Skip to main content

Advertisement

African trypanosomes

Abstract

African trypanosomes cause human African trypanosomiasis and animal African trypanosomiasis. They are transmitted by tsetse flies in sub-Saharan Africa. Although most famous for their mechanisms of immune evasion by antigenic variation, there have been recent important studies that illuminate important aspects of the biology of these parasites both in their mammalian host and during passage through their tsetse fly vector. This Primer overviews current research themes focused on these parasites and discusses how these biological insights and the development of new technologies to interrogate gene function are being used in the search for new approaches to control the parasite. The new insights into the biology of trypanosomes in their host and vector highlight that we are in a ‘golden age’ of discovery for these fascinating parasites.

What are trypanosomes?

The Trypanosomatidae (Phylum Euglenozoa) are obligate parasitic protozoans, which infect all vertebrate classes, in addition to insects and plants. Trypanosomatids are uniflagellated, and like other organisms in the order Kinetoplastida, are characterised by a modified mitochondrial genome known as the kinetoplast [1, 2]. This is composed of DNA ‘mini’- and ‘maxicircles’ [3, 4], which can vary in number and catenation depending on the particular species [5]. Among the trypanosomatids, Trypanosoma is a genus of particular medical and veterinary concern [6, 7]. The Salivaria group of trypanosomes, so named for being transmitted in the infected saliva of a tsetse fly vector (Glossina spp.), is represented by Trypanosoma brucei, T. congolense and T. vivax. The former is the most well-studied of the salivarian trypanosomes, with subspecies T. b. gambiense and T. b. rhodesiense being the causative agents of human African trypanosomiases (HAT). The dixenous life-cycle of T. brucei comprises vertebrate stages and tsetse stages, which involves differentiation through a number of morphological forms, accompanied by remodelling of gene expression and metabolic processes [8, 9] (Additional file 1). In the bloodstream, two morphotypes exist, slender and stumpy forms [10, 11]. Slender forms evade the mammalian antibody response through antigenic variation, entailing expression of a variable surface glycoprotein (VSG) monolayer which covers the trypanosome cell. Changes in the expression of VSG genes generate antigenically distinct subpopulations, which can evade the host immune response, and so sustain the parasite infection [12, 13]. The transition from slender to stumpy forms is regulated by a quorum sensing (QS) type process which prolongs host survival and promotes transmission because stumpy forms are non-proliferative in the blood, but preferentially survive in and colonise the tsetse midgut.

While T. brucei is certainly a model for trypanosome biology, and indeed that of other kinetoplastids, significant variation in morphology, transmission, and life-cycle, exists between different trypanosome species. Neither T. vivax nor T. congolense, for example, (both causing animal African trypanosomiasis, AAT), show morphological differentiation in the bloodstream, though both exhibit density-dependent growth control [14, 15]. Also, T. b. equiperdum has circumvented the need for an arthropod vector altogether, and instead is sexually transmitted between equines. Exciting advances in our understanding of trypanosome biology continue to be made, in addition to the development of new tools for studying these organisms. This Primer will highlight important recent advances in this field, as well as identifying areas that warrant investigation in future.

Why study the biology of trypanosomes?

Trypanosomatids diverged from other eukaryotic lineages at least 500 million years ago and so provide a valuable model for evolutionarily distinct eukaryotic processes [16, 17]. Despite their divergence, they exhibit tractable and fascinating biology in terms of their nuclear and nucleosome architecture/substructures, transcription, organellar segregation, DNA replication, developmental biology and flagellar motility, for example [18, 19]. This tractability has been greatly assisted by the capacity to culture trypanosomatids, to generate different life-cycle stages and by the development of advanced genetic tools. Inducible systems for the regulated ectopic expression of genes, or RNAi mediated knockdown have been widely exploited, with CRISPR-Cas9 genome editing [20, 21] recently added to the already large toolbox available to manipulate trypanosomatid genomes and gene expression. As well as their intrinsically interesting biology, trypanosomes remain important causes of disease. Human cases have reduced in recent years, currently at approximately 2000 cases per year, although under-reporting and the discovery of asymptomatic carriers [22] may require revaluation of the prospects for disease elimination. For the animal disease, trypanosomosis remains prevalent and devastating, generating significant restrictions on animal productivity in the tsetse belt of sub-Saharan Africa.

The genome of T. brucei has 11 megabase-chromosomes (~of 35 Mb total) as well as 5 intermediate (200–300 kb) and ~60–100 mini-chromosomes (30–150 kb) [23,24,25]. The megabase chromosomes are segregated via a kinetochore machinery that appears highly divergent with respect to the canonical eukaryotic model [26]. Genes are organised and co-transcribed in large multi-gene (polycistronic) units [27]. There is little evidence of clustering of genes according to life-cycle regulation in T. brucei, such that post transcriptional mechanisms control gene expression (RNA stability, translation, codon usage) [18]. Most protein-coding genes are transcribed by RNA polymerase II (RNAP II) with the mRNA for all protein-coding genes being capped by addition via trans splicing of a 39 nt ‘spliced leader’ (SL) sequence, this assisting translation and stability. Only two genes have been identified with an intron that requires cis splicing [28]. The promoter for the SL array, consisting of 100–200 tandem SL RNA mini-exons [29], possesses a very divergent RNAP II initiation complex [30, 31]. Sites of RNA polymerase II initiation in the genome are epigenetically marked, being enriched in histone modifications H4K10ac and H3K4me3 and four different histone variants that are also found at convergent transcription strand switch regions [32].

Trypanosoma brucei also possesses a uniquely evolved multifunctional RNAP I system, responsible for the transcription of ribosomal gene units, as well as the major surface proteins procyclins (expressed on tsetse midgut forms) and bloodstream VSG genes [33, 34]. It is estimated that there are around 10 million copies of VSGs on the surface of a parasite shielding invariant surface proteins [35,36,37]. Trypanosoma brucei possesses ~2500 VSG genes or pseudogenes [24], located in subtelomeric regions of the chromosomes (comprising ~30% of the genome) [12, 38]. It is now understood that during the course of an infection there are up to 100 VSGs that are expressed in the population at a peak of infection with a few variants dominating at each peak, allowing immune evasion [13, 39]. In the bloodstream stage, there are ~15 telomeric VSG expression sites (ES), each containing 8–9 ES associated genes (ESAGs), 70 bp repeats and a VSG gene [40, 41]. Trypanosoma brucei exhibits monoallelic expression of one ES at a time allowing the expression of one VSG variant per parasite [34, 42, 43]. ES transcription does not occur in the nucleolus, unlike other RNAP I transcripts (rRNA, procyclin genes [44, 45]), but at an extranucleolar focus of RNAP I (the ‘expression site body’) [44, 46, 47]. The switch between different ES is controlled epigenetically [48] and occurs early during the time course of an infection [13, 39]. Later, but still early during the infection, DNA recombination of entire VSG genes occurs into the active ES, from mainly telomeric regions, and is mediated via the 70 bp repeats. When this repertoire is exhausted, recombination occurs from subtelomeric array VSGs, with segmental VSGs conversion producing mosaic VSGs [13, 23, 39, 41, 49,50,51]. These VSG recombinations are dependent on a DNA break [40].

Cytological organisation and flagellar motility have also proved fascinating areas of trypanosome biology of relevance across eukaryotes. Trypanosomes are highly ordered, with their single copy organelles precisely positioned and their movements carefully orchestrated during cell division [52]. Ultrastructural studies using electron microscopy and electron tomography have revealed how the microtubule corset of the trypanosome is co-ordinated with the parasite flagellar basal body and flagellum close to the flagellar pocket [53,54,55,56]. This organisation and the experimental tractability of trypanosomes has made these organisms invaluable as a model for flagellar and ciliary biology in eukaryotic organisms [57, 58].

Three advances in the last decade

High-throughput phenotyping

With the development of Illumina sequencing, a method to survey the representation of genetically distinct cells in a complex population was provided. This has proved immensely powerful in trypanosome research, allowing a genome-wide RNA interference-based phenotyping approach (RNAi target sequencing, RIT-seq) [59]. Using doxycycline-regulated RNAi induction, cytocidal or cytostatic phenotypes that diminish the representation of the RNAi-target sequence relative to the population, reveals “cold spots” that correspond to predicted mRNA sequences. Thus, by combining RNAi and Illumina sequencing, it has been possible to identify genes important for trypanosome growth in the bloodstream and procyclic form as well as during differentiation between these forms.

The use of RIT-seq was also applied in positive selective screens for the identification of genes that contribute to drug resistance [60]. Screens were performed using all current HAT drugs (Eflornithine, Suramin, Nifurtimox, Pentamidine and Melarsoprol) and each yielded a population of cells displaying an RNAi-inducible drug resistance phenotype, associated with the downregulation of specific genes. These have provided molecular insight into the trafficking pathways and resistance mechanisms for anti-trypanosomal drugs.

RIT-seq had also been used effectively to explore trypanosome biology, an example being its use to dissect the signal-response pathway that promotes stumpy formation [61]. Although the quorum sensing signal SIF (stumpy induction factor) was unidentified, the screen exploited the ability of cell-permeable analogues of cAMP or AMP to mimic SIF in laboratory adapted trypanosome lines [62]. By using cAMP analogue-induced cell cycle arrest as the selection, genes involved in the signal-response pathway that promotes stumpy formation were identified from the RNAi inserts enriched in outgrowing proliferative parasites.

Environmental and cell-cell communication

Akin to many other single-celled microbes, it has become clear that African trypanosomes exhibit social behaviours throughout their life-cycle. One example is the coordinated swarming or social motility of T. brucei procyclic forms observed on culture plates [63, 64]. cAMP plays a role in this phenomenon, with knockdown of the cAMP-specific phosphodiesterase PDEB1 found to inhibit social motility [65], and knockdown of certain adenylate cyclases resulting in a ‘hypersocial’ phenotype [66]. This is not dissimilar to the role of cyclic-diGMP in bacterial swarming motility [67].

Social behavior in bloodstream T. brucei in the form of quorum sensing was described in the 1990s [10, 68], with the demonstration that differentiation from the slender to the stumpy bloodstream form occurred in a density-dependent manner resulting from accumulation of an unidentified parasite-derived factor. As detailed above, the signaling components and RNA regulators that drive the generation of the stumpy form, have begun to be revealed in the last decade [61]. Interestingly, despite their lack of morphological differentiation in the bloodstream, the signaling components required to perceive and respond to the density-signal in T. brucei are conserved in T. congolense. Moreover, T. congolense produces a QS signal that causes T. brucei to differentiate to stumpy forms in mixed infections [69], revealing that inter-species QS is possible, just as occurs in bacterial communities.

Consistent with the trypanosome’s reliance on post-transcriptional gene regulation, a number of RNA-binding proteins have been found to play important roles in trypanosome biology [70]. For instance, overexpression of a single RNA-binding protein, RBP6, causes the spontaneous progression from procyclic forms to epimastigote and metacyclic forms in culture, mimicking the transitions occurring in the tsetse fly vector [71]. Another RNA-binding protein was found to be involved in maintenance of the bloodstream form developmental state, with RBP10 binding certain procyclic-specific transcripts and targeting them for repression [72]. Interestingly, regulatory RNA-binding proteins seem to dominate during the control of developmental gene expression as the parasite transitions between life-cycle stages; levels of constitutively expressed transcripts are, in contrast, apparently governed by codon usage bias [73, 74].

A more nuanced picture of infection

Several developments have reshaped our view of the trypanosome infection dynamic and its interactions in a host context.

First, the perception of trypanosome infections as comprising cyclical ‘waves’ of parasitaemia in the bloodstream is now recognised as overly simplistic. Thus, rather than a procession of antigenic variants arising sequentially over time, it is now clear that there is considerable complexity in VSG expression dynamics, with many simultaneously expressed VSGs present within chronic infections albeit comprised of major and minor types [13, 39]. Chronic infections also do not involve an alternating fluctuation between mainly slender and mainly stumpy forms dependent on the overall parasitaemia, with instead, transmissible stumpy forms found to be prevalent throughout long-term infections in mice [75]. It was suggested that by limiting the number of slender forms, which can generate new antigenic variants, the parasites preserve their repertoire of available VSG types whilst also prolonging host survival and so maximizing the probability of transmission.

Secondly, we now understand how human infectivity is possible for some African trypanosomes. Subspecies of T. brucei, T. b. gambiense and T. b. rhodesiense, are the only African trypanosomes able to successfully establish in a human host, because other species are killed by the trypanolytic serum component, apolipoprotein L1 (APOL1). Trypanosoma b. rhodesiense has bypassed this defense through the evolution of Serum resistance associated (SRA) protein, derived from truncation of a VSG [76, 77]. The mechanisms by which T. b. gambiense outmaneuvers human defenses have taken longer to resolve but involve a role for T. b. gambiense-specific glycoprotein (TgsGP), deletion of which restored parasite sensitivity to human serum [78, 79]. Development of a recombinant Papio papio APOL1 mutant that could inhibit T. b. gambiense infection of mice illustrates how an increased knowledge of host-parasite interactions can open up new therapeutic possibilities [80].

Thirdly, the perception of trypanosomiasis as largely being a bloodstream infection has been revised. Trypanosoma brucei, for instance, can be found in abundance within adipose tissue. Adipose tissue parasites are transcriptionally distinct from those in the bloodstream, and may utilize fatty acids as a carbon source, indicating adaptation to their environment [81]. Pockets of T. brucei have also been found in the skin [22, 82]. The blood-brain barrier makes it challenging to effectively target parasites residing in the brain during drug treatment of late stage trypanosomiasis and parasites in the adipose tissue and skin may present a similar challenge for drug delivery. Drug efficacy may also be influenced by pharmacokinetic parameters which differ during the circadian cycle. Although hosts exhibit well defined circadian rhythms, trypanosome gene expression has also been shown to demonstrate circadian oscillation [83]. These oscillations can be observed in culture, following a period of entrainment, generating periodic peaks and troughs in the transcript levels of genes linked to metabolic processes. Interestingly, T. brucei infection can also disrupt the normal circadian rhythm of a murine host, leading to the proposition that sleeping sickness in humans represents a parasite-induced circadian disease [84].

Three areas ripe for development

Coherent assembly of signalling pathways and complexes

The ability of trypanosomes to sense and respond to changes in their environment is essential for their survival. However, the mechanisms underlying the transduction of extracellular signals is poorly understood, with most studies focusing on the function of individual molecules or molecular classes (e.g. kinases). Nonetheless, the development of high throughput technologies has allowed the analysis of signalling pathways at a genome-wide scale in trypanosomes. For example, in the QS response generating stumpy forms, components that fell into a potential hierarchical organisation were identified [61, 85]. To assemble these into a coherent pathway, a concerted knock-out and overexpression approach has been used, revealing dependencies between individual components [86]. With systematic gene ablation and mutation now possible via CRISPR/Cas9 [20], this approach can be extended to understand how components connect to transduce external cues. Refinements in transcriptome analysis, proteomics and phosphoproteomics can further enhance the resolution of signal network analysis and extend the understanding of the associated molecular interactions. Any external cue that generates a defined phenotypic response is accessible to dissection by an integration of these selection and analytical approaches, promising a more coherent understanding of how parasites interact with and respond to their environment to alter life history traits.

The comparative biology of African trypanosome species

Although T. brucei has been the long-established model for African trypanosome biology, the reduced importance of human infections and the recognition of the distinct biology of T. congolense and T. vivax has generated renewed interest in these parasites. Research articles focused on T. congolense and T. vivax infections in the field have provided important insight into the dynamics of infection, host resistance and pathology, as well as prevalence of individual species in livestock, wild animals and tsetse flies. In contrast, knowledge of the molecular biology of T. congolense and T. vivax has lagged behind the wealth of information accumulated for T. brucei. The recent advances in culture techniques and genetic manipulation of T. congolense [87, 88] and T. vivax [89] will enable similar strides to be made in the understanding of the distinct cell biology of these important veterinary pathogens. Each African trypanosome species has developed significant and surprisingly distinct adaptations to their hosts, for example in their predicted surface-expressed proteins [90] and VSG gene archive and expression architecture [91, 92]. Each also shows a distinct developmental path in their tsetse fly vector [9]. It is now clear that we will not be able to fully rely on extrapolating information from T. brucei when it comes to investigating the biology of other trypanosome species.

Host-parasite-vector interactions

Like other parasites, the fate of African trypanosomes is tied to interactions with their host, at least until they can escape to another host via their tsetse fly vector. Whilst immune evasion is one mechanism, trypanosomes also interact with the host immune system with the potential to regulate the course of infection, either through antibody clearance [93] or immune suppression [94]. These host-parasite interactions create the potential for genotype-by-genotype (G×G) interactions which can influence the virulence (parasite numbers) and pathology (disease) generated by parasites in the field. Virulence is often counterbalanced by the transmission benefits of infection chronicity, with the potential to generate evolutionary interactions between trypanosomes and their hosts. This is particularly complex for African trypanosomes which, unlike many parasites, have the ability to infect many mammalian host species, and can also exist in mixed coinfections with other trypanosome species. The tools and information to explore these interactions are emerging as different trypanosome species become genetically tractable and there is an expanding dataset of genomic sequences for field isolates. There is also a new focus on the interaction between the parasite and innate and acquired immune mechanisms [94, 95].

The passage of the trypanosome through the tsetse fly is also increasingly accessible, using sophisticated live cell microscopy [96], the development of labelled trypanosome lines which can be tracked during their journey in the insect vector [97] and the establishment of tools for the manipulation of tsetse gene expression using RNAi [98]. Exploring the tsetse-parasite interaction has also become tractable through increased sensitivity in transcriptome analyses using RNA-seq [99,100,101] and biochemical and proteomic analysis of the parasite metabolic pathways in the fly midgut [102,103,104]. All of these developments can also be deployed to understand the comparative transmission biology of different trypanosome species, given the distinct developmental paths of T. brucei, T. congolense and T. vivax through their vector [105]. Analyses of the interface between the trypanosome in the host and in their vector represent important studies because targeting parasite transmission remains the most effective and cost-efficient route to disease control [106].

Finally, the interaction between the trypanosome and its host in different compartments must be understood. The recent exciting discoveries of the novel niches exploited by trypanosomes in their mammalian host raises questions about the parasite’s biochemistry, drug sensitivity and immune susceptibility in these compartments, and their kinetics of exchange with the bloodstream population [107, 108]. These adaptations may also differ between different trypanosome species with consequences for drug efficacy in cattle populations with mixed infections, in particular.

Conclusions

Trypanosomes continue to provide important insights to the evolution of eukaryotic cells, while remaining significant pathogens in sub-Saharan Africa. In combination, this makes the continued analysis of trypanosome biology compelling. With respect to fundamental biology, the unusual kinetochore composition in trypanosomatids has gained significant interest in the wider cell biology community, as has their unique mitochondrial biology and flagellar dynamics. Significant progress has also been made in therapeutics for these parasites, with several compounds at different stages of the drug development pipeline, most notably fexinidazole [109] and oxaboroles [110]. This has resulted from significant investment in the discovery of small molecule therapeutics which offer oral delivery and reduced toxicity. Similarly, the innovations in gene technology, including high throughput genetic screens, comprehensive protein localisation approaches ([111] http://tryptag.org) and the application of CRISPR technology to laboratory adapted and developmentally competent trypanosome lines have accelerated understanding of the parasite’s biology whilst helping the prediction of how drug resistance might emerge to new therapies. The first golden age of trypanosome biology was in the 1900s when the disease agent and vector was identified, and the second was in the 1980s and early 1990s where the unusual molecular mechanisms of trypanosomes were uncovered, particularly in relation to antigenic variation. We are currently in the third golden age, where this molecular understanding is applied to the fascinating biology of the parasite in the host and in the field, and for the discovery of new therapies.

References

  1. 1.

    Vickerman K. Polymorphism and mitochondrial activity in sleeping sickness. Nature. 1965;208:762–6.

  2. 2.

    Lukes J, Hashimi H, Zikova A. Unexplained complexity of the mitochondrial genome and transcriptome in kinetoplastid flagellates. Curr Genet. 2005;48:277–99.

  3. 3.

    Shapiro TA, Englund PT. The structure and replication of kinetoplast DNA. Annu Rev Microbiol. 1995;49:117–43.

  4. 4.

    Ryan KA, Shapiro TA, Rauch CA, Englund PT. Replication of kinetoplast DNA in trypanosomes. Annu Rev Microbiol. 1988;42:339–58.

  5. 5.

    Borst P, Fase F-F, Weijers PJ, Barry JD, Tetley L, Vickerman K. Kinetoplast DNA from Trypanosoma vivax and T. congolense. Mol Biochem Parasitol. 1985;15:129–42.

  6. 6.

    Giordani F, Morrison LJ, Rowan TG, De Koning HP, Barrett MP. The animal trypanosomiases and their chemotherapy: a review. Parasitology. 2016;143:1862–89.

  7. 7.

    Buscher P, Cecchi G, Jamonneau V, Priotto G. Human African trypanosomiasis. Lancet. 2017;390:2397–409.

  8. 8.

    Silvester E, McWilliam KR, Matthews KR. The cytological events and molecular control of life cycle development of Trypanosoma brucei in the mammalian bloodstream. Pathogens. 2017;6:29.

  9. 9.

    Rotureau B, Van Den Abbeele J. Through the dark continent: African trypanosome development in the tsetse fly. Front Cell Infect Microbiol. 2013;3:53.

  10. 10.

    Vassella E, Reuner B, Yutzy B, Boshart M. Differentiation of African trypanosomes is controlled by a density sensing mechanism which signals cell cycle arrest via the cAMP pathway. J Cell Sci. 1997;110:2661–71.

  11. 11.

    Matthews KR, Gull K. Evidence for an interplay between cell cycle progression and the initiation of differentiation between life cycle forms of African trypanosomes. J Cell Biol. 1994;125:1147–56.

  12. 12.

    Horn D. Antigenic variation in African trypanosomes. Mol Biochem Parasitol. 2014;195:123–9.

  13. 13.

    Mugnier MR, Cross GA, Papavasiliou FN. The in vivo dynamics of antigenic variation in Trypanosoma brucei. Science. 2015;347:1470–3.

  14. 14.

    Silvester E, Young J, Ivens A, Matthews KR. Interspecies quorum-sensing in co-infections can manipulate trypanosome transmission potential. Nature Microbiol. 2017;2:1471–9.

  15. 15.

    Shapiro SZ, Naessen J, Liesegang B, Moloo SK, Magondu J. Analysis by flow cytometry of DNA synthesis during the life cycle of African trypanosomes. Acta Trop. 1984;41:313–23.

  16. 16.

    Dacks JB, Walker G, Field MC. Implications of the new eukaryotic systematics for parasitologists. Parasitol Int. 2008;57:97–104.

  17. 17.

    Lukes J, Skalicky T, Tyc J, Votypka J, Yurchenko V. Evolution of parasitism in kinetoplastid flagellates. Mol Biochem Parasitol. 2014;195:115–22.

  18. 18.

    Daniels JP, Gull K, Wickstead B. Cell biology of the trypanosome genome. Microbiol Mol Biol Rev. 2010;74:552–69.

  19. 19.

    Hodges ME, Scheumann N, Wickstead B, Langdale JA, Gull K. Reconstructing the evolutionary history of the centriole from protein components. J Cell Sci. 2010;123:1407–13.

  20. 20.

    Beneke T, Madden R, Makin L, Valli J, Sunter J, Gluenz E. A CRISPR Cas9 high-throughput genome editing toolkit for kinetoplastids. R Soc Open Sci. 2017;4:170095.

  21. 21.

    Rico E, Jeacock L, Kovarova J, Horn D. Inducible high-efficiency CRISPR-Cas9-targeted gene editing and precision base editing in African trypanosomes. Sci Rep. 2018;8:7960.

  22. 22.

    Capewell P, Cren-Travaille C, Marchesi F, Johnston P, Clucas C, Benson RA, et al. The skin is a significant but overlooked anatomical reservoir for vector-borne African trypanosomes. Elife. 2016;5:e17716.

  23. 23.

    Berriman M, Ghedin E, Hertz-Fowler C, Blandin G, Renauld H, Bartholomeu DC, et al. The genome of the African trypanosome Trypanosoma brucei. Science. 2005;309:416–22.

  24. 24.

    Cross GA, Kim HS, Wickstead B. Capturing the variant surface glycoprotein repertoire (the VSGnome) of Trypanosoma brucei Lister 427. Mol Biochem Parasitol. 2014;195:59–73.

  25. 25.

    Hertz-Fowler C, Renauld H, Berriman M. Trypanosomes: after the genome. Glasgow: Wellcome Centre for Molecular Parasitology, University of Glasgow, Horizon Bioscience; 2007.

  26. 26.

    Akiyoshi B, Gull K. Discovery of unconventional kinetochores in kinetoplastids. Cell. 2014;156:1247–58.

  27. 27.

    Kolev NG, Franklin JB, Carmi S, Shi H, Michaeli S, Tschudi C. The transcriptome of the human pathogen Trypanosoma brucei at single-nucleotide resolution. PLoS Pathog. 2010;6:e1001090.

  28. 28.

    Mair G, Shi H, Li H, Djikeng A, Aviles HO, Bishop JR, et al. A new twist in trypanosome RNA metabolism: cis-splicing of pre-mRNA. RNA. 2000;6:163–9.

  29. 29.

    Campbell DA, Thomas S, Sturm NR. Transcription in kinetoplastid protozoa: why be normal? Microbes Infect. 2003;5:1231–40.

  30. 30.

    Gilinger G, Bellofatto V. Trypanosome spliced leader RNA genes contain the first identified RNA polymerase II gene promoter in these organisms. Nucleic Acids Res. 2001;29:1556–64.

  31. 31.

    Günzl AVL, Myler PJ. Trypanosomes: after the genome. Horizon Bioscience: Wellcome Centre for Molecular Parasitology, University of Glasgow, Glasgow, UK; 2007.

  32. 32.

    Wright JR, Siegel TN, Cross GA. Histone H3 trimethylated at lysine 4 is enriched at probable transcription start sites in Trypanosoma brucei. Mol Biochem Parasitol. 2010;172:141–4.

  33. 33.

    Gunzl A, Bruderer T, Laufer G, Schimanski B, Tu LC, Chung HM, et al. RNA polymerase I transcribes procyclin genes and variant surface glycoprotein gene expression sites in Trypanosoma brucei. Eukaryot Cell. 2003;2:542–51.

  34. 34.

    Johnson PJ, Kooter JM, Borst P. Inactivation of transcription by UV irradiation of T. brucei provides evidence for a multicistronic transcription unit including a VSG gene. Cell. 1987;51:273–81.

  35. 35.

    Cross GA. Identification, purification and properties of clone-specific glycoprotein antigens constituting the surface coat of Trypanosoma brucei. Parasitology. 1975;71:393–417.

  36. 36.

    Schwede A, Jones N, Engstler M, Carrington M. The VSG C-terminal domain is inaccessible to antibodies on live trypanosomes. Mol Biochem Parasitol. 2011;175:201–4.

  37. 37.

    Schwede A, Macleod OJ, MacGregor P, Carrington M. How does the VSG coat of bloodstream form African trypanosomes interact with external proteins? PLoS Pathog. 2015;11:e1005259.

  38. 38.

    Ersfeld K. Nuclear architecture, genome and chromatin organisation in Trypanosoma brucei. Res Microbiol. 2011;162:626–36.

  39. 39.

    Hall JP, Wang H, Barry JD. Mosaic VSGs and the scale of Trypanosoma brucei antigenic variation. PLoS Pathog. 2013;9:e1003502.

  40. 40.

    Glover L, Hutchinson S, Alsford S, McCulloch R, Field MC, Horn D. Antigenic variation in African trypanosomes: the importance of chromosomal and nuclear context in VSG expression control. Cell Microbiol. 2013;15:1984–93.

  41. 41.

    Hertz-Fowler C, Figueiredo LM, Quail MA, Becker M, Jackson A, Bason N, et al. Telomeric expression sites are highly conserved in Trypanosoma brucei. PLoS One. 2008;3:e3527.

  42. 42.

    Borst P. Antigenic variation and allelic exclusion. Cell. 2002;109:5–8.

  43. 43.

    Pays E. Regulation of antigen gene expression in Trypanosoma brucei. Trends Parasitol. 2005;21:517–20.

  44. 44.

    Landeira D, Navarro M. Nuclear repositioning of the VSG promoter during developmental silencing in Trypanosoma brucei. J Cell Biol. 2007;176:133–9.

  45. 45.

    Nazer E, Sanchez DO. Nucleolar accumulation of RNA binding proteins induced by Actinomycin D is functional in Trypanosoma cruzi and Leishmania mexicana but not in T. brucei. PLoS One. 2011;6(8):e24184.

  46. 46.

    Chaves I, Zomerdijk J, Dirks-Mulder A, Dirks RW, Raap AK, Borst P. Subnuclear localization of the active variant surface glycoprotein gene expression site in Trypanosoma brucei. Proc Natl Acad Sci USA. 1998;95:12328–33.

  47. 47.

    Navarro M, Gull K. A pol I transcriptional body associated with VSG mono-allelic expression in Trypanosoma brucei. Nature. 2001;414:759–63.

  48. 48.

    Zomerdijk JC, Kieft R, Duyndam M, Shiels PG, Borst P. Antigenic variation in Trypanosoma brucei: a telomeric expression site for variant-specific surface glycoprotein genes with novel features. Nucleic Acids Res. 1991;19:1359–68.

  49. 49.

    Barry JD, Marcello L, Morrison LJ, Read AF, Lythgoe K, Jones N, et al. What the genome sequence is revealing about trypanosome antigenic variation. Biochem Soc Trans. 2005;33:986–9.

  50. 50.

    Marcello L, Barry JD. Analysis of the VSG gene silent archive in Trypanosoma brucei reveals that mosaic gene expression is prominent in antigenic variation and is favored by archive substructure. Genome Res. 2007;17:1344–52.

  51. 51.

    Young R, Taylor JE, Kurioka A, Becker M, Louis EJ, Rudenko G. Isolation and analysis of the genetic diversity of repertoires of VSG expression site containing telomeres from Trypanosoma brucei gambiense, T. brucei and T. equiperdum. BMC Genomics. 2008;9:385.

  52. 52.

    Gull K. The cytoskeleton of trypanosomatid parasites. Annu Rev Microbiol. 1999;53:629–55.

  53. 53.

    Vaughan S. Assembly of the flagellum and its role in cell morphogenesis in Trypanosoma brucei. Curr Opin Microbiol. 2010;13:453–8.

  54. 54.

    Lacomble S, Vaughan S, Gadelha C, Morphew MK, Shaw MK, McIntosh JR, et al. Basal body movements orchestrate membrane organelle division and cell morphogenesis in Trypanosoma brucei. J Cell Sci. 2010;123:2884–91.

  55. 55.

    Robinson DR, Gull K. Basal body movements as a mechanism for mitochondrial genome segregation in the trypanosome cell cycle. Nature. 1991;352:731–3.

  56. 56.

    Hoog JL, Bouchet-Marquis C, McIntosh JR, Hoenger A, Gull K. Cryo-electron tomography and 3-D analysis of the intact flagellum in Trypanosoma brucei. J Struct Biol. 2012;178:189–98.

  57. 57.

    Broadhead R, Dawe HR, Farr H, Griffiths S, Hart SR, Portman N, et al. Flagellar motility is required for the viability of the bloodstream trypanosome. Nature. 2006;440:224–7.

  58. 58.

    Morga B, Bastin P. Getting to the heart of intraflagellar transport using Trypanosoma and Chlamydomonas models: the strength is in their differences. Cilia. 2013;2:16.

  59. 59.

    Alsford S, Turner DJ, Obado SO, Sanchez-Flores A, Glover L, Berriman M, et al. High-throughput phenotyping using parallel sequencing of RNA interference targets in the African trypanosome. Genome Res. 2011;21:915–24.

  60. 60.

    Alsford S, Glover L, Leung KF, Field MC, Horn D. High-throughput decoding of antitrypanosomal drug efficacy and resistance. Nature. 2012;482:232–6.

  61. 61.

    Mony BM, Macgregor P, Ivens A, Rojas F, Cowton A, Young J, et al. Genome-wide dissection of the quorum sensing signalling pathway in Trypanosoma brucei. Nature. 2014;505:681–5.

  62. 62.

    Laxman S, Riechers A, Sadilek M, Schwede F, Beavo JA. Hydrolysis products of cAMP analogs cause transformation of Trypanosoma brucei from slender to stumpy-like forms. Proc Natl Acad Sci USA. 2006;103:19194–9.

  63. 63.

    Oberholzer M, Lopez MA, McLelland BT, Hill KL. Social motility in African trypanosomes. PLoS Pathog. 2010;6:e1000739.

  64. 64.

    Imhof S, Knüsel S, Gunasekera K, Vu XL, Roditi I. Social motility of African trypanosomes is a property of a distinct life-cycle stage that occurs early in tsetse fly transmission. PLoS Pathog. 2014;10:e1004493.

  65. 65.

    Oberholzer M, Saada EA, Hill KL. Cyclic AMP regulates social behavior in African trypanosomes. mBio. 2015;6:e01954.

  66. 66.

    Lopez MA, Saada EA, Hill KL. Insect stage-specific adenylate cyclases regulate social motility in African trypanosomes. Eukaryot Cell. 2015;14:104–12.

  67. 67.

    Simm R, Morr M, Kader A, Nimtz M, Römling U. GGDEF and EAL domains inversely regulate cyclic di-GMP levels and transition from sessility to motility. Mol Microbiol. 2004;53:1123–34.

  68. 68.

    Reuner B, Vassella E, Yutzy B, Boshart M. Cell density triggers slender to stumpy differentiation of Trypanosoma brucei bloodstream forms in culture. Mol Biochem Parasitol. 1997;90:269–80.

  69. 69.

    Silvester E, Young J, Ivens A, Matthews KR. Interspecies quorum sensing in co-infections can manipulate trypanosome transmission potential. Nat Microbiol. 2017;2:1471–9.

  70. 70.

    Clayton C. The regulation of trypanosome gene expression by RNA-binding proteins. PLoS Pathog. 2013;9:e1003680.

  71. 71.

    Kolev NG, Ramey-Butler K, Cross GAM, Ullu E, Tschudi C. Developmental progression to infectivity in Trypanosoma brucei triggered by an RNA-binding protein. Science. 2012;338:1352–3.

  72. 72.

    Mugo E, Clayton C. Expression of the RNA-binding protein RBP10 promotes the bloodstream-form differentiation state in Trypanosoma brucei. PLoS Pathog. 2017;13:e1006560.

  73. 73.

    Jeacock L, Faria J, Horn D. Codon usage bias controls mRNA and protein abundance in trypanosomatids. eLife. 2018;7:e32496.

  74. 74.

    de Freitas Nascimento J, Kelly S, Sunter J, Carrington M. Codon choice directs constitutive mRNA levels in trypanosomes. eLife. 2018;7:e32467.

  75. 75.

    MacGregor P, Savill NJ, Hall D, Matthews KR. Transmission stages dominate trypanosome within-host dynamics during chronic infections. Cell Host Microbe. 2011;9:310–8.

  76. 76.

    Xong HV, Vanhamme L, Chamekh M, Chimfwembe CE, Van Den Abbeele J, Pays A, et al. A VSG expression site-associated gene confers resistance to human serum in Trypanosoma rhodesiense. Cell. 1998;95:839–46.

  77. 77.

    Vanhamme L, Paturiaux-Hanocq F, Poelvoorde P, Nolan DP, Lins L, Van Den Abbeele J, et al. Apolipoprotein L-I is the trypanosome lytic factor of human serum. Nature. 2003;422:83–7.

  78. 78.

    Capewell P, Clucas C, DeJesus E, Kieft R, Hajduk S, Veitch N, et al. The TgsGP gene is essential for resistance to human serum in Trypanosoma brucei gambiense. PLoS Pathog. 2013;9:e1003686.

  79. 79.

    Uzureau P, Uzureau S, Lecordier L, Fontaine F, Tebabi P, Homble F, et al. Mechanism of Trypanosoma brucei gambiense resistance to human serum. Nature. 2013;501:430–4.

  80. 80.

    Fontaine F, Lecordier L, Vanwalleghem G, Uzureau P, Van Reet N, Fontaine M, et al. APOLs with low pH dependence can kill all African trypanosomes. Nat Microbiol. 2017;2:1500–6.

  81. 81.

    Trindade S, Rijo-Ferreira F, Carvalho T, Pinto-Neves D, Guegan F, Aresta-Branco F, et al. Trypanosoma brucei parasites occupy and functionally adapt to the adipose tissue in mice. Cell Host Microbe. 2016;19:837–48.

  82. 82.

    Caljon G, Van Reet N, De Trez C, Vermeersch M, Pérez-Morga D, Van Den Abbeele J. The dermis as a delivery site of Trypanosoma brucei for tsetse flies. PLoS Pathog. 2016;12:e1005744.

  83. 83.

    Rijo-Ferreira F, Pinto-Neves D, Barbosa-Morais NL, Takahashi JS, Figueiredo LM. Trypanosoma brucei metabolism is under circadian control. Nat Microbiol. 2017;2:17032.

  84. 84.

    Rijo-Ferreira F, Carvalho T, Afonso C, Sanches-Vaz M, Costa RM, Figueiredo LM, et al. Sleeping sickness is a circadian disorder. Nat Commun. 2018;9:62.

  85. 85.

    Mony BM, Matthews KR. Assembling the components of the quorum sensing pathway in African trypanosomes. Mol Microbiol. 2015;96:220–32.

  86. 86.

    McDonald L, Cayla M, Ivens A, Mony BM, MacGregor P, Silvester E, et al. Non-linear hierarchy of the quorum sensing signalling pathway in bloodstream form African trypanosomes. PLoS Pathog. 2018;14:e1007145.

  87. 87.

    Gibson W, Kay C, Peacock L. Trypanosoma congolense: molecular toolkit and resources for studying a major livestock pathogen and model trypanosome. Adv Parasitol. 2017;98:283–309.

  88. 88.

    Coustou V, Guegan F, Plazolles N, Baltz T. Complete in vitro life cycle of Trypanosoma congolense: development of genetic tools. PLoS Negl Trop Dis. 2010;4:e618.

  89. 89.

    D’Archivio S, Medina M, Cosson A, Chamond N, Rotureau B, Minoprio P, et al. Genetic engineering of Trypanosoma (Dutonella) vivax and in vitro differentiation under axenic conditions. PLoS Negl Trop Dis. 2011;5:e1461.

  90. 90.

    Jackson AP, Allison HC, Barry JD, Field MC, Hertz-Fowler C, Berriman M. A Cell-surface phylome for african trypanosomes. PLoS Negl Trop Dis. 2013;7:e2121.

  91. 91.

    Abbas AH, Silva Pereira S, D’Archivio S, Wickstead B, Morrison LJ, Hall N, et al. The structure of a conserved telomeric region associated with variant antigen loci in the blood parasite Trypanosoma congolense. Genome Biol Evol. 2018;10:2458–73.

  92. 92.

    Jackson AP, Berry A, Aslett M, Allison HC, Burton P, Vavrova-Anderson J, et al. Antigenic diversity is generated by distinct evolutionary mechanisms in African trypanosome species. Proc Natl Acad Sci USA. 2012;109:3416–21.

  93. 93.

    Engstler M, Pfohl P, Herminghaus S, Boshart M, Wiegerttjes G, Heddergott N, et al. Hydrodynamic flow-mediated protein sorting on the cell surface of trypanosomes. Cell. 2007;131:505–15.

  94. 94.

    Frenkel D, Zhang F, Guirnalda P, Haynes C, Bockstal V, Radwanska M, et al. Trypanosoma brucei co-opts NK cells to kill splenic B2 B cells. PLoS Pathog. 2016;12:e1005733.

  95. 95.

    Stijlemans B, Caljon G, Van Den Abbeele J, Van Ginderachter JA, Magez S, De Trez C. Immune evasion strategies of Trypanosoma brucei within the mammalian host: progression to pathogenicity. Front Immunol. 2016;7:233.

  96. 96.

    Schuster S, Kruger T, Subota I, Thusek S, Rotureau B, Beilhack A, et al. Developmental adaptations of trypanosome motility to the tsetse fly host environments unravel a multifaceted in vivo microswimmer system. Elife. 2017;6:e27656.

  97. 97.

    Peacock L, Ferris V, Sharma R, Sunter J, Bailey M, Carrington M, et al. Identification of the meiotic life cycle stage of Trypanosoma brucei in the tsetse fly. Proc Natl Acad Sci USA. 2011;108:3671–6.

  98. 98.

    Walshe DP, Lehane SM, Lehane MJ, Haines LR. Prolonged gene knockdown in the tsetse fly Glossina by feeding double stranded RNA. Insect Mol Biol. 2009;18:11–9.

  99. 99.

    Savage AF, Kolev NG, Franklin JB, Vigneron A, Aksoy S, Tschudi C. Transcriptome profiling of Trypanosoma brucei development in the tsetse fly vector Glossina morsitans. PLoS One. 2016;11:e0168877.

  100. 100.

    Awuoche EO, Weiss BL, Mireji PO, Vigneron A, Nyambega B, Murilla G, et al. Expression profiling of Trypanosoma congolense genes during development in the tsetse fly vector Glossina morsitans morsitans. Parasit Vectors. 2018;11:380.

  101. 101.

    Helm JR, Hertz-Fowler C, Aslett M, Berriman M, Sanders M, Quail MA, et al. Analysis of expressed sequence tags from the four main developmental stages of Trypanosoma congolense. Mol Biochem Parasitol. 2009;168:34–42.

  102. 102.

    Mantilla BS, Marchese L, Casas-Sanchez A, Dyer NA, Ejeh N, Biran M, et al. Proline metabolism is essential for Trypanosoma brucei brucei survival in the tsetse vector. PLoS Pathog. 2017;13:e1006158.

  103. 103.

    Millerioux Y, Mazet M, Bouyssou G, Allmann S, Kiema TR, Bertiaux E, et al. De novo biosynthesis of sterols and fatty acids in the Trypanosoma brucei procyclic form: carbon source preferences and metabolic flux redistributions. PLoS Pathog. 2018;14:e1007116.

  104. 104.

    Eyford BA, Sakurai T, Smith D, Loveless B, Hertz-Fowler C, Donelson JE, et al. Differential protein expression throughout the life cycle of Trypanosoma congolense, a major parasite of cattle in Africa. Mol Biochem Parasitol. 2011;177:116–25.

  105. 105.

    Van Den Abbeele J, Rotureau B. New insights in the interactions between African trypanosomes and tsetse flies. Front Cell Infect Microbiol. 2013;3:63.

  106. 106.

    Aksoy S, Buscher P, Lehane M, Solano P, Van Den Abbeele J. Human African trypanosomiasis control: achievements and challenges. PLoS Negl Trop Dis. 2017;11:e0005454.

  107. 107.

    Tanowitz HB, Scherer PE, Mota MM, Figueiredo LM. Adipose tissue: a safe haven for parasites? Trends Parasitol. 2017;33:276–84.

  108. 108.

    McCulloch R, Cobbold CA, Figueiredo L, Jackson A, Morrison LJ, Mugnier MR, et al. Emerging challenges in understanding trypanosome antigenic variation. Emerg Top Life Sci. 2017;1:585–92.

  109. 109.

    Mesu V, Kalonji WM, Bardonneau C, Mordt OV, Blesson S, Simon F, et al. Oral fexinidazole for late-stage African Trypanosoma brucei gambiense trypanosomiasis: a pivotal multicentre, randomised, non-inferiority trial. Lancet. 2018;391:144–54.

  110. 110.

    Field MC, Horn D, Fairlamb AH, Ferguson MA, Gray DW, Read KD, et al. Anti-trypanosomatid drug discovery: an ongoing challenge and a continuing need. Nat Rev Microbiol. 2017;15:217–31.

  111. 111.

    Dean S, Sunter JD, Wheeler RJ. TrypTag.org: A trypanosome genome-wide protein localisation resource. Trends Parasitol. 2017;33:80–2.

Download references

Acknowledgements

We thank members of KMʼs laboratory for discussions.

Funding

Work in KMʼs laboratory is funded by a Wellcome Trust Investigator Award (103740/Z14/Z) and a Wolfson Research Merit Award (WM140045). The funders had no role in the preparation of the manuscript.

Availability of data and materials

Not applicable.

Authorsʼ contributions

MC, FR, ES, FV and KM contributed equally to preparation of the manuscript. The poster was designed by ES, with input from KM. All authors read and approved the final manuscript.

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Author information

Correspondence to Keith R. Matthews.

Additional file

13071_2019_3355_MOESM1_ESM.tif

Additional file 1. Poster on recent developments in the biology of African trypanosomes depicting the life-cycle of Trypanosoma brucei. In the left call out box are shown developments in the biology of trypanosomes in their mammalian host discussed in the text. In the right call out box are shown relevant features of the biology of trypanosomes in their arthropod vector, the tsetse fly. The bottom box highlights recent technological developments for dissecting gene function or location in trypanosomes.

Rights and permissions

Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Keywords

  • Trypanosoma brucei
  • Sleeping sickness
  • Trypanosome

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate. Please note that comments may be removed without notice if they are flagged by another user or do not comply with our community guidelines.