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Toxoplasma gondii: Entry, association, and physiological influence on the central nervous system

  • Oscar A. Mendez,

    Affiliations Graduate Interdisciplinary Program in Neuroscience, University of Arizona, Tucson, Arizona, United States of America, BIO5 Institute, University of Arizona, Tucson, Arizona, United States of America

    ORCID http://orcid.org/0000-0003-3476-5864

  • Anita A. Koshy

    akoshy@email.arizona.edu

    Affiliations BIO5 Institute, University of Arizona, Tucson, Arizona, United States of America, Department of Immunobiology, University of Arizona, Tucson, Arizona, United States of America, Department of Neurology, University of Arizona, Tucson, Arizona, United States of America

Toxoplasma gondii: Entry, association, and physiological influence on the central nervous system

  • Oscar A. Mendez, 
  • Anita A. Koshy
PLOS
x

Abstract

Toxoplasma gondii is one of the world’s most successful parasites, in part because of its ability to infect and persist in most warm-blooded animals. A unique characteristic of T. gondii is its ability to persist in the central nervous system (CNS) of a variety of hosts, including humans and rodents. How, what, and why T. gondii encysts in the CNS has been the topic of study for decades. In this review, we will discuss recent work on how T. gondii is able to traverse the unique barrier surrounding the CNS, what cells of the CNS play host to T. gondii, and finally, how T. gondii infection may influence global and cellular physiology of the CNS.

Introduction

Toxoplasma gondii is an obligate intracellular parasite of the phyla Apicomplexa. Felids are the only definitive host for T. gondii, but T. gondii has a wide intermediate host range and has been documented to naturally infect most warm-blooded animals including birds, rodents, and humans [1]. In most hosts, T. gondii establishes a life-long, latent infection in tissues such as skeletal muscle, cardiac muscle, or the central nervous system (CNS), which includes the brain, the spinal cord, and the retina. In this review, we will often use CNS interchangeably with the brain, in which the majority of the work on T. gondii has been done.

Humans primarily acquire T. gondii through contaminated food or water or via vertical transmission. T. gondii is found worldwide and seroprevalence rates range from <10% to >60% [2]. During acute infection, T. gondii disseminates throughout the host as a tachyzoite, the fast-replicating form of the parasite, which is targeted by the host immune response [3,4]. As infection proceeds, T. gondii transitions into the chronic stage of infection via conversion to the slowly replicating bradyzoite, which encysts. A number of transitions occur between tachyzoites and bradyzoites/cysts, some of which are thought to enable the bradyzoite/cyst to escape immune detection, thereby leading to persistent infection [5]. In humans and rodents, the brain is the major organ of encystment and persistence [1,6].

The ability of T. gondii to asymptomatically persist in the CNS of immunocompetent individuals is highly unusual, as when most microbes cross into the CNS, symptomatic and often lethal disease ensues. This tropism for the CNS underlies the devastating disease T. gondii causes in those with deficient immune responses—e.g., developing fetuses or AIDS patients. Finally, T. gondii’s predilection for the CNS has been linked to a number of behavioral deficits in rodents [711]. Thus, the T. gondii–CNS interaction is of particular interest for understanding symptomatic toxoplasmosis as well as rodent behavioral changes. As a summary of the behavioral deficits and possible causes has been reviewed recently [1214], here we will focus on work exploring how T. gondii enters the CNS, establishes a persistent infection, and affects CNS physiology.

Brief overview of the CNS

The CNS is a complex organ composed of multiple cells that include neurons and glia. Glia were originally defined as neuronal support cells and consist of oligodendrocytes, astrocytes, and microglia, which are tissue-resident macrophages that have a hematopoietic origin as opposed to neuroectodermal origin [15,16]. What percentage of glia are astrocytes, oligodendrocytes, or microglia will vary by location and state (e.g., baseline versus infected). Neurons are the signaling cells of the CNS and comprise a vast array of subtypes distributed across the CNS in a heterogeneous manner. Regional and long-distance connections between neurons underlie the intricate processing network that allows complex integration of information from external stimuli to initiate and generate movements and complex behaviors. Oligodendrocytes insulate the axons of neurons with myelin, allowing for proper signal conduction. Astrocytes are support units responsible for promoting efficient signaling between neurons, release of growth factors, and separation of neuronal groups [17]. When these cell populations are discussed, the number of glia to neurons is often overestimated. Careful counts suggest that glia and neurons are approximately equivalent in number across a range of mammals, including humans [1820]. Astrocyte and oligodendrocyte turnover does occur in the CNS, but this turnover becomes attenuated in adulthood, with a vast majority of glia becoming oligodendrocytes [2123]. For neurons, even in adulthood, replication occurs but only in distinct regions of the brain and in limited amounts [24]. While microglia and infiltrating macrophages are essential for controlling T. gondii in the brain, neurons and astrocytes are the parenchymal cells that have been most implicated in playing a role in CNS toxoplasmosis.

Entering the CNS

For any pathogen to enter the CNS, it must cross the blood-brain barrier (BBB). The BBB is a selective barrier that is composed of endothelial cells that line microvessels in the brain. The basal lamina and pericytes surround endothelial cells, followed by enclosing astrocytic endfeet (Fig 1A). These cellular interactions allow the BBB to exclude large peptides and proteins and only allow free diffusion of small gaseous molecules like oxygen and carbon dioxide. At baseline, the presence of tight junctions and adherence junctions excludes paracellular movement of hydrophilic molecules and the migration of cells past the endothelial barrier [25]. Though originally conceived of as a highly impermeable barrier through which most organisms and cells could not pass, it is now recognized that microbes have developed multiple mechanisms for crossing the BBB, and in the right context, immune cells cross into the CNS for immune surveillance [2629]. For T. gondii, 3 mechanisms have been proposed for CNS entry: paracellular crossing, transcellular crossing, or the so-called “Trojan horse” mechanism, whereby an infected immune cell crosses the BBB, bringing the intracellular parasite with it (Fig 1B).

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Fig 1. Diagram of the physical and cellular interactions that compose the blood-brain barrier.

(A) The blood-brain barrier (BBB) is composed of microvessels surrounded by an endothelial cell layer with tight junctions; pericytes surround the tight junctions and then astrocytic processes or endfeet provide the final layer. (B) Toxoplasma gondii has been postulated to have 3 mechanisms for crossing the BBB. (1) Paracellular entry, in which T. gondii migrates directly through the tight junctions of the endothelial cell layer, (2) transcellular entry, in which free parasites in the vascular compartment infect endothelial cells, replicate, and then egress out of the basolateral side of endothelial cell (lysing the host cell), (3) the “Trojan horse” method, whereby an infected immune cell infiltrates the CNS, after which the parasite egresses out of the immune cell and into the brain parenchyma.

https://doi.org/10.1371/journal.ppat.1006351.g001

Paracellular entry

Several lines of evidence suggest that T. gondii could cross into the CNS via the paracellular method (Fig 1B, part 1). Though T. gondii lacks cilia and flagella, the parasite is able to propel itself using actin-myosin motors, generating movement termed “gliding motility” [30]. This gliding motility is thought to aid T. gondii across the first barrier it encounters: the epithelium of the small intestine [31]. The gut epithelial barrier shares many features with the BBB including tight junctions, paracellular junctions, regulation of barrier properties, and an immune barrier [32,33]. Parasites are able to cross polarized cell monolayers and extracellular matrix, which mimic both the BBB and gut epithelial barrier [34]. In addition, parasites are able to transmigrate across the intestinal epithelium ex vivo [34]. Importantly, recent work using physiological shear force applied to live-cell microfluidic chambers has shown that tachyzoites are capable of adhering to and migrating on vascular endothelium in these semiphysiologic conditions [35], though actual paracellular crossing by tachyzoites was not observed in these conditions (personal communication, M. Lodoen to A. Koshy).

Transcellular migration

While Plasmodium sporozoites have been observed to migrate through cells [31], for T. gondii, transcellular migration refers to infection of a cell followed by replication then lyses or egress from the basolateral side (Fig 1B, part 2). This concept has been primarily conceptualized for crossing from the gut epithelium into the intestinal lamina propria [36,37]. However, a recent study utilizing transgenic reporter systems and multiphoton in vivo imaging suggests such a mechanism may also have a role in how T. gondii crosses the BBB. In this study, Konradt et al. found infected endothelial cells in multiple organs including the brain. Further work showed that free tachyzoites in the bloodstream were able to adhere to CNS endothelial cells, invade, replicate, and eventually egress from these cells, potentially depositing these egressing parasites into the CNS parenchyma [38]. This work also used in vitro assays to show that, in shear stress conditions, parasites were able to attach and invade endothelial cells, especially at lower shear forces, consistent with the in vivo observation that parasite-infected endothelial cells were primarily found in smaller diameter vessels [38]. Finally, this study noted a lack of infected infiltrating immune cells early in CNS infection, consistent with prior work that also noted CNS parasite infection preceding immune-cell infiltration into the CNS [39].

Infected immune cells

The final mechanism proposed for T. gondii entry into the CNS is via the “Trojan horse” mechanism (infected immune cells) (Fig 1B, part 3). Several studies using in vitro models have shown that infected immune cells have increased motility and are capable of crossing endothelial barriers, including during shear stress [4042]. Additionally, intravenous inoculation of mice with infected macrophages or dendritic cells led to a more rapid appearance of parasites in the CNS compared to inoculation with free parasites [40,43], though multiple mechanisms (not just increased BBB crossing by infected immune cells) might explain these results.

In summary, T. gondii may enter the CNS through multiple mechanisms. Outstanding questions include the relative importance of each mode of entry and how different mechanisms of entry might affect which regions of the CNS are infected or even which CNS cells directly interact with T. gondii.

CNS host cell interactions

Given the importance of CNS persistence to clinical disease, where and how T. gondii persists in the CNS in the immunocompetent host has long been an area of interest. Human data on CNS regions “susceptible” to T. gondii primarily come from autopsies of AIDS patients. In these studies of severely immunocompromised patients, T. gondii lesions were consistently found in the cerebral cortex and basal ganglia, with fewer lesions in the cerebellum, brainstem, and spinal cord [4446]. These data are consistent with the localization of cysts observed in rodents [4749].

Host cell–T. gondii interactions of immunocompetent humans are not well characterized. One recent report from primarily immunocompetent patients states that T. gondii was seen in both neurons and astrocytes, but it is unclear from the methodology how this observation was determined [50]. Given the lack of human data, our understanding of CNS host cell–T. gondii interactions has, by necessity, come from in vitro and rodent models of CNS toxoplasmosis. In mouse models of toxoplasmosis, some controversy existed over whether T. gondii cysts were intra- or extracellular [51,52], but the earliest reports using electron microscopy (EM) suggested that cysts were within cells, though this work did not identify the specific cell type [53,54]. Subsequent improvements in EM led to the identification of synapses near cysts, suggesting that cysts persisted within neurons [55]. The finding that cysts are intracellular and primarily within neurons was further supported by additional studies, including 1 that utilized parasites recently isolated from AIDS patients and a common lab-passaged strain [56,57]. A recent study utilizing immunofluorescence and confocal microscopy also found that cysts were primarily associated with antineuronal labeling [58].

The recognition that T. gondii persists primarily in neurons raised the question of why T. gondii—a parasite that can invade most nucleated cells in vitro [59] and that brings its own invasion machinery [60]—predominantly persists in neurons. In vitro studies showed that T. gondii readily infects and encysts in both astrocytes and neurons [61,62], but astrocytes were capable of using multiple mechanisms to clear intracellular parasites [6366]. Conversely, neurons were not capable of such clearance [67], a finding in line with other evidence that neurons lacked full immune-response capabilities [68]. Based upon these studies and the previously reviewed in vivo work, the working model for T. gondii–CNS host cell interactions was that after entering the CNS, parasites invaded both astrocytes and neurons. Astrocytes then cleared the intracellular parasites while neurons could not, leaving neurons as the host cell for persistent infection.

Until recently, this model could not be tested in vivo as there was no way to identify host cells that had been invaded but cleared the intracellular parasite. The advent of the T. gondii–Cre system, an in vivo system in which parasites trigger permanent host-cell expression of green fluorescent protein (GFP) even when the cell is not productively infected, offered a platform to test this model (Fig 2) [69,70].

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Fig 2. Using the Toxoplasma gondii–Cre system to test 2 models of why T. gondii primarily persists in neurons in vivo.

(A) After entering the CNS, T. gondii should be able to interact with different cells in the brain, including both astrocytes (orange) and neurons (beige). As the T. gondii–Cre system leads to the expression of a green fluorescent protein (GFP) in host cells injected with T. gondii protein regardless of infection status, it can help distinguish between 2 likely models for T. gondii persistence in neurons. (B) Model 1: after infiltration of the CNS, T. gondii interacts with and invades both astrocytes and neurons, causing both cell types to express GFP (green). Astrocytes clear the intracellular parasite while neurons cannot, leaving neurons as the primary host cell for persistent infection. (C) Model 2: after infiltration of the CNS, T. gondii almost exclusively interacts with and invades neurons, leading to GFP expression primarily in neurons. Neurons potentially clear some but not all invading parasites, leaving neurons as the primary host cell for persistent infection.

https://doi.org/10.1371/journal.ppat.1006351.g002

Studies using this system suggest that (1) T. gondii interacts with far more CNS cells than previously described, (2) the majority of these interactions do not lead to productive infection, and (3) throughout CNS infection, T. gondii predominantly and almost exclusively interacts with neurons [70,71]. These data suggest a new model for T. gondii–CNS host cells, one in which T. gondii persists in neurons because parasites primarily interact with and invade neurons.

Many outstanding questions remain, which include (1) determining what factors drive the T. gondii–neuron interaction, (2) if the injected but uninfected neurons arise through aborted invasion and/or neuron clearance of intracellular parasites, and (3) how uninfected, injected neurons differ from (or are the same as) infected neurons, especially in terms of neuroanatomic localization, gene expression, and electrophysiology.

Physiological changes and effects during T. gondii infection

Until recently, little work had been done on how T. gondii changes CNS physiology, but in the last several years, a number of important in vivo studies have begun to address this question. A major mechanism for affecting CNS physiology would be through changes in neurotransmitters, the molecules that neurons use for interneuronal communication. In vivo measurements of neurotransmitters have primarily measured global changes within the brain, not cell-specific changes. The dopaminergic system has been of major interest because dopamine is essential for locomotor activity (movement) and various forms of learning, including fear [72]. As such, investigators have sought to implicate the dopaminergic system in infection-induced behavioral changes [9]. Several studies that directly measured CNS dopamine levels or dopamine metabolites suggest that postinfection, CNS dopamine levels increase [73,74], though another group was unable to confirm these changes [75]. One explanation for these contradictory findings is that each group used different mouse strains, which can affect the immune response to T. gondii [76,77]. As immune cells have been shown to make dopamine [78] and none of the groups determined the cellular source of the measured dopamine or metabolites, these differences might simply reflect differing levels of immune infiltration into the CNS rather than changes in the CNS dopaminergic cells or pathways.

Two recent papers have focused on the neuron neurotransmitter–physiology link and collectively found that the infected CNS shows altered excitability. David et al. found that glutamate, the major excitatory neurotransmitter of the CNS, was increased in the CNS of infected mice secondary to disruption of the normal reuptake of extracellular glutamate by astrocytes via the glutamate transporter (GLT-1) (Fig 3). They further showed that these changes were correlated with decreases in neuronal dendritic spine density (an early sign of distressed neurons) as well as depth-electrode electroencephalogram (EEG) changes and that most of these changes could be reversed with a brief treatment with Ceftriaxone, an antibiotic that increases GLT-1 levels [79]. Brooks et al. examined the effect of T. gondii infection on γ-aminobutyric acid (GABA), the major inhibitory neurotransmitter in the brain. Within the hippocampus, they showed that glutamate decarboxylase (GAD), a key enzyme responsible for the conversion of glutamate to GABA and the most common marker for identifying GABAergic neurons, was mislocalized within neurons in this region, though the absolute levels of GAD were unchanged compared to uninfected mice. Though the authors did not directly measure GABA levels, they did use skull EEG recordings to show that infected mice had an increase in spontaneous seizures and sensitivity to drug-induced seizures, both of which would be expected consequences of the loss of neuronal inhibition [80]. In addition to effects on neurotransmitters, infection has been noted to cause physical changes to the CNS. One study that evaluated mice infected with T. gondii for 8 months found that infected mice had both an increase in the size of the ventricles, fluid-filled spaces in the center of the brain, as well as some asymmetry of the brain parenchyma. While both findings could be explained by loss of parenchymal cells in the brain, histologic studies showed no neuronal loss, axonal injury, or “extensive” demyelination [11]. However, another study using diffusion tensor imaging, an MRI method used to evaluate white matter projections (the axons of neurons and the oligodendrocytes that produce the myelin that ensheaths axons) found mistargeting of discrete neuronal projections in mice chronically infected for 4–5 months compared to control animals. This study specifically focused on the somatosensory cortex (SSC), the part of the brain involved in tactile modes of sensation, as the authors had noted abnormalities in the SSC in full brain evaluations. Following up with immunofluorescence and Western blots, the authors observed reductions in dendritic arborization, spine number, and essential synaptic proteins, suggesting that the synaptic connections of the SSC had been disrupted [81]. This latter study suggests that T. gondii infection causes discrete areas of disruption and neuronal loss, which may be missed with global surveys rather than focal studies.

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Fig 3. Schematic of Toxoplasma gondii effects on glutamate and glutamate decarboxylase.

(A) Under normal conditions, glutamate, an excitatory neurotransmitter, is released into the synaptic cleft from the presynaptic neuron. Glutamate then diffuses across the synaptic cleft to act on glutamate receptors on the postsynaptic neuron, leading to excitation of the postsynaptic neuron. This glutamatergic signaling is terminated by uptake and recycling of synaptic glutamate by the glutamate transporter GLT-1 on surrounding astrocytes. Glutamate uptake by GLT-1 is essential to avoid excessive glutamate signaling, which can lead to postsynaptic excitotoxicity and neuronal death. After an infection, GLT-1–dependent transport into astrocytes is impaired, allowing for an increase in glutamate accumulation in the synaptic cleft, which is expected to lead to more excitation of the postsynaptic neuron. (B) Glutamate decarboxylase (GAD) is primarily found on presynaptic terminals, where it will process glutamate into γ-aminobutyric acid (GABA), the major inhibitory neurotransmitter in the brain. Once GABA is released into the synaptic cleft, it will bind onto GABA receptors on the postsynaptic neuron, which decreases the excitability of the postsynaptic neuron. In the case of infection, GAD is redistributed into the cytosol of the presynaptic neuron, which would be expected to cause improper GABA localization at synapses, leading to decreased inhibition of the postsynaptic neuron.

https://doi.org/10.1371/journal.ppat.1006351.g003

In summary, what has become apparent is that the form and physiology of the rodent brain are affected during both acute and chronic infection. The mechanisms that cause these changes in CNS physiology may be due to (1) direct parasite interactions via injection of effector proteins or persistence within a host neuron, (2) indirect effects of the CNS immune response to control the infection, or (3) both. Additionally, as these studies focused on relatively acute infections, the duration of these changes has yet to be determined. In addition, it remains unclear how these findings translate to human outcomes, though both congenitally infected patients and AIDS patients with toxoplasmic encephalitis are known to have seizures [82,83].

Conclusion

In the last decade, important work has been done to better define many molecular and cellular aspects of the T. gondii–CNS interaction. While these studies leave a number of outstanding questions as noted above, they form the foundation of an exciting time in the evolution of our understanding of CNS toxoplasmosis.

Acknowledgments

We wish to thank the entire Koshy lab for helpful discussion and review.

References

  1. 1. Dubey J. Toxoplasmosis of Animals and Humans, Second Edition [Internet]. Parasites & Vectors. CRC Press; 2009.
  2. 2. Pappas G, Roussos N, Falagas ME. Toxoplasmosis snapshots: Global status of Toxoplasma gondii seroprevalence and implications for pregnancy and congenital toxoplasmosis. Int J Parasitol. Australian Society for Parasitology Inc.; 2009;39: 1385–1394. pmid:19433092
  3. 3. Partanen P, Turunen HJ, Paasivuo RA, Leinikki PO. Immunoblot analysis of Toxoplasma gondii antigens by human immunoglobulins G, M, and A antibodies at different stages of infection. J Clin Microbiol. 1984;20: 133–135. pmid:6746885
  4. 4. Bessieres MH, Le Breton S, Seguela JP. Analysis by immunoblotting of Toxoplasma gondii exo-antigens and comparison with somatic antigens. Parasitol Res. 1992;78: 222–228. Available: http://www.ncbi.nlm.nih.gov/pubmed/1589430 pmid:1589430
  5. 5. Kim S- K, Boothroyd JC. Stage-specific expression of surface antigens by Toxoplasma gondii as a mechanism to facilitate parasite persistence. J Immunol. 2005;174: 8038–8048. pmid:15944311
  6. 6. Remington JS, Cavanaugh EN. Isolation of the encysted form of Toxoplasma gondii from human skeletal muscle and brain. N Engl J Med. Massachusetts Medical Society; 1965;273: 1308–1310. pmid:5852454
  7. 7. Berdoy M, Webster JP, Macdonald DW. Parasite-altered behavior: is the effect of Toxoplasma gondiion Rattus norvegicus specific? Parasitology. 1995;111: 403–409. pmid:11023404
  8. 8. Berdoy M, Webster JP, Macdonald DW. Fatal attraction in rats infected with Toxoplasma gondii. Proc R Soc B. 2000;267: 1591–1594. pmid:11007336
  9. 9. Vyas A, Kim S-K, 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: 6442–7. pmid:17404235
  10. 10. Tan D, Vyas A. Infection of male rats with Toxoplasma gondii induces effort-aversion in a T-maze decision-making task. Brain Behav Immun. Elsevier Inc.; 2016;53: 273–277. pmid:26783701
  11. 11. Hermes G, Ajioka JW, Kelly KA, Mui E, Roberts F, Kasza K, et al. Neurological and behavioral abnormalities, ventricular dilatation, altered cellular functions, inflammation, and neuronal injury in brains of mice due to common, persistent, parasitic infection. J Neuroinflammation. BioMed Central; 2008;5: 48. pmid:18947414
  12. 12. da Silva RC, Langoni H. Toxoplasma gondii: host–parasite interaction and behavior manipulation. Parasitol Res. 2009;105: 893–898. pmid:19548003
  13. 13. Vyas A. Mechanisms of Host Behavioral Change in Toxoplasma gondii Rodent Association. PLoS Pathog. 2015;11: e1004935. pmid:26203656
  14. 14. Kannan G, Pletnikov M V. Toxoplasma gondii and cognitive deficits in schizophrenia: An animal model perspective. Schizophr Bull. 2012;38: 1155–1161. pmid:22941742
  15. 15. Ginhoux F, Lim S, Hoeffel G, Low D, Huber T. Origin and differentiation of microglia. Front Cell Neurosci. 2013;7: 45. pmid:23616747
  16. 16. Prinz M, Priller J. Microglia and brain macrophages in the molecular age: from origin to neuropsychiatric disease. Nat Rev Neurosci. 2014;15: 300–312. pmid:24713688
  17. 17. Kandel ER. Principles of neural science. 5th ed. 2013.
  18. 18. Azevedo FAC, Carvalho LRB, Grinberg LT, Farfel JM, Ferretti REL, Leite REP, et al. Equal numbers of neuronal and nonneuronal cells make the human brain an isometrically scaled-up primate brain. J Comp Neurol. 2009;513: 532–41. pmid:19226510
  19. 19. Herculano-Houzel S, Mota B, Lent R. Cellular scaling rules for rodent brains. Proc Natl Acad Sci. 2006;103: 12138–12143. pmid:16880386
  20. 20. Herculano-Houzel S. The remarkable, yet not extraordinary, human brain as a scaled-up primate brain and its associated cost. Proc Natl Acad Sci. 2012;109: 10661–10668. pmid:22723358
  21. 21. McCarthy GF, Leblond CP. Radioautographic evidence for slow astrocyte turnover and modest oligodendrocyte production in the corpus callosum of adult mice infused with 3H-thymidine. J Comp Neurol. 1988;271: 589–603. pmid:3385018
  22. 22. Horner PJ, Palmer TD. New roles for astrocytes: The nightlife of an “astrocyte”. La vida loca! Trends Neurosci. 2003;26: 597–603. pmid:14585599
  23. 23. Marshall CAG, Suzuki SO, Goldman JE. Gliogenic and neurogenic progenitors of the subventricular zone: who are they, where did they come from, and where are they going? Glia. 2003;43: 52–61. pmid:12761867
  24. 24. Ming G-L, Song H. Adult neurogenesis in the mammalian brain: significant answers and significant questions. Neuron. 2011;70: 687–702. pmid:21609825
  25. 25. Abbott NJ, Rönnbäck L, Hansson E. Astrocyte–endothelial interactions at the blood–brain barrier. Nat Rev Neurosci. 2006;7: 41–53. pmid:16371949
  26. 26. Hickey WF, Kimura H. Graft-vs.-host disease elicits expression of class I and class II histocompatibility antigens and the presence of scattered T lymphocytes in rat central nervous system. Proc Natl Acad Sci U S A. 1987;84: 2082–6. Available: http://www.ncbi.nlm.nih.gov/pubmed/3550805 pmid:3550805
  27. 27. Boerman RH, Peters ACB, Bloem BR, Raap AK, van der Ploeg M. Spread of herpes simplex virus to the cerebrospinal fluid and the meninges in experimental mouse encephalitis. Acta Neuropathol. 1992;83: 300–307. pmid:1313632
  28. 28. Verma S, Lo Y, Chapagain M, Lum S, Kumar M, Gurjav U, et al. West Nile virus infection modulates human brain microvascular endothelial cells tight junction proteins and cell adhesion molecules: Transmigration across the in vitro blood-brain barrier. Virology. 2009;385: 425–433. pmid:19135695
  29. 29. Chai Q, He WQ, Zhou M, Lu H, Fu ZF. Enhancement of blood-brain barrier permeability and reduction of tight junction protein expression are modulated by chemokines/cytokines induced by rabies virus infection. J Virol. 2014;88: 4698–710. pmid:24522913
  30. 30. Dobrowolski JM, Sibley LD. Toxoplasma Invasion of Mammalian Cells Is Powered by the Actin Cytoskeleton of the Parasite. Cell. 1996. pp. 933–939. pmid:8601316
  31. 31. Tardieux I, Ménard R. Migration of Apicomplexa across biological barriers: The Toxoplasma and Plasmodium rides. Traffic. 2008;9: 627–635. pmid:18194412
  32. 32. Daneman R, Rescigno M. The Gut Immune Barrier and the Blood-Brain Barrier: Are They So Different? Immunity. 2009;31: 722–735. pmid:19836264
  33. 33. Daneman R, Prat A. The blood-brain barrier. Cold Spring Harb Perspect Biol. Cold Spring Harbor Laboratory Press; 2015;7: a020412. pmid:25561720
  34. 34. Barragan A, Sibley LD. Transepithelial migration of Toxoplasma gondii is linked to parasite motility and virulence. J Exp Med. 2002;195: 1625–33. Available: http://www.ncbi.nlm.nih.gov/pubmed/12070289 pmid:12070289
  35. 35. Harker KS, Jivan E, McWhorter FY, Liu WF, Lodoen MB. Shear forces enhance Toxoplasma gondii tachyzoite motility on vascular endothelium. MBio. 2014;5: e01111–13. pmid:24692639
  36. 36. Dubey JP. Bradyzoite-Induced Murine Toxoplasmosis: Stage Conversion, Pathogenesis, and Tissue Cyst Formation in Mice Fed Bradvzoites of Different Strains of Toxoplasma gondii. J Euk Microbiol. 1997;44: 592–602. pmid:9435131
  37. 37. Gregg B, Taylor BC, John B, Tait-Wojno ED, Girgis NM, Miller N, et al. Replication and distribution of Toxoplasma gondii in the small intestine after oral infection with tissue cysts. Infect Immun. 2013;81: 1635–43. pmid:23460516
  38. 38. Konradt C, Ueno N, Christian DA, Delong JH, Pritchard GH, Herz J, et al. Endothelial cells are a replicative niche for entry of Toxoplasma gondii to the central nervous system. Nat Microbiol. 2016;1: 16001. pmid:27572166
  39. 39. Conley FK, Jenkins KA. Immunohistological study of the anatomic relationship of toxoplasma antigens to the inflammatory response in the brains of mice chronically infected with Toxoplasma gondii. Infect Immun. 1981;31: 1184–1192. pmid:7228401
  40. 40. 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: 1611–1623. pmid:16984416
  41. 41. Lachenmaier SM, Deli MA, Meissner M, Liesenfeld O. Intracellular transport of Toxoplasma gondii through the blood-brain barrier. J Neuroimmunol. 2011;232: 119–30. pmid:21106256
  42. 42. Ueno N, Harker KS, Clarke E V., McWhorter FY, Liu WF, Tenner AJ, et al. Real-time imaging of Toxoplasma-infected human monocytes under fluidic shear stress reveals rapid translocation of intracellular parasites across endothelial barriers. Cell Microbiol. 2014;16: 580–595. pmid:24245749
  43. 43. 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: 309–316. pmid:16051744
  44. 44. Lang W, Miklossy J, Deruaz JP, Pizzolato GP, Probst A, Schaffner T, et al. Neuropathology of the acquired immune deficiency syndrome (AIDS): a report of 135 consecutive autopsy cases from Switzerland. Acta Neuropathol. 1989;77: 379–90. Available: http://www.ncbi.nlm.nih.gov/pubmed/2540610 pmid:2540610
  45. 45. Strittmatter C, Lang W, Wiestler OD, Kleihues P. The changing pattern of human immunodeficiency virus-associated cerebral toxoplasmosis: a study of 46 postmortem cases. Acta Neuropathol. 1992;83: 475–481. pmid:1621505
  46. 46. Laing RBS, Flegg PJ, Brettle RP, Leen CLS, Burns SM. Clinical features, outcome and survival from cerebral toxoplasmosis in Edinburgh AIDS patients. Int J STD AIDS. 1996;7: 258–264. pmid:8876356
  47. 47. Berenreiterová M, Flegr J, Kuběna AA, Němec P. The distribution of Toxoplasma gondii cysts in the brain of a mouse with latent toxoplasmosis: Implications for the behavioral manipulation hypothesis. PLoS ONE. 2011;6. pmid:22194951
  48. 48. Dubey JP, Ferreira LR, Alsaad M, Verma SK, Alves DA, Holland GN, et al. Experimental Toxoplasmosis in Rats Induced Orally with Eleven Strains of Toxoplasma gondii of Seven Genotypes: Tissue Tropism, Tissue Cyst Size, Neural Lesions, Tissue Cyst Rupture without Reactivation, and Ocular Lesions. PLoS ONE. 2016;11: e0156255. pmid:27228262
  49. 49. Evans AK, Strassmann PS, Lee I-P, Sapolsky RM. Patterns of Toxoplasma gondii cyst distribution in the forebrain associate with individual variation in predator odor avoidance and anxiety-related behavior in male Long–Evans rats. Brain Behav Immun. 2014;37: 122–133. pmid:24269877
  50. 50. Alvarado-Esquivel C, Sánchez-Anguiano LF, Mendoza-Larios A, Hernández-Tinoco J, Pérez-Ochoa JF, Antuna-Salcido EI, et al. Prevalence of Toxoplasma gondii infection in brain and heart by immunohistochemistry in a hospital-based autopsy series in Durango, Mexico. Eur J Microbiol Immunol. 2015;5: 143–149. pmid:26185682
  51. 51. Ghatak NR, Zimmerman HM. Fine structure of Toxoplasma in the human brain. Arch Pathol. 1973;95: 276–83. Available: http://www.ncbi.nlm.nih.gov/pubmed/4348725 pmid:4348725
  52. 52. Pavesio CE, Chiappino ML, Setzer PY, Nichols BA. Toxoplasma gondii: differentiation and death of bradyzoites. Parasitol Res. 1992;78: 1–9. Available: http://www.ncbi.nlm.nih.gov/pubmed/1584739 pmid:1584739
  53. 53. Wanko T, Jacobs L, Gavin MA. Electron Microscope Study of Toxoplasma Cysts in Mouse Brain. J Protozool. 1962;9: 235–242. pmid:14004896
  54. 54. Sims TA, Hay J, Talbot IC. An electron microscope and immunohistochemical study of the intracellular location of Toxoplasma tissue cysts within the brains of mice with congenital toxoplasmosis. Br J Exp Pathol. 1989;70: 317–25. Available: http://www.ncbi.nlm.nih.gov/pubmed/2504268 pmid:2504268
  55. 55. Ferguson DJP, Hutchison WM. The host-parasite relationship of Toxoplasma gondiiin the brains of chronically infected mice. Virchows Arch A Pathol Anat Histopathol. 1987;411.
  56. 56. Ferguson DJP, 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. pmid:3422976
  57. 57. Ferguson DJ, Huskinson-Mark J, Araujo FG, Remington JS. A morphological study of chronic cerebral toxoplasmosis in mice: comparison of four different strains of Toxoplasma gondii. Parasitol Res. 1994;80: 493–501. pmid:7808999
  58. 58. 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: pmid:21625284
  59. 59. Werk R. How does toxoplasma gondii enter host cells? Rev Infect Dis. 1985;7: 449–457. pmid:3898305
  60. 60. Blader IJ, Coleman BI, Chen C-T, Gubbels M-J. Lytic Cycle of Toxoplasma gondii: 15 Years Later. Annu Rev Microbiol. Annual Reviews; 2015;69: 463–85. pmid:26332089
  61. 61. 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: 637–41. Available: http://www.ncbi.nlm.nih.gov/pubmed/9272550 pmid:9272550
  62. 62. Halonen SK, Lyman WD, Chiu FC. Growth and development of Toxoplasma gondii in human neurons and astrocytes. J Neuropathol Exp Neurol. 1996;55: 1150–6. Available: http://www.ncbi.nlm.nih.gov/pubmed/8939198 pmid:8939198
  63. 63. 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: 516–8. Available: http://www.ncbi.nlm.nih.gov/pubmed/7844409 pmid:7844409
  64. 64. Halonen SK, Taylor GA, Weiss LM. Gamma interferon-induced inhibition of Toxoplasma gondii in astrocytes is mediated by IGTP. Infect Immun. 2001;69: 5573–5576. pmid:11500431
  65. 65. Martens S, Parvanova I, Zerrahn J, Griffiths G, Schell G, Reichmann G, et al. Disruption of Toxoplasma gondii parasitophorous vacuoles by the mouse p47-resistance GTPases. PLoS Pathog. 2005;1: 0187–0201. pmid:16304607
  66. 66. Degrandi D, Kravets E, Konermann C, Beuter-Gunia C, Klümpers V, Lahme S, et al. Murine guanylate binding protein 2 (mGBP2) controls Toxoplasma gondii replication. Proc Natl Acad Sci U S A. 2013;110: 294–9. pmid:23248289
  67. 67. Schlüter D, Deckert M, Hof H, Frei K. Toxoplasma gondii infection of neurons induces neuronal cytokine and chemokine production, but gamma interferon- and tumor necrosis factor-stimulated neurons fail to inhibit the invasion and growth of T. gondii. Infect Immun. 2001;69: 7889–7893. pmid:11705972
  68. 68. Rall GF, Mucke L, Oldstone. Consequences of cytotoxic T lymphocyte interaction with major histocompatibility complex class I-expressing neurons in vivo. J Exp Med. 1995;182: 1201–1212. pmid:7595191
  69. 69. Koshy AA, Fouts AE, Lodoen MB, Alkan O, Blau HM, Boothroyd JC. Toxoplasma secreting Cre recombinase for analysis of host-parasite interactions. Nat Methods. 2010;7: 307–9. pmid:20208532
  70. 70. Koshy AA, Dietrich HK, Christian DA, Melehani JH, Shastri AJ, Hunter CA, et al. Toxoplasma Co-opts Host Cells It Does Not Invade. PLoS Pathog. 2012;8: e1002825. pmid:22910631
  71. 71. Cabral CM, Tuladhar S, Dietrich HK, Nguyen E, MacDonald WR, Trivedi T, et al. Neurons are the Primary Target Cell for the Brain-Tropic Intracellular Parasite Toxoplasma gondii. PLoS Pathog. 2016;12: e1005447. pmid:26895155
  72. 72. Beninger RJ. The role of dopamine in locomotor activity and learning. Brain Research Reviews. Elsevier; 1983. pp. 173–196.
  73. 73. Stibbs HH. Changes in brain concentrations of catecholamines and indoleamines in Toxoplasma gondii infected mice. Ann Trop Med Parasitol. 1985;79: 153–7. Available: http://www.ncbi.nlm.nih.gov/pubmed/2420295 pmid:2420295
  74. 74. Ihara F, Nishimura M, Muroi Y, Mahmoud ME, Yokoyama N, Nagamune K, et al. Toxoplasma gondii Infection in Mice Impairs Long-Term Fear Memory Consolidation through Dysfunction of the Cortex and Amygdala. Infect Immun. 2016;84: 2861–70. pmid:27456832
  75. 75. 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: 1039–47. pmid:25547791
  76. 76. Brown CR, McLeod R. Class I MHC genes and CD8+ T cells determine cyst number in Toxoplasma gondii infection. J Immunol. 1990;145: 3438–41. Available: http://www.ncbi.nlm.nih.gov/pubmed/2121825 pmid:2121825
  77. 77. Suzuki Y, Orellana MA, Wong SY, Conley FK, Remington JS. Susceptibility to chronic infection with Toxoplasma gondii does not correlate with susceptibility to acute infection in mice. Infect Immun. 1993;61: 2284–8. Available: http://www.ncbi.nlm.nih.gov/pubmed/8500870 pmid:8500870
  78. 78. Pacheco R, Contreras F, Zouali M. The dopaminergic system in autoimmune diseases. Front Immunol. Frontiers; 2014;5: 117. pmid:24711809
  79. 79. David CN, Frias ES, Szu JI, Vieira PA, Hubbard JA, Lovelace J, et al. GLT-1-Dependent Disruption of CNS Glutamate Homeostasis and Neuronal Function by the Protozoan Parasite Toxoplasma gondii. PLoS Pathog. 2016;12: e1005643. pmid:27281462
  80. 80. Brooks JM, Carrillo GL, Su J, Lindsay DS, Fox MA, Blader IJ. Toxoplasma gondii infections alter GABAergic synapses and signaling in the central nervous system. MBio. 2015;6: 1–9. pmid:26507232
  81. 81. Parlog A, Harsan L-A, Zagrebelsky M, Weller M, von Elverfeldt D, Mawrin C, et al. Chronic murine toxoplasmosis is defined by subtle changes in neuronal connectivity. Dis Model Mech. 2014;7: 459–69. pmid:24524910
  82. 82. Luft BJ, Remington JS. Toxoplasmic encephalitis in AIDS. Clin Infect Dis. 1992;15: 211–222. pmid:1520757
  83. 83. Neu N, Duchon J, Zachariah P. TORCH Infections. Clin Perinatol. 2015;42: 77–103. pmid:25677998