). The group has recently released a p53 mutant function database that can be downloaded and also searched online (http://www-p53.iarc.fr/p53MUTfunction.html). Data from overexpression of p53 mutant proteins in human cells and from yeast assays have been included. Results have been grouped into five categories: (1) retained wildtype activity; (2) loss-of-function; (3) gain of function; (4) dominant-negative effect; and (5) temperature sensitivity. Once all the available data are deposited and standardized, this database will be a valuable tool to assess whether functionality data contribute prognostic value to the analysis of p53 mutation status in cancer. Soussi et al., have instead used only the data set from the library of 2314 p53 mutants [224] to compare the functional impact of the mutations with the frequency of p53 mutations in the tumor database (http://p53.curie.fr) [226]. They concluded that more than 50% of the rare p53 mutants in the database display significant activity, while the most frequent mutations have lost function. The apparent heterogeneity of the functional consequences of
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p53 mutations associated with cancer further emphasizes the value of a comprehensive, accessible, and fully searchable p53 mutant function database to address the clinical relevance of p53 status in tumors.
REFERENCES 222. Resnick, M. A., and A. Inga. 2003. Functional mutants of the sequence-specific transcription factor p53 and implications for master genes of diversity. Proc. Natl. Acad. Sci. USA 100:9935–9939. 223. Tomso, D. J., A. Inga, D. Menendez, G. S. Pittman, M. R. Campbell, F. Storici, D. A. Bell, and M. A. Resnick. 2005. Functionally distinct polymorphic sequences in the human genome that are targets for p53 transactivation. Submitted. 224. Kato, S., S.Y. Han, W. Liu, K. Otsuka, H. Shibata, R. Kanamaru, and C. Ishioka 2003. Understanding the function–structure and function–mutation relationships of p53 tumor suppressor protein by high-resolution missense mutation analysis. Proc. Natl. Acad. Sci. USA 100: 8424–8429. 225. Shiraishi, K., S. Kato, S. Y. Han, W. Liu, K. Otsuka, M. Sakayori, T. Ishida, M. Takeda, R. Kanamaru, N. Ohuchi, and C. Ishioka. 2004. J. Biol. Chem. 272:348–355. 226. Soussi, T., S. Kato, P. P. Levy, and C. Ishioka. 2005. Reassessment of the tp53 mutation database in human disease by data mining with a library of tp53 missense mutations. Hum. Mut. 25:6–17. 227. Zacchi, P., M. Gostissa, T. Uchida, C. Salvagno, F. Avolio, S. Volinia, Z. Ronai, G. Blandino, C. Schneider, and G. Del Sal. 2002. The prolyl isomerase Pin1 reveals a mechanism to control p53 functions after genotoxic insults. Nature 419: 853–857.
Chapter 11 ABC TRANSPORTERS IN YEAST – DRUG RESISTANCE AND STRESS RESPONSE IN A NUTSHELL
Karl Kuchler and Christoph Schüller Max F. Perutz Laboratories, Department of Medical Biochemistry; Division of Molecular Genetics, Medical University Vienna, Campus Vienna Biocenter, Vienna, Austria
Key words:
1
Yeast, ABC protein, antifungal resistance, drug screening, detoxification, stress response
SUMMARY
ATP-binding cassette (ABC) proteins have been implicated in multidrug resistance phenomena and some are also intimately connected to prominent genetic diseases. Thus, many members of the ubiquitous superfamily of ABC proteins act at crossroads of vital cellular processes. For example, mammalian ABC proteins such as P-glycoproteins (P-gp) or multidrug resistance-associated proteins (MRPs) are associated with multidrug resistance (MDR) in various cancers. Likewise, homologues of mammalian ABC transporters in bacteria, fungi, or parasites are tightly linked to pleiotropic drug and multiple antibiotic resistance phenomena. Because yeast orthologues of mammalian MDR genes operate in this unicellular eukaryote, it has been acclaimed as a prime model system for studies on mammalian ABC proteins. Moreover, many efforts have been made during the past decade to exploit yeast as a model system for drug development and drug screening. In this chapter, we shall provide a comprehensive overview on yeast ABC transporters implicated in drug resistance and cellular detoxification. We shall discuss the regulatory network of pleiotropic drug resistance (PDR) in the budding yeast Saccharomyces cerevisiae. Finally, we 289 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 289–314. © 2007 Springer.
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shall reiterate the advantages and pitfalls associated with using baker’s yeast as a model system for the molecular analysis of mammalian ABC transporters, in particular its suitability and amenability for drug screening or even drug discovery related to the function of ABC transporters with medical importance.
2
INTRODUCTION
Although intrinsic drug resistance has been known for quite some time [15], MDR may develop under selective conditions such as those caused by drug exposure or stepwise selection of cells for resistance to xenobiotics or toxic small molecules [84]. MDR is often characterized by cross-resistance to many unrelated compounds [14], and while this is a matter of intense discussion, MDR of tumor cells or tissues may impair efficacy or even prohibit anticancer therapy in vivo. At least in great many in vitro studies using cultured cells, MDR phenotypes clearly cause cells to become refractory to chemotherapy [14, 84]. Complex MDR phenotypes may result from several mechanisms operating simultaneously or consecutively in a developmental process [84]. Mechanisms of MDR include decreased drug uptake, plasma membrane permeability changes, intracellular drug inactivation, transcriptional activation/deactivation, gene amplification, drug target gene mutations, or even compartmentspecific sequestration of certain drugs (Figure 1). Notably, similar mechanisms may counteract inhibitory effects of endogenous metabolites in case they are toxic or detrimental for growth. Importantly, ATP-dependent drug efflux from cells mediated by dedicated membrane transporters is considered a major cause of MDR [14]. Such drug efflux can be accomplished by socalled ABC transporters, which comprise one of the largest protein families with more than 3,000 known representatives operating in all living cells from bacteria to man [24]. Most ABC proteins are membrane-associated and display transport activities for great many different cytotoxic drugs or antibiotics. Substrate transport is powered by ATP hydrolysis, but ATP may also have important roles in regulating ABC protein function or substrate specificity. Although the domain organization of most if not all ABC proteins is conserved throughout evolution, the molecular basis for their broad drug substrate specificity has remained an unsolved mystery [24].
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Figure 1. Principle mechanisms of pleiotropic drug resistance in living cells. Complex PDR phenotypes can arise from several molecular mechanisms. PDR is typically characterized by cross-resistance to many structurally and functionally unrelated drugs. Each mechanism may operate on its own or in combination with another one at the same time or in a developmental process. Yellow balls indicate environmental xenobiotics or drugs; green balls, endogenous metabolites; N, nucleus; V, vacuole.
Similar to the situation in mammals, overexpression of ABC transporters in yeast or fungal pathogens leads to PDR [11]. MDR phenomena have also been described in parasites [106] and even in bacteria [156]. In fact, PDR in fungi [152] is at least in principle quite similar to MDR, since it can be invoked by imposing drug selection on S. cerevisiae cells [6]. Again, parasites [106] and bacteria [81] exposed to sublethal doses of xenobiotics will eventually aquire a multiple antibiotic resistance phenotype in vitro and in vivo. Genome sequencing uncovered 30 ABC proteins in yeast [26, 127, 138], some of which are structural and functional homologues of mammalian ABC pumps. The molecular basis and mechanisms of ABC-mediated drug resistance have thus been under intense investigation for many years. Studies in numerous laboratories disclosed the functions of most yeast ABC proteins, but also uncovered a complex network of transcription factors required for the dynamic regulation of ABC genes in response to many environmental cues. Because certain fungal ABC transporters are also implicated in drug resistance, stress response, as well as other cellular processes [10], yeast has been considered a well-suited model system for studies on ABC proteins of medical importance [11, 138]. Yeast strains lacking endogenous ABC pumps have been utilized as “test tubes” in functional cloning approaches. Closely related yeasts such as Pichia pastoris were also successfully exploited as eukaryotic hosts for the high-level
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expression of certain ABC transporters. Cross-complementation studies yielded important insights on mammalian, plant and parasite ABC transporters of the P-gp and MRP families. In this chapter, we provide a comprehensive discussion of how yeast has been beneficial for studies on drug resistance mechanisms mediated by ABC efflux pumps, and how these studies may aid future attempts to overcome ABC-mediated diseases.
3
FUNCTIONS OF YEAST ABC TRANSPORTERS IN DRUG RESISTANCE AND STRESS RESPONSE
According to their evolutionary relationships, yeast ABC proteins are classified into five distinct groups represented by the MDR, PDR, MRP/CFTR, ALDP, and YEF3/RLI families [127]. In this chapter, we shall limit our discussion on yeast ABC proteins (Table 1) implicated in cellular detoxification, stress response and drug resistance, and refer the reader to a widely available review literature on the function other yeast ABC proteins or non-ABC genes [68] of the PDR network [11, 68, 138]. Table 1. Membrane-associated yeast ABC transporters and some relevant transport substrates ABC protein Pdr5p/Sts1p
Vmr1p
Relevant substrates Antifungal drugs, steroids, myco-toxins, peptides Drugs, steroids, mutagens, 4-NQO Detergents, chloramphenicol, phenolic herbicides Membrane detergents? Oligomycin, reveromycin A, drugs C3-C7 weak acid anions, benzoate, sorbate GSH-conjugates, cadmium, DNB-GS Cadmium, DNB-GS, glucuronides Taurocholate, UCB, bilirubin Unknown
Nft1p
Unknown
Snq2p Pdr15p
Pdr10p Yor1p Pdr12p
Ycf1p Bpt1p Ybt1p/Bat1p
Length 1,511
Topology (ABC-TMS6)2
Location PM
References [13, 9]
1,501
(ABC-TMS6)2
PM
[129]
1,529
(ABC-TMS6)2
PM
[150]
1,511
(ABC-TMS6)2
PM
[151]
1,477
TMS11-ABC-TMS6ABC (ABC-TMS6)2
PM
[62]
PM
[111]
TMS11-ABC-TMS6ABC TMS11-ABC-TMS6ABC TMS11-ABC-TMS6ABC TMS11-ABC-TMS6ABC TMS11-ABC-TMS6ABC
VAC
[137]
VAC
[66, 132]
VAC
[105]
Unknown
SGD
Unknown
SGD [96]
1,511
1,515 1,515 1,515 1,592 1,524
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ABC, ATP-binding cassette; TMS, transmembrane segment; PM, plasma membrane; VAC, Vacuole; GS, glutathione S; UCB, unconjugated bilirubin; PM, plasma membrane; 4-NQO, 4-nitroguinoline-A-oxide; DNB-GS, (2,4-dinitrobenzene)-glutathione; UCB, unconjugated bilirubin; SGD, Saccharomyces Genome Database (http://www.yeastgenome.org/).
3.1
The plasma membrane ABC transporters – a first defense
Although most PDR transporters are found in the plasma membrane, most intracellular compartment membranes also harbor at least one or two ABC transporters (Figure 2). Well-known ABC pumps of the PDR family are the plasma membrane-localized efflux pumps Pdr5p [9, 13] and Snq2p [129], whose overexpression mediates pronounced resistance phenotypes. These transporters recognize a remarkable broad spectrum of xenobiotics and hydrophobic drugs, comprising hundreds of different compounds that are extruded to the extracellular space [13, 21, 35, 48, 57, 69, 88]. The first yeast drug pump identified was Snq2p [129], causing tolerance to mutagens such as 4-nitroquinoline-N-oxide (4-NQO), although Snq2p also modulates sensitivity to Na+, Li+, and Mn2+ ions [100]. PDR5/STS1/YDR1/LEM1 was then identified independently by several groups through its ability to mediate resistance to cycloheximide [9], cerulenenin and staurosporine [57], mycotoxins [13] or as glucocorticoid transporter [70]. Other closely related members of the PDR family include Pdr10p and Pdr15p, sharing more than 70% primary sequence identity with Pdr5p [151]. Their transport functions or substrates have remained obscure for quite some time. However, recent evidence suggests that Pdr15p is a stress response gene displaying signifycant transport capacity for chloramphenicol [153] and phenolic herbicides (Mamnun and Kuchler, unpublished data). Notably, a distinct expression regulation of Pdr15p and Pdr5p suggests a normal role of Pdr15p [153] and Pdr5p [91] in detoxification under adverse conditions and in actively growing cells, respectively. Another ABC transporter mediating drug efflux is represented by Yor1p, a plasma membrane pump of the MRP/CFTR family [62]. Yor1p was originally identified as a oligomycin transporter, but it can also transport a large spectrum of different drugs [21, 69]. Therefore, certain plasma membrane ABC transporters constitute a first defense line against environmental toxins. Some others may contribute to endogenous detoxification because they can extrude toxic metabolites that accumulate during cell growth. For instance, the plasma membrane pump Pdr12p, which is most closely related to Snq2p, is essential for adaptation to weak organic acid stress exerted by benzoate or sorbate [111]. Pdr12p also effluxes metabolites such as short chain carboxylic acid anions, and it is essential for
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weak acid stress adaptation [111]. Finally, the PDR transporters Aus1p and Pdr11p may be involved in sterol uptake under conditions of impaired ergosterol biogenesis [150], but a distinct drug efflux activity has not been established for them.
Figure 2. Membrane ABC transporters in yeast. Graphical presentation of characterized ABC transporters and their cellular localization in yeast. Membrane-associated yeast ABC transporters of the vacuolar membrane, the plasma membrane, the inner mitochondrial membrane, and the peroxisome are depicted. Only those membrane-associated ABC transporters are depicted for which literature data on their function is available. ER, endoplasmic reticulum; N, nucleus; VB, vacuole; M, mitochondrium; GV, Golgi vesicles; P. peroxisomes.
A second important mechanism of PDR involves the intracellular sequestration of metabolites or xenobiotics into the vacuole (Figure 2). Hence, the MRP/CFTR-like ABC pumps Ycf1p, Ybt1p, and Bpt1p perhaps constitute a second and equally important intracellular defense line in the vacuolar membrane against cytotoxic compounds or even stress-derived breakdown products [127]. Ycf1p mediates vacuolar detoxification of heavy metals, as well as glutathione S-conjugates [82, 137]. This explains the drug and heavy metal tolerance phenotypes of strains with increased YCF1 gene dosage, but also the hypersensitivity of cells lacking Ycf1p. Loss of Ycf1p also causes defects in vacuolar uptake of As(GS)3 [46, 137]. Similarly, a lack of Yor1p also causes cadmium hypersensitivity, indicating a functional overlap of plasma membrane Yor1p with vacuolar Ycf1p [21, 62]. Two additional MRP/CFTR members exist that have been implicated in vacuolar transport processes. The Ybt1p/Bat1p pump transports taurocholate [105] and magnetic resonance contrast agents across vacuolar membranes [109]. The closest Ycf1p homologue Bpt1p accounts for one third of the
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total vacuolar uptake of glutathione conjugates. Furthermore, Bpt1p mediates vacuolar delivery of cadmium and unconjugated bile pigments [66, 110, 132]. Interestingly, Bpt1p levels are strongly elevated in early stationary phase, suggesting that both Bpt1p and Ycf1p represent key players in vacuolar sequestration of glutathione conjugates, which is important during later stages of cell growth [66]. A new MRP/CFTR transporter of as yet unknown substrate specificity may be encoded by NFT1 [96]. This gene exists in some strain backgrounds as a fusion of the YKR103 and YKR104 ORFs. In the sequenced strain S288c, however, these ORFs are separated by a single stop codon, but this may in fact represent mutation or genomic polymorphism [96]. Finally, Vmr1p (YHL035c) could represent another MRP/CFTR-like transporter of the vacuolar membrane, but a cellular transport function remains to be disclosed (http://www.yeastgenome.org/). Notably, vacuolar detoxification of conjugated metabolites, xenobitics, or toxins by MRP/CFTR-like transporters, in addition to their cellular extrusion across the plasma membrane, represents a fundamental detoxification mechanism in plants [95]. Likewise, xenobiotic and drug detoxification in mammals is accomplished by their conjugation and subsequent removal from organs such as liver, kidney, or intestine by several MRP-like transporters [14]. The yeast MDR family also harbors mitochondrial ABC transporters, Mdl1p, Mdl2p [25], and Atm1p [79], all of which localize to the inner mitochondrial membrane [83]. None of these transporters has been associated with typical drug transport. However, the peptide exporter Mdl1p [155] may also serve a function in oxidative stress response as its overexpression suppresses a loss of Atm1p [19]. Atm1p is otherwise mediating mitochondrial export of Fe/S cluster precursors [65]. The ALD-family members Pxa1p/Pxa2p [131, 136] are found in the peroxisomal membrane [5], where they may mediate peroxisomal uptake of very long chain fatty acids. Interestingly, Mdl1p, Mdl2p, Pxa1p and Pxa2p, and Atm1p, are homologues of human ABC genes such as the TAP peptide transporters required for MHC class I antigen presentation [20], the adrenoleukodystrophy genes [5], and ABC7 involved in X-linked sideroblastic anemia and ataxia [4], respectively. What then is the physiological role of PDR-genes apart from their ability to confer basal drug tolerance and hyperresistance upon increasing their gene dosage? A hypothetical function of yeast ABC pumps might be a role in the asymmetric distribution of phospholipids in membranes or the membrane removal of damaged or oxidized lipids. The fact that membranedamaging agents such as herbicides, lysophospholipids, and other detergents strongly induce Pdr15p seems to further support this idea (Mamnun and Kuchler, in preparation). At least some fungal pumps [134] and their
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regulators [63] have indeed been implicated in membrane lipid transport or even phospholipid flipping (see also section 4). In fact, lipid homeostasis in the bilayer must undergo rapid and dynamic changes to form transient regions with distinct functions such as lipid rafts or even non bilayer phases [33]. However, the current evidence for a role of yeast ABC pumps in membrane lipid homeostasis, lipid-remodeling, or trans-bilayer transport [23] is vague at best, due to a lack of related phenotypes of cells lacking these pumps. Nevertheless, several MRP and MDR-like ABC transporters in the liver are tightly linked to membrane transport of sterols or phospholipids, since phenotypes of associated human diseases, mouse models lacking corresponding ABC transporters [67] or in vitro studies [114] strongly suggest a role of mammalian ABC transporters in lipid transport [14]. The most prominent and abundant yeast PDR pumps Pdr5p and Snq2p have the ability to transport hundreds of different compounds, including lipid-like substrates or even detergents. However, their physiological substrates, apart from detoxification of environmental xenobiotics, remain obscure. Notably, the expression regulation of the PDR5-family genes is quite distinct. Pdr5p is mainly expressed during logarithmic growth [91], whereas Pdr15p is strongly induced when cells approach stationary phase [153], and both Pdr10p (Wolfger and Kuchler, unpublished data) and Pdr15p [153] are highly induced by a variety of adverse conditions or metabolic stress. Further, Pdr12p effluxes weak organic acids such as C3–C7 carboxylate anions including toxic metabolites such as propionate [111], thus participating in the defense against metabolic stress. Taken together, a normal function of PDR transporters in detoxification and stress adaptation remains as the most plausible one, particularly considering the environmental cues yeast cells encounter in nature. In fact, it is feasible that yeast ABC transporters have a limited number of endogenous “drug-like” substrates, but have rather evolved to serve defensive functions in the everongoing “biological” warfare of various microbial species, including plants and their fungal pathogens that have to share similar ecological niches [143].
4
THE PLEIOTROPIC DRUG RESISTANCE (PDR) NETWORK – THE TRANSCRIPTION CONNECTION
Transcription must play a key role in modulating PDR, since cells ought to be able to discriminate between cellular defense against metabolites or if there is a need to combat environmental challenges. Indeed, most yeast PDR and MRP/CFTR genes (Table 1) form a complex genetic network known
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as the PDR network. Several related and to some extent promiscuous transcription factors (Table 2) control expression of target genes to allow for control of ABC pumps under a variety of growth conditions and environmental cues (Figure 3). For simplicity, Figure 3 only depicts ABC genes present in the PDR network, while it also includes several non-ABC genes such as major facilitator permeases and others [10, 30, 32, 68]. Table 2. Transcriptional Regulators of Yeast ABC transporters in the PDR Network Protein
TF family
Function Regulator of PDR network
Promoter motif PDRE
Pdr1p
Zn(II)2Cys6 - TF
Pdr3p
Zn(II)2Cys6 - TF
Pdr8p
Zn(II)2Cys6 - TF
Reference [8]
Regulator of PDR network
PDRE
[29]
Regulation of PDR15,
PDRE
[56]
YOR1, SNQ2 Yrr1p
Zn(II)2Cys6 - TF
Regulation of SNQ2, YOR1
PDRE
[22]
Yrm1p
Zn(II)2Cys6 - TF
Regulation of PDR, Yrr1p
Unknown
[86]
SRE
[145]
SRE
[145]
competitor? Ucp22p
Zn(II)2Cys6 - TF
Regulator of PDR11 and AUS1
Ecm22pp
Zn(II)2Cys6 - TF
Regulator of PDR11 and AUS1
Rdr1p
Zn(II)2Cys6 - TF
Negative regulator of PDR5
PDRE??
[54]
Ngg1p
Coactivator
Pdr1p modulator, HAT
Not applicable
[94]
PDRE??
[2]
WARE
[71]
YRE
[101]
YRE
[99]
STRE
[36]
complex Stb5p
Zn(II)2Cys6 - TF
Regulation of PDR
War1p
Zn(II)2Cys6 - TF
Required for stress
transporters induction of PDR12 Yap1p
bZip - TF
Regulation of YFC1, cadmium resistance
Cad1p/Yap2p
bZip - TF
Regulator of cadmium resistance by Ycf1p
Msn2p/Msn4p
Cys2His2 - TF
General stress response regulators
PDR, pleiotropic drug resistance; PDRE, pleiotropic drug resistance element; STRE, stress response element; YRE, Yeast AP-1 response element; TF, transcription factor; SRE, sterol regulatory element.
The first yeast gene capable of conferring PDR was in fact PDR1 that encoded a predicted Zn(II)2Cys6 transcription factor [8] rather than a membrane efflux pump. Further work showed that additional Zn(II)2Cys6
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regulators (Table 2), including Pdr3p [29], Yrr1p [22, 157], Pdr8p [56], Ucm22p and Ecm22p [145, 150], and War1p [71] play crucial roles in the regulation of yeast ABC pumps during normal cellular growth but also in response to adverse conditions. While several factors contribute to the regulation of ABC transporters in the network (Figure 3), Pdr1p and Pdr3p are still considered the master regulators of PDR [11, 68]. Pdr1p and Pdr3p constitutively localize to the nucleus, and control PDR genes as homo- and heterodimers [90] through so-called PDR elements (PDREs, 5′-YCCYCCGR-3′) in target promoters [30, 60]. A single cis-acting PDRE is necessary and sufficient to attract control by Pdr1p/Pdr3p [60, 61]. Deletion of both genes causes pronounced drug hypersensitivity phenotypes, since they regulate basal expression of several ABC transporters, including PDR5 [60], PDR10, PDR15 [151], SNQ2 [89], YOR1 [52], as well as other non-ABC genes implicated in the PDR phenomenon [30, 68]. In addition, Yrr1p as well as Pdr8p are involved in the regulation of Snq2p, Pdr15p, and Yor1p [22, 56, 157]. The presence of two PDREs in the promoters of PDR3 and YRR1 suggest autoregulatory loops in their control [21, 27, 157]. The scenario is even more complex, since Yrm1p appears to compete with Yrr1p for promoter occupancy on certain target genes, but a physiological relevance for such a competition is unclear [86]. Strikingly, Pdr1p and Pdr3p can exert opposing regulatory effects on target ABC genes that even encode highly homologous transporters. For example, a lack of PDR1 reduces PDR5 expression, but increases PDR15 mRNA levels. In turn, only Pdr3p, but not Pdr1p, is required for basal PDR15 expression [151].
Figure 3. ABC gene regulation within the yeast PDR network. The gene names in the ellipses in the center represent ABC target genes of transcriptional regulators depicted about and below. Transcription factors exerting regulatory effects are connected with target genes by an arrow. An arrow can indicate positive or negative control. Note that the cartoon only
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includes ABC genes for clarity and simplicity. Several non-ABC genes such as membrane transporters of the major facilitator family are also regulated by some of the transcription factors depicted in the cartoon. For details see text and references.
All GAL4-like and fungal-specific Zn(II)2Cys6 transcription factors share a rather simple molecular architecture: the N-terminal zinc cluster mediates DNA binding, while the regulatory and activation domains are located in the large C-terminal part [93]. Constitutive active, and sometimes dominant alleles of Pdr1p/Pdr3p as well as other Zn(II)2Cys6 regulators arise from mutations in the activation domain or in the middle homology region that is considered an inhibitory domain [93]. This is consistent with the fact that many gain-of-function mutations in PDR1/PDR3 are found in this region or in the C-terminal activation domains [17, 29, 68, 104, 133]. Hyperactive alleles of PDR1, PDR3, or YRR1 cause a pronounced PDR phenotype, since target pumps such as Pdr5p [98] or Snq2p [89] and Yor1p [157] are dramatically overexpressed. Most constitutive alleles of PDR1 and PDR3 were identified during genetic screens for increased drug tolerance [17, 63, 104, 117, 149]. Equivalent Pdr8p variants were generated by analogy to Pdr1p/Pdr3p, and in combination with microarray profiling, Pdr15p and Yor1p were identified as regulatory targets for Pdr8p [56]. Interestingly, dominant PDR1 and PDR3 alleles can also be specifically obtained by longterm selection of yeast cultures with the antifungal drug fluconazole [6]. Moreover, mutations of the fluconazole target gene ERG3 are also preferentially selected during the in vitro evolution of antifungal resistance [6]. Gain-of-function Pdr1p/Pdr3p variants have also been obtained in a mutant screen for genes defective in phosphatidylethanolamine endocytosis, implying a role of some Pdr1p targets in maintaining the phospholipid asymmetry at the plasma membrane [63]. Pdr1p/Pdr3p have also been implicated in the control of spingholipid biosynthesis through the regulation of Ipt1p [51]. Furthermore, the ABC transporters Aus1p and Pdr11p are regulated by Ucp2p and Ecm22p, two Zn(II)2Cys6 ergosterol pathway regulators that drive gene expression through sterol regulatory elements (SRE) in target promoters [145]. However, in all cases, the molecular signaling events leading to the activation of Zn(II)2Cys6 PDR regulators (Table 2) have remained enigmatic, and additional components have escaped genetic identification until now. It is possible that the PDR1/3 system represents a genetic sensing tool allowing for the rapid selection of hyperactive transcription factor genes as an alternative to complex drug recognition and signal transduction pathways. The advantage of such a general drug-sensing tool might be that it is not restricted to certain molecule classes, thereby providing versatility that could be beneficial for efficient xenobiotic defense. It is interesting to note,
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that several transcription factors of this family seem to respond directly to small molecules, perhaps inducing conformational changes that lead to their activation and recruitment of the mediator complex. For instance, Gal4p [154], Leu3p [50, 147], or Put3p [31] are activated by their cognate ligands via direct binding. Target pumps of the bacterial mar operon are also activated to induce antibiotic resistance after binding of several different antibiotics and chemicals to the Mar regulator [81]. By analogy, several mammalian MRP and MDR-like ABC genes are tightly regulated by heteroand homodimeric members of the RXR, FXR family, which are activated by ligand-binding. RXR and FXR regulators play important roles in lipid homeostasis, sterol transport, and hormonal response [39, 92, 124]. Although Zn(II)2Cys6 genes are restricted to fungal species, they may be viewed at least as functionally related to nuclear orphan receptors in mammals. However, activating drug or stress ligands have not been identified for fungal PDR regulators as yet. Additional transcription factors increase the complexity of the PDR network. For example, Ngg1p seems to interact with the C-terminal activation domains of Pdr1p thus reducing its regulatory activity [94, 121]. Further, Rdr1p appears to repress PDR5 transcription [54], while Sbt5p seems to engage PDREs to act as a positive regulator of PDR5 and SNQ2 expression [2]. Since transcription factors regulating the environmental stress response are targeting many genes [43], it is not surprising that some overlaps exist between PDR and stress response. For example, PDR15 regulation involves Pdr1p and Pdr3p, as well as the general stress response regulators Msn2p and Msn4p [153]. Moreover, PDR12 is regulated by the stress-specific transcription factor War1p through cis-acting WARE motifs in PDR12 [71]. Weak acid stress also activates the Msn2p/Msn4p general stress response [128]. However, Pdr12p induction does not need Msn2p or Msn4p, but only the War1p regulator, whose posttranslational activation involves a phosphorylation cycle [71]. Further, the bZip transcription factor Yap1p is a prime regulator of oxidative stress adaptation [28, 29, 101], and it also induces Snq2p in response to oxidative stress (Mahé and Kuchler, unpublished data). This Yap1p-mediated cadmium resistance requires Ycf1p, suggesting a direct regulation of YCF1 by Yap1p [148]. Notably, Ycf1p-mediated cadmium tolerance is also controlled through another bZip regulator, Cad1p/Yap2p [37]. In summary, the above-said illustrates the intense molecular crosstalk operating between ABC transporter regulation and stress response. Even mitochondrial dysfunction impacts gene expression within the PDR network [32, 53], but the physiological relevance needs further clarification. Fact is that the regulation of the PDR network utilizes a complex array of functionally distinct but structurally related and sometimes redundant
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transcription factors, all of which provide a dynamic expression control under normal and many adverse conditions. This system might have evolved to cope with rapidly changing growth conditions, so as to maintain the most appropriate transporter composition at the right cellular membrane at any given moment of the yeast life cycle.
5
FUNCTIONAL EXPRESSION OF MAMMALIAN ABC GENES AND DRUG DISCOVERY – A YEAST TOOLBOX
The pronounced drug hypersensitivity of certain pump deletion strains indicated that such strains are useful for susceptibility testing, drug identification, as well as functional cloning approaches. Simple susceptibility testing indeed revealed hundreds of potential substrates for Pdr5p, Snq2p, and Yor1p [69]. Similarly, drug profiling of strains with multiple deletions of ABC genes (YOR1, SNQ2, PDR5, YCF1, PDR10, PDR11, and PDR15), as well as the PDR1 and PDR3 regulators, disclosed the extraordinary hypersensitivity of such strains [119]. Our laboratory has constructed numerous isogenic strains lacking all plasma membrane as well as vacuolar ABC pumps (Schuetzer-Muehlbauer and Kuchler, unpublished data). We have also generated many combinations of ABC deletions, yielding dozens of strains with qualitative and quantitative distinct drug susceptibility spectra (Schuetzer-Muehlbauer and Kuchler, unpublished data). These host strains may in fact be useful for setting up cell-based high-throughput screening (HTS) assays in cases were compound libraries are limited in terms of drug concentrations. Interestingly, we have generated a strain lacking Pdr5p, Snq2p, the a-factor mating pheromone transporter Ste6p [73, 97] as well as the ERG6 gene [41]. This tester strain display an almost three orders of magnitude increased susceptibility to a vast range of hydrophobic drugs when compared to its wild-type parent (Pandjaitan and Kuchler, unpublished data). ERG6 was removed in this pump deletion background, because it has been known for quite some time that cells lacking Erg6p have abnormal plasma membrane ergosterol levels, which causes dramatic changes in overall cell permeability to ions and many drugs [41, 113, 130]. This supersensitive strain somehow “diffused” into the yeast community and it has indeed been successfully exploited for drug discovery approaches by HTS, leading to the identification of novel CDK inhibitors [49]. Because PDR results from efflux pump overexpression, simple gene dosage approaches allow for cloning of new ABC drug resistance genes.
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Indeed, this strategy identified the first Candida albicans ABC transporters, Cdr1p [112, 123] and Cdr2p [122], both of which mediate clinical antifungal resistance. The functional expression of C. albicans ABC transporters in baker’s yeast also uncovered a hitherto unknown caspofungin transport capacity of Cdr2p [126]. Importantly, this defined system also established that antifungal drug resistance can be efficiently reversed through the use of ABC efflux pump inhibitors [125] or it can be used for the molecular analysis of fungal drug pump function [103]. Similarly, MDR reversal has also been attempted in cancer therapy in a clinical setting as a means to restore anticancer drug response, but the success has been limited up to now [84]. To study functional conservation and to clone and identify novel drug transporters, several mammalian ABC genes have been functionally expressed in yeast. Cross-complementation studies yielded important information about the function mammalian, parasite, plant, and other fungal ABC genes. The first functional expression demonstrated that human Mdr1 P-gp is functional when expressed in yeast, since it confers resistance to valinomycin and actinomycin D [74, 142]. Interestingly, mouse mdr3, when expressed in yeast cells lacking the Ste6p a-factor transporter [72], rescues the mating defect [115], whereas human Mdr1 is unable to extrude the mating pheromone [74]. Likewise, pfmdr1 from Plasmodium falciparum appears to complement a loss of Ste6p, at least in certain strain backgrounds [146]. Mouse mdr3 in yeast also confers resistance to immunosuppressive drugs such as FK520 [116] or the anticancer drug dactinomycin [55]. Yeast expression of human Mdr2 demonstrated its physiological function as a phospholipid translocase [120]. Further, expression of MRPs, homologues of Ycf1p, revealed that human MRP1 as well as plant AtMRPs can complement the heavy metal sensitivity of cells lacking Ycf1p [42, 85, 140]. Nevertheless, the signals for correct trafficking of MRPs appear to work inefficiently in yeast, since MRPs and Mdr1 are also found in other cellular membranes than their normal cellular location in mammalian cells [74, 140]. Yeast has also been used for the analysis and expression of human ABC genes implicated in genetic diseases, including the CFTR [118] cystic fibrosis transmembrane conductance regulator [64, 107, 139], and the TAP transporters of antigen presentation, which are also functional as peptide transporters in yeast [144]. Furthermore, the human ABC7 gene [4], which is mutated in sideroblastic anemia with cerebellar ataxia (XLSA/A), was produced in yeast and complemented the defect of the yeast homologue ATM1 [12]. Baker’s yeast and other yeasts have also been exploited as single cell “bags” for the high-level expression and purification of endogenous [38] and
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heterologous ABC pumps [16, 59, 80]. For example, the PMA1 promoter of the plasma membrane proton pump, significantly boosted MRP1 expression, allowing for the production of milligram amounts of purified MRP1 [76, 77]. The related species P. pastoris is also suited for production of mammalian ABC proteins or individual nucleotide binding domains. Purified proteins can be reconstituted into in vitro systems, enabling studies on the catalytic cycle of ABC transporters, structural analyses [16, 80] and perhaps even crystallization trials [38]. Although expression of certain mammalian ABC proteins has been accomplished in yeast, many attempts to express mammalian or human transporters in yeast failed or gave inconsistent results. This may be because of nonfunctional or different trafficking signals, improper protein folding, aberrant cellular localization, lack of appropriate in vitro or in vivo assays, unknown drug substrates or just inappropriate strain backgrounds. Furthermore, the different lipid environment of yeast membranes may change or even inhibit ABC function. In fact, this is the case for the retinal ABC transporter, whose activity is even modulated by lipids in the native membrane environment [1]. Moreover, other ABC pumps even mediate membrane transport of lipids or sterols [75, 114, 120]. Changes in the normal lipid composition, like those caused by defects in ergosterol biosynthesis, dramatically modulate cell permeability [41, 113, 130] and may therefore also affect transport activity, substrate specificity, or even the proper folding of ABC pumps [135]. However, once expression is achieved and verified by appropriate functional assays, yeast does represent a useful host for the molecular genetic analysis of heterologous ABC transporters.
6
CONCLUSIONS AND PERSPECTIVES
Of all eukaryotic model organisms, yeast is perhaps the most versatile one, since a wealth of genetic and biochemical tools are available. For instance, a gene deletion collection is widely available (EUROSCARF, Germany; http://www.uni-frankfurt.de/fb15/mikro/euroscarf/), all yeast genes have been tagged with GFP [45], GST fusions are available, and gene arrays have become almost routine tools by now. Even protein arrays have been recently described [158, 159]. The yeast community benefits from highly developed gene ontology at the Saccharomyces Genome Database (http:// www.yeastgenome.org/) that comes along with most up-to-date online databases [34]. Of course, above all is the genetic tractability of yeast with its elegant genetics [40] and gene-replacement methods. This makes yeast a fascinating and irresistible system, enabling functional studies like in no other eukaryotic model system. In fact, yeast perhaps is the most promising
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model for systems biology approaches that are currently receiving considerable attention in the global science community [18]. After entering the post-genomic era, a series of elegant genome-wide approaches further sparked the general interest in yeast as a tool for drug discovery. For instance, genome-wide drug profiling [108], haplo-insufficiency screening [47, 87], and global metabolomics [3] indicated that yeast can be very useful for drug target identification. Haplo-insufficiency screening may even provide clues about targets of orphane drugs and disclose possible mechanisms of drug action [87]. The global description of transcription factor promoter-binding sites undoubtedly indicates incredible complexity, but at the same time suggest meaningful genetic and biochemical approaches to better understand transcription in general [78]. We are just beginning to understand the genetic interactions by exploring genome-wide synthetic lethality relationships of otherwise nonessential genes, which might be beneficial for future strategies to dissect complex genetic diseases in humans [141]. The knowledge about global protein interaction maps [7, 44] and perhaps their dynamic changes during cell growth is closing the circle. Nevertheless, as elegant as genome-wide approaches seem, one must not forget that they do not alleviate the need for mechanistic approaches to deepen the understanding of complex biological phenomena such as PDR development or ABC protein function both at the molecular level, and in a whole-cell context. To come to full circle, yeast is unlikely to loose its attraction as a valuable test tube for the functional characterization of other eukaryotic or mammalian ABC proteins. Once physiological substrates of ABC pumps have been identified, one will be able to produce tailor-made yeast strains with desired drug susceptibility, facilitating drug discovery through HTS approaches [49, 102]. The disadvantage of a generally reduced drug susceptibility of yeast cells compared to mammalian cells can be overcome by using supersensitive strains lacking certain ABC pumps [49, 58]. Another argument stems from the financial aspect, since whole cell-based assays in yeast are more than order of magnitude cheaper than comparable assays in mammalian cell culture. This way, yeast will perhaps contribute to a better understanding of other medically relevant ABC proteins, since the closed nutshell opens a door to a defined genetic system for analysis at the molecular level. Therefore, yeast must not be neglected as a model system for functional approaches on eukaryotic ABC transporters of medical importance, even in cases where no cellular function is known.
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ACKNOWLEDGMENTS Our research is supported by grants from the Austrian Science Foundation (FWF-15934-B08), by funds from the Austrian National Bank (OENB #9985), the “Herzfelder Family Foundation”, the Vienna Science and Technology Funds from the City of Vienna (WWTF-Project LS113), and in part by COST Action B16 action from the European Commission.
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Chapter 12 THE FHCRC/NCI YEAST ANTICANCER DRUG SCREEN Susan L. Holbeck1 and Julian Simon2 1
National Cancer Institute, Developmental Therapeutics Program, Information Technology Brach, Rockville, MD; 2Clinical Research Division, Fred Hutchinson Cancer Research Center, Seattle, WA
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INTRODUCTION
Anticancer drug discovery has historically been done empirically, screening for compounds that inhibit the growth of tumor cells in culture, or that are effective against implanted tumors in mice. This approach is analogous to the highly effective screens carried out in the 1950s and 1960s for antimicrobial agents and has had its successes – indeed, many of the anticancer drugs currently in clinical use were developed in this fashion. However, as our understanding of the molecular mechanisms underlying the development and growth of cancer cells has improved, new approaches to drug discovery have begun to build on this knowledge. The recognition that not all tumors arising in the same tissue are due to the same underlying defects calls for treatments regimes that will be tailored to these molecular alterations. The drug screen described in this chapter was initiated with the hypothesis that single gene changes that are often associated with particular hereditary and sporadic forms of cancer may serve as determinants of drug sensitivity [27]. In principle, there are two possible mechanisms by which a drug can be more toxic to a cell containing a particular genetic alteration. First, by a mechanism in which damage caused by the drug is normally repaired in a wild-type cell by a protein that has been deleted or altered in the mutant. An example of this is the sensitivity of mutants defective in recombinational repair of DNA double-strand breaks (DSBs) (e.g., yeast rad50 mutants) to agents that cause DSBs (e.g., the topoisomerase poison 315 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 315–346. © 2007 Springer.
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etoposide). The second mechanism is more interesting and potentially more important as a anticancer therapy strategy. In this scenario, genes that are normally nonessential are made essential by the deletion of the cancerrelated gene. Inhibition of the normally nonessential gene product by a small molecule leads to cell death in the mutant background. This phenomenon is called synthetic lethality and is an example of context-specific cytotoxicity. By screening for agents that elicit synthetic lethality in cells harboring cancer-related mutations, we may be able to identify a new armamentarium of context-specific anticancer agents. The drug discovery effort described here began in 1997 as a collaboration between the Developmental Therapeutics Program (DTP) of the National Cancer Institute (NCI) and the Fred Hutchinson Cancer Research Center (FHCRC) and used the yeast Saccharomyces cerevisiae to screen a large number of compounds for those that are selectively toxic to cells with alterations in defined genes. This screen utilized a panel of yeast strains, each with defined alterations in DNA repair genes or in cell cycle control genes. Differential sensitivity of various DNA repair mutant strains to a compound can identify pathways important for repair of damage caused by that agent. DNA damage repair pathways represented in this screen included DNA DSB repair (rad50 and rad52), ultraviolet (UV)-excision repair (rad14), DNA mismatch repair (mlh1), removal of O6-methylguanine (mgt1), and post-replication repair (rad18). Alterations in human homologs of these genes have been identified in human tumors. The DNA mismatch repair gene hMLH1 is mutated or silenced in hereditary and sporadic forms of colon cancer, and is one source of the “Microsatellite Instability” (MIN) phenotype. Defects in UV-excision repair lead to the cancer-prone syndrome Xeroderma pigmentosum. The breast cancer predisposing gene BRCA1 associates with components of the DSB repair pathway. The screen also included strains altered in cell cycle control – again, these reflect alterations observed in human tumors. Bub3p is a component of the mitotic spindle checkpoint, which ensures that chromosomes are attached to the mitotic spindle before allowing passage through mitosis. Defects in a component of this checkpoint have been seen in some human tumors, giving rise to the “Chromosome Instability” (CIN) phenotype. The RAD53 gene is part of two checkpoints, monitoring completeness of S-phase DNA synthesis, and also arresting cell growth in response to DNA damage in G2. The screen also includes a strain overexpressing the G1 cyclin, CLN2, which controls entry into S-phase. Finally, the SGS1 gene is involved in an ever-growing list of DNA transactions, including DNA replication, recombination, and telomere function. There are several human homologs of SGS1, two of which, BLM and WRN, lead to cancer-prone diseases, Bloom’s and Werner’s syndromes, respectively.
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Nearly 100,000 compounds from the repository of the NCI-DTP were screened for their ability to inhibit the growth of this panel of yeast strains. The following sections describe the design of the screen, the compounds used, more detailed information on the genes altered in the screen, general results, and some specific examples. The data generated by this effort are freely available to the public on a web site maintained by the NCI-DTP at http://dtp.nci.nih.gov/yacds/index.html. The results of the screen – the selective activity of agents in particular genetic contexts – have been interpreted by comparing the sensitivity of a panel of mutant yeast strains to Food and Drug Administration (FDA)approved anticancer agents [46] and other literature sources. Compounds showing structural similarity to agents known to exhibit increased toxicity in specific mutant backgrounds were segregated from structurally unique compounds.
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DESIGN OF THE SCREEN
The screen consisted of 3 stages. In the prescreen, Stage 0, compounds were screened in duplicate at a single dose (50 µM) against a panel of 6 strains, each harboring 1 or 2 gene alterations of interest. In addition, all strains were deleted for 3 additional genes. PDR1 and PDR3 encode transcription factors that control the expression of drug efflux pumps. ERG6 alters the ergosterol composition of the plasma membrane, and increases sensitivity to certain drugs. A complete list of the genotypes of the strains used can be found at http://dtp.nci.nih.gov/yacds/index.html. Compounds showing sufficient activity against at least one of the strains in Stage 0 testing were selected for testing in the next stage (Stage 1). Here compounds were tested in duplicate at 2 doses (5 and 50 µM) against the same 6 strains used in Stage 0. In addition to confirmation of activity, compounds were also examined for some degree of specificity. Compounds that met a quantitative criterion of selectivity – greater than fivefold difference in growth inhibition between the most and least sensitive strain – were selected for Stage 2 testing. Stage 2 used an expanded panel of yeast strains, with a single gene alteration of interest in each strain. As in the earlier stages, all strains (except one) were also deleted for the pdr1, pdr3, and erg6 genes. The single exception, rad50, ERG6, PDR1, PDR3 was included to assess the effect of drug uptake and efflux on growth inhibitory activity. Compounds were tested in duplicate at 5 doses (1.2, 3.6, 11, 33, and 100 µM). Data for Stage 0, Stage 1, and Stage 2 testing is available through the NCI-DTP web site at http://dtp.nci.nih.gov/yacds/index.html.
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GENES ALTERED IN THE SCREEN
3.1
DNA repair/DNA metabolism genes
3.1.1
MGT1
MGT1 encodes O6-methylguanine DNA methyltransferase, which removes the methyl group from guanine residues in modified DNA by covalently transferring the methyl group of either O6-methylguanine or O4methylthymine to a cysteine residue of Mgt1p [53, 56]. This residue also reacts with free 06-benzyl guanine, which is used therapeutically to inactivate the human homolog (MGMT) prior to chemotherapy with alkylating agents such as N-methyl-N′-nitro-N-nitrosoguanidine (MNNG) [38]. O6-methylguanine lesions, if unrepaired, lead to insertion of T rather than C opposite the altered base during DNA synthesis, which can then be acted on by mismatch repair activities [41]. This is mutagenic, as seen by the increased spontaneous mutation rate in mgt1 mutants [54]. Such lesions occur following exposure to certain environmental agents (e.g., methyl bromide) [39]. Mutations in the yeast gene can be complemented using the human homolog [54]. mgt1 mutants show increased sensitivity to streptozocin [46], MNNG [53] and other alkylating agents [55]. 3.1.2
MLH1
MLH1 is involved in repair of DNA mismatches (both single base mismatches and frameshifts) [19], in concert with MSH2 and PMS1. In addition to repairing DNA mismatches, mismatch repair prevents recombination between DNA sequences that have diverged [11]. Mlh1p binds ATP and has a weak ATPase activity – ATP binding may facilitate its interaction with other mismatch repair proteins [24]. MutLalpha, a heterodimer of Mlh1p and Pms1p, has DNA-binding activity [25]. Mlh1p was demonstrated to bind to the RecQ-like helicase Sgs1p [37] (which is mutated in other strains in the NCI yeast anticancer drug screen). This association is also seen with the human Mlh1 protein and the BLM gene, a RecQ-like helicase mutated in the cancer-prone Bloom’s syndrome. Human Mlh1 is mutated in many patients with the hereditary cancer-prone HNPCC, characterized by microsatellite instability (the so-called MIN phenotype). There are currently no compounds identified that specifically target mlh1 mutants. In fact, mismatch repair mutants show decreased sensitivity to some DNA-damaging agents used in anticancer chemotherapy [14].
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RAD14
RAD14 is one of a group of genes needed for excision repair of DNA damaged by UV and other lesions that distort the helix. This process involves recognition of damaged DNA (the step in which Rad14p participates), followed by removal of a short stretch of the damaged DNA strand and resynthesis, using the undamaged strand as a template [40]. RAD14 is the homolog of the human xeroderma pigmentosum complementation group A (XPA) gene [3]. Individuals with mutations in XPA exhibit increased sensitivity to UV light and an increased incidence of skin cancer. Rad14p and XPA each contain a zinc finger [22]. Rad14p is part of a large protein complex, which also contains Rad1p, Rad7p, Rad10p, Rad16p, Rad23p, RPA, RPB1, and TFIIH [43], although there is some controversy over whether the complex is preformed in the absence of DNA damage [48], or assembles in response to a lesion [23]. Strains defective in UV excision repair have increased sensitivity to nitrogen mustard [33]. The null mutant shows wild-type growth rates [3]. A rad14 null mutant shows increased sensitivity to the alkylating agents cisplatin, thiotepa, mechlorethamine and mitomycin C, to DNA synthesis inhibitors hydroxyurea, and actinomycin D, to compounds that induce DNA breaks, such as bleomycin and doxorubicin, and to pentostatin [46]. 3.1.4
RAD18
RAD18 is needed for post-replication repair of DNA damage, and mutants are hypersensitive to DNA alkylation. The Rad18 protein contains a ring-finger motif, through which it interacts with Rad6p, a ubiquitinconjugating enzyme. Mutations in the ring-finger motif in the human Rad18 homolog cause sensitivity to UV light, methyl methanesulfonate, and mitomycin C. In addition to ubiquitin-conjugating activity, the Rad18pRad6p heterodimer binds DNA and hydrolyzes ATP [2]. 3.1.5
RAD50
Rad50p, in concert with Mre11p and Xrs2p, forms a nuclease that is needed for repair of double-stranded DNA breaks using either homologous recombination [47] or nonhomologous end-joining (NHEJ) pathways [10, 15, 50]. The nuclease is manganese-dependent with both 3′–5′ exonuclease and single-stranded DNA endonuclease activities [18, 50]. Rad50p is required for activation of the Rad53 checkpoint in response to gammairradiation, which induces DSBs [20]. The Rad50p/Mre11p/Xrs2p complex is evolutionarily conserved – the analogous nuclease in humans is composed
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of Rad50, Mre11, and NBS1. Patients with defects in NBS1 have the cancerprone Nijmegen breakage syndrome and are hypersensitive to ionizing radiation. The breast cancer-associated gene BRCA1 associates with human Rad50 [57]. Rad50p also has a role in telomere maintenance [8, 12]. Structural data on Rad50p suggest that Zn++ coordinated hooks on 2 Rad50p molecules may function to keep the DNA strands involved in repair of a DSB connected with one another [28]. 3.1.6
RAD52
Like Rad50p, Rad52p is needed for repair of double-stranded DNA breaks by homologous recombination. However, Rad52p is not need for NHEJ and does not have a role in telomere maintenance. Mutants lacking RAD52 are extremely sensitive to DSBs, and are much more sensitive to Mitomycin C than is the rad50 mutant. The role of Rad52p appears to be to facilitate the interaction of the RecA homolog Rad51p with RPA and singlestranded DNA, thereby promoting DNA strand exchange [35]. Rad52p is needed for formation of Holliday junctions, an important recombination intermediate [58]. 3.1.7
SGS1
SGS1 is emerging as a player in many aspects of DNA metabolism, including telomere maintenance, DNA recombination, and DNA replication. It has recently been identified as a component of the intra-S checkpoint, which senses DNA damage during S-phase, although it has no role in the G1 and G2 DNA damage checkpoints [17]. An sgs1 mutant has a several 100fold increase in genome rearrangements following mild DNA damage [34]. Sgs1p is a member of the RecQ-like helicase family, which includes two human genes that cause cancer-prone syndromes when mutated. Defects in the human RecQ-like BLM gene cause Bloom’s syndrome, characterized by genome instability, sun-sensitivity, and increased incidence of cancer. Defects in a second human RecQ-like gene, WRN, cause Werner’s syndrome, characterized by premature aging, telomere shortening and increased incidence of tumors. A third homolog, RECQ4, has been linked to the premature aging and cancer-prone Rothmund–Thomson syndrome [31]. Two additional human homologs, of unknown function, have been identified. RecQ-like helicases may be required to prevent improper recombination at stalled DNA replication forks, in association with topoisomerase III [4, 29]. While deletion mutants of sgs1 are viable on their own, they are not able to tolerate deletion of certain other genes (synthetic lethality), including DNA repair or metabolism genes rad50, sae2, hpr5,
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mms4, top1, and pol32 [49]. Sgs1p interacts with the mismatch repair protein Mlh1p – this interaction is also seen with the human BLM and MLH1 proteins [37]. Sgs1p can unwind G-quadraplexes, a structure that can be formed by the G-rich single strand found at telomeres. This unwinding is inhibited by compounds that bind to G-quadraplexes [26].
3.2
Cell cycle control genes
3.2.1
BUB3
Bub3p is a component of the mitotic spindle checkpoint, together with Bub1p, Mad1p, Mad2p, Mad3p, and Mps1p. Bub3 mutants, lacking checkpoint function, fail to arrest following treatment with spindledestabilizing drugs, such as the tubulin-binding benzimidazoles, leading to chromosome mis-segregation and cell death. In the absence of spindle damage, mutations in the mitotic spindle assembly checkpoint may contribute to aneuploidy. The function of these checkpoint proteins is conserved, with higher eukaryotic homologs binding to kinetochores. Bub3p has WD40 repeats which mediate interactions with Mad2, Mad3, and Cdc20 (anaphase promoting complex activator) [16]. Bub3p is phosphorylated by Bub1p kinase. Mutations in human Bub1 have been associated with the CIN phenotype in colon carcinomas and other types of cancer [9]. 3.2.2
CLN2
CLN2 encodes a G1/S-specific cyclin that regulates the activity of the cyclin-dependent kinase (CDK) Cdc28p [42, 52]. Cln2p accumulates through G1, becomes phosphorylated and ubiquitinated, then is rapidly degraded by the proteasome, allowing cells to enter S phase. The strains used in the screen overexpress this cyclin using the strong inducible GAL1 promoter. Overexpression of G1 cyclins or loss of CDK inhibitors is thought to drive progression from the G1 to S phase in cancer cells, and high levels of cyclin E have been associated with a poor prognosis in breast cancer [30]. 3.2.3
RAD53 (mec2-1)
RAD53 is an essential gene in yeast, therefore the screen utilizes a point mutant in this gene, rather than a gene disruption. RAD53 was identified independently by several groups, giving rise to a host of synonyms (MEC2, SPK1, SAD1). Rad53p is a component of two cell cycle checkpoints in yeast.
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It is activated by the phosphatidylinositol-(3′) kinase-like protein kinases Mec1p and Tel1p in response to DNA damage (which also activates Chk1p), as well as by inhibition of DNA synthesis (which does not signal Chk1p) [45]. The human homolog, CHEK2, is a serine/threonine kinase that phosphorylates Cdc25C, preventing entry into mitosis [5]. CHEK2 is autophosphorylated in response to DNA damage, which upregulates its kinase activity.
4
COMPOUNDS
Compounds for the screen came from the repository of the NCI–DTP. This collection contains compounds submitted for screening as possible anticancer or antiviral compounds by a large number of suppliers. The suppliers include academic groups, government scientists, small biotechnology companies, and large pharmaceutical companies from all over the globe. The acquistion of compounds began in 1956 and continues to this day, with over 500,000 compounds currently registered. Interested parties can find details about submitting compounds at http://dtp.nci.nih.gov/. About half of the compounds are covered by a confidentiality agreement, which precludes the NCI from disseminating data derived from these compounds. Data derived from “open” compounds not covered by a confidentiality agreement is publicly available through the DTP web site, and includes compound structures, data from the NCI 60 human tumor cell line screen, the NCI AIDS screen, and the NCI yeast anticancer drug screen. In addition, if sufficient inventory is available, the NCI may be able to supply qualified researchers with small quantities of “open” compounds from the DTP repository for nonhuman use. The NCI does not routinely verify the structures of submitted compounds. In addition, the stability of different chemotypes varies widely. Thus, researchers are encouraged to verify the structure of any compounds before investing significant resources in lead development. Indeed, one of the compounds identified as a specific inhibitor of the DSB repair mutants rad50 and rad52 [13] had a structure different than that indicated by the supplier. Compounds submitted to the NCI–DTP may or may not have an identified mechanism of action. Many compounds were synthesized as chemically interesting structures, or were submitted as purified natural products, with cellular targets yet to be determined. Other submitted compounds have been rationally designed with specific targets in mind. Finally, many compounds are structural analogs of anticancer agents with known mechanisms of action. The end result is that the compounds in the
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NCI–DTP repository represent a broad range of chemotypes with a broad range of biological activities. To facilitate drug discovery in novel molecularly targeted screens, the DTP has put together several plated sets of compounds for distribution to qualified researchers. The diversity set is a group of 1,990 compounds selected to represent a broad range of chemical structures. All of the compounds in the set have been tested in both the NCI 60 human tumor cell line screen, as well as the NCI yeast anticancer drug screen. The mechanistic diversity set is somewhat smaller, with 879 compounds, and was selected for diversity of behavior in the NCI 60 human tumor cell line screen. Most of these compounds [662] have also been tested in the NCI yeast anticancer drug screen. Many of the compounds have been tested in the standard NCI–DTP anticancer drug screen, which employs a panel of 60 human tumor cell lines. Compounds are screened across a 5-log dose range for their ability to inhibit the growth of, or to kill, these human tumor cell lines. The pattern of which tumor cell lines are more sensitive, and which lines less so, provides a fingerprint for each compound. The comparison of these fingerprints between different compounds led to the development of the COMPARE algorithm [36], which allows users to search for compounds with similar patterns to a compound of interest. Compounds with similar mechanisms of action tend to have similar fingerprints. The COMPARE algorithm has led to the identification of numerous tubulin-binding agents, as well as a novel structural class of cyclin-dependent kinase inhibitors. NCI–DTP databases also include measurement of numerous “molecular targets” in the 60 cell line panel. The pattern of molecular target expression can be compared to compound sensitivity data, as a means of generating hypotheses about how a compound of unknown mechanism might function. The 60 cell line screen began in it’s current form in 1992. Compounds submitted in prior years may have been tested against P388 or L1210 mouse tumor models [51]. This data, structures of the compounds and access to the COMPARE algorithm, is freely available through the NCI–DTP web site at http://dtp.nci.nih.gov/. Some of the compounds showing selective activity in Stage 2 of the yeast screen had not been through the 60 cell line screen. Many of these were sent for 60 cell line testing, to allow users of the yeast screen data to link into other NCI–DTP data on these compounds.
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5
RESULTS
5.1
Stage 0 results
Compounds were screened in duplicate against a panel of 6 strains, each harboring 1 or 2 gene alterations of interest, plus mutations in pdr1, pdr3, and erg6. Stage 0 screening began in 1997 and was completed in 2001, with a total of 97,261 compounds screened (some multiple times). Data for 87,264 “open” compounds tested in Stage 0 are available on the DTP web site (http://dtp.nci.nih.gov/yacds/index.html). The remaining 10,003 compounds are covered by a confidentiality agreement with the NCI, and were screened while the Seattle Project was an NCI field station. Stage 0 was designed as a prescreen, to eliminate inactive compounds and determine whether compounds merited further multidose testing. Inhibition of the growth of at least one of the strains by at least 70% was required to trigger further testing in Stage 1. Nearly (14,466) compounds (15% of those tested) met this criterion. While the cutoff of 70% growth inhibition was selected for the purposes of this assay, this is somewhat arbitrary, and does not mean that compounds below the cut-off are inactive. Figure 1 displays the fraction of compounds with at least one strain meeting various growth inhibition cutoffs.
Figure 1. A total of 97,261 compounds were tested in Stage 0. The figure indicates the percentage of compounds where at least one of the strains exhibited growth inhibition equal to or greater than the indicated cutoff. 70% growth inhibition in one of more strains was chosen as the criterion for further testing.
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Stage 0 was done in duplicate. The median difference between measurements was 0.096 (based on fraction growth inhibition, with 1.0 denoting 100% growth inhibition), with the median of the average of duplicates 0.05. For measurements where the average of the duplicates was >0.7 growth inhibition (i.e., the criterion for further testing), the median difference between duplicate measurements was 0.014, with most (90%) of the duplicates having a difference of less than 0.11. There were instances where there were large differences between measurements. Compounds were chosen for further testing if any single measurement exceeded 70% growth inhibition.
5.2
Stage 1 results
Compounds meeting the selection criterion in Stage 0 screening were selected for rescreening in Stage 1. The same 6 strains as in Stage 0 were used, but compounds were tested in duplicate at 2 doses: 50 µ (the concentration used in Stage 0) and 5 µM. A total of 16,885 compounds were screened. Data for 14,837 “open” compounds are publicly available through the DTP web site. Stage 1 was designed to confirm activity of compounds, and to weed out generally cytotoxic compounds. Almost 5,270 compounds (31%) failed to reconfirm activity, with no strain at either dose showing growth inhibition of greater than 60%. Of these, 3,137 (19% of Stage 1 tested compounds) failed to inhibit any strain at any dose by greater than 30%. Compounds with low activity were not tested further. About 1,283 compounds (7.6%) inhibited the growth of all strains at both doses by at least 60%, and 773 compounds (4.6%) inhibited all strains at both doses by at least 90%. A subset of approximately 100 of these compounds were retested at sufficiently lower concentrations to allow IC50 determination. Most were found to be generally toxic, inhibiting the growth of all strains at approximately the same concentration. Because of this finding, the remaining potent compounds were excluded from further study, however these could also be candidate antifungal agents. Several are natural products that target essential processes such as protein synthesis. Those compounds that exhibited at least a fivefold differential between the most and least sensitive strains were selected for testing in Stage 2. The majority of the Stage 0 and Stage 1 (50 µM) measurements agreed fairly well with one another. There were 583,614 measurements in common (same compound tested on the same strain at the same dose). Of these 524,307 (90%) differed from one another by less than 10% growth inhibition (i.e., if one measurement was 60%, the other was between 50% and 70%).
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Nearly 541,823 (93%) differed by less than 20% growth inhibition, 15,447 (2.6%) had values that differed by more than 50% growth inhibition, and 5,574 (1%) had values that differed by more than 80% growth inhibition. 5.2.1
Selective Stage 1 compounds
Three of the Stage 1 strains contained single mutations of interest. For these, a brief summary of compounds selective for each follows. 5.2.1.1 Compounds specific for the bub3 strain in Stage 1 The bub3 mutant is deficient in the mitotic spindle checkpoint. It has been shown to be sensitive to compounds that interact with tubulin (such as benzimidazoles) as well as other components of the mitotic spindle. Approximately two hundred eighty-four compounds selectively inhibited the growth of the bub3 mutant (selectivity defined here as the bub3 strain exhibiting at least 40% growth inhibition, that the bub3 inhibition be at least 5 times greater than that of the least sensitive strain, and that all other strains were at least one-third less sensitive than the bub3 strain). Somewhat surprisingly, the majority fall into structural classes without an identified mechanism of action. Only 4% of the bub3-selective compounds were structural analogs of compounds known to interfere with tubulin (including steroid analogs), 4% contained heavy metals, 3% were analogs of compounds that cause DNA damage, and 2% were analogs of compounds that affect cell cycle or signal transduction. bub3-selective compounds will be discussed in more detail in the Stage 2 results (section 5.3.1). We believe that there are two reasons why more spindle poisons, which are abundant in the NCI collection [36], were not identified as bub3-selective agents in the screen. First, although yeast tubulin is highly homologous to human tubulin, subtle differences in ligand-binding sites confer resistance to several antimitotic agents (e.g., Paclitaxel) [21] in yeast. Second, the liquid growth assay used in the drug screen relies on growth differences, as measured by changes in optical density of the culture, over a small number of cell divisions, typically 7–8 divisions, whereas colony formation assays require up to 20 divisions. Cells arrested by spindle poisons in mitosis continue to increase in size and therefore optical density leading to an underestimation of growth inhibition in checkpoint proficient (BUB3+) strains in the liquid assay. This is actually an advantage, allowing us to identify agents with other mechanisms of action. 5.2.1.2 Compounds specific for the rad50 strain in Stage 1 The rad50 mutant is defective in DSB repair, so is expected to be sensitive to compounds that introduce DSBs into DNA. This class would
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include topoisomerase I (Topo I) and topoisomerase II (Topo II) poisons, as well as bleomycins. In addition, agents that cause DNA damage (e.g., cisplatin) are more toxic to the rad50 mutant than to wild type. For these agents, however, the rad18 mutant is the most sensitive. 311 compounds showed selectivity for the rad50 mutant in Stage 1 testing (analogous to that defined for the bub3 strain in section 5.2.1.1). Many of these compounds (58%) were known Topo I or Topo II poisons, structural analogs of topoisomerase poisons, or compounds with planar ring structures that might plausibly intercalate into DNA. While 3% of the rad50-selective compounds contained heavy metals. The remainder of the rad50-selective compounds did not fall into structural classes expected to cause DSBs. rad50-selective compounds will be discussed in more detail in the Stage 2 results (section 5.3.7). 5.2.1.3 Compounds specific for the rad53 (mec2-1) strain in Stage 1 The RAD53 gene is needed for both the S-phase DNA synthesis checkpoint and the G2 DNA damage checkpoint. Fifty-six compounds exhibited selectivity for the mec2-1 strain (analogous to that defined for the bub3 strain in section 5.2.1.1.). And 13% of these compounds were structural analogs of compounds known to interfere with DNA synthesis (e.g., antifolates). Eleven percent of the mec2-1 selective compounds contained heavy metals, a further 9% have structures suggesting they may be DNA intercalators. The majority of the Stage 1 mec2-1 selective compounds do not fall into structural classes expected to trigger either of the RAD53dependent checkpoints. rad53-selective compounds will be discussed in more detail in the Stage 2 results (section 5.3.8).
5.3
Stage 2 results
Compounds that showed selective activity in Stage 1, with the strain with the maximum growth inhibition at least five times greater than the strain with the least growth inhibition, were selected for testing in Stage 2. A total of 3,498 compounds were tested in duplicate at 5 doses (1.2, 3.7, 11, 33, and 100 µM). Compounds were screened against a panel of 13 strains, 2 of which serve as wild-type controls. The remainder contained a single gene alteration of interest. All but one of the strains was also deleted for the pdr1, pdr3, and erg6 genes. The rad50 EPP+ strain, wild-type for ERG6, PDR1 and PDR3 genes, was included to determine whether further studies with these compounds would be affected by these drug sensitivity mutations. Data are publicly available for the 3,137 compounds that are not covered by a confidentiality agreement. Stage 2 data, including bar graphs, dose-response
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curves, links to compound structures and other screening data on these compounds may be accessed at http://dtp.nci.nih.gov/yacds/index.html. Nearly 703 of the Stage 2 tested compounds (20%) failed to inhibit the growth of any of the strains by 70%. While active, 1,091 compounds (31%) did not show much selectivity between the strains. The remaining 49% of the tested compounds showed some degree of selectivity for one or more strains. These compounds are discussed in more detail below. 5.3.1
Compounds selective for the bub3 strain in Stage 2
Using the criteria of a minimum 70% growth inhibition, 29 compounds were found to be highly specific for bub3, inhibiting only the bub3 strain at 2 or more of the tested doses. A further 14 compounds inhibited the growth of the bub3 mutant, plus one additional strain at 2 or more doses. A total of 286 compounds inhibited bub3, either alone or in combination with several other strains, for at least one dose.
Figure 2. Structures are shown of representative compounds selective for the spindle checkpoint mutant bub3 in Stage 2 testing. NSC 1331 was selective for bub3 at all 5 doses. Two compounds with related structures were also bub3-selective. NSC 4000206 and two related compounds were bub3-specific at multiple doses.
Figure 2 contains structures for selected compounds highly specific for bub3. As stated previously, the bub3 strain is expected to be sensitive to microtubule-targeting compounds. None of the major spindle poison compound classes (i.e., vinca alkaloids, taxanes, benzimidazoles, or colchicine analogs) scored as bub3-selective hits. The 28 bub3-selective agents are a structurally diverse set of compounds, although there are some scaffolds represented multiple times in this group. The 3 most highly specific compounds (NSCs 1331, 16706, and 8513) may be structurally related to Pedicin (NSC 255993), which is reported to inhibit microtubule assembly [1]. Three bub3-selective compounds (NSCs 16461, 400206, and 30548) share a planar tricyclic anthrone structure, which could plausibly intercalate into DNA. A dose-response curve for one bub3-specific
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compound, NSC 1331, is shown in Figure 3. Very few of the bub3-selective compounds were related to known tubulin-interacting agents, which are highly represented in the DTP compound database. However, many of the compounds that bind mammalian tubulin do not interact with yeast tubulin [6].
Figure 3. Growth inhibition as a function of dose is plotted for cells treated with NSC 1331. This compound specifically inhibited the growth of the bub3 spindle checkpoint mutant at all doses tested in Stage 2, without significant inhibition of any of the other strains.
A number of the bub3-selective compounds were tested further in Rat1a fibroblasts and human tumor cell lines. The following compounds had effects in mammalian cells consistent with triggering of the spindle checkpoint. NSC 5062, while having no effect on tubulin polymerization, did cause an accumulation of mammalian fibroblasts in G2/M, with a defect in chromosome segregation. NSC 142496-treated mammalian cells accumulate in G2, with the formation of tubulin rings. NSC 319447 treatment led to mono-attached chromosomes in mammalian cells. NSC 672053 disrupted microtubules in mammalian cells. 5.3.2
Compounds selective for the CLN2 overexpressing strain in Stage 2
Two compounds were moderately specific for the strain overexpressing the G1 cyclin CLN2, inhibiting only CLN2oe at 2 doses. Structures of these compounds are displayed in Figure 4. NSC 180198 is related to the carboxaldehyde thiosemicarbazone (e.g., triapine) class of ribonucleotide reductase inhibitors. NSC 659206 is a bis-trihalocarbinol derivative of cyclohexanone, unrelated to known biologically active agents. A total of 64 compounds inhibit CLN2oe +/– several other strains at least 1 dose.
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Figure 4. These two compounds were selectively toxic to the G1 cyclin CLN2-overexpressing strain at two of the doses tested in Stage 2. NSC 180198 is related to ribonucleotide reductase inhibitors.
5.3.3
Compounds selective for the mgt1 strain in Stage 2
There were no compounds that inhibited only the mgt1 strain at more than one dose. A total of 139 compounds inhibited mgt1 +/– 1 or more strains at least 1 dose. The mgt1 mutant strain is known to be sensitive to alkylative DNA damage in general and methylation at guanine residues at the O6 position in particular. Even methylating agents, however, are more toxic to the rad6 and rad18 post-replication repair mutant strains than to the mgt1 strain. It is therefore expected that the mgt1 mutant strain would be most sensitive among the panel of strains to few, if any, compounds. 5.3.4
V.3.4. Compounds selective for the mlh1 strain in Stage 2
There were no compounds that inhibited the growth of only the mlh1 mutant at 2 or more doses. A total of 164 inhibited the mlh1 strain +/– 1 or more other strains at least 1 dose. Mlh1p is a component of mismatch repair in yeast. There are no known agents that are selectively toxic to the mlh1 deletion strain. 5.3.5
Compounds selective for the rad14 strain in Stage 2
One compound, haloperidol (NSC 615296), inhibited the growth of just the rad14 mutant at 3 doses. Haloperidol is used clinically as an antipsychotic and is reported to bind to dopamine receptors [44]. Rad14p is part of the nucleotide excision pathway for repair of bulky DNA adducts and cross-links. Haloperidol does not contain reactive chemical groups that would suggest DNA damage as the mechanism of action. Among the FDAapproved agents, cross-linking agents mechlorethamine (nitrogen mustard)
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and cisplatin are among the most active agents against the rad14 deletion mutant [46]. As with the mgt1 mutant however, even with these agents, the rad14 strain is not the most sensitive strain in the panel: the most sensitive strains to mechlorethamine and cisplatin are rad6 and rad18 post-replication repair mutants. Structures of two rad14-selective agents are shown in Figure 5.
Figure 5. Structures are shown for two compounds that selectively inhibited the growth of the rad14 excision repair mutant.
5.3.6
Compounds selective for the rad18 strain in Stage 2
A large proportion of the Stage 2 tested compounds were highly specific for the rad18 mutant. About 94 compounds (3% of all Stage 2 compounds) inhibited only the rad18 mutant at 2 or more doses. Figure 6 presents structures for 4 representative compounds.
Figure 6. Structures are given for representative compounds selective for the post-replication repair mutant rad18 in Stage 2 testing. NSC 86324 is a nitrogen mustard, as were a number of other rad18-seletive compounds. NSC 142226, an ---halocarbonyl, and NSC 3750, a nitroaromatic, are representative of compound classes found multiple times among the rad18selective agents.
As already mentioned, the rad18 strain represents a loss-of-function in post-replication, or daughter-strand gap repair. Loss of either rad6 and rad18 confers sensitivity toward a broad range of DNA damaging agents. Compounds such as cisplatin, thiotepa, carmustine, hydroxyurea, bleomycin
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as well as ionizing radiation are more toxic to rad18 mutant strain than to wild type yeast. It is not surprising then that the vast majority of compounds showing rad18 selectivity are electrophiles capable of damaging DNA. The most highly represented classes of electrophiles include: nitrogen mustards (e.g., NSC 86324) (i.e., bis-[2-chloroethyl]-amines) related to the FDAapproved agents mechlorethamine, melphalan, chlorambucil, and cyclophosphamide; α-halocarbonyl compounds (e.g., NSC 142226); nitroaromatics (e.g., NSC 3750) and aza-aromatic N-oxides (e.g., NSC 5094). Interestingly, several compounds that appear to have low potential for causing DNA damage (e.g., NSC 136325, a bromo-fluorene derivative) also score as rad18 selectives. 5.3.7
Compounds selective for the rad50 and rad52 strains in Stage 2
A large number of Stage 2 tested compounds exhibited specificity for the DSB repair mutants rad50 and rad52. Almost 121 compounds (3% of Stage 2 tested compounds) were highly specific for these strains, inhibiting only the DSB repair mutants at 2 or more doses. Representative data is shown in Table 3. Three of the compounds were exceedingly specific, inhibiting only the DSB repair mutants at all 5 doses. This included NSCs 295501 and 603071, analogs of the Topo I poison camptothecin; and NSC 669380, a synthetic epi-podophylotoxin (i.e., related to etoposide) derivative.
Figure 7. Representative structures of rad50/rad52-specific compounds are shown. NSC 295501 is an analog of the topoisomerase I poison Camptothecin – this class is highly represented among the agents selective for the DSB repair mutants. The remaining three structures represent classes of known topoisomerase II poisons – NSC 669380 is an epipodophyllotoxin, NSC 5159 is Chartreusin, and NSC 637992 in a pyrazoloacridine. All of these compounds have planar ring structures that intercalate into DNA.
An additional 10 compounds inhibited the growth of only DSB repair mutants at 4 of the 5 doses. Half of these have structures that might be expected to interfere with topoisomerases. Chartreusin (NSC 5159) is reported to bind DNA and inhibit the catalytic activity of Topo II [32]. NSC 302991 is a camptothecin analog, NSC 343501 is an analog of Amsacrine,
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a Topo II poison. NSCs 129364, 637992 (pyrazoloacridone), 669965 (pyrimidoacridone) have planar ring structure with the potential to intercalate into DNA. Of the remaining 108 rad50 rad52 specific compounds, 92 are DNA intercalaters or Topo II poisons (or analogs thereof ), and 13 are analogs of camptothecin. The remaining 16 compounds are structurally unrelated to known topoisomerase poisons. A total of 953 compounds inhibit rad50 or rad52, either alone or in combination with a few more strains, in at least one of doses. 5.3.8
V.3.8. compounds selective for the rad53 (mec2-1) strain in Stage 2
Six compounds were highly specific for the rad53 mutant strain, inhibiting only this strain in 2 of the doses. Of the 5 not covered by a confidentiality agreement, 2 are structurally related to the antifolate methotrexate, which is expected to trigger the S-phase DNA synthesis checkpoint. The remaining 3 contain heavy metals (Au, Ru, Os). A total of 195 compounds inhibit the rad53 strain, either alone or in combination with other strains, in at least 1 of the tested doses. 18 compounds (9%) (including the 2 already mentioned) are analogs of nucleosides, antifolates, or other compounds expected to interfere with DNA synthesis. 21 compounds (11%) are related to topoisomerase poisons, alkylating agents, or other compounds that introduce DNA damage. 5.3.9
Compounds selective for the sgs1 strain in Stage 2
There were no compounds highly selective for the sgs1 mutant. A total of 436 compounds inhibited sgs1, alone or in combination with a few more strains in at least one dose. Of these 12 (3%) are analogs of compounds that interfere with DNA synthesis, 14 (3%) are DNA damaging agents (alkylating, topoisomerase poisons, etc.). 5.3.10
Compounds selective for several strains in Stage 2
A number of compounds were highly selective for multiple strains, inhibiting the growth of only those strains by at least 70% at 2 or more doses. For the purposes of this section, rad50 and rad52 will be counted together, as both are needed for DSB repair, although there are a few compounds that show differential activity between these two DSB repair mutant strains. The predominant pattern observed for compounds that target more than one strain is the activation of DNA damage responses. This is perhaps not surprising given the high number of DNA damage response
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mutants in the yeast strain panel and the abundance of DNA damaging agents in the NCI compound collection.
Figure 8. Growth inhibition as a function of dose is plotted for cells treated with NSC 16258. This compound was selectively toxic to the DSB repair mutants rad50 and rad52. It is structurally unrelated to known topoisomerase poisons, which make up the majority of the rad50/rad52 selective agents, and thus represents a novel structure selective for the DSB repair mutants.
Figure 9. The structure of NSC 368252, a compound with modest selectivity for the rad53 checkpoint mutant is shown. In contrast to the more highly selective rad53 compounds, this agent is not structurally related to known antimetabolites of DNA damaging agents.
In general, the rad18 mutant is the single most sensitive strain to DNA damage and any compound that elicits a multistrain sensitivity profile that includes rad18 has to be viewed as a potential DNA alkylator, cross-linker, or cutter. Table 1 lists the NSC numbers and strain selectivity for compounds that target multiple strains.
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Table 1. Compounds listed in this table exhibited selectivity for multiple strains at two or more doses. The strains inhibited, and the number of doses with this profile are indicated. For this table, the rad50EP+ strain was not included. NSC
Strains inhibited
Number of Structural characteristics doses with this pattern
613609
bub3 rad50
3
Planar ring
607088
bub3 sgs1
3
Mn++
400655
mec2 mlh1
3
Planar ring
21276
mec2 rad50 rad52
2
2-Chloro-1,3-dinitrobenzene
117619
rad14 rad50 rad52
2
1,2-Dibromo-1,2-diphenylcyclopropane
134981
rad14 rad50 rad52
2
1,2-Diphenyl-cyclopropene
645457
rad14 sgs1
2
665606
rad14 sgs1
2
1510
rad18 rad50 rad52
2
Proflavine (Planar ring)
6208
rad18 rad50 rad52
2
Alkylator
21662
rad18 rad50 rad52
2
Phenazine bis-N-oxide (Planar ring)
81052
rad18 rad50 rad52
2
Streptonigrin related (Planar ring)
141586
rad18 rad50 rad52
2
Planar ring
400244
rad18 rad50 rad52
3
Planar ring
402910
rad18 rad50 rad52
3
Phenazine bis-N-oxide (Planar
405789
rad18 rad50 rad52
2
(Planar ring)
409304
rad18 rad50 rad52
2
Phenazine bis-N-oxide (Planar
ring)
ring) 507478
rad18 rad50
3
622143
rad18 rad52
2
24047
rad18 sgs1
4
bis-aromatic (Planar ring)
669264
rad50 sgs1
2
Guanine diphosphate ether
403546
bub3 rad18 rad50
3
683910
bub3 rad18 rad50
2
609959
mec2 rad50 rad52 sgs1
3
Planar ring Camptothecin analog (Planar ring)
50076
rad14 rad18 rad50 rad52
2
Nitrogen mustard
668523
rad14 rad18 rad50 rad52
2
Nitrogen mustard
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Number of Structural characteristics doses with this pattern
632
rad18 rad50 rad52 sgs1
2
Aromatic propylene oxide ether (Planar ring)
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rad18 rad50 rad52 sgs1
2
Aromatic propylene oxide ether
338423
rad18 rad50 rad52 sgs1
2
Aromatic propylene oxide ether
664867
rad18 rad50 rad52 sgs1
2
612071
wt1 mgt1 mlh1 sgs1
3
Weak uracil analog
612072
wt1 mgt1 mlh1 sgs1
2
Weak uracil analog
612074
wt1 mgt1 mlh1 sgs1
2
Weak uracil analog
658900
CLN2oe bub3 rad18 rad50
2
665773
CLN2oe rad14 mgt1 sgs1
2
108708
mec2 rad18 rad50 rad52 sgs1
2
154894
rad14 rad18 rad50 rad52 sgs1
2
329226
rad14 rad18 rad50 rad52 sgs1
4
Planar ring
373980
rad14 rad18 rad50 rad52 sgs1
2
Planar ring
652888
rad14 rad18 rad50 rad52 sgs1
2
Nitro-acridine (Planar ring)
270346
CLN2oe bub3 mec2 mgt1 mlh1
2
665727
CLN2oe rad14 rad18 rad52 mlh1 sgs1
2
621482
mec2 rad14 rad18 rad50 rad52 mgt1 mlh1
3
sgs1 wt2 78572
mec2 rad14 rad18 rad50 rad52 sgs1
3
Planar ring
373981
mec2 rad14 rad18 rad50 rad52 sgs1
2
Planar ring
382770
mec2 rad14 rad18 rad50 rad52 sgs1
2
610190
mec2 rad14 rad18 rad50 rad52 sgs1
3
Ellipticine analog (Planar ring)
667644
mec2 rad14 rad18 rad50 rad52 sgs1
2
Nitro-acridine (Planar ring)
673793
mec2 rad14 rad18 rad50 rad52 sgs1
2
Nitro-acridine (Planar ring)
673795
mec2 rad14 rad18 rad50 rad52 sgs1
2
Nitro-acridine (Planar ring)
673799
mec2 rad14 rad18 rad50 rad52 sgs1
2
Nitro-acridine (Planar ring)
673805
mec2 rad14 rad18 rad50 rad52 sgs1
2
Nitro-acridine (Planar ring)
404027
rad14 rad18 rad50 rad52 mgt1 mlh1 sgs1 wt2
2
(Planar ring)
5.3.10.1 Compounds selective for 2 strains Ten compounds inhibited the rad18, rad50, and rad52 strains, but none of the other strains, in at least 2 of the doses. Three of these, NSCs 21662,
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402910, and 409304 are phenazine bis-N-oxides. These compounds undergo bioreductive activation to generate radical species. A fourth compound, proflavine (NSC 1510), has a somewhat related acridine structure. NSC 81052, a quinoline quinone that is related to the left - hand portion of Streptonigrin, exhibits specificity for these three strains. Streptonigrin is reported to cause multiple types of DNA damage, including DNA breaks and DNA adducts [7]. The remaining rad18, rad50, rad52 selective compounds (NSCs 6208, 141586, 400244, and 405789) are structurally unrelated. While NSC 6208 is an alkylator, the remaining compounds appear to be of low reactivity. A single compound, NSC 507478, was selective for the rad18 and rad50 strains, inhibiting just these two at three of the tested doses. This is one of the few compounds that showed inhibition of the rad50 mutant without affecting the growth of the other DSB repair mutant, rad52. There is no information available on mechanism of this compound. A single compound, NSC 622143, was selective for the rad18 and rad52 mutants, while not inhibiting the rad50 mutant or any of the other strains. Nothing is known regarding the mechanism of action of this compound. Given the high concordance in sensitivity of rad50 and rad52 strains to both topoisomerase poisons and DNA alkylating agents, compounds that affect these strains differentially are of potential interest in dissecting the functions of Rad50p and Rad52p. Two compounds were selective for the rad50 and sgs1 mutants. One of these is covered by a confidentiality agreement. The other compound (NSC 669264) is a guanine diphosphate ether containing a long aliphatic tail. Unlike most compounds that inhibit the rad50 strain, these compounds were much less active against the other DSB repair mutant, rad52. Both rad50 and sgs1 have been implicated in telomere function. Two compounds (NSCs 645457 and 665606) were selective for rad14 and sgs1 at two of the doses. These compounds are unrelated to one another. Neither contains electrophilic groups and both were relatively inactive in the 60 cell line screen. Since Rad14p is normally involved in nucleotide excision repair of DNA damage, these compounds may highlight an additional function of this protein. Two compounds, NSCs 117619 and 134981, inhibited the growth of the rad14, rad50, and rad52 strains, but no others, at two doses. These compounds, a 1,2-dibromo-1,2-diphenyl-cyclopropane and a 1,2-diphenylcyclopropene, share somewhat related structures and may behave as electrophiles. A single compound (NSC 21276) was selective for rad53 and rad50 rad52 at two doses. NSC 24047 was highly selective for the rad18 and sgs1 mutants, inhibiting the growth of just these strains in four of the tested doses.
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This bis-aromatic appears to have little potential to cause DNA damage. NSC 400655 exhibited specificity for both the rad53 and mlh1 mutants at three of the doses. Very few compounds showed this degree of selectivity for the mismatch repair mutant. One compound, NSC 607088, was selective for the bub3 and sgs1 strains at three of the doses. A single compound (NSC 613609) selectively inhibited the growth of the bub3 and rad50 strains at 3 doses. This compound is one of the few seen that distinguishes between the 2 DSB repair mutants. Since, with the exception of the DNA damage profile of some or all of the rad18, rad14, rad50, rad52, rad53, and sgs1 strains, there is little literature precedent for overlapping function of the mutants in our panel, it is difficult to make definitive statements about the mechanism of action of compounds that target multiple strains. It is impressive, however, to note the seemingly large number of cellular responses demonstrating subtle differences in the compounds’ mechanisms of action. Subtle differences such as these may be the deciding factor between compounds that make it to the clinic and those that do not. 5.3.10.2 Compounds selective for 3 strains Two compounds, NSCs 50076 and 668523, were selective for the rad14, rad18, rad50, and rad52 strains at 2 doses. These compounds share a related structure related to the nitrogen mustard mechlorethamine (bis-[2chloroethyl]-amine) and are likely DNA alkylating and cross-linking agents. Four compounds exhibited selectivity for the rad18, rad50 rad52, and sgs1 strains. Three of these (NSCs 632, 634, and 338423) are structurally related, highly reactive aromatic propylene oxide ethers and are also likely DNA damaging agents. The fourth compound (NSC 664867) is structurally unrelated but contains several electrophilic functionalities and may damage DNA as well. Two compounds (NSCs 403546 and 683910) were selective for bub3, rad18, and rad50. While not close structural analogs, these two do share a carbon double-bonded to a nitrogen. NSC 180198 inhibited the growth of only the bub3, CLN2oe, and rad50 strains at two doses. A single compound, NSC 609959 was selective for the rad53, sgs1, and rad50 rad52 mutants at three doses. 5.3.10.3 Compounds selective for more than 3 strains Nine compounds were selective for the rad53, rad14, rad18, rad50 rad52, and sgs1 strains. This can be interpreted as a broad DNA damage response. Five of these (NSCs 667644, 673793, 673795, 673799, and 673805) share similar structures, and demonstrated quite potent activity in the NCI 60 human tumor cell line screen, with GI50 values of 10–7 to 10–8 M.
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These compounds are called nitracrines (nitro-acridines) and are DNAintercalating alkylating agents. Although the exact mechanism of action is undefined, these compounds have received attention in Europe, where several have undergone clinical trials. The remaining 4 compounds (NSCs 78572, 373981, 382770, and 610190) are structurally unrelated, and were about 2 logs less active in the 60 cell line screen. Interestingly, NSC 610190 is a synthetic ellipticine derivative. Several of these DNA-interactive agents have been evaluated in pre-clinical studies. Four compounds were selective for rad14, rad18, rad50, rad52, and sgs1 in at least two of the doses. Two of these compounds (NSCs 329226 and 373980) have structures similar to one another, while the other two (NSCs 154894 and 652888) are structurally unrelated. NSC 652888 is related to the nitracrines mentioned in the previous section. One compound, NSC 665773 showed specificity for the CLN2oe, rad14, mgt1, and sgs1 strains at 2 doses. A single compound (NSC 658900) was selective for bub3, rad18, CLN2oe, and rad50 rad52 at 2 doses.
5.4
Correlation of Stage 2 and Stage 1 results
For the 3 strains with a single mutation of interest in Stage 1 testing, we can ask whether the Stage 2 data are consistent with that from Stage 1 testing. The discussion here pertains to those compounds that were highly specific (i.e., that inhibited only the given strain in at least two of the Stage 2 doses) for the bub3 and rad53 strains. In addition, those compounds highly selective for rad50 (with or without rad52) are compared to the Stage 1 rad50 results. For the remaining strains, the use of multiple alterations in the Stage 1 strains would confound this type of analysis. Most of the 29 compounds highly specific for bub3 in Stage 2 testing were also specific for the bub3 mutant in Stage 1 tests. For 21 of these compounds, the bub3 mutant was the most sensitive strain in Stage 1 testing. One of the Stage 2 bub3-selective compounds was not selective for bub3 in Stage 1. The remainder of these Stage 2 bub3-selective compounds inhibited the bub3 mutant in Stage 1 testing, but inhibited the growth of some of the other strains to the same extent. Five compounds were highly selective for the rad53 mutant in the Stage 2 assay. For four of these, the rad53 mutant was the most sensitive strain in Stage 1. The fifth compound inhibited the growth of the rad53 stain in Stage 1, but inhibited other strains to the same degree. 121 compounds were highly selective for the rad50 strain (and rad52) in Stage 2 testing. For 112 of these, the rad50 mutant strain was the most sensitive in Stage 1. The other nine compounds inhibited the rad50 mutant, as well as other strains in Stage 1.
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Thus for the 155 compounds for which the Stage 1 and Stage 2 results can be readily compared, the vast majority (88%) were selective for the same strain in both stages. A further 11% were somewhat less selective in Stage 1 assays, inhibiting other strains in addition to that seen in Stage 2. Only one of the these had discordant Stage 1 and Stage 2 results.
6
EFFECT OF ERG6, PDR1, AND PDR3 MUTATIONS ON DRUG SENSITIVITY
The erg6, pdr1, pdr3 (epp) mutations were included in all of the strains used in Stage 0 and Stage 1 and in all but one strain in Stage 2 to increase drug uptake (erg6) and decrease drug efflux (pdr1 and pdr3). By including both rad50, erg6, pdr1, pdr3 (rad50epp-) and rad50, ERG6, PDR1, PDR3 (rad50EPP+) strains in the Stage 2 panel, we were able to quantify the effect of the “drug-sensitizing” mutations. We stratified rad50-selective compounds based on the following criterion. For each compound, if there were at least two doses at which the rad50epp- strain but not rad50EPP+ had a 70% or greater growth inhibition, then the compound was counted as having increased activity in the mutant “drug uptake and efflux” background. Of the 116 rad50-selective compounds, 66 compounds, or 57% had increased toxicity due to the epp mutations. For all Stage 2-tested compounds, about one-third were aided by the drug-sensitizing mutations, with the rad50eppstrain half a log more sensitive than the rad50EPP+ strain.
7
PDR AND MDR
Pleiotropic drug resistance (Pdr), genes in yeast encode a number of ATP-binding cassette (ABC) transporter proteins that function as detoxifying efflux pumps in a manner analogous to the mammalian multiple drug resistance, or MDR proteins. Since multidrug resistance is a major problem in cancer chemotherapy, we wondered whether sensitivity to PDR1 and PDR3 in yeast status correlated with MDR sensitivity in human cells. Of the compounds where rad50epp- was at least tenfold more sensitive than the rad50EPP+ strain, 161 have been tested in the 60 human tumor cell line screen. Only 6% of these showed a significant negative correlation with MDR activity in the 60 cell line panel.
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SUMMARY
A major goal of anticancer drug discovery and development in the new century is the identification of compounds that kill cancer cells selectively without causing the catastrophic damage to normal tissues that is the hallmark of current cytotoxic chemotherapy agents. Such selective agents will arise in a number of ways. First, through rational selection of drug targets. For example, the BCR-ABL translocation that is typical of chronic myelogenous leukemia (CML) creates a hyper-active form of the ABL tyrosine kinase. Inhibition of ABL, and BCR-ABL by analogy, is a reasonable strategy to inhibit the growth and possibly kill BCR-ABL expressing cells. Since normal cells have low levels of ABL activity, inhibition of ABL in normal tissues is not expected to be a problem. This approach led to the identification of Gleevec (imatinib mesylate), which has proven to be exceptionally effective and free of side effects, validating the rational target selection strategy. Other tumors, however, lack obvious targets. While many breast cancers overexpress cyclin E, leading to increased CDK activity, inhibition of that activity has, to date, failed to result in impressive clinical responses. Another approach for the development of selective anticancer agents is through phenotypic, cell-based screening for compounds that selectively target specific cancer-related contexts and take advantage of synthetic lethality. This is the approach outlined in this chapter. We have assembled a panel of isogenic yeast strains containing alterations in DNA damage response pathways that are homologous to those often mutated in human cancers. This panel of strains was screened against nearly 100,000 compounds from the NCI–DTP repository for differential growth inhibition. The ultimate aim of this project is to identify agents that will target tumors harboring the DNA damage mutations represented in the yeast strain panel. If the major hurdle of the rational approach is the selection of “valid” anticancer drug targets, then the major hurdle of our unbiased approach is the identification of the drug targets. Many of the compounds identified in these screens have very promising selectivity for a specific mutation but it will take more time until we know the molecular targets and mechanisms of action for these compounds. The yeast data provides a very different metric of biological activity, a metric that complements the NCI’s human cancer cell line screen. The panel of strains was assembled based on a vast literature on the drug sensitivity of various DNA response mutant yeast strains as well as our preliminary evaluation of FDA-approved, small molecule anticancer agents. We were able to evaluate agents and make rational choices as to whether a particular agent was expected to target a mutant strain and, therefore, represented a known mechanism of action, or whether the agent was
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structurally unrelated to known compounds and potentially represented a new mechanism of action and a new target. It is the latter class of compounds that are the most interesting and will be studied in more detail in the future. The screen yielded over 1,500 compounds that showed selective growth inhibition for one or more of the strains in the panel. Overall, the screen design yielded highly reproducible results with 99% of the Stage 2 selective compounds showing similar activity in Stage 1 screens. Selectivity for individual strains was assessed using both strict numerical criteria as well as a less quantitative, experience-based criteria. For example, several compounds selective for the rad18, rad50, or bub3 strains showed selective growth inhibition over the entire concentration range, with IC50s for mutant and wild type strains differing by more than 100-fold. On the other hand, the mlh1 and CLN2 overexpression strains yielded only mildly selective compounds having at best a modest twofold difference in sensitivity. Thus for each strain in the panel we applied slightly different criteria for the selection of compounds. The results presented in this chapter highlight the value of anticancer drug screening using cell-based assays in genetically- defined model organisms. This represents a unique resource, with data for nearly 90,000 compounds freely available through a public web site at http://dtp.nci.nih. gov/yacds/index.html.
ACKNOWLEDGMENTS The authors wish to thank Richard Klausner, Leland Hartwell, and Stephen Friend for initiating this project; Heather Dunstan, John Lamb, Philippe Szankasi, David Evans, and Michele Cronk for implementing the screen; and Edward Sausville, Jill Johnson, Tim Myers, Daniel Zaharevitz and David Segal for database and web site development, and for project oversight.
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50. Usui, T., T. Ohta, H. Oshiumi, J. Tomizawa, H. Ogawa, and T. Ogawa. 1998. Complex formation and functional versatility of Mre11 of budding yeast in recombination. Cell 95:705–716. 51. Venditti, J. M., R. A. Wesley, and J. Plowman. 1984. Current NCI preclinical antitumor screening in vivo: results of tumor panel screening, 1976–1982, and future directions. Adv. Pharmacol. Chemother. 20:1–20. 52. Wittenberg, C., K. Sugimoto, and S. I. Reed. 1990. G1-specific cyclins of S. cerevisiae: cell cycle periodicity, regulation by mating pheromone, and association with the p34CDC28 protein kinase. Cell 62:225–237. 53. Xiao, W., B. Derfler, J. Chen, and L. Samson. 1991. Primary sequence and biological functions of a Saccharomyces cerevisiae O6-methylguanine/O4-methylthymine DNA repair methyltransferase gene. EMBO J. 10:2179–2186. 54. Xiao, W., and T. Fontanie. 1995. Expression of the human MGMT O6-methylguanine DNA methyltransferase gene in a yeast alkylation-sensitive mutant: its effects on both exogenous and endogenous DNA alkylation damage. Mutat. Res. 336:133–142. 55. Xiao, W., and L. Samson. 1992. The Saccharomyces cerevisiae MGT1 DNA repair methyltransferase gene: its promoter and entire coding sequence, regulation and in vivo biological functions. Nucleic Acids Res. 20:3599–3606. 56. Xiao, W., and L. Samson. 1993. In vivo evidence for endogenous DNA alkylation damage as a source of spontaneous mutation in eukaryotic cells. Proc. Natl. Acad. Sci. USA 90:2117–2121. 57. Zhong, Q., C. F. Chen, S. Li, Y. Chen, C. C. Wang, J. Xiao, P. L. Chen, Z. D. Sharp, and W. H. Lee. 1999. Association of BRCA1 with the hRad50-hMre11-p95 complex and the DNA damage response. Science 285:747–750. 58. Zou, H., and R. Rothstein. 1997. Holliday junctions accumulate in replication mutants via a RecA homolog-independent mechanism. Cell 90:87–96.
Chapter 13 YEAST AS A MODEL TO STUDY THE IMMUNOSUPPRESSIVE AND CHEMOTHERAPEUTIC DRUG RAPAMYCIN
John R. Rohde, Sara A. Zurita-Martinez and Maria E. Cardenas Department of Molecular Genetics and Microbiology, Duke University Medical Center, Durham, NC 27710, USA
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INTRODUCTION
Rapamycin is a natural product of a soil bacterium and has potent immunosuppressive, and antiproliferative actions. First identified in 1975 as an antifungal drug, rapamycin languished in obscurity after it was found to cause bone marrow suppression [112]. Interest in rapamycin was later rekindled when it was discovered to be structurally related to the potent T-cell inhibitor FK506 [65]. Subsequent studies, conducted first in yeast and then in mammalian cells, revealed the molecular basis of therapeutic action. Rapamycin diffuses into the cell and binds to a small cellular protein, FKBP12, forming an FKBP12–rapamycin protein–drug complex that is the active intracellular agent. This complex then binds to and inhibits the Tor kinases, which function in nutrient sensing pathways that control cell growth and differentiation (Figure 1) (reviewed in [101]). The Tor kinases and FKBP12 are conserved from yeast to worms, flies, plants, and humans. Rapamycin has received FDA approval as an immunosuppressant, and more recently as an antiproliferative agent to inhibit restenosis following cardiac stenting [84]. Recent studies indicate that rapamycin and its analogs will find additional clinical applications as chemotherapeutic agents, topical
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immunosuppressive therapies in dermatology, and novel antifungal agents [33, 45, 83, 88, 116]. We review here the use of yeast as a model to elucidate the molecular basis of therapeutics for this exciting natural product. The history of the budding yeast Saccharomyces cerevisiae as a model to identify the targets and molecular basis of therapeutic action of rapamycin and other immunosuppressive drugs began with the discovery of rapamycin (sirolimus), cyclosporine A (CsA), and FK506 (tracrolimus) themselves. These immunosuppressive drugs were identified in three independent screens conducted at three different pharmaceutical companies. CsA was discovered at Sandoz Pharmaceuticals in a screen of soil samples for inhibitors of a mixed lymphocyte response (MLR) assay. CsA is produced by a fungus, Tolypocladium inflatum, which was isolated from a soil sample
Figure 1. TOR responds to diverse signals to regulate cell growth. TOR is activated by compounds associated with a favorable energy status including ATP, amino acids, and phosphatidic acid. TOR is inactivated in response to nutrient depletion, mitochondrial dysfunction as well as rapamycin–FKBP12. Active TOR promotes cell growth while inactivation of TOR results in growth arrest and autophagy in yeast and growth arrest and apoptosis in higher eukaryotes.
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from northern Norway [13]. CsA received FDA approval in 1983 as an immunosuppressant to prevent and treat graft rejection in organ transplant recipients, revolutionized organ transplant therapy, and became the gold standard for immunosuppressive therapies. FK506 was subsequently discovered at Fujisawa Pharmaceuticals in 1987 in a soil sample taken from the Tsukuba region of northern Japan and found to be a macrolide produced by the soil bacterium S. tsukubaensis [65]. FK506 (tacrolimus) received FDA approval in 1994 and has gone on to have considerable impact in transplant medicine. Rapamycin was discovered from a screen for novel natural products in 1975 at Wyeth–Ayerst Pharmaceuticals [112, 125]. In this case, the screen was for antifungal activity, and the rapamycin producing bacterium, S. hygroscopicus, was discovered in an isolate from the beaches of Easter Island (Rapa nui). Rapamycin remains one of the most potent anti-Candida drugs ever discovered [5], however the early finding that it caused bone marrow suppression halted development as an antimicrobial agent. It was only following the discovery of FK506 in 1987, and the appreciation that FK506 and rapamycin are structurally related, that rapamycin was resurrected from the shelf and studies began again in earnest to understand its molecular targets and possible therapeutic applications. Studies in yeast that began in the late 1980s defined the molecular targets of rapamycin, contributed to elucidate the mechanism of action of FK506 and CsA, and fueled comparable studies in mammalian T-cells that led to considerable insights into the molecular basis of therapeutic action (reviewed in [20]). Early studies from Merck demonstrated that FK506 and rapamycin inhibit T-cell proliferation by blocking different signaling pathways [38, 39]. FK506, like CsA, blocked the T-cell antigen response pathway necessary for signaling cascades to drive expression of hundreds of genes required for T-cell activation. In contrast, rapamycin had no effect on the T-cell antigen response pathway, but instead blocked T-cell proliferation in response to interleukin-2 (IL-2). FK506 and rapamycin were found to act as reciprocal antagonists [38, 39], suggesting that the two exerted their actions via a common target. Concurrently, biochemical studies led to the identification of a small abundant cellular binding protein, FK506 binding protein of 12 kDa (FKBP12), which is bound with high affinity to either FK506 or to rapamycin [49, 117]. However, given that FKBP12 is an abundant, ubiquitous protein expressed in all cells in the human body, a general view was that its drug-binding activity might not be involved in a very specific action in lymphocytes. Studies in yeast resolved this dilemma, and established unequivocally the central role of the FKBP12 protein in rapamycin action. Around this time, an FKBP12 homolog that shared 54% amino acid sequence identity with the human FKBP12 protein was identified in the
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yeast S. cerevisiae [52, 132]. Subsequently the crystal structures were solved for both the yeast and the human protein and found to be essentially superimposable [104, 123]. The gene encoding the yeast FKBP12 protein was cloned and disrupted. Whereas wild-type yeast cells are exquisitely sensitive to growth inhibition by rapamycin, with a minimum inhibitory concentration (MIC) of 25 ng/ml, yeast cells lacking FKBP12 were completely viable and only slightly reduced in growth rate [52, 67, 132]. Thus inhibition of an essential function of FKBP12 could not explain the potent toxic action of FKBP12. Taken together, these findings support a model in which both the FKBP12 protein and its ligand rapamycin are both required to exert a toxic effect in yeast. These findings also established unequivocally that FKBP12 plays a central role in the action of rapamycin, and the next challenge then was to identify the molecular target of the FKBP12–rapamycin complex. The targets of the FKBP12–rapamycin complex, the products of the TOR1 and TOR2 genes (target of rapamycin), were discovered in a genetic screen in yeast searching for rapamycin resistant mutants [51]. Mutations in three different genes were identified. First, mutations in the FPR1 gene encoding FKBP12 were found to be recessive, and resulted from amino acid substitutions that based on structural studies of the FKBP12–rapamycin complex were predicted to be critical for rapamycin binding. Importantly, mutations in two other genes identified were genetically distinct from FPR1, mapped to two different genomic locations and conferred dominant, or semidominant, drug resistance in genetic crosses. Based on an unusual genetic behavior between alleles of these three genes, known as nonallelic noncomplementation, it was proposed that the three might form a physical complex. The TOR1 and TOR2 genes were subsequently cloned by the Hall and Livi laboratories, revealing that they encode extremely large proteins, ~280 kDa, that Tor1 and Tor2 are homologs of each other, and that both share a C-terminal domain with homology to lipid and protein kinases [18, 54, 71]. Further studies demonstrated that FKBP12–rapamycin forms a physical complex with the yeast Tor1 and Tor2 proteins [22, 76, 118]. Later, work from five different groups converged to identify the mammalian Tor homolog (mTor) via its ability to bind the FKBP12–rapamycin complex [15, 27, 105, 106]. In addition, it was demonstrated that yeast Tor–mTor hybrid genes are capable of providing Tor function in yeast cells [2]. Tor homologs were later identified in other organisms; similar to S. cerevisiae two Tor proteins have been characterized in S. pombe, and a single Tor homolog has been identified in C. albicans, C. neoformans, D. melanogaster, A. thaliana, and H. sapiens [15, 32, 51, 71, 82, 92, 105, 131, 136].
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STRUCTURE OF THE TOR KINASES
The TOR proteins belong to a family of phosphatidylinositol kinase (PIK)-related kinases which include the mammalian phosphatidylinositol 3-kinase itself, ataxia telangiectasia mutated (ATM), ataxia telangiectasia related (ATR), DNA-dependent protein kinase, and the yeast MEC1, Rad53, and TEL1 proteins, all of which are involved in regulating cell cycle progression in response to exogenous or endogenous signals [62]. A hallmark of the PIK-related family is a C-terminal kinase domain with identity to both protein and lipid kinases. In fact, a specific protein kinase activity has been demonstrated for both the mammalian and the yeast Tor homologs [3, 16]. In addition to the kinase domain, the TOR proteins contain an FKBP12–rapamycin binding (FRB) domain and a toxic effector domain. Certain mutations in the FRB domain render Tor unable to bind FKBP12–rapamycin and, as a consequence, confer dominant rapamycin resistance [25, 76, 118]. Moreover, microinjection of the FRB domain into human cells arrests cell growth in the G1 phase of the cell cycle [126]. Although, rapamycin blocks most Tor functions in living cells, the Tor kinase activity is only partially inhibited by this drug in vitro, suggesting that the FRB domain plays an important role in intact cells [3, 16]. Interestingly, it has been suggested that phosphatidic acid interacts with the FRB domain, blocking FRB and activating Tor [41]. However these studies await further confirmation or a demonstration that they are conserved in other organisms. Extensive gene deletion analysis revealed that both the integrity of the Tor protein as well as the kinase activity are required to provide Tor function in yeast cells. Importantly, these studies led to the characterization of a 500 amino acid toxic domain in the central portion of the Tor protein that inhibits cell growth when overexpressed [3]. BLAST searches with the yeast Tor toxic domain revealed limited identity over a 240 amino acid region with the PIK family members Atr, Rad3, Mei-41, and Atm [3]. In accord with these results, recent sequence alignment analysis with multiple PIK family members identified a shared domain named FAT (for Frap, Atm, Trrap) that completely overlaps with the Tor toxic domain [14]. The dominant negative effects of the FRB and toxic effector domains suggest that these domains interact with important upstream regulators or downstream effectors of the Tor signaling cascade; however, their exact in vivo functions remain to be elucidated. The N-terminal region of the Tor proteins, amino acids 71–1147 in Tor1, contains multiple HEAT motifs and mediates association of Tor with downstream targets and with cellular membranes [4, 11, 72]. Interestingly, in mTor deletion of a C-terminal 30 amino acid region, which is absent in the yeast Tor proteins, enhances kinase activity and signaling
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[115]. This region includes Ser-2448, which was shown to be a target for phosphorylation by protein kinase B, raising the possibility that in mammalian cells Tor is subject to signaling by the PI3K/AKT pathway.
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INITIATION OF TRANSLATION IS GOVERNED BY TOR SIGNALING
The Tor pathway plays a central role in promoting translation in both yeast and mammalian cells (Figure 2) [6, 9]. In mammalian cells, the Tor kinase has been implicated in phosphorylating two proteins that function in translational control: PHAS-I
Figure 2. TOR controls translation in eukaryotes. In mammals and drosophila, mitogens and nutrients inactivate Tsc1/Tsc2 to favor the GTP-bound form of Rheb. Rheb activates TOR which in turn favors translation initiation by activating p70 S6 kinase as well as inactivating the translational repressor PHAS-I. In yeast cells, TOR responds to nutrients and acts on Eap1, a homolog of PHAS-I. Inactivation of TOR by nutrient limitation or rapamycin treatment activates the Gcn2 kinase, resulting in eIF2α phosphorylation and inhibition of overall translation. At this point it is not known if the yeast homolog of Rheb functions in the TOR pathway or if the TOR–Gcn2 cross talk exists in mammals and drosophila.
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and p70-S6 kinase [16, 19, 73, 98]. The PHAS-I protein forms a physical complex with eIF4E, thereby blocking the ability of eIF4E to mediate recognition of the 5′ CAP on mRNA and initiate recruitment of ribosomes to the mRNA. Thus, PHAS-I is an inhibitor of translation initiation, and the Tor pathway antagonizes the function of PHAS-I during culture conditions that favor cell growth and division (reviewed in [44]). When nutrients are limited, or rapamycin is present, the Tor signaling pathway is inactive or less active, and PHAS-I forms a stable complex with eIF4E to prevent CAP recognition and translation of the message. A central question in the field has been whether or not the Tor kinase directly phosphorylates PHAS-I or p70-S6 kinase in vivo [17]. This issue has been in part resolved by the discovery of Tor interacting proteins that facilitate interaction with and modification of these key substrate molecules (see below). In yeast cells, rapamycin also leads to an inhibition of translation initiation. Early studies implicated a different molecular target in yeast cells as the translation initiation factor eIF4G, which physically interacts with the 5′ CAP-binding factor eIF4E and assists in the recruitment of additional factors to the initiation complex. Rapamycin leads to degradation of eIF4G in yeast cells and a profound block to translational initiation [10]. Two possible distant homologs of the mammalian PHAS-I regulatory protein have been identified in yeast and are called p20 and Eap1. Both proteins physically interact with yeast eIF4E and thereby inhibit CAP dependent translation initiation in a fashion analogous to the PHAS-I protein in mammalian cells. While neither has been implicated as a direct substrate or regulatory target of Tor in yeast cells, deletion of the EAP1 gene does confer a modest level of resistance to rapamycin, indicating that liberating eIF4E from this negative regulatory factor can enhance cell survival in the presence of rapamycin [31]. It is thus quite remarkable that the role of the Tor signaling pathway in translational control has been in large part conserved from yeast to humans, even given that the direct molecular targets may be distinct in these divergent organisms. Recent studies have revealed a new and intriguing role for the Tor pathway in regulating translation during amino acid limitation in yeast cells. Amino acid limitation is normally sensed by the general control response, in which the Gcn2 protein senses uncharged tRNAs and controls translation of the mRNA encoding the global transcription factor Gcn4 [55]. The Gcn2 protein has two functional domains; a protein kinase and a domain that shares identity with tRNA synthetases. Amino acid limitation leads to an excess of uncharged tRNA molecules in the cell, which is sensed by the Gcn2 kinase, leading to its dephosphorylation and activation. Gcn2 in turn phosphorylates eIF2α, altering its activity in the cell to limit translation of the vast majority of mRNA molecules but leads to enhanced translation of
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the Gcn4 mRNA. The mRNA encoding Gcn4 is unique in that it contains four upstream open reading frames, and during normal nutrient conditions, ribosomes initiate translation at these upstream ORFs and rarely reach the start codon for Gcn4 itself. In cells limited for amino acids, the action of Gcn2 on eIF2α slows overall translation initiation allowing an increased scanning rate and initiation to produce Gcn4. In turn, Gcn4 then activates the expression of genes involved in amino acid biosynthesis, and a plethora of other genes. The first hints that the Tor pathway might impinge on the general control response were studies demonstrating that rapamycin can stimulate expression of a GCN4-lacZ reporter gene [122]. However, given previous studies that implicate the Tor pathway in the control of amino acid transporter stability in yeast, an alternative explanation was that rapamycin was inducing the general control response via an indirect effect on amino acid uptake. More recently, two studies have converged to implicate the Tor pathway in a more direct action on the general control response [26, 70]. The Gcn2 kinase is regulated by both uncharged tRNAs, which stimulate its activity, and by a kinase that phosphorylates and inactivates Gcn2 in rich medium. Inhibition of Tor by rapamycin leads to Ser 577 dephosphorylation in Gcn2, activation of Gcn2, and increased eIF2α phosphorylation. The net effect of rapamycin is then an increased level of Gcn4 expression and activation of the general control response. The power of rapamycin to activate Gcn2 also requires the ability of Gcn2 to bind uncharged tRNAs, and thus provides an additional regulatory input to Gcn2 that allows the Tor signaling cascade and limiting amino acids to be coordinately detected. The physiological significance of this expanded regulatory network is at present not clear, but it may allow enhanced induction of the general control response when cells are more severely starved than simple amino acid limitation affords. Alternatively, under conditions of amino acid starvation Tor also regulates the general control response [87], especially since Tor responds to amino acids in mammalian cells. These mechanisms allow coordination between two global regulatory networks, the Tor and Gcn pathways, which together function as both translational and transcriptional regulatory cascades. Whether or not the cross talk between these signaling pathways occurs in mammalian cells is not yet known. It is intriguing however that the mammalian Gcn2 protein has been identified as a key sensor to many signaling inputs and controls the integrated stress response. Ron and coworkers have proposed that mammalian cells utilize this ancient pathway to integrate diverse stress signals to regulate gene expression of the targets of the transcription factor ATF4, which is regulated in a manner analogous to the yeast Gcn4 factor [48].
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TOR REGULATES A COMPLEX TRANSCRIPTIONAL PROGRAM
The recent development of genome wide arrays and the high specificity of rapamycin to inhibit Tor function have aided the unraveling of a highly complex transcriptional program regulated by the TOR pathway in yeast cells (Figure 3). Earlier work had established a role for TOR signaling in the synthesis of rRNA and tRNA [80, 135]. Although the targets of this regulation are currently unknown, mutations that affect protein phosphatase 2A (PP2A) function result in alteration of both rRNA and tRNA gene expression [119, 124]. Thus, an intriguing possibility is that regulation of these genes by the TOR pathway is accomplished via PP2A. Studies employing genome wide arrays revealed that ribosomal protein (RP) genes expressed by PolII are repressed by the addition of rapamycin in a manner that closely resembles nutritional limitation [21, 50, 68, 97]. The
Figure 3. TOR controls transcription of genes involved in growth and adaptation to starvation and other stresses. In response to nutrients TOR activates the transcription of genes required for ribosome biogenesis. At the same time TOR negatively regulates genes required for adaptation to stress and nutrient scavenging. A common strategy for the repression of this latter class of genes is through nuclear exclusion of the transcription factors. TOR inactivates protein phosphatase 2A, which is required for nuclear localization of Msn2, Msn4, and Gln3 as well as translation of the message for the transactivator Gcn4. TOR also excludes the bZIP transcription factors Rtg1 and Rtg3.
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regulation of the RP genes by Tor signaling appears to be achieved via an still unknown effector as two independent studies have demonstrated that PP2A have little effect on regulating RP gene expression [40]. Most RP gene promoters feature binding sites for the activator/repressor protein, Rap1, and also for Abf1. Other transactivators, including Reb1, Gcr1, and Gcr2 are proposed to cooperate with Rap1 and Abf1 in the regulation of these genes (reviewed in [129]). In addition, the forkhead like transcription factor Fhl1 and its activator Ifh1 as well as Yap5 have recently been shown to bind to the majority of RP gene promoters and are likely to contribute to RP gene expression [74]. Earlier work linked Rap1 mediated regulation of RP genes to the cAMP and cell integrity pathways, however the precise events in this regulation have not been elucidated [66, 89]. Recent studies have identified components of the chromatin remodeling machinery that, presumably in connection with Rap1 and Abf1, contribute to RP gene expression in response to stress and nutrient signals and offer an opportunity to fine tune expression of these genes by signal transduction cascades [100]. The histone acetylase Esa1 has been recently shown to be recruited to RP gene promoters under conditions of active transcription [100]. We demonstrated that rapamycin, like nutrient limitation, results in loss of Esa1 from the RP gene promoters, rapid histone deacetylation by the Rpd3–Sin3 complex, and decreased transcription [102]. In addition, our studies reveal that rapamycin does not alter the occupancy of the Rap1 and Abf1 factors at RP gene promoters [102] (see Figure 4). Evidently, the elucidation of the detailed mechanisms by which Tor acts to promote occupancy of Esa1 and associated proteins to the RP genes will require extensive analysis. Further dissection of the regulatory events governing the RP genes is a challenging problem and undoubtedly will unravel the molecular mechanism by which nutrient and stress signals sensed by three signaling pathways converge at RP promoters to regulate gene expression. Recently, the Tor pathway was shown to control the expression of the nitrogen catabolite repressed (NCR) genes, underscoring the central role of Tor in nitrogen sensing [7, 11, 21, 50, 68]. The NCR genes are repressed by the availability of preferred nitrogen sources, such as glutamine or ammonia, and derepressed in the presence of limiting or poor nitrogen sources, such as proline or urea (for review see [79]). Many NCR genes are regulated by a set of GATA transcription factors, including the transactivators Gln3 and Nil1, and by their inhibitor Ure2 [30, 79]. Under rich growth conditions both Gln3 and Ure2 are phosphoproteins and upon shift to poor nitrogen sources or treatment with rapamycin, these factors are rapidly dephosphorylated [7, 21, 50]. More recent evidence suggests that Tor1, Gln3, and Ure2 form a complex. Accordingly, it was shown that Gln3 binds to the HEAT repeats on
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Figure 4. TOR signaling links nutrient sensing to histone acetylation to regulate gene expression.
Tor1, and that Ure2 is recruited to the complex indirectly by its interaction with Gln3 [11]. In the same study evidence was presented that Tor1 is capable of directly phosphorylating Gln3 in vitro [11]. However, these studies await independent confirmation. In the phosphorylated state, Ure2 and Gln3 form a complex and dephosphorylation of this complex by Tor inhibition promotes complex dissociation and nuclear translocation of Gln3 [7, 11]. Indirect evidence indicates that the rapid dephosphorylation of Ure2 and Gln3 is regulated by Tor via protein phosphatase type 2A. Thus, rapamycin treatment or entry into stationary growth phase induces the dissociation of the PP2A catalytic subunit Sit4 from its regulatory subunit Tap42 [36]. Although earlier studies suggested that Tap42 ejected an inhibitory role on Sit4 for the expression of the NCR genes, two independent studies have concluded that both of these factors are required for expression of these genes [7, 22, 41]. Finally it was shown that nuclear import of Gln3 requires the importin Srp1 while Crm1/Xpo is necessary for the nuclear export [23].
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A prominent set of genes that responds to nitrogen catabolite repression encodes enzymes that operate the glyoxalate and TCA cycles. Expression of these genes is directed by the Rtg1–Rtg3 heterodimeric transcription factor and their regulators Rtg2 and Mks1. When yeast cells are grown in poor nitrogen sources, such as urea, these cycles are upregulated to afford production of α-ketoglutarate that in turn is utilized in the de novo synthesis of glutamate and glutamine. The TOR signaling pathway controls the activity of the Rtg proteins in a manner strikingly similar to that of the Gln3 protein [68]. It has been shown that Rtg2 and Mks1 form a complex in the cytosol which negatively regulates the activity of the Rtg1–Rtg3 dimer [37, 113, 114]. Tor inhibition by glutamate or glutamine depletion, or by rapamycin, results in dephosphorylation of Mks1 and Rtg3 and nuclear translocation of Rtg1 and Rtg3 [37, 113]. The importin Msn5 was shown to be required for the export of Rtg1 and Rtg3 as msn5 mutations result in permanent nuclear localization of these factors. Interestingly, an msn5 mutation does not cause activation of the target genes for Rtg1 and Rtg3; instead, addition of rapamycin is still required for Rtg directed gene expression. These studies illustrate that TOR signaling functions at two levels to regulate Rtg directed transcription: first, to trigger the nuclear localization of the transactivators, and second at a later step to achieve proper gene expression. A growing body of data supports the notion that the control of nuclear localization may be a general mechanism by which the TOR kinases regulate transcription. Tor signaling also influences the mechanisms by which the transcription factors Msn2 and Msn4 are sequestered in the cytoplasm through interaction with the negative regulators Bmh1 and Bmh2 [7]. Treatment with rapamycin results in expression of the STRE genes by 2 h postexposure, although in this case the activation of the target genes is not precisely correlated with the rapid nuclear localization of Msn2 and Msn4 that was reported by Beck and Hall, suggesting additional levels of control for these genes (as is the case with Rtg controlled genes) [21, 50]. In a more recent study, rapamycin was found to have no effect on nuclear import of Msn2, but instead delayed nuclear export [40]. In addition, loss of Tap42 function in temperature sensitive mutant strains shifted to the nonpermissive temperature led to an induction of the stress response genes, implicating a role for Tap42 in the pathway [40]. This finding is again in contrast to an earlier report based on studies with the tap42-11 mutant allele that were interpreted as excluding a role for Tap42 in the Tor pathway leading to control of the stress response genes [8]. Taken together, the current view is that the Tor–Tap42–Sit4 pathway controls the nuclear export of the Msn2 and Msn4 transcription factors that regulate expression of the stress response genes. These results may help explain the marked delay in induction of the
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STRE genes by rapamycin. Taken together these studies reveal that the Tor pathway controls both nuclear import and nuclear export of key regulatory transcription factors. The Tor pathway has also been shown to control gene silencing at subtelomeric regions. These studies capitalized on the Saccharomyces toolbox-ordered sets of gene deletions. Zheng and coworkers screened ~4000 viable gene deletions for altered sensitivity to rapamycin [24]. One of these mutants, sir3, displayed an increased resistance to rapamycin. Studies of this phenomenon revealed that Tor negatively regulates MPK1, the MAP kinase of the cell integrity pathway. MPK1 in turn phosphorylates Sir3 to result in increased expression of subtelomeric genes involved in the repair of cell wall damage [1]. Once again the powerful tools available in yeast were able to quickly identify an important mode of cross talk between two different signal transduction pathways. Finally, observations in mammalian cells suggest that mTOR and PP2A control the activity of the STAT3 protein in a fashion similar to the control of Gln3 activity in yeast. STAT3 is retained in the cytoplasm until activation by the JAK kinases that act by phosphorylating tyrosine residues in response to cytokines. Phosphorylation of serine and threonine residues (most notably Ser727) also occurs but it is not yet clear if phosphorylation positively or negatively regulates the ability of STAT3 to activate transcription. Inhibition of PP2A activity increases STAT3 phosphorylation on serine and threonine residues and results in a concominant exclusion of STAT3 from the nucleus [133]. Reportedly STAT3 directed transcription is blocked by rapamycin treatment and in vitro phosphorylation studies indicate that mTOR may phosphorylate this protein directly [134]. While these studies convincingly show the involvement of mTOR and PP2A in regulating the Ser727 phosphorylation status of STAT3, the physiological consequences of this phosphorylation remain unknown. Importantly, constitutive activation of STAT3 results in malignant transformation and cancer.
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TOR SIGNALS VIA A CONSERVED PROTEIN PHOSPHATASE COMPLEX
The Tor signaling cascade functions to control the activity of the PP2A-related catalytic subunits Sit4, Pph21, and Pph22 and their associated regulatory factor Tap42. Studies by Arndt and colleagues first demonstrated that the Sit4 catalytic subunit and related proteins physically interact with the essential Tap42 protein [36, 77]. Most interestingly, the Sit4–Tap42 complex disassociates when cells are limited for nutrients or exposed to rapamycin, and certain TAP42 mutant alleles, such as tap42-11, confer
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partial resistance to rapamycin in yeast cells. Subsequent studies have demonstrated that Sit4 plays a key role in controlling the Ure2–Gln3 regulatory complex that governs expression of the NCR genes [7], and also more recently in the stress response pathway governed by Msn2/4 [40]. Although Tap42 was originally thought to only function as a negative factor, studies implicate Tap42 as playing both positive and negative signaling roles downstream of the Tor kinase. Earlier studies demonstrated that shift of the tap42-11 conditional allele to the nonpermissive temperature arrested cell growth but did not lead to an induction of the NCR gene response, indicating that Tap42 must play a role in addition to simply inhibiting Sit4 in the signaling cascade [21]. More recently, this finding was confirmed employing additional temperature sensitive Tap42 conditional mutants [40]. Tap42 may play a role in targeting Sit4 to different locations in the cell, or to different substrates. One report has suggested that Tap42 might be a direct substrate of the Tor kinases [60], but this finding has not yet been independently confirmed. An additional layer of regulation on the Tap42–Sit4 complex was introduced by the identification of a Tap42 interacting protein, named Tip41 [59]. Tor signaling promotes the phosphorylation of Tip41 and stabilization of the Tap42–Sit4 complex. Rapamycin treatment results in Tip41 dephosphorylation, mediated at least in part by Sit4, which favors Tip41 association with Tap42 and presumably leads to the release of an activated form of Sit4. In mammalian cells, the Tap42 homolog is the α4 protein, which forms physical complexes with protein phosphatase catalytic subunits related to the yeast Sit4 protein. The role of the Tor pathway in controlling PP2A activity in mammalian cells is at present controversial. Moreover, it is not clear that the stability of the PP2A–α4 complex is regulated by Tor signaling in the same fashion as the yeast Sit4–Tap42 complex and some findings consistent with a direct effect of mTor on PP2A itself have been reported [85, 86, 95]. A tempting model then is that the Tor proteins which contain a Cterminal protein kinase domain signals in conjunction with the associated phosphatases Sit4–Tap42, Pph21–Tap42, and Pph22–Tap42. This regulatory network may therefore function to more rapidly and dynamically control the phosphorylation state of target substrates than either the kinase or the phosphatase alone could achieve. By coupling the activity of the two, the Tor cascade could rapidly mediate the dephosphorylation of proteins that are also substrates of the Tor kinase activity. This would lead to very rapid alterations in cell physiology in response to changing nutrient conditions, much more rapid than could be achieved by inhibiting only the activity of the kinase alone since protein turnover and new synthesis would then be required to lead to the production of dephosphorylated target molecules.
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REGULATION OF PROTEIN DEGRADATION AND AUTOPHAGY
Autophagy is a response to nitrogen, carbon, or sulfur starvation in which indiscriminate portions of the cytosol are engulfed by a membrane to form autophagic bodies. These autophagic bodies then fuse with lysosomes or, in yeast, with the vacuole, and proteins are degraded, resulting in the release of free amino acids which are in turn recycled by the cell. Tor inactivation by rapamycin treatment induces autophagy and the evidence indicates that Tor regulation is exerted at the transcriptional as well as a posttranscriptional level [90]. The pathway that mediates autophagy is operated by the APG genes [42, 90]. Among the genes activated by Tor inhibition and regulated via activation of Gln3 are the APG genes: APG1, AGP3, AGP5, APG7, APG8, APG12, and APG13 (reviewed in [99]). In addition, Tor regulates the interaction of Apg13 with Apg1 to result in activation of the Apg1 protein kinase required for autophagy [61]. Thus, nutrient starvation or rapamycin treatment results in dephosphorylation of Apg13, leading to Apg13–Apg1 complex formation, Apg1 activation, and autophagy [61]. In addition to the Tor pathway two other nutrient signaling pathways operated by the Snf1 and Pho85 kinases have recently been shown to regulate autophagy [128]. It will be of considerable interest to establish if these regulatory cascades converge at the Apg1 activation step.
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TOR REGULATES POLARIZATION OF THE ACTIN CYTOSKELETON
In yeast, Tor1 and Tor2 have overlapping rapamycin-sensitive functions in the control of cell growth, translation, and transcription. In addition, Tor2 has a unique and essential rapamycin-insensitive function involving polarization of the actin cytoskeleton [110, 111, 138]. Establishment and regulation of the actin cytoskeleton is controlled in part by small GTPbinding proteins encoded by the RHO1, RHO2, and CDC42 genes which are highly conserved throughout evolution. Activity of these proteins is governed by a switch between the GDP-bound inactive, and GTP-bound active states. Genetic analysis of the Tor2-unique function has led to a model whereby Tor2 controls this switch. Lethal tor2 mutations can be suppressed by mutations in the SAC7 gene, which encodes a GTPase activating protein (GAP) for Rho1 and Rho2, by overexpression of Rho1 or Rho2 themselves, or by Rom2, a GTP exchange factor (GEF) that promotes the GTP (active) state of the Rho proteins [110]. Furthermore, alleles of TOR2 that are defective for the Tor2-unique function have been
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characterized and were found to cause a G2/M cell cycle arrest consisting of cells with unpolarized actin [53]. Importantly, Rho1 is known to activate the PKC signaling pathway in yeast and overexpression of Pkc1 rescues alleles of tor2 defective in the Tor2-unique function. In mammalian cells, mTOR regulates the activity of PKC but it is as yet unclear how similar this connection will be to that of yeast. A link between the actin cytoskeleton and mTOR has not yet been established in mammalian cells and PKC activation by mTOR is rapamycin-sensitive in mammals, whereas the Tor2unique function is rapamycin-insensitive in yeast [138]. Recently, careful studies have revealed a rapamycin-sensitive and cell cycle dependent polarization of the yeast actin cytoskeleton [127]. These studies suggest that Tor1 may have a greater role in actin polarization than previously appreciated.
8
TOR REGULATES MICROTUBULE FUNCTION
A second function for the Tor kinases at the G2/M phase of the cell cycle is beginning to emerge. Recent reports suggest that the Tor kinases play a role in the assembly or function of microtubules [12, 29]. This appears to be a rapamycin-sensitive function that is shared between yeast Tor1 and Tor2 and that is conserved from yeast to mammals [12]. Genetic and biochemical evidence in yeast suggests that this process involves the kinesin-related proteins (KRPs), which are important for spindle–pole separation at anaphase [29]. Consistent with this, rapamycin treatment causes defects in chromosome and nuclear segregation in both yeast and mammalian cells [12, 29].
9
TOR INTERACTING PROTEINS AND NUTRIENT SENSING
The downstream elements of the Tor signaling pathway likely involve the recently identified Tor partner proteins, Raptor/Kog1, Avo1/2/3, and Lst8/GβL [47, 63, 64, 75, 130]. Kog1/Raptor is conserved from yeast to humans and plays an essential function in yeast as part of a complex of proteins (called the Torc1 complex) involving the Tor1 [130] or Tor2 kinases [75] and Lst8. Interestingly, Lst8 contains 7 WD-40 repeats and is also conserved in mammals, where it is known as G • L. Some evidence suggests that nutrient limitation promotes the formation of the mTor–raptor complex in mammalian cells resulting on mTOR inhibition. Furthermore, it has been proposed that raptor facilitates Tor association with substrates
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[63]; Hara, 2002 #3227]. In particular, the mammalian raptor protein can bind to the Tor substrates p70 S6 kinase and PHAS-I via a previously defined domain known as the TOR signaling motif (TOS) [91, 108, 109]. The TOS domain allows PHAS-I to dock onto the mTor/Raptor protein complex, and PHAS-I is then phosphorylated to promote release and activation of eIF4E and cell growth. In yeast cells, a second complex, called the Torc2 complex, can be detected containing Tor2, Lst8, and the Avo1, Avo2, and Avo3 proteins [75, 130]. FKBP12–rapamycin can physically associate with the Torc1 complex but not with the Torc2 complex, suggesting that this second complex mediates the rapamycin-insensitive role of Tor2 in controlling polarization of the actin cytoskeleton. If and how any of these newly discovered Tor associated proteins communicate nutrient sensing via Tor to PP 2A or any of its associated subunits is as yet unknown.
10
TOR, THE TUBEROUS SCLEROSIS PROTEINS, RHEB, AND NUTRIENT SENSING
A new regulatory cascade that is conserved from budding and fission yeast to humans involving the tuberous sclerosis gene products and the small G protein Rheb has emerged that may link activation of the Tor pathway to nutrients. The mammalian TSC1 and TSC2 genes encode the proteins hamartin and tuberin, which physically interact to form a protein complex that functions as a tumor suppressor [69]. Mutations in the TSC1 or TSC2 genes are associated with both renal cell carcinomas and the formation of benign hamartomas. Studies conducted concurrently in drosophila and mammalian cells first linked the functions of both the mammalian and the fly Tsc1/2 homologs to the activation of the p70 S6 kinase cascade that is controlled by Tor [43, 58]. The Tsc1 and Tsc2 proteins were found to negatively regulate Tor signaling, and to form a physical complex with the drosophila Tor homolog known as dTOR. Mutants lacking either Tsc1/2 protein exhibited a Tor-dependent increase in p70 S6 kinase activity, and the Tsc1/2 proteins also control the activity of the Tor regulated protein PHAS-I. A series of recent studies converged to identify the small G protein Rheb as the target of the Tsc2 GTPase activating protein (GAP) function. These findings suggests that Rheb is an intermediary in the control of Tor activity by the Tsc1/2 protein complex [57, 107, 120, 137]. Earlier studies in budding and fission yeast and pathogenic fungi have implicated the Rheb signaling in nitrogen sensing and the control of amino acid transport. Budding yeast mutants lacking Rheb were found to be viable but hypersensitive to amino acid analogs, such as canavanine, and also exhibited an increase in amino acid uptake [121]. Fission yeast mutants
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lacking the Rheb homolog Rhb1 were also viable but exhibited defects in nutrient sensing and arrested in G1 more rapidly than wild-type cells in response to nitrogen limitation. Furthermore, in the rhb1 mutant cells several nitrogen starvation induced genes were found to be expressed under rich nitrogen conditions [78]. Homologs of the mammalian Tsc1 and Tsc2 proteins in fission yeast are known to regulate the proper targeting and function of amino acid permeases, a role consistent with related studies on the Rheb homolog [81]. The Aspergillus fumigatus Rheb homolog, rhbA, is induced in response to nitrogen starvation. The rhbA mutant cells are viable but their growth was compromised in poor nitrogen sources, nitrogen rich conditions failed to repress their asexual development, and they were hypersensitive to rapamycin [93, 94]. Taken together, these findings are consistent with a role for the small G protein Rheb in nutrient sensing in budding yeast, fission yeast, and pathogenic fungi and suggest Rheb might play a globally conserved role in activating the Tor kinases in response to certain nutritional cues. Rheb is not essential for growth in any of these fungi, in contrast to Tor, and thus Rheb likely represents one of several redundant regulatory inputs. The key questions that remain are how the Tor kinases sense nutrients and how they communicate this information to their downstream effectors. Two recent studies suggest that the mTOR kinase is activated in response to two small molecules: ATP and PA [35, 41]. Phosphatidic acid is produced by phospholipase D and has been reported to bind mTOR and thereby stimulate its kinase activity, in vitro and in vivo [41]. Phospholipase D is conserved in yeasts, where it plays a known role in promoting morphological changes to pheromone and in meiosis [46, 103]. As yet, no role has been adduced for phospholipase D in controlling Tor signaling in yeasts. Recent studies reveal that Pld1 is required for filamentous growth in Candida albicans [56], and our studies have shown that the Tor pathway is necessary for filamentous growth in S. cerevisiae, C. albicans, and C. neoformans [34]. Finally, Thomas and colleagues have suggested that the low intrinsic Km of mTOR for ATP poises the enzyme to serve as a general ATP sensor for the cell [35]. Thus PA and ATP may contribute to the activation of mTOR via or in addition to nutrient sensing mechanisms.
11
SUMMARY REMARKS AND PERSPECTIVES
From its original discovery in 1975 as an antifungal natural product with an undesired immunosuppressive effect, rapamycin emerged as a powerful immunosuppressive drug that received FDA approval in 1999. Rapamycin also inhibits growth and proliferation of a variety of other cells types,
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including smooth muscle cells and several neoplastic cell lines. The recent finding that cardiac stents impregnated with rapamycin can significantly reduce restenosis, and that rapamycin can inhibit growth of a variety of different tumors, will lead to new clinical indications for this novel natural product. In light of these significant clinical advances, a more detailed understanding of the targets of rapamycin and their function in growth promoting pathways should provide considerable insight into the molecular basis of therapeutic action and support the development of additional and novel targets for therapeutic intervention. Of particular interest and importance is the clinical utility of rapamycin as a novel chemotherapy agent. A flurry of recent reports have identified unique tumor cell lines that are exquisitely sensitive to growth inhibition by the mTOR inhibitor rapamycin and its analogs. Particularly, in cells and mice lacking the tumour suppressor PTEN, a lipid phosphatase that controls signaling via the PI3 kinase pathway, rapamycin inhibits p70 S6 kinase activity and oncogenesis [28, 45, 83, 88, 96, 116]. These findings apply to a broad range of tumor types, including tumors of the immune system (multiple myeloma), common solid organ tumors (breast, prostate), and less common but devastating solid organ tumors (glioblastoma). There are a variety of phase II and phase III trials that are ongoing now, and the years ahead hold excitement and promise of a significant impact for rapamycin in a multitude of diverse clinical applications. Much of our current understanding of rapamycin targets and mechanism of action stems directly from genetic and molecular studies conducted in the model yeast S. cerevisiae. In summary, these contributions include: 1. 2.
Identification of the rapamycin binding protein FKBP12 Demonstration that FKBP12 is not essential for growth but is required for rapamycin action 3. Identification of Tor1 and Tor2 as the targets of the FKBP12–rapamycin complex 4. Definition of functional domains of the Tor kinases 5. Elucidation of the role of the Tor pathway in translation 6. Finding that Tor controls PP 2A 7. Illumination of the Tor pathway role in nutrient sensing 8. Discovery that the Tor pathway controls transcriptional programs for cell growth 9. Application of a “chemical genomics” approach to identify novel Tor functions 10. Identification of novel Tor binding proteins conserved from yeast to humans (Kog1/raptor and Lst8/GβL)
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These findings highlight the vast potential of simple, genetically tractable model systems to inform our understanding of drug targets and mechanisms of action in more complex systems such as mammalian cells. These advances also highlight the power of yeast genetics to define the direct targets of rapamycin as the conserved FKBP12 and Tor proteins. The challenges that lie ahead are: (1) to define the upstream regulatory pathways that allow Tor to sense nutrients, (2) to elucidate the detailed mechanisms by which Tor controls the translational and transcriptional machineries to effect developmental and transcriptional programs required to respond to altered growth conditions to promote cell growth and proliferation; and (3) to bring these molecular insights to bear on clinical applications that will improve health and prolong life.
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79. Magasanik, B., and C. A. Kaiser. 2002. Nitrogen regulation in Saccharomyces cerevisiae. Gene 290:1–18. 80. Mahajan, P. B. 1994. Modulation of transcription of rRNA genes by rapamycin. Int. J. Immunopharmacol. 16:711–721. 81. Matsumoto, S., A. Bandyopadhyay, D. J. Kwiatkowski, U. Maitra, and T. Matsumoto. 2002. Role of the Tsc1-Tsc2 complex in signaling and transport across the cell membrane in the fission yeast Schizosaccharomyces pombe. Genetics 161:1053– 1063. 82. Menand, B., T. Desnos, L. Nussaume, F. Berger, D. Bouchez, C. Meyer, and C. Robaglia. 2002. Expression and disruption of the Arabidopsis TOR (target of rapamycin) gene. Proc. Natl. Acad. Sci. USA 99:6422–6427. 83. Mita, M. M., A. Mita, and E. K. Rowinsky. 2003. Mammalian target of rapamycin: a new molecular target for breast cancer. Clin. Breast Cancer 4:126–137. 84. Morice, M. C., P. W. Serruys, J. E. Sousa, J. Fajadet, E. Ban Hayashi, M. Perin, A. Colombo, G. Schuler, P. Barragan, G. Guagliumi, F. Molnar, and R. Falotico. 2002. A randomized comparison of a sirolimus-eluting stent with a standard stent for coronary revascularization. N. Engl. J. Med. 346:1773–1780. 85. Murata, K., J. Wu, and D. L. Brautigan. 1997. B cell receptor-associated protein alpha4 displays rapamycin-sensitive binding directly to the catalytic subunit of protein phosphatase 2A. Proc. Natl. Acad. Sci. USA 94:10624–10629. 86. Nanahoshi, M., T. Nishiuma, Y. Tsujishita, K. Hara, S. Inui, N. Sakaguchi, and K. Yonezawa. 1998. Regulation of protein phosphatase 2A catalytic activity by alpha4 protein and its yeast homolog Tap42. Biochem. Biophys. Res. Commun. 251:520–526. 87. Natarajan, K., M. R. Meyer, B. M. Jackson, D. Slade, C. Roberts, A. G. Hinnebusch, and M. J. Marton. 2001. Transcriptional profiling shows that Gcn4p is a master regulator of gene expression during amino acid starvation in yeast. Mol. Cell Biol. 21:4347–4368. 88. Neshat, M. S., I. K. Mellinghoff, C. Tran, B. Stiles, G. Thomas, R. Petersen, P. Frost, J. J. Gibbons, H. Wu, and C. L. Sawyers. 2001. Enhanced sensitivity of PTEN-deficient tumors to inhibition of FRAP/mTOR. Proc. Natl. Acad. Sci. USA 98:10314–10319. 89. Neuman-Silberberg, F. S., S. Bhattacharya, and J. R. Broach. 1995. Nutrient availability and the RAS/cyclic AMP pathway both induce expression of ribosomal protein genes in Saccharomyces cerevisiae but by different mechanisms. Mol. Cell Biol. 15:3187–3196. 90. Noda, T., and Y. Ohsumi. 1998. Tor, a phosphatidylinositol kinase homologue, controls autophagy in yeast. J. Biol. Chem. 273:3963–3966. 91. Nojima, H., C. Tokunaga, S. Eguchi, N. Oshiro, S. Hidayat, K. Yoshino, K. Hara, N. Tanaka, J. Avruch, and K. Yonezawa. 2003. The mammalian target of rapamycin (mTOR) partner, raptor, binds the mTOR substrates p70 S6 kinase and 4E-BP1 through their TOR signaling (TOS) motif. J. Biol. Chem. 278:15461–15464. 92. Oldham, S., J. Montagne, T. Radimerski, G. Thomas, and E. Hafen. 2000. Genetic and biochemical characterization of dTOR, the Drosophila homolog of the target of rapamycin. Genes Dev. 14:2689–2694. 93. Panepinto, J. C., B. G. Oliver, T. W. Amlung, D. S. Askew, and J. C. Rhodes. 2002. Expression of the Aspergillus fumigatus rheb homologue, rhbA, is induced by nitrogen starvation. Fungal Genet. Biol. 36:207–214. 94. Panepinto, J. C., B. G. Oliver, J. R. Fortwendel, D. L. Smith, D. S. Askew, and J. C. Rhodes. 2003. Deletion of the Aspergillus fumigatus gene encoding the Ras-related
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Chapter 14 USE OF YEAST AS A MODEL SYSTEM FOR IDENTIFYING AND STUDYING ANTICANCER DRUGS
Jun O. Liu1 and Julian A. Simon2 1
Departments of Pharmacology and Neuroscience, Johns Hopkins School of Medicine, Baltimore, MD 21205; 2Divisions of Clinical Research, Basic Sciences and Human Biology, Fred Hutchinson Cancer Research Center, Seattle, WA 98109
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INTRODUCTION
As signified in part by the publication of the current book, yeast is becoming a more widely used system for studying small molecule–protein interactions in general and anticancer agents in particular. There are a number of reasons for this surging popularity. As a eukaryote, it contains many, if not most, of the essential genes involved in the control of cell cycle, cellular structure, metabolism, and stress responses that are conserved from yeast to humans, making it possible to use yeast to study the function of those genes. For this reason, what is learned in a yeast cell can often been extrapolated into a mammalian counterpart. Unlike mammalian cells, yeast is relatively easy to grow and manipulate, with its natural habitat being the harsh environment of the wild. Moreover, decades of research on both the biochemistry and genetics of the budding yeast has accumulated a wealth of information about yeast and made available a number of tools that allows for the manipulation of genes in yeast, including knockout and ectopic expression of both endogenous and exogenous genes. Several recent advances have greatly facilitated the use of yeast as a powerful system to study small molecules. They include the advent of the yeast two-hybrid system, the sequencing of the genome of the budding yeast Saccharomyces cerevisiae and the development of DNA chip and 375 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 375–391. © 2007 Springer.
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microarrays that allow for the monitoring of the expression of the complete yeast genome. In this chapter, we will cover three different ways yeast has been used to facilitate the identification, characterization, and mechanistic studies of anticancer drugs. These include (1) the use of yeast as a surrogate system to identify and study anticancer drugs; (2) the use of yeast as selfcontained microvessles to perform large-scale parallel analysis of interactions between proteins and small ligands; and (3) the use of the yeast transcriptional profiling for validating the molecular target for given drugs. For the most part, we have selected examples from our own laboratories. We thus have attempted to be more illustrative than comprehensive in this chapter.
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USE OF YEAST AS A SURROGATE SYSTEM TO IDENTIFY AND STUDY ANTICANCER DRUGS
Many of the essential genes, including those involved in the control of cell cycle, cell structure, transcription and basic metabolism, are conserved between yeast and humans. It is those essential genes involved in the regulation of cell proliferation that are often mutated in human cancer cells, leading to uncontrolled growth. For those genes, yeast represents an ideal surrogate system to identify and study anticancer drugs. In some instances, a conserved gene between yeast and human can be studied even if the fundamental physiological functions differ, as long as there is a suitable phenotypic readout upon inhibition of the gene function. We will use two examples to illustrate this point, one being a family of anti-angiogenic natural products, fumagillin and their common protein target, the type 2 methionine aminopeptidase (MetAP2) and the other being the identification of the NAD+-dependent histone deacetylase Sir2p in yeast involved in silencing, aging, and cell cycle control.
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Studies of the fumagillin family of anti-angiogenicinhibitors for the type 2 methionine aminopeptidase (MetAP2)
Fumagillin and ovalicin (Figure 1) are a family of structurally related natural products that were serendipitously discovered as potent angiogenesis inhibitors [7, 25]. A structural analog of fumagillin, known as TNP-470 or AGM-1470, has been undergoing testing for treatment of a variety of cancers. The molecular mechanism of inhibition of angiogenesis by this class of inhibitors was elucidated through the detection, isolation, and identification of the molecular target for these inhibitors. A 67 kDa protein was detected by photoaffinity labeling using photosensitive probes derived
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from fumagillin. The target protein was isolated using affinity chromatography and identified by mass spectrometry as the MetAP2 [16, 41].
Figure 1. The chemical structures of fumagillin, TNP-470, and ovalicin.
Methionine aminopeptidases represent a family of highly conserved proteins from bacterial to humans [26, 33]. Their primary function is to remove the N-terminal methionine from newly synthesized proteins for subsequent posttranslational modifications. In prokaryotes, there is only one copy of methionine aminopeptidase gene. Knockout of the gene has a lethal phenotype, indicating that methionine aminopeptidase is an essential gene in bacteria [4]. In eukaryotes, however, there are two different genes for methionine aminopeptidases, hence the names type 1 (MetAP1) and type 2 (MetAP2) [1, 28]. Despite their different N-terminal extension sequences, the catalytic domains of MetAP1 and MetAP2, including the five amino acid residues involved in the formation of a bimetallic active site, are highly conserved (Figure 2). Genetic analyses in yeast revealed that the two methionine aminopeptidase genes can be deleted individually without affecting viability, although they lead to a slow growth phenotype [5, 28]. Double knockout of the two MetAP genes, however, is lethal, suggesting that the two genes encoding methionine aminopeptidases are functionally redundant and together they are essential for yeast growth [28].
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Figure 2. Schematic comparison of the domain structures of eukaryotic MetAP1 and MetAP2. The five amino acid residues in the catalytic domain are shown with single letter code.
Since the MetAP2 was isolated from mouse embryo extract via affinity chromatography using the fumagillin or ovalicin as an affinity ligand, a major question was the specificity of fumagillin for MetAP2, given the extremely high sequence similarity between MetAP1 and MetAP2 [1, 28]. Using the wild type and map1 and map2 mutant strain of yeast, it was found that only the map1 mutant strain of yeast is sensitive to inhibition by fumagillin or ovalicin, providing strong evidence that the drugs are highly specific for MetAP2 (Figure 3) [16, 41]. In addition to demonstrating the extraordinary specificity of fumagillin for MetAP2, these studies also raised the interesting question of how inhibition of MetAP2 alone led to cell cycle inhibition of human endothelial cells. Although the map1 mutant has not been further exploited, it is apparent that the mutant can be used for screening of small molecule libraries to identify novel inhibitors of MetAP2.
Figure 3. Specific inhibition of MetAP2 by TNP-470 in yeast. The wild type, map1 (yeast MetAP1) and map2 (yeast MetAP2) mutant strains were plated on YEPD plates containing either dimethylsulfoxide as a control or 50 nM of either TNP-470 or ovalicin. The plates were incubated at 30ºC for 4 days before they were photographed.
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Identification and studies of Sir2p inhibitors in yeast
Sir2p (Sir stands for Silent Information Regulator) is a founding member of a family of NAD+-dependent protein deacetylases that are conserved from bacteria to humans [11]. Eukaryotic Sir2 homologs have been shown to be involved in the regulation of transcription, cell cycle control, repair of DNA damage, and aging [13, 18]. At the molecular level, Sir2p and its family members regulate protein–protein interactions through removal of acetyl groups from lysine residues. Regulation of chromatin structure is accomplished by deacetylation of histone proteins within targeted chromatin regions [18]. In yeast, Sir2p is known to be responsible for silencing transcription of the ribosomal DNA array (rDNA), the silent mating-type loci HMR and HML as well as regions near telomeres [24, 27, 42]. A human homolog of the yeast Sir2p, hSIR2, was recently shown to deacetylate the tumor suppressor gene p53, downregulating the transcriptional activity of p53 and repressing p53-dependent apoptosis of tumor cells in response to DNA damage and oxidative stress [34, 43]. Thus, inhibitors of the human Sir2p homologs may be useful in potentiating cytotoxic chemotherapy by enhancing the p53-depenent apoptosis and cell cycle arrest. That Sir2p in yeast is responsible for silencing the chromatin in proximity of telomeres was demonstrated by the repression of a reporter gene integrated into the telomere proximal region. It was shown that the uracil biosynthsesis gene URA3 is silenced when it is in close proximity of a telomere [15]. The deletion of the yeast SIR2 gene leads to the derepression of the transcription of the URA3 gene, as well as other genes in normally silent loci. To identify small molecule inhibitors of the yeast Sir2p as a first step toward identifying inhibitors for its human counterpart, two similar approaches were taken, both involving the exploitation of the URA3 gene as a reporter placed within a Sir2p-dependent silent locus [2, 17]. It was expected that inhibition of yeast Sir2p would lead to the derepression of the silenced URA3. The two groups used slightly different readouts, one exploiting the URA3 auxotroph [2] and the other utilizing the URA3dependent toxicity by a small molecule 5-fluoroorotic acid [17]. The advantage of the use of the yeast URA3 auxotroph is that it is a positive selection, eliminating potential false positive hits that may be toxic to yeast by mechanisms other than inhibition of Sir2p. Upon screening a library of 6,000 chemical compounds for those that enabled the growth of the yeast with a telomeric URA3 gene in the absence of uracil, 11 hits emerged. One out of the 11 hits was confirmed with a second reporter yeast strain with the TRP1 gene integrated at another silent locus growing in the absence of tryptophan. The hit was named splitomicin (Figure 4).
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Figure 4. Structure of splitomicin (A) and its effect on yeast responsiveness to α factor (B). The helo of cells indicates that splitomicin induced loss of sensitivity of yeast to α factorinduced silencing in the mating type loci.
The yeast system not only enabled the identification of hits, but also facilitated the subsequent verification of splitomicin as a relatively selective inhibitor for yeast Sir2p and identification of residues in Sir2p that are critical for its binding to splitomicin [2]. Yeast contains four SIR2 homologs, HST1-4 (HST stands for Homologue of Sir Two). To determine whether splitomicin is specific for Sir2p, the transcriptional profiles of wild- type yeast cells treated with splitomicin and the isogenic yeast strains with deletion of SIR2 and each of its four homologs were obtained and compared to each other. It was found that the majority of the changes (88%) in transcription as a consequence of splitomicin treatment are seen in the sir2 mutant yeast strain. Interestingly, a small number of changes in transcription (9%) were identical to those seen in the hst1 mutant strain. In contrast, there was no overlap in transcriptional profile changes between splitomicin-treated yeast cells and the hst2-4 mutant strains. These results clearly indicated that splitomicin is selective toward Sir2p and Hst1p, but does not interact with the other members of the Sir2p family of proteins. PCR-mediated mutagenesis of the SIR2 open reading frame was used to identify three drug-resistant mutants of Sir2p using similar reporter screens in yeast. The drug-resistant mutations all occurred in a region of the protein that is part of the putative active site for Sir2 and is highly conserved between yeast Sir2p and Hst1p, suggesting that splitomicin is likely to interact with the active sites of both Sir2p and Hst1p. In an independent study, a similar approach was used to screen for the yeast Sir2p inhibitors with the yeast strain containing a telomeric copy of URA3 gene [17]. Three small molecule inhibitors of Sir2p were identified, which also inhibited the human SIRT2 protein with similar potencies. It is noteworthy that the screens employed in the studies would not have been possible without the use of yeast. Even though the human SIRT2 is known, its celluar function is largely unknown and it is much more difficult
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to devise a similar reporter system that would allow for a phenotypic screen for the human SIRT2 inhibitors directly.
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YEAST CELLS AS MINIATURIZED BIOCHEMICAL REACTION VESSELS FOR ASSESSING DRUG–PROTEIN INTERACTIONS IN LARGE NUMBERS
The ability of yeast cells to take up one or more plasmids and express the specific proteins encoded by the plasmids makes it possible to use individual yeast cells as independent “containers” for analyzing the interaction between the proteins expressed in the yeast and the small molecules administered. Given the ease of culturing yeast cells and the ease with which exogenous plasmids can be introduced into yeast, large-scale parallel analyses are feasible. Thus, each yeast cell can be turned into a self-contained reaction vessel to assay the interaction between a ligand and its target. One of the major challenges is to develop a suitable readout that allows for quick and easy detection of the binding between a drug and its protein target in yeast cells. One type of reporter system couples a ligand-binding event with the transcription of a specific reporter gene introduced into the yeast. Another reporter system links the ligand–protein binding event with the reconstitution of an enzyme that can be easily detected. In this chapter, we have chosen a few systems that have the potential of being adapted for such an application or have been used to assess drug-protein interactions.
3.1
Yeast two-hybrid system and its variants
Since its development, the yeast two-hybrid system has become one of the most powerful tools for uncovering new protein–protein interactions [6, 8]. The yeast two-hybrid system exploits an intrinsic property of eukaryotic transcription factors to function with two separable and portable domains, a DNA-binding domain and a transcriptional activation domain. By splitting a transcription factor into two separate proteins and fusing into each two interacting proteins, it is possible to reconstitute the transcription activity. In theory, the yeast two-hybrid system is ideal for identifying inhibitors for protein–protein interactions. Unfortunately, due to the relatively large binding interface often associated with protein–protein interactions [32], it is in general difficult to discover small molecules that directly compete for a
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protein–protein interaction. Nevertheless, there do exist a few successful examples of small molecule antagonists for protein–protein interactions identified through the two-hybrid system [23, 44]. Some protein–protein interactions are mediated by defined protein modules and a short, contigous oligopeptide. These include the binding of SH2 and SH3 domains to phosphotyrosine-containing peptides or prolinerich peptides, respectively [37]. The detection of an SH3 domain and its target sequence can be easily established using the yeast two-hybrid system. The binding of SH2 domains to their target requires a phosphorylation step, which usually occurs in mammalian cells upon activation of specific signaling pathways but is absent in yeast. To circumvent the problem, a new version of the yeast two-hybrid system was developed in which a tyrosine kinase is coexpressed in yeast, along with the fusion proteins containing the SH2 domain and the phosphorylated peptide and the system is dubbed the yeast tribrid system [36]. Thus, the expressed tyrosine-containing peptide will be phosphorylated by the expressed kinase, enabling it to form a complex with the SH2 domain fusion protein. This system seems to be ideally suited for high-throughput screen to identify inhibitors of the SH2 domain–ligand interaction as well as inhibitors for the kinase. It remains to be seen how successful such screening system is for identifying inhibitors for specific SH2 domains.
3.2
Yeast three-hybrid system
The yeast three-hybrid system is adapted from the yeast two-hybrid system, except that the DNA-binding domain and the transactivation domain is bridged by two ligand–receptor pairs, instead of two interacting proteins in the two-hybrid system (Figure 5). Thus, the receptor for one ligand is fused to the DNA-binding domain and the receptor for a second ligand is fused to the transcriptional activation domain. In addition to the two hybrid fusion proteins, the system requires a third hybrid small molecule, a dimer of two ligands. Thus, when the two hybrid fusion proteins are expressed in yeast, the small hybrid ligand would enter the cells by diffusion and form a ternary complex with the two fusion proteins, reconstituting a functional transcription factor to drive reporter gene expression, similar to the yeast two-hybrid system.
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Figure 5. The basic components of yeast three-hybrid system. The triangle and semicircle represent two different chemical ligands. UAS, upstreat activating sequences.
The three-hybrid system can be conceptually divided into two ligand– receptor pairs. One ligand–receptor pair can be prefixed, consisting of a known ligand and a known receptor. For the second ligand–receptor pair, either the ligand or the receptor can be unknown, which can be identified through screening with the three-hybrid system. In the case of a known ligand with unknown receptor, the ligand can be covalently linked to the prefixed known ligand and the resulting hybrid ligand can be used in screen a cDNA library fused to the transcriptional activation domain to identify the potential protein target for the ligand. By the same token, if the receptor is known but a suitable ligand is needed, a library of ligands all tethered to the prefixed known ligand can be screened to identify a lead that is capable of binding to the receptor target. These two complementary applications should significantly facilitate both ligand and receptor discovery processes. The idea was first reduced to practice by employing the hormone-binding domain of the rat glucocorticoid receptor and human FKBP, along with a synthetic dimer of their respective ligands, dexamethasone and FK506 [29]. The hormone-binding domain of the glucocorticoid receptor was fused with the LexA DNA-binding domain and the human FKBP was fused with the B42 transcriptional activation domain [19]. When the two fusion proteins were expressed in yeast, the addition of the dexamethasone-FK506 hybrid ligand did not lead to activation of the LacZ reporter gene. It turned out that the wild-type rat glucocorticoid recceptor, when expressed in yeast, suffers from a significant decrease in binding affinity for dexamethasone. To
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increase the affinity of the hormone-binding domain of the rat glucocorticoid receptor for dexamethasone, several mutants were created which were known to enhance the affinity of the glucocorticoid receptor expressed in yeast for dexamethasone [3, 12]. One of the double mutants, a C656G and F620S double mutant, when employed to the three-hybrid setting, yielded strong reporter gene activity in the presence of the dexamethasone-FK506 dimer. The reporter gene activation was shown to be completely dependent on the interaction between the two ligands and the receptors, as the threehybrid reporter gene activation is sensitive to inhibition by free FK506. When the expression of the FKBP-B42 fusion protein was turned off by switching the carbon source from galactose to glucose, the activation of the reporter gene was also abrogated. These control experiments unambiguously demonstrated that the activation of reporter gene was a result of the threehybrid ligand–protein interactions. The three-hybrid system was used to screen a cDNA library to isolate cDNAs that encode for FK506-binding proteins with the dexamethasoneFK506 as a bait. Similar to the yeast two-hybrid screen, the three-hybrid screen is rapid and convenient. Unlike the typical two-hybrid screen, additional controls are available for the three-hybrid screen that ensures the elimination of false positive clones (Figure 6). One control is the dependence of the reporter gene activation on the presence of the hybrid ligand. In fact, the majority of the clones identified in the initial three-hybrid screen were eliminated by this control. Another critical control is the sensitivity of the three-hybrid interaction to competition by free FK506, which enabled the elimination of another six of the nine potential positive clones. All three remaining clones were found to harbor plasmids encoding variants of cDNAs for human FKBP12. The rapid isolation of cDNAs for human FKBP12 by the three-hybrid system (in 2–3 weeks) clearly demonstrated the power of this approach for identifying protein targets for small “orphan” ligands.
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Figure 6. Summary of the yeast three-hybrid screen for target proteins for FK506.
The yeast three-hybrid system does have its limitations, including the requirement of the protein receptors to exist as a fusion protein with either the DNA-binding domain or transcription activation domain of a transcription factor. The ability of hybrid ligands to penetrate the yeast membranes is another barrier to its general application. Solutions to these potential problems are beginning to be addressed. The three-hybrid system based on the glucocorticoid receptor– dexamethasone pair has been shown to work when the other ligand–receptor pair is wild-type FKBP and FK506. It has been found that when the affinity between FKBP and FK506 is further decreased by replacing FKBP12 with other members of the FKBP family, such as FKBP13 and FKB P25 or by generating FKBP12 mutants that have decreased affinity for FK506, the reporter gene activation can no longer be detected (E. Griffith and J. O. Liu, unpublished results). This is understandable as the three-hybrid system involves the formation of a ternary complex from three partners, the two hybrid fusion proteins and the hybrid ligand, in contrast to the two individual fusion proteins required for two-hybrid system. In addition to the dexamethasone-FK506 as the hybrid ligand, two alternative hybrid ligands were recently developed and tested in a three-hybrid
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setting, which further validated the concept of the three-hybrid system. Both groups employed dihydrofolate reductase and its potent inhibitor methotrexate as one receptor–ligand pair to replace FKBP12-FK506 [22, 30]. Cornish and colleagues used the LexA DNA-binding domain and B42 transactivation domain [30] while Henthorn and coworkers employed the Gal4 DNA-binding domain and Gal4 transactivation domain [22]. In both cases, robust LacZ reporter gene activity was observed in yeast expressing appropriate fusion proteins in the presence of the dexamethasonemethotrexate hybrid ligands. Cornish and colleagues found that at the same concentration of the hybrid ligand, the reporter gene activity was about 150- fold higher for the dexamethasone-methotrexate system than for the dexamethasone-FK506-based system [30]. Assuming that the reporter gene activation is proportional to the overall affinity of the hybrid ligand for the two fusion receptors, this is expected as the affinity of methotrexate for dihydrofolate reductase is in the picomolar range [38] while the affinity of FK506 for FKBP12 is in the low nanomolar range [20, 40]. Using the Gal4based system, Henthorn and coworkers further demonstrated the utility of the system to screen a cDNA library to identify clones encoding dihydrofolate reductase [22]. Similar to the three-hybrid screen for FKBPs, no false positive clones were found after their elimination using the competition assay with excess amount of free methotrexate. It will be interesting to see whether there is a significant improvement in sensitivity of the three-hybrid system when the dihydrofolate reductase–methotrexate pair is used to replace the glucocorticoid receptor–dexamethasone pair.
4
YEAST WHOLE GENOME GENE EXPRESSION PROFILES AS FINGERPRINTS FOR TARGET VALIDATION, TARGET IDENTIFICATION, AND TOXICITY PROFILING
The expression of genes in yeast, like in any other organisms, is highly coordinated. Perturbation of one pathway or interference with the function of one protein often leads to the changes in the levels of expression of many other genes in a defined manner. The completion of the sequencing of the yeast genome allows for the comparison of expression profile of the entire yeast genome by either DNA chip or microarrays [9, 39]. The change in the expression of each gene, or lack of it, represents a point in a one-dimensional space. With nearly 6,000 yeast genes, the expression profiles in yeast represents a 6,000-dimentional space that allows for extremely precise comparison of gene expression patterns. This, together with the collection of
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the yeast knockout mutants, offers a great opportunity to validate drug targets that are conserved between yeast and humans. A pioneering set of experiments validating the concept with microarrays was carried out with the immunosuppressant drugs cyclosporin A (CsA), FK506 and yeast mutants in which the drug-binding proteins are deleted [35]. CsA and FK506 represent a new class of drugs that act by forming ternary complexes with their respective immunophilins and the ultimate target, a calcium-dependent protein phosphatase known as calcineurin [10, 31]. Thus, CsA binds to cyclophilins while FK506 binds to FKBPs. It is the binary CsA–cyclophilin complex and the FK506–FKBP complex that then binds to calcineurin, inhibiting its protein phosphatase activity. As calcineurin and immunophilins are conserved from yeast to humans, yeast has been exploited as a model system to gain insight into the molecular mechanism of inhibition of calcineurin by the immunosuppressants [21]. Although calcineurin has been shown to be a target for these immunosuppressants, how specific they are remains unknown. Using microarrays, the “signatures” in gene expression patterns associated with the treatment of wild-type yeast with the immunosuppressants and those associated with the deletion of calcineurin and immunophilins were determined. It was found that treatment of wild-type yeast with FK506 caused a greater than twofold change in expression of 36 genes. A strikingly similar signature was observed when an isogenic calcineurin mutant strain was compared with the wild-type yeast or when the wild-type yeast was treated with CsA with high correlation coefficients (Figure 7). These signatures, however, showed no statistically significant correlation with 40 other perturbations in the form of either genetic mutations or drug treatment that are known to be unrelated to calcineurin pathway. These results gave indication that calcineurin is a likely target for both CsA and FK506.
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Figure 7. Correlation coefficients for gene expression profiles as a result of the different types of perturbation of yeast using microarray.
To validate that the convergence of the gene profile signatures of calcineurin mutant and those upon treatment of yeast with CsA and FK506 did not result from coincidence, the signatures of the calcineurin mutant upon treatment with both drugs were obtained. The profile of calcineurin mutant treated with CsA or FK506 is completely different from those of wild-type yeast treated with the drugs or calcineurin mutant, indicating that calcineurin is necessary for the changes in gene expression in wild-type yeast cells. Since the inhibition of calcineurin by FK506 required the preformation of the FK506–FKBP complex, the FK506 treatment signature was also obtained with fpr1 mutant strain of yeast. Similar to calcineurin mutant, the gene expression profile for fpr1 mutant is also different from the wild-type FK506 treatment signature, indicating that FKBP in yeast is also required for FK506 activity in yeast. In contrast, treatment of a mutant strain that is deficient in cyclophilin, the cph1 mutant, with FK506 yielded a signature that is similar to the wild type, consistent with the fact that cyclophilin does not mediate the action of FK506. These results confirm that calcineurin is a common target for both CsA and FK506 while FKBP a target that is specific for FK506, but not CsA. Each of the more than 6,000 individual genes and putative open reading frames has been deleted in yeast [14]. The time will come when a compendium of signatures of gene expression will be obtained for all the viable mutants of yeast and possibly the inviable mutants with conditional knockouts. Once such a database exists and experimental conditions are standardized to allow direct comparisons of newly obtained experimental data and the archival database, it is conceivable that every drug whose target has a yeast counterpart can be subjected to the same validation process, which will not only help to validate the putative target, but will also provide information about the specificity of the drug toward the intended target.
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It is worth noting, however, this approach, while powerful, does have its limitations. The major limitation lies at the limited size of the yeast genome in comparison to that of humans. As a consequence, many genes present in mammalian cells, including those involved in cell–cell communications and intracelluar signaling such as protein tyrosine kinases, are absent in yeast. The collection of the compendium of the yeast mutants is not applicable to studying drugs that interact with those proteins. Another limitation will be those genes that are essential to yeast growth. More sophisticated methods of downregulating those genes such as reduction in the expression level without complete knockout will be important for studying drugs that affect those genes. Nevertheless, the existing mutants and their signatures should take us a long way toward facilitating the study of many types of drugs already.
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Chapter 15 GENETIC ANALYSIS OF CISPLATIN RESISTANCE IN YEAST AND MAMMALS
Seiko Ishida and Ira Herskowitz * Department of Biochemistry and Biophysics, University of California, San Francisco, CA
Cisplatin is one of the most widely prescribed anticancer drugs. Its cytotoxic effect was discovered serendipitously in a study designed to test the effects of electric fields on the growth of the bacterium Escherichia coli [49]. In the presence of electric fields, bacterial growth continues but the cells do not divide. This inhibition was found to derive from the presence of a platinum complex, cis-diamminedichloroplatinum [II] (cisplatin), formed by electrolysis at the platinum electrodes [50]. The observation that cisplatin inhibits bacterial cell division raised the possibility that it might have anticancer activity, which was first demonstrated against a mouse sarcoma [51]. Since its approval by the FDA in 1979, cisplatin has been included in chemotherapeutic regimens for many different types of cancers. Notably, cisplatin-based chemotherapy is curative for the majority of patients with advanced testicular cancer, even with metastases, which was fatal in the precisplatin era [43]. Cisplatin is also effective for ovarian, bladder, cervical, head and neck, and small-cell lung cancers. Many patients with these cancers, however, eventually relapse and become refractory to cisplatin. In addition, cisplatin has minimal activity against some common cancers such as colorectal cancer. Increasing dosage to overcome resistance can cause serious side effects in the kidneys and the inner ear. Understanding the mechanism of intrinsic and acquired resistance to cisplatin is critical in developing a more effective cure for cancer. In this chapter, we review mammalian genes that have been genetically shown to be involved in cisplatin resistance and discuss use of bakers’ yeast, Saccharomyces *
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cerevisiae, for identifying genes and understanding their functions in cisplatin resistance.
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CISPLATIN – ITS MODE OF ACTION
Intravenously administered cisplatin retains its neutral structure in plasma, where the chloride concentration is ~100 mM [52]. The mechanism by which cisplatin enters cells is not known [22]. Once cisplatin passes through the plasma membrane into the cytoplasm of a cell, where the chloride concentration drops to ~3 mM, its chlorides are displaced by water molecules. This aquated product is the reactive form, a potent electrophile that can react with any nucleophile, including the sulfhydryl groups on proteins and nitrogen donor atoms on nucleic acids.
Figure 1. An overview of cisplatin uptake and drug action.
Work from many laboratories has implicated DNA as a critical target for cisplatin cytotoxicity, the most revealing evidence being the hypersensitivity to cisplatin of both prokaryotic and eukaryotic cells deficient in DNA repair [3, 20, 45]. The most prevalent cisplatin-induced DNA adduct is the intrastrand cross-link in which the platinum is covalently bound to the N7 positions of the imidazole ring of two adjacent guanines [15]. The intrastrand cross-links are repaired by the nucleotide excision repair pathway [4]. Unrepaired DNA damage triggers cell cycle arrest, and cells eventually undergo programmed cell death (apoptosis) [13].
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GENES THAT GOVERN CISPLATIN RESISTANCE IN MAMMALIAN CELLS
In principle, increased cisplatin resistance could result from decreasing the number of cisplatin adducts or inhibiting apoptosis in response to cisplatin-induced DNA damage. A reduction in the number of cisplatin adducts can be achieved by reducing the cellular level of cisplatin by a decrease in drug uptake or an increase in drug efflux, reducing the reactivity of the drug, reducing the accessibility of DNA to the drug, or enhancing removal of the adducts from DNA. Factors that antagonize activation of apoptotic pathways in response to DNA damage could increase cell survival to cisplatin treatment. In mammalian systems, genes that affect cellular sensitivity to cisplatin have been identified using a candidate-gene approaches described below (see Table 1).
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Mammalian genes that inhibit cisplatin accumulation in cells – cMOAT/MRP2 and ATP7B
Canalicular multispecific organic anion transporter (cMOAT), also known as apical mulitdrug resistant protein (MRP2), is predominantly localized to the canalicular membrane of hepatocytes and to the apical membrane of proximal tubules in the kidney. Because of its structural and functional similarities with MRP1, an integral membrane protein associated with multidrug resistance, a potential involvement of cMOAT/MRP2 in cisplatin resistance was tested. Human hepatic cancer cell lines transfected with antisense cMOAT/MRP2 cDNA exhibit an increase in cisplatin sensitivity accompanied by an increase in cellular accumulation of cisplatin [33]. In addition, overexpression of human cMOAT/MRP2 increases cisplatin resistance in various mammalian cell lines [10, 31] and decreases drug accumulation in Chinese hamster ovary cells [31]. These observations demonstrate that cMOAT/MRP2 inhibits cisplatin accumulation in cells, possibly by enhancing drug efflux. ATP7B is a copper-transporting P-type ATPase that transports copper into the lumen of the Golgi and into secretory vesicles. It belongs to a class of heavy metal transporting P-type ATPases that pump cadmium, zinc, silver, and lead in addition to copper. In a study designed to investigate a role of ATP7B in resistance to various heavy metals including the platinum compound cisplatin, it was noted that overexpression of human ATP7B in human epidermoid carcinoma cells conferred increased cisplatin resistance, accompanied by an increase in cisplatin efflux [34]. These findings suggest that ATP7B protein may serve as a pump to assist cisplatin efflux.
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2.2
Mammalian genes that inactivate cisplatin by increasing thiols – γ -glutamylcysteine synthetase and metallothionein
Cisplatin is a potent electrophile inside cells and reacts with any nucleophile, including the sulfhydryl groups on proteins and nucleophilic groups on nucleic acids. Glutathione (γ-glutamylcysteinyl glycine, GSH), one of the most abundant thiols in cells, binds to cisplatin and the GSH– cisplatin complex is transported across the plasma membrane [27]. The ratelimiting step of GSH synthesis is catalyzed by γ-glutamylcysteine synthetase (γ-GCS). Overexpression of human γ-GCS in a human small-cell lung cancer cell line increases the cellular GSH content, accompanied by a decrease in cellular cisplatin level and an increase in cisplatin resistance [37]. Furthermore, human colon cancer cells transfected with a hammerhead ribozyme against γ-GCS mRNA exhibit a reduction in the level of γ-GCS transcripts, an increase in cisplatin sensitivity [48], and a decrease in cisplatin efflux [25], suggesting that γ-GCS protein protects cells from cisplatin toxicity by sequestering intracellular cisplatin and exporting it. Metallothioneins (MTs) are cysteine-rich proteins that constitute a major fraction of intracellular protein thiols. Cisplatin has been shown to covalently bind MT [36]. Overexpression of human MT–IIA in mouse cells results in increased cisplatin resistance [32]. Cells derived from MT-null mice lacking both MTI and MTII exhibit enhanced sensitivity to cisplatin [35]. The MT-null mice manifest more severe kidney and liver damage after cisplatin treatment and exhibit reduced survival after cisplatin injection compared to wild-type mice [42]. Taken together, these results demonstrate that MTs play a role in detoxification of cisplatin.
2.3
Mammalian genes that mediate cytotoxic response to cisplatin adducts – MSH2 and MLH1
Mismatch repair is a post-replication repair system that corrects unpaired or mispaired nucleotides. Deficiency in this repair system predisposes cells to genomic instability. A defect in mismatch repair plays an important role in hereditary nonpolyposis colon cancer and may play a role in a variety of sporadic tumors as well. In addition to its role in processing mismatched bases, mismatch repair in E. coli has been implicated in cytotoxic effects of DNA-damaging agents including cisplatin [19]. A mismatch recognition complex in human consists of Msh2 and GTBP (hMsh6), whose binding to mismatched bases is further stabilized by the association of Mlh1 and Pms2 proteins. Purified human Msh2 binds to cisplatin G–G intrastrand adducts [12, 47], and both hMsh2 and hMlh1 are detected on cisplatin-
Chapter 15: Genetic Analysis of Cisplatin Resistance in Yeast and Mammals 397 treated DNA after incubation with nuclear extract [16]. Human cancer cell lines deficient in mismatch repair due to mutations in hMSH2 or hMLH1 are more resistant to cisplatin treatment than cells whose repair deficiency is complemented by introduction of chromosome 3 (containing hMSH2) or chromosome 2 (containing hMLH1) [1]. No difference is observed in the level of cisplatin bound to DNA or in the rate of removal of cisplatin-DNA adducts between these mismatch repair-deficient and -proficient cells [9]. Furthermore, cell lines deleted for the MSH2 gene demonstrate increased resistance to cisplatin compared to wild-type cell lines in vitro and when implanted in nude mice [17]. It is hypothesized that mismatch repair proteins sense the cisplatin-induced damage and initiate a series of events that results in cell cycle arrest and death. Two models have been proposed to explain the connection between the mismatch repair pathway and sensitivity to DNA damage caused by anticancer drugs such as cisplatin. In one model, DNA lesions are recognized and processed by the mismatch repair pathway, but because mismatch repair excises a tract from the newly incorporated strand, damage in the parental strand is not removed. Repeated attempts at processing are proposed to create persistent gaps that trigger cell cycle arrest or cell death. This model is consistent with the observations that mismatch repair-proficient cells show a reduced level of nascent DNA synthesis as measured by incorporation of radiolabeled nucleotides into chromosomal DNA after treatment of cells with cisplatin [56] or into cisplatinated plasmids after incubation in vitro with cell extracts [14]. In addition, mismatch repair-proficient cells are less efficient in reactivation of cisplatin-damaged plasmid DNA as measured by luciferase activity from a cisplatin-damaged reporter plasmid [9]. It is possible that a more direct signaling pathway links the mismatch repair pathway and cisplatin cytotoxicity. Assembly of mismatch repair proteins on cisplatinDNA adducts, whose DNA structure is different from that of unpaired or mispaired bases [55], could initiate a signaling cascade that leads to cell death.
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3
UNDERSTANDING MECHANISMS OF CISPLATIN RESISTANCE IN THE BUDDING YEAST SACCHAROMYCES CEREVISIAE
3.1
Yeast as an experimental system to understand mechanisms of cisplatin resistance
Yeast provides excellent opportunities to identify genes and functions of their products in cisplatin resistance for three reasons. First, yeast cells are sensitive to cisplatin treatment. This enables isolation of mutants with increased cisplatin resistance. Second, at least 30% of the yeast genome has mammalian homologs [5]. Genes identified in yeast may thus have mammalian counterparts that function in a similar manner. Third, genetic manipulations are much easier and faster in yeast compared to mammalian systems. The doubling time is ~90 min compared to >20 h in cultured mammalian cell lines [23]. The fact that yeast can exist as a haploid with only one copy of each chromosome makes it possible to carry out random mutagenesis or targeted deletion of a gene and observe a loss-of-function phenotype. Mutations in different genetic loci can be combined by crossing haploid mutants and sporulating them. Two genetic approaches can be taken to identify genes involved in cisplatin resistance. One is to look for genes that confer increased cisplatin resistance when overexpressed. The other is to mutagenize cells and look for mutants that show increased cisplatin resistance and identify genes altered in the mutants. The two methods are complementary. In overexpression screens, one may miss genes whose products act in a complex with other proteins. In this case, overexpressing one component of the complex may not result in a phenotype. Such genes may be uncovered by mutagenesis screens, since disruption of any one of the components may inactivate the entire complex. In mutagenesis screens, genes that have redundant functions will be missed. Such genes, however, may be identified in overexpression screens.
3.2
Identification of yeast genes that govern cisplatin resistance
Three methods have been undertaken to identify yeast genes which could potentially govern cisplatin resistance – searching for genes whose products bind to platinated DNA and genes whose overexpression or loss of function allows cells to grow in the presence of a toxic level of cisplatin (Table 1).
Chapter 15: Genetic Analysis of Cisplatin Resistance in Yeast and Mammals 399 Table 1. Genes that govern cisplatin resistance. Protein Genes that govern cisplatin resistance Human cMOAT/MRP2 Canalicular multispecific anion transporter/apical multidrug resistant protein Human ATP7B Copper-transporting P-type ATPase on Golgi Human γ-glutamylcysteine Enzyme involved in synthtase glutathione synthesis Human MT Metallothionein Human MSH2 Mispatch repair protein Human MLH1 Mispatch repair protein Human SRPK1 Kinase Mouse CTR1 Copper transporter Yeast 1XR1 HMG-domain protein Yeast PHR1 Yeast CIN5 Yeast YDR259c Yeast PDE2 Yeast ZDS2 Yeast SKY1 Yeast CTR1
Photolyase Unknown Unknown cAMP phosphodiesterase Unknown Kinase Copper transporter
Suggested role in cisplatin resistance Cisplatin efflux
Cisplatin efflux Inactivation of cisplatin Inactivation of cisplatin Enhancing cell death Enhancing cell death Unknown Cisplatin uptake Shielding adducts from repair Unknown Unknown Unknown Unknown Unknown Unknown Cisplatin uptake
In an effort to identify genes whose products bind to platinated DNA, a λ-gt11 yeast expression library was screened with radiolabeled platinated DNA, leading to the identification of IXR1 [6]. IXR1 encodes a protein with an high mobility group (HMG)-box motif known to bind irregular DNA structures. Ixr1p specifically binds to platinated DNA but not to UVdamaged DNA. Disruption of IXR1 results in a twofold increase in resistance to cisplatin. In another study, Phr1p, a yeast photolyase that binds to UV-induced cyclobutane dimers and repairs the damage was shown to bind to DNA damage caused by cisplatin [18]. Cisplatin sensitivity correlates with gene dosage of PHR1. Two screens have been carried out to search for genes that confer resistance to cisplatin when overexpressed in wild-type cells or in cells sensitized to cisplatin by inactivation of nucleotide excision repair. Wild-type cells transformed with multicopy plasmids carrying CIN5 or YDR259c, both encoding proteins with bZIP motifs commonly present in AP-1 transcription factors, exhibit increased resistance to cisplatin [21]. Cellular accumulation of cisplatin in these cells is not affected. Deletion of either CIN5 or YDR259c, however, does not make cells more sensitive to cisplatin. rad4 mutants, which are defective in nucleotide excision repair, become more resistant to cisplatin when transformed with multicopy plasmids carrying PDE2, a cAMP
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phosphodiesterase, or ZDS2, which is implicated in cell cycle control and resistance to multiple drugs [7]. The increase in cisplatin resistance is not accompanied by a reduction in drug accumulation. The effect of overexpression of these genes in wild-type cells has not been reported. A transposon library was used by two groups to mutagenize cells and to isolate mutants with increased cisplatin resistance. In this method, cells are transformed with yeast genomic fragments whose sequence is interupted at random places by a transposon [8]. Transposon-mediated mutagenesis allows rapid identification of the sequence disrupted by the transposon in the mutant and subsequent identification of the affected gene. Screening of transposon-mutagenized rad4∆ mutants led to the identification of the SKY1 gene, whose deletion in a rad4∆ background and in wild-type cells causes an increase in cisplatin resistance [53]. Overexpression of SKY1 in rad4∆ results in increased sensitivity to the drug. Sky1p is a kinase thought to phosphorylate regulators of RNA processing. How SKY1 affects cisplatin resistance is not known. rad4∆ sky1∆ mutants display similar levels of cisplatin in cells and in DNA compared to rad4∆ mutants [54]. In another study, transposon-mediated mutagenesis was performed in wild-type cells, in which case inactivation of the MAC1 gene resulted in increased cisplatin resistance [26]. MAC1 encodes a transcription factor [29, 57, 38, 44]. In subsequent studies, it was shown that deletion of the high-affinity copper transporter gene CTR1, one of the genes whose transcription is activated by Mac1p, causes cisplatin resistance similar to that of mac1∆ mutants [26]. Both mac1∆ and ctr1∆ mutants show a decreased level of platinum in cells and in DNA.
3.3
Understanding mechanisms of increased cisplatin resistance in yeast mutants and implications in mammals
Measurement of platinum levels in cells and in DNA is a powerful method for understanding the basis for cisplatin resistance in mutant strains. For many genes, increased resistance is not accompanied by changes in cellular accumulation of the drug or the cisplatin adduct level. Overexpression of CIN5 or YDR259c in wild-type cells or overexpression of PDE2 or ZDS2 in rad4∆ mutants confers an increase in cisplatin resistance, but the level of cellular cisplatin is unchanged [21, 7]. Deletion of SKY1 in rad4∆ mutants causes an increase in resistance, but no change is observed in the levels of cisplatin in cells or in DNA [54]. Among the yeast genes implicated in cisplatin resistance, IXR1 and CTR1 are the most characterized for their roles in governing cisplatin sensitivity, which appear to be conserved in mammals as well.
Chapter 15: Genetic Analysis of Cisplatin Resistance in Yeast and Mammals 401 3.3.1
Ixr1p shields cisplatin adducts from repair
Ixr1p is a protein that contains HMG domain and represses transcription of the COX5b gene by binding to its upstream regulatory region [40]. One of the common features of HMG-domain proteins is their ability to bind to irregular DNA structures, and several of them including Ixr1 have been demonstrated to bind to cisplatin adducts [30]. Deletion of IXR1 results in a twofold increase in cisplatin resistance, accompanied by a threefold decrease in the level of adduct [6]. No change in cisplatin resistance is observed in cells deleted for ROX1, another repressor of COX5b, indicating that the increase in resistance observed in the ixr1∆ mutant is not due to overexpression of COX5b [46]. The increased resistance observed in an ixr1∆ mutant is dependent on intact excision repair:deletion of IXR1 in cells defective in nucleotide excision repair, the major repair mechanism that removes cisplatin adducts, does not make them more resistant to cisplatin, i.e., a rad2∆ ixr1∆ strain is as sensitive to cisplatin as a rad2∆ strain [45]. It has thus been proposed that binding of Ixr1p to cisplatin adducts shields them from being repaired by the excision repair. In mammals, human HMG2 confers increased cisplatin sensitivity when overexpressed in a human non-small cell lung cancer cell line [2]. In these cells, the level of cellular platinum is increased. Two HMG proteins, HMG1 and the human mitochondrial transcription factor (mtTFA), inhibit nucleotide excision repair of cisplatin-damaged DNA fragments in vitro [24]. HMG-domain proteins thus appear to play an important role in both yeast and mammals in limiting access to cisplatin adducts by the excision repair machinery. 3.3.2
III.3.2 Ctr1p mediates cisplatin uptake
Ctr1p is a high-affinity copper transporter residing in the plasma membrane. Deletion of CTR1 results in a twofold increase in cisplatin resistance and a twofold decrease in cellular accumulation of cisplatin [26]. The increase in resistance in the ctr1∆ mutant is not likely to be a secondary effect of intracellular copper deficiency due to the ctr1∆ mutation, since deletions of other copper transporter genes or genes involved in intracellular copper trafficking and utilization do not result in increased cisplatin resistance. Furthermore, addition of copper to the ctr1∆ mutant, at a concentration that restores copper uptake and growth to this mutant, does not increase sensitivity to cisplatin, suggesting that the resistance observed in the ctr1∆ mutant is not due to copper starvation. Addition of copper to wildtype cells increases cisplatin resistance and decreases cisplatin accumulation,
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both of which are not observed in the ctr1∆ mutant, indicating that the effects of copper on reducing cisplatin uptake are mediated by the copper transporter Ctr1p [26]. Interestingly, cisplatin treatment results in delocalization and degradation of Ctr1p [26]. Taken together, these findings support a hypothesis that the copper transporter Ctr1p mediates uptake of cisplatin in yeast. Mammals contain functional homologs of the yeast CTR1 gene, hCTR1 in humans and mCTR1 in mice, which complement a yeast ctr1 mutant for intracellular copper deficiency [41, 58]. Mouse cell lines that are wild-type, heterozygous, or homozygous for a knockout allele of mCTR1 display a graded increase in cisplatin resistance and decrease in cisplatin accumulation [26]: heterozygous cells are fourfold more resistant to cisplatin and exhibit a 35% decrease in cisplatin accumulation compared to wild-type cells; homozygous cells show an eightfold increase in resistance and a 70% decrease in drug accumulation. These observations in mice suggest that hCtr1 in humans, which is 92% identical to mCtr1, may play a key role in cisplatin uptake and resistance. The mechanism of cisplatin uptake has been unclear. Inability to saturate the rate of cisplatin uptake supports a simple diffusion model, whereas the existence of agents and conditions that modulate cisplatin accumulation, such as pH, osmolarity, sodium, potassium, protein kinase C (PKC), protein kinase A (PKA), and the calcium/calmodulin pathway, suggests that some component of cisplatin uptake is mediated by a transport mechanism [22]. It has been proposed that about 50% of the intial rate of uptake is due to passive diffusion and that the remaining 50% is due to facilitated diffusion through an as yet unidentified gated channel. Ctr1p might be responsible for the facilitated uptake of cisplatin. Cisplatin uptake is not completely abolished in the ctr1∆ mutants in both yeast and mice [26]. The residual cisplatin uptake observed in the ctr1∆ mutants might be due to diffusion across the plasma membrane or to additional transport proteins. 3.3.3
Other yeast genes whose mammalian homologs are implicated in cisplatin resistance
All of the yeast genes identified in the screens have mammalian homologs or are involved in functions that are present in mammals. In addition to IXR1 and CTR1, mammalian homologs of the yeast PDE2 and SKY1 genes may have roles in cisplatin resistance in mammals. Overproduction in yeast of Pde2p, a phosphodiesterase that functions in the PKA pathway, confers increased cisplatin resistance to mutants defective in excision repair [7]. Studies in mammalian cells also suggest potential links between the PKA pathway and cisplatin sensitivity [11]. Deletion of the
Chapter 15: Genetic Analysis of Cisplatin Resistance in Yeast and Mammals 403 SKY1 gene in the excision repair-deficient rad4∆ mutant increases cisplatin resistance [53]. Antisense experiments in human ovarian carcinoma cell lines demonstrate that inactivation of its human homolog, SRPK1, leads to increased cisplatin resistance [53]. The basis for increased cisplatin resistance in cells overexpressing PDE2 or in cells deleted for SKY1 is not known. Having yeast models should facilitate elucidation of the functions of these mammalian gene products in mediating cisplatin sensitivity.
3.4
Limitations of the use of yeast in understanding cisplatin resistance in mammals
Although mismatch repair deficiency has been associated with cisplatin resistance in mammalian cells [17], deletion of mismatch repair genes in yeast – MSH2, MSH3, MSH6, MLH1, or PMS1 – does not result in increased cisplatin resistance [26]. In mammals, mismatch repair deficiency is also associated with resistance to other DNA-damaging agents [39]. It has been postulated that detection of DNA damage by the mismatch repair system triggers a pathway that leads to apoptosis. Cisplatin-induced death of mammalian cells exhibits certain features of apoptosis, such as chromatin condensation, DNA fragmentation, and activation of key regulators of apoptosis such as p53 and p73 [13, 28]. In yeast, however, there has been no clear evidence for the existence of an active cell death pathway such as apoptosis. Lack of involvement of mismatch repair genes in cisplatin resistance in yeast may reflect differences in the way mammalian cells and yeast cells die after exposure to toxic doses of cispatin.
4
CONCLUSIONS AND PERSPECTIVES
Cisplatin is one of the most widely used anticancer drugs effective in the treatment of a variety of cancers. Intrinsic or acquired resistance to cisplatin reduces its efficacy, however, which limits its curative potential. Considerable effort has been made to define the cellular and molecular mechanisms responsible for cisplatin resistance in mammals. Most of the mammalian genes governing cisplatin resistance were identified due to their biochemical properties – cMOAT/MRP2 and ATP7B are involved in efflux of drugs and metals, γ-glutamylcysteine synthetase, and metallothionein increase intracellular thiols that react with cisplatin, and mismatch repair proteins and HMG domain proteins bind to platinated DNA. Genetic selections in yeast have identified a number of genes that affect cisplatin resistance, most of which either have mammalian homologs or are involved in functions that are also present in mammals. Genetic analyses of
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the yeast ixr1∆ mutants indicated that HMG-domain proteins confer sensitivity to cisplatin by shielding cisplatin adducts from repair. These findings complement biochemical studies of the role of human HMG proteins in cisplatin resistance. Increased cisplatin resistance and decreased cisplatin uptake of yeast ctr1∆ mutants led to the identification of the mammalian copper transporter Ctr1 as a mediator in cisplatin uptake. Although deletion and overexpression studies in yeast and mammalian cells have clearly demonstrated the role of some genes in cisplatin resistance, whether these genes are involved in cisplatin resistance of tumors in patients remains to be determined. To understand clinical mechanisms of resistance, analysis of specimens from patients is necessary. Studies of genetically modified cell lines and mice will provide complementary information. Studies with yeast have identified additional candidate genes that should be closely monitored in clinical settings. Mutations in these genes or changes in their expression levels may influence the efficacy of cisplatin and thus could be a crucial factor that may determine the prognosis of the patient. Because Ctr1 is a cell-surface protein, it may be a good target for developing drugs that would antagonize cisplatin uptake to protect normal tissues from damage or that would facilitate cisplatin uptake to sensitize tumor cells to cisplatin. Localized administration of such Ctr1 modifiers might greatly enhance the efficacy of cisplatin against tumors and minimize its side effects.
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S. Ishida and I. Herskowitz modification of the yeast anticancer drug sensitivity profile accompanied by a mutator phenotype. Mol. Pharmacol. 61:659–666. Takahara, P. M., A. C. Rosenzweig, C. A. Frederick, and S. J. Lippard. 1995. Crystal structure of double-stranded DNA containing the major adduct of the anticancer drug cisplatin. Nature 377:649–652. Vaisman, A., M. Varchenko, A. Umar, T. A. Kunkel, J. I. Risinger, J. C. Barrett, T. C. Hamilton, and S. G. Chaney. 1998. The role of hMLH1, hMSH3, and hMSH6 defects in cisplatin and oxaliplatin resistance:correlation with replicative bypass of platinum-DNA adducts. Cancer Res. 58:3579–3585. Yamaguchi-Iwai, Y., M. Serpe, D. Haile, W. Yang, D. J. Kosman, R. D. Klausner, and A. Dancis. 1997. Homeostatic regulation of copper uptake in yeast via direct binding of MAC1 protein to upstream regulatory sequences of FRE1 and CTR1. J. Biol. Chem. 272:17711–17718. Zhou, B., and J. Gitschier. 1997. hCTR1:a human gene for copper uptake identified by complementation in yeast. Proc. Natl. Acad. Sci. USA 94:7481–7486.
Chapter 16 USING YEAST TOOLS TO DISSECT THE ACTION OF ANTICANCER DRUGS: MECHANISMS OF ENZYME INHIBITION AND CELL KILLING BY AGENTS TARGETING DNA TOPOISOMERASES
Anna T. Rogojina, Zhengsheng Li, Karin C. Nitiss and John L. Nitiss Department of Molecular Pharmacology, St. Jude Children’s Research Hospital, Memphis, TN 38105
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INTRODUCTION
Resistance to anticancer agents is one of the defining problems of cancer pharmacology. Drug resistance is clearly a major obstacle to the cure of many neoplastic diseases. Through the years we have come to understand that anticancer drug resistance has different biochemical and molecular mechanisms depending on the class of drug. Early studies centered on trying to understand “acquired drug resistance”, alterations in cancer cells that were originally sensitive to one or more chemotherapeutic agents, but which acquired changes that attenuated the cell cytotoxicity of the original agent, and also frequently resulted in “multidrug resistance”, resistance to multiple classes of anticancer agents. While prevention of acquired drug resistance, and a detailed understanding of mechanisms leading to drug resistance are still critical concerns of cancer pharmacology, we now understand that many cancer cells accumulate changes that result in inherent drug resistance. Alternatively, some of the changes leading to the development of neoplastic disease can make cancer cells inherently more sensitive to specific classes of anticancer agents. Current experimental approaches are providing a detailed description of the molecular changes that occur in many types of cancers. 409 J.L. Nitiss et al. (eds.), Yeast as Tool in Cancer Research, 409–427. © 2007 Springer.
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One approach to converting this description of the molecular changes that occur in cancer into effective therapeutic strategies entails the design of new agents that specifically inhibit the proteins that have become essential for the survival and proliferation of cancer cells. A second approach is to optimize drugs acting against established anticancer targets. Both approaches require a detailed understanding of how anticancer drugs can lead to cell death, starting with a precise understanding of the target(s) of a drug, the cellular process affected by drug action, and the consequences of target inhibition. DNA topoisomerases are an important target of many clinically active anticancer agents. Both type I and type II topoisomerases are targeted by various anticancer drugs. Type I topoisomerases are targeted by the camptothecins, which have demonstrated substantial clinical activity in a wide range of tumors. Topoisomerase II is the target of many chemically diverse agents including epipodophyllotoxins such as etoposide [74] and anthracyclines such as doxorubicin [64]. While these agents have substantial antitumor activity, a wide range of questions remains to be answered. For example, while it is well established that drugs such as etoposide target topoisomerase II, the biochemical mechanisms leading to enzyme inhibition remain to be identified. This is an important issue for topoisomerase II-targeting agents such as mAMSA that have been clinically disappointing. While many properties of a drug can influence its clinical activity, differences in biochemical mechanisms of drugs affecting the same target may be of decisive importance. Since drugs targeting topoisomerases are typically used in combination with drugs affecting other targets, a comprehensive understanding of cell-killing mechanisms is an important tool for devising optimal multidrug schedules. Finally, as is the case for most types of cytotoxic chemotherapy, drugs targeting topoisomerases can lead to a wide range of undesirable side effects. For topoisomerase II-targeting drugs, this includes the induction of translocations that can cause secondary malignancies [38]. Yeast has been a powerful model system for studying both the biological roles of DNA topoisomerases, and for studying the mechanism of action of drugs targeting these enzymes [41, 42]. Since topoisomerases play roles in so many different DNA transactions, higher eukaryotes do not readily tolerate mutations that impair topoisomerase activity. By contrast, introduction of defined topoisomerase alterations has proven to be a productive approach to understanding topoisomerase biology as well as drug action. This review concentrates on using yeast to study antitopoisomerase drug action. Several recent reviews have provided a detailed description of the roles of topoisomerases in different biological processes [41, 42, 71].
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HOW TOPOISOMERASE-TARGETING DRUGS KILL CELLS: USING YEAST FOR MECHANISM-BASED STUDIES OF DRUG ACTION AND DRUG SPECIFICITY
The reaction mechanism of all topoisomerases includes the transient cleavage of DNA by a tyrosine participating in a transesterification reaction, resulting in the formation of a covalent phosphotyrosine linkage with DNA, and concomitant breaking of the phosphodiester backbone of DNA. Type I topoisomerases make single-strand breaks in DNA while type II enzymes in eukaryotic cells are homodimeric enzymes that introduce double-strand breaks using one tyrosine from each subunit. DNA topoisomerases use these strand breaks as DNA gates for carrying out strand passage and altering DNA topology. After strand passage has occurred, a reverse reaction reseals the DNA break, using the energy of the phosphotyrosine bond. A full description of mechanistic and structural aspects of topoisomerase biochemistry can be found in recent reviews [11, 72]. Most drugs targeting DNA topoisomerases act by increasing the level of the cleaved intermediate where the enzyme is covalently bound to DNA. Typically, such agents reversibly inhibit the DNA ligation reaction that seals DNA breaks after strand passage. Because the enzyme remains trapped on the DNA, drugs targeting topoisomerases will inhibit enzyme activity, as well as generating enzyme-mediated DNA damage. Although topoisomerases are critical for many DNA transactions that take place during replication, transcription, and chromosome segregation, the major killing mechanism for these agents arises from the generation of enzyme-mediated DNA damage. This hypothesis was demonstrated using yeast strains with alterations in DNA topoisomerases. The idea behind the experiments is as follows: if topoisomerase-targeting drugs kill cells by depriving cells of an activity important for survival, then decreasing the level of the targeted topoisomerase should lead to enhanced drug sensitivity. By contrast, if enzyme-mediated DNA damage is responsible for cell killing, then loss of topoisomerase activity will decrease the amount of enzymemediated DNA damage, and cell survival will be enhanced. The first experiments testing this experimental scheme were applied to topoisomerase I. Topoisomerase I is not required for viability in Saccharomyces cerevisiae, and cells carrying a deletion of the TOP1 gene have subtle phenotypes, with minimal effects on cell viability [22, 65, 66]. Therefore, a drug targeting topoisomerase I such as camptothecin could result in cell killing by two possible mechanisms. The first mechanism is enzymemediated DNA damage, which would be abolished upon deletion of the TOP1 gene. The second mechanism would occur if camptothecin affects (an essential) target besides topoisomerase I. In the latter case, deletion of
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topoisomerase I would not prevent camptothecin-induced cell killing. When yeast cells lacking top1 were treated with camptothecin, drug-induced cytotoxicity was completely eliminated [18, 39]. This result eliminated the possibility that camptothecin targets some other important function, and established that the sole cytotoxic target of camptothecin (in yeast) is topoisomerase I. A similar series of experiments were carried out for drugs that target DNA topoisomerase II. Since yeast topoisomerase II is essential for viability [15, 23], the approach described above required modification. First, a simple test for poisoning of topoisomerase II could be carried out by assessing the effect of enzyme overexpression on drug sensitivity [45] This type of experiment can show that poisoning, i.e., generation of enzyme-mediated DNA damage plays a role in drug sensitivity. However, it cannot demonstrate whether inhibition of enzyme activity also contributes to cell killing. To test this point, drug sensitivity can be examined in contexts where enzyme activity is reduced, but not completely eliminated. Since several temperature-sensitive alleles of TOP2 have been described, one way to reduce enzyme activity is to grow strains at a semi-permissive temperature. For example, the top2-1 allele, has normal enzyme activity at 25°, greatly reduced activity (∼10% of wild type) at 30°, and minimal activity at 35° [15]. Importantly, cells are still able to grow at 30°, albeit with a substantially reduced growth rate. Thus, if cell killing depends entirely on generation of enzyme-mediated DNA damage, the reduced enzyme activity at 30° should result in drug resistance. Conversely, if reduction of enzyme activity also plays a role in cell killing, loss-of-enzyme activity due to the thermolabile enzyme should enhance the effects of drug-mediated enzyme inhibition. Growing top2-1 cells at 30° clearly leads to high levels of resistance to both etoposide and mAMSA, indicating that generation of DNA damage is the most important determinant of cell killing [46]. Furthermore, since changing Top2 activity abolished cell killing, cytotoxicity by these agents in yeast depends only on the drugs action against Top2. While this experimental strategy for testing drug specificity can work for any drug causing cell killing in yeast, the identification of alleles conferring resistance to multiple classes of Top2 poisons has proven to be a simpler experimental strategy. In practice, one compares the sensitivity of a compound in an isogenic pair of yeast strains that differ only at the Top2 gene, with one strain carrying a wild-type Top2 gene, and the other a drug resistant allele [29, 50]. The experimental systems described above have clearly demonstrated the importance of generation of enzyme-mediated DNA damage in cell killing by drugs targeting either topoisomerase I or topoisomerase II. One implication of these results is that since cell killing does not depend significantly on inhibition of enzyme activity, there may not be a strong rationale for combining drugs targeting both topoisomerases. This combination has been
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investigated in both preclinical models and in several clinical trials. Frequently the combination of drugs targeting topoisomerase I and topoisomerase II leads to high levels of toxicity, but little clinical efficacy [24, 40, 56]. While elevated levels of DNA damage may be achievable by combining topoisomerase I and topoisomerase II targeting agents, there is no other mechanistic rationale for this type of combination. In addition to testing the general mechanism of action of drugs targeting topoisomerases, the strains described above have been very useful in demonstrating the targets of novel agents. For example, fluoroquinolones targeting eukaryotic topoisomerase II were shown to be very specific for topoisomerase II [17]. In some cases, the yeast model systems clarified mechanisms of drug action in cases where biochemical data were ambiguous or contradictory [47, 50]. Finally, this distinct pattern of sensitivity can be used for screens for identifying new topoisomerase-targeting agents. Another important aspect of drugs targeting DNA topoisomerases is that drug action is typically freely reversible. Removal of drug leads to a rapid reversal of DNA cleavage both in cells, and with in vitro reactions with purified topoisomerases [10]. Liu and colleagues proposed a model, illustrated in Figure 1, that can explain how reversible topoisomerase cleavage can be converted into irreversible DNA damage [13, 76]. Proteins tracking along DNA, such as a replication fork, can collide with a covalent complex. A collision between a replication fork and a Top1 covalent complex can lead directly to a double strand break (that does not have protein covalently attached) and a gapped double helix that has protein covalently bound at the 3′ end of the gap (see Figure 1). The fate of a collision between a replication fork and a Top2 covalent complex is less clear. Since Top2 forms a very stable dimer [63], it is unclear whether the collision might break the dimer apart, leading to a double strand break with protein attached at each 5′ end. One possibility is that the stalling of the replication fork provokes a processing reaction (discussed in section 4), leading to conversion of the reversible covalent complex into an irreversible lesion. Figure 1. Topoisomerase - mediated DNA damage is responsible for cell killing. Covalent Top1 or Top2 complexes acting in front of replication forks can be converted into irreversible DNA damage. Top1 covalent complexes can be converted into double-strand breaks without additional processing, while the mechanism of generating double-strand breaks by Top2 covalent complexes remains poorly understood.
It is also instructive to consider the fate of collisions between covalent complexes and tracking proteins besides replication forks. In the case of Top2 covalent complexes, a protein tracking along DNA such as RNA
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polymerase may provoke similar effects as a replication fork; stalling of the tracking process, and breaking of the covalent complex or stimulating of processing reactions. By contrast, a collision of a tracking RNA polymerase with a Top1 covalent complex may lead to a single-strand break, but will not readily generate a double-strand break. These considerations would predict that ongoing DNA replication would be required for double-strand break generation with Top1-targeting drugs, but not required for Top2-targeting agents. Indeed, arresting replication prevents cell killing by camptothecin both in yeast and in mammalian cells, while ongoing DNA replication is not required for cell killing by Top2-targeting agents.
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RESISTANCE TO TOPOISOMERASETARGETING DRUGS: IDENTIFYING CHANGES IN TOPOISOMERASE PROTEINS THAT ALTER DRUG SENSITIVITY
A general question in cancer pharmacology concerns how cancer cells can become resistant to anticancer agents. It is appreciated that substantial information can be obtained by identifying and characterizing mutations in drug targets that confer resistance. For drugs that act as inhibitors of enzyme activity, resistance-conferring mutations are typically dominant. Therefore, the resistant mutations can be readily isolated even in mammalian cells, and proof that the mutation in the gene encoding the drug target is causally related to drug resistance is usually straightforward. Mutations leading to resistance typically occur close to drug-binding sites. Therefore, identifycation of a spectrum of drug-resistant mutants can be valuable for understanding drug mechanisms. The unique mechanism of topoisomerase poisons complicates the identification of drug-resistant forms of topoisomerases, and makes it much more difficult to demonstrate that the identified mutation causes drug resistance. As described in the previous section, resistance to topoisomerase poisons is recessive [46, 48]. Interpretation of mutations leading to drug resistance is also more complicated, since as noted above, reduction of enzyme activity is sufficient to confer drug resistance. Because of these technical challenges, yeast has proven to be uniquely useful for identifying mutant forms of topoisomerases with altered drug sensitivity, and for demonstrating whether specific mutations cause altered drug sensitivity. The first camptothecin resistant mutant in topoisomerase I was characterized by Andoh and colleagues. This mutation was present in the coding sequence of topoisomerase I in a mammalian cell line that had been selected for camptothecin resistance [2]. One strategy for demonstrating that the mutation is responsible for resistance is to reconstitute the identified
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mutation in the cDNA of Top1, and following expression in E. coli, showing that the mutation resulted in a protein that is camptothecin resistant [21]. This laborious approach showed that the resistance in the cell line (at least in part) arose from mutation(s) in Top1. Similar types of experiments were carried out using mammalian cell lines selected for resistance to either etoposide or mAMSA. While several mutations were identified in the coding sequence of Top2α [6, 25, 32], the inability to easily express eukaryotic Top2 in E. coli prevented a direct demonstration that the identified mutations resulted in drug-resistant proteins. Direct demonstration that the identified mutations altered drug sensitivity first took advantage of the strong homology between yeast and human topoisomerases. Introduction of the mutations seen in mammalian cells into the coding sequence of yeast Top2 was sufficient to confer resistance to Top2-targeting drugs [48]. Subsequently, a more rigorous demonstration that the mutations confer drug resistance was achieved by complementing yeast top2 mutations by expression of human topoisomerase II [27]. Expression of alleles of human Top2 carrying mutations identified in drug-resistant mammalian cell lines still complemented a deficiency of yeast Top2, indicating that the alleles encoded active enzymes. Furthermore, expression of these alleles in yeast also resulted in resistance to Top2-targeting drugs, efficiently demonstrating that the identified mutations alter the drug sensitivity of topoisomerase II [27]. Since the mutations confer recessive drug resistance, simple introduction of the drug-resistant Top2 allele will not confer drug resistance. Extinguishing expression of wild-type drug-sensitive alleles of Top2, along with expression of a drug-resistant allele of Top2 is required to reconstitute drug resistance. While RNA interference technologies now make such (difficult) experiments plausible, an additional issue is that ectopic expression of topoisomerases is normally deleterious [36], further complicating the use of mammalian cells for this type of experiment. In addition to characterizing mutations selected in mammalian cells, yeast represents an ideal system for selecting a broad spectrum of mutants in topoisomerase genes that alter drug sensitivity (reviewed in [5, 43, 44, 58]. In practice, the experiments in yeast have involved introduction of libraries carrying a topoisomerase gene that has been subjected to in vitro mutagenesis, followed by screening or selection for drug-resistant (or hypersensitive) clones. As suggested above, a common class of mutations will be mutations that reduce topoisomerase activity. To reduce this concern, a common approach has been to express the topoisomerase gene from a strong promoter [30, 35]. This approach can also be readily adapted to the examination of human topoisomerases by mutagenizing a human topoisomerase gene, followed by selection for altered drug sensitivity in yeast [3, 52]. Following selection for mutations with alterations in drug sensitivity, a detailed biochemical analysis of the mutant proteins to establish the
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mechanisms of resistance is required. The details of biochemical changes associated with topoisomerases with altered drug sensitivities is beyond the scope of this review (see [20, 55, 62, 72] for a discussion of the molecular mechanisms of drug sensitivity as revealed by biochemical analysis). However, it is important to note that using this approach to identify mutations conferring drug resistance has significant shortcomings. Notably, mutations conferring resistance can arise from reduced enzyme activity. Identification of some of the determinants of binding of topoisomerase Itargeting drugs has been made largely because of the determination of three dimensional x-ray structures of the ternary complex of topotecan (a camptothecin derivative), DNA, and topoisomerase I [9, 62]. Structures of Top2:DNA:drug have not yet been reported, nor have plausible detailed models for drug:protein been proposed. An alternate approach to understanding drug-protein interactions has been to identify mutations in Top2 that lead to enhanced drug sensitivity [16, 26]. Mutations conferring drug hypersensitivity probably do not occur solely due to reduced enzyme activity. Several mutations have been identified leading to high levels of hypersensitivity to Top2-targeting agents (Rogojina and Nitiss, unpublished data). This approach may be particularly useful to generate models for drug binding, and potentially to generate reagents that can be used to obtain details of drug-protein interactions.
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EVENTS DOWNSTREAM OF TOPOISOMERASE INHIBITION: GENES ALTERING SENSITIVITY TO TOPOISOMERASE-TARGETING DRUGS
An important strength of yeast genetic systems is the ability to identify genes that are required for survival under stress conditions. Yeast systems have been particularly important in identifying and characterizing DNA repair and tolerance pathways. Importantly, much of the work in DNA repair pathways in yeast has proven to be directly relevant to mammalian repair pathways [60]. There have been two broad approaches to studying the genetic control of sensitivity to topoisomerase-targeting agents in yeast. The first approach takes advantage of knowledge of the type of lesion that is generated by trapping a topoisomerase covalent complex, and assesses the roles of relevant repair pathways. More recently, it has been possible to undertake genome-wide examination of genes required for surviving topoisomerase-mediated damage. Since camptothecin generates lethal damage preferentially during S phase [49], and since sensitivity to camptothecin can be observed without additional mutations that enhance drug accumulation, several genome-wide screens have identified camptothecin hypersensitive mutants [4, 8, 14]. The poor accumulation of most topoisomerase II-targeting
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agents in yeast has made genome-wide screens more difficult for this class of drugs. These two approaches are clearly complementary, and both approaches have led to the identification of important pathways for repairing topoisomerase-mediated DNA damage. This section will first discuss some of the targeted approaches, and then summarize some of the interesting results from genome-wide approaches. The repair of topoisomerase-mediated DNA damage requires the collaboration of several distinct DNA repair pathways. As shown in Figure 1, replication in the presence of Top1 poisons leads directly to double-strand breaks. As expected, cells lacking double-strand break repair pathways such as homologous recombination are hypersensitive to camptothecin [39]. The repair of camptothecin-induced double-strand breaks includes recruitment of the histone variant H2AX to the site of double-strand breaks, with subsequent recruitment of other repair proteins [7, 53, 57]. Since the generation of double-strand breaks following collisions with topoisomerase II may require additional processing steps, the role of H2AX and other signaling processes are poorly understood with topoisomerase II poisons. Nonetheless, double-strand break repair pathways also play essential roles in cell survival following exposure to topoisomerase II poisons. Interestingly, the nonhomologous end-joining pathway of double-strand break repair is important for cell survival following exposure to the clinically important Top2 poison etoposide [31], which may relate to the genesis of translocations leading to secondary cancers. By contrast, yeast cells lacking nonhomologous end-joining are not hypersensitive to camptothecin (unpublished data), consistent with results obtained with DT-40 (chicken) cells [1]. In addition for the need to repair double-strand breaks, there are other aspects of topoisomerase/DNA covalent complexes that require other DNA repair and stress tolerance pathways. For example, in addition to doublestrand break repair, recombination pathways and replication functions are also required for restarting of blocked or collapsed replication forks [12, 37]. Topoisomerase: DNA covalent complexes have been shown to impede replication fork progression, leading to a requirement for functions needed for fork restart. Mutants have been isolated that are hypersensitive to Top1 poisons and are defective in functions that are important for lagging strand DNA synthesis [59]. These results indicate that the aberrant DNA replication that occurs in the presence of Top1 poisons results in an enhanced requirement for specific DNA replication functions that may play roles in restarting blocked or broken replication forks. In addition to replication restart mechanisms, a variety of checkpoint functions are also needed to ensure that compete and accurate completion of replication occurs. Mutants defective in both S-phase and DNA damage checkpoints are hypersensitive to camptothecin [70] (see below).
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A third aspect of repair of topoisomerase-mediated DNA damage is the repair of DNA that has protein covalently bound to a DNA end. In principle, a variety of repair nucleases, possibly in conjunction with proteolytic enzymes could repair this type of damage. Nash and colleagues have identified a specific repair enzyme that participates in the repair of Top1-mediated DNA damage [54, 75]. This enzyme, termed tyrosyl-DNA phosphodiesterase (Tdp1) can remove peptides covalently bound to DNA via a 3′ phosphortyrosine linkage. Tdp1 was originally isolated from yeast by looking directly for a biochemical activity able to disjoin a 3′ phosphotyrosine linkage from an oligonucleotide [75]. The yeast gene was subsequently cloned based on its ability to confer sensitivity to camptothecin. Interestingly, tdp1– single mutants have only modest sensitivity to camptothecin, but show synergistic sensitivity in combinations with mutations in other repair genes (such as the RAD9 checkpoint function) [54]. The observation that tdp1– single mutants have modest camptothecin sensitivity has led to a search for other proteins that may participate in the removal of proteins covalently bound to DNA. Several groups have shown that the mutation in either component of the Rad1–Rad10 nuclease greatly enhances the camptothecin sensitivity of tdp1– mutants [14, 34, 68]. The nuclease activity of Rad1–Rad10 complex is consistent with removal of a protein covalently bound to the 3′ end of DNA. Similarly, loss-of-function of the Mms4–Mus81 nuclease also enhances the sensitivity of tdp1– mutants to camptothecin [14, 34]. However, both Rad1–Rad10 and Mms4–Mus81 also play critical roles in homologous recombination. Therefore, it will be important to directly demonstrate that these nuclease complexes are able to process 3′ phosphotyrosine adducts on DNA. Removal of Top1 covalent complexes by Tdp1-dependent pathways is likely to be more complicated than the simple action of the Tdp1 protein. Removal of Top1 protein by Tdp1p is highly inefficient unless Top1p is denatured or proteolyzed prior to Tdp1 action. Furthermore, it is unclear how Tdp1p is recruited to Top1 covalent complexes. One plausible model is that covalent complexes are first recognized by unknown proteins, leading to ubiquitination and proteolysis of Top1p which would create a substrate for Tdp1 or other proteins that can complete the removal of the phosphotyrosyl peptide from DNA. As indicated above, double-strand break repair pathways play critical roles in the repair of Top2-mediated DNA damage [61]. There is less information concerning possible roles of other repair pathways. As indicated below, some of the genomic approaches are beginning to yield information concerning pathways important for surviving Top2-mediated DNA damage. Since Top2p forms a 5′ phosphotyrosyl linkage with DNA, it was predicted that Tdp1p would not be active against lesions involving Top2p. Recently, we reported that deletion of the TDP1 gene in yeast confers
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hypersensitivity to Top2-targeting agents. Combining tdp1 mutations with deletions of genes involved in nonhomologous end-joining, excision repair, or post-replication repair enhanced sensitivity to Top2-targeting drugs over the level seen with single mutants, suggesting that Tdp1 may function in collaboration with multiple repair pathways. Deletion of tdp1 can sensitize yeast cells to drugs targeting Top2 even when TOP1 is deleted, excluding models where Top2 covalent complexes can lead to trapping of Top1 (with a requirement that Tdp1 function to repair consequences of Top1 being trapped). A critical experiment showed that bacterially expressed yeast Tdp1p is able to remove a peptide derived from yTop2 that is covalently bound to DNA by a 5′ phosphotyrosyl linkage. Additionally, mutation of one of the essential histidines of Tdp1 to alanine completely ablated activity against 5′ phosphotyrosyl substrates, demonstrating that the enzyme mechanism against 5′ and 3′ modified substrates is probably identical [51]. These results establish Tdp1 as part of a pathway for removing both Top1and Top2-mediated DNA damage. Furthermore, some excision repair functions (such as the Rad2p) may also define other pathways for removing Top2 covalently bound to DNA. The development of a complete set of yeast strains that carries deletions of each yeast open reading frames represents an outstanding tool for genetic analysis on a genome-wide scale [67]. This resource allows identification of all (nonessential) genes that confer a particular phenotype, and eliminates much of the tedious mutant purification required in conventional genetic screens. Several genomic screens have been carried out for yeast mutants that are hypersensitive to DNA damaging agents. Some of these screens have used camptothecin sensitivity as part of a secondary characterization [4, 8, 73]. These screens led to the identification of more than 100 genes that confer hypersensitivity to camptothecin. These include many repair proteins described above, such as recombination and checkpoint functions. They also include proteins required for a variety of other processes including transcription, and chromatin structure that may plausibly function in repair processes. An intriguing set of mutants has no obvious connection to DNA metabolism. Further experiments will be needed to determine the genetic pathways that these proteins represent. There have also been efforts to use the deletion collection to identify mutants required for repairing Top2 mediated damage. Osheroff and colleagues screened the yeast deletion collection using high concentrations of etoposide. This screen yielded mutants required for homologous recombination as well as a novel gene, MMS22 that is also required for survival following exposure to alkylating agents and ionizing radiation. The small number of genes identified probably stems from the poor accumulation of etoposide in wild-type yeast cells. To overcome this difficulty, we have taken advantage of mutations in the TOP2 coding sequence (described in section 3) that confer hypersensitivity to mAMSA. A useful property of
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these mutants is that they confer dominant hypersensitivity to mAMSA; hence they can be readily transformed into yeast cells. In a preliminary screen, we analyzed the mAMSA sensitivity of the approximately 300 deletion strains previously shown to be sensitive to either MMS or ionizing radiation. Approximately 100 strains also showed hypersensitivity to mAMSA (Jeffrey Berk and JL Nitiss, in preparation). These experiments demonstrate that the genetic control of sensitivity to Top2-targeting drugs is also complex, with many of the same proteins participating in the repair of both Top1 and Top2 damage. Interestingly, several deletions confer camptothecin sensitivity but do not affect sensitivity to mAMSA. Other deletions show hypersensitivity to mAMSA, but wild-type sensitivity to camptothecin. These results indicate that repair of Top1 and Top2 damage differ in some details in yeast, as would be expected from the differences in the lesions generated. While the genomic approaches using the yeast deletion strains are very powerful, there are some shortcomings. First, the screens that have been carried out identify only genes that are not essential for viability. Several genes that participate in DNA repair also carry out essential functions. Second, it is unclear how many relevant mutants have been missed in the screens that have been carried out. While this is a shortcoming of any genetic screen, the fact that limited amounts of camptothecin are accumulated in wild-type yeast cells suggests that mutants that are sensitive to small amounts of damage are most likely to be isolated. An alternate way of dealing with the issues described above is to devise a way of mimicking drug action by mutations in target proteins. Mutations in topoisomerases that result in enzymes that are partly defective in religation or that exhibit elevated levels of DNA cleavage would generate the same effects in cells as treatment of cells with a topoisomerase-targeting agent [33]. Levin et al. isolated this type of mutation in yeast Top1. Bjornsti and colleagues have subsequently used these Top1 mutants to identify temperature-sensitive mutants that are inviable at a nonpermissive temperature only when the cells express an elevated cleavage mutant of Top1 [33, 59]. Importantly, they showed that this type of screen could identify mutations in genes that are essential for viability [19]. For example, the mutant screen identified mutations in CDC45, a protein involved in both replication initiation and elongation [59]. Mutations were also identified in UBC9, an E2 specifically involved in conjugation of the ubiquitin homolog SUMO to a variety of proteins [28]. The approach taken by Bjornsti and colleagues may also be applicable to studying Top2-targeting drugs. We identified a mutation in human Top2α changing Asp48 to Asn that also has the property of failing to transform yeast cells deficient in recombination repair [69]. In repair proficient strains, the Asp48Asn mutant can be expressed and complements a temperature
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sensitive top2 mutation. Purified Asp48Asn Top2α has relaxation activity similar to the wild-type enzyme, but the purified protein exhibited approximately fivefold increase in levels of drug-independent cleavage compared to the wild-type enzyme [69]. The lethality exhibited by the mutant in recombination deficient mutants is likely due to its enhanced drug independent cleavage. Since the mutant can introduce Top2-mediated DNA damage in the absence of drug, it can be exploited in screens for mutants that are deficient in repairing Top2-mediated DNA damage. The genetic approaches outlined above provide important tools for understanding the repair of DNA damage by both Top1- and Top2-targeting drugs. There are additional tools that are needed to construct biochemically testable models of repair of topoisomerase mediated DNA damage. We still lack the capability to accurately measure topoisomerase mediated damage in yeast cells. Efficient measurement of topoisomerase covalent complexes in yeast cells could be used to determine the processes that convert covalent complexes into irreversible damage, and then to measure the rates of disappearance of the protein that is covalently bound to DNA.
Figure 2. Pathways for the repair of Top2 mediated DNA damage. Following recognition of Top2 covalent complexes (perhaps by collision with replication forks as shown in Figure 1), repair can be initiated either by proteolysis or by nucleolytic processing. Proteolysis will not completely remove the protein, since the phosphotyrosyl linkage to DNA cannot be removed by proteases. Therefore, after proteolysis, a nucleolytic processing step is still required. The product of nucleolytic processing is a DNA molecule containing a double-strand break.
Figure 2 shows a conceptual model for repairing Top2 mediated damage; similar models can be proposed for the repair of Top1 damage. Trapped covalent complexes are recognized by (unknown) proteins, leading to initial
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processing. The first step may be proteolysis of the bound protein, or nucleolytic removal of the protein, and we suggest that there may separate pathways depending on proteolysis or nucleolytic removal. Proteolysis is insufficient to completely remove the protein, so a nucleolytic processing step is required after proteolysis. The advantage of proteolytic pathways is likely the more efficient removal of a small peptide bound to DNA compared to a large protein. For example, Top2 generates a footprint on DNA of about 30 nucleotides, which may substantially hinder access of nucleolytic repair activities. The product of complete removal of Top2 is a DNA containing a double-strand break. Pathways such as homologous recombination or nonhomologous end-joining would then process this broken DNA. An interesting speculation is that specific processing pathways may preferentially use specific double-strand break repair pathways. For example, a proteolysis-dependent pathway may be preferentially commit substrates to homologous recombination.
5
FUTURE PROSPECTS FOR UNDERSTANDING THE ACTION OF DRUGS TARGETING TOPOISOMERASES USING YEAST
Drugs targeting DNA topoisomerases are highly active against a variety of tumors. This provides a strong incentive for continuing investigations into the mechanisms of drug action. As this review highlights, most of the critical questions regarding the action of topoisomerase-targeting drugs have not yet been answered. The following questions continue to fascinate us, and it is likely that yeast will continue to be an important tool in answering the questions. First, how does the binding of a topoisomerase targeting drug lead to trapping of covalent complexes? Can this information be used to rationally design new topoisomerase targeting drugs with improved spectra of activity? Clearly, these questions will require many different approaches, and the tools described in section 3 will certainly play a role Second, what are the different ways that topoisomerase-mediated damage can be repaired? For topoisomerase II, the repair pathways are particularly critical, since treatment of patients with drugs such as etoposide can lead to oncogenic translocations. Presumably, topoisomerase-mediated translocations arise from one or more repair pathways. Knowledge of the different repair pathways may indicate which repair pathways are error-prone (i.e., prone to generating interchromosomal translocations). This may lead to approaches that minimize the translocations without compromising clinical efficacy.
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Finally, there are a broad range of specific biochemical and pharmacological questions about currently used topoisomerase targeting agents, and new experimental drugs. The clear ability of yeast systems to answer questions not accessible with other experimental systems will hopefully make it easier to bring novel, safe therapies to a clinical setting more rapidly and efficiently.
ACKNOWLEDGMENTS Work in our laboratory is supported by grants CA 52814 and CA82313 from the National Cancer Institute, and by the American Lebanese Syrian Associate Charities (ALSAC).
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INDEX ATPases 5, 114, 142–144, 152, 157, 158, 162, 163, 165, 318, 395, 399 AtMRPs 302 Atr 18, 33, 34, 53–55, 59, 182, 187, 351 Atr/Atrip 53 Atrip 33, 53, 182 acquisition 233, adozelesin 17 alkylative 330 aminoacylated 87 angio-genicinhibitors 376 ansamycins 157–159, 165 antifolates 327, 333 aquated 394 ars 2 aureobasidin 195, 197 autophosphorylated 322 aza 332
bisubstrate 103 bleomycins 319, 327, 331 CaaX 101–112, 114, 115 Caenorhabditis 155, 179, 196 Cdk 3, 4, 6–8, 17, 38–40, 42–44, 46–49, 55–57, 92, 142, 160–161, 165, 301, 321, 341 CHIP 142, 161, 239–241, 260, 375, 386 CKI’s 44, 43 Claspin 18 Clbs 44–46, 49, 55, 56, 61, 76, 88, 89, 91 Clns 44 CsA 348, 349, 387, 388 Cter 235, 236, 238, 240, 248, 255, 262, 263, 287 camptothecin 332, 333, 335, 410–412, 414–420 carboxylmethylation 101 caspase 45 cDNAs 130, 184, 214, 215, 220, 221, 223, 244, 245, 247, 248, 249, 252, 255, 264, 265, 286, 383, 384, 386, 395, 415 cdc 4, 6–10, 14, 15, 17–19, 33, 39 40, 41, 43–46, 49, 50, 52, 55, 60 61, 62, 76, 88–92, 102, 104, 106 132, 142, 146, 149, 159, 160–162, 164, 165, 179–184, 199, 200, 321, 322, 361, 420
bax 242, 250, 252, 253, 259, 266, 267 benzimidazoles 58, 321, 326, 328 benzoquinoid 157, 158 bHLH 146 bilayer 296 bioreductive 337 biosynthesis 102, 109, 192–198, 200, 201, 203, 246, 299, 303, 354 bis-trihalocarbinol 329
429
430 centromeres 1, 14, 20, 42, 248 centromeric 2, 10, 245, 264, 266 ceramidases 195, 198 cerulenenin 293 chemotypes 322, 323 chromatinized 240, 256, 260 cisplatin 319, 331, 393–404 cMOAT 395, 399, 403 cohesins 14, 20, 42, 45 counterselection 212, 213, 215, 246 cross-linking 330, 338 cyclin 3, 6, 7, 38–40, 42–46, 48, 49, 55, 57, 76, 88–92, 161, 165, 180, 316, 321, 323, 329, 330, 341 cyclohexanone 329 cyclophilin 151, 153, 154, 162, 387, 388 cyclosome 44, 55, 89 DnaJ 107–109, 158 DnaK 158 DSBs 13, 186, 187, 257, 315, 319, 320, 322, 326, 327, 332, 333, 337, 411, 413, 414, 418, 421, 422 dideoxynucleotides 241 dihydroceramidase 198 dihydroceramide 195 dimer 5, 144, 154, 155, 160, 234, 237, 239, 259, 358, 382–384, 399, 413 dissociators 211–218, 220, 221, 223–226, 256 downregulated 18, 20, 90, 92, 257 downregulation 199 dTOR 363 Euplotes 128, 129 Exportin 79–81, 87 ellipticine 336, 339 enzymology 9 epipodophyllotoxin 332, 410
Index etoposide 201, 316, 410, 412, 415, 417, 419, 422 exocyst 107 FKBPs 142, 144, 147, 151, 153, 154, 161–163, 347–350, 362, 365, 383–388 FOA 213–217, 246, 251, 256, 257, 265 Fraumeni 19, 235 farnesylated 103, 105–110, 115, 116 farnesylation 101–109, 115–116 farnesyltransferase 101–105, 110, 115 forkhead 356 fumonisins 195, 196 GeneChip 241, 249 GGTase 102–105 Gleevec 341 GPCRs 203 GTPase 46, 60, 80–84, 109, 110, 361, 363 geldanamycin 143, 148, 164 gemciytabine 201 geminin 4 geranylgeranylation 102 geranylgeranyltransferase 102 glutamylcysteine 396, 399, 403 glutamylcysteinyl 396 glycerophospholipid 197 hyperrecombination 19 helicase 5, 9, 11, 33, 87, 318, 320 hematopoetic 125 heterodimers 192, 195, 298, 318, 319 hexadecanal 194 hnRNPs 76, 87, 88 homodimeric 300, 411 hTERT 129–132, 149 hus 183
Index hydroxyurea 8, 10, 182, 183, 317, 331 hyperacetylation 256 hyperphosphorylated 56, 60, 90 hyperrecombinant 11 imatinib 341 immunophilins 147, 151, 153, 154, 156, 161, 164, 387 immunosupressant 192 importin/karyopherin 78, 80, 81 importins 78, 79, 81–83, 87, 89–91, 93, 357 inositolphosphoceramides 192, 197, 198 interactors 257 intercalaters 333 intercalators 327 intergenic 2 initiation 3, 4, 6–9, 14–16, 18, 33, 34, 39, 41, 47, 233, 352–354 intrastrand 394, 396 intronic 239 KRPs 362 Kss/Fus 105 karyopherins 78–83, 86, 87 kinases 3, 4, 6–8, 10, 11, 14, 17–19, 33, 34, 38–40, 43, 44, 46, 47, 51–55, 57–64, 77, 78, 89–91, 105, 106, 142, 145, 147–149, 152, 159–162, 164, 165, 182, 184, 185, 194–197, 199, 203, 235, 236, 238, 240, 244, 258, 321–323, 341, 347, 351–354, 358–366, 382, 389, 399, 400, 402 kinetochores 13, 42, 60, 61, 321 LacZ 152, 309, 353, 383 LexA 152, 383, 386 ligase 9, 12, 18, 44, 89, 183, 238 lysophospholipids 295
431 mAMSA 410, 412, 415, 419, 420 Microsatellite 316, 318 MRP 289, 292–296, 300, 302 manumycin 105 mcm 4–6, 9, 12, 15–19, 33 mec 51, 52 metabolomics 304 metastatic 20 microarray 299, 376, 386–388 microdomains 201 microsequence 128 minichromosome 4 misexpression 91 mislocalization 91, 110, 219 mislocalized 220 mispaired 396, 397 mTOR 350, 351, 359–362, 364, 365, 366 mulitdrug 114, 289, 340, 395, 399, 409, 410 multicellular 179, 181, 243 multichaperone 156 mutator 12 myriocin 192, 195 Nibrin 52, 53 nitroaromatic 331, 332 nitrosoguanidine 183, 318 noncomplementation 350 nucleoporins 76, 77, 81, 85, 86 overexpression 18, 58, 114, 145, 149, 153, 154, 159, 160, 186, 212, 219, 221, 245, 263, 287, 291, 293, 295, 301, 321, 342, 361, 395, 396, 398, 400, 401, 404, 412 oncogene 17, 19, 78, 106, 161, 165, 199, 212, 233 orthologue 45, 53, 55, 56, 63, 165, 289 ovalicin 376–378 overexpress 321, 341
432 p53 75, 126, 142, 145, 146, 148, 157, 165, 211–226, 233–268, 284–288, 379, 403 PDREs 297, 298, 300 Pdr 289, 291–301, 304, 340 Pedicin 328 PolII 355 Polymorhphism 286, 295 PxxP 236 palmitaldehyde 194 palmitoyltranferase 192 pastoris 291, 303 peptidomimetic 103, 104 perfetto 264, 265 phosphorylation 3, 4, 6–8, 11, 17, 20, 33, 40, 41, 43, 46, 48, 51–57, 60, 61, 63, 78, 89–93, 147, 157, 165, 180, 184, 185, 194, 201, 212, 218, 236, 238, 300, 351–354, 359, 360, 382 phytoceramidase 198 phytoceramide 195, 198 polymerase, DNA 7–9, 12, 14, 15, 19, 47–49, 59, 125, 155, 184, 185, 341 premeiotic 15, 16 prenyltransferases 102 preRC 3–5, 7, 8, 18 prereplicative 3, 33 primase 9, 54, 59 processivity 9, 184 proteasome 89, 90, 147, 238, 321 Rab 103 Rac 103 RAD52 51, 128, 129, 316, 320, 322, 332–339 Raf 112, 113, 148, 160, 162 Ral 107 RanGAP 80, 82–84 RanGDP 82–84, 86, 87, 90 RanGEF 80, 82, 84 RanGTP 82–87, 90 Rapamycin 347–366
Index RecQ 11, 318, 320 RFC 8, 9, 14, 53, 54, 57, 59, 182 Rheb 103, 107, 109, 110–116, 352, 363, 364 RhoA 102 Rnr 50, 51, 56, 58 Rothmund 18, 19, 320 Rpa 8, 9, 18, 54, 59, 319, 320 rad 10, 19, 49–51, 64, 182, 183, 184, 185, 186 rDNA 10, 379 recombinogenic 124 reductase 10, 15, 50, 56, 193, 329, 330, 386 reductional 15 reinitiation 8, 47, 62 relocalized 89 replicative 5, 233 replication 7, 33 reveromycin 292 rhbA 364 rheostatable 245, 253, 259, 261 ribonucleotide 10, 15, 50, 56, 185, 329, 330 rRNAs 86, 87, 355 Schizosaccharomyces pombe 1–20, 33, 39, 43, 52, 53, 56, 64, 76, 80, 88, 91, 92, 103, 104, 108–116, 154, 155, 157, 165, 179, 181, 182, 184, 186, 195, 215, 257, 350 ScRheb 111, 113–115 SpRas 109 SpRheb 111, 113–115 securins 19, 42, 45, 46, 49, 56, 57 separase 42, 45, 49 separin 56 sevenless 160 signaling 46, 53–55, 60, 63, 64, 92, 141, 145–149, 151–153, 155, 158–160, 181, 185, 186, 191, 199, 201, 203, 241–243, 299,
Index 349, 351, 352, 353, 354–365, 382, 389, 397, 417 snRNPs 87 sphingomyelinases 195, 198 spingholipid 299 splitomicin 379, 380 sporulate 15 staurosporine 293 subcomplexes 5 subtelomeric 359 sumolation 236, 238, supertrans 253, 259, 268 synaptobrevin 108 synthase 195–198 syringomycin 193, 195, 351–366 Tor 113, 347, 348, 350, 351, 352, 353, 354, 355, 356, 357, 358, 359, 360, 361, 362, 363, 364, 365, 366 TPRs 77, 142, 144, 147, 151–157, 161, 164 TRanscription/EXport 88 Transportin 80, 395, 399 Trrap 351 taxanes 328 telangiectasia 18, 19, 53, 186, 351 tetramerization 220, 222, 235, 236, 255, 262, 263, 287 tetratricopeptide 144 thialysine 107, 113, 114 topoisomerase 315, 332–334, 337, 409–414, 416–418, 420–423 topoisomerase I 327, 410–416 topoisomerase II 327, 410, 412, 413, 415, 417, 422 tracrolimus 348 transactivate 249, 250, 263 transactivational 158 transactivators 356, 358 transactive 250 transcriptional 50, 56, 58, 87, 92, 127, 153, 155, 202, 217, 218,
433 220, 222, 223, 224, 239, 240, 244, 250, 256, 261, 284, 286, 290, 297, 298, 354, 355, 361, 365, 366, 376, 379–383 triapine 329 tribrid 382 transactivation 213, 214, 217, 218, 222, 234–237, 239–242, 244–253, 255–268, 284–287, 382, 386 tumourigenesis 20, 124, 127, 129, 130, 133, 141, 145, 155, 165 tumourigenic 123, 130, 131, 148, 165, 166 tumours 7, 17–20, 63, 75, 82, 88, 91, 103, 105, 113, 124, 127, 130, 131, 148, 155, 161, 165, 187, 203, 211, 221, 225, 226, 233–238, 240–245, 247–254, 257, 259, 260, 262–264, 266– 268, 290, 315, 316, 320, 322, 323, 329, 338, 340, 341, 363– 365, 379, 396, 404, 410, 422 Ub 142 ubiquitin 4, 43–45, 48, 55, 57, 89, 201, 212, 214–217, 238, 319, 420 ubiquitinate 45 ubiquitinated 321 ubiquitination (Ubi) 41, 45, 46, 49, 61, 108, 236, 418 ubiquitylation 202 Xenopus 4, 6, 9, 38, 43, 44, 48, 54, 91, 110, 162, 179, 181 XPA 260, 319 XPC 260 xeroderma 18, 19, 316, 319 YACs 124 zebrafish 110
