Characterization of the DNA Binding Properties of CST (CTC1-STN1-TEN1) and Their

Characterization of the DNA Binding Properties of CST (CTC1-STN1-TEN1) And Their

Importance for CST Function in Telomeric as well as Genome-wide Replication

A dissertation submitted to the

Division of Graduate Studies and Research

Of the University of Cincinnati

In partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY (Ph.D.)

In the Department of Molecular Genetics, Biochemistry and Microbiology

in College of Medicine

at University of Cincinnati

2017

Anukana Bhattacharjee

B.S., University of Calcutta, India, 2008

M.S., University of Calcutta, India, 2010

Committee Chair: Carolyn M. Price, Ph.D

Committee Members:

Anil Menon, Ph.D

Iain Cartwright, Ph.D

Rhett Kovall, Ph.D

Satoshi Namekawa, Ph.D

Abstract: Telomeres are the end of chromosomes that protect DNA ends from being recognized as DNA damage and act as a buffer for loss of DNA at the chromosome terminus. Telomeric DNA has a unique structure as it is composed of kilobases of double-stranded DNA with a tandem repetitive sequence

(TTAGGG. AATCCC) followed by a short single-stranded overhang region. Telomeres are bound by a number of proteins that help in protection of telomeres from damage signaling and chromosomal fusions as well as help in telomere replication and functions. In vertebrates, the primary telomere binding protein complex is shelterin, which is composed of six subunits, that bind to both double strand and single strand regions of telomere and bridges between them. Shelterin is important for protecting telomeres from damage and also brings in telomerase for telomere extension. The other major telomere binding protein complex is CST (CTC1-STN1-TEN1) which has been shown to localize at telomeres (1). Human CST is a ssDNA-binding complex that was originally identified as a DNA polymerase α stimulatory factor. CST functions in telomere replication first by aiding passage of the replication machinery through the telomere duplex and then enabling fill-in synthesis of the telomeric C-strand following telomerase action. CST also binds to ssDNA other than telomeres and has genome-wide roles in the resolution of replication stress.

CST bears striking resemblance to RPA, the ssDNA binding protein responsible for moderating key transactions in DNA replication, recombination and repair. STN1 and TEN1 contain OB fold domains and are structurally similar to RPA2 and RPA3 respectively. While CTC1 is much larger than RPA1, the

C-terminus is predicted to harbor three OB folds with high structural similarity to the three DNA binding motifs of RPA1 (OB folds A-C). The similarities between CST and RPA suggested that the various functions of CST might utilize subsets of OB folds for different modes of DNA binding. To address this possibility, we generated a CST DNA binding mutant by altering three residues in the STN1 OB fold

(STN1-OBM). The equivalent residues in RPA2 contact or lie close to DNA in the crystal structure. In vitro studies indicated that STN1-OBM moderately decreases CST binding to short G-strand oligonucleotides; however, binding to long telomeric or non-telomeric oligonucleotides is largely unaffected. These results indicate that the STN1 OB fold is responsible for high affinity binding to short stretches of telomeric G-strand DNA. Moreover, CST appears to resemble RPA in exhibiting different

DNA binding modes but the trajectory of DNA engagement is different. Our data suggest STN1, TEN1

OB-folds lie close to the 3’ end of ssDNA even for the shortest oligonucleotide CST binds to, in contrast to RPA where only the longest oligonucleotide contacts RPA2. To determine the in vivo effect of altered

DNA binding, we asked if STN1-OBM expression alters telomere replication or genome-wide replication rescue. Interestingly, we found STN1-OBM to be a separation of function mutant. The STN1-OBM cells had increased anaphase bridges and multiple telomeric FISH signals (MTS). However, the length of the telomeric G-overhang and the rate of C-strand fill-in were normal. Likewise, the cells showed wild type sensitivity to hydroxyurea (HU) and the level of new origin firing after release from HU was unaffected.

Thus, the ability to bind short stretches of ssDNA appears to be important for replication through natural barriers such as telomeres but is less critical for C-strand fill-in or stress-induced origin firing. Overall our work suggests that CST binds DNA dynamically via multiple OB folds and mediates different transactions via specific DNA binding modes.

Although the architecture or modes of DNA binding differ for RPA and CST, their overall structural similarity motivated us to use RPA as a model to investigate DNA binding properties underlying CST function. RPA binds to ssDNA with high affinity via OB-fold domains. Yet individual OB-folds of RPA can micro-dissociate from the DNA promoting sliding of RPA on the DNA, melting of dsDNA or secondary structures, as well as loading or unloading of interaction partners. This dynamic binding underlies the various roles of RPA in replication, repair and recombination. By using single molecule fluorescence assays, we show that in contrast to RPA, CST cannot melt dsDNA but it can resolve secondary structures such as G4. The efficiency of G4 unfolding by CST, and its known abundance in G- rich regions genome-wide could explain its role in resolution of replication stress. Our work has also shown that CST can recognize ss-dsDNA junction. Previous studies have shown that during telomere replication, the C-strand fill-in reaction occurs via incremental extension of the 5’ terminus by lagging strand synthesis. The ss-dsDNA junction recognition explains how CST could promote this incremental

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DNA synthesis. Overall, our work provides insight into the mechanism by which CST might resolve replication issues at the telomere and genome wide.

Acknowledgements:

My long and fruitful journey of science would not have been possible without the love and support of some amazing people in my life. I always believe that they taught me to learn, appreciate science and develop into a scientist. First, I would like to convey my sincere gratitude to Dr. Carolyn Price, my graduate advisor. She has been an incredible mentor to me. When I look back now, I realize that she taught me not only to conduct lab experiments, write scientific manuscripts and critically think about science, but she also taught me to deal with life and face challenges. I would also like to thank my committee members Dr. Rhett Kovall, Dr. Iain Cartwright, Dr. Anil Menon and Dr. Satoshi Namekawa for their insightful comments and guiding me throughout my graduate school. Today I appreciate the tough love you showed towards me. It was a pleasure learning science with you all.

Next, I would like to thank the past and present members of Price lab. I have made friends who were there for me during my ups and downs. Their suggestions, lab-help and listening to me during tough times, got me going. I would like to specially thank Dr. Jason Stewart who mentored me and taught me to grow up, both as a scientist and as a person. I would also like to thank all the people in our department who had always been there for me when I needed them the most. I greatly appreciate the help from Dr. Sandra

Degen and Dr. Edmund Choi who arranged financial support for me under exceptional circumstances.

They made it possible for me to join Price lab for the second time when there was no funding available.

On a slightly different note, a special thanks to Henrietta Lacks, whose immortal Hela cells was the fundamental cell system throughout the first part of my research.

Finally, I would like to take time to thank my family- my parents and my fiancé, the strongest support and foundation of my life. The biggest influence of my scientific career are my parents, who have always encouraged me to ask questions, stimulated me to understand the “why and how” of everything. I always look up to them and if I could be half as successful as my parents are in their professional and personal life, I would consider myself to be well achieved. And last but not the least, I would like to thank my fiancé, Dr. Soumitra Ghosh, without whose support, I might have given up a long time ago. He has

vi always been there to listen to me whether I am happy or frustrated, give me the right advice and push me to achieve my best. Thanks for being so patient with all my tantrums, madness and still believing in me. It all paid off well. I couldn’t have made it this far without you.

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Table of contents

1. Introduction…………………………………………………………………………………….…1

1.1 Structure of telomeric DNA………………………………………………………………..….1

1.2 Replication………………………………………………………………………………...... 2

1.2.1 General eukaryotic replication regulation and replication stress………..…………....2

1.2.2 The end replication problem………………………………………….…………….…4

1.3 Telomerase………………………………………………………………………...……….….5

1.4 Telomeric replication stress……………………………………………………….….………..8

1.5 Telomeric proteins…………………………………………………………………….……….8

1.5.1 Vertebrate Shelterin………………………………………………...………..………12

1.5.2 Mammalian CST…………………………………………………...………….…….14

1.6 Telomeres and Diseases………………………………………...……………………….…...17

1.6.1 Telomere and Cancer ………………………………………………………….…….17

1.6.2 Disease associated with short telomeres …………….……………………………...18

1.7 Perspectives and conclusions…………………………………..…………………………….19

1.8 Dissertation goals ……………………………………………...………………………….…20

2. Material and Methods ……………………………………………...…………………….…….21

2.1 Generation of STN1-OB cells and verification of cell lines (HeLa)……….……………...21

2.2 Western Blots………………………………………………………………...…………….22

2.3 Anaphase bridge analysis…………………………………………………..………………22

2.4 Telomere FISH……………………………………………………………………………..22

2.5 Genomic DNA isolation……………………………………………………………………23

2.6 Cell cycle synchronization and G-overhang analysis…………………………….………..23

2.7 MTT assay………………………………………………………………………………….23

2.8 DNA fiber analysis……………………………………………………………...………….24

2.9 Co-immunoprecipitation…………………………………………………...………………24

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2.10 Isolation of bacmid DNA…………………………………………………………………..24

2.11 Sf9 cell culture……………………………………………………..………..…………….25

2.12 Transfection of Sf9 cells with bacmid DNA for baculo-virus production………….…….25

2.13 Baculo-virus amplification………………………………………………………..………25

2.14 CST protein expression in insect cells………………………………………….…………26

2.15 CST purification from insect cells……………………………………………..………….26

2.16 CST expression and purification from human cells………………………………………27

2.17 Preparation of DNA constructs for in vitro binding assays……………………………….27

2.18 Electrophoretic mobility shift assays and UV crosslinking………………………………28

2.19 Filter binding assay to determine Kd,app and t1/2 ……………………………………28

2.20 Strand melting assay………………………………………………………………29

2.21 Single molecule fluorescence resonance energy transfer (smFRET) assay………29

2.22 smFRET data acquisition…………………………………………………………30

2.23 smFRET data analysis…………………………………….………………………30

3. STN1 OB Fold Mutation Alters DNA Binding and Affects Selective Aspects of CST

Function……………………………………………………………………………...…………..32

3.1 Abstract………………………………………………………………………..……………..32

3.2 Author summary………………………………………………………………………...……33

3.3 Introduction…………………………………………………………………………………..33

3.4 Results………………………………………………………………………………..………36

3.4.1 Alteration of DNA binding by STN1 OB-fold mutation……………………………36

3.4.2 STN1-OBM fails to rescue anaphase bridges after endogenous STN1 depletion.…39

3.4.3 STN1-OBM affects telomere duplex replication but not C-strand fill-in………..….41

3.4.4 STN1-OBM does not disrupt interaction with TPP1 or DNA pol α…………..…….46

3.4.5 STN1-OBM can function in replication rescue after genome wide replication fork

stalling…………………………………………………………………...……….….47

3.4.6 Effects of STN1-OBM on binding affinity and stability…………….………………49

3.4.7 CST subunit interaction with ssDNA substrates……………………….……………52

3.5 Discussion………………………………………………………………..……..……………56

3.6 Supplemental Figures…………………………………………………...……………………60

4. CST DNA-binding dynamics melt G4 structure and regulate protein association with partial duplex and ssDNA………………………………………………..……………………..66 4.1 Abstract……………………………………………………………..………………………..66

4.2 Introduction………………………………………………………..…………………………66

4.3 Results…………………………………………………………………..……………………68

4.3.1 CST recognizes ss-dsDNA junctions but does not melt extended stretches of duplex DNA…………………………………………………………...…………………….68 4.3.2 CST can bind and unfold G4 structures…………………….……………………….73

4.3.3 CST exhibits facilitated displacement……………………………………………….76

4.4 Discussion……………………………………………………………..……………………..80

4.5 Supplemental figures…………………………………………………………………………83

5. Importance of CST as a Complex………………………………………………...…………….86

5.1 Introduction…………………………………………………………………………………..87

5.2 Results………………………………………………………………………..………………87

5.2.1 ST shows significant reduction in binding affinity compared to CST but not CS…. 87

5.2.2 Absence of the subunits changes the DNA binding architecture of the complex…90

5.3 Discussion………………………………………………………………………………..…...92

6. Discussion and Future Directions…………………………………...………………………….94

6.1 Diverse roles of CST in resolution of replication issues………………………..…………….94

6.2 Delving into Mechanism of action of CST……………………………..…………………….95

6.2.1 Proposed mechanism of CST action at C-strand Fill-in……………………………………97

6.2.2 Mechanism of action of CST in resolution of replication at G-rich DNA……………...…..99

6.3 Proposed model for CST action……………………………………………………..………100

6.4 Does CST need to work as a complex?...... 100

7. References…………………………………………………………………………...………….102

1 Introduction: In contrast to the circular genomes of the prokaryotes and the eukaryotic organelles, the natural ends of nuclear genomic eukaryotic chromosomes are linear and capped with nucleoprotein structures called telomeres. Long before the basic features of DNA and replication were known, telomeres were recognized (Herman Muller, 1946;

Barbara McClintock, 1983) as being essential for the stability of chromosomes. Variations in telomere state underlie cell senescence, stem cell biology, and the development of many diseases like bone marrow failure syndromes, leukemia, and cancer development(2,3). Telomeres protect the Figure 1: The end of chromosomes, chromosomes from breakage and prevent the terminal known as telomeres are composed of double stranded and single stranded DNA from being recognized as a DNA double-strand DNA that forms a t-loop structure. (From Carolyn Price) break. A defective telomeric cap can result in a damage response leading to a cell cycle check point and attempts to repair the chromosome end by non- homologous end joining (NHEJ). NHEJ leads to end-to-end fusion of chromosomes(4). Telomeres are also important because they provide a mechanism to compensate for the inability of DNA polymerases to completely replicate the very end of chromosome, a problem known as the end replication problem (4,5).

1.1 Structure of Telomere DNA: Telomeric DNAs from most eukaryotes and plants are composed of short tandem repeat DNA sequences (TTAGGG in humans). Most of the telomere is double-stranded (2-

20 kb in humans, ~100 kb in rodents), but the 3’ ends are single stranded forming an overhang region (50-

500 nt in humans)(6). In almost all organisms, the 3' strand of the telomere is rich in guanosine. The resulting 3' overhang is known as the G-overhang or the G-tail. Because of the G/C bias, the two strands of the telomeric DNA are also known as the G- and C-strands.(4,7) Electron microscopy revealed that

1 mouse and human telomeres are organized into large lariat-like t-loops (Figure 1) that are presumed to form through strand invasion of the duplex telomeric repeat by the 3’ overhang. The overhang then base pairs with the complementary C-strand replacing the G-strand in that region to form a displacement loop

(D-loop). This structure sequesters the telomere end and facilitates chromosome end protection by hiding the telomere terminus from the DNA damage repair machinery (8,9). It is not clear whether t-loops

persist throughout the cell cycle or require prolonged

resolution during DNA replication. Because of its enriched G

content, the single-stranded telomeric DNA can also form G-

quadruplexes (Figure 2), where each G base serves as both a

donor and an acceptor for hydrogen bond formation leading

to Hoogstein base pairing instead of the classic Watson-Crick

Figure 2: Schematic outline of a G- base pairing(10). A central cation stabilizes the stacking of quadruplex, showing tetrads (left) G-quadruplexes by counteracting the repulsions from the stacking to form an intramolecular structure (right) (From: Julian inwardly facing oxygen molecules. In humans, telomeric G- Huppert) quadruplex structures have been implicated both as a

mechanism for telomere protection and suppression of recombination. They also pose a challenge to telomere replication.

1.2 Replication: DNA replication is the process of producing two identical copies from one original DNA molecule. Most of the telomeric dsDNA is replicated by the conventional replication machinery in a semi-conservative manner where a replication fork moves from a sub-telomeric origin towards the DNA terminus. However telomeres possess intrinsic replication barriers that induce replication fork stalling and/or collapse of the replication machinery. Failure of proper telomere replication can induce genomic instability and eventual cell death or senescence(5).

1.2.1 General eukaryotic replication regulation and replication stress: Regulation and accuracy of eukaryotic replication depends on strict temporal separation of origin licensing and origin firing followed

2 by DNA synthesis. In G1 phase a pre-replication complex is formed by the recruitment of inactive MCM helicase complex, origin recognition complex (ORC), CDC6 and DNA replication licensing factor Cdt1

(11). Upon entry into S phase, several kinases (DDK, CDKs) trigger recruitment of additional proteins to the origins to form a replication initiation complex, which induces association of DNA polymerases and the replicative helicases, followed by initiation of DNA synthesis at bidirectional replication forks. The

MCM complex loading is inhibited after early onset of S-phase to prevent re-initiation and re-replication of the genome. The DNA primase/ polymerase α complex initiates DNA synthesis. DNA polymerase δ continues synthesizing of the lagging strand with formation of Okazaki fragments whereas DNA polymerase ε synthesizes the leading strand in a continuous manner. Bidirectional replication forks continue replicating until they meet another fork/ newly synthesized DNA or they reach chromosomal end. In cells, replication forks often stall and collapse because they encounter varied endogenous and exogenous replication barriers. Endogenous obstacles might include damaged or broken DNA, interstrand DNA crosslinks, DNA secondary structures such as G-quadruplexes, DNA-RNA hybrids forming

R-loops which are difficult to resolve by helicases, exhaustion of dNTPs or proteins necessary for replication such as RPA.

These natural replication barriers are intrinsically recombinogenic and mutagenic and thus may promote Figure 3: End Replication chromosomal instability. Exogenous replication protein problem: Removal of primer at the very end after replication inhibitors and irradiation leading to DNA damage also impede generates a gap, losing some DNA information from the replication. To rescue stalled replication in S-phase, DNA- parental strand. (By Carolyn replication checkpoint pathways co-ordinate cell cycle arrest Price) and rescue of stalled forks by protecting and reactivating the

3 replication machinery or by firing dormant origins.

1.2.2 The end replication problem: The end replication, or the end shortening problem, was first put forward by Olovnikov in 1971. Because all DNA polymerases synthesize DNA only in the 5’3’ direction, the lagging strand is replicated in a discontinuous fashion beginning with synthesis of short

RNA/DNA primers and synthesizing the complementary DNA in installments as the parental strand unwinds, known as Okazaki fragments. Subsequent removal of the RNA from the most distal of these fragments leaves a short gap of 8-12 nucleotides at the chromosome terminus. This gap results in loss of a small amount of DNA at each round of replication (Figure 3). The gap can be even larger if the Okazaki fragment is not placed right at the 3’ end. In the absence of complementary machinery, the resulting loss of DNA with each replication cycle causes the daughter chromosomes to gradually shorten (12,13). An additional cause for recurrent telomere shortening is the exonucleolytic degradation of the leading strand telomere (14). This occurs in order to produce an overhang on both 3' terminal ends. The loss of telomeric

DNA and the DNA binding proteins that form a protective cap structure, leads to activation of a p53/Rb mediated cell cycle checkpoint arrest at G1 followed by senescence (15). Loss of the check point can result in further rounds of replication, more chromosome shortening and eventually end-to-end fusion of chromosomes leading to genome instability.

Telomeric shortening is counteracted by telomerase, a ribonucleoprotein reverse transcriptase.

Telomerase is capable of telomere extension/ elongation by adding new telomeric repeats at the end of the overhang by using its own RNA moiety as a template (16). After the extension of the 3’ end by telomerase, the complimentary C-rich strand can be filled in (C-strand fill-in) by the conventional replication machinery mediated by DNA polymerase α. Although telomerase-mediated repeat synthesis can balance the loss of DNA at chromosome ends, in humans, telomerase is expressed in most tissues only during the first week of embryogenesis. This leads to a gradual telomere shortening over a human life-span. Repression of telomerase in somatic cells is thought to have a powerful tumor suppressive function (17). Short telomeres that accumulate following an excessive number of cell division cycles

4 induce senescence, which counteracts the formation of pre-malignant lesions. This growth barrier is overcome in most cancer cells by the re-expression of telomerase (18,19) which results in cell

immortality.

1.3 Telomerase: Telomerase was discovered in 1985 in the

holotrichous ciliate Tetrahymena thermophila by Carol

Greider and Elizabeth Blackburn (20). Two main components

of this enzyme are the telomerase RNA component (TR) and

the telomerase reverse transcriptase (TERT) (Figure 4). The

Figure 4: Schematic reverse transcriptase activity of TERT utilizes TR as a representation of telomerase components showing telomerase template to elongate the 3’ overhangs of telomeric ends. TR RNA (TR) and TERT. (by Shelley contains a short internal template region that hybridizes to the Schlender) telomeric G-overhang for telomeric repeat synthesis. The

template region in all TRs includes a 5' portion that is copied and a 3' portion that is typically used for the alignment of primer DNA (21). Despite the divergence in sequence and size of TR, some structural features appear to be conserved through evolution and their features are important for the catalytic cycle. The conserved regions are the template, a pseudo-knot and a stem-loop or bulged stem junction with conserved paired and unpaired nucleotides, and a 3’ element required for RNA stability (22). The template of telomerase is single stranded, allowing base pairing with the telomere 3’ end within the active site of TERT. The length of the RNA template of TR is approximately 1.5 to 2 times the telomeric repeat length, thus enabling both annealing of the telomeric 3’ end with the template and addition of one telomeric repeat per elongation. After addition of a single repeat, chromosome ends are repositioned for telomerase to add additional repeats, a process referred as repeat addition processivity (RAP). TERT proteins share sequence homology in their carboxy-terminal half with reverse transcriptases (RT) from retro-elements and retroviruses. All seven defined RT motifs

(1, 2, A, B’, C, D and E) are present in TERT. Phylogenetic analysis suggests that TERT is most related to RTs encoded by non-long terminal repeat (LTR) retro-transposons. A TERT-specific domain adjacent

5 to the RT motifs, the telomeric RNA binding domain (TRBD), is necessary and sufficient for the high biological specificity of TERT-TR interaction (23,24). The four functional domains of TERT are

Telomerase N-terminal domain (TEN), TRBD, RT domain and C-terminal extension (CTE). Human telomerase ribonucleoproteins (RNPs) are enriched in Cajal bodies, nuclear domains of concentrated RNP biogenesis and recycling. In most human cells, Cajal bodies appear to function in telomerase trafficking to the telomere. Cajal bodies are not detectable in all cell types and are disrupted by some forms of cell stress, raising the question of whether TR distribution and its access to telomeres also vary with the state of the cell (25).

Multiple accessory factors associate with the protein and RNA components of telomerase to make up the complete holoenzyme. These factors help in proper folding of TER, TER and TERT assembly as well as intracellular trafficking, recruitment of the ribonucleoprotein complex to chromosomal ends, and activation of the enzyme to elongate telomeres (26). Telomerase and the accessory proteins have undergone rapid evolution in different species. In some organisms, the accessory factors involved in the recruitment and activation are part of the telomerase complex whereas in other organisms they exist as telomere-binding proteins. Although the telomerase holoenzyme is best characterized in yeast and

Tetrahymena, a lot is now known about human telomerase. Telomerase biogenesis: In humans, the 3’ stabilizing element in hTR is an H/ACA motif that co-transcriptionally associates with cofactors dyskerin,

NOP10, NHP2 and the chaperone NAF1, leading to RNA stabilization and nuclear localization. The CAB box domain at the 3’ hairpin of hTR is necessary for its trafficking to Cajal bodies where the NAF1 is replaced by GAR1. Loss or mutation in dyskerin leads to the disease dyskeratosis congenita. hTERT is synthesized in the cytoplasm where association with chaperons HSP90 and p23 takes place. AAA+

ATPases Reptin and Pontin facilitate the association of hTR and hTERT to form the catalytically active enzyme although the exact mechanism and subcellular localization of this process is still under investigation. A crucial interaction between telomerase and TCAB1 is needed for the localization of active telomerase to Cajal bodies. In S phase of the cell cycle, telomerase is recruited to telomeres by interaction with TPP1, a telomere binding protein. In contrast, in budding yeast, telomerase assembly

6 takes place outside nucleus and therefore requires export of TR. The telomerase holoenzyme is then transported back to nucleus by assembling with the Ku 70/80 heterodimer and TLC1. Est3, which is a structural homolog of human TPP1, associates with the preformed Est2/TERT-Est1-TLC1complex at the

G2/M phase. In Tetrahymena, the accessory factors are p65, p50, Teb1 and 7-4-1 complex (p75-p45-p19). p50 stabilizes the interaction of Teb1with telomerase and 7-1-4. Both p50 and Teb1 stabilize interaction with the telomere and are important for RAP and telomerase activity.

Telomerase trafficking: In human cells, telomerase localizes to Cajal bodies by interaction with

TCAB1. During S-phase, telomeres transiently associates with Cajal bodies, potentially to deliver telomerase to telomeres by a hand-off mechanism. Depletion of Cajal bodies leads to defect in telomerase recruitment to telomeres, but this phenotype can be rescues by overexpression of telomerase. A similar phenotype is observed after loss of TCAB1, but interestingly cannot be rescued by telomerase overexpression. In human, TPP1 is the key telomere binding protein for recruitment of telomerase by interaction with the TEN domain of hTERT. In yeast, the telomerase subunit Est1 interacts with Cdc13, a telomerase binding protein that helps in its recruitment to telomeres.

Telomerase activation: In humans, RAP and telomerase activation are regulated by TPP1 in association with POT1, another telomere binding protein. In addition to the positive regulators of telomerase, there are factors that counteract telomerase activity after S-phase. In humans, CST, a telomere binding protein complex has been implicated in terminating telomerase activity.

To prevent critical telomere shortening and hence to achieve unlimited replicative lifespan, most cancer cells reactivate telomerase. However a minority use a different mechanism called alternative lengthening of telomeres (ALT), a recombination-based lengthening of telomeres (27,28). ALT cells exhibit an increased abundance of extra-chromosomal circles of double-stranded telomeric DNA (t- circles), derived from deleterious homologous recombination (HR) events at the T-loop.

1.4 Telomeric replication stress: Telomeres resemble fragile sites, which are observed in genomic regions that challenge replication, especially when replication is stressed by dNTP depletion or DNA polymerase inhibition (29). Replication stress and replication fork stalling leads to appearance of multi- telomeric signals (MTS) in metaphase chromosomes where multiple telomere FISH signals, instead of only one, are seen at each chromosome arm. Although the underlying causes of MTS are not fully understood, they share common features with fragile sites which cytogenetically appear as chromosome gaps and breaks. Telomeres pose problems during replication because they harbor several obstacles such as heterochromatin or telomere compaction, attachment to the nuclear envelop, (T)-loop lariat-like structure. Telomeres transcribe a long noncoding RNA from telomeric repeat that forms integral part of telomere heterochromatin, known as TERRA(telomere repeat containing RNA) (30). TERRA can form

R-loop at telomeres by forming RNA: DNA hybrids which is also one of the endogenous obstacles that impede telomere replication fork progression. Telomeres are guanine (G) rich and thus are capable of forming stable secondary structures such as guanine quadruplex (G4) DNA structures in vitro (Figure2).

In vivo, G4-DNA could form at the single-strand telomeric overhang, the base of the T-loop, or, more generally, during replication, repair, and transcription of telomeric DNA. At telomeres, replication forks appear to occasionally stall due to various above-mentioned forms of replication stress. The lack of downstream replication origins at telomeres makes it harder to rescue the stalled replication forks. Thus if not properly resolved, the replication stalling results in DNA breaks and sporadic loss of chunks of telomeric DNA (12). Several helicases and telomere binding proteins help in resolving all these replication barriers and ensure proper replication of telomeres.

1.5 Telomeric Proteins: Telomeres are bound by multiple proteins that are important for telomere replication and maintenance. These proteins regulate formation of a protective cap structure that prevents

DNA damage signaling, inappropriate recombination and degradation by exonucleases. They are also important for replication of the telomeric double-strand DNA, telomerase recruitment and C-strand fill-in to maintain telomere length. Although telomeres and telomeric proteins from diverse organisms share functional similarity, the amino acid sequence initially suggested that telomeric proteins are very diverse

8 and not evolutionary conserved. However subsequent structural analysis and functional studies have revealed a conservation of key structural domain, indicating that the functions of telomeric proteins are quite conserved even though their sequences have evolved rapidly.

Vertebrate telomeres are bound by a protein complex called shelterin that bridge the double-stranded region to the single-stranded G-overhang (Figure 4) (31). Shelterin is composed of six proteins. The myb domain-containing TRF1 and TRF2 (Telomeric Repeat Binding Factor 1 and 2) that bind the double- stranded DNA, the OB-fold containing protein POT1 (Protection of Telomeres 1) that binds the G- overhang and TIN2 (TRF1 Interacting Nuclear protein 2) and TPP1 which bridges them to link TRF1/2 to

POT1. RAP1 (human ortholog of the yeast Repressor/ Activator Protein 1), another myb domain containing protein binds to the TRF1-TRF2. Shelterin protects natural chromosomal ends from being recognized as DNA double-strand breaks by sequestering the

DNA terminus from the DNA repair machinery. It also regulates telomerase-dependent telomere maintenance. The components of shelterin specifically recognize and bind to the telomeric repeats and are present on telomeres throughout the cell cycle. It has been demonstrated, by its isolation from Figure 4: Schematic nuclear cell extracts, that shelterin can form a stable complex representation of Shelterin components bound to telomeric even in the absence of telomeric DNA (32). Several DNA. (By Carolyn Price) components of shelterin are highly conserved in other eukaryotes. Vertebrate POT1 and TPP1 are orthologs of the telomeric protein TEBPα and TEBPβ found in the ciliate Oxytricha. TPP1 and POT1 are also conserved in Saccharomyces pombe (31). As in vertebrate shelterin, where the TPP1/POT1 dimer is connected to

TRF/RAP1 by TIN2, in S. pombe this connection is mediated by Poz1, which binds both Rap1 and the

TPP1 ortholog Tpz1 (33).

In contrast to vertebrates and S. pombe, in Saccharomyces cerevisiae telomeres have no shelterin-like complex and instead are bound by two separate complexes. Rap1/Rif1/Rif2 complex binds to the double- strand telomeric DNA and Cdc13/Stn1/Ten1 complex binds to the G-overhang (Figure 5) (34).

Interestingly, there is no direct bridging between these complexes as with the vertebrate shelterin complex. Cdc13 binds sequence-specifically to the G-overhang and recruits two other proteins, Stn1 and Ten1, to form a trimeric complex known as the CST complex (1). The CST complex from budding yeast is extremely important for chromosome end-protection and defects in any component of the complex Figure 5: Model for CST in lead to C-strand degradation, accumulation of ss G-rich telomeric DNA replication in budding yeast and vertebrates. S. telomeric DNA, a DNA damage response and late S/G2 cell cerevisiae CST interacts with the Est1 component of telomerase to cycle arrest. Phosphorylation of Cdc13 bound to the G- promote telomeric DNA synthesis overhang leads to interaction with Est1. This interaction on the G-overhang, and with Pol α/primase to facilitate lagging mediates the recruitment of telomerase to chromosome ends strand replication of the C-strand. Vertebrate CST associates with in late S-phase. Stn1 has been shown to down regulate Pol α/primase and stimulates its priming activity. The shelterin telomerase by competing with Est1 for binding to Cdc13. component TPP1 contacts telomerase and is postulated to Following telomerase extension, Cdc13 and Stn1 interact to recruit it to the chromosome coordinate fill-in of the complimentary C-strand. This is terminus. TPP1 may also recruit CST to the telomere via proposed to occur by recruitment of DNA polα/primase (pol interactions with STN1. (From (1))

α) through direct interactions between CST and the Pol1 and

Pol12 subunits of pol α (35). Stn1 interacts with Ten1 through its N-terminus and to Cdc13 and Pol12 (subunit of polα-primase complex) through its C- terminus. Notably, point mutations in either CDC13 or POL1 that reduce the Cdc13-Pol1 interaction, also resulted in telomerase mediated G-strand lengthening. This suggests that while Cdc13 interacts with Pol1, the formation of longer G-strands are prevented by inhibiting telomerase activity. Within the CST

10 complex, Stn1 and Ten1 modulate the function of Cdc13 by facilitating the interaction with pol α. Given the interactions with pol α, CST appears to not only restrict telomerase mediated extension of G-strand but also couple this to the priming of the complimentary C-strand by facilitating DNA polymerase α as well as primase to polymerase switch of DNA polymerase α. Thus in budding yeast, CST is responsible for regulating multiple steps in telomere replication in addition to end protection (36). Although none of the scCST components show any obvious sequence similarity with POT1, TPP1 or any other shelterin component, the DNA binding domain of Cdc13 contains an OB-fold that is structurally similar to the OB- fold present in the DNA binding domain of POT1 (1). Moreover depletion of POT1 or TPP1 from human cells shows a similar telomere uncapping to that seen after removal of the yeast CST complex. This led to the hypothesis that scCdc13 and mammalian POT1 were functional homologs. However, further structural and genetic studies have revealed that CST has a higher resemblance to RPA (Replication

Protein A).

RPA is a heterotrimer (RPA70-RPA32-RPA14) that binds ssDNA through a series of OB-folds in eukaryotic cells. It is the primary single-strand DNA binding protein in eukaryotes and is important for

DNA replication, repair and recombination. RPA70 (Rpa1) contains four OB-folds, three of which binds ssDNA. RPA 32 (Rpa2) contains one OB-fold that binds DNA and a C-terminal winged helix domain, which is important for protein-protein interactions. RPA14 (Rpa3), the smallest member of this complex, also contains an OB-fold that mainly involved in protein interactions (37). Crystal structures have revealed that budding yeast Stn1-Ten1 and Rpa2-Rpa3 complexes have substantial structural similarity in their OB-fold motifs, subunit interaction surfaces and in the Stn1 N- and C-terminal extension regions

(38). In addition to structural similarity, like budding yeast CST, RPA also seems to be involved in the stimulation of DNA polymerases. However, significant differences in the relative orientation of the subunits and in the structure of most of the connecting loop regions indicate that, although CST shares significant structural similarity with RPA, it is tailored to perform a different biological function.

The CST complex is conserved in many organisms. Given that fission yeast are more related to budding yeast than to mammals, it was perplexing that fission yeast seemed lack a CST-like complex.

However, genome analysis of S. pombe then revealed a gene similar to budding yeast STN1 (1).

Subsequent analysis revealed that S. pombe has a Stn1-Ten1 complex present at telomeres although no homolog to Cdc13 has been found. This complex has an essential role in end protection. In Arabidopsis, identification of a STN1 ortholog and evidence of co-localization of the encoded protein with telomeres, gave the initial hint of the presence of CST complex in higher eukaryotes. Additional information about

Arabidopsis CST complex was obtained when a genetic screen for mutants with dysfunctional telomeres uncovered a novel gene, CTC1 (Conserved Telomere maintenance Component 1). Disruption of either of

CTC1 or STN1 led to telomere deprotection phenotypes, which included rapid telomere shortening, a large increase in G-overhang and frequent telomere fusions (39). Subsequent studies indicated that CTC1 and STN1 form a complex with TEN1 and this CST complex is responsible for telomere protection in

Arabidopsis, much like the scCST complex in budding yeast. Although CTC1 has no sequence homology with Cdc13, both contain multiple OB-fold domains. Vertebrate CST complex were identified by database search using PSI-BLAST and HHpred to identify CTC1 and analysis of a putative STN1 ortholog. Mass spectrometry analysis and subsequent characterization lead to the discovery of this trimeric complex. Given the known functions of shelterin it was a surprise to find that mammalian cells contained a second telomere binding protein complex, raising questions about a more complex mechanism of action for vertebrate telomeric proteins.

1.5.1 Vertebrate Shelterin: Individual shelterin components have very specific roles in telomere protection and/ or telomere replication. Both TRF1 and TRF2 bind DNA as homodimers, which improves their affinity for the DNA binding. Dimerization is also responsible for their ability to act as architectural factors by changing the higher-order structure of their DNA substrate (40). TRF1 and TRF2 share a common domain structure consisting of the TRF homology (TRFH) domain. This domain functions as the dimerization domain and it also contains a versatile peptide docking site through which TRF1/2 recruit

12 other proteins to telomeres. TRF1 and TRF2 are both extremely abundant and are thought to cover each telomere with thousands of dimers. Both proteins undergo posttranslational modifications including phosphorylation, sumoylation and/or PARsylation. Although the precise functional significances of these modifications are yet to be elucidated, they can regulate their activation/ interaction with other proteins

(31). Conditional deletion of either of TRF1 or TRF2 destabilizes shelterin, which leads to telomeric deprotection, eventually causing a DNA damage response. However the roles of TRF1 and TRF2 are not identical. TRF1 helps to promote replication through the telomere duplex, possibly by recruiting helicases such as BLM and RTEL (41). Removal of TRF1 leads to replication fork stalling and ultimately leads to defect in packaging of telomere tracts, which can be visualized as multi-telomeric signals (MTS) in a

FISH assay using telomeric probes (42). In contrast, TRF2 maintains the G-overhang and plays a key role in G-overhang generation following DNA replication by recruiting the Apollo nuclease. Removal of

TRF2 leads to rapid G-overhang degradation and telomere fusions via NHEJ. TRF2 can also form t-loop- like structures when provided with a model telomere substrate (43) suggesting it is responsible for t-loop formation. Rap1 is an essential but poorly characterized 1:1 constitutive binding partner of TRF2. Unlike its budding yeast counterpart, RAP1 lacks its own DNA binding domain and is lost upon TRF2 deletion

(44). Deletion of mouse RAP1 leads to increased telomere recombination and in human it seems to function in tandem with TRF2 to prevent NHEJ and chromosome fusions (45). As mentioned earlier,

TIN2, that forms a bridge between the shelterin components that bind the ds- and ss- telomeric DNA by binding to TRF1, TRF2 and TPP1, is essential for stabilization of shelterin (46). Although TRF1 and

TRF2 bind to DNA independently with high affinity, loss of TIN2 leads to significant loss of both the proteins. Both TIN2 and TRF1 interact with the cohesion subunit SA1 and this interaction is important to maintain cohesion between sister chromatids along telomeric region and along the chromosome arms.

Certain TIN2 mutations have been found in patients with short telomeres and the pathology associated with dyskeratosis congenita (47). TIN2 also recruits TPP1 (and therefore POT1) to the complex (48).

POT1 and TPP1 function together by forming a heterodimer that regulates telomerase activity and general access to G-overhang. The N-terminus of TPP1 harbors an OB-fold domain that interacts with and

13 recruits telomerase to telomeres. In contrast, POT1 sequesters the DNA 3’ end to make it inaccessible to telomerase. Thus TPP1 and POT1 have opposing effects on telomerase activity. POT1 binds to telomeric

G-strand DNA via two OB-fold domains. Although TPP1 does not bind DNA directly, it increases the binding affinity of POT1 by 5-10 folds. TPP1 is also required to localize POT1 to telomeres, as POT1 lacks a nuclear localization sequence. POT1-TPP1 prevents binding of RPA to G-overhang and hence activation of an ATR mediated DNA damage response. Although POT1 alone inhibits telomerase, when TPP1 and POT1 form a complex with telomeric DNA, this increases the activity and processivity of the human telomerase core enzyme. Studies show that POT1-TPP1 switches from inhibiting telomerase access to the telomere, as a component of shelterin, to serving as a processivity factor for telomerase during telomere extension (49).

1.5.2 Mammalian CST: The rapid evolution of telomere proteins and limited sequence similarity have made it hard to identify orthologs in different species. Nevertheless, homologs of Arabidopsis CTC1 were identified in many vertebrates by database searches. Subsequently mass spectrometry and bioinformatic analyses confirmed the presence of STN1 and TEN1 along with CTC1 as a complex. These tools revealed genes harboring low sequence identity but higher similarity in predicted secondary structures. Structure predictions also clearly showed presence of OB-folds in all three subunits indicating similarity with RPA.

CTC1, STN1 and TEN1 co-purify from cell extracts and co-localize at telomeres. However, immunolocalization studies indicated that only a fraction of mammalian CTC1 is present at telomeres

(~20%). In fact, depletion of either subunit results in a variety of telomeric and genome-wide defects.

Early insights into the basis of CST function came from a prior biochemical analysis of Pol α-primase published in 1990 (50,51). In this work, two components described as AAFs (Alpha Accessory Factors), were identified during DNA polymerase α purification and shown to enhance Pol α primase activity and association with a DNA template. In 2009, the AAF subunits were sequenced and found to correspond to

CTC1 and STN1. AAF/CTC1-STN1 co-purified with the core enzyme of the polymerase complex and allowed synthesis of long RNA-DNA Okazaki fragments in vitro. Although the Pol α stimulation

14 resembles an increase in the polymerase processivity, AAF/CTC1-STN1 allowed numerous termination- reinitiation events during DNA synthesis by facilitating binding of the polymerase to the template. And thus AAF was referred to as template affinity protein. This activity is DNA polymerase α specific and shows no specificity to any other polymerases tested. This work, along with the structural similarity to

RPA, indicated functions of CST in replication.

Knock-down of CTC1, STN1 or TEN1 was then found to cause an increase in multiple telomeric signals

(MTS) suggesting defects in telomere duplex replication (52,53). Moreover, a CsCl density centrifugation assay, it has been shown that knockdown of STN1 leads to delay in the telomere duplex replication (54).

All these evidences clearly indicate an important role of CST in telomere duplex replication. CST depletion also leads to accumulation of G-overhang ssDNA. This phenotype is also a defect associated with telomere replication and could arise from increased telomerase action or G-overhang processing or decreased C-strand fill-in. STN1 has been shown to interact with TPP1, suggesting possible coordination between CST and shelterin to regulate telomerase activity and promote C-strand fill-in (55). Recent studies indicate that the interaction of CST with TPP1 may help in terminating telomerase activity at telomeres (56). Given the possible association of CST in telomeric C-strand synthesis after the telomerase mediated telomeric G-overhang extension, one attractive model for mammalian CST function is that it facilitates Polα-primase activity to fill-in the C-strand. Analysis of G-overhang structure after

STN1 or TEN1 depletion revealed normal kinetics for telomerase-mediated extension but a delay in subsequent overhang shortening, that resulted from a defect in C-strand fill-in (54). On the other hand,

CTC1 knockout has been shown to mediate both an increase and a decrease in telomere length (57). The increase arises from G -strand elongation indicating its importance in terminating telomerase activity.

Defect in the C-strand fill-in leads to an overall shortening of telomeres emphasizing the importance of this complex in maintaining overall telomere length.

In addition to its role at telomeres, CST has also been shown to have genome-wide roles. As mentioned earlier, only a fraction of CST localizes at telomeres and recent studies have shown that CST binds to GC

15 rich regions or fragile sites under stress (58). CST depletion from cells results in an elevated level of

γH2AX phosphorylations indicating increased DNA damage and formation of anaphase bridges that has been associated with defect in non-telomeric replication (53,59,60). The γH2AX does not colocalize with telomeres and no increases in telomere fusions are observed. This indicates an overall genomic instability caused by the loss of CST rather than just a telomere deprotection. Conditions promoting replication stress such as DNA damage or repetitive DNA sequences may cause the polymerases to uncouple from

MCM helicase and accumulation of ssDNA. CST could act as a recruiting factor of DNA polymerase α to initiate DNA replication under stress. The Price lab has shown that CST helps in dormant or late origin firing under hydroxyurea induced stress to rescue stalled replication (53). Under stress, CST also interacts with Rad51, which could be a possible mechanism for rescuing stalled replication or CST functioning in

HR mediated DNA damage repair (58).

Like yeast and plant CST, mammalian CST also binds to ssDNA and contains multiple predicted OB- folds (59). OB-folds are multifunctional domains that are implicated in recognition and binding to single- stranded DNA and RNA and are also important for protein-protein interactions. The structure of an OB- fold is a simple framework of five-stranded β-barrels connected by loops and helices forming a platform for polymer recognition and tailored in different ways for differential specificity (61). An NMR study of mouse STN1 showed significant structural similarity to RPA2 in the OB-fold domains. Recent crystallographic high-resolution structure of the human STN1-TEN1 complex revealed that hSTN1 consists of an OB domain and tandem C-terminal winged Helix-Turn-Helix motifs while hTEN1 consists of a single OB-fold (62). Contacts between the STN1 and TEN1 OB domains facilitate formation of a complex that is strikingly similar to RPA 2/3 and the yeast Stn1-Ten1 complex. Full length STN1 and

TEN1 form a stable heterodimer via the N-terminal portion of STN1. Contacts between these two proteins are mediated by extensive interactions between the C-terminal α3 helix of STN1 and α2 helix of TEN1 and the β-barrels (part of OB-domains) of the two proteins. Currently no high-resolution structure exists for CTC1 due to problems in protein expression and purification. However, Phyre2 protein threading

16 predicts six OB-fold DNA binding domains (DBD) in CTC1 of which three OB-folds in the C-terminal shows close structural similarity with RPA whereas the two OB-folds close to the N-terminal are more like those of POT1. Multiple studies have shown CST binds to a three repeat G-rich telomeric sequence

(18 nt) stably with nano-molar affinity and with sigh stability whereas the binding to the C-strand telomeric sequence is very unstable. STN1-TEN1 alone binds to the telomeric sequence with micro-molar affinity. In vitro binding studies indicate CST can bind to both telomeric and non-telomeric sequences with a preference for G-rich sequences. Binding of CST to oligonucleotides devoid of guanine, e.g. dT or

C-strand of telomeres is unstable. Although the sequence specificity of CST is not well defined, its preference towards G-rich sequences indicate possible of role of CST in resolving the DNA secondary structures formed in G-rich DNA such as G-quadruplexes.

1.6 Telomeres and Disease: Telomere dysfunction has long been associated with diseases, including cancer, bone marrow failure, dyskeratosis congenita, pulmonary fibrosis and also aging (2,3).

1.6.1 Telomere and Cancer: Despite being highly efficient, the cellular replication machinery cannot fully replicate each chromosomal end. In human cells that lack telomerase, this leads to a loss of 50-200 bps of telomeric DNA at each human somatic cell division. Eventually the shortened telomeres block further cell proliferation and the cell enters replicative senescence. Thus, telomere length works as an intracellular timer that can limit cell replication and the number of mitotic cell cycles. The telomere shortening works as a protection against cancer, because after 50 populations of cell doubling sufficient

DNA is lost to trigger a p53-mediated permanent cell cycle arrest in G1. However, if the cell can bypass the cell cycle checkpoint arrest by inactivation of p53 or Rb, the cell keeps on dividing leading to a crisis stage with severe attrition of telomeres. In some cases, cells can activate telomerase to maintain telomere length at this stage and the cells become immortal, leading to cancer. Carcinogenesis is a multistep process where the tumor cells acquire various genetic and molecular abnormalities, hallmarks of the carcinogenic selection process. One of the major features is cell immortality, involving telomere length stabilization. In the majority of cancer cells (~80%), this is achieved by activation of telomerase, while in

17 some others homologous recombination-based ALT is activated. In somatic cells hTER is constitutively active and so activation of telomerase requires re-expression of the hTERT gene. Several constitutive

TERT mutations have been associated with development of several cancers including myeloid leukemia, glioma, and renal carcinoma.

In the cells without telomerase activation, the telomere shortening causes loss of function. The critically shortened telomeres lose the protective cap and thus lose function in chromosome protection. Uncapped telomeres accumulate DNA damage signals leading to NHEJ mediated chromosomal fusions. Increased amounts of chromosomal breaks take place leading to chromosome imbalance, non-reciprocal translocation, altered gene expression and overall genomic instability, which all are hallmarks of cancer cells. Although telomere shortening leads to cancer suppression by the senescence mechanism, critically short telomeres promoting genomic instability can contribute to the enhanced chances of carcinogenesis and development of malignancy. Thus, telomeres act as both tumor suppressor and tumor promoting factors. A significant proportion of patients in colorectal cancer and esophageal cancer have been reported with short telomeres and chromosomal instability. Short telomere in leukocyte has been implicated as a risk factor and is considered as a biomarker for many solid tumors. Shelterin components that has direct role in telomere maintenance such as TRF2, POT1 are also getting increased attention in correlation to development of cancer. In fact, several somatic mutations in the OB-fold domain of POT1 have been associated with chronic lymphocytic leukemia (CLL).

1.6.2 Diseases associated with short telomeres: Telomeres naturally vary in length in an inherited way.

But some individuals have abnormally short telomeres that eventually may lead into a stem cell failure phenotype. The stem cell failure becomes evident in highly proliferative tissues such as the bone marrow and intestinal tract where the replicative potential of stem cells is critical for homeostasis. Short telomeres in early childhood or even in young adults pose higher severity of diseases. The most common manifestation is bone marrow failure. Affected individuals may develop intestinal villous atrophy, immunodeficiency and infertility with a varying severity. The first disorder to be linked to telomerase

18 mutations and short telomeres was Dyskeratosis congenita (DC), which is characterized by dystrophic nails, oral leukoplakia and patchy skin hyperpigmentation. It is a progressive disease where severity increases with age. Mucocutaneous defects are detected in infants, followed by bone marrow failure in the first or second decade. Development of aplastic anemia or involvement of other organs, such as the pulmonary systems can be lethal. Pedigree analysis led to the association of X-linked mutations in DKC1, a gene that encodes dyskerin, a small RNA binding protein that binds to ribosomal RNAs and TER. Very short telomeres have been found in all patients and mutations in TER, TERT, NOP10 and NHP2 have also been associated with DC. Shorter telomeres are linked with higher severity of the disease. A particularly severe variant of DC is Hoyerraal-Hridarsson syndrome (HHS), a disease that shows progressive pancytopenia, various neurological manifestations, ataxia and growth retardation.

The recent discovery of bone marrow failure and telomere defects in patients with an autosomal recessive disorder known as Coats plus (CP) provides yet another instance of disruption of telomere homeostasis leading to human disease pathology. Although CP shares some overlapping phenotypes with DC patients such as nail dystrophy, anemia or hair greying, patients develop more severe phenotypes than DC. These include retinal telangiectasia, intracranial calcification, gastrointestinal bleeding and osteopenia. CP patients possess very short telomeres. Several mutations in CTC1 and STN1 have been associated with

Coats plus. Knowing the various functions of CST it can be predicted that not only disruption in telomere homeostasis but the down-regulation of genome-wide functions of CST might contribute significantly to the severity of this disease. Many Coats plus patients showing overlapping phenotypes with dyskeratosis congenita carry bi-allelic CTC1 mutations, which include both point mutations and more severe mutations that presumably lead to a complete loss of function (63,64). Such CTC1 mutations are clearly highly deleterious as patients tend to die by the age of 30.

1.7 Perspectives and conclusions: It is now widely accepted that telomere length acts as an intracellular timer, limiting cell replication and cell proliferation. Critically short or unprotected telomeres are recognized as double strand breaks and activate the H2AX–ATM–Chk2 pathway, inducing subsequent

19 senescence via P53/Rb surveillance. Overall, these observations highlight the need for improved understanding of telomere-driven senescence and of the mechanisms involved when this process fails to protect against cancer progression in the early stages of disease onset, or prevent bone marrow failures.

Both telomerase and telomeric proteins are clearly essential to maintain telomeres and hence, chromosomes in a fully functional state. Therefore, it is important to understand the underlying mechanisms of their action and the detailed interactions among them.

1.8 Dissertation Goals:

Overall I characterized DNA binding properties of CST that are relevant to its diverse functions in replication at telomeres as well as genome-wide. There are three major goals of my dissertation work.

First is to study the role and the importance of the OB-fold domain of STN1 by introducing three putative mutations affecting DNA binding. Second aim is to investigate if DNA binding of CST is dynamic similar to RPA and its interaction with various DNA secondary structures. Third goal is to investigate importance of individual components of CST in DNA binding. Overall the work in this dissertation is our attempt to take a mechanistic approach to understand roles of CST in diverse steps of replication both at telomeres and genome-wide.

2. Materials and Methods:

2.1 Generation of STN1-OB cells and verification of cell lines

HeLa 1.2.11 STN1 knockdown (clone shSTN1-1), shSTN1 rescue (STN1-Res), control non-target (clone shNT-2) and TEN1 overexpressing cell lines were as described previously (53,54,65). Cells were cultured in RPMI-1640 with 10% FBS, antibiotics and glutamine. The STN1 OB fold mutant (STN1-OBM) was made by PCR-mediated site-directed mutagenesis of STN1 cDNA using the following primers (mutations are underlined):

W89A, 5’-TGCATCTGCGCCAAAAAGTTGAATACTGAGTCTGTATCAGC and

5’-CAACTTTTTGGCGCAGATGCAGTTTATAACTCCAGTGC;

R139L and Y141A, 5’-TCCTCACAGCCAGAGAAGAGCGAGAGATTCATGCCACC and 5’-

CTCTTCTCTGGCTGTGAGGATACTGCCTCTGACTCGG. The resulting sh-RNA resistant STN1-

OBM gene was cloned into pMSCV-IThy1-1 retroviral vector, upstream of IRES and the gene encoding the Thy1-1 cell surface protein. Retrovirus was produced and used to infect HeLa shSTN1-7 cells. Cells were harvested 48 hours after infection, incubated with APC-conjugated anti-Thy1-1 antibody (BD-

Pharmingen) and sorted by FACS to isolate Thy1-1/STN1 expressing cells. Pools of sorted cells were expanded and tested for STN1 expression by Western blot. A PCR-based assay was used to monitor the integrity of the STN1-OBM and STN1-Res cell lines because the two cDNAs only differed in sequence at the site of the STN1-OBM point mutations. To avoid amplification and sequencing of the endogenous gene locus, genomic DNA was amplified with primers directed to the FLAG tag and the junction between exons 6 and 7 of the STN1 cDNA (5’-AGCTGGTACCATGGATTATAAAGATGATG

ATGATAAACAGCCTGGATCCAGCCG-3’ and 5’-CAGGGCGCCTGGATTGCT-3’). The products were then sequenced to verify the presence or absence of the mutant allele. The sequencing primer hybridized to the junction between exons 2 and 3 (5’-GCCAGGTGCCAGGTGTAT-3’). The shSTN1 cells were monitored regularly for STN1 depletion by Western blotting with antibody to STN1 and all the modified cells were re-sorted periodically to maintain expression.

2.2 Western blots: Proteins were separated by SDS-PAGE and transferred to nitrocellulose membrane.

The membrane was blocked with 5% milk and incubated with antibodies to STN1 (1:2,000) made by immunizing rabbits with full-length purified STN1 (65,66), Actinin (1:10,000) from Santa Cruz (sc-

17829), HA (1:2000) from Cell signaling (3724S), FLAG (1:2000) rabbit from Sigma (F7425), purified

TEN1 antibody (1:1000) (66), DNA polα (1:1000) goat polyclonal from Santa Cruz (sc-5920), Goat-α-

Mouse-HRP from Thermo Scientific (32430), Goat-α-Rabbit-HRP from Thermo Scientific (32460) and

Donkey- α-Goat-HRP from Santa Cruz (sc-2020).

2.3 Anaphase bridge analysis: HeLa cells were grown on coverslips overnight prior to addition of nocodazole (50 ng/ml). After 4 hours the cells were released into fresh media for 45-60 min then fixed in

3% formaldehyde. The coverslips were washed with PBS, dehydrated with cold ethanol (70%, 90%,

100%) and mounted with fluoro-gel (Electron Microscopy Sciences) and 0.2 μg/ml DAPI. The slides were viewed under 100x with a Nikon Eclipse E400 fluorescent microscope equipped with a Spot 2 digital camera (Diagnostic instruments Inc.). The number of anaphase cells with bridges out of 200 total anaphase cells was scored for each sample.

2.4 Telomere FISH: HeLa cells were grown overnight to 40-50% confluency. Colcemid (0.5 μg/ml) was added for 1.5 hour, the cells were harvested, fixed in methanol acetic acid and used to make metaphase spreads. Telomere FISH was performed essentially as described (53), using FITC-(TTAGGG)3 probe

(Biosynthesis). To amplify the FISH signal the slides were blocked with PBG (0.5% BSA and 0.2% cold water fish gelatin in 1x PBS) for 20 min after the final hybridization step. Slides were then incubated with

6 μg/ml biotinylated anti-fluorescein for 1-2 hour at room temperature followed by 16 μg/ml fluorescein avidin (Vector laboratories) for 1 hour at 37C in a humidified chamber. The slides were washed with

PBS, dehydrated with ethanol (70%,90%,100%) and mounted with Fluro-gel (Electron Microscopy

Sciences) with 0.5 μg/ml DAPI. Telomere FISH images were taken at 100x. MTS (Multi-telomere signals) were scored blindly. At least 200 chromosomes were analyzed per independent experiment.

2.5 Genomic DNA isolation: ~5x106 cells were washed with PBS, lysed in 1 ml nuclei lysis buffer

(Promega Wizard kit) and treated with RNAse (10 ng/ml) for 30 min at 37C for 30 min. Samples were then digested with Proteinase K (10 ng/ml) for 4-5 hours at 37C, cooled on ice for 15 min prior to addition of 350 µl protein precipitation buffer (Promega Wizard kit). Samples were incubated on ice for a further 15 min then centrifuged at 13,000 rpm for 30 min. DNA was precipitated with isopropanol and re- suspended in TE buffer (10mM Tris-HCL pH8.0, 1mM EDTA).

2.6 Cell cycle synchronization and G-overhang analysis: Cells were synchronized at the G1/S boundary by double thymidine block as previously described (53). Synchrony was monitored by FACS analysis of DNA content. Cells were released into fresh media after the second thymidine treatment and harvested 0, 6, 8, 10 and 12 hours later for DNA isolation. To analyze G-overhang abundance, samples were digested or mock-digested overnight with Exo1, followed by restriction digestion with HinfI and

MspI and separated briefly in 1% agarose gels to keep the telomeric restriction fragments in a tight band.

Gels were dried and hybridized with (TA2C3)4 probe under non-denaturing conditions. The DNA was then denatured, and the gel was rehybridized with the same probe. Bands were quantified by PhosphorImager using Image Quant software. For each sample, band intensity from the native gel was normalized to that of the denatured gel to control for differences in loading. To assess changes in overhang abundance rather than internal ssDNA, the normalized band intensity of the Exo1 digested sample was subtracted from the mock digested counterpart.

2.7 MTT assay: Cell viability and proliferation were monitored using the tetrazolium based MTT colorimetric assay (67). Cells were grown overnight in 24 well plates to 60-70% confluency.

The culture medium was then replaced with DMEM containing 1 mg/ml MTT and left for 40 min at

37oC. The medium was removed and cells were washed with PBS 3 times. DMSO was added to dissolve the formazan crystals and left for 15 min with shaking at room temperature. The reaction intensity was measured with a multi-well scanning spectrophotometer (Synergy MX, BioTek) at 570nm in triplicates in a 96-well plate.

2.8 DNA-fiber analysis: Cells were labeled with 50 uM IdU for 15 mins. Control cells were then labeled with 100 um CldU for 20 mins. The remaining cells were treated for 2 mM Hu for 2 hr. The HU was then removed and the cells labeled with 100 m CldU for 60 min. Cells were harvested, lysed and spread on slides by hydrodynamic flow as described (68). Slides were incubated with mouse α‐BrdU (1:500, Becton

Dickson) and rat α‐BrdU (1:500, Accurate Chemical) to detect IdU and CldU respectively. The signal was then amplified and visualized by incubation with AlexaFluor 594 rabbit α‐mouse (1:1000) and

AlexaFluor 488 chicken α‐rat (1:750) (Invitrogen) secondary antibodies and AlexaFluor 594 goat α‐rabbit

(1:1000) and AlexaFluor 488 (1:750) tertiary antibodies. Confocal images of DNA fibers were acquired using a Zeiss LSM710 microscope. The fibers were scored using previously described software (54).

2.9 Co-immunoprecipitation: To detect TPP1 interaction with STN1, HEK293T cells were transfected with mCherry-HA-TPP1 (69) and FLAG-STN1 (WT or OBM) expression constructs. To detect interaction between CST and DNA polα, cells were transfected with Flag-CTC1, Flag-STN1 (WT or

OBM) and TEN1 expression pcDNA constructs. Cells were extracted 72 hours later with 20 mM Tris pH

8.0, 100 mM NaCl, 1 mM MgCl2, 0.1% NP-40 (Igepal). TPP1 was precipitated with HA antibody, STN1 and CTC1 were precipitated with FLAG beads.

2.10 Isolation of Bacmid DNA: To construct bacmid DNA, the pFastBac1 DNAs were transformed into competent DH10Bac E. coli cells. DH10Bac cells contain a bacmid DNA that has a lacZ expression system and a mini-attTn7 transposition site which allows integration of the Tn7R/Tn7L sites found on the pFastBac1 plasmid as well as a helper plasmid which contains genes necessary for transposition of the target gene into the bacmid DNA. Transformed bacteria were selected on LB plates with 50 µg/ml kanamycin, 10 µg/ ml tetracycline, and 7 µg/ml gentamycin for 48 hours. In the presence of 40 µg/ml

IPTG and 100 µg/ml X-gal, the colonies on the plate that were white, indicated correct integration of the gene construct as integration at the attTn7 transposition site interrupts lacZ expression on the bacmid

DNA and thereby no blue coloration. For a second step purity screening, to ensure zero contamination in the picked colonies, at least four apparent white colonies were picked and streaked on a similar LB plate

24 as before for 48 hours. Next day, two correct colonies were picked and grown in 3ml LB with 50 µg/mL kanamycin, 10 µg/mL tetracycline, and 7 µg/mL gentamycin for 16 hours in the shaker @ 37°C. The bacmid DNA was isolated using the ZR BAC DNA mini-prep Kit from ZYMO Research (D4048). To check the correct insert, PCR was performed using M13F (5' TGTAAAACGACGGCCAGT 3’) and

M13R (5’CAGGAAACAGCTATGAC 3’) primers.

2.11 Sf9 Cell Culture: Sf9 cells were grown in suspension culture @27°C at 130rpm in SF900II serum free media (Invitrogen) at a concentration of 1-3 x 106cells/ml. For initial virus production (P0) and viral titering, cells were allowed to adhere to plates at about 60% confluence and cultured at 27°C.

2.12 Transfection of SF9 cells with Bacmid DNA for virus production: 9 x 105 Sf9 cells from the suspension culture were seeded in a 6-well plate and allowed to attach for 1 hour at 27°C with 150 µg/ ml

Penicillin/ Streptomycin (Penn/Strep). Each of 1 µg of bacmid DNA and 6 µl of Cellfectin transfection reagent (Life Technologies) was diluted in 100 µL of unsupplemented Grace's Medium from (Gibco).

The diluted cellfectin was added to the bacmid DNA followed by 30 minutes incubation at room temperature. The cells were washed with 2 mL of unsupplemented Grace's Medium. 800 µL of unsupplemented Grace's Medium was added to the DNA: lipid complexes that contained the bacmid

DNA and the Cellfectin, and the total mixture were added to the cells. The cells were incubated for 5 hours at 27°C before the transfection mixture was removed and 2 mL of SF900II media with 150 µg/ ml

Penn/Strep was added to the cells. The cells were then allowed to incubate for ~72 hours before the first viral stock was collected.

2.13 Baculo-virus amplification: 72 hours after the transfection of SF9 cells with the bacmid DNA, the cells were monitored every hour for signs of viral infection. When the cell viability began to drop to about

80%, the media was collected containing the first viral stock (P1). The media was centrifuged at 500 x g for 5 minutes to pellet down cells and other debris. The supernatant containing the virus was collected and stored at 4°C covered with foil to protect the media from light from one well. Virus from the other wells,

25 were flash frozen and stored at -80°C for future use. To amplify the viral stock and obtain higher titer viral stocks (P2 and subsequent P3, P4), SF9 cells were grown as a suspension culture in 100 ml SF900II media with a density of 1-2 x 106 cells/mL. The P1 stock was assumed to have a titer of 5 x 106 pfu/ml.

Viral stock was added at an MOI of 0.1 pfu/cell and the cells were allowed to grow and incubate until they again reached a cell viability of ~80% in about 72 hours. This P2 viral stock was then collected and stored in the same way as the P1 stock. A P3 viral stock was made in the same way as the P2 stock but assuming a titer of 5 x 107 pfu/ml. After production of viral stocks, the P3 stock titer was determined by infecting EZ Titer cells with a serial dilution of virus. These modified insect cells contain a GFP expression construct under control of the polyhedron promoter, so that GFP is expressed only in cells that are infected with virus. These cells were plated and virus was added in a serial dilution, and allowed to infect for 72 hours before the cells were viewed under the inverted fluorescence microscope and GFP- expressing cells were counted to determine the approximate viral titer.

2.14 CST Protein expression in insect cells: For protein expression, SF9 cells were grown to 1-5 x 106 cells/ ml in a suspension cultures same as before. They were co-infected with CTC1, STN1 (WT or

OBM) and TEN1 viruses at an MOI of 1-2 and allowed to grow at 30°C and 130 rpm for between 52-60 hours. When the cell viability was between 80-85% the cells were collected by centrifugation at 200 x g for 5 minutes. The cells were washed in 1X PBS and the pellet was then flash frozen in liquid nitrogen and stored at -80°C prior to purification.

2.15 CST expression and purification from insect cells: SF9 cells were co-infected with the baculovirus encoding FLAG-CTC1, His-STN1/STN1-OBM or TEN1 (all tags were N-terminal). Infected cells were lysed in 25 mM Tris-HCL pH 7.5, 500 mM NaCl, 0.5% NP-40, 1 mM PMSF and protease inhibitor cocktail. The supernatant was supplemented with 1 mM DTT and 25 mM imidazole and applied to nickel-sepharose beads (GE healthcare 17-5268-01). The beads were washed twice with 25 mM Tris pH 7.5, 500 mM NaCl, 0.5% NP40, 25 mM imidazole, 1 mM DTT and once with 25 mM Tris pH 7.5,

500 mM NaCl, 25 mM imidazole, 1mM DTT. The protein was eluted with 25 mM Tris 7.5, 500 mM

NaCl, 100 mM imidazole, 10% glycerol, 1 mM DTT and then diluted 1:4 with 25 mM Tris pH 7.5, 175 mM NaCl, 10% glycerol, 1 mM DTT to bring the imidazole to 25 mM and NaCl to 300 mM. FLAG beads (Sigma, A2220) were added to the diluted protein and incubated for 1 hr at 4°C. Beads were washed with 25 mM Tris pH 7.5, 200 mM NaCl, 10% glycerol, 1mM DTT. The protein was eluted with

3X FLAG peptide and stored either at 4°C or at -80°C (with 0.1 mg/ml BSA and 15% glycerol) after flash freezing in liquid nitrogen. The concentration was determined by PAGE and silver staining using a BSA standard.

2.16 CST expression and purification from human cells: HEK293T cells were co-transfected with pMIG3-Flag-Strep-CTC1, pcDNA-Flag-STN1 or pcDNA-Myc-STN1 and pMIT-TEN1. Cells were lysed with lysis buffer (25mM Tris-HCl pH7.5, 300mM NaCl, 1mM PMSF, 0.5x protease inhibitor cocktail).

Lysate was adjusted to 1mM DTT before adding Flag beads (Sigma A2220) followed by 1 hour incubation at 4°C. Beads were washed with wash buffer (25mM Tris pH7.5, 300mM NaCl, 10% glycerol,

1mM DTT) and protein was eluted from the beads with 3XFlag peptide. The protein was stored either at

4°C or at -80°C (with 0.1 mg/ml BSA and 15% glycerol) after flash freezing in liquid nitrogen. The concentration was determined by PAGE and silver staining using a BSA standard.

2.17 Preparation of DNA constructs: Oligonucleotides used for electrophoretic mobility shift assays

(EMSA) were radiolabeled with 32P-ATP (Parkin Elmer BLU002Z250). Single-stranded and fold back oligonucleotides were boiled for 2 min and fast cooled on ice for 15 min in STE buffer (20 mM Tris-HCl pH 8.0, 100 mM NaCl, 1 mM EDTA). To prepare partial duplex substrates, the long and short single- stranded oligonucleotides were mixed in a 1:1.1 molar ratio (5 µM long oligonucleotide) in STE buffer, boiled for 2 min and slow cooled to room temperature 2 hrs. Partial duplex substrates for smFRET assays were prepared by mixing Cy3 and Cy5-labelled oligonucleotides (IDT) in a 1:1.5 molar ratio in

T50 buffer (10 mM Tris-HCL pH 8.0, 50 mM NaCl), boiled for 2 min and slow cooled to room temperature over 2 hrs. To ensure formation of G4, substrates (ssDNA for EMSA and partial duplex for smFRET) were further diluted to desired concentrations (1fmol/ul for EMSA and 5 pM for smFRET) in

G4 buffer (20 mM Tris-HCl pH 8.0, 3 mM MgCl2, 150 mM NaCl (G4 buffer Na) or 100 mM KCl (G4 buffer K)) and incubated for 10 min before addition to the reaction mix for EMSA or immobilization on slides for smFRET.

2.18 Electrophoretic mobility shift assays and UV crosslinking. CST(WT) or CST(STN1-OBM) (0.5-

20 nM) was incubated with 32P-labeled oligonucleotide (0.1 nM) in 25 mM Tris pH 8.0, 1 mM DTT, 150 mM NaCl for 30 min at RT. For competition assays, binding was for 1 hour at 4°C, cold competitor was then added and samples were incubated for a further 16 hours. Samples were separated in 0.7% agarose gels with 1xTAE at 100V for 1 hr. Gels were dried on Hybond XL membrane and quantified by

PhosphorImager. For UV cross-linking 0.1 nM CST was incubated with 0.1 nm oligonucleotide for 30 min and then subject to cross-linking in a Stratagene Hybridization Oven for 30 min on ice at full power

(7200mJ/cm2) using 6 UVA bulbs.

2.19 Filter Binding Assay to determine Kd,app and t½: Double-filter binding assays were performed as described (70,71) using a 72-well minifold vacuum manifold slot blot apparatus and nitrocellulose and

HyBond XL filters (GE Healthcare). Phosphorimager screens were scanned on a TyphoonTrio phosphorimager (GE Healthcare) and the amount of bound versus free DNA was quantified using

ImageQuantTL software. Graphpad prism software was used to for plotting and curve fitting (one site specific saturation binding equation for Kd and one phase exponential decay equation for t½). To determine Kd,app, CST (0.01-20 nM) was incubated with 32P-labeled oligonucleotide (0.01 nM) in binding buffer (25 mM Tris pH 8.0, 1 mM DTT, 150 mM NaCl) for 18 hours at 4°C to reach reaction equilibrium. To determine the dissociation rate (t½), 20 nM CST was incubated with 0.1 nM 32P-labeled oligonucleotide for 1 hour at 4°C. 0.5 µM of the corresponding unlabeled oligonucleotide was then added.

Samples were analyzed by filter binding assay after 0, 30, 60, 180, 360, 540 or 1080 min incubation with cold competitor.

2.20 Strand melting assay: 20 nM CST were incubated with 0.1 nM 32P-labeled ds-oligonucleotide in 25 mM Tris-HCl pH 8.0, 1 mM DTT, 150 mM NaCl for 30 minutes at RT. To monitor strand displacement, binding reactions were terminated by addition of 6 × helicase dye (30% glycerol, 50 mM EDTA, 0.9%

SDS, 0.25% bromophenol blue, and 0.25% xylene cyanol) that denatures the protein. Reactions were then loaded on 15% native polyacrylamide gel in 1× TBE and run at 200 V for 1 hour. To ensure that melted duplex could not re-anneal after reaction termination, a 500x molar excess of unlabeled short (22 nt) oligonucleotide was added to control reactions in addition to the 6 x helicase buffers. Other samples were boiled in the helicase buffer prior to gel loading and monitored for duplex formation (control).

2.21 Single molecule fluorescence resonance energy transfer (smFRET) assays:

Single molecule fluorescence assays were carried out as described before (72). Briefly, quartz slides and coverslips were treated with methanol, sonicated in acetone and then in potassium hydroxide, burned each side using a propane torch (to burn away any florescent molecule), incubated in amino silane solution

(150 ml of methanol, 7.5 ml of acetic acid and 1.5 ml of amino silane), and coated with a mixture of 97% mPEG (to minimize surface interaction with the protein) and 3% biotin PEG. After preparation of PEG slides, the flow chamber was assembled using strips of double-sided tape and epoxy(73). We flowed in 30

μl of 0.2 mg/ml NeutrAvidin in T50 into each empty flow channel and for incubation. Then, 5 minutes later, the excess NeutrAvidin was washed out with T50. Partial duplex DNA molecules were immobilized on the slides by biotin-neutravidin interaction. Excess oligos were washed away either with T50 buffer or with G4 buffer. To detect binding/ melting of G4, 2nM CST in binding buffer (25mM Tris-HCl pH 8.0,

1mM DTT, 150mM NaCl) were added and incubated for 10 minutes at RT. Excess CST were washed away with T50 buffer and to avoid rapid photobleaching of fluorescent dyes, the channels were supplemented with an image buffer with an oxygen scavenging system (Binding buffer + 0.8 mg/ml glucose oxidase, 0.625% glucose, 3mM Trolox and 0.03mg/ml catalase) before taking images. For real time images, 2nM CST in image buffer were added to the oligo bound channels and imaging was initiated immediately. For the real time images with different concentrations of CST, 2nM CST in image buffer

29 were added to the channels and incubated for 1 minute followed by either washout of excess CST with image buffer of addition of 5nM CST in image buffer before prompt initiation of imaging. Whole procedure was carried out at room temperature.

2.22 smFRET Data Acquisition.

Prism-type total internal reflection fluorescence (TIRF) microscopy was used to acquire single-molecule

FRET. The excitation beam was focused into a pellin broca prism (Altos Photonics), which was placed on top of a quartz slide with a thin layer of immersion oil in between to match the index of refraction. Cy3

(donor) and Cy5 (acceptor) dyes were excited through the dual-laser excitation system (532 and 640 nm,

CrystaLaser) via TIRF. The fluorescence signals from Cy3 and Cy5 that were collected by a water immersion objective lens (60×, 1.2 N.A. Nikon) and then passed through a notch filter to block out excitation beams. The emission signals of Cy5 dyes were separated by a dichroic mirror (FF662-FDi01;

Semrock) and detected by the electron-multiplying charge-coupled device camera (iXon 897; Andor

Technology). Data were recorded with a time resolution of 100 ms as a stream of imaging frames and analyzed with scripts written in interactive data language to give fluorescence intensity time trajectories of individual molecules.

2.23 smFRET Data Analysis

Basic data analysis was carried out by the smCamera software written in C++ (Microsoft). with FRET efficiency, E, calculated as the intensity of the acceptor channel divided by the sum of the donor and acceptor intensities. Leakages from the donor channel to the acceptor channel and vice versa were corrected. FRET histograms were generated by using over 4,000 molecules and were fitted to Gaussian distributions with an unrestrained peak center position in Prism 7 (GraphPad Software). Traces with or without CST fall off were collected from more than 2000 traces for each individual experiment. Dwell times were collected by measuring the time the molecule spends in a particular FRET state. The dwell-

30 time histograms were generated from more than 300 dynamic events. All of the experiments were conducted at least three separate repeats.

3. STN1 OB Fold Mutation Alters DNA Binding and Affects Selective Aspects of CST Function

3.1 ABSTRACT

Mammalian CST (CTC1-STN1-TEN1) participates in multiple aspects of telomere replication and genome-wide recovery from replication stress. CST resembles Replication Protein A (RPA) in that it binds ssDNA and STN1 and TEN1 are structurally similar to RPA2 and RPA3. Conservation between

CTC1 and RPA1 is less apparent. Currently the mechanism underlying CST action is largely unknown.

Here we address CST mechanism by using a DNA-binding mutant, (STN1 OB-fold mutant, STN1-OBM) to examine the relationship between DNA binding and CST function. In vivo, STN1-OBM affects resolution of endogenous replication stress and telomere duplex replication but telomeric C-strand fill-in and new origin firing after exogenous replication stress are unaffected. These selective effects indicate mechanistic differences in CST action during resolution of different replication problems. In vitro binding studies show that STN1 directly engages both short and long ssDNA oligonucleotides, however STN1-

OBM preferentially destabilizes binding to short substrates. The finding that STN1-OBM affects binding to only certain substrates starts to explain the in vivo separation of function observed in STN1-OBM expressing cells. CST is expected to engage DNA substrates of varied length and structure as it acts to resolve different replication problems. Since STN1-OBM will alter CST binding to only some of these substrates, the mutant should affect resolution of only a subset of replication problems, as was observed in the STN1-OBM cells. The in vitro studies also provide insight into CST binding mechanism. Like

RPA, CST likely contacts DNA via multiple OB folds. However, the importance of STN1 for binding short substrates indicates differences in the architecture of CST and RPA DNA-protein complexes. Based on our results, we propose a dynamic DNA binding model that provides a general mechanism for CST action at diverse forms of replication stress.

3.2 Author Summary

Mammalian CST (CTC1/STN1/TEN1) is a three protein complex that aids in several steps during telomere replication and has genome-wide roles during recovery from replication fork stalling. Loss of

CST leads to abnormalities in telomere structure, genomic instability and defects in chromosome segregation. Currently, we do not understand how CST acts to ensure the resolution of very diverse types of replication problem. We set out to address this question by studying a mutant form of CST that was predicted to alter DNA binding. The mutations are in the STN1 subunit. In vivo, the STN1 mutant (STN1-

OBM) affects some aspects of CST function while others are normal. The effects of STN1-OBM do not align with the telomeric versus non-telomeric roles of CST but instead separate out different aspects of

CST function at telomeres and genome-wide. In vitro binding studies indicate that STN1-OBM disrupts binding to only short DNA substrates. Since CST is likely to encounter DNA substrates of varied length and structure in vivo as it helps resolve different replication problems, this finding starts to explain why

STN1-OBM affects only certain aspects of CST function. Our in vitro binding studies also shed light on how CST actually binds to DNA and they suggest a novel “dynamic binding model” that provides a mechanistic explanation for how CST helps resolve a diverse array of replication problems to preserve genome stability.

3.3 INTRODUCTION

Although DNA replication must occur rapidly and with high fidelity, the replisome frequently encounters obstacles such as DNA damage or repetitive sequence that cause the replication fork to stall.

Since stalled forks can lead to double strand breaks and genomic instability, multiple pathways exist to ensure their resolution (74,75). Telomeres pose a particular challenge to the replication machinery due to their repetitive G-rich sequence and the inability of DNA polymerase to completely replicate the DNA 5’ terminus (76-78). To ensure telomeres are duplicated efficiently, the replication process occurs in several distinct steps (7,76,79) and involves a number of ancillary proteins (80-83). First, the repetitive dsDNA

33 is duplicated by the replisome with assistance from various accessory factors. Next, the chromosome ends are processed by nucleases to form a single-stranded overhang on the 3’ G-rich strand (termed the G- overhang). In telomerase positive cells, the G-overhang is then extended by telomerase. Finally, much of the elongated overhang is converted to duplex DNA by DNA polymerase alpha (pol α) in a process known as C-strand fill-in. This leaves a short G-overhang that is then bound by telomere proteins.

CST is a protein complex that binds ssDNA and promotes telomere replication in a wide range of eukaryotes (36,84-87). Budding yeast CST (Cdc13-Stn1-Ten1) binds the G-overhang where it protects the telomere, recruits telomerase and mediates C-strand fill-in (35,88-90). Mammalian CST (CTC1-

STN1-TEN1) is less important for telomere-end protection but it functions both in telomere duplex replication and C-strand fill-in (39,52-54,60). It is also proposed to limit telomerase action, perhaps by competing for binding to the telomere protein TPP1 (56,66).

CST has additional genome-wide roles that are just starting to be appreciated (53,58,65,84,91,92).

In humans, CST facilitates recovery from various forms of replication stress throughout the genome. It promotes activation of dormant or late firing origins in response to replication fork stalling (53) and enhances viability when cells are treated with drugs that block replication fork progression (65).

Mutations in CTC1 cause the diseases Coats plus and dyskeratosis congenita (93-95). The telomeric and non-telomeric roles of CST are likely to underlie the severity of these diseases.

Although the mechanism of CST action is still unclear, multiple studies indicate a link to pol .

Mammalian CTC1 and STN1 were originally identified as Alpha Accessory Factor (AAF), a factor that co-purified with pol  and enhanced its processivity and affinity for ssDNA templates (50,51). CST and pol  have since been shown to interact in yeast, plants and mammals (1,35,52,96). Xenopus CST stimulates DNA priming by pol  on ssDNA (97) while Candida CST enhances primase activity and primase to polymerase switching (98).

CST exhibits notable structural similarities to Replication Protein A (RPA) the eukaryotic ssDNA binding protein that directs the assembly of multi-protein complexes needed for DNA replication, recombination and repair (Fig. 1A)(60). RPA has three subunits (RPA1, RPA2 and RPA3) that together harbor six OB (oligonucleotide-oligosaccharide binding) folds (Fig. 1A) (99,100). Four of the OB folds participate in DNA binding. Because RPA has multiple DNA binding sites, individual OB folds can undergo rapid dissociation and re-association without causing the protein to fall off the DNA (101,102).

This dissociation and re-association of OB folds underlies RPA function as it makes binding dynamic and enables RPA to diffuse along DNA to melt DNA structure or load and unload proteins needed for replication, recombination or repair (37).

Like RPA, CST appears to harbor OB folds in all three subunits and X-ray crystallography indicates striking structural similarity between STN1-TEN1 and RPA2-RPA3 dimers (38,62,103). The structural conservation encompasses the OB-fold and winged helix domains and the dimerization interface. The large subunits of RPA and CST appear less well conserved. Although RPA1 and Cdc13 from budding yeast each harbor 4 OB folds, Cdc13 needs only one OB fold for high affinity binding

(100,104). Moreover, Cdc13 dimerizes through its N-terminal OB fold to form a DNA pol α binding surface, whereas RPA1 does not self-associate (105). In mammalian cells, Cdc13 is replaced by CTC1 but the two proteins share little sequence identity and the extent of structural or functional conservation is unclear (1). Protein threading programs (Phyr2 and HHpred) predict 5-6 OB folds in human CTC1 with the three most C-terminal folds resembling those of RPA1 (Fig. 1A). In vitro studies have revealed an additional parallel between RPA and human CST as in each case high affinity DNA binding requires formation of the three protein complex (56,60,99,100). The structural similarities between RPA and CST raise the possibility that dynamic DNA binding through multiple OB folds may also contribute to CST function.

Since so little is known about CST mechanism of action and the relationship between DNA binding and CST function, we set out to analyze how reduced DNA binding affects CST activity at

35 telomeres and elsewhere in the genome. We describe a STN1 OB fold mutant that preferentially affects in vitro binding to short DNA substrates. In vivo, the mutant can substitute for wild type STN1 in some aspects of CST function but other aspects are impaired. DNA binding studies indicate that, like RPA,

CST appears to contact DNA via multiple OB folds and to have distinct modes of binding. However, we also provide evidence that the organization of OB-fold engagement by CST is quite different.

3.4 RESULTS

3.4.1 Alteration of DNA-binding by STN1 OB-fold mutation

To generate the STN1 OB-fold mutant (STN1-OBM) we changed three residues (W89A, R139L, Y141A) that are conserved between STN1 and the OB fold of RPA2 and which either directly contact, or lie very close to DNA in RPA crystal structures (Fig. 1B) (99,106). The W89A and Y141A mutations were chosen because the equivalent mutations in mouse STN1/AAF-44 reduced DNA binding by ~60% in pull-down assays with biotin-labeled poly-dC (50)

Co-immunoprecipitation and tandem affinity purification experiments verified that the STN1 mutant retained the ability to form a complex with CTC1 and TEN1 (Fig. 1C & D). In initial experiments, we co-expressed FLAG-tagged STN1 or STN1-OBM with HA-CTC1 in a previously characterized HeLa cell line over-expressing TEN1 (65). When STN1-OBM was immunoprecipitated from whole cell lysate, both CTC1 and TEN1 co-purified (Figure 1C). We also generated recombinant CST complexes containing wild type STN1 (CST (WT)) or STN1-OBM (CST(STN1-OBM)) by co-infecting insect cells with baculovirus encoding FLAG-tagged CTC1, untagged TEN1 and His-tagged STN1 or STN1-OBM.

Protein complexes were affinity purified on nickel resin followed by FLAG beads (Fig. 1D) and again

CTC1 and TEN1 co-purified with STN1-OBM.

Fig 1. STN1 OB fold mutation affects DNA binding but not CST complex formation.

(A) Cartoon depicting known and predicted OB fold domains in RPA and CST, white boxes indicate OB folds. Black bars mark OB folds in CST (known for STN1, predicted for CTC1, see Materials and

Methods) where mutations alter DNA binding (50,86). (B) Alignment of human STN1 and RPA32 OB

37 folds showing identical and similar (+) residues (107). Arrows indicate mutations in the STN1 OB fold mutant (STN1-OBM), arrowheads mark corresponding RPA residues that contact DNA directly in the U. maydis crystal structure (99). (C) Western blots showing co-immunoprecipitation of CTC1 and TEN1 with STN1 or STN1-OBM using extracts from TEN1 overexpressing HeLa cells transiently transfected with HA-CTC1 and FLAG-STN1 or FLAG-STN1-OBM. (D) Silver-stained gels showing co-purification of FLAG-CTC1 and TEN1 with His-STN1 or His-STN1-OBM expressed in insect cells. Left, a full 12% gel showing all three subunits. Right, portion of a more heavily stained 15% gel showing STN1 and

TEN1. (E-F) EMSAs showing CST(WT) or CST(STN1-OBM) binding to non-telomeric (NonTel) or telomeric G-strand (TelG) oligonucleotides or dsDNA (DS) of the indicated lengths. Increasing concentrations of CST were incubated with 0.1 nM labeled DNA for 30 min prior to separation in agarose gels. (E) The same CST(WT) or CST(STN1-OBM) preparation was used in all the EMSAs with ssDNA

We next examined how STN1-OBM affects the ability of CST to bind a range of DNA substrates.

As the affinity of CST for short versus long substrates seems to depend on DNA sequence (56), we monitored binding of CST(WT) and CST(STN1-OBM) to telomeric and non-telomeric oligonucleotides of various lengths (Fig. 1E-F, Table 1). When we used electrophoretic mobility shift assays (EMSA) to compare binding of CST(WT) and CST(STN1-OBM) to 48 nt substrates, the two complexes appeared to bind both non-telomeric (NonTel-48) and telomeric G-strand (TelG-48) DNA with similar affinity.

However relative to CST(WT), the CST(STN1-OBM)) bound less efficiently to the 36 nt non-telomeric

(NonTel-36) and the 18 nt telomeric G-strand (TelG-18) substrates. Neither complex bound equivalent concentrations of the 18 nt non-telomeric oligonucleotide (NonTel-18) or dsDNA (Fig. 1E)(60). These results suggest that the STN1-OBM preferentially affects binding to short substrates. Note, we refer to

TelG-18 as a short substrate because CST has very low affinity for DNA with fewer telomeric repeats, e.g. TelG-12 (56). Our results also confirm that CST binds both telomeric and non-telomeric DNA but that telomeric DNA is preferred when substrate length is short (56).

Table 1. Sequence of oligonucleotides used in DNA binding assays.

Oligo Sequence (5’-3’)

TelG-18 GGTTAGGGTTAGGGTTAG

TelG-36 GGTTAGGGTTAGGGTTAGGGTTAGGGTTAGGGTTAG

TelG-48 GGTTAGGGTTAGGGTTAGGGTTAGGGTTAGGGTTAGGGTTAGGGTTAG

NonTel-18 AGCGTATCCGTTCAGTTG

NonTel-36 AGCGTATCCGTTCAGTTGAGCGTATCCGTTCAGTTG

NonTel-48 AGCGTATCCGTTCAGTTGAGCGTATCCGTTCAGTTGAGCGTATCCGTT ds 15 TTTCGATCTACGTCAGCA 5’ ...... TTGCTAGATGCAGTCGT 3’ ds 40 TTTTACGTCAGCACGATCTACGTCAGCACGATCTACGTCAGCA 5’ ...... TTATGCAGTCGTGCTAGATGCAGTCGTGCTAGATGCAGTCGT 3’

3.4.2 STN1-OBM fails to rescue to anaphase bridges after endogenous STN1 depletion

To examine the in vivo effects of STN1 OB fold mutation, we generated HeLa cells that stably express FLAG-tagged STN1-OBM (Fig. 2A and Appendix Fig. S1A) by introducing an shRNA-resistant

STN1-OBM cDNA into a previously characterized HeLa cell line expressing STN1 shRNA (shSTN1)

(53,54). A cell line expressing FLAG-tagged shRNA-resistant wild type STN1 (STN1-Res) was previously made in the same manner (53).

Fig 2. In vivo expression of STN1-OBM causes anaphase bridges.

(A) Western blot showing levels of STN1 in HeLa cells expressing non-target shRNA (shNT) or STN1 shRNA (shSTN1) and shSTN1 cells with sh-resistant mutant STN1 (STN1-OBM) or wild type STN1

(STN1-Res). Blot was probed with antibody to STN1 or to actinin for a loading control. (B) Left; representative images of DAPI-stained anaphase cells with/ without bridges. Anaphase cells with no bridge in shNT (top) and with bridges in shSTN1 and STN1-OBM cells (middle and bottom). Right; quantification of bridges (200 anaphases counted per cell line per experiment. n = 3 experiments, mean +

S.E.M, p-values are indicated above bars).

In initial experiments, we asked if STN1-OBM could rescue the increase in anaphase bridges that occurs after STN1 depletion (53). STN1-OBM cells and a series of control cells (shSTN1, STN1-Res and shNT, a non-target shRNA control) were arrested in mitosis with nocadazole, released for 45-60 min,

40 fixed and scored for the number of anaphase cells with DAPI-stained bridges (Fig. 2B). As previously described, depletion of STN1 caused an increase in bridges and this was rescued by expression of sh- resistant wild type STN1 (53). In contrast, expression of sh-resistant STN1-OBM did not rescue bridge formation but instead further increased the fraction of anaphase cells with bridges. The reason for the higher level of bridges in the STN1-OBM cells relative to the shSTN1 cells is unclear but a possible cause is that STN1-OBM replaces residual endogenous STN1 in CST complexes. Overall, the inability of

STN1-OBM to rescue the anaphase bridge phenotype indicates that STN1-OBM cannot substitute for wild type STN1 in some aspects of CST function.

3.4.3 STN1-OBM affects telomere duplex replication but not C-strand fill-in

Anaphase bridges can have a number of causes including telomere-to-telomere fusion and the presence of unresolved replication intermediates either at telomeres or elsewhere in the genome

(108,109). Thus, to ask more specifically whether STN1-OBM affects the telomeric roles of CST, we looked for changes in telomere structure. Metaphase spreads from STN1-OBM and control cells were hybridized with telomere probe and examined for telomere loss, telomere fusions or other abnormal telomere signals. As previously reported, we did not observe an increase in telomere loss or telomere fusions in the shSTN1 and STN1-Res cells (53)(Appendix Fig. S1B). This was also true for the STN1-

OBM cells (Appendix Fig. S1B), indicating that the anaphase bridges caused by STN1-OBM expression are unlikely to be caused by telomere fusions. However, relative to the STN1-Res control, the STN1-

OBM cells showed a large increase in individual chromatids exhibiting Multiple Telomeric FISH Signals

(MTS) (Fig. 3A). As expected, the shSTN1 cells also showed an increase in MTS but it was lower than in the STN1-OBM cells. Again this may reflect the displacement of residual endogenous STN1 with STN1-

OBM in CST complexes. Past studies have shown that MTS arise after fork stalling during replication of the telomere duplex (80) and they occur after depletion of the various factors needed for telomere

41 replication, including CST (54,80). In particular, STN1 depletion slows replication through the telomere duplex and causes the appearance of MTS (54). We therefore conclude that STN1-OBM is unable to rescue the deficiency in telomere duplex replication caused by STN1 depletion.

We next asked if STN1-OBM affects telomere length or G-overhang structure. Genomic DNA was isolated from STN1-OBM or control cells and telomere restriction fragments were examined by

Southern blotting or in-gel hybridization to monitor telomere length (Appendix Fig. S1C). This analysis revealed that telomeres of shSTN1, STN1-Res and STN1-OBM cells were very similar in length, thus confirming our previous finding that telomere length in HeLa cells is largely unaffected by STN1 knockdown (54) and indicating that STN1-OBM has dominant negative effect. We then used an in-gel hybridization assay to ask if STN1-OBM affects G-overhang structure. Restriction digested DNA was separated briefly in agarose gels and hybridized with a probe to the telomeric G-strand under non- denaturing conditions (Fig. 3B). The DNA was then denatured and re-hybridized with the same probe.

Quantification of the overhang signal relative to total telomeric DNA revealed the expected increase in overhang amount in the STN1-depleted cells (Fig. 3C). This increase has previously been shown to result from inefficient C-strand fill-in following telomerase extension (52,54). Given that STN1-OBM affects binding to telomeric G-strand DNA in vitro (Fig. 1E) and telomere duplex replication in vivo (Fig. 2), we anticipated that the STN1-OBM cells would also have a deficiency in C-strand fill-in. However, to our surprise we found that STN1-OBM cells had normal length overhangs (Fig. 3C) implying that the mutant

STN1 was able to rescue C-strand fill-in.

Fig 3. STN1-OBM causes multiple telomere signals (MTS) but does not affect G-overhang maintenance.

(A) Telomere FISH of STN1-OBM or shSTN1 cells. Left; representative images of single metaphase chromosomes. White arrows, MTS; green, FITC-(C3TA2)3 probe; blue, DAPI. Right; Quantification of

MTS (n = 4 experiments mean + S.E.M.). Individual chromosomes were scored positive for MTS if they had MTS at one or more telomeres. (B-C) G-overhang abundance in asynchronous cells monitored by in- gel hybridization with (A2TC3)4 probe. (B) Representative gels showing hybridization to genomic DNA from the indicated cells under native or denaturing conditions. (C) Quantification of G-overhang abundance in asynchronous cells (n = 3 experiments, mean + S.E.M., p-values are indicated above bars).

G-overhang length is determined by a number of activities that occur at specific stages in the cell cycle. Overhangs are elongated in S-phase as a result of G-strand synthesis by telomerase and C-strand resection by nuclease (7,79). They are then returned to their original length in late S/G2 via C-strand fill- in by DNA pol α (54). Given this balance between activities, it was possible that the normal length overhangs in the STN1-OBM cells result from decreased G-strand extension in S-phase in combination with decreased C-strand fill-in during late S/G2. To investigate this possibility, we examined G-overhang length dynamics during the cell cycle. Cells were synchronized in G1/S with a double thymidine block, released into S-phase and harvested at intervals as they passed through mid S-phase, G2/M and back into

G1 (Fig. 4A and Appendix Fig. S2A). Following DNA isolation, relative overhang length was examined by in-gel hybridization as described above (Figure 4B-C). Quantification of the overhang signal indicated that the STN1-Res cells showed the expected increase in overhang abundance as they transitioned from

G1 (0 hr) into mid S-phase (6 hr) (7,54,79). The overhang signal then declined due to C-strand fill-in as the cells transitioned into G2 (8 hrs) and G1 of the next cell cycle (10-12 hrs) (10,54). Interestingly, the pattern of overhang elongation and shortening in the STN1-OBM cells was indistinguishable from that seen with the control STN1-Res cells indicating that STN1-OBM does not affect overhang elongation or

C-strand fill-in. In contrast, the shSTN1 cells exhibited the expected delay in overhang shortening in late

S/G2 reflecting the deficiency in C-strand fill-in (54). Thus although STN1-OBM affects telomere duplex replication, it does not appear to affect C-strand fill-in by DNA pol α.

Fig 4. STN1-OBM is competent for C-strand fill-in and TPP1 and pol α interaction.

(A-C) Analysis of C-strand fill-in. (A) FACS analysis of DNA content showing synchrony of STN1-

OBM cells used in (B). (B) Representative gels showing in-gel hybridization of (TA2C3)4 probe to DNA from cells harvested at the indicated times after release from G1/S block. (C) Quantification of G- overhang abundance. Cell types were analyzed in pairs, n = 3 experiments for shSTN1 + STN1-OBM,

45 mean + S.E.M.; n = 2 experiments for STN1-OBM + STN1-Res, error bars show min/max values). (D)

Western blot showing co-immunoprecipitation of TPP1 with STN1 or STN1-OBM. Cells were transfected with FLAG-STN1 or FLAG-STN1-OBM plus HA-mCherry-TPP1 expression constructs.

TPP1 was precipitated with antibody to HA. (E) Co-immunoprecipitation of DNA pol α with CST. Cells were transfected with FLAG-STN1 or FLAG-STN1-OBM, Myc-CTC1 and TEN1. CST was precipitated with FLAG beads.

3.4.4 STN1-OBM does not disrupt interaction with TPP1 or DNA pol α

Several studies have shown that STN1 can interact with the shelterin protein TPP1 (55,56), suggesting that this interaction might be important for recruiting CST or stabilizing CST binding at the telomere. Given that OB folds can mediate protein-protein interactions as well as DNA binding (100), we considered the possibility that the in vivo effects of STN1-OBM expression might reflect decreased binding of CST to TPP1. To test for a disruption in the TPP1-STN1 interaction, we transfected 293T cells with constructs encoding FLAG-tagged STN1 or STN1-OBM and HA-mCherry-tagged TPP1 (69) and monitored co-immunoprecipitation of TPP1 with STN1. When TPP1 was precipitated with antibody to

HA, Western blot analysis showed that the STN1 and STN1-OBM co-precipitated with equivalent efficiency (Fig. 4D). We therefore conclude that STN1-OBM retains the ability to bind TPP1. We also tested whether STN1-OBM disrupts binding to DNA pol α, the only other known CST binding partner

(52,110). 293T cells were transfected with constructs encoding TEN1, FLAG or Myc-tagged CTC1 and

FLAG-STN1 or FLAG-STN1-OBM, and CST was then precipitated from extracts with FLAG antibody.

Western blot analysis of the immunoprecipitates showed that pol α co-precipitated with FLAG-STN1 only if both CTC1 and STN1 were overexpressed (Appendix Fig. S2B). However, the level of pol α precipitation was similar with CST(WT) and CST(STN1-OBM), indicating that STN1-OBM does not prevent CST from binding to pol α (Fig. 4E and Appendix Fig. S2B). Our finding that C-strand fill-in is

46 unaffected by STN1-OBM (Fig. 3B and Fig. 4C) provides further support for a functional interaction between pol α and CST(STN1-OBM),

3.4.5 STN1-OBM can function in replication rescue after genome wide replication fork stalling

Since the above studies indicate that STN1-OBM has selective effects on CST function, we next examined whether the mutant affects the response to genome-wide replication fork stalling. In initial experiments, we asked if STN1-OBM could substitute for wild type STN1 to maintain cell viability after

HU (hydroxyurea) treatment. STN1-OBM and control cells were treated with 2 mM HU for 0-24 hrs, allowed to recover for 24 hrs then cell viability was monitored by MTT assay (Fig. 5A). As observed previously, STN1 depletion increased sensitivity to HU (65). However, wild type STN1 (STN1-Res) and

STN1-OBM rescued this sensitivity to an equal extent, indicating the mutant was sufficient to allow CST function in recovery from prolonged fork stalling.

To further explore the effect of STN1-OBM on recovery from fork stalling, we performed DNA fiber analysis to determine if the mutant can substitute for endogenous STN1 to promote origin firing after HU treatment. Cells were labeled with IdU (iododeoxyuridine) for 15 minutes, treated with HU for two hours then released into media containing CldU (chlorodeoxyuridine) for 60 min (Fig. 5B). The cells were then collected, lysed and the DNA fibers spread on silanized slides by hydrodynamic flow (68). The fibers were stained with antibody to IdU and CldU then visualized by confocal microscopy to score the replication events (Fig. 5B-D, Appendix Fig. S3). As observed previously, the HU-treated shSTN1 cells exhibited fewer green-only (CldU-only) tracks than the shNT and STN1-Res control cells (53,65), indicating that STN1 depletion caused a decrease in new origin firing after HU release. In contrast, the

HU-treated STN1-OBM cells exhibited a similar number of green-only tracks to the control cells. The frequency of other replication events was also similar (Appendix Fig. S3). We therefore conclude that the

STN1-OBM can substitute for wild type STN1 to promote new origin firing. Overall, our results indicate that STN1-OBM does not affect the capacity of CST to aid in the restart of replication following

47 exogenous replication stress. This is in direct contrast to the inability of STN1-OBM to rescue the effects of endogenous stress as seen by the increase in anaphase bridges and MTS in unchallenged STN1-OBM cells.

Fig 5. STN1-OBM rescues viability and restores origin firing after HU treatment.

(A) MTT assay showing viability after HU treatment. Cells were treated with 2 mM HU for the indicated times and harvested for MTT assay 24 hrs later. Values are relative to untreated cells of the same cell

48 type. Each time point was assayed in triplicate and the data are shown as the mean ± S.D from 3 independent experiments. For each cell line, the value of the untreated sample was set at 1. (B-D) DNA fiber analysis of origin firing following release from 2 mM HU. (B) Left: schematic showing timing of

IdU and CldU labeling relative to HU treatment. Right: types of replication event scored. (C)

Representative images of DNA tracks. Red, IdU; Green, CldU. (D) Graph indicating the percent of DNA tracks showing new origin firing (green-only tracks) (n = 7 experiments, mean + S.E.M, p-values are indicated above bars).

3.4.6 Effects of STN1-OBM on binding affinity and stability

Our finding that STN-OBM affects only specific aspects of CST function is analogous to what has been observed for certain RPA OB-fold mutants, which support DNA replication but are defective for

DNA repair (111,112). These mutants cause only a small decrease in overall affinity of RPA for ssDNA and the deficit in repair is thought to result from a change in the dynamics of RPA binding through its multiple OB folds (37,112). The structural similarities between CST and RPA suggest that CST function could also rely on dynamic binding using multiple OB folds. We therefore set out to explore the extent to which RPA binding can be used as a paradigm for understanding CST activity and the in vivo separation of function observed with STN1-OBM.

As a first step, we revisited the effect of STN1-OBM on DNA binding by using filter binding assays to quantify the affinity of CST(WT) and CST(STN1-OBM) for telomeric and non-telomeric substrates of various lengths (Fig. 6A & C, Appendix Fig. S4). CST purified from insect cells was incubated with 32P-labeled DNA then the DNA-protein complexes were separated from free DNA by filtration through a sandwich of nitrocellulose and HyBond membrane. The bound versus free DNA was quantified and used to calculate the apparent dissociation constant (Kd,app). Despite the different approach used to separate bound from free DNA in the filter binding and the original gel shift assay (Fig.

1E), the results of the two assays were qualitatively similar. The filter binding indicated that CST(WT) and CST(STN1-OBM) bound the 48 nt telomeric and non-telomeric substrates with a similar Kd,app while binding to TelG-18 was decreased for CST(STN1-OBM) relative to CST(WT) (Fig. 6A & C).

Thus, the filter binding analysis again indicated that STN1-OBM preferentially affects binding to short

DNA substrates. However the analysis also revealed that the overall decrease in Kd,app for CST(STN1-

OBM) binding to TelG-18 was only 2-3 fold.

When gel shift assays were used to examine CST(STN1-OBM) binding to the TelG-18 and

NonTel-36 substrates a substantial amount of DNA was seen to migrate between the bands corresponding to free DNA and CST-bound DNA (Fig. 1E). This observation suggested that the DNA-protein complexes were dissociating and hence the decrease in Kd,app for CST(STN1-OBM) might reflect less stable binding. To test this possibility, we measured the rate of CST(WT) and CST(STN1-OBM) dissociation (t½) from selected substrates. CST complexes were bound to 32P-labeled oligonucleotide, challenged with an excess of the corresponding cold oligonucleotide for various times and the remaining labeled DNA/protein complex was quantified by filter binding. This experiment revealed that CST(STN1-

OBM) dissociated from the labeled TelG-18 and NonTel-36 1.6-2.6-fold faster than CST(WT) whereas dissociation from TelG-48 and NonTel-48 was essentially the same (Fig. 6B & C). We therefore conclude that the STN1-OB fold acts to stabilize CST binding to short ssDNA substrates.

The 2-3 fold decrease in affinity of CST(STN1-OBM) for TelG-18 resembles the modest decrease in RPA affinity for ssDNA that has been observed after mutation of individual OB folds

(111,113). In the case of RPA, the small effect on overall binding affinity reflects the presence of multiple

DNA binding domains within the complex such that disruption of one binding domain has a small effect on the macroscopic affinity constant. Thus, the observed decrease in CST(STN1-OBM) binding fits with the model that CST also engages DNA via multiple DNA binding domains. Given the six predicted OB folds in CTC1, we anticipate that these multiple DNA binding domains correspond to the STN1 OB fold plus some or all of the OB folds in CTC1.

Fig 6. Analysis of CST DNA binding parameters.

(A) Binding isotherms used to determine apparent dissociation constants for CST(WT) or CST(STN1-

OBM) and the indicated ssDNA substrates. Data were obtained by filter binding assay and fit to a one site

51 specific binding model. Mean ± SEM, n = 3 independent experiments each with a different protein preparation. (B) Dissociation kinetics for CST bound to the indicated substrates The fraction of labeled

DNA remaining bound was determined by filter binding at the indicated times. Data were fit to a one phase exponential decay equation to obtain the dissociation rate (t½)). Mean ± SEM, n = 3 independent experiments. (C) Table summarizing Kd(app) and t½ for CST(WT) or CST(STN1-OBM) and the indicated ssDNA substrates.  undetectable binding, ND: not determined.

3.4.7 CST subunit interactions with ssDNA substrates

While CST appears to resemble RPA in terms of subunit composition and utilization of multiple

OB folds for DNA binding, our finding that CST(STN1-OBM) destabilizes binding to short oligonucleotides suggested a significant difference in how the two complexes bind short DNA substrates

(Fig. 6C). RPA binds DNA in a 5’ to 3’ direction with the OB-folds of RPA1 contacting DNA towards the 5’ end and providing the highest affinity binding sites (99,113). As a result, OB-A and OB-B of RPA1 provide the only contacts to an 8 nt substrate. OB-A, -B and –C of RPA1 contact substrates of 12-23 nt but RPA2 (the STN1 equivalent) only contacts longer substrates of 30 nt (99,100). Consequently, mutations in RPA2 OB-D affect binding to long rather than short ssDNA (111,113). Our finding that

STN1-OBM destabilizes binding to short (e.g. TelG-18) but not long (TelG-48 & NonTel-48) substrates

(Fig. 6C) suggested that, unlike RPA2, STN1 directly engages the DNA of short substrates to stabilize binding.

To further explore this possibility, we used photo-crosslinking to explore the proximity of individual CST subunits to the 5’ or 3’ ends of 18 or 48 nt TelG oligonucleotides. CST(WT) and

CST(STN1-OBM) were incubated with 32P-labeled TelG-18 or TelG-48 that had a photoactivatable 4- thiothymidine (s4T) at the third nucleotide from the 5’ or 3’ end (Fig. 7A). The DNA-protein complexes were cross-linked by irradiation with UVA and then separated in a SDS-polyacrylamide gel. The gel was

52 scanned by phosphorimager to determine whether CTC1, STN1 or TEN1 had been cross-linked to the labeled DNA. Equivalent UV-irradiated samples separated in the same gel were used for Western blot analysis to determine the positions of uncross-linked CTC1, STN1 and TEN1. The low level of cross- linking precluded detection of the cross-linked DNA-protein complexes by Western blot. Additional reactions that had not been subject to cross-linking were analyzed by EMSA to monitor DNA binding. As shown in Fig.7B, the s4T residues did not significantly alter CST(WT) or CST(STN1-OBM) binding to either substrate.

Analysis of the crosslinking products obtained with CST(WT) and 5’- or 3’-s4T TelG-18 revealed labeled bands that migrated at positions expected for CTC1 (130 kD) and STN1 (>43 kD) (Fig. 7C) indicating cross-linking to either substrate. However, cross-linking of STN1 relative to CTC1 was less efficient with the 5’-s4T TelG-18, suggesting that STN1 was positioned closer to the DNA 3’ end. It was not possible to tell if TEN1 was cross-linked to either substrate because TEN1 migrated in the same region of the gel as the uncross-linked DNA. Thus, bands corresponding to TEN1-TelG-18 may be obscured by the heavy signal from the uncross-linked DNA. Overall, the results indicate that binding of

CST to a short 18 nt substrate positions the DNA in close proximity to STN1. Comparison of the cross- linking products obtained with the 3’ modified TelG-18 and CST(WT) or CST(STN1-OBM) revealed that cross-linking to STN1-OBM was reduced relative to wild type STN1. This finding indicates that the contacts between STN1 and DNA are altered by STN1-OBM.

Analysis of the products obtained with CST(WT) bound to TelG-48 revealed that only CTC1 was reproducibly cross-linked to the 5’-s4T substrate. In contrast, the 3’-s4T substrate crosslinked to all three

CST subunits. CTC1 cross-linked more efficiently than STN1 or TEN1 and the level of TEN1 cross- linking was somewhat variable (Fig. 7D) (note: cross-linking of TEN1 to TelG-48 retards TEN1 migration enough for the band from the cross-linked product to become visible above the uncross-linked

DNA). The cross-linking of STN1 and TEN1 to the 3’-s4T substrate but not the 5’-s4T substrate indicates that both subunits must be in close proximity to the DNA 3’ terminus but not the 5’ terminus.

Fig 7. Photo-crosslinking of CST subunits to thiothymidine substituted ssDNA.

(A) Positions of s4T substitutions in TelG-18 and TelG-48 DNA substrates. (B) EMSA showing the s4T substitutions do not affect CST(WT) or CST(STN1-OBM) binding to TelG-18 or TelG-48. (C-D)

Products obtained after photo-crosslinking. Left and right panels: Phosphorimager scans showing 32P- labeled cross-linking products. Central panels: Western blots from the same gels showing positions of uncross-linked CTC1, STN1 and TEN1. (C) Products obtained with 3’ (left) or 5’ (right) modified TelG-

18. (D) Products obtained with 3’ (left) or 5’ (right) modified TelG-48. * indicates cross-linking products observed only in some experiments. They may represent CTC1 or STN1 degradation products. Markers on the phosphorimager scans were obtained by laying the gels on nitrocellulose membrane and marking the positions of the marker bands with radioactive ink. For the Western blots, the membrane was cut into pieces, probed with antibody to CTC1, STN1 or TEN1, reassembled and exposed to film. The film was laid over the membrane and photographed to visualize both the markers and the CST bands. (E) Dynamic binding model of CST showing micro-dissociation of an individual OB fold (blue) to allow binding of an alternative protein (yellow).

Examination of the photo-products obtained with of CST(STN1-OBM) bound to 3’-s4T TelG-48 revealed less cross-linking to STN1 and TEN1 relative to CST(WT) but CTC1 photoproducts were still formed, again indicating that STN1-OBM alters how STN1 contacts DNA.

Taken together the above results demonstrate that CST(WT) binds long substrates with the DNA

3’ end positioned close to the CTC1-STN1-TEN1 interface while the 5’ end only contacts CTC1. We therefore infer that, CST binds DNA in a similar orientation to RPA: i.e. with the large subunits of each complex contacting DNA near the 5’ end and the two smaller subunits positioned at the 3’ end. However, our data indicate that the identity of the binding sites used to engage short DNA substrates differs between CST and RPA. For CST, the binding sites lie close to the interface between CTC1 STN1 and

TEN1, they engage DNA toward the 3’ end, and STN1 plays an important role in stabilizing the interaction. For RPA, the primary binding sites for short substrates are OB-A and OB-B of RPA1 and these engage DNA at the 5’ end. Thus, despite sharing some common structural features CST and RPA engage DNA quite differently.

In addition to addressing the architecture of CST-DNA complexes, the in vitro cross-linking studies combine with the analysis of DNA binding affinity start to explain the in vivo separation of function observed with STN1-OBM cells. Our results show that the STN1 OB-fold mutation alters the interaction between STN1 and ssDNA and this translates into altered binding of CST to some but not all

DNA substrates. In vivo, CST is likely to encounter DNA substrates of varied length and structure as the complex helps resolve a wide range of replication problems at telomeres and genome-wide. Thus, similar to what has been observed for RPA OB-fold mutants (111,112), the altered DNA binding caused by

STN1-OBM is likely to impair the ability of CST to bind and mediate the resolution of only a subset of replication intermediates.

3.5 Discussion

Here we describe a series of in vivo and in vitro experiments that address the mechanism of CST action at telomeres and elsewhere in the genome. We show that a STN1-OB-fold mutant (STN1-OBM) which preferentially decreases affinity of CST for short ssDNA substrates is competent for some aspects of CST function but deficient in others. The effects of STN1-OBM do not align with the telomeric versus non-telomeric roles of CST, but instead separate out the different aspects of CST function both during telomere replication and in genome-wide replication rescue. At telomeres, STN1-OBM cells are competent for C-strand fill-in following telomerase action but they exhibit increased MTS which are indicative of deficiencies in the earlier process of telomere duplex replication. STN1-OBM cells are also competent to restart replication via new origin firing following exogenous genome-wide replication stress. However, STN1-OBM is not able to prevent the accumulation of anaphase bridges during mitosis.

The latter finding indicates a deficiency in genome-wide resolution of endogenous replication stress because the anaphase bridges caused by CST depletion occur at both telomeric and non-telomeric loci

(65,66). Our findings underscore the importance of CST for multiple processes associated with telomere replication and genome-wide replication rescue. They also strongly suggest that different DNA binding transactions are needed for CST to resolve different forms of replication stress with a subset of these transactions being disrupted by STN1-OBM. While STN-OBM did not inhibit interactions with TPP1 or pol α (Fig 4), it is possible that STN1-OBM disrupts CST interaction with as yet unidentified partner proteins. If this is the case, the interaction of STN1 with such proteins might provide an additional mechanism to target CST to its various sites of action within the genome.

Our in vitro DNA binding studies using CST(WT) and CST(STN1-OBM) provide new insight into the mechanism of mammalian CST binding to ssDNA. Past studies provided conflicting information concerning the sequence specificity of CST binding (56,60). We now confirm that human CST binds long

(48 nt) substrates with little sequence specificity, however sequence identity is important for binding to short (18 nt) substrates as the telomeric G-strand substrate TelG-18 is bound with high affinity while binding to the non-telomeric substrate NT-18 is undetectable. We also provide evidence that human CST harbors multiple DNA binding domains. The STN1-OB fold comprises one of these domains and based on structure prediction, we suggest that OB folds in CTC1 comprise the others. Since CST only bound the

18 nt substrate that had the sequence of telomeric G-strand DNA (Fig. 1), the domain(s) that bind short oligonucleotides (i.e. the STN1 OB fold or an adjacent OB fold in CTC1) must provide important determinants for sequence-specific binding. Given that long substrates (telomeric and non-telomeric) are bound with higher affinity than short substrates and their binding is less affected by STN1-OBM, it seems likely that these substrates contact additional DNA-binding domains beyond those used to contact short substrates.

The known structural similarity between STN1-TEN1 and RPA2-RPA3, together with the likely presence of multiple OB folds in CTC1, had previously suggested an RPA-like binding mechanism

57 whereby mammalian CST contacts DNA via multiple OB folds. However, this was not a foregone conclusion because S. cerevisae CST binds DNA through one high affinity binding site in Cdc13 (104).

While our work supports the multiple OB fold binding mechanism for mammalian CST, it also reveals significant differences between CST and RPA in the contributions made by individual subunits during binding to ssDNA. For RPA, the only binding sites for short substrates correspond to the OB folds of

RPA1 that bind proximal to the DNA 5’ end (99,100). These OB folds also comprise the highest affinity binding sites. However, for CST, both CTC1 and STN1 contact short DNA substrates and STN1, which binds near the DNA 3’ end, is necessary to stabilize binding. These findings imply that the high affinity binding sites in CST are contributed by STN1 and CTC1 and they interact with DNA towards the 3’ end.

While this architecture differs from that of RPA, it is well suited for CST to bind a telomeric 3’ overhang.

Despite the above differences between the two protein complexes, RPA can still be used as a model to help us understand the relationship between CST function and its mechanism of DNA binding.

The ability of RPA to act as a hub that directs the sequential loading and unloading of partners such as

Rad51 and Rad52 or SV40 T-antigen and pol α stems from the dynamic nature of RPA binding to ssDNA (99,102,114). Because RPA utilizes 4 OB-folds to bind DNA, individual OB folds can undergo rapid microscopic dissociation and re-association from the DNA without causing the whole protein to dissociate (101,102). Instead the rapid dissociation and re-association of individual OB folds is what enables RPA to diffuse along DNA to melt DNA structure or load and unload partner proteins (37).

Given that mammalian CST is likely to bind DNA via a similar number of OB-folds, it is possible that CST binding is also dynamic. If so, microscopic dissociation of individual OB folds from ssDNA could enable CST to engage or disengage interaction partners from the DNA (Fig. 7E). Like the CST complex from Candida glabrata, mammalian CST might also be able to resolve unwanted DNA structure such as G quadruplexes (G4) (115). The dynamic binding model for CST action is appealing because it can explain why CST is involved in multiple steps during telomere replication and in the resolution of diverse forms of replication stress. It can also explain many of the phenotypes of CST depletion. For

58 example, during telomere replication, CST might aid in removal of G4 structure from the lagging strand during replication of the duplex DNA and it may engage pol α on the G-overhang to initiate C-strand fill- in following telomerase action. The role in G4 structure removal could explain why STN1 depletion leads to a slowing of telomere duplex replication with formation of MTS. Likewise, the role in pol α engagement could explain why C-strand fill-in is disrupted despite pol α remaining associated with the telomere (52,54). The ability of CST to engage pol α to initiate DNA synthesis at dormant or late firing origins could also explain why STN1 depletion inhibits replication restart after genome-wide replication fork stalling. Moreover, resolution of DNA structure at G-rich or regions of repetitive sequence could underlie the role of CST in resolving endogenous replication stress at non telomeric loci (58,65,66).

While current models for CST function have focused on the regulation of DNA pol α, the large size of CTC1 suggests that CST will have many interaction partners. Thus, mammalian CST may well direct the actions of additional proteins involved in the resolution of replication stress. A broader understanding CST function will require the identification of these proteins and analysis of how CST modifies their ability to engage with stalled forks, replication origins or other replication intermediates.

If having multiple DNA binding domains and a dynamic DNA binding mechanism is so important for CST function in mammals, one has to ask why S. cerevisiae Cdc13 uses only one OB fold to bind DNA (104) and S. pombe appears to lack a Cdc13/CTC1 subunit (116). One possibility is that the multiple DNA binding domains necessary for dynamic binding are provided through dimerization or alternative subunit stoichiometries such as those found in S. cerevisiae and C. glabrata (105,115). An alternative answer could lie in the division of labor between CST and RPA and how this has evolved between organisms. In S. pombe, RPA cooperates with the helicase Pif1 to help resolve G4 structures at lagging strand telomeres (117,118) and a simple Stn1/Ten1 complex appears sufficient to regulate telomerase to pol α switching for C-strand fill-in (116,119). Thus, a full CST complex with dynamic

DNA binding properties may be unnecessary for telomere maintenance. Perhaps a CST complex is also

59 superfluous for genome-wide replication rescue because S. pombe RPA has adapted to function in these processes.

3.6 Supplemental Figures:

Fig S1. (A) PCR and sequencing strategy to monitor cells for presence of the wild type sh-resistant STN1 allele versus STN1-OBM. The cartoon indicates relative location of exons in endogenous STN1 mRNA.

Arrowhead indicates exon with mutations. Arrows indicate locations of primers used for PCR (black) or sequencing (dotted). (B) Telomere FISH of metaphase spreads from shSTN1, shNT, STN1-Res and

STN1-OBM cells. Representative images show that STN1-OBM does not cause increased telomere

60 fusion or telomere loss. White arrows, MTS; green, FITC-(C3TA2)3 probe; blue, DAPI. (C) Non- denaturing in-gel hybridization showing telomeric restriction fragments from the indicated cell lines.

Mean telomere lengths are shown at the bottom. Values represent the weight averaged mean from 3 or 4 independent experiments ± SD.

Fig S2. (A) FACS analysis showing cell synchronization of shSTN1, STN1-OBM and STN1-Res cells

62 used to analyze G-overhang length. (B) Co-immunoprecipitation of DNA pol α with CST. Extracts were from cells transfected with the indicated constructs. CST was precipitated with FLAG beads, these were then heated to 50C and loaded on the gel. Western blots were performed with antibody to Pol α, STN1,

TEN1 or FLAG. The Western blots with STN1 and TEN1 antibody show only the overexpressed protein because the levels of endogenous protein are too low to detect with the exposures that are shown.

Fig S3. Quantification of tracks scored during DNA fiber analysis with the indicated cell lines. The table shows total number of tracks scored for each replication event. Number in brackets indicates the percent of total tracks.

Fig S4. (A) Representative slot blots used to determine DNA binding affinity (Kd) for CST(WT) and

CST(STN1-OBM) binding to NonTel-36 or TelG-18. DNA concentrations are shown in brackets. (B)

Representative slot blot used to determine t½ for CST(WT) and CST(STN1-OBM) binding to NonTel-36 or TelG-18. Time of incubation with cold competitor DNA is shown in brackets.

Fig S5. Photocrosslinking of CST subunits to unmodified or 3’ thiothymidine substituted TelG-18.

CST(WT) or CST(STN1-OBM) was were incubated with unmodified or modified TelG-18, samples were irradiated with UV, separated in SDS gels and analyzed by phosphorimaging. * indicates cross-linking products observed only in some experiments. Markers on the phosphorimager scans were obtained by laying the gels on nitrocellulose membrane and marking the positions of the marker bands with radioactive ink.

4. CST DNA-binding dynamics melt G4 structure and regulate protein association with partial duplex and ssDNA

4.1 ABSTRACT

Human CST (CTC1-STN1-TEN1) is a ssDNA-binding complex that helps resolve replication problems both at telomeres and genome-wide. CST resembles Replication Protein A (RPA) in that the two complexes harbor comparable arrays of OB-folds and have structurally similar small subunits. However, the overall architecture and functions of CST and RPA are distinct. Currently, the mechanism underlying

CST action at diverse replication issues remains unclear. To clarify CST mechanism, we examined the capacity of CST to bind and resolve DNA structures found at sites of CST activity. We show that CST binds preferentially to ss-dsDNA junctions, an activity that can explain the incremental nature of telomeric C-strand synthesis following telomerase action. We also show that CST unfolds G-quadruplex structures, thus providing a mechanism for CST to facilitate replication through telomeres and other GC- rich regions. Finally, smFRET analysis indicates that CST binding to ssDNA is dynamic with CST complexes undergoing concentration-dependent self-displacement. These findings support an RPA-based model where dissociation and re-association of individual OB-folds allow CST to mediate loading and unloading of partner proteins to facilitate various aspects of DNA replication.

4.2 INTRODUCTION

Mammalian telomeres are bound by a protein complex called shelterin, which sequesters the DNA terminus to prevent it from being sensed as DNA damage and activating repair activities(76,120). The shelterin components TRF1 and TRF2 bind the telomeric dsDNA, which is composed of

TTAGGGAATCCC repeats, while POT1 binds to the ssDNA overhang on the G-rich strand. TPP1 links POT1 to TRF1/2 via TIN2. Mammalian cells also contain a second telomere-associated complex called CST (CTC1-STN1-TEN1) that is essential for telomere replication(39,60). Like its yeast counterpart (Cdc13-Stn1-Ten1), mammalian CST helps maintain telomere length by ensuring dsDNA is

66 formed after telomerase elongates the 3’ G-rich strand(36,54,56,57,121,122). Both yeast and mammalian

CST perform this role by enabling DNA polymerase to synthesize the complementary C-rich strand(36,54,98). This process is known as C-strand fill-in. In mammalian cells, CST-mediated C-strand fill-in is absolutely required for telomere length maintenance and cells lacking CST exhibit progressive telomere shortening similar to what is observed in cells that lack telomerase(57).

Mammalian CST has additional roles in replication both at telomeres and elsewhere in the genome. At telomeres, CST aids in passage of the replication fork through the telomere duplex. Removal of CST slows replication through this region(54,121) and leads to sudden telomere loss and/or a fragile telomere phenotype(53,66,121). The genome-wide roles of CST include facilitating replication through

GC-rich regions(58) and promoting firing of late or dormant replication origins after replication fork stalling(53,65). Loss of CST causes an increase in non-telomere associated anaphase bridges, fragile site expression and chromosome breaks, and a decrease in origin firing after HU treatment(53,58,66). Exactly how CST resolves all these replication-issues remains unclear. However multiple studies indicate a link to

DNA polymerase α (Pol α) as CST interacts with Pol α and stimulates its activity(50,59,121,123). During telomeric C-strand fill-in, CST appears unnecessary for telomeric localization of Pol α(121), suggesting that the complex acts to engage Pol α on the ssDNA rather than in Pol α recruitment. As CST preferentially localizes to GC-rich regions of the genome, it has also been suggested that CST may prevent replication fork stalling by melting G-quadruplex (G4) structure(58,115).

Given that CST harbors multiple OB-folds (one each in STN1 and TEN1 and 5-6 predicted in

CTC1)(59,62), we previously suggested that CST might perform its varied roles in replication by utilizing a dynamic binding mechanism similar to that observed for RPA(37,59,101,124). The role of RPA is to direct assembly and disassembly of the multi-protein complexes needed for DNA replication, repair and recombination(37,100). RPA can perform these actions by virtue of having multiple OB-folds, which individually release and rebind ssDNA without causing the entire complex to dissociate(37,124). As a result, RPA can diffuse along ssDNA to melt DNA structure or provide sites to load or unload partner

67 proteins(101,125,126). A similar dynamic binding mechanism could also enable CST to melt DNA structures at GC-rich regions or load Pol α for C-strand fill-in.

To clarify the mechanism of CST action, we have performed a series of DNA binding studies using substrates that mimic structures commonly found at telomeres or GC-rich regions. Our analysis uncovered ss-dsDNA junction binding and G4 melting activities that are likely to underlie specific aspects of CST function in telomere replication and genome-wide resolution of replication stress. Moreover, smFRET studies provide support for a dynamic binding mechanism that may provide a general foundation for many aspects of CST action.

4.3 RESULTS

4.3.1 CST recognizes ss-dsDNA junctions but does not melt extended stretches of duplex DNA

To better understand the role of CST in telomeric C-strand fill-in, we set out to examine CST binding to substrates mimicking the junction between the telomeric dsDNA and the 3’ overhang. During telomere replication, extension of the G-strand by telomerase occurs processively(7,127). However, the complimentary C-strand is synthesized more gradually(7) by repeated addition of DNA segments to the 5’ terminus in a process similar Okazaki fragment synthesis. It is not known how DNA polymerase is positioned to generate each new DNA fragment, but one possible mechanism is that CST recognizes the ss-dsDNA junction to direct DNA polymerase to the adjacent region on the overhang.

To explore whether CST recognizes ss-dsDNA junctions, we performed in vitro binding studies using recombinant CST and a mock telomere substrate that harbored both double and single-stranded telomeric DNA. CST was purified from insect cells (supplementary Fig. 1a) that had been co-infected with baculovirus encoding CTC1, STN1 and TEN1(59). The mock telomere substrate was generated from a single (fold-back) oligonucleotide that self-hybridized to form 15 nt dsDNA and an 18 nt 3’ telomeric

G-strand overhang (Fig. 1a, Table 1). We also generated control ss-dsDNA junction substrates that consisted of all non-telomeric sequence or contained telomeric sequence only in the ssDNA. Individual

68 substrates were incubated with various concentrations of CST and binding was analyzed by electrophoretic gel mobility shift assay (EMSA).

The analysis revealed that CST bound not only the mock telomere substrate but also the non- telomeric and partial telomeric substrates with equal efficiency. This result was unexpected because although CST binding to long (35-50 nt) ssDNA substrates is sequence independent, substrates in the 18-

30 nt range are only recognized if they resemble telomeric G-strand sequence(56,59,60) (supplementary

Fig. 1b). We next examined whether CST recognizes a fully non-telomeric junction substrate that had only a 10 nt 3’ overhang. Interestingly, CST also bound this DNA (Fig. 1b) although with lower efficiency than the substrate with the 18 nt overhang. This result was surprising because CST is unable to bind fully single-stranded DNA ≤12 nt in length even if it consists of TTAGGG repeats(56). Since CST is also unable to bind dsDNA(56,59,60) (supplementary Fig. 1c), our results suggested that the ss-dsDNA junction stabilized binding.

Unlike telomeres, DNA replication and repair intermediates often consist of partial duplex DNA with a 5’ overhang. We therefore asked whether CST exhibits a preference for junction substrates with 3’ versus 5’ overhangs. We found CST bound substrates with either overhang orientation with similar efficiency (Fig. 1a). This result was again unexpected as CST binds ssDNA with 5’ to 3’ directionality with the highest affinity binding sites positioned towards the DNA 3’ end(59). Overall, our results suggested that CST preferentially binds ss-dsDNA junctions regardless of sequence or DNA orientation.

However as all the junction substrates tested were formed from a single fold-back oligonucleotide, an alternative possibility was that, like RPA(128,129), CST has helix destabilizing activity and hence is able to melt the DNA duplex to give a ssDNA substrate of 53 nt. This would provide sufficient length of non- telomeric ssDNA for CST to bind.

To test if CST can destabilize and melt dsDNA, we performed a strand-melting assay using a junction substrate formed from two separate 32P-labeled oligonucleotides. The substrate consisted of 32 nt and 22 nt non-telomeric oligonucleotides annealed to form a 22 bp duplex with a 10 nt 3’ overhang (Fig. b-c, Table 1). This substrate is comparable to those that can be destabilized and melted by RPA(128). To

69 test for strand melting, the DNA was incubated with 20 nM CST for 30 minutes at RT, SDS was then added to denature the protein and samples were analyzed in native acrylamide gels (Fig. 1b).

Figure 1. CST binds to ss-dsDNA junctions without melting the DNA duplex

(a) EMSAs showing CST binding to junction substrates with an 18 nt overhang. Substrates were generated using fold-back oligonucleotides with: (i) Telomeric sequence at junction, (ii) mixed telomeric and non-telomeric sequence, (iii)-(iv) non-telomeric sequence with 3’ overhang (iii) or 5’ overhang (iv).

Black lines represent non-telomeric DNA, grey lines indicate telomeric sequence. Reactions contained the indicated concentrations of CST and 0.1 nM labeled DNA. (b) EMSAs showing CST binding to non- telomeric junction substrates with a 10 nt overhang. (i) Junction substrate was formed from a fold-back oligonucleotide or (ii) by annealing 32 and 22 nt oligonucleotides. (iii) Binding to 32 nt oligonucleotide alone. (c) Strand-melting assays with junction substrate formed by annealing 32 and 22 nt non-telomeric

70 oligonucleotides. Boiled: samples were boiled to melt DNA just prior to gel loading. Expected positions of partial duplex and ssDNA are shown to the left. All experiments were repeated with three independent protein preparations.

The assay provided no evidence for strand melting as the amount of annealed substrate remained unchanged and the amount of unannealed 32 nt or 22 nt DNA did not increase after CST addition (Fig.

1c). To control for possible re-annealing of separated DNA strands during gel loading, some reactions were terminated by the addition of both SDS and an excess of unlabeled 22 nt ssDNA to serve as a trap for any free 32P-labeled 32 nt oligonucleotide (this 22 nt DNA is not bound by CST (supplementary Fig.

1b). However, there was still no evidence of strand melting as again there was no increase in the amount of unannealed 32P-labeled 22 nt ssDNA. The lack of strand-melting activity did not reflect failure of CST to bind the annealed substrate because gel shift assays indicated that CST bound the substrate formed from the two separate oligonucleotides as efficiently as the fold-back substrate with the 10 nt overhang

(Fig. 1b). It therefore appears that CST is unable to melt extended stretches of duplex DNA. In this respect, CST differs from RPA, which exhibits robust helix destabilizing/strand-melting activity(128,129).

To verify that CST recognizes ss-dsDNA junctions but lacks the ability to melt DNA duplex we turned to a single molecule FRET (smFRET) assay. We prepared non-telomeric partial duplex FRET substrate with an 18 nt 3’ overhang by annealing a 3’ Cy3 (donor dye)-labeled 36 nt oligonucleotide with an 18 nt 5’ Cy5 (acceptor dye)-labeled oligonucleotide(130) (Fig. 2a). The substrate was then anchored to neutravidin-coated slides through a 3’ biotin located on the 18 nt oligonucleotide. The conformational state of the DNA was then monitored by smFRET using a prism-type total-internal-reflection configured microscope. The FRET efficiency (E) between Cy3 and Cy5 dyes can report the conformation of ssDNA before and after protein binding. FRET efficiency data were collected from >4,000 individual molecules obtained from from approximately 15 fields of view and plotted as a FRET histogram (Fig. 2b). As expected, a high FRET signal was observed in the absence of CST due to the flexibility of the 18 nt

71 ssDNA overhang bringing the Cy3 and Cy5 labels into close proximity. To examine CST binding activity, the protein was added to the immobilized DNA for 10 min and then removed by flushing the slide with imaging buffer prior to data aquisition. Data analysis reavealed that CST addition resulted in a

FRET efficiency switch from ~0.72 to ~0.12, indicating an increased time-averaged distance between

Cy3 and Cy5 dyes as a result of protein binding.

Figure 2. Single molecule FRET showing CST binds to ss-dsDNA junctions

(a) Cartoon showing DNA substrate design and anticipated FRET signals in the presence or absence of

CST. If CST binds without melting the anchoring DNA duplex, the high FRET signal will be lost but emission from the Cy3 donor (green) will be retained. If CST melts the DNA duplex, the Cy3 donor signal will also be lost. (b) FRET histograms generated from FRET measurements of >4,000 individual molecules. Left: DNA alone, right: DNA + 2 nM CST (c) Representative smFRET real-time trace showing change in FRET (bottom) and individual Cy3 and Cy5 signals (top) with time.

We next asked whether the shift in FRET signal was due to CST destabilizing and melting the DNA duplex leading to loss of the Cy3-labeled oligonucleotide from the slide (Fig 2a). To look for loss of the

Cy3-labeled DNA, we performed a time course analysis to monitor the change in Cy3 and Cy5 signals for individual substrate molecules over time right after CST addition (Fig. 2c). These single-molecule real- time courses showed retention of the Cy3 oligonucleotide with the Cy3 and Cy5 signals exhibiting an anti-corrolated FRET change upon protein binding. These results confirm that CST binds to non- telomeric ss-dsDNA junctions without extensive destabilization of the associated DNA dupex.

It remains possible that CST can destabilize a few base pairs of dsDNA at the junction region as such limited strand melting would not have been detected by the above experiments. However, generation of a few extra nt of ssDNA as a result of limted strand melting cannot explain why CST binds efficiently to junction substrates with a 10-18 nt overhang but binds poorly, or not at all, to ssDNA in the ≤32 nt (Fig

1b and supplementary Fig. 1b). Thus, the above experiments indicate that CST specifically recognizes the ss-dsDNA junction in addition to the adjacent tract of ssDNA. This junction binding activity may be important for CST to engage proteins such as DNA polymerase near primer template junctions during telomere replication and genome-wide resolution of replication stress.

4.3.2 CST can bind and unfold G4 structures

Regions of the genome harboring GC-rich repeats appear to act as barriers to DNA replication due to the single-stranded template DNA forming G-quadruplex (G4) structures which block DNA polymerase(83,131,132). Since CST binds preferentially to GC-rich DNA(56,58-60) and aids in replication through GC-rich regions of the genome(54,58), we next asked whether CST can bind and unfold G4 DNA. To test for G4 binding, we performed EMSAs using a selection of four repeat telomeric oligonucleotides with well characterized G4-forming potential(130). Substrate oligonucleotides were designed to have 0-6 nt ssDNA located 5’ and/or 3’ of the G4 structure (Table 1). The binding analysis revealed that CST bound all of the G4 substrates regardless of whether they had ssDNA 3’ or 5’ of the G4 structure (Fig. 3a, 3G4, G5, 3G46)). However, the substrate that contained only the G4 structure and no 5’

73 or 3’ ssDNA (G4) was bound somewhat less efficiently, suggesting that a short stretch of ssDNA enhances binding. Our findings imply that CST can either unfold a G4 structure to bind the resulting ssDNA, or CST can bind a G4 structure without unfolding it. The later would be unexpected as OB folds usually bind single-stranded rather than base-paired DNA/ RNA(61,133).

Figure 3. CST binds and unfolds G4 DNA

(a) EMSAs showing CST can bind to telomeric DNA oligonucleotides that form G4 structures. Substrates had 3’ ssDNA (3G4), 3’ or 5’ ssDNA (G5 and 3G46) or no 3’ or 5’ ssDNA (G4). Binding reactions contained 10 nM CST, 0.1 nM DNA in 150 mM NaCl, 3 mM MgCl2. (b) Schematic of the G4 smFRET substrates showing expected FRET signals with/without G4 unfolding. (c-e) Representative FRET

Histograms showing CST can bind and unfold G4 DNA. Top, DNA alone; bottom, DNA +2 nM CST.

CST was added for 10 min, then excess protein was washed out prior to FRET measurement. (c-d)

Unfolding of 3G4 and 3G46 in 150 mM NaCl + 3 mM MgCl2. (e) Unfolding of 3G4 in 100 mM KCl + 3 mM MgCl2. (f) Representative smFRET real-time trace showing a decrease in FRET after CST binding.

Top, Cy3 and Cy5 signals showing complementary transition upon CST binding. Bottom, FRET signal.

Measurement was performed in the presence of 2 nM CST (no washout).

To determine if CST can unfold G4 DNA, we again turned to smFRET. The setup was as described above (Fig 2a), except the Cy3-labeled oligonucleotide harbored 4 telomeric repeats adjacent to the region annealed to the Cy5-labeled anchor oligonucleotide (Fig. 3b). The Cy3 and Cy5 labels were positioned on either side of the telomeric repeats so that unfolding of G4 DNA would induce a decrease in

FRET efficiency. Two different Cy3-labeled substrates were tested: 3G4-Cy3 which terminated with a G4 and 3G46–Cy3 which had an additional 6 nt of non-telomeric ssDNA at the 3’ end (Fig 3c-d, Table 1). To ensure G4 stability, the buffers used during slide preparation, CST binding and data acquisition contained

150 mM NaCl. In initial experiments, FRET signals from >4,000 individual molecules were quantified and plotted as a FRET histogram. In the absence of CST, we observed high FRET signals characteristic of

G4 DNA (Fig. 3c). To confirm G4 formation, the 150 mM NaCl was replaced with 0 mM NaCl to destabilize the G4(130) (Supplementary Fig. 2a).This caused a decrease in FRET efficiency, as expected for G4 unfolding, with appearance of an intermediate FRET signal characteristic of flexible ssDNA.

To examine the effect of CST binding, protein was added to the flow chamber and allowed to bind the immobilized DNA for 10 min, and then excess CST was flushed out with binding buffer

75 followed by imaging buffer. Imaging of the slides revealed that addition of CST to either the 3G4-Cy3 or the 3G46–Cy3 substrate caused almost complete loss of the high FRET signal (Fig.3c-d). This result indicates CST can unfold G4 DNA and bind the resulting ssDNA. The G4 unfolding and DNA binding appeared to be stable as the loss of high FRET signal persisted throughout the 10-20 min time period taken to acquire sufficient images to build the FRET histograms. Similar results were obtained when the experiment was repeated using buffers containing 100 mM KCl + 3 mM MgCl2 instead of 150 mM NaCl, conditions which stabilize G4 and favor formation of a mix of parallel and antiparallel structures (Fig.

3e)(134). We therefore conclude that CST efficiently unfolds G4 DNA. Given this capability, we surmise that CST aids in recovery from replication stress by removing G4 structures that form during replication through GC-rich regions of the genome.

To further characterize the G4 unfolding process, we performed a real-time analysis to examine the change in FRET signals from individual DNA molecules with time after CST addition (Fig. 3f). The resulting 90 sec real-time traces revealed a sharp one-step FRET decrease from the high FRET to the low

FRET state. Thus, a single CST binding event causes rapid and complete G4 unfolding. The rapid one step change in FRET efficiency is similar to that observed when RPA or the C. glaubrata Cdc13-Stn1-

Ten1 complex unfold G4 structures(115,135) but it differs from the mechanism of G4 unfolding by POT1 which occurs in four steps and requires two POT1 molecules(130). The four FRET transitions seen with

POT1 reflect sequential binding of the 4 OB folds (two from each POT1 molecule) to cause gradual G4 unfolding. The more efficient G4 unfolding by human CST, CgCST and RPA may reflect the larger number OB folds in these complexes and/or a smoother DNA binding trajectory(59,99,115,136).

4.3.3 CST exhibits facilitated displacement

While analyzing real-time traces showing the change in FRET from individual DNA molecules, we noticed that only 75% of the traces showed stable G4 unfolding and CST binding during the 90 sec time course. In contrast, the remaining 25% of traces showed reformation of the high FRET configuration

(Fig. 4a-b) indicating subsequent CST dissociation. This dissociation was observed for both the junction

76 substrate with the 18 nt non-telomeric 3’ overhang (Fig. 4a) and the 3G4-Cy3 substrate (Fig. 4b). In some traces, multiple binding and dissociation events were apparent during a single time course. The frequent dissociation of CST from the two substrates was striking because filter-binding assays have previously shown that CST binding to telomeric and many non-telomeric substrates is very stable(56,59). For telomeric oligonucleotides, t½ for CST dissociation ranges from 4.5-8 hours for 18-48 nt substrates.

The discrepancy in CST binding stability revealed by the smFRET real-time analysis as compared to filter-binding assays reminded us of findings with RPA where the stability of RPA binding to ssDNA depends on the concentration of excess unbound RPA(126) (see below). In our smFRET experiments, unbound CST was present during acquisition of the real-time data. In contrast, unbound

CST was essentially absent from the filter-binding experiments used to measure CST dissociation rates because an excess of cold competitor DNA was added to samples following the initial binding reaction(56,59). This difference in amount of unbound CST present in the two experiments, suggested to us that the rapid dissociation of CST seen in the smFRET real-time analysis might be directly related to the excess CST present in the flow chamber during data acquisition.

To test whether CST dissociation depends on the concentration of free protein, we obtained real- time traces from slides harboring three separate flow chambers that each contained a different amount of free CST during data acquisition. Following immobilization of the 3G4 DNA substrate, 2 nM CST added to each chamber for 1 min. The chambers were then washed with imaging buffer containing 0 nM, 2 nM or 5 nM CST and 90 sec real-time traces were captured from each portion of the slide. The result was striking as the fraction of traces showing one or more CST dissociation and G4 refolding events was directly proportional to the CST concentration present during data acquisition, with the most dissociation observed at the highest CST concentration (Fig. 4c). Correspondingly, the dwell time for CST association with the DNA was inversely proportional to protein concentration, with shorter dwell times seen as CST concentration increased (Fig. 4d). Based on these observations, we conclude that the presence of excess unbound CST increases the likelihood that a bound CST complex will dissociate, i.e. unbound CST facilitates the displacement of bound CST.

Figure 4. Dissociation of CST from DNA is concentration dependent

(a-b) Representative smFRET real-time traces showing CST dissociation from junction substrate with 18 nt overhang (a) or 3G4 G4-forming substrate (b). PB, photobleaching. (c) Fraction of traces showing one or more dissociation events after CST removal (0 nM), in the continued presence of 2 nM CST, or after

78 addition of 5 nM CST (n=3 independent experiments ± S.E.M.). (d) Dwell times for CST binding before dissociation. Traces analyzed as were in (c). (e-g) Models illustrating how unbound CST could cause facilitated displacement of bound CST (e), how junction recognition and dynamic binding could lead to

Pol α loading on the G-strand overhang to achieve C-strand fill-in (f), or G4 unfolding due to CST binding. Oblongs represent individual OB folds in CTC1, STN1 and TEN1 (g).

This facilitated displacement of CST is reminiscent of the facilitated exchange that has been observed with RPA(126). Although RPA binds tightly to ssDNA, it dissociates rapidly when free RPA is present thus allowing another RPA molecule to bind. This concentration-dependent protein turnover is thought to reflect the dynamic mechanism of RPA binding where individual OB folds undergo microscopic dissociation from the DNA(37,101). Dissociation of individual OB folds leads to macroscopic RPA dissociation only if free RPA is available to occupy the exposed ssDNA. Like RPA,

CST harbors multiple ssDNA binding sites including the OB-fold in STN1 and likely some of the 5-6 predicted OB-folds in CTC1(59,62). We therefore propose that CST also binds DNA dynamically and the observed concentration-dependent self-displacement of CST is caused by microscopic dissociation of individual DNA binding sites/OB folds from the DNA substrate. As with RPA, the microscopic dissociation would allow a second CST complex (or another protein) to initiate binding. In some cases, the outcome would be unstable binding and dissociation of both complexes from the DNA (Fig. 4e). We term this process “facilitated displacement”. In our experiments, this displacement would be observed as reappearance of the high FRET signal.

It may be that CST also undergoes facilitated exchange where a bound CST complex dissociates and is simultaneously replaced by a second complex. We would not expect to detect these events with our experimental set up as the DNA would not be fully released so the high FRET signal would not reappear. The ability of RPA to undergo facilitated exchange is thought to be extremely important for

RPA function because it provides a mechanism to load downstream ssDNA binding proteins during DNA replication and repair(37,124,126). We suggest that facilitated displacement and possibly facilitated

79 exchange also underlie CST function as it would allow CST to load proteins needed to resolve various forms of replication stress (Fig. 4f).

4.4 DISCUSSION

Here we demonstrate that human CST exhibits a series of DNA binding activities that are directly relevant to the various roles of CST genome-wide. These activities include specific recognition of ss- dsDNA junctions, removal of G4 structure and facilitated displacement of protein molecules from a DNA substrate. Together or individually these activities provide mechanisms for CST to enable DNA replication through GC-rich sequence, regulate C-strand synthesis at telomeres, and support priming of

DNA synthesis down-stream of DNA lesions and possibly at late-firing or dormant origins(53,97,98). Our finding that free CST can promote the release of bound CST from ssDNA is particularly interesting because it provides evidence for dynamic DNA binding through dissociation and re-association of individual DNA-binding domains/OB-folds. The dynamic binding is likely to provide a mechanism for engaging partner proteins such as Pol α by exposing stretches of ssDNA to nucleate protein association.

It is striking that CST and RPA share many of the same DNA binding activities despite having distinct complex architecture(59) and quite separate roles in DNA replication(54,137). Given that CST is used to resolve replication-related issues that are guaranteed to arise each cell cycle (e.g. telomeric C- strand fill-in) it may be that cells have evolved this second multi-OB fold ssDNA binding complex to avoid the complication of ATR activation by RPA-coated ssDNA(37,138). For example, CST binding to the G-overhang after telomerase extension prevents RPA association and ATR activation(57) while simultaneously providing a mechanism to engage DNA polymerase for C-strand fill-in. Likewise, CST binding at sites of G4 structure could provide a way to remove the structure and rapidly reinitiate DNA synthesis without affecting the overall replication program through ATR activation.

While relatively few CST interactions partners have been identified thus far (only Pol α, TPP1 and Rad51)(55,58,121), the large size of CTC1 suggests there are likely to be many others. Hence, the

80 dynamic nature of CST binding to ssDNA may well allow CST to regulate the DNA-association of an array of different proteins depending on the replication issue to be resolved. Currently it is unclear whether CST regulation is restricted to protein displacement or if CST can also load proteins on ssDNA.

Additional studies are required to clarify this point. Further studies are also needed to determine whether

CST resembles RPA (101) in being able to diffuse along ssDNA. However, such work is best performed with fluorescently labeled protein (101,126) and it has not yet been possible to prepare preparations of labeled CST with good DNA binding activity (data not shown).

The combined ability of CST to recognize ss-dsDNA junctions and bind DNA dynamically provide a ready explanation for why the process of telomeric C-strand fill-in occurs via incremental extension of the DNA 5’ terminus rather than a single primed DNA synthesis reaction (Fig. 4f). While

CST appears to coat the newly extended overhang(56,57), it may be that only the complex adjacent to the ss-dsDNA junction is capable of engaging DNA polymerase for C-strand synthesis. This might be because junction recognition somehow alters CST binding dynamics to favor partial CST dissociation from the ssDNA with concomitant DNA polymerase loading. Upon synthesis of one segment of C-strand, a new ss-dsDNA junction would be generated and the adjacent CST complex would become competent to engage DNA polymerase to synthesize the next segment of C-strand DNA. It is possible that the junction recognition on partial duplex DNA also favors loading of CST binding partners at DNA lesions and replication blocks elsewhere in the genome.

The G4-unfolding activity of CST is likely to be important for CST function both during C-strand fill-in and conventional replication of GC-rich dsDNA. At the time of C-strand fill-in, the elongated 3’ overhang can form G4 structures which must be removed to prevent them from impeding DNA synthesis by Pol α. During replication of dsDNA, G4 structures also need to be removed to prevent obstruction of

DNA polymerase(132) (Fig. 4g). It is now apparent that in addition to CST, cells harbor a plethora of factors capable of resolving G4 DNA (e.g. RPA, POT1, BLM, WRN, RTEL and other helicases)(83,139,140), raising the question as to why so many apparently redundant activities are needed.

Presumably each activity has specific advantages and disadvantages. We suggest that CST is well suited

81 for efficient G4 removal in situations where RPA-mediated ATR activation is undesirable and helicases are unable to act. Such a situation may occur during leading strand replication through GC-rich regions. If the leading strand polymerase is blocked by G4 DNA, there may not be room to load BLM or WRN upstream of the G4 and, as these helicases only translocate in a 3’ to 5’ direction, it would not be useful to load them down stream of the G4(83,141). In this situation, binding of CST may be an efficient way to remove the G4 without causing ATR activation. Moreover, if the G4 causes the leading strand polymerase to disengage, CST may enable a new polymerase to load and re-prime the leading strand (Fig.

4g). At telomeres, G4 structures are likely to form on the lagging strand template. In this situation BLM,

WRN and POT1 may all contribute to G4 removal (83,130,141). However, CST unfolds G4 structures more efficiently than POT1, which could explain why the presence of CST makes replication through the telomere duplex more efficient.

Mutations in the CTC1 and STN1 subunits of CST cause a severe disease called Coats plus(93-

95). Patients suffer from pleiotropic symptoms that include neurological disorders due to brain calcifications, GI and retinal bleeding, and bone marrow failure. In some cases, symptoms overlap those seen in patients with the short telomere disorder Dyskeratosis congenita(93). The patient mutations are always biallelic and they result in partial loss of CST function. It is striking that many of the CTC1 mutations lie in the predicted OB fold domains (59,110). This positioning suggests that the mutations are likely to affect DNA binding activities such as junction recognition, G4 unfolding or overall binding dynamics. If this is the case, loss of the CST-based pathway to resolve replication issues may require use of alternative, less benign pathways that result in cellular damage and hence the clinical symptoms of

Coats plus. Understanding which activities of CST are altered by specific CTC1 mutations could lead to better disease management in Coats plus patients.

4.5 Supplementary Figures

Supplemental Figure 1. CST binds to ss-dsDNA junctions

(a) Silver stained polyacrylamide gel (15% on the bottom, 12% on top) showing co-purified CST subunits. (b-c) EMSAs showing CST binding to various substrates. Reactions contained the indicated amounts of CST and 0.1 nM DNA. (b) EMSAs with18 nt, 22 nt, 26 nt or 32 nt non-telomeric ssDNA to determine minimum substrate length required for CST binding. (c) EMSA showing lack of CST binding to dsDNA corresponding to duplex region of the non-telomeric fold-back substrates.

Supplementary Figure 2. smFRET controls

(a) FRET histogram demonstrating the sensitivity of G4-forming substrate to salt concentration.

Anchored 3G4-Cy3 was incubated with the indicated concentrations of NaCl prior to data acquisition. (b)

FRET histogram showing that BSA cannot disrupt G4 DNA; top, 3G4 alone; bottom, 3G4 + 0.1 mg/ml

BSA.

Table: Oligonucleotides used for EMSA, Strand-melting assay and smFRET

Underline indicates regions of dsDNA

EMSA 5’3’

Non-Tel 18 AGCGTATCCGTTCAGTTG

Telomere fold-back CTAACCGCATCTAGCTTTTTGCTAGATGCGGTTAGGGTTAGGGTTAGGGTTAG (3’ overhang)

Mixed fold-back ACGACTGCATCTAGCTTTTTGCTAGATGCAGTCGTGGTTAGGGTTAGGGTTAG (3’ overhang)

Non-Tel fold-back ACGACTGCATCTAGCTTTTTGCTAGATGCAGTCGT (3’ overhang) AGCGTATCCGTTCAGTTG

Non-Tel fold-back AGCGTATCCGTTCAGTTG (5’ overhang) ACGACTGCATCTAGCTTTTTGCTAGATGCAGTCGT

Non-Tel fold-back ACGACTGCATCTAGCTTTTTGCTAGATGCAGTCGT (dsDNA)

EMSA & strand- 5’3’ melting assay

Non-Tel 22 GTCGATCTGAGTCACTGAGTAC

Non-Tel 26 GTACTCAGTGACTCAGATCGACAGCG

Non-Tel 32 GTACTCAGTGACTCAGATCGACAGCGTATCCG

EMSA G4 5’3’

3G4 TTAGGGTTAGGGTTAGGGTTAGGG

3G5 TTAGGGTTAGGGTTAGGGTTAGGGTTAGGG

3G46 TTAGGGTTAGGGTTAGGGTTAGGGAGCGTA

G4 GGGTTAGGGTTAGGGTTAGGG

FRET 5’3’

Cy5 18 nt anchor Cy5-GCCTCGCTGCCGTCGCCA-Bio

Cy3 10 nt overhang TGGCGACGGCAGCGAGGCAGCGTATCCG-Cy3

Cy3 3G4 TGGCGACGGCAGCGAGGCTTAGGGTTAGGGTTAGGGTTAGGG-Cy3

Cy3 3G46 TGGCGACGGCAGCGAGGCTTAGGGTTAGGGTTAGGGTTAGGGAGCGTA-Cy3

5. Importance of CST as a complex

5.1 Introduction: To better understand CST function in human cells, our lab has generated conditional

CTC1 or TEN1 knockout cells to investigate if individual subunits play specific roles in facilitating discrete steps in replication. We have found that CTC1 knockout cells show gradual elongation of the G- overhang due to elongation of the G-strand by telomerase. They also show a deficiency in C-strand fill- in. These findings indicate that CTC1 is important both for terminating the telomerase activity and facilitating C-strand fill-in. It likely functions in both reactions by engaging DNA polymerase-α. The lack of maintenance of the C-strand leads to gradual shortening of overall telomere length, similar to that observed in cells lacking telomerase. The telomere shortening indicates C-strand fill-in is as important as telomeric G-strand extension by telomerase in maintaining telomere length. The increase in length of the single-stranded G-overhang leads to accumulation of RPA, which then triggers DNA damage signaling

(ɤH2AX accumulation) at the telomeres. Interestingly, TEN1 knock-out cells show only a modest increase in G-overhang length: ~2x compared to wild type cells, similar to STN1 knock-down cells. Moreover this increase in length arises specifically from the lack of C-strand fill-in. We do not see any net increase in the absolute length of G-strand in a qFISH assay. This lack of G-strand elongation indicates that dimeric

CS complex alone is sufficient for telomerase displacement but the whole complex is required for C- strand fill-in synthesis. There are two obvious and testable possibilities for the effect of the TEN1 K/O.

First, lack of TEN1 could cause a defect in interaction with DNA polymerase α, which could in turn explain the problem with C-strand fill-in. Second, there could be a defect in DNA binding of the remaining dimeric CS complex. The first possibility is negated as our lab has also shown that CS alone is sufficient to interact with DNA polymerase α in an in-vitro co-immunoprecipitation assay. To test if the effect of TEN1 knockout on G-overhang maintenance comes from a difference in DNA binding in the absence of either CTC1 or TEN1, I performed several different DNA binding assays. I did not observe a difference in binding to any ssDNA, G-quadruplex or telomeric and non-telomeric junction substrates in the absence of the TEN1 subunit in electrophoretic mobility shift assays (EMSA). However, smFRET assay failed to detect binding/ stable binding of CS to either junction substrate or G4 tracts. This

86 discrepancy between the EMSA and the smFRET data can be explained by a difference in the DNA binding configurations of the CS and CST complex. In contrast, in the absence of CTC1, the dimeric ST complex does not bind to any of these substrates in a stable manner. These data are consistent with chromatin immunoprecipitation (ChIP) results performed in the lab (data not shown) that shows STN1 localization to telomeres is decreased in the absence of CTC1 but not in the absence of TEN1.

5.2 Results:

5.2.1 The CTC1-STN1 (CS) complex binds DNA with similar affinity to CST but the affinity of STN1-

TEN1 (ST) complex is lower

We examined how the absence of either CTC1 (ST) or TEN1 (CS) affects the ability of CST to bind a range of DNA substrates. As the affinity of CST for short versus long substrates seems to depend on

DNA sequence (56), we monitored binding of CST, CS and ST to telomeric and non-telomeric oligonucleotides (Fig.1, A-B). When we used electrophoretic mobility shift assays (EMSA) to compare binding of CST(WT), CS and ST to telomeric and non-telomeric single-stranded oligonucleotides, CST and CS bound to DNA with similar affinity whereas binding of ST to the DNA appeared as a smear / faint band at a similar concentration of protein, indicating a significant reduction in binding. This finding indicates that in the absence of CTC1, the complex cannot bind to DNA in a stable manner, but that the absence of TEN1 does not affect binding to ssDNA. Next, we examined if the binding of CST to the ss- dsDNA junctions is disrupted in the absence of TEN1 or CTC1 as we think this binding activity could be important for C-strand fill-in synthesis. To test for possible differences in binding to the junction substrate compared to CST, we used EMSA assays to compare binding of CS versus CST to a non-telomeric fold- back oligonucleotide with an 18nt overhang and a partial double strand oligo with a 10 nucleotide telomeric overhang and telomeric sequence in the junction (Fig. 1, C-D). The overhang lengths were chosen to be shorter than the length of ssDNA of the same sequence normally recognized by. Again we observed no difference in binding of CS and CST to either of these substrates but ST again showed a smeary appearance in the gel. The similarity in DNA binding of CS and CST indicates either the defects in C-strand fill-in in the CTC1 and TEN1 knockout cells are for different reasons or that DNA binding is

87 not sufficient to facilitate the C-strand fill-in. Another possibility would be that the whole CST complex is needed to melt G-quadruplex structures which are abundant in the telomeric overhang and these could get in the way of the C-strand fill-in. To test binding to a G4 tract, we performed an EMSA assay with

CST, CS and ST using a telomeric four repeat oligonucleotide. The oligonucleotide was first incubated in 150mM NaCl, 3mM MgCl2 to facilitate G4 formation. Again both CST and CS bound to this substrate and ST gave a smeary appearance (Fig 1, E). All these results are consistent with the ChIP data confirming that absence of TEN1 does not affect binding or localization of CS to telomeres. We concluded that CTC1 is important and CS is sufficient for binding to any DNA substrates that CST as a complex can normally bind to. We were not able to purify CTC1 alone (CTC1 is unstable in the absence of STN1 and TEN1) but previously it has been shown that CTC1 alone is not sufficient to bind to single stranded DNA or for interaction with DNA polymerase α (110).

Figure 1: CST and CS binds to ssDNA and ss-dsDNA junctions but not ST alone. (A-B) CST and

CS but not ST binds to both telomeric (A) and non-telomeric (B) ssDNA with equal efficiency. (C) CST and CS but not ST binds to non-telomeric junction substrate. (D) CST and CS binds to telomeric junction with a 10nt overhang with equal efficiency (top); EMSA with a telomeric 10nt oligonucleotide used as a control (bottom). ST does not bind to any of these substrates. (E) CST and CS but not ST binds to G4 structure DNA. In each case 0.1nM oligonucleotide was incubated with indicated amount of protein at room temperature for 30 minutes before running in a 0.7% agarose gel.

5.2.2 Absence of the TEN1 subunit changes the DNA binding architecture of the CTC1-STN1 complex

As EMSA can only detect binding and cannot provide any information about changes in the DNA conformation upon protein binding, we analyzed DNA conformation using smFRET assays as described previously. We used both an 18nt overhang junction substrate and a G4 forming substrate. We prepared partial duplex FRET substrate with an 18 nt 3’ overhang or a telomeric 4 repeat sequence oligonucleotide by annealing 3’ Cy3 (donor dye)-labeled oligonucleotide with an 18 nt 5’ Cy5 (acceptor dye)-labeled oligonucleotide(130). The substrate was then anchored to neutravidin-coated slides through a 3’ biotin located on the 18 nt oligonucleotide. The conformational state of the DNA was then monitored by smFRET using a prism-type total-internal-reflection configured microscope. The FRET efficiency (E) between Cy3 and Cy5 dyes can report the conformation of ssDNA before and after protein binding. FRET efficiency data were collected from >4,000 individual molecules obtained from from approximately 15 fields of view and plotted as a FRET histogram. As expected, a high FRET signal was observed in the absence of CST due to the flexibility of the 18 nt ssDNA overhang or from the ordered G4 structure bringing the Cy3 and Cy5 labels into close proximity. To examine CST binding activity, the protein was added to the immobilized DNA for 10 min and then removed by flushing the slide with imaging buffer prior to data aquisition. We detected a high FRET peak in the histogram in the absence of protein. Upon addition of CST, the FRET peak reduced to 0.0-0.1 as seen previously. However, upon addition of either

CS or ST there was no change in the high FRET peak (Fig. 2). This indicated that CS or ST either cannot bind at all or bind unstably to both the junction substrate and the G4 substrate.

A possible explanation for the conflicting results obtained with CTC1-STN1 in the EMSAs and the smFRET assay using the ds-ssDNA junction substrate and G4 substrates could be that these two approaches are designed to detect different aspects of protein-DNA binding. EMSAs detect the direct binding affinity and show a molecular weight shift in electrophoresis no matter how the DNA was bound, while smFRET can detect a more dynamic binding process/ binding architecture at a given point. The

FRET signals are given by the distance between the two fluorophores at the ends of two oligonucleotides, so that smFRET is able to detect if the ssDNA is completely stretched out upon binding of the protein.

The binding of CS seems to be stable as we get sharp bands in EMSA in a situation where the binding reaction was much longer than smFRET. Therefore, the absence of a loss in the FRET efficiency shift for

CTC1-STN1 may derive from binding only to the ss-dsDNA junction so that only part of the DNA is covered. In contrast, CST can totally cover the DNA to stretch the fluorophores far apart. This inappropriate binding in absence of TEN1 might cause loss of loading or activation of Polα at the ds-ss telomere DNA junction structure, resulting in the failure of C-strand fill-in.

Figure 2: Single molecule FRET showing CST binds to ss-dsDNA junctions and G4 substrate but

CS and ST do not (A-B) FRET histograms generated from FRET measurements of >4,000 individual molecules. Top: DNA alone, Junction substrate with 18nt NonTel overhang (A), G4 substrate (150mM

NaCl +3mM MgCl2) (B); Bottom: DNA + BSA/ 2 nM CST/ 2 nM CS/ 2 nM ST (from left to right). CST was added for 10 min, and then excess protein was washed out prior to FRET measurement.

5.3 Discussion: Experiments performed by other lab members have shown that CST as a complex is not needed to terminate telomerase activity, but that CS is sufficient. Also CS alone can localize and bind to telomeres (Price lab, data not shown). Moreover, CS can bind to nontelomeric DNA efficiently, but ST binds to DNA with much lower efficiency. The efficient DNA binding by CS but not ST, explains why certain telomeric and non-telomeric defects are associated with the CTC1 knockout but are not affected by the TEN1 knockout. Nonetheless we still saw a defect in C-strand fill-in upon depletion of TEN1. This functioning indicates CST as a complex is necessary for C-strand fill-in. Because CS alone can bind to telomeric DNA and can also interact with DNA polymerase α, this result was puzzling. As our previous results suggested that an ss-ds junction binding activity is necessary for the incremental C-strand fill-in process, we monitored the junction binding activity of these sub-complexes. Although CS appeared to bind to the junction substrates with similar efficiently to CST by EMSA, our smFRET results suggests a differential binding architecture or stabilization in the presence of CST. Similar results were obtained when we monitored binding of CS to the G4 structure. Overall this result implies that TEN1 enables the

CST complex to bind telomeric ss-dsDNA junctions in a conformation necessary to engage or activate

DNA polymerase α for C-strand fill-in. ST did not appear to bind to any of these substrates indicating the importance of multi-OB folds present in CTC1 with regard to DNA binding.

Figure 3: Model for mechanism of function of CST at telomeres. CST and CS can both bind to telomeric overhang and terminate telomere elongation by telomerase. Knockdown of CTC1 leads to inefficient binding by ST alone and so the complex can no longer terminate telomerase activity. But binding of CST and CS to ss-dsDNA junctions differ in architecture such that CS cannot facilitate proper

C-strand fill-in.

6. Conclusions and future directions

6.1 Diverse roles of CST in resolution of replication issues

For many years, the perception of telomere biologists regarding capping complexes was that two distinct telomere capping complexes had evolved, shelterin in vertebrates and CST (Cdc13, Stn1, Ten1) in yeast

(1). The discovery of a separate ssDNA binding complex, CST (CTC1-STN1-TEN1) in Arabidopsis and humans, raised the possibility of a more complex telomere regulatory system in eukaryotes. Early insights into the roles of CST came from the original studies of AAF (DNA polymerase α accessory factor) and the apparent structural similarity to RPA, which together indicated roles in replication (60). Since then, we and others have shown that human CST plays diverse roles in replication (1,36,39,53,54,56-

58,65,86,98). At telomeres, it helps the replication machinery pass through the double strand region overcoming various replication barriers (54). In telomerase positive cells, studies suggest that CST might be important for terminating telomerase activity after elongation of G-strand by interacting with TPP1-

POT1 (56). After telomerase mediated G-strand extension, CST has been shown to be essential in facilitating complementary C-strand fill-in at late S/G2 phase. Originally the CST complex was identified as DNA polymerase α accessory factors (AAF) (50,123) which led to the hypothesis that CST helps in C- strand fill-in, presumably via interaction with DNA polymerase α. Evidence from yeast that Stn1 promotes DNA primase to polymerase switching and mediates polymerase α activity further supports this concept (98). Unpublished data from the Price lab now shows that dimeric CTC1-STN1 complex alone is sufficient to terminate telomerase activity. Interestingly however, the whole complex is needed to facilitate C-strand fill-in even though CTC1-STN1 alone can interact with DNA polymerase α (59). In addition to its telomeric roles, CST is important for non-telomeric replication by virtue of its capacity to rescue replication after fork stalling due to various endogenous and exogenous stresses (53,65). Depletion of CST subunits leads to accumulation of non-telomeric DNA damage signals and anaphase bridges as well as loss of sister chromatid cohesion, all of which indicates genomic instability in cells. Depletion of

CST subunits also leads to more cell death after hydroxyurea, camptothecin, aphidicolin or MMS

94 treatment, providing further evidence for the importance of CST in rescuing cells from replication stress.

One of the mechanisms by which CST helps cells to recover from fork stalling after exogenous replication stress is by promoting firing of late or dormant replication origin. Although the underlying mechanism of this late origin firing is unknown, this may also involve association of CST with DNA polymerase α. Although both C-strand fill-in at telomeres and genome-wide late origin firing may involve

CST mediated DNA polymerase α activation, they must involve different interaction partners/ mechanism of action. C-strand fill-in only takes place after telomerase mediated G-strand extension where the conventional DNA replication machinery is no longer available. In contrast, the late origin firing in duplex genomic DNA may involve conventional initiation/ re-initiation factors different from C-strand fill-in. How one complex can facilitate so many mechanistically different reactions associated with replication is intriguing. The broader disease phenotypes and greater disease severity in Coats plus patients compared to dyskeratosis congenita, indicates additional underlying causes of the diseased state.

Thus it seems likely that genome-wide instability caused by the point mutations in CTC1 and STN1 is responsible for the further complexity/ severity of the Coats plus patients. Hence, understanding the detailed mechanism by which CST regulates genomic stability and prevents telomere loss is extremely important with great therapeutic potentials.

The studies described in this thesis are our attempt to unravel the mechanism of action of CST related to various aspects of replication, both at telomeres and at non-telomeres genome-wide. To investigate the mechanism in further detail, we tried to understand the DNA binding properties of CST.

6.2 Delving into the mechanism of action of CST

Prior studies along with ours made it clear that CST binds preferentially to telomeric G-strand DNA and binding to the telomeric C-strand is unstable. These data are consistent with binding of CST to the telomeric G-overhang. Also CST has been shown to be enriched in GC-rich regions genome-wide.

Together these data suggest preference of CST towards binding guanine. The high structural similarity of

CST with RPA, especially in the DNA binding OB-fold domains, suggested that CST may resemble RPA in having multiple DNA binding modes involving varied numbers of OB-folds. It has been shown that these multiple DNA binding modes play a very important role in RPA function during DNA replication, repair and recombination. To dissect the importance of DNA binding properties of CST in its mechanism, we investigated the interaction of CST with various lengths, sequences and structures of oligonucleotides that CST might encounter during replication. We have shown that CST does not bind to dsDNA or a ssDNA shorter than 18 nt. CST binds efficiently to a 18 nt long oligonucleotides only with a telomeric sequence repeat but not to a scrambled non-telomeric sequence of same length. The shortest non- telomeric oligonucleotide that CST can bind to is 32 nt but it does so with a higher off-rate (unstable binding). As the oligonucleotide length gets longer (~48 nt), CST does not distinguish between telomeric vs. non-telomeric sequences, and binds to both with equal efficiency. Our DNA-protein crosslinking assay also indicates that the 3’ end of the ssDNA always lies closer to the junction of all three subunits of

CST and the 5’ end lies further away. This shows that the DNA binding architecture of CST is very different than RPA, despite their structural similarity. These differences in DNA binding along with their distinct roles in replication emphasize that CST is not just a redundant version of RPA.

Although RPA and CST bind to DNA with different architecture, we still cannot ignore their structural similarity; the involvement of multiple OB-folds in DNA binding leading to different DNA binding modes as well as the orientations of binding are very similar. To understand the specific functions of CST compared to RPA and the specific roles of CST in DNA replication, further experiments need to be done.

One obvious follow up experiment is to test if CST has a strong preference for guanines in the oligonucleotide, and what percentage of guanines are optimal for CST binding even in a scrambled sequence. Previous studies have shown that CST binding to telomeric C-strand, which has no guanine, is very weak. As mentioned before, genome-wide ChIP-seq data from previous studies have shown that

CST is enriched in GC rich regions. These studies, in conjunction with ours indicate that CST has a preference for guanines. This preferential binding of CST to guanine rich sequences is interesting as RPA

96 shows a preference for pyrimidine over purines, indicating one more difference with CST. On the same line of thought, one could also do a competition assay or a facilitated exchange assay with telomeric DNA for CST and RPA to dissect if CST has higher affinity for telomeric DNA. This could potentially explain evolution of CST to prevent RPA binding at telomeres and triggering damage signaling.

Last, but not least, we should consider the fact that sequence specificity of a protein motif should always be considered in conjunction with its functional aspects as there is no cutoff for biologically relevant affinity for a protein or a domain. A protein binds to varied DNA sequences with a continuum of affinities and should not be strictly regarded as “binding” or “not binding”. Therefore, an in vitro difference in binding affinity of CST to different length and sequence of oligonucleotides may indicate a moderate difference in activity but may not represent an actual difference in function in vivo.

In addition to our work showing CST’s affinity for different sequence and lengths of ssDNA, we have also shown that CST recognizes and binds to ss-dsDNA junctions without any sequence or orientation specificity of the oligonucleotide. This could be important for the incremental C-strand fill-in at telomeres and also at stalled replication forks either in the leading or lagging strand.

6.2.1 Proposed mechanism of CST action at C-strand Fill-in

At telomeres, the mechanism of C-strand fill-in after telomerase mediated G-strand elongation has always been a puzzle. Although it has been established that DNA polymerase α plays a major role in this fill-in step, the process by which polymerase is recruited without a conventional replication fork and participation of replication origin proteins (And1, Cdk) is not clear. It has been shown that in the absence of CST there is more DNA polymerase α at telomeres (121), which indicates CST does not play a required role in recruitment of DNA polymerase α. However, in the absence of CST, telomeres accumulate excess G-overhang generated only, or partially, from defects in C-strand fill-in (54,57). The defect in C-strand fill-in even at the presence of DNA polymerase α supports the idea that CST facilitates the polymerase activity and is absolutely necessary for proper C-strand fill-in. the enhanced polymerase

97 activity may reflect the ability of CST to facilitate primase to polymerase switching (98) as described for yeast. There is no other step in cell cycle that can compensate for the loss of CST in terms of this defect in

C-strand fill-in which eventually can lead into overall telomere shortening. The telomere shortening shows that C-strand fill-in and so the CST function is as important for telomere length maintenance as the elongation of G-strand by telomerase.

Previous studies indicate that C-strand fill-in is an incremental process where the fill-in starts from close to the 5’ end of the C-strand or the ss-dsDNA junction (7). Only about 70 nucleotides are needed to fill-in the C-strand and a single Okazaki fragment could achieve this. Thus it is puzzling that the C-strand fill-in takes place in gradual smaller pieces. The process of fill-in starts only after telomerase has finished extending G-strand and the absence of conventional replication proteins appear to make this process unique. The underlying mechanism of this process has been quite unclear for several reasons. First, CST does not bind only at the ss-dsDNA junctions but instead covers the entire G-overhang after telomerase extension. Secondly, DNA polymerase α can also bind to the telomere independently of presence of CST.

So how polymerase α starts the fill-in from close to the junction and how these two protein complexes coordinate, remained unclear. Because our study indicates that CST binds to the ss-dsDNA junctions, we propose that the junction-bound CST probably undergoes some conformational change that facilitates the fill-in synthesis by DNA polymerase α.

These pieces of information about C-strand fill-in, only begins to highlight the complexity of the process.

To address the underlying mechanism of C-strand fill-in and to understand the role of CST in this process, several follow up experiments need to be done. Although binding of CST to the ss-dsDNA junction could be important for C-strand fill-in, nothing is known about how CST affects DNA polymerase/ primase activity at the junction. In vitro studies to examine this issue could also illuminate the necessity of the complete CST complex to fill-in the C-strand and why CS alone fails to do so. In addition, in vitro, CST does not need a junction substrate to enhance the primase activity or primase-to- polymerase switch of DNA polymerase α (98). Therefore, some other protein interaction/ post

98 translational modification in-vivo must be present to enforce this stringency. A mass-spec analysis with

CST would be worthwhile, to understand the involvement of other proteins in this process.

Additional questions resolve around the interplay between CST and POT1. As POT1 also binds to the ssDNA, being a part of the shelterin complex that bridges between the ds-ssDNA at telomeres, questions remain as to whether CST displaces POT1 to bind to the junction. A competition assay between CST and

POT1 could be performed with a junction substrate to answer this question. Also, if we eliminate the junction binding activity of CST, we could confirm the necessity of this property of CST in the fill-in synthesis. This could be achieved by mutating different domains of CST and figuring out which domain of CST is responsible for junction binding.

6.2.2 Mechanism of action of CST in resolution of replication at G-rich DNA

Telomeres possess many different secondary structures such as T-loops, D-loops, R-loops and G- quadruplexes, which makes it a difficult region to replicate. Not only at telomeres, but at G-rich regions in the genome, G-quadruplex formation blocks passage of the replication machinery. If not resolved and replicated properly, these endogenous replication barriers can lead to loss of large chunks of telomeric

DNA. The highly stable G-quadruplex structures can also regulate transcription, depending on their site of formation, such as at promoters, or at coding or non-coding strands. Similar to many other ssDNA binding proteins, we have shown CST can bind and melt a G4 structure. The melting of G4 by CST is much more efficient (faster and stable) than that achieved by the ssDNA telomeric binding protein POT1

(130). We think this higher efficiency could be one of the potential mechanisms how CST can rescue stalled replication, and it provides an insight as to why it is enriched in GC rich DNA genome-wide.

It would be interesting to dissect the mechanism how CST unfolds G4. One can test the directionality by smFRET assay with internal labeling of oligonucleotide. It would be interesting to check if CST can melt a G4 at the ss-dsDNA junction of telomeres in a 5’-3’ direction. This could emphasize the importance of

99 junction binding of CST. It would be also interesting to check if CST has a sliding activity similar to RPA or a reeling activity similar to BLM that eventually unfolds the G4 structures.

6.3 Proposed model for CST action:

We think that although RPA and CST have quite different roles and DNA binding architecture, their striking structural similarity could still enlighten us about many DNA binding properties of CST.

Although RPA binds to a single stranded DNA stably, but its individual OB-folds micro-dissociate and re-bind to the DNA. This allows RPA to use its OB-folds as feet and helps it to diffuse along single stranded DNA. This helps RPA to destabilize duplex DNA, melt DNA secondary structures and perhaps even load or unload different interaction partners. We think CST might use a similar mechanism to unfold

G4 tracts or for the facilitated displacement of protein. The possession of many OB-folds might hinder the ability of CST to diffuse through a longer stretch of DNA which could be the reason why CST cannot facilitate helix destabilization or dsDNA melting.

Given that CST can undergo facilitated displacement, we have also wondered if CST undergoes facilitated exchange similar to RPA. Although it was beyond the scope of our experimental setup, it would be interesting to label CST with different fluorophores and perform a DNA curtains experiment similar to RPA and monitor exchange reaction (125). This could be also done with CST and POT1 or

CST and DNA polymerase α to investigate if CST can load or unload different interaction partners on ssDNA.

6.4 Does CST need to work as a complex?

We have also investigated if individual subunits play an important role in DNA binding or whether the whole complex is important. We monitored DNA binding of dimeric CS and ST complexes, and we made mutations in CTC1 and STN1 and examined binding with the mutated CST complex. We used the same oligonucleotides described above for either electrophoretic mobility shift assay (EMSA) or single molecule FRET (smFRET). We concluded that CTC1 is absolutely necessary for DNA binding. We do

100 not see efficient DNA binding with dimeric ST complex either with EMSA to ssDNA, with smFRET to ss-dsDNA junctions, or to G4, or at telomeres by ChIP in CTC1 knockout cells. Interestingly CS binds to all these oligonucleotides when monitored by EMSA or ChIP but smFRET data indicate a differential conformation of binding and suggest that the binding is either not efficient or somehow CST cannot stretch out the DNA. This starts to explain why dimeric CS complex is sufficient to terminate telomerase activity but not for C-strand fill-in. Mutations in the OB-fold or DNA binding domain of STN1 (STN1-

OBM) gave us an interesting perspective on the importance of DNA binding of CST. Our experiments indicated that the OB-fold of STN1 is responsible for sequence specificity, and also that the junction of three subunits of CST lies close to the 3’ end of the oligonucleotide regardless of the length of the substrate. As the oligonucleotide gets longer it engages more OB-folds of CTC1, CST loses the ability to differentiate between telomeric or non telomeric sequence. However, other studies indicate CST still prefers guanine-rich sequences. Mutation in STN1-OBM thus affected binding to only shorter oligonucleotides (18 nt telomeric or 36 nt non-telomeric). Most importantly, this STN1-OB mutant showed separation of function, in that it rescued defects in C-strand fill-in and reduction in dormant origin firing in STN1 knockdown cells, but we could still see significant amount of anaphase bridge formation or MTS indicating defects in double strand DNA replication both at telomeres and genome-wide.

Recently a member of the Price lab made CTC1 mutants with each of 6 predicted OB-fold deletions.

These mutants could be used individually to investigate the DNA binding and allow correlation with their in-vivo phenotypes to better understand the functions of each domain.

101

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