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The malaria parasite Plasmodium falciparum apicoplast indirect aminoacylation pathway utilizes a non-discriminating glutamyl-tRNA synthetase to synthesize Glu-tRNAGln and a glutaminyl-tRNA amidotransferase to convert Glu-tRNAGln to Gln-tRNAGln. Here, we show that Plasmodium falciparum and other apicomplexans possess a unique heterodimeric glutamyl-tRNA amidotransferase consisting of GatA and GatB subunits (GatAB). We localized the P. falciparum GatA and GatB subunits to the apicoplast in blood stage parasites and demonstrated that recombinant GatAB converts Glu-tRNAGln to Gln-tRNAGln in vitro. We demonstrate that the apicoplast GatAB-catalyzed reaction is essential to the parasite blood stages because we could not delete the Plasmodium berghei gene encoding GatA in blood stage parasites in vivo. A phylogenetic analysis placed the split between Plasmodium GatB, archaeal GatE, and bacterial GatB prior to the phylogenetic divide between bacteria and archaea. Moreover, Plasmodium GatA also appears to have emerged prior to the bacterial-archaeal phylogenetic divide. Thus, although GatAB is found in Plasmodium, it emerged prior to the phylogenetic separation of archaea and bacteria.
The human malaria parasite Plasmodium falciparum is responsible for 124–283 million cases of malaria and an estimated 0.6 million deaths every year (1). It contains a relict plastid, the remnant of an ancient secondary endosymbiotic event in which the eukaryotic progenitor of the malaria parasite engulfed a photosynthetic eukaryote known as the apicoplast (2). The Plasmodium apicoplast possesses a 35-kb circular genome with 60 genes (3) that encode components of the apicoplast transcriptional and translational apparatus such as RNA polymerase subunits, the elongation factor EF-Tu, several ribosomal proteins, rRNAs, and tRNAs (4,–8), as well as the SufB protein thought to play a role in FeS cluster formation (9). Most apicoplast proteins, however, are encoded by the nuclear genome and are imported into the organelle post-translationally (10). Over 500 apicoplast-targeted proteins were identified in P. falciparum (11, 12), revealing apicoplast biosynthetic pathways for fatty acids (13, 14), isoprenoid precursors (15), and heme (16), as well as enzymes for tRNA modification (12) and lipoylation (17). Several of these pathways exhibit prokaryote-like features and are potential drug targets (12, 15, 18). Recent studies have shown that apicoplast isoprenoid precursor biosynthesis is essential in P. falciparum asexual stages (19), indicating that the pathway cannot be bypassed by salvaging lipids from the host and may be a good drug target in asexual stages. The type II fatty acid and heme biosynthetic pathways, however, are not essential in the asexual stages (18), and although not good targets for asexual stage chemotherapy, they may prove to be valuable drug targets in liver stages (2).
Translational accuracy is required to properly decipher the genetic code during protein synthesis. The fidelity of protein synthesis largely depends on the formation of correct aminoacyl-tRNAs by aminoacyl-tRNA synthetases (aaRSs).2 In the classic model, each species of aaRS strictly discriminates one amino acid from among the 20 canonical amino acids, as well as its cognate tRNA isoacceptor from the non-cognate tRNAs. However, genomic and biochemical analyses have revealed that the full complement of 20 aaRSs is used only in the eukaryotic cytoplasm and a minority of bacteria, whereas a majority of bacterial and archaeal genomes lack genes encoding glutaminyl-tRNA synthetase (GlnRS) and or asparaginyl-tRNA synthetase (AsnRS) (20). In these organisms, Gln-tRNAGln and/or Asn-tRNAAsn are synthesized via an indirect pathway (21). In most bacteria and archaea lacking GlnRS, tRNAGln is first misaminoacylated with Glu in a reaction catalyzed by a non-discriminating GluRS (ND-GluRS) that can glutamylate both tRNAGlu and tRNAGln (see Reactions 1 and 2).
The glutamyl residue of Glu-tRNAGln is then transamidated by a glutamyl-tRNAGln amidotransferase (Glu-AdT) in the presence of ATP using Gln as an amide donor, producing Gln-tRNAGln. Similarly, in the case of Asn-tRNAAsn formation in organisms lacking AsnRS, Asn-tRNAAsn is synthesized by a non-discriminating aspartyl-tRNA synthetase and an aspartyl-tRNAAsn amidotransferase (Asp-AdT). Two types of tRNA-dependent amidotransferases are known as follows: the heterotrimeric GatCAB (22) and the heterodimeric GatDE (20). Bacterial GatCAB functions as a Glu-AdT or an Asp-AdT in a species-specific manner. In some bacteria lacking both AsnRS and GlnRS, GatCAB acts as both a Glu-AdT and an Asp-AdT (21). As each subunit of GatCAB is encoded only in archaeal genomes lacking a gene for AsnRS, archaeal GatCAB seems to function as an Asp-AdT. GatDE is only found in archaea and functions as a Glu-AdT (21).
Aminoacyl-tRNA formation is essential for protein synthesis. Despite the central importance of this process in all living organisms, it remains unknown how Plasmodium synthesizes Gln-tRNAGln in the apicoplast. The Plasmodium apicoplast genome does not encode any tRNA synthetases, and the nuclear genome does not contain an apicoplast-targeted GlnRS. We recently reported that a nucleus-encoded non-discriminating GluRS that is imported into the apicoplast is responsible for the formation of misacylated Glu-tRNAGln and is essential in the erythrocytic stages (23). In this study, we aimed to further clarify the formation of Gln-tRNAGln in the Plasmodium apicoplast by a unique Plasmodium signature protein GatAB.
A list of putative apicoplast-targeted proteins conserved in P. falciparum, Theileria parva, and Toxoplasma gondii was obtained from the supplementary information in Gardner et al. (24). Nucleotide or amino acid sequences of Plasmodium genes or proteins (11, 25) and those from other species were obtained from PlasmoDB (26), the Wellcome Trust Sanger Institute GeneDB website, or UniProt (27).
The structure of Plasmodium GatA and GatB was modeled according to the crystal structures of Staphylococcus aureus GatCAB (2g5h and 3ip4) (28, 29) using the Swiss-Model automated comparative protein-modeling server (30). In the modeled range, the sequence identity of PfGatA and 2g5h was 32.2%, and the sequence identity of PfGatB and 3ip4 was 24.8%. No suitable structural template could be found for amino acid regions 1–180, 425–539 and 708–744 in PfGatA and amino acid regions 1–350 and 461–530 in PfGatB. The spatial arrangement of PfGatA and PfGatB was determined based on the crystal structures of S. aureus GatCAB. Attempts to model GatA and GatB according to the crystal structure of Pyrococcus abyssi archaeal GatDE (1zq1) (31) resulted in unusable models. All figures were prepared with Chimera (32).
The phylogenetic trees were constructed using the tools provided on line (33). The “one-click” mode was used, employing ClustalW (34) for sequence alignment, and Gblocks (35, 36), PhyML (36), and TreeDyn (37) for curation of the multiple sequence alignment, tree construction, and rendering, respectively. The final tree was constructed using 100 bootstraps.
P. falciparum clone 3D7 parasites (MRA-102, MR4, BEI Resources, Manassas, VA) were grown in human O+ red blood cells at 4% hematocrit in RPMI 1640 medium supplemented with Albumax (Life Technologies, Inc.) to a final concentration of 0.5% and gassed with 5% CO2 and 0.5% O2 in N2 at 37 °C as described previously (38). To generate an episomal transfection vector, the bipartite apicoplast targeting sequences of PfGatA (amino acids 1–102) and PfGatB (amino acids 1–98) were amplified from P. falciparum 3D7 genomic DNA and cloned into the BglII and AvrII sites of a pCHD-GFP vector (39). Synchronized P. falciparum 3D7 ring-stage parasites were transfected with 100 μg of purified plasmid DNA (plasmid maxi kit, Qiagen) via electroporation (40, 41), and the transfected parasites were selected using 2.5 nm WR99210. The transfection experiments were performed in duplicate and repeated once.
Subcellular localization of cloned P. falciparum transfectants expressing GFP-tagged PfGatA, PfGatB, or Myc-tagged PbGatA was performed as outlined in Ref. 42. Double staining was performed using a rabbit polyclonal anti-acyl carrier protein (ACP) primary antibody (diluted 1:500) (10) as an apicoplast marker and a mouse monoclonal anti-Myc antibody (Santa Cruz Biotechnology, diluted 1:500) to detect PbGluRS-Myc. Fluorescent staining was achieved using Alexa Fluor-conjugated secondary antibodies (Invitrogen) specific to rabbit (Alexa Fluor 594, red) or mouse (Alexa Fluor 488, green) IgGs. DAPI was used to stain nucleic acids, and the mitochondrion was stained by incubating the parasites in culture media for 30 min with 20 nm MitoTracker Red (Invitrogen) and fixing the cells as outlined above. Images were acquired using an Olympus Delta Vision imaging system (Applied Precision) with a ×100 objective and deconvolved using the SoftWoRx package (Applied Precision) with the default parameters.
A 4× myc tag was appended to the 3′ end of the gene encoding the putative apicoplast-targeted P. berghei GatA (PlasmoDB ID PBANKA_071810) as described previously (18). A 3.5-kb fragment of the 3′ end of the gene without the stop codon was amplified from P. berghei ANKA genomic DNA using primers PbGatA_F2 (TACCGCGGATAATATACAACCAATAACATTATAG) and PbGatA_R (ATACTAGTACTAGCCTTATTTTCCAAATTGTGAAC); the SacII and SpeI restriction sites are underlined. Polymerase chain reactions (PCR) (50 μl) contained 50 ng of genomic DNA, 0.1 μm of each primer, 5 μl of buffer, and 1 μl of Advantage 2 polymerase (Clontech). The PCR product was digested with SacII and SpeI and cloned into the b3D myc vector (18). P. berghei ANKA parasites were transfected, and parasites with myc insertions in the gatA gene were selected and cloned as described (18). Plasmid integration at the 5′- and 3′-insertion sites was confirmed via PCR using primer pairs PbGatA_For2 (TACCGCGGATAATATACAACCAATAACATTATAG) and PbGatA_int5_Rev2 (GAGACAGCTCAATTCTTTATGTCC) for the 5′ integration test and PbGatA_int3_For2 (CCTCTTCGCTATTACGCCAGCT) and PbGatA_Rev4 (GAACCACCAGATGACCCACCACATG) for the 3′ integration test. The strategy described previously (42) was used to attempt deletion of the PbGatA gene via double-crossover recombination. The primers used to amplify the genomic regions were as follows: PbGatA_KO.Pr1For (GCCCGCGGGCATGAGTTGTTAAAAGTTGCC) and PbGatA_KO.Pr2Rev (AGTTCTACTGGGCCCAAATTTAAGCATACAGAAAGTGAC); PbGatA_KO.Pr3For (CTTAAATTTGGGCCCAGTAGAACTAGAACATGAGGG) and PbGluRS_KO.Pr4Rev (GCCCGCGGTTTGTCCTTACAACTTTCTTACC). Primers 1 and 2 were designed to amplify a 942-bp fragment containing the last 102 bp of the PbGatA coding sequence and 840 bp of the 3′-UTR. Primers 3 and 4 were designed to amplify a 734-nucleotide fragment containing the first 62 bp of the PbGluRS coding sequence and 672 bp of the 5′-UTR. Primer 1 contained an added 5′-terminal GC dinucleotide and a SacII site. Primer 4 contained an added 5′-terminal GC dinucleotide and a SacII site. Primers 2 and 3 contained an ApaI site flanked by complementary sequences (underlined) for recombinatorial PCR. The two genomic fragments were first amplified in separate reactions using the cycling parameters described above, and the resulting products were combined in a second PCR to form a single product, which was digested with SacII cloned into the B3D KO Red vector. This construct was linearized with ApaI, and P. berghei ANKA parasites were transfected as described (43). The transfection experiments were performed in duplicate and repeated once.
PfGluRS, PfGatA, and PfGatB were expressed in Escherichia coli KRX cells (Promega) containing the pRARE2 plasmid (EMD4BioSciences) and the pET-29a vector encoding either the predicted mature PfGluRS coding sequence (Pf3D7_1357200, amino acids 78–574) (23), PfGatA (Pf3D7_0416100, amino acids 101–827), and PfGatB (Pf3D7_0628800, amino acids 96–884). PfGluRS was codon-optimized for wheat (23) and PfGatA and PfGatB for E. coli (GeneArt, Inc.). All constructs included N-terminal His6 tags.
Synthetic genes encoding P. falciparum apicoplast tRNAGlu and tRNAGln (both from GenBankTM accession number X95276) were expressed and purified from E. coli and 32P-labeled on their 3′-OH termini using the E. coli CCA-adding enzyme as described (23). The aminoacylation assay using recombinant PfGluRS was performed and quantified as described (23). Glutamylated tRNAs were phenol (Tris-buffered, pH 7.9)/chloroform-extracted, and unincorporated [α-32P]ATP was removed using Bio-Spin 30 columns (Bio-Rad).
Transamidation assays were carried out in 1× AdT buffer (100 mm Hepes·KOH, pH 7.2, 30 mm KCl, 12 mm MgCl2, and 5 mm DTT) with 2.6 mm l-glutamine, 4 mm ATP, 500 nm 32P-labeled Glu-tRNAGln, and 50 nm each of GatA and GatB. Reactions were carried out at 37 °C for 5 min. Aliquots (4 μl) were quenched on ice with 4 μl of 100 mm sodium citrate, pH 4.74, and 0.66 mg/ml of nuclease P1 (Sigma) and incubated at room temperature for 35 min. To separate glutaminyl-AMP (Gln-AMP) from Glu-AMP and AMP, 2.0 μl of the digested samples were separated on 20 × 20-cm PEI cellulose TLC plates. The plates were then developed in 100 mm ammonium acetate, 5% acetic acid and air-dried. Spot positions and intensities were measured by phosphorimaging, as described (44). To test whether PfGluRS and PfGluAdT together could sequentially form Gln-tRNAGln from precursors in a single reaction, we initiated some reactions by adding premixed PfGluRS, PfGatA, and GatB (50 nm each) in the presence of 2.6 mm l-Glu and l-Gln at 37 °C for 5 min and quantified as described above. To test whether PfGatAB could use an alternative amide donor, 2.6 mm l-asparagine was used instead of l-glutamine.
The formation of aminoacyl-tRNAs is a crucial step in protein synthesis. Despite the central importance of this process in all living organisms, it has been unclear how Plasmodium synthesizes Gln-tRNAGln in the apicoplast. We recently reported that Plasmodium apicoplast glutamyl-tRNA synthetase is a non-discriminating enzyme that forms both Glu-tRNAGlu and Glu-tRNAGln and is essential in erythrocytic stages of the parasite life cycle (23). Amidation of Glu-tRNAGln to Gln-tRNAGln requires a tRNA-dependent amidotransferase (AdT). The P. falciparum genome contains two single-exon genes that encode putative orthologs of the GatA and GatB subunits of the bacterial glutamyl-tRNA amidotransferase (Glu-AdT) as follows: GatA (PF3D7_0416100, 96 kDa, 826 amino acids) and GatB (PF3D7_0628800, 102 kDa, 882 amino acids) (Table 1). Both P. falciparum proteins possess predicted N-terminal bipartite apicoplast targeting sequences, suggesting that they are the subunits of an amidotransferase that participates in an apicoplast indirect aminoacylation pathway. Orthologs of PfGatA and PfGatB are conserved in apicomplexans that possess an apicoplast (e.g. Theileria (24, 45)) but not in apicomplexans that lack an apicoplast (e.g. Cryptosporidium). Although bacterial Glu-AdTs have the subunit composition GatCAB (28), we did not find a Plasmodium gene that encoded a GatC homolog nor did we find a GatF ortholog of the GatFAB yeast mitochondrial Glu-AdT (46, 47). In archaea, GatDE is used for Gln-tRNAGln formation. It is notable that the cradle domain in archaeal GatE and bacterial GatB share a similar fold, but the overall architectures of archaeal GatD and bacterial GatA are completely different.
Bacterial GatCAB includes three subunits as follows: the glutaminase GatA; the transamidase GatB; and GatC, which is a small protein (12 kDa) that appears to perform the role of a structural stabilizer at the interface between the GatA and GatB subunits (28, 29). GatB contains a transamidase domain called the “cradle” and a helical domain that binds to tRNA.
Plasmodium GatB (Fig. 1) includes an N-terminal apicoplast targeting sequence (residues 1–183), two unique inserts (residues 184–362 and 461–524) of unknown functions, and two domains typical for bacterial GatB, the cradle domain (residues 350–460 and 526–700), and a helical domain (residues 704–880). Instead of GatB, archaea use GatE, which contains an AspRS-like insertion in its cradle domain (31). No similarity could be found between the Plasmodium and archaeal insertions. Furthermore, we could not find a structural template for the Plasmodium insertions, which is the reason why they were not modeled. Most of the amino acids that are important for substrate recognition, Mg2+/Mn2+ coordination, and the ammonia channel are either conserved in PfGatB, bacterial GatB (28, 29), and archaeal GatE (31) proteins or replaced by conservative substitutions (Fig. 1).
In S. aureus GatB (28, 29), the conserved residues His12, Glu124, and Glu150 and three water molecules coordinate the Mg2+ ion; the equivalent residues in PfGatB are Glu451 and Glu539. The corresponding residue to His12 of S. aureus is probably situated in the histidine-rich unmodeled N-terminal region of PfGatB. Most likely residues Asp579 and Glu595 are part of the second transient binding site of Mg2+/Mn2+ ions; the corresponding residues in S. aureus GatB are Glu10, Asp192, and Glu210 (28, 29). Furthermore all residues forming the ADP-binding site are conserved or functionally replaced in Plasmodium. In P. falciparum, the phosphate moiety could interact with the conserved residues Asn581, Ser583, and Arg593, which is also an arginine in archaeal GatE (Arg239, P. abyssi) (31) but a lysine in bacterial GatB (Lys208, S. aureus) (28, 29). The hydrophobic adenosine binding pocket is less conserved between the species; in S. aureus it is formed by Val152, Pro155, and Phe205 (28, 29) and in P. abyssi by Ser191, Pro194, and Gly236 (31). The corresponding residues in Plasmodium are Val541 and two charged residues Lys544 and Lys590. However, the lysine residues interact with their hydrophobic side chains as well as with the adenosine, so that the plasmodial adenosine binding pocket is also hydrophobic. Inspection of the different GatB structures revealed that the C-terminal region in PfGatB (amino acids 805–880, Fig. 1) is highly mobile and could be involved in tRNA recognition, as seen in bacterial GatBs (28, 29).
Bacterial GatA consists of a single domain (28, 29), which is homologous to other amidases in its catalytic core (22, 28, 29, 48), whereas archaeal GatD consists of three domains as follows: an N-terminal domain and two AnsA-like domains (31). The predicted PfGatA (Fig. 1) contains an apicoplast targeting sequence (amino acids 1–110) and two inserts of unknown function, which are 115 and 37 amino acids long and found only in Plasmodium (residues 425–539 and 708–744, respectively). The active site of bacterial GatA (28, 29) includes a conserved Arg and a conserved Asp residue, which interact with the carboxyl and amide groups of bound glutamine. The corresponding residues in PfGatA are Arg647 and Glu755. Furthermore the Ser-cis-Ser-Lys catalytic scissors (Ser322, Ser346,and Lys251) that are involved in glutaminase- and glutamine-dependent transamidase activity in bacterial GatA (28, 29, 49) and the oxyanion hole (28, 29) that stabilizes the tetrahedral covalent intermediate (Thr343, Gly344, Gly345, and Ser346) are conserved in PfGatA (Fig. 1). In contrast to PfGatB, which has only 21% sequence identity to a potential human analog, PfGatA exhibited 40% identity to the human homolog, primarily due to the conservation of residues in the vicinity of the glutaminase domain (Fig. 1).
Nakamura et al. (28) propose that a hydrophilic tunnel channels NH3 from the glutaminase site in GatA to the transamidase site of GatB. Almost all residues that line this tunnel in S. aureus GatCAB (21 residues) (28) are strictly conserved in Plasmodium (17 residues), including a conserved Thr (343PfGatA and 175S. aureus, GatA) at the entrance and a conserved Lys (410PfGatB and 79S. aureus, GatB) at the tunnel exit.
GatC, a small protein, is the third subunit in bacterial Glu-AdTs (28, 29) and performs a structural stabilizer role at the interface between the GatA and GatB subunits. A homolog of the gatC gene could not be identified in the P. falciparum genome, but components of the insertions found in GatA (residues 425–539 and 708–744) and in GatB (residues 1–350 and 461–525) could perform the task of bacterial GatC. This hypothesis is supported by the presence of two conserved residues (Asn52, S. aureus and Arg268, S. aureus), which form hydrogen bonds between the bacterial GatB and GatC subunits are also present in PfGatB (Asn383, PfGatB and Asn669, PfGatB) (Fig. 1).
To determine the evolutionary origin of plasmodial GatB and GatA proteins, we constructed phylogenetic trees with bacterial GatB and archaeal GatE proteins and with bacterial GatA and archaeal GatD proteins, respectively. If P. falciparum GatB or GatA evolved from either the bacterial GatB/GatA or archaeal GatE/GatD lineages, one would expect P. falciparum GatB/GatA to appear in either the GatB/GatA or GatE/GatD clades, much like GlnRS enzymes grouping with the eukaryotic glutamyl-tRNA synthetases (GluRS) (50, 51). Instead, our phylogenetic trees had three branches representing distinct groupings, one for the P. falciparum GatB or GatA subunits, a second containing the bacterial GatB/GatA subunits, and a third for the archaeal GatE/GatD subunits. Suspecting an artifact because of long-branch attraction (52), we reanalyzed our dataset by including GatA and GatB sequences from other Plasmodium species, the cyanobacterium Anabaena variabilis, the vascular plant Arabidopsis thaliana, and the chromerids Chromera velia and Vitrella brassicaformis. This phylogenetic tree also had three branches representing distinct groupings, one for the apicomplexan/Plasmodium GatB or GatA, a second of the bacterial and green plastid GatB/GatA, and a third of the archaeal GatE/GatD subunits (Fig. 2, A and B) with the newly added sequences of chromerids (C. velia and V. brassicaformis) and plants (A. variabilis and A. thaliana) nesting deeply among the bacterial sequences, away from the Plasmodium branch. The Plasmodium branch of the tree was also consistent with the following previously established phylogenetic relationships within Plasmodium spp.: the monophyly of rodent parasites (53); the close affinity of Plasmodium vivax with Plasmodium cynomolgi and Plasmodium knowlesi, each with bootstrap values > = 0.79; and the separate clustering of P. falciparum from the clades containing the rodent plasmodial parasites or the clade that contains P. vivax (53, 54). Furthermore, the GatA and GatB sequences from two other Apicomplexa, the piroplasms Theileria (24, 45) and Babesia (55), which like Plasmodium lack gatC or gatF in their nuclear genomes, were placed in the same clade as the Plasmodium spp. (Fig. 2, A and B), indicating that the placement of Plasmodium GatA and GatB proteins on a separate branch was unlikely to be an artifact.
These results indicate that Plasmodium GatA and GatB belong to subfamilies that are distinctly different from those of known GatA, GatB, GatD, or GatE subunits. These findings imply that Plasmodium GatA and GatB co-evolved and that Plasmodium GatAB is a paralog of GatCAB, GatFAB, and GatDE.
The P. falciparum GatA (PF3D7_0416100) and GatB (PF3D7_0628800) contain a predicted apicoplast bipartite targeting sequence (11, 25, 56), but their subcellular localization has never been established experimentally. To determine whether the enzymes are targeted to the apicoplast, P. falciparum parasites were transfected with episomal constructs in which the predicted apicoplast targeting sequences of either PfGatA and PfGatB were fused to the N terminus of green fluorescent protein (GFP) and expressed in transfected parasites under the control of the P. falciparum calmodulin promoter (39). Fixed blood stage parasites expressing PfGatA-GFP or PfGatB-GFP were stained with anti-GFP and anti-ACP antibodies (14) to detect the fusion proteins and mark the apicoplast, respectively. Deconvolution fluorescence microscopic examination revealed distinct subcellular GFP localization for all constructs. As with other cell lines expressing GFP in the apicoplast (39), PfGatA-GFP and PfGatB-GFP were immunolocalized within a characteristically small and round compartment in ring stage parasites (Fig. 3, A, panel i, and B, panel i), which then elongated and developed into a complex branched form at the trophozoite stage (Fig. 3, A, panel ii, and B, panel ii) prior to splitting into numerous individual structures in schizonts, one for each daughter merozoite. Furthermore, the anti-GFP and anti-ACP signals were colocalized, confirming that the GatA- and GatB-GFP fusion proteins were present in the apicoplast.
To investigate potential dual localization to the mitochondrion, the same transgenic PfGatA- and PfGatB-GFP P. falciparum clones were incubated with MitoTracker Red and then fixed, stained with anti-GFP monoclonal antibody, and examined via deconvolution fluorescence microscopy. In ring stages transfected with PfGatA-GFP, the anti-GFP antibody and MitoTracker Red marked closely apposed organelles that were clearly distinct (Fig. 3C, panel i) and which began to enlarge in early trophozoites (Fig. 3C, panel ii). Little co-localization between the mitochondrion and PfGatB-GFP was observed in late trophozoites (Fig. 3D, panel i) or late schizont stages with the mitochondrial staining largely distinct from that of PfGatB-GFP (Fig. 3D, panel ii).
To confirm apicoplast localization that was determined via transfection of the episomal constructs in P. falciparum, we tagged the endogenous gene encoding the P. berghei ortholog of PfGatA (PBANKA_071810) with a quadruple Myc tag (Fig. 4). We tagged the endogenous PbGatA coding sequence to reduce the possibility that the fusion protein would be mistargeted due to inappropriate timing or intensity of expression. Fixed blood stage parasites were stained with anti-Myc antibody to detect PbGatA and ACP antisera (14) to detect the apicoplast, and the samples were observed using Deltavision deconvolution fluorescence microscopy. Structures containing the Myc-tagged PbGatA (PbGatA-myc) exhibited a typical apicoplast appearance (Fig. 4C) similar to that observed using the episosomal PfGatA-GFP and PfGatB-GFP constructs in P. falciparum in vitro.
Bioinformatic analyses strongly suggest that Plasmodium lacks an apicoplast-targeted GlnRS (57, 58), implying that indirect aminoacylation is probably the sole route for Gln-tRNAGln formation in the apicoplast and that GatA and GatB are essential components of the protein biosynthetic pathway in Plasmodium. To test whether PbGatA was required for blood stage growth, we transfected P. berghei parasites with a construct (Fig. 5) designed to delete the endogenous PbGatA gene via double-crossover recombination. As a control, we transfected parasites from the same batch with the construct used previously to generate the PbGatA-myc transgenic parasites (Fig. 4). In three independent experiments, the PbGatA genomic locus was refractory to gene deletion via double-crossover recombination. Transgenic PbGatA-myc parasites, however, were readily obtained (data not shown). These two experiments indicate that although the PbGatA locus is accessible to recombination, the PbGatA gene is refractory to deletion, strongly suggesting that the apicoplast-targeted GatA is essential in blood stage parasites.
To show biochemically that PfGatAB encodes the apicoplast glutamyl-tRNA amidotransferase, we independently expressed PfGatA and PfGatB in E. coli. A two-step purification procedure combining Ni-NTA affinity and size exclusion chromatography allowed purification of 10 mg of PfGatA and PfGatB per liter of culture. SDS-PAGE analysis of the purified enzymes corroborated the predicted molecular masses of the two open reading frames, GatA and GatB (85.0 and 94.5 kDa, respectively, Fig. 6, A and B), indicating that both P. falciparum subunits could be expressed and purified independently. This was also the case with Helicobacter pylori GatCAB (22), but not with Bacillus subtilis GatCAB, in which the GatA subunit could not be expressed in E. coli in the absence of GatB (59).
Before assaying PfGatAB for amidotransferase activity, we first performed control reactions to verify that it would not glutamylate 32P-labeled apicoplast tRNAGlu or tRNAGln. The control reactions showed that PfGatAB did not exhibit any glutamylation activity (Fig. 6C, lanes 1 and 2). We then tested PfGatAB for transamidation activity in a reaction containing l-glutamate as the amide donor and either P. falciparum 32P-labeled Glu-tRNAGln or Glu-tRNAGlu that had been produced using recombinant PfGluRS (23). PfGatAB transamidated neither of these glutamylated tRNA substrates (Fig. 6C, lanes 3 and 4), implying that PfGatAB does not utilize l-glutamate as an amide donor. However, PfGatAB transamidated Glu-tRNAGln to form Gln-tRNAGln in the presence of l-glutamine (Fig. 6C, lane 6) but as expected did not utilize Glu-tRNAGlu as an amide acceptor (Fig. 6C, lane 5). These data show that PfGatAB has a high specificity for Glu-tRNAGln and utilizes l-glutamine as the amide donor in the transamidation reaction.
Next, we explored the possibility that PfGluRS and PfGluAdT could act together in a single reaction to sequentially form Gln-tRNAGln from precursors. We performed an experiment that allowed the transamidation reaction to take place immediately after aminoacylation by premixing equimolar amounts of PfGluRS, PfGatA, and PfGatB for 3 min on ice and then incubating them at 37 °C with all substrates required for the glutamylation and amidation reactions. Formation of Glu-tRNAGln and its conversion to Gln-tRNAGln were observed (Fig. 6D, lane 3). Similar results were obtained when we used l-asparagine as the amide donor for the transamidation reaction (Fig. 6D, lane 4), implying that PfGatAB can utilize both l-asparagine and l-glutamine as amide donors. We also noted that the Glu-AMP spot in the TLC assay was less intense than the Gln-AMP (Fig. 6D, lanes 3 and 4) and control Glu-AMP spots (Fig. 6D, lane 2), indicating that the sequential glutamylation and amidotransferase reactions proceeded rapidly. This could be due to the sequestration and immediate substrate channeling (60) of the misacylated Glu-tRNAGln from the ND-GluRS to GatAB to prevent the release of misacylated Glu-tRNAGln and subsequent misincorporation of l-Glu in place of l-Gln during protein synthesis (61). The different amidotransferase assay approaches used here and the results obtained demonstrate that PfGatAB does not require the GatC subunit for transamidation reaction.
We did not test whether PfGatAB possesses Asp-AdT activity as bacterial GatCABs do because the Plasmodium genome possesses two predicted AspRSs, one cytoplasmic and the other apicoplast-targeted (11, 25, 56, 62), as well as an apicoplast-targeted AsnRS. Thus, the apicoplast appears to possess all components required for asparaginylation of apicoplast tRNAAsn via direct aminoacylation. Together, these data are consistent with the conclusion that PfGatAB is a Glu-AdT.
To date, two different tRNA-dependent AdTs are known: the heterotrimeric GatCAB (22) and the heterodimeric GatDE (20) enzymes. The latter is an archaeal signature enzyme and serves as the Glu-AdT for Gln-tRNAGln biosynthesis in archaea (20). GatCAB is found in both bacteria and archaea (20, 59). In archaeal genomes, GatCAB is encoded only when an AsnRS is not (63). All bacterial GatCAB enzymes studied to date are able to serve as both a Glu-AdT and an Asp-AdT in vitro (64,–69). The activity/ies actually performed by bacterial GatCABs in vivo is/are determined by the non-discriminating aaRS (ND-GluRS and/or ND-AspRS) possessed by each organism. For example, bacteria such as B. subtilis (70) that have an ND-GluRS but lack an ND-AspRS use their GatCAB solely as a Glu-AdT (22). In bacteria possessing an ND-AspRS but lacking an ND-GluRS (e.g. Pseudomonas aeruginosa, Neisseria meningitidis, Thermus thermophilus, and Deinococcus radiodurans), GatCAB serves only as an Asp-AdT (64, 65, 71,–76). In bacteria carrying both non-discriminating aaRSs (ND-GluRS and ND-AspRS) such as Chlamydia trachomatis (66) and H. pylori (77,–79), GatCAB serves as a Glu/Asp-AdT (66, 68, 69). The P. falciparum genome (11) encodes two putative d-AspRS enzymes, one of which (PF3D7_0514300) possesses a predicted N-terminal apicoplast-targeting sequence, suggesting that it is imported into the apicoplast (56, 62). The other AspRS (PF3D7_0102900) lacks an apicoplast-targeting signal and is probably cytoplasmic (57, 58, 80). Similarly, Plasmodium contains two putative AsnRSs as follows: one apicoplast-targeted (PF3D7_0509600) and the other cytoplasmic (PF3D7_0211800) (11, 24, 25). Thus, in the cytoplasm and apicoplast of Plasmodium, Asn-tRNAAsn is formed via direct aminoacylation.
We previously demonstrated that the P. falciparum nuclear genome (23) encodes two putative GluRS enzymes. One, Pf3D7_1349200, appears to be cytoplasmic (58). The other, Pf3D7_1357200, possesses a predicted N-terminal apicoplast-targeting sequence (56, 62), was localized to the apicoplast, and exhibits non-discriminating glutamylation activity in vitro, producing both Glu-tRNAGlu and Glu-tRNAGln. It is the first enzyme in the apicoplast's indirect aminoacylation pathway (23). In this study we have further dissected the apicoplast's indirect aminoacylation pathway by identifying the PfGatA and PfGatB subunits of the apicoplast aminoacyl-tRNA AdT. Using episomal constructs in which GFP was fused to the predicted PfGatA and GatB apicoplast targeting sequences, we demonstrated that GFP was trafficked to the apicoplast in erythrocytic stage parasites. Minor overlaps between the anti-GFP and MitoTracker Red signals were observed where the apicoplast and mitochondrion appeared to contact one another (Fig. 3, C, panel ii, and D, panel ii), a phenomenon observed with other apicoplast-targeted proteins (81, 82). Furthermore, we myc-tagged the endogenous P. berghei chromosomal gatA gene, and we observed that the tagged protein was trafficked to the apicoplast, confirming the results obtained in P. falciparum with episomal constructs. Localization of P. falciparum GatAB solely in the apicoplast differs from the situation in Arabidopsis, where the GatCAB amidotransferase is targeted to both the plastid and the mitochondrion (83). Dual targeting of PfGatAB to the plastid and mitochondrion is also unlikely because, as in the related parasite T. gondii, the Plasmodium mitochondrion probably imports aminoacylated tRNAs from the cytoplasm (58).
We expressed recombinant PfGatA and PfGatB independently in E. coli (Fig. 6, A and B), and we combined them in vitro to examine their ability to transamidate apicoplast Glu-tRNAGlu or Glu-tRNAGln that had been previously glutamylated using recombinant PfGluRS (23). PfGatAB demonstrated a remarkable tRNA substrate specificity by converting Glu-tRNAGln to Gln-tRNAGln but did not transamidate Glu-tRNAGlu (Fig. 6C). The Plasmodium apicoplast possesses an ND-GluRS (23) but lacks an ND-AspRS (57, 58), and therefore it almost certainly utilizes the PfGatAB as a Glu-AdT. As in Plasmodium, bacteria such as B. subtilis (70) that possess an ND-GluRS but lack an ND-AspRS use their GatCAB only as a Glu-AdT (64). Because genes encoding GatA and GatB are present in all known Plasmodium genomes but not in any known organism in the other two domains, we concluded that GatAB is the Plasmodium glutamyl-tRNAGln amidotransferase (GatAB/Glu-AdT).
We also showed that PfGluRS, PfGatA, and PfGatB, when briefly pre-mixed, can glutamylate apicoplast tRNAGln and transamidate it to form Gln-tRNAGln in vitro (Fig. 6D). This suggests that the Plasmodium apicoplast may contain a tRNAGln·ND-GluRS·GatA·GatB complex, akin to the transamidosomes described in archaea (61) and the eubacterium T. thermophilus (84). Such a complex would prevent challenging the genetic code integrity as demonstrated for tRNA-dependent Asn formation (61).
We used different approaches to test for the PfGatAB-catalyzed amidotransferase reaction. Our findings consistently showed that the Plasmodium parasite has established an evolved glutamyl-tRNA amidotransferase reaction that takes into account the absence of the gatC gene in the parasite genome and therefore does not require it for the transamidation reaction, contrary to the amidotransferase activity of the GatCAB paralog where the three subunits are required for enzyme activity (22). This is the first biochemical evidence for Glu-tRNAGln transamidation by a GatAB in the absence of a GatC subunit.
We investigated the evolution of Plasmodium GatA and GatB subunits in comparison with the human GatA and GatB, the GatA and GatB subunits of bacterial GatCABs, the GatA and GatB subunits of other plastids, and the GatD and GatE subunits of archaeal GatDE. In the unrooted phylogeny of GatB and GatE proteins, Plasmodium GatBs were not placed within the bacterial GatB, archaeal GatE, or the plastid GatB clades but were placed on a separate branch (Fig. 3A). The divide is well supported with a bootstrap value of 100. In a similar fashion, the unrooted phylogeny of GatA and GatD proteins, Plasmodium GatA was not placed within the bacterial GatA, the plastid GatA, or the archaeal GatD clades but was placed on a separate branch (bootstrap value of 100) (Fig. 3B). Additionally the GatB and GatA sequences of T. parva and Babesia bovis, which like Plasmodium lack gatC in their genomes, were placed in the same clade as the Plasmodium orthologs. Taken in total, the results strongly suggest that GatAB, which is uniquely found in Apicomplexa that possess an apicoplast, is a paralog to GatCAB, GatFAB, and GatDE.
To identify conserved and divergent features of PfGatB and PfGatA, homology modeling was performed in comparison with the known S. aureus GatCAB structure (Fig. 1) (28, 29). All functional features of bacterial GatCAB and archaeal GatDE are conserved in PfGatAB, but we found that PfGatAB had unique inserts that could not be fitted into the GatCAB model (Fig. 1). In archaeal GatDE, the unstructured N-terminal insert of GatD has been reported to play a structural role that gives it the ability to associate to GatE (31). GatF found only in fungal genomes (46, 47) and GatC found in bacterial, plant, and mammalian AdTs have similar unstructured characteristics and play a similar structural role where they reinforce the interaction between the GatA and GatB subunits by encircling their interface (28, 29). We were not able to identify a homolog of the gatC or gatF genes in the P. falciparum genome, leading us to speculate that the unique inserts present in PfGatAB could be playing a yet undetermined structural role of holding PfGatA and PfGatB together, similar to that of the insertion found in GatD. Crystallization studies on the PfGatAB enzyme will help determine the roles of these inserts in catalysis.
Here, we have shown that besides possessing an apicoplast-targeted, non-discriminating GluRS (23), malaria parasites contain a unique apicoplast-targeted glutamyl-tRNA amidotransferase that amidates Glu-tRNAGln in the absence of GatC. The apicoplast is an attractive drug target because it is essential to both blood and liver stage parasites (18, 19), harbors several metabolic pathways absent in the host, and its transcriptional and translational machinery are of bacterial origin. The apicoplast's indirect aminoacylation pathway is probably essential in malaria parasites because the parasite genome does not encode an apicoplast-targeted GlnRS (57, 58, 85). PfGluRS, the first enzyme in the apicoplast's indirect aminoacylation pathway, was refractory to gene deletion and is therefore essential for blood stage development (23). In this study, the gene encoding for apicoplast PbGatA was refractory to gene deletion, despite the fact that it was accessible to myc tagging. Thus, both PfGluRS and PfGlu-AdT are potentially good drug targets.
The emergence of resistant strains of P. falciparum continues to fuel an urgent need to develop new antimalarials. Besides our findings in this study, the Plasmodium GatAB amidotransferase has also been identified to be an important antimalarial drug target using in silico molecular modeling approaches (86). However, despite the chemical diversity of compound collections, robust inhibitors that act specifically against Plasmodium GatAB amidotransferases and not other amidotransferases have yet to be identified. Further investigations examining the requirements for apicoplast indirect aminoacylation across the Plasmodium life cycle and screening of compounds to identify specific inhibitors of plasmodial GluRS and Glu-AdT enzymes may lead to novel ways to target this pathway for chemotherapy.
M. J. G. conceived and directed the project. M. J. G. and B. M. designed the research; B. M.M., K. F. W., K. B., G. R., L. L., J. A., and T. M. N. performed the experiments; and B. M. M., M. J. G., K. F. W., and K. B. wrote the manuscript. All authors approved the final version of the manuscript.
We thank Geoffrey McFadden for providing anti-ACP antibody, J. Lapointe and A. Weiner for plasmids expressing CCA-adding enzyme, and Y. M. Hou for the pGFIB vector. We thank MR4 for providing P. falciparum 3D7 parasites contributed by Dan Carucci and Alister Craig. The unpublished P. berghei genome sequence data were produced by the Pathogen Sequencing Group at the Wellcome Trust Sanger Institute and can be obtained on line. The unpublished C. velia and V. brassicaformis ortholog sequences were graciously provided by Arnab Pain and Hifzur Ansari of King Abdullah University of Science and Technology. The Chromera and Vitrella sequencing project was funded by a King Abdullah University of Science and Technology OCRF (GCR) award (FIC/2010/09) to Arnab Pain. We also thank Jonathan Eisen and Dongying Wu for their consulting on the phylogenetic analyses.
*This work was supported, in whole or in part, by National Institutes of Health Grant R21AI81097 from the NIAID (to M. J. G.). This work was supported by the M. J. Murdock Charitable Trust, the Center for Infectious Disease Research, and Puget Sound Partners for Global Health (to M. J. G.) and by Deutsche Forschungsgemeinschaft Award BE 1540/11-2 (to K. B.). The authors declare that they have no conflicts of interest with the contents of this article.
2The abbreviations used are: