Identification of candidate apicoplast proteins based on their gene expression profile

To more fully understand the many functions of the apicoplast, we wished to establish a comprehensive catalog of organellar proteins. To this end, we developed a multi-step pipeline to prioritize potential candidates. In the first step, we paid particular attention to the timing of gene expression. Behnke and colleagues have recently discovered common periodic profiles of mRNA abundance for distinct organellar proteins in Toxoplasma. This cell cycle timing appears to coincide with the formation of the organelles. Similar time-ordered relationships leading to shared transcription patterns had previously been observed for organellar biogenesis during replication of Plasmodium merozoites in the red blood cell [34]. Thus, co-regulation of transcription might be used as one criterion to identify unknown components of organelles [35], [36]. To explore whether timing of mRNA expression could be used to identify new apicoplast genes, we compiled a list of 20 proteins, for which apicoplast localization had been confirmed in T. gondii and Plasmodium (Table S1). The mRNAs encoding these proteins have similar cell cycle profiles with peak levels during the G1 to S phase transition [36]. A group of 369 candidate genes (Figure 1B, Table S2) matched this specific periodic pattern with high correlation (GeneSpring 11.0, Pearson correlation at 0.05 FDR). In addition to the original 20 apicoplast proteins the expanded list captured at least 8 previously confirmed apicoplast proteins and proteins from secondary plastids of other organisms (Table S1) demonstrating the predictive value of this approach. To assess the efficiency of this filter, we selected two sets of control genes. One set contained 200 genes whose periodic pattern is inverse to the median expression profile of the 369 genes (R3 and R4 in [36]), none of the confirmed apicoplast genes was found here. Additionally, 200 genes were selected at random from ToxoDB in 3 independent experiments, those included 0, 0 and 2 of the confirmed apicoplast proteins. This suggested significant enrichment in the list used.

In order to further assess candidates and to derive a more focused list suitable for experimental validation, we applied a secondary screen based on molecular phylogeny. The rationale behind this second filter is the hypothesis that genes encoding apicoplast proteins follow a phyletic distribution that is consistent with the evolutionary path that led to the acquisition of the organelle (Figure 1A). The predicted protein sequences of the first 200 candidates in the expression list were retrieved from the T. gondii genome database (http://toxodb.org/toxo/) and used to perform BLAST searches against the NCBI nucleotide collection database (http://blast.ncbi.nlm.nih.gov/), and the results were systematically analyzed. A set of rules was developed by which we scored the presence or absence of homologs in different clades, and the degree of statistical confidence associated with that identification. Based on these rules we assigned positive or negative points to candidates (see Materials and Methods for details). The parameters were iteratively adjusted such that a positive score (1 or more) was obtained for all the 28 known apicoplast proteins (Table S1) mentioned above. Accordingly only genes with positive scores were considered candidates for further characterization. Half (49.5%) of the genes analyzed met this second requirement, and the first 57 genes (Table S3) were selected as a manageable number to further test experimentally.

Identification of 11 novel apicoplast proteins

The next step in our pipeline was to determine experimentally whether a candidate gene product indeed targets to the plastid. We introduced a DNA cassette encoding an epitope-tag and a selectable marker directly into the native gene locus via homologous recombination. The efficiency of in-locus tagging is dramatically increased in T. gondii mutants that lack TgKu80, a key component of the non-homologous end-joining repair machinery [37], [38]. In this mutant, stable transformation occurs predominantly through homologous recombination. Using the approach described by Huynh and coworkers we attempted 3' endogenous tagging for each of the 57 candidates.

After manually reassessing the predicted gene models (see Materials and Methods) we designed primers to generate targeting plasmids through ligation independent cloning for all 57 genes (Table S3). Sequences for 48 genes were successfully amplified and cloned into p3HA.LIC.CATΔpac (see Materials and Methods). Each of the resulting vectors was linearized using a unique restriction site contained within the gene specific sequence (Table S3). DNA was transfected into the newly engineered TATiΔKu80 line (see below for details) and chloramphenicol resistant parasites were established with all 48 constructs. In 30 of these polyclonal pools of stably transfected parasites, we detected unambiguous expression of the epitope-tag by immunofluorescence assay (IFA) (Table S3). 11 candidates (Table 1) were localized to the apicoplast or its immediate proximity (Figure 2A,B and Figure S1). Other localizations observed included mitochondrion (2 gene products), nucleus (5), Golgi apparatus (3), cytosol (2), endoplasmic reticulum (1), vesicular pattern (4) and unknown compartments (2) (Table S3, Figure 2C and Figure S2). Apicoplast-localized gene products therefore make up 36% of the total candidates localized in this study.

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Figure 2. Localization of 30 previously uncharacterized proteins by endogenous epitope tagging.

Fluorescence microscopy analysis of parasites expressing endogenously HA-tagged proteins. (A) shows examples of a luminal apicoplast protein (top panel), a protein localizing to the periphery of the apicoplast (middle panel), and an example for the second patchy peripheral pattern observed (bottom panel). Numbers on the left represent the numbers of proteins showing the described pattern. (B) shows representative image of ATrx2 (top panel) and of PPP1 (bottom panel) localizations. (C) Representative images of tags that resulted in the identification of non-apicoplast proteins. Likely target organelles are indicated. Scale bar = 5 µm. HA in green, CPN60 in red, numbers reflect the ToxoDB gene ID as detailed in the Results section.

https://doi.org/10.1371/journal.ppat.1002392.g002

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Table 1. Summary of the bioinformatics and experimental data for each of the eleven new apicoplast proteins found in this work.

https://doi.org/10.1371/journal.ppat.1002392.t001

Note that some of the non-apicoplast localized proteins, may nevertheless be involved in apicoplast biogenesis. Such may be the case for vesicular proteins. In that context TGME49_070070 is of particular interest. The predicted protein encoded by this gene contains a C-terminal transmembrane domain and share several domains with proteins known for their role in vesicle trafficking. The protein does not overlap with markers of the Golgi apparatus or the apicoplast (Figure S2), and its precise subcellular localization is yet to be determined.

Six new luminal apicoplast proteins have likely roles in genome maintenance and metabolism

As documented by IFA, the 11 new apicoplast proteins showed variation in their staining pattern, suggesting that they might target to different sub-compartments within the apicoplast. We divided them into two initial groups: the first group consists of proteins that fully co-localize with the luminal apicoplast marker CPN60 [13] indicating that they likely are also luminal proteins. The second group consists of proteins that, while consistently in proximity, have little or no direct overlap with staining from CPN60 (Figure 2A,B Figure S1, Table 1).

To gain insight into the function of these proteins we subjected their amino acid sequences to a variety of bioinformatic analyses (Table 1). We found three to be predicted membrane proteins (110770, 059230 and 039320; we refer here to genes and proteins by the number component of the corresponding gene ID in ToxoDB, so that TGME49_110770 becomes 110770). We then identified several conserved functional domains (Table 1). The first of those is encoded by 021330, a homolog of the subunit A of the bacterial type II topoisomerase, DNA-gyrase. The genomes of T. gondii and of the Plasmodium spp. each encode two gyrase subunits (GyrA and GyrB), and both were studied in Plasmodium [39], [40]. While the apicoplast targeting of 021330 was not surprising based on its Plasmodium homolog, other proteins found in the luminal group were less readily anticipated. Three of those are also likely involved in plastid genome maintenance: 00884 encodes a presumptive ATP-dependent DNA helicase that shows significant sequence similarity with the RecG protein of Gram-negative bacteria. RecG is a double-stranded DNA translocase that is involved in DNA recombination and repair [41]. 059230 encodes a domain found in phage-integrases and DNA break-rejoining enzymes, suggesting that this protein is also part of the machinery that mediates apicoplast DNA recombination events. The third protein, 091670, contains domains typically found in RNA-helicases.

The luminal group also includes 039320, which encodes a protein with a Bol-A like domain at its C-terminus. The exact molecular function of BolA proteins is still under study. Escherichia coli BolA is suggested to be implicated in the tolerance towards different environmental pressures, since its expression is induced under stress conditions. This leads to the reduction of cell surface area, via switching between elongation and septation systems during cell division [42]. While these observations propose a potential role in transcription regulation, a recent study raises a second interesting possibility in the context of plastids. A component of the Arabidopsis thaliana plastid Iron-Sulfur cluster pathway, SufE, was reported that also contain a BolA domain in its C-terminus [43]. A second protein found in our screen is proposed to be involved in the same pathway: whereas the product of 021920 is annotated as a hypothetical protein, more careful recent analysis suggests its involvement in iron-sulfur cluster biogenesis as a NifU-like scaffold protein [2].

Taken together proteins from the first group show predicted domains that tie them to luminal functions of the plastid consistent with their observed localization. The last luminal apicoplast protein, 039680 (Figure S1), has no predicted domain from which a putative function could be inferred.

Four new proteins with non-luminal apicoplast localizations

The staining pattern of four of the new apicoplast proteins suggested a potential non-luminal localization within the apicoplast. The protein encoded by 002440 is found in close proximity to CPN60 but shows additional punctate staining around the plastid that has not been previously reported (Figure 2A). The product of 001270 shows staining that is very similar to the previously described circum-plastid localization of ATrx [23] (Figure 2A). Finally the proteins encoded by 110770 (Figure 2B) and 087270 (Figures 2B, 3A) show a patchy peripheral apicoplast localization reminiscent of the localization of Der1, UfD1 and CDC48 proteins that are believed to reside in the periplastid compartment (PPC) – a compartment equivalent to the cytoplasm of the algal endosymbiont [13]. To follow this further we performed cryo-electron microscopy using the endogenously HA-tagged 087270 cell-line, as this protein appeared to be expressed at the highest level. Cryosections were stained with HA specific antibodies followed by gold labeled anti-immunoglobulin. We observed strong labeling of the periphery of the apicoplast, while gold beads were absent from the lumen of the organelle (Figure 3B). Interestingly, we noticed a polarized distribution of the protein in the apicoplast periphery.

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Figure 3. PPP1 is a resident of the apicoplast periphery and most likely the periplastid compartment.

(A) Fluorescence microscopy analysis of parasites expressing endogenously HA-tagged PPP1 (in green in both panels), co-stained with the luminal marker CPN60 (red in upper panel) and peripheral marker ATrx1 (red in lower panel). Note that in both cases the signals do not overlap completely. Scale bar = 5 µM, and scale bar for magnified insert = 1 µM. (B) Electronmicrographs of parasites expressing endogenously HA-tagged PPP1 using immuno-gold staining after cryo-sectioning. The images on the right show only the apicoplast. Scale bar - 1 µm. (C) T. pseudonana cells were transfected with a 267353–GFP encoding plasmid and stable transgenics were established by drug selection. Expression of the transgene was induced by nitrate. Note GFP fluorescence (green) at the center of the chloroplast (imaging chlorophyll autofluorescence in red). Scale bar 2 µm.

https://doi.org/10.1371/journal.ppat.1002392.g003

Two proteins conserved in and unique to the red lineage

Studying the two presumptive PPC proteins in greater detail we noticed a restricted distribution across the tree of life: tBLASTn searches against GenBank and various additional nucleotide databases (including recent EST, GSS and environmental sampling efforts) revealed homologs of these genes only in a relatively small collection of organisms. Specifically, these proteins are restricted to chromalveolates that maintain a plastid, and to red algae. We further noted that the homologs found in the cryptomonads Guillardia theta and Hemiselmis andersenii are encoded in the nucleomorph, the relict nucleus of the algal symbiont that resides in the PPC of some secondary plastids [44]. Both these nucleomorphs are of red algal origin, while the nucleomorph of the chlorarachniophyte Bigelowiella natans is of green algal origin, and does not encode a homolog of this protein.

We performed multiple sequence alignments of the putative orthologs of each of the two putative T. gondii PPC proteins. In both cases we observed a core region conserved among all sequences (Figure 4A and Figure S5B). For 110770, this conserved region corresponded to a predicted thioredoxine-like domain (Trx, superfamily 52833, Figure S5B). The statistical support for a Trx-like domain is not equally strong in all homologs (the E-value for the T. gondii protein is 4e–03). However, we note the invariant presence of a C-X-X-C motif, the critical signature of Trx-domains [45], in all sequences (Figure S5B). These observations suggest 110770 to be a second plastid-resident thioredoxin-like protein in T. gondii, which we here name ATrx2.

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Figure 4. PPP1 is conserved specifically among members of the red algal lineage.

(A) Multiple protein alignment of the putative orthologues of PPP1. Color gradient corresponds to sequence similarity, deep blue represents 100% identity. Size of black bars indicates the level of conservation across the consensus. (B) Neighbor joining tree of PPP1. Bootstrap values are indicated for each node.

https://doi.org/10.1371/journal.ppat.1002392.g004

For 087270 the high similarity block was found at the carboxy-terminal end and revealed a conserved glutamine (Q) rich patch (Figure 4A). We subjected both protein families to phylogentic analyses (see Materials and Methods). Figure 4B shows a neighbor-joining tree derived from 140 unambiguously aligned residues of 087270. The apicomplexan proteins form a well-supported clade, diatoms and brown algae form a sister clade to the apicomplexans, and cryptomonads and red algae are the members most distant to apicomplexans.

We note that while electron microscopy supported localization of 087270 to the apicoplast periphery, its resolution is generally not sufficient to distinguish between the various peripheral compartments. As described previously, the distances between these compartments are similar to the size of the antibodies used for labeling [13], and this makes a firm assignment of a precise subcompartment in apicomplexans difficult. However, in diatoms, periplastid proteins appear to localize to a distinct and identifiable compartment referred to as the blob-like structure at the center of the usually bilobed chloroplast [46]. We identified a diatom homolog for 087270, and decided to localize it in Thalassiosira pseudonana, a genetically tractable diatom. The coding sequence of the corresponding gene (protein ID 267353, JGI: Joint Genome Institute: www.jgi.doe.gov/)) was amplified from cDNA and introduced into a diatom expression vector under the nitrate reductase (NR) promoter, resulting in a C-terminal GFP fusion [47]. The resulting construct was then used to transform T. pseudonana by microparticle bombardment. Stable transformants were selected in the presence of CloneNAT and cloned on agar plates. Clones were propagated under nitrogen starvation and were tested for expression upon induction by growth in nitrate medium for 24 h. As shown in Figure 3C we detected GFP fluorescence mainly in discrete spots at the center of the chloroplast just outside of the plastid lumen (as detected by the red autofluorescence of chlorophyll). This localization pattern is consistent with the previously described “blob-like” structure in diatoms [46], [48]. Taken together our analyses suggest that 087270 likely resides in the apicoplast PPC, we will subsequently refer to this protein as peripheral plastid protein 1 (PPP1).

A robust system for the isolation of conditional mutants

We next wanted to define the biological role of PPP1 using a reverse-genetic loss-of-function approach. We first attempted a direct gene knock-out. We engineered a targeting cosmid by recombineering [6] (Figure S3), replacing the coding sequence within TOXOW30 with a chloramphenicol acetyl transferase marker. The modified cosmid was transfected into a TgKu80 knock-out background (generously shared by Huynh and Carruthers) [38]. Whereas stable drug resistant parasites were established from each of 4 independent transfections, the PPP1 locus was not disrupted in any of the 43 clones that were screened by PCR (Figure S3). These observations suggested that PPP1 may be indispensable for parasite survival. Similarly, we targeted the loci of four additional newly identified apicoplast proteins, two of which we could disrupt (Table 1 and discussion). We therefore next elected to construct conditional mutants. Several approaches have been developed to isolate conditional mutants [49]-[53]. Out of these the tetracycline-regulated transactivator system has been well suited for our studies of apicoplast biology [3], [6], [9], [13], [32]. We reasoned that we could further streamline this approach by engineering a parasite strain that combines high efficiency of homologous recombination with regulated gene expression.

We recombineered cosmid TOXOY50 to replace the coding sequence of the TgKu80 locus with the phleomycin resistance marker BLE [6]. The resulting modified cosmid was then transfected into a line expressing a tetracycline-regulated transactivator (TATi) [51] and stable clones were established by phleomycin selection. A TgKu80 deletion clone was identified by PCR (Figure 5A) and successful deletion of the gene was further verified by Southern blot (Figure 5C).

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Figure 5. Deletion of the TgKu80 gene in TATi transactivator parasite line.

(A) PCR analysis using primers specific for the TgKu80 coding region (left), targeting construct integration (middle), or an unrelated control region (right) using genomic DNA from RHΔHX, TATIΔTgKu80 and RHΔTgKu80 [38] as template. Position of primers is indicated in panel B. (B) Schematic representations of homologous recombination between the BLE modified TOXOW30 cosmid and the TgKu80 locus. The probe used for the southern blot presented in panel C is indicated and sizes expected after BglII digest are shown. (C) Southern blot analysis of the TATiΔTgKu80 clone and its parental TATi line demonstrating TgKu80 disruption.

https://doi.org/10.1371/journal.ppat.1002392.g005

In this background we tested the efficiency of an endogenous promoter replacement strategy. In this strategy, the native promoter is replaced with the tetracycline-regulated promoter [51] via double homologous recombination, resulting in a conditional knock-down mutant in a single step (see scheme in Figure 6A).

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Figure 6. Isolation of a conditional PPP1 mutant by promoter replacement.

(A) Schematic representation of the two-step manipulation of the PPP1 locus: First the gene is endogenously tagged by single cross-over (cassette in green), followed by promoter replacement by double-homologous recombination (cassette in blue). A legend for the tag and promoters used is show in the upper right corner. (B) Southern blot analysis before and after tagging and promoter replacement. Strains, restriction enzymes and expected band-sizes are indicated to the right. Asterisks indicate bands resulting from additional non-homologous integration of the pLIC_3HA plasmid.

https://doi.org/10.1371/journal.ppat.1002392.g006

We assessed PPP1's gene model experimentally and identified its true start site (at position 1,435,362 on chromosome TGME49_chrV). We then engineered a targeting plasmid that flanked a pyrimethamine resistance marker and the regulated promoter with sequences from the PPP1 locus to replace the native promoter with this cassette (see Materials and Methods, and Figure 6A). The resulting vector was linearized with AvrII and transfected into a TATiΔKu80 background, where PPP1 was already endogenously tagged at the C-terminus as described above (Figure 6A). Clones were isolated after pyrimethamine selection and tested for promoter replacement by PCR. Both the C-terminal tagging and the promoter replacement in this (Tet)PPP1(HA) line were confirmed by Southern blot (Figure 6B, and Figure S4).

We next tested whether the inserted promoter can be used to control PPP1 expression by addition of anhydrous tetracycline (ATc) to the growth medium. We measured PPP1 levels using the endogenous HA tag by Western blot and IFA (Figure 7A,B). In the absence of ATc, we detected the two forms of the protein at a level comparable to the tagged parental strain in which expression is driven by the native promoter (Figure 7A). ATc addition resulted in marked reduction of expression, and 48 hours after addition of ATc the protein was no longer detectable (Figure 7A).

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Figure 7. PPP1 is essential for parasites growth.

(A) Western blot analysis was of proteins extracts from PPP1(HA) and (Tet)PPP1(HA) using anti HA antibody. Note comparable levels of PPP1 when expressed from the native promoter (N) or the inducible T7S4 promoter (I). Protein levels decline swiftly upon ATc treatment, numbers indicate days of culture in 0.5 µg/ml ATc. Tubulin (lower panel) serves as a loading control. (B) Fluorescence microscopy analysis of (Tet)PPP1(HA parasites using anti-HA antibody grown in the presence and absence of ATc. (C) Plaque assays performed in the absence (-) or presence (+) of ATc. Parasite line name is indicated to the left. (D) Fluorescence microscopy of parasites expressing a second copy of PPP1, which is labeled using an anti-Ty antibody (red). The apicoplast marker used is the endogenously tagged ACP-YFP (AY, green). Scale bar = 5 µm. (E) Fluorescence growth assays show growth deficiency in ATc treated (Tet)PPP1(HA). Circles, -ATc; squares, +ATc. Each data point represents the mean of six wells, and the error bar gives the standard deviation. (F) Fluorescence microscopic analysis of endogenously tagged 039680 using anti-HA (green) and the luminal marker CPN60 (red) show that 039680 is likely a luminal apicoplast protein. (G) PCR analysis of the promoter replacement clone of 039680 with diagnostic primer sets is consistent with locus modification. The size of amplicons matches the prediction for integration at both the 5' and the 3' insertion site (900 and 1800 bp respectively). (H) Plaque assays performed with the promoter replaced 039680 cell line in the presence or absence ATc show that this gene is required for growth.

https://doi.org/10.1371/journal.ppat.1002392.g007

PPP1 is essential for parasite survival and apicoplast biogenesis

In order to establish whether PPP1 is essential for T. gondii survival, parasite growth was examined by plaque assay. (Tet)PPP1(HA) parasites, their parental ((PPP1(HA)) and grand-parental (TATiΔKu80) lines were all capable of forming plaques when grown in normal medium (Figure 7C). However, in the presence of ATc, no plaques were observed in the (Tet)PPP1(HA) line after incubation times as long as 12 days (Figure 7C), while both parental lines form plaques readily under these conditions. To confirm that this growth defect is solely due to depletion of the targeted gene, we re-introduced PPP1 as a Ty tagged minigene under the control of a constitutive promoter. We constructed a vector to target and disrupt the uracil phosphoribosyl-transferase (UPRT) locus and used 5-fluorodeoxyuridine (FUDR) to isolate stable transgenic clones. Integration of PPP1Ty was confirmed by PCR (not shown), as well as by IFA using antiTy antibody (Figure 7D). Complementation of (Tet)PPP1(HA) with PPP1Ty rescued the growth phenotype observed by plaque assay upon ATc treatment (Figure 7C). To time the onset of growth inhibition, an RFP-RFP transgene [32] was introduced into the (Tet)PPP1(HA) line and a clone was isolated by cell sorting [3]. We measured the growth of parasites in the presence and absence of ATc using a real-time fluorescence assay [54]. Parasites cultured in the presence of ATc showed a pronounced growth defect compared with parasites grown without ATc, and this defect became detectable on day 3 (Figure 7E). Taken together these observations demonstrate that PPP1 is essential for parasite growth, and that tightly regulated mutants can be constructed in the TATiΔKu80 strain.

To test the reproducibility of the TATiΔKu80 approach, we next attempted a promoter replacement for 039680, one of the new luminal plastid proteins (Figure S1 and Figure 7F) that we identified (Table 1). This is a protein of unknown function, yet is conserved among apicomplexans. After testing the gene model and identifying its start site (at position 831330 on chromosome TGME49_chrVI), we targeted the promoter as described above for PPP1, and the corresponding (Tet)039680 clones were isolated after selection with pyrimethamine. Integration of the construct into the locus, and promoter replacement, were confirmed by PCR (Figure 7G). We tested the effect of ATc on the growth of these parasites by plaque assay. (Tet)039680 parasites formed plaques in the absence of ATc, but no plaques were observed upon growth in ATc (Figure 7H). We conclude that 039680 has a critical role in the lumen of the apicoplast, and further that the TATiΔKu80 provides a reliable background for the rapid construction of conditional mutants.

PPP1 is required for import into the apicoplast

Having demonstrated that PPP1 is essential for growth, we were next interested to examine its specific biological role. Based on the putative PPC location we hypothesized that it may act in apicoplast biogenesis. To be able to follow organelle biogenesis, the luminal acyl carrier protein (ACP) was tagged with YFP directly in its locus in the mutant line, (Tet)PPP1(HA), to establish a native marker. Our western blot analysis suggested complete depletion of PPP1 between 24 h and 48 h of ATc treatment (Figure 7A). We therefore imaged ACP-YFP parasites at 48 h of growth in ATc. We observed intact plastids of typical shape in essentially all parasites (Figure 8A). This finding argues against a role of PPP1 in apicoplast division which produces aberrant plastid morphology and immediate loss of apicoplasts due to unequal segregation [55]. However, we noted that the intensity of the ACP-YFP signal in the organelle appeared to be reduced, and we observed an accumulation of the reporter protein outside of the plastid (Figure 8A arrowhead). This phenotype was observed previously in mutants of import components and suggested a protein import defect. We therefore next studied the maturation of ACP in this mutant in the presence and absence of ATc. Like many apicoplast proteins, ACP is synthesized with a bipartite N-terminal signal, which is cleaved upon reaching the organelle lumen, and this modification depends on import of the protein into the lumen of the organelle [13], [21], [32]. In cells grown in the absence of ATc, two bands are detected by western blot corresponding to the premature and mature protein forms (Figure 8B -ATc). In contrast, cells grown for 48 hours in ATc show a pronounced reduction of the mature form with concomitant accumulation of premature ACP-YFP (Figure 8B,C 48 h). We no longer detected the mature form after 72 hours of ATc treatment (Figure 8B,C 72 h). These observations suggested that loss of PPP1 results in an import defect.

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Figure 8. Loss of PPP1 results in apicoplast protein import defect.

(A) To study apicoplast biogenesis in the PPP1 mutant we engineered a reporter by tagging the ACP locus with YFP. No overt apicoplast division defects were observed upon 48 h ATc treatment. However, note accumulation of fluorescence outside of the apicoplast in treated parasites after 48 h (arrowheads). (B) Western blot analysis following the maturation of ACP-YFP in the (Tet)PPP1(HA) mutant. Parasites were grown in ATc for the times indicated. Anti-GFP antibody was used to detect ACP-YFP. Note the pronounced loss of the mature form of the protein. (C) Bands in B were quantified by densitometry and the ratio of mature to premature protein is shown for each time point. (D) Pulse-chase analysis of protein synthesis and post-translational lipoylation of apicoplast (single band labeled PDH-E2) and mitochondrial proteins (two bands marked mito-E2). Parasites were metabolically labeled as detailed in [13] and lipoylated proteins were isolated by immunoprecipitation using a specific antibody. One hour of labeling (Pulse, P) was followed by a two hour of chase (C) resulting in import and lipoylation of PDH-E2 in non treated parasites. Lipoylation is lost upon ATc treatment. Data shown are representative of four independent experiments. Bands labeled with an asterisk represent lipoylated host cell proteins (note that these bands are observed in host-only controls (Host, C). (E) Bands in D were quantified by densitometry and the ratio of lipoylated PDH-E2 over mito-E2 (upper mitochondrial band) is shown. (F) (Tet)PPP1(HA) parasite were grown in ATc as indicated and plastids were counted in 100 four-parasites vacuoles for each sample. Y-axis shows the percent of parasites that show a clearly identifiable apicoplast.

https://doi.org/10.1371/journal.ppat.1002392.g008

As an independent measure of protein import, we determined the level of lipoylation of the apicoplast pyruvate dehydrogenase E2 subunit (PDH-E2). This was measured in real time by pulse-chase labeling after ATc treatment. Proteins are then immunoprecipitated with an antibody specific for lipoic acid. Protein import is a prerequisite for lipoylation of PDH-E2, as the process requires two apicoplast resident enzymes, and the precursor molecule octanoyl-ACP, which is synthesized de novo in the lumen of the organelle by the type II fatty acid synthesis system [3], [13], [32], [56]. We observed lipoylated PDH-E2 in lysate of untreated cells (Figure 8D, -ATc, C). After 24 hours under ATc treatment, the level of lipoylated PDH-E2 was dramatically reduced, and after 2 days was no longer detected (Figure 8C, +ATc, C and 8D 24 h/48 h). At the same time, lipoylation of mitochondrial proteins remained unchanged (Figure 8D mito-E2 bands).

It was previously documented that ablation of proteins involved in apicoplast import will ultimately result in organelle loss [13], [32]. Using the ACP-YFP marker, we scored the number of plastids at different time points of ATc treatment. At 24 hour and 48 hour time points, apicoplast numbers are comparable to the untreated control (over 92%). In contrast, at the 72 hour time point, we detect plastids in 61.25% of parasites (Figure 8E). This timing is similar to our previous observation for the TgDer1 mutant, a periplastid component of the organellar protein import machinery [13]. Comparing the timing of our various measurements we find that loss of import precedes the loss of the organelle and parasite death. We note that after 72 h growth in the presence of ATc, organelles other than the plastid (such as the mitochondrion, micronemes and rhoptries) were indistinguishable from untreated or parental parasites (data not shown).

Parasite and diatom culture and genetic manipulation

Parasites were grown in hTERT-BJ1 (clontech) cells in supplemented Dulbecco's modified Eagle's medium [67]). Parasite cloning and plaque assays were performed in human foreskin fibroblasts (HFF). For the selection of stable transgenic lines, drugs were added as follow: 1 µM pyrimethamine added one day after transfection for one week, 20 µM chloramphenicol added the day of transfection for three weeks, 5 µM FUDR added two days after transfection for one week. To repress the regulated promoter, parasites were grown in the presence of 0.5 µM anhydrotetracycline (ATc).

Thalassiosira pseudonana (Hustedt) Hasle et Heimdal CCMP1335 was grown in an artificial seawater medium (EASW) according to the North East Pacific Culture Collection protocol (http://www3.botany.ubc.ca/cccm/NEPCC/esaw.html) at 18°C under constant light. Where indicated, NaNO₃ was omitted from the medium (nitrogen-free medium) or replaced by 0.55 mM NH₄Cl (ammonium medium).

Identifying apicoplast protein candidates using bioinformatics resources

The mRNA abundance dataset generated by [36] was interrogated using GeneSpring GX 11..5 (Agilent). We searched for genes with periodic expression patterns that matched each of the 20 “baits” (Table S1), using Pearson correlation at 0.05 FDR. A group of 369 candidate genes (Table S2) was composed where each gene showed high correlation to at least one of the genes in the training group.

We next screened the first 200 genes in this list using a scoring system to filter candidates based on molecular phylogeny. We used tBLASTn to compare each of our candidate amino acid sequences against translated nucleotide databases. Only hits with E-value scores smaller than 1×10⁻⁴ were considered, and the following scores were given to candidates whose homologs were found in the following organisms (organellar genomes): Cyanobacteria (10); Cryptosporidium (−1); No Cryptosporidium (1); Plasmodium (1); distribution of homologs among eukaryotes outside Archaeplastida (−1 to -5, depending on number of organisms and level of similarity and coverage); none of the latter (1); distribution among Archaeplastida (1 to 5); nucleomorph genome (5).

Multiple alignment and phylogenetic analyses

We generated multiple sequence alignments using ClustalX. The pairwise alignment parameters included a gap opening penalty of 35, and gap extension penalty of 0.75. Sequences that were used for alignments were identified on publicly available databases (see results section), and their accession numbers are provided in Table S5. 140 (PPP1) or 99 (ATrx2) unambiguously aligned amino acid positions were used for analysis using Jalview (full alignments available on request). For PPP1 the alignment presented in Figure 4 was used to generate a bootstrapped neighbor joining in ClustalX with 999 repetitions. The tree was visualized with TreeView.

Sequence analyses and gene cloning

Prior to tagging, the computationally predicted gene models for each of the candidates were re-assessed manually, with particular attention paid to the predicted stop codon. We scrutinized expressed sequence tags (ESTs), peptides identified through proteome-analyses, and the presumptive position of active promoters as identified by chromatin modification (all accessed through ToxoDB). We determined alternative gene models for 4 of the 57 candidates. No in silico evidence was available at the time of assessment to dispute the predicted gene models for the remaining 54 genes. Data from deep sequencing mRNA became available at later stages of our study, and supported the predicted C-terminal assignment for 53 of our models (Table S3),

As the size and processing pattern of internally tagged PPP1 detected by western blots was inconsistent with the gene model, we experimentally determined its N-terminus. We tested three alternative translation start sites for which the respective coding sequences were amplified from cDNA, cloned into an expression vector that fuses a C-terminal myc epitope tag [32] and transfected into parasites. Immunofluorescence showed that only the start codon at position 1,435,362 on chromosome TGME49_chrV resulted in apicoplast targeting (data not shown). Western blot analysis of this strain resulted in bands of sizes indistinguishable to those detected from the endogenously HA-tagged protein (not shown). Similarly, for 039680 we experimentally established an alternative start site from the predicted gene model at position 831330 on chromosome TGME49_chrVI. The new gene models are reported in user comments on the corresponding gene pages in ToxoDB and were submitted to genbank (JN053049, JN053050). In both cases, the coding sequence was amplified by PCR from T. gondii RH cDNA using primers introducing flanking BglII or BclII and AvrII or XbaI restriction sites, and products were cloned into BglII and AvrII sites in pDT7S4myc [32].

To modify loci with a triple HA epitope tag, the vector p3HA.LIC.DHFR (generously shared by Huynh and Carruthers [38]) was modified to replace the DHFR (Dihydrofolate reductase) cassette (HindIII and NotI) with a chloramphenicol acetyl-transferase (CAT) expression cassette excised with the same enzymes from pTUB5CAT [68]. Next, the PacI site found within the CAT-ORF was mutated by site directed mutagenesis (Stratagene) to render the PacI site found within the LIC (ligation independent cloning) sequence unique, generating p3HA.LIC.CATΔpac. Targeting sequences were amplified by PCR from T. gondii RH genomic DNA and products were LIC-cloned into p3HA.LIC.CATΔpac as described previously [38]. Positive clones were isolated using primer 1595 in combination with the specific forward primer by PCR screen and confirmed by sequencing. pLIC ACP YFP was generated similarly in pLIC YFP DHFR [38].

For complementation and UPRT targeting, PPP1 was first sub-cloned from pDT7S4PPP1.1myc into pBTTgTPTty [6] using BglII and AvrII sites. Then the entire TUB_PPP1_Ty cassette was excised with BamHI and SpeI and cloned into the BglII and AvrII sites in pUPRT-KO (a gift of Brooke and Gubbels). The resulting vector, pUPRT_(TUB)PPP1Ty was transfected into (Tet)PPP1(HA) and clones were isolated after FUDR (5 µM) selection, and tested for expression of PPP1Ty by IFA.

For promoter replacements pDT7S4myc [32] was modified such that the 3' targeting flank (the coding sequence beginning with the initiation codon) was cloned between BglII and AvrII, and the 5' flank (upstream of predicted promoter region) was cloned between two NdeI sites found in the 5'UTR of the DHFR selection cassette.

Promoter regions were predicted based on the ChIP on chip data [69] viewed through ToxoDB. Accordingly, the 1.7 kb fragment found in positions 1,434,003 to 1,435,362 on TGME49_chrV was amplified with NdeI sites for PPP1, and the 0.7 Kb upstream of position 831330 on chromosome TGME49_chrVI was amplified with MseI sites for 039680. The resulting vectors were linearized with AvrII (PPP1) and BstBI (039680).

All primers used are found in Tables S3 and S4.

Microscopy

Immunofluorescence assays were performed as previously described [70]. We used anti-HA antibodies (Roche) at a dilution of 1∶200, anti-CPN60 [13] at 1∶1000, anti-ATrx1 [23] at 1∶1000, anti-Ty BB2 hybridoma supernatant [71] at 1∶5, anti–Myc (Pierce) at 1∶100, anti-GFP (Roche) at 1∶200, anti-TgMys [72] at 1∶1000, antiROM4 [70] at 1∶100, anti-IMC3 [73] at 1∶500. Fluorescence images were acquired using a Delta Vision microscope as described [13].

For cryo-electron microscopy, infected cells were fixed in 4% paraformaldehyde/0.05% glutaraldehyde (Polysciences Inc.) in 100 mM PIPES buffer. Samples were then embedded in 10% gelatin and infiltrated overnight with 2.3 M sucrose/20% polyvinyl pyrrolidone in PIPES at 4°C. Samples were frozen in liquid nitrogen and sectioned with a cryo-ultramicrotome. Sections were probed with anti-HA antibody followed by a rat secondary antibody conjugated to 18 nm colloidal gold, stained with uranyl acetate/methylcellulose, and analyzed by transmission EM as described previously [74].

Confocal fluorescence microscopy of T. pseudonana was performed using an inverted Zeiss LSM 510 laser scanning microscope (Jena, Germany). Fluorescent signals were detected for GFP (Argon laser, 488 nm) using a 505/550-nm bandpass filter and chloroplast auto-fluorescence (HeNe laser, 543 nm) using a 585 nm long pass filter in the multitrack mode of the microscope. Cells were immobilized for microscopy by a thin slice of 1% Agarose dissolved in EASW medium.

Western blotting and pulse-chase analyses

Western blotting was performed as previously described [32]. We used anti-HA antibodies (Roche) at a dilution of 1∶100, anti-Tubulin [75] at 1∶1000, anti-GFP (Roche) 1∶200 , anti-Myc (Pierce) 1∶100.

Pulse-chase analyses were performed as described previously [13], [32] with the modification that experiments were performed in T25 flasks.