The molecular biology and immune control of chronic Toxoplasma gondii infection

Toxoplasma gondii is an incredibly successful parasite owing in part to its ability to persist within cells for the life of the host. Remarkably, at least 350 host species of T. gondii have been described to date, and it is estimated that 30% of the global human population is chronically infected. The importance of T. gondii in human health was made clear with the first reports of congenital toxoplasmosis in the 1940s. However, the AIDS crisis in the 1980s revealed the prevalence of chronic infection, as patients presented with reactivated chronic toxoplasmosis, underscoring the importance of an intact immune system for parasite control. In the last 40 years, there has been tremendous progress toward understanding the biology of T. gondii infection using rodent models, human cell experimental systems, and clinical data. However, there are still major holes in our understanding of T. gondii biology, including the genes controlling parasite development, the mechanisms of cell-intrinsic immunity to T. gondii in the brain and muscle, and the long-term effects of infection on host homeostasis. The need to better understand the biology of chronic infection is underscored by the recent rise in ocular disease associated with emerging haplotypes of T. gondii and our lack of effective treatments to sterilize chronic infection. This Review discusses the cell types and molecular mediators, both host and parasite, that facilitate persistent T. gondii infection. We highlight the consequences of chronic infection for tissue-specific pathology and identify open questions in this area of host-Toxoplasma interactions.

In North America and Europe, environmental isolates of T. gondii predominantly belong to three major strains or types: type I, type II, and type III. These types are notable in that virulence, measured by lethal dose (LD), differs by several logs in inbred strains of mice (e.g., C57BL/6, CBA/J, BALB/c). Type I is the most virulent (LD₁₀₀ of 1–10 tachyzoites), compared with type II (LD₅₀ of 100–1000) and type III (LD₅₀ of ~100,000 to 1 million), which are substantially less aggressive in vivo (31, 32). Human infection is dominated by type II in North America and Europe; however, a fourth type, haplogroup 12, was recently isolated from North American patients and wild animals (33). In Asia and Africa, region-specific clonal lineages have also been isolated (34). Strains that do not fit the pattern of clonal lineage expansion have been identified in South America belonging to haplogroups 4 through 15 (35–37). Whole genome sequencing indicates that genomic admixture and recombination between a limited number of ancestral strains account for T. gondii’s genetic diversity. The relationship between strains was determined by comparison of the inheritance pattern of large gene haploblocks. These haploblocks encode virulence-associated, secreted parasite effector proteins, suggesting that the unique assortment of effector alleles may also control pathogenesis and/or transmission rates (38). Importantly, there is evidence of coevolution between virulent haplotypes and mouse IFN-inducible immunity-related GTPase (IRG) genes, which are critical mediators of cell-intrinsic parasite killing in mouse models (39, 40).

Activating a robust immune response is critical to both host and parasite survival. Bradyzoite cysts are resistant to peptic proteases, but tachyzoites are not: if the host dies before chronic infection is established, parasite transmission does not occur. In keeping with this paradigm, the parasite has evolved effectors that selectively activate host immune cell signaling in addition to strategies to avoid sterilizing immunity. In acute and chronic infection, T. gondii grows and persists within a parasitophorous vacuole membrane (PVM). The PVM is generated from the host plasma membrane as the parasite ratchets its way into the cell using injected parasite effector proteins. This process avoids the lytic environment of the endo/lysosomal compartments and accounts for the parasite’s remarkable capacity to infect almost any nucleated cell type in vitro (41). In vivo, however, the cell types harboring the parasite are more limited. After a parasite cyst is ingested, T. gondii invades the distal jejunum of the small intestine in mice (42, 43). The precise host cell types mediating invasion (e.g., M cells, epithelial cells) are not clear; however, T. gondii sporozoites and tachyzoites have been observed in intestinal epithelial cells (44). Tachyzoites are also observed within infiltrating immune cells (43–45). The CCL2/CCR2 chemokine axis is a conserved mechanism of monocyte recruitment in mice and humans (46). The parasite effectors Tg14-3-3 and TgWIP have been shown to promote hypermotility in infected human and murine dendritic cells (47, 48). Dendritic cell hypermotility has been observed in vivo, suggesting that it may be a stealth mechanism that facilitates parasite dissemination while avoiding detection by circulating immune effectors (49, 50).

The long evolutionary relationship between T. gondii and mammalian hosts is evident in analysis of the pathways infected cells use to detect and destroy the parasite (39). Three main arms of innate immune sensing have been described in T. gondii infection: the Toll-like receptors (TLRs), the IFN-inducible GTPases, and the inflammasomes. TLR and IL-1 receptor (IL-1R) signaling through MyD88 is a central mediator of IL-12 secretion and the protective Th1 response to T. gondii (51). In mice that are intraperitoneally infected with T. gondii, the parasite protein profilin directly binds and activates TLR11, contributing to IL-12 production and parasite restriction (52). However, profilin is an actin-modifying protein sequestered within parasites and TLR11 signals from endosomal compartments, suggesting that this pathway may be mostly activated by phagocytosed, dead, or dysfunctional parasites (Figure 1). Consistent with this model, after oral infection, TLR11-deficient mice had minimal defects in their Th1 response compared with mice deficient in MyD88 or TLR2, TLR4, and TLR9; however, treating TLR11-deficient mice with antibiotics phenocopied MyD88-deficient mice (53, 54). These data indicate that gut commensal microbiota can prime a protective immune response to T. gondii independent of parasite recognition by TLR11. It is notable that human TLR11 is a pseudogene, indicating alternative innate sensing mechanisms to detect and destroy T. gondii in human cells.

Figure 1

Innate immune signaling and the influence of parasite effectors. T. gondii grows within a parasitophorous vacuole membrane (PVM) that protects the parasite from cytosolic immune sensors and avoids fusion with the endolysosomal compartments containing Toll-like receptors (TLRs). In the mouse, TLR11 recognizes Tg profilin, an actin-modifying protein that is exposed once dead or damaged parasites are phagocytosed. TLR11 is a pseudogene in humans. TgGRA15 can promote host NF-κB phosphorylation and nuclear translocation. In mice, NF-κB stimulation is necessary for transcriptional regulation of the inflammasome components NLRP1, NLRP3, and IL-1; however, human monocytes can engage an NLRP3 inflammasome independent of NF-κB prestimulation. A detailed mechanism of inflammasome activation, parasite killing, and host cell death remains elusive, particularly in regard to signal integration with IFN-γ. IFN-γ signaling induces STAT1 translocation to the nucleus and upregulation of IFN-responsive genes, including immunity-related GTPases (IRGs, mouse) and guanylate-binding proteins (GBPs, human and mouse), which functions to attack parasite vacuole, leading to parasite killing and host cell death. In human cells, GBP1 is necessary for this process, which leads to AIM2 activation of an alternative apoptosis pathway. The type I parasite rhoptry proteins, TgROP5, 17, and 18, can dismantle the function of the mouse IRGs IRGa6 and IRGb6 at the PVM, inactivating GBP attack and parasite killing. The parasite dense granule effector TgIST is a nuclear repressor of STAT1 transcription. TgROP16 is a kinase that phosphorylates and activates host STAT3 and STAT6. TgEGGR affects host gene expression through E2F3- and E2F4-mediated epigenetic modifications. In infected monocytes and dendritic cells (DCs), TgWIP and 14-3-3 proteins promote cell mobility, a putative mechanism of intracellular parasite dissemination in vivo.

The inflammasome links detection of microbial components or cell damage associated with infection to the release of IL-1 family cytokines and, often, inflammatory cell death. While the inflammasome response to protozoa is understudied in comparison with bacterial and viral pathogens, what is known about T. gondii recognition suggests major differences (55, 56). The inflammasome sensors NLRP1 (in mice) and NLRP3 (in mice and humans) have been shown to process and release IL-1β in response to T. gondii infection (Figure 1 and refs. 57–59). Mice deficient in NLRP3, caspase-1, and/or caspase-11 have a higher parasite burden in vivo (57, 58, 60). However, unlike better-studied inflammasome triggers (e.g., the NLRP1 protease anthrax lethal toxin or bacterial pathogens that activate NLRP3), pyroptotic host cell death is not observed in the mouse or human cells (57–59). Unlike murine macrophages, the NLRP3 inflammasome in human monocytes is activated independent of the TLR pathway via Syk and CARD9 signaling; and IL-1β release is independent of the pore-forming gasdermin D (59, 61–63). Open questions in the molecular mechanism of the inflammasome response to T. gondii include what parasite signals activate the inflammasome, why proptosis is not engaged, and whether this is the result of active parasite manipulation.

Recent data also suggest crosstalk between the inflammasome and the IFN-inducible GTPases, a pathway that surveys the cell for foreign or damaged membranes and targets them for clearance downstream of IFN-γ. In human cells the dynamin-superfamily guanylate-binding protein 1 (GBP1) localizes to the PVM; this triggers release of parasite DNA into the host cell cytosol, where it is detected by the inflammasome sensor AIM2 (Figure 1 and refs. 63, 64). For reasons that are still unclear, a pyroptotic inflammasome response is not engaged; instead, an alternative apoptotic pathway of host cell death is activated (63, 64). Unlike the human system, mice rely on an expanded family of p47 IRGs to detect the parasite vacuole downstream of IFN-γ. Mouse IRGM1 and IRGM3 regulate the interaction between IRGa6, IRGb6, and phospholipids at the PVM (65). A broader range of mouse GBPs have been implicated in T. gondii clearance; however, the mechanism of parasite killing and host cell death in mouse cells is not known (66–68). The importance of the IRG system in parasite clearance is underscored by the observation that type I parasites express a triad of secreted effectors, rhoptry protein 5 (ROP5), ROP17, and ROP18, which bind to and inactivate the GTPase function of IRGa6 and IRGb6 (69, 70). This inactivation is a major mechanism of type-specific virulence, as type II and type III parasites express alleles of Rop5 or Rop18, respectively, that cannot effectively subvert IRG attack. Although there has been tremendous progress toward identifying the classes of cell-autonomous immune signaling in response to T. gondii, the field lacks an integrated model of cell-autonomous sensing across these pathways for both mouse and human systems, particularly in cell types other than fibroblast, monocyte, and macrophage.

T.gondii recognition by innate immune sensors triggers a Th1-polarized, CD8⁺ T cell–dependent immune response that is necessary for host survival. There are many excellent reviews on the immunobiology of infection (7, 8, 71, 72), so we will touch briefly on aspects of the acute immune response that are necessary for the progression to chronic infection. Mice deficient in IL-12, TNF-α, and IFN-γ or their signaling pathways die of parasite overgrowth in acute infection (73–76). IFN-γ and IL-12 deficiency is rare in humans and has not been correlated with increased susceptibility to toxoplasmosis; however, monocyte-derived macrophages from IFNGR1-deficient patients fail to restrict T. gondii after IFN-γ stimulation compared with healthy-donor macrophages (77, 78). Mice deficient in the IL-6 pathway fail to mount a protective B cell response and die in early chronic infection (79). IL-10 and regulatory T cells play an equally important role in host survival by limiting the magnitude of the inflammatory response and bystander damage (80–85).

A growing number of T. gondii effectors have been identified that are secreted into the host cell to control immune signaling. These effectors are released from secretory organelles known as the rhoptries (ROP) and the dense granules (GRA), and many of the effectors are polymorphic across strains and play a role in virulence (Figure 1). GRA15 activates NF-κB, and GRA24 activates the p38 MAPK pathway to promote expression of IL-12 and IL-18, the upstream regulators of IFN-γ and T cell activation (86, 87). ROP16 is a serine-threonine kinase that directly phosphorylates STAT3 and STAT6 and dampens IL-12 production, which may be consistent with the concept that fine-tuning immune response is necessary for host survival and parasite transmission (88, 89). However, the effector TgIST was recently identified as an inhibitor of IFN receptor signaling. TgIST binds STAT1 and forms an inhibitory complex with the nucleosome remodeling deacetylase (Mi-2/NuRD) complex. This suppresses transcription of IRF1-dependent cytokines, MHC class II expression and antigen presentation, and inducible nitric oxide synthase (iNOS) expression, which kills parasites by producing reactive nitrogen species (90, 91). Similarly, TgTEEGR interacts with E2F3 and E2F4 transcription factors, and forms a nuclear complex with a catalytic subunit of polycomb repressor complex to block NF-κB–mediated expression of proinflammatory cytokines like IL-1β and IL-6 (92). These effectors are among 200–300 predicted secreted effector proteins in the parasite genome, the majority of which have not been characterized, particularly in the context of chronic infection.

In chronic infection, the central nervous system contains the highest frequency of parasites per gram of tissue. The potential implications of neural infection for host behavior and homeostasis have led to great interest in understanding the biology of T. gondii infection in the brain. Our understanding of chronic central nervous system infection is almost exclusively based on murine models of infection. There are many open questions, beginning with how the parasite traverses the blood-brain barrier (BBB). Using intravital microscopy, T. gondii has been imaged replicating within brain endothelial cells and then directly entering the brain (93). Mice infected intravenously with the T. gondii RH strain had a higher brain parasite load than mice infected with the CPS strain, which cannot replicate in vivo, suggesting that T. gondii growth within vascular endothelial cells may be an important stopover before direct entry into the brain (Figure 2A and ref. 93). Perfusion of Evans blue dye shows increased BBB permeability during chronic T. gondii infection, accompanied by reduced blood flow and capillary rarefication which may permit immune cell entry into the brain (94). Using intravital microscopy, CCR2⁺ monocytes are found to accumulate, exhibiting rolling and cradling behavior at the BBB (95). This observation, coupled with the high frequency of infection of dendritic cells and their hypermotility phenotype, has led to the Trojan horse hypothesis: that parasites traverse the BBB within immune cells (Figure 2A and refs. 49, 96). Although direct evidence for this model is lacking, antibody depletion of CD11b⁺ leukocytes correlated with reduced brain parasite load; and adoptive transfer of T. gondii–infected CD11c⁺ or CD11b⁺ cells into naive mice led to neural infection (97).

Figure 2

T. gondii entry and control of persistent infection in the brain. (A) In acute infection, T. gondii is frequently observed in immune cells, including monocytes and dendritic cells, with hypermigratory behavior. During infection, blood-brain barrier (BBB) permeability increases and monocytes accumulate in the endothelial lumen, interacting with endothelial cells. These observations have led to the hypothesis that migratory immune cells deliver T. gondii to the BBB and, perhaps, smuggle them into the brain. Replicating parasites are also observed in brain endothelial cells, whose subsequent lysis may be a mechanism of T. gondii entry into the brain. (B) During acute infection parasites are observed infecting neurons, astrocytes, microglia, and infiltrating immune cells. Astrocytes and microglia as well as peripheral monocytes can clear parasites with cell-autonomous immune pathways. (C) As chronic infection progresses, infected astrocytes and microglia or the parasites within them are cleared and cysts are primarily observed within neurons. Most parasite cysts are not associated with immune infiltrate; however, individual parasites or parasite debris can be observed colocalizing with immune infiltrate.

Analysis of mouse brain sections and an extremely limited number of healthy human brain samples indicates that most intracellular cysts are not associated with immune infiltration (98). However, within the same brain section, inflammatory foci can be observed containing parasites or parasite debris, activated microglia, macrophages, and T cells (99). Depleting IFN-γ or CD4⁺ and CD8⁺ T cells leads to parasite recrudescence (100). Taken together these data suggest that intracellular cysts are relatively immunologically silent; however, cysts that lyse (spontaneously or through recrudescence) are recognized and quickly contained by infiltrating immune cells (Figure 2C).

Experiments using parasites engineered to secrete Cre recombinase in Cre reporter mice have demonstrated that neurons are the major cell type interacting with the parasite in the brain, although T cells, monocytes or macrophages, microglia, and astrocytes are reporter-positive early in brain infection (Figure 2B and refs. 101–103). These data also suggest that rather than having a tropism for neurons, T. gondii is cleared from non-neuronal cell types in the brain. Consistent with this model, disabling the IFN-γ signaling in mouse astrocytes by knocking out the transcription factor STAT1 led to greater incidence of cysts within astrocytes (104); and IFN-γ depletion increased the percentage of infected astrocytes (102). Subsequently, the ability of mouse astrocytes to restrict T. gondii growth in response to IFN-γ was shown to depend on IRGM3 (IGTP), not iNOS; IRGM3 and IRGa6 disrupted the PVM and, in one study, led to parasite egress (105–107). In human astrocytes, IL-1β in combination with IFN-γ induced iNOS-dependent killing of T. gondii (108), whereas TNF- and IFN-γ limited T. gondii growth by tryptophan starvation via upregulated indoleamine 2,3-dioxygenase (IDO) (109). The parasite effector TgGRA15 has been shown to limit IDO-mediated parasite restriction in cultured glioblastoma and neuroblastoma cell lines (110). IFN-γ in combination with TNF-α or LPS has also been shown to activate parasite killing functions of human and murine microglia through iNOS-dependent and -independent mechanisms (111–113). Microglia can also produce IFN-γ and TNF-α, which are critical for central nervous system restriction of the infection (114); IFN-γ, in particular, has been shown to induce adhesion molecule expression on vascular endothelial cells and promote the expression of CXCL9, CXCL10, and CCL5, which recruit peripheral immune cells to the brain (115, 116). Most of these data are from in vitro experiments, and better tools to study microglia and astrocyte function in vivo will be important to clarify which pathways control central nervous system infection.

While brain-resident immune cells contribute to T. gondii restriction, the role of cell-autonomous immunity in neurons is less clear. A recent study using OVA-expressing parasites and conditional MHC class I–deficient mice demonstrated that neurons can present T. gondii–derived antigens to initiate a CD8⁺ T cell response (117); however, whether endogenous parasite epitopes are efficiently presented on neurons is yet to be examined. Brain-infiltrating CD8⁺ T cells have been shown to control cyst burden indirectly through IFN-γ secretion, and, to a lesser extent, via perforin-dependent killing of infected cells (118–120). It is notable that perforin has been shown to trigger parasite egress in vitro, suggesting that perforin may limit parasite growth but other cells are responsible for parasite killing, potentially through cell-autonomous immunity (121). A minimal reliance on perforin-mediated T. gondii clearance also fits a model wherein cellular cytotoxicity should be limited in the brain to promote survival of neurons, which have an extremely limited regenerative capacity.

Currently there are no therapeutic tools that effectively target bradyzoite cysts and sterilize chronic infection. Our understanding of bradyzoite biology is weaker than our understanding of tachyzoite biology. This is linked to long-standing technical challenges associated with genetic manipulation of bradyzoite-specific genes that are required to perform “necessary and sufficient” experiments. However, the recent bloom in CRISPR/Cas9 tools has led to gains in this arena (122, 123).

The transition between tachyzoite and bradyzoite has commonly been referred to as “switching”; however, recent studies suggest that stage conversion is a continuum under epigenetic and transcriptional regulation rather than a finite life stage. Bradyzoite polarization can be induced by cell stressors including alkaline media, heat shock, and oxidative stress (refs. 124–126 and Figure 3). IFN-γ treatment has been shown to induce bradyzoite gene expression in infected macrophages but not fibroblasts, suggesting that cell type–specific differentiation signals may also exist (127). Compared with fibroblasts, infected neuronal or skeletal muscle cells support a stronger expression of bradyzoite markers and a higher frequency of cyst development (128). It is worth noting that neurons and muscle cell types are historically difficult to culture, suggesting that cell stress signals may be relevant to bradyzoite development in these models. However, terminally differentiated myotubes are reported to support a higher frequency of bradyzoites compared with dividing myoblast progenitor cells, suggesting that cell cycle may provide developmental cues for the parasite development as well (129).

Figure 3

Environmental and host cell–specific pressures driving the T. gondiitachyzoite to bradyzoite transition. Left: T. gondii tachyzoites can invade almost any nucleated host cell type and grow within the PVM formed from host plasma membrane. In vitro, a range of tissue culture stress conditions can upregulate bradyzoite-specific genes. As parasites polarize to a bradyzoite transcriptional profile, they synthesize a heavily glycosylated cyst wall beneath the PVM. The frequency and rate of bradyzoite differentiation are also influenced by the host cell type, cell cycle status, the host cell lifespan, and inflammatory signals in vitro. In vivo, cysts are most frequently observed in neurons, cardiac muscle, skeletal muscle, and retinal pigment epithelial cells. If the host is immune-suppressed, parasites shift toward a replicative tachyzoite form in a process referred to as recrudescence, which is associated with tissue damage, particularly in the eye.

Although the precise signals are unclear, histone methylation and acetylation are important epigenetic regulators of bradyzoite differentiation. Treating tachyzoites with arginine methyltransferase inhibitor, AMI-I, induces a reduction of histone H3R17 methylation and bradyzoite differentiation in vitro (130). The T. gondii histone acetyltransferase TgGCN5a is enriched at promoter regions of bradyzoite-specific genes, and TgGCN5a-deficient parasites fail to upregulate the bradyzoite markers Bag1 and Ldh2 under stress (131). Treating infected cells with the histone deacetylase inhibitor FR235222 induces bradyzoite differentiation through inhibiting TgHDAC3 (132). Phosphorylation of the T. gondii eukaryotic initiation factor 2 α subunit (TgeIF2α) is enhanced under stress conditions and is necessary for bradyzoite differentiation (133). Guanabenz, an eIF2α dephosphorylation inhibitor, has been shown to impair tachyzoite proliferation and promote bradyzoite differentiation in vitro (134).

The ApiAP2 family of transcription factors are emerging as central regulators of bradyzoite differentiation. This family consists of 67 genes, many of which are associated with bradyzoite stage–specific expression. Specifically, AP2XI-4– and AP2IV-3–knockout parasites have reduced expression of bradyzoite-specific genes after in vitro switch; and AP2XI-4–null T. gondii forms fewer cysts in mice (135, 136). AP2IV-4 knockouts express some bradyzoite-specific genes under tachyzoite culture, but had fewer brain cysts in mice (137). Using a CRISPR/Cas9 guide RNA library targeting mostly AP2 domain–containing proteins and predicted nucleic acid–binding proteins, bradyzoite formation deficient 1 (BFD1), a Myb-like transcription factor, was recently identified as a key regulator of bradyzoite differentiation in vitro and in vivo in mice. Interestingly, Bfd1 mRNA is expressed in tachyzoites; however, protein expression is only induced by stress conditions (138). It remains to be seen whether immunosuppression induces any parasite recrudescence in mice infected with BFD1-deficient parasites and how BFD1- and AP2-family proteins coordinate bradyzoite differentiation.

T.gondii bradyzoite cysts are often defined by formation of a cyst wall consisting of heavily glycosylated proteins underneath the PVM (139). The cyst wall is essential for transmission, protecting the parasite from gastric proteases and the low pH of the stomach. Parasites deficient in the cyst wall–localized bradyzoite pseudokinase 1 (BPK1) were more sensitive to pepsin digestion and less orally infectious than WT parasites (140). Parasites that were rendered genetically deficient in cyst glycoproteins, including loss of the nucleotide-sugar transporter TgNST1 or the heavily glycosylated cyst wall protein TgCST1, have defects in cyst number, cyst stability, and infectivity during oral infection (141–143). The cyst wall may also protect bradyzoites from enzymatic attack during chronic infection. The Wilson laboratory demonstrated that chitinase-expressing, alternatively activated (M2) macrophages were able to recognize and degrade chitin-like polysaccharides in the cyst wall (144). Consistent with this observation, a GWAS identified single-nucleotide polymorphisms in the intergenic region of the human CHIA locus, which expresses chitinase, that were significantly associated with T. gondii infection (145).

T.gondii infection is the most frequent cause of posterior uveitis, also referred to as chorioretinitis or inflammation of the retina and choroid (pigmented vascular coat of the eye) (30). This is one area of T. gondii infection that has been more extensively studied in patients than in animal models, which have been limited until recently. Type II strains, most frequently associated with infection in Europe and North America, are associated with chorioretinitis (146, 147). Historically, ocular toxoplasmosis was associated with congenital infection; however, rates of disease associated with postnatal infection are rising and associated with new T. gondii strains (148). Over 70% of patients presenting with acute ocular toxoplasmosis already have ocular scars, suggesting that disease progression is driven by the inflammatory response to recrudescent T. gondii leading to the accumulation of tissue damage over time (149). Immune-competent individuals are able to control ocular infection, but early antiparasitic treatment is critical to limit the extent of retinal damage (150). Human retinal vascular endothelial cells are more sensitive to infection than other endothelial cell types, suggesting a potential mechanism of entry into the eye (151). T. gondii cysts have been observed in retinal pigmented epithelial cells (152). In a mouse model of ocular toxoplasmosis, retinal pigment epithelial cells and infiltrating immune cells expressed the T cell inhibitory ligand PD-L1 (153). This may be an important mechanism to limit tissue pathology, although the parasites may exploit this axis for persistence. IFN-γ and IL-6, which are both critical in restricting systematic parasitemia (79, 100), were elevated in the vitreous humor of mice with ocular lesions. However, intraocular injection of an IFN-γ–blocking antibody impaired parasite control and worsened tissue damage, while, perhaps counterintuitively, injection of an IL-6–blocking antibody improved parasite control and minimized ocular damage (154–156). Patients infected with virulent South American haplotypes of T. gondii, which have been associated with aggressive chorioretinitis, had less IFN-γ and IL-17 but higher IL-13 and IL-6 levels in the eye compared with European patients infected with virulent type I (157). However, it is currently not clear whether these differences in immune regulation control ocular disease severity.

In mice and rats, infection with T. gondii leads to a well-established loss of innate aversion behavior to felines, which has been proposed to benefit the parasite by facilitating transmission via predation (11, 158, 159). Whether these behavioral phenotypes are driven by specific changes in neural activity or a more general effect of inflammation is an open question. The observation that T. gondii expresses two aromatic amino acid hydrolases that produce l-DOPA, AAH1, and AAH2 led to the hypothesis that the parasite could modulate dopaminergic neuron function. However, deletion of AAH2 failed to alter brain dopamine levels, neuroinflammation, or behavioral alterations in T. gondii–infected mice (160, 161), although these genes are necessary for oocyst development in the cat (162). Notably, mice infected with an avirulent mutant of type I T. gondii or the related organism Neospora caninum, which are cleared before establishing chronic infection, exhibit loss of aversion behavior even though chronic infection is not sustained (163). These data suggest that acute inflammation may be sufficient to trigger sustained behavioral changes, although the molecular bases for behavioral changes in T. gondii infection are unclear. Recently, the olfactory GPCR trace amine-associated receptor 4 (TAAR4) was shown to recognize 2-phenylethylamine, a metabolite enriched in urine of predators, including feline species. There is no homolog of TAAR4 in humans, but mice deficient in TAAR4 do not engage in avoidance behavior to bobcat and mountain lion urine (164). That the olfactory neurons expressing TAAR4 are altered or damaged during T. gondii infection is a compelling hypothesis that remains to be tested.

Sustained interaction with the immune system is a hallmark of T. gondii infection: throughout chronic infection humans and mice have high titers of T. gondii–specific IgG and sera cytokines. There is growing evidence that T. gondii infection is associated with cachexia in mice, an immune-metabolic disease of sustained muscle wasting. Cachexia positively correlates with parasite load and inflammation severity; however, hypermetabolic weight loss cannot be rescued by diet supplementation (13, 165–167). In oral infection, intestinal barrier inflammation resolves during chronic infection, but commensal dysbiosis does not (14, 168); however, dysbiosis is not sufficient for cachexia, as uninfected cage mates experienced a similar microbial shift but did not develop cachexia (14). Chronically infected mice have sustained changes in splenic and lymph node architecture and are more susceptible to acute viral challenge (169). Moreover, cachectic mice were more susceptible to LPS challenge than mice that recovered weight (170). Recently, mice deficient in the IL-1R axis were shown to recover from acute cachectic weight loss, although chronic parasite burden was similar to that in wild-type mice (171). A study from the Wohlfert laboratory showed that infection-induced myositis could be reversed by depletion of regulatory T cells, which were enriched in skeletal muscle (172). Parasite biology that promotes behavior modification and cachexia in rodent hosts may provide a selective advantage to T. gondii by increasing the likelihood of predation and transmission to feline hosts. It is important to note that there is currently no evidence of cachexia in immune-competent humans with chronic T. gondii infection. However, cachexia is a predictor of mortality in almost every chronic human disease with limited experimental tools to probe sustained disease. The interaction between T. gondii and mice is proving an informative model to understand the pathophysiology of cachexia, which can be applied to understand other disease settings.

T.gondii’s ability to establish a persistent chronic infection is essential for parasite transmission. However, there is much to learn about this stage of infection in animal and human hosts. Deep sequencing has unraveled a far greater diversity in T. gondii gene assortment than originally thought, which has opened the door to understanding how parasite genetics influences pathology associated with chronic infection. CRISPR/Cas9 tools are expanding our ability to manipulate the T. gondii genome to understand how gene expression in bradyzoites controls differentiation, cyst stability, and oral infectivity of the parasite. Bradyzoite biology is intimately linked to the immune response during chronic infection. The coming decades will likely reveal mechanisms of cell-autonomous immunity to chronic T. gondii infection in the brain and other chronically infected tissues, as well as reveal the costs of the chronic inflammatory response for host homeostasis. A better understanding of this biology is needed to develop therapeutic strategies that effectively target bradyzoite cysts. Given the long evolutionary relationship between mammalian hosts and T. gondii, such studies are likely to discover important information about the regulation of immune functions during chronic inflammation more broadly.

Conflict of interest: The authors have declared that no conflict of interest exists.

Copyright: © 2020, American Society for Clinical Investigation.

Reference information: J Clin Invest. 2020;130(7):3370–3380.https://doi.org/10.1172/JCI136226.

1. Dubey JP, Miller NL, Frenkel JK. The Toxoplasma gondii oocyst from cat feces. J Exp Med. 1970;132(4):636–662.
   View this article via: PubMed CrossRef Google Scholar
2. Black MW, Boothroyd JC. Lytic cycle of Toxoplasma gondii. Microbiol Mol Biol Rev. 2000;64(3):607–623.
   View this article via: PubMed CrossRef Google Scholar
3. Jones JL, Dargelas V, Roberts J, Press C, Remington JS, Montoya JG. Risk factors for Toxoplasma gondii infection in the United States. Clin Infect Dis. 2009;49(6):878–884.
   View this article via: PubMed CrossRef Google Scholar
4. Dubey JP, Speer CA, Shen SK, Kwok OC, Blixt JA. Oocyst-induced murine toxoplasmosis: life cycle, pathogenicity, and stage conversion in mice fed Toxoplasma gondii oocysts. J Parasitol. 1997;83(5):870–882.
   View this article via: PubMed CrossRef Google Scholar
5. Speer CA, Dubey JP. Ultrastructure of early stages of infections in mice fed Toxoplasma gondii oocysts. Parasitology. 1998;116(pt 1):35–42.
   View this article via: PubMed Google Scholar
6. Martorelli Di Genova B, Wilson SK, Dubey JP, Knoll LJ. Intestinal delta-6-desaturase activity determines host range for Toxoplasma sexual reproduction. PLoS Biol. 2019;17(8):e3000364.
   View this article via: PubMed CrossRef Google Scholar
7. Yarovinsky F. Innate immunity to Toxoplasma gondii infection. Nat Rev Immunol. 2014;14(2):109–121.
   View this article via: PubMed CrossRef Google Scholar
8. Sasai M, Pradipta A, Yamamoto M. Host immune responses to Toxoplasma gondii. Int Immunol. 2018;30(3):113–119.
   View this article via: PubMed CrossRef Google Scholar
9. Coombes JL, Hunter CA. Immunity to Toxoplasma gondii—into the 21st century. Parasite Immunol. 2015;37(3):105–107.
   View this article via: PubMed CrossRef Google Scholar
10. Dewan A, Pacifico R, Zhan R, Rinberg D, Bozza T. Non-redundant coding of aversive odours in the main olfactory pathway. Nature. 2013;497(7450):486–489.
    View this article via: PubMed CrossRef Google Scholar
11. Boillat M, et al. Neuroinflammation-associated aspecific manipulation of mouse predator fear by Toxoplasma gondii. Cell Rep. 2020;30(2):320–334.e6.
    View this article via: PubMed CrossRef Google Scholar
12. Jin RM, Blair SJ, Warunek J, Heffner RR, Blader IJ, Wohlfert EA. Regulatory T cells promote myositis and muscle damage in Toxoplasma gondii infection. J Immunol. 2017;198(1):352–362.
    View this article via: PubMed CrossRef Google Scholar
13. Arsenijevic D, Girardier L, Seydoux J, Chang HR, Dulloo AG. Altered energy balance and cytokine gene expression in a murine model of chronic infection with Toxoplasma gondii. Am J Physiol. 1997;272(5 pt 1):E908–E917.
    View this article via: PubMed Google Scholar
14. Hatter JA, et al. Toxoplasma gondii infection triggers chronic cachexia and sustained commensal dysbiosis in mice. PLoS One. 2018;13(10):e0204895.
    View this article via: PubMed CrossRef Google Scholar
15. Cañón-Franco WA, López-Orozco N, Gómez-Marín JE, Dubey JP. An overview of seventy years of research (1944-2014) on toxoplasmosis in Colombia, South America. Parasit Vectors. 2014;7:427.
    View this article via: PubMed Google Scholar
16. Guigue N, et al. Continuous decline of Toxoplasma gondii seroprevalence in hospital: a 1997-2014 longitudinal study in Paris, France. Front Microbiol. 2018;9:2369.
    View this article via: PubMed Google Scholar
17. Dubey JP, Lago EG, Gennari SM, Su C, Jones JL. Toxoplasmosis in humans and animals in Brazil: high prevalence, high burden of disease, and epidemiology. Parasitology. 2012;139(11):1375–1424.
    View this article via: PubMed CrossRef Google Scholar
18. 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(2):267–299.
    View this article via: PubMed CrossRef Google Scholar
19. Wong SY, Remington JS. Biology of Toxoplasma gondii. AIDS. 1993;7(3):299–316.
    View this article via: PubMed CrossRef Google Scholar
20. Assi MA, Rosenblatt JE, Marshall WF. Donor-transmitted toxoplasmosis in liver transplant recipients: a case report and literature review. Transpl Infect Dis. 2007;9(2):132–136.
    View this article via: PubMed CrossRef Google Scholar
21. Galván-Ramírez ML, Sánchez-Orozco LV, Gutiérrez-Maldonado AF, Rodriguez Pérez LR. Does Toxoplasma gondii infection impact liver transplantation outcomes? A systematic review. J Med Microbiol. 2018;67(4):499–506.
    View this article via: PubMed CrossRef Google Scholar
22. Renoult E, et al. Toxoplasmosis in kidney transplant recipients: report of six cases and review. Clin Infect Dis. 1997;24(4):625–634.
    View this article via: PubMed CrossRef Google Scholar
23. Montoya JG, et al. Infectious complications among 620 consecutive heart transplant patients at Stanford University Medical Center. Clin Infect Dis. 2001;33(5):629–640.
    View this article via: PubMed CrossRef Google Scholar
24. Wreghitt TG, et al. Toxoplasmosis in heart and heart and lung transplant recipients. J Clin Pathol. 1989;42(2):194–199.
    View this article via: PubMed CrossRef Google Scholar
25. Sturge CR, Yarovinsky F. Complex immune cell interplay in the gamma interferon response during Toxoplasma gondii infection. Infect Immun. 2014;82(8):3090–3097.
    View this article via: PubMed CrossRef Google Scholar
26. Montoya JG, Liesenfeld O. Toxoplasmosis. Lancet. 2004;363(9425):1965–1976.
    View this article via: PubMed CrossRef Google Scholar
27. Luft BJ, Remington JS. Toxoplasmic encephalitis in AIDS. Clin Infect Dis. 1992;15(2):211–222.
    View this article via: PubMed CrossRef Google Scholar
28. Dunay IR, Gajurel K, Dhakal R, Liesenfeld O, Montoya JG. Treatment of Toxoplasmosis: historical perspective, animal models, and current clinical practice. Clin Microbiol Rev. 2018;31(4):e00057-17.
    View this article via: PubMed Google Scholar
29. Khan A, et al. Genetic divergence of Toxoplasma gondii strains associated with ocular toxoplasmosis, Brazil. Emerging Infect Dis. 2006;12(6):942–949.
    View this article via: PubMed CrossRef Google Scholar
30. Maenz M, Schlüter D, Liesenfeld O, Schares G, Gross U, Pleyer U. Ocular toxoplasmosis past, present and new aspects of an old disease. Prog Retin Eye Res. 2014;39:77–106.
    View this article via: PubMed CrossRef Google Scholar
31. Sibley LD, Boothroyd JC. Virulent strains of Toxoplasma gondii comprise a single clonal lineage. Nature. 1992;359(6390):82–85.
    View this article via: PubMed CrossRef Google Scholar
32. Su C, Howe DK, Dubey JP, Ajioka JW, Sibley LD. Identification of quantitative trait loci controlling acute virulence in Toxoplasma gondii. Proc Natl Acad Sci U S A. 2002;99(16):10753–10758.
    View this article via: PubMed CrossRef Google Scholar
33. Khan A, Dubey JP, Su C, Ajioka JW, Rosenthal BM, Sibley LD. Genetic analyses of atypical Toxoplasma gondii strains reveal a fourth clonal lineage in North America. Int J Parasitol. 2011;41(6):645–655.
    View this article via: PubMed CrossRef Google Scholar
34. Galal L, Hamidović A, Dardé ML, Mercier M. Diversity of Toxoplasma gondii strains at the global level and its determinants. Food Waterborne Parasitol. 2019;15:e00052.
    View this article via: PubMed Google Scholar
35. Su C, et al. Globally diverse Toxoplasma gondii isolates comprise six major clades originating from a small number of distinct ancestral lineages. Proc Natl Acad Sci U S A. 2012;109(15):5844–5849.
    View this article via: PubMed CrossRef Google Scholar
36. Minot S, et al. Admixture and recombination among Toxoplasma gondii lineages explain global genome diversity. Proc Natl Acad Sci U S A. 2012;109(33):13458–13463.
    View this article via: PubMed CrossRef Google Scholar
37. Shwab EK, et al. Geographical patterns of Toxoplasma gondii genetic diversity revealed by multilocus PCR-RFLP genotyping. Parasitology. 2014;141(4):453–461.
    View this article via: PubMed CrossRef Google Scholar
38. Lorenzi H, et al. Local admixture of amplified and diversified secreted pathogenesis determinants shapes mosaic Toxoplasma gondii genomes. Nat Commun. 2016;7:10147.
    View this article via: PubMed Google Scholar
39. Müller UB, Howard JC. The impact of Toxoplasma gondii on the mammalian genome. Curr Opin Microbiol. 2016;32:19–25.
    View this article via: PubMed CrossRef Google Scholar
40. Hassan MA, Olijnik AA, Frickel EM, Saeij JP. Clonal and atypical Toxoplasma strain differences in virulence vary with mouse sub-species. Int J Parasitol. 2019;49(1):63–70.
    View this article via: PubMed CrossRef Google Scholar
41. Mordue DG, Håkansson S, Niesman I, Sibley LD. Toxoplasma gondii resides in a vacuole that avoids fusion with host cell endocytic and exocytic vesicular trafficking pathways. Exp Parasitol. 1999;92(2):87–99.
    View this article via: PubMed CrossRef Google Scholar
42. Molloy MJ, et al. Intraluminal containment of commensal outgrowth in the gut during infection-induced dysbiosis. Cell Host Microbe. 2013;14(3):318–328.
    View this article via: PubMed CrossRef Google Scholar
43. Gregg B, et al. Replication and distribution of Toxoplasma gondii in the small intestine after oral infection with tissue cysts. Infect Immun. 2013;81(5):1635–1643.
    View this article via: PubMed CrossRef Google Scholar
44. Coombes JL, et al. Motile invaded neutrophils in the small intestine of Toxoplasma gondii-infected mice reveal a potential mechanism for parasite spread. Proc Natl Acad Sci U S A. 2013;110(21):E1913–E1922.
    View this article via: PubMed CrossRef Google Scholar
45. Courret N, Darche S, Sonigo P, Milon G, Buzoni-Gâtel D, Tardieux I. CD11c- and CD11b-expressing mouse leukocytes transport single Toxoplasma gondii tachyzoites to the brain. Blood. 2006;107(1):309–316.
    View this article via: PubMed CrossRef Google Scholar
46. Safronova A, et al. Alarmin S100A11 initiates a chemokine response to the human pathogen Toxoplasma gondii. Nat Immunol. 2019;20(1):64–72.
    View this article via: PubMed CrossRef Google Scholar
47. Weidner JM, et al. Migratory activation of parasitized dendritic cells by the protozoan Toxoplasma gondii 14-3-3 protein. Cell Microbiol. 2016;18(11):1537–1550.
    View this article via: PubMed CrossRef Google Scholar
48. Sangaré LO, et al. In vivo CRISPR screen identifies TgWIP as a Toxoplasma modulator of dendritic cell migration. Cell Host Microbe. 2019;26(4):478–492.e8.
    View this article via: PubMed CrossRef Google Scholar
49. Lambert H, Hitziger N, Dellacasa I, Svensson M, Barragan A. Induction of dendritic cell migration upon Toxoplasma gondii infection potentiates parasite dissemination. Cell Microbiol. 2006;8(10):1611–1623.
    View this article via: PubMed CrossRef Google Scholar
50. Ólafsson EB, Ross EC, Varas-Godoy M, Barragan A. TIMP-1 promotes hypermigration of Toxoplasma-infected primary dendritic cells via CD63-ITGB1-FAK signaling. J Cell Sci. 2019;132(3):jcs225193.
    View this article via: PubMed CrossRef Google Scholar
51. Scanga CA, et al. Cutting edge: MyD88 is required for resistance to Toxoplasma gondii infection and regulates parasite-induced IL-12 production by dendritic cells. J Immunol. 2002;168(12):5997–6001.
    View this article via: PubMed CrossRef Google Scholar
52. Yarovinsky F, et al. TLR11 activation of dendritic cells by a protozoan profilin-like protein. Science. 2005;308(5728):1626–1629.
    View this article via: PubMed CrossRef Google Scholar
53. Denkers EY. A gut feeling for microbes: getting it going between a parasite and its host. Cell Host Microbe. 2009;6(2):104–106.
    View this article via: PubMed CrossRef Google Scholar
54. Benson A, Pifer R, Behrendt CL, Hooper LV, Yarovinsky F. Gut commensal bacteria direct a protective immune response against Toxoplasma gondii. Cell Host Microbe. 2009;6(2):187–196.
    View this article via: PubMed CrossRef Google Scholar
55. Broz P, Dixit VM. Inflammasomes: mechanism of assembly, regulation and signalling. Nat Rev Immunol. 2016;16(7):407–420.
    View this article via: PubMed CrossRef Google Scholar
56. Swanson KV, Deng M, Ting JP. The NLRP3 inflammasome: molecular activation and regulation to therapeutics. Nat Rev Immunol. 2019;19(8):477–489.
    View this article via: PubMed CrossRef Google Scholar
57. Ewald SE, Chavarria-Smith J, Boothroyd JC. NLRP1 is an inflammasome sensor for Toxoplasma gondii. Infect Immun. 2014;82(1):460–468.
    View this article via: PubMed CrossRef Google Scholar
58. Gorfu G, et al. Dual role for inflammasome sensors NLRP1 and NLRP3 in murine resistance to Toxoplasma gondii. mBio. 2014;5(1):e01117-13.
    View this article via: PubMed Google Scholar
59. Gov L, Schneider CA, Lima TS, Pandori W, Lodoen MB. NLRP3 and potassium efflux drive rapid IL-1β release from primary human monocytes during Toxoplasma gondii infection. J Immunol. 2017;199(8):2855–2864.
    View this article via: PubMed CrossRef Google Scholar
60. Coutermarsh-Ott SL, Doran JT, Campbell C, Williams TM, Lindsay DS, Allen IC. Caspase-11 modulates inflammation and attenuates Toxoplasma gondii pathogenesis. Mediators Inflamm. 2016;2016:9848263.
    View this article via: PubMed Google Scholar
61. Pandori WJ, Lima TS, Mallya S, Kao TH, Gov L, Lodoen MB. Toxoplasma gondii activates a Syk-CARD9-NF-κB signaling axis and gasdermin D-independent release of IL-1β during infection of primary human monocytes. PLoS Pathog. 2019;15(8):e1007923.
    View this article via: PubMed CrossRef Google Scholar
62. Gov L, Karimzadeh A, Ueno N, Lodoen MB. Human innate immunity to Toxoplasma gondii is mediated by host caspase-1 and ASC and parasite GRA15. mBio. 2013;4(4):e00255-13.
    View this article via: PubMed Google Scholar
63. Fisch D, et al. Human GBP1 is a microbe-specific gatekeeper of macrophage apoptosis and pyroptosis. EMBO J. 2019;38(13):e100926.
    View this article via: PubMed Google Scholar
64. Fisch D, et al. Human GBP1 differentially targets Salmonella and Toxoplasma to license recognition of microbial ligands and caspase-mediated death [pulished online Apr 23, 2020. bioRxiv. https://doi.org/10.1101/792804.
65. Lee Y, et al. Initial phospholipid-dependent Irgb6 targeting to Toxoplasma gondii vacuoles mediates host defense. Life Sci Alliance. 2020;3(1):e201900549.
    View this article via: PubMed CrossRef Google Scholar
66. Yamamoto M, et al. A cluster of interferon-γ-inducible p65 GTPases plays a critical role in host defense against Toxoplasma gondii. Immunity. 2012;37(2):302–313.
    View this article via: PubMed CrossRef Google Scholar
67. Collazo CM, et al. Inactivation of LRG-47 and IRG-47 reveals a family of interferon gamma-inducible genes with essential, pathogen-specific roles in resistance to infection. J Exp Med. 2001;194(2):181–188.
    View this article via: PubMed CrossRef Google Scholar
68. Saeij JP, Frickel EM. Exposing Toxoplasma gondii hiding inside the vacuole: a role for GBPs, autophagy and host cell death. Curr Opin Microbiol. 2017;40:72–80.
    View this article via: PubMed CrossRef Google Scholar
69. Fleckenstein MC, Reese ML, Könen-Waisman S, Boothroyd JC, Howard JC, Steinfeldt T. A Toxoplasma gondii pseudokinase inhibits host IRG resistance proteins. PLoS Biol. 2012;10(7):e1001358.
    View this article via: PubMed CrossRef Google Scholar
70. Etheridge RD, Alaganan A, Tang K, Lou HJ, Turk BE, Sibley LD. The Toxoplasma pseudokinase ROP5 forms complexes with ROP18 and ROP17 kinases that synergize to control acute virulence in mice. Cell Host Microbe. 2014;15(5):537–550.
    View this article via: PubMed CrossRef Google Scholar
71. Lima TS, Lodoen MB. Mechanisms of human innate immune evasion by Toxoplasma gondii. Front Cell Infect Microbiol. 2019;9:103.
    View this article via: PubMed Google Scholar
72. Tsitsiklis A, Bangs DJ, Robey EA. CD8⁺ T cell responses to Toxoplasma gondii: lessons from a successful parasite. Trends Parasitol. 2019;35(11):887–898.
    View this article via: PubMed CrossRef Google Scholar
73. Yap G, Pesin M, Sher A. Cutting edge: IL-12 is required for the maintenance of IFN-gamma production in T cells mediating chronic resistance to the intracellular pathogen, Toxoplasma gondii. J Immunol. 2000;165(2):628–631.
    View this article via: PubMed CrossRef Google Scholar
74. Suzuki Y, Orellana MA, Schreiber RD, Remington JS. Interferon-gamma: the major mediator of resistance against Toxoplasma gondii. Science. 1988;240(4851):516–518.
    View this article via: PubMed CrossRef Google Scholar
75. Deckert-Schlüter M, et al. Interferon-gamma receptor-deficiency renders mice highly susceptible to toxoplasmosis by decreased macrophage activation. Lab Invest. 1996;75(6):827–841.
    View this article via: PubMed Google Scholar
76. Schlüter D, et al. Both lymphotoxin-alpha and TNF are crucial for control of Toxoplasma gondii in the central nervous system. J Immunol. 2003;170(12):6172–6182.
    View this article via: PubMed CrossRef Google Scholar
77. Bustamante J, Boisson-Dupuis S, Abel L, Casanova JL. Mendelian susceptibility to mycobacterial disease: genetic, immunological, and clinical features of inborn errors of IFN-γ immunity. Semin Immunol. 2014;26(6):454–470.
    View this article via: PubMed CrossRef Google Scholar
78. Janssen R, et al. Divergent role for TNF-alpha in IFN-gamma-induced killing of Toxoplasma gondii and Salmonella typhimurium contributes to selective susceptibility of patients with partial IFN-gamma receptor 1 deficiency. J Immunol. 2002;169(7):3900–3907.
    View this article via: PubMed CrossRef Google Scholar
79. Jebbari H, Roberts CW, Ferguson DJ, Bluethmann H, Alexander J. A protective role for IL-6 during early infection with Toxoplasma gondii. Parasite Immunol. 1998;20(5):231–239.
    View this article via: PubMed CrossRef Google Scholar
80. Wilson EH, Wille-Reece U, Dzierszinski F, Hunter CA. A critical role for IL-10 in limiting inflammation during toxoplasmic encephalitis. J Neuroimmunol. 2005;165(1-2):63–74.
    View this article via: PubMed CrossRef Google Scholar
81. Neyer LE, Grunig G, Fort M, Remington JS, Rennick D, Hunter CA. Role of interleukin-10 in regulation of T-cell-dependent and T-cell-independent mechanisms of resistance to Toxoplasma gondii. Infect Immun. 1997;65(5):1675–1682.
    View this article via: PubMed CrossRef Google Scholar
82. Gazzinelli RT, et al. In the absence of endogenous IL-10, mice acutely infected with Toxoplasma gondii succumb to a lethal immune response dependent on CD4+ T cells and accompanied by overproduction of IL-12, IFN-gamma and TNF-alpha. J Immunol. 1996;157(2):798–805.
    View this article via: PubMed Google Scholar
83. Jankovic D, et al. Conventional T-bet(+)Foxp3(-) Th1 cells are the major source of host-protective regulatory IL-10 during intracellular protozoan infection. J Exp Med. 2007;204(2):273–283.
    View this article via: PubMed CrossRef Google Scholar
84. Hall AO, et al. The cytokines interleukin 27 and interferon-γ promote distinct Treg cell populations required to limit infection-induced pathology. Immunity. 2012;37(3):511–523.
    View this article via: PubMed CrossRef Google Scholar
85. Oldenhove G, et al. Decrease of Foxp3+ Treg cell number and acquisition of effector cell phenotype during lethal infection. Immunity. 2009;31(5):772–786.
    View this article via: PubMed CrossRef Google Scholar
86. Rosowski EE, et al. Strain-specific activation of the NF-kappaB pathway by GRA15, a novel Toxoplasma gondii dense granule protein. J Exp Med. 2011;208(1):195–212.
    View this article via: PubMed CrossRef Google Scholar
87. Braun L, et al. A Toxoplasma dense granule protein, GRA24, modulates the early immune response to infection by promoting a direct and sustained host p38 MAPK activation. J Exp Med. 2013;210(10):2071–2086.
    View this article via: PubMed CrossRef Google Scholar
88. Butcher BA, et al. Toxoplasma gondii rhoptry kinase ROP16 activates STAT3 and STAT6 resulting in cytokine inhibition and arginase-1-dependent growth control. PLoS Pathog. 2011;7(9):e1002236.
    View this article via: PubMed CrossRef Google Scholar
89. Ong YC, Reese ML, Boothroyd JC. Toxoplasma rhoptry protein 16 (ROP16) subverts host function by direct tyrosine phosphorylation of STAT6. J Biol Chem. 2010;285(37):28731–28740.
    View this article via: PubMed CrossRef Google Scholar
90. Olias P, Etheridge RD, Zhang Y, Holtzman MJ, Sibley LD. Toxoplasma effector recruits the Mi-2/NuRD complex to repress STAT1 transcription and block IFN-γ-dependent gene expression. Cell Host Microbe. 2016;20(1):72–82.
    View this article via: PubMed CrossRef Google Scholar
91. Gay G, et al. Toxoplasma gondii TgIST co-opts host chromatin repressors dampening STAT1-dependent gene regulation and IFN-γ-mediated host defenses. J Exp Med. 2016;213(9):1779–1798.
    View this article via: PubMed CrossRef Google Scholar
92. Braun L, et al. The Toxoplasma effector TEEGR promotes parasite persistence by modulating NF-κB signalling via EZH2. Nat Microbiol. 2019;4(7):1208–1220.
    View this article via: PubMed CrossRef Google Scholar
93. Konradt C, et al. Endothelial cells are a replicative niche for entry of Toxoplasma gondii to the central nervous system. Nat Microbiol. 2016;1:16001.
    View this article via: PubMed Google Scholar
94. Estato V, et al. The neurotropic parasite Toxoplasma gondii induces sustained neuroinflammation with microvascular dysfunction in infected mice. Am J Pathol. 2018;188(11):2674–2687.
    View this article via: PubMed CrossRef Google Scholar
95. Schneider CA, et al. Imaging the dynamic recruitment of monocytes to the blood-brain barrier and specific brain regions during Toxoplasma gondii infection. Proc Natl Acad Sci U S A. 2019;116(49):24796–24807.
    View this article via: PubMed CrossRef Google Scholar
96. Bierly AL, Shufesky WJ, Sukhumavasi W, Morelli AE, Denkers EY. Dendritic cells expressing plasmacytoid marker PDCA-1 are Trojan horses during Toxoplasma gondii infection. J Immunol. 2008;181(12):8485–8491.
    View this article via: PubMed CrossRef Google Scholar
97. Courret N, Darche S, Sonigo P, Milon G, Buzoni-Gâtel D, Tardieux I. CD11c- and CD11b-expressing mouse leukocytes transport single Toxoplasma gondii tachyzoites to the brain. Blood. 2006;107(1):309–316.
    View this article via: PubMed CrossRef Google Scholar
98. Pusch L, Romeike B, Deckert M, Mawrin C. Persistent toxoplasma bradyzoite cysts in the brain: incidental finding in an immunocompetent patient without evidence of a toxoplasmosis. Clin Neuropathol. 2009;28(3):210–212.
    View this article via: PubMed Google Scholar
99. Mendez OA, Koshy AA. Toxoplasma gondii: entry, association, and physiological influence on the central nervous system. PLoS Pathog. 2017;13(7):e1006351.
    View this article via: PubMed CrossRef Google Scholar
100. 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(1):175–180.
     View this article via: PubMed Google Scholar
101. Fischer HG, Nitzgen B, Reichmann G, Gross U, Hadding U. Host cells of Toxoplasma gondii encystation in infected primary culture from mouse brain. Parasitol Res. 1997;83(7):637–641.
     View this article via: PubMed CrossRef Google Scholar
102. Cabral CM, et al. Neurons are the primary target cell for the brain-tropic intracellular parasite Toxoplasma gondii. PLoS Pathog. 2016;12(2):e1005447.
     View this article via: PubMed CrossRef Google Scholar
103. Melzer TC, Cranston HJ, Weiss LM, Halonen SK. Host cell preference of Toxoplasma gondii cysts in murine brain: a confocal study. J Neuroparasitology. 2010;1:N100505.
     View this article via: PubMed Google Scholar
104. Hidano S, et al. STAT1 signaling in astrocytes is essential for control of infection in the central nervous system. mBio. 2016;7(6):e01881-16.
     View this article via: PubMed Google Scholar
105. Melzer T, Duffy A, Weiss LM, Halonen SK. The gamma interferon (IFN-gamma)-inducible GTP-binding protein IGTP is necessary for toxoplasma vacuolar disruption and induces parasite egression in IFN-gamma-stimulated astrocytes. Infect Immun. 2008;76(11):4883–4894.
     View this article via: PubMed CrossRef Google Scholar
106. Halonen SK, Weiss LM. Investigation into the mechanism of gamma interferon-mediated inhibition of Toxoplasma gondii in murine astrocytes. Infect Immun. 2000;68(6):3426–3430.
     View this article via: PubMed CrossRef Google Scholar
107. Martens S, et al. Disruption of Toxoplasma gondii parasitophorous vacuoles by the mouse p47-resistance GTPases. PLoS Pathog. 2005;1(3):e24.
     View this article via: PubMed CrossRef Google Scholar
108. Peterson PK, Gekker G, Hu S, Chao CC. Human astrocytes inhibit intracellular multiplication of Toxoplasma gondii by a nitric oxide-mediated mechanism. J Infect Dis. 1995;171(2):516–518.
     View this article via: PubMed CrossRef Google Scholar
109. Däubener W, et al. Anti-parasitic effector mechanisms in human brain tumor cells: role of interferon-gamma and tumor necrosis factor-alpha. Eur J Immunol. 1996;26(2):487–492.
     View this article via: PubMed CrossRef Google Scholar
110. Bando H, et al. Toxoplasma effector GRA15-dependent suppression of IFN-γ-induced antiparasitic response in human neurons. Front Cell Infect Microbiol. 2019;9:140.
     View this article via: PubMed Google Scholar
111. Chao CC, Hu S, Gekker G, Novick WJ, Remington JS, Peterson PK. Effects of cytokines on multiplication of Toxoplasma gondii in microglial cells. J Immunol. 1993;150(8 pt 1):3404–3410.
     View this article via: PubMed Google Scholar
112. Chao CC, Gekker G, Hu S, Peterson PK. Human microglial cell defense against Toxoplasma gondii. The role of cytokines. J Immunol. 1994;152(3):1246–1252.
     View this article via: PubMed Google Scholar
113. Freund YR, Zaveri NT, Javitz HS. In vitro investigation of host resistance to Toxoplasma gondii infection in microglia of BALB/c and CBA/Ca mice. Infect Immun. 2001;69(2):765–772.
     View this article via: PubMed CrossRef Google Scholar
114. Sa Q, et al. Cutting Edge: IFN-γ produced by brain-resident cells is crucial to control cerebral infection with Toxoplasma gondii. J Immunol. 2015;195(3):796–800.
     View this article via: PubMed CrossRef Google Scholar
115. Wen X, Kudo T, Payne L, Wang X, Rodgers L, Suzuki Y. Predominant interferon-γ-mediated expression of CXCL9, CXCL10, and CCL5 proteins in the brain during chronic infection with Toxoplasma gondii in BALB/c mice resistant to development of toxoplasmic encephalitis. J Interferon Cytokine Res. 2010;30(9):653–660.
     View this article via: PubMed CrossRef Google Scholar
116. Ochiai E, et al. CXCL9 is important for recruiting immune T cells into the brain and inducing an accumulation of the T cells to the areas of tachyzoite proliferation to prevent reactivation of chronic cerebral infection with Toxoplasma gondii. Am J Pathol. 2015;185(2):314–324.
     View this article via: PubMed CrossRef Google Scholar
117. Salvioni A, et al. Robust control of a brain-persisting parasite through MHC I presentation by infected neurons. Cell Rep. 2019;27(11):3254–3268.e8.
     View this article via: PubMed CrossRef Google Scholar
118. Denkers EY, et al. Perforin-mediated cytolysis plays a limited role in host resistance to Toxoplasma gondii. J Immunol. 1997;159(4):1903–1908.
     View this article via: PubMed Google Scholar
119. Suzuki Y, et al. Removal of Toxoplasma gondii cysts from the brain by perforin-mediated activity of CD8+ T cells. Am J Pathol. 2010;176(4):1607–1613.
     View this article via: PubMed CrossRef Google Scholar
120. Tiwari A, Hannah R, Lutshumba J, Ochiai E, Weiss LM, Suzuki Y. Penetration of CD8⁺ cytotoxic T cells into large target, tissue cysts of Toxoplasma gondii, leads to its elimination. Am J Pathol. 2019;189(8):1594–1607.
     View this article via: PubMed CrossRef Google Scholar
121. Persson EK, et al. Death receptor ligation or exposure to perforin trigger rapid egress of the intracellular parasite Toxoplasma gondii. J Immunol. 2007;179(12):8357–8365.
     View this article via: PubMed CrossRef Google Scholar
122. Sidik SM, et al. A genome-wide CRISPR screen in Toxoplasma identifies essential apicomplexan genes. Cell. 2016;166(6):1423–1435.e12.
     View this article via: PubMed CrossRef Google Scholar
123. Shen B, Brown KM, Lee TD, Sibley LD. Efficient gene disruption in diverse strains of Toxoplasma gondii using CRISPR/CAS9. mBio. 2014;5(3):e01114–e01114.
     View this article via: PubMed Google Scholar
124. Soête 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(4):361–370.
     View this article via: PubMed CrossRef Google Scholar
125. 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(3):323–331.
     View this article via: PubMed CrossRef Google Scholar
126. Donald RG, et al. Toxoplasma gondii cyclic GMP-dependent kinase: chemotherapeutic targeting of an essential parasite protein kinase. Eukaryotic Cell. 2002;1(3):317–328.
     View this article via: PubMed CrossRef Google Scholar
127. Bohne W, Heesemann J, Gross U. Induction of bradyzoite-specific Toxoplasma gondii antigens in gamma interferon-treated mouse macrophages. Infect Immun. 1993;61(3):1141–1145.
     View this article via: PubMed CrossRef Google Scholar
128. Swierzy IJ, et al. Divergent co-transcriptomes of different host cells infected with Toxoplasma gondii reveal cell type-specific host-parasite interactions. Sci Rep. 2017;7(1):7229.
     View this article via: PubMed CrossRef Google Scholar
129. Swierzy IJ, Lüder CG. Withdrawal of skeletal muscle cells from cell cycle progression triggers differentiation of Toxoplasma gondii towards the bradyzoite stage. Cell Microbiol. 2015;17(1):2–17.
     View this article via: PubMed CrossRef Google Scholar
130. Saksouk N, et al. Histone-modifying complexes regulate gene expression pertinent to the differentiation of the protozoan parasite Toxoplasma gondii. Mol Cell Biol. 2005;25(23):10301–10314.
     View this article via: PubMed CrossRef Google Scholar
131. Naguleswaran A, Elias EV, McClintick J, Edenberg HJ, Sullivan WJ. Toxoplasma gondii lysine acetyltransferase GCN5-A functions in the cellular response to alkaline stress and expression of cyst genes. PLoS Pathog. 2010;6(12):e1001232.
     View this article via: PubMed CrossRef Google Scholar
132. Bougdour A, et al. Drug inhibition of HDAC3 and epigenetic control of differentiation in Apicomplexa parasites. J Exp Med. 2009;206(4):953–966.
     View this article via: PubMed CrossRef Google Scholar
133. Sullivan WJ, Narasimhan J, Bhatti MM, Wek RC. Parasite-specific eIF2 (eukaryotic initiation factor-2) kinase required for stress-induced translation control. Biochem J. 2004;380(pt 2):523–531.
     View this article via: PubMed Google Scholar
134. Konrad C, Queener SF, Wek RC, Sullivan WJ. Inhibitors of eIF2α dephosphorylation slow replication and stabilize latency in Toxoplasma gondii. Antimicrob Agents Chemother. 2013;57(4):1815–1822.
     View this article via: PubMed CrossRef Google Scholar
135. Walker R, et al. The Toxoplasma nuclear factor TgAP2XI-4 controls bradyzoite gene expression and cyst formation. Mol Microbiol. 2013;87(3):641–655.
     View this article via: PubMed CrossRef Google Scholar
136. Hong DP, Radke JB, White MW. Opposing transcriptional mechanisms regulate Toxoplasma development. mSphere. 2017;2(1):e00347-16.
     View this article via: PubMed Google Scholar
137. Radke JB, et al. Transcriptional repression by ApiAP2 factors is central to chronic toxoplasmosis. PLoS Pathog. 2018;14(5):e1007035.
     View this article via: PubMed CrossRef Google Scholar
138. Waldman BS, Schwarz D, Wadsworth MH, Saeij JP, Shalek AK, Lourido S. Identification of a master regulator of differentiation in Toxoplasma. Cell. 2020;180(2):359–372.e16.
     View this article via: PubMed CrossRef Google Scholar
139. Skariah S, McIntyre MK, Mordue DG. Toxoplasma gondii: determinants of tachyzoite to bradyzoite conversion. Parasitol Res. 2010;107(2):253–260.
     View this article via: PubMed CrossRef Google Scholar
140. Buchholz KR, Bowyer PW, Boothroyd JC. Bradyzoite pseudokinase 1 is crucial for efficient oral infectivity of the Toxoplasma gondii tissue cyst. Eukaryotic Cell. 2013;12(3):399–410.
     View this article via: PubMed CrossRef Google Scholar
141. Zhang YW, Halonen SK, Ma YF, Wittner M, Weiss LM. Initial characterization of CST1, a Toxoplasma gondii cyst wall glycoprotein. Infect Immun. 2001;69(1):501–507.
     View this article via: PubMed CrossRef Google Scholar
142. Tomita T, et al. The Toxoplasma gondii cyst wall protein CST1 is critical for cyst wall integrity and promotes bradyzoite persistence. PLoS Pathog. 2013;9(12):e1003823.
     View this article via: PubMed CrossRef Google Scholar
143. Caffaro CE, Koshy AA, Liu L, Zeiner GM, Hirschberg CB, Boothroyd JC. A nucleotide sugar transporter involved in glycosylation of the Toxoplasma tissue cyst wall is required for efficient persistence of bradyzoites. PLoS Pathog. 2013;9(5):e1003331.
     View this article via: PubMed CrossRef Google Scholar
144. Nance JP, et al. Chitinase dependent control of protozoan cyst burden in the brain. PLoS Pathog. 2012;8(11):e1002990.
     View this article via: PubMed CrossRef Google Scholar
145. Wang AW, et al. Genome-wide association study in two populations to determine genetic variants associated with Toxoplasma gondii infection and relationship to schizophrenia risk. Prog Neuropsychopharmacol Biol Psychiatry. 2019;92:133–147.
     View this article via: PubMed CrossRef Google Scholar
146. Fekkar A, et al. Direct genotyping of Toxoplasma gondii in ocular fluid samples from 20 patients with ocular toxoplasmosis: predominance of type II in France. J Clin Microbiol. 2011;49(4):1513–1517.
     View this article via: PubMed CrossRef Google Scholar
147. Herrmann DC, et al. Genotyping of samples from German patients with ocular, cerebral and systemic toxoplasmosis reveals a predominance of Toxoplasma gondii type II. Int J Med Microbiol. 2014;304(7):911–916.
     View this article via: PubMed CrossRef Google Scholar
148. Holland GN. Ocular toxoplasmosis: a global reassessment. Part I: epidemiology and course of disease. Am J Ophthalmol. 2003;136(6):973–988.
     View this article via: PubMed CrossRef Google Scholar
149. Bosch-Driessen LE, Berendschot TT, Ongkosuwito JV, Rothova A. Ocular toxoplasmosis: clinical features and prognosis of 154 patients. Ophthalmology. 2002;109(5):869–878.
     View this article via: PubMed CrossRef Google Scholar
150. Vishnevskia-Dai V, et al. Chorio-retinal toxoplasmosis: treatment outcomes, lesion evolution and long-term follow-up in a single tertiary center. Int Ophthalmol. 2020;40(4):811–821.
     View this article via: PubMed CrossRef Google Scholar
151. Smith JR, Franc DT, Carter NS, Zamora D, Planck SR, Rosenbaum JT. Susceptibility of retinal vascular endothelium to infection with Toxoplasma gondii tachyzoites. Invest Ophthalmol Vis Sci. 2004;45(4):1157–1161.
     View this article via: PubMed CrossRef Google Scholar
152. Tedesco RC, Smith RL, Corte-Real S, Calabrese KS. Ocular toxoplasmosis: the role of retinal pigment epithelium migration in infection. Parasitol Res. 2004;92(6):467–472.
     View this article via: PubMed CrossRef Google Scholar
153. Charles E, et al. CD4 T-cell suppression by cells from Toxoplasma gondii-infected retinas is mediated by surface protein PD-L1. Infect Immun. 2010;78(8):3484–3492.
     View this article via: PubMed CrossRef Google Scholar
154. Calabrese KS, Tedesco RC, Zaverucha do Valle T, Barbosa HS. Serum and aqueous humour cytokine response and histopathological alterations during ocular Toxoplasma gondii infection in C57BL/6 mice. Micron. 2008;39(8):1335–1341.
     View this article via: PubMed CrossRef Google Scholar
155. Rochet É, et al. Interleukin-6-driven inflammatory response induces retinal pathology in a model of ocular toxoplasmosis reactivation. Infect Immun. 2015;83(5):2109–2117.
     View this article via: PubMed CrossRef Google Scholar
156. Iqbal J, Al-Awadhi MA, Raghupathy RG. TGF-β1 levels and intraocular tissue alterations in mice infected with a virulent type I RH Toxoplasma gondii strain. Exp Parasitol. 2016;162:57–63.
     View this article via: PubMed CrossRef Google Scholar
157. de-la-Torre A, et al. Severe South American ocular toxoplasmosis is associated with decreased Ifn-γ/Il-17a and increased Il-6/Il-13 intraocular levels. PLoS Negl Trop Dis. 2013;7(11):e2541.
     View this article via: PubMed CrossRef Google Scholar
158. 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(15):6442–6447.
     View this article via: PubMed CrossRef Google Scholar
159. Berdoy M, Webster JP, Macdonald DW. Fatal attraction in rats infected with Toxoplasma gondii. Proc Biol Sci. 2000;267(1452):1591–1594.
     View this article via: PubMed CrossRef Google Scholar
160. Wang ZT, Harmon S, O’Malley KL, Sibley LD. Reassessment of the role of aromatic amino acid hydroxylases and the effect of infection by Toxoplasma gondii on host dopamine. Infect Immun. 2015;83(3):1039–1047.
     View this article via: PubMed CrossRef Google Scholar
161. McFarland R, et al. AAH2 gene is not required for dopamine-dependent neurochemical and behavioral abnormalities produced by Toxoplasma infection in mouse. Behav Brain Res. 2018;347:193–200.
     View this article via: PubMed CrossRef Google Scholar
162. Wang ZT, Verma SK, Dubey JP, Sibley LD. The aromatic amino acid hydroxylase genes AAH1 and AAH2 in Toxoplasma gondii contribute to transmission in the cat. PLoS Pathog. 2017;13(3):e1006272.
     View this article via: PubMed CrossRef Google Scholar
163. Ingram WM, Goodrich LM, Robey EA, Eisen MB. Mice infected with low-virulence strains of Toxoplasma gondii lose their innate aversion to cat urine, even after extensive parasite clearance. PLoS One. 2013;8(9):e75246.
     View this article via: PubMed CrossRef Google Scholar
164. Ferrero DM, et al. Detection and avoidance of a carnivore odor by prey. Proc Natl Acad Sci U S A. 2011;108(27):11235–11240.
     View this article via: PubMed CrossRef Google Scholar
165. Arsenijevic D, Bilbao FD, Giannakopoulos P, Girardier L, Samec S, Richard D. A role for interferon-gamma in the hypermetabolic response to murine toxoplasmosis. Eur Cytokine Netw. 2001;12(3):518–527.
     View this article via: PubMed Google Scholar
166. Zhou CX, Zhou DH, Elsheikha HM, Liu GX, Suo X, Zhu XQ. Global metabolomic profiling of mice brains following experimental infection with the cyst-forming Toxoplasma gondii. PLoS One. 2015;10(10):e0139635.
     View this article via: PubMed CrossRef Google Scholar
167. Zhou CX, Zhou DH, Elsheikha HM, Zhao Y, Suo X, Zhu XQ. Metabolomic profiling of mice serum during toxoplasmosis progression using liquid chromatography-mass spectrometry. Sci Rep. 2016;6:19557.
     View this article via: PubMed Google Scholar
168. Melchor SJ, Ewald SE. Disease tolerance in Toxoplasma infection. Front Cell Infect Microbiol. 2019;9:185.
     View this article via: PubMed Google Scholar
169. Kugler DG, et al. Systemic toxoplasma infection triggers a long-term defect in the generation and function of naive T lymphocytes. J Exp Med. 2016;213(13):3041–3056.
     View this article via: PubMed CrossRef Google Scholar
170. Arsenijevic D, et al. Metabolic-cytokine responses to a second immunological challenge with LPS in mice with T. gondii infection. Am J Physiol. 1998;274(3):E439–E445.
     View this article via: PubMed Google Scholar
171. Melchor SJ, et al. IL-1R regulates disease tolerance and cachexia in toxoplasma gondii infection. J Immunol. 2020;204(12):3329–3338.
     View this article via: PubMed CrossRef Google Scholar
172. Jin RM, Blair SJ, Warunek J, Heffner RR, Blader IJ, Wohlfert EA. Regulatory T cells promote myositis and muscle damage in Toxoplasma gondii infection. J Immunol. 2017;198(1):352–362.
     View this article via: PubMed CrossRef Google Scholar