Cellular prion protein (PrPC) is an ubiquitously expressed glycoprotein whose roles are still widely discussed, particularly in the field of immunology. Using TgA20- and Tg33-transgenic mice overexpressing PrPC, we investigated the consequences of this overexpression on T cell development. In both models, overexpression of PrPC induces strong alterations at different steps of T cell maturation. On TgA20 mice, we observed that these alterations are cell autonomous and lead to a decrease of αβ T cells and a concomitant increase of γδ T cell numbers. PrPC has been shown to bind and chelate copper and, interestingly, under a copper supplementation diet, TgA20 mice presented a partial restoration of the αβ T cell development, suggesting that PrPC overexpression, by chelating copper, generates an antioxidant context differentially impacting on αβ and γδ T cell lineage.

Prion diseases form a group of fatal neurodegenerative disorders including Creutzfeldt-Jakob disease, kuru and fatal familial insomnia in humans, scrapie, and bovine spongiform encephalopathy in animals (1). Cellular prion protein (PrPC)5 is a surface GPI-anchored glycoprotein mainly expressed on neurons (2, 3). The amino-terminal region of PrPC contains five repeating, highly conserved octapeptide domains that bind Cu2+ and other metal ion such as Ni2+, Zn2+, or Mn2+ with lower affinity (4, 5, 6, 7). The physiologic function(s) of PrPC remain(s) to be fully defined. One of the many hypotheses supports a protective function against oxidative stress (4, 7, 8, 9, 10). PrPC may also play a role in signal transduction through members of the Fyn tyrosine kinase family (11, 12, 13).

Most studies about cellular and scrapie pathological prion protein (PrPsc) are focused on the brain. Pathological lesions associated with prion diseases are found in the CNS, but several studies implicate the lymphoid system in the pathogenesis of the disease. PrPSc was found in lymph nodes, spleen, and Peyer’s patches of infected mice (14, 15). Then, it was reported that SCID mice, lacking both B and T cells, are no longer able to support PrPSc replication in the spleen (16). Recently, an essential role in PrPSc infection was attributed to follicular dendritic cells in the spleen (17, 18) and circulating dendritic cells (19). Finally, precursors, immature thymocytes, and activated mature T cells express moderate level of PrPC, whereas most of the mature lymphocytes are PrPC negative (20, 21). The outcome of differential expression of the PrPC among various T cell populations remains to be fully analyzed during the differentiation of functional T lymphocytes.

αβ T cell development in the thymus is characterized by successive differentiation steps (22): T lymphocyte precursors begin as CD4CD8, double-negative (DN) cells. Development of this DN population is divided into four cell subpopulations based on the expression of CD25 and CD44: CD44+CD25 (DN1), CD44+CD25+ (DN2), CD44CD25+ (DN3), and CD44CD25 (DN4). Then, DN4 thymocytes progress to CD4+CD8+ double-positive (DP) cells and begin to express low levels of αβTCR. A fraction of the DP cells is positively selected to become either CD4+ or CD8+ single-positive (SP) cells, which finally immigrate to the periphery. In the thymus, a weak proportion of the T cells bear at their surface a γδTCR. These T cells differentiate along the DN stage and most of them never express the CD4 or CD8 molecules (23).

Several lines of evidence support that the immune system is affected by copper levels (24). For instance, the immune response is impaired in case of copper deficiency (25, 26, 27, 28); furthermore, reduction of the copper amount decreases both IL-2 mRNA and IL-2 production in peripheral T cells (29, 30). At last, the lack of copper can also promote oxidative stress and damage in T cells, compromising the antioxidant defense system (31, 32).

It is now well documented that stress affects the function of B and T cells and more particularly that TCR signaling pathways are affected by oxidative stress (33). Using an ex vivo experimental approach, it was observed that antioxidants affect thymus development by decreasing the number of αβ T cells (34). Taking into account the protective action of PrPC against oxidative stress (4, 7, 8, 9, 10) and the alteration of T cell development by antioxidants, we wondered whether expression of PrPC could modulate T cell differentiation. To address this point, we analyzed mice that overexpress PrPC. The TgA20 mouse strain was created by introducing a transgene encoding WT murine PrPC under the control of its own promoter into a PrP0/0 mouse (35), whereas Tg33 mice have a transgene driven by the lck promoter, limiting the expression of PrPC to T cells (36). No immunological disorder is evident in PrP0/0 mice (37), but they do have a reduced ability to accommodate oxidative stress (38).

Examination of T cell development in TgA20 mice revealed a reduced thymus size, a profound alteration of αβ T cell differentiation, and an expansion of γδ T cells in the thymus. In 10.5-wk-old Tg33 mice, we also observed a reduced thymus size, along with a reduction of the percentage of DP cells. In TgA20, the defects in αβ T cell development were partially restored by dietary copper supplementation, suggesting that the overexpression of PrPC interferes with T cell redox activity that crucially impacts development of normal αβ/γδ T cell balance.

TgA20 mice (35), PrP−/− (37), Sv129, and C57BL/6 were obtained from the Centre National de la Recherche Scientifique/Centre de Distribution, Typafe et Archivage Animal (Orléans, France) and maintained under specific pathogen-free conditions in the animal facility of the Commissariat à l’Energie Atomique-Grenoble in accordance with institutional guidelines. Tg33 mice (36) were bred and maintained in the animal facility of the Neuropathology Institute of Zurich (Zurich, Switzerland). TgA20, Tg33, and PrP-knockout mice are under a Sv129 × C57BL/6 background.

BM-chimeric mice were reconstituted by injecting 5 × 106 Thy-1-depleted BM cells into lethally irradiated (10 Gy) recipient mice. Mixed chimeras were performed by mixing 50% of C57BL/6 BM precursor cells (Ly5.1+) with 50% of C57BL/6 cells or 50% of TgA20 cells bearing the Ly5.2 allele. Thymocytes were analyzed 2 mo after reconstitution. FACS analyses were performed on Ly5.2+-gated cells. Maturation of Ly5.1 cells was normal in all situations (data not shown).

Supplemented TgA20 or WT mice were given water with 250 mg/L CuSO4 and 50 mg/L sucrose, whereas the control group received the same water but without copper. The diet was maintained in the course of pregnancy and lactation. After weaning, TgA20 and WT mice were watered for 1 more wk with the same diet as their mothers and sacrificed for analysis.

For flow cytometry analysis, single-cell suspensions were prepared in PBS, 3% FCS, 0.16% sodium azide (FACS wash), stained with saturating concentrations of conjugated Abs for 30 min at 4°C, and washed in FACS wash. Data acquisition and analysis were performed with a FACSCalibur flow cytometer equipped with CellQuest software (BD Biosciences). The conjugated Abs used for staining were all obtained from BD Pharmingen except for the anti-PrP Abs: SAF32 was obtained from SPI-Bio, then Alexa-488-conjugated (Molecular Probes) in our laboratory, and the 6H4-Cy5 Ab was obtained from Prionics.

Electrothermal atomic absorption spectrophotometry (PerkinElmer) was used to quantify the wet weight copper and zinc ions in brain and thymus from WT and TgA20 mice as described previously (39). Their levels were normalized to the protein content and measured with a protein assay kit (Pierce).

When needed, results are presented as mean ± SD of several experiments (n ≥3). Statistical significance was determined using Student’s t test.

Anatomic examination revealed a dramatic thymic atrophy in TgA20 mice as compared with control WT mice (SV129 × C57BL/6) and PrP0/0 mice (Fig. 1,A), whereas their spleen size was normal (data not shown). Although their size and cell number were similar, the spleen of the TgA20 mice contained many more cells expressing high levels of PrPC compared with WT splenocytes. Similarly, in TgA20 thymocytes, the level of expression of PrPC was ∼50 times that of WT mice thymocytes (Fig. 1,B). Hence, despite their atrophied thymus, TgA20 mice overexpress the PrPC protein. To examine whether this overexpression regulates T cell differentiation, cell suspensions from thymi of TgA20 and WT or PrP0/0 control mice were analyzed by FACS. The total number of thymocytes in TgA20 mice was 10.1 ± 1.3 million (n > 10), whereas the number of cells observed in WT mice was 151 ± 36 million (n > 10) and 141 ± 19 million in the PrP0/0 mice (n = 8). Since the FACS profiles of C57BL/6, Sv129, Sv129 × C57BL/6, and PrP0/0 strains and the proportion of each thymic subpopulation (DN, DP, SP, γδTCR+… ) were found to be similar (data not shown), we chose to present only the data obtained with the Sv129 × C57BL/6 mice, which corresponds to the background of the PrP0/0 and TgA20 mice. Analyses of thymocytes for cell surface expression of CD4 and CD8 indicated that TgA20 mice contain the four cell subsets but with dramatic changes in their relative proportions (Fig. 2,A). First, the major subset, namely, DP cells, which represented 85% of the total thymocytes in WT mice, was reduced to 32% in TgA20 mice. SP proportions were also decreased, especially CD4+ SP cells, from 9 to 4%, whereas CD8+ SP cells were only slightly affected. Second, DN cells representing <2% in WT thymus accounted for up to 61% of the TgA20 thymocytes. Because DN cells are defined by absence of CD4 and CD8, they may encompass various kinds of cells, including true T cell precursors, differentiated T cells expressing γδ TCR, and circulating cells such as B lymphocytes, NK, and dendritic cells. We further sought for the nature of the DN cells by conducting a detailed phenotypic analysis. Strikingly, although the percentage of cells bearing a γδTCR was <0.2% of the total thymocytes in WT mice, it reached up to 26% in TgA20 mice, corresponding to an increase of 9-fold in the absolute number of cells (Fig. 2,B). The proportion of B cells expressing CD19 and B220 markers increased in TgA20, although the actual numbers of B cells were identical in both mice (Fig. 2 C). Taken together, these data suggest that overexpression of PrPC promotes γδ T cells and has a major negative impact on αβ T cell differentiation.

FIGURE 1.

Thymic atrophy of the TgA20 and Tg33 thymi and PrP expression. A, Thymus from 10.5-wk-old WT, PrP−/−, TgA20, and Tg33 mice. The data are representative of >10 animals analyzed. B and C, Thymocytes and splenocytes from TgA20 (B) or Tg33 (C) (solid lines) and control (gray histograms) mice were stained with the SAF-32 (B) or 6H4 (C) mAb.

FIGURE 1.

Thymic atrophy of the TgA20 and Tg33 thymi and PrP expression. A, Thymus from 10.5-wk-old WT, PrP−/−, TgA20, and Tg33 mice. The data are representative of >10 animals analyzed. B and C, Thymocytes and splenocytes from TgA20 (B) or Tg33 (C) (solid lines) and control (gray histograms) mice were stained with the SAF-32 (B) or 6H4 (C) mAb.

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FIGURE 2.

Thymic subpopulations. A–C, Thymic cells from WT and TgA20 mice were stained with the indicated mAb and analyzed by FACS. D, DN subpopulations: cells gated for CD4, CD8, CD11c, CD19, NK1.1, and γδTCR were analyzed for CD44 and CD25 expression. Data are representative of 10 TgA20 and 10 control mice.

FIGURE 2.

Thymic subpopulations. A–C, Thymic cells from WT and TgA20 mice were stained with the indicated mAb and analyzed by FACS. D, DN subpopulations: cells gated for CD4, CD8, CD11c, CD19, NK1.1, and γδTCR were analyzed for CD44 and CD25 expression. Data are representative of 10 TgA20 and 10 control mice.

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To further check for a putative impairment in the early stages of T cell differentiation, we analyzed the expression of CD25 and CD44 on DN T cells after depletion of CD4, CD8, CD11c, CD19, γδTCR, and NK1.1 markers (Fig. 2 D). These DN cells represented 2.7 × 106 and 4.1 × 106 cells in WT and TgA20, respectively. Consequently, the population of T cell precursors represented 41% of the total thymocytes in TgA20 but only 1.8% in the control mice. The partition of the four main DN subpopulations was disturbed: TgA20 mice had an increased percentage of DN3 (from 57 to 85%) and, reciprocally, the percentage of DN2 and DN4 decreased. Considering the absolute number of cells, the DN3 compartment contained 1.5 × 106 cells in control and 3.5 × 106 in TgA20 thymus. These data demonstrate that thymocyte differentiation is partially arrested at the immature DN3 stage in TgA20 mice and suggests again that overexpression of PrPC negatively interferes with αβ T cell development.

To rule out the possibility that our results could result from a transgene position effect, we performed additional experiments with another mouse strain overexpressing PrPC. Tg33 mice also derive from PrP0/0 mice, but unlike TgA20 mice, the transgene encoding WT murine PrPC is driven by the lck promoter, limiting the expression of PrPC to T cells (36). Unlike TgA20 mice, at 3.5 wk we observed no difference in thymic size or cellularity between WT and Tg33 mice. But with 10.5-wk-old TgA33 mice, significant PrPC expression on thymocytes correlated to decreased thymic size and cellularity (56 ± 5 million) (Fig. 1,A) without affecting the size or cellularity of the spleen. FACS analysis of PrPC expression in the thymus revealed the presence of one population expressing a high level of PrPC (Fig. 1,C), which has been shown to represent CD3+ cells (36). The thymi of these mice showed a reduction of the percentage of DP cells, but no significant increase in the percentage of DN cells was observed (Fig. 3). In contrast, an increase in immature CD8 SP cells (ISP) expressing an intermediate level of CD8, not underscored in TgA20, was noticed (Fig. 3). Finally, we observed a slight increase in the percentage (0.9%) and in the absolute number (0.51 × 106) of γδ+ T cells as compared with WT mice (0.2%–0.3 × 106) (data not shown).

FIGURE 3.

Thymic subpopulations in Tg33 mice. FACS analysis of thymocytes from Sv129 × C57BL/6, TgA20, and Tg33 mice. The gate shows the ISP cells. Percentages of each subpopulation are indicated

FIGURE 3.

Thymic subpopulations in Tg33 mice. FACS analysis of thymocytes from Sv129 × C57BL/6, TgA20, and Tg33 mice. The gate shows the ISP cells. Percentages of each subpopulation are indicated

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Therefore, in these mice also, the thymic differentiation process is altered. All of these data argue against a transgene or insertional artifact and confirm an effect of PrPC overexpression in thymic development. We have then focused the following experiments on the Tga20 line.

In the course of thymus differentiation, during the DN to DP transition, thymocytes down-regulate CD25 and up-regulate CD5 (40). In TgA20 mice, these transitory expressions were affected (Fig. 4): CD25 was still present on the surface of a large proportion of DP, CD4+ SP (44%), and CD8+ SP cells (18%), and the majority of DP and 48% of CD4+ SP cells expressed CD5 at lower levels. Again, CD8+ SP cells were less affected, with only 33% of them failing to up-regulate CD5. In WT mice, high expression of αβTCR was observed in all of the CD4+ SP and >70% of the CD8+ SP cells. This expression was altered in TgA20 mice, where only 53% of CD4+ and 57% of CD8+ cells expressed high levels of αβTCR (Fig. 4). Finally, one of the hallmarks of the differentiation process is the expression of the activation marker CD69 that first appears on DP thymocytes as they begin positive selection (41). In TgA20 mice, we observed a reduction of CD69 expression in CD4+ SP, suggesting a partially impaired positive selection. Taken together, our results show that the overexpression of PrPC has an impact on the major checkpoints of the αβ T cell differentiation pathway.

FIGURE 4.

Phenotypic analysis of cell subsets in the TgA20 and WT mice. Thymocytes from TgA20 (solid lines) and WT (gray histograms) mice were analyzed by FACS. Cells were gated on the DN, DP, CD4+, or CD8+ SP compartments and expression of CD25, CD5, αβ TCR, or CD69 was analyzed.

FIGURE 4.

Phenotypic analysis of cell subsets in the TgA20 and WT mice. Thymocytes from TgA20 (solid lines) and WT (gray histograms) mice were analyzed by FACS. Cells were gated on the DN, DP, CD4+, or CD8+ SP compartments and expression of CD25, CD5, αβ TCR, or CD69 was analyzed.

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We next investigated whether a correlation exists between the level of PrPC expression and the partial arrest in T cell differentiation observed in TgA20 mice. To address this question, FACS analysis was performed using the classical set of T lymphocyte-specific markers in combination with anti-PrPC Ab. In TgA20 mice, almost all DP thymocytes expressed high levels of PrPC (Fig. 5,A), whereas SP thymocytes can be subdivided into two distinct populations based on the expression of PrPC: one expressing high levels (35 and 13% for CD4+ and CD8+ cells, respectively) and the other expressing intermediate levels of PrPC in CD4+ or very low levels in CD8+ cells. In the DN subset of WT mice, ∼5% of the cells expressed PrPC. In contrast, in TgA20 mice, we found >50% of PrPC-positive cells in this subset. To further identify the phenotype of the PrPC-positive cells in the DN population, we used CD44 and CD25 markers in combination with PrPC. In TgA20 mice, the expression of PrPC was first seen on >15% of DN2 thymocytes, 89% of DN3 cells, and only 15% of the DN4 cells (Fig. 5 B). Therefore, in TgA20 mice, the expression of PrPC starts with the expression of CD25 marker and most of DN3 cells, which correlate to the stage of development arrest, are PrPC+. The loss of the DN4 population (as mentioned above) in TgA20 mice could be the consequence of the blockade of the DN3 PrPC high cells at this stage of development. Interestingly, we observed that γδ+ T cells, NK and B cells do not express PrPC (data not shown).

FIGURE 5.

Expression of PrPC. A, Thymocytes from TgA20 (solid lines) and WT (gray histograms) mice were stained and cells were gated on the DN, DP, CD4+, or CD8+ SP compartments. The expression of PrP (SAF 32 mAb) in each subset was analyzed. B, Cells were gated for CD4, CD8, CD11c, CD19, NK1.1, and γδTCR and PrP expression was investigated in WT and TgA20 within the DN1, DN2, DN3, and DN4 subpopulations. Staining control is shown as dotted line.

FIGURE 5.

Expression of PrPC. A, Thymocytes from TgA20 (solid lines) and WT (gray histograms) mice were stained and cells were gated on the DN, DP, CD4+, or CD8+ SP compartments. The expression of PrP (SAF 32 mAb) in each subset was analyzed. B, Cells were gated for CD4, CD8, CD11c, CD19, NK1.1, and γδTCR and PrP expression was investigated in WT and TgA20 within the DN1, DN2, DN3, and DN4 subpopulations. Staining control is shown as dotted line.

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Because SP thymocytes were heterogeneous in regard to their PrPC expression, we re-examined in more detail the phenotype of SP-PrPC-positive vs SP-PrPC-negative cells and found that most SP thymocytes do not express at the same time high levels of PrPC and αβTCR: in the CD4+TCRhigh population, <25% expressed high levels of PrPC, whereas 76% were PrPlow/−. Similarly, in the CD8+TCRhigh thymocytes only 9.9% were PrPC+. Conversely, in the CD4+ and CD8+TCRlow cells, 70 and 53% of the cells expressed high levels of PrPC, respectively (data not shown). This may be explained by two not mutually exclusive hypotheses. PrPC+ DP thymocytes may down-regulate their PrPC, giving rise to PrPint or PrPlow SP cells expressing high levels of αβTCR as in WT. Or some immature DN thymocytes may further proceed with their development, generating some SP progeny before the expression of PrPC could have started and blocked the appearance of the αβTCR.

Lymphoid development in thymus relies on constant interactions with epithelial cells, which have been reported to express PrPC (42). To evaluate whether the alterations we observed were due to PrPC+ epithelial cells or to lymphocytes themselves, we performed hemopoietic cell reconstitution experiments in lethally irradiated hosts and analyzed thymic reconstitution. As shown in Table I, the retardation in T cell development was detected only when cells of the hemopoietic lineage were from TgA20 BM. In this case, the thymic cellularity was drastically reduced (60- to 70-fold) with an increase of thymocytes at the DN stage. However, cells able to pass the DP stage became mature thymocytes at a normal cell ratio. We observed the same phenotype in competitive reconstitution experiments using a mixture of TgA20 and C57BL/6 BM precursor cells, differing in the allogenic markers Ly5.2 and Ly5.1, respectively (Table II). When BM cells were all from C57BL/6, the partition of Ly5.2 cells followed a normal distribution (DN/DP = 2/90), whereas this partition was altered when Ly5.2 cells were from TgA20 mice (DN/DP = 15/70) and similar to that observed in chimeras with a TgA20 donor (see Table I). Moreover, in the latter experiments, only BM TgA20-derived cells gave rise to high percentages of γδ T cells. Those results demonstrate that the defect is carried by TgA20 thymocytes themselves.

Table I.

The defect in thymocyte maturation is cell autonomous in TgA20 mice

BM ChimerasCellularitya% Thymocyte Subpopulationsb
Donor Recipient  DN DP CD4 CD8 
C57BL/6 C57BL/6 147 ± 5 90 
C57BL/6 TgA20 121 ± 20 90 
TgA20 TgA20 2 ± 1 15 78 
TgA20 C57BL/6 2 ± 1 36 56 
BM ChimerasCellularitya% Thymocyte Subpopulationsb
Donor Recipient  DN DP CD4 CD8 
C57BL/6 C57BL/6 147 ± 5 90 
C57BL/6 TgA20 121 ± 20 90 
TgA20 TgA20 2 ± 1 15 78 
TgA20 C57BL/6 2 ± 1 36 56 
a

Cellularity is expressed as 106 cells and is a mean of three independent reconstituted mice.

b

Partition of thymocyte subpopulations is a representative experiment of three independent ones.

Table II.

Competitive reconstitution with BM precursor cells

Mixed ChimerasaThymus% among Ly5.2+ Cellsb
Cellularityc % Ly5.2 DN DP CD4 CD8 γδT cellsd 
C57BL/6 [LY5.1]/C57BL/6 [Ly5.2] 158 ± 18 30 ± 10 90 
C57BL/6 [Ly5.1]/TgA20 [Ly5.2] 85 ± 16 2 ± 1 15 70 41 
Mixed ChimerasaThymus% among Ly5.2+ Cellsb
Cellularityc % Ly5.2 DN DP CD4 CD8 γδT cellsd 
C57BL/6 [LY5.1]/C57BL/6 [Ly5.2] 158 ± 18 30 ± 10 90 
C57BL/6 [Ly5.1]/TgA20 [Ly5.2] 85 ± 16 2 ± 1 15 70 41 
a

Mixed chimeras were performed in C57BL/6 Ly5.1 recipients by injections of 50% Ly5.1 C57BL/6 mixed with either 50% Ly5.2 C57BL/6 or TgA20 BM precursor cells.

b

Partition of thymocyte subpopulations is a representative experiment of three independent ones.

c

Cellularity is expressed as 106 cells and is a mean of three independent reconstituted mice.

d

Represent cells bearing a γδ TCR among CD3 high thymocytes.

Because it has been proposed that PrPC binds and chelates copper (4, 43), we investigated whether PrPC overexpression is linked to an increase in copper content. Compared with WT mice, TgA20 mice displayed an increase of copper in the brain and thymus (Fig. 6). Such an increase was not detected in the spleen and no difference was observed for zinc content in the different tissues. Total copper was measured, i.e., intra- and extracellular copper, free or bound to cell surface proteins. Knowing that PrPC chelates copper and that, in TgA20, PrPC is overexpressed, this suggests that the amount of free copper is decreased, creating a deficit in this cation which is known to play a crucial role in the regulation of the redox balance. To test this hypothesis, we raised the pool of free copper by a dietary copper supplementation. Whereas the increase of cells is only 1.7-fold with WT mice (189 vs 110.4 million cells), we observed a 9.7-fold increase in TgA20 mice (33 vs 3.4 million cells) (Fig. 7,A). To determine whether this dramatic increase is associated with a resumption of thymocyte maturation, we analyzed the partition of the DN vs DP compartments of control or supplemented TgA20 and WT mice (Fig. 7 B). With copper-supplemented TgA20 mice, we systematically observed an increase in the percentage of DP cells and a decrease in DN cells compared with nonsupplemented mice. The SP subpopulations remained more or less stable. With WT-supplemented mice, no significant changes were noted as compared with control mice. Thus, copper-supplemented TgA20 mice show a phenotype that comes closer to that of WT mice. γδ TCR+ cells were also affected by copper supplementation, decreasing from 26.8 to 21.8% (data not shown). These results suggest that the partial blockage observed in TgA20 could be linked to a deficit in free copper since thymic differentiation restarts after copper addition.

FIGURE 6.

Metal concentration in brain, thymus and spleen. Copper or zinc concentration measurements were performed by electrothermal atomic absorption spectrophotometry. ∗, Correspond to p < 0.05.

FIGURE 6.

Metal concentration in brain, thymus and spleen. Copper or zinc concentration measurements were performed by electrothermal atomic absorption spectrophotometry. ∗, Correspond to p < 0.05.

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FIGURE 7.

Partial reversion of the TgA20 phenotype under copper supplementation. A, Thymus cellularity of TgA20 and WT 4-wk-old mice either supplemented or not with copper is expressed in millions cells. Each plot represents an individual mouse. The value of the mean cellularity is indicated and delineated by a horizontal line. B, FACS analysis of the thymocytes partition among the DN, DP, and SP compartments in TgA20 and WT mice. The profiles are representative of five mice analyzed for control groups and or nine mice analyzed for supplemented groups.

FIGURE 7.

Partial reversion of the TgA20 phenotype under copper supplementation. A, Thymus cellularity of TgA20 and WT 4-wk-old mice either supplemented or not with copper is expressed in millions cells. Each plot represents an individual mouse. The value of the mean cellularity is indicated and delineated by a horizontal line. B, FACS analysis of the thymocytes partition among the DN, DP, and SP compartments in TgA20 and WT mice. The profiles are representative of five mice analyzed for control groups and or nine mice analyzed for supplemented groups.

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In this study, we have shown that in TgA20, overexpression of PrPC in the thymus hampers the different checkpoints essential to the complete differentiation pathway of αβ T cells. The observed effect is cell autonomous and involves the majority of immature thymocytes developmentally blocked at the DN3 stage and the DP to SP transition, as demonstrated by the delay in CD5 expression and CD25 down-regulation in DP and SP compartments. It is noteworthy that DN3 and DP thymocytes are cells that express the highest level of PrPC. At the end of the thymic differentiation, a high level of αβ TCR is normally observed at the surface of thymocytes but the presence of PrPC seems to exclude a correct expression of the receptor. In Tg33 mice, the consequences of PrPC overexpression on T cell differentiation appear from 10.5 wk and, at this time, the overexpression also leads to a reduced thymus size and cellularity and to a reduction of DP cells. We also observed an increase in the percentage of ISP cells in Tg33 mice, reinforcing the idea that overexpression of PrPC alters, in this second model of mouse also, the differentiation of αβ T cells.

Keeping in mind that in the brain, PrPC has been implicated in signal transduction and/or oxidative stress regulation (11, 39), we explored these two functions of PrPC to explain the profound alterations of αβ T cell development.

Although in a normal thymus, more CD4+ than CD8+ cells are generated after CD4/CD8 lineage commitment (44), this difference is not observed in TgA20 mice, essentially due to a reduction in the CD4+ subset. For all of the markers analyzed in Fig. 4, the CD4+ compartment is the most affected: about one-half the CD4+ population has an altered phenotype (CD25+, CD5, TCRβ, and CD69), whereas CD8+ cells present a phenotype very close to that of the WT mice. It is commonly admitted that positive selection and CD4/CD8 T lineage commitment take place at DP stage of development. In the TgA20 mice, DP cells present a defect in CD5 expression, reflecting a less advanced maturation stage. This defect is amplified in the CD4+ population after commitment. It has been proposed that a strong activation of p56lck, a Src family tyrosine kinase, leads to CD4+ cells differentiation, whereas weak or no signaling through this kinase leads to CD8+ cells differentiation (45, 46, 47). The maturation delay observed in the TgA20 DP cells could induce an inability of these cells to correctly transduce this signal. Consequently, the low signal remaining would not be sufficient to lead to a correct CD4 commitment without impairing the CD8 progeny. To reinforce the putative implication of lck in the blockade, it is interesting to note that in lck−/− mice, the thymocytes are also blocked at the DN to DP transition (48). Furthermore, it has been proposed that the PrPC, attached to the outer cell membrane via its GPI-anchor, may act inside the cell by transducing a signal through the fyn tyrosine kinases (11, 12). Finally, it has been recently shown that cross-linking of PrPC leads to recruitment of Thy-1 and of molecules important for T cell activation, i.e., TCR/CD3, Fyn, Lck, and linker for activation of T cell (49).

Fyn is known to be implicated in a lck/fyn-Zap70/syk tyrosine kinase cascade initiated by a CD3-associated TCR or pre-TCR complex (50). The pre-TCR expression is followed by an activation cascade leading to the DN3 to DN4 transition. pTα−/− mice present a phenotype similar to the one observed in TgA20 (51, 52) in which we have seen a block in thymocyte development at this transition. Therefore, in TgA20 mice, the defect in signal transduction may also be driven at the pre-TCR stage. It is noteworthy that at this stage, the cells express high amounts of PrPC. The overexpression of PrPC could function as a negative regulator of αβ T cell development, leading to signals interfering with a correct DN to DP transition and DP to SP maturation. Interestingly, a delay (increase in ISP and decrease in DP cells) in the developmental process of thymocytes is also observed in Tg33 mice.

In the view of our results, we favor a second hypothesis, not mutually exclusive with the first one, that overexpression of PrPC could generate an antioxidant state, altering αβT lymphocyte maturation. It is known that copper deprivation impairs both innate and adaptative immunity. For example, copper deficiency reduces the level of IL-2 mRNA and the secretion of this cytokine (30, 53). Also, it has been clearly shown that murine PrPC might serve as a copper chelating or buffering agent in the outer side of the cell membrane (39, 43). In the thymus, circulating copper and/or copper from exocytosis could be chelated by PrPC at the cell surface and lead to a reduction of free copper, creating an antioxidant environment that is harmful to the αβ T cell development. This hypothesis is supported by the observation that addition of copper partially reversed the phenotype found in the TgA20 mice. This also correlates with the fact that addition of antioxidant agents to fetal thymus organ cultures arrests the αβ, but not γδ T cell development in a dose-dependent manner (34). Of particular note is the fact that mature TgA20 thymocytes express less PrPC than their Tg33 counterparts, but in addition to thymocytes, their epithelial cells also express PrPC, possibly creating a more antioxidative environment for T cell development. This could explain why the defects due to PrPC overexpression are observed earlier in TgA20 than in Tg33. Thus, little change in the global redox balance in the thymus has dramatic consequences on the αβ T cell development.

In the various invalidated gene models where αβ T cell development is impaired (pTα−/−, TCRβ−/−, lck−/−, fyn−/−), an increase in γδ T cell numbers was never observed (50). In TgA20 mice (and to a lesser extent in Tg33 mice), the percentage and the absolute number of γδ+ thymocytes increase. As already shown by Ivanov et al. (34), αβ and γδ T cells differ in their ability to adapt to the antioxidant environment. We propose that the great number of PrP molecules at the surface of thymus cells in overexpressing mice (either from Tga20 or Tg33 mice) promotes such an antioxidant environment and thus favors γδ T cell development and/or survival. Moreover, since γδ T cells do not express PrPC at their surface, they are therefore protected from the deleterious effects of PrPC-copper complexes.

The alteration of αβ T cell development and the expansion of γδ T cells observed in this model of PrPC overexpression could be explained by the interference of PrPC with signaling pathways but more probably by the antioxidant effect due to its copper-chelating properties. Although the absence of PrPC has no direct consequence on the differentiation of T cells, we show that overexpression of this protein has a major impact on αβ and γδ T cell development. As the immune system is destabilized, overexpression of PrPC may have a dramatic effect in case of an infection by the scrapie form.

We thank Dr. Candéias and Dr. Viret for helpful discussion; A.-M. Laharie, V. Collin, N. Brient, E. Borel, and L. Pouyet for technical assistance.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by specific grants from the Groupement d’Intérêt Scientifique-Prion (Grants A42 and A108), the Commissariat à l’Energie Atomique ToxNuc Program (Grant 13-10), and the European Union, Project Number QLK5-CT-2002-01044. V.A.-A. is a recipient of a GIS-Prion Fellowship.

5

Abbreviations used in this paper: PrPC, cellular prion protein; PrP, prion protein; DN, double positive; DP, double positive; SP, single positive; WT, wild type; BM, bone marrow; ISP, immature SP; PrPsc, scrapie pathological PrP.

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