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Characterizing canonical and non-canonical roles of telomerase reverse transcriptase in transformed human… Thompson, Connor Alexander 2016

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  !!! CHARACTERIZING CANONICAL AND NON-CANONICAL ROLES OF TELOMERASE REVERSE TRANSCRIPTASE IN TRANSFORMED HUMAN CELLS AND CANCER   by   Connor Alexander Thompson   BSc, McGill University, 2012   A THESIS SUBMITTED IN PARTIAL FULLFILLMENT OF  THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE   in   The Faculty of Graduate and Postdoctoral Studies  (Pharmaceutical Sciences)  The University of British Columbia  (Vancouver)  August 2016  © Connor Alexander Thompson, 2016 ! ii!ABSTRACT  Telomerase is the ribonucleoprotein reverse transcriptase that catalyzes the synthesis of TTAGGG nucleotide repeats at the ends of linear chromosomes, contributing to proper telomeric structure and cap formation. Most human somatic cells have low or undetectable telomerase expression. In contrast, telomerase overexpression is found in over 85% of human cancers allowing cancer cells to replicate indefinitely. Telomerase inhibition by GRN163L (Imetelstat) has previously been observed to potentiate genotoxic stress in a cell-cycle (S/G2) specific manner, through an unknown mechanism. We hypothesized that GRN163L treatment alters cell-cycle kinetics and that this effect depends upon active signaling through ataxia telangiectasia mutated (ATM). Here we tested the effects of combining GRN163L and the topoisomerase II inhibitor etoposide, together with pharmacological ATM inhibition on MCF-7 breast cancer cells, to assess dependence of telomerase’s cyto-protective function on this DNA-damage repair transducer. Additive increased cytotoxicity and cell-cycle profile alterations depended upon the order of treatment addition. Investigating possible causes of these cell-cycle distribution changes we observed that telomerase inhibition alone induces γH2AX DNA-damage foci in a subset of telomerase-positive cells but not telomerase-negative primary human fibroblasts. Additional FACS and immunocytochemistry experiments indicate that ! iii!GRN163L-treated cells were reversibly stalled but not arrested at G2/M. Our results suggest that treatment with GRN163L sensitizes telomerase-positive cells to cell-cycle specific DNA-damaging agents through the engagement of an ATM-dependent DNA-damage signal, which may represent a separate mechanism by which telomerase inhibition could affect DNA repair homeostasis in telomerase-positive cancer cells. ! In addition to its telomere-maintenance function telomerase has recently been reported to participate in non-canonical activities such as protection from DNA-damaging agents, apoptosis, cellular proliferation rate, and resistance to oxidative stress. In a separate study, we hypothesized that overexpression of telomerase in transformed human cells would increase their survival following exposure to DNA-damaging agents. Our results indicate that telomerase expression protects cells from a variety of DNA-damaging drugs by improving the kinetics of DNA-repair. Telomerase expression also allows surviving cells to tolerate increased levels of chromosomal instability following drug exposure. This work has implications on improving the design of future telomerase inhibition strategies to also target non-canonical effects of this enzyme.!!! iv!PREFACE Chapter 2: This material is from a manuscript in preparation. Jialin Xu provided the TRAP data (Supplementary Fig. 2-3). Arthur Chen assisted in the quantification of immunocytochemistry data (Fig. 8-10). I performed all other experiments shown. Helen Fleisig contributed to early stages of concept formation. Judy Wong and I conceived the experiments and wrote the manuscript. None of this text is taken from previously published articles.  Chapter 3: A version of this material has previously been published as Fleisig HB, Hukezalie KR, Thompson CA, Au-Yeung TT, Ludlow AT, Zhao CR, Wong JM. 2016. Telomerase reverse transcriptase expression protects transformed human cells against DNA-damaging agents, and increases tolerance to chromosomal instability. Oncogene 35(2):218-27. Some content may briefly appear in the introduction as well (Ch.1), though not verbatim. I have limited the scope of this manuscript to primarily focus on those experiments that I developed and worked on directly but have summarized the work of others from our lab in order to put the results in context. Kyle Hukezalie provided the data for the viability and FACS experiments outlined (Fig. 14). Kyle Hukezalie and Judy Wong provided the data for the chromosome-healing assay. Helen Fleisig prepared the metaphase spreads (Fig. 16) and I imaged and quantified them. I developed the assay and performed additional experiments covered here (Fig. 15). Judy Wong and Helen Fleisig conceived the experiments and Judy Wong and Kyle Hukezalie wrote the manuscript for the published paper.         ! v!TABLE OF CONTENTS  ABSTRACT ........................................................................................................................... ii PREFACE ............................................................................................................................. iv TABLE OF CONTENTS ........................................................................................................ v LIST OF TABLES ............................................................................................................... viii LIST OF FIGURES ............................................................................................................... ix LIST OF ABBREVIATIONS ................................................................................................. xi ACKNOWLEDGMENTS ..................................................................................................... xiv CHAPTER 1: INTRODUCTION ............................................................................................. 1 1.1. TELOMERES ........................................................................................................................... 1 1.1.1. Telomere structure and function ....................................................................................... 1 1.1.2. Consequences of telomere shortening ............................................................................. 5 1.2. TELOMERASE ........................................................................................................................ 7 1.2.1. Telomerase reverse transcriptase (TERT) ........................................................................ 7 1.2.2. Telomerase RNA (TER) and H/ACA proteins ................................................................... 9 1.3. TELOMERE MAINTENANCE BY TELOMERASE ............................................................... 10 1.4. TELOMERASE AND TELOMERES IN CANCER ................................................................. 14 1.4.1. Roles of telomeres in carcinogenesis and cancer progression ....................................... 14 1.4.2. Telomerase expression in cancer ................................................................................... 16 1.4.3. Telomere maintenance by alternative lengthening of telomeres (ALT) .......................... 17 CHAPTER 2: ....................................................................................................................... 19 2.1. INTRODUCTION ................................................................................................................... 19 2.1.1. Anti-telomerase agents for chemotherapy ...................................................................... 19 2.1.2. DNA-damaging chemotherapeutic drugs ........................................................................ 25 2.1.3. GRN163L and DNA-damaging drug combinations ......................................................... 26 2.1.4. DNA-damage response pathways and telomerase inhibition ......................................... 28 2.2. HYPOTHESIS ........................................................................................................................ 29 2.2.1. Specific aims ................................................................................................................... 29 2.3. METHODS ............................................................................................................................. 30 2.4. RESULTS .............................................................................................................................. 37 ! vi!2.4.1. Potentiation of Etoposide by GRN163L depends upon active ATM signaling ................ 37 2.4.2. Telomerase inhibition in telomerase-positive but not telomerase-negative cell lines increases the proportion of cells in G2/M phases of the cell cycle ........................................... 41 2.4.3. GRN163L treatment causes an increase in the proportion of cells that have 4N DNA content and this effect is removed by pre-treatment with an ATM inhibitor .............................. 47 2.4.4. GRN163L treatment induces G2 phase DNA damage foci only in telomerase-positive but not telomerase-negative cells ............................................................................................. 50 2.5. DISCUSSION ......................................................................................................................... 60 CHAPTER 3 ......................................................................................................................... 67 3.1. INTRODUCTION ................................................................................................................... 67 3.1.1. Non-canonical roles of telomerase in cancer .................................................................. 67 3.1.2. Role of telomerase at the mitochondria .......................................................................... 69 3.1.3. TERT domains and non-canonical functions .................................................................. 71 3.1.4. Strategies for studying non-canonical functions of telomerase ....................................... 71 3.1.5. Recombinant telomerase expression in ALT cells confers protection against cytotoxicity from DNA damage and increases kinetics of ds-break repair ................................................... 73 3.1.6. TERT’s telomere-repeat-synthesis activity is dispensable to DNA-damage tolerance ... 75 3.1.7. Cells with ~8N DNA content were detected in TERT-positive, but not vector control VA13 ALT cells, after induction of DNA damage ...................................................................... 76 3.2. HYPOTHESIS ........................................................................................................................ 78 3.2.1. Specific aims ................................................................................................................... 78 3.3. METHODS ............................................................................................................................. 78 3.4. RESULTS .............................................................................................................................. 81 3.4.1. Cytological abnormalities in VA13 cells following the induction of double-strand DNA damage are partially rescued with recombinant TERT expression ........................................... 81 3.4.2. Karyotype analysis of TERT-positive ALT cells with 8N DNA suggests that bypass of mitosis results in gross genomic instability ............................................................................... 84 3.5. DISCUSSION ......................................................................................................................... 86 CHAPTER 4 ......................................................................................................................... 88 4.1. CONCLUSIONS AND FUTURE DIRECTIONS ..................................................................... 88 4.1.1. Brief summary ................................................................................................................. 88 4.1.2. Telomerase inhibition and potentiation of DNA-damaging agents .................................. 89 4.1.3. ATM signaling and telomerase inhibition ........................................................................ 92 4.1.4. Non-canonical protective roles of TERT against DNA damage ...................................... 94 4.1.5. Future directions for telomerase inhibition in human anti-cancer chemotherapy ............ 96 ! vii!REFERENCES .................................................................................................................... 99 APPENDIX: Supplementary Figures .............................................................................. 117          !!    ! viii!LIST OF TABLES Table 1: Profiles of DNA-damaging drugs tested in combination with GRN163L………...27              !!! ix!LIST OF FIGURES Figure 1: Telomere cap structure in humans…………………………………………………..4 Figure 2: Modeling telomere length and telomerase expression in carcinogenesis……...15 Figure 3: Telomerase inhibition by GRN163L (Imetelstat)…………………………….…....24 Figure 4: GRN163L potentiation of etoposide cytotoxicity requires functional ATM signaling…………………………………………………………………………………………...39 Figure 5: Telomerase inhibition increases proportion of MCF-7, HT29, LS180 and MDA-MB 231 cells with 4N DNA content…………………………………………………………..…44 Figure 6: GRN163L reduces cell number in telomerase-positive cell lines following 7-day incubation in a dose-dependent manner………………………………………...………........45 Figure 7: ATMi and GRN163L treatments affect cell cycle population distributions……..49 Figure 8: Telomerase inhibition induces DNA damage foci in telomerase-positive, but not telomerase-negative cells…………………………………………………………………….....52 Figure 9: Cyclin B1/γH2AX ICC and FACS experiments indicate that GRN163L treatment increases the population of cells in late S/G2 phases……………………………………..…54 Figure 10: Functional ATM signaling is required for GRN163L-induced DDR foci formation and G2/M checkpoint stall…………………………………………………………………........57 ! x!Figure 11: Increased mitotic defects following release of cells from G2 stall by ATM inhibition…………………………………………………………………………………………...59 Figure 12: Suggested model of effects of GRN163L on telomere cap formation………...65 Figure 13: Canonical and non-canonical roles of telomerase reverse transcriptase…….70 Figure 14: Telomerase expression confers survival advantages against double-strand DNA-damaging agents in ALT cells…………………………………………………………….74 Figure 15. ALT cells exhibited high levels of nuclear morphological abnormalities following the induction of DNA damage; these abnormalities are partially rescued with telomerase expression………………………………………………………………………………………...83 Figure 16. Cytogenetic analysis of chromosome numbers with metaphase spreads……85   !   !!!! xi!LIST OF ABBREVIATIONS  ALT  alternative lengthening of telomeres ANOVA analysis of variance ATM  ataxia telangiectasia mutated  ATR  ataxia telangiectasia and Rad3 related ATP  adenosine triphosphate bp  base pair  CFU   colony forming unit assay CTE  C-terminal extension d-loop  displacement loop  DMEM Dulbecco’s Modified Eagle’s Medium DMSO Dimethyl sulfoxide dNTP  deoxynucleotide triphosphate DSB  double-strand break  EC50  effective concentration 50% FISH  fluorescence in situ hybridization H/ACA hinge ACA domain HAART highly active anti-retroviral therapy ! xii!ICC  immunocytochemistry kb  kilobase KO  knock out LD50  lethal dose 50%  mtDNA mitochondrial DNA NES  nuclear export signal NLS   nuclear localization signal  NNRTI  non-nucleoside reverse transcriptase inhibitor NRTI  nucleoside reverse transcriptase inhibitor nt   nucleotide(s) PD  population doubling PBS  phosphate-buffered saline PCR   polymerase chain reaction POT1  protection of telomeres 1 Rb  retinoblastoma protein RNP   ribonucleoprotein ROS   reactive oxygen species RPA  replication protein A ! xiii!RT  reverse transcriptase siRNA  small interfering RNA SSB  single-strand break ssDNA  single-stranded DNA T-loop  telomere loop TCAB1 telomerase Cajal body protein 1 TEN  telomerase essential N-terminus TER   telomerase RNA  TERT  telomerase reverse transcriptase  TIN2  TRF2- and TRF1-interacting nuclear protein 2 TRBD   telomerase RNA binding domain TRF1   telomere repeat binding factor 1 TRF2   telomere repeat binding factor 2 WST-1 water-soluble tetrazolium salt-1 WT   wild type WCL  whole cell lysate ! xiv!ACKNOWLEDGMENTS I would like to thank all those who have helped me and contributed to the completion of this thesis. Firstly, my supervisor Dr. Judy Wong who has taught me so much and serves as an excellent role model for what a good scientist should be like. I would also like to express my gratitude to the other members of the Wong Lab, particularly Jialin Xu for her assistance in the lab and Kyle Hukezalie for teaching me many techniques. I also thank my family for their support during the last few years. Finally, I want to thank Emily Lostchuck without whom I would not have made it through either my undergraduate or graduate degrees. I appreciate everyone’s help and wish you luck in the future. ! 1!CHAPTER 1: INTRODUCTION 1.1. TELOMERES 1.1.1. Telomere structure and function  Telomeres are nucleoprotein structures present at the ends of eukaryotic linear chromosomes. In mammals they are composed of repeated sequences of (TTAGGG)n nucleotides, their complementary strand, and associated proteins (1). Telomeres solve the problem of the lagging strand gap left by the RNA primers of DNA polymerase and associated chromatin degradation by acting as disposable non-coding buffers. This process termed the “end replication problem” contributes to the loss of 50-100 bp of 3’ terminal telomeric DNA with each replication cycle (2). Telomeres also distinguish natural chromosome ends from DNA damage or double-strand breaks (DSBs), and protect against erroneous recombination and random fusion events (3).  Chromosome ends were first recognized as important for cellular replication by McClintock as early as 1941 (4). The initial discovery of the sequence of telomeres was almost four decades later, first in Tetrahymena in 1978 (5) and eventually the human sequence was reported in 1988 (6). The telomeric (TTAGGG)n sequence is conserved across vertebrate species suggesting a shared origin in a common ancestor over 400 ! 2!million years ago (7). Telomeres are now universally recognized as crucial structures for maintenance of genome stability in humans and most higher eukaryotes. In humans telomeres typically range from 5-15 kb in length (8).  Telomeres are known to associate with sequence-specific single and double-stranded DNA binding proteins termed the shelterin complex, which support proper telomere function (Fig. 1 a) (9). Shelterin is composed of six subunits: the ds-telomeric DNA binding proteins telomere repeat binding factor 1 (TRF1) and telomeric repeat binding factor 2 (TRF2); the ss-telomeric DNA binding protein protection of telomeres 1 (POT1); and the other stabilizing/regulatory members of the complex repressor/activator protein 1 (RAP1), TRF1- and TRF2- interacting nuclear protein 2 (TIN2), and TPP1 (1, 10). Components of shelterin also play a role in regulating the recruitment and activity of the telomere maintenance enzyme telomerase (2, 3, 11, 12). Disruption of normal shelterin levels and stoichiometry, either through depletion or overexpression of its subunits can impair telomere and telomerase functions (10, 13).  Telomeres end in a G-rich ssDNA overhang that is the site of elongation by the telomere maintenance enzyme telomerase and must be distinguished from DNA damage by intracellular DNA-damage sensors (5, 14, 15). This overhang is sequestered by the formation of a higher-order capping structure composed of a displacement loop (d-loop) ! 3!caused by the G-rich ssDNA overhang looping back for invasion of the duplex DNA, to form the telomere loops or “T-loops” (Fig. 1 b) (6, 13). These structures were first observed by electron microscopy and their removal was found to result in activation of p53, inappropriate repair, and chromosomal fusions (7, 16). Shelterin proteins are also necessary for T-loop stabilization and proper function. For instance inhibition of TRF2 expression leads to loss of the 3’ DNA overhang due to the unrestrained actions of endonucleases ERCC1/XPF (8, 17). Telomere structure is complex and dynamically regulated during different phases of the cell cycle to facilitate optimal protection of chromosome ends and faithful inheritance of genetic information (9, 10, 13, 18).           ! 4!      Figure 1. Telomere cap structure in humans a) The six proteins composing the human shelterin complex bind specifically to telomeric DNA. Also depicted is the G-rich 3’ telomeric DNA overhang that is the site of nucleotide repeat addition by telomerase. b) Telomeric loops (T-loops) form by displacement of dsDNA by the ssDNA 3’-overhang. The G-rich overhang must be long enough for this function to take place. Thus critically short telomeres cannot successfully form T-loops leading to activation of an intracellular DNA-damage response (DDR). Shelterin proteins help stabilize these structures and are crucial for telomere protection. Figure adapted from work by Judy Wong, Helen Fleisig, and Raina Tamakawa. ! 5!1.1.2. Consequences of telomere shortening   Without the action of telomerase, telomeres are continually eroded with successive cellular replications; eventually reaching a short telomere checkpoint that activates cellular surveillance mechanisms, causing either arrest and senescence or apoptosis (19, 20). This is due to the requirement of RNA priming for synthesis of the Okazaki fragments complementary to the lagging strand during DNA-replication, which therefore cannot copy the 3’ telomeric termini where the last segment of RNA primer is removed (21, 22). Each time a cell divides this “the end replication problem” along with telomere cap processing after completion of replication contributes to the loss of 50-100 bp of telomeric DNA (23, 24). Telomere attrition has been determined to limit the replicative lifespan of cells in culture, a phenomenon known as the Hayflick limit, and may serve an evolutionarily adaptive anti-cancer function by preventing uncontrolled cellular growth (25, 26). When telomeres become critically short they can no longer form the T-loops necessary for proper capping function. Although it has also been suggested that some in vitro senescence is likely due to persistent DNA-damage response from inadequate culture conditions rather than strictly telomere attrition per se (27), telomeres play an important role in the induction of senescence as they are the primary sites of DDR foci accumulation over many rounds of cellular replication both in vitro and in vivo (28, 29, 30).  ! 6! Though telomeres progressively shorten with chronological age in most somatic tissues there is controversy with respect to the utility of using telomere length as a biomarker of organismal aging (31, 32). There is some established correlation between age, mortality, and telomere length, but as telomere length is highly variable within the human population and the direct contribution of short-telomere-induced senescence to the aging phenotype is uncertain, the predictive power of this marker is inconsistent (3, 33, 34, 35). Senescent cells in vivo often secrete inflammatory molecules in a process termed the senescence-associated secretory phenotype (SASP) (36). This process may promote loss of tissue homeostasis and cancer formation, thereby adding another possible mechanistic contribution of short telomeres to the diseases of aging (19, 37). Peto’s longstanding paradox observing that rates of cancer formation do not scale with body size and cell number in large long-lived mammals (38), may be partially explained by the presence of on average shorter inherited telomeres and lower telomerase activity in large long-lived mammals compared to that found in smaller short-lived mammals (39). Thus telomeres may function as a form of “mitotic clock” to limit the number of divisions possible for any given cell and thereby limit the accumulation of cancer-causing mutations (3). Furthermore, genetic lesions in telomerase subunits that lead to sub-optimal telomere ! 7!length maintenance underlie several premature aging syndromes such as dyskeratosis congenita, aplastic anemia, and idiopathic pulmonary fibrosis (40).  1.2. TELOMERASE 1.2.1. Telomerase reverse transcriptase (TERT)   Telomeres are maintained by the ribonucleoprotein (RNP) complex telomerase. The core catalytic components of the telomerase holoenzyme in vitro minimally consist of the telomerase RNA (TER) and the telomerase reverse transcriptase (TERT) protein (41, 42). Telomerase was first discovered in Tetrahymena by Greider and Blackburn in 1985, for which they were awarded the 2009 Nobel Prize in Physiology and Medicine (43, 44). The catalytic component of telomerase was first isolated in yeast (45), then in humans (46) and its structure and functional domains have been well characterized. Human TERT is a specialized and unique reverse transcriptase of 1132 amino acids encoded by the hTERT gene located at chromosome 5p15.33 (47). The three-dimensional architecture of TERT (and other eukaryotic TERT-like proteins) has been defined through single-particle electron microscopy, NMR, and comparative crystallographic studies across multiple species (48, 49, 50). TERT can be subdivided into ! 8!three domains with distinct functions; the N-terminal extension, the reverse transcriptase (RT) domain, and the C-terminal extension (CTE) (9). The N-terminus contains the telomerase essential N-terminal (TEN) domain which binds the ss-telomeric overhang during nucleotide addition, and the telomerase RNA binding domain (TRBD) that recognizes and interacts directly with TER (51, 52). The RT domain is the site of catalysis and shares structural and functional homology with retroviral reverse transcriptases (48,  53). The CTE is less well defined but TERT variants missing large portions of this region are defective for TER binding and are catalytically inactive (54).    Telomerase activity is regulated through atypical alternative splicing of TERT, with some splice variants unable to form functional holoenzyme complexes (55). This phenomenon has only recently been recognized likely due to the low levels of TERT expression found in most cells. There have been 22 distinct splice variants of TERT identified to date (56). The most common alternatively spliced TERT product, β-deletion-TERT (β-del-TERT), is found in a high ratio to full-length transcripts in human embryonic stem cells and cancer models (57, 58). β-del-TERT retains the N-terminal domains, but lacks the reverse transcriptase motifs, and is confirmed to not harbor any polymerase activity (58). The high levels of expression of TERT variants present at the protein level is ! 9!an interesting phenomenon and may suggest yet unexplored functional roles in cancer progression.   1.2.2. Telomerase RNA (TER) and H/ACA proteins   The telomerase RNA is a crucial component of the telomerase holoenzyme complex with roles in assembly, localization, structural stabilization, and as the template for hexanucleotide repeat addition by TERT (9, 42). TER is a 451 nt non-coding RNA consisting of the core domain (containing the ssRNA template used for repeat addition and the pseudoknot motif essential for holoenzyme assembly), conserved regions 4 and 5 (CR4/CR5), the template boundary element (TBE), and the box H/ACA domain, which contains the Cajal body localization sequence (CAB box) and biogenesis promoting box (BIO box) elements (9, 59, 60, 61). TER binds to telomerase at two independent sites on TERT creating the minimal core of the enzyme capable of generating telomerase activity in vitro (62).  In vivo, the H/ACA proteins dyskerin, Nhp2, Nop10 and Gar1 also associate with active telomerase, contributing to stabilization, cellular localization and proper assembly of the enzyme (9). The Cajal body RNA chaperone protein TCAB1 binds directly to TER and is essential for endogenous telomerase recruitment to telomeres (63, 64). Other proteins ! 10!associated with regulating the biogenesis, cellular localization and catalytic activity of telomerase include Hsp90, 14-3-3, pontin, reptin, NAT10, hnRNPs (subtypes C and U), and GNL3L though they are not permanent members of the RNP complex (65). TER like other small cytoplasmic RNAs (scRNAs) is extensively processed prior to performing its telomere length maintenance function as part of the mature telomerase holoenzyme complex (61, 66).    1.3. TELOMERE MAINTENANCE BY TELOMERASE   As a ribonucleoprotein reverse transcriptase, telomerase catalyzes the synthesis of (TTAGGG)n nucleotide repeats at the ends of linear chromosomes, maintaining the non-coding “buffer” necessary to distinguish natural chromosome ends from dsDNA breaks and preventing inappropriate repair and fusion events (67, 68). Telomerase activity is primarily regulated at the transcriptional level by the production of TERT, which is transcribed only in embryonic cell stages and in a few adult cell types such as germ line and stem/progenitor cells in humans (9). TERT protein and mRNA have been detected only transiently at low levels in telomerase-negative cells, while TER in contrast is expressed ubiquitously (69). Due to this abundance of RNA template even in adult human cells, recombinant TERT expression alone restores telomerase activity. Telomerase may ! 11!be generated in a “just in time” regulatory process as suggested by the presence of incomplete complexes (sometimes lacking either TER or TERT and their binding partners) detectable in various stages of assembly (70).    Both TER and TERT are known primarily to localize to the nucleus, transit through the nucleolus and arrive in the Cajal bodies to perform their canonical function of synthesizing telomeric hexanucleotide repeats (9, 64). TERT has also been observed in the cytoplasm where its function is less well characterized (63, 68). TERT has recently been reported to localize to the mitochondria under conditions of oxidative stress where it may play a role in protecting mtDNA from ROS (71, 72). TERT contains both a nuclear localization signal (NLS) and a nuclear export signal (NES) and the sequence of TERT nuclear import and its localization pathway are known (72-75).    Very little telomerase is needed to maintain telomeres and prevent senescence.  This has been estimated to be as low as ~1% of the TERT expression levels that are commonly found in cancers (76). The shortest telomeres within an individual cell appear to be preferentially elongated by telomerase (77). As the number of shelterin complexes is proportional to telomere length, this advances the possibility of a protein counting mechanism for the regulation of telomerase activity and recruitment (78). At telomere ends, telomerase also associates with members of the telomeric DNA-binding proteins of ! 12!the shelterin complex for proper alignment with the telomere terminal and efficient catalysis (13). TPP1 and TIN2 have been reported to interact directly with, recruit, and position telomerase on its telomeric substrate (11, 79). Additionally, the trimeric CST complex composed of Ctc1, Stn1, and Ten1 binds the G-rich overhang and is important for telomere cap formation and regulating telomere repeat addition (80). Also, ssDNA-binding protein 1 (SSB1) accumulation on the G-rich strand may further mediate recruitment of TERT (14) Telomerase is recruited to telomeres during the DNA-synthesis phase of the cell cycle where the actions of multiple RecQ helicases (WRN, BLM), exonucleases (Exo1, Apollo), homologous recombination pathway effectors (ATM, ATR, 53BP1), and histone aceltylation/methylation enzymes (DNMT1, SUV39H1/2, SIRT1) coordinate to allow the passage of DNA polymerase (and telomerase) to copy through the chromosomal end structures (T-loops) and restore the heterochromatin state of newly replicated telomeres (18, 65, 81-84). As previously mentioned, these higher-order capping structures are essential for proper telomere function and cell cycle progression. The current two-stage model of telomere shortening proposes the idea that when telomeres become critically short, T-loops are lost. This causes senescence and in the absence of adequate tumor suppressor functions, senescence bypass that leads to further telomere attrition. At this ! 13!second stage, TRF2 binding becomes sufficiently depleted and cells will undergo erroneous repair leading to massive genome instability or “crisis” (18, 85). A DNA-damage response mediated by the serine-threonine protein kinases ataxia telangiectasia mutated (ATM) and ataxia telangiectasia and Rad3-related (ATR) caused by dysfunctional telomeres has been reported to regulate telomerase recruitment at S phase (86).  Carol Greider recently proposed an alternate model for telomerase recruitment in which the enzyme travels with DNA polymerase along the moving replication fork. As the replication machinery copies through telomeres and more shelterin proteins are encountered, the tailgating telomerase experiences increasing likelihood of falling off of this entourage (87), explaining why shorter telomeres (with fewer bound shelterin units) are more likely to be extended through the action of telomerase.  Although untested, the strength of this model is in explaining a plausible mechanism for protein counting that distinguishes long telomeres from short telomeres, as direct mediators of signaling between short but functional telomeres and telomerase have not yet been identified.      ! 14!1.4. TELOMERASE AND TELOMERES IN CANCER 1.4.1. Roles of telomeres in carcinogenesis and cancer progression   Rarely, in the context of inactivated genomic surveillance machinery such as p53 and retinoblastoma protein (Rb), telomere shortening may progress to a crisis stage where critically short telomeres are recognized as DSBs leading to fusions, breakage-fusion-bridge cycles, and widespread genomic instability (3, 88). Massive cell death occurs at this point, and in order for the rare cell lineage to survive this stage and progress to malignancy, they must stabilize their genome through telomerase (1, 2, 89). This process of achieving stable telomere maintenance is often the last stage preceding the emergence of tumorigenic transformation (Fig. 2). Telomerase-positive cancer cells are thereafter dependent upon continued telomerase activity to maintain limitless replicative potential.        ! 15!        Figure 2. Modeling telomere length and telomerase expression in carcinogenesis    In telomerase-negative human somatic cells telomere length shortens with successive cellular divisions, eventually reaching a checkpoint stage where genomic surveillance mechanisms initiate proliferative arrest (senescence) or apoptosis. Loss of p53 and Rb tumor suppressor functions may allow further cell divisions to occur and severe telomere erosion. During this dysfunctional telomere state, known as “crisis” rampant genomic rearrangement can result in aneuploidy and cellular transformations. A subset of these cells will reactivate telomerase allowing for cellular immortalization. Telomerase activity provides telomere length maintenance and possible non-telomere-related utility to cancer cell fitness and survival.  In late stages of cancer development, this telomerase “addiction” provides a possible target for therapeutic intervention. Figure provided by Judy Wong and Kyle Hukezalie.  ! 16!1.4.2. Telomerase expression in cancer   Changes in telomerase expression and activity profile are associated with different human pathologies but are most prevalent in cancer. Telomerase activation is observed in 85-90% of all human cancers surveyed (90). This is a notable majority, despite the existence of an alternative telomere maintenance mechanism (alternative lengthening of telomeres or ALT) (91) (Section 1.5.3.). Human cancer cells express telomerase constitutively, which allow them to replicate indefinitely, at a level higher than that needed for telomere length maintenance. All cancer cells must find a mechanism to stabilize their telomeres for long-term growth; cancers without detectable telomerase activity have been observed to undergo spontaneous remission (92, 93).    Overwhelmingly, the most common mechanism of telomerase activation is through promoter mutations of the hTERT gene, which are among the most common non-coding mutations found across all cancer types (94-96). Active chromatin marks are found at these altered sites in cells with hTERT promoter mutations and have been observed correlate with levels of TERT expression (97). Although common, hTERT promoter mutations are not a universal feature of telomerase-positive cancers and the principle determinants of whether or not telomerase is expressed in these cells is unknown (98). Cancer cells can activate expression of the hTERT gene through a variety of mechanisms ! 17!including changes in epigenetic marks (97), alternative splicing (99), and possibly loss of auto-regulation due to telomere shortening as the gene is located in close proximity to the telomere at the end of chromosome 5 (100).  The reasons for the strong preference of cancer cells for telomerase expression over the ALT mechanism for telomere length maintenance remain largely unknown.  1.4.3. Telomere maintenance by alternative lengthening of telomeres (ALT) A subset of human cancers and transformed human fibroblasts in culture (10-15%) maintain their telomere length using the non-reciprocal homologous recombination-based mechanism termed alternative lengthening of telomeres (101, 102). ALT cancers exhibit several features consistent with maintenance by a recombination-based mechanism including high levels of circular telomeric DNA (103), heterogeneous telomere length (101), and elevated frequency of sister chromatid exchange (104). ALT cells also display high levels of non-TTAGGG variant repeats (94). These degenerate sequences are possibly generated due to the fact that the donor sequence for telomere elongation can be either a sister chromosome, a non-homologous chromosome, or possibly extrachromosomal circular telomeric DNA copied at non-fixed boundaries. (101, 105, 106). ALT telomeres have a unique epigenetic landscape and their non-canonical ! 18!telomeric sequences may not bind shelterin proteins with normal fidelity, leading to possible deficiency in function (107). A marker of ALT cells are the ALT-associated promyelocytic leukemia bodies (APBs) that contain telomeric DNA and shelterin proteins (108). These structures may facilitate recombination by bringing heterologous chromosomes into close proximity with each other (109). ALT is most commonly found in some sarcomas (osteosarcomas and liposarcomas) and sometimes tumors of the central nervous system where its prognostic significance is uncertain (91, 110). One potential consequence of ALT-positive status in cancers is that they are unlikely to respond to anti-telomerase therapies, as they do not depend on telomerase for telomere-length maintenance. Interestingly, removing telomerase activity in murine lymphomas using an inducible allele results in the emergence of ALT and metabolic changes (111). This has not been demonstrated in the context of long-term telomerase inhibition but may occur and warrants further study. Currently there are no therapies specific to ALT (91).  !!!! 19!CHAPTER 2:  !This material is from a manuscript in preparation. Running title: Transient telomerase inhibition alters cell cycle kinetics and sensitizes human breast cancer cells to G2-specific DNA-damaging agents  Thompson CA, Fleisig HB, Wong JMY.   Jialin Xu provided the TRAP data (Supplementary Fig. 2-3). Arthur Chen assisted in the quantification of immunocytochemistry data (Fig. 8-10). I performed all other experiments shown. Helen Fleisig contributed to early stages of concept formation and FACS experiments (Fig. 5). Judy Wong and I conceived the experiments and wrote the manuscript. None of this text is taken from previously published articles.   2.1. INTRODUCTION 2.1.1. Anti-telomerase agents for chemotherapy Strategies for telomerase inhibition   Telomerase reverse transcriptase (TERT), the catalytic subunit of telomerase, is constitutively expressed in cancer and transformed human cells but repressed or only transiently expressed in somatic human cells. As such, telomerase inhibition is an attractive therapeutic target (3). The goal in anti-telomerase therapy is to selectively drive cancer cells to senescence or apoptosis while leaving normal cells unharmed. Since cancer cells generally have shorter telomeres than normal cells the danger of transiently ! 20!inhibiting telomerase should be minimal (112). There have been many diverse approaches tested for extinguishing telomerase activity, but to date (July, 2016) none have been approved for clinical use. Telomerase inhibition by different interventions has been demonstrated to slow proliferation and force cancer cells to apoptosis in multiple studies (113, 114).    As the structure and mode of action of the catalytic subunit TERT is well characterized, a number of strategies have focused on inhibiting the activity of TERT in order to selectively target cancer cells as a therapeutic approach (48, 115). The non-nucleoside reverse transcriptase inhibitor (NNRTI) BIBR1532 binds to the thumb domain of TERT impairing its binding to TER and has been demonstrated to effectively inhibit telomerase activity (116). As TERT is structurally and mechanistically similar to viral reverse transcriptases, inhibitors of viral reverse transcriptases including all nucleoside reverse transcriptases (NRTIs) commonly used in highly active anti-retroviral therapy (HAART) for HIV (zidovudine, stavudine, tenofovir, didanosine, abacavir), inhibit telomerase activity in vitro (117). This has relevance for possible accelerated aging in patients undergoing HAART but is unlikely to be very useful for anti-cancer chemotherapy as evidenced by the high incidence of cancer in this group (118). No drug directly targeting TERT has progressed to clinical trials yet. TER as the other core component of ! 21!telomerase has also attracted interest as a possible therapeutic target and the anti-sense oligonucleotide GRN163L (Imetelstat) developed by Geron Corporation is the most promising telomerase inhibitor developed to date (See    Indirect approaches to target telomeres or reduce telomerase activity have also been investigated. Telomeres harbor high levels of the tertiary planar DNA structures called G-quadruplexes formed by non-Watson-Crick Hoogsteen base pairing between tracts of guanine repeats (119, 120). These structures present a barrier to the replication machinery and can kill rapidly proliferating cancer cells when treated with stabilizing ligands (121, 122).  Preferentially stabilization at telomeres, using G-quadruplex targeting ligands specific to topologies found at telomeres may have scientific and therapeutic applications (123, 124). Targeting factors involved in telomere homeostasis such as tankyrase, TERT protein stability (HSP90), or the 3’-overhang of telomeres with complementary oligos (so called “T-oligos”) that cause a persistent DDR have also been investigated (114). Instead of inhibiting telomerase, a new approach being explored takes advantage of existing telomerase activity in cancer cells by incorporating nucleoside analogs that do not bind shelterin proteins, such as 6-thio-2’-deoxyguanosine, in order to more rapidly induce telomere dysfunction (125, 126). Immunotherapies against TERT fragments presented on the cell membrane of cancer cells by the human leukocyte ! 22!antigen (HLA) class I pathways are another promising approach to cancer treatment and possibly prevention. There are three anti-TERT vaccines (GV1001, GRNVAC1, and Vx001) that have been tested in stage II and III clinical trials examining efficacy against non-small-cell lung cancer, acute myelogenous leukemia, and metastatic pancreatic cancer (114, 127). Although well tolerated, it is currently unclear if these vaccines will progress to larger multi-center trials. GRN163L (Imetelstat)   The most advanced clinical strategy in telomerase inhibition is Imetelstat (GRN163L) (114, 128). GRN163L is a synthetic, lipid-conjugated, 13-mer oligonucleotide N3’ P5’ thio-phosphoramidate complementary to the template of the telomerase RNA component, where it acts as a competitive antagonist (Fig. 3 a) (129). This structure binds RNA with high efficiency, is stable in in vivo conditions, is resistant to nucleases, and can readily pass through cell membranes due to the fatty palmitoyl group. GRN163L binding blocks the normal association of TER with the chromosomal substrate, efficiently inhibiting its ability to synthesize de novo telomeric repeats (Fig. 3 b). While successful in multiple cell culture and xenograft models (130-132), clinical trials have yielded mixed results with several early study terminations due to safety and efficacy concerns (see comprehensive ! 23!summary of trials to date here (114)). The specific cause of the high incidence of reported side effects has not been demonstrated but has been suggested to involve interference with putative roles of telomerase in hematopoietic progenitor cell maturation (114). Two recent trials for the hematological proliferative disorders essential thrombocythemia and myelofibrosis show more promising results (128,133). In late 2015 the FDA assigned GRN163L orphan drug status for myelofibrosis and myelodysplastic syndromes affording it certain developmental incentives (134). GRN163L also displays some off-target effects on cell adhesion independent of telomerase activity that may have anti-metastatic potential (135, 136). This could conceivably be due to GRN163L forming intermolecular G-quadruplexes and is an avenue our lab will likely examine in the near future.      ! 24!  Figure 3. Telomerase inhibition by GRN163L (Imetelstat) a) GRN163L is a synthetic lipid-conjugated oligo complementary to the template region of TER. Its thio-phosphoramidate backbone allows it to bind RNA with high affinity, avoid inducing RNAse H activity, and increases resistance to cleavage by cellular nucleases (137)  b) GRN163L blocks substrate recognition and binding by telomerase, which is the first step of telomere elongation. Telomerase activity is efficiently abolished by GRN163L treatment.   ! 25!2.1.2. DNA-damaging chemotherapeutic drugs   Mainstays of current cancer treatment include surgery, radiation, and chemotherapy. DNA-damaging agents were historically some of the first anti-cancer chemotherapeutic drugs developed, such as the World War I chemical warfare-inspired nitrogen mustard compounds β-chlorethyl amines (138) and the paradigm-shifting folate analog aminopterin used for the first ever treatment of childhood acute lymphoblastic leukemia (139). Cancer cells are especially vulnerable to DNA damage due to selection for fast proliferation, which often results in reduced stringency of DNA damage-sensing pathways and repair, as well as the ability to bypass cell cycle checkpoints. DNA double-strand breaks (DSBs) are among the most toxic types of DNA lesions to cells and can form due to direct actions of chemical compounds causing DSBs; or other types of damage leading to replication errors during S-phase DNA-replication (140). The major categories of DNA-damaging agents are: those that directly interact with DNA such as alkylating-like platinum agents (141), antimetabolites that impair some aspect of nucleotide metabolism or DNA synthesis like 5-fluorouracil (5FU) or methotrexate (142), and topoisomerase poisons that cause DSBs by affecting interactions between proteins and DNA (143). The primary limiting factors of chemotherapeutic drug treatments for ! 26!cancer therapy are toxicity to the patient and acquired adaptive resistance obtained by cancer cells (144).  Topoisomerases release the DNA double helix from torsional strain generated by unwinding during DNA replication or transcription by creating transient DNA breaks that allow the DNA to relax and pass through the gap created and then re-ligating the strands. Topoisomerases come in two types: topoisomerase I, which creates a single-stranded break (SSB) allowing the DNA strand to relax before being reannealed; and topoisomerase II, which creates a DSB before passing the supercoiled DNA through it (140, 145). Topoisomerase poisons bind to the cleaved-DNA-topoisomerase complex and block the re-ligation step. This later results in the formation of DSBs when encountered by a moving replication fork during cell division (146). Due to this mechanism of action topoisomerase inhibitors are most active during and immediately following the cell division phase of the cell cycle (S/G2 phases) and cytotoxic activity correlates with intracellular topoisomerase levels (147).  2.1.3. GRN163L and DNA-damaging drug combinations   Previous work in our lab demonstrated that combining telomerase inhibition by GRN163L with genotoxic agents potentiated the cytotoxic effects of some, but not all, ! 27!types of DNA-damaging agents tested (148). The cytotoxic responses to the topoisomerase inhibitors irinotecan and etoposide (with preferential action against topoisomerase I and II respectively) were augmented by telomerase inhibition in MCF-7 breast cancer cells and the colon cancer cell lines HT29 and LS180. In contrast, telomerase inhibition had no potentiating effect on the non-cell-cycle-specific crosslinking agent oxaliplatin or the direct DNA-break inducer bleomycin. This study was the first to show short-term potentiation of DNA-damaging agents by pharmacological telomerase inhibition.  Drug Mechanism of action Cell cycle specificity Potentiated by GRN163L Bleomycin Direct DNA break inducer Non-specific No Oxaliplatin Crosslinking agent Non-specific No Irinotecan Topoisomerase I inhibitor S/G2 Yes Etoposide Topoisomerase II inhibitor S/G2 Yes  Table 1: Profiles of DNA-damaging drugs tested in combination with GRN163L DNA-damaging drug profiles and results from viability experiments in our previous study combining these drugs with the telomerase inhibitor GRN163L. The agents potentiated by GRN163L are both topoisomerase inhibitors with primary activity during S/G2 phases of the cell cycle (149). ! 28!2.1.4. DNA-damage response pathways and telomerase inhibition  In the same study, combining GRN163L with chemical inhibition by the specific inhibitor KU55933 of the serine/threonine protein kinase Ataxia telangiectasia mutated (ATM) further enhanced cytotoxicity of etoposide in two out of three cell lines tested (increased cytotoxicity was observed in LS180 colorectal carcinoma and MCF-7 breast adenocarcinoma but not in HT29 colorectal carcinoma line) (148). These observations are interesting in light of current models of telomerase recruitment involving active DDR signaling through ATM and ATR (86). Activation of specific DNA damage response is found transiently in S phase during normal DNA replication at unstructured open telomere ends and has been detected in the form of ATM-MRN and ATR-ATRIP complexes (150, 151).  Also it has recently been reported that ATM activity is required for telomere elongation in mouse and human cells (152).   In the study outlined in this chapter, we aimed to deduce how ATM signaling interacts with telomerase inhibition by GRN163L and results in the increased sensitivity to cytotoxicity by the S/G2-specific DNA-damaging agent etoposide. We also set out to characterize the cytotoxic mechanism and possible DDR pathway effectors or regulators through which telomerase inhibition potentiates select DNA-damaging agents. Our data demonstrate that the presence of telomerase activity affects cell cycle kinetics by ! 29!promoting clearance of DDR signals at G2 phase, coinciding with the timing of telomere replication and cap formation. This advantage in cell-cycle progression kinetics provides a strong impetus for oncogenic transformed cells to choose telomerase over other telomere length maintenance mechanisms. Our data provide the incitement to redesign clinical use of telomerase inhibition strategies in a telomere length-independent manner.  2.2. HYPOTHESIS  Transient telomerase inhibition induces changes in cell cycle kinetics, which are dependent upon active ATM signaling. T-loop formation and clearance of cell cycle checkpoints will be delayed creating telomere structural vulnerability. 2.2.1. Specific aims (1) Study the interactions between telomerase inhibition, ATM signaling and the potentiation of etoposide cytotoxicity in MCF7 breast cancer cells. (2) Investigate the relationship between telomerase inhibition by GRN163L treatment and changes in cell cycle kinetics, through the actions of the ATM signaling pathway in cancer cell models  ! 30!2.3. METHODS Cell lines and Reagents   MCF-7, MDA-MB 231, HT29, LS180, BJ fibroblasts, and WI-38 VA-13 were obtained from the American Type Culture Collection (ATCC). MDA-MB 231 (NucLight Red) cells were obtained from Essen Bioscience. Cell culture media, antibiotics and other cell culture reagents are commercially available from Invitrogen/Life Technologies unless otherwise noted. Cells were maintained under standard culture conditions of 37°C and 5% CO2 with 100U each of the antibiotics penicillin and streptomycin, in the presence of appropriate Fetal Bovine Serum (FBS) concentrations ranging from 5-15% as indicated by ATCC. Etoposide was obtained from Sigma/Aldrich. ATM inhibitor KU55933 was obtained from Calbiochem. KU55933 treatments were administered as 10 µM in DMSO, as previously determined to efficiently inhibit ATM function (148). GRN163L (Imetelstat) and GRN mismatch (GRN-MM) control were obtained from Geron, resuspended in PBS and vortexed for 15 min before determining the stock concentration for each experiment using UV-spectrophotometer absorbance. At high concentrations, GRN163L is known to have solubility issues and requires oligonucleotide concentration measurements before each experiment. GRN163L/GRN-MM treatment was administered at 10 µM final concentrations, a dose previously determined to reduce telomerase activity at least 100-! 31!fold in these cell lines (148). Replacement of media took place 2-3 times per week and cells were passaged upon reaching 80-90% confluency.  Colony forming unit assay (CFU)   Assay was performed essentially as previously described (148). 3 x 105 MCF-7 cells per well were seeded in 6-well plates, allowed to settle and pretreated with GRN163L or ATMi (KU55933) for 24 hours prior to addition of a serial dilution of etoposide (2-fold 12-points, 12.8 µM highest dose) in combination with the previously used inhibitor or in combination with both inhibitors present. Five treatment conditions were tested no pre-treatment (NT), GRN163L for 48 h (GG), KU55933 for 48 h (AA), GRN163L for 24 h followed by both GRN163L and KU55933 for 24 h (GB), and KU55933 for 24 h followed by both GRN163L and KU55933 for 24 h (AB) were the conditions tested. After incubation the cells were harvested with trypsin/EDTA, counted, seeded at 3 x 105 cells per well in 6-well plates in 0.95 ml 0.4% agar, 1x DMEM (45%), 2x DMEM (20%), and 10% FBS over a 2 ml base layer consisting of 0.53% agar (final concentrations of 1x DMEM and 10% FBS were the same as the top layer) and allowed to grow into individual colonies at 37°C under 5% CO2 for 2 weeks. Colonies were scored after 14 days and counted as positive if their size exceeded 50 µm. Duplicate wells were counted for each point and each experiment ! 32!was repeated four times. Dose response curves of MCF-7 cells pre/co-treated with indicated inhibitors were constructed and LD50s generated for comparison. Each count was normalized to the inhibitor only no-etoposide wells as an internal control and expressed as a percentage loss for curve construction. Dose-response analysis was undertaken using GraphPad Prism (v6.0b).  Immunocytochemistry (ICC)   2.75 x 104 cells per well were seeded in 6-well plates and allowed to settle on glass coverslips overnight before 24-48 h incubation with indicated treatments. Cells were fixed with 3.7% Pfa in CSK buffer for 15 min at room temperature, washed twice with 1x PBS, and either stored at 4°C or stained immediately. Fixed cells were permeabilized with PBS-t (PBS + 0.1% Triton X-100) for 5 min and blocked with 4% BSA in TBS for 1 h at room temperature. Primary antibodies were sourced and diluted in blocking solution as follows: anti-phospho-histone H2AX (Ser139) 1:500 (JBW301 EMD Millipore, stock 1mg/ml), phospho-H3 1:500 (06-570 EMD Millipore, stock 1mg/ml), cyclin B1 1:100 (Santa Cruz H-433, stock 200 µg/ml) and incubated overnight with agitation at 4°C. Cells were then washed 3x for 10 min with PBS-t. Secondary antibodies (AF488 and Texas Red, Abcam) were incubated for 1 h at room temperature in blocking solution. Cells were washed 3x ! 33!with PBS-t then incubated for 5 min with DAPI (1:125000, stock 5 mg/ml). Cells were washed again 2x with PBS then mounted on slides with Fluoromount G (SouthernBiotech). Images were collected using a Zeiss LSM 700 confocal microscope and Zen 2012 (Zeiss) software. For quantification of DDR foci and nuclear abnormalities, images were collected, blinded and scored with >400 cells analyzed for each condition. Nuclear abnormalities were scored manually based on standard binning criteria (179).   Fluorescence-activated cell sorting (FACS)   Cells were plated at 3 x 105 cells per well in 6-well plates and allowed to settle. After treatment for 24-48 h with the described drug regimes, cells were harvested by trypsinization and fixed with EtOH (cells suspended in 1 ml 1% FBS/PBS added drop wise to 4 ml 100% EtOH). Cells were then stained immediately or stored at -20°C. FITC-Anti-Cyclin B1 (BD Biosciences) immunostaining and propidium iodide nucleic acid labeling were conducted according to manufacturer’s protocol (Becton Dickinson), followed by 15 min RNAse treatment at 37°C to remove non-specific signals. Labeled cells were sorted using the BD LSRII flow cytometer (UBC Life Sciences Institute). Data was analyzed using Flow Jo (Tree Star Inc.) software.  ! 34!Whole cell lysate preparation     Cells were harvested and washed 2x with ice-cold PBS and centrifuged for 5 min at 1,500 rpm at 4°C. Cell lysate was prepared in 500 µl of hypotonic cell lysis buffer (20 mM Hepes, 2 mM MgCl, 21.0 mM DTT, 0.2 mM EGTA, 0.1 mM PMSF, 10% glycerol). Cell suspensions were then snap frozen in liquid nitrogen for 30 sec, thawed at 37°C in a water bath and vortexed well. This freeze/thaw procedure was repeated three times. 5.0 M NaCl was then added to the lysate in two parts to a final concentration of 400 mM and incubated on ice for 15 min for soluble nuclear protein extraction. Lysates were cleared by centrifugation at 13,200 rpm for 15 min and supernatant (whole cell lysate fraction) was saved in aliquots for subsequent analysis. Assay was conducted essentially as previously described (149, 153).   Bradford protein assay     Protein concentrations of whole cell lysate fractions were measured according to manufacturer’s instructions using the Bradford protein assay (Bio-Rad Laboratories Inc., Hercules, CA) essentially as previously described (149, 153). Dye reagent was prepared by dilution of one part Dye Reagent Concentrate 1:5 in deionized water. A protein standard series was prepared using a 2-fold dilution series of BSA (0.0625-0.5 mg/ml). ! 35!Whole cell lysate samples were prepared by diluting 2 µl of each sample with 38 µl of lysis buffer. Each standard and diluted sample were then pipetted (10 µl) into separate wells in a 96-well plate before 200l of the diluted dye reagent was added to each well. The plate was incubated at room temperature for 5 min and absorbance was measured at 595 nm. The linear equation generated from the sample standards was used to determine protein concentration.  Telomerase Repeat Amplification Protocol (TRAP)   Telomerase activity analysis was conducted essentially as previously described (149). Whole cell lysate from MCF-7 or MDA-MB 231 treated with indicated drug regime was used as the source of telomerase. TRAP assay was performed using 5, 1, 0.2, and 0.08 µg (5-fold dilution series) of.WCL. The forward primer used was M2 (5’-AATCCGTCGAGCAGAGTT-3’), and addition by telomerase of the 6-nt telomeric repeats to the 3’-end of the primer were carried out at 30° C for 1 h, in the presence of 2 ng/l M2 primer, 1x PCR buffer (20 mM Tris-Cl (pH = 8.0), 15 mM MgCl2, 680 mM KCL, 10 mM EDTA, 0.5% Tween 20), and 50 M dNTPs. Reactions were heat-inactivated at 95°C for 30 sec. Primer extension products were then amplified by PCR using 1.25U Pfu DNA polymerase and 0.1 g of a Cy5-fluorophor-labeled reverse primer CX3 (CY5-5’-! 36!CCGCGCCCTAACCCTAACCT-3’) in 1x PCR buffer with 5.0 x 103 copies of ITAS internal control. Resulting products were analyzed by gel electrophoresis using 10% nondenaturing polyacrylamide gel (10% bis:acryl (19:1), 0.5x TBE, 0.1% APS, 0.1% TEMED) and visualized using the TyphoonTM scanner fitted with the appropriate laser and filters (GE HealthSciences).   Cell proliferation assays   MCF-7 and VA-13 (vector control) cells were seeded at 5 x 104 cells per well in 6-well plates and treated as indicated for 7 days for coulter counter experiments. At indicated time points, cells were harvested with trypsin/EDTA and 200 µL counted according to manufacturer’s instructions in order to calculate total cell number (Beckman Coulter). Nuclear counts for MDA-MB 231 NucLight Red cells were obtained by seeding at 5 x 103 cells per well in 96-well plates and imaged every 2 h using the Essen Bioscience IncuCyte Zoom™ in an incubator at 37°C. Media with added drug was replaced every 3 days.  ! 37!Data analysis   GraphPad Prism version 6 (GraphPad Software Inc., San Diego, CA) was used for statistical analysis and data presentation. Student’s t-test was used to generate P-values or where appropriate ANOVA followed by Fisher’s LSD test. P < 0.05 were considered statistically significant.   2.4. RESULTS  2.4.1. Potentiation of Etoposide by GRN163L depends upon active ATM signaling   In order to clarify the role of ATM signaling in GRN163L-mediated potentiation of etoposide cytotoxicity, we performed order-of-treatment-addition experiments using GRN163L (10 µM) and ATM inhibition by KU55933 (10 µM) with an optimized colony forming unit assay (148). This GRN163L dose had previously been determined to inhibit telomerase activity by >100 fold in multiple cancer cell types. MCF-7 breast cancer cells were pre-treated for 24 h with GRN163L or KU55933 before addition of the S/G2-specific DNA-damaging drug etoposide with continued inhibitor treatment or in combination with both GRN163L and KU55933 for 24 h. After 48 h treatment, cells were harvested and set ! 38!in soft agar medium to recover for 2 weeks (Fig. 4 a). In agreement with previous results GRN163L (GG) or KU55933 (AA) treatment alone significantly potentiated etoposide toxicity (Fig. 4 b, c) (Supplementary Fig. 1) (148). The potentiation effects of ATM inhibition on etoposide toxicity were consistently higher than that from GRN163L treatments. This observation is attributed to the major roles of ATM in the signal transduction regulation of cellular responses to dsDNA damages, in addition to its role in telomere maintenance and structural homeostasis. In agreement with our previous data, treatment with both inhibitors in combination had an additive effect on etoposide cytotoxicity when the cells were pre-treated with GRN163L first (GB). In contrast, this additive effect was lost when cells were pre-treated with KU55933 before GRN163L and etoposide addition (AB), suggesting that functional ATM-dependent DDR signaling is required for potentiation of etoposide’s genotoxicity by GRN163L.        ! 39!a)    b)    MCF-7 GRN163L/ATMi  (n=4)10-3 10-2 10-1 100 1010255075100CFU loss (%) NTEtoposide (µM)GGAAGBAB! 40! c)       Figure 4. GRN163L potentiation of etoposide cytotoxicity requires functional ATM signaling a) MCF-7 breast adenocarcinoma cells were pretreated with 10 µM GRN163L or ATMi (KU55933) for 24 h prior to addition of a serial dilution (12.8 – 0.0125 µM) of etoposide (E) in combination with the previously used inhibitor or in combination with both inhibitors. After 24 h incubation the cells were counted, seeded in soft agar, and colonies scored after 14 days. b) Dose-response curves of MCF-7 cells pre/co-treated with indicated inhibitors. No pre-treatment (NT), GRN163L, two doses at T = 0 h and T = 24 h for a total of 48 h (GG), KU55933, two doses at T = 0 h and T = 24 h for a total of 48 h, (AA), GRN163L for 24 h then both GRN163L and KU55933 for 24 h (GB), KU55933 for 24h then both GRN163L and KU55933 for 24 h (AB). Error bars represent SEM c) Calculated LD50s from CFU data. ANOVA and Fisher’s LSD test were used to generate P-values (* = P ≤0.05, ** = P ≤ 0.01, *** = P ≤ 0.001, **** = P ≤ 0.0001). Error bars represent SD.  NT GG AA GB AB0. LD50sLD50 Etoposide [ ] µM********ns**! 41!2.4.2. Telomerase inhibition in telomerase-positive but not telomerase-negative cell lines increases the proportion of cells in G2/M phases of the cell cycle   To better understand how telomerase inhibition contributes to the increased genotoxicity of S/G2 chemotherapeutic agents, we measured the effect of GRN163L treatment on cell cycle progression through analysis of DNA content by fluorescence-activated cell sorting (FACS). Cells were treated with GRN163L or its mismatch control (GRN-MM) oligo for 24 h and cell cycle profiles at the time of harvest were visualized with propidium iodide (PI) staining.    In telomerase-positive MCF-7 and MDA-MB 231 (mammary adenocarcinomas) cells and HT29 and LS180 (colorectal carcinomas) cells, we observed a significant increase in cell population with 4N DNA content following GRN163L but not with GRN-MM treatment (Fig. 5 a-d). In contrast, the cell cycle profiles of telomerase-negative transformed cells (VA-13, which maintain their telomeres with the alternative lengthening of telomere mechanism) and primary human foreskin fibroblasts (BJ) were unaffected by both GRN163L treatment, and its mismatched control (Fig. 5 e-f).    GRN163L treatment induced a small but reproducible change in cell cycle profiles. We reasoned that the observed increase in 4N cell population may represent a delay in ! 42!clearance from G2 stage of the cell cycle rather than a cell-cycle arrest, as cell growth recovered following the removal of GRN163L, as observed in previous CFU assays. Accordingly, if GRN163L treatment were causing a delayed progression at G2/M of the cell cycle we would predict that these effects would be apparent in long-term proliferation assays. We used the IncuCyte Zoom live cell imaging system and red fluorescent NucLight tagged MDA-MB 231 cells to measure the effects of continuous GRN163L treatment on cell counts over 7 days. Consistent with a small but accumulative growth disadvantage conferred by delayed clearance from G2 phase, we observed lower nuclear object counts with 10 µM GRN163L treatment (Fig. 6 a). A 2 µM GRN163L treatment appeared to display a trend toward an intermediate effect on growth, in agreement with the partial inhibition of cellular telomerase activity (TRAP) by this intermediate dose of GRN163L.   In order to measure growth in our parental MCF-7 cells we employed the same treatment regime and measured absolute cell counts by Coulter counting. Similarly we saw a small growth deficit in MCF-7 apparent with 10 µM GRN163L treatment (Fig. 6 b). In contrast, in telomerase-negative VA-13 cells GRN163L treatment had no effect on cell counts (Fig. 6 c). Telomerase activity was effectively inhibited by 10 µM and partially inhibited by 2 µM GRN163L in MDA-MB 231 as measured by telomeric repeat ! 43!amplification protocol (TRAP) (Supplementary Fig. 2). These results are consistent with a small accumulative growth disadvantage apparent over long time courses of continuous treatment with GRN163L. Cell density also appeared to plateau earlier with 10 µM GRN163L treatment in both MCF-7 and MDA-MB 231. This is possibly related to effects of GRN163L on adhesion as previously described (135, 136)    Together, our data suggest that treatments with GRN163L cause changes in cell cycle kinetics.  Conceivably, the delayed clearance from G2 phase induced by treatments with GRN163L is engaged through the activation of telomeric DDR pathways. Further activation of the same DDR pathways at G2 (by topoisomerase inhibitors) may contribute to an increase in cellular stress in this particular cell cycle phase beyond a tolerable threshold.      ! 44! ! 45!Figure 5. Telomerase inhibition increases proportion of MCF-7, HT29, LS180 and MDA-MB 231 cells with 4N DNA content No effect was observed in non-telomerase expressing cell lines VA-13 (vector control) and BJ. Indicated cell lines were treated for 24h with 10 µM GRN163L or the mismatch oligo (GRN-MM) control before harvesting and PI staining for FACS analysis. Representative cell cycle profiles are shown for indicated treatments and cell lines. Histograms indicate ratio of cells with 4N DNA content compared to 2N DNA content. ANOVA and Fisher’s LSD test were used to generate P-values (* = P ≤0.05, ** = P ≤ 0.01, *** = P ≤ 0.001).   a)  0 20 40 60 80 100 120 140 160050010001500Time (hr)Red Object Count  (1/mm2 )MDA-MB 231 Red (n=3)NT2 µM GRN163L10 µM GRN163L*! 46!b)                             c)                  0 2 4 6 8010203040DayCell Count (Normalized to Day 0)MCF-7 (n=3)NT2 µM GRN163L10 µM GRN163L********0 2 4 6 80510152025DayCell Count (Normalized to Day 0)VA-13 (n=3)NT2 µM GRN163L10 µM GRN163L! 47! Figure 6. GRN163L reduces cell number in telomerase-positive cell lines following 7-day incubation in a dose-dependent manner a) MDA-MB 231 (NucLight Red) cells were continuously treated with GRN163L (feeding every 3 days) for 7 days. Red object counts (representing nuclei expressing mKate2 red fluorescent protein) were automatically collected every 2 h using the Essen IncuCyte Zoom. b-c) Telomerase-positive MCF-7 cells and telomerase-negative VA-13 (vector control) cell numbers were assessed by Coulter counter counts at indicated time points under the same treatment regime. ANOVA and Fisher’s LSD test were used to generate P-values comparing 10 µM GRN163L to the no treatment condition at same time point (* = P ≤0.05, ** = P, *** = P ≤ 0.001).    2.4.3. GRN163L treatment causes an increase in the proportion of cells that have 4N DNA content and this effect is removed by pre-treatment with an ATM inhibitor     Next, we examined the effects of GRN163L/KU55933 order-of-addition treatment regimens on cell cycle progression with FACS to see whether these GRN163L-dependent cell cycle profile changes are contingent upon active ATM signaling. Again we observed an increase in cell population with 4N DNA content following GRN163L treatment in MCF-7 cells (Fig. 7 a), indicating a likely phase-stalling/delayed clearance at G2. This effect on cell cycle progression was abrogated by KU55933 pretreatment but not by treatment with KU55933 following 24 h of GRN163L exposure. These order-of-addition effects mirror the ! 48!rank order of increased sensitivities towards etoposide treatment observed in our previous CFU experiments.    We analyzed the sub-G1 populations as a functional read-out of treatment toxicities. The proportion of sub-G1 cell populations were not significantly affected by GRN163L or KU55933 treatments alone, or in any treatment combinations, indicating that these changes in cell cycle profiles are likely not due to increased rates of apoptosis or cell death (Fig. 7 b). Additionally, we tested whether KU55933 treatment may be interfering with the efficiency of telomerase inhibition by GRN163L. Telomerase activity was assayed using TRAP and we observed no impact of ATMi treatment on GRN163L’s telomerase inhibition efficiency (Supplementary Fig. 3). Together, these results indicate that cell cycle alterations induced by GRN163L are dependent on functional ATM signaling.       ! 49!a)                b)     Figure 7. ATMi and GRN163L treatments affect cell cycle population distributions a) MCF-7 cells were treated with the indicated inhibitors in the same scheme as Fig. 4 for 48 h before being fixed and analyzed for DNA content by PI FACS. b) Possible contributions of apoptosis and cell death to altered cell cycle profiles were assayed by comparing sub-G1 populations of different treatment regimes. ANOVA and Fisher’s LSD test used to generate P-values (* = P ≤0.05).  NT GG AA GB AB0. ATMi/GRN163L (n=6)4N/2N RatioNTGGAAGBAB* ***NT GG AA GB AB02468MCF-7 ATMi/GRN163L (n=6)% Sub G1NTGGAAGBAB! 50!2.4.4. GRN163L treatment induces G2 phase DNA damage foci only in telomerase-positive but not telomerase-negative cells   GRN163L binds to the template region of TER and blocks telomerase’s access to the telomeric substrate. Even though G-rich overhangs can be generated through the actions of exonucleases Apollo and EXO1 (18, 83), de novo synthesis of telomeric overhangs by telomerase could be a more efficient process and blocking telomerase access may delay clearance from the telomere capping checkpoint.  Conceivably, following GRN163L treatments, we should be able to visualize the delayed clearance of ATM/ATR-induced DDR signals by immunocytochemistry and confocal microscopy. We used antibodies against the DDR marker γH2AX (Ser139), which is primarily phosphorylated by ATM but also by ATR and DNA-PKcs and is used as a standard marker of uncapped or dysfunctional telomeres (150, 151, 154).    GRN163L treatment for 24 h resulted in increased formation of γH2AX DNA damage foci in a subset of MCF-7 and HT29 cells (Fig. 8 a-d) but not telomerase-negative primary human BJ fibroblasts (Fig. 8 e-f). The mismatched control oligomer (GRN-MM) had no significant effect on increasing foci accumulation when images were quantified for foci-positive cells. Secondary antibody-alone controls indicate that this increase in foci number is not due to increased non-specific background staining in GRN163L-treated ! 51!cells (Supplementary Fig. 4). Accumulations of these DDR foci, following treatments with GRN163L but not its mismatched control, are consistent with an increase in uncapped telomeres in the absence of telomerase activity.         ! 52!  Figure 8. Telomerase inhibition induces DNA damage foci in telomerase-positive but not telomerase-negative cells a-d) Telomerase-positive MCF-7 and HT29 cells treated for 24 h with 10 µM GRN163L display increased γH2AX DNA damage foci in a subset of cells when analyzed by immunocytochemistry. Treatment for 24 h with 10 µM GRN-MM control does not induce this effect. Etoposide treatment at 1.4 µM was used as a positive control. e-f) Treatment with GRN163L did not induce these DDR foci in telomerase-negative primary human fibroblasts (BJ). Arthur Chen performed quantification of foci-positive cells blinded for each treatment conditions and cell line. Histogram showed accumulation of cells with ≥10 foci, normalized to the numbers obtained from untreated (NT) controls. Student's t-test was used to generate P-values (* = P ≤0.05, ** = P ≤ 0.01, n ≥ 3).   ! 53!  In order to better define the population of DDR foci-positive cells and their cell cycle phase characteristics, we co-stained GRN163L-treated MCF-7 and HT29 cells with DDR markers and cell cycle phase markers. γH2AX foci-positive cells co-stain with cytoplasmic cyclin B1, a marker of late S/G2 phase (Fig. 9 a-d). γH2AX foci-positive cells do not co-stain with phospho-histone H3 (H3P) an M phase marker in MCF-7, indicating that the DDR foci-positive cells are likely in G2 phase (Fig. 9 e-f).    In support of the immunocytochemistry data, we observed an increase in the proportion of cells that stained positive for Cyclin B1 and had 4N DNA content, as measured by FACS (Fig. 9 g-h). Consistent with a delayed clearance from (rather than an arrest at) G2 phase, the magnitude of this effect is small (~5%) but is not observed with GRN-MM treatment. Overall, this suggests an accumulation of cells at the G2 checkpoint due to persistent DNA damage foci following GRN163L treatment. We conclude that GRN163L treatments correlate with transient accumulation of DDR marker γH2AX, coinciding with a delayed clearance of the telomere checkpoint at late G2 phase of the cell cycle.     ! 54! Figure 9. Cyclin B1/γH2AX ICC and FACS experiments indicate that GRN163L treatment increases the population of cells in late S/G2 phases a-f) MCF-7 and HT29 cells were treated for 24 h with GRN163L and stained for cytoplasmic cyclin B1, a marker of late S/G2, and phospho-histone H3 (H3P) a marker of M phase in combination with γH2AX. Foci-positive cells (≥10 foci) co-staining with cyclin B1 or H3P were quantified with >400 cells analyzed for each condition. Arthur Chen performed quantification of foci-positive cells blinded for each treatment conditions and cell line g-h) MCF-7 cells were treated with GRN163L for 24 h then stained with FITC-conjugated cyclin B1 and PI. ANOVA and Fisher’s LSD test were used to generate P-values (* = P ≤0.05, ** = P ≤ 0.01). ! 55!2.4.5. GRN163L-induced DDR foci formation depends on active ATM signaling    We next tested whether ATM signaling is required for the persistent DNA-damage foci induced by GRN163L. We again used our order-of-addition experimental scheme (Fig. 4 a) to treat MCF-7 cells with KU55933 and GRN163L for 48 h either alone or in different combinations, and quantify results of ICC experiments staining for γH2AX. The untreated (NT) negative control, 1.4 µM Etoposide (ET) positive control, and GRN163L alone treatment conditions behaved as in our previous experiments (Fig. 10 a-b). Also as expected, we observed that treatment with KU55933 alone (AA) or before GRN163L addition (AB) abolished γH2AX foci formation. Interestingly, treatment with KU55933 after 24 h incubation with GRN163L appeared to only reduce (although this effect is not statistically significant) but not eliminate γH2AX DDR foci. We reason that blocking the activity of ATM prevents the phosphorylation and formation of new γH2AX foci but does not remove the phosphorylated proteins that have already formed.  To provide further insight into the cell cycle dynamics present under the different treatment regimes, we stained cells from different treatment groups for phospho-histone H3P and quantified the proportion of cells progressing to M phase (Fig. 10 a, c). Under non-treated conditions we found that ~5% of MCF-7 cells stained positive for H3P. ! 56!Treatment with GRN163L alone decreases the proportion of cells in M phase, consistent with a stall at the G2 checkpoint due to persistent DDR signaling. Treatment with KU55933 treatment alone or prior to GRN163L addition further reduces the number of M-phase cells. This may be due to either faster passage through G2/M or a temporary arrest in G1 as suggested by our earlier FACS experiments (Fig. 8 a). In contrast, cells that were treated with GRN163L followed by KU55933 + GRN163L showed an increase in the proportion of cells in M phase. These data suggested that continuous ATM signaling is essential for the G2 stall following GRN-163L-induced DDR foci formation. Blocking ATM signaling following GRN163L treatment released MCF-7 cells from the G2 cell cycle checkpoint and allowed progression to the next phase of the cell cycle. This release of previously stalled cells manifests as an increase in cells entering M phase. Consistent with improperly capped telomeres proceeding to mitosis, we see an increase in micronuclei formation in the GRN163L 24 h/KU55933 + GRN163L 24 h conditions indicating mitotic defects and increased genomic instability (Fig. 11).      ! 57!a)  ! 58!b)        c)   Figure 10. Functional ATM signaling is required for GRN163L-induced DDR foci formation and G2/M checkpoint stall a) MCF-7 cells were treated with the indicated inhibitor in the same scheme as Fig. 4 for 48 h before being fixed and stained for γH2AX and H3P. b-c) Images were blinded and scored to determine co-staining quantification with >400 cells scored per condition. Arthur Chen performed quantification of foci/H3P-positive cells blinded for each treatment conditions and cell line. ANOVA and Fisher’s LSD test were used to generate P-values (* = P ≤0.05, ** = P ≤ 0.01, *** = P ≤ 0.001). Error bars represent SD. NT GG AA GB AB0. γH2AX Positive (n=3)Proportion of cells with ≥10 γH2AX fociNTGGAAGBAB*nsNT GG AA GB AB0. H3P Positive (n=3)Proportion H3P Positivens***** NTGGAAGBABns! 59!a)  b)  !!Figure 11. Increased mitotic defects following release of cells from G2 stall by ATM inhibition MCF-7 were treated with GRN163L for 48 h, GRN163L 24 h/GRN163L + KU55933 24 h, or vehicle control before fixation and DAPI nuclear staining. a) Representative images of treatments examined, arrows indicate micronuclei. b) Nuclear abnormalities were quantified on the same sets of images as in Figure 10. ANOVA and Fisher’s LSD test were used to generate P-values (* = P ≤0.05, ** = P ≤ 0.01, *** = P ≤ 0.001). Error bars represent SD.!NT GG GB0. Nuclear Abnormalities CountProportion of NucleiMicronucleatedMultinucleatedBudding***! 60!2.5. DISCUSSION   Telomeres differentiate the natural ends of chromosomes from random breaks through the formation of structurally distinct heterochromatin caps. During DNA replication, these telomere-loops are dismantled and the G-rich strand overhangs are synthesized by recruited telomerase or processed through nuclease actions (11, 12, 18, 83, 121). Finally, the loop structure is reformed via the coordinated actions of helicases and epigenetic modifiers (18, 81). ATM activation is concurrent with exposed chromosome termini and is crucial for the correct rebuilding of these chromosomal end-structures prior to the cell-division phase (151). Accordingly, the involvement of dysfunctional or defective helicases (BLM, RTEL1), nucleases (Apollo), or telomere factors (TRF1, TRF2) will result in telomere deprotection, cell cycle arrest, and the loss of cell viability (18, 121).    The role of telomerase in telomeric chromatin formation is inferred through its de novo telomere synthesis function but has never been directly demonstrated. Our data suggest that, in telomerase-positive cells, the actions of telomerase improve the kinetics of G-rich overhang formation, thus facilitating efficient higher-order telomeric chromatin reformation and clearance from the G2 checkpoint. In telomerase-positive cells, GRN163L blocks the access of the reverse transcriptase complex to its telomeric substrate, thereby preventing de novo synthesis of G-rich telomere repeats (129). Accordingly, ATM-! 61!mediated DDR signals at unstructured telomeres take longer to resolve, resulting in delayed passage through cell cycle checkpoints and an accumulation of cells in G2 (Fig. 11). The lack of telomerase action stalls, but does not entirely arrest cells at G2. This is because G-rich overhangs can be produced, albeit with reduced efficiency, through the actions of multiple nucleases (18, 121). Thus, the inhibition of telomerase will delay, but not stop, the progression of the cell cycle.    Improving the kinetics of telomere-cap formation with telomerase expression is consistent with previous work in our lab showing that overexpression of telomerase in telomerase-negative ALT cells conferred a growth advantage and faster passage through S/G2 phases of the cell cycle (155). This selective advantage may cause cancer cells to become “addicted” to telomerase activity. Notably, telomerase-negative cells are still able to reform telomere cap structures in the absence of telomerase due to the presence of C-strand-specific nucleases (156, 157). This is also supported by the lack of significant decreases in cell counts for GRN163L-treated telomerase-negative cell lines. Therefore somatic tissues with low telomerase expression should be unaffected, leading to a tolerable clinical toxicity profile (128, 129).    We previously observed that short-term treatment with GRN163L potentiated the effects of DNA-damaging agents in a non-telomere-length-dependent manner (148). Data ! 62!from our current study indicates that this effect is due to stalling via an ATM-dependent DNA damage signal at the telomere induced by telomerase inhibition. ATM inhibition prior to GRN163L addition reduces the DDR signal induced by GRN163L and its effects on cell cycle progression and potentiation possibly by preventing recruitment of telomerase to the telomere or via epistatic effects of KU55933 on cell cycle progression. Conversely, cytotoxicity is exacerbated with the addition of ATMi after 24 h of GRN163L treatment as the G2/M checkpoint has already been engaged. The increased etoposide cytotoxicity potentiation observed under these conditions may be due to the removal of ATM-signaling, allowing progression of previously stalled cells to progress to M phase where unrepaired chromosomal structure defects resulted in abnormal cell division and/or mitotic catastrophe. The higher levels of micronuclei formation observed following this treatment regime support this and are of indicative possible mitotic defects and chromosomal instability due to errors in telomere processing. The differences in total exposure time between the GRN163L/KU55933 24 h then GRN163L + KU55933 24 h treatments will be addressed with similar follow-up experiments using both inhibitors for 48 h.    The sustained DDR signal at unstructured telomere ends may act in an additive manner with DNA-damaging agents, pushing a greater proportion of cells over the apoptotic threshold. This order-of-addition effect may be explained by a previously-! 63!reported role for ATM in regulating recruitment of telomerase to telomeres (86). Inhibiting ATM before GRN163L treatment prevents engagement of the G2 checkpoint and may accelerate passage through G2 by reducing inhibitory phosphorylation of C-strand-specific EXO1 nuclease activity, leading to faster overhang formation and less topoisomerase exposure (158). This is consistent with the observed trend toward reduced populations of cells with 4N DNA content following KU55933 treatment. Alternatively, ATM activity may be necessary for efficient progression through S phase preventing KU55933-treated cells from entering G2. Topoisomerase inhibitors and other drugs that cause replicative stress may synergize particularly well with GRN163L; telomerase depletion in yeast has been observed to cause chronic replication stress and stalled replication forks (159). It is also possible that these changes in cell cycle distribution result in more cells in S/G2 (when topoisomerases are most active), thereby sensitizing them to topoisomerase inhibition (147). Additionally, proper T-loop formation may involve topoisomerase activity, as previously suggested (68). Off-target effects of GRN163L on adhesion (possibly through interactions between intermolecular G-quadruplexes and extracellular adhesion proteins) may also contribute to cellular stress signals and the differences we see in proliferation rates observed, though these are likely minor as telomerase-negative cell lines appear unaffected. ! 64!  The long lag time associated with multiple rounds of cellular replication, required for telomeres to shorten significantly, has been a major theoretical barrier to the utilization of telomerase inhibitors for anti-cancer chemotherapy (3). Recent clinical trials of GRN163L in myelofibrosis and thrombocythemia have cast doubt on this premise, as telomere length did not seem to change in response to therapy and baseline telomere length was not found to be predictive of positive therapeutic response (128, 133). In this context, our data provide an alternate explanation to the observed clinical effects of GRN163L: telomerase inhibition in these hematopoietic cell types may induce similar distortions in cell cycle kinetics without parallel observable effects on telomere-length regulation. Telomeric-DNA-replication stress could be partially relieved by an increase in the dNTP (purine) pool (159), which is a particular vulnerability in hematopoietic cancers as they frequently display dysregulated dNTP metabolism (160). This model predicts existing therapeutic agents that target the available dNTP pools, such as mycophenolic acids, will have synergistic effects with GRN163L treatments in vulnerable hematopoietic cell types. Our study broadens the conceptual scope of targeting telomerase beyond its telomere-length-maintenance activities in cancer to include combined therapies with DNA-damaging agents to take advantage of vulnerabilities presented by inhibiting its roles in higher-order telomere structure regulation. ! 65!  ! 66! Figure 12. Suggested model of effects of GRN163L on telomere cap formation In the presence of telomerase kinetics of telomere cap formation are optimal and involve a transient ATM-dependent DNA damage signal for telomerase recruitment (green circles = γH2AX). Telomerase extends the G-rich overhang facilitating T-loop formation. Treatment with GRN163L blocks the access of telomerase preventing overhang extension and slowing kinetics of telomere cap formation. Cell cycle stalling at G2 is caused by DNA damage responses at the unstructured telomere ends and telomere-specific replication stress. KU55933 treatment alone or KU55933 before GRN163L results in a delayed exit from S phase due to disrupted function of ATM during replication thereby abrogating the effect of GRN163L-induced G2 stalling. Treatment with GRN163L for 24 h before KU55933 causes a stall due to delayed telomere cap formation but as ATM is required to maintain the cell cycle checkpoint at the G2/M boundary it’s inhibition allows the release of stalled cells and procession to M phase. Since the telomere cap structures are not properly formed in this scenario there is an increased incidence of mitotic defects and cell death. ! !!!!!!!!!!!!!!!! 67!CHAPTER 3 !A version of this material has previously been published as: Fleisig HB, Hukezalie KR, Thompson CA, Au-Yeung TT, Ludlow AT, Zhao CR, Wong JM. 2016. Telomerase reverse transcriptase expression protects transformed human cells against DNA-damaging agents, and increases tolerance to chromosomal instability. Oncogene 35(2):218-27.   Kyle Hukezalie provided the data for the viability and FACS experiments outlined (Fig. 14). Kyle Hukezalie and Judy Wong provided the data for the chromosome-healing assay. Helen Fleisig prepared the metaphase spreads (Fig. 16) and I imaged and quantified them. I developed the nuclear abnormality-scoring assay and performed the experiments covered here (Fig. 15). Judy Wong and Helen Fleisig conceived the experiments and Judy Wong and Kyle Hukezalie wrote the manuscript for the published paper.  3.1. INTRODUCTION 3.1.1. Non-canonical roles of telomerase in cancer   In addition to its telomere length maintenance function telomerase has also been implicated in other roles. These include regulation of non-telomeric DNA damage responses (69), promotion of cell growth and proliferation (161), acceleration of cell cycle kinetics (155), and influences on mitochondrial DNA (mtDNA) damage and mitochondrial integrity following oxidative stress (72). These extra-telomeric functions of telomerase may be advantageous in the context of increased proliferation and metabolic stress such as that found in rapidly-dividing cancer cells. TERT has also been reported to act as a ! 68!transcriptional modulator of the Wnt/β-catenin signaling pathways, though this remains controversial (162, 163). Additionally, when complexed with the RNA component of the mitochondrial endoribonuclease MRP, TERT displays properties of an RNA-dependent RNA polymerase, suggesting it may play a role in regulating gene expression (164). Nuclear TERT has been shown to protect against apoptosis independent of telomere length (165).    The delineation between telomere-dependent and telomere-independent effects of telomerase can be difficult to distinguish, since effects can be co-dependent.  The possibility of cellular impact through contributions from both telomere-related and telomere-independent roles is high, especially if the read-out is measuring simple growth advantages. Additionally, changes in telomerase activity, whether by recombinant protein expression or genetic/chemical inhibition, are expected to impact telomere synthesis. As subtle changes in telomere-structure maintenance could indirectly affect DNA-damage responses, it is also possible that telomerase expression promotes DNA repair through its telomere-synthesis activity.   !! 69!3.1.2. Role of telomerase at the mitochondria  Recent studies have shown that TERT is transported to the mitochondria under conditions of oxidative stress. There it appears to have a protective effect on mtDNA and positively impact mitochondrial function (71, 72, 166) Genetic extinction of telomerase in murine cancer cells results in downregulation of peroxisome proliferator-activated receptor gamma coactivator 1- β (PGC-1β) ) (111), the master regulator of mitochondrial biogenesis, function, and oxidative stress resistance, and its targets. As well, telomerase gene extinction increases the sensitivity of murine cancer cells to the knockdown of the major mitochondrial ROS antioxidant superoxide dismutase 2 (SOD2) (111). Similarly, telomerase-null mice exhibit mitochondrial defects, impaired gluconeogenesis, cardiomyopathy, and increased ROS production in the setting of telomere dysfunction (167). This has led to a model in which short telomeres cause activation of p53, which binds to and represses PGC-1α and PGC-1 β promoters, directly linking telomere integrity to mitochondrial function (168). An important caveat to this model is that these irregularities also begin to manifest in the first generation of mice before telomeres have had time to significantly shorten, suggesting that telomere length may not be the sole determinant of mitochondrial dysfunction. It is unclear whether telomerase’s effects on ! 70!mitochondrial integrity and function are primarily mediated by its activities in the nucleus or at the mitochondria.     !    Figure 13. Canonical and non-canonical roles of telomerase reverse transcriptase Telomerase activities at the telomere can be conceptually classified as canonical activities while non-telomere length maintenance functions can be considered non-canonical. Non-canonical activities may be beneficial to cancer cells via increasing stress resistance capabilities and invasive/metastatic growth potential as well as through possible other functions.     ! 71!3.1.3. TERT domains and non-canonical functions   There is conflicting evidence with regard to the necessity of telomerase’s catalytic activity for its non-canonical functions (166, 169). In mouse neurons treated with the excitotoxin N-methyl-D-aspartic acid (NMDA), TERT overexpression was shown to be beneficial for protection of mitochondrial function in the absence of detectable TER or telomerase activity (170). Similarly, removal of TERT but not TER reduces survival and stress resistance in mouse embryonic fibroblasts and neurons exposed to NMDA or the ATP-competitive kinase inhibitor staurosporine (171). Conditional induction of TERT, without the requirement of TER has been observed to increase proliferation of hair follicle stem cells in mice (172). It is possible that telomerase activity may be required for some but not all non-canonical effects observed.  3.1.4. Strategies for studying non-canonical functions of telomerase    As previously mentioned there is an alternative mechanism for telomere length maintenance termed alternative lengthening of telomeres (ALT) that does not require telomerase. We can take advantage of this feature and use it as a system to study TERT’s putative non-canonical functions by expressing telomerase and its functional variants in ALT cells without interfering with ALT telomere maintenance capacity through the ! 72!recombination-based mechanism (173, 174). Recombinant expression of telomerase in transformed human cells that utilize the ALT mechanism for telomere maintenance confers growth advantages to these cells (155). Retroviral-vector-directed telomerase expression in human ALT cells promotes proliferation in normal culture and in anchorage-independent conditions, a common in vitro test for cellular transformation. Telomerase expression promotes progression through G2/M phases of the cell cycle, resulting in the accelerated growth of telomerase-positive ALT cells. This ALT-cell growth promotion phenotype requires the fully active telomerase holoenzyme, as the expression of dominant-negative TERT or other partial loss-of-function TERT mutants, such as the dissociation-of-telomerase-activities mutants, does not lead to the same accelerated growth phenotype. Stable expression of TERT (TERT/TER in the cases of VA13 and the osteosarcoma U2OS ALT model) reconstitutes telomerase activity without the abrogation of molecular characteristics of ALT (175, 176) The same ALT cell models expressing recombinant telomerase components were used here to assess the tolerance of TERT-positive ALT cells to treatment with DNA-damaging agents and to compare their sensitivities to those of TERT-negative isogenic cell models.    ! 73!Note: 3.1.5-3.1.7. is a condensed summary of work appearing in (175).   3.1.5. Recombinant telomerase expression in ALT cells confers protection against cytotoxicity from DNA damage and increases kinetics of ds-break repair    TERT+/- ALT cells were transiently treated with a panel of DNA-damaging agents and monitored for growth using the colony-forming-unit (CFU) assay over a two-week period. Dose-response experiments with VA13 and GM00847 isogenic cell models showed a clear telomerase-dependent protective effect against the cytotoxicity of all DNA-damaging agents tested (Fig. 13). Constitutive expression of telomerase in ALT cells conferred survival advantages against dsDNA breaks. In contrast with our prior data, with small-molecule-directed transient inhibition of telomerase in breast/colorectal cancer cells, this protection was not restricted by the cell-cycle-specificity/mechanism of the chemotherapeutic agents tested. Telomerase+ ALT also showed improvement in cell viability immediately after exposure to dsDNA-damaging agents, as measured by the acute WST-1 metabolic assay where we also observed increased short-term survival.  ! 74!    Figure 14. Telomerase expression confers survival advantages against double-strand DNA-damaging agents in ALT cells Cytotoxicity dose-response experiments against double-strand DNA-damaging agents were performed with CFU assay. a) Recombinant expression of TERT and TER in VA13 ALT cells confers survival advantages against dsDNA-damaging agent etoposide and b) irinotecan. c) TERT expression in GM00847 cells confers protection against cell cycle non-specific agent oxaliplatin. Error bars represent standard deviations, which may not be visible on the dose-response curves at higher drug concentrations due to the near complete loss of viability. Data are representative of at least three independent experiments. ***P<0.01 Unpaired Student’s T-test. Performed by Kyle Hukezalie. ! 75!  To investigate the mechanism behind the protection against dsDNA-damaging agents, the kinetics of dsDNA repair in TERT+/- VA13 cells were examined following transient exposure to bleocin using the chromosome-healing assay. Paired comparison of residual dsDNA-break signals between telomerase-positive and vector control VA13 cells showed significant differences in the removal of fragmented chromosomes at each of the measured time points (175). In contrast to the efficient removal of fragmented chromosomes in TERT+VA13 cells, at the conclusion of the timed collection experiment (T=48 h), substantial amounts of dsDNA breaks persisted in VA13 vector control cells, correlating to decreased cellular viability.   3.1.6. TERT’s telomere-repeat-synthesis activity is dispensable to DNA-damage tolerance    To discern whether telomere-repeat-synthesis activity is necessary for the observed increase in tolerance to genotoxic insults, we expressed catalytically-defective telomerase variants in VA13 ALT cells and subjected these cell models to the same cytotoxicity measurements (CFU proliferation and WST-1 viability assays). Expression of a TER-binding defective variant, RBD1-TER (155, 177), protected against genotoxic stimuli in VA13, GM00847, and U2OS ALT cells at a level similar to WT-TERT. We ! 76!repeated the RBD1 study in the absence of co-expressed recombinant TER to confirm that TER is not involved in the DNA damage tolerance with similar results. The most common alternatively spliced TERT product, β-deletion-TERT (β-del-TERT), is found in a high ratio to full-length transcripts in human embryonic stem cells, and cancer models where it lacks telomerase activity (57,58). In agreement with a recent study (178), expression of β-del-TERT with TER increased EC50 values for bleocin compared to cells expressing empty vector. Together, our cytotoxicity measurements in ALT cells expressing RBD1- and β-del-TERT support the existence of TERT functions that are not related to telomere synthesis, but are important to the chemotherapy-resistant properties of transformed cells.  3.1.7. Cells with ~8N DNA content were detected in TERT-positive, but not vector control VA13 ALT cells, after induction of DNA damage   Next, we decided to examine the chromosomal content of VA13 vector and WT-TERT positive cells following treatment with irinotecan or the DNA-polymerase inhibitor aphidicolin. We observed the accumulation of an extra population of cells with DNA content of approximately 8N in WT-TERT-positive and RBD1-TERT VA13 cells (both possessing the DNA-damage survival phenotype), but not in VA13 vector control cells. ! 77!The “new” aneuploid state was maintained for several generations after the genotoxic event when sorted from the mass culture by flow cytometry and allowed to recover under normal culturing conditions.    We reasoned that the induction of DNA damage prevents the successful completion of the first cycle of cell division. In the absence of TERT, VA13 ALT cells whose cell division was suspended due to mitotic catastrophe were eliminated by apoptosis or other cytotoxic means. In contrast, TERT expression increased dsDNA-damage repair, albeit with an error-prone mechanism, and allowed the bypass of programmed cell death. In this case, cell division was still suspended, resulting in the accumulation of replicated genetic material and DNA content in these cells approaching 8N. Subsequently, the TERT-positive VA13 cells recovered from suspension in cell division and resumed cycling with stochastic chromosome division in the presence of imperfect DNA repair. This was followed by several generations of cell division in the presence of continuous chromosome-number instability until a new median was reached.        ! 78!3.2. HYPOTHESIS Overexpression of TERT in ALT cells increases survival following DNA damage and promotes tolerance against genomic instability. This TERT function is distinct from its role in telomere length maintenance.   3.2.1. Specific aims  (1) Quantify the non-canonical effects of TERT on cell division and morphology, following treatment with DNA-damaging agents in ALT cells (2) Quantify the non-canonical effects of TERT on tolerance against chromosomal instability in ALT cells  3.3. METHODS  !Cell culture and reagents The construction of ALT cell models expressing recombinant TERT and TER has been described previously (155). The catalytically-defective TERT mutants include DN (TERT DV710-711AI), RBD1 (TERT FY561-562AA), RBD3 (NAAIRS substitution of TERTaa386-391), and RBD5 (NAAIRS substitution of TERTaa512-517). Cells were grown ! 79!in DMEM, supplemented with 5% fetal bovine serum and penicillin/streptomycin at 37°C under 5% CO2 as described above (Ch. 2.3.). Double-strand DNA damaging chemicals were obtained through Fisher BioReagents (aphidicolin), Tocris Bioscience (etoposide, oxaliplatin), EMD Millipore (bleocin), Sigma-Aldrich (irinotecan), and Calbiochem (cisplatin).  Construction of TERT retroviral vectors   Cloning of the retroviral vectors coding for TERT mutants was described previously (155). Beta-deletion TERT was obtained from RT-PCR and subcloned into pBabehygro vector. All retroviral vectors were sequenced to confirm cloning fidelity.  Flow-cytometry based DNA content analysis and live-cell fluorescence-assisted cell sorting   Following the induction of DNA damage, cells were harvested at designated time points and analyzed for DNA content with propidium iodide staining followed by flow cytometry, as previously described (155) (Ch. 2.3.). Labeled cells were sorted using the BD Calibur flow cytometer (UBC Biomedical Research Center). FACS data was analyzed with FlowJo software (FlowJo LLC, Ashland, OR, USA). ! 80!  FACS recovery of ALT cells based on DNA content was performed following a 10 h recovery period after a 24h DNA-damage induction treatment. Following cell harvest by trypsinization and Hoechst 33342 labeling, ALT cells with an apparent 8N DNA content were collected by FACS. Karyotype of the 8N populations of ALT cells were examined after recovery as an adherent culture for at least 6 PDs.  Nuclear Abnormalities: Staining and Quantifications   Cells were seeded onto glass cover slips, treated with genotoxin for 24 h, and then allowed to recover for 9 h in the absence of drugs. Harvested cells were fixed as described  above (Ch. 2.3.), stained with DAPI (1:125,000), then mounted with Fluoromount-G (Southern Biotech, Birmingham, AL). Images were collected using a Zeiss LSM 700 confocal laser scanning microscope and ZEN software (Carl Zeiss Microimaging, Gottingen, Germany). Collected images were blinded using R (R Foundation for Statistical Computing, Vienna, Austria), and scored based on criteria (179) with >400 nuclei scored for each treatment condition.     ! 81!Metaphase spreads and telomere fluorescent in situ hybridization    Metaphase spread preparation and PNA probe hybridization were performed as described (180). Confocal images were collected as described above (Ch. 2.3.).  3.4. RESULTS  3.4.1. Cytological abnormalities in VA13 cells following the induction of double-strand DNA damage are partially rescued with recombinant TERT expression   Previous characterization of ALT cells revealed a higher sensitivity to DNA damage, a reduction in IR-induced DNA repair capacity, and a defective G2/M checkpoint (181). Together, these characteristics may reflect the compensatory changes in the DNA-damage surveillance threshold, particularly in G2 of the cell cycle when homologous recombination dominates, to accommodate the recombination-based mechanism of telomere-length maintenance. In corroboration of this theory, our cytological analysis of VA13 ALT cells revealed high levels of nuclear morphological abnormalities that were exacerbated by the chemical treatments with aphidicolin and irinotecan (Fig. 15).  ! 82!  We counted the percentage of abnormal nuclei according to published guidelines (179). Under log-phase growth conditions, the proportion of VA13 vector cells exhibiting abnormal nuclear morphology exceeded 30% and this proportion was similarly observed in VA13 cells stably expressing WT- and RBD1-TERT. Treatments with aphidicolin and irinotecan induced a robust increase in VA13 cells with abnormal nuclear morphologies, with cells exhibiting abnormal nuclei phenotype exceeding 50% of the counted cell population, in the case of irinotecan treatment. These abnormalities were partially rescued with the recombinant expression of WT- as well as RBD1-TERT (P<0.05) (Figure 15 b-c). This nuclear morphology protection was observed in VA13-TERT cells treated with both aphidicolin and irinotecan, which may be related to the overall improvement of cell survival following DDR induction.  ! 83!  Figure 15. ALT cells exhibit high levels of nuclear morphological abnormalities following the induction of DNA damage; these abnormalities are partially rescued with telomerase expression a) We observed a high level of cytogenetic abnormalities in resting VA13 ALT cells, b) with increasing amounts following exposure to DNA-damaging agents irinotecan, c) and aphidicolin. Recombinant expression of telomerase in VA13 ALT cells did not reduce cytogenetic abnormalities in resting cell populations, but reduced the proportion of cells with abnormal nuclei following exposure to the two DNA-damaging chemicals. Data are representative of at least three independent experiments. *P<0.1 **P<0.05 ****P<0.01 Two-Way ANOVA and Tukey’s test.  ! 84!3.4.2. Karyotype analysis of TERT-positive ALT cells with 8N DNA suggests that bypass of mitosis results in gross genomic instability    We performed metaphase analysis to characterize the drug-exposed 8N-DNA TERT-positive VA13 cells isolated by fluorescence-assisted cell sorting. Following sorting, cells were allowed to recover in normal growth medium and metaphase spreads were prepared after 6-8 population doublings. In agreement with the PI staining pattern, we observed an increase in chromosome numbers compared to both the parental TERT-positive VA13 cells and the VA13 ALT vector cells.    We found that the 8N-DNA TERT-positive VA13 population exhibited the widest range of chromosome numbers (Fig. 16). The median chromosome number from this cell line, however, was less than double the chromosome number of the parental cell line (TERT-positive VA13), indicating that a stochastic loss of chromosomes from the approximate 8N content had occurred. While it is possible that the counting of the complete set of chromosomes in the 8N-DNA TERT-positive VA13 cell lines has low accuracy, we contend that the change in chromosome numbers was a function of genomic instability following the telomerase-expressing ALT cells’ exposure to genotoxic agents. ! 85!        Figure 16. Cytogenetic analysis of chromosome numbers with metaphase spread Telomeres were labeled with a peptide-nucleic-acid probe against the telomeric DNA repeat sequence, conjugated to a fluorophore (FiTC). DNA is counterstained with DAPI. Quantification of median chromosome number suggests the sorted 8N population may be unstable, and a mixed population of survived cells, with new median chromosome numbers, exists in the same culture following timed selection. Metaphase spread prepared by Helen Fleisig and Judy Wong, microscopy and chromosome quantification done by Connor Thompson.  ! 86!3.5. DISCUSSION    Over 85% of human cancers prefer telomerase reactivation to the ALT mechanism of recombination-dependent telomere synthesis. We contend that this “telomerase addiction” could partly stem from advantages provided by TERT’s non-telomere related activity, which facilitate cell growth and proliferation after genotoxic stress. Our data show that constitutive TERT expression provides oncogenic transformed cells with survival advantages following chemically induced double-strand DNA damage, in a manner functionally distinct from its role in telomere synthesis and structure maintenance.    Many ALT tumors are highly sensitivity to DNA damage, have slow DNA repair kinetics, and are defective in the G2/M checkpoint (181). Our data demonstrated that chemotherapy-induced dsDNA breaks are repaired with improved kinetics by the recombinant expression of TERT, leading to an improvement in cell survival following genotoxin exposure. In agreement, our cytologic and cytogenetic analysis of VA13 vector control cells revealed a high degree of abnormality; a large percentage of cells exhibiting abnormal nuclear morphologies in the form of micronuclei and multi-nucleation. The high percentage of ALT cells exhibiting nuclear abnormalities confirmed published reports of the silencing of the G2/M checkpoint, leading to mitotic catastrophe and abnormal nuclear divisions. Accumulation of abnormal nuclear morphologies is exacerbated by chemical ! 87!treatments with DNA-damaging agents, a function of the reported deficiency in G2-phase-associated DNA damage response in ALT tumors.    Recombinant WT-TERT expression promotes dsDNA repair in ALT cells and improves survival against genotoxic stimuli. However, improved survival against genotoxic stress in telomerase-positive ALT cells does not equate to proper cell division. In the absence of correct DNA repair mechanisms, error-prone repair of the ALT genome leads to a temporary suspension of cell division, avoiding mitotic catastrophe but resulting in the accumulation of cells with double the genomic content (8N). TERT-positive ALT cells with 8N DNA content were able to re-enter the cell cycle after a period of recovery in culture. Thus, TERT-expression promotes tolerance of gross genomic instability and the resultant aneuploidy. Our data indicate that TERT expression promotes oncogenic growth of transformed cells in multiple ways: TERT expression is associated with increased tolerance to the cytotoxic effects of DNA-damaging events, and resolution of the damaged DNA in surviving cells promotes genomic instability, which further drives the process of aneuploidy.       ! 88!CHAPTER 4 !4.1. CONCLUSIONS AND FUTURE DIRECTIONS !4.1.1. Brief summary ! Telomerase activity is present in the overwhelming majority of cancer cells where it maintains telomere length and allows for unlimited replicative capacity. Telomerase inhibition by GRN163L (Imetelstat) has previously been demonstrated to have a potentiating effect on S/G2 cell-cycle-specific DNA-damaging agents. Our work shows that GRN163L treatment causes a transient DNA-damage response signal in telomerase-positive cells that leads to a greater accumulation of cells in G2 phase, which may explain this selective potentiation. We observed that the potentiating effect of GRN163L depends upon an active ATM-mediated DDR signal. In other work, we also showed that high levels of exogenous telomerase expression in transformed human cells increases tolerance to DNA-damaging agents and chromosomal instability. Our data support further investigation of therapies targeting (either directly or indirectly) other functions of telomerase besides telomere length maintenance for anti-cancer chemotherapy.   ! 89!4.1.2. Telomerase inhibition and potentiation of DNA-damaging agents   Previous work from our lab showed that short-term treatment with GRN163L potentiated the cytotoxic effect of some but not all DNA-damaging drugs tested (148). This effect was only observed with the topoisomerase inhibitors etoposide and irinotecan that act primarily during S/G2 phases of the cells cycle but not with the non-cell-cycle-specific crosslinking agent oxaliplatin or direct DNA-break-inducer bleomycin. In the work outlined in Chapter 2 of this manuscript we aimed to better understand why inhibition by GRN163L seems to have a cell-cycle-specific mode of action for preferential potentiation of DNA-damaging agents.    We observed that treatments with GRN163L sensitized telomerase-positive cells to cell-cycle-specific DNA-damaging agents through delayed resolution of an ATM-dependent DNA-damage signal. Treatment of telomerase-positive but not telomerase-negative cell lines with GRN163L caused an increase in cell population with 4N DNA content as measured by FACS. Telomerase inhibition alone induces γH2AX DNA-damage foci in a subset of telomerase-positive cells. We reason that this may be due to the loss of telomerase actions in optimal T-loop formation. Additional FACS and immunocytochemistry experiments show that the 4N DNA content and foci-positive cells co-stain for the G2/M phase marker Cyclin B1 but not for the M-phase marker phospho-! 90!histone H3. Treatment-related cytotoxicity was minimally observed, suggesting that GRN163L-treated cells were reversibly stalled but not arrested at G2.    This G2 stall may increase the cytotoxicity of DNA-damaging agents that work preferentially during this phase of the cell cycle via an additive effect. DNA damage signaling caused by GRN163L treatments could push cells that would otherwise survive the topoisomerase inhibition to apoptosis. Alternatively, GRN163L treatment may act synergistically with topoisomerase inhibitors to create telomere dysfunctional stress. Topoisomerases have been suggested to play a role in telomere structure resolution following DNA replication (68). In the absence of C-rich overhang elongation and other possible structural facilitations by telomerase, it is possible that topoisomerase activity becomes more important for proper telomere processing. The loss of both telomerase and topoisomerase actions at the time of telomere replication may therefore be acting in a synergistic manner to create unfavorable survival conditions.     The data in this study represent a separate mechanism by which telomerase inhibition could affect telomere-maintenance kinetics and homeostasis. As multiple cellular replication cycles are expected in order for telomeres to become critically short following telomerase inhibition, the telomere-attrition paradigm predicts that any short-term telomerase inhibition effect is unexpected. Our data contradict this paradigm by ! 91!demonstrating that transient (24-48 h) exposure to GRN163L is enough to alter the progression of the cell cycle and potentiate the effects of DNA-damaging chemicals.  Our proposal of telomere-length-independent effects from telomerase inhibition is also supported by recent clinical trials for hematopoietic proliferative disorders myelofibrosis and thrombocythemia, which showed that telomere length does not appear to determine clinical response (128, 133). Additionally, our previous work testing the combination effects of telomerase inhibition in human breast and colon cancer cell lines with topoisomerase inhibitors (148) and a recent study combining telomerase inhibition with the purine analog fludarabine in primary human leukemic leukocytes (182) show that short-term GRN163L treatment can have a potentiating effect on cell-cycle-specific DNA-damaging drugs. Our data is relevant to understanding telomerase’s roles in telomere structural maintenance and cell cycle progression, thereby presenting new testable hypotheses and possibilities for combination drug regimens.    These ideas are conceptually distinct from our work in Chapter 3 of this thesis implicating telomerase in roles other than its telomere-length maintenance function. Reported non-canonical functions of telomerase include regulation of non-telomeric DNA-damage responses (175), increasing apoptosis resistance (178), promotion of cell growth and proliferation-regulating genes (161), and managing mitochondrial DNA (mtDNA) ! 92!damage and mitochondrial integrity following oxidative stress (183). These effects do not require functional telomerase activity and may act outside of the canonical holoenzyme complex (175, 184). !4.1.3. ATM signaling and telomerase inhibition    Additive cytotoxic effects of KU55933 were observed when combined with GRN163L telomerase inhibition and the cell cycle specific DNA-damaging agent etoposide in MCF-7. When we performed order-of-addition experiments ATMi was found to block the GRN163L-mediated formation of DDR foci and cell cycle profile changes that we saw with GRN163L treatment alone. Interestingly, additive increased cytotoxicity and GRN163L mediated cell cycle profile alterations depended upon the order of treatment addition. When KU55933 was added before but not after GRN163L we lost the effects of GRN163L on cell cycle kinetics and potentiation. This suggests that the effects of GRN163L on potentiation of cytotoxicity and cell cycle alteration depend upon functional ATM signaling. Continuous activity of ATM may be required to hold the “stalled” cells at G2 as indicated by our immunocytochemistry experiments using the M phase marker phospho-histone H3, which displayed a large increase when KU55933 treatment followed incubation with GRN163L. This increase could represent cells being released from G2, where in the ! 93!context of concurrent etoposide treatment causes cells with damaged DNA to undergo mitotic catastrophe (Fig. 10). These cells die at a higher rate than cells where cell cycle checkpoints are intact and DNA-damage-repair systems have a chance to resolve lesions that might otherwise result in aberrant chromosome segregation and cell death.   ATM is known to be an important mediator of telomerase recruitment to telomeres (86, 152). An ATM-dependent DDR response signal is present at normal telomeres during S-phase directly before telomere elongation by telomerase (150). A full DDR response leading to senescence or apoptosis is not engaged here as several parallel mechanisms including epigenetic regulation and inhibition by shelterin coordinate prevent propagation of this signal (18, 185). Disruption of ATM as found in ataxia telangiectasia fibroblasts results in rapid loss of telomere length (186). Chemical inhibition of ATM by KU55933 may repress telomerase recruitment, this explains the lack of additive cytotoxicity observed when KU55933 treatment is followed by GRN163L treatment.   The presence of telomerase may improve the kinetics of T-loop formation and thereby reduce the duration of DDR signaling at unstructured chromosome ends, allowing for faster clearance of the G2/M checkpoint. The ssDNA G-rich overhang that forms the T-loop can be generated by telomerase repeat synthesis or the less-efficient action of various nucleases on the C-rich strand (18, 121) GRN163L blocks access of the ! 94!telomerase holoenzyme complex to the telomeric substrate preventing synthesis of the G-rich strand and so causing increased reliance on the slower mechanism of overhang generation (129). Telomerase expression improving the kinetics of telomere-cap formation is consistent with previous work in our lab showing that overexpression of telomerase in telomerase-negative ALT cells confers a growth advantage and faster passage through S/G2 phases of the cell cycle (155). Additionally, as suggested by our FACS data showing a trend toward reduced populations of cells with 4N DNA content following KU55933 treatment, inhibiting ATM activity may stall cells in G1 phase or expedite passage through G2 by reducing inhibitory phosphorylation of C-strand-specific EXO1 nuclease activity (158)  4.1.4. Non-canonical protective roles of TERT against DNA damage  In a separate study described in Chapter 3 of this thesis, we demonstrated that overexpression of TERT in ALT cells results in increased tolerance to DNA-damaging agents and chromosomal instability. Interestingly, this effect did not depend on telomerase functionality for repeat addition as demonstrated by the experiments using of RBD1 TERT with and without TER (175). This indicated that the protective activity of TERT is non-telomere catalysis related and is separate from the telomere-length-maintenance role of ! 95!TERT at telomeres. We showed that TERT-positive ALT cells surviving treatments with DNA-damaging agents had increased levels of nuclear abnormalities as observed by confocal microscopy indicating defects in replication. Treatments with aphidicolin or irinotecan induced an increase in TERT-positive ALT cells with duplicated genomic content (8N). FACS-selected 8N-DNA cells experienced a few cycles of genomic evolution as evidenced by a variable PI staining pattern. Increased chromosomal content (but less than 8N content) was verified after 6-8 population doublings via metaphase spreads and manual counting. Our data demonstrated that TERT expression aids cancer progression by promoting resistance to DNA damage induced through replication stress or by chemotherapy. Additionally, not only does TERT protect cells from genotoxic stress, but those cells that do survive have better tolerance against chromosomal instability, fueling tumor evolution.   This data contrasts with results of our other study, showing protective effects of telomerase expression on various anti-cancer treatments, including both cell-cycle-specific (etoposide, irinotecan) and non-specific agents (bleocin, oxaliplatin, cisplatin). We contend that increased general stress tolerance with stable recombinant telomerase expression is directly related to telomere-independent non-canonical telomerase activities. The use of the telomerase-specific inhibitor GRN163L, a PNA-oligonucleotide inhibitor that binds to ! 96!TER with high affinity, may have hindered our ability to detect this non-canonical activity. Given that the mechanism of TERT function responsible for the general increase in cell survival does not rely on telomere-telomerase interactions, it is thus not surprising that telomerase inhibition by GRN163L, which competitively displaces telomere-telomerase binding, might not reveal its non-canonical mode of action.   4.1.5. Future directions for telomerase inhibition in human anti-cancer chemotherapy  Our work confirms that transient telomerase inhibition increases the cytotoxicity of DNA-damaging agents in a cell cycle phase-specific manner (148) (Chapter. 2). Clinically, this observation could translate to testing telomerase inhibition with DNA-damaging agents that work primarily during S/G2 phases of the cell cycle. The potentiation effect of GRN163L does not depend upon significant telomere shortening based on the short time courses of experiments being insufficient for telomere attrition to occur. This is an advantage for potential therapeutic applications. The DDR induced by GRN163L treatment and treatment with other DNA-damaging drugs may work in an additive fashion to push cells past an apoptotic threshold, resulting in increased cytotoxicity. As telomerase depletion in yeast has been observed to cause chronic replication stress and stalled ! 97!replication forks (159), combining GRN163L with topoisomerase inhibitors and other drugs that cause persistent replicative stress may synergize particularly well.  In terms of pharmacodynamics, our study of telomere maintenance by telomerase, DDR signaling, and the effects on cell cycle progression are intrinsic tumor properties, which are unlikely to be directly influenced by the microenvironment. Accordingly, we believe 2-D culture models are representative in this case. However, we are cognizant that the pharmacokinetic properties of drug treatments are dependent on microenvironment and organismal parameters, which 2-D models cannot begin to approximate. These properties are beyond the scope of our current studies, but will be (or already have been) addressed in pre-clinical studies of the respective inhibitors we used in our experiments. The impact of pharmacokinetic characteristics of GRN163L on cell cycle progression of solid tumors requires further investigation.   Current clinical development of telomerase inhibition is focused primarily on TERT’s telomere synthesis activity and has found limited therapeutic success. Our study suggests that this strategy likely limits the scope of the effect that telomerase inhibitor treatment has on tumor growth. Targeting non-canonical TERT activities, particularly with co-administration of DNA-damaging agents, may provide new therapeutic interventions for telomerase-positive cancers. Drugs predicted above to synergize with GRN163L have not ! 98!been tested, as most clinical trials so far using multiple drugs have been small phase I trials focused primarily on safety and minimal toxic dose determination. Paclitaxel (microtubule stabilizer, M phase specific), bevacizumab (VEGF mAb), bortezomib (proteasome inhibitor), paclitaxel/carboplatin (non-cell-cycle-specific DNA crosslinker), and trastuzumab (HER2 mAb) have been combined in trials with GRN163L with no short-term potentiation observed, though some trials were stopped prematurely due to hematological toxicity (114). The recent trials in essential thrombocythemia and myelofibrosis take advantage of these hematopoietic effects and have had promising results that do not appear to be dependent upon patient telomere length (128, 133).   The results of our study overexpressing TERT in ALT cells also has implications for anti-telomerase therapies. Since the protective effects of TERT expression against DNA-damaging agents did not depend upon functional telomerase catalytic activity, approaches designed to block telomere repeat synthesis such as GRN163L may be missing important aspects of TERT’s activity in cancer cells. Future studies in our lab will map the regions of TERT required for its protective functions and isolate other interacting partners of TERT giving us new information for more efficiently designing therapeutic interventions. ! 99!REFERENCES  1. Shay JW, Wright WE. Role of telomeres and telomerase in cancer. Semin. Cancer Biol. 2011 Dec;21(6):349–53.  2. Wong JMY, Collins K. Telomere maintenance and disease. Lancet. 2003 Sep 20;362(9388):983–8.  3. Shay JW. Role of Telomeres and Telomerase in Aging and Cancer. Cancer Discov. 2016 Jun;6(6):584–93.  4. McClintock B. The Stability of Broken Ends of Chromosomes in Zea Mays. Genetics. 1941 Mar;26(2):234–82.  5. Blackburn EH, Gall JG. A tandemly repeated sequence at the termini of the extrachromosomal ribosomal RNA genes in Tetrahymena. J. Mol. Biol. 1978 Mar 25;120(1):33–53.  6. Moyzis RK, Buckingham JM, Cram LS, Dani M, Deaven LL, Jones MD, et al. A highly conserved repetitive DNA sequence, (TTAGGG)n, present at the telomeres of human chromosomes. Proc. Natl. Acad. Sci. U.S.A. 1988 Sep;85(18):6622–6.  7. Meyne J, Ratliff RL, Moyzis RK. Conservation of the human telomere sequence (TTAGGG)n among vertebrates. Proc. Natl. Acad. Sci. U.S.A. 1989 Sep;86(18):7049–53.  8. Allsopp RC, Harley CB. Evidence for a critical telomere length in senescent human fibroblasts. Exp. Cell Res. 1995 Jul;219(1):130–6.  9. Hukezalie KR, Wong JM. Structure-Function Relationship and Biogenesis Regulation of the Human Telomerase Holoenzyme. FEBS J. 2013 Apr 3.  10. de Lange T. Shelterin: the protein complex that shapes and safeguards human telomeres. Genes Dev. 2005 Sep 15;19(18):2100–10.  11. Zhong FL, Batista LFZ, Freund A, Pech MF, Venteicher AS, Artandi SE. TPP1 OB-fold domain controls telomere maintenance by recruiting telomerase to chromosome ends. Cell. 2012 Aug 3;150(3):481–94.  ! 100!12. Nandakumar J, Cech TR. Finding the end: recruitment of telomerase to telomeres. Nat. Rev. Mol. Cell Biol. 2013 Feb;14(2):69–82.  13. Schmutz I, de Lange T. Shelterin. Curr. Biol. 2016 May 23;26(10):R397–9.  14. Pandita RK, Chow TT, Udayakumar D, Bain AL, Cubeddu L, Hunt CR, et al. Single-strand DNA-binding protein SSB1 facilitates TERT recruitment to telomeres and maintains telomere G-overhangs. Cancer Res. 2015 Mar 1;75(5):858–69.  15. Blackburn EH, Epel ES, Lin J. Human telomere biology: A contributory and interactive factor in aging, disease risks, and protection. Science. 2015 Dec 4;350(6265):1193–8.  16. Griffith JD, Comeau L, Rosenfield S, Stansel RM, Bianchi A, Moss H, et al. Mammalian telomeres end in a large duplex loop. Cell. 1999 May 14;97(4):503–14.  17. Yang Q, Zheng Y-L, Harris CC. POT1 and TRF2 cooperate to maintain telomeric integrity. Mol. Cell. Biol. 2005 Feb;25(3):1070–80.  18. Arnoult N, Karlseder J. Complex interactions between the DNA-damage response and mammalian telomeres. Nat. Struct. Mol. Biol. 2015 Nov;22(11):859–66.  19. Campisi J, Andersen JK, Kapahi P, Melov S. Cellular senescence: a link between cancer and age-related degenerative disease? Semin. Cancer Biol. 2011 Dec;21(6):354–9.  20. Rodier F, Campisi J. Four faces of cellular senescence. J. Cell Biol. 2011 Feb 21;192(4):547–56.  21. Okazaki R, Okazaki T, Sakabe K, Sugimoto K. Mechanism of DNA replication possible discontinuity of DNA chain growth. Jpn. J. Med. Sci. Biol. 1967 Jun;20(3):255–60.  22. Zhao Y, Shay JW, Wright WE. Telomere terminal G/C strand synthesis: measuring telomerase action and C-rich fill-in. Methods Mol. Biol. 2011;735:63–75.  23. Makarov VL, Hirose Y, Langmore JP. Long G tails at both ends of human chromosomes suggest a C strand degradation mechanism for telomere shortening. Cell. 1997 Mar 7;88(5):657–66.  ! 101!24. Aubert G, Lansdorp PM. Telomeres and aging. Physiol. Rev. 2008 Apr;88(2):557–79.  25. HAYFLICK L. THE LIMITED IN VITRO LIFETIME OF HUMAN DIPLOID CELL STRAINS. Exp. Cell Res. 1965 Mar;37:614–36.  26. Wright WE, Shay JW. Cellular senescence as a tumor-protection mechanism: the essential role of counting. Curr. Opin. Genet. Dev. 2001 Feb;11(1):98–103.  27. Ramirez RD, Morales CP, Herbert BS, Rohde JM, Passons C, Shay JW, et al. Putative telomere-independent mechanisms of replicative aging reflect inadequate growth conditions. Genes Dev. 2001 Feb 15;15(4):398–403.  28. Suram A, Kaplunov J, Patel PL, Ruan H, Cerutti A, Boccardi V, et al. Oncogene-induced telomere dysfunction enforces cellular senescence in human cancer precursor lesions. EMBO J. 2012 Jun 29;31(13):2839–51.  29. Suram A, Herbig U. The replicometer is broken: telomeres activate cellular senescence in response to genotoxic stresses. Aging Cell. 2014 Oct;13(5):780–6.  30. Hewitt G, Jurk D, Marques FDM, Correia-Melo C, Hardy T, Gackowska A, et al. Telomeres are favoured targets of a persistent DNA damage response in ageing and stress-induced senescence. Nat Commun. 2012;3:708.  31. Mather KA, Jorm AF, Parslow RA, Christensen H. Is telomere length a biomarker of aging? A review. J. Gerontol. A Biol. Sci. Med. Sci. 2011 Feb;66(2):202–13.  32. Mather KA, Jorm AF, Milburn PJ, Tan X, Easteal S, Christensen H. No associations between telomere length and age-sensitive indicators of physical function in mid and later life. J. Gerontol. A Biol. Sci. Med. Sci. 2010 Aug;65(8):792–9.  33. Aubert G, Hills M, Lansdorp PM. Telomere length measurement-caveats and a critical assessment of the available technologies and tools. Mutat. Res. 2012 Feb 1;730(1-2):59–67.  34. Bakaysa SL, Mucci LA, Slagboom PE, Boomsma DI, McClearn GE, Johansson B, et al. Telomere length predicts survival independent of genetic influences. Aging Cell. 2007 Dec;6(6):769–74.  ! 102!35. Bischoff C, Petersen HC, Graakjaer J, Andersen-Ranberg K, Vaupel JW, Bohr VA, et al. No association between telomere length and survival among the elderly and oldest old. Epidemiology. 2006 Mar;17(2):190–4.  36. Velarde MC, Demaria M, Campisi J. Senescent cells and their secretory phenotype as targets for cancer therapy. Interdiscip Top Gerontol. 2013;38:17–27.  37. Tchkonia T, Zhu Y, van Deursen J, Campisi J, Kirkland JL. Cellular senescence and the senescent secretory phenotype: therapeutic opportunities. J. Clin. Invest. 2013 Mar 1;123(3):966–72.  38. Peto R. Quantitative implications of the approximate irrelevance of mammalian body size and lifespan to lifelong cancer risk. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 2015 Jul 19;370(1673).  39. Gomes NMV, Ryder OA, Houck ML, Charter SJ, Walker W, Forsyth NR, et al. Comparative biology of mammalian telomeres: hypotheses on ancestral states and the roles of telomeres in longevity determination. Aging Cell. 2011 Oct;10(5):761–8.  40. Trudeau MA, Wong JMY. Genetic Variations in Telomere Maintenance, with Implications on Tissue Renewal Capacity and Chronic Disease Pathologies. Curr Pharmacogenomics Person Med. 2010 Mar 1;8(1):7–24.  41. Bodnar AG, Ouellette M, Frolkis M, Holt SE, Chiu CP, Morin GB, et al. Extension of life-span by introduction of telomerase into normal human cells. Science. 1998 Jan 16;279(5349):349–52.  42. Weinrich SL, Pruzan R, Ma L, Ouellette M, Tesmer VM, Holt SE, et al. Reconstitution of human telomerase with the template RNA component hTR and the catalytic protein subunit hTRT. Nat. Genet. 1997 Dec;17(4):498–502.  43. Greider CW, Blackburn EH. Identification of a specific telomere terminal transferase activity in Tetrahymena extracts. Cell. 1985.  44. Zakian VA. The ends have arrived. Cell. 2009. pp. 1038–40.  45. Lingner J, Hughes TR, Shevchenko A, Mann M, Lundblad V, Cech TR. Reverse transcriptase motifs in the catalytic subunit of telomerase. Science. 1997 Apr 25;276(5312):561–7.  ! 103!46. Nakamura TM, Morin GB, Chapman KB, Weinrich SL, Andrews WH, Lingner J, et al. Telomerase catalytic subunit homologs from fission yeast and human. Science. 1997 Aug 15;277(5328):955–9.  47. Harrington L, McPhail T, Mar V, Zhou W, Oulton R, Bass MB, et al. A mammalian telomerase-associated protein. Science. 1997 Feb 14;275(5302):973–7.  48. Sandin S, Rhodes D. Telomerase structure. Current opinion in structural biology. 2014.  49. Jiang J, Chan H, Cash DD, Miracco EJ, Ogorzalek Loo RR, Upton HE, et al. Structure of Tetrahymena telomerase reveals previously unknown subunits, functions, and interactions. Science. 2015 Oct 30;350(6260):aab4070.  50. Mitchell M, Gillis A, Futahashi M, Fujiwara H, Skordalakes E. Structural basis for telomerase catalytic subunit TERT binding to RNA template and telomeric DNA. Nat. Struct. Mol. Biol. 2010 Apr;17(4):513–8.  51. Sealey D, Zheng L, Taboski M. The N-terminus of hTERT contains a DNA-binding domain and is required for telomerase activity and cellular immortalization. Nucleic acids …. 2010.  52. Lai CK, Mitchell JR, Collins K. RNA binding domain of telomerase reverse transcriptase. Mol. Cell. Biol. 2001 Feb;21(4):990–1000.  53. Gillis AJ, Schuller AP, Skordalakes E. Structure of the Tribolium castaneum telomerase catalytic subunit TERT. Nature. 2008 Oct 2;455(7213):633–7.  54. Zhu S, Rousseau P, Lauzon C, Gandin V, Topisirovic I, Autexier C. Inactive C-terminal telomerase reverse transcriptase insertion splicing variants are dominant-negative inhibitors of telomerase. Biochimie. 2014 Jun;101:93–103.  55. Wong MS, Wright WE, Shay JW. Alternative splicing regulation of telomerase: a new paradigm? Trends Genet. 2014 Oct;30(10):430–8.  56. Hrdlicková R, Nehyba J, Bose HR. Alternatively spliced telomerase reverse transcriptase variants lacking telomerase activity stimulate cell proliferation. Mol. Cell. Biol. 2012 Nov;32(21):4283–96.  ! 104!57. Ulaner GA, Hu JF, Vu TH, Giudice LC, Hoffman AR. Telomerase activity in human development is regulated by human telomerase reverse transcriptase (hTERT) transcription and by alternate splicing of hTERT transcripts. Cancer Res. 1998 Sep 15;58(18):4168–72.  58. Yi X, White DM, Aisner DL, Baur JA, Wright WE, Shay JW. An alternate splicing variant of the human telomerase catalytic subunit inhibits telomerase activity. Neoplasia. 2000 Sep;2(5):433–40.  59. Feng J, Funk WD, Wang SS, Weinrich SL, Avilion AA, Chiu CP, et al. The RNA component of human telomerase. Science. 1995 Sep 1;269(5228):1236–41.  60. Ly H, Blackburn EH, Parslow TG. Comprehensive structure-function analysis of the core domain of human telomerase RNA. Mol. Cell. Biol. 2003 Oct;23(19):6849–56.  61. Egan ED, Collins K. An enhanced H/ACA RNP assembly mechanism for human telomerase RNA. Mol. Cell. Biol. 2012 Jul;32(13):2428–39.  62. Mitchell JR, Collins K. Human telomerase activation requires two independent interactions between telomerase RNA and telomerase reverse transcriptase. Mol. Cell. 2000 Aug;6(2):361–71.  63. Venteicher AS, Abreu EB, Meng Z, McCann KE, Terns RM, Veenstra TD, et al. A human telomerase holoenzyme protein required for Cajal body localization and telomere synthesis. Science. 2009 Jan 30;323(5914):644–8.  64. Stern JL, Zyner KG, Pickett HA, Cohen SB, Bryan TM. Telomerase recruitment requires both TCAB1 and Cajal bodies independently. Mol. Cell. Biol. 2012 Jul;32(13):2384–95.  65. Collins K. The biogenesis and regulation of telomerase holoenzymes. Nat. Rev. Mol. Cell Biol. 2006 Jul;7(7):484–94.  66. Kiss T, Fayet E, Jády BE, Richard P, Weber M. Biogenesis and intranuclear trafficking of human box C/D and H/ACA RNPs. Cold Spring Harb. Symp. Quant. Biol. 2006;71:407–17.  67. Martínez P, Blasco MA. Telomeric and extra-telomeric roles for telomerase and the telomere-binding proteins. Nat. Rev. Cancer. 2011 Mar;11(3):161–76.  ! 105!68. Giraud-Panis M-J, Pisano S, Benarroch-Popivker D, Pei B, Le Du M-H, Gilson E. One identity or more for telomeres? Front Oncol. 2013;3:48.  69. Masutomi K, Yu EY, Khurts S, Ben-Porath I, Currier JL, Metz GB, et al. Telomerase maintains telomere structure in normal human cells. Cell. 2003 Jul 25;114(2):241–53.  70. Fu D, Collins K. Purification of human telomerase complexes identifies factors involved in telomerase biogenesis and telomere length regulation. Mol. Cell. 2007 Dec 14;28(5):773–85.  71. Saretzki G. Telomerase, mitochondria and oxidative stress. Exp. Gerontol. 2009 Aug;44(8):485–92.  72. Kovalenko OA, Caron MJ, Ulema P, Medrano C, Thomas AP, Kimura M, et al. A mutant telomerase defective in nuclear-cytoplasmic shuttling fails to immortalize cells and is associated with mitochondrial dysfunction. Aging Cell. 2010 Apr;9(2):203–19.  73. Jakob S, Schroeder P, Lukosz M, Büchner N, Spyridopoulos I, Altschmied J, et al. Nuclear protein tyrosine phosphatase Shp-2 is one important negative regulator of nuclear export of telomerase reverse transcriptase. J. Biol. Chem. 2008 Nov 28;283(48):33155–61.  74. Chung J, Khadka P, Chung IK. Nuclear import of hTERT requires a bipartite nuclear localization signal and Akt-mediated phosphorylation. J. Cell. Sci. 2012 Jun 1;125(Pt 11):2684–97.  75. Seimiya H, Sawada H, Muramatsu Y, Shimizu M, Ohko K, Yamane K, et al. Involvement of 14-3-3 proteins in nuclear localization of telomerase. EMBO J. 2000 Jun 1;19(11):2652–61.  76. Ouellette MM, Liao M, Herbert BS, Johnson M. Subsenescent telomere lengths in fibroblasts immortalized by limiting amounts of telomerase. Journal of Biological …. 2000.  77. Zou Y, Sfeir A, Gryaznov SM, Shay JW. Does a sentinel or a subset of short telomeres determine replicative senescence? Molecular biology of …. 2004.  ! 106!78. Loayza D, de Lange T. POT1 as a terminal transducer of TRF1 telomere length control. Nature. 2003 Jun 26;423(6943):1013–8.  79. Abreu E, Aritonovska E, Reichenbach P, Cristofari G, Culp B, Terns RM, et al. TIN2-tethered TPP1 recruits human telomerase to telomeres in vivo. Mol. Cell. Biol. 2010 Jun;30(12):2971–82.  80. Rice C, Skordalakes E. Structure and function of the telomeric CST complex. Comput Struct Biotechnol J. 2016;14:161–7.  81. García-Cao M, O'Sullivan R, Peters AHFM, Jenuwein T, Blasco MA. Epigenetic regulation of telomere length in mammalian cells by the Suv39h1 and Suv39h2 histone methyltransferases. Nat. Genet. 2004 Jan;36(1):94–9.  82. Chan SS, Chang S. Defending the end zone: studying the players involved in protecting chromosome ends. FEBS Lett. 2010 Sep 10;584(17):3773–8.  83. Lenain C, Bauwens S, Amiard S, Brunori M, Giraud-Panis M-J, Gilson E. The Apollo 5' exonuclease functions together with TRF2 to protect telomeres from DNA repair. Curr. Biol. 2006 Jul 11;16(13):1303–10.  84. Singh DK, Ghosh AK, Croteau DL, Bohr VA. RecQ helicases in DNA double strand break repair and telomere maintenance. Mutat. Res. 2012 Aug 1;736(1-2):15–24.  85. Cesare AJ, Karlseder J. A three-state model of telomere control over human proliferative boundaries. Curr. Opin. Cell Biol. 2012 Dec;24(6):731–8.  86. Tong AS, Stern JL, Sfeir A, Kartawinata M, de Lange T, Zhu X-D, et al. ATM and ATR Signaling Regulate the Recruitment of Human Telomerase to Telomeres. Cell Rep. 2015 Nov 24;13(8):1633–46.  87. Greider CW. Regulating telomere length from the inside out: The replication fork model. bioRxiv. 2016.  88. Shay JW, Pereira-Smith OM, Wright WE. A role for both RB and p53 in the regulation of human cellular senescence. Exp. Cell Res. 1991 Sep;196(1):33–9.  89. Savage SA, Gadalla SM, Chanock SJ. The long and short of telomeres and cancer association studies. J. Natl. Cancer Inst. 2013 Apr 3;105(7):448–9.  ! 107!90. Kim NW, Piatyszek MA, Prowse KR, Harley CB, West MD, Ho PL, et al. Specific association of human telomerase activity with immortal cells and cancer. Science. 1994 Dec 23;266(5193):2011–5.  91. Shay JW, Reddel RR, Wright WE. Cancer. Cancer and telomeres--an ALTernative to telomerase. Science. 2012 Jun 15;336(6087):1388–90.  92. Hiyama E, Hiyama K, Yokoyama T, Matsuura Y, Piatyszek MA, Shay JW. Correlating telomerase activity levels with human neuroblastoma outcomes. Nat. Med. 1995 Mar;1(3):249–55.  93. Tabori U, Vukovic B, Zielenska M, Hawkins C, Braude I, Rutka J, et al. The role of telomere maintenance in the spontaneous growth arrest of pediatric low-grade gliomas. Neoplasia. 2006 Feb;8(2):136–42.  94. Bojesen SE, Pooley KA, Johnatty SE, Beesley J, Michailidou K, Tyrer JP, et al. Multiple independent variants at the TERT locus are associated with telomere length and risks of breast and ovarian cancer. Nat. Genet. 2013 Apr;45(4):371–84–384e1–2.  95. Borah S, Xi L, Zaug AJ, Powell NM, Dancik GM, Cohen SB, et al. Cancer. TERT promoter mutations and telomerase reactivation in urothelial cancer. Science. 2015 Feb 27;347(6225):1006–10.  96. Heidenreich B, Rachakonda PS, Hemminki K, Kumar R. TERT promoter mutations in cancer development. Curr. Opin. Genet. Dev. 2014 Feb;24:30–7.  97. Stern JL, Theodorescu D, Vogelstein B, Papadopoulos N, Cech TR. Mutation of the TERT promoter, switch to active chromatin, and monoallelic TERT expression in multiple cancers. Genes Dev. 2015 Nov 1;29(21):2219–24.  98. Vinagre J, Almeida A, Pópulo H, Batista R, Lyra J, Pinto V, et al. Frequency of TERT promoter mutations in human cancers. Nat Commun. 2013;4:2185.  99. Wong MS, Shay JW, Wright WE. Regulation of human telomerase splicing by RNA:RNA pairing. Nat Commun. 2014;5:3306.  100. Shay JW, Wright WE. Implications of mapping the human telomerase gene (hTERT) as the most distal gene on chromosome 5p. Neoplasia. 2000 ! 108!May;2(3):195–6.  101. Bryan TM, Englezou A, Dalla-Pozza L, Dunham MA, Reddel RR. Evidence for an alternative mechanism for maintaining telomere length in human tumors and tumor-derived cell lines. Nat. Med. 1997 Nov;3(11):1271–4.  102. Fasching CL, Bower K, Reddel RR. Telomerase-independent telomere length maintenance in the absence of alternative lengthening of telomeres-associated promyelocytic leukemia bodies. Cancer Res. 2005 Apr 1;65(7):2722–9.  103. Ogino H, Nakabayashi K, Suzuki M, Takahashi E, Fujii M, Suzuki T, et al. Release of telomeric DNA from chromosomes in immortal human cells lacking telomerase activity. Biochem. Biophys. Res. Commun. 1998 Jul 20;248(2):223–7.  104. Londoño-Vallejo JA, Der-Sarkissian H, Cazes L, Bacchetti S, Reddel RR. Alternative lengthening of telomeres is characterized by high rates of telomeric exchange. Cancer Res. 2004 Apr 1;64(7):2324–7.  105. Muntoni A, Neumann AA, Hills M, Reddel RR. Telomere elongation involves intra-molecular DNA replication in cells utilizing alternative lengthening of telomeres. Hum. Mol. Genet. 2009 Mar 15;18(6):1017–27.  106. Henson JD, Neumann AA, Yeager TR, Reddel RR. Alternative lengthening of telomeres in mammalian cells. Oncogene. 2002 Jan 21;21(4):598–610.  107. O'Sullivan RJ, Almouzni G. Assembly of telomeric chromatin to create ALTernative endings. Trends Cell Biol. 2014 Nov;24(11):675–85.  108. Yeager TR, Neumann AA, Englezou A, Huschtscha LI, Noble JR, Reddel RR. Telomerase-negative immortalized human cells contain a novel type of promyelocytic leukemia (PML) body. Cancer Res. 1999 Sep 1;59(17):4175–9.  109. Draskovic I, Arnoult N, Steiner V, Bacchetti S, Lomonte P, Londoño-Vallejo A. Probing PML body function in ALT cells reveals spatiotemporal requirements for telomere recombination. Proc. Natl. Acad. Sci. U.S.A. 2009 Sep 15;106(37):15726–31.  110. Heaphy CM, Subhawong AP, Hong S-M, Goggins MG, Montgomery EA, Gabrielson E, et al. Prevalence of the alternative lengthening of telomeres telomere ! 109!maintenance mechanism in human cancer subtypes. Am. J. Pathol. 2011 Oct;179(4):1608–15.  111. Hu J, Hwang SS, Liesa M, Gan B, Sahin E, Jaskelioff M, et al. Antitelomerase therapy provokes ALT and mitochondrial adaptive mechanisms in cancer. Cell. 2012 Feb 17;148(4):651–63.  112. Buseman CM, Wright WE, Shay JW. Is telomerase a viable target in cancer? Mutat. Res. 2012 Feb 1;730(1-2):90–7.  113. Ahmed S, Passos JF, Birket MJ, Beckmann T. Telomerase does not counteract telomere shortening but protects mitochondrial function under oxidative stress. Journal of cell …. 2008.  114. Jafri MA, Ansari SA, Alqahtani MH, Shay JW. Roles of telomeres and telomerase in cancer, and advances in telomerase-targeted therapies. Genome Med. 2016;8(1):69.  115. Wu RA, Dagdas YS, Yilmaz ST, Yildiz A, Collins K. Single-molecule imaging of telomerase reverse transcriptase in human telomerase holoenzyme and minimal RNP complexes. Elife. 2015;4.  116. Bryan C, Rice C, Hoffman H, Harkisheimer M, Sweeney M, Skordalakes E. Structural Basis of Telomerase Inhibition by the Highly Specific BIBR1532. Structure. 2015 Oct 6;23(10):1934–42.  117. Hukezalie KR, Thumati NR, Côté HCF, Wong JMY. In vitro and ex vivo inhibition of human telomerase by anti-HIV nucleoside reverse transcriptase inhibitors (NRTIs) but not by non-NRTIs. PLoS ONE. 2012;7(11):e47505.  118. Vogel M, Friedrich O, Lüchters G, Holleczek B, Wasmuth JC, Anadol E, et al. Cancer risk in HIV-infected individuals on HAART is largely attributed to oncogenic infections and state of immunocompetence. Eur. J. Med. Res. 2011 Mar 28;16(3):101–7.  119. Lam EYN, Beraldi D, Tannahill D, Balasubramanian S. G-quadruplex structures are stable and detectable in human genomic DNA. Nat Commun. 2013;4:1796.  120. Chambers VS, Marsico G, Boutell JM, Di Antonio M, Smith GP, Balasubramanian ! 110!S. High-throughput sequencing of DNA G-quadruplex structures in the human genome. Nat. Biotechnol. 2015 Aug;33(8):877–81.  121. Martínez P, Blasco MA. Replicating through telomeres: a means to an end. Trends Biochem. Sci. 2015 Sep;40(9):504–15.  122. Balasubramanian S, Hurley LH, Neidle S. Targeting G-quadruplexes in gene promoters: a novel anticancer strategy? Nat Rev Drug Discov. 2011 Apr;10(4):261–75.  123. Spinelli N, Labbé P, Monchaud D. Template‐Assembled Synthetic G‐Quadruplex (TASQ): A Useful System for Investigating the Interactions of Ligands with Constrained Quadruplex Topologies. … A European Journal. 2010.  124. Laguerre A, Hukezalie K, Winckler P, Katranji F, Chanteloup G, Pirrotta M, et al. Visualization of RNA-Quadruplexes in Live Cells. J. Am. Chem. Soc. 2015 Jul 8;137(26):8521–5.  125. Mender I, Gryaznov S, Shay JW. A novel telomerase substrate precursor rapidly induces telomere dysfunction in telomerase positive cancer cells but not telomerase silent normal cells. Oncoscience. 2015;2(8):693–5.  126. Mender I, Gryaznov S, Dikmen ZG, Wright WE, Shay JW. Induction of telomere dysfunction mediated by the telomerase substrate precursor 6-thio-2'-deoxyguanosine. Cancer Discov. 2015 Jan;5(1):82–95.  127. Kyte JA. Cancer vaccination with telomerase peptide GV1001. Expert Opin Investig Drugs. 2009 May;18(5):687–94.  128. Tefferi A, Lasho TL, Begna KH, Patnaik MM, Zblewski DL, Finke CM, et al. A Pilot Study of the Telomerase Inhibitor Imetelstat for Myelofibrosis. N. Engl. J. Med. 2015 Sep 3;373(10):908–19.  129. Herbert B-S, Gellert GC, Hochreiter A, Pongracz K, Wright WE, Zielinska D, et al. Lipid modification of GRN163, an N3“-->P5” thio-phosphoramidate oligonucleotide, enhances the potency of telomerase inhibition. Oncogene. 2005 Aug 4;24(33):5262–8.  130. Dikmen ZG, Gellert GC, Jackson S, Gryaznov S, Tressler R, Dogan P, et al. In vivo ! 111!inhibition of lung cancer by GRN163L: a novel human telomerase inhibitor. Cancer Res. 2005 Sep 1;65(17):7866–73.  131. Hu Y, Bobb D, He J, Hill DA, Dome JS. The HSP90 inhibitor alvespimycin enhances the potency of telomerase inhibition by imetelstat in human osteosarcoma. Cancer Biol. Ther. 2015;16(6):949–57.  132. Lu R, Pal J, Buon L, Nanjappa P, Shi J, Fulciniti M, et al. Targeting homologous recombination and telomerase in Barrett's adenocarcinoma: impact on telomere maintenance, genomic instability and tumor growth. Oncogene. 2014 Mar 20;33(12):1495–505.  133. Baerlocher GM, Oppliger Leibundgut E, Ottmann OG, Spitzer G, Odenike O, McDevitt MA, et al. Telomerase Inhibitor Imetelstat in Patients with Essential Thrombocythemia. N. Engl. J. Med. 2015 Sep 3;373(10):920–8.  134. Search Orphan Drug Designations and Approvals.  135. Jackson SR, Zhu CH, Paulson V, Watkins L. Antiadhesive effects of GRN163L—an oligonucleotide N3′→ P5′ thio-phosphoramidate targeting telomerase. Cancer Res. 2007.  136. Mender I, Senturk S, Ozgunes N, Akcali KC, Kletsas D, Gryaznov S, et al. Imetelstat (a telomerase antagonist) exerts off‑target effects on the cytoskeleton. Int. J. Oncol. 2013 May;42(5):1709–15.  137. Faria M, Spiller DG, Dubertret C, Nelson JS, White MR, Scherman D, et al. Phosphoramidate oligonucleotides as potent antisense molecules in cells and in vivo. Nat. Biotechnol. 2001 Jan;19(1):40–4.  138. GOODMAN LS, WINTROBE MM. Nitrogen mustard therapy; use of methyl-bis (beta-chloroethyl) amine hydrochloride and tris (beta-chloroethyl) amine hydrochloride for Hodgkin's disease, lymphosarcoma, leukemia and certain allied and miscellaneous disorders. J Am Med Assoc. 1946 Sep 21;132:126–32.  139. FARBER S, DIAMOND LK. Temporary remissions in acute leukemia in children produced by folic acid antagonist, 4-aminopteroyl-glutamic acid. N. Engl. J. Med. 1948 Jun 3;238(23):787–93.  ! 112!140. Cheung-Ong K, Giaever G, Nislow C. DNA-damaging agents in cancer chemotherapy: serendipity and chemical biology. Chem. Biol. 2013 May 23;20(5):648–59.  141. Kelland L. The resurgence of platinum-based cancer chemotherapy. Nat. Rev. Cancer. 2007 Aug;7(8):573–84.  142. Parker WB, Cheng YC. Metabolism and mechanism of action of 5-fluorouracil. Pharmacol. Ther. 1990;48(3):381–95.  143. Pommier Y. Drugging topoisomerases: lessons and challenges. ACS Chem. Biol. 2013 Jan 18;8(1):82–95.  144. Sawicka M, Kalinowska M, Skierski J, Lewandowski W. A review of selected anti-tumour therapeutic agents and reasons for multidrug resistance occurrence. J. Pharm. Pharmacol. 2004 Sep;56(9):1067–81.  145. Forterre P, Gribaldo S, Gadelle D, Serre M-C. Origin and evolution of DNA topoisomerases. Biochimie. 2007 Apr;89(4):427–46.  146. Froelich-Ammon SJ, Osheroff N. Topoisomerase poisons: harnessing the dark side of enzyme mechanism. J. Biol. Chem. 1995 Sep 15;270(37):21429–32.  147. Burgess DJ, Doles J, Zender L, Xue W, Ma B, McCombie WR, et al. Topoisomerase levels determine chemotherapy response in vitro and in vivo. Proc. Natl. Acad. Sci. U.S.A. 2008 Jul 1;105(26):9053–8.  148. Tamakawa RA, Fleisig HB, Wong JMY. Telomerase inhibition potentiates the effects of genotoxic agents in breast and colorectal cancer cells in a cell cycle-specific manner. Cancer Res. 2010 Nov 1;70(21):8684–94.  149. Tamakawa RA. Combination chemotherapy with telomerase inhibitors and genotoxic compounds against breast and colorectal cancers. 2010.  150. Verdun RE, Crabbe L, Haggblom C, Karlseder J. Functional human telomeres are recognized as DNA damage in G2 of the cell cycle. Mol. Cell. 2005 Nov 23;20(4):551–61.  151. Verdun RE, Karlseder J. The DNA damage machinery and homologous ! 113!recombination pathway act consecutively to protect human telomeres. Cell. 2006 Nov 17;127(4):709–20.  152. Lee SS, Bohrson C, Pike AM, Wheelan SJ, Greider CW. ATM Kinase Is Required for Telomere Elongation in Mouse and Human Cells. Cell Rep. 2015 Nov 24;13(8):1623–32.  153. Hukezalie KR. Characterizing the effects of N/NRTIs on human telomerase activity in vitro and telomere maintenance in a transformed human cell model. 2011.  154. Takai H, Smogorzewska A, de Lange T. DNA damage foci at dysfunctional telomeres. Curr. Biol. 2003 Sep 2;13(17):1549–56.  155. Fleisig HB, Wong JMY. Telomerase promotes efficient cell cycle kinetics and confers growth advantage to telomerase-negative transformed human cells. Oncogene. 2012 Feb 23;31(8):954–65.  156. Wu P, Takai H, de Lange T. Telomeric 3' overhangs derive from resection by Exo1 and Apollo and fill-in by POT1b-associated CST. Cell. 2012 Jul 6;150(1):39–52.  157. Kibe T, Zimmermann M, de Lange T. TPP1 Blocks an ATR-Mediated Resection Mechanism at Telomeres. Mol. Cell. 2016 Jan 21;61(2):236–46.  158. Bolderson E, Tomimatsu N, Richard DJ, Boucher D, Kumar R, Pandita TK, et al. Phosphorylation of Exo1 modulates homologous recombination repair of DNA double-strand breaks. Nucleic Acids Res. 2010 Apr;38(6):1821–31.  159. Xie Z, Jay KA, Smith DL, Zhang Y, Liu Z, Zheng J, et al. Early telomerase inactivation accelerates aging independently of telomere length. Cell. 2015 Feb 26;160(5):928–39.  160. Kohnken R, Kodigepalli KM, Wu L. Regulation of deoxynucleotide metabolism in cancer: novel mechanisms and therapeutic implications. Mol. Cancer. 2015;14:176.  161. Smith LL, Coller HA, Roberts JM. Telomerase modulates expression of growth-controlling genes and enhances cell proliferation. Nat. Cell Biol. 2003 May;5(5):474–9.  162. Park J-I, Venteicher AS, Hong JY, Choi J, Jun S, Shkreli M, et al. Telomerase ! 114!modulates Wnt signalling by association with target gene chromatin. Nature. 2009 Jul 2;460(7251):66–72.  163. Listerman I, Gazzaniga FS, Blackburn EH. An investigation of the effects of the core protein telomerase reverse transcriptase on Wnt signaling in breast cancer cells. Mol. Cell. Biol. 2014 Jan;34(2):280–9.  164. Maida Y, Yasukawa M, Furuuchi M, Lassmann T, Possemato R, Okamoto N, et al. An RNA-dependent RNA polymerase formed by TERT and the RMRP RNA. Nature. 2009 Sep 10;461(7261):230–5.  165. Werner C, Gensch C, Pöss J, Haendeler J, Böhm M, Laufs U. Pioglitazone activates aortic telomerase and prevents stress-induced endothelial apoptosis. Atherosclerosis. 2011 May;216(1):23–34.  166. Santos JH, Meyer JN, Van Houten B. Mitochondrial localization of telomerase as a determinant for hydrogen peroxide-induced mitochondrial DNA damage and apoptosis. Hum. Mol. Genet. 2006 Jun 1;15(11):1757–68.  167. Sahin E, Colla S, Liesa M, Moslehi J, Muller FL, Guo M, et al. Telomere dysfunction induces metabolic and mitochondrial compromise. Nature. 2011 Feb 17;470(7334):359–65.  168. Sahin E, Depinho RA. Axis of ageing: telomeres, p53 and mitochondria. Nat. Rev. Mol. Cell Biol. 2012 Jun;13(6):397–404.  169. Del Bufalo D, Rizzo A, Trisciuoglio D, Cardinali G, Torrisi MR, Zangemeister-Wittke U, et al. Involvement of hTERT in apoptosis induced by interference with Bcl-2 expression and function. Cell Death Differ. 2005 Nov;12(11):1429–38.  170. Kang HJ, Choi YS, Hong S-B, Kim K-W, Woo R-S, Won SJ, et al. Ectopic expression of the catalytic subunit of telomerase protects against brain injury resulting from ischemia and NMDA-induced neurotoxicity. J. Neurosci. 2004 Feb 11;24(6):1280–7.  171. Lee J, Sung YH, Cheong C, Choi YS, Jeon HK, Sun W, et al. TERT promotes cellular and organismal survival independently of telomerase activity. Oncogene. 2008 Jun 12;27(26):3754–60.  ! 115!172. Sarin KY, Cheung P, Gilison D, Lee E, Tennen RI, Wang E, et al. Conditional telomerase induction causes proliferation of hair follicle stem cells. Nature. 2005 Aug 18;436(7053):1048–52.  173. Fleisig HB, Wong JMY. Telomerase as a clinical target: current strategies and potential applications. Exp. Gerontol. 2007 Jan;42(1-2):102–12.  174. Perrem K, Colgin LM, Neumann AA, Yeager TR, Reddel RR. Coexistence of alternative lengthening of telomeres and telomerase in hTERT-transfected GM847 cells. Mol. Cell. Biol. 2001 Jun;21(12):3862–75.  175. Fleisig HB, Hukezalie KR, Thompson CAH, Au-Yeung TTT, Ludlow AT, Zhao CR, et al. Telomerase reverse transcriptase expression protects transformed human cells against DNA-damaging agents, and increases tolerance to chromosomal instability. Oncogene. 2016 Jan 14;35(2):218–27.  176. Fleisig H, Wong J. Measuring cell cycle progression kinetics with metabolic labeling and flow cytometry. J Vis Exp. 2012;(63):e4045.  177. Armbruster BN, Etheridge KT, Broccoli D, Counter CM. Putative telomere-recruiting domain in the catalytic subunit of human telomerase. Mol. Cell. Biol. 2003 May;23(9):3237–46.  178. Listerman I, Sun J, Gazzaniga FS, Lukas JL, Blackburn EH. The major reverse transcriptase-incompetent splice variant of the human telomerase protein inhibits telomerase activity but protects from apoptosis. Cancer Res. 2013 May 1;73(9):2817–28.  179. Fenech M. Cytokinesis-block micronucleus cytome assay. Nat Protoc. 2007;2(5):1084–104.  180. Poon SS, Lansdorp PM. Measurements of telomere length on individual chromosomes by image cytometry. Methods Cell Biol. 2001;64:69–96.  181. Lovejoy CA, Li W, Reisenweber S, Thongthip S, Bruno J, de Lange T, et al. Loss of ATRX, genome instability, and an altered DNA damage response are hallmarks of the alternative lengthening of telomeres pathway. PLoS Genet. 2012;8(7):e1002772.  ! 116!182. Shawi M, Chu TW, Martinez-Marignac V, Yu Y, Gryaznov SM, Johnston JB, et al. Telomerase contributes to fludarabine resistance in primary human leukemic lymphocytes. PLoS ONE. 2013;8(7):e70428.  183. Gordon DM, Santos JH. The emerging role of telomerase reverse transcriptase in mitochondrial DNA metabolism. J Nucleic Acids. 2010;2010.  184. Ding D, Zhou J, Wang M, Cong Y-S. Implications of telomere-independent activities of telomerase reverse transcriptase in human cancer. FEBS J. 2013 Jul;280(14):3205–11.  185. Carneiro T, Khair L, Reis CC, Borges V, Moser BA, Nakamura TM, et al. Telomeres avoid end detection by severing the checkpoint signal transduction pathway. Nature. 2010 Sep 9;467(7312):228–32.  186. Vaziri H, West MD, Allsopp RC, Davison TS, Wu YS, Arrowsmith CH, et al. ATM-dependent telomere loss in aging human diploid fibroblasts and DNA damage lead to the post-translational activation of p53 protein involving poly(ADP-ribose) polymerase. EMBO J. 1997 Oct 1;16(19):6018–33.             ! 117!APPENDIX: Supplementary Figures !!Treatment 0-24 h Treatment 24-48 h LD50 (nM) No treatment Etoposide alone 86.7 ± 5.8 GRN163L  Etoposide + GRN163L  40.6 ± 6.2 KU55933  Etoposide + KU55933  24.7 ± 1.1 KU55933  Etoposide + KU55933 + GRN163L  26.6 ± 6.0 GRN163L  Etoposide + KU55933 + GRN163L  15.6 ± 2.6  Supplementary Figure 1. Calculated LD50s for order-of-addition CFU experiments    Supplementary Figure 2. GRN163L inhibits telomerase activity in a dose-dependent manner MCF-7 and MDA-MB 231 cells undergoing GRN163L treatments at concentrations observed to reduce cellular proliferation were assessed for telomerase activity using TRAP. GRN163L at 10 µM efficiently inhibits telomerase catalytic activity, 2 µM shows an intermediate effect. (with Jialin Xu) ! 118!!   Supplementary Figure 3. KU55933 does not effect telomerase inhibition by GRN163L Telomerase Repeat Amplification Protocol (TRAP) assay was used on extracts from cells undergoing the same treatments to assess whether ATMi affects the efficiency of telomerase inhibition by GRN163L. (with Jialin Xu)!! 119!! Supplementary Figure 4. Secondary Alone Controls for GRN163L induced DDR foci GRN163L treatment does not increase non-specific γH2AX antibody staining in MCF-7 or HT29 cell lines.  !!


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