Search tips
Search criteria 


Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
FEMS Microbiol Rev. Author manuscript; available in PMC 2013 May 1.
Published in final edited form as:
PMCID: PMC3319474

Mechanisms of Toxoplasma gondii persistence and latency


Toxoplasma gondii is an obligate intracellular protozoan parasite that causes opportunistic disease, particularly in immunocompromised individuals. Central to its transmission and pathogenesis is the ability of the proliferative stage (tachyzoite) to convert into latent tissue cysts (bradyzoites). Encystment allows Toxoplasma to persist in the host, and affords the parasite a unique opportunity to spread to new hosts without proceeding through its sexual stage, which is restricted to felids. Bradyzoite tissue cysts can cause reactivated toxoplasmosis if host immunity becomes impaired. A greater understanding of the molecular mechanisms orchestrating bradyzoite development is needed to better manage the disease. Here we will review key studies that have contributed to our knowledge about this persistent form of the parasite and how to study it, with a focus on how cellular stress can signal for the reprogramming of gene expression needed during bradyzoite development.

Keywords: parasite, Apicomplexa, differentiation, eukaryotic pathogen, microbial persistence, stress response


Toxoplasma gondii is one of the most successful parasites on Earth. Despite its obligate intracellular lifestyle, this protozoan parasite has remarkable transmissibility, and has permanently infected most species of mammals and birds around the world. As we will discuss, the ability of Toxoplasma to encyst inside the cells of host tissues and reconvert back into its proliferative stage was an evolutionary paradigm shift, circumventing the need for the parasite to undergo its sexual stage in order to be transmitted to a new host. This unusual property affords Toxoplasma a second major route of transmission: the capacity to disseminate clonally through intermediate hosts.

Toxoplasma belongs to phylum Apicomplexa, which contains many other protozoan pathogens of human and veterinary importance, such as Plasmodium spp. (malaria), Cryptosporidium spp. (cryptosporidiosis), and Eimeria spp. (poultry coccidiosis). Toxoplasmosis is a notorious opportunistic disease in AIDS patients and other immunocompromised individuals that most commonly results from reactivation of a previous infection (Luft, et al., 1984). Ocular infection by Toxoplasma is a major cause of retinochoroiditis in both immunocompromised and immunocompetent individuals (Wallace & Stanford, 2008). Additionally, Toxoplasma can cross the placental barrier and cause abortion or congenital birth defects if the mother becomes infected for the first time shortly before or during pregnancy (Jones, et al., 2003). In utero infection is also associated with an elevated risk of ocular toxoplasmosis due to spontaneous reactivation of disease (Wallace & Stanford, 2008).

Toxoplasma has a complex life cycle consisting of multiple stages that fluctuate between proliferative and latent stages (Fig. 1). The proliferative stage, known as the tachyzoite, replicates exponentially by endodyogeny (asexual division whereby two daughter cells form within a single mother cell) inside of host cells with a doubling time of ~7 hours (Radke & White, 1998). The intracellular tachyzoites, housed in a membrane-bound compartment called the parasitophorous vacuole (PV), can convert into virtually quiescent forms known as bradyzoites, which transform the parasitophorous vacuole membrane (PVM) into a cyst wall in the process. Bradyzoite cysts remain infectious and can form in skeletal, heart and CNS tissues, granting Toxoplasma the ability to spread to a new host following predation of its former host. Felines serve as the definitive host for Toxoplasma, whose intestinal milieu is the only known environment suitable for the sexual stage of the parasite life cycle. Due to factors that have yet to be identified, the cat gut induces bradyzoites to differentiate into macrogametes and flagellated microgametes that fuse and lead to the formation of oocysts. Dissemination of the stable, highly infectious oocysts into the environment provides another major route of transmission through the food and water chain (Fig. 1). While it is not possible to distinguish infection from meat versus oocysts, it has been proposed that the surge of infection in teenagers and low prevalence in younger children indicates that transmission through cysts in undercooked meat is important (Dubey & Jones, 2008). As a consequence of these multiple routes of transmission, Toxoplasma has made itself at home in at least one in three people. Seroprevalence varies widely between geographical regions, but estimates suggest that between 30%–65% of humans worldwide are infected (Tenter, et al., 2000).

Figure 1
Life cycle of Toxoplasma gondii

The ability to differentiate into bradyzoites and form these impenetrable cysts makes it currently impossible to eradicate Toxoplasma from the host. While several drugs are available that control acute toxoplasmosis, such as pyrimethamine plus sulfadiazine, no short-term treatment exists that can eliminate the cysts, which also appear impervious to the immune response. The presence of latent bradyzoite cysts makes Toxoplasma a chronic infection, and the ability of bradyzoites to reconvert into rapidly growing tachyzoites explains the high frequencies of acute toxoplasmosis often observed in immunocompromised individuals (Wong & Remington, 1993, Montoya & Liesenfeld, 2004). Toxoplasmic encephalitis (TE), characterized by intracerebral lesions, is a significant CNS complication in AIDS patients (Fig. 2).

Figure 2
Reactivated Toxoplasma infection

Microbial persistence and latency are conserved strategies that numerous pathogens use to their advantage. The ability of Toxoplasma to form latent cysts is an evolutionary trade off that reduces virulence while increasing transmission – a win-win situation for the parasite. In terms of human and veterinary health, understanding bradyzoite conversion is the key to minimizing transmission and managing chronic infection. In this review, we will discuss bradyzoite development and physiology, with a focus on the molecular mechanisms orchestrating differentiation. We will also highlight methods used to study bradyzoite cysts in vitro and in vivo and present important questions that require further investigation.

Biology of the bradyzoite cyst

Morphological features of bradyzoites and cysts

Bradyzoite tissue cysts develop and remain inside of nucleated cells, exhibiting tropism similar to tachyzoites, but are more frequently found in brain, eye, skeletal muscle, and cardiac tissue. Within the brain, bradyzoite cysts appear to form in a wide variety of regions, although a subtle tropism for the medial and basolateral amygdala has been reported (Vyas, et al., 2007). Mature tissue cysts can be detected within 7–10 days post-infection, but hallmarks of bradyzoite differentiation can be detected within 3 days post-induction in vitro. Recent studies using bioluminescence imaging technology also show that conversion to bradyzoites begins rapidly in vivo, as early as one day post-infection (Di Cristina, et al., 2008). Tissue cysts vary in size and shape. Young cysts can be as small as 5 μm in diameter with as few as two bradyzoites while mature cysts can be up to 70 μm with ~1000 bradyzoites (Dubey, et al., 1998) (Fig. 3A). Individual bradyzoites (7 × 1.5 μm) are slender relative to tachyzoites (6 × 2 μm), with a more pronounced bow or crescent shape (Mehlhorn & Frenkel, 1980). Also different from tachyzoites, the nucleus in mature bradyzoites resides at the parasite’s posterior, rhoptry organelles are electron dense, and there is a marked increase in micronemes and amylopectin (glucose storage) granules (Dubey, et al., 1998) (Fig. 3B).

Figure 3
Bradyzoite cyst and morphology

The bradyzoite PVM/cyst wall is <0.5 μm thick and elastic to accommodate expansion. The cyst wall is built from chitin and glycoproteins secreted by the parasites (Boothroyd, et al., 1997, Zhang, et al., 2001). The sugars in the cyst wall are capable of binding lectins Concanavalin A, wheat germ agglutinin, soy bean agglutinin, and Dolichos biflorus agglutinin (Sethi, et al., 1977), the latter of which is commonly used as a diagnostic tool for cyst wall staining (Fig. 4). In mature cysts, a matrix material fills the PV space between bradyzoites (Ferguson & Hutchison, 1987). The number of cyst wall/matrix proteins that have been characterized to date is limited. CST1 is a 116 kDa cyst wall glycoprotein that is the major binding constituent of Dolichos lectin (Zhang, et al., 2001). MAG1 is a 65 kDa protein that localizes to the cyst wall and matrix (Parmley, et al., 1994). At least one dense granule protein, GRA5, has been reported to be highly concentrated in the cyst wall; less intense staining of GRA1, GRA3, and GRA6 can also be observed (Lane, et al., 1996, Ferguson, 2004). Upon ingestion by a host, the cyst wall is degraded by acid-pepsin digestion, releasing the bradyzoites. Unlike tachyzoites, bradyzoites are resistant to proteolytic enzymes (Jacobs, et al., 1960), a key feature that assures survival in the host stomach. The bradyzoites proceed to invade intestinal epithelial cells to begin infection and dissemination throughout the host.

Figure 4
Dolichos lectin stains bradyzoite cyst wall

Studies have demonstrated that mature bradyzoites isolated from the brains of mice infected with Toxoplasma oocysts represent a growth-arrested stage with parasites suspended in G0 with uniform 1N DNA content (Radke, et al., 2003). To provide insights into the dynamics of early stage conversion, an in vitro system using type II (Prugniaud) parasites subjected to CO2 deprivation was developed to maintain bradyzoites for up to 2 weeks. Using this system, it was observed that immature bradyzoites replicate slowly (requiring >12 hours compared to the ~7 hours for tachyzoites) and asynchronously, using a combination of endodyogeny and endopolygeny (asexual replication whereby more than two daughter cells form within the mother cell) (Dzierszinski, et al., 2004). Interestingly, within the context of this system, 10–20% of developing bradyzoite vacuoles contained individual parasites lacking an apicoplast, a non-photosynthetic plastid-like organelle acquired from an algal endosymbiont (Dzierszinski, et al., 2004, McFadden, 2010). Despite their reputation for quiescence, the developing bradyzoites generated under these conditions were highly motile and capable of exiting a host cell without destroying it to invade a neighboring cell; bradyzoites within mature cysts purified from mouse brains also exhibit intravacuolar motility (Dzierszinski, et al., 2004). Bradyzoite motility within and between host cells may explain why increases in cyst burden are observed in chronically infected mice despite the low frequency of cyst rupture.

Metabolism in the latent stage

Changes in parasite metabolism accompany the switch to a latent lifestyle. A number of metabolic enzymes have tachyzoite and bradyzoite-specific isoforms (e.g. ENO2/ENO1 and LDH1/LDH2), suggesting a fine tuning of metabolism between these two life cycle stages. The abundance of polysaccharide in the form of amylopectin granules in the bradyzoites reflects a major shift in carbohydrate metabolism. This idea is supported by biochemical analyses that concluded bradyzoites lack a functional TCA cycle and respiratory chain, suggesting a predominant role for anaerobic glycolysis during this stage (Denton, et al., 1996). In contrast, tachyzoites likely use both mitochondrial oxidative phosphorylation and glycolysis to generate ATP. Pyruvate kinase and lactate dehydrogenase activities are markedly higher in bradyzoites, suggesting that lactate production is particularly important during latency (Denton, et al., 1996). A bradyzoite-specific isoform of lactate dehydrogenase (LDH2) has been identified that is likely to be specialized for the latent stage (Yang & Parmley, 1997). LDH2 is resistant to acidic pH, and would continue to function during the acidification that would occur during the catabolism of amylopectin to lactate in bradyzoites. Knockdown of LDH2 resulted in parasites that were unable to establish significant cyst burdens in the brains of infected mice (Al-Anouti, et al., 2004).

Two other bradyzoite-specific isoforms of key glycolytic enzymes have been identified: glucose-6-phosphate isomerase (G6-PI) and enolase-1 (ENO1) (Dzierszinski, et al., 1999). The enolase isoforms display distinct enzymatic properties and differ in their stability. The tachyzoite-specific enolase (ENO2) exhibits a similar Km value for the 2-phosphoglycerate substrate, but has a threefold higher specific activity at Vmax compared to ENO1 (Dzierszinski, et al., 2001). Considered together, these studies are consistent with the idea that stage-specific enzymes exist that are designed to tune glycolysis to accommodate proliferation or dormancy. Alternatively, or in addition to tailoring glycolysis, the enolases may play a role in transcriptional regulation. ENO1 and ENO2 have been found not only in the cytoplasm but also in the parasite nucleus of bradyzoites and tachyzoites (Ferguson, et al., 2002). An alternatively translated form of human ENO1 also localizes to the nucleus and can repress transcription by binding to the c-myc promoter (Feo, et al., 2000).

A number of enzymes with roles in the metabolism of oxygen radicals appear to be up-regulated in bradyzoites, suggesting that the encysted forms are equipped to deal with long-term exposure to reactive metabolites (Manger, et al., 1998). Consistent with this idea, it has been reported that mRNAs encoding various DNA repair enzymes increase in bradyzoites (Manger, et al., 1998, Yahiaoui, et al., 1999).

Triggers of bradyzoite formation

Strain differences

Three predominating strains of Toxoplasma (designated type I, II, and III) have been documented in North America and Europe that differ in growth rate, virulence in mice, and ability to form cysts (Howe & Sibley, 1995). Genetic polymorphism analyses indicate that these clonal lineages emerged after a single genetic cross and their swift expansion resulted from the acquisition of direct oral infectivity (Su, et al., 2003). It is estimated this genetic cross occurred ~10,000 years ago, coincident with the establishment of agriculture and domestication of animals such as the cat in the Fertile Crescent. Oral infectivity is a critical factor linked with the ability of some strains to efficiently differentiate into bradyzoites (Fux, et al., 2007).

Type I strains grow rapidly and are hypervirulent in mice (LD50 ~ 1 parasite). The commonly used type I laboratory strain is designated RH, after the initials of the boy from which it was isolated in 1939 (Sabin, 1941). RH strain is generally considered to have lost the ability to develop into mature cysts at high frequency, perhaps as a consequence of prolonged propagation in vitro. To what degree RH strain has lost this developmental competency is a matter of debate and seems to vary between individual laboratory strains. During stress, type I strains including RH will activate gene expression of some bradyzoite marker genes, and in some cases bradyzoite surface antigens and cyst wall proteins have been detected (Soete, et al., 1994, Bohne & Roos, 1997, Lescault, et al., 2010). Isolated reports claim RH parasites can form cysts in mice, via treatment with atovaquone and pyrrolidine dithiocarbamate (Djurkovic-Djakovic, et al., 2005) or prior vaccination with soluble tachyzoite antigen and IL-12, although the bradyzoites formed by RH differ ultrastructurally and in their sensitivity to pepsin-HCl (Yap, et al., 1998). In addition, RH strain parasites attenuated in virulence due to disruption of dense granule gene GRA2 are capable of establishing chronic infection in mice (Mercier, et al., 1998). Other type I strains that have not been subject to extensive in vitro culture, such as GT1, are able to form normal cyst walls in vitro during stress (Khan, et al., 2009). Interestingly, it has been suggested that the phenotypic differences between type I strains are unlikely to be due to sequence variation (Khan, et al., 2009). Indeed, both epigenetic and translational control mechanisms have been linked with stage conversion (discussed below).

Type II (e.g. Prugniuad or Pru, ME49) and type III strains (e.g. VEG) have lower replication rates and readily form cysts in vitro and in vivo; consequently, they are hypovirulent in mice (LD50 ~104). The most commonly isolated strain from clinical toxoplasmosis is type II (Howe, et al., 1997), although type I strains are typically responsible for acute outbreaks. The genomes of representatives from all three lineages have been sequenced (data is available at, but which genes account for the differences reported in these strains have only begun to be resolved.

In terms of studying bradyzoite differentiation, strain choice is a crucial factor. While type I strains, particularly RH, grow faster and are easier to genetically manipulate, they are limiting in the study of cyst formation. However, several groups have made important discoveries regarding bradyzoite development using type I RH strain (discussed below). There are also more transgenic parasites made in the RH background that facilitate gene tagging/disruption, or conditional gene expression (Meissner, et al., 2007, Fox, et al., 2009, Huynh & Carruthers, 2009). Type II and III strains form mature cysts readily, but grow slowly and are more difficult to manipulate. Consequently, there are presently fewer tools available to study parasite physiology in the hypovirulent strains.

In vitro stresses induce stage conversion

It is well documented that conversion to the latent stage is a stress-mediated response, coupled with a slowing of the parasite cell cycle. One of the most commonly used in vitro methods to prompt bradyzoite differentiation is alkaline pH 8.0–8.2 (Soete, et al., 1994). A wide variety of other stress agents have since been reported, including sodium nitroprusside, which acts as a source of exogenous nitric oxide (NO) and also inhibits proteins involved in the parasite mitochondrial respiratory chain (Bohne, et al., 1994). Similarly, drugs that interfere with the parasite mitochondria also induce differentiation to bradyzoites (Bohne, et al., 1994, Tomavo & Boothroyd, 1995). Heat shock and treatment with sodium arsenite also trigger expression of bradyzoite antigens (Soete, et al., 1994). Nutrient deprivation is a potent inducer of bradyzoite formation and can be achieved through arginine starvation (Fox, et al., 2004), axenic incubation (Yahiaoui, et al., 1999), or pyrimidine depletion in uracil phosphoribosyltransferase (UPRT)-deficient parasites subjected to ambient (0.03%) CO2 (Bohne & Roos, 1997, Dzierszinski, et al., 2004). More recently, insults that cause endoplasmic reticulum (ER) stress or that interfere with calcium-induced egress have also been found to induce bradyzoite cyst formation (Nagamune, et al., 2008, Narasimhan, et al., 2008). Treatment with IFN-γ does not induce conversion to bradyzoites cultured in human fibroblasts (Soete, et al., 1994), but will do so in murine macrophages, presumably due to the stimulated releases of NO (Bohne, et al., 1993). Long-term culturing of cysts was accomplished in vitro using murine astrocytes and intermittent inclusion of IFN-γ in the culture media (Jones, et al., 1986). Methods used to convert tachyzoites to bradyzoites in vitro are summarized in Box 1.

Box 1

Induction of bradyzoites in vitro

  • Acidic or alkaline pH
  • Sodium nitroprusside
  • Atovaquone
  • Heat shock (43°C)
  • Sodium arsenite
  • Arginine starvation
  • Axenic incubation
  • Tunicamycin
  • Pyrimidine deprivation (ΔUPRT parasites, 0.03% CO2)
  • Fluridone (disruption of abscisic acid-mediated calcium signaling)

Whether these stress treatments act on the parasite directly (while they are extracellular), and/or if they act indirectly on intracellular parasites by stressing the host cell, is unclear. Weiss et al. has shown that when extracellular parasites are exposed to alkaline stress for 1 hr and allowed to re-infect host cells, bradyzoite differentiation is observed, albeit at a lower frequency than stressed intracellular parasites (Weiss, et al., 1998). Additionally, extracellular tachyzoites deprived of host cells for 12 hrs converted to bradyzoites upon reinfection of host cells (Yahiaoui, et al., 1999). These studies suggest that extracellular parasites not only sense their environment, but are marked in some way to initiate the bradyzoite development program upon re-entry into a host cell. Epigenetic-mediated gene regulation offers a mechanism by which this form of “short term memory” of the parasite’s environment could occur.

Conversion to bradyzoite cysts may be driven by physiological factors other than exogenous stresses. Tachyzoites can spontaneously convert into bradyzoites, and low MOIs and/or frequent removal of egressed tachyzoites from cultures of infected cells enriches for cysts (McHugh, et al., 1993). The proclivity towards spontaneous differentiation is influenced by the type of parasite strain and host cell background (Ferreira da Silva, et al., 2008), and cysts are more frequently detected in differentiated host cells that are long-lived (Dubey, et al., 1998). Further evidence that the host cell environment is a determinant in parasite differentiation comes from studies of a trisubstituted pyrrole known as Compound 1 (Donald, et al., 2002). Compound 1 was shown to act directly on human host cells to slow tachyzoite replication and induce bradyzoite-specific gene expression in type II and III strain parasites (Radke, et al., 2006). In these studies, human cell division autoantigen-1 (CDA1) was associated with promoting parasite differentiation, as tachyzoites infecting host cells overexpressing CDA1 underwent bradyzoite conversion (Radke, et al., 2006). Pre-stressing human foreskin fibroblasts (HFF) host cells prior to infection can also stimulate bradyzoite formation (Radke, et al., 2006). Collectively, these studies argue that tachyzoites are capable of assessing not only the type of host cell they invade, but also whether those host cells are under strain. These studies also suggest that the trigger(s) to differentiate are complex and multifactorial, consisting of both endogenous and exogenous factors.

In vivo factors relevant to the maintenance of bradyzoite cysts

The primary host response controlling Toxoplasma infection is mediated by CD8+ T-cells with synergistic action from CD4+ T-cells (Gazzinelli, et al., 1991, Gazzinelli, et al., 1992). While CD8+ T-cells have been shown to be cytotoxic to Toxoplasma-infected peritoneal macrophages (Kasper, et al., 1992), it is their production of interferon gamma (IFN-γ) that likely has a greater impact on controlling infection. IFN-γ is well documented as a key cytokine in the host immune response to Toxoplasma infection that limits intracellular replication of the parasite. Infected mice that are immunodepleted of IFN-γ succumb to toxoplasmosis instead of developing chronic infection (Suzuki, et al., 1988). Similarly, administration of IFN-γ protects against lethal infection (McCabe, et al., 1984). With regards to the mechanism of IFN-γ, it has been reported that in human brain microvascular endothelial cells (HBMEC), IFN-γ induces the tryptophan-degrading enzyme indoleamine 2,3-dioxygenase (Daubener, et al., 2001), which in turn would starve parasites of tryptophan (Pfefferkorn, et al., 1986). Starvation for key amino acids like tryptophan is likely to slow parasite growth and trigger cyst formation. Further support for this idea comes from studies demonstrating that arginine starvation induces bradyzoite differentiation (Fox, et al., 2004). IFN-γ also induces an oxidative burst and the production of NO, which may act as a direct trigger for bradyzoite differentiation (Bohne, et al., 1994). IFN-γ-inducible immunity-related GTPases (IRG proteins/p47 GTPases), such as IGTP and LRG-47, have also been linked to reduced parasite viability in activated macrophages (Butcher, et al., 2005). Most recently, the IRG protein Irga6 (IIGP1), which participates in the disruption of the PVM, is phosphorylated and inactivated by the type I ROP18 kinase as an immune evasion strategy that promotes virulence (Fentress, et al., 2010, Steinfeldt, et al., 2010). In short, numerous mechanisms exist that account for how IFN-γ controls tachyzoite proliferation in vivo, but whether IFN-γ acts directly on parasites to induce or maintain bradyzoites in cyst forms is less clear.

Other cytokines have been linked to promoting bradyzoite differentiation. In murine macrophages, tumor necrosis factor (TNF)α synergizes with IFN-γ to facilitate conversion to bradyzoites (Bohne, et al., 1994). Another study reported that the proinflammatory interleukin IL-6 favors the formation of bradyzoites in vitro in human fibroblasts (Weiss, et al., 1995). Given that heat shock is a potent in vitro trigger for bradyzoite differentiation, it is conceivable that fever could contribute to stage conversion during acute toxoplasmosis in vivo.

As discussed in the preceding section, host cell background appears to be a key factor influencing bradyzoite formation. Bradyzoite cysts have a clear predilection to form in brain (specifically in neurons, astrocytes, and microglia) and muscle tissue rather than lungs, liver, spleen, or kidneys (Dubey, et al., 1998, Luder, et al., 1999). Cysts have been found in several regions of the brain, including the cerebral cortex, superior and inferior colliculus, cerebellum, olfactory bulbs, and medulla oblongata, and have also been found in the spinal cord (Di Cristina, et al., 2008). It remains to be determined if there is something special about the intracellular environments of these host cells that favors spontaneous conversion to latent cysts in addition to being immunologically privileged sites.

The pathogenesis of Toxoplasma infection also relies in part on host genetics. Studies in humans and mice have revealed varying abilities to control infection and cyst burden (Brown, et al., 1995, Suzuki, et al., 1996, Mack, et al., 1999, Suzuki, 2002). For example, BALB/c mice are genetically resistant to Toxoplasma infection, harboring fewer brain cysts compared to other infected mouse strains. It was recently determined that BALB/c CD8+ T-cells can eliminate brain cysts through their perforin-mediated activity (Suzuki, et al., 2010). This study provides evidence that certain cytotoxic T-cells harbor a capacity to recognize and destroy intracellular brain cysts, which have been previously believed to be impervious to host immunity.

Reactivation of acute infection

In vitro generated bradyzoites will quickly revert to proliferative tachyzoites upon removal of the stress agent used to differentiate the parasites. These studies support the idea that cellular stress is a key factor not only in prompting development of bradyzoites, but also in maintaining the encysted form. In immunocompromised patients, reactivation of Toxoplasma infection is typically seen after the CD4+ T-cell count drops below 100–200 cells/mm3 (Pereira-Chioccola, et al., 2009) (Fig. 2). Ferguson et al. have reported that on rare occasion bradyzoite cysts will rupture in immunocompetent mice, leading to rapid recruitment of inflammatory cells to the site (Ferguson, et al., 1989). In the absence of a normal immune response to induce the newly released tachyzoites to convert into bradyzoites, the tachyzoites will continue to replicate and disseminate throughout the host, leading to serious complications and possibly death. Mouse models for reactivated toxoplasmosis have been developed that further underscore the relevance of IFN-γ in controlling latency (Suzuki & Joh, 1994, Dunay, et al., 2009). The steroid immunosuppressant dexamethasone has also been used to reactivate toxoplasmosis in chronically infected mice (Nicoll, et al., 1997, Djurkovic-Djakovic & Milenkovic, 2001). It is not currently known if host immunity directly prevents bradyzoites from reactivating and/or functions to quickly kill or trigger differentiation of reactivated tachyzoites escaping cysts.

Role of stress signaling in bradyzoite development and maintenance

As described earlier, virtually all types of stress can induce cyst development in vitro. Common denominators between the stresses that induce conversion to bradyzoites include the slowing of parasite proliferation, the induction of heat shock proteins, and translational control mediated by eIF2α phosphorylation (Fig. 5).

Figure 5
Stage differentiation of Toxoplasma

Bradyzoite development and the parasite cell cycle

Cell cycle blocks do not trigger bradyzoite differentiation, demonstrating that cell cycle progression is required for cyst formation (Gubbels, et al., 2008). Virtually all of the stress conditions that promote bradyzoite differentiation mentioned above reduce the proliferation of tachyzoites. Slowing of the parasite cell cycle has been linked to the initiation of the bradyzoite developmental program from a late-S/G2 subpopulation containing 1.8—2N DNA content (Jerome, et al., 1998, Radke, et al., 2003). During bradyzoite differentiation, these parasites proceed through M phase then arrest in G1/G0 with uniform 1N DNA content. The unique lateS/G2 stage represents a premitotic cell cycle checkpoint for the commitment to bradyzoite formation and growth arrest following mitosis. The identification of cyclical expression of several bradyzoite-specific mRNAs, which exhibit peak expression in the late mitotic period, lends support to this model (Behnke, et al., 2010). Curiously, one of the mRNAs with peak expression in tachyzoite cytokinesis is also significantly increased during bradyzoite differentiation; this mRNA encodes AP2VIIa-1, a likely transcription factor (discussed below). It is enticing to speculate that AP2VIIa-1 is a master regulator that coordinates changes in the transcriptome that lead to bradyzoite conversion. The nature of the signaling mechanisms that accompany slowed growth and ultimately lead to bradyzoite differentiation has yet to be fully elucidated; the following sections detail what has been determined to date.

Heat shock proteins and signaling pathways

One of the earliest described bradyzoite-specific genes is Hsp30/BAG1, related to the small heat shock proteins found in plants (Bohne, et al., 1995, Parmley, et al., 1995). BAG1 appears ~2–3 days after bradyzoite induction, but its relevance in differentiation was clouded when disruption of the gene in a type II strain failed to block tissue cyst formation (Bohne, et al., 1998). A similar study by Zhang et al. confirmed that BAG1 knockouts still form cysts in mice, but do so at a significantly lower frequency (Zhang, et al., 1999). Weiss et al. have demonstrated that Hsp70 is induced during alkaline-mediated bradyzoite differentiation, and that an inhibitor of Hsp90, Hsp70, and Hsp27 synthesis can suppress bradyzoite development in vitro (Weiss, et al., 1998). Another study also implicated a role for Hsp70 during reactivation of chronic toxoplasmosis in vivo (Silva, et al., 1998). Hsp90, in complex with p23 co-chaperone, has also been implicated in bradyzoite development (Echeverria, et al., 2010), and exhibits different localization patterns, being found in both the nucleus and cytoplasm in bradyzoites rather than just the cytoplasm (Echeverria, et al., 2005). Consistent with these data, SAGE libraries indicate that Hsp90 is detected early in bradyzoite differentiation (Radke, et al., 2005). Intriguingly, geldanamycin, an antibiotic that perturbs the normal function of Hsp90, blocks conversion of tachyzoites to bradyzoites as well as the reversion of bradyzoites to tachyzoites (Echeverria, et al., 2005). The mitochondrial chaperone Hsp60 exists as two alternatively spliced transcripts, both of which are up-regulated in bradyzoites (Toursel, et al., 2000). Upon bradyzoite induction, TgHsp60 localizes to two unknown vesicular bodies distinct from the parasite mitochondrion. Another heat shock protein, DnaK-tetratricopeptide repeat (DnaK-TPR), which also interacts with p23, was recently found to be expressed predominantly in bradyzoites (Ueno, et al., 2011). Three Hsp40/DnaJ family members were also found to be up-regulated in alkaline-stressed tachyzoites: TGME49_115690, TGME49_010430 and TGME49_023950 (Angel and Sullivan, unpublished).

Other conventional stress response signaling pathways well characterized in higher eukaryotes may also function in Toxoplasma, and possibly contribute to bradyzoite development. Homologues of mitogen activated protein kinases (MAPKs), which regulate diverse biological processes including cellular stress responses, have been identified in Toxoplasma (Lacey, et al., 2007). During in vitro differentiation to bradyzoites, mRNA levels for TgMAPK-1 increase, suggesting a possible role in stage conversion (Brumlik, et al., 2004). Increasing levels of cGMP and cAMP, as well as inhibiting downstream cGMP- and c-AMP-dependent kinases, have also been linked to bradyzoite differentiation (Kirkman, et al., 2001, Eaton, et al., 2006). The precise roles of Toxoplasma HSPs and signaling pathways during stage conversion warrant more detailed investigation.

It was recently discovered that stress conditions also induce the synthesis of the phytohormone abscisic acid (ABA) by the apicoplast (Nagamune, et al., 2008). ABA leads to the production of cyclic ADP ribose (cADPR), which controls release of intracellular calcium stores in Toxoplasma and induces egress. Pharmacological inhibition of ABA synthesis with fluridone blocked egress and triggered bradyzoite differentiation, suggesting ABA-mediated calcium signaling is an important factor in whether the parasite is lytic or latent (Nagamune, et al., 2008).

Translational control

Translational repression appears to operate in both tachyzoites and bradyzoites. Appreciable mRNAs encoding the bradyzoite-specific proteins G6-PI and MAG1 are readily detected in tachyzoites as well, suggesting that such messages are translationally repressed during the tachyzoite stage (Dzierszinski, et al., 1999, Weiss & Kim, 2000). Similarly, LDH1 protein is only expressed in bradyzoites, yet LDH1 mRNAs are equally detectable in both stages, suggesting that LDH1 is translationally repressed in bradyzoites (Yang & Parmley, 1997). The Toxoplasma eukaryotic translation initiation factor 4A (eIF4), which facilitates the binding of capped mRNA to the 40S ribosomal subunit, is down-regulated in bradyzoites, possibly reflective of the global reduction in protein synthesis during latency (Gastens & Fischer, 2002).

Recent studies have expanded and clarified the role of translational control in the parasite stress response and differentiation. Cellular stresses are well known inducers of protein translation control through phosphorylation of the alpha subunit of eukaryotic translation initiation factor-2 (eIF2α). When phosphorylated, eIF2α dampens global translation initiation, thereby favoring the preferential translation of a subset of mRNAs that encode proteins geared towards alleviating the stress (Wek, et al., 2006). As in other species, Toxoplasma phosphorylates its eIF2α orthologue (TgIF2α) in response to numerous stresses, including those that trigger bradyzoite differentiation (Sullivan Jr, et al., 2004). Alkaline pH, heat shock, sodium nitroprusside, sodium arsenite, and tunicamycin have all been shown to cause phosphorylation of TgIF2α (Sullivan Jr, et al., 2004, Narasimhan, et al., 2008). Moreover, TgIF2α remains phosphorylated in mature bradyzoites generated in vitro (Narasimhan, et al., 2008). We have also found that salubrinal, a specific inhibitor of eIF2α dephosphorylation (Boyce, et al., 2005), induces bradyzoite development in vitro (Narasimhan, et al., 2008). Collectively, these studies suggest a major role for TgIF2α phosphorylation (TgIF2α~P) in the development and maintenance of bradyzoite cysts. This idea has been supported in studies performed in the related apicomplexan parasite Plasmodium. Zhang et al. demonstrated that a Plasmodium eIF2 kinase designated IK2 controls the latency of sporozoites in the mosquito salivary gland (Zhang, et al., 2010).

TgIF2α~P may also facilitate Toxoplasma dissemination to immune privileged sites where bradyzoites tend to form. A greater capacity to survive outside host cells may help explain the increased ability of type I strains to disseminate throughout the host organism, particularly to the CNS. Two prevailing models describing how tachyzoites traverse biological barriers include the “Trojan horse” mechanism and paracellular transmigration (Elsheikha & Khan, 2010). Type I parasites are not as efficient as type II and III strains at inducing migratory phenotypes in infected dendritic cells (Lambert, et al., 2009), suggesting type I strains may rely more heavily on dissemination as extracellular parasites (Barragan & Sibley, 2002). Being ill-equipped to respond robustly to the stress of an extracellular environment, type II and III strains must remain confined to the intracellular environment, a limitation that is likely to contribute to their decreased virulence. Through generation of a non-phosphorylatable mutant, we have found that TgIF2α~P promotes survival of extracellular tachyzoites and does so in a strain-dependent manner (Joyce, et al., 2010). Type I RH strain shows rapid and robust TgIF2α~P within hours after egress, whereas the type II strain is slower to phosphorylate TgIF2α (Joyce, et al., 2010). It is tempting to speculate that the translational control induced by TgIF2α reprograms gene expression to protect extracellular parasites, and the increased viability of RH strain may stem from its increased capacity to phosphorylate TgIF2α for cytoprotection.

The relevance of eIF2α~P in Toxoplasma and Plasmodium latent forms underscores the need to characterize the parasite family of eIF2α kinases. Higher eukaryotes possess four eIF2α kinases, each appearing to respond to specific stress arrangements: GCN2 is activated by nutrient deprivation, PKR is activated by viral infection, PEK/Perk is activated by ER stress, and HRI is activated by heme deficiency and heat shock (Wek, et al., 2006). Toxoplasma also possesses four eIF2α kinases (TgIF2K-A through –D), some of which contain motifs suggestive of functional equivalency to mammalian enzymes. For example, based on the presence of a transmembrane domain, we hypothesized that TgIF2K-A may be equivalent to PEK/Perk. Indeed, TgIF2K-A localizes to the parasite ER and associates with the ER-resident chaperone BiP/GRP in a stress-dependent fashion (Narasimhan, et al., 2008). TgIF2K-D is most similar to GCN2, possessing a histidyl-tRNA synthetases (HisRS)-related domain that stimulates kinase activity by binding to uncharged tRNAs that accumulate during nutrient starvation. We have recently demonstrated that TgIF2K-D enhances the survival of extracellular tachyzoites deprived of host cell resources (Sullivan, unpublished). The roles of the other parasite eIF2 kinases, and how they become activated during different stress conditions, are important areas for future study.

Reprogramming the genome for bradyzoite development

In other species, stress-induced eIF2α~P reduces global protein synthesis in favor of a subset of mRNAs encoding factors needed to respond to the cellular insult or signal. Master regulator transcription factors such as GCN4/ATF4 are preferentially translated during stress, which subsequently reprogram the genome for stress remedy. Using polyribosome profiling and [35S]-Met/Cys labeling, we have verified that TgIF2α~P significantly curtails protein production (Narasimhan, et al., 2008). These data align with an earlier study showing that eIF4A, a DEAD-box RNA helicase that facilitates binding of capped mRNA to the 40S ribosomal subunit, is strongly down-regulated in bradyzoites (Gastens & Fischer, 2002). Recently, we have used Toxoplasma microarrays to identify mRNAs that are preferentially translated in the polyribosome fraction during ER stress (Sullivan, unpublished). Among these genes were several so-called AP2 proteins, which contain plant-like DNA-binding domains and represent a major lineage of transcription factors in Apicomplexa (Painter, et al., 2011). In the absence of GCN4/ATF4 homologues in Apicomplexa, it is plausible to speculate that AP2 proteins may be functional equivalents of these well-conserved master regulators.

Transcriptional regulation clearly plays a major role in bradyzoite development as evidenced by numerous studies showing stage-specific gene expression (Manger, et al., 1998, Cleary, et al., 2002, Radke, et al., 2005, Sullivan, et al., 2009). A seminal study generated multiple serial analysis of gene expression (SAGE) libraries to construct progressive “snapshots” of the developmental transition from sporozoite to tachyzoite to bradyzoite in different Toxoplasma strains (Radke, et al., 2005). Results from this study showed that distinct gene expression cascades occur through developmental transitions, underscoring the importance of transcriptional regulation throughout these events. Promoters that regulate BAG1 and a bradyzoite-specific NTPase during bradyzoite development were fine mapped to a 6–8 bp resolution and these minimal cis-elements were capable of converting a constitutive promoter to one that is induced by bradyzoite conditions (Behnke, et al., 2008). Together, these studies reveal that conventional eukaryotic promoter mechanisms are fundamentally at work to coordinate gene expression driving stage differentiation, but the usage of AP2 proteins as transcriptional regulators is different than higher eukaryotic counterparts. Details on the roles of AP2 factors, 11 of which are induced in bradyzoites, are emerging in ongoing studies.

Recently, the bradyzoite-specific promoter ENO1 was used as “bait” in an affinity-based strategy to isolate DNA-associating factors that may repress this gene in tachyzoites. Thirty-nine putative nuclear factors were found and divided into three categories: (i) 11 proteins with significant similarity to known nuclear factors, including a protein with a RNA-specific DEAD/DEAH box helicase domain, a pinin domain, and an FK506BP homologue, (ii) 7 proteins corresponding to kinases, phosphatases, and HSPs, and (iii) 21 hypothetical proteins, two of which have significant homology to Alba proteins, which are chromatin-associated silencing factors best characterized in Archaea. The FK506BP homologue (TgNF3) was extensively characterized. While not a direct binder of DNA, TgNF3 physically interacts with free core and nucleosome-associated histones to exert a gene silencing function at ENO1 and 18S ribosomal RNA genes, consistent with the yeast homologue known to be a histone chaperone that regulates rDNA silencing. Interestingly, TgNF3 is in the nucleus (enriched in the nucleolus) of tachyzoites but in the cytoplasm of bradyzoites (Olguin-Lamas, et al., 2011), suggesting that differential compartmentalization may influence activation of the bradyzoite ENO1 gene.

Another important factor implicated in contributing to bradyzoite gene expression is histone modification and chromatin remodeling (Bougdour, et al., 2010, Dixon, et al., 2010). The first indication that histone modifications play a role in Toxoplasma differentiation emerged when it was reported that histones at promoter regions of bradyzoite genes are hypoacetylated in tachyzoites and become acetylated after bradyzoite induction. Conversely, tachyzoite-specific genes that are hyperacetylated in the tachyzoite stage display decreased acetylation levels upon induction of differentiation (Saksouk, et al., 2005). The lysine (K) acetyltransferase (KAT) TgGCN5-A was localized to promoters of active developmentally regulated genes while the lysine deacetylase TgHDAC3 was situated at the promoters of inactive genes (Saksouk, et al., 2005). Further study of a TgGCN5-A knockout parasite has revealed a vital role for this KAT in expression of bradyzoite-specific genes during differentiation. Although TgGCN5-A appears dispensable in type I tachyzoites during normal culture conditions, the TgGCN5-A knockout parasites fail to upregulate bradyzoite marker genes upon induction of differentiation by alkaline stress (Naguleswaran, et al., 2010). The importance of histone acetylation for control of differentiation is underscored by the finding that chemical inhibition of TgHDAC3 with low doses of the compound FR235222 caused conversion to bradyzoites. The conversion was accompanied by hyperacetylation of the upstream regions of >350 genes, one third of which are specific to bradyzoites (Bougdour, et al., 2009).

While acetylation of histone lysine residues leads to gene activation, methylation of histone residues can result in either gene activation or repression. Monomethylation of lysine 20 of histone H4 (H4K20) by the methyltransferase TgSET8 promotes heterochromatin formation and subsequent gene silencing (Sautel, et al., 2007). Concurrent with the global downregulation of gene expression in the quiescent bradyzoite, monomethylation of H4K20 is significantly enriched (Sautel, et al., 2007), suggesting that TgSET8 is involved in maintenance of this transcriptionally suppressed state. In contrast, arginine methylation of histones has been linked to gene activation. Chemical inhibition of the arginine methyltransferase TgCARM1 triggers bradyzoite differentiation (Saksouk, et al., 2005). Consistent with this observation, TgCARM1 associates with the promoter regions of tachyzoite-specific genes in tachyzoites but is enriched at bradyzoite-specific genes upon induction of differentiation with alkaline pH (Saksouk, et al., 2005).

The Toxoplasma genome contains 17 predicted members of the SWI/SNF family of ATP-dependent nucleosome remodeling complexes (Dixon, et al., 2010), a number of which may be involved in stage conversion. An EST analysis of bradyzoites formed in vivo by Manger et al. identified a SNF2-like protein (TGME49_073870) that is upregulated during differentiation (Manger, et al., 1998). Message levels for another SNF2-like homologue, TgSRCAP, were shown to be upregulated during in vitro bradyzoite differentiation (Sullivan, et al., 2003).

Finally, alterations of nucleosome composition are another likely method of gene expression control in bradyzoite differentiation. Toxoplasma possesses novel H2A and H2B histone variants, substitutions of which can modulate gene expression (Sullivan, et al., 2006). The H2AZ and H2Bv variants are enriched at active genes, however the H2AX variant present at inactive genes is upregulated in the bradyzoite stage (Dalmasso, et al., 2009). Toxoplasma expresses many more chromatin remodelers that have yet to be fully characterized that are candidates contributing to the reprogramming of the genome during stage conversion (Dixon, et al., 2010).

Bradyzoite mutants

The haploid nature of tachyzoite/bradyzoite stages facilitates the generation of gene knockouts and disruptions for mutational analysis. The power of Toxoplasma as a molecular genetics system has been exploited to determine genes involved in stage conversion. A number of genes have been linked to having a role in bradyzoite differentiation using this approach, including the aforementioned BAG1 gene, the P-type H+-ATPase PMA1 (Holpert, et al., 2006), and the bradyzoite surface antigen SAG1-related sequence SRS9 (Kim, et al., 2007). Recently, knockouts of dense granule proteins GRA4 and GRA6 were shown to have dramatically reduced cyst burdens at 3 weeks post-infection in C57BL/6 mice, especially as a dual knockout (Fox, et al., 2011). A significant defect in cyst burden was also reported at 5 weeks post-infection for mutants lacking the entire 14 kb ROP4/7 locus (Fox, et al., 2011). It should be mentioned that mutations resulting in decreased cyst numbers in vivo may not directly correlate to effects on tachyzoite to bradyzoite conversion, as it is difficult to rule out effects on tachyzoite viability and dissemination in vivo.

To discover novel genes involved in bradyzoite development, various groups have generated developmental mutants. Singh et al. developed a transgenic parasite clone in type II Pru strain that contained GFP fused to the bradyzoite-specific promoter LDH2. After exposing these parasites to a chemical mutagen, variants defective in bradyzoite conversion (GFP negative) could be isolated using FACS following culture in bradyzoite inducing conditions (Singh, et al., 2002). Microarray analysis of the mutants following various stresses revealed a hierarchy of gene activation, supporting the idea that multiple induction conditions lead into a common pathway that reprograms for bradyzoite gene expression. It was noted that a 14-3-3 homologue, PITSLRE kinase, and a vacuolar ATPase exhibited decreased expression levels in most of the bradyzoite differentiation mutants (Singh, et al., 2002). A similar approach was followed to develop insertional mutants impaired in bradyzoite gene activation, leading to the discovery that a nucleolar CCHC zinc finger protein (ZFP1) and a pseudouridine synthase homologue (PUS1) are involved in bradyzoite differentiation (Vanchinathan, et al., 2005, Anderson, et al., 2009).

Matrajt et al. applied insertional mutagenesis on type I RH parasites lacking UPRT, which facilitates differentiation to bradyzoites following CO2 starvation (Matrajt, et al., 2002). Using a selectable marker driven by a bradyzoite-specific promoter, insertional mutants were isolated that have deficiencies in BAG1 expression and cyst wall formation. Like the mutants generated in the Singh study, these mutants replicated well under bradyzoite differentiation conditions, and microarray analysis revealed a gene expression profile more aligned with that seen for tachyzoites. Among the genes rescued and implicated as having a role in the differentiation process include a splicing factor, an oocyst wall protein, and AP2XII-6 (Lescault, et al., 2010). AP2XII-6 is particularly intriguing as AP2 domain proteins are potential transcription factors. One mutant contained an insert just upstream of a gene prediction for a DNA replication factor, but the mapped disrupted locus may be a non-coding RNA (Matrajt, 2010). Moreover, state modeling was used to capture hidden variation between parasite lines to reveal additional genes likely to be involved in bradyzoite differentiation, including transcription elongation factor Spt5, DNA primase subunit, TBC domain protein, HECT type ubiquitin ligase, and 6 hypothetical genes of unknown function (Lescault, et al., 2010).

The Knoll laboratory has generated and screened over 8000 insertional mutants by immunofluorescence microscopy for defects in bradyzoite cyst formation. Nine mutants were identified as defective in both cyst wall formation and expression of BAG1. One of these mutants contained an insertion in a gene encoding a serine/proline-rich proteophosphoglycan. This proteophosphoglycan is upregulated in bradyzoites and enhances cyst wall component expression and assembly through an unknown mechanism (Craver, et al., 2010). Another insertional mutant indicates that TgRSC8, which has homology to the catalytic component of the SWI/SNF and RSC (Remodel the Structure of Chromatin) complexes found in S. cerevisiae, is important for the upregulation of a subset of bradyzoite genes during in vitro differentiation (Rooney, et al., 2011). The latter further highlights the importance of chromatin remodeling during stage conversion.

How these gene products identified from the mutants contribute to bradyzoite conversion awaits further investigation. It is important to note that in each of these studies, the bradyzoite deficient mutants are leaky, suggesting that control of bradyzoite differentiation is a complex process with redundancies, and no single gene appears to be able to completely ablate bradyzoite formation.

Summary and Future Outlook

A schematic diagram highlighting recent key discoveries in the mechanisms of stage conversion is presented in Figure 5. Despite our advances, there are still many fundamental questions concerning bradyzoite differentiation to address, such as the composition of the cyst wall and matrix, how multiple signals are integrated into a common differentiation response, and which AP2 factors (or other transcription factors) operate as master regulators to coordinate the bradyzoite gene expression program.

As the molecular toolbox for parasite investigation expands, so will our understanding of bradyzoite differentiation. A major impediment to interrogating bradyzoite development is the lack of tools developed specifically for type II strains. The recent generation of Ku80 knockouts in type II Pru strain will greatly facilitate the study of genes linked to bradyzoite conversion (Fox, et al., 2011). Conditional knockout systems also need to be developed in type II backgrounds for the study of essential genes. Like EST and SAGE library predecessors, Toxoplasma microarrays (ToxoGeneChiP) continue to be utilized to resolve the bradyzoite transcriptome further, and ChIP-chip approaches can be applied to elucidate the bradyzoite epigenome; high-throughput sequencing promises to increase the resolution of these datasets even further. Methods that need improvement include techniques to purify tissue cysts from host cells, which would provide a clearer picture of the bradyzoite proteome and metabolome. Another important advance is the application of bioluminescence imaging to study the course of Toxoplasma infection, including the reactivation of chronic infection, in real time in living mice (Saeij, et al., 2005).

Future studies must also begin to focus on targeting bradyzoite formation and/or eliminating tissue cysts. Is it possible to develop avirulent, non-encysting Toxoplasma mutants for potential vaccines, particularly in livestock to reduce zoonotic transmission? The study of cytokines relevant to cyst formation and stability, and the observation that certain CD8+ T-cells can directly target cysts (Suzuki, et al., 2010), opens the door for immunomodulation approaches to eliminate cysts. More traditional pharmacological methods may still be useful in eradicating cysts if the drugs are conjugated to arginine oligomers, which have been shown to penetrate cyst walls (Samuel, et al., 2003). D-luciferin, used during bioluminescent imaging of infection, was also noted as being able to access and be metabolically processed by bradyzoites within tissue cysts (Di Cristina, et al., 2008). The latest discoveries illuminating the mechanics of how the bradyzoite gene program unfolds offer a rich repository of potential novel drug targets.

Numerous recent studies suggest that the persistence of Toxoplasma cysts in the brain may have a significant impact on the health of immunocompetent hosts. It is now well documented that infected rats undergo an exquisite change in behavior that would work to enhance transmission of the parasite back to its definitive host (Webster, 2007). Remarkably, rodents with latent Toxoplasma infection converted their aversion to feline odors into attraction, suggesting they would be more easily preyed upon in the field (Vyas, et al., 2007). Whether bradyzoite cysts within human brains affect behavior is less clear, and we are far from understanding the mechanisms (Fekadu, et al., 2010, Webster & McConkey, 2010).

A great deal of progress has been made since Toxoplasma tissue cysts were first observed in 1928, but much work remains to be completed to fully understand the molecular mechanisms driving latency and the impact of chronic infection on host behavior. The significance for continued study of latency in Toxoplasma is underscored by parallels between bradyzoites and other eukaryotic pathogens with dormant stages, exemplified by the discovery that translational control through eIF2α phosphorylation is a critical determinant in latent malarial sporozoites as well as Toxoplasma bradyzoites.


Research in the Sullivan laboratory is supported by grants from the National Institutes of Health (R01 AI077502, R21 AI084031, and R21 AI083732). We thank Drs. Louis Weiss (Albert Einstein College of Medicine) and David Ferguson (University of Oxford) for supplying images. Illustrations in Figures 1 and and55 were completed by Christopher M. Brown (Office of Visual Media, Indiana University School of Medicine). We also thank Drs. Louis Weiss and Sherry Queener (Indiana University School of Medicine) for critically reading the manuscript and providing helpful suggestions.


1. Al-Anouti F, Tomavo S, Parmley S, Ananvoranich S. The expression of lactate dehydrogenase is important for the cell cycle of Toxoplasma gondii. J Biol Chem. 2004;279:52300–52311. [PubMed]
2. Anderson MZ, Brewer J, Singh U, Boothroyd JC. A pseudouridine synthase homologue is critical to cellular differentiation in Toxoplasma gondii. Eukaryot Cell. 2009;8:398–409. [PMC free article] [PubMed]
3. Barragan A, Sibley LD. Transepithelial migration of Toxoplasma gondii is linked to parasite motility and virulence. J Exp Med. 2002;195:1625–1633. [PMC free article] [PubMed]
4. Behnke MS, Radke JB, Smith AT, Sullivan WJ, Jr, White MW. The transcription of bradyzoite genes in Toxoplasma gondii is controlled by autonomous promoter elements. Mol Microbiol. 2008;68:1502–1518. [PMC free article] [PubMed]
5. Behnke MS, Wootton JC, Lehmann MM, et al. Coordinated progression through two subtranscriptomes underlies the tachyzoite cycle of Toxoplasma gondii. PLoS One. 2010;5:e12354. [PMC free article] [PubMed]
6. Bohne W, Roos DS. Stage-specific expression of a selectable marker in Toxoplasma gondii permits selective inhibition of either tachyzoites or bradyzoites. Mol Biochem Parasitol. 1997;88:115–126. [PubMed]
7. Bohne W, Heesemann J, Gross U. Induction of bradyzoite-specific Toxoplasma gondii antigens in gamma interferon-treated mouse macrophages. Infect Immun. 1993;61:1141–1145. [PMC free article] [PubMed]
8. Bohne W, Heesemann J, Gross U. Reduced replication of Toxoplasma gondii is necessary for induction of bradyzoite-specific antigens: a possible role for nitric oxide in triggering stage conversion. Infect Immun. 1994;62:1761–1767. [PMC free article] [PubMed]
9. Bohne W, Gross U, Ferguson DJ, Heesemann J. Cloning and characterization of a bradyzoite-specifically expressed gene (hsp30/bag1) of Toxoplasma gondii, related to genes encoding small heat-shock proteins of plants. Mol Microbiol. 1995;16:1221–1230. [PubMed]
10. Bohne W, Hunter CA, White MW, Ferguson DJ, Gross U, Roos DS. Targeted disruption of the bradyzoite-specific gene BAG1 does not prevent tissue cyst formation in Toxoplasma gondii. Mol Biochem Parasitol. 1998;92:291–301. [PubMed]
11. Boothroyd JC, Black M, Bonnefoy S, et al. Genetic and biochemical analysis of development in Toxoplasma gondii. Philos Trans R Soc Lond B Biol Sci. 1997;352:1347–1354. [PMC free article] [PubMed]
12. Bougdour A, Braun L, Cannella D, Hakimi MA. Chromatin Modifications: Implications in the Regulation of Gene Expression in Toxoplasma gondii. Cell Microbiol. 2010 In press. [PubMed]
13. Bougdour A, Maubon D, Baldacci P, et al. Drug inhibition of HDAC3 and epigenetic control of differentiation in Apicomplexa parasites. J Exp Med. 2009;206:953–966. [PMC free article] [PubMed]
14. Boyce M, Bryant KF, Jousse C, et al. A selective inhibitor of eIF2alpha dephosphorylation protects cells from ER stress. Science. 2005;307:935–939. [PubMed]
15. Brown CR, Hunter CA, Estes RG, et al. Definitive identification of a gene that confers resistance against Toxoplasma cyst burden and encephalitis. Immunology. 1995;85:419–428. [PubMed]
16. Brumlik MJ, Wei S, Finstad K, et al. Identification of a novel mitogen-activated protein kinase in Toxoplasma gondii. Int J Parasitol. 2004;34:1245–1254. [PubMed]
17. Butcher BA, Greene RI, Henry SC, et al. p47 GTPases regulate Toxoplasma gondii survival in activated macrophages. Infect Immun. 2005;73:3278–3286. [PMC free article] [PubMed]
18. Cleary MD, Singh U, Blader IJ, Brewer JL, Boothroyd JC. Toxoplasma gondii asexual development: identification of developmentally regulated genes and distinct patterns of gene expression. Eukaryot Cell. 2002;1:329–340. [PMC free article] [PubMed]
19. Craver MP, Rooney PJ, Knoll LJ. Isolation of Toxoplasma gondii development mutants identifies a potential proteophosphogylcan that enhances cyst wall formation. Mol Biochem Parasitol. 2010;169:120–123. [PMC free article] [PubMed]
20. Dalmasso MC, Onyango DO, Naguleswaran A, Sullivan WJ, Jr, Angel SO. Toxoplasma H2A variants reveal novel insights into nucleosome composition and functions for this histone family. J Mol Biol. 2009;392:33–47. [PMC free article] [PubMed]
21. Daubener W, Spors B, Hucke C, Adam R, Stins M, Kim KS, Schroten H. Restriction of Toxoplasma gondii growth in human brain microvascular endothelial cells by activation of indoleamine 2,3-dioxygenase. Infect Immun. 2001;69:6527–6531. [PMC free article] [PubMed]
22. Denton H, Roberts CW, Alexander J, Thong KW, Coombs GH. Enzymes of energy metabolism in the bradyzoites and tachyzoites of Toxoplasma gondii. FEMS Microbiol Lett. 1996;137:103–108. [PubMed]
23. Di Cristina M, Marocco D, Galizi R, Proietti C, Spaccapelo R, Crisanti A. Temporal and spatial distribution of Toxoplasma gondii differentiation into Bradyzoites and tissue cyst formation in vivo. Infect Immun. 2008;76:3491–3501. [PMC free article] [PubMed]
24. Dixon SE, Stilger KL, Elias EV, Naguleswaran A, Sullivan WJ., Jr A decade of epigenetic research in Toxoplasma gondii. Mol Biochem Parasitol. 2010;173:1–9. [PMC free article] [PubMed]
25. Djurkovic-Djakovic O, Milenkovic V. Murine model of drug-induced reactivation of Toxoplasma gondii. Acta Protozool. 2001;40:99–106.
26. Djurkovic-Djakovic O, Nikolic A, Bobic B, Klun I, Aleksic A. Stage conversion of Toxoplasma gondii RH parasites in mice by treatment with atovaquone and pyrrolidine dithiocarbamate. Microbes Infect. 2005;7:49–54. [PubMed]
27. Donald RG, Allocco J, Singh SB, Nare B, Salowe SP, Wiltsie J, Liberator PA. Toxoplasma gondii cyclic GMP-dependent kinase: chemotherapeutic targeting of an essential parasite protein kinase. Eukaryot Cell. 2002;1:317–328. [PMC free article] [PubMed]
28. Dubey JP, Jones JL. Toxoplasma gondii infection in humans and animals in the United States. Int J Parasitol. 2008;38:1257–1278. [PubMed]
29. Dubey JP, Lindsay DS, Speer CA. Structures of Toxoplasma gondii tachyzoites, bradyzoites, and sporozoites and biology and development of tissue cysts. Clin Microbiol Rev. 1998;11:267–299. [PMC free article] [PubMed]
30. Dunay IR, Chan WC, Haynes RK, Sibley LD. Artemisone and artemiside control acute and reactivated toxoplasmosis in a murine model. Antimicrob Agents Chemother. 2009;53:4450–4456. [PMC free article] [PubMed]
31. Dzierszinski F, Nishi M, Ouko L, Roos DS. Dynamics of Toxoplasma gondii differentiation. Eukaryot Cell. 2004;3:992–1003. [PMC free article] [PubMed]
32. Dzierszinski F, Mortuaire M, Dendouga N, Popescu O, Tomavo S. Differential expression of two plant-like enolases with distinct enzymatic and antigenic properties during stage conversion of the protozoan parasite Toxoplasma gondii. J Mol Biol. 2001;309:1017–1027. [PubMed]
33. Dzierszinski F, Popescu O, Toursel C, Slomianny C, Yahiaoui B, Tomavo S. The protozoan parasite Toxoplasma gondii expresses two functional plant-like glycolytic enzymes. Implications for evolutionary origin of apicomplexans. J Biol Chem. 1999;274:24888–24895. [PubMed]
34. Eaton MS, Weiss LM, Kim K. Cyclic nucleotide kinases and tachyzoite-bradyzoite transition in Toxoplasma gondii. Int J Parasitol. 2006;36:107–114. [PMC free article] [PubMed]
35. Echeverria PC, Matrajt M, Harb OS, et al. Toxoplasma gondii Hsp90 is a potential drug target whose expression and subcellular localization are developmentally regulated. J Mol Biol. 2005;350:723–734. [PubMed]
36. Echeverria PC, Figueras MJ, Vogler M, et al. The Hsp90 co-chaperone p23 of Toxoplasma gondii: Identification, functional analysis and dynamic interactome determination. Mol Biochem Parasitol. 2010;172:129–140. [PMC free article] [PubMed]
37. Elsheikha HM, Khan NA. Protozoa traversal of the blood-brain barrier to invade the central nervous system. FEMS Microbiol Rev. 2010;34:532–553. [PubMed]
38. Fekadu A, Shibre T, Cleare AJ. Toxoplasmosis as a cause for behaviour disorders--overview of evidence and mechanisms. Folia Parasitol (Praha) 2010;57:105–113. [PubMed]
39. Fentress SJ, Behnke MS, Dunay IR, et al. Phosphorylation of immunity-related GTPases by a Toxoplasma gondii-secreted kinase promotes macrophage survival and virulence. Cell Host Microbe. 2010;8:484–495. [PMC free article] [PubMed]
40. Feo S, Arcuri D, Piddini E, Passantino R, Giallongo A. ENO1 gene product binds to the c-myc promoter and acts as a transcriptional repressor: relationship with Myc promoter-binding protein 1 (MBP-1) FEBS Lett. 2000;473:47–52. [PubMed]
41. Ferguson DJ. Use of molecular and ultrastructural markers to evaluate stage conversion of Toxoplasma gondii in both the intermediate and definitive host. Int J Parasitol. 2004;34:347–360. [PubMed]
42. Ferguson DJ, Hutchison WM. An ultrastructural study of the early development and tissue cyst formation of Toxoplasma gondii in the brains of mice. Parasitol Res. 1987;73:483–491. [PubMed]
43. Ferguson DJ, Hutchison WM, Pettersen E. Tissue cyst rupture in mice chronically infected with Toxoplasma gondii. An immunocytochemical and ultrastructural study. Parasitol Res. 1989;75:599–603. [PubMed]
44. Ferguson DJ, Parmley SF, Tomavo S. Evidence for nuclear localisation of two stage-specific isoenzymes of enolase in Toxoplasma gondii correlates with active parasite replication. Int J Parasitol. 2002;32:1399–1410. [PubMed]
45. Ferreira da Silva MD, Barbosa HS, Gross U, Luder CG. Stress-related and spontaneous stage differentiation of Toxoplasma gondii. Mol Biosyst. 2008;4:824–834. [PubMed]
46. Fox BA, Gigley JP, Bzik DJ. Toxoplasma gondii lacks the enzymes required for de novo arginine biosynthesis and arginine starvation triggers cyst formation. Int J Parasitol. 2004;34:323–331. [PubMed]
47. Fox BA, Ristuccia JG, Gigley JP, Bzik DJ. Efficient gene replacements in Toxoplasma gondii strains deficient for nonhomologous end joining. Eukaryot Cell. 2009;8:520–529. [PMC free article] [PubMed]
48. Fox BA, Falla A, Rommereim LM, et al. Type II Toxoplasma gondii KU80 Knockout Strains Enable Functional Analysis of Genes Required for Cyst Development and Latent Infection. Eukaryotic cell 2011 [PMC free article] [PubMed]
49. Fux B, Nawas J, Khan A, Gill DB, Su C, Sibley LD. Toxoplasma gondii strains defective in oral transmission are also defective in developmental stage differentiation. Infect Immun. 2007;75:2580–2590. [PMC free article] [PubMed]
50. Gastens MH, Fischer HG. Toxoplasma gondii eukaryotic translation initiation factor 4A associated with tachyzoite virulence is down-regulated in the bradyzoite stage. Int J Parasitol. 2002;32:1225–1234. [PubMed]
51. Gazzinelli R, Xu Y, Hieny S, Cheever A, Sher A. Simultaneous depletion of CD4+ and CD8+ T lymphocytes is required to reactivate chronic infection with Toxoplasma gondii. J Immunol. 1992;149:175–180. [PubMed]
52. Gazzinelli RT, Hakim FT, Hieny S, Shearer GM, Sher A. Synergistic role of CD4+ and CD8+ T lymphocytes in IFN-gamma production and protective immunity induced by an attenuated Toxoplasma gondii vaccine. J Immunol. 1991;146:286–292. [PubMed]
53. Gubbels MJ, White M, Szatanek T. The cell cycle and Toxoplasma gondii cell division: tightly knit or loosely stitched? Int J Parasitol. 2008;38:1343–1358. [PubMed]
54. Holpert M, Gross U, Bohne W. Disruption of the bradyzoite-specific P-type (H+)-ATPase PMA1 in Toxoplasma gondii leads to decreased bradyzoite differentiation after stress stimuli but does not interfere with mature tissue cyst formation. Mol Biochem Parasitol. 2006;146:129–133. [PubMed]
55. Howe DK, Sibley LD. Toxoplasma gondii comprises three clonal lineages: correlation of parasite genotype with human disease. J Infect Dis. 1995;172:1561–1566. [PubMed]
56. Howe DK, Honore S, Derouin F, Sibley LD. Determination of genotypes of Toxoplasma gondii strains isolated from patients with toxoplasmosis. J Clin Microbiol. 1997;35:1411–1414. [PMC free article] [PubMed]
57. Huynh MH, Carruthers VB. Tagging of endogenous genes in a Toxoplasma gondii strain lacking Ku80. Eukaryot Cell. 2009;8:530–539. [PMC free article] [PubMed]
58. Jacobs L, Remington JS, Melton ML. The resistance of the encysted form of Toxoplasma gondii. J Parasitol. 1960;46:11–21. [PubMed]
59. Jerome ME, Radke JR, Bohne W, Roos DS, White MW. Toxoplasma gondii bradyzoites form spontaneously during sporozoite-initiated development. Infect Immun. 1998;66:4838–4844. [PMC free article] [PubMed]
60. Jones J, Lopez A, Wilson M. Congenital toxoplasmosis. Am Fam Physician. 2003;67:2131–2138. [PubMed]
61. Jones TC, Bienz KA, Erb P. In vitro cultivation of Toxoplasma gondii cysts in astrocytes in the presence of gamma interferon. Infection and immunity. 1986;51:147–156. [PMC free article] [PubMed]
62. Joyce BR, Queener SF, Wek RC, Sullivan WJ., Jr Phosphorylation of eukaryotic initiation factor-2{alpha} promotes the extracellular survival of obligate intracellular parasite Toxoplasma gondii. Proc Natl Acad Sci U S A. 2010;107:17200–17205. [PubMed]
63. Kasper LH, Khan IA, Ely KH, Buelow R, Boothroyd JC. Antigen-specific (p30) mouse CD8+ T cells are cytotoxic against Toxoplasma gondii-infected peritoneal macrophages. J Immunol. 1992;148:1493–1498. [PubMed]
64. Khan A, Behnke MS, Dunay IR, White MW, Sibley LD. Phenotypic and gene expression changes among clonal type I strains of Toxoplasma gondii. Eukaryot Cell. 2009;8:1828–1836. [PMC free article] [PubMed]
65. Kim SK, Karasov A, Boothroyd JC. Bradyzoite-specific surface antigen SRS9 plays a role in maintaining Toxoplasma gondii persistence in the brain and in host control of parasite replication in the intestine. Infect Immun. 2007;75:1626–1634. [PMC free article] [PubMed]
66. Kirkman LA, Weiss LM, Kim K. Cyclic nucleotide signaling in Toxoplasma gondii bradyzoite differentiation. Infect Immun. 2001;69:148–153. [PMC free article] [PubMed]
67. Lacey MR, Brumlik MJ, Yenni RE, Burow ME, Curiel TJ. Toxoplasma gondii expresses two mitogen-activated protein kinase genes that represent distinct protozoan subfamilies. J Mol Evol. 2007;64:4–14. [PubMed]
68. Lambert H, Vutova PP, Adams WC, Lore K, Barragan A. The Toxoplasma gondii-shuttling function of dendritic cells is linked to the parasite genotype. Infect Immun. 2009;77:1679–1688. [PMC free article] [PubMed]
69. Lane A, Soete M, Dubremetz JF, Smith JE. Toxoplasma gondii: appearance of specific markers during the development of tissue cysts in vitro. Parasitol Res. 1996;82:340–346. [PubMed]
70. Lescault PJ, Thompson AB, Patil V, et al. Genomic data reveal Toxoplasma gondii differentiation mutants are also impaired with respect to switching into a novel extracellular tachyzoite state. PLoS One. 2010;5:e14463. [PMC free article] [PubMed]
71. Luder CG, Giraldo-Velasquez M, Sendtner M, Gross U. Toxoplasma gondii in primary rat CNS cells: differential contribution of neurons, astrocytes, and microglial cells for the intracerebral development and stage differentiation. Exp Parasitol. 1999;93:23–32. [PubMed]
72. Luft BJ, Brooks RG, Conley FK, McCabe RE, Remington JS. Toxoplasmic encephalitis in patients with acquired immune deficiency syndrome. JAMA. 1984;252:913–917. [PubMed]
73. Mack DG, Johnson JJ, Roberts F, et al. HLA-class II genes modify outcome of Toxoplasma gondii infection. Int J Parasitol. 1999;29:1351–1358. [PubMed]
74. Manger ID, Hehl A, Parmley S, et al. Expressed sequence tag analysis of the bradyzoite stage of Toxoplasma gondii: identification of developmentally regulated genes. Infect Immun. 1998;66:1632–1637. [PMC free article] [PubMed]
75. Matrajt M. Non-coding RNA in apicomplexan parasites. Mol Biochem Parasitol. 2010;174:1–7. [PMC free article] [PubMed]
76. Matrajt M, Donald RG, Singh U, Roos DS. Identification and characterization of differentiation mutants in the protozoan parasite Toxoplasma gondii. Mol Microbiol. 2002;44:735–747. [PubMed]
77. McCabe RE, Luft BJ, Remington JS. Effect of murine interferon gamma on murine toxoplasmosis. J Infect Dis. 1984;150:961–962. [PubMed]
78. McFadden GI. The apicoplast. Protoplasma 2010
79. McHugh TD, Gbewonyo A, Johnson JD, Holliman RE, Butcher PD. Development of an in vitro model of Toxoplasma gondii cyst formation. FEMS Microbiol Lett. 1993;114:325–332. [PubMed]
80. Mehlhorn H, Frenkel JK. Ultrastructural comparison of cysts and zoites of Toxoplasma gondii, Sarcocystis muris, and Hammondia hammondi in skeletal muscle of mice. J Parasitol. 1980;66:59–67. [PubMed]
81. Meissner M, Agop-Nersesian C, Sullivan WJ., Jr Molecular tools for analysis of gene function in parasitic microorganisms. Appl Microbiol Biotechnol. 2007;75:963–975. [PubMed]
82. Mercier C, Howe DK, Mordue D, Lingnau M, Sibley LD. Targeted disruption of the GRA2 locus in Toxoplasma gondii decreases acute virulence in mice. Infect Immun. 1998;66:4176–4182. [PMC free article] [PubMed]
83. Montoya JG, Liesenfeld O. Toxoplasmosis. Lancet. 2004;363:1965–1976. [PubMed]
84. Nagamune K, Hicks LM, Fux B, Brossier F, Chini EN, Sibley LD. Abscisic acid controls calcium-dependent egress and development in Toxoplasma gondii. Nature. 2008;451:207–210. [PMC free article] [PubMed]
85. Naguleswaran A, Elias EV, McClintick J, Edenberg HJ, Sullivan WJ., Jr Toxoplasma gondii lysine acetyltransferase GCN5-A functions in the cellular response to alkaline stress and expression of cyst genes. PLoS Pathog. 2010;6:e1001232. [PMC free article] [PubMed]
86. Narasimhan J, Joyce BR, Naguleswaran A, et al. Translation Regulation by Eukaryotic Initiation Factor-2 Kinases in the Development of Latent Cysts in Toxoplasma gondii. J Biol Chem. 2008;283:16591–16601. [PMC free article] [PubMed]
87. Nicoll S, Wright S, Maley SW, Burns S, Buxton D. A mouse model of recrudescence of Toxoplasma gondii infection. J Med Microbiol. 1997;46:263–266. [PubMed]
88. Olguin-Lamas A, Madec E, Hovasse A, et al. A Novel Toxoplasma gondii Nuclear Factor TgNF3 Is a Dynamic Chromatin-Associated Component, Modulator of Nucleolar Architecture and Parasite Virulence. PLoS pathogens. 2011;7:e1001328. [PMC free article] [PubMed]
89. Painter HJ, Campbell TL, Llinas M. The Apicomplexan AP2 family: integral factors regulating Plasmodium development. Mol Biochem Parasitol. 2011;176:1–7. [PMC free article] [PubMed]
90. Parmley SF, Weiss LM, Yang S. Cloning of a bradyzoite-specific gene of Toxoplasma gondii encoding a cytoplasmic antigen. Mol Biochem Parasitol. 1995;73:253–257. [PubMed]
91. Parmley SF, Yang S, Harth G, Sibley LD, Sucharczuk A, Remington JS. Molecular characterization of a 65-kilodalton Toxoplasma gondii antigen expressed abundantly in the matrix of tissue cysts. Mol Biochem Parasitol. 1994;66:283–296. [PubMed]
92. Pereira-Chioccola VL, Vidal JE, Su C. Toxoplasma gondii infection and cerebral toxoplasmosis in HIV-infected patients. Future Microbiol. 2009;4:1363–1379. [PubMed]
93. Pfefferkorn ER, Eckel M, Rebhun S. Interferon-gamma suppresses the growth of Toxoplasma gondii in human fibroblasts through starvation for tryptophan. Mol Biochem Parasitol. 1986;20:215–224. [PubMed]
94. Radke JR, White MW. A cell cycle model for the tachyzoite of Toxoplasma gondii using the Herpes simplex virus thymidine kinase. Mol Biochem Parasitol. 1998;94:237–247. [PubMed]
95. Radke JR, Guerini MN, Jerome M, White MW. A change in the premitotic period of the cell cycle is associated with bradyzoite differentiation in Toxoplasma gondii. Mol Biochem Parasitol. 2003;131:119–127. [PubMed]
96. Radke JR, Behnke MS, Mackey AJ, Radke JB, Roos DS, White MW. The transcriptome of Toxoplasma gondii. BMC Biol. 2005;3:26. [PMC free article] [PubMed]
97. Radke JR, Donald RG, Eibs A, Jerome ME, Behnke MS, Liberator P, White MW. Changes in the expression of human cell division autoantigen-1 influence Toxoplasma gondii growth and development. PLoS Pathog. 2006;2:e105. [PMC free article] [PubMed]
98. Rooney PJ, Neal LM, Knoll LJ. Involvement of an RSC complex ortholog in developmental regulation in the parasite Toxoplasma gondii. PLoS One. 2011 In press. [PMC free article] [PubMed]
99. Sabin A. Toxoplasmic encephalitis in children. J Am Med Assoc. 1941;116:801–807.
100. Saeij JP, Boyle JP, Grigg ME, Arrizabalaga G, Boothroyd JC. Bioluminescence imaging of Toxoplasma gondii infection in living mice reveals dramatic differences between strains. Infect Immun. 2005;73:695–702. [PMC free article] [PubMed]
101. Saksouk N, Bhatti MM, Kieffer S, et al. Histone-modifying complexes regulate gene expression pertinent to the differentiation of the protozoan parasite Toxoplasma gondii. Mol Cell Biol. 2005;25:10301–10314. [PMC free article] [PubMed]
102. Samuel BU, Hearn B, Mack D, et al. Delivery of antimicrobials into parasites. Proc Natl Acad Sci U S A. 2003;100:14281–14286. [PubMed]
103. Sautel CF, Cannella D, Bastien O, et al. SET8-mediated methylations of histone H4 lysine 20 mark silent heterochromatic domains in apicomplexan genomes. Mol Cell Biol. 2007;27:5711–5724. [PMC free article] [PubMed]
104. Sethi KK, Rahman A, Pelster B, Brandis H. Search for the presence of lectin-binding sites on Toxoplasma gondii. J Parasitol. 1977;63:1076–1080. [PubMed]
105. Silva NM, Gazzinelli RT, Silva DA, Ferro EA, Kasper LH, Mineo JR. Expression of Toxoplasma gondii-specific heat shock protein 70 during In vivo conversion of bradyzoites to tachyzoites. Infect Immun. 1998;66:3959–3963. [PMC free article] [PubMed]
106. Singh U, Brewer JL, Boothroyd JC. Genetic analysis of tachyzoite to bradyzoite differentiation mutants in Toxoplasma gondii reveals a hierarchy of gene induction. Mol Microbiol. 2002;44:721–733. [PubMed]
107. Soete M, Camus D, Dubremetz JF. Experimental induction of bradyzoite-specific antigen expression and cyst formation by the RH strain of Toxoplasma gondii in vitro. Exp Parasitol. 1994;78:361–370. [PubMed]
108. Steinfeldt T, Konen-Waisman S, Tong L, et al. Phosphorylation of mouse immunity-related GTPase (IRG) resistance proteins is an evasion strategy for virulent Toxoplasma gondii. PLoS Biol. 2010;8:e1000576. [PMC free article] [PubMed]
109. Su C, Evans D, Cole RH, Kissinger JC, Ajioka JW, Sibley LD. Recent expansion of Toxoplasma through enhanced oral transmission. Science. 2003;299:414–416. [PubMed]
110. Sullivan WJ, Jr, Narasimhan J, Bhatti MM, Wek RC. Parasite-specific eukaryotic initiation factor -2 (eIF2) kinase required for stress-induced translation control. Biochem J. 2004;380:523–531. [PubMed]
111. Sullivan WJ, Jr, Naguleswaran A, Angel SO. Histones and histone modifications in protozoan parasites. Cell Microbiol. 2006;8:1850–1861. [PubMed]
112. Sullivan WJ, Jr, Smith AT, Joyce BR. Understanding mechanisms and the role of differentiation in pathogenesis of Toxoplasma gondii: a review. Mem Inst Oswaldo Cruz. 2009;104:155–161. [PMC free article] [PubMed]
113. Sullivan WJ, Jr, Monroy MA, Bohne W, et al. Molecular cloning and characterization of an SRCAP chromatin remodeling homologue in Toxoplasma gondii. Parasitol Res. 2003;90:1–8. [PubMed]
114. Suzuki Y. Host resistance in the brain against Toxoplasma gondii. J Infect Dis. 2002;185(Suppl 1):S58–65. [PubMed]
115. Suzuki Y, Joh K. Effect of the strain of Toxoplasma gondii on the development of toxoplasmic encephalitis in mice treated with antibody to interferon-gamma. Parasitol Res. 1994;80:125–130. [PubMed]
116. Suzuki Y, Orellana MA, Schreiber RD, Remington JS. Interferon-gamma: the major mediator of resistance against Toxoplasma gondii. Science. 1988;240:516–518. [PubMed]
117. Suzuki Y, Wang X, Jortner BS, et al. Removal of Toxoplasma gondii cysts from the brain by perforin-mediated activity of CD8+ T cells. Am J Pathol. 2010;176:1607–1613. [PubMed]
118. Suzuki Y, Wong SY, Grumet FC, et al. Evidence for genetic regulation of susceptibility to toxoplasmic encephalitis in AIDS patients. J Infect Dis. 1996;173:265–268. [PubMed]
119. Tenter AM, Heckeroth AR, Weiss LM. Toxoplasma gondii: from animals to humans. Int J Parasitol. 2000;30:1217–1258. [PMC free article] [PubMed]
120. Tomavo S, Boothroyd JC. Interconnection between organellar functions, development and drug resistance in the protozoan parasite, Toxoplasma gondii. Int J Parasitol. 1995;25:1293–1299. [PubMed]
121. Toursel C, Dzierszinski F, Bernigaud A, Mortuaire M, Tomavo S. Molecular cloning, organellar targeting and developmental expression of mitochondrial chaperone HSP60 in Toxoplasma gondii. Mol Biochem Parasitol. 2000;111:319–332. [PubMed]
122. Ueno A, Dautu G, Haga K, Munyaka B, Carmen G, Kobayashi Y, Igarashi M. Toxoplasma gondii: A bradyzoite-specific DnaK-tetratricopeptide repeat (DnaK-TPR) protein interacts with p23 co-chaperone protein. Exp Parasitol 2011 [PubMed]
123. Vanchinathan P, Brewer JL, Harb OS, Boothroyd JC, Singh U. Disruption of a locus encoding a nucleolar zinc finger protein decreases tachyzoite-to-bradyzoite differentiation in Toxoplasma gondii. Infect Immun. 2005;73:6680–6688. [PMC free article] [PubMed]
124. Vyas A, Kim SK, Giacomini N, Boothroyd JC, Sapolsky RM. Behavioral changes induced by Toxoplasma infection of rodents are highly specific to aversion of cat odors. Proc Natl Acad Sci U S A. 2007;104:6442–6447. [PubMed]
125. Wallace GR, Stanford MR. Immunity and Toxoplasma retinochoroiditis. Clinical and experimental immunology. 2008;153:309–315. [PubMed]
126. Webster JP. The effect of Toxoplasma gondii on animal behavior: playing cat and mouse. Schizophr Bull. 2007;33:752–756. [PMC free article] [PubMed]
127. Webster JP, McConkey GA. Toxoplasma gondii-altered host behaviour: clues as to mechanism of action. Folia Parasitol (Praha) 2010;57:95–104. [PubMed]
128. Weiss LM, Kim K. The development and biology of bradyzoites of Toxoplasma gondii. Front Biosci. 2000;5:D391–405. [PMC free article] [PubMed]
129. Weiss LM, Ma YF, Takvorian PM, Tanowitz HB, Wittner M. Bradyzoite development in Toxoplasma gondii and the hsp70 stress response. Infect Immun. 1998;66:3295–3302. [PMC free article] [PubMed]
130. Weiss LM, Laplace D, Takvorian PM, Tanowitz HB, Cali A, Wittner M. A cell culture system for study of the development of Toxoplasma gondii bradyzoites. J Eukaryot Microbiol. 1995;42:150–157. [PubMed]
131. Wek RC, Jiang HY, Anthony TG. Coping with stress: eIF2 kinases and translational control. Biochem Soc Trans. 2006;34:7–11. [PubMed]
132. Wong SY, Remington JS. Biology of Toxoplasma gondii. AIDS. 1993;7:299–316. [PubMed]
133. Yahiaoui B, Dzierszinski F, Bernigaud A, Slomianny C, Camus D, Tomavo S. Isolation and characterization of a subtractive library enriched for developmentally regulated transcripts expressed during encystation of Toxoplasma gondii. Mol Biochem Parasitol. 1999;99:223–235. [PubMed]
134. Yang S, Parmley SF. Toxoplasma gondii expresses two distinct lactate dehydrogenase homologous genes during its life cycle in intermediate hosts. Gene. 1997;184:1–12. [PubMed]
135. Yap GS, Scharton-Kersten T, Ferguson DJ, Howe D, Suzuki Y, Sher A. Partially protective vaccination permits the development of latency in a normally virulent strain of Toxoplasma gondii. Infect Immun. 1998;66:4382–4388. [PMC free article] [PubMed]
136. Zhang M, Fennell C, Ranford-Cartwright L, et al. The Plasmodium eukaryotic initiation factor-2alpha kinase IK2 controls the latency of sporozoites in the mosquito salivary glands. J Exp Med. 2010;207:1465–1474. [PMC free article] [PubMed]
137. Zhang YW, Halonen SK, Ma YF, Wittner M, Weiss LM. Initial characterization of CST1, a Toxoplasma gondii cyst wall glycoprotein. Infect Immun. 2001;69:501–507. [PMC free article] [PubMed]
138. Zhang YW, Kim K, Ma YF, Wittner M, Tanowitz HB, Weiss LM. Disruption of the Toxoplasma gondii bradyzoite-specific gene BAG1 decreases in vivo cyst formation. Mol Microbiol. 1999;31:691–701. [PMC free article] [PubMed]