Two C. parvum strains, i.e., the commercially available Iowa C. parvum strain P102 (Waterborne, New Orleans, USA) and the Leipzig strain (gp60 IIaA15G2RI subtype, University of Leipzig, Germany), were used to investigate the interaction of C. parvum with human DCs in vitro. The Leipzig strain corresponds to the most commonly reported strain of C. parvum in Germany and other developed countries. C. parvum was maintained by serial passage of oocysts in 1-day-old calves at the Faculty of Veterinary Medicine of the University of Leipzig, as reported elsewhere (13). Oocysts of the Leipzig strain were isolated from feces of infected calves using a sedimentation/flotation method (27, 28) and maintained at 4°C in sterile phosphate-buffered saline (PBS) (pH 7.4) with penicillin/streptomycin (200 µg/mL; Sigma-Aldrich, Darmstadt, Germany) and amphotericin B (5 µg/mL; Sigma-Aldrich, Darmstadt, Germany). The C. parvum oocyst storage medium was exchanged at monthly intervals. Therefore, oocysts were pelleted (5,000 ×g for 10 min) and supplemented with fresh medium containing penicillin/streptomycin (13, 27). Before excystation, C. parvum oocysts were pelleted at 5,000 ×g for 5 min at 4°C to isolate sporozoites (13, 27). Then, acidified (pH 2.0) and sterile pre-warmed (37°C) 1× Hank’s balanced salt solution (HBSS; Sigma-Aldrich, Darmstadt, Germany) was used as oocyst excystation medium (10 min at 37°C). Thereafter, free-released sporozoites were pelleted and incubated at 37°C for 10 min in 1× HBSS with pH 7 (Sigma-Aldrich, Darmstadt, Germany). Once again pelleted, the sporozoites were resuspended in phenol red-free Roswell Park Memorial Institute (RPMI) 1640 cell culture media (Gibco, Darmstadt, Germany) supplemented with 0.3 g/mL l-glutamine, 10% fetal bovine serum (FBS; Gibco, Darmstadt, Germany), and 100 UI penicillin and 0.1 mg streptomycin/mL (both Sigma-Aldrich, Darmstadt, Germany). As the Leipzig strain was no longer available to us, we validated by flow cytometry that the Iowa strain activated MO-DCs to a similar extent. In phagocytosis assays, bona fide identical phagocytosis rates were obtained. The effect on DC mobility was comparable as well.

As a source of immature human DCs, we isolated CD14⁺ monocytes from human blood and generated monocyte-derived dendritic cells (MO-DC) using granulocyte-macrophage colony-stimulating factor (GM-CSF) and interleukin (IL)-4 as described (29). Briefly, buffy coats from healthy donors (n = 15) were obtained from the Center of Transfusion Medicine and Hemotherapy of the University Hospital of Giessen and Marburg (UKGM) in Giessen, Germany. The use of buffy coats for research purposes was approved by the local ethics committee of the Justus Liebig University Giessen and UKGM. For the entire isolation procedure and cell culture, exclusively pyrogen-free media, buffers, and equipment were used to exclude unintentional pre-activation and maturation of human DCs; 15 mL aliquots of resuspended buffy coats were diluted 1:1 with DC medium RPMI 1640 (Gibco, Darmstadt, Germany), supplemented with 10% fetal calf serum (FCS) (Sigma-Aldrich, Darmstadt, Germany) and 2 mM l-glutamine and 1 mM sodium pyruvate (both Sigma-Aldrich, Darmstadt, Germany) and overlaid on 15 mL of Biocoll^® (Biochrom GmbH, Berlin, Germany) with a density of 1,077 g/mL. Samples were centrifuged for 30 min at 2,500 ×g without break. Peripheral blood mononuclear cells (PBMCs) were harvested at the interface of Biocoll^® and supernatant, washed several times in MACS^® separation buffer with 2% bovine serum albumin (BSA; Miltenyi Biotec GmbH, Bergisch Gladbach, Germany), and subjected to positive selection of CD14⁺ monocytes using immunomagnetic beads and reagents from Miltenyi Biotec GmbH. Briefly, 2.5 × 10⁸ PBMCs and 500 µL Miltenyi human CD14⁺ MicroBeads^® were incubated in a total volume of 2.5 mL MACS^® buffer for 15 min at 4°C, and CD14⁺ monocytes were isolated using LS Columns and QuadroMACS™ separator according to the recommendations of the manufacturer. A purity of CD14⁺ monocytes of >95% was obtained (data not shown). A total of 3 × 10⁶ monocytes suspended in 3 mL DC medium supplemented with 1,000 U/mL human GM-CSF (ImmunoTools GmbH, Friesoythe, Germany) and 1,000 U/mL human IL-4 (ImmunoTools GmbH) were incubated in 6-well plates (day 0). On day 3, half of the medium was replaced with fresh DC medium with double-concentrated cytokines. Immature MO-DCs were harvested on day 6 of culture and further cultured for the following assays in DC medium either with or without further stimulation. Assays were performed in 24-well plates (Thermo Fisher Scientific, Braunschweig, Germany) in a total volume of 500 µL DC medium with 1 × 10⁵ DCs/well or in 96-well plates (Thermo Fisher Scientific, Braunschweig, Germany) in a total volume of 200 µL with 2 × 10⁴ DCs/well.

In order to visualize DC–parasite interactions at ultrastructural level via scanning electron microscopy (SEM), viable C. parvum oocysts/sporozoites (ratio 1:3) were co-cultured with human DCs for 120 min on coverslips pre-coated with poly-l-lysine 0.01% (10 mm/diameter; Thermo Fisher Scientific, Braunschweig, Germany). In brief, co-cultures were performed at 37°C and 5% CO₂ for the entire experimental period. In each well, 2 × 10⁵ DCs and 6 × 10⁵ of the parasite stages were seeded. The cells were then fixed in 2.5% glutaraldehyde (Merck, Darmstadt, Germany) and post-fixed with 1% osmium tetroxide (Merck, Darmstadt, Germany), rinsed in distilled water, dried, CO₂-treated to the critical point, and lastly sputtered with gold particles. SEM analysis was conducted at the Institute of Anatomy and Cell Biology at the Justus Liebig University Giessen, Germany, utilizing a scanning electron microscope (Philips XL30^®, Eindhoven, The Netherlands) equipped with a digital camera.

Immature human myeloid DCs from day 6 of culture were harvested, resuspended in fresh DC medium, and exposed to 2 × 10⁵ C. parvum oocysts or to lipopolysaccharide (LPS; Sigma-Aldrich, Darmstadt, Germany) or left without further stimulation (24-well plates, total volume of 500 µL DC medium, 1 × 10⁵ DCs/well overnight). For flow cytometry analysis, differently treated DCs were all harvested on day 7 of culture. Aliquots of approximately 5 × 10⁵ DCs were washed in PBS/2% FCS and incubated for 15 min in 10% heat-inactivated human serum to block Fc receptors. After washing, cells were resuspended in sterile 50 µL PBS/2% FCS with 0.5 µg fluorescence-conjugated monoclonal antibodies (mAbs) directed against the CD surface markers CD1a, CD3, CD14, CD19, CD83, and HLA-DR (BioLegend, Amsterdam, Netherlands) or CD11b, CD58, CD40, and CD86 (ImmunoTools GmbH, Friesoythe, Germany) or with isotype control mAbs labeled with the corresponding fluorescence conjugates (BioLegend, Amsterdam, Netherlands). Some samples were fixed and rendered mAb-permeable using FoxP3^® fixation (Miltenyi Biotec GmbH, Bergisch Gladbach, Germany) before staining. After incubation for 30 min in the dark, stained cells were washed twice in 1 mL PBS/2% FCS, resuspended after centrifugation in 0.5 mL PBS/2% FCS, and analyzed using a BD Accuri™ C6 flow cytometer (BD Biosciences, San Jose, CA, USA).

To assess potential DC-derived extracellular traps (ETs) after parasite exposure, human DCs on day 6 of culture and C. parvum sporozoites/oocysts (DC:C. parvum ratio 1:3) were co-cultured [37°C, 5% CO₂, 120 min, on sterile glass coverslips (15-mm diameter; Thermo Fisher Scientific, Braunschweig, Germany) that had been pre-treated with 0.01% poly-l-lysine (Sigma-Aldrich, Darmstadt, Germany)]. Cell fixation was conducted with 4% paraformaldehyde (Merck, Darmstadt, Germany), and samples were kept at 4°C until final experimentation. To analyze whether DCs are able to release ETs against parasite stages, cells were treated with specific antibodies raised against typical ET components, i.e., anti-myeloperoxidase (MPO) antibodies (1:500; Abcam-ab208670, Darmstadt, Germany) and anti-global histone antibodies (H1, H2A/H2B, H3, and H4; 1:500; Merck, Darmstadt, Germany) [overnight at room temperature (RT)]. Samples were, therefore, subjected to three washes with sterile PBS (Sigma-Aldrich, Darmstadt, Germany) and a 60-min incubation in 2% BSA (Sigma-Aldrich, Darmstadt, Germany) supplemented with 0.3% Triton X100 (Thermo Fisher Scientific, Braunschweig, Germany). Samples were then incubated with primary antibodies overnight followed by incubation in secondary antibody solutions [1:500: IgG #A11005-Alexa 594 goat anti-mouse and IgG #A11008-Alexa 488 goat anti-mouse (Thermo Fisher Scientific, Braunschweig, Germany)] (60 min, RT, total darkness) after three washing steps with sterile PBS (Sigma-Aldrich, Darmstadt, Germany). Anti-fading buffer with DAPI (Thermo Fisher Scientific, Braunschweig, Germany) was used to mount the samples after they had been rinsed three times with sterile PBS. A confocal microscope (Nikon ECLIPSE Ti2, Melville, NY, USA) and an inverted epifluorescence microscope IX81^® (Olympus, Tokyo, Japan) equipped with an XM10^® digital camera (Olympus, Tokyo, Japan) were both used for analysis.

Immature human MO-DCs were harvested on day 6 of culture, resuspended in fresh medium, stimulated with either LPS (Sigma-Aldrich, Darmstadt, Germany) or C. parvum stages as indicated, or left without further stimulation. Assays were performed in 96-well plates (Thermo Fisher Scientific, Braunschweig, Germany) in triplicates. Independent ELISAs were performed with MO-DCs generated from three different donors (n = 3), yielding comparable results. Commercial ELISA kits were purchased from BioLegend (IL-6) and ImmunoTools (IL-8) and used according to the recommendations of manufacturers. First, capture antibodies were transferred to each well and incubated overnight at RT. After completely removing capture antibodies and washing, a blocking buffer was added and incubated for 1 h at RT. Then, the blocking buffer was removed, and standard and samples were diluted in reagent diluent as indicated and transferred into respective wells. After washing five times with washing buffer, detection antibodies were added, and samples were incubated for 2 h at RT. After washing, poly-HRP-streptavidin (poly-HRP-streptavidin; Hamburg, Germany) was added to each well and incubated for 30 min at RT. After washing, pre-warmed TMB (Sigma-Aldrich, Darmstadt, Germany) was added to each well and incubated for up to 60 min at RT under visual control. Samples were finally analyzed in a Tristar^® microplate reader (Berthold Technologies, Bad Wildbad, Germany) at 450 nm.

To distinguish between C. parvum stages attached to the surface of DCs and phagocytosed parasites, a phagocytosis assay was performed (30), which allows for the discrimination of intra- and extracellular fluorescein-labeled particles by trypan blue quenching. Therefore, C. parvum oocysts and sporozoites were labeled with NHS-fluorescein (Sigma-Aldrich, Darmstadt, Germany). Saccharomyces cerevisiae specimens were equally processed and used for positive controls as described elsewhere (30). However, in contrast to yeast, C. parvum oocysts/sporozoites were not subjected to a 99°C heat-inactivation step and were treated with reduced centrifugation forces to keep them alive. Parts of the C. parvum samples were incubated for 30 min at 55°C to kill them before labeling, thereby representing a further control to assess whether sporozoites actively invade human DCs. After visual control of equal fluorescent dye labeling, parasites were added to DCs for 3–4 h to allow phagocytosis. In general, sporozoites and oocysts were distinguishable from one another as well as from DCs by size in FSC/SSC. Moreover, they were detectable in the fluorescence channel FL-1 due to labeling with NHS-fluorescein. After incubation for phagocytosis, cells were harvested and analyzed with and without the addition of trypan blue (Sigma-Aldrich, Darmstadt, Germany). Trypan blue quenches the green fluorescence of free parasites but not of phagocytosed ones, as living cells remain negative for trypan blue, and direct contact with trypan blue is needed for quenching. Green fluorescence was detected in FL-1, while red fluorescence due to quenching was detected in FL-4. As a further control, DC-mediated phagocytosis was blocked with cytochalasin D (10 µg/mL; Merck, Darmstadt, Germany).

Human DCs on day 6 of culture were either subjected to phagocytosis without pre-stimulation or in some experiments incubated with LPS overnight before C. parvum oocysts/sporozoites were added on day 7. In total, samples were monitored for 3–5 h by fluorescence microscopy. Therefore, DCs were resuspended in an imaging medium including 20 μM Hoechst (Thermo Fisher Scientific, Braunschweig, Germany) and 0.1% BSA (Sigma-Aldrich, Darmstadt, Germany). Sterile ibidi^® plastic cell culture plates (dish 35 mm² with low profile, ibidi^®, Gräfelfing, Germany) were used to seed 1 mL of this cell solution (0.5 × 10⁶ DCs) before incubation at 5% CO₂ and 37°C in an incubation cell chamber (ibidi^® dish 35 mm²). For DC settling and to prevent condensation on the ibidi^® incubation chamber lid, a 30-min resting period at 37°C was employed. Subsequently, 2 × 10⁵ NHS-fluorescein-labeled oocysts/sporozoites were added into the chamber and incubated for 3–5 h. Using a Fluo-3D Cell Explorer^® (Nanolive, Lausanne, Switzerland) equipped with a ×60 magnification (λ = 520 nm, sample exposure 0.2 mW/mm²) and a field depth of 30 µm, the samples were analyzed. Image capture was configured for refractive index (RI) and Hoechst (blue channel) detection (Thermo Fisher Scientific, Braunschweig, Germany) acquiring images every minute for 3–5 h. Thereafter, each channel was independently exported using Steve^® software v.2.6 (Nanolive, Lausanne, Switzerland) and processed using Image J^® software (Fiji version 1.7; NIH, USA). Using RI values from the images, digital staining was accomplished thereafter to achieve 3D holotomographic imaging.

To estimate glycolytic and oxidative responses, ECAR and OCR values of human MO-DCs were quantified using the metabolic extracellular flux analyzer Seahorse XFp^® (Agilent, Ratingen, Germany) according to the recommendations of the manufacturer (31); 1 × 10⁵ cells in 50 µL were carefully transferred into each well of an 8-well XF^® Seahorse analyzer plate (Agilent, Ratingen, Germany), which had been coated previously with 0.001% poly-l-lysine for 30 min (Sigma-Aldrich, Darmstadt, Germany). Then, 50 μL of XF^® assay medium (Agilent, Ratingen, Germany) was administered to control wells. Subsequent to a 45-min incubation period at 37°C without CO₂ supplementation, 130 µL of XF^® assay medium was supplemented to each well to achieve a final volume of 180 µL per well. After 6 cycles of baseline measurement in the Seahorse XF^®, a suspension of C. parvum (3 × 10⁵ oocysts/20 μL) or 20 µL aliquots containing other microbial components or pure XF^® assay medium were added via instrument-own injection ports, and ECAR/OCR values were continuously detected for additional 30 cycles. OCR and ECAR registries were evaluated using Wave^® software (Desktop Version, Agilent, Ratingen, Germany) as previously stated elsewhere (31, 32).

For this study, human MO-DCs were generated in sufficient purity and viability in an immature state to allow for subsequent maturation by co-culture with C. parvum. To this end, pyrogen-free disposable equipment and media were employed for the entire isolation procedure and cell culture to exclude unintentional pre-activation and maturation of the MO-DCs. CD14⁺ monocytes were enriched from PBMCs by positive selection with paramagnetic beads attached to anti-CD14 antibodies. CD14 is part of the LPS receptor of monocytes and macrophages but cannot signal by itself, as it has no intracellular domain. The purity of positive selected CD14⁺ monocytes is higher, fewer particles like thrombocytes are included, which may interfere, and truly immature DCs can be generated as frequently and reliably as shown in numerous publications including our own (28) and this study. As shown in Supplementary Figure 1A , the purity of CD14⁺ monocytes was >95% in the positive fraction. MO-DCs harvested on day 6 of culture following cultivation with GM-CSF and IL-4 by gentle rinsing of the non-adherent MO-DCs again yielded purity > 95% by flow cytometry. Stimulation with LPS indicated upregulation of all activation markers, as expected ( Supplementary Figure 1B ). The viability of monocytes and MO-DCs as indicated by trypan blue staining was >95% as well. If not mentioned otherwise in the text, all immature DCs were harvested on day 6 of culture, incubated in fresh medium with or without C. parvum or other stimuli, and analyzed depending on the assay either the same day or on day 7. Regardless of stimulation, all samples of the same assay were incubated under identical conditions for exactly the same time.

The elaborated structure of DCs is of functional relevance. Veiled structures are involved in migration and interaction with antigens (see Supplementary Videos ), and ultrathin dendrites enlarge the surface and enhance the chance to interact with naïve T cells, detecting fitting peptides in context with MHC molecules in the T-cell areas of the draining lymph nodes. The SEM-based ultrastructural analysis identified both types of dendrites upon MO-DCs cultured alone or co-cultivated with C. parvum oocysts. MO-DCs are morphologically indistinguishable from non-exposed control cells ( Figure 1A ). C. parvum oocysts interacting with DCs are indicated by arrows.

As sentinel cells of the immune system, DCs express an extraordinarily high number of pattern recognition receptors (PRRs) to identify invasive pathogens (14, 15) including prototypic molecules of major groups of microorganisms like yeasts and gram-positive or gram-negative bacteria. Work on human cell lines and mouse models hint at a potential role of TLR-3 or TLR-4 in the detection of C. parvum (33, 34), but even immune-suppressive mechanisms of C. parvum could not be ruled out. Thus, we used the gold standard TLR-4 ligand LPS and C. parvum oocysts either alone or in combination in the first experiment to stimulate MO-DCs and analyzed the effect on the expression of HLA-DR antigen-presentation molecules on MO-DCs. Both stimuli either alone or in combination enhanced the surface expression of HLA-DR ( Supplementary Figure 2A ), indicating maturation of DCs and not inhibition.

C. parvum-driven stimulation of MO-DCs was also apparent microscopically by boosted cluster formation ( Figure 1B and Supplementary Figure 2B ). During the generation of DCs from monocytes by culture with GM-CSF and IL-4, cells typically build dendrites and form clusters of cells. When DCs are harvested, these aggregates disintegrate due to resuspension. Without further stimulation, DCs then remain more or less evenly distributed in the wells. In current experiments, both LPS stimulation and C. parvum exposure equally induced significant cluster formation of MO-DCs ( Figure 1B , Supplementary Figure 2B ). In Figure 1B , single oocysts, single MO-DCs, and MO-DC clusters are illustrated and indicated by arrows. Strikingly, a concentration as low as 2.5 pg/mL of LPS was sufficient to induce DC aggregation ( Supplementary Figure 1A ), indicating that human DCs effectively responded to such trace amounts of microbial substances with maturation. Moreover, DC aggregation after C. parvum exposure was direct proof of parasite-driven MO-DC movements, interactions, and activation. Consistent with this finding, we found that parasite exposure led to an enhanced expression of the adhesion molecules CD58 (LFA-3) and CD11b (integrin α-M) on the surface of MO-DCs ( Figure 1C ). An increase of adhesion molecules is required to improve interaction with T cells in vivo, and clustering indicates active movement as required in vivo to move to the place of infection to take up antigens, then to the lymph vessels, and finally to the T-cell areas in the lymph nodes.

Additionally, we used live cell 3D holotomographic microscopy (3D Cell Explorer^®, Nanolive, Lausanne, Switzerland) to better illustrate the early effects of C. parvum oocyst/sporozoite confrontation on MO-DC shape and motility. Overall, migratory activities of DCs toward C. parvum were monitored either with or without LPS pre-stimulation overnight on day 7 of culture. As visible in Videos 1 and 2 (see the Supplementary Material ), several DCs were highly motile also in the case of double stimulation (C. parvum + LPS), indicating that motility was not suppressed by C. parvum contact. While DCs moved over comparable distances with or without LPS pre-stimulation, LPS-treated cells seemed more adherent and changed direction more frequently, while cells without LPS pre-stimulation moved straight on for longer distances. The morphology of DCs and tracks of selected cells with calculated distances covered per time are depicted ( Figure 1D ).

A typical activation-driven expression pattern was observed following stimulation of immature MO-DCs on day 6 with or without C. parvum stages or LPS overnight followed by analysis on day 7 of culture. Unstimulated and stimulated DCs had already downregulated CD14 ( Figure 2A ). The antigen-presenting molecule CD1a, which allows DCs to present microbial lipid antigens to T cells, was already upregulated ( Figure 2B ). The latter marker indicated that the cells were healthy and survived quantitatively during cultivation. Human HLA-DR molecules are required to present microbial peptides to (naïve) T cells. Since exclusively professional antigen-presenting cells (APCs) express MHC class II molecules while nearly all cells express MHC class I molecules, human MHC class II (HLA-DR) was selected for the current analysis. As expected, HLA-DR was already expressed on isolated monocytes and enhanced on day 6 of culture (not shown). This molecule was further upregulated on day 7 by both LPS and C. parvum stimulation ( Figure 2C ). However, the expression levels of HLA-DR were quite heterogeneous, which may be based on a high number of molecules still being located on intracellular membranes or within intracellular vesicles and not yet loaded with peptides. Therefore, permeabilized MO-DCs were additionally tested under identical culture conditions. Indeed, permeabilization resulted in more focused staining, thereby confirming parasite-driven HLA-DR upregulation ( Figure 2D ). Of note, both processes, i.e., increased synthesis and peptide loading of new HLA molecules and peptide loading of intracellularly stored but unloaded HLA molecules at the time point of pathogen exposure, increase the likelihood that sufficient numbers of microbial peptides are presented among a vast majority of irrelevant peptides. This process is essential to induce a proper adaptive immune response against pathogenic microorganisms. Obviously, in the case of C. parvum stage exposure, these reactions were efficiently driven, finally resulting in MO-DC activation and maturation.

Moreover, CD83, a reliable frequently used maturation marker of human DCs, was upregulated upon stimulation with either LPS or C. parvum ( Figure 2E ). The same was true for the costimulatory molecules CD40 ( Figure 2F ) and CD86 ( Figure 2G ) when compared to unstimulated control cells. Together with CD80, they represent the most important costimulatory molecules of DCs, which are especially needed at high densities to successfully stimulate naïve T cells. In contrast to CD40 and CD86, CD80 was already highly expressed on immature DCs (data not shown). Taken together, all markers displayed indicated the maturation of MO-DCs upon parasite exposure ( Figure 2 ). The data are representative of three independent experiments using DCs generated from different blood donors (n = 3).

DCs secrete a number of cytokines and chemokines to induce and regulate adaptive immune responses (15). In the current study, MO-DCs were exposed to C. parvum stages, and IL-6- and IL-8-based responses were quantified. Overall, IL-6 is an important molecule for the induction of Th-1 cells, Th-17 cells, and regulatory T cells. IL-8 represents a chemokine that is relevant for attracting other effector cells of the innate immune system like PMNs, which were already shown to hamper C. parvum host cell invasion by phagocytosis or NETosis (35, 36). In preliminary assays, the functionality of the tests was confirmed using LPS stimulation as positive controls ( Figure 3 , left panels). While the expression level of both factors was low without stimulation, trace amounts of LPS (2.5 pg/mL) were sufficient to yield almost maximal secretion levels. Thus, the culture supernatants had to be diluted fivefold (IL-6) or 10-fold (IL-8) compared to the recommendations of the manufacturers to avoid exceeding the upper detection limit. Overall, LPS-induced IL-6 and IL-8 secretion was induced at least 30- and 70-fold, respectively ( Figure 3 , right panels). Referring to C. parvum exposure, MO-DCs responded with high significance (C. parvum vs. control: p ≤ 0.0001) with IL-6 ( Figure 3A ) and IL-8 secretion ( Figure 3B ). When stimulating MO-DCs with different numbers of parasites, a slight dose dependency of cytokine production was apparent. The different donors showed typical individual quantitative variations in their responses, but in principle, all reacted by strong cytokine production. Of note, the strength of parasite-driven reactions was considerably high since the total cytokine/chemokine levels upon C. parvum stimulation following dilution of the supernatants even exceeded cytokine levels obtained from LPS-stimulated DCs and as well the highest values of the standard curve, so the cytokine concentration of IL-6 and IL-8 is not displayed in the figure. C. parvum stimulation induced, however, several thousand pg/mL of IL-6 and IL-8, while unstimulated cells expressed only less than 10 pg/mL of these cytokines. Moreover, oocysts and excystation material from the same number of oocysts induced high levels of IL-6 and IL-8 secretion, indicating that protein suspensions from C. parvum were sufficient for human MO-DC activation ( Supplementary Figure 3 ).

C. parvum oocyst and sporozoite stages are reported to induce NET formation in human and bovine PMNs (33), and a recent publication suggested ET formation by CD123⁺ plasmacytoid DCs (37). Here, we studied, therefore, whether human MO-DCs may perform ETosis upon exposure to C. parvum stages. Overall, both DNA and histones were exclusively detected in the nuclear compartment of MPO-labeled DCs. Stimulation with neither C. parvum nor PMA induced ET formation in human MO-DCs ( Supplementary Figure 4 ).

In general, a robust immune response is driven when DCs directly phagocytize microorganisms. DCs may phagocytize any particles <1–5 µm, but uptake efficiency is markedly enhanced if DCs sense microbial antigens by PAMP receptors or based on complement/antibody-mediated opsonization. As phagocytosis is a fundamental and efficient mechanism of the human immune defense, we here intended to assess DC phagocytic capacities upon C. parvum confrontation (38, 39).

Light microscopy implicated that C. parvum oocysts were efficiently phagocytized by activated human DCs (not shown). For direct proof of phagocytosis and quantification, we applied a phagocytosis assay (29), which allows the discrimination between intracellular and extracellular fluorescein-labeled particles. For positive controls, fluorescein-labeled S. cerevisiae was included in the experiments. A mixture of fluorescence-labeled C. parvum sporozoites and oocysts was used to expose MO-DCs on day 6 of culture ( Figures 4A , 5A ). When assessing parasite fractions, flow cytometry analysis unveiled two distinct populations in FL-1/FL-4 with fluorescence in FL-1 (gates R2 and R3). Particles with higher FL-1 fluorescence (gate R3) were smaller than particles from gate R2 as shown in FSC/SSC plots, indicating that C. parvum sporozoites were labeled with higher fluorescence intensity than oocysts ( Figure 5B ). This was in line with fluorescence microscopic images of sporozoites/oocysts. However, a small proportion (approximately 10%–15%) of oocysts displayed a higher fluorescence intensity than other oocysts in microscopy ( Supplementary Figure 5 ). They may contribute to the signals between gates R2 and R3 and may represent the less abundant thin-walled type of oocysts. To distinguish between DC-adherent and DC-internalized parasites, trypan blue was added to the medium and adjusted to a pH of 4.0 using a citrate buffer just before analysis. Consequently, the green fluorescence of external parasites was quenched ( Figure 4B , arrowheads). Intracellular parasites, in contrast, were protected from trypan blue staining and, therefore, still displayed a green fluorescence ( Figure 4B , arrows). Uptake of either C. parvum oocysts or sporozoites was clearly detectable ( Figure 4C ). The general gating strategy is depicted in Figures 5A, B . In Figure 5C , cells were treated with trypan blue, and DCs were gated by size in FSC/SSC. Consequently, all cells are in the upper left quadrant because DCs were not labeled (left panel). Some DCs previously exposed to fluorescein-labeled C. parvum (middle panel) or S. cerevisiae (right panel) for 3–4 h, however, displayed a green fluorescence (FL-1, upper right) due to phagocytized microbes.

Overall, the current data indicated that ~25% of DCs indeed phagocytized C. parvum stages ( Figure 5C ). In contrast, nearly all DCs took up S. cerevisiae (96.3%, Figure 5C ) during the same time period, though S. cerevisiae is bigger and uptake is more difficult. Maybe DCs were activated more efficiently by S. cerevisiae based on yeast-derived PAMPs (e.g., TLR-2-ligand zymosan). The phagocytosis inhibitor cytochalasin D blocked the uptake of C. parvum, while heat killing of C. parvum (55°C for 30 min) did not influence this effector mechanism, indicating that uptake was truly by DC-derived phagocytosis and not by sporozoite invasion ( Figure 5D ).

Energy for human DC maturation is provided by different metabolic pathways including glycolysis, detectable by an increase in ECAR, and/or oxidative phosphorylation, detectable by an increase in OCR. Stimulation of immature MO-DCs on day 6 of culture with different stimuli like LPS, S. cerevisiae, flagellin, or PMA + ionomycin all increased ECAR, thereby indicating enhanced glycolytic responses ( Supplementary Figure 6 ). In contrast, exposure of MO-DCs to C. parvum stages failed to induce glycolytic responses ( Figure 6 ). However, the combination of LPS and C. parvum triggered equal glycolytic responses, and LPS alone denied any inhibitory effect of the parasite. In contrast to ECAR, oxidative responses of MO-DCs were indeed induced by C. parvum stages since OCR slowly but steadily increased following parasite exposure, to a similar extent as for LPS stimulation. In line with this, double stimulation with C. parvum + LPS showed additive effects on OCR values ( Figure 6 ).