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Chapter 3 : Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages

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Abstract:

This chapter briefly outlines the cellular architecture of the blood stages of , for which there is a considerable body of molecular data. The asexual forms, which dominate the relationship between the human host and parasite in terms of time spent and pathological impact, are traditionally classified by their detailed light microscopic features as seen in Giemsa-stained blood films. As the parasite continues to grow and differentiate, it exports membranes and other structures into the surrounding RBC. In , these include membranes-the clefts of Maurer, circular clefts, small vesicles, and dense protein-containing projections from the surface of the RBC, termed knobs. Similar structures exist in other species, such as Schüffner’s dots (cleft-like membranes) in and caveolae analogous to knobs in and , although these are invaginations of the membrane rather than protrusions. In this chapter, the authors follow the terminology of Atkinson and Aikawa (1990) to distinguish the circular clefts from the (short) clefts of Maurer, avoiding the name term tubulovesicular network (TVN) because of its alternative usage to describe the complete system of exported membranes. The brief description of the morphological changes occurring within the parasite during the asexual and sexual erythrocytic periods indicates the great complexity and continual modulation of cellular form typical of , a statement which can be repeated for the other species of malaria parasite and other stages not considered here.

Citation: Bannister L, Margos G, Hopkins J. 2005. Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages, p 24-49. In Sherman I (ed), Molecular Approaches to Malaria. ASM Press, Washington, DC. doi: 10.1128/9781555817558.ch3

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Figures

Image of FIGURES 1 TO 8
FIGURES 1 TO 8

Merozoite ultrastructure of P. falciparum. Figure 1 is a scanning EM of a merozoite attached to an RBC, demonstrating the relative sizes of the two cells. Figure 2 is a transmission EM of a longitudinal section through a free merozoite, showing a number of organelles. In Fig. 3, typical mature rhoptries are present within merozoites shortly to be released from a schizont. Figure 4 and insets within it show the apical region of a merozoite and the three types of apical organelles (magnifications in all images are the same). In Fig. 5, a merozoite has been freeze fractured to show numerous intramembranous particles in the two rhoptry membrane faces, representing intramembranous protein domains. In Fig. 6 and 7, the merozoite pellicle and adjacent structures are shown at a higher magnification; in Fig. 6, the section includes the nuclear envelope as well as the two membranes of the IMC, the merozoite plasma membrane. In Fig. 7, the close relation of the apicoplast to the subpellicular microtubules and mitochondrion is seen in transverse section. Figure 8 shows a freeze-fracture preparation of a merozoite, revealing the different membrane faces and intramembranous particle distributions of the pellicle including the plasma membrane and IMC. Mitoch, mitochondrion; Mt, microtubules; Mz, merozoite; PM, plasma membrane.

Citation: Bannister L, Margos G, Hopkins J. 2005. Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages, p 24-49. In Sherman I (ed), Molecular Approaches to Malaria. ASM Press, Washington, DC. doi: 10.1128/9781555817558.ch3
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Image of FIGURES 9 TO 14
FIGURES 9 TO 14

Ring-stage ultrastructure. Figure 9 shows an early ring and RBC at low magnification, the parasite curved into a cup-like form in this example. In Fig. 10, a larger ring shows the development of ribosomes and RER and the accumulation of hemozoin-containing vacuoles. In Fig. 11, a ring stage situated adjacent to the RBC membrane contains more highly developed endoplasmic reticulum, a mitochondrion, and a large vacuole. Figure 12 details part of the ring surface at high magnification, showing three double-membraned vesicles (arrows). In Fig. 13, the zone of coated vesicle budding from the nuclear envelope is seen on the left and a series of double-membrane vesicles lies close to the parasite's surface (white arrows). In Fig. 14, detail of a maturing ring, with a complex network of RER, a mitochondrion in transverse section, and a dense hemozoin-containing vacuole are shown. Hz, hemozoin; Mitoch, mitochondrion; vac, large vacuole.

Citation: Bannister L, Margos G, Hopkins J. 2005. Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages, p 24-49. In Sherman I (ed), Molecular Approaches to Malaria. ASM Press, Washington, DC. doi: 10.1128/9781555817558.ch3
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Image of FIGURES 15 TO 17
FIGURES 15 TO 17

Trophozoite stage ultrastructure. Figure 15 shows the arrangement of organelles in a mature trophozoite. The parasite contains a large pigment vacuole with lysing food vacuoles and hemozoin within it and other structures as detailed in Fig. 18 to 26. Figure 16 is a scanning EM of a trophozoite-infected RBC, showing the characteristic irregular surface, including small knob-like protruberances. In Fig. 17, a more mature trophozoite is shown, with a thin layer of cytoplasm superficially resembling a circular cleft delimiting a large mass of partially enclosed RBC cytosol (white arrow). Hz, hemozoin; Mitoch, mitochondrion.

Citation: Bannister L, Margos G, Hopkins J. 2005. Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages, p 24-49. In Sherman I (ed), Molecular Approaches to Malaria. ASM Press, Washington, DC. doi: 10.1128/9781555817558.ch3
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Image of FIGURES 18 TO 26
FIGURES 18 TO 26

Details of trophozoite ultrastructure. In Fig. 18, a branched complex series of RER cisternae are seen to be continuous with the nuclear envelope. Two Maurer's clefts are also visible. Figure 19 shows the Golgi complex of a mature trophozoite, where an extension of the nuclear envelope is the site of multiple coated vesicle budding. A tubular structure, corresponding to a Golgi cisterna, is closely associated with this mass of vesicles. In Fig. 20, the close association between the two-membrane mitochondrion and three-membrane apicoplast is depicted. Figure 21 shows a cytostomal vacuole in the process of formation through the cytostomal ring at the trophozoite's surface. The cytosol of the RBC, with the PVM and parasite's plasma membrane, is seen to be invaginated into the vacuole. In Fig. 22, the pigment vacuole of a maturing trophozoite contains angular crystals of hemozoin. Figures 23 to 26 depict structures exported from the parasite into the RBC. In Fig. 23, a typical Maurer's cleft with an external dense coating in transverse section is present in the RBC cytosol; two surface knobs are shown. In Fig. 24, a Maurer's cleft is visible in oblique section, revealing an irregular plate-like form. The insert shows a small exported vesicle with a spiky surface, present in the RBC cytosol. Figure 25 shows a circular cleft, continuous with the PVM (arrows).This example is more complex internally than usual, because of the presence of smaller circular cleft membranes within it. Figure 26 depicts a tangential section through the surface of an infected RBC, showing a high density of knobs associated with its membrane, a configuration typical of a multiply infected RBC. Mitoch, mitochondrion; PM, parasite plasma membrane.

Citation: Bannister L, Margos G, Hopkins J. 2005. Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages, p 24-49. In Sherman I (ed), Molecular Approaches to Malaria. ASM Press, Washington, DC. doi: 10.1128/9781555817558.ch3
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Image of FIGURES 27 TO 33
FIGURES 27 TO 33

Ultrastructure of schizont development. Figure 27 shows an early, two-nucleus schizont, one of the nuclei with a part of a mitotic spindle. Also visible are a cytostomal vacuole formed at the end of a deep intrusion of RBC cytoplasm into the parasite's surface (asterisk) and a large pigmented vacuole containing a small cluster of hemozoin crystals. The surface of the infected RBC shows numerous knobs. Figure 28 shows a more advanced schizont at approximately the eight-nucleus stage with spheroidal rhoptries in pairs around the periphery indicating the positions of future merozoite apices. Also present is a prominent lipid body. Note the absence of knobs from the surface of the RBC, as expected from this knobless strain (C10) of parasite, contrasting with the IT04 strain depicted in Fig. 27. Figure 29 shows a schizont immediately after the end of nuclear division, with the beginning of merozoite budding from the central mass (residual body) containing the hemozoin. Note that the rhoptries are now elongated club-like forms approaching their mature state. Figure 30 shows merozoites, here elongated and connected only by narrow stalks to the residual body, but the PVM and RBC hemoglobin are still intact. Figure 31 shows a schizont with the merozoites now separate from the residual body, surrounded by only a single membrane after the hemoglobin has been released. In Fig. 32 and 33, scanning EMs of schizonts from which the membranes of the RBC and PV have been mechanically removed during specimen preparation are shown. Figure 32 shows an early budding stage, with merozoite apices protruding from the residual body; Fig. 33 depicts a rather later stage with two more mature clusters of merozoites, probably representing a double RBC infection. Circl, circular cleft; Hz, hemozoin crystals; Pvac, pigment vacuole.

Citation: Bannister L, Margos G, Hopkins J. 2005. Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages, p 24-49. In Sherman I (ed), Molecular Approaches to Malaria. ASM Press, Washington, DC. doi: 10.1128/9781555817558.ch3
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Image of FIGURES 34 TO 37
FIGURES 34 TO 37

Details of merozoite development. In Fig. 34, the mitotic spindle and associated SPBs and early rhoptry centers (black arrows) are situated in an early (four-nucleus) schizont. In Fig. 35, at a late final divisional state (8 nuclei, progressing to 16) merozoite apices with pairs of rhoptries are developing at either end of a dividing nucleus (the spindle is not visible in this specimen). In each pair, one rhoptry is typically more mature than the other. Pellicles are assembled around the merozoite apices, giving their membranes a dense multilayered appearance (white arrows). Figure 36 shows two merozoites at a late schizont stage, although they are still attached to the residual body. Coated vesicles are budding from the nuclear envelope close to the SPBs, and a Golgi cisterna lies more apically near each, enlarged on the right to form a rounded vesicle close to a developing rhoptry. In Fig. 37, the apical migration of micronemes along a subpellicular microtubule is shown in a longitudinal section of a late merozoite. The section also passes through the edge of the apical prominence and its polar rings. Mt, subpellicular microtubule; PolR, polar rings. Figures 35 and 36 are reproduced from Bannister et al., 2000, by kind permission of Cambridge University Press. Figure 37 is reproduced from Bannister et al., 2003, by kind permission of the Journal of Cell Science.

Citation: Bannister L, Margos G, Hopkins J. 2005. Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages, p 24-49. In Sherman I (ed), Molecular Approaches to Malaria. ASM Press, Washington, DC. doi: 10.1128/9781555817558.ch3
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Image of FIGURES 38 TO 48
FIGURES 38 TO 48

Gametocyte ultrastructure. Figure 38 shows a stage V macrogametocyte in longitudinal section. Hemozoin crystals (Hz) are present around the nucleus, and osmiophilic bodies (Osb) situated around the parasite's periphery. Clusters of RER are present at either end. Figure 39 depicts a transverse section through an RBC containing a mature gametocyte; RER is present in the parasite's cytoplasm. On the left, the RBC is flattened into a flap-like extension (Laveran's bib) to one side; around the parasite, the hemoglobin is beginning to disappear. Figure 40 details the surface region of a mature gametocyte and adjacent RBC. Visible is the PVM closely adhering to the three-membraned pellicle of the gametocyte. An osmiophilic body (Osb) with a narrow stalk is in contact with the pellicle, which also contains a multilamellar membranous structure (Lam). Note that a hemoglobin-free zone is present immediately outside the PVM. In Fig. 41, a higher magnification of a group of osmiophilic bodies, one of them with a duct-like extension, is shown. Figure 42 shows a mitochondrion in longitudinal section with tubular cristae. Figure 43 shows the transverse section of an apicoplast. In Fig. 44, a multilamellar membranous inclusion in the nuclear envelope is shown. Figure 45 shows the transverse section of a macrogametocyte nucleus containing a nucleolus-like body. In Fig. 46, the transverse section of a stage IV gametocyte is shown with a band of subpellicular microtubules (Mt) and vacuolar structures. Gametocyte-specific membranous clefts are also visible in the RBC. In Fig. 47, higher magnification through the surface region of a stage IV gametocyte shows the pellicle with a band of subpellicular microtubules (Mt), some of them doublets or triplets, attached to the pellicle. Two transversely sectioned mitochondrial profiles are also present. The PVM encloses the gametocyte. In Fig. 48, a higher magnification of part of Fig. 47 shows details of the subpellicular microtubules and their pellicle attachments. Mitoch, mitochondrion; Nlb, nucleolus-like body; Stic, sexual-stage tubular intraerythrocytic compartment.

Citation: Bannister L, Margos G, Hopkins J. 2005. Making a Home For Post-Genomics: Ultrastructural Organization of the Blood Stages, p 24-49. In Sherman I (ed), Molecular Approaches to Malaria. ASM Press, Washington, DC. doi: 10.1128/9781555817558.ch3
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References

/content/book/10.1128/9781555817558.chap3
1. Adisa, A.,, M. Rug,, M. Foley,, and L. Tilley. 2002. Characterisation of a delta-COP homologue in the malaria parasite, Plasmodium falciparum. Mol. Biochem. Parasitol. 123: 11- 21.
2. Aikawa, M. 1967. Ultrastructure of the pellicular complex of Plasmodium fallax. J. Cell Biol. 35: 103 113.
3. Aikawa, M. 1971. Plasmodium: the fine structure of malarial parasites. Exp. Parasitol. 30: 284 320.
4. Aikawa, M.,, and T. M. Seed,. 1980. Morphology of plasmodia, p. 285 344. In J. P. Kreier (ed.), Malaria, vol. 1. Academic Press, New York, N.Y.
5. Aikawa, M.,, C. G. Huff,, and H. Sprinz. 1969. Comparative fine structure study of the gametocytes of avian, reptilian, and mammalian malarial parasites. J. Ultrastruct. Res. 26: 316 331.
6. Aikawa, M.,, L. H. Miller,, J. Johnson,, and J. Rabbege. 1978. Erythrocyte entry by malarial parasites. A moving junction between erythrocyte and parasite. J. Cell Biol. 77: 72 82.
7. Aikawa, M.,, L. H. Miller,, J. R. Rabbege,, and N. Epstein. 1981. Freeze-fracture study on the erythrocyte membrane during malarial parasite invasion. J. Cell Biol. 91: 55 62.
8. Aikawa, M.,, J. R. Rabbege,, I. Udeinya,, and L. H. Miller. 1983. Electron microscopy of knobs in Plasmodium falciparum-infected erythrocytes. J. Parasitol. 69: 435 437.
9. Aikawa, M.,, R. Carter,, Y. Ito,, and M. M. Nijhout. 1984. New observations on gametogenesis, fertilization, and zygote transformation in Plasmodium gallinaceum. J. Protozool. 31: 403 413.
10. Aikawa, M.,, K. Kamanura,, S. Shiraishi,, Y. Matsumoto,, H. Arwati,, M. Torii,, Y. Ito,, T. Takeuchi,, and B. Tandler. 1996. Membrane knobs of unfixed Plasmodium falciparum infected erythrocytes: new findings as revealed by atomic force microscopy and surface potential spectroscopy. Exp. Parasitol. 84: 339 343.
11. Akaki, M.,, E. Nagayasu,, Y. Nakano,, and M. Aikawa. 2002. Surface charge of Plasmodium falciparum merozoites as revealed by atomic force microscopy with surface potential spectroscopy. Parasitol. Res. 88: 16 20.
12. Alano, P.,, D. Read,, M. Bruce,, M. Aikawa,, T. Kaido,, T. Tegoshi,, S. Bhatti,, D. K. Smith,, C. Luo,, S. Hansra,, R. Carter,, and J. F. Elliott. 1995. COS cell expression cloning of Pfg377, a Plasmodium falciparum gametocyte antigen associated with osmiophilic bodies. Mol. Biochem. Parasitol. 74: 143 156.
13. Atkinson, C.T.,, and M. Aikawa. 1990. Ultrastructure of malaria-infected erythrocytes. Blood Cells 16: 351 368.
14. Bannister, L. H.,, and G. H. Mitchell. 1986. Lipidic vacuoles in Plasmodium knowlesi erythrocytic schizonts. J. Protozool. 33: 271 275.
15. Bannister, L. H.,, and A. R. Dluzewski. 1990. The ultrastructure of red cell invasion in malaria infections: a review. Blood Cells 16: 257 292.
16. Bannister, L. H.,, and G. H. Mitchell. 1995. The role of the cytoskeleton in Plasmodium falciparum merozoite biology: an electron-microscopic view. Ann. Trop. Med. Parasitol. 89: 105 111.
17. Bannister, L. H.,, G. A. Butcher,, E.D. Dennis,, and G. H. Mitchell. 1975. Structure and invasive behaviour of Plasmodium knowlesi merozoites in vitro. Parasitology 71: 483 491.
18. Bannister, L. H.,, J. M. Hopkins,, R. E. Fowler,, S. Krishna,, and G. H. Mitchell. 2000. Ultrastructure of rhoptry development in Plasmodium falciparum erythrocytic merozoites. Parasitology 121: 273 287.
19. Bannister, L. H.,, J. M. Hopkins,, R. E. Fowler,, S. Krishna,, and G. H. Mitchell. 2001. A brief illustrated guide to the ultrastructure of Plasmodium falciparum asexual blood stages. Parasitol. Today 16: 427 433.
20. Bannister, L. H.,, J. M. Hopkins,, A. R. Dluzewski,, G. Margos,, I. T. Williams,, M. J. Blackman,, C. H. Kocken,, A. W. Thomas,, and G. H. Mitchell. 2003. Plasmodium falciparum apical membrane antigen 1 (PfAMA-1) is translocated within micronemes along subpellicular microtubules during merozoite development. J. Cell Sci. 116: 3825 3834.
21. Bannister, L. H.,, J. M. Hopkins, , G. Margos,, A. R. Dluzewski,, and G. H. Mitchell. 2004. Three-dimensional ultrastructure of the ring stage of Plasmodium falciparum: evidence for export pathways. Microsc. Microanal. 10: 551 562.
22. Bushell, G. R.,, L. T. Ingram,, C. A. Fardoulys,, and J. A. Cooper. 1988. An antigenic complex in the rhoptries of Plasmodium falciparum. Mol. Biochem. Parasitol. 28: 105 112.
23. Chishti, A. H.,, K. I. Andrabi,, L. H. Derick,, J. Palek,, and S. C. Liu. 1992. Isolation of skeleton-associated knobs from human red blood cells infected with malaria parasite Plasmodium falciparum. Mol. Biochem. Parasitol. 52: 283 287.
24. Cooke, B. M.,, K. M. Lingelbach,, L. H. Bannister,, and L. Tilley. 2004. Protein trafficking in Plasmodium falciparum-infected red blood cells. Trends Parasitol. 20: 581 589.
25. Culvenor, J. G.,, K. P. Day,, and R. F. Anders. 1991. Plasmodium falciparum ring-infected erythrocyte surface antigen is released from merozoite dense granules after erythrocyte invasion. Infect. Immun. 59: 1183 1187.
26. Divo, A. A.,, T. G. Geary,, J. B. Jensen,, and H. Ginsburg. 1985. The mitochondrion of Plasmodium falciparum visualized by rhodamine 123 fluorescence. J. Protozool . 32: 442 446.
27. Eksi, S.,, A. Stump,, S. L. Fanning,, M. I. Shenouda,, H. Fujioka,, and K. C. Williamson. 2002. Targeting and sequestration of truncated Pfs230 in an intraerythrocytic compartment during Plasmodium falciparum gametocytogenesis. Mol. Microbiol. 44: 1507 1516.
28. el Shoura, S. M.,, and O. M. al Amari. 1993. Falciparum malaria in naturally infected human patients. I. Ultrastructural differences between malaria pigments in intraerythrocytic asexual and sexual forms. J. Morphol. 215: 201 206.
29. Elford, B. C.,, G. M. Cowan,, and D. J. P. Ferguson. 2004. Parasite-regulated membrane transport processes and metabolic control in malaria-infected erythrocytes. Biochem. J. 308: 361 374.
30. Etzion, Z.,, M. C. Murray,, and M. E. Perkins. 1991. Isolation and characterization of rhoptries of Plasmodium falciparum. Mol. Biochem. Parasitol. 47: 51 61.
31. Florens, L.,, M. P. Washburn,, J. D. Raine,, R. M. Anthony,, M. Grainger,, J. D. Haynes,, J. K. Moch,, N. Muster,, J. B. Sacci,, D. L. Tabb,, A. A. Witney,, D. Wolters,, Y. Wu,, M. J. Gardner,, A. A. Holder,, R. E. Sinden,, J. R. Yates,, and D. J. Carucci. 2002. A proteomic view of the Plasmodium falciparum life cycle. Nature 419: 520 526.
32. Florent, I.,, S. Charneau,, and P. Grellier. 2004. Plasmodium falciparum genes differentially expressed during merozoite morphogenesis. Mol. Biochem.Parasitol. 135: 143 148.
33. Fowler, R. E.,, A. M. Smith,, J. Whitehorn,, I. T. Williams,, L. H. Bannister,, and G. H. Mitchell. 2001. Microtubule associated motor proteins of Plasmodium falciparum merozoites. Mol. Biochem. Parasitol. 117: 187 200.
34. Garcia, C. R.,, M. Takeuschi,, K. Yoshioka,, H. Miyamoto. 1997. Imaging Plasmodium falciparum-infected ghost and parasite by atomic force microscopy. J. Struct. Biol. 119: 92 98.
35. Garnham, P. C. C. 1966. Malaria Parasites and Other Haemosporidia. Blackwell, Oxford, United Kingdom.
36. Guinet, F.,, J. A. Dvorak,, H. Fujioka,, D. B. Keister,, O. Muratova,, D. C. Kaslow,, M. Aikawa,, A. B. Vaidya,, and T. E. Wellems. 1996. A developmental defect in Plasmodium falciparum male gametogenesis. J. Cell Biol. 135: 269 278.
37. Haldar, K.,, B. U. Samuel,, N. Mohandas,, T. Harrison,, and N. L. Hiller. 2001. Transport mechanisms in Plasmodium-infected erythrocytes: lipid rafts and a tubovesicular network. Int. J. Parasitol. 31: 1393 1401.
38. Healer, J.,, S. Crawford,, S. Ralph,, G. McFadden,, and A. F. Cowman. 2002. Independent translocation of two micronemal proteins in developing Plasmodium falciparum merozoites. Infect. Immun. 70: 5751 5758.
39. Heidrich, H. G.,, M. Matzner,, A. Miettinen-Baumann,, and W. Strych. 1986. Immunoelectron microscopy shows that the 80,000-dalton antigen of Plasmodium falciparum merozoites is localized in the surface coat. Z. Parasitenkd. 72: 681 683.
40. Hempelmann, E.,, C. Motta,, R. Hughes,, S. A. Ward,, and P. Bray. 2004. Plasmodium falciparum: sacrificing membrane to grow crystals? Trends Parasitol. 19: 23 26.
41. Hopkins, J. M.,, R. E. Fowler,, S. Krishna,, I. Wilson,, G. H. Mitchell,, and L. H. Bannister. 1999. The plastid in Plasmodium falciparum asexual blood stages: a three-dimensional ultrastructural analysis. Protista. 150: 283 295.
42. Jackson, K. E.,, N. Klonis,, D. J. P. Ferguson,, A. Adisa,, C. Dogovski,, and L. Tilley. 2004. Food vacuole-associated lipid bodies and heterogeneous lipid environments in the malaria parasite, Plasmodium falciparum. Mol. Microbiol. 54: 109 122.
43. Jaikaria, N. S.,, C. Rozario,, R. G. Ridley,, and M. E. Perkins. 1993. Biogenesis of rhoptry organelles in Plasmodium falciparum. Mol. Biochem. Parasitol. 57: 269 279.
44. Johnson, D.,, K. Gunther,, I. Ansorge,, J. Benting,, A. Kent,, L. Bannister,, R. Ridley,, and K. Lingelbach. 1994. Characterization of membrane proteins exported from Plasmodium falciparum into the host erythrocyte. Parasitology 109: 1 9.
45. Kaidoh, T.,, J. Nath,, H. Fujioka,, V. Okoye,, and M. Aikawa. 1995. Effect and localization of trifluralin in Plasmodium falciparum gametocytes: an electron microscopic study. J. Eukaryot. Microbiol. 42: 61 64.
46. Kaidoh, T.,, J. Nath,, V. Okoye,, and M. Aikawa. 1993. Novel structure in the pellicular complex of Plasmodium falciparum gametocytes. J. Eukaryot. Microbiol. 40: 269 271.
47. Kass, L.,, D. Willerson, Jr.,, K. H. Rieckmann,, P. E. Carson,, and R. P. Becker. 1971. Plasmodium falciparum gametocytes. Electron microscopic observations on material obtained by a new method. Am. J. Trop. Med. Hyg. 20: 187 194.
48. Kohler, S.,, C. F. Delwiche,, P. D. Denny,, L. G. Tilney,, P. Webster,, R. J. M. Wilson,, J. D. Palmer,, and D. S. Roos. 1997. A plastid of probable green algal origin in apicomplexan parasites. Science 275: 1485 1487.
49. Kriek, N.,, L. Tilley,, P. Horrocks,, R. Pinches,, B. C. Elford,, D. J. P. Ferguson,, K. Lingelbach,, and C. Newbold. 2003. Characterization of the pathway for transport of the cytoadherence-mediating protein, PfEMP1, to the host cell surface in malaria parasite-infected erythrocytes. Mol. Microbiol. 50: 1215 1227.
50. Krungkrai, J.,, P. Prapunwattana,, and S. R. Krungkrai. 2000. Ultrastructure and function of mitochondria in gametocytic stage of Plasmodium falciparum. Parasite 7: 19 26.
51. Ladda, R. L. 1969. New insights into the fine structure of rodent malarial parasites. Mil. Med. 134: 825 865.
52. Langreth, S. G.,, J. B. Jensen,, R. T. Reese,, and W. Trager. 1978. Fine structure of human malaria in vitro. J. Protozool. 25: 443 452.
53. Ling, I. T.,, L. Florens,, A. R. Dluzewski,, O. Kaneko,, M. Grainger,, B. Y. S. Yim Lim,, T. Tsuboi,, J. M. Hopkins,, J. R. Johnson,, M. Torii,, L. H. Bannister,, I. I. I. Yates, Jr.,, A. A. Holder,, and D. Mattei. 2004. The Plasmodium falciparum clag9 gene encodes a rhoptry protein that is transferred to the host erythrocyte upon invasion. Mol. Microbiol. 52: 107 118.
54. Lingelbach, K.,, and K. A. Joiner. 1998. The parasitophorous vacuole membrane surrounding Plasmodium and Toxoplasma: an unusual compartment in infected cells. J. Cell Sci. 111: 1467 1475.
55. Margos, G.,, L. H. Bannister,, A. R. Dluzewski,, J. M. Hopkins,, I. T. Williams,, and G. H. Mitchell. 2004. Correlation of structural development and differential expression of invasion-related molecules in schizonts of Plasmodium falciparum. Parasitology 129: 273 287.
56. McLaren, D. J.,, L. H. Bannister,, P. I. Trigg,, and G. A. Butcher. 1979. Freeze fracture studies on the interaction between the malaria parasite and the host erythrocyte in Plasmodium knowlesi infections. Parasitology 79: 125 139.
57. Meszoely, C. A.,, E. F. Erbe,, R. L. Steere,, J. Trosper,, and R. L. Beaudoin. 1987. Plasmodium falciparum: freeze-fracture of the gametocyte pellicular complex. Exp. Parasitol. 64: 300 309.
58. Mitchell, G. H.,, and L. H. Bannister. 1988. Malaria parasite invasion: interactions with the red cell membrane. Crit. Rev.Oncol. Hematol. 8: 225 310.
59. Mitchell, G. H.,, A. W. Thomas,, G. Margos,, A. R. Dluzewski,, and L. H. Bannister. 2004. Apical membrane antigen 1, a major malaria vaccine candidate, mediates the close attachment of invasive merozoites to host red blood cells. Infect. Immun. 72: 154 158.
60. Morrissette, N. S.,, J. M. Murray,, and D. S. Roos. 1997. Subpellicular microtubules associate with an intramembranous particle lattice in the protozoan parasite Toxoplasma gondii. J. Cell Sci. 110: 35 42.
61. Nagao, E.,, O. Kaneko,, and J. A. Dvorak. 2000. Plasmodium falciparum-infected erythrocytes: qualitative and quantitative analyses of parasite-induced knobs by atomic force microscopy. J. Struct. Biol. 130: 34 44.
62. Noe, A. R.,, D. J. Fishkind,, and J. H. Adams. 2000. Spatial and temporal dynamics of the secretory pathway during differentiation of the Plasmodium yoelii schizont. Mol. Biochem. Parasitol. 108: 169 185.
63. Noland, G. S.,, N. Briones,, and D. J. Sullivan, Jr. 2003. The shape and size of hemozoin crystals distinguishes diverse Plasmodium species. Mol. Biochem. Parasitol. 130: 91 99.
64. Ponnudurai, T.,, A. H. Lensen,, J. F. Meis,, and J. H. Meuwissen. 1986. Synchronization of Plasmodium falciparum gametocytes using an automated suspension culture system. Parasitology 93: 263 274.
65. Prensier, G.,, and C. Slomianny. 1986. The karyotype of Plasmodium falciparum determined by ultrastructural serial sectioning and 3D reconstruction. J. Parasitol. 72: 731 736.
66. Przyborski, J. M.,, H. Wickert,, G. Krohne,, and M. Lanzer. 2003. Maurer's clefts—a novel secretory organelle? Mol. Biochem. Parasitol. 132: 17 26.
67. Roger, N.,, J. F. Dubremetz,, P. Delplace,, B. Fortier,, G. Tronchin,, and A. Vernes. 1988. Characterization of a 225 kilodalton rhoptry protein of Plasmodium falciparum. Mol. Biochem. Parasitol. 27: 135 141.
68. Ruangjirachuporn, W.,, B. A. Afzelius,, H. Helmby,, A. V. S. Hill,, B. M. Greenwood,, J. Carlson,, K. Berzins,, P. Perlmann,, and M. Wahlgren. 1992. Ultrastructural analysis of fresh Plasmodium falciparum- infected erythrocytes and their cytoadherence to human leukocytes. Am. J. Trop. Med. Hyg. 46: 511 519.
69. Ruiz, F. A.,, S. Luo,, S. N. Moreno,, and R. Docampo,. 2004. Polyphosphate content and fine structure of acidocalcisomes of Plasmodium falciparum. Microsc. Microanal. 10: 563 567.
70. Russell, D. G.,, and R. G. Burns. 1984. The polar ring of coccidian protozoans: a unique microtubule-organizing centre. J. Cell Sci. 65: 193 207.
71. Salmon, B. L.,, A. Oksman,, and D. E. Goldberg. 2001. Malaria parasite exit from the host erythrocyte: a two-step process requiring extraerythrocytic proteolysis. Proc. Nat. Acad. Sci. USA 98: 271 276.
72. Sam-Yellowe T. Y.,,, L. Florens,, T. Wang,, J. D. Raine,, D. J. Carucci,, R. Sinden,, and I. I. I. Yates, Jr. 2004. Proteome analysis of rhoptry-enriched fractions isolated from plasmodium merozoites. J. Proteome Res. 3: 995 1001.
73. Scherf, A.,, R. Carter,, C. Petersen,, P. Alano,, R. Nelson,, M. Aikawa,, D. Mattei,, D. S. Pereira,, and J. Leech. 1992. Gene inactivation of Pf11–1 of Plasmodium falciparum by chromosome breakage and healing: identification of a gametocyte-specific protein with a potential role in gametogenesis. EMBO J 11: 2293 2301.
74. Shaw, M. K.,, J. Thompson,, and R. E. Sinden. 1996. Localization of ribosomal RNA and Pbs21- mRNA in the sexual stages of Plasmodium berghei using electron microscope in situ hybridization. Eur. J. Cell Biol. 71: 270 276.
75. Sinden, R. E. 1982a. Gametocytogenesis of Plasmodium falciparum in vitro: an electron microscopic study. Parasitology. 84: 1 11.
76. Sinden, R. E. 1982b. Gametocytogenesis of Plasmodium falciparum in vitro: ultrastructural observations on the lethal action of chloroquine. Ann. Trop. Med. Parasitol. 76: 15 23.
77. Sinden, R. E.,, E. U. Canning,, and B. J. Spain. 1976. Gametogenesis and fertilization in Plasmodium yoelii nigeriensis: a transmission electron microscope study. Proc. R. Soc. Lond. B Biol. Sci. 193: 55 76.
78. Sinden, R. E.,, R. H. Hartley,, and N. J. King. 1985. Gametogenesis in Plasmodium; the inhibitory effects of anticytoskeletal agents. Int. J. Parasitol. 15: 211 217.
79. Slomianny, C. 1990. Three-dimensional reconstruction of the feeding process of the malaria parasite. Blood Cells 16: 669 378.
80. Slomianny, C.,, G. Prensier,, and P. Charet. 1985. Ingestion of erythrocytic stroma by Plasmodium chabaudi trophozoites: ultrastructural study by serial sectioning and 3-dimensional reconstruction. Parasitology 90: 579 588.
81. Taraschi, T. F. 1999. Macromolecular transport in malaria-infected erythrocytes. Novartis Found. Symp. 226: 114 120.
82. Taraschi, T. F.,, D. Trelka,, T. Schneider,, and I. Matthews. 1998. Plasmodium falciparum: characterization of organelle migration during merozoite morphogenesis in asexual malaria infections. Exp. Parasitol. 88: 184 193.
83. Thomas, A. W.,, L. H. Bannister,, and A. P. Waters. 1990. Sixty-six kilodalton-related antigens of Plasmodium knowlesi are merozoite surface antigens associated with the apical prominence. Parasite Immunol. 12: 105 113.
84. Topolska, A. E.,, L. Wang,, C. G. Black,, and R. L. Coppel,. 2004. Merozoite cell biology, p. 200 215. In A. P. Waters, and C. J. Janse (ed.), Malaria Parasites: Genomes and Molecular Biology. Caister Academic Press, New York, N.Y.
85. Torii, M.,, J. H. Adams,, L. H. Miller,, and M. Aikawa. 1989. Release of merozoite dense granules during erythrocyte invasion by Plasmodium knowlesi. Infect. Immun. 57: 3230 3233.
86. Van Wye, J.,, N. Ghori,, P. Webster,, R. R. Mitschler,, H. G. Elmendorf,, and K. Haldar. 1996. Identification and localization of rab6, separation of rab6 from ERD2 and implications for an ‘unstacked’ Golgi, in Plasmodium falciparum. Mol. Biochem. Parasitol. 83: 107 120.
87. Vickerman, K.,, and F. E. G. Cox. 1967. Merozoite formation in the erythrocytic stages of the malaria parasite Plasmodium vinckei. Trans. R. Soc.Trop. Med. Hyg. 61: 303 312.
88. Vielemeyer, O.,, M. T. McIntosh,, K. A. Joiner,, and I. Coppens. 2004. Neutral lipid synthesis and storage in the intraerythrocytic stages of Plasmodium falciparum. Mol. Biochem. Parasitol. 135: 197 209.
89. Waller, R. F.,, and G. I. McFadden. 2005. The apicoplast: a review of the derived plastid of apicomplexan parasites. Curr. Issues Mol. Biol. 7: 57 80.
90. Wickert, H.,, F. Wissing,, K. T. Andrews,, A. Stich,, G. Krohne,, and M. Lanzer. 2003. Evidence for trafficking of PfEMP1 to the surface of P. falciparum-infected erythrocytes via a complex membrane network. Eur. J. Cell Biol. 82: 271 284.
91. Wickham, M. E.,, J. G. Culvenor,, and A. F. Cowman. 2003. Selective inhibition of a two-step egress of malaria parasites from the host erythrocyte. J. Biol. Chem. 278: 37658 37663.
92. Winograd, E.,, C. A. Clavijo,, L. Y. Bustamante,, and M. Jaramillo. 1999. Release of merozoites from Plasmodium falciparum-infected erythrocytes could be mediated by a non-explosive event. Parasitol. Res. 85: 621 624.
93. Wunderlich, F.,, H. Stubig,, and E. Konigk. 1982. Development of Plasmodium chabaudi in mouse red blood cells: structural properties of the host and parasite membranes. J. Protozool. 29: 60 66.
94. Yayon, A.,, R. Timberg,, S. Friedman,, and H. Ginsburg. 1984. Effects of chloroquine on the feeding mechanism of the intraerythrocytic human malarial parasite Plasmodium falciparum. J. Protozool. 31: 367 372.

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