FIGURE 1 | PD-L1/PD-L2 surface expression on hMDM and hMDDC after *Lm* infection. (A) hMDM1, hMDM2, or hMDDC were incubated with/without *Lm* (MOI 10) for 24 h and expression levels of PD-L1 or PD-L2 were determined by flow cytometry, respectively. hMDM and hMDDCs were gated using their FSC/SSC properties. Data are presented as mean ± SD RFI (ratio of the mean fluorescence intensity of specific markers to the mean fluorescence intensity of isotype controls). Representative histograms for a surface marker staining of PD-L1 and PD-L2 on infected (black filled histogram) or non-infected (black non-filled histogram) hMDMs and hMDDCs, compared to the respective isotype (black dashed line) control. Statistics were calculated by Wilcoxon matched-pairs signed rank test, *P* < 0.05 is considered statistically significant (\**P* < 0.05; \*\**P* < 0.01). At least three independent experiments were performed (*n* = 7–9). Expression of exhaustion marker after PHA-stimulation of PBLs. (B) PBLs were incubated for 6 days with/without PHA (0.5 µg/mL) and expression levels of the indicated surface molecules were determined *via* flow cytometry. T-cells were gated *via* their FSC/SSC properties and CD3. Data are presented as RFI (ratio of the mean fluorescence intensity of specific markers to the mean fluorescence intensity of isotype controls) or percentage of CD3+ cells. Statistics were calculated using the parametric paired *t*-test (CD3, CD28) or Wilcoxon matched-pairs signed rank test; *P* < 0.05 is considered statistically significant (\**P* < 0.05; \*\**P* < 0.01). At least three independent experiments were performed (*N* = 6–9). hMDM1, proinflammatory human monocyte-derived macrophages type 1; hMDM2, anti-inflammatory human monocyte-derived macrophages type 2; hMDDC, human monocyte-derived dendritic cells; *Lm*, *Leishmania major*; PD-L1, programmed death-1 ligand 1; PD-L2, programmed death-1 ligand 2; MFI, mean fluorescence intensity; PHA, phytohemagglutinin; PBL, peripheral blood lymphocytes; LAG-3, lymphocyte-activation gene 3; TIM-3, T-cell immunoglobulin and mucin-domain containing-3.

#### PD-1/PD-L Blockade-Induced T-Cell Proliferation and *Lm* Infection Rate Are Dependent on the Initial Host Cell Phenotype

First, we addressed, whether T-cell effector functions of PBLsPHA can be reinvigorated by the therapeutic anti-PD-1 antibody nivolumab and whether this is influenced by the type of antigen presenting cell. Therefore, we compared hMDM1, hMDM2 and hMDDC, respectively, as *Lm* antigen presenting cells in cocultures with autologous PBLsPHA. Cocultivation of *Lm*-infected hMDM1 with PBLsPHA significantly increased T-cell proliferation (12.32 ± 9.49%), compared to the uninfected control (4.63 ± 3.33%) (**Figure 2A**). This indicates that PHA-prestimulated T-cells have residual effector functions. Furthermore, by blocking PD-1, T-cell proliferation (25.96 ± 15.07%) was significantly enhanced (**Figure 2A**). Concomitantly, we observed a significant reduction in *Lm* infection in the sample cocultivated with PBLPHA (44.62 ± 6.87%) compared to infected hMDM1 only (69.66% ± 5.07%), which consequently was reduced to a stronger extent upon PD-1 blockade (39.47 ± 9.48%) (**Figure 2A**). In cocultures of *Lm*-infected hMDM2 with autologous PBLPHA, no significant difference in T-cell proliferation was observed (3.84 ± 1.74%) compared to the uninfected control (3.21 ± 2.05%) (**Figure 2B**). However, PD-1 blockade enhanced T-cell proliferation significantly in the infected sample (9.71 ± 5.13%) (**Figure 2B**). Consistently, *Lm* infection rate of hMDM2 in the presence of PBLsPHA was not decreased significantly (49.33 ± 10.90%) compared to infected hMDM2 only (63.49 ± 23.31%). Surprisingly, no significant effect on *Lm* infection could be detected when PD-1 was blocked (45.99 ± 8.94%), even though T-cell proliferation was increased (**Figure 2B**). Compared to hMDM1 and hMDM2 *Lm*-infected hMDDC induced the highest T-cell proliferation in the PBLsPHA coculture (17.71 ± 10.73%) and, after PD-1 blockade, enhanced T-cell proliferation the most (49.30 ± 13.81%) (**Figure 2C**). Consequently, less *Lm*-infected hMDDCs were observed in the presence of PBLsPHA (42.02 ± 9.17%) and PD-1 blockade dampened the infection rate of hMDDCs the most (24.68 ± 8.15%), compared to hMDM1 and hMDM2 (**Figure 2C**). PD-1 blockade in the absence of PBLsPHA did not influence *Lm* infection rate in hMDM1, hMDM2 or hMDDC (Figure S1 in Supplementary Material). Furthermore, we showed that PD-1 is blocked on T-cells throughout the whole coculture experiment, indicating that the used amount of nivolumab is sufficient (Figure S2 in Supplementary Material). Our data demonstrate that T-cell function is enhanced by PD-1 blockade, however, the extent of the effects strongly depends on the *Lm* host cell phenotype.

#### TNF**α** and IFN**γ** Release Is Strongly Enhanced after PD-1 Blockade in the *Lm*-Infected hMDDC Coculture

The proinflammatory cytokines TNFα and IFNγ play a critical role in shaping the immune response against *Leishmania* infection (31). Thus, we analyzed TNFα and IFNγ levels in the coculture supernatants by ELISA. PD-1 blockade significantly increased TNFα levels in the infected hMDM1 samples (216.60 ± 114.90 pg/mL; 88.57 ± 52.52 pg/mL) (**Figure 2D**). Also IFNγ release was augmented by PD-1 blockade in the infected hMDM1 samples (80.61 ± 59.73 pg/mL; 16.95 ± 25.68 pg/mL) (**Figure 2D**). TNFα levels in the hMDM2 samples were similar to the hMDM1 samples. In contrast to hMDM1, IFNγ was barely detectable in the PD-1-blocked infected hMDM2 sample (17.62 ± 24.18 pg/mL) (**Figure 2E**). Regarding *Lm*-infected hMDDC, PD-1 blockade significantly increased TNFα levels in the supernatants (374.50 ± 156.60 pg/mL; 167.20 ± 89.77 pg/mL). IFNγ levels in the supernatant of *Lm*-infected hMDDC were low (25.58 ± 25.54 pg/mL), but highly enhanced by PD-1 blockade (288.40 ± 103.20 pg/mL) (**Figure 2F**). This suggests that effects induced by PD-1 blockade may be mediated by TNFα and IFNγ. To investigate this idea, we performed a correlative analysis and neutralized both cytokines.

#### TNF**α** but Not IFN**γ** Neutralization Partially Reversed Effects of Nivolumab Treatment

To assess whether cytokine release correlates with T-cell proliferation or *Lm* infection rate, we performed a correlative analysis. To this end, we focused on *Lm*-infected hMDDC, because these host cells displayed the strongest effects mediated by PD-1 blockade in the coculture experiment (**Figures 2C,F**). We only used the datasets of infected samples in presence and absence of nivolumab (**Figure 3A**). TNFα release correlated with T-cell proliferation (*r* = 0.90). TNFα release tendentially correlated

were cocultivated with *Lm*-infected host cells and blocked with nivolumab (αPD-1). After 5 days, cells were harvested and analyzed by flow cytometry (hMDM/ hMDDC gated *via* their FSC/SSC properties, T-cells additionally *via* CD3). Data are presented as mean percentage of CFSElow proliferating T-cells ± SD or *Lm* dsRED-infected hMDDC ± SD, respectively. (D–F) IFNγ and TNFα were measured in the coculture supernatants by ELISA as described. Data are presented as mean pg/mL ± SD. For statistics, a parametric paired *t*-test was performed, *P* < 0.05 is considered statistically significant (\**P* < 0.05, \*\**P* < 0.01, \*\*\**P* < 0.001, \*\*\*\**P* < 0.0001). At least three independent experiments were performed (*N* = 5–13). hMDM1, proinflammatory human monocyte-derived macrophages type 1; hMDM2: anti-inflammatory human monocyte-derived macrophages type 2; hMDDC, human monocyte-derived dendritic cells; *Lm*, *Leishmania major*; PD-1, programmed death-1; PBL, peripheral blood lymphocytes; TNFα, tumor necrosis factor α; IFNγ, interferon-γ; CFSE, 5(6)-carboxyfluorescein diacetate N-succinimidyl ester.

inversely with infection rate (*r* = −0.63). There was a positive correlation between IFNγ release and T-cell proliferation (*r* = 0.83). Also infection rate tendentially correlated inversely with IFNγ

release (*r* = −0.79). To investigate a causal connection between increased TNFα/IFNγ release and T-cell proliferation/parasite survival, we neutralized TNFα and IFNγ. Neutralization of IFNγ

FIGURE 3 | Neutralization of TNFα but not IFNγ partially reversed PD-1 blockade-mediated effects. (A) TNFα- and IFNγ-release correlates with T-cell proliferation. All *Lm*-infected hMDDC samples cocultured with PBLPHA in presence or absence of anti-PD-1 (αPD-1) were considered (TNFα and IFNγ: *n* = 12) and Pearson correlation coefficients (*r*), the coefficient of determination (*R2* ) plus the significance level (*P*) were calculated for the indicated datasets. *r* ≥ 0.7 = positive correlation; *r* ≤ −0.7 = inverse correlation. (B) CFSE-labeled autologous PBLsPHA were cocultivated with *Lm*-infected hMDDC, nivolumab (αPD-1), IFNγ neutralizing antibody (αIFNγ) and TNFα neutralizing antibody (αTNFα) as indicated. After 5 days, cells were harvested and analyzed by flow cytometry (hMDM/hMDDC gated *via* their FSC/SSC properties, T-cells additionally *via* CD3). Data are presented as mean percentage of CFSElow proliferating T-cells ± SD or *Lm* dsRED-infected hMDDC ± SD, respectively. For statistics, a parametric paired *t*-test was performed, *P* < 0.05 is considered statistically significant, \**P* < 0.05, \*\**P* < 0.01. At least three independent experiments were performed (*N* = 5–6). hMDDC, human monocyte-derived dendritic cells; *Lm*, *Leishmania major*; PD-1, programmed death-1; PBL, peripheral blood lymphocytes; TNFα, tumor necrosis factor α; IFNγ, interferon-γ; CFSE, 5(6)-carboxyfluorescein diacetate N-succinimidyl ester.

in the infected sample had no significant effect on T-cell proliferation (**Figure 3B**). Neutralization of TNFα tendentially reduced T-cell proliferation compared to *Lm* infection only (2.48 ± 0.80%; 5.02 ± 2.36%). Simultaneous neutralization of both cytokines reduced T-cell proliferation significantly (2.89 ± 1.74%) compared to the untreated control. PD-1 blockade increased T-cell proliferation (26.18 ± 18.11%). Neutralization of IFNγ did not decrease PD-1 blockade-induced T-cell proliferation (20.29 ± 15.08%), whereas anti-TNFα treatment had a significant reducing effect (9.37 ± 7.72%). Simultaneous neutralization of both cytokines did not enhance this effect (8.30 ± 6.25%). In the absence of anti-PD-1 antibody, *Lm* infection rate (47.42 ± 19.11%) was not increased in the IFNγ neutralized sample (51.55 ± 18.48%). Interestingly, TNFα neutralization highly increased parasite survival (65.85 ± 9.31%), which was not further enhanced by additional neutralization of IFNγ (61.87 ± 13.05%). Infection rate in hMDDCs decreased in the presence of anti-PD-1 (23.12 ± 15.78%). Neutralizing single or both cytokines revealed the same effect, as observed for the samples without nivolumab. Again, only TNFα neutralization significantly increased parasite survival (43.40 ± 24.17%) (**Figure 3B**). Blocking with an isotype control did not alter proliferation or parasite survival (data not shown). Even though the correlative analysis pointed toward a role for IFNγ in T-cell proliferation and parasite killing, only neutralization of TNFα partially reversed PD-1 blockade-mediated effects. On the other hand, TNFα neutralization already improved parasite survival in the absence of PD-1 blockade, indicating that TNFα might act independently of PD-1.

#### PD-1 Blockade Shifts *Lm*-Induced CD4**+**-T-Cells toward a TH1- or TH1/TH2 Phenotype

We sought to examine the T-cell response after PD-1 blockade in more detail. Therefore, we characterized the proliferating CFSElow and the non-proliferating CFSEhi CD4+ and CD8+-T-cells in the hMDDC coculture (**Figure 4A**). T-cells were composed of ~60% CD4+ cells and ~40% CD8+ (uninfected control); in the PD-1 blocked *Lm*-infected sample almost exclusively CD4+-T-cells proliferated (33.89 ± 17.95% CFSElow CD4+; 8.52 ± 5.73% CFSElow CD8+) (**Figure 4A**). We confirmed this finding in a coculture experiment using purified CD4+ PBLsPHA or CD8+ PBLsPHA and detected T-cell proliferation only in the presence of CD4+ PBLsPHA (**Figure 4B**). Hence, PD-1 blockade mainly enhanced effector functions of CD4+ T-cells. As mentioned earlier, in the classical mouse model of leishmaniasis the induction of a TH1 response leads to healing, whereas induction of TH2 cells promotes disease (2). In human leishmaniasis such a TH1/TH2 dichotomy is not that evident (5, 6). Thus, we examined in our model whether the TH1-specific transcription-factor Tbet and the TH2-specific transcription-factor GATA3 are differentially expressed upon PD-1 blockade. The representative dot-blots on the left of **Figure 4C** indicate differential intra-nuclear expression of Tbet and GATA3 in presence or absence of nivolumab in the *Lm*-infected sample. Compared to the untreated infected sample, a significantly higher number of Tbet+ (13.06 ± 8.35%; 26.06 ± 9.46%) and Tbet+/GATA3+ (8.68 ± 9.22%; 14.41 ± 11.33%) proliferating CD4+ T-cells was measured in the presence of nivolumab, whereas the percentage of GATA3+ T-cells was tendentially lower (18.77 ± 16.68; 12.68 ± 10.54) (**Figure 4C**). We conclude that PD-1 blockade shifts *Lm*-induced CD4+ T-cells more toward a TH1- or TH1/TH2 phenotype. The higher percentage of TH1 T-cells might be implicated in the improved parasite killing.

#### Intracellular Expression of Perforin, Granzymes, and Granulysin (GNLY) Is Increased in PD-1-Blocked *Lm*-Induced CD4**+** T-Cells

CD4+ T-cells can release an extensive repertoire of cytolytic effector molecules, used either to help or to initiate apoptosis of a target cell (32). To examine, whether PD-1 blockade increases expression of such cytolytic effector molecules, we analyzed intracellular expression of perforin (PRF1), granulysin (GNLY), granzyme A (GZMA), and granzyme B (GZMB) in CFSElowCD4+-T-cells in the hMDDC coculture using flow cytometry (**Figure 4D**). Intracellular expression of PRF1 was significantly higher (1.13– 4.26-fold increase) in the proliferating CD4+-T-cells after PD-1 blockade relative to the untreated infected sample (**Figure 4D**). Like for PRF1, we measured a higher GZMA (1.53–8.17-fold increase), GZMB (1.31–4.3-fold increase) and GNLY expression (1.01–1.93-fold increase) in the infected samples when PD-1 was blocked (**Figure 4D**). To conclude, we determined intracellular PRF1, GNLY, GZMA, and GZMB to be increased in the *Lm*-induced proliferating CD4+ T-cells upon PD-1 blockade. This strongly suggests that reduced parasite survival upon PD-1 blockade is mediated in part by PRF1, GZMA, GZMB, and GNLY.

# DISCUSSION

The function of the coinhibitory PD-1/PD-L pathway and the phenomenon of T-cell exhaustion have been well defined in the chronic LCMV mouse model (18, 28, 33). Moreover, T-cell exhaustion was demonstrated to occur in many human diseases caused by viruses (34, 35), bacteria (36), fungi (37), cancer (38, 39) and protozoan infections including leishmaniasis (24, 40, 41). Remarkably, effector functions of exhausted T-cells can be reinvigorated by blocking the PD-1/PD-L interaction, thereby reducing LCMV loads in the chronic LCMV mouse model (18). This principle has been extended to other applications, like human cancers (42) and animal infection models (21, 23). Experimental animal models of leishmaniasis indicate that the use of PD-1/ PD-L blocking antibodies might be beneficial for treatment of chronic forms of leishmaniasis (21–23).

Previously we reported that *Lm-*infected macrophages generated from monocytes of *Leishmania*-naive German blood donors induce a MHC class II-dependent T-cell response in autologous *in vitro* cocultures. This early T-cell response reduced parasite load in the infected macrophages (11, 27). PD-1/PD-L interactions are not prominent during the early T-cell priming phase, but they regulate the T-cell response during the effector phase (43). During chronic infections and cancer, PD-1/PD-L interactions play an important role in induction of T-cell exhaustion (44).

To investigate PD-1/PD-L interactions in *Lm* infection of human myeloid and lymphoid cells, we used a newly established

immunostained (A) (CD3, CD4, CD8) (C,D) (CD3, CD4, PRF1, GZMA, GZMB, GNLY, Tbet, GATA3) and analyzed by flow cytometry. T-cells were gated *via* their FSC/ SSC properties and the labeled antibodies. Data are presented as (A) percentage of total T-cells, (B,C) mean ± SD percentage or (D) mean ± SD *x*-fold change (MFI) relative to the untreated infected sample. At least two independent experiments were performed (*N* = 4–8). (B) CFSE-labeled autologous CD4+ or CD8+ PBLsPHA were cocultivated with *Lm*-infected hMDDC and nivolumab (αPD-1). After 5 days, cells were harvested and analyzed by flow cytometry. T-cells were gated *via* their FSC/SSC properties and CD3. At least two independent experiments were performed (*N* = 4). Statistics were calculated (B) using the parametric paired *t*-test or (C,D) the Wilcoxon matched-pairs signed rank test; *P* < 0.05 is considered statistically significant (\**P* < 0.05; \*\**P* < 0.01). hMDDC, human monocytederived dendritic cells; *Lm*, *Leishmania major*; PHA, phytohemagglutinin; PBL, peripheral blood lymphocytes; PRF1, perforin; GZMA, granzyme A; GZMB, granzyme B; GNLY, granulysin; Tbet, T-box transcription factor; GATA3, GATA-binding protein 3; CFSE, 5(6)-carboxyfluorescein diacetate N-succinimidyl ester; MFI, mean fluorescence intensity.

*in vitro* model. In this model, we mimicked exhausted T-cells by stimulating PBLs with PHA prior to cocultivation with autologous *Lm*-infected host cells. PHA is a mitogen used for polyclonal activation and expansion of T-cells (45). Furthermore, it is reported to lead to functionally impaired T-lymphocytes in contrast to CD3/CD28 stimulation (30). We confirmed PHA-stimulated T-cells express high levels of PD-1, PD-L1, LAG-3, TIM-3, and 2B4, and thus resemble exhausted T-cells.

In the current study, we found the anti-PD-1 antibody nivolumab to enhance T-cell effector functions and reduce parasite survival differently depending on the *Lm* host cell phenotype that was initially infected. Our data suggest *Lm*-infected hMDM2 not to be target of PD-1/PD-L-mediated inhibition, whereas PD-1/ PD-L interactions strongly inhibit T-cell effector functions on *Lm*-infected hMDDC. A reason for this might be that both PD-1 ligands are higher expressed on hMDDC compared to hMDM2. Furthermore, antigen-presenting dendritic cells are much more potent in inducing T-cell responses compared to macrophages. Arnold et al. compared differently polarized monocyte-derived macrophages and dendritic cells in their ability to induce autologous T-cells by using mycobacterial purified protein derivate as recall antigen or keyhole limpet hemocyanin as a primary antigen for naive T-cell responses. In this context dendritic cells (hMDDC) induced the highest levels of T-cell proliferation to both antigens, whereas LPS + IFNγ-treated macrophages (comparable to hMDM1) were less effective in inducing antigen-specific T-cell responses followed by IL-4-treated macrophages (comparable to hMDM2) (46).

Focusing on hMDDC, we could partially reverse effects of PD-1 blockade by neutralization of soluble TNFα. Chiku et al. demonstrated elevated levels of TNFα and a concomitant decrease of parasite load after PD-1/PD-L blockade in the canine model of visceral leishmaniasis (47).

Furthermore, we found that neutralization of IFNγ, which was induced after PD-1 blockade in the hMDDC samples, had no significant effect on T-cell proliferation or parasite load. IFNγ is reported to mediate resistance to *Lm* in mice by inducing iNOS expression (2). In human myeloid cells iNOS (or NOS2) expression and function is controversially debated (48, 49). *In vitro* generated human myeloid cells are unable to express functional iNOS (NOS2) because the essential iNOS cofactor BH4 is missing (48). In contrast to mice, ROS play an important role in parasite control during human Leishmaniasis. IFNγ was shown to induce ROS in *Leishmania*-infected human monocytes, which dampened overall parasite load (8). Furthermore, ROSdependent parasite control was rather evident in monocytes from CL patients than in monocytes from healthy individuals (7). Thus, it could be that IFNγ-induced ROS production in *Lm*-infected hMDDC was to low to see significant effects of IFNγ neutralization on infection rate in our *in vitro* model. One reason for why we did not observe significant differences in T-cell proliferation might be the general increased survival of T-cells upon IFNγ. However, IFNγ does not increase the number of antigen-specific T-cell divisions. This was shown, e.g., for murine OVA-specific CD4+-T-cells (50).

In our *in vitro* model, preferentially CD4+-T-cells proliferated upon *Lm* infection and nivolumab treatment. In the classical mouse model of *Lm*, induction of TH1 response leads to resistance whereas the induction of a TH2 response promotes disease (2). Human cutaneous leishmaniasis patients with moderate disease symptoms show a balanced TH1/TH2 response, whereas an imbalance of TH1/TH2 is associated with disease severity (51). Experiments with blood of prostate and advanced melanoma cancer patients revealed that PD-1 blockade augments TH1 responses und suppresses TH2 responses (52). By analyzing intranuclear expression of the TH1-specific Tbet and TH2-specific GATA3, we found PD-1 blockade to shift *Lm*-induced CD4+ T-cells more toward a TH1 or TH1/TH2 phenotype, respectively. Thus, the higher abundance of TH1 T-cells and their effector functions may be implicated in reduction of *Lm* infection.

Specific killing of intracellular parasites in a concerted action of perforin, granzymes, and granulysin (expressed by T-cells) with minimal collateral damage to the host cell was demonstrated in transgenic mouse models (53). PD-1 blockade is described to increase perforin, granzyme B, and granulysin expression in T-cells of tuberculosis and cancer patients (54). Focusing on *Lm*-induced CD4+ T-cells, we detected increased intracellular levels of perforin, granzyme A and B, and granulysin after PD-1 blockade. This suggests that cytolytic molecules might contribute to the reduction of *Lm* in hMDDC.

Altogether, our data suggest how the PD-1/PD-L axis could modulate *Lm* host cells and CD4+ T-cells in patients suffering from chronic forms of leishmaniasis. So far, the focus of research groups is mostly on CD8+ T-cell exhaustion, which was observed in diffuse cutaneous (40) and visceral human leishmaniasis (24). In case of the cutaneous forms of human leishmaniasis, CD8+ T-cells have a dual role (55) but their contribution in resolving primary cutaneous *Leishmania* infection might be negligible (56, 57). CD4+ T-cells activate leishmanicidal functions of infected macrophages and dendritic cells. In our experiments infected dendritic cells benefit the most from PD-1 blockade, as this strongly enhanced CD4+ T-cell effector functions and parasite killing. Additionally we found increased levels of maturation markers on *Lm*-infected dendritic cells after PD-1 blockade (Figure S3 in Supplementary Material). To induce a strong cellmediated immunity, e.g., after vaccination, adequate maturation of dendritic cells is important. In animal models, it was demonstrated that vaccination using *L. mexicana* LPG induced PD-1/ PD-L2 expression on several immune cells in a dose dependent fashion (58). Blocking the PD-1/PD-L interaction could be a valuable approach to enhance efficacy of *Leishmania* vaccine candidates. Dendritic cell-based immunotherapy in combination with antimonials has been shown to significantly reduce parasite burden in experimental models of visceral leishmaniasis (59, 60). This approach might also benefit from PD-1 checkpoint inhibitors.

Collectively, by using a limited reductionist approach the present work provides new insights regarding the PD-1/PD-L axis in *Lm* infection of primary human cells and its consequence for adaptive immunity. Further experiments using material obtained from chronic leishmaniasis patients can contribute to a better understanding of PD-1 blockade-mediated effects.

#### AUTHOR CONTRIBUTIONS

CF, PC, ZW, GR, and GVZ contributed to conception and design of the study; CF, KA, and GANN performed and analyzed experiments; CF wrote the first draft of the manuscript; all authors contributed to manuscript revision, read, and approved the submitted version.

#### ACKNOWLEDGMENTS

We thank Wijnand Helfrich for carefully reading and commenting on this manuscript. We thank Bianca Walber and Silvia Isabel Rösser for the technical support.

#### REFERENCES

#### FUNDING

This Project was funded by Paul-Ehrlich-Institut internal funding.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at http://www.frontiersin.org/articles/10.3389/fimmu.2017.01880/ full#supplementary-material.

mononuclear phagocytes. *J Infect Dis* (1993) 167(6):1358–63. doi:10.1093/ infdis/167.6.1358

**Conflict of Interest Statement:** 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.

*Copyright © 2017 Filippis, Arens, Noubissi Nzeteu, Reichmann, Waibler, Crauwels and van Zandbergen. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Absence of Tumor Necrosis Factor Supports Alternative Activation of Macrophages in the Liver after Infection with *Leishmania major*

*Shanshan Hu1,2, Cameron Marshall1 , Jocelyn Darby1 , Wei Wei2 , Alan Bruce Lyons3 and Heinrich Körner1,2\**

*1Menzies Institute for Medical Research, University of Tasmania, Hobart, TAS, Australia, 2 Institute of Clinical Pharmacology, Anhui Medical University, Key Laboratory of Anti-Inflammatory and Immunopharmacology, Ministry of Education, Engineering Technology Research Centre of Anti-Inflammatory and Immunodrugs in Anhui Province, Hefei, China, 3School of Medicine, University of Tasmania, Hobart, TAS, Australia*

#### *Edited by:*

*Christoph Hölscher, Forschungszentrum Borstel (LG), Germany*

#### *Reviewed by:*

*Frank Brombacher, International Centre for Genetic Engineering and Biotechnology (ICGEB), South Africa Elmarie Myburgh, University of York, United Kingdom*

> *\*Correspondence: Heinrich Körner<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Microbial Immunology, a section of the journal Frontiers in Immunology*

*Received: 18 September 2017 Accepted: 03 January 2018 Published: 19 January 2018*

#### *Citation:*

*Hu S, Marshall C, Darby J, Wei W, Lyons AB and Körner H (2018) Absence of Tumor Necrosis Factor Supports Alternative Activation of Macrophages in the Liver after Infection with Leishmania major. Front. Immunol. 9:1. doi: 10.3389/fimmu.2018.00001*

The absence of tumor necrosis factor (TNF) causes lethal infection by *Leishmania major* in normally resistant C57BL/6J (B6.WT) mice. The underlying pathogenic mechanism of this fatal disease has so far remained elusive. We found that B6.WT mice deficient for the *tnf* gene (B6.TNF−/−) displayed not only a non-healing cutaneous lesion but also a serious infection of the liver upon *L. major* inoculation. Infected B6.TNF−/− mice developed an enlarged liver that showed increased inflammation. Furthermore, we detected an accumulating monocyte-derived macrophage population (CD45+F4/80+CD11bhiLy6Clow) that displayed a M2 macrophage phenotype with high expression of CD206, arginase-1, and IL-6, supporting the notion that IL-6 could be involved in M2 differentiation. In *in vitro* experiments, we demonstrated that IL-6 upregulated M-CSF receptor expression and skewed monocyte differentiation from dendritic cells to macrophages. This was countered by the addition of TNF. Furthermore, TNF interfered with the activation of IL-6 induced gp130-signal transducer and activator of transcription (STAT) 3 and IL-4-STAT6 signaling, thereby abrogating IL-6-facilitated M2 macrophage polarization. Therefore, our results support the notion of a general role of TNF in the inflammatory activation of macrophages and define a new role of IL-6 signaling in macrophage polarization downstream of TNF.

#### Keywords: *Leishmania major*, liver, tumor necrosis factor, monocytes, IL-6

#### INTRODUCTION

The infectious disease leishmaniasis has been identified as a "neglected tropical disease" by the WHO and affects a significant number of people worldwide. The infection is caused by members of the intracellular protozoan parasite genus *Leishmania* spp. (1) and occurs when parasites are transmitted to their human hosts by *Phlebotomus* spp. or *Lutzomyia* spp. sandfly vectors during their blood-meal (2). Depending on the specific parasite species and the immune response of the host, infection with *Leishmania* spp. results in a variety of clinical manifestations that can range from self-healing skin lesions to progressive and ultimately fatal infections of visceral organ such as the spleen and liver (3). In the mammalian host, the parasites reside in their amastigote form within macrophages and dendritic cells (DCs) (4). The experimental model of cutaneous leishmaniasis, which is based on a subcutaneous inoculation with isolates of the species *Leishmania major* features a strong genetic dichotomy. Mice of a C57BL/6 background (B6.WT) display a localized, self-healing infection characterized by interferon-γ (IFN-γ) production, while mice of the BALB/c background respond to infection with a preferential production of Th2 cytokines, such as IL-4, IL-10, and IL-13. These mice succumb to the infection after a progressive course of disease and a marked visceralization of the pathogen. This genetic dichotomy as a basis of resistance or susceptibility was instrumental in the development of the T helper (Th)1/Th2 paradigm (1).

The early expression of these respective cytokines has a profound effect on the activation state of macrophages, the predominant host cells of these pathogens. The T cell and NK cell cytokine IFN-γ triggers a classical activation that equips macrophages located in skin and local draining lymph node with mechanisms such as an upregulation of inducible nitric oxide synthase (iNOS) that allow an effective response to the challenge (5). In contrast, the cytokines IL-4, IL-10, and IL-13 result in an alternative activation in macrophages (6) that allows them to contribute to tissue remodeling and wound healing (7). While these cytokines have clearly accepted general roles in the establishment of an immune response to pathogens the proinflammatory cytokine tumor necrosis factor (TNF) has been seen as a cofactor that had an important but limited role in the immune defense (8). Nevertheless, infection experiments using *L. major*, specifically the BNI substrain, and other intracellular pathogens demonstrated a strong protective effect of TNF since *L. major* BNI infected TNF-deficient mice succumbed rapidly to the parasites despite a strong Th1-type response (9, 10). Recent observations have allowed an insight in the underlying deficiencies that cause this susceptibility. It could be demonstrated that in the *L. major* model TNF is necessary to prevent an ill-timed accumulation of alternatively activated macrophages concurrently to classically activated macrophages thus indicating a new and unique role for TNF (11). In the absence of TNF an elevated accessibility of arginase-1 (Arg-1) promoter and enhancer structures permitted a hyper-expression of Arg-1 and caused a subsequent lack of nitric oxide (NO) production presumably due to competition between the two enzymes iNOS and Arg-1, that depleted their common substrate l-arginine (12).

This mechanistic model of TNF-dependent restriction of alternative activation and the consequences for a host that lacks TNF has been established in skin and draining lymph nodes of *L. major* BNI infected mice (12). Other organs such as spleen and liver also show significant reproduction of parasites which is not detectable in TNF-positive hosts. The immune response in these organs has not yet been investigated in more detail and we hypothesized that we would also find increased alternative activation of macrophages. By analyzing *L. major* BNI infection in the liver, we found a comparatively low iNOS expression in B6.TNF−/− macrophages and an accumulation of a myeloid population that exhibited an alternatively activated-like macrophage phenotype, with high expression of Arg-1, CD206 and IL-6. In an *in vitro* assay, we demonstrated that bone marrow-derived DCs treated with IL-6 increased the expression of M-CSFR and generated less CD11c+ cells, while adding TNF reinstated CD11c+ cell generation and concurrently inhibited M-CSFR expression. Furthermore, we could show that both TNF and IL-6 had a regulatory effect on M2 macrophage differentiation which depends on modulation of mIL-6/gp130/signal transducer and activator of transcription (STAT) 3 or IL-4-STAT6. These findings of our study are emphasizing again that expression of TNF is critical to preventing a spread of parasites to visceral organs.

# RESULTS

#### Progressive Liver Infection by *L. major* BNI in B6.TNF**−**/**−** Mice

Infection of C57BL/6 mice that lack an expression of TNF with *L. major* BNI results in a progressive course of disease and visceralization while B6.WT mice contain the infection and recover spontaneously (10). In our infection experiments, a significant lesion was observed in both B6.WT and B6.TNF−/− mice from day 21 after infection. While the footpads swelling remained moderate in B6.WT mice and subsided after day 35, B6.TNF−/− mice showed a progressively increasing footpad swelling (**Figure 1A**). Additionally, in B6.TNF−/− mice the infection spread to visceral organs such as the liver resulting in a mild hepatomegaly (**Figure 1B**) and a significant increase in liver weight (**Figure 1C**). Parasites in the liver were detected from day 21 after infection increasing in number to more than 3 × 105 parasites per gram liver tissue at day 42 postinfection (p.i.), while the liver of infected B6.WT mice remained essentially parasite free (**Figure 1D**).

#### Discrete Inflammatory Foci in the Liver of B6.TNF**−**/**−** Mice during Cutaneous Leishmaniasis

In both B6.WT and B6.TNF−/− mice normal, uninfected liver tissue consists of hexagonal hepatocytes radiating from the region of the central vein toward the periphery. From day 35 after infection, a point in time coinciding with a significant increase of liver weight and parasitic burden (**Figures 1B,C**), abnormal liver structures such as swelling of hepatocytes and diffusely infiltrating inflammatory cells could be detected in *L. major* BNIinfected B6.TNF−/− mice (**Figure 2A**). Additionally, inflammatory foci appeared almost exclusively in infected gene-deficient mice (**Figure 2A**). The generic macrophage marker CD68 was used to determine the phenotype of the cell focus (**Figure 2B**). Weak hepatic expression of CD68 was observed in B6.WT and B6.TNF−/− control groups, which was upregulated at day 42 after infection compared to its B6.WT counterpart. Cells that were CD68+ were mainly detected in areas around the borders of inflammatory foci. Taken together, these results suggested that infiltration of inflammatory cells and hepatic inflammation were significantly elevated in B6.TNF−/− mice, compared to the corresponding B6.WT mice. Interestingly, the number of inflammatory foci was significantly higher in B6.TNF−/− mice as compared to that in B6.WT mice (**Figure 2C**).

#### Cytokines Levels in Serum of B6.WT and B6.TNF**−**/**−** Mice after Infection

Cytokine levels in the serum are an indication of the type of immune response to *L. major* BNI (9). The titers of monocyte chemoattractant protein (MCP)-1, IL-6, IFN-γ, IL-10, TNF, and

\*\**p* < 0.01).

IL-12 (p70) were determined in the serum of *L. major* BNIinfected B6.WT and B6.TNF−/− mice using a cytokine bead array (CBA) and compared to uninfected controls.

Serum levels of the proinflammatory cytokines MCP-1, IL-6, and IFN-γ increased significantly in B6.TNF−/− mice over the course of disease (**Figure 3**). High levels of MCP-1 were detected in the B6.TNF−/− mice after day 28 after infection (**Figure 3A**) indicating that *L. major* BNI-induced inflammation increased the potential to recruit monocytes. IL-6 was found to be increased by 17 times at the same point in the course of infection and 9 times at day 35 and day 42 after infection when compared to infected B6.WT mice (**Figure 3B**). Finally, a high concentration of IFN-γ is characteristic for a Th1-type immune response in both genotypes and was determined to validate our data. Similar to previously published results (9), B6.TNF−/− mice showed a significant increase of IFN-γ (**Figure 3C**) compared to that in B6.WT (day 28 after infection: mean ± SD, 269.45 ± 163.56 versus mean ± SD, 3.50 ± 1.49; day 35 after infection: mean ± SD, 668.39 ± 424.14 versus mean ± SD, 3.50 ± 2.46; day 42 after infection: mean ± SD, 766.03 ± 622.3 versus mean ± SD, 6.14 ± 4.11). Serum concentration of IL-10 (limit of detection 17.5 pg/ml) and IL-12p70 (limit of detection 10.7 pg/ml) remained below the detection limit for the assay in more than half of the serum samples throughout the course of infection (data not shown). TNF was only observed in B6.WT mice (data not shown).

#### A Monocyte-Derived Macrophage (Mo-M) Population Accumulates in the Liver of *L. major* BNI-Infected B6.TNF**−**/**−** Mice

A strong accumulation of inflammatory myeloid cells has been described in skin and draining lymph node in *L. major* BNI infection (11). Furthermore, it has been demonstrated that TNF is necessary for classically activated macrophage phenotype (12). Therefore, we analyzed the quantity and composition of infiltrating inflammatory cells in the liver of B6.WT and B6.TNF−/− mice using comprehensive flow cytometric analysis with two different panels over the course of infection.

Three distinct subsets were characterized based on the expression of CD11b, Ly6C, CD45, and F4/80 as described previously (13). The liver-resident Kupffer cells (KCs) were defined as CD45+F4/80+CD11b−Ly6C− population. Recruited

TNF−/− mice compared to the B6.WT group. Bar = 25 μm for all the pictures. (C) The number of inflammatory foci per area is shown. The *p*-values were calculated using two tailed Mann–Whitney *U*-test (\*\*\**p* < 0.001).

B6.WT and B6.TNF−/− mice (uninfected and 42 days after infection). At day 42 after infection, CD68 immunostaining is markedly increased in B6.

inflammatory monocytes (Mo) which differentiate to Mo-M and potentially to inflammatory DCs (Mo-DC) were defined as CD45+F4/80+CD11blowLy6Chi. Finally, Mo-M displayed the

phenotype of CD45+F4/80+CD11bhiLy6Clow (13–15) (#or gating strategy and flow cytometric identification of subpopulations see Figure S1 in Supplementary Material).

experiments and each experiment was performed in five mice. Error bars denote SD. The *p*-values were calculated using two tailed Mann–Whitney

*U*-test (\*\**p* < 0.01, \*\*\**p* < 0.001).

The number of KCs was not significantly different between the genotypes over the course of a *L. major* BNI infection except for day 21 after infection. At this point in time, the CD45+F4/80hiCD11b−Ly6C− population was around twofold higher in B6.WT mice as compared to B6.TNF−/− mice (**Figures 4A,B**). The number of inflammatory monocytes increased in correlation with the footpad swelling irrespective of the genotype. At day 42, the infiltrating monocytes started to decrease (B6.WT) or reached a plateau (B6.TNF−/−) (**Figures 4A,C**). At the same time, Mo-M which represented a small population in B6.WT mice were elevated significantly in B6.TNF−/− mice (**Figures 4A,D**). To analyze the specific role of TNF in the accumulation of Mo-M, we employed TNFcompetent mice of the BALB/c strain which are highly susceptible to *L. major* BNI infection and display progressive visceralization (Figures S2A,B in Supplementary Material). This control experiment showed a Mo-M population comparable to B6.WT mice and indicated that the accumulation of the Mo-M population depends on TNF.

Infection leads to a strong increase of CD11c+iNOS+TNF+ inflammatory DCs (here termed Mo-DC) at the infection site. These cells are also derived from inflammatory monocytes and act as effector and antigen-presenting cells. Therefore, we investigated whether the absence of TNF affects the differentiation process of Mo-DCs during *L. major*-BNI induced liver infection and we examined the expression of CD11c based on CD45+CD11bhiLy6Chi population. Mice of both genotypes had a very distinct CD11c+ CD11bhi Ly6Chi population of comparable size. In B6.TNF−/− mice the populations displayed a comparatively

by two independent experiments. The *p*-values were calculated using two tailed Mann–Whitney *U*-test (\*\**p* < 0.01).

lower expression of CD11b, Ly6C, and CD11c than B6.WT mice (Figures S3A,B in Supplementary Material), which could indicate differences of inflammatory status or maturity of Mo-DCs.

#### The Mo-M Population in TNF**−**/**−** Mice Display an Alternatively Activated Phenotype with High IL-6 Expression

As shown above, B6.TNF−/− mice fail to clear the parasites from the liver and display an accumulation of a TNF-specific, unique Mo-M accumulation. To further investigate this Mo-M population in B6.TNF−/− mice during murine leishmaniasis, we followed the previously used gating strategy and combined both CD11b+Ly6C+ populations since CD11bhighLy6Clow are missing in B6.WT mice (**Figure 5A**). We characterized the phenotypes using intracellular flow cytometry of the markers IL-6, CD206 and, as control for the quality of the sort, IFN-γ. The intracellular antigens were depicted against SSC (**Figure 5B**). Corresponding to the increased IL-6 secretion in serum of infected B6.TNF−/− mice, an increased level of IL-6 expression was detected in liver macrophages. Although IL-6 is regarded as proinflammatory cytokine, it could be involved in the establishment of a Th2

response which in turn, could modulate the activation pathway of the macrophage differentiation. Additionally, the macrophage mannose receptor CD206, was strongly upregulated. As expected, there was no difference in the presence of IFN-γ between B6.WT and B6.TNF−/− because IFN-γ is not produced by Mo or Mo-M. In summary, we found SSC, IL-6 and CD206 increased in the combined Mo and Mo-M of B6.TNF−/−mice, indicating that this population comprises are large proportion of alternatively activated macrophages in B6.TNF−/− mice during *L. major* BNI infection. The detection of SSChigh cells in B6.TNF−/− mice was striking and indicates a marked presence of cells with high granularity that could represent infected cells.

Previously, it had been shown that a CD11bhi Ly6Clow myeloid population harbored a markedly increased number of parasites in skin and draining lymph nodes (11). Because infected cells with a large burden of parasites are fragile and difficult to detect using flow cytometry, we sorted the distinct Mo and Mo-M populations from B6.WT and B6.TNF−/− mice and conducted a Romanowsky stain (Diff-Quik). There was no distinctive visible difference between Mo and Mo-M cells. In B6.WT mice there were no visible parasite associated with macrophages. In contrast, in both CD11b+Ly6Chi and CD11b+Ly6Clow macrophage populations isolated from B6.TNF−/− mice parasites could be detected inside and on the surface of macrophages (Figure S4 in Supplementary Material).

To further analyze the Mo-M population in the liver during *L. major* BNI infection, we isolated these cells and characterized their phenotype with regard to the gene expression of these marker molecules using qPCR. We confirmed our flow cytometry results and showed that IL-6 and the alternative activation markers including Arg-1 and CD206 were significantly higher expressed in B6.TNF−/− mice as compared to B6.WT mice. In contrast to previous results, in the liver iNOS expression was decreased significantly in B6.TNF−/− mice (**Figure 5C**).

#### Macrophage Activation Is Correlated with Parasitic Burden in Liver

The regulation of myeloid iNOS expression after challenge *in vivo* is complex and influenced by factors such as pathogen, genetic background and cytokine environment. Therefore, we examined the response of CD11b+ cells to *L. major* BNI in the livers of B6.WT and B6.TNF−/− mice (**Figures 6A,B**). Mice from both genotypes showed a strong presence of CD11b+ cells at day 42 after infection while *L. major* amastigotes could only be detected directly in B6.TNF−/− mice (**Figures 6A,B**). The absence of *L. major* in the livers of B6.WT mice was correlated with a strong iNOS expression, while in the absence of TNF, iNOS expression was decreased (**Figure 6A**) similar to the situation in susceptible BALB/c mice (Figure S2A in Supplementary Material) while CD206 expression was increased and associated with a strong presence of *L. major* (**Figure 6B**). Thus, these results suggested a presence of alternatively activated macrophages during leishmanial infection in the liver in B6.TNF−/− mice.

#### TNF and IL-6 Modulate the Process of Monocyte Differentiation to Macrophage or DC *In Vitro*

One of the cytokines which was upregulated significantly in the liver is IL-6 (compare **Figure 3**). The presence of IL-6 has been shown to modulate the differentiation of monocytes and skew the developmental outcome to macrophages rather than DC, which can be reversed by TNF (16). The observed overexpression of IL-6 in B6.TNF−/− mice could facilitate the increased presence of alternatively activated macrophages in the liver. To test this hypothesis, we decided to use mouse bone marrow-derived macrophages to analyze their differentiation in the context of TNF and IL-6 *in vitro*. Bone marrow cells of B6.WT mice were cultured with IL-4 and granulocyte macrophage colony-stimulating factor (GM-CSF) resulting in 64.2% CD11b+CD11c+ DC. The addition of IL-6 at day 3 of culture reduced the generation of cells with this

FIGURE 6 | Inducible nitric oxide synthase (iNOS) and CD206 expression in B6.TNF−/− mice. Immunofluorescence staining of (A) CD11b, iNOS, and *Leishmania major* and (B) CD11b, CD206, and *L. major* in liver tissue of B6.WT (upper panel) and B6.TNF−/− mice (lower panel) 42 days after infection. Single color staining is shown for *L. major* (green), CD206 and iNOS (red), CD11b (gray), and DAPI (blue). The single colors have been merged. The figure represents one of three independent experiments.

phenotype to 51% (**Figure 7**). In contrast, culturing cells in the presence of a combination of IL-6 and 50 ng/ml TNF allowed the expression of CD11c to be restored to previous levels, indicating TNF reversed the differentiation process from monocytes to DCs which was inhibited by IL-6 (**Figures 7A,B**). The combination of GM-CSF and IL-4 potentially caused ectodomain shedding of the M-CSF receptor, inhibiting the differentiation of macrophages from monocytes (17). Thus, we next examined the level of M-CSFR expression when treated with IL-6 and TNF using flow cytometry, immunofluorescence and qPCR (**Figures 7B,C**). Adding IL-6 resulted in an increased level of M-CSFR expression compared to the control group which was consistent with the increased accumulation of DCs (**Figure 7A**). After an additional exposure to TNF, IL-6-treated DCs showed a markedly reduced M-CSFR level (**Figure 7A**). Using immunofluorescence DCs showed increased expression of M-CSFR when treated with IL-6 alone, but its CD11c expression was reduced compared to the control group (**Figure 7B**). An exposure of cells to IL-6 and TNF-α together reduced the expression of M-CSFR and CD11c was elevated. These results were confirmed using qPCR (**Figure 7C**).

#### A Regulatory Balance of TNF and IL-6 Mediate Alternative Macrophage Polarization

Since TNF skewed IL-6-driven differentiation from macrophage to DC, we next examined whether TNF and IL-6 also affected alternative activation as represented by comparing F/80 and CD206 expression patterns. Treatment of the three different types of macrophages with IL-6 alone did not change M0 and M1 phenotype, based on the expression of F4/80+CD206+ compared to the control group (**Figure 8A**). However, a significant increase of F4/80+CD206+ population was observed after IL-4 treatment (**Figures 8A,B**). An analysis using qRT-PCR also revealed increased expression of CD206 and Arg-1 mRNA, and a reduced iNOS mRNA expression (**Figure 8C**), which indicated IL-6 only interfered with IL-4-induced alternative activation. Furthermore, when applied as cotreatment with IL-6 and TNF, M0 and M1 were still not affected based on the parameters assessed, compared to IL-6-treated group. The size of the F4/80+CD206+ population as well as the expression of CD206 and Arg-1 mRNA were significantly reduced with increased concentration of TNF but the iNOS mRNA expression was upregulated in the presence of TNF-α and IL-6 (**Figure 8C**).

#### Effect of a Blockade of the Interaction of TNF and IL-6

To further dissect the regulatory effect between TNF-α and IL-6 in M2 macrophage activation, we used a human TNF-receptor:Fc fusion protein (Enbrel) which is a TNF inhibitor with human/ mouse cross-reactivity (18). After 8 days in M-CSF-containing medium the bone marrow derived undifferentiated macrophages expressed IL-6 (**Figures 9A,G**). Additional exposure to IL-4 for 48 h induced macrophages from both B6.WT and B6.TNF−/− mice to display CD206 (**Figures 9B,H**). The level of IL-6 was largely unchanged in B6.WT macrophages but strongly upregulated in B6.TNF−/− macrophages. Interestingly, macrophages pretreated with increasing concentrations of Enbrel (5, 10, 25, 50 µg/ml) and then exposed to IL-4 not only upregulated the expression of CD206 but also increased the level of IL-6 (**Figures 9C–F**). Together, these data showed that IL-6 was associated with IL-4-driven alternative macrophage activation. The presence of TNF inhibited this process as indicated by a downregulation of Arg-1 and CD206. In contrast, blockade of TNF was facilitating a CD206 and IL-6 expression, supporting recent data obtained in TNF−/− mice (12) and suggesting a balancing effect between TNF and IL-6 in alternative macrophage activation.

#### The Regulatory Effect of the Balance between TNF and IL-6 Affects gp130/ STAT3 and IL-4/STAT6 Signaling

Using qPCR, we examined the expression of the receptor molecules specific for IL-6, IL-6 receptor (IL-6R), and gp130, to determine which signaling pathway is active after stimulation with TNF and IL-6 in M2 macrophages. There was no significant change of IL-6R mRNA while gp130 was increased significantly in the IL-6-treated group but its expression was reduced when cells were additionally exposed to TNF (**Figure 10A**). A Western Blot analysis of IL-6R and gp130 showed an upregulation of these molecules in the presence of IL-6 and a significant downregulation by TNF (**Figures 10B–D**). Further analysis of the levels of STAT3 and 6 demonstrated that the protein expression of these transcription factors was unchanged *per se* but that the level of phosphorylation decreased in the presence of TNF (**Figures 10B,E,F**).

# MATERIALS AND METHODS

#### Mouse Strains

C57BL/6J mice (B6.WT), BALB/c mice (both Jackson Laboratories, Bar Harbor, ME, USA) and C57BL/6J mice genetically deficient for the *tnf* gene (B6.TNF−/−) (19) were used at 6–12 weeks of age. All mice were kept under specific pathogen-free conditions at the animal facilities of the Menzies Institute for Medical Research, Hobart, Australia. Infection was performed with sex- and age-matched B6.WT and B6.TNF−/− mice. Animal care and experiments were approved by the animal ethics committee of the University of Tasmania, Hobart, Australia (Animal Ethics Number: A13934 and A13935).

# Culture of *L. major* and Infection

The virulent *L. major* strain BNI (MHOM/IL/81/FE/BNI) was kindly provided by Prof. Christian Bogdan (Institute of Microbiology, Hygiene and Immunology, Erlangen, Germany). The infectivity of the parasites was maintained by passage through the tissue of susceptible BALB/c mice as described (20). Prior to infection, the parasites were cultured *in vitro* in Novy-MacNeal-Nicolle blood agar slants in RPMI 1640 medium (Thermo Fisher Scientific, VIC, Australia) containing 10% rabbit serum (Applied Biological Products Management, SA, Australia), penicillin/streptomycin, nonessential amino acids, and 10 mM HEPES (Thermo Fisher Scientific) as described (10, 20). Mice were injected with 3 × 106 stationary-phase *L. major* promastigotes bilaterally into the skin of the hind footpads. The progress

FIGURE 7 | The effect of tumor necrosis factor (TNF) and IL-6 in monocyte differentiation. (A) Bone marrow monocytes were cultured for 4 days with granulocyte macrophage colony-stimulating factor and IL-4. IL-6 and/or TNF were added after day 4. At day 7, differentiation was monitored using the expression of CD11b, CD11c, and M-CSFR (CD115) using flow cytometry. A representative result is shown. (B) Quantification the results of a CD11b versus M-CSFR (CD115) staining of three independent differentiation experiments. The *p*-values were calculated using one-way ANOVA (\**p* < 0.05). (C) Immunofluorescence photographs represent sections of cells from each culture condition labeled for M-CSFR (red) CD11c (green) and DAPI (blue). (D) qPCR data revealed M-CSFR gene expression in cells treated with IL-6 and TNF. The results are representative of three independent experiments. Means ± SD were calibrated to median values of three experiments. The *p*-values were calculated using one-way ANOVA and Tukey's comparison test (\**p* < 0.05 and ##*p* < 0.01).

FIGURE 8 | The effect of tumor necrosis factor (TNF) and IL-6 in macrophage differentiation. Bone marrow-derived macrophages were cultivated in the presence of macrophage colony-stimulating factor and harvested after 8 days. Subsequently, macrophages were exposed to LPS and interferon-γ or IL-4 for 24 h to generate M1 and M2 macrophages. The upregulation of the marker molecules F/80 and CD206 was quantified. Unchanged macrophages were considered as M0 phenotype. (A) M0, M1, and M2 macrophages were incubated with IL-6 or IL-6/TNF, and differentiation was analyzed. All the data are presented as means ± SD after normalization to control group values of three experiments. The *p*-values were calculated using one-way ANOVA and Tukey's comparison test (\**p* < 0.05 when compared to the control group, and #*p* < 0.05 when comparing to the IL-6-treated group). (B) Flow cytometry of cells from each culture condition using the macrophage markers F4/80 and CD206. (C) Quantification of the expression of inducible nitric oxide synthase (iNOS), arginase-1 (Arg-1), and CD206 in M2 macrophages treated with IL-6 and TNF using qPCR. Results are representative of three independent experiments, and presented as mean ± SD normalized with regard to the control group. The *p*-values were calculated using one-way ANOVA and Tukey's comparison test (\*\**p* < 0.01 when compared to control group, and #*p* < 0.05, ##*p* < 0.01 when compared to the IL-6-treated group).

of the infection was monitored by measuring the footpad swelling (lesion size) using a metric caliper (10).

One of three independent experiments is shown.

Limiting dilution experiments were performed to determine the parasite burden in the infected liver. Single cell suspensions were prepared in supplemented Schneider's media (Thermo Fisher Scientific) and serial dilutions (threefold) were pipetted across a 96-well plate with 12 replicates in an end-point titration. The plates were incubated for 10–14 days at 27°C before the number of *Leishmania*-positive wells were determined using both a light microscope Olympus CKX31 (Olympus, VIC, Australia) and a spectra Max/M2 Microplate reader (Molecular Devices, Sunnyvale, CA, USA). The parasitic burden was calculated as described (10).

#### CBA Assay

Mouse whole blood was collected by cheek bleeding. Mouse serum was stored at −80°C for further use. Cytokine titers of IL-6, IL-10, MCP-1, IFN-γ, TNF, and IL-12p70 were determined in serum using the mouse inflammation CBA kit (BD Biosciences, NSW, Australia) following the manufacturer's instructions. Samples were acquired on a BD FACSCanto II using FACSDiva 6.1 software and analyzed with FCAP Array version 1.0 software (BD Biosciences).

#### Liver Mononuclear Cell Isolation

Mice were sacrificed by CO2 and then perfused slowly *via* the ascending aorta with 30 ml PBS and EDTA (Thermo Fisher

FIGURE 10 | The regulatory effects of tumor necrosis factor (TNF) and IL-6 affect the expression of gp130 and IL-6R and the phosphorylation of signal transducer and activator of transcription (STAT) 3 and STAT6. (A) qRT-PCR analysis of different signaling molecules in M2 macrophages treated with IL-4 and TNF. Results were normalized to the control group and represent means ± SD of three experiments. Statistical analysis was performing using ANOVA with Tukey's posttest, and the results were considered significant with a \**p* < 0.05, \*\**p* < 0.01 when compared to the control group, and # *p* < 0.05 and ##*p* < 0.01 when compared to the IL-6-treated group. (B) Western Blot analysis of IL-6R, gp130 and b-actin as loading control. Furthermore, the transcription factors signal transducer and activator of transcription (STAT) 3 and STAT6 were analyzed as phosphorylated and non-phosphorylated proteins. One of three experiments is shown. (C,D) Changes of IL-6R and gp130 relative to β-actin are shown. (E,F) The ratio of as phosphorylated and non-phosphorylated STAT3 and 6 is depicted. Results were calibrated to the control group value and represent means ± SD of three experiments. \**p* < 0.05, \*\**p* < 0.01 when compared to the control group, # *p* < 0.05 and ##*p* < 0.01 when compared to the IL-6-treated group. One-way ANOVA analysis with Tukey's postcomparison.

Scientific). Livers were then removed and stored as required. Isolation of liver mononuclear cell was achieved by cutting the organ in small pieces and then digesting it in Hank's Balanced Salt solution (Thermo Fisher Scientific) containing collagenase II (100 U/ml, Thermo Fisher Scientific) and DNase I (1 U/μl, Sigma-Aldrich, NSW, Australia) for 30 min at 37°C shaking at 200 rpm. The suspension was filtered through a 100 µm strainer (Thermo Fisher Scientific) to remove tissue debris. Cells were resuspended in PBS/BSA and mononuclear cell were isolated using a Histopaque 1083 gradient (Sigma-Aldrich). The gradient was initially centrifuged at 80 g for 3 min followed by centrifugation at 1,400 *g* for 15 min at 4°C. The cells at the interface were harvested, resuspended in 10 ml PBS and centrifuged at 600 *g* for 10 min at 4°C. The pellet was resuspended in PBS and the cell number determined in a Neubauer hemocytometer (Australian Scientific, NSW, Australia).

#### Generation and Tissue Culture of Bone Marrow-Derived Macrophages and DC

BM cells were flushed from femur and tibia of uninfected B6.WT and B6.TNF−/− mice and cultured in RPMI 1640 media (Thermo Fisher Scientific) supplemented as described with penicillin/streptomycin, nonessential amino acids, and 10 mM HEPES (Thermo Fisher Scientific) and either 10% of L929 tissue culture supernatant containing macrophage colony stimulating factor (M-CSF) for 7 days (21) or 10% tissue culture supernatant of GM-CSF-transfected X63-AG8 cells for 10 days (22). Recombinant mouse IFN-γ (20 ng/ml; Peprotech, Lonza, VIC, Australia) and IL-4 (10 ng/ml; Peprotech) were added into the medium for 24 h to differentiate M1 and M2, respectively. For the final 24 h, recombinant mouse IL-6 (10 ng/ml, Peprotech), TNF (Peprotech) were added. In blocking experiments increasing concentrations of a human TNFR:Fc (Enbrel; Amgen, North Ryde, NSW, USA) were added for 4 h before the addition of IL-4.

#### Flow Cytometry and Cell Sorting

Multicolor flow cytometry was performed following an established protocol (23).  For intracellular flow cytometry the cells were fixed with FOXP3 Fix/Perm buffer and permeabilized with FOXP3 Perm buffer (BioLegend) according to the manufacturer's protocol. Intracellular proteins were targeted with rat antimouse CD206 (PE; clone C068C2; BioLegend), rat antimouse IL-6 (PE; clone MP5-20F3; BD Biosciences), rat antimouse IFN-γ (PE; clone XMGI-2; BD Biosciences), rabbit anti-*L. major* [clone V121 (11)], and mouse antimouse Arg-1 (PE; polyclonal antiserum; R&D Systems, Sydney, NSW, Australia). Streptavidin conjugated to V500 (BD Biosciences) was used to reveal biotinylated primary mAbs. Cells were acquired on a BD FACSCanto II flow cytometer using BD FACSDiva version 6.1.3 (BD Biosciences) and analyzed with FlowJo software version 10.1(Tree Star Inc., Ashland, OR, USA). For flow cytometric cell sorting, two populations defined by CD45+F4/80+CD11b+Ly6Clow and CD45+F4/80+CD11b+Ly6Chi were sorted using a Beckman Coulter Astrios MoFlo. For liver DC marker comparison, CD11c (PE-Cy7; clone HL3; BD Biosciences) was used. Bone marrow-derived cells were stained with CD11b (FITC; clone M1/70; BD Biosciences), CD11c (PE-Cy7; clone HL3; BD Biosciences), F4/80 (APC-Cy7; clone BM8; eBioscience), CD206 (PE; clone C068C2; BioLegend), and M-CSFR (APC; clone AFS98; Biolegend) as experiments required.

#### Immunohistochemistry

Liver tissue specimen was fixed in formalin and embedded in paraffin. Histological sections of 4 µm thickness were stained with hematoxylin and eosin using a standard protocol. The histopathological changes before and after *L. major* infection were observed using a Leica DM2500 (North Ryde, Australia). To assess the degree of inflammation, three representative inflammatory foci were imaged at 100× magnification. The number of inflammatory foci in each image was quantified and the average for each of the three representative areas per animal was then calculated.

For immunohistochemical staining, tissue sections were deparaffinized in xylene and rehydrated. The antigens were retrieved in 10 mmol/l sodium citrate buffer (pH 9.0) for 10 min at 100°C and then cooled to room temperature before being stained. Endogenous peroxidase activity was quenched by treatment with 3% H2O2 in methanol for 10 min. The sections were blocked in protein block (X0909, Dako, VIC, Australia) for 30 min at room temperature and then incubated with a primary mAb to CD68 (ab31630, Abcam, VIC, Australia) for 1 h at 37°C. Immunoreactivity was visualized with diaminobenzidine (Dako) using the Envision system (Dako) according to the manufacturer's protocol. The nuclei were lightly counterstained with hematoxylin solution. A negative control was prepared using the same staining procedure but was not incubated with the abovementioned primary antibodies. Images were obtained using an Olympus BX53 microscope (Olympus), and semiquantitative analysis was conducted using Image-Pro Plus software (ImageJ, USA).

Flow cytometrically sorted cells were centrifuged on slides in a Cytospin™ 4 Cytocentrifuge (Thermo Fisher Scientific) at 500 *g* for 10 min and underwent microscopical analysis after air-drying and staining with Diff-Quik reagent.

#### Immunofluorescence

Liver tissue samples were dissected and rapidly frozen in Tissue-Tek optimal cutting temperature medium (VWR, QLD, Australia) in liquid nitrogen vapor and stored at −80°C. Sections (8 µm) were cut using a cryotome (Thermo Fisher Scientific), air-dried, and fixed in acetone at −20°C. Prior to staining, sections were rehydrated in PBS/1% BSA for 60 min. Sections were incubated for 60 min with a biotinylated or purified antibody, washed three times with PBS/BSA and labeled for 60 min with a secondary reagent. The primary antibodies and the second labeling reagents are described in **Table 1**. Sections were mounted with polyvinyl alcohol mounting media with DABCO (Sigma-Aldrich) to prevent fading. Anti-*Leishmania* antibodies (clone V121, MHOM/IL/67/Jericho II) were purified from rabbit serum (a generous gift from Emanuela Handman, WEHI, Australia) (11). Immunofluorescence images were visualized using UltraView Spinning disk confocal microscope with Velocity Software (Perkin Elmer, MA, USA). All the images were processed using ImageJ version 1.50i (ImageJ, USA).

#### Quantitative Real-Time PCR

RNA was extracted from sorted liver macrophages and from tissue culture by using Tri-Reagent (Sigma-Aldrich) and RNA

TABLE 1 | Primary and secondary antibodies used in confocal microscopy.

isolation mini kit (Bioline, Alexandria, Australia) according to the manufacturer's instructions. RNA was stored in RNase-free water at −80°C. The QuantiTect Reverse Transcription Kit (Qiagen, Melbourne, VIC, Australia) was used to reverse-transcribe up to 1,000 ng total RNA. cDNA (2 µL) was amplified by Quantitative real-time PCR on the Rotor-Gene Q qPCR instrument (Qiagen) with 10 µl reactions using the SensiFAST™ SYBR No-Rox Kit (Bioline). The appropriate oligonucleotide primers were listed in **Table 2**, and the reaction was performed under the following conditions: samples were heated at 95°C for 3 min and amplified with 40 cycles of 95°C for 10 s, 60°C for 15 s, and 72°C for 30 s. Reactions were performed in duplicate and gene expression levels were normalized to β-actin. Relative gene expression between samples was calculated using the 2−ΔΔCt calculation method.

#### Immunoblotting

Cells were lysed in RIPA lysis buffer (50 mmol/l Tris-HCl, pH 7.4, 150 mmol/l NaCl, 10 mmol/l phenylmethylsulfonylfluoride, 1 mmol/l EDTA, 0.1% SDS, 1% Triton X-100, 1% sodium deoxycholate) for 30–40 min on ice. Protein concentrations were determined using Pierce™ BCA protein assay kit (Thermo Scientific). Proteins were resolved by SDS-PAGE and then transferred to polyvinylidene fluoride membranes (Millipore, VIC, Australia). Membranes were blocked for 1 h at room temperature in 5% skim milk in 0.1% TBS/0.1% Tween20 and then incubated overnight with rabbit polyclonal antibodies to IL-6Rα (Sino Biological, Beijing, China), gp130 (R&D Systems), phospho-STAT6 (R&D Systems) and phospho-STAT3 (Cell Signaling Technology), STAT-3 (Cell Signaling Technology, QLD, Australia), and goat polyclonal antibodies to STAT6 (R&D systems). β-Actin (Abcam) was used as loading control.

Membranes were incubated with appropriate antigoat or antirabbit secondary antibodies (Santa Cruz, QLD, Australia) for 1 h at room temperature. Membranes were washed, incubated with Western Lightning Plus Enhanced Chemiluminescence Solution (PerkinElmer, Woodbridge, ON, Canada) for 1 min and exposed to Amersham Imager 600 (GE Healthcare Life Sciences, NSW, Australia) for 5 s to 10 min. For the analysis of the ratio of phosphorylated/non-phosphorylated transcription factors blots were stripped using mild conditions (Thermo Fisher Scientific) and reprobed with the appropriate antibodies. The density of the specific bands was quantified using Image J software (ImageJ, USA).

TABLE 2 | Primers used for qPCR characterizing monocytes.

#### Statistical Analysis

Statistical evaluation of liver weight, limiting dilution, RT-PCR, and immunohistochemical analysis results were presented as mean ± SD, as appropriate. Results were analyzed using GraphPad prism 5 software (Graphpad Software, San Diego, CA, USA) by Mann–Whitney tests for samples with unknown and potentially disparate variances, or by one-way ANOVA followed by *post hoc* analysis with Tukey's test with *p* < 0.05 accepted as a level of statistical significance.

#### DISCUSSION

Tumor necrosis factor is a pleiotropic cytokine originally named after its proposed tumoricidal effects that has since been identified as a central effector cytokine with a broad range of biological activities such as induction of cell death, modification of cell migration, and regulation of DCs differentiation *in vitro* (24). Interestingly, its presence has been shown to be irreplaceable for effective immune responses to the bacterial or parasitic intracellular pathogens such as *Mycobacterium tuberculosis*, *Listeria monocytogenes*, or *L. major* but the underlying mechanisms that lead to this susceptibility are still not clear (10, 25–27).

After deletion of the *tnf* gene, normally resistant B6.WT mice are unable to control a cutaneous infection with *L. major* BNI (10). They develop a progressive infection that ultimately spreads to visceral organs including the liver. In our detailed analysis of this organ during *L. major* infection, we found that after day 21 after infection the size in B6.TNF−/− mice was enlarged significantly and increasing numbers of viable *L. major* parasites could be detected while B6.WT controls essentially remained parasite free. The progressive infection that was caused by the absence of TNF was accompanied by a strong increase in the proportion of Mo-Ms that showed clear signs of alternative activation. The marked expression of IL-6 prompted us to analyze the interaction of IL-6 and TNF during macrophage differentiation and the signaling pathways *via* IL-6Ra, gp130, and STAT3 and 6 *in vitro*. Our results point to a role of IL-6 facilitating macrophage differentiation downstream of TNF.

Macrophages, while acting as a major reservoir for *L. major* parasites (28), are also potent effector cells that kill parasites *in vivo* with a strong production of NO (29, 30). In the skin and draining local lymph nodes, the tissues predominantly analyzed in *L. major* infection, resident macrophages are initially relevant for the immune response but are quickly outnumbered by inflammatory monocyte derived DCs (31–33). In the liver, which constitutes a major target organ of a visceralized *L. major* infection in immune-incompetent mice, the inflammatory infiltration of monocytic cells has not yet been addressed in detail. In our study, we defined tissue resident KC, recruited inflammatory monocytes (Mo) and the cell types Mo differentiate into, Mo-DCs and Mo-M according to their expression of the marker molecules CD45, F4/80, CD11b, and Ly6C (13–15) and analyzed the dynamic of these cell populations in response to *L. major* infection in the absence of TNF. Fate-mapping has demonstrated that liver-resident F4/80+ KC are derived from embryonic cells and self-maintain independently from hematopoietic input under non-inflammatory steady state conditions (34). These cells have been shown to have an anti-inflammatory function that is abrogated by inflammation (35). Only hours after inoculation with *L. monocytogenes,* KC become infected and rapidly undergo necroptosis, thereby triggering the recruitment of monocytes (13). Unsurprisingly, in the slower moving *L. major* infection we did not see a significant decline of the numbers of KC nor did we detect a significant genotype-dependent changes between B6.WT and B6.TNF−/− during the course of infection (with the exception of day 21 after infection). Interestingly, while the number of liver Mo increased in correlation with the lesion size but independent of the mouse genotype, the population of Mo-M saw a significant increase in TNF-deficient mice but was hardly detectable in B6.WT mice. Since Mo-M in the liver differentiate out of the population of Mo in response to inflammatory signals (13) this observation could point to an absence of these signals in B6.WT mice due to an infection that is contained in skin and draining LN or potentially, to a contributing role of TNF in the differentiation (12).

Monocytes are present in form of steady-state precursors in the peripheral circulation and are recruited to organs such as the liver during inflammation (34). Here they differentiate to Mo-M and Mo-DCs in response to inflammatory cues. In the experiments presented here, the population of Mo increased after day 35 p.i. in line with the increased footpad swelling in both groups of mice. Numbers of Mo began to decrease at day 42 p.i. in B6.WT indicating that during the resolution phase of infection the recruitment of Mo ceased. However, in B6.TNF−/− mice Mo continued to be recruited and to differentiate into Mo-M resulting in a strong presence of this cell type in the late phase of infection. Like TNF-deficient mice the highly susceptible BALB/c mice fail to control leishmaniaisis. However, Mo-M are not found in *L major* infected BALB/c mice, which supports the notion that an accumulation of Mo-M is due specifically to the absence TNF. In previous experiments, a similar Mo-M population was observed in the skin and the draining lymph node of B6.TNF−/− mice during *L. major* BNI infection. It exhibited a phenotype that was CD11b+Ly-6ClowCCR2lowiNOSlow and harbored a large number of parasites (11). In further detailed mechanistic analyzes it could be demonstrated that these cells coexpressed iNOS with high levels of CD206 and Arg-1, which indicated an M2-like phenotype (12). This finding was seemingly contradicting a long line of publications that showed that TNF was supporting the expression of iNOS. The role of TNF in the induction of iNOS and the effector molecule NO during the innate immune response to *L. major* was investigated initially in *in vitro* models (36). It could be shown that it activated macrophages and synergized with IFN-γ to induce effector functions (8, 37). However, in more complex *in vivo* infection models, the well-defined role of TNF in the iNOS-inducing cytokine network determined *in vitro* became controversial (38) while the central effector role of NO remained undisputed (29, 30). A *L. major* infection of a TNFR1-negative mouse strain showed that in absence of this proinflammatory signaling pathway these mice developed persistent lesions but controlled the pathogen (38). Further investigations, using TNFR2- and TNFR1/2-deficient mice clearly demonstrated that the TNFR2 signaling pathway lacked a clear inflammatory function while the outcome of the infection of mice deficient for both TNFR1 and TNFR2 was similar to the TNFR1-negative strain (39). Unexpectedly, it became clear that the expression of iNOS was sustained in these mice (39). Finally, infection experiments using genetically pure TNF-negative C57BL/6 mice resulted in a progressive and ultimately fatal, infection (10) despite a strong Th1 response which was characterized by a hyper-expression of IFN-γ and the presence of iNOS (9, 11). The differences in the published clinical outcomes of these infection experiments were probably due to variations of the genetic background of the parasite strains (40) in combination with a contaminating presence of congenic regions (41) in the genomes of the TNFR1- and 2-deficient mice (39).

Recently, the apparent contradiction of the presence of iNOS in TNF-deficient mice and their concurrent susceptibility to *L. major* BNI infection could be explained with the observation that TNF caused a direct suppression of Arg-1 expression and of other molecules associated with an alternative activation of myeloid cells. In TNF-negative mice the number of Arg1+ cells was increased in skin and draining LN and in the absence of TNF a coexpression of Arg-1 and iNOS could be detected. Since both enzymes share l-arginine as substrate a coexpression in macrophages caused competition for the substrate. Consequently, loss of TNF significantly reduced the production of NO, resulting in fatal leishmaniasis (12). Interestingly, in this model the CD11b+ Arg1+ cells isolated from skin and draining LN of *L. major* BNIinfected B6.WT and B6.TNF−/− mice were predominately coexpressing CD11c+ (12) and had therefore a phenotype that had been described earlier in the leishmanial model (42). This aspect was not addressed in the present study but it should be noted that the concept of inflammatory versus alternative activation can also be observed in DC (43). Taken together, our conclusive detection of M2 macrophages in the liver has strengthened the concept that the M2-suppressing role of TNF is not organ- or tissue-specific and, together with the observation of M2-like cells in the spleen of *L. monocytogenes* infected TNF-deficient mice, supports the notion of a fundamental, yet so far undescribed biological activity of this cytokine (12, 44, 45).

In the present study, we confirmed that IFN-γ was detectable at a higher level in B6.TNF−/− mice than in B6.WT mice (9). Similarly, MCP-1 (46), a pivotal cytokine implicated in recruitment and activation of monocytes (47) was overexpressed in the serum of TNF-deficient mice after *L. major* BNI infection. However, most interestingly, we could show that the major proinflammatory cytokine IL-6 was significantly overexpressed. Historically, after it had been demonstrated that these cytokines were involved in the acute phase reaction (48) this pleiotropic cytokine has been grouped with IL-1 and TNF as a classical proinflammatory mediator that appears early in the immune response (49). The interplay of TNF and IL-6 has been addressed in *in vitro* experiments using DC and macrophages. An activation of TNF-negative bone marrow derived DC resulted in the secretion of a decreased amount of IL-10 but IL-6 production remained unchanged (50). An infection with *L. major* or *L. donovani* with subsequent activation with LPS resulted in a strong IL-6 expression (51). Finally, IL-6 can downregulate the expression of proinflammatory cytokines including TNF (52) and it has been described to skew the differentiation of monocytes to macrophages (53). This action can be reversed by the additional presence of TNF (16) and this activity is caused by opposite effects of these cytokines on M-CSF receptor expression and internalization. One major activities of IL-6 could potentially be important *in vivo* during the immune response to *L. major* BNI. The presence of IL-6 has been reported to block the differentiation of regulatory T cells (Tregs) and to support the generation of IL-17+ Th cells (Th17) (54). However, an infection of mice which were genetically deficient for IL-6 with *L. major* demonstrated an effective and protective antileishmanial response (55) despite originally having been described to have an impaired antibacterial, antiviral, and acute phase response (56); a more detailed analysis of the adaptive and the innate branches of the immune system could not detect major deficiencies in these mice (57).

In the classical IL-6 signaling pathway, IL-6 signal transduction requires the formation of a trimer consisting of ligand, the IL-6 receptor (IL-6R) α-chain and the signal transducing membrane glycoprotein gp130. IL-6 binds to membrane-bound IL-6R (mIL-6Rα) which recruits its signaling component gp130. This combination contributes to the plethora of pro- and anti-inflammatory functions (58). Unlike the ubiquitously distributed gp130, mIL-6Rα is expressed exclusively on the surface of hepatocytes and some myeloid and lymphoid cell populations such as monocytes and T cells (59). However, most proinflammatory functions of IL-6 are mediated through a second IL-6-dependent signaling pathway *via* the soluble IL-6R in the IL-6 trans-signaling pathway which can trigger IL-6-mediated response in cells negative for mIL-6R (58).

In our study, stimulation with IL-6 increased the expression of mIL-6R and gp130 but the additional presence of TNF caused a rapid downregulation of these receptor molecules and a significant reduction of STAT3 and 6 phosphorylation. In the context a *L. major* infection of TNF-competent animals, the role of IL-6 in the determination of the outcome of macrophage differentiation seems to be negligible (57). This changes however, in the absence of TNF. Now a strong expression of IL-6 contributes significantly to the dysregulation that leads to a fatal presence of M2 macrophages in skin (12), lymphoid organs and visceral organs such as the liver. Potentially our finding could have ramifications for the meanwhile ubiquitous anti-TNF and IL-6 therapy in human chronic inflammatory disorders. Therefore, the complex regulatory interactions of IL-4, IL-6, and TNF in macrophage differentiation need to be investigated at the molecular level in more detail (12, 16, 53).

Taken together, we show that in the absence of TNF an infection with *L. major* BNI quickly spreads to the liver. The infection causes a strong infiltration of monocytes and leads to the presence of an accumulating number of Mo-Ms which display an M2 phenotype, express IL-6 and harbor parasites which contributes to the fatal outcome of leishmaniasis in these mice. Furthermore, we show that in the absence of the M1 determining cytokine TNF, anti-inflammatory function of IL-6 can dominate the differentiation of monocytes and provide a first insight that this cytokine can contribute to the skewing of monocytes to a M2 phenotype.

# ETHICS STATEMENT

Animal care and experiments were approved by the animal ethics committee of the University of Tasmania, Hobart, Australia (Animal Ethics Numbers: A13934 and A13935).

# AUTHOR CONTRIBUTIONS

SH: designed experiments, carried out experiments, acquired data, analyzed data, and edited the manuscript. CM and JD: carried out experiments, analyzed data, and edited the manuscript. WW: edited the manuscript and revised the final version. AL: designed experiments, edited the manuscript, and revised the final version. HK: conception of the project, designed experiments, analyzed data, wrote the manuscript, edited the manuscript, and revised the final version.

# ACKNOWLEDGMENTS

We thank Sarah Kane and Drs. Terry Pinfold and David Steele for their technical support and Paul Scowen and his team for animal husbandry. The work was funded by the Menzies Institute of Medical Research. SH was supported by an AMU/UTAS PhD scholarship.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at http://www.frontiersin.org/articles/10.3389/fimmu.2018.00001/ full#supplementary-material.

FIGURE S1 | Macrophages derive from recruited monocytes. Flow cytometric gating strategy and analysis of the three distinct subsets of CD45+ F4/80+ liver macrophages after *L. major* BNI infection.

FIGURE S2 | Analysis by immunofluorescence and flow cytometry in *L. major*-infected BALB/c mice. (A) Immunofluorescence staining of CD11b (gray), iNOS (green), *L. major* (red), and DAPI (blue) in liver tissue of BALB/c mice postinfection. Results represent one of three biological repeats. (B) Flow cytometric analysis revealed the changes of three different CD45+F4/80+ liver

# REFERENCES

macrophage populations based on identical gates as in Figure 3 over the course of *L. major* BNI infection.

FIGURE S3 | The expression of CD11c on monocytic cells from B6.WT and B6. TNF−/− mice. (A) Flow cytometric analysis of liver Ly6C+CD11b+ cells obtained from *L. major* BNI*-*infected B6.WT and B6.TNF−/− mice. The populations of Ly6C+CD11b+ cells were gated and the CD11c expression of these cells is shown in a representative example. (B) Quantification by flow cytometry of the expression of CD11c on the Ly6C+CD11b+ cells upon *L. major* BNI infection. The data represent the median of mean intensity of fluorescence (MIF) of CD11c expression by Ly6C+CD11b+ cells upon *L. major* BNI infection. Results represent means ± SD, *n* = 15. \**p* < 0.05 comparing to B6.WT group, two-tailed Mann– Whitney *U*-test.

FIGURE S4 | The morphology of Mo and Mo-M from B6.WT and B6. TNF−/− mice. (A) Representative Romanowsky (Diff-Quik) stain of liver sections are shown (*n* = 3 per group). After day 42 p.i., *L. major* BNI parasites were found inside and outside of the cells (arrowheads) in the liver of B6.WT and B6. TNF−/− mice (magnification 400×).

kupffer cells display different ontogeny and functions in acute liver injury. *J Immunol* (2014) 193(1):344–53. doi:10.4049/jimmunol.1400574

to endotoxic shock, yet succumb to *L. monocytogenes* infection. *Cell* (1993) 73(3):457–67. doi:10.1016/0092-8674(93)90134-C

**Conflict of Interest Statement:** 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.

*Copyright © 2018 Hu, Marshall, Darby, Wei, Lyons and Körner. This is an openaccess article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Monocyte-Derived signals activate human natural Killer cells in response to *Leishmania* Parasites

*Helena Messlinger1 , Heidi Sebald1 , Lukas Heger 2 , Diana Dudziak2,3, Christian Bogdan1,3\*† and Ulrike Schleicher1,3\*†*

*1Mikrobiologisches Institut – Klinische Mikrobiologie, Immunologie und Hygiene, Universitätsklinikum Erlangen, Friedrich-Alexander-Universität (FAU) Erlangen-Nürnberg, Erlangen, Germany, 2 Laboratory of DC Biology, Department of Dermatology, Universitätsklinikum Erlangen, Friedrich-Alexander-Universität (FAU) Erlangen-Nürnberg, Erlangen, Germany, 3Medical Immunology Campus Erlangen, Friedrich-Alexander-Universität (FAU) Erlangen-Nürnberg, Erlangen, Germany,*

#### *Edited by:*

*Uwe Ritter, University of Regensburg, Germany*

#### *Reviewed by:*

*Birgit Strobl, Veterinärmedizinische Universität Wien, Austria Hira Nakhasi, Center for Biologics Evaluation and Research (FDA), United States*

#### *\*Correspondence:*

*Christian Bogdan<EMAIL_ADDRESS>Ulrike Schleicher<EMAIL_ADDRESS>
> *† Shared senior authorship.*

#### *Specialty section:*

*This article was submitted to Microbial Immunology, a section of the journal Frontiers in Immunology*

*Received: 03 October 2017 Accepted: 04 January 2018 Published: 24 January 2018*

#### *Citation:*

*Messlinger H, Sebald H, Heger L, Dudziak D, Bogdan C and Schleicher U (2018) Monocyte-Derived Signals Activate Human Natural Killer Cells in Response to Leishmania Parasites. Front. Immunol. 9:24. doi: 10.3389/fimmu.2018.00024*

Activated natural killer (NK) cells release interferon (IFN)-γ, which is crucial for the control of intracellular pathogens such as *Leishmania*. In contrast to experimental murine leishmaniasis, the human NK cell response to *Leishmania* is still poorly characterized. Here, we investigated the interaction of human blood NK cells with promastigotes of different *Leishmania* species (*Leishmania major*, *Leishmania mexicana*, *Leishmania infantum*, and *Leishmania donovani*). When peripheral blood mononuclear cells or purified NK cells and monocytes (all derived from healthy blood donors from Germany without a history of leishmaniasis) were exposed to promastigotes, NK cells showed increased surface expression of the activation marker CD69. The extent of this effect varied depending on the *Leishmania* species; differences between dermotropic and viscerotropic *L. infantum* strains were not observed. Upregulation of CD69 required direct contact between monocytes and *Leishmania* and was partly inhibitable by anti-interleukin (IL)-18. Unexpectedly, IL-18 was undetectable in most of the supernatants (SNs) of monocyte/parasite cocultures. Confocal fluorescence microscopy of non-permeabilized cells revealed that *Leishmania*-infected monocytes trans-presented IL-18 to NK cells. Native, but not heat-treated SNs of monocyte/*Leishmania* cocultures also induced CD69 on NK cells, indicating the involvement of a soluble heat-labile factor other than IL-18. A role for the NK cell-activating cytokines IL-1β, IL-2, IL-12, IL-15, IL-21, and IFN-α/β was excluded. The increase of CD69 was not paralleled by NK cell IFN-γ production or enhanced cytotoxicity. However, prior exposure of NK cells to *Leishmania* parasites synergistically increased their IFN-γ release in response to IL-12, which was dependent on endogenous IL-18. CD1c+ dendritic cells were identified as possible source of *Leishmania*-induced IL-12. Finally, we observed that direct contact between *Leishmania* and NK cells reduced the expression of CD56 mRNA and protein on NK cells. We conclude that *Leishmania* activate NK cells *via* trans-presentation of IL-18 by monocytes and by a monocyte-derived soluble factor. IL-12 is needed to elicit the IFN-γ-response of NK cells, which is likely to be an important component of the innate control of the parasite.

Keywords: *Leishmania*, natural killer cells, monocytes, innate immunity, human cutaneous and visceral leishmaniasis

**Abbreviations:** DC, dendritic cell; NK, natural killer; IFN, interferon; ILC, innate lymphoid cell; ft-lysate, freeze–thaw lysate; PBMC, peripheral blood mononuclear cell; Pfa, paraformaldehyde; Th, T helper lymphocytes; TW, transwell.

# INTRODUCTION

Natural killer (NK) cells are members of the innate lymphoid cells (ILCs), which do not express rearranged antigen receptors and are characterized by an absent or only slow clonal expansion. Based on their ability to rapidly release the T helper (Th) 1 signature cytokine interferon (IFN)-γ upon stimulation, NK cells belong to the type 1 ILCs. However, in contrast to other ILC1s NK cells are developmentally dependent on eomesodermin (eomes), require interleukin (IL)-15 instead of IL-7 for cell survival, and kill virally infected or tumor cells by exocytosis of cytotoxic granules [reviewed in Ref. (1)].

As a first sign of activation mouse and human NK cells upregulate the expression of surface CD69, a type II C-type lectin absent from resting NK cells (2–5). CD69 is a costimulatory molecule which is able to enhance NK cell effector functions (6). Depending on the activation signal NK cells can also produce soluble mediators other than IFN-γ including pro- [e.g., tumor necrosis factor (TNF)] or anti-inflammatory cytokines (e.g., IL-10), growth factors (e.g. granulocyte-monocyte colony-stimulating factor), and chemokines (e.g., CCL2-5 and CXCL8) [reviewed in Ref. (7)]. Based on these properties, mouse and human NK cells exert various immunoregulatory functions and contribute not only to the antitumor response, but also to the defense against viruses, bacteria, fungi, and parasites (8–15).

To acquire full effector capacity NK cells require priming by cytokines and accessory cells such as dendritic cells (DCs) (16–19). Cytokines that activate human NK cells include IFN-α/β (20, 21), IL-1 (22), IL-2 (23), IL-12 (24), IL-15, IL-18 (25), IL-21 (26), and IL-27 (27). In most cases, a combination of at least two cytokines is needed to achieve a full NK cell response. In addition, NK cells can also be activated by ligation of pattern recognition receptors such as toll-like receptors (TLRs) or by differential engagement of activating and inhibitory NK cell receptors [reviewed in Ref. (7, 28)]. One of the NK activating receptors, NKp46 [*syn.* natural cytotoxicity triggering receptor (NCR)1] represents the most specific NK cell marker in mammalian organisms (29). Besides NKp46, CD56, also known as neural cell adhesion molecule 1 (NCAM1), is commonly used to define human NK cells as CD56+CD3− cells. In humans, the two main NK cell effector functions, cytotoxicity and cytokine production, have been associated with two distinct NK cell subsets: CD56brightCD16− NK cells that predominate in lymphatic tissues and are specialized in IFN-γ secretion, and CD56dimCD16+ NK cells that are mainly present in peripheral blood and show cytotoxic activity (30, 31). However, dependent on the mode of activation, both NK cell subpopulations may also exhibit the "non-specialized" NK cell effector function (32–34). The function of CD56 on NK cells is largely unknown, but published data indicate a relationship between the height of CD56 expression and the degree of activation (35).

*Leishmania* are protozoan parasites with a dimorphic cell cycle. The flagellated, promastigote form of *Leishmania* is transmitted by the bites of sand flies. In the mammalian host, the promastigotes are endocytosed by phagocytic cells and transform into the aflagellated stage (amastigotes) that replicates within phago(lyso) somes (36). Depending on the *Leishmania* species and strain and the immune response and genetic background of the host, infections can be asymptomatic, lead to self-healing or chronic cutaneous leishmaniasis (CL; e.g., *Leishmania major*, *Leishmania mexicana*) or non-healing, progressive mucocutaneous leishmaniasis (e.g., *Leishmania braziliensis*), or can cause visceral leishmaniasis (VL; *Leishmania infantum* and *Leishmania donovani*) due to systemic spreading of the parasites (37). Experimental animal models of CL and VL led to the identification of the key immune mechanisms required for the control of infection, which include the generation of IL-12 and TNF, the expansion of IFN-γproducing CD4+ and CD8+ T cells and the induction of antileishmanial effector pathways such as inducible nitric oxide synthase (iNOS). By contrast, induction of macrophage-deactivating cytokines such as IL-10 and transforming growth factor (TGF) β as well as overshooting production of Th2 cytokines were associated with disease progression [reviewed in Ref. (38–40)]. Many of the above-mentioned mechanisms also hold true in human leishmaniasis, as biopsies of chronic CL lesions and leukocytes of VL patients displayed high IL-10 and TGFβ content, whereas cells of cured patients produced IL-12 and IFN-γ (41–46).

Natural killer cells were found to participate in the innate control of *Leishmania* in infected mice but were not essential for generating a Th1 response and ultimate healing of the disease [reviewed in Ref. (13)]. During later stages of VL, mouse NK cells showed adverse effects and inhibited protective immunity in an IL-10-dependent manner (47). The protective function of NK cells in murine leishmaniasis is largely due to their release of IFN-γ and subsequent stimulation of iNOS-dependent killing of parasites, as they were not able to recognize *Leishmania*-infected host cells as targets for direct cytolysis *in vitro* and *in vivo* (48). During the early phase of infection, NK cell activation in *Leishmania*infected mice required DC- and TLR9-dependent production of IL-12, T cell-mediated release of IL-2, and the presence of IL-18 (18, 49, 50). In *L. major* infections of mice, IFN-α/β was necessary for full NK cell activation (51). *Leishmania* parasites failed to directly activate mouse NK cells (18).

Several observations argue for a protective role of NK cells also in human leishmaniasis. These include (a) a reduced NK cell number in the blood of patients with acute VL that was restored after successful chemotherapy; (b) the influx of NK cells into lesions of CL patients, who showed suppressed NK cell cytotoxicity during active disease, but positive response to treatment (52–54); and (c) a reduced number, TLR expression and IFN-γ and TNF-production by NK cells in patients with diffuse as compared with localized CL due to *L. mexicana* infection (55, 56). Unlike to murine NK cells, mechanisms of human NK cell activation are less clear. Some studies claimed indirect stimulation of human blood NK cells by accessory cells releasing cytokines after contact to *Leishmania* (57–59). Other reports suggested direct activation of NK cells by *Leishmania* in a lipophosphoglycan (LPG)/TLR2-dependent or LPG-independent manner (60, 61) or even excluded a NK cell IFN-γ response in *Leishmania*- or *Leishmania* antigen-stimulated peripheral blood mononuclear cells (PBMCs) (62, 63).

To define the activation signals required for a human NK cell effector response to *Leishmania* parasites and to address the question whether there are differences between *Leishmania* species, we performed cocultures of human PBMCs or highly purified cell populations from healthy German volunteers with four different *Leishmania* species and analyzed the NK cell response. The data obtained show that NK cells cannot be directly activated by *Leishmania* promastigotes but require cytokine signals from monocytes.

# MATERIALS AND METHODS

#### *Leishmania* Parasites

Promastigotes of the following *Leishmania* species and strains were used: *L. infantum* MHOM/DE/98/LUB1 [isolated in our laboratory from bone marrow (BM) of a German patient with VL] (64), *L. infantum* MHOM/DE/2012/VA21737 (isolated in our laboratory from BM of a German patient with VL), *L. infantum* MHOM/DE/2014/VA20763 (isolated in our laboratory from the skin lesion of a Croatian patient with CL), *L. infantum* MCAN/ ES/2010/BON (isolated in our laboratory from peripheral blood of a Swiss dog with VL), *L. major* MHOM/IL/1981/FEBNI (isolated from the skin lesion of an Israeli patient with CL) (65), *L. mexicana* MNYC/BZ/1962/M379 [isolated from a vesper rat (ATCC® 50156™); kindly provided by Sigrid Roberts, Hillsboro, OR, USA] and *L. donovani* (MHOM/SD/1962/1S-CL2D clonal line LdBob; originally isolated from a Sudanese patient with VL; kindly provided by Steve Beverley, St. Louis, MO, USA) (66). In case of *L. infantum,* the strain MHOM/DE/98/LUB1 was used unless otherwise stated. For all strains, aliquots of a promastigote culture (derived from amastigotes isolated from experimentally infected mice) were frozen after only two to three *in vitro* passages for expansion. All experiments were performed with freshly thawed aliquots of these promastigotes which were grown at 28°C/5% CO2/95% humidified air in modified Schneider's *Drosophila* insect medium as described (67) for a maximum of six *in vitro* passages. For fixation of promastigotes, parasites were incubated for 10 min in 4% paraformaldehyde (Pfa) at room temperature (RT) followed by three washes with PBS. Freeze–thaw (ft) lysates of *Leishmania* promastigotes were generated by four cycles of freezing at −80°C and thawing at RT.

#### PBMC Preparation and Purification of Different Cell Populations from the Blood

Mononuclear cells from EDTA-anticoagulated human peripheral blood (PBMCs) of healthy human volunteers living in Erlangen and without any history of leishmaniasis were isolated using density centrifugation (1.077 g/ml Biocoll, Biochrom). For generation of autologous plasma, blood was first centrifuged, and the resulting plasma supernatant (SN) was heat inactivated (56°C, 30 min) and filtered, while the remaining cell pellet was resuspended in PBS to proceed with PBMC preparation. Different cell populations of PBMCs including CD3+CD56− T cells, CD19+ B cells, CD56+CD3− NK cells, CD14+ monocytes, CD14highCD16−CCR2high classical monocytes, CD14lowCD16highCCR2− non-classical monocytes, and CD14highCD16lowCCR2+ intermediate monocytes were purified by cell sorting using the FACS Aria II instrument (BD Biosciences). Purification of CD1c+ DCs was performed as described with slight modifications (68). Briefly, leukocyte reduction cones retrieved from anonymous healthy adult donors were used as source for PBMCs. CD1c+ DCs were then enriched with the EasySep Pan-DC Pre-Enrichment Kit (Stemcell Technologies) and isolated by cell sorting using an FACS Aria II (BD Bioscience) as CD3−CD14−CD19−CD20−CD56−HLA-DR+CD1c+CD11c+ cells. All sorted cell populations showed a purity of >96%. For immunofluorescence analysis, monocytes were purified by negative selection using the Monocyte Isolation Kit II (Miltenyi Biotech) following the manufacturer's instructions.

# Culture and Stimulation of Cells

Human leukocytes with or without *Leishmania* promastigotes were cultured in 96-well plates (PBMCs: 5 × 105 cells/well, 200 µl; purified NK cells: 0.5–1.5 × 105 /well depending on the total recovery, 200 µl), 48-well plates (PBMCs: 106 cells/well, total volume 500 µl; purified monocytes: 5 × 105 /well, 500 µl), or in 24-well plates with a transwell (TW) insert (0.4 µm pore size; Corning, Wiesbaden, Germany; 5 × 105 NK cells/insert, 0.5–1 × 106 monocytes/bottom well) at 37°C and 5% CO2/95% humidified air for 20 h using RPMI1640 (Gibco™ Life Technologies; ThermoFisher Scientific, cat. no. 21875-034) supplemented with 10 mM HEPES (ThermoFisher Scientific), 50 µM 2-mercaptoethanol (Sigma-Aldrich), 100 U/ml penicillin and 100 µg/ml streptomycin (ThermoFisher Scientific), and 10% heat-inactivated autologous plasma. *Leishmania* promastigotes were added at different parasite/host cell ratios [multiplicity of infection (MOI) 0.2, 1, 5, 10, or 33]. Pfa-fixed *Leishmania* promastigotes or *Leishmania* freeze– thaw lysate were used in analogy to the MOI of viable parasites. When different purified leukocyte populations (e.g., NK cell and monocytes) were cocultured with *Leishmania* promastigotes, cell populations were used at the same ratios as present in nonseparated PBMCs of this donor, unless otherwise stated. In some of the experiments, leukocyte/promastigote cocultures were incubated in the presence of specific blocking antibodies (Abs) against different cytokines [sheep-anti-IFN-α (1:350; 10,000 neutralizing units/ml) or sheep-anti-IFN-β antiserum (1:3; 1,000 neutralizing units/ml), obtained from the NIAID Repository, Braton Biotech Inc., Rockville, MD, USA; mouse-anti-IL-1β, 10 µg/ml, CRM56, eBioscience/ThermoFisher Scientific; mouse-anti-IL-2, 1 µg/ml, AB12-3G4, eBioscience/ThermoFisher Scientific; rat-anti-IL-6, 5 µg/ml, MQ2-13A5, BioLegend; mouse-anti-IL-12/IL-23p40, 20 µg/ml, C11.5, BioLegend; mouse-anti-IL-15, 1 µg/ml, ct2n, eBioscience/ThermoFisher Scientific; and mouse-anti-IL-18, 1.5 µg/ml, 125-2H, MBL] or the respective control sera or isotype control Abs. To verify the efficacy of the Ab-treatment, cells were stimulated with the appropriate recombinant cytokine. Cytokines/chemokines used were as follows: huIFN-α and huIFN-β (100 U/ml; NIAID Repository, Braton Biotech Inc., Rockville, MD, USA), rhuIL-1β (20 ng/ml, PeproTech), rhuIL-2 (200 U/ml; Chiron, Emeryville, CA, USA), rhuIL-4 (250 U/ml, PeproTech), rhuIL-6 (10 ng/ml, BioLegend), rhuIL-8 (10 ng/ ml, BioLegend), rhuIL-12p70 (10 ng/ml, PeproTech), rhuIL-15 (12 ng/ml, PeproTech), rhuIL-18 (10 ng/ml, MBL), and rhuMIP1α (20 ng/ml, BioLegend). In addition, PBMCs or purified NK cells were stimulated with cell culture SNs (vol/vol 20–80%) of previous leukocyte/*Leishmania* cocultures of the same donor. In some cases, the SNs were incubated with blocking Abs to cytokines (see above) for 1–2 h at 37°C and 5% CO2/95% humidified air before being added to freshly isolated cells.

As a positive control for the stimulation of DCs resiquimod (R848, 5 µg/ml, InvivoGen), a TLR7/8 agonist, was used.

#### Cytokine Measurements in Cell Culture SNs

Commercial ELISA kits were used for determining the content of human IL-2, IL-6, IL-18, or IL-12p35/70 (eBioscience/ThermoFisher Scientific) and IL-10 or IL-12/IL-23p40 (BioLegend) in culture SNs. Multiplex ELISA was performed using the ProcartaPlex® Multiplex Immunoassay (Human Cytokine/Chemokine/Growth Factor Panel 1, 45 plex; eBioscience/ThermoFisher Scientific), which was analyzed with a MAGPIX® instrument and the xPONENT® software (eBioscience/ThermoFisher Scientific).

# Flow Cytometry

For surface phenotyping and cell sorting of freshly isolated or cultured leukocytes, fluorochrome-labeled or biotinylated Abs against the following antigens were used (all from eBioscience/ ThermoFisher Scientific, unless otherwise stated): CD3ε (OKT3, FITC, PerCP-Cy5.5, biotinylated), CD11b (ICRF44, V450, BD Biosciences), CD11c (3.9, PerCP-eflour®710), CD14 (61D3, FITC), CD16 (CB16, PerCP-eflour®710, eflour®450), CD19 (HIB19, PE, eflour®450), CD25 (BC96, PE-Cy7), CD56 (CMSSB, PE-Cy7, APC), CD69 (FN50, PerCP-Cy5.5, BioLegend), CD192/CCR2 (K036C2, PerCP-Cy5.5, PE-Cy7; BioLegend), and CD335/Nkp46 (9E2, PE, Miltenyi Biotech). Staining with biotinylated Abs was followed by incubation with fluorochrome (FITC or APC)-labeled streptavidin (BD Biosciences) to allow detection.

Staining of blood CD1c+ DCs was done as described with minor modifications (68). Briefly, after enrichment of human DCs with the EasySep Pan-DC Pre-Enrichment Kit (Stemcell Technologies), cells were stained with fluorochrome-coupled Abs against CD1c (L161, APC/Cy7, BioLegend), CD3 (UCHT1, BUV395, BD Bioscience), CD11b (M1/70, Alexa Fluor 700, BioLegend), CD11c (3.9, PE/Cy7, BioLegend), CD14 (HCD14, Alexa Fluor 700, BioLegend), CD19 (HIB19, V450, BD Bioscience), CD20 (2H7, eflour®450), CD56 (5.1H11, Brilliant Violet 421, BioLegend), CD123 (6H6, BV605, BioLegend), CD141 (1A4, Brilliant Violet 711, BD Bioscience), CD303a (201A, PerCP-Cy5.5, BioLegend), and HLA-DR (L243, Brilliant Violet 510, BioLegend) for 30 min on ice.

The specificity of the stainings was verified by use of isotype control Abs. Cells were analyzed with an FACS Canto II system and Diva 6.1.2 (both BD Biosciences) and FlowJo 10.0.7 (FlowJo LLC, Ashland, OR, USA) software. DCs were analyzed with an FACS LSRFortessa™. For intracellular staining of IFN-γ, GolgiStop™ (1:1,500 µg/ml, BD Biosciences) was added during the final 6–10 h of cell culture to prevent secretion of cytokines. After staining of surface molecules, cells were fixed by Cytofix/ Cytoperm™ (BD Biosciences), washed twice with a saponincontaining buffer, and stained for intracellular accumulated IFN-γ (α-huIFN-γ, 4S.B3, APC) (18).

# Confocal Laser Scanning Fluorescence Microscopy (CLSFM) of Infected Monocytes

After coculture of untouched purified monocytes with *Leishmania* promastigotes (MOI 10) for 20 h, 2.5 × 105 cells in 30 µl were transferred to the marked reaction field of adhesion slides (Marienfeld Laboratory Glassware) prepared as recommended by the manufacturer. After cell adhesion, slides were washed twice in PBS buffer and cells were fixed with 4% Pfa. Fixed monocytes were either directly stained or additionally permeabilized with methanol (−20°C) before staining. For IL-18 staining, non-specific binding sites were blocked with PBS/2% BSA/10% normal goat serum and cells were stained with mouse-anti-IL-18 monoclonal Ab (125-2H, MBL) overnight at 4°C. As specificity control, the mouse-anti-IL-18 mAb was pretreated with rhuIL-18 (1.2 µg/ml, 30 min, 37°C). All Abs were diluted in PBS/0.5% BSA/0.5% normal goat serum. After washing with PBS/0.1% Tween Alexa Fluor 568-conjugated goat anti-mouse secondary Abs (ThermoFisher Scientific) were added for 30 min at RT. Cell nuclei were visualized by DAPI staining. Slides were mounted in Vectashield (Vector laboratories) and cover slips, dried in the dark for at least 12 h at 4°C, and analyzed by CLSFM (LSM700, Zeiss) using a 63× objective. Image processing was performed using the ZEN software 2009 (Zeiss).

# Cytotoxicity Assay

Peripheral blood mononuclear cells (with the percentage of Nkp46+CD3− NK cells determined by flow cytometry) were added to K562 tumor target cells in NK cell/target cell ratios of 20:1, 10:1, 5:1, and 2.5:1. A standard chromium-51 release assay was performed (48). Briefly, K562 tumor cells were labeled with ~150 μCi 51Cr (Perkin-Elmer) for 90 min. Cocultures of effector and target cells were incubated in complete RPMI1640 medium containing 10% heat-inactivated fetal calf serum (Sigma-Aldrich, cat. no. F-7524, lot. no. 036K3397) for 4 h. The release of 51Cr into the SNs was measured as counts per minute (cpm) using a TopCount NXT microplate gamma counter (Perkin-Elmer). Based on the spontaneous (target cells alone) and the maximum release (51Cr-labeled cells directly added to the LUMA measurement plate) % specific lysis was calculated as (cpmsample − cpmspontaneous)/ (cpmmaximum − cpmspontaneous) × 100.

# RNA Preparation and Quantitative RT-PCR

Total RNA was prepared with the RNeasy Mini Kit (Qiagen). cDNA synthesis and quantitative RT-PCR analysis were performed (49) using the following assays: NCAM1 (CD56) (Hs00941830\_m1), GAPDH (Hs02758991\_g1).

# Statistical Analysis

Results were displayed as mean ± SEM or as median and were statistically analyzed by the Mann–Whitney *U* test using GraphPad Prism software v.4. Significant differences between unstimulated and stimulated samples were marked by asterisks, significant differences between stimulated samples by diamonds. Significant *p* values are indicated as follows: \*,#*p* < 0.05; \*\*,##*p* < 0.01; \*\*\*,###*p* < 0.001.

#### RESULTS

#### NK Cells within PBMCs, but Not NK Cells Alone Upregulate CD69 in Response to *Leishmania*

To investigate whether *Leishmania* parasites themselves and/ or host cell-derived factors activate human NK cells, PBMCs of healthy volunteers without history of leishmaniasis were cultured in the presence of promastigotes of different *Leishmania* species. Blood NK cells gated as NKp46+CD3− viable single cells were analyzed for surface expression of the early activation marker CD69 by flow cytometry after 6, 12, and 20 h of incubation. As induction of CD69 was most prominent after 20 h (Figure S1 in Supplementary Material) and the viability of NK cells decreased thereafter, this time point was chosen for further analyses. Stimulation with promastigotes of all *Leishmania* species tested induced upregulation of CD69 on human NK cells in a large number of different blood donors (total of 36), most of which were tested several times in independent experiments (**Figure 1A**). The average induction of CD69+ NK cells by *L. infantum* was lower than by *L. major*, *L. mexicana*, and *L. donovani*. There were no differences observed between viscerotropic and dermotropic human strains or a canine strain of *L. infantum* (**Figure 1B**). The percentage of CD69+ NK cells increased with the parasite/host cell ratio used (**Figure 1C**). Fixed or lysed parasites still caused an induction of CD69 on NK cells, which, however, was tentatively or significantly reduced when compared with viable parasites (**Figure 1D**). By contrast, CD69 induction on NK cells was abolished when *Leishmania* promastigotes were separated from PBMCs by a membrane (pore size 0.4 µm) (**Figure 1E**).

Having seen that direct contact between *Leishmania* and NK cells or other cell types within the PBMC culture was necessary to upregulate CD69, we next investigated whether the parasite was able to directly activate human NK cells. To this end, highly purified CD3−NKp46+ NK cells as well as whole PBMCs of the very same donor were stimulated by promastigotes (**Figure 2A**). Whereas NK cells within the PBMC/*Leishmania* coculture readily upregulated CD69, purified NK cells failed to do so, irrespective of the *Leishmania* species used. From these

Figure 1 | CD69 is upregulated on natural killer (NK) cells after contact with *Leishmania*. Human peripheral blood mononuclear cells were cocultured for 20 h with *Leishmania* promastigotes of different species (A), strains (B), amounts (C), or integrity (D), and with or without host cell/parasite contact (E) or interleukin (IL)-12 and IL-18 (10 ng/ml) (A,B), followed by surface expression analysis of CD69 on NKp46+CD3− NK cells by flow cytometry. Unless otherwise indicated the multiplicity of infection (MOI) was 10. (A) Results of 129/117/107/83/21/88 blood samples for the six stimulations. Medians are indicated by red lines. FACS plots show results of NKp46+CD3− NK cells of one representative experiment. (B) *Leishmania infantum* strains isolated from human patients with visceral leishmaniasis (VL) (VL1, VL2) or CL or a dog [canine leishmaniasis (CanL)]. Mean ± SEM of 9/9/9/9/7/9 donors. (C) Mean ± SEM of 119/43/17/119 donors for the four stimulations. (D) Mean ± SEM of 22/22/18/18 (viable), 8/9/6 [paraformaldehyde (Pfa)-fixed], and 17/15/14 [freeze–thaw lysate (Ft-lysate)] donors. (E) Mean ± SEM of six donors. Abbreviation: TW, transwell. \*,#*p* < 0.05; \*\*,##*p* < 0.01; and \*\*\*,###*p* < 0.001 two-tailed Mann–Whitney *U* test.

data we conclude that the upregulation of CD69 on NK cells in response to *Leishmania* is dependent on the presence of accessory cells.

#### Upregulation of CD69 on NK Cells Requires Infected Monocytes Trans-Presenting IL-18

To elucidate which additional cell population is needed to activate human NK cells in response to promastigotes, CD3+Nkp46− T cells, CD19+ B cells, and CD14+ monocytes were sorted and added separately to a coculture of *Leishmania* and purified NK cells of the same donor. Addition of monocytes to the NK cell/parasite culture restored induction of CD69 on NK cells to a similar level as observed in whole PBMC cocultures, whereas addition of T or B cells did not support the expression of CD69 on NK cells (**Figure 2A**). As human CD14+ monocytes are subdivided in classical (cMo, CD14highCD16−CCR2high), intermediate (intMo, CD14highCD16lowCCR2+), and nonclassical monocytes (ncMo, CD14lowCD16highCCR2−) (69), the three subpopulations were purified and evaluated for their capacity to induce CD69 on NK cells after a 20 h coculture with *Leishmania* and purified NK cells. All three types of monocytes were able to induce CD69 on NK cells in response to *Leishmania* (**Figure 2B**).

To define whether infected monocytes stimulated NK cells *via* a soluble factor or by a cell contact-dependent mechanism, a TW system was used. Whereas NK cells of a mixed NK cell/monocyte/*Leishmania* culture showed an increase in CD69 expression after 20 h of incubation, NK cells that were separated from infected monocytes by a membrane did not (**Figure 3**). Likewise, NK cells did not become activated in TW experiments, in which NK cells and *Leishmania* were separated from uninfected monocytes (two independent experiments, data not shown). Thus, direct contact between infected human monocytes and NK cells is essential to upregulate CD69 on NK cells.

In murine leishmaniasis NK cell activation is mediated by cytokines [reviewed in Ref. (13)]. We therefore hypothesized that this might also apply for human NK cells and screened for NK cell-activating cytokines that are trans-presented by myeloid cells to the respective receptor on NK cells without being necessarily secreted. As both IL-15 and IL-18 were reported to be trans-presented by human monocytes (70, 71), we tested whether they are involved in the induction of CD69. Using neutralizing Abs, we found that the expression of CD69 on NK cells in infected PBMCs was partially dependent on IL-18 (**Figure 4A**). In the case of NK cell/monocyte/*Leishmania* cocultures, a similar effect was observed, which, however, did not quite reach the level of significance (**Figure 4A**). By contrast, neutralization of IL-15 did not affect the expression of CD69 on NK cells (**Figure 4B**). Two observations argue for monocyte-mediated trans-presentation rather than secretion of IL-18: first, in most of the SNs of PBMC/*Leishmania*, NK cell/monocyte/*Leishmania* or monocyte/*Leishmania* cocultures IL-18 was not measurable by ELISA (detection limit was 20 pg/ml); only in few of them (mostly after *L. major* stimulation) low levels of IL-18 (≤500 pg/ ml) were found (Table S1 in Supplementary Material). Second, IL-18 was visualized on the surface of purified monocytes which had been in contact with *L. major* for 20 h and were stained for IL-18 after fixation with Pfa without permeabilization (**Figure 5**). Permeabilization of infected monocytes intensified the IL-18 staining, because intracellular IL-18 became additionally detectable (**Figure 5**). Together, these data suggest that *Leishmania*-infected monocytes trans-present IL-18 to human NK cells.

#### A Heat-Labile Soluble Factor of Monocyte/*Leishmania* Cocultures Contributes to the Upregulation of CD69 on Human NK Cells

To investigate whether soluble factors released by infected monocytes are additionally involved in human NK cell activation, freshly isolated PBMCs or purified NK cells were incubated with culture SNs of *Leishmania*-stimulated (a) PBMCs, (b) purified NK cells (with or without monocytes), or (c) purified

monocytes, all from the same blood donor. SNs of host cell-free *Leishmania* cultures were included as control. In the presence of SNs from previous PBMC/*Leishmania* or monocyte/*Leishmania* cultures, both NK cells within PBMCs and purified NK cells showed an upregulation of CD69, indicating that monocytes and/ or *Leishmania* release a soluble factor after contact to each other that activates NK cells; SNs of promastigotes cultured without host cells had no effect (**Figure 6A**, upper and lower panels). The stimulatory effect of SNs derived from PBMC/*Leishmania* cocultures was concentration-dependent and, except for *L. mexicana*, as strong as direct stimulation of PBMCs by the parasite (**Figures 6B,C**). SNs of PBMCs incubated with dead parasites did not upregulate CD69 on NK cells (**Figure 6D**). The activity of the SNs was also lost, when they were boiled before addition to the PBMCs. By contrast, filtration (pore size 0.22 µm) did not influence their activity excluding host cell or parasite debris as stimuli (**Figure 6E**). Together, these data indicate that the NK cell stimulating activity of the SNs presumably results from a heatlabile protein released by infected monocytes.

Next, we analyzed the spectrum of cytokines and chemokines secreted by *Leishmania*-activated monocyte/NK cell cultures of three different donors. We focused on *L. major*, because SNs from *L. major*-stimulated cell cultures were on average most potent in upregulating CD69 on NK cells. Using a Procarta® Multiplex Immunoassay, substantial amounts of several cytokines and chemokines were measured in the SNs of all three tested individuals (Table S2 in Supplementary Material). In further experiments, we concentrated on those factors that were strongly induced and had already been linked to NK cell activation (IL-1β, IL-6, IL-8, IL-18, and MIP-1α) (72–78). In addition, IL-2, IL-12, IL-15, IL-21, and IFN-α/β, all known as NK cell stimulatory cytokines (7), were included in the analysis. IL-12p40 protein was detected in low amounts in the SNs of *Leishmania*-stimulated PBMCs but was absent when monocytes had been depleted (**Figure 6F**),

monocytes after exposure to *Leishmania*. Purified CD14+ monocytes attached to an adhesion slide were incubated with or without *Leishmania major* promastigotes (multiplicity of infection 10) for 20 h. Thereafter, monocytes were either fixed with paraformaldehyde (Pfa) or fixed with Pfa and permeabilized with methanol, before being stained for IL-18 (white) and with DAPI (blue). As controls, the mouse-anti-IL-18 antibody (Ab) was pre-absorbed with rhuL-18, or the cells were incubated with the secondary Ab alone. Representative images of one of three independent experiments are shown.

indicating that it is released by *Leishmania*-triggered monocytes. The reason for the differential induction of IL-12p40 by the three *Leishmania* species (**Figure 6F**) tested is currently unknown, but is in line with previous observations that parasite species causing self-healing CL (*L. major*) elicit higher production of IL-12 as compared to parasite species associated with visceral disease (*L. donovani*) (79). Upregulation of CD69 on NK cells was clearly seen after stimulation of PBMCs with IL-2, IL-15, IL-18, or IFN-α/β, to a minor extent also with IL-1β and IL-6, but was not detectable following exposure of PBMCs to IL-8, IL-12, IL-21, or MIP-1α (**Figure 7A**). To determine whether one or several of the CD69-inducing cytokines represent the crucial NK cellstimulatory component within the SNs of PBMC/*Leishmania* cocultures, SNs were preincubated with neutralizing Abs or respective isotype controls. We did not observe any alteration in NK cell CD69 expression upon addition of individual neutralizing Abs or combinations thereof, although the stimulatory effect of the respective recombinant cytokine was clearly abrogated by the Ab treatment (**Figure 7B**). We conclude that none of the CD69-inducing cytokines (IL-1β, IL-2, IL-12, IL-15, IL-18, and

Messlinger et al. *Leishmania* and Human NK Cells

IFN-α/β) represents the soluble NK cell stimulating factor in supernatants of PBMC/*Leishmania* cocultures.

#### IL-12 Is Required to Elicit NK Cell IFN-**γ** Release in Response to *Leishmania* in PBMC or NK/Monocyte Cultures

Upregulation of CD69 is a first sign of NK cell activation but does not automatically entail the production of IFN-γ required for parasite control in mouse and human leishmaniasis (13). We therefore analyzed, whether IFN-γ was expressed by *Leishmania*stimulated PBMCs. IFN-γ was neither detectable in culture SNs by ELISA (**Figure 8A**) nor in NK cells by intracellular cytokine staining (**Figure 8B**), whereas stimulation with IL-12/IL-18 elicited a clear IFN-γ response of NK cells (**Figures 8A,B**). Likewise, IL-12/IL-18, but not exposure to *Leishmania* enhanced the cytotoxic activity of NK cells (**Figure 8C**). Thus, stimulation of PBMCs by *Leishmania* was not sufficient to induce NK cell effector functions. Interestingly, stimulation of PBMCs with IL-12/18 and *Leishmania* promastigotes further increased NK cell cytotoxicity as compared with cells activated by IL-12/IL-18 alone (**Figure 8C**).

To determine whether exogenously added IL-12 and IL-18 differentially contributed to the NK cell effector response, PBMCs were cocultured with parasites in the presence of either IL-12 or IL-18. Whereas IL-18 was largely ineffective (**Figure 9A**), IL-12 and *Leishmania*, but not IL-12 alone, triggered the release of IFNγ in PBMC cultures (**Figure 9B**). Titrating IL-12, a concentration of 300 pg/ml was sufficient to induce IFN-γ in the presence of *Leishmania* (**Figure 9C**). Blockade of IL-18 abolished the IL-12/*Leishmania*-induced IFN-γ response (**Figure 9D**).

In the mouse system, DCs are IL-12 producers during the early phase of *Leishmania* infection (18, 80). To investigate whether primary human DCs are capable to respond to *Leishmania* parasites by secreting IL-12, CD1c+ DCs of human PBMCs were purified by cell sorting and cultured in the presence of *Leishmania* promastigotes. After 20 h, an average of 800 (±116) pg/ml IL-12p40 (mean ± SEM of four donors) was detected in the SNs of the DC cultures (**Figure 10**). By contrast, CD14+ monocytes sorted in parallel did not release measurable amounts of IL-12p40 in response to *Leishmania* (**Figure 10**). Interestingly, when CD1c+ DCs were incubated together with parasites and sorted monocytes and NK cells, the 20 h culture SNs contained an increased concentration of IL-12p40 (1,485 ± 213 pg/ml, mean ± SEM of four donors). Also, in the presence of all three host cells and *L. major* parasites low amounts of bioactive IL-12p70 (*ca.* 90 pg/ ml) became detectable in two of four analyzed donors, whereas CD1c+ blood DCs or monocytes alone or cultures of monocytes and NK cells failed to generate IL-12p70 in response to *Leishmania* (data not shown).

Taken together, these data demonstrate that the absent NK cell effector response in PBMC/*Leishmania* cocultures is most likely due to an insufficient IL-12 production that presumably results from the low number of DCs in human PBMCs (68). Once IL-12 is added (or released by DCs after contact with NK cells and monocytes), endogenous IL-18 (produced and trans-presented by monocytes) acts synergistically with the IL-12 to elicit the expression of IFN-γ in NK cells.

concentration of interleukin (IL)-12p40 in culture SNs was determined by ELISA (values below detection limit are marked by triangles). Mean ± SEM of 6/6/5/5 (PBMC) and 6/5/5/6 (PBMC w/o monocytes) donors for the four stimulations. \*,#*p* < 0.05, \*\**p* < 0.01, and \*\*\**p* < 0.001 two-tailed Mann–Whitney *U* test.

#### Expression of CD56 on Human NK Cells Is Downregulated after Contact with *Leishmania*