RosaiDTR mice (19) (kindly provided by Ari Waisman, Mainz, Germany) and CD11c-cre mice (20) were crossed and maintained under specific pathogen-free conditions at the Animal Facility of the University Hospital Essen, Germany. Cryopreserved Plasmodium yoelii 17XNL (non-lethal) infected red blood cells (iRBCs) were passaged once through WT mice before being used in experimental animals. For infection 1 × 10⁵ iRBCs were injected i.v. The frequency of iRBCs (parasitemia) was determined by microscopic examination of Giemsa-stained blood films. For CD11c^high cell depletion, RosaiDTR/CD11c-cre mice were injected i.p. with 12 ng/g body weight of diphtheria toxin (DT; Merck, Darmstadt, Germany) starting 1 day before or at day 4 of P. yoelii infection and subsequently every day. Alternatively, mice were treated with DT only once 1 day prior to infection. The study was carried out in accordance with the guidelines of the German Animal Protection Law and the state authority for nature, environment and customer protection, North Rhine-Westphalia, Germany. The protocol was approved by the state authority for nature, environment and customer protection, North Rhine-Westphalia, Germany.

The spleen is the key site for removal of parasitized red blood cells and generation of immunity (22). Therefore, splenocytes were analyzed in all experiments performed in this study. Single-cell suspensions of splenocytes were generated by rinsing spleens with erythrocyte lysis buffer and washing with PBS supplemented with 2% FCS and 2 mM EDTA. Anti-CD3, anti-CD4, anti-CD8, anti-CD11c, anti-CD49d, anti-CD335, anti-CD19, anti-B220, anti-Ly6C, anti-MHC-II, and anti-IFN-γ (all BD Biosciences, Heidelberg Germany); anti-TNF-α and anti-Foxp3 (all eBioscience, Frankfurt, Germany); anti-CD317, anti-CD69, anti-CD64, anti-FcεRI, anti-γδTCR, anti-granzyme B, and anti-CD11a (all Biolegend, London, UK) were used as fluorescein isothiocyanate, pacific blue, phycoerythrin, BD Horizon V450, allophycocyanin or peridinin-chlorophyll protein conjugates. Dead cells were identified by staining with the fixable viability dye eFlour 780 (FVD) (eBioscience, Frankfurt, Germany). Intracellular staining for Foxp3 and granzyme B was performed with the Foxp3 staining kit (eBiocience, Frankfurt, Germany) according to the manufacturer’s recommendations. Cytokine production from freshly isolated splenocytes was measured by stimulating cells with 10 ng/ml phorbol 12-myristate 13-acetate (PMA, Sigma-Aldrich, München, Germany) and 100 µg/ml ionomycin (Sigma-Aldrich, München, Germany) in the presence of 5 µg/ml Brefeldin A and 5 µg/ml Monensin (eBioscience, Frankfurt, Germany) for 4 h at 37°C, followed by intracellular staining with the respective antibody cocktail using the Foxp3 staining kit. Flow cytometric expression analyses were performed with a LSR II instrument using DIVA software (BD Biosciences, Heidelberg Germany).

Blood samples were collected, incubated at room temperature, and centrifuged at 6,797 × g. Cytokines were quantified in sera by using a Luminex Screening assay (R&D Systems, Wiesbaden, Germany) and a Luminex 200 system with Luminex IS software (Luminex Corporation, MV’s-Hertogenbosch, Netherlands) according to the manufacturer’s instructions.

Statistical analyses were performed with One-Way ANOVA, Student’s t-test for parametric and Mann–Whitney test for non-parametric distributed data with significance set at the levels of *p < 0.05, **p < 0.01, and ***p < 0.001. Normality was tested using the D’Agostino–Pearson omnibus and Kolmogorov–Smirnov test. All analyses were calculated with GraphPad Prism 5.0 Software (GraphPad Software, La Jolla, CA, USA).

To study the impact of conventional CD11c^high DCs on the T cell response elicited by infection with P. yoelii, we made use of RosaiDTR^tg/tg/CD11c-cre^tg/wt mice (further referred as RosaiDTR/CD11c-cre mice). These mice express the DTR in CD11c⁺ cells due to CD11c driven cre-mediated excision of the floxed STOP cassette located upstream of the transgenic DTR. Daily treatment of double-transgenic mice with DT resulted in almost complete loss of splenic CD11c^highCD317⁻ DCs (Figures 1B–D) as analyzed by flow cytometry (Figure 1A). As controls RosaiDTR^tg/tg/CD11c-cre^wt/wt littermates (further referred to as RosaiDTR mice) were used which do not express the DTR. Injection of DT had no influence on the frequency and absolute numbers of CD11c^high DCs in these mice (Figures 1B–D). In addition to depletion of CD11c^high conventional DCs (CD11c^highCD8⁺ and CD11c^highCD8⁻ DCs), DT treatment also partially affects the CD11c^int cell compartment. We detected significant reduced frequencies of CD11c^intCD11b⁻Ly6C⁺B220⁺ plasmacytoid DCs (pDCs) in DT-treated non-infected and infected RosaiDTR/CD11c-cre mice in contrast to RosaiDTR littermates (Figure 1E). CD11c-cre mice express the cre recombinase in more than 90% of cDCs and also in 86% pDCs (20), which might explain depletion of pDCs in RosaiDTR/CD11c-cre double-transgenic mice after DT treatment. Moreover, we detected elevated frequencies of CD11c^intCD11b⁺CD64⁺FcεRI⁺MHCII⁺ cells in the spleen of P. yoelli-infected mice, which likely represent monocyte-derived DCs (moDCs) (23). Interestingly, DT treatment resulted in significant higher percentages of CD11c^intCD11b⁺CD64⁺FcεRI⁺MHCII⁺ cells in RosaiDTR/CD11c-cre mice than in control littermates at day 7 after P. yoelli infection (Figure 1F). The absolute number of splenocytes increased during P. yoelii infection, but did not significantly differ between cDC-depleted and control mice (Figure S1A in Supplementary Material) and percentages of NK cells as well as B cells were also similar between DT-treated RosaiDTR/CD11c-cre mice and RosaiDTR littermates (Figures S1B,C in Supplementary Material).

First, we investigated the effect of long-term DC deficiency on T cell activation in P. yoelii-infected mice. For this purpose, we treated RosaiDTR/CD11c-cre and RosaiDTR control mice with DT 1 day before infection and subsequently every day. At days 3, 7, and 10 post infection (p.i.), the phenotype of gated CD8⁺ and CD4⁺ T cells (Figures 2A and 3A) was analyzed by flow cytometry. IFN-γ production of splenic T cells was determined after re-stimulation with PMA and Ionomycin, due to the lack of identified endogenous T cell epitopes within the blood-stage P. yoelii parasites. As depicted in Figure 2, long-term depletion of CD11c^high DCs resulted in significantly reduced frequencies of IFN-γ-producing CD8⁺ T cells at day 7 p.i. (Figure 2B). Moreover, we detected lower percentages of CD8⁺ T cells expressing the early activation marker CD69 at day 3 p.i. (Figure 2C) and granzyme B (GzmB) at days 3 and 7 p.i. (Figure 2D). To investigate whether changes in the activation status of CD8⁺ T cells during long-term CD11c^high DC depletion is due to bystander effects or a direct result from antigen-specific stimulation, we analyzed the frequencies of CD11a-expressing CD8⁺ T cells in P. yoelii-infected mice. This approach was already described for the detection of antigen-specific CD8⁺ T cells during bacterial, viral, and also Plasmodium infections (24–26). The percentage of CD11a⁺ CD8⁺ T cells was very low in non-infected mice, while increased in P. yoelii-infected mice at day 7 p.i. (Figure 2E). Moreover, CD11c^high depletion resulted in significantly reduced frequencies of CD11a-expressing CD8⁺ T cells at day 7 after P. yoelii infection compared to controls (Figure 2E).

Phenotypic analysis of CD4⁺ T cells from P. yoelii-infected mice revealed that long-term DC deficiency interferes with IFN-γ production at day 7 p.i. (Figure 3B). In addition, the frequency of CD69-expressing CD4⁺Foxp3⁻ T cells was significantly decreased in DC-depleted mice at days 3, 7, and 10 after P. yoelii infection (Figure 3C). CD11a⁺CD49d⁺ staining has been shown to delineate previously activated CD4⁺ T cells from naive cells in Plasmodium-infected mice (27) and, therefore, represent T cells that were likely to be responding to a variety of Plasmodium antigens. Interestingly, long-term depletion of CD11c^high DCs led only to a slightly, albeit not significantly, reduced percentage of antigen-experienced CD11a⁺CD49d⁺ CD4⁺Foxp3⁻ T cells in P. yoelii-infected DT-treated RosaiDTR/CD11c-cre mice (Figure 3D).

Our previous studies revealed an expansion and activation of Foxp3⁺ naturally occurring Tregs in the course of P. yoelii infection (6, 7). To analyze whether CD11c^high DCs also influence the Treg compartment, we determined the frequency and activation status of CD4⁺Foxp3⁺ Tregs in DT-treated RosaiDTR/CD11c-cre and control mice at different time points after P. yoelii infection by flow cytometry. Interestingly, long-term depletion of CD11c^high DCs did not change the percentage of Foxp3⁺ Tregs (Figure 4A), but resulted in decreased frequencies of CD69-expressing CD4⁺Foxp3⁺ Tregs in non-infected mice and at day 3 p.i. (Figure 4B). These results suggest that CD11c^high DCs are not only important for the activation of CD8⁺ and CD4⁺ effector T cells but also for CD4⁺Foxp3⁺ Tregs.

In addition to analyzing the impact of DCs on the T cell response, we asked whether depletion of CD11c^high DCs modulates the systemic cytokine profile and parasite clearance. Therefore, we determined the level of pro-inflammatory cytokines in the serum of P. yoelii-infected DT-treated RosaiDTR/CD11c-cre and RosaiDTR controls at different time points p.i. by Luminex technology. As depicted in Figure 5, depletion of DCs resulted in significant lower production of IFN-γ (Figure 5A) and TNF-α (Figure 5B) at day 3 post P. yoelii infection. In addition, we detected decreased IFN-γ mRNA expression in the spleen of cDC-depleted mice in comparison to RosaiDTR control mice at day 3 p.i. (Figure S2A in Supplementary Material). To gain further insights into the cellular source of this cytokine, we determined the frequencies of IFN-γ expressing NK, NKT, and γδ T cells, because these cells have been described to produce elevated levels of IFN-γ at early time points during Plasmodium infection (28, 29). Well in line with these studies, we detected increased IFN-γ production by all of these cell types in P. yoelii-infected mice (Figures S2B–D in Supplementary Material). However, the frequencies of IFN-γ⁺CD335⁺CD3⁻ NK cells were not decreased, but the percentages of IFN-γ⁺CD335⁺CD3⁺ NKT cells and IFN-γ⁺γδTCR⁺ T cells were reduced in cDC-depleted mice compared to control littermates after infection (Figures S2B–D in Supplementary Material). These results suggest that reduced IFN-γ level in serum and spleen of cDC-depleted P. yoelii-infected mice are at least in part a result of lower frequencies of IFN-γ producing NKT cells and γδ T cells. In addition, we have recently shown that also CD11c⁺ DCs themselves produce elevated levels of IFN-γ and might, therefore, also contribute to higher IFN-γ production in control mice than in cDC-depleted mice during the early phase of P. yoelii infection (14).

Although DCs promote T cell activation and contribute to systemic production of pro-inflammatory cytokines during the early phase of P. yoelii infection, we detected no significant differences in parasite clearance between DT-treated RosaiDTR/CD11c-cre mice and control mice (Figure 5C). Hence, differences in T cell activation between long-term DC-depleted and control mice can directly be attributed to the presence of CD11c^high DCs and are not an indirect effect of lower antigen load.

Next, we asked whether the presence of DCs is critical at the onset of infection or whether they are able to provoke efficient T cell responses also during an established infection. Therefore, we depleted CD11c^high DCs from P. yoelii-infected RosaiDTR/CD11c-cre mice by DT treatment starting at day 4 p.i. As control RosaiDTR littermates were also treated with DT from day 4 until day 14 post P. yoelii infection. At day 7 and day 14 p.i., we analyzed the T cell phenotype. We did not observe any differences in the frequency of IFN-γ producing CD8⁺ T cells and CD4⁺ T cells (Figure 6A), GzmB-expressing CD8⁺ T cells and CD69-expressing CD4⁺Foxp3⁻ T cells (Figure 6B) at days 7 and 14 p.i. between mice depleted from CD11c^high DCs and controls. This indicates that effector T cell activation by CD11c^high DCs takes place during the initial phase of infection. However, DC-depletion at day 4 of P. yoelii infection resulted in significantly reduced frequencies of CD4⁺Foxp3⁺ Tregs (Figure 6C) and had no impact on parasite burden (Figure 6D) compared to DT-treated controls.

These results imply that long-term depletion of CD11c^high DCs during the entire P. yoelii infection modulates T cell activation and systemic production of pro-inflammatory cytokines, whereas depletion after establishment of infection has no impact on the effector T cell phenotype and parasite clearance.

To further corroborate our finding that DC-mediated priming of T cells at early time points during infection is required to provoke efficient T cell activation, we depleted CD11c^high DCs by a single injection of DT to RosaiDTR/CD11c-cre mice and RosaiDTR littermates only the day before P. yoelii infection. This approach resulted in DC deficiency at the time point of infection lasting until 48–72 h p.i. (Figure 7A). As depicted in Figure 7B, a single injection of DT to RosaiDTR/CD11c-cre mice 1 day before P. yoelii infection was sufficient to significantly reduce the frequency of CD69-expressing CD8⁺ and CD4⁺Foxp3⁻ T cells as well as CD4⁺Foxp3⁺ Tregs at day 3 p.i. (Figure 7B). In addition, we detected lower serum levels of IFN-γ and TNF-α in P. yoelii-infected DC-depleted mice in comparison to control littermates (Figure 7C) and observed no differences in parasite burden at day 3 p.i. (Figure 7D). Strikingly, these data reflect our results from analysis of mice with long-term depletion of CD11c^high DCs (Figures 1–5) underlining the importance of CD11c^high DCs in T cell priming at early time points during P. yoelii infection.