CHARACTERIZATION OF AN EB1 HOMOLOGUE IN TRYPANOSOMA BRUCEI

END-BINDING PROTEIN 1 (EB1): CHARACTERIZATION OF AN EB1 HOMOLOGUE IN TRYPANOSOMA BRUCEI LIM LI FERN (B.Sc.(Hons.), NTU) A THESIS SUBMITTED FOR THE DEGREE OF MASTER OF SCIENCE DEPARTMENT OF BIOLOGICAL SCIENCES NATIONAL UNIVERSITY OF SINGAPORE 2012 DECLARATION I hereby declare that the thesis is my original work and it has been written by me in its entirety. I have duly acknowledged all the sources of information which have been used in this thesis. This thesis has also not been submitted for any degree in any university previously. ____________________________ LIM LI FERN 14 AUGUST 2012 ACKNOWLEDGEMENTS Nani gigantum humeris insidentes. The 12th-century Latin quotation attributed to Bernard of Chartres and made famous by Isaac Newton ("If I have seen further, it is by standing on the shoulders of giants,") recognizes two things: one, a person's achievements is never merely the sum of the work of his own hands; two, a person can never be an island who has achieved anything worth recording. Both truths have been self-evident to me in my time working on this thesis, and it is only right that these giants, in every sense of the word, be gratefully given their due recognition in enabling me to put together this humble report. Dr. Cynthia He deserves more than just a word of thanks or a grateful mention; she deserves a medal for her unending patience, remarkable insight, and generous guidance throughout my years of working under her. No graduate student could claim to have a better supervisor; her steady direction in the early days of raw unfocused enthusiasm, her infectious excitement in the (oh so frequent!) periods of experimental doldrums, her unwavering support through the final days of rushing the last experiments while painfully hammering out the thesis word by word -this report is a testament to her gift of supervision, and I only wish that I could have done more to render to her the credit she deserves. My awesome colleagues, past and present, made the lab a wonderful place to be in even on the occasional Saturday (and Sunday) evening; there is no way to adequately thank all these i fantastic people who have laughed, cried, eaten chocolate after chocolate and argued pseudo-philosophy with me throughout these years -Dr. Li, Dr. Zhou and Zhang Yu have on numerous occasions advised me on the finer points of experimental techniques, Sun Ying mentored me when I first joined the lab as an absolute greenhorn and Wang Min was my guide in preparing for motherhood as she was in operating the Guava cell counting machine. Dulani mothered me throughout her stay in the lab, Shima made my day, every day, by winking her welcome and helping out with my cells when I was away, while Foong Mei and Shen Qian have made the lab a much cheerier place with their ready smiles and helping hands. (Shen Qian, keep drawing those awesome pictures!) But it is really this special group of people -Omar, Ladan and Anaïs -who have been the best friends any person could ask for. You guys widened my perspectives, challenged my assumptions, went the extra mile then stuck around for the next two (under the pretext of having to stay late anyway), turned boring minipreps and IFs into random philosophical battlegrounds and made lunchtimes into so much more than just shovelling food. If I made it through the course alive and sane, it is really because you guys were around; thank you, from the bottom of my heart. I also need to extend a grateful word of thanks to Wang Chao from Dr. Adam Yuan's lab at the NUS Centre for BioImaging Sciences for his assistance in purifying His-EB1 protein for antibody generation; his expertise was invaluable, and his kindness in answering my generally numerous and sometimes inane questions is not soon forgotten. I would also like to render my appreciation to the National University of Singapore for providing me with monetary support and the opportunity to experience a season of exploration and research. ii My deepest thanks must now be expressed to my family, who has been a constant source of every good thing in every possible sense of the word; it will truly have not been possible without you all. Jauh di mata, dekat di hati, as the Malay saying goes, but thank God for technology, and thank you for being so understanding all these years when I forgot birthdays and went AWOL for months; I've been the recipient of so much grace it's bordering on ridiculous (not that I'm complaining!) and I only hope that this work brings you joy, as it is but a testament to your unwavering love and care. Last but far from least, I gratefully thank my husband, Yong Jie, for his constant love and support throughout the years. We've progressed from being attached to being engaged, and from being engaged to being married, and from being married to being expectant parents... all within the time it took to finish this thesis. And what a ride it's been! Thank you for the journey, and thank you for being around. This humble work is dedicated to you. Fern 13 August 2012 iii TABLE OF CONTENTS ACKNOWLEDGEMENTS i TABLE OF CONTENTS iv SUMMARY vi LIST OF TABLES 1 LIST OF FIGURES 2 LIST OF ABBREVIATIONS 4 CHAPTER 1. Introduction 6 1.1 6 1.2 1.3 An overview of Trypanosoma brucei 1.1.1 T. brucei: Ecological, economic and political impact 6 1.1.2 T. brucei life cycle 6 1.1.3 T. brucei as a model organism 8 1.1.4 Overview on the major ultrastructure features 8 1.1.5 T. brucei cell division 10 An overview of End-binding protein 1 (EB1) 14 1.2.1 EB1 homologues 14 1.2.2 EB1 domain organization 14 1.2.3 EB1 cellular localization 16 1.2.4 EB1 as a keystone +TIP protein 16 1.2.5 Role of EB1 in mitosis 17 1.2.6 Putative mechanisms of EB1 cellular interaction 18 Why study EB1 in T. brucei? 20 CHAPTER 2. Materials and Methods 23 2.1 Molecular cloning 23 2.2 Cell lines, cultivation conditions and plasmid transfection 24 2.3 Clonal selection of stable transformants by limiting dilution 25 2.4 RNAi assay 25 2.5 Anti-TbEB1 antibody 25 2.6 Affinity selection of anti-TbEB1 polyclonal antibody 26 iv 2.7 Immunofluorescence microscopy 27 2.8 Immunoblot analysis 28 CHAPTER 3. Results 3.1 31 T. brucei putative EB1: establishing sequence homology and functional conservation 31 3.1.1 Bioinformatics 31 3.1.2 Establishing cellular localization of TbEB1 39 3.2 Tracking TbEB1 cellular localization throughout the T. brucei cell cycle 42 3.3 Functional study on TbEB1 44 3.4 Production and characterization of anti-TbEB1 antibody 48 3.4.1 Immunoblot analysis 49 3.4.2 Immunofluorescence assay 50 CHAPTER 4. Discussion 54 4.1 TbEB1: a putative EB1 homologue? 54 4.2 Tracking TbEB1 localization throughout the cell cycle 57 4.3 Characterizing the anti-TbEB1 antibody 58 4.3 Functional study of EB1 62 CHAPTER 5. Conclusion and future direction 63 BIBLIOGRAPHY 66 v SUMMARY The African trypanosome Trypanosoma brucei is a protozoan parasite that causes human African trypanosomiasis in 36 countries spanning sub-Saharan Africa -a major cause of human mortality as well as a major barrier to sustainable economic growth in these primarily agrarian societies. T. brucei relies primarily on an extended microtubule-based cytoskeletal network to define cell shape and regulate its cellular processes, with a significantly reduced dependency on other traditional elements of the eukaryotic cytoskeleton. Indeed, actin depletion is non-lethal in procyclic trypanosome cells, and there is no known trypanosome homologue of intermediate filaments. Given the parasite's reliance on its microtubule-based network, it was a natural step in the same direction to search for proteins that regulate microtubule dynamics throughout the trypanosomal cell cycle. Such a protein was already well-known in many eukaryotes, spanning organisms as evolutionarily diverse as yeast, sea urchins, plants and humans. This conservation of function argued for a fundamentally important role for End-Binding Protein 1 (EB1); indeed, EB1 has gathered recognition as a master regulator of microtubule plus-end dynamics in light of its independent localization and +TIP recruitment to the growing tips of microtubules. In terms of domain organization, EB1 comprises only two major domains connected by a flexible, poorly-conserved linker -an amino-terminus Calponin homology (CH) domain crucial for microtubule-binding, and a carboxyl-terminus EB1-like homology (EBH) domain which mediates interaction with various +TIPs. Interestingly, there has been only one study on EB1 conducted thus far on a protozoan parasite and none at all in a trypanosome system. This study aimed to identify and briefly vi characterize a putative EB1 homologue in T. brucei. Sequence alignment of the putative trypanosome EB1 homologue TbEB1 against established EB1 homologues revealed strong CH and EBH domain sequence conservation despite poor overall sequence homology. TbEB1 also seemed to retain traditional EB1 localization to microtubule plus-ends; when attached to a fluorescent tag, this resulted in a distinct fluorescence signal at the posterior tip of the cell body. To facilitate further study on TbEB1 function and localization, an anti-TbEB1 antibody was raised in rabbit and affinity-purified with His-EB1 protein before use in subsequent immunofluorescence assays and immunoblot analysis. Anti-TbEB1 antibody staining revealed two specific TbEB1 localization sites -a signal at the cell posterior tip that elongated towards the cell anterior as the cell cycle progressed, and a second localization that closely shadowed the nascent FAZ structure but not the older, existing FAZ. Both sites appeared to be temporally-regulated and closely associated with cell cycle progress, resulting in a unique localization pattern. Attempts were also made to characterize TbEB1 function via EB1-RNAi induction; however, all attempts at completely depleting TbEB1 has to date been unsuccessful. Partial TbEB1 depletion in a YFP-EB1 over-expression background exerted no adverse effect on cell morphology nor on cell fitness, as evidenced by the comparative growth curve plotted against control cells. This may be due TbEB1's ability to function at low cellular levels, but this observation warrants further investigation, especially since a previous study utilizing high-throughput screening indicated that TbEB1 RNAi-mediated depletion resulted in a significant loss of fitness. vii LIST OF TABLES TABLE 1. List of plasmids used in study. 29 Table 2. List of antibodies used in study. 29 TABLE 3. List of constructs and primers used in this study. 30 1 LIST OF FIGURES FIGURE 1. Cartoon representation of the major cell cycle stages in T. brucei. 13 FIGURE 2. Schematic diagram of human EB1 domain organization. 15 FIGURE 3. Schematic diagram of the major EB1 domains in T. brucei. 31 FIGURE 4. Sequence alignment of full length EB1 homologues from different species. 33 FIGURE 5a. Sequence alignment of the Calponin Homology (CH) domains. 35 FIGURE 5b. Sequence alignment of the EB-like homology (EBH) domains. 35 FIGURE 6. Sequence alignment of full length putative trypanosomatid EB1 homologues. 37 FIGURE 7a. Sequence alignment of the trypanosomatid putative EB1 Calponin Homology (CH) domains. 38 FIGURE 7b. Sequence alignment of the trypanosomatid putative EB-like homology (EBH) domains. 38 FIGURE 8. TbEB1 localized to the posterior tip of the cell. 39 FIGURE 9. Co-staining YFP-EB1 with other cellular markers affirmed TbEB1 localization at microtubule plus ends. 41 FIGURE 10. TbEB1 localization at the posterior end of T. brucei exhibited a temporal modulation that correlated closely with cell cycle progress. 43 FIGURE 11. Immunoblot analysis of EB1 RNAi up to 5 days post-induction. 45 FIGURE 12. Growth curve of TbEB1-RNAi induced cultures. 45 FIGURE 13. Immunofluorescence analysis of EB1 RNAi induction. 46 FIGURE 14. Anti-TbEB1 immune serum is non-specific in its detection of TbEB1. 49 2 FIGURE 15. Western blot analysis of 29.13 (control) cells and PXS2YFPEB1 cells using affinity-purified anti-TbEB1. 50 FIGURE 16. Co-staining YTAT cells with anti-TbEB1 and YL1/2 antibody confirmed a temporally-modulated EB1 localization at the posterior tip of the cell body. 51 FIGURE 17. Co-staining YTAT cells with anti-TbEB1 and FAZ/flagellum markers confirmed a temporally-sensitive labelling pattern which closely traced the nascent FAZ structure. 53 FIGURE 18. Detergent treatment resulted in punctate-like EB1 localization throughout the cell body. 57 FIGURE 19. Comparative fluorescence labelling of EB1 in a YFP-EB1 over-expressing cell line confirmed that the anti-TbEB1 antibody labelling pattern was identical to YFP-EB1 localization. 61 FIGURE 20. Anti-TbEB1 antibody labelling 24 hours post RNAi induction in a YFP-EB1 over-expression background. 61 3 LIST OF ABBREVIATIONS +TIP plus-end tracking protein aa amino acids ABS actin binding site APC adenomatous polyposis coli CAP-Gly glycine-rich cytoskeleton-associated protein CH calponin homology (domain) DAPI 4, 6-diamidino-2-phenylindole EB end-binding protein EBH end-binding homology (domain) FAZ flagellum attachment zone HAT human African trypanosomiasis IPTG isopropyl β-D-1-thiogalactopyranoside LB Luria broth MAP microtubule associated protein MT microtubule MtQ FAZ microtubule quartet PCR polymerase chain reaction RNAi RNA interference 4 PVDF polyvinylidene difluoride SDS-PAGE sodium dodecyl sulphate polyacrylamide gel electrophoresis TBST Tris-buffered saline - Tween 20 buffer 5 INTRODUCTION 1.1 An overview of Trypanosoma brucei 1.1.1 T. brucei: Ecological, Economic and Political impact The African trypanosome, Trypanosoma brucei, is the protozoan parasite responsible for the African sleeping sickness in 36 countries of sub-Saharan Africa, many of which fall in the category of the poorest developing nations in the world. Many affected populations live beyond the reach of accessible health services in areas where health systems are either weakened or non-existent due to political upheaval and rampant poverty -a contributing factor to the alarming mortality rate of this tropical disease, already fatal in the absence of treatment. Sleeping sickness, clinically known as human African trypanosomiasis (HAT), leaves a devastating impact upon the socio-economic profiles of these communities; most recent conservative estimates place the number of new cases at 30,000 yearly, although during epidemic periods sleeping sickness surpassed even HIV/AIDS as the greatest cause of mortality in several villages in the Democratic Republic of Congo, Angola and Southern Sudan (http://www.who.int/en/). A sub-species, T. brucei brucei, has also been shown to infect cattle and game animals with the disease 'nagana', curtailing agricultural progress and thereby reinforcing persisting poverty in afflicted areas (Simarro et al., 2008). There is currently no vaccine available and the four existing drug treatments are old (Suramin, a primary treatment for acute human trypanosomiasis, was discovered in 1917 and patented in 1924), difficult to apply in the field and have toxic side effects (Brun et al., 2010) . 1.1.2 T. brucei life cycle The trypanosome shuttles between mammalian hosts via a specific arthropod vector, the tsetse fly (Glossina spp) which is found only in sub-Saharan Africa. A fly is infected when it 6 takes a blood meal on an animal or human harbouring the human-pathogenic parasites. The parasite then proceeds to establishes itself in the fly midgut, proliferating and transitioning through several intermediate stages (in strict chronological order) in different locations before transforming into the infectious metacyclic stage in the salivary glands of the fly (Roditi and Lehane, 2008; Vickerman et al., 1988). This process necessitates highly- coordinated modulation of many basic biological processes (Fenn and Matthews, 2007), which suggests that trypanosomes are not only capable of adapting to rapidly changing environments, but that they also possess the capacity for rigorously programmed differentiation. Once the trypanosomes gain entry into the bloodstream of a new mammalian host, they proliferate as morphologically slender forms, which later give rise to stumpy, non-proliferative forms as parasite numbers increase (Matthews et al., 2004). This not only limits parasite density, thereby prolonging host survival (and therefore increasing the probability of disease transmission); interestingly, it also results in a uniform cell cycle arrest of stumpy forms in G1 phase (Shapiro et al., 1984), a process that ensures that reentry into the cell cycle is coordinated with the morphological changes that occur upon parasite retransmission into the tsetse vector (Matthews and Gull, 1994; Vassella et al., 1997; Ziegelbauer et al., 1990). This is of particular importance because the successful completion of the parasite procyclic (or insect-form) cell cycle relies on correct organelle positioning (Matthews, 2005). Indeed, since trypanosomes morph into at least five morphologically distinct cell types throughout its transition from vector to host (Sharma et al., 2009; Van Den Abbeele et al., 1999; Vickerman, 1985) while its microtubule cytoskeleton remains largely intact throughout the process, accurate spatial and temporal duplication and segregation of its many single-copy organelles is paramount to the survival of the parasite (Sherwin and Gull, 1989b). 7 1.1.3 T. brucei as a model organism The species most used in laboratory studies to date is T. b. brucei -an animal-infectious species, although it is not pathogenic for humans. Studies thus far have focused on the procyclic and bloodstream form of the parasite, the two proliferative stages, mainly because these stages are readily cultured in vitro (Gull, 1999). Several other factors lend themselves to the recommendation of this ancient eukaryote as an excellent model for addressing fundamental biological questions of broad interest and applicability, not least the fact that parasite is genetically tractable -targeted gene knockouts via homologous recombination, tetracycline-inducible ectopic gene expression of recombinant proteins and interference RNA (RNAi) as well as systems for forward genetics (Cross, 2001; Kelly et al., 2007; Meissner et al., 2007; Motyka and Englund, 2004) have since become routine. Reverse genetics and post-genomic work has also been further expedited by the release of the complete genome sequence in 2005 (Aslett et al., 2010; Berriman et al., 2005), while production of large scale RNAi libraries have been efficient and informative launching pads for the study of potentially interesting genes that have hitherto been overlooked (Alsford et al., 2011; Morris et al., 2002; Schumann Burkard et al., 2011). 1.1.4 Overview on the major ultrastructure features The African trypanosoma has a slender, elongated shape measuring about 15µm in length and 8µm at its widest girth. It possesses a single flagellum that propels the parasite forward, thus establishing the anterior-posterior axis of the cell. The flagellum, comprising a canonical 9+2 microtubule axoneme and a fibrillar structure known as the paraflagellar rod (PFR), is laterally attached to the cell body in a left-handed helix, beginning from where it exits the cell body via the flagella pocket near the posterior end of the cell along to the anterior 8 (Sherwin and Gull, 1989a). This characteristic, polarized shape, which remains intact throughout much of the cell cycle, is defined by a highly stable, highly cross-linked and intrinsically polarized sub-pellicular microtubule cytoskeleton (Angelopoulos, 1970). The microtubules are equally spaced (18-22 nm) and are uniformly arrayed with their plus ends at the posterior end of the cell (Robinson et al., 1995), with the exception of a microtubule quartet (MtQ), which is part of a specialized ultrastructure known as the flagellum attachment zone (FAZ). The FAZ structure, which undergirds and tethers the flagellum along most of the length of the cell body, is composed of a filament structure which connects the cell body with the PFR in the flagellum, and the specialized MtQ, which originates close to the basal bodies and thus possesses a polarity opposite to that of the microtubules in the sub-pellicular corset (Robinson et al., 1995; Sherwin and Gull, 1989a; Vaughan et al., 2008). The FAZ thus forms a "seam" in the microtubule corset, and from observation of procyclic cells in culture, it is believed to define the axis and direction of the cleavage furrow during cytokinesis (Robinson et al., 1995). The FAZ has also been proposed to control basal body and flagellar pocket positioning (Absalon et al., 2007; Bonhivers et al., 2008a). T. brucei also possesses a single copy of many organelles such as the mitochondrion and Golgi which are precisely positioned within the microtubule corset, resulting in a highly reproducible and polarized cell. They have been shown to be generally concentrated between the posterior end and centre of the cell. Many of them also physically tethered together -the kinetoplast (a mass of catenated DNA which forms the mitochondrial genome) is physically connected to the proximal end of the two basal bodies (Ogbadoyi et al., 2003; Robinson and Gull, 1991), while the mature basal body subtends the single flagellum, which exits the cell via the flagellar pocket (Lacomble et al., 2010). Since the flagellum is tethered to the cell body via the FAZ, which has also been shown to be in close contact with the 9 flagellar pocket (Lacomble et al., 2009), it is not surprising that correct organelle segregation during cell division are dependent upon proper FAZ formation and flagellum elongation (Absalon et al., 2007; Bonhivers et al., 2008b). The single Golgi is also precisely positioned within the cell, and while there has been no evidence on its physical interaction with other cytoskeletal structures, it has been shown to share the same spatial-temporal dynamics of duplication and segregation (Field et al., 2000; He et al., 2004). 1.1.5 T. brucei cell division T. brucei cell division is a rigorous, spatiotemporally-coordinated and highly reproducible process -a fact that has significantly aided analysis on the regulation of the cell cycle and other cellular processes (Robinson and Gull, 1991; Sherwin and Gull, 1989a; Woodward and Gull, 1990). This has enabled the cell cycle progression to be monitored simply by using a DNA dye to visualize the nucleus and kinetoplast, the G1 and S phases of which are closely related to their relative stages of division and segregation; unlike other eukaryotic cells, the trypanosome coordinates the S-phases of both its DNA masses, namely nuclear DNA, and the mitochondrial DNA within the kinetoplast (Woodward and Gull, 1990). Cells with one kinetoplast and one nucleus (1K1N) are in the G1/S phase, while those with two kinetoplasts and a single nucleus (2K1N) indicate that the cells are in the G2/M phase. Cells bearing segregated kinetoplasts and nuclei (2K2N) are on the verge of cytokinesis (Sherwin and Gull, 1989a; Woodward and Gull, 1990). In the same manner, antibodies have been raised against several key parasite organelles and proteins, and immunostaining using these antibodies to complement DNA staining has opened up even more insight into the coordinated dynamics of many key cellular processes (Sherwin and Gull, 1989a). 10 The start of the cell cycle begins with the S-phase of mitochondrial DNA, closely followed by basal body maturation and duplication during the G1-S transition. The maturing pro-basal body then seeds the new flagellum, which invades the existing flagellar pocket to form the new flagellar axoneme (Sherwin and Gull, 1989a) (Figure 1B). In the T. brucei procyclic stage, the new flagellum tip is physically connected to the old flagellum via a mobile transmembrane junction known as the flagella connector; in a novel example of cytotaxic inheritance, transmission of cell polarity and axis in cell shape, cell division as well as the direction of motility of daughter cells takes place as the new flagellum elongates along the old flagellum, guided by the flagellar connector (Beisson and Sonneborn, 1965; Briggs et al., 2004; Moreira-Leite et al., 2001). Interestingly, disruption of the new flagellum extension was shown to result in a shorter FAZ construction, whose length correlates with that of the new flagellum. Accordingly, the progeny that inherits the new flagellum during cell division is shorter, thus establishing a direct correlation between the flagellum, FAZ and cell length (Kohl et al., 2003). Basal body migration is also affected (Absalon et al., 2007; Davidge et al., 2006); ultimately, disturbance to flagellum growth or flagellum attachment to the cell body is lethal to the cell (LaCount et al., 2000; Nozaki et al., 1996). Indeed, experiments generating morphometric measurements have also affirmed that cell length is more closely related to flagellum length rather than to cell volume (Rotureau et al., 2011). The Golgi apparatus also duplicates at this time and segregates together with the duplicated kinetoplasts and flagella, powered by the movement of the segregating basal bodies (Field et al., 2000; He et al., 2004) (Figure 1C). This process is followed by nuclear mitosis; nuclear DNA divide within an intact nuclear membrane which then also segregate (Sherwin and Gull, 1989a) (Figure 1D). The completion of mitosis leaves one of the two nuclei positioned between the two kinetoplasts, thus 11 ensuring that the ensuing cleavage leaves both daughter cells with a full complement of organelles (Robinson et al., 1995) (Figure 1E). Interestingly, although mitotic checkpoints in eukaryotes traditionally track the progress of chromosomal duplication and segregation, the progress of cytokinesis in T. brucei seems to depend on the completion of kinetoplast segregation rather than nuclear mitosis; indeed, cytokinesis still occurs during mitotic spindle disruption, generating zoids -daughter cells with a kinetoplast but no nucleus (Hammarton et al., 2003; Li and Wang, 2003; Ploubidou et al., 1999). The cleavage furrow ingresses in a unidirectional manner from the anterior to the posterior of the cell, passing between the old and the new flagella. Exactly how the site of ingression is determined is still unknown, although it has been postulated that the FAZ provides the structural information necessary to position the cleavage furrow. Indeed, the FAZ forms a unique "seam" in the corset microtubules due to the reverse polarity of the MtQ, and its elongation is concomitant with the growth of the new flagellum towards the anterior end of the cell where cleavage initiates (Robinson et al., 1995). Extension of the new flagellum is accompanied by a concomitant elongation of the subpellicular microtubules at the posterior end and intercalation of new microtubules within the existing cortical network, resulting in a significant increase in total cell volume (Rotureau et al., 2011; Sherwin and Gull, 1989b; Sherwin et al., 1987). It has been suggested that the intercalation of new microtubules indicates that the sub-pellicular microtubules are distributed semi-conservatively to the daughter cells (Sherwin and Gull, 1989b; Sherwin et al., 1987), although exactly how this happens at a single microtubule level, or how the cell coordinates the duplication and segregation of both its microtubules and organelles during cell division is still unknown. As in mammalian cells, the resulting daughter cells remain attached for a short period of time after cytokinesis before the final abscission. 12 FIGURE 1. Cartoon representation of the major cell cycle stages in T. brucei. (A) A single copy of the major organelles (nucleus, kinetoplast, basal body and the golgi apparatus) are present in an interphase cell. The single flagellum is tethered to the cell body via the FAZ structure. (B,C,D,E) As the cell undergoes cell division, the organelles duplicate and segregate in strict chronological and temporal order, which culminates in cytokinesis. This allows parasite cell cycle stages to also be categorized according to the division and segregation state of the nucleus and kinetoplast (1K1N, 2K1N, 2K2N), readily visualized with DAPI staining using fluorescence microscopy. Green minicircles, basal bodies; blue minicircles, kinetoplasts; red dots, Golgi; blue circles, nuclei. The pink line marks the older, existing flagellum while the yellow one represents the new flagellum. Both new and old FAZ structures are represented by a series of faint lines undergirding the flagella. 13 1.2 An overview of End-Binding protein 1 (EB1) 1.2.1 EB1 homologues The EB family comprises a group of microtubule plus-end tracking proteins (+TIPs) which have been evolutionarily conserved and studied in organisms ranging from yeast to humans. The first characterized member of the family, human EB1, was identified in a yeast-twohybrid screen for interacting partners of the adenomatous polyposis coli (APC) tumor suppressor protein COOH terminus; hence the name End-Binding protein (Su et al., 1995). Since then, EB1 has been found in nearly every organism and cell type studied, even in unicellular organisms that lack APC, arguing for a more primitive role that predates the appearance of APC in evolution (Tirnauer and Bierer, 2000). Mammals have three EBs (EB1, EB2 and EB3) which share 57-66% amino acid identity; although similar in structure, they are encoded by different genes (Su and Qi, 2001). Single EB homologues have also been identified in Botryllus schlosseri (EB1-BOTSC) (Pancer et al., 1996), fission yeast Schizosaccharomyces pombe (Mal3) (Beinhauer et al., 1997) and budding yeast Saccharomyces cerevisiae (Bim1p) (Schwartz et al., 1997). EB is also conserved in plants Arabidopsis thaliana has been reported to harbour at least 3 EB homologues (Chan et al., 2003; Mathur et al., 2003). 1.2.2 EB1 domain organization At a structural level, EB proteins are small, globular dimers which typically contain highly conserved N- and C-terminal domains that are connected by a less conserved linker sequence (Figure 2). The N-terminal domain, since determined to be both necessary and sufficient for microtubule binding, contains a calponin homology (CH) domain, the crystal structure of which has been shown to be a highly conserved fold (Hayashi and Ikura, 2003; 14 Slep and Vale, 2007). The C-terminal region contains a coiled-coil domain, which mediates the parallel dimerization of EB protein monomers (Honnappa et al., 2005). The coiled-coil domain of EB proteins partially overlaps with a unique EB1-like motif known as the endbinding homology (EBH) domain (Honnappa et al., 2005), which is implicated in EB1 interaction with numerous binding partners carrying the SxIP sequence motif (Akhmanova and Steinmetz, 2008; Honnappa et al., 2009). X-ray crystallography of the EB1 C-terminal showed that the EBH domain forms a coiled-coil that, in its homodimeric structure, folds back upon itself to form a 4-helix bundle, with its most invariant and conserved residues either buried in a deep hydrophobic cavity or forming a polar rim (Honnappa et al., 2005; Slep et al., 2005). The flexible C-terminal 20-30 residue tail of EB proteins mostly comprises a low complexity sequence, and is believed to play a role in EB1 self-inhibition. Most EB proteins, however, also harbour within this region a highly conserved acidic-aromatic EEY/F sequence motif similar to that found in α-tubulin and CAP-Gly proteins, the latter of which has been documented to interact with EB1 at this very site (Honnappa et al., 2005; Komarova et al., 2005; Weisbrich et al., 2007). FIGURE 2. Schematic diagram of human EB1 domain organization. EB1 comprises two highly conserved functional modules (the Calponin homology (CH) domain and EB1 homology (EBH) domain) separated by a more variable linker sequence. The carboxyl-terminus acidic tail is composed of low complexity sequence. The CH domain and linker sequence (indicated in blue) are positively charged, while the presence of acidic residues in the EBH domain and disordered tail region (indicated in red) results in the EB1 C-terminal being negatively charged. Domain boundaries are indicated by residue positions directly below the diagram. 15 1.2.3 EB1 cellular localization While many proteins localize to the microtubule cytoskeleton, specific localization to microtubule plus ends is a characteristic belonging to relatively few. EB1 appears to belong to this category, exhibiting the propensity to localize with a higher concentration to plus ends of both cytoskeletal and mitotic microtubules. For example, although over-expressed GFP-Bim1p in budding yeast was shown to localize to the entire microtubule cytoskeleton, native levels of GFP-Bim1p expression resulted in a selective localization to microtubule plus ends and the spindle pole body (Tirnauer et al., 1999). This pattern in also seen in higher organisms; in mammalian tissue culture cells, EB1 has been shown to localize to the distal tips of cytoskeletal microtubules, centrosomes, spindle poles as well as the midbody at different stages of the cell cycle (Morrison et al., 1998). There has been, however, controversy surrounding the mechanism by which EB1 localizes and attaches to microtubules; while EB1 has been shown to track growing ends of microtubules independently of other +TIPs (Bieling et al., 2007) and bind directly to microtubule filaments (Hayashi and Ikura, 2003), multiple studies have debated whether EB1 first copolymerizes with tubulin dimers and therefore preferentially accumulates at microtubule plus ends (Juwana et al., 1999; Slep and Vale, 2007) or whether EB1 specifically recognizes and binds with increased affinity to microtubule plus ends directly because of its distinct biochemical and/or structural state (Bieling et al., 2007; Dragestein et al., 2008). 1.2.4 EB1 as a keystone +TIP protein Despite disagreements on the exact mechanism employed, there is an increasing perception of EB1 as a master plus-end tracking protein which recruits multiple distinct +TIPs and itself forms the core for various protein complexes that form at dynamic microtubule plus ends 16 (Lansbergen and Akhmanova, 2006). The budding yeast EB1 homologue Bim1p, for instance, binds a protein complex containing Kar9 and Myo2p, resulting in the cortical capture of microtubules which facilitates orientation of the spindle towards the yeast bud site (Korinek et al., 2000; Lee et al., 2000). Studies have also shown that EB1 activity is crucial for recruitment of +TIP CLIP-170 to microtubule plus ends in fission yeast (Bieling et al., 2007), an observation consistent with RNAi studies in mammalian cells, suggesting that EB1 was pivotal in localizing CLIP-170 to the dynamic ends of microtubules (Komarova et al., 2005). In vertebrate cells, EB1 is also attributed with the ability to bind APC and target it to the growing ends of microtubules (Mimori-Kiyosue et al., 2000). The functional significance of this is still uncertain, although it has been previously shown that ablations of the APC Cterminal EB1 binding domain are frequently associated with familial and sporadic colorectal cancers (Polakis, 1997). 1.2.5 Role of EB1 in mitosis EB1's ability to localize independently to plus ends of mitotic and cortical microtubules also allows it to modulate their dynamic behaviour throughout the cell cycle; shedding light on EB1's ability interaction with various microtubule structures as well as binding partners would offer new perspectives on cell cycle progression and cellular processes. For instance, EB1 has been shown to localize to the interface between kinetochores and growing microtubules, suggesting that EB1 may modulate microtubule dynamicity during mitosis. Several experiments seem to support this; deletion of Bim1 results in aberrant spindles and nuclear migration defects (Schwartz et al., 1997), while loss of Mal3 caused an increase in the number of cells exhibiting condensed chromosomes and displaced nuclei. While no gross morphological abnormalities were observed in the spindles of Mal3-deficient cells, overexpression of Mal3, however, resulted in compromised spindle formation, severe growth 17 inhibition and abnormal cell morphology (Beinhauer et al., 1997). Generation of an EB1 null mutant in Dictyostelium confirms that EB1 is required for proper mitotic spindle formation (Rehberg and Graf, 2002), an observation that agrees with the RNAi studies carried out in Drosophila, which also resulted in defective chromosomal segregation (Rogers et al., 2002). EB1 has also been reported to localize to centrosomes in a process independent of microtubule association (Louie et al., 2004). Localization of EB1 at the centriole/basal body of fibroblasts is implicated in the assembly of its primary cilia (Schroder et al., 2007). The role of EB1 seems to extend beyond its involvement in cilia/flagella assembly; interestingly, EB1 has also been shown to not only localize to the basal body but also to the flagella tip of Chlamydomonas reinhardtii, and depletion of EB1 is accompanied by accumulation of intraflagellar transport (IFT) particles near the flagella tip (Pedersen et al., 2003). 1.2.6 Putative mechanisms of EB1 cellular interaction Years of study have made it clear that EB1 plays a major role in regulating microtubule dynamics both in vivo and in vitro systems, although opinions differ as to EB1's precise influence on the different parameters which govern the dynamic instability of microtubules. The controversy is exacerbated by differing (and sometimes seemingly conflicting) results from experiments carried out in different organisms and in various experimental systems. Bim1p, the budding yeast homologue of EB1, for example, has been shown to promote microtubule dynamicity (Schwartz et al., 1997; Tirnauer et al., 1999). Indeed, microtubules in bimI-null cells are considerably less dynamic compared to their wild-type counterparts (Tirnauer et al., 1999), an observation that agrees with RNAi studies carried out in Drosophila, which shows that the loss of EB1 causes most of the microtubules to enter a 'paused' state, in which they neither grow nor shrink, although this does not alter overall microtubule organization in interphase cells (Rogers et al., 2002). This observation mirrors 18 the results obtained from RNAi studies in mouse fibroblast cells, where EB1 depletion caused microtubules to spend more time pausing and less time in growth (Kita et al., 2006). Other studies, however, suggest that EB1 stabilizes microtubules through various mechanisms; readdition of EB1 to EB1-immunodepleted Xenopus egg extracts decreases microtubule catastrophes and promotes rescues, leading to increased microtubule polymerization and decreased pausing (Tirnauer et al., 2002). Similar results were reported for Mal3 in fission yeast (Busch and Brunner, 2004), while in Arabidopsis, over-expression of AtEB1a-GFP resulted in microtubule stabilization (Chan et al., 2003). EB1 has also been shown to promote microtubule stabilization in mammalian cell cultures (Wen et al., 2004), even if it exerts little effect on microtubule growth rates or rescues (Komarova et al., 2009). Results are equally varied in in vitro studies and experiments in purified systems. One in vitro study in fission yeast suggested that while Mal3 neither stabilizes nor destabilizes microtubule tips, it acts to stabilize the microtubule lattice, effectively inhibiting shrinkage via microtubule depolymerization and increasing the frequency of rescues (Katsuki et al., 2009). This study seems to affirm previous findings that Mal3 stabilizes the microtubule lattice seam (Sandblad et al., 2006). Mal3 has also been shown to induce initial formation of tubulin sheets at growing microtubule ends (Vitre et al., 2008) and then promoting microtubule assembly into 13-protofilament microtubules with a high proportion of A-lattice protofilament contact (des Georges et al., 2008), thereby stimulating microtubule nucleation, sheet growth and closure. Other studies on Mal3 and mammalian EB1, however, seem to indicate that EB1 actually stimulates microtubule dynamics by increasing the frequency of both catastrophes and rescues, suggesting instead that in cells EB1 prevents catastrophes by counteracting other microtubule regulators (Bieling et al., 2007; Komarova et al., 2009). Still further in vitro experiments described microtubule catastrophe 19 suppression by EB1 (Manna et al., 2008) or asserted that EB1 does not significantly alter microtubule dynamic instability parameters in the presence of tubulin alone, suggesting that other cellular factors may modulate EB1 behaviour within the cell (Dixit et al., 2009). Much of this experimental variation may be due to differences in EB1 concentration used, tubulin preparations, purification or visualization tags (Zhu, 2011) and other assay conditions, but the mechanisms employed by EB1 in its role as a regulator of microtubule dynamics still remain the subject of intense discussion. 1.3 Why study EB1 in T. brucei? EB1 is known to localize directly to the plus-ends tips of growing microtubules, recruiting other +TIPs in the process and itself forming the core of fast-changing +TIP complexes (Akhmanova and Steinmetz, 2008) which dynamically modulate microtubule behaviour. Coupled with the unidirectional arrangement of microtubules forming the sub-pellicular corset in which the growing ends point towards the posterior tip of the cell, the trypanosome microtubule cytoskeleton has been shown to be highly polarized, which in turn enforces cellular polarity as it directs and coordinates major cellular events such as cell division. In addition, the FAZ structure (which comprises a microtubular quartet that collectively possess a polarity opposite to that of the sub-pellicular corset) is also thought to define the axis and direction of cytokinesis during cell division. As such, it is obvious that precise modulation and proper regulation of the T. brucei microtubule cytoskeleton is essential for parasite survival. T. brucei possesses a cytoskeletal organization unlike no other; its heavy reliance on an extensively developed microtubule cytoskeletal network, coupled with a reduced dependence on other eukaryotic cytoskeletal elements such as actin makes it an ideal model in which to study the effects of EB1 and the mechanisms by which they are exerted on the highly-regulated dynamics of the microtubule network -knowledge 20 crucial for a deeper understanding of parasite behaviour as well as of the mechanisms underlying EB1 function, form and interaction. It is interesting to note that although EB1 has been discovered and studied in organisms as diverse as humans, plants and sea urchins, there has not been a single published attempt to study EB1 in a trypanosomal system. Indeed, although trypanosomatids rely heavily on a microtubule-based cytoskeletal system for survival and have been noted to possess a putative EB1 homologue, there has been to date no published work on the characterization of trypanosomal EB1 homologues. Remarkably, this also holds true for protozoan parasites in general, a diverse group of unicellular eukaryotic organisms that have collectively caused a great number of diseases with devastating economic and socio-political ramifications, of which T. brucei is a major representative. To date, there has only been one study on EB1 in protozoan parasites, which was conducted in Giardia lamblia (Kim et al., 2008). T. brucei, which is fast gaining acceptance as experimentally tractable, attractive model organism due to the tight spatiotemporal coordination of its cell cycle, the complete sequencing of its genome, the subsequent rapid development of molecular tools and experimental techniques, is potentially a good representative model in which to study EB1 -knowledge of which would be a step forward towards better understanding the molecular workings of these parasites, and thus possibly pave the way toward better parasitic disease management and new drug development. 21 In this study, I aim to take the first steps towards verifying and characterizing the putative T. brucei EB1 homologue; namely, to test and establish sequence and domain homology as well as to confirm its functional conservation as a +TIP. To this end, 1. I utilized bioinformatics tools to test sequence conservation and domain preservation, 2. Established EB1 localization within the T. brucei cell, 3. Scrutinized EB1 localization within the context of the T. brucei cell cycle via ectopic introduction of the YFP-EB1 fusion gene, 4. Attempted to characterize the EB1 RNAi phenotype in order to better understand EB1 function in the parasite, and 5. Obtained and purified an anti-TbEB1 antibody specifically raised against the putative T. brucei EB1 homologue, which facilitated further study on EB1 function via immunofluorescence assays and immunoblot analysis. 22 MATERIALS AND METHODS 2.1 Molecular cloning The coding sequence of the putative T. brucei EB1 homologue, TbEB1 (gene ID Tb09.160.1440) was found following a DNA sequence blast search of the T. brucei genome (a service kindly provided by the TriTryp database at http://tritrypdb.org/tritrypdb/) using the human EB1 sequence as search input. Desired fragments were then amplified via polymerase chain reaction (PCR). Annealing temperatures used depended on the size of the amplified fragment (Table 1), but standard PCR was generally performed in 50µl reactions using purified T. brucei genome as template, and amplified by either Taq DNA polymerase (Fermentas) or Advantage2 polymerase (Clontech), depending on the level of desired accuracy and length of the amplified fragment. All reactions were carried out on DNA Engine® Peltier Thermal Cycler or My Cycler™ Thermal Cycler (Bio-Rad, USA). PCR reactions were then subjected to DNA gel electrophoresis (1% agarose gel, run at 10V/cm), after which amplified fragments were identified and excised for purification using QIAquick PCR Purification Kit (QIAGEN). Purified DNA fragments and their designated plasmids vectors were subsequently digested with the suitable restriction enzymes, according to the protocol recommended by the manufacturers of the restriction enzymes used, before they were incubated together overnight at 16°C at a molar ratio of 1:3 respectively to facilitate ligation of corresponding restriction sites. Ligated constructs were then transformed into competent E. coli TOP10 cells via heat shock which were subsequently spread onto LB agar plates containing the necessary antibiotics and incubated for 12-16 hours at 37°C. This enabled selection of single-clone colonies. Harvested plasmid 23 constructs were then checked for sequence integrity before being reintroduced into E. coli cells for the purpose of construct amplification. 2.2 Cell lines, cultivation conditions and plasmid transfection All experiments highlighted in this thesis were conducted in either one of two procyclic cell lines - the YTat1.1 cell line (Ruben et al., 1983) or the 29.13 cell line (Wirtz et al., 1999). The YTat1.1 cell line was cultivated at 28°C in Cunningham's medium supplemented with 15% heat-inactivated fetal bovine serum (Clontech). The 29.13 cell line was maintained at 28°C in Cunningham medium containing 15% heat-inactivated, tetracycline-free fetal bovine serum (Clontech) in the presence of 15μg/ml G418 and 50μg/ml hygromycin. Plasmids could be transfected either transiently or stably into parasite procyclic cells. 3050µg of plasmid was used in a transient transfection, while a stable transfection required at least 15µg of linearlized plasmid. Approximately 5x107 log-phase cells were mixed with the required amount of plasmid and subjected to 2 pulses of electroporation (at 1500V) with an interval of 10 seconds between pulses. All electroporation experiments were carried out on a BioRad Gene Pulser (1500 V, 25 μF, ∞ Ω) (Biorad). Transiently transfected cells were checked for ectopic gene expression between 16-28 hours post transfection, while cells which underwent stable transfection were typically cloned and subjected to antibiotic selection 6 hours post transfection. 24 2.3 Clonal selection of stable transformants by limiting dilution In order to obtain clonal cell lines of stably-transfected cells, parasite cultures were serially diluted in a 96-well microtiter plate such that the parasites were eventually cultured at dilutions below one cell per well (Rosario, 1981). To accomplish this, parasite cultures growing in mid log-phase were diluted two fold in each subsequent column of wells, resulting in a maximum dilution of 211 times the original culture concentration. Plates were then sealed and incubated at 28°C, 5% CO2 for approximately 2 weeks until clonal cultures were obtained. 2.4 RNAi assay A suitable EB1 RNAi sequence was selected using the online program RNAit (http://trypanofan.path.cam.ac.uk/software/RNAit.html) (Redmond et al., 2003). A 593 bp length of EB1 coding sequence was introduced into T. brucei in a pZJM vector linearized with SacII, after which stable transfectants were obtained via cloning by limiting dilution. Production of double-stranded RNA was induced via addition of 10 µg/ml tetracycline; in order to ascertain the degree of RNAi penetration, cultures were sampled every 24 hours cells were immunoblotted to check for EB1 protein concentration and examined for YFP-EB1 fluorescence. Cell concentration was also measured every 24 hours (up to 5 days) postinduction. 2.5 Anti-TbEB1 antibody The His-tagged EB1 (His-EB1) construct was generated by cloning the full-length TbEB1 DNA sequence in-frame into the E.coli vector pET30a+ vector (Novagen) (Table 3), generating a 25 fusion protein with a N-terminus six histidine residue tag. BL21 E.coli cells were transformed with the His-EB1 construct and plated on LB agar plates with suitable antibiotics to obtain a clonal population. Transformed cells were cultured at 37°C to an OD600 of 0.4 before induction with 0.4 mM isopropyl-beta-D-thiogalactoside (IPTG) overnight at 20°C. His-EB1 recombinant protein was then affinity-purified using a nickel column (Sigma) and eluted from the column with several rounds of Equilibrium buffer (0.1 M Tris [pH7.4], 500 mM NaCl and 10% glycerol) supplemented with increasing concentrations of imidazole. The pooled fractions containing His-EB1 were then exchanged into a gel filtration buffer (25 mM Tris [pH7.4], 500 mM NaCl) by running the fractions through a Superdex 200 gel filtration column (GE Healthcare) to prevent protein precipitation by imidazole. Purity of the purified His-EB1 was assessed using sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE); most His-EB1 protein was recovered in the soluble fraction. Purified protein was used for polyclonal antibody production in rabbits, and the affinity purified immune serum of one rabbit was used in all subsequent experiments. 2.6 Affinity purification of anti-TbEB1 polyclonal antibody A purified fraction of His-EB1 protein was run on an SDS-PAGE, transferred onto a polyvinylidene difluoride (PVDF) membrane (Biorad). Protein bands were visualized with Ponceau S dye to facilitate identification and excision of the His-EB1 band. The protein strip was then blocked with 5% milk in TBS-Tween 20 (TBST) for 20 minutes, followed by two washes in TBST before incubation with crude antibody serum overnight at 4°C. After three washes with TBST, bound antibodies were eluted with 0.1 M glycine-HCl buffer (pH2.7) twice; 5 minutes with gentle mixing for the first fraction, and 3 minutes for the second. 1/10 26 the glycine buffer volume of 2 M Tris (pH8.0) was then used to neutralize the antibody fractions, which would prevent antibody denaturation as a result of low pH. The resulting affinity-purified immune serum was then used in all subsequent immunoblotting and immunofluorescence experiments. 2.7 Immunofluorescence microscopy Concentrated T. brucei cell suspension was spread on sterile glass coverslips and allowed to settle. Cells were then permeabilized and fixed in methanol chilled to -20°C for 5-7 minutes. To better visualize anti-TbEB1 antibody labelling of the parasite cytoskeleton, cells were first extracted with 1% Nonidet P40 in PBS, then fixed in 4% paraformaldehyde (PFA) for an additional 20 minutes at 4°C. All coverslips were blocked in 3% BSA for 45 minutes before antibody labelling to prevent non-specific antibody binding. Fixed cells were then incubated with primary antibody in blocking solution for an hour at room temperature, then washed briefly in PBS before incubation with secondary antibody at room temperature for half an hour. Coverslips with attached cells were then counterstained with 2µg/ml 4, 6-diamidino-2phenylindole (DAPI) for 20 minutes, followed by washing in PBS before a final wash with Milli-Q water. Coverslips were mounted with Fluorescence Mounting medium (SouthernBiotech Fluoromount-GTM) and allowed to dry before observation. All primary antibodies used in this study and their relevant dilutions are listed in Table 3. All fluorescein-conjugated secondary antibodies (Sigma) were used at 1:2000 dilution. Fixed cells were observed under a fluorescence microscope (model Axio Observer Z1, Zeiss) equipped with a CCD camera (model CoolSNAP HQ2, Photometrics). Images were processed with Adobe Photoshop CS5. 27 2.8 Immunoblot analysis Parasite cells were washed in PBS and lysed by boiling the samples at 100°C for 5 minutes in 3X Loading Buffer (150 mM Tris-HCl [pH6.8], 6% SDS, 30% glycerol, 2.5% 2-mercaptoethanol, 0.06% Bromophenol Blue). Proteins in cell lysate were resolved by electrophoresis at for 1.5 hours at 120V on a 12% polyacrylamide gel, then electrophoretically transferred onto a methanol-activated PVDF membrane for an hour at 70V. The membranes were then blocked with 5% milk in TBST for an hour prior to incubation with their respective primary antibodies diluted in blocking solution for an hour, after which they were washed briefly in TBST. Membranes were then incubated with secondary antibodies conjugated with horseradish peroxidase for half an hour. After a few final washes in TBST, desired protein bands were visualized with SuperSignal® West Dura Extended Duration Substrate solution (Thermo Scientific) using a chemiluminescence detector (model ImageQuant LAS 4000, GE Healthcare). Should there be a need to reprobe the membrane with different antibodies, the membrane was stripped with stripping buffer (2% SDS, 62.5 mM Tris-HCl [pH6.8], 100 mM 2mercaptoethanol) for 30 minutes at 60°C, followed by a few brief washes with TBST. Membranes were then blocked again with 5% milk in TBST before incubation with the desired primary antibody. 28 Vector Purpose Vector origin pZJM Constitutive overexpression Endogenous replacement via homologous recombination RNAi Modified pCR4Blunt-TOPO vector (Morriswood et al., 2009) (Wang et al., 2000) pET30a+ Protein expression Novagen pXS2 TOPO (Bangs et al., 1996) Antibiotic resistance Cell type (expression) Visualization tag Linearization site for stable transfection E. coli T. brucei Ampicillin Blasticidin T. brucei YFP or BB2 NsiI Ampicillin Blasticidin T. brucei YFP or BB2 NsiI or PacI Ampicillin Phleomycin - SacII Kanamycin - T. brucei E. coli (strain BL21) - - TABLE 1. List of plasmids used in study. Antigen YFP CC2D YL1/2 PFR PAR L3B2 BB2 α-tubulin EB1 Host Organism Rabbit (pAb) Rabbit (pAb) Rat (pAb) Rat (pAb) Mouse (mAb) Mouse (mAb) Mouse (mAb) Mouse (mAb) Rabbit (pAb) Antibody Origin Abnova (Zhou et al., 2011) (Kilmartin et al., 1982) Abnova (Ismach et al., 1989) (Kohl et al., 1999) (Bastin et al., 1996) Santacruz biotechnology B512 This study Staining Pattern FAZ, basal bodies Tyrosinolated tubulin Paraflagellar rod Paraflagellar rod FAZ filament Cytoskeleton Under investigation Dilution for IB 1:1000 1:2000 1:100 Dilution for IF 1:500 1:500 1:1000 1:500 1:1000 1:25 1:100 1:1000 1:50 TABLE 2. List of antibodies used in study. Staining patterns for YFP and BB2 are dependent upon the protein to which they are fused. Only anti-YFP, antiTbEB1 and anti-α-tubulin antibodies were used for immunoblot analysis in this study. 29 29 Construct pXS2YFP-EB1 pXS2BB2-EB1 TOPOYFP-EB1 Insert Full-length EB1 500 bp of 5' UTR and 500 bp of 5' end of coding sequence Primers Forward: CGGGATCCGACCATCGCAATACCCATGG Reverse: CGGAATTCTTACTCTGCAGCGTACAATAC Restriction sites Forward: BamHI Reverse: EcoRI UTR Forward: CCTTAATTAACGAGGAATGTAATGTTGGGG Reverse: CCCAAGCTTCGGTAACGATAATAACGGGG UTR Forward: PacI Reverse: HindIII 500 bp 5' coding sequence Forward: CGGGATCCATGGACCATCGCAATACCC Reverse: TGCATGCATATATCCCGTCTCACCACTGT 500 bp 5' coding Forward: BamHI Reverse: NsiI pZJM-EB1 RNAi fragment as identified by the RNAit program (Redmond et al., 2003) Forward: GCTCTAGAGGCCTTGGTGATGTGCTTAT Reverse: GCTCTAGAGTCTGCTTGTCCTCTACGGC Forward: BamHI Reverse: EcoRI pET30a-EB1 Full-length EB1 Forward: GAAGATCTGACCATCGCAATACCCATGG Reverse: CGGAATTCTTACTCTGCAGCGTACAATAC Forward: BglII Reverse: EcoRI TABLE 3. List of constructs and primers used in this study. 30 30 RESULTS 3.1. T. brucei putative EB1: establishing sequence homology and functional conservation 3.1.1 Bioinformatics A quick blast of the T. brucei genome (available on the TriTryp database) uncovered a single putative homologue (gene ID: Tb09.160.1440), a predicted coding gene on the T. brucei chromosome 9. It spans 1611 bases and has a predicted protein molecular weight of approximately 57 kDa. 2 major domains were identified by the TriTryp blast engine -a Nterminal calponin homology (CH) domain (residues 19-147), and a C-terminal EB1-like homology (EBH) domain (residues 489-534) with significant E-values (3.5 x 10-20 and 3.2 x 10-14 respectively) (Figure 3). FIGURE 3. Schematic diagram of the major EB1 domains. EB1 generally comprises 2 major domains which are linked by a flexible intermediate domain (I). The amino-terminus calponin-homology (CH) domain is implicated in microtubule-binding, while the carboxyl-terminus EB1-like homology (EBH) domain (which possesses the propensity to adopt a α-helical coiled coil structure) encompasses the unique EB1-like sequence motif, and has been shown to be crucial in +TIP interaction. Relative domain positions in the 57kDa putative T. brucei EB1 homologue, TbEB1, are indicated directly below the schematic. However, aligning the T. brucei putative homologue (hereafter referred to as TbEB1) with sequences of other established EB1 homologues indicated that they shared poor overall sequence homology, with the alignments returning significantly fewer matches towards the carboxy terminus (Figure 4). Indeed, human EB1 (268 aa) only shared 8% sequence identity with TbEB1, while Dictyostelium EB1 (a much longer sequence than human EB1 at 506 aa) scored a 12% sequence identity. Suspecting that this may be due to (i) the vastly different 31 lengths of different EB1 homologues, (ii) the presence of a poorly-conserved linker sequence between the two major EB1 domains (Bu and Su, 2003), which prevented alignment programs from picking up sequence similarities found at the protein C-terminus, and/or (iii) great evolutionary distances between the homologues (which intuitively suggested an inherently poor overall sequence homology), conservation of the major protein domains and motifs were then examined to establish at least a reasonable chance of functional conservation in TbEB1. Interestingly, alignment of the N-terminal CH domains and Cterminal EBH domains showed decent sequence similarity (NCBI's Blast program's estimate of sequence identities range from 24% to 38% for CH domains and 25% to 48% for EBH domains), of which the specific motifs of each domain showed the highest level of conservation. The N-terminal CH domain, for example, is traditionally defined by a number of almost invariant core residues, which are likely to be the major residues dictating the eventual folding of the domain's 3D-structure (Gimona et al., 2002). These residues also mainly conserved in TbEB1, as are the residues in the actin-binding sites (ABS) which characterize the CH domain (Figure 5a). Alignment of the C-terminal EBH domains also indicated decent sequence conservation, particularly at the EB1-like sequence motif which straddles two α-helices crucial for EBH domain function (Figure 5b) (Honnappa et al., 2005). It is interesting to note, however, that the slightly less conserved C-terminal EEY/F motif, shown to be vital for EB1 interaction with several important CAP-Gly +TIPs such as CLIP-170, is conspicuously absent from the TbEB1 sequence, which terminates prematurely compared to human EB1 C-terminal sequence. 32 33 FIGURE 4. Sequence alignment of full length EB1 homologues from different species. Accession numbers are as follows: Homo sapiens (human), SwissProt: Q15691; Xenopus laevis, GenBank: AAH68630; Drosophila melanogaster, TrEMBL: Q9V9A6; Caenorhabditis elegans, GenBank: NP_507526; Arabidopsis thaliana, GenBank: BAB11500; Dictyostelium discoideum, TrEMBL: Q8WQ86; Saccharomyces cerevisiae, SwissProt: P40013; Trypanosoma brucei, TriTrypt gene ID: Tb09.160.1440. 34 FIGURE 5a.. Sequence alignment of the Calponin Homology (CH) domains. Residues shaded in red (and marked with an asterisk) are invariant, while residues shaded in blue (and double-dotted) are highly conserved. The red lines indicate actin-binding actin binding sites (ABS). FIGURE 5b.. Sequence alignment of the EB-like EB homology (EBH) domains. Residues shaded in red (and marked with an asterisk) are invariant nt while those in blue (and marked with a colon)) are highly conserved. The red lines indicate the EB1-like EB1 sequence ence motif. 35 Having established that TbEB1 shared decent EB1 domain sequence identity with established EB1 homologues, the TbEB1 sequence was then blasted on the TriTryp database to recover potential EB1 homologues in other kinetoplastids. The search turned up serveral putative homologues from other trypanosomatid species, many of which are agents of diseases with tremendous socio-political and economic impact. However, no genes with significant sequence homology with TbEB1 was found in Leishmania, as previously reported (Berriman et al., 2005). As with the previous alignment attempts between TbEB1 and established EB1 homologues, aligning the putative trypanosomatid homologues also resulted in the same trend -the number of matches significantly decreased towards the carboxy-terminus, resulting in poor overall sequence homology (Figure 6). As before, aligning only the CH and EBH domains also returned highly-matching alignments (Figures 7a and 7b), which supports the existence of a very poorly conserved linker sequence . It also indicates that TbEB1 is, at least at a sequence level, a good representative of trypanosome EB1 homologues, and could potentially be a good experimental model in which to further study trypanosome EB1 form and function. 36 FIGURE 6.. Sequence alignment of full length putative trypanosomatid EB1 homologues. Species identities are as follows: tbru, Trypanosoma brucei; tcon, Trypanosoma congolense; congolense tviv, Trypanosoma vivax; tcru, Trypanosoma cruzi strain CL Brener.. Their respective gene IDs are as shown above. All sequences are declared conserved and hypothetical by the TriTrp database. 37 FIGURE 7a. Sequence alignment of the trypanosomatid putative EB1 Calponin Homology (CH) domains. Residues shaded in red (and marked with an asterisk) are invariant, while residues shaded in blue (and double-dotted) dotted) are highly conserved. The red lines indicate actin-binding actin binding sites (ABS). FIGURE 7b.. Sequence alignment of the trypanosomatid putative EB-like EB like homology (EBH) domains. Residues shaded in red (and marked with an asterisk) are invariant while those in blue (and marked with a colon) are highly conserved. The red lines indicate the EB1-like EB1 sequence equence motif. 38 3.1.2 Establishing cellular localization of TbEB1 In order to establish TbEB1's localization within the cell, TbEB1 constructs bearing either YFP or BB2 N-terminal visualization tags were introduced into 29.13 procyclic cells. Upon observation using fluorescence microscopy, the labelling pattern of over-expressed BB2- or YFP-tagged EB1 was established to be virtually identical (Figure 8A,C). Cells with a YFP-EB1 construct knock-in at a native locus that replaced the original putative EB1 gene (thus expressing endogenous levels of YFP-EB1) also appeared to exhibit the same localization pattern (Figure 8B); however, given the relatively weak YFP signal in these cells, subsequent localization experiments were conducted using the YFP-EB1 over-expressing cell line. FIGURE 8. TbEB1 localized to the posterior tip of the cell. TbEB1 exhibited identical localization patterns regardless of visualization tag (YFP or BB2) or expression level (endogenous or overexpression). Immunofluorescence pictures are arranged above their corresponding DIC picures. (A) Over-expression YFP-EB1, (B) endogenous-level expression YFP-EB1, (C) over-expression BB2-EB1. 39 Over-expression of YFP-tagged TbEB1 (YFP-EB1) resulted in specific localization to the posterior tip of the cell, widely accepted to be where the collective plus ends of the unidirectional corset microtubules converge (Robinson et al., 1995). A weaker signal at the tapered anterior tip of the cell was also generally observed. Double-labelling of fixed cells with other cellular markers affirmed the observation; co-staining YFP-EB1-expressing cells with YL1/2, an antibody which recognizes growing microtubule ends and the basal bodies (Sherwin et al., 1987), clearly showed TbEB1 localizing as an intense dot at the very tip of microtubule ends labelled with YL1/2 in interphase cells (Figure 9). The YFP-EB1 fluorescence pattern was also replicated in cells co-labelled with anti-α-tubulin monoclonal antibody raised against an epitope which does not undergo post-translational modification (Figure 9). Interestingly, TbEB1 labelling at the posterior tip of many interphase parasite cells did not seem to overlap with the ends of the microtubules labelled with either YL1/2 or anti-αtubulin antibody, suggesting that TbEB1 may selectively localize directly to the microtubule ends rather than processively track the microtubules to the growing ends of the filament. Co-staining with anti-α-tubulin antibody also showed that YFP-EB1 localization did not extend into either the existing flagellum or the daughter flagellum (if present), which remained unlabelled in cells ectopically expressing YFP-EB1 despite the presence of the microtubular axoneme contained within the flagellum. A low-level general fluorescence was also observed in the cell body throughout the cell cycle, although fluorescence intensity varied at different stages of cell division. 40 The results thus far suggest that TbEB1 not only retains decent domain homology when compared to other established EB1 homologues, but it also localizes to the plus ends of microtubules in T. brucei, thus suggesting its ability to function as a classical +TIP. FIGURE 9. Co-staining YFP-EB1 with other cellular markers affirmed TbEB1 localization at microtubule plus ends. Methanol-fixed interphase cells over-expressing YFP-EB1 co-labelled with (A) YL1/2 and (B) anti-α-Tubulin (a-Tub) antibodies showed a bright spot at the posterior tip of the cell body (closed arrows) which generally did not overlap with either YL1/2 or anti-α-Tubulin staining. A fainter YFP signal was also sometimes observed at the anterior tip of the cell body (open arrows), believed to be where the FAZ MtQ plus ends are located. YL1/2 labels tyrosinolated tubulin, found at the growing ends of microtubules and basal bodies (arrowheads). 41 3.2 Tracking TbEB1 cellular localization throughout the T. brucei cell cycle Interestingly, further immunofluorescence studies of fixed parasite cells suggest that TbEB1 localization is subject to a temporal modulation which strongly correlates with progress through the T. brucei cell cycle, stages of which are easily identified by taking into consideration the presence or absence of the daughter flagellum and the length thereof, the position of the basal bodies, and the position (as well as division state) of the nucleus and kinetoplast (Figure 10). In cells ectopically expressing YFP-EB1 recombinant protein, TbEB1 localized as an intense dot at the posterior tip of the parasite cell throughout the cell G1 and S phase, which coincides with the 1K1N stage of cell division. The intense dot then elongated into a line that stretched towards the anterior from the posterior tip of the cell throughout the G2/M phase. The line appeared to lengthen in tandem with the nuclear duplication and segregation the cells progressed from the 2K1N to the 2K2N stage. Interestingly, the line was generally found on the side of the cell opposing the nascent FAZ structure, although it is still unclear whether this is an experimental artifact that arose due to flattening a cell and imaging it on a 2D plane. The specific localization of TbEB1 generally disappeared as the nucleus completed its mitotic division and the cell progressed into cytokinesis, although the intensity of general fluorescence in the cell also increased in tandem with cell cycle progression. Specific TbEB1 localization reappeared at the tip of the new daughter cells after the completion of cytokinesis. 42 FIGURE 10. TbEB1 localization at the posterior end of T. brucei exhibited a temporal modulation that correlated closely with cell cycle progress. In 1K1N cells, TbEB1 localized as a specific dot to the tip of the cell posterior end (closed arrow) (A), and remained there throughout basal body duplication (B). The segregating basal bodies exert a mechanical force on the physically linked kinetoplast (C), which then also divides and segregates together with the separating basal bodies (D). Note that the nascent FAZ structure (visualized here with anti-CC2D antibody) begins to form at this stage as well. The nucleus then undergoes mitosis and separates, leaving one daughter nucleus in between the separated kinetoplasts. At this point, TbEB1 localization elongated from a dot into a line that stretched towards the cell anterior (open arrow points the direction of elongation) (E). Once the organelles are in place, the cell undergoes cytokinesis (F), and TbEB1 relocalized to the posterior tips of the new daughter cells. 43 3.3 Functional study on TbEB1 Previous attempts at shedding light on EB1 function in other experimental models and systems via EB1 knockdown/knockout have resulted in differing outcomes (Komarova et al., 2009; Tirnauer et al., 1999; Wen et al., 2004). Therefore, in order to study the role of EB1 in T. brucei, multiple attempts were made to generate a EB1 RNAi cell line. Unfortunately, several culminating factors (such as the lack of an anti-TbEB1 antibody at this point, the absence of any obvious phenotype following EB1-RNAi induction and the resulting difficulty in mass-testing clones for complete RNAi penetration) ultimately made it unfeasible at this point to generate a RNAi cell line in which TbEB1 was completely depleted. To circumvent the problem, the EB1-RNAi construct was introduced into a YFP-EB1 over-expression background. Preliminary results were encouraging; there was substantial decrease in the fluorescence intensity of YFP-EB1 over the course of 48 hours post RNAi induction, suggesting that the EB1-RNAi construct was capable of knocking down TbEB1. Unfortunately, when the experiment was later repeated and allowed to run for a longer period of time, although cells observed for 5 consecutive days post-induction showed a further slight decrease in YFP-EB1 intensity after 48 hours, RNAi induction still failed to completely deplete TbEB1 after 120 hours of RNAi induction (Figure 13). This observation was supported by a parallel immunoblot experiment; membranes loaded with cell lysate from induced cells and probed with anti-TbEB1-antibody also registered a decrease in YFPEB1 and endogenous TbEB1 levels (Figure 11). Unfortunately, other than a decrease in the levels of general fluorescence and, to a lesser extent in the intensity of specific EB1 localization at the posterior tip, no other obvious phenotypes were observed following partial TbEB1 knockdown; induced EB1-RNAi cells exhibited no gross abnormality in cell morphology when scrutinized under fluorescence microscopy and the duplication rate of the induced EB1-RNAi cell line closely paralleled that of the control population (Figure 12). 44 With the present availability of anti-TbEB1 antibody, renewed attempts to obtain a clonal EB1 RNAi cell line are currently underway. FIGURE 11. Immunoblot analysis of EB1 RNAi up to 5 days post-induction. EB1 RNAi was induced in a YFP-EB1 over-expression background. YFP-EB1 levels decreased noticeably after 48 hours, while levels of endogenous EB1 also decreased gradually over time. However, EB1 RNAi failed to completely deplete cellular EB1 after 5 days of induction. FIGURE 12. Growth curve of TbEB1-RNAi induced cultures. Partial depletion of TbEB1 did not appear to have an adverse effect on T. brucei fitness at 120 hours post-induction, as evidenced by the minimal difference in doubling index between RNAi-induced and control cultures. 45 FIGURE 13. Immunofluorescence analysis of EB1 RNAi induction. (A) EB1 RNAi induction in a YFP-EB1 over-expression background resulted in a significant decrease in YFP intensity after 24 hours, followed by a further slight decrease until the end of the experiment. (B) Live-cell imaging of RNAi-induced and control populations 120 hours following RNAi induction confirmed significant, if incomplete, EB1 depletion in RNAi cells, particularly at the posterior tip of the cell (indicated with closed arrows). (C) Labelling induced cells with antiTbEB1 antibody revealed a corresponding decrease in endogenous EB1 levels throughout RNAi induction. Closed arrows indicate the posterior tip of the T. brucei cell body. 46 47 3.4 Production and characterization of anti-TbEB1 antibody Previous studies in other organisms have raised the possibility that visualization and purification tags may affect EB1 localization (Landgraf et al., 2012; Zhu, 2011). Moreover, at higher concentrations EB1 has been shown to bind along the microtubule lattice both in vitro (Vitre et al., 2008) and in mammalian cells (Ligon et al., 2003) instead of localizing specifically to the microtubule tips. In order to test the results obtained thus far in the cell lines over-expressing YFP-EB1 and BB2-EB1, attempts have been made to raise a polyclonal sera against TbEB1 that could be used in Western blot and immunofluorescence assays. Full length TbEB1 cDNA was fused to 6xHIS tag and expressed in E. coli. Affinity purified recombinant protein was then used for rabbit immunization to produce polyclonal antibodies. Initial immunofluorescence tests of the immune serum and protein-A purified immune serum yielded perplexing results. Similar to the localization pattern of YFP-EB1, anti-TbEB1 serum also stained a distinct dot at the posterior tip of the cell. Interestingly, the antibody also distinctly stained the nucleus/nuclear envelope and localized as a generally punctate line that seemed to closely follow the FAZ structure (Figure 14A). A corresponding immunoblot of T. brucei cell lysate probed with anti-TbEB1 antibody resulted in multiple bands, suggesting that the polyclonal serum may not be altogether specific against TbEB1 (Figure 14B). Immunostaining with protein-A purified immune serum yielded similar results. 48 FIGURE 14. Anti-TbEB1 immune serum is non-specific in its detection of TbEB1. (A) The fluorescence localization pattern of anti-TbEB1 immune serum from a rabbit immunized with purified His-EB1 protein showed a strong signal at the posterior tip of the cell body, as expected (arrow). However, the antibody also picked up a strong signal in the area closely following the FAZ structure, as well as a slightly weaker signal in the area around the nucleus. (B) Immunoblot analysis of 29.13 cell lysate probed with anti-TbEB1 immune serum resulted in multiple bands beside the expected TbEB1 band at ~57kDa. 3.4.1 Immunoblot analysis The serum was then affinity-purified to harvest anti-TbEB1 antibody fractions which most specifically recognized purified His-tagged TbEB1. Specificity of the affinity-selected serum fraction was demonstrated by immunoblotting against endogenous TbEB1 protein and YFPtagged EB1 in both 29.13 cells and YFP-EB1 over-expressing cells. The antibody recognized a single band at approximately 57 kDa that corresponded to the expected size of endogenous TbEB1 in 29.13 cells, as well as an additional band at approximately 84 kDa when probed against total cell lysate of YFP-EB1 over-expressing cells, which corresponded to the expected size of YFP-tagged EB1 (Figure 15). In contrast, probing total cell lysate of YFP-EB1 49 over-expressing cells with anti-GFP antibody resulted only in a single band at about 84 kDa. Collectively, these results affirm anti-TbEB1 antibody specificity for TbEB1. FIGURE 15. Western blot analysis of 29.13 (control) cells and PXS2YFPEB1 cells using affinity-purified anti-TbEB1. The His-EB1 affinity-purified serum specifically recognized TbEB1 in both endogenous wild-type and YFP-tagged form. As control, the membrane was also probed with anti-GFP antibody, which yielded a single band corresponding to YFP-EB1 on the PXS2YFPEB1 lane. 3.4.2 Immunoflorescence assay Immunofluorescent staining with the purified anti-TbEB1 antibody revealed a pattern that differed from the staining pattern of the original serum as well as the localization pattern of over-expressed YFP-EB1. The anti-TbEB1 antibody staining pattern labelled two specific areas within the trypanosomatid cell with varying levels of intensity: the posterior tip of the cell and a punctate line which closely followed the nascent daughter FAZ in dividing cells. Similar to the previous immunofluorescence assays, anti-TbEB1 antibody staining was localized as a bright dot at the posterior tip of the cell, which elongated concomitant with nuclear division and segregation (Figure 16). 50 FIGURE 16. Co-staining YTAT cells with anti-TbEB1 and YL1/2 antibody confirmed a temporallymodulated EB1 localization at the posterior tip of the cell body. The staining pattern of anti-TbEB1 antibody confirmed the TbEB1 localization pattern observed in YFP-EB1 over-expressing cells, which was temporally-regulated and closely correlated to the progress of the cell cycle. 51 Co-staining with anti-L3B2 antibody (FAZ specific) (Kohl et al., 1999) and anti-PAR antibody (flagellum specific) (Ismach et al., 1989) also revealed a selective staining that closely traced the newly-forming FAZ, especially in duplicating cells (Figure 17). Similar to the specific staining at the posterior tip of the cell, immunostaining at this location also exhibited a temporal modulation throughout the parasite cell cycle -although there was little to no antiTbEB1 antibody staining of the area surrounding the FAZ structure in 1K1N cells, the staining intensity increased dramatically at the 2K1N stage and was maintained throughout the 2K2N stage until the parasite cell underwent cytokinesis. Interestingly, there were two prominent places where the anti-TbEB1 antibody did not label: the area surrounding the older FAZ structure (Figure 17A) and the flagella, both old and new (Figure 17B). It was also noted that co-localization of anti-TbEB1 and L3B2 staining at the FAZ area was also not a 100% exact, as there is generally a diversion in localization pattern near the kinetoplast where the FAZ begins. The persistent, low level of general fluorescence seen in YFP-EB1 over-expressing cells was also observed with anti-TbEB1 antibody staining. It is, however, interesting to note that unlike the even wash of YFP signal in the YFP-EB1 over-expression cell line, the pattern of antibody staining was often punctate, suggesting that it may reflect a distribution of EB1 to the growing ends of individual microtubules in the sub-pellicular corset. 52 FIGURE 17. Co-staining YTAT cells with anti-TbEB1 and FAZ/flagellum markers confirmed a temporally-sensitive labelling pattern which closely traced the new growing FAZ structure. (A) Anti-TbEB1 and L3B2 (FAZ cytoplasmic-filament specific) co-labelling exhibited a colocalization pattern that shadowed the newly-forming FAZ, beginning from the 2K1N stage (new FAZ labelled with open arrows). (B) Co-staining with anti-TbEB1 and PAR (flagellum specific) antibodies confirmed that TbEB1 did not localize to either of the flagella, staining instead a punctate line that closely shadowed the FAZ undergirding the newer flagellum, indicated with open arrows. 53 DISCUSSION EB1 has long been studied in various eukaryotes since it was first discovered in 1995, ranging from single-celled yeast to evolutionarily advanced mammals. In the face of accumulating evidence, there is increasing acceptance of EB1 as a key regulator of microtubule dynamics as well as a master integrator of the +TIP network, which has resulted in a concerted and sustained attempt at understanding the exact role of EB1 in its cellular context. It is against this backdrop that T. brucei emerges as an attractive organism for the study of EB1. Its cytoskeleton is primarily dependent on an elaborated tubulin-based network that forms its major ultrastructures, such as the FAZ component, the sub-pellicular microtubule corset and its single flagella. In addition, microtubule dynamics are also implicated in the parasites major cellular events, such as mitosis, organelle segregation and cell division. Moreover, T. brucei also appears to have a reduced dependence on the other filament classes generally found in eukaryotic cells; there are presently no known T. brucei homologues of intermediate filaments, and although homologues for actin-myosin network components have been identified, the loss of actin in the procyclic form is not lethal at this stage (Shi et al., 2000). 4.1 TbEB1: a putative EB1 homologue? Given the parasite's heavy reliance on its microtubule cytoskeleton, it was only natural that a T. brucei EB1 homologue was hypothesized to exist. Blasting the human EB1 sequence on the TriTryp database returned a single plausible T. brucei candidate, with an E-value of 3.4 x 10-6, which denotes a significant sequence homology given the difference in sequence length 54 (human EB1, 268aa; TbEB1, 536aa) and evolutionary distance between the two putative homologues. The additional presence of a fairly well-conserved N-terminal CH domain and a C-terminal EBH domain, characteristic of established EB1 homologues, argues that TbEB1 may possess similar molecular interactions (and thus possibly similar cellular function), which serves to further strengthen the plausibility of TbEB1 being a true EB1 homologue. However, the premature truncation of the TbEB1 C-terminal domain in which the EEY/F motif (consistently found in existing EB1 homologues) appears to be deleted, raises questions as to how TbEB1 then interacts with CAP-Gly proteins which traditionally recognize and bind EB1 at this particular motif. Interestingly, a T. brucei appears to possess at least one putative CLIP170 homologue (E-value 1.2 x 10-23), a CAP-Gly protein which also localizes to the growing ends of microtubules; further study into the possibility and nature of TbEB1-CLIP170 interaction would certainly offer new perspective on microtubule plus-end dynamics in T. brucei. At any rate, it has been shown that the absence of the EEY/F motif does not compromise TbEB1's ability to accumulate at microtubule plus ends (Komarova et al., 2005), although it is also interesting to note that the EEY/F domain has been implicated in EB1 autoinhibition, which negatively regulates EB1's ability to suppress microtubule shortening (Hayashi et al., 2005; Manna et al., 2008). It has also been shown in at least two in vitro studies that a constitutively active mutant exerts a stronger effect on microtubule dynamics even at a very low EB1:tubulin ratio compared to full length EB1 (Hayashi et al., 2005; Manna et al., 2008), which suggests that TbEB1 dynamics may differ from standards which have been previously established for other EB1 homologues. 55 Results from the fluorescence studies also seem to corroborate with existing bioinformatics data on supporting TbEB1 as a possible EB1 homologue. Following the localization pattern of established EB1 homologues, YFP- tagged TbEB1 seems to localize to the plus ends of cortical microtubules which converge at the posterior tip of the trypanosome cell, resulting in a brightly fluorescent dot which forms at the very tips of the microtubules. This suggests that TbEB1, like most EB1 homologues, may selectively recognize and associate with growing microtubule plus ends (either due to its distinct biochemical/structural state) instead of undergoing processive transport to the microtubule tips (Maurer et al., 2012). It is precisely this distinct ability to localize independently to the microtubule plus ends that allows EB1 to act as a key recruiter of modulating +TIPs to the growing tips of microtubules. Over-expression of YFP-EB1 also resulted in a persistent, low level of fluorescence throughout the cell body. This supports the findings of a previous study which showed that new, short microtubules started invading the existing cytoskeletal array from very early on in the cell cycle by intercalating the old microtubules of the sub-pellicular corset in preparation of eventual cell division (Sherwin and Gull, 1989b). Localization of EB1 to the plus ends of individual microtubules, then, would result in the punctate-like distribution of TbEB1 localization, which is clearly and fairly consistently observed in anti-TbEB1 antibody staining. In addition, another study in budding yeast also showed that over-expressed GFP-Bim1 localized to the entire microtubule skeleton instead of preferentially to the microtubule plus ends, which may provide an alternative explanation for uniformed fluorescence in the cell body (Tirnauer et al., 1999). Cellular fluorescence could also be due to the presence of a cytoplasmic pool of YFP-EB1; interestingly, treating cells with detergent (which strips off most of the parasite membranes while leaving an intact cytoskeleton) before staining with anti-TbEB1 antibody resulted in a marked decrease in the intensity of cellular fluorescence, 56 resulting instead in a punctate-like staining of the remaining cytoskeleton (Figure 18). This observation argues for the presence of free cytoplasmic YFP-EB1, although whether TbEB1 exists in a soluble complex with other proteins or as free molecules remains to be seen; EB1's ability to bind to free tubulin dimers, for instance, has long been a subject of debate (Diamantopoulos et al., 1999; Ligon et al., 2006). The combined results seem to suggest that the low level of cellular fluorescence in cells over-expressing YFP-EB1 might be due to a combination of several factors discussed above. FIGURE 18. Detergent treatment resulted in punctate-like EB1 localization throughout the cell body. Co-labelling fixed YTAT cells extracted with 1% NP40 showed that EB1 localized throughout the subpellicular corset (labelled with anti-α-tubulin antibody) in a punctate-like pattern. The absence of the typically vivid anti-TbEB1 staining at the posterior tip of the cell body is probably due to the loss of posterior-end microtubule integrity, resulting in a gaping hole -a feature commonly observed in detergent-treated cells (indicated by open arrows). 4.2 Tracking TbEB1 localization throughout the cell cycle TbEB1's temporally-sensitive localization in the dividing trypanosoma cell offers a fresh perspective on the modulation of microtubule dynamics throughout the cell cycle. Such cell cycle specificity, an unusual feature among microtubule-associated proteins (MAPs), not only adds to the unique appeal of studying EB1 as an informative component of microtubule network regulation but also suggests that EB1 could be used as a new marker for tracking T. brucei cell cycle progression. It has been postulated that the microtubule corset duplicates in a semi-conservative manner, with new short microtubules growing in between existing ones and using them as templates to preserve the shape of the newly-forming sub-pellicular 57 corset (Sherwin and Gull, 1989b). The results obtained in this study seem to agree with this model of semi-conservative inheritance, not least because the steady fluorescence signal of YFP-EB1 at the posterior end of the cell (combined with data from co-staining experiments with YL1/2) suggests that at no time does the existing sub-pellicular corset break down to reform into two new sets of daughter corsets during cell division. Moreover, the localization of YFP-EB1 (which elongates from a prominent dot into a line stretching from the posterior tip of the cell opposite the flagellum) presumably indicates that growing microtubule ends are concentrated in a strictly-defined area at the posterior end of the parasite cell, which extends as the cell cycle progresses. Interestingly, the extension of this area as the parasite moves through the cell cycle (visualized as an elongation of the YFP-EB1 signal) also shows a strong temporal correlation with nuclear division and segregation, prompting consideration of the possibility that the two events may also be co-regulated. 4.3 Characterizing the anti-TbEB1 antibody The discovery of TbEB1's temporally-regulated cellular localization has raised the possibility that TbEB1 could possibly be used as a new T. brucei cellular marker for the purpose of tracking the parasite cell cycle progression, or simply for establishing parasite microtubule cytoskeleton polarity -a potentially useful indicator in assays where cytoskeletal perturbations are expected. Thus far, only one other trypanosomal protein, known as Gb4, has been reported to localize to the posterior tip of the parasite. Similar to EB1, this 28kb protein has been postulated to cap the ends of microtubules and possibly connect them to the cell membrane (Rindisbacher et al., 1993); however, unlike EB1 Gb4 has not been shown to exhibit a temporally regulated localization pattern, nor has further work on this protein been thus far reported. Moreover, EB1 has never been studied in kinetoplastids; given that TbEB1 shares excellent sequence homology with the putative EB1 homologues in other 58 trypanosomatid species (many of which are direct causes of a wide array of diseases), it is our hope that an anti-TbEB1 antibody would eventually also be potentially useful in the study of other trypanosomatids. The anti-TbEB1 antibody exhibited a complex localization pattern when examined using immunofluorescence microscopy. While the anti-TbEB1 antibody picked up a similar signal at the posterior tip of the cell body when compared to the localization pattern of overexpressed YFP-EB1, co-staining experiments with L3B2, a marker that localizes to the cytoplasmic filament of the FAZ structure (Kohl et al., 1999), also surprisingly showed an additional prominent signal which closely follows the nascent FAZ in dividing cells. EB1's unique localization pattern, which presumably traces the plus-ends of growing microtubules, is reminiscent of the staining pattern of ɣ-tubulin, which instead labels the minus ends of microtubules. Scott et al., in their 1997 paper, reported that ɣ-tubulin was observed to localize as a bright dot to the tapered anterior tip of the cell body (where the minus ends of many of the sub-pellicular microtubules converge), along the cytoplasmic filaments of the FAZ structure, and as a general, low-level punctate fluorescence throughout the cell body (Scott et al., 1997). In many ways, this seeming coincidence is not unexpected, since ɣtubulin and EB1 are both microtubule end-tracking proteins (albeit at different ends), and electron microscopy images of the trypanosome sub-pellicular microtubule array seems to suggest that the corset is fairly uniform in its distribution of cortical microtubules in spite of the consistent direction of microtubule polarity (Sherwin and Gull, 1989a). However, unlike ɣ-tubulin's indiscriminate labelling of both FAZ structures in the cell, TbEB1's selectively staining on/near the newly-forming FAZ (a pattern which is, as a rule, much weaker or absent around the older FAZ) is fairly surprising, and indicates a temporally-sensitive facet to the localization of TbEB1. In a sense, this staining pattern is also reminiscent of the YL1/2 59 antibody staining pattern, which not only stains newly-formed microtubules but also the nascent daughter flagella in the cell cycle stages preceding kinetoplast segregation (Sherwin et al., 1987). Like YL1/2, anti-TbEB1 antibody also exhibits a localization that is closely associated with microtubule growth, subjected to temporal modulation but is at the same time closely connected to the progress of the parasite cell cycle (Sherwin et al., 1987). Despite the wealth of information afforded by the initial localization experiments utilizing constitutive over-expression of YFP-EB1 in T. brucei, it is clear the localization pattern of over-expressed YFP-EB1 does not entirely match the staining pattern of the anti-TbEB1 antibody. It is still unclear whether the persistent absence of staining along the newlyforming FAZ cytoplasmic filament in YFP-EB1 over-expressing cells is due to the signal being masked by the increased intensity of background fluorescence, or to the possibility that the visualization tag may hinder TbEB1 localization to certain parts of the parasite cell. Interestingly, staining an over-expressing YFP-EB1 cell with anti-TbEB1 antibody resulted in a staining pattern that was identical with YFP-EB1 localization, suggesting that specific FAZassociated staining could be masked by the increased background fluorescence intensity in YFP-EB1 over-expressing cells (Figure 19). This possibility is supported by the observation in cells expressing EB1 RNAi in a YFP-EB1 over-expression background; the FAZ labelling is sometimes observed 24 hours post-RNAi induction when the cells are probed with antiTbEB1 antibody, although the signal rapidly drops in intensity as the RNAi experiment progressed (Figure 20). In contrast, no such labelling was seen when YFP fluorescence was tracked during the same RNAi experiment, supporting the possibility that absence of FAZassociated labelling in YFP-EB1 cells could also be caused by YFP's inability to enter or label the FAZ structure. 60 FIGURE 19. Comparative fluorescence labelling of EB1 in a YFP-EB1 over-expressing cell line confirmed that the anti-TbEB1 antibody labelling pattern was virtually identical to YFP-EB1 localization. Both over-expressed YFP-EB1 and anti-TbEB1 staining exhibited the typical, temporallymodulating localization to the posterior tip of the cell (indicated with closed arrows in the 1K1N and 2K2N cells; the direction of signal elongation is indicated in the 2K2N cell by an open arrow). The FAZassociated staining seen in anti-TbEB1 labelling of wild-type YTAT cells was conspicuously absent. FIGURE 20. Anti-TbEB1 antibody labelling 24 hours post RNAi induction in a YFP-EB1 overexpression background. Partial depletion of YFP-EB1 reduced the YFP signal intensity sufficiently to allow anti-TbEB1 FAZ-associated staining to be visible (closed arrow). 61 4.4 Functional study of EB1 Although RNAi experiments were attempted in order to characterize the function of the putative T. brucei EB1 homologue, failure to completely deplete EB1 in T. brucei despite numerous attempts have unfortunately rendered the results inconclusive. Partial depletion of EB1 in a YFP-EB1 over-expression background did not appear to exert a drastic negative impact on cell fitness; a result that seems to contradict a previous high-throughput RNAi phenotyping report by Alsford et al. which indicated that TbEB1 depletion resulted in significant loss of fitness in procyclic cells (Alsford et al., 2011). The apparent contradiction may be due to the possibility that TbEB1's cellular function necessitates only a very small amount of EB1 to be present. Previous reports on EB1 dynamics offer support to this possibility; EB1 has been postulated to form the core of the microtubule plus-end complex, thus requiring only a few EB1 molecules to recruit a complement of other +TIPs which then exert a cooperative effect on microtubule dynamics (Lansbergen and Akhmanova, 2006). Moreover, EB1 has also been shown to exhibit rapid turnover at microtubule plus ends, averaging a dwell-time of 0.81 ± 0.06 seconds and undergoing multiple rounds of microtubule association/dissociation over the lifetime of a microtubule plus-end structure (Dixit et al., 2009). Coupled with EB1's ability to bind multiple distinct +TIPs with relatively weak binding affinities, it has been suggested that +TIPs function in rapidly changing networks of interaction (Akhmanova and Steinmetz, 2008) -a scenario well suited for a very low concentration of EB1 to exert a wide area and great variation of effect. Furthermore as mentioned earlier, the lack of the auto-inhibiting motif EEY/F at the TbEB1 C-terminal may also possibly endow TbEB1 with a constitutively active phenotype, allowing TbEB1 to exert a stronger effect even at low EB1:tubulin ratios compared to other EB1 homologues which possess a C-terminal EEY/F motif. 62 CONCLUSION AND FUTURE DIRECTION EB1 is a ubiquitous microtubule plus end binding protein that has been conserved across eukaryotic organisms of remarkably diverse evolutionary progress. EB1 homologues have since been studied in organisms as varied as yeast, sea urchins, plants and humans, suggesting a functionally crucial role for EB1 in the regulation of microtubule dynamics. The highly reproducible spatiotemporal regulation of the T. brucei cell cycle (not only wellunderstood as a result of intense research on this topic but also encountered time and again in this study) has served to underscore the suitability of the trypanosome as a model organism for the study of organelle biogenesis, organelle inheritance and microtubule regulation. Further exploitation of T. brucei's heavy reliance on its microtubule cytoskeletal network would certainly offer new perspectives on the function and mechanisms of EB1, a master regulator of +TIP activity and microtubule plus-end dynamics. The results gathered thus far in this study has been encouraging; not only does the putative EB1 homologue show significant domain sequence conservation when compared with other established EB1 homologues, TbEB1 also seems to retain its traditional proclivity for microtubule plus ends. The implications of this are two-fold; one, the unidirectional arrangement of microtubules in the sub-pellicular corset translates into a distinct EB1 localization at the posterior tip of the parasite cell body, which automatically positions the new anti-TbEB1 antibody as a potential cell polarity marker -a great help, for instance, in experiments that result in disruption of polarity indicators such as the cell shape, organelle arrangement and flagellum attachment. Second, the temporal modulation of EB1 localization at the posterior end of the cell adds a spatial dimension to the existing 63 knowledge on trypanosome microtubule cytoskeleton inheritance, widely believed to follow a semi-conservative model. It would be interesting to follow up on this point with 3Dmodelling studies in order to establish a spatially accurate picture of EB1's involvement in the microtubule dynamics involved in cytoskeleton inheritance. In addition, TbEB1 also shows a unique, temporally-sensitive localization pattern that closely shadows the nascent FAZ structure in duplicating cells; a pattern that interestingly does not extend to the older existing FAZ structure. This unique localization pattern is intriguing, and warrants further investigation in order to draw more accurate conclusions on EB1's exact localization and purpose in that locality; immuno-electron microscopy experiments, for example, would provide invaluable information on the detailed localization of TbEB1. The present results gleaned from this study have affirmed that further research on the trypanosomal EB1 homologue would be advantageous to the advancement of knowledge in both fields of EB1 and T. brucei research. A RNAi-mediated, complete depletion of EB1 in the trypanosomal cell would provide an invaluable first step towards a more thorough characterization of EB1 function in T. brucei. Domain truncation studies would then shed further light on the function of individual domains and help confirm TbEB1 as a true EB1 homologue, as would complementation experiments in BIM1-null yeast and other organisms with EB1 homologues. Given EB1's widely accepted role as a master regulator of +TIP activity, it is eventually also of paramount importance to the fuller elucidation of T. brucei microtubule dynamics to identify, characterize and further study the binding partners of TbEB1. 64 In the end, however, it is important never to lose sight of the forest for the trees. In the light of decades of weakened economies and ravaged lives that are the direct result of a rampant spread of protozoan parasitic diseases like African trypanosomiasis, the need for new treatment is not only urgent, but also fast becoming necessary. A deeper, more thorough understanding of the molecular mechanisms underlying parasite survival -such as the role of EB1 in the regulation of the trypanosome's microtubule dynamics- is key to the control and, hopefully, eventual eradication of the dreaded disease. 65 BIBLIOGRAPHY Absalon, S., L. Kohl, C. Branche, T. Blisnick, G. Toutirais, F. Rusconi, J. Cosson, M. Bonhivers, D. Robinson, and P. Bastin. 2007. Basal body positioning is controlled by flagellum formation in Trypanosoma brucei. PloS one. 2:e437. Akhmanova, A., and M.O. Steinmetz. 2008. Tracking the ends: a dynamic protein network controls the fate of microtubule tips. Nature reviews. Molecular cell biology. 9:309322. Alsford, S., D.J. Turner, S.O. Obado, A. Sanchez-Flores, L. Glover, M. Berriman, C. HertzFowler, and D. Horn. 2011. High-throughput phenotyping using parallel sequencing of RNA interference targets in the African trypanosome. Genome research. 21:915924. Angelopoulos, E. 1970. Pellicular microtubules in the family Trypanosomatidae. The Journal of protozoology. 17:39-51. Aslett, M., C. Aurrecoechea, M. Berriman, J. Brestelli, B.P. Brunk, M. Carrington, D.P. Depledge, S. Fischer, B. Gajria, X. Gao, M.J. Gardner, A. Gingle, G. Grant, O.S. Harb, M. Heiges, C. Hertz-Fowler, R. Houston, F. Innamorato, J. Iodice, J.C. Kissinger, E. Kraemer, W. Li, F.J. Logan, J.A. Miller, S. Mitra, P.J. Myler, V. Nayak, C. Pennington, I. Phan, D.F. Pinney, G. Ramasamy, M.B. Rogers, D.S. Roos, C. Ross, D. Sivam, D.F. Smith, G. Srinivasamoorthy, C.J. Stoeckert, Jr., S. Subramanian, R. Thibodeau, A. Tivey, C. Treatman, G. Velarde, and H. Wang. 2010. TriTrypDB: a functional genomic resource for the Trypanosomatidae. Nucleic acids research. 38:D457-462. Bangs, J.D., E.M. Brouch, D.M. Ransom, and J.L. Roggy. 1996. A soluble secretory reporter system in Trypanosoma brucei. Studies on endoplasmic reticulum targeting. The Journal of biological chemistry. 271:18387-18393. Bastin, P., Z. Bagherzadeh, K.R. Matthews, and K. Gull. 1996. A novel epitope tag system to study protein targeting and organelle biogenesis in Trypanosoma brucei. Molecular and biochemical parasitology. 77:235-239. Beinhauer, J.D., I.M. Hagan, J.H. Hegemann, and U. Fleig. 1997. Mal3, the fission yeast homologue of the human APC-interacting protein EB-1 is required for microtubule integrity and the maintenance of cell form. The Journal of cell biology. 139:717-728. Beisson, J., and T.M. Sonneborn. 1965. CYTOPLASMIC INHERITANCE OF THE ORGANIZATION OF THE CELL CORTEX IN PARAMECIUM AURELIA. Proc Natl Acad Sci U S A. 53:275282. Berriman, M., E. Ghedin, C. Hertz-Fowler, G. Blandin, H. Renauld, D.C. Bartholomeu, N.J. Lennard, E. Caler, N.E. Hamlin, B. Haas, U. Bohme, L. Hannick, M.A. Aslett, J. Shallom, L. Marcello, L. Hou, B. Wickstead, U.C. Alsmark, C. Arrowsmith, R.J. Atkin, A.J. Barron, F. Bringaud, K. Brooks, M. Carrington, I. Cherevach, T.J. Chillingworth, C. Churcher, L.N. Clark, C.H. Corton, A. Cronin, R.M. Davies, J. Doggett, A. Djikeng, T. Feldblyum, M.C. Field, A. Fraser, I. Goodhead, Z. Hance, D. Harper, B.R. Harris, H. Hauser, J. Hostetler, A. Ivens, K. Jagels, D. Johnson, J. Johnson, K. Jones, A.X. Kerhornou, H. Koo, N. Larke, S. Landfear, C. Larkin, V. Leech, A. Line, A. Lord, A. Macleod, P.J. Mooney, S. Moule, D.M. Martin, G.W. Morgan, K. Mungall, H. Norbertczak, D. Ormond, G. Pai, C.S. Peacock, J. Peterson, M.A. Quail, E. Rabbinowitsch, M.A. Rajandream, C. Reitter, S.L. Salzberg, M. Sanders, S. Schobel, S. Sharp, M. Simmonds, A.J. Simpson, L. Tallon, C.M. Turner, A. Tait, A.R. Tivey, S. Van Aken, D. Walker, D. Wanless, S. Wang, B. White, O. White, S. Whitehead, J. Woodward, J. Wortman, M.D. Adams, T.M. Embley, K. Gull, E. Ullu, J.D. Barry, A.H. Fairlamb, F. Opperdoes, B.G. Barrell, J.E. Donelson, N. Hall, C.M. Fraser, et al. 2005. The genome of the African trypanosome Trypanosoma brucei. Science (New York, N.Y.). 309:416-422. 66 Bieling, P., L. Laan, H. Schek, E.L. Munteanu, L. Sandblad, M. Dogterom, D. Brunner, and T. Surrey. 2007. Reconstitution of a microtubule plus-end tracking system in vitro. Nature. 450:1100-1105. Bonhivers, M., N. Landrein, M. Decossas, and D.R. Robinson. 2008a. A monoclonal antibody marker for the exclusion-zone filaments of Trypanosoma brucei. Parasites & vectors. 1:21. Bonhivers, M., S. Nowacki, N. Landrein, and D.R. Robinson. 2008b. Biogenesis of the trypanosome endo-exocytotic organelle is cytoskeleton mediated. PLoS biology. 6:e105. Briggs, L.J., P.G. McKean, A. Baines, F. Moreira-Leite, J. Davidge, S. Vaughan, and K. Gull. 2004. The flagella connector of Trypanosoma brucei: an unusual mobile transmembrane junction. Journal of cell science. 117:1641-1651. Brun, R., J. Blum, F. Chappuis, and C. Burri. 2010. Human African trypanosomiasis. Lancet. 375:148-159. Bu, W., and L.K. Su. 2003. Characterization of functional domains of human EB1 family proteins. The Journal of biological chemistry. 278:49721-49731. Busch, K.E., and D. Brunner. 2004. The microtubule plus end-tracking proteins mal3p and tip1p cooperate for cell-end targeting of interphase microtubules. Current biology : CB. 14:548-559. Chan, J., G.M. Calder, J.H. Doonan, and C.W. Lloyd. 2003. EB1 reveals mobile microtubule nucleation sites in Arabidopsis. Nature cell biology. 5:967-971. Cross, G.A. 2001. African trypanosomes in the 21st century: what is their future in science and in health? International journal for parasitology. 31:427-433. Davidge, J.A., E. Chambers, H.A. Dickinson, K. Towers, M.L. Ginger, P.G. McKean, and K. Gull. 2006. Trypanosome IFT mutants provide insight into the motor location for mobility of the flagella connector and flagellar membrane formation. Journal of cell science. 119:3935-3943. des Georges, A., M. Katsuki, D.R. Drummond, M. Osei, R.A. Cross, and L.A. Amos. 2008. Mal3, the Schizosaccharomyces pombe homolog of EB1, changes the microtubule lattice. Nature structural & molecular biology. 15:1102-1108. Diamantopoulos, G.S., F. Perez, H.V. Goodson, G. Batelier, R. Melki, T.E. Kreis, and J.E. Rickard. 1999. Dynamic localization of CLIP-170 to microtubule plus ends is coupled to microtubule assembly. The Journal of cell biology. 144:99-112. Dixit, R., B. Barnett, J.E. Lazarus, M. Tokito, Y.E. Goldman, and E.L. Holzbaur. 2009. Microtubule plus-end tracking by CLIP-170 requires EB1. Proc Natl Acad Sci U S A. 106:492-497. Dragestein, K.A., W.A. van Cappellen, J. van Haren, G.D. Tsibidis, A. Akhmanova, T.A. Knoch, F. Grosveld, and N. Galjart. 2008. Dynamic behavior of GFP-CLIP-170 reveals fast protein turnover on microtubule plus ends. The Journal of cell biology. 180:729-737. Fenn, K., and K.R. Matthews. 2007. The cell biology of Trypanosoma brucei differentiation. Current opinion in microbiology. 10:539-546. Field, H., T. Sherwin, A.C. Smith, K. Gull, and M.C. Field. 2000. Cell-cycle and developmental regulation of TbRAB31 localisation, a GTP-locked Rab protein from Trypanosoma brucei. Molecular and biochemical parasitology. 106:21-35. Gimona, M., K. Djinovic-Carugo, W.J. Kranewitter, and S.J. Winder. 2002. Functional plasticity of CH domains. FEBS letters. 513:98-106. Gull, K. 1999. The cytoskeleton of trypanosomatid parasites. Annual review of microbiology. 53:629-655. Hammarton, T.C., J. Clark, F. Douglas, M. Boshart, and J.C. Mottram. 2003. Stage-specific differences in cell cycle control in Trypanosoma brucei revealed by RNA interference of a mitotic cyclin. The Journal of biological chemistry. 278:22877-22886. 67 Hayashi, I., and M. Ikura. 2003. Crystal structure of the amino-terminal microtubule-binding domain of end-binding protein 1 (EB1). The Journal of biological chemistry. 278:36430-36434. Hayashi, I., A. Wilde, T.K. Mal, and M. Ikura. 2005. Structural basis for the activation of microtubule assembly by the EB1 and p150Glued complex. Molecular cell. 19:449460. He, C.Y., H.H. Ho, J. Malsam, C. Chalouni, C.M. West, E. Ullu, D. Toomre, and G. Warren. 2004. Golgi duplication in Trypanosoma brucei. The Journal of cell biology. 165:313321. Honnappa, S., S.M. Gouveia, A. Weisbrich, F.F. Damberger, N.S. Bhavesh, H. Jawhari, I. Grigoriev, F.J. van Rijssel, R.M. Buey, A. Lawera, I. Jelesarov, F.K. Winkler, K. Wuthrich, A. Akhmanova, and M.O. Steinmetz. 2009. An EB1-binding motif acts as a microtubule tip localization signal. Cell. 138:366-376. Honnappa, S., C.M. John, D. Kostrewa, F.K. Winkler, and M.O. Steinmetz. 2005. Structural insights into the EB1-APC interaction. The EMBO journal. 24:261-269. Ismach, R., C.M. Cianci, J.P. Caulfield, P.J. Langer, A. Hein, and D. McMahon-Pratt. 1989. Flagellar membrane and paraxial rod proteins of Leishmania: characterization employing monoclonal antibodies. The Journal of protozoology. 36:617-624. Juwana, J.P., P. Henderikx, A. Mischo, A. Wadle, N. Fadle, K. Gerlach, J.W. Arends, H. Hoogenboom, M. Pfreundschuh, and C. Renner. 1999. EB/RP gene family encodes tubulin binding proteins. International journal of cancer. Journal international du cancer. 81:275-284. Katsuki, M., D.R. Drummond, M. Osei, and R.A. Cross. 2009. Mal3 masks catastrophe events in Schizosaccharomyces pombe microtubules by inhibiting shrinkage and promoting rescue. The Journal of biological chemistry. 284:29246-29250. Kelly, S., J. Reed, S. Kramer, L. Ellis, H. Webb, J. Sunter, J. Salje, N. Marinsek, K. Gull, B. Wickstead, and M. Carrington. 2007. Functional genomics in Trypanosoma brucei: a collection of vectors for the expression of tagged proteins from endogenous and ectopic gene loci. Molecular and biochemical parasitology. 154:103-109. Kilmartin, J.V., B. Wright, and C. Milstein. 1982. Rat monoclonal antitubulin antibodies derived by using a new nonsecreting rat cell line. The Journal of cell biology. 93:576582. Kim, J., S. Sim, K. Song, T.S. Yong, and S.J. Park. 2008. Giardia lamblia EB1 is a functional homolog of yeast Bim1p that binds to microtubules. Parasitology international. 57:465-471. Kita, K., T. Wittmann, I.S. Nathke, and C.M. Waterman-Storer. 2006. Adenomatous polyposis coli on microtubule plus ends in cell extensions can promote microtubule net growth with or without EB1. Molecular biology of the cell. 17:2331-2345. Kohl, L., D. Robinson, and P. Bastin. 2003. Novel roles for the flagellum in cell morphogenesis and cytokinesis of trypanosomes. The EMBO journal. 22:5336-5346. Kohl, L., T. Sherwin, and K. Gull. 1999. Assembly of the paraflagellar rod and the flagellum attachment zone complex during the Trypanosoma brucei cell cycle. The Journal of eukaryotic microbiology. 46:105-109. Komarova, Y., C.O. De Groot, I. Grigoriev, S.M. Gouveia, E.L. Munteanu, J.M. Schober, S. Honnappa, R.M. Buey, C.C. Hoogenraad, M. Dogterom, G.G. Borisy, M.O. Steinmetz, and A. Akhmanova. 2009. Mammalian end binding proteins control persistent microtubule growth. The Journal of cell biology. 184:691-706. Komarova, Y., G. Lansbergen, N. Galjart, F. Grosveld, G.G. Borisy, and A. Akhmanova. 2005. EB1 and EB3 control CLIP dissociation from the ends of growing microtubules. Molecular biology of the cell. 16:5334-5345. 68 Korinek, W.S., M.J. Copeland, A. Chaudhuri, and J. Chant. 2000. Molecular linkage underlying microtubule orientation toward cortical sites in yeast. Science (New York, N.Y.). 287:2257-2259. Lacomble, S., S. Vaughan, C. Gadelha, M.K. Morphew, M.K. Shaw, J.R. McIntosh, and K. Gull. 2009. Three-dimensional cellular architecture of the flagellar pocket and associated cytoskeleton in trypanosomes revealed by electron microscope tomography. Journal of cell science. 122:1081-1090. Lacomble, S., S. Vaughan, C. Gadelha, M.K. Morphew, M.K. Shaw, J.R. McIntosh, and K. Gull. 2010. Basal body movements orchestrate membrane organelle division and cell morphogenesis in Trypanosoma brucei. Journal of cell science. 123:2884-2891. LaCount, D.J., S. Bruse, K.L. Hill, and J.E. Donelson. 2000. Double-stranded RNA interference in Trypanosoma brucei using head-to-head promoters. Molecular and biochemical parasitology. 111:67-76. Landgraf, D., B. Okumus, P. Chien, T.A. Baker, and J. Paulsson. 2012. Segregation of molecules at cell division reveals native protein localization. Nature methods. 9:480482. Lansbergen, G., and A. Akhmanova. 2006. Microtubule plus end: a hub of cellular activities. Traffic (Copenhagen, Denmark). 7:499-507. Lee, L., J.S. Tirnauer, J. Li, S.C. Schuyler, J.Y. Liu, and D. Pellman. 2000. Positioning of the mitotic spindle by a cortical-microtubule capture mechanism. Science (New York, N.Y.). 287:2260-2262. Li, Z., and C.C. Wang. 2003. A PHO80-like cyclin and a B-type cyclin control the cell cycle of the procyclic form of Trypanosoma brucei. The Journal of biological chemistry. 278:20652-20658. Ligon, L.A., S.S. Shelly, M. Tokito, and E.L. Holzbaur. 2003. The microtubule plus-end proteins EB1 and dynactin have differential effects on microtubule polymerization. Molecular biology of the cell. 14:1405-1417. Ligon, L.A., S.S. Shelly, M.K. Tokito, and E.L. Holzbaur. 2006. Microtubule binding proteins CLIP-170, EB1, and p150Glued form distinct plus-end complexes. FEBS letters. 580:1327-1332. Louie, R.K., S. Bahmanyar, K.A. Siemers, V. Votin, P. Chang, T. Stearns, W.J. Nelson, and A.I. Barth. 2004. Adenomatous polyposis coli and EB1 localize in close proximity of the mother centriole and EB1 is a functional component of centrosomes. Journal of cell science. 117:1117-1128. Manna, T., S. Honnappa, M.O. Steinmetz, and L. Wilson. 2008. Suppression of microtubule dynamic instability by the +TIP protein EB1 and its modulation by the CAP-Gly domain of p150glued. Biochemistry. 47:779-786. Mathur, J., N. Mathur, B. Kernebeck, B.P. Srinivas, and M. Hulskamp. 2003. A novel localization pattern for an EB1-like protein links microtubule dynamics to endomembrane organization. Current biology : CB. 13:1991-1997. Matthews, K.R. 2005. The developmental cell biology of Trypanosoma brucei. Journal of cell science. 118:283-290. Matthews, K.R., J.R. Ellis, and A. Paterou. 2004. Molecular regulation of the life cycle of African trypanosomes. Trends in parasitology. 20:40-47. Matthews, K.R., and K. Gull. 1994. Evidence for an interplay between cell cycle progression and the initiation of differentiation between life cycle forms of African trypanosomes. The Journal of cell biology. 125:1147-1156. Maurer, S.P., F.J. Fourniol, G. Bohner, C.A. Moores, and T. Surrey. 2012. EBs recognize a nucleotide-dependent structural cap at growing microtubule ends. Cell. 149:371382. 69 Meissner, M., C. Agop-Nersesian, and W.J. Sullivan, Jr. 2007. Molecular tools for analysis of gene function in parasitic microorganisms. Applied microbiology and biotechnology. 75:963-975. Mimori-Kiyosue, Y., N. Shiina, and S. Tsukita. 2000. The dynamic behavior of the APC-binding protein EB1 on the distal ends of microtubules. Current biology : CB. 10:865-868. Moreira-Leite, F.F., T. Sherwin, L. Kohl, and K. Gull. 2001. A trypanosome structure involved in transmitting cytoplasmic information during cell division. Science (New York, N.Y.). 294:610-612. Morris, J.C., Z. Wang, M.E. Drew, and P.T. Englund. 2002. Glycolysis modulates trypanosome glycoprotein expression as revealed by an RNAi library. The EMBO journal. 21:44294438. Morrison, E.E., B.N. Wardleworth, J.M. Askham, A.F. Markham, and D.M. Meredith. 1998. EB1, a protein which interacts with the APC tumour suppressor, is associated with the microtubule cytoskeleton throughout the cell cycle. Oncogene. 17:3471-3477. Morriswood, B., C.Y. He, M. Sealey-Cardona, J. Yelinek, M. Pypaert, and G. Warren. 2009. The bilobe structure of Trypanosoma brucei contains a MORN-repeat protein. Molecular and biochemical parasitology. 167:95-103. Motyka, S.A., and P.T. Englund. 2004. RNA interference for analysis of gene function in trypanosomatids. Current opinion in microbiology. 7:362-368. Nozaki, T., P.A. Haynes, and G.A. Cross. 1996. Characterization of the Trypanosoma brucei homologue of a Trypanosoma cruzi flagellum-adhesion glycoprotein. Molecular and biochemical parasitology. 82:245-255. Ogbadoyi, E.O., D.R. Robinson, and K. Gull. 2003. A high-order trans-membrane structural linkage is responsible for mitochondrial genome positioning and segregation by flagellar basal bodies in trypanosomes. Molecular biology of the cell. 14:1769-1779. Pancer, Z., E.L. Cooper, and W.E. Muller. 1996. A urochordate putative homolog of human EB1, the protein which binds APC1. Cancer letters. 109:155-160. Pedersen, L.B., S. Geimer, R.D. Sloboda, and J.L. Rosenbaum. 2003. The Microtubule plus end-tracking protein EB1 is localized to the flagellar tip and basal bodies in Chlamydomonas reinhardtii. Current biology : CB. 13:1969-1974. Ploubidou, A., D.R. Robinson, R.C. Docherty, E.O. Ogbadoyi, and K. Gull. 1999. Evidence for novel cell cycle checkpoints in trypanosomes: kinetoplast segregation and cytokinesis in the absence of mitosis. Journal of cell science. 112 ( Pt 24):4641-4650. Polakis, P. 1997. The adenomatous polyposis coli (APC) tumor suppressor. Biochimica et biophysica acta. 1332:F127-147. Redmond, S., J. Vadivelu, and M.C. Field. 2003. RNAit: an automated web-based tool for the selection of RNAi targets in Trypanosoma brucei. Molecular and biochemical parasitology. 128:115-118. Rehberg, M., and R. Graf. 2002. Dictyostelium EB1 is a genuine centrosomal component required for proper spindle formation. Molecular biology of the cell. 13:2301-2310. Rindisbacher, L., A. Hemphill, and T. Seebeck. 1993. A repetitive protein from Trypanosoma brucei which caps the microtubules at the posterior end of the cytoskeleton. Molecular and biochemical parasitology. 58:83-96. Robinson, D.R., and K. Gull. 1991. Basal body movements as a mechanism for mitochondrial genome segregation in the trypanosome cell cycle. Nature. 352:731-733. Robinson, D.R., T. Sherwin, A. Ploubidou, E.H. Byard, and K. Gull. 1995. Microtubule polarity and dynamics in the control of organelle positioning, segregation, and cytokinesis in the trypanosome cell cycle. The Journal of cell biology. 128:1163-1172. Roditi, I., and M.J. Lehane. 2008. Interactions between trypanosomes and tsetse flies. Current opinion in microbiology. 11:345-351. 70 Rogers, S.L., G.C. Rogers, D.J. Sharp, and R.D. Vale. 2002. Drosophila EB1 is important for proper assembly, dynamics, and positioning of the mitotic spindle. The Journal of cell biology. 158:873-884. Rosario, V. 1981. Cloning of naturally occurring mixed infections of malaria parasites. Science (New York, N.Y.). 212:1037-1038. Rotureau, B., I. Subota, and P. Bastin. 2011. Molecular bases of cytoskeleton plasticity during the Trypanosoma brucei parasite cycle. Cellular microbiology. 13:705-716. Ruben, L., C. Egwuagu, and C.L. Patton. 1983. African trypanosomes contain calmodulin which is distinct from host calmodulin. Biochimica et biophysica acta. 758:104-113. Sandblad, L., K.E. Busch, P. Tittmann, H. Gross, D. Brunner, and A. Hoenger. 2006. The Schizosaccharomyces pombe EB1 homolog Mal3p binds and stabilizes the microtubule lattice seam. Cell. 127:1415-1424. Schroder, J.M., L. Schneider, S.T. Christensen, and L.B. Pedersen. 2007. EB1 is required for primary cilia assembly in fibroblasts. Current biology : CB. 17:1134-1139. Schumann Burkard, G., P. Jutzi, and I. Roditi. 2011. Genome-wide RNAi screens in bloodstream form trypanosomes identify drug transporters. Molecular and biochemical parasitology. 175:91-94. Schwartz, K., K. Richards, and D. Botstein. 1997. BIM1 encodes a microtubule-binding protein in yeast. Molecular biology of the cell. 8:2677-2691. Scott, V., T. Sherwin, and K. Gull. 1997. gamma-tubulin in trypanosomes: molecular characterisation and localisation to multiple and diverse microtubule organising centres. Journal of cell science. 110 ( Pt 2):157-168. Shapiro, S.Z., J. Naessens, B. Liesegang, S.K. Moloo, and J. Magondu. 1984. Analysis by flow cytometry of DNA synthesis during the life cycle of African trypanosomes. Acta tropica. 41:313-323. Sharma, R., E. Gluenz, L. Peacock, W. Gibson, K. Gull, and M. Carrington. 2009. The heart of darkness: growth and form of Trypanosoma brucei in the tsetse fly. Trends in parasitology. 25:517-524. Sherwin, T., and K. Gull. 1989a. The cell division cycle of Trypanosoma brucei brucei: timing of event markers and cytoskeletal modulations. Philosophical transactions of the Royal Society of London. Series B, Biological sciences. 323:573-588. Sherwin, T., and K. Gull. 1989b. Visualization of detyrosination along single microtubules reveals novel mechanisms of assembly during cytoskeletal duplication in trypanosomes. Cell. 57:211-221. Sherwin, T., A. Schneider, R. Sasse, T. Seebeck, and K. Gull. 1987. Distinct localization and cell cycle dependence of COOH terminally tyrosinolated alpha-tubulin in the microtubules of Trypanosoma brucei brucei. The Journal of cell biology. 104:439446. Shi, H., A. Djikeng, T. Mark, E. Wirtz, C. Tschudi, and E. Ullu. 2000. Genetic interference in Trypanosoma brucei by heritable and inducible double-stranded RNA. RNA (New York, N.Y.). 6:1069-1076. Simarro, P.P., J. Jannin, and P. Cattand. 2008. Eliminating human African trypanosomiasis: where do we stand and what comes next? PLoS medicine. 5:e55. Slep, K.C., S.L. Rogers, S.L. Elliott, H. Ohkura, P.A. Kolodziej, and R.D. Vale. 2005. Structural determinants for EB1-mediated recruitment of APC and spectraplakins to the microtubule plus end. The Journal of cell biology. 168:587-598. Slep, K.C., and R.D. Vale. 2007. Structural basis of microtubule plus end tracking by XMAP215, CLIP-170, and EB1. Molecular cell. 27:976-991. Su, L.K., M. Burrell, D.E. Hill, J. Gyuris, R. Brent, R. Wiltshire, J. Trent, B. Vogelstein, and K.W. Kinzler. 1995. APC binds to the novel protein EB1. Cancer research. 55:2972-2977. 71 Su, L.K., and Y. Qi. 2001. Characterization of human MAPRE genes and their proteins. Genomics. 71:142-149. Tirnauer, J.S., and B.E. Bierer. 2000. EB1 proteins regulate microtubule dynamics, cell polarity, and chromosome stability. The Journal of cell biology. 149:761-766. Tirnauer, J.S., S. Grego, E.D. Salmon, and T.J. Mitchison. 2002. EB1-microtubule interactions in Xenopus egg extracts: role of EB1 in microtubule stabilization and mechanisms of targeting to microtubules. Molecular biology of the cell. 13:3614-3626. Tirnauer, J.S., E. O'Toole, L. Berrueta, B.E. Bierer, and D. Pellman. 1999. Yeast Bim1p promotes the G1-specific dynamics of microtubules. The Journal of cell biology. 145:993-1007. Van Den Abbeele, J., Y. Claes, D. van Bockstaele, D. Le Ray, and M. Coosemans. 1999. Trypanosoma brucei spp. development in the tsetse fly: characterization of the postmesocyclic stages in the foregut and proboscis. Parasitology. 118 ( Pt 5):469-478. Vassella, E., B. Reuner, B. Yutzy, and M. Boshart. 1997. Differentiation of African trypanosomes is controlled by a density sensing mechanism which signals cell cycle arrest via the cAMP pathway. Journal of cell science. 110 ( Pt 21):2661-2671. Vaughan, S., L. Kohl, I. Ngai, R.J. Wheeler, and K. Gull. 2008. A repetitive protein essential for the flagellum attachment zone filament structure and function in Trypanosoma brucei. Protist. 159:127-136. Vickerman, K. 1985. Developmental cycles and biology of pathogenic trypanosomes. British medical bulletin. 41:105-114. Vickerman, K., L. Tetley, K.A. Hendry, and C.M. Turner. 1988. Biology of African trypanosomes in the tsetse fly. Biology of the cell / under the auspices of the European Cell Biology Organization. 64:109-119. Vitre, B., F.M. Coquelle, C. Heichette, C. Garnier, D. Chretien, and I. Arnal. 2008. EB1 regulates microtubule dynamics and tubulin sheet closure in vitro. Nature cell biology. 10:415-421. Wang, Z., J.C. Morris, M.E. Drew, and P.T. Englund. 2000. Inhibition of Trypanosoma brucei gene expression by RNA interference using an integratable vector with opposing T7 promoters. The Journal of biological chemistry. 275:40174-40179. Weisbrich, A., S. Honnappa, R. Jaussi, O. Okhrimenko, D. Frey, I. Jelesarov, A. Akhmanova, and M.O. Steinmetz. 2007. Structure-function relationship of CAP-Gly domains. Nature structural & molecular biology. 14:959-967. Wen, Y., C.H. Eng, J. Schmoranzer, N. Cabrera-Poch, E.J. Morris, M. Chen, B.J. Wallar, A.S. Alberts, and G.G. Gundersen. 2004. EB1 and APC bind to mDia to stabilize microtubules downstream of Rho and promote cell migration. Nature cell biology. 6:820-830. Wirtz, E., S. Leal, C. Ochatt, and G.A. Cross. 1999. A tightly regulated inducible expression system for conditional gene knock-outs and dominant-negative genetics in Trypanosoma brucei. Molecular and biochemical parasitology. 99:89-101. Woodward, R., and K. Gull. 1990. Timing of nuclear and kinetoplast DNA replication and early morphological events in the cell cycle of Trypanosoma brucei. Journal of cell science. 95 ( Pt 1):49-57. Zhou, Q., B. Liu, Y. Sun, and C.Y. He. 2011. A coiled-coil- and C2-domain-containing protein is required for FAZ assembly and cell morphology in Trypanosoma brucei. Journal of cell science. 124:3848-3858. Zhu, Z. 2011. Probing Interactions Between Eb1, Microtubules and Actin. In Chemistry and Biochemistry. Vol. Doctor of Philosophy. University of Notre Dame. Ziegelbauer, K., M. Quinten, H. Schwarz, T.W. Pearson, and P. Overath. 1990. Synchronous differentiation of Trypanosoma brucei from bloodstream to procyclic forms in vitro. European journal of biochemistry / FEBS. 192:373-378. 72 73 [...]... buffer 5 INTRODUCTION 1.1 An overview of Trypanosoma brucei 1.1.1 T brucei: Ecological, Economic and Political impact The African trypanosome, Trypanosoma brucei, is the protozoan parasite responsible for the African sleeping sickness in 36 countries of sub-Saharan Africa, many of which fall in the category of the poorest developing nations in the world Many affected populations live beyond the reach of. .. analysis of EB1 RNAi induction 46 FIGURE 14 Anti-TbEB1 immune serum is non-specific in its detection of TbEB1 49 2 FIGURE 15 Western blot analysis of 29.13 (control) cells and PXS2YFPEB1 cells using affinity-purified anti-TbEB1 50 FIGURE 16 Co-staining YTAT cells with anti-TbEB1 and YL1/2 antibody confirmed a temporally-modulated EB1 localization at the posterior tip of the cell body 51 FIGURE 17 Co-staining... are in the G1/S phase, while those with two kinetoplasts and a single nucleus (2K1N) indicate that the cells are in the G2/M phase Cells bearing segregated kinetoplasts and nuclei (2K2N) are on the verge of cytokinesis (Sherwin and Gull, 1989a; Woodward and Gull, 1990) In the same manner, antibodies have been raised against several key parasite organelles and proteins, and immunostaining using these antibodies... series of faint lines undergirding the flagella 13 1.2 An overview of End-Binding protein 1 (EB1) 1.2.1 EB1 homologues The EB family comprises a group of microtubule plus-end tracking proteins (+TIPs) which have been evolutionarily conserved and studied in organisms ranging from yeast to humans The first characterized member of the family, human EB1, was identified in a yeast-twohybrid screen for interacting... conservation and domain preservation, 2 Established EB1 localization within the T brucei cell, 3 Scrutinized EB1 localization within the context of the T brucei cell cycle via ectopic introduction of the YFP -EB1 fusion gene, 4 Attempted to characterize the EB1 RNAi phenotype in order to better understand EB1 function in the parasite, and 5 Obtained and purified an anti-TbEB1 antibody specifically raised against... YFP -EB1 localization 61 FIGURE 20 Anti-TbEB1 antibody labelling 24 hours post RNAi induction in a YFP -EB1 over-expression background 61 3 LIST OF ABBREVIATIONS +TIP plus-end tracking protein aa amino acids ABS actin binding site APC adenomatous polyposis coli CAP-Gly glycine-rich cytoskeleton-associated protein CH calponin homology (domain) DAPI 4, 6-diamidino-2-phenylindole EB end-binding protein EBH... assay conditions, but the mechanisms employed by EB1 in its role as a regulator of microtubule dynamics still remain the subject of intense discussion 1.3 Why study EB1 in T brucei? EB1 is known to localize directly to the plus-ends tips of growing microtubules, recruiting other +TIPs in the process and itself forming the core of fast-changing +TIP complexes (Akhmanova and Steinmetz, 2008) which dynamically... actin makes it an ideal model in which to study the effects of EB1 and the mechanisms by which they are exerted on the highly-regulated dynamics of the microtubule network -knowledge 20 crucial for a deeper understanding of parasite behaviour as well as of the mechanisms underlying EB1 function, form and interaction It is interesting to note that although EB1 has been discovered and studied in organisms... plus-end tracking protein which recruits multiple distinct +TIPs and itself forms the core for various protein complexes that form at dynamic microtubule plus ends 16 (Lansbergen and Akhmanova, 2006) The budding yeast EB1 homologue Bim1p, for instance, binds a protein complex containing Kar9 and Myo2p, resulting in the cortical capture of microtubules which facilitates orientation of the spindle towards... flagella tip of Chlamydomonas reinhardtii, and depletion of EB1 is accompanied by accumulation of intraflagellar transport (IFT) particles near the flagella tip (Pedersen et al., 2003) 1.2.6 Putative mechanisms of EB1 cellular interaction Years of study have made it clear that EB1 plays a major role in regulating microtubule dynamics both in vivo and in vitro systems, although opinions differ as to EB1' s ... BioImaging Sciences for his assistance in purifying His -EB1 protein for antibody generation; his expertise was invaluable, and his kindness in answering my generally numerous and sometimes inane... An overview of End-binding protein (EB1) 14 1.2.1 EB1 homologues 14 1.2.2 EB1 domain organization 14 1.2.3 EB1 cellular localization 16 1.2.4 EB1 as a keystone +TIP protein 16 1.2.5 Role of EB1. .. labelling of EB1 in a YFP -EB1 over-expressing cell line confirmed that the anti-TbEB1 antibody labelling pattern was identical to YFP -EB1 localization 61 FIGURE 20 Anti-TbEB1 antibody labelling