Telomere Structure and Maintenance In

Home , ORC1, POLE (gene)

Telomere structure and maintenance in

Trypanosoma brucei

Ranjodh Singh Sandhu

Bachelor of Science

Punjabi University, 2004

Master of Science (Biotechnology)

Punjabi University, 2006

submitted in partial fulfillment of requirements for the degree

Doctor of Philosophy in Regulatory Biology

at the

Cleveland State University

October, 2014

We hereby approve this dissertation for Ranjodh Singh Sandhu Candidate for the Doctor of Philosophy in Regulatory Biology degree for the Department of Biological, Geological and Environmental Sciences and the CLEVELAND STATE UNIVERSITY College of Graduate Studies

______Date -______Dr. Bibo Li, BGES, Cleveland State University Major Advisor

______Date -______Dr. Valentin Börner, BGES, Cleveland State University Advisory Committee Member

______Date -______Dr. Alexandru Almasan, Lerner Research Institute, Cleveland Clinic Advisory Committee Member

______Date -______Dr. Sailen Barik, BGES, Cleveland State University Advisory Committee Member

______Date -______Dr. Aaron Severson, BGES, Cleveland State University Internal Examiner

______Date -______Dr. Kausik Chakrabarti, Carnegie Mellon University External Examiner

Student’s Date of Defense: October 28th 2014

Dedicated to my family

Acknowledgements

Working on my dissertation has been both challenging and interesting.

The hurdles I met during this study were somehow made interesting by all those who inspired and guided me throughout this work.

Most importantly, I would like to express my deepest gratitude to my advisor Dr. Bibo Li for her excellent guidance, patience, motivation and providing me the opportunity to work in her laboratory. She has been a great mentor, providing freedom that allowed me to think and work independently, yet always reachable and helpful when I needed guidance.

I am grateful to my graduate advisory committee members Drs. Valentin

Börner, Alexandru Almasan and Sailen Barik for their time, constructive criticism and invaluable suggestions. It has been a great honor to have leaders in their field show interest in my work. I would also like to thank members of my dissertation defense committee Dr. Aaron Severson and Dr. Kausik Chakrabarti for providing their valuable time and their willingness to be members of my dissertation defense committee. Ph. D. was impossible without guidance and persistent help from all the departmental faculty and staff members.

I am thankful to my lab mates and friends Sanaa Jehi, Unnati Pandya,

Vishal Nanavaty, Nicole Kresak, Imaan Benmerzouga and all the undergraduate students. Without their friendship, advice and encouragement I would not have succeeded in completing my work. I am deeply thankful to Jasvinder Singh Ahuja for his constant encouragement, help and invaluable friendship. I would like to thank my wife Rima for her support and patience at all times. This thesis would not have been possible without the love, support and constant encouragement of my family. They have always been there for me and gave me strength surpass difficult times.

Telomere structure and maintenance in Trypanosoma brucei

Ranjodh Singh Sandhu

1ABSTRACT

Trypanosoma brucei is a protozoan parasite that causes sleeping sickness in humans and nagana in animals. The main reason for persistent infection of T. brucei is that in mammalian hosts, T. brucei undergoes antigenic variation and regularly switches its major surface antigen, Variant Surface

Glycoproteins (VSG), to evade the host's immune response. VSGs are exclusively expressed in a monoallelic manner from VSG expression sites (ESs) located at subtelomeric loci. We and others have shown that telomeres play important roles in VSG expression and switching regulation.

In most eukaryotes, telomere maintenance mainly relies on telomerase, a specialized reverse transcriptase. In T. brucei, the protein component of telomerase (TbTERT) has been identified previously. In this study, we identified and characterized the RNA component of T. brucei telomerase (TbTR). We established that TbTR interacts with TbTERT and is critical for telomere maintenance. We also provided insights of biogenesis of RNA component and predicted its native folding.

Telomere DNA consists of TG-rich sequences, and there is a single- stranded 3’ G-rich overhang at the very end of the telomere called G-overhang.

The G-overhang serves as a primer for telomerase-mediated telomeric DNA synthesis and also participates in formation of the T-loop structure that helps protect telomere termini from illicit DNA repair activities. The G-overhang vii

structure in T. brucei was poorly understood. We employed various methods to characterize the structure of G-overhang in T. brucei. We show that the terminal nucleotides on both G-rich and C-rich strands are specific and are regulated by telomeric proteins.

TbRAP1, an intrinsic component of the T. brucei telomere complex, is important for VSG silencing. However, the mechanism by which TbRAP1 regulates VSG silencing is unclear. We have established a number of TbRAP1 conditional knockout strains expressing various TbRAP1 mutants and provided preliminary data about functions of different TbRAP1 domains.

We have provided new insights into telomere maintenance in T. brucei, which will help better understand how telomeres contribute to antigenic variation and develop T. brucei as a model system for telomere biology research.

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2Table of Contents

ABSTRACT ...... VII LIST OF FIGURES ...... XII LIST OF TABLES ...... XV CHAPTER I INTRODUCTION ...... 1 1.1 AFRICAN TRYPANOSOMIASIS ...... 1 1.2 HUMAN AFRICAN TRYPANOSOMIASIS ...... 3 1.3 DISEASE DIAGNOSIS ...... 4 1.4 TREATMENT OF HUMAN AFRICAN TRYPANOSOMIASIS ...... 6 1.5 THE LIFE CYCLE OF T. BRUCEI ...... 8 1.6 THE T. BRUCEI GENOME ...... 10 1.7 GENE EXPRESSION AND GENOME ORGANIZATION IN T. BRUCEI ...... 12 1.8 SURFACE COAT PROTEINS AND THEIR EXPRESSION ...... 15 1.9 ANTIGENIC VARIATION IN T. BRUCEI ...... 18 1.9.1 VSG switching ...... 19 1.9.2 Monoallelic expression ...... 22 1.10 TELOMERES ...... 26 1.10.1 The telomere G-overhang structure ...... 28 1.10.2 The telomere protein complex ...... 30 1.11 TELOMERASE ...... 37 1.11.1 Telomerase reverse transcriptase (TERT) ...... 38 1.11.2 Telomerase RNA (TR) ...... 39 1.11.3 Recruitment of telomerase to telomeres ...... 42 1.12 TELOMERE AND TELOMERE MAINTENANCE IN T. BRUCEI ...... 43 1.13 SIGNIFICANCE OF STUDY ...... 46 CHAPTER II MATERIALS AND METHODS ...... 48 2.1 TRYPANOSOME STRAINS AND CULTURE CONDITIONS ...... 48 2.2 TRANSFECTION ...... 49 2.3 GENOMIC DNA ISOLATION ...... 49 2.4 ADAPTOR LIGATION ASSAY ...... 49 2.5 EXO-T TREATMENT OF GENOMIC DNA ...... 51 2.6 SINGLE TELOMERE LENGTH ANALYSIS (STELA) ...... 51 2.7 T7 EXO NUCLEASE TREATMENT OF GENOMIC DNA ...... 52 2.8 LIGATION MEDIATED PRIMER EXTENSION (LMPE) ...... 52 2.9 PREPARATION OF ARTIFICIAL TELOMERE SUBSTRATE FOR LMPE ...... 54 2.10 SOUTHERN BLOT ANALYSIS ...... 54 2.11 NORTHERN BLOT ANALYSIS ...... 55 2.12 REVERSE TRANSCRIPTION AND QUANTITATIVE REAL TIME PCR ...... 56 2.13 RNA IMMUNOPRECIPITATION (RNA IP) ...... 56 2.14 CLONING OF TERMINAL FRAGMENTS OF TELOMERES ...... 57 2.15 TELOMERASE ACTIVITY ASSAY ...... 58 2.16 PREPARING RADIOACTIVE PROBE ...... 58 ix

CHAPTER III A TRANS-SPLICED TELOMERASE RNA DICTATES TELOMERE SYNTHESIS IN TRYPANOSOMA BRUCEI ...... 60 3.1 INTRODUCTION ...... 60 3.2 RESULTS...... 64 3.2.1 Identification of a putative TbTR gene ...... 64 3.2.2 Association of TbTR and TbTERT in vivo ...... 68 3.2.3 Deletion of TbTR leads to progressive telomere shortening ...... 69 3.2.4 TbTR DKO cells lack telomerase activity ...... 76 3.2.5 Mutation in the TbTR template region resulted in altered telomere sequences ...... 80 3.2.6 Biogenesis of TbTR ...... 82 3.2.7 Secondary structure of TbTR ...... 85 3.3 DISCUSSION ...... 89 CHAPTER IV G-OVERHANG STRUCTURE OF TRYPANOSOMA BRUCEI ... 96 4.1 INTRODUCTION ...... 96 4.1.1 G-overhang function ...... 97 4.1.2 Generation of G-overhang ...... 99 4.1.3 G-overhang binding Proteins ...... 103 4.1.4 G-overhang processing ...... 105 4.1.5 Molecular steps of G-overhang generation ...... 108 4.1.6 Exception to the rule ...... 114 4.1.7 Methods to study the telomere G-overhang ...... 115 4.1.8 G-overhang in T. brucei ...... 119 4.2 RESULTS:...... 120 4.2.1 The Terminal Nucleotide at the G-rich strand of T. brucei telomeres is tightly regulated ...... 120 4.2.2 Telomerase activity is required to maintain the telomere G- overhang in T. brucei ...... 128 4.2.3 Deletion of TbKu results in a change in G-overhang dynamics. .. 130 4.2.4 Continuous propagation of the telomerase null T. brucei strain leads to a change in G-overhang dynamics...... 134 4.2.5 MRE11 deletion has no effect on telomere G-rich strand terminal sequence...... 138 4.2.6 The C-rich strand terminal nucleotide sequence of T. brucei telomere is tightly regulated...... 139 4.2.7 Telomerase and TbKu have no effect on the C-strand terminal nucleotide sequence...... 143 4.2.8 Ligation mediated primer extension assay for measuring the length of G-overhang ...... 145 4.2.9 The G-overhang length of T. brucei telomere ...... 149 4.3 DISCUSSION ...... 150 4.3.1 The molecular mechanism of overhangs processing ...... 165 4.4 CONCLUSION ...... 168

CHAPTER V FUNCTIONAL ANALYSIS OF DOMAINS OF THE TELOMERIC PROTEIN TBRAP1 ...... 169 5.1 INTRODUCTION ...... 169 5.1.1 The domain structure of RAP1 homologues ...... 173 5.2 RESULTS...... 179 5.2.1 The conditional knockout system ...... 179 5.2.2 Confirmation of conditional knockout system ...... 182 5.2.3 Conditional knockout of TbRAP1 results in several hundred folds of derepression of silent VSGs ...... 185 5.2.4 The TbRAP1 BRCT domain is required for VSG silencing ...... 187 5.2.5 The TbRAP1 Myb domain deletion leads to a null phenotype ..... 192 5.2.6 The Myb-like domain of TbRAP1 is essential for T. brucei survival...... 194 5.2.7 The RCT domain is essential for TbRAP1 function...... 195 5.3 DISCUSSION ...... 198 5.4 CONCLUSION ...... 202 CHAPTER VI SUMMARY AND FUTURE PERSPECTIVE ...... 203 6.1 TELOMERASE REGULATION IN T. BRUCEI ...... 203 6.2 FUNCTIONS OF TBTR DOMAINS ...... 207 6.3 TBTR AS A POTENTIAL DRUG TARGET ...... 208 6.4 G-OVERHANG STRUCTURE MAINTENANCE ...... 211 BIBLIOGRAPHY ...... 214

3List of Figures

Figure 1.1 Life cycle of T. brucei ...... 9 Figure 1.2 Genome organization in T. brucei...... 11 Figure 1.3 Expression of cell surface proteins ...... 21 Figure 1.4 Mechanism of VSG switching ...... 23 Figure 1.5 T-loop structure ...... 29 Figure 1.6 Telomere protein complex in different organisms ...... 35 Figure 3.1 Identification of T. brucei telomerase RNA ...... 63 Figure 3.2 Analysis of TbTR expression ...... 66 Figure 3.3 Full length TbTR ...... 68 Figure 3.4 Molecular validation of TbTR ...... 69 Figure 3.5 Confirmation of TbTR deletion ...... 70 Figure 3.6 Telomere length changes in WT and TbTR DKO BF cells ...... 72 Figure 3.7 Progressive telomere shortening at silent expression sites ...... 73 Figure 3.8 Telomere length changes in WT and TbTR DKO PF cells ...... 74 Figure 3.9 The telomere shortening phenotype in TbTR DKO cells is complemented by an ectopically expressed TbTR WT allele ...... 75 Figure 3.10 TbTR DKO cells do not have growth defects ...... 76 Figure 3.11 Optimizing TRAP assays to examine telomerase activities in T. brucei cells...... 77 Figure 3.12 TRAP assay to compare the telomerase ...... 79 Figure 3.13 Cloning of telomeres ...... 80 Figure 3.14 Biogenesis of TbTR ...... 83 Figure 3.15 Chemical probing of TbTR and secondary structure prediction ...... 88 Figure 4.1 End replication problem ...... 98 Figure 4.2 G overhang generation ...... 101 Figure 4.3 Processing of G overhang in S. cerevisiae ...... 102 Figure 4.4 Generation of the telomeric overhang in mammalian cells ...... 103 Figure 4.5 Comparison of G-overhang end structures ...... 114 Figure 4.6 Schematic of adaptor ligation assay ...... 122 Figure 4.7 G-overhang ending in bloodstream form T. brucei: ...... 123 Figure 4.8 Signal in adaptor mediated end ligation assay depends on ligation. 124 Figure 4.9 Quantification of G-overhang signals of bloodstream form T. brucei125 Figure 4.10 G-overhang ending in procyclic form T. brucei ...... 126 Figure 4.11 Quantification of G-overhang signals of procyclic form T. brucei. .. 127 Figure 4.12 TbTERT deletion leads to loss of 5’ TTAGGG 3’ G-overhang endings...... 129 Figure 4.13 Loss of TbTERT results in loss of 5’ TTAGGG 3’-ending G- overhangs...... 130 Figure 4.14 TERT deletion leads to loss of 5’ TTAGGG 3’ G-overhang endings...... 131 Figure 4.15 TbKU80 deletion leads to decrease in 5’ TTAGGG 3’ overhang endings and increase in 5’ TAGGGT3’ overhang endings...... 132 Figure 4.16 Loss of TbKU80 results in loss 5’TTAGGG3’ ending ...... 133 xii

Figure 4.17 TbTR deletion leads to a change of G-overhang end sequence dynamics...... 135 Figure 4.18 Loss of TbTR results in a change in G-overhang ending sequence...... 136 Figure 4.19 Complementation of TbTR was able to restore G-overhang terminal sequence...... 137 Figure 4.20 TbMRE11 is not required for G-overhang end sequence specificity...... 138 Figure 4.21 Schematic showing ligation mediated PCR assay ...... 140 Figure 4.22 Ending of C-Rich strand is tightly regulated ...... 141 Figure 4.23 Quantitation of terminal signals of the telomere C-rich strand in bloodstream form of T. brucei...... 142 Figure 4.24 Deletion of TbKu or TbTR has no effect on terminal sequence of the telomere C-rich strand ...... 144 Figure 4.25 Schematic showing steps of ligation mediated primer extension assay...... 146 Figure 4.26 Ligation mediated primer extension is a sensitive method to detect the telomere length of G-overhang...... 147 Figure 4.27 Complementarity of the guide oligonucleotide with the terminal G- overhang sequence is a prerequisite primer extension ...... 148 Figure 4.28 T. brucei telomeres have short G-overhangs ...... 151 Figure 4.29 Schematic showing steps of telomerase dependent telomere elongation...... 153 Figure 4.30 Telomere terminal sequence in T. brucei ...... 155 Figure 4.31 Proposed mechanism for the the G-overhang maintenance ...... 166 Figure 4.32 Proposed mechanism of the telomere C-strand terminal nucleotide specification ...... 168 Figure 5.1 Schematic drawing of domains of Rap1 homologs...... 174 Figure 5.2 Schematic drawing of the LoxP system...... 177 Figure 5.3 Generating a single floxed allele of TbRAP1...... 178 Figure 5.4 Induction of Cre leads to the efficient removal of the floxed TbRAP1 allele ...... 181 Figure 5.5 TbRAP1 is haploinsufficient ...... 184 Figure 5.6 Conditional knockout of TbRAP1 leads to derepression of BES-linked and MES-linked VSGs ...... 186 Figure 5.7 Graphical representation of TbRAP1 mutants ...... 187 Figure 5.8 Deletion of the BRCT domain of TbRAP1 leads to a severe growth arrest ...... 188 Figure 5.9 BRCT domain deletion of TbRAP1 leads to several thousand folds of derepression of BES-linked VSGs ...... 191 Figure 5.10 Deletion of the Myb domain of TbRAP1 leads to a severe growth arrest ...... 192 Figure 5.11 Myb domain deletion of TbRAP1 leads to several hundred folds of derepression of BES-linked VSGs...... 196

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Figure 5.12 Deletion of the Myb like domain of TbRAP1 leads to a severe growth arrest...... 197 Figure 5.13 Deletion of the RCT domain of TbRAP1 leads to a TbRAP1 null phenotype ...... 199 Figure 6.1 TbKU80 deletion leads to a decrease in telomerase activity ...... 205 Figure 6.2 TbKU80 deletion does not affect TbTR levels ...... 206 Figure 6.3 Mutation in TbTR template leads to severe growth defects ...... 209

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4List of Tables

Table 2.1 Guide oligonucleotide sequence ...... 51 Table 2.2 Sequence of Top oligonucleotides...... 53 Table 3.1 Telomere sequences in several representative telomere clones ...... 94

1CHAPTER I

Introduction

1.1 African trypanosomiasis:

Trypanosomes are obligate parasites belonging to the class Kinetoplastida and genus Trypanosoma. These unicellular and flagellated parasites live and multiply mostly extracellularly in the blood and tissue fluids of their hosts. Most of the trypanosomes are heteroxenous, i.e. they require more than one host to complete their life cycle (Yao, 2010). Some trypanosomes, such as T. brucei are highly pathogenic and infect humans as well as animals. Infection of humans by

T. brucei results in human African trypanosomiasis (HAT) or ‘sleeping sickness’, while infection in animals results in animal African trypanosomiasis (AAT) or

‘Nagana’. T. brucei is transmitted by the bite of a blood sucking fly known as Tse-

Tse (Glossina spp.) that acts as a vector host. The sub-Sahara region of Africa that is affected by trypanosomiasis corresponds to the range of tsetse (Simarro et al., 2012).

T. congolense, T. vivax and T. brucei spp. are the major causative agents of nagana. In domestic animals, nagana is often fatal and is one of the most

economically important livestock diseases in Africa. It has a devastating impact on rural areas. Approximately 50 million cattle are threatened by trypanosomiasis

(Holmes, 2013). Severe outbreaks of this disease in cattle lead to low productivity and stunted economical development. Moreover, domestic livestock acts as a reservoir of parasites for human infections (Shaw, 2004).

HAT is caused by two subspecies of T. brucei, T. brucei gambiense and T. brucei rhodesiense. HAT is widespread and covers an area of approximately

1.55 million km2 in Africa (Simarro et al., 2012). If untreated, HAT can be fatal. T. b. gambiense is the causative agent of chronic and more prevalent form (97% of the cases) of HAT in West and Central Africa while T. b. rhodesiense causes a rare (3% of the cases), acute and more severe infection in East and South regions of Africa (Simarro et al., 2008). If untreated, patients infected with T. b. rhodesiense die within months after infection while patients with T. b. gambiense infection die within few years of infection (Pepin, 2010). In the early stage of

HAT, trypanosomes are limited to the blood and lymphatic system of the host leading to the development of symptoms such as headache, fever, fatigue, etc. In the second stage of this disease, trypanosomes are detectable in cerebrospinal fluid leading to development of symptoms such as confusion; disturbed sleep pattern, sensory disturbances, extreme lethargy, poor condition and coma

(Kennedy, 2013; Kennedy, 2004; Lundkvist et al., 2004). Trypanosome species causing AAT are unable to infect humans due to the presence of trypanosome lytic factors (TLFs) in human serum (Capewell et al., 2011). T. b. rhodesiense that causes severe form of HAT expresses a serum resistance protein SRA (Van 2

Xong et al., 1998). Instead, T. b. gambiense reduces the expression of a receptor that is involved in uptake of lytic factor in trypanosome cells (Kieft et al., 2010). T. b. brucei is very closely related to T. b. rhodesiense and T. b. gambiense, but it is sensitive to human lytic factors. This makes T. b. brucei a very useful model organism to study in the lab.

1.2 Human African Trypanosomiasis

Cases of HAT have declined in recent years. The latest epidemic of HAT was reported in 1970s and it lasted untill late 1990s (Berrang-Ford et al., 2006).

The upward rise of HAT has been finally decreased due to better treatment and more efficient control of the insect vector. In 2009, World Health Organization

(WHO) reported less than 10,000 cases of HAT. This decline in the number of cases has continued as in 2012, only 7216 cases were reported by WHO.

However, this is just a fraction of the actual number of cases, as many cases are not recognized or reported due to incomplete screening (Chappuis et al., 2010).

According to WHO, the true number of cases is believed to be approximately

30,000 per year.

Symptoms of HAT are divided into two stages: early stage and late stage.

These stages are mostly difficult to differentiate as these often merge with each other. The early stage is called the hemolymphatic stage. The onset of this disease is variable and it usually takes 1 to 3 weeks after a tsetse bite for symptoms to appear. A painful swelling called Chancres appears at the site of the bite. The main symptoms of early hemolymphatic stage are episodes of fever

that last for 1 to 7 days, lymphadenopathy, malaise, headache, weight loss, and painful joints, etc. Patients may also develop clinical features, such as enlarged spleen and liver, cardiovascular, endocrine and ophthalmic abnormalities.

(Kennedy, 2004) (Kennedy, 2013). The late stage of HAT is called the encephalitic stage. In this stage trypanosomes cross the blood brain barrier and enter the nervous system. A wide variety of symptoms occur at this stage that include various psychiatric, reflex, and sleep abnormalities. Other common features of this stage are meningoencephalitis, tremors, and speech abnormality

(Blum et al., 2006) (MacLean et al., 2012). Almost all parts of the nervous system are affected. Without proper treatment, patient’s condition worsens and symptoms such as seizures, cerebral edema, systemic organ failure, and severe somnolence appear. This is followed by an inevitable death (Bouteille and

Buguet, 2012).

1.3 Disease diagnosis

One of the key diagnostic clues is the presentation of clinical symptoms in a dwelling where the disease is endemic. However many other diseases exhibit similar symptoms as HAT. Therefore it is important to exclude the presence of other diseases, such as malaria, toxoplasmosis, HIV infection, leishmaniasis, and typhoid (Atouguia et al., 2000) (Kennedy, 2004). Diagnosis at the early hemolymphatic stage involves demonstrating the presence of trypanosomes in the peripheral blood of infected individuals. This type of diagnosis is successful, particularly in case of the T. b. rhodesiense infection, as this type of HAT

includes presence of high number of parasites in the blood (Kennedy, 2006).

However, detecting the parasite in the blood of a patient infected with T. b. gambiense is more challenging due to the cyclical nature and therefore rarer occurrence of high parasitaemia (Meltzer et al., 2012). Therefore, another diagnostic test used for T. b. gambiense is based on the detection of variable surface antigen LiTat 1.3. This test is called the card agglutination trypanosomiasis test (CATT). It is a simple and quick test that is widely used in population screening. However, limited sensitivity and high frequency of dubious results are some of the limitations of CATT (Truc et al., 2002). Another diagnostic tool used to detect early stage trypanosome infection is PCR. PCR is a highly sensitive and specific technique to detect HAT. In spite of its high efficiency, advanced techniques such as PCR are not available in the field conditions

(Mugasa et al., 2012) (Kennedy, 2004).

To successfully treat HAT, it is imperative to distinguish the early hemolymphatic stage of infection from the late encephalitic stage. The accurate stage identification of HAT patients is necessary, as failure to appropriately treat a patient with CNS drugs in the encephalitic stage of infection will lead to eventual death, while inaccurate treatment of patients in early stage of infection with CNS drugs will lead to unnecessary drug toxicity (Kennedy, 2008). The encephalitic stage of HAT is detected by analyzing the CNS fluid for the presence of trypanosomes and also by detecting elevated levels of white blood cells (Kennedy, 2008). In summary, quick, easy to undertake, and efficient

diagnostic tools that have applicability in affected regions of Africa are required for detection of HAT.

1.4 Treatment of Human African Trypanosomiasis

Four main drugs, namely suramin, pentamidine, melarsoprol, and eflornithine (difluoromethylornithine, or DFMO) have been used for the treatment of HAT (Brun et al., 2010). Treatment of HAT with these drugs is not very satisfactory as these drugs are very toxic and difficult to administer (Steverding,

2010). For early stage HAT infection by T. b. gambiense, a drug called pentamidine is administered intramuscularly or intravenously, while early stage T. b. rhodesiense infection is treated by administering suramin intravenously (Brun et al., 2010). These treatments are usually effective in preventing disease progression but they also have some side effects, such as hypotension, hyperglycemia or hypoglycemia and gastrointestinal disturbances in case of treatment with pentamidine, and anaphylactic shock, neurological complications upon treatment with suramin (Kennedy, 2004) (Kennedy, 2013).

While these drugs are used to treat early stages of two forms of HAT, treating late stages of HAT is much more difficult as in late stage of infection trypanosomes have crossed the blood brain barrier. Only one drug, melarsoprol, is used to treat late stages of both forms of HAT, but it is most commonly used in

T. b. rhodesiense infection (Kuepfer et al., 2012). Although melarsoprol is an effective treatment for late stage of HAT, the main drawback of this drug is that it cannot be administered orally. This drug is administered intravenously by

propylene glycol, as it is insoluble in water, making drug injections very painful for the patient (Kennedy, 2004). Melarsoprol is also very toxic. The overall mortality rate due to melarsoprol is really high, at 5-10%, due to development of post treatment reactive encephalopathy (Barrett et al., 2007). There is also evidence of drug resistance with this drug, and treatment failure rates of 30% are reported in patients (Baker et al., 2013).

Eflonithine is used to treat late stage HAT caused by T. b. gambiense.

Nifurtimox - eflornithine combination therapy (NECT) is used as a first line of treatment for patients in late stages of T. b. gambiense infection (Franco et al.,

2012). This combination drug treatment has a significantly reduced mortality rate to 0.7% as compared to a treatment using eflornithine alone (with a mortality rate of 2.1%) and melarsorprol (with a mortality rate of 5-10%) (Kennedy, 2004)

(Barrett et al., 2007). However NECT is not effective against T. b. rhodesiense infections (Willert and Phillips, 2012). It still needs to be administered intravenously and NECT treatment results in many side effects such as seizures, gastrointestinal problems, alopecia, and bone marrow toxicity (Kennedy, 2004).

Several new drugs for treatment of HAT are under various stages of development and are also being tested on animal models, such as fexinidazole, which is effective and non toxic in animal models (Kaiser et al., 2011), and

SCYX-7158 that has the ability to cure CNS infection of trypanosomes in murine models (Jacobs et al., 2011).

1.5 The Life Cycle of T. brucei:

T. brucei needs two hosts to complete its life cycle, an insect vector

(tsetse) and a mammalian host. In tsetse, the parasite lives and proliferates in the midgut. This form of T. brucei is called procyclic form. Later, the parasite migrates to the salivary glands of the fly and differentiates into metacyclic form.

Here it acquires infectivity and is transmitted to a new mammalian host upon biting of the infected tsetse. Upon entry into the new mammalian host, T. brucei lives freely in the bloodstream and this stage is called bloodstream form. T. brucei lives as a proliferative slender form in the bloodstream of the mammalian host. The slender form is later replaced by the short stumpy form (Robertson,

1912) (Vickerman, 1985). This metamorphosis is important for two reasons. First, the slender form is replaced by the stumpy form when the number of parasites in the host is increased (Matthews, 2005). The stumpy form is non proliferative, thus the number of parasites is not increased any further after conversion to the stumpy form and survival of the host is prolonged. Second, the stumpy form is arrested in G1 phase of the cell cycle. This arrest ensures that morphological changes that occur after transmission of the stumpy form into tsetse can be coordinated with re-entry into the cell cycle. Once the stumpy form is transmitted back to the vector, it is metamorphosed into the procyclic form in the midgut of the insect (Figure 1.1).

There are several morphological differences between procyclic, metacyclic, and bloodstream forms of the T. brucei. One such difference is in the cell surface glycoprotein coat. In procyclic form, T. brucei expresses procyclin as 8

the surface coat, while in metacyclic and bloodstream forms, variant surface glycoproteins (VSGs) are the major components of T. brucei surface coat

(Vickerman, 1969) (Cross, 1975) (Cross, 1977) (Roditi et al., 1989) (Roditi et al.,

1998). This change in surface coat is very important for survival and proliferation of T. brucei in the insect as well as in the mammalian host (Figure 1.1).

Figure 1.1 Life cycle of T. brucei Schematic representation of the T. brucei life cycle (Dreesen et al. 2007).

1.6 The T. brucei Genome

The haploid genome of T. brucei is estimated to be about 35 Mb in size and contains the mitochondrial genome and the nuclear genome (Acosta-

Serrano et al., 2007). The mitochondrial genome of T. brucei or kDNA

(kinetoplastid DNA) is present within a single mitochondrion of T. brucei. kDNA is arranged in interlocked circular DNA made up of a few dozens of maxi-circles that are about 25 kb and several thousands of mini-circles that are about 1 kb

(Simpson, 1987; Liu et al., 2005). Maxi-circles encode for rRNAs and mitochondrial proteins, while mini-circles mainly encode for guide RNAs that serve as templates for RNA editing (Liu et al., 2005). The nuclear genome of T. brucei is divided into three types of chromosomes based on their sizes: 11 pairs of megabase chromosomes (0.9-5.7 Mb), 2-4 intermediate chromosomes (300-

900 kb), and ~ 100 minichromosomes (50-100 kb) (Melville et al., 1998)

(Wickstead et al., 2004) (Berriman et al., 2005). All of these chromosomes are linear and possess TTAGGG telomeric repeats at both ends (Figure 1.2).

Most of the coding DNA of T. brucei genome is distributed among 11- megabase chromosomes that are present as homologous pairs (Berriman et al.,

2005). Interestingly, these homologous chromosomes are not exactly identical as they vary significantly in size (Melville et al., 1999). Heterogeneity in size of homologous chromosomes is largely attributed to the subtelomeric differences mostly due to the duplication of VSG gene arrays (Callejas et al., 2006). These variations in size result in haploid like sequences on homologous chromosomes and thus lead to hemizygous condition. Sequencing of 11 megabase 10

Figure 1.2 Genome organization in T. brucei Three different types of chromosome are present in T. brucei. 11 megabase chromosomes encode for essentially all housekeeping genes in form of polycistronic units. 2-4 intermediate chromosomes contain Bloodstream form expression site (BES) in subtelomeric regions. Complete sequence of intermediate chromosomes is not known, but 177 bp repeats are present. ~ 100 minichromosomes are present in T. brucei genome. The core of minichromosomes is made up of 177 bp repeats and about one third of minichromosomes contain VSG genes at subtelomeric regions (Adopted from Akiyoshi and Gull, 2013). chromosomes of T. brucei predicted that there are approximately 9,000 protein- coding genes on these chromosomes (Berriman et al., 2005). Out of these, approximately 2500 genes represent VSG genes and pseudogenes (Cross et al.,

2014,) .

Not much is known about the intermediate chromosomes of T. brucei and their ploidy is unclear. Intermediate chromosomes harbor a core of non-repetitive sequence and 177 bp repeats at their subtelomeric regions (Van der Ploeg et al.,

1984a) (Wickstead et al., 2004). Some intermediate chromosomes contain VSG expression sites at their subtelomeres, however no housekeeping genes are reported on these chromosomes so far (Van der Ploeg et al., 1984a).

Minichromosomes are thought to be reservoirs of genes encoding for

VSGs (Borst, 1986). Each minichromosome is made up of 177 bp repeats of unknown function at its core and about one third of minichromosomes ends contain a VSG gene (Gibson and Borst, 1986) (Weiden et al., 1991) (Alsford et al., 2001) (Wickstead et al., 2004) (Cross et al., 2014).

1.7 Gene expression and genome organization in T. brucei

T. brucei and related trypanosomes have developed an unusual way of gene expression and mRNA maturation. Most of the genes in T. brucei are within polycistronic units each of which contains multiple genes (Tschudi and Ullu,

1988) (Daniels et al., 2010). Although the polycistronic transcription is known to take place in prokaryotes and eukaryotes, the scale at which T. brucei utilizes this process is exceptional. Most of the genes in the genome of T. brucei are organized into large directional gene clusters that contain genes in head to tail orientation (Daniels et al., 2010) (Figure 1.2). This is reminiscent of bacterial operons except that in the case of T. brucei, genes in a polycistron may not be functionally related. Recent data from RNA-seq and ChIP-seq experiments

combined with whole genome organization data suggest that the genome of T. brucei (excluding subtelomeric regions) is divided into approximately 150 polycistronic units transcribed by RNA polymerase II (Kolev et al., 2010) (Siegel et al., 2009) (Daniels et al., 2010). These polycistronic units have an average size of 153 kb and contain about 55 genes each (Daniels et al., 2010). Genes within a polycistronic unit are transcribed from the same strand of DNA and the region that separates two neighboring polycistronic units is called strand switch region (SSR). SSRs can be divergent or convergent. Transcription initiates at divergent SSRs, thus they are believed to contain promoter and transcription start sites. Convergent SSRs are thought to be transcription termination sites.

Although recent studies have provided some evidence that chromatin composition is involved in transcription initiation and termination, much remains unclear (Siegel et al., 2009). As the promoter requirements for RNA polymerase

II dependent transcription are unknown in T. brucei, it is unclear from where transcription begins and terminates (Gunzl et al., 2007) (Martínez-Calvillo et al.,

2010).

Once a polycistronic unit is transcribed, trans-splicing is then used to resolve the polycistronic transcription unit into monocistronic translatable units

(Huang and Van der Ploeg, 1991) (LeBowitz et al., 1993). During the process of trans-splicing, a 39-nt spliced leader (SL) RNA is added to the individual mRNAs at the 5’ end (Parsons et al., 1984). SL RNA provides an unusual 5’ cap structure called ‘cap 4’ (Perry et al., 1987). ‘Cap 4’ differs from the typical 5’ cap of eukaryotic mRNA due to additional four methyl groups that are present at the first 13

4 nucleotides of SL RNA (Bangs et al., 1992). Tandem mRNAs present on the polycistronic units are separated by intergenic sequence and the addition of the

SL RNA on the downstream gene is coupled with polyadenylation of the upstream gene (LeBowitz et al., 1993).

Despite of the differences in transcription and RNA processing, T. brucei has highly conserved copies of all three RNA polymerases (Palenchar and

Bellofatto, 2006). However they have clearly acquired new functions to fulfill some parasite specific demands. RNA polymerase II transcribes polycistronic units of protein coding genes (as discussed above) and monocistronic transcripts of SL RNA (Gilinger and Bellofatto, 2001). RNA polymerase II also transcribes polycistronic clusters of snoRNAs present throughout the megabase chromosomes. RNA polymerase III transcribes tRNAs and the U–rich snRNAs

(Fantoni et al., 1994). Surprisingly, the RNA polymerase I in T. brucei not only transcribes pre-rRNAs (18S, 5.8S, and 28S) but also transcribes mRNAs.

Importantly, mRNAs transcribed by RNA polymerase I include those that code for major cell surface proteins of the parasite at different life stages (Günzl et al.,

2003) (Pays, 2005). RNA polymerase I transcribes polycistronic units that contain

VSG genes at subtelomeric loci of megabase and intermediate chromosomes at the bloodstream form stage (Günzl et al., 2003). In procyclic form, it transcribes polycistronic units containing procyclin genes located internally on megabase chromosomes 6 and 10 (Günzl et al., 2003). Lastly, in the metacyclic form, RNA polymerase I transcribes the monocistronic unit of metacyclic VSGs (mVSGs)

located at subtelomeric regions of megabase and intermediate chromosomes

(Günzl et al., 2003) (Ginger et al., 2002).

Genome sequence of three Trypanosomatid species, namely T. brucei, T. cruzi, and Leishmania major (also called as tritryps), not only confirmed the unusual organization of genes in polycistronic transcription units but also revealed a high degree of synteny among these three Trypanosomatid species

(El-Sayed et al., 2005). Genome organization maps based on this synteny have revealed that the chromosomes can be divided into two units based on the types of gene clusters (El-Sayed et al., 2005). First is the central core of the chromosomes that contains housekeeping genes and is conserved across the tritryps. Second is the species specific subtelomeric proximal region, and genes located at subtelomeres are mostly devoted to parasite-host interactions and evasion of host’s immune system.

1.8 Surface coat proteins and their expression

During various life stages, T. brucei expresses distinct cell surface coat proteins. In bloodstream form, the bloodstream form VSGs covers the surface of parasite while metacyclic VSGs cover its surface during the metacyclic stage. In the procyclic form, procyclin is the major surface protein. As parasite survival depends upon the right surface coat, the expression of these different surface coat proteins is tightly regulated (McCulloch, 2004).

In the bloodstream form, T. brucei cell surface is covered by a 15 nm thick coat of ~ 10 million identical molecules of VSGs (Cross, 1975). These VSGs are

present on the extracellular surface of the parasite and are attached to the membrane by a glycosylphosphatidylinositol anchor (GPI anchor) (Ferguson et al., 1988) (Ferguson, 1999) (Schwede and Carrington, 2010). VSGs are densely packed and are highly immunogenic (David Barry and McCulloch, 2001).

Although T. brucei genome contains more than 2,500 VSG genes and pseudogenes, only one is expressed at any given time. These VSGs are expressed exclusively from the bloodstream form VSG expression sites (BESs) at subtelomeric loci (De Lange and Borst, 1982) (El-Sayed et al., 2000). BESs are ~40–60 kb polycistronic transcription units transcribed by RNA polymerase I

(Günzl et al., 2003) (Hertz-Fowler et al., 2008).

Each BES contains a set of expression site associated genes (ESAGs),

TAA rich sequence of imperfect repeats called 70 base pair repeats, and a VSG gene (Pays et al., 2001) (Hertz-Fowler et al., 2008). The VSG gene is located at the 3’ end of eacg BES within 2 kb of the telomere repeat. So far, fifteen highly conserved BESs have been identified in a widely studied T. brucei strain, Lister

427 strain (Hertz-Fowler et al., 2008). At any given time, only a single BES is fully actively transcribed while all other BESs are reversibly repressed (Bernards et al., 1984). Apart from BES-linked VSG genes, large numbers of VSG genes are present as long tandem arrays on the hemizygous subtelomeric loci of megabase chromosomes and also as single copy genes on the subtelomeric regions of minichromosomes. These VSG genes lack a promoter and thus are normally not expressed.

Metacyclic VSGs (mVSGs) are expressed from metacyclic VSG expression sites (MESs) located at the subtelomeric region of megabase chromosomes (Lenardo et al., 1984). MESs are monocistronic units that are transcribed by RNA polymerase I from a promoter located approximately 5 kb upstream of the mVSG gene (Alarcon et al., 1994) (Ginger et al., 2002).

Metacyclic population can express 15-20 mVSGs collectively in tsetse salivary glands, however each parasite cell is coated by only one type of mVSG (Barry et al., 1998).

In the insect host, a less dense coat of procyclins replaces the VSG coat

(Mowatt and Clayton, 1987). Procyclins are resistant to the proteases present in the mid-gut of tsetse thus allowing proliferation of the parasite. Unlike the bloodstream form VSG and mVSG genes, the procyclin genes are expressed from chromosome internal sites (Mowatt et al., 1989) (Clayton et al., 1990).

Procyclin genes are organized as tandem repeats of two to three copies present on chromosomes 6 and 10 and are transcribed by RNA polymerase I. Four types of procyclin genes are present in the T. brucei genome, three of these encode for

EP proteins [glutamic acid (E)-proline (P) dipeptide repeats] (Roditi et al., 1998).

These EP proteins can further be distinguished by the presence or absence of the N-linked glycan. The fourth gene encodes for the GPEET protein (a pentapeptide repeat protein). GPEET does not contain any glycosylation site

(Roditi et al., 1998). Two or more types of procyclin proteins can be expressed simultaneously on the surface of the parasite (Mowatt et al., 1989) (Figure 1.3).

1.9 Antigenic Variation in T. brucei

As mentioned earlier, T. brucei lives and multiplies in the bloodstream and tissue spaces of mammalian host; this renders T. brucei fully exposed to the immune surveillance of the host. In order to evade the host immune response and survive in this hostile environment, T. brucei constantly changes its surface coat. This infamous ability to change the surface coat is called antigenic variation. First reported by Ronald Ross and Thomson, this regular switching of the coat leads to characteristic remitting and relapsing waves of low and high parasitemia eventually exhausting the immune system of the host (Ross and

Thomson, 1910). Antigenic variation allows the T. brucei to establish a persistent infection in the host.The m olecular mechanism responsible for antigenic variation has been a focus of intense research for many years, but the details of this mechanism are still not clear. Although antigenic variation is a widely used strategy to evade host immune system, T. brucei masters it (David Barry and

McCulloch, 2001).

There are three main requirements to successfully use antigenic variation to evade the host immune response (Borst and Genest, 2006). First, a large pool of genes coding for antigenically distinct surface proteins should be present.

Approximately 30% of the T. brucei genome is dedicated to store more than

2,500 VSG genes and pseudogenes (Cross et al., 2014) (Horn, 2014). These

VSG genes can recombine to form novel VSG genes, making the pool of antigenically distinct VSGs essentially limitless (Marcello and Barry, 2007).

Second, the parasite should be able to switch from expression of one variant 18

surface protein to another. T. brucei exploits both transcription and recombination based mechanisms to switch from one VSG to another. Third, only one type of

VSG is expressed at a given time (monoallelic expression) to ensure that after each VSG switching, the previously expressed VSG is no longer present on the cell surface. The T. brucei genome contains approximately 15 BESs but only one is fully transcribed at any time, resulting in a single type of VSG being expressed at any given time.

1.9.1 VSG switching

Wild type isolates of the bloodstream form T. brucei spontaneously switches from one VSG to another at the rate of 10-2 to 10-4 /doubling time of about 6 hrs (Turner and Barry, 1989) (Turner, 1997). These switching events can be broadly classified into two types: transcriptional (or in situ switch) and recombination based switch (McCulloch, 2004) (Horn and McCulloch, 2010)

(Horn, 2014). In case of transcriptional switch, a new BES is expressed and the originally active BES is silenced (Borst and Ulbert, 2001). Not much is known about how this is achieved and regulated. In recombination-based switch, one of the silent VSG genes is recombined into an active BES (David Barry and

McCulloch, 2001). Different mechanisms are used to remove the old and introduce the new VSG gene in the active BES.

One of the best-studied and most frequently used mechanisms is gene conversion, in which a silent donor VSG gene replaces the originally active VSG gene. In this process, the originally active VSG gene is lost while the donor VSG

gene is duplicated (Robinson et al., 1999). Another mechanism of VSG switching is the reciprocal telomeric exchange in which the recombination between two chromosome ends results in a reciprocal transfer of new VSG to the active BES and the previously active VSG to a silent BES (Pays et al., 1985) (Rudenko et al.,

1996). Finally, recombination can copy segments of multiple VSG genes into the active BES, resulting in a new chimeric VSG gene (Marcello and Barry, 2007).

Many factors have been shown to be important for VSG switching.

As most of the switching events are based on homologous recombination

(HR), a conserved HR protein RAD51 is clearly important for VSG switching

(McCulloch and Barry, 1999), Deletion of other HR proteins such as BRCA2 and

RAD51 paralogs (RAD52 and RAD53) also leads to lowering of switching frequency (McCulloch and Barry, 1999) (Proudfoot and McCulloch, 2005)

(Hartley and McCulloch, 2008). Although much reduced, recombination based switching also occurs in the absence of RAD51, suggesting that RAD51- independent pathways are also involved in recombination based switching (Kim and Cross, 2011). Microhomology-mediated end joining (MMEJ) is implicated for these residual-switching events in the absence of HR (Glover et al., 2010).

Although MMEJ is shown to mediate DNA recombination, factors required for

MMEJ are still unknown in T. brucei (Glover et al., 2010).

Figure 1.3 Expression of cell surface proteins Schematic representing loci of VSGs and procyclin expression sites in T. brucei genome (see text for details) (Adopted from Navarro et. al. 2007).

Factors that block aberrant recombination, such as Topo3 and RMI1

(RecQ-mediated genome instability) prevent switching (Kim and Cross, 2010)

(Kim and Cross, 2011). Deletion of these factors results in an increased frequency of switching. RNAi mediated depletion of origin recognition complex 1

(ORC1) displayed an elevated frequency of VSG switching (Benmerzouga et al.,

2013). Finally, as most of the recombination required for switching takes place in proximity of telomeres, both telomere length and telomeric proteins play important roles in regulating VSG switching events. Telomerase protein component (TbTERT) deletion cell lines harboring extremely small telomeres have a higher frequency of VSG switching, highlighting the importance of telomere maintenance in regulation of antigenic variation (Hovel-Miner et al.,

2012). Moreover, depletion of any known telomeric proteins (TbTRF, TbTIF2, and TbRAP1) results in an elevated frequency of VSG switching (Jehi et al.,

2014b) (Jehi et al., 2014a) (Nanavaty and Li unpublished data).

1.9.1 Monoallelic expression

To ensure effectiveness of VSG switching, it is important to express only one VSG at any time, therefore monoallelic expression of VSG is critical for antigenic variation. As mentioned earlier, T. brucei cells have about 15 BESs that can express VSGs but only one BES is fully active at any given time. BESs are very similar in gene organization and have ~ 90% sequence identity, but T. brucei cells are able to fully transcribe only one BES in a mutually exclusive

fashion. Studies have suggested that monoallelic expression is controlled at multiple levels.

Figure 1.4 Mechanism of VSG switching A single VSG is expressed from a subtelomeric expression site. The promoter of expression site is indicated with a flag and transcription is indicated by arrow under the expression site. Active VSG gene before switching is indicated by red colored rectangle. Silent VSG genes are either present in long tandom arrays in subtelomeric region (shown as different colored rectangular boxes) or as single gene within expression sites. Three modes of VSG switching are shown. In gene conversion, silent VSG is duplicated into active VSG expression site and originally active VSG gene is lost. In telomere exchange, DNA between two-chromosome ends is exchanged. In in situ switch transcription at new expression site is activated and transcription from previously active expression site is turned off (Adapted from Taylor and Rudenko 2006).

Sequestering one BES: Allelic exclusion of the actively transcribing BES is a very important aspect of monoallelic expression and this is achieved by compartmentalization of the active BES (Navarro and Gull, 2001) (Chaves et al.,

1999). Transcription of the active BES takes place in a special nuclear compartment called expression site body (ESB) (Navarro and Gull, 2001). ESB is located outside of the nucleolus. In nucleolus, RNA polymerase I transcribes rRNA and procyclin genes while ESB is exclusively used by RNA polymerase I for transcribing the active BES. ESB only contains the active BES while all the inactive BESs are present elsewhere in the nucleoplasm (Chaves et al., 1998)

(Navarro and Gull, 2001). Not much is known about the nature and composition of ESB, but it is proposed to be a privileged location in the nucleus where RNA polymerase I is present at a high concentration and this location can be occupied by only one BES at any given time (Borst, 2002) (Navarro and Gull, 2001).

Moreover, it is suggested that the limitation of factors required for proper ESB assembly restricts the number of ESB to one per cell thus ensuring expression of only one BES (Vanhamme et al., 2001) (Stockdale et al., 2008).

Silencing of BESs: Proper silencing of BES outside the ESB is very important to maintain monoallelic VSG expression. Studies have suggested that multiple mechanisms are important for this regulation. Earlier studies suggested that transcription at the silent BESs is completely turned off to ensure monoallelic expression (Chaves et al., 1998) (Chaves et al., 1999). Contrary to this, later studies confirmed that the promoters of silent BESs are active because substantial and productive transcription of silent BESs could be detected in the 24

bloodstream form (Vanhamme et al., 2000). Closer analysis of the transcripts from silent BESs established that transcription from these silent BESs is aborted before it can transcribe the VSG gene (Vanhamme et al., 2000) (Kassem et al.,

2014). This suggested tight transcription control of VSG gene expression but not on transcription initiation at these silent BESs. Based of these studies, it is proposed that promoters of silent BESs could efficiently recruit the transcriptional machinery but the transcription from these silent BESs is not efficiently elongated and thus is prone to get aborted before it could transcribe the VSG gene

(Kassem et al., 2014). It is not clear if attenuation of these silent transcripts is the consequence of lack of elongation factors sequestered in ESB or some unknown factors force this attenuation.

Recent studies have established that chromatin structure plays an important role in regulating BES expression. Chromatin of silent BESs is densely packed with nucleosome while chromatin of the active BES is very relaxed and devoid of nucleosomes (Figureueiredo and Cross, 2010) (Stanne and Rudenko,

2010). It is not understood if the depletion of nucleosomes from the active BES is the consequence of active transcription or if it is a prerequisite for active transcription.

Studies have revealed that chromatin remodelers play an important role in regulating expression at silent VSGs. Depletion of the histone methyltransferase

TbDOT1B results in derepression of the silent VSGs by approximately 10-fold while the depletion of many other chromatin remodelers, such as histone 3, histone 1, and histone chaperons FACT (facilitates chromatin transcription), NLP 25

(nucleoplasmin-like protein), CAF-1b (chromatin assembly factor 1b) and ASF1a

(anti-silenceing factor 1a) resulted in derepression of silent BES promoters without effecting silent VSG expression (Alsford and Horn, 2012) (Povelones et al., 2012) (Denninger et al., 2010) (Narayanan et al., 2011). Moreover, recent study suggested that activation of a new BES is sufficient to attenuate the transcription of formerly active BES by an unknown feedback loop to ensure monoallelic expression (Batram et al., 2014).

Lastly, as all BESs are present at subtelomeric regions, telomere structure plays an important role in regulating VSG expression. Depletion of telomeric protein TbRAP1 leads to 10-100 folds of depression of all the silent BESs, underscoring the importance of telomeres in VSG expression regulation (Yang et al., 2009) (Pandya et al., 2013).

1.10 Telomeres

Telomeres are the physical ends of linear chromosomes that are required to maintain stability of the genome. Hermann Muller, while working on

Drosophila, first established the existence of telomeres when he noted that deletion of ends of the chromosomes led to the loss of chromosomes. Thus, he concluded that the natural ends of linear chromosomes are special and are required for maintaining chromosome stability. He termed the ends of chromosomes as telomeres (greek words “telo”, meaning "end," and mere, meaning "part) (Muller, 1938). 50 years later, sequencing of the chromosome ends of Tetrahymena revealed the actual structure of telomeres (Blackburn and

Gall, 1978) (Yao et al., 1981). We now know that telomeres are tandem arrays of simple DNA repeats of varying length. In most organisms, telomeric DNA consists of GT-rich repeat sequence. For example, telomere repeats of

Tetrahymena are 5’ TTGGGG 3’ while in case of human and T. brucei they are 5’

TTAGGG 3’ repeats (Blackburn and Gall, 1978) (Blackburn and Challoner,

1984). The number of telomeric repeats differs between species but is maintained within a certain range for a given species. For example, in budding yeast, telomere length is maintained between 270-300 bp while the length of human telomeres ranges from 2-15 kb (Shampay et al., 1984) (Moyzis et al.,

1988; Lansdorp et al., 1996). Although the bulk of telomeric repeats are composed of double stranded DNA, in most organisms, telomeres end with a protrusion of single stranded DNA at the 3’ end that is called G-overhang or 3’ overhang (Henderson and Blackburn, 1989). Sequence-specific proteins that can bind to the double-stranded and single-stranded DNA bind to telomeres to form a nucleoprotein complex. Studies have established that both DNA sequence and various telomere-binding proteins contribute for proper telomere functions.

Telomeres perform two main functions: 1. Solve the end protection problem. Telomeres provide a protective cap to chromosome ends and preserve genome stability by shielding natural chromosome ends from being recognized as double strand breaks (DSBs) (de Lange, 2009) (Stewart et al., 2012) 2. Solve the end replication problem. The terminal sequence of a linear eukaryotic chromosome is unable to be fully replicated by conventional DNA polymerases, which leads to the progressive loss of terminal sequences at chromosome ends. 27

This is called the end replication problem (Watson, 1972; Olovnikov, 1973). This problem stems from the use of small 5’ RNA primers by DNA polymerase to initiate synthesis of daughter DNA strand. These short RNA primers are removed at the end of the replication, creating a gap of 8-10 nucleotides at the end of the chromosome that cannot be filled by DNA polymerase. Most of the eukaryotes use telomerase, a ribonucleoprotein complex (RNP), to compensate for the loss of telomere sequence due to the end replication problem.The telomerase is a highly specialized reverse transcriptase enzyme that uses a short stretch of its integral RNA component as a template to synthesize a telomere DNA de novo.

1.10.1 The telomere G-overhang structure

As mentioned earlier, the termini of telomeres are not blunt-ended but rather have a single-stranded 3ʹ protrusion of the G-rich strand known as G- overhang or 3’-overhang. The G-overhang is a conserved feature of the telomere structure and is observed in mammals, yeast, ciliates, and trypanosomes. G- overhangs are present on both ends of the chromosome and their length is tightly regulated (Henderson and Blackburn, 1989). In Tetrahymena, the length of the

G-overhang is 14-21 nt, while in most of the vertebrates the G-overhang is much longer, ranging from 50-400 nt (Jacob et al., 2001) (Makarov et al., 1997). Loss of G-overhang results in dysfunctional telomeres, underscoring the importance of the G-overhang.

The G-overhang can fold back to invade the double-stranded region of the telomere resulting in a lasso like structure at the chromosome end called T-loop

(telomeric loop) (Figure 1.5). T-loops were first observed in vitro by performing electron microscope analysis of human telomeres (Griffith et al., 1999). The T- loop structure is now believed to be a conserved feature of eukaryotic chromosomes and has been observed in ciliates, trypanosomes, and plants (de

Lange, 2004) (Murti and Prescott, 1999)(Muñoz‐Jordán et al., 2001)(Cesare et al., 2003). The T-loop structure is proposed to be important for telomere capping function (De Lange, 2004)(de Lange, 2009). Formation of the T-loop results in folding back and tucking in of the very end of telomere and thus sealing the end of the chromosome. This sealing is believed to prevent the chromosome end from being recognized as DSB by DNA damage response machinery (Palm and

Figure 1.5 T-loop structure (a) Telomeres terminate in a 3´ G-rich single-stranded extension called the G- overhang. The G-overhang invades the duplex region of telomeres and assembles into a t-loop structure. (b) Electron microscopy image showing t-loop at both ends of the T. brucei chromosome (Muñoz‐Jordán et al., 2001).

de Lange, 2008). Moreover, it is proposed that T-loops negatively regulate telomerase activity at telomeres as the 3’ overhang required for the telomerase action is engaged in base pairing in T-loops.

Although T-loops provide an elegant model to explain the capping function of telomeres, T-loops are not observed in many species. In the ciliate Oxytricha nova, the telomere 3’ overhang is hidden by a complex of two proteins (On

TEBPα and On TEBPβ) that binds to 3’ overhang tenuously (Horvath et al.,

1998). In yeast, the T-loop structure is yet to be reported (de Lange, 2004).

Moreover, in yeast telomeres are proposed to loop-back on themselves via protein-protein interactions and thus provide a protection function. In summary, the G-overhang structure is a conserved feature and appears to contribute to chromosome end protection (discussed in more detail in chapter IV).

1.10.2 The telomere protein complex

Both duplex and single-stranded telomere DNA binding proteins have been identified to bind telomere DNA. Studies in different species have established that double-stranded telomere DNA binding proteins bind to DNA by a conserved myb domain while the single-stranded telomere DNA binding proteins use a conserved OB (oligonucleotide/oligosaccharide binding) fold to bind with 3’ overhang (Horvath, 2008). In addition to the proteins that directly bind to telomeric DNA, many other proteins are recruited to telomeres by protein- protein interactions. Together with telomeric DNA, these proteins make a

nucleoprotein complex that is critical for telomere function and length maintenance.

In S. cerevisiae, the duplex region of telomeric DNA is bound by scRAP1

(repressor activator protein 1). scRAP1 binds to duplex DNA using its central myb and myb-like domains and facilitates recruitment of many other telomeric proteins (Longtine et al., 1989) (König et al., 1996) (Figure 1.6). ScRAP1 recruits

Rif proteins (RAP1 interacting factors), Rif1 and Rif2, to the telomeres (Hardy et al., 1992) (Wotton and Shore, 1997). This protein complex of scRAP1, Rif1, and

Rif2 negatively regulates the telomere length (Wotton and Shore, 1997). ScRAP1 also recruits Sir proteins (Silent Information Regulator), Sir3 and Sir4, which are required for heterochromatin formation at the telomere (Hardy et al., 1992)

(Moretti et al., 1994) (Cockell et al., 1995) (Moretti and Shore, 2001).

Cdc13 binds to the 3’ overhang of telomeres and interacts with Stn1,

Ten1, and EST1 (ever shorter telomeres) (Lin and Zakian, 1996) (Bourns et al.,

1998) (Qi and Zakian, 2000) (Grandin et al., 1997) (Grandin et al., 2001b)(Figure

1.6). Cdc13 regulates recruitment of telomerase to the telomere thereby regulating telomere length (Pennock et al., 2001) (Chandra et al., 2001). Another protein that binds to S. cerevisiae telomeres is yeast Ku (yKu). yKu is a heterodimer composed of yKu70 and yKu80 and is an essential component of the non homologous end joining (NHEJ) machinery (Boulton and Jackson, 1996)

(Gravel et al., 1998) (Boulton and Jackson, 1998). It is unclear where yKu associates with telomeric DNA but its association with telomeres is critical for telomere function. Moreover, yKu is proposed to regulate recruitment of 31

telomerase to the telomeres through its interaction with telomerase RNA component (TLC1) (Peterson et al., 2001) (Stellwagen et al., 2003,).

A protein complex called Shelterin associates with the mammalian telomere (de Lange, 2005) (Figure 1.6). Shelterin is composed of TTAGGG

Repeat binding Factors 1 and 2 (TRF1 and TRF2), TRF1 Interacting Nuclear protein 2 (TIN2), RAP1, TPP1, and Protection Of Telomere 1 (POT1) (Palm and de Lange, 2008). Three of the Shelterin proteins directly interact with telomeric

DNA. TRF1 and TRF2 bind to duplex telomeric DNA as homodimers while POT1 binds to the 3’ overhang (Broccoli et al., 1997) (Bilaud et al., 1997) (Baumann and Cech, 2001). TRF1 and TRF2 recruit four Shelterin components to telomeres: RAP1, TIN2, TPP1 and POT1 (Li et al., 2000; Kim et al) (1999; Ye et al., 2004b) (Takai et al., 2011).

TRF1 and TRF2 both utilize the C-terminal DNA binding myb domain to bind to the telomeric duplex DNA and an internal TRF Homology domain (TRFH) to form a homodimer (Broccoli et al., 1997) (Bianchi et al., 1997) (Chapman et al., 2005) (Fairall et al., 2001). They do not directly interact with each other but share a common binding partner TIN2 (Broccoli et al., 1997) (Fairall et al., 2001)

(Kim et al., 1999) (Kim et al., 2004) (Ye et al., 2004a) (Houghtaling et al., 2004).

TRF1 plays an important role in telomere replication, presumably by recruiting

BLM (bloom syndrome) and RTEL1 (regulator of telomere length1) helicases, to remove replication blocks from the telomeric DNA (Figure 1.7) (Sfeir et al., 2009).

TRF2 is required for telomere protection (Sfeir and de Lange, 2012) (de Lange,

2010) (Celli and de Lange, 2005). Removal of TRF2 from the telomeres leads to 32

loss of the G-overhang and accumulation of MRN complex (Mre11, Rad50, and

Nbs1) (Dimitrova and de Lange, 2009). Accumulation of MRN complex leads to activation of ATM (Ataxia Telangiectasia Mutated) that triggers DNA damage response pathway at telomeres (Dimitrova and de Lange, 2009) (Denchi and de

Lange, 2007). Moreover, inhibition of TRF2 results in telomere fusions mediated by NHEJ (van Steensel et al., 1998) (Celli and de Lange, 2005) (Sfeir and de

Lange, 2012). Although the mechanism by which TRF2 blocks ATM activation and NHEJ at the telomeres is not clear, it is speculated that TRF2 may mediate these functions by facilitating T-loop formation (de Lange, 2010). Two independent studies support this hypothesis. Firstly, an in vitro experiment suggested that TRF2 could help to generate the T-loop on a telomeric substrate

(Stansel et al., 2001). Secondly, a recent finding suggests that TRF2 is also required for T-loop formation in vivo (Doksani et al., 2013). TRF2 is also important for the G-overhang generation by recruiting and regulating the activity of Apollo nuclease (Wu et al., 2010).

In contrast to scRAP1, RAP1 in mammalian cells does not bind to telomeric DNA directly rather it is recruited to telomeres by TRF2 (Li et al., 2000).

Overexpression of full length or truncated mutants of human RAP1 (hRAP1) results in telomere elongation (Li and de Lange, 2003). Based on this result, it is proposed that hRAP1, similarly to scRAP1, negatively regulates telomere length in cis (Li and de Lange, 2003). Overexpression and nucleoplasmic accumulation of RAP1 may lead to titration of a limiting RAP1 interacting factor away from the telomere bound RAP1 (Li and de Lange, 2003). Binding of this factor to telomeric 33

RAP1 may be required for proper telomere length homeostasis.In humans, RAP1 blocks NHEJ at telomeres while in mice, it blocks homology directed repair

(HDR) (Sarthy et al., 2009) (Sfeir et al., 2010) (Sfeir and de Lange, 2012).

TIN2 is recruited to the telomere by interacting with TRF1 and TRF2 (Kim et al., 1999). TIN2 also interacts with TPP1 and thus helps to bridge the Shelterin components that bind to duplex DNA (TRF1 and TRF2) and components that bind to single stranded (TPP1 and POT1) telomeric DNA. TIN2 is required to stabilize TRF1 and TRF2 at telomeres (Ye et al., 2004a) (Kim et al., 2004)

(O'Connor et al., 2006). TIN2 contributes to TRF2’s function of suppressing ATM and TIN2 is also required for efficient recruitment of TPP1/POT1 to the telomeres

(Abreu et al., 2010).

TPP1 forms a heterodimer with POT1 and is essential for the function of

POT1 at telomeres (Chen et al., 2007) (Wang et al., 2007). TPP1 interacts with

TIN2 and thus bridges POT1 to the central components of the Shelterin complex.

POT1 binds to the G-overhang and protects telomere ends from nucleolytic degradation. In complex with TPP1, POT1 prevents RPA mediated activation of

ATR (Ataxia Telangiectasia Mutated and Rad3 related protein) at telomeres

(Denchi and de Lange, 2007). Moreover, the TPP1/POT1 complex also regulates telomerase activity at the telomere end (Ye et al., 2004b) (Xin et al., 2007).

Figure 1.6 Telomere protein complex in different organisms Schematic repersentaion of telomeric proteins bound to telomere sequence.Sequence and length of telomeres vary among different species In S.cerevisiae and S.pombe telomeres are usally less than 0.5kb and are composed of imperfect reapeats of G-rich sequence. In vertebrates and T. brucie telomeres range from 2-50kb and are composed of TTAGGG repeats Similar to the duplex region of telomere sequence the length of single stranded region at the end of telomeres is specific for a species and varies among different spcecies. Conserved protein complex binds to single and double straned region of telomeres. Proteins shown in similar colors are close homologs.

Although POT1 can bind to the G-overhang by its conserved OB folds, studies have suggested that the interaction of POT1 with TIN2 through TPP1 is required for its localization to the G-overhang (Hockemeyer et al., 2007).

Disruption of the interaction between TIN2 and the TPP1/POT1 complex results in the removal of TPP1/POT1 from the G-overhang. Based on these data it has been proposed that the interaction of TIN2 with TPP1/POT1 is critical to block

RPA loading on to the G-overhang (Palm and de Lange, 2008) (Denchi and de

Lange, 2007).

In Schizosaccharomyces pombe, the telomere protein complex closely resembles that of the mammals. S. pombe telomeric proteins consist of TAZ1

(TRF1/TRF2 ortholog) that binds to the duplex telomere DNA, while spPOT1 and

TPZ1 (POT1/TPP1 ortholog) bind to the 3’ overhang (Cooper et al., 1997)

(Miyoshi et al., 2008). Similar to mammalian, spRAP1 (RAP1 ortholog) and POZ1

(that has similar functions of TIN2) connect TPZ1/spPOT1 to TAZ1 (Kanoh and

Ishikawa, 2001) (Miyoshi et al., 2008) (Dehé and Cooper, 2010) (Figure 1.6).

In addition to the above-mentioned proteins, another evolutionarily conserved complex, known as the CST complex has been implicated in the telomere maintenance. In S. cerevisiae, this complex is composed of Cdc13,

Stn1, and Ten1. Cdc13 binds to the 3’ overhang and recruits Stn1 and Ten1 to the telomere end. All three proteins of CST are required for telomere end protection and loss of any of these proteins results in telomere dysfunction

(Grandin et al., 2001b). Similarly to budding yeast, mammals have a conserved

CST complex (CTC1-Stn1-Ten1) that localizes to telomeres and is required for 36

proper telomere replication and maintenance (Miyake et al., 2009). In S. pombe,

Stn1 and Ten1 have been identified so far and their function at the telomeres seems conserved (Gao et al., 2007). Furthermore, several proteins that are known for their nontelomeric functions are also found associated with telomeres.

These include DNA damage repair proteins such as the Ku70/80 heterodimer,

MRN, poly ADP ribose polymerase (PARP) 1 and 2, Xeroderma pigmentosum group F (XPF/ERCC1) and DNA helicases and nucleases like BLM, RTEL1,

Werner syndrome (WRN), exonuclease (EXO1), and Apollo (Zhu et al., 2000)

(Dantzer et al., 2004) (Gomez et al., 2006) (Zhu et al., 2003) (Opresko et al.,

2002) (Crabbe et al., 2004) (van Overbeek and de Lange, 2006) (Wu et al.,

2010) (Wu et al., 2012).

1.11 Telomerase

The conventional replication apparatus is unable to completely replicate the chromosome ends (end replication problem), resulting in a gradual erosion of telomere DNA during each cell cycle. To compensate for this loss, most eukaryotic cells use telomerase (Flores and Blasco, 2010). Telomerase is a ribonucleoprotein enzyme that contains a highly conserved reverse transcriptase

(TERT) component and an associated RNA component (TR) used as a template for de novo telomere DNA synthesis.

Apart from these core factors, the telomerase requires several auxiliary factors to be functional in vivo. Some of these factors are required for proper processing of the telomerase complex, while others are required for telomerase

recruitment and activity at the telomere end. In S. cerevisiae, the telomerase is composed of three proteins EST1, EST2 (TERT), and EST3. Absence of any of these proteins results in the loss of telomerase activity in vivo while only EST2 and TR are required for in vitro activity (Lendvay et al., 1996). These results imply that EST1 and EST3 are auxiliary factors that are required for telomerase recruitment and activity at telomeres (Cohn and Blackburn, 1995) (Lingner et al.,

1997a). Studies of EST1 function suggest that it is required for the telomerase recruitment and activation. EST1 interacts with Cdc13 and this interaction is proposed to mediate EST2 recruitment to telomeres. (Evans and Lundblad,

2002) (Evans and Lundblad, 1999). In vitro studies suggest that the purified

EST1 protein can stimulate telomerase activity, implying that EST1 plays a role in telomerase activation (DeZwaan and Freeman, 2009). Role of EST3 is not known, although studies suggest that part of EST1 function may be mediated through its interaction with EST3 (Talley et al., 2011) (Tuzon et al., 2011).

TPP1/POT1 is implicated in both recruitment and activation of telomerase in vertebrates (Nandakumar et al., 2012) (Wang et al., 2007).

1.11.1 Telomerase reverse transcriptase (TERT)

TERT was first identified in the ciliate Euplotes aediculatus and was called p123 (Lingner and Cech, 1996). Sequencing of p123 revealed the presence of conserved reverse transcriptase motifs, which were closely related to the reverse transcriptase of non-LTR retrotransposons (Lingner et al., 1997b). By using this conserved motifs information, TERT was identified in different organisms

(mammals, worms, yeast trypanosomes, and plants) (Nakamura et al., 1997)

(Meier et al., 2006) (Counter et al., 1997) (Dreesen et al., 2005) (Fitzgerald et al.,

1999). TERT contains four conserved domains: N-terminal TEN domain, high- affinity telomerase RNA binding domain (TRBD), reverse transcriptase active site domain (RT) that contains 5 conserved motifs, and a C-terminal extension (TEC)

(Autexier and Lue, 2006). The process by which these different domains coordinate to form an active telomerase complex is not clear.

1.11.2 Telomerase RNA (TR)

In different organisms (protozoa, yeasts, vertebrates), TR molecules differ considerably in size and sequence, but harbor conserved structural elements

(Smekalova et al., 2012). In addition to the template for telomere extension, TR contains several structural elements necessary for the catalytic activity, processivity, stability, maturation, and localization of the telomerase. All known

TRs contain a pseudoknot, the template, and a template 5’-boundary element,

TBE (Smekalova et al., 2012).

In ciliates, TR is transcribed by RNA polymerase III. The mature TR molecule lacks posttranscriptional modifications such as a 5’ cap and a 3’ poly A tail (Lingner et al., 1994) (Greider and Blackburn, 1989). Moreover, due to its transcription by RNA polymerase III, the 3’ end of TR has heterogeneous tracts of uracil (Lingner et al., 1994). TR in ciliates contains a conserved pseudoknot and a TBE (Lingner et al., 1994). In Tetrahymena, interaction of TERT with TR is dependent on p65. The TRBD domain of TERT interacts with the TBE of TR

while p65 interacts with the 3’ stem loop structure (P4) of TR. Both these interactions are required for proper binding of TERT and TR (O'Connor and

Collins, 2006).

In S. cerevisiae, TR (TLC1) is transcribed by RNA polymerase II. Unlike ciliates, TLC1 contains both 5’ m7G (monomethylguanosine) cap and a poly A tail. During maturation, the 5’ m7G cap is replaced by a TMG (2,2,7- trimethylguanosine) cap and the poly A tail is removed (Singer and Gottschling,

1994) (Chapon et al., 1997) (Seto et al., 1999). Co-immunoprecipitation data suggested that TLC1 interacts with Sm protein complex and these proteins are required for proper biogenesis of TR (Egan and Collins, 2012). In addition to the pseudoknot, TLC1 also contains two stem–loop structures that are crucial for

EST1 and yKu binding (Osterhage and Friedman, 2009) (Stellwagen et al., 2003)

(Fisher et al., 2004).

Similar to the budding yeast counterpart, human TR (hTR) is transcribed by RNA polymerase II and has a 5’ m7G cap, although polyadenylation of hTR has not been verified so far (Fu and Collins, 2003). During maturation, hTR acquires a TMG cap at the 5’ end (Jády et al., 2004). hTR contains TBE, pseudoknot, hairpin-hinge-hairpin-ACA (H/ACA) loop and two conserved regions called CR4/5 loop and CR7 loop (Chen et al., 2000). The H/ACA loop is further characterized into two motifs: BIO box and CAB box. While the CAB box is conserved among different H/ACA containing non-coding RNAs, the BIO box is hTR specific (Egan and Collins, 2012). The TRBD region of TERT interacts with the template, pseudoknot, and CR4/5, while the TEN domain of TERT interacts 40

with template/pseudoknot region of hTR. These interactions are required for the telomerase activity (Lai et al., 2001) (Robart and Collins, 2011). The H/ACA loop of hTR is recognized by a heterotrimer of Dyskerin, NOP10, and NHP2, which are essential for maturation of hTR (Wang and Meier, 2004). H/ACA binding proteins transport hTR to Cajal bodies via the transport factors PHAX and

Nopp140 (Yang et al., 2000) (Boulon et al., 2004). In Cajal bodies hTR associates with TCAB1/WDR79 by its CAB box and this interaction is required for proper maturation of the telomerase RNP in Cajal body (Venteicher et al., 2009)

(Egan and Collins, 2012). The importance of proper hTR maturation is highlighted by the fact that mutations in the genes required for hTR biogenesis are responsible for dyskeratosis congenita (DC, a human disease due to telomerase deficiency) (Armanios, 2009) (Zhong et al., 2011).

In summary, maturation of TR is essential for proper localization of the telomerase. A number of proteins (i.e. dyskerin, NHP2, and NOP10) bind to and stabilize the newly transcribed TR. Loss of function of these proteins causes mislocalization of telomerase in the nucleus, thereby crippling the ability of telomerase to maintain telomeres. In addition, both yeast TLC1 and human hTR interact with Ku; this interaction has been proposed to be critical for recruiting telomerase to telomeres, as mentioned above. Apart from being used as a template to elongate teloemeres, TR is also required for the proper recruitment of telomerase holoenzyme to the telomeres.

In Chapter III, I will describe the identification and characterization of the telomerase RNA component of T. brucei. We found that it is transcribed by RNA 41

polymerase II and contains a poly A tail. TbTR contains an SL RNA sequence at its 5’ end and acquires a TMG cap during its maturation. Surprisingly, the TMG cap is not retained on the TbTR that makes a complex with TbTERT, suggesting additional steps in TbTR maturation that are absent in humans and in yeast (see chapter III for more details).

1.11.3 Recruitment of telomerase to telomeres

In humans, telomere-binding proteins exert positive and negative effects on the function of telomerase. It was shown that TRF1 inhibits telomerase function in vivo. Telomerase is recruited to telomeres by TPP1, which in turn is recruited to the telomere by TIN2 (Abreu et al., 2010) (Tejera et al., 2010). Once at telomeres, POT1-TPP1 facilitates telomerase action at the 3’ overhang by increasing the processivity of telomerase (Wang et al., 2007) (Latrick and Cech,

2010) (Nandakumar et al., 2012). Moreover, based on the studies that DC patients have mutations in TIN2 outside its TPP1-binding domain, it is proposed that TIN2 can directly affect telomerase action at the telomere ends (Walne et al.,

2008) (Sasa et al., 2012). This hypothesis is further supported by the finding that

TIN2 DC mutations do not cause telomere end protection dysfunction but only affect telomerase activity (Yang et al., 2011). Additionally, Ku has also been shown to interact with hTERT and hTR, however the functional relevance of this interaction is not known (Chai et al., 2002) (Ting et al., 2005).

In budding yeast, Cdc13 positively regulates the recruitment of telomerase by interacting with EST1 (Nugent et al., 1996) (Bianchi et al., 2004). In addition,

the Ku heterodimer has also been shown to play an important role in the recruitment of telomerase. Ku interacts with TLC1 directly, and has been proposed to promote the recruitment of telomerase to the telomeres (Stellwagen et al., 2003,) (Fisher et al., 2004). Although recent studies suggest that Ku is incapable of binding to both telomeric DNA and TLC1 simultaneously, making it harder to decipher the exact role of Ku in the telomerase recruitment (Pfingsten et al., 2012).

Both protein and RNA components of telomerase RNP are essential for the recruitment of telomerase. Certain N- and C-terminal regions of TERT, when mutated, cause loss of telomerase activity in vivo, with the reverse transcriptase activity retained in vitro. These regions are proposed to be important for recruitment of telomerase (Armbruster et al., 2001) (Banik et al., 2002). Although recent studies have provided some insights on the mechanism by which telomerase is recruited to and activated at the telomeres, the exact mechanism is unclear.

1.12 Telomere and telomere maintenance in T. brucei

T. brucei telomeres, similar to their human counterparts, are composed of

TTAGGG repeats (Blackburn and Challoner, 1984). The length of T. brucei telomeres ranges from 3-20 kb with an average size of 15 kb in the Lister 427 strain commonly used in our lab. Electron microscopic analysis confirmed the presence of the T-loop at the telomeres of T. brucei. Despite of the similar telomere length, T. brucei T-loops are much smaller (median size 1.1 kb) than

those of human telomeres (median size 14 kb) (Muñoz‐Jordán et al., 2001). In this study we characterized the G-overhang structure in T. brucei that is essential for T-loop formation.

So far, three conserved telomeric proteins have been identified in T. brucei. TbTRF (ortholog of TRF2), TbRAP1 (ortholog of RAP1), and TbTIF2

(ortholog of TIN2). TbTRF was the first telomeric protein to be identified in T. brucei (Li et al., 2005). Depletion of TbTRF results in an accute growth arrest.

TbTRF directly binds to duplex telomeric DNA in a sequence specific manner and is required for the G-overhang maintenance (Li et al., 2005). Moreover, loss of TbTRF results in an elevated frequency of VSG switching (Jehi et. al. 2014b)

TbRAP1 was identified in a yeast two-hybrid screen as a TbTRF interacting factor (Yang et al., 2009). TbRAP1 localizes to the telomeres and is also present in other nuclear compartments. This is reminiscent of scRAP1 that not only localizes to the telomeres but also is present at 5% of the promoters in the yeast genome (Yang et al., 2009). TbRAP1 plays an important role in the regulation of VSG silencing and switching (Yang et al., 2009) (Pandya et al.,

2013) (Nanavaty and Li unpublished data). Lastly, TbTIF2 (TRF interacting factor

2) is a telomeric protein important for maintaining subtelomeric and telomeric integrity (Jehi et al., 2014a). Loss of TbTIF2 results in an elevated frequency of

VSG switching, presumably due to the increased DSBs in the subtelomeric regions (Jehi et al., 2014a). Roles of TbRAP1 and TbTIF2 in telomere maintenance and protection are not well understood.

Unlike other organisms, telomeres in T. brucei grow at a constant rate of

6-8 bp/ population doubling (Bernards et al., 1983) (Van der Ploeg et al., 1984b).

Although the mechanism by which telomere homeostasis is regulated in T. brucei is not clear, studies have identified factors that are required for proper telomere length regulation. One of these factors is the Ku70/80 heterodimer. Ku70/80 is a conserved component of the NHEJ pathway and is important for telomere functions in both S. cerevisiae and humans. In S. cerevisiae, loss of yKu results in telomere shortening and extended G-overhangs outside of S phase. Deletion of yKu also leads to the inhibition of subtelomeric transcriptional silencing (TPE)

(Polotnianka et al., 1998) (Boulton and Jackson, 1998).

In humans Ku is essential and conditional knockout of Ku results in massive loss of telomeric DNA (Wang et al., 2009). In T. brucei Ku is non- essential. Deletion of TbKu80 had no effect on TPE but resulted in progressive shortening of telomeres, suggesting the telomere maintenance function of Ku is conserved in T. brucei (Conway et al., 2002a) (Janzen et al., 2004). The mechanism by which TbKu80 regulates telomere length is not known. In this study, we present data suggesting that similarly to yKu, TbKu is required for the activity of telomerase at telomeres. In T. brucei, TbTERT was identified on the basis of a conserved reverse transcriptase domain. Deletion of TbTERT leads to progressive shortening of telomeres (Dreesen et al., 2005). In this study we identify the telomerase RNA of T. brucei and confirmed that it serves as the template for telomere synthesis.

1.13 Significance of study

T. brucei uses antigenic variation of VSGs to circumvent host immune response. Several studies have now established that both telomeric proteins and telomere length play important roles in regulating antigenic variation. In order to better delineate the molecular mechanism by which antigenic variation is regulated, understanding the telomere structure is of paramount importance. In this study we uncovered several aspects of telomere structure and maintenance in T. brucei, thus advancing our current understanding about the telomere biology of this parasite.

Most of our knowledge about telomere biology is gained from the organisms that are closely related on the evolutionary scale (human, mice, yeast all belong to the same eukaryotic super-clade of Opisthokonta) (Walker et al.,

2011) (Adl et al., 2012) (Katz, 2012). In order to understand the convergence or divergence of different aspects of telomere biology, it is very important to study evolutionary distant organisms (Akiyoshi and Gull, 2013). These studies will not only help us to understand the fundamentals of telomere function and maintenance, but will also reveal the evolutionary history of telomere structure and function. Trypanosomes are one of the earliest organisms that branched from eukaryotic evolutionary tree and are evolutionary distant from most commonly studied eukaryotes (Akiyoshi and Gull, 2013). Among trypanosomes,

T. brucei is the most tractable member and is thus most suitable as a model organism. 10% of T. brucei genomic DNA is telomere DNA, making it convenient to study telomere functions through biochemical approaches. In addition, to the 46

genome sequence and genome wide RNAi library, a number of molecular tools are available in T. brucei, including highly efficient homologous recombination based gene targeting (gene knockout and endogenous gene tagging), inducible expression systems, and RNAi. Furthermore, a number of T. brucei telomeres are marked by unique subtelomeric virulence genes, allowing for examination of telomere lengths at a high resolution of individual chromosome end, making T. brucei a very attractive model system. Our study has not only provided new insights into the factors that contribute to telomere integrity in T. brucei but also developed several molecular assays to monitor the telomere structure. Thus our findings will further develop T. brucei as a model for telomere biology.

2CHAPTER II

Materials and Methods

2.1 Trypanosome strains and culture conditions

T. brucei bloodstream form Lister 427 strain expressing VSG2 were cultured in HMI-9 media at 37°C with 7.5% CO2. The SM strain is engineered to express T7 RNA polymerase and TET repressor in the WT 427 background

{Wirtz et al., 1999,}. The neomycin resistant gene is used for selection of T7 polymerase and TET repressor expression. SM is cultured in HMI-9 containing

2.5 μg/ml G418 (Sigma) and is served as the parent for other bloodstream form strains generated in this study. Wild-type T. brucei 427 procyclic form is cultured in SDM-79 media at 27ºC and is the parent for TbTR deletion procyclic form strain. Procyclic strain engineered to express T7 RNA polymerase and TET repressor (strain 29-13) is served as the parental for TbKU80 deletion procyclic form strain.

2.2 Transfection

T. brucei cell transfection was done using a Nucleofector with Basic

Parasites Buffer 1 provided by the manufacturer (Lonza). Program X-001 and W-

14 were used to transfect BF and PF cells, respectively.

2.3 Genomic DNA isolation:

Total genomic DNA was isolated as previously described (Muñoz‐Jordán et al., 2001). Brifly 200 million BF or PF cells were washed and resuspended in 1 ml of TNE (10 mM Tris pH 7.4, 10 mM EDTA, 100 mM NaCl). 1 ml of

TNES/proteinase K (10 mM Tris pH 7.4, 100 mM NaCl, 10 mM EDTA, 1% SDS +

100 µg/ml proteinase K (Roche)) was added and samples were incubated for 16 hours at 37°C. After overnight incubation DNA was extracted using phenol/chloroform. DNA was precipitated with sodium acetate and isopropanol and gently spooled out and was resuspended in 300 ul TNE+100 mg/ml RNase.

After 2 hour incubation with RNase at 37°C, 300 ul of TNES/protrinase K was added and reaction samples were incubated for 1 hour at 37°C. Following incubation, DNA was extracted and precipitated as mentioned above and resuspended in 100 ul of TE (10 mM Tris pH 7.5/1 mM EDTA).

2.4 Adaptor Ligation Assay

The adaptor ligation assay was performed as previously described (Jacob et al., 2001).with some modifications. The guide oligonucleotides were separately annealed to the unique oligonucleotide and ligated onto genomic DNA. 70 pmol of unique oligonucleotide 5’ CCCTATAGTGAGTCGTATTA 3’ was radiolabeled

with 30 uci of γ32P-[ATP] using 10 U T4 polynucleotide kinase (NEB) in 1 x T4

PNK buffer in a 30 μL reaction for 1 hour at 37ºC. The reaction products were purified using a Qiagen nucleotide removal column and eluted in 75 μL of EB.

Radiolabeled unique oligonucleotides were annealed separately to guide oligonucleotides to form different adaptors. Seven different annealing reactions consisted of 10 μL of radiolabeled unique oligonucleotides (approximately 9 pmol) added to 10 pmol of one of seven guide oligonucleotides. Annealing was performed by boiling unique/guide oligonucleotide combinations for 5 min and allowing the reaction to cool to room temperature slowly. 1.5-2 μg genomic DNA was ligated using 0.5 U DNA ligase (NEB) to each annealed adaptor overnight at

16ºC. Next day restriction digestion of ligated products was performed using frequent cutting enzymes AluI and MboI. Digestion is kept overnight at 37ºC.

Digested product is separated by electrophoresis in a 0.7% agarose gel. 20X20 cm gel cast is used to prepare the gel and electrophoresis is done for 1,000

Volt•hours. After electrophoresis, the ETBR-stained DNA is visualized under UV light, and a picture of the gel is taken. This picture was served as a loading control. Subsequently, the gel is dried using a Gel dryer, and the dried gel was wrapped in plastic wrap and exposed to a phosphorimager screen for overnight.

Quant J software was used to quantify the signal.

The final signal for each adaptor is calculated by dividing normalized signal for each adaptor by the normalized signal of NS adaptor (where normalized signal for each adaptor equals the Signal intensity of the adaptor from phosphorimager divided by the signal intensity on the ETBR gel). 50

2.5 EXO-T treatment of genomic DNA

DNA from 200 million cells (30-50 µg) of genomic DNA was incubated with 50 U of EXO-T (NEB) for 16 hours. After incubation, DNA was extracted by phenol/chloroform-spooling method. Control DNA without EXO-T was incubated and extracted same way as EXO-T treated samples.

Table 2.1 Guide oligonucleotide sequence (Permutations in bold)

Guide oligo1 TG1 5’ ACGACTCACTATAGGGCCCTAA

Guide oligo2 TG2 5’ ACGACTCACTATAGGGCCTAAC

Guide oligo3 TG3 5’ ACGACTCACTATAGGGCTAACC

Guide oligo4 TG4 5’ ACGACTCACTATAGGGTAACCC

Guide oligo5 TG5 5’ ACGACTCACTATAGGGAACCCT

Guide oligo6 TG6 5’ ACGACTCACTATAGGGACCCTA

Guide oligoNS TGNS 5’ ACGACTCACTATAGGGTGGGTG

2.6 Single telomere length analysis (STELA)

STELA was adapted from (Sfeir et al, 2005) with modifications. Briefly,

100 ng genomic DNA was ligated to each guide oligonucleotide (10 pmole) at

16°C for 16 hrs in 10 μl ligation reaction containing 1 × ligase buffer and 200 U

T4 ligase (NEB). 2 ul of the ligated DNA was used for subsequent PCRs. PCR reactions (16 cycles of 95°C for 15 s, 50°C for 20 s, and 68°C for 30 s) were carried out in 25 μl containing 0.2 μM primers. Forward primer (5’

TTAGGGTTAGGGTT 3’) and common guide primer (5’ ACGACTCACTATAGGG 51

3’) are used, and 2U of TAQ polymerase (NEB) was used in the reaction. PCR products were resolved on 0.7% agarose gels in separate lanes. After electrophoresis, DNA was denatured and transferred nylon membranes (Hybond

N, GE healthcare), followed by UV cross-linking and hybridization with a telomeric probe. Signals were detected by Phosphorimager screen (GE healthcare). Quant J software was used to quantify the signals, and the signal from NS oligonucleotide was used to normalize all other signals. Local average option was selected as a background in quantification.

2.7 T7 EXO nuclease treatment of Genomic DNA

1.3 µg of genomic DNA was incubated with 10 U of T7 exo (NEB) for 35 min at 25ºC. After incubation, DNA was precipitated and used for STELA reaction.

2.8 Ligation mediated primer extension (LMPE)

LMPE was performed as previously described (Jacob et al., 2001). with some modifications. Unique oligonucleotide was phosphorylated with non- radioactive ATP while all 7 guide oligonucleotides were separately end-labeled with P32. Unique oligonucleotide was annealed separately to each radiolabeled guide oligonucleotide. 10 pmol of each unique and guide oligonucleotide was used to generate adaptor similarly as described for adaptor ligation assay. The adaptor was ligated to 1-2.5 µg of genomic DNA. Ligation was performed at 16°C overnight with 0.5 U of T4 DNA ligase. Next day DNA was extracted with phenol/chloroform and after ethanol precipitation DNA was resuspended in 10 ul

ddH2O. Primer extension was performed at 30°C for 90 min in 20 µl reactions containing 10 µl of eluted DNA, 0.5 mM dATP, 0.5 mM dCTP, 0.5mM dTTP, 50 mg/ml bovine serum albumin (BSA), 1 X NEB buffer 2, and 3 U of T4 DNA polymerase. The samples were then denatured and separated on 12% polyacrylamide denaturing sequencing gels.

Table 2.2 Sequence of Top oligonucleotides.

Top-TG1-36nt

5’GTGTCACGAATGGCTACAACGTTTGAGTATGTAGTTGGATGTAGTTAGT

ATTAGGG

Top-TG1-26nt

5’ GTGTCACGAATGGCTACAACGTTTGAGTATGTAGTTGGATTTAGGG

Top-TG1-16nt

5’ GTGTCACGAATGGCTACAACGTTTGAGTATTTAGGG

Top-TG4-26nt

5’ GTGTCACGAATGGCTACAACGTTTGAGTATGTAGTTGGATGGGTTA

Top-TG4-16nt

5’ GTGTCACGAATGGCTACAACGTTTGAGTATGGGTTA

(Sequence in bold represents overhang after annealing to the bottom oligonucleotide and sequence in red represents the complementary sequence for unique/guide adaptor)

2.9 Preparation of artificial telomere substrate for LMPE

Multiple oligonucleotides were designed to produce different lengths 3’ protrusion upon annealing with complementary sequence. The oligonucleotide whose sequence produced the 3’ protrusion was called as ‘Top oligo’ (see Table

2.2 for sequence detail) while the complementary sequence oligonucleotide was called ‘Bottom oligo’: 5’ GTTGTAGCCATTCGTGACAC 3’. To make a telomeric overhang substrate 10 pmole of ‘Top oligo’ was mixed with 10 pmole of ‘Bottom oligo’. The mixture of Top/Bottom oligonucleotide was boiled for 5 min and allowed to cool slowing to room temperature. 1 ul of the duplex was used to perform LMPE instead of genomic DNA.

2.10 Southern blot analysis

Genomic DNA was isolated using DNAzol method. 2-5 µg of genomic

DNA was digested overnight with desired endonuclease in not more than 40 µl of reaction volume. DNA was separated on 0.7% agarose gel prepared in 0.5xTAE at 100 V until the orange G dye reaches the bottom of the gel. The gel was scanned on Typhoon FLA 9140 by placing a ruler next to it. DNA was depurinated (0.25 M HCL) for 30 minutes followed by denaturation (1.5 M NaCl;

0.5 M NaOH) for one hour (buffer was changed after 30 minutes). Finally the

DNA was neutralized (1 M Tris 7.4; 1.5 M NaCl) for one hour and blotted onto hybond nylon membrane (GE healthcare life sciences) in 20 x SSC (3 M NaCl,

0.3 M sodium citrate). After overnight blotting, the membrane was cross-linked

(Stratalinker UV crosslinker) and pre-hybridized for ~1 hour in CHURCH mix (0.5

M NaPi pH 7.2, 4 mM EDTA pH 8.0, 7% SDS, 1% BSA) at 65ºC. Hybridization was done at 65ºC overnight using the CHURCH mix and appropriate radiolabeled probe. The blot was washed three times using CHURCH wash (40 mM NaPi pH 7.2, 1 mM EDTA pH 8.0, 1% SDS) for ~15 minutes each at 65ºC, wrapped, and exposed to a phosphorimager screen (GE Healthcare Life

Sciences).

2.11 Northern blot analysis

Approximately 100 million T. brucei cells were used to isolate total RNA using STAT-60. RNA was separated on 1.5% agarose gel in 1 x MOPS and formaldehyde. RNA samples were added to loading buffer (1.5 μl 10 x MOPS,

2.6 μl 37% formaldehyde, 7.5 μl formamide, 3 μl 5 x loading buffer and ethidium bromide) and heated for 10 min at 65ºC. Gel was run in a fume hood in 1 x

MOPS buffer at 50V for 2-3 hrs. Quality and loading of RNA was determined by ethidium bromide staining. Gel was washed in ddH2O for 30 min to remove formaldehyde. Following the wash, gel was blotted onto a nylon hybond membrane (GE healthcare life sciences) for overnight in 20 x SSC (3 M NaCl, 0.3

M sodium citrate). RNA was crosslinked to the membrane and prehybridized at

55ºC for 50-60 min using the CHURCH mix. Blot was hybridized with appropriate probe at 55ºC for overnight. Following hybridization blot was washed three times for 30 minutes each at 55ºC using the CHURCH wash. After washing, the blot was exposed 1 hr to overnight depending upon the signal to a Phosphorimager screen (GE healthcare life sciences).

2.12 Reverse transcription and quantitative Real time PCR

Approximately 100 million T. brucei cells were used to isolate total RNA using STAT-60. QIAGEN RNAeasy kit was used according to the manufacturer recommendation to clean and treat RNA samples with DNAase. M-MLV

(Promega) reverse transcriptase kit was used according to the manufacturer’s protocol to synthesize cDNA. Bio-Rad iTaq SYBR Green supermix with ROX was used according to the manufacturer’s protocol to perform Quantitative RT-PCR.

DNA Engine Opticon 2 (Bio-Rad) system was used to perform real time PCR. To quantify the levels of mRNA the following formula was used, and the input amount of mRNA was normalized by using β-tubulin mRNA as a reference.

Ct-control-Ct-induced Ct value for loading control = Ct control/ Ct induced sample = 2

Next enrichment or decrease in mRNA level of the target gene is calculated by

Ct method.

Ct-control target gene-Ct-induced target gene Ct value for target gene = 2

Ct-control target gene-Ct-induced target gene Ct-control-Ct-induced Ctvalue for target gene=2 /2

Control samples are without induction (-Dox) and induced samples are induced for 24 hrs or 36 hrs as described (+Dox).

2.13 RNA Immunoprecipitation (RNA IP)

All steps in this protocol are performed at 4ºC unless indicated otherwise.

Dynabeads were prepared according to manufacturer recommendations. 200 million bloodstream form parasite cells were harvested and washed once with

TDB. Cell pellet was suspended in 1 ml of NET-5 buffer (40 mM Tris-HCl, pH 7.5,

420 mM NaCl, 0.5% Nonidet P-40,) containing 2 mg/ml aprotinin A, 1 mg/ml leupeptin, 1mg/ml pepstatin A and 10 units of RNAsin. Freezing and thawing of cells suspension was repeated 4 times to lyse the cells. Cells were froze at -80ºC and thawed on ice. Lysate was cleared by centrifugation at 15,000 g for 30 mins.

Cleared lysate is added to a fresh eppendorf tube and equal amount of NET1.5

(40 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.5% Nonidet P-40) containing 2 mg/ml aprotinin A, 1 mg/ml leupeptin, 1mg/ml pepstatin A and 10 units of RNAsin is added to bring down salt concentration. 100 μl of lysate is taken out as input and the rest of the lysate was divided into two equal volumes. RNA was extracted by phenol/chloroform and precipitated with 1/10 volume of sodium acetate and 2.5 volumes of chilled ethanol. 1 μl of 20 mg/ml glycogen was added to increase the recovery of RNA and to help visualize the pallet. Samples were stored overnight at -20ºC. Samples were cleaned using QIAGEN RNeasy kit was used according to the manufacturer recommendation to clean and treat RNA samples with

DNAase. qRT-PCR was performed as described earlier.

2.14 Cloning of terminal fragments of telomeres

The unique oligo 5’-CCCTATAGTGAGTCGTATTA-3’ was treated with polynucleotide kinase and ATP and annealed to the Guide oligo 5’-

ACGACTCACTATAGGGACCC-3’ to generate the adaptor. 10 pmole of adaptor was ligated with 1 µg of genomic DNA followed by PCR using 5’-

TTAGGGTTAGGGTTAGGGTTAGGG-3’ and the Guide oligo as primers. The

PCR product was then inserted into pGEM-T-easy vector (Promega) and the resulting plasmid was sequenced with T7 and Sp6 primers.

2.15 Telomerase activity assay

Direct telomerase activity assays were performed according to published procedure using (TTAGGG)3 as the substrate. 5 μl of the sample was electrophoresed over a 10% urea-polyacrylamide sequencing gel.

For the PCR-based TRAP assay, 10 million T. brucei cells were lyzed in

100 μl of CHAPS buffer from the TRAPeze© Kit (Millipore). In each reaction, 0.1–

0.5 μg of protein extract, 40 U of RNasin, 2 pmole of γ-32P end-labeled TS primers, and reverse primers for PCR amplification were mixed with 0.05 mM of dNTP, 20 mM TrisCl pH 8.3, 1.5 mM MgCl2, 63 mM KCl, 0.05% Tween 20 and 1 mM EGTA. 20 ng of RNase A was added together with the cell lysate in reactions treated with RNase. Telomerase-mediated primer extension was carried out at

30°C for 30 min followed by PCR amplification using the TS and reverse primers.

Products from TRAP assays were analyzed on 12.5% nondenaturing PAGE.

2.16 Preparing radioactive probe

To radiolabel the probe, 70 ng-100 ng of the probe DNA was mixed with 5 ng of random hexamer oligonucleotide (in a total volume of 41 μl). DNA was denatured and annealed to random hexamer oligonucleotides by heating the reaction to 100ºC for 5 min and then immediately kept on ice. Nucleotide mixture containing dATP, dGTP, and dTTP is added to the final concentration of 0.25 mM and 3 μl of 32P-alpha-dCTP (3000 Ci/mmol) was added to a final volume of 49

μl. Random primers were then extended using DNA polymerase (Klenow fragment New England Biolabs) in the final volume of 50 μl. The reaction was incubated at room temperature for 90 minutes. Reaction was terminated by adding 50 μl of TNES buffer (10 mM Tris pH7.4, 10 mM EDTA, 100 mM NaCl,

1%SDS). The probe was column purified using G-50 beads Sepharose (in 3 ml syringe) and eluted using TNES buffer. The probe was denatured at 100ºC for 5 min and filtered through a 0.22 μm syringe filter.

For in-gel hybridization of telomeric probe, 50 ng of TELG (TTAGGG)4 was radiolabeled with 30 µci of γ32P-[ATP] using 10 U T4 polynucleotide kinase

(NEB) in 1 X T4 PNK buffer in a 30 μL reaction for 1 hour at 37ºC. After incubation, radiolabelled oligonucleotide was purified using Qiagen nucleotide removal kit. Purified probe was mixed with 25 ml of CHURCH mix and added to the blot for hybridization.

3CHAPTER III

A Trans-spliced Telomerase RNA Dictates Telomere Synthesis in

Trypanosoma brucei

3.1 Introduction

Telomeres, the nucleoprotein complexes located at the ends of linear chromosomes, are essential for genome stability. Because conventional DNA polymerases are incapable of replicating the linear DNA ends completely, most eukaryotic cells use telomerase, an RNA-protein complex, to maintain telomeres

(Greider and Blackburn, 1987) (Lingner and Cech, 1998) (de Lange, 2009). The telomerase RNA is an integral component of telomerase that dictates the RNA template-dependent telomere synthesis (Greider and Blackburn, 1989) (Collins,

2008). Lack of telomerase activity leads to telomere shortening in a number of eukaryotic organisms including human, mouse, plant, worm, budding and fission yeasts, and protozoan parasites such as Plasmodium falciparum and

Trypanosoma brucei (Harley et al., 1990) (Blasco et al., 1997) (Fitzgerald et al.,

1999) (Meier et al., 2006) (Lendvay et al., 1996) (Nakamura et al., 1997) (Bottius et al., 1998) (Dreesen et al., 2005). Telomerase contains a core protein

component, Telomerase Reverse Transcriptase (TERT) that provides the catalytic activity and an RNA component (TR) that provides the template for telomere synthesis (Cech, 2004). TERT belongs to the reverse transcriptase

(RT) family and contains an RT domain with all RT-specific motifs (1, 2, and A–E motifs) (Podlevsky and Chen, 2012). Unlike classical RTs, telomerase requires an obligatory internal RNA component as a template for DNA synthesis and can copy the template sequence repetitively through efficient translocation. TR can also contribute greatly to the processivity of the telomerase (Berman et al., 2011)

(Qi et al., 2012). Hence, dysfunctional TR leads to telomere length shortening.

Although the domain structures of TERT are conserved across different species, the TR molecules from different organisms vary greatly in sequence and size (from ~150 nt in ciliates to ~1,800 nt in P. falciparum (Greider and

Blackburn, 1989) (Feng et al., 1995) (Chen et al., 2000) (Singer and Gottschling,

1994) (Leonardi et al., 2007) (Chakrabarti et al., 2007). However, a common theme in TR secondary structure has been observed (Theimer and Feigon,

2006), which consists of a single stranded template, a pseudoknot, a Template

Boundary Element (TBE), and a telomerase-interacting domain. Mutations in the

TR template can introduce mutations into the telomere sequences. This can affect the active-site functions of telomerase due to altered enzyme-substrate interaction (Gilley et al., 1995). Mutated telomere sequences may not be recognized by telomere DNA binding factors, which can also lead to telomere dysfunctions.

T. brucei is a protozoan parasite that causes African trypanosomiasis in humans. One major reason for persistent T. brucei infection is that it undergoes antigenic variation and regularly switches its surface antigen (VSG) to evade the host’s immune responses (David Barry and McCulloch, 2001). There are more than 2,500 VSG genes in T. brucei genome (Berriman et al., 2005), but only one

VSG is expressed exclusively from one of multiple, nearly identical subtelomeric

VSG expression sites at any time (Cross, 1975). In T. brucei, telomerase activity imparts the predominant mechanism for telomere maintenance (Cano et al.,

1999), and telomere elongates at an average rate of 4–6 bp per population doubling (PD) during continuous cell growth (Bernards et al., 1983). The TbTERT gene has been identified, and its deletion led to a steady telomere shortening at a rate of 3–4 bp/PD (Dreesen et al., 2005). Interestingly, when the active VSG- marked telomere is shortened to ~1 kb, the VSG switching rate is increased ~10 fold (Hovel-Miner et al., 2012). In addition, TbRAP1, an intrinsic T. brucei telomere component, is essential for regulation of subtelomeric VSG gene expression (Yang et al., 2009). Therefore, understanding the telomerase- mediated telomere maintenance in T. brucei can help to elucidate the mechanisms of T. brucei pathogenesis.

Figure 3.1 Identification of T. brucei telomerase RNA (a) Comparison of telomerase RNA template sequences among human (host), mouse, non-pathogenic and pathogenic protozoa. (b) Schematic representation of the computational search method used to identify putative TbTR gene (see text for details). E1 and E2 represent exons of a protein- coding gene. Inset shows sequence conservation, identified by multiple sequence alignment program ‘Multalin’ (http://multalin.toulouse.inra.fr/multalin/), in the putative TbTR ‘template’ adjacent region among 5 different Trypanosoma species. (Tbr, T. brucei brucei; Tbg, T. brucei gambiense; Tco, T. congolense; Tvi, T. vivax; Tcr, T. cruzi).(Experiment & analysis performed by Dr. Kausik Chakrabarti’s lab). 63

Among all the parasitic protozoa for which genome sequences are now available, T. brucei is the most amenable to genetic and biochemical studies due to their rapid growth (as short as 6 hrs/PD), simple in vitro culturing procedures, efficient knockout/knockin mediated by homologous recombination, and availability of useful molecular tools such as inducible expression (Wirtz et al.,

1999) and RNAi (Kolev et al., 2011). Particularly, T. brucei telomere sequence

(Blackburn and Challoner, 1984) and T-loop structure (Muñoz‐Jordán et al.,

2001) (De Lange, 2004) are identical to those in humans, and the telomere protein complex is largely conserved between T. brucei and vertebrates (Yang et al., 2009) (Li et al., 2005) (Jehi et al 2014a). Therefore, T. brucei can serve as a useful model system for dissecting the structure and function of telomerase in pathogenic protozoa and for comparative analysis of telomerase function and evolution in general.

In this study, we identify the putative T. brucei TR gene through an in silico approach and further provide in vivo characterization of TbTR’s native folding and activity. Additionally, we determine TbTR’s in vivo interaction with TbTERT and its critical role in telomere maintenance in T. brucei.

3.2 Results

3.2.1 Identification of a putative TbTR gene

Telomerase RNA length and sequence is highly divergent across different spices, hindering the use of sequence-based homology to identify telomerase

RNA. In contrast the template region of telomerase RNA is generally

corresponds to 1.5 telomeric repeats with some exceptions (human, mouse and

K. lactis, whose templating regions are 11). Cloning of T. brucei telomeres and in vitro studies suggested that the T. brucei telomerase RNA component template region consists of 5´-CCCTAACCC-3 (Figure.3.1a). To identify the telomerase

RNA gene, Dr. Kausik Chakarborty’s lab performed a BLAST search for short, nearly exact matching sequences in the T. brucei genome. In this process he identified a number of initial TR candidates (Figure.3.1b), many of which overlapped with annotated ORFs, structural RNAs (rRNAs, tRNAs or snoRNAs) and sequences at the telomeric ends of T. brucei genome (Figure.3.2b). Dr.

Chakrabarti discarded those initial candidates overlapping annotated protein or

RNA-coding regions or spanning telomeric repeat sequences and narrowed the list down to a few hits that showed a high degree of sequence similarity around the putative TbTR template region (Figure.3.1b). Dr. Chakrabarti then extended the comparative genome analysis in both directions flanking the TbTR template for all hits to search for syntenic regions among five related Trypanosoma subspecies (Figure.3.1b). Eventually he identified a unique sequence of ~200 nt in T. brucei genome that gave the highest scoring hits when used as BLAST query against each of the other Trypanosoma genomes. These hits are highly conserved in multiple regions including the putative template domain among five

Trypanosoma sub-species (Figure.3.1b, inset).

I performed Northern analysis using a putative TbTR specific probe. We identified a transcript of ~900 nt (Figure.3.2a), which we propose to be the putative TbTR. We found that TbTR is expressed at similar levels in the 65

Figure 3.2 Analysis of TbTR expression (a) Northern analysis of TbTR expression in WT BF and PF cells. Top blot hybridized with a 900 bp TbTR specific probe. Bottom, Ethidium bromide- stained RNA gels showing rRNA species, served as a loading control. (b) Northern analysis of TbTERT expression in WT BF and PF cells. Top blot hybridized with TbTERT specific probe. Bottom, Ethidium bromide-stained RNA gels showing rRNA species, served as a loading control.

bloodstream form (BF, when T. brucei is inside a mammalian host) and procyclic form (PF, when T. brucei is inside the midgut of its insect vector) stages

(Figure.3.2a). Likewise, TbTERT appears to be expressed at similar levels in both BF and PF cells (Figure.3.2b).

To map both ends of TbTR, Dr. Chakrabarti’s lab used RNA Ligase-

Mediated RACE (RLM-RACE), which selects full-length mRNAs from total RNA by enzymatic treatments followed by identification of cDNA ends via adapter mediated PCR (Schaefer, 1995). One prominent PCR product was identified from the 5’ RACE reaction in addition to two minor amplicons of larger sizes

(Figure.3.3a). Sequencing analysis of the RACE fragments identified the major 5’ end nucleotides of the TbTR transcript (Figure.3.3b). The 3’ end of TbTR also showed ambiguity in end processing, generating three different 3’ RACE

products (330, 280 and 150 bp). Cloning and sequencing these RACE products identified alternative 3’ ends of TbTR (Figure.3.3c).

3.2.2 Association of TbTR and TbTERT in vivo

To examine the interaction between TbTR and TbTERT, Dr. Chakrabarti’s lab raised peptide antibodies against TbTERT using three TbTERT N-terminal peptides. Immunoprecipitation (IP) with one of these antibodies allowed us to obtain a major protein species of 110 kDa and a minor one of 130 kDa

(Figure.3.4a, arrows). In the subsequent western analysis of the immunoprecipitated products, ~40% of the input sample (the 130 kDa band) was pulled down by anti-TbTERT antibody but not by the pre-immune serum

(Figure.3.4b, asterisk). MALDI-TOF mass spectrometry analysis of this IP product identified several peptides that match the published TbTERT sequence, confirming that this is indeed TbTERT. At this point the relationship between the two different-sized bands detected on the SDS-PAGE is unclear.

Subsequently, Dr. Chakrabarti’s lab isolated RNA from the DNase-treated

IP product and performed two different reverse transcription reactions, one with random primers (RP) and the other with Oligo dT primers (OP). RT-PCR analysis with TbTR internal primer pairs spanning the RNA template domain identified a band of expected size of about 300 bp (Figure.3.4c, lanes 2 & 4) from both RP and OP samples but not from a control reaction lacking RT (Figure.3.4c, lane 3).

Therefore, this data suggests that TbTR is polyadenylated, which associates with

TbTERT in vivo. Immunopurified T. brucei telomerase displayed excellent

Figure 3.3 Full length TbTR 5’ RLM-RACE determined the possible transcription start sites of TbTR after sequencing the RT-PCR product from RACE reaction. A solid line marks the major PCR product, while two faint bands are marked with dotted lines, which could represent the minor species with alternative. (b) Sequence alignment of the major 5’ RLM-RACE product in (a) and the TbTR gene. (c) TbTR encoding DNA sequence on chromosome 11, Plus (+) strand. ψTbTR ends determined by 5’ and 3’ RLM RACE. * denotes high frequency SL-containing reads mapping to a particular position of trans-splicing at the 5’ end OR poly A containing reads mapping to a particular position of polyadenylation at the 3’ end determined by RNA-seq or Oligo dT RT-PCR. # IP and RTPCR sequencing (single experiment). Minimum TbTR transcribing region is shown in gray shade. M represents molecular weight marker. (Experiment & analysis performed by Dr. Kausik Chakrabarti’s lab).

stability and was able to elongate T. brucei telomere substrate in a direct telomerase activity assay (Figure.3.4d). This activity is sensitive to RNase, but addition of RNase inhibitor, RNasin, can restore the activity during RNase treatment, indicating that this activity depends on an essential RNA component.

Figure 3.4 Molecular validation of TbTR (a) SDS-PAGE profile showing a ~110 kDa and ~130 kDa band in TbTERT IP sample. (b) IP and western blotting using an anti-TbTERT peptide antibody. The pre-immune serum was used as a negative control (Pre). Asterisk represents the TbTERT protein. (c) TbTR is associated with TbTERT. T. brucei cell extract was immunoprecipitated with anti-TbTERT peptide antibody, and the IP product was reverse transcribed after DNase treatment with (+) or without (-) RT using either Random Primer (RP) or Oligo dT primer (OP), followed by PCR amplification with TbTR specific primers. Genomic DNA (gDNA) was used as a control. Extra spaces between the marker and lane 2 on the same gel were deleted from the image to save space. (d) Telomerase activity assay using TbTERT IP product pretreated with RNase A or RNasin. The periodicity of repeat was determined by radiolabeled input oligos of 18 and 24 nt (marked on the left). The product with one TTAGGG repeats addition is marked as (+6). M represents molecular weight marker. (Experiment & analysis performed by Dr. Kausik Chakrabarti’s lab).

3.2.3 Deletion of TbTR leads to progressive telomere shortening

As an independent approach to validate the identity of TbTR, I established

BF and PF TbTR null cells by replacing the TbTR endogenous alleles with

Hygromycin resistance (HYGRO) and Blasticidin S resistance (BSD) genes sequentially. I verified the genotype of TbTR double knockout (DKO) cell lines by

PCR analysis (Figure.3.5b) and Southern blotting (Figure.3.5d). Northern

Figure 3.5 Confirmation of TbTR deletion (a) Genomic map of the TbTR locus for the WT allele (top) and the deletion allele (bottom). The P1 primers are specific to a DNA region located upstream of TbTR and P2 primers are specific to the TbTR gene. Restriction enzyme sites are indicated. (b) Confirmation of the endogenous WT alleles in WT cells and the deleted alleles in TbTR DKO candidates by PCR. Top, BF cell lines; Bottom, PF cell lines. C1, C2, C6, and C21 are independent TbTR DKO clones (c) Northern analysis using a TbTR- specific probe confirmed that TbTR is not expressed in DKO cells. Top, BF cells; Bottom, PF cells (d) Southern analysis of the WT and TbTR DKO candidates. PF, left; BF, right. The position of the Southern probe is shown in (a).

analysis also showed that TbTR was not detectable in the TbTR DKO cells

(Figure.3.5c).

Telomeres grow at a 6–10 bp/PD rate on average in WT BF T. brucei cells when they are continuously propagated (Bernards et al., 1983) (Van der Ploeg et al., 1984b). As a control, I cultured the WT BF cells for more than 130 PDs and observed the same telomere elongation phenotype (Figure.3.6a). In contrast, in

BF TbTR DKO clones, telomeres shorten progressively (Figure.3.6b). In telomere

Southern analyses using a TTAGGG repeat probe; all major telomere fragments can be detected (Figure.3.6b). By calculating the sizes of 5 individual telomere bands in clone C1 (Figure.3.6b, left panel, asterisks), I estimated a telomere- shortening rate of 3–5 bp/PD during the culturing period. To estimate the rate more accurately, I examined the lengths of telomeres that are specifically marked by a subtelomeric VSG11 (aka VSG BR2) gene by Southern blotting (Figure.3.7, left panel). Our T. brucei strain has more than one copy of the VSG11 gene, among which two are telomeric (Dreesen et al., 2005) (Dreesen and Cross,

2006) and are shown as the 20 kb and 7.8 kb bands. I followed the smaller telomeric fragment for more accurate size measurement (Figure.3.7, panel, asterisk) and calculated the telomere-shortening rate to be ~3 bp/PD. In addition,

I examined three VSG8 (aka VSG OD1)-marked bands that are apparently telomeric in the C1 cells (Figure.3.7, left panel, asterisks), and the telomere- shortening rate is estimated to be ~5 bp/PD. Similarly, by calculating the sizes of

6 individual telomere bands in clone C2 (Figure.3.6b, right, asterisks), I estimated a telomere-shortening rate of ~5 bp/PD. Therefore, telomere-shortening rate in 71

TbTR DKO cells is similar to that in TbTERT null cells (3–4 bp/PD) (Dreesen et al., 2005) (Dreesen and Cross, 2006).

Figure 3.6 Telomere length changes in WT and TbTR DKO BF cells Genome DNA was prepared from WT (a) and TbTR DKO (b) BF T. brucei cells at regular time intervals. A (TTAGGG)4 oligo probe was used in (a) and (b).C1 and C2 are independent clones. PD number at each time point are indicated on the top, and telomere bands used for telomere length shortening or elongation rate calculations were marked with asterisks.

Telomere length changes have not been carefully studied in WT or

TbTERT null PF cells. I therefore cultured the PF WT cells continuously for over

150 PDs and estimated the telomere elongation rate to be on average ~5 bp/PD

(Figure.3.8a). In contrast, telomeres shorten at a rate of 5–6 bp/PD in both C6 and C21 TbTR DKO clones (Figure.3.8b). Therefore, in both BF and PF cells 72

deleted of TbTR, telomere maintenance is defective, strongly arguing that TbTR encodes the RNA component of telomerase.

Although deletion of TbTR led to telomere shortening, we did not observe any growth defects after a long-term (more than 350 PDs) continuous cell culture

(Figure.3.9), which is the same as TbTERT-deleted cells (Dreesen and Cross,

2006). To further confirm that the telomere shortening phenotype is caused by

Figure 3.7 Progressive telomere shortening at silent expression sites The VSG8 and VSG11-specific probes reveal both chromosome internal and telomeric DNA fragments. C1 and C2 are independent clones. PD numbers at each time point are indicated on the top, and telomere bands used for telomere length shortening rate calculations were marked with asterisks.

the lack of TbTR transcript, I introduced a complementary ectopic WT allele of

TbTR into the BF TbTR DKO cells. A Tet-inducible WT TbTR allele together with its original 5' and 3' flanking sequences (~500 bp) is targeted to the rDNA array.

Figure 3.8 Telomere length changes in WT and TbTR DKO PF cells Genomic DNAs were prepared at regular time intervals from WT (a) or TbTR DKO (b) PF cells. C6 and C21 are independent clones. Southern analyses were carried out using the (TTAGGG)4 oligo probe.

Upon induction, I observed expression of TbTR at a level higher than that in WT cells (Figure.3.10a). Examination of telomere lengths in two independent complementary clones by Southern analysis showed that telomeres elongate at a steady rate of 8 bp/PD (Figure.3.10b). Similarly, I targeted a WT allele of TbTR with its original 5’ and 3’ flanking sequences into the tubulin array in PF TbTR

Figure 3.9 The telomere shortening phenotype in TbTR DKO cells is complemented by an ectopically expressed TbTR WT allele Northern blots showing expression of the ectopic TbTR allele in BF (a) or PF (c) TbTR DKO cells: Top, blot hybridized with a TbTR-specific DNA probe. Bottom: rRNA species are shown as a loading control. Genomic DNA was isolated at regular intervals and telomere southern was carried out using the (TTAGGG)4 oligo probe on DNA isolated from BF (b) and PF (d) complementation cell lines. B1 and B2 are independent clones of BF cells.

DKO cells. This ectopic allele of TbTR is expected to be expressed constitutively, and northern analysis showed that it is expressed, although at a level lower than

WT (Figure.3.10c). Telomere Southern analysis indicated that telomeres did elongate in these cells, but quantification showed a telomere growth rate of only

~2 bp/PD, suggesting that the weak expression of TbTR led to a slow telomere growth (Figure.3.10d). Therefore, the telomere-shortening phenotype in TbTR

null cells is complemented by a WT allele of TbTR, indicating that the lack of

TbTR function is responsible for the telomere maintenance defect in T. brucei.

3.2.4 TbTR DKO cells lack telomerase activity

I adopted a PCR-based TRAP assay (Kim et al., 1994) to determine if deletion of TbTR abolished telomerase activity. Telomerase activity is expected to add is expected to add 6 nt (TTAGGG) repeats onto TS 3’ end in an RNase-

Figure 3.10 TbTR DKO cells do not have growth defects BF (left) and PF (right) WT and TbTR DKO cells were cultured continuously for over six months. Seven (BF/TbTR DKO C1) or five (BF/TbTR DKO C2) periods of 14 days or four periods of 42 days (PF/TbTR DKO C6) were randomly picked from this long-term culture (with no overlap in the time frame) and the average PD verses time (days) were calculated and plotted. Error bars represent standard deviation.

Figure 3.11 Optimizing TRAP assays to examine telomerase activities in T. brucei cells TRAP assays were carried out according to (Kim et al., 1994) (a) TRAP assay with 0.15 µg T. brucei PF cell extract using TS primer as substrate. The TS oligo, reverse primer, or the cell extract was omitted in lanes 2, 3, or 4, respectively. Lane 6 represents RNase treated sample. Optimizing TRAP assays to examine telomerase activities in T. brucei cells. TRAP assays were carried out according to (Kim et al., 1994). (a) 0.3 μg of T. brucei cell extract (lanes 1 – 6) or 0.4 μg of cell extract from an hTERT over-expression colon cancer cell line (lanes 7 & 8) was used in each reaction. 10 μl (lanes 1–4, 7 & 8) or 20 μl (lanes 5 & 6) of product was loaded in each lane, respectively.

sensitive manner. When T. brucei cell extract was used, a similar RNase- sensitive periodical amplification profile was observed (Figure.3.11a lanes 5 & 6).

No product was detected when the TS or reverse primer was omitted

(Figure.3.11a, lanes 2 & 3, respectively).

When cell extract was omitted from the reaction, we did not observe the ladderof products except one faint band of ~ 40 bp (Figure.3.11a lane 4), indicating that this band did not result from telomerase activity but is most likely a primer-dimer product. Careful examination of the sequences of the TS (5’

AATCCGTCGAGCAGAGTT 3’) and the reverse primer (5’

CCCTTACCCTTACCCTTACCCTAA 3’) also revealed that the two primers can anneal with two T:A base pairs and the primer-dimer product should have a size of 40 bp. WT cell lysate gave a same sized band with stronger intensity because any telomerase extended product will be amplified with the same TS and reverse primer pair in the TRAP assay.

Utilizing this TRAP assay, I compared the telomerase activities from WT and TbTR DKO cells and TbTR DKO cells containing a TbTR complementation allele using equal amount of cell extracts (Figure.3.11b). No telomerase extension products were detected in TbTR DKO cell lines (Figure.3.11b, lane 3), while a weak telomerase activity in the complementation line is observed.

(Figure.3.11b, lane 5, compared to WT in lane 1). To confirm detectable telomerase activity in the complementation line, I repeated the same experiment but used double amount of cell extract from the complementation line and loaded twice as much of its product on gel (Figure.3.12a). In this case, I clearly observed telomerase extension products from the complementation line (Figure.3.12a, lane

6) but not from the DKO cells (Figure.3.12a, lane 4). Even when double amount of cell extract from TbTR DKO cells were used, we still did not observe any telomerase extension products (Figure.3.12b). Therefore, TbTR DKO cells lack 78

Figure 3.12 TRAP assay to compare the telomerase (a) TRAP assay to compare the telomerase activities in WT (lanes 2 & 3), TbTR DKO (lanes 4 & 5), and TbTR DKO cells with a complementation TbTR allele (lanes 6 & 7) with or without RNase A treatment. 0.25 µg (lanes 2 & 3), 0.28 µg (lanes 4 & 5), or 0.43 µg (lanes 6 & 7) of cell extracts were used in each TRAP reaction, and 20 µl (lanes 2–5) or 40 µl (lanes 6 & 7) of final products were loaded in each lane, respectively. (b) 0.44 μg of TbTR DKO cell extract was used in the TRAP assay.

the telomerase activity, and the complementation line restored the telomerase activity. The apparent low telomerase activity in the complementation cells is presumably due to the fact that the TbTR complementation allele was expressed at a lower level than in WT cells (Figure.3.10c).

Figure 3.13 Cloning of telomeres (a) TbTRt mutant allele is expressed upon induction in BF/TbTR DKO background. Northern analysis was carried out using a TbTR-specific probe (top). Ethidium bromide-stained rRNA was shown as a loading control (bottom). (b) Adaptor-ligation mediated PCR amplification of the telomere terminal fragment. Telomeres with the WT (TTAGGG) or mutant (TAAGGG) sequences were shown on the left. Theadaptor with a duplex region bearing unique sequence and a 4 nt 3’ overhang is shown on the right. After annealing of the adaptor overhang and the telomere G-overhang and ligation of the two, the terminal telomere fragment can be amplified with a forward primer (TTAGGG)4 and a backward primer specific to the adaptor.

3.2.5 Mutation in the TbTR template region resulted in altered telomere sequences

To further prove that TbTR is used as the template in telomere DNA synthesis, I mutated the TbTR template 5’-CCCTAACCCTA-3’ into 5’-

CCCTTACCCTA-3’ (termed TbTRt) and expressed the mutant allele in TbTR null cells. An incorporation of the mutant telomere sequence at the chromosome ends would most definitively confirm the identity of TbTR.

Since our TbTR null clones have only been cultured for a short period of time, most telomeres are still relatively long. Therefore, it will take a considerable length of time to observe a significant change in telomere length after the ectopic

TbTRt is introduced into these cells (so as to verify that TbTRt is functional). It has been shown that the active VSG-marked telomere is more prone to large- sized telomere breaks than other telomeres (Bernards et al., 1983) (Van der

Ploeg et al., 1984b) (Pays et al., 1983). Therefore, I subcloned TbTR null cells and picked a clone in which the active VSG2 (aka VSG221)-marked telomere shortened to ~1.5 kb. I transfected the TbTRt-expressing construct into these cells, and northern analysis showed that TbTRt was expressed at a slightly higher than WT level in all three resulting independent clones (Figure.3.13a).

Within a week, I was able to detect VSG2-marked telomere growth in TbTRt transfectants, which were used for subsequent telomere cloning.

To clone the newly elongated telomeres in TbTRt expressing cells, I took advantage of the G-rich 3’ overhang structure at the very end of T. brucei telomeres (Sandhu and Li, 2011). I have found that at least some of the telomere

G-overhangs end in 5’-TAGGGT. Therefore, I was able to ligate an adaptor with a 16 bp duplex DNA of unique sequence and a 4 nt 3' overhang that matches with the terminal 5'-GGGT telomere sequence at chromosome ends

(Figure.3.13b). Subsequently, the terminal few hundred bps of telomere DNA was amplified by one backward primer specific to the unique region of the adaptor and a forward primer containing 4 repeats of TTAGGG. The amplified

DNA fragments were then inserted into pGEM-T-easy (Promega) by TA cloning followed by sequencing analysis.

From three independent TbTRt-expressing clones, several individual telomere-containing plasmids were cloned and sequenced. Telomere sequences of six representative clones are shown in Table 3.1. They all contain a long stretch of TTAGGG WT repeats and a short stretch of TAAGGG mutant repeats at the very end. As a control, telomere fragments cloned from WT cells all contain TTAGGG repeats. Therefore, the mutated TbTRt template sequence was indeed incorporated into telomeres, confirming that TbTR provides the RNA template for T. brucei telomerase.

3.2.6 Biogenesis of TbTR

Most transcription in T. brucei is polycistronic and few genes have been associated with conventional RNA Pol II promoters. Trans-splicing is a major mechanism of transcript maturation in T. brucei that differs greatly from that of mammals and insects. Dr. Chakrabarti’s lab therefore examined if TbTR requires trans-splicing for maturation. RNA sample isolated from immunoprecipitated

TbTERT fraction was reverse-transcribed using both RP and OP primers.

Subsequent PCR analysis using one primer specific to the 5’ end of TbTR and another specific to the 35 nt Spliced Leader (SL) RNA common to all T. brucei trans-spliced mRNAs (Parsons et al., 1984) resulted in products with the expected size (Figure.3.14a). A mock reaction without RT failed to yield any

Figure 3.14 Biogenesis of TbTR (a) Detecting spliced leader (SL) sequence at the 5’ end of immunopurified TbTR. RT-PCR was done with (+) or without (-) RT using a primer specific to TbTR and a primer specific to the SL sequence. Lanes 2 and 4, RT with Random Primer (RP) and Oligo dT Primer (OP), respectively. Lane 3 represents control reaction without RT. (b) Determination of 5’ cap status of TbTR from total RNA and TbTERT-IP RNA. Total or TbTERT antibody- immunopurified RNA fraction from WT cells were immunoprecipitated with TMG antibody, and the IP products were treated with (+) or without (-) RT followed by amplification with TbTR specific primers. The supernatant fraction of TbTERT-IP (sup) and the total RNA extract from TbTR DKO cells were examined the same way. Genomic DNA (gDNA) was amplified with the same TbTR primers as a control. (c) Schematic drawing of the upstream area of the TbTR gene. Potential RNA Pol II specific transcription factor binding sites are shown. Arrow indicates transcription start site (downstream of SL sequence (e) PF T. brucei cells were treated with Indazolo-sulfonamide RNAP III inhibitor for 24 hrs followed by RNA extraction. Northern blotting was done with TbTR- or U2 snRNA-specific oligonucleotide probes (top). Densitometry-quantified TbTR and U2 snRNA levels were normalized against untreated samples, and the relative changes are shown at the bottom (Experiment & analysis performed by Dr. Kausik Chakrabarti’s lab)

product. These data suggest that trans-splicing is involved in the maturation of

TbTR, which is different from all other known telomerase RNA species.

T. brucei SL RNA methylations are required for trans-splicing (McNALLY and Agabian, 1992). However, in contrast to U1, U2, and U4 snRNAs and the U3 snoRNA that contain a 2,2,7-trimethyl guanosine cap (Tschudi and Ullu, 2002),

T. brucei SL RNAs do not contain the characteristic U snRNA cap structure.

Instead of a TMG cap, SL RNAs provide an unusual cap determination of the cap

4 structure, Dr. Chakrabarti decided to examine whether TbTR contains a TMG cap. He immunoprecipitated total T. brucei RNA with a monoclonal antibody specific for TMG cap. The resulting IP product gave rise to the expected band after RT-PCR amplification using TbTR specific primers when WT cells but not

TbTR DKO cells were used (Figure.3.14b). However, when RNA was first immunoprecipitated with TbTERT-specific antibody followed by precipitation with the anti-TMG antibody, no RT-PCR product was detected using the same set of primers (Figure.3.14b, pellet). Interestingly, TMG signal was detected in the supernatant fraction of the TbTERT IP (Figure.3.14b, sup), suggesting that the mature TbTR incorporated into telomerase does not possess a canonical cap structure that can be recognized by the anti-TMG antibody. Therefore, we speculate that TbTR might acquire a cap 4 structure (Lenardo et al., 1985)

(Freistadt et al., 1988) that was not detected by the TMG antibody, which is consistent with our previous finding that the TbTERT-associated TbTR contains the SL sequence (Figure.3.14a).

Because both human telomerase RNA and SL RNAs in trypanosomes are transcribed by RNA polymerase II (RNAP II), we were curious to know which

RNA polymerase transcribes TbTR. Dr. Chakrabarti’s lab identified a putative

‘TATA’ motif, multiple GATA type transcription factor binding sites (Figure.3.14c), and a C/EBP1 type motif upstream of the TbTR transcription start site using

Emboss: Tfscan (http://helixweb.nih.gov/emboss/html/tfscan.html), suggesting that TbTR is transcribed by RNAP II. In the absence of a good T. brucei cell permeable inhibitor for RNAP II, Dr. Chakrabarti turned to a RNAP III inhibitor that has a broad effect on RNAP III transcription in eukaryotic cells (IC50 = 27 and 32 µM for human and S. cerevisiae RNAP III, EMD Biosciences). Dr.

Chakrabarti’s lab cultured PF T. brucei cells in the presence (10 µM and 50 µM) or the absence of this inhibitor for 24 hours followed by total RNA isolation and northern analysis. Since U2 spliceosomal RNA in T. brucei is transcribed by

RNAP III (Fantoni et al., 1994), Dr. Chakrabarti used U2 snRNA as a positive control. Northern analysis revealed 16-23% reduction in the U2 mRNA level when cells were treated with the RNAP III inhibitor, whereas the TbTR level did not change significantly (Figure.3.14d), further suggesting that TbTR is transcribed by RNAP II.

3.2.7 Secondary structure of TbTR

TbTR folding was first analyzed using parameters that consider sequence conservation information from multiple Trypanosoma subspecies. Dr. Chakrabarti recorded nucleotide changes consistent with RNA evolution (covariations) rather

than third position variation common to protein coding regions. The 5’ and 3’

RLM-RACE generated minimal TbTR sequence was used to generate alignments for modeling a consensus secondary structure for Trypanosoma TR molecules. Initially, using Turbofold program (Harmanci et al., 2011), Dr.

Chakrabarti was able to compare the common domain architecture of individually folded Trypanosoma TR molecules to derive a consensus structure. This structure was then verified by RNAalifold (Bernhart et al., 2008), and a potential region to form a pseudoknot was identified by Pknots program (Rivas and Eddy,

1999). Finally, covariations were checked manually within conserved domains.

To verify this model, Dr. Chakrabarti’s lab performed in vivo RNA footprinting. To determine how this RNA folds in vivo, RNA was isolated from T. brucei nuclei that were exposed to DMS and CMCT (Xu and Culver, 2009). Dr.

Chakrabarti found that the majority of the nucleotides in template domain of

TbTR is exposed and is modified by DMS or CMCT (Figure.3.15a), suggesting that the TbTR template is generally accessible to single strand-specific hybridization to telomeric repeats and to incoming nucleotides for telomere synthesis. In contrast, very few nucleobases in TBE, which could form a helical structure, are exposed (Figure.3.15b).

Thus, phylogenetic analysis, supported by in vivo chemical footprinting data, showed a strong correlation in developing a TbTR secondary structure model (Figure.3.15c). In this model, a 5’ proximal stem loop (Helix-I) is present in all Trypanosoma TR analyzed, which is shorter than the yeast telomerase RNA

(TLC1) hairpin that interacts with Ku (Stellwagen et al., 2003). We can also 86

detect a conserved template proximal hairpin (Helix-II), which is likely the TBE.

Our iterative folding process always detected the template domain to be single- stranded in optimal physiological temperature and salt conditions, which is embedded as a highly conserved domain between TBE and the pseudoknot. The possibility of forming a pseudoknot within the TbTR core 5’ domain was highlighted by the presence of a pair of compensatory mutations identified in this structure. A smaller, alternative, minimum free energy pseudoknot structure within 50 nt of the template domain was identified by iterative folding process which is not supported by conservation or covariation. In addition, a 3’ end multiloop structure with a long hairpin (Helix-IV) was identified mostly in the proximal stem resembling the terminal structure of hTR, which is supported by the presence of covariation. Dr. Chakrabarti was not able to identify any typical

Sm binding sequence or verify any hTR like H/ACA motif in TbTR by computational analysis. Our analysis provides a strong but conservative model for native folding of TbTR, which requires further validation with mutational analysis and functional studies. However, the overall similarities among phylogenetic and in vivo structure probing data provide a convincing paradigm that can help to understand telomerase functions in a genetically tractable human pathogen.

Figure 3.15 Chemical probing of TbTR and secondary structure prediction Chemical probing of TbTR and secondary structure prediction. Chemical probing of (a) the template region and (b) TBE of TbTR with DMS and CMCT in vivo. Control (con) represents RNAs not exposed to DMS. Lanes G, U, A and C represent dideoxynucleotide sequencing results. Solid black circles represent full accessibility to chemicals, while open circles represent moderate accessibility. The same symbols are used to mark sites of chemical accessibility along RNA template and helices I-IV on the side in (c). (c) Secondary structure model of TbTR RNA. Between two paired RNA strands (marked in the center), solid lines represent Watson-Crick base pairing, open circles represent Wobble pairs, and long dotted lines represent putative long distance/tertiary interactions. Asterisks mark consensus base pairs on both sides. Compensatory mutations/covariations are shaded with grey circles, and single base changes from phylogenetic sequence analysis are circled. Helical domains are numbered I to IV from 5′ to 3. (Experiment &Analysis performed by Dr. Kausik Chakrabarti lab) 88

3.3 Discussion

Several lines of evidence indicate that the TbTR we identified is indeed the RNA component of T. brucei telomerase. The telomere complex has been shown to play an important role in T. brucei pathogenesis (Yang et al., 2009).

Therefore, altering the TbTR template sequence can have profound effect on the telomere structure, which may have therapeutic implications.

In addition, previous studies proposed a putative template domain for

Trypanosoma telomerase RNA (Cano et al., 1999) (Munoz and Collins, 2004), which has a unique cytosine rich motif distinctive from any other known TR molecules except another pathogenic protozoa Plasmodium falciparum

(Figure.3.1a). Through mutation analysis we confirmed that this motif indeed is part of TbTR template. This motif may play a novel role in substrate recognition

(Munoz and Collins, 2004). Therefore, understanding the functions of TbTR can provide clues to a novel telomere maintenance mechanism in kinetoplastid parasites.

The loss of telomeric repeats after TbTR deletion and restoration of telomere elongation in the presence of an exogenous WT copy indicate that a single-copy gene located on chromosome 11 encodes TbTR. The major TbTR transcript is a 900 nt RNA, as shown in northern analyses. Reverse transcription of TbTERT-associated TbTR (isolated through TbTERT IP) with Oligo dT primer followed by PCR with TbTR gene specific and SL-specific primer pairs gave a product of expected size, suggesting that the mature TbTR contains a poly A tail and a 5’ SL cap. Capping of mRNAs by SL sequence through trans-splicing is a 89

unique RNA maturation process characteristic of Trypanosoma subspecies.

Although different than cis-splicing, trans-splicing in T. brucei still requires canonical spliceosomal U RNA machinery (Tschudi and Ullu, 1990) and Sm component proteins (Palfi et al., 1991) (Mandelboim et al., 2003). Telomerase

RNA maturation in fission yeast involves mechanism akin to cis-splicing (Box et al., 2008), which requires sequential binding of Sm proteins to telomerase RNA

(Tang et al., 2012). Although we were unable to detect any typical Sm binding sequence in the TbTR, we cannot rule out the possibility of TbTR interacting with

Sm proteins since it could be required in trans-splicing for TbTR maturation. It is interesting to note that splicing appears to be a common mechanism for maturation of larger telomerase RNAs, such as those encoded by fission yeast

(Box et al., 2008), Plasmodium sp. (Chakrabarti et al., 2007) (unpublished data), or Trypanosoma, and could be linked to polyadenylation as seen in budding yeast (Chapon et al., 1997). Oligo dT primed RT-PCR analysis was previously used to determine the polyadenylation status of human telomerase RNA (Kim et al., 2001) and to measure the cellular concentration of yeast telomerase RNA,

TLC1 (Mozdy and Cech, 2006). Using a similar approach, we were able to identify polyadenylated TbTR transcript not only in total RNA fraction but also from immunopurified telomerase complex (Figure.3.14a). Therefore, TbTR appears to exist in multiple trans-spliced and/or polyadenylated isoforms, as a part of the active telomerase RNP complex or processing intermediates

(Figure.3.2c). Their functions in telomere synthesis or developmental regulation in insect and vertebrate hosts, if any, remain to be established. 90

Like yeast and human telomerase RNA (Feng et al., 1995), TbTR is likely transcribed by RNAP II. However, the 5’ cap modification of RNAP II transcribed

RNAs in trypanosomes is different from that in other eukaryotes. Usually the SL

RNAs possess a complex cap structure, called ‘cap 4’, which is essential for trans-splicing (McNALLY and Agabian, 1992) (Bangs et al., 1992) (Ullu and

Tschudi, 1991). The RNAP III encoded spliceosomal RNAs, such as U2 and U4, possess a TMG cap, whereas U5 lacks it (Xu et al., 1997). We did detect TMG capped TbTR in the total RNA fraction. However, TbTR does not appear to be

TMG capped when associates with TbTERT, thus, based on the cap requirements for trans-spliced products, there is a possibility that trans-spliced

TbTR contains the cap 4 structure. However, we are uncertain at this stage if and why TbTR carries a TMG during primary processing, although mRNAs in C. elegans can keep their TMG caps even after trans-splicing (Liou and Blumenthal,

1990).

The telomere elongation rate in WT cells is very similar in BF and PF cells, which is consistent with our observation that both TbTR and TbTERT are expressed at very similar levels. In TbTR DKO cells expressing TbTRt mutant, most of the mutant telomeres only had a short stretch of mutant sequence incorporated, suggesting low processivity of T. brucei telomerase. This is consistent with our telomerase primer extension result and previous observations

(Cano et al., 1999).

TR’s function of serving as a template goes beyond simple substrate recognition and hybridization. Trypanosome telomerase is semi-processive in 91

nature (Cano et al., 1999; Munoz and Collins, 2004), although it makes telomeric repeats identical to those synthesized by human telomerase, a highly processive enzyme. Such difference in enzyme processivity can be due to the differences in core template sequences or template adjacent region between host and pathogen. For example, there is a ‘UAA’ duplication in human TR template that could render enhanced processivity (Chen and Greider, 2003). In contrast, there is a ‘CCC’ duplication in TbTR template, which could provide high affinity substrate hybridization (Munoz and Collins, 2004). However, this could also affect the telomere elongation rate due to stronger binding of template to telomeric substrate.

In TbTR complementation cell lines, the telomere elongation rate appears to depend on the level of TbTR expression. This observation strikes a possibility of TbTR being the rate-limiting factor for telomere elongation, although more careful analyses are necessary to reveal the relationship between TbTR/TbTERT expression level, telomerase activity, and in vivo telomere elongation rates.

Interestingly, hTERT is usually only expressed in telomerase positive cells while hTR expression appears to be ubiquitous (Feng et al., 1995), suggesting that hTERT might be the rate-limiting factor in human cells for telomere elongation.

However, expression of ectopic hTERT and hTR both appear to be able to increase telomerase activity and lead to telomere elongation, and their effects are additive (Cristofari and Lingner, 2006). Therefore it would be necessary to further investigate in details the telomerase regulation in T. brucei to reveal any

similar or different underlying mechanisms, which should contribute to anti- parasite drug development in the future.

Given major differences in molecular properties and activity of T. brucei and human TR, establishment of a secondary structure model for TbTR is important for future genetic and functional analysis. Our in vivo structure probing data strongly supports the phylogenetics-based and computer-developed model.

First, iterative folding of predicted TR sequences from five different Trypanosoma subspecies identified the template region to be single-stranded in an unbiased, independent process, which perfectly correlated with in vivo chemical probing data. Thus, the majority of the TbTR template residues are unlikely to be occupied in any hydrogen bonding or tertiary or RNA–protein interactions.

Second, the differences in chemical accessibility between the template domain and adjacent 5’ helix-II in in vivo assays suggested a helix forming potential for

TBE.

Finally and most importantly, conservation among functional domain sequences or structures provided additional confidence for the structure prediction. For example, the NMR structure of the hTR showed extensive tertiary interactions between the loop and stem nucleotides, rendering an essential triple helix in the pseudoknot structure (Theimer et al., 2005) (Shefer et al., 2007). A similar structure could possibly form in TbTR pseudoknot (Figure.3.15c).

Additionally, the minimal CR4/CR5 domain sequence of hTR required for reconstitution of active telomerase in vitro is >70% conserved with TbTR helix IV sequence (Figure.3.15c), including the P6.1 loop which is shown to be essential 93

for TERT binding and telomerase activity (Mitchell and Collins, 2000). Thus, these findings provide a testable model for further analysis.

In TbTERT null cells, when silent ES-marked telomeres are extremely short (~ 40 bp), it was observed that the short telomeres can be stably maintained for several tens of generations without significant changes, indicating the existence of an unusual telomerase-independent telomere maintenance mechanism in T. brucei (Dreesen and Cross, 2006). It will be interesting to examine whether the same mechanism can be activated in TbTR DKO cells that have lost most telomere DNA.

Table 3.1 Telomere sequences in several representative telomere clones Cloned telomeres from BF/TbTR DKO + TbTRt cells

Clone 3 5’ (CCCTTA)6(CCCTAA)73 Clone 7 5’ (CCCTTA)3(CCCTAA)95 Clone 15 5’ (CCCTTA)4(CCCTAA)84 Clone 18 5’ (CCCTTA)3(CCCTAA)105 Clone 19 5’ (CCCTTA)5(CCCTAA)49 Clone 20 5’ (CCCTTA)5(CCCTAA)121 Cloned telomeres from BF/WT cells

Clone 9 5’ (CCCTAA)92

Telomeres and subtelomeres are recombination hotspots for parasitic protozoa such as T. brucei that undergo antigenic variation. In TbTERT null cells, when the active VSG-marked telomeres are extremely short, an elevated VSG switching frequency was observed (Hovel-Miner et al., 2012). It has been hypothesized that extremely short telomeres allow more frequent chromosome end breaks that damage the active VSG gene, which forces the cell to undergo antigenic variation (Cohen et al., 2007). Whether a similar effect can be seen in 94

TbTR DKO cells remains to be investigated. Since genes encoding virulence factors such as VSGs are expressed from subtelomeric regions, our study lays a foundation for the thorough understanding of TbTR’s role in telomere maintenance and virulence gene regulation in a genetically tractable model pathogen.

4CHAPTER IV

G-overhang Structure of Trypanosoma brucei

4.1 Introduction

Telomeres are specialized nucleoprotein complexes present at the ends of chromosomes. In most species, telomeric DNA consists of tandem arrays of TG- rich sequence bound by a protein complex called telosome. Telomeric repeats can be prefect such as the TTAGGG repeat in vertebrate cells and a protozoan parasite T. brucei, or they can be imperfect repeats such as (TG1-3)n in budding yeast (Moyzis et al., 1988) (Shampay et al., 1984). Telomeres typically consist of long tracts of duplex telomeric DNA and terminate in a 3’ single-strand G-rich overhang (Henderson and Blackburn, 1989). This 3’ G-rich overhang structure has been observed in many species and is critical for telomere maintenance and function. G-overhang serves as a primer for telomerase-dependent synthesis of telomere DNA. It can also invade the duplex telomere region to form a higher- order structure called T-loop, which has been proposed to protect telomere terminus against illicit DNA processes including degradation, repair, and recombination (Greider and Blackburn, 1987) (Griffith et al., 1999). Similar to the

duplex region of telomeres, the length and structure of the telomere G-overhang is tightly regulated and maintained. In S. cerevisiae, the G-overhang is 12-14 nucleotide long with a transit increase in length in S phase, during which 30-100 nucleotide long G-overhangs can be detected (Wellinger et al., 1993). Similarly, mammalian cells have 12-300 nucleotide long G-overhangs at their telomere ends (Makarov et al., 1997) (Chai et al., 2006) (Dai et al., 2010). The mechanism for generating overhang has been a subject of intense research for many years.

A common theme that has emerged from these studies is that the G-overhang is the net result of the semiconservative replication of linear DNA molecules, telomerase activity on pre-existing 3’ overhangs, and tightly controlled nucleolytic processing of linear DNA ends (Wellinger et al., 1996) (Makarov et al., 1997).

4.1.1 G-overhang function

The G-overhang structure is important for telomere maintenance and function. First, it serves as a primer for telomere DNA synthesis by telomerase.

An in vitro assay confirmed that DNA primers containing G-rich telomeric repeats as well as double stranded DNA containing telomeric 3' overhangs were extended by telomerase. In contrast, blunt ended DNA and double stranded DNA containing 5’ overhangs were not used as substrates by telomerase (Greider and

Blackburn, 1987). These results underscored the importance of G-overhang in telomerase dependent telomere maintenance. Second, G-overhangs are

Figure 4.1 End replication problem A linear chromosome replication: Origin of replication (circle) and the telomere sequence is shown. The replication bubble is extended as the replication fork travels along the DNA strand as indicated by the bidirectional arrows. RNA primers are indicated in red; newly synthesized DNA strands are shown. Upon extension of the replication bubbles to the chromosome termini, the replication machinery can extend to the end of the chromosome generating a blunt‐end at the leading strand of the chromosome, which is processed to generate a G- overhang. At the lagging strand after degradation of the last RNA primer a short G‐overhang remains. Multiple round of DNA replication without telomerase action leads to telomere shortening.

essential for formation of the protective t-loop structure (Murti and Prescott,

1999) (Griffith et al., 1999) (Muñoz‐Jordán et al., 2001) (Cesare et al., 2003).

Recently it was shown that TRF2 suppresses ATM signaling and NHEJ at telomeres by facilitating the formation of T-loop (Doksani et al., 2013). Thus the

T-loop provides the critical means by which telomeres prevent chromosome ends from signaling illicit DNA damage responses. Moreover, removal of proteins that bind to G-overhang leads to massive loss of telomere sequences (Booth et al.,

2001) (Grandin et al., 2001a) (Grandin et al., 1997) (Baumann and Cech, 2001)

(Hockemeyer et al., 2006). Exposure of the G-overhang has been also shown to be associated with senescence of human primary cells (Stewart et al., 2003). In summary, the telomere G-overhang structure is a conserved feature of eukaryotic cells and is required for telomere functions.

4.1.2 Generation of G-overhang

The conventional semiconservative replication is unable to fully replicate linear DNA molecules. The inherent property of DNA polymerase dictates that it requires a primer with available 3’-OH for DNA synthesize. After semiconservative DNA replication, the RNA primer used to initiate the replication at the extreme 5’ end is removed, resulting in loss of the terminal sequences.

This inability to copy the very end of linear DNA molecules by the replication machinery is termed ‘end replication problem’ (Watson, 1972) (Olovnikov, 1973)

(Figure.4.1). When telomeric DNA is replicated by DNA polymerase, the

TTAGGG (G-rich) strand serves as the template for discontinuous or lagging-

strand synthesis, while the CCCTAA (C-rich) strand serves as the template for continuous or leading-strand synthesis. After replication two different types of telomeric ends are generated. Lagging strand has 3’ single stranded DNA due to the removal of the RNA primer from the last Okazaki fragment, making G- overhang a natural consequence of the end replication problem. On the other hand, leading strand is blunt ended after replication. Intriguingly, G-overhangs are also observed at the leading strand, suggesting that resection of leading or C rich strand in 5’ to 3’ direction is responsible for generation of these overhangs

(Makarov et al., 1997) (LeBel and Wellinger, 2005) (Figure.4.2).

In S. cerevisiae, G-overhang maturation is dependent on cell cycle and overhangs are processed differently on leading and lagging strand. Removal of the last primer from the lagging strand results in an overhang on the lagging strand whereas a 5’ end resection and strand filling lead to generation of the mature overhang on the leading strand (Soudet et al., 2014) (Figure 3.3). In accordance with this, MRX (Mre11, Rad50, and Xrs2) and Sae2 nucleases preferentially localize to the leading strand and are required for 5’ end resection of the C-rich strand (Bonetti et al., 2009). Activity of MRX complex at the telomere depends on its phosphorylation by Tel1 (ATM homolog) in S phase

(Martina et al., 2012). Similarly, Cyclin dependent kinase Cdk1 phosphorylates

Sae2 in S phase, which leads to the recruitment of Sae2 at the telomere end

(Bonetti et al., 2009). Lack of Mre11 or Sae2 dramatically impairs 5’ end resection, but the C-rich strand is still partially resected in these deletion mutants, suggesting that 5’ end resection does not solely depend on the MRX complex 100

Figure 4.2 G overhang generation The G-overhang is generated at both leading strand and lagging strand. Newly replicated stands are denoted in blue and cyan. Removal of the last RNA primer (shown in red) results in a G overhang on the 3’ end resulting from lagging strand synthesis. Replication results in blunt end at leading strand. A 5’ to 3’ nuclease MRX/Sae2 in yeast and Apollo/Exo1 in mammals (shown as packman) digestion of the C-strand creates a 3’ overhang on the G-strand.

(Larrivée et al., 2004). In agreement with this, exonucleases including Exo1,

Dna2, and Sgs1 have been shown to act on telomere ends to generate 3’ overhang (Bonetti et al., 2009). Nonetheless, studies suggest that MRX and

Sae2 play a major role in telomere resection while Dna2, Sgs1, and Exo1 may provide a backup mechanism for generation of 3’ overhang when MRX/Sae2 activity is compromised (Figure4.3).

101

Figure 4.3 Processing of G overhang in S. cerevisiae Generation of G-overhang in S. cerevisiae: Single-stranded overhangs originate at the lagging-strand telomeres because of the last RNA primer removal from last Okazaki fragment at the end of a linear DNA molecule. CST/Ku bind to these overhang and prevent further processing. In contrast, leading-strand telomere is blunt-ended after replication and the C-strand must be processed by nucleases to generate 3’-ended single-strand overhangs. Telomerase can extend the 3’ single-strand overhangs, and the C-strand can be filled in by the polα–primase complex.

In mammals, different nucleases such as Apollo and Exo1 process telomere ends to generate G-overhangs (Wu et al., 2012a). TRF2, a Shelterin component, recruits Apollo to the leading strand, where Apollo initiates resection

102

of 5’ end using its nuclease activity (Wu et al., 2012). Unlike Apollo, Exo1 acts on both leading and lagging strand of the newly synthesized telomere (Wu et al.,

2012) (Figure.4.4). In T. brucei, evidence for the presence of the telomere G- overhang on both ends of chromosomes came from the observation that T-loop is present on both ends of chromosomes (Muñoz‐Jordán et al., 2001). So far no nuclease has been identified in T. brucei to be responsible for G-overhang generation at the leading strand 5’ end.

Figure 4.4 Generation of the telomeric overhang in mammalian cells Last primer removal results in overhang at lagging strand which in turn is bound by POT1b.POT1b inhibits Apollo at lagging strand ends but have limited affect on EXO1 activity. EXO1 activity results in long G-overhang at both the ends. At leading strand, TRF2, recruits Apollo that initiates 5’ strand resection- generating G-overhang. POT1b gets loaded on the terminal overhang and inhibits resection by Apollo. POT1b then recruits CST, which facilitates fill-in synthesis of the C-rich strand at both ends, this results in mature overhang generation (Wu et al., 2012).

4.1.3 G-overhang binding Proteins

Telomeric DNA is bound by specialized proteins, those that can bind to double stranded DNA and those that can bind to single stranded DNA. These 103

proteins function in telomere length maintenance as well as chromosome end capping. In S. cerevisiae, the CST complex (telomeric RPA) binds to G-overhang at least in S phase. CST is a heterotrimeric complex composed of Cdc13, Stn1, and Ten1 (Gao et al., 2007) (Tsukamoto et al., 2001) (Grandin et al., 1997). CST plays an important role in regulating G-overhang length, as loss of CST results in

C-strand degradation and longer G-overhangs as compared to wild type cells

(Maringele and Lydall, 2002) (Ngo and Lydall, 2010). Studies have revealed two main functions of the CST complex. Firstly, CST inhibits nuclease action at the telomere end. Secondly, it recruits DNA Polymerase alpha (Polα) to the telomere end to fill-in the C-strand sequence (Qi and Zakian, 2000) (Grossi et al., 2004)

(Casteel et al., 2009) (Sun et al., 2010). In S. pombe, and in higher eukaryotes, single strand DNA binding protein Pot1 binds to G-overhang (Baumann and

Cech, 2001) (Churikov et al., 2006). In S. pombe, loss of Pot1 results in an increase in G-overhang length by two to three folds (Pitt and Cooper, 2010).

In humans, Pot1 knockdown results in 20-30% reduction in G-overhang signals (Hockemeyer et al., 2005). Unlike humans, mouse has two Pot1 proteins,

Pot1a and Pot1b, both of which bind to G-overhang (Hockemeyer et al., 2006).

Pot1a suppresses the ATR signaling and Pot1b regulates the G-overhang length

(Hockemeyer et al., 2006). Loss of Pot1b in mouse results in an ~two-fold increase in the G-overhang signal at the leading strand and an ~three-fold increase at the lagging strand, indicating that Pot1b plays a role at both newly synthesized telomere ends (Hockemeyer et al., 2008). Arabidopsis thaliana encodes two Pot1 paralogs, but none of them exhibit DNA binding activity in vitro 104

and are proposed to have evolved to mediate telomerase regulation instead of chromosome end protection (Shakirov et al., 2005) (Surovtseva et al., 2007). In contrast, Physcomitrella patens (moss) harbors only one Pot1 protein, which binds to G-overhang, and Pot1 mutant shows an ~eight-fold increase in G- overhang signals (Shakirov et al., 2010). In summary, functions of single strand telomere binding proteins (Cdc13 and POT1) in G-overhang maintenance and telomere capping are conserved in evolution. In vitro studies have suggested that

T. brucei cell lysate has a complex that specifically binds to the telomere single stranded G-rich sequence (Cano et al., 2002). But identity of this activity remains elusive so far.

4.1.4 G-overhang processing

Several proteins regulate the G-overhang processing. One such protein is

Ku, which is a key player in the non-homologous end-joining (NHEJ) process. Ku is a heterodimer composed of Ku70 and Ku80 (Mimori and Hardin, 1986)

(Feldmann and Winnacker, 1993) (Taccioli et al., 1994) (Boulton and Jackson,

1996). In S. cerevisiae, deletion of Ku leads to an increase in G- overhang length in G1 phase of the cell cycle and this increase depends on the activity of EXO1

(Polotnianka et al., 1998) (Bonetti et al., 2010) (Vodenicharov et al., 2010)

(Maringele and Lydall, 2002). Thus Ku is proposed to block the 5’ to 3’ resection of telomeric ends by EXO1 in G1 phase (Bonetti et al., 2014). In mammals, Ku, along with TRF2, protects telomere ends from homologous recombination.

Conditional knockout of Ku86 in human leads to massive and rapid loss of

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telomeric DNA in form of extrachromosomal t-circles (Wang et al., 2009). In mouse cells Ku act parallel with TRF2 to block recombination at telomere ends

(Celli et al., 2006). However, its role, if any, in blocking the resection of telomere ends is not known. In plants, Ku is proposed to protect the C-rich strand from being resected, as deletion of Ku in A. thaliana leads to a 3-fold increase in G- overhang length (Kazda et al., 2012). In T. brucei, deletion of Ku leads to telomere shortening (Conway et al., 2002a) (Janzen et al., 2004). However, it is still not clear if Ku protects resection of the C-rich strand and/or is required for recruitment of telomerase to the telomere. Besides Ku, the CST complex plays an important role in regulating G-overhang in budding yeast as well as in mammalian cells.

In S. cerevisiae, CST has been proposed to work differently at the lagging and leading strand. Cdc13, a CST complex component, regulates timely replication of the lagging strand but not the leading strand (Soudet et al., 2014).

Furthermore, it has been suggested that Cdc13 is also involved in positioning

RNA primer on the most distal Okazaki fragment, thereby regulating the G- overhang length at the lagging strand (Soudet et al., 2014). On the leading strand, Cdc13 is involved in regulating the nuclease activity and block 5’ resection (Garvik et al., 1995) (Lydall and Weinert, 1995) (Vodenicharov and

Wellinger, 2006). CST binds to both leading and lagging telomeres and is involved in C-rich strand fill in by interacting with the Polα/ Primase complex (Qi and Zakian, 2000; Grossi et al., 2004).

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In S. pombe, only STN1 and TEN1 have been identified so far, but the function of this complex in maintaining G-overhang seems conserved (Martín et al., 2007). In mammalian cells, The CST complex is composed of CTC1, STN1, and TEN1 (Miyake et al., 2009) Mammalian CST interacts with telomeres and is implicated in telomere replication and the C-rich strand fill-in by recruitment of the

Polα/ Primase complex. In mouse, CST interacts with Pot1b, and this interaction is required for the recruitment of CST complex to the telomere (Wu et al., 2012).

Cells lacking Pot1b or expressing mutant Pot1b that lacks the CST binding domain (Pot1bΔCST) had significantly reduced amount of telomere-associated

CST, which in turn resulted in long G-overhangs (Wu et al., 2012). This defect was mainly due to the loss of C-rich strand fill-in (Wu et al., 2012). Similarly in human cells, CST is localized to G-overhang, and loss of CST results in longer

G-overhangs arguably due to loss of C-rich strand fill in (Wang et al., 2012)

(Huang et al., 2012). Surprisingly in A. thaliana CST functions as a major telomere-capping complex in addition to regulating C-rich strand fill-in (Song et al., 2008) (Surovtseva et al., 2009). In summary, the CST complex is conserved during the evolution (Sun et al., 2009) (Price et al., 2010) (Chen and Lingner,

2013). It acts as a specialized complex to recruit DNA-polα to the chromosome end to maintain proper G-overhang length and prevent excessive loss of C-rich strand. In T. brucei, the CST complex is yet to be identified. However it is also possible that CST-independent mechanism is used to maintain G-overhang in T. brucei.

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4.1.5 Molecular steps of G-overhang generation

Several studies have shown that generation of G-overhang is tightly linked with DNA replication and is regulated by cell cycle. In S. cerevisiae, G-overhang is extended in S phase and is shortened in G2 and G1 phases (Wellinger et al.,

1993) (Larrivée et al., 2004) (Soudet et al., 2014). During DNA replication the lagging strand G-overhang is generated due to the end replication problem

(Soudet et al., 2014). In the absence of telomerase no further processing of lagging strand G-overhang takes place, therefore the removal of last RNA primer from the lagging strand results in a mature G-overhang (Soudet et al., 2014). In telomerase positive cells Cdc13 dependent recruitment of telomerase to the G- overhang in late S phase results in extension of G-overhang (Li et al., 2009b).

Whereas in late G2 phase, Cdc13 forms a complex with Stn1 and Ten1, which in turn recruits the Polα/Primase complex to fill-in the C-rich strand, resulting in shortening of G-overhang (Hang et al., 2011). On the leading strand, G-overhang formation is initiated by nuclease action of MRX and Sae2 (Faure et al., 2010).

During late S phase, MRX and Sae2 are recruited to the leading strand and resect C-rich strand (Faure et al., 2010) (Soudet et al., 2014). This resection results in G-overhang formation, which in turn is bound by Cdc13. Binding of

Cdc13 inhibits nuclease degradation of the C-rich strand (Xu et al., 2009)

(Anbalagan et al., 2011). Following binding of Cdc13 to the newly synthesized overhangs, similar steps at both leading and lagging strand lead to maturation of

G-overhang (Soudet et al., 2014). Cdc13 forms two separate complexes with

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different functions at G-overhang, and post-translational modifications of Cdc13 regulated by cell cycle control the balance between these complexes.

The interaction between Cdc13 and telomerase-associated Est1 depends upon phosphorylation of Cdc13 at T308 by Cdk1 (Li et al., 2009b). This modification occurs in late S phase, and it favors the interaction between Cdc13 and Est1, thus promoting telomerase recruitment to the telomeres and resulting in G-overhang elongation (Li et al., 2009b). When telomerase action is accomplished, Cdc13 limits continuous telomerase activity by interacting with

Stn1 and Ten1 and promoting the synthesis of the C-rich strand by DNA Polα.

Recently SUMOylation of Cdc13 at K909 has been shown to promote the interaction between Cdc13 and Stn1 to form the CST complex thereby inhibiting interaction of Cdc13 with telomerase (Hang et al., 2011). SUMOylation of Cdc13 starts at early S phase and peaks at late S phase. In contrast, phosphorylation of

Cdc13 and telomerase recruitment peak at late S to G2 phases, suggesting that

SUMOylation of Cdc13 may restrict telomerase activity at telomeres after DNA replication is completed. As Cdc13 also inhibits telomerase in late G2 phase, it is proposed that Cdc13 may exert a dual-phase inhibition of telomerase (Wu et al.,

2012b) (Churikov et al., 2013). However the switch that decreases the interaction of Cdc13 with telomerase while increases its interaction with Stn1 and Ten1 in late G2 phase is still not clear.

In higher eukaryotes, the steps of mature G-overhang formation resemble those in S. cerevisiae. In human cells, the end replication problem leads to 70-

100 nucleotide overhangs at the lagging strand (Chow et al., 2012). In contrast to 109

S. cerevisiae, human replication machinery fails to initiate the replication from very end of the chromosome and thus results in long overhangs (Chow et al.,

2012). In the absence of telomerase, no further processing take place at lagging strand telomeres (Chow et al., 2012). However, in mouse cells, Exo1 has been implicated in C-rich strand resection at the lagging strand, generating long G- overhangs at the lagging strand. Positioning of last RNA primer during replication of telomere is not studied in mouse (Wu et al., 2012). Replication of leading strand is proposed to result in blunt end or probably an end with a 5’ overhang o

1-2 nt long due to dissociation of the replication apparatus from the extreme chromosome termini prior to incorporating the final 1 or 2 nucleotide (Chow et al.,

2012). This is followed by resection of C-rich strand (Chow et al., 2012). The leading strand G-overhang of nearly the mature size can be detected in late S/G2 phase (Chow et al., 2012). In human cells the C-rich strand terminal nucleotides are tightly regulated and end in 5’ CTAACC 3’ (Sfeir et al., 2005) (Hockemeyer et al., 2005)(Figure4.5). The precise molecular mechanism required to generate this specific sequence is not known. However, positioning of the last RNA primer during replication and the C-rich strand fill-in are proposed to contribute to this sequence specificity (Dai et al., 2010) (Chow et al., 2012).

Given the fact that G-overhang of nearly the mature size can be detected at the lagging strand before the removal of the RNA primer argues that no further processing of lagging strand takes place (Chow et al., 2012). Thus removal of the last primer leads to sequence specificity of the C-rich strand. In contrast to the lagging strand, sequence of terminal nucleotides of the leading strand telomeres 110

remains random until late S to G2 phase (Chow et al., 2012). In mouse cells after the passage of replication fork, TRF2 recruits Apollo to the leading end (Wu et al., 2012). Apollo initiates the 5’ end resection, which is further extended by

EXO1 (Wu et al., 2012). Subsequently, POT1 binds to single strand G-overhang and inhibits the action of nucleases on both the leading and lagging strand (Wu et al., 2012). Furthermore, POT1 and Tpp1 (a POT1 binding protein) recruit the

CST complex for C-rich strand fill-in (Wu et al., 2012). Based on these data, it has been suggested that POT1 may play a major role in regulating C-rich strand terminal nucleotide sequence specificity. Indeed, depletion of POT1 results in loss of terminal nucleotide sequence specificity.

In telomerase positive cells, POT1-Tpp1 complex can also recruit telomerase to telomeres (Wang et al., 2007) (Abreu et al., 2010) (Zaug et al.,

2010) (Nandakumar et al., 2012). Upon recruitment to the telomere, telomerase activity leads to G-overhang extension. Therefore, telomerase may play a role in specifying G-rich strand terminal nucleotide. In agreement with this, in telomerase positive cells G-rich strand preferentially terminates with the sequence GGTTAG as compared to telomerase null cells that have a random terminal sequence (Sfeir et al., 2005) (Figure.4.5). However the exact mechanism by which the terminal sequence of C-rich or G-rich strand is specified is not clear.

In ciliate Tetrahymena, the sequence specificity of G-overhang end is tightly regulated and is independent of the telomerase activity (Jacob et al.,

2001). The C-rich strands end with the sequence 3’ AACCC as well as with 3’ 111

AACC while most G-rich strands terminate with the sequence 5’ TGGGGT

(Jacob et al., 2001). In telomerase mutants a slight shift was observed to the 5’

GGGGTT permutation, but the majority of G-overhangs still ended in 5’ TGGGGT

(Jacob et al., 2003). This indicates that although telomerase may have a slight influence on G-overhang processing, it is not a determining factor in specifying the terminal sequence (Jacob et al., 2003) (Figure.4.5).

In S. pombe, Tel1 (ATM) and Rad3 (ATR) dependent phosphorylation of

Ccq1 (no homolog has been identified so far in S. cerevisiae and mammals) results in recruitment of telomerase to telomeres (Moser et al., 2011) (Yamazaki et al., 2012). SUMOylation of TPZ1 (the Tpp1 homolog) on K242 prevents telomerase accumulation at telomeres by promoting recruitment of Stn1-Ten1 to telomeres (Miyagawa et al., 2014) (Garg et al., 2014). These findings highlight the evolutionary conserved mechanism by which phosphorylation and

SUMOylation provide a switch that regulates G-overhang extension and C-rich strand fill-in (Garg et al., 2014). In agreement with this notion it was recently shown that human Tpp1 is phosphorylated at multiple sites during late S/G2 phases and this modification of Tpp1 is associated with higher telomerase activity (Zhang et al., 2013). Phosphorylation of Tpp1 at S111 promotes telomerase recruitment whereas disruption of this phosphorylation results in decreased telomerase activity at telomeres (Zhang et al., 2013). In S. cerevisiae, the telomerase inhibition is proposed to be due to decreased interaction of Cdc13 with the telomerase complex and increased interaction with Stn1 and Ten1 in late

S/G2 phases (Churikov et al., 2013). By contrast, in humans the CST complex is 112

thought to directly terminate telomerase activity at telomeres. CST is proposed to compete with POT1–Tpp1 for telomere single stranded DNA (Chen et al., 2012)

(Chen and Lingner, 2013). Binding of CST increases during late S/G2 phases

(Chen et al., 2012) (Chen and Lingner, 2013). This coincides with telomerase inactivation and Polα/Primase recruitment for C-rich strand fill-in (Chen et al.,

2012) (Chen and Lingner, 2013). In agreement with this, depletion of CST leads to excessive telomerase activity and thus promotes telomere elongation (Chen et al., 2012).

In conclusion, at least four molecular events contribute to G-overhang sequence specificity. First, removal of the last RNA primer in DNA replication creates a 3’ overhang at the lagging-strand telomere, which in turn is bound by single strand DNA binding proteins (Cdc13 in yeast and POT1 in many other eukaryotes). Second, the 5' end of the C-rich strand of the leading-strand daughter telomere is resected by nucleases/helicases such as the MRX complex in yeast and Apollo and EXO1 in mammalian cells, generating a G-overhang that is essential for binding of single strand DNA binding proteins and subsequent recruitment of telomerase. After being recruited to the telomere, telomerase synthesizes the G-rich strand telomere DNA at both telomere ends, further extending G-overhang and possibly influencing the terminal nucleotide sequence. Finally, the CST complex mediates the C-rich strand fill-in and more

C-rich strand repeats are synthesized in the late S/G2 phase, shortening G- overhang length and possibly specifying the terminal nucleotide on the C-rich strand. Surprisingly, the C-rich strand still ends with 3’ CCAATC upon Stn1 113

Figure 4.5 Comparison of G-overhang end structures G-overhang size in human varies from 35-600nt while ciliates (Tetrahymena and Euplotes) have short and precise overhang size. Terminal nucleotides for both G and C strand are very precise for both ciliates while in human only C- strand is specified. G strand terminal in humans is less precise (Fan and Price, 1997) (Jacob et al., 2001) (Sfeir et al., 2005).

depletion in human cells, suggesting that some unknown activity may be involved in the regulation of the C-rich strand processing Deciphering the precise mechanism that governs the C-rich strand terminal sequence specificity will help to better understand how G-overhang length is regulated.

4.1.6 Exception to the rule

Single-stranded telomere G-overhang has been proposed to be a universal feature of eukaryotic chromosomes and is central to the telomere end protection function (Oganesian and Karlseder, 2009). However, study of nematode telomeres challenged this hypothesis by demonstrating the presence of 5’ C-rich overhangs (Raices et al., 2008). In C. elegans, telomeres carry 3’ G- rich overhang as well as 5’ C-rich overhang. Furthermore, it was demonstrated that specific single stranded DNA binding proteins bind to these overhangs

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(CeOB1 binds to single-stranded G-rich overhang, whereas CeOB2 has a preference for the C-rich overhang) and provide telomere protection function

(Raices et al., 2008). However, it is unclear whether these C-rich overhangs are generated by 3’ to 5’ resection of the G-rich strand, by incomplete replication, or by some unknown mechanism.

Recently the presence of a 5’-C-rich overhang in human ALT cells as well as mouse cells that lack telomerase was demonstrated (Oganesian and

Karlseder, 2011). However, how these C-rich overhangs are generated and maintained is not clear. So far no factor, if any, has been identified that can bind to the C-rich strand in mammalian cells. Telomere homologous recombination

(HR) has been proposed to be involved in C-rich overhang maintenance, as C- rich overhang levels are greatly perturbed in response to depletion of HR proteins (RAD51, RAD52, and XRCC3) (Oganesian and Karlseder, 2011).

Finally, blunt ended telomeres have been observed in plants. These blunt telomere ends are proposed to be the result of leading strand replication and are protected by Ku (Kazda et al., 2012).

4.1.7 Methods to study the telomere G-overhang

Several assays have been employed to monitor the telomere G-overhang status. The earliest method used to examine G-overhang was the use of osmium tetroxide, which selectively reacts with thymidine nucleotide present in the single stranded DNA. The osmium tetroxide-modified thymidine is cleaved by piperidine treatment and the cleaved product is resolved on a 12% denaturing

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polyacrylamide gel (Henderson and Blackburn, 1989). Although this method is very sensitive to detect the presence of telomere G-overhangs, it failed to provide precise overhang length.

Another method used to examine the G-overhang is Primer-Extension

Nick Translation Reaction (PENT) (Makarov et al., 1997) (Riha et al., 2000). In this method, a C-rich primer is hybridized to G-overhang and primer extension is performed using DNA pol I. DNA pol I fills the gap between the primer and 5’ end of the G-overhang. As DNA pol I possesses 5’ to 3’ exonuclease activity, it will propagate into double stranded telomeric DNA. Primer extension is performed without dGTP so that extension will stop once non-telomeric sequence is reached, producing a nick. Extension products are analyzed by electrophoresis in alkalic gel followed by hybridizion with a G-rich telomeric probe. This method allows the newly synthesized telomere strands that contain contain sequence complementary to G-overhangs to be separated from chromosomal bulk. PENT has been used to show that telomeres of human cells and A. thaliana contain G- overhangs (Makarov et al., 1997) (Riha et al., 2000). One drawback of this assay is that it could not be used to detect G-overhangs that are less than 30 nt long.

One of the most common methods used to detect G-overhangs is native in-gel hybridization (McElligott and Wellinger, 1997). This method employs a radiolabeled C-rich oligonucleotide probe to detect G-overhang. As this assay is performed in non-denaturing conditions, C-rich oligonucleotide can only anneal to the G-rich overhang region. The intensity of hybridization signal in this assay provides a good estimation of the G-overhang length. 116

To precisely measure the length of G-overhang, following four methods have been developed: telomere oligonucleotide ligation assay (T-OLA) (Chai et al., 2005), telomere oligonucleotide primer extension (TOPE), ligation-mediated primer extension (LMPE) (Jacob et al., 2001), and digestion with duplex-specific nuclease (DSN) (Zhao et al., 2008).

In T-OLA, radiolabeled oligonucleotides complementary to the G-overhang are annealed to the single-stranded overhang. Annealed oligonucleotides in exact juxtaposition are then ligated by DNA ligase into long arrays. The ligation products are then separated on a denaturing gel, which will separate the oligonucleotide array from the chromosome DNA. The G-overhang length is therefore proportional to the size of ligated oligonucleotide array detected on the gel. TOPE relies on extension of a radiolabeled oligonucleotide primer that is annealed to the G-overhang. Primer extension is performed using T4 DNA polymerase, which does not have and strand displacement activity. Therefore, it will extend the primer until the single strand and double strand DNA junction.

Extension products are separated on a denaturing gel and their sizes represent the G-overhang length.

In LMPE a double stranded DNA adaptor bearing a 3’ overhang that is complementary to G-overhang terminal sequence is used. This adaptor is ligated to G-overhang. After ligation, T4 DNA polymerase is used to extend the adaptor and the product is separated on a denaturing gel. The size of the product on the gel represents the length of G-overhang (this will be discussed in detail below). In

DNS digestion, a nuclease specific for double strand DNA is used to digest the 117

bulk of genomic DNA into small fragments (less 10 bp). The digested product is resolved on a native gel and G-overhang is detected by probing with radiolabelled C-rich oligonucleotide probes.

Apart from determination of G-overhang length, various methods have been employed to accurately determine the terminal nucleotide of both G-rich and C-rich strands. Adaptor ligation is used to determine the terminal nucleotide of the G-rich strand (Jacob et al., 2001). In this assay a radiolabelled adaptor with a 3’ overhang is used. Ligation of the adaptor to the G overhang depends on the perfect complementarity between the adaptor and G-overhang. Ligated product is separated on a native gel and only adaptors that are ligated to the telomere are retained in the gel.

Single telomere length analysis (STELA) is used to determine the terminal nucleotide of the C-rich strand (Sfeir et al., 2005). In this assay oligonucleotide including a 5’ unique sequence and a 3’ C-rich telomeric sequence is annealed to the G-overhang. If the oligonucleotide is juxtaposed next to the terminal nucleotide of the C-rich strand, it can be ligated to the C-rich strand DNA. PCR using a forward primer specific to a subtelomeric locus and a reverse primer specific to the unique region of the oligonucleotide is performed and the product is separated by gel electrophoresis and hybridized with telomeric sequence specific probe. Successful ligation of the oligonucleotide to the telomere depends on the precise annealing of the oligonucleotide to the G-overhang at single strand and double strand DNA junction. The terminal sequence of the C-rich

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strand telomere DNA can be determined by the sequence of the primer that was able to be ligated and give PCR product.

4.1.8 G-overhang in T. brucei

The first evidence of telomere G-overhang in T. brucei came from the electron microscopy analysis of T. brucei chromosomes. This analysis confirmed the presence of T-loop on chromosome ends, suggesting the existence of G- overhang (Muñoz‐Jordán et al., 2001). This study further suggested that the length of G-overhang in T. brucei is 75-225 nucleotides (Muñoz‐Jordán et al.,

2001). However, a follow up study using native in-gel hybridization failed to detect any overhang signals at T. brucei telomeres (Dreesen et al., 2005). As the native in-gel hybridization technique is only capable of detecting G-overhang of more than 30 nt long, it was proposed that T. brucei G-overhang is shorter than

30 nt (Dreesen et al., 2005). So far no study has confirmed the length and terminal nucleotide of G-overhang in T. brucei.

In this study, we employed ligation-mediated primer extension to measure the length of T. brucei telomere G-overhang. We also employed the adaptor ligation assay and modified STELA to determine the terminal nucleotide at both the 5’ and 3’ telomere ends in T. brucei. We developed higher resolution assays to study the G-overhang structure in T. brucei. This will help us to better understand the structure and function of telomere in T. brucei, which play important roles in T. brucei pathogenesis. Using our protocols we studied functions of several telomere proteins in G-overhang maintenance. Our results

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further validated that T. brucei is a good model for studying mechanisms of telomere maintenance.

4.2 Results:

4.2.1 The Terminal Nucleotide at the G-rich strand of T. brucei telomeres is tightly regulated

To better understand the terminal structure of T. brucei telomere we employed the adaptor ligation assay published previously (Jacob et al., 2001). In this assay a radiolabelled adaptor with a one-sided 3’ overhang is ligated to telomere ends. The adaptor is obtained by annealing a unique oligonucleotide and a guide oligonucleotide, so that the double stand region of the resulting adaptor has a unique sequence while the 3’ overhang region has telomeric sequences. Six guide oligonucleotides were designed such that their overhangs cover all the possible permutations of TTAGGG. This ensures that at least one of these six adaptors would have an overhang that is the exact complement of the telomere G-overhang terminus. Therefore the 6 nt overhang will provide specificity to the adaptor for ligation of unique oligonucleotide pairs to chromosome ends. Following ligation, genomic DNA is digested with frequent- cutting restriction endonucleases and resolved on a agarose gel. Only the adaptors that are ligated to the chromosome DNA will be retained in the gel

(Figure.4.6).

I performed adaptor ligation to determine the terminal sequence of T. brucei telomere G-overhang. As shown in Figure.4.7a, the adaptor with the 3’

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AATCCC overhang (TG1) was retained in the gel in a large amount compared to other adaptors. This suggests that the TG1 adaptor was able to ligate to the telomere DNA and that most of the T. brucei G-overhangs have a terminal sequence 5’ TTAGGG 3’. Signals of ligation products using other adaptors were very weak except for TG6, whose overhang sequence is 3’ ATCCCA, indicating that a minor proportion of T. brucei telomere G-overhangs have a terminal nucleotide sequence of 5’ TAGGGT. To confirm that the signals are indeed due to ligation of adaptors to the telomere G-overhang, we used EXO-T nuclease as a control. EXO-T is a single strand specific 3’ to 5’ exonuclease that can remove

3’ G-overhang from the chromosome end. As shown in Figure.4.7b, the signals from adaptors were almost all lost when the genomic DNA was treated with EXO-

T. Thus, successful ligation of the adaptor to the chromosome DNA depends on

G -overhang. As a control, we omitted the ligation step, and no signal was detected (Figure.4.8, no ligase lane), indicating that the adaptor needs to be ligated to G-overhang to be retained in the gel end telomere overhang structure specifically.

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Figure 4.6 Schematic of adaptor ligation assay Telomeres with TTAGGG sequences are shown at left. The radiolabeled adaptor with a duplex region bearing unique sequence and a 6nt 3’ overhang is shown on the right. Perfect alignment of the adapter and telomere overhangs allows the ligation of the adapter to the telomere end. Subsequently, genomic DNA was digested with frequent cutting enzyme and separated on agarose gel. The radioactive telomere end can be visualized by exposing the dried gel to autoradiograph films. Signal intensity represents amount of telomere G-overhang. Six different adapters with permutations of the 5’-CCCTAA-3’ overhang were prepared to detect all possible telomere G– overhang signals.

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Figure 4.7 G-overhang ending in bloodstream form T. brucei: (a) Left top Schematic of adaptor ligation assay representing the permuations to which adaptors can anneal. Left, bottom Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. Ligation with TG1 with a 5’- CCCTAA-3’ overhang gives a strong signal, indicating that a large amount of TG1 adaptor has been ligated to the telomere ends. Hence, most telomere G- overhangs end in 5’-TTAGGG-3’. Ligation with TG6 with a 5’-ACCCTA-3’ overhang yields a signal higher than background, suggesting that some of the telomere G-overhangs end in 5’-TAGGGT-3’. (b) Schematic showing genomic DNA after digestion with Exo.T. Exo-T has a 3’–5’ specific single-strand DNA nuclease activity. Treating the genomic DNA with Exo-T prior to adapter ligation removed the signal. 123

Figure 4.8 Signal in adaptor mediated end ligation assay depends on ligation. Genomic DNA of bloodstream form T. brucei was used to perform the assay. Left, Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. No signal was detected reaction ligation step was omitted (No Ligase lane) although TG1 adaptor was incubated with genomic DNA.

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Figure 4.9 Quantification of G-overhang signals of bloodstream form T. brucei (a)Signals were calculated by dividing the autoradiograph signals by the ethidium bromide staining signals. Signals from ligation experiments using different adapters were normalized against that using a non-specific adapter (TGNS) (b) Signals were calculated by dividing the autoradiograph signals by the ethidium bromide staining signals. Signals from ligation experiments using different adapters were normalized against that using a non-specific adapter (TGNS). Percentage of each adaptor signal was calculated from EXO-T sensitive signal. Standard deviation from three independent experiments was calculated.

We performed multiple independent experiments and detected reproducible levels of signals when using each adaptor. To quantify the signals obtained for individual adaptor, we used a nonspecific adaptor (TGNS) whose overhang bears a nontelomeric sequence as a control for nonspecific ligation background. The signals obtained in an autoradiograph were normalized with the amount of DNA used for each adaptor. These signals were further normalized with the signal obtained using the nonspecific adaptor (TGNS) (Figure.4.9a).

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Levels of normalized signals obtained for TG1 and TG6 adaptors were significantly higher than that for TGNS, and EXO-T treatment reduced these signals to nearly background level (Figure.4.9a). These data suggest that T. brucei has two types of telomere G-overhang. The majority of G-overhangs in T. brucei have a sequence complementary to the TG1 overhang while a minor proportion of G-overhangs have a sequence complementary to the TG6 overhang. Thus approximately 70% of telomere G-overhangs end in 5’ TTAGGG

3’, while ~20% end in 5’ TAGGGT 3’ (Figure.4.9b).

Figure 4.10 G-overhang ending in procyclic form T. brucei Left, Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. Genomic DNA treated with Exo-T prior to adapter ligation was used as control.

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Figure 4.11 Quantification of G-overhang signals of procyclic form T. brucei. (a) Signals were calculated by dividing the autoradiograph signals by the ethidium bromide staining signals. Signals from ligation experiments using different adapters were normalized against that using a non-specific adapter (TGNS). (b) Distribution of G-overhang end signals in procyclic form T. brucei. Signals were calculated by dividing the autoradiograph signals by the ethidium bromide staining signals. Signals from ligation experiments using different adapters were normalized against that using a non-specific adapter (TGNS). Percentage of each adaptor signal was calculated from EXO-T sensitive signal. Standard deviation from three independent experiments was calculated.

The same adaptor ligation assay was used to determine the 3’ terminal sequence of G-overhang in procyclic form T. brucei cells. We could not detect any difference in the 3’ terminal sequence of G-overhangs between procyclic and bloodstream form stages (Figure.4.10, 4.11a & 4.11b), suggesting that 3’ end of

G-overhang is similarly regulated in both life stages of T. brucei.

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4.2.2 Telomerase activity is required to maintain the telomere G-overhang in T. brucei

Studies have suggested that telomeric proteins play a major role in maintaining G-overhang at telomeres by regulating telomerase activity and exonuclease activity at telomeres. To determine if any known telomere protein plays a role in G-overhang maintenance in T. brucei we examined the G- overhang structure using the adaptor ligation assay in telomere protein mutants.

Telomerase is a key factor involved in telomere maintenance (Dreesen et al.,

2005). Telomerase uses the telomere G-overhang as a substrate to synthesize the G-rich strands telomeric DNA and may have a capability to alter the terminal sequence of G-overhang.

In T. brucei, deletion of the protein subunit of telomerase (TbTERT) results in progressive shortening of telomeres at a constant rate of 3-5 bp/population doubling (Dreesen et al., 2005). To determine if telomerase activity alters the G- overhang terminal sequence, we performed the adaptor ligation assay in

TbTERT null cells (BF/TERTDKO). As shown in Figure.4.12 and 4.13, deletion of

TbTERT resulted in loss of the 5’-TTAGGG overhang signal complementary to the TG1 adaptor. In contrast, deletion of TbTERT seemed to have no effect on the 5’ TAGGGT overhang signal complementary to the TG6 adaptor. Therefore,

G-overhangs ending in 5’-TTAGGG depend on the telomerase activity, while G- overhangs ending in 5’-TAGGGT do not.

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To confirm if the loss of TG1 adaptor signal is indeed due to the loss of telomerase activity, we ectopically expressed TbTERT-GFP in TbTERT null cells

(BF/TERTDKO/TERTGFP) and performed the adaptor ligation assay. As shown in Figure.4.14 upon expression of TbTERT-GFP, TG1 signal was restore to wild type level. Taken together these results confirm that in T. brucei, telomerase plays an important role in deciding the terminal sequence of G-overhang.

Figure 4.12 TbTERT deletion leads to loss of 5’ TTAGGG 3’ G-overhang endings. Genomic DNA from wild type and TbTERT deleted bloodstream form cells was used to perform adaptor ligation assay. Left, Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. Clear decrease in signal due to ligation of TG1 adaptor was observed upon TbTERT deletion.

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12 BF/Genomic DNA BF/TERTDKO 10

0 Siganl intensity relative to TGNS relative to intensity Siganl

Figure 4.13 Loss of TbTERT results in loss of 5’ TTAGGG 3’-ending G- overhangs. Quantification of adaptor ligation signals in wild type and tbTERT null cells. The telomere G-overhang signals were quantified and normalized the same way as in Figure.4.9. Standard deviation from three independent experiments was calculated.

4.2.3 Deletion of TbKu results in a change in G-overhang dynamics.

Ku is a heterodimer that consists of Ku70 and Ku80. Ku is best known for its essential role in NHEJ, an important mechanism of DNA damage repair. In addition, Ku has also been shown to play an important role in telomere maintenance. In S. cerevisiae and human cells, Ku interacts with both the protein and RNA components of the telomerase complex.

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Figure 4.14 TERT deletion leads to loss of 5’ TTAGGG 3’ G-overhang endings. Genomic DNA from wild type and TbTERT deleted bloodstream form cells was used to perform adaptor ligation assay. Left, Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. Clear decrease in signal due ligation of TG1 adaptor was observed upon TbTERT deletion.

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Figure 4.15 TbKU80 deletion leads to decrease in 5’ TTAGGG 3’ overhang endings and increase in 5’ TAGGGT3’ overhang endings. Genomic DNA from TbKU80 deleted (BF/KU80DKO) and TbKU80 deleted with ectopic expression of GFP-KU80 cells (BF/KU80DKO/GFPKU80) was used to perform adaptor ligation assay. Left, Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. TbKU80 deletion leads to clear decrease in TG1 signal and an increase in TG6 signal, while ectopic expression of TbGFPKU80 leads to increase in TG1 signal and decrease in TG6 signal.

In S. cerevisiae, Ku deletion results in long G-overhangs throughout the cell cycle (Gravel et al., 1998) (Gravel et al., 1998). Similarly in human cells, conditional deletion of Ku86 (Ku80 homolog) leads to a loss of telomere sequence (Wang et al., 2009). In T. brucei, loss of TbKu results in progressive shortening of telomeres at the same rate as TbTERT null mutant. Thus it was proposed that TbKu is involved in telomerase recruitment or activation. To

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determine if loss of TbKu mimics TbTERT’s effect on G-overhang terminal sequence specificity, we perform the adaptor ligation assay in TbKu null cells

(BF/KU80DKO).

Figure 4.16 Loss of TbKU80 results in loss 5’TTAGGG3’ ending Loss of TbKU80 results in loss of G-overhang signals ending in sequence 5’TTAGGG3’ and gain in signal for 5’ TAGGGT 3’ sequence at telomeric ends. The telomere G-overhang signals were quantified and normalized the same way as in Figure.4.9. Standard deviation from three independent experiments was calculated.

As expected (Figure.4.15, 4.16), deletion of TbKu80 resulted in decreased

TG1 signal levels in a manner similar to TbTERT deletion. However, in contrast to TbTERT deletion that had no effect on TG6 signal, TbKu80 deletion resulted in increased TG6 signal. This result indicates that in TbKu null cells, G-overhang terminal sequence is processed into 5’ TAGGGT-ending structure instead of 5’

TTAGGG. Expression of an ectopic allele of TbKu in Ku null cells

(BF/KU80DKO/GFP-KU80) resulted in restoration of the wild type levels of TG1 and TG6 signals (Figure.4.15 right panel and 4.16). 133

4.2.4 Continuous propagation of the telomerase null T. brucei strain leads to a change in G-overhang dynamics.

In budding yeast, Ku is required not only for the recruitment of telomerase but also to protect telomere termini from nuclease activity. Deletion of yeast Ku leads to excessive 5’ resection at telomeres, leading to longer G-overhangs

(Gravel et al., 1998) (Gravel et al., 1998). In contrast to budding yeast, deletion of

Ku in T. brucei has no detectable effect on G-overhang length (Janzen et al.,

2004). Therefore it was proposed that in T. brucei Ku is only required for telomerase recruitment (Janzen et al., 2004). Given these data, we were surprised to observe that telomerase null and Ku null cells have different preference for G-overhang terminal sequences. We propose two possibilities to explain this observation. First, similar to yeast Ku, TbKu also has two functions at telomere ends: it not only recruits telomerase but also protects the telomere end from nuclease activity. Second, TbKu null cells and TbTERT null cells were examined at different times after the deletion was established, and this could be the reason for different G-overhang structures observed in these cells. TbTERT null cells were cultured over a long period of time (~ 2 yrs), resulting in extensive telomere shortening as compared to TbKu null cells.

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Figure 4.17 TbTR deletion leads to a change of G-overhang end sequence dynamics. Genomic DNA from TbTR deletion (BF/TRDKO) was used to perform adaptor ligation assay. Genomic DNA was isolated at different time points i.e.week 4, week 11, and 2.5years. Left, Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. TbTR deletion leads to gradual decrease in TG1 signal and an increase in TG6 signal at early time points (week 4 and week 11) while at later time point most of the signals from adaptors were lost.

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WT/BF

BF/TR-DKO/TR 10 BF/TR-DKO-WEEK4

8 BF/TR-DKO-WEEK11

BF/TR-DKO-2years 6

2 Signal intensity relative to TGNS relative to intensity Signal

Figure 4.18 Loss of TbTR results in a change in G-overhang ending sequence. The telomere G-overhang signals were quantified and normalized the same way as in Figure.4.9. Standard deviation from three independent experiments was calculated.

To differentiate between these possibilities, we established a strain

(BF/TRDKO) that deleted both alleles of the RNA component of telomerase

(TbTR) and compared the G-overhang terminal nucleotide sequence at different time points after deletion of TbTR. As shown in Figure.4.17 and 4.18, upon TbTR deletion the signal from TG1 adaptor decreased significantly, confirming that G- overhang terminal sequence 5’ TTAGGG depends on the telomerase activity. In 136

contrast, signal from TG6 adaptor was increased in TbTR deletion, similarly to the scenario observed shortly after TbKu was deleted (week 4 and 11). However, after a long-term culture (2 yrs), this increase in TG6 signal was reduced

(Figure.4.17 and 4.18). Ectopic expression of TbTR in TbTR null cells

(BF/TRDKO/TR) resulted in reappearance of TG1 signal, which is the same as in wild type cells (Figure.4.19).

Figure 4.19 Complementation of TbTR was able to restore G-overhang terminal sequence. Genomic DNA from TbTR deleted (BF/TRDKO) and TbTR deleted with ectopic expression of TbTR cells (BF/TRDKO/TR) was used to perform ligation mediated end ligation assay. Left, Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. TbTR expression leads clear increase in signal due ligation of TG1 adaptor in TbTR deletion cells.

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4.2.5 MRE11 deletion has no effect on telomere G-rich strand terminal sequence.

Deletion of Mre11, a protein of the MRX complex leads to a decrease in telomere length and G overhang length in budding yeast, suggesting that MRX complex is important for telomere maintenance (Larrivée et al., 2004). In contrast, deletion of Mre11 in T. brucei has no effect on telomere length

Figure 4.20 TbMRE11 is not required for G-overhang end sequence specificity. Genomic DNA from TbMRE11 deleted bloodstream form cells (BF/MRE11DKO) was used to perform adaptor ligation assay. Left, Ethidium bromide stained gel represents the approximately equal amount of DNA loaded in each lane. Right, exposure result of the same gel after drying. No change in G-overhang end sequence was detected in TbMRE11.

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maintenance. In addition, we could not detect any alteration in terminal nucleotide sequence of the telomere G-rich strand upon deletion of Mre11 in T. brucei (BF/MRE11DKO) (Figure.4.20)

4.2.6 The C-rich strand terminal nucleotide sequence of T. brucei telomere is tightly regulated.

Previous studies in Tetrahymena, Euplotes, and human cells have suggested that overhang generation results in a very specific terminal nucleotide sequence at the C-rich strand (Fan and Price, 1997) (Jacob et al., 2001) (Sfeir et al., 2005). To test the regulation of terminal nucleotide sequence of the telomere

C-rich strand in T. brucei, we designed a ligation based PCR assay (Figure.4.21).

In this assay, we ligated guide oligonucleotides to the genomic DNA. PCR amplification was performed using reverse primer complementary to the unique sequence of the guide oligonucleotide and forward primer complementary to telomeric repeats. Southern blot analysis of amplified product was performed using TTAGGG probe to visualize telomere sequence specific signal. Guide oligonucleotides are designed to have 6 nt complementary sequence at its 3’ end that will anneal to the telomere sequence. Six different guide oligonucleotides complementary to the six permutations of telomere sequence 5’ TTAGGG were used to anneal with the G overhang. Ligation of the guide oligonucleotide to the

C-rich strand will take place only if the guide oligonucleotide is placed immediately next to the end of the C-rich strand. Ligation of the guide oligonucleotide with the C-rich strand is a prerequisite for subsequent PCR

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amplification. Relative intensity of telomere specific signals from Southern blot for each guide oligonucleotide can be used to calculate the popularity of a particular terminal sequence of the C-rich strand.

Figure 4.21 Schematic showing ligation mediated PCR assay A guide oligonucleotide containing telomeric repeats at 3’ end was annealed to the telomeric ends followed by ligation. PCR is performed on ligated product using a forward primer specific for telomeric sequence and a reverse primer specific for guide oligonucleotide. Southern blot was performed on the PCR product using a TTAGGG probe.

As shown in Figure.4.22a and 4.23, ~70% of the PCR products was generated using the guide oligonucleotide bearing the 5’ CCTAAC sequence

This result suggests that most of the C-rich strands terminate in 5’ CCTAAC 3’. In order to test specificity of our assay, we randomized the C-rich strand terminal sequence by using T7 exonuclease (T7 EXO) that has a 5’-3’ exonuclease activity. This resulted in the loss of preference for 5’ CCTAAC 3’ (Figure.4.22 b

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and 4.23), suggesting that the assay is reliable. From this we concluded that the terminal

Figure 4.22 Ending of C-Rich strand is tightly regulated (a). Schematic showing ligation mediated PCR based assay. Seven guide oligonucleotides were incubated with genomic DNA in separate reactions, followed by ligation. PCR was performed on ligated product and Southern blot was performed. Signal from second guide oligonucleotide (G2) is significantly higher than other guide oligonucleotides. (b). Genomic DNA was treated with T7 exonuclease that has 5’ to 3’ activity. Treatment with T7 exonuclease results in randomization of C-rich strand terminal nucleotides leading to equal strength of signal detected for all guide oligonucleotides.

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nucleotide of the telomere C-rich strand is tightly regulated in T. brucei and

terminates preferably in 5’ CCTAAC 3’ sequence.

100 BF/Genomic DNA 80

0 Signal intensity relative to TGNS relative to intensity Signal

Figure 4.23 Quantitation of terminal signals of the telomere C-rich strand in bloodstream form of T. brucei. Signals were calculated by normalizing signal obtained from telomere specific guide oligonucleotides with signal obtained from nonspecific guide oligonucleotides. Graph shows the comparative signals obtained from each of the guide oligonucleotides. Signal from guide oligonucleotide two was significantly higher suggesting that about 65% of C-rich strands have terminal sequence of 3’ CAATCC. Signal obtained from guide oligonucleotide three was also higher suggesting that about 25% of the C-rich strands have a terminal nucleotide sequence of 3’ CCAATC. T7 exonuclease treatment resulted in almost equal signal strength obtained from all the guide oligonucleotides. Standard deviation from three independent experiments was calculated.

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4.2.7 Telomerase and TbKu have no effect on the C-strand terminal nucleotide sequence.

As our ligation mediated PCR based assay confirmed that terminal sequence of the C-rich strand is tightly regulated in T. brucei, we next asked if telomere maintenance proteins somehow regulate this terminal sequence.

Telomerase activity is not expected to have any effect on the C-rich terminal sequence as telomerase acts on the G-rich strand to elongate telomeres. As expected (Figure.4.24b and 4.24d), deletion of TbTR had no effect on the C-rich strand terminal sequence. In contrast to telomerase, Ku has been shown to affect the C-rich strand dynamics. In budding yeast, Ku is important for the C-rich strand protection. In the absence of yeast Ku, the C-rich strand is resected by 5’-

3’ exonucleases MRX/Sae2 and Exo1 (Gravel et al., 1998) (Polotnianka et al.,

1998) (Vodenicharov et al., 2010). As deletion of Ku in T. brucei has telomere shortening phenotype, we wondered if TbKu also has a similar role as yeast Ku in telomere maintenance by protecting the C-rich strand from nuclease activity.

To this end, we performed ligation mediated PCR based assay using Ku80 null cells (BF/KU80DKO). As shown in Figure.4.24a and 4.24c, a guide oligonucleotide with the telomere sequence 5’ CCTAAC 3’ produced maximum level of PCR products, similar to that in wild type cells. These results indicate that unlike budding yeast Ku, which protects the C-rich strand from 5’-3’ exonuclease rection, Ku in T. brucei is not required for the C-rich strand protection.

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Figure 4.24 Deletion of TbKu or TbTR has no effect on terminal sequence of the telomere C-rich strand (a, b) TbKU80 null cells as well as TbTR null cells were used to perform ligation mediated PCR based assay. As shown above, no change in terminal nucleotide sequence of C-rich strand was detected upon deletion of TbKU80 or TbTR. (c) Graph showing quantitation of signals obtained for terminal nucleotide sequence of C-rich strand in WT and TbKU80 null cells. Signal detected for guide oligonucleotide 2 (G2) were most prominent in WT as well as TbKU80 null cells. (d) Graph showing quantitation of signals obtained for terminal nucleotide sequence of C-rich strand in WT and TbTR null cells. Standard deviation from three independent experiments was calculated. Signal detected for guide oligonucleotide 2 (G2) were most prominent in WT as well as TbTR null cells.

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4.2.8 Ligation mediated primer extension assay for measuring the length of G-overhang

Length of the telomere G-overhang typically varies among different eukaryotic species. In human cells, it is approximately 12-300 nt long, whereas in budding yeast, it ranges from 10-30 nt (Chai et al., 2006) (Chow et al., 2012)

(Soudet et al., 2014). In T. brucei, G-overhang is thought to be very small but the exact length is not known (Dreesen et al., 2005). To address this question, we set out to measure the length of G-overhang at a resolution of single nucleotide.

We employed ligation mediated primer extension assay previously used to measure the G-overhang length in Tetrahymena (Jacob et al., 2001). In this method, an oligonucleotide with a unique sequence is annealed to a radiolabelled guide oligonucleotide that has an additional 6 nt of telomere sequence at its 3’ end. After annealing the guide oligonucleotide with the unique oligonucleotide, the resulting 6 nt 3’ overhang acts as an adaptor that can base pair with the G-overhang terminal sequence. Following ligation, primer extension is performed using T4 DNA polymerase that lacks strand displacement and 5’–3’ exonuclease activities. As a result, the guide oligonucleotide is extended to the single strand–double strand DNA junction. The extension product is resolved on a polyacrylamide denaturing gel. The length of the guide oligonucleotide region bearing the unique sequence is subtracted from the length of the final primer extension product to give the length of the telomere G-overhang (Figure.4.25).

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Figure 4.25 Schematic showing steps of ligation mediated primer extension assay. Telomeres with TTAGGG sequences are shown at left. Seven radiolabelled guide oligonucleotides (red triangle) were annealed to unique oligonucleotides to make an adaptor with a duplex region bearing unique sequence and a 6nt 3’ overhang is shown on the right. Perfect alignment of the adapter and telomere overhangs allow the ligation of the adapter to the telomere end. Following ligation, guide oligonucleotide was extended using T4 DNA polymerase until single stranded–double stranded DNA junction was reached. Extended product was separated on polyacrylamide gel and length of the overhang was calculated by subtracting the length of unique sequence guide oligonucleotide from length of final extension product.

In order to optimize the protocol, first we designed artificial overhangs of known length and used them as a substrate for our primer extension assay.

Three different artificial substrates of different overhang lengths i.e. 16 nt, 26 nt, and 36 nt were used for the primer extension assay.

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Figure 4.26 Ligation mediated primer extension is a sensitive method to detect the telomere length of G-overhang. (a) Three artificial overhangs of different sizes (36nt, 26nt and 16nt) were designed bearing a sequence complementary to TG1 adaptor. Primer extension was performed using these overhangs as substrates. (b) Extension products were resolved on a polyacrylamide gel. The expected sizes of extension products were obtained. 147

Figure 4.27 Complementarity of the guide oligonucleotide with the terminal G-overhang sequence is a prerequisite primer extension Adaptors used in ligation mediated primer extension assay are equally competent to anneal with complementary overhangs and also for subsequent extension. Two artificial overhangs of different sizes (26nt and 16nt) were designed bearing a sequence complementary to TG4 adaptor. Primer extension was performed using these overhangs as substrates. Extension 148

products were resolved on a polyacrylamide gel. The expected sizes of extension products were obtained.

As shown in Figure.4.26, we were able to measure the overhang length at the resolution of single nucleotide. Also, we used two types of artificial overhangs that have different terminal sequences to confirm that all the adaptors used in this assay are competent for extension by T4 DNA polymerase. As shown in

Figure.4.26, when an artificial overhang complementary to adaptor TG1 (5’

CCCTAA 3’) was used, extension product was detected only in TG1 lane.

Consistently, when an artificial overhang complementary to TG4 adaptor (5’

TACCCT) was used, extension product was detected in TG4 lane (Figure.4.27).

These results suggest that this assay is highly sensitive for measuring length of the overhang accurately. Again, the complementarity of the guide oligonucleotide with the terminal G-overhang sequence is a prerequisite for a successful ligation and subsequent primer extension. Our data confirmed that all guide oligonucleotides are equally competent in annealing with their complementary sequences and subsequent extension by T4 DNA polymerase.

4.2.9 The G-overhang length of T. brucei telomere

We employed the ligation-mediated primer extension assay to measure the length of the telomere G-overhang in T. brucei. As shown in Figure.4.28, an extension product (lane 2) of approximately 28 nt was obtained when the guide oligonucleotide TG1 was used. No extension product was detected when other guide oligonucleotides were used. The G-overhang length was calculated by subtracting the 16 nt (length of the guide oligonucleotide region with the unique 149

sequence) from the final extension product. Therefore the length of G overhang in T. brucei is approximately 12 nt. Using adaptor ligation, we detected two types of G-overhangs complementary to TG1 and TG6 adaptors. However using the ligation mediated primer extension assay we could not detect any extension product with the TG6 adaptor (lane 7). It is possible that the G-overhang ending with 5’-TAGGGT (corresponding to TG6 adaptor) are very short (~6-7 nt), therefore adaptors with six nucleotide overhang can only anneal to G-overhang but is not able to extend it.

4.3 Discussion

In first part of the study we adopted a couple of reliable and sensitive methods to determine the terminal nucleotide sequence of the telomere G- overhang in T. brucei. We modified the adaptor ligation assay and used it to examine the structure of T. brucei G-overhang. We optimized the adaptor ligation assay so that it is sensitive to detect minor alterations in the terminal nucleotide sequence of G-overhang. Different concentrations of oligonucleotides and genomic DNA were used to obtain a ratio that resulted in reproducible results.

We used different controls to validate our assay. To verify that the adaptors used in this assay were specific for the telomere G overhang, we used an EXO-T exonuclease as a control. EXO-T has a 3’ to 5’ exonuclease activity specifically for single-strand DNA, which can remove G-overhang. As expected, treating genomic DNA with EXO-T prevented most of adaptor ligation to the telomere,

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Figure 4.28 T. brucei telomeres have short G-overhangs Genomic DNA was ligated to six different adaptors followed by primer extension using T4 DNA polymerase. End-labeled TG1 oligonucleotide was loaded as a negative control (lane 2). The oligonucleotide itself runs at 22 nt. Only TG1 adaptor yielded extension products significantly longer than the guide oligonucleotide itself (red arrow head, lane 3).

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thereby confirming that this assay specifically detects the telomere G-overhang signal.

We also used an adaptor bearing an overhang with nontelomeric sequence (TGNS) to exclude the possibility of random annealing and ligation of adaptors to genomic DNA. Also, to confirm that ligation of adaptors to genomic

DNA is a prerequisite for their retention in the gel and subsequent detection; we omitted the ligation step from the assay as a negative control. As expected, no adaptor signal was detected in the absence of ligation. Thus, the adaptor ligation assay that we adopted and modified is a sensitive and accurate method to detect the terminal nucleotide sequence of G-overhang.

We employed the adaptor ligation assay to determine the terminal nucleotide sequence of T. brucei G-overhang. In telomerase positive cells, most telomeres end with the sequence 5’ TTAGGG, while some telomeres have the 5’

TAGGGT terminal sequence. A possible explanation for this is that the telomerase copies the RNA template (5’ CCCUAACCC 3’) completely before it dissociates from the telomere end. These results are consistent with the in vitro telomerase primer extension assay performed using T.brucei cell lysate (Cano et al., 1999). In this assay, various primers with telomere sequence permutations were used as the substrates for telomerase. Result obtained from this assay not only suggested that T. brucei telomerase can extend any telomere sequence permutation but also confirmed that T. brucei telomerase translocates only after it copies complete RNA template (Figure.4.29). Extrapolating these results in vivo would mean telomerase activity on telomere ends should result in 5’TTAGGG3’ 152

permutation. Indeed results from our adaptor mediated ligation assay confirmed that in wild type cells, the majority of telomeres end with 5’ TTAGGG 3’ while this preference was lost when telomerase activity was compromised. This result further suggests that telomerase in T. brucei acts on the majority of telomeres and maintains their ends as 5’ TTAGGG 3’.

Figure 4.29 Schematic showing steps of telomerase dependent telomere elongation. TERT copies nucleotides onto the telomere end and dissociates upon reaching the 5' end of the template thereby specifying terminal nucleotide of the telomere.

This situation is different from budding yeast in which telomerase acts only on a subset of telomeres in each cell cycle. Although detailed mechanism of this regulation is not clear, studies have revealed some interesting insights. In budding yeast, short telomeres are preferentially elongated by telomerase while telomerase activity at long telomeres is restricted. It is estimated that under wild type condition (telomere length about 300bp), only 6-8% of telomere ends are elongated by telomerase in each cell cycle (Teixeira et al., 2004). Studies in human cells have suggested that unlike yeast telomerase, mammalian telomerase acts on most of the telomere ends in each cell cycle (Zhao et al.,

2009). Similarly it is possible that in T. brucei, telomerase acts on most of the

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telomere ends and thereby counteracts different processes (for e.g.. end replication problem or nuclease activity) that may alter the 3’ end of the telomeres. Moreover studies in both yeast and mammalian cells have demonstrated that telomerase activity is restricted to S phase of the cell cycle

This regulation is proposed to be important for maintaining telomere length homeostasis (Diede and Gottschling, 1999) (Marcand et al., 2000) (Ten Hagen et al., 1990) (Wright et al., 1999) (Jády et al., 2006) (Zhao et al., 2009). It is not clear if such a restriction is also exists on T. brucei telomerase. In contrast to other organisms where telomere length is maintained within a certain range, in T. brucei, telomeres grow at a constant rate of 6-8bp per population doubling

(Bernards et al., 1983; Dreesen et al., 2005) (Sandhu et al., 2013). These results indicate that telomere length homeostasis is not maintained in T. brucei.

Therefore it is possible for telomerase to act on telomere ends without any restriction and thereby maintaining the 5’ TTAGGG 3’ ends (Figure.4.30).

Interestingly, some telomere G-overhangs end in 5’ TAGGGT 3’ in the wild type cells. We suggest two possible explanations for this scenario. First, telomerase may not be able to fully copy the RNA template all the time before it dissociates from the telomere substrate. Second, this type of G-overhang may be processed independently of the telomerase activity. To better understand the role of telomerase in regulating G-overhang terminal sequence, we determined the terminal sequence of G-overhang in cells that lack the telomerase activity

(TbTERTDKO and TbTRDKO). We observed that upon deletion of the telomerase protein component (TbTERTDKO), the amount of G-overhangs 154

ending in 5’ TTAGGG 3’ was significantly reduced. This supports our previous hypothesis that continuous activity of telomerase is required to maintain 5’

TTAGGG 3’-ends. No change was detected in the amount of G-overhangs that end in 5’ TAGGGT 3’ in these cell lines. These data indicate that the 5’ TAGGGT

3’ ending of G-overhangs is not dependent on the telomerase activity

(Figure.4.30).

Figure 4.30 Telomere terminal sequence in T. brucei (a) G-overhang length is small in T.brucei. Terminal nucleotide of both C and G strand is very precise and telomerase is the main determinant of G strand terminus (b) Unknown activity becomes a major determinant of G-strand terminal nucleotide in the absence of telomerase while C strand terminal nucleotide is maintained independently of telomerase.

We were surprised to observe that loss of 5’ TTAGGG 3’ ends didn’t result in gain of any other types of G-overhang ends in TbTERTDKO cells. We propose two scenarios in which this could happen. First, it is possible that G-overhang ends were randomized upon telomerase deletion but overhangs were too small to be detected by our assay. Second, it is also possible that extensive culturing of

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telomerase null cells resulted in activation of telomere maintenance pathway that leads to an overhang structure that is not detected by our assay. Cancer cells that lack telomerase activity and rely on recombination to maintain telomeres have 5’ as well as 3’ overhangs (Oganesian and Karlseder, 2011). Similar mechanism of telomere maintenance could also exist in T. brucei cells lacking telomerase. Such a mechanism would result in G-overhang structure that is not be recognized by our adaptor based end ligation assay and will result in overall decrease of G-overhang end signals. Studies in yeast have established that the propagation of yeast strains without telomerase results in a dynamic change in telomere structure and maintenance (Lundblad and Blackburn, 1993). Culturing of yeast strains without telomerase results in telomere shortening in early population doublings (Lundblad and Szostak, 1989) (Lundblad and Blackburn,

1993). Once telomeres fall below a critical length, recombination based telomere maintenance is triggered (Lundblad and Blackburn, 1993). As our TbTERT cell lines are extensively cultured, it is possible that these cell lines have established recombination or some other unknown mechanism to maintain telomeres.

To exclude this possibility and to examine the kinetics of the change in G- overhang terminal sequence,, we performed the adaptor based end ligation assay at different population doublings after deletion of telomerase RNA component (TbTRDKO). We observed a clear transition from 5’ TTAGGG-ending

G-overhangs to 5’ TAGGGT-ending G-overhangs soon after deletion of TbTR

(week 4 to week 11). These data suggest that after telomerase deletion other overhangs do accumulate. The fact that we do not detect randomization of G- 156

overhang ending in the absence of telomerase suggests that an unknown mechanism exists that prefers 5’TAGGGT 3’ ending of G-overhangs. At early population doublings, the total amount of G-overhang signal remains comparable to the wild type. This indicates that most of the telomeres still possess G- overhangs, however these overhangs now end with 5’ TAGGGT 3’ instead of 5’

TTAGGG 3’.

Upon extensive culture of TbTR null cells (2 yrs), the level of 5’ TAGGGT- ending G-overhangs is decreased to the wild type value resulting in an overall reduction of the G-overhang signal. It is possible that overhangs at this stage are too small to detect or telomerase independent stabilization of telomeres results in an overhang structure that is not detected by our assay. T. brucei genome consists of three types of chromosomes : 11 base of mega base chromosomes,

4-5 intermediate chromosomes , and ~100 minichromosomes (Van der Ploeg et al., 1984c) (Melville et al., 1998). At this stage we are not sure if the overhangs at these different types of chromosomes are processed differently or similarly.

Although speculative, it is possible that telomere ends are processed differently at different types of chromosomes. In this scenario, it is possible that G- overhangs with 5’ TAGGGT 3’ ends are preferentially maintained on a certain type of chromosomes. As we detect the wild type level of 5’ TAGGGT 3’ signal in extensively cultured telomerase null cells, this would mean that the type of chromosomes with 5’ TAGGGT 3’ ending are able to protect their G-overhangs.

Therefore these chromosomes retain a similar to the wild type level of 5’

TAGGGT 3’ signal. 157

In Tetrahymena, telomerase-independent processing of G-overhang terminal sequence has been proposed (Jacob et al., 2003). It has been suggested that a 3’ to 5’ nuclease activity is responsible for generating a specific terminal sequence at G-overhang (Jacob et al., 2003). However the nuclease responsible for such activity has not been identified. Recently in human cells,

WRN (Werner syndrome protein), a RecQ helicase was shown to have 3’ to 5’ exonuclease activity on the telomeric substrate in vitro (Li et al., 2009a). The activity of WRN was strictly dependent on the presence of telomeric sequences in both duplex DNA as well as in 3' overhang (Li et al., 2009a). This suggests that

WRN processes telomere DNA with a 3' single-strand overhang with high specificity (Reddy et al., 2010). In T. brucei, an exonuclease activity could also be responsible for processing G-overhang termini from 5’ TTAGGG to 5’

TAGGGT in the absence of the telomerase activity. Alternately, the terminal sequence 5’ TAGGGT could also result from the removal of the last RNA primer during replication (end replication problem). In WT cells, the majority of these ends are extended by telomerase leading to a terminal sequence of 5’ TTAGGG.

In contrast, when telomerase activity is absent, 5’ TAGGGT ends accumulate.

Given our data of C-rich strand terminal sequence the latter scenario does not seem likely. As C-rich strand in T. brucei ends in 5’CCTAAC 3’, it would mean that the G-rich strand of the telomere should end in 5’GTTAGG 3’ after DNA replication. As in our assay we detect 5’ TAGGGT 3’ ending instead of 5’

GTTAGG 3’, therefore processing of the 3’ end of the G-rich strand seems to be required. Nonetheless, these findings warrant further analysis of G-overhang 158

structure to identify potential factors that may play a role in specifying terminal sequence of G-overhang in the absence of telomerase.

The Ku70/80 heterodimer is a conserved DNA binding protein with several cellular functions. Apart from being a major component of DNA double strand break repair machinery; Ku also plays an important role in telomere maintenance. At telomeres, Ku is involved in recruitment of telomerase and also protects the C-rich strand. We speculated that TbKu could affect G-overhang status as deletion of tbKu80 results in telomere shortening in a manner similar to telomerase mutant. As expected, the deletion of Ku80 resulted in an increase in

5’ TAGGGT terminal sequence, similarly to that in telomerase null cells. These findings further suggest that Ku is required for recruiting telomerase to the telomere ends, but further investigation is necessary to confirm this hypothesis.

Lastly, we determined the G-overhang terminal sequence in TbMre11 null cells. In budding yeast, deletion of Mre11 leads to a defect in G-overhang formation. This defect is mainly due to the role of Mre11 in resection of C-rich strand to generate G-overhang (Bonetti et al., 2009). In contrast, deletion of

Mre11 in vertebrates has no detectable effect on G-overhang maintenance (Wu et al., 2012). Using the adaptor ligation assay, we could not detect any change in

G-overhang terminal sequence in TbMre11 null cells when compared to WT cells. These finding are in agreement with the previous report that T. brucei

Mre11 is not required for telomere maintenance.

In the second part of this study, we optimized a ligation mediated PCR assay to examine the terminal sequence of the C-rich strand. We modified 159

previously published assay STELA and implemented it to determine the C-rich strand terminal sequence and its regulation in T. brucei. Different ligation conditions and number of PCR cycles were tested to increase reproducibility and sensitivity of the assay. Using ligation mediated PCR assay, we observed that the terminal sequence of the C-rich strand is tightly regulated and ends with5’

CCTAAC 3’. We used T7 EXO as a control to verify the specificity of our assay.

T7 EXO has a 5’ to 3’ specific exonuclease activity and treating genomic DNA with T7 EXO resulted in resection of the C-rich strand so that it terminates with a random permutation of the telomere repeat. We observed the expected phenotype, suggesting that this assay is very sensitive to detect alterations in the

C-rich strand terminal sequence.

Our data suggests that the vast majority of C-rich strands in T. brucei end with the sequence 5’ CCTAAC 3’ Thus, C-rich strand processing is stringently regulated. This observation is in agreement with results from ciliates (Euplotes and Tetrahymena) and humans (Fan and Price, 1997) (Jacob et al., 2001) (Sfeir et al., 2005). In Euplotes, the terminal sequence of the C-rich strand is always 5’

CCCCAAAA 3’ while in Tetrahymena the C-rich strand ends with either 5’CCAA3’ or 5’ CCCAA 3’(Fan and Price, 1997) (Jacob et al., 2001). In human cells, most of C-rich strands bear a terminal sequence 5’ CTAACC 3’(Sfeir et al., 2005).

These observations suggest that the mechanism of specifying terminal sequence of the C-rich strand is conserved. So far no factor has been identified that plays a role in specifying terminal sequence of the C-rich strand. As telomerase can only act on the 3’ end of G-overhang, its activity should not affect the terminal 160

sequence of the C-rich strand. In agreement with this, deletion of TbTR had no effect on the terminal sequence of the C-rich strand. In S. cerevisiae, Ku is proposed to protect the C-rich strand from resection by exonucleases (Gravel et al., 1998) (Polotnianka et al., 1998) (Boulton and Jackson, 1996). This prompted us to analyze the terminal sequence of the C-rich strand in the absence of

TbKu80. However, we observed no alteration in the C-rich strand terminal sequence in TbKU80 null cells, suggesting that in T. brucei, Ku may only be required for telomerase recruitment but not for C-rich strand protection.

Several models have been proposed for the regulation of the terminal sequence of the C-rich strand. Given the fact that the G-overhangs of lagging strand and leading strand are generated by different mechanisms, it is possible that entirely different mechanisms are used to generate terminal sequence specification of the C-rich strand on leading versus lagging strand, which suggests the involvement of two separate complexes. Recently, in humans it was shown that the lagging strand C-rich terminus is specified immediately after replication, while the specification of the leading strand C-rich terminus is delayed several hours after replication (Chow et al., 2012). These observations clearly indicate two independent steps to specify the terminal nucleotide of the C- rich strand at leading and lagging strand. Another explanation could be that the same complex works at the last step of maturation of leading and lagging C-rich strands. In humans, lagging C-rich strand is proposed to mature immediately after removal of the primer from last Okazaki fragment, which would be responsible for specifying the terminal nucleotide (Chow et al., 2012). 161

On the other hand, resection of the leading daughter C-rich strand leads to the generation of long G-overhangs that is subsequently filled in by the CST- dependent DNA replication by conventional DNA polymerase (Chow et al.,

2012). In this fill-in process, RNA primers are also required at their C-rich strand termini, which would be removed following the fill-in. This would mean that the specification of the terminal sequence of both C-rich strands (leading and lagging) could occur by the same mechanism i.e. by removal of the last RNA primer. On the contrary, it was reported that depletion of human Stn1 does not affect specification of the terminal nucleotide of the C-rich strand (Dai et al.,

2010) (Huang et al., 2012). As Stn1 is required for proper activation of polymerase to fill in the C-rich strand, it was concluded that C-rich strand fill-in might not be responsible for specifying the terminal nucleotide of the C-rich strand (Dai et al., 2010) (Huang et al., 2012). So far, the only known factor that affects the C-rich strand terminal nucleotide is the G-overhang binding protein

POT1. Depletion of hPOT1 results in randomization of C-rich strand termini, suggesting that binding of POT1 to the G-overhang regulates the C-rich strand process (Hockemeyer et al., 2005) (Dai et al., 2010). However the mechanism how POT1 regulates the C-strand ending is not clear. Here we show that the terminal nucleotide of the C-rich strand in T. brucei is also regulated, underscoring the importance of the precise telomere terminal structure.

Currently, the mechanism responsible for specifying the C-rich strand terminal nucleotide is elusive; our studies in T. brucei have provided the necessary tools

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to examine the terminal structure of telomere, which will help to identify factors that contribute to telomere C-rich strand process.

In the third part of our study, we aimed to accurately measure the telomere G-overhang length. We employed LMPE, previously used to accurately measure the G-overhang length in Tetrahymena (Jacob et al., 2001). Artificial substrates containing telomere overhangs were used to standardize different steps of this assay. We confirmed that ligation and subsequent extension of the adaptor was dependent on the presence of complementarity between the adaptor overhang and the artificial substrate overhang. Using this assay we predicted that G-overhang length at T. brucei telomeres is about 12nt.

To test the sensitivity of LMPE, we treated the T. brucei genomic DNA with EXO-T. As EXO-T has a 3’ to 5’ exonuclease activity specific for single- strand DNA, we expected to see no extension product after EXO-T treatment.

However, we saw no difference in the extension product whether the genomic

DNA was treated with EXO-T or not. We tried many different conditions but failed to eliminate the extension product with the EXO-T treatment. Different adaptor concentrations (from 0.1 picomole to 10 picomole) were tested. Different ligation conditions (from 30-minute incubation at room temperature to 16ºC incubation overnight) were used. Primer extension conditions (temperature and time) were modified. In summary, we were unable to standardize a condition in which the primer extension products were detected using untreated genomic DNA and abolished using the EXO-T treated genomic DNA.

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Many factors can lead to this observation. First, the signal that we are detecting in LMPE assay is non-specific and is generated from ligation of adaptors to random DNA breaks or nicks in the genomic DNA. We rule out this possibility because the signal was reproducibly detected only when the TG1 adaptor was used. Second, it could be an intrinsic property of the TG1 adaptor

(with an overhang that is compatible to the telomeric overhang sequence 5’

TTAGGG 3’) to produce an extension product. However, when an artificial substrate with an overhang of 5’ GGGTTA 3’ was used in LMPE, the TG4 instead of the TG1 adaptor was able to produce an extension product, arguing that all adaptors used are equally competent to be ligated and produce extension products. Third, it is possible that a small amount of adaptors can still be ligated to the residual overhangs that survived the EXO-T exonuclease activity. In

LMPE, the guide oligonucleotide was radiolabeled. Only the primer extension products were detectable in the end result, while the total amount of telomere G- overhang is not measured in this assay. Therefore, if a few G-overhangs survive the EXO-T treatment, they will be ligated to the adaptor and result in extension products. In contrast, the adaptor ligation assay detects the total amount of G- overhangs that can be ligated to the adaptor. Treating genomic DNA with EXO-T greatly decreased the telomere ends that can be ligated with the adaptor.

However, residue ligated products can still be detected in the gel, supporting the hypothesis that EXO-T does not completely eliminate all G-overhangs. In summary, since EXO-T treatment cannot eliminate all primer extension products in LMPE, we cannot rule out the possibility of some non-specificity of the assay. 164

Nonetheless our data suggest that the primer extension product likely reflects G- overhang length in T. brucei. Further studies are necessary to confirm these findings.

4.3.1 The molecular mechanism of overhangs processing

Results from our study suggest that the terminal nucleotide of both G and

C-rich strands is tightly regulated. The specificity of the G-rich strand terminal nucleotide seems to depend on two independent processes. One is the telomerase activity that defines the terminal sequence of the majority of G-rich strands as 5’ TTAGGG 3’. Another is an unknown mechanism most likely involving a 3’ to 5’ exonuclease activity that acts on residual telomere ends in the presence of telomerase to produce a terminal sequence 5’-TAGGGT-3'. This 3’ to 5’ exonuclease activity is prominent in telomerase null cells. Specification of C- rich strand terminal sequence can be achieved by removal of the last RNA primer at the lagging strand and by 5’ to 3’ exonuclease activity at the leading strand.

One mechanism to generate the exact nucleotide at the termini of both G and C- rich strands is to involve activities of 3’ to 5’ and 5’ to 3’ exonucleases along with telomere binding proteins (Figure.4.31).

After DNA replication, the 5’ to 3’ exonuclease activity processes the C- rich strand and generates sufficiently long overhang that is subsequently bound by a single-strand DNA binding protein. Upon binding to the G-overhang, this protein will prevent further resection of the C-rich strand and will also provide a boundary for the G-rich strand processing. At C-rich strand, this G-overhang

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Figure 4.31 Proposed mechanism for the the G-overhang maintenance (1) After telomere replication, leading and lagging telomere ends are generated. (2) Leading strand replication results in blunt ended telomeres. Red color represents daughter strand of DNA and while black color represents parent strand of DNA. (3) Putative 5’ to 3’ exonuclease activity (red packman) results in overhang generation at the leading strand and this overhang can be extended by telomerase. (4) Extended overhangs are bound by single stranded DNA (ssDNA) binding protein (green circle). This binding leads to the protection of overhang and possibly specification of last nucleotide on C strand. (5) Lagging strand replication results in generation of overhang due to the removal of last RNA primer. Blue color represents daughter strand of DNA while black color represent parent strand of DNA (6) Overhang at the lagging strand can be further extended by telomerase (7) Overhang bound by ssDNA binding protein and C strand is specified. (8) 3’ to 5’ exonuclease activity (orange packman) can process G-overhang and this activity is blocked by ssDNA binding protein. (9) 3’ end of G-overhang is specified by ssDNA binding protein, which protects 5’ TAGGGT 3’ ends from 5’ to 3’ exonuclease activity. Telomerase activity can extend these endings to 5’TTAGGG’3, thus in the presence of telomerase activity, majority of the telomeres end with 5’ TTAGGG 3’. In contrast, in the absence of telomerase activity, 5’ TAGGGT 3’ are more prevalent due to the exonuclease activity.

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binding protein can perform two functions. First, it blocks 5’ to 3’ exonuclease activity and second, it recruits C-strand fill-in factors. Thus G-overhang binding protein can regulate C-rich strand ending by either blocking exonuclease activity or by positioning that last RNA primer used for strand fill-in. Indeed, study in mouse cells suggests that POT1b performs both of these activities at telomere ends (Wu et al., 2012). Moreover in humans depletion of POT1 results in randomized C-strand termini suggesting the role of single strand binding protein in regulating C-strand ending (Hockemeyer et al., 2005).

Studies have also shown that proteins that bind to duplex telomeric sequence play an important role in G-overhang maintenance. Study in budding yeast suggests that RAP1 along with Rif1 and Rif2 plays an important role in C-rich strand protection. Removal of this complex from the telomere ends results in extensive C-rich strand degradation (Bonetti et al., 2010) (Vodenicharov et al.,

2010). Moreover, in vitro study of human RAP1 suggests that RAP1 along with

TRF2 binds to single strand double strand junction of telomeric sequence (Arat and Griffith, 2012). Although relevance of this binding in vivo is not known, it is possible that this complex is also involved in telomere C-rich strand protection.

Similar mechanism of C-rich strand protection could also define terminal nucleotide of C-rich strand in T. brucei. At G-overhang, single strand binding protein will prevent the 3’ to 5’ exonuclease activity and thus define 5’ TAGGGT

3’ overhang endings. In telomerase positive cells, most of these ends will be extended by the telomerase and will have the sequence 5’ TTAGGG 3’.In

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contrast, upon loss of telomerase activity majority of G-overhangs end in 5’

TAGGGT 3’ sequence (Figure.4.32).

Figure 4.32 Proposed mechanism of the telomere C-strand terminal nucleotide specification Binding of double strand DNA binding protein(s) will inhibit the action of nuclease most probably by limiting the access of exonuclease creating a boundary for exonuclease action. Second, a single strand DNA binding protein possibly limits the access of exonuclease at both G and C strand.

4.4 Conclusion

Much remains unknown about the mechanism of how telomere ends are maintained in eukaryotes. Using several molecular approaches, we provide new insight into factors that contribute to telomere integrity in T. brucei. We successfully developed techniques to examine the dynamics of telomere G- overhang that could be used to better understand molecular steps that lead to maturation of the telomere terminal structure. Our study has laid the groundwork for understanding fundamental mechanisms of telomere G-overhang maintenance in T. brucei thus developing T. brucei as a model for telomere biology.

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5CHAPTER V

Functional Analysis of Domains of the Telomeric Protein TbRAP1

5.1 Introduction

Telomeres are bound by specialized protein complex that is required for proper telomere function and maintenance. Over the past 20 years, many telomere-associated proteins have been identified and characterized. One of the first protein to be identified as telomere-associated factor was RAP1. After its identification in S. cerevisiae, it was later shown that RAP1 is highly conserved and is present in T. brucei, S. pombe, and vertebrates (Longtine et al., 1989)

(Yang et al., 2009) (Kanoh and Ishikawa, 2001) (Chikashige and Hiraoka, 2001)

(Li et al., 2000). At telomeres RAP1 is required for many different aspects of telomere function and maintenance including telomere length regulation, protection of telomere ends from resection, inhibition of DNA damage checkpoint, and telomeric silencing (Marcand et al., 1997) (Kanoh and Ishikawa, 2001)

(Vodenicharov et al., 2010) (Miller et al., 2005)

In addition, Rap1 also acts as a transcription factor and controls the expression of various genes in S. cerevisiae and vertebrates (Shore and

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Nasmyth, 1987) (Lieb et al., 2001) (Martinez et al., 2010) (Lickwar et al., 2012).

ScRAP1 regulates the telomere length by recruiting Rif1 and Rif2 to telomeres

(Wotton and Shore, 1997). C-terminal truncation of scRAP1 or deletion of

Rif1/Rif2 proteins results in telomere elongation, suggesting that scRAP1 is a negative regulator of telomere length (Hardy et al., 1992) (Wotton and Shore,

1997). Consistent with its role as a negative regulator, overexpression of scRAP1 led to telomere shortening. Based on scRAP1 tethering experiments a ‘protein counting’ model has been proposed (Marcand et al., 1997). According to this model, the number of scRAP1 molecules bound to telomeres serves as a gauge for the telomere length sensing mechanism. Although it is not clear how scRAP1 exerts its role in telomere length regulation, recent studies have suggested two possible mechanisms. First, scRAP1, together with Rif1 and Rif2, restricts telomerase activity in G1 phase (Gallardo et al., 2011) thereby preventing telomere elongation. Second, scRAP1 in complex with Rif2 and, to a less extent, with Rif1 regulates telomere end processing by the MRX complex (Bonetti et al.,

2010) (Vodenicharov et al., 2010). A study in cdc13 temperature sensitive mutant background revealed that scRAP1, alongwith Rif1 is important for telomere capping and is thus required for telomere protection function (Anbalagan et al.,

2011) (Xue et al., 2011). Also, scRAP1 inhibits NHEJ at telomeres by multiple pathways (Pardo and Marcand, 2005). Along with Rif2, ScRAP1 prevents NHEJ at telomeres most likely by preventing the activation of MRX complex (Marcand et al., 2008). Moreover, studies targeting scRAP1 and Rif2 to non-telomeric DSB sites revealed that the scRAP1-Rif2 complex acts as NHEJ inhibitor independent 170

of telomere sequence. At telomeres, scRAP1 inhibits NHEJ also by recruiting

Sir4 (Marcand et al., 2008). However the mechanism by which Sir4 blocks NHEJ at telomeres is not clear. Lastly the DNA binding domain of scRAP1 inhibits

NHEJ independently of Rif2 and Sir4 (Marcand et al., 2008).

In S. pombe, spRAP1 along with Taz1 maintains telomere length homeostasis by inhibiting telomere elongation and controlling the telomere 3’ overhang generation (Kanoh and Ishikawa, 2001) (Miller et al., 2005). SpRap1 prevents NHEJ at telomeres but is shown to promote homologous recombination in the absence of Taz1 (Subramanian et al., 2008). Also spRAP1 is important for telomere clustering towards the spindle pole body at the premeiotic horsetail stage (Chikashige and Hiraoka, 2001).

In mammals, RAP1’s function at telomeres is complicated. In humans,

RAP1 is an essential protein, while in mice it is non-essential (Li et al., 2000)

(Martinez et al., 2010). In humans, RAP1 negatively regulates telomere length and is required for inhibition of NHEJ at the telomere (Li et al., 2000) (Bae and

Baumann, 2007) (Sarthy et al., 2009). In contrast, mice have mildly shortened telomeres when RAP1 is deleted (Sfeir et al., 2010). Intriguingly, mouse RAP1 is not required to inhibit NHEJ but is important for suppressing homology directed repair (Sfeir et al., 2010). Regardless of the species-specific differences in the functions of RAP1, it is evident that RAP1 is important for telomere length maintenance and protection.

Besides its role in telomere length maintenance and protection, RAP1 represses transcription of subtelomeric genes by promoting telomere position effect TPE. 171

C-terminal deletion of scRAP1 leads to a complete loss of TPE, underscoring the importance of RAP1 in establishment of a repressed chromatin structure

(Aparicio et al., 1991). Deletion studies in fission yeast and mice further confirmed the role of RAP1 in establishing TPE in these organisms (Kanoh and

Ishikawa, 2001) (Martinez et al., 2010). Apart from its role at telomeres, RAP1 also acts as a transcription factor. In S. cerevisiae, scRAP1 directly binds to about 300 genomic loci in a sequence specific manner (Lieb et al., 2001). Here, it acts as a general transcription activator or repressor. In mice, studies have revealed that mammalian RAP1, similar to its yeast counterpart, also controls the expression of several genes (Martinez et al., 2010) (Teo et al., 2010) (Yeung et al., 2013).

In T. brucei, RAP1 was identified as an interacting partner of TbTRF (Li et al., 2005) (Yang et al., 2009). TbRAP1 is an essential protein and it seems to localizes to telomeres by interacting with TbTRF (Yang et al., 2009). TbRAP1 is required for proper silencing of ES linked VSGs to ensure monoallelic expression of VSGs (Yang et al., 2009). Depletion of TbRAP1 leads to expression of multiple

VSGs on T. brucei cell surface (Yang et al., 2009) (Pandya et al., 2013).

Moreover, depletion of TbRAP1 led to stronger derepression of telomere proximal genes than genes located more distal to telomeres (Yang et al., 2009)

(Pandya et al., 2013). These observations suggest that TbRAP1 promotes TPE similarly as its yeast and mammalian counterparts. However the mechanism of how TbRAP1 establishes/maintaines silencing is not fully understood. Potential roles of TbRAP1 in telomere length regulation and chromosome end protection 172

are not clear. In other organisms, RAP1 plays an important role in inhibiting

NHEJ. Since no homolog of ligase IV has been identified in T. brucei, it is proposed that NHEJ is either absent or unique in this ancient organism (Burton et al., 2007). Moreover, in vitro study confirmed that T. brucei cell extract has DNA end ligating activity that is independent of the Ku heterodimer (that is a major player in NHEJ), suggesting that NHEJ might be absent in T. brucei (Burton et al., 2007). However, T. brucei has a very efficient homologous repair pathway

(Conway et al., 2002b). Recent unpublished work from our lab suggests that depletion of TbRAP1 in BF cells results in an elevated VSG switching frequency, where most VSG switchers arose through DNA recombination events. Thus, like its mammalian homolog, TbRAP1 might prevent homologous recombination at telomeres. These observations indicate that RAP1 is involved in multiple telomeric and nontelomeric functions.

5.1.1 The domain structure of RAP1 homologues

RAP1 is one of the most conserved telomeric proteins (Chen et al., 2011).

It contains three conserved domains namely BRCT, Myb, and RCT (Figure.5.1).

BRCT (BRCA1 C-terminal homology domain) is located at the N-terminus of

RAP1s, Myb is a central domain that is involved in DNA binding (in budding yeast) or possibly protein interactions (in vertebrates), and RCT is RAP1 C- terminal domain (König et al., 1996) (Kabir et al., 2010). BRCT is a phosphopeptide-binding domain, most commonly present in proteins involved in cell cycle regulation and DNA damage response (Yu et al., 2003) (Manke et al.,

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2003). Although in RAP1, this domain is highly conserved from yeast to mammals, the exact functional relevance of this domain is not known.

Figure 5.1 Schematic drawing of domains of Rap1 homologs.

In S. cerevisiae and S. pombe, the BRCT domain seems to have no essential function as deletion of this domain has no effect on cell viability or known telomeric and non-telomeric functions of RAP1 (Graham et al., 1999)

(Fujita et al., 2012). In humans, The RAP1 BRCT domain is required for telomere length maintenance as overexpression of BRCT-deletion RAP1 mutant results in significantly longer telomeres with much narrower size distribution (Li and de

Lange, 2003). However, the molecular mechanism by which the RAP1 BRCT 174

domain controls the telomere length is largely unknown. Recently, an in vitro study analyzing the DNA binding ability of human RAP1 suggested that the

BRCT domain might be important for the DNA binding activity of RAP1 (Arat and

Griffith, 2012). In T. brucei, RAP1 contains a conserved BRCT domain towards the N-terminus and this domain along with N terminal region of RAP1 interacts with TbTRF (Yang et al., 2009). Any additional functions attributable to this domain of TbRAP1 remain to be determined.

The central region of RAP1 contains a Myb domain. The Myb domain is mostly present in transcription factors and is important for sequence specific binding of proteins to DNA (Biedenkapp et al., 1988). ScRap1 has an additional

Myb-like domain that cooperates with Myb to bind DNA in a sequence-specific manner (Conrad et al., 1990) (König et al., 1996). Apart from their DNA binding ability, The Myb and Myb-like domains in ScRap1 are thought to be important for repression of NHEJ at telomeres (Marcand et al., 2008). Despite the presence of a Myb and a Myb-like domain in RAP1 of T. brucei and S. pombe, these domains do not seem to play any role in DNA binding. In T. brucei, TRF recruits RAP1 to telomeres and in S. pombe, Taz1 performs a similar function (Kanoh and

Ishikawa, 2001) (Yang et al., 2009). In mammals, TRF2 recruits RAP1 to telomeres. Mammalian RAP1 contains a single Myb domain (Li et al., 2000).

Because usually at least two Myb domains are required to bind DNA, it was hypothesized that human RAP1 does not bind telomere DNA directly (Li et al.,

2000). This was supported by the fact that mutations that abolish interaction between hTRF2 and hRAP1 led to the removal of hRAP1 from telomeres (Chen 175

et al., 2011). However, a recent in vitro assay showed that human RAP1 can binds telomere DNA (Arat and Griffith, 2012).

RCT is a conserved protein interacting domain present at the C-terminal region of RAP1 homologues (Chen et al., 2011). The RCT domain in ScRap1 interacts with Sir proteins (Sir3 and Sir4), which are important for subtelomeric transcription silencing. ScRap1 directly recruits Sir3 and Sir4 to telomeres and establishes the heterochromatic structure by indirect recruitment of histone deacetylase Sir2 via its interaction with Sir4 (Moretti et al., 1994) (Moretti and

Shore, 2001) (Chen et al., 2011). The RCT domain of ScRap1 also recruits Rif1 and Rif2 to telomeres for telomere length regulation and telomere protection function (Hardy et al., 1992) (Wotton and Shore, 1997). In fission yeast, the

RAP1 RCT domain interacts with Taz1 and this interaction facilitates recruitment of RAP1 to the telomeres (Kanoh and Ishikawa, 2001) (Chen et al., 2011). In mammals RAP1 is recruited to telomeres through the interaction between the

RCT domain and TRF2 (Li et al., 2000) (Chen et al., 2011). T. brucei RAP1 has a conserved RCT domain but the functional relevance of this domain is still elusive.

In summary, RAP1 domains are conserved among different organisms but functions of these domains are diverse in different organisms.

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Figure 5.2 Schematic drawing of the LoxP system. The upper panel shows the LoxP site consisting of 13 bp inverted repeat sequences and a core of 8 bp nonpalindromic sequence (underlined). Cre mediates recombination between two direct repeats of LoxP. Following recombination, a covalently closed circle of DNA containing the sequence located between two LoxP sites is excised out of the chromosome and a product containing one repeat of LoxP is retained within the chromosome.

The role of TbRAP11 in silencing ES linked VSGs to ensure monoallelic expression is well established (Yang et al., 2009) (Pandya et al., 2013). Recent studies from our lab demonstrated that TbRAP1 knockdown leads to a change in chromatin structure independent of the transcription status (Pandya et al., 2013).

However, the molecular mechanism by which TbRAP1 regulates chromatin structure and establishes transcriptional silencing is not well known. Moreover, not much is known about potential roles of TbRAP1 in telomere maintenance.

RAP1 is reported to act as an adaptor protein that performs diverse functions by interacting with different factors (Kabir et al., 2010). Conserved domains of RAP1

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Figure 5.3 Generating a single floxed allele of TbRAP1. (a) Schematic representation of targeting of LoxP sites flanking the single TbRAP1 allele. (b) Southern blot analysis showing the insertion of both LoxP sites in a single allele of TbRAP1. Genomic DNA was digested with BamHI and XbaI. Southern blot was performed and probed with a TbRAP1 specific sequence (blue rectangle in (a)). BF/WT represents the size of WT fragment after restriction digestion. BF/RAP1Flox/+C1 and BF/RAP1Flox/+C2 represent the clones in which single TbRAP1 allele is floxed. (c) Severe growth defect was observed in the C1 and C2 clones containing LoxP and TK cassettes upon treatment with GCV while no growth defect was observed in the WT.

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act as interaction modules that can bind with various proteins, thereby performing different functions of RAP1. We sought out to establish a system to delineate the roles of these domains in mediating different functions of TbRAP1.

5.2 Results

5.2.1 The conditional knockout system

To study functions of various TbRAP1 domains, we established a conditional knockout system. We used a well-characterized Cre-loxP system to generate a conditional knockout cell line for TbRAP1. This system relies on the site-specific activity of Cre recombinase (cyclization recombinase). Cre is a site- specific DNA recombinase that can recognize the 34 bp loxP site and catalyze

DNA recombination between loxP repeats through both inter and intra molecular events (Abremski and Hoess, 1984). Cre mediated recombination between two direct loxP repeats results in excision of the DNA sequence within these repeats as covalently closed circle (Figure.5.2). Cre is highly efficient and requires no additional factors for its activity, thus making it an ideal enzyme for a variety of genetic manipulations. The Cre-loxP system has been previously used in T. brucei for generating gene-knockouts and is well accepted (Barrett et al., 2004)

(Scahill et al., 2008) (Kim et al., 2013). To generate a conditional knockout system for TbRAP1, we designed two targeting plasmids containing loxP sites.

These plasmids were designed to integrate within intergenic sequences that flank the TbRAP1 gene. A cassette containing loxP sequence followed by a sequence encoding for a fusion protein HYG-GFP-TK was targeted at the region

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immediately upstream of the TbRAP1 gene. The fusion protein contained a

Hygromycin B (HYG) resistance protein (for positive selection), a green fluorescent protein (GFP) and a thymidine kinase protein (TK) (for negative selection by Gancyclovir (GCV)). GFP served as an epitope to test the efficiency of excision of DNA sequence following the induction of Cre. Similarly, a cassette containing a sequence encoding for a fusion protein BSD-GFP-TK followed by the loxP site was targeted at the region immediately downstream of the TbRAP1 gene. The fusion protein contained a Blasticidin (BSD) resistance protein (for positive selection), GFP, and TK (Figure.5.3a). These loxP targeting constructs were transfected into T. brucei BF strain that constitutive expresses the T7 polymerase and the Tet repressor, allowing for inducible expression of genes driven by Tet-regulated promoters at later steps. Southern blot analysis was performed to confirm that both loxP sites flanked the same allele of TbRAP1

(Figure.5.3b). This strain was named BF/RAP1flox/+. It is resistant to HYG and

BSD, expresses the GFP epitope, and is sensitive to GCV due to the expression of TK. GCV sensitivity was confirmed by adding GCV to the growth medium. As shown in Figure.5.3c, addition of 30 ug/ml of GCV led to a severe growth arrest of BF/RAPflox/+ cells (both C1 and C2 clones) within 24 hrs, while no growth defect was observed in wild-type cells. Having confirmed the correct integration of loxP sites, we next introduced an inducible expression construct of Cre in these cells. We used pLEW100Cre-EP1 (Scahill et al., 2008) plasmid to target

Cre into one of the rDNA spacer regions. Expression of Cre is under the control

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Figure 5.4 Induction of Cre leads to the efficient removal of the floxed TbRAP1 allele (a) Schematic showing the genomic map of WT and floxed TbRAP1 alleles. (b) Growth curve of the cells containing floxed TbRAP1 allele in the Cre induced (green line, +Dox) and uninduced (blue line, -Dox) conditions. In the presence of selection drugs, Cre induction leads to severe growth arrest (green line, +Dox) while Cre induction in the absence of drug selection has no growth defect (pink line, +Dox). (c) Western blot analysis showing the efficient removal of HYG-GFP-TK and BSD-GFP-TK upon Cre induction. Whole cell lysate was prepared before and after 24 hours of Cre induction. Western blot was performed using α GFP antibody. (d) Southern blot analysis before and after induction of Cre shows efficient excision of floxed TbRAP1 allele. Left panel shows the schematic and expected band size of WT TbRAP1 allele (7.9kb) and of floxed TbRAP1 allele (4.7kb). B1and B2 are two independent clones.

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of a GPEET promoter and two tetracycline operators are positioned between the

GPEET promoter and the Cre-coding sequence, allowing tetracycline-dependent control of Cre expression. This construct is expected to have essentially no Cre expression under un-induced condition (Scahill et al., 2008). pLEW100Cre-EP1 also harbors a bleomycin resistance gene (BLE) driven by a T7 promoter to aid in the selection of transfected cells. This strain was named BF/RAP1flox/+/Cre and was cultured under continuous drug selection (HYG and BSD for loxP sites and

BLE for Cre).

5.2.2 Confirmation of conditional knockout system

To determine if induction of Cre expression leads to efficient removal of the TbRAP1 floxed allele, we induced Cre expression by adding doxycycline

(Dox). As shown in Figure.5.4b, no growth defect was observed upon induction of Cre expression (compare –Dox +BLE+HYG+BSD with +DOX +BLE). This suggests that the expression of Cre is not toxic to the cells. As the loxP sites flank the drug resistance gene-cassettes, induction of Cre should lead to the removal of these drug resistance genes. As shown in Figure.5.4b, in the presence of hygromycin and blasticidine, Cre induction leads to severe growth arrest (+DOX +BLE+HYG+BSD) due to toxic effects of these drugs. Western blot analysis confirmed the loss of GFP expression within 24 hrs upon Cre induction

(Figure.5.4c). Southern blot analysis was also performed to further confirm the efficient excision of the floxed TbRAP1 allele. As shown in Figure.5.4d, the floxed

TbRAP1 allele was completely lost upon induction of Cre. Next, we determined

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the mRNA level of TbRAP1 upon induction of Cre. Total RNA was extracted and the steady state level of TbRAP1 mRNA was determined by performing quantitative RT-PCR (qRT-PCR). As shown in Figure.5.5c, an ~50% reduction in

TbRAP1 mRNA level was observed after 24 hrs of Cre induction. This suggests that the floxed TbRAP1 allele was efficiently excised following the Cre expression. Western blot analysis also confirmed a decrease in the protein level of TbRAP1 (Figure.5.5b). Our previous studies have shown that TbRAP1 plays a very important role in mediating VSG silencing. As induction of Cre in

BF/RAP1flox/+/Cre cells results in the removal of one TbRAP1 allele, leaving only one functional TbRAP1 allele in the genome, we were curious to test if TbRAP1 is haploid sufficient for silencing VSGs. We therefore performed qRT-PCR to compare steady state mRNA levels of several silent BES-linked VSGs and metacyclic VSGs (mVSGs) in cre-induced and uninduced cells. As shown in

Figure.5.5d, we observed a 2 to 2.3-fold derepression of BES linked VSGs

(VSG3 and VSG11) and 1.5 to 2.5-fold derepression of mVSGs (mVSG397 and mVSG639). This result suggests that TbRAP1 is haploid-insufficient as deletion of one TbRAP1 allele results in mild derepression of silent VSGs.

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Figure 5.5 TbRAP1 is haploinsufficient (a) Graphical representation showing that floxed TbRAP1 allele is excised out of the chromosome upon induction of Cre. N1N2, B1B2 and M1M2 are three sets of primers specific to NT, BRCT and Myb domain sequences of TbRAP1 respectively. (b) Western blot analysis showing reduction in TbRAP1 protein level upon Cre induction. Whole cell lysate was prepared before (0 hours) and after (24hours) Cre induction. α RAP1 antibody was used to perform Western blot. EF-2 was used as a loading control. (c) qRT-PCR was performed using primer sets mentioned in (a). Fold change in mRNA level of TbRAP1 was calculated after 24 hours of Cre induction by normalizing against 0 hour of Cre induction. Samples were also normalized with tubulin which was used as an internal control. Steady state level of TbRAP1 was reduced by ~50% within 24 hours of Cre. (d) qRT-PCR was performed to compare the steady state level of BES-linked VSGs (6 and 11) and MES-linked mVSGs (397 and 639) before (0 hours) and after (24 hours) of Cre induction. Normalization was done as described for (c). Removal of floxed TbRAP1 allele leads to ~2.0 to 2.3 folds of derepression of BES-linked VSGs and ~1.5 to 2.5 folds of derepression of MES-linked mVSGs.

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5.2.3 Conditional knockout of TbRAP1 results in several hundred folds of derepression of silent VSGs

Previous studies have utilized the RNAi method to deplete TbRAP1. One of the major drawbacks of this approach is that RNAi-induced depletion of

TbRAP1 may not be complete. To compare the efficiency of conditional knockout and the RNAi, we replaced the unfloxed allele of TbRAP1 with a drug resistance gene Puromycin, resulting in the BF/RAP1flox/-/Cre train (Figure.5.6a). As shown in Figure.5.6b, induction of Cre led to a severe growth arrest by 24 hrs in

BF/RAP1flox/-/Cre cells but not in BF/RAP1flox/+/Cre cells lie. This is expected, as

TbRAP1 is essential for T. brucei survival and induction of Cre results in TbRAP1 null phenotype. Western blot analysis confirmed that most TbRAP1 protein is lost by 24 hrs of Cre induction (Figure.5.6c). To further confirm the efficiency of the condition knockout system, we compared the steady state mRNA level of

TbRAP1 in BF/RAP1flox/-/Cre and BF/RAP1flox/+/Cre cells before and after induction of Cre. As shown in Figure.5.6d, induction of Cre led to 80-94% reduction of TbRAP1 mRNA in BF/RAP1flox/-/Cre while BF/RAP1flox/+/Cre had a reduction of 50% in TbRAP1 mRNA. We next compared the steady state mRNA level of BES-linked VSGs (VSG6 and VSG11) and mVSG639 before and after

Cre induction. Several hundred-fold increase in VSG mRNA was observed after the induction of Cre in BF/RAP1flox/-/Cre cells (Figure.5.6e).

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Figure 5.6 Conditional knockout of TbRAP1 leads to derepression of BES-linked and MES-linked VSGs (a) WT TbRAP1 allele is replaced by Puromycin gene and floxed allele of TbRAP1 is removed using inducible Cre expression. N1N2 and M1M2 are the two sets of primers specific to NT and Myb domain sequences of TbRAP1 respectively. (b) Growth curve showing severe growth defect upon induction of Cre (+ Dox). Error bars represent the standard deviation obtained from three independent experiments. (c) Western blot analysis showing reduction in TbRAP1 protein level upon Cre induction. Whole cell lysate was prepared before (0 hours) and after (24hours) Cre induction. α RAP1 antibody was used to perform Western blot. EF-2 was used as a loading control. (d) qRT-PCR was performed using primer sets mentioned in (a). Fold change in mRNA level of TbRAP1 was calculated for control cells (BF/RAP1Flox/+) and conditional knockout cells (BF/RAP1Flox/-) as described in Figure.5.5c. Steady state level of TbRAP1 in control cells was reduced by ~50% within 24 hours of Cre induction while it was reduced by 80-94% in conditional knockout cells. (e) qRT-PCR was performed to compare the steady state level of BES-linked VSGs (6 and 11) and MES-linked mVSGs (639) before (0 hours) and after (24 hours) Cre induction. Normalization was done as described in Figure.5.5c. Conditional knockout of TbRAP1 leads to several hundred folds of derepression. 186

5.2.4 The TbRAP1 BRCT domain is required for VSG silencing

Studies in other organisms suggest that the ability of RAP1 to perform different roles depends on its ability to interact with various partners. Moreover, it is proposed that different domains of RAP1 interact with distinct factors and these interactions mediate different functions of RAP1. To understand the role of

Figure 5.7 Graphical representation of TbRAP1 mutants HA represents the FLAG-HA-HA epitope

each domain of TbRAP1, we generated TbRAP1 mutants with deletions of different domains (Figure.5.7) and introduced these mutants into the conditional

TbRAP1 knockout strain (BF/RAP1flox/+/Cre) to replace the remaining WT

TbRAP1 allele. The TbRAP1 BRCT domain is likely a protein interaction domain.

This domain is present in proteins that function in DNA damage repair and cell cycle regulation, such as BRCA1, 53BP1 etc. The BRCT domain interacts with

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other proteins by recognizing peptides containing a phospho-SXXF motif (X is any amino acids).

Figure 5.8 Deletion of the BRCT domain of TbRAP1 leads to a severe growth arrest (a) Schematic illustration of the genotype of TbRAP1 before and after induction of Cre. WT TbRAP1 allele is replaced by a mutant TbRAP1 allele bearing a deletion of BRCT domain and floxed allele of TbRAP1 is removed using inducible Cre expression. (b) Western blot analysis showing reduction in TbRAP1 protein level upon Cre induction in two independent clones, 4 and 5. Whole cell lysate was prepared before (0 hours) and after (24hours) Cre induction. α RAP1 antibody was used to detect endogenous TbRAP1 protein. α HA antibody was used to detect the level of mutant TbRAP1 protein bearing BRCT domain deletion. EF-2 was used as a loading control. (c) Growth curve showing severe growth arrest upon induction of Cre (+ Dox). Error bars represent the standard deviation obtained from three independent experiments from two independent clones, 4 and 5.

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The role of the BRCT domain in RAP1 homologues is not known. Using yeast two hybrid approach, it was shown that the N-terminal two thirds of

TbRAP1 including the BRCT domain interacts with TbTRF. However the functional relevance of this interaction is not clear. To better understand the importance of TbRAP1BRCT domain, we replaced the unfloxed TbRAP1 allele in

BF/RAP1flox/+/Cre with a TbRAP1 mutant allele whose BRCT domain was deleted

(resulting in strain BF/RAP1flox/ΔBRCT/Cre) (Figure.5.8a). Cre was induced to pop out the floxed TbRAP1 allele, leaving a mutant TbRAP1 with the BRCT domain deletion as the only TbRAP1 allele in the genome. Following the induction of Cre expression, effects of BRCT deletion on the growth of T. brucei were assessed.

As shown in Figure.5.8c, induction of Cre resulted in a severe growth arrest within 24 hrs (clone 4 and clone 5, + Dox). Western blot analysis was also performed to compare the TbRAP1 protein level before and after induction of Cre

(Figure.5.8b). The mutant TbRAP1 protein (ΔBRCT) contains a Flag-HA-HA

(F2H) tag, which allowed us to differentiate between the wild-type and the mutant proteins. As shown in Figure.5.8b, a decrease in the TbRAP1 protein level was observed when antibody against the endogenous TbRAP1 protein was used

(αRap1), indicating that the floxed TbRAP1 allele is lost upon induction of Cre. In contrast, no change in the level of TbRAP1 ΔBRCT mutant protein was detected when using an antibody specific for the HA epitope. To compare the steady state mRNA level of TbRAP1, we designed two sets of primers for qRT-PCR. The first set of primers (N1 and N2) was specific for an N terminal region of TbRAP1. This primer set will amplify both the floxed and the mutant alleles of TbRAP1. The 189

second set of primers (B1 and B2) was specific for the BRCT domain of TbRAP1 and thus will only amplify the floxed allele of TbRAP1 (Figure.5.9a). Following induction of Cre for 24 hrs, total RNA was collected from induced and uninduced cells and qRT-PCR was performed. As shown in Figure.5.9b, approximately 50% reduction in TbRAP1 mRNA was observed when N1 and N2 primers were used.

On the other hand, 70-80% reduction in TbRAP1 mRNA was observed when B1 and B2 primers were used. As N1 and N2 primers anneal to both floxed and mutant TbRAP1 alleles and as Cre induction results in the removal of the floxed

TbRAP1 allele, a maximum 50% decrease in the TbRAP1 mRNA is expected.

On the contrary, B1 and B2 primers only anneal to the floxed TbRAP1 allele that is removed upon Cre induction, a more severe decrease in the qRT-PCR result is expected. Derepression of silent VSGs is a well-characterized defect of TbRAP1 dysfunction. To examine the role that the TbRAP1 BRCT domain plays in silencing of VSGs, we performed qRT-PCR to compare steady state VSG mRNA levels. As shown in Figure.5.9 c & d, upon induction of Cre, we observed several hundred- to several thousand-fold of derepression in BES linked VSGs (VSG3 and VSG8 in clone 4, VSG8 and VSG11 in clone 5). This defect in VSG silencing is much more severe than the phenotypes observed in conditional deletion of

TbRAP1. These data suggest that the BRCT domain of TbRAP1 is critical for cell survival and VSG silencing.

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Figure 5.9 BRCT domain deletion of TbRAP1 leads to several thousand folds of derepression of BES-linked VSGs (a) Schematic of TbRAP1 genotype before and after induction of Cre. N1N2 and B1B2 are the two sets of primers specific to NT and BRCT domain sequences of TbRAP1 respectively. (b) qRT-PCR was performed using primer sets mentioned in (a). Fold change in mRNA level of TbRAP1 was calculated for two independent clones, 4 and 5. Steady state level of TbRAP1 was reduced by ~70 to 80% within 24 hours of Cre induction as detected by primer set B1B2. (e) qRT-PCR was performed to compare the steady state level of BES-linked VSGs (3, 6, 8 and 11) before (0 hours) and after (24 hours) Cre induction. Normalization was done as described in Figure.5.5c. Deletion of BRCT domain of TbRAP1 leads to several thousand folds of derepression.

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Figure 5.10 Deletion of the Myb domain of TbRAP1 leads to a severe growth arrest (a) Schematic illustration of the genotype of TbRAP1 before and after induction of Cre. WT TbRAP1 allele is replaced by a mutant TbRAP1 allele bearing a deletion of Myb domain and floxed allele of TbRAP1 is removed using inducible Cre expression. (b) Western blot analysis showing reduction in TbRAP1 protein level upon Cre induction. Whole cell lysate was prepared before (0 hours) and after (24 and 40 hours) Cre induction. α RAP1 antibody was used to detect endogenous TbRAP1 protein.α HA antibody was used to detect the level of mutant TbRAP1 protein bearing Myb domain deletion. EF-2 was used as a loading control. (c) Growth curve showing severe growth arrest upon induction of Cre (+ Dox).

5.2.5 The TbRAP1 Myb domain deletion leads to a null phenotype

Not much is known about the role of the Myb domain in RAP1 orthologs.

In budding yeast, RAP1 Myb domain is required for DNA binding (König et al.,

1996). However the function of this domain in other RAP1 homologues is not well uderstood. To examine the role of TbRAP1 Myb domain, the unfloxed WT

TbRAP1 allele in BF/RAP1flox/+/Cre cells was replaced with a mutant TbRAP1 192

allele lacking the Myb domain (ΔMyb) (BF/RAP1flox/ΔMyb/Cre) (Figure.5.10a.) The mutant ∆Myb TbRAP1 protein contains a F2H epitope at its N-terminus, which allowed us to differentiate between the wild type and the mutant proteins.

Western blot analysis performed using antibody against the HA epitope confirmed that the mutant TbRAP1 is expressed (Figure.5.10b αHA). To analyze the effect of TbRAP1∆Myb mutant, we induced the expression of Cre to remove the floxed WT TbRAP1 allele. Upon induction of Cre, a severe growth defect was observed, suggesting that the Myb domain is required for proper growth of T. brucei (Figure.5.10c). We confirmed the removal of the floxed TbRAP1 allele by performing western blot analysis. As shown in Figure.5.10b (αTbRAP1), induction of Cre led to a significant decrease of the wild-type TbRAP1 protein within 24 hrs while the protein level of mutant TbRAP1 (αHA) had no significant change. We next examined TbRAP1 mRNA level by performing qRT-PCR. Two sets of primers were again used to quantify TbRAP1 mRNA level (N1 and N2 that anneal to the beginning of the TbRAP1 gene, and M1 and M2 that anneal to the TbRAP1 region encoding the Myb domain) (Figure.5.11a). As shown in

Figure.5.11b, we observed a significant reduction of the steady state mRNA level of the wild-type TbRAP1 allele in two independent experiments upon Cre induction (about 65% within 32 hrs in experiment 1 and about 55% in experiment

2, compare M1 and M2 primer set values that only anneal with floxed allele).

Next, we examined the role of the Myb domain in silencing of VSGs. We compared steady state mRNA levels of several ES linked and metacyclic VSGs.

As shown in Figure.5.11c, mRNA levels of all BES linked VSGs (VSG 3, 6, 9, 193

and 11) were increased by several hundred-fold in the absence of the TbRAP1

Myb domain. Similarly, mRNA levels of mVSGs (mVSG 397 and mVSG639) were increased by 10 to several hundred folds in the absence of the TbRAP1

Myb domain. These results indicate that the Myb domain of TbRAP1 plays an important role in silencing ES-linked as well as metacyclic VSGs.

5.2.6 The Myb-like domain of TbRAP1 is essential for T. brucei survival.

In ScRap1, the Myb-like domain along with the Myb domain binds to specific DNA sequences. This domain is also present in RAP1 homologues of S. pombe and T. brucei, but not much is known about the functional importance of this domain in these organisms. To better understand the role of the Myb-like domain of TbRAP1, we transfected a mutant allele of TbRAP1 harboring the deletion of Myb-like domain (ΔMYBL) in BF/RAP1flox/+/Cre cells resulting in

BF/RAP1flox/ΔMYBL/Cre. We confirmed the expression of mutant TbRAP1 ∆MybL protein by performing western analysis using HA antibody (Figure.5.12c). Upon induction of Cre, TbRAP1 mRNA level was decreased by 40% (Figure.5.12d) within 24 hrs as assessed by q-RT-PCR using N1 and N2 primers. This suggests that the floxed TbRAP1 allele is removed efficiently. Induction of Cre led to a severe growth defect of BF/RAP1flox/ΔMYBL/Cre cells suggesting that the Myb-like domain is required for the survival of T. brucei (Figure.5.12b). We also examined the role of the Myb-like domain in VSG silencing by comparing steady state levels of VSG mRNA before and after induction of Cre. As shown in

Figure.5.12e, we observed approximately 20-30 fold increase in steady state

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level of mRNA of BES linked VSGs (VSG8 and VSG11) and approximately 18 fold increase in mVSG397 within 24 hrs of Cre induction.

5.2.7 The RCT domain is essential for TbRAP1 function.

The RCT domain of RAP1 is an evolutionarily conserved protein-protein interaction module. In S. pombe and mammalian cells, the RCT domain interacts with Taz1 and TRF2, respectively, while in S. cerevisiae, the RCT domain interacts with Rif1/2 and Sir3/4 proteins. In order to understand the role of

TbRAP1 RCT domain, we used our conditional knockout system. The RCT domain of TbRAP1 contains a putative nuclear localization signal (NLS). In order to facilitate the nuclear localization of TbRAP1 that lacks the RCT domain, we fused the NLS at the C-terminus of this TbRAP1 ΔRCT mutant (ΔRCT+NLS). We transfected a mutant TbRAP1 ∆RCT+NLS allele in BF/RAP1flox/+/Cre cells to generate BF/RAP1flox/ΔRCT+NLS/Cre (Figure.5.13a). Western analysis confirmed the expression of the mutant TbRAP1 protein (Figure.5.13c). As shown in

Figure.5.13b, addition of NLS to the TbRAP1 ΔRCT mutant enabled nuclear localization of this mutant. Next, we determined the effect of ∆RCT mutant on the function of TbRAP1. Induction of Cre led to a severe growth arrest in

BF/RAP1flox/ΔRCT+NLS/Cre cells (Figure.5.13d). Removal of the floxed WT TbRAP1 allele upon induction of Cre was confirmed by performing qRT-PCR. Steady state level of TbRAP1 mRNA was compared before and after 24 hrs of Cre induction.

As shown in Figure.5.13e, induction of Cre led to an approximately 40% reduction in total TbRAP1 mRNA as assessed by qRT-PCR using N1 and N2,

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Figure 5.11 Myb domain deletion of TbRAP1 leads to several hundred folds of derepression of BES-linked VSGs. (a) Schematic of TbRAP1 genotype before and after induction of Cre. N1N2 and M1M2 are the two sets of primers specific to NT and Myb domain sequences of TbRAP1 respectively. (b) qRT-PCR was performed using primer sets mentioned in (a). Fold change in mRNA level of TbRAP1 was calculated for two independent inductions. Steady state level of TbRAP1 was reduced by ~65% in experiment I and ~55% in experiment II as detected by primer set M1M2. (c) qRT-PCR was performed to compare the steady state level of BES- linked VSGs (3, 6, 9 and 11) and MES-linked mVSGs (3, 6, 397 and 639) before (0 hours) and after (24 and 36 hours) Cre induction. Normalization was done as described in Figure.5.5c. Two independent experiments were performed. Deletion of Myb domain of TbRAP1 leads to several hundred folds of derepression.

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Figure 5.12 Deletion of the Myb like domain of TbRAP1 leads to a severe growth arrest. (a) Schematic illustration of the genotype of TbRAP1 before and after induction of Cre. WT TbRAP1 allele is replaced by a mutant TbRAP1 allele bearing a deletion of Myb like domain and floxed allele of TbRAP1 is removed using inducible Cre expression. N1N2 and M1M2 are the two sets of primers specific to NT and Myb domain sequences of TbRAP1 respectively (b) Growth curve showing severe growth arrest upon induction of Cre (+Dox) in absence of Myb like domain. Error bars represent the standard deviation obtained from three independent experiments. (c) Western blot analysis was performed to confirm the expression of mutant TbRAP1 protein. Whole cell lysate was prepared and Western blot was performed using α RAP1 antibody to detect endogenous TbRAP1 protein and α HA antibody to detect the level of mutant TbRAP1 protein bearing Myb like domain deletion. EF-2 was used as a loading control. (d) qRT- PCR was performed using primer sets mentioned in (a). Fold change in mRNA level of TbRAP1 was calculated as described before. Steady state level of TbRAP1 was reduced by ~40% as detected by primer sets N1N2 and M1M2. (e) 197

qRT-PCR was performed to compare the steady state level of BES-linked VSGs (8 and 11) and MES-linked mVSGs 397 before (0 hours) and after (24 hours) Cre induction. Normalization was done as described in Figure.5.5c. Deletion of Myb like domain of TbRAP1 leads to mild derepression and M1 and M2 primers, indicating that the floxed WT allele is efficiently deleted.

We also analyzed the effect of TbRAP1 ∆RCT mutant on VSG silencing by determining the mRNA level of VSGs before and after induction of Cre inBF/RAP1flox/ΔRCT+NLS/Cre cells. As shown in Figure.5.13f, the Cre induction led to a derepression of the ES linked VSGs (VSG6 and VSG11) and mVSG639.

5.3 Discussion

Understanding the regulation of VSG silencing has been an area of active research in T. brucei. So far only a few factors have been identified that affect the

VSG silencing. TbRAP1 is by far the most important protein that regulates VSG silencing and thereby ensuring the monoallelic expression. However, the detailed mechanism by which TbRAP1 regulates VSG silencing is unknown. Studies of

RAP1 orthologs in other organisms have suggested that RAP1 is a multifunctional protein. These studies exhibit a common theme that RAP1 interacts with various proteins through its various conserved domains to perform specific functions. In T. brucei, RAP1 consists of all the conserved domains but how these domains modulate the function of TbRAP1 is not clear. In this study we established a conditional knockout system to study functional importance of

TbRAP1 domains. Our preliminary data suggest that this system is very efficient to study the functions of TbRAP1 domains.

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Figure 5.13 Deletion of the RCT domain of TbRAP1 leads to a TbRAP1 null phenotype (a) Schematic illustration of the genotype of TbRAP1 before and after induction of Cre. WT TbRAP1 allele is replaced by a mutant TbRAP1 allele bearing a deletion of RCT domain and carrying a NLS and floxed allele of TbRAP1 is removed using inducible Cre expression. N1N2 and M1M2 are the two sets of primers specific to NT and Myb domain sequences of TbRAP1 respectively (b) Immunocytology was performed to confirm the nuclear localization of RCT domain deleted and NLS tagged mutant of Tb RAP1. α HA antibody was used to stain for TbRAP1 and is shown in red channel, HYG-GFP-TK or BSD-GFP- TK are shown in green channel. Chromatin was stained with DAPI and is shown in blue channel. (c) Western blot analysis was performed to confirm the level of 199

TbRAP1 protein before and after Cre induction. Whole cell lysate was prepared and Western blot was performed using α RAP1 antibody to detect endogenous TbRAP1 protein and α HA antibody to detect the level of mutant TbRAP1 protein bearing RCT domain deletion. Tubulin was used as a loading control. (d) Growth curve showing severe growth arrest upon induction of Cre (+Dox) in absence of RCT domain. (e) qRT-PCR was performed using primer sets mentioned in (a). Fold change in mRNA level of TbRAP1 was calculated as described before. Steady state level of TbRAP1 was reduced by ~40% as detected by primer sets N1N2 and M1M2. (f) qRT-PCR was performed to compare the steady state level of BES-linked VSGs (6 and 11) and MES-linked mVSGs 639 before (0 hours) and after (24 hours) Cre induction. Normalization was done as described in Figure.5.5c. Deletion of RCT domain of TbRAP1 leads to derepression.

We employed the Cre-loxP system to generate strains of conditional knockout of TbRAP1. This system has several advantages over other genetic manipulations to study the function of essential proteins. First, induction of Cre results in removal of the floxed allele with high specificity and efficiency. This makes it easy to study the immediate effects of the loss of function phenotype of the target gene and avoid off target effects and incomplete depletion often associated with RNAi silencing. Second, conditional knockout yields high reproducibility. In our study, we used the same parent cell line (BF/RAP1flox/+/Cre) to study the effect of TbRAP1 mutants. This reduced the variation due to the different efficiencies with which the wild type RAP1 is depleted.

Our preliminary data provides some interesting insights into the function of

TbRAP1 domains. Upon deletion of the BRCT domain, we observed a higher level of derepression of VSGs than a simple conditional deletion of TbRAP1, suggesting that the BRCT domain may have a dominant negative effect. In humans, overexpression of the RAP1 mutant lacking BRCT resulted in

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elongation of telomeres (Li et al., 2000) (Li and de Lange, 2003). It was hypothesized that the BRCT domain may interact with factor(s) responsible for telomere length maintenance, and mutant RAP1 protein that lacks BRCT competes with the wild-type protein for telomere binding but fails to recruit factor(s) required for proper telomere length regulation and thus causes deregulation of telomere length homeostasis (Li and de Lange, 2003). In T. brucei, RAP1 interacts with TbTRF and this interaction presumably results in subsequent recruitment of TbRAP1 to the telomere. Yeast two hybrid analysis suggests that the BRCT domain may be important for TbRAP1/TbTRF interaction

(Yang et al., 2009). Hence, mutant TbRAP1 that lacks BRCT may fail to localize to the telomere. Although we do not have a direct evidence that BRCT deletion mutant of TbRAP1 is defective in telomere association, a high level of derepression of silent VSGs can be explained if BRCT deletion TbRAP1 mutant is no longer located at the telomere. When Cre is not induced, both mutant and wild-type TbRAP1 proteins are present in roughly equal amounts. Once Cre is induced, the floxed WT allele is removed, leaving only the mutant TbRAP1 allele.

This results in a situation similar to overexpression of mutant allele that results in titration of interacting factor(s). This speculation is supported by a study in budding yeast, overexpression of the N-terminal deletion mutant (ΔBB) that lacks

DNA binding ability results in telomere elongation by titration of Rif1 and Rif2 in the nucleoplasm(Conrad et al., 1990). In the future, overexpression of BRCT deletion mutant can be carried out to test whether there is any derepression of

VSGs. 201

5.4 Conclusion

In this study we aimed to identify separation of function mutants of

TbRAP1 by deleting different domains of RAP1 but all the tested deletion mutants resulted in severe growth arrest. This obscured us from understanding the mechanism by which TbRAP1 acts as a pleiotropic factor. As the derepression of silent VSGs is unlikely to be the cause of severe growth arrest, it is possible to screen for the mutants that are viable but are defective for VSG derepression. In the future, studies can be carried out with smaller truncations or even point mutations of TbRAP1 to identify separation of function mutants.

Conditional knockout system contains a selectable marker to select for the clones that lose the floxed WT allele. This makes this system ideal for high throughput screening for mutants that are able to sustain the cell viability in the absence of the wild-type allele. Moreover, in our study, we analyzed only one specific function of TbRAP1, which is silencing of VSGs. As RAP1 is a multifunctional telomeric protein, it is important to analyze the effects of deletions of different domains of RAP1 on telomere maintenance.

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6CHAPTER VI

Summary and Future Perspective

6.1 Telomerase regulation in T. brucei

Telomeres play a key role in regulating antigenic variation in T. brucei.

However not much is known about the maintenance of telomeres in this organism. The protein component of telomerase (TbTERT) was previously identified and its role in telomere maintenance was established (Dreesen et al.,

2005). In this study, we identified the other component of telomerase holoenzyme, telomerase RNA. We confirmed its association with telomerase and its role in telomere maintenance. In most of the eukaryotic systems, telomerase is recruited to the telomeres by factors that can interact with protein component of the telomerase or with the RNA component. As now both the components of telomerase have been identified in T. brucei, further studies can be carried out to identify the factors required for its regulation at telomeres.

The Ku heterodimer is a multifunctional complex. Famous for its role in NHEJ pathway, Ku also helps to recruit telomerase to the telomere ends. In budding yeast, it is proposed that Ku performs its telomere maintenance function by

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enhancing the nuclear localization of TLC1 (telomerase RNA) and also by promoting recruitment of EST2 (TERT) to the telomere. However, a recent study found that the tethering of EST2 to the telomeres or enhancing the nuclear localization of TLC1 does not compensate for Ku loss (Williams et al., 2014). In human cells, Ku is known to interact with hTR but the functional significance of this interaction is not known. Ku deletion in T. brucei has a similar phenotypic defect at telomeres as telomerase deletion, suggesting a conserved role of Ku in telomere maintenance. To better understand the role of Ku in telomerase recruitment and its activity we analyzed its interaction with TbTR.

To determine if TbTR interacts with TbKu in vivo we tagged TbKu80 with

GFP and performed RNA Immunoprecipitation using an antibody against GFP.

Co-Immunoprecipitated RNA was quantified by qRT-PCR. We used cells expression GFP tagged TbTERT as a positive control and cells expressing only

GFP as a negative control. As shown in Figure.6.1a, approximately 50-fold enrichment of TbTR was observed in TbTERT-GFP pull down when compared to the negative control. We also observed a significant enrichment of TbTR in GFP-

KU80 pull down experiment (Figure.6.1b). This is expected as TbTERT has a strong affinity for TbTR while the interaction between TbKu80 and TbTR might be much weaker and transient. We also observe a mild enrichment of tubulin RNA in

TbKu80 pull down, suggesting that TbKu80 may interact with other cellular RNAs the same as in yeast or the conditions we used for the experiment were not specific enough to exclude non-specific bindings.

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Figure 6.1 TbKU80 deletion leads to a decrease in telomerase activity (a) TbTERT and TbTR interact strongly. qRT PCR was performed to detect TbTR and tubulin (control) co-immunoprecipitated with TbTERT and TbKU80. Cell line expressing only GFP was used as negative control. Quantification of qRT PCR is shown. The average enrichment of pulldown sample over input (%) was calculated from three independent experiments. Error bars represent standard deviation (b) Same qRT PCR values from (a) are plotted for empty vector control and TbKU80 (c) TbKU80 deletion affects telomerase activity in vitro: 0.25ug of T. brucei cell extract was used in each reaction and loaded on to the gel as labeled. Blue line on the left marks the decrease in activity detected in TbKU80 null cells. Three independent experiments confirmed the similar decrease in activity 205

More careful analysis will be carried out in the future to validate this interaction.

Our current observations suggest that the function of Ku is to recruit telomerase to the telomeres and it is not known whether Ku plays any role in telomerase activity regulation. To test if TbKu may regulate telomerase activity per se, we performed the TRAP assay to compare the activity of telomerase in the presence

Figure 6.2 TbKU80 deletion does not affect TbTR levels Northern blot analysis was performed to compare TbTR level in wild type (WT/PF) and TbKU80 mutants.In KU80+/- cell line one allele of TbKU80 is replaced by drug marker. In KU80-/- cell lines both the alleles of TbKU80 are replaced by drug markers resulting in TbKU80 null cell lines. C1 and C2 are two indepentdent clones Top panel hybridized with TbTR specific probe. Bottom rRNA species shown as loading control.

and absence of TbKu. Cell lysates of wild type (PF/WT) and TbKu80 (PF/KU80-/-) null cells were used to perform TRAP assay. Surprisingly, we observed a reduction in the telomerase activity in the absence of TbKu80 (Figure.6.1c). We also performed a Northern analysis to determine whether the reduction in telomerase activity was due to a decrease in the TbTR level and found that TbTR level is not affected by TbKu80 deletion (Figure.6.2). This is very different from previous observations in yeasts and mammalian cells, where Ku is only 206

implicated in the recruitment of telomerase. Therefore, it is possible that Ku is required for both proper telomerase recruitment and stimulation of telomerase activity. This hypothesis is supported by a recent finding suggesting that in the budding yeast, Ku is not only required for telomerase recruitment but also plays an important role in stabilization and activation of telomerase holoenzyme

(Williams et al., 2014). Moreover, in humans, Ku heterodimer interacts with protein component of telomerase (hTERT) and this interaction is independent of telomerase RNA (Chai et al., 2002). However the functional significance of this interaction is unclear. In the future, potential interaction between TbTERT and

TbKu will be determined. This will help us to understand the function of TbKu in telomerase recruitment and activity stimulation. So far we have used the TRAP assay to determine the activity of telomerase. This assay may underestimate the differences in the processivity of telomerase due to the amplification of product signal in the PCR step. In the future, another assay called as telomerase primer extension assay will be utilized to visualize the actual difference in telomerase processivity, as it is a more sensitive assay to measure subtle differences in the telomerase processivity.

6.2 Functions of TbTR domains

Despite the remarkable diversity in TR size and sequence in different organisms, all TRs including TbTR share similar core domains such as the pseudoknot and TBE. However the exact functions of these domains are not completely understood. Therefore structure function analysis of TbTR will be

207

carried out by deleting or mutating conserved domains of TbTR and analyzing the effect of these deletions on the telomerase complex formation, its recruitment to the telomeres and its activity. T. brucei is a very good model organism to perform such kind of structure function analysis, as telomerase is not essential in

T. brucei and telomerase activity is readily detectable both in vivo and in vitro.

These studies can provide insights to the role that TbTR plays in telomerase activity and will help to elucidate the process by which telomerase is recruited to the telomeres. Results from these experiments performed in T. brucei are easily translatable to human telomerase complex as the structure and function of telomerase complex is largely conserved across these species.

6.3 TbTR as a potential drug target

In T. brucei, deletion of telomerase results in a low rate of telomere shortening. However there is no adverse effect on its growth (Dreesen et al.,

2005). Based on these results, it was proposed that the telomerase could not act as a potential drug target against T. brucei. However, a recent study argued against it by showing that the lack of telomerase activity in T. brucei results in an elevated rate of VSG switching due shortening of telomeres (Hovel-Miner et al.,

2012). The uncontrolled switching of antigenic variants could make the parasite vulnerable to the host immune system and thus can be used as a potential target against the parasite. Moreover, in our study, we noticed that a change in the template sequence of an ectopically expressing TbTR results in a growth defect in TbTR null cells. Based on these observations, we hypothesized that a change

208

in the telomeric sequence of T. brucei due to the template alteration in TbTR results in the loss of telomere protection function. To determine if the ectopic expression of a mutant TbTR with an altered template could also induce growth defect in the wild type TbTR background, we transfected a mutant TbTR with an altered template (5’ CCCTAACCC 3’ wild type template was altered to 5’

GGCTAACCC 3’) in the wild type strain. As shown in Figure.6.3, we observed a severe growth arrest in two independent cell lines harboring altered TbTR. These studies can be further extended to validate TbTR as a potential drug target.

Figure 6.3 Mutation in TbTR template leads to severe growth defects Mutated TbTR was ectopically expressed in wild type bloodstream form cells. Induction of the mutant TbTR expression results in severe growth defect in two idependent clones. Standard deviation between experiments is shown as error bar.

209

Telomeric proteins bind to telomeres in a sequence specific manner.

Mutations in the TbTR template will result in an altered telomeric sequence. This altered telomeric sequence may lead to the inability of the telomeric proteins to bind to the telomeres. Although never tested, it is generally assumed that a limited number of mutations in the sequence of telomeres is sufficient to alter the structure and function of telomeres (Guiducci et al., 2001). Intriguingly, cells that maintain telomere length independent of telomerase are also sensitive to the mutations in TR template. Although speculative, it is possible that the presence of an active telomerase complex has the ability to inhibit telomerase independent telomere maintenance mechanisms. T. brucei is a unique model to understand the effect of altered TR template on the telomere maintenance and protection. As the structure of the DNA binding domain of TbTRF was recently resolved (Jehi et. al., 2014b), TbTR mutations can be designed that will have a minimal effect on the binding of TbTRF to telomeres. These studies will help us to decipher if the deleterious effect of mutations in telomeric sequence is due to the inability of the telomeric proteins to bind to the telomeres or if the telomeric sequence is important by itself. Moreover, mutations in TbTR can be designed in a way that they only affect the activity of telomerase without affecting its recruitment to the telomeres. This study will help us to understand if the telomerase plays any role in the inhibition of telomerase independent telomere maintenance pathway.

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6.4 G-overhang structure maintenance

Maintenance of the proper G-overhang structure is very crucial for telomere functions. In chapter IV, I discussed various tools that can be used to analyze G-overhang structure in T. brucei. By using these tools, we have also determined the terminal nucleotide of both the G-rich and C-rich strand of the telomere in T. brucei. We also studied the effects of deletions of several telomeric proteins on G-overhang maintenance.

One of the obvious unanswered questions raised by our study is: what is the mechanism of end specification and why it is important? Our study has provided the necessary tools to determine the nature of processing that happens at the ends of telomeres. Future studies will be designed to identify the nucleases or the regulatory proteins that are involved in specifying the terminal nucleotide of G-overhang. In human cells, C-rich strand specification requires

POT1, however the mechanism by which POT1 specifies the terminal nucleotide is not known. As genome-wide RNAi library is available for T. brucei, a screen can be carried out to identify potential factors that function in C-rich and G-rich terminal nucleotide specification.

In most organisms, G-overhang length is cell cycle regulated. In budding yeast, G-overhang length is increased in late S and G2 followed by a decrease in late G2 and G1 (Bonetti et al., 2009) (Soudet et al., 2014). Similarly, in humans,

G-overhang undergoes dynamic changes during the cell cycle (Chai et al., 2006)

(Chow et al., 2012). Using the G-overhang assays described in chapter IV, such

211

studies can be carried out in T. brucei to determine if its G-overhang also undergoes cell cycle regulated changes.

At this point, we are not sure about the exact length of T. brucei G-overhang.

Knowledge of G-overhang length is very crucial for our understanding of G- overhang maintenance. The LMPE assay described in chapter IV has been successfully used in Tetrahymena to measure the G-overhang length (Jacob et al., 2001). In Tetrahymena, \ telomeres are short and have a uniform length of about 300 bp. Uniformity of telomere length facilitates structural analysis of the

G-overhang. On the contrary, telomeres in T. brucei are long and are very heterogeneous in length. This could be one of the reasons that the LMPE assay is not very efficient in detecting the G-overhang length in T. brucei. In the future, other methods can be employed to validate our results obtained by LMPE assay.

One recent method used to detect blunt ended telomeres in plant cells can be modified to measure the G-overhang length in T. brucei. This assay relies on ligation of a DNA hairpin to the end of the telomere and thus covalently linking C- rich and G-rich complementary strands (Kazda et al., 2012). This results in the electrophoretic mobility shift of telomeres to double-sized fragments under denaturing conditions. According to the LMPE results, telomere overhangs in wild type T. brucei cells should be from six to twelve nucleotides long. DNA hairpins with six to twelve nucleotides long 3’ single strand protrusions can be designed and their ligation to telomere ends can be analyzed by running denaturing gels.

Collectively, data presented in this thesis provides new insights into the mechanism of telomere maintenance in T. brucei. Using a combination of 212

molecular and genetic tools, I discovered the roles of different factors in regulating telomere G-overhang in T. brucei. I optimized new methods to examine the dynamics of the telomere ends in T. brucei, which can be used to identify factors required for proper telomere maintenance. Understanding the processing of telomere ends will enhance our knowledge about telomere replication and telomere protection in T. brucei. This knowledge will help us to use telomere biology to eradicate this parasite.

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