Apicomplexan parasites invade host cells by forming a ring-like junction with the cell surface and actively sliding through the junction inside an intracellular vacuole. Apical membrane antigen 1 is conserved in apicomplexans and a long-standing malaria vaccine candidate. It is considered to have multiple important roles during host cell penetration, primarily in structuring the junction by interacting with the rhoptry neck 2 protein and transducing the force generated by the parasite motor during internalization. Here, we generate Plasmodium sporozoites and merozoites and Toxoplasma tachyzoites lacking apical membrane antigen 1, and find that the latter two are impaired in host cell attachment but the three display normal host cell penetration through the junction. Therefore, apical membrane antigen 1, rather than an essential invasin, is a dispensable adhesin of apicomplexan zoites. These genetic data have implications on the use of apical membrane antigen 1 or the apical membrane antigen 1–rhoptry neck 2 interaction as targets of intervention strategies against malaria or other diseases caused by apicomplexans.
Most apicomplexans, including the agents of malaria (Plasmodium) and toxoplasmosis (Toxoplasma), are obligate intracellular parasites. Many invade host cells by a conserved mechanism involving the formation of a zone of tight attachment between the parasite apex and the host cell1, called tight junction (TJ). The current view is that the TJ is primarily a molecular bridge between the parasite sub-membrane actin-myosin motor and a stable and stationary2 anchor associated with the host cell surface/cortex, which allows parasite traction into a parasitophorous vacuole (PV) within the host cell3,4.
The transmembrane protein apical membrane antigen 1 (AMA1), arguably the most studied protein in the Apicomplexa phylum and a long-standing malaria vaccine candidate, is thought to shape the TJ on the parasite side5,6,7,8,9,10,11. The cytoplasmic tail of AMA1 was reported to bind aldolase8 in vitro, considered a signature of proteins that bind parasite actin and the motor12,13. The ectodomain of AMA1 tightly binds in parasite extracts to the rhoptry neck 2 (RON2) protein9,10, a protein secreted from the parasite rhoptries that specifically localizes at the TJ, where it inserts in the host cell membrane and is presumably linked to the cell cortical cytoskeleton via other RON proteins, like RON4. Moreover, antibodies or peptides that inhibit the AMA1–RON2 interaction drastically reduce host cell invasion by Plasmodium merozoites14,15,16 and Toxoplasma tachyzoites9,10. T. gondii or P. falciparum AMA1 bound to RON2 peptide were co-crystalized, revealing a conserved RON2 loop inserting deep into an AMA1 hydrophobic groove17,18. This reinforced the model of this interaction constituting the traction point used by the parasite to power the active internalization inside the PV19,20,21, and led to the proposal of developing broad-spectrum small-molecule inhibitors of apicomplexan invasion targeting the AMA1–RON2 interaction22,23,24. In addition, AMA1 has been reported to be involved in rhoptry secretion15,25 as well as for providing a signal initiating intracellular replication26.
Recently, P. berghei and T. gondii parasites in which AMA1 was silenced (AMA1 knockdown, AMA1KD) were found to remain competent for shaping a TJ and invading host cells27. However, as AMA1KD parasites might still express residual levels of AMA1, these data were not considered as challenging any of the proposed roles of AMA1 in invasion18,23,28,29. In agreement with an essential role of AMA1 at some stage of the parasite invasion process, all attempts to inactivate AMA1 in both Plasmodium30,31 and Toxoplasma32 have failed so far.
Here, we report the inactivation of AMA1 in the tachyzoite of T. gondii, which invades virtually any cell type in the host, and in the merozoite and sporozoite stages of P. berghei, which invade erythrocytes and hepatocytes, respectively. AMA1 was deleted from the parasite genome by the diCre-loxP recombination approach in T. gondii and by direct homologous recombination in P. berghei. All three AMA1 knockout (AMA1KO) zoites are still capable of penetrating the respective host cell like the wild type (WT). Tachyzoites and merozoites, however, display a defect in host cell binding. These genetic data indicate that AMA1 and the RON proteins act separately during apicomplexan invasion, and that the AMA1–RON2 interaction does not have an essential role at the TJ.
Role of AMA1 in T. gondii tachyzoite infection of host cells
To investigate the role of AMA1 in T. gondii tachyzoites, we generated AMA1KO parasites using the diCre-loxP site-specific recombination system33. The loxPAMA1loxP-YFP-HXGPRT construct (Fig. 1a) was inserted in the ku80::diCre strain, which encodes two inactive fragments of Cre fused to rapamycin-binding proteins. Upon rapamycin addition and Cre reconstitution, recombinant parasites excise AMA1 (Fig. 1b) and express YFP. An excised clone, called TgAMA1KO, which does not produce AMA1 as shown by western blot (Fig. 1c) and immunofluorescence (Fig. 2a) analysis, was selected and maintained using routine culture procedures. When measuring parasite infectivity by plaque assay, the plaque size is ~2 to 2.5 times smaller with TgAMA1KO than control parasites (Fig. 2a,b), a rather mild phenotype. Reintroduction of internally tagged AMA1FLAG in TgAMA1KO parasites fully restores overall growth, demonstrating the specificity of the observed phenotype (Fig. 2a). Importantly, no significant difference can be noticed in the replicative rates (Fig. 2c,d), egress efficiency (Fig. 2e) or motility patterns (Fig. 2f) between TgAMA1KO and control tachyzoites. Therefore, AMA1 is not involved in tachyzoite gliding, intravacuolar replication or egress from host cells.
We then investigated AMA1KO tachyzoite invasion of host cells in more detail. When measured by fluorescence microscopy, TgAMA1KO tachyzoite invasion efficiency is 30–40% that of the parental ku80::diCre strain (Fig. 3a). To investigate whether the pattern of TgAMA1KO tachyzoite invasion of host cells is normal or altered, TgAMA1KO tachyzoites invading human foreskin fibroblasts (HFFs) or normal rat kidney (NRK) fibroblasts were captured by confocal microscopy and analysed after three-dimensional reconstruction (Fig. 3b). Entering mutant zoites (n=53) constantly display a typical RON4 circular staining around the parasite constriction site after cell permeabilization, indicating normal rhoptry secretion and TJ formation during invasion. In line with the normal gliding capacity of TgAMA1KO tachyzoites, micronemal protein 2 (MIC2)34, which is secreted like AMA1 from the microneme organelles, is normally exposed on the surface of invading TgAMA1KO tachyzoites (Fig. 3b). Moreover, video microscopy of invading TgAMA1KO zoites shows that successful invasion follows a one go and smooth process with similar kinetics as controls (Fig. 3c,d and Supplementary Movies 1–3), and in all cases (n=20) a clear constriction site, suggestive of normal TJ, is observed. We conclude that in the tachyzoite AMA1 is not necessary for structuring a fully functional TJ, in which the RON proteins act independently of AMA1.
Next, we assessed tachyzoite adhesion to host cells. After 15 or 60 min incubation with live HFF cell monolayers, approximately two- to threefold fewer mutant versus control tachyzoites associate with host cells (total: extracellular attached and internalized combined), whereas three- to fourfold fewer mutants are located inside host cells, suggesting a primary defect of mutant zoites in host cell attachment (Fig. 3a). Importantly, about a third of the TgAMA1KO tachyzoite population adopts a distinct position relative to the host cell than controls, by binding only via the apical end rather than throughout their length (Fig. 3e), like previously observed for AMA1KD tachyzoites27. This confirms that AMA1 has an important role in tachyzoite adhesion to/positioning onto host cells before TJ formation, an event that favours, but is not required for, host cell invasion.
Role of AMA1 in P. berghei merozoite infection of erythrocytes
To inactivate AMA1 in P. berghei, WT ANKA blood stages were transfected with a construct designed to replace endogenous AMA1 by pyrimethamine-resistance and green-fluorescence cassettes (Fig. 4a). Dedicated in vivo selection protocols with several days of drug pressure reproducibly generated mixtures of targeted green fluorescent protein (GFP+) AMA1− parasites, that is, AMA1KO, and non-targeted GFP− AMA1+ parasites, presumably spontaneous pyrimethamine-resistant mutants that typically emerge after long selection times. Southern blot analysis indicates the presence in the selected population of both the WT AMA1 locus and the expected allelic replacement (Fig. 4b). In agreement with this, immunofluorescence assays reveal erythrocytes infected by either GFP+ AMA1− or GFP− AMA1+ parasites (Fig. 4c). The multiplication rate of AMA1KO parasites, assessed by co-injection with control red fluorescent protein (RFP+) parasites35 in mice and monitoring parasite multiplication by fluorescence-activated cell sorting (FACS), is ~35% that of RFP+ parasites (Fig. 4d). As internalized AMA1KO parasites generate normal numbers of progeny merozoites after a normal developmental cycle (Fig. 4e), that is, AMA1 is not important for merozoite replication inside erythrocytes, the decreased multiplication rate of AMA1KO parasites reflects a defect in merozoite entry into erythrocytes.
We next characterized interactions between AMA1KO merozoites and erythrocytes using imaging flow cytometry (IFC), which combines microscopy and flow cytometry and provides quantitative and functional information using imaging algorithms. Briefly (see Methods), after mixing mouse erythrocytes pre-stained with the lipid dye PKH26 with P. berghei GFP+ merozoites36 collected from synchronized schizont cultures, parasites interacting with a host cell are identified as GFP signals in a gated PKH26+ population (Fig. 5a, left panel), and internalized parasites are further recognized by co-localization of GFP with an increased PKH26 signal relative to the rest of the cell (Fig. 5b), a labelling suggestive of merozoites surrounded by a tight-fitting vacuole membrane37 (Fig. 5a, right panel). Using control GFP+ merozoites incubated for 10 min with PKH26-stained erythrocytes before fixation, ~43.8% score as ‘associated’ with erythrocytes (EryA; Fig. 5a, left panel), whereas ~3.8% score as ‘internalized’ inside erythrocytes (EryI; Fig. 5a, right panel). Importantly, cytochalasin D, which prevents merozoite internalization but not attachment to erythrocytes38, does not significantly affect the EryA but drastically reduces the EryI population (Fig. 5a,c), which validates the EryI population algorithm. Using merozoites of the AMA1KO-containing population, ~8.6% of the GFP+ AMA1KO merozoites score as EryA and ~0.48% as EryI (Fig. 5d). A similar reduction relative to control merozoites is obtained when samples are fixed after 3 min incubation (Fig. 5e), indicating a primary defect in adhesion of AMA1KO merozoites. Like AMA1KO tachyzoites, AMA1KO merozoites form a normal RON (RON2) ring during host cell invasion (Fig. 4f).
As additional mutations, compensatory or adverse, might accumulate in the AMA1KO parasites propagated for extended times (up to 30 days) before IFC analysis, we next characterized AMA1KD merozoites generated by Flippase (Flp)/Flp Recombination Target (FRT)-mediated recombination27 immediately before IFC (Fig. 6a). In this approach, AMA1KD mosquito-stage sporozoites normally invade hepatocytes and transform into AMA1KD hepatic merozoites27. The latter cannot accumulate compensatory mutations before IFC, as they are generated in the absence of selection pressure and following a single invasion/multiplication cycle. We first analysed control hepatic merozoites. IFC analysis shows that ~44.3% and ~4.1% of control GFP+ hepatic merozoites score as EryA and EryI, respectively, indicating that erythrocytic and hepatic merozoites bind and invade erythrocytes with similar efficiency in this assay. We then used AMA1KD hepatic merozoites, composed of ~85% of excised AMA1− parasites lacking any detectable AMA1 and ~15% of non-excised AMA1+ individuals used as internal controls (Fig. 6b). IFC analysis after AMA1 immunostaining (Fig. 7a) shows that ~48.8% and ~5.3% of AMA1+ controls score as EryA and EryI, respectively, indicating that they behave like the WT (Fig. 7b). In contrast, only ~3.3% of AMA1− merozoites score as EryA (Fig. 7b), that is, ~15-fold less than internal controls, demonstrating a major role of AMA1 in merozoite attachment. As expected, AMA1− merozoites also generate EryI events after 10 min (Figs. 6c and 7b) or 3 min incubation (Fig. 7c). Remarkably, EryI events are approximately fivefold less frequent in AMA1− than AMA1+ merozoites when normalized to input merozoites, but approximately threefold more frequent in AMA1− merozoites when normalized to attached parasites (Fig. 7b, P<0.01, two-tailed t-test). Therefore, as with the Toxoplasma tachyzoite, AMA1 favours Plasmodium merozoite attachment to, but not internalization into, the host cell.
AMA1 has no role in P. berghei sporozoite infection of hepatocytes
Recent work using P. berghei AMA1KD and RON4KD sporozoites revealed strikingly distinct phenotypes, with essential and dispensable roles for RON4 and AMA1, respectively, during sporozoite invasion of hepatocytes27. To test AMA1KO sporozoite capacity to invade hepatocytes, populations of GFP+ AMA1KO/GFP− AMA1+ parasites were transferred to mosquitoes. The same ratio of GFP+ versus GFP− sporozoites is found in the blood fed to mosquitoes and in the mosquito salivary glands, indicating that AMA1 has no detectable effect on parasite development in the mosquitoes (Fig. 8a). The capacity of these salivary gland sporozoites to invade cultured hepatocytes was then tested. After sporozoite incubation with HepG2 cells in vitro, a similar proportion of AMA1KO versus GFP− AMA1+ parasites is found in the input sporozoites and in hepatic schizonts developing inside HepG2 cells 60 h post infection (Fig. 8b). Likewise, in co-infection experiments of HepG2 cells with RFP+ AMA1+ as control, AMA1KO sporozoites display similar invasive capacity as the control (Fig. 8c).
Finally, the infectivity of AMA1KO sporozoites was tested in vivo. We found that intravenous injection into mice of as few as 500 AMA1KO/AMA1+ sporozoites (Fig. 8d) or HepG2 cell-released hepatic merozoites (not shown) is sufficient to generate blood-stage parasite populations containing AMA1KO parasites, demonstrating that parasites can complete a life cycle without producing AMA1. Moreover, injection into mice of only 50 AMA1KO/AMA1+ infected erythrocytes is also sufficient to produce AMA1KO-containing blood-stage populations (not shown). However, attempts of cloning AMA1KO parasites were unsuccessful. This is likely due to the slower increase in parasitemia of AMA1KO parasites, delaying the emergence of an AMA1KO population that is eventually cleared by the mouse immune system before being detectable. Nonetheless, we cannot rule out the formal hypothesis that AMA1KO parasites cannot be cloned because they require soluble AMA1 secreted from the AMA1+ counterparts. However, this hypothesis of AMA1 as an essential diffusible factor appears unlikely, as AMA1KO growth is observed after co-injection of less than 50 blood stages and 500 sporozoites in the whole animal.
We have inactivated AMA1 both in Toxoplasma and Plasmodium using diCre-loxP-mediated recombination and direct gene targeting, respectively, and found that AMA1-deficient T. gondii tachyzoites and P. berghei merozoites and sporozoites were still invasive and displayed a normal host cell penetration step. The most striking phenotype is that of AMA1KO sporozoites, which showed no defect in hepatocyte invasion, confirming prior data obtained with AMA1KD sporozoites that invaded hepatocytes even better than the WT27. This now demonstrates that AMA1 is dispensable for hepatocyte invasion and that, given the essential role of RON4 in the process27, the RON complex acts in an AMA1-independent manner. The lack of an invasion phenotype of AMA1KO sporozoites strongly suggests that AMA1 is not involved in TJ function.
In contrast to sporozoites, AMA1-deficient merozoites and tachyzoites displayed an approximately three- to fivefold decrease in overall invasion efficiency. However, like sporozoites, they penetrated host cells like the WT. They formed a normal constriction and a normal RON ring at the TJ, and tachyzoites were internalized at the normal average speed of ~20 s. Moreover, quantitative IFC analysis indicated that AMA1KD merozoites invaded erythrocytes better than controls when normalized to adherent merozoites, reminiscent of the increased infectivity of AMA1KD sporozoites.
The decrease in invasion efficiency of AMA1-deficient tachyzoites and merozoites, which showed no defect in host cell penetration, was associated with altered zoite adhesion to host cells. Fewer AMA1-deficient merozoites bound to erythrocytes in IFC experiments, including in 3′ adhesion assays. Lack of AMA1 only modestly reduced the numbers of bound tachyzoites but affected their positioning onto cells, with AMA1-deficient tachyzoites more frequently adopting an upward position when compared with controls. AMA1 might thus be important in a pre-invasive zoite orientation step, as earlier proposed for merozoites39. A gradient of AMA1 on the zoite surface might create a gradient of interaction forces in a Velcro-like mechanism that might either apically reorient a zoite-expressing AMA1 mostly at its front end (merozoite) or flatten a zoite-expressing AMA1 all over its surface (tachyzoite). Why AMA1 has zoite-dependent contributions is unclear but might be related to zoite shape. A zoite-specific optimal positioning step, possibly involved in inducing rhoptry secretion, might be useful for the pear-shaped tachyzoites and merozoites and dispensable or even inhibitory for the naturally flattened sporozoites.
Therefore, genetic data indicate a model where AMA1 and the RON proteins have separate roles during apicomplexan invasion. AMA1 acts in a host cell-binding step that impacts the frequency but not quality of RON-dependent TJ formation, and the AMA1–RON2 interaction is not involved in the transduction of the force generated by the zoite motor during invasion. It can be argued that our data are still compatible with an essential function of AMA1 at the TJ, if the residual invasion capacity of AMA1 mutants is ensured by an AMA1-like, functionally redundant protein. This hypothesis is highly unlikely for several reasons. First, the invasive AMA1 mutants displayed a normal entry phenotype including a fully functional TJ. This implies that any compensatory mechanism would need to be of optimal efficiency but expressed in only a subset of mutants (those that invade), a situation different from classical compensation by a suboptimal homolog that affects phenotype quality in all mutant parasites. Second, P. berghei AMA1KD sporozoites generated by Flp/FRT-mediated 3′-untranslated region (UTR) excision and T. gondii AMA1KD tachyzoites generated by Tet-mediated transcriptional repression were silenced immediately before phenotype analysis, thus precluding any selection of compensatory mechanism(s). Interestingly, AMA1KO T. gondii tachyzoites grown in continuous culture, which adapted by overexpressing the AMA1 homologue TgME49_300130 by ~15-fold (Fig. 1d), displayed a significantly milder adhesion phenotype, suggesting that the AMA1 homologue indeed compensated the adhesion defect of AMA1 mutants. The hypothesis of compensation at the TJ is also highly improbable in Plasmodium, which contains a single AMA1 gene. The parasite product most closely related to AMA1 is the transmembrane protein MAEBL40, which in P. berghei is only detected in oocyst sporozoites where it confers binding to the mosquito salivary glands but not invasion of hepatocytes41. Therefore, rather than AMA1-complementing TJ components, AMA1-related proteins in both Toxoplasma and Plasmodium appear to function in zoite adhesion, like AMA1.
One question raised by the model in which AMA1 and the RON proteins have dissociated functions is the role of the AMA1–RON2 interaction. The interaction is not essential but is important, being evolutionarily conserved. It might be required for processing/cleavage of surface AMA1 passing the TJ, perhaps allowing the disengagement of interaction of AMA1 with its host cell receptor and facilitating zoite sliding free into the PV. Interestingly, AMA1 undergoes a conformational change upon RON2 binding17, which could lead to loss of adhesive function or exposure of cleavage sites. This would reconcile the genetic data and the fact that antibodies or small molecules that inhibit the interaction can reduce zoite invasion9,10,14,15,16,18,24,42. The increased frequencies of Plasmodium merozoite (relative to adhesive parasites) and sporozoite invasion might also point to a modulatory/inhibitory role, possibly in preventing other interactions important for TJ formation. More work is needed to understand the exact contribution of the AMA1–RON2 interaction, which appears to impact AMA1 but not the TJ per se.
The demonstration of the dispensability of AMA1 in any step of host cell invasion by apicomplexan zoites does not question the potential efficacy of AMA1 as target of malaria prevention measures. A large body of work shows the efficacy of antibodies to AMA1 in blocking erythrocyte infection43,44,45, which might also reduce sporozoite invasion of hepatocytes46. Likewise, although the AMA1–RON2 interaction might not have any positive role in invasion, its inhibition by small molecules might still efficiently perturb invasion. Nonetheless, our finding that AMA1-less variants would be only partially impaired in adhesion while retaining normal if not increased invasive capacity, raises the possibility of rapid parasite adaptation to intervention strategies targeting only AMA1 or the AMA1–RON2 interaction.
P. berghei WT ANKA strain GFP fluorescent (GFP@HSP70)36, RFP fluorescent (L733)35, AMA1/Cond27 or AMA1KO were maintained in 3-week-old female Wistar rats or 3-week-old female Swiss mice. Mice or rats were infected with P. berghei parasites by intraperitoneal or intravenous injections. Parasitemia was followed daily by blood smears and FACS analysis. Anopheles stephensi (Sda500 strain) mosquitoes were reared at the Centre for Production and Infection of Anopheles (CEPIA) at the Pasteur Institute as described47. HepG2 cell for sporozoite infection were cultured in Dulbecco’s modified Eagle’s medium (DMEM) or McCoy’s 5A medium supplemented with 10% fetal calf serum and neomycin (50 μg ml−1, Sigma).
T. gondii tachyzoites were cultured in human HFF cells maintained in DMEM supplemented with 10% fetal calf serum, 2 mM glutamine and 25 μg ml−1 gentamicin.
All experiments using rodents were performed in accordance with the guidelines and regulations of the Pasteur Institute and are approved by the Ethical Committee for Animal Experimentation.
Cloning of DNA constructs
To generate the plasmid pGFP-hDHFR-PbAMA1KO, the 3′UTR of Pbama1 was amplified from P.berghei genomic DNA (gDNA) with primers 3′UTR PbAMA1 fw/rv and cloned in sites PstI and XhoI in a modified pUC18 plasmid containing a new multiple cloning site and a human dihydrofolate reductase (hDHFR) cassette48 (plasmid BGP-F). The 5′UTR of Pbama1 was amplified with primers 5′UTR PbAMA1 fw/rv and cloned in sites SacI and EcoRI in a pUC18 plasmid containing GFP@HSP70 cassette36 in sites SalI and SacI. Finally, 3′UTR-hDHFR was removed from the previous plasmid and cloned in the latter in sites PstI and SalI.
To generate p5RT70-loxPAMA1loxP-YFP-HXGPRT, the TgAMA1 open reading frame (ORF) was amplified from T. gondii gDNA using primers TgAMA1 ORF fw/rv. In addition, 5′UTR and 3′UTR of ama1 was amplified from T. gondii gDNA using 5′UTR TgAMA1 fw/rv or 3′UTR TgAMA1 fw/rv, respectively. First, the 5′UTR of TgAMA1 was inserted upstream of p5RT70 in p5RT70-loxPKillerRedloxP-YFP-HXGPRT plasmid33 in ApaI restriction sites. Then, the killerRed ORF was exchanged by TgAMA1 ORF using EcoRI and PacI. Finally, the 3′UTR of TgAMA1 was cloned in after HXGPRT selection cassette using SacI restriction sites.
Transfections and selection
P. berghei genetic manipulation was performed as described49. P. berghei AMA1KO were generated by double homologous recombination to replace the endogenous ama1-coding sequence by a hDHFR cassette48 and a GFP fluorescence cassette36. The targeting sequence with the two homologous regions flanking the selection cassettes was PCR amplified from plasmid pGFP-hDHFR-PbAMA1KO using primers 5′UTR PbAMA1 fw and 3′UTR PbAMA1 rv, and gel purified using the NucleoSpin Gel and PCR Clean-up kit (Macherey-Nagel) following kit instructions. After transfection of an enriched preparation of P. berghei ANKA schizonts and re-injection into mice, mutants were selected with constant treatment with pyrimethamine in drinking water until green fluorescent parasitemia was detected. Drugs were used as described49. The presence of AMA1KO was confirmed by PCR analysis with primers Pa/Pb, specific for the WT ama1 locus, and Pb/Pc, specific for integration at the ama1 locus, and by Southern blotting of total gDNA after digestion with the restriction enzymes MfeI or NdeI, with a probe hybridizing at the 5′UTR of ama1, amplified with primers 5′Pbama1-probe fw/rv, to recognize the WT or the mutant loci with different sizes.
For T. gondii genetic manipulation, ca 1 × 107 of freshly lysed parasites were transfected with 60 μg linearized DNA by electroporation. Selection was performed with mycophenolic acid (12.5 mg ml−1 in MeOH) and xanthine (20 mg ml−1 in 1 M KOH)50, or phleomycin (50 μg ml−1)51.
The TgAMA1IoxP strain was generated by replacement of the endogenous ama1 by floxed ama1 via homologous recombination. The targeting sequence p5RT70-loxPAMA1loxP-YFP-HXGPRT was removed from plasmid by digestion with NsiI and XmaI restriction enzymes and transfected into ku80::diCre recipient strain33. Parasites with stable integration were selected by the treatment with xanthine and mycophenolic acid. Integration by homologous recombination was confirmed by 5′TgAMA1out fw (P1) and p5RT70 rv (P1′) primers. In addition, a PCR with TgAMA1int fw (P2) and TgAMA1 ORF rv (P2′) primers was conducted to discriminate the presence of genomic or coding sequence of ama1 ORF.
To generate TgAMA1KO, amal ORF was excised from the genome by activation of diCre with 50 nM rapamycin for 16 h. Subsequent limited dilution of induced TgAMA1loxP pool led to a clonal TgAMA1KO population, which was confirmed by genomic PCR using primers upstream and downstream of the loxP sites, 5′UTR TgAMA1 fw (P3) and YFP rv (P3′). The loss of ama1 ORF was further verified by a PCR in the ORF with primers TgAMA1int fw (P2) and TgAMA1 ORF rv (P2′).
For complementation studies, TgAMA1FLAG 52 was used.
Southern and western blotting
gDNA from P. berghei and T. gondii to use as a PCR template and for Southern blotting was extracted using Qiagen dneasy blood and tissue kit according to manufacturer’s protocol.
For Southern blotting of P. berghei gDNA, samples were digested with MfeI or NdeI restriction enzymes overnight, precipitated with ethanol, washed and separated in agarose gel. The gel was transferred to a Hybond-XL membrane (GE-Healthcare) and blotting was performed using the DIG easy Hyb kit and DIG wash and block buffer kit from Roche according to manufacturer’s protocol. The probe was amplified with primers 5′Pbama1-probe fw/rv using the DIG Probe Synthesis kit from Roche.
Tachyzoite western blot samples were obtained by spinning down extracellular parasites and incubating with RIPA buffer (50 mM Tris-HCl pH 8; 150 mM NaCl; 1% Triton X-100; 0.5% sodium deoxycholate; 0.1% SDS; 1 mM EDTA) for 20 min on ice. Unless indicated otherwise 106 parasites were loaded onto a SDS acrylamide gel and immunoblot was performed as previously described53. Briefly, proteins were transferred onto a nitrocellulose membrane, after blocking the membranes were incubated with primary antibody for 1 h (mouse anti-AMA1 1:1,000; rabbit anti-aldolase 1:10,000) followed by incubation with horseradish peroxidase-labelled secondary antibodies (1:50,000; Jackson ImmunoResearch) for 2 h.
RNA from freshly egressed parasites was purified using Trizol followed by chloroform extraction. For cDNA synthesis, 2.5 μg of total RNA were retrotranscribed using the SuperScript VILO (Invitrogen, Life Technology). Quantitative real-time PCR was performed on a LightCycler 480 (Roche) using the LightCycler 480 SYBR Green I Master Mix (Roche). PCR primers were designed to amplify a 100-bp target gene fragment: AMA1 Fw: 5′-TGGAGAGAACCCAGATGCGTTCCT-3′; AMA1 Rv: 5′-CAGTGTAGTCGAGGCAACGGCC-3′; TgME49_300130 Fw: 5′-CCAGGACACGATGCCGCTCG-3′; TgME49_300130 Rv: 5′-AACCCCTCCGCCTCGTCCTT-3′. cDNA levels were normalized to α-tubulin levels measured with primers: Fw: 5′-GCATGATCAGCAACAGCACT-3′; Rv: 5′-ACATACCAGTGGACGAAGGC-3′. Experiments were performed four times with two different RNA preparations.
For P. berghei merozoites, sporozoites and infected HepG2 cells immunofluorescence, samples were fixed with 4% paraformaldehyde, 0.0075% glutaraldehyde in PBS for 1 h54, permeabilized with 0.1% Triton X-100 in PBS, blocked with BSA 3% in PBS, and stained with primary rabbit polyclonal antibodies to the P. berghei AMA1 peptide CRASHTTPVLMQKPYY (Eurogentec, 1:500 dilution), or primary polyclonal antibodies to the P. berghei RON2 peptide KKLGKLREKIVNGLFKKRGK (Thermo Scientific, 1:500 dilution), followed by secondary Alexa-Fluor-conjugated antibodies (Molecular Probes, 1:500 dilution). Images were acquired using an Axiovert II fluorescence microscope (Zeiss) or the ImageStreamX from AMNIS.
For T. gondii immunofluorescence analysis, infected HFF monolayers grown on coverslips were fixed in 4% paraformaldehyde for 20 min at room temperature, followed by permeabilization (0.2% Triton X-100 in PBS) and blocking (2% BSA and 0.2% Triton X-100 in PBS). The staining was performed using primary antibody (mouse anti-AMA1, 1:1,000; mouse anti-SAG1, 1:1,000; rabbit anti-MIC2, 1:500; rabbit anti-IMC1, 1:1,500; rabbit anti-GAP45 1:1,000) followed by secondary Alexa-Fluor-conjugated antibodies (Molecular Probes, 1:3,000). Images were acquired with CCD camera under Deltavision Core or confocal Nikon Ti eclipse microscopes (z–stacks of 0.2–0.3 μm, × 100 immersion objective), deconvolved using SoftWoRx Suite 2.0 (Applied Precision, GE) when needed and further processed using ImageJ 1.34r and Photoshop (Adobe Systems) software.
Production of merozoites and ImageStream analysis
Erythrocytic merozoites were obtained by culturing infected rat or mouse blood for 16 h, at 37 °C, 5% CO2 and 10% O2, under shaking (90 r.p.m.), in RPMI 1,640 medium (Gibco) supplemented with 20% fetal calf serum and 50 μg ml−1 neomycin. Mature schizonts were separated in a Nycodenz gradient and merozoites were isolated by filtration of schizonts through a 5-μm filter, followed by another filtration through a 1.2-μm filter.
The GFP fluorescent AMA1KD strain conditionally knocks down AMA1 expression in mosquito stages, producing sporozoite populations in mosquito salivary glands in which up to 95% of the parasites express undetectable levels of AMA1 (ref. 27). AMA1KD sporozoites were used to infect HepG2 cell in vitro and 62 h post infection the emerging merosomes were collected, and hepatic merozoites were obtained by filtration through a 5-μm filter.
The purified merozoites were cultured in vitro with PKH26-stained rat red blood cells for 3 or 10 min under agitation (400 r.p.m.), and cultures were fixed with 4% paraformaldehyde, 0.0075% glutaraldehyde in PBS for further permeabilization and staining with anti-AMA1 (Eurogentec, peptide CRASHTTPVLMQKPYY) and secondary Alexa-Fluor 647. Cells were acquired in an ImageStreamX using a × 60 objective, excitation lasers 488, 561 and 642 nm, and analysed using the software IDEAS, from AMNIS.
Merozoites attached to red blood cells were assessed by double fluorescence (PKH26 and GFP), and invaded cells were assessed with a sequence of algorithms that identify PKH26 duplication because of PV formation (R3 Bright Similarity Channels 2 and 4, Intensity Weighted).
T. gondii invasion/attachment assay
To investigate the attachment and invasion rates of the TgAMA1KO parasites, a red/green invasion assay was performed as described earlier55. HFFs were grown on coverslips of a 24-well plate and infected with 5 × 106 freshly collected parasites. Plates were centrifuged for 2 min at 200 g and incubated at 37 °C, 5% CO2 for 15 min. Subsequently, cells were fixed in 4% paraformaldehyde (PFA) for 15 min followed by immunostaining with α-SAG-1 primary and Alexa-Fluor secondary antibodies before Triton X-100 permeabilization (0.2% in PBS) and immunostaining with α-IMC1 primary and Alexa-Fluor secondary antibodies. Extracellular and intracellular parasites were counted in ten fields of view (× 100 objectives) and calculated as a percentage value of RH Δhxgprt parasites normalized to 100%.
To measure T. gondii tachyzoite position relative to the host cell, we adapted the assay previously described27. HeLa cells were plated on poly-lysine-coated glass coverslips in a 6-well plate, transfected with 1 μg of plasmid-encoding mCherry in the pDisplay Vector (Invitrogen) and used 20 h later for a 5-min invasion assay. Cells were fixed in PBS-4% PFA (20 min, room temperature) and stained with anti-SAG-1 antibodies followed by Alexa-Fluor anti-mouse antibodies to label extracellular parasites. Samples were scanned on the confocal Nikon Ti Eclipse microscope and images were captured and analysed with Metamorph software (using the 4D viewer application). For each zoite, an ellipsoid was fit to measure the longitudinal axis, whereas the cell surface contacting the tachyzoite centres of mass was affected to all the isosurfaces. The plane of the cell was reconstructed and angle values between the longer axis of the parasite and the host cell plane were generated by Metamorph.
T. gondii replication assay
1 × 105 ku80::diCre or TgAMA1FLAG or 5 × 105 TgAMA1KO were inoculated onto a confluent monolayer of HFFs grown on coverslips (24-well plate) and incubated in normal growth conditions. One hour post inoculation, coverslips were washed in PBS to remove extracellular parasites and thus synchronize the cell cycle. Cells were further grown in normal growth conditions until as indicated, fixed and immunostained. The number of parasites per vacuole was determined for 100 vacuoles.
T. gondii plaque assay
200 RH Δhxgprt or TgAMA1FLAG parasites or 1,000 TgAMA1KO parasites were added onto a confluent monolayer of HFF cells of a six-well plate. After incubating for 6 days, the HFF monolayer was washed in PBS and fixed in ice-cold methanol for 20 min. Afterwards, the HFF cells were stained with Giemsa. The area of ten plaques was assessed using Image J 1.34r software.
T. gondii egress assay
4 × 105 parasites were grown in HFF monolayers on coverslips for 36 h. Media were exchanged for pre-warmed, serum-free DMEM supplemented with calcium ionophore 2 μM (A23187 in DMSO)56. After incubation for 5 min at normal growth conditions (37 °C; 5% CO2), cells were fixed and stained with anti-GAP45 primary antibody and Alexa-Fluor secondary antibody. Two hundred vacuoles were scored in each experiment.
T. gondii motility assay
Freshly egressed tachyzoites were allowed to glide for 30 min on glass coverslips coated with 50 μg ml−1 heparin in PBS. Parasites and trails were then stained with anti-P30 antibodies and visualized with an inverted laser scanning microscope (Eclipse Ti, Nikon). Images were analysed using Metamorph and ImageJ software. Numbers of helical and circular trails associated with parasites were scored in 30 fields.
T. gondii video microscopy
Time-lapse video microscopy was conducted with the DeltaVision® Core microscope using a × 40 immersion lens. Freshly lysed RH Δhxgprt, ku80::diCre or TgAMA1KO were added onto HFF, HeLa or U373 monolayer grown in glass dishes (ibidi; μ-Dish35 mm, high). Forty hours post inoculation, invasion of freshly egressed parasites was observed. Normal growth conditions were maintained throughout the experiment (37 °C; 5% CO2). Images were recorded at one frame per second. Further image processing was performed using ImageJ 1.34r software and with Photoshop (Adobe Systems).
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We thank the Center for Human Immunology at Institut Pasteur for support in conducting this study, the Center of Production and Infection of Anopheles (CEPIA-Institut Pasteur) for mosquito rearing, the ISO 9001 qualified CYBIO FACS facilities of the Institut Cochin for support, Eloise Galtier for FACS acquisition, and Ming Kalanon for critical reading of the manuscript. This work was supported by funds from the Pasteur Institute, the French National Research Agency, the EviMalar, the Wellcome Trust, and the Fondation pour la Recherche Médicale (FRM team support, Barriers and Pathogens). D.Y.B. was a recipient of a Manlio Cantarini Fellowship, N.A. is supported by an EviMalaR PhD fellowship (European FP7/2007-2013, grant number 242095), V.L. is supported by an EC Marie Curie integration grant (PIRG05-GA-2009-249158), M.M. is funded by a Wellcome Trust Senior Fellowship (087582/Z/08/Z). The Wellcome Trust Centre for Molecular Parasitology is supported by core funding from the Wellcome Trust (085349).
The authors declare no competing financial interests.
Time-lapse series showing a RH ku80::diCre tachyzoite invading a HeLa cell. Frames were recorded every 1s and the video is played at 5 frames s-1. The white arrow indicates the TJ. (AVI 366 kb)
Overlay of DIC + GFP time-lapse series showing an AMA1KO tachyzoite invading a HeLa cell. Frames were recorded every 1s and the video is played at 5 frames s-1. The white arrow indicates the TJ. (AVI 1167 kb)
Overlay of DIC + GFP time-lapse series showing an AMA1KO tachyzoite invading a U373 cell. Frames were recorded every 1s and the video is played at 5 frames s-1. The white arrow indicates the TJ. (AVI 268 kb)
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Bargieri, D., Andenmatten, N., Lagal, V. et al. Apical membrane antigen 1 mediates apicomplexan parasite attachment but is dispensable for host cell invasion. Nat Commun 4, 2552 (2013). https://doi.org/10.1038/ncomms3552
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