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© The Rockefeller University Press,
0021-9525/1999//937 $5.00
The Journal of Cell Biology, Volume 147, Number 5,
, 1999 937-944
Brief Report |
Conservation of a Gliding Motility and Cell Invasion Machinery in Apicomplexan Parasites
menarr01{at}mcrcr6.med.nyu.edu
Most Apicomplexan parasites, including the human pathogens Plasmodium, Toxoplasma, and Cryptosporidium, actively invade host cells and display gliding motility, both actions powered by parasite microfilaments. In Plasmodium sporozoites, thrombospondin-related anonymous protein (TRAP), a member of a group of Apicomplexan transmembrane proteins that have common adhesion domains, is necessary for gliding motility and infection of the vertebrate host. Here, we provide genetic evidence that TRAP is directly involved in a capping process that drives both sporozoite gliding and cell invasion. We also demonstrate that TRAP-related proteins in other Apicomplexa fulfill the same function and that their cytoplasmic tails interact with homologous partners in the respective parasite. Therefore, a mechanism of surface redistribution of TRAP-related proteins driving gliding locomotion and cell invasion is conserved among Apicomplexan parasites.
Key Words: gliding motility • cell invasion • Apicomplexan parasites • thrombospondin-related anonymous protein • micronemal protein 2
© 1999 The Rockefeller University Press
MANY Apicomplexan protozoan parasites have life cycle stages that actively invade target cells. Most of these invasive stages are also mobile by means of gliding motility, a form of substrate-dependent locomotion during which, in contrast to crawling motility, the moving cell maintains a fixed shape. Gliding motility and cell invasion, which depend on microfilaments in the parasite (Dobrowolski and Sibley 1996), are thought to result from a capping activity of parasite surface ligands. After parasite attachment to the host cell, a tight junction initially forms between the apical tip of the parasite and the host cell membrane and is progressively redistributed to the posterior pole of the parasite as it moves into the target cell (Aikawa et al. 1978; Jensen and Edgar 1978; Pimenta et al. 1994). In addition, a capping activity is suggested by the observation that invasive stages are able to redistribute molecules bound to their surface, such as antibodies or cationized ferritin, and shed them from their posterior pole (Vanderberg 1974; Dubremetz et al. 1985; Speer et al. 1985). Based on these phenomena, it has been proposed that forward locomotion on the substrate and penetration into a host cell result from the backward translocation of parasite ligands bound to their substrate/cell receptors (Russell and Sinden 1981; Russell 1983; King 1988).
In Plasmodium (malaria) sporozoites, the parasite form that infects the salivary glands of the mosquito vector and the liver of the mammalian host, thrombospondin-related anonymous protein (TRAP) (Robson et al. 1988) is a candidate ligand for interaction with host cell or substrate receptors. TRAP is a type 1 transmembrane protein that carries two adhesive domains in its extracellular portion, an A-type domain first described in von Willebrand factor and a motif similar to the type 1 repeat of thrombospondin (TSR). As shown in Table , a putative TRAP paralog has been identified in the ookinete stage of Plasmodium (CTRP) (Trottein et al. 1995; Yuda et al. 1999) and putative orthologs have been identified in the Apicomplexan parasites Toxoplasma (micronemal protein 2, MIC2) (Wan et al. 1997), Eimeria (Etp100) (Tomley et al. 1991; Pasamontes et al. 1993) and Cryptosporidium (TRAPC1) (Spano et al. 1998). These proteins carry various numbers of A-type and TSR domains, but their cytoplasmic tails do not exhibit primary sequence homology. Some of these proteins (TRAP, MIC2, and Etp100) have been localized to the parasite micronemes, secretory vesicles of the apical complex that release their content at the anterior pole of the parasite. Gene disruption experiments have shown that TRAP(–) sporozoites do not display gliding motility and do not infect the salivary glands of the mosquito vector and the liver of the mammalian host (Sultan et al. 1997).
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L deletion was generated by PCR using 5' primer P008 (5' CGCGAAGCTTCTGAATGTTCTACTACATGTGACAATG 3'), which hybridizes from nt 736 of TRAP onwards, and 3' primer P011 (5' CGCTTAATTAACGCTACTTCCTGCTATAAAATTATAACC 3'), which hybridizes at the 3' end of TRAP and introduces a stop codon as well as a PacI restriction site (bolded). The resulting PCR product was then cloned into plasmid pCRScriptSK, yielding plasmid pL1. A linker encompassing the first 24 bp of the TRAP 3' UTR, including the natural XbaI site located 18 bp from the stop codon, was then cloned downstream from the TRAP coding sequence in plasmid pL1 using PacI and EcoRI adaptors, creating plasmid pL2. Further 3' UTR, borne by a XbaI–XmnI 1.4-kb fragment, was then cloned downstream from the linker in plasmid pL2 digested with XbaI and EcoRV, yielding plasmid pMutL. The HincII–AflII internal portion of plasmid pMutL, which extends from nt 1150 of TRAP to 0.6 kb 3' to its stop codon, was then used to replace its wild-type counterpart in plasmid pINCO, giving rise to plasmid pTL. The 3' end of TRAP bearing the TS deletion was generated by PCR using 5' primer P017 (5' GAATGGAGTGAATGTTCTACTACATGTG 3'), which hybridized from nt 738 of TRAP onwards, and 3' primer P012 (5' CGCTTAATTAACAACAATACCCTTTTCATCATCTGC 3') that hybridizes at the 3' end of TRAP and introduces a stop codon as well as a PacI restriction site (bolded). The resulting PCR product was then cloned into plasmid pCRScriptSK, yielding plasmid pS1. The 3' UTR of TRAP, borne by the PacI-KpnI fragment of plasmid pMutL, was further cloned into plasmid pS1, yielding plasmid pMutS. The HincII-AflII internal portion of plasmid pMutS was then used to replace its wild-type counterpart in plasmid pINCO, giving rise to plasmid pTS. The exchanged fragments in plasmids pTS and pTL were sequenced and confirmed to differ from the corresponding wild-type fragment only by the desired mutation.
The DNA encoding the cytoplasmic tail of MIC2 was amplified from the XhoI fragment of BAC G11-11 (Wan et al. 1997) using 5' primer P27 (5' AAAACTGCAGGATCCCCATCCGCGGAGATAG 3') and 3' primer P28 (5' TGCTCTAGATATATATGTTTATTAAAATTACTCCATCCACATATCACTATCG 3'), which contain a PstI and a XbaI site, respectively (bolded). The resulting PCR fragment was digested with PstI and XbaI and cloned into plasmid pMutS digested with the same enzymes, yielding plasmid pMut-MIC2. The AgeI-AflII internal portion of plasmid pMut-MIC2 was sequenced, confirmed to contain the expected sequence, and used to replace its wild-type counterpart in plasmid pINCO, giving rise to plasmid pTMIC.
Both TRYP and ACID mutations were generated using 5' primer P017 (5' GAATGGAGTGAATGTTCTACTACATGTG 3') and 3' primer SK01 for the TRYP mutation (5' TCATCTAGATATATATGTTTATTAAAATTAGCTAGCGTCATTATCTTCAGGTAATTTAAACT- GCTC 3') or SK02 for the ACID mutation (5' TCATCTAGATATATATGTTTATTAAAATTAGTTCCAGGCATTGCTAGCAGG- TAATTTAAACTGCTC 3'). Both 3' primers contain an XbaI site, and a mutation-tagging NheI site (bolded). PCR fragments were cloned into plasmid pCRScript, excised as an AgeI-XbaI insert and cloned into plasmid pMutS digested with AgeI and XbaI. The AgeI-AflII fragments of the resulting plasmids that encompassed the mutations were sequenced, verified to differ from their wild-type counterpart only by the desired mutations, and used to replace the corresponding fragment in plasmid pINCO, yielding plasmids pTRYP and pACID.
Parasite Transfection and Phenotypic Analysis
Parasite transformation and selection was performed as described (Ménard and Janse 1997; Waters et al. 1997). Anopheles stephensi mosquitoes were fed on infected young rats and sporozoites dissected out at days 14–18 postfeeding. Preparation of sporozoites from the various mosquito compartments was as described (Sultan et al. 1997). For immunofluorescence assays, sporozoites were incubated in RPMI–3% BSA on ice for 3 h, pelleted, and resuspended in 0.5% BSA/PBS containing primary antibody at 1:50. After 30 min at 37°C, sporozoites were pelleted, washed three times with PBS, and air dried in wells on glass IFA slides. For permeabilized staining, some wells were incubated again with primary antibody after drying. Revelation was performed with anti–rabbit IgG-FITC (Kirkegaard & Perry Laboratories, Inc.) at 1:40 in 0.5% BSA/PBS for 30 min at 37°C, slides were washed three times with PBS and mounted. For cell invasion assays, 
105 HepG2 cells were seeded in eight chamber slides and grown to semiconfluency. Sporozoites (15,000/well) were added, incubated 2 h at 37°C, and washed off. After 48 h, parasite exoerythrocytic forms (EEF) were revealed as described (Sultan et al. 1999) using primary antibody against parasite HSP70, which is expressed only in maturing liver stages.
Immunoelectron Microscopy
Samples were fixed for 30 min at 4°C with 1% formaldehyde, 0.5% glutaraldehyde in 0.1 M phosphate buffer, pH 7.4. Fixed samples were washed, dehydrated, and embedded in LR White resin (Polysciences Inc.) as described previously (Aikawa and Atkinson 1990). Thin sections were blocked in PBS containing 5% wt/vol nonfat dry milk and 0.01% vol/vol Tween 20 (PBTM). Grids were then incubated with primary antibodies diluted 1:50 to 1:300 in PBTM for 2 h at room temperature. After washing, grids were incubated for 1 h in 15-nm gold-conjugated goat anti–rabbit IgG (Amersham Life Sciences), diluted in 1:20 in PBS containing 1% wt/vol bovine serum albumin and 0.01% vol/vol Tween 20 (PBTB), rinsed with PBTB, and fixed with glutaraldehyde to stabilize the gold particles. Samples were stained with uranyl acetate and lead citrate, and examined in an electron microscope (CEM902; Carl Zeiss, Inc.).
Antibodies
Polyclonal antibodies against the TRAP repeats (antirepeats) were obtained using a recombinant polypeptide corresponding to residues 263–428 of TRAP, and polyclonal antibodies against the TRAP cytoplasmic tail (antitail) using the synthetic peptide corresponding to D586DE to DND604 of the TRAP tail.
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S and TL, respectively. For this, modifications in the single-copy TRAP gene were introduced via a single recombination event promoted by targeting insertion plasmids (Nunes et al. 1999). As shown in Fig. 1 B, homologous integration of these targeting plasmids generate two TRAP copies: the first is full-length, bears the mutation, and is flanked by expression sequences, whereas the second lacks the 5' part of the gene and promoter sequences.
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S- or the TL-encoding mutations (plasmids pINCO, pTS, and pTL, respectively), were independently transformed into WT merozoites. One parasite clone from each transformation experiment, named INCO (integration control), TS, and TL, was selected in rats by limiting dilution. Southern hybridization demonstrated that parasites in each clone had a single copy of the corresponding plasmid integrated into chromosomal TRAP, as depicted in Fig. 1 B (data not shown). The first TRAP duplicate, which should contain the mutation present in the targeting plasmid, was specifically amplified by PCR using primers O1 and T7 (Fig. 1 B) and analyzed by restriction digestion. The PCR products obtained from TS and TL parasites contained the corresponding deletion tagged with a PacI site, as shown in Fig. 1 C.
Mutant parasite clones, created at the red blood cell stage, were then transmitted to Anopheles stephensi mosquitoes. Sporozoites are formed inside oocysts in mosquito midguts and are released in the hemolymph, the fluid that bathes the mosquito body cavity. TRAP production was assessed in sporozoites collected from mosquito midguts; i.e., freshly released from, or still within, oocysts. Sporozoite extracts were subjected to Western blot using antibodies directed against the TRAP extracellular repeats (antirepeats) or the TRAP cytoplasmic tail (antitail). As shown in Fig. 2 A, wild-type and INCO sporozoites produced similar amounts of TRAP. TS and TL sporozoites expressed comparable amounts of their TRAP truncate that, as expected, did not react with antitail antibodies.
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S and TL sporozoites did not invade the salivary glands. In the WT and INCO clone, virtually all sporozoites permeabilized before staining strongly reacted with antirepeats as well as antitail antibodies, in a typical patchy pattern (Fig. 2 B). However, when staining was performed with live parasites, only 5% of sporozoites in both populations reacted with antirepeats, but not antitail, antibodies. Fluorescence patterns mostly resembled caps covering the sporozoite body to various extents, frequently over less than half its length. In addition, a ring-type pattern could be observed in a small number of WT sporozoites. In the TS and TL clones, 5% of the live sporozoites also strongly reacted with antirepeats antibodies and displayed the cap-type fluorescence pattern (Fig. 2 B), indicating that the cytoplasmic tail of TRAP is dispensable for surface exposure of the protein. To determine the subcellular localization of TRAP, WT and TL sporozoites were examined by immunoelectron microscopy with antirepeats and antitail antibodies (Fig. 2 C). Using antitail antibodies, WT, but not TL, sporozoites were labeled. However, micronemal localization was not unambiguous with these antibodies. Labeling with antirepeats antibodies showed association of TRAP with the micronemes in both TL (Fig. 2 C) and WT (data not shown) sporozoites. This subcellular localization was similar to that previously described for Plasmodium falciparum TRAP/SSP-2 (Rogers et al. 1992a), indicating that the TL truncate was correctly targeted to the parasite micronemes.
We then examined the phenotype of recombinant sporozoites (Table ). Similar numbers of sporozoites were found in the midguts of mosquitoes infected with the three parasite populations. However, dramatically fewer sporozoites were found associated with the salivary glands of mosquitoes infected with TS or TL parasites than with INCO or WT parasites. This indicated that the former, like TRAP(–) sporozoites (Sultan et al. 1997), did not infect salivary glands. Sporozoite infectivity to the vertebrate host was estimated by injecting sporozoites intravenously into rats and measuring the prepatent period of erythrocytic infection. Whereas WT and INCO sporozoites induced erythrocytic infections with similar prepatent periods, TS and TL sporozoites were not infective. Microscopic examination of sporozoites deposited on glass slides revealed that WT and INCO sporozoites glided with similar speed (2 µm/s) and pattern, but that TS or TL sporozoites did not exhibit typical gliding (TS sporozoites displayed a limited locomotion described below). Sporozoite invasion into mammalian cells was examined by immunostaining of EEF of the parasite that developed within hepatoma HepG2 cells. Whereas WT and INCO sporozoites generated similar numbers of EEF, no EEF was detected in cells incubated with either TS or TL sporozoites. Therefore, the cytoplasmic tail of TRAP is dispensable for surface presentation of the protein, but is essential for sporozoite gliding motility and cell invasion.
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L deletion site (Fig. 1 A). The insertion plasmid that contained the sequence encoding the tail switch, pTMIC, was transformed into WT merozoites, and one resistant clone, TMIC, was selected. Southern blot hybridization confirmed that a single copy of the plasmid was integrated at the TRAP locus in TMIC parasites (data not shown), and PCR amplification of the first TRAP duplicate confirmed the presence of the additional BamHI site tagging the exchange (Fig. 1 C). To rule out the possibility that a discontinuous gene conversion event had preserved the BamHI site (located at the junction of the TRAP and MIC2 sequences; see Fig. 1 B) but corrected downstream heterologies, the sequence of two independent PCR products generated with primers O1 and T7 was determined. Both sequences contained the desired exchange. As shown in Table , TMIC sporozoites behaved similarly to WT or INCO sporozoites in all tests performed. Most strikingly, TMIC sporozoites glided at the same average speed and followed a similar circular pattern than WT or INCO sporozoites (Fig. 3 A). In addition, TMIC sporozoites were as infectious to the rodent host as WT sporozoites. In all rodent infection experiments, Southern hybridization indicated that the blood stages of the parasite induced by TMIC sporozoites still contained the TMIC recombinant locus (data not shown), with no trace of WT TRAP that could have been recreated via plasmid excision. We conclude that the cytoplasmic tail of MIC2, despite little primary amino acid sequence similarity to the TRAP cytoplasmic tail, can function in its place during gliding locomotion and cell invasion by malaria sporozoites. The cytoplasmic tails of these proteins must then interact with homologous partners in the respective Apicomplexan host.
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S, and TL sporozoites. Ring-type patterns, however, seemed less intense in TRYP and ACID sporozoites than in the WT. However, since the ring-type patterns showed some variability in WT sporozoites themselves (Fig. 2 B and 3 B), the significance of this difference remains questionable.
As shown in Table , both TRYP and ACID sporozoites were not infective to the mosquito salivary glands or the rodent liver, nor did they invade HepG2 cells in vitro. Surprisingly, the gliding phenotype of TRYP and ACID sporozoites was not abolished but drastically modified, and identical to the gliding pattern of TS sporozoites. Whereas 10% of WT hemolymph sporozoites typically described circles at a constant speed, the same proportion of hemolymph sporozoites in the TRYP, ACID, and TS clones all displayed an identical phenotype of "pendulum" gliding. This new phenotype consisted in repeated cycles of (a) gliding over one third of a circle, (b) stopping for usually 1–2 s, and (c) moving back to the original position (Fig. 3 A). This phenotype was not observed with TL or TRAP knock-out sporozoites, indicating that it depends on the cytoplasmic tail of TRAP. Importantly, it was not due to low-level expression (as verified with TS sporozoites) or mistargeting of the TRAP variant (since their surface presentation was not affected), and thus appeared to be a direct consequence of the modifications.
In WT sporozoites, TRAP is released on the substrate during gliding locomotion (Fig. 3 B). Labeling of TRAP on the substrate revealed a periodic intensity pattern reminiscent of the nonuniform expression of TRAP on the sporozoite surface. Also, TRAP released on the substrate is detected by antirepeats, but not antitail, antibodies (data not shown), suggesting that TRAP release occurs after cleavage of the protein. Therefore, the pendulum phenotype may be explained by the sporozoite capacity to translocate TRAP to its posterior tip, making it glide over one third of a circle (corresponding approximately to the sporozoite length), and incapacity to release TRAP accumulated at its posterior pole, making it stop. The TRAP cytoplasmic tail could therefore have a bifunctional character; the membrane proximal part would be required for protein translocation along the cortical microfilaments, whereas the distal part would contain a signal for TRAP release at the posterior pole, probably by proteolytic cleavage.
Whatever the molecular basis for the pendulum phenotype, it demonstrates a direct role of TRAP as a transmembrane link in gliding motility (and cell invasion). The critical role of the conserved tryptophan and acidic residues also indicates a similar role for TRAP-related proteins.
Concluding Remarks
The results presented here show that TRAP and structurally related proteins constitute a family of functionally homologous bridging proteins involved in parasite gliding motility and cell penetration. The interchangeability of the TRAP and MIC2 cytoplasmic tails suggests the conservation of the molecular machinery that drives protein redistribution. The cytoplasmic tails of these proteins may interact with some component of the cortical microfilament system, possibly a motor protein. Recent evidence suggests that myosin colocalizes with actin underneath the parasite plasma membrane in a circumferential pattern and powers motility and cell invasion by Toxoplasma gondii tachyzoites (Dobrowolski et al. 1997). Favorable signals on the parasite surface may trigger interaction between the cytoplasmic tail of the TRAP-related protein with myosin, and their translocation along submembranous actin filaments. In addition, if our interpretation of the pendulum phenotype is correct (i.e., if the distal part of the TRAP cytoplasmic tail and the conserved tryptophan are necessary for protein release), then the product(s) necessary for releasing the bridging protein may also be conserved in these parasites.
Members of the TRAP family of proteins have been found in the malaria sporozoite and ookinete, as well as in invasive stages of Toxoplasma, Cryptosporidium, and Eimeria. An invasion system dependent on the redistribution of TRAP-related proteins, which contain various associations of A-type and TSR domains, may be restricted to penetration of Apicomplexan parasites into epithelial cells. For example, the bridging protein of the capping machinery used by the malaria merozoite for invading a red blood cell, which remains unknown, probably requires specialized cell-binding modules. However, the cytoplasmic tail of such protein might carry the features shared by members of the TRAP protein family. Further unraveling of the mechanisms of gliding motility and cell invasion by Apicomplexa should reveal new features of cell adhesion associated with force transduction and may identify common targets for blocking their infectivity.
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This work was supported by grants from Burroughs Wellcome Fund (New Initiative in Malaria Research), United Nations Development Programme/World Bank/World Health Organization Special Programme, the Karl-Enigk Foundation, and the National Institutes of Health (AI-43052 and AI-35827). S. Kappe is a recipient of the B. Levine fellowship in malaria vaccinology. R. Ménard is a recipient of the Burroughs Wellcome Fund Career Award in the Biomedical Sciences.
Submitted: 30 September 1999
Revised: 18 October 1999
Accepted: 18 October 1999
Stefan Kappe and Thomas Bruderer contributed equally to this work and should be considered co-first authors.
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