Figures
Abstract
Apicomplexans are pathogens responsible for malaria, toxoplasmosis, and crytposporidiosis in humans, and a wide range of livestock diseases. These unicellular eukaryotes are stealthy invaders, sheltering from the immune response in the cells of their hosts, while at the same time tapping into these cells as source of nutrients. The complexity and beauty of the structures formed during their intracellular development have made apicomplexans the darling of electron microscopists. Dramatic technological progress over the last decade has transformed apicomplexans into respectable genetic model organisms. Extensive genomic resources are now available for many apicomplexan species. At the same time, parasite transfection has enabled researchers to test the function of specific genes through reverse and forward genetic approaches with increasing sophistication. Transfection also introduced the use of fluorescent reporters, opening the field to dynamic real time microscopic observation. Parasite cell biologists have used these tools to take a fresh look at a classic problem: how do apicomplexans build the perfect invasion machine, the zoite, and how is this process fine-tuned to fit the specific niche of each pathogen in this ancient and very diverse group? This work has unearthed a treasure trove of novel structures and mechanisms that are the focus of this review.
Citation: Striepen B, Jordan CN, Reiff S, van Dooren GG (2007) Building the Perfect Parasite: Cell Division in Apicomplexa. PLoS Pathog 3(6): e78. https://doi.org/10.1371/journal.ppat.0030078
Editor: B. Brett Finlay, University of British Columbia, Canada
Published: June 29, 2007
Copyright: © 2007 Striepen et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: Our work on apicomplexans is currently supported by grants from the US National Institutes of Health to BS (AI 55268 & 64671), University of Georgia graduate fellowships to SR and CNJ, and a CJ Martin Fellowship from the Australian National Health and Medical Research Council to GGvD.
Competing interests: The authors have declared that no competing interests exist.
Abbreviation: IMC, inner membrane complex
A Lean and Mean Invasion Machine
A wide variety of prokaryotic and eukaryotic pathogens have evolved the ability to invade and replicate within the cells of their hosts. Few have developed the level of sophistication and control exerted by the members of the Apicomplexa [1]. Upon contact with a suitable host cell, apicomplexans can invade within seconds, with minimal apparent disturbance of the infected cell (Figure 1). This process is dependent on actin and myosin and is driven by parasite and not host motility [2,3]. Tightly associated with host cell penetration is the secretion of three distinct parasite organelles: rhoptries, micronemes, and dense granules. Secretion is timed in succession, and secreted proteins play key roles in adhesion, motility and formation, and elaboration of the parasitophorous vacuole, a new cellular compartment established during invasion that the parasite occupies during its intracellular development (see [4,5] for detailed reviews of this process in Toxoplasma and Plasmodium, respectively).
(A) Highly simplified apicomplexan life cycle. Apicomplexans are haplonts, and meiosis (sporogony) immediately follows fertilization. Fertilization might occur within a host cell or extracellularly, giving rise to an oocyst or, less frequently, an invasive stage zygote (ookinete).
(B) Schematic representation of a zoite (not all structures are present in all apicomplexans). AP, apicoplast; AR, apical rings; CC, centrocone; CE, centrosome; CO, conoid; DG, dense granule; ER, endoplasmic reticulum; G, Golgi; IMC, inner membrane complex; MI, mitochondrion; MN, microneme; MT, subpellicular microtubule; NU, nucleus; RH, rhoptry.
(C) Zoites actively invade the cells of their hosts, establishing a specialized parasitophorous vacuole (PV) (in some species the parasite lyses the vacuole and develops freely in the cytoplasm).
The cellular structure of the zoite, the non-replicative extracellular stage, appears streamlined towards one goal: finding and invading the next host cell. Zoites are found at various stages of the apicomplexan life cycle and are the product of asexual as well as sexual replication processes (see Figure 1A for a simplified apicomplexan life cycle). The zoite is highly polarized, with the apical tip containing the organizing center for the subpellicular microtubles that run along the longitudinal axis of the parasite [6]. This axis also polarizes the cell's motility, driving the parasite into host cells with its apex first. In some species, the tip is further elaborated by the conoid, a cytoskeletal structure that is built from a unique, tightly wound tubulin polymer and is extended during invasion and motility [7]. Importantly, the apical end is also the site for rhoptry and microneme secretion, with these organelles tightly packed into the anterior portion of the cell. While the anterior of the zoite is focused on invasion, the rest of cell carries the genetic material and tools to grow and develop once in the host cell, including a nucleus and a single mitochondrion, plastid, and Golgi.
Divide and Conquer
While invasive zoites are similar across the phylum, intracellular stages differ dramatically in size, shape, and architecture (see Figure 2 for a selection of micrographs). The basis for this diversity lies in the flexibility of the apicomplexan cell cycle. Apicomplexans are able to dissociate and variably mix and match three elements that follow each other invariably in most other cells: DNA replication and chromosome segregation, nuclear division, and, lastly, cytokinesis or budding (see Figure 3 for a schematic). While Toxoplasma completes all elements of the cycle after each round of DNA replication, Plasmodium and Sarcocystis forgo cytokinesis and/or nuclear divisions for multiple cycles, forming stages that are multinucleate or contain a single polyploid nucleus (these division modes are also known as endodyogeny, schizogony, and endopolyogeny [8–10]). Dramatic differences in the division mode also occur between different life cycle stages in a single species; asexual stages of Toxoplasma in the cat intestine, for example, divide by endodyogeny and endopolygeny [11]. In each case, however, the development will culminate in the emergence of multiple invasive zoites, which seek new host cells to invade. Apicomplexans of the genus Theileria are a surprising exception to this divide and conquer scenario. Theileria sporozoites remain in the lymphocyte that they initially invade, where they amplify in numbers without resorting to leaving the shelter of the host cell. The key to this trick lies in this parasite's ability to transform the host cell through manipulation of the NFκB pathway. The parasite assembles and activates a mammalian IKK signalosome on its surface, promoting unchecked host cell replication [12,13]. Theileria also interacts with host cell microtubules, enabling these parasites to migrate to, and apparently latch onto, host cell centrosomes. This results in partitioning of parasites into forming daughter cells of the host, exploiting the host's mitotic spindle (see Figures 2 and 3; [12,14]; and D. Dobbelaere, personal communication).
(A) In T. gondii, two daughters are formed during budding. IMC1, red; MORN1, green (reproduced with permission from [32]).
(B) T. gondii. Histone H2, red; IMC3, green (reproduced from [71]).
(C) In Plasmodium falciparum liver schizont, budding results in massive numbers of zoites. Image courtesy of Volker Heussler.
(D) T. gondii, phase contrast image of parasitophorous vacuole harboring multiple tachyzoites.
(E and F) P. falciparum late erythrocyte schizont. Acyl carrier protein (plastid), green. RBC, red blood cell.
(G–I) Sarcocystis neurona. Two intracellular stages with polyploid nuclei, one in interphase and one during mitosis. Tubulin, red.
(J) S. neurona budding. IMC3, green.
(K) A Theileria schizont divides in association with its host cell. Polymorphic immunodominant molecule (parasite surface), green; γ-tubulin (host centrosomes), red. HN, host nucleus. Image courtesy of Dirk Dobbelaere. The DNA dye DAPI is shown in blue throughout. Not to scale.
Schematic outline of cell division by Toxoplasma (endodyogeny), Plasmodium (schizogony), and Sarcocystis (endopolygeny). The Theileria schizont is divided in association with host cell division (HN, host nucleus). DNA, grey; IMC, purple; centrosome, red. Note that a centriole as center of the spindle plaque body has not been clearly demonstrated in P. falciparum. Both Sarcocystis and Theileria develop directly in the host cell cytoplasm, while Toxoplasma and Plasmodium are contained within a parasitophorous vacuole (light blue).
Checkpoints and Master Switches
Initial work using inhibitors of DNA synthesis (e.g., aphidocolin) and microtuble disrupting agents suggested that classical cell cycle checkpoints might be lacking in apicomplexans [15,16], pointing to potentially novel mechanisms of control over their complex cell cycles. However, studies using different blocking agents (thymidine, pyrrolidine dithiocarbamate) and characterization of a series of temperature-sensitive mutants have found that the Toxoplasma cell cycle can be halted at what appear to be specific points, including the G1/S and S/M boundaries [17–19]. Furthermore, genomic and experimental surveys for proteins commonly associated with cell cycle checkpoints have identified numerous candidates, including cyclins and cyclin-dependent kinases in Plasmodium and Toxoplasma [20–23]. An attractive model could suggest the presence of developmentally regulated sets of cell cycle factors resulting in different cell division types, which are in turn controlled by master switches. For example, we could hypothesize that Toxoplasma tachyzoites contain master switches to promote nuclear division following DNA synthesis, and cell division following mitosis. Down-regulation of the nuclear division master switch would result in the multiple rounds of DNA synthesis observed during Sarcocystis endopolygeny, while down-regulation of the cytokinesis master switch would lead to the multinucleated schizonts observed in other stages of the Toxoplasma life cycle and in Plasmodium blood stages. Some initial support for this idea has begun to emerge. A series of homologs of the centrosome-associated NIMA kinase (which, in fungi, controls entry into mitosis and spindle formation) have been shown to be essential for cell cycle progression and survival in Plasmodium by gene targeting studies [24–27] and in Toxoplasma by analysis of temperature-sensitive parasite mutants (M. Gubbels and B. Striepen, unpublished data). NIMA genes appear to be differentially expressed over the Plasmodium life cycle. Nek4, for example, is specifically expressed in the female gametocyte and is required for the initial chromosome duplication in the ookinete (zygote) preceding meiosis [25,26], but is dispensable in other stages.
Counting Chromosomes
A fascinating question when considering the various forms of apicomplexan cell division is, how do parasites keep track of their chromosomes in polyploid stages, and how do they know how many zoites to make upon cytokinesis? The following two observations might be important to consider: the final budding of zoites is invariably associated with a last round of DNA replication and nuclear division, and studies that have used high doses of microtubule disrupting agents have found this to lead to a catastrophic breakdown of the coordination of nuclear division and budding in a variety of species [15,28–30]. This suggests that the mitotic spindle, or its organizing center, controls the number of daughter cells and the site where they are to be formed. Apicomplexans use an intranuclear spindle and maintain the nuclear envelope throughout mitosis. The spindle resides in a dedicated elaboration of the nuclear envelope, the centrocone ([31]; Figure 4A), and interacts with the cytoplasmatic centrosome through an opening of the envelope. Interestingly, recent studies in Toxoplasma and Sarcocystis using antibodies to tubulin and MORN1 (a protein that localizes to the centrocone; see below) have shown that the centrocone is maintained throughout the cell cycle [29,32]. Persistence of the spindle, and persistent kinetochore attachment of chromosomes to the spindle microtubules, would provide a mechanism to maintain the integrity of chromosomal sets through polyploid stages [29]; however, this hypothesis requires experimental validation. While centrocone-like structures have been identified in Plasmodium during mitosis and budding [9,33], it is currently not clear if these persist (developing reagents to the Plasmodium homolog of the MORN1 protein should quickly resolve this question).
(A–C) Schematic representation of the nucleus during interphase (A), mitosis (B), and mid-stage budding (C). Smaller type abbreviations refer to organelle-specific marker proteins in T. gondii (most are available as fluorescent protein in vivo tags, see text for further details and references). AP, apicoplast; AR, apical rings; CC, centrocone; CH, chromosome; CO, conoid; CT, centromere; EX, ER exit site; MT, subpellicular microtubule; NE, nuclear envelope; PR, posterior ring; SP, spindle.
(D–K) Time lapse series of nuclear division in T. gondii reproduced from [32]. The nucleus is labeled in red (Histone H2b-RFP) and MORN1 in green (MORN1-YFP).
Building the Zoite Scaffold
Apicomplexans preassemble zoites as buds either internally in the cytoplasm (Toxoplasma) or directly under the surface membrane (Plasmodium). The scaffold for bud assembly and the outline of the new daughter cells is provided by the pellicle, which consists of subpellicular microtubules and the inner membrane complex (IMC). The subpellicular microtubules emerge from an apical microtubule organizing center associated with the polar rings and run along the longitudinal axis of the cell [34,35]. The IMC is a system of flattened membrane cisternae stabilized by a membrane-associated protein meshwork facing the cytoplasma. Several of the protein components of this meshwork have been characterized and they share weak similarity with articulins, filament proteins found in ciliates [36–38]. Several IMC proteins show dynamic regulation, with their expression timed to coincide with budding [29]. Some IMC proteins also undergo proteolytic processing, and it has been suggested that this process confers increased rigidity to the IMC following its deposition [38,39]. Recently, proteins integral or tightly associated with the outer IMC membrane have been identified. GAP50, together with GAP45, serve as internal anchors of myosin A and the associated gliding motility machinery [40,41]. However, the function of PHIL1, which forms a ring structure at the apical tip of the bud, remains to be elucidated [42].
Following mitotic separation of the chromosomes, budding initiates in the direct vicinity of the centrosomes. The first identifiable sign of the bud is a flattened vesicle associated with a small number of evenly spaced microtubules [8,31,43,44]. This structure is further elaborated into a cup, with the conoid at its apex and microtubules extending from the conoid to posterior ring, delimiting the bud. Genetic and proteomic studies in Toxoplasma have identified a number of proteins associated with these early processes, and fluorescent protein tagging and live cell microscopy has painted a highly dynamic picture of their localization and function. The Toxoplasma gondii genome encodes several centrin genes, with centrin 1, 2, and 3 having been localized by GFP fusion [45–47]. While centrin 1 and 3 appear to be focused at the centrosome, centrin 2 additionally labels the conoid and a peculiar group of punctate structures in the apex of the cell [45,46]. Dynein light chain, a component of the minus end-directed microtubular motor dynein, has been detected near the centrosome and the conoid, and may be involved in conoid and centrosomal movements. MORN1 has been particularly informative as a marker for budding, as it labels both the centrocone and spindle and the apical and posterior ends of the bud (Figure 4; [32,46]). The precise chronology of assembly—especially in the very early phase of bud development—remains to be elucidated, and would benefit from the generation of mutants for the various steps involved. Early electron microscopic studies have implicated a striated fiber as an organizing element [44]; interestingly, proteins similar to algal-striated fiber assemblins have been identified recently in apicomplexans and have been shown to localize to the centrosomal region during budding [48]. Once the bud is assembled it grows rapidly, most likely driven by microtubule growth. This process runs opposite to spindle extension and effectively partitions the nucleus and much of the cytoplasm. Toward the end of bud development, the MORN1 ring at the posterior end of the bud shows pronounced contraction (see Figure 4J and 4K), which likely aids in organellar division (see below) and cytokinesis. Several observations are consistent with an association of this ring with myosin B/C [32,49]; however, the actin-destabilizing drug cytochalasin D does not interfere with parasite division [15].
Completing Parasite Assembly
A fully formed apicomplexan parasite requires a multitude of organelles and intracellular structures that will enable it to carry out the next task of its life cycle—to egress from the host cell and invade a new one. Rhoptries, micronemes, and dense granules form de novo during budding, anterior to the nucleus, endowing each daughter cell with the apical secretory organelles necessary for invasion. Expression of rhoptry and microneme proteins is regulated at the transcriptional level and timed to conincide with budding [50–53]. The apicomplexan secretory pathway is highly polarized, with an endoplasmic reticulum (ER) exit site localized on the apical face of the nucleus adjacent to the centrocone [43,54,55]. Here, proteins are loaded into coated vesicles that travel to the Golgi and on to several (still poorly characterized) trans-Golgi, pre-rhoptry, and pre-miconeme compartments [54,56,57]. The Golgi is associated with the centrosome(s), which play an important role in its duplication [58]. Golgi duplication is among the earliest events of budding [47,59]. In Plasmodium, the Golgi divides multiple times during intracellular development, and upon zoite formation, a single Golgi is associated with each bud [60]. The spatially fixed line-up of ER exit site and Golgi and their association with the nucleus and centrosome likely acts as a highly effective cellular “funnel”, directing the flow of proteins and membranes into the growing buds. IMC proteins, including the N-glycosylated GAP50 [40], probably derive from the Golgi, suggesting that membranes of the IMC form from Golgi-derived vesicles. This would explain the necessity for early division of the Golgi during budding, and suggests that Golgi positioning by the centrosome is critical in mediating deposition of the IMC.
Apicomplexan Parasites
Apicomplexans harbor two endosymbiont-derived organelles, the mitochondrion and the apicoplast, both of which perform a broad array of metabolic functions and are essential for intracellular parasite development [61–64]. These organelles carry their own genomes [65–69] and therefore can not be formed de novo, but must undergo division followed by segregation into buds. Genomic analyses in apicomplexans have identified proteins commonly involved in mitochondrial division, like dynamin-related proteins [62]. However, the FtsZ-based division machine found in a wide variety of chloroplasts has been lost in apicomplexans [70,71]. Instead of relying on their ancestral prokaryotic division ring, it would appear that apicoplasts have developed novel means of division. One model suggests that the force for apicoplast division is provided by association of the apicoplast with the mitotic spindle [72]. Dynamic association between the centrosome(s) and the apicoplast has been demonstrated in Toxoplasma and Sarcocystis and provides a likely means by which these organelles are properly segregated into forming buds [29,72]. In both organisms, fission of the organelle into daughter plastids is tightly associated with budding, and the constrictive MORN1 ring found at the posterior end of each bud provides an attractive candidiate for a fission mechanism (see Figure 4C; [32,71]). A second model suggests that apicoplast fission is independent of cytokinesis and relies on a medial division ring formed by yet-to-be identified components [73]. The development of the plastid in organisms dividing by schizogony, like Plasmodium and Eimeria, is not fully understood [74,75]. While centrosome association is likely to be involved in the segregation into daughters, it is unclear if such association occurs in earlier stages. In Plasmodium, mitochondria and apicoplasts form a physical association shortly before budding [74], suggesting that segregation of these organelles into daughter buds is tightly linked. Nevertheless, better in vivo markers (especially for the centrosome) are needed to identify mechanisms of organellar division and segregation in these organisms.
Outlook
The advent of reverse genetics for a variety of apicomplexans has led to a renaissance in the study of the cell biology of these parasites. A number of exciting new structures and mechanisms have been discovered in this process. Not unlike the study of host cell invasion, exploring the intracellular development of apicomplexans has brought out conserved themes at the mechanistic level, suggesting significant similarity between different species within the phylum. The “post-genomic” era of apicomplexan cell biology offers powerful experimental avenues that will undoubtedly drive our understanding of cell division and zoite formation. Gene expression profiling using microarrays, now available for several systems, has identified large groups of candidate genes that are expressed during budding. Comparative genomic analysis can be used to further narrow the list of candidates. The ever improving forward and reverse genetics tool box offers robust experimental avenues to test the function of essential genes, and genetic analysis will be critical for establishing the sequence of events during budding [76]. The coming years will likely reveal an increasingly detailed and mechanistic picture of these tiny diabolical, yet fascinating, invasion machines.
Acknowledgments
We thank Dirk Dobbelaere, Volker Heussler, Lawrence Bannister, and Marc-Jan Gubbles for images and discussion.
References
- 1. Sibley LD (2004) Intracellular parasite invasion strategies. Science 304: 248–253.
- 2. Dobrowolski JM, Sibley LD (1996) Toxoplasma invasion of mammalian cells is powered by the actin cytoskeleton of the parasite. Cell 84: 933–939.
- 3. Meissner M, Schluter D, Soldati D (2002) Role of Toxoplasma gondii myosin A in powering parasite gliding and host cell invasion. Science 298: 837–840.
- 4. Carruthers V, Boothroyd JC (2006) Pulling together: An integrated model of Toxoplasma cell invasion. Curr Opin Microbiol 10: 83–89.
- 5. Cowman AF, Crabb BS (2006) Invasion of red blood cells by malaria parasites. Cell 124: 755–766.
- 6. Morrissette NS, Sibley LD (2002) Cytoskeleton of apicomplexan parasites. Microbiol Mol Biol Rev 66: 21–38. table of contents.
- 7. Hu K, Roos DS, Murray JM (2002) A novel polymer of tubulin forms the conoid of Toxoplasma gondii. J Cell Biol 156: 1039–1050.
- 8. Sheffield HG, Melton ML (1968) The fine structure and reproduction of Toxoplasma gondii. J Parasitol 54: 209–226.
- 9. Bannister LH, Hopkins JM, Fowler RE, Krishna S, Mitchell GH (2000) A brief illustrated guide to the ultrastructure of Plasmodium falciparum asexual blood stages. Parasitol Today 16: 427–433.
- 10. Speer CA, Dubey JP (1999) Ultrastructure of shizonts and merozoites of Sarcocystis falcatula in the lungs of budgerigars (Melopsittacus undulatus). J Parasitol 85: 630–637.
- 11. Speer CA, Dubey JP (2005) Ultrastructural differentiation of Toxoplasma gondii schizonts (types B to E) and gamonts in the intestines of cats fed bradyzoites. Int J Parasitol 35: 193–206.
- 12. Dobbelaere DA, Kuenzi P (2004) The strategies of the Theileria parasite: A new twist in host-pathogen interactions. Curr Opin Immunol 16: 524–530.
- 13. Heussler VT, Rottenberg S, Schwab R, Kuenzi P, Fernandez PC, et al. (2002) Hijacking of host cell IKK signalosomes by the transforming parasite Theileria. Science 298: 1033–1036.
- 14. Shaw MK, Tilney LG, Musoke AJ (1991) The entry of Theileria parva sporozoites into bovine lymphocytes: Evidence for MHC class I involvement. J Cell Biol 113: 87–101.
- 15. Shaw MK, Compton HL, Roos DS, Tilney LG (2000) Microtubules, but not actin filaments, drive daughter cell budding and cell division in Toxoplasma gondii. J Cell Sci 113: 1241–1254.
- 16. Shaw MK, Roos DS, Tilney LG (2001) DNA replication and daughter cell budding are not tightly linked in the protozoan parasite Toxoplasma gondii. Microbes Infect 3: 351–362.
- 17. Radke JR, Striepen B, Guerini MN, Jerome ME, Roos DS, et al. (2001) Defining the cell cycle for the tachyzoite stage of Toxoplasma gondii. Mol Biochem Parasitol 115: 165–175.
- 18. White MW, Jerome ME, Vaishnava S, Guernin MN, Behnke M, et al. (2005) Genetic rescue of a Toxoplasma gondii conditional cell cycle mutant. Mol Microbiol 55: 1060–1067.
- 19.
White MW, Radke JA, Conde de Felipe M, Lehmann M (2007) Cell cycle control/parasite division. In: Aijoka JW, Soldati D, editors. Toxoplasma: Molecular and cellular biology. Norwich (United Kingdom): Horizon Scientific Press. pp. 263–282. pp.
- 20. Chen Y, Jirage D, Caridha D, Kathcart AK, Cortes EA, et al. (2006) Identification of an effector protein and gain-of-function mutants that activate Pfmrk, a malarial cyclin-dependent protein kinase. Mol Biochem Parasitol 149: 48–57.
- 21. Khan F, Tang J, Qin CL, Kim K (2002) Cyclin-dependent kinase TPK2 is a critical cell cycle regulator in Toxoplasma gondii. Mol Microbiol 45: 321–332.
- 22. Kvaal CA, Radke JR, Guerini MN, White MW (2002) Isolation of a Toxoplasma gondii cyclin by yeast two-hybrid interactive screen. Mol Biochem Parasitol 120: 187–194.
- 23. Ward P, Equinet L, Packer J, Doerig C (2004) Protein kinases of the human malaria parasite Plasmodium falciparum: The kinome of a divergent eukaryote. BMC Genomics 5: 79.
- 24. Lye YM, Chan M, Sim TS (2006) Pfnek3: An atypical activator of a MAP kinase in Plasmodium falciparum. FEBS Lett 580: 6083–6092.
- 25. Reininger L, Billker O, Tewari R, Mukhopadhyay A, Fennell C, et al. (2005) A NIMA-related protein kinase is essential for completion of the sexual cycle of malaria parasites. J Biol Chem 280: 31957–31964.
- 26. Khan SM, Franke-Fayard B, Mair GR, Lasonder E, Janse CJ, et al. (2005) Proteome analysis of separated male and female gametocytes reveals novel sex-specific Plasmodium biology. Cell 121: 675–687.
- 27. Dorin D, Le Roch K, Sallicandro P, Alano P, Parzy D, et al. (2001) Pfnek-1, a NIMA-related kinase from the human malaria parasite Plasmodium falciparum Biochemical properties and possible involvement in MAPK regulation. Eur J Biochem 268: 2600–2608.
- 28. Morrissette NS, Sibley LD (2002) Disruption of microtubules uncouples budding and nuclear division in Toxoplasma gondii. J Cell Sci 115: 1017–1025.
- 29. Vaishnava S, Morrison DP, Gaji RY, Murray JM, Entzeroth R, et al. (2005) Plastid segregation and cell division in the apicomplexan parasite Sarcocystis neurona. J Cell Sci 118: 3397–3407.
- 30. Fennell BJ, Naughton JA, Dempsey E, Bell A (2006) Cellular and molecular actions of dinitroaniline and phosphorothioamidate herbicides on Plasmodium falciparum: Tubulin as a specific antimalarial target. Mol Biochem Parasitol 145: 226–238.
- 31. Dubremetz JF (1973) Ultrastructural study of schizogonic mitosis in the coccidian, Eimeria necatrix (Johnson 1930). J Ultrastruct Res 42: 354–376.
- 32. Gubbels MJ, Vaishnava S, Boot N, Dubremetz JF, Striepen B (2006) A MORN-repeat protein is a dynamic component of the Toxoplasma gondii cell division apparatus. J Cell Sci 119: 2236–2245.
- 33. Bannister LH, Hopkins JM, Margos G, Dluzewski AR, Mitchell GH (2004) Three-dimensional ultrastructure of the ring stage of Plasmodium falciparum: Evidence for export pathways. Microsc Microanal 10: 551–562.
- 34. Russell DG, Burns RG (1984) The polar ring of coccidian sporozoites: A unique microtubule-organizing centre. J Cell Sci 65: 193–207.
- 35. Morrissette NS, Murray JM, Roos DS (1997) Subpellicular microtubules associate with an intramembranous particle lattice in the protozoan parasite Toxoplasma gondii. J Cell Sci 110: 35–42.
- 36. Gubbels MJ, Wieffer M, Striepen B (2004) Fluorescent protein tagging in Toxoplasma gondii: Identification of a novel inner membrane complex component conserved among Apicomplexa. Mol Biochem Parasitol 137: 99–110.
- 37. Mann T, Beckers C (2001) Characterization of the subpellicular network, a filamentous membrane skeletal component in the parasite Toxoplasma gondii. Mol Biochem Parasitol 115: 257–268.
- 38. Mann T, Gaskins E, Beckers C (2002) Proteolytic processing of TgIMC1 during maturation of the membrane skeleton of Toxoplasma gondii. J Biol Chem 277: 41240–41246.
- 39. Hu K, Mann T, Striepen B, Beckers CJ, Roos DS, et al. (2002) Daughter cell assembly in the protozoan parasite Toxoplasma gondii. Mol Biol Cell 13: 593–606.
- 40. Gaskins E, Gilk S, DeVore N, Mann T, Ward G, et al. (2004) Identification of the membrane receptor of a class XIV myosin in Toxoplasma gondii. J Cell Biol 165: 383–393.
- 41. Baum J, Richard D, Healer J, Rug M, Krnajski Z, et al. (2006) A conserved molecular motor drives cell invasion and gliding motility across malaria life cycle stages and other apicomplexan parasites. J Biol Chem 281: 5197–5208.
- 42. Gilk SD, Raviv Y, Hu K, Murray JM, Beckers CJ, et al. (2006) Identification of PhIL1, a novel cytoskeletal protein of the Toxoplasma gondii pellicle, through photosensitized labeling with 5-[125I]iodonaphthalene-1-azide. Eukaryot Cell 5: 1622–1634.
- 43. Bannister LH, Hopkins JM, Fowler RE, Krishna S, Mitchell GH (2000) Ultrastructure of rhoptry development in Plasmodium falciparum erythrocytic schizonts. Parasitology 121(Part 3): 273–287.
- 44. Dubremetz JF (1975) Genesis of merozoites in the coccidia, Eimeria necatrix. Ultrastructural study. J Protozool 22: 71–84.
- 45. Nagamune K, Sibley LD (2006) Comparative genomic and phylogenetic analyses of calcium ATPases and calcium-regulated proteins in the apicomplexa. Mol Biol Evol 23: 1613–1627.
- 46. Hu K, Johnson J, Florens L, Fraunholz M, Suravajjala S, et al. (2006) Cytoskeletal components of an invasion machine—The apical complex of Toxoplasma gondii. PLoS Pathog 2: e13..
- 47. Hartmann J, Hu K, He CY, Pelletier L, Roos DS, et al. (2006) Golgi and centrosome cycles in Toxoplasma gondii. Mol Biochem Parasitol 145: 125–127.
- 48. Lechtreck KF (2003) Striated fiber assemblin in apicomplexan parasites. Mol Biochem Parasitol 128: 95–99.
- 49. Delbac F, Sanger A, Neuhaus EM, Stratmann R, Ajioka JW, et al. (2001) Toxoplasma gondii myosins B/C: One gene, two tails, two localizations, and a role in parasite division. J Cell Biol 155: 613–623.
- 50. Baldi DL, Andrews KT, Waller RF, Roos DS, Howard RF, et al. (2000) RAP1 controls rhoptry targeting of RAP2 in the malaria parasite Plasmodium falciparum. EMBO J 19: 2435–2443.
- 51. Brown PJ, Billington KJ, Bumstead JM, Clark JD, Tomley FM (2000) A microneme protein from Eimeria tenella with homology to the Apple domains of coagulation factor XI and plasma pre-kallikrein. Mol Biochem Parasitol 107: 91–102.
- 52. Hoane JS, Carruthers VB, Striepen B, Morrison DP, Entzeroth R, et al. (2003) Analysis of the Sarcocystis neurona microneme protein SnMIC10: protein characteristics and expression during intracellular development. Int J Parasitol 33: 671–679.
- 53. Bozdech Z, Llinas M, Pulliam BL, Wong ED, Zhu J, et al. (2003) The transcriptome of the intraerythrocytic developmental cycle of Plasmodium falciparum. PLoS Biol 1: e5..
- 54. Hager KM, Striepen B, Tilney LG, Roos DS (1999) The nuclear envelope serves as an intermediary between the ER and golgi complex in the intracellular parasite Toxoplasma gondii. J Cell Sci 112: 2631–2638.
- 55. Pfluger SL, Goodson HV, Moran JM, Ruggiero CJ, Ye X, et al. (2005) Receptor for retrograde transport in the apicomplexan parasite Toxoplasma gondii. Eukaryot Cell 4: 432–442.
- 56. Ngo HM, Hoppe HC, Joiner KA (2000) Differential sorting and post-secretory targeting of proteins in parasitic invasion. Trends Cell Biol 10: 67–72.
- 57. Harper JM, Huynh MH, Coppens I, Parussini F, Moreno S, et al. (2006) A cleavable propeptide influences Toxoplasma infection by facilitating the trafficking and secretion of the TgMIC2-M2AP invasion complex. Mol Biol Cell 17: 4551–4563.
- 58. He CY (2007) Golgi biogenesis in simple eukaryotes. Cell Microbiol 9: 566–572.
- 59. Pelletier L, Stern CA, Pypaert M, Sheff D, Ngo HM, et al. (2002) Golgi biogenesis in Toxoplasma gondii. Nature 418: 548–552.
- 60. Struck NS, de Souza Dias S, Langer C, Marti M, Pearce JA, et al. (2005) Re-defining the Golgi complex in Plasmodium falciparum using the novel Golgi marker PfGRASP. J Cell Sci 118: 5603–5613.
- 61. Ralph SA, Van Dooren GG, Waller RF, Crawford MJ, Fraunholz MJ, et al. (2004) Tropical infectious diseases: Metabolic maps and functions of the Plasmodium falciparum apicoplast. Nat Rev Microbiol 2: 203–216.
- 62. van Dooren GG, Stimmler LM, McFadden GI (2006) Metabolic maps and functions of the Plasmodium mitochondrion. FEMS Microbiol Rev 30: 596–630.
- 63. Mazumdar J, Wilson E, Masarek K, Hunter C, Striepen B (2006) Apicoplast fatty acid synthesis is essential for organelle biogenesis and parasite survival in Toxoplasma gondii. Proc Natl Acad Sci U S A 103: 13192–13197.
- 64. Jomaa H, Wiesner J, Sanderbrand S, Altincicek B, Weidemeyer C, et al. (1999) Inhibitors of the nonmevalonate pathway of isoprenoid biosynthesis as antimalarial drugs. Science 285: 1573–1576.
- 65. Kohler S, Delwiche CF, Denny PW, Tilney LG, Webster P, et al. (1997) A plastid of probable green algal origin in Apicomplexan parasites. Science 275: 1485–1489.
- 66. McFadden GI, Reith ME, Munholland J, Lang-Unnasch N (1996) Plastid in human parasites. Nature 381: 482.
- 67. Feagin JE (1992) The 6-kb element of Plasmodium falciparum encodes mitochondrial cytochrome genes. Mol Biochem Parasitol 52: 145–148.
- 68. Wilson RJ, Denny PW, Preiser PR, Rangachari K, Roberts K, et al. (1996) Complete gene map of the plastid-like DNA of the malaria parasite Plasmodium falciparum. J Mol Biol 261: 155–172.
- 69. Vaidya AB, Akella R, Suplick K (1989) Sequences similar to genes for two mitochondrial proteins and portions of ribosomal RNA in tandemly arrayed 6-kilobase-pair DNA of a malarial parasite. Mol Biochem Parasitol 35: 97–107.
- 70. Osteryoung KW, Nunnari J (2003) The division of endosymbiotic organelles. Science 302: 1698–1704.
- 71. Vaishnava S, Striepen B (2006) The cell biology of secondary endosymbiosis—How parasites build, divide and segregate the apicoplast. Mol Microbiol 61: 1380–1387.
- 72. Striepen B, Crawford MJ, Shaw MK, Tilney LG, Seeber F, et al. (2000) The plastid of Toxoplasma gondii is divided by association with the centrosomes. J Cell Biol 151: 1423–1434.
- 73. Ferguson DJ, Henriquez FL, Kirisits MJ, Muench SP, Prigge ST, et al. (2005) Maternal inheritance and stage-specific variation of the apicoplast in Toxoplasma gondii during development in the intermediate and definitive host. Eukaryot Cell 4: 814–826.
- 74. van Dooren GG, Marti M, Tonkin CJ, Stimmler LM, Cowman AF, et al. (2005) Development of the endoplasmic reticulum, mitochondrion and apicoplast during the asexual life cycle of Plasmodium falciparum. Mol Microbiol 57: 405–419.
- 75. Ferguson DJ, Campbell SA, Henriquez FL, Phan L, Mui E, et al. (2007) Enzymes of type II fatty acid synthesis and apicoplast differentiation and division in Eimeria tenella. Int J Parasitol 37: 33–51.
- 76.
Striepen B, Soldati D (2007) Genetic manipulation of Toxoplasma gondii. In: Weiss LM, Kim K, editors. Toxoplasma gondii: The model Apicomplexan. Perspective and methods. Oxford: Elsevier. pp. 391–418. pp.