Similar inflammatory DC maturation signatures induced by TNF or Trypanosoma brucei antigens instruct default Th2-cell responses



This article is corrected by:

  1. Errata: Corrections Volume 42, Issue 6, 1649, Article first published online: 8 June 2012


DCs represent the major cell type leading to polarized T-helper (Th) cell responses in vivo. Here, we asked whether the instruction of murine Th2 responses by DCs matured with the proinflammatory cytokine TNF is qualitatively different from maturation by different types of TLR4/MyD88-dependent variant-specific surface glycoproteins (VSGs) of Trypanosoma brucei (T. brucei). The results obtained by analyzing DC surface markers, Notch ligand mRNA, cytokines, asthma, and experimental autoimmune encephalomyelitis (EAE) models as well as performing microarrays indicate that both types of stimuli induce similar inflammatory, semi-mature DC profiles. DCs matured by TNF or VSG treatment expressed a common inflammatory signature of 24 genes correlating with their Th2-polarization capacity. However, the same 24 genes and 4498 additional genes were expressed by DCs treated with LPS that went on to induce Th1 cells. These findings support the concept of a default pathway for Th2-cell induction in DCs matured under suboptimal or inflammatory conditions, independent of the surface receptors and signaling pathways involved. Our data also indicate that quantitative differences in DC maturation might direct Th2- vs Th1-cell responses, since suboptimally matured inflammatory DCs induce default Th2-cell maturation, whereas fully mature DCs induce Th1-cell maturation.


DCs play a fundamental role in the induction of adaptive immune responses as well as in the maintenance of peripheral tolerance 1–3. Through the expression of pattern-recognition receptors (PPRs) such as Toll-like receptors (TLRs), DCs are able to sense a wide array of pathogens and mount an appropriate T-helper (Th) cell response 4. Naïve CD4+ T-cell precursors can differentiate into a variety of Th-cell lineages characterized by the cytokines produced: Th1 cells secrete predominately IFN-γ, Th2 cells release IL-4, IL-5, and IL-13 and Th17 cells typically produce IL-17 5. Although the contribution of DCs for CD4+ Th-cell polarization is under debate 6, several DC-derived mechanisms have been described to significantly direct Th-cell phenotypes. DCs change their maturation status by upregulating surface expression of MHC class II and costimulatory molecules and by producing a defined set of cytokines to optimally induce distinct Th-cell responses 7–9. Due to their immunostimulatory function, DCs are of particular interest in immunotherapy settings, such as cancer therapy and infectious disease intervention 10, 11. Thus, the Th-cell polarizing profile defined by the maturation signature of DCs is of vital interest.

Several membrane markers on DCs and soluble factors secreted by DCs have been described to polarize toward Th2 responses. These include costimulatory molecules such as OX40 12, ICOS-L 13, the Notch family member Jagged-2 14, the cytokine IL-6 15, or arachidonic acid metabolites such as PGE216–18. Much less is known about the factors that induce such Th2-instructing DC. Helminth infections such as Schistosoma mansoni or Nippostrongylus brasiliensis (reviewed in 19, 20) give rise to Th2-cell polarizing DCs, which appear largely immature or only partially mature. Helminth-derived secretory products seem to evoke only mild transcriptional programming and maturation of DCs 21, 22. Interestingly, also proinflammatory cytokines such as TNF or IL-6 23, 24 or tissue disruption induce a similar partially mature phenotype and in the latter case has been attributed to a limited DC activation through the Wnt signaling pathway 25, 26. We and others have demonstrated that DCs conditioned by the inflammatory mediator TNF show a particular maturation phenotype characterized by upregulation of MHC II and costimulatory molecules but no production of cytokines 23, 25, 27. Others suggested that IL-6, induced by low TLR2 and TLR4 triggering, functions as an autocrine/paracrine signaling loop on DCs which itself drives partial maturation of DCs but does not promote Th1-cell responses 24, 28. Thus, partially matured DCs conditioned by inflammatory mediators or low concentrations of TLR ligands have been shown to instruct Th2-cell responses.

However, this raises the question whether endogenous proinflammatory signals and pathogenic signals from parasites trigger the same quality of partial DC maturation leading to Th2-cell responses. Understanding these differences and similarities will be valuable to understand parasitic immune evasion but also for immunotherapy settings where Th2-cell responses act tolerogenic. This has been observed before, especially upon repetitive stimulation of Th2-cell responses characterized by increasing numbers of regulatory IL-10-producing T (Tr1) cells as a tolerance mechanism 29, 30.

Indeed, repetitive injections of TNF-matured DCs prevented the induction of the autoimmune disease EAE mediated at least in part by IL-10+ CD4+ T cells 23. Later, other autoimmune diseases such as thyroiditis and arthritis were also prevented by the application of TNF-matured DCs 31, 32. The protective response as induced by three injections of TNF-conditioned DCs in the EAE setting was controlled by the simultaneous activation of CD1d-dependent NKT cells, generating a rapid type 2 cytokine environment 33. However, DCs partially matured by TNF were not able to prevent footpad swelling of mice in the leishmaniasis model, further contributing to the hypothesis that a Th2-cell immune deviation mechanism is responsible for the tolerance induction in the EAE model 34. Again, the differences among the similar Th2/Tr1-inducing DC maturation profiles by inflammation or pathogens remained poorly investigated.

Sleeping sickness is caused by Trypanosoma brucei, a single-cell protozoan transmitted to humans by bites of an infected tsetse fly. Studies with resistant mouse models revealed that mice mount an early IFN-γ response during trypanosoma infection followed by a late cytokine switch to the anti-inflammatory IL-10, IL-13, and IL-4 35. This remarkable cytokine shift was also described in helminths infection models such as S. mansoni and Echinococcus multilocularis, which rather protects the host from extensive tissue damage under unrestricted Th1-cell mediated inflammation 36. As helminths are experts in modulating the immune system, their antigens are extensively studied to define how they trigger antigen-presenting cells such as macrophages and DCs to induce Th2-cell responses 19. Trypanosomes are extracellular protozoa, which adapt their protective surface coat consisting of 107 identical densely packed glycoproteins known as variant-specific surface glycoproteins (VSGs) to continuously evade the immune system 37. Hereby, vast amounts of VSG are periodically released into the bloodstream triggering an effective immune response. Earlier reports demonstrated that both soluble VSG (sVSG) and membrane-bound VSG (mfVSG) are the predominant T. brucei components, eliciting differential macrophage activation dependent on MyD88 signaling 38, 39.

In this report, we compared the Th1/Th2-cell inducing pathogenic T. brucei antigens with the Th2-cell inducing inflammatory stimulus TNF for their DC stimulatory capacity. Therefore, sVSG and mfVSG both derived from the T. brucei AnTat1.1 strain and sVSG derived from the T. brucei MiTat1.5 strain were compared. The major difference between the two sVSG proteins used resides in the fact that the MiTat1.5 sVSG lacks GPI-linked galactose moieties and has two additional carbohydrate chains in the protein core as compared with the AnTat1.1 sVSG 38. Our results demonstrate that both T. brucei antigens or TNF induce partial DC maturation signatures defined by upregulation of surface markers but limited or no cytokine production with a strikingly similar gene expression signature. All partial maturation signatures induced the differentiation of Th2-cell responses in vitro and in vivo. These differential Th2-cell profiles showed similar protective effects in the autoimmune disease EAE but no effect in an allergic asthma model. Our data suggest that pathogenic MyD88-dependent VSG antigens and the inflammatory stimulus TNF program for a largely overlapping inflammatory, semi-mature DC signature, inducing default Th2-cell immune responses based on quantitative DC maturation differences.


Partial maturation of DCs by TNF and T. brucei-derived VSG antigens

We compared different T. brucei-derived antigens (AnTat1.1-derived sVSG and mfVSG and MiTat1.5-derived sVSG) with TNF and LPS to induce surface marker expression, cytokine secretion, and differential expression of Notch ligands on DCs. All stimuli upregulated the expression of MHC II, CD40, CD80, and CD86 surface markers compared with untreated DCs (Fig. 1A and B and Supporting Information Fig. 1B). The induction by TNF and T. brucei antigens AnTat1.1-derived mfVSG and MiTat1.5-derived sVSG was, however, below the expression levels achieved by LPS- or sVSG-conditioned DCs (Fig. 1A and B and Supporting Information Fig. 1B). Cytokine analysis revealed that TNF-conditioned DCs do not secrete cytokines or only at very minor levels IL-12p40 or IL-6 (Fig. 1C, Supporting Information. Fig. 1D) as shown previously 23. The T. brucei AnTat1.1-derived mfVSG induced minor amounts of IL-12p40 in DCs as compared with TNF-matured DCs. Interestingly, MiTat1.5-derived sVSG induced substantial IL-6 cytokine release in the presence of IL-1β. None of the stimuli induced IL-12p70 in contrast with LPS-matured and AnTat1.1-derived sVSG-stimulated DCs, which secreted high amounts of all cytokines tested (Fig. 1C, Supporting Information Fig. 1D). Furthermore, LPS or AnTat1.1-derived sVSG stimulation of DCs showed a higher relative mRNA expression of the Th1-cell instructive Notch ligand Delta4 and of Jagged1 but downregulated Jagged2 (Fig. 1D). In contrast, the T. brucei antigens mfVSG and MiTat1.5-derived sVSG induced high expression of the Th2-cell associated Jagged2 but showed only low levels of Delta4 and this to a similar extent as TNF stimulation (Fig. 1D).

Figure 1.

TNF and the T. brucei antigens AnTat1.1-derived sVSG and mfVSG, and Mitat1.5 sVSG induce upregulation of surface marker expression but different levels of cytokine production in DCs. (A) DCs generated from C57BL/6 mice were stimulated for 24 h with the indicated reagents, stained for surface marker expression, and analyzed by flow cytometry. Cells were all gated for high CD11c expression. Numbers indicate the percentage of cells in each quadrant. Data are representative of up to seven independent experiments. (B) As in (A) but figures combine the mean fluorescence intensities (MFIs) of individual DC cultures from different mice of up to seven independent experiments. (C) Supernatants were harvested and tested for their cytokine content by ELISA. Results are combined means+SD of duplicate samples of up to five independent experiments. (D) DCs were matured for 6 h with the indicated stimuli followed by RNA isolation. Real-time quantitative PCR for Jagged1, Jagged2, and Delta4 was performed on cDNA and expression normalized to β-actin levels. Relative expression is represented as levels to untreated DC controls (indicated by dashed line). Results shown are combined mean data of two independent experiments. *p<0.05, **p<0.01, and ***p<0.001, one-way ANOVA followed by Bonferroni post-testing.

Together, TNF and the T. brucei antigens AnTat1.1-derived mfVSG and MiTat1.5-derived sVSG only partially mature DCs as detected by upregulation of surface markers, no or low cytokine production and high relative expression of the Notch ligand Jagged2. In contrast, the AnTat1.1-derived sVSG resembles more LPS-matured DCs. Therefore, and within the major scope of this study, subsequent experiments were conducted with the T. brucei-derived mfVSG and MiTat1.5 sVSG antigens.

In addition, we prepared BM cells from mice deficient in TLR4 and/or MyD88 adaptor protein signaling to define which pattern recognition receptor cascade is required for the observed partial maturation phenotypes. DCs defective in TLR4 signaling still upregulated MHC II and CD86 upon mfVSG exposure, but largely failed to increase surface markers expression in TLR4/MyD88−/− DCs (Supporting Information Fig. 1C). Surprisingly, maturation by MiTat was almost completely blocked in DCs insensitive for TLR4-mediated stimuli and this to a similar extent as LPS-treated DCs. In contrast, MHC II and CD86 upregulation remained unimpaired upon TNF conditioning of TLR4 insensitive or TLR4/MyD88−/− DCs. Together, these data indicate that T. brucei-derived antigens induce distinct partial maturation stages in DCs dependent on MyD88 signaling.

DC conditioning by TNF or T. brucei-derived VSG antigens share similar gene expression profiles

Since the previous experiments did not reveal major differences in the maturation profiles of TNF-, LPS-, or VSG-stimulated DCs, we performed microarray analyses with the differentially stimulated DCs to cover a broader spectrum of gene regulation. After 24 h, treatment cells were prepared for the arrays. The data indicated that LPS stimulation was very different from that by TNF, mfVSG, and sVSG (MiTat1.5) and the latter were highly similar to each other and not so different from untreated DCs (Fig. 2A). More detailed analyses of differentially expressed genes indicated that only 175 genes were induced after TNF, 160 with mfVSG, 466 with MiTat1.5 sVSG but 4969 with LPS were changed more than two-fold over untreated DCs (Fig. 2B). The whole microarray array data are accessible under GEO ( These data also indicate that a common inflammatory signature of 24 genes is shared among all four conditions (Fig. 2C). These genes are largely overlapping with those reported by others before as a typical inflammatory pattern for DCs 40 and thereby indicate the reliability of our microarray approach. The different intensities of induction between TNF/mfVSG/MiTat1.5 sVSG and LPS further strengthen the semi-mature state of the former group (Fig. 2C). Remarkably, this common signature is also completely shared among the stimuli TNF, mfVSG, and MiTat1.5 sVSG, since no different or additional genes were induced (triple field with zero genes, Fig. 2B). Thus, the semi-mature DC signature was represented by upregulation of CD40, CD72, IL-1α, IL-1β, IL-6, CXCL2, SOCS3, Jagged-1, Pleckstrin-2 (Plek2), serum amyloid 3(Saa3), ladinin (Lad), follistatin, (FST), activin (Inhba), and downregulation of PGE-receptor 3 (Ptger3), CD62L (Sell) SIGNR2 (CD209c). In contrast, the fully matured DC signature of genes induced by LPS include the common 24 genes, but regulated additional 4498 genes that were not shared with the other stimuli (Fig. 2B). The exclusive gene signatures induced by TNF alone (Supporting Information Fig. 2) or the comparisons of mfVSG with MiTat1.5 sVSG (Supporting Information Fig. 3) were not marked by a strong immunological signature of gene regulation. Taken together, the common signature of DCs matured by TNF, mfVSG, and MiTat1.5 sVSG induces far fewer genes than LPS, which are mainly characterized by a common signature of 24 mostly inflammatory genes.

Figure 2.

Similar gene expression signatures at partial DC maturation stages. Microarray analysis was performed with BMDCs after treatment for 24 h with the indicated stimuli and compared with untreated DCs. (A) Hierarchical cluster analysis of gene expression profiles of TNF/mfVSG/MiTat1.5 sVSG-stimulated DCs as compared with LPS-stimulated DCs. (B) VENN diagram indicating the number of differentially regulated genes after the differential treatments. Induction was considered when expression was greater than two-fold over untreated DCs. Twenty four genes comprising an inflammatory pattern are commonly regulated. (C) Heat map of signal intensities of DCs after TNF, mfVSG, or MiTat1.5 sVSG stimulation as compared with untreated DCs. Data show single time point values of a single microarray (see Materials and methods).

Partial DC maturation stages induce Th2-cell responses

To dissect the importance of the partial DC maturation phenotypes in directing distinct Th cell differentiation patterns, we cocultured DCs with OVA-specific TCR-transgenic CD4+ OT-II T cells and checked the Th-cell profile by intracellular cytokine staining. Polarizing by LPS showed a clear shift toward IFN-γ, indicating a Th1-cell profile. DC maturation with TNF and mfVSG shifted the T cells toward a Th2/Th9-cell pattern and DC stimulation with MiTat1.5 sVSG heavily reduced Th2-cell and Th9-cell but left the Th1-cell background profile unaltered (Fig. 3A and B and Supporting Information Fig. 4). Furthermore, induction of IL-17 production in T cells was negligible under all conditions (Supporting Information Fig. 4) and T cells did not produce anti-inflammatory IL-10 after one round of DC stimulation (data not shown) and as we reported previously 23.

Figure 3.

Partially matured DCs direct Th2-cell differentiation. (A) DCs were matured with the indicated stimuli and cocultured in the presence of OVA peptide with naïve CD4+ OT-II T cells in vitro. At day 5 of coculture, T cells were restimulated with PMA and ionomycin and intracellular cytokine levels were determined by flow cytometry. Numbers in gates indicate the percentage of cytokine-positive cells gated on CD4+ cells. One representative experiment of four experiments is shown. (B) Levels of cytokine-positive cells as in shown in (A) but normalized to untreated samples. Data presented are pooled means of single sample analyses up to four independent experiments+SDs. *p<0.05, **p<0.01, one-way ANOVA followed by Bonferroni post-testing.

Earlier reports demonstrated that BM-derived DCs efficiently induced CD4+CD25+FoxP3+ Treg cells in vitro predominately in the presence of exogenously supplied TGF-β 41. Indeed, hardly any Treg-cell generation could be detected in the absence of exogenous TGF-β irrespective of the maturation phenotype of the DCs (Supporting Information Fig. 5A). Nevertheless, DCs matured with TNF, mfVSG, or MiTat1.5 sVSG induced high levels of FoxP3+ Treg cells in vitro to a similar extent as untreated DCs but superior to LPS-conditioned DCs when exogenous TGF-β was supplied to the culture, further supporting the observation that DCs require TGF-β for the efficient induction of FoxP3+ Treg cells in vitro as described previously 41.

In summary, our data identify Th2-cell differentiation patterns linked to partial DC maturation stages with quantitative differences between pathogen-derived, TLR-dependent VSG antigens, and non-TLR-dependent TNF stimulation in vitro. No induction of FoxP3+ Treg cells could be observed by any of our DCs in the absence of exogenous TGF-β in vitro.

Partially mature DCs show similar priming and Th-cell polarizing capacity in vivo

To assess how these DC maturation signatures prime T-cell responses in vivo, we injected differentially matured and OVA-loaded DCs together with OVA-specific TCR-transgenic OT-II T cells i.v. and determined proliferation and cytokine production of injected T cells. DCs matured with TNF, mfVSG, or MiTat1.5 sVSG all induced proliferation of CFSE-labeled T cells (Fig. 4A). The most profound priming in T cells was detected upon injection of LPS-matured DCs as determined by flow cytometry (Fig. 4A) or calculated as the division index (Fig. 4B). Furthermore, one single injection of DCs conditioned with TNF, mfVSG, or MiTat1.5 sVSG increased intracellular IL-13 and IL-5 release by ex vivo restimulated OVA-TCR-specific T cells (Fig. 4C and D), in contrast to mice which received LPS-matured DCs which showed only background levels of IL-13- or IL-5-producing OVA-TCR-specific T cells (Fig. 4C and D). Similar to our in vitro findings (Supporting Information Fig. 4B), a low frequency of IFN-γ-releasing T cells was detectable after a single injection, irrespective of the DC maturation regimen. Clearly polarized Th1-cell responses resulted only after injection of LPS-matured DCs (data not shown and Fig. 4C and D). Furthermore, injection of DC conditioned with TNF, mfVSG, or MiTat1.5 sVSG did not raise the frequency or total cellular amounts of FoxP3+ Treg cells among OVA-TCR-specific T cells in vivo similar to LPS-matured DCs (Supporting Information Fig. 5B and C) further strengthening the observation that partially mature DCs efficiently induce proliferation and priming of (CFSE labeled) OVA-TCR-specific T cells in vivo (Fig. 4A).

Figure 4.

DCs conditioned by TNF or T. brucei antigens show similar T-cell priming capacities in vivo. (A) Mice received an intravenous injection of CFSE-labeled DO11.10 T-cell suspension followed by DCs matured with the indicated stimuli and loaded with OVA peptide. Untreated animals did not receive any DC injections. Histogram plots show CFSE dilution of CFSE+ KJ1-26+ T cells recovered from spleens of mice 4 days after final injection. One representative experiment of two is shown. (B) Level of CFSE dilution was calculated as division indices. Figure depicts pooled mean data+SD of two individual cultures/mice/independent experiments. (C) Mice received CD4+ CD25 OT-II T cells intravenously followed by a single injection of DCs matured with indicated stimuli and loaded with OVA peptide. Intracellular cytokine content of CD4+ Vβ5+ T cells was determined 6 days post-injection after peptide restimulation of splenocytes. Numbers in plots represent percentage of cells gated on CD4+ Vβ5+ cells. One representative experiment is shown. (D) As in (C) but graph represents pooled mean data of up to three individual mice/experiments per condition. Cells were gated on CD4+ Vβ5+ cells. *p<0.05, **p<0.01, and ***p<0.001, one-way ANOVA followed by Bonferroni post-testing.

Together, DCs conditioned by TNF- or T. brucei-derived VSG antigens induce profound and comparable Th2-cell priming in vivo.

Partially matured DCs do not affect asthma in mice

Asthma induced by alum-guided immunization of mice with OVA is a widely used model for a Th2-cell mediated disease characterized by proinflammatory lung infiltrates of eosinophilic granulocytes and a subsequent Th2-cell dependent production of OVA-specific IgG1 and IgE 42. Mice subjected to repeated sensitization and antigen challenges showed a profound influx of total cells, in particular eosinophils in the bronchoalveolar lavage (BAL) as a major parameter for asthma (Fig. 5A). Three repetitive injections of OVA-loaded TNF, mfVSG, or MiTat1.5 sVSG-matured DCs did not change the total cellular influx in the lungs compared with noninjected animals. In addition, the influx of eosinophils, macrophages, neutrophils, and lymphocytes was not differentially regulated upon injection of partially matured DCs and this independently of the inflammatory vs TLR-dependent quality of the maturation stimuli (Fig. 5A and data not shown).

Figure 5.

No influence on the allergic asthma model by all types of partially mature DCs. (A) Mice received three repetitive injections of DCs loaded with OVA protein and matured with the indicated stimuli at days −7, −5, and −3 before asthma induction. Control mice received no DC treatment. Cell numbers recovered from BAL 6 days after final antigen challenge are shown. Each dot represents data obtained from one mouse. Data shown are pooled from all mice used in up to four independent experiments. (B) Graphs show OVA-specific IgG1 or IgE levels in sera of mice as detected by ELISA. Mice received DC treatments as described in (A). Each dot depicts OVA-specific IgG or IgE levels of serum of one mouse used in up to four different experiments. *p<0.05, **p<0.01, ***p<0.001, one-way ANOVA followed by Bonferroni post-testing.

OVA-specific Th2-cell dependent IgG1 and IgE were detected in the serum of mice upon alum/OVA sensitization and antigen challenges. Surprisingly, no change was detectable at the OVA-specific Ig levels when mice were pretreated with the differentially matured and OVA-loaded DCs (Fig. 5B). Together, MyD88-dependent T. brucei-derived VSG antigens or nonTLR-dependent TNF conditioning of DCs did not alter subsequent Th2-cell driven allergic asthma.

Similar protective potential by partially mature DCs in the autoimmune model EAE

EAE serves as a common murine model for the early phases of multiple sclerosis, which can be achieved by immunizing mice with the auto-antigen MOG in CFA. Mice develop MOG-reactive pathogenic Th1 and Th17 cells, which then infiltrate into the CNS and cause inflammatory edema leading to the reversible paralysis symptoms 43. Previously, we have shown that repeated injections of DCs stimulated with TNF and loaded with MOG-peptide suppressed EAE, partially by creating a Th2/Tr1 cytokine environment including immune deviation and IL-10-mediated suppression 23, 33. We therefore wanted to analyze how the partial DC maturation stages induced by TLR-dependent or independent stimuli would modulate the autoimmune disease EAE.

To detect whether the DC injections ameliorate or worsen the disease, we switched the amounts per DC injection from 3 to 3.5×106 cells, which is the fully protective protocol 23, 33, 44 to 2–2.5×106 cells, which leads to about 50% reduced clinical score 44. Three i.v. injections of suboptimal amounts of MOG-loaded TNF-matured DCs protected mice partially from EAE as 10 out of 15 mice developed clinical symptoms and mice only reached a mean maximum score of 1.850±0.944 (Fig. 6A and B). Surprisingly, mice which received three injections of DCs matured with the T. brucei antigens mfVSG or MiTat1.5 sVSG were also partially protected from EAE as 8 out of 12 and 13 out of 19 mice developed signs of EAE, respectively (Fig. 6A and B). Together, our data indicate that all partially mature DCs protected mice to a similar extent from EAE.

Figure 6.

Similar protection efficiency by all types of partially mature DCs in the autoimmune model EAE. (A) Mice received three repetitive injections of DCs loaded with MOG peptide and matured with the indicated stimuli at days −7, −5, and −3 before EAE induction. Control mice received no DC treatment. Mice were monitored daily for disease score. Results represent the average disease score of all mice used in up to six independent experiments (numbers of mice indicated in (B)). (B) Statistical overview showing incidence, mean maximum score, and mean day of onset of all EAE experiments performed. Mice were treated with three i.v. injections of MOG-loaded DCs stimulated with indicated stimuli before EAE induction. (C) Mice received three consecutive injections of MOG peptide-loaded DCs generated from C57BL/6 mice and matured with indicated stimuli before EAE induction. At day 30 after EAE induction, spleens were removed and restimulated with graded concentrations of MOG peptide. Supernatants were analyzed for their cytokine content by ELISA. Graphs show relative cytokine production normalized to the cytokine content of untreated controls. Each dot represents the relative cytokine production of splenocytes of pooled mice for one condition within one experiment restimulated at 20 or 2 μg/mL MOG peptide. *p<0.05 and **p<0.01 one-way ANOVA followed by Bonferroni post-testing.

DCs programmed by TNF or T. brucei VSG antigens mount a protective Th2/Tr1-cell response in EAE

As published previously 33, protection from EAE by TNF-matured DCs required activation of IL-10+ IL-13+ cytokine-producing CD4+ Th2/Tr1 cells. IL-4 is also produced but immediately consumed in normal mice and only detectable in IL-4R-deficient mice 33. We therefore assessed how the differentially matured DCs influenced the T-cell cytokine profile of the spleens as detected after MOG peptide restimulation and cytokine analysis.

The cytokine profile of T cells from untreated mice typically consists of high amounts of proinflammatory IFN-γ and IL-17 but low amounts of IL-10 and IL-13. In contrast, this pattern becomes inverted in mice, which received repetitive injections of TNF-matured DCs 23, 33. Under the partially protective conditions, we observed the same shifts, although, due to the suboptimal settings, not always reaching statistical significance (Fig. 6C). Nevertheless, splenocytes from mice injected with DCs matured with the VSGs significantly downregulated IL-17 production comparable to the T-cell cytokine profile of TNF-DC-treated animals. Mice treated with MiTat-matured DCs, however, were not able to block the nonprotective IFN-γ production as TNF-DC-treated animals, but in addition, retained high production of the disease-preventing cytokines IL-13 and IL-10 (Fig. 6C). Moreover, repetitive injections of differentially matured DCs did not alter the frequencies of FoxP3-expressing Treg cells in spleens of EAE-diseased mice (Supporting Information Fig. 5D). This suggests that semi-mature DCs regulate EAE by protective mechanisms other than CD25+ FoxP3+ Treg-cell induction. In sum, the partial DC maturation stages were all equally effective in creating a protective Th2/Tr1-cell environment, which was able to block the Th1/Th17-cell mediated EAE.


In this study, we showed that similar partial maturation stages of DCs can be achieved with the proinflammatory cytokine TNF and the T. brucei antigens mfVSG and MiTat1.5 sVSG. Our data further indicate that low concentrations of pathogen-derived TLR-mediated stimuli program DCs similarly to the inflammatory cytokine TNF for the differentiation toward an inflammatory, semi-mature DC phenotype. These partial DC maturation stages were able to induce Th2-cell priming in vitro and in vivo and induced only quantitative differences in the extent of Th2-cell differentiation. Moreover, these Th2-cell signatures did not differ in their intrinsic quality to heal autoimmune diseases such as EAE and had no influence on allergic asthma. These data have important implications for the understanding of parasitic immune evasion, the design of vaccines and provide further insights how DC maturation signatures critically contribute to the differentiation of defined Th-cell subsets.

The stimulus LPS triggers DC maturation through TLR4 ligation and directs Th-cell differentiation toward Th1-cells. Less is known which PRRs drive Th2-cell associated immune responses. Recent reports suggest that house dust mite allergens initiate asthmatic inflammation by signaling through the TLR4 receptor complex in part by LPS contamination 45, 46. Our data show that the T. brucei antigen MiTat1.5 sVSG-conditioned DCs to produce IL-6 and IL-1β, which is dependent on TLR4 and the adaptor molecule MyD88. A novel TLR4-mediated signaling pathway was identified in which TLR4 stimuli trigger a rapid increase in intracellular cAMP followed by translocation of the transcription factor CREB and IL-6 production 47. Further investigation is needed to address whether MiTat1.5 sVSG activation of DCs is accompanied with an intracellular cAMP rise and CREB transcription factor translocation.

The T. brucei AnTat1.1 antigen mfVSG triggers the activation of DCs mainly independent of the TLR4 receptor complex but dependent on the signaling adaptor MyD88. Earlier reports showed that mfVSG triggers macrophage activation through a MyD88-dependent signaling cascade 38, 39. Heating VSG antigens for 15 min at 95°C did not abrogate the DC maturation activity (data not shown), indicating that the glycosyl-inositol-phosphate (GIP) moieties of the GPI anchor are the DC-activating factors as suggested previously for macrophages 38. In analogy with other parasitic protozoa such as Leishmania major, Plasmodium falciparum, and T. cruzi, GPI anchors of T. brucei are believed to form the most prominent inflammation and disease-inducing component 48. Indeed, recent reports showed that the GPI anchors of P. falciparum mainly trigger MyD88-dependent TLR2 and to a lesser extent TLR4 signaling in macrophages 49.

DCs sense different pathogens and respond by upregulating MHC and costimulatory molecules as well as cytokine production to mount an appropriate T-cell response. However, it appears that DCs release substantial amounts of cytokines only upon strong TLR activation 23, 27, 29, 50. We found that mfVSG and Mitat1.5 sVSG act on DCs through MyD88 to mediate maturation but result in a TNF-like inflammation-induced partial maturation profile, leading to Th2-cell polarization. Although we also detected some IL-9-producing T cells in vitro, this was not observed after injection. Since there is ongoing debate whether IL-9 is part of Th2 cells or belonging to an own Th9 subset 51, we did not further address IL-9 in our Th2-cell studies. Inflammatory mediators can activate DCs also in vivo, which are similarly unable to produce IL-6 or IL-12p40 27, 50, 52. Sporri and Reis e Sousa have shown that DCs activated by inflammatory mediators in vivo induced Th cells but these were unable to support immunoglobulin isotype switching 27. Similarly, in this study all partially matured DCs types were unable to alter IgG1 and IgE levels in the asthma model. Recent reports also suggest that IL-6 by triggering IL-21 secretion in T cells drives the differentiation of Th cells that acquire the ability to provide B-cell help for isotype switching 53. Here, DCs matured with MiTat1.5 sVSG showed substantial production of IL-6, but DC treatment did not modify the isotype switch compared with other maturation stimuli in the allergic asthma model. However, it remains to be determined whether DCs conditioned by MiTat1.5 sVSG can induce B-lymphocyte helper T cells in the absence of any additional adjuvant activity. The capacity to provide efficient B-cell help might further delineate distinct functions of the Th2-cell subsets induced by inflammatory mediators or TLR agonists as identified in this study.

In our study, the nonpathogen-derived inflammatory stimulus TNF and type 2 pathogen-derived antigens show remarkable similarities for the maturation of BM-derived DCs, i.e. the in vitro counterpart of the so-called TNF/iNOS-producing DCs (Tip-DCs) 54. It has been suggested that DCs develop by means of the growth factor fms-like tyrosine kinase 3 (FLT-3L) in the steady state, whereas under inflammation DCs are additionally generated from monocytes by GM-CSF 54. Initial encounter with a pathogen and, hence, initial Th-cell polarization will most likely occur solely by the tissue-resident DCs or, in case of tse-tse fly-mediated blood infection with trypanosomes, steady-state DCs. Tip-DCs develop later during infection from recruited monocytes and by GM-CSF secreted from T cells at the site of inflammation. Others reported that the steady-state occurring splenic DC subsets (CD8α, CD8α+ or plasmacytoid DCs) show intrinsic differences to mount preferentially a Th1- or Th2-cell biased response 8, 55, 56. Thus, our BM-DC equivalents to Tip-DCs might play a decisive role in dampening or modulating the initially mounted Th-cell response to effectively eliminate the invading pathogen, a process also referred to as “success-driven” Th-cell modulation 57. The functional difference of inflammatory vs steady-state occurring DCs might explain the reason why DCs indirectly activated by inflammatory mediators in vivo failed to mount Th2-cell responses, but inflammation drives Th2-cell differentiation at the Tip-DC level 27, 52.

The analyses of our microarray data indicated that (i) TNF, the AnTat1.1 mfVSG and the MiTat1.5 sVSG regulated only a limited set of genes in DCs as compared with LPS, (ii) the regulation patterns of TNF, AnTat1.1 mfVSG, and the MiTat1.5 sVSG are widely overlapping, and (iii) the differences between TNF (only proinflammatory) and AnTat1.1 mfVSG or the MiTat1.5 sVSG (presumed antiparasitic Th2-cell immunity) are remarkably few.

Our findings that TNF induces less gene regulation as compared with LPS is in agreement with the findings using a DC line 58 and also the general inflammatory pattern of 24 genes we found, shared remarkable overlap with the 44 genes that have been found by others 40, sharing key factors such as CD40, IL-1β, and IL-6. While LPS induced the same 24 genes, it regulated many more others, suggesting that inflammatory semi-maturation may represent more a quantitatively different state of maturation, rather than a completely different quality. One marked difference is the absence of IL-12p40 in our general inflammatory profile of 24 genes, which appeared only after LPS stimulation. This may be due to the fact that in the studies with the D1 line only pathogens but not inflammatory mediators were included and IL-12p40 thereby reflects pathogen stimulation. In addition, the lack of genes specifically regulated by mfVSG and MiTat1.5 sVSG would indicate an immune response against T. brucei is missing. The Th2-cell response generated by mfVSG and MiTat1.5 sVSG-matured DCs was expected to result in an enhanced isotype switches and IgG1 and IgE production in the asthma model. However, here the two VSG antigens behaved like TNF, i.e. “only inflammatory.” This may point to some VSG antigens as a means of immune evasion, mimicking an inflammatory reaction in the absence of pathogens.

The fact that TNF and the mfVSG and Mitat1.5 sVSG regulate only few genes, whereas LPS regulates the same but almost 5000 genes in addition, argues for predominantly quantitative differences between the two types of DC maturation. However, since these quantitative changes led to qualitatively different Th1 or Th2-cell polarization, this may reflect another DC-based aspect of the “strength of signal” theory where peptide titrations and affinities heavily influenced the Th-cell skewing potential 59, 60. The peptide dose dependency has been shown to be independent of the DC subtype but strong LPS or CpG stimulation clearly shifted toward Th1-cell 61. As a mechanism how this could be regulated, others proposed that weak T-cell stimulation prevents CD40L upregulation, which in turn was required to trigger CD40 on DCs for their IL-12 production and Th1-cell immunity 62. Thus, weak DC stimulation would then result in a Th2-cell response, whereas strong DC stimulation, i.e. by DC maturation with LPS or weak maturation but presenting high doses of peptide, would result in a Th1-cell polarization.

The three signal models as initially proposed by Kapsenberg 7 explain how DCs mediate Th-cell differentiation: peptide-MHC ligation (signal 1), costimulatory signaling (signal 2), and a selective cytokine set initiate the differential Th-cell commitment (signal 3). For Th1-cell polarization, IL-12p70 production by DCs is, besides the recently described CD70-dependent pathway 63, a clear signal toward Th1-cell polarization but signal 3 for Th2 cells remains less clear.

Previous reports have shown that the Th2-cell promoting mediator PGE2 induces the secretion of IL-12p40 in DCs thereby inhibiting the production of the Th1-cell driving cytokine IL-12p70 16–18. It has been proposed that blocking or washing out IL-12p70 production is sufficient to drive the differentiation of Th2-cell responses by the so-called default or exhaustion pathway 64, 65. The elimination of IL-12p70 from the context of antigen presentation by mature DCs would result in a similar phenotype of inflammatory semi-mature DCs as we have generated them here. The differences in the production of low levels of IL-6 or IL-12p40 by DCs matured with TNF, mfVSG, or MiTat1.5 sVSG do not seem to shift the qualitative Th2-cell profile but only result in minor quantitatively different amounts of Th2 cells. In addition, these differences did not have functional consequences after injection on asthma or EAE.

Due to the fact that VSG-mediated semi-maturation of DCs is dependent on MyD88 signaling, we may have to consider these Th2-cell inducing antigens as weak TLR agonists. Others have shown that especially TLR2 triggering of DCs can lead to a Th2-cell priming with or without coinduction of Th17 cells 66, 67 although there are also other results for Schistosoma antigens that induce Th2-cell responses without the involvement of TLR2, TLR4, or MyD88 68. This indicates that quantitatively low TLR signaling but also TLR- and MyD88-independent DC maturation can lead to Th2-cell polarization.

One would expect that if DCs conditioned by TNF or VSG antigens induce preferentially immunogenic Th2-cell responses, they should increase the severity of asthma symptoms when pulsed with the allergens and injected before disease induction. Alternatively, if these DCs prime Th1-cell responses, the disease should ameliorate. We did not test LPS-matured DCs in this context. Others have addressed this question before by using CpG-matured BM-DCs, which are similar to LPS for the instruction of Th1-cell responses, but without effects on asthma 69. Lambrecht's group has shown that rather plasmacytoid DCs may be able to control asthma 70, 71.

Semi-mature DCs prevented the paralyzing symptoms in the EAE model by immune deviating toward a Th2/Tr1 protective response, whereas LPS-matured DCs were not protective 33, 72, 73. However, the application of semi-mature DCs in Th2-cell associated asthma model neither ameliorated nor worsened the disease symptoms, similarly to the previous data obtained for the murine L. major Th2-cell infection model 34. These data suggest that the Th2/Tr1 differentiation as induced by semi-mature DCs in Th2-cell models results in a balance between an intrinsic inflammation-limiting Tr1 response and the active asthma-promoting Th2-cell response. Interestingly, the upcoming role of such balanced Th2-cell responses in limiting tissue pathology and inflammation has been discussed previously in several infection models and especially for macrophages 74–76.

Collectively, the observations described in this study indicate that DCs induced Th2-cell differentiation at a partial maturation stage. TNF and T. brucei-derived mfVSG and Mitat1.5 sVSG antigens induce similar maturation signatures of inflammatory semi-mature DCs leading to Th2-cell induction. This inflammatory Th2-cell inducing signature is, however, shared with the Th1-cell inducing stimulus LPS, which regulates additional genes for Th1-cell induction. Our data support an inflammatory DC-induced Th2-cell default pathway that is predominantly marked by quantitative maturation differences as compared with Th1-cell inducing DCs.

Materials and methods


C57BL/6 and BALB/C mice were bred in our own animal breeding facilities or purchased from Harlan. OT-2 mice (C57BL/6 background, F. Carbone, Melbourne), DO11.10 TCR-transgenic mice (BALB/C background, generated by K. Murphy, New York), TLR4-mutated C3H/HeJ (JAX mice), and TLR4/MyD88−/− mice (on a 129Sv x C3H/HeN genetic background, originally generated by S. Akira, Osaka and provided by A. Gessner, Erlangen) were all bred under specific pathogen-free conditions. All animal experiments were performed in accordance with the guidelines of the local authorities.

VSG preparations and LPS quantification

Trypanosomes (T. brucei Antat1.1 and MiTat1.5) were harvested from infected blood by DE52 chromatography, using sterile PBS (pH 8.0) supplemented with 1.6% glucose for equilibration and elution 77. sVSG was prepared by osmotic shock as described previously 78. Briefly, the DE52-purified parasites were resuspended in Balts-buffer (50 mM sodium phosphate buffer, pH 5.5) and incubation on ice for 30 min followed by a 5-min incubation at 37°C. The solution was subsequently centrifuged (1400 rpm, 7 min, 4°C) and the supernatant treated with benzonase (VWR) to remove potential DNA/RNA contamination (as described by the supplier). The supernatant was dialyzed against 10 mM Tris, pH 7.4, and the sVSG was purified using ion-exchange chromatography and gel filtration as described previously 79, 80. mfVSG was prepared as described previously 81. Prior to performing a size exclusion chromatography (equilibrated against 10 mM Tris, pH 7.4, containing 0.02% N-octylglucoside, Sigma-Aldrich), the mfVSG was treated with benzonase (similar as for sVSG) to remove potential nucleic acid contamination. The protein concentration of both VSGs was estimated spectrofotometrically by a detergent-compatible protein assay kit (Bio-Rad) using BSA as a standard. The purity of both sVSG and mfVSG was checked in SDS-PAGE and found to be >95%. In addition, Western blot analysis, using rabbit polyclonal anti-VSG and anti-cross-reacting determinant Abs confirmed the presence of the GPI anchor on mfVSG 82. Finally, the endotoxin levels were determined using the Limulus amebocyte lysate (LAL) test (Cambrex) according to the manufacturers' instructions and found to be <0.5 pg/μg VSG.

Generation and maturation of BM-DCs

BM-DCs were generated as described previously 83. Briefly, BM-precursor cells were isolated from the hind limbs and seeded out in petri dishes (10 cm, Greiner) at 3×106 cells per dish. For microarray analysis, BM-precursor cells were depleted of B and T cells by using anti-CD19 and anti-CD90 magnetic beads (Miltenyi Biotec), respectively. Cells were cultured in RPMI 1640 (PAA) supplemented with 10% heat-inactivated fetal calf serum (FCS, PAA), penicillin (100 U/mL; PAA), streptomycin (100 mg/mL; PAA), L-glutamine (2 mM; PAA) and β-mercaptoethanol (50 mM; Sigma-Aldrich). Culture medium was additionally supplemented with 10% supernatant from a GM-CSF-transfected cell line 84. At d7 or d8, BM-derived DCs were harvested and replated at a density of 106 cells/mL in a 24-well plate (nontissue culture treated; Greiner). For maturation analysis of cytokine production and surface marker expression, BM-DCs were cultured for 20–24 h in the presence of TNF (500 U/mL; PeproTech), LPS (Escherichia coli 0127:B8 0.1 μg/mL; Sigma-Aldrich), sVSG or mfVSG from clone AnTat1.1 (2 μg/mL), or sVSG MiTat1.5 (2 μg/mL). For in vivo polarization assays, BM-DCs were seeded at a density of up to 5×106 cells/mL, matured for 4 h only with different maturation stimuli and additionally loaded with 40 μg/mL MOG35–55-peptide (synthesized and HPLC purified by R. Volkmer, Charité, Berlin, Germany), 10 μM OVA-peptide327–339 (Activotec) or 50–100 μg/mL OVA protein (endotoxin-free; Hyglos) as indicated.

Cytokine detection by ELISA

DC culture supernatants were analyzed for their cytokine content by commercially available ELISA kits for IL-1β, IL-6, IL-10, IL-12p40, IL-12p70, TNF (all BD), and IL-23 (eBiosciences). Supernatants of T cells were analyzed for IL-4, IL-10, IFN-γ (BD), IL-13, and IL-17 (eBiosciences).

Flow cytometry

Cells were stained in ice-cold PBS supplemented with 0.1% BSA and 0.1% sodium azide. To avoid unspecific Ab binding, cells were incubated with 2.4G2 (hybridoma supernatant) or medium supplemented with 10% FCS and were stained with the following Abs: anti-CD11c-PerCP-Cy5.5 (N418; Caltag), anti-CD11c-APC (HL3; BD), anti-CD25-FITC (7D4; BD), anti-CD25-PE or allophycocyamin (APC) (PC61; BD), anti-CD40-PE (3/23; BD), anti-CD80-FITC (16-10A1; BD), anti-CD86-FITC (GL1; BD), anti-MHCII-PE (M5/114.15.2; BD), anti-Vβ5.1 and 5.2 TCR-biotin or FITC (MR9-4; BD), anti-DO11.10-TCR-TriColor (KJ1-26; Caltag), anti-CD4-APC or PerCP (RM4-5; BD). Isotype control Abs were used at the same concentration. Intracellular FoxP3 was stained using the eBioscience® anti-mouse FoxP3 staining set and anti-FoxP3-PE or APC Abs (FJK-16s; eBioscience) according to the manufacturer's instructions (eBioscience). For intracellular cytokine detection, cells were stained for surface markers followed by fixation in 2% formaldehyde and permeabilization in perm buffer (0.5% saponin in PBS) and then stained in perm buffer for the following Abs: anti-IL-4-PE or APC (11B11; BD), anti-IL-5-PE (TRFK5; BD), anti-IL-9-PE (RM9A4, Biolegend), anti-IL-10-FITC or APC (JES5-16E3; BD), anti-IL-13-PE (eBio13A, eBioscience), anti-IL-17-PE or PerCP-Cy5.5 (TC11-18H10.1; BD) and anti-IFN-γ-FITC or PE (XMG1.2; BD). Samples were measured at a FACScan or FACScalibur flow cytometer (BD) and data were analyzed with FlowJo software (TreeStar).

Real-time PCR of Notch ligands

Total RNA was extracted from DC lysates using Trizol® reagent (Invitrogen) and performed according to the manufacturer's instructions. cDNA was synthesized using Superscript III Reverse Transcriptase (Invitrogen). Quantitative expression of the Notch ligands Jagged1, Jagged2, and Delta4 was determined with a Biorad iCycler iQ (Biorad) using primers described previously 14. Real-Time PCR was run for 40 cycles and performed in 25 μL volume containing 0.5× Absolute QPCR SYBR Green mix (Thermo Fisher Scientific), 1 μL of 1:10 diluted cDNA sample and 0.2 μM of each primer. Quantifications of the samples were determined by the ΔΔ cycle threshold (Ct) method. The housekeeping gene β-actin was used for normalization of the samples.

Microarray hybridization and data analysis

Total RNA from DCs treated for 24 h with LPS (E. coli 0127:B8 0.1 μg/mL), Antat1.1 sVSG, mfVSG, MiTat1.5 (2 μg/mL), TNF (500 U/mL; PeproTech) or without a stimulus, was extracted using the Trizol® reagent according to the manufacturer's instructions (Invitrogen). RNA integrity and comparability between samples was tested using a BioAnalyzer (Agilent, Santa Clara, CA). RNA integrity numbers were between 9, 8, and 10. Samples were prepared and microarray analysis was performed as we described previously 85.

In vitro T-cell priming and polarization

Purified naïve CD4+ T cells were isolated from the spleens and lymph nodes of OT-2 mice by depleting non-CD4+ cells using the CD4+ T-cell enrichment kit (StemCell Technologies). Then, CD4+ T cells were further enriched by negative selection using MACS technology with anti-CD25 PE and anti-PE magnetic beads (Miltenyi Biotech). For T-cell differentiation assays, purified CD4+CD25 OT-II T cells (5×104) were cultured with day 8 BM-derived DCs (104–105) and 50 nM OVA-peptide327–339 (Activotec) in the presence or absence of maturation stimuli. Cultures were restimulated at day 5 by PMA (10 ng/mL) and ionomycin (1 μg/mL) (both Sigma-Aldrich) in the presence of Golgistop as indicated by the manufacturer (BD).

In vitro Treg-cell conversion assay

Treg-cell assays were set up as described previously 41 with minor modifications. Briefly, purified CD4+ CD25 OT-II T cells (2×104) were cultured with day 8 BM-DCs (6×103) matured with various maturation stimuli for 4–6 h prior to coculture and 100 ng/mL OVA-peptide327–339 (Activotec) in 96-well round-bottom plates (Greiner Bio-One). Additional recombinant porcine TGF-β1 (R&D systems) was added to the culture at a concentration of 2 ng/mL when indicated. Cultures were analyzed on day 5 by flow cytometry staining of surface markers CD4 and CD25 and the transcription factor FoxP3 as described in the previous section.

In vivo T-cell priming

For in vivo proliferation assays, spleens and lymph nodes were isolated from DO11.10 mice and labeled with CFSE (Invitrogen) according to the manufacturer's instructions. Mice received 107 CFSE-labeled cells injected in the tail vein in addition to 2–2.5×106 DC matured and loaded with OVA-peptide327–339 (Activotec) as described in the previous section. In total, 96 h after the final injection, CFSE dilution of splenocytes was analyzed. Division index was calculated as the mean number of divisions among cells, which divided at least once. For in vivo polarization assays, 106-purified CD4+ CD25 OT-II or DO11.10T cells were injected i.v. followed 24 h later by injection of 2–2.5×106 DCs matured and loaded with OVA-peptide327–339 (Activotec). Transferred T cells were analyzed for their cytokine content by restimulation of splenocytes 6 days after final injection with 10 μM OVA-peptide327–339 (Activotec) during 72 h. Brefeldin A (5 μg/mL; Sigma) was added during final 6 h of restimulation followed by intracellular cytokine staining as described.


EAE induction was performed as described previously 23. Briefly, C57BL/6 mice were injected s.c. with 200 μg MOG35–55 peptide emulsified in CFA (Sigma-Aldrich) further enriched with Mycobacterium tuberculosis H37RA (5 mg/mL) (Difco). Additionally, mice were injected with 400 ng Pertussis toxin i.p. (List Biological Laboratories) at days 0 and 2 of EAE induction. Mice were scored daily for clinical disease symptoms according to the following scale: 0, no disease; 1, limp tail weakness; 2, hind limp weakness; 3, hind limp paralysis; 4, hind and fore limp paralysis; and 5, moribund or death. DCs were injected at days −7, −5, and −3 before EAE induction in the tail-vein of mice and for a total of 2–2.5×106 DCs. Thirty days after EAE-induction, spleens were removed for restimulation and seeded out as triplicates of 4×105 cells per well in a flat-bottomed 96-well plate (Greiner) in the presence of graded concentrations of MOG35–55 peptide. After 72 h of restimulation, supernatant was harvested and analyzed for its cytokine content by ELISA.

Asthma induction

BALB/C mice were sensitized by i.p. injections of 10 μg OVA protein (Hyglos) mixed in aluminum hydroxide at days 0 and 14 of asthma induction. Mice treated as negative controls received injections of aluminum hydroxide only. DCs were injected at day −7, −5, and −3 before asthma induction in the tail-vein of mice and for a total of 2–2.5×106 cells. Then, mice were challenged by intranasal administrations of 100 μg OVA protein in 50 μL PBS at days 22, 23, and 24 of asthma induction. Six days after the last OVA challenge, mice were lethally anesthetized followed by bleeding of the axillary veins for serum immunoglobulin analysis. Blood was coagulated for 2 h at room temperature and centrifuged on 3000×g for 5 min to recover the serum. Circulating OVA-specific IgG subclasses were determined by ELISA. For this, 96-well plates (♯353279; BD) were coated overnight at 4°C with OVA protein (Sigma; 100 μg/mL) in 0.1 M NaHCO3 coating buffer. Sera were loaded as serial dilutions in 1% FCS in PBS. OVA-bound Abs in the sera were detected by horseradish peroxidase-conjugated mouse heavy chain-specific Abs: anti-mouse IgG1-HRP (Serotec), or IgE-biotin and streptavidin-HRP (BD) followed by the substrate tetramethylbenzidine (BD). Absorbance was detected using an ELISA microplate reader (Vmax; Molecular Devices). Serum titers were calculated from the serial dilution, which was 1.5-fold increased compared with baseline (optical density of negative control mice). BAL was performed by flushing the lungs through an opening in the trachea with PBS from a syringe. Differential cell count of the BAL was determined by recording total cell amount and spinning cells on microscope glass slides using a Cytospin Universal centrifuge (Hettich, Germany). Cytospins were stained with hematoxylin-eosin solution (Diff-Quick staining set; Medion Diagnostic) and cells were classified using standard morphologic criteria.

Statistical analysis

Data are represented as mean data±SD. Statistical significance was analyzed with GraphPad Prism software using one-way ANOVA followed by Bonferroni post-testing and significance accepted if p<0.05. Data of EAE and asthma experiments were validated using Kruskal–Wallis test followed by Dunn's post-test and considered as significant if p<0.05.


This work was supported by the German Research Council (DFG) through the Sonderforschungsbereich SFB581, International Research Training Grant IRTG1522 and the Transregio Collaborative Research Centre TR52. The authors thank A. Gessner for providing the C3H/HeJ and TLR4/MyD88−/− mice.

Conflict of interest: The authors declare no financial or commercial conflict of interest.