Adult F. hepatica flukes were collected from the liver of infected sheep, as previously described (31). The whole F. hepatica tissue extract containing a mixture of somatic antigens (FhSA) was obtained by homogenizing two flukes in sterile PBS using a glass homogenizer, followed by a sonification step (4 x 30 sec with intervals of 20 sec on ice) and centrifugation for 25 min (18 000 x g at 4°C). fhESPs were collected as described previously (31). Briefly, adult flukes were first washed in PBS and incubated in RPMI 1640 at 37°C for 2 hours, then incubated for ~32 hours at 37°C in RPMI 1640 supplemented with 100 U/ml of penicillin and 0.1 mg/mlof streptomycin. The media was collected and immediately frozen at -80°C, and replaced with fresh media every 2-3 hours. Later, batches were combined, spun down (8,000 × g, 10 min, 4°C) and concentrated until ~1.5 mg/ml using Amicon^® Ultra-15 Centrifugal Filter Unit with a 3 kDa cut-off (Millipore). Protein concentration was determined by BCA assay (Pierce/Thermo Scientific).

Total RNA was isolated from adult F. hepatica fluke using a Total RNA mini Plus kit (A&A Biotechnology) and reverse transcribed into cDNAs using a Maxima™ First Strand cDNA Synthesis Kit for RT-qPCR (Thermo Scientific) according to the manufacturer’s protocol. The total cDNA was used as a template for FABP1 (GenBank: M95291.1) and FABP5 (GenBank: KJ713302.1) amplification, respectively. Full coding sequences were cloned in frame into the pPICZαB vector (Invitrogen) using PstI and XbaI restriction sites. The primer sequences used were: [F_FhFABP1_PstI] ATT CTG CAG GGA TGG CTG ACT TTG TGG, [R_FhFABP1_XbaI] TCT CTA GAC TCG CTT TGA GCA GAG TGG, [F_FhFABP5_PstI] CCG CTG CAG TCA TGT CTG GAT TTA TCG G, [R_FhFABP5_XbaI] GGC TCT AGA TCT TTG ATG CGT TGG TAT CTC C. The integrity of the constructs was verified by nucleotide sequencing. Recombinant plasmids were then linearized with PmeI and transformed into Pichia pastoris X33 strain using electroporation method. The expression of recombinant FABPs was performed in Buffered complex medium containing 0.5% methanol at 29°C for 96 h. The protein was purified on a Ni-NTA resin column (Macherey-Nagel) and the eluted fractions were concentrated and dialyzed against PBS using the same Amicon system as described above. Endotoxins were removed using Endotoxin Removal Spin Columns (Pierce) and the final eluate was passed through a 0.22 µm filter. The presence of purified recombinant FABPs was confirmed by SDS-PAGE and Western blotting analysis. Cathepsin L5 (fhCL5) was produced as previously described (32). Protein concentrations were determined by BCA assay (Pierce/Thermo).

Human monocytes were isolated from concentrated peripheral blood (buffy coat) donated with informed consent by healthy anonymous volunteers at Sanquin bloodbank (Amsterdam, The Netherlands). Following gradient centrifugation and positive selection using CD14 MACS beads (Miltenyi Biotec, Germany), as previously described (33), monocytes were next differentiated to moDCs using human rGM-CSF (20ng/ml, BioCource/Life technologies) and rIL-4 (0.86ng/ml, R&D Systems) for 5-6 days, with media refreshment on day 3. Immature moDC (0.5x10⁶/ml) were stimulated with 100 ng/ml LPS (E. coli 0111 B4 strain, InvivoGen, San Diego) on day 5-6 in the presence or absence of F. hepatica molecules fhSA (50 μg/ml), fhESPs (25 μg/ml), recombinant molecules fhFABP1 (10, 25, 50 μg/ml), fhFABP5 and fhCL5 (25 μg/ml each). After 48 hours of stimulation, cell supernatants were collected for cytokine/chemokine analyses and cells were stained with Fixable Aqua Dead Cell Stain Kit (Invitrogen/Thermo Scientific) for determination of cell surface expression of co-stimulatory molecules by FACS (FACSCanto II, BD Bioscience) using various antibodies ( Table S1 ). In addition, primed moDC (1x10⁴ cells) were co-cultured with CD40L-expressing J558 cells (1:1) for 24 hours, followed by supernatant collection. Cytokine (IL-12p70, IL-6, IL-10) and chemokine (CXCL11, TSP-1) concentrations were determined using commercial ELISA kits ( Table S1 ).

For analysis of T cell polarization, moDCs primed with LPS and F. hepatica molecules were co-cultured with allogenic naïve CD4+ T cells for 6-7 days in the presence of staphylococcal enterotoxin B (10 pg/ml) with or without the addition of a chemokine mix (CCL19, CXCL9, CXCL11; 0.33ng/ml of each (Peprotech), α-TSP1 (clone 133, Sigma-Aldrich) or isotype control IgG2b (BioLegend, Ultra LEAF Purified) antibodies (10 µg/ml). On day 7, cells were harvested and transferred to a 24 well plate and cultured in the presence of human rIL-2 (10U/ml, R&D Systems) for 4 additional days. Intracellular cytokine production was analyzed after 6 hours restimulation with phorbol myristate acetate (100 ng/ml) and ionomycin (1 µg/ml) with addition of brefeldin A (10 µg/ml) during the last 4 hours. Subsequently, the cells were stained with Fixable Aqua Dead Cell Stain Kit (Invitrogen) for 15 min at RT, fixed with 1.9% formaldehyde (Sigma-Aldrich), permeabilized (eBioscience™ Permeabilization Buffer) and stained with anti-IL-4, anti-IL-10 and anti-IFNγ for 30 min at RT. Alternatively, 1x10⁵ T cells were restimulated using anti-CD3 and anti-CD28 (BD Biosciences). 24 hours after restimulation, supernatants were collected, and IL-10 production by T cells was measured by ELISA (BD Bioscience). For analysis of transcription factors, cells were first stained with Fixable Aqua Dead Cell Stain Kit (Invitrogen) for 15 min at RT and fixed with FoxP3/Transcription Factor Staining Buffer Set (Invitrogen, for FOXP3 detection) for 1 hour at 4°C. Then, cells were washed twice with permeabilization buffer (eBioscience) and stained with CD3, CD4, FOXP3, GATA3, and T-bet antibodies ( Table S1 ). All samples were run on a FACSCanto II or BD LSR II and analyzed using FlowJo (Version vX.0.7 TreeStar).

To analyze the suppressive capacity of moDCprimed T cells, 5 x 10⁴ moDCs were co-cultured with 5 x 10⁵ naïve CD4⁺ T cells for 6 days. These T cells (test T cells) were harvested, washed, counted and irradiated (3000 RAD) to prevent expansion. Bystander target T cells (responder T cells), which were allogeneic memory T cells from the same donor as the test T cells, were labeled with 0.5 μM cell tracking dye 5,6-carboxy fluorescein diacetate succinimidyl ester (CFSE). Subsequently, 5 x 10⁴ test T cells, 2.5 x 10⁴ responder T cells, and 1 x 10³ LPS-stimulated DC were co-cultured for an additional 6 days. Proliferation was determined by flow cytometry, by co-staining with CD3, CD4 and CD25 antibodies ( Table S1 ).

The supernatants collected from LPS-stimulated moDCs with or without fhFABP1 (25 μg/ml) were pooled from 4 donors in equal volume and subjected to the Proteome Profiler Human XL Cytokine Kit (R&D Systems) according to the manufacturer’s instructions. Pre-blocked nitrocellulose membranes of the Human Cytokine Array were incubated with 1 ml of pooled supernatants and detection antibody cocktail overnight at 4°C on a rocking platform. The membranes were washed three times with 1× Wash Buffer (R&D Systems) to remove unbound proteins. Chemiluminescent detection reagents were applied to detect spot densities. Membranes were exposed to X-ray film and analyzed using the manufacturer’s image analysis software (Quantity One).

Immature moDC (0.5 x 10⁶/ml) were stimulated with 100 ng/ml LPS on day 5-6 in the presence or absence of fhFABP1 (25μg/ml). After 5 h of stimulation, cells were harvested, washed two times with ice-cold PBS and snap-frozen. Total RNA was isolated with an RNA purification kit (Macherey-Nagel) and was subjected to poly(A) enrichment using NEBNext Poly(A) mRNA Magnetic Isolation Module (New England Biolabs). Strand-specific cDNA libraries were constructed with NEBNext^® Ultra™ II Directional RNA Library Prep Kit for Illumina^® (New England Biolabs). Sequencing was performed using NovaSeq6000 (Illumina) sequencer in PE150 mode. The subsequent quality control steps and computational analyses are described in details in the Supplementary Method . Data are publicly available on NCBI (GSE197462).

All mouse experiments were performed in accordance with the Guide for the Care and Use of Laboratory Animals of the Institute for Laboratory Animal Research and have received approval from the university Ethical Review Board (Leiden University Medical Center, Leiden, The Netherlands; DEC12199 & PE.18.025.010). All mice were purchased from Envigo (Horst, The Netherlands) and housed at Leiden University Medical Center in a temperature-controlled room with a 12 hour light-dark cycle and ad libitum access to food and tap water.

8-10 weeks-old C57BL6/J male mice were injected in either left or right hind hock with 30 µl of an emulsion containing 25 µg of fhFABP1 or PBS with or without 200 ng of LPS. The draining popliteal lymph nodes (LNs) were collected after 6 days in 500 μl HBSS medium for subsequent analysis. LNs were mashed up with the back of a syringe and incubated with digestion buffer (1 μg/ml of Collagenase D and 2000 U/ml of RNAse) for 20 min at 37°C. Next, samples were filtered through a 100 μm cell strainer and washed 3 times with MACS buffer (PBS, 0.5% BSA, 2 mM EDTA). Cells were ultimately centrifuged at 1500 rpm for 5 min at 4°C, counted and plated for further experiments.

8-10 weeks-old C57BL/6JOlaHsd male mice were fed a high-fat diet (HFD, 45% energy derived from fat, D12451) for 12 weeks. At the end of this run-in period, mice were randomized in two groups based on body weight, fat mass, and fasting plasma glucose levels, and next received intraperitoneal (i.p.) injection of either fhFABP1 (25 μg) or vehicle control (sterile-filtered PBS) every 3 days for 4 weeks. Body weight was monitored throughout the experiment and body composition was measured at week 4 of treatment using an EchoMRI (Echo Medical Systems, Houston, TX, USA). Whole-body insulin tolerance (ipITT) and glucose tolerance (ipGTT) tests were performed at week 3-4 of treatment in 4 hours- or 6 hours-fasted mice, respectively, as described previously (34). At the end of the experiment, mice were sacrificed through an overdose of ketamine/xylazine and both visceral white adipose tissue (epidydimal; eWAT) and liver were weighed and collected for further processing and analyses.

1.5 x 10⁶ popliteal LN cells/ml from individual animals were restimulated with 50 ng/ml PMA plus 2 µg/ml ionomycin and 10 µg/ml brefeldin A for 4 hours. Then cells were first stained with Fixable Aqua Dead Cell Stain Kit (Invitrogen/Thermo) and fixed with 1.9% PFA, and next permeabilized with permeabilization buffer (eBioscience). To analyze CD4⁺ T cells, LN cells were stained for CD11b, CD11c, GR-1, B220, NK1.1 as a lineage cocktail. CD4⁺ T cells were identified as CD3, CD4 and CD44 positive and analyzed for IL-4, IFNγ and IL-10 intracellular expression with specific antibodies ( Table S1 ). Alternatively, 1 x 10⁶ popliteal LN cells/ml were stimulated with 10 µg/ml of PHA for 4 days and cytokines levels were measured by FACS using Cytometric Bead Assay (CBA; BD bioscience).

eWAT and liver were collected at sacrifice after a 1 min transcardial perfusion with PBS through the left ventricle and processed as described previously, with a modified digestion mix for eWAT containing 1 mg/ml collagenase type I from Clostridium histolyticum, 2% (w/v) dialyzed bovine serum albumin (BSA, fraction V) and 6 mM D-Glucose in HEPES-buffered Krebs (34, 35). After isolation, leukocytes were counted using a hemocytometer, incubated with the live/dead marker Zombie-UV, fixed using either 1.85% formaldehyde or the FOXP3 Transcription Factor Staining Buffer kit (Invitrogen) and stained as described below. Cells were measured within 4 days post fixation. For analysis of macrophages, cells were permeabilized and incubated with an antibody against YM1 conjugated to biotin and rabbit-anti-RELMα, washed, and stained with streptavidin-PerCP-Cy5.5, anti-rabbit-Alexa Fluor 647 and antibodies directed against CD45, CD64, F4/80, Ly6C and Siglec-F. For analysis of CD4⁺ T cells, cells were stained with antibodies against CD3, CD4, CD8, CD44, CD45, and a lineage cocktail containing B220, CD11b, CD11c, GR-1 and NK1.1 targeting antibodies. For intracellular cytokine detection, isolated cells were restimulated with 100 ng/ml PMA and 1 μg/ml ionomycin in the presence of 10 μg/ml Brefeldin A for 4 hours at 37°C prior to live/dead staining and fixation. CD4⁺ T helper cell subsets were identified using the antibodies listed above and the addition of anti-IL-5 (for Th2) and anti-IFNγ (for Th1) after permeabilization (Invitrogen). Gates were set according to Fluorescence Minus One (FMO) controls. Representative gating strategies are shown in Figure S1 and antibody information is listed in Table S1 .

All data are presented as mean ± standard error of the mean (SEM). Statistical analysis was performed using GraphPad Prism 8.0 (GraphPad Software, La Jolla, CA, USA) with unpaired t-test or one-way analysis of variance (ANOVA) followed by Fisher’s post-hoc test. Differences between groups were considered statistically significant at P < 0.05. Outliers were identified according to the two-standard deviation method.

In order to determine the immunomodulatory effects of F. hepatica-derived native molecules, we first tested the impact of both somatic antigens (fhSA) and excretory-secretory products (fhESPs) using an in vitro model of human monocyte-derived dendritic cells (moDCs, Figure 1A ). We showed that stimulation with both fhSA and fhESPs potently induced IL-10 production in LPS-activated moDCs while neither IL-12p70 and IL-6 production nor the cell-surface expression of the main co-stimulatory markers were affected ( Figures 1B, C, E, F ). In addition, we found that fhSA, and to a lesser extent fhESPs, conditioned moDCs to promote Th2 skewing and concomitantly reduce Th1 polarization, ultimately leading to an increased Th2-on-Th1 ratio ( Figures 1D, G ). We next aimed to identify the specific molecule(s) present in fhESPs that drive this effect. We focused on cathepsin and Fatty Acid Binding Protein (FABP) families as they are the most abundant molecules present in adult fhESPs (27). Using a yeast expression system, we generated recombinant molecules for three of the fhESP molecules, namely fhCL5, fhFABP1 and fhFABP5, and tested their immunomodulatory properties in moDCs ( Figure 2A ). Although fhCL5 and fhFABP5 had only a marginal impact on moDC cytokine production and T cell skewing, fhFABP1 reduced production of inflammatory cytokines and potently induced stimulation of IL-10 secretion by moDCs ( Figure 2B ). Furthermore, fhFABP1 instructed moDCs to promote Th2 over Th1 polarization, leading to an increase in the Th2-on-Th1 ratio ( Figure 2C ).

We next explored in more depth the impact of fhFABP1 on human moDCs, first showing that the molecule can bind ( Figure 3A ) and then be rapidly internalized by the cell ( Figures 3B and S2 ). LPS-stimulated moDCs were next treated with increasing concentrations of fhFABP1 followed by determination of cytokine secretion, following either direct stimulation or secondary restimulation with CD40L, and of T cell skewing capacity ( Figure 3C ). We confirmed that fhFABP1 neither induced apoptosis/cell death ( Figure S3 ), nor affected the expression of canonical DC co-stimulatory markers ( Figure 3D ), but significantly reduced the production of pro-inflammatory cytokines IL-12p70 and IL-6, and increased IL-10 secretion by moDCs ( Figure 3E ). When co-cultured with a CD40L-expressing cell line to mimic the interaction with CD4⁺ T cells, we also found a potent and dose-dependent inhibition of the pro-inflammatory cytokines while IL-10 secretion was not significantly changed ( Figure 3F ). In addition, naïve CD4⁺ T cells co-cultured with fhFABP1-conditioned moDCs increased IL-4 and decreased IFNγ expression relative to controls, indicating an upward shift in the Th2-on-Th1 ratio ( Figure 3G ). Furthermore, a significant increase in T cell IL-10 secretion was observed after re-stimulation with anti-CD3 and anti-CD28, suggesting that fhFABP1 may also enhance DC-mediated priming of regulatory T cells (Tregs) ( Figure 3H ). However, we did not find changes in the T cell expression of the canonical Treg transcription factor FOXP3, while a significant increase in the Th2 transcription factor GATA3 was observed ( Figure 3I ). Altogether, our data suggest that fhFABP1 may induce tolerogenic DCs (TolDCs), which are known to differ from immunogenic DCs through their expression of costimulatory molecules, pro-inflammatory cytokine secretion, and ability to prime Tregs (36). We therefore compared the effects of fhFABP1 to those of Vitamin D3 (VitD3) - a well-known inducer of TolDCs (37–39) - on DC expression of several co-stimulatory and tolerogenic markers. As expected, VitD3-conditioned moDCs exhibited reduced expression of co-stimulatory markers CD80 and CD83, and a concomitant increase in CD163 and ILT3, both tolerogenic markers, while PD-L1 and CD103 remained unchanged ( Figures 4A, B ). By contrast, fhFABP1-conditioned moDCs did not exhibit increased expression of CD163 and ILT3 but displayed significant upregulation of CD103, a surface co-stimulatory molecule shown to be involved in induction of DC tolerance (40) ( Figures 4A, B ). Furthermore, IL-10 secretion was significantly increased by fhFABP1 compared to VitD3-conditioned moDCs, whereas TGFβ production was not affected by either stimulus ( Figures 4C, D ). Of note, similar features were observed when moDCs were stimulated with TNF and IL1β ( Figure S4 ), indicating that the immunomodulatory effects of fhFABP1 are not only mediated by LPS-induced TLR4 signalling.

In order to elucidate the underlying molecular mechanism(s) by which fhFABP1 modulates DC-mediated T cell polarization, we performed an in-depth DC phenotyping through unbiased analyses of cell transcriptome, lipidome and secretome, as well as intrinsic core metabolic pathways. Using RNA sequencing, we found that fhFABP1 induced a significant up- and downregulation of 167 and 68 transcripts, respectively, in LPS-stimulated moDCs ( Figure 5A ). Gene ontology analysis indicated an over-representation of upregulated genes involved in immune response, cell proliferation and leukocyte chemotaxis, and of downregulated genes belonging to various type of inflammatory response pathways ( Figure 5B ). Among the genes that were both highly expressed and the most significantly regulated by fhFABP1, we found some encoding secreted factors known to be involved in DC-mediated Th1 and Treg priming (41–45), notably IL10, CXCL9, CXCL11, CCL19, and THBS1 (Thrombospondin 1, TSP-1) ( Figures 5C , S5A ). Using a cytokine array covering more than 100 secreted proteins, we confirmed that fhFABP1 potently induces TSP-1 secretion by moDCs and a concomitant decrease in CXCL9, CXCL11 and CCL19 production ( Figure 5D ). The effects of fhFABP1 on the two most up- (TSP-1) and downregulated (CXCL11) secreted proteins by moDCs were also confirmed by ELISA ( Figures 5E , S5B ). Since changes in cellular metabolism underpins DC immune functions (36), we also assessed the impact of fhFABP1 on DC intrinsic core metabolic pathways using different approaches. At the transcriptional level, gene set enrichment analysis did not reveal any significant overrepresentation of glucose, fatty acid and amino acid metabolic pathways in fhFABP1-conditioned moDCs ( Figure S6A ). In line with this, mitochondrial mass assessed by Mitotracker Green staining ( Figure S6C ) and both mitochondrial oxygen consumption and glycolytic rates measured by Seahorse flux analyser ( Figure S6B ) were not changed by fhFABP1 in LPS-stimulated moDCs. Furthermore, neither neutral lipids ( Figure S6D ) or intracellular levels of a broad range of lipid species ( Figure S6D ) were affected, indicating that fhFABP1 immunomodulatory effects are unlikely to be explained by modulation of DC metabolism.

We next investigated the role played by the secreted factors identified using our transcriptomic and proteomic approaches in T cell polarization by fhFABP1-conditioned moDCs. For this purpose, we either replenished the chemokines found to be reduced by fhFABP1 (CXCL11, CCL9 and CXCL9) or neutralized DC-secreted TSP-1 using a blocking antibody during DC-T cell co-culture ( Figures 6A, B ). Although the chemokines mixture did not affect the capacity of fhFABP1-conditioned moDCs to modulate the Th2/Th1 balance and T cell IL-10 production ( Figure 6C ), neutralizing TSP-1 abolished the effects of fhFABP1 on DC-mediated T cell polarization ( Figure 6D ). Altogether, we demonstrate that fhFABP1 modulates T cell polarization by promoting TSP-1 secretion in moDCs. Finally, we assessed whether fhFABP1-conditioned moDCs could prime regulatory T cells with suppressive capacity, by measuring their respective ability to suppress proliferation of bystander T cells. We found that in contrast to VitD3, T cells primed by fhFABP1-conditioned moDCs did not inhibit the proliferation of bystander T cells ( Figure 6E ), indicating that despite inducing IL-10-secreting T cells, fhFABP1 did not promote DC-mediated skewing of classical Tregs with suppressive capacity.

To assess whether fhFABP1 can also modulate T cell polarization in vivo, we administered the recombinant molecule, with or without LPS, via hock injection in the mouse, and then analysed T cells from the draining popliteal lymph nodes (pLNs) after 6 days ( Figure 7A ). As expected, LPS induced a significant increase in both total pLN immune cells and CD44⁺CD4⁺ T cells ( Figures 7B, D ), while the percentages of total CD4⁺ T cells in pLN cells remained unchanged regardless of the conditions ( Figure 7C ). Remarkably, fhFABP1 significantly increased the percentage of IL-4-expressing CD4⁺ T cells as well as IL-4 secretion by pLN cells after ex vivo restimulation, while a trend for decreased IFNγ-expressing CD4⁺ T cells and IFNγ secretion by pLN cells was observed ( Figures 7E, F ). Altogether, and in line with our in vitro data, fhFABP1 can also affect the Th2/Th1 balance towards an enhanced Th2 immune response in vivo ( Figures 7E, F ).

Infection with various helminth species or treatment with their immunomodulatory molecules have been reported to restore type 2 immunity in metabolic organs and improve insulin sensitivity and glucose homeostasis in obese mice (6). Therefore, we investigated whether fhFABP1-induced Th2 immune response may alleviate obesity-induced metabolic dysfunctions. For this purpose, HFD-fed mice received biweekly intraperitoneal injections with fhFABP1 for four weeks ( Figure 8A ), a treatment that did not affect body weight gain and body composition in obese mice ( Figures 8B–D ). Immune cell phenotyping in metabolic tissues showed that although no differences in Kupffer cell and CD4⁺ T cells numbers were observed in the liver ( Figure S7 ), fhFABP1 markedly increased both total leukocytes and CD4⁺ T cells in white adipose tissue (WAT) ( Figures 8E, F ). Strikingly, fhFABP1 potently increased the numbers of CD44⁺ CD4⁺ T cells (data not shown), which were characterized by increased frequencies of IL-5⁺ Th2 cells, and by a trend for decreased IFNγ⁺ Th1 cells ( Figures 8G, H ). This was in line with what was observed in draining pLNs after hock injection of fhFABP1. Furthermore, the numbers of eosinophils ( Figure 8I ) and YM1⁺ macrophages were also increased while total macrophages and RELMα⁺ macrophages were unchanged ( Figures 8J, K ). However, this potent WAT type 2 immune response was not associated with any beneficial effects on HFD-induced metabolic dysfunctions, since neither fasting glycemia, nor whole-body insulin sensitivity and glucose tolerance were improved by fhFABP1 treatment ( Figures 8L–N ).

Altogether, we showed that although fhFABP1 modulates T cell polarization, notably by promoting DC TSP-1 secretion in vitro, it does not affect metabolic homeostasis in a mouse model of obesity and type 2 diabetes.