Telomeres and Telomerase in Cancer [1 ed.] 1603273069, 9781603273060, 9781603278799

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Telomeres and Telomerase in Cancer

Keiko Hiyama Editor

Telomeres and Telomerase in Cancer

Editor Keiko Hiyama Hiroshima University Dept. Translational Cancer Research 1-2-3 Kasumi Hiroshima Minami-Ku 734-8551 Japan [email protected]

ISBN 978-1-60327-306-0 e-ISBN 978-1-60327-879-9 DOI: 10.1007/978-1-60327-879-9 Library of Congress Control Number: 2008940944 # Humana Press, a part of Springer Science + Business Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science + Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Cover design was arranged from a fluorescence in situ hibrydization (FISH) photo of a pancreatic tumor with intratumoral heterogeneity in telomere lenghts and an illustration provided by Dr. Y. Hashimoto, Dr. E. Hiyama, and Ms. Y. Hiyama. Printed on acid-free paper. springer.com

Preface

Telomerase, an enzyme that elongates telomeres and endows eukaryotic cells with immortality, was first discovered in Tetrahymena in 1985 and studied among basic scientists in the 1980s. In the 1990s, it was proven that this enzyme also plays a key role in the development of human cancers and many clinical researchers became involved in this field, and in the twenty-first century, telomeres and telomerase are becoming key factors in “stem cell” research including cancer stem cells, regenerative medicine, and congenital diseases with “stem cell dysfunction.” Since telomeres/telomerase biology on ciliates, yeasts, and model mice were studied ahead of humans by basic researchers, existing monographs on telomeres and telomerase have devoted much space to biology in such well-studied species, fundamentally important but somewhat different from humans. They are very informative but sometimes confusing for clinical doctors. Now clinical trials and molecular diagnosis targeting telomeres and telomerase in cancer have been started, and all medical oncologists and medical students are required to have knowledge of telomeres and telomerase biology in humans. So, this book focuses on the telomeres and telomerase in human cancers and may provide a basic understanding of up-todate topics of these unique and fascinating molecules. I have been enamored with the scientific mystery of telomeres and telomerase along with my husband Eiso since 1990, and been supported by Dr. Jerry W. Shay and my colleagues and friends, many of them kindly contributed to this book as chapter authors. Our study has been encouraged by the Radiation Effects Research Foundation, Hiroshima University Graduate School of Biomedical Sciences, and Hiroshima University 21st Century COE Program-Radiation Casualty Medical Research Center. I would like to express my sincere gratitude to all contributors in this book, Ms. Rachel R. Warren and Mr. Michael Taylor for editorial support, and Dr. Mieczyslaw A. Piatyszek and all my colleagues, friends, and staff for their valuable suggestions and assistance. This book is dedicated with gratitude to my family, for their love and encouragement. Hiroshima Japan

Keiko Hiyama

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Contents

Part I: Basic Background 1

Telomeres and Telomerase in Humans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Keiko Hiyama, Eiso Hiyama, and Jerry W. Shay

2

Telomere-Binding Proteins in Humans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 Nadya Dimitrova

3

Regulation of Telomerase Through Transcriptional and Posttranslational Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 Amy N. Depcrynski, Patrick C. Sachs, Lynne W. Elmore, and Shawn E. Holt

4

Telomere Dysfunction and the DNA Damage Response . . . . . . . . . . . . . . . . . 87 Malissa C. Diehl, Lynne W. Elmore, and Shawn E. Holt

5

Alternative Lengthening of Telomeres in Human Cells . . . . . . . . . . . . . . . 127 Hilda A. Pickett and Roger R. Reddel

6

Mouse Model: Telomeres and Telomerase in Stem Cell and Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149 Xin Huang and Zhenyu Ju

Part II: Telomeres and Telomerase in Human Cancers 7

Role of Telomeres and Telomerase in Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . 171 Keiko Hiyama, Eiso Hiyama, Keiji Tanimoto, and Masahiko Nishiyama

8

Diagnostic Value I: Solid Tumors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 Eiso Hiyama and Keiko Hiyama

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Contents

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Diagnostic Value II: Hematopoietic Malignancies . . . . . . . . . . . . . . . . . . . . 211 Junko H. Ohyashiki and Kazuma Ohyashiki

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Therapeutic Targets and Drugs I: Telomerase and Telomerase Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225 Brittney-Shea Herbert and Erin M. Goldblatt

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Therapeutic Targets and Drugs II: G-Quadruplex and G-Quadruplex Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . 251 Chandanamali Punchihewa and Danzhou Yang

12

Therapeutic Targets and Drugs III: Tankyrase 1, Telomere-Binding Proteins, and Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 281 Hiroyuki Seimiya and Takashi Tsuruo

13

Therapeutic Targets and Drugs IV: Telomerase-Specific Gene and Vector-Based Therapies for Human Cancer . . . . . . . . . . . . . . . . . . . . . 293 Toshiyoshi Fujiwara, Yasuo Urata, and Noriaki Tanaka

Part III: Experimental Protocols 14

Protocol I: Telomerase Activity and Telomerase Expression . . . . . . . . 315 Eiso Hiyama

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Protocol II: Importance and Methods of Telomere G-Tail Length Quantification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 337 Akira Shimamoto, Eriko Aoki, Angie M. Sera, and Hidetoshi Tahara

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Protocol III: Detection of Alternative Lengthening of Telomeres . . . 351 Wei-Qin Jiang, Jeremy D. Henson, Axel A. Neumann, and Roger R. Reddel

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

Contributors

Eriko Aoki Department of Cellular and Molecular Biology, Division of Integrated Medical Science, Program for Biomedical Research, Graduate School of Biomedical Sciences, Hiroshima University, Hiroshima, Japan Amy N. Depcrynski Department of Human Genetics, Medical College of Virginia at Virginia Commonwealth University, Richmond, VA, USA Malissa C. Diehl Department of Human Genetics, Medical College of Virginia at Virginia Commonwealth University, Richmond, VA, USA Nadya Dimitrova The Rockefeller University, New York, NY, USA Lynne W. Elmore Department of Pathology and Massey Cancer Center, Medical College of Virginia at Virginia Commonwealth University, Richmond, VA, USA Toshiyoshi Fujiwara Center for Gene and Cell Therapy, Okayama University Hospital, Okayama, Japan Division of Surgical Oncology, Department of Surgery, Okayama University Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama, Japan Erin M. Goldblatt Department of Medical and Molecular Genetics, Indiana University Melvin and Bren Simon Cancer Center, Indiana University School of Medicine, Indianapolis, IN, USA Jeremy D. Henson Children’s Medical Research Institute and University of Sydney, Sydney, NSW, Australia

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Contributors

Brittney-Shea Herbert Department of Medical and Molecular Genetics, Indiana University Melvin and Bren Simon Cancer Center, Indiana University School of Medicine, Indianapolis, IN, USA Eiso Hiyama Natural Science Center for Basic Research and Development, Hiroshima University, Hiroshima, Japan Keiko Hiyama Department of Translational Cancer Research, Research Institute for Radiation Biology and Medicine, Hiroshima University, Hiroshima, Japan Shawn E. Holt Department of Human Genetics, Department of Pathology, Department of Pharmacology and Toxicology, Massey Cancer Center, Medical College of Virginia at Virginia Commonwealth University, Richmond, VA, USA Xin Huang Institute of Laboratory Animal Sciences, Max Planck Partner Group Program on Stem Cell and Aging, Chinese Academy of Medical Sciences, Beijing, China Wei-Qin Jiang Children’s Medical Research Institute and University of Sydney, Sydney, NSW, Australia Zhenyu Ju Institute of Laboratory Animal Sciences, Max Planck Partner Group Program on Stem Cell and Aging, Chinese Academy of Medical Sciences, Beijing, China Axel A. Neumann Children’s Medical Research Institute and University of Sydney, Sydney, NSW, Australia Masahiko Nishiyama Saitama Medical University International Medical Center, Saitama, Japan Junko H. Ohyashiki Intractable Diseases Research Center, Tokyo Medical University, Tokyo, Japan Kazuma Ohyashiki First Department of Internal Medicine, Tokyo Medical University, Tokyo, Japan Hilda A. Pickett Children’s Medical Research Institute and University of Sydney, Sydney, NSW, Australia

Contributors

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Chandanamali Punchihewa College of Pharmacy, The University of Arizona, Tucson, AZ, USA Roger R. Reddel Children’s Medical Research Institute and University of Sydney, Sydney, NSW, Australia Patrick C. Sachs Department of Human Genetics, Medical College of Virginia at Virginia Commonwealth University, Richmond, VA, USA Hiroyuki Seimiya Division of Molecular Biotherapy, Cancer Chemotherapy Center, Japanese Foundation for Cancer Research, Tokyo, Japan Angie M. Sera Department of Cellular and Molecular Biology, Division of Integrated Medical Science, Program for Biomedical Research, Graduate School of Biomedical Sciences, Hiroshima University, Hiroshima, Japan Jerry W. Shay Department of Cell Biology, University of Texas Southwestern Medical Center at Dallas, Dallas, TX, USA Akira Shimamoto Department of Cellular and Molecular Biology, Division of Integrated Medical Science, Program for Biomedical Research, Graduate School of Biomedical Sciences, Hiroshima University, Hiroshima, Japan Hidetoshi Tahara Department of Cellular and Molecular Biology, Division of Integrated Medical Science, Program for Biomedical Research, Graduate School of Biomedical Sciences, Hiroshima University, Hiroshima, Japan Noriaki Tanaka Division of Surgical Oncology, Department of Surgery, Okayama University Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama, Japan Keiji Tanimoto Department of Translational Cancer Research, Research Institute for Radiation Biology and Medicine, Hiroshima University, Hiroshima, Japan Takashi Tsuruo Cancer Chemotherapy Center, Japanese Foundation for Cancer Research, Tokyo, Japan

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Yasuo Urata Oncolys BioPharma, Inc., Tokyo, Japan Danzhou Yang College of Pharmacy, The University of Arizona, Tucson, AZ, USA and Arizona Cancer Center, Tucson, AZ, USA and BIO5 Institute, The University of Arizona, Tucson, AZ, USA

Contributors

Chapter 1

Telomeres and Telomerase in Humans Keiko Hiyama, Eiso Hiyama, and Jerry W. Shay

Abstract Telomerase can compensate for telomere shortening and helps prevent cellular senescence in eukaryotic cells. In humans, only specific germline cells and the vast majority of cancer cells have sufficient activity for indefinite proliferation. Lymphocytes and stem/progenitor cells in self-renewal tissues have weak activity for extension of their lifespan, but they still undergo replicative senescence. In contrast, most somatic cells do not have telomerase activity and display a finite replicative lifespan. Heterozygous mutations in either of principal telomerase components, TERT or TERC, cause telomere dysfunction and unexpectedly early senescence to stem cells of renewal tissues. Thus, restoration of telomere function in regenerative medicine via telomerase expression and inhibition of telomerase as an anticancer strategy is a double-edged sword of telomeres and telomerase in clinical medicine. Keywords: Telomere, Telomerase, Germline cell, Cancer cell, Stem cell, Cellular immortalization, Telomere dysfunction, End-replication problem, Mortality stage, Hayflick limit, TERT, TERC.

1.1

Introduction

Somatic cells explanted into tissue culture do not divide indefinitely (1) because of lack or low levels of telomerase and by progressive telomere shortening each time a cell divides. In contrast, some cells, such as male germline cells, have a greatly extended capacity to divide because of expression of the ribonucleoprotein enzyme telomerase, the sole cellular enzyme that can elongates telomeres (Fig. 1.1). DNA sequences of human daughter cells are not completely identical with those of their parent cell: During DNA synthesis prior to cell division, both ends of each chromosome, ‘‘telomeres’’, are not replicated completely because of the ‘‘end-replication problem’’ (2, 3), oxidative damage, and other poorly defined end processing events. K. Hiyama(*) Department of Translational Cancer Research, Research Institute for Radiation Biology and Medicine, Hiroshima University, 1-2-3 Kasumi, Minami-ku, Hiroshima, 734-8551 Japan, e-mail: [email protected]

K. Hiyama (ed.), Telomeres and Telomerase in Cancer. DOI: 10.1007/978-1-60327-879-9_1, # Humana Press, a part of Springer Science + Business Media, LLC 2009 3

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Germline cell (telomerase ++)

Somatic cell (telomerase − / + )

senescence apoptosis

Immortal cancer cells (telomerase activation or ALT)

Fig. 1.1 Telomere, telomerase, and cellular lifespan. Telomerase, or much less commonly alternative lengthening telomeres (ALT) mechanism, can compensate telomere shortening and prevent cellular senescence

This progressive telomere shortening is the cellular fate of eukaryotes that have linear chromosomes. The research on telomeres and telomerase was started by a small group of basic researchers in the 1980s, but many clinical researchers and pathologists came in this field after development of the telomerase detection ‘‘TRAP’’ assay in 1994, which enabled scientists to detect telomerase activity in clinical materials. Now we are in the era when the biology of telomeres and telomerase are required knowledge for clinicians, especially for medical oncologists. To encourage newcomers in the field who are not familiar with the milestone discoveries on telomeres and telomerase, we list them in Table 1.1, especially focusing on human research. In addition, we provide a brief overview of some of the key historical findings. We apologize for any major contributions omitted from this table.

1.2

Telomere Structure

Both ends of all chromosomes, ‘‘telomeres’’, end with G-rich repeats in 50 –30 strand in every eukaryotes (4, 8, 10). Every vertebrate has (TTAGGG)n repeats, while other species have different G-rich sequences, e.g., Tetrahymena has (TTGGGG)n and Schizosaccharomyces pombe has GGTTAC(A)(C)(G0-6). In humans, (TTAGGG)n repeats are about 15–20 kb in length at birth and about 810 kb in adults, but the length varies among individuals, organs, cells, and even among chromosomes. The extreme end of each telomere is not blunt (Chap. 15): the 30 single-strand overhang is about 200 nucleotides and loops back with some of the double-stranded telomeric DNA to make a telomere loop called ‘‘T loop’’ (66), so

1 Telomeres and Telomerase in Humans

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Table 1.1 Milestones of telomeres and telomerase research focusing on human 1961.12. Hayflick L and Moorhead PS (1) proposed a limitation of replicative lifespan of human diploid cells 1971.12. Olovnikov AM (2); 1972.10. Watson JD (3) proposed ‘‘End-replication problem’’ hypothesis 1978.3. Blackburn EH and Gall JG (4) identified telomeric repeats in Tetrahymena 1981.5. Klobutcher LA et al. (5) found 30 single-stranded overhang of the G-rich strand in ciliates 1984.2. Ide T et al. (6) showed lifespan elongation by SV40 mediated transformation in normal human diploid cells 1985.12. Greider C and Blackburn EH (7) identified telomerase activity in Tetrahymena 1986. Cooke HJ and Smith BA (8) showed longer telomeres in germ cells than in somatic cells 1988.8. Pereira-Smith OM and Smith JR (9) proposed 4 genes that regulate cellular immortalization of human cells 1988.9. Moyzis RK et al. (10) determined human telomeric repeat sequences as ‘‘TTAGGG’’ 1989.1. Greider C and Blackburn EH (11) cloned telomerase RNA component in Tetrahymena 1989.6. Allshire RC et al. (12) identified 3 types of repeat at subtelomeres and found TTAGGG repeats longer in sperm than in blood 1989.7. Wright WE et al. (13) proposed ‘‘two-stage model’’ for the escape from human cellular senescence 1989.11. Morin GB (14) identified telomerase activity in human cells (HeLa) 1990.2. de Lange T et al. (15) demonstrated structure of human chromosome ends and telomere shortening in tumors 1990.5. Harley CB et al. (16) demonstrated shortening of telomeres during ageing in cultured human fibroblasts 1990.8. Hastie ND et al. (17) found shortening of telomeres in colorectal cancers and with aging 1991.4. Zahler AM et al. (18) found that telomeric G-quartet structure is a negative regulator of elongation by telomerase in Oxytricha 1991.11. Harley CB (19) proposed ‘‘Telomere hypothesis’’ as mitotic clock 1992.2. Hiyama E et al. (20) proposed clinical association of telomere length in neuroblastoma 1992.5. Counter CM et al. (21) demonstrated experimental evidence of ‘‘Telomere hypothesis’’ 1994.7. Shirotani Y et al. (22) proposed clinical association of telomere length in lung cancer 1994.12. Kim NW et al. (23) developed ‘‘TRAP assay’’ and demonstrated telomerase activity in all cancer cell lines and 90% of cancerous tissues examined as well as the first evidence for the alternative lengthening of telomeres (ALT) pathway 1995.3. Hiyama E et al. (24) proposed association of telomerase activity with pathogenesis and prognosis of neuroblastoma 1995.3. Piatyszek MA et al. (25) showed telomerase activity in peripheral blood mononuclear cells 1995. 5. Counter CM et al. (26) showed upregulation of telomerase activity in leukemia cells 1995.6. Hiyama K et al. (27) proposed a clonal selection model of telomerase positive cancer cells in lung cancer development 1995.6. Collins K et al. (28) cloned telomerase protein components ‘‘p80’’ and ‘‘p95’’ in Tetrahymena 1995.6. Chadeneau C et al. (29) found telomerase activity in colorectal carcinoma but not in adenomatous polyps 1995.7. Tahara H et al. (30) showed telomerase activity in hepatitis and cirrhotic tissues in addition to hepatocellular carcinomas 1995.8. Hiyama E et al. (31) found associations of telomerase activity with stage, prognosis, telomere length alteration, and aneuploidy in gastric cancer 1995.9. Bryan TM et al. (32) identified alternative lengthening of telomeres (ALT) in human cultured immortal cells (continued)

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Table 1.1 (continued) 1995. 9. Feng J et al. (33) cloned human telomerase RNA component’’TERC (hTR)’’ 1995. 9. Lingner J et al. (34) proposed ‘‘leading strand problem’’ instead of ‘‘ragging strand problem’’ as ‘‘end-replication problem’’ 1995.10. Ohmura H et al. (35); 1998.10. Tanaka H, et al. (36) proposed existence of telomerase repressor gene in human chromosome 3 1995.10. Hiyama K et al. (37) identified activation of telomerase upon proliferation in normal human lymphocytes and hematopoietic progenitor cells 1995.11. Langford LA et al. (38) found telomerase activity in a distinct subgroup of brain tumors and reported telomerase correlated with the stage of disease 1995.12. Sharma HW et al. (39) showed downregulation of telomerase activity upon differentiation of immortal leukemia cells 1995.12. Chong L et al. (40) cloned human telomere binding protein ‘‘hTRF1’’ 1996.1. Hiyama E et al. (41) proposed a diagnostic usefulness of detecting telomerase activity in cytologic specimens of breast cancer 1996.1. Holt SE et al. (42) found that telomerase is active throughout the cell cycle but repressed in G0 1996.4. Taylor RS et al. (43) detected telomerase activity in normal epidermis and inflammatory skin lesions 1996.9. Hiyama E et al. (44) detected telomerase activity in normal human intestinal crypts 1996.10. Bodnar AG et al. (45) found that increase in telomerase activity during T cell activation is transient and does not prevent telomere shortening in long-term culture 1996.11. Tatematsu K et al. (46) developed ‘‘Stretch PCR’’ for quantitative evaluation of telomerase activity 1997.2. Kyo S et al. (47) detected telomerase activity in human proliferative-phase endometrium 1997.2. Harrington L et al. (48) cloned human telomerase-associated protein ‘‘hTEP1 (TP1)’’ 1997.2. van Steensel B and de Lange T (49) identified control of telomere length by TRF1 1997.4. Sun D et al. (50) demonstrated inhibition of human telomerase by a synthetic G-quadruplex interactive compound. 1997.6. Ohyashiki K et al. (51) developed ‘‘in situ TRAP assay’’ 1997.8. Nakamura TM et al. (52); 1997.8. Meyerson M et al. (53) cloned human telomerase reverse transcriptase ‘‘hTERT (hEST2)’’ 1997.10. Blasco MA et al. (54) developed mTR/ mice and found viable up to 6th generation 1997.10. Broccoli D et al. (55); 1997.10. Bilaud T et al. (56) cloned telomere binding protein ‘‘hTRF2’’ 1997.12. Weinrich SL et al. (57) showed reconstitution of in vitro telomerase activity only by TERC (hTR) and TERT (hTRT) 1997.11. Bryan TM et al. (58) identified alternative lengthening of telomeres (ALT) in human tumors (1771) 1998.1. Bodnar AG et al. (59) showed extension of cellular life-span by expression of hTERT in normal human cells 1998.2. van Steensel B et al. (60) identified protection of human telomere end-to-end fusion by TRF2 1998.4. Lee HW et al. (61) found telomere dysfunction in highly proliferative organs in lategeneration mTR‐/‐ mice 1998.9. Ulaner GA et al. (62) found alternate splicing of hTERT 1999.1. Cong YS et al. (63); 1991.2. Takakura M et al. (64) cloned and characterized hTERT promoter region 1999.1. Morales CP et al. (65) showed immortalization without malignant transformation of normal human fibroblasts by expression of hTERT (continued)

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Table 1.1 (continued) 1999.5. Griffith JD et al. (66) found that mammalian telomeres end in a large duplex loop, ‘‘T-loop’’ 1999.7. Hahn WC et al. (67) created human tumor cells with defined genetic elements: TERT, SV40 large-T, and oncogenic H-ras 1999.9. Yeager TR et al. (68) found a novel type of PML body in ALT cells. 1999.10. Hahn WC et al. (69) showed that dominant negative form of hTERT inhibits telomerase activity and tumorigenicity of immortal cancer cells 1999.12. Kim SH et al. (70) identified TIN2 as a new regulator of telomere length. 1999.12. Herbert B et al. (71) showed that PNA and 20 -O-MeRNA oligomers reversibly inhibit telomerase activity and induce telomere shortening 1999.12. Mitchell JR et al. (72) found dysfunction of telomerase in X-linked dyskeratosis congenita with mutations in dyskerin 2000.1. Thomas M et al. (73) demonstrated elongation of bovine adrenocortical cell function with hTERT expression in experimental xenotransplantation 2000.10. Hooijberg E et al. (74) established immortal CD8 + T cell clones by hTERT expression 2000.12. Dunham MA et al. (75) demonstrated that ALT occurs by means of homologous recombination and copy switching 2001.2. Shin-ya K et al. (76); 2002. 3. Kim MY et al. (77) demonstrated the effects of telomestatin as a telomerase inhibitor 2001.6. Baur JA et al. (78) demonstrated telomere position effect in human cells 2001.7. Hemann MT et al. (79) found that telomere dysfunction is recognized at the onset of meiosis and triggers germ cell apoptosis in mice 2001.9. Vulliamy T et al. (80) found mutations in hTR in autosomal dominant DKC 2002.6. Vulliamy T et al. (81) found mutations in hTR in aplastic anemia 2002.7. Yatabe N et al. (82) demonstrated the effects of 2–5A antisense therapy directed against hTR in cervical cancer cells 2002.7. Seimiya H et al. (83) demonstrated the effects of telomerase inhibitors MST-312, -295, and -1991 in human cancer cells 2002.10. Stewart SA et al. (84) demonstrated that telomerase contributes to tumorigenesis by a telomere length-independent mechanism 2002.11. Seger YR et al. (85) transformed a normal human cell by adenovirus E1A, Ha-RasV12, and MDM2 expression without telomerase activation 2003.3. Hakin-Smith V et al. (86) found that ALT phenotype is a good prognosis indicator in glioblastoma multiform 2003.4. Zhang A et al. (87) found deletion of hTERT and haploinsufficiency of telomere maintenance in Cri du chat syndrome 2003.4. Stewart SA et al. (88) proposed that erosion of single-strand telomeric overhang, rather than overall telomere length, serves to trigger replicative senescence 2003.4. Ulaner GA et al. (89) found that telomerase activation and ALT are comparably poor prognosis indicators in osteosarcoma 2003.5. Colqin LM et al. (90) proposed that human POT1 protein can act as a telomerasedependent positive regulator of telomere length 2003.6. Loayza D and de Lange T (91) proposed that POT1 interacts with TRF1 complex and transmits information of telomere length to the telomere terminus 2003.6. Lin SY et al. (92) proposed 3 tumor suppressor pathways involved in hTERT repression: Mad1/c-Myc, SIP1, and Menin 2003.7. Masutomi K et al. (93) proposed that hTERT is expressed even in normal human somatic cells maintaining telomere structure such as 30 single-stranded overhang 2003.7. Asai A et al. (94) developed a telomerase template antagonist GRN163 and demonstrated its anticancer effects in vitro and in xenograft model (continued)

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Table 1.1 (continued) 2003.8. Tauchi T et al. (95) demonstrated effects of G-quadruplex-interactive telomerase inhibitor telomestatin (SOT-095) in leukemia cells 2004.2. der-Sarkissian H et al. (96) proposed that the chromosomes with shortest telomeres are the first to become unstable in telomerase-negative-transformed cells 2004.5. Preto A et al. (97) demonstrated that telomere erosion triggers growth arrest in thyroid cancer cells with wild p53 while it cause crisis by abrogation of p53 2005.1. Seimiya H et al. (98) demonstrated that tankyrase 1 inhibition enhances telomere shortening by telomerase inhibitor 2005.4. Fasching CL et al. (99); Marciniak RA et al. (100); 2005.11. Cerone MA, et al. (101) proposed that ALT-associated promyelocytic leukemia bodies (APBs) are not always essential for ALT-mediated telomere maintenance 2005.5. Yamaguchi H et al. (102) found mutations in TERT in aplastic anemia 2005.6. Sun B et al. (103) demonstrated a minimal set of genetic alterations required for fibroblast transformation 2005.7. Nakamura M et al. (104) demonstrated that hTERT KO by siRNAs sensitizes cervical cancer cells to ionizing radiation and chemotherapy 2005.8. Herbert BS et al. (105) demonstrated a superiority of lipid modification of GRN163 (GRN163L) in telomerase inhibition 2005.8. Zaug AJ et al. (106) found that human POT1 disrupts telomeric G-quadruplexes allowing telomerase extension 2005.8. Flores I et al. (107) found that mobilization of stem cells out of their niche was inhibited by telomere shortening and promoted by Tert overexpression in mice 2005.9. de Lange T (108) proposed a concept of ‘‘Shelterin’’ as telomere binding proteins consisting of TRF1, TRF2, POT1, TIN2, TPP1, and RAP1 2005.11. Djojosubroto MW et al. (109) demonstrated in vitro and in vivo effects of hTR antagonist GRN163 and GRN163L on hepatoma cells 2005.11. Armaninos M et al. (110); Goldman F et al. (111) demonstrated TERC haploinsufficiency on the inheritance of telomere length in autosomal dominant dyskeratosis congenital 2005.11. Verdun RE et al. (112) found that telomeres of telomerase-negative cells recruit Mre11, phosphorylated NBS1, and ATM in every G2 phase of the cell cycle 2005.12. Horikawa I et al. (113) found that a GC-box within the hTERT promoter is responsible for the human-specific TERT repression 2006.1. Anderson CJ et al. (114) found that hypoxia induces the transcriptional activity of both hTR and hTERT gene promoters and increase of active hTERT splice variant 2006.2. Compton SA et al. (115) found NOS-dependent telomere shortening and apoptosis of prostate cancer cells by inhibition of Hsp90 2006.2. Chai W et al. (116) demonstrated different overhang sizes at leading versus lagging strands of human telomeres 2006.3. Tahara H et al. (117) found that telomestatin induces loss of 30 telomeric overhang through TRF2 protein dissociation from telomeres in cancer cells 2006.3. Gellert GC et al. (118); 2006.5. Hochreiter AE et al. (119) demonstrated in vitro and in vivo effects of hTR antagonist GRN163L on breast cancer cells 2006.5. Trapp S et al. (120) demonstrated tumor-promoting effects of vTR in a chicken natural virus-host infection model 2006.5. Ambrus A et al. (121) proposed ‘‘mixed parallel/antiparallel G-strands’’ as intact telomeric G-quadruplex structure 2007.2. Xin H et al. (122); 2007.2. Wang F et al. (123) proposed POT1-TPP1 complex as a processivity factor for telomerase 2007.3. Cohen SB et al. (124) showed that human telomerase exists as a complex of two molecules each of hTERT, hTR, and dyskerin (continued)

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Table 1.1 (continued) 2007.3. Armanios MY et al. (125); 2007.5. Tsakiri KD et al (126) identified mutations in TERT/ TERC and short telomeres as etiology of familial and/or adult-onset pulmonary fibrosis 2007.7. Jiang WO et al. (127) identified 8 candidate ALT genes as PML, TRF1, TRF2, TIN2, RAP1, MRE11, RAD50, and NBS1 2007.10. Xu L and Blackburn EH (128) proposed ‘‘T-stumps’’ in immortal cancer cells as the minimal telomeric unit that can be protected by telomere binding proteins 2007.11. Azzalin CM et al. (129); 2008. 2. Schoeftner S et al. (130) identified active transcription of human telomeres into ‘‘telomeric repeat-containing RNA (TERRA or TelRNAs)’’ that regulate telomerase activity 2007.11. Takahashi K et al. (131) found that human iPS cells derived from fibroblasts activated intrinsic telomerase 2008.3. Venteicher AS et al. (132) found that additional enzymes (ATPases pontin and reptin) are required for telomerase assembly 2008.3. Stadtfeld M et al. (133) demonstrated that activation of endogenous telomerase is one of late events during fibroblast reprogramming to iPS cells in mouse

that the chromosome end is distinguished from bona fide dsDNA breaks and protected from exposure to DNA repair system (Chap. 4).

1.3

Why do Telomeres Gradually Shortened?

Until the early 1960s, cultured normal human cells were believed to be able to replicate indefinitely as long as good culture conditions were maintained. In 1961, Hayflick reversed this concept and demonstrated convincingly that normal human cells have a limit in the number of possible cell divisions: lung fibroblast 55 times, heart 26, kidney 40, and skin 43 (1). So, this phenomenon of replicative senescence in normal cells is often called the ‘‘Hayflick limit’’. In 1971 and 1972, Olovnikov and Watson reported the mechanism of this limit in Russian and English, respectively, as an ‘‘end-replication problem’’. When they originally proposed this problem, ‘‘lagging (discontinuous) strand’’ was considered to be responsible for telomere shortening, because DNA polymerase replicates only in the 50 –30 direction, and requires an RNA primer in starting DNA replication and a complementary strand for replication. Then, after removal of the RNA primer, the 50 end of the lagging strand locates inside of the extreme 30 end of the complement strand, i.e., ‘‘lagging strand problem’’ (134). However, considering the structure of 30 telomere overhang at the end of telomeres, the ‘‘end-replication problem’’ mechanism was then proposed as a ‘‘leading strand problem’’, i.e., inability of leading strand DNA synthesis to produce the 30 overhang (Fig. 1.2), and then ‘‘lagging strand problem’’ may occur in the next round of replication (34). The ‘‘end-replication problem’’ and resulting limit of cellular lifespan exists in all eukaryotes that have linear chromosomes but not in prokaryotes that have circular chromosomes without chromosomal ‘‘ends’’.

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Fig. 1.2 Renewed ‘‘end-replication problem.’’ Incomplete replication of leading strand due to formation of 30 overhang, as well as that of lagging strand due to RNA primer as a prerequisite for DNA synthesis, is responsible for telomere shortening

1.4

Functions of Telomeres and Telomerase

Telomeres consist of noncoding TTAGGG repeats (but the reader should be aware that it was recently reported that telomeric repeats are transcribed and this telomeric RNA may regulate telomerase activity (129)). Telomeres protect chromosome ends from DNA degradation, DNA repair mechanisms, and fusion. Uncapped telomeres activate the DNA damage response and cause end-to-end fusions resulting in cellular senescence, apoptosis, and further chromosomal instability (Table 1.2). Since genes near telomeres may be reversibly silenced in a telomere lengthdependent manner by telomere position effect (TPE) (78), telomere shortening may result in restoration of expression of such silenced genes. Moreover, telomeres appear to play an important role in ‘‘bouquet’’ formation at the beginning of meiosis, and telomere dysfunction results in germ cell death (79). The well known function of telomerase is elongation of telomeres, so that cells can increase their replicative capacity, sometimes indefinitely (Table 1.3). However, it may be that even in some normal fibroblasts, telomerase is expressed at low levels. However, this amount of telomerase cannot maintain telomere length but during each S phase may play a role in maintaining chromosomal structure (93).

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Table 1.2 Function of telomeres and dysfunction due to telomere shortening Function Consequence of telomere shortening/dysfunction Prevention of erosion of genes in Cellular senescence, cell death, and/or carcinogenesis subtelomeres Telomere position effects (TPE) Reactivation of the silenced genes near telomeres Protection of chromosomal Chromosome fusion, anaphase bridge end-to-end fusion T-loop formation and Disruption of T-loop inducing p53 mediated cellular chromosome stability senescence and apoptosis, chromosome fusion ‘‘Bouquet’’ formation at the Impaired meiosis and germ cell apoptosis beginning of meiosis

Table 1.3 Function of telomerase and consequence of its activation Function Consequence examples of telomerase activation Elongation of telomeres Elongation of cellular lifespan or immortalization Maintenance of chromosomal Telomerase is transiently expressed in each S phase structure in normal cells Addition of malignant potential Tumor formation with nontumorigenic ALT cells Promotion of stem cell e.g., increased hair growth proliferation DNA repair? Required to form DNA damage foci following irradiation Self-renewal capacity? Required to reprogram fibroblasts to iPS cells

Telomerase may also have roles in stem cell proliferation (107), and reprogramming of iPS cells (131, 133). The mechanisms of these functions of telomerase may or may not be related to maintenance of telomere length.

1.5

Two Mortality Stage Mechanisms and Telomere Hypothesis

Two mortality stage mechanisms, ‘‘M1’’ and ‘‘M2’’, must be overcome for normal cells to escape from cellular senescence and become immortal (13, 19). Normal cells stop dividing at the ‘‘Hayflick limit,’’ i.e., mortality stage 1 (M1), where p16/pRb and TP53 recognize perhaps a single uncapped telomere as broken or damage DNA. To bypass this potent tumor suppressor mechanism, cells can divide beyond M1 and continue replication by inactivating these tumor suppressor genes (termed extended lifespan). However, the cells again stop dividing at the mortality stage 2 (M2), also called ‘‘crisis’’. At this stage, many telomeres are critically shortened, end-end fusions occur, and cells stop dividing. The escape from M2 in human cells is extremely rare and almost universally involves the upregulation or reactivation of telomerase as a telomere-maintenance mechanism (19). Much less commonly other telomere maintenance mechanisms such as alternative lengthening telomeres (ALT, See Chap. 5) (58) are engaged. The cells that have activated telomerase can overcome the M2, and become immortal (Fig. 1.3).

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Fig. 1.3 Telomere hypothesis and two independent mortality mechanisms controlling cellular senescence and immortalization. Normal stem cells have elongated lifespan but are not immortal. Cancer stem cells may arise from cells that have bypassed M2 as well as from normal stem cells. The origin of cancer stem cells is still an hypothesis and it is possible that they may have the same, shorter, or longer telomeres compared to the bulk population of tumor cells

Thus both M1 and M2 may be thought of as initial anti-cancer protection mechanisms and only when both have been bypassed are cells immortal and then can progress to advanced malignancies. Thus some preneoplastic lesions may be arrested at M1, some early stage clinical cancers may have overcome M1, but not M2 (transformed but mortal cancer cells), while advanced cancers probably have overcome M2 (transformed and immortal cancer cells). All cancer cell lines and ‘‘cancer stem cells’’ have likely overcome both M1 and M2. Meanwhile, normal lymphocytes and stem/progenitor cells in self-renewal tissues have highly regulated telomerase activity, but gradually senesce and thus while telomerase may partially extend their lifespan, the cells are mortal, since they have not overcome M1 nor M2 (Fig. 1.3, Table 1.2) (37, 43, 44, 47).

1.6

Telomerase is a Conserved Reverse-Transcriptase

Human telomerase is a ribonucleoprotein enzyme composed of catalytic component TERT, telomerase reverse-transcriptase (52, 53), and RNA template TERC (or hTR), telomerase RNA component (or human telomerase RNA) (33). Telomerase can elongate the G-rich 30 telomere overhang using the TERC as template. Since telomeric DNA is synthesized according to the complementary RNA sequence, telomerase is a reverse-transcriptase. The catalytic component gene TERT is evolutionally conserved (52), and has been alternatively called as hTERT (meaning

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‘‘human TERT’’), hTRT (52), hEST2 (53), hTCS1 (135), or TP2 (136) by researchers who independently cloned this gene in 1997. In vivo, Cohen SB et al. purified the catalytically active human telomerase complex (650–670 kD) and proposed that telomerase may exist as a dimer consisting of two molecules each of hTERT (127 kD), hTR (153 kD), and dyskerin (57 kD) (124). The telomerase core enzyme may complex with additional telomerase binding proteins (Chap. 2), and its function is regulated transcriptionally and posttranscriptionally (Chap. 3).

1.7

Telomerase Activity and Cellular Immortalization

Expression of TERT is a prerequisite but not always sufficient for cellular immortalization in most human cells (especially when the cell culture conditions are not optimized). Among the core components of telomerase, TERC is constitutively expressed in most cells regardless of their telomerase activity level, and TERT expression levels determine the telomerase activity qualitatively and quantitatively. Ectopic expression of TERT induces telomerase activity in telomerase-negative somatic cells in vitro (57), but cellular immortalization is still a relatively rare event with many clones expressing telomerase potentially remaining mortal with or without elongation of lifespan (59, 65). Thus, activation of telomerase is a prerequisite, except for rare ALT cells, but not necessarily a sufficient condition for cellular immortalization.

1.8

Telomerase is Activated in >80% of Human Malignancies

Every type of human malignancy examined to the present time has evidence of telomerase activation with the average being detected in >80% of overall cancer tissues (137) (Chap. 8). In general, the incidence and level of telomerase activation are higher in advanced stages than early stages, in metastatic lesions than primary lesions, in poor prognosis cases than good prognosis cases, and in malignant lesions than in precancerous lesions, indicating that continuous progression of cancers may ultimately depend on telomerase in all human malignancies. Thus, telomerase components and associated proteins are becoming not only a diagnostic marker of cancer but also the molecular targets of anticancer strategies and some of them are under clinical trials (see Part II).

1.9

Telomerase Activity in Normal Somatic Cells and Stem Cells

In most human normal somatic cells, telomerase activity is undetectable. However, lymphocytes and most, but not all, stem/progenitor cells in self-renewal tissues can express telomerase upon mitogenic stimulation (37). These cells have elongated lifespan so that humans can retain immune reactivity to each antigen and maintain

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Table 1.4 Telomere length, telomerase expression, and cellular lifespan in human cells Telomere length Telomerase expression (detectable level by usual analysis)a (cellular lifespan) +  Stable (immortal)

Germline cells, immortal cancer cells

Slowly shortened (mortal with elongated lifespan)

Part of in vitro immortalized cellsb, a few cancers (e.g. part of sarcoma)a Mesenchymal stem cells

Lymphocytes (activated), renewing stem/progenitor cells (e.g. hematopoietic, intestinal, epidermal, hair follicle), endometrial cells (proliferative) Shortened (mortal) Stem cells in progeria, stem cells Most somatic cells, part of with dysfunction (e.g. a part of cancer cells aplastic anemia, dyskeratosis congenital, IPF c) a At very low levels, telomerase is expressed during each S phase even in normal somatic cells (93) b ALT (alternative lengthening of telomeres) cells c IPF idiopathic pulmonary fibrosis

the function of important organs, such as the bone marrow, intestine, skin, etc. However, when these cells stop dividing and/or are differentiated, telomerase activity disappears. Thus, normal somatic cells never become immortal in vivo, even those with regulated telomerase activation (Table 1.4) unless there is loss of tumor suppressor genes or activation of oncogenes. In contrast, human mesenchymal stem cells (hMSCs) may have very low or no detectable telomerase activity (138–140), whereas they can maintain longer telomeres than those in usual somatic cells. hMSCs lack characteristics of ALT cells such as PML bodies and may have unique telomere maintaining mechanism (141). In the mouse, telomerase activity is detectable in normal somatic cells in addition to stem cells, and telomere length is around fivefold longer than human (50 kb vs. 10 kb). Since their lifespan is much shorter than human, telomeres in murine cells are not shortened to reach the Hayflick limit within a lifetime even without telomerase, and mTR/ (telomerase RNA knockout) mice can survive until the sixth generation (54). At this late generation, telomere dysfunction is manifested as stem cell dysfunction and infertility in mTR/ mice (Chap. 6).

1.10

Telomere-Binding Proteins

In addition to the core telomere-binding proteins ‘‘Shelterin’’ (TRF1, TRF2, POT1, TIN2, TPP1, and RAP1), DNA repair proteins are also involved in telomere maintenance: Ku complex, MRN complex (MRE11, RAD50, NBS1), XPF/ ERCC1, ATM, BLM/WRN, RAD51D, and RAD54 (108). Mutations or absence of these genes cause short telomeres, end-to-end chromosomal fusions, premature aging phenotypes, and/or cancer predisposition (Chap. 2).

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Telomere Dysfunction and Human Diseases

Mutations in telomerase component genes, TERT and TERC, cause stem cell dysfunction, resulting in dyskeratosis congenita (80), aplastic anemia (81), and idiopathic pulmonary fibrosis (125, 126). This suggests that individuals with heterozygous mutations in TERT or TERC have reduced telomerase activity in their stem cells and suffer accelerated telomere shortening possibly due to haploinsufficiency, except for mutations in the template domain of TERC, which can show dominant negative effects (142). Stem cell dysfunction due to accelerated telomere shortening is caused also by mutations in telomerase binding proteins, such as dyskerin (72). This suggests that telomerase is not in excess (e.g., individuals need long telomeres for a full lifespan). Importantly these telomere-associated genetic diseases suggest that inhibition of only 50% of telomerase in human cancer may be sufficient to drive cancer cells with short telomeres into apoptotic cell death leading to durable cancer responses prior to affecting normal stem cells.

1.12

Concluding Remarks

Telomeres and telomerase dysfunction causes unexpected early senescence to stem cells in renewal tissues or graft tissues, while maintenance of telomeres via activation of telomerase is the critical offender for the indefinite proliferation of immortal cancer cells. Restoration of telomere function in regenerative medicine and inhibition of telomerase (e.g., induction of telomere dysfunction) as an anticancer strategy, respectively, is the double-edged sword in clinical medicine.

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103. Sun B, Chen M, Hawks CL, et al. The minimal set of genetic alterations required for conversion of primary human fibroblasts to cancer cells in the subrenal capsule assay. Neoplasia 2005;7:585–93. 104. Nakamura M, Masutomi K, Kyo S, et al. Efficient inhibition of human telomerase reverse transcriptase expression by RNA interference sensitizes cancer cells to ionizing radiation and chemotherapy. Hum Gene Ther 2005;16:859–68. 105. Herbert BS, Gellert GC, Hochreiter A, et al. Lipid modification of GRN163, an N30 – > P50 thio-phosphoramidate oligonucleotide, enhances the potency of telomerase inhibition. Oncogene 2005;24:5262–8. 106. Zaug AJ, Podell ER, Cech TR. Human POT1 disrupts telomeric G-quadruplexes allowing telomerase extension in vitro. Proc Natl Acad Sci USA 2005;102:10864–9. 107. Flores I, Cayuela ML, Blasco MA. Effects of telomerase and telomere length on epidermal stem cell behavior. Science 2005;309:1253–6. 108. de Lange T. Shelterin: the protein complex that shapes and safeguards human telomeres. Genes Dev 2005;19:2100–10. 109. Djojosubroto MW, Chin AC, Go N, et al. Telomerase antagonists GRN163 and GRN163L inhibit tumor growth and increase chemosensitivity of human hepatoma. Hepatology 2005;42:1127–36. 110. Armanios M, Chen JL, Chang YP, et al. Haploinsufficiency of telomerase reverse transcriptase leads to anticipation in autosomal dominant dyskeratosis congenita. Proc Natl Acad Sci USA 2005;102:15960–4. 111. Goldman F, Bouarich R, Kulkarni S, et al. The effect of TERC haploinsufficiency on the inheritance of telomere length. Proc Natl Acad Sci USA 2005;102:17119–24. 112. Verdun RE, Crabbe L, Haggblom C, et al. Functional human telomeres are recognized as DNA damage in G2 of the cell cycle. Mol Cell 2005;20:551–61. 113. Horikawa I, Chiang YJ, Patterson T, et al. Differential cis-regulation of human versus mouse TERT gene expression in vivo: identification of a human-specific repressive element. Proc Natl Acad Sci USA 2005;102:18437–42. 114. Anderson CJ, Hoare SF, Ashcroft M, et al. Hypoxic regulation of telomerase gene expression by transcriptional and post-transcriptional mechanisms. Oncogene 2006;25:61–9. 115. Compton SA, Elmore LW, Haydu K, et al. Induction of nitric oxide synthase-dependent telomere shortening after functional inhibition of Hsp90 in human tumor cells. Mol Cell Biol 2006;26:1452–62. 116. Chai W, Du Q, Shay JW, et al. Human telomeres have different overhang sizes at leading versus lagging strands. Mol Cell 2006;21:427–35. 117. Tahara H, Shin-Ya K, Seimiya H, et al. G-Quadruplex stabilization by telomestatin induces TRF2 protein dissociation from telomeres and anaphase bridge formation accompanied by loss of the 30 telomeric overhang in cancer cells. Oncogene 2006;25:1955–66. 118. Gellert GC, Dikmen ZG, Wright WE, et al. Effects of a novel telomerase inhibitor, GRN163L, in human breast cancer. Breast Cancer Res Treat 2006;96:73–81. 119. Hochreiter AE, Xiao H, Goldblatt EM, et al. Telomerase template antagonist GRN163L disrupts telomere maintenance, tumor growth, and metastasis of breast cancer. Clin Cancer Res 2006;12:3184–92. 120. Trapp S, Parcells MS, Kamil JP, et al. A virus-encoded telomerase RNA promotes malignant T cell lymphomagenesis. J Exp Med 2006;203:1307–17. 121. Ambrus A, Chen D, Dai J, et al. Human telomeric sequence forms a hybrid-type intramolecular G-quadruplex structure with mixed parallel/antiparallel strands in potassium solution. Nucleic Acids Res 2006;34:2723–35. 122. Xin H, Liu D, Wan M, et al. TPP1 is a homologue of ciliate TEBP-beta and interacts with POT1 to recruit telomerase. Nature 2007;445:559–62. 123. Wang F, Podell ER, Zaug AJ, et al. The POT1-TPP1 telomere complex is a telomerase processivity factor. Nature 2007;445:506–10.

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124. Cohen SB, Graham ME, Lovrecz GO, et al. Protein composition of catalytically active human telomerase from immortal cells. Science 2007;315:1850–3. 125. Armanios MY, Chen JJ, Cogan JD, et al. Telomerase mutations in families with idiopathic pulmonary fibrosis. N Engl J Med 2007;356:1317–26. 126. Tsakiri KD, Cronkhite JT, Kuan PJ, et al. Adult-onset pulmonary fibrosis caused by mutations in telomerase. Proc Natl Acad Sci U S A 2007;104:7552–7. 127. Jiang WQ, Zhong ZH, Henson JD, et al. Identification of candidate alternative lengthening of telomeres genes by methionine restriction and RNA interference. Oncogene 2007;26:4635–47. 128. Xu L, Blackburn EH. Human cancer cells harbor T-stumps, a distinct class of extremely short telomeres. Mol Cell 2007;28:315–27. 129. Azzalin CM, Reichenbach P, Khoriauli L, et al. Telomeric repeat containing RNA and RNA surveillance factors at mammalian chromosome ends. Science 2007;318:798–801. 130. Schoeftner S, Blasco MA. Developmentally regulated transcription of mammalian telomeres by DNA-dependent RNA polymerase II. Nat Cell Biol 2008;10:228–36. 131. Takahashi K, Tanabe K, Ohnuki M, et al. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 2007;131:861–72. 132. Venteicher AS, Meng Z, Mason PJ, et al. Identification of ATPases pontin and reptin as telomerase components essential for holoenzyme assembly. Cell 2008;132:945–57. 133. Stadtfeld M, Maherali N, Breault DT, et al. Defining molecular cornerstones during fibroblast to iPS cell reprogramming in mouse. Cell Stem Cell 2008;2:230–40. 134. Levy MZ, Allsopp RC, Futcher AB, et al. Telomere end-replication problem and cell aging. J Mol Biol 1992;225:951–60. 135. Kilian A, Bowtell DD, Abud HE, et al. Isolation of a candidate human telomerase catalytic subunit gene, which reveals complex splicing patterns in different cell types. Hum Mol Genet 1997;6:2011–9. 136. Harrington L, Zhou W, McPhail T, et al. Human telomerase contains evolutionarily conserved catalytic and structural subunits. Genes Dev 1997;11:3109–15. 137. Shay JW, Bacchetti S. A survey of telomerase activity in human cancer. Eur J Cancer 1997;33:787–91. 138. Hiyama E, Hiyama K. Telomere and telomerase in stem cells. Br J Cancer 2007;96:1020–4. 139. Yanada S, Ochi M, Kojima K, et al. Possibility of selection of chondrogenic progenitor cells by telomere length in FGF-2-expanded mesenchymal stromal cells. Cell Prolif 2006;39:575–84. 140. Zimmermann S, Voss M, Kaiser S, et al. Lack of telomerase activity in human mesenchymal stem cells. Leukemia 2003;17:1146–9. 141. Zhao YM, Li JY, Lan JP, et al. Cell cycle dependent telomere regulation by telomerase in human bone marrow mesenchymal stem cells. Biochem Biophys Res Commun 2008;369:1114–9. 142. Garcia CK, Wright WE, Shay JW. Human diseases of telomerase dysfunction: insights into tissue aging. Nucleic Acids Res 2007;35:7406–16.

Chapter 2

Telomere-Binding Proteins in Humans Nadya Dimitrova

Abstract Shelterin, the telomere-secific protein complex, is essential for genome stability and cell viability. Shelterin accumulates at telomeres and transforms chromosome ends into specialized structures that evade recognition by the DNA damage signaling and repair machineries and are maintained through consecutive cell divisions. Shelterin accomplishes these tasks through its ability to remodel the telomeric DNA into a protected structure and to locally inhibit the activation of the DNA damage response. Furthermore, shelterin plays an essential role in controlling telomere length homeostasis by suppressing excessive nuclease activity at the chromosome terminus and by regulating telomerase. The capacities of the telomere-binding proteins to prevent genome instability and to influence telomere length make shelterin an essential factor in both normal cell growth and tumorigenesis. Keywords: Shelterin, T-loop, ATM, ATR, NHEJ, HR.

2.1

Introduction

In the 1940s, the special qualities of ‘‘natural’’ ends of linear chromosomes were first recognized. Barbara McClintock observed that in contrast to ‘‘broken’’ ends, which tended to fuse and create dicentric chromosomes, ‘‘natural’’ chromosome ends were stably maintained (1). We now know that chromosome ends are stable because they are capped by telomeres, dynamic and complex nucleoprotein machineries that protect the integrity of chromosomes and are essential for cellular survival. The telomeric DNA is composed of a long array of double-stranded TTAGGG repeats that extend into a single-stranded overhang on the G-rich strand (2, 3). The repetitive and highly defined sequence of human telomeres prompted the search for factors that bind specifically to the telomeric repeats and could give an insight into how telomeres protect chromosome integrity. TRF1 (TTAGGG-repeat binding factor 1) was the first protein to be found due to its specific association with duplex N. Dimitrova The Rockefeller University, Box 159, New York, NY 10065, e‐mail: [email protected] K. Hiyama (ed.), Telomeres and Telomerase in Cancer. DOI: 10.1007/978-1-60327-879-9_2, # Humana Press, a part of Springer Science + Business Media, LLC 2009 23

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TTAGGG repeats in HeLa cell nuclear extract (4). Since then, five more telomere binding proteins have been identified – TRF2, POT1, RAP1, TIN2, and TPP1. Together, these six factors form the shelterin complex, which coats specifically the telomeric DNA and is essential for the prevention of detrimental genome instability (5). It is thought that shelterin has the ability to remodel the telomeric DNA into a protected structure and to locally inhibit the activation of the DNA damage response machinery. Furthermore, shelterin plays an essential role in determining telomere length by suppressing excessive nuclease activity at the chromosome terminus and by regulating telomerase, the enzyme that elongates telomeres by adding TTAGGG repeats to the 30 end (reviewed in (5)) (Fig. 2.1). In addition to maintaining chromosome integrity at the cellular level, telomeres have been proposed to play an important role as a tumor suppressor mechanism that limits the replicative lifespan of human somatic cells (reviewed in (6)). The

Fig. 2.1 Overview of the multiple roles of shelterin at human telomeres. (a) Shelterin protects chromosome ends. Telomeric DNA consists of 2–30 Kb double-stranded TTAGGG repeats that extent into 50–300 nt single-stranded TTAGGG overhang on the 30 strand. Shelterin complex specifically coats both the double-stranded portion of the telomere and the single-stranded extension. The presence of shelterin at telomeres promotes the formation of a protective structure at chromosome ends and also suppresses the activation of DNA damage signaling and repair pathways. (b) Shelterin regulates telomere length. Telomere ends are subject to degradation by unknown 50 –30 nuclease(s) that resects the 50 -strand to generate the telomere overhang. Shelterin regulates the activity and/or recruitment of this nuclease and thereby prevents excessive nuclease degradation and telomere shortening. At the same time, telomerase can elongate telomeres by adding TTAGGG repeats to the 30 end, an activity that is positively and negatively regulated by shelterin

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chromosome terminus is shortened each cell division by two additive processes. First, the DNA replication machinery requires a primer to initiate 50 –30 replication. Therefore, in each round of DNA replication, as the primer is removed, the newly synthesized strand lacks the first 10–15 base pairs at the 50 end. This is known as the end-replication problem (7, 8). Second, chromosome ends are also shortened by nuclease activity that generates the telomeric overhang (3, 9). The generation of a single-stranded 30 extension is a structural requirement for the telomere-mediated protection of the chromosome terminus from further degradation. As a result, telomeres shorten at an average of 100 base pairs at each cell division (10, 11). This is a regulated process and when telomeres become critically short, cells enter a terminal growth arrest state called senescence (12). Experimentally, the limited replicative potential of human cells was first described as the Hayflick limit and came from the observation that human somatic cells could only be maintained in culture for about 50 cell divisions before they stopped dividing (13). In the context of the organism, this limit to cellular proliferation is predicted to be a powerful tumor suppressor mechanism. The evidence that exogenous introduction of telomerase could overcome that barrier and extend the replicative lifespan indefinitely, quickly put telomeres and in particular telomerase, in the spotlight of the cancer biology field (14, 15). Indeed, more than 80% of human tumors have inappropriate reactivation of telomerase and virtually all tumor cells have established telomere maintenance mechanisms that allow for unlimited proliferation (16). In contrast to telomerase, the role of shelterin in the tumor suppressor function of telomeres has been more difficult to define. Disruption of the shelterin complex impairs both its telomere length regulation and telomere protection functions, leading to acute genome instability and cell cycle arrest (reviewed in (5)). However, it is conceivable that subtle alterations of one or more of the shelterin components could deregulate the telomere length homeostasis or, alternatively, cause transient genome instability that in the absence of functional p53 and Rb pathways would accelerate carcinogenesis (17). In this chapter, each shelterin member is described individually and in the context of the shelterin complex. I will then discuss insights gained from different experimental model systems on how shelterin functions to preserve telomere integrity and regulate telomere length.

2.2

Telomere-Binding Proteins in Human Cells

2.2.1

Shelterin

2.2.1.1

TRF1 and TRF2

The shelterin complex consists of six members (5) (Fig. 2.2). The first two factors, TRF1 and TRF2, bind to the double-stranded portion of the telomere and are essential for the recruitment and stabilization of the other shelterin members

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Fig. 2.2 The shelterin complex consists of six subunits. TRF1 and TRF2 are dimers that specifically recognize and bind to double-stranded telomeric DNA with their Myb domains. The binding site for each TRF1 or TRF2 dimer can be overlapping or separate sequences as indicated. RAP1 is a TRF2-binding partner. TIN2, on the other hand, interacts with both TRF1 and TRF2 and in turn recruits TPP1 and POT1 to the double-stranded portion of the telomere. The ability of TIN2 to bind to both TRF1 and TRF2, independently or simultaneously, creates the possibility for different shelterin subcomplexes as shown. The TRF1/TIN2, TRF2/TIN2, and TRF1/TRF2/TIN2 shelterin subcomplexes could all potentially play roles in the enrichment of TPP1/POT1 at single-stranded DNA. The POT1 binding sequence at single stranded DNA can be located at an internal site or at the 30 end as indicated

(4, 18, 19). The high specificity and affinity of TRF1 and TRF2 for telomeric DNA is achieved by two complementary mechanisms. First, both proteins contain homologous carboxy-terminal DNA-binding (SANT/Myb-type) domains that recognize 50 -YTAGGGTTR-30 sequence in double-stranded DNA with high specificity (20–22). In addition, both TRF1 and TRF2 contain structurally similar dimerization (TRFH) domains and exist as homodimers in solution, with TRF2 having a propensity to form higher-order oligomers (19, 23, 24). In the case of TRF1, it has been shown that upon dimerization, the simultaneous binding of two Myb domains increases its affinity for DNA approximately tenfold (20). Both TRF1 and TRF2 are essential for cell viability, and deletion of TRF1 and TRF2 genes leads to early lethality in mouse development (25–27). 2.2.1.2

TIN2

Interestingly, TRF1 and TRF2 are bridged by another shelterin factor, TIN2. TIN2 can interact simultaneously with TRF1 and TRF2 through its central region and amino-terminal half, respectively, and in turn recruits to the telomere two other shelterin components, TPP1 and its binding partner POT1 (28–32). TIN2 plays a core role in the sheltering complex. First, TIN2 protects TRF1 from the factor tankyrase 1, which has the ability to PARsylate TRF1 in human cells, a modification that strongly reduces its binding to DNA (33, 34). Second, TIN2 promotes the stable association of TRF2 at telomeres by tethering it to TRF1 (29, 31, 32). In vivo data indicate that upon downregulation of TRF1 or TIN2 by RNAi, the localization of TRF2 at telomeres is significantly diminished (29, 31). Finally, TIN2 is essential for the recruitment of POT1 to double-stranded telomeric DNA and, therefore,

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plays a crucial role in the loading of POT1 onto the single-stranded telomeric DNA (see later). Thus, TIN2 is necessary both for the stabilization of the TRF1/TRF2 scaffold on the double-stranded DNA and for the coating of the single-stranded DNA with POT1 (Fig. 2.2).

2.2.1.3

TPP1 and POT1

The main function of TPP1, the most recently identified shelterin component, is to link TIN2 and POT1 (30, 32, 35) (Fig. 2.2). In the absence of TPP1, POT1 is not recruited to chromosome ends and the phenotypes mirror POT1 loss. At first glance, it is surprising that the association of POT1 with telomeres is not dependent on its ability to bind DNA. POT1 contains two oligonucleotide/oligosaccharide-binding (OB) folds that are highly specific for single-stranded 50 -(T)TAGGGTTAG-30 sequence, both when the sequence is located at an internal site and when it is at a 30 terminus (36–39). Instead, the recruitment and stabilization of POT1 at telomeres relies on the bridge that links POT1 through TPP1 and TIN2 to TRF1/TRF2 bound at the double-stranded repeat array (30, 35, 40, 41). This is supported by chromatin immunoprecipitation data showing that longer telomeres recruit more POT1, although the length of the single-stranded telomeric DNA is not significantly altered. In addition, POT1 truncation mutants that lack the DNA-binding OB folds still localize efficiently at telomeres (37). The model that has emerged argues that the local enrichment of POT1 molecules in the vicinity of the chromosome terminus complements the high specificity of POT1 for single-stranded telomeric DNA and promotes more efficient loading. Human cells express two forms of POT1 from alternatively spliced mRNAs. The abundance of the short form, POT1–55, which lacks the first OB-fold required for DNA binding, is approximately tenfold lower than the abundance of full-length POT1 (42). The function of the truncated protein remains to be established. Interestingly, mouse cells have two POT1 genes, POT1a and POT1b (43, 44). Both proteins associate with telomeres and share similar sequences and domain structures. However, they are not functionally redundant. POT1a is an essential gene, as its deletion leads to early embryonic lethality, while POT1b deficient mice are viable (43). The roles of the two mouse POT1 proteins are discussed below. Studies of the functionally divergent POT1s in rodents have provided interesting implications for their human counterpart.

2.2.1.4

RAP1

The sixth shelterin component, RAP1 is a binding partner of TRF2 (45). The interaction between the two factors is required for the recruitment of RAP1 to telomeric DNA and is essential for the stability of RAP1 protein levels (45). The exact function of RAP1 remains to be determined but mice lacking RAP1 are not viable suggesting that it plays an important role in telomere protection

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(van Overbeek M. and de Lange T., unpublished data). Although there are not any known direct binding partners of RAP1, it is likely that its BRCT and Myb domains are involved in protein–protein interactions of functional importance.

2.2.1.5

Perspective

Taken as a whole, the intricate interconnections between the different members of shelterin ensure that the complex has high affinity and specificity for telomeric DNA. Indeed, shelterin is highly abundant exclusively at telomeres and its known functions are restricted to telomere maintenance. At the same time, the redundancy of some of the recruitment mechanisms allows for flexibility. It remains to be determined whether different shelterin subcomplexes might be required to execute diverse functions and whether plasticity in the shelterin complex might be essential for structural remodeling of the telomeric DNA as cells progress through the cell cycle.

2.2.2

Telomere-Associated Proteins in Human Cells

In addition to shelterin, a number of other proteins have been detected at human telomeres. Most of these factors are DNA damage signaling and repair molecules that have been implicated to associate transiently with telomeres and to perform essential accessory functions in telomere maintenance. However, all of these proteins have primary functions that are independent from telomere biology. Examples of such factors include the Mre11 complex (46), which is thought to sense the presence of double-strand breaks and to participate in the homologous recombination pathway of DNA repair; XPF/ERCC1 (47), a component of the nucleotide-excision repair pathway; Apollo (48, 49), a putative 50 exonuclease; DNA-PKcs (50, 51), the PIKK kinase involved the nonhomologous end-joining (NHEJ) pathway; Ku70/80 (50, 52, 53), also involved at the first step of the NHEJ pathway; BLM and WRN RecQ helicases (54–57), implicated in branch migration of recombination structures; Rad51D (58), a factor with a potential role in homologous recombination; and others. Deficiency in some of these factors leads to telomere phenotypes and has also been independently implicated in human diseases. For example, Apollo knockdown leads to extensive DNA damage signaling at chromosome ends and aberrant telomere structures (48, 49). DNA-PKcs and Rad51D deficiencies, on the other hand, result in mild fusion phenotypes (58, 59). In the case of WRN helicase, its deficiency causes loss of lagging-strand telomeres (54), while mutations in the protein have been associated with Werner’s syndrome (reviewed in (60)). Similarly, BLM helicase is mutated in Bloom’s syndrome patients. Whether or not some of the symptoms in Werner’s and Bloom’s patients are caused by telomere dysfunction has not been conclusively determined. As described below, the main protective function of shelterin is to mask the chromosome ends from recognition by the DNA damage signaling and repair

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machineries. Therefore, it seems paradoxical that factors involved in these pathways are specifically recruited to telomeres. It is possible that their function is tailored in the context of shelterin to service telomeres without activating their respective signaling and repair pathways.

2.3

Shelterin Shapes Telomeric DNA into a Protected Structure

To maintain genome integrity, cells have evolved extensive mechanisms for immediate detection and repair of DNA damage. Double-strand breaks (DSBs), in particular, are highly toxic lesions. A single unrepaired or incorrectly repaired DSB can lead to significant loss of genetic information as a portion of the chromosome is detached from the centromere and might be missegregated in the following round of cell divisions. This can cause cell death, mutations, and chromosomal translocations, and can lead to diseases such as cancer. Unavoidably, in all eukaryotic cells, chromosome ends resemble DSBs and need to be masked to prevent recognition by the DNA damage signaling and repair machineries. Hiding chromosome ends into protected and veiled structures is the essential function of shelterin (5). This is achieved in several steps, which closely resemble the initial reactions of DNA repair mediated by the homologous recombination (HR) pathway. Normally, the HR machinery is involved in the error-free repair of DSBs. The first step in HR is the nucleolytic generation of a 30 overhang, which then strand invades a homologous region on the sister chromatid. In the following step, a Holliday junction intermediate is formed as a result of branch migration. Finally, upon completion of the repair reaction, the Holliday junction is resolved by resolvases to separate the two sister chromatids.

2.3.1

The Role of Shelterin in Generation of the 30 Overhang

An important requirement for telomere protection is the generation of a 30 overhang (Fig. 2.3a). Upon completion of DNA replication, the chromosome ends are either blunt or have a short 10–15 base pairs 30 extension on the lagging-strand chromosome. In most human cells, the average length of the telomeric overhang is 50–300 nucleotides (10, 61). This long single-stranded extension is generated by the action of an unknown nuclease(s) that resects the 50 strand (3, 9). Recent data suggest that the activity of the nuclease(s) might be regulated by POT1 (Fig. 2.3b). Human telomeres terminate precisely on CCAATC-50 on the resected strand but the exact end is randomized upon downregulation of POT1 protein levels by RNAi (42, 62). It is not known whether human POT1 interacts directly with the nuclease(s) to control its processivity or prevents excessive resection by binding to the DNA. One piece of evidence in favor of the second

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Fig. 2.3 Shelterin remodels chromosome ends into a protective structure. (a) The generation of the 30 telomere overhang is nuclease-dependent. Upon completion of DNA replication, chromosome ends are either blunt (leading-strand chromosome, bottom) or have a short 10–15 nt 30 extension due to the end-replication problem (lagging-strand chromosome, top). Unknown nuclease(s) resects the 50 strand to generate the overhang. (b) Regulation of nuclease activity by POT1. POT1 determines the precise CCAATC-50 end of human telomeres and regulates overhang length. As the preferred binding site of POT1 is located only 2 nt away from the precise 50 end, it is possible that POT1 determines the extent of 50 strand degradation by binding to the single-stranded DNA and directly inhibiting nuclease activity. (c) TRF1 and TRF2 mediate the stand invasion event, when the single-stranded overhang invades the double-stranded portion of the telomere and base pairs with the complimentary strand to generate the t-loop. (d) T-loop. The t-loop configuration is a protective structure, in which the 30 end of the chromosome is tucked away, possibly as a mechanism to evade recognition by the DNA damage signaling and repair machineries. The t-loop most likely recruits shelterin as shown. The displaced strand (D loop) is presumably coated with POT1, recruited by TRF1/TRF2 shelterin complexes bound to the double-stranded portions of the t-loop

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model is that the preferred POT1 DNA-binding site is located only 2 nucleotides away from the 50 end (39). Strikingly, deletion of mouse POT1b, one of the two POT1 genes in rodents, leads to extreme overhang elongation (43). This is seen both in mouse embryonic fibroblasts (MEFs) isolated from POT1b-deficient embryos and in liver samples taken from adult POT1b-deficient mice. The increase in single-stranded TTAGGG repeats is attributed to excessive 50 –30 nucleolytic activity and the resulting degradation of the C-rich 50 strand. Although POT1b-deficient mice are viable, the excessive loss at the 50 strand leads to progressive telomere shortening. Upon prolonged culture of POT1b-deficient MEFs, as telomere length erodes to a critical level, there is also evidence for significant genome instability and chromosome endto-end fusions (63). It is not known yet whether human POT1, which sets the 50 end sequence, has a similar role in preventing excessive nucleolytic degradation at the telomere terminus. On the contrary, experimental evidence suggests that downregulation of POT1 in human cells by RNAi leads to a slight but reproducible overhang loss (42). However, this result could be an artifact of incomplete knockdown of POT1 protein levels and the discrepancy can only be addressed in complete absence of human POT1. Theoretically, if human POT1 played a role in regulating the nucleolytic generation of the telomere overhang, then mutations in human POT1 that prevent control over the nuclease(s) would lead to progressive telomere erosion. The existence of such POT1 mutations would have profound implications for genome instability and tumorigenesis.

2.3.2

T-Loop Formation

Once the overhang is generated, the next step in telomere protection is thought to be the formation of a lariat structure at the chromosome terminus, referred to as telomeric loop (t-loop). In the t-loop configuration, the single-stranded telomeric DNA invades the double-stranded portion of the telomere, displaces the G-rich strand, and base pairs with the complementary strand (Fig. 2.3c, d). The predicted role of t-loops is to effectively shield the chromosome end from nucleolytic attack and from recognition by DNA damage factors (reviewed in (64) ). T-loops of purified human telomeres, which have been cross-linked to maintain structure integrity, can be directly visualized by electron microscopy (65). Analysis of the structural features confirms a strand invasion event, including the presence of single-stranded G-rich strand, which forms a displacement loop (D loop). The size range of t-loops is heterogeneous and roughly correlates with the total telomere length, suggesting that the strand invasion takes place at a random site along the telomere duplex array (65). Recently, electron microscopy analysis of whole telomere chromatin isolated from chicken erythrocytes and mouse splenocytes further revealed the presence of intact nucleosome arrays along the t-loop structures (66).

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So far, no assay has been developed to test for the presence of t-loops in vivo in human cells. The current model argues that t-loops are probably present at all chromosome ends, throughout the cell cycle, except perhaps temporarily during S-phase when the passage of the replication machinery would release the invading strand, thereby revealing a naked or POT1-bound single-stranded DNA end. Both TRF1 and TRF2 have been implicated to play roles in t-loop formation. In vitro data suggest that TRF2 has the ability to remodel DNA. To begin with, when recombinant TRF2 is incubated with a duplex TTAGGG repeat array containing a 30 single-stranded overhang, 10–15% of the resulting complexes resemble t-looplike structures generated by a strand invasion event (65, 67). Biochemical analysis further suggests that TRF2 has the ability to modify DNA topology and more specifically, to induce untwisting of neighboring DNA, thereby promoting strand invasion (68). TRF1 also has in vitro DNA remodeling capacity including ability to bend, loop, and pair distant regions containing telomeric repeats (20, 23, 24). The versatility of TRF1 in modifying telomeric DNA topology is attributed to the structural flexibility of the region between its dimerization domain and DNAbinding domain, which allows two TRF1 molecules to bind to spatially distant telomeric repeats while maintaining a dimerized core (20). It is also likely that some of the telomere-associated factors described above may also participate in t-loop assembly. In particular, the Mre11 complex and BLM helicase have the functional requirements to promote t-loop formation and/or resolution but experimental evidence in support of this hypothesis is lacking.

2.3.3

Prevention of Inappropriate T-Loop Deletion

The final product of t-loop formation is a structure that closely resembles an HR intermediate. On the basis of the predicted structure, the t-loop contains a Holliday junction-like configurations. If resolved, the result would be the circularization of the t-loop and significant shortening of the remaining telomere (69). This phenotype is similar to the telomere rapid depletion (TRD) events observed in yeast (70). Evidence strongly argues that TRF2 is involved in preventing inappropriate resolution of the t-loop. Overexpression of a mutant allele of TRF2 lacking the aminoterminal basic domain (TRF2DB) leads to excessive telomere loss and increased appearance of telomere sequence-containing circles that are approximately the size of t-loops (71). The telomere loss phenotype, referred to as t-loop HR, is abrogated in XRCC3 and NBS1-deficient cells, implicating the involvement of the HR pathway (71). It remains to be established why introduction of TRF2DB leads to t-loop deletion. One possibility is that the overexpressed protein titrates a factor that plays a role in maintaining t-loops. Another possibility is that the basic domain of TRF2 is directly involved. This model is strongly supported by biochemical experiments demonstrating that TRF2 but not TRF2DB binds to replication forks

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and four-way junctions in vitro in a structure-specific although sequence-independent manner (72). These data predict that TRF2 robustly interacts with a t-loopspecific structure and either stabilizes it to prevent branch migration or spatially prevents the access of a resolvase. T-loop deletion involves inappropriate processing of the chromosome terminus that can lead to significant telomere loss. Interestingly, telomeric circles have also been detected at low frequency in a variety of human cells, suggesting that they can occur spontaneously (71). It remains to be determined whether sporadic t-loop deletion contributes to stochastic shortening of human telomeres. If t-loop deletion led to complete telomere erosion, such events, even if extremely rare, would significantly promote genome instability. In addition, increased presence of telomere circles has been detected in alternative lengthening of telomeres (ALT) cells (71, 73). The recombination-based ALT pathway is active in many tumor cell lines that have undetectable telomerase activity and is an alternative mechanism for maintaining telomere length (reviewed in (74)).

2.4

Suppression of the ATR and ATM DNA Damage Response Pathways

In summary, the t-loop effectively solves the problem of having an exposed DNA end at the chromosome terminus by masking it into a circular structure. However, t-loops still contain features that would be recognized by the DNA damage surveillance machinery – the single-stranded DNA on the D loop and several regions with single-stranded double-stranded transitions. Again, the shelterin complex is responsible for suppressing the activation of the DNA damage surveillance machinery at those sites.

2.4.1

Suppression of ATR Pathway

Stretches of single-stranded DNA are recognized as sites of damage by the ATR pathway. The ATRIP/ATR complex is recruited to RPA-coated single-stranded DNA (75). The ATR kinase induces cell cycle arrest through the phosphorylation and activation of the downstream checkpoint kinase Chk1 (76). The ATR pathway responds primarily to single-stranded breaks such as generated by replication fork stalling or UV-induced damage. Telomeres also contain stretches of single-stranded DNA even when the 30 overhang is base-paired as in the t-loop configuration, because the D loop is exposed and can potentially recruit RPA and activate the ATR pathway (65). Recent data on mouse and human cells suggest that POT1 is the shelterin component that prevents the activation of the ATR pathway at telomeres (77) (Fig. 2.4). Conditional deletion of POT1a or knockdown of its recruiter, TPP1, in

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Fig. 2.4 Shelterin protects chromosome ends from recognition by the DNA damage signaling and repair machineries. TRF2 complex inhibits the activation of the ATM pathway and prevents nonhomologous end-joining (NHEJ) of telomeres (left). In the absence of TRF2, ATM is activated and phosphorylates Chk2 kinase, which in turn promotes the p53/p21 pathway, leading to senescence or apoptosis, depending on the cell type. Active ATM kinase also leads to the accumulation of multiple DNA damage response factors, including g-H2AX and MDC1, at chromosome ends, which can promote the NHEJ pathway at telomeres (middle panel). The overhang is cleaved in a reaction dependent on the XPF/ERCC1 endonuclease, the Ku70/80 complex is loaded on the resulting ends and DNA ligase IV executes the fusion reaction. Chromosome end-to-end fusions can be deleterious as they lead to the formation of dicentric chromosomes that cannot segregate properly during mitosis. On the other hand, POT1 (bound along the overhang or on the D loop) suppresses the ATR signaling pathway (right). Upon loss of POT1, ATR is activated and phosphorylates and activates the downstream Chk1 kinase. In the absence of TRF2, ATM, and POT1 in mouse cells, ATR activation also promotes the NHEJ pathway

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MEFs leads to acute activation of the DNA damage response at chromosome termini (40, 43, 44). This involves the recruitment of the DNA damage response factors g-H2AX and 53BP1 to telomeres. In addition, Chk1 is phosphorylated suggesting that the downstream signaling pathway is triggered. The activation of the DNA damage pathway in response to TPP1/POT1a loss is dependent on the ATR kinase (77). When ATR protein levels are downregulated by RNAi, the damage response is abrogated. One possibility is that POT1 prevents the activation of damage signaling by competing with RPA for the binding to the single-stranded telomeric DNA. The high specificity of POT1 for telomeric sequence combined with its high abundance at the chromosome end might be important factors that allow POT1 to efficiently coat the single-stranded DNA on the D loop or along the overhang. Human POT1 plays a similar role in preventing ATR activation as mouse POT1a. On the other hand, human cells that overexpress mutant POT1, which lacks the first OB-fold required for DNA binding, POT1DOB, proliferate normally (37). This contradicts the model that POT1 coats single-stranded DNA to mask it from recognition by the DNA damage machinery and competes directly with RPA for binding at those sites. However, the presence of endogenous POT1 in these experiments might be enough to suppress ATR activation.

2.4.2

Suppression of ATM Pathway

TRF2 is responsible for the suppression of the ATM pathway (77) (Fig. 2.4). ATM kinase, in contrast to ATR, responds primarily to the presence of DSBs and its principal downstream effector is the Chk2 kinase (78). In TRF2-deficient MEFs, telomeres activate the ATM-dependent pathway, including the recruitment of a number of DNA damage response factors, such as g-H2AX, MDC1, 53BP1 and the Mre11 complex, to chromosome ends (27). Loss of TRF2 also leads to the autophosphorylation of the ATM kinase, which in turn phosphorylates and activates Chk2. Telomere dysfunction can also be induced in human cells, when the function of human TRF2 is suppressed as a result of the overexpression of a dominant negative allele of TRF2, which lacks the amino-terminal basic and the carboxy-terminal Myb domains (TRF2DBDM) (79). The dominant negative allele dimerizes with the endogenous protein but since it lacks the DNA-binding domain, the resulting heterodimer does not localize to telomeres. In addition to activating the canonical ATM pathway (80), as described above, exogenous introduction of TRF2DBDM results in p53-dependent cell cycle arrest or apoptosis, depending on the cell type (81, 82). There are several models as to how TRF2 prevents the activation of ATM kinase at functional telomeres. First, TRF2 might be required to maintain the terminal structure, which in trun may repress DNA damage signaling by preventing the binding of the sensor in the ATM pathway. It is likely that the role of TRF2 in promoting t-loop formation may be required to prevent activation of the ATM

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pathway. A complementary model suggests that TRF2 directly inhibits the activation of ATM kinase. TRF2 binds weakly to ATM in a region that contains the residue Serine 1981, one of the autophosphorylation sites that promote ATM activation (85). Furthermore, overexpression of TRF2 reduces the ATM-dependent response to irradiation-induced damage. As a result, both the activation of downstream effectors and the induction of cell cycle arrest are significantly diminished (85). Presumably, since TRF2 is exclusively enriched at chromosome termini and not elsewhere in the cell, ATM activation would be specifically dampened in the vicinity of telomeres. Therefore, even if telomeres present DNA structures that would normally signal to the ATM pathway, TRF2 locally suppresses any downstream propagation.

2.5

Prevention of Inappropriate NHEJ and HR Repair at Chromosome Ends

In addition to suppressing the activation of ATR and ATM signaling, shelterin efficiently prevents inappropriate repair reactions at chromosome ends (Fig. 2.4). The consequences of aberrant repair processing of telomeres in human cells can be deleterious. In particular, fused chromosomes, which have been joined end-to-end, are dicentric and cannot properly segregate in mitosis. Instead, they propagate the bridge-breakage-fusion cycle (86), which can lead to extensive genomic instability as chromosomes are broken and rejoined at random places each cell division. The role of shelterin in suppressing inappropriate repair at telomeres can be best appreciated in the setting when loss of TRF2 function leads to uncapping of chromosome ends. Upon inhibition of TRF2 – both in the TRF2 conditional knockout MEFs and upon exogenous overexpression of the TRF2DBDM allele in human cell – telomere-mediated protection is lost and chromosome ends undergo extensive processing (27, 79). The consequences are striking. Metaphase spreads collected five days after deletion of TRF2 reveal that virtually all chromosomes have fused to one another, creating long trains, with the telomeric DNA retained at the sites of fusion (27). Evidence for the involvement of the NHEJ pathway came from genetic experiments, which showed that Ku70 and DNA ligase IV are essential factors executing the end-joining of dysfunctional telomeres (27, 83, 84). In addition, the 30 telomere overhangs are removed by the XPF/ERCC1 endonuclease promoting the NHEJ reaction (47). Interestingly, in mouse cells overhang cleavage and end-joining are coupled, while in human cells the two processes can occur independently (27, 87). As overhang loss is a prerequisite for the execution of the NHEJ reaction, it is possible that TRF2 prevents inappropriate repair by hiding the overhang into the t-loop structure. Furthermore, the circular configuration of t-loops would prevent the first step of NHEJ – loading of the Ku70/80 complex on a free double-stranded DNA end. In addition, the binding partner of TRF2, RAP1 may be directly involved

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in preventing end-joining of short telomere arrays in an in vitro reconstituted system, even in the absence of t-loop formation (88). It has not been established yet whether RAP1 inhibits the NHEJ pathway in vivo. Finally, it is possible that TRF2 prevents the activation of repair in part by suppressing ATM-mediated DNA damage signaling. Although DNA damage signaling and repair have been viewed as largely separate processes but recent data suggest that they are intrinsically connected. More specifically, phosphorylation of H2AX by ATM kinase and recruitment of MDC1 to g-H2AX-coated chromatin at TRF2-depleted telomeres significantly accelerates NHEJ (87). In the absence of H2AX and MDC1, the joining of dysfunctional telomeres is notably delayed. Therefore, TRF2 might partially suppress repair by preventing activation of the ATM pathway. In support, TRF2- and ATM-deficient MEFs, which fail to activate the telomere damage response, also do not contain telomere end-to-end fusions (77). On the other hand, NHEJ is not a consequence of POT1 loss, despite the activation of DNA damage signaling at telomeres, suggesting that the presence of functional TRF2 is crucial for the repression of inappropriate repair (42, 43, 77). There is evidence that telomere fusions can also occur naturally, as cells approach senescence. Recent study documents fusions of critically short telomeres in a fibroblast population that is ongoing cell divisions past the senescence setpoint (89), underscoring the importance of understanding the role of shelterin in suppressing inappropriate repair processes. Interestingly, TRF2 plays a role in the repression of the HR pathway as well. As described above, dysfunctional telomeres resulting from TRF2 loss are repaired primarily through the NHEJ pathway. However, in the absence of Ku70 and a functional NHEJ pathway, deletion of TRF2 does not lead to overhang loss and does not result in telomere fusions. Instead, extensive HR between sister telomeres takes place, leading to numerous telomeric sister-chromatid exchanges (T-SCEs) (84). One interpretation of this data is that a TRF2 and Ku-dependent mechanism exists that suppresses T-SCEs to prevent drastic telomere length changes that would be an inevitable consequence of unequal exchanges.

2.6 2.6.1

Telomere Length Regulation by the Shelterin Complex Shelterin-Mediated Control of Telomerase

Telomerase is active in the germ line and inappropriately activated in the majority of human cancers. In those cases, telomerase functions to counteract telomere shortening and to maintain a stable telomere length setting. It has been firmly established that telomerase is regulated by a negative feedback loop that involves shelterin (reviewed in (90)) (Fig. 2.5). In short, addition of more telomeric repeats by telomerase leads to the recruitment of more shelterin, which in turn has the ability to inhibit telomerase recruitment and/or activity.

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Fig. 2.5 Shelterin regulates telomere length homeostasis. Long telomeres contain more shelterin (top panel). The increased amount of shelterin bound at the double-stranded portion of the telomere increases the chance that POT1/ TPP1 will be loaded at the 30 terminus and block the access of telomerase to its substrate. Short telomeres or inhibition of shelterin (bottom panel) results in inefficient loading of POT1/ TPP1 on the overhang or leads to loading of POT1/ TPP1 at an internal site. POT1/ TPP1 complex that is not located at the 30 terminus might have a positive effect on telomerase recruitment and/or processivity, thus promoting elongation of short telomeres

2.6.1.1

TRF1

TRF1 was the first shelterin component implicated in telomere length homeostasis in human cells. Two initial experiments suggested that TRF1 is a negative regulator of telomerase (91). On the one hand, long-term overexpression of TRF1 resulted in gradual and progressive telomere shortening, even in the presence of telomerase. On the other hand, significant telomere elongation was induced when the binding of endogenous TRF1 to telomeric repeats was prevented. Next, it was shown that more TRF1 is recruited to longer telomeres (37, 92) and that TRF1 exercises its inhibitory effect on telomerase only in cis (93). Taken together, these experiments led to the conclusion that TRF1 plays a role in a ‘‘protein counting’’ mechanism, which presumes that the length of the telomeric array is translated into number of TRF1 molecules bound to telomere repeats (91, 94).

2.6.1.2

POT1 and TPP1

In turn, TRF1 transmits the signal through TIN2 and TPP1 to the terminal effector, POT1 (37). POT1 binds to single-stranded DNA, the site where telomerase acts, and therefore, has the potential to negatively affect the recruitment of the enzyme.

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Evidence in support of this model came from experiments in which POT1DOB, a mutant allele of POT1, which lacks the crucial domain for DNA binding, is introduced into telomerase-positive cells (37). The outcome is rapid and extensive telomere elongation that occurs in the absence of any other significant alterations to the shelterin complex. The interpretation of these experiments is that POT1 has the ability to convey the ‘‘protein counting’’ signal from TRF1 to the telomere terminus, where, by binding to the single-stranded DNA end, POT1 directly inhibits the access of telomerase to its substrate (95, 96). Recent structural data further suggested that the POT1–TPP1 complex is conserved and analogous to the ciliate TEBPa/b complex, which is responsible for overhang protection (41, 97). This correlation led to the intriguing possibility that POT1–TPP1 might act as a unit to regulate telomere length homeostasis. Strikingly, POT1 and TPP1 complexed with telomeric DNA demonstrated an ability to significantly increase the activity and processivity of the human telomerase core enzyme (97). In addition, although TPP1 enhanced the binding of POT1 to single-stranded DNA, thereby inhibiting the access of telomerase to its substrate, TPP1 also directly interacted with telomerase, possibly playing a role in its recruitment to chromosome ends (41). Therefore, it is conceivable that POT1–TPP1 complex acts both as a negative and as a positive regulator of telomerase (41). The current model argues that POT1–TPP1 complex switches from inhibiting the access of telomerase to the telomere to serving as a processivity factor for telomerase during telomere extension (97). It remains to be determined what circumstances promote one state vs. the other.

2.6.1.3

TRF2 and RAP1

Other shelterin factors are also involved in the regulation of telomere length. Overexpression of TRF2 leads to telomere shortening (98), while expression of RAP1 mutants affects the heterogeneity of telomere length (99). Although the role of these two factors in telomere length homeostasis is less well defined, they might affect indirectly the TRF1-TPP1-POT1 axis within the context of the shelterin complex. It is also possible that TRF2 and RAP1 function independently by recruiting an unknown factor(s) involved in telomerase regulation.

2.6.1.4

Tankylase 1

Another level of telomere length control is exercised through an enzyme called tankyrase 1, which modifies TRF1 (33). Tankyrase 1 is one of the telomere-associated proteins that are recruited to telomeres but whose abundance is much less compared with that of the shelterin subunits. Tankyrase 1 is of particular interest because in human cells it has the ability to add poly-ADP-ribose

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(PAR) chains to a motif located in the amino-terminal part of TRF1. As a consequence of this modification (referred to as PARsylation), TRF1 detaches from telomeric DNA (33). Thus, when tankyrase 1 is overexpressed, telomeres undergo progressive elongation (100). This is explained by decreased recruitment of TRF1 to telomeres and the resulting relief of negative regulation on telomerase activity. Conversely, when tankyrase 1 is inhibited, telomeres shorten, consistent with a situation when in the absence of tankyrase the binding of TRF1 to telomeres is promoted (101, 102). Interestingly, TIN2 is also involved in this pathway and has the capacity to protect TRF1 from tankyrase 1-dependent modification (34). As tankyrase 1 has additional roles in other cellular processes, including mitosis (103), it would be interesting to dissect the pathway that regulates its activity at telomeres.

2.6.1.5

Perspective

Most of the studies in the past have focused on the negative regulation exerted by shelterin on telomerase activity. However, a recent report demonstrated that the protein levels of telomerase within the nucleus are very limited (around 20 molecules per cell) (104). An independent study provided evidence that limiting telomerase levels are, in fact, required for the maintenance of stable telomere length (105). These new concepts opened the possibility that telomerase might need help from shelterin to localize to chromosome ends that require elongation. The data that POT1–TPP1 complex plays a role in positively regulating the processivity of the telomerase enzyme and may promote its recruitment to telomeres fit with that model (41, 97).

2.6.2

Telomere Length Homeostasis in the Absence of Telomerase

Telomerase is not active in most human somatic cells. Therefore, in the absence of telomere elongation, the length homeostasis in these cells depends primarily on the rate of telomere shortening. As mentioned earlier, telomeres erode as a result of the end-replication problem and as a consequence of nucleolytic degradation each cell division. In the absence of telomerase, shelterin controls the extent of the nuclease activity and thereby is the main factor involved in telomere length regulation. As telomeres gradually shorten to a critical length, the progressively diminishing levels of shelterin at telomeres remain the primary factor maintaining chromosome ends. This is supported by evidence that even though overexpression of TRF2 in primary human cells increases the rate of telomere shortening, the telomere length at senescence, defined as senescence setpoint, is reduced from 7 to 4 kilobases (98). In this setting, overexpressed TRF2 is capable of protecting critically short telomeres and preventing chromosome end fusions. Thus, senescence is induced by a

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change in the protected status of shortened telomeres rather than by a complete loss of telomeric DNA and increased presence of shelterin can delay this event (98). Conversely, reduced shelterin levels might have the opposite effect and induce senescence earlier, even in the presence of relatively long telomeres. Recently, it was determined that the minimum telomere array detected in human presenescent cells spans between 7 and 14 TTAGGG repeats (89). Telomeres that have eroded beyond that level may no longer recruit sufficient shelterin to maintain integrity and are likely to activate the DNA damage machinery and become fusogenic (106–108). As a consequence of excessive telomere shortening, their genomes may become highly unstable. It is precisely at that moment that inappropriate activation of telomerase would significantly promote tumorigenesis by stabilizing the telomere length of cells that have already acquired genomic alterations (reviewed in (109)). Under normal circumstances, this is avoided by the activation of the senescence program, which leads to permanent cell cycle arrest (reviewed in (110)). It would be of particular significance to determine whether shelterin participates in this process. For example if, as described above, shelterin plays a dual role in the regulation of telomerase – then the absence of shelterin at eroded telomeres might also preclude inappropriate telomerase activity at those chromosome ends. Alternatively, elevated levels of shelterin, TRF2 in particular, might prevent chromosome instability even when telomeres have become critically short. This would be especially important in situations when additional mutations in the DNA damage signaling pathway delay or compromise the execution of the senescence program. Dissecting how and at what point cycling cells with a functional shelterin complex at telomeres transition to senescence is essential for our understanding of tumorigenesis.

2.7

Concluding Remarks

The role of shelterin at human telomeres is multifaceted and intricate. It is truly amazing that a single complex can accomplish such a variety of different tasks – from structural remodeling of the chromosome terminus, to suppression of the DNA damage signaling and repair machineries, to regulation of telomere-length homeostasis. In all of these cases, however, it seems that shelterin has to carry out seemingly contradictory functions. For example, shelterin allows for plasticity as cells progress through the cell cycle and chromosome termini undergo DNA replication, but at all times shelterin maintains its robust protective function. As discussed, a failure of shelterin to shield telomeres would have deleterious consequences for the cell as chromosome ends are recognized as sites of DNA damage and activate checkpoint signaling. In addition, shelterin prevents inappropriate signaling and repair but at the same time recruits various DNA damage signaling and repair factors to chromosome ends. It remains to be established how the activities of these factors are regulated at functional telomeres. Finally, shelterin seems to simultaneously promote and inhibit telomerase activity. It is crucial to

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comprehend the contrasting roles of shelterin in telomerase regulation, as defects in telomere-length maintenance can lead to severe disorders at the level of the organism. Dyskeratosis congenita is an example of a syndrome resulting from significant telomere loss, while the opposite, inappropriate maintenance of telomere length, is a hallmark of cancer. Therefore, a deeper understanding of shelterin will not only expand our knowledge of cell biology, but will also provide crucial implications for human health.

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68. Amiard S, Doudeau M, Pinte S et al. A topological mechanism for trf2-enhanced strand invasion. Nat Struct Mol Biol 2007; 14:147–54. 69. Haber J. Telomeres thrown for a loop. Mol Cell 2004; 16:502–03. 70. Bucholc M, Park Y, Lustig AJ. Intrachromatid excision of telomeric DNA as a mechanism for telomere size control in saccharomyces cerevisiae. Mol Cell Biol 2001; 21:6559–73. 71. Wang RC, Smogorzewska A, De Lange T. Homologous recombination generates t-loopsized deletions at human telomeres. Cell 2004; 119:355–68. 72. Fouche N, Cesare AJ, Willcox S, Ozgur S, Compton S, Griffith J. The basic domain of trf2 directs binding to DNA junctions irrespective of the presence of ttaggg repeats. J Biol Chem 2006; 281:37486–95. 73. Cesare AJ, Griffith JD. Telomeric DNA in alt cells is characterized by free telomeric circles and heterogeneous t-loops. Mol Cell Biol 2004; 24:9948–57. 74. Reddel RR. Alternative lengthening of telomeres, telomerase, and cancer. Cancer Lett 2003; 194:155–62. 75. Zou L, Elledge SJ. Sensing DNA damage through atrip recognition of rpa-ssdna complexes. Science 2003; 300:1542–48. 76. Liu Q, Guntuku S, Cui XS et al. Chk1 is an essential kinase that is regulated by atr and required for the g(2)/m DNA damage checkpoint. Genes Dev 2000; 14:1448–59. 77. Lazzerini Denchi E, De Lange T. Protection of telomeres through independent control of atm and atr by trf2 and pot1. Nature 2007; 448:1068–71. 78. Chaturvedi P, Eng WK, Zhu Y et al. Mammalian chk2 is a downstream effector of the atmdependent DNA damage checkpoint pathway. Oncogene 1999; 18:4047–54. 79. Van Steensel B, Smogorzewska A, De Lange T. Trf2 protects human telomeres from end-toend fusions. Cell 1998; 92:401–13. 80. Takai H, Smogorzewska A, De Lange T. DNA damage foci at dysfunctional telomeres. Curr Biol 2003; 13:1549–56. 81. Karlseder J, Broccoli D, Dai Y, Hardy S, De Lange T. P53- and atm-dependent apoptosis induced by telomeres lacking trf2. Science 1999; 283:1321–25. 82. Smogorzewska A, De Lange T. Different telomere damage signaling pathways in human and mouse cells. Embo J 2002; 21:4338–48. 83. Smogorzewska A, Karlseder J, Holtgreve-Grez H, Jauch A, De Lange T. DNA ligase ivdependent nhej of deprotected mammalian telomeres in g1 and g2. Curr Biol 2002; 12:1635. 84. Celli G, Lazzerini Denchi E, De Lange T. Ku70 stimulates fusion of dysfunctional telomeres yet protects chromosome ends from homologous recombination. Nat Cell Biol 2006; 8: 885–90. 85. Karlseder J, Hoke K, Mirzoeva OK et al. The telomeric protein trf2 binds the atm kinase and can inhibit the atm-dependent DNA damage response. PLoS Biol 2004; 2:E240. 86. McClintock B, in The fusion of broken ends of sister half-chromatids following chromatid breakage at meiotic anaphase, Ed. (Garland Publishing, Inc, New York and London, 1938), pp. 1–48. 87. Dimitrova N, De Lange T. Mdc1 accelerates nonhomologous end-joining of dysfunctional telomeres. Genes Dev 2006; 20:3238–43. 88. Bae N, Baumann P. A rap1/trf2 complex inhibits nonhomologous end-joining at human telomeric DNA ends. Mol Cell 2007; 26:323–34. 89. Capper R, Britt-Compton B, Tankimanova M et al. The nature of telomere fusion and a definition of the critical telomere length in human cells. Genes Dev 2007; 21:2495–508. 90. Smogorzewska A, De Lange T. Regulation of telomerase by telomeric proteins. Ann Rev Biochem 2004; 73:177–208. 91. Van Steensel B, De Lange T. Control of telomere length by the human telomeric protein trf1. Nature 1997; 385:740–43. 92. Smogorzewska A, Van Steensel B, Bianchi A et al. Control of human telomere length by trf1 and trf2. Mol Cell Biol 2000; 20:1659–68.

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93. Ancelin K, Brunori M, Bauwens S et al. Targeting assay to study the cis functions of human telomeric proteins: Evidence for inhibition of telomerase by trf1 and for activation of telomere degradation by trf2. Mol Cell Biol 2002; 22:3474–87. 94. Marcand S, Gilson E, Shore D. A protein-counting mechanism for telomere length regulation in yeast. Science 1997; 275:986–90. 95. Kelleher C, Kurth I, Lingner J. Human protection of telomeres 1 (pot1) is a negative regulator of telomerase activity in vitro. Mol Cell Biol 2005; 25:808–18. 96. Lei M, Zaug AJ, Podell ER, Cech TR. Switching human telomerase on and off with hpot1 protein in vitro. J Biol Chem 2005; 280:20449–56. 97. Wang F, Podell E, Zaug AJ et al. The pot1-tpp1 telomere complex is a telomerase processivity factor. Nature 2007; 445:506–10. 98. Karlseder J, Smogorzewska A, De Lange T. Senescence induced by altered telomere state, not telomere loss. Science 2002; 295:2446–49. 99. Li B, De Lange T. Rap1 affects the length and heterogeneity of human telomeres. Mol Biol Cell 2003; 14:5060–68. 100. Smith S, De Lange T. Tankyrase promotes telomere elongation in human cells. Curr Biol 2000; 10:1299–302. 101. Seimiya H, Muramatsu Y, Smith S, Tsuruo T. Functional subdomain in the ankyrin domain of tankyrase 1 required for poly(adp-ribosyl)ation of trf1 and telomere elongation. Mol Cell Biol 2004; 24:1944–55. 102. Donigian J, De Lange T. The role of the poly(adp-ribose) polymerase tankyrase1 in telomere length control by the trf1 component of the shelterin complex. J Biol Chem 2007; 282:22662–67. 103. Dynek JN, Smith S. Resolution of sister telomere association is required for progression through mitosis. Science 2004; 304:97–100. 104. Cohen S, Me G, Lovrecz G, Bache N, Robinson P, Reddel RR. Protein composition of catalytically active human telomerase from immortal cells. Science 2007; 315:1850–53. 105. Cristofari G, Lingner J. Telomere length homeostasis requires that telomerase levels are limiting. EMBO J 2006; 25:565–74. 106. Bakkenist CJ, Drissi R, Wu J, Kastan MB, Dome JS. Disappearance of the telomere dysfunction-induced stress response in fully senescent cells. Cancer Res 2004; 64:3748–52. 107. D’adda Di Fagagna F, Reaper PM, Clay-Farrace L et al. A DNA damage checkpoint response in telomere-initiated senescence. Nature 2003; 426:194–98. 108. Herbig U, Jobling WA, Chen BP, Chen DJ, Sedivy JM. Telomere shortening triggers senescence of human cells through a pathway involving atm, p53, and p21(cip1), but not p16(ink4a). Mol Cell 2004; 14:501–13. 109. Artandi SE, Depinho RA. A critical role for telomeres in suppressing and facilitating carcinogenesis. Curr Opin Genet Dev 2000; 10:39–46. 110. Campisi J. Cellular senescence as a tumor-suppressor mechanism. Trends Cell Biol 2001; 11:S27–31.

Chapter 3

Regulation of Telomerase Through Transcriptional and Posttranslational Mechanisms Amy N. Depcrynski, Patrick C. Sachs, Lynne W. Elmore, and Shawn E. Holt

Abstract The enzyme telomerase is associated with nearly 90% of human cancers. To better understand telomerase at the molecular level, a number of proteins involved in its regulation, either directly or indirectly, have been identified. This chapter aims to give a broad overview of both transcriptional and posttranslational telomeraseregulating proteins. Telomerase is transcriptionally repressed and activated by proteins acting on the promoter region, such as the Mad/Max heterodimer, c-Myc, p53, and Rb. Various kinases and ubiquitin ligases interact with telomerase, suggesting that phosphorylation and ubiquitination play important roles in inhibiting and activating the enzyme. Also included in this chapter are proteins that regulate localization of hTERT, assembly of hTERT, hTR regulators, and telomere-binding proteins that associate with telomerase. By gaining a better understanding of how telomerase is regulated, we can identify ways to block the enzyme in cancer cells or to activate the enzyme in normal cells as a means of modifying the cellular aging process. Keywords: Telomerase, Aging, Chemotherapeutics, Transcription, Telomere.

3.1

Introduction

In normal human cells, DNA polymerases are unable to replicate the very end of the chromosome, resulting in an inability to maintain telomeres. This phenomenon is known as the ‘‘end-replication problem,’’ which is characterized by cell division and gradual telomere shortening, ultimately leading to the growth-arrest state known as senescence. In cells with unlimited proliferative capacity, the enzyme telomerase maintains chromosome lengths by providing a template and a catalytic subunit to add telomeric sequences. Telomerase is associated with almost 90% of human cancers and nearly 99% of advanced malignancies, making it an obvious S.E. Holt(*) Department of Pathology, at Virginia Commonwealth University, Richmond, VA 23298-0662, e-mail: [email protected]

K. Hiyama (ed.), Telomeres and Telomerase in Cancer. DOI: 10.1007/978-1-60327-879-9_3, # Humana Press, a part of Springer Science + Business Media, LLC 2009 47

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diagnostic and therapeutic target (1). Most normal human cells lack expression of the catalytic subunit of telomerase, hTERT. Therefore, understanding the regulation of telomerase at the molecular level is critical for defining alternatives to blocking the enzyme in tumor cells to treat or prevent cancer, or activating it in normal cells to provide alternatives to cellular aging. In this chapter, we focus on telomerase-regulating proteins in humans, to provide a broad overview of the proteins whose functions, through either direct or indirect interactions with telomerase, have implications in the fields of oncology and aging.

3.2

Transcriptional Regulators

In 1999, the cloning and characterization of the hTERT promoter structure was published (2–5). Regulation of the hTERT subunit seems to be extremely important for carcinogenesis, as the RNA subunit (hTR) is constitutively expressed in most cells (6, 7), while the catalytic subunit (hTERT) is suppressed in normal somatic cells but expressed in tumor cell types (8–10). Ectopic expression of hTERT results in functional telomerase, telomere elongation, and extension of lifespan in a variety of cell types (11). Since then, many groups have undertaken the task of elucidating the genes involved in hTERT activation and regulation to provide a better understanding of telomerase’s role in tumorigenesis and extension of cellular life span. This field of research has led to the discovery of many promising transcription factors; however, none of the proposed factors alone have proven to be clear on/off switches for hTERT expression. The confounding problem is mainly due to the intrinsically complicated regulation of the hTERT gene. The transcriptional regulators discussed here, both positive and negative, include a variety of proteins that could potentially serve as chemotherapeutic targets, as many are oncogenes or involved in tumor suppressor pathways. The fact that so many of the proteins that participate in hTERT also participate tumor formation (or suppression) further implicates telomerase in tumor pathogenesis. The GC-rich hTERT promoter, unlike most promoters, lacks TATA or CCAAT boxes. There is an initiator-like sequence (CCTCTCC), which aids in RNA polymerase II locating the transcription start site in the absence of the TATA box. The promoter also contains Sp1 binding sites, two c-Myc binding sites (E boxes, at nucleotide numbers
34 and
242), AP binding sites (at nucleotide numbers
718 and
1,655), and a CCAC box (Fig. 3.1) (2–5). These binding sites interact with a variety of regulatory genes, which will be discussed later.

3.2.1

Negative Transcriptional Regulators

3.2.1.1

Mad/Max

The most extensively studied repressor is the Mad/Max heterodimer, which binds to the E-box domains of the hTERT core promoter region. The binding of

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Fig. 3.1 Structure and regulation of the hTERT promoter. Proteins that activate hTERT are located above the line, repressors are located below the line. Proteins that bind directly to the promoter are indicated on the line, in approximate locations as they occur along the promoter. Those proteins labeled in boxes above and below the line indicate proteins found at the promoter, acting indirectly through interactions with proteins binding directly to the promoter (See Color Insert)

the Mad/Max complex to the hTERT promoter prevents Myc/Max complex formation, which prevents the subsequent binding and activation at the E-boxes, thus repressing hTERT’s activity (12). The dynamic interactions between Myc/Max and Mad/Max have been implicated in many different cellular tissues as one of the most critical regulators for telomerase. The deregulation and overexpression of c-Myc that occur in cancer progression often correlates with the upregulation of hTERT activity (13–16), suggesting hTERT as a target for c-Myc regulation (discussed in detail later). Binding of Myc/Max and Mad/Max complexes to the hTERT promoter was initially discovered through the downregulation of hTERT mRNA by drug-induced differentiation of U937 cells, which resulted in decreased transcription of hTERT

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through reduced promoter activity. By examining the transcription factors present, Mad was found to bind as a repressor (17). Ectopic expression of c-Myc counteracted the Mad-mediated repression of hTERT, leading to the conclusion that Mad is a direct negative regulator of hTERT and can be inhibited by the presence of c-Myc (18). Mad transcriptionally regulates hTERT through an interaction with c-Myc in hTERT-positive cells (bladder cancer cells) and normal cells (19). Mad/Max may be in a ‘‘switched’’ role where the functions of Myc/Max are in competition for the Mad/Max complex to regulate telomerase activity. This function has direct implications for both tumorigenesis and regulation of cellular differentiation (12). Specifically, in proliferating cells, c-Myc is bound to hTERT at the E boxes, whereas in differentiated cells, Mad is bound at the promoter. This switch may be the key to the on/off regulation of telomerase in somatic cells (20). 3.2.1.2

Other Repressors Targeting c-Myc-Dependent Telomerase Activation

Further studies revealed that there is a full gamut of proteins capable of downstream targeting c-Myc-dependent telomerase activation. Receptor Ck, one such signaling protein, represses hTERT through the transcriptional repression of c-Myc, a function thought to be mediated by the inhibition of Protein Kinase C (21). Other regulatory genes involved with repression of c-Myc, and therefore repression of hTERT, include the HTLV-1 oncogene Tax, which downregulates the activity of telomerase directly through inhibition of hTERT transcription. The repression of hTERT is related to the competitive binding of Tax at the E-boxes domains in the core promoter region, which inhibits the binding of the c-Myc activator, thus preventing the transcription of hTERT. To date, Tax expression has only been reported during adult T cell leukemia/lymphoma (ATLL) pre-leukemic cell proliferation coupled with p53 inactivation (22). This disruption of telomere elongation appears to allow for the dividing cells to undergo fusions and subsequent ploidy changes that are associated with the process of leukemia progression (22). Because this is a cell type-specific regulation, it suggests a direct therapeutic target specifically for ATLL leukemia cells. 3.2.1.3

BRCA1

In breast cancer-related studies, a complex of the tumor suppressor BRCA1, Nmi (N-Myc and c-Myc interacting protein), and c-Myc inhibit the activation of the hTERT promoter activity through c-Myc. Co-expression of mutant Nmi with wild type BRCA1 blocked the hTERT activity, while expression of Nmi and mutated BRCA1 failed to inhibit c-Myc induction. The interaction of Nmi in vivo and in vitro with BRCA1 and c-Myc results in negative regulation of hTERT (23). Further, Nmi acts as an adaptor and recruiter of BRCA1 attachments to c-Myc. Therefore, BRCA1 and Nmi may be regulating c-Myc’s ability to induce tumorigenesis, while BRCA1 may act as a tumor suppressor by targeting c-Myc’s transcriptional

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activation of hTERT (23). Additionally, in cultured breast cancer cells, the interaction between BRCA1 and c-Myc inhibited activation of hTERT (24). Although indirect, the regulation of hTERT through BRCA1 (already a promising candidate for the development of targeted chemotherapy) may provide an additional target for adjuvant breast cancer therapy. Alternatively, inhibition of BRCA1 may result in an off-target effect of inhibiting telomerase, thereby providing an additional mode of action from a single drug directed against BRCA1.

3.2.1.4

p53

p53 is a potent tumor suppressor, which would naturally correlate with its interaction in hTERT regulation. Expression of ectopic p53 inhibits cancer cell growth and induces apoptosis or G1 arrest, which results in complete inhibition of telomerase activity in cancer cell lines. Pancreatic cells with mutant p53 showed high telomerase activity while ectopic p53 gene expression caused a decline in hTERT mRNA expression. These results suggest p53 as a negative transcriptional regulator of telomerase (25). In a cervical carcinoma cell line, p53 expression also repressed telomerase activity, presumably through p53’s interaction with multiple Sp1 binding sites on the hTERT promoter (although other factors may also be involved) (26). When p53 interacts with Sp1, a known activator of transcription, Sp1 loses its ability to activate hTERT expression (26). This interaction occurs through formation of an Sp1–p53 complex that is unable to bind the promoter (27). hTERT regulation by p53 also occurs in other tumor-derived cell lines, including those from prostate and uterine malignancies (26). Expression of wild type p53 in immortalized fibroblasts and lung cancer cells also causes a downregulation of telomerase activity (28, 29). However, in immortalized endothelial cells, p53 had no affect on hTERT activity (30), suggesting that p53’s effect on telomerase is celltype specific. In fact, the p53 target, p21waf1, when expressed in pancreatic cancer cell lines, also had no effect on telomerase, yet like p53, p21waf1 expression resulted in cell growth arrest and apoptosis (25). p53 is able to maintain its cellular functions while simultaneously inhibiting telomerase in certain cell types (25).Through the negative regulation of telomerase, p53 maintains its role as a tumor suppressor, but may only function through telomerase in a cell-type specific manner.

3.2.1.5

Rb

Another tumor suppressor protein, the retinoblastoma (Rb) protein, is able to induce senescence of tumor cells and inhibit telomerase activity, a function that is independent of p53. Certainly, the artificial in vitro setting does not closely recapitulate what occurs in a clinical setting; nonetheless, telomerase regulation through Rb without wild type p53 dependency suggests a role in inhibiting tumor growth and immortalization (31). In a squamous cell carcinoma line, reduction of telomerase activity correlated with an increase in Rb protein levels in the G0 and S phases of

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the cell cycle. Full length Rb, when overexpressed, caused a significant decrease in telomerase activity, further implicating its role in the regulation of telomerase activation in cancer cells (32). However, this does not necessarily implicate Rb as a direct regulator of telomerase transcriptional regulation. When Rb is overexpressed in carcinoma cells, it alone is sufficient to downregulate telomerase promoter activity, which can then be rescued by cdk2 and cdk4 expression. This implicates Rb in regulation of hTERT but does not characterize the exact mode of the regulation; whether it is directly acting on the hTERT promoter or acting secondarily to some other primary pathway still needs to be resolved (33). 3.2.1.6

IRF1 and p27Kip1

Interferon-g/interferon regulatory factor-1 (IRF1) signaling has previously been implicated in the downregulation of hTERT expression (34). Downstream of Interferon-g/IRF1 is the tumor suppressor protein, p27kip1, whose upregulation is dependent of IRF1. p27kip1 itself is able to inhibit transcriptional activation of hTERT. It has been hypothesized that p27kip1 acts by inhibiting the ability of the HPV E7 protein to upregulate the hTERT promoter activity, because Human Papillomavirus (HPV) E7 increases hTERT-promoter activity (35). p27kip1 is also a regulator of cyclin/CDK complexes and the cell cycle, and may be acting via a different pathway, through interactions with Rb or E2F, for example. The purported relationship between HPV E7, p27kip1, and hTERT seems to be far-reaching and may be better explained through interactions more closely related to cell cycle regulators, although this needs to be examined further. 3.2.1.7

TGFb, Smad3, and SIP1

TGFb decreases the activity of hTERT (36) with Smad3 and SIP1 contributing to its regulation (37–39). SIP1 (Smad interacting protein 1) is a downstream target of TGFb (40) and is required for TGFb to repress hTERT, although SIP1 alone does not regulate hTERT in vitro (39). Upon the activation of TGFb, an increase in Smad3 promotes binding to its own sequence adjacent in the hTERT promoter to the E-box/c-Myc binding site (at nucleotide numbers
262 and
284) (Fig. 3.1), which appears to be the main effect of TGFb inhibition on hTERT expression (38). 3.2.1.8

AP1

Binding sites for activator protein 1 (AP1), a heterodimeric complex of the Jun and Fos families, as well as sites for AP2 and AP4, have been identified within the hTERT promoter (2–5). AP1 over-expression studies in HeLa cells suggest that these binding sites do indeed have a suppressive effect on telomerase activity. This suppression mainly depends on the binding regions found between
2,000 and
378 on the hTERT promoter (Fig. 3.1); mutations of these sites resulted in a derepressive effect and elevated hTERT transactivation (41).

3 Transcriptional Regulators

3.2.1.9

53

WT1

Wilms’ tumor 1 (WT1) is one of the least studied binding sites on the hTERT promoter. This binding element is found on the fringe of the core promoter region at
352 (Fig. 3.1). WT1 has the ability to repress the activity of hTERT in 293T cells but not in HeLa cells, suggesting that WT1’s repressive effects are cell-type specific (e.g., those expressing endogenous WT1) (42). Although WT1 appears to be a repressor of telomerase, its limited actions may only allow for targeted therapy in certain tumor types.

3.2.1.10

Other Tumor Suppressors

A genetic screen identified a number of other tumor suppressors implicated in hTERT repression. This screen involved expression of a GFP reporter driven by the hTERT promoter in HeLa cells, with negative regulators of hTERT expression being identified through enhanced retroviral mutagens (ERM) (39). Candidate negative transcriptional regulators of hTERT included hSir2 and the cell signaling regulators Rak and BRCT-repeat inhibitor (BRIT1). Although RAK and BRIT1 are potential tumor suppressors identified in this screen, their direct roles in the regulation of telomerase have not been determined (39). Menin, a tumor suppressor, physically associates with the hTERT promoter region to inhibit transactivation. The primary study of the repression of hTERT via Menin cursorily examined possible repressive elements using the same general genetic screen (39). A transcriptional silencer of the hTERT promoter was identified in a variety of cervical and prostate cancer cell lines, and the inhibitory effects were enhanced with cellular differentiation. An homology search resulted in the identification of binding motifs for the myeloid-specific zinc finger protein 2 (MZF-2), Table 3.1 Negative transcriptional regulators Protein Regulatory role Mad1 Repressor, E-box binding Max heterodimer Repressor, through protein kinase C Receptor Ck Tax Repressor, E-box binding BRCA1/Nmi Repressor, c-Myc binding p53 Repressor, Sp1 binding Rb Repressor Repressor, with IRF1 p27kip1 MZF2 Repressor, direct promoter binding AP1 Repressor, direct promoter binding WT1 Repressor, direct promoter binding TGFb Repressor, with Smad3 BRIT/RAK Repressor, unknown mode of action Menin Repressor, direct promoter binding

Reference (12, 17–20) (21) (22) (23, 24) (25–30) (31, 32) (34, 35) (43) (2–5, 41) (42) (36–39) (39) (39)

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which when mutated, resulted in activation of hTERT transcription (43). Gel shift assays showed that MZF-2 associates at specific hTERT promoter binding sites at nucleotides
514,
543,
620, and
696, and when MZF-2 is overexpressed, it results in downregulation of hTERT and telomerase activity, suggesting MZF-2 is a negative regulator of hTERT (43). Unfortunately, these general screening techniques provide little insight into the mechanisms of repression (Table 3.1).

3.2.1.11

Perspective

A number of the repressors discussed here act directly at the hTERT promoter to suppress its activation. Those that act indirectly, such as the c-Myc and p27kip1 examples described above, warrant further study. Determining how both direct and indirect acting repressors inhibit telomerase will likely provide new targets in the treatment of cancer. Alternatively, repressing the negative regulators may provide means to extend lifespan in normal human cells.

3.2.2

Positive Transcriptional Regulators

3.2.2.1

c-Myc

The protooncogene c-Myc has been implicated in the activation of telomerase in a variety of normal human cells (12, 42, 44) and is one of the most well-studied hTERT regulatory components. As discussed in the previous sections, inhibiting c-Myc’s ability to bind to the hTERT promoter is a major mechanism of many hTERT repressors. c-Myc transcriptionally transactivates the catalytic subunit hTERT in normal human mammary epithelial cells. c-Myc activation overlaps with telomerase expression in normal tissues, which may implicate telomerase in tumors where c-Myc is activated. This may mean one of two things: (1) telomerase activity reflects oncogene activation (i.e., correlative but not mechanistically related) or (2) telomerase activation by oncogenes contributes to the formation of the tumor (13). In cervical cancer lines, further correlation was drawn between c-Myc expression and telomerase activity: hTERT and c-Myc expression were concordant in most malignant samples, although there was no correlation between histopathology or prognosis (14). c-Myc’s role in transcriptional activation of hTERT and increased expression of telomerase may be a key factor in c-Myc’s ability to immortalize and transform cells. Activation of hTERT transcription by c-Myc is a direct effect. The hTERT promoter contains two primary E-box-binding sites at positions
34 and
242, (Fig. 3.1) both of which are related to c-Myc binding (15). Although c-Myc induces hTERT in rat embryo fibroblast cells, hTERT cannot induce c-Myc’s transformation abilities, a result that does not exclude the possibility that the interaction helps to maintain immortalization in tumorigenic cells (16). The interaction between c-Myc

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and telomerase is likely to be important in tumorigenesis, yet, it may not be a direct result of c-Myc’s oncogenic activities.

3.2.2.2

Survivin

Survivin, a member of the apoptosis inhibitor family, upregulates hTERT expression, therefore positively regulating telomerase activity. By observing its effects on Sp1 and c-Myc, Survivin increases the DNA-binding abilities of the two proteins to the hTERT promoter and increased their phosphorylation. It was concluded that Survivin also increases Sp1- and c-Myc-dependent hTERT transcription (45). Through an indirect action on the hTERT promoter, Survivin is able to enhance the transcriptional activation of hTERT through Sp1 and c-Myc.

3.2.2.3

hALP

A cDNA library derived from HeLa cells was screened for hTERT transcriptional regulators using an hTERT promoter-based yeast one-hybrid assay (46). This genetic screen resulted in the identification of a clone that resembled a GNAT family protein (N-acetyltransferase domain) and was named hALP (human Nacetyltransferase-like protein). Because it is able to specifically acetylate free histones in vitro, it was hypothesized that hALP regulates the activity of histone acetylation, which involves transcriptional regulation of hTERT. It is also thought that hALP may interact with Sp1, which may also contribute to its regulation of hTERT, but this remains to be experimentally determined (46).

3.2.2.4

TEIF

Another novel gene named telomerase transcriptional elements-interacting factor (TEIF) was found to interact with the hTERT promoter at nucleotide number
2 (Fig. 3.1) (47). When transfected into HeLa cells, TEIF caused transactivation of the hTERT promoter together with elevated telomerase activity. Antisense expression of TEIF downregulated telomerase activity in HeLa cells and suppressed tumor formation in nude mice (47). Together, these data suggest a role for TEIF as a positive regulator of hTERT transcription.

3.2.2.5

JNK Pathway

In ovarian epithelial cells, the transcription factor c-Jun activated hTERT transcription via the JNK pathway. JNK, a downstream target of PI3K, likely activates transcription of the hTERT promoter through phosphorylation and activation of cJun, whereby c-Jun can functionally bind the AP1 sites in the hTERT promoter

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(48). However, this is in stark contrast to studies done with distal AP1 sites, which showed a repressive effect when c-Jun is bound to the hTERT promoter (41). When JNK was expressed in telomerase-negative cells, it induced telomerase activity. This was further supported by the introduction of the JNK antagonist JIP (JNK inhibitor protein), which when expressed in the same cells with JNK suppressed the activation of telomerase by JNK. Even though the authors concluded that c-Jun is a downstream target for hTERT activation, it seems likely that this effect is also mediated by another JNK-activated transcriptional protein (48). This effect may also be cell type-specific, as AKT (another kinase in the PI3 Kinase pathway) has been found to posttranslationally regulate hTERT in a number of systems, which will be discussed in more detail below.

3.2.2.6

HPV 16 E6

One of the major effects of HPV Type 16 E6 infection on cells is immortalization via hTERT activity (49–51). To accomplish this, the E6 oncoprotein was shown to bind to an associated protein E6-AP, which forms the E3 ubiquitin ligase. This ligase then binds and targets p53 for proteasome degradation, while increasing the expression of the catalytic component of telomerase (52–55). To define the mechanism of the E6-mediated telomerase increase, a yeast-two hybrid screen identified the protein NFX1. There are two isoforms of NFX1: NFX1-123, which coactivates hTERT with c-Myc, and NFX1-91, which may potentially act as a destabilized repressor (49). This finding was further supported by a synergistic assay of both knockdown and overexpression of the NFX1 activating protein, correlating with a respective decrease or increase in hTERT activity (56).

3.2.2.7

STAT3

STAT3 was first examined as a possible marker of tumor immortalization (57). Downregulation of STAT3 by siRNA in telomerase-expressing tumor cells results in a decrease in hTERT expression independent of c-Myc function. Stimulation of STAT3 with growth factor PDGF and cytokine IL-6 both have a positive effect on STAT3 and hTERT’s expression. STAT3 directly binds to the hTERT promoter at two locations (
1,587 and
3,308) to regulate its expression in a positive manner (Fig. 3.1) (58).

3.2.2.8

EWS-ETS

Another specific regulator of hTERT was found in Ewing’s Sarcoma through the EWS-ETS oncoproteins, which were shown to activate hTERT (59). hTERT is highly expressed in Ewing’s Sarcoma tissue and the hTERT promoter was found to be directly activated by the Ewing’s Sarcoma-associated fusion protein, EWS-ETS

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(EWS-ER81) at
23 (Fig. 3.1) (59). This regulator, although specific to hTERT, is also specific to Ewing’s Sarcoma patients, and therefore will be limited in its therapeutic implications.

3.2.2.9

HIF1a

Hypoxia inducible factor 1 (HIF1a) regulates oxygen homeostasis in cells (60) and is linked to tumor development in cells that are exposed to a hypoxic microenvironment (61–63). An examination of the cellular response to hypoxic conditions of the human placenta during the first trimester suggests HIF1a is involved in activation of hTERT, predominantly via HIF-1a binding directly to two putative responsive elements in the hTERT promoter (at nucleotide numbers +44 and
165) (Fig. 3.1) (64). Implicated in response to trophoblast growth, HIF1a could also be associated with resistance to chemotherapeutic treatment when a hypoxic condition exists in cancer cells. A choriocarcinoma cell line grown under hypoxic conditions resulted in high HIF1a levels, correlating with an upregulation of hTERT (65). In cervical cancer cells, hypoxia activates telomerase, again, correlating with an increase in HIF1a protein (64). HIF1a is a key factor in tumor progression; its role in the activation of hTERT further implicates telomerase in the tumorigenesis pathway and provides an additional therapeutic target, although to date the data are predominantly corollary (Table 3.2).

3.2.3

Both Positive and Negative Regulators

3.2.3.1

E2F

Some binding elements, such as E2F, both activate and repress hTERT activity. E2F-1 was implicated in the repression of the hTERT promoter through the identification and mutation of putative E2F-1 binding sites proximal to the transcriptional start site of the hTERT promoter (at
68,
98,
174, and
251) (Fig. 3.1). Also, overexpression of E2F-1 repressed hTERT promoter activity in a Table 3.2 Positive transcriptional regulators Protein Regulatory role c-Myc Activator, E-box binding, Max heterodimer Survivin Activator, Sp1/c-Myc binding hALP Activator, direct promoter interaction TEIF Activator, direct promoter interaction c-Jun Activator, direct promoter interaction E6/NFX1 Activator, p53 degradation/promoter interaction STAT3 Activator, direct promoter interaction EWS-ETS Activator, direct promoter interaction HIF1a Activator, direct promoter interaction

Reference (12–16, 42, 44) (45) (46) (47) (41, 48) (49–56) (58) (59) (64, 65)

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variety of human tumor and immortalized cell lines. In carcinoma cells, E2F-1 can rescue the downregulation of hTERT caused by Rb, enhanced by cyclin-dependent kinases (33). E2F-1 could be inhibiting the binding and activation of hTERT though direct competition for binding with Sp1, a known activator. In normal somatic cells, however, E2F-1 was found to activate hTERT transcription through a noncanonical E2F site present in the hTERT promoter. Additional E2F proteins (E2F2 and 3) repressed hTERT activity in 293, HeLa, and U2OS tumor cell lines, while E2F4 and E2F5 did not. Yet all were found to activate the hTERT promoter in normal human somatic cells (IMR90, HFF, and WI38) (66). E2F-1 can promote and inhibit tumorigenesis, activating and repressing gene targets, respectively, similar to its role in hTERT expression, the regulation of which may provide an explanation of E2F-1’s regulation of other genes (67, 68). 3.2.3.2

Sp1

Another protein implicated in the dual regulation of hTERT is Sp1. Interactions with specific accessory factors guide Sp1’s regulatory effect on hTERT (12, 26, 45, 46, 69–72). The Sp1 transcription factor positively regulates hTERT transcription by binding two canonical and three degenerate sites located between the two E-boxes, 110 bp upstream of the transcription start site in the hTERT promoter (at
82,
112,
132,
161,
179, and
950) (Fig. 3.1). Sp1 interacts with c-Myc to activate hTERT. The E-box of the core promoter of hTERT that binds Myc/Max and the GC-box, which binds Sp1, are required for transactivation. When Sp1 sites were mutated, Myc/Max had little effect on transactivation, indicating that there is a positive correlation between c-Myc and Sp1’s effect on the transcriptional regulation of hTERT (12). In addition, Sp1 positively regulates hTERT in the latency-associated nuclear antigen of Kaposi’s sarcoma-associated herpes virus (LANA-KSHV). hTERT promoter activity is upregulated in KSHV by an interaction of Sp1 with LANA. LANA acts as an oncoprotein in part because it interacts with Sp1 and aids in cellular immortalization (69). When bound to Sp3, Sp1 negatively regulates hTERT expression by binding tightly to the hTERT promoter and interacting with histone deacetylase (HDAC) (70). This hetero-dimerization functions as a recruitment method of HDAC to the hTERT promoter, which results in histone deacetylation, chromatin condensation, and transcriptional silencing of the hTERT gene. Analysis of deletion constructs suggests a region of the hTERT promoter that uses HDAC-mediated transcriptional repression in IMR90, WI38, and HFF cells. Proteins Sp1 and Sp3 bind to this repressive element and subsequent mutations of their binding sites results in an increase in hTERT promoter activity (70). 3.2.3.3

USF1 and USF2

The upstream stimulatory factor (USF) 1 and 2 proteins are yet another set of proteins found to regulate hTERT, but through different mechanisms (73, 74). They associate with the two E-boxes located in the hTERT core promoter (Fig. 3.1);

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however, their regulatory actions are highly cell‐type specific. In immortalized 293T cells, USF1 and USF2 activate the hTERT promoter, where expression of both proteins enhances hTERT promoter activity, possibly acting as a heterodimer. In immortalized ovarian cancer cells, USF1 and USF2 have stronger binding affinity than Myc/Max. Further testing in additional immortalized and nonimmortalized cell lines indicates that USF1 and USF2 bind the hTERT E-boxes regardless of cell line and bind the upstream and downstream E-boxes both in vitro and in vivo (73). However, USF1 and USF2 only activate hTERT transcription in immortalized cells. This difference may indicate that USF1 and USF2 are basal repressors of hTERT expression in normal somatic cells by physically blocking c-Myc access to the E-boxes (73). USF1 and USF2 are negative transcriptional repressors of hTERT in oral cancer cells, similarly through binding of the E-box rather than direct interaction with c-Myc or Mad (74). USF is expressed at lower levels in cancer cells than in normal cells, suggesting that its involvement in telomerase regulation is less in tumorigenic cells, thereby potentially contributing to elevated telomerase activity in cancer cells.

3.2.3.4

p73

The p73 protein is similar to p53 in its ability to suppress both tumor formation and hTERT transactivation through direct interaction with Sp1 (71, 72). However, one conflicting study identified an interesting relationship between p53 and p73. The coexpression of both a and b isoforms of p73, with p53, relieves p53’s repressive ability, allowing for activation of hTERT. This p53 suppression requires p73’s DNA-binding ability and p73’s activation of the E3 ligase, HDM2 (75). In breast tumor cells, p73 inhibition leads to decreased hTERT expression and telomerase activity. It is possible that p73 acts to abrogate p53’s activity and therefore regulates hTERT expression (75). Unfortunately, this study is largely corollary with little direct experimental evidence for p73 involvement in hTERT gene regulation (Table 3.3).

Table 3.3 Both positive and negative transcription regulators Protein Regulatory role E2F1 Repressor, GC-box binding (cancer) Activator, non-canonical binding (normal) Sp1 Repressor, heterodimerize with Sp3 Activator, GC-box binding/LANA interaction USF1/2 Repressor, E-box binding (normal) Activator, E-box binding (cancer) p73 Repressor, Sp1 interaction Activator, p73b expression

Reference (33, 66–68) (12, 69, 70) (73, 74) (71, 72, 75)

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3.2.3.5

Perspective

The identification of transcriptional regulators of telomerase provides numerous potential targets for chemotherapeutic and anti-aging interventions. The ability to activate or silence hTERT expression suggests that a key component of cellular immortalization can be controlled. In parallel, many of the genes found to regulate hTERT have other major cellular targets, making the inhibition or activation of these genes nearly impossible, while maintaining normal cellular function. Clearly, understanding the transcriptional regulation of telomerase will yield more appropriate targets for blocking telomerase in cancer cells (inhibit an activator or activate a repressor) and/or turning on telomerase in normal cells (inhibit a repressor or activate an activator).

3.3

Posttranslational Regulators

Although many of the proteins that regulate telomerase affect its transcriptional expression, there are a number of proteins that act posttranscriptionally or posttranslationally on telomerase. These proteins act through modifications, such as phosphorylation or ubiquitination, complex assembly (with hTR and TP1), allowance of telomere access, or altered subcellular localization. The posttranscriptional regulators may allow telomerase to be reversibly regulated: inactive in normal somatic cells but activated in cancer or immortalized cells. These mechanisms may be in response to DNA damage, genetic instability, or induced by a variety of responses to the tumor microenvironment (including hypoxia and vascularization). Much remains to be elucidated about the mechanisms for regulation of telomerase, but the regulators discussed here give initial clues. One of the proteins found to specifically interact with the hTR component is TP1, which was initially thought to eventually provide a direct means of inhibition by blocking its interaction with hTR (76). However, to date, no regulatory function has been identified for TP1. The proteins discussed in the remainder of this chapter provide other means for regulating telomerase which, although indirect, may prove effective.

3.3.1

Kinases

3.3.1.1

PP2A and Akt

As is the case with most proteins, hTERT is subject to both positive and negative regulation through phosphorylation. Through the discovery of a number of proteins involved in phosphorylation and dephosphorylation of telomerase, many pathways involved in the activation of telomerase have been identified, such as the Akt/PI3 Kinase-mediated signaling pathway. Initially, protein phosphatase 2A (PP2A) was

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found to inhibit telomerase activity in the nuclear fraction of human cancer cells, which was reversed by endogenous protein kinases, suggesting that telomerase phosphorylation and dephosphorylation may act as a switch to regulate telomerase activity in the nucleus of cancer cells (77). Putative Akt kinase consensus sequences (at nucleotides 220–229 and 817–826) are located in hTERT (at amino acids 220– 229 and 817–826). Treatment of a melanoma cell line with okadaic acid (a PP2A inhibitor) or growth factor deprivation resulted in activation of Akt and enhanced telomerase activity, suggesting that Akt kinase activates, either directly or indirectly, hTERT through phosphorylation. Treatment with wortmannin, a PI3KAkt kinase inhibitor, resulted in downregulation of hTERT phosphorylation and telomerase activity (78). The PI3K pathway (i.e., involving Akt-regulating telomerase activity) has been identified as a regulatory factor in other cell lines as well (79, 80). In a human multiple-myeloma cell line, cytokines such as interleukin-6 (IL-6) and IGF-1 upregulate telomerase activity without altering the hTERT expression, likely as a result of P13k/Akt/NFkB signaling where NFkB regulates transcription and Akt regulates posttranscriptional phosphorylation to protect against apoptosis (81). Akt may also influence other telomerase regulatory proteins including the association of the chaperone protein hsp90 with Akt and hTERT, suggesting that the complex formation of hTERT and hsp90 (82) includes Akt. Inactivation of telomerase by the hsp90 inhibitor novobiocin also disrupted the Akt/hsp90 interaction, causing inactivation of Akt and telomerase (83). The same Akt effect was not observed with treatment of geldanamycin, which binds the N-terminal ATPase domain of hsp90, inhibiting its chaperone function (84), although telomerase activity is inhibited (82). The explanation for the difference between novobiocin and geldanamycin may be as simple as cell-type differences or more likely, that the C-terminal hsp90 binding by novobiocin disrupts the Akt/Hsp90 interactions while the N-terminal geldanamycin binding does not.

3.3.1.2

PKC

The PKC inhibitors bisindolylmaleimide I and H7 inhibit telomerase activity through an action specific to PKCa, b, g, d, e, and z, thereby implicating PKCs in the negative regulation of telomerase activity (85–88). PKCa interacts with both the hTEP1 peptide and hTERT. When dephosphorylated by PP2A, hTEP1 was rephosphorylated by PKCa, and the hTERT interaction with hTEP1 increased, suggesting that PKC also mediates the phosphorylation of hTERT through hTEP1 (89). However, because of the indirect nature of these findings, these results warrant further examination. If real, they, along with the evidence from Akt interactions, suggest that telomerase exists in two forms: phosphorylated and dephosphorylated, relating to its active and inactive form, respectively. PKCy, in human T lymphocytes, acts through the NFkB pathway to activate hTERT expression (90). This pathway has been linked to hTERT regulation through TNFa (discussed later), as well as other protein kinase pathways (e.g., the c-Jun pathway discussed earlier).

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c-Abl and KIP

The c-Abl protein tyrosine kinase, which is activated by DNA double strand breaks, associates directly with hTERT (91). Exposure to ionizing radiation resulted in increased tyrosine phosphorylation of hTERT through a c-Abl-dependent mechanism in MCF-7 breast cancer cells, whereas cells expressing a kinase inactive form of c-Abl had no effect on hTERT phosphorylation. Telomerase function was inhibited in MEF cells expressing functional c-Abl, with a significant increase in telomerase activity in inactive c-Abl cells (91). This finding points to DNA damage signals regulating telomerase activity. KIP (kinase interacting protein), a DNAPKcs-interacting protein that binds to the upstream kinase domain of DNA-PKcs (92), interacts with hTERT in vitro and in vivo. Presently, it is unknown whether KIP acts directly or indirectly to regulate hTERT, but it appears that KIP stimulates telomerase activity and telomere-length maintenance (93). 3.3.1.4

MAPK

In solid tumors, Hypoxia, a classic characteristic of solid tumor microenvironment, upregulates telomerase activity in a serum and pH-dependent manner. This upregulation correlates with activation of MAPK expression (94). Hypoxia has also been identified as a factor in the regulation of hTERT transcription through HIF1a (as discussed earlier) (64, 65). The involvement of the MAPK cascade in hypoxia and the subsequent upregulation of telomerase suggest that there are multiple factors within the hypoxic environment of solid tumors that regulate telomerase activity (Table 3.4). 3.3.1.5

Perspective

It is clear that in a variety of cell types, the phosphorylation and status of hTERT regulates telomerase activity both directly and indirectly. As described here, a variety of different kinases physically associate with hTERT, some of which cause phosphorylation. Unfortunately, not all the kinases that bind hTERT result in phosphorylation. Understanding the mechanisms of how kinases regulate hTERT

Table 3.4 Posttranslational regulators of telomerase: kinases Protein Regulatory role PP2A Repressor, dephosphorylation PKC Activator, phosphorylation, different isoforms Akt Activator, phosphorylation, through PI3K pathway c-Abl Repressor, dephosphorylation KIP Activator c-Jun Activator, through JNK pathway MAPK Activator, response to hypoxia

Reference (77) (85–90) (78–81, 83) (91) (93) (41, 48) (94)

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function will be critical in identifying important telomerase inhibitory pathways relevant to phosphorylation.

3.3.2

Ligases

3.3.2.1

MKRN1 and Smurf2

Ubiquitin ligases including MKRN1 and Smurf2 are negative regulators of hTERT. The Makorin RING finger protein 1 (MKRN1) binds hTERT and mediates the ubiquitination of hTERT, acting as an E3 ligase (95). When the hTERT and hsp90 complex is altered (as with the treatment of geldanamycin), hTERT is ubiquitinated and degraded in a proteasome-mediated manner. MKRN1 enhances hTERT ubiquitination in the absence of geldanamycin treatment, suggesting that MKRN1 functions as an E3 ligase to aid in ubiquitination of hTERT in the nucleus when hsp90 is intact. MKRN1 may also regulate hTERT by causing changes in its expression and activity. For example, continued expression of MKRN1 decreased the expression of telomerase (95). Smurf2 is also an E3 ubiquitin ligase, which has previously been implicated in ubiquitination of Smad-mediated TGF-b signaling (96) that, when upregulated, produces a telomere-dependent senescence phenotype. In fibroblasts, Smurf2 is upregulated by telomere shortening that occurs as cells enter replicative senescence, When Smurf 2 is overexpressed at similar levels observed during replicative senescence, hTERT immortalization of fibroblasts is reversed. Smurf2 induces senescence in early passage fibroblasts, not by its E3 ligase activity, but by a novel protein–protein interaction with one or more proteins (not explored in this study), acting through either the p53 or pRb pathway (97).

3.3.2.2

E6/E6AP

The induction of the hTERT promoter requires both E6 and the E6AP ubiquitin ligase proteins, which may reflect a need for the ubiquitination of other associated proteins in the core promoter region of hTERT (98). The role of E6AP at the hTERT promoter is presently unknown, but it is hypothesized that it cooperates with E6 and directs specific proteins to be ubiquitinated (49, 98). As mentioned earlier, E6AP ubiquitin ligase represses telomerase through an interaction with NFX1-123 and is required for transactivation of the hTERT promoter through an interaction with NFX1-91 (49). NFX1-123 is coactivated with c-Myc at the hTERT promoter while NFX1-91 represses the hTERT promoter. The fact that NFX1-91 acts as a repressor of hTERT by binding the proximal promoter led to the examination of whether E6 targets NFX1-91 for degradation to relieve repression of hTERT. It was subsequently shown that derepression of hTERT transcription was

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Table 3.5 Posttranslational regulators of telomerase: ligases Protein Regulatory role Smurf2 Repressor, through telomere shortening MKRN1 Repressor, ubiquitination E6/E6AP Repressor with NFX1-91, Activator with NFX1-123

Reference (97) (95) (49, 98)

stimulated by the ability of E6 to target NFX1-91 to the ubiquitin pathway. NFX191 stability was significantly decreased in the presence of E6, with the reduction of NFX1-91 protein levels being proteasome-dependent. NFX1-123 remained stable in the presence of E6, suggesting a preferential binding and destabilization of NFX1-91 by E6. NFX1-91 was highly ubiquitinated in the presence of E6 whereas NFX1-123 was not. In primary human epithelial cells, E6/E6AP mediated ubiquitination and subsequent degradation of NFX1-91 induced hTERT expression and delay of senescence (49) (Table 3.5).

3.3.2.3

Perspective

It is likely that there is a dynamic interaction of telomerase-specific degradation pathways, including the E3 ligases described here. Because of its specific association with the hsp90 chaperone complex, hTERT ubiquitination and subsequent degradation is mediated by Hsp90 and could be an important mechanism for inhibiting telomerase function. In fact, activation of proteasomal pathways directed at telomerase may be critical for reprogramming cellular senescence or inducing apoptosis in cancer cells.

3.3.3

Polymerases

The family of poly (ADP-ribose) polymerases, or PARPs, negatively regulate telomerase and telomere extension through interaction with TRF1, TRF2, Tankyrase1, Tankyrase2, PARP1 and PARP2. Particularly PARP1 and PARP2, catalyze poly(ADP-ribosyl)ation as a result of DNA strand breaks from ionizing radiation, oxidative stress, and alkylating agents. Along with this, PARP1, PARP2, and the tankyrases TANK1 and TANK2 are involved in telomere regulation (100). These four members of the PARP family associate with telomeric DNA and poly (ADP-ribosyl)ate telomeric-associated proteins TRF1 and TRF2 and block their DNA-binding ability (99–102). This regulates the ability of telomerase to control telomere extension. Through its poly(ADP-ribosyl)ation activity, Tankyrase1 and Tankyrase2 are able to release TRF1 from the telomere, possibly resulting in allowance of telomerase to access the telomere (99, 100). PARP1 and PARP2 both functionally interact with TRF2 and aid in maintaining TRF2’s function at the telomere. PARP2 interacts with TRF2 and regulates its function at the telomere

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through poly(ADP-ribosyl)ation (101). PARP1 specifically localizes with TRF2 to damaged telomeric DNA, most often correlating with critically short telomeres. Once there, PARP poly(ADP-ribosyl)ates TRF2, dissociating it from the telomere so that DNA damage repair proteins can repair the telomeric DNA (102). The association of PARP1 and PARP2 with TRF2 protects the dysfunctional telomeres by maintaining TRF2’s function at the t-loop, allowing DNA damage repair proteins to fix the telomeric DNA and likely preventing access by telomerase. Although specific studies of telomerase and PARP have not been conducted, the action of TRF2 in preventing telomerase access can be extrapolated to PARP’s additional protection at the telomere.

3.4

Regulators of Assembly

The high molecular weight of telomerase suggests that it is composed of not only the hTERT and hTR components, but also additional proteins that may contribute to its functionality (103). Although TP1 has been shown to interact with hTERT, there is no indication that it plays any role in the function of telomerase. However, it has been found that telomerase requires a number of molecular chaperones, or heat shock proteins, to complex with the catalytic component of telomerase to be functionally active (82, 104, 105). Heat shock proteins, associate with numerous client proteins, including protein kinases and steroid hormone receptors, and play roles in folding, assembly, stabilization, and even degradation (106). An interaction between hsp90, the co-chaperone p23, and hTERT was defined initially in vitro (82) and then validated in human cell lysates (82). This interaction appears independent of the template RNA, hTR, but directs the proper assembly of the catalytic hTERT subunit with the hTR component of telomerase. Blocking hsp90 pharmacologically results in inhibition of the assembly of telomerase, further indicating an important role for hsp90 and p23 in the formation of the telomerase holoenzyme. Elevated hsp90-associated chaperone levels (including hsp70, hsp40, p23, hsp27, and hsf-1) were observed in tumor tissue in parallel with telomerase regulation during cancer cell transformation and progression suggesting that chaperones may aid in assembly of telomerase and may also stabilize and prevent degradation of telomerase in cancer cells (107). Interestingly, hsp70 associates only with inactive hTERT in the absence of hTR, dissociating from hTERT in its active form, while hsp90 and p23 remained associated (104). Our original model for the assembly of active telomerase (104) has been modified and is shown in Fig. 3.2. In this revised model, the template subunit hTR and the catalytic subunit hTERT are not bound together. hTERT is bound to hsp90, p23, hsp70, hsp40, and HOP. Through an ATP-dependent reaction, hsp70, hsp40, and HOP dissociate from hTERT and hTERT is then bound to hTR. hsp90 and p23 remain bound to hTERT in the functional telomerase holoenzyme. The dependence of telomerase on hsp90 provides a possible target for cancer therapy. There are inhibitors of hsp90 already in use or in clinical trials as possible therapies for breast cancer and multiple myeloma, including geldanamycin

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Fig. 3.2 Proposed model for assembly of functional telomerase. Telomerase is assembled in an ATP-dependent manner through interactions with several chaperone proteins. Inactive hTERT binds hsp90 and its co-chaperone p23, along with hsp70, hsp40, and HOP. hsp70, hsp40, and HOP associate transiently and are removed when hTR and hTERT assemble, forming the active telomerase complex. hsp90 and p23 remain associated with the functional complex (See Color Insert)

Table 3.6 Posttranslational regulators of telomerase: chaperones Protein Regulatory role Hsp90 Required for telomerase assembly Hsp70 Transient association with hTERT Hsp40/ydj Transient association with hTERT HOP Transient association with hTERT p23 Required for telomerase assembly

Reference (82, 104, 107) (82, 104, 107) (82, 104, 107) (82, 104, 107) (82, 104, 107)

and radicicol (reviewed in: 108). Inhibition of hsp90 interferes with the assembly of functional telomerase by preventing proper protein folding. Geldanamycin prevents p23’s association with hsp90 by binding in the ATP pocket rendering hsp90 nonfunctional (84). Treatment of cancers in which telomerase is upregulated (virtually all cancers) may benefit from disruption of the hsp90/hTERT interaction (Table 3.6).

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67

Additional Regulators

3.5.1

Telomere-Binding Proteins

3.5.1.1

Shelterin and Associated Proteins

Telomerase must be able to access the telomere ends to promote elongation and provide unlimited proliferative potential to germ, stem, and cancer cells. The telomere is bound by a number of proteins critical for protecting the telomere ends from being recognized as a double strand breaks, and some of the same telomere binding proteins also regulate telomerase access in tumor cells. These proteins include the Shelterin complex: TRF1, TRF2, hPOT1, hRAP1, TPP1, TIN2, (109) and also Tankyrase1, Tankyrase2, Ku 70/80, Ku86, and the MRE11 complex, some of which are thought to have a regulatory effect on telomerase at the telomere (99, 100, 110–113). Proteins such as TRF1 and TRF2 specifically interact with the telomere, by coating the telomere and sequestering the ends in a t-loop formation (114–116). Others interact with TRF1 and TRF2 by blocking telomere elongation (Tankyrase1 and Tankyrase2), inhibiting telomerase (PINX1), or altering telomere structure (hRAP1, hPOT1), (Fig. 3.3) (reviewed in: 109 and 117).

3.5.1.2

PINX1

PINX1 is a negative regulator of hTERT, binding to both hTERT and TRF1 (114, 118). In vitro, PINX1 binds directly to the hTERT protein subunit, mostly at the hTR-binding domain, but also with the hTR subunit, suggesting that this regulation is not through competitive binding. In cells, the association of PINX1 and hTR is dependent on hTERT, resulting in telomerase repression (119). PINX1 binds hTERT through its TID domain. Overexpression of PINX1 or TID inhibits telomerase and induces crisis, while depletion of endogenous PINX1 elongates the telomeres and enhances telomerase activity (118). PINX1, when depleted, has also been shown to increase tumorigenicity in nude mice leading to the conclusion that not only is PINX1 a negative regulator of telomerase, but it is also a potential tumor suppressor (118).

3.5.1.3

TopoIIa

Topoisomerase IIa has been described as a telomerase-associated protein, although a functional consequence of this interaction is unknown. TopoIIa associates with telomeres and cleaves telomeric sequences in its normal function during DNA unwinding. The interaction is not mediated by DNA binding and may act to allow the t-loop to unwind so that telomerase may access the end of the telomere (Aisner and White, UT Southwestern – Dallas, personal communication).

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Fig. 3.3 Structure of the telomere and t-loop. (a) Proteins associated with the telomere and telomerase. Most of the proteins found at the telomere bind to telomeric DNA or interact directly with proteins that bind telomeric DNA. Most make up the Shelterin complex, including TRF1, TRF2, Rap1, POT1, TPP1, and TIN2. Also found at the telomere are the DNA damage‐associated proteins, MRE11, RAD50 and NBS1. The telomere-binding proteins function to protect the telomere from telomerase access but do not physically associate with telomerase. Proteins that interact directly with telomerase include Tankyrase1, Tankyrase2, Ku 70/80 and Ku 86, PINX1, and TOPOII (which has also been suggested to bind at the telomere). (b) The t-loop structure and inhibition of telomerase. The t-loop allows sequestration of the end of the telomere, preventing the end from being recognized as damaged DNA. It also prevents telomerase from acting at the telomere and adding telomeric repeats. TRF2 has a major role in forming the t-loop, as it allows the 30 overhang to invade the duplex DNA and create the t-loop structure (See Color Insert)

3.5.1.4

HP1

Overexpression of heterochromatin protein 1 (HP1) and the isoforms HP1HSa and HP1HSb results in decreased association of hTERT with the telomere because of an alteration in the telomeric chromatin. Under these conditions, telomerase is no longer able to access the telomere, resulting in a negative effect on the binding of other proteins and reduced cell growth (120).

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69

Perspective

Clearly, access to the telomere is crucial for telomerase to function properly in the cell, and critical interactions between telomerase and telomeric proteins may provide new targets for chemotherapeutics. A better understanding of telomere binding proteins may also provide insight into cellular aging: as the cell loses telomeric bases and, therefore, substrate for the binding of telomeric proteins, the accessibility of telomerase to the telomere should increase, along with cellular proliferation. Unfortunately, the ability for telomerase to access the telomere ends without regulation is also what results in the uncontrolled proliferation seen in tumor cells. A complete understanding of the mechanisms of telomeric proteins and their interactions with telomerase is imperative for understanding, and controlling tumor growth.

3.5.2

Localization

The localization of telomerase is a major factor in its regulation. For telomerase to elongate telomeres, it must be recruited to telomeric DNA. In normal cells, a large portion of telomerase colocalizes with nucleoli, but there is an intranuclear redistribution of telomerase in a cell cycle-dependent manner, likely coinciding with telomere elongation (121). Transformed cells express increased telomerase over a longer time, which may allow telomerase to act on telomeres in an environment where telomeres are critically short or there is a limit to telomerase activation, allowing for stabilization of genomic breaks or escape from crisis (121). hTR and hTERT colocalize to the nucleolus (hTERT (122, 123); hTR (124–126)), where the components likely assemble to form the active telomerase complex, which is then translocated to the nucleoplasm to act on the telomere (123). 3.5.2.1

Nucleolin

Nucleolin, a nucleolar phosphoprotein, interacts with telomerase and modifies its subcellular localization (127). Nucleolin binds hTERT in both the nucleolus and the nucleoplasm in an hTR-dependent manner at two distinct regions on nucleolin. The interaction with nucleolin did not inhibit telomerase activity but it caused hTERT and hTR to localize to the nucleoplasm and may aid in assembly of telomerase and to maintain telomerase in the nucleoplasm to be translocated to the telomere (127). There appear to be multiple ways telomerase can translocate to the nucleus, some of which will be discussed later.

3.5.2.2

TNFa

Telomerase was identified as a downstream target of NFkB (81), and hTERT interacts directly with the NFkB p65 protein (128). TNFa, a member of the NFkB pathway, can induce nuclear translocation of the hTERT-p65 complex (128). TNFa induces telomerase activity in the cytoplasm and translocates

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activated telomerase to the nucleus. Through treatment with NFkB translocation inhibitors and PI3K inhibitors, TNFa’s ability to translocate telomerase to the nucleus and induce telomerase activity was blocked, suggesting that both the activation and translocation of telomerase are regulated by PI3K/Akt/NFkB signaling pathways through TNFa (129). 3.5.2.3

14-3-3 Family

The family of 14-3-3 signaling proteins, specifically 14-3-3s (130), has been identified as hTERT-binding partners (131), and it appears that binding is required for the accumulation of hTERT in the nucleus. Ectopic expression of a dominant negative 14-3-3 resulted in hTERT accumulating predominantly in the cytoplasm instead of the nucleus. A mutant hTERT-3A that could not bind 14-3-3 also was localized to the cytoplasm. 14-3-3 facilitates hTERT nuclear localization, while CRM1/exportin-1 mediates nuclear export, which when disrupted, the localization of hTERT to the cytoplasm was impaired. The 14-3-3 interaction had no effect on telomerase activity in vitro or in cells. Other mechanisms for 14-3-3-mediated telomerase regulation (i.e., affecting telomerase’s ability to bind the telomere or facilitating the binding of other 14-3-3 binding proteins to telomerase) have been postulated but no data have been generated to support such telomerase regulation (131). 3.5.2.4

PINX1

PINX1, previously mentioned as a telomere-binding protein, has recently been identified as having an effect on the localization of hTERT (132). PINX1 cotransfected with GFP-hTERT into cancer cells resulted in a dramatic nucleolar localization of the fusion protein (as opposed to the typical diffuse pattern throughout the nucleoplasm) and colocalization of hTERT with PINX1. Importantly, nucleolar colocalization was found with endogenous hTERT, suggesting PINX1 aids in localization of hTERT. When a mutant form of PINX1, found in a large number of hepatocarcinoma patients, was coexpressed with GFP-hTERT, it no longer sequestered hTERT in the nucleolus of cancer cells. Both the wild type and the cancer-associated mutant form of PINX1 were found to cause a modest decrease in telomerase activity, suggesting the sequestering of hTERT and the inhibition on telomerase may be two distinct functions of PINX1 (132).

3.5.3

Viral Proteins

3.5.3.1

HPV E6 and E2

The HPV proteins, particularly the E6 and E7 oncoproteins, have been associated with certain anogenital cancers including cervical carcinomas. These genes in high risk HPVs (such as types 16 and 18) are necessary and sufficient to immortalize many normal cell types through suppression of p53 and Rb pathways (133, 134). In

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HPV-16, E6’s ability to induce telomerase activity occurs through its upregulation of the hTERT promoter. As discussed earlier, E6, along with the E6AP E3 ligase, has been implicated in the activation of hTERT expression, although the exact mechanism is currently unknown (49, 50). Independent studies (135–137) have explored the mechanism of activation of the hTERT gene by HPV-16 E6 oncoprotein. A 251bp promoter region of hTERT is essential for E6 to activate its transcription without altering c-Myc or Mad protein expression (135). c-Myc and Sp1 were initially thought to be necessary for the activation of hTERT by E6 (136), yet subsequent evidence shows that the E boxes in the hTERT promoter, and not c-Myc, are required for E6 activation (137). This does not necessarily mean that c-Myc is not involved in some way, but that E6 may make the promoter more accessible to c-Myc (138). Mutation of hTERT’s E box, which is bound specifically by c-Myc, USF1, and USF2 also leads to decreased activation of hTERT by E6, possibly through USF1 and USF2 blocking E6’s ability to bind to the E box. E6 allows c-Myc to replace these factors and activate hTERT (55). The HPV E2 protein also has been implicated in repression of hTERT transcription through SP1 binding sites at the hTERT promoter (139). 3.5.3.2

LMP2A

The latent membrane protein (LMP)2A is encoded by the Epstein Barr Virus (EBV) and is thought to be involved in EBV-mediated tumorigenesis (reviewed in: 140). In carcinoma cells expressing (LMP)2A, a constant reduction of hTERT mRNA coincided with decreased telomerase activity (141). Activating the (LMP)2A pathway to regulate hTERT may aid in the control of EBV‐induced malignancy. Further studies need to be conducted on endogenous hTERT to ensure the findings are biologically relevant (141).

3.5.3.3

LANA and E1A

As mentioned previously, LANA of Kaposi’s Sarcoma-associated Herpes virus targets the Sp1 binding sites of the hTERT promoter to positively regulate transcription, resulting in enhanced telomerase activity (69, 142). Another viral protein that interacts with the Sp1 sites on hTERT’s promoter is the adenoviral protein, E1A, which may have an effect on activation of the hTERT promoter, possibly through the Sp1 sites or through chromatin modification of the corepressor CtBP (143, 144). 3.5.3.4

HBV

The Hepatitis B virus (HBV) genome can integrate into the promoter region of hTERT in hepatocellular carcinoma cell lines (145–147), resulting in activation of hTERT. In vitro, the Hepatitis B virus X (HBX) gene induced hTERT transcription

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(148), while in cancer cells, there was a correlation between HBX and hTERT expression (suggesting HBX upregulated the expression and activity of telomerase) (149). This suggests a preferential integration site in the genome for HBV, possibly resulting in an induction in tumorigenesis through the activation of hTERT. 3.5.3.5

Perspective

The induction of hTERT in response to viral infection gives a clue as to the mechanism of oncogenesis associated with these viruses. There are few viruses that give rise to cancer, and findings that HPV and HBV also regulate telomerase activation explain, at least in part, how telomerase can contribute to viral-induced cancer.

3.5.4

Hormone Receptors

In cancers, particularly breast and prostate, the function of the hormones estrogen, androgen, and progesterone often have an impact on the development and subsequent treatment of cancer. Telomerase is also upregulated in these cancers, and although the increase in telomerase and hormone may only be correlative, it may also be suggestive that there is some interaction between telomerase and hormones. 3.5.4.1

ER

Estrogen activates telomerase, acting directly and indirectly on the hTERT promoter at nucleotide numbers
940 and
2,754 (Fig. 3.1), with a putative estrogen response element (ERE) in the promoter and an Sp1/ER site (through c-Myc) having less of an effect (150). In the endometrium, telomerase activity switches from a weakly active form to being highly upregulated during the menstrual cycle, suggesting estrogen may be the key regulator for telomerase in this cycle, as well as having a role in endometrial cancer (151). hTERT expression is activated through hormone treatment of telomerase-negative human ovarian epithelium cells, indicating a direct physiological stimulus to activate telomerase in normal cells (150). Estrogen acts in an Akt/PI3K-dependent pathway to phosphorylate telomerase through the estrogen receptor. In ovarian cancer cells, which express estrogen receptors (as in MCF-7 breast cancer cells), Akt is involved in the estradiol (E2) induction of hTERT expression (152).

3.5.4.2

PR

Alternatively, progesterone, which normally acts as an antagonist to estrogen’s actions in the reproductive organs, induces hTERT expression in a time-dependent manner: at 3 h after exposure to progesterone, hTERT expression in breast and endometrial cancer cell lines was induced and peaked at 12 h and then decreased rapidly (153).

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Over a 72-h period, however, progesterone inhibited estrogen’s induction of hTERT expression at the transcriptional level through a p21Waf1/Cip1-mediated pathway. Progesterone’s rapid induction of hTERT seems to be related to the MAPK cascade, although further study is necessary (153). 3.5.4.3

AR

The androgen receptor (AR) is a major target for therapy in prostate cancer. When AR is mutated, it loses its role as a protective agent, with reduced levels found in men with prostate cancer (154). Wild‐type AR blocks hTERT expression by inhibiting transactivation with endogenous wild‐type AR being recruited to the promoter in vivo. In a mutant AR cell line, hTERT is no longer repressed (155), providing yet another mode of action for chemotherapeutics directed at telomerase in prostate cancer.

3.5.5

hTR Binders

3.5.5.1

pRb and Small Nucleolar RNA-Binding Proteins

hTR, the RNA template component of the telomerase complex, interacts with pRB (156) and a number of small nucleolar RNA-binding proteins, including dyskerin (125), GAR1 (157), NHP2, and NOP10 (158). However, the effects of these interactions on telomerase regulation have yet to be elucidated. Other hTR binding proteins and their effects on telomerase and its activity are discussed below. 3.5.5.2

NF-Y and MDM2

The hTR promoter has four Sp1 sites and a CCAAT box (159). Nuclear factor Y (NF-Y) is a regulator of the hTR promoter (156) through binding at the CCAAT box, as is Sp1 and Sp3 which depend on NF-Y (160). MDM2 also binds at the promoter and may repress activation by Sp1 or pRb or NF‐Y interactions, although further studies into its direct action must be conducted (161). The hTR promoter has not been studied as extensively as the hTERT promoter and fewer proteins have been identified as regulators, mostly because hTR is ubiquitously expressed in most normal and cancer cells. 3.5.5.3

hnRNPs

The heterogeneous nuclear ribonucleoproteins (hnRNPs) are implicated in positive regulation of telomerase and telomere length. hnRNPs bind telomeric repeats and the hTR component of telomerase, and include hnRNP A1, C1/C2 and D, E, and K (162). hnRNP C1 and C2 associate with hTR and binding of C1/C2 correlates with telomerase’s access at the telomere (163–166), presumably through C1/C2’s ability

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to interact with the telomeres. This may occur through interactions with TRF1 and TRF2, or through protein–protein interaction with other hnRNPs at the telomeric DNA. hnRNP A1 is implicated in binding to the telomeric repeats in vitro, thereby preventing extension by telomerase, and it appears to act as a telomere end-binding protein that maintains the 30 overhang (167). A model proposed by Ford et al. (162) suggests how telomerase is recruited to the telomere through both telomerase and telomere-associated hnRNP proteins. The single strand-displaced region of the t-loop, known as the d-loop, associates with hnRNP A1 and may also associate with A2-B1, D, and E. Telomerase is proposed to be recruited to the telomere by a multimeric complex of hnRNPs based on whether the d-loop or hnRNPs bind directly to the 30 G-rich overhang. A model system that does not express hTR and hTERT was tested with expressed variants of hTR and hTERT that specifically affect hnRNP binding sites, further implicating these hnRNPs in regulation of telomerase (168). hnRNP A2 binds both the first 71 nucleotides of hTR and hnRNP A1, which ultimately associates with the telomeric DNA repeat sequence in vivo to aid in telomeric maintenance, along with the telomere-binding proteins TRF1 and TRF2 (169). It was proposed that hnRNP A1 simultaneously binds hTR and telomeric DNA sequence repeats, which allows for recruitment of telomerase to the telomere (170). This may also occur with hnRNP A2, though the mechanisms appear different and remain to be explored, as hnRNP A2 binds preferentially to TRF2 rather than telomerase RNA or telomeric DNA (169). hnRNP A1, when depleted in human embryonic kidney cell extracts, results in reduced telomerase activity in addition to disrupting the structure of the telomere ends. This activity was recovered after addition of hnRNP A1 and A2. hnRNP A1 and A2 are not required for assembly, as the chaperones hsp90 and p23 are, though they are required for telomerase activity. Nor are they required for recruiting telomerase to the telomere, as was suggested earlier, but rather for elongation (171). Further study of the mechanisms of hTR targeting may lead to a better understanding of telomere elongation and maintenance (Tables 3.7–3.11).

3.6

Concluding Remarks

The telomerase regulating proteins discussed in this review represent potential targets for inhibiting telomerase in the treatment of cancer. Aberrant expression of oncoproteins, tumor suppressors, and mediators of cell survival have long been implicated in tumorigenesis. However, their interactions and/or effects on telomerase provide novel telomerase inhibition strategies while simultaneously blocking additional regulatory pathways essential for cancer cell growth and survival. The dependency of the telomerase holoenzyme on the hsp90 complex also allows for targeting telomerase via chaperone inhibition, as is proving useful in on-going clinical trials. Although the activation of telomerase is a critical step in providing cancer cells with unlimited proliferative potential, expression of telomerase in normal cells

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Table 3.7 Posttranslational regulators of telomerase: telomere associated proteins Protein Regulatory role Reference TRF2 Repressor, prevent telomerase access to telomere, (115, 172, 173) sequesters T-loop TRF1 Repressor, prevent telomerase access to telomere (114, 172–174) hPOT1 Repressor, telomere elongation (175, 176) hRAP1 Repressor, telomere elongation (177) TPP1 Repressor, binds telomeric proteins (178–181) TIN2 Repressor, telomere elongation (182, 183) Tankyrase1/2 Repressor, regulates telomerase access at telomere (99, 100, 184) Ku70/80 Activator, interacts with hTR and hTERT (113, 185) Ku86 Repressor, telomere elongation (111, 112) MRE11 Telomere protection/capping (185, 186) complex PINX1 Repressor, interacts with hTERT (118, 119) PARP1/2 Repressor, interacts with TRF1/2 (187) TOPOIIa Repressor, interacts with hTERT Aisner and White, personal comm. HP1HSa/b Repressor, chromatin remodeling (120) Rad51D/54 Telomere capping (188, 189) ATM Telomere protection (190)

Table 3.8 Posttranscriptional regulators of telomerase: localization Protein Regulatory role nucleolin Alters telomerase subcellular localization PINX1 Repressor, sequesters hTERT 14-3-3 family CRM1/exportin-1 TNFa

Activator, enhances nuclear localization Mediates hTERT nuclear export Activator, translocates telomerase to nucleus, with NFkB-p65

Table 3.9 Regulators of telomerase: viral proteins Protein Regulatory role KHSV-LANA Activator, transcriptional, binds Sp1on hTERT promoter E1A Activator, through SP1 sites and CtBP LMP2A Repressor, transcriptional HPV E6 Activator, transcriptional HPV E2 Repressor, transcriptional HBV Activator HBX Activator

Reference (127) (118, 119, 132, 191) (130, 131) (131) (128, 129)

Reference (69, 142) (143, 144) (141) (49, 50, 55, 133–139) (139) (145–147) (148, 149)

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Table 3.10 Regulators of telomerase: hormone receptors Protein Regulatory role ER Activator, direct and indirect, transcriptional AR Repressor, transcriptional PR Activator and repressor, time-dependent

Table 3.11 Posttranslational regulators of telomerase: hTR-binding proteins Protein Regulatory role NF-Y Regulator at hTR promoter dyskerin Interacts with hTR GAR1 Interacts with hTR NHP2 Interacts with hTR NOP10 Interacts with hTR pRb Interacts with hTR MDM2 Repressor at hTR promoter hnRNP A1 Repressor, binds telomere, prevents access hnRNP A2 hnRNPC1/2

Repressor, binds hnRNPA1 and telomere, prevents access Repressor, associate with hTR

hnRNP B1,D,E,K

Repressor, binds telomere, prevents access

Reference (150–152) (155) (153)

Reference (156, 160) (125) (157) (158) (158) (156) (161) (167, 168, 170, 171) (165, 169) (163, 164, 166, 168) (162)

prevents cellular senescence. Therefore, induction of telomerase activators or conversely inhibition of repressors in normal cells could prevent cellular aging. Additional studies are now necessary to gain a better understanding of the transcriptional and posttranscriptional pathways regulating telomerase, which will ultimately allow for the development of novel targeted therapies in the treatment of cancer and other age-associated diseases.

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Chapter 4

Telomere Dysfunction and the DNA Damage Response Malissa C. Diehl, Lynne W. Elmore, and Shawn E. Holt

Abstract In the absence of a protective state at telomeres, chromosome ends become dysfunctional and may ultimately contribute to genomic instability. To avoid the disruption of telomere function, a host of telomere-binding and associated proteins are critically involved in maintaining structure and preventing subsequent activation of a DNA damage response. In addition to these factors, a multitude of DNA damage response proteins also normally localize to the telomere, without triggering a repair response or cell cycle arrest. Their involvement suggests that telomere maintenance and the damage response are highly interdependent for ensuring genomic integrity. In this regard, a paradigm emerges in which telomeres may actually act as sentinels for monitoring damaged DNA and mediating repair. The close cooperativity between telomere-binding proteins and DNA damage proteins is extremely important, since a compromised DNA damage response in the context of damaged telomeres leads to significant clinical manifestations, including normal aging, genome instability and premature aging syndromes, neurodegenerative diseases, and cancer. Therapeutic possibilities that have been developed for these diseases target telomeres, telomerase, or DNA damage response mediators. The interdependence of telomere maintenance and DNA damage signaling pathways is clearly evident since both entities converge toward a common goal of maintaining genomic integrity. Keywords: Telomere-binding proteins, Telomere maintenance, Dysfunction, DNA damage, Genomic instability.

S.E. Holt(*) Departments of Pathology, at Virginia Commonwealth University, 1101 E. Marshall Street, Richmond, VA 23298-0662, USA, e-mail: [email protected]

K. Hiyama (ed.), Telomeres and Telomerase in Cancer. DOI: 10.1007/978-1-60327-879-9_4, # Humana Press, a part of Springer Science + Business Media, LLC 2009 87

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Introduction

From initial work with yeast and Tetrahymena to the more modern day trend of studying mammalian telomeres, a wealth of information has been amassed on telomere biology and function. The protection provided by chromosome ends is essential for maintaining chromosomal and genomic stability. In the absence of such protective mechanisms, molecular and cellular responses trigger the activation of DNA surveillance and repair programs. To this end, the coexistence of DNA damage factors and telomere-binding proteins at the telomere may superficially appear as a paradox. However, further inspection suggests that this localization is more than mere coincidence. Proper telomere structure, including the presence of telomere-binding proteins, is crucial for avoiding dysfunction. Since DNA damage response proteins are also associated with the telomere, a complete discussion of telomere dysfunction necessitates their inclusion. The close interplay between the seemingly separate entities, telomere-binding proteins and DNA damage proteins, is extremely important, since a compromised DNA damage response in the context of damaged telomeres has significant clinical implications. Many cancers, premature aging and genome instability syndromes, and neurodegenerative diseases reflect deficiencies in telomerebased and DNA damage proteins. Because of this connection, it may be possible to develop and utilize specifically targeted therapies as treatment. The list of novel telomere-associated proteins will continue to expand and the definitive role of each protein must be elucidated accordingly. Although the players may vary, it is nevertheless evident that telomere maintenance and DNA damage signaling pathways converge toward a common goal of maintaining genomic integrity. The interdependence of telomere function and the DNA damage response remains a source of intense scrutiny as the layers of complexity begin to be appreciated.

4.2 4.2.1

Telomeres Structure, Function, and Telomere-Associated Proteins

Telomeres are specialized structures found at the ends of linear chromosomes that are distinct from the remainder of the genomic chromatin in many ways, both structurally and functionally. These nucleoprotein complexes contain noncoding DNA distinguished by the highly conserved 50 -d(TTAGGG)-30 sequence in humans. The presence of multiple guanine residues allows the telomere to inherently form G-quadruplex structures under physiological conditions (1). The telomeric region consists of an area of double-stranded DNA followed by a stretch of single-stranded DNA to produce a G0 -overhang at the 30 end. Telomeres exist in variable lengths among different organisms or cells of different origins. For instance, the average length ranges from 3 to 20 kb in humans, while inbred strains of mice have been shown to have telomeres as long as 150 kb (2).

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As dynamic structures, the primary function of telomeres is to provide a capping mechanism, and to this end, telomeres fulfill three roles (3). First, intact telomeres protect natural DNA ends from being recognized as double-stranded breaks and consequently activating a DNA damage response (4). In other words, telomeres assure that the ends of normal linear chromosomes are not subjected to unwarranted mechanisms of repair. Second, telomeres provide protection from inappropriate exonuclease degradation and third, from end-to-end fusions (4, 5). Normal cells without a capping mechanism are also vulnerable to recombination due to their highly repetitive sequence (3). Other downstream chromosomal instabilities including translocations, nondisjunction, and aneuploidy may occur at later rounds of cell division, which likely contribute to tumorigenesis (6). Thus, maintaining telomere integrity and avoiding dysfunction are critical for genomic stability. A host of telomere-binding proteins assure that telomeres do not trigger a DNA damage response, since an unfolded telomere could be sensed as a double-strand DNA break (Fig. 4.1). Some of these capping proteins include telomeric repeat binding protein factor 1 and 2 (TRF1 and 2), TIN2 (TRF1-interacting nuclear factor 2), POT1 (protection of telomeres), RAP1 (repressor/activator protein), MRE11 complex, Ku, PTOP/PIP1, and tankyrase 1/2 (7). These proteins cooperatively establish TRF2 Interactors: Mre11 /Rad50/NBS1 RAD51 WRN BLM PARP Ku70 XRCC1/XPF Apollo

Others: ATM RAD51 Ku86 DNA-PKcs tankyrase

hRAP1 TRF1

TRF1

TRF2

TIN2

TRF2

TRF2

TRF1 interactors: Ku70 Ku 80

TPP1 TRF1 POT1

TRF1

POT1

5’ TRF2 POT1

5’

3’

TRF2

3’

nucleosomes D-loop

Others: telomerase

Fig. 4.1 Proteins localized to the human telomere. Formation of the T-loop and D-loop require the assistance of several telomere-binding proteins. Of these, the shelterin complex, consisting of TRF1, TRF2, TIN2, TPP1, RAP1, and POT1, collectively contains five DNA-binding domains (two each in TRF1 and TRF2 and one in POT1), thus making it uniquely suited to recognize telomeric DNA. The role of other telomere-associated proteins in homologous recombination or nonhomologous end joining implicates DNA damage sensing and repair to be intimately connected to telomere biology. The absence or deficiency in telomere-based or DNA damage and repair proteins may lead to telomere dysfunction and ultimately, genomic instability (See Color Insert)

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the telomere loop, or T-loop, in which the single-stranded 30 overhang folds back and invades duplex DNA (8, 9). The region of double-strand invasion is referred to as the displacement loop or D-loop (8, 10). The existence of apparently evolutionarily conserved T-loops supports the presence of higher-order structure in telomeres (1, 10). Of these telomere-binding proteins, six integral proteins, TRF1, TRF2, TIN2, RAP1, TPP1, and POT1, constitute the Shelterin complex (8). Unlike other telomere-associated proteins, this complex is abundant only at chromosome ends and remains associated at the telomere throughout the cell cycle (8). Two of the shelterin components, TRF1 and TRF2, contain DNA-binding domains that recognize the double-stranded portion of telomeric DNA, and thus are necessary for the proper formation and stabilization of the T/D-loop (1). TRF2 is also found at the junction of duplex DNA invasion, around the D-loop (11). These proteins bind as preformed homodimers and are able to form higher order oligomers (8). Homodimerization of TRF1 and TRF2 is essential to their proper functioning since loss of this ability results in a failure to localize to the telomere (1). Although TRF1 and TRF2 localize to the same region of the telomere, they serve different functions. TRF1 mainly acts as a negative regulator of telomere length, partially through inhibition of telomerase. Overexpression of TRF1 results in gradual telomere shortening, while dominant-negative TRF1 causes elongation in the presence of telomerase (2). In contrast, TRF2 has emerged as the major protective factor at chromosome ends, acting as a positive regulator of telomere length (6). In support of this role, overexpression of TRF2 results in increased telomere shortening, without an associated increase in replicative senescence rate (12). This finding indicates that TRF2 acts to stabilize and protect shortened telomeres and prevents the induction of senescence. Functional inactivation of TRF2 via a dominant-negative mutant results in a loss of T-loop formation, leading to the production of nonhomologous end joining (NHEJ)-mediated end-to-end fusions. These fusions presumably arise from the cell’s inability to distinguish natural ends and broken DNA (6, 13). Additionally, recent data indicate that TRF2 may assume other (nontelomeric) cellular roles. For instance, TRF2 has been implicated in sensing and responding to irradiation-induced nontelomeric interstitial DNA damage (14). In this context, the interaction of TRF2 with numerous mediators of DNA repair implicates this protein in serving multifunctional roles. Other factors involved in telomere capping include TIN2 and tankyrase (a poly (ADP-ribose) polymerase), which act as negative and positive regulators of telomere length, respectively (10). TIN2 is believed to provide a scaffolding unit for other proteins to dock and modulate TRF1 function (2). Tankyrase is required for resolution and complete separation of sister telomeres during mitosis. Tankyrase is recruited by TRF1 to the telomere where it interacts with the acidic N-terminal domain of TRF1 (9, 15). Human RAP1 indirectly binds to the telomere via TRF2 and has been reported to have both negative and positive roles (2, 10). POT1 specifically binds single-stranded telomeric sequences of both the 30 overhang portion of the telomere and internally displaced regions in order to regulate telomere elongation (8, 16). Loss of POT1 results in loss of telomeric sequences

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and the appearance of fusions that may solicit a damage response. TPP1 negatively regulates length by interacting with POT1 and TIN2 using different domains (2) and in doing so, it holds all the components of the shelterin complex together (8). In addition to members of the shelterin complex, the lariat T-loop structure is also associated with proteins involved in DNA repair, DNA processing, and proteins that specifically bind single-stranded DNA, which will be discussed later (2). Taken together, information about both linear and three-dimensional structure and associated proteins has provided valuable insight into telomere biology and function. The plethora of mediators that aid in preserving this function and the complexity of the telomeric structure indicate that telomere dysfunction is intolerable and must be prevented. Therefore, ensuring the presence of a protected state at chromosome ends is necessary for upholding chromosomal and genomic integrity.

4.2.2

Telomerase and Telomere Maintenance

During normal replication, the discontinuous property of lagging-strand synthesis produces a stretch of unreplicated DNA between the final RNA priming event and the terminus due to DNA polymerase inaccessibility. This phenomenon, termed the end replication problem, effectively shortens telomeres by 20–200 bases with each round of cell division until a critically short length is reached (10). The progressive accumulation of short telomeres directs cells with functional tumor suppressors into senescence, characterized as an irreversible G1 growth arrested state (17, 18). To counteract the end replication problem in an attempt to avoid telomere dysfunction, the processive ribonucleoprotein enzyme, telomerase, extends these continuously shortened ends. First identified in Tetrahymena by Greider and Blackburn (19), telomerase consists of an internal RNA template (hTR in humans) that recognizes the single stranded overhang produced as a result of DNA replication and a catalytic reverse transcriptase (hTERT in humans) that uses the template to add on telomeric DNA (20, 21). Access of telomerase to the telomere may be regulated by telomeric structure as determined by the presence of telomere-binding proteins (22). Proper telomerase assembly requires the association of various proteins including the heat shock protein 90 multichaperone complex, consisting of Hsp90, p23, Hsp70, p60, and Hsp40/ydj. However, only Hsp90 and p23 have been shown to stably associate with active enzyme (23). Many client proteins, such as kinases, transcription factors, hormone receptors, and telomerase, undergo posttranslational maturation via the Hsp90 chaperone cycle to determine their cellular fate (24, 25). Because Hsp90 expression is increased in many human malignancies, Hsp90 inhibition has become a promising approach for cancer therapy since many client proteins can be simultaneously targeted and disrupted (25, 26). Although issues of drug specificity exist, Hsp90 in cancer cells have been found predominantly in a form that is bound to its client proteins, while most normal cells have uncomplexed Hsp90. This heightened formation of multiclient complexes is therefore not due to increased levels of Hsp90 itself, but rather an increased association with Hsp90

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(27). Furthermore, Hsp90 in cancer cells have a higher affinity for anti-Hsp90 compounds compared to that in normal cells, thus providing tumor selectivity via, as yet, an unknown manner (27). In addition to higher levels of Hsp90 multiclient complexes, tumor, stem, and germ cells also activate telomerase during late S phase (10). Unlike what occurs in normal somatic cells, telomerase maintains telomeres in these cells, thus providing an unlimited proliferative potential (7, 28). However, despite high levels of telomerase activity in cancer cells, telomeres are nevertheless maintained at a relatively short length. In fact, telomere length abnormalities occur early in the initiation of epithelial carcinogenesis (29), for example, in ductal carcinoma in situ (DCIS) of the breast (30). Since its activity has been detected in more than 85% of all malignant human cancers, telomerase is a logical therapeutic target and molecular marker as a measure for human tumorigenic conversion.

4.2.3

Telomere Dysfunction and Genomic Instability

The role of structure in terms of both telomeric DNA and associated proteins, as well as its inherent capping function, support the notion that maintaining telomere integrity is critical for genomic stability (5, 31). Based on the seminal work of McClintock and Muller in the 1930s and 1940s, it is clear now that broken or damaged chromosome ends are unstable and may predispose a cell to genomic rearrangements (32, 33). Since those early studies, telomeres have been identified as highly specialized structures that provide stability to DNA ends (34). Maintenance of telomere length homeostasis is essential for cell viability, without which, telomere dysfunction inevitably ensues. Telomeric DNA is maintained at a defined equilibrium, although repeats vary in number between different chromosomes and so telomeric length is quite heterogeneous. In mammalian cells, telomere-binding proteins establish this equilibrium (34). Telomere dysfunction is induced by three main mechanisms, all of which share the common theme of deregulation or displacement of telomere-binding proteins. In the first mechanism, inactivation of telomere-binding proteins disrupts telomere function and leads to telomere length deregulation, as previously discussed. In this unprotected state, chromosome ends are sensed as double-strand breaks and processed accordingly, resulting in dysfunctional telomeres (34). In the second mechanism, alterations in the telomeric sequence also induce dysfunction, since changes in the repetitive tracts render these regions incapable of recruiting telomere-binding proteins (34). This is supported by studies with Tetrahymena thermophila in which the telomerase RNA component was mutated such that abnormal telomere repeats were synthesized (35). Although telomere elongation was able to occur, these mutants displayed abnormal cell morphology, reduced cell division due to blockage during anaphase, and induction of senescence (35, 36). Similarly, yeast telomerase RNA template mutants also show aberrant telomere length maintenance due to essentially the removal of RAP1-binding sites on the telomere (37). Telomeres in these mutants either were elongated, but poorly regulated in length, or shortened until senescence was reached (37). In this scenario,

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alterations in the telomeric sequence effectively act in the same manner as inactivating telomere-binding proteins. In the third mechanism, telomere erosion leads to telomere dysfunction. The binding of telomere-associated proteins is a dynamic process in which the availability of any one binding site is a central concern (34). If proteins bind independently of each other, fewer numbers of repeats leave fewer binding sites, and thus, less protein is able to bind. On the other hand, cooperative binding would mean that the presence of fewer available tracts makes it less likely that any telomeric repeat would be bound (34). In either situation, telomeric DNA contains such few bases that telomere-binding proteins cannot associate. Yeast without normal telomerase activity undergoes progressive telomere shortening and loss of viability in the form of senescence, albeit as a delayed response (38). Mice with defective telomerase activity also share this phenotype, as well as delayed apoptosis and chromosome instability (39). These findings indicate that loss of telomerase is not the major driving factor in cell fate, but rather the resulting loss of telomere length. In accordance with the delayed phenotypes, telomere length must reach a certain critically short threshold before dysfunction ensues (34). Collectively, these mechanisms of telomere dysfunction share the property of loss of telomere-binding proteins as the underlying mechanistic cause. When checkpoint mechanisms are intact, cells respond to dysfunction by undergoing senescence or apoptosis. In the absence of such checkpoints, cells continue to proliferate leading to increased genomic instability (40). One form of instability is the end-to-end joining of chromosome ends via NHEJ, which produces circular chromosomes or dicentric chromosomes and contribute to extensive genomic instability due to breakage–fusion–bridge cycles (41). Alternatively, homologous recombination may occur between telomeric or subtelomeric regions as a secondary mechanism of telomere maintenance. In yeast, type I recombination involves amplification of subtelomeric regions and requires the assistance of Rad 54, 51, 55/57, 52, and Exo1 (5). Type II recombination involves telomeric repeat amplification and is associated with many regulators of DNA damage and repair responses, such as Mre11, Rad50, Xrs2, ATM, and human Werner’s (WRN) and Bloom’s (BLM) syndromes helicase orthologs (5). Given that telomere dysfunction ultimately leads to chromosomal and genomic instability, it is not surprising that deleterious deletions and amplifications also drive carcinogenesis (41). As a corollary, it has been shown that both telomere shortening and cancer incidence increase over time (17). Thus, telomere dysfunction is likely associated with many cellular consequences due to the fact that chromosome ends essentially maintain chromosomal integrity in the form of DNA damage protection.

4.3

DNA Damage Response

Proper cellular functioning and survival assurance depend on the ability to detect and correct DNA errors by the DNA damage response. Although straightforward in theory, in reality, it is a highly conserved complex process involving the

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coordination of many factors involved in a variety of signaling pathways to ensure integrity of the genome. DNA is under constant attack by genotoxic agents, some of which include ultraviolet light, ionizing radiation, environmental mutagens, and endogenous reactive oxygen species (ROS) (42). These ROS species preferentially target guanine residues, and since telomeres have abundant guanine bases, chromosome ends tend to be attacked resulting in DNA breaks and telomere shortening (43, 44). In this way, ROS-induced telomere shortening may contribute to the suspension of DNA polymerases during the cell cycle and elicit a similar damage response as double-strand breaks (43). Regardless of the source of damage, replication of DNA is not always completely faithful. Without a mechanism in place to counteract these insults, any part of normal DNA replication or transcription may be blocked; mutagenesis may occur; and cells may succumb to cytotoxicity in the form of death, senescence, or malignant transformation. In accordance with this, defects in this process are extremely detrimental to the cell, resulting in mutations and chromosomal aberrations, which may contribute to malignant transformation (41).

4.3.1

Components of the Response and Signaling Pathways

In its most basic format, the DNA damage response pathway consists of three main components: sensors, transducers, and effectors (42). Upon sensing DNA damage, signal transduction of the damage signal to a variety of pathways occurs, including those involved in cell cycle checkpoints, DNA repair, apoptosis, and telomere maintenance (41). Two types of kinases, phosphoinositide-3-kinase-related kinases (PIKKs) and checkpoint kinases, act to mediate signal transduction (42). These serine-threonine kinases sense DNA damage or stalled replication forks and initiate a signaling cascade by phosphorylating factors in cell cycle control and DNA repair (45). PIKKs can be further categorized into ATM (ataxia telangiectasia mutated protein or Sc and Sp Tel1), ATR (ATM and Rad3-related protein or Sc Mec1 and Sp Rad3) (42, 46), and DNA-PK (involved in nonhomologous end joining DNA repair). ATM primarily responds to ionizing radiation, which produces doublestrand breaks that modify chromatin structure (Fig. 4.2) (47). Once these lesions are sensed, ATM is activated via phosphorylation at Ser1981, Ser367, and Ser1893, and initiates a damage signal (42, 48, 49). ATM then phosphorylates Chk2 proteins localized to the area around the lesion and p53, releasing it from MDM2 (48). The multifunctional MRN complex, consisting of Mre11, Rad50, and Nbs1, is also phosphorylated by ATM. However, this complex may also play an earlier role as a sensor since it binds damaged DNA via Rad50 independently of ATM (42, 49). In this scenario, MRN and ATM may associate via protein–protein interactions, followed by recruitment to sites of double-strand breaks (50). Also, cells deficient in MRN have reduced ATM autophosphorylation (51), since interaction between these two factors is thought to stimulate a conformational change that activates ATM. The second major player, ATR, mainly responds to ultraviolet light exposure, single-stranded DNA, and stalled replication forks (46). ATR is found to be stably

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Fig. 4.2 Telomere dysfunction activates the DNA damage response. A variety of DNA insults can attack chromosome ends, leading to telomere dysfunction. These damaged ends are then recognized as DNA damage and processed accordingly. Intact checkpoint mechanisms activate a cascade of responses mediated by DNA damage proteins. The culmination of these activities contributes to cell cycle arrest followed by resolution of the damage via DNA repair or other cellular fates, including senescence and apoptosis. Replicative senescence may have evolved as part of an antitumor protective mechanism, whereby bypassing this checkpoint may lead to neoplastic transformation. Apoptosis may have evolved as a mechanism of ensuring survival of only undamaged cells and thus maintaining genomic integrity (See Color Insert)

associated with ATR interacting protein (ATRIP), which interacts with replication protein A (RPA) with high affinity (42). This is supported by observations in which RPA and the ATR–ATRIP complex colocalize to nuclear foci upon DNA damage (52). Since RPA binds single-stranded DNA, the recruitment of ATR–ATRIP signifies that these regions of DNA are sensed as damaged sites. Before effectors are activated, mediators and adaptors of the DNA damage response act as signal amplifiers by promoting the interaction between transducing kinases and effector kinases, which are especially important when low levels of DNA damage are detected (42). In terms of the MRN–ATM complex, recruitment to double-strand breaks induces phosphorylation of histone H2AX, which serves as a docking site for the Mdc1 adapter protein. This in turn quickly modifies the DSB-flanking chromatin, facilitating H2AX phosphorylation (50).

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The collective outcome of stimulation of sensors, signal transducers, and effectors is to assure an effective cellular response to DNA damage. Early acting components not only activate cell cycle checkpoints, such that DNA replication and transcription are obstructed, but also trigger DNA repair and induction of apoptotic mechanisms. Cells that arrest in the G1 phase respond to elevated levels of p53 by upregulating p21Waf1/Cip1, a cyclin-dependent kinase inhibitor that acts to suppress the kinase activity of cyclinE/cdk2 (53). A functional G1 checkpoint ensures that damaged DNA is not replicated, whereas S phase checkpoints generally monitor cell cycle progression. ATM-phosphorylated Chk2 acts upon Cdc25A phosphatase by targeting it for ubiquitin-mediated degradation. The normal Cdc25A substrates, cyclinE/cdk2 and cyclinA/cdk2, then remain inactive (54). Additionally, ATM also phosphorylates Nbs1, as well as the breast cancer susceptibility gene product, BRCA1, and SMC1. The combination of these events halts S phase progression such that DNA synthesis is delayed. Finally, damage in the G2 phase results in ATR-mediated activation of Chk1, which acts upon both Cdc25A and Cdc25C phosphatases. Phosphorylation of Cdc25C results in sequestration by the 14-3-3 protein, effectively preventing Cdc25C from activating cyclinB1/cdc2 (55). Once checkpoints have been successfully activated, cell cycle progression halts until mechanisms to relieve these obstructions are resolved. Functional checkpoints ensure cellular integrity by allowing time for the damaged DNA to be repaired. Assuming DNA repair is successful, cells may either undergo checkpoint recovery or undergo adaptation (50). In recovery, progression into the M phase is associated with inactivation of Chk1 via the ubiquitin/proteasome-mediated degradation of Claspin and Wee1, a mitosis inhibiting kinase (50), which is closely followed by accumulation of Cdc25A, subsequent activation of Cdc25C, and entry into mitosis. Recovery may be mediated by the removal or dephosphorylation of gH2AX, a phosphorylated histone tightly associated with DNA damage-induced foci (56). Adaptation, on the other hand, involves entering mitosis even in the presence of unaddressed checkpoints and unrepaired DNA damage (57). This mechanism is not completely understood and may entail elimination of damaged cells via mitotic catastrophe (58). Regardless of the final cellular outcome, it is evident that cell cycle checkpoints ensure that appropriate signals are elicited and transduced in response to DNA damage.

4.3.2

Mechanisms of DNA Repair

Cell cycle checkpoints in and of themselves are essential in detecting damage and initiating an appropriate response. Although this cellular defense mechanism is indispensable, repair programs must also be available and functional to correct these DNA lesions. Abnormal or loss of DNA repair results in the accumulation of mutations and chromosomal rearrangements and instability. The significance of such repair mechanisms is extremely relevant to normal cellular functioning since defects in these pathways are associated with malignancy or other human diseases characterized by DNA damage protein deficiencies. DNA repair in its broadest sense encompasses direct

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Fig. 4.3 Mechanisms of DNA repair implicated at the telomere. Telomere dysfunction is associated with the involvement of two major mechanisms of repair, homologous recombination (HR) and nonhomologous end joining (NHEJ). (a) Homologous recombination is error free and requires a homologous template. In the case of repetitive sequences, such as that found at telomeres, singlestrand annealing may occur in which these sequences on each strand anneal to each other. Alternatively, RAD51-mediated strand invasion may produce recombined products upon DNA synthesis, ligation, and resolution of junctions. Blue strand: strand containing DSB. Red strand: homologous template. (b) Nonhomologous end joining is usually error prone and does not necessarily involve ligation of homologous chromosomes. The annealing of nonhomologous chromosomes contributes to a cell’s mutagenic potential, as translocations are a frequent finding in cancer. Blue and red strands represent different chromosomes. See text for detailed explanation (See Color Insert)

reversal, nucleotide excision repair (NER), base excision repair (BER), mismatch repair (MMR), single-strand annealing (SSA), homologous recombination (HR), and nonhomologous end joining (NHEJ) (59, 60). For the purposes of this chapter, only the latter two mechanisms (HR and NHEJ) will be considered since they are integral components of a cell’s response to telomere dysfunction.

4.3.2.1

Homologous Recombination

Unlike yeast, both HR and NHEJ are equally important for double-strand break correction in mammals (Fig. 4.3) (61). These repair pathways are evolutionarily conserved, since homologues of the relevant genes can be found in various species

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from yeast to humans (62). A complete understanding of HR requires a discussion of the role of each protein in the repair machinery. HR in mammals requires replication protein A (RPA), DNA polymerases, and the RAD52 gene family, which includes RAD50, RAD51, RAD52, RAD54, RAD54B, NBS, and MRE11 (60, 63). Additionally, a family of RAD51-related genes exists and can further be subdivided into XRCC2, XRCC3, RAD51B, RAD51C, and RAD51D genes (60, 63). As seen in cells with the DSB repair‐defective disease Nijmegen break syndrome, there is a complete lack of Rad50 foci, implying that loss of this protein is associated with a lack of cell cycle checkpoint activation and therefore has a direct role in DNA repair (64). Human Rad51 has pairing and strand exchange activities, which are stimulated by the addition of RPA and Rad52 (65). In studies with mice, a deficiency in Rad51 is associated with chromosome loss and radiation sensitivity, highlighting its importance in normal repair processing (66). Human Rad52 directly binds DSBs, protecting exposed ends and promoting reparative ligation (60). Similar to Rad51 deficiency, defects in Rad52 confer sensitivity to ionizing radiation and are linked to abnormal meiotic recombination (60). Furthermore, Rad54, XRCC2, and XRCC3 defects lead to reduced recombinatorial rates, supporting the involvement of these proteins in HR (67, 68). Interestingly, the association of the breast cancer susceptibility gene products BRCA1 and BRCA2 with Rad51 also facilitates HR (69, 70). BRCA1 defective cells likewise show reduced recombination, suggesting that a connection between altered recombination and tumorigenesis may exist (71). Based on the collective evidence presented here, it is clear that these proteins respond to DNA damage and are essential for mediating properly timed and legitimate repair and for preventing malignant transformation. The evolutionary conservation of key regulators of the DNA damage response underscores the significance of their roles in repair. Repair of DSBs via HR, as its name suggests, requires the presence of a homologous DNA template, which makes HR a very conservative, essentially error-free process (72). Briefly, the donor template sequence is copied into the region to be repaired, thus creating an exact copy of the undamaged sequence (Fig. 4.3). The initial step is to process DSBs by a 50 to 30 exonuclease, possibly mediated by the MRN complex, such that single-stranded 30 overhangs are produced at each break (73). Following this preparatory phase, RAD52 and RPA then coat the 30 single-stranded region (74, 75). At this point, two different scenarios of DSB correction can ensue. For repetitive sequences, such as those found at the telomere, single-strand annealing is a frequent occurrence in which the overhang regions are displaced as repeat tracts from each strand anneal (76). Alternatively, strand invasion ensues in which RPA is displaced by RAD51, and single-stranded regions are subsequently coated with RAD51 to form a nucleoprotein complex (77). RAD51-mediated initiation of homology search, strand pairing, and strand exchange may be aided by RAD51B, RAD51C, RAD51D, XRCC2, and BRCA2 (65, 78, 79). In addition to these proteins, RAD54 also helps in strand invasion (80). The ends are then elongated by DNA polymerase until sufficiently long to base pair with the remaining end and joined by ligases to produce a fully repaired double strand with no or minimal errors (72).

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Nonhomologous End Joining

In contrast to HR, NHEJ does not utilize a homologous template, occurs quickly, and produces deletions, and thus is normally error prone (Fig. 4.3) (72). Like its role in HR, the MRN complex may be involved initially to prepare or clean up DSBs for further processing (59, 81). The ends of the double-strand break are recognized and bound by a heterodimer of Ku70 and Ku80, which has high affinity for exposed ends and aligns the broken DNA (60). Binding of this heterodimer in the form of a ring around the damaged DNA provides structural support during repair (82). DNAdependent protein kinase catalytic subunit (DNA-PKcs), which may act as a sensor, is then recruited to this heterodimer and establishes a synapse within the double-strand break to facilitate rejoining of the two DNA ends (83). DNA-PKcs phosphorylates DNA-binding proteins that modulate cellular responses to damage (60). The interaction of Artemis, which has endonuclease activity, with DNA-PKcs allows inaccessible structures like hairpins to be opened and further processes the ends (84). Nucleases remove, insert, or substitute a few bases at the double-strand break to make the region more amenable for ligation by the complex of ligase IV and its associated protein XRCC4 (60, 61, 85). In order for efficient ligation to occur, 30 OH and 50 -phosphate entities are added to aligned ends by polynucleotide kinase (PNK) (86). Although NHEJ usually rejoins segments of the same chromosome, ligation of broken ends from different chromosomes is also possible. This results in translocation and when combined with the alteration of bases, contributes to the cell’s mutagenic potential (61) since it could lead to activation of oncogenes or loss of tumor suppressor genes (60). The selective utilization of either mechanism may be related to when in the cell cycle the damage occurs. For instance, HR may be favored in late S and G2 phases when sister chromatids are in close proximity, while NHEJ is more prominent during the G1 phase when homologous chromosomes are farther apart (87). Thus, although HR and NHEJ operate through distinct mechanisms and differ temporally, they have an overlapping role in repair of DSBs and share a common goal. 4.3.2.3

Perspective

The ability to distinguish damaged DNA from natural ends and to faithfully replicate the entire genome is an extraordinary task. The DNA damage surveillance and repair responses are essential for detecting errors and combating insults to the genome to maintain fidelity. The coordinated actions of damage sensors, checkpoint systems, and repair mechanisms are closely intertwined as indicated by a number of common mediators, some of which exhibit interdependence. It is apparent that regulation of DNA damage and modulation of a response is a highly orchestrated process that has serious repercussions on cellular survival when defects in any of the mediators are present. Future studies on both established and novel factors in sensing, transducing, and effecting responses will only contribute to furthering the global understanding of their functions and interactions.

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4.4

Linking DNA Damage Response and Telomere Dysfunction

4.4.1

DNA Damage and Telomere-Binding Proteins: A Role in Telomere Maintenance

Based on the previous discussion of mediators of the DNA damage and repair response, it is evident that a multitude of proteins with various functions acting at different stages are required. Not only do these proteins localize to double-strand breaks, but some are also found either directly bound to or indirectly associated with the telomere. Such telomere-based DNA repair proteins include ATM, DNA-PKcs, RAD51, Ku70, Ku86, and the RAD50-MRE11-NBS1 complex (2). Furthermore, DNA processing enzymes that interact at the telomere include WRN, BLM, ERCC1-XPF1, and Apollo (2). Interestingly, the normal localization of these DNA damage and repair proteins does not trigger a DNA damage response or cell cycle arrest (6). The fact that these DNA damage response proteins and telomerebinding proteins reside at chromosome ends and their involvement in both telomere maintenance and the damage response suggests interdependency for ensuring genomic integrity (88, 89). Loss of any telomere binding or telomere-localized proteins could lead to compromised structure, function, and protective failure. Given that most of these proteins participate in repair mechanisms, the convergence of the coexistence of the DNA damage response and telomere homeostasis has been a subject of great interest. As a dynamic structure, unfolding of the T-loop is required for passing of the replication fork. At this point, the telomere may share similarities with a doublestrand break since it is in an unprotected state due to exposure of chromosome ends (2). Loss of the protective capping function leads to a variety of DNA attacks, including nucleolytic or enzymatic degradation, oxidative metabolism, and interaction with other chromosomal segments (88). In 1996, two separate groups observed that cells of patients with ataxia telangiectasia (AT), in which the ATM gene is mutated, as well as yeast cells with a disrupted open reading frame in Ku80, displayed a telomere dysfunction phenotype, including accelerated telomere shortening and increased frequency of end fusions (90, 91). Since then, major headway has been made in further elucidating the role of DNA damage and repair mediators in telomere biology, but this task is far from complete. Nevertheless, a global picture is emerging that intrinsically implicates repair mechanisms as contributors of telomere maintenance. Since telomere homeostasis is such an integral part of normal cell functioning, mechanisms that maintain appropriate length are indispensable. As previously discussed, telomere dysfunction may be brought about by disruption of telomerebinding proteins, alteration of telomere sequence, or telomere erosion. One method to counteract this dysfunction is via the reactivation of telomerase. However, telomerase is tightly regulated, expressed during development and in stem and germ cells but repressed in most normal somatic cells. Also, telomerase simply

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elongates chromosome ends without dictating or maintaining a certain telomere length (34). Survival and proliferation in the absence of telomerase suggests that there must be telomerase-independent mechanisms that act to maintain telomere integrity (92). Since the two main mechanisms of DNA repair in mammalian cells are NHEJ and HR, it is not surprising that there is a functional linkage between telomere integrity and this subclass of DNA damage response proteins. Specifically, DNAPKcs plays an important role in NHEJ but is also essential for proper telomere capping (93). Severe combined immunodeficiency (SCID) mice that have mutant DNA-PKcs exhibit end-to-end chromosome fusions and elongated telomeres (94). Likewise, the NHEJ-acting Ku70/Ku80 heterodimer affects telomere capping via its interaction with TRF1 and/or TRF2 (95, 96). Although telomere shortening has been reported in Ku86-deficient mouse models and human cell lines (97, 98), telomere elongation has been seen in Arabidopsis plants and mouse models with deletion of both telomerase and Ku70 or Ku86, respectively, suggesting that Ku acts as a negative regulator of telomere length by preventing access of telomerase to the telomeres (99, 100). Despite these differences, it is apparent that this protein plays a role in modulating telomere capping and length deregulation. Furthermore, Ku70 is believed to interact with heterochromatin protein 1a (HP1a), which is associated with heterochromatin as well as telomeres (101). This may implicate Ku70 in modulating a telomere position effect in subtelomeric regions. Besides DNA-PKcs and Ku, other NHEJ proteins such as ligase IV are associated with telomere maintenance but to a lesser degree. Ligase IV deficiency produces telomeric fusions, since it is involved in joining double-strand breaks, without signs of telomere length abnormalities (97). Also, overexpression of TRF2 and its direct interaction with ATM has been shown to effectively inactivate this kinase and abrogate an ATM-dependent damage response (102). Finally, the MRN complex, which participates in both NHEJ and HR, associates with TRF2 such that loss of this complex results in accelerated telomere shortening, as seen in patients with Nijmegen break syndrome (103). This suggests that MRN is not simply a mediator of the DNA damage response, but is also involved in maintaining a protected state at the telomere along with TRF2. Although HR is another main form of double-strand break repair in response to replication fork stalling or DNA damaging agents, it may also be an alternative mechanism of mammalian telomere maintenance (92, 104). In terms of protein expression, the colocalization of the HR protein RAD51 and TRF2 to the telomere can be visualized via immunofluorescence, where it is believed to play a role in preventing telomere dysfunction (105). A deficiency in RAD51, as well as another HR protein RAD54, in mice is associated with radiation sensitivity and poor double-strand break repair, as well as telomere shortening and increased frequency of end-to-end chromosome fusions (92, 106). In the context of HR, telomere elongation would proceed following one of the two scenarios. In the first mechanism, intertelomere HR, the 30 single-stranded end of one telomere pairs with and invades the duplex region of another telomere aided by RAD51, RAD52, and RPA. This action produces a D-loop-like structure that is stabilized by RAD54

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and undergoes branch migration and subsequent replication by DNA polymerase (92, 107). Alternatively in the second mechanism, intratelomere HR involves invasion of the 30 overhang end into the duplex region of the same telomere, forming a T-loop structure (92). Both methods use the same proteins and polymerase to catalyze invasion and elongate telomeres, respectively, but intratelomere HR may also provide a capping function through the formation of the T-loop. Evidence in support of HR in elongation and protection of telomeres is provided by the existence of an alternative lengthening of telomeres (ALT) pathway in immortalized human cell lines, approximately 10% of human tumors, and telomerase-null mouse cell lines (108–110). ALT cells display telomere dynamics in which telomeres with various critically short lengths are targeted and elongated to various lengths, which is suggestive of a more intertelomeric recombination method in mammalian cells (111). Recombinatorial proteins characteristically found in cells with undetectable telomerase activity, but exhibiting ALT activity, include RAD51, RAD52, MRN, RPA, WRN, and BLM (111). Additionally, poly (ADP-ribose) polymerase (PARP), which binds single- and double-stranded breaks and interacts with p53 to modulate BER, may also be involved in ALT since loss of PARP induces an ALT-like telomere phenotype (112). Moreover, PARP binds with high affinity to TRF2, making the telomere connection to DNA repair even more sound (10). PARP-deficient cells display increased numbers of chromosomes with undetectable telomeric signal, thus implicating this protein in telomere maintenance (113). The presence of these proteins at the telomere suggests that the ALT pathway is another form of telomere dysfunction. Although ALT is more of a secondary pathway of telomere maintenance, it nevertheless needs to be considered as a potential alternative DNA damage response. In addition to proteins involved in NHEJ and HR, other repair proteins are implicated in telomere function. ATM-defective cells exhibit telomere dysfunction in the form of extrachromosomal telomeric fragments and accelerated telomere shortening (114). TRF2 also interacts with the ERCC1-XPF endonuclease, which processes the 30 overhang during nucleotide excision repair and recombination (115). Mutations in XPF are associated with UV sensitivity and ERCC1-XPF deficient cells show increased levels of telomeric DNA-containing double-minute chromosomes (TDMs), a relatively new marker for telomere dysfunction (116). Additionally, TRF2 interacts with the Apollo 50 -to-30 exonuclease that aids in telomere protection (117). Although the list of relevant proteins discussed herein is not exhaustive, it nevertheless provides indisputable data on the linkage between telomere biology and the DNA damage response.

4.4.2

Resolution of the Paradox

Telomere-binding proteins and DNA damage proteins both reside at telomeric DNA, but the exact purpose of this coexistence is unclear. It is possible that mechanisms of repair act in two different capacities: first, to repair DSBs at internal

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chromosome sites, and second, to repair at telomeres. However, it seems more likely that repair mechanisms, its associated proteins, and telomere-binding proteins act cooperatively to establish a balance in chromosomal integrity. This could involve a competition between repair proteins and, for example, telomerase action at the telomere (89, 118). In this manner, telomerase elongates telomeres as a way to ‘‘heal’’ shortened or ‘‘broken’’ DNA ends. Unregulated telomerase activity could lead to karyotypic instability, since it may respond to internal DSBs as shortened telomeres. At this point, repair proteins are signaled to restrict telomerase action to chromosome ends (89). In support of DNA damage and telomere maintenance linkage, TRF2 has been shown to translocate to sites of double-strand breaks and act in a telomereindependent fashion (14). This finding suggests that TRF2, as perhaps other telomere-binding proteins may by a part of a global cellular response to damage. Additionally, unlike mammalian cells, Drosophila telomeres are not maintained by telomerase and do not contain structures such as T-loops. Rather, Drosophila utilizes DNA damage proteins in order to maintain telomeres, many of which have orthologs in mammalian systems, such as Ku, ATM, and the MRN complex (119). Finally, mice with defective TERT present with dysfunctional telomeres, are sensitive to ionizing radiation, and repair double-strand breaks with slower kinetics (120). In a similar fashion, telomere dysfunction in cells of cancer patients may predispose these individuals to sensitivity to DNA damaging agents (121). Although the primary function of telomeres is to provide protection for chromosome ends, it has been suggested that telomeres may actually act as sentinels for monitoring damaged DNA and mediating repair (122). It may be possible that telomeres endure the majority of induced damage so that the remainder of the genome maintains fidelity. Taken together, a relationship between telomere maintenance and the DNA damage and repair response emerges that serves dual purposes. In one capacity, this relationship ensures the formation of a functional telomere structure, which is dependent on the presence of elements of the DNA repair machinery. In this manner, damage-associated proteins are required for telomere protection and legitimate replication. In a second capacity, this interrelationship reflects a surveillance system in which dysfunctional telomeres are recognized by the DNA damage response and subsequently processed in the same manner as double-strand breaks (123). Collectively, these observations support an integrative model of telomere function and DNA repair in which telomere maintenance is intimately connected with components of the DNA checkpoint and damage machinery to ensure genomic integrity (124).

4.4.3

Cellular Fates

Aside from deficient telomere-binding or associated proteins, normal cell cycling can also lead to telomere erosion and subsequent instigation of a damage response. The cooperation of telomere structure and function and the DNA damage response have significant implications for growth arrest, senescence, and apoptosis. Progressive

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telomere shortening destabilizes the T-loop structure and predisposes the chromosome to deleterious uncapping events and dysfunction (125). Although other forms of senescence exist, loss of protection from telomeres can lead to replicative senescence in which cell growth is irreversibly arrested, after a certain amount of population doublings, i.e., once the cell has reached the Hayflick limit (126–129). This supports the thought that cellular senescence may have evolved as part of an antitumor protective mechanism, and that bypassing this checkpoint leads to neoplastic transformation (130). In addition to telomere-dependent growth arrest, stress-induced senescence is triggered by DNA damaging agents, such as irradiation, chemicals, and oxidative stress, all generating double-strand breaks, some of which occur at the telomere (131–133). In this way, DNA damage responders can sense aberrations in telomeres and halt proliferation to avoid the accumulation of detrimental mutations (134). Activation of the DNA damage response via either growth arrest-inducing mechanism triggers a cascade of signals mediated by ATM, ATR, and DNAPKcs (as discussed above). These kinases also phosphorylate histone gH2AX on Ser139 at the site of DNA damage, followed by the recruitment of 53BP1, the damage checkpoint protein MDC1/NFBD1, and NBS1 to the locus of damage (129). Collectively, these molecules assemble into DNA damage-induced foci that initiate senescence (129). In support of this, the fibroblastic cell strains MRC5 and BJ display increased staining of gH2AX and 53BP1 as cells enter a senescent state, while those same cells immortalized by hTERT show relatively low signal intensity (135). In addition to these two markers of DNA damage, MDC1 and NBS1 are also detectable and phosphorylation of p53, Chk1, and Chk2 were observed (135). ATM-activated p53 acts to induce cell cycle arrest in response to senescence‐associated DNA damage foci. The level of p21, a cyclin-dependent kinase inhibitor, also increases upon senescence and may contribute to cell cycle arrest (Fig. 4.2) (129). Given that DNA damage foci accumulate at double-strand breaks, it is necessary to address whether these foci also localize to dysfunctional telomeres. The creation of a fibrosarcoma cells using the dominant negative TRF2DBDM allele produces uncapped telomeres that induce growth arrest (136). Telomeric DNA, as well as TRF1, was bound to gH2AX, 53BP1, and NBS1, suggesting that unprotected telomeres may activate a DNA damage response without actual chromosomal breakage (135, 137). However, DNA damage-induced foci, as measured by gH2AX, did not colocalize with TRF2 (138), which is in agreement with a model in which loss of TRF2 produces uncapped telomeres and the subsequent formation of damage-induced foci at chromosome ends (139). While illegitimate telomere exposure is necessary for assembly of these foci, the mechanism by which short telomeres induce foci formation is less clear. It is possible that telomeric overhangs are first processed into blunt ends before DNA damage/senescence signaling occurs. Alternatively, telomeric overhang processing could be a consequence of the DNA damage/senescence response (129, 140). Whichever the case, these data nevertheless indicate that production of DNA damage-induced foci at uncapped or dysfunctional telomeres trigger the replicative senescence response.

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As an alternative to senescence, short unprotected telomeres may also trigger apoptosis, depending on signaling by the p53-induced DNA damage cascade (2, 123). p53-dependent apoptosis is mediated by cell cycle checkpoints and the transcriptional activation of DNA repair factors and proapoptotic proteins, such as Apaf1 and Bcl-2 (141). On the other hand, p53-independent mechanisms of apoptosis also exist. The transcription of the p53 family member, p73, is stimulated by the transcription factor, E2F1, which is implicated in inducing cell death in the absence of p53 (142). In support of the role of p73 in apoptosis, high levels of this protein are found in cells with inactivated pRb (or unrepressed E2F1) and undergoing malignant transformation (143). Furthermore, reactivation of telomerase or inactivation of p53 and pRb (or aberrant expression of p16) allows continued proliferation by bypassing senescence (6, 144). In this scenario, cells continue to divide beyond their normal replicative capacity, producing telomeres that completely lack protective DNA. At this point, the cell enters a crisis stage marked by severe chromosomal instability and cell death (123, 134). Cells that achieve immortalization, most likely via reactivation of telomerase, also display properties of transformation, regardless of whether telomerase is activated (145). The decision to follow the apoptotic route has most likely evolved as a way to preserve genomic integrity via selective killing of damaged cells (141). In summary, cellular responses to DNA damage are triggered by various signals, but ultimately contribute to cell cycle arrest, repair, and replicative senescence or apoptosis (Fig. 4.2).

4.5

Clinical Implications

From model organisms to humans, defects in the DNA damage/repair response and telomere maintenance are associated with significant clinical defects (40). In addition to normal aging, genome instability and premature aging syndromes, neurodegenerative diseases, and especially cancer all share defects in components of the DNA damage response (Table 4.1). These disorders also exhibit a significant relationship to telomere function since dysfunctional telomeres are not an uncommon finding. Although the culpable genes or proteins have been identified in these diseases, more research is necessary to translate these findings to a clinical therapeutic setting. Table 4.1 Summary of human syndromes with direct or indirect telomere involvement Disease Defect Cellular outcome Reference AT ATM IR sensitivity, chr. fusions (40, 146, 147) ATLD Mre11 Defective MRN complex (148, 149) NBS NBS1 Defective MRN complex (40, 150) BS BLM Hyper recombination as SCEs (151, 152) WS WRN Illegitimate recombination (153–155) HGPS HA(?) Defective DNA repair (156, 157) DC Dyskerin Deficient telomerase activity (158, 159) hTR (160) AD b-amyloid Decreased mental capacity (161, 162)

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Aging

Life expectancy has increased remarkably in developed nations, but still remains a serious concern, and as such, continues to be an area of intense interest (163). Although damage to DNA has long been attributed to the aging process, recently more emphasis is being placed on telomere shortening as an adjunct component (164, 165). As such, telomere length may be used as a marker of aging and to assess the accumulation of age-related DNA damage (166). For instance, UV exposure induces DNA repair at damaged telomeres, the rate and the efficiency of which declines with age (165). Furthermore, since telomerase maintains telomeres in immortal and cancer cells, its absence in normal cells has been related to cellular aging (167, 168). The role of telomerase in maintaining proliferative capacity is evident in somatic cells transfected with telomerase, since these cells remain youthful and proliferate indefinitely without undergoing malignant transformation (167, 169–171). These data highlight the presence of a strong relationship between aging, telomeres, and telomerase (164). Even with the identification of telomerase as a promising regulator of cellular aging, defining the molecular mechanisms underlying the aging process is still in its infancy and likely involves both genetic and environmental factors.

4.5.2

Genome Instability and Premature Aging Syndromes

In addition to the normal aging process, premature aging and genomic instability syndromes also involve deficient DNA damage repair with disruption of telomere length or function. These rare syndromes, as its name suggests, are characterized by the onset of age-associated symptoms much earlier than in the nonaffected individual (172). Additionally, other age-related diseases, such as cancer, diabetes, and cardiovascular disease, are found in these patients. Although there is a multitude of genome instability syndromes with defects in DNA repair, only those with specific implications for telomere-binding or associated proteins will be discussed. Specifically, Ataxia Telangiectasia (AT), Ataxia Telangiectasia-like disorder (ATLD), Nijmegen break syndrome (NBS), Bloom syndrome (BS), Werner syndrome (WS), and Hutchinson–Guilford progeria syndrome (HGPS) all present with alterations in the aging process and are associated either directly or indirectly with telomere biology (40, 164).

4.5.2.1

Ataxia Telangiectasia and Ataxia Telangiectasia-Like Disorder

Patients with AT and ATLD display premature aging, short stature, neuronal degeneration in the form of upper and lower limb ataxia, and extreme sensitivity to ionizing radiation (146, 173). Although ATLD individuals share clinical features that are indistinguishable from AT, ATLD is considered a milder subset of AT, with

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symptoms appearing later and disease progressing slower (174). Genomic instability is prevalent on the molecular level as chromosomal translocations, mostly involving breakpoints in the T cell receptor and immunoglobulin genes (only in AT cases), and compromised cell cycle checkpoints (40, 175, 176). Since sensitivity to IR is a major feature observed in AT patients, it was found that the IR-responsive gene, ATM, is defective in AT cells. Without ATM, p53 cannot be appropriately regulated and HR is disrupted due to ineffective recruitment of RAD51 and RAD52 to sites of repair (147, 177). AT cells show lack of telomere maintenance in the form of high levels of ROS and end-to-end fusions, mostly arising from translocation events and likely contributing to the development of lymphomas and leukemias (40, 91, 173, 176). These phenomena are all linked to telomere-associated proteins, suggesting that ATM may provide protection at telomeres by preventing accelerated telomere shortening (91). In contrast to AT, ATLD arises due to a mutated Mre11 gene, in which the Mre11 protein transcript is degraded (148, 178). It is noteworthy that the levels of Nbs1 and Rad50 proteins are also reduced in ATLD cells, indicating that MRN complex assembly is affected by Mre11 deficiency (149). The localization of this complex to sites of DNA damage is important for repair since ATLD cells fail to form irradiation-induced foci (174, 179). Unlike in AT, it is not clear whether there is a predisposition to cancer in ATLD, since so few patients have been identified (174). Nevertheless, Mre11 is indispensable in the formation of a properly functioning MRN complex that elicits cellular responses to DNA damage.

4.5.2.2

Nijmegen Break Syndrome

On a cellular level, NBS shares similarities with AT and ATLD since NBS is also associated with a high frequency of translocations and characteristic impairment of response to double-strand breaks (173). A deficiency of the NBS1 gene product, nibrin, results in a failure to assemble IR-induced complex with Mre11 and Rad50 (40, 150, 180). This deficiency may affect telomere maintenance since fibroblasts from NBS individuals show accelerated telomere shortening that cannot be rescued by the introduction of telomerase (150). Furthermore, NBS1 colocalizes with TRF2 and promyelocytic leukemia (PML) bodies, which are found in telomerase-negative ALT cells (139, 181). NBS1 deficiency also results in an increase in telomere associations, in which telomeres of the same or different chromosomes are found in close proximity in metaphase spreads (182). Collectively, these data suggest a role for NBS1 in modulating telomere length to maintain chromosomal stability.

4.5.2.3

Bloom and Werner Syndromes

Both Bloom and Werner syndromes are well characterized disorders of genomic instability and premature aging, characterized by growth retardation without mental

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retardation, diabetes, and a predisposition to cancer (sarcomas in the case of WS) (183, 184). WS patients also display other age-related clinical indicators, including premature cataract, skin wrinkling, osteoporosis, and atherosclerotic heart disease, which appear in adolescence, thus decreasing maximum lifespan to less than 50 years (153, 164, 185–187). BS patients typically exhibit sensitivity to chemical agents and UV sunlight manifesting as pigmentation changes in the skin and immunodeficiency (188, 189). Both BLM and WRN have sequence similarity to the highly conserved RecQ family of ATP-dependent DNA helicases (151, 153, 154, 190). These helicases preferentially unwind G-quadruplex DNA that forms in G-rich regions like telomeric DNA (191, 192). As such, BLM and WRN bind to and colocalize with TRF2, supporting the finding that BLM is found in the nucleus as both discrete foci and in diffuse distribution, and that WRN cells show defective repair at telomeres (165, 193, 194). Although BLM is found localized to telomeric sequences, it has not been shown to actively affect telomere length (194). In contrast, WS cells also show accelerated telomere shortening with reduced doubling times and fewer cell divisions, leading to a shortened lifespan (195–197). Since telomerase is able to rescue this shortening (without fully correcting chromosomal instability), WRN could act in a protective manner by mediating the onset of senescence triggered by exposed ends, or by mediating telomerase access (40, 153, 198). While BLM and WRN additionally interact with RPA to stimulate its activity, they also associate with RAD51 and Ku70/80, respectively (10, 199–201). As such, BS cells display hyper‐recombination between homologous chromosomes and telomere associations, while WS cells show inefficient recombinatorial repair of DNA crosslinks (152, 153, 188, 190, 202–205). These aberrations lead to chromosomal instability, such as translocations and rearrangements (155). Taken together, it is evident that BLM and WRN have multifunctional roles in mediating recombination and maintaining chromosomal integrity (203).

4.5.2.4

Hutchinson–Guilford Progeria Syndrome

Premature aging is not evident outright in HGPS syndrome, but develops by the first 2 years of life (164, 206). Although the exact pathophysiology is unknown, it is thought that excessive hyaluronic acid (HA) excretion from the bladder or defective DNA repair may contribute to the disorder (156, 157, 207, 208). On a cellular level, accumulation of senescent cells has been proposed in the acceleration of the aging process in these individuals (209). As such, telomere length maintenance and telomerase activity have been implicated as regulatory factors in the pathogenesis of HGPS. In support of this, fibroblasts derived from HGPS patients are shorter than those from age-matched controls (210). Despite the uncertainty in the causative mechanisms of premature aging in HGPS, telomere maintenance is nevertheless a relevant component of cellular lifespan and must be considered in the pathogenesis of this disease.

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Dyskeratosis Congenita

Although Dyskeratosis Congenita (DC) may share features with premature aging syndromes, it is mainly a disorder of telomere maintenance (211). Since this disease affects highly regenerative cells, it is characterized by skin lesions and bone marrow failure (158, 212). Three forms of the disease exist: X-linked, autosomal dominant, and autosomal recessive, with the autosomal dominant form being surprisingly less severe (158). The culpable gene, DKC1, of the X-linked form encodes for dyskerin, a highly conserved nucleolar protein that is involved in rRNA synthesis, ribosomal subunit assembly, and centromere or microtubule binding (158, 159, 213). Dyskerin has been shown to be a component of the telomerase complex (214). In support of this role, DC fibroblasts display reduced telomerase RNA and activity with a concordant decrease in telomere lengths (215). The autosomal dominant form of DC most likely involves a defect in the hTR gene, which causes improper secondary structure formation of hTR (160, 216, 217). The observation that both forms of DC are caused by defects in telomerase strongly indicates that defective telomere maintenance is the underlying basis (158). Although the culpable gene has not yet been identified in the autosomal recessive form of DC, evidence based on the other two forms suggests that it likely is also involved in telomerase assembly or activity (158). Taken together, the existence of a disease of dysfunctional telomerase highlights the significance of telomere biology in cellular homeostasis.

4.5.4

Cancer

Cancer in its vast array of forms is predominantly a disease linked with advancing age (4). Over time, the effect of accumulation of DNA damage produces extremely complex cytogenetic profiles, with especially high frequencies of translocations (4). Continual chromosomal and genomic instability are amenable environments for increased mutagenesis in oncogenes and tumor suppressor genes, allowing tumorigenesis to proceed unhindered (17, 40). Since telomerase is elevated in the majority of cancers, including breast, colorectal, and nonsmall cell lung cancer, its activity is a good indicator of overall survival, and may be used as confirmation of malignancy (218–221). Defects in DNA damage response components, such as p53, ATM/ATR, and BRCA1/2, contribute to cell cycle checkpoint malfunction and continued proliferation in the context of unrepaired DNA (53). For example, deficiencies in HR and NHEJ are associated with an increase in chromosomal aberrations, instability, and a predisposition to malignant transformation (68, 222–224). Also, ATM- and p53mediated formation of DNA damage foci appear at shortened or dysfunctional telomeres, targeting them for DNA repair (13, 137).

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Furthermore, highly complex abnormal karyotypes tend to be associated with shorter telomeres (225). Based on this observation, it is important to address whether telomere shortening and subsequent dysfunction directly cause instability, rather than being simply the result of excessive proliferation. Evidence in support of the former role shows that in pancreatic tumor cells, telomere attrition is the earliest detectable change (226). However, this does not exclude the possibility of a high proliferative rate as the source of shortened telomeres. Furthermore, cells that escape senescence continue to shorten their telomeres resulting in massive chromosomal instability, especially the appearance of chromosomal fusions (227). Aberrant resolution of these fusions can lead to deletion of tumor suppressor genes or amplification of oncogenes, all contributing to malignancy (4). For instance, p53 null mice that lack checkpoint control and thus enter senescence have dysfunctional telomeres and are more likely to possess an unstable genome (228). Although the exact mechanisms of tumorigenesis are variable and require further clarification, dysfunctional telomeres are nevertheless key players in contributing to genomic instability (229).

4.5.5

Neurodegenerative Diseases

Although the role of telomeres in cancer and aging has been intensively studied, telomere function in postmitotic neuronal cells is less clear. Nondividing neurons only constitute 10% of brain cells, but a disruption in telomere homeostasis has significant pathologic consequences, including mental retardation, developmental delay, and neurological disorders such as Alzheimer’s disease (AD) and dementia (161, 230–233). Similar to adult somatic cells, differentiated neurons do not normally express telomerase activity. However, upon oxidative or hypoxic stress, or excess neurotransmitter receptor stimulation, telomerase is reactivated in neurons (234, 235). Specifically, lymphocytes of AD patients have telomere shortening and aberrant telomerase activity, such that hTERT may impart some protection from amyloid b-peptide accumulation and DNA damaging agents (162, 236). These observations suggest that telomerase may be a mediator of DNA repair, ensuring cell survival upon cytotoxic stress (237). Furthermore, mutations in the RNA template component hTR leads to telomere dysfunction and is associated with reduced proliferation in adult neural stem cells (238). In addition to an association with telomerase, overexpression of a dominant negative form of TRF2 results in activation of ATM and gH2AX in both mitotic astrocytes and postmitotic neurons, but only mitotic cells undergo senescence upon p53 stabilization (239). Developing neuronal cells may recruit TRF2 to sites of damage to aid in repair and to relieve any telomere position effect on telomere-proximal genes involved in neurogenesis (240). Collectively, these findings indicate a role for telomere maintenance in neuronal survival.

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Therapeutic Possibilities Telomerase and Telomere-Based Therapies

Most studies on the development of drug targeting and therapeutics for disease involving telomerase inhibition is most often with special emphasis on implications for cancer treatment. Telomerase inhibition should be effective and specific, while normal cells lacking telomerase should not be significantly affected by anti-telomerase therapy. Since telomerase expression in cancer cells can be used as a novel marker for screening, early detection, and prognosis, clinical trials are already underway to assess this potential (241, 242). Inhibition of telomerase may either be direct by targeting the essential components required for assembly or activity, or be indirect via targeting of its telomeric DNA substrate (Table 4.2). Targeting of the RNA template and nontemplate regions would restrict hTERT accessibility and prevent telomerase assembly, inducing telomere shortening and limiting proliferation (245, 253). Most strategies rely on antisense oligonucleotides, hammerhead ribozymes, or small molecule inhibitors to modulate telomerase expression or activity (254). Modifications in the sugar phosphosdiester backbone of short DNA/RNA antisense molecules aimed at hTR allow these agents to have high penetration, binding affinity and specificity, and resistance to nuclease degradation (254). Such molecules include peptide nucleic acids (PNA); DNA oligomers with phosphorothiate (PS-DNA) and phosphoramidite (PN-DNA) linkages; RNA oligomers with methyl-substituted (2-OMe RNA) and methoxyethyl-substituted (2-MOE RNA) ribose rings; and variations thereof (253, 255–258). For instance, GRN163L is a lipidated PN-DNA oligonucleotide that is complementary to hTR (243, 244). The potency of this inhibitor has been demonstrated in rodent xenograph models of lung cancer, hepatoma, and glioblastoma, in which administration of GRN163L inhibited tumor progression and shortened telomeres (243, 259, 260). The efficacy of GRN163L in preclinical studies has prompted phase I/II clinical trials for chronic lymphocytic leukemia (4).

Table 4.2 Telomere and telomerase targeting compounds Target Compound Direct hTR GRN163L hTERT AZT TDG-TP BIBR1532 Indirect G-quadruplex 9-anilino proflavine triazine fluoroquinophenoxazine telomestatin pentacyclic acridines

Reference (243, 244) (245) (246) (247) (248) (249) (250) (251) (252)

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Although hTR-directed ribozymes have been shown to reduce telomerase activity, the effect is more pronounced when targeted against hTERT. Nucleoside analogs inhibiting reverse transcriptase activity, such as 30 -azido-30 -deoxythymidine (AZT) and 6-thio-20 -deoxyguanosine 50 -triphosphate (TDG-TP), significantly reduce telomerase activity but only transiently affect telomere length (245, 246). On the other hand, 2-((E)-3-naphtalen-2-yl-but-2-enolyamino)-benzoic acid (BIBR1532) leads to telomere shortening and senescence in cancer cells (247). A main consideration of telomerase-directed therapy is the persistence of telomerase in stem cell-like normal cells. Inhibition of telomerase in these cells may elicit adverse physiological effects upon long-term treatment. Despite these drawbacks, there is great potential in their utilization and further investigation is warranted. Indirect targeting of telomerase via disruption of telomere structure takes advantage of the presence of repetitive sequences and secondary structure found in the T-loop (261). Disruption in telomere maintenance can also be achieved by targeting tankyrase. As a positive regulator of telomere length, tankyrase catalyzes the inhibition of TRF1 binding to or the dissociation of TRF1 from telomeres, thus disassembling the T-loop and allowing an opportunity for elongation (15, 261). In addition to T-loop disruption, the formation of G-quadruplexes sequesters telomeric DNA ends into intramolecular structures of G-tetrads and shields it from telomerase activity (262). The presence of G-quadruplex interacting compounds, such as 9-anilino proflavine, triazine, fluoroquinophenoxazine, telomestatin, and pentacyclic acridines, interferes with telomere structure to effectively inhibit telomerase access, causing telomere shortening and senescence (248–252). Regardless of the mechanism of action of these agents or specific targets, they all negatively affect telomere integrity. It is important to note that telomerase inhibition is not immediately effective since significant telomere shortening only occurs after many cell divisions. In the meantime, the cancer could considerably heighten in clinical complications. Also, telomerase-positive cells, such as germ and other proliferative cells, may be adversely affected by telomerase inhibition (263). However, these cells naturally have longer telomeres and are often in a quiescent state, so the effect will not be immediate as renewal of stem cells occurs transiently (242). Another consideration is the activation of alternative methods of telomere elongation, such as the ALT pathway. Thus, telomerase- and telomere-based therapies will likely be most useful when used as adjuvant therapy in conjunction with traditional cancer treatments, such as resection, radiation, and chemotherapy (242).

4.6.2

Targeting DNA Damage Response and Repair

Inhibition of DNA damage proteins or repair mechanisms has been widely studied in the context of anticancer applications. Most radio- and chemotherapeutic compounds can be directed to produce a broad range of DNA damage, and therefore address the involvement of multiple pathways in repair of lesions. As a method of

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Table 4.3 DNA damage and repair-based compounds Target Compound Reference DNA-PKcs NU7026 (264) Nu7441 (265) IC87102 (266) IC87361 (267) Wortmannin (268) OK-1035 (269) LY294002 (270) PARP NU1025 (271) ATM KU-0055933 (272)

sensitization to existing therapies, DNA repair proteins can be inhibited by small molecules targeting various key players to elicit a more effective cytotoxic response (Table 4.3). Cells deficient in DNA-PKcs show hypersensitivity to ionizing radiation due to defective DNA repair (273). Since inhibition of DNA-PKcs is hypothesized to induce the same response, NU7026 has recently been developed as a DNA-PKcs inhibitor, selectively acting on DNA-PKcs-containing cells to potentiate cytotoxicity (264, 274). Additional DNA-PKcs inhibitors, such as Nu7441 (265), IC87102 (266), IC87361 (267), wortmannin (268), OK-1035 (269), and LY294002 (270), are all potent radiosensitizers. Furthermore, PARP inhibitors like NU1025 and ATM kinase inhibitors like KU-0055933, which blocks phosphorylation of ATM targets, are also associated with increased cytotoxicity and chemo- and radiosensitization (271–273). Finally, the direct involvement of WRN and BLM helicases in HRmediated repair and their interaction with the MRN complex provides a strong incentive for developing targeted therapies against these proteins (275). Components of other repair pathways also exist as targets for drug inhibition and may be implicated in cancer therapy. An exhaustive compilation of all these targets is beyond the scope of this chapter, but suffice is to say that the number of potential targets continues to grow. The fact that many of these proteins are localized or associated at the telomere suggests that their inhibition may be an indirect method of telomere disruption.

4.7

Concluding Remarks

The field of telomere biology has seen significant advances in understanding telomere function and interaction on a global scale. Gone is the simplistic view of telomeres as simply noncoding disposable DNA. It is now apparent that telomeres are highly regulated and complex structures that play important roles in assuring genomic fidelity. Although the roles of all the players in this process are not fully understood, current data nevertheless provide a multifaceted representation of

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normal telomere behavior. It is especially apparent that telomere maintenance and the DNA damage response are interdependent, and one cannot fully explain either process without mentioning the other with respect to telomeres. As the knowledge base increases with each new study, the potential for development of treatments for diseases including, but not limited to, cancer will simultaneously grow. Although much progress has been made toward elucidating telomere dysfunction and the DNA damage response, many areas still need to be explored. The future will surely see exciting advances in understanding telomere biology and its potential application into clinical settings.

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Chapter 5

Alternative Lengthening of Telomeres in Human Cells Hilda A. Pickett and Roger R. Reddel

Abstract Telomere renewal is a prerequisite for cellular immortalisation. Some cells maintain their telomeres by a telomerase-independent alternative lengthening of telomeres (ALT) mechanism. Characteristic features of most ALT-positive human cells include highly heterogeneous telomere lengths, PML nuclear bodies containing telomeric DNA and telomere-binding proteins, a high frequency of telomeric exchange events, and the presence of extrachromosomal telomeric DNA circles. Numerous proteins involved in DNA recombination, repair and replication also associate with APBs, and proteins involved in homologous recombination are necessary for ALT. These and other data indicate that the mechanism of telomere lengthening in ALT-positive cells may involve recombinationmediated replication of telomeric sequences. Keywords: Telomere, Telomere maintenance mechanism, Alternative lengthening of telomeres, Homologous recombination, ALT-associated PML bodies.

5.1

Introduction

Telomeric DNA is cumulatively lost from the ends of linear chromosomes with each round of cell division due to the end replication problem, ultimately resulting in a limitation of cell proliferative capacity (1). In order to circumvent this limitation, immortal cells with unlimited proliferative potential (including immortal cell lines and tumour cells) require an active telomere maintenance mechanism (TMM). The majority of immortal cells, as well as germline and stem cells, activate the ribonucleoprotein holoenzyme complex telomerase, which comprises the RNA subunit hTR, the reverse transcriptase hTERT and dyskerin (2). Approximately 10 –15% of human tumours utilise a telomerase-independent TMM known as alternative lengthening of telomeres (ALT) (3–5). R.R. Reddel(*) Children’s Medical Research Institute, 214 Hawkesbury Road, Westmead, NSW 2145, and University of Sydney, NSW 2006, Australia, e-mail: [email protected]

K. Hiyama (ed.), Telomeres and Telomerase in Cancer. DOI: 10.1007/978-1-60327-879-9_5, # Humana Press, a part of Springer Science + Business Media, LLC 2009 127

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The existence of ALT in human cells was deduced from the observation that telomerase-negative cell lines were able to maintain the length of their telomeres for many hundreds of population doublings (3, 5, 6). ALT can occur in many types of cancers, but is particularly prevalent in tumours of mesenchymal origin, such as sarcomas and astrocytomas (7). Further molecular characteristics of the ALT mechanism have since been identified, consistent with a homologous recombination (HR)-dependent repair mechanism. Nevertheless, many of the molecular details of ALT remain elusive. This chapter will describe the characteristics of ALT cells, and what is currently known about the mechanisms responsible for telomerase-independent telomere length maintenance.

5.2

Telomere Length Phenotype

Telomere lengths in the human germline are normally maintained by telomerase at around 15 kb (8, 9). The telomeres of cultured normal human somatic cells shorten at a rate of 40–200 bp per cell division (1). Cells eventually reach a state known as cellular senescence, during which the cells remain metabolically active but cease to divide, and at which point terminal restriction fragment (TRF) lengths are typically between 5 and 8 kb in length (10–12). Telomere shortening and the onset of cellular senescence is thought to be a major barrier to tumorigenesis. Cells which bypass cellular senescence and are ultimately immortal have an active TMM, be it telomerase or ALT. Most of the immortal cell populations utilising an ALT TMM that have been characterised to date show a heterogeneous telomere length phenotype by TRF length analysis (Fig. 5.1), with telomere lengths distributed from very short (50 kb) (3, 4, 13–15), in contrast to the much more homogeneous telomere lengths of 100 mM) of these compounds to induce a senescence-like phenotype, despite telomere shortening (25, 36, 46, 50, 56). However, little data have been recently reported on the use of nucleoside analogs as telomerase inhibitors, as these original findings on the nonspecific inhibition of other polymerases and general cytotoxicity proved troublesome for cancer therapeutic development (Table 10.2).

10.3.1.2

Nonnucleoside Agents

A variety of nonnucleoside agents have also been shown to have antitelomerase activity, result in decreased proliferation, and induce a senescence-like phenotype

hTERT vaccine

hTERT epitopes on cancer cells; cancer remission/ prevention of relapse

Breast (NCT00573495)

Prostate, AML (NCT00510133)

Yes

Yes

NSCLC: combination with paclitaxel and carboplatin (NCT00510445)

Yes

Good tolerability, favorable pharmacokinetics, no measurable change in telomerase activity or telomere length due to short treatment times, beneficial outcome on tumor growth Ongoing; well-tolerated, no dose-limiting toxicities or adverse side effects Early stages of recruitment; determine the safety and maximum tolerated dose when given in combination with a standard paclitaxel/ carboplatin regimen Ongoing; in the tumor microenvironment, infiltration of T cells and widespread tumor necrosis, longer median overall survival Ongoing; telomerasespecific immune responses, no significant toxicity, clearance of circulating cancer cells, prolonged PSA doubling times

Chronic lymphocytic leukemia (NCT00124189); multiple myeloma

Solid tumors (NCT00310895)

Current status/results

Cancer type

Yes

Table 10.2 Status of telomerase inhibitors in the clinic Agent Target/expected Therapies in trial? outcome Telomerase template hTR/hTERC Yes antagonist, oligonucleotides competitive (GRN163L) enzyme inhibitor, progressive telomere shortening

Johannes Vieweg, Duke University Medical Center (http://www. mc.duke.edu/) and Geron Corporation (http://www.geron. com/)

Robert Vonderheide, University of Penn Medical (http://www. med.upenn.edu/)

Geron Corporation (http://www.geron. com/) Geron Corporation (http://www.geron. com/)

Developer, investigators Geron Corporation (http://www.geron. com/)

230 B.-S. Herbert, E.M. Goldblatt

Telomeres/telomerase accessibility to telomeres; cell death due to telomere uncapping, dysfunction hTERT inhibition; Reverse transcriptase inhibitor, used in HIV infection treatment hTERT inhibition No

Not specifically for targeting telomerase

No

hTERT/hTERC Inhibit hTERT No (trials exist for inhibitors expression or HSP90 assembly, inhibitors) (siRNA, synthesize MT-hTer, mutant telomeres hammerhead ribozymes, HSP90 inhibitors) With clinicaltrials.gov ID, not including gene therapy trials

Nonnucleoside compounds (BIBR1532)

Nucleoside analog (AZT)

G-Quadruplex stabilizers (Telomestatin, BRACO19, RHPS4)

Yes

Multiple

Multiple

Multiple

Pancreatic (NCT00425360), lung (NCT00509457), melanoma (NCT00021164) Multiple

Preclinical testing

Preclinical testing

None

Ongoing; no adverse side effects, no autoimmune response, no effect on bone marrow stem cells, immune responserelated benefit Preclinical testing

Gustav Gaudernack, Norwegian Radium Hospital (http://www. radium.no/), Cell Genesis (http://www. cellgenesys.com/)

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after long-term treatment [Table 10.1; (25, 50, 55)]. Green tea epicatechin derivatives (EGCG) and synthetic compounds with EGCG-related moieties (MST-312, MST-295, and MST-199) have been demonstrated as effective hTERT inhibitors via biochemical or genetic/epigenetic mechanisms not completely understood (57– 59). The most widely studied compound of this class targeting hTERT is BIBR1532 (2-[(E)-3-naphthalen-2-yl-but-2-enoylamino]-benzoic acid), a highly specific, noncompetitive catalytic inhibitor that is mechanistically similar to inhibitors of HIV reverse transcriptase (46, 60). This compound has a high specificity for telomerase and has been shown to inhibit telomerase in a multiple number of tumor cell types with effective concentrations to inhibit 50% of activity (EC50) in the nanomolar (nM) range (61). In vivo studies have shown that long-term treatment with this compound can significantly decrease tumor growth. The long lag phase associated with this type of inhibition, taking up to 100 days of treatment prior to seeing the beneficial effects of telomerase inhibition, is expected for a classical telomerase inhibitor but may not be clinically relevant for effective cancer treatment (25, 46, 50, 60, 61). However, telomerase inhibition by BIBR1532 can sensitize drugresistant cancer cells, as well as drug-sensitive cancer cells, to other chemotherapeutics (62). Currently, none of these hTERT inhibitors are under clinical investigation, although preclinical testing is ongoing (Table 10.2).

10.3.1.3

Other Direct and Indirect Methods

As the regulation of telomerase is multifaceted and complex, so are the opportunities to target telomerase. Other direct and indirect methods for inhibiting hTERT have been reported, and the reader is directed to recent reviews on other potential telomerase inhibitors not described in this chapter (46, 55). Agents such as ceramides, irradiation, chemotherapy, Gleevec, tamoxifen, retinoids, epicatechins (e.g., epigallocatechin gallate or EGCG), and protein kinase C (PKC) inhibitors have been shown to affect hTERT expression, phosphorylation status, or cellular localization (18, 46, 55). hTERT can be also prone to degradation by ubiquitin ligases. For example, overexpression of the recently reported ubiquitin ligase MKRN1 in H1299 lung carcinoma cells promotes the degradation of hTERT and decreases telomerase activity (63). Many of the mechanisms for these agents on telomerase inhibition remain unclear, may be nonspecific, or be due to cytoxocity, which itself reduces telomerase activity. Furthermore, understanding the mechanisms of hTERT inhibitors can be complicated by the recent suggestion that hTERT may exhibit other activities beside the maintenance of telomeres (51, 64). For instance, Masutomi et al. suggested that the stable expression of retroviral vectors encoding hTERT-specific short hairpin RNA (shRNA) renders normal human BJ foreskin fibroblasts cells sensitive to DNA damage (52). The state of various histones in cell extracts was measured biochemically, and the loss of hTERT was reported to alter the overall chromatin state, but not the short-term telomere integrity (52). However, there is no clear evidence of whether any of the small molecule hTERT inhibitors also affect normal cellular function and hence extracurricular activities of telomerase.

10 Therapeutic Targets and Drugs I: Telomerase and Telomerase Inhibitors

10.3.2

Targeting hTERC (hTR or hTER)

10.3.2.1

Antisense Oligonucleotides

233

Telomerase requires its RNA component hTERC for its reverse transcriptase function. In 1995, using an antisense RNA to hTERC, Feng et al. provided one of the first reports demonstrating that the RNA component of telomerase could be effectively targeted in human cancer cells, resulting in inhibition of telomerase activity, progressive telomere shortening, and subsequent cell death (6). Standard antisense oligonucleotides (ODNs) contain DNA bases that form DNA–RNA hybrids with their target mRNA. RNase H then recognizes these hybrids and cleaves them, resulting in reduced protein expression. However, telomerase is not a typical antisense target because RNase H cleavage of hTERC is not necessary. In addition, hTERC is a good target for oligonucleotide-based mechanisms for several other reasons (65). First, for telomerase to function, the template region of hTERC must be able to pair with the telomere by Watson–Crick base pairing and thus can be exposed to oligonucleotides. Second, the known sequence of the 11-nucleotidelong hTERC template region (50 -CUAACCCUAAC-30 ) simplifies the design of oligonucleotide-based inhibitors. Many of the oligonucleotides analyzed contain sequences that are complementary to a 13-nucleotide-long region, which partially overlaps and extends by four nucleotides beyond the 50 -boundary of the template region of hTERC. Third, oligonucleotides that contain mismatch or scrambled bases allow for intrinsic experimental controls. Finally, oligonucleotides already have supportive evidence for being applicable within the clinic, and certain ODNs can be spontaneously taken up by some animal and human tissues (65). Unlike traditional antisense ODNs that inhibit translation by binding to mRNA, ODNs complementary to the hTERC template region actually act as competitive enzyme inhibitors (or template antagonists) because these agents block the active site of hTERT reverse transcriptase (i.e., hTERC is not translated into peptides). The early investigations into targeting the hTERC component using traditional antisense ODNs validated it as a good target; however, unmodified ODNs are readily degraded by nucleases and are unstable. Therefore, investigators have focused on the development of modified ODNs to improve the use of oligonucleotide-based targeting of hTERC (Table 10.1). This includes ODNs composed of chemically modified bases, as opposed to DNA bases, since RNase H cleavage is not necessary for telomerase inhibition.

10.3.2.2

Chemically Modified Oligonucleotides

Peptide nucleic acids (PNAs) were among the first chemically modified oligonucleotides targeting hTERC to be reported as effective in inhibiting telomerase activity with EC50s in the pico- to nanomolar range (66). PNAs are DNA mimics with a neutral amide backbone, but are not well distributed in vivo (65, 67). Therefore, subsequent investigations have utilized well-characterized chemistries

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that could be optimal for translational research in vivo (i.e., in animals and humans). This led to research in the development of 20 -O-alkyl RNAs, such as 20 -O-methyl RNA (20 -O-MeRNA) and 20 -O-methoxyethyl RNA (MOE RNA), as effective telomerase inhibitors with high affinity for hTERC binding (41, 68–70). Indeed, the study by Herbert et al. using a 13-mer 20 -O-MeRNA targeting the hTERC template region was one of the first reports to support the rationale for targeting telomerase in cancer by demonstrating telomere shortening and cell growth inhibition after telomerase inhibition in human cancer cell lines (39–41). ODNs tagged with 20 ,50 -oligoadenylate (2-5A) have also been shown to inhibit telomerase activity and induce apoptosis in many different types of cancer cells in vitro and in vivo (71–77). However, these effects were seen within 3–6 days, without telomere shortening, and may be due to a mechanism of action of degrading hTERC by RNase L (78) or one that is not specific to telomerase inhibition and progressive telomere erosion (42). Recent studies using 2-5A ODNs have suggested that 2-5A can act synergistically with conventional radiation- and chemotherapies in inhibiting glioma cancer cell growth in vitro and in vivo (79).

10.3.2.3

PS-, NP-, and NPS-ODNs

Additional development of oligonucleotides as telomerase template antagonists has involved phosphorothioate (PS), N30 -P50 phosphoramidate, and N30 -P50 thiophosphoramidate DNA (46, 65, 80–83). DNA oligonucleotides containing PS linkages between the bases have enhanced stability against nuclease digestion as well as enhanced binding to proteins (65). This latter property may result in cellular phenotypes unrelated to the effect on the target RNAs when these PS-ODNs are introduced into cells. N30 -P50 phosphoramidates (NP) designed to be complementary to the template region of hTERC provided further optimization of telomerase template oligonucleotide antagonists compared with a mismatch control (80, 81). Finally, N30 -P50 thio-phosphoramidates (NPS) take advantage of the PS and NP oligonucleotide properties (65, 82, 83). NPS ODNs targeting the telomerase template region (e.g., GRN163) have been shown to be effective telomerase inhibitors and anticancer agents through the shortening of telomeres to a critical length for cancer cell survival (82, 84–87). The 13-mer, lipid-conjugated NPS telomerase template antagonist GRN163L represents one of the latest generation of modified oligonucleotides targeting hTERC (Table 10.1). GRN163L contains a lipophilic, palmitoyl tail at the 50 terminal end of its sequence (50 -Palm-TAGGGTTAGACAA-30 ), which allows for efficient cellular uptake in vitro and in vivo without the use of transfection reagents (e.g., lipid carriers) or electroporation (88). The lipid modification of GRN163L enhances the stability and potency of telomerase inhibition (EC50 in subnanomolar concentrations) and inhibition of cell growth compared with that of GRN163, a nonconjugated NP with the same sequence, without the use of transfection reagents (88). This potency was not due to the palmitoyl moiety alone, as a lipid-modified mismatch NPS had no effect on telomerase inhibition and cell growth at the same

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concentration. GRN163L has been extensively studied in many cancer cell types in vitro and in preclinical in vivo xenograft models with encouraging results, all of which have provided support for the use of these antagonists as targeted cancer therapeutics (89–94). Interestingly, recent reports suggest dramatic effects of GRN163L in cancer cell growth inhibition well before the bulk of the telomeres within a population of cells have reached critical shortening (90–92, 94). Jackson et al. reported that when cancer cells were given GRN163L before they were allowed to attach to culture dishes, GRN163L prevented the cells from adhering to the dishes (94). These effects were shown to be correlated to the sequence (i.e., mismatch controls did not have an antiadhesive effect), length, backbone chemistry, and lipid modification of the oligonucleotide; however, GRN163L also had the same effect on immortalized cells that use the ALT pathway and are telomerase deficient (94). These findings may help explain the in vivo observations where GRN163L inhibited tumor growth and metastasis of human cancer cells within a month of treatment (one to three times per week at pharmacological doses). Research into the mechanisms of action for GRN163L is ongoing and should be useful for further preclinical and clinical testing. As described in more detail later in this chapter, these telomerase template antagonists are the first telomerase inhibitors to be translated to the clinic and are currently undergoing testing in Phase I/II clinical trials in different cancer types.

10.3.2.4

RNAi, Ribozymes, and Mutant hTERCs

It has also been shown that depleting the endogenous wild-type hTERC in cells through the use of RNA interference, ribozymes, or mutant hTERCs can lead to reduced telomerase activity (Table 10.1). For instance, hTERC-targeting siRNAs have been shown to decrease telomerase activity and can inhibit xenograft tumor formation without the shortening of the overall telomere population (46, 95–97). The second technique for depleting wild-type hTERC uses a mutated template hTERC (mutant template hTer or MT-hTer; Table 10.1) to allow for synthesis and incorporation of ‘‘mutant’’ telomeric DNA, which can subsequently affect the binding of protective telomere-associated proteins and telomere structure (6, 98– 102). Goldkorn and Blackburn showed that the effects of MT-hTer/hTERC siRNA required a functional and catalytically active telomerase (102). The MT-hTer has also been shown to decrease cell viability and increase apoptosis independently of initial telomere length. In addition, MT-hTer inhibited tumorigenesis in mice (50, 101, 102). The use of lentiviral vectors for the delivery of MT-hTer and hTERC siRNA has been shown to rapidly inhibit cancer cell growth within days, well before the predicted lag phase seen with other classical hTERC telomerase inhibitors (101). More recently, MT-hTer was reported to increase the sensitivity of cancer cells to chemotherapeutic agents without requiring global telomere shortening (103).

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Summary

Taken together, targeting the hTERC component of telomerase has received a great amount of attention in terms of cancer therapeutic potential. Approaches targeting hTERC can provide an advantage over targeting hTERT, particularly if more evidence comes forward regarding other roles for hTERT in normal cell survival, since the expression levels of the proteins/RNA within the telomerase complex are not specifically altered (except when using 2-5A oligonucleotides or siRNA). Using these applications for targeting telomerase may provide not only potential clinical benefit, but also scientific insight into the role of telomerase in telomere and cancer biology.

10.3.3

Targeting Telomerase-Associated Proteins

In vitro, hTERC and hTERT are enough to reconstitute telomerase activity (1, 6, 8–10, 104, 105). However, a multitude of other proteins have been implicated to be associated with telomerase in vivo (1). Of this long list of proteins, dyskerin, hTEP1, and the chaperones HSP90/p23 have been shown to play a major role in telomerase assembly, stability, and catalytic activity since defects in these proteins can have negative effects on these properties. As described earlier, mutations or defects in dyskerin have been shown to cause bone marrow diseases such as anemia and dyskeratosis congenita. The role of hTEP1 in telomerase function remains unclear, but poly(ADP-ribose) polymerase (PARP) inhibitors have been shown to downregulate hTEP1 expression and telomerase activity in leukemia cells (46, 106, 107). Chaperone inhibitors, particularly for HSP90, have been the most extensively studied within the class of telomerase-associated protein inhibitors, which is the focus of discussion for the rest of this section [Table 10.1; (108) for review]. In general, the heat shock protein (HSP) HSP90 is a chaperone protein that plays a central role in the stability and proper folding of a select number of other proteins such as HER2, androgen and estrogen steroid receptors, AKT/PKB, C-RAF, CDK4, survivin, HIF-1a, mutant TP53, and BCR/ABL [see (109) for review]. The critical role of HSP90 in the stability of these proteins, which are typically active in cancer, has led to the rationale for the development of HSP90 inhibitors as anticancer agents. Furthermore, cancer cells may have a high-affinity, activated form HSP90 compared with an inactive form in normal cells (108, 110). Blocking HSP90 activity has been achieved through the use of pharmacological inhibitors such as geldanamycin (GA) and the geldanamycin analog 17-allylamino-17demethoxy-geldanamycin (17-AAG). Further studies of 17-AAG analogs, such as 17-DMAG [see (109) for a more complete review], have shown continued improvement in the metabolism and antiproliferative capability of these HSP90 inhibitors.

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HSP90 has also been shown to associate with functional human telomerase (111). In addition, the cochaperone p23 has been shown to associate with active telomerase, while another chaperone, HSP70, associates with hTERT in a transient fashion (111, 112). The role of chaperones in telomerase assembly and function is not completely understood and may be more complex than previously expected (113); however, it has been suggested that chaperones may aid in the reverse transcriptase process of telomerase (112). As hTERT generates de novo hexameric telomeric repeats, it must translocate to the next available position for further processing, changing its conformation during telomere processing. Although the assembly of telomerase occurs in the cytoplasm with the aid of most of the available chaperones, the few molecules of available nuclear HSP90/p23 may aid in adjusting telomerase conformation while functioning on the telomere (112). Studies on the ability of HSP90 inhibitors to affect telomerase activity have resulted in novel approaches toward targeting telomerase (Table 10.1). Downregulation of HSP90 in various cancer cells by ODNs resulted in inhibition of telomerase activity (114). Pharmacological HSP90 agents (such as GA, 17-AAG, and novobiocin) have been shown to block the assembly of active telomerase as well as the association of p23 to the telomerase complex in vitro as well as in vivo (111, 113, 115). Furthermore, Harvey et al. reported that chronic treatment of prostate cancer cells with subtoxic concentrations of another HSP90 pharmacological inhibitor, radiciol, resulted in blocking telomerase assembly and telomerase activity (116). The concentrations used in all of these studies (100 ng/ml or 0.3 mM) were not cytotoxic. Villa et al. also reported that the basal level of telomerase in cancer cells may influence the sensitivity to HSP90 inhibitors (115). Compton et al. demonstrated that long-term treatment (60 days) with HSP90 inhibitors, by either pharmacological (e.g., radiciol) or genetic (siRNA knockdown) approaches resulted in dramatic telomere shortening and cell death via nitric oxide synthase (NOS)-induced free radical production (117). However, the low concentration of radiciol used in this study (0.3 mM) only transiently reduced telomerase activity (50 kB, in contrast to the much more homogeneous telomere lengths in normal and telomerase-positive cells (Fig. 16.1a).

16.2.1

Buffers and Solutions

1. CHAPS (3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulphonate) lysis buffer: 10 mM Tris-HCl pH 7.5, 1 mM MgCl2, 1 mM EGTA, 0.5% (v/v) CHAPS, 10% (v/v) glycerol, 5 mM beta-mercaptoethanol, 0.1 mM 4-(2-aminoethyl)-benzenesulphonyl fluoride hydrochloride (AEBSF). 2. 50
Denhardt’s solution: 1% (w/v) Ficoll-400, 1% (w/v) BSA (fraction V), 1% (w/v) polyvinylpyrrolidone 40 (PVP-40). 3. DNA lysis solution: 50 mM Tris-HCl pH 8.0, 20 mM EDTA, 2% sodium dodecylsulphate (SDS). 4. 20
SSC buffer: 3 M NaCl, 0.3 M tri-sodium citrate. 5. 10
TBE buffer: 0.9 M Tris-HCl pH 8.0, 0.9 M borate acid, 20 mM EDTA. 6. TE buffer: 10 mM Tris-HCl pH 8.0, 1 mM EDTA.

16.2.2

DNA Isolation

For tumour samples, approximately 100 mg of frozen tissue is homogenised at 4 C in 200 mL of CHAPS lysis buffer, incubated on ice for 20 min and centrifuged at 18,000
g at 4 C for 20 min (with CHAPS lysis buffer, both DNA and protein can be collected from the same sample). Genomic DNA is extracted from the pellet by homogenising lightly in 5.5 mL of DNA lysis solution containing 100 mg/mL Pronase protease (Sigma) and incubating for 16 h at 37 C with occasional gentle inversion. For cultured cells, DNA are isolated from cell pellets, as described for tumour samples, except that the homogenisation steps are replaced by gentle mixing by pipetting. Lysates for genomic DNA extraction from tumour and cell culture samples are cooled on ice for 5 min, 2 mL of saturated NaCl is added and the mixture is incubated at 4 C for 8–16 h. The precipitate is removed by repeated centrifugation (2,000
g at 4 C for 15 min)

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and the supernatant is transferred to a clean 15 mL centrifuge tube. The DNA is precipitated by adding 2.5 volumes of 100% ethanol and storing at 20 C for at least 12 h. When the genomic DNA is required for further analyses, the sample is centrifuged at 2,000
g, 4 C for 15 min, washed with 70% ethanol, resuspended in 50 mL of TE buffer and stored at 4 C.

16.2.3 Southern Analysis of TRF Lengths TRFs are generated from genomic DNA by digesting for 12–16 h at 37 C with 4 U/mg each of Hinf I and RsaI restriction enzymes (Roche) and 25 ng/mg RNAse (DNAse free; Roche), then heat inactivating at 80 C for 20 min and storing at 4 C. The digested DNA is quantitated in a fluorescence spectrophotometer. The digested genomic DNA samples (1.5 mg/well) are loaded onto a 1% agarose gel in 0.5
TBE buffer and TRFs separated by pulsed-field gel electrophoresis using a CHEF-DR II apparatus (BioRad), in recirculating 0.5
TBE buffer at 14 C and with a ramped pulse speed of 1–6 s at 200 V for 14 h. An appropriate DNA molecular weight marker such as Low Range PFG Marker (New England BioLabs) is included in each gel. The gel is then stained in 0.5 mg/mL ethidium bromide for 30 min and photographed on a UV transilluminator to confirm equal loading and record marker positions. The gel is dried under a vacuum at 65 C until it is approximately 0.5 mm thick and has just turned translucent, washed in denaturing solution (0.5 M NaOH, 1.5 M NaCl) for 45 min, then washed in neutralising solution (1 M Tris-Cl pH 8.0, 1.5 M NaCl) for 45 min. The gel is prehybridised in 30 mL containing 5
SSC, 5
Denhardt’s solution, 0.5 mM tetrasodium pyrophosphate, and 10 mM disodium hydrogen orthophosphate at 37 C for 2–6 h. An oligonucleotide (TTAGGG)3 probe (150 ng; Sigma) is 50 end-labelled with 50 mCi of [g32P]-dATP (3,000 Ci/mmol; New England Nuclear, Dupont) and 10U T4 kinase (Promega) in a volume of 10 mL at 37 C for 30 min. The probe is purified by ethanol precipitation and resuspended in 30 mL of TE. 3
106 counts per minute (cpm) of probe is added directly to the prehybridising gel, and incubated at 37 C overnight in a rotating oven (Hybaid). The gel is then washed three times in 0.1
SSC for 7 min at 37 C, exposed to a phosphor screen and scanned with a STORM 860 optical scanner with ImageQuant software (Molecular Dynamics). Molecular weights of telomeric bands are determined by constructing a standard curve from the DNA markers run on the same gel.

16.3

Detection of APBs by Immunostaining and Telomere Fluorescence in Situ Hybridisation (FISH)

APB-staining is a well-established assay specific for differentiating ALT cancer cells from non-ALT cells by the presence of APBs, a unique hallmark of ALT. This assay is based on immunofluorescence/telomere FISH to detect APBs–colocalisation

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of PML protein with one of the telomere-binding proteins such as TRF2 or telomeric DNA in a bright nuclear focus of characteristic morphology. This is a relatively rapid and easy assay when compared with the TRF analysis, and thus quite suitable for clinical diagnostic application.

16.3.1 Buffers and Solutions 1. Antifade mounting medium: 90% glycerol buffered with 20 mM Tris-HCl pH 8.0, 2.33% (w/v) 1,4-diazabicyclo[2.2.2]octane (DABCO). 2. Blocking solution: 2% (w/v) BSA (fraction V), 0.2% (v/v) Tween 20, 5% (v/v) glycerol in PBS. 3. Hybridisation buffer: 70% formamide (deionised), 10 mM Tris-HCl pH 7.5, 1% Blocking Reagent (Roche), 5% MgCl2 buffer (82 mM Na2HPO4, 9 mM citric acid, 25 mM MgCl2). 4. Washing solution A: 70% formamide, 10 mM Tris-HCl pH 7.2, 0.1% BSA. 5. Washing solution B: 0.05 M Tris-HCl pH 7.5, 0.15 M NaCl, 0.05% Tween 20.

16.3.2 APBs in Cell Culture Monolayers Cells grown in four-well or two-well chamber slides (Nunc) are washed twice with PBS and fixed for 15 min in 2% paraformaldehyde at room temperature (RT), followed by two washes with PBS, and then permeated with methanol/acetone (1:1) at 20 C for 15 min. The fixed cells are washed and rehydrated in PBS for >30 min, and then incubated with primary antibodies either for 1 h at RT or overnight at 4 C, followed by the incubation of fluorescently conjugated secondary antibodies at RT for 30–40 min. Both primary and secondary antibodies are diluted in the blocking solution. To visualise DNA, slides are incubated for 3 min in PBS with 20 mg/mL of 4,6 diamidino-2-phenylindole (DAPI). Three washes after each staining step are carried out by agitating in PBS. Finally, the preparations are mounted in the antifade mounting medium. Primary antibodies used preferentially for detection of APBs in monolayers are anti-TRF2 mouse antibody (1:200 dilution; Upstate Biotechnology) and anti-PML rabbit antibody (1:500 dilution; Chemicon). Other primary antibodies used for detecting APBs are anti-TRF1 rabbit antibody (1:400 dilution; (30)) and antiPML mouse antibody (1:200 dilution; Santa Cruz). The secondary antibodies used are as follows: FITC- or Texas Red-conjugated goat anti-mouse and goat anti-rabbit (Jackson ImmunoResearch).

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16.3.3

APBs in Tumour Specimens

16.3.3.1

Preparation of Tumour Sections

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Frozen sections are cut 5–7 mm thick and fixed in 1:1 methanol/acetone at 20 C for 12 min. After air drying, slides are rehydrated in PBS and stained immediately. For paraffin-embedded specimens, sections are cut 8-mm thick, and are then baked on Superfrost1 Plus microscope slides (Menzel-Glaser) at 65 C for 20 min and dewaxed in xylene for 5 min. Surface decalcification is needed for some osteosarcoma paraffin sections, in which case the paraffin-embedded specimen is pretreated with RDO Rapid Decalcifier, according to the manufacturer’s instructions (Apex Engineering Products Corporation). Slides are rehydrated and prepared for immunofluorescence and FISH by microwave heating to 120 C in 90% glycerol (1 mM EDTA) buffered with 10 mM Tris at pH 10.5, and maintained at 110–120 C for 15 min. The slides are cooled and rinsed in PBS before immunostaining/FISH. The staining procedures used for frozen and paraffin sections are identical. 16.3.3.2

Staining Procedures for Tumour Sections

For detection of APBs with double immunofluorescence, the staining procedure is essentially the same as that for the monolayer cultured cells (described in Sect. 16.3.2). Briefly, mouse anti-TRF2 and rabbit anti-PML antibodies are incubated for 1 h at RT or overnight at 4 C, followed by the incubation of Texas Red-conjugated goat anti-mouse and FITC-conjugated goat anti-rabbit FITC for 30–40 min at RT. The preparations are mounted in the antifade mounting medium. Detection of APBs in tissue samples, especially in paraffin-embedded specimens, is usually achieved by a combination of telomere FISH and immunostaining for PML protein (Fig. 16.2). Immunofluorescence is performed with anti-PML rabbit antibody (Chemicon) and anti-rabbit FITC goat antibody (Sigma). Sections are then cross-linked with 4% formaldehyde for 10 min and dehydrated with increasing concentrations of ethanol for telomere FISH with a 50 -labelled Cy3(50 -CCCTAA-30 )3 peptide nucleic acid (PNA) probe (Panagene). 0.5 mg/mL PNA probe in hybridisation buffer is applied to the slides, which are then denatured at 80 C for 3 min. After a 3 h hybridisation in the dark at RT, slides are washed in washing solution A for 10 min, counterstained with DAPI in washing solution B for 5–10 min and mounted in the antifade mounting medium. The slides are examined on a Leica DMLB epifluorescence microscope with a cooled charge-coupled device camera (SPOT2; Diagnostic Instruments). 16.3.3.3

APB-Counting for Tumour Specimens

A set of criteria is used to determine the APB status of tumour sections (9). Briefly, an APB is considered to be present if a focus of telomeric DNA is localised within (not adjacent or overlapping) a PML focus in the nucleus. To avoid false positives, the

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Fig. 16.2 APB assay in soft tissue sarcomas (STS). Examples of combined PML immunofluorescence and telomere FISH in (a) frozen and (b) paraffin-embedded STS tissue sections. Indirect immunofluorescence with fluorescein (FITC) label was used for PML bodies and telomere FISH was performed using a Cy3-conjugated telomeric peptide nucleic acid (PNA) probe (See Color Insert)

telomeric DNA component of the APB must have a more intense fluorescence than the telomeres on that slide (for practical purposes, the working criterion used with the Cy3 conjugated telomeric probe is to require that with the appropriate camera exposure for the telomeric DNA component of the APB, the telomeres are not visible). The section is scored as positive for APBs if they are detected in 10 or more nuclei and in 0.5% or more of the cells in the section (Fig. 16.2a). To avoid artefacts, a cell is not considered to contain APBs if more than 25% of the colocalised foci occur outside nuclei (correcting for ratio of nuclear area to nonnuclear area). Slides are not scored as negative unless >2,000 tumour cell nuclei are examined.

16.3.4

Identification of ALT Genes by APB-Screening Assay

APBs are usually found in