Motile Systems in Malaria Merozoites: How is the Red Blood Cell Invaded?

Reviews 40 Freedman, L.P. (1999) Increasing the complexity of coactivation in nuclear receptor signalling. Cell 97, 5–8 41 Kliewer, S.A. et al. (1999) Orphan nuclear receptors: shifting endocrinology into reverse. Science 284, 757–760 42 Laudet, V. (1997) Evolution of the nuclear receptor superfamily: early diversification from an ancestral orphan receptor. J. Mol. Endocrinol. 19, 207–226 43 Escriva, H. et al. (1997) Ligand binding was acquired during evolution of nuclear receptors. Proc. Natl. Acad. Sci. U. S. A. 94, 6803–6808 44 Sluder, A.E. et al. (1999) The nuclear receptor superfamily has undergone extensive proliferation and diversification in nematodes. Genome Res. 9, 103–120 45 Yao, T.P. et al. (1992) Drosophila ultraspiracle modulates ecdysone receptor function via heterodimer formation. Cell 71, 63–72 46 Freebern, W.J. et al. (1999) Identification of a cDNA encoding a retinoid-X-receptor homologue from Schistosoma mansoni. J. Biol. Chem. 274, 4577–4585 47 Mangelsdorf, D.J. and Evans, R.M. (1995) The RXR heterodimers and orphan receptors. Cell 83, 841–850 48 Mangelsdorf, D.J. et al. (1991) A direct repeat in the cellular retinolbinding protein type II gene confers differential regulation by RXR and RAR. Cell 66, 555–561

49 Kostrouch, Z. et al. (1998) Retinoic acid X receptor in the diploblast, Tripedalia cystophora. Proc. Natl. Acad. Sci. U. S. A. 95, 13442–13447 50 Biesalski, H.K. et al. (1992) Modulation of myb gene expression in sponges by retinoic acid. Oncogene 7, 1765–1774 51 Wolff, K.M. and Scott, A.L. (1995) Brugia malayi: retinoic acid uptake and localization. Exp. Parasitol. 80, 282–290 52 Lala, D.S. et al. (1997) Activation of the orphan nuclear receptor steroidogenic factor 1 by oxysterols. Proc. Natl. Acad. Sci. U. S. A. 94, 4895–4900 53 Wurtz, J.M. et al. (1996) A canonical structure for the ligand-binding domain of nuclear receptors. Nat. Struct. Biol. 3, 87–94 54 Bobek, L.A. et al. (1988) Small gene family encoding an eggshell (chorion) protein of the human parasite Schistosoma mansoni. Mol. Cell. Biol. 8, 3008–3016 55 Davis, P.J. and Davis, F.B. (1996) Non-genomic actions of thyroid hormone. Thyroid 6, 497–504 56 Haas, W. and Schmidt, R. (1982) Characterization of chemical stimuli for the penetration of Schistosoma mansoni cercariae. 1. Effective substances, host specificity. Z. Parasitenkd. 66, 293–307 57 Prins, A. et al. (1993) Schistosoma mansoni and Trichobilharzia ocellata: comparison of secreted cercarial eicosanoids. J. Parasitol. 79, 130–133

Motile Systems in Malaria Merozoites: How is the Red Blood Cell Invaded? J.C. Pinder, R.E. Fowler, L.H. Bannister, A.R. Dluzewski and G.H. Mitchell The ability of the malaria parasite to invade erythrocytes is central to the disease process, but is not thoroughly understood. In particular, little attention has been paid to the motor systems driving invasion. Here, Jennifer Pinder, Ruth Fowler and colleagues review motility in the merozoite. The components of an actomyosin motor are present, including a novel unconventional class XIV myosin, now called Pfmyo-A, which, because of its time of synthesis and location, is likely to generate the force required for invasion. In addition, there is a subpellicular microtubule assemblage in falciparum merozoites, the f-MAST, the integrity of which is necessary for invasion. Form and function in the invasive zoites of malaria parasites (merozoites, sporozoites, ookinetes) clearly place Plasmodium in the phylum Apicomplexa. Like others in this taxon (eg. Toxoplasma, Eimeria and Gregarina), these zoites possess the characteristic apical organelles used to invade or migrate through host cells. Typical apicomplexans also show an unusual form of motility, called gliding motility, where movement is achieved while maintaining anteroposterior polarity and without cellular deformation1,2. Gliding motility is distinct from cell crawling, as seen in amoebae or neutrophils. Amoeboid crawling shows no constant polarity of movement and deforms the cell membrane by generating filopodia and Jennifer Pinder and Anton Dluzewski are at the MRC Muscle and Cell Motility Unit, Randall Institute, KCL, 26–29 Drury Lane, London, UK WC2B 5RL. Lawrence Bannister is at the Centre for Neuroscience, Hodgkin Building, Guy’s, King’s and St Thomas’ School of Basic Medical Science, KCL, Guy’s Campus, London, UK SE1 1UL. Ruth Fowler and Graham Mitchell are in the Malaria Laboratory, Department of Immunobiology, New Guy’s House, Guy’s, King’s College and St Thomas’ Hospitals’ School of Medicine, KCL, Guy’s Hospital, London, UK SE1 9RT. Tel: +44 207 955 4421, Fax: +44 207 955 8894, e-mail: [email protected] 240

lamellipodia through actin polymerization. The cell body is then pulled forward by tractive effort, generated by myosins interacting with actin filaments. For apicomplexan gliding, a linear motor model has been proposed2, which can also explain the observations of retrograde translocation of latex beads over their surface. In this model, actin filaments anchored to a microtubule, or some other cytoskeletal structure beneath the plasma membrane (PM), interact with myosin heads. The myosin tails interact with transmembrane components, which, in turn, bind the substrate (or bead) externally. The membrane-spanning molecule in other motile stages might be a member of the thrombospondin-related protein/ micronemal 2 (TRAP/MIC2) family of secretory adhesins3,4, but none of this family is known to be present in P. falciparum merozoites. Merozoite invasion The merozoite stage of Plasmodium is adapted to erythrocyte invasion. Unlike sporozoites, the merozoite has not been seen to glide, although it is not known how it behaves in the confines of a venule crowded with adherent schizonts. In vitro, merozoites do not move across a substrate at all, until contacting a red blood cell (RBC) surface. Can invasion, then, truly be a motile process? It is reasonable to expect motor activity in the invading parasite, as the RBC, although capable of endocytosis, is not phagocytic, yet the merozoite must translocate across the plane of the RBC membrane. It is notable that Toxoplasma gondii, which routinely invades cells that can phagocytose, does not exploit the phagocytic behaviour of the cell, but instead uses its own motor system for invasion5. Several steps in RBC invasion can be recognized (reviewed in Ref. 6). The initial attachment of the merozoite to the RBC surface is random and reversible; comparable

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Reviews Box 1. Myosin

I

Myosin is a motor protein that walks along actin filaments towards the barbed end using energy derived from the actinactivated hydrolysis of myosin-bound ATP (Fig. I). Myosins have three structural regions: the head and neck, which are highly conserved, and the tail which confers the specific functions of the various myosin classes. Myosins form a vast, diverse super-family, which is grouped into classes (at present 15), according to the structure of the tail domain (Fig. II). Conventional myosins (skeletal and smooth muscle types belonging to class II), are two-headed filament-forming proteins whereas all other classes, the unconventional myosins, may have two or only a single head. Variously spliced myosins of several classes can be expressed within a single cell. Many myosins have not been isolated, their existence only being known from a molecular biology approach, but some are now being synthesized in a functional form, using eukaryotic protein expression systems.

Rebinding Actin Myosin

Translocation

1

Dissociation

3

2

II

Class XIV

Class I

Toxoplasma gondii TgM-A (AF006626)

Rat myosin 1a (X68199)

III

6 Putative calmodulin/light chain binding sites Membrane-binding tail Class II

Chicken skeletal myosin (P13538) Essential light chains Regulatory }

Head

Neck

Globular Long a helix Binds ATP Composed of 1–6 Binds actin IQ motifs Highly conserved Binds regulatory Motor domain proteins Might phosphorylate Controls calcium sensitivity

Tail which might:

Form a coiled coil Self-associate Contain SH3 motif Bind actin Bind membrane Bind G proteins

Parasitology Today

Coiled coil tail Class VIIa

Human myosin VIIa (U39226)

Kinesin tail homology

attachment occurs, unproductively, on other surfaces. However, on the RBC surface, attachment is followed by an apparently organized motile event: the reorientation of the parasite to bring its apical prominence into contact with the host cell. This reorientation could be driven by a backwards-running motor1. Alternatively, it could depend on receptors for the host cell increasing in avidity towards the merozoite apex (as suggested for TRAP in sporozoite invasion4), or it might, in fact, depend on random Brownian motion. After apical orientation, there is a tight interaction of parasite receptors with specific ligands on the RBC membrane. The relative importance of the roles played in this process by merozoite coat proteins [eg. merozoite surface protein 1 (MSP1)] or apically secreted components [eg. apical merozoite antigen 1 (AMA1), erythrocyte-binding antigen 175 kDa (EBA175)] remains unclear; antibodies against appropriate determinants of numerous merozoite proteins will block invasion. Crucially, the tight attachment step can be uncoupled from the subsequent steps in invasion by pretreatment of the parasites with the actin-capping and -depolymerizing agent cytochalasin B. This was first shown for P. knowlesi7 and later confirmed for P. falciparum8. Merozoites treated with cytochalasin B remain attached to the RBC. With the use of electron microscopy, an electron-dense undercoating of the RBC membrane can Parasitology Today, vol. 16, no. 6, 2000

be seen at the junction of merozoite and RBC. One merozoite submembranous protein has been described uniquely at this junction: merozoite capping protein 1 (MCP1), with a predicted oxidoreductase activity9. The attached merozoites secrete material from their rhoptries, engendering local vacuolation of the RBC membrane7,10. Cytochalasin treatment halts the process at this point where, in uninterrupted invasion, the merozoite generates an invagination (the invasion pit) in the RBC beneath the region of contact. In normal invasion, the merozoite moves into the deepening invagination, while the electron-dense junction travels as an annulus over its surface11. The membrane of this invagination eventually seals at the posterior end of the merozoite to form the parasitophorous vacuole (PV). The PV membrane (PVM) is probably a product both of the secretions of the rhoptries and of the host cell membrane lipid bilayer, augmented soon after invasion by the discharge of the dense bodies into the PV (see, for example, Refs 6,12). It is difficult to conceive that the generation of the PVM can itself be responsible for pulling the merozoite into the RBC; hydrostatic pressure of the haemosol surely militates against this, although insertion of lipophilic components into the cytosolic leaflet of the RBC membrane will cause it to invaginate13. One may suppose, then, that during invasion, true merozoite motility is required twice: first for reorientation and second, for movement into the PV. 241

Reviews

Fig. 1. The location of actomyosin and microtubule systems in Plasmodium falciparum. Confocal microscope images (a) of a mature schizont containing merozoites surrounding a residual body labelled with Pf-myoA antiserum (i) and rhodamine-conjugated phalloidin (ii). The colocalization of actin and myosin along the merozoite membranes is also shown where the two images have been superimposed (iii). Immunofluorescence microscopy of merozoites (b). The localization of cytoskeletal proteins in merozoites is shown by colour codes (i), corresponding to actin labelled with fluorescent phalloidin (green) and myosin labelled with Pfmyo-A antiserum (red) (ii) and the microtubules of the P. falciparum merozoite assemblage of subpellicular microtubules (green) (iii). Merozoite nuclei were stained with DAPI (blue). Immuno-electron microscopy showing distribution of Pfmyo-A in the malaria parasite (c). Immunogoldlabelled merozoites before their release from the red blood cell, labelled with anti-Pfmyo-A (i). The myosin is located around the periphery of the merozoite. The apical prominence (ma) overlies the large pear-shaped secretory rhoptry and elsewhere in the parasite a flat cisterna (cis) underlies the plasma membrane (PM). The nucleus of the parasite (nu) is visible at the base of the merozoite. A longitudinal/oblique section through the apical end of a free merozoite (ii), showing labelling associated with the cortical cytoplasm between the PM (pla) and the outer membrane of the subplasmalemmal cisterna (cis). (Reproduced from Ref. 21, with permission from the Company of Biologists.) Scale bars 5 5 mm (a), 1 mm (b), 200 nm (c i) and 100 nm (c ii). 242

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Reviews Two cisternal Merozoite plasma membrane membranes Merozoite coat RBC receptors

f-MAST (1 of 2/3 microtubules)

Capped actin

Membrane flows past myosin-membrane attachment site?

Microneme

X

X

{

Pa organ rasite elles X etc X X X

Rhoptry

Attachment zone

Ubiquitinated-G-actin

Polar rings ?

Parasitophorous vacuole membrane

−

Actin flow ?

Invasion pit Discharge of rhoptry and microneme contents

+

P

Myosin (Pfmyo-A)

Receptor-driven trans-membrane signal? (G-protein/phosphoinositide link?) Leads to myosin phosphorylation and thence activation of myosin motor

Parasitology Today

Fig. 2. Diagrammatic representation of the apical region of a Plasmodium falciparum merozoite invading a red blood cell. The lefthand half of the diagram shows ultrastructural features, whereas the right-hand half shows the molecular relationships discussed in the text.

Cytoskeletal and motor proteins in cell motility Classic structural molecules with roles in cell motility are actin and tubulin, which are both able to polymerize to form linear arrays: microfilaments and microtubules, respectively. The controlled assembly and disassembly of these structures can generate force. Actin is an extremely highly conserved and very abundant protein, which exists in two interconvertible states: monomeric G-actin, and polymerized F-actin. F-actin polymers are double-helical filaments capable of extension or shortening at either end, which are remodelled and regulated by actin-binding proteins (ABPs) that end-cap, sever and crosslink them. In P. falciparum, ubiquitination might also regulate the polymerization state of actin8. Myosins are ATPases, whose head domains bind to and are activated by filamentous actin, and whose tails bind other structures. When an ATP molecule is hydrolysed, a conformational change of the myosin head occurs such that force and movement are generated, with the actin and other partner moving relative to one another (Box 1). Microtubules play key roles in regulating cell shape and polarity and the distribution of organelles. They also generate motility, notably in stable structures, such as cilia and flagella, and in mitotic and meiotic spindles. Like microfilaments, many cytoplasmic microtubules are highly dynamic, so generating a plastic structural framework. This is regulated by several cellular effectors: microtubule-associated proteins (MAPs) and related proteins14, including kinesin and dynein ATPase motors, which shuttle on cytoplasmic microtubules, and have been shown to mediate many intracellular transport processes. Parasitology Today, vol. 16, no. 6, 2000

Actin in invasion Plasmodium falciparum has two actin genes: (pf)-actinI is expressed throughout the life cycle, but (pf)-actinII is expressed only in sexual stages15. A role for actin in merozoite invasion is suggested by the blocking action of cytochalasin B7. There is good evidence for F-actin in P. falciparum merozoites8,16, but microfilaments have not yet been resolved unequivocally by electron microscopy, which suggests that they might be rather short or unstable. Tardieux et al.17 have characterized a family of actin-capping and -uncapping proteins in P. knowlesi merozoites, including a 70 kDa protein cognate with the heat-shock protein (HSP) of P. falciparum. The presence of this heat shock complex HSC70/34/ 32 kDa complex shortened the average length of rabbit muscle actin filaments in vitro. Tardieux et al. proposed that capping and uncapping of actin could provide the mechanism for localized actin filament growth to generate movement of the merozoite into the host cell17. In P. falciparum schizonts, Tardieux et al.18 have detected a coronin, which can be expected to promote rapid polymerization and crosslinking of actin. Among other apicomplexans, Toxoplasma invasion depends on the actin of the parasite and can be inhibited by cytochalasin5, but Theileria invasion does not, depending instead on modified host endocytosis19. Myosin: Pfmyo-A in invasion Eukaryotic cells generally contain myosins of several classes20. Hence, we can expect that the P. falciparum genome contains genes for several myosins. A search for P. falciparum myosin genes was made21 using the method of Bement et al.22 – ie. the cloning and sequencing of products of amplification obtained by PCR on 243

Reviews genomic DNA using highly degenerate primers to two conserved sequences, both present on all known myosins. One of these sites, the phosphate-binding P-loop, GESGAGKT, is absolutely conserved (it is also present, although not always fully conserved, in GTP-binding proteins). The other site is extremely highly conserved, LEAFGNAKT, and binds nucleotide. Following amplification at low stringency, sequence analysis of 20 clones revealed evidence of only a single myosin gene, Pf-myo1, whereas the same approach had yielded three myosin sequences in Toxoplasma gondii (TgM-A, TgM-B and TgM-C)23. Pf-myo1, like TgM-A, is a very small (~90 kDa) class XIV single-headed myosin, with no recognizable neck region and a very short C-terminal tail (Box 1). Very recently, two further myosin genes have been identified using the facilities of the P. falciparum genome sequencing project24. These were named PfM-B (first identified by Dame and colleagues25) and PfM-C. Both lie within class XIV, although they are considerably larger than Pf-myo1. Hettman and colleagues24 also completed the sequence of Pf-myo1, but referred to it as PfM-A, because of its 63% identity with TgM-A. Subsequently, the authors concerned with describing them21,24 have agreed that the best nomenclature for these myosins, to be used henceforth, is Pfmyo-A (instead of Pf-myo1 or PfM-A), Pfmyo-B and Pfmyo-C (to replace PfM-B and PfM-C, respectively). In the asexual blood stages of P. falciparum, Pfmyo-A is expressed essentially only in mature merozoites21 (Fig. 1). It is peripherally located with a predominance towards the apex (Fig. 1b), and electron microscopy indicates that it is located between the PM and the cisternal membranes (Fig. 1c). It is not detected in young ring stages after invasion, and its first appearance in the schizont is very late in merogony (Fig. 1a). It is labile21, and it partitions with the membrane fraction from merozoites, from which it cannot readily be dissociated, suggesting the presence of a membrane protein or lipid receptor21. A dibasic motif in the myosin tail is thought to be an essential determinant of membrane localization in class XIV myosins24. Thus, Pfmyo-A is in the right place and occurs at the right time to be a component of the merozoite invasion motor. Invasion is also inhibited by the specific myosin ATPase inhibitor 2,3-butanedione monoxime (BDM)21,26. Similarly, the motility of Toxoplasma is blocked by BDM, without affecting microneme secretion27 (whereas myosin light chain kinase inhibitors do not inhibit Toxoplasma motility, but do inhibit microneme secretion in that species27). Microtubules: the f-MAST in invasion Plasmodium falciparum has four single-copy tubulin genes (reviewed in Ref. 28). Of these, those encoding a-I, b and g tubulin are transcribed throughout the life cycle, and the gene encoding a-II tubulin is transcribed predominantly in gametocytes, when unique b tubulin transcripts are also upregulated. The P. falciparum a and b tubulins show great homology with other sequenced tubulins28, but for P. falciparum g tubulin (as for other g tubulins), homology is comparatively low. In Plasmodium, microtubules are abundant in mitotic spindles in schizonts (for details, see Ref. 29). Subpellicular microtubules are found in sporozoites, gametocytes, ookinetes and merozoites. A single narrow band of microtubules has been observed in P. falciparum merozoites, by immunofluorescence29,30 (Fig. 1b), and by electron microscopy31. It consists of two (sometimes 244

three) microtubules, running in parallel along one side of the merozoite from the third polar ring towards the posterior, and lies just beneath the cisternal membranes (Fig. 1b). We have called this structure the P. falciparum merozoite assemblage of subpellicular microtubules (f-MAST)32. Antimicrotubule drugs were used to assess the role of the f-MAST in the invasion process30. Shortening the f-MAST reduced the number of resultant rings, whether using dinitroanilines, which disassemble/fragment plant and protozoan microtubules33,34, or colchicine, which prevents further tubulin polymerization and, at high concentrations, depolymerizes microtubules35. Treatment with Taxol®, which promotes polymerization and stabilizes microtubules36, did not inhibit invasion30,37,38. By exploiting the antithetical activities of colchicine and Taxol®, it was shown that microtubules were the colchicine target and that other effects of colchicine were unimportant here30,32,39. We conclude from these experiments that the integrity of the f-MAST is important, but that invasion does not require dynamic microtubules. There are several functions that the f-MAST might subserve as a stable structure. First, it may act as a directional guide, indicating the apical polarity of the merozoite, or act as a structural brace, perhaps facilitating a cortical flow of F-actin over myosin in the peripheral membranes. Second, Morrisette et al.40 suggest that apicomplexans might move by a novel microtubulebased mechanism, creating cell surface distortions via a filamentous network sandwiched in the subpellicular membranes. Microtubule-based motors would be necessary for this motility, and are implicated in transport processes, including organelle and protein trafficking. Clearly, the f-MAST role might involve microtubulebased motors, such as kinesins, kinesin-like proteins (KIFs) and dyneins. Kinesin-like genes have been identified in the P. falciparum genome and both merozoite dynein and kinesin have been demonstrated by fluorescent imaging and immunoblotting (R.E. Fowler et al., unpublished). Third, it is possible that components of the cytoplasmic dynein complex serve to coordinate the actomyosin and tubulin based motility systems. Finally, merozoite orientation on the RBC surface might be attributable to microtubule motor protein activity. Considerations in modelling invasion Figure 2 shows some possible relationships between structures and events in invasion. We consider that the actin-Pfmyo-A motor is the principal component in P. falciparum invasion motility, but is unlikely to drive the reorientation step (because this occurs in the presence of cytochalasin). Similarly, apical secretion is insensitive to cytochalasin (and in Toxoplasma is separable from BDMsensitive invasion27) and might not involve Pfmyo-A. If Pfmyo-A drives the gliding motility of invasion, then several questions arise. First, we do not know how the motor is controlled. Regulation of Pfmyo-A activity might involve a mechanism similar to that found in amoeboid, yeast and Aspergillus myosins I. They are regulated by phosphorylation at a consensus site: the TEDS site (derived from a single letter amino acid sequence)41. In Pfmyo-A, the TEDS site is a threonine residue, capable of phosphorylation. In Fig. 2, we suggest linkage via G protein from surface receptors; elsewhere we have pointed out that microtubules might also have a signalling role38. Second, we do not know how traction is Parasitology Today, vol. 16, no. 6, 2000

Reviews brought to bear on the host cell. MCP1 in the junction does not span the merozoite membrane, although it might interact with a merozoite cytoskeletal component9. Third, we are not sure how purchase is obtained against the intracellular structures of the merozoite. One possibility is that the f-MAST provides this, either directly or indirectly, perhaps by stiffening the cisternal membranes. This question of purchase might be answered in part if the positioning of actin filaments was such that they could generate ‘cortical tension’. The total F-actin per merozoite can only provide for some 20 microfilaments, each 1 mm in length, but it is likely that the actual filaments are shorter. Staining patterns show that F-actin is distributed in the cell more widely than is myosin (Fig. 1b), and viscosity caused by its polymerization and the crosslinking of short microfilaments might endow the merozoite with the stiffness necessary for invasion. We have modelled both the carriage of merozoite constituents on microfilaments and the flow of membrane forward, while myosin is anchored at the attachment site, but this is not the only possible scheme. The relative positions of actin and myosin might be different21, and the control of the polymerization state of actin by ubiquitin, or coronin and other ABPs, might be crucial8,17,18. The role of f-MAST is enigmatic. The necessity for its presence (but not for microtubule dynamism after late merogony) excludes one aspect of microtubule function, but leaves open the questions of its importance in transport, motility, the determination of polarity and shape or transmission of force. Finally, motile apicomplexans share polar rings. Do these play an important role, for example in determining the direction of movement, as well as anchoring cytoskeletal structures and defining the shape of the apical prominence? Acknowledgements We are very grateful for the support in carrying out this work of the Wellcome Trust (Grant nos 048244 and 045828), The Special Trustees of Guy’s and St Thomas’ Hospitals, The Stanley Thomas Johnson Foundation and The Royal Society. We are greatly indebted to Kate Kirwan for the preparation of the illustrations, and thank Dominique Soldati for providing a manuscript before publication. References 1 King, C.A. (1988) Cell motility of sporozoan protozoa. Parasitol. Today 4, 315–319 2 Russell, D.G. (1983) Host cell invasion by Apicomplexa: expression of the parasite’s contractile system? Parasitology 87, 199–209 3 Sibley, L.D. et al. (1998) Gliding motility: an efficient mechanism for cell penetration. Curr. Biol. 8, R12–R14 4 Naitza, S. et al. (1998) The thrombospondin-related protein family of Apicomplexan parasites: the gears of the cell invasion machinery. Parasitol. Today 14, 479–484 5 Sibley, L.D. (1995) Invasion of vertebrate cells by Toxoplasma gondii. Trends Cell Biol. 5, 129–132 6 Gratzer, W.B. and Dluzewski, A.R. (1993) The red blood cell and malaria parasite invasion. Semin. Hematol. 30, 232–247 7 Miller, L.H. et al. (1979) Interaction between cytochalasin B treated malarial parasites and erythrocytes. Attachment and junction formation. J. Exp. Med. 149, 172–184 8 Field, S.J. et al. (1993) Actin in the merozoite of the malaria parasite, Plasmodium falciparum. Cell Motil. Cytoskeleton 25, 43–48 9 Hudson-Taylor, D.E. et al. (1995) Plasmodium falciparum protein associated with the invasion junction contains a conserved oxidoreductase domain. Mol. Microbiol. 15, 463–471 10 Dluzewski, A.R. et al. (1989) Red cell membrane protein distribution during malarial invasion. J. Cell Sci. 92, 691–699 11 Aikawa, M. et al. (1978) Erythrocyte entry by malaria parasites. A moving junction between erythrocyte and parasite. J. Cell Biol. 77, 72–82 12 Ward G.E. et al. (1993) The origins of parasitophorous vacuole membrane lipids in malaria-infected erythrocytes. J. Cell Sci. 106, 237–248

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