The human pathogenic parasite Trypanosoma brucei possess both de novo and salvage routes for the biosynthesis of pyrimidine nucleotides. Consequently, they do not require salvageable pyrimidines for growth. Thymidine kinase (TK) catalyzes the formation of dTMP and dUMP and is one of several salvage enzymes that appear redundant to the de novo pathway. Surprisingly, we show through analysis of TK conditional null and RNAi cells that TK is essential for growth and for infectivity in a mouse model, and that a catalytically active enzyme is required for its function. Unlike humans, T. brucei and all other kinetoplastids lack dCMP deaminase (DCTD), which provides an alternative route to dUMP formation. Ectopic expression of human DCTD resulted in full rescue of the RNAi growth phenotype and allowed for selection of viable TK null cells. Metabolite profiling by LC-MS/MS revealed a buildup of deoxypyrimidine nucleosides in TK depleted cells. Knockout of cytidine deaminase (CDA), which converts deoxycytidine to deoxyuridine led to thymidine/deoxyuridine auxotrophy. These unexpected results suggested that T. brucei encodes an unidentified 5' -nucleotidase that converts deoxypyrimidine nucleotides to their corresponding nucleosides, leading to their dead-end buildup in TK depleted cells at the expense of dTTP pools. Bioinformatics analysis identified several potential candidate genes that could encode 5’-nucleotidase activity including an HD-domain protein that we show catalyzes dephosphorylation of deoxyribonucleotide 5’-monophosphates. We conclude that TK is essential for synthesis of thymine nucleotides regardless of whether the nucleoside precursors originate from the de novo pathway or through salvage. Reliance on TK in the absence of DCTD may be a shared vulnerability among trypanosomatids and may provide a unique opportunity to selectively target a diverse group of pathogenic single-celled eukaryotes with a single drug.
The parasitic trypanosomatids are vector-borne single-celled eukaryotic pathogens that cause significant disease and mortality in tropical and subtropical countries [1]. Human African trypanosomiasis (HAT), Leishmaniasis and Chagas disease affect 20 million people combined, but control is hampered by lack of good drugs, drug resistance and challenges in drug administration [2,3]. New drugs for the treatment of all three diseases are badly needed. HAT, also known as sleeping sickness, is caused by Trypanosoma brucei, an extracellular parasite that replicates in the blood in early stages of infection. It crosses the blood-brain barrier in later stages leading to progressive neurological complications that disrupt the sleep/wake cycle and which eventually progress to coma and death [4]. The WHO estimates that yearly cases have dropped below 10,000 [5]; however, attempts to fully eliminate the disease face the difficulties of a substantial animal reservoir and problematic drug therapies that have high toxicity or are difficult to administer in a rural setting. Pyrimidine and purine biosynthesis is essential in trypanosomatids to generate precursors needed for the biosynthesis of DNA, RNA and sugar nucleotides [6,7]. Purines are obtained entirely by salvage routes through an array of interconnected and seemingly redundant pathways [8]. However, despite this redundancy, enzymes from the purine pathway including GMP synthase have been shown to be essential for pathogenicity in vivo [9]. In contrast, trypanosomatids are able to synthesize pyrimidines either through the de novo biosynthetic pathway or through salvage of preformed nucleosides and bases [7,10,11]. Genes have been identified for the complete de novo pyrimidine biosynthetic pathway, for several key salvage enzymes and for a number of interconversion enzymes [12] (Fig 1). Genetic knockout studies have shown that loss of various de novo pyrimidine biosynthetic enzymes leads to pyrimidine auxotrophy that can be rescued by exogenous uracil [13–15]. These findings are consistent with reports that uracil transport is the primary route for pyrimidine salvage [10]. However knockout of UMP synthase lead to avirulence in mice suggesting that in vivo pyrimidine salvage may be insufficient to completely overcome loss of the de novo pathway [15]. These studies have shown that despite apparent redundancy, enzymes in both the pyrimidine and purine biosynthetic pathways can be essential, especially for in vivo virulence of T. brucei, which as an extracellular parasite lacks access to high intracellular concentrations of metabolites. T. brucei lack several transporters and enzymes found in higher eukaryotes that may make them more vulnerable to disruption of the pyrimidine biosynthetic pathway. The primary pyrimidine transporter in T. brucei preferentially takes up uracil, whereas transport of uridine, 2’-deoxyuridine, thymidine and cytidine is either non-existent or inefficient requiring high nucleoside concentrations [10]. Trypanosomatids lack dCMP deaminase (DCTD), an important contributor to dTTP biosynthesis through deamination of dCMP to dUMP in many higher eukaryotes [17,18]. Instead trypanosomatids were thought to rely on dUTPase to convert uracil nucleotides synthesized by the de novo pathway into the thymine nucleotide pools [19]. T. brucei encodes three pyrimidine salvage enzymes: uracil phosphoribosyltransferase (UPRT), thymidine kinase (TK), and uridine phosphorylase (UPP), and additionally a cytidine deaminase (CDA) that can convert deoxycytidine to deoxyuridine (Fig 1). T. brucei UPP was shown to prefer uridine and deoxyuridine as substrates, and the enzyme was reported not to be essential based on RNAi knockdown studies [20]. In contrast to mammalian cells, which encode both cytosolic TK1 and mitochondrial TK2 [21–23], trypanosomatids possess only TK1. TbTK is however a unique fusion of two TK domains that function as a pseudodimer. The N-terminal domain is catalytically inactive, while the C-terminal domain exhibits canonical TK activity [24]. Recent RNAi studies have suggested that TK is essential but a mechanistic understanding for why TK would be required has not emerged [25,26]. Herein we report the results of genetic studies to determine the roles of pyrimidine salvage enzymes in pyrimidine nucleotide biosynthesis, growth and virulence in T. brucei. We found that knockdown of either TK or CDA led to cell death despite the finding that T. brucei does not require exogenous pyrimidines for growth. The TK conditional null mutant (c-null) could not be rescued by exogenous pyrimidines, whereas the CDA null was auxotrophic for thymidine or deoxyuridine. We were able to abolish the parasite’s reliance on TK by providing an alternative metabolic route to dUMP formation via DCTD. Metabolomic analysis showed that a significant cellular response to TK depletion was dead-end formation of pyrimidine nucleosides resulting in depletion of dTTP. These data taken together with the phenotype of the CDA null suggested the presence of an unidentified 5' -nucleotidase that promotes interconversion between the cytosine and thymine deoxynucleotide pools. Bioinformatics analysis suggested that T. brucei encodes a number of potential 5’-nucleotidases and we show that an HD-domain protein related to a well-characterized bacterial 5’-nucleotidase is able to catalyze this reaction. Thus our data show that TK is essential for the de novo biosynthesis of dTTP and that together with CDA and the 5’-nucelotidase these enzymes play a key role in maintaining balanced levels of the various deoxyribonucleotide pools. We propose that the absence of DCTD in trypanosomes, and other protists, may be a shared vulnerability, presenting a potential opportunity to develop a pan-trypanosomatid TK inhibitor to exploit this unique feature of the protozoan pyrimidine pathway.
To evaluate the essentiality of TK, we attempted to generate a TK null cell line in T. brucei BSF single marker (SM) cells. T. brucei is diploid, therefore minimally two alleles are present for each gene. Allelic replacement via homologous recombination was achieved by transfecting parasites with a PCR product containing a resistance marker flanked by the TK 5’ and 3’ UTRs. Removal of the first allele was successful; however we were unable to replace the last allele after repeated attempts, suggesting essentiality. To further evaluate this hypothesis, a TK conditional null (TK c-null) cell line was generated. The single allele knockout cells (SKO) were transfected with a vector conferring tetracycline (Tet) regulated expression of N-terminally FLAG tagged T. brucei TK (FLAG-TbTK). Expression of FLAG-TbTK was induced by addition of Tet and the remaining TK allele was successfully removed generating the TK c-null cell line. PCR amplification of the TK locus confirmed replacement by the two selectable markers (S1 Fig). To determine the effects caused by the loss of TK expression on cell growth, Tet was removed from the medium, which led to rapid growth arrest and near total cell death by day 3 (Fig 2A). Coincident with this growth arrest, TK transcript (by qPCR) and the TK protein (western blot) were depleted within 24 h after Tet removal, confirming good regulatory control of the ectopic TK copy (Fig 2A and 2B). Parasites reemerged several days later, likely due to the loss of Tet regulation. This has been reported to be a common phenomenon in T. brucei, likely due to mutations resulting in the loss of Tet regulation, e. g. [9,27–29]. The TK c-null cells grew normally in pyrimidine-free medium (containing dialyzed fetal bovine serum) in the presence of Tet confirming that T. brucei is not auxotrophic for pyrimidines (Fig 2A). Upon subsequent removal of Tet, TK depletion led to cell death with a similar time course to medium containing non-dialyzed (normal) serum. In vivo studies were performed to determine if TK was essential to support T. brucei infection in mice. In parallel to T. brucei SM infected mice, two groups (n = 3) of TK c-null infected mice were given either doxycycline (Dox) treated water or water only. As expected mice infected with SM cells in either condition (+/- Dox) had detectable levels of parasitemia by 72 h post infection, with fatalities occurring in all mice in both groups by day 6 (Fig 2C and S2 Fig). Similarly, all TK c-null infected mice treated with Dox to maintain expression of the Tet-regulated TK copy eventually died within the timeframe of the study. One mouse in this group showed a delayed time before succumbing to parasitemia suggesting some variability in TK expression levels in the TK c-null cells. Mice infected with the TK c-null strain treated only with water remained healthy and had no detectable levels of parasitemia past 30 days. Thus we conclude that TK is essential for T. brucei virulence and infectivity in vivo. The finding that TK is essential in T. brucei is puzzling as no clear mechanistic role for TK in parasite fitness is apparent. T. brucei requires TK, which is a pyrimidine salvage enzyme; yet there is no requirement for salvageable pyrimidines for growth. To gain further mechanistic insight into this conundrum, we sought to address three possible explanations for the essentiality of TK: 1) parasites require an active TK enzyme, but it makes a novel product; 2) the TK protein, but not its catalytic activity is needed in some regulatory capacity; 3) parasites require formation of dUMP/dTMP by TK to balance pathway flux even under conditions where all pyrimidine precursors originate from the de novo pathway. To provide additional mechanistic insight, an inducible RNAi cell line targeting TK mRNA was created so that we could easily introduce various rescue plasmids to address our mechanistic hypotheses. A Tet-regulated vector capable of producing a hairpin transcript targeting the 3’UTR of TK was generated and transfected into the TK SKO cell line. Induction of TK RNAi by addition of Tet led to a significant growth defect, although the growth defect was not as severe as observed for TK c-null cells (Fig 3A and 3B). TK mRNA expression was reduced to 20–25% of wild-type control levels by RNAi targeting the TK transcript (Fig 3C). However, the reduction in TK transcript levels was less in comparison to that observed in the TK c-null cells, explaining why the effect on cell growth was less pronounced. To shed light on whether a novel TK product was being formed we transfected parasites with plasmids encoding rescue proteins from three different sources: AU1-tagged T. brucei TK, FLAG-tagged human HsTK, and Herpes simplex TK (HsvTK). HsTK has been shown to have more stringent substrate specificity than the T. brucei enzyme [24], whereas the viral HsvTK possesses broader substrate specificity than the human enzyme [30]. Rescue protein expression was also under control of Tet promoter. Thus, addition of Tet to these cells induces simultaneous knockdown of endogenous TK and expression of the tagged rescue protein. We found that the growth phenotype was reversed by expression of TK from all three species: T. brucei (TbTK) (Fig 3A), human (HsTK) (Fig 3B) and viral TK (HsvTK) (S3 Fig). Expression of TbTK and HsTK was confirmed by western blot (Fig 3A and 3B) and knockdown of endogenous TK was monitored by qPCR (Fig 3C). Viral TK expression was confirmed by the observance of ganciclovir sensitivity that was less apparent in cells expressing T. brucei TK (S3 Fig). Ganciclovir is a subversive substrate of HsvTK leading to premature chain termination of newly synthesized DNA [31]. The ability of both HsTK and HsvTK to rescue the TK RNAi growth phenotype shows that T. brucei TK is unlikely to catalyze a novel reaction, as the required activity is present in enzymes from other species that are known to have a range of substrate specificities. To confirm that TbTK’s essential function is dependent on catalytic activity, mutations in the active site of both TbTK and HsTK were created. We targeted two conserved residues (S4 Fig) with described roles in the TK catalytic mechanism: T. brucei E286, which is reported to function as a proton acceptor [32] and human K32 which is an essential ATP binding residue [33]. Rescue plasmids were constructed as described above with the mutant TKs under the control of the Tet promoter and transfected into the TK RNAi line. In contrast to the wild type enzymes, neither the TbTK E286A nor HsTK K32I active site mutants were able to reverse the RNAi induced growth phenotype (Fig 3D and 3E). These data demonstrate that TK catalytic activity is required for its role in T. brucei cell survival. While our data clearly show that T. brucei is not a pyrimidine auxotroph, we exploited the fact that the TK RNAi line retains partial TK activity (the knockdown is only 75–80% effective by RNAi (Fig 3C) ) to assess whether we could use pyrimidine rescue to determine which TK product was needed for T. brucei growth. We found that high concentrations (significantly above physiological levels) of deoxyuridine (dUrd) resulted in the partial rescue of the RNAi growth phenotype (Fig 4A and S5A Fig) (5 mM rescued but 1 mM did not). However, similar levels of uridine (Urd) (Fig 4B and S5B Fig) or thymidine (dThd) (Fig 4C and S5C Fig) did not restore growth and dThd (0. 15–1. 0 mM) was in fact growth inhibitory to the TK RNAi +Tet induced cells but not to cells that expressed TK (-Tet), suggesting some type of feedback regulation. In contrast, the addition of dUrd or uracil to TK c-null cells, which are >99% depleted of TbTK, were unable to circumvent lethality of the TK knockout showing that TK activity is required for dUrd rescue (Fig 4C and S6 Fig). These data confirm that TK plays an essential role in maintaining dUMP pools. The ability of dUrd to partially reverse the RNAi growth phenotype highlights an interesting feature in T. brucei pyrimidine metabolism. In most mammals, it has been suggested that a significant portion of dTTP is derived from dUMP produced by dCMP deaminase (DCTD) [17,18]. Trypanosomatids lack this enzyme, restricting the number of metabolic routes dedicated to dUMP formation. We hypothesized that due to the lack of DCTD, trypanosomatids require TK to supplement dUMP pools. To test this hypothesis a Tet-regulated vector encoding human DCTD (HsDCTD) was transfected into the TK RNAi cell line to drive simultaneous expression of HsDCTD and knockdown of endogenous TbTK. The expression of FLAG-tagged HsDCTD completely rescued the TK RNAi-induced growth phenotype (Fig 5A). Expression of FLAG-tagged HsDCTD was confirmed by western blot and qPCR analysis confirmed that TK mRNA expression was simultaneously reduced (Fig 5B). To further demonstrate that HsDCTD can functionally replace TK, a TK null cell line was created in the background of FLAG-tagged HsDCTD Tet-regulated expression plasmid. Both TK alleles were replaced by selectable markers through homologous recombination in the presence of Tet to maintain expression of HsDCTD. PCR amplification of the region flanking the TK 5’ and 3’ UTRs confirmed replacement of TK with the selectable markers (S1 Fig). PCR analysis also confirmed that the TK gene was no longer detectable in genomic DNA from the TK null cells. Removal of Tet from this cell line led to depletion of FLAG-tagged HsDCTD and resulted in a severe growth phenotype by day 2 after Tet removal (Fig 5C). Cell growth of the HsDCTD TK null line was less severely impacted than the TK c-null line expressing Tet-regulated TbTK from the rescue plasmid, perhaps reflecting a higher residual expression level of HsDCTD (Fig 5C). By day 5 after Tet withdrawal cells began growing again coincident with re-expression of HsDCTD, again suggestive of emergence of cells that have mutations leading to loss of Tet regulation (Fig 5C). Thus, the data support the hypothesis that TK is essential in T. brucei and that its role is to contribute to the formation of dUMP in the absence of DCTD. Analysis of ~130 soluble metabolites from TK c-null cells was performed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) to determine the impact of TK depletion on metabolite pools (Fig 6 and S7–S9 Figs and S1 Appendix). Extracts were collected from TK c-null cells grown in medium containing normal non-dialyzed serum at 24 h ± Tet. An early time point was selected so that metabolite pools would be less affected by non-specific changes resulting from cell death later in the time course. Significant changes in the measured metabolite levels were mostly confined to pyrimidine nucleosides: there was a 15-30-fold accumulation of the TK substrates dUrd and dThd, a 3-fold accumulation of dCtd, and a 70-fold increase in thymine levels (Fig 6A). In contrast, the pyrimidine nucleotides CMP and UMP were not significantly changed by TK depletion, suggesting that the de novo pathway was able to maintain the uridine nucleotide pools. In further support, HPLC analysis of UDP-sugars was performed (Fig 6B). Nucleotide sugars are formed from UTP, thus their measurement provides a read-out of effects on intracellular UTP concentrations. The relative abundance of UDP-GlcNAc, UDP-Galactose, and UDP-Glucose were not significantly changed by TK depletion, confirming that UTP pools are not linked to TK activity. Thymidine nucleotide pools were not detected by the LC-MS/MS analysis so instead we quantitated dTTP levels using an enzymatic assay and found that dTTP levels were reduced to 30% of control levels 48 h after Tet removal, confirming TK is essential for synthesis of dTTP (Fig 6C). To better understand the consequences of TK depletion in a pyrimidine-free medium the LC-MS/MS analysis was repeated for TK c-null cells grown in medium supplemented with dialyzed serum (Fig 6C and S7–S9 Figs and S1 Appendix). Similar to the results for cells grown in normal serum based medium, we observed a statistically significant buildup of TK substrates dUrd and dThd, and of the dUrd precursor dCyd, though the increases (3-6-fold) were less than observed for cells grown in normal serum based medium. We also observed changes in the levels of TCA intermediates including decreases in citrate and aconitate and an increase in α-ketoglutarate, and perturbations in other metabolites related to pyrimidine biosynthesis including decreased carbamoylphoshate (CP) and acetyl-ornithine (30 and 80%, respectively) and a 2-fold increase in homocysteine (Fig 6C and 6D). Not all of these latter changes reached statistical significance. Some modest effects were also observed on several purines: an increase in xanthine and/or hypoxanthine was observed in both medium conditions, which may suggest some type of cross-regulation between pyrimidine and purine biosynthetic pathways. Thymine was only observed in cells grown in medium supplemented with non-dialyzed serum, which could supply a source for thymine, confirming genomic analysis that suggests T. brucei does not encode thymine biosynthetic enzymes. The finding that the deoxypyrimidine nucleoside pools buildup upon TK depletion for cells grown in pyrimidine-free (dialyzed serum based) medium suggests that an undiscovered biosynthetic route for formation of deoxypyrimidine nucleosides must be present in T. brucei. To investigate if deoxycytidine could be involved in formation of dUMP and dTMP we decided to characterize the effects of depleting cytidine deaminase (CDA) on parasite growth. Based on the current annotation of the pyrimidine biosynthetic pathway, CDA like TK should catalyze a redundant reaction, since the deoxynucleotide pools can be supplied from the de novo pathway. However, attempts to generate a CDA null were unsuccessful, and indeed the CDA null could only be obtained in the presence of 500 μM dThd (Fig 7 and S10 Fig). Upon removal of dThd, CDA null cells exhibited a severe growth defect (Fig 7A). Cells cultured for longer eventually died around day 7–10 after dThd removal. Additional pyrimidine rescue studies revealed that both dThd and dUrd were capable of rescuing growth (EC50 of 6–20 μM), whereas uracil (up to 250 μM) was not (Fig 7C–7E). The concentrations of dThd and dUrd required for rescue are significantly above reported human blood levels (range of 0. 2–0. 6 μM; http: //www. hmdb. ca). CDA null cells were grown ± dThd for 12 h in media supplemented with dialyzed FBS and the metabolite pools were analyzed by LC-MS/MS as described above. Overall the effects of CDA depletion on cellular metabolism were very similar to the effects observed after depletion of TK, confirming a link between the roles of the two enzymes (S11 Fig, S12 Fig and S1 Appendix). We observed a significant buildup (~7-fold) in the CDA substrate, dCtd, for the CDA null cells grown in the absence of Thd that was accompanied by changes in TCA intermediates and other metabolites that serve as precursors for de novo pyrimidine biosynthesis. These changes include increases in α-ketoglutarate (11-fold), glutamate (7-fold) and homocysteine (2-fold), and decreases in carbamoyl phosphate (2-fold). We also observed decreased levels of several amino acids, of polyamines, particularly N-acetyl putrescine and decreased levels of several purine mono-phosphate nucleotides. Neither dUrd nor dThd were detected in CDA null cells in the presence or absence of added dThd, suggesting they were either not formed (dUrd) or rapidly metabolized (dThd). In the absence of CDA, the buildup of dCtd suggests that the pyrimidine deoxynucleotides have been sequestered into a dead-end product that would be expected to lead to depletion of key nucleotides and to cell death as was observed for the TK null cells. These data support a role for CDA in the interconversion of deoxycytidine, deoxyuridine and thymidine pools, which is needed to balance these pools in a cell like T. brucei that expresses TK but not DCTD. In order to identify the potential enzyme (s) responsible for converting deoxypyrimidine nucleotides into their corresponding nucleosides we undertook a bioinformatics analysis of the T. brucei genome. 5' -nucleotidase activities that catalyze formation of deoxyuridine, deoxycytidine, or thymidine from their respective mono-phosphate nucleotides are ascribed to two EC numbers (EC3. 1. 3. 5 and EC3. 1. 3. 89). Inspection of domain types performing these activities revealed nine protein families defined by PFAM, with all having homologous structure representatives in the PDB (Table 1). The families further merge into five different homologous fold types; including three different α/β sandwich folds (phosphoglycerate mutase-like, SurE-like, and HAD domain-related), one α+β four-layer sandwich fold (metallo-dependent phosphatases), and one all-α fold (HD-domain). The phosphoglycerate mutase-like representatives are limited to mammalian acid phosphatases, prostate (ACPP) enzymes that convert extracellular AMP to adenosine (i. e. ecto 5' nucleotidase activity) [34]. An additional identified enzyme (NT5E) from the metallo-dependent phosphatase fold group exhibits a similar ecto 5' -nucleotidase activity [35]. Examples of this fold group in both eukaryotes and bacteria contain signal peptides and are extracellular. In contrast, the remaining SurE, HAD domain-related, and HD-domain representatives appear cytosolic. We found evidence for all of the 5' -nucleotidase homologous fold types in the T. brucei genome with the exception of the SurE fold class (S1 Table). We identified nine phosphoglycerate mutase-like sequences, 33 metallo-dependent phosphatase sequences, 16 HAD domain-related sequences, and one HD-domain sequence (S1 Table). The presence of numerous examples of potential 5' -nucleotidase domains, many of which are annotated as hypothetical proteins, suggests multiple possible proteins that T. brucei could use to form deoxyuridine, deoxycytidine, or thymidine de novo. However, three of the identified genes possess specific PFAM domains described as having 5’-nucleotidase activity (EC 3. 1. 3. 5 or EC 3. 1. 3. 89) and thus are the highest ranked candidates. One encodes a HD domain protein: hypothetical protein (Tb09. 211. 2190) and two encode HAD-like domains: a putative p-nitrophenylphosphatase (Tb927. 8. 7510) and a hypothetical protein (Tb09. 211. 1880). The T. brucei HD domain protein is a homolog of E. coli 5’-nucleotidase YfbR (S13 Fig) while the T. brucei HAD-domain proteins are related to enzymes shown to have 5’-nucleotidase activity in both yeast and E. coli. [36–38]. These enzymes have been reported to have broad substrate specificity functioning on all three pyrimidine deoxyribose monophosphates. To provide support for our hypothesis that T. brucei encodes a 5’-nucleotidase we cloned, expressed and purified the recombinant T. brucei HD domain homolog (Tb09. 211. 2190) of bacterial 5’-nucleotidase YfbR (S14 Fig). We found that the T. brucei YfbR-like HD protein showed a metal dependent 5’-nucleotidase activity (Fig 8A). Similar to the bacterial enzyme it was most active in the presence of Co+2 (0. 5 mM), but activities within 2-4-fold of levels observed for Co+2 were also obtained using Mn+2 (0. 5 mM) and physiological levels of Mg+2 (10 mM). No activity was observed in the presence of Zn+2 or EDTA. The specific activity of the T. brucei HD domain 5’-nucleotidase was very similar to the reported activity of E. coli YfbR [38]. The T. brucei enzyme showed a broad substrate range functioning on both pyrimidine and purine deoxyribonucleoside and ribonucleoside 5’-monophosphates, though it was most active on the deoxypyrimidine nucleotides (dCMP, dUMP and dTMP) (Fig 8B). It showed no activity towards diphosphate nucleotides. The T. brucei enzyme was somewhat more promiscuous then E. coli YfbR, which was unable to catalyze hydrolysis of ribonucleoside 5-monophosphates [38]. Both T. cruzi and Leishmania encode homologs of the T. brucei HD-domain 5’-nucleotidase (S1 Table and S13 Fig) suggesting they both also will be able to convert 5’-deoxyribonucleotide monophosphates to their respective nucleosides. To assess if TK essentiality was likely to extend to other pathogenic protozoa we utilized the KEGG pathway database to determine the distribution of TK and DCTD throughout eukaryotes (Fig 9). A striking disparity was observed within protists when compared to higher eukaryotes. The vast majority of higher eukaryotes possess both TK and DCTD, which may explain TK’s non-essential role in these organisms. In contrast, the kinetoplastids and a number of other protozoan human pathogens such as Giardia encode only TK, suggesting that TK may be essential in these organisms as well. We also note that several of the protists such as Entamoeba histolytica, which lack DCTD, instead encode dCTP deaminase, an enzyme found almost exclusively in bacteria. Similar to DCTD, the ability to deaminate dCTP to dUTP offers an alternative path from cytosine to thymine nucleotide pools and thus we would predict that TK would not be essential in these species. Interestingly, these organisms represent the only eukaryotic KEGG organisms that have dCTP deaminase.
T. brucei encodes a complete de novo pyrimidine biosynthetic pathway, as well as a number of pyrimidine salvage enzymes that were thought to be redundant based on the presence of the de novo pathway. Herein we describe the first comprehensive analysis of the role of the pyrimidine salvage enzymes in T. brucei and we show that while T. brucei is not auxotrophic for pyrimidines, both TK and CDA are essential for in vitro growth and TK is essential for infectivity in vivo as well. The finding that these enzymes are essential could not be explained by the current annotation of the pyrimidine pathway in T. brucei. Our mechanistic analysis of the TK and CDA null cell lines uncovered the existence of an interconversion network between the deoxypyrimidine nucleoside and nucleotide pools, including the presence of a previously unknown 5’-nucleotidase that converts deoxycytidine, deoxyuridine and thymidine nucleotides to their respective nucleosides. In the absence of TK or CDA to balance this 5’-nucleotidase activity, the metabolic cycle breaks down leading to dead-end buildup of deoxypyrimidine nucleosides and to cell death. The existence of this recycling pathway provides a mechanism for the parasite to interconvert and balance the relative levels of the deoxyuridine, deoxycytidine and thymidine pools whether they originate from the de novo pathway or through salvage. Our conclusions are supported by the following arguments. Firstly, TK is essential for both in vitro growth and infectivity in a mouse model of T. brucei infection and for formation of dTTP despite the fact that T. brucei is not a pyrimidine auxotroph. Thus the essential role of TK is not to salvage externally acquired pyrimidine precursors. Our data clearly show that TK activity is required for its function and that it plays a key role in the synthesis of dUMP, even for cells grown in a pyrimidine-free environment. We found that the function of TK can be replaced by expression of human DCTD, which provides an alternative route to dUMP formation from dCMP in many higher eukaryotes [17,18]. DCTD has been shown to be essential for cell cycle progression and formation of dTTP pools in eukaryotes that lack TK (e. g. Schizosaccharomyces pombe) [39]. These data suggest that DCTD and TK can have functionally redundant roles in contributing to dTTP pools, supporting our observation that TK is essential for formation of thymine nucleotides in T. brucei. The next significant key to the puzzle came from analysis of metabolomic data from the TK c-null cell line. These data showed that even in the absence of external pyrimidines the TK substrates dUrd and dThd, as well as the dUrd precursor dCtd buildup, leading to a dead-end accumulation of these precursors away from the essential deoxynucleotide pools resulting in depletion of dTTP. In the absence of an exogenous supply of these nucleosides the current annotation of the T. brucei genome does not provide a mechanism for these nucleosides to be synthesized, suggesting the presence of a missing enzyme that catalyzes conversion of deoxynucleotides into deoxynucleosides. The findings that CDA null cells are auxotrophic for dThd or dUrd further support this hypothesis since based on redundancy in the pathway, CDA should not be essential under any conditions. Furthermore the CDA null data support the presence of an enzymatic link between the deoxycytidine-containing nucleotide pools and dCtd/dUrd, since either dUrd or Thd are required for growth of CDA null cells. These data are consistent with previous published untargeted metabolomics data showing that isotope-labeled glucose was incorporated into both dUrd and dThd, and thus that T. brucei was capable of synthesizing these nucleosides de novo [40]. Lastly, the inability of uracil to rescue the growth deficit of the CDA null cells shows that uridine phosphorylase is not able to efficiently convert uracil to dUrd, eliminating the only known potential source for dUrd biosynthesis in T. brucei. Uridine phosphorylase was previously suggested to be the source of dUrd, based on the isotope-labeled glucose study [40], but our result is instead consistent with previous reports that 5-fluorouracil and 5-fluoro-Urd are not substrates for this enzyme [10]. Thus taken together, our data lead to the conclusion that T. brucei encodes an unidentified 5' -nucleotidase that converts dCMP and dTMP to dCtd and dThd, respectively. We identified a number of potential candidate genes in T. brucei that could encode this activity, including a homolog of the E. coli HD protein YfbR and two strong candidates from the HAD-domain related family. Notably we showed that the T. brucei YfbR homolog encodes a metal dependent HD domain 5’-nucleotidase with broad substrate specificity functioning on all three pyrimidine deoxy-mononucleotides. Whether or not the T. brucei YfbR homolog is the only 5’-nucleotidase in T. brucei, or whether it is even the dominant enzyme with this capability remain open questions. Mammalian cells encode at least seven 5’-nucleotidases with overlapping specificities [41–43] and E. coli encodes minimally three, one each from the HD, HAD and SurE superfamilies [38]. Thus it is likely that other candidate T. brucei genes identified in our bioinformatics analysis will also display activity. In mammalian cells the 5’-nucleotidases have been shown to be required for regulation of cellular dNTP levels and to provide a mechanism to maintain balanced ratios between the pools, which is essential for high fidelity DNA synthesis [43]. Like the T. brucei HD-domain 5’-nucleotidase, all described nucleotidases from the various families exhibit broad substrate specificity. The broad specificity allows these enzymes to function in a ubiquitous capacity for interconversion of the nucleotide pools. The finding of 5’-nucleotidase activity in T. brucei leads directly to the essentiality of both TK and CDA, as in their absence the dead-end buildup of pyrimidine nucleosides leads to depletion of pyrimidine deoxynucleotides and to cell death. Within this context, the ability of DCTD to rescue the TK null cell line suggests that DCTD is able to effectively compete with the 5' -nucleotidase for the dCMP pools, converting sufficient amounts to dUMP where it can be efficiently shunted to dTMP even in the absence of TK. The existence of the metabolic cycle involving TK, CDA and 5’-nucleotidase provides the cell with a mechanism to interconvert between the deoxyuridine, deoxycytidine and deoxythymidine pools allowing presumably for better regulation and balance of their relative levels. While dUMP can also be formed from UDP, this pathway is apparently not sufficient to keep up with dUMP needs in the face of the dead-end accumulation of the TK substrates in the absence of TK. However this pathway remains an important additional source of dUMP as null mutants of dUTPase have been reported to be thymidine auxotrophs [19]. Our metabolomic analysis also uncovered some additional insights into T. brucei metabolism and regulation. In the presence of an outside source of pyrimidines (non-dialyzed serum), the pyrimidine nucleosides dUrd, dCyd, dThd and thymine accumulated in the TK c-null cells to higher levels than for cells grown in pyrimidine-free medium (dialyzed serum). These data confirm that in the absence of TK there is dead-end accumulation of these nucleosides but they also suggest that uridine phosphorylase is not a significant drainage point for these pools. Thus T. brucei uridine phosphorylase primarily catalyzes conversion of Urd to uracil, while it is not capable of synthesizing dUrd (as described above), or using it efficiently as a substrate. This hypothesis is consistent with previous reports that the recombinant T. brucei enzyme is 10-fold more active on Urd than dUrd [20]. Our metabolomic data also suggest that one response of T. brucei to TK depletion is increased nucleoside transport despite the fact that the upregulated transport was unable to relieve the growth block. A similar accumulation in dUrd in the presence of normal serum was previously reported for T. brucei BSF treated with thymidylate synthase inhibitors [10] suggesting this is a common response to starvation of thymine nucleotides. Finally, we noted that the levels of TCA intermediates were significantly perturbed in both the TK c-null and CDA null cells, including significant increases in α-ketoglutarate and homocysteine upon loss of TK expression or removal of thymidine from the CDA null cells. α-ketoglutarate is formed in the transamination reaction that generates L-Asp, which in turn is required for the first step in de novo pyrimidine biosynthesis, while homocysteine leads to formation of methionine then 5,10-methylene tetrahydrofolate, needed to convert dUMP to dTMP. Taken together with an observed decrease in carbamoyl phosphate, another precursor of the de novo pathway, the data suggest the cells may attempt to compensate for the loss of TK by increasing flux through the de novo pathway. Finally we also observed a significant decrease in acetyl-ornithine/acetyl-putrescine. It is not immediately apparent how these metabolites are synthesized, but their presence in T. brucei has been previously noted [40]. It is also not immediately clear what role they may play in pyrimidine biosynthesis, but both the synthesis and degradation of acetyl-ornithine can be catalyzed by aminotransferases, and in the case of its degradation this pathway links back to glutamate pools, and thus potentially to pyrimidine biosynthesis. The specific aminotransferases that catalyze these reactions are not annotated in the T. brucei genome, but aminotransferases have been reported to have broad and redundant substrate specificities in E. coli [44]. While a key aspect of our work was to elucidate the role of TK and CDA in linking the de novo pathway to synthesis of the deoxynucleotide pools, we have also validated TK as a drug target in T. brucei by showing that it is essential both in vitro and in vivo. The finding that the TK c-null cells cannot be rescued by exogenous pyrimidines shows it would not be possible for even an intracellular parasite to get around the block. Our work additionally showed that CDA is essential for in vitro growth of blood form T. brucei. While we did not determine if CDA is required for virulence in vivo, it remains a possibility provided that blood thymidine levels are below those required for rescue. Pyrimidine deoxynucleosides levels in human blood are reported to be ~10-fold below the EC50 that we measured for efficient rescue of CDA null growth. Furthermore, T. brucei has low affinity and/or poor efficiency transporters for deoxynucleosides [10], suggesting that CDA may be essential for infection in humans. However, additional studies will be needed to address this question conclusively. The presence of multiple pathways to synthesize dUMP appears to be an important shared characteristic amongst many eukaryotic cells, with the data suggesting that some organisms require either TK or DCTD to link de novo biosynthesis to the thymine deoxynucleotides. Interestingly, our bioinformatics analysis shows that other single-celled eukaryotic pathogens, including all three disease-causing trypanosomatids, encode TK but lack DCTD. These data suggest that TK may be essential in these other pathogens and may potentially provide a path forward to develop drugs that have pan-activity against a range of human pathogens. However the essentiality in other organisms would be dependent on the presence of the 5’-nucleotidase activity and likely also on limited catabolism of dUrd back to uracil by uridine phosphorylase. In support, a homolog of the T. brucei HD domain 5’nucleotidase is present in Leishmania and T. cruzi. Furthermore Leishmania major TK null cell line showed severely reduced growth rates [45]. In contrast, deletion of TK from Cryptosporidium parvum was not lethal, which is predicted by the presence of both TK and DCTD [46]. The finding that human cells contain both TK and DCTD, and that TK is not essential in human cells [47] supports the potential for selectively targeting TK from the eukaryotic pathogens that lack DCTD. Thus in conclusion, the unexpected finding that TK is essential in T. brucei and its mechanistic role in supporting de novo pyrimidine biosynthesis has uncovered a unique opportunity for the potential development of a pan-trypanosomatid therapy.
All animal care and experimental procedures were in accordance with the office of Laboratory Animal welfare (OLAW) guidelines provided by the National Institute of Health (NIH) USA (Assurance Number D16-00296) and the USDA (Registration Number 74-R-0072) as specified by the University of Texas Southwestern Medical Center Animal Care and Use Committee (IACUC) guidelines. Animal experiments were approved by the Ethical Review Committee at the University of Southwestern Medical Center and performed under the IACUC-2012-0021 protocol. All mice were housed in an animal facility with barrier SPF conditions. T. brucei gene sequences were obtained from TriTrypDB and the gene accession numbers are as follows: TK (Tb927. 10. 880), CDA (Tb927. 9. 3000), HD-domain 5’-nucleotidase (Tb09. 211. 2190), TERT (Tb927. 11. 10190). The HsTK1 (P04183) and HsDCTD (P32321) amino acid sequences were obtained from UniProt, and HsvTK was derived from Addgene plasmid #48356. Experiments were performed using T. brucei BSF SM cells genetically manipulated to express T7 RNA polymerase and the Tet repressor (TetR) [29]. Cells were grown in HMI-19 medium supplemented with 10% fetal bovine serum (FBS) at 37°C in 5% CO2. HMI-19 is a modified medium that we previously reported [9]. It was designed to contain more physiologically relevant purine and pyrimidine levels and it is supplemented with only 10 μM hypoxanthine and no added thymidine, except that present in FBS. To obtain completely pyrimidine-free conditions, normal FBS was replaced with dialyzed FBS in media where indicated. All cells were maintained in exponential growth (105−106 cells/mL). SM cells were maintained in G418 (2. 5 μg/mL) to retain the T7 polymerase and TetR. TK and CDA RNAi and knockout lines were cultured in the appropriate antibiotic depending on the transfected plasmid at the following concentrations unless otherwise stated: 2. 5 μg/mL G418 (Life Technologies), 2. 5 μg/mL blasticidin (InvivoGen), 2. 5 μg/mL phleomycin (InvivoGen), 1–2 μg/mL hygromycin (Sigma), 0. 1 μg/mL puromycin (Sigma), and 1 μg/mL Tet (RPI). For all nucleoside supplementation experiments 100 mM stocks of sterile filtered deoxyuridine (Sigma), thymidine (Sigma), uridine (Sigma), and uracil (Sigma) were added to cultures at concentrations indicated. All c-null lines were supplemented with 1 μg/mL Tet daily to maintain steady expression of Tet-regulated proteins. For pyrimidine and Tet free conditions, cells were washed (3 x 20 mL) with the appropriate media prior to beginning the growth experiments. For evaluation of growth rates, cells were washed and replated in media containing no antibiotics at a density of 20,000 cells/mL and diluted over the course of the study to maintain exponential growth. Cell density was determined using a hemocytometer (Bright-Line) with a lower limit of detection of 104cells/ml. Two technical replicates were averaged for each counted sample. Total cell numbers were calculated by multiplying cell density by the dilution factor and volume [48]. For each transfection, parasites (107) were suspended in Human T Cell Nucleofector Buffer (Lonzo) (100 μL) containing NotI linearized vector (5 μg) or purified PCR product (1 μg) as described [49]. All transfected DNA was confirmed by sequencing prior to transfection. Negative controls cells transfected with buffer only were prepared alongside samples to optimize selection conditions. Cells were transfected using protocol X-001 on the Amaxa Nucleofector (Lonza) and then transferred to media (25 mL) and allowed to recover 8 h prior to addition of selection antibiotics. Two dilutions (1: 20 and 1: 40) of culture, containing selection antibiotics, were plated in 24-well plates at 2 mL/well. Negative control plates were monitored throughout the experiment to ensure selection was achieved. After several days, wells containing a cell density of about 106 cells/ml were selected for generation of clonal lines by limiting dilutions. The T. brucei TK and human (Hs) DCTD expressing TK c-null cell lines were generated utilizing the fusion PCR method [49,50]. Cloning primers are shown in S2 Table. The first TK allele was replaced by the HYG resistance gene by PCR fusion of TK 5’ and 3’ UTRs to HYG. The HYG resistance gene was derived from the pLew90 vector [29] (a gift from George A. M. Cross). UTRs were amplified from genomic DNA isolated from SM cells. To generate the TK single allele knockout (SKO) line the purified PCR product was transfected into SM cells and hygromycin resistant cells were selected in medium containing G418 and hygromycin. The TK SKO line was then used to generate the remaining cell lines. A Tet-regulated vector containing either FLAG-tagged TbTK or FLAG-tagged HsDCTD was cloned as follows. The T. brucei TK or the human DCTD genes were PCR amplified from T. brucei SM genomic DNA or from human cDNA synthesized from RNA extracted from a human breast adenocarcinoma cell line (MDA-MB-231), respectively. For both constructs, the forward direction PCR primer contained a flanking 5’ HindIII restriction site and an N-terminal FLAG tag; the reverse direction primer contained a flanking 3’ BamHI restriction site. The restriction digested PCR products were ligated into the pLew100v5-phleo vector (a gift from George A. M. Cross). The pLew100v5-phleo vector was linearized by the NotI restriction enzyme to facilitate integration into the rRNA spacer region. Linearized vector (5 μg) was transfected into the TK SKO cell line and selected for resistance to phleomycin. The resulting clones were screened to identify those with the tightest level of Tet regulation of the ectopically expressed protein. Finally, the remaining TK allele was replaced by a PAC resistance gene synthesized by GenScript. The PAC TK UTR fusion product was generated as described above and transfected into TK SKO cells expressing either T. brucei TK or HsDCTD grown in Tet containing medium for 2 days prior to transfection. TK c-null cells were selected and maintained in G418, phleomycin, hygromycin, puromycin, and Tet (added daily). PCR primers flanking the 5’ and 3’UTRs were used to confirm that the TK gene had been replaced by the selectable markers. RNAit (http: //trypanofan. bioc. cam. ac. uk/software/RNAit. html) was used to identify a suitable 566 bp region located in the TK 3’UTR. The 3’UTR was targeted to allow compatibility with TbTK rescue plasmids (described below), that utilize instead the ALD 3’UTR. Genomic DNA isolated from SM cells was used as template for PCR amplification of the target region and TA cloned into the Gateway vector pCR8/GW/TOPO (Life Technologies). The Tet inducible stem loop was created by addition of Gateway LR Clonase to a reaction containing both the Gateway vector (100 ng) and pTrypRNAiGate vector (100 ng) [51]. TK SKO (hyg) cells were transfected with the vector and integration into the rRNA spacer region was selected using phleomycin. For studies of the effects of TK knockdown, Tet was added daily to induce formation of the hairpin leading to knockdown of TK mRNA. Cells were grown in the absence of other antibiotics for these studies. The TbTK rescue construct under control of the Tet promoter was generated using the same approach described above for the c-null cell line except that an N-terminal AU1-tag was included instead of a FLAG-tag to allow detection of the expressed protein. The gene encoding the HsTK open reading frame was synthesized by GenScript and cloned into the pUC57 vector, which was used for subsequent PCR amplification to generate the FLAG-tagged HsTK rescue construct. HsvTK was amplified from the pHJ17 Hyg-TK-loxP vector (Addgene). To generate the TbTK E286A and HsTK K32I mutants, both wild-type genes were subcloned into the pCR2. 1-TOPO TA vector (Invitrogen). Complimentary PCR primers containing the desired point mutation were synthesized. Phusion polymerase (NEB) was used to amplify the entire vector according the to the following parameters: initial denaturation at 95°C for 30 s followed by 18 cycles of denaturation at 95°C for 30 s, annealing at 68°C for 1 min, and amplification at 72°C for 5 min. Each reaction (50 μL) was treated with DpnI (NEB) (1 μL) overnight at 37°C followed by transformation into T10 cells and selection with ampicillin (100 μg/ml). Clones were sequenced using M13 primers. All constructs contained flanking 5’ HindIII and 3’ BamHI restriction sites that permitted ligation into the pLew100v5-bsd vector[49]. The vectors were linearized with NotI and transfected into TK RNAi cells as described above. SM cells were transfected with the HYG resistance gene flanked by the CDA 5’ and 3’ UTRs, generated by fusion PCR as described above, to generate the SKO in medium containing G418 and hygromycin. For the remaining allele, a fusion PCR product containing the PAC resistance gene was transfected into the SKO cells. Null cells were selected in growth medium containing G418, hygromycin, puromycin, and Thd (500 μM). PCR primers flanking the CDA 5’ and 3’ UTRs were used to confirm replacement of the CDA alleles. T. brucei TK expressing TK c-null cells were grown with or without Tet for 24 h and CDA null cells were grown with or without thymidine (0. 5 mM) for 12 h. Cells (108) were harvested by centrifugation (3500 RPM, 5 min) and then washed in cold PBS (50 mL). Washed pellet was resuspended in 1 mL pre-chilled (-80°C) 80% methanol and incubated on ice for 10 min. The cell extract was centrifuged (16,000 x g, 4°C, 20 min) to remove insoluble debris and 0. 9 mL of supernatant was dried using a vacuum centrifuge. Samples were stored at -80°C prior to analysis. For pyrimidine-free studies, a starter culture was washed, as described above, and grown in pyrimidine-free medium for 48 h prior to the start of the experiment. Targeted metabolite profiling by LC-MS/MS was performed as previously described allowing for detection of ~ 130 standard metabolites [52]. While this method allowed for quantitation of many key nucleosides and bases, the deoxynucleotides were not profiled as they are not part of the trained set of the facility. In order to attempt to identify these metabolites we isolated a larger cell number (5×108cells) and again used targeted LC-MS/MS for detection as described [53]. Levels of deoxynucleotides in wild-type control SM cells were barely detectable so null lines were not analyzed. Because we were unable to quantitate dTTP by LC-MS/MS approaches we employed a previously reported enzymatic assay that monitors Klenow DNA polymerase catalyzed incorporation of 3H-dATP into synthetic oligonucleotides in a reaction that is proportional to the amount of dNTP [54,55]. Through use of a standard curve the targeted dNTP concentration in the sample was determined. The oligo template that was used for the assay was as reported [54]. To prepare cell extracts, TK c-null cells were grown with or without Tet for 24 or 48 h in standard HMI-19 media supplemented with normal FBS and were harvested by centrifugation (3500 RPM, 5 min) and washed once with PBS. The pellets were resuspended in 60% methanol (250 μL) and incubated at -20°C overnight. Cell extracts were placed in a boiling water bath for 5 min, centrifuged (16,000xg, 20 min, 4°C), and then the soluble fraction was dried by vacuum centrifugation. Dried extracts were dissolved in 100 μL sample buffer (40 mM Tris-HCl pH 7. 4,10 mM MgCl2). Each reaction (100 μL) contained 40 mM Tris-HCl (pH 7. 4), 10 mM MgCl2,5 mM DTT, 0. 25 μM oligonucleotide template, 1. 5 μg RNase A, 0. 25 μM 3H-dATP (ARC- 17. 2 Ci/mmol), 0. 3 units Klenow Fragment (NEB), and cell extract (10 μL) or dTTP standard. Reactions were incubated at 37°C for 1 hr before spotting (85 μL of reaction) onto DE81 paper disks (23 mm-GE Healthcare), which were then air dried. Disks were washed (3 x 10 min) with 25 mL 5% Na2HPO4, rinsed once with water (25 mL) and absolute ethanol (15 mL). Dried disks were placed in scintillation liquid and radioactivity was measured by scintillation counting. A standard curve was generated with 0–4 pmol dTTP (New England BioLabs). T. brucei cells (2x107) were pelleted by centrifugation (3500 RPM, 5 min), washed in cold PBS once and then resuspended in 1 mL pre-chilled methanol. Samples were freeze-thawed in liquid nitrogen, placed on ice for 10 min, and centrifuged (16,000 x g for 10 min at 4°C) to remove insoluble debris. A portion of the supernatant (0. 8 mL) was vacuum dried and the residue resuspended in 40 mM sodium phosphate, pH 7. 4. Samples were syringe filtered (Millex GV 0. 22 μM–Millipore) to remove fine particles. High-Performance Anion Exchange Chromatography (HPAEC) analysis was performed as published [56] but with a modified elution gradient that was optimized for the separation of UDP-GalNAc, UDP-GlcNAc, UDP-Gal and UDP-Glc. Briefly, chromatography was performed on a Dionex ICS3000 HPAEC system with a CarboPac PA1 analytical column (4 mm Å~ 250 mm) and guard column (4 mm Å~ 50 mm). The method was performed with eluents 1 mM NaOH (E1) and 1 M NaOAc, 1 mM NaOH (E2) as follows: 0 min—20% E2,10 min—45% E2,25 min—45% E2,35 min—45% E2,40 min—100% E2,50 min—100% E2,55 min—20% E2,65 min—20% E2. Equal amounts of sample (approximately 50 Mio cells) were analyzed within each experiment and the signal divided by cell count. Synthetic standards were acquired from Promega. DNAzol (Molecular Research Center) was used to isolate genomic DNA from T. brucei cells. Typically, 5x107 cells were collected for DNA extraction using guidelines recommended by the manufacturer. Total RNA was extracted from samples (3x107cells) using TRIzol (Invitrogen), following the manufacturer’s protocol. As described above, total RNA was isolated from samples and treated with DNaseI (Invitrogen) to eliminate gDNA contamination. A cDNA reverse transcription kit (Applied Biosystems) was used to synthesize cDNA for downstream analysis. Relative mRNA abundance was quantified using iTaq SYBR Green Supermix with ROX (Bio-Rad) utilizing a standard curve for each set of primers per experiment. For all experiments, TERT was used as a reference gene [57]. Data was collected on the CFX96 (Bio-Rad) and analysis was performed using the Pfaffl method [58]. Mice (C57BL/6J) were purchased from Wakeland Laboratory (UT Southwestern) and were group-housed in filter-top cages. The animal facility has standard laboratory conditions: 21 to 22°C ambient temperature and a 12 h light/12 h dark cycle. Chow and water were available ad libitum. Both doxycycline (Dox) water and water only (controls), were supplemented with 0. 1% saccharin to ensure animals drank the Dox supplemented water. Mice from each group were introduced to the study drinking water 2 days prior to infection. Water bottles were protected from light and replaced every 2–3 days. Mice drank approximately 12. 5 mL of water daily. Mice (8 weeks old, n = 6) (12 in total, 3 per study arm) were infected intraperitoneally with 103 SM or TK c-null parasites ± Dox. Prior to the infection TK c-null parasites were propagated in +Tet conditions to ensure parasite viability at the start of the study. Mice were monitored for parasitemia starting three days post-infection by collecting 1 μL of blood from the tail in a 1: 150 dilution of medium and counted using a hemocytometer as described [59]. Mice were monitored for 30 days post infection. Cells (5x107) were harvested by centrifugation (3500 RPM, 5 min), washed twice in cold PBS (10 mM Na2HPO4,1. 8 mM KH2PO4,137 mM NaCl, 2. 7 mM KCl, pH 7. 4), and resuspended (50 uL) in trypanosome lysis buffer (50 mM Hepes, pH 8. 0,100 mM NaCl, 5 mM, β-mercaptoethanol, 2 mM PMSF, 1 mg/mL leupeptin, 2 mg/mL antipain, 10 mg/mL benzamidine, 1mg/mL pepstatin, 1 mg/mL chymostatin). Cells were lysed by 3 freeze/thaw cycles. Insoluble debris was pelleted by centrifugation (16,000 x g, 20 min, 4°C) and the soluble fraction was collected. The Bio-Rad Protein Assay reagent was used to determine protein concentration with bovine serum albumin (BSA) used to generate a standard curve. Total protein (20 μg) was resolved by 12% SDS-PAGE and transferred to a PVDF membrane using the Mini Trans-Blot Cell (BioRad). Membranes were blocked by 5% non-fat dry milk in Tris-buffered saline (TBS) (20 mM Tris (pH 7. 6), 150 mM NaCl) for 1 h then incubated with a primary antibody in 5% milk and TBS-T (TBS + 0. 1% Tween-20) overnight at 4°C. The following dilutions were used for each primary antibody: αFLAG 1: 1000 (Rabbit polyclonal-Thermo Fisher), αAU1 1: 1000 (Mouse monoclonal – Covance), αHsTK 1: 500 (Mouse monoclonal—Thermo Fisher), αTbBiP 1: 100,000 (Covance). For detection of the primary antibody, the membrane was incubated in a 1: 10,000 dilution of a Protein A-HRP conjugate (Abcam) in TBS-T (5% milk) for 1 hour at room temperature. The membrane was washed 5 x 5min with TBS-T and incubated in SuperSignal West Pico Chemiluminescent Substrate (Thermo Scientific) for 5 minutes. The membrane was visualized by the ImageQuant LAS 4000 (GE). Graphs were generated in GraphPad Prism version 7. 0a for Mac, GraphPad Software, San Diego California USA (www. graphpad. com), and statistical analysis was performed as indicated in the figure legends. The KEGG (Kyoto Encyclopedia of Genes and Genomes[60,61]) pyrimidine metabolic pathway highlights two enzymatic reactions (EC 3. 1. 3. 5 and EC 3. 1. 3. 89) that perform the 5' - (deoxy) nucleotidase activity required to produce pyrimidine deoxynucleosides in other organisms. We searched all reviewed UniProtKB [62] entries with these two described enzyme activities, identifying 561 genes with EC 3. 1. 3. 5 activity and 70 genes with EC 3. 1. 3. 89 activity. We sorted the identified genes according to their assigned PFAM domains [63], keeping nine representative sequences from each unique PFAM, which correspond to five different fold groups (Table 1). For stringent identification of T. brucei protein sequences with potential 5' nucleotidase activity, the representative sequences were used as queries to search the NCBI NR database using PSI-BLAST [64] (5 iterations, E-value cutoff 0. 001), storing the resulting position-specific scoring matrix as a checkpoint file for re-initiating BLAST against a database of protein sequences from the T. brucei genome (E-value cutoff 1). Identified T. brucei protein sequences were assigned PFAM domains using batch CD search [65], keeping those sequences with PFAM domains described as possessing 5’-nucleotidase activity. To identify all potential T. brucei protein sequences with domains related to those described as having 5' -nucleotidase activity, we queried the T. brucei genome using RPS-BLAST (E-value cutoff 0. 006) with a library of sequence profiles downloaded from the conserved domain database (CDD ID in S1 Table) corresponding to each described 5' -nucleotidase PFAM in Table 1. Identified sequences were crosschecked for the presence of the query domain using batch CD search [65] or HHPRED [66,67], reporting the positive hits using the initial RPS-BLAST E-values in S1 Table. Additional methods details are provided in S1 Text. The DNA sequence for the T. brucei 5’-nucleotidase (HD-fold, YfbR-like) was obtained from TriTryDB (Tb09. 211. 2190). The E. coli codon optimized gene was synthesized by GenScript. PCR was used to generate flanking BsaI and XbaI restriction sites that allowed for cloning into the pE-SUMO (KAN) vector (LifeSensors, Malvern, PA) and expression as a N-terminal His6-SUMO fusion protein. The nucleotidase pE-SUMO vector was transformed into BL21 cells. Cells were cultured in 2L LB-KAN (50 μg/mL) media at 37°C until OD600 0. 7, then cooled to 16°C and induced by 500 μM IPTG (Isopropyl β-D-1-thiogalactopyranoside) for 16 h. Cells were collected by centrifugation and suspended in buffer A (500 mM NaCl, 50 mM HEPES pH 7. 5,5 mM imidazole, 5% glycerol, 5 mM 2-mercaptoethanol) supplemented with 2 mM PMSF (phenylmethane sulfonyl fluoride) and a protease inhibitor cocktail (1 mg/mL leupeptin, 2 mg/mL antipain, 10 mg/mL benzamidine, 1 mg/mL pepstatin, 1 mg/mL chymostatin). Cells were lysed by cell disruptor and the cell debris removed by centrifugation. Supernatant was applied to a HisTrap HP column (GE Healthcare) and washed with buffer A. The His6-SUMO nucleotidase fusion was eluted using a gradient of 5–45% buffer B (500 mM NaCl, 50 mM HEPES pH 7. 5,5% glycerol, 5 mM 2-mercaptoethanol, 500 mM imidazole). Fractions were analyzed by SDS-PAGE and those containing the T. brucei HD domain 5’-nucelotidase were pooled, concentrated (10 kDa MWCO Millipore) and dialyzed against buffer A. The His6-Sumo tag was removed by overnight incubation at 4°C with His6-tagged Ubiquitin-like-specific protease 1 (ULP1) as described [68]. After incubation the mixture was applied to a second HisTrap HP column and cleaved 5’-nucelotidase was collected in the flow-through, while His6-tagged ULP-1 and impurities were retained on the resin. The flow through was concentrated and protein purity was assessed to be >98% by SDS-PAGE analysis (S14 Fig). This tagless protein was used to collect the data shown in Fig 8. To provide an alternative purification method, His6-Sumo tagged 5’-nucleotidase eluted from the first Ni+2 column was further purified by Gel Filtration chromatography on a SuperDex 200 Prep Grade column (GE Healthcare) using buffer A. The His6-Sumo tagged 5’-nucleotidase showed similar activity (within 2-fold) of the untagged version. Protein concentrations were determined by Bradford Assay (Bio-Rad). T. brucei 5’-nucleotidase activity was assessed using an endpoint assay for released inorganic phosphate (Pi) using Malachite Green as the detection reagent as described [38,69]. Briefly, each reaction (160 μL) contained 50 mM HEPES pH 7. 5,0. 5 mM CoCl2,1 mM substrate, and enzyme. The reactions were incubated for 10–30 min (to confirm linearity) at 37°C and then 50 uL was treated with 100 mM EDTA. Malachite green reagent (150 μL) was added to each quenched reaction and incubated at room temperature for 5 min prior to measurement at 650 nm. A range of substrate (0. 1–1. 0 mM), metal (0. 5–10 mM) and enzyme (25–100 nM) concentrations were tested to confirm linear dependence on enzyme concentration and to confirm that the reaction rate versus substrate curve was a saturable process. The production of Pi was measured at 650 nm. Absorbance was converted to μmoles of Pi using a standard curve generated using phosphate standard (Cayman Chemical) ranging from 0–100 μM. For the substrate and metal ion specificity studies shown in Fig 8, assays were run with 100 nM enzyme for 20 min using a substrate concentration of 1 mM. Metal ion concentrations are indicated on the graph. All data were collected in triplicate.
Human pathogenic trypanosomatids are responsible for several life threatening diseases, together infecting 20 million people, while treatment is complicated by poor drug therapies. The unique biology of these organisms has led to the need for different drug therapies to be developed for each. Identification of enzymatic targets that could be used to develop a single drug capable of treating multiple parasites would be revolutionary. Herein we show that the trypanosomatid biosynthetic pathway used to synthesize key precursors for DNA biosynthesis is unexpectedly vulnerable. We find that seemingly redundant enzymes thymidine kinase and cytidine deaminase are required, not for their typical role in salvaging exogenous precursors, but are instead essential for de novo synthesis of thymine nucleotides. Trypanosomatids lack alternative routes to synthesize these nucleotides, which are found in other eukaryotic cells, while encoding a previously unknown activity that degrades them. For these reasons thymidine kinase is essential to support infection in mice and has strong potential as a new drug target. While our work focused on the causative agent of African sleeping sickness, the impact of our findings may extend to other pathogenic trypanosomatids and potentially to additional single-celled eukaryotic human pathogens.
blood cells
cell physiology
glycosylamines
medicine and health sciences
rna interference
immune cells
chemical compounds
immunology
cell metabolism
organic compounds
parasitic protozoans
protozoans
pyrimidines
epigenetics
molecular biology techniques
research and analysis methods
thymidines
white blood cells
artificial gene amplification and extension
genetic interference
animal cells
gene expression
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biochemistry
rna
trypanosoma
organic chemistry
nucleic acids
cell biology
k cells
polymerase chain reaction
nucleosides
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trypanosoma brucei gambiense
glycobiology
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Pyrimidine Salvage Enzymes Are Essential for De Novo Biosynthesis of Deoxypyrimidine Nucleotides in Trypanosoma brucei