Institute of Immunology and Infection Research, School of Biological Sciences, University of Edinburgh, West Mains Road, Edinburgh, EH9 3JT, UK
(e-mail: keith.matthews{at}ed.ac.uk)
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The organization of cell structure is fundamental to the trypanosome throughout its life cycle. Modulations of the overall cell morphology and positioning of the specialized mitochondrial genome, flagellum and associated basal body provide the classical descriptions of the different life cycle stages of the parasite. The dependency relationships that govern these morphological changes are now beginning to be understood and their molecular basis identified. The overall picture emerging is of a highly organized cell in which the rules established for cell division and morphogenesis in organisms such as yeast and mammalian cells do not necessarily apply. Therefore, understanding the developmental cell biology of the African trypanosome is providing insight into both fundamentally conserved and fundamentally different aspects of the organization of the eukaryotic cell.
Key words: Morphology, Cell division, Differentiation, Life cycle
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The parasitic protozoa are excellent examples of organisms that display extreme adaptation to their environment, in many cases because they must evade the immune response of a mammalian host. They also exhibit complex life cycles, frequently being transmitted between mammalian hosts by arthropod vectors, in which they often also face similarly hostile conditions. The variety of different conditions encountered means that a single organism (or its offspring) must show both adaptability and the capacity for rigorously programmed differentiation. Occurring within a single cell, these developmental steps require coordinated modulation of many basic biological processes. Here, I discuss recent advances in our understanding of the basic aspects of the cell biology of the parasite Trypanosoma brucei and place these into the context of its life cycle.
![]() |
Background |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The trypanosome is transmitted between mammalian hosts by the tsetse fly, Glossina spp, in which it initially establishes in the midgut after a bloodmeal but then migrates to the salivary glands in preparation for transmission to a new mammalian host (Fig. 1). In mammals, the parasite survives free in the bloodstream, being able to evade antibody responses through antigenic variation (McCulloch, 2004; Pays et al., 2004
). This entails the sequential expression of antigenically distinct variable surface glycoproteins (VSGs), which are linked to the surface membrane by a glycosylphosphatidylinositol (GPI) anchor. Trypanosomes proliferate in the mammalian bloodstream as morphologically slender forms, these being replaced by non-proliferative stumpy forms as parasite numbers increase (Matthews et al., 2004
). This serves two purposes. First, the accumulation of division-arrested forms limits the increase in parasite numbers and thereby prolongs host survival (and hence the probability of disease transmission). Second, the uniform arrest of stumpy forms in G1 phase of the cell cycle ensures that the morphological changes that occur upon transmission to the tsetse fly can be coordinated with re-entry into the cell cycle. This is important because correct organelle positioning is crucial for successful completion of the cell cycle of tsetse midgut procyclic forms.
|
Upon uptake by the tsetse, bloodstream trypanosomes replace the VSG coat with a less-dense surface coat composed of procyclins, which are also GPI anchored (Roditi and Liniger, 2002). Energy generation also switches from being exclusively based on glycolysis to a mitochondrion-based respiratory system, which requires structural elaboration and metabolic activation of the organelle. After proliferation in the tsetse midgut, the parasite migrates to the salivary gland. The epimastigote forms generated there attach to the gland wall through elaborations of the flagellar membrane. After further multiplication, the parasite undergoes division arrest, re-acquires a VSG coat and is released into the salivary gland lumen, in preparation for inoculation into a new mammalian host.
Most studies on the trypanosome have focused on the bloodstream and procyclic forms of the parasite. This is because these stages can be readily cultured in vitro. They are also genetically tractable: knockouts can be generated through homologous recombination, and tetracycline-regulatable ectopic expression and RNA interference (RNAi) approaches are routine. Forward genetic approaches have been more limited since trypanosomes are diploid and the generation of sexual crosses is experimentally challenging; however, the recent production of large-scale RNAi libraries (Morris et al., 2002) and the development of a mariner-based transposon mutagenesis strategy (coupled with frequent loss of heterozygosity; Leal et al., 2004
) offer exciting prospects for informative mutant screens.
![]() |
Trypanosome cell architecture |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
The most posterior structure is the mouth of the flagellar pocket. This is the exit point for the flagellum, which is tethered along the exterior length of the parasite. The flagellar pocket is the only site of endo- and exo-cytosis (Overath and Engstler, 2004); this is important in bloodstream forms, in which the surface membrane is densely packed with VSG to protect against the alternative pathway of complement activation (Ferrante and Allison, 1983
) and to shield common antigenic determinants from immune recognition. This dense packing requires that the GPI-anchored VSG is significantly concentrated during trafficking from the endoplasmic reticulum (ER) to the surface (Grunfelder et al., 2002
). Indeed, overall membrane uptake and VSG recycling at the flagellar pocket occur at a phenomenal rate in bloodstream cells, considering the restricted area of the flagellar pocket (the surface VSG pool turns over entirely within 12 minutes despite the flagellar pocket occupying only 5% of the cell surface area; Engstler et al., 2004
). Endocytosis of the VSG and other molecules in the flagellar pocket is clathrin dependent: RNAi directed against the clathrin heavy chain causes rapid death of cells after massive enlargement of the flagellar pocket (Allen et al., 2003
). A similar phenotype is seen when actin is targeted by RNAi in bloodstream forms, implicating this protein in endocytosis and intracellular trafficking (Garcia-Salcedo et al., 2004
). Endocytosis in procyclic cells occurs at a lower rate, but is also clathrin dependent (Allen et al., 2003
; Hung et al., 2004
). Interestingly, however, RNAi directed against actin is not lethal in this life cycle stage (Garcia-Salcedo et al., 2004
).
The small size of the trypanosome, its high rate of trafficking of GPI-anchored VSG to the surface and the concentration of the endocytic apparatus into the posterior end of the cell have made bloodstream forms an excellent system for analysis of protein trafficking using fluorescence and electron microscopy. Moreover, the single Golgi stack of the parasite and its restricted positioning between the nucleus and flagellar pocket has enabled T. brucei to become a model for the biogenesis of this organelle in eukaryotes. Thus, He et al. have derived a new marker for the T. brucei Golgi (TbGRASP) similar to mammalian GRASP-55 and used a photobleaching approach to demonstrate that, during cell division, the new Golgi forms de novo from material sourced directly from the old Golgi, rather than from the ER (He et al., 2004).
The motility of the trypanosome is dependent upon its single flagellum, which has a conventional axonemal structure plus an associated paraflagellar rod (Vaughan and Gull, 2003). This is a semi-rigid structure found in the kinetoplastids and euglenoids that contributes to parasite motility (Bastin et al., 1998
), perhaps assisting trypanosome flagellar beat efficiency in the viscous mammalian bloodstream. Trypanin (Hutchings et al., 2002
), a trypanosome protein related to a subunit of the dynein motor regulatory complex (PF2 in Chlamydomonas and Gas11/Gas 8 in mammalian cells), also contributes to motility (Rupp and Porter, 2003
). Trypanin contains a microtubule-binding domain and appears to stabilize the interaction between the flagellum and the subpellicular cytoskeleton, in a region called the flagellum-attachment zone (Kohl et al., 1999
). This binding seems to impart directional motility on the parasite, since RNAi directed against trypanin causes uncontrolled tumbling, which contrasts with the paralysis induced by disruption of the paraflagellar rod. As well as assisting an understanding of motility, the study of flagellar proteins is also proving valuable for the dissection of the intraflagellar transport machinery, for which the tools available for gene function analysis make T. brucei an excellent model (Ersfeld and Gull, 2001
; Kohl et al., 2003
).
During cell division, the growing daughter flagellum precisely tracks the old flagellum, such that structural information is transferred from the old to the new flagellum through a novel example of cytotaxic inheritance (Moreira-Leite et al., 2001). This information is imparted through the flagellar connector, a mobile structure that connects the tip of the new flagellum with three of the doublet microtubules of the axoneme of the old flagellum (Briggs et al., 2004
). Although it is apparently fundamental to the division of the procyclic form, there is no evidence for the existence of a flagellar connector in the bloodstream stage of T. brucei, nor is any related structure obvious in other kinetoplastids (Briggs et al., 2004
). This surprising finding highlights very basic differences in cell-cycle control for different life cycle stages of trypanosomes. Perhaps the procyclic trypanosome population can afford fewer mistakes in cell division as it struggles to establish a foothold in the tsetse fly.
The trypanosome flagellum originates in a basal body that is, in turn, linked through the mitochondrial membrane to the mitochondrial genome, which comprises a mass of catenated DNA termed the kinetoplast. The kinetoplast and basal body are linked by a tripartite attachment complex that must traverse both the cell and the mitochondrial membranes (Fig. 3) (Ogbadoyi et al., 2003). This comprises a series of filaments providing guide ropes through which mitochondrial genome segregation is linked to replication and segregation of the basal body and flagellum. This, in turn, is linked to the microtubule cytoskeleton, such that drugs that disrupt the cytoskeleton prevent both basal body and kinetoplast segregation (reviewed by Gull, 2003
).
|
The mitochondrion itself is a single elongated structure that runs from the posterior to the anterior of the cell. In bloodstream forms, the mitochondrion is a simple tubular structure devoid of cristae. This reflects the absence of mitochondrial respiration during this stage, energy generation being dependent on glycolytic reactions compartmentalized within specialized organelles termed glycosomes (Parsons, 2004). However, the procyclic form does not have the luxury of blood glucose as an abundant energy source and has a highly active mitochondrion. This is atypical in that an acetate:succinate CoA transferase and succinyl-CoA synthetase cycle is present (Bochud-Allemann and Schneider, 2002
; van Weelden et al., 2003
), in addition to the components of the Krebs cycle and the electron transport chain. Acetate:succinate CoA transferase, which also operates in the anaerobic mitochondria of some metazoa and anaerobic protists but not mammals, generates ATP by the conversion of acetyl coA to acetate. As in the case of other single-copy organelles, the mitochondrion must segregate with fidelity during cell division, and one contributor to this appears to be a dynamin-like protein (TbDLP), which is involved in the division of the mitochondrial membrane either directly or through recruitment of other effectors of membrane scission (Morgan et al., 2004
). Interestingly, this protein is encoded by the only gene of this family identified in the T. brucei genome, which excludes a role for T. brucei dynamin-like proteins in intracellular trafficking - a common role for such GTPases in other eukaryotic cells.
![]() |
Cell division and organelle positioning |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The most obvious morphological difference between the different life cycle stages of the trypanosome is the position of the kinetoplast relative to the posterior end of the cell. In bloodstream forms, the kinetoplast lies close to the posterior end of the cell, and each daughter kinetoplast remains in this region throughout the cell cycle until cytokinesis. By contrast, the kinetoplast lies midway between the cell nucleus and posterior in procyclic forms; in epimastigote forms (in the tsetse salivary gland), the kinetoplast is anterior to the central nucleus (Fig. 4). The reasons for these shape changes during the life cycle are completely unknown, although they are clearly required to establish the cell architecture necessary for cell division of each life cycle stage. Perhaps an increased length of attached flagellum along the cell body assists motility in bloodstream forms, whereas the longer anterior flagellum aids substrate attachment of the epimastigote stage.
|
The mechanics of kinetoplast repositioning have been studied during differentiation between bloodstream and procyclic forms and have been found to comprise two components. Most importantly, the posterior end of the cell grows out by 3 µm through polar extension of the microtubule cytoskeleton. The kinetoplast itself also moves towards the nucleus as DNA replicates during differentiation of non-proliferative stumpy forms to procyclic forms (Matthews et al., 1995
).
Several molecules that can influence this aspect of the trypanosome cell in a stage-specific manner have been identified. The first, TbZFP2 (Hendriks et al., 2001), was identified as a member of a novel family of small proteins possessing a CX8CX5CX3H zinc finger, which is predicted to bind RNA. Although uniformly expressed during the life cycle, TbZFP2 generates a procyclic, stage-specific morphological phenotype - `nozzle formation' - when ectopically overexpressed (Fig. 5). This displays expansion of the kinetoplast-posterior dimension of the cell, in some cases leading to cells twice their normal length. The cells also accumulate in G1 phase, either because TbZFP2 overexpression perturbs the cell-cycle machinery, or because the resulting cytoskeletal disruption prevents cell-cycle progression. Overexpression of a related small CCCH zinc-finger protein (TbZFP3) produces a similar procyclic-specific phenotype (A. Paterou and K.R.M., unpublished), as does RNAi directed against CYC2, a trypanosome cyclin (Hammarton et al., 2004
; Li and Wang, 2003
). However, in these cases, the cell-cycle arrest is less stringent, and nozzled cells are able to segregate their kinetoplasts.
|
Although it is likely that some mutants demonstrating the nozzle phenotype are a consequence of cell-cycle defects or secondary perturbations in the cell, recent evidence suggests that there is a relationship between nozzle formation and the normal process of kinetoplast repositioning during differentiation. Thus, a third small CCCH molecule, TbZFP1, is transiently expressed during synchronous differentiation to procyclic forms when the kinetoplast is normally repositioned (Hendriks et al., 2001). Significantly, knocking out this gene specifically compromises kinetoplast repositioning during differentiation, whereas ectopic expression of a rescue copy restores it (E. Hendriks and K.R.M., unpublished). The TbZFP proteins are small molecules (<150 residues in length) that contain both RNA- and protein-binding motifs. It is expected, therefore, that these proteins act in concert with other (currently unidentified) proteins to govern mRNA abundance or translational capacity. Their stage-specific effects indicate a role in differential gene expression.
Stage-specific components of the cytoskeleton have also been identified, including cytoskeleton-associated protein (CAP)15, which is enriched in bloodstream forms, and CAP17 (Vedrenne et al., 2002) and CAP5.5 (Hertz-Fowler et al., 2001
), which are both procyclic specific. These proteins do not label microtubules uniformly, each being absent on the flagellum, and CAP15 and CAP17 show reduced abundance at the posterior end of the cell, where the cytoskeleton is most dynamic. This might indicate an association with microtubule stability. Supporting this, expression of CAP15 in mammalian cells promotes resistance to microtubule depolymerization by nocodazole, whereas ectopic overexpression in trypanosomes perturbs normal cell structure and division. Molecules important in stage-specific morphology are therefore emerging, and it will be interesting to discover how their roles differ in distinct stages of the life cycle.
![]() |
Complexity in two genomes |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Ablation of one component of the editing complex, RNA ligase, reveals that RNA editing (and hence mitochondrial DNA) is required in both bloodstream and procyclic forms (Schnaufer et al., 2001). This was a surprising finding, since several bloodstream-form akinetoplastic T. brucei mutants have been selected by parasite growth in the presence of DNA-intercalating agents. Nonetheless, it is supported by the observation that RNAi directed against a component of the kinetoplast replication machinery, topoisomerase II, is lethal in bloodstream-form parasites (Timms et al., 2002
) as well as procyclic forms (Wang and Englund, 2001
). Other components of the mitochondrial DNA replication apparatus are unique to kinetoplastids, including at least six DNA polymerases (Klingbeil et al., 2002
) and a novel DNA ligase (Sinha et al., 2004
).
The nuclear genome of trypanosomes shows an equivalent complexity: T. brucei contains 11 `megabase' chromosomes and >100 50 kb minichromosomes. These minichromosomes harbour a repertoire of VSG genes, each flanked by a 177 bp repeat sequence (Wickstead et al., 2004
). Although the minichromosomes segregate with fidelity (Wickstead et al., 2003
), this is not through association with the microtubule spindle via kinetochore attachment but rather results from tracking along the microtubules that run from pole to pole (Ersfeld and Gull, 1997
). The megabase chromosomes have a more conventional organization and segregation, although their gene organization is not. The genes are arranged in polycistronic arrays that have distant upstream (as yet unidentified) promoters. These gene arrays are not organized into operons, but instead genes that are differentially expressed through the life cycle can be adjacent. Stage-regulated gene expression is almost exclusively controlled at the post-transcriptional level, through differential RNA processing (all genes are trans-spliced; only one gene that has a cis intron has been identified; Mair et al., 2000
), mRNA stability and translation (Clayton, 2002
).
Uniquely, the genes that encode the major surface antigens of the bloodstream and procyclic form are transcribed by RNA polymerase I (Gunzl et al., 2003). Moreover, VSG genes are always transcribed from expression sites at chromosome ends. Only one of these is active at any one time and this is apparently due to association of the telomere with an expression site body, a discrete and structurally stable RNA polymerase 1 transcription factory located at the nuclear envelope (Navarro and Gull, 2001
). VSG expression control breaks down in stumpy forms as the trypanosome prepares for transmission to the procyclic form (Amiguet-Vercher et al., 2004
). In the procyclic form, two forms of procyclin are expressed: EP and GPEET. These are named for the amino acid repeat that is contained within each protein and each is functionally distinct. GPEET is essential in procyclic forms, whereas EP knockouts are viable as procyclic culture forms but infect tsetse flies poorly. Both forms are rapidly induced during differentiation; however, GPEET expression is downregulated as the differentiating cells become established. This is mediated through a response element contained within secondary structure in the 3'-untranslated region of the GPEET mRNA. This regulates mRNA stability in response to glycerol and glucose, presumably through interaction with RNA-binding proteins (Vassella et al., 2000
). Interestingly, this control operates through the activity of mitochondrial enzymes (Vassella et al., 2004
), thereby linking developmental surface antigen expression with environment sensing and metabolic activity (Morris et al., 2002
). Dissecting the chain of interactions between the external environment and gene expression will be key to understanding the control of the life cycle. The limited importance of transcription initiation and transcription factors in regulated gene expression suggests that there is a novel complexity to the post-transcriptional mechanisms governing developmental processes in trypanosomes.
![]() |
The trypanosome life cycle: interconnections and integration of the whole cell |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Much less understood is the development of the trypanosome in the mammalian bloodstream and in the tsetse fly. Most experiments on bloodstream forms have focused on laboratory-adapted strains that grow uncontrolled in mammalian hosts (or in culture) without developing division-arrested stumpy forms. However, the transition between slender and stumpy forms poses many interesting problems of trypanosome biology that are still to be addressed. For example, cell division arrest and the transition from slender to stumpy forms are believed to be triggered by a parasite-derived signalling molecule, stumpy-induction factor (SIF; Vassella et al., 1997). However, although this is known to operate through a cyclic-AMP-dependent pathway, the identity of this molecule is unknown. Similarly, why do all trypanosomes not undergo irreversible transition to stumpy forms in the blood when the vast majority have been triggered to do so? This might reflect the fact that the bloodstream pool of trypanosomes is not uniformly exposed to SIF (for example, as a consequence of being sequestered in the tissues or lymphatic system) or that a subpopulation of parasites is refractory to SIF. These questions are interesting not only from the standpoint of trypanosome biology and the biology of cell fate and developmental decisions but also because understanding the generation of transmission-competent stumpy forms could provide new strategies for blocking trypanosome spread.
![]() |
Conclusions and perspectives |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Clearly, T. brucei continues to provide interesting insights into the basic and unusual workings of an evolutionarily ancient eukaryote. However, its contributions to cell biology are not merely interesting oddities or evolutionary fossils of the processes found in higher eukaryotes. Rather, the currently available armoury of genetic tools, molecular markers and tractable biology makes these organisms an excellent model in their own right for addressing fundamental questions of broad interest and applicability. Notwithstanding this, the potential for using this knowledge to tackle a hugely important disease gives this work special relevance.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Allen, C. L., Goulding, D. and Field, M. C. (2003). Clathrin-mediated endocytosis is essential in Trypanosoma brucei. EMBO J. 22, 4991-5002.
Amiguet-Vercher, A., Perez-Morga, D., Pays, A., Poelvoorde, P., van Xong, H., Tebabi, P., Vanhamme, L. and Pays, E. (2004). Loss of the mono-allelic control of the VSG expression sites during the development of Trypanosoma brucei in the bloodstream. Mol. Microbiol. 51, 1577-1588.[CrossRef][Medline]
Bastin, P., Sherwin, T. and Gull, K. (1998). Paraflagellar rod is vital for trypanosome motility. Nature 391, 548.[CrossRef][Medline]
Bochud-Allemann, N. and Schneider, A. (2002). Mitochondrial substrate level phosphorylation is essential for growth of procyclic Trypanosoma brucei. J. Biol. Chem 277, 32849-32854.
Briggs, L. J., McKean, P. G., Baines, A., Moreira-Leite, F., Davidge, J., Vaughan, S. and Gull, K. (2004). The flagella connector of Trypanosoma brucei: an unusual mobile transmembrane junction. J. Cell Sci. 117, 1641-1651.
Clayton, C. E. (2002). Life without transcriptional control? From fly to man and back again. EMBO J. 21, 1881-1888.
Engstler, M., Thilo, L., Weise, F., Grunfelder, C. G., Schwarz, H., Boshart, M. and Overath, P. (2004). Kinetics of endocytosis and recycling of the GPI-anchored variant surface glycoprotein in Trypanosoma brucei. J. Cell Sci. 117, 1105-1115.
Ersfeld, K. and Gull, K. (1997). Partitioning of large and minichromosomes in Trypanosoma brucei. Science 276, 611-614.
Ersfeld, K. and Gull, K. (2001). Targeting of cytoskeletal proteins to the flagellum of Trypanosoma brucei. J. Cell Sci. 114, 141-148.
Feagin, J. E. and Stuart, K. (1988). Developmental aspects of uridine addition within mitochondrial transcripts of Trypanosoma brucei. Mol. Cell. Biol. 8, 1259-1265.[Medline]
Ferrante, A. and Allison, A. C. (1983). Alternative pathway activation of complement by African trypanosomes. Parasite Immunol. 5, 491-498.[Medline]
Garcia-Salcedo, J. A., Perez-Morga, D., Gijon, P., Dilbeck, V., Pays, E. and Nolan, D. P. (2004). A differential role for actin during the life cycle of Trypanosoma brucei. EMBO J. 23, 780-789.
Gibson, W. (2001). Sex and evolution in trypanosomes. Int. J. Parasitol. 31, 643-647.[Medline]
Grunfelder, C. G., Engstler, M., Weise, F., Schwarz, H., Stierhof, Y. D., Boshart, M. and Overath, P. (2002). Accumulation of a GPI-anchored protein at the cell surface requires sorting at multiple intracellular levels. Traffic 3, 547-559.[CrossRef][Medline]
Grunfelder, C. G., Engstler, M., Weise, F., Schwarz, H., Stierhof, Y. D., Morgan, G. W., Field, M. C. and Overath, P. (2003). Endocytosis of a glycosylphosphatidylinositol-anchored protein via clathrin-coated vesicles, sorting by default in endosomes, and exocytosis via RAB11-positive carriers. Mol. Biol. Cell 14, 2029-2040.
Gull, K. (2003). Host-parasite interactions and trypanosome morphogenesis: a flagellar pocketful of goodies. Curr. Opin. Microbiol. 6, 365-370.[CrossRef][Medline]
Gunzl, A., Bruderer, T., Laufer, G., Schimanski, B., Tu, L. C., Chung, H. M., Lee, P. T. and Lee, M. G. (2003). RNA polymerase I transcribes procyclin genes and variant surface glycoprotein gene expression sites in Trypanosoma brucei. Eukaryot. Cell 2, 542-551.
Hammarton, T. C., Clark, J., Douglas, F., Boshart, M. and Mottram, J. C. (2003). Stage-specific differences in cell cycle control in Trypanosoma brucei revealed by RNA interference of a mitotic cyclin. J. Biol. Chem. 278, 22877-22886.
Hammarton, T. C., Engstler, M. and Mottram, J. C. (2004). The Trypanosoma brucei cyclin, CYC2, is required for cell cycle progression through G1 phase and for maintenance of procyclic form cell morphology. J. Biol. Chem 279, 24757-24764.
He, C. Y., Ho, H. H., Malsam, J., Chalouni, C., West, C. M., Ullu, E., Toomre, D. and Warren, G. (2004). Golgi duplication in Trypanosoma brucei. J. Cell Biol. 165, 313-321.
Hendriks, E. F., Robinson, D. R., Hinkins, M. and Matthews, K. R. (2001). A novel CCCH protein which modulates differentiation of Trypanosoma brucei to its procyclic form. EMBO J. 20, 6700-6711.
Hertz-Fowler, C., Ersfeld, K. and Gull, K. (2001). CAP5.5, a life-cycle-regulated, cytoskeleton-associated protein is a member of a novel family of calpain-related proteins in Trypanosoma brucei. Mol. Biochem. Parasitol. 116, 25-34.[CrossRef][Medline]
Hung, C. H., Qiao, X., Lee, P. T. and Lee, M. G. (2004). Clathrin-dependent targeting of receptors to the flagellar pocket of procyclic-form Trypanosoma brucei. Eukaryot. Cell 3, 1004-1014.
Hutchings, N. R., Donelson, J. E. and Hill, K. L. (2002). Trypanin is a cytoskeletal linker protein and is required for cell motility in African trypanosomes. J. Cell Biol. 156, 867-877.
Kilmartin, J. V., Wright, B. and Milstein, C. (1982). Rat monoclonal antitubulin antibodies derived by using a new nonsecreting rat cell line. J. Cell Biol. 93, 576-582.[Abstract]
Klingbeil, M. M. and Englund, P. T. (2004). Closing the gaps in kinetoplast DNA network replication. Proc. Natl. Acad. Sci. USA 101, 4333-4334.
Klingbeil, M. M., Motyka, S. A. and Englund, P. T. (2002). Multiple mitochondrial DNA polymerases in Trypanosoma brucei. Mol. Cell 10, 175-186.[Medline]
Kohl, L., Sherwin, T. and Gull, K. (1999). Assembly of the paraflagellar rod and the flagellum attachment zone complex during the Trypanosoma brucei cell cycle. J. Eukaryot. Microbiol. 46, 105-109.[Medline]
Kohl, L., Robinson, D. and Bastin, P. (2003). Novel roles for the flagellum in cell morphogenesis and cytokinesis of trypanosomes. EMBO J. 22, 5336-5346.
Leal, S., Acosta-Serrano, A., Morris, J. and Cross, G. A. (2004). Transposon mutagenesis of Trypanosoma brucei identifies glycosylation mutants resistant to concanavalin A. J. Biol. Chem 279, 28979-28988.
Li, Z. and Wang, C. C. (2003). A PHO80-like cyclin and a B-type cyclin control the cell cycle of the procyclic form of Trypanosoma brucei. J. Biol. Chem 278, 20652-20658.
Madison-Antenucci, S., Grams, J. and Hajduk, S. L. (2002). Editing machines: the complexities of trypanosome RNA editing. Cell 108, 435-438.[Medline]
Mair, G., Shi, H. F., Li, H. J., Djikeng, A., Aviles, H. O., Bishop, J. R., Falcone, F. H., Gavrilescu, C., Montgomery, J. L., Santori, M. I. et al. (2000). A new twist in trypanosome RNA metabolism: cis-splicing of pre-mRNA. RNA 6, 163-169.
Matthews, K. R. (1999). Developments in differentiation of T. brucei. Parasitol. Today 15, 76-80.[CrossRef][Medline]
Matthews, K. R., Sherwin, T. and Gull, K. (1995). Mitochondrial genome repositioning during the differentiation of the African trypanosome between life cycle forms is microtubule mediated. J. Cell Sci. 108, 2231-2239.
Matthews, K. R., Ellis, J. R. and Paterou, A. (2004). Molecular regulation of the life cycle of African trypanosomes. Trends Parasitol. 20, 40-47.[CrossRef][Medline]
McCulloch, R. (2004). Antigenic variation in African trypanosomes: monitoring progress. Trends Parasitol. 20, 117-121.[CrossRef][Medline]
Moreira-Leite, F. F., Sherwin, T., Kohl, L. and Gull, K. (2001). A trypanosome structure involved in transmitting cytoplasmic information during cell division. Science 294, 610-612.
Morgan, G. W., Goulding, D. and Field, M. C. (2004). The single dynamin-like protein of Trypanosoma brucei regulates mitochondrial division and is not required for endocytosis. J. Biol. Chem. 279, 10692-10701.
Morris, J. C., Wang, Z., Drew, M. E. and Englund, P. T. (2002). Glycolysis modulates trypanosome glycoprotein expression as revealed by an RNAi library. EMBO J. 21, 4429-4438.
Navarro, M. and Gull, K. (2001). A pol I transcriptional body associated with VSG mono-allelic expression in Trypanosoma brucei. Nature 414, 759-763.[CrossRef][Medline]
Ogbadoyi, E. O., Robinson, D. R. and Gull, K. (2003). A high-order transmembrane structural linkage is responsible for mitochondrial genome positioning and segregation by flagellar basal bodies in trypanosomes. Mol. Biol. Cell 14, 1769-1779.
Overath, P. and Engstler, M. (2004). Endocytosis, membrane recycling and sorting of GPI-anchored proteins: Trypanosoma brucei as a model system. Mol. Microbiol. 53, 735-744.[CrossRef][Medline]
Parsons, M. (2004). Glycosomes: parasites and the divergence of peroxisomal purpose. Mol. Microbiol. 53, 717-724.[CrossRef][Medline]
Pays, E., Vanhamme, L. and Perez-Morga, D. (2004). Antigenic variation in Trypanosoma brucei: facts, challenges and mysteries. Curr. Opin. Microbiol. 7, 369-374.[CrossRef][Medline]
Ploubidou, A., Robinson, D. R., Docherty, R. C., Ogbadoyi, E. O. and Gull, K. (1999). Evidence for novel cell cycle checkpoints in trypanosomes: kinetoplast segregation and cytokinesis in the absence of mitosis. J. Cell Sci. 112, 4641-4650.
Robinson, D. R., Sherwin, T., Ploubidou, A., Byard, E. H. and Gull, K. (1995). Microtubule polarity and dynamics in the control of organelle positioning, segregation, and cytokinesis in the trypanosome cell cycle. J. Cell Biol. 128, 1163-1172.[Abstract]
Roditi, I. and Liniger, M. (2002). Dressed for success: the surface coats of insect-borne protozoan parasites. Trends Microbiol. 10, 128-134.[CrossRef][Medline]
Rupp, G. and Porter, M. E. (2003). A subunit of the dynein regulatory complex in Chlamydomonas is a homologue of a growth arrest-specific gene product. J. Cell Biol. 162, 47-57.
Schnaufer, A., Panigrahi, A. K., Panicucci, B., Igo, R. P., Jr, Wirtz, E., Salavati, R. and Stuart, K. (2001). An RNA ligase essential for RNA editing and survival of the bloodstream form of Trypanosoma brucei. Science 291, 2159-2162.
Sherwin, T. and Gull, K. (1989a). The cell division cycle of Trypanosoma brucei brucei: timing of event markers and cytoskeletal modulations. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 323, 573-588.[Medline]
Sherwin, T. and Gull, K. (1989b). Visualization of detyrosination along single microtubules reveals novel mechanisms of assembly during cytoskeletal duplication in trypanosomes. Cell 57, 211-221.[Medline]
Sinha, K. M., Hines, J. C., Downey, N. and Ray, D. S. (2004). Mitochondrial DNA ligase in Crithidia fasciculata. Proc. Natl. Acad. Sci. USA 101, 4361-4366.
Timms, M. W., van Deursen, F. J., Hendriks, E. F. and Matthews, K. R. (2002). Mitochondrial development during life cycle differentiation of African trypanosomes: evidence for a kinetoplast-dependent differentiation control point. Mol. Biol. Cell 13, 3747-3759.
van Weelden, S. W., Fast, B., Vogt, A., van der Meer, P., Saas, J., van Hellemond, J. J., Tielens, A. G. and Boshart, M. (2003). Procyclic Trypanosoma brucei do not use Krebs cycle activity for energy generation. J. Biol. Chem. 278, 12854-12863.
Vassella, E., Reuner, B., Yutzy, B. and Boshart, M. (1997). Differentiation of African trypanosomes is controlled by a density sensing mechanism which signals cell cycle arrest via the cAMP pathway. J. Cell Sci. 110, 2661-2671.
Vassella, E., den Abbeele, J. V., Butikofer, P., Renggli, C. K., Furger, A., Brun, R. and Roditi, I. (2000). A major surface glycoprotein of Trypanosoma brucei is expressed transiently during development and can be regulated post-transcriptionally by glycerol or hypoxia. Genes Dev. 14, 615-626.
Vassella, E., Probst, M., Schneider, A., Studer, E., Renggli, C. K. and Roditi, I. (2004). Expression of a major surface protein of Trypanosoma brucei insect forms is controlled by the activity of mitochondrial enzymes. Mol. Biol. Cell 16, 16.
Vaughan, S. and Gull, K. (2003). The trypanosome flagellum. J. Cell Sci. 116, 757-759.
Vedrenne, C., Giroud, C., Robinson, D. R., Besteiro, S., Bosc, C., Bringaud, F. and Baltz, T. (2002). Two related subpellicular cytoskeleton-associated proteins in Trypanosoma brucei stabilize microtubules. Mol. Biol. Cell 13, 1058-1070.
Wang, Z. and Englund, P. T. (2001). RNA interference of a trypanosome topoisomerase II causes progressive loss of mitochondrial DNA. EMBO J. 20, 4674-4683.
Wickstead, B., Ersfeld, K. and Gull, K. (2003). The mitotic stability of the minichromosomes of Trypanosoma brucei. Mol. Biochem. Parasitol. 132, 97-100.[CrossRef][Medline]
Wickstead, B., Ersfeld, K. and Gull, K. (2004). The small chromosomes of Trypanosoma brucei involved in antigenic variation are constructed around repetitive palindromes. Genome Res. 14, 1014-1024.