Networks of gene expression regulation in Trypanosoma brucei.

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Contents lists available at ScienceDirect

Molecular & Biochemical Parasitology

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Review

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Networks of gene expression regulation in Trypanosoma brucei C.E. Clayton ∗

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Q2 Universität Heidelberg, Zentrum für Molekulare Biologie der Universität Heidelberg, DKFZ-ZMBH Alliance, Im Neuenheimer Feld 282, 69120 Heidelberg, Q3 Germany

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a r t i c l e

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Article history: Available online xxx

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Keywords: Trypanosoma mRNA decay mRNA processing Splicing Translation

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Contents

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Regulation of gene expression in Kinetoplastids relies mainly on post-transcriptional mechanisms. Recent high-throughput analyses, combined with mathematical modelling, have demonstrated possibilities for transcript-specific regulation at every stage: trans splicing, polyadenylation, translation, and degradation of both the precursor and the mature mRNA. Different mRNA degradation pathways result in different types of degradation kinetics. The original idea that the fate of an mRNA – or even just its degradation kinetics – can be defined by a single “regulatory element” is an over-simplification. It is now clear that every mRNA can bind many different proteins, some of which may compete with each other. Superimposed upon this complexity are the interactions of those proteins with effectors of gene expression. The amount of protein that is made from a gene is therefore determined by a complex network of interactions. © 2014 Published by Elsevier B.V.

Trypanosome transcription . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The contributions of nuclear processes to mRNA regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Degradation of the mature mRNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Degradation pathways for cytosolic mRNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of translation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Granules, degradation and translation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of mRNA decay and translation: the roles of RNA binding proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Finding regulatory proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Trypanosome transcription Gene expression in Kinetoplastids is remarkable because transcription is polycistronic (reviewed in [1]). The 5 end of each individual mRNA is generated by trans splicing of a capped “spliced leader” (SL, 39nt in Trypanosoma brucei) at the 5 end [2]. The trans spliceosome is linked, in a way that is not yet understood, to the polyadenylation machinery, such that each splicing reaction directs – and is coupled to – polyadenylation of the preceding mRNA [3]. For individual mRNAs that are synthesised by RNA polymerase II, this set-up results in a complete lack of transcriptional regulation: almost the entire burden of determining how much of each mRNA is

∗ Tel.: +49 6221 546876; fax: +49 6221 545894. E-mail address: [email protected]

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present in the cell falls on post-transcriptional processes. Possible points of regulation are trans splicing, polyadenylation, degradation of the mRNA within the nucleus, export from the nucleus, and degradation of the mRNA in the cytosol. The amount of protein that is made might also be regulated though control of translation initiation or elongation. T. brucei grows in the blood and tissue fluids of mammals, and in the midgut, foregut, proventriculus and salivary glands of Tsetse flies [4]. In the mammal, the bloodstream-form trypomastigotes rely on substrate-level phosphorylation for ATP generation, with glucose as the main energy source [5,6]; and upon antigenic variation of variant surface glycoprotein (VSG) for defence against the humoral immune system [7]. The procyclic trypomastigotes in the Tsetse midgut have a much more active mitochondrion [6], and have procyclins on the surface [8]; while epimastigotes in the proventriculus and salivary glands have a third type of

http://dx.doi.org/10.1016/j.molbiopara.2014.06.005 0166-6851/© 2014 Published by Elsevier B.V.

Please cite this article in press as: Clayton CE. Networks of gene expression regulation in Trypanosoma brucei. Mol Biochem Parasitol (2014), http://dx.doi.org/10.1016/j.molbiopara.2014.06.005

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surface protein called BARP [9]. Finally, there are non-dividing forms called metacyclics, with surface VSG, in the salivary glands. A bloodstream-form trypanosome (growing at 37 ◦ C, with a division time of 6–8 h) has about 20,000 mRNAs [10]. (There are about 7000 different open reading frames (ORFs) in the genome [11].) Procyclic forms are grown at 27 ◦ C; they divide slower than bloodstream forms, are about twice the size, and correspondingly have twice as many mRNA molecules [10]. The transcriptomes of bloodstream and procyclic forms have been analysed by several laboratories, initially with the use of microarrays, and later by high-throughout sequencing of cDNA [11–14]. In bloodstream forms, the mRNAs from most genes are present at 1–2 copies per cell, but for some, over 200 are present [14,15]. Genes in the latter category are always present in multiple copies, often as tandem repeats. It appears that the rate of constitutive polymerase II transcription is simply inadequate to supply more than about 50 mRNAs per ORF. An alternative mechanism to escape the limitations of the constitutive pol II transcription is unique to salivarian trypanosomes: the genes encoding the variant surface glycoprotein (VSG) and the procyclins are transcribed by RNA polymerase I. RNA polymerase I has about ten-fold more active initiation than for polymerase II, and may also show faster elongation [10,16]. In addition, polymerase I initiation can be regulated by epigenetic means, allowing both stage-specific transcription and the exclusive expression of one VSG at a time – the prerequisite for antigenic variation [7]. 2. The contributions of nuclear processes to mRNA regulation Fig. 1 illustrates transcription and processing for an imaginary 10 kb region, in the form of a time-lapse image with the position of RNA pol II indicated at 1-min intervals. The processes that determine the amount of mature mRNA that is made and can be exported to the cytoplasm are as follows: 1) Transcription. The transcription rate has not been measured. We do know, though, that the parasite has to be making enough

mRNA all the time in order to both balance mRNA degradation, and to double the amount of mRNA before the cell divides. This means that we can calculate the average rate of RNA synthesis. If we assume that the rate of transcription elongation is similar to average estimates from Opisthokonts – about 20 nt/s – we can calculate how often initiation has to happen. This gives an average distance between pol II elongation complexes of 40 kb [10]. 2) Trans splicing. Careful measurements on the PGKB mRNA in vivo [10], and the HSP70 [17] and tubulin mRNAs [18] in an in vitro system, indicated that splicing happens 1–2 min after the splice acceptor site was synthesised. For technical and theoretical reasons, it is not possible to measure splicing rates across the entire transcriptome, but results from measurements for a few thousand genes suggested that trans splicing half-times might be between 1 and 5 min [14] (which translates into average splicing times of 1.5–7 min). Assuming the elongation rate of 20 nt/s, his would mean that a splice acceptor site will be processed when the pol II complex that made it is 1–6 kb downstream. In the figure, RNAs A. C and D are all trans spliced about 1 min after their splice acceptor sites were synthesised, while RNA B is has a poor splicing signal so its splicing is delayed by a further 3 min. 3) Polyadenylation. Since polyadenylation is coupled to trans splicing of the next mRNA downstream, it is expected to show the same kinetics [18] – perhaps with a slight delay. In Fig. 1, therefore, RNAs B, C and D are polyadenylated 1–2 min after the poly(A) site was made, but polyadenylation of RNA A is delayed. 4) Instinctively, one would think that if an mRNA were processed really slowly, there should be less of it than if it were processed fast. Indeed, if mRNA synthesis has only just been turned on, the processing rate has a critical influence of the rate at which the mRNA accumulates. However, once steady-state has been reached, the processing rate is irrelevant, since each mRNA always gets made eventually. Fig. 1 shows this: there is just as much of RNA B as there is of RNAs A, C and D. For mRNA processing to have any effect on steady-state abundances, processing has to be competing with degradation. Indeed, degradation does occur in the absence of processing. The only

Fig. 1. Pathway of mRNA formation. Four genes – A (blue), B (pink), C (purple) and D (green) – are in a polycistronic transcription unit. They are preceded by another gene that is not shown – mRNA in orange. The coding regions are darker-coloured than the untranslated regions. The figure is a time-lapse snapshot of RNA polymerase II taken at 1-min intervals, assuming an elongation rate of about 20 nt/s. The black star is the spliced leader. In this hypothetical example, 5 -trans splicing of mRNAs A, C, and D happens after 1–2 min of splicing acceptor synthesis, but splicing of B is slower (5 min) – for example, it might have a weak polypyrimidine tract. Note that the slow splicing of “B” does not affect the abundance of RNAs “A” or “B”, but causes slow polyadenylation of “A” and thus retention of mRNA “A” at the chromatin. The black “chompers” represent co-transcriptional degradation; the one at the 5 kb mark is currently active. RNAs “A” and “B” are much more likely to be “chomped” than RNAs “C and “D” because they are attached to the chromatin for longer.


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measurement available – for the PGKB gene – gives precursor degradation a half-life of about 8 min [10]. This would be degradation of mRNA that is still attached to the elongating pol II complex. Possibilities include an endonuclease associated with pol II or chromatin, or abortive cleavage by the processing machinery. An activity that cut as indicated in Fig. 1 (black head with open mouth) would attack the A–B precursor, with subsequent exonucleolytic destruction of the pieces in processes analogous to the nuclear quality control machineries of Opisthokonts [19–21]. 3. Degradation of the mature mRNA Once the mRNA has been completed it is exported from the nucleus, using mechanisms related to those in other eukaryotes [22,23]. Since nuclear export rates are determined primarily by the time taken to diffuse to the nuclear pore [24], we expect export to take less than a minute for the relatively small trypanosome nucleus. On its way to the nuclear pore, a defective mRNA might be degraded by the quality control machinery. Standard measurements of mature mRNA decay do not distinguish between nuclear and cytosolic degradation. The rates of decay of mature mRNAs have been studied on a transcriptome-wide scale [14]. This was done by inhibiting transcription, then characterising transcriptomes by high-throughput sequencing (RNASeq). The median half-life for bloodstream forms was 12 min, with 20 min for procyclic forms, but the range was huge: some mRNAs did not appear to be degraded at all, while others had half-lives of as little 5 min. Many developmentally regulated mRNAs also had different decay rates in bloodstream and procyclic forms. Using the numbers above, one can ask how much each process contributes to overall regulation of gene expression. If decay of the mature mRNA were the only factor affecting mRNA abundance, the number of mRNAs per gene should be directly proportional to the mRNA half-life. In practice, a large proportion of mRNAs is less abundant that would be expected from decay rates alone [14]. The easiest interpretation of this result is that many transcripts are destroyed as precursors. Since it is polyadenylation that finally detaches the mRNA from its template, this would lead to preferential loss of mRNAs that are polyadenylated slowly, either because polyadenylation is inefficient, or because the mRNA is very long. For example, in Fig. 1, RNA A is attached to the template for 5 min because it is polyadenylated slowly, and RNA B is attached for the same time because it is an over 5 kb long. RNA C, in contrast, is attached for less than 2 min. Support for this idea comes from the fact that for any given half-life, long mRNAs are less abundant than short ones [14]. If the mRNA abundance were indeed decreased by longer association with chromatin, then inhibition of transcription elongation should lead to disproportionate loss of long mRNAs. This was indeed was seen after RNAi targeting the putative elongation factor CTR9 [25,26]. It is therefore possible that mRNA abundance could be affected by transcriptional pausing. Such pausing has been documented in mammalian cells [27,28] and yeast [29,30], often depending on cross-talk between RNA polymerase II and the processing machinery. There is as yet no evidence for such cross-talk in Kinetoplastids [31], but transcriptional pausing might nevertheless be caused by specific DNA sequences, secondary structures or chromatin modification. 4. Degradation pathways for cytosolic mRNAs Fig. 2 shows the likely pathways of mRNA degradation in trypanosomes [32]. The mRNA is being translated, and the poly(A)

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tail is bound by poly(A) binding protein (PABP) (Fig. 2A). There are two PABPs in trypanosomes, PABP1 and PABP2, which seem to have slightly different functions [33]. The 5 cap of the mRNA is bound by the translation initiation factor eIF4E, which recruits eIF4G. In other eukaryotes, eIF4G binds to the helicase eiF4A, and also to PABP – thus effectively circularising the mRNA [34,35]. The figure also shows various RNA-binding proteins associated with the 3 -untranslated region (3 -UTR) of the mRNA. These are shown because in nearly all cases examined so far, the half-lives of trypanosome mRNAs have been shown to be determined by the sequence of their 3 -UTR [36–38]. For most trypanosome mRNAs, as in other eukaryotes [39], the first step in degradation is removal of the poly(A) tail (Fig. 2B). The CAF1–NOT complex is responsible for most deadenylation in trypanosomes [40], as in other eukaryotes examined so far [41]. It is built around a large scaffold protein called NOT1. CAF1 has the exoribonuclease activity [42]. The functions of the other subunits – NOT2, NOT5, NOT10, NOT11 and CAF40 – are not yet understood, although the mutual interactions have been characterised, and some structural information is available for the yeast complex. There is also a helicase called DHH1, which is involved in multiple processes. In trypanosomes, but not human cells, NOT10 is needed for stable attachment of CAF1 to the complex [43]. RNAi against either CAF1 or NOT10 almost completely blocks deadenylation and simultaneously delays degradation of most mRNAs [43]. Thus, although CAF1 is active as a monomer on naked mRNAs in vitro, it can only deadenylate mRNAs in vivo when it is associated with the complex. The difference might be caused by the presence of PABP. One possibility is that different subunits of the complex serve to recruit CAF1 to its mRNA substrates for long enough to dislodge PABP. PAN2 is an additional deadenylase, associated with a second protein called PAN3; the trypanosome PAN2/PAN3 complex has some role in deadenylation, since depletion of PAN2 prolongs the half-lives of mRNAs which normally have intermediate stability [44]. It has been postulated that in Opisthokonts, PAN2/PAN3 has a role in initiating deadenylation, or in maintaining correct poly(A) tail lengths [45]. The roles of three trypanosome poly(A) ribonuclease (PARN) homologues are unclear [46]. Once the poly(A) tail length has been reduced to less than 20 nt, PABP can no longer bind and the interaction of the 3 and 5 ends via the translation initiation complex is correspondingly lost (Fig. 2C). Now, the cap can be removed (Fig. 2D). The cytosolic decapping enzymes of Opisthokonts are usually Nudix hydrolases [47]. Trypanosomes have five proteins with Nudix hydrolase domains, but these either have obvious alternative functions, or are in organelles, or are not essential for survival as judged by RNAi experiments. DHH1 is involved in Opisthokont decapping, but affinity purification experiments with trypanosome DHH1 have revealed no proteins that can be linked to decapping. At present there seem to be three options: (a) two of the Nudix hydrolases are involved in decapping, with mutually redundant functions; (b) there is a specific decapping enzyme that is completely unrelated to that of Opisthokonts (could this be a guanylyl transferase operating in reverse?); (c) decapping is effected by a sequencespecific endonuclease that cuts within the spliced leader. One thing that is absolutely certain is that decapping must happen, since we know that mRNAs are subject to 5 –3 degradation by XRNA (Fig. 2E) [15,44]. XRNA is normally a highly processive enzyme: that is, once it has started degrading an mRNA it continues rapidly through it unless a very strong secondary structure gets in the way [44]. In addition to 5 –3 degradation, the deadenylated mRNA body is subject to 3 –5 degradation by a complex called the exosome (Fig. 2E). Nine exosome subunits form a ring-shaped complex, which binds to RNA and disrupts secondary structures, while the


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Fig. 2. Pathways of mRNA degradation in the cytosol. (A) The dark blue open reading frame is being translated. It has a pale green 5 -UTR and a longer 3 -UTR which is able to bind several RBPs. The capped spliced leader is bound by the translation initiation complex, which interacts with PABP. (B) Deadenylation by the CAF1/NOT complex leads to (C) loss of PABP and the interaction with the cap-binding complex. (D) The RNA can now be decapped (DCP) and (E) digested by the exosome and XRNA. During steps B and C, translation initiation can continue. (F) An alternative rapid pathway is initiated by decapping followed by (G) 5 –3 degradation. In this second case, if translation had already initiated, the ribosomes would be able to run off but further initiation would not be possible. A given mRNA sequence might sometimes follow one pathway and sometimes another.

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tenth has the exoribonuclease activity. In Opisthokonts there are two alternative exoribonucleases, Rrp44 and Rrp6 (yeast nomenclature). Rrp44 is cytosolic while Rrp6 is predominantly in the nucleus. The trypanosome exosome is stably associated only with RRP6 and there is no evidence for interaction with RRP44 [48,49]. The function of trypanosome RRP44 – which is an essential enzyme – is currently unclear [50]. There is also a trypanosome gene encoding a homologue of DIS3L2, a 3 –5 exoribonuclease that is related to RRP44; in fission yeast, this enzyme is involved in the 3 –5 degradation of mRNAs that are tagged with oligo (U) tails [51]. The trypanosome genome encodes two more predicted 3 –5 exoribonuclease III proteins that are not predicted to be in the mitochondria; their functions are not known. An alternative pathway proceeds without deadenylation: this involves direct decapping (Fig. 2F) followed by 5 –3 degradation (Fig. 2G). This pathway is important in rapid mRNA destruction, since depletion of XRNA delays degradation of many mRNAs that normally have short half-lives [14,15]. The functions of two additional 5 –3 exoribonucleases, XRNB and XRNC, are not known.

XRNB is a pseudogene in the “genome strain” TREU927, but is intact in the commonly-used Lister 427 strain [15,52]. The kinetics of mRNA decay are often shown as a simple exponential curve. However, for many mRNAs, both in budding yeast [53] and trypanosomes [14], this is not the best fit to the data. In some cases, there is an initial very slow phase, followed by more rapid decay (Fig. 3A). This pattern is shown mainly by mRNAs with relatively long half-lives [14]. The most likely explanation is that at the time of transcription inhibition, most of those mRNAs have rather long poly(A) tails, with only a few being on the verge of losing PABP (Fig. 3A). After transcription inhibition, the tails are gradually shortened. During this time, very few molecules lose their tails, so few are degraded. After a certain interval, larger numbers of mRNAs start to lose PABP; as soon as they have done so, degradation proceeds apace. RNAs with the slow-fast pattern tend to be long-lived; the exponential pattern of decay is mostly also dependent on deadenylation, but with rather faster kinetics. The deadenylation-dependent pathway of mRNA decay can be initiated


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Fig. 3. Possible explanations for complex mRNA degradation kinetics. (A) The slow-fast pathway. Ten mRNAs are shown (cyan), at various times after transcription inhibition. The number of mRNAs remaining at each time point is plotted on the upper left graph. The black line joins the points and the blue line is an exponential decay curve. The plot on the right is real data for histone H4 [14]. At the onset of transcription inhibition, most of them have long poly(A) tails. After 30 min, one has lost the tail completely (pale cyan) and will be destroyed rapidly by decapping, XRNA, and the exosome. As the RNA population gets “older” relative to the start point, the poly(A) tails get progressively shorter. At 90 min, suddenly, 5 of the ten mRNAs have lost the poly(A) tails and the amount of mRNA declines precipitously. (B) The fast-slow pathway. The figure is like (A) except that the real data are for an mRNA with the EP procyclin 3 UTR. In addition, the calculated contributions of the 5 and 3 pathways are indicated, based on data obtained after RNAi [44]. This time, out of the 10 RNAs, 4 have already been decapped after 15 min although none has lost the poly(A) tail. Note that the graphs actually are looking at the cell population. For a low abundance mRNA, there would be less than one mRNA per cell. At any one time, the “choice” between the 5 and 3 pathways is different in individual cells.

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during active translation [54], since only decapping will completely prevent initiation. The second non-exponential decay pattern is very rapid initial degradation, followed by a slower phase (Fig. 3B). This pattern

has been investigated in detail for a reporter mRNA with the EP procyclin 3 -UTR, expressed in bloodstream forms (see for example [44]). It is easiest explained by the existence of two mRNA populations. One population is subject to the rapid, deadenylation


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independent 5 –3 pathway shown in Fig. 2 (F, G), while the remainder is subject to the slower, deadenylation-dependent route. Messenger RNAs with this fast-slow degradation pattern mostly have short half-lives, and are stabilised by depletion of both XRNA and components of the deadenylation machinery (Fig. 3B). This mode of decay could result in degradation of an mRNA before any translation at all can occur.

5. Regulation of translation Trypanosome proteome studies have shown that, as in other eukaryotes, protein levels are only partially correlated with mRNA levels [55–57]. This could reflect differences in translation or protein stability. To find out whether mRNAs also differ in their translation efficiency, Vasquez et al. counted RNA fragments that were bound to ribosomes – a technique called “ribosomal profiling [58]. They found that mRNAs had ribosome densities that varied over 2 orders of magnitude. For an mRNA present at the median value of about 2 copies per cell, that could mean – for example – a difference between 10 ribosomes per mRNA and just 1 ribosome in every 10 mRNAs. Quantitation of polysome-associated mRNAs also showed differences between long slender and growth-arrested stumpy forms [59]. Numerous mechanisms of translation regulation have been documented in eukaryotes, most of which operate at the level of translation initiation [60]. The list below does not mention miRNAmediated mechanisms since these are unlikely to be present in trypanosomes [61].

a) Activation of tRNA is the earliest step, and requires GDP on eIF2␣ to be exchanged for GTP. Phosphorylation of eIF2␣ sequesters eIF2-alpha in the inactive GDP-bound form. Activation of the kinases responsible thus stops all translation. This is an indiscriminate form of regulation that will affect all mRNAs. The existence of eIF2␣ phosphorylation has been documented in Leishmania [62] and T. cruzi [63]. b) Meanwhile, the cap is bound by the eIF4E, which in turn interacts with the eIF4G-eIF4A complex. Activate tRNA binds to a free 40S subunit in the presence of additional translation factors (eIFs 1,2,3 and 5). The resulting “43S complex” binds to eIF4G via eIF3. This step can be prevented by 4E-binding proteins (4E-BP), which bind to eIF4E and prevent its binding to eIF4G. This is – in principle – an indiscriminate form of regulation, but could in theory be made specific by association of 4E-BP with a particular mRNA. Trypanosomes have six different version of eIF4E and eIF4G, but only one cytosolic eIF4A [64,65]. Inconveniently, eIF4E3 interacts with eIF4G4, and eIF4E4 interacts with eiF4G3 [64]. Polysomal proteomes from bloodstream and procyclic forms contain all four of these proteins, while the other eIF4Es and Gs were not detected (C. Klein and C. Clayton, unpublished). The roles of the different eIF4Es have not really been sorted out. They show differing abilities to bind to the highly methylated trypanosome cap, and, confusingly, results for the Leishmania homologues cannot always be extrapolated to trypanosomes. In Leishmania a protein called 4E-IP interacts with eIF4E1; it is not related to 4E-BP [66]. There is evidence that Leishmania eIF4E1 is involved in translation initiation in the amastigote stage and during thermal stress. The interaction with 4E-IP might inhibit eIF4E1 activity in the promastigote stage [66]. eIF4E6 is found in a complex with eIF4G5, which interacts with a third protein, 4G5-IP. 4G5-IP has domains whose secondary structures resemble those of two domains found in capping enzymes: nucleotide triphosphate hydrolase and guanylyltransferase [67].

c) Once the 43S complex has bound the mRNA, it adds additional factors and becomes the 48S complex. The small subunit starts to move along the mRNA, and as soon as it encounters an AUG, the 60S subunit binds and translation is initiated. The progress along the mRNA can be inhibited by the presence of secondary structures or bound proteins in the 5 -UTR. The classic example of an RBP binding to the 5 -UTR is the binding of human iron regulatory protein to the ferritin mRNA [60]. Riboswitches are RNA sequences whose conformation depends on the presence of a small ligand [68]. One can also imagine that tight folding of the 5 -UTR might inhibit scanning. No riboswitches have yet been demonstrated in Kinetoplastids. d) About 20% of all trypanosome mRNAs have 5 -UTRs that contain an AUG upstream of the main coding region, giving a short “upstream open reading frame”. Translation of the main open reading frame depends either on this upstream AUG being ignored by the scanning complex, or on re-initiation after termination. In either case, the efficiency of translation initiation on the main ORF might be decreased. From the ribosome profiling results, on average the presence of an upstream ORF in a trypanosome mRNA decreases the translation efficiency by 25% [58]. In mammals, yeast and plants, the presence of a translated upstream ORF can lead to degradation by the nonsense-mediated decay pathway [69–71]. It is not clear whether a classical nonsense-mediated pathway exists in trypanosomes [72]. e) Recent calculations suggest that in yeast, at least, the rate of translation elongation is as important as initiation in controlling protein synthesis [73]. The activity of the eEF1 complex, which delivers the amino-acylated tRNA to the ribosome, is increased by phosphorylation [74], which will have a global effect on translation. One instance of transcript-specific translational arrest via effects on eEF1A has also been documented [75]. Nothing is known about this in trypanosomes.

6. Granules, degradation and translation In response to stress, mRNAs can be sequestered from the translation and degradation machineries in “stress granules”, from which the mRNA can return to translation upon relief from the stress condition. In metazoan cells, the granules formed after exposure to arsenite involve mechanism (a) above [76]. Heat shock causes elongation pausing and formation of stress granules in mammalian cells [77]. Trypanosomes also form granules in response to starvation or heat shock; and heat shock collapses polysomes [78]. As for metazoan heat shock granules [76], eIF2␣ phosphorylation is not involved [78]. Interestingly, PABP1 and PABP2 show different responses to stress. PABP2 localises much more efficiently to starvation-induced stress granules, and only PABP2 accumulates in granules at the nuclear periphery in response to Sinefungin [33]. The various eIF4E isoforms also differ in their localisation to stress granules but the functional implications are not yet clear [33]. One aspect that has as yet been poorly investigated in trypanosomes is the relationship between translation and mRNA decay. Does translation inhibition result in mRNA decay and if so, does it matter how translation is inhibited? In mammalian cells and yeast, degradation enzymes are concentrated in small foci called P-bodies. P-bodies are smaller than, and distinct from, stress granules although they share some protein components, including SCD6/RAP55. It is thought that mRNAs might be able to be transferred between the two granule types. Inhibition of translation with cycloheximide, which freezes ribosomes on mRNA, prevents formation of P-bodies and stress granules, while treatment with puromycin, which releases the mRNAs from the ribosomes, enhances P-body formation. The relevance of this is


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unclear, though, because the formation of visible P-bodies is not necessary for fully functional mRNA decay. Depletion of SCD6 in trypanosomes results in a two-fold global increase in translation [79], while its over-expression results in stress independent aggregation [80]. So far, the only experiment that addressed the translationdegradation question directly in trypanosomes concerned the GPI-PLC mRNA, which is abundant in bloodstream forms and absent in procyclic forms. Insertion of a translation-inhibitory stem-loop in the 5 -UTR had no effect on the mRNA level in either form [81]. In contrast to this result, cycloheximide treatment has been shown to inhibit developmentally-regulated degradation of several mRNAs, including EP and GPI-PLC. Since EP mRNA degradation was also inhibited by puromycin treatment [82], which dissociates ribosomes from the mRNA, this effect cannot be due to ribosomes protecting the mRNA from degradation. The standard interpretation has therefore been that mRNA stabilisation by cycloheximide is due to “inhibition of synthesis of an unstable regulatory protein” [81]; similar inferences have been made for other trypanosomatids (e.g. [83,84]) However, we have tested six different mRNAs that are not developmentally regulated, and preliminary results suggest that for all of them, both cycloheximide and puromycin increase abundance (C. Helbig, ZMBH, unpublished). This suggests that if the effect is indeed caused by the disappearance of an unstable protein, that protein is likely to be a component of the general RNA degradation machinery. The apparent specificity that has been documented could be explained as follows. If an mRNA has a half-life of under 30 min, inhibition of decay for 60 min should cause an increase in the steady-state level, whereas if an RNA has a half-life of over 60 min, the steady-state level should not be much affected. For example, in bloodstream forms, the mRNA levels of 6 unstable mRNAs encoding the cytochrome oxidase complex were increased after cycloheximide treatment; but the mRNA with the longest half-life, encoding cox IV, showed the smallest effect and that encoding the RNA-binding protein ZFP3 (half-life about 70 min) stayed constant [85]. An alternative to the “unstable protein” hypothesis is that protein synthesis inhibition triggers a stress response that includes a cessation of mRNA decay.

7. Regulation of mRNA decay and translation: the roles of RNA binding proteins The responsibility for normal mRNA homeostasis in trypanosomes resides mainly with RNA-binding proteins (RBPs). Their roles in controlling gene expression have been thoroughly reviewed very recently [36–38]. Here, I will therefore consider only a few examples, concentrating on mechanisms. Some ways in which proteins could influence mRNA decay and translation are illustrated in Fig. 4. Parts A–C each show a polysomal mRNA which is circularised via the PABP–eIF4G interaction; Fig. 4A and B shows various options for endogenous proteins and Fig. 4C is an experimental scenario. Protein (A) destabilises the mRNA by recruiting deadenylases; here I show the CAF1/NOT complex but recruitment of other deadenylases is an option. Protein (B) can stabilise the mRNA by competing with protein (A) for RNA binding. In contrast, protein (C) has an active stabilising role. It could act by recruiting other proteins that can stabilise the mRNA, such as PABP, as shown here; alternatively, it might stabilise the mRNA indirectly, by increasing the efficiency of translation initiation. Proteins (D–F) enhance degradation by recruiting the exosome, the decapping machinery and XRNA, respectively, and protein (G) blocks translation initiation by recruiting 4E-IP. Protein (H) is an abundant protein that is bound to the RNA with low affinity and specificity; it has no

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other interactions and would fall off if a protein with higher affinity were present. The best-characterised example of the scenarios in the figure is the degradation of mammalian mRNAs that have AU-rich elements in their 3 -UTRs [86]. Binding of TTP, Brf-1, KSRP and some isoforms of AUF1 to the sequence results in mRNA decay, and proteins of the HuR family stabilise the mRNAs by preventing binding of the destabilising RBPs. TTP is able to recruit both the Ccr4-Caf1-Not and the decapping complexes, and its activity and interactions are regulated by phosphorylation [86]. More generally, experiments with yeast RBPs have shown that they bind many RNAs, and conversely, each RNA binds several RBPs [87]. Fig. 4 vastly understates the complexity present in real regulation. The median 3 -UTR length in T. brucei is around 400 nt [11–13]; RNA recognition motifs (RRM) and CCCH zinc fingers (CCCH) each bind to only 4 nt, while a pumilio protein with 8 Puf domains binds to 8 nt. Allowing for the presence of multiple motifs in some proteins, and for steric hindrance between bound proteins, and the effects of RNA secondary structure, one could estimate that one protein might bind every 20 nt, so the median 3 -UTR would be able to bind at least 20 RBPs. Very often, several proteins will be competing for binding. For example, no fewer than 8 trypanosome proteins have so far been shown to bind preferentially to U- or AUrich elements: UBP1 and UBP2 [88], ZFP3 [89], ZC3H11 [90] DRBD3 [91,92], DRBD4 [91], DRBD12 and DRBD13 [93]. On top of competition between cytosolic RBPs, and possible post-transcriptional modification of those RBPs, the nuclear history of the mRNA might affect its fate in the cytoplasm, since some proteins that bind in the nucleus might form part of the mRNP. Thus a potential splice site that is not used could carry associated splicing factors with it into the cytosol, where the latter could influence mRNA fate by preventing the binding of cytosolic proteins; HnRNPF/H and DRBD33 are examples [91,94]. Extrapolating the earlier calculation to the 20,000 mRNAs of a bloodstream-form trypanosome, there would be space for 4 × 105 RBPs on the 3 -UTRs. The abundances of very few RBPs are known; estimates include >106 per cell for UBP1 and UBP2 [95], 5 × 104 /cell for PUF5 [96], and about 104 /cell for PUF2 [26]. Even these sparse estimates are enough to make it clear that together, RBPs are present in vast excess relative to their potential binding sites. An abundant, relatively non-specific protein that binds a sequence with low affinity may compete effectively with a more specific, high-affinity binder that is less well expressed. We therefore have to imagine that each UTR carries a multitude of proteins that are all jockeying for position in a dynamic fashion. This complicates interpretation of experimental results: over-expression of one mRNA might divert RBPs from their normal targets, and over-expression of an RBP could result in it binding to an mRNA with which it normally never interacts. Conversely, depletion of one RBP could result in totally abnormal binding of its cognate RNA sequence by some other RBP. Given this scenario, it is not surprising that so far, attempts to identify “the regulatory element” in mRNAs have met with limited success. Often, the sequences required for regulation cannot be narrowed down to less than 200 nt. Deletions in regulatory 3 -UTRs can also give paradoxical results: for example, for a reporter mRNA with the EP 3 -UTR, expressed in procyclic forms, a short deletion abolished expression, but a more extended deletion restored expression again [97]. Interactions of several different proteins are apparently required to get the regulatory flexibility that is optimal for trypanosome survival. Moreover, protein binding could be affected by (or itself change) the RNA conformation (for a possible example see [98]). Even where a relatively short regulatory domain can be identified, it is unlikely that an RBP with a single RNA binding domain can do the job. Puf proteins, or single RRM or CCCH domains,


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have RNA-protein dissociation constants in the 1–2 digit nM range [99–101]. In contrast, a target mRNA present at 30 molecules per trypanosome (which is higher than most) is at a concentration of 0.001 pM. Binding of an RBP with a single domain to the mRNAs is therefore kinetically impossible unless affinities are increased at least 1000-fold by cooperative interactions. One possibility is the presence of more than one RNA-binding domain; otherwise, protein–protein interactions are essential. The mechanism of action of one trypanosome protein, ZC3H11, is at least partially understood. In procyclic forms, ZC3H11 is required for survival after heat shock because it is required for preservation and translation of chaperone mRNAs [90]. In bloodstream forms, it is essential, and its depletion leads to decreased levels of some of the same mRNAs. The single zinc finger of ZC3H11 is specific for the sequence UAUU, and binds in vitro to (UAU) repeats with an apparent dissociation constant of about 10 nM [90]. In vivo, ZC3H11 shows specific binding only to mRNAs with multiple copies of a UAUU sequence, implying that cooperativity is essential [90]. Although ZC3H11 can interact with itself, this is not enough for its activity. Instead, function requires a complex set of protein–protein interactions [102]. In bloodstream forms, ZC3H11 recruits two additional proteins called MKT1 and PBP1, which also mutually interact. Yeast PBP1 interacts with itself, so polymerisation of trypanosome PBP1 might enhance the specificity of the ZC3H11–MKT1–PBP1 complex for (UAU) repeats. In addition, PBP1 also interacts both with another protein of unknown function called LSM12, and the PABPs. ZC3H11 therefore serves as a platform to recruit a complex of MKT1, PBP1, LSM12 and PABP to target mRNAs. To investigate the functions of each protein we used “tethering”. The principle of the method is illustrated for protein “X” in Fig. 4C. It employs a reporter RNA which contains, in its 3 -UTR, a short sequence capable of very high-affinity binding to a protein. In the figure, this is the “boxB” sequence from bacteriophage lambda. The protein of interest (X in the figure) is expressed as a fusion with the protein that binds to the RNA bait: in the figure, this is the lambdaN peptide. Using this method, PABP is a positive regulator in all species tested so far, including trypanosomes. The recruitment of PABP could therefore explain the ability of ZC3H11 to stabilise its targets [102]. We showed that tethered ZC3H11, MKT1, LSM12 and PBP1 are all activators at the normal growth temperature, but that ZC3H11 has to recruit both MKT1 and PBP1 in order to function in bloodstream forms [102]. Whether MKT1 and PBP1 are also required for the heat shock response is not yet known.

8. Finding regulatory proteins Work to identify regulatory RBPs in trypanosomes has so far relied on two approaches. One is to start from a protein that has one of the known RNA binding domains and to try to find out what it does, and the other is to start with the regulated RNA, and to try to find the proteins that regulate it. Candidate RBPs can be chosen based on previous knowledge. The results of a high-throughput RNAi screen [103] are a useful (though not infallible) starting point. For example, one might look for a protein that is only expressed in, and is also essential in, one life-cycle stage [11,58]. Another starting point could be the presence of a conserved RNA-binding domain [90,93]. The most important experimental task – apart from confirming the highthroughput results, and checking that the protein is in the cytosol, is to find the RNA targets. The ideal method is to cross-link it to RNA in vivo, using UV light, affinity purify the protein, then sequence the attached RNA. The binding sites can be identified by addition of RNase to digest any RNA that is not protein-protected. This method has worked well for some proteins that stabilise mRNAs (e.g. [90]), and in one case, binding sites were mapped [104]. Success depends on the presence of sufficient protein-bound RNA, so it is most likely for proteins that stablise the mRNA targets. Another approach is to deplete the protein (usually by RNAi) then examine the transcriptome. If the protein is essential, it is vital to harvest the cells well before growth effects are seen, otherwise secondary effects confound the results [26]. The problem with this RBP-centred approach is that often, it proves very difficult to find out the protein’s function. Also, rsome egulators may not have classical RNA-binding domains. The alternative approach starts at the other end, with a regulated RNA. The regulatory sequence is defined with reporter gene experiments, then one tries to find proteins that bind to it. This can be done by monitoring purification by gel shift analysis, and/or by coupling the sequence to a column, and using it for affinity purification. This latter method can only be used if the sequence is relatively short – ideally, less than 30 nt, but has the advantage of being capable of identifying proteins that do not have classical RNA-binding domains [105]. On the downside, when a cytosolic extract is presented with naked RNA in vitro the spectrum of bound protein will be dictated only by the abundance and binding constants of the proteins for the bait sequence, and not by the history of the mRNA or the multiple protein–protein interactions that can occur on the intact transcript. Indeed, so far the method it has resulted in purification only of abundant RNA-binding proteins, and not

Fig. 4. Possible modes of action of RBPs. The cartoon of the RNA is similar to that in Fig. 3. RNA binding proteins are in black. (A) Protein A recruits the CAF1–NOT complex; B competes with A for RNA binding; C recruits and intermediate protein and PABP; D recruits the exosome; (B) E recruits the decapping enzyme, and F recruits XRNA, G recruits 4E-IP and inhibits translation initiation. Protein H is not doing anything except binding the RNA (and perhaps influencing secondary structure). (C) This cartoon illustrates tethering: protein “X” is fused to the lambda N peptide (␭N), which is has been tethered to a reporter RNA containing the boxB sequence.


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proteins that are actually involved in regulation of the sequence under investigation (e.g. [92]). The ideal strategy to identify proteins that bind a particular mRNA would be to cross-link proteins to RNA in vivo, purify the mRNA, then identify the proteins by mass spectrometry Various strategies for this are possible (discussed in [106]):

(a) One could use an affinity column bearing oligonucleotides that hybridise to the mRNA. (b) One could add specific sequence tags that allow for high affinity purification. This could be done using the tethering approach, but using an easily-purified protein such as streptavidin as the tag protein “X” in Fig. 4C. (c) An RNA “aptamer” sequence that directly binds to a ligand can be used [106].

Each of these methods works for abundant, stable RNA–protein complexes, and each can be used at an analytical scale to demonstrate (by western blotting) that a known protein of interest is present on an mRNA. A strategy of type (a) was used to find all proteins associated with mammalian and yeast poly(A)+ RNA [107–109]; it would be very useful to do this for trypanosomes. For specific mRNAs, at preparative scale, no reliable method is yet available despite attempts in several laboratories (e.g. [106]). This is probably because the number of mRNA molecules available to be purified is very low compared to the number of protein molecules in the cell. For trypanosomes, a well expressed “bait” mRNA might be present at 100 copies in a cell containing at least 108 proteins. If we assume a 20% yield from purification, and that quantitative mass spectrometry can detect 500 proteins, we would need 104 fold purification to have any hope of finding a specific protein. That is a challenging prospect. If the mRNA were unstable, and so in fewer copies, the outlook would be worse. To obtain a list of potential post-transcriptional regulators, we exploited the tethering approach [110]. We used host cells expressing a boxB-containing mRNA encoding selectable marker, and transfected them with a library of plasmids encoding trypanosome lambdaN fusions. The screen identified about 300 potential posttranscriptional regulators from trypanosomes. As in all screens, some regulators will have been missed and some of the 300 candidates are bound to be false positives. Nevertheless, the results did identify known regulators: not only RNA-binding proteins, but also intermediates and direct effectors. Three components of the NOT complex and 15 proteins with RNA-binding motifs were identified as suppressors; the latter included RBP10, which had previously been shown to inhibit translation [111]. Notably, the T. brucei homologue of Leishmania 4E-IP had really strong suppressive activity. The activators of expression included 28 proteins with RNA binding domains, including ZC3H11 and DRBD3, the two PABPs, and PBP1. There were also some translation factors, including eIF4Es 3 and 4, and eiF4Gs 3,4, and 5. We found regulatory activity for several dozen proteins of previously unknown function, several of which are essential for trypanosome survival [112]. Interestingly, the regulators included a few metabolic enzymes, which might be able to form a link between metabolism and RNA degradation. Post-transcriptional regulation involves extensive networks of protein–protein interactions [113,114]. In the future, phenotypic screens with the known effectors of mRNA degradation and translation, together with identification of protein-protein and protein–RNA interactions, are likely to provide information about the mechanisms of regulatory RBPs. Recent technical developments such as high efficiency trypanosome transformation [115], sensitive quantitative mass spectrometry, and the ability to score the

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results of screens using high-throughput sequencing [102,103] will greatly accelerate this process. Acknowledgements Work in my laboratory is funded mainly by the Land of Baden- Q4 Württemberg and the Deutsche Forschungsgemeinschaft. Some results described in this review were from the Sysmo Project “The silicon trypanosome”, funded by the Bundesministerium für Bildung und Forschung. I thank Shula Michaeli (Bar-Ilan University) and Jean Beggs (Edinburgh University) for useful discussions. References [1] Daniels J, Gull K, Wickstead B. Cell biology of the trypanosome genome. Microbiol Mol Biol Rev 2010;74:552–69. [2] Michaeli S. Trans-splicing in trypanosomes: machinery and its impact on the parasite transcriptome. Future Microbiol 2011;6:459–74. [3] Clayton C, Michaeli S. 3 processing in protists. Wiley Interdiscip Rev RNA 2011;2:247–55. [4] Rotureau B, Van Den Abbeele J. Through the dark continent: African trypanosome development in the tsetse fly. Front Cell Infect Microbiol 2013;3:53. [5] Michels PA, Bringaud F, Herman M, Hannaert V. Metabolic functions of glycosomes in trypanosomatids. Biochim Biophys Acta 2006;1763:1463–77. [6] Bringaud F, Riviere L, Coustou V. Energy metabolism of trypanosomatids: adaptation to available carbon sources. Mol Biochem Parasitol 2006;149:1–9. [7] Rudenko G. Epigenetics and transcriptional control in African trypanosomes. Essays Biochem 2010;48:201–19. [8] Roditi I, Liniger M. Dressed for success: the surface coats of insect-borne protozoan parasites. Trends Microbiol 2002;10:128–34. [9] Urwyler S, Studer E, Renggli CK, Roditi I. A family of stage-specific alaninerich proteins on the surface of epimastigote forms of Trypanosoma brucei. Mol Microbiol 2007;63:218–28. [10] Haanstra J, Stewart M, Luu V-D, van Tuijl A, Westerhoff H, Clayton C, et al. Control and regulation of gene expression: quantitative analysis of the expression of phosphoglycerate kinase in bloodstream form Trypanosoma brucei. J Biol Chem 2008;283:2495–507. [11] Siegel T, Hekstra D, Wang X, Dewell S, Cross G. Genome-wide analysis of mRNA abundance in two life-cycle stages of Trypanosoma brucei and identification of splicing and polyadenylation sites. Nucleic Acids Res 2010;38:4946–57. [12] Nilsson D, Gunasekera K, Mani J, Osteras M, Farinelli L, Baerlocher L, et al. Spliced leader trapping reveals widespread alternative splicing patterns in the highly dynamic transcriptome of Trypanosoma brucei. PLoS Pathogens 2010;6:e1001037. [13] Kolev N, Franklin J, Carmi S, Shi H, Michaeli S, Tschudi C. The transcriptome of the human pathogen Trypanosoma brucei at single-nucleotide resolution. PLoS Pathogens 2010;6:e1001090. [14] Fadda A, Ryten M, Droll D, Rojas F, Färber V, Haanstra J, et al. Transcriptomewide analysis of mRNA decay reveals complex degradation kinetics and suggests a role for co-transcriptional degradation in determining mRNA lev- Q5 els. (submitted for publication). [15] Manful T, Fadda A, Clayton C. The role of the 5 –3 exoribonuclease XRNA in transcriptome-wide mRNA degradation. RNA 2011;17:2039–47. ˜ J, Wirtz LE, [16] Biebinger S, Rettenmaier S, Flaspohler J, Hartmann C, Pena-Diaz et al. The PARP promoter of Trypanosoma brucei is developmentally regulated in a chromosomal context. Nucleic Acids Res 1996;24:1202–11. [17] Huang J, van der Ploeg L. Maturation of polycistronic pre-mRNA in Trypanosoma brucei: analysis of trans splicing and poly(A) addition at nascent RNA transcripts from the hsp70 locus. Mol Cell Biol 1991;11: 3180–90. [18] Ullu E, Matthews KR, Tschudi C. Temporal order of RNA-processing reactions in trypanosomes: rapid trans splicing precedes polyadenylation of newly synthesized tubulin transcripts. Mol Cell Biol 1993;13:720–5. [19] Kilchert C, Vasiljeva L. mRNA quality control goes transcriptional. Biochem Soc Trans 2013;41:1666–72. [20] Cristodero M, Clayton C. Trypanosome MTR4 in involved in ribosomal RNA processing. Nucleic Acids Res 2007;35:7023–30. [21] Etheridge RD, Clemens DM, Gershon PD, Aphasizhev R. Identification and characterization of nuclear non-canonical poly(A) polymerases from Trypanosoma brucei. Mol Biochem Parasitol 2009;164:66–73. [22] Serpeloni M, Moraes CB, Muniz JR, Motta MC, Ramos AS, Kessler RL, et al. An essential nuclear protein in trypanosomes is a component of mRNA transcription/export pathway. PLoS ONE 2011;6:e20730. [23] Dostalova A, Kaser S, Cristodero M, Schimanski B. The nuclear mRNA export receptor Mex67-Mtr2 of Trypanosoma brucei contains a unique and essential zinc finger motif. Mol Microbiol 2013. [24] Mor A, Suliman S, Ben-Yishay R, Yunger S, Brody Y, Shav-Tal Y. Dynamics of single mRNP nucleocytoplasmic transport and export through the nuclear pore in living cells. Nat Cell Biol 2010;12:543–52.


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Developmental regulation of nuclear gene expression in Trypanosoma brucei.

High-throughput Gene Tagging in Trypanosoma brucei.

The actin gene promoter of Trypanosoma brucei.

Transcript-specific developmental regulation of polyadenylation in Trypanosoma brucei mitochondria.

Sequence of a tRNA gene cluster in Trypanosoma brucei.

SUMOylation in Trypanosoma brucei.

Localization of serum resistance-associated protein in Trypanosoma brucei rhodesiense and transgenic Trypanosoma brucei brucei.

Differential expression of the oligomycin-sensitive ATPase in bloodstream forms of Trypanosoma brucei brucei.

Anatomy of the parp gene promoter of Trypanosoma brucei.

Allelic polymorphism of the Trypanosoma brucei polyubiquitin gene.

Bloodstream and metacyclic variant surface glycoprotein gene expression sites of Trypanosoma brucei gambiense.

De Novo Genome Assembly Shows Genome Wide Similarity between Trypanosoma brucei brucei and Trypanosoma brucei rhodesiense.

The promoter for a variant surface glycoprotein gene expression site in Trypanosoma brucei.

Nitric oxide-mediated cytostatic activity on Trypanosoma brucei gambiense and Trypanosoma brucei brucei.

Trypanosoma evansi is alike to Trypanosoma brucei brucei in the subcellular localisation of glycolytic enzymes.

A vanillic acid inducible expression system for Trypanosoma brucei.

Characterization of different proteolytic activities in Trypanosoma brucei brucei.

Integrative analysis of the Trypanosoma brucei gene expression cascade predicts differential regulation of mRNA processing and unusual control of ribosomal protein expression.

Cure of Trypanosoma brucei brucei and Trypanosoma brucei rhodesiense infections in mice with an irreversible inhibitor of S-adenosylmethionine decarboxylase.

Diminazene aceturate residues in the tissues of healthy, Trypanosoma congolense and Trypanosoma brucei brucei infected dogs.

Cultivation of bloodstream Trypanosoma brucei.

Structures of adenosine kinase from Trypanosoma brucei brucei.

Characterization of an endopeptidase of Trypanosoma brucei brucei.

Transient activity assays of the Trypanosoma brucei variant surface glycoprotein gene promoter: control of gene expression at the posttranscriptional level.