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J Biol Chem
J Biol Chem
The Journal of Biological Chemistry
0021-9258
1083-351X
American Society for Biochemistry and Molecular Biology

S0021-9258(24)02109-4
10.1016/j.jbc.2024.107608
107608
Research Article
Functionality of the V-type ATPase during asexual growth and development of Plasmodium falciparum
Shadija Neeta
Dass Swati
Xu Wei
Wang Liying
Ke Hangjun hk84@drexel.edu
∗
Center for Molecular Parasitology, Department of Microbiology and Immunology, Drexel University College of Medicine, Philadelphia, Pennsylvania, USA
∗ For correspondence: Hangjun Ke hk84@drexel.edu
29 7 2024
9 2024
29 7 2024
300 9 10760824 12 2023
2 7 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Vacuolar type ATPases (V-type ATPases) are highly conserved hetero-multisubunit proton pumping machineries found in all eukaryotes. They utilize ATP hydrolysis to pump protons, acidifying intracellular or extracellular compartments, and are thus crucial for various biological processes. Despite their evolutionary conservation in malaria parasites, this proton pump remains understudied. To understand the localization and biological functions of Plasmodium falciparum V-type ATPase, we employed CRISPR/Cas9 to endogenously tag the subunit A of the V1 domain. V1A (PF3D7_1311900) was tagged with a triple hemagglutinin epitope and the TetR-DOZI-aptamer system for conditional expression under the regulation of anhydrotetracycline. Via immunofluorescence assays, we identified that V-type ATPase is expressed throughout the intraerythrocytic developmental cycle and is mainly localized to the digestive vacuole and parasite plasma membrane. Immuno-electron microscopy further revealed that V-type ATPase is also localized on secretory organelles in merozoites. Knockdown of V1A led to cytosolic pH imbalance and blockage of hemoglobin digestion in the digestive vacuole, resulting in an arrest of parasite development in the trophozoite-stage and, ultimately, parasite demise. Using bafilomycin A1, a specific inhibitor of V-type ATPases, we found that the P. falciparum V-type ATPase is likely involved in parasite invasion but is not critical for ring-stage development. Further, we detected a large molecular weight complex in blue native-PAGE (∼1.0 MDa), corresponding to the total molecular weights of V1 and Vo domains. Together, we show that V-type ATPase is localized to multiple subcellular compartments in P. falciparum, and its functionality throughout the asexual cycle varies depending on the parasite developmental stages.

Keywords

malaria
Plasmodium
Plasmodium falciparum
V-type ATPase
proton pump
pH regulation
digestive vacuole
hemoglobin digestion
ring stage
trophozoite stage
schizont stage
Abbreviations

3HA triple hemagglutinin epitope

BCECF-AM 2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein, acetoxymethyl ester

BN-PAGE blue native-PAGE

DV digestive vacuole

gRNA guide RNA

HCv hemoglobin-containing vesicle

HR homologous region

IDC intraerythrocytic developmental cycle

IFA immunofluorescence assay

Immuno-EM immuno-electron microscopy

PfVP1 P. falciparum vacuolar pyrophosphatase 1

PPi pyrophosphate

PPM parasite plasma membrane

PVM parasitophorous vacuolar membrane

RBC red blood cell

TEM transmission electron microscopy

V-type ATPase vacuolar type ATPase

Reviewed by members of the JBC Editorial Board. Edited by Ronald Wek
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pmcMalaria remains one of the deadliest infectious diseases worldwide. In 2021, the World Health Organization reported 247,000,000 malaria cases and 619, 000 deaths (1). The causative agent of this disease is the protozoan parasite called Plasmodium, belonging to the phylum Apicomplexa. Among the five Plasmodium species infecting humans (Plasmodium falciparum, Plasmodium vivax, Plasmodium ovale, Plasmodium malariae, and Plasmodium knowlesi), P. falciparum is responsible for the majority of malaria cases and deaths worldwide. All clinical symptoms of malaria arise due to the repetitive growth of the parasites in red blood cells (RBCs); hence, the intraerythrocytic developmental cycle (IDC) or the asexual blood stage has long been recognized as a crucial target for developing antimalarial interventions.

Throughout the 48-h IDC of P. falciparum, parasites reside in a parasitophorous vacuole and contain many intracellular compartments or organelles, including the cytosol, the digestive vacuole (DV) for hemoglobin digestion, the nucleus/endoplasmic reticulum/Golgi apparatus, the mitochondrion, the apicoplast, and secretory organelles in the mature schizont-stage and merozoites. Some of these subcellular compartments are known to have distinct pH, which is the key for sustaining the biological processes inside them. For instance, the parasite cytosol maintains a static pH near 7.3 (2), whereas the DV requires a lower pH near 5.0 for proper hemoglobin digestion and heme detoxification (3, 4, 5, 6). The mitochondrion has a slightly basic pH (7.37 ± 0.09), whereas the apicoplast is nearly neutral (7.12 ± 0.40) (7).

The regulation of pH in distinct subcellular compartments involves multiple players, including buffers, enzymes, transporters, and proton pumps (8). Proton pumps are critical because they can directly transport protons against the concentration gradient, actively regulating subcellular pH (9). In P. falciparum, two types of proton pumps are present (10), including the pyrophosphate (PPi)-driven H+-pumping pyrophosphatases or vacuolar pyrophosphatases (11, 12, 13), and the ATP-driven vacuolar type ATPases (V-type ATPases) (14). VP1 (vacuolar pyrophosphatase 1) is an example of PPi-driven proton pumping pyrophosphatase. In P. falciparum, we have shown that P. falciparum vacuolar pyrophosphatase 1 (PfVP1) is mainly localized to the parasite plasma membrane (PPM), regulates cytosolic pH, and plays essential roles in the ring-stage and the ring-to-trophozoite transition (15). On the other hand, V-type ATPase is a large molecular weight ATP hydrolyzing proton pumping machinery (16), consisting of a V1 domain containing subunits of A-H and a Vo domain containing subunits of a, c, c’, c’’, d, and e. The cytosolic V1 domain holds the ATP hydrolysis hexamer made of A and B subunits, whereas the membrane bound Vo domain contains the H+ channel, which is involved in proton transport across the membrane. A unique feature of this machinery lies in the reversible assembly and disassembly of the V1Vo domains (17), regulating the pump’s function in response to various extracellular conditions (18). The P. falciparum genome encodes all V1 domain subunits and most Vo domain subunits that are present in other eukaryotes (www.PlasmoDB.org). The recent genome-wide mutagenesis study in P. falciparum (19) and the knockout survey in Plasmodium berghei (20) revealed that nearly all V-type ATPase subunits are essential in the asexual blood stage. Despite its significance, our understanding of the physiological role of V-type ATPase in malaria parasites remains rather limited.

In classical model organisms, including yeast, humans, and plants, V-type ATPase is localized to the membranes of multiple subcellular compartments such as endosomes, lysosomes, vacuoles, Golgi, and the plasma membrane (21, 22, 23). The acidification of these organelles by V-type ATPase is crucial to maintaining normal physiological functions in the cells. In marine phytoplankton such as diatoms, dinoflagellates, and coccolithophores, V-type ATPase is localized to chloroplast membranes and can significantly enhance carbon fixation and oxygen production (24), which are crucial processes for producing organic compounds and maintaining the atmospheric oxygen levels on earth. In P. falciparum, previous studies have suggested that V-type ATPase is a major proton pump responsible for maintaining the cytosolic pH (2) and DV pH (5) in trophozoite-stage parasites. However, its localization has remained elusive or somewhat controversial. An early study indicated that V-type ATPase has heterogenous but not well-defined localization (25). In another study, immunofluorescence assays (IFAs) and immuno-electron microscopy (immuno-EM) suggested that V-type ATPase is localized to the parasite periphery, DV, and small clear vesicles (26). However, the use of antibodies against the subunits of the bovine V-type ATPase in that study (26) raised concerns about antibody specificity toward the Plasmodium counterpart. Furthermore, another report revealed that V-type ATPase is also localized to the RBC membrane, apparently being exported to the RBC cytosol (27). Thus, studies of the P. falciparum V-type ATPase have not provided a consensus on its localization. Therefore, further investigation is needed to clarify the localization and function of this important proton pump in malaria parasites.

In this study, we employed multiple approaches to investigate the localization and functionality of V-type ATPase in P. falciparum throughout the asexual lifecycle. Our data suggest that V-type ATPase exhibits dynamic localization patterns that vary depending on the developmental stages of the parasite. Moreover, our results underscore the essentiality of V-type ATPase for trophozoite development and parasite invasion but likely not for ring-stage development.

Results

Endogenous tagging of V1A via CRISPR/Cas9 for localization and conditional knockdown

The highly conserved V-type ATPase has two domains, the V1 domain, responsible for hydrolyzing ATP, and the Vo domain, facilitating proton translocation using the free energy released from ATP hydrolysis. In P. falciparum, all the necessary subunits to form the V1 subunit are encoded, including subunits of A (PF3D7_1311900), B (PF3D7_0406100), C (PF3D7_0106100), D (PF3D7_1341900), E (PF3D7_0934500), F (PF3D7_1140100), G (PF3D7_1323200), and H (PF3D7_1306600). The Vo domain comprises subunits a (PF3D7_0806800), c (PF3D7_0519200), c’’ (PF3D7_1354400), and e (PF3D7_0721900). Subunits A and B together form the ATP hydrolyzing hexamer, powering the rotational movement of the entire proton pump. To study V-type ATPase in P. falciparum, we successfully tagged the endogenous locus of subunit A (V1A) at the C terminus with the triple hemagglutinin epitope (3HA) and elements required for conditional knockdown using the TetR-DOZI-aptamer system (28, 29). This was achieved via CRISPR/Cas9 (30, 31) with two different guide RNA (gRNA) plasmids in a WT P. falciparum line, NF54attB (32) (Fig. S1A). We termed the resulting transgenic parasite line NF54attB-V1A-3HAapt and performed limiting dilution to obtain pure parasite clones. The D11 clone was chosen for all subsequent experiments. The genotype of the NF54attB-V1A-3HAapt line was verified by diagnostic PCR (Fig. S1B). The expression of V1A-3HA proteins at the expected molecular weight was confirmed by Western blot (Fig. S1C). In addition to the full-length protein, a smaller band around 55 kDa was also detected, likely due to partial degradation of V1A-3HA proteins.

To verify the localization of V-type ATPase in P. falciparum over the 48-h lifecycle, we tightly synchronized the parasites with alanine/Hepes and performed IFAs. Starting from the early ring-stage (T0), samples were harvested and examined every few hours throughout the asexual cycle. We used PfExp2 (33), the marker of the parasitophorous vacuolar membrane (PVM), as the indicator of the parasite periphery. V1A was seen to be expressed in all asexual blood stages (Fig. 1A). In the ring-stage, V1A was localized to the parasite periphery, exhibiting partial colocalization with Exp2. In the trophozoite-stage, V1A was predominantly localized to both the PPM and the DV. This localization pattern was also observed in the early schizont-stage. To study mature schizont-stage parasites, we performed IFA in parasites that were pretreated with a reversible protein kinase G (pKG) inhibitor, ML10 (34). Treatment of ML10 inhibits parasite egress, leading to an increased number of mature schizont-stage parasites. In the ML10-treated schizonts, V1A’s localization appeared punctate, lacking confined signals showing circular or membranous structures (Fig. 1B). To further verify this observation at a higher resolution, we performed immuno-EM on the ML10-treated samples. Indeed, immuno-EM revealed a specific localization of V1A on rhoptries (Fig. 2, black arrows) and some small vesicles near them (Fig. 2, red arrows), indicating that V-type ATPase is also present on the secretory organelles in the merozoites. Further, quantification of 55 immuno-EM images from two independent experiments revealed the distribution of gold particles on various subcellular structures, including secretory organelles (56.12 ± 8.22%), rhoptries (17.35  ± 4.46%), PPM (15.47 ± 2.72%), and cytosol (11.06 ± 1.05%). Altogether, we have used IFA and immuno-EM to uncover the dynamic localization of V-type ATPase in different asexual blood stages of P. falciparum. The dynamic localization of V-type ATPase may be associated with its multifunctionality throughout the 48-h lifecycle.Figure 1 Localization of V1A throughout the asexual blood stages.A, immunofluorescence of the V1A protein in the ring, trophozoite, and schizont stages. In the NF54attB-V1A-3HAapt line, parasites were tightly synchronized and fixed every few hours starting from the early ring-stage (T0). V1A was detected using anti-HA and FITC-labeled secondary antibodies. Exp2, a marker for the parasitophorous vacuolar membrane (PVM), was detected using anti-Exp2 (52) and tetramethylrhodamine-labeled secondary antibodies. B, immunofluorescence of V1A in mature schizont-stage parasites. The synchronized late trophozoite-stage parasites were treated with 25 nM of ML10 for 14 h to reach the mature schizont-stage (34). V1A was detected by anti HA and FITC-labeled secondary antibodies. In A-B, DNA was stained with 4′,6-diamidino-2-phenylindole. 3HA, triple hemagglutinin epitope.

Figure 2 Localization of V1A on secretory organelles in mature schizont-stage parasites.A–F, immuno-EM images of NF54attB-V1A-3HAapt in the mature schizont-stage. The synchronized late trophozoite-stage parasites were treated with 25 nM of ML10 for 14 h to reach the mature schizont-stage (34). Parasites were then fixed and subjected to immuno-EM analysis. V1A signals were detected on rhoptries (black arrows) and secretory organelles near rhoptries (red arrows). G and H, negative control images with the primary antibody omitted. Bars in A–H, 500 nm. 3HA, triple hemagglutinin epitope; Dv, digestive vacuole; Immuno-EM, immuno-electron microscopy; N, nucleus; PPM, parasite plasma membrane; PVM, parasitophorous vacuolar membrane; R, rhoptries; RBCM, red blood cell membrane; sv, secretory organelles.

V-type ATPase is essential for trophozoite development and hemoglobin digestion

To understand the essentiality of V-type ATPase in P. falciparum, we conducted knockdown studies using two different methods. In the first method, we synchronized the parasites multiple times and initiated aTc removal in the early ring-stage (Fig. 3A). The knockdown parasites appeared healthy at 24 h post aTc removal but failed to produce any new ring-stage parasites in the subsequent 24 h. By 48 h of knockdown, the parasites were arrested in the trophozoite-stage and eventually succumbed to lysis. Therefore, V1A knockdown from the early ring-stage resulted in trophozoite arrestment and parasite demise within one cycle. In the second method, aTc removal began with synchronized trophozoite-stage parasites (Fig. 3D). The knockdown culture was able to produce new ring-stage parasites, but these parasites were arrested again in the trophozoite-stage at 48 h post aTc removal. The arrested parasites did not initiate another IDC but remained arrested or lysed. In additon to morphological examinations, Figure 3, B and E showed the quantification of parasite growth in the aTc (±) conditions in each knockdown experiment. Figure 3, C and F presented the V1A protein levels via Western blot. The band intensities were analyzed by ImageJ (https://imagej.net/ij/download.html) and shown in Fig. S2. The V1A protein level was observed to decrease by ∼ 80% at 24 h post knockdown and by >95% at 48 h post knockdown. Thus, we revealed that V-type ATPase is essential for parasite maturation and replication.Figure 3 Essentiality of V-type ATPase in the asexual blood stages. Giemsa-stained images showing parasite morphologies resulting from V1A knockdown initiated from the ring-stage (A) or the trophozoite-stage (D). Quantification of growth index, the product of parasitemia and splitting factors, in the aTc (±) cultures of the knockdown experiment initiated from the ring-stage (B), or the trophozoite sage (E). At each time point, parasitemia was determined from microscopic counting. Mean ± SD of growth index from three biological replications are shown. Western blot showing the V1A protein levels in the knockdown experiment initiated from the ring-stage (C) or the trophozoite-sage (F). The same blots were reprobed with anti-Exp2 to show loading controls. G, Giemsa-stained images showing parasite morphological changes over the detailed time-course experiment. Experiments of A–F were repeated three times. Panel G was repeated two times. The scale bars represents 5 μm. V-type ATPase, vacuolar type ATPase.

To better understand parasite developmental blockage upon V1A knockdown, we observed the morphological changes of the parasites in a detailed time-course experiment (Fig. 3G). In tightly synchronized early ring-stage parasites, aTc removal was initiated, and thin blood smears were collected every 8 h during the ring-stage and every 4 h during the trophozoite and schizont stages. Unlike the aTc (+) parasites that completed their development normally, the knockdown parasites progressed through the ring-stage but were unable to proceed beyond the mid-trophozoite stage. They were arrested as early as 32 h post aTc removal and failed to exhibit any growth progression in the remaining hours of the IDC, resulting in nearly 100% growth inhibition (Fig. S3). Altogether, we have shown that knockdown of V-type ATPase led to the arrest of parasite development in the trophozoite-stage within one IDC. Overall, our results confirm that V-type ATPase is essential for P. falciparum, with the trophozoite-stage being the most vulnerable to the loss of this proton pump.

V-type ATPase has been suggested to regulate the pH of DV in trophozoite-stage parasites (3, 4, 5). A low pH (∼5.0) is essential for hemoglobin digestion, which is extremely important for malaria parasites to obtain nutrients and gain space to grow inside RBCs (35). To get a better understanding of the knockdown effects on the DV, we performed transmission electron microscopy (TEM) studies. The knockdown experiment was initiated from Percoll-enriched schizont-stage parasites, and parasites were enriched by a magnetic column at 36 h post aTc removal. The parasitemia of both aTc (±) cultures reached nearly 100%. In agreement with Figure 3, the aTc (−) culture displayed noticeable abnormalities compared to the control (Fig. 4A). Under TEM, the knockdown parasites exhibited striking features (Fig. 4B). Their DV showed significantly higher electron density than the control and contained many hemoglobin positive vesicles. Hemoglobin uptake is initiated from a specialized structure called cytostome (36), and hemoglobin-containing vesicles (HCvs) are pinched off from the membranes and fused with the DV to release their contents for proper digestion and heme detoxification. The HCvs are thus surrounded by two membranes, the PPM on the outside and the PVM on the inside (37). In the knockdown parasites, the HCvs inside the DV were apparently surrounded by a single membrane (Fig. 4B, red arrows). This indicated that fusion of HCVs’ outer membrane with the DV was likely not affected upon depletion of V-type ATPase, but degradation of the inner membrane of HCvs was blocked. To date, it remains entirely unknown how the HCv’s inner membrane degrades inside the DV. Our data suggests that this process is extremely pH-dependent. Interestingly, we also observed that some vesicles in the DV of the knockdown parasites had much reduced electron density, likely due to partial hemoglobin digestion occurring before the DV pH increased to a malfunctioning level (Fig. 4B, blue arrow). Overall, our results show that V-type ATPase is critical for hemoglobin release and digestion in the DV.Figure 4 V-type ATPase is essential for hemoglobin digestion in the digestive vacuole.A, Giemsa-stained images of control and V1A knockdown parasites. aTc removal was initiated from schizont-stage parasites, and parasites were enriched by magnetic columns at 36 h post aTc removal. The scale bar represents 5 μm. B, Transmission electron microscopic images of control and V1A knockdown parasites. In the aTc (−) parasites, many vesicles inside the digestive vacuole had electron dense signals and were surrounded by a single membrane (red arrows). Some vesicles had translucent appearance (blue arrow). The scale bar represents 500 nm. V-type ATPase, vacuolar type ATPase.

Apart from its role in the DV, V-type ATPase is known to regulate cytosolic pH by pumping protons across the PPM in P. falciparum (2). This understanding was based on the observation that bafilomycin A1, an inhibitor of V-type ATPase, caused a dramatic decline of cytosolic pH (2). A recent study suggests that bafilomycin A1 causes a stearic hinderance and blocks the c-ring rotation by preventing the interaction between subunit a and the c-ring (38). The effect of V-type ATPase on cytosolic pH, however, has not been confirmed by genetic studies in Plasmodium. To address this, we utilized the well-established pH measurement technique with 2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein, acetoxymethyl ester (BCECF-AM) in saponin-treated trophozoites (2). We quantified cytosolic pH of aTc (±) cultures at 36 h post aTc removal initiated from Percoll-enriched schizont-stage parasites. We included bafilomycin A1 as a positive control. Our assays revealed that cytosolic pH was 7.24 ± 0.08 in the control parasites but decreased to 6.01 ± 0.28 upon knockdown of V-type ATPase (Fig. 5). This result confirmed that V-type ATPase is the major proton pump for regulating cytosolic pH in trophozoite-stage parasites. Being consistent with the published data (2), we also observed that bafilomycin A1 caused a profound decrease of cytosolic pH from 7.24 to 6.51 ± 0.1 within 5 min of incubation (Fig. 5). Interestingly, the combination of V1A knockdown and bafilomycin A1 resulted in an even lower level of cytosolic pH (5.77 ± 0.15), although this effect was not statistically significant compared to knockdown alone (p > 0.05). Overall, our findings confirm that V-type ATPase plays a major role in regulating cytosolic pH in the trophozoite-stage parasites.Figure 5 V-type ATPase regulates cytosolic pH in trophozoite-stage parasites. At 36 h post knockdown, aTc (±) cultures of the NF54attB-V1A-3HAapt line were subjected to pH measurement using the BCECF-AM method in the absence or presence of bafilomycin A1 (100 nM) for 5 min. pH measurements were conducted in a spectrofluorometer (Hitachi F-7000). Error bars indicated the SD of 3 to 4 measurements in each condition. Statistical analysis was done by student t test (unpaired, two-tailed). This experiment was repeated two times. V-type ATPase, vacuolar type ATPase; BCECF-AM, 2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein, acetoxymethyl ester.

Our data thus far have shown that knockdown of V-type ATPase led to the arrest of trophozoite-stage parasites that contained large amounts of undigested hemoglobin in the DV and reduced cytosolic pH. To understand the viability of the arrested “trophozoite-looking” parasites, we tried to rescue them by adding back aTc, which is a convenient way to restore the expression of V1A proteins in the knockdown culture. We initiated knockdown of V1A from tightly synchronized early ring-stage parasites and performed addback at different time points post aTc removal (28 h, 36 h, 40 h, and 48 h) (Fig. S4). In agreement with Figure 3G, the knockdown parasites at 28 h did not display obvious morphological defects in Giemsa-stained smears. Upon aTc addback, they grew normally as expected. At 36 h post aTc removal, the knockdown parasites displayed severe growth arrestment, showing minimal expansion in their size compared to the control. However, upon aTc addback, they were able to restore the progression of the asexual cycle in the next 48 h. This result suggested that the arrested parasites at 36 h post knockdown remained largely viable, despite having severe defects in hemoglobin digestion (Fig. 4) and pH regulation (Fig. 5). At 40 h post knockdown, some arrested parasites were still rescued by aTc addback. However, a few hours later at 48 h post knockdown, aTc addback failed to rescue any parasite growth. The knockdown parasites either displayed abnormal morphologies or lysed. Altogether, these results imply that P. falciparum parasites can sustain viability in a severely stressed condition for a period of several hours.

Bafilomycin A1 likely affects parasite invasion

The localization of V-type ATPase on rhoptries and other secretory organelles prompted us to further investigate the potential role of this proton pump in parasite egress and invasion. However, technical difficulties arose when using V1A knockdown parasites for this purpose. In the knockdown studies initiated from the ring, trophozoite, or schizont stages (Figure 3, Figure 4, Figure 5), an extended period of ∼30 h was required before phenotypic defects could be detected. We reasoned that a pool of pre-existing V1A proteins, present before aTc removal, permitted parasite survival and maturation over an extended period until the proteins were fully exhausted. Additionally, all knockdown approaches resulted in the arrest of trophozoites, consistently proving that the trophozoite-stage is the most vulnerable, but it rendered examination of schizonts technically challenging. To circumvent these issues, we utilized bafilomycin A1 to rapidly block V-type ATPase in the late schizont-stage parasites and observed its effect on parasite egress and invasion. As shown above, bafilomycin A1 is a rapid V-type ATPase inhibitor, reducing cytosolic pH from ∼7.3 to ∼6.5 in a few minutes’ incubation (Fig. 5) and (2). We also performed a 72-h SYBR Green assay and determined bafilomycin A1’s efficacy in parasite culture (EC50, 8.1 ± 0.5 nM).

Next, we tightly synchronized the parasites with alanine/Hepes and used ML10 to block egress, thereby increasing the percentage of mature schizonts in the culture. At the mature schizont-stage (∼0 h), we added bafilomycin A1 (100 nM, ∼10xEC50) for 2 h, washed it out, and allowed parasites to invade new RBCs for 4 h. At 6 h post the initiation of the experiment, we counted the parasitemia of newly formed ring-stage parasites and used it as a rough readout for estimating successful egress and invasion (Fig. 6A). Parasite growth and morphology were also examined at 24 h and 48 h post the initiation of the experiment. At 6 h, while the control culture had a 4.77% ± 0.25% of new ring-stage parasites, the bafilomycin A1–treated culture had a significantly reduced parasitemia (0.24% ± 0.07%) (Fig. 6B). In both the treated and control cultures, mature schizonts were rarely present at this point, and merozoites contacting RBCs were readily observed (Fig. 6C). In the next 24 h and 48 h, the control culture followed the normal progression of the lifecycle. In contrast, parasitemia of the treated culture barely increased (Fig. 6B). The parasites failed to invade RBCs, and “merozoite-like” parasites appeared to stick to the RBC surface even at 48 h post the initiation of the experiment (Fig. 6C). Quantification of parasite morphologies of Figure 6C was shown in Fig. S5. Collectively, these results indicated that bafilomycin A1 treatment leads to a severe blockage of parasite invasion (see Discussion).Figure 6 Effects of bafilomycin A1 on parasite egress and invasion.A, schematic illustration of the experimental design (made with BioRender). Mature schizont-stage parasites (NF54attB-V1A-3HAapt) were treated with bafilomycin A1 (100 nM) for 2 h. The drug was washed out and parasites were allowed to invade new RBCs for 4 h. At 6 h, the parasitemia of newly formed ring-stage parasites was monitored. Parasite morphology and number were further examined at 24 h and 48 h posttreatment. B, quantification of parasite growth throughout the experiment. Parasitemia was determined by microscopic counting. Mean ± SD of five biological replicates are shown. Note: total parasitemia was presented at 24 h and 48 h posttreatment, whereas only new ring-stage parasites were counted at 6 h. C, Giemsa-stained images showing parasite morphologies in the control and treated cultures. The scale bar represents 5 μm. A–C, these experiments were repeated five times. RBC, red blood cell.

V-type ATPase is not critical for ring-stage development

So far, we have demonstrated the critical role of V-type ATPase in the trophozoite-stage (Figure 3, Figure 4, Figure 5) and its potential involvement in parasite invasion (Fig. 6). To understand the significance of V-type ATPase for ring-stage development, we conducted an aTc pulse-chase experiment. Briefly, on day 0, we started the knockdown experiment in tightly synchronized mid-trophozoite stage parasites by removing aTc. The cultures were separated into four aliquots and maintained under aTc (+) (one aliquot) or aTc (−) (three aliquots) conditions. On day 1, aTc was added back to two aliquots of aTc (−) cultures for 2 h and 4 h, respectively, and was thoroughly washed out. The aTc pulse-chase cultures were then maintained in aTc (−) media for the rest of the experiment. On days 1 to 4, protein samples were harvested from each condition and subjected to Western blot (Fig. 7, A and B). Parasite morphologies and quantities were examined in Giemsa-stained thin blood smears (Fig. 7, A and C). On day 1 (24 h post knockdown), the aTc pulse-chase cultures significantly restored the V1A expression level. Parasites of all four conditions were in the ring-stage, and no abnormality was observed. On day 2, while the aTc (+) culture was in the trophozoite-stage, the aTc (−) culture was arrested as expected. However, the pulse-chase cultures had healthy looking trophozoites, indicating that a short aTc addback on day 1 facilitated the parasites to fulfill trophozoite-stage development without showing discernible defects on day 2. On day 3, while the aTc (+) culture initiated a new ring stage, the aTc (−) culture died or lysed as expected. However, the pulse-chase cultures had healthy looking ring-stage parasites, and their V1A proteins were barely detectable. This result indicated that V-type ATPase did not play a major role in the ring-stage because parasites were healthy without V1A expression in this stage. On day 4, the pulse-chase cultures were again arrested in the trophozoite-stage due to the lack of V1A proteins. Altogether, this experiment not only confirmed the essentiality of V-type ATPase for trophozoite-stage development but also suggested that this proton pump is likely not essential for the ring-stage.Figure 7 V-type ATPase is not critical for ring-stage development.A, V1A protein levels and parasite morphologies of the aTc pulse-chase experiment. On each day, protein samples were collected from all conditions, including aTc (+), aTc (−), aTc (−)/2 h (2 h of aTc addback and washout on day 1), and aTc (−)/4 h (4 h of aTc addback and washout on day 1), and analyzed by Western blot. V1A was detected by anti-HA antibody. The same blots were reprobed with Exp2 antibody to show loading controls. Parasite morphologies of each condition from days 1 to 4 were shown beneath the blots. The scale bar represents 5 μm. B, quantification of band intensities in A with ImageJ. In each condition on each day, the band intensity of V1A was adjusted according to the Exp2 loading control, and the adjusted band intensity was normalized to that of the aTc (+) lane to achieve a ratio. The normalized ratios were presented. C, parasitemia of the aTc pulse-chase experiment. In each condition on each day, parasitemia was determined by microscopic counting. Growth index is the product of parasitemia and splitting factors. A–C, these experiments were repeated four times. V-type ATPase, vacuolar type ATPase.

To further verify the role of V-type ATPase for ring-stage development, we took advantage of bafilomycin A1 and assessed its toxicity in the ring versus trophozoite stages. In highly synchronized ring-stage versus trophozoite-stage parasites, we treated them with 10 nM (∼1xEC50) or 100 nM (∼10xEC50) of bafilomycin A1 for 2 h and 4 h, washed out the drug, and examined parasitemia and parasite morphologies at 24 h and 48 h posttreatment. Starting in the ring-stage, treatment of bafilomycin A1 in all conditions (10 nM for 2 h, 100 nM for 2 h, 10 nM for 4 h, and 100 nM for 4 h) did not cause a significant decline in parasitemia at 24 h or 48 h later (Fig. 8A). This indicated that ring-stage parasites were not susceptible to bafilomycin A1. In contrast, in the trophozoite-stage, treatment of bafilomycin A1 in all conditions led to a significant decrease in parasitemia at 24 h and 48 h (Fig. 8B). Parasite morphologies post the treatment were shown in Giemsa-stained thin blood smears (Fig. S6). Starting in the ring-stage, the treated parasites in all conditions progressed normally. In contrast, right after bafilomycin A1 treatment in the trophozoite-stage, the parasites exhibited discernible growth defects as their sizes were much smaller than those of control parasites. The treated trophozoites also produced fewer ring-stage parasites in the following 24 h. Altogether, the insusceptibility of ring-stage parasites to bafilomycin A1 suggests that the V-type ATPase is likely not essential for ring-stage development.Figure 8 Toxicity of bafilomycin A1 in the ring-stage versus the trophozoite-stage. Parasitemia of the cultures at 24 h or 48 h posttreatment of bafilomycin (10 nM, 100 nM) for 2 h or 4 h initiated from the ring-stage (A) or the trophozoite-stage (B). In each condition, parasitemia was determined by counting 1000 RBCs under a light microscope. Statistical analysis was done by one-way ANOVA. This experiment was repeated two times. RBC, red blood cell.

Detection of the V-type ATPase complex by blue native-PAGE/Western blot

Finally, we employed blue native electrophoresis and Western Blot to estimate the molecular weight of the P. falciparum V-type ATPase. In mammalian cells, the complex has been detected by blue native-PAGE (BN-PAGE) and its molecular weight is ∼1.0 MDa (39). The estimated molecular mass of the yeast V-type ATPase is also 1.0 MDa (40). To access the native conformation of the P. falciparum V-type ATPase, we set up knockdown experiments in the early ring-stage and collected parasite pellets at 32 h post aTc removal. The samples were solubilized in a mild detergent, 1% glyco-diosgenin, and the supernatants were separated on BN-PAGE followed by Western blot. In the aTc (+) sample, we observed a band near the molecular marker of 1026 kDa, suggesting that it is the intact V-type ATPase complex in P. falciparum (Fig. 9A). Unexpectedly, we also observed multiple bands around the molecular markers of 720 kDa, 480 kDa, and 240 kDa (Fig. 9A). The identities of these bands are unknown. But all bands detected in the aTc (+) sample were specific to V1A, as their intensities were greatly diminished in the knockdown sample. Along with BN-PAGE, we also subjected aTc (±) parasite samples in regular SDS/PAGE and Western blot. As expected, the level of V1A was markedly reduced in the knockdown sample, whereas the control protein, EXP2, did not show significant changes (Fig. 9B). These results implied that the V-type ATPase forms a large molecular weight complex in P. falciparum, and formation of the complex likely requires a dynamic and complex process.Figure 9 Detection of the V-type ATPase complex in BN/PAGE.A, V-type ATPase forms a large molecular weight complex near the marker of 1026 kDa. The knockdown experiment began in the early ring-stage. Samples of aTc (±) cultures were collected at 32 h post aTc removal and analyzed by BN-PAGE. V1A was detected by anti-HA antibody. The black arrow indicates the potentially intact V1Vo complex. Blue arrows indicate bands with unknown identities. B, confirmation of V1A knockdown at 32 h post aTc removal via SDS-PAGE/Western blot. V1A was detected by anti-HA antibody. Exp2 shows the loading control. A and B, these experiments were repeated two times. BN-PAGE, blue native-PAGE; V-type ATPase, vacuolar type ATPase.

Discussion

In this study, we have characterized the essentiality, localization, and biological functions of V-type ATPase in P. falciparum throughout the asexual blood stage. By tagging subunit A of the V1 domain via CRISPR/Cas9, we generated a conditional knockdown parasite line under aTc regulation and obtained several interesting results.

Firstly, we found that localization of the P. falciparum V-type ATPase is highly dynamic throughout the 48-h asexual lifecycle (Figs. 1 and 2). In the trophozoite-stage, V-type ATPase localizes to the PPM and DV, agreeing with its expected roles in regulating pH of the cytosol (2) and the DV (5). When the parasite progresses from the trophozoite-stage to the early schizont-stage, the V-type ATPase retains its localization on the membranes of PPM and DV. However, in the late schizont-stage, distribution of V1A becomes punctate, with no clear membranous patterns (Fig. 1B). Immuno-EM further revealed V1A’s localization on rhoptries and nearby secretory organelles (Fig. 2), indicating the presence of the V-type ATPase on merozoites’ secretary organelles. To our best knowledge, this is the first report showing localization of V-type ATPase on mature secretory organelles in Apicomplexa, the phylum to which Plasmodium belongs. A previous study highlighted the function of V-type ATPase in facilitating the maturation of secretory organelles in Toxoplasma gondii, an apicomplexan parasite distinctly related to Plasmodium (41). However, the presence of V-type ATPase on mature rhoptries or micronemes was not detected in T. gondii (41). Thus, the localization of V-type ATPase on rhoptries and secretory organelles in Plasmodium is a unique feature. Moreover, our data suggest that V-type ATPase is also expressed and localized to the PPM of the ring-stage parasite, but not to the plasma membrane of RBCs.

Secondly, among the three consecutive asexual blood stages, we uncovered that the trophozoite-stage is the most vulnerable to the loss of this proton pump (Figure 3, Figure 4, Figure 5). Our knockdown experiments show that trophozoite development is inevitably arrested regardless of when aTc removal is initiated from the 48-h asexual lifecycle. This is because V-type ATPase plays a plethora of essential functions in trophozoites including regulating cytosolic pH (Fig. 5) and facilitating hemoglobin digestion in the DV. In the fast-growing trophozoite, acidification of the cytosol upon the loss of V-type ATPase would shut down numerous biochemical reactions occurring in both the cytosol and the nucleus, leading to severe parasite growth arrest. Further, hemoglobin maldigestion in the DV would render the parasite lacking sufficient nutrients and space for rapid growth. We were astonished to observe undigested hemoglobin in the DV of the V1A knockdown parasites (Fig. 4). To date, the processes of hemoglobin uptake from cytostomes and the delivery of HCvs to the DV remain enigmatic in P. falciparum, as these processes are highly dynamic, transient, and difficult to capture. A previous study detected one fusion event of HCv and the DV membrane among >600 thin parasite sections examined under TEM (42), suggesting that the outer membrane of HCvs fuses with the DV membrane. Pinching off from the cytostome, the HCvs are enclosed by the PPM on the outside (outer membrane) and the PVM on the inside (inner membrane). Our data suggests that the hemoglobin-containing structures in the DV of V1A knockdown parasites contain one membrane, likely the inner membrane of HCvs (Fig. 4). Therefore, upon the loss of V-type ATPase, fusion of the HCv’s outer membrane (originated from PPM) was not affected but cleavage of the HCv’s inner membrane (originated from PVM) was blocked. The enzyme(s) responsible for digesting the PVM on HCvs remains entirely unknown at present, but it is evidently pH-dependent. Further, since PVM is a lipid bilayer, a lipase is likely evolved in this process. However, identification of such lipase(s) might be a challenging task. A recent study identified only one out of 19 phospholipases is essential for asexual stage development (43). That essential phospholipase, PI-PLC (PF3D7_1013500), is not likely responsible for cleaving HCvs’ inner membrane because it is localized to the cytosol, not to the DV (43). Interestingly, knockdown of V-type ATPase blocked hemoglobin digestion in the DV, but the upstream trafficking of HCvs enroute to the DV was seemingly not affected because no HCvs were accumulating in the parasite cytosol. This differed from a previous study that showed accumulation of HCvs inside the parasite cytosol upon conditional inactivation of PfVPS45 (44), a protein involved in vesicular trafficking.

Thirdly, our immuno-EM data (Fig. 2) prompted us to explore the function of V-type ATPases in the late schizont-stage. We identified a potentially essential role of V-type ATPase for parasite invasion. To alleviate the slow effect of genetic knockdown, we used bafilomycin A1 to chemically inhibit V-type ATPase in a short time (2 h) in mature schizonts and quantified newly formed ring-stage parasites post the treatment (Fig. 6). Bafilomycin A1’s specificity as a V-type ATPase blocker has been well documented and it has been used as a standard reagent in P. falciparum research (2, 5). We also confirmed that bafilomycin A1 executes its effects very rapidly (Fig. 5). Using a short treatment of bafilomycin A1 in mature schizonts, we observed a drastic decline in the parasite’s ability to form new ring-stage parasites (Fig. 6). The 6-h window of our experiment covers a series of highly complicated processes from the end phase of schizogony, egress, to invasion. We admit that these processes are rapid and challenging to study. We believe that V-type ATPase is essential for parasite invasion because at 6 h post the initiation of the experiment, the treated culture had no mature schizonts anymore, and merozoites were readily detectable and appeared to be attached to RBC surfaces. An obvious limitation of this study lies in our qualitative estimation of bafilomycin A1’s effects on these highly dynamic biological processes occurring from the end of schizogony to the initiation of a new ring-stage. Thus, our data so far have not firmly defined the role of V-type ATPase in parasite egress, invasion, or both. Future detailed investigations are required to better understand the roles of V-type ATPase in these highly dynamic biological processes.

We believe that V-type ATPase is not critical for ring-stage development. In our regular knockdown experiments initiated from the trophozoite-stage, we consistently observed healthy ring-stage parasites 24 h later (Fig. 3D). This result indicated that V-type ATPase is not critical for ring-stage development. But the possibility that a small amount of protein present at 24 h post knockdown was sufficient to support ring-stage growth has not been ruled out. To address this issue, we developed an aTc pulse-chase experiment (Fig. 7). We started the knockdown experiment in the trophozoite-stage on day 0 and added aTc for a very short time (2 h and 4 h) on day 1 to help parasites fulfill development on day 2. On day 3, healthy ring-stage parasites were formed but V1A protein levels were undetectable. At this point, the small amounts of proteins added on day 1 should be fully exhausted. Yet, the ring-stage parasites appeared healthy and progressed to the next stage on day 4. Without V-type ATPase, parasites were again arrested in the trophozoite-stage on day 4. These results reinforced the essentiality of V-type ATPase for trophozoite-stage development but not for ring-stage development. To further validate this, we treated highly synchronized cultures in the ring-stage versus trophozoite-stage with bafilomycin A1 (Figs. 8 and S6). Treatment of bafilomycin A1 in the ring-stage at concentrations of 10 nM (1xEC50) or 100 nM (10xEC50) for 2 h or 4 h did not cause noticeable defects. In contrast, the same treatments in the trophozoite-stage parasites led to significant parasite growth arrestment and parasitemia decline. Together, using two approaches, we have shown that V-type ATPase is not essential for ring-stage development. We have recently discovered that the PPi-driven proton pump, PfVP1, is essential for the ring-stage (45). Thus, our data suggests that P. falciparum utilizes different proton pumping mechanisms at different stages of the asexual cycle. In the ring-stage, parasites prefer to use the PPi-driven proton pump (PfVP1) rather than the ATP consuming V-type ATPase, which could facilitate the immature parasite to adapt to its unique bioenergetic environment under a low level of glycolysis.

Lastly but not least, as we prepared this manuscript for publication (46), a report by Alder and colleagues have also unveiled the localization and function of the P. falciparum V-type ATPase using an inducible KO approach (47). They have conditionally knocked out subunits B, a, c, and discovered V-type ATPase’s role for acidifying the DV and controlling the DV’s morphogenesis. Interestingly, our findings from knockdown of V1A align well with those obtained from inducible knockout of V1B. Both we and Alder et al. revealed the essentiality of V-type ATPase for trophozoite-stage development and hemoglobin digestion. Given that subunits A and B work together to form the ATP hydrolysis hexamer in the V1 domain, it is not surprising that genetic knockdown (or knockout) of either V1A or V1B leads to common downstream defects. Therefore, we and Alder et al. have collectively confirmed the essentiality of V-type ATPase for asexual stage development. While Alder and colleagues primarily focused on the functions of V-type ATPase in the DV of trophozoites, our work has expanded the understanding of this proton pump in other asexual blood stages beyond the trophozoite-stage. We have found that V-type ATPase is likely essential for parasite invasion, but it is not critical for ring-stage development. We also show that V-type ATPase localizes to various subcellular compartments, and its localization is stage specific.

In summary, this study has yielded valuable insights into the localization and functions of V-type ATPase in P. falciparum, highlighting its critical roles in regulating pH in various subcellular compartments in different stages of the asexual developmental cycle. Moreover, results of our work and Alder et al. (47) could provide valuable guidance for drug development strategies since V-type ATPase was implicated as an antimalaria drug target (48, 49).

Experimental procedures

Plasmid construction

Subunit A of P. falciparum V-Type ATPase (V1A, PF3D7_1311900) was endogenously tagged with a 3xHA epitope and the TetR-DOZI-Aptamer system at the C terminus via CRISPR/Cas9. Two NFCas9/gRNA constructs were made according to our cloning procedures published earlier (50). The DNA oligos used for cloning and sequencing were listed in Table S1 (P1–P7). To create the pMG75 template plasmid for double crossover recombination, the 5′ and 3′ homologous regions (5′HR and 3′HR) of V1A were PCR amplified from WT genomic DNA using primers P8 and P9 for 5′HR and P10 and P11 for 3′HR. The PCR product was purified and then digested with SacII/BstEII for 5′HR and BssHII/SacII for 3′HR. The digested fragments were sequentially cloned into the pMG75 vector bearing blasticidin deaminase (BSD) by T4 DNA ligase. The clones were screened by proper restriction digestion. The positive clones were further confirmed by Sanger sequencing using primers P12 to P13. The final maxi-prep plasmid of pMG75-BSD-V1A-3HA was linearized with EcoRV before being transfected into parasites together with gRNA plasmids.

Parasite culture and transfection

P. falciparum WT strain NF54attB was generously provided by Dr Joshua Beck (Iowa State University) (32). This parasite line is compatible with all three transfection markers that are commonly used, including blasticidin-S deaminase (BSD), human dihydrofolate reductase (hDHFR), and yeast dihydroorotate dehydrogenase (yDHODH). P. falciparum was cultured in human O+ RBCs (5% hematocrit, Interstate Blood Bank, Inc) in the standard RPMI 1640 medium containing 0.5% AlbuMAX II (Invitrogen), 15 mM Hepes, 10 mg/L hypoxanthine, 25 mM NaHCO3, and 50 mg/L gentamicin. Transfections were performed using the standard methods in the ring-stage with details shown previously (50, 51). The concentration of aTc was 250 nM in all aTc (+) cultures. All cultures were free of mycoplasma examined by routine PCR testing.

Immunofluorescence assays

Parasites were tightly synchronized with alanine (0.5 M)/Hepes (10 mM) several times. At each time point of the experiment, ∼50 μl parasitized RBCs were collected and fixed with 4% paraformaldehyde/0.0075% glutaraldehyde in 1× PBS. Blocking was done by 3% bovine serum albumin/PBS for overnight at 4 °C. Fixed cells were permeabilized by Triton X-100 (0.25%)/PBS and incubated with primary antibodies overnight at 4 °C, including α-HA (1:300, mouse, sc-7392, Santa Cruz Biotechnology) and α-Exp2 (1:500, rabbit, Dr Burns, Drexel University) (52). Following thorough washing, the samples were incubated with fluorescein isothiocyanate- or tetramethylrhodamine-labeled secondary antibodies for 3 h at room temperature. Other procedures followed standard IFA protocols. Images were captured using the Nikon Ti microscope and analyzed using Nikon NIS elements software (https://www.microscope.healthcare.nikon.com/products/software/nis-elements).

Western Blot for BN-PAGE and SDS-PAGE

Parasite cultures were lysed by saponin (0.05%)/PBS and host hemoglobin was removed by several washes with 1× PBS. In each culture condition, two aliquots of samples were obtained for BN-PAGE and SDS-PAGE, respectively. For BN-PAGE, the saponin lysed pellet was solubilized with 1% glyco-diosgenin in MESH buffer (250 mM sucrose, 10 mM Hepes, 1 mM EDTA, 1× protease inhibitor cocktail) for overnight at 4 °C. The supernatant after a high-speed centrifugation (13,000 rpm, 10 min) was used for blue-native gel electrophoresis and Western blot. For SDS-PAGE, the saponin lysed pellet was solubilized by 2% SDS/Tris (65 mM, pH 6.8) for overnight at 4 °C. The supernatant after a high-speed centrifugation (13,000 rpm, 10 min) was used for denatured electrophoresis. For all samples, the protein concentrations were determined by the Pierce bicinchoninic acid Protein Assay Kit (Thermo Fisher Scientific) to ensure equal loading. Antibodies used for Western blot included α-HA (1:10,000, mouse, sc-7392, Santa Cruz Biotechnology), α-Exp2 (1:10,000, rabbit) (52), horseradish peroxidase-conjugated goat anti-mouse secondary (1:10,000, A16078, Thermo Fisher Scientific), and horseradish peroxidase-conjugated goat anti-rabbit secondary (1:10,000, 31,460, Thermo Fisher Scientific). The membranes were incubated with Pierce ECL Western Blot Substrates and developed using the Bio-Rad chemidoc imaging system.

Transmission electron microscopy

Parasites were synchronized with alanine (0.5 M)/Hepes (10 mM) several times and the schizont-stage parasites were enriched by Percoll. After several washes with 1× PBS, the schizonts were added with new RBCs and cultured in aTc (±) conditions for 36 h. Both cultures were then passed through a MACS magnetic column (MiltenyiBiotec) to enrich trophozoite-stage parasites, which were fixed with freshly made 2% paraformaldehyde/2.5% glutaraldehyde in 100 mM sodium cacodylate buffer for overnight at 4 °C. The samples were shipped to Dr Wandy Beatty from the Molecular Microbiology Imaging Facility at Washington University for further processing and imaging. Other procedures were carried out as shown previously (51).

Immuno-electron microscopy

Parasites were synchronized with alanine (0.5 M)/Hepes (10 mM) several times and the trophozoite-stage parasites were enriched using a MACS magnetic column (MiltenyiBiotec). The enriched parasites were treated with 25 nM ML10 (34) for 14 h and then fixed with freshly made 4% paraformaldehyde/0.05% glutaraldehyde/100 mM Pipes buffer for overnight at 4 °C. The samples were shipped to Dr Wandy Beatty from the Molecular Microbiology Imaging Facility at Washington University for further processing and imaging. Other procedures were carried out as previously published (50). Antibodies used in the study included α-HA (1:300, mouse, sc-7392, Santa Cruz Biotechnology) and goat anti-mouse IgG 18 nm colloidal gold-conjugated secondary antibody (1:100, Jackson ImmunoResearch Laboratories). A parallel sample with the primary antibody omitted was used as the negative control, which did not react to the secondary antibody.

pH measurement with BCECF-AM, a ratio-metric pH indicator

pH measurement was carried out according to the published protocols (2). Post 36 h of aTc removal, the control and knockdown cultures were incubated with 4 μM BCECF-AM (Thermo Fisher Scientific) for 30 min at 37 °C. Pluronic at 1:1000 dilution was added to increase cellular permeability to BCECF-AM. The parasites were treated with rapid saponin lysis to remove hemoglobin. The pellet was washed two times with saline/glucose buffer (NaCl 125 mM, KCl 5 mM, MgCl2 1 mM, glucose 20 mM, Hepes 25 mM, pH 7.4) and resuspended in 1 ml of warm saline/glucose buffer. The sample’s fluorescence was monitored using a spectrofluorometer (Hitachi F-7000) at 535 nm with excitation wavelengths set at 490/440 nm. When bafilomycin A1 (100 nM) was used in aTc (±) conditions, the drug was incubated with saponin-lysed and washed parasites for 5 min before they were further analyzed by Hitachi F-7000. The fluorescence ratios were back calculated to pH values according to a linear regression curve generated from several known pH standards (pH 6.8, 7.1, and 7.4), as shown previously (2).

Data availability

All data are presented in the main manuscript and Supporting information.

Supporting information

This article contains supporting information.

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.

Supporting information

Table S1

Supplemental figure

Figure S1

Figure S4

Figure S5

Figure S4

Figure S5

Figure S6

Acknowledgments

We are grateful for the technical assistance from members of the Center for Molecular Parasitology at Drexel University College of Medicine and intellectual input from Dr Akhil B. Vaidya. We thank Dr Joshua Beck (Iowa State University) for providing the parasite line, Drs Jaquin Niles (MIT) and Sean Prigge (Johns Hopkins University) for providing the TetR-DOZI-aptamer plasmids, Drs Baker and Ooij for providing the compound, ML10. We thank Dr Wandy Beatty at Washington University in St Louis for performing TEM and immuno-EM studies. We thank ChatGPT3.5 for correcting minor grammatic issues throughout the manuscript.

Author contributions

N. S., S. D., and H. K. conceptualization; N. S., S. D., and H. K. formal analysis; H. K. funding acquisition; N. S., S. D., W. X., L. W., and H. K. investigation; N. S., S. D., W. X., and H. K. methodology; H. K. project administration; N. S., S. D., W. X., and H. K. resources; H. K. supervision; N. S., S. D., W. X., and H. K. validation; N. S. and H. K. visualization; H. K. writing–original draft; N. S., S. D., W. X., L. W., and H. K. writing–review and editing.

Funding and additional information

This work was supported by grants (R21AI156735 and R01AI184855 ) from 10.13039/100000002 National Institutes of Health /10.13039/100000060 NIAID (National Institute of Allergy and Infectious Diseases) to H. K. S. D. was supported by R01AI028398 from 10.13039/100000002 NIH /10.13039/100000060 NIAID  to A. B. V. The funders have no roles in study design, data collection, data analysis, and manuscript preparation. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.
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