The Development of Unconventional Extrathymic Activated CD4<sup>+</sup>CD8<sup>+</sup> T Cells in Chagas Disease

Morrot, Alexandre

doi:https://doi.org/10.5402/2013/801975

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Volume 2013 | Article ID 801975 | https://doi.org/10.5402/2013/801975

The Development of Unconventional Extrathymic Activated CD4⁺CD8⁺ T Cells in Chagas Disease

Alexandre Morrot¹

Academic Editor: Y.-H. Gan, H. Hisaeda, Y. Lai

Received10 Apr 2013

Accepted02 Jun 2013

Published19 Jun 2013

Abstract

The numbers of extrathymic CD4⁺CD8⁺ double-positive (DP) T cells are augmented in various pathophysiological conditions, such as infectious diseases caused by intracellular pathogens, organs subjected to autoimmune attack and malignant tumors. The roles performed by extrathymic DP T cells are not clear, and it is not known how they are distributed in the body. In animals they have been considered memory cells involved in adaptive immune responses against virus infections or participating in pathological responses. In experimental Trypanosoma cruzi infections, there is a severe thymic atrophy and this results in the release of activated DP T cells to peripheral organs. In severe cardiac forms of human chronic Chagas disease activated HLA-DR⁺ DP T cells are present in the blood. In investigating the basis of premature thymocyte release during chagasic thymic atrophy we found that the parasite trans-sialidase (TS) altered intrathymic thymocyte maturation and was associated with increased numbers of recent T cells in peripheral lymphoid organs. In what follows we propose to describe what is known about the origin of the extrathymic DP T cells in human Chagas disease and animal models of the disease.

1. T Cell Development in the Thymus

The formation of mature lineage-committed T cells requires the specialized environment of the thymus, whereas B cells generally develop while still in the bone marrow [1, 2]. The thymus, located in the thoracic cavity, comprises an outer cortex and an inner medulla. T cells are continuously generated in the thymus and released to the periphery. The complex developmental process responsible for generating T cells within the thymus depends on signals from stromal cells that direct the maturation, expansion, and selection of T cell precursors [3, 4]. The stromal cells thus provide a thymic microenvironment appropriate for T cell development [5, 6]. Thymus function is defective in a variety of clinical situations. Such a defective thymic microenvironment can prevent the development of the adaptive immune system, cause life-threatening immunodeficiencies and autoimmune reactions, and inhibit the process of immunological surveillance [7–12], which is thought to be responsible for recognizing and removing malignant cells as they arise.

Although the thymus does not contain self-renewing hematopoietic precursor cells, progenitor cells are constantly recruited from the blood. The recruited cells have a triple-negative CD3⁻CD4⁻CD8⁻ phenotype and penetrate the vascularized microenvironment of the thymus at the boundary between the cortex and medulla. Once in the thymus, they designated early T lineage progenitors [4, 5, 13], and engagement of their Notch1 receptors with the delta-4-like ligands on cortical thymic epithelial cells induces proliferation of the cells and results in their commitment to the T cell lineage [14–17]. After an ordered sequence of developmental steps (Figure 1), they eventually express both the CD4 and CD8 antigens and intact antigen-specific T cell receptors [18, 19]. These cells referred, to as DP thymocytes, are submitted to two successive selection steps that lead to the elimination of cells whose randomly assigned T cell receptor (TCR) specificities are undesirable [18].

Figure 1

Schema of T cell development in the thymus and of the proposed role of extrathymic DP T cells. Left figure: progenitors of CD4⁻CD8⁻ (DN) T cells enter the thymus from bone marrow via the large blood vessels at the border between cortex and medulla and move towards the capsule of the organ. Rearrangement at the T cell receptor (TCRβ): γ and δ gene loci commence, and strong TCR signaling favors development of the γ δ T cell lineage, while weak TCR signals lead to α β commitment [20]. Only cells that rearrange their T cell receptor β (TCRβ) chain proliferate extensively and give rise to CD4⁺CD8⁺ double-positive (DP) thymocytes. These cells occupy much of the cortex and move around slowly while their TCRα chains are rearranged. During this phase, weak interaction in the cortex between the newly formed T cell receptors (TCRs) on cortical thymocytes and self-peptides bound to major histocompatibility complex (MHC) proteins on cortical thymic epithelial cells (TECs) leads to survival and maturation of the thymocytes. Thereafter only T cells bearing TCR with a low avidity for any of the self-peptide-MHC class I or II molecules presented on thymic epithelial cells (TECs) are subjected to selection, becoming CD8⁺ or CD4⁺ single-positive (SP) T cells [18]. Other cells with high avidity for any of the self-peptide-MHC class I or II molecules die by apoptosis via negative selection in the central medullary region of the thymus or are alternatively differentiated into regulatory T cells [21, 22]. Some DP thymocytes express CD1a, which binds self-glycolipids, and other DP thymocytes that are able to interact with these cells enter the NKT lineage. Most NKT cells that exit the thymus become mature in the periphery. Mature cells within the thymus can either emigrate from the thymus or remain for a period in that organ [23]. Right figure: in a pathological setting, the severe thymic atrophy leads to release of activated DP T cells to the periphery. In Chagas disease, it has been well demonstrated that the T. cruzi trans-sialidase affects the T cell intrathymic development resulting in an increase in the number of CD4⁺8⁺ double-positive recent thymic emigrants to the secondary lymphoid tissues. In the periphery, the coexpression of CD4 and CD8 molecules on the DP T cells released from the thymus may increase the affinity of the TCR during priming and so may reduce the threshold for costimulatory signals at T cell-dendritic cell immunological synapses [24, 25]. Since the DP T cells express TCR complex with CD4 and CD8 molecules, they would be able to recognize both peptide-class I and II MHC receptors at the same time, and these cells could rapidly eliminate APCs before other single-positive T cells could even be activated [26, 27]. Thus the DP T cells might have an immunodominant effect that could increase adaptive immune responses. The prematurely released activated DP thymocyte cells might also mature into functionally competent SP cells able to recognize pathogen antigens. Alternatively, they could express forbidden α βTCR with self-antigen specificities and could thus induce autoimmune responses.

Many of the TCRs on DP thymocytes generated by TCR gene rearrangement are unable to bind self major histocompatibility complex (MHC) molecules, and such cells will die within 3-4 days. Therefore an initial selection process in the thymus will only retain those T cells that bear a αβ TCR that is able to still recognise self MHC. This process is known as positive selection [4, 28] and occurs when the T cells bind cortical epithelial cells expressing Class I or Class II MHC plus self-peptides with a sufficiently high affinity to receive a survival signal [29, 30]. The remaining cells die within a few days from “death by neglect” [31, 32]. The surviving thymocytes undergo in the medulla of the thymus a second, negative selection process, which leads programmed cell death of thymocytes whose TCRs have an above-threshold affinity for self-peptide/MHC complexes. In that way thymocytes capable of generating elicit autoimmune responses are removed, and only T cells able to recognize nonself-antigens in the context of self-MHC molecules are released into the periphery [33, 34].

During T cell development, thymocytes that produce a TCR restricted to class I MHC molecules adopt a CD4⁻CD8⁺ phenotype while those with an MHC class II-restricted TCR specificity turn into CD4⁺CD8⁻ cells [14, 35, 36]. After a time required for maturation, these naïve single-positive (SP) T cells (i.e., CD4⁻CD8⁺ and CD4⁺CD8⁻ thymocytes) exit the thymic medulla and migrate to the periphery in a process that requires signaling via sphingosine 1-phosphate receptor type 1 [37, 38]. In this way, thymocyte development requires signals emanating from a number of different types of stromal cells, particularly thymic epithelial cells (TEC); these influence their entry, division, maturation, and survival [39, 40]. The TECs form a three-dimensional network held together by contacts via dendrite-like processes, and the TEC elements occupying the thymic cortex and medulla are different. The processes involved in TEC multiplication and maintenance have been extensively studied since the functioning of these cells is affected by aging and by anticancer treatments [41–44].

2. Infectious Diseases Can Result in Thymic Atrophy

Several infectious pathogens for example, viruses (HIV, HCV, rabies virus), parasites (Trypanosoma cruzi, Plasmodium berghei, Schistosoma mansoni, and Trichinella spiralis), and fungi (Paracoccidioides brasiliensis and Histoplasma capsulatum) can cause atrophy of the thymus. It is not completely clear how this atrophy occurs and the mechanism may vary [45–53]. However, there are common histological features, such as lower numbers of cortical thymocytes and breakdown of the clear-cut distinction between cortex and medulla. In some cases atrophy may be transient, for example, in experimental infection with Histoplasma capsulatum and Toxoplasma gondii [53–55].

A number of not mutually exclusive factors may be involved in infection-induced thymic atrophy: fewer precursor cells may enter the thymus, these cells may have a lower ability to divide, they may die more frequently, and the rate at which they leave the thymus may be higher. The proliferative ability of thymocytes from mice acutely infected with T. cruzi was observed to be reduced due to lower production of interleukin-2 (IL-2), and this in turn was linked to elevated levels of IL-10 and interferon-γ (IFN-γ) [56]. There is also evidence that T. spiralis infection can affect the proportions of different thymocyte subsets, which is seen as a reduced responsiveness of the thymocytes to the T cell mitogen, concanavalin A. On the other hand, the proliferative response of thymocytes from S. mansoni-infected mice, to concanavalin A is unaffected [57]. It appears therefore that some, but not all, parasites bring about a reduction in the proliferative ability of thymocytes that could account at least in part for the thymic atrophy.

In most infectious diseases causing thymic atrophy, the major biological event associated with thymocyte loss is cell death by apoptosis. This is seen, for example, in experimental models of Trypanosoma cruzi [58, 59] and Plasmodium berghei infection [51, 60]. Most of the cells that die are CD4⁺CD8⁺ thymocytes, but other subtypes such as double-negative (DN) T cells and SP cells are also reduced in number [53]. Glucocorticoid hormones appear to be responsible for the atrophy and thymocyte death during parasitic infections. In acute infections serum glucocorticoid levels rise and thymocyte caspases-8 and -9 are activated [55]. Higher levels of serum glucocorticoids have also been found in experimentally induced malaria, American trypanosomiases or Chagas disease, African trypanosomiases or sleeping sickness, toxoplasmosis, leishmaniasis, and schistosomiasis. Both tumor necrosis factor (TNF-α) and glucocorticoid serum levels rise in acute experimental T. cruzi infections and are associated with thymic atrophy and thymocyte depletion [53, 55, 61–64]. However additional mechanisms have been implicated, at least, in T. cruzi infections. Some of the factors that may be responsible for the apoptotic cell death are the T. cruzi-derived trans-sialidase [65], host-derived galectin-3 [66–69], and androgens and extracellular ATP [70].

3. Premature Release of Immature Thymocytes Bypasses Negative Selection During Trypanosoma cruzi-Induced Thymic Atrophy

In both murine and human models it has been shown that infection with the intracellular protozoan Trypanosoma cruzi results in thymic atrophy and release of undifferentiated DP T cells to periphery [61, 71, 72]. These events appear to be linked to a particular pathogen-host relationship formed during infection. In T. cruzi infections, the inflammatory syndrome induced by TNF-α in the acute phase of infection activates the hypothalamus-pituitary-adrenal (HPA) axis leading to release of corticosterone, and this appears to be related to the changes in both lymphoid and nonlymphoid thymus compartments and to disease outcome [59, 73, 74]. However, other host-derived molecules play a part in the thymocyte depletion. Thymic atrophy, for example, is not seen in T. cruzi-infected galectin-3 knockout mice [55, 66]. On the other hand, parasite-derived factors have also been shown to affect the thymic homeostasis. This is the case for the parasite trans-sialidase [75–77], which is implicated in the death of intrathymic T cells [78]. Moreover, the dramatic changes in the thymic microenvironment occurring after acute infection are also associated with enhanced expression of extracellular matrix ligands and receptors, which correlates with the fibronectin-driven migration of DP thymocytes out of the thymus and the abnormal increase of immature DP T cells in lymph nodes [72].

Until recently it was not clear whether intrathymic negative selection was affected by the alterations in the thymic microenvironment caused by T. cruzi infection. Clonal deletion normally appears to occur late in thymocyte development at the transition from DP to SP cells and happens at the junction between cortex and medulla [79]. In this connection, thymic metallophilic macrophages present precisely at this boundary have been shown to be involved in thymocyte maturation [80]. They differ in a number of ways from cortical macrophages and from medullary interdigitating cells and, in particular, contain large endocytic compartments devoted to the processing and presentation of antigens by MHC class II molecules [81]. The topological distribution of these metallophilic macrophages is altered by infection; their number increases, and while some remain in the corticomedullary region, many more are spread throughout the cortex. The thymic localization of these cells may indicate that clonal selection switches from the corticomedullary junction to the cortex [82].

It is generally accepted that T cell development is controlled by interactions between TEC and thymocytes [83, 84]. Thus, when the thymic architecture is disturbed, the pattern of expression of autoantigens by TECs is altered, as well as the functioning of the thymus [85, 86]. Atrophy of the medulla and reduced levels of Aire and tissue-restricted self-antigens (TRA) transcripts are seen in mouse models deficient in a number of genes of the NFκB pathway, such as TRAF6, NIK, RelB, or p52, suggesting that this pathway is important for development of the thymic medulla [87, 88]. We measured the expression of Aire and of highly selective tissue-restricted antigens [89] by real-time PCR in whole thymuses of T. cruzi-infected mice and detected levels that were quite similar to control levels. These findings suggest that the expression of peripheral antigens in the infected thymuses is sufficient to modulate the induction of tolerance by negative selection [82].

Later in the course of infection, thymic atrophy becomes apparent and there is a rise in the number of apoptotic intrathymic DP T cells [90]. Although this increase may be due to the changes observed in the organ, we found that expression of Bim, a proapoptotic factor essential for negative selection [78, 91], was maintained while the number of DPs declined [82]. Furthermore, using the OTII TCR transgenic system which recognizes a CD4⁺ epitope from the chicken ovalbumin (OVA) protein, we observed that apoptosis of TCR-stimulated semimature thymocytes was provoked when we gave the cognate OVA peptide to acutely infected mice undergoing thymic atrophy. Taken together, these data show that DP T cells can be negatively selected during infection-induced thymic atrophy, suggesting that negative selection operates normally. This is consistent with the finding that mature single-positive CD4⁺ or CD8⁺ T cells within the thymus do not harbor forbidden TCR genes, unlike their DP counterparts [82].

4. The Trypanosoma cruzi trans-Sialidase Promotes the Escape of Immature Double-Positive CD4⁺CD8⁺ Thymocytes in Chagas Disease

T. cruzi cannot synthesize sialic acid de novo, and the trans-sialidase allows the parasite to scavenge α-(2,3)-linked sialyl residues from exogenous glycoconjugates and to transfer them to acceptors resembling mucin that covers the surface of the parasite [75, 77, 92, 93]. When the sialylation profile of cell surface receptors is altered by the TS this can induce apoptosis of cells of the immune system in vivo, including those in the thymus [94, 95], and this apoptosis is associated with cell death in the thymic nurse cell complex [96, 97]. Several mouse thymocyte cell surface proteins that accept sialyl residues in TS-catalyzed reactions have been identified by mass spectrometry [97]. Moreover we have observed that the TS, is able to activate the c-Jun N-terminal kinases (JNK) MAPK pathway in thymocytes [98]. JNK controls the migration of a variety of cell types and cell adhesion and migration depend on cytoskeletal structures involving the actin network and actin-binding proteins, which are known to be phosphorylated by JNK [63, 99, 100]. In fact, we have shown that the cell-signaling processes involving the TS mobilize actin filaments [98], and others have shown that these signaling processes depend on interactions between TEC and thymocytes, which are required for migration of thymocytes during intrathymic development [101].

We have recently tested whether the trans-sialidase can itself alter the migratory responses of thymocytes, which could account for the release of DP T cells from the thymus during infection. We found that the T. cruzi TS increased the adhesion of thymocytes to TEC [98]. Interestingly, in order to migrate, thymocytes must adhere to their microenvironment and then dissociate from it so that molecules with adhesive and deadhesive properties may affect their migration [59, 102]. In fact, we found that treatment of thymocytes with T. cruzi TS in vitro stimulated their migration towards fibronectin, indicating a role for this enzyme in the migration of these cells [98]. The migration of thymocytes is a key event in their intrathymic differentiation [59, 103].

Our findings indicated that the migratory properties of intrathymic thymocytes are modulated by the parasite trans-sialidase (TS). Several steps involve migration, including entry of the precursor cells into the thymus, movement of immature thymocytes within the cortex and from there to the medulla, and, finally, exit from the thymus [59, 72]. All these events depend upon initial adhesion to the extracellular matrix (ECM) elements followed by dissociation and binding to TEC or microenvironmental components [67, 72, 104]. Interestingly, we found that trans-sialidase treatment of CD4⁺CD8⁺ cells within the thymus affected their maturation, resulting in a premature thymic emigration and increase in the number of recent DP thymic emigrants in the periphery, which ultimately account for the thymic atrophy process. In humans, we detected a possible correlation between the presence of the trans-sialidase and the number of peripheral blood DP T cells in chronic chagasic patients with the cardiac clinical forms of Chagas disease [98]. The presence of peripheral activated DP T cells with potentially autoreactive TCR may contribute to the immunopathological events found in this disease, and our data support the potential utility of chemotherapy against the parasite TS in Chagas disease.

5. Peripheral Differentiated CD4⁺CD8⁺ T Cells with Activation Phenotype Correlate with Severity of Chagas Disease

It is clear that DP T cells are released prematurely from infected thymuses into the periphery, where their maturation into functionally competent single-positive cells continues [105] even though intrathymic checkpoints remain active in the acute phase of murine Chagas disease [53, 82]. This unconventional and rare (<5%) lymphocyte population is also seen in secondary lymph nodes, as we demonstrated in an experimental model of Chagas disease in which a DP T cell subset increased up to 16-fold in subcutaneous lymph nodes [82, 106]. Release during both acute and chronic T. cruzi infections may be promoted by the high level of CD62L (L-selectin), which controls lymphocyte homing to lymph nodes [107]. We have examined whether DP T cells exhibit activated properties similar to effector/memory single positive T cells [82].

Interestingly, we found that these cells do indeed develop an activated phenotype, with high levels of the activation markers CD44 and CD69, which are tightly linked to the differentiation status of T cells [82]. Moreover, when we purified (>98%) DP cell obtained by cell sorting from infected mice they were found to produce high levels of IFN-γ mRNA [82]. Furthermore, we observed that these DP T cells had higher cytotoxic activity than naïve single-positive CD4⁺ or CD8⁺ T cells [108, 109], in agreement with previous observations on the extrathymic DP T cells in cynomolgus monkeys and chimpanzees infected with hepatitis C and SIV virus [110–113], as well as in patients infected with cytomegalovirus (HCMV) and human immunodeficiency virus (HIV) [114–119]. Most significantly, patients with the cardiac form of Chagas disease had increased numbers of peripheral blood HLA-DR⁺ DP T cells [82]. These findings suggest that this DP T cell subset is associated with the development of the cardiac form of the disease and probably represent activated cells [82, 106].

The studies summarized above suggest that the DP T cell subset plays a role in the inflammatory processes generated by parasite-driven immune responses. It may be that coexpression of CD4 and CD8 molecules on the T cell membrane increases the affinity of the TCR for its target cell. If so, the simultaneous triggering of CD4 and CD8 coreceptors by MHC-antigen complexes would lower the threshold for antigen signaling and reduce the need for costimulatory signals in order to achieve activation of the T cells [120, 121]. As a result, the DP T cells could be activated by low antigen concentrations and might be activated at the onset of infection when parasite-derived antigens are limiting. It could be therefore that these cells promote adaptive immune responses by secreting cytokines which could stimulate dendritic cells, so providing a link between innate and adaptive immune responses. On the other hand they might eliminate activated antigen presenting cells, since theoretically they recognize both class I and class II MHC complexes, and this would favor the infecting parasite (Figure 1).

In conclusion, despite the fact that the key intrathymic checkpoints required for negative selection are maintained during acute chagasic thymic atrophy, the peripheral DP T cells have an activated phenotype like that ascribed to activated and memory SP T cells: they produce IFN-γ, express CD44 and CD69, and have cytotoxic activity [82]. This is of special significance since fully activated peripheral DP T cells are seen in the cardiac form of chronic Chagas disease [82, 106]. The presence of these cells challenges current views concerning the T cell populations involved in adaptive immune responses during T. cruzi infections: it suggests that DP T cells participate from early on in the immune response against such infections. The in vivo role of these cells remains unclear, but the fact that they have potentially autoreactive TCR may influence the immunopathological events that occur in both murine and human Chagas disease. If it turns out that the level of DP T cells is correlated with the extent of myocardial damage in the cardiac form of the disease, these cells could act as clinical markers of disease progression and might contribute to the design of alternative treatments. These studies of the role of the DP T cells should provide fundamental insight into virus-host relationship established during T. cruzi infections.

6. Peripheral Activated CD4⁺CD8⁺ T Cells May Be a Common Feature in Infectious Diseases

Although most T cells are either CD4⁺ or CD8⁺, a few peripheral CD4⁺CD8⁺ T cells have been found in both humans and animals and higher numbers of such cells are seen in patients with acute and chronic viral infections [113, 114, 117, 118, 122–125]. Unlike immature thymic thymocytes, peripheral DP T cells have the properties of mature T cells including antigen-dependent cytokine production and cytolytic activity, and they express markers involved in immunological memory. In pigs CD4⁺CD8⁺ T cells (Th/memory) possess memory and T-helper and cytolytic properties, and they secrete IFN-γ [114, 125–127]. This T cell subset was associated with protection in pigs vaccinated against pseudorabies virus [126, 127]. In intranasally vaccinated and virulent porcine reproductive and respiratory syndrome virus, an arterivirus challenged pigs, there is an enhanced frequency of CD4⁺CD8⁺ T cells [128]. Such cells appear to be the progeny of normal antigen-stimulated single-positive T cells. Human DP T cells can be easily produced from SP T cells in vitro, and adoptive transfer experiments in animals indicate that at least some DP cells can arise from CD4⁺ SP precursors in vivo [129].

The expression of CD4 in DP T cells has a bearing on HIV-1 disease because such cells should be infectable with HIV-1. Indeed, studies indicate that DP T cells isolated from HIV-1-infected patients have been shown to contain HIV-1 provirus [130, 131]. Moreover intestinal DP T cells are at least susceptible to SIV/HIV-1-mediated depletion as conventional CD4 T cells; similarly, numbers of peripheral blood DP T cells are lower in SIV-infected macaques than in uninfected animals [132, 133]. On the other hand circulating levels of DP T cells were not found to be reduced in HIV-1-infected patients and have even been found to be increased in some patients [114, 116]. Interestingly, examination of a number of HIV-1 subjects receiving a therapeutic vaccine showed that the DP T cells were polyfunctional in that they produced substantial levels of cytokines and also had cytotoxic T lymphocyte (CTL) activity, and this behavior has also been observed in other antigen systems. Importantly, the polyfunctionality of the DP T cells in HIV-1-infected patients was correlated with lower virus loads and nonprogressive disease. However, the antigen-specific properties of DP T cells have not been studied in large numbers of unvaccinated HIV-1-infected individuals [108, 115, 117].

Most studies of HIV-1 pathology have concentrated on chronically infected patients mainly because few patients are identified soon after infection. Examination of macaque models and naturally infected humans has revealed a dramatic depletion of CD4 T cells in the gut mucosa early in infection [117, 134]. It is well established that the virus set point emerging soon after HIV-1 infection predicts later disease progression; hence study of the immune mechanisms in early infected patients, including the T cell polyfunctionality, should reveal important features related to virus set point and thus to disease progression [135, 136]. This may be the case for HCV infections, in which the intrahepatic and circulating DP T cells are populations of activated, central, and effector memory cells, with heterogeneous differentiation patterns [109, 113]. Differences in the measured proportions of circulating DP T cells could depend on the clinical phase of HCV infection or patient populations. Interestingly, it has also been shown that double-positive CD4⁺CD8⁺ T cells really exist in extrathymic sites in vivo, as these cells were detected in situ at sites of inflammation and viral replication and could be cloned from chimpanzee liver biopsies during HCV infections. These findings therefore suggest that CD4⁺CD8⁺ T cells contribute early to the immune response to virus infections [113].

There is no doubt that the presence of mature, functional CD4⁺CD8⁺ T cells in peripheral blood and tissues in the infectious diseases challenges our view of the T cell populations involved in adaptive immune responses [113]. Studies on HIV infection model have shown evidences that DP T cells have a memory phenotype and produce cytokines in response to viral antigens. Also it appears that DP T cells can be infected with HIV-1 both in vitro and in vivo [114]. The most significant findings of this study were that HIV-1-reactive cells could be identified within the two previously defined subpopulations of DP T cells based on their levels of CD4 and CD8 expression, and these sharing properties of both conventional CD4 and CD8 SP T cells and therefore could be considered polyfunctional. These HIV-1-specific IFN-γ-producing DP T cells were present at higher frequencies in chronically infected patients than in early infection, and a greater fraction expressed LAMP markers [114, 118, 119].

7. Concluding Remarks

DP T cells are released from the thymus along with other lymphocytes, and their number nonspecifically varies with upon infection, inflammation, and autoimmune diseases [55, 129, 137–139]. Such cells could be generated outside the thymus and/or result from de novo expression of CD8 or CD4 in single-positive CD4⁺/CD8⁺ T cells. An alternative possibility is that they arise specifically in the microenvironment of the thymus (which includes thymic epithelial cells, L-Ti-like cells, local macrophages, and DCs), under the influence of pathogens, and the consequences of infections and other physiological events causing thymic involution [53, 55, 129]. In fact the properties of extrathymic DP T cells indicate that they are closely related developmentally to conventional CD8⁺ and CD4⁺ T cells [82]. A key question is whether activation of the peripheral DP T cells during infections depends on interaction with the cognate peptide/MHC complex (pMHC) on antigen-presenting cells (APC). Activation of those cells, which expresses nonselected αβTCR, would avoid the need for stimulation of specific T cell receptors. However it is difficult to identify the basis of the nonspecific activation of the DP T cells and its possible relevance to cellular immune response to specific pathogens because of the low frequency of DP T cells. In this connection, a correlation between numbers of DP T cell subsets and the extent of inflammatory pathology could provide a clinical marker of disease progression in pathogen infections and may facilitate the design of novel therapeutic approaches to infectious diseases.

Acknowledgments

This work was supported by grants from Conselho Nacional de Desenvolvimento Científico e Tecnológico do Brasil (CNPq), Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES), and Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ).

References

A. Y. Lai and M. Kondo, “T and B lymphocyte differentiation from hematopoietic stem cell,” Seminars in Immunology, vol. 20, no. 4, pp. 207–212, 2008.
View at: Publisher Site | Google Scholar
T. Okada, S. Moriyama, and M. Kitano, “Differentiation of germinal center B cells and follicular helper T cells as viewed by tracking Bcl6 expression dynamics,” Immunological Reviews, vol. 247, no. 1, pp. 120–132, 2012.
View at: Publisher Site | Google Scholar
D. J. Izon, R. L. Boyd, G. A. Waanders, and A. Kelso, “The myelopoietic inducing potential of mouse thymic stromal cells,” Cellular Immunology, vol. 124, no. 2, pp. 264–277, 1989.
View at: Publisher Site | Google Scholar
W. van Ewijk, “The thymus: ‘interactive teaching during lymphopoiesis’,” Immunology Letters, vol. 138, no. 1, pp. 7–8, 2011.
View at: Publisher Site | Google Scholar
H. T. Petrie and J. C. Zúñiga-Pflücker, “Zoned out: functional mapping of stromal signaling microenvironments in the thymus,” Annual Review of Immunology, vol. 25, pp. 649–679, 2007.
View at: Publisher Site | Google Scholar
S. Zuklys, C. E. Mayer, S. Zhanybekova et al., “MicroRNAs control the maintenance of thymic epithelia and their competence for T lineage commitment and thymocyte selection,” Journal of Immunology, vol. 189, no. 8, pp. 3894–3904, 2012.
View at: Publisher Site | Google Scholar
K. W. Pyke, H. M. Dosch, M. M. Ipp, and E. W. Gelfand, “Demonstration of an intrathymic defect in a case of severe combined immunodeficiency disease,” The New England Journal of Medicine, vol. 293, no. 9, pp. 424–428, 1975.
View at: Google Scholar
K. S. Landreth, K. McCoy, and J. Clagett, “Deficiency in cells expressing terminal transferase in autoimmune (motheaten) mice,” Nature, vol. 290, no. 5805, pp. 409–411, 1981.
View at: Google Scholar
T. Katagiri, P. L. Cohen, and R. A. Eisenberg, “The lpr gene causes an intrinsic T cell abnormality that is required for hyperproliferation,” Journal of Experimental Medicine, vol. 167, no. 3, pp. 741–751, 1988.
View at: Google Scholar
H. M. Georgiou and T. E. Mandel, “Induction of insulitis in athymic (nude) mice: the effect of NOD thymus and pancreas transplantation,” Diabetes, vol. 44, no. 1, pp. 49–59, 1995.
View at: Google Scholar
H. S. Scott, M. Heino, P. Peterson et al., “Common mutations in autoimmune polyendocrinopathy-candidiasis-ectodermal dystrophy patients of different origins,” Molecular Endocrinology, vol. 12, no. 8, pp. 1112–1119, 1998.
View at: Google Scholar
A. L. Fletcher, A. Calder, M. N. Hince, R. L. Boyd, and A. P. Chidgey, “The contribution of thymic stromal abnormalities to autoimmune disease,” Critical Reviews in Immunology, vol. 31, no. 3, pp. 171–187, 2011.
View at: Google Scholar
C. Schmitt, S. Ktorza, S. Sarun, C. Blanc, R. de Jong, and P. Debre, “CD34-expressing human thymocyte precursors proliferate in response to interleukin-7 but have lost myeloid differentiation potential,” Blood, vol. 82, no. 12, pp. 3675–3685, 1993.
View at: Google Scholar
I. Visan, J. S. Yuan, J. B. Tan, K. Cretegny, and C. J. Guidos, “Regulation of intrathymic T-cell development by lunatic fringe- Notch1 interactions,” Immunological Reviews, vol. 209, pp. 76–94, 2006.
View at: Publisher Site | Google Scholar
I. Visan, J. S. Yuan, Y. Liu, P. Stanley, and C. J. Guidos, “Lunatic Fringe enhances competition for Delta-like Notch ligands but does not overcome defective pre-TCR signaling during thymocyte β-selection in vivo,” Journal of Immunology, vol. 185, no. 8, pp. 4609–4617, 2010.
View at: Publisher Site | Google Scholar
P. P. Roozen, M. H. Brugman, and F. J. Staal, “Differential requirements for Wnt and Notch signaling in hematopoietic versus thymic niches,” Annals of the New York Academy of Sciences, vol. 1266, pp. 78–93, 2012.
View at: Publisher Site | Google Scholar
I. van de Walle, E. Waegemans, J. de Medts et al., “Specific Notch receptor-ligand interactions control human TCR-αβ/γδ development by inducing differential Notch signal strength,” Journal of Experimental Medicine, vol. 210, no. 4, pp. 683–697, 2013.
View at: Publisher Site | Google Scholar
P. E. Love and A. Bhandoola, “Signal integration and crosstalk during thymocyte migration and emigration,” Nature Reviews Immunology, vol. 11, no. 7, pp. 469–477, 2011.
View at: Publisher Site | Google Scholar
V. C. Seitan, B. Hao, K. Tachibana-Konwalski et al., “A role for cohesin in T-cell-receptor rearrangement and thymocyte differentiation,” Nature, vol. 476, no. 7361, pp. 467–471, 2011.
View at: Publisher Site | Google Scholar
D. A. Witherden and W. L. Havran, “Molecular aspects of epithelial γδ T cell regulation,” Trends in Immunology, vol. 32, no. 6, pp. 265–271, 2011.
View at: Publisher Site | Google Scholar
L. Klein and K. Jovanovic, “Regulatory T cell lineage commitment in the thymus,” Seminars in Immunology, vol. 23, no. 6, pp. 401–409, 2011.
View at: Publisher Site | Google Scholar
J. M. Coquet, J. C. Ribot, N. Babala et al., “Epithelial and dendritic cells in the thymic medulla promote CD4⁺Foxp3⁺ regulatory T cell development via the CD27-CD70 pathway,” Journal of Experimental Medicine, vol. 210, no. 4, pp. 715–728, 2013.
View at: Publisher Site | Google Scholar
D. I. Godfrey and S. P. Berzins, “Control points in NKT-cell development,” Nature Reviews Immunology, vol. 7, no. 7, pp. 505–518, 2007.
View at: Publisher Site | Google Scholar
J. Banchereau, F. Briere, C. Caux et al., “Immunobiology of dendritic cells,” Annual Review of Immunology, vol. 18, pp. 767–811, 2000.
View at: Publisher Site | Google Scholar
N. Garbi and T. Kreutzberg, “Dendritic cells enhance the antigen sensitivity of T cells,” Frontiers in Immunology, vol. 3, article 389, 2012.
View at: Google Scholar
S. C. Knight, B. A. Askonas, and S. E. Macatonia, “Dendritic cells as targets for cytotoxic T lymphocytes,” Advances in Experimental Medicine and Biology, vol. 417, pp. 389–394, 1997.
View at: Google Scholar
R. A. Willis, J. W. Kappler, and P. C. Marrack, “CD8 T cell competition for dendritic cells in vivo is an early event in activation,” Proceedings of the National Academy of Sciences of the United States of America, vol. 103, no. 32, pp. 12063–12068, 2006.
View at: Publisher Site | Google Scholar
J. C. Howard and D. B. Wilson, “Specific positive selection of lymphocytes reactive to strong histocompatibility antigens,” Journal of Experimental Medicine, vol. 140, no. 3, pp. 660–672, 1974.
View at: Google Scholar
B. A. Kyewski, C. G. Fathman, and H. S. Kaplan, “Intrathymic presentation of circulating non-major histocompatibility complex antigens,” Nature, vol. 308, no. 5955, pp. 196–199, 1984.
View at: Google Scholar
S. M. Hedrick, “Positive selection in the thymus: an enigma wrapped in a mystery,” Journal of Immunology, vol. 188, no. 5, pp. 2043–2045, 2012.
View at: Publisher Site | Google Scholar
Y. Zilberman, E. Yefenof, S. Katzav, A. Dorogin, N. Rosenheimer-Goudsmid, and R. Guy, “Apoptosis of thymic lymphoma clones by thymic epithelial cells: a putative model for ‘death by neglect’,” Immunology Letters, vol. 67, no. 2, pp. 95–104, 1999.
View at: Publisher Site | Google Scholar
O. Cohen, E. Ish-Shalom, S. Kfir-Erenfeld, I. Herr, and E. Yefenof, “Nitric oxide and glucocorticoids synergize in inducing apoptosis of CD4⁺8⁺ thymocytes: implications for “death by neglect” and T-cell function,” International Immunology, vol. 24, no. 12, pp. 783–791, 2012.
View at: Publisher Site | Google Scholar
H. Hengartner, B. Odermat, R. Schneider et al., “Deletion of self-reactive T cells before entry into the thymus medulla,” Nature, vol. 336, no. 6197, pp. 388–390, 1988.
View at: Google Scholar
Y. Takahama, “T lymphocyte development and selection in the thymic microenvironments,” Nature Reviews Immunology, vol. 84, no. 3, pp. 177–1782, 2012.
View at: Google Scholar
C. J. Guidos, J. S. Danska, C. G. Fathman, and I. L. Weissman, “T cell receptor-mediated negative selection of autoreactive T lymphocyte precursors occurs after commitment to the CD4 or CD8 lineages,” Journal of Experimental Medicine, vol. 172, no. 3, pp. 835–845, 1990.
View at: Publisher Site | Google Scholar
E. Fiorini, I. Ferrero, E. Merck et al., “Cutting edge: thymic crosstalk regulates delta-like 4 expression on cortical epithelial cells,” Journal of Immunology, vol. 181, no. 12, pp. 8199–8203, 2008.
View at: Google Scholar
M. L. Allende, J. L. Dreier, S. Mandala, and R. L. Proia, “Expression of the sphingosine 1-phosphate receptor, S1P1, on T-cells controls thymic emigration,” The Journal of Biological Chemistry, vol. 279, no. 15, pp. 15396–15401, 2004.
View at: Publisher Site | Google Scholar
Y. Maeda, N. Seki, N. Sato, K. Sugahara, and K. Chiba, “Sphingosine 1-phosphate receptor type 1 regulates egress of mature T cells from mouse bone marrow,” International Immunology, vol. 22, no. 6, pp. 515–525, 2010.
View at: Publisher Site | Google Scholar
M. A. Ritter and D. B. Palmer, “The human thymic microenvironment: new approaches to functional analysis,” Seminars in Immunology, vol. 11, no. 1, pp. 13–21, 1999.
View at: Publisher Site | Google Scholar
N. R. Manley, E. R. Richie, C. C. Blackburn, B. G. Condie, and J. Sage, “Structure and function of the thymic microenvironment,” Frontiers in Bioscience, vol. 16, no. 7, pp. 2461–2477, 2011.
View at: Publisher Site | Google Scholar
K. Hirokawa and T. Makinodan, “Thymic involution: effect on T cell differentiation,” Journal of Immunology, vol. 114, no. 6, pp. 1659–1664, 1975.
View at: Google Scholar
L. Henry and G. Anderson, “Epithelial-cell architecture during involution of the human thymus,” Journal of Pathology, vol. 152, no. 3, pp. 149–155, 1987.
View at: Google Scholar
N. Yajima, K. Sakamaki, and S. Yonehara, “Age-related thymic involution is mediated by Fas on thymic epithelial cells,” International Immunology, vol. 16, no. 7, pp. 1027–1035, 2004.
View at: Publisher Site | Google Scholar
G. Anderson and Y. Takahama, “Thymic epithelial cells: working class heroes for T cell development and repertoire selection,” Trends in Immunology, vol. 33, no. 6, pp. 256–263, 2012.
View at: Publisher Site | Google Scholar
S. R. Wellhausen and D. L. Boros, “Atrophy of the thymic cortex in mice with granulomatous Schistosomiasis mansoni,” Infection and Immunity, vol. 35, no. 3, pp. 1063–1069, 1982.
View at: Google Scholar
P. G. Auwaerter, H. Kaneshima, J. M. McCune, G. Wiegand, and D. E. Griffin, “Measles virus infection of thymic epithelium in the SCID-hu mouse leads to thymocyte apoptosis,” Journal of Virology, vol. 70, no. 6, pp. 3734–3740, 1996.
View at: Google Scholar
P. C. S. Souto, V. N. Brito, J. Gameiro, M. A. da Cruz-Höfling, and L. Verinaud, “Programmed cell death in thymus during experimental paracoccidioidomycosis,” Medical Microbiology and Immunology, vol. 192, no. 4, pp. 225–229, 2003.
View at: Publisher Site | Google Scholar
V. N. Brito, P. C. S. Souto, M. A. Cruz-Höfling, L. C. Ricci, and L. Verinaud, “Thymus invasion and atrophy induced by Paracoccidioides brasiliensis in BALB/c mice,” Medical Mycology, vol. 41, no. 2, pp. 83–87, 2003.
View at: Google Scholar
W. Chen, R. Kuolee, J. W. Austin, H. Shen, Y. Che, and J. W. Conlan, “Low dose aerosol infection of mice with virulent type A Francisella tularensis induces severe thymus atrophy and CD4⁺CD8⁺ thymocyte depletion,” Microbial Pathogenesis, vol. 39, no. 5-6, pp. 189–196, 2005.
View at: Publisher Site | Google Scholar
W. Savino, “The thymus is a common target organ in infectious diseases,” PLoS Pathogens, vol. 2, no. 6, article e62, 2006.
View at: Publisher Site | Google Scholar
J. Gameiro, P. R. A. Nagib, C. F. Andrade et al., “Changes in cell migration-related molecules expressed by thymic microenvironment during experimental Plasmodium berghei infection: consequences on thymocyte development,” Immunology, vol. 129, no. 2, pp. 248–256, 2010.
View at: Publisher Site | Google Scholar
C. Francelin, L. C. Paulino, J. Gameiro, and L. Verinaud, “Effects of Plasmodium berghei on thymus: high levels of apoptosis and premature egress of CD4⁺CD8⁺ thymocytes in experimentally infected mice,” Immunobiology, vol. 216, no. 10, pp. 1148–1154, 2011.
View at: Publisher Site | Google Scholar
A. Morrot, J. B. de Albuquerque, L. R. Berbert, C. E. de Carvalho Pinto, J. de Meis, and W. Savino, “Dynamics of lymphocyte populations during Trypanosoma cruzi infection: from thymocyte depletion to differential cell expansion/contraction in peripheral lymphoid organs,” Journal of Tropical Medicine, vol. 2012, Article ID 747185, 7 pages, 2012.
View at: Publisher Site | Google Scholar
L. Verinaud, M. A. da Cruz-Höfling, J. K. Sakurada et al., “Immunodepression induced by Trypanosoma cruzi and mouse hepatitis virus type 3 is associated with thymus apoptosis,” Clinical and Diagnostic Laboratory Immunology, vol. 5, no. 2, pp. 186–191, 1998.
View at: Google Scholar
J. de Meis, D. A. Farias-de-Oliveira, P. H. N. Panzenhagen et al., “Thymus atrophy and double-positive escape are common features in infectious diseases,” Journal of Parasitology Research, vol. 2012, Article ID 574020, 9 pages, 2012.
View at: Publisher Site | Google Scholar
M. D. C. L. de Moraes, P. Minoprio, M. Dy, M. Dardenne, W. Savino, and M. Hontebeyrie-Joskowicz, “Endogenous IL-10 and IFN-γ production controls thymic cell proliferation in mice acutely infected by Trypanosoma cruzi,” Scandinavian Journal of Immunology, vol. 39, no. 1, pp. 51–58, 1994.
View at: Publisher Site | Google Scholar
B. M. Greene and D. G. Colley, “Eosinophils and immune mechanisms. III. Production of the lymphokine eosinophil stimulation promotor by mouse T lymphocytes,” Journal of Immunology, vol. 116, no. 4, pp. 1078–1083, 1976.
View at: Google Scholar
A. Henriques-Pons, J. DeMeis, V. Cotta-de-Almeida, W. Savino, and T. C. Araújo-Jorge, “Fas and perforin are not required for thymus atrophy induced by Trypanosoma cruzi infection,” Experimental Parasitology, vol. 107, no. 1-2, pp. 1–4, 2004.
View at: Publisher Site | Google Scholar
E. Roggero, A. R. Pérez, O. A. Bottasso, H. O. Besedovsky, and A. del Rey, “Neuroendocrine-immunology of experimental Chagas' disease,” Annals of the New York Academy of Sciences, vol. 1153, pp. 264–271, 2009.
View at: Publisher Site | Google Scholar
L. J. M. Carvalho, M. F. Ferreira-da-Cruz, C. T. Daniel-Ribeiro, M. Pelajo-Machado, and H. L. Lenzi, “Plasmodium berghei ANKA infection induces thymocyte apoptosis and thymocyte depletion in CBA mice,” Memórias do Instituto Oswaldo Cruz, vol. 101, no. 5, pp. 523–528, 2006.
View at: Google Scholar
M. C. L. de Moraes, M. Hontebeyrie-Joskowicz, F. Leboulenger, W. Savino, M. Dardenne, and F. Lepault, “Studies on the thymus in Chagas' disease. II. Thymocyte subset fluctuations in Trypanosoma cruzi-infected mice: relationship to stress,” Scandinavian Journal of Immunology, vol. 33, no. 3, pp. 267–275, 1991.
View at: Google Scholar
V. N. Ivanov and J. Nikolić-Žugić, “Biochemical and kinetic characterization of the glucocorticoid-induced apoptosis of immature CD4⁺CD8⁺ thymocytes,” International Immunology, vol. 10, no. 12, pp. 1807–1817, 1998.
View at: Google Scholar
E. Corrêa-De-Santana, M. Paez-Pereda, M. Theodoropoulou et al., “Hypothalamus-pituitary-adrenal axis during Trypanosoma cruzi acute infection in mice,” Journal of Neuroimmunology, vol. 173, no. 1-2, pp. 12–22, 2006.
View at: Publisher Site | Google Scholar
A. Lepletier, C. V. de Frias, A. Morrot, and W. Savino, “Thymic atrophy in acute experimental Chagas disease is associated with an imbalance of stress hormones,” Annals of the New York Academy of Sciences, vol. 1262, no. 1, pp. 45–50, 2012.
View at: Publisher Site | Google Scholar
C. S. Rosenberg, D. L. Martin, and R. L. Tarleton, “CD8⁺ T cells specific for immunodominant trans-sialidase epitopes contribute to control of Trypanosoma cruzi infection but are not required for resistance,” Journal of Immunology, vol. 185, no. 1, pp. 560–568, 2010.
View at: Publisher Site | Google Scholar
G. A. Rabinovich, L. G. Baum, N. Tinari et al., “Galectins and their ligands: amplifiers, silencers or tuners of the inflammatory response?” Trends in Immunology, vol. 23, no. 6, pp. 313–320, 2002.
View at: Publisher Site | Google Scholar
W. Savino, D. A. Mendes-da-Cruz, S. Smaniotto, E. Silva-Monteiro, and D. M. S. Villa-Verde, “Molecular mechanisms governing thymocyte migration: combined role of chemokines and extracellular matrix,” Journal of Leukocyte Biology, vol. 75, no. 6, pp. 951–961, 2004.
View at: Publisher Site | Google Scholar
B. N. Stillman, D. K. Hsu, M. Pang et al., “Galectin-3 and galectin-1 bind distinct cell surface glycoprotein receptors to induce T cell death,” Journal of Immunology, vol. 176, no. 2, pp. 778–789, 2006.
View at: Google Scholar
E. Silva-Monteiro, L. R. Lorenzato, O. K. Nihei et al., “Altered expression of galectin-3 induces cortical thymocyte depletion and premature exit of immature thymocytes during Trypanosoma cruzi infection,” The American Journal of Pathology, vol. 170, no. 2, pp. 546–556, 2007.
View at: Publisher Site | Google Scholar
M. Mantuano-Barradas, A. Henriques-Pons, T. C. Araújo-Jorge, F. Di Virgilio, R. Coutinho-Silva, and P. M. Persechini, “Extracellular ATP induces cell death in CD4⁺/CD8⁺ double-positive thymocytes in mice infected with Trypanosoma cruzi,” Microbes and Infection, vol. 5, no. 15, pp. 1363–1371, 2003.
View at: Publisher Site | Google Scholar
W. Savino, M. D. C. L. de Moraes, M. Hontebeyrie-Joskowicz, and M. Dardenne, “Studies on the thymus in Chagas' disease. I. Changes in the thymic microenvironment in mice acutely infected with Trypanosoma cruzi,” European Journal of Immunology, vol. 19, no. 9, pp. 1727–1733, 1989.
View at: Google Scholar
V. Cotta-de-Almeida, A. Bonomo, D. A. Mendes-da-Cruz et al., “Trypanosoma cruzi infection modulates intrathymic contents of extracellular matrix ligands and receptors and alters thymocyte migration,” European Journal of Immunology, vol. 33, no. 9, pp. 2439–2448, 2003.
View at: Publisher Site | Google Scholar
E. Roggero, A. R. Pérez, M. Tamae-Kakazu et al., “Edogenous glucocorticoids cause thymus atrophy but are protective during acute Trypanosoma cruzi infection,” Journal of Endocrinology, vol. 190, no. 2, pp. 495–503, 2006.
View at: Publisher Site | Google Scholar
A. R. Pérez, E. Roggero, A. Nicora et al., “Thymus atrophy during Trypanosoma cruzi infection is caused by an immuno-endocrine imbalance,” Brain, Behavior, and Immunity, vol. 21, no. 7, pp. 890–900, 2007.
View at: Publisher Site | Google Scholar
S. Schenkman, M. Jiang, G. W. Hart, and V. Nussenzweig, “A novel cell surface trans-sialidase of Trypanosoma cruzi generates a stage-specific epitope required for invasion of mammalian cells,” Cell, vol. 65, no. 7, pp. 1117–1125, 1991.
View at: Google Scholar
S. Schenkman, L. P. de Carvalho, and V. Nussenzweig, “Trypanosoma cruzi trans-sialidase and neuraminidase activities can be mediated by the same enzymes,” Journal of Experimental Medicine, vol. 175, no. 2, pp. 567–575, 1992.
View at: Google Scholar
S. S. Dc-Rubin and S. Schenkman, “Trypanosoma cruzi trans-sialidase as a multifunctional enzyme in Chagas' disease,” Cellular Microbiology, vol. 14, no. 10, pp. 1522–1530, 2012.
View at: Publisher Site | Google Scholar
D. H. Gray, F. Kupresanin, S. P. Berzins et al., “The BH3-only proteins Bim and Puma cooperate to impose deletional tolerance of organ-specific antigens,” Immunity, vol. 37, no. 3, pp. 451–462, 2012.
View at: Publisher Site | Google Scholar
J. Sprent and H. Kishimoto, “The thymus and negative selection,” Immunological Reviews, vol. 185, pp. 126–135, 2002.
View at: Publisher Site | Google Scholar
N. M. Milićević and Z. Milićević, “Thymus cell-cell interactions,” International Review of Cytology, vol. 235, pp. 1–52, 2004.
View at: Publisher Site | Google Scholar
N. M. Milićević and Z. Milićević, “Ultrastructural identification of specialized endocytic compartments in macrophages of the thymic cortico-medullary zone and germinal centers of peripheral lymphatic organs of the rat,” Annals of Anatomy, vol. 182, no. 5, pp. 471–478, 2000.
View at: Google Scholar
A. Morrot, E. Terra-Granado, A. R. Pérez et al., “Chagasic thymic atrophy does not affect negative selection but results in the export of activated CD4⁺CD8⁺ T cells in severe forms of human disease,” PLoS Neglected Tropical Diseases, vol. 5, no. 8, Article ID e1268, 2011.
View at: Publisher Site | Google Scholar
E. Fiorini, P. C. Marchisio, M. T. Scupoli et al., “Adhesion of immature and mature T cells induces in human thymic epithelial cells (TEC) activation of IL-6 gene trascription factors (NF-κB and NF-IL6) and IL-6 gene expression: Role of α3β1 and α6β4 integrins,” Developmental Immunology, vol. 7, no. 2–4, pp. 195–208, 2000.
View at: Google Scholar
T. Cejalvo, J. J. Munoz, E. Tobajas et al., “Ephrin-B-dependent thymic epithelial cell-thymocyte interactions are necessary for correct T cell differentiation and thymus histology organization: relevance for thymic cortex development,” Journal of Immunology, vol. 190, no. 6, pp. 2670–2681, 2013.
View at: Publisher Site | Google Scholar
G. A. Hollander, B. Wang, A. Nichogiannopoulou et al., “Developmental control point in induction of thymic cortex regulated by a subpopulation of prothymocytes,” Nature, vol. 373, no. 6512, pp. 350–353, 1995.
View at: Google Scholar
S. Zuklys, G. Balciunaite, A. Agarwal, E. Fasler-Kan, E. Palmer, and G. A. Hollander, “Normal thymic architecture and negative selection are associated with Aire expression, the gene defective in the autoimmune-polyendocrinopathy-candidiasis-ectodermal dystrophy (APECED),” Journal of Immunology, vol. 165, no. 4, pp. 1976–1983, 2000.
View at: Google Scholar
P. Peterson, T. Org, and A. Rebane, “Transcriptional regulation by AIRE: molecular mechanisms of central tolerance,” Nature Reviews Immunology, vol. 8, no. 12, pp. 948–957, 2008.
View at: Publisher Site | Google Scholar
M. Zhu, R. K. Chin, P. A. Christiansen et al., “NF-κB2 is required for the establishment of central tolerance through an Aire-dependent pathway,” Journal of Clinical Investigation, vol. 116, no. 11, pp. 2964–2971, 2006.
View at: Publisher Site | Google Scholar
R. T. Taniguchi and M. S. Anderson, “The role of Aire in clonal selection,” Immunology and Cell Biology, vol. 89, no. 1, pp. 40–44, 2011.
View at: Publisher Site | Google Scholar
M. C. L. de Moraes, M. Hontebeyrie-Joskowicz, M. Dardenne, and W. Savino, “Modulation of thymocyte subsets during acute and chronic phases of experimental Trypanosoma cruzi infection,” Immunology, vol. 77, no. 1, pp. 95–98, 1992.
View at: Google Scholar
X. Liu, S. Dai, Y. Zhu, P. Marrack, and J. W. Kappler, “The structure of a Bcl-xL/Bim fragment complex: implications for Bim function,” Immunity, vol. 19, no. 3, pp. 341–352, 2003.
View at: Publisher Site | Google Scholar
S. Schenkman, M. A. J. Ferguson, N. Heise, M. L. C. de Almeida, R. A. Mortara, and N. Yoshida, “Mucin-like glycoproteins linked to the membrane by glycosylphosphatidylinositol anchor are the major acceptors of sialic acid in a reaction catalyzed by trans-sialidase in metacyclic forms of Trypanosoma cruzi,” Molecular and Biochemical Parasitology, vol. 59, no. 2, pp. 293–303, 1993.
View at: Publisher Site | Google Scholar
A. R. Todeschini, W. B. Dias, M. F. Girard, J. Wieruszeski, L. Mendonça-Previato, and J. O. Previato, “Enzymatically inactive trans-sialidase from Trypanosoma cruzi binds sialyl and β-galactopyranosyl residues in a sequential ordered mechanism,” The Journal of Biological Chemistry, vol. 279, no. 7, pp. 5323–5328, 2004.
View at: Publisher Site | Google Scholar
M. S. Leguizamon, O. E. Campetella, S. M. G. Cappa, and A. C. C. Frasch, “Mice infected with Trypanosoma cruzi produce antibodies against the enzymatic domain of trans-sialidase that inhibit its activity,” Infection and Immunity, vol. 62, no. 8, pp. 3441–3446, 1994.
View at: Google Scholar
J. Mucci, E. Mocetti, M. S. Leguizamón, and O. Campetella, “A sexual dimorphism in intrathymic sialylation survey is revealed by the trans-sialidase from Trypanosoma cruzi,” Journal of Immunology, vol. 174, no. 8, pp. 4545–4550, 2005.
View at: Google Scholar
J. Mucci, A. Hidalgo, E. Mocetti, P. F. Argibay, M. S. Leguizamón, and O. Campetella, “Thymocyte depletion in Trypanosoma cruzi infection is mediated by trans-sialidase-induced apoptosis on nurse cells complex,” Proceedings of the National Academy of Sciences of the United States of America, vol. 99, no. 6, pp. 3896–3901, 2002.
View at: Publisher Site | Google Scholar
R. P. Muiá, H. Yu, J. A. Prescher et al., “Identification of glycoproteins targeted by Trypanosoma cruzi trans-sialidase, a virulence factor that disturbs lymphocyte glycosylation,” Glycobiology, vol. 20, no. 7, pp. 833–842, 2010.
View at: Publisher Site | Google Scholar
A. F. Nardy, J. L. da Silva Filho, A. R. Pérez et al., “Trans-sialidase from Trypanosoma cruzi enhances the adhesion properties and fibronectin-driven migration of thymocytes,” Microbes and Infection, vol. 15, no. 5, pp. 365–374, 2013.
View at: Publisher Site | Google Scholar
S. C. da Costa, K. S. Calabrese, P. G. Bauer, W. Savino, and P. H. Lagrange, “Studies of the thymus in Chagas' disease: III. Colonization of the thymus and other lymphoid organs of adult and newborn mice by Trypanosoma cruzi,” Pathologie Biologie, vol. 39, no. 2, pp. 91–97, 1991.
View at: Google Scholar
K. Utsumi, M. Sawada, S. Narumiya et al., “Adhesion of immature thymocytes to thymic stromal cells through fibronectin molecules and its significance for the induction of thymocyte differentiation,” Proceedings of the National Academy of Sciences of the United States of America, vol. 88, no. 13, pp. 5685–5689, 1991.
View at: Publisher Site | Google Scholar
W. Savino, D. A. Mendes-da-Cruz, J. S. Silva, M. Dardenne, and V. Cotta-de-Almeida, “Intrathymic T-cell migration: a combinatorial interplay of extracellular matrix and chemokines?” Trends in Immunology, vol. 23, no. 6, pp. 305–313, 2002.
View at: Publisher Site | Google Scholar
W. Gillespie, J. C. Paulson, S. Kelm, M. Pang, and L. G. Baum, “Regulation of α2,3-sialyltransferase expression correlates with conversion of peanut agglutinin (PNA)+ to PNA- phenotype in developing thymocytes,” The Journal of Biological Chemistry, vol. 268, no. 6, pp. 3801–3804, 1993.
View at: Google Scholar
L. Linhares-Lacerda, M. Ribeiro-Alves, A. C. M. D. A. Nogueira et al., “RNA interference-mediated knockdown of CD49e (α5 integrin chain) in human thymic epithelial cells modulates the expression of multiple genes and decreases thymocyte adhesion,” BMC Genomics, vol. 11, supplement 5, article S2, 2010.
View at: Publisher Site | Google Scholar
I. M. Otto, T. Raabe, U. E. E. Rennefahrt, P. Bork, U. R. Rapp, and E. Kerkhoff, “The p150-Spir protein provides a link between c-Jun N-terminal kinase function and actin reorganization,” Current Biology, vol. 10, no. 6, pp. 345–348, 2000.
View at: Publisher Site | Google Scholar
D. A. Mendes-da-Cruz, J. de Meis, V. Cotta-de-Almeida, and W. Savino, “Experimental Trypanosoma cruzi infection alters the shaping of the central and peripheral T-cell repertoire,” Microbes and Infection, vol. 5, no. 10, pp. 825–832, 2003.
View at: Publisher Site | Google Scholar
N. A. Giraldo, N. I. Bolaños, A. Cuellar et al., “Increased CD4⁺/CD8⁺ double-positive T cells in chronic chagasic patients,” PLoS Neglected Tropical Diseases, vol. 5, no. 8, Article ID e1294, 2011.
View at: Publisher Site | Google Scholar
S. D. Rosen, “Ligands for L-selectin: homing, inflammation, and beyond,” Annual Review of Immunology, vol. 22, pp. 129–156, 2004.
View at: Publisher Site | Google Scholar
K. Nam, H. Akari, K. Terao, H. Shibata, S. Kawamura, and Y. Yoshikawa, “Peripheral blood extrathymic CD4⁺ CD8⁺ T cells with high cytotoxic activity are from the same lineage as CD4⁺ CD8^- T cells in cynomolgus monkeys,” International Immunology, vol. 12, no. 7, pp. 1095–1103, 2000.
View at: Google Scholar
M. Nascimbeni, E. Shin, L. Chiriboga, D. E. Kleiner, and B. Rehermann, “Peripheral CD4⁺CD8⁺ T cells are differentiated effector memory cells with antiviral functions,” Blood, vol. 104, no. 2, pp. 478–486, 2004.
View at: Publisher Site | Google Scholar
H. Akari, K. Terao, Y. Murayama, K. Nam, and Y. Yoshikawa, “Peripheral blood CD4⁺CD8⁺ lymphocytes in cynomolgus monkeys are of resting memory T lineage,” International Immunology, vol. 9, no. 4, pp. 591–597, 1997.
View at: Publisher Site | Google Scholar
H. Akari, K. Nam, K. Mori et al., “Effects of SIVmac infection on peripheral blood CD4⁺CD8⁺ T lymphocytes in cynomolgus macaques,” Clinical Immunology, vol. 91, no. 3, pp. 321–329, 1999.
View at: Publisher Site | Google Scholar
L. Lamontagne, E. Massicotte, and C. Page, “Mouse hepatitis viral infection induces an extrathymic differentiation of the specific intrahepatic αβ-TCR(intermediate) LFA-1(high) T-cell population,” Immunology, vol. 90, no. 3, pp. 402–411, 1997.
View at: Google Scholar
M. Nascimbeni, S. Pol, and B. Saunier, “Distinct CD4⁺CD8⁺ double-positive T cells in the blood and liver of patients during chronic hepatitis B and C,” PLoS ONE, vol. 6, no. 5, Article ID e20145, 2011.
View at: Publisher Site | Google Scholar
L. Weiss, A. Roux, S. Garcia et al., “Persistent expansion, in a human immunodeficiency virus-infected person, of Vβ-restricted CD4⁺CD8⁺ T lymphocytes that express cytotoxicity-associated molecules and are committed to produce interferon-γ and tumor necrosis factor-α,” Journal of Infectious Diseases, vol. 178, no. 4, pp. 1158–1162, 1998.
View at: Google Scholar
M. A. Suni, S. A. Ghanekar, D. W. Houck et al., “CD4⁺CD8^dim T lymphocyte exhit enhanced cytokine expression, proliferation and cytotoxic activity in response to HCMV and HIV-1 antigens,” European Journal of Immunology, vol. 31, no. 8, pp. 2512–2520, 2001.
View at: Publisher Site | Google Scholar
A. Zloza, Y. B. Sullivan, E. Connick, A. L. Landay, and L. Al-Harthi, “CD8⁺ T cells that express CD4 on their surface (CD4 ^dimCD8^bright T cells) recognize an antigen-specific target, are detected in vivo, and can be productively infected by T-tropic HIV,” Blood, vol. 102, no. 6, pp. 2156–2164, 2003.
View at: Publisher Site | Google Scholar
R. Howe, S. Dillon, L. Rogers et al., “Phenotypic and functional characterization of HIV-1-specific CD4⁺CD8⁺ double-positive T cells in early and chronic HIV-1 infection,” Journal of Acquired Immune Deficiency Syndromes, vol. 50, no. 5, pp. 444–456, 2009.
View at: Publisher Site | Google Scholar
N. K. Chauhan, M. Vajpayee, K. Mojumdar, R. Singh, and A. Singh, “Study of CD4⁺CD8⁺ double positive T-lymphocyte phenotype and function in Indian patients infected with HIV-1,” Journal of Medical Virology, vol. 84, no. 6, pp. 845–856, 2012.
View at: Publisher Site | Google Scholar
M. A. Frahm, R. A. Picking, J. D. Kuruc et al., “CD4⁺CD8⁺ T cells represent a significant portion of the anti-HIV T cell response to acute HIV infection,” Journal of Immunology, vol. 188, no. 9, pp. 4289–4296, 2012.
View at: Publisher Site | Google Scholar
L. D. Barber and P. Parham, “Peptide binding to major histocompatibility complex molecules,” Annual Review of Cell Biology, vol. 9, pp. 163–206, 1993.
View at: Google Scholar
R. Brink, T. G. Phan, D. Paus, and T. D. Chan, “Visualizing the effects of antigen affinity on T-dependent B-cell differentiation,” Immunology and Cell Biology, vol. 86, no. 1, pp. 31–39, 2008.
View at: Publisher Site | Google Scholar
J. Hernández, Y. Garfias, A. Nieto, C. Mercado, L. F. Montaño, and E. Zenteno, “Comparative evaluation of the CD4⁺CD8⁺ and CD4⁺CD8^- lymphocytes in the immune response to porcine rubulavirus,” Veterinary Immunology and Immunopathology, vol. 79, no. 3-4, pp. 249–259, 2001.
View at: Publisher Site | Google Scholar
P. Hillemeyer, M. D. White, and D. W. Pascual, “Development of a transient CD4⁺CD8⁺ T cell subset in the cervical lymph nodes following intratracheal instillation with an adenovirus vector,” Cellular Immunology, vol. 215, no. 2, pp. 173–185, 2002.
View at: Publisher Site | Google Scholar
S. Raza, S. Naik, V. P. Kancharla, F. Tafera, and M. R. Kalavar, “Dual-positive (CD4⁺/CD8⁺) acute adult T-cell leukemia/lymphoma associated with complex karyotype and refractory hypercalcemia: case report and literature review,” Case Reports in Oncology, vol. 3, no. 3, pp. 489–494, 2010.
View at: Publisher Site | Google Scholar
D. Bismarck, N. Schütze, P. Moore, M. Büttner, G. Alber, and H. V. Buttlar, “Canine CD4⁺CD8⁺ double positive T cells in peripheral blood have features of activated T cells,” Veterinary Immunology and Immunopathology, vol. 149, no. 3-4, pp. 157–166, 2012.
View at: Publisher Site | Google Scholar
B. T. Ober, A. Summerfield, C. Mattlinger et al., “Vaccine-induced, pseudorabies virus-specific, extrathymic CD4⁺CD8⁺ memory T-helper cells in swine,” Journal of Virology, vol. 72, no. 6, pp. 4866–4873, 1998.
View at: Google Scholar
T. G. M. de Bruin, E. M. A. van Rooij, Y. E. de Visser, and A. T. J. Bianchi, “Cytolytic function for pseudorabies virus-stimulated porcine CD4⁺CD8dull+ lymphocytes,” Viral Immunology, vol. 13, no. 4, pp. 511–520, 2000.
View at: Google Scholar
V. Dwivedi, C. Manickam, R. Patterson et al., “Cross-protective immunity to porcine reproductive and respiratory syndrome virus by intranasal delivery of a live virus vaccine with a potent adjuvant,” Vaccine, vol. 29, no. 23, pp. 4058–4066, 2011.
View at: Publisher Site | Google Scholar
Y. Parel and C. Chizzolini, “CD4⁺ CD8⁺ double positive (DP) T cells in health and disease,” Autoimmunity Reviews, vol. 3, no. 3, pp. 215–220, 2004.
View at: Publisher Site | Google Scholar
S. G. Kitchen, Y. D. Korin, M. D. Roth, A. Landay, and J. A. Zack, “Costimulation of naive CD8⁺ lymphocytes induces CD4 expression and allows human immunodeficiency virus type 1 infection,” Journal of Virology, vol. 72, no. 11, pp. 9054–9060, 1998.
View at: Google Scholar
G. J. Hughes, A. Cochrane, C. Leen, S. Morris, J. E. Bell, and P. Simmonds, “HIV-1-infected CD8⁺CD4⁺ T cells decay in vivo at a similar rate to infected CD4 T cells during HAART,” AIDS, vol. 22, no. 1, pp. 57–65, 2008.
View at: Publisher Site | Google Scholar
J. J. Mattapallil, E. Reay, and S. Dandekar, “An early expansion of CD8αβ T cells, but depletion of resident CD8αα T cells, occurs in the intestinal epithelium during primary simian immunodeficiency virus infection,” AIDS, vol. 14, no. 6, pp. 637–646, 2000.
View at: Publisher Site | Google Scholar
R. S. Veazey, K. G. Mansfield, I. C. Tham et al., “Dynamics of CCR5 expression by CD4⁺ T cells in lymphoid tissues during simian immunodeficiency virus infection,” Journal of Virology, vol. 74, no. 23, pp. 11001–11007, 2000.
View at: Publisher Site | Google Scholar
V. M. Hirsch, “What can natural infection of African monkeys with simian immunodeficiency virus tell us about the pathogenesis of AIDS?” AIDS Reviews, vol. 6, no. 1, pp. 40–53, 2004.
View at: Google Scholar
J. W. Mellors, C. R. Rinaldo Jr., P. Gupta, R. M. White, J. A. Todd, and L. A. Kingsley, “Prognosis in HIV-1 infection predicted by the quantity of virus in plasma,” Science, vol. 272, no. 5265, pp. 1167–1170, 1996.
View at: Google Scholar
R. Gautam, T. Gaufin, I. Butler et al., “Simian immunodeficiency virus SIVrcm, a unique CCR2-tropic virus, selectively depletes memory CD4⁺ T cells in pigtailed macaques through expanded coreceptor usage in vivo,” Journal of Virology, vol. 83, no. 16, pp. 7894–7908, 2009.
View at: Publisher Site | Google Scholar
V. Ribrag, D. Salmon, F. Picard, M. Guesnu, D. Sicard, and F. Dreyfus, “Increase in double-positive CD4⁺CD8⁺ peripheral T-cell subsets in an HIV-infected patient,” AIDS, vol. 7, no. 11, article 1530, 1993.
View at: Google Scholar
G. F. Ferraccioli, E. Tonutti, L. Casatta et al., “CD4 cytopenia and occasional expansion of CD4⁺CD8⁺ lymphocytes in Sjogren's syndrome,” Clinical and Experimental Rheumatology, vol. 14, no. 2, pp. 125–130, 1996.
View at: Google Scholar
J. Hirao and K. Sugita, “Circulating CD4⁺CD8⁺ T lymphocytes in patients with Kawasaki disease,” Clinical and Experimental Immunology, vol. 111, no. 2, pp. 397–401, 1998.
View at: Publisher Site | Google Scholar

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Copyright © 2013 Alexandre Morrot. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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