We initially broadly defined the targets of CD4 T cell reactivity to PLP in “non-tolerant” Plp^KO BL/6 mice through re-stimulating lymph node T cells from immunized mice with a set of overlapping 25-mer peptides corresponding to the entire PLP protein (Figure 1A) (8). The PLPko mouse used in our study has been characterized extensively with regards to a verification of the absence of PLP protein or any truncated version thereof (20). Overlapping 12-mer peptides spanning these regions were then used to precisely fine-map the respective I-A^b-restricted epitopes (Figure 1B). This revealed three core 9-mer epitopes spanning amino acids 11–19, 174–182, and 240–248, in the following referred to as epitopes #1, #2, and #3, respectively. The experimental epitope definition was retrospectively combined with an in silico prediction of T cell epitopes using the Immune Epitope Database and Analysis Resource (IEDB) (21, 22). The IEDB algorithm predicts and ranks the relative binding strengths of all 15-mer peptides that can be generated from a given protein. For PLP, the seven 15-mer peptides containing epitope #3 were among the top eight predicted I-A^b binders, and all of the 15-mers harboring epitope #1 were ranked between positions 10 and 20 (Figure S1 in Supplementary Material). Epitope #2-containing 15-mers had the weakest binding scores and ranked between positions 33 and 57. Consistent with this relative ranking, an in silico prediction of MHC-binding affinities using the SSM-align algorithm (23) yielded mean IC₅₀ values of 168 ± 61 nM for epitope #3-containing peptides and 715 ± 262 or 1,533 ± 498 nM for peptides containing epitopes #1 or #2, respectively.

Compared to the pronounced CD4 T cell responses to all three epitopes in Plp^KO mice, there was essentially no recall response to epitopes #1 and #3 with cells from immunized Plp^WT mice (Figure 1C). However, a robust response to epitope #2 was observed, although the magnitude of this response was somewhat attenuated as compared to Plp^KO mice. To assess how the absence or presence of CD4 T cell responsiveness to these PLP epitopes correlated with EAE susceptibility, we immunized Plp^WT mice with epitopes #1, #2, or #3 according to the EAE protocol. Whereas “EAE-immunization” with epitope #1 and epitope #3 failed to promote disease symptoms, epitope #2 elicited EAE with a typical acute disease course, that is, onset of symptoms at around day 8, followed by a disease peak at days 14–16 and subsequent remission (Figure 1D, and data not shown).

Together, these observations reveal a differential impact of self-tolerance on CD4 T cell responses to three distinct epitopes of PLP in BL/6 mice. Tolerance was “leaky” for the epitope with the lowest predicted MHC II-binding capacity. Moreover, the absence or presence of recall responsiveness to a given epitope tightly correlated with resistance or susceptibility to EAE, respectively.

In order to follow the fate of epitope #2-specific CD4 T cells and characterize the apparent “failure” of tolerance induction to this epitope, we generated transgenic mice (TCR-PLP2) expressing a PLP_172–183-specific TCR that was derived from immunized Plp^KO mice. In TCR-PLP2 Plp^KO mice, i.e., in the absence of cognate self-antigen, there was a pronounced skewing toward the CD4 lineage in the thymus, and the vast majority of CD4 single-positive (SP) thymocytes expressed the transgenic TCR chains (Vα2 and Vβ14) (Figure 2A). A very small fraction of Vα2⁺Vβ14⁺ CD4 SP cells expressed Foxp3. Importantly, none of these parameters was altered in TCR-PLP2 Plp^WT mice, indicating that PLP epitope #2-specific CD4 T cells were not subject to central tolerance induction (Figure 2A). Also in the periphery, neither the frequency of CD4 T cells nor their expression of Vα2⁺Vβ14⁺ was different between TCR-PLP2 mice on Plp-deficient or -sufficient background, suggesting that epitope #2-specific T cells in both settings persisted as naïve cells (Figure 2B). Consistent with this, purified CD4 T cells from TCR-PLP2 Plp^WT displayed an identical in vitro proliferative response to stimulation with titrated amounts of PLP_172–183 as cells from TCR-PLP2 Plp^KO controls (Figure 2C). Adoptively transferred naïve TCR-PLP2 cells from Plp^KO mice persisted in a non-dividing state in Plp^WT recipients, whereas cells from both TCR-PLP2 Plp^KO or TCR-PLP2 Plp^WT origin proliferated upon challenge with cognate peptide, irrespective of being parked in a Plp-deficient or -sufficient recipient (Figure 2D). These findings further substantiated the idea that PLP epitope #2 of natural origin is not sufficiently processed and presented from physiological sources to become “visible” to peripheral CD4 T cells. However, “EAE-immunization” with peptide PLP_172–183 elicited EAE in TCR-PLP2 Plp^WT mice, suggesting that these TCR transgenic cells can turn into pathogenic effectors upon appropriate stimulation and recognize the physiologically processed epitope (Figure 2E).

Thus, PLP_172–183-specific TCR transgenic CD4 T cells with encephalitogenic potential were not subject to tolerance induction. Instead, they persisted in a state of “ignorance” that was broken upon immunogenic delivery of cognate antigen, in line with the capacity of PLP_172–183 immunization to elicit EAE in a polyclonal setting.

The expression of an endogenously rearranged TCRα chain together with the transgenic TCR elements may result in a dual specificity and thereby complicate the interpretation of TCR instructed cell fate decisions. For instance, the emergence of a small population of Foxp3⁺ cells among Vα2⁺Vβ14⁺ cells in both thymus and periphery of TCR-PLP2 mice was PLP independent, and thus likely to reflect a “cryptic autoreactivity” conveyed by endogenous TCR rearrangements. To assess the consequences of excluding endogenous TCR rearrangements, we generated Rag1-deficient TCR-PLP2 mice. Irrespective of an intact Plp gene, this resulted in the virtual absence of Foxp3⁺ cells from thymus and periphery. Importantly, as in Rag1^+/+ mice, the frequency and absolute number TCR-PLP2⁺ cells was indistinguishable between Plp-deficient or –sufficient mice, confirming that their cognate antigen PLP_172–183 is essentially “invisible” to these cells (Figures 3A,B). However, in contrast to mice on Rag1^+/+ background, PLP-sufficient TCR-PLP2 Rag1^–/– mice developed “spontaneous” EAE starting at around 5 weeks of age (Figures 3C,D).

These observations confirmed that TCR-PLP2⁺ CD4 T cells can be pathogenic, yet do not encounter cognate antigen during thymic differentiation or in the periphery, so that they initially remain in a state of “ignorance.” However, in a truly monoclonal setting their latent autoimmune potential can with increasing age be “spontaneously” unleashed, possibly facilitated by the lack of T_reg cells.

In contrast to our observations with the “EAE-susceptibility-associated” epitope #2, BL/6 mice did not display a CD4 T cell recall response to epitope #1 and were resistant to EAE induction by immunization with this peptide (see Figure 1). In order to characterize the mode of CD4 T cell tolerance induction to the “EAE-resistance-associated” epitope #1, we generated a PLP_11–19-specific TCR transgenic mouse strain (TCR-PLP1). Compared to TCR-PLP1 Plp^KO controls, the frequency and number of CD4 SP cells in the thymus of TCR-PLP1 Plp^WT mice were strongly diminished (Figure 4A). Moreover, whereas in PLP-deficient mice, around 85% of CD4 SP cells expressed both transgenic TCR chains, only about 50% of CD4 SP cells were Vα3.2⁺Vβ6⁺ in PLP-sufficient mice. Among the remaining TCR-PLP1⁺ CD4 SP cells in PLP^WT mice, an elevated frequency of cells was Foxp3⁺, whereby in terms of absolute numbers, this reflected an only moderate, yet significant, increase (Figure 4A). In peripheral lymphoid organs of TCR-PLP1 Plp^WT mice, the proportion of CD4⁺ T cells was reduced by about fourfold when compared to Plp^KO controls (Figure 4B). Whereas in PLP-deficient mice, around 75% of peripheral CD4 T cells expressed the transgenic TCR, less than 10% of peripheral CD4⁺ T cells did so in TCR-PLP1 Plp^WT mice. TCR-PLP1⁺ CD4 T cells contained only a minute fraction of Foxp3⁺ cells in PLP-deficient mice, while Foxp3⁺ cells constituted more than a third of TCR-PLP1⁺ cells in Plp^WT mice (Figure 4B).

To ask whether the segregation of peripheral TCR-PLP1⁺ cells in PLP-sufficient mice into Foxp3⁺ and Foxp3^– cells reflected the co-existence of T_reg cells and bona fide naïve cells, we assessed the functional properties of both subsets. Purified CD25⁺ cells indeed suppressed the proliferation of naïve TCR-PLP1 T_conv cells in an antigen-specific manner (Figure 4C). However, whereas naïve TCR-PLP1 cells from PLP-deficient donors vigorously proliferated upon adoptive transfer into Plp^WT recipients, illustrating that I-A^b/PLP_11–19 ligands of physiological origin are constitutively “visible” in peripheral lymphoid tissues, co-transferred CD25^– TCR-PLP1⁺ cells from TCR-PLP1 Plp^WT mice did not proliferate, indicating that these cells were in a cell intrinsic refractory state of anergy (Figure 4D).

We did not observe spontaneous EAE in TCR-PLP1 Plp^WT mice. In order to assess the “autoimmune potential” of TCR-PLP1 CD4 T cells, we generated TCR-PLP1 Rag1^–/– Plp^KO mice. Monoclonal TCR-PLP1 CD4 T cells from these donors were differentiated in vitro into Th1 or Th17 effectors and subsequently transferred into Rag1^–/– Plp^WT recipients, revealing that both Th1- or Th17-polarized cells induced EAE, albeit with a slightly lower maximal disease score in the case of Th1 cells (Figure 4E).

In sum, these data showed that PLP epitope #1-specific CD4 T cells bear autoimmune potential, which was at least in part harnessed through tolerance induction during intrathymic T cell differentiation. The predominant mode of central tolerance induction among TCR-PLP1 CD4 T cells appeared to be clonal deletion. At the same time, a fraction of cells differentiated into T_reg cells and efficiently seeded peripheral lymphoid organs, were they co-existed with anergic Foxp3^–CD25^– TCR-PLP1 cells.

We previously reported that Plp^WT → Plp^KO thymus chimeras, where TECs are the sole potential source of tolerizing antigen, recapitulated the “operational tolerance” of Plp^WT mice, as evident from a similar reduction of the CD4 T cell recall response after immunization compared to Plp^KO mice (8). This indicated that PLP expression in TECs was sufficient for CD4 T cell tolerance. Given that PLP transcripts are also detectable in dendritic cells (DCs), albeit at substantially lower levels (Figure 5A), we wondered whether TEC-derived PLP was in fact essential for central tolerance, or whether there was a degree of redundancy with other potential intra- or extrathymic sources of tolerizing antigen. To address this, we generated mice with a conditional deletion of the Plp gene in TECs (Foxn1-Cre Plp^floxed), in the following referred to as Plp^ΔTEC. Strikingly, all manifestations of central tolerance in the TCR-PLP1 system were abolished in Plp^ΔTEC mice (Figures 5B,C). Thus, both the frequency of CD4 SP cells as well as the proportion of Vα3.2⁺Vβ6⁺ among CD4 SP cells were indistinguishable from those observed in the complete absence of PLP in Plp^KO mice. In the same vein, the frequency of Foxp3⁺ cells among TCR-PLP1 CD4 SP cells of Plp^ΔTEC mice was identical to that in “antigen-free” Plp^KO mice.

The “promiscuous” transcription of peripheral antigens in the thymus is mostly confined to “mature” medullary thymic epithelial cells (mTECs) expressing high levels of MHCII, and this reflects the predominant expression of the transcriptional activator Aire in this cell type. PLP’s intrathymic expression pattern is rather untypical for a promiscuously expressed tissue-restricted antigen in that it is also detectable in “immature” (MHCII^lo) mTECs and at comparable levels also in cortical thymic epithelial cells (cTECs) (Figure 5A). Since MHCII^lo mTECs and cTECs barely, or not at all, express Aire, respectively, we hypothesized that central tolerance to PLP was Aire independent. However, both negative selection and T_reg induction were abrogated in Aire-deficient TCR-PLP1 Plp^WT mice, so that the thymocyte composition in these mice phenocopied that in TCR-PLP1 Plp^KO and TCR-PLP1 Plp^ΔTEC mice (Figure 5D).

Together, these data showed that PLP expression in TECs was necessary and sufficient for central tolerance induction in PLP epitope #1-specific CD4 T cells. Intriguingly, although PLP transcripts were not confined to Aire-expressing mTEC^hi cells, intrathymic tolerance induction to PLP depended upon Aire.

Self-antigens that are expressed in TECs may be presented to developing CD4 T cells via two distinct, yet mutually not exclusive routes (24). On the one hand, tolerogenic encounter of such antigens by CD4 T cells may depend upon “antigen handover” and presentation by thymic DCs. On the other hand, mTECs, or TECs in general, may autonomously present endogenously expressed antigen to CD4 T cells via unconventional MHC class II-loading pathways (25).

Two experimental systems were employed to address this issue in the TCR-PLP1 model. First, we generated TCR-PLP1 Plp^WT mice lacking DCs as a result of diphtheria toxin expression in CD11c⁺ cells (“ΔDC” mice) (26). Despite the lack of DCs, the thymic phenotype of these mice was essentially identical to that of DC-sufficient TCR-PLP1 Plp^WT mice (Figures 6A,B). Second, we generated TCR-PLP1 mice with an impaired capacity to directly present antigen on mTECs as a consequence of mTEC-specific knock-down of MHC class II (“C2TAkd” mice) (27). All manifestations of central tolerance induction to PLP epitope #1 were significantly attenuated in these mice. Specifically, the proportion of CD4 SP cells, the percentage of Vα3.2⁺Vβ6⁺ among CD4 SP cells and the relative abundance of TCR-PLP1⁺ Foxp3⁺ CD4 SP cells were all partially “rescued,” i.e., they ranged in between TCR-PLP1 Plp^WT and TCR-PLP1 Plp^KO mice (Figure 6C).

Collectively, these findings documented a non-redundant role of mTECs for central tolerance induction to PLP epitope #1 through direct presentation of endogenously derived antigen. Intrathymic presentation by DCs, if occurring at all, was dispensable for central tolerance.

Although the conditional deletion of PLP in TECs abolished central tolerance to PLP epitope #1, neither Plp^ΔTEC mice nor TCR-PLP1 transgenic Plp^ΔTEC mice developed spontaneous EAE (data not shown). Even the active immunization of these mice with the epitope #1-containing peptide PLP_11–20 according to the “EAE protocol” failed to promote disease symptoms (data not shown), suggesting that peripheral tolerance mechanisms were able to compensate for the absence of intrathymic censorship.

Indeed, the CD4 T cell compartment in secondary lymphoid tissues of TCR-PLP1 Plp^ΔTEC mice exhibited clear indications of peripheral antigen encounter. Thus, despite an identical thymocyte composition as in TCR-PLP1 Plp^KO mice (see Figures 5B,C), there was a marked reduction in the overall size of the peripheral CD4 T cell population in TCR-PLP1 Plp^ΔTEC mice (Figure 7A). Although this resembled a similar CD4 T cell reduction in TCR-PLP1 Plp^WT mice, Plp^ΔTEC mice had a substantially higher percentage of TCR-PLP1⁺ cells among peripheral CD4 T cells (47.9 ± 5.1 versus 8.9 ± 0.8%; P < 0.001). The proportion of Foxp3⁺ T_reg cells among TCR-PLP1⁺ CD4 T cells in Plp^ΔTEC mice was as low as in fully PLP-deficient mice, suggesting that PLP expression in TECs was crucial for the establishment of the prominent peripheral TCR-PLP1⁺Foxp3⁺ T_reg population in TCR-PLP1 Plp^WT mice. By contrast, the CD25^– subset of TCR-PLP1⁺ cells from Plp^ΔTEC mice functionally resembled the corresponding population in Plp^WT mice with respect to being refractory to antigenic stimulation in vivo (Figure 7B), indicating that anergy induction occurred independent of thymic PLP encounter. Consistent with this, the anergy marker FR4 was similarly elevated on Foxp3^–CD25^–TCR-PLP1⁺ cells from both Plp^ΔTEC and Plp^WT mice (Figure S2 in Supplementary Material). These data showed that in the absence of PLP expression in TECs, peripheral tolerance toward PLP epitope #1 appeared to operate via deletional mechanisms and anergy induction rather than T_reg conversion.

Ablation of thymic PLP expression in Plp^ΔTEC mice revealed that “downstream” of the thymus, peripheral antigen encounter could shape the fate of TCR-PLP1 CD4 T cells. To ask whether presentation of I-A^b/PLP_11–19 ligands in the periphery was DC dependent, we transferred CFSE-labeled TCR-PLP1 cells into DC-sufficient or DC-deficient Plp^WT mice. Whereas transferred cells vigorously proliferated in control recipients, they failed to do so in ΔDC mice, indicating that antigen presentation by DCs was crucially involved (Figure 7C).

To address whether the display of PLP epitope #1 by peripheral DCs reflected the presentation of endogenously derived PLP or required the uptake and presentation of PLP from a non-hematopoietic source, we performed analogous experiments with bone marrow (BM) chimeric recipients (Figure 7D). Whereas TCR-PLP1 cells underwent proliferation when transferred into MHCII^+/+ → Plp^WT control chimeras, they remained quiescent in MHCII^–/– → Plp^WT chimeras, consistent with a requirement for antigen presentation by DCs. Importantly, transferred cells likewise failed to proliferate in MHCII^+/+ → Plp^KO chimeras, establishing that PLP from a non-hematopoietic source was crucially required.

Considering that in Plp^WT mice, PLP was concomitantly presented intrathymically by mTECs as well as in the periphery by DCs, we wondered in how far the peripheral composition and functionality of TCR-PLP1⁺ cells was shaped by thymic output versus peripheral tolerance induction. The striking phenotypic similarity between DC-sufficient and DC-deficient TCR-PLP1 Plp^WT mice (compare Figures 6A,B) also applied to the periphery (Figure 7A). Specifically, total CD4 cells were reduced to the same extent, and the comparably small populations of TCR-PLP1⁺ cells in both genotypes similarly segregated into Foxp3⁺ T_regs and Foxp3^–CD25^– cells, suggesting that the overall composition of the TCR-PLP1⁺ CD4 T cell pool was mostly shaped by thymic output. Interestingly, though, CD25^– cells from ΔDC mice proliferated upon adoptive transfer into Plp^WT mice (Figure 7B), indicating that the anergic state of these cells in DC-sufficient mice is induced and/or maintained by peripheral antigen encounter on DCs.

Our findings suggested that the lack of thymic negative selection in TCR-PLP1 Plp^ΔTEC mice could be largely compensated by DC-mediated peripheral deletion and anergy induction, while the “normal” formation of a PLP-specific peripheral T_reg compartment appeared to depend on thymic T_reg induction by mTECs. Given this differential degree of redundancy between thymic and peripheral antigen encounter with respect to recessive versus dominant tolerance mechanisms (deletion and anergy versus T_reg induction, respectively), we sought evidence for a defect in dominant tolerance to PLP in the polyclonal repertoire of non-TCR transgenic Plp^ΔTEC mice. To do so, we adoptively transferred naïve TCR-PLP1 cells into Plp^ΔTEC or Plp^WT mice simultaneously to “EAE-immunization” with PLP_11–20. This resulted in the development of EAE in both genotypes. However, the disease course was more severe in mice lacking thymic PLP expression, consistent with a “less robust” state of tolerance, possibly owing to lack or diminution of PLP-specific T_reg cells (Figure 7E).

MHC class II^–/– (H2-Ab1^–/–) (56), C2TAkd (27), ΔDC (26), Plp^KO (20), and Aire^–/– (57) mice have been described previously. The gene encoding for PLP is located on the X chromosome. We did not observe any differences in the composition and phenotype of lymphoid compartments between TCR-PLP1 or TCR-PLP2 transgenic PLP-sufficient females (Plp^+/+) and males (Plp^+/y) or between PLP-deficient females (Plp^–/–) or males (Plp^–/y). Therefore, female and male mice were included into the PLP-sufficient or -deficient groups and their genotype indicated as Plp^WT or Plp^KO. Plp^ΔTEC mice were obtained by crossing Foxn1-Cre (58) mice to mice carrying a conditional Plp allele in which exon 3 is flanked by loxP sites (Plp^fl) (59). C57BL/6 mice were purchased from Charles River. Mice were maintained under specific pathogen-free conditions in individually ventilated cages. All phenotypic analyses were performed in young adult mice at 4–5 weeks of age. Animal studies and procedures were approved by local authorities (Az 7-08 and 142-13).

To generate TCR transgenic mice, we used rearranged V(D)J regions of TCRs from PLP epitope #1 or #2-specific T cell hybridomas (D9-119-2 or A43-11-5, respectively). Hybridoma D9-119-2 (TCR-PLP1) was obtained by immunizing Plp^KO mice with peptide PLP_1–25, repeated re-stimulation of draining LN cells with PLP_9–20 and subsequent fusion with BW5147 cells. Its TCRα chain contains a rearrangement of Trav9d-3.201 (Vα3.2) and Traj34-201, the TCRβ chains harbors a Trbv19-201 (Vβ6), Trbd1, and Trbj1-1 rearrangement. Hybridoma A43-11-5 (TCR-PLP2) was obtained by immunizing Plp^KO mice with peptide PLP_160–184, repeated re-stimulation of draining LN cells with PLP_172–183, and subsequent fusion with BW5147 cells. Its TCRα chain contains a Trav14-1-201 (Vα2) and Traj23-201 rearrangement, the TCRβ chain harbors a Trbv31-01 (Vβ14), Trbd1, and Trbj1-1 rearrangement. TCRα and TCRβ rearrangements from the D9-119-2 and A43-11-5 hybridomas were PCR amplified and cloned into the cassette vectors pTαcass and pTβcass as described before (60). Transgenic mice were generated by injection of linearized DNA into pronuclei of C57BL/6 zygotes.

FITC-conjugated monoclonal antibodies (mAbs) to Vα3.2 (RR3-16); PE-conjugated mAbs to Vβ6 (RR4-7), Vα2 (B20.1), and Vβ14 (14-2); CyChrome-conjugated mAbs to CD8 (53-6.7), PE-Cy7-conjugated streptavidin and mAbs to CD25 (PC61) and FR4 (12A5); APC-Cy7-conjugated mAbs to CD4 (GK1.5); biotin-conjugated mAbs to CD8 (53-6.7), CD4 (GK 1.5), B220 (RA3-6B2) and Vα2 (B20.1); APC-conjugated mAbs to Foxp3 (FJK-16s), CD45.2 (104); Pacific Blue-conjugated mAbs to CD45.1 (A20) were from BioLegend or eBiosciences, respectively; flow cytometric analyses were performed on a FACSCanto II (BD) using FACSDiva (BD) and FlowJo (Tree Star) software.

Purified human PLP (100% identical to mouse PLP) was prepared as described (7). 25-mer peptides were synthesized in the central core facility of the DKFZ (8). 12-mer peptides were synthesized in the central core facility of the DKFZ or purchased from BioTrend.

The IEDB tool (http://www.iedb.org/) combines various machine learning algorithms for peptide epitope prediction in the context of a given MHC haplotype. The full-length PLP amino acid sequence was analyzed using the IEDB tool in the default “consensus” setting for prediction methods (21, 22). A lower score indicates better binding to MHCII. The theoretical IC₅₀ values were obtained with the SMM-align predictor (23).

Mice were immunized with 100 µg purified PLP or 50 µg peptide emulsified in complete Freund’s adjuvant (volume/volume, 1/1). After 9 days, popliteal and inguinal lymph node cells were harvested and cultured for 72 h in triplicate at a concentration of 4 × 10⁵ cells/well in round-bottomed, 96-well plates in serum-free medium (HL-1; BioWhittaker) in the presence or absence of titrated amounts of peptide or 5 µg/ml of the respective peptide for epitope mapping. Proliferation was measured by incorporation of 3H-thymidine, added for the last 24 h of culture at a concentration of 1 μCi/well. Only mice with comparable control responses to the complete Freund’s adjuvant component PPD were included in the analyses. Proliferation indices were calculated as ³H-Thymidin-incorporation in the presence of specific peptide divided by background proliferation w/o peptide. For direct ex vivo proliferation assays, 4 × 10⁵ T cells and 3 × 10⁴ irradiated (3,000 rad) BM-derived DCs were cultured in the presence of increasing peptide concentrations.

Bone marrow was depleted of B and T cells using biotinylated mAbs to B220, CD8, and CD4 and streptavidin MACS beads (Miltenyi Biotech). Female recipient mice were irradiated with 2 × 550 rad and reconstituted with 5 × 10⁶ BM cells.

TCR-PLP1 or TCR-PLP2 CD4⁺ T cells were isolated from pooled lymph nodes and spleen by cell sorting or by depletion of CD25⁺ cells with biotin-conjugated anti-CD25 and anti-SAV MicroBeads followed by positive enrichment with anti-CD4 microbeads (Miltenyi Biotech). T cells were labeled with CFSE (Life Technologies) and 5 × 10⁶ T cells were injected i.v. into the lateral tail vein of congenic recipient mice. After 4 days, spleen or lymph node cells of recipient mice were analyzed by flow cytometry.

Naive CD4⁺ T cells were prepared from TCR-PLP1 Plp^KO mice using the mouse CD4 T cell isolation kit (Miltenyi Biotech). For Th1 polarization, cells were incubated for 5 days in anti CD3 (2C11) (5 µg/ml) coated 12-well plates (1 × 10⁶ cells/well) in the presence of recombinant IL-2 (5 ng/ml), recombinant IL-12 (10 ng/ml), anti-CD28 mAb (37.51) (3 µg/ml), and anti-IL-4 mAb (11B11) (10 µg/ml). For Th17 polarization, cells were stimulated in the presence of IL-6 (50 ng/ml), IL-1β (10 ng/ml), TGF-β1 (1 ng/ml), anti-CD28 (5 µg/ml), anti-IL-4 (10 µg/ml), anti-IFN-γ (XMG1.2) (10 µg/ml), and anti-IL-2 (JES6-1A2) (10 µg/ml). To verify successful polarization, cells were re-stimulated on day 5 with PMA and ionomycin for 5 h and stained intracellularly for expression of lineage-specific effector cytokines (IFN-γ and IL-17).

TCR-PLP1⁺CD25⁺ cells were sorted from TCR-PLP1 Plp^WT mice (CD45.2) and co-cultured at different ratios with 2 × 10⁴ CFSE-labeled TCR-PLP1⁺CD25^– responder cells (CD45.1) from TCR-PLP1 Plp^KO mice in the presence of 2 × 10⁵ irradiated spleen APCs and 5 µg/ml peptide PLP_9–20. After 4 days, CFSE dilution in gated CD45.1⁺ cells was analyzed by flow cytometry.

For “active EAE” induction, mice were immunized subcutaneously at three sites on the flanks with 200 µg peptide PLP_9–20, PLP_172–183, or PLP_238–249 in a total of 200 µl PBS/CFA emulsion (1:1). Pertussis toxin (400 ng) was given on days 0 and 2. In some experiments, mice received 5 × 10⁶ naïve TCR-PLP1 cells 6 h prior to the immunization. Signs of EAE were scored as follows: 0, none; 0.5, partial loss of tail tonus; 1, limp tail; 1.5, wobbly gait and limb tail; 2.5, paralysis of one hind leg; 3, hind-limb paralysis on both sides; 3.5, hind-limb and partial front limbparalysis; 4, complete hind-limb and forelimb paralysis; 5, moribund. Disease incidence and scores were measured daily. For “passive EAE” induction with Th1 or Th17 cells, Rag1^–/– Plp^WT recipients were injected i.v. with 5 × 10⁶ in vitro polarized TCR-PLP1 cells. Scoring was carried out as described above.

Thymic stromal cells were isolated as described (61). TEC subtypes were classified according to the following surface phenotypes: cTECs, CD45⁻EpCAM⁺Ly51⁺CD80⁻; mTEC^hi, CD45⁻EpCAM⁺Ly51⁻CD80^hiMHCII^hi; mTEC^lo, and CD45⁻EpCAM⁺Ly51⁻CD80^loMHCII^lo. RNA was isolated from sorted cells using the miRNAeasy kit including a DNase digest (Qiagen) and reverse transcribed using the iScript cDNA synthesis kit (Bio-Rad). PCR reactions were performed in duplicates on a CFX96 real-time thermal cycler (Bio-Rad) using the Ssofast EvaGreen Supermix (Bio-Rad). Fluorescence was recorded at the annealing step and relative expression levels were calculated with the comparative Ct method, using HPRT as housekeeping gene for normalization. Primers: Plp-forward, GGGCTTGTTAGAGTGTTGTGC; Plp-reverse, GAAGAAGAAAGAGGCAGTTCCA; Hprt-forward, TGAAGAGCTACTGTAATGATCAGTCAAC; Hprt-reverse, AGCAAGCTTGCAACCTTACCA.

Unless indicated otherwise, statistical significance was assessed using the two-tailed unpaired Student’s t-test with Welch’s correction for unequal variances.