Biochemical Journal

Research article

Overexpression of a cytosolic pyrophosphatase (TgPPase) reveals a regulatory role of PPi in glycolysis for Toxoplasma gondii

Douglas A. Pace , Jianmin Fang , Roxana Cintron , Melissa D. Docampo , Silvia N. J. Moreno


PPi is a critical element of cellular metabolism as both an energy donor and as an allosteric regulator of several metabolic pathways. The apicomplexan parasite Toxoplasma gondii uses PPi in place of ATP as an energy donor in at least two reactions: the glycolytic PPi-dependent PFK (phosphofructokinase) and V-H+-PPase [vacuolar H+-translocating PPase (pyrophosphatase)]. In the present study, we report the cloning, expression and characterization of cytosolic TgPPase (T. gondii soluble PPase). Amino acid sequence alignment and phylogenetic analysis indicates that the gene encodes a family I soluble PPase. Overexpression of the enzyme in extracellular tachyzoites led to a 6-fold decrease in the cytosolic concentration of PPi relative to wild-type strain RH tachyzoites. Unexpectedly, this subsequent reduction in PPi was associated with a higher glycolytic flux in the overexpressing mutants, as evidenced by higher rates of proton and lactate extrusion. In addition to elevated glycolytic flux, TgPPase-overexpressing tachyzoites also possessed higher ATP concentrations relative to wild-type RH parasites. These results implicate PPi as having a significant regulatory role in glycolysis and, potentially, other downstream processes that regulate growth and cell division.

  • Apicomplexa
  • cellular metabolism
  • glycolysis
  • parasite
  • PPi
  • pyrophosphatase (PPase)
  • Toxoplasma gondii


PPi is a byproduct of many biosynthetic reactions (the synthesis of nucleic acids, coenzymes, proteins and isoprenoids, and the activation of fatty acids), and it has been proposed that the removal of PPi by PPases (pyrophosphatases) makes biosynthetic reactions thermodynamically favourable [1]. In addition, bioenergetic and regulatory roles of PPi have been suggested [2]. PPi can be generated by photophosphorylation, oxidative phosphorylation and glycolysis, and can be used in a number of reactions to replace ATP [3].

The cytosolic concentration of PPi is regulated in higher organisms, predominantly through the activity of soluble cytosolic PPases [4]. Inorganic PPases include membrane-bound V-H+-PPases (vacuolar H+-translocating PPases) and soluble-form PPases. The membrane-bound V-H+-PPases utilize the energy released by hydrolysis of PPi to transport protons across the membrane of cells or organelles [58]. The soluble inorganic PPases that hydrolyse PPi to Pi are essential enzymes and have high activity in the cytoplasm. The absence of these PPases would lead to the build-up of toxic levels of PPi, accounting for the essential nature of the enzymes. Two families of non-homologous soluble inorganic PPases have been described: family I PPases, which are widespread in all types of organisms and prefer Mg2+ as a cofactor [9,10], and family II PPases, which are exclusive to bacteria and prefer Mn2+ as a cofactor [911]. One of the most studied family I PPases is that from Saccharomyces cerevisiae [12]. In addition to its PPase activity, this enzyme displays polyphosphatase activity in the presence of transition metal ions such as Zn2+, Mn2+ and Co2+ as cofactors [1316], and it also can hydrolyse organic tri- and di-phosphates, such as ATP and ADP [1618].

An unusual characteristic of Toxoplasma gondii, a major opportunistic pathogen of fetuses from recently infected mothers and of patients with AIDS, is that it possesses cellular levels of PPi that are higher than those of ATP [19]. In addition, it stores PPi and poly P (polyphosphate) in acidic organelles named acidocalcisomes [2022]. Acidocalcisomes have been found in a number of organisms from bacteria to humans [23]. Acidocalcisomes from T. gondii are characterized by their electron density, high content of cations bound to PPi and poly P, and a number of pumps in their membranes, among them a V-H+-PPase, which contributes to their acidification [2022,24]. Incubation of fixed Trypanosoma cruzi [25] or Trypanosoma evansi [26] cells with a PPase removes the electron-dense matrix of acidocalcisomes, which indicates that PPi is an important component of this organelle's structure. In addition to its use by the acidocalcisomal V-H+-PPase [21,27], T. gondii PPi can also be used in place of ATP as an energy donor in the PPi-dependent PFK (phosphofructokinase) reaction [28].

In the present study, we characterized biochemically a soluble PPase and named it TgPPase (T. gondii PPase). By overexpressing this enzyme in T. gondii tachyzoites we were able to isolate clones with up to ten times higher enzymatic activity than wild-type cells. This high cytosolic PPase activity altered the cytosolic concentration of PPi, which was significantly reduced when compared with the cytosolic level in wild-type RH tachyzoites. These mutant cells showed alterations in their glycolytic pathway, leading us to propose a regulatory role for PPi in the glycolytic pathway of these parasites.


Chemicals and reagents

AMDP (aminomethylenediphosphonate) was synthesized by Professor Michael Martin (Department of Biophysics, University of Illinois at Urbana-Champaign, Urbana, IL, U.S.A.). Restriction enzymes, T4 DNA ligase, reverse transcriptase, Taq polymerase, DNA ladder, TRIzol® reagent and goat serum were from Gibco. The pET28a+ expression system, Ni-NTA (Ni2+-nitrilotriacetate) His·Bind® resin and benzonase nuclease were from Novagen. The pCR2.1-TOPO cloning kit, secondary antibodies, BCECF [2′,7′-bis-(2-carboxyethyl)-5(6)-carboxyfluorescein] and BCECF-AM (where AM is acetoxymethyl ester) were from Invitrogen. Hybond-N nylon membrane, HiTrap desalting columns and the ECL (enhanced chemiluminescence) kit were obtained from Amersham Pharmacia Biotech. All other reagents were of analytical grade.

Culture methods

T. gondii tachyzoites (RH) were grown in hTERT (human telomerase reverse transcriptase) host cells using protocols described previously [29]. Transgenic fluorescent T. gondii tachyzoites expressing a YFP (yellow fluorescent protein)–YFP fusion gene were a gift from Dr Boris Striepen (Department of Cellular Biology, University of Georgia, Athens, GA, U.S.A.) [30].

T. gondii growth measurements

[3H]Uracil incorporation was conducted in hTert cells that were cultured in 12-well plates for 24 h before they were challenged with 1×105 tachyzoites per well. [3H]Uracil incorporation was measured 24 h later by measuring the amount of [3H]uracil incorporated into each well during the last 4 h [31,32]. T. gondii plaque assays were performed as described previously [33]. Assays were conducted in six-well plates each containing a confluent layer of hTERT host cells. Parasites (200 per well) were incubated for 9 days to allow invasion and replication (formation of plaques). Plaque number and relative plaque area (i.e. the percentage of total area occupied by a plaque-forming unit) were determined using ImageJ software (

TgPPase cDNA cloning by 5′ and 3′ RACE (rapid amplification of cDNA ends)

The protein sequence of the Trypanosoma brucei soluble inorganic PPase (GenBank® accession number AAP74702) was used to search the T. gondii genome database, revealing three polypeptides sharing high similarity to known soluble inorganic PPases. The 5′ and 3′ RACE technique was used to sequence the 5′ and 3′ ends of the TgPPase cDNA, using information from known contigs and EST (expressed sequence tag) sequences. The Invitrogen Life Technologies kit for 5′ and 3′ RACE was used.

Database search and phylogenetic analysis

BLAST analysis of the TgPPase protein against the OrthoMCL database (version 2) ( was performed to search for putative PPase orthologues. A multiple sequence alignment from the sequences obtained was created by ClustalW [34] and optimized using GeneDoc (version 2.6.002) [35] and MacClade (version 4.06) [36].

Phylogenetic trees were built on the basis of the optimized alignment using the parsimony and distance methods from the Felsenstein PHYLIP package (version 3.65). Briefly, 100 alignment replicates were created by the bootstrap method using the SEQBOOT program [37,38]. The bootstrapped dataset was used to generate trees by the parsimony method using the PROTPARS program. For calculating distance, PROTDIST was used to generate 100 matrices from the bootstrapped dataset, by applying the Dayhoff PAM (point accepted mutation) evolutionary model of amino acid substitutions [39]. A total of 100 distance trees were built by the neighbour-joining method [40]. The final consensus trees were built using the majority rule and were visualized using TreeViewX (version 0.4) and Mega 3.1 (version 3.1) [41].

Isolation of cells overexpressing TgPPase

The TgPPase gene was amplified using primers (5′-AGATCTATGCAGTCTGCACCTCTGGC-3′ and 5′-CCTAGGCGGTAGCCACAACTTTTG-3′) containing BglII and AvrII restriction enzyme sites (underlined) by RT (reverse transcription)–PCR. The PCR products were cloned into the expression vectors pTubP30-FLAG/sagCAT (a gift from Dr William Sullivan, Department of Pharmacology and Toxicology, Indiana University School of Medicine, Indianapolis, IN, U.S.A.) [42], replacing the P30 gene. The resulting construct pTub-TgPPase–FLAG/sagCAT was sequenced and used for transfection of T. gondii tachyzoites using published protocols [33]. Selection was performed with 20 μM chloramphenicol. Cloning was performed by limited dilution and one clone was selected for further analysis and named TgPPase-OE (TgPPase-overexpressing cells). A clone expressing two copies of the gene encodingYFP was used for immunofluorescence co-localization assays [30].

Preparation of recombinant TgPPase

The whole open reading frame of TgPPase was amplified by RT–PCR with the primers 5′-CATATGCAGTCTGCACCTCTGGC-3′ and 5′-GTCGACCGGTAGCCACAACTTTTGC-3′, containing the restriction sites for NdeI and SalI (underlined). The PCR product was cloned into the pCR 2.1-TOPO TA vector and subcloned into the expression vector pET28a+. The recombinant construct TgPPase/pET28a was transformed into Escherichia coli BL21-CodonPlus(DE3)-RIPL (Stratagene), and protein expression was induced with 1 mM IPTG (isopropyl β-D-thiogalactopyranoside). rTgPPase (recombinant TgPPase) was purified with a His·Bind Quick 900 cartridge according to the manufacturer's instructions (Novagen). The eluted fractions were pooled and desalted using a HiTrap column.

Antibody generation and purification

The purified rTgPPase protein was sent to Cocalico Biologicals for production of guinea pig polyclonal antiserum. The antiserum was affinity-purified with the cyanogen-bromide-activated resin [43].

SDS/PAGE and Western blot analysis

Proteins were separated by SDS/PAGE using standard protocols. Western blot analysis was performed as described previously [44] using an affinity-purified anti-TgPPase polyclonal antibody (1:10000 dilution) or anti-(α-tubulin) monoclonal antibody (1:15000 dilution).

Fluorescence microscopy

Immunofluorescence assays were performed as described previously [44] by using as primary antibody the purified anti-TgPPase at 1:100 dilution. The secondary antibody was Alexa Fluor® 555-conjugated goat anti-guinea pig IgG (H+L) (1:1000 dilution). Images were collected using an Olympus IX-71 inverted fluorescence microscope with a Photometrix-cooled charge-coupled-device camera (CoolSnapHQ), and deconvolved for 15 cycles using Softworx deconvolution software (Applied Precision).

PPase activity measurement

rTgPPase activity was assayed by measuring the release of Pi using a method described previously [45]. The standard assay mixtures varied based on what was specifically being tested. For pH-optimum experiments, assay conditions were 20 mM Tris/Hepes, 1.36 ng of purified rTgPPase protein and 23 μM PPi with 3 mM MgCl2 (as a cofactor) for experiments using PPi as a substrate. The same buffer and amount of protein were used with 9 μM poly P3 (tripolyphosphate) and 3 mM CoCl2 (as a cofactor) for experiments using poly P3 as a substrate. Experiments testing inhibition of rTgPPase activity used 50 mM Tris/HCl (pH 8.5) when PPi was the substrate and 50 mM Mes (pH 6.0) when poly P3 was the substrate. For experiments testing divalent cation cofactor preference, the assay conditions were similar, but cofactors were used as stated in the Figure legends at a final concentration of 3 mM. Experiments testing alternative substrates [100 μM GP4 (guanosine tetraphosphate), 100 μM ATP or 100 μM poly P75] used assay mixtures containing 50 mM Tris/HCl (pH 8.5) or 50 mM Mes (pH 6.5) and 3 mM MnCl2 and 45 ng of purified rTgPPase. After a 10 min incubation at 37°C, reactions were stopped by the addition of an equal volume of a mixture of three parts of 0.045% Malachite Green and one part of 4.2% ammonium molybdate [46]. The absorbance at 660 nm was measured with a SpectraMax M2e plate reader (Molecular Devices). The specific activity of rTgPPase was defined as μmol of Pi released·min−1 per mg of protein. For determination of optimal pH conditions, a 20 mM Tris/Hepes mixed buffering system was employed to ensure differences in activity were only due to pH and not ionic conditions. Inhibitor experiments were conducted at Km concentrations for both PPi and poly P3.

To measure PPase activity in total cell lysates, purified tachyzoites were washed twice with BAG [buffer A plus glucose: 116 mM NaCl, 5.4 mM KCl, 0.8 mM MgSO4, 50 mM Hepes, (pH 7.2) and 5.5 mM glucose], and resuspended in a small amount of the same buffer. The cells were broken by sonication in a Branson Sonifier 450 instrument (10% amplitude, two 5 s pulses).

The activity was also measured in subcellular fractions obtained after lysis of tachyzoites with silicon carbide as described previously [24]. The homogenate was centrifuged at 100 000 g for 60 min at 4°C to obtain a supernatant and a pellet fraction. The enzymatic activity was measured as described above.

Extraction and determination of PPi and ATP

PPi and ATP were extracted with ice-cold 0.5 M HClO4 as described previously [24]. PPi measurements were obtained by measuring the amount of Pi released upon treatment with TIPP (Thermostable Inorganic Pyrophosphatase; NEB). The reactions were performed in 50 mM Tris/HCl (pH 7.5) and 6 mM MgCl2, using 0.04 units/ml of inorganic PPase. After a 20 min incubation at 30°C, the Pi released was measured with Malachite Green as described previously [45]. Using the same HClO4 extraction, samples were used for the determination of short-chain poly P by quantifying the amount of phosphate after treatment with S. cerevisiae exopolyphosphatase.

For the determination of cytosolic and organellar PPi, purified tachyzoites (4–8×108 cells) were washed once in a hypo-osmotic lysis buffer [20 mM Hepes (pH 7.3), 100 mM sucrose and 10 mM sodium fluoride] by centrifugation at 1000 g. The pellet was subjected to three cycles of freezing and thawing using a solid CO2/ethanol bath (5 min) and a 37°C water bath (1 min), resuspended in hypo-osmotic lysis buffer and centrifuged at 14000 g for 20 min at 4°C. The supernatant was taken as the cytosolic fraction and the pellet as the organellar fraction containing PPi stored predominantly in acidocalcisomes. The efficiency of this fractionation was assessed by testing for LDH (lactate dehydrogenase) activity, a cytosolic enzyme in T. gondii [47] that therefore should not be detected in the organellar fraction. LDH activity was monitored fluorimetrically by using the LDH present in the experimental samples to oxidize NADH to NAD+ as the enzyme converted pyruvate (supplied in the reaction buffer) into lactate (excitation at 340 nm, emission at 460 nm). Both fractions were analysed as described above for PPi and short-chain poly P. The intracellular and organellar concentrations of PPi were calculated from the respective cell volume of 16.5±3 μl for 109 cells [24], assuming that PPi is mainly located in acidocalcisomes and these occupy 1% of the total cell volume. Acidocalcisome volume was estimated assuming a spherical shape with a diameter of 200 nm and an average of ten acidocalcisomes per extracellular tachyzoite.

ATP was measured using an ATP determination kit (Molecular Probes). For calculation of the intracellular concentrations of ATP in tachyzoites, a cell volume of 16.5±3 μl for 109 cells was used [24].

Measurement of proton extrusion in wild-type (RH) and TgPPase-OE cells

Rates of proton extrusion were measured in RH and TgPPase-OE cells following basic procedures [48]. Freshly egressed tachyzoites were washed with BAG and split into two portions: cells washed further and suspended in BAG, and cells washed further and suspended in buffer A without glucose. Both cell preparations were kept on ice at a concentration of 1×109 cells·ml−1 until analysis. Proton extrusion was measured as described previously [48]. The fluorescence was measured using a FL-4500 fluorescence spectrophotometer (Hitachi). Calibration of fluorescence measurements to specific pH values was performed as described previously [48].

Measurement of intracellular pH in wild-type (RH) and TgPPase-OE cells

Loading of RH and TgPPase-OE cells with BCECF-AM for pH measurements was performed as described previously [49]. Intracellular pH was measured in a SpectraMax M2e plate reader at excitation wavelengths of 505/440 nm and emission at 530 nm. The calibration of the pH units/fluorescence ratio was as described previously [49,50].

Measurement of lactate extrusion in wild-type (RH) and TgPPase-OE cells

Freshly egressed tachyzoites were washed three times with BAG and resuspended in the same buffer at a cell concentration of 5×108 cells·ml−1. Reactions were terminated at different times (0–10 min) by spinning the cells down and removing the supernatant for analysis of the extruded lactate. This metabolite was quantified by the enzymatic conversion of lactate into pyruvate using LDH (Sigma) and measurement of the concomitant production of NADH. Enzymatic assay conditions were set up following published conditions [51,52]. A 70 μl aliquot equivalent to 3.5×107 cells was added to 200 μl of reaction mixture (600 mM glycine, 346 mM hydrazine, 17 mM EDTA, 15 mM NAD and 10 units·ml−1 LDH; pH=9.2) and fluorescence was monitored on the SpectraMax M2e plate reader for 20 min. NADH was measured using excitation and emission wavelengths of 340 nm and 460 nm respectively. Fluorescence values were converted into units of lactate using a standard curve where known amounts of L-lactate were added to the reaction mixture and NADH production was monitored.


Cloning the gene encodingTgPPase

The gene encoding a soluble PPase was identified by a BLASTN search with the T. brucei soluble inorganic PPase sequence (GenBank® accession number AAP74702). The T. gondii gene encodes a protein of 381 amino acids (TgPPase) with a predicted molecular mass of 42 kDa. The TgPPase sequence has an overall identity with Y-PPase (yeast PPase) of only 28%, but the 13 essential amino acids that form the active-site structure of the Y-PPase [inferred from the crystal structure (PDB code 1WGJ)] are conserved [53] (see Supplementary Figure S1 at The gene is expressed in both tachyzoites and bradyzoites (, TGME49_083830).

Phylogenetic analysis of the TgPPase gene

The TgPPase amino acid sequence (AAU88181) was used as a query to search for orthologues in the OrthoMCL database (version 2). Our phylogenetic analysis shows two main branches with high bootstrap support (100%/72%) when kinetoplastid inorganic PPases are used as the outgroup (Figure 1). The two groups are (i) bacterial and bacterial-like plant soluble PPases, and (ii) plant chloroplast and fungal/animal cytosolic and mitochondrial PPases (Figure 1). In addition, our phylogenetic tree supports the notion that the T. gondii enzyme is a type I inorganic PPase since it clusters with fungal/animal and plant chloroplast soluble inorganic PPases with high bootstrap confidence levels using both distance and parsimony methods (74%/74%) (see Supplementary Figure S2 at for the alignment of the sequences used for the tree in Figure 1).

Figure 1 Molecular phylogenetic analysis of type I inorganic PPases

The consensus phylogenetic tree was built from 100 replicates using the distance and parsimony methods as described in the Experimental section. Bootstrap values from 100 replicates are shown in bold and italics as obtained by the distance and parsimonius methods respectively. The 0.2 bar represents amino acid substitutions per site. Sequence accession numbers as provided by GenBank® are indicated in parentheses as follows: Aquifex aeolicus VF5 (NP_214066.1), Arabidopsis thaliana chloroplast PPase (At5g09650.1), A. thaliana (At1g01050.1), Chlamydomonas reinhardtii mitochondrial PPase (AJ298232), C. reinhardtii chloroplast PPase (AJ298231), Cryptosporidium parvum (cgd4_1400), E. coli W3110 (NP_418647.1), Homo sapiens cytosolic PPase (ENSP00000317687), H. sapiens mitochondrial PPase (ENSP00000343885), Kluyveromyces lactis cytosolic PPase (KLLA0E17721g), K. lactis mitochondrial PPase (KLLA0E11055g), Leishmania major putative mitochondrial PPase (LmjF03.0910), Mus musculus cytosolic PPase (ENSMUSP00000020286), M. musculus mitochondrial PPase (BAB22922), Nostoc sp. PCC 7120 (P80562), Oryza sativa chloroplast PPase (3698.m00155), O. sativa (12001.m13461), Plasmodium falciparum 3D7 (PFC0710w), Rhodospirillum rubrum (AF115341_1), S. cerevisiae cytosolic PPase (YBR011C), S. cerevisiae mitochondrial PPase (YMR267W), Solanum tuberosum (CAA12415), Synechocystis PCC 6803 (P80507), Thermoplasma acidophilum (P37981), Thermus thermophilus (BAA24521), T. gondii (AAU88181), T. brucei putative mitochondrial PPase (Tb927.3.2840/Tb03.27C5.190), T. cruzi putative mitochondrial PPase (Tc00.1047053508181.140) and Zea mays (O48556). CHL, chloroplastic, CYT, cytosolic, MIT, mitochondrial.

Expression, purification and biochemical characterization of TgPPase

The expression of TgPPase and purification by affinity chromatography produced a single protein band (see Supplementary Figure S3 at The purified protein was characterized biochemically.

The enzymatic activity of the rTgPPase showed an absolute requirement for divalent cation cofactors, with Mg2+ being the most efficient at a concentration of 1 mM (238.17±6.51 μmol Pi·min−1 per mg of protein). The optimum pH for the hydrolysis of PPi was 8.5 (Figure 2A). At this pH, other cations such as Co2+, Zn2+, Ca2+ and Mn2+ marginally stimulated the activity over basal levels (Figure 2C). The enzyme was also able to hydrolyse poly P3, although with lower efficiency (Table 1). For this polyphosphatase activity the preferred cofactor was Co2+ (Figures 2B and 2D), as occurs with Y-PPase [13,16], and the optimum pH range was 6–6.5. The enzyme had low hydrolysing activity of long-chain poly P (poly P75), ATP and GP4 in the presence of Mn2+ as the divalent cation cofactor (see Supplementary Figures S4A and S4C at

Figure 2 Enzymatic characterization of rTgPPase

PPase activity was determined as described in the Experimental section using 23 μM PPi (A and C) or 9 μM poly P3 (B and D). (A and B) Optimum pH measurements with PPi (A) or poly P3 (B) as substrate. A 20 mM Tris/Hepes mixed buffering system was used to manipulate the pH of the reaction buffer. (C and D) Optimum cation concentration measurements with PPi (C) or poly P3 (D) as substrate. Results are expressed as percentage of maximum activity, taken as 100%. Where indicated, 5 mM EDTA was used. (E and F) Enzymatic activity measured with different concentrations of PPi (E) or poly P3 (F). Experiments were performed using 3.0 mM MgCl2, (E) or 3 mM CoCl2 (F). Insets represent the linear transformation, by double reciprocal plot, of each curve. Experiments were repeated three times, each one in triplicate, with similar results. Results are means±S.E.M. for three experiments.

View this table:
Table 1 Kinetic parameters of rTgPPase with different substrates

Km and Vmax were calculated using SigmaPlot 10.0.

Table 1, Figures 2(E) and 2(F) and Supplementary Figures S4(B) and S4(D) show the kinetic properties of the enzyme for the hydrolysis of PPi, poly P3, poly P75, ATP and GP4. At the optimal conditions, it is clear that the catalytic efficiency for the hydrolysis of PPi (measured as kcat/Km) is significantly higher than for the hydrolysis of the other substrates.

Inhibition of the rTgPPase

AMDP, a specific inhibitor of V-H+-PPase, did not show any effect on the activity of this enzyme at a concentration of 40 μM (results not shown). However, NaF, a known inhibitor of soluble PPases, inhibited the enzyme with an IC50 of 223±31.8 μM (see Supplementary Figure S5A and Supplementary Table S1 at Poly P3 hydrolytic activity was also significantly inhibited by fluoride (Supplementary Figure S5C). Sodium molybdate caused minor inhibition of PPi hydrolysis (maximum inhibition=27%, Supplementary Figure S5B). The small amount of activity for poly P3 hydrolysis was effectively inhibited by sodium molybdate, a general phosphatase inhibitor. IDP (imidodiphosphate; 40 μM), a PPi analogue, did not have any effect on the activity of rTgPPase (results not shown).

Subcellular localization of TgPPase

To investigate the localization of TgPPase, we performed indirect IFA (immunofluorescence analysis) in tachyzoites expressing cytosolic YFP using antibodies against the TgPPase enzyme. As evidenced by the colocalization with YPF, the endogenous localization of the TgPPase enzyme was determined to be in the cytosol in tachyzoites of T. gondii (Figure 3A). Western blot analysis of cytosolic and membrane fractions of tachyzoites confirmed the cytosolic localization of the protein (Figure 3B). Both the specific activity and the total activity of the enzyme were higher in the soluble fractions (Figures 3C and 3D).

Figure 3 TgPPase localizes to the cytosol of wild-type RH T. gondii tachyzoites

RH (wild-type) tachyzoites expressing YFP were used to test for colocalization of endogenous expression of TgPPase. (A) TgPPase localized to the cytoplasm of RH tachyzoites. Parasites were fixed and stained with an anti-TgPPase antibody (1:100 dilution) or observed by direct YFP fluorescence where indicated. The overlay image shows co-localization of both proteins in the cytosol. Scale bars, 5 μm. (B) Western blot analysis of subcellular fractions of T. gondii tachyzoites. Equal protein amounts (5 μg) from supernatant (S) and pellet (P) fractions were loaded. The molecular mass standards (in kDa) are shown on the left-hand side. The blue arrow indicates the band corresponding to the TgPPase. (C and D). Specific and total enzymatic activity, respectively, in supernatant (S) and pellet (P) fractions after centrifugation of lysates at 100 000 g for 1 h. Other experimental details are as described in the Experimental section. DIC, differential interference contrast.

Effect of overexpression of TgPPase on cell growth and metabolism

A clone of T. gondii tachyzoites was isolated after limited dilution of cells transfected with the pTubTgPPase-FLAG/sagCAT plasmid and selection with chloramphenicol. These cells showed significantly higher levels of TgPPase protein (Figure 4A) and enzymatic activity (Figure 4B; P<0.05). Further IFA localization studies with cytosolic YFP determined that the TgPPase protein in OE (overexpressing) mutants maintained a cytosolic localization (results not shown). Rates of growth in OE mutants using uracil incorporation were found to be significantly lower (P<0.05) when compared with the growth rate of wild-type RH parasites (Figure 4C). The lower growth rates were confirmed using plaque assays to measure growth/invasion efficiency in wild-type and OE mutants (Figures 4D–4F). OE mutants had significantly lower plaque numbers (P<0.001) as well as lower relative size of plaque-forming units (P<0.05).

Figure 4 Cells overexpressing TgPPase show higher levels of TgPPase protein and activity, but grow at a slower rate

(A) Western blot analysis using RH cells and TgPPase-OE cells transfected with the expression vector ptubTgPPase-FLAG/sagCAT and selected with chloramphenicol as indicated in the Experimental section. The blot was sequentially probed with anti-TgPPase antibody (upper panel) and anti-(α-tubulin) antibody as loading control (lower panel). (B) PPase activity of crude cell lysates obtained as described in the Experimental section. AMDP, a specific inhibitor of V-H+-PPase at 40 μM, was added to the lysate. (C) OE mutant tachyzoites had significantly lower [3H]uracil incorporation than wild-type cells after a 24 h inoculation of hTERT host cells. The results are representative of three independent experiments, each one performed in triplicate. Results are means±S.D. *P<0.05 (Student's t test). (DF) Plaque assays comparing growth and invasion ability of wild-type cells (RH) and TgPPase overexpressing mutants (OE). (D) Determination of plaque-forming units in a monolayer of hTERT host cells. A total of 200 parasites were incubated in each well for 9 days then washed away; remaining host cells were stained to determine the number of zones cleared of host cells due to parasite invasion and replication (i.e. plaques). Results are averages±S.E.M., n=6. (E) Determination of relative area of plaques. Data taken from the same experiments shown in (D). The area represents the percentage of the total available area occupied by the average plaque. Single-factor ANOVA (df=1,11) showed significantly lower plaque numbers and plaque area in OE mutant cells. *P<0.05; ***P<0.001. (F) Representative wells used to determine plaquing efficiency for RH and TgPPase OE mutant parasites.

To understand the possible consequences of elevated PPase activity in OE mutants, we investigated the cellular amounts of phosphate-related molecules. Cellular levels of PPi were 40% lower than in wild-type RH cells (Figure 5A; P<0.01). Cellular concentrations of short-chain poly P in OE mutants and RH cells were similar at ~5 mM in phosphate equivalents (Figure 5B). Under identical conditions as above, cellular concentrations of ATP were significantly elevated (P<0.05) by 75% in extracellular tachyzoites overexpressing TgPPase (Figure 5C). However, under conditions in which no glucose was present, the difference in ATP concentration was abolished.

Figure 5 T. gondii parasites overexpressing TgPPase contain lower PPi and higher ATP levels

(A) Overexpression of TgPPase leads to a significant reduction in total PPi levels as compared with wild-type RH tachyzoites (ANOVA: df=1,16; *P<0.01). Results are averages±S.E.M of nine quantifications for each cell type. (B) No difference in short-chain poly P levels between wild-type RH and OE mutants for TgPPase. Results are averages of nine determinations for each cell type. Values are in phosphate equivalents of hydrolysed short-chain poly P groups. (C) Overexpression of TgPPase leads to an increase in the intracellular levels of ATP in the presence of glucose. Newly released tachyzoites were filtered and washed twice with BAG with or without glucose and incubated in the same buffer for 1 h. ATP was extracted and measured as described in the Experimental section. The results are representative of three independent experiments, each one performed in triplicate. Results are means±S.D. *P<0.05 (Student's t test).

In order to rule out the possibility that the PPase OE phenotype was only a consequence of genetic manipulation and therefore non-specific in nature, we measured cellular PPi concentration and growth in mutant parasites expressing an unrelated gene, the fluorescent tomato protein (hereafter referred to as Tomato OE mutants). The Tomato OE mutants were derived from the same genetic background as the PPase OE mutants and underwent similar genetic manipulation. The cellular PPi concentration in the Tomato OE parasites was similar to that measured in wild-type RH parasites (see Supplementary Figure S6A at Likewise, growth rates in these unrelated expression mutants were similar to those of wild-type RH parasites as measured by plaque assays (Supplementary Figure S6B). These results indicate that the phenotype observed in the PPase OE mutants is not an artefact of genetic manipulation and bolster the relationship between PPi concentration and ATP and growth.

Since most PPi in T. gondii is localized in acidocalcisomes [24], we investigated whether cytosolic or organellar PPi was affected in cells overexpressing the PPase. The distribution of PPi in the cytosolic and organellar fractions of extracellular tachyzoites was determined by lysis of the plasma membrane by freeze–thaw cycling in a hypotonic buffer. In order to determine the concentration of PPi in these fractions, we assumed that the cellular volume and the size and number of acidocalcisomes in the OE mutants were similar to those of wild-type parasites (i.e. 1×109 parasites=16.5 μl and acidocalcisomes occupy 1% of the cellular volume). This assumption was confirmed in our microscopic analyses and in our indirect immunofluorescence assays examining acidocalcisome markers (e.g. TgVP1; results not shown). The majority of PPi was found in the organellar fraction (~ 360 mM; Figure 6B). A significant difference, however, was observed in the soluble fraction (Figure 6A). RH cells had more than six times the concentration of PPi than OE mutants (0.55 and 0.09 mM respectively; P<0.01). The efficiency of this fractionation was verified by measuring LDH activity in both the supernatant (i.e. cytosolic) and the pellet (i.e. organellar) fractions. Over 95% of measured LDH activity was found in the cytosolic fraction (results not shown). In conjunction with the PPi distribution, these results demonstrate effective lysis of the plasma membrane without significant disruption of organelles containing PPi (i.e. acidocalcisomes). These results indicate that the total cellular differences in PPi are a result of different concentrations of PPi in the cytosol, which is in agreement with the cellular location of the TgPPase enzyme (Figure 3).

Figure 6 PPi content of cytosolic and acidocalcisome fractions

PPi was determined in cytosol (A) and 14 000 g pellet (acidocalcisome) (B) fractions obtained as described in the Experimental section from wild-type (RH) and TgPPase-OE parasites. Results represent the average concentration±S.E.M. of three quantifications for each cell type. Concentration was determined by dividing the total PPi (nmol) in the cytosolic and organellar fractions by their respective estimated volumes, as described in the Experimental section. The cytosolic PPi concentration was significantly lower in OE mutants (ANOVA: df=1,4; *P<0.01).

Metabolic changes in TgPPase-OE cells as measured by proton extrusion, lactic acid release and intracellular pH

T. gondii is known to preferentially metabolize glucose to lactic and acetic acids and excrete them into the medium, accompanied by protons [54]. The large glucose-dependent differences observed in ATP concentration in conjunction with the cytosolic differences in PPi concentration between RH and OE mutant parasites prompted us to investigate the glycolytic flux of extracellular tachyzoites. Proton extrusion and lactate extrusion were measured in extracellular tachyzoites in the presence and absence of glucose. In cells that rely on glycolysis for energy production, these processes have been shown to be reliable metrics of glycolytic flux [5557].

To investigate whether the increase in ATP levels resulted as a consequence of an increased glycolytic rate, proton extrusion was measured in RH parasites and OE mutants, using the free-acid form of BCECF and quantifying the rate at which the pHe (pH of extracellular medium) changed. Proton extrusion was measured in RH cells (Figures 7A, 7C and 7E) and OE mutants (Figures 7B, 7D and 7F) in the presence of glucose (5 mM) and compared with cells deprived of glucose for ~2 h (Figures 7A and 7C, RH cells; Figures 7B and 7D, OE mutants). Furthermore, 5 mM glucose was added to the cells deprived of glucose during the proton extrusion measurement to evaluate the immediate response in cellular metabolism (Figures 7A, trace b, and 7E, RH cells; Figures 7B, trace b, and 7F, OE mutants). RH cells showed a small, but significant, increase (ANOVA: df=1,11; P<0.01) in proton extrusion when glucose was present (Figure 7A, trace c) or when glucose was added during the experimental incubation period (Figure 7A, trace b) relative to rates when cells were glucose-deprived (Figure 7A, trace a). Proton extrusion rates in OE mutants exhibited a greater sensitivity to the presence or absence of glucose. In the absence of glucose, (Figure 7B, trace a) rates of proton extrusion in OE mutants were slightly lower, although statistically similar to rates in glucose-deprived RH cells (average glucose-deprived rates were −4.3×10−4 and −3.1×10−4 pHe units·s−1 for RH and OE cells respectively). When glucose was present in the experimental buffer, rates of proton extrusion in OE cells exhibited a significant 3.2-fold increase (ANOVA: df=1,11; P<0.001) from −3.1×10−4 to −9.8×10−4 pHe units·s−1 (Figure 7B, traces b and c). Average rates of proton extrusion while glucose was present were significantly higher (ANOVA: df=1,11; P<0.001) in OE cells by 1.6-fold, relative to RH cells (−9.8×10−4 and −6.0×10−4 pHe units·s−1 for TgPPase-OE and RH cells respectively; compare Figures 7D and 7F with 7C and 7E, grey columns). Despite the large differences in proton extrusion, there was no significant difference in pHi (intracellular pH) between RH parasites and OE mutants (ANOVA: df=1,5; P>0.05; results not shown). The average pHi was 7.18 and 7.23 for RH and OE cells respectively. Rates of proton extrusion in the unrelated Tomato OE mutants were similar to those of wild-type parasites (Supplementary Figure S7 at, providing further support for the relationship between cellular PPi concentration and glycolytic flux in T. gondii.

Figure 7 Proton extrusion by wild-type (RH) and TgPPase-OE parasites

Rates of proton extrusion were measured using the free-acid form of BCECF. (A, C and E) Summary of proton extrusion in RH cells. (B, D and F) Summary of proton extrusion in OE mutant cells. (A) Representative tracings depicting changes in pHe in RH tachyzoites in the presence of 5 mM glucose (black, trace c), no glucose (light grey, trace a) and in cells where 5 mM glucose was added 300 s into recording (dark grey, trace b). (B) Same tracing as in (A) using OE mutants. (C) Resultant slope values (change in pHe per s) in the presence (grey) or absence (white) of glucose for proton extrusion in RH cells using data represented in traces a and c in (A). (D) Resultant slope values (change in pHe per s) in the presence (grey) or absence (white) of glucose for proton extrusion in OE mutant cells using data represented in traces a and c in (B). (E) Rate of pHe change before (white) and after (grey) addition of 5 mM glucose to medium in RH cells using data represented in trace b in (A). (F) Rate of pHe change before (white) and after (grey) addition of 5 mM glucose to medium in OE mutant cells using data represented in trace b in (B). All estimates shown in panels (CF) were determined from three independent experiments, and error bars represent the S.E. of the slope.

To better assess the link between PPi and glycolytic flux, rates of lactate extrusion were measured in RH and soluble PPase OE mutants as an independent verification of the rate of glycolysis. Rates of lactate extrusion were measured in RH cells and OE mutant cells in which glucose had been added to the experimental buffer (5 mM). Duplicate measurements of the amount of lactate extruded to the extracellular medium over the course of 10 min (measurements were taken at 0, 1, 2, 5 and 10 min) showed a significant linear increase with time for both cells types (Figure 8). The lactate extrusion rate for RH cells was 1.07±0.03 (S.E.M.) fmol·h−1 per cell, and for the OE cells (Figure 8, inset) it was 1.90±0.04 (S.E.M.) fmol·h−1 per cell (ANOVA: df=1,18; P<0.001). The difference in lactate extrusion between RH and OE cells was 1.8-fold, a value similar to the difference in proton extrusion rates for both cell types (1.6-fold, see above).

Figure 8 Lactate extrusion in wild-type (RH) and TgPPase-OE parasites

Lactate extrusion rates were measured using an enzyme-coupled reaction (LDH) fluorescently monitoring the production of NADH as lactate is converted into pyruvate. Lactate extrusion was monitored in RH (white circles) and OE mutant (grey circles) tachyzoites with duplicate samples taken at 0, 1, 2, 5 and 10 min for each cell type. Regression analysis was performed on each cell type, and the resultant slope, depicting the rate of lactate extrusion, is shown in the inset as the cell-specific rate of lactate extrusion. Error bars are the S.E. of the slope. *P<0.001; ANOVA comparison of slopes between RH and TgPPase-OE cells (df=1,18; F value=253).

The difference in the proton and lactate extrusion rates between RH and OE cells might be due to an inhibitory effect of PPi on the glycolytic pathway. The T. gondii PK (pyruvate kinase) is an allosteric enzyme and is activated by glucose 6-phosphate [58]. We investigated whether PPi could be inhibiting this enzyme by measuring its activity in a T. gondii lysate in the presence of various PPi concentrations. For the range of PPi concentrations that we measured in the cytosol (i.e. up to 1 mM), we found no detectable inhibition of PK (results not shown).


The goals of the present study were to characterize the enzyme that directly regulates cytosolic levels of PPi in the apicomplexan parasite T. gondii and employ its expression as a means to manipulate cytosolic PPi concentrations in this medically relevant parasite. Bioinformatic searching (e.g. BLASTN searching and querying for soluble PPase yielded TgPPase as the only soluble PPase found in the T. gondii genome. PPi is both an important substrate and product for several metabolic reactions, and its generation in the cytosol is highly regulated so as to avoid a build-up of toxic concentrations. For these reasons, we chose to use the overexpression of TgPPase as a means to manipulate the cytosolic concentration of PPi. It was our expectation that, given the placement of PPi in so many key metabolic reactions, its manipulation would have far-reaching consequences on the metabolism of T. gondii tachyzoites.

The alignment of the predicted TgPPase protein sequence shows conservation of amino acids present in other PPases. Considering its phylogenetic position, its Mg2+ requirement and its fluoride sensitivity, the TgPPase belongs to the family I PPases. Phylogenetic analysis shows that apicomplexan PPases cluster with the sequences belonging to mitochondria and chloroplasts, suggesting a proteobacterial origin. The cytosolic localization of TgPPase could be due to specific requirements of this enzyme in T. gondii. One hypothesis could be that the TgPPase gene was acquired through horizontal gene transfer, a mechanism that has been shown to occur in apicomplexan parasites [5962]. Since the kinetoplastid PPases are highly divergent but still related to other type I inorganic PPases, we selected them as an outgroup (Figure 1). In a study by Gómez-García et al. [63], the Leishmania major inorganic PPase was suggested to be an ancestral type I eukaryotic soluble PPase with a calcium-dependent activity.

The catalytic efficiency of the purified recombinant TgPPase enzyme, as shown by the kcat/Km values in Table 1, indicates that the enzyme is highly efficient when hydrolysing PPi. The enzyme is also able to hydrolyse poly P3, poly P75, ATP and GP4 with lower efficiency (Table 1). It is surprising that the recombinant enzyme shows an optimum pH of 8.5 and at the same time is cytosolic (Figure 3). On analysing Figure 2(A) it can be observed that the activity is still high at cytosolic pH 7.1–7.4. The enzyme preference for Mg2+, when hydrolysing PPi, and for other non-physiological cations such as Co2+ or Mn2+, when hydrolysing other substrates, is similar to what has been found for Y-PPase [16].

Given that the phylogenetic analysis and biochemical characterization fully supported the identity of TgPPase as a Type I PPase with a high specificity for PPi and only much lower affinities for alternative substrates, we employed the overexpression of this enzyme as a means to decrease the cytosolic concentration of PPi. Overexpression successfully increased the production of TgPPase protein as well as its activity. This increased activity was still maintained in the cytosol of OE mutants, as measured by fractionation experiments and indirect immunofluorescence assays. In conjunction with this, we observed a significant decrease in the substrate of TgPPase, PPi, which was localized to the cytosol of OE mutants. These results confirm that the overexpression of TgPPase is a reliable tool for understanding the metabolic significance of PPi.

The phenotype of the T. gondii clone overexpressing the gene encoding TgPPase (OE mutant) was somewhat unexpected. Relative to the wild-type parasites, OE mutants had higher levels of ATP. T. gondii tachyzoites rely on glycolysis for energy production [64,65]. Unlike glycolysis in mammalian cells, T. gondii possesses a PFK that utilizes PPi to drive the rate-limiting conversion of fructose 6-phosphate into fructose 1,6-bisphosphate [28]. Our initial expectation was that the decrease in cellular PPi concentration would cause a decrease in glycolysis, since a key energetic substrate had been significantly reduced. Measurements of ATP concentration showed that, contrary to this expectation, energy metabolism had increased in the OE mutants. To ensure that the increase in ATP was a specific outcome of increased energy metabolism, we measured rates of glycolytic flux.

TgPPase OE mutants displayed a significant increase in glycolysis, as measured by both proton and lactate extrusion from extracellular tachyzoites. Rates of proton and lactate extrusion have been shown to be tightly linked to glycolytic flux in a wide range of prokaryotes and eukaryotes, including bacteria [56,66], yeast [67] and human platelets [55]. T. gondii, like other apicomplexans, obtains most of its energy through glycolysis [64,65], with lactate representing a dominant end-product of glycolysis [54,68] Recently, Lin et al. [65] used lactate extrusion to measure glycolysis in extracellular tachyzoites of T. gondii. These reasons compelled us to use rates of proton and lactate extrusion as metrics of glycolytic flux for direct comparison between RH parasites and OE mutants. Both cell types responded to the addition of 5 mM glucose with increased rates of proton extrusion. However, the rate of extrusion was ~60% higher in OE mutants when in the presence of glucose. Similarly, rates of lactate extrusion in the presence of 5 mM glucose were significantly higher (~80%) in the OE mutants. The high sensitivity and immediate response to the addition of glucose in both cell types is not surprising given that the localization of glycolytic enzymes in extracellular tachyzoites is at the pellicle region just below the plasma membrane [47]. This stimulation in glycolytic flux resulted in a quantitatively similar increase in ATP concentration in the OE mutants (i.e. ATP concentration increased by 75%). To show that these phenotypic traits observed in the TgPPase OE mutants are specific to the reduction in PPi, we compared our results from wild-type RH cells and TgPPase OE mutants with a completely unrelated expression mutant. Genetically modified parasites expressing the fluorescent tomato protein had similar levels of PPi to RH and, likewise, similar rates of growth and glycolysis (as measured by proton extrusion).

The regulatory mechanism by which cytosolic concentrations of PPi can be related to higher glycolytic flux in T. gondii remains undetermined. Regulation of glycolysis has been studied in several organisms from bacteria to higher eukaryotes. It has been demonstrated in other cell types that PPi inhibits several glycolytic enzymes, such as hexokinase in T. brucei [69] and PEPCK (phosphoenolpyruvate carboxykinase) in T. cruzi [70]. In animal cells, PPi, either administered directly or produced intracellularly by activation of short-chain fatty acids, produces effects that mimic glucagon injection in that liver glucose is increased, and there is a change in the 3-phosphoglycerate/pyruvate ratio, suggestive of PK inhibition [4]. Three enzymes that typically are observed to be critical regulators of glycolytic flux are hexokinase, PFK and PK. In T. gondii, however, very little is known about the regulation of glycolysis [71]. T. gondii possesses one annotated hexokinase, but it does not use PPi as a substrate and is not inhibited by it either [71]. Our present analysis of PK in T. gondii wild-type and TgPPase-overexpressing cells found no significant difference in PK activity for the range of PPi differences between the two cell types.

As mentioned earlier, T. gondii tachyzoites possess a plant-like PFK that uses PPi as the high-energy phosphoryl donor rather than ATP (PPi-PFK). Therefore a decrease in PPi would be expected to result in a corresponding decrease in the rate-limiting conversion of fructose 6-phosphate into fructose 1,6-bisphosphate by PPi-PFK. However, the Km for PPi of PPi-PFK is 33 μM [28]. The cytosolic concentrations of both wild-type and OE mutants may be high enough (550 and 90 μM respectively) to result in only minimal differences in PPi-PFK activity. An alternative explanation that has been proposed in plant cells [72] is that the increase in Pi from the breakdown of PPi results in activation of fructose-6-phosphate 2-kinase, the product of which is fructose 2,6-bisphosphate, a strong activator of PPi-PFK [72]. However, unlike plant cells, the PPi-PFK of T. gondii does not appear to be activated by fructose 2,6-bisphosphate [28].

The up-regulation of glycolysis and increase in ATP levels due to decreases in cytosolic PPi has also been observed in plant cells [72,73]. In developing tubers of potato, cytosolic overexpression of PPase causes a decrease in cytosolic PPi as well as an unexpected increase in sucrose degradation and subsequent increases in glycolysis and starch production [73]. The up-regulation of these pathways is thought to be due to the PPi-dependent sucrose-degradation pathway and downstream activation of starch synthesis. The cytosol of T. gondii possesses crystalline, water-soluble, polysaccharide particles called amylopectin granules, which are similar to those of plant starch [74]. Although these granules are typically associated with the bradyzoite and sporozoite stages, it has been shown that acidification of the medium for tachyzoites can cause a significant increase in the presence of amylopectin granules [74]. The increase that we observe in glycolysis and concomitant proton extrusion may be related to the activation of amylopectin synthesis and therefore analogous to stimulation of starch synthesis seen in plant cells. In addition, this effect could lead to the observed decrease in uracil incorporation and plaque formation, demonstrating a decrease in growth rates in overexpressing parasites.

It is a widely held concept that soluble PPases play an important metabolic role by driving reactions such as protein, RNA and DNA synthesis to completion [1] and that the PPi produced in anabolic reactions is instantly hydrolysed. However, a potential role of PPi as a regulator of metabolic processes has also been proposed [4]. Our results support this assertion. Our findings highlight the regulatory role of PPi on a critical metabolic pathway of T. gondii. The regulatory role of PPi requires further study. Metabolomic analysis in conjunction with the use of these overexpressing mutants to modify the cytosolic concentration of PPi should provide insights into further downstream consequences resulting from changing cytosolic PPi concentrations. Such analysis may have far-reaching implications for the role of PPi in metabolic regulation in a variety of cell types.


Douglas Pace assisted in experimental design, performed most of the experiments, and was a primary designer and writer of the manuscript. Jianmin Fang performed experiments pertaining to enzymatic characterization of TgPPase and immunofluorescent localization of TgPPase in T. gondii tachyzoites, and assisted with writing the manuscript. Roxana Cintrón performed the phylogenetic analyses of TgPPase and assisted with writing the manuscript. Melissa Docampo assisted with enzyme characterization experiments. Silvia Moreno designed experiments and was a primary designer and writer of the manuscript.


This work was supported by the National Institutes of Health (NIH) [grant number AI-079625 (to S.N.J.M)]. D.A.P. was partially supported by an NIH T32 training grant [grant number AI-60546] to the Center for Tropical and Emerging Global Diseases. R.C. was supported in part by an NIH research supplement [grant number 3R01AI068647-04S1].


We thank Professor Boris Striepen (University of Georgia, Athens, GA, U.S.A.) for transgenic fluorescent T. gondii tachyzoites and the ptubYFP-YFP/sagCAT expression vector, Dr William Sullivan (Indiana University School of Medicine, Indianapolis, IN, U.S.A.) for the pTubP30-GFP-FLAG/sagCAT expression vector and Takashi Asai for useful discussions. Nhu-y Nguyen provided assistance with proton extrusion measurements as part of an undergraduate research project. Joanna Cox, supervised by Dr Shuhong Luo, participated in the initial cloning of the TgPPase gene as part of an undergraduate project. Cuiying Jiang and Allysa Smith provided excellent technical help.


  • The nucleotide and protein sequence data reported will appear in GenBank®, EMBL, DDBJ and GSDB Nucleotide Sequence Databases under the accession numbers AY619688 and AAU88181 respectively.

Abbreviations: AM, acetoxymethyl ester; AMDP, aminomethylenediphosphonate; BAG, buffer A plus glucose [116 mM NaCl, 5.4 mM KCl, 0.8 mM MgSO4, 50 mM Hepes, (pH 7.2) and 5.5 mM glucose]; BCECF, 2′,7′-bis-(2-carboxyethyl)-5(6)-carboxyfluorescein; GP4, guanosine tetraphosphate; hTERT, cells, human telomerase reverse transcriptase cells; IFA, immunofluorescence analysis; LDH, lactate dehydrogenase; OE, mutant, overexpressing mutant; pHe, pH of extracellular medium; pHi, intracellular pH; poly, P, polyphosphate; poly, P3, tripolyphosphate; PPase, pyrophosphatase; PK, pyruvate kinase; PFK, phosphofructokinase; RACE, rapid amplification of cDNA ends; RT, reverse transcription; TgPPase, Toxoplasma gondii soluble PPase; rTgPPase, recombinant TgPPase; TgPPase-OE, cells, TgPPase overexpressing cells; V-H+-PPase, vacuolar H+-translocating PPase; YFP, yellow fluorescent protein; Y-PPase, yeast PPase


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