Female C57BL/6 (wild type; WT) and C57BL/6-TLR2^−/− (TLR2^−/−) mice weighing 18–22 g were housed in the animal facility of the Department of Biochemistry and Immunology, School of Medicine of Ribeirão Preto, University of São Paulo (Brazil), in temperature-controlled rooms (22–25°C) and received water and food ad libitum. All experiments were conducted in accordance with the National Institutes of Health (NIH) guidelines for the welfare of experimental animals and with the approval of the Ethics Committee of the School of Medicine of Ribeirão Preto.

L. infantum (isolate HU-UFS14) was cultured in Schneider medium with 20% heat-inactivated fetal bovine serum, 5% penicillin and streptomycin (both from Sigma-Aldrich, Saint Louis, MO, USA), and 2% male human urine. Parasite virulence was maintained by serial passages in BALB/c mice. Mice were injected in their retro-orbital plexus with 10⁷ stationary-phase L. infantum promastigotes in 100 μL of PBS. Hepatic and splenic parasite burdens were determined using a quantitative limiting dilution assay.

Generation of bone marrow-derived cells (BMDCs) was performed as previously described (Carregaro et al., 2008). For the DC activation assay, BMDCs (1 × 10⁶ cells/mL) cultured in RPMI-1640 supplemented with 10% FBS were infected with L. infantum promastigotes at a 1:5 ratio (cells/parasites) for 24 h. The supernatants were collected to measure IL-12p40 (BD Biosciences, catalog number 555165) and IL-10 (R&D Systems, catalog number DY417-05) production by an ELISA kit, according to the manufacturer's instructions. BMDCs were harvested from some cultures, and their surface expression was characterized by flow cytometry using antibodies against CD11c, TLR2, MHCII, CD40, and CD86 conjugated to PECy7, PE, FITC, PerCP, APC, respectively, as well as control isotypes.

Single-cell suspensions of spleen tissue samples from TLR2^−/− or WT mice at the 4th wpi were aseptically prepared, diluted to a concentration of 2 × 10⁶ cells/mL, and dispensed into 48-well plates in a total volume of 500 μl of complete RPMI-1640 medium (1 × 10⁶ cells/well; Gibco) with or without L. infantum crude antigen (50 μg/mL). The cell culture supernatants were harvested after 72 h of culture at 37°C in 5% CO₂ and the IL-17, IFN-γ, and IL-10 levels in the supernatants were determined using commercial ELISA kits. For detection of IFN-γ and IL-17 in the liver, tissue samples were harvested by tissue trimmer, weighed, and titered in 1 mL of PBS Complete (Roche Diagnostics, Mannheim, Germany) containing protease inhibitor cocktail. The levels of cytokines were determined using commercial ELISA kits from R&D Systems for IL-10 and IL-17(catalog number DY417 and DY421, respectively) and BD Biosciences for IFN-γ (catalog number DY485).

For leukocyte identification, spleeny cells isolated from WT and TLR2^−/− were stained with fluorescent labeled monoclonal antibody that recognizes a target feature on or in the cell. The inflammatory cells were gated based on their characteristic size (FSC) and granularity (SSC), and the T lymphocytes (CD4⁺CD3⁺), dendritic cell activation markers (CD11c^highCD40⁺, CD11c^highCD86⁺, and CD11c^highMHC-II⁺) and neutrophils (Ly6G^highMHCII⁻) were identified individually. For intracellular staining, the cells were cultured with PMA (50 ng/mL) and ionomycin for 4 h in order to obtain the maximum cytokine production, permeabilized with a Cytofix/Cytoperm Kit (BD Biosciences) according to the manufacturer's guidelines and stained with anti-IFN-γ or anti-IL-17 conjugated to APC-Cy7 and Alexa Fluor 700 and with anti-CD3 and anti-CD4 for surface staining with FITC and PerCP, respectively. Rat IgG2b and IgG2a were used as the isotype controls. All the antibodies were supplied from BD Biosciences and eBiosciences (San Diego, CA, USA). Cell acquisition was performed using a FACSort flow cytometer. The data were plotted and analyzed using FlowJo software (Tree Star, Ashland, OR, USA). The total leucocyte counts were determined by measuring the relative expression of the leucocyte subpopulations stained with specific antibody in 300,000 acquired events proportional to the leukocyte number obtained in a Neubauer chamber.

The mice were euthanized 0 and 4 weeks after infection, and their livers were removed. The tissues were fixed in formalin, dehydrated in graded ethanol, and embedded in paraffin. Serial sections (5 μm) were cut and mounted on glass slides precoated with 0.1% poly-l-lysine (Sigma-Aldrich). Histological assessment was performed after routine hematoxylin-eosin staining. The area of the liver lesion was determined using Leica Qwin software (Mannheim, Germany).

For immunohistochemical reactions, the paraffin was removed from the tissues, and antigenic recovery was performed by heating the sections in citrate buffer (pH 6.0) for 30 min at 37°C. Endogenous peroxidase was blocked using 3% H₂O₂, cells were permeabilized with Triton 0.5%, and non-specific reactions were blocked with 1% bovine serum albumin (BSA). The sections were incubated overnight with a monoclonal rat anti-7/4 antibody (Abcam Plc, Cambridge, UK) or control isotype antibodies (Abcam, Cambridge, MA, USA), followed by incubation with a biotinylated secondary antibody and avidin-biotin complex (Vector Laboratories, Ontario, Canada). The reaction was detected with diaminobenzidine, and the sections were counterstained with Mayer's hematoxylin. For intracellular staining for iNOS (Santa Cruz Biotechnologies, Dallas, TX, USA), liver sections were permeabilized with 0.01% saponin. Afterward, the sections were incubated with an avidin-biotin-peroxidase complex (Vector Laboratories, Ontario, Canada), and the color was developed using 3,3′-diaminobenzidine (Vector Laboratories). The slides were counterstained with Mayer's hematoxylin.

Total RNA was isolated from neutrophil cultures or from the spleens and livers of WT and TLR2^−/− mice at the 4th wpi and uninfected using an SV Total RNA Isolation System Kit (Promega, Madison, WI, USA). Gene expression was normalized to hypoxanthine-guanine phosphoribosyltransferase (HPRT) expression for spleen and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) expression for liver and neutrophil culture. The primer sequences used were as follows: HPRT forward primer, 5′-TGGAAAAGCCAAATACAAAGC-3′, and reverse primer, 5′-CAACATCAACAGGACTCCTCG-3′; GAPDH forward primer, 5′-TGCAGTGGCAAAGTGGAGAT-3′, and reverse primer, 5′-CGTGAGTGGAGTCATACTGGAA-3′; TLR2 forward primer, 5′-AAGTCTCCGGAATTATCAGTCC-3′, and reverse primer, 5′-TGATGGATGTCGCGGAT-3′; and iNOS forward primer, 5′-CGAAACGCTTCACTTCCAA-3′, and reverse primer, 3′-TGAGCCTATATTGCTGTGGCT-5′.

For bone marrow neutrophil purification, femurs and tibias were removed and flushed. The bone marrow cells were suspended in Hanks' balanced salt solution (Sigma-Aldrich, St. Louis, MO, USA) and laid on top of a two-layer Percol (Sigma-Aldrich, St. Louis, MO, USA) gradient (72 and 65% in Hanks' balanced salt solution). Mature neutrophils were recovered at the interface of the gradient fraction. For NO, TNF, and IL-10 quantification, neutrophils (5 × 10⁵) were cultured in RPMI-1640 supplemented with 10% FBS in the presence of L. infantum promastigote forms at a 1:5 ratio (cells:parasites) for 24 h at 37°C in an atmosphere of 5% CO₂. The supernatants were harvested to measure NO by the Griess method (Martins et al., 2001) and TNF and IL-10 by an ELISA kit according to the manufacturer's instructions (BD Biosciences, catalog number DY410-05 and DY417, respectively). For neutrophil activation, 1 × 10⁶ neutrophils/well were distributed in 48-well plates in 300 μL of complete RPMI medium in the presence of medium only or L. infantum promastigote forms at a 1:5 ratio for 4 h. For the L. infantum uptake assay, parasites were previously stained with 1.25 mM carboxyfluorescein succinimidyl ester (CFSE; Sigma-Aldrich, St. Louis, MO, USA) in PBS. The neutrophils were primed with recombinant murine IFN-γ (100 ng/mL) for 1 h and cultured with CFSE-stained L. infantum promastigotes for 4 h. Then, the cells were fixed and stained with anti-α-Ly6G and anti-α-CD11b antibodies that were conjugated with APC and PEcy7, respectively (BD Bioscience and eBioscience, San Diego, CA, USA). Cell acquisition was performed using a FACSort flow cytometer. The data were plotted and analyzed using FlowJo software (Tree Star, Ashland, OR, USA).

Data are expressed as the mean ± SEM and are representative of 2–4 independent experiments. The results from individual experiments were not combined because they were analyzed individually. The means from the different groups were compared by ANOVA followed by Tukey's honest significant difference (HSD) test. Comparisons between two groups were determined using Student's t-test. Analyses were performed using Prism 5.0 software (GraphPad). Statistical significance was set at P < 0.05.

It has been suggested that TLR2 interacts with Leishmania species, including L. infantum, and triggers the host immune response against the parasite (de Veer et al., 2003; Assis et al., 2012). First, we investigated whether TLR2 expression was altered in BMDCs infected with promastigote forms of L. infantum. During infection, both the percentage and the mean fluorescence intensity (MFI; Figure 1B) of TLR2 expression on CD11c⁺ cells were increased. Furthermore, tlr2 mRNA expression in the spleen and liver from infected WT mice at the 4th week was significantly increased, p = 0.002 and p = 0.001, respectively (Figure 1C), demonstrating that L. infantum infection modulates TLR2 expression both in vitro and in vivo.

To assess the role of TLR2 in the control of parasite growth, TLR2^−/− mice and control littermates were infected with 10⁷ promastigote forms of L. infantum, and parasite loads in their spleens and livers were quantified after different time-points of infection. TLR2^−/− mice harbored more parasites in both target organs during weeks 4 and 6 after infection than WT animals (Figure 1D).

The control of infection during experimental VL is related to leukocyte infiltration of the liver, generating granulomas (Stanley and Engwerda, 2006). Thus, in order to determine the factors involved in the susceptibility of infected TLR2^−/− mice, the amount of inflammatory cell infiltration in the liver of WT and TLR2^−/− mice at the 6th wpi and naïve mice, as a control, was determined. Histologic examination of livers demonstrated that WT mice exhibited large areas of inflammatory tissue at the 4th wpi. In the absence of TLR2, the inflammatory infiltrate was dispersed in the hepatic parenchyma, which presented reduced areas of leucocyte infiltration (Figure 1E). Confirming the histopathology analyses, the quantification of inflamed areas in hepatic tissue demonstrates smaller areas of inflammatory infiltrate in the TLR2^−/− mice. While the average size of inflammatory areas in the WT group was 8,000 μm², the average size of the inflammatory areas in TLR2^−/− mice was 5,000 μm², p < 0.05 (Figure 1F), suggesting that TLR2 contributes to inflammatory infiltration during VL.

We have previously shown that TLR9 pathway signaling, in addition to modulating innate immunity, drives the adaptive immune response controlling parasitic spread (Sacramento et al., 2015). To determine whether this activation mechanism induced by TLR9 is shared with other TLR ligands, we determined both Th1 and Th17 responses in target organs of infection. Spleen cells from naïve and infected WT and TLR2^−/− mice were restimulated in vitro with polyclonal PMA plus ionomycin, and intracellular cytokine production was analyzed. L. infantum infection promoted both Th1 and Th17 patterns of immune response in WT and TLR2^−/− mice compared with respective littermate controls. However, significant reductions in the frequencies and absolute numbers of both IFN-γ (Figures 2A,C,D)- and IL-17 (Figures 2B,E,F)-producing CD4-T cells were observed in infected TLR2^−/− mice compared with those in infected WT mice. In addition, liver samples from infected TLR2^−/− mice contained significantly lower levels of IFN-γ (Figure 2G) and IL-17 (Figure 2H) compared with those of WT mice. IL-10 production was measured in cultured spleen cells in response to stimulation with parasite antigen and, consistent with a reduction in pro-inflammatory cytokines, was significantly increased in the supernatant of cultured cells lacking TLR2 compared with the littermate control (Figure 2I), suggesting that TLR2 signaling modulates cytokines during VL.

DC maturation and inflammatory cytokine secretion are important for modulating CD4⁺ T cell polarization (von Stebut et al., 2000). Therefore, we evaluated the activation profile of DCs from the spleens of WT and TLR2^−/− mice at the 6th wpi or of naïve mice. Our data demonstrated impairment of MHCII (Figures 3A,B), CD40 (Figures 3A,C), and CD86 (Figures 3A,D) expression on CD11c⁺ cells recovered from infected TLR2^−/− mice compared to infected WT mice. As a consequence of the failure of DC maturation, in vitro-infected BMDCs derived from TLR2^−/− mice infected with L. infantum presented reduced amounts of IL-12p40 (Figure 3E) and elevated secretion of IL-10 (Figure 3F) into the supernatant compared to WT BMDCs. Taken together, these results suggest that the absence of TLR2 committed the DC activation profile, affecting both Th1 and Th17 immune response development possibly due to a detrimental effect on IL-10 production.

Neutrophils are recruited to Leishmania inoculation foci (Thalhofer et al., 2011) and participate in the restriction of parasites in target organs of VL (Smelt et al., 2000; Rousseau et al., 2001; McFarlane et al., 2008; Sacramento et al., 2015). To evaluate the contribution of TLR2 in the recruitment of neutrophils during L. infantum infection, we characterized the neutrophils present in spleens and livers from infected WT or TLR2^−/− mice. Based on the characteristic size (FSC) and granularity (SSC), we observed a significant reduction in the percentage of granulocytes gated from infected TLR2^−/− mice compared to infected WT mice (Figure 4A). We also observed relatively minor differences in terms of the percentage of Ly6G⁺MHCII⁻ cells (Figures 4B,C), most notably, a significant reduction in the total number of this population in infected TLR2^−/− mice relative to the littermate controls (Figure 4D).

Regarding neutrophils in the liver, immunohistochemical analysis demonstrated less 7/4-stained cells in the liver sections from infected TLR2^−/− mice compared to infected WT mice (Figure 4E). We quantified the staining and identified a drastic drop, ~60%, in the number of neutrophils in the liver sections from infected TLR2^−/− mice compared to those from infected WT mice (Figure 4F). The reduction in the number of neutrophils in the absence of TLR2 was concomitant with reduced production of CXCL1, a neutrophil chemotactic mediator, by splenic cells restimulated with parasite antigen (Figure 4G). Consistent with the failure of neutrophil migration and the reduced DC activation profile (Figures 3B–D), we found that bone marrow-derived dendritic cells (BMDCs) from TLR2^−/− mice presented a deficiency in the secretion of CXCL1 when infected with L. infantum (Figure 4H). All together, these data indicate the participation of TLR2 signaling in DC-mediated neutrophil recruitment to inflammatory foci of VL.

Interleukin 17A acts synergistically with Interferon-γ to promote protection against L. infantum infection through iNOS expression and NO production (Nascimento et al., 2014). Because the absence of TLR2 affected both Th1 and Th17 development, we therefore investigated iNOS expression in target organs of VL from infected TLR2^−/− mice and littermate controls. The immunohistochemical qualitative analysis demonstrated that L. infantum infection induces iNOS expression in the livers of both WT and TLR2^−/− mice; however, decreased staining areas for iNOS were observed in the liver sections of infected TLR2^−/− mice relative to infected WT mice (Figure 5A). Similar effects were also observed in the spleens of mice that lacked TLR2 that presented reduced expression of mRNA for inos compared to infected WT mice (Figure 5B), indicating the involvement of TLR2 in the induction of iNOS expression during VL.

During Leishmania infection, different cell types such as macrophages, inflammatory monocytes, and neutrophils produce NO-dependent iNOS expression, participating actively in the control of parasites (Liew et al., 1991; Wei et al., 1995; Olekhnovitch and Bousso, 2015). To determine which cellular subtype is the primary source of NO in a TLR2-dependent manner, we evaluated iNOS expression in splenic macrophages (CD11b⁺CD11c⁻Ly6G⁻ cells) and neutrophils (CD11b⁺Ly6G⁺MHCII⁻ cells) from infected WT and TLR2^−/− mice (Figures 6A,D). Interestingly, the absence of TLR2 on macrophages affected neither the percentage (Figure 6B) nor MFI (Figure 6C) of iNOS expression in macrophages. However, we found a reduction in iNOS expression in neutrophils from mice that lacked TLR2 compared with WT neutrophils. The reduction in iNOS expression was ~50% (Figure 6E) in terms of percentage and 20% in terms of MFI (Figure 6F).

Having determined that TLR2 plays an important role in iNOS expression by neutrophils, we evaluated whether, similar to DCs (Figure 1A), L. infantum infection may induce TLR2 expression on neutrophils. Thus, isolated bone marrow-derived neutrophils from WT mice were cultured with promastigote forms of L. infantum for 4 h, and the cells were harvested for tlr2 mRNA expression by quantitative PCR. We observed that the infection induces an ~7.4-fold increase in tlr2 mRNA expression (Figure 6G). Confirming the importance of TLR2 in iNOS expression, we observed that in vitro neutrophils from TLR2^−/− mice infected with L. infantum presented a reduction in mRNA expression for inos compared to WT neutrophils. Even in the presence of IFN-γ, the expression of inos mRNA was not induced (Figure 6H). Consistently, the amounts of NO measured in supernatant cultures from TLR2^−/− neutrophils were significantly reduced by ~70% compared with WT neutrophils (Figure 6I). In addition, infected TLR2^−/− neutrophils presented a significant reduction in the amount of TNF-α compared to WT neutrophils (Figure 6J). Because stimulation of TLRs modulates neutrophil activation by up-regulating integrins (Sabroe et al., 2003), we evaluated the expression of CD11b in infected WT and TLR2^−/− neutrophils. Flow cytometric analysis of Ly6G⁺MHCII⁻ cells demonstrated an ~50% reduction in the MFI of CD11b expression in TLR2^−/− neutrophils relative to WT neutrophils (Figure 6K).

To extend these findings, the phagocytosis capacity of TLR2^−/− neutrophils was evaluated. For this purpose, WT and TLR2^−/− bone marrow neutrophils were previously primed with IFN-γ and then infected with CFSE-stained L. infantum promastigote forms for 4 h. The TLR2^−/− neutrophils presented a reduction in parasite uptake compared to WT neutrophils (P < 0.01), even when primed with IFN-γ (P < 0.003; Figure 6L). In agreement with these findings, TLR2^−/− neutrophils presented increased amounts of IL-10 (Figure 6M). These data demonstrate the participation of TLR2 in neutrophil activation, TNF-α production and NO release, consequently interfering with the parasitic uptake ability.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.