Human sample collection was in accordance with Good Clinical Practice guidelines. The study was approved by the Ethics Committee of the University Hospital of Fuenlabrada (Madrid, Spain). All subjects provided their written informed consent.

Animal experiments were performed in accordance with the IBMC.INEB Animal Ethics Committees and the Portuguese National Authorities for Animal Health guidelines (directive 2010/63/EU). Begoña Pérez-Cabezas and Anabela Cordeiro-da-Silva are accredited for animal research (Portuguese Veterinary Direction, Ministerial Directive 113/2013).

Diagnosed VL patients were treated with liposomal Amphotericin B (21 mg/kg). Cure was 100% in 3 months after diagnosis. Subjects without previous VL symptomology were included as negative controls. Plasma was recovered from heparinized blood and stored at −20°C for posterior IL-27 determination using MILLIPLEX MAP (Millipore, Germany) and analyzed on a Bio-Plex-200 Luminex (Bio-Rad, CA, USA) (minimum detectable concentration 0.063 ng/mL).

A clone of virulent L. infantum (MHOM/MA/67/ITMAP-263) was maintained by weekly subpassages at 26°C in RPMI 1640 medium supplemented with 10% heat-inactivated fetal bovine serum (FBS), 2 mM l-glutamine, 100 U/mL penicillin, 100 microgram/mL streptomycin, and 20 mM HEPES buffer (all from BioWhittaker, Lonza, Switzerland). Promastigotes from 4 to 10 passages were used in these experiments.

Six- to eight-week-old male BALB/c and C57BL/6 mice (Charles River Laboratories, France) were maintained under specific pathogen-free conditions at the IBMC facilities. Animals were infected i.p. with 1 × 10⁸ stationary promastigotes from 5 culture days.

Blood from mice was collected through intracardiac puncture under isoflurane anesthesia. Serum was collected and stored at −80°C for posterior analysis.

Spleen and Liver were aseptically collected from 24 h infected mice. Splenocytes were obtained and labeled to distinguish DCs (CD11b⁺/CD11c^high) and macrophages (CD11b⁺/Ly6G⁻/Ly6C⁺/F4-80⁺), and sorted using a FACSAria and the FACSDiva software (BD Biosciences, NJ, USA) for posterior RNA analysis. Sorted cells purity was >95%.

Enriched Kupffer cells were obtained after hepatic collagenase digestion, followed by a Percoll density gradient centrifugation (Sigma-Aldrich, MO, USA) and a brief adhesion step, as described elsewhere (14). The percentage of Kupffer cells was determined by flow cytometry (F4/80⁺/CD11b^variable). Cells were resuspended in lysis buffer and frozen for posterior RNA analysis.

Total RNA was extracted (RNeasy Mini Kit, Qiagen, CA, USA), quantified (NanoDrop1000, Thermo Scientific, MA, USA), and reversely transcribed (NZY-First-Strand cDNA synthesis Kit, NZYTech, Portugal). Quantitative RT-PCRs were run on an iQ5 iCycler thermal cycler (Bio-Rad) (15). Results were analyzed (built-in iQ5 v2, Bio-Rad) and normalized using the reference gene GAPDH. The sequences of the primers used were IL-27p28 Forward TCGATTGCCAGGAGTGAACC, Reverse CGAAGTGGTAGCGAGGAAG; EBI3 Forward AGCAGCAGCCTCCTAGCCT, Reverse ACGCCTTCCGGAGGGTC; and GAPDH Forward CTGGTCCTGAGTGTAGCCCAA, Reverse CATGGCCTTCCGTGTTCCTA.

For bone marrow-derived DCs (BMDCs) differentiation, 6 × 10⁶ bone marrow cells were seeded in 25 mL of complete RPMI supplemented with 50 μM 2-mercaptoethanol (Sigma-Aldrich) and 10% of GM-CSF-containing supernatant from J558 cell conditioned medium (DC medium) (16). Cells were cultured at 37°C and 5% CO₂ for 3 days, after which the same amount of DC medium was added to each flask. At day 6, half of the culture supernatant was replaced with fresh DC medium. At day 8, cells were thoroughly resuspended, recovered, counted, and, finally, plated in 96 round-bottom culture plates at 1 × 10⁵ cells/well in 200 μL of DC medium. Bone marrow-derived macrophages (BMMØ) were obtained from non-adherent bone marrow cells collected after an initial overnight incubation (37°C and 5% CO₂) in complete Dulbecco’s modified Eagle’s medium (DMEM) (Lonza). Non-adherent cells were counted and distributed in 96 flat-bottom well culture plates at 1 × 10⁵ cells/well in 100 μL of complete DMEM supplemented with 5% of l-929 cell conditioned medium (LCCM). After 3 days of culture, 100 μL of DMEM + 5% LCCM were added per well. Half of the media volume was renewed at day 6 of culture. BMMØ acquired a definitive differentiation status at day 8.

Stationary promastigotes from 5 culture days (cultured as described in Section “Parasites and Mice”) were added to BMDC or BMMØ at a 10:1 ratio. Non-internalized parasites were washed after 4 h incubation (37°C and 5% CO₂). Non-infected cells were stimulated with 1 μg/mL LPS (Sigma-Aldrich) as positive control, or received complete medium as negative control. Supernatants were recovered 24 h post-infection for cytokine quantification.

Twenty-four hours after infection, BALB/c mice were i.p. treated with 20 μg of goat anti-mouse IL-27p28 neutralizing antibody (a-IL-27) and C57BL/6 with 1 μg of mouse recombinant IL-27 (rIL-27) (both from R&D Systems), as previously reported (17, 18). As controls, BALB/c and C57BL/6 received 20 μg of goat IgG (isotype control) and PBS (mock), respectively.

Spleen and liver were aseptically collected, weighted, and disrupted. Splenic and hepatic parasite burdens were assessed by the limit dilution method. The parasite titer was considered as the last dilution with >1 motile parasite. The number of parasites per gram of organ was calculated, as discussed previously (19). Peritoneal cells were recovered and 2 × 10⁵ cells from each exudate were subjected to cytospin in 200 μL of PBS during 5 min at 1000 rpm using a Shandon Cytospin II (GMI, MN, USA). Cell preparations were fixed with 2% paraformaldehyde (PFA) for 20 min. Afterward, an adapted staining protocol was performed (20) by 2 min immersion in Kaplow modified solution, followed by 45 s immersions in Hemacolor reagent 1 and reagent 2 (Merk Millipore, Germany). Finally, slides were washed with distilled water, air dried, and observed by optical microscopy (100× magnification). For determination of the percentage of infected cells, 200 consecutive cells were differentially counted (infected versus non-infected) in 3 different areas of the same preparation and the mean of the 3 areas was calculated. The number of parasites per infected cells was assessed by counting 100 different infected cells from which the mean was calculated.

The anti-mouse monoclonal antibodies used to perform this study were all purchased to BioLegend (CA, USA) except if otherwise stated: FITC-labeled anti-IgM (R6-60.2, BD Biosciences, NJ, USA), anti-MHC-II(I-Ad) (AMS-32.1, BD), anti-MHC-II(I-A/I-E) (M5114.15.2), anti-IFN-γ (XMG1.2), and anti-IL-17A (TC11-18H10.1); PE-labeled anti-CD8 (53-6.7, BD), anti-CD11b (M1/70), anti-Siglec-F (E50-2440, BD), anti-F4/80 (BM8), anti-IL-4 (11B11), and anti-IL-6 (MP5-20F3); PerCP-labeled anti-CD11b (M1/70); PerCP-Cy5.5-labeled anti-Ly6C (HK1.4), anti-F4-80 (BM8), and anti-TNFα (MP6-XT22); PE-Cy7-labeled anti-CD3 (HA2) and anti-CD11b (M1/70); APC-Cy7-labeled anti-CD11c (N418); APC-labeled anti-CD19 (6D5), anti-IL-5 (TRFK5), and anti-IL-10 (JES5-16E3); BV510-labeled anti-CD4 (RM4-5); and Pacific Blue™-labeled anti-Ly6G (1A8).

To analyze lymphoid and myeloid cell populations, two panels of antibodies were designed. The lymphoid panel was composed of anti-CD8, -CD3, -CD4, and -CD19. The Myeloid panel comprised anti-CD11b, -CD11c, -Siglec-F, -Ly6C, -Ly6G, and -MHC-II. Surface staining of peritoneal and splenic cells was performed in PBS + 0.5% BSA (20min, 4°C) followed by 15 min fixation using 1% PFA. For intracellular staining, splenocytes were cultured for 2 h with PMA/Ionomycin (50/500 ng/mL) and then for 2 h with Brefeldin A (10 μg/mL). Cells were surface stained and then intracellularly after fixation and permeabilization with 1% saponin (all from Sigma) (21). Samples were acquired in a FACSCanto (BD) and analyzed using the FlowJo software v10 (TreeStar, OR, USA).

An initial gate plotting FSC-A versus SSC-A was performed. Afterward, singlets were selected by plotting FSC-A versus FSC-H and the remaining cell populations were resolved. T lymphoid cell populations were defined as CD3⁺/CD4⁺ and CD3⁺/CD8⁺, while B cells were defined as CD19⁺. Cytokine production by T cells was assessed within CD3⁺/CD4⁺ and CD3⁺/CD8⁺cells. Myeloid cell populations were gated as eosinophils (Siglec-F⁺/SSC-H^int/high), neutrophils (CD11b^high/Ly6G^high/Siglec-F⁻), DCs (CD11c⁺/MHC-II^int/high), and macrophages (CD11b⁺/CD11c⁻/Ly6G⁻/Siglec-F⁻).

Cytokines were quantified, according to the manufacturer’s instructions, using the commercial kits: IL-27 ELISA Ready-SET-Go!^® (eBiosciences, CA, USA) (detection limit 16 pg/mL); IL-27p28/IL-30 and IL-10 DuoSet ELISA (R&D Systems, MN, USA) (detection limits 15.6 and 31.2 pg/mL, respectively); IL-12p70 and IFN-γ ELISA MAX Deluxe (BioLegend, CA, USA) (detection limit 4 pg/mL for both cytokines).

Results are expressed as mean ± SEM. Statistical differences were analyzed using GraphPad Prism v6.01 (CA, USA). Comparisons between human samples were performed using Mann–Whitney test for unpaired samples and Wilcoxon test for paired data. Mice experimental groups were compared using the unpaired t-test unless otherwise stated.

To address if the increase of systemic IL-27 is a common fact of VL among different endemic areas (7, 8), we determined this cytokine in the plasma of active L. infantum infected individuals from a current outbreak in Spain. Patients with active disease presented higher IL-27 levels than both cured (p ≤ 0.001) and negative controls (p ≤ 0.001) (Figure 1A). Some patients were re-sampled after cure, which confirmed that IL-27 decreases after successful treatment (p ≤ 0.01) (Figure 1A).

The role of IL-27 during Leishmania infection was addressed using mouse models by several authors (10–13, 22). However, whether IL-27 is also increased in the serum of Leishmania-infected mice has never been shown. Therefore, we addressed if BALB/c and C57BL/6, two susceptible VL mouse species, displayed a similar systemic increase of IL-27 as observed in L. infantum-infected humans. While IL-27 remained unchanged in the serum of infected C57BL/6 mice, infected BALB/c showed an early increase that was significant 4 days after infection (p ≤ 0.05) and always higher than the IL-27 levels of C57BL/6 mice (p ≤ 0.05 comparing the day 2 and p ≤ 0.01 the day 4 after infection) (Figure 1B).

As the main sources of IL-27 are myeloid cell populations, and macrophages and DCs play critical roles during Leishmania infection, we decided to discriminate their contribution to IL-27 response after L. infantum infection. For that, the expression of IL-27 subunits in splenic DCs and macrophages sorted from infected BALB/c and C57BL/6 was evaluated. While the expression of EBI3 remained always comparable to basal levels (Figure S1 in Supplementary Material), IL-27p28 expression was significantly upregulated but only in DCs from BALB/c 24 h after infection (p ≤ 0.05) (Figure 2A). The contribution of Kupffer cells, the resident liver macrophages, for the IL-27 response to L. infantum was also addressed. The expression of IL-27 subunits was analyzed in Kupffer cells from non-infected and infected BALB/c mice. However, no increase in the RNA levels of IL-27p28 and EBI3 was detected in these cells 24 h after infection (Figure S2 in Supplementary Material), suggesting that DCs, and not macrophages, are the main cell source responsible for the increase of IL-27 in our model.

To confirm our data at protein level, we determined IL-27 and IL-27p28 in supernatants from 24 h L. infantum-infected BMMØ and BMDCs. Although after infection the production of IL-27p28 was greater in BMMØ from C57BL/6 than from BALB/c (p ≤ 0.05), IL-27 concentration was similar in both mouse strains. No difference was found after LPS activation (Figure 2B). In line with ex vivo results, both IL-27p28 and IL-27 secretion by BMDCs were significantly higher in cells from BALB/c when compared to C57BL/6 (p ≤ 0.001 and p ≤ 0.01, respectively) (Figure 2B). Interestingly, the ability of BMDCs to produce both cytokines after LPS stimulation was higher in C57BL/6 than in BALB/c, indicating that these cells are really capable of producing IL-27, but not in response to L. infantum infection. These results suggest that the parasite can actively modulate the secretion of IL-27 and IL-27p28 in BMDCs obtained from BALB/c, but not from C57BL/6 mice.

To determine whether IL-27 contribute to the infection outcome, we neutralized IL-27 in BALB/c and administered rIL-27 to C57BL/6 mice 24 h after infection, coinciding with the IL-27p28 RNA peak in the spleen and before the IL-27 increase observed in the serum, 2 days after the infection, in BALB/c mice. Seventy-two hours later, these treatments significantly decreased in BALB/c and increased in C57BL/6 splenic and hepatic parasite burdens (Figure 3A). We also observed that IL-27 significantly affected the percentage of infected cells and the number of parasites per infected cells in the peritoneal cavity (Figure 3B). These results revealed a relation between the levels of IL-27 and the establishment of the infection.

To elucidate how IL-27 interferes at the early stages of L. infantum infection, we performed a kinetic study of the peritoneal and splenic compartments after IL-27 modulation. Interestingly, IL-27 neutralization in infected BALB/c mice transiently decreased IL-10 in the peritoneal cavity (p ≤ 0.01), and the administration of rIL-27 increased the presence of this cytokine in infected C57BL/6 mice 24 h (p ≤ 0.01) and 72 h (p ≤ 0.05) after treatment, always comparing with control infected animals (Figure 4). In addition, rIL-27 in C57BL/6 promoted a reduction of IFN-γ (p ≤ 0.05) and IL-12p70 (p ≤ 0.01) 24 h after treatment in comparison with infected non-treated animals (Figure 4). However, this shift in the cytokine profile determined by IL-27 almost did not alter cell recruitment to the site of infection, as when comparing treated and non-treated animals only the presence of B cells in the peritoneal cavity significantly increased 72 h after IL-27 blockage in BALB/c mice (Figure 5). In the spleen, the number of CD4⁺ T cells increased in BALB/c mice 24 h after IL-27 blockage (p ≤ 0.05), and the supply of the cytokine decreased the presence of the same cells in C57BL/6 mice after 72 h (p ≤ 0.05) (Figure 5). The administration of rIL-27 also prevented the infiltration of neutrophils in the spleen of C57BL/6 mice (p ≤ 0.01) (Figure 6). Analysis of the cytokine production revealed that IL-27 neutralization increased the numbers of IFN-γ producing CD4⁺ and CD8⁺ T cells 24 h post-treatment (Figure 7) (p ≤ 0.01). These differences were not detected at 72 h, likely indicating a reversible effect (Figure 7). The potential secretion of other cytokines was also analyzed but no differences between α-IL-27 and IgG receiving mice were observed (Figure S3 in Supplementary Material). In contrast, rIL-27 did not alter the cytokine response of C57BL/6 splenic T cells (Figure 7; Figure S3 in Supplementary Material).