Foam cell formation and caseation are hallmarks of human tuberculosis, yet their genesis, lipid composition, and significance for outcome remain poorly understood. Caseation could be due to cell death, resulting from mycobacterial toxicity and/or an exuberant host immune response, as well as altered lipid metabolism. Possible mechanisms include mobilization from lipid stores, endocytosis and deposition in macrophages, enhanced synthesis, locally and systemically, and/or failure to degrade accumulated lipids in macrophages. Macrophage foam cells form readily after uptake of various protein-bound lipids by a range of scavenger receptors, e.g., SR-A, CD36 and LDL receptors (Neyen and Gordon, 2014). Lipid droplets arise by synthesis and export from the endoplasmic reticulum to the cytosol, before becoming bound by vesicle membranes (**Figure 2**). Macrophage synthesis, uptake and exchange of lipids are regulated by transcription factors such as PPAR gamma and cytokines released by T lymphocytes and adipocytes (Daniel et al., 2011; Ouimet et al., 2016; Cambier et al., 2017). Foam cells in mouse granuloma models have been shown to express Wnt 6, involved in macrophage arginase1 induction and proliferation (Schaale et al., 2013). In addition to MTB-macrophage interactions, there seems to be an axis of interactions among MTB, macrophages and adipocytes, eg multinucleated macrophages are closely associated with adipocytes in white adipose tissue crown-like structures (Touton cells) and there is evidence that MTB can infect adipocytes (Neyrolles et al., 2006; Agarwal et al., 2016). Lipids derived from intracellular bacteria and/or plasma are potentially fusogenic, contributing in addition to host cytokines to Langhans giant cell formation. At a systemic level there is an association between TB and diabetes, cachexia is a characteristic feature of advanced TB, possibly mediated by TNF (originally named cachectin) and IL-6, and there is more than a superficial resemblance between caseation and atheroma, a hallmark of noninfective atherosclerosis. Recent studies have drawn attention to cholesterol (Huang et al., 2018) and oxysterol metabolism (Lu et al., 2017) in immune cells and of phenolic glycolipids in experimental mycobacterial infection (Cambier et al., 2017). Huang et al. have stressed the capacity of MTB to access and degrade fatty acids and sterols in order to survive within macrophages. To gain a better understanding of prolonged hostpathogen interactions in chronic TB, it will be necessary to perform lipidomic analysis of the heterogeneous macrophage populations as well as MTB, ideally, in vivo, and at the single cell level.

### CONCLUSIONS

Tuberculosis has been well documented as a clinical and pathological entity, yet many aspects of the disease remain unclear; while its rich natural history provides unique opportunities for fundamental and applied research. Since the nineteenth century it has been known that macrophages are an integral component of its pathogenesis and sequelae. In this review we have shown how cellular and molecular advances have generated new insights into the role of macrophages and, conversely how tuberculosis research continues to provide a wealth of theoretical and practical problems for study of macrophage immunobiology. Although we are beginning to understand the phenotypic distinction between tissue- resident yolk sac-derived and inflammatory bone marrow- derived macrophages in the mouse, this concept needs validation in humans; moreover, we do not know the impact of specific organ localization as well as origin on macrophage- MTB interactions. The extensive heterogeneity of macrophage populations in vivo, revealed by population and single cell analysis, mandates exquisite selectivity to target particular subsets of infected cells, without collateral injury to the host, as emphasized by Srivastava et al. (2014). Furthermore, the associated caseation and fibrosis, hallmark complications of the human disease, are poorly expressed in experimental models and in vitro; distinctions between the immune response to TB in humans and in animal models are emphasized by Scriba et al. (2017). Further progress will depend on access to human tissue from patients, the development of sensitive methods to study small amounts of tissue in situ, ideally by non-invasive methods over the course of the disease, and validation with in vitro models of human cells as well as development of new animal models that mimic the disease in humans. Shorter courses and more effective drug treatment of patients will be achieved by high throughput screening within macrophages to identify novel mycobacterial targets. The role of macrophages in granuloma and giant cell formation, epitheliodisation, caseation and fibrosis, all associated with TB, requires further investigation.

Study of tuberculous granulomas should promote understanding of other granulomatous diseases such as sarcoidosis and autoimmune destructive conditions such as Granulomatosis with polyangiitis, formerly called Wegener's granulomatosis. The co-epidemic of TB with HIV-1/AIDS, characteristically treated as a T cell failure, also involves macrophage dysfunction, as a viral reservoir, but especially when successful HAART restores CD4 T lymphocyte function, resulting in the Immune reconstitution inflammatory syndrome (IRIS) (Bell et al., 2017). Corticosteroid treatment, usually associated with increased risk in TB treatment, may be of benefit in preventing this outcome.

Apart from its remaining, but limited value as a vaccine, BCG offers a useful starting point to probe macrophage functions as an immunological adjuvant, in non-tuberculous intracellular infections and in selected cancer therapy. Recent benefits from BCG administration have been ascribed to imprinting haematopoietic stem cells in bone marrow to generate daughter macrophages with enhanced antimycobacterial effector function (Kaufmann et al., 2018). Dissection of the BCG genome should reveal new immunomodulatory prospects, without side effects resulting from unbridled macrophage activation. Perhaps most exciting will be to utilize such information in host-directed and combination therapy of drug-resistant Tuberculosis (Reiche et al., 2017), for example, targeting DNA replication and repair for the development of novel therapeutics against TB (Guler and Brombacher, 2015).

### AUTHOR CONTRIBUTIONS

SG conceived and wrote the manuscript. MM wrote the manuscript and produced the figure. FM wrote the manuscript. AP reviewed and revised the manuscript.

## FUNDING

Wellcome Trust (MM, 206751/Z/17/Z), SA Medical Research Council (SAMRC) SIR grant (MM), The SA National Research Foundation (MM), SAMRC with funding from SA Department of Health (MM).

# ACKNOWLEDGMENTS

We are grateful for stimulating discussions with colleagues.

### REFERENCES

antimicrobial activity of human macrophages. Sci. Transl. Med. 3:104ra102. doi: 10.1126/scitranslmed.3003045

intracellular burden of Mycobacterium tuberculosis in lymphatic endothelial cells. BMC Biol. 16:1. doi: 10.1186/s12915-017-0471-6

inflammatory response to mycobacterial infections. Cell 148, 434–446. doi: 10.1016/j.cell.2011.12.023

**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 Marakalala, Martinez, Plüddemann and Gordon. 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) and the copyright owner 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.

# The c-Type lectin receptor Dc-sign has an anti-inflammatory role in human M(il-4) Macrophages in response to *Mycobacterium tuberculosis*

*Geanncarlo Lugo-Villarino1,2,3\*† , Anthony Troegeler1,2,3†, Luciana Balboa2,3,4, Claire Lastrucci 1,2,3,5, Carine Duval1 , Ingrid Mercier1 , Alan Bénard1,6, Florence Capilla7 , Talal Al Saati 7 , Renaud Poincloux1 , Ivanela Kondova8 , Frank A. W. Verreck8 , Céline Cougoule1,2,3, Isabelle Maridonneau-Parini 1,2,3, Maria del Carmen Sasiain2,3,4 and Olivier Neyrolles1,2,3*

*1 Institut de Pharmacologie et de Biologie Structurale, IPBS, Université de Toulouse, CNRS, UPS, Toulouse, France, 2 International Associated Laboratory (LIA) CNRS "IM–TB/HIV" (1167), Toulouse, France, 3 International Associated Laboratory (LIA) CNRS "IM–TB/HIV" (1167), Buenos Aires, Argentina, 4 IMEX-CONICET, Academia Nacional de Medicina, Buenos Aires, Argentina, 5Centre for Genomic Regulation, Barcelona, Spain, 6Department of Surgery, University, Hospital Erlangen, Friedrich-Alexander, University Erlangen-Nürnberg, Erlangen, Germany, 7 INSERM/UPS/US006 CREFRE, CHU Purpan, Toulouse, France, 8Biomedical Primate Research Centre, Rijswijk, Netherlands*

#### *Edited by:*

*Steffen Stenger, Universitätsklinikum Ulm, Germany*

#### *Reviewed by:*

*Hridayesh Prakash, All India Institute of Medical Sciences, India Masaaki Miyazawa, Kindai University, Japan*

#### *\*Correspondence:*

*Geanncarlo Lugo-Villarino<EMAIL_ADDRESS>
*†These authors have contributed equally to this work.*

#### *Specialty section:*

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

*Received: 06 December 2017 Accepted: 03 May 2018 Published: 12 June 2018*

#### *Citation:*

*Lugo-Villarino G, Troegeler A, Balboa L, Lastrucci C, Duval C, Mercier I, Bénard A, Capilla F, Al Saati T, Poincloux R, Kondova I, Verreck FAW, Cougoule C, Maridonneau-Parini I, Sasiain MdC and Neyrolles O (2018) The C-Type Lectin Receptor DC-SIGN Has an Anti-Inflammatory Role in Human M(IL-4) Macrophages in Response to Mycobacterium tuberculosis. Front. Immunol. 9:1123. doi: 10.3389/fimmu.2018.01123*

DC-SIGN (CD209/CLEC4L) is a C-type lectin receptor (CLR) that serves as a reliable cell-surface marker of interleukin 4 (IL-4)-activated human macrophages [M(IL-4)], which historically represent the most studied subset within the M2 spectrum of macrophage activation. Although DC-SIGN plays important roles in *Mycobacterium tuberculosis* (Mtb) interactions with dendritic cells, its contribution to the Mtb–macrophage interaction remains poorly understood. Since high levels of IL-4 are correlated with tuberculosis (TB) susceptibility and progression, we investigated the role of DC-SIGN in M(IL-4) macrophages in the TB context. First, we demonstrate that DC-SIGN expression is present both in CD68+ macrophages found in tuberculous pulmonary lesions of non-human primates, and in the CD14+ cell population isolated from pleural effusions obtained from TB patients (TB-PE). Likewise, we show that DC-SIGN expression is accentuated in M(IL-4) macrophages derived from peripheral blood CD14+ monocytes isolated from TB patients, or in macrophages stimulated with acellular TB-PE, arguing for the pertinence of DC-SIGN-expressing macrophages in TB. Second, using a siRNA-mediated gene silencing approach, we performed a transcriptomic analysis of DC-SIGN-depleted M(IL-4) macrophages and revealed the upregulation of pro-inflammatory signals in response to challenge with Mtb, as compared to control cells. This pro-inflammatory gene signature was confirmed by RT-qPCR, cytokine/chemokine-based protein array, and ELISA analyses. We also found that inactivation of DC-SIGN renders M(IL-4) macrophages less permissive to Mtb intracellular growth compared to control cells, despite the equal level of bacteria uptake. Last, at the molecular level, we show that DC-SIGN interferes negatively with the pro-inflammatory response and control of Mtb intracellular growth mediated by another CLR, Dectin-1 (CLEC7A). Collectively, this study highlights a dual role for DC-SIGN as, on the one hand, being a host factor granting advantage for Mtb to parasitize macrophages and, on the other hand, representing a molecular switch to turn off the pro-inflammatory response in these cells to prevent potential immunopathology associated to TB.

Keywords: macrophages, *Mycobacterium tuberculosis*, DC-SIGN, C-type lectin receptors, anti-inflammatory

## INTRODUCTION

According to the latest World Health Organization (WHO) report, tuberculosis (TB) is the largest killer among communicable diseases (WHO Annual report 2017). In 2016, there were an estimated 1.7 million deaths due to TB, making it the leading cause of death worldwide due to a single infectious agent, *Mycobacterium tuberculosis* (Mtb). In general, it is estimated that one quarter of the human population could be latently infected with Mtb (1). The bacillus may be active either after infection or through the reactivation of latent infection, which occurs in approximately 5% of infected people. During latency, for which there are no pathological or contagious conditions, Mtb is contained within elaborated aggregates of immune cells that are called granulomas, the hallmark of TB (2, 3). It is thought that a dedicated immune response is responsible for the formation and maintenance of granulomas, which will ultimately determine the outcome of the disease (2, 4). However, there is a strong need to better understand the factors that define an efficient immune response both during the early and late phases of Mtb infection in order to facilitate potential targets for preventive and therapeutic purposes.

Macrophages are considered key players during the early and late stages of Mtb infection (5). These sentinel cells are strategically located in secondary lymphoid organs and multiple mucosal sites, such as lung alveolar and interstitial space. At such, macrophages recognize and internalize Mtb and, consequently, modulate the inflammatory response to shape their microenvironment (e.g., granulomas) and the adaptive immune response against this pathogen. Interestingly, these cells display a high degree of tissue heterogeneity within the broad spectrum of pro- (M1) and anti-inflammatory (M2) programs of activation that manifest intracellular pathogen resistance and permissiveness, respectively (6). Macrophages may also serve as long-lived pathogen tissue reservoirs and contribute to TB pathogenesis (6–9). Remarkably, Mtb influences the differentiation, maturation, and activation of macrophages, resulting in the circumvention of the immune system and augmented persistence in the host (6–8, 10). This capacity of Mtb to modulate the host pro-inflammatory response and seize the anti-inflammatory mechanisms has generated a keen interest to investigate how this pathogen manipulates the process of macrophage activation.

The initial interaction with Mtb is thought to be crucial for macrophage activation and the eventual disease outcome. Pattern recognition receptors (PRRs) expressed in macrophages determine the binding, internalization, and fate of the bacillus' intracellular lifestyle. Among the various PRR families that recognize Mtb, the C-type lectin receptors (CLR) are known to contribute to the control or persistence of this pathogen within macrophages (11–13). The CLR family includes collectins, selectins, endocytic and phagocytic receptors, and proteoglycans. CLRs are calciumdependent glycan-binding proteins exhibiting similarities in the structures of the carbohydrate-recognition domain (CRD), which in turn recognize the carbohydrates expressed on the surface of Mtb including glycolipids [e.g., phosphatidyl-*myo*-inositol mannoside (PIM)], glycoglycans [e.g., lipoarabinomannan (LAM)], polysaccharides (e.g., α-glucan) and glycoproteins (e.g.,19 kDa antigen). In recent years, our understanding of the interactions of mycobacterial ligands with CLRs has advanced considerably, specifically in membrane-anchored phagocytic receptors such as the mannose receptor MRC1 (CD206/CLEC13D), Dectin-1 (CLEC7A) and Dectin-2 (CLEC6A), Mincle (CLEC4E), and Dendritic Cell-Specific Intercellular adhesion molecule-3-Grabbing Non-integrin (DC-SIGN/CD209/CLEC4L) (11–15). In general terms, the activation of these receptors by Mtb leads to downstream effects like endocytosis, oligomerization, intracellular trafficking, and signal transduction. Emerging evidence points out that signaling pathways triggered by these CLRs converge in a limited set of synergistic or antagonistic interactions with other PRRs and with each other, giving rise to a phenomenon known as signaling crosstalk. Interestingly, Mtb has evolved the capacity to subvert CLR signaling crosstalk to increase its survival and fitness within macrophages (5, 11, 15).

Studies by others and us identified and characterized DC-SIGN as a key receptor for Mtb in human dendritic cells and alveolar macrophages (14, 16–20). This CLR is a transmembrane receptor that possesses one single extracellular CRD (at the C-terminus) capable of recognizing mannose-containing molecules such as those present in mycobacterial Man-LAM (mannose-capped LAM), lipomannan (LM), arabinomannan, glycoproteins (e.g., 19, 38, and 45 kDa antigens), PIMs, and α-glucan, among others. Its functional cytoplasmic domain (at the N-terminus) contains different motifs that are crucial for endocytosis/phagocytosis, intracellular trafficking, and signal transduction (11). In the case of Mtb, it was shown that DC-SIGN is targeted by the mycobacterial ManLAM to induce the immunosupressive mediator interleukin-10 (IL-10) and counteract the toll-like receptor-4 (TLR-4)-dependent pro-inflammatory response (14). In this manner, the bacillus prevents the proper activation of dendritic cells given that IL-10 inhibits the expression of co-stimulatory molecules (e.g., CD86) and production of IL-12, which are essential for the activation of type-1 immunity best represented by T-helper 1 (Th1) cells. Of note, the hijacking of IL-10 production *via* DC-SIGN in dendritic cells seems to be a general evasion strategy by various pathogens like *Mycobacterium leprae*, *Helicobacter pylori,* and *Candida albicans*, among others (14).

Until recently, the expression of DC-SIGN was thought to be exclusive to human dendritic cells. In fact, this CLR is also expressed in human alveolar macrophages and some lymphocyte populations (19, 21). In particular, DC-SIGN is a reliable cellsurface marker in human macrophages activated with interleukin 4 (IL-4) [M(IL-4)], which is historically the most representative subset within the M2 spectrum of macrophage activation (22, 23). While the role of M(IL-4) macrophages has not been explored in in the TB context, high levels of type-2 inflammatory signals, such as IL-4, are correlated to TB susceptibility and progression (10). Reciprocally, the predominant type-2 inflammatory environment shifts toward type-1 immune signals [e.g., interferon-γ (IFNγ)] upon successful treatment of pulmonary TB. This is important because, in comparison to IFNγ, IL-4 renders macrophages less microbicidal against intracellular bacterial infections (6, 24). Furthermore, our group has also reported that DC-SIGN expression is induced specifically in alveolar macrophages of patients with active TB (19), suggesting a major role for this CLR in the interaction between macrophages and Mtb.

In this study, we provide for the first-time evidence supporting the anti-inflammatory role of DC-SIGN in the M(IL-4) macrophage response to Mtb. We report that DC-SIGN expression is accentuated in macrophages under different contexts using samples from TB patients, or in tuberculous pulmonary lesions of non-human primates (NHP), arguing for the pertinence of DC-SIGN-expressing macrophages in TB pathology. In the absence of this CLR, M(IL-4) macrophages displayed a proinflammatory signature upon challenge with Mtb, and acquired a better ability to control the intracellular growth of this pathogen. Finally, while there are no major changes in the production of IL-10, we demonstrate that DC-SIGN interferes negatively with the activation of M(IL-4) macrophages triggered by Dectin-1.

#### MATERIALS AND METHODS

#### Ethics Statement for Non-Human Primate Samples

The NHP study protocol was done to comply with the EC Directive 86/609/EEC, approved by the local independent ethics committee prior to the start of the study, and executed under Dutch law on animal experiments (agreement number DEC#579). The endpoint for any particular animal was based either by signs of severe disease (humane endpoint criteria, referring to animal condition by adverse body weight development, respiratory capacity, and animal behavior) or by protocol, which limited the follow-up time to 1-year postinfection.

#### Non-Human Primate Handling

The NHP samples were prepared from animals that were used for vaccine research and development purposes, as previously described (25). Briefly, healthy young adult female rhesus macaques (*Macaca mulatta*), all captive-bred for research purposes and of homogeneous breeding background, were challenged with 500 colony forming units (CFU) of Mtb strain Erdman K01 (prepared and provided under an agreement between WHO and CBER/FDA with assistance of Aeras), which was administered by intra-bronchial instillation under sedation. At endpoint, the animals were sedated, euthanized and submitted to macroscopic lung pathology scoring, as previously published (26).

#### Histological Analyses on Non-Human Primate Samples

The gross pathological findings were assessed and described by an experienced veterinary pathologist while blinded for treatment as previously described (25). Representative lung biopsies were collected and fixed in 10% neutral buffered formalin and embedded in paraffin for long-term storage. Tissue sections were stained with hematoxylin and eosin (HE) for histomorphological analysis. Histopathological scoring of TB lesions in NHP was determined using a worksheet in which TB disease from lung biopsies was described (25, 26). Immunohistochemical staining was performed on paraffin-embedded tissue sections using the following monoclonal primary antibodies: CD68 (clone:KP1, Dako), DC-SIGN (clone 120612, R&D System), CD163 (clone 10D6, Leica/Novocastra), and MerTK (clone: Y323, Abcam). After incubation with primary antibodies, sections were stained with biotin-conjugated polyclonal anti-mouse or -rabbit immunoglobulin antibodies followed by the streptavidin-biotinperoxidase complex (ABC) method (Vector Laboratories), and then were counterstained with hematoxylin. Slides were scanned with the Panoramic 250 Flash II (3DHISTECH). For confocal microscopy, samples were stained with primary antibodies as described above and followed by anti-mouse IgG isotype specific or anti-rabbit IgG antibodies labeled with Alexa488 and Alexa555 (Molecular Probes). Samples were mounted with Prolong® Antifade reagent (Molecular Probes) and examined using a 40×/0.95N.A. objective of an Olympus FV1000 confocal microscope.

#### Ethics Statement for Human Samples

Blood samples from healthy subjects (HS) or TB patients were provided by the Blood Transfusion Service, Hospital Fernandez, Buenos Aires (agreement number CEIANM-52-5-2012), or the Hospital F. J. Muñiz, Buenos Aires (protocol number: NIN-1671- 12). Pleural effusions (PE) were obtained by therapeutic thoracentesis by physicians at the Hospital F. J. Muñiz (Buenos Aires). The research was carried out in accordance with the Declaration of Helsinki (2013) of the World Medical Association and was approved by the Ethics Committees of the Hospital F. J. Muñiz and the Academia Nacional de Medicina de Buenos Aires (protocol number: NIN-1671-12). Written informed consent was obtained before sample collection. The diagnosis of TB pleurisy was based on a positive Ziehl–Nielsen staining or Lowestein–Jensen culture from PE and/or histopathology of pleural biopsy and was further confirmed by an Mtb-induced IFN-γ response and an ADApositive test (27). Mononuclear cells from peripheral blood (PB) and PE were isolated by Ficoll-Hypaque gradient centrifugation (Pharmacia, Uppsala, Sweden), as described previously (25, 28).

Likewise, monocytes from HS were isolated from buffy coat provided by Etablissement Français du Sang, Toulouse, under contract 21/PLER/TOU/IPBS01/2013-0042. According to articles L1243-4 and R1243-61 of the French Public Health Code, the contract was approved by the French Ministry of Science and Technology (agreement number AC 2009-921). Donors signed and provided written informed consents before sample collection.

### Preparation of Pool of Sera and Tuberculous PE

The cell-free supernatant from tuberculous PE and sera was transferred into new plastic tubes, further centrifuged at 12,000 *g* for 10 min and aliquots were stored at −80°C. Pools were prepared by mixing same amounts of eight individual PE or serum. The pools were de-complemented at 56°C for 30 min, and filtered by 0.22 µm in order to remove any remaining debris or residual bacteria.

#### Preparation of Human Monocyte-Derived Macrophages From HS and TB Patients

Monocytes from HS or TB patients were isolated and differentiated into macrophages as previously described (25, 28). Briefly, purified CD14+ monocytes from HS were differentiated for 5–7 days in RPMI-1640 medium (GIBCO), 10% fetal bovine serum (FBS, Sigma-Aldrich), and human recombinant macrophage colony-stimulating factor (M-CSF, Peprotech) at 10 ng/ mL. The cell medium was renewed every 3 or 4 days. Thereafter, macrophages were treated with IL-4 (Peprotech) at 20 ng/mL to induce the M(IL-4) program, 10% v/v of a pool of sera from HS or TB patients, or of acellular fraction of TB PE for 48 h. Untreated cells under differentiation with M-CSF only were considered as part of the M(M-CSF) program.

Alternatively, monocyte isolation and purification was done as previously published (29). Succinctly, following Ficoll gradient enrichment, monocytes were purified using positive selection with anti-CD14 microbeads and MACS separation columns (Miltenyi Biotec), according to manufacturer's instructions. For macrophage differentiation, monocytes were allowed to adhere to glass coverslips (VWR international) in 6-well or 24-well plates (Thermo Scientific), at 1.5 × 106 and 3 × 105 cells/well, respectively, for 2 h at 37°C in warm RPMI-1640 medium (GIBCO). The medium was then supplemented to a final concentration of 10% FBS (Sigma-Aldrich) and human recombinant M-CSF (Peprotech) at 20 ng/ml, and the cells were allowed to differentiate for 5 days. The cell medium was renewed at day 3 or 4 of culture. At day 5, macrophages were activated for 48 h with 1 µg/ml of lipopolysaccharide (LPS, Invivogen) and 2.5 ng/ml of IFN-γ (Miltenyi Biotec) to induce the M(LPS + IFN-γ) program, and with 20 ng/ml of IL-4 (Miltenyi Biotec) for the M(IL-4) program.

#### Flow Cytometry

Cells from TB patients and related controls from HS (2 × 105 cells) were labeled as described above and acquired in a FACSAria II cytometer (BD Biosciences). The monocyte–macrophage population was gated according to its forward scatter (FSC) and size scatter (SSC) proprieties. The percentage of positive cells and the median fluorescence intensity (MFI) were analyzed using FCS Express V3 software (De Novo Software, Los Angeles, CA, USA). In the context of macrophage activation programs, the CLR expression was analyzed by flow cytometry using antibodies (general dilution of 1:400) against DC-SIGN and Dectin-1 from R&D Systems, and MRC1 from BD Pharmingen (San Diego, CA, USA). Concerning the cellular content in PE fluid from TB patients, DC-SIGN expression was also analyzed by flow cytometry in mononuclear cells, gating within the CD14+ population and using human anti-DC-SIGN (R&D Systems).

Alternatively, the antibody staining of macrophages was performed as previously described (29). Adherent cells were collected using the Cell Dissociation Buffer according to manufacturer's instructions (Life Technologies), centrifuged for 5 min at 340 *g* at 4°C, and then stained in cold FACS buffer (PBS pH 7.2, 5% BSA) for 25 min with the indicated fluorophore-conjugated antibodies using a general dilution of 1:400. The antibodies were the following: CD16 (FCGR3A), PD-L1 (CD274), CD11b (MAC-1) and CD14 from Biolegend; CD163 (SCARI1), CD86 (B7-2), CD64 (FCGR1A), CD36 (SCARB3), CD11c (ITGAX), IL-7R (CD127), MRC1, and DC-SIGN from BD Biosciences; MerTK (Tyro12) from R&D system, and HLA-DR from Santa Cruz Biotechnology. In parallel, we also performed the staining with the corresponding isotype control antibody. Afterward, the cells were washed with cold FACS buffer, centrifuged for 5 min at 340 *g* at 4°C, and analyzed by flow cytometry using LSR-II flow cytometer (BD Biosciences). Data was then acquired and analyzed using the FlowJo 7.6.5 software.

#### siRNA Silencing of DC-SIGN and Dectin-1

The siRNA gene silencing in human primary macrophages was performed using the forward transfection approach, as previously described (29). Briefly, macrophages differentiated at day 5 were transfected using the lipid-based HiPerfect system (Qiagen) and an ON-TARGETplus SMARTpool siRNA targeting DC-SIGN (siDC-SIGN) and Dectin-1 (siDectin-1), or a non-targeting siRNA (siControl) (Dharmacon), at a final concentration of siRNA at 200 nM. Of note, for the simultaneous inactivation of two genes, the working siRNA concentration combined for DC-SIGN and Dectin-1 did not exceed the final concentration of 200 nM, as previously published (29, 30). After 6 h, RPMI-1640 medium supplemented with M-CSF (10 ng/ml) was added to each well, and the cells were allowed to recuperate overnight. The following day, IL-4 (20 ng/ml) was added to the transfected macrophages in order to induce the M(IL-4) program along with the CLR expression for an additional 72 h (or as indicated). The inactivation for DC-SIGN and Dectin-1 was confirmed by flow cytometry in non-permeabilized cells at the indicated time points post-transfection. Cell viability was determined by using the Annexin-V-FITC kit designed for flow cytometry. Cell viability was defined as cells negative for either Annexin-V-FITC and/or propidium iodide. For the upregulation of co-stimulatory molecules, control and DC-SIGN-depleted M(IL-4) macrophages were stimulated with LPS (1 µg/ml) for 24 h, and the cell-surface marker expression was assessed by flow cytometry.

### Blocking and Stimulation of DC-SIGN and Dectin-1 Receptors

Human macrophages were differentiated at day 5, and activated with IL-4 (20 ng/ml) for additional 48 h. At day 7, the M(IL-4) macrophages were washed with warmed RPMI-1640 medium supplemented with M-CSF (10 ng/ml) and pre-incubated for 30 min with 10 µg/ml blocking antibodies specific to either Dectin-1 (MAB1859, R&D Systems) or DC-SIGN (ab13847, abcam), or both. As a control, M(IL-4) macrophages were incubated with an irrevelevant antibody (Ab-Control). For efficient stimulation of Dectin-1, M(IL-4) macrophages were treated with cytochalasin D (1 µg/ml, C8273, SIGMA) in combination with purified β-glucan from Saccharomyces cerevisiae (10 µg/ml, G5011, SIGMA), as previously described (31); for DC-SIGN, cells were stimulated with 10 µg/ml of ManLAM (kindly provided by Dr. Jerome Nigou, IPBS/CNRS); and for TLR-4, cells were stimulated with 1 µg/ml of LPS. After 24 h, the supernatants were collected and stored at −80°C until further use for ELISA analysis.

### Mtb Strain, Culture, and Preparation for Infection

All manipulation with Mtb (H37Rv strain) was performed in a dedicated BSL-3 laboratory. Mtb was cultured at 37°C in Middlebrook 7H9 medium (Difco) supplemented with 10% albumin-dextrose-catalase (Difco) and 0.05% Tween-80 (Sigma-Aldrich). For infection, exponentially growing Mtb was centrifuged (2,000 *g*) for 15 min, and resuspended in 1x phosphate buffered saline (PBS). Clumps were dissociated by passages through a 26-G needle, and then resuspended in RPMI-1640 containing 10% FBS. The mycobacterial concentration was determined by measuring optical density at 600 nm [OD600]. For binding experiments, the GFP-expressing Mtb (H37Rv) strain was generated and cultivated as previously published (20).

#### RNA Extraction and Transcriptomic Analysis

Control and DC-SIGN-depleted M(IL-4) macrophages (approximately 1.5 million cells) were infected with Mtb at a multiplicity of infection (MOI) of 3 bacteria to 1 cell in RPMI-1640 with 10% FBS for 4 h. The cells were then washed twice with 1x PBS. At this point, the cells were either treated with TRIzol Reagent (Invitrogen) to harvest at 4 h postinfection (*p.i*.) and stored at −80°C, or cultured with RPMI-1640 with 10% FBS overnight. The procedure with the TRIzol reagent was repeated the following day to harvest at 18 h *p.i*. and stored at −80°C. Total RNA was extracted from the TRIzol samples using the RNeasy mini kit (Qiagen). The amount and purity of RNA (absorbance at 260/280 nm) was measured with the Nanodrop ND-1000 apparatus (Thermo Scientific). Complementary DNA was reverse transcribed from 1 µg total RNA with Moloney murine leukemia virus reverse transcriptase (Invitrogen) using random hexamer oligonucleotides for priming, according to the manufacturer's protocol. The microarray analysis was done using the Agilent Human GE 4 × 44 v2 (single color), as previously described (32). Briefly, the hybridization was performed with 2 µg Cy3-cDNA and the hybridization kit (Roche NimbleGen). According to manufacturer's protocol, the samples were incubated for 5 min at 65°C, and 5 min at 42°C before loading for 17 h at 42°C. After washing, the microarrays were scanned with MS200 microarray scanner (Roche NimbleGen). Using Feature Extraction software, the Agilent raw files were extracted and then processed through Bioconductor (version 3.1) in the R statistical environment (version 3.2.0). A careful assessment of the quality of the hybridization, evaluation of the sampling method and normalization of the expression values was done as previously published (32). We then obtained the differentially expressed genes (DEGs) between control and DC-SIGN-depleted M(IL-4) macrophages at each time point after Mtb infection based on false discovery rate (*t*-test, *P* < 0.1) and a threshold of twofold change in the comparison between the two conditions. The normalized values for the entire microarray analysis and the determined DEGs are provided in Tables S1–S3 in Supplementary Material. Finally, DEGs were analyzed with QIAGEN's Ingenuity® Pathway Analysis (IPA®, QIAGEN Redwood City, www.qiagen.com/ ingenuity).

#### qRT-PCR Analysis

The amplification of RNA was performed with an ABI Prism 7500 Sequence Detector (Applied Biosystems) using the PCR SYBR Green sequence detection system (Eurogentec, Seraing, Belgium). Primers are listed in Table S4 in Supplementary Material. Data were analyzed using the software supplied with the Sequence Detector (Applied Biosystems). The mRNA content was normalized to the metastatic lymph node protein 51 (MLN51) mRNA and quantified using the ΔΔCt method.

#### Semi-Quantitative Cytokine/Chemokine Antibody Array Assay and Quantitative ELISA

Control and DC-SIGN-depleted M(IL-4) macrophages were infected with Mtb at MOI of 3. Supernatants were harvested at 4 and 18 h p.i., centrifuged (2,000 *g*) for 10 min, passed through filters (0.22 µm pores) and stored at −80°C. With the use of the Human Cytokine Antibody Array C Series 1000 (RayBiotech), we assessed the supernatant content in terms of cytokine and chemokines following the manufacturer's instructions, as previously described (33). We used Amersham Hyperfilm ECL (GE Healthcare) to detect individual signals, and the GS-800 calibrated densitometer (Bio-Rad) to quantify these signals. As instructed, the positive control signal on each array was used to normalize the rest of the detected signals. In parallel, cytokine quantification was measured in cell supernatants by ELISA using kits from BD Bioscience (TNFα, IL-6, and IL-10), according to manufacturer's instructions.

#### Binding and Phagocytosis of Mtb

As previously described (19), control and DC-SIGN-depleted M(IL-4) macrophages were infected with the GFP-expressing Mtb strain, at a MOI of 5 for 4 h, at either 4°C or 37°C, in order to assess binding and phagocytosis, respectively. Cells were then washed with 1× PBS (without calcium and magnesium), collected using the Cell Dissociation Buffer, centrifuged for 5 min at 340 *g* at 4°C, fixed with 4% PFA for 2 h, washed with 1× PBS and resuspended in FACS buffer. Cells were then analyzed by flow cytometry.

#### Measurement of Mtb Intracellular Growth

Macrophages were washed with PBS and then infected with Mtb at a MOI of 0.2 bacteria/cell in RPMI-1640 with 10% FBS for 4 h. Cells were then washed twice with 1x PBS before addition of RPMI-1640/10% FBS. At the indicated time points, the cells were lysed in 0.01% Triton X-100 (Sigma-Aldrich), and serial dilutions of the lysates were plated onto 7H11-Oleic Albumin Dextrose Catalase (Difco) agar medium for CFU scoring.

#### Statistical Analyses

Two-tailed Wilcoxon (matched-paired test) was applied to compare the M(IL-4) macrophage population under two conditions (siControl versus siDC-SIGN). Two-tailed unpaired *t*-test (parametric) was applied on data sets with a normal distribution, whereas one-tailed unpaired Mann–Whitney (nonparametric) was done for data not showing a normal distribution and where the outcome was already expected, as indicated for each figure legend. *P* < 0.05 was considered as the level of statistical significance.

# RESULTS

### DC-SIGN-Expressing Macrophages Are Present in Pulmonary Lesions of NHP Infected With Mtb

Tuberculous granulomas have organized microenvironments that presumably balance the antimicrobial functions to control bacteria growth and anti-inflammatory properties to limit pathology in the lung (34). Recently, others and we have detected the high abundance of an M2-like macrophage population in tuberculous granulomas in the context of pulmonary lesions of NHPs with severe TB (25, 35). Here, we further investigated whether DC-SIGN-expressing macrophages are present in these samples derived from Mtb-infected rhesus macaques, which displayed different levels of lung pathology. In samples from NHPs exhibiting a low pathological score in lungs, immunohistochemical analyses revealed the low presence of CD68+ macrophages and DC-SIGN-expressing cells in areas where leukocyte infiltrate was also detected by HE staining (**Figure 1A**). By contrast, in samples from animals characterized by a high lung pathological score, we observed not only the characteristic necrotic lung granuloma and CD68+ macrophages abundantly infiltrating the interstitial lung tissue, but also the increased number of DC-SIGN-expressing cells in these areas (**Figure 1A**). To determine whether DC-SIGN is expressed by M2 macrophages in the context of tuberculous granulomas, we also performed an additional staining for CD163 and MerTK, two strong markers of M2 macrophages. As shown for **Figure 1B**, DC-SIGN-expressing cells localized at the granuloma periphery and alveoli, where CD163+ and MerTK+ macrophages are also found. Importantly, co-localization analysis revealed the presence of cells positive for DC-SIGN within CD68+ (41 ± 20%, *n* = 893 counted cells) and CD163+ (27 ± 3%, *n* = 754 counted cells) macrophage populations (**Figure 1C**). These analyses demonstrate that the environment generated during pulmonary TB is associated with enhanced DC-SIGN expression in CD68+ and CD163+ macrophages, which become accentuated in NHPs with severe TB as previously described (25).

### Human Tuberculosis-Associated Microenvironment Induces DC-SIGN Expression in Macrophages

Our group previously reported that, in lungs, DC-SIGN expression is induced specifically in alveolar macrophages of patients with active TB (19). Using samples obtained from patients with active TB, we expanded our assessment of DC-SIGN expression in the context of macrophage activation. First, we isolated and differentiated monocytes from HS or TB patients into macrophages. Macrophages were activated toward the M(IL-4) program and the expression of DC-SIGN was measured by flow cytometry analysis. Our results revealed that, unlike unactivated macrophages [M(M-CSF)], DC-SIGN expression was accentuated in M(IL-4) macrophages and with a higher tendency (albeit not significant) in TB patients (**Figure 2A**). Second, we induced the activation of macrophages (derived

row) and DC-SIGN (bottom row), among areas where leukocyte infiltration (top row) is detected by hematoxylin and eosin (HE), in pulmonary tissue and granulomas of NHP with very mild (left columns) and severe (right columns) pathology. (B) Representative immunohistochemical images illustrating the distribution of DC-SIGN, CD163, and MerTK in areas where leukocyte infiltration is detected by HE, such as in granulomas of NHP with severe pathology. (C) *Left panel*: immunostaining of DC-SIGN (green: Alexa-488) and CD68 (red: Alexa-555) in lung tissue of NHP with severe pathology; *right panel*: immunostaining of DC-SIGN (green: Alexa-488) and CD163 (red: Alexa-555) in lung tissue of NHP with severe pathology. Green arrow points out a cell positive for DC-SIGN only; red arrow a cell positive for CD68 or CD163 only; and yellow arrow for a cell positive for both DC-SIGN and CD68/CD163. Scale bar = 10 µm.

from HS monocytes) with cell-free supernatants from sera or PE from TB patients (TB-S and TB-PE, respectively). We noticed that while sera from either HS or TB patients failed to induce DC-SIGN expression, TB-PE significantly increased the cell-surface level for this CLR (**Figure 2B**). Last, we measured

Figure 2 | Human tuberculosis-associated microenvironment induces DC-SIGN expression in macrophages. (A) DC-SIGN expression is induced in M(IL-4) macrophages from TB patients. Freshly isolated monocytes from healthy subjects (HS, white) and TB patients (TB, black) were differentiated into macrophages using M-CSF. At day 5, the cells were activated with IL-4 (20 ng/ ml) for 48 h to induce the M(IL-4) macrophage program. Otherwise, macrophages were kept under M-CSF to fully establish the M(M-CSF) program. The cells were harvested and the DC-SIGN expression was analyzed by flow cytometry. Vertical bar graphs depicting the median fluorescent intensity (MFI) of DC-SIGN expression in the different cell populations. Results are expressed as mean ± SD (*n* = 10 donors). (B) DC-SIGN expression is induced by pleural fluid from TB patients. Freshly isolated monocytes from HS were differentiated into macrophages using M-CSF. At day 5, the cells were activated for 48 h with sera from HS (HS-S, black) and TB patients (TB-S, gray), acellular pleural fluid from TB patients (TB-PE, white), and IL-4 (20 ng/ml, vertical stripes). The cells were harvested and the DC-SIGN expression was analyzed by flow cytometry. Vertical bar graphs depicting the MFI of DC-SIGN expression in the different cell populations. Results are expressed as mean ± SD (*n* = 13 donors). (C) DC-SIGN-expressing macrophages are present in the pleural cavity of TB patients. Mononuclear cells were isolated either from peripheral blood from HS (HS-PB) and TB patients (TB-PB), or from the pleural effusions from TB patients (TB-PE), and the expression of DC-SIGN was analyzed on the CD14+ population by flow cytometry. Results are expressed as vertical scatter plots showing the MFI of DC-SIGN for each population; each individual symbol represents a single donor. Two-tailed *t*-test (unpaired/parametric): *\*P* < 0.05, \*\**P < 0.01, \*\*\*P* < 0.001, \*\*\*\**P* < 0.0001, NS = not significant*.*

DC-SIGN expression directly in the CD14+ cell population present in the circulation or PE of TB patients. We failed to observe the presence of DC-SIGN-expressing CD14+ blood monocytes in either HS or TB patients. Strikingly, however, we detected DC-SIGN highly expressed in all CD14+ monocyte/ macrophages collected in TB-PE (**Figure 2C**). Altogether, these results demonstrate that TB-associated environments are capable to induce DC-SIGN expression in the context of human macrophage activation.

### Depletion of DC-SIGN Expression Does Not Affect Cell Viability nor the Establishment of the M(IL-4) Macrophage Program

In the context of the most commonly studied *in vitro* programs of human macrophage activation [i.e., M(LPS + IFNγ) and M(IL-4)], DC-SIGN is a reliable cell-surface marker for M(IL-4) macrophages (22, 23). To confirm this, we differentiated freshly isolated CD14+ monocytes from healthy donors into macrophages. We activated macrophages toward M(IL-4) and M(LPS + IFNγ), and then assessed the cell-surface marker profile of these cell populations by flow cytometry. While we noticed that expression of DC-SIGN and other markers varies dramatically between donors (in terms of MFI), we still detected a significantly higher cell-surface expression of DC-SIGN in M(IL-4) in comparison to M(LPS + IFNγ) macrophages (Figure S1A in Supplementary Material). Its expression was accompanied by that of Dectin-1 and MRC1, a *bona fide* M(IL-4) macrophage marker (22). We also observed a higher expression (albeit not significant) of CD11b and CD36 in M(IL-4) compared to M(LPS + IFNγ) cells (Figure S1A in Supplementary Material). By contrast, M(LPS + IFNγ) were distinguished by significant elevated levels of CD64, CD86, and PD-L1 (Figure S1A in Supplementary Material). These results confirm that DC-SIGN is exclusively expressed in M(IL-4) macrophages, thus making it an ideal cell model to study the role of this CLR in human macrophages.

With the aim to examine the role of DC-SIGN in the activation of the M(IL-4) program, we made use of a lipid-based siRNAmediated gene silencing protocol we recently developed (29, 30). Macrophages were transfected with siRNA targeting this CLR (siDC-SIGN) or with non-targeting scrambled siRNA control (siControl), and these cells were then stimulated with IL-4 to activate the M(IL-4) program. As expected, while the MFI of DC-SIGN expression augmented under 72 h treatment with IL-4 in control cells, the induction of DC-SIGN was minimal in cells transfected with siRNA targeting this CLR (Figure S1B in Supplementary Material). In addition, cell viability was not affected after depletion of DC-SIGN for 72 h post-transfection (Figure S1B in Supplementary Material). Similar results in terms of DC-SIGN depletion and cell viability were obtained at 96, 120, and 144 h post-transfection (data not shown). Importantly, we noticed that DC-SIGN-depleted macrophages differentiate normally toward the M(IL-4) program (Figure S2A in Supplementary Material), and up-regulate co-stimulatory receptors when challenged with LPS (Figure S2B in Supplementary Material), as compared to control cells. Therefore, the inactivation of DC-SIGN in M(IL-4) macrophages appears to have no major consequences at steady state conditions or in response to LPS challenge.

#### The Pro-Inflammatory Response of M(IL-4) Macrophages Against Mtb Is Regulated by DC-SIGN

To explore the role of DC-SIGN in the M(IL-4) macrophage response to Mtb, we performed a genome-wide transcriptome analysis of cells expressing DC-SIGN (siControl) or not (siDC-SIGN) challenged with Mtb at different time points. In general, DC-SIGN-depleted M(IL-4) macrophages displayed a broad DEG profile when compared to control cells. At 4 h postinfection (*p.i.*), we observed that the majority of genes that are upregulated in DC-SIGN-depleted M(IL-4) macrophages are of proinflammatory nature, such as interferon alpha inducible protein (*IFI27*), *IL-6*, Oncostatin M (*OSM*), *CXCL1*, *IL-17RB,* and *IL-1β*, among others (Tables S1 and S2 in Supplementary Material). By contrast, we observed very few genes downregulated in DC-SIGN-depleted M(IL-4) macrophages, including *FCER1A* (*FcERI*), *CD1B*, *CCL17* (*TARC*), and DC-SIGN, among others (Tables S1 and S2 in Supplementary Material). At 18 h *p.i.*, the pro-inflammatory tendency is further accentuated in the DEG that becomes upregulated in DC-SIGN-depleted M(IL-4) macrophages (Figure S3A and Tables S1 and S3 in Supplementary Material). For example, Ingenuity pathway analysis indicated that the DEG in these cells formed a network centered on NF-κB, a master transcription factor that regulates the proinflammatory response against infection (Figure S3A and Table S1 in Supplementary Material). Similar to 4 h *p.i.*, there were few genes that were downregulated in in DC-SIGN-depleted M(IL-4) macrophages when compared to control cells at 18 h *p.i*., including DC-SIGN (Figure S3A and Tables S1 and S3 in Supplementary Material). Importantly, we confirmed the higher expression of the pro-inflammatory genes*,* and the downregulation of DC-SIGN, at 18 h *p.i.* by qPCR analysis (**Figure 3A**). Of note, the mRNA expression of *IL-10* was not affected significantly in the absence of DC-SIGN (**Figure 3A**).

At the protein level, we performed a dedicated antibody-based membrane array to assess the cytokine and chemokine content in the culture medium collected from DC-SIGN-depleted and control M(IL-4) macrophages at 18 h *p.i.* with Mtb. Again, we observed the shift toward a pro-inflammatory profile in DC-SIGNdepleted M(IL-4) macrophages as they secreted more IL-6, TNF, and CXCL1, among other cytokines and chemokines, compared to the supernatant content from siControl cells (**Figure 3B**). By contrast, the secretion of anti-inflammatory factors such as vascular endothelial growth factor A (VEGFA), thrombopoietin (THPO), TNF receptor superfamily member 10C (TNFRSF10C/ TRAIL-R3) and 10D (TNFRSF10D/TRAIL-R4), among others, was downregulated in the absence of DC-SIGN (**Figure 3B**). Finally, we validated by ELISA analysis the high level of both IL-6 and TNF in the supernatant content of DC-SIGN-depleted M(IL-4) macrophages in comparison to that of control cells (**Figure 3C**). Similar to RNA levels, we observed no differences in the secretion of IL-10 in the absence of DC-SIGN, as measured both by the antibody-based membrane arrays and ELISA (**Figures 3B,C**).

Figure 3 | DC-SIGN regulates the pro-inflammatory response by M(IL-4) macrophages against Mtb. Human monocytes were differentiated into macrophages using M-CSF. At day 5, macrophages were transfected with siRNA targeting DC-SIGN (siDC-SIGN) or a non-targeting siRNA (siControl). The following day, the cells were activated with IL-4 (20 ng/ml) for 48 h to induce M(IL-4) program. The cells were then infected with Mtb at multiplicity of infection (MOI) of 3 bacteria to 1 cell. At 18 h *p.i*., the cells were harvested and their supernatant collected. Assessment of (A) gene mRNA expression by qRT-PCR analysis, and (B) cytokine and chemokine content by semi-quantitative antibody array. Vertical bar graphs illustrate the fold change of mRNA/protein levels in siDC-SIGN over siControl macrophages; "0" was set arbitrarily to represent no change. Results are expressed as mean ± SD (*n* = 4 donors). One-tailed Mann–Whitney (unpaired/nonparametric): *\*P* < 0.05. (C) Assessment of cytokines by ELISA analysis. Results are expressed as before-and-after plot for the indicated genes (*n* = 11 donors). Each circle within the plots represents a single donor. Two-tailed Wilcoxon (matched-paired/nonparametric): *P* < 0.05 was considered as the level of statistical significance.

Collectively, these results suggest that the expression of DC-SIGN restrain the pro-inflammatory capacity of M(IL-4) macrophages in response to Mtb.

#### Inactivation of DC-SIGN Expression in M(IL-4) Macrophages Reduces the Intracellular Mtb Burden

While macrophages from the M2 spectrum of activation are associated with intracellular pathogen permissivity (6), it was previously shown that activation of human macrophages with IL-4 enhances the microbicidal capacity against Mtb in a dosedependent manner (24). To assess the role of DC-SIGN in the control Mtb intracellular growth, we first conducted flow cytometry-based experiments using a fluorescent Mtb strain (H37RveGFP) to determine the capacity of M(IL-4) macrophages to bind and internalize the bacillus when DC-SIGN is inactivated. To our surprise, we noticed that both parameters were equally performed by control and DC-SIGN-depleted M(IL-4) macrophages (**Figure 4A**). Next, we carried out the CFU assays. At 4 h *p.i.,* we confirmed that DC-SIGN-depleted M(IL-4) macrophages displayed normal levels of bacteria internalization compared to control cells (**Figure 4B**). However, when DC-SIGN is depleted, Mtb intracellular growth is better restricted at day 3, 5, and 7 *p.i.* compared to control M(IL-4) macrophages (**Figure 4B**). These results indicate that the expression of DC-SIGN renders M(IL-4) more permissive to intracellular growth by Mtb.

#### DC-SIGN Interferes Negatively With the Activation of M(IL-4) Macrophages Triggered by Dectin-1 in Response to Mtb

Dectin-1 is an important CLR expressed in human dendritic cells that activates a pro-inflammatory response distinguished by IL-6, TNF, IL-23, and IL-1β, against Mtb (36). Using our siRNA-mediated gene silencing method (29), we inactivated the expression of Dectin-1 in M(IL-4) macrophages and confirmed its pro-inflammatory role in the response to challenge with Mtb, as reflected by the decrease of IL-6 secretion (as an example of the pro-inflammatory response) at 18 *p.i.* (**Figure 5A**; Figure S4 in Supplementary Material). Importantly, simultaneous inactivation of Dectin-1 and DC-SIGN resulted in a similar IL-6 secretion as Dectin-1 inactivation alone, indicating that the high levels observed for this cytokine in DC-SIGN-depleted

Mtb. Control and DC-SIGN-depleted macrophages were tested for the capacity to bind (at 4°C, left) or phagocytose (at 37°C, right) the Mtb strain expressing GFP during 4 h of challenge. Bar graphs illustrate the median fluorescent intensity (MFI) of cells positive for GFP, as measured by flow cytometry analysis. Results are expressed as mean ± SD (*n* = 4 donors). NS, not significant. (B) DC-SIGN influences the capacity of M(IL-4) macrophages to control the Mtb intracellular burden. The cells were infected with Mtb (MOI of 0.2 bacteria to 1 cell) and the intracellular growth of the bacteria was followed at 4 h (day 0), 72 h (day 3), 120 h (day 5), and 168 h (day 7), as measured by colony forming unit (CFU) assays. Results are expressed as vertical scatter plots showing the CFU scoring per ml; each circle represents a single donor. Two-tailed Wilcoxon (matched-paired/nonparametric): *\*P* < 0.05; NS, not significant.

Figure 5 | DC-SIGN interferes negatively with the activation of M(IL-4) macrophages triggered by Dectin-1. Human monocytes were differentiated into macrophages using M-CSF. At day 5, the macrophages were transfected with siRNA targeting DC-SIGN (siDC-SIGN, squares) and Dectin-1 (siDectin-1, upward triangle), both siRNAs (siDKO, downward triangle), or a non-targeting siRNA (siControl, circles). The following day, the cells were activated with IL-4 (20 ng/ml) for 48 h to induce M(IL-4) program and the C-type lectin receptor expression. (A) Assessment of IL-6 secretion by ELISA analysis. The cells were infected with Mtb at MOI of 3 bacteria to 1 cell. At 18 h *p.i*., the supernatant from these cells was collected. Results are expressed as vertical scatter plots, and as mean ± SD (*n* = 7 donors). (B) Assessment of Mtb intracellular growth by colony forming unit (CFU) assay. The cells were infected with Mtb (MOI of 0.2 bacteria to 1 cell), and the intracellular growth of the bacteria was followed at 4 h (day 0, left) or 120 h (day 5, right). Results are expressed as vertical scatter plots, and as mean ± SD (*n* = 8 donors). Two-tailed Wilcoxon (matched-paired/nonparametric): *\*P* < 0.05, \*\**P* < 0.01; NS, not significant. (C,D) Upon differentiation until day 5, macrophages were then activated with IL-4 for 48 h. Prior to stimulation, M(IL-4) macrophages were pre-treated for 30 min with blocking antibodies for either DC-SIGN or Dectin-1, or both. An irrelevant antibody was used as a control. M(IL-4) macrophages were then treated with either (C) cytochalasin D (1 μg/ml), β-glucan (10 µg/ml) and ManLAM (10 µg/ml), or (D) LPS (1 µg/ml) and ManLAM (10 µg/ml). After 24 h, the supernatants were collected and the production of IL-6 was measured by for ELISA analysis.

M(IL-4) macrophages are dependent on the Dectin-1 signaling pathway (**Figure 5A**; Figure S4 in Supplementary Material). By contrast, the production of IL-10 was not affected by the inactivation of either DC-SIGN or Dectin-1 (Figure S5A in Supplementary Material). In terms of bacterial burden, we found that Dectin-1 depletion rendered the M(IL-4) macrophages more susceptible to Mtb growth (**Figure 5B**; Figure S4 in Supplementary Material). Moreover, simultaneous inactivation of both CLRs fully reversed the lower CFU counts observed when DC-SIGN is inactivated alone, suggesting that Dectin-1 depletion is dominant over that of DC-SIGN (**Figure 5B**; Figure S4 in Supplementary Material).

To better understand the potential crosstalk between these receptors in the generation of a pro-inflammatory response (e.g., IL-6 production), we employed the use of the ligands β-glucan and ManLAM to stimulate specifically Dectin-1 and DC-SIGN, respectively. As a control for macrophage activation, we used LPS as the ligand for TLR-4. As illustrated in the **Figures 5C,D**, we noticed that the stimulation of M(IL-4) macrophages with β-glucan and ManLAM induces the production of IL-6, albeit to a lower level than that obtained with LPS and ManLAM. Interestingly, while the use of a blocking antibody targeting DC-SIGN resulted in the upregulation of IL-6 after stimulation with β-glucan and ManLAM (**Figure 5C**), it failed to do so upon stimulation with LPS and ManLAM (**Figure 5D**). As expected, the use of a blocking antibody targeting the Dectin-1 receptor diminished the production of IL-6 induced by β-glucan and ManLAM (**Figure 5C**), but it had no effect on the production driven by the stimulation with LPS and ManLAM (**Figure 5D**). Likewise, simultaneous blocking of DC-SIGN and Dectin-1 resulted in a diminished production of IL-6 upon stimulation only with by β-glucan and ManLAM (**Figure 5C**), and not with LPS and ManLAM (**Figure 5D**). Of note, the crosstalk between DC-SIGN and Dectin-1 is specific for IL-6 because the production of the anti-inflammatory IL-10 was not affected (Figures S5B,C in Supplementary Material).

Altogether, these findings support a role of DC-SIGN in interfering negatively with the pro-inflammatory response triggered by Dectin-1 during Mtb infection of M(IL-4) macrophages.

#### DISCUSSION

This study highlights a dual role for DC-SIGN as, on the one hand, being a host factor granting advantage for Mtb to parasitize macrophages and, on the other hand, representing a molecular switch to turn off the pro-inflammatory response in these cells to potentially prevent potential immunopathology associated to TB. Notwithstanding, there are some limitations to this study that should be considered in the interpretation of results and derived conclusions. First, while our siRNA-mediated gene silencing protocol is effective in primary macrophages, without altering cell viability or biological functions (29), the expression of DC-SIGN is not totally abrogated. Prior to infection with Mtb, macrophages transfected with the siRNA targeting DC-SIGN still continue to express this CLR (albeit at minimal levels) upon activation with IL-4, ranging between 5 and 25% compared to control cells. Second, working with human primary macrophages introduces a high degree of biological variance compared to experiments done with human cell lines (e.g., THP-1) or bone marrow derived macrophages from congenic mice. Third, among membrane-anchored CLRs, there is a well-known functional redundancy that might cooperate in a coordinated immune response in favor or against Mtb infection (11, 37). Last, the context for which DC-SIGN is activated should be taken into account, as this CLR can mediate both immunosuppression and immunity (38). Indeed, there are multiple studies supporting the immunosuppressive role of DC-SIGN in macrophages during inflammatory contexts, including in autoimmunity and, just recently, in allotransplantation acceptance (38–40). Likewise, DC-SIGN is well known to play a pro-inflammatory role by enhancing the antigen-presentation process in human dendritic cells and inducing a strong activation of CD4+ and CD8+ T cells (41, 42). In the case of M(IL-4) macrophages, our findings on DC-SIGN should be taken within the immunosuppression context given that the nature of these cells is that of wound healers and tissue remodelers (22). In spite of these limitations, our study dealing with primary human cells in a TB context provides relevant data improving our knowledge on DC-SIGN.

We first described the presence of DC-SIGN-expressing macrophages in the pleural cavity of patients and in the lung tissue of NHP with active TB. The identification of a humanCD14+ macrophage population displaying high cell-surface levels of DC-SIGN, MRC1, and CD163, is supported by our previous finding describing the presence of the CD16hiCD163hiMerTKhi immunosuppressive macrophages in the pleural cavity of patients with active TB (25), and by our first report on DC-SIGN-expressing alveolar macrophages isolated from the bronchoalveolar lavage in patients with pulmonary TB (19). This suggests that a TB-associated environment favors the presence of macrophages with an M2-marker signature, including DC-SIGN expression. Interestingly, IL-4 is one of the most detected cytokines in TB PE, and thus making probably the responsible signal to induce DC-SIGN expression in cells at this site (43). IL-13 is another cytokine that is present in the pleural cavity of active TB patients (44), and that is probably responsible for the mediating DC-SIGN expression in macrophages since its signaling pathway is considered to be equivalent to that of IL-4 (22). At the pulmonary tissue level, while we detected DC-SIGN-expressing cells with a morphology typical of dendritic cells and lymphocytes (likely B cells), there were also numerous DC-SIGN-expressing cells which also co-express the macrophage marker CD68 and CD163. This is supported by previous reports describing the presence of M2-like cells in the context of tuberculous granulomas structures (25, 34, 35). Beyond the TB context, CD68+ macrophages expressing DC-SIGN were preferentially detected in granuloma lesions in lepromatous leprosy (45), as well as in granuloma-like structures in pathological conditions of dermatological diseases, like granuloma annulare and necrobiosis lipoidica (46). Altogether, these findings confirmed the presence of DC-SIGN-expressing macrophages in different contexts of active TB, which may not be able to mount an appropriate type-1 immune response against Mtb infection, and thus may likely contribute to the pathogenesis of this disease. While limited, these findings provide an important association between the abundance of DC-SIGN expressing macrophages and active TB, and they highlight the need to establish whether these cells actually play a pathophysiological role in this disease.

We demonstrated that DC-SIGN regulates the pro-inflammatory response of M(IL-4) macrophages during Mtb infection. Indeed, this inflammatory profile closely resembles the common response of macrophages to bacterial infections involving the upregulation of genes typical within the spectrum of M1 macrophages (6). If excessive or prolonged, the M1 macrophage response could then be deleterious for the host in terms of tissue damage or organ failure, as demonstrated during *E. coli* infection in baboon experimental peritonitis (47). In addition, a recent study in human microglia supports the immunosuppressive role for DC-SIGN in macrophages against inflammatory insults (40). Garcia-Vallejo et al. elegantly demonstrated that this CLR interacts with fucosylated glycans on myelin oligodendrocyte glycoprotein resulting in the synergistic upregulation of the TLR4-dependent production of the anti-inflammatory cytokine IL-10 in these macrophages. However, to our surprise, the IL-10 production appears not to be responsible for the dysregulation of pro-inflammatory signals in M(IL-4) macrophages. In human dendritic cells, the activation of DC-SIGN with mycobacterial ManLAM (or agonist antibodies) leads to the synergetic increase of IL-10 if it coincides with TLR4 stimulation with bacterial LPS (17, 48). Based on this well-established crosstalk, we expected the levels of IL-10 to be significantly lower in DC-SIGN-depleted M(IL-4) macrophages compared to control cells, and thus explaining the tilting toward a pro-inflammatory nature. It is likely that human M(IL-4) macrophages differ from dendritic cells (and other cells) in this respect. This is supported by our previous study showing that DC-SIGN does not potentiate IL-10 secretion in LPS-stimulated alveolar macrophages from TB patients, which highly expressed this CLR along with TLR4 (19). Moreover, another key study using humanized DC-SIGN mice demonstrated the anti-inflammatory role of this CLR in macrophages conferring protection against autoimmunity in intravenous immunoglobulin therapy, which is dependent on IL-4 and IL-33 but not IL-10 (39). It is worth mentioning that the physiological role of TLR4 remains to be proven in the context of TB. While Mtb-derived compounds can activate TLR4 *in vitro* (49, 50), it does not appear to affect the *in vivo* immune response against Mtb, as demonstrated in the mouse model (51, 52). Beyond IL-10, we showed a downregulation of protein levels involved in angiogenesis and vascularization (e.g., VEGFA), thrombopoiesis (e.g., THPO), and anti-apoptosis factors (e.g., TRAIL-R3, TRAIL-R4), in the absence of DC-SIGN during Mtb infection. These results infer a decrease of homeostatic functions such as tissue repair and remodeling, which are hallmark functional properties of M(IL-4) macrophages (22, 53, 54), and they support an immunosuppressive role for DC-SIGN in M(IL-4) macrophages that seems independent on the potentiation of IL-10 production, which may represent yet another way for Mtb to control NF-κB-driven pro-inflammatory signals.

We determined that DC-SIGN expression in M(IL-4) macrophages affects their capacity to control the Mtb burden. To our knowledge, this is the first time that deficiency of DC-SIGN in human cells is shown to directly improve the control of bacterial load in the context of infection. This improved control of Mtb burden is neither due to defects in the differentiation or activation of M(IL-4) macrophages, nor due to deficient bacterial recognition or intake by these cells, in the absence of DC-SIGN. In terms of binding, we were greatly surprised that DC-SIGN-depleted M(IL-4) macrophages were able to bind and phagocytose Mtb at the same level as control cells. In the context of DC-SIGN-expressing alveolar macrophages from TB patients, and of the human monocytic cell line expressing DC-SIGN (THP-1:DC-SIGN), we convincingly demonstrated in the past that this CLR contributed majorly to the binding and infection of these cells by Mtb (19). However, one postulate governing M(IL-4) macrophages is the acquisition of an entirely different phagocytic receptor repertoire compared to other macrophages and cell types (22). In addition to DC-SIGN, these cells are characterized by MRC1, MSR1 (macrophage scavenger receptor 1), Dectin-1, DCIR (CLEC4A), DCL-1 (CLEC13A), MGL (CLEC10A), CD36, MS4A4A (CD20-L1), and CD23 (CLEC4J), among others. Thus, it is most likely that the absence of DC-SIGN is compensated by the high abundance of these receptors in M(IL-4) macrophages unlike other cells. Recently, we described that human DC-SIGN-depleted M(IL-4) macrophages became resistant to *Leishmania infantum* infection (30). This improved control of parasite infection was dependent on the high production of IL-1β in macrophages lacking DC-SIGN. In fact, we demonstrated that DC-SIGN downregulated the mRNA expression of *LTA4H* (leukotriene A4 hydrolase), whose enzymatic activity is critical for LTB4 (leukotriene B4) synthesis, and consequently, for the caspase-1-dependent production of IL-1β. In the TB context, this is not the case for DC-SIGN. While we confirmed a significant augmentation of the *IL-1β* mRNA, we did not observe a change in the levels of IL-1β and LTB4 proteins, nor in the *LTA4H* mRNA expression, in DC-SIGN-depleted M(IL-4) macrophages at 18 h *p.i.* (data not shown). Furthermore, there were no changes in mRNA expression of antimicrobial peptides (based on the transcriptomic data), the production of reactive oxygen species, nor in the autophagy flux, between DC-SIGN-depleted and control M(IL-4) macrophages at 18 h *p.i.* (data not shown). Therefore, we inferred there might be alternative microbicidal mechanisms affected by DC-SIGN expression in M(IL-4) macrophages to control the bacilli intracellular burden.

Finally, we shed light on the capacity of DC-SIGN to regulate the pro-inflammatory response and control of bacterial burden driven by Dectin-1. Both CLRs are expressed favorably in M(IL-4) macrophages. Our results demonstrate that Dectin-1 plays a pro-inflammatory and microbicidal role against Mtb. This is in agreement with the fact that Dectin-1 is an ITAM (immunoreceptor tyrosine-based activation motif) immunoreceptor with the capacity to engage a pro-inflammatory response upon engagement with its ligand, including through the activation NFκβ-dependent transcription profile (13). In the context of TB, Dectin-1-depleted mice exhibit reduction of bacterial burden in the lungs, but there are no differences in lung pathology score, cytokine levels, or survival (55). However, in human dendritic cells, Dectin-1 is clearly partially responsible for the pro-inflammatory response against Mtb infection, specifically in the induction of the Th1/Th17 immune response (36, 56). In the case of DC-SIGN, our results propose a role for this CLR to interfere negatively in the pro-inflammatory response to Mtb that is dependent on Dectin-1. The signaling crosstalk between Dectin-1 and DC-SIGN has been reported in the recent past (30, 36, 57). While this crosstalk can be synergistic such as in the case of prostaglandin 2 (PGE2) production in human dendritic cells, where these receptors are shown to even bind together (57), it can also be the opposite. In Mtb and *L. infantum* infection, DC-SIGN interferes negatively with the production of proinflammatory signals triggered by Dectin-1 in human dendritic cells and M(IL-4) macrophages, respectively (30, 36). In this study, while we cannot exclude contribution of other PRRs (e.g., TLR-2), we propose that the pro-inflammatory signature and improved control of Mtb burden in the absence of DC-SIGN must be mainly contributed by Dectin-1. Whether this signaling crosstalk can be translated to other human macrophage subsets expressing both CLRs remains to be confirmed.

Lugo-Villarino et al. DC-SIGN Anti-Inflammatory Role in TB

In conclusion, most of what we know about DC-SIGN in the field of host–pathogen interaction mainly derives from the work done in human dendritic cells. While both dendritic cells and macrophages perform similar functions, these cells are thought to have different roles in the context of TB. On the one hand, dendritic cells are known as the professional antigen-presenting cell because they possess a great capacity to pick up and process antigens, migrate to secondary lymphoid organs, and present antigenic information to tailor an adaptive immune response against Mtb. On the other hand, macrophages are considered to be the premier effector because they are the first leukocyte to encounter Mtb in the alveolar space and activate the innate immune response to contain and eliminate this pathogen at the site of infection. Our present study brings into focus the anti-inflammatory role of DC-SIGN in M(IL-4) macrophages. Since type-2 inflammatory signals (e.g., IL-4, IL-13) are correlated with TB susceptibility and progression, we believe that DC-SIGN expression in these macrophages is of pertinence and consequence to TB pathogenesis. Indeed, these findings support the notion that Mtb hijacks the immunosuppressive aspects of DC-SIGN to invade and persist in M(IL-4) macrophages and, at the same time, modulate the local inflammatory response by these cells in its favor. Future studies shall focus on the identification of endogenous ligands targeting DC-SIGN to trigger the wound healing and tissue remodeling activity of M(IL-4) macrophages for the benefit of the patient in lung inflammatory disease. Furthermore, we believe these results can be extrapolated from the context of TB into parasitology, as M(IL-4) macrophages are considered as essential effector cells in parasite eradication (22). All things considered, beyond the role of DC-SIGN in macrophages, this study also points toward the need to investigate the pathophysiological impact of IL4 and other type-2 immune signals in the TB context, which remains unknown.

### ETHICS STATEMENT

Ethics Statement for Non-Human Primate Samples: In the Netherlands, the NHP study protocol was done to comply with the EC Directive 86/609/EEC, approved by the local independent ethics committee prior to the start of the study, and executed under Dutch law on animal experiments (agreement number DEC#579). The endpoint for any particular animal was based either by signs of severe disease (human endpoint criteria, referring to animal condition by adverse body weight development, respiratory capacity and animal behavior) or by protocol, which limited the follow-up time to 1-year postinfection. Ethic Statement for Human Samples: In Argentina, blood samples from HS or TB patients were provided by the Blood Transfusion Service, Hospital Fernandez, Buenos Aires (agreement number CEIANM-52-5-2012), or the Hospital F. J. Muñiz, Buenos Aires (protocol number: NIN-1671-12). PE were obtained by therapeutic thoracentesis by physicians at the Hospital F. J. Muñiz (Buenos Aires). The research was carried out in accordance with the Declaration of Helsinki (2013) of the World Medical Association, and was approved by the Ethics Committees of the Hospital F. J. Muñiz and the Academia Nacional de Medicina de Buenos Aires (protocol number: NIN-1671-12). Written informed consent was obtained before sample collection. The diagnosis of TB pleurisy was based on a positive Ziehl–Nielsen staining or Lowestein–Jensen culture from PE and/or histopathology of pleural biopsy, and was further confirmed by an Mtb-induced IFN-γ response and an ADA-positive test (27). Mononuclear cells from PB and PE were isolated by Ficoll-Hypaque gradient centrifugation (Pharmacia, Uppsala, Sweden), as described previously (25, 28). In France, monocytes from HS were isolated from buffy coat provided by Etablissement Français du Sang, Toulouse, under contract 21/PLER/TOU/IPBS01/2013-0042. According to articles L1243-4 and R1243-61 of the French Public Health Code, the contract was approved by the French Ministry of Science and Technology (agreement number AC 2009-921). Donors signed and provided written informed consents before sample collection.

# AUTHOR CONTRIBUTIONS

Conceptualization and methodology: GL-V, AT, LB, CL, CC, and ON. Software: IM. Investigation: GL-V, AT, LB, CL, CD, IM, AB, FC, TS, RP, IK, and CC. Resources: FV, IM-P, MS, and ON. Writing: GL-V and ON. Visualization: TS, CC, and RP. Supervision: GL-V and ON. Corresponding author: GL-V.

### ACKNOWLEDGMENTS

We acknowledge P. Constant and F. Levillan, and the BSL3 facilities at IPBS including the newly created multi-pathogen laboratory; the flow cytometry facility and personnel at TRI-IPBS, in particular C. Bordier; J. Nigou for providing reagents; and I. Vergne for expertise and technical support provided in the assessment of autophagy. We greatly thank D. Hudrisier, C. Gutierrez, C. Vérollet, B. Raynaud, M. Dupont, L. Bernard, and C. A. Spinner, for critical reading of the manuscript and helpful comments.

# FUNDING

This work was supported in part by ECOS-Sud (A14S01), *Laboratoire International Associé* (LIA, No. 1167), the *Centre National de la Recherche Scientifique*, the *Université Paul Sabatier*, the *Agence Nationale de la Recherche* (ANR 2010–01301, ANR14–CE11–0020–02, ANR16–CE13–0005–01, ANR–11–EQUIPEX–0003), the *Fondation pour la Recherche Médicale* (DEQ2016 0334894; DEQ2016 0334902), the *Fondation Bettencourt–Schueller*, and the Argentinean National Agency of Promotion of Science and Technology (PICT-2015-0055). FV acknowledges support from Aeras (Rockville, MD, USA). GL-V acknowledges support by FRM (SPF20110421334).

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at https://www.frontiersin.org/articles/10.3389/fimmu.2018.01123/ full#supplementary-material.

## REFERENCES

**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 Lugo-Villarino, Troegeler, Balboa, Lastrucci, Duval, Mercier, Bénard, Capilla, Al Saati, Poincloux, Kondova, Verreck, Cougoule, Maridonneau-Parini, Sasiain and Neyrolles. 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) and the copyright owner 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.*

#### *Edited by:*

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

#### *Reviewed by:*

*Peter Murray, Max Planck Institute of Biochemistry (MPG), Germany Joseph E. Qualls, Cincinnati Children's Research Foundation, United States Robin James Flynn, University of Liverpool, United Kingdom*

#### *\*Correspondence:*

*Luciana Balboa<EMAIL_ADDRESS>
*† These authors have contributed equally to this work.*

#### *Specialty section:*

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

*Received: 30 November 2017 Accepted: 20 February 2018 Published: 09 March 2018*

#### *Citation:*

*Genoula M, Marín Franco JL, Dupont M, Kviatcovsky D, Milillo A, Schierloh P, Moraña EJ, Poggi S, Palmero D, Mata-Espinosa D, González-Domínguez E, León Contreras JC, Barrionuevo P, Rearte B, Córdoba Moreno MO, Fontanals A, Crotta Asis A, Gago G, Cougoule C, Neyrolles O, Maridonneau-Parini I, Sánchez-Torres C, Hernández-Pando R, Vérollet C, Lugo-Villarino G, Sasiain MdC and Balboa L (2018) Formation of Foamy Macrophages by Tuberculous Pleural Effusions Is Triggered by the Interleukin-10/Signal Transducer and Activator of Transcription 3 Axis through ACAT Upregulation. Front. Immunol. 9:459. doi: 10.3389/fimmu.2018.00459*

# Formation of Foamy Macrophages by Tuberculous Pleural Effusions Is Triggered by the Interleukin-10/ Signal Transducer and Activator of Transcription 3 Axis through ACAT Upregulation

*Melanie Genoula1,2,3, José Luis Marín Franco1,2,3†, Maeva Dupont 2,3,4†, Denise Kviatcovsky1,2,3, Ayelén Milillo5 , Pablo Schierloh1,2,3, Eduardo Jose Moraña6 , Susana Poggi 6 , Domingo Palmero6 , Dulce Mata-Espinosa7 , Erika González-Domínguez 8 , Juan Carlos León Contreras7 , Paula Barrionuevo5 , Bárbara Rearte5 , Marlina Olyissa Córdoba Moreno5 , Adriana Fontanals9 , Agostina Crotta Asis10, Gabriela Gago10, Céline Cougoule2,3,4, Olivier Neyrolles2,3,4, Isabelle Maridonneau-Parini 2,3,4, Carmen Sánchez-Torres 8 , Rogelio Hernández-Pando7 , Christel Vérollet 2,3,4†, Geanncarlo Lugo-Villarino2,3,4†, María del Carmen Sasiain1,2,3† and Luciana Balboa1,2,3\**

*1 Laboratorio de Inmunología de Enfermedades Respiratorias, Instituto de Medicina Experimental (IMEX)-CONICET, Academia Nacional de Medicina, Buenos Aires, Argentina, 2 International Associated Laboratory (LIA) CNRS IM-TB/HIV (1167), Toulouse, France, 3 International Associated Laboratory (LIA) CNRS IM-TB/HIV (1167), Buenos Aires, Argentina, 4 Institut de Pharmacologie et de Biologie Structurale, Université de Toulouse, CNRS, UPS, Toulouse, France, 5 Laboratorio de Fisiología de los Procesos Inflamatorios, Instituto de Medicina Experimental (IMEX)-CONICET, Academia Nacional de Medicina, Buenos Aires, Argentina, 6 Instituto Prof. Dr. Raúl Vaccarezza, Hospital de Infecciosas Dr. F. J. Muñiz, Buenos Aires, Argentina, 7Sección de Patología Experimental, Departamento de Patología, Instituto Nacional de Ciencias Médicas y Nutrición Salvador Zubirán, Mexico City, Mexico, 8Departamento de Biomedicina Molecular, Centro de Investigación y de Estudios Avanzados del Instituto Politécnico Nacional, Mexico City, Mexico, 9 Fundación Instituto Leloir, CABA, Buenos Aires, Argentina, 10 Laboratory of Physiology and Genetics of Actinomycetes, Instituto de Biología Molecular y Celular de Rosario (IBR-CONICET), Facultad de Ciencias Bioquímicas y Farmacéuticas, Universidad Nacional de Rosario, Rosario, Argentina*

The ability of *Mycobacterium tuberculosis* (Mtb) to persist in its human host relies on numerous immune evasion strategies, such as the deregulation of the lipid metabolism leading to the formation of foamy macrophages (FM). Yet, the specific host factors leading to the foamy phenotype of Mtb-infected macrophages remain unknown. Herein, we aimed to address whether host cytokines contribute to FM formation in the context of Mtb infection. Our approach is based on the use of an acellular fraction of tuberculous pleural effusions (TB-PE) as a physiological source of local factors released during Mtb infection. We found that TB-PE induced FM differentiation as observed by the increase in lipid bodies, intracellular cholesterol, and expression of the scavenger receptor CD36, as well as the enzyme acyl CoA:cholesterol acyl transferase (ACAT). Importantly, interleukin-10 (IL-10) depletion from TB-PE prevented the augmentation of all these parameters. Moreover, we observed a positive correlation between the levels of IL-10 and the number of lipid-laden CD14+ cells among the pleural cells in TB patients, demonstrating that FM differentiation occurs within the pleural environment. Downstream of IL-10 signaling, we noticed that the transcription factor signal transducer and activator of transcription 3 was activated by TB-PE, and its chemical inhibition prevented the accumulation of lipid bodies and ACAT expression in macrophages. In terms of the host immune response, TB-PE-treated macrophages displayed immunosuppressive properties and bore higher bacillary loads. Finally, we confirmed our results using bone marrow-derived macrophage from IL-10−/− mice demonstrating that IL-10 deficiency partially prevented foamy phenotype induction after Mtb lipids exposure. In conclusion, our results evidence a role of IL-10 in promoting the differentiation of FM in the context of Mtb infection, contributing to our understanding of how alterations of the host metabolic factors may favor pathogen persistence.

Keywords: ACAT, interleukin-10, foamy macrophages, lipids, signal transducer and activator of transcription 3, tuberculosis

#### INTRODUCTION

Tuberculosis (TB) is a highly contagious disease caused by *Mycobacterium tuberculosis* (Mtb) infection. Even though the treatment of the disease has been standardized for a while, TB still remains one of the top 10 causes of death worldwide with 10.4 million new cases and 1.3 million deaths from TB among HIV-negative people in 2016 (1). Chronic host– pathogen interaction in TB leads to extensive metabolic remodeling in both the host and the pathogen (2). In fact, the success of Mtb as a pathogen derives from its efficient adaptation to the intracellular milieu of human macrophages. An important strategy to reach this metabolic adaptation is the promotion of lipid body accumulation by the host macrophage leading to foamy macrophages (FM) differentiation. The formation of lipid-laden macrophages is caused by infectious agents through deregulation in the balance between the influx and efflux of lipids. Key for the biogenesis of lipid bodies is the enzyme acyl CoA:cholesterol acyltransferase (ACAT), which represents an ideal target for pathogens (3).

Lipid body accumulation within leukocytes is a common feature in both clinical and experimental infections, especially in mycobacterial infections (4, 5). Mtb infection leads to the induction of FM, a process which is promoted by several mycobacterial lipids (6–8). This event enables the fusion between Mtb-containing phagosomes and lipid bodies resulting in an abundant supply of lipids for the pathogen (7), allowing Mtb to switch into a dormancy phenotype and to become tolerant to several front-line antibiotics (9). For this reason, lipid bodies are considered to be a secure niche for Mtb conferring protection from bactericidal mechanisms, such as respiratory burst (10). Moreover, the presence of FM within granulomatous structures was demonstrated in both experimentally infected animals and patients, especially in individuals developing secondary TB (5, 11). Therefore, FM may play a central role in mycobacterial persistence and reactivation (12, 13).

Concerning the impact of FM on the host immunity against Mtb, it was shown that human macrophages exposed to lipids prior to Mtb infection failed to produce TNF-α and to clear the infection (14, 15). Taking into account that FM generated prior to Mtb infection impair the host immune response, there is a keen interest to identify the host-derived cytokines released at the site of Mtb infection, and to understand how these signals contribute to FM differentiation and alter host defense against Mtb. In this regard, it is well known that different activation programs in macrophages driven by pro or anti-inflammatory cytokines are associated to changes in the lipid metabolism (16). Therefore, it is likely that host cytokines produced in response to Mtb infection contribute to lipids turnover promoting FM formation, and consequently lead to Mtb persistence.

In this work, we report that a TB-associated microenvironment induces FM differentiation program dependent partially on the interleukin-10 (IL-10)/signal transducer and activator of transcription 3 (STAT3) axis through ACAT upregulation. Our approach was to model a genuine TB-associated microenvironment by employing a physiological relevant sample derived from active TB patients, such as the acellular fraction of tuberculous pleural effusions (TB-PE). Indeed, TB-PE are manifested in up to 30% of patients with TB, and they are caused by the spread of Mtb into the pleural space and subsequent local inflammation and recruitment of leukocytes (17). Based on this, we show that the acquisition of the foamy phenotype with immunosuppressive properties involves high IL-10 release, low TNF-α production, impaired Th1 activation, and high bacillary loads. We also confirmed that IL-10-deficiency in bone marrow-derived macrophages (BMDM) prevented partially foamy phenotype induction upon exposure to Mtb lipids. In conclusion, our results provide evidence for a role of IL-10 in promoting foamy differentiation of macrophages in the context of Mtb infection.

#### MATERIALS AND METHODS

#### Bacterial Strain and Antigens

*Mycobacterium tuberculosis* H37Rv strain was grown at 37°C in Middlebrook 7H9 medium (Difco) supplemented with 10% albumin-dextrose-catalase (Difco) and 0.05% Tween-80 (Sigma-Aldrich). The Mtb γ-irradiated H37Rv strain (NR-49098) and its total lipids' preparation (NR-14837) were obtained from BEI Resource, USA.

## Preparation of Human Monocyte-Derived Macrophages (MDM)

Buffy coats from healthy donors were prepared at *Centro Regional de Hemoterapia Garrahan* (Buenos Aires, Argentina) according to institutional guidelines (resolution number CEIANM-664/07). Informed consent was obtained from each donor before blood collection. Peripheral blood mononuclear cells were obtained by Ficoll gradient separation on Ficoll-Paque (GE Healthcare). Then, monocytes were purified by centrifugation on a discontinuous Percoll gradient (Amersham) as previously described (18). After that, monocytes were allowed to adhere to 24-well plates (Costar) at 5 × 105 cells/well for 1 h at 37°C in warm RPMI-1640 medium (GIBCO). The cells were then washed with warm PBS twice. The final purity was checked by fluorescence-activated cell sorting analysis using an anti-CD14 monoclonal antibody (mAb) and was found to be >90%. The medium was then supplemented to a final concentration of 10% fetal bovine serum (FBS, Sigma-Aldrich) and human recombinant Macrophage Colony-Stimulating Factor (M-CSF, Peprotech) at 10 ng/ml. Cells were allowed to differentiate for 5–7 days.

### Preparation of Pleural Effusion (PE) Pools

Pleural effusions were obtained by therapeutic thoracentesis by physicians at the *Hospital F. J Muñiz* (Buenos Aires, Argentina). The research was carried out in accordance with the Declaration of Helsinki (2013) of the World Medical Association, and was approved by the Ethics Committees of the *Hospital F. J Muñiz* and the *Academia Nacional de Medicina de Buenos Aires* (protocol number: NIN-1671-12). Written informed consent was obtained before sample collection. We collected a total of 43 PE samples which were classified according to their etiology, being 38 of them associated to TB (TB-PE) and 5 to heart failure (HF-PE). Individual samples of confirmed TB patients were used for correlation analysis. A group of samples (*n* = 10) were pooled and used for *in vitro* assays to treat macrophages. The selection of these samples was based merely on practical reasons, involving those samples that have been collected earlier through the course of this study. The diagnosis of TB pleurisy was based on a positive Ziehl–Nielsen stain or Lowestein–Jensen culture from PE and/or histopathology of pleural biopsy, and was further confirmed by an Mtb-induced IFN-γ response and an ADA-positive test (19). Exclusion criteria included a positive HIV test, and the presence of concurrent infectious diseases or non-infectious conditions (cancer, diabetes, or steroid therapy). None of the patients had multidrug-resistant TB. Those PE samples derived from patients with pleural transudates secondary to heart failure (HF-PE, *n* = 5) were employed to prepare a second pool of PE, used as control of non-infectious inflammatory PE. The PE were collected in heparin tubes and centrifuged at 300 *g* for 10 min at room temperature without brake. The cell-free supernatant was transferred into new plastic tubes, further centrifuged at 12,000 *g* for 10 min and aliquots were stored at −80°C. After having the diagnosis of the PE, pools were prepared by mixing same amounts of individual PE associated to a specific etiology. The pools were decomplemented at 56°C for 30 min, and filtered by 0.22 µm in order to remove any remaining debris or residual bacteria.

# FM Induction

Macrophages were plated on glass coverslips within a 24-well tissue culture plate (Costar) at a density of 5 × 105 cells/ml per well with or without 20% v/v of PE, 10 µg/ml of Mtb lipids (BEI resources) or infected with Mtb (MOI 2:1) for 24 h. When indicated, cells were pre-incubated with either Cucurbitacin I (50–100 nM, Sigma-Aldrich), or the STAT3 inhibitor Stattic (1–20 µM, Sigma-Aldrich), or the ACAT inhibitor Sandoz 58-035 (5–50 µg/ml, Sigma-Aldrich) for 2 h prior TB-PE addition and for further 24 h during TB-PE incubation. DMSO alone was used as control. Alternatively, cells were treated with recombinant human IL-10 (Peprotech) at the indicated doses. Foam cell formation was followed by Oil Red O (ORO) staining (Sigma-Aldrich) as previously described (20) at 37°C for 1–5 min, and washed with water three times. For the visualization of the lipid bodies, slides were prepared using the aqueous mounting medium Poly-Mount (Polysciences), observed *via* light microscope (Leica) and finally photographed using the Leica Application Suite software. Alternatively, after fixation, cells were labeled with 1 µg/ml of BODIPY 493/503 (Life technologies) for 15 min in order to visualize the lipid bodies by green fluorescence emission using a confocal microscope (Olympus BX51).

### Infection of Human Macrophages with Mtb

Infections were performed in the biosafety level 3 laboratory at the *Unidad Operativa Centro de Contención Biológica, ANLIS-MALBRAN* (Buenos Aires), according to the biosafety institutional guidelines. Macrophages seeded on glass coverslips within a 24-well tissue culture plate (Costar) at a density of 5 × 105 cells/ml were infected with Mtb H37Rv strain at a MOI of 2:1 during 1 h at 37°C. Then, extracellular bacteria were removed gently by washing with pre-warmed PBS, and cells were cultured in RPMI-1640 medium supplemented with 10% FBS and gentamicin (50 µg/ml) for 24 h. The glass coverslips were fixed with PFA 4% and stained with ORO, as was previously described.

# Quantification of Total Cholesterol

Total cholesterol was determined in TB-PE or cell lysates using the *Colestat Enzimatico kit* according to manufacturer instructions (Wiener Lab, Argentina). This assay is based in Trinder reaction in which cholesterol in the sample is quantified by enzymatic hydrolysis of cholesterol esters (21).

# Lipid Analysis

Total lipids were extracted from the same number of macrophages with methanol/chloroform (2:1 v/v) as described by Bligh and Dyer (22). After extraction, lipids were dried and analyzed by thin layer chromatography on silica gel 60 F254 plates (Merck), using hexane/diethyl ether/acetic acid (75:25:1, v/v/v) as the developing solvent. Chemical staining with Cu-phosphoric was used for detection.

### Phenotypic Characterization by Flow Cytometry

Macrophages were centrifuged for 7 min at 1,200 rpm and then stained for 40 min at 4°C with fluorophore-conjugated antibodies FITC-anti-CD36 (clone 5-271), PerCP.Cy5.5-anti-CD14 (clone HCD14), PE-anti-CD163 (clone GHI/61), FITC-anti-CD206 (clone C068C2) (Biolegend), FITC-anti-HLA-DR (clone G46-6), or PE-anti-CD274 (clone MIH1) (BD Biosciences), and in parallel, with the corresponding isotype control antibody. After staining, the cells were washed with PBS 1×, centrifuged and analyzed by flow cytometry using FACSCalibur cytometer (BD Biosciences). The median fluorescence intensity were analyzed using FCS Express V3 software (*De Novo* Software, Los Angeles, CA, USA). The mean of the median fluorescence intensities (MFI) of the independent assays were used for comparisons between experimental conditions.

#### Soluble Cytokine Determinations

The amounts of human TNF-α, IL-1β, IL-10, IL-6, IFN-γ, and IL-4 were measured by ELISA, according to manufacturers' instructions kits (TNF-α, IL-4, and IFN-γ Ready-SET-Go!™ Kits from eBioscience; IL-10, IL-1β, and IL-6 ELISA MAX™ Deluxe Kits from Biolegend). The detection limit was 3 pg/ml for TNF-α; 8 pg/ml for IL-10 and IL-6; 6.25 pg/ml for IFN-γ and IL-4; and 15.6 pg/ml for IL-1β. Murine TNF-α and IL-10 were measured by ELISA, according to manufacturer's instructions (OptEIA™ Set kits from BD Bioscience) with a detection limit at 30 pg/ml.

#### Cytokine Depletions of PE

Tuberculosis-PE were incubated with 10 µg/ml of the following neutralizing antibodies for 1 h at 4°C for the specific depletion of IL-10 (αIL-10, clone 19F1; Biolegend), IL-6 (αIL-6, clone MQ2- 13A5; BD Bioscience), IL-1β (αIL-1β, clone 8516.311; SIGMA), or TNF-α (αTNF-α, clone MAb1; Biolegend). Then, 100 µl/ml of Protein G Sepharose beads (Amersham) were added and incubated for 1 h at 4°C. Finally, TB-PE were centrifuged at 12,000 *g* to remove antibody-bead complexes and then filtered (0.22-µm pores) before use. IFN-γ depletion was performed by incubating TB-PE for 2 h in sterile 96-well plates that had been coated with the capture antibody provided by the Human IFN-γ ELISA Kit (BD Bioscience). In all cases, depletions were controlled by ELISA.

#### Electron Microscopy

Macrophages exposed (or not) to TB-PE, and depleted (or not) for IL-10, were prepared for transmission electron microscopy study. For this purpose, cells were centrifuged at 6,000 rpm for 1 min, fixed in 1% glutaraldehyde dissolved in 0.1 M cacodylate buffer (pH 7); post-fixed in 2% osmium tetroxide; dehydrated with increasing concentrations of ethanol and gradually infiltrated with Epon resin (Pelco). Thin sections were contrasted with uranyl acetate and lead citrate (Electron Microscopy Sciences, Fort Washington, PA, USA) and examined with a FEI Tecnai transmission electron microscope (Hillsboro, OR, USA). For morphometry, 30 cells from each condition were randomly selected and digitalized at 40,000× magnification. Then the area of lipid electron dense vacuoles per cell were measured and compared in each experimental group by automated morphometry.

# Proliferation of Antimycobacterial CD4 T Cells Induced by Macrophages

We purified and maintained CD4 T cells from blood derived from healthy PPD+ donors as previously demonstrated (23). In parallel, we cultured autologous monocytes to generate macrophages and exposed them or not to TB-PE 20% v/v for 24 h. The next day, the medium was replaced, and macrophages were loaded with mycobacterial antigens by adding γ-irradiated Mtb at 2 bacteria to 1 macrophage ratio for further 24 h. Thereafter, the medium was replaced and autologous carboxyfluorescein succinimidyl ester (CFSE)-labeled CD4 T cells (Invitrogen) were added at a ratio of 10 lymphocytes to 1 macrophage for 5 days, as detailed previously (24). To determine IFN-γ production among proliferating CD4 T cells (CFSElow), brefeldin A (5 µg/ml; Sigma Chemical Co.) was added 4 h prior the end of the coculture to block cytokine secretion. Thereafter, CFSE-labeled lymphocytes were first stained with anti-CD4-PerCP-Cy5.5 mAbs (eBioscience), then fixed with 0.5% PFA for 15 min. Second, cells were permeabilized with 500 µl Perm2 (Becton Dickinson, Cockysville, MD, USA) for 10 min and incubated with anti-IFN-γ-PE mAb (Invitrogen, CA, USA). Cells were gated according to its FSC and side scatter (SSC) properties analyzed on FACScan (Becton Dickinson). In order to gate out dead lymphocytes, the gate of CD4 T cells with increased SSC and low FSC was excluded (25). Isotype matched controls were used to determine auto-fluorescence and non-specific staining. Analysis was performed using the FCS Express (*De Novo* Software) and results were expressed as percentages of IFN-γpos/CFSElow/ CD4pos T cells induced by the different macrophage populations. To complement these results, the amounts of IFN-γ released in the supernatant throughout the coculture were determined by ELISA according to the manufacturer's kit indications (BD Bioscience).

### Measurement of Bacterial Intracellular Growth in Macrophages by Colony Forming Unit (CFU) Assay

Macrophages exposed (or not) to TB-PE, were infected with H37Rv Mtb strain at a MOI of 1 bacteria/cell in triplicates. After 2 h, extracellular bacteria were removed by gently washing four times with pre-warmed PBS. At 4 h and days 3, 6, and 10, cells were lysed in 0.1% SDS and neutralized with 20% Bovine Serum Albumine in Middlebrook 7H9 broth. Serial dilutions of the lysates were plated in triplicate, onto 7H11-Oleic Albumin Dextrose Catalase (OADC, Difco) agar medium for CFU scoring at 21 days later.

#### Western Blots

Macrophages were treated (or not) with TB-PE. Following the different experimental treatments, cells were lysed in ice-cold buffer consisting of 150 mM NaCl, 10 mM Tris, 5 mM EDTA, 1% SDS, 1% Triton X-100, 1% sodium deoxycholate, gentamicin/streptomycin, 0.2% azide plus a cocktail of protease inhibitors (Sigma-Aldrich). Lysates were incubated on ice for 3 h and cleared by centrifugation for 15 min at 14,000 rpm at 4°C. Protein concentrations were determined using the BCA protein assay (Pierce). Equal amounts of protein (40 µg) were then resolved on a 10% SDS-PAGE. Proteins were then transferred to Hybond-ECL nitrocellulose membranes (Amersham) for 2 h at 100 V and blocked with 1% BSA-0.05% Tween-20 for 1 h at room temperature. Membranes were then probed with primary anti-human ACAT (1:200 dilution, SOAT; Santa Cruz) or anti-human pY705-STAT3 (1:1,000 dilution, Cell Signaling Technology, clone D3A7) overnight at 4°C. After extensive washing, blots were incubated with a HRP-conjugated goat anti-rabbit IgG Ab (1:5,000 dilution; Santa Cruz Biotechnology) or HRP-conjugated goat antimouse IgG Ab (1:2,000 dilution; Santa Cruz Biotechnology) for 1 h at room temperature. Immunoreactivity was detected using ECL Western Blotting Substrate (Pierce). Protein bands were visualized using Kodak Medical X-Ray General Purpose Film. For internal loading controls, membranes were stripped by incubating in buffer consisting of 1.5% Glycine, 0.1% SDS, 1% Tween-20, pH 2.2 for 10 min twice, extensively washed and then reprobed with anti-β-actin (1:2,000 dilution; ThermoFisher, clone AC-15) or anti-STAT3 Ab (1:1,000 dilution; Cell Signaling Technology, clone D1A5). Results from Western blot were analyzed by densitometric analysis (Image J software).

#### Immunostaining

Macrophages were plated on glass coverslips and were treated with or without 20% v/v of TB-PE for 24 h. Cells were fixed with PFA 4% for 20 min at room temperature and then PFA was quenched with 50 mM NH4Cl for 2 min. Cells were rinsed in PBS once and then were labeled with 1 µg/ml of BODIPY 493/503 (Life technologies) for 15 min before permeabilization with PBS-Triton X-100 0.1% for 10 min. Cells were then incubated with PBS-BSA 3% w/v for 30 min prior to overnight incubation at 4°C with primary anti-human pY705-STAT3 (dilution 1/100, Cell Signaling Technology, Clone D3A7). Cells were then washed and incubated with Goat anti-Mouse IgG, AlexaFluor 555 (dilution 1/1,000, Cell Signaling Technology) for 1 h at room temperature. Cells were extensively washed and then incubated for 10 min with DAPI in PBS-BSA 1% (500 ng/ml, Sigma-Aldrich). Finally, slides were mounted and visualized with a Leica DM-RB fluorescence microscope.

#### Mice

Wild-type (WT) BALB/c and IL-10 knockout (IL-10 KO) (C.129P2(B6)-IL-10 tm1Cgn/J) BALB/c male mice (8–12 weeks old), were obtained from the Leloir Institute Foundation. Animals were bred and housed in accordance with the guidelines established by the Institutional Animal Care and Use Committee of Institute of Experimental Medicine (IMEX)-CONICET-ANM. All animal procedures were shaped to the principles set forth in the Guide for the Care and Use of Laboratory Animals (26).

### Generation of BMDMs

Femurs and tibia from WT and IL-10 KO mice were removed after euthanasia and the bones were flushed with RPMI-1640 medium by using syringes and 25-Gauge needles. The cellular suspension was centrifuged and the red blood cells were removed. The BMDMs were obtained by culturing the cells with RPMI-1640 medium containing l-glutamine, pyruvate, β-mercaptoethanol (all from Sigma-Aldrich), 10% FCS and 20 ng/ml of murine recombinant M-CSF (Biolegend) at 37°C in a humidified incubator for 7–8 days. Differentiated BMDMs were re-plated into 24-well tissue culture plates in complete medium prior to cell stimulation with Mtb lipids for 24 h. Alternatively, recombinant murine IL-10 (Peprotech) at 10 ng/ml was added to the cultures for 24 h.

### Statistical Analysis

All values are presented as mean and SEM of a number of independent experiments. Independent experiments are defined as those performed with macrophages derived from monocytes isolated independently from different donors. As most of our datasets did not pass the normality tests, non-parametric tests were applied. Comparisons between more than two paired data sets were made using the Friedman test followed by Dunn's Multiple Comparison Test. Comparisons between two paired experimental conditions were made using the two-tailed Wilcoxon Signed Rank. Correlation analyses were determined using the Spearman's rank test. For all statistical comparisons, a *p* value < 0.05 was considered significant.

# RESULTS

### Tuberculous PE Fluids Induce Lipid Bodies Accumulation in Macrophages

It has been demonstrated that Mtb induces a lipid-rich foam cell phenotype in host macrophages (6–8, 27), possibly *via* TLR2 and TLR6 activation (28). Moreover, merely isolated lipids from Mtb were able to induce the foamy phenotype (7, 27). Based on this knowledge, we aimed to determine whether soluble factors found in physiological samples derived from active TB patients could promote the generation of FM. To this end, we treated human M-CSF-driven macrophages with cell-free preparations of PE derived from patients with tuberculosis (TB-PE) in order to mimic a genuine microenvironment derived during Mtb infection. According to the pattern of staining of neutral lipids with ORO, we observed that TB-PE treatment induced lipid bodies accumulation in macrophages to the same extent as Mtb infection or exposure to Mtb-derived lipids (**Figure 1A**). We also demonstrated that the formation of lipid bodies was specific for TB-PE treatment in comparison to PE from patients with heart failure (HF-PE) (**Figure 1B**). Moreover, biochemical analysis of the lipids recovered from macrophages identified higher abundance of cholesteryl esters and triacylglycerols in TB-PE-treated macrophages in comparison to control or HF-PE-treated macrophages (**Figure 1C**). Additionally, unlike HF-PE, the treatment with TB-PE resulted in an increase of the total intracellular cholesterol content (**Figure 1D**) and a twofold increase of CD36 expression (**Figure 1E**). Therefore, soluble factors released during pleural Mtb infection induced lipid body accumulation in human macrophages that typically confirm the FM phenotype.

Figure 1 | Tuberculous pleural effusions (TB-PE) induce lipid bodies accumulation in macrophages. (A) Human monocyte-derived macrophages (MDM) were treated either with TB-PE or with *Mycobacterium tuberculosis* (Mtb) lipids, or infected with Mtb H37Rv for 24 h and then stained with Oil Red O (ORO). Representative images are shown in left panels (40× magnification) and the integrated density of ORO staining is shown in right panels. Values are expressed as means ± SEM of six independent experiments, considering five microphotographs per experiment. Wilcoxon signed rank test: \**p* < 0.05. (B) MDM were treated with TB-PE or PE from heart failure patients (HF-PE) and stained with ORO (*n* = 5). (C) Left panel shows a representative thin layer chromatographic analysis of lipids from MDM treated or not with either TB-PE or HF-PE. Total lipids were extracted from untreated MDM (lane 1), TB-PE-treated MDM (lane 2), HF-PE-treated MDM (lane 3), and the standard lipids triacylglycerol (TAG, lane 4) and free cholesterol (CHO, lane 5). Cholesterol esters (CE) are also indicated. Right panels depict the area of spots for CE, TAG, and CHO in control and TB-PE or HF-PE treated MDM (*n* = 4). (D) Intracellular total cholesterol (CHO) content in MDM exposed or not to TB-PE or HF-PE measured by an enzymatic method (*n* = 5). (E) Mean fluorescence intensity (MFI) of CD36 cell-surface expression in MDM exposed or not to TB-PE or HF-PE measured by flow cytometry (*n* = 5). (B–E) Friedman test followed by Dunn's Multiple Comparison Test: \**p* < 0.05; \*\**p* < 0.01 for TB-PE vs Ctl or as depicted by lines.

### FM Induced by Tuberculous PE Display Immunosuppressive Properties

In order to assess whether the differentiation into FM induced by TB-PE may have a negative impact on the development of the antimycobacterial response, we determined phenotype and functions of TB-PE-induced FM. As observed in **Figure 2A**, macrophages treated with TB-PE displayed a higher expression of anti-inflammatory markers such as CD163, mannose receptor or CD206 (MRC1), and PD-L1, and a lower expression of the MHC class II cell-surface receptor, HLA-DR. In accordance with the acquisition of this anti-inflammatory profile, TB-PEinduced FM secreted higher levels of IL-10 and lower levels of TNF-α upon stimulation with irradiated *Mycobacterium tuberculosis* (iMtb) than untreated cells (**Figure 2B**). In order to assess the effect of TB-PE treatment on the presentation of mycobacterial antigens, macrophages were treated with or without TB-PE for 24 h, washed and stimulated with iMtb for further 24 h. Thereafter, cells were washed and cocultured with autologous CFSE-labeled CD4 T cells derived from healthy PPD positive subjects for 5 days. Based on the CFSE labeling of CD4 T cells, while TB-PE-treated macrophages did not induce differential levels of proliferation of antimycobacterial CD4 T cells (**Figure 2C**), they promoted differential distribution of the IFN-γ producing clones (**Figure 2D**). In particular, those macrophages treated with TB-PE induced lower percentages of IFN-γ producing clones (**Figure 2D**) and a lower release of IFN-γ (**Figure 2E**), as compared to untreated macrophages in response to iMtb stimulation. Of note, IL-4 and IL-17 contents were undetectable in these assays (data not shown). Therefore, the activation of Ag-specific IFN-γ-producing CD4 T cells was impaired when macrophages were exposed to TB-PE, suggesting that the accumulation of lipid bodies is accompanied by a reduced capacity to activate antimycobacterial Th1 cells.

Next, we assessed whether the treatment with TB-PE had an impact on the control of the bacillary load. To accomplish this, macrophages were treated with TB-PE for 24 h, washed and infected with Mtb. Although no differences were observed in the uptake of the mycobacteria as judged by the scoring of the CFU at 4 h post-infection (**Figure 2F**), a significant increase in the bacillary load was observed in TB-PE-treated macrophages at later time points (**Figure 2G**). Therefore, FM induced by TB-PE are more susceptible to Mtb intracellular growth.

#### IL-10 Promotes Accumulation of Lipid Bodies in Macrophages under TB-PE Treatment

In order to evaluate the host factors involved in promoting the accumulation of lipid bodies by TB-PE, we depleted different cytokines known to be highly present in this fluids (29, 30), including IL-10, TNF-α, IL-1β, IL-6, and IFN-γ. We then evaluated the ability of these depleted-PE to induce the foamy phenotype in macrophages. The depletion of each individual cytokine was confirmed by ELISA (Figure S1A in Supplementary Material). Interestingly, only the depletion of IL-10 from TB-PE was capable of preventing lipid bodies accumulation (**Figure 3A**). This result was also confirmed by labeling the cells with Bodipy staining (Figure S1B in Supplementary Material). In addition, macrophages exposed to IL-10-depleted TB-PE showed a reduction of intracellular cholesterol content and CD36 cell-surface expression in comparison to those cells treated with non-depleted or depleted of any other cytokines (**Figures 3B,C**). In line with these results, the addition of recombinant IL-10 to the IL-10 depleted TB-PE restored the acquisition of the foamy phenotype in a dose-dependent manner (**Figure 3D**). Of note, the addition of IL-10 in the absence of TB-PE did not induce the foamy phenotype (**Figure 3D**). Thereafter, we determined the presence of the lipidic vacuoles by electron microscopy in macrophages treated with TB-PE depleted (or not) of IL-10. As it is shown in **Figure 3E**, while untreated macrophages showed numerous pseudopodia and empty cytoplasmic vacuoles, TB-PE-treated macrophages displayed electron dense lipid osmiophilic vacuoles, which correspond to lipid bodies. Interestingly, macrophages treated with IL-10-depleted TB-PE showed smaller-sized lipidic vacuoles than those exposed to non-depleted TB-PE. These results indicate that the IL-10 present in TB-PE is a key host factor promoting the lipid-laden phenotype in the presence of exogenous lipids.

### IL-10 Levels Correlate with FM Abundance in PE

In order to evaluate whether IL-10 level is associated to the acquisition of the foamy phenotype in the course of a Mtb natural infection, we assessed the levels of IL-10 and total cholesterol in individual preparations of TB-PE, and the numbers of FM found

Figure 2 | Macrophages treated with tuberculous pleural effusion (TB-PE) display immunosuppressive properties. (A) Mean fluorescence intensity (MFI) of CD163, PDL-1, MR, and HLA-DR measured by flow cytometry in human monocyte-derived macrophages (MDM) exposed or not to TB-PE (*n* = 5). Wilcoxon signed rank test: \**p* < 0.05. (B) Levels of secreted IL-10 and TNF-α by MDM exposed or not to TB-PE in response to irradiated *Mycobacterium tuberculosis* (iMtb) measured by ELISA (*n* = 5). Friedman test followed by Dunn's Multiple Comparison Test: \**p* < 0.05; \*\**p* < 0.01, for iMtb-stimulated vs unstimulated, or as indicated in the graph. (C–E) MDM from healthy PPD+ donors exposed or not to TB-PE for 24 h, were stimulated or not with iMtb for 24 h and then used as antigen presenting cells (APC) in autologous proliferation assays. Cocultures were performed with autologous carboxyfluorescein succinimidyl ester (CFSE)-labeled CD4 T cells at a ratio of 10 T cells: 1MDM. (C) Representative histograms showing CFSE labeling in CD4pos T cells activated by macrophages exposed (or not) to TB-PE, loaded (or not) with iMtb. Right panel shows the quantification of the percentages of proliferating CD4pos T cells (CFSElow/CD4pos cells) in each condition. Loaded vs unloaded with iMtb: \**p* ≤ 0.05 (*n* = 4). (D) Representative dot plots showing CD4 and IFN-γ expression among CFSElow/CD4pos T cells activated by MDM exposed (or not) to TB-PE, loaded (or not) with iMtb. Right panel shows the quantification of the percentages of proliferating IFN-γ-producing CD4pos T cells (IFN-γpos/CFSElow/ CD4pos T cells) in each condition (*n* = 4). Friedman test followed by Dunn's Multiple Comparison Test: \**p* < 0.05, for iMtb-stimulated vs unstimulated, or as indicated in the graph. (E) The amounts of IFN-γ released throughout the coculture were determined by ELISA. Friedman test followed by Dunn's Multiple Comparison Test: \**p* < 0.05; \*\**p* < 0.01, for iMtb-stimulated vs unstimulated, or as indicated in the graph. (F) Bacillary loads in MDM treated (or not) with TB-PE, washed, and infected with *Mycobacterium tuberculosis* (Mtb) strain H37Rv for 4 h (*n* = 6). Wilcoxon signed rank test. (G) Intracellular colony forming units were determined at different time points in MDM treated with TB-PE for 24 h, washed and infected with Mtb (*n* = 10). Wilcoxon signed rank test: \**p* < 0.05; \*\**p* < 0.01; \*\*\**p* < 0.001 for TB-PE treated vs Ctl.

within the pleural fluids mononuclear cells (PFMC). We found that levels of IL-10 and total cholesterol were positively correlated (**Figure 4A**), unlike other cytokines such as IFN-γ, IL-6, TNF-α, and IL-1β (Figure S2 in Supplementary Material). Noticeably,

Test: \**p* < 0.05; \*\**p* < 0.01; \*\*\**p* < 0.001 for experimental condition vs Ctl or as depicted by lines.

there was also a positive correlation between the level of IL-10 and the number of FM among the PFMC (**Figure 4B**). Moreover, lipid-laden CD14+ cells were found in the pleural compartment but not in paired peripheral blood (**Figure 4C**). These results

small sized ones. Morphometric study of the area per vacuole in each condition is shown (*n* = 14). (A–E) Friedman test followed by Dunn's Multiple Comparison

support the idea that IL-10 potentiates the acquisition of the foamy program in human macrophages in the context of a natural infection.

### IL-10 Deficiency Prevents the Foamy Phenotype in BMDM

To confirm the role of IL-10 in the differentiation of FM, we evaluated whether BMDM derived from IL-10-knock out (KO) mice could indeed become FM after the treatment with lipids from Mtb. First, we determined IL-10 production in M-CSF-derived BMDM from WT mice stimulated with mycobacterial lipids for 24 h. As shown in **Figure 5A**, unlike the undetectable level of TNF-α (data not shown), BMDM secreted IL-10 in response to mycobacterial lipid-stimulation. Second, we compared whether BMDM derived from WT or IL-10-KO mice differed in their propensity to accumulate lipid bodies in response to Mtb-derived lipids. As illustrated in **Figures 5B,C**, IL-10-deficiency prevented the foamy phenotype induced by Mtb lipids, which in turn could be partially reverted by the addition of exogenous IL-10. In agreement with our previous results, exogenous IL-10 did not induce the foamy phenotype in the absence of the source of lipids in BMDM. These results obtained in murine macrophages confirm those obtained in human TB-PE-treated macrophages, demonstrating the key role of IL-10 in favor of the differentiation program toward foamy cells.

# The IL-10/STAT3 Axis Is Involved in the FM Differentiation Induced by TB-PE

Considering the role of IL-10 in promoting the differentiation of macrophages into FM, and that STAT3 is a pivotal transcriptional factor induced by IL-10 (31, 32), we studied the contribution of STAT3 activation in the induction of the foamy phenotype by measuring its phosphorylated form (pSTAT3). First, we determined that STAT3 was activated in TB-PE-treated macrophages detecting its phosphorylated form by western blot and immunofluorescence microscopy (**Figures 6A,B**). As depicted in **Figure 6B** FM induced by TB-PE showed nuclear localization of STAT3 phosphorylated on tyrosine 705, reflecting STAT3 activation. Importantly, pharmacological inhibition of STAT3 with Stattic (or with cucurbitacin I) prevented the accumulation of lipid bodies in macrophages in a dose-dependent manner (**Figure 6C**; Figures S3A,B in Supplementary Material). Therefore, the enhancement of FM differentiation driven by IL-10 is mediated by STAT3.

### IL-10 Enhances ACAT Expression in TB-PE-Treated Macrophages Leading to FM Differentiation

In order to elucidate the mechanism by which IL-10 promotes FM differentiation, we assessed the expression of ACAT, which is central for the biogenesis of lipid bodies by converting free- into

esterified-cholesterol that are eventually packed inside the lipid droplets (33). We first showed that the foamy phenotype induced by TB-PE was dependent on ACAT activity, as judged both by the increase of ACAT expression after TB-PE treatment (**Figure 7A**), and by the blocking of the accumulation of lipid bodies in the presence of Sandoz, a specific inhibitor of ACAT (**Figure 7B**). Noticeably, the inhibition of ACAT lead to reduced amounts of both cholesteryl esters and triacylglycerols in TB-PE-treated macrophages (**Figure 7C**), confirming the involvement of ACAT in FM formation upon TB-PE treatment. We then assessed ACAT expression in IL-10-depleted TB-PE and we found that its expression was reduced in the absence of IL-10 (**Figure 7A**). In agreement with this result, ACAT induction by TB-PE was abolished when STAT3 activity was inhibited (**Figure 7D**). Therefore, the IL-10/STAT3 axis activated by TB-PE enhances FM differentiation through the upregulation of ACAT expression, leading to an increased biogenesis and accumulation of lipid bodies.

Based on our findings, we propose a model for the modulation of FM in the context of a physiologically relevant microenvironment promoted by Mtb infection for which the axis IL-10/ STAT3 induces the accumulation of lipid bodies *via* ACAT upregulation. This in turn is accompanied by an increase of CD36 and the acquisition of immunosuppressive properties, such as a reduced induction of antimycobacterial Th1 clones, an enhanced production of IL-10 and a more permissive phenotype for bacillary growth (**Figure 8**).

#### DISCUSSION

Tuberculosis, as a chronic condition, entails the establishment of extensive metabolic remodeling in both host and pathogen. One of the consequences of this adaptation is the formation of FM. Since FM have been associated with the bacilli persistence and tissue pathology (6, 8, 12, 27, 34), we aimed to determine which host factors may contribute to enlarge the pool of FM in TB. In this sense, we used the acellular fraction of TB-PE to mimic those soluble factors released locally during Mtb infection. Although pleural disease due to *Mtb* is generally categorized as extra-pulmonary, there is an intimate anatomic relationship between the pleura and the pulmonary parenchyma (35, 36). Current literature supports the notion that TB-PE is the consequence of a direct local infection with a cascade of events, including an immunological response, instead of the result of a pure delayed hypersensitivity reaction, as previously thought (37). Even though we cannot state that macrophages infiltrating the pleural cavity will reproduce with fidelity those macrophages in the infected alveolar space or lung interstitial tissue, we can affirm that TB-PE represents microenvironment from a human respiratory cavity that is impacted by the infection. To our knowledge, this is the first time that such a complex but physiologically relevant human sample has been used in order to study the biology of FM. Using this *in vitro* model, we demonstrated that the acellular fraction of TB-PE induces the accumulation of lipid bodies in human macrophages. Moreover, our finding was validated by the detection of lipid-laden CD14+ cells isolated directly from the mononuclear cells of the PE, providing physiological relevance to our *in vitro* model. Taking into account that tuberculous PE has, if any, very few bacilli content (38), and that it displays high cholesterol content, we infer that in our model the source of lipids feeding the lipid bodies in the macrophages are likely host-derived components instead of mycobacterial ones.

24 h. Lipid bodies' content was determined by Oil red O (ORO) staining (*n* = 6). Representative images are shown and the integrated density of ORO staining is shown. Friedman test followed by Dunn's Multiple Comparison Test: \**p* < 0.05; \*\**p* < 0.01 for TB-PE treated vs Ctl, or as depicted by lines.

Although lipid bodies were qualified as passive organelles involved in lipid storage, it became clear that these organelles play a central role in several inflammatory (e.g., atherosclerosis) or chronic infectious diseases (e.g., TB) (8, 13). In this work, we provide evidence that FM differentiation induced by TB-PE is potentiated by IL-10 in association with the acquisition immunosuppressive properties, impairing the activation of antimycobacterial Th1 clones, producing the anti-inflammatory cytokine IL-10 and bearing higher bacillary loads. This is in line with previous reports characterizing the immunosuppressive profile in macrophages activated by the IL-10/STAT3 signaling pathway (39, 40). In this study, we show that FM formation depends (albeit partially) on the IL-10/STAT3 axis, and thus establishing an association between both processes (accumulation of lipid bodies and immunosuppression). Within this context, we observed that both processes are enhanced by the IL-10/STAT3 signaling pathway, arguing for likelihood of two independent outcomes emanating from the same signaling axis (**Figure 8**). Of note, it has been reported that the synthetic glucocorticoid dexamethasone can upregulate ACAT expression promoting formation of FM (41). Although there are extensive reports demonstrating transcriptional interactions between STAT3 and glucocorticoids leading to repression or synergism of target genes (42), it is interesting to notice that both IL-10 and glucocorticoids can polarize macrophages into an immunoregulatory profile (43). Based on that, we propose that the establishment of a foamy phenotype is accompanied by the acquisition of immunosuppressive properties.

It should be mentioned that Stattic is not an inhibitor completely specific for STAT3 (44). In this regard, a desirable goal is the discrimination between STAT3 and STAT1 involvement because both transcription factors can be activated by IL-10 (albeit at different levels), and because of their high degree of homology, particularly in their SH2 domains (44). Under our experimental conditions, unlike the strong activation of STAT3, we did not observe the phosphorylation of STAT1

Figure 7 | The interleukin-10 (IL-10)/signal transducer and activator of transcription 3 (STAT3) axis promotes foamy macrophage formation through ACAT upregulation. (A) Analysis of ACAT and β-actin protein expression level by western Blot (left panel) and quantification (right panel; *n* = 5) in human monocyte-derived macrophages (MDM) treated with tuberculous pleural effusion (TB-PE) for 24 h depleted or not for IL-10. (B) MDM were treated or not with different concentrations of Sandoz (Sz, ACAT inhibitor) for 1 h, exposed or not to TB-PE for 24 h, and then stained with Oil Red O (ORO). The Integrated density and representative images are shown (*n* = 6). (A,B) Friedman test followed by Dunn's Multiple Comparison Test: \**p* < 0.05; \*\**p* < 0.01 for TB-PE treated vs Ctl, or as depicted by lines. (C) Thin layer chromatographic analysis of lipids from MDM treated or not with TB-PE in the presence or not of Sz. Total lipids were extracted from untreated MDM (lane 1), TB-PE-treated MDM (lane 2), TB-PE-treated MDM in the presence of Sz (lane 3), and the standard lipids triacylglycerol (TAG, lane 4) and cholesterol (CHO, lane 5). Cholesterol esters (CE) are also indicated. Right panels depict the area of spots for CE, TAG, and CHO in control and TB-PE in the presence or not of Sz treated MDM (*n* = 2). (D) Analysis of ACAT and β-actin protein expression level by Western Blot (left panel) and quantification (right panel; *n* = 3) in MDM treated or not with different concentrations of Stattic (ST) for 2 h and then exposed or not to TB-PE for 24 h. Friedman test followed by Dunn's Multiple Comparison Test: \**p* < 0.05 for TB-PE treated vs Ctl.

in macrophages treated with TB-PE (data not shown and **Figure 6**). Likewise, the depletion of IFN-γ (a STAT1 activating cytokine) from TB-PE did not prevent the accumulation lipid bodies in macrophages (**Figures 3A–C**). Therefore, we consider that our experimental model is dependent of STAT3 instead of STAT1.