PPAR-α Activation Mediates Innate Host Defense through Induction of TFEB and Lipid Catabolism

Yi Sak Kim, Hye-Mi Lee, Jin Kyung Kim, Chul-Su Yang, Tae Sung Kim, Mingyu Jung, Hyo Sun Jin, Sup Kim, Jichan Jang, Goo Taeg Oh, Jin-Man Kim and Eun-Kyeong Jo
J Immunol April 15, 2017, 198 (8) 3283-3295; DOI: https://doi.org/10.4049/jimmunol.1601920

Abstract

The role of peroxisome proliferator–activated receptor α (PPAR-α) in innate host defense is largely unknown. In this study, we show that PPAR-α is essential for antimycobacterial responses via activation of transcription factor EB (TFEB) transcription and inhibition of lipid body formation. PPAR-α deficiency resulted in an increased bacterial load and exaggerated inflammatory responses during mycobacterial infection. PPAR-α agonists promoted autophagy, lysosomal biogenesis, phagosomal maturation, and antimicrobial defense against Mycobacterium tuberculosis or M. bovis bacillus Calmette–Guérin. PPAR-α agonists regulated multiple genes involved in autophagy and lysosomal biogenesis, including Lamp2, Rab7, and Tfeb in bone marrow–derived macrophages. Silencing of TFEB reduced phagosomal maturation and antimicrobial responses, but increased macrophage inflammatory responses during mycobacterial infection. Moreover, PPAR-α activation promoted lipid catabolism and fatty acid β-oxidation in macrophages during mycobacterial infection. Taken together, our data indicate that PPAR-α mediates antimicrobial responses to mycobacterial infection by inducing TFEB and lipid catabolism.

This article is featured in In This Issue, p.3003

Introduction

Nuclear receptor peroxisome proliferator–activated receptor α (PPAR-α) is a PPAR isoform that regulates the expression of genes involved in glucose and lipid metabolism as well as inflammatory processes (1–3). All PPAR members belong to the adopted orphan nuclear receptor family and are activated by various endogenous ligands, such as steroids, retinoids, and cholesterol metabolites, saturated and unsaturated fatty acids, as well as pharmacologic agents (4, 5). PPAR-α is a transcription factor that binds to cis-acting DNA elements, i.e., PPAR response elements in the promoter region of numerous target genes (6). Through the regulation of these target genes, PPAR-α acts as a key regulator of energy metabolism, and mitochondrial and peroxisomal function (2, 4, 7). In addition, PPAR-α is responsible for activation of fatty acid β-oxidation (FAO) and ketogenesis and simultaneous inhibition of glycolysis and fatty acid synthesis (2, 8). However, the function of PPAR-α in innate host defense during mycobacterial infection is largely unknown.

Mycobacterium tuberculosis is the major causative pathogen of human tuberculosis (TB), an important infectious disease that causes two million deaths per year worldwide (9). M. tuberculosis is a highly successful pathogen that is capable of surviving and persisting within host phagocytes by inhibiting phagosomal maturation and escape from innate immune effectors (10, 11). In addition, pathogenic mycobacteria use host lipids to enter and multiply within non-maturing parasitophorous vacuoles in host cells (12). Host macrophages use several strategies to clear virulent mycobacteria. Autophagy, an intracellular lysosomal degradation process, is an important cell-autonomous defense system involved in innate and adaptive immune responses and thus contributes to host defense against various intracellular microbes including M. tuberculosis and M. bovis bacillus Calmette–Guérin (BCG) (13). Among the transcription factors implicated in autophagy, transcription factor EB (TFEB) is a critical regulator of autophagic activation (14, 15). TFEB transcriptionally regulates numerous genes involved in multiple steps of autophagy, including autophagosomal and lysosomal biogenesis and substrate targeting and degradation (15). Furthermore, TFEB is required for the expression of genes involved in lipid degradation and FAO in mitochondria (16).

In this study we identified a critical role for PPAR-α in host defense against M. tuberculosis and BCG mycobacterial infections, mediated by the ability of PPAR-α to directly activate the expression of genes involved in autophagosomal and lysosomal biogenesis and function. In addition, PPAR-α activation led to upregulation and nuclear translocation of TFEB, which is required for lysosomal biogenesis and antimicrobial responses against M. tuberculosis infection. Furthermore, PPAR-α activation inhibited lipid body formation and promoted the expression of genes involved in FAO in macrophages, suggesting a role for PPAR-α in lipid catabolism during mycobacterial infection.

Materials and Methods

Mice

Ppara−/− mice were kindly provided by Dr. Gonzalez (17). We backcrossed with C57BL/6 mice more than 10 times to establish C57BL/6 congenic Ppara−/− line. The littermate control mice were used for all animal experiments (control groups; Ppara+/+). Mice were housed in specific pathogen free conditions, and maintained on a 12 h light/dark cycle. The littermate control mice used for experiments (control groups; Ppara+/+) were obtained from our inbred C57BL/6 strains. The mice used in the experiments were all 8–10 wk old males. Animal-related procedures were reviewed and approved by the Institutional Animal Care and Use Committee, Chungnam National University School of Medicine (CNUH-014-A0006-1; Daejeon, Korea).

Bone marrow–derived macrophage culture

Bone marrow–derived macrophages (BMDM) were isolated and differentiated after culture for 4–5 d in medium containing 25 ng/ml M-CSF (416-ML; R&D Systems), as described previously (18). The culture medium consisted of DMEM (12-604F; Lonza) supplemented with 10% heat-inactivated FBS (35-015-CV; Lonza), and penicillin–streptomycin-amphotericin B (17-745E; Lonza).

Mycobacterial strains

M. tuberculosis H37Rv was kindly provided by R.L. Friedmann (University of Arizona, Tucson, AZ). M. bovis BCG was obtained from the Korean Institute of Tuberculosis (Osong, Korea). All mycobacterial strain cultures were performed as described previously (18). M. tuberculosis or BCG were grown in Middlebrook 7H9 (Difco, 271310) medium supplemented with 0.5% glycerol, 0.05% Tween-80 (Sigma), and 10% oleic acid, albumin, dextrose, and catalase (Difco, 212240).

For M. tuberculosis– or BCG-expressing enhanced red fluorescent protein (ERFP) strains, we used Escherichia coli–BCG shuttle expression plasmid pMV262-RFP, under the control of the heat shock protein 60 promoter. BCG was transformed with the recombinant plasmid pMV262-RFP by electroporation, then kanamycin-resistant clones were selected in 7H9 broth supplemented with kanamycin (50 μg/ml) for further experiments (19). M. tuberculosis– or BCG-ERFP strains were grown in Middlebrook 7H9 medium supplemented with OADC and kanamycin. All mycobacterial suspensions were aliquoted and stored at −80°C. Midlogarithmic-phase bacteria (OD = 0.6) were used in all assays. The CFU were enumerated on Middlebrook 7H10 agar (BD Biosciences).

Mycobacterial infection in vivo

Mycobacterial infection in vivo was performed as described previously (18). Frozen M. tuberculosis and BCG were thawed and centrifuged, and then their pellets were resuspended in PBS plus 0.05% Tween-80 (PBST) before infection. Ppara+/+ and Ppara−/− mice were intravenously injected with M. tuberculosis (1 × 106 CFU per mouse) or BCG (1 × 107 CFU per mouse) for 14 or 21 d. Mice were maintained in biosafety level 3 laboratory facilities. The experimental procedures were reviewed and approved by the Institutional Animal Care and Use Committee of Bioleaders (Daejeon, Korea). At 14 or 21 d, mice were euthanized and the lungs, spleens, and livers harvested for CFU assay, immunohistochemistry (IHC) staining, and quantitative real-time PCR (qPCR) analysis.

For measurement of the bacterial burden in the lung, spleen, and liver of mice sacrificed at 14 or 21 d after M. tuberculosis or BCG infection, tissues were homogenized in PBST, and serial dilutions of the homogenates were plated on 7H10 agar plates, with colonies counted 14 d later.

Histology and immunohistochemistry

Lung samples were fixed in 10% formalin and embedded in paraffin wax. Paraffin sections (4 μm) were then cut and stained with H&E. Inflammation in lung sections was graded for severity by scanning multiple random fields in 10 sections of each tissue per mouse. An overall histopathological score was assigned to each tissue in each animal based on the extent of granulomatous inflammation as follows: 0 = no lesion, 1 = minimal lesion (1–10% of tissue in section involved), 2 = mild lesion (11–30% involved), 3 = moderate lesion (30–50% involved), 4 = marked lesion (50–80% involved), and 5 = severe lesion (>80% involved), as described previously (20).

For IHC staining, lung paraffin sections (4 μm) were cut and immunostained with Abs specific for cyclooxygenase 2 (COX-2) (ab15191; Abcam), anti-Ly-6G6C Ab (ab2557, NIMP-R14; Abcam), and TNF-α (sc-52746; Santa Cruz). IHC staining tissue slides for COX-2 and neutrophils were imaged using an Aperio ScanScope CS System and confocal images were taken with a Leica TCS SP8 confocal system and processed with the Leica LAS AF Lite program. The histopathological score data used for this study were provided by the Biobank of Chungnam University Hospital, a member of the Korea Biobank Network.

CFU assays

For quantification of intracellular bacteria, CFU assays were performed as described previously (21). M. tuberculosis– or BCG-infected BMDM were then lysed with 0.3% saponin to release intracellular bacteria. The infected lysates were then resuspended vigorously, transferred to screw-capped tubes, and sonicated in a preheated 37°C water bath sonicator (Elma) for 5 min. Aliquots of the sonicates were diluted and each sample was plated separately on 7H10 agar plates and incubated at 37°C in 5% CO2 for 14 to 21 d.

Immunofluorescence of autophagy analysis

Autophagosome formation was assayed by LC3 punctate staining, as described previously (21). For endogenous LC3 puncta or phagosomal maturation analysis, the stimulated cells on coverslips were washed twice with PBS, and fixed with 4% paraformaldehyde for 15 min after permeabilization with 0.25% Triton X-100 for 10 min. The cells were incubated with primary Abs [anti-LC3; MBL International, PM036, anti-LAMP2; Santa Cruz, sc-5571, and anti- Ras-related proteins in brain (Rab) 7; Santa Cruz, sc-6563] at 4°C overnight. Cells were washed twice with PBS to remove excess primary Abs and incubated with fluorescently labeled secondary Abs at room temperature (RT) for 2 h. Nuclei were stained with DAPI (Sigma) for 1 min. After mounting, fluorescence images were acquired using a Leica TCS SP8 confocal system, and MetaMorph Advanced Imaging acquisition software version 7.8 (Molecular Devices).

Autophagic flux analysis using a tandem fluorescent-tagged LC3

For autophagic flux analysis, tandem-tagged tandem fluorescent-tagged LC3 (mCherry-EGFP-LC3) retrovirus was generated. Phoenix amphotropic cells were seeded at 70–80% confluence into a six-well plate and transfected with 1 μg of pBabe-mCherry-GFP-LC3 (22418; Addgene), 0.75 μg of pCL-Eco (12371; Addgene), and 0.25 μg of pDM2.G (12259; Addgene) using Lipofectamine 2000 (Invitrogen). After 18 h, transfection cell culture medium was removed and replaced with fresh medium. The retrovirus-containing medium was harvested at 60 h posttransfection and filtered through a 0.45 μm syringe filter. For autophagic flux analysis, BMDM were transduced with tandem fluorescent-tagged LC3 (mCherry-EGFP-LC3B) retrovirus for 24 h and stimulated with GW7647 (1008613; Cayman Chemical) or Wy14643 (C7081; Sigma). The stimulated cells were placed on coverslips, washed twice with PBS, and fixed with 4% paraformaldehyde for 15 min. After mounting, fluorescence images were acquired using a confocal laser scanning microscope.

Flow cytometry

Flow cytometry analysis of LC3B was performed as described previously (21). GW7647- or Wy14643- stimulated BMDM were washed in fixation and permeabilization solution (BD Biosciences) for 10 min. Cells were incubated with an anti-LC3B Ab (2775; Cell Signaling) for 1 h and a secondary anti-rabbit IgG-Alexa 488 Ab (A11034; Invitrogen) for 30 min on ice. Cells were then washed in PBS, resuspended in FACS buffer and analyzed immediately. The samples were examined using a FACSCanto II flow cytometer (Becton Dickinson). Flow cytometry data were analyzed using the FlowJo software (Tree Star).

Immunoblotting

Immunoblotting was performed as described previously (22). For immunoblotting, cells were lysed in RIPA (10 mM Tris-HCl at pH 8, 1 mM EDTA, 140 mM NaCl, 0.1% SDS, 0.1% sodium deoxycholate, and 1% Triton X-100) and a protease inhibitor mixture (Roche). The cell suspension was incubation and centrifuged at 14,000 × g for 10 min at 4°C. The protein supernatant was collected and concentration was measured by BCA assay (Pierce). The protein extracts were boiled in 1× SDS sample buffer, loaded onto SDS-PAGE, and then transferred to polyvinylidene difluoride membranes (IPVH00010; Millipore). Specific Abs were used in this study, as follows: LC3 (L8918; Sigma), PPAR-α (ab45859; Abcam), UVRAG (ab70807; Abcam), TFEB (A303-673A; Bethyl Laboratories), and Actin (sc-1616-R; Santa Cruz). Protein signals were visualized using ECL solution (WBKL S0500; Millipore) and detected with an UVitec Alliance mini-chemiluminescence device (UVitec, Rugby, U.K.).

RNA extraction, semiquantitative RT-PCR, and qPCR

RNA extraction, semiquantitative RT-PCR, and qPCR were performed as described previously (18). Briefly, total RNA from cells was isolated using TRIzol reagent (15596-026; Thermo Fisher Scientific) and used for synthesis of cDNA with Superscript II reverse transcriptase (Invitrogen). For semiquantitative RT-PCR reactions involved 30 cycles of annealing, extension, and denaturation as described previously (22). Reactions were run on a Veriti 96-Well Thermal Cycler (Thermo Fisher). The PCR products were analyzed by electrophoresis on 1.5% agarose gel.

qPCR was carried out using cDNA, primers, and SYBR Green master mix (Qiagen). Reactions were run on a Rotor-Gene Q 2plex system (Qiagen, Germany). To analyze qPCR data, we performed relative quantification using the 2ΔΔ threshold cycle (Ct) method with Gapdh as an internal control gene. Data were expressed as relative fold changes. The following primers were used: Tnf forward 5′-GGTGCCTATGTCTCAGCCTCTT-3′, reverse 5′-GCCATAGAACTGATGAGAGGGAG-3′; Il6 5′-TACCACTTCACAAGTCGGAGGC-3′, reverse 5′-CTGCAAGTGCATCATCGTTGTTC-3′; Il1b forward 5′-TGGACCTTCCAGGATGAGGACA-3′, reverse 5′-GTTCATCTCGGAGCCTGTAGTG-3′; Becn1 forward 5′-AGGCGAAACCAGGAGAGAC-3′, reverse 5′-CCTCCCCGATCAGAGTGAA-3′; Atg7 forward 5′-CCTGTGAGCTTGGATCAAAGGC-3′, reverse 5′-GAGCAAGGAGACCAGAACAGTG-3′; Lc3b forward 5′-TTATAGAGCGATACAAGGGGGAG-3′, reverse 5′-CGCCGTCTGATTATCTTGATGAG-3′; Lamp1 forward 5′-CAGCACTCTTTGAGGTGAAAAAC-3′, reverse 5′-CCATTCGCAGTCTCGTAGGTG-3′; Lamp2 forward 5′-GAGCAGGTGCTTTCTGTGTCTAG-3′, reverse 5′-GCCTGAAAGACCAGCACCAACT-3′; Uvrag forward 5′-GACTTTGGAATAATGCCGGATCG-3′, reverse 5′-CAGCCCATCCAGGTAGACTTT-3′, Vps11 forward 5′-AAAAGAGAGACGGTGGCAATC-3′, reverse 5′-AGCCCAGTAACGGGATAGTTG-3′; Vps34 forward 5′-CCTGGACATCAACGTGCAG-3′, reverse 5′-TGTCTCTTGGTATAGCCCAGAAA-3′; Tfeb forward 5′-CGCCTGGAGATGACTAACAAGC-3′, reverse 5′-GGCAACTCTTGCTTCACCACCT-3′; Lipa forward 5′-TGTTCGTTTTCACCATTGGGA-3′, reverse 5′-CGCATGATTATCTCGGTCACA-3′; Cd36 forward 5′-CCCTTGGCAACCAACCACAA-3′, reverse 5′-AACCATCCACCAGTTGCTCC-3′; Cpt1a forward 5′-CTCCGCTCGCTCATTCCG-3′, reverse 5′-CACACCCACCACCACGATAA-3′; Acadl forward 5′-CATCGCAGAGAAACATGGCG-3′, reverse 5′-TGGCTATGGCACCGATACAC-3′; Actb forward 5′-CATTGCTGACAGGATGCAGAAGG-3′ reverse 5′-TGCTGGAAGGTGGACAGTGAGG-3′; Gapdh forward 5′-CATCACTGCCACCCAGAAGACTG-3′, reverse 5′-ATGCCAGTGAGCTTCCCGTTCAG-3′.

ELISA analysis

BMDM were treated as indicated and processed for analysis by sandwich ELISA. Cell culture supernatants were analyzed using a Mouse BD OptEIA Set ELISA Kit (BD Biosciences) to detect TNF-α (558534), IL-6 (555240), and IL-1β (559603). All assays were performed as recommended by the manufacturers.

Lentiviral short hairpin RNA generation and transduction

Generation of lentiviral short hairpin RNA (shRNA) and titration was performed as described previously (22). For silencing target genes in BMDM, the pLKO.1-based lentiviral shRNA construct, TFEB (sc-38510-SH; Santa Cruz), was used. Briefly, the lentivirus was generated by transfection of HEK293T cells with pLKO puro.1 or target shRNA plasmids and packaging plasmids (pMDLg/pRRE, pRSV-REV, and pMD2.VSV-g; Addgene) using Lipofectamine 2000 (Invitrogen). After 72 h, lentivirus-containing supernatants were collected and a number of lentiviruses titrated. For lentivirus transduction, BMDM in medium containing 10% FBS were infected with lentiviral vectors in the presence of 8 μg/ml polybrene (Sigma). After 2 d, cells were harvested and target gene knockdown analyzed.

Immunofluorescence microscopy of nuclear translocation of TFEB

Translocation of TFEB into the nucleus was detected using immunofluorescence staining. Briefly, Ppara+/+ and Ppara−/− BMDM were stimulated with GW7647 or Wy14643 for 1 h and then fixed with 4% paraformaldehyde in PBS for 10 min. Cells were permeabilized with 0.25% Triton X-100 in PBS for 10 min and stained with anti-TFEB (A303-673A; Bethyl Laboratories) at 4°C overnight. The cells were then washed in PBS, and incubated with anti-rabbit Alexa Fluro 488 (A-11008, for 2 h; Invitrogen) at RT. Nuclei were stained by incubation with DAPI for 1 min. After mounting, fluorescence images were acquired using a confocal laser-scanning microscope.

Lipid body staining

Ppara+/+ and Ppara−/− BMDM were infected with M. tuberculosis– or BCG-ERFP and stimulated with GW7647 or Wy14643 for 24 h. For neutral lipids were stained using BODIPY 493/503, as previously described (23). Cells were fixed in 4% (v) paraformaldehyde and neutral lipids were stained using 5 μg/ml of BODIPY 493/503 (D-3922; Molecular Probes) for 30 min at RT. Nuclei were stained by incubation with DAPI for 1 min. After mounting, fluorescence images were acquired using a confocal laser–scanning microscope.

Real-time measurement of oxygen consumption rate assay

Real-time analysis of the oxygen consumption rate (OCR) was performed as described previously (22). Briefly, BMDM were plated in XF-24 cell culture microplates (2–3 × 105 cells per well in 200 μl) and then stimulated with GW7647 or Wy14643. After 18 h, the medium was removed and cells were washed then analyzed in XF Running Buffer according to the manufacturer’s instructions to analyze real-time values of OCR. Where indicated, OCR was analyzed in response to 2 mg/ml oligomycin A (75351; Sigma), 5 mM carbonyl cyanide 3-chlorophenylhydrazone (C2759; Sigma), and 2 mM rotenone (R8875; Sigma). The cells were analyzed using an XF-24 Extracellular Flux Analyzer (Seahorse Bioscience).

NF-κB–luciferase reporter assays

NF-κB–luciferase reporter assays were performed as described previously (24). Briefly, BMDM were transduced with NF-κB–luciferase adenovirus (Genetransfer Vector Core) for 36 h, and infected with M. tuberculosis for 6 h. M. tuberculosis–infected cells were washed three times in PBS and cell extracts were prepared by adding 100 μl of 1× Passive Reporter Lysis Buffer (Promega, Madison, WI). Luciferase activity was measured using the Luciferase Assay System (Promega), according to the manufacturer’s instructions.

Statistical analysis

Data are represented as the mean ± SD. All analyses were performed using SPSS 22.0 and GraphPad Prism 5.0. Statistical analysis was performed using an unpaired two-tailed Student t test indicated for comparison between groups. A p value <0.05 was considered significant.

Results

PPAR-α is essential for antimicrobial responses and inhibition of excessive inflammatory responses to mycobacterial infection in macrophages and in vivo

The function of PPAR-α in innate host defense is largely unknown. To evaluate its role in host defense against mycobacterial infection, we first investigated whether PPAR-α is essential for antimicrobial effects against M. tuberculosis and BCG infection in BMDM. To this end, wild-type (Ppara+/+) and PPAR-α–knockout (Ppara−/−) BMDM were infected with M. tuberculosis or BCG at various multiplicities of infection (MOIs). Macrophage PPAR-α gene expression was induced after M. tuberculosis infection (data not shown). As shown in Fig. 1A and 1B, the intracellular survivals of M. tuberculosis and BCG were significantly increased in BMDM from Ppara−/− mice, compared with those from Ppara+/+ mice, in an MOI-dependent manner.

FIGURE 1.
FIGURE 1.

PPAR-α is required for antimicrobial host defense against mycobacterial infection. (A and B) Ppara+/+ and Ppara−/− BMDM were infected with M. tuberculosis [MOI = 1, 5, or 10; (A)] or BCG [MOI = 1, 5, or 10; (B)] for 3 d and then intracellular bacterial loads determined. (C) Ppara+/+ and Ppara−/− mice (n = 8 per group) were infected intravenously with M. tuberculosis (1 × 106 CFU). Mice were sacrificed after 21 d of M. tuberculosis infection, and the bacterial loads in lung, liver, and spleen were determined by CFU assay. (D) Ppara+/+ and Ppara−/− mice (n = 6–8 per group) were infected intravenously with BCG (1 × 107 CFU). Mice were sacrificed after 14 or 21 d of BCG infection, and the bacterial loads in the lung were determined by CFU assay. (E and F) Representative H&E stained images in lung tissue from Ppara+/+ and Ppara−/− mice (n = 7 per group) infected with BCG or PBST (n = 3) for 21 d. (E) Representative images. Scale bar, 200 μm. (F) Quantitative analysis of histopathology scores in 10 random fields. (G and H) Neutrophil infiltration (G) and COX-2 expression (H) in lung tissues from Ppara+/+ and Ppara−/− mice (n = 8 per group) infected with BCG or PBST (n = 3) for 21 d. Representative images. Scale bars, 50 μm (G) or 100 μm (H). (I and J) Quantitative scoring of brown color neutrophils (I) and COX-2+ cells (J) were counted from eight random fields. (K and L) Ppara+/+ and Ppara−/− mice (n = 8 per group) were infected intravenously with M. tuberculosis (K) or BCG (L) for 21 d. qPCR analysis of Tnf and Il6 mRNA levels in lung (left) and spleen (right) tissue samples. (M) Ppara+/+ and Ppara−/− mice (n = 8 per group) were infected intravenously with BCG for 21 d. Representative immunofluorescence images of TNF-α in lung tissues. Scale bar, 50 μm. Data shown are mean ± SD of three independent experiments. **p < 0.01, ***p < 0.001, compared with the control condition (two-tailed Student t test). Mtb, M. tuberculosis.

To further examine the effect of PPAR-α deficiency on antimycobacterial immunity in vivo, Ppara+/+ and Ppara−/− mice were infected intravenously with M. tuberculosis or BCG. As shown in Fig. 1C and 1D, the mycobacterial loads were significantly increased in the lung (Fig. 1D), liver, and spleen in Ppara−/− mice relative to Ppara+/+ mice at days 14 and 21 after M. tuberculosis or BCG infection. In addition, Ppara−/− mice responded to i.v. BCG challenge with significantly increased granuloma-like lesions in lungs compared with Ppara+/+ mice (Fig. 1E, 1F). In lung granulomas after BCG challenge, neutrophil infiltration and COX-2 levels were increased in Ppara−/− mice compared with Ppara+/+ mice (Fig. 1G–J). We then assessed expression of the proinflammatory cytokines TNF-α and IL-6 in lung tissues from Ppara+/+ and Ppara−/− mice. Tnf and Il6 mRNA levels were markedly elevated in the lung and spleen tissues of Ppara−/− mice compared with the corresponding tissues of Ppara+/+ mice (Fig. 1K, 1L). TNF-α production was also significantly elevated in the lung tissues of Ppara−/− mice, compared with Ppara+/+ mice, after mycobacterial infection (Fig. 1M). These results indicate that PPAR-α is required for antimicrobial host defense and attenuates systemic inflammation during mycobacterial infection.

PPAR-α agonists enhance antimicrobial defense, but suppress inflammatory responses against mycobacterial infection in BMDM in a PPAR-α–dependent manner

We assessed whether PPAR-α agonists enhance antimicrobial responses in macrophages from Ppara+/+ and Ppara−/− mice. Ppara+/+ and Ppara−/− BMDM were infected with M. tuberculosis or BCG for 3 d with or without various PPAR-α agonists (GW7647 or Wy14643), and the survival of intracellular mycobacteria was determined by a CFU assay. PPAR-α agonists (GW7647 and Wy14643) (3, 25) exhibited significant killing effects against M. tuberculosis (Fig. 2A, 2B) and BCG (Fig. 2C, 2D) in Ppara+/+ BMDM in a dose-dependent manner. The intracellular survival levels of M. tuberculosis and BCG were similarly inhibited in Ppara+/+ BMDM by PPAR-α agonists 2 d postinfection (data not shown). In contrast, the PPAR-α agonists did not exert antimycobacterial effects in Ppara−/− BMDM (Fig. 2A–D), suggesting that the antimicrobial activities were dependent on PPAR-α activation.

FIGURE 2.
FIGURE 2.

PPAR-α agonists activate antimicrobial responses but inhibit inflammatory responses against mycobacterial infection in macrophages. (A–F) Ppara+/+ and Ppara−/− BMDM were infected with M. tuberculosis (MOI = 5) or BCG (MOI = 5) for 4 h and then stimulated with GW7647 (2, 10, or 20 μM) or Wy14643 (20, 50, or 100 μM). (A–D) Intracellular bacterial loads were determined by CFU assay after 3 d of infection. (E) Cells were lysed and subjected to qPCR analysis (for 6 h) of Tnf and Il6. (F) TNF-α and IL-6 in supernatants were determined by ELISA (for 18 h). Data shown are mean ± SD of three independent experiments. **p < 0.01, ***p < 0.001, compared with the control condition (two-tailed Student t test). GW, GW7647; ns, nonsignificant; SC, solvent control (0.1% DMSO); Wy, Wy14643.

PPAR-α has been reported to control inflammatory responses (26, 27). However, the effects of PPAR-α in inflammatory responses during mycobacterial infection remain unclear. We thus compared the expression of three proinflammatory cytokines (TNF-α, IL-6, and IL-1β) between Ppara+/+ and Ppara−/− BMDM. Tnf, Il6, and Il1b mRNA levels were significantly increased in Ppara−/− BMDM after M. tuberculosis infection (Supplemental Fig. 1A). Consistent with these findings, the reporter gene activities of NF-κB were markedly increased in Ppara−/− BMDM after M. tuberculosis infection (Supplemental Fig. 1B). In addition, the production of proinflammatory cytokines was markedly increased in Ppara−/− compared with Ppara+/+ BMDM after M. tuberculosis infection (Supplemental Fig. 1C).

We then examined the effects of PPAR-α agonists on proinflammatory cytokine production in BMDM after mycobacterial infection. As shown in Fig. 2E, PPAR-α agonists (GW7647 and Wy14643) caused a dose-dependent inhibition of Tnf and Il6 mRNA levels in Ppara+/+ after 6 h of infection with M. tuberculosis or BCG. Similar to these findings, proinflammatory cytokine production in response to M. tuberculosis or BCG was dose-dependently decreased by PPAR-α agonists (GW7647 and Wy14643) in BMDM after 18 h of M. tuberculosis or BCG infection (Fig. 2F). However, PPAR-α agonists did not modulate proinflammatory cytokine mRNA and protein levels in Ppara−/− BMDM after M. tuberculosis or BCG infection (Fig. 2E, 2F). These data suggest that PPAR-α agonists promote innate antimicrobial defense, but negatively regulate M. tuberculosis– or BCG-induced production of proinflammatory cytokines in BMDM in a PPAR-α–dependent manner.

PPAR-α activation enhances autophagy and expression of autophagy-related genes in macrophages

Previous studies showed that pharmacological activation of PPAR-α promotes hepatic autophagy through transcriptional activation of autophagy-related genes (Atgs) (25). To determine the autophagy-inducing capacity of PPAR-α agonists in macrophages, Ppara+/+ and Ppara−/− BMDM were treated with PPAR-α agonists, and autophagic activation was evaluated by assessing LC3 punctae formation by immunofluorescence analysis. As shown in Fig. 3A and 3B, the number of LC3 punctae was significantly increased in Ppara+/+ BMDM after treatment with PPAR-α agonists (GW7647 and Wy14643), when compared with that in Ppara−/− BMDM. In addition, PPAR-α agonists (GW7647 and Wy14643) robustly upregulated the lipidated form of LC3 in Ppara+/+ BMDM, as determined by immunoblotting of autophagosomal membrane–associated LC3-II fractions (Fig. 3C, 3D). In contrast, the level of lipidated LC3 was not significantly upregulated in Ppara−/− BMDM (Fig. 3C, 3D). Flow cytometric analysis using an LC3B-specific Ab revealed an increase in LC3B expression, which is correlated with autophagosomal formation (21), by PPAR-α agonists in Ppara+/+ BMDM (Fig. 3E, 3F). However, PPAR-α–induced LC3B expression was not significantly upregulated in Ppara−/− BMDM, when compared with that in Ppara+/+ BMDM (Fig. 3E, 3F). We next monitored autophagic flux in macrophages using a mCherry-EGFP-LC3B and measured mRFP-GFP-LC3 delivery to lysosomes in BMDM treated with the PPAR-α agonists. As shown in Fig. 3G, the number of red punctae was increased in Ppara+/+ BMDM, whereas only the yellow punctae were observed in Ppara−/− BMDM, in response to PPAR-α agonists.

FIGURE 3.
FIGURE 3.

PPAR-α activation enhances autophagy and increases autophagy-related gene expression in macrophages. (A–F) Ppara+/+ and Ppara−/− BMDM were stimulated with GW7647 (10 μM) or Wy14643 (50 μM). (A and B) Alexa488-conjugated LC3 (green) and DAPI (blue) were detected by confocal microscopic analysis at 8 h of GW7647 or Wy14643 stimulation. (A) Representative images. Scale bar, 5 μm. (B) Statistical analysis of LC3 punctate foci per cell. (C and D) Cells were stimulated with GW7647 or Wy14643 for the time as indicated. Cell lysates were subjected to immunoblot analysis of LC3 and Actin. The densitometry values for LC3 II were normalized to Actin in each lower panel. (E and F) Flow cytometric analysis of LC3B expression at 24 h of GW7647 or Wy14643 treatment (E). (F) Statistical analysis of LC3B by flow cytometry. (G) Ppara+/+ and Ppara−/− BMDM were transduced with a retrovirus expressing a tandem LC3 plasmid (mCherry-EGFP-LC3B) and stimulated with GW7647 or Wy14643 for 24 h. Cells were fixed, and mCherry- or EGFP-expressing LC3 was detected by confocal microscopy. Scale bar, 5 μm. (H) Ppara+/+ and Ppara−/− BMDM were stimulated with GW7647 or Wy14643 for 6 or 18 h. Becn1, Atg7, Lc3b, Lamp1, Lamp2, and Rab7 mRNA levels were determined by qPCR analysis. Representative confocal microscopic images from three independent samples are shown, with each experiment including at least 50 cells scored from seven random fields (A and G). Data shown are mean ± SD of three independent experiments. **p < 0.01, ***p < 0.001, compared with the control condition (two-tailed Student t test). GW, GW7647; SC, solvent control (0.1% DMSO); Wy, Wy14643.

A number of Atgs identified in yeasts and mammals are associated with key functions of the autophagy pathway (28). In addition, PPAR-α activation leads to transcriptional activation of numerous Atgs by directly binding to their promoter regions (25). We thus examined whether PPAR-α activation increases transcriptional activation of Atgs in macrophages. Application of the PPAR-α agonists significantly increased Becn1, Atg7, Lc3b, Rab7, lysosomal-associated membrane protein (Lamp)1, and Lamp2 mRNA levels in Ppara+/+ BMDM (Fig. 3H). In addition, PPAR-α activation was required for this induction of expression, because the transcriptional activation of these genes was not significantly upregulated in Ppara−/− BMDM, when compared with that in Ppara+/+ BMDM (Fig. 3H). Thus, these data indicate that PPAR-α activation robustly enhances autophagosome formation and autophagic flux in BMDM.

PPAR-α is required for phagosomal maturation in macrophages during mycobacterial infection

To determine the importance of PPAR-α–induced autophagic activation during mycobacterial infection, we assessed the effects of PPAR-α on the colocalization of M. tuberculosis phagosomes with autophagosomes or lysosomes in macrophages infected with M. tuberculosis–ERFP. As shown in Fig. 4A, treatment of M. tuberculosis–infected BMDM with PPAR-α agonists resulted in increased colocalization of the autophagosomal marker LC3 with M. tuberculosis phagosomes. Immunofluorescence analyses using an anti-LAMP-2A Ab also showed an increase in the colocalization of M. tuberculosis and BCG phagosomes with LAMP-2A-positive lysosomes in Ppara+/+ BMDM after treatment with the PPAR-α agonists (Fig. 4B, Supplemental Fig. 2). The PPAR-α agonists led to markedly increased amounts of LC3+ autophagosomes and lysosomes in Ppara+/+ BMDM infected with M. tuberculosis or BCG (Fig. 4C, 4D, Supplemental Fig. 2). However, these responses were not significantly up-regulated in Ppara−/− BMDM (Fig. 4C, 4D, Supplemental Fig. 2). In addition, the fusion of mycobacterial phagosomes containing M. tuberculosis with late endosomes and lysosomes was arrested (29). Rab7 functions as a checkpoint in the regulation of phagosomal maturation upon recruitment into M. tuberculosis– or BCG-containing phagosomes (30, 31). Because application of PPAR-α agonists led to an increase in Rab7 expression (Fig. 3H), we determined whether PPAR-α agonists also enhance the recruitment of Rab7 into phagosomes containing M. tuberculosis. As shown in Fig. 4E and 4F treatment of BMDM with PPAR-α agonists increased Rab7 recruitment into M. tuberculosis–containing phagosomes. Therefore, PPAR-α is required for phagosomal maturation and recruitment of Rab7 into M. tuberculosis phagosomes in macrophages during mycobacterial infection.

FIGURE 4.
FIGURE 4.

PPAR-α is required for phagosomal maturation in macrophages during mycobacterial infection. (A–D) Ppara+/+ and Ppara−/− BMDM were infected with M. tuberculosis–ERFP (MOI = 5) for 4 h and then stimulated with GW7647 (10 μM) or Wy14643 (50 μM) for 24 h. (A) M. tuberculosis–ERFP (red), Alexa-488 conjuated-LC3 (green), and DAPI (blue) were detected by confocal microscopy. Scale bar, 5 μm. (C) Statistical analysis of M. tuberculosis–ERFP and LC3 colocalization per cell containing 200 internalized mycobacteria. (B) M. tuberculosis-ERFP (red), Alexa488-conjuated LAMP2 (green), and DAPI (blue) were detected by confocal microscopy. Scale bar, 5 μm. Representative three-dimensional reconstructed Z-stack immunofluorescence images are shown in each lower panel. (D) Statistical analysis of M. tuberculosis and LAMP2 colocalization per cell containing 200 internalized mycobacteria. Representative confocal microscopic images from three independent samples are shown, with each experiment including at least 100 cells scored from seven random fields (A–D). (E and F) Ppara+/+ and Ppara−/− BMDM were infected with M. tuberculosis–ERFP (MOI = 5, for 4 h) and then stimulated with GW7647 (10 μM) or Wy14643 (50 μM) for 24 h. (E) M. tuberculosis–ERFP (red), Alexa488-conjugated LAMP2 (green), and DAPI (blue) were detected by confocal microscopy. Scale bar, 5 μm. (F) Statistical analysis of M. tuberculosis–ERFP and Rab7 colocalization. Representative confocal microscopic images from three independent samples are shown, with each experiment including at least 50 cells scored from seven random fields. Data shown are mean ± SD of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, compared with the control condition (two-tailed Student t test). GW, GW7647; SC, solvent control (0.1% DMSO); Wy, Wy14643; ns, nonsignificant.

PPAR-α agonists enhance the expression and nuclear translocation of TFEB, which is essential for induction of autophagy-related genes and lysosomal biogenesis in macrophages

Because PPAR-α was found to be essential for the expression of multiple Atgs in BMDM (Fig. 3H), we evaluated the effects of PPAR-α on the transcriptional activation and nuclear translocation of TFEB. As shown in Fig. 5A–C, treatment with PPAR-α agonists led to robust expression (Fig. 5A) and rapid nuclear translocation of TFEB (Fig. 5B, 5C). Although PPAR-α stimulation significantly increased translocation of cytosolic TFEB to the nucleus in Ppara+/+ BMDM, this was markedly decreased in Ppara−/− BMDM (Fig. 5B, 5C). We next examined the role of TFEB in PPAR-α–mediated induction of genes involved in autophagosomal formation and maturation into autolysosomes. As expected, PPAR-α activation markedly increased the transcriptional activation of several genes involved in autophagosomal and autolysosomal formation (Uvrag, Lc3b, Vps11, and Vps34) in BMDM at 6 and 18 h. Importantly, lentiviral-mediated knockdown of TFEB significantly decreased the transcriptional activation of these genes (Fig. 5D), suggesting that TFEB is required for the expression of Atgs in response to PPAR-α activation.

FIGURE 5.
FIGURE 5.

PPAR-α-induced TFEB expression is essential for autophagy-related gene induction and lysosomal biogenesis in macrophages. (A–C) Ppara+/+ and Ppara−/− BMDM were stimulated with GW7647 (10 μM) or Wy14643 (50 μM) for 6 (A) or 18 h (A–C). (A) Tfeb expression level was measured by qPCR analysis. (B) Immunofluorescence microscopic analysis of TFEB nuclear translocation. Alexa488-conjugated TFEB (green) and DAPI (blue) were detected by confocal microscopic analysis. Scale bar, 10 μm. (C) Quantitative analysis of TFEB nuclear translocation. (D–H) BMDM were transduced with shTFEB–expressing lentivirus and then stimulated with GW7647 or Wy14643 for 6 (D) or 18 h (D, G, and H). (D) Uvrag, Lc3b, Vps11, and Vps34 expression levels were detected by qPCR analysis. (E and F) Alexa 488-conjugated LC3B was analyzed by flow cytometry. Representative histograms (E) and statistical analysis of LC3B expression (F). (G and H) Alexa488-conjugated LAMP2 (green) and DAPI (blue) were detected by confocal microscopic analysis. (G) Representative of images. Scale bar, 5 μm. (H) Statistical analysis of LAMP2 punctate foci per cell. Representative confocal microscopic images from three independent samples are shown, with each experiment including at least 50 cells scored from seven random fields (B and G). Data shown are mean ± SD of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, compared with the control condition (two-tailed Student t test). GW, GW7647; SC, solvent control (0.1% DMSO); Wy, Wy14643.

We next investigated the effect of TFEB on PPAR-α–induced autophagic activation in BMDM. As shown in Fig. 5E and 5F, knockdown of TFEB using lentiviral shRNA targeting TFEB significantly decreased the LC3B expression in BMDM after treatment with GW7647 or Wy14643. Moreover, UVRAG protein levels in BMDM decreased in response to treatment with GW7647 or Wy14643 (data not shown). We further assessed the role of TFEB in PPAR-α–induced LAMP2 expression and puncta formation in BMDM. Silencing of TFEB led to significant inhibition of LAMP2 expression and puncta formation in BMDM (Fig. 5G, 5H). These data collectively suggest that PPAR-α activation results in the induction and nuclear translocation of TFEB, which are required for expression of autolysosome-related genes, autophagic activation, and lysosomal biogenesis, in BMDM.

PPAR-α–induced TFEB expression enhances phagosomal maturation and antimicrobial responses but attenuates inflammatory responses during mycobacterial infection

We elucidated the role of TFEB in PPAR-α–mediated phagosomal maturation and antimicrobial responses in BMDM during mycobacterial infection. Similar to the findings described above (Fig. 4A, 4B), PPAR-α stimulation resulted in a significant increase in colocalization of M. tuberculosis phagosomes with the autophagosomal marker LC3 (Fig. 6A, 6B) and lysosomal marker LAMP2 (Fig. 6C, 6D) in BMDM transduced with nonspecific lentiviral shRNA. Silencing of TFEB by lentiviral shRNA specific to Tfeb (shTFEB) markedly inhibited PPAR-α–induced colocalization of M. tuberculosis phagosomes with autophagosomes and lysosomes in BMDM (Fig. 6A–D). It was noted that the PPAR-α-mediated increases in LC3 and LAMP2 levels, representing autophagosomal and lysosomal biogenesis, respectively, in nonspecific lentiviral shRNA–transduced/M. tuberculosis–infected BMDM were significantly decreased in shTFEB-transduced BMDM (Fig. 6A, 6C). Additionally, lentiviral knockdown of TFEB significantly suppressed the PPAR-α–induced antimicrobial responses against M. tuberculosis infection in BMDM (Fig. 6E, 6F). We further investigated the role of TFEB in the regulation of inflammatory responses in BMDM during mycobacterial infection. As shown in Fig. 6G and 6H, silencing of TFEB increased production of the proinflammatory cytokines TNF-α and IL-6 in BMDM in response to M. tuberculosis or BCG. These data suggest that TFEB is required for PPAR-α–mediated phagosomal maturation and killing of mycobacteria as well as the suppression of proinflammatory responses in BMDM during M. tuberculosis infection.

FIGURE 6.
FIGURE 6.

PPAR-α-induced TFEB enhances phagosomal maturation and antimicrobial responses but decreases inflammatory responses during M. tuberculosis infection. (A–D) BMDM were transduced with shTFEB-expressing lentivirus, infected with M. tuberculosis–ERFP (MOI = 5, for 4 h), and then stimulated with GW7647 (10 μM) or Wy14643 (50 μM) for 24 h. (A and B) M. tuberculosis-ERFP, Alexa488-conjugated LC3 and DAPI were detected by confocal microscopy. (A) Representative of images. Scale bar, 5 μm. (B) Statistical analysis of M. tuberculosis-ERFP and LC3 colocalization per cell containing 200 internalized mycobacteria. (C and D) M. tuberculosis-ERFP (red), Alexa488-conjugated LAMP2 (green) and DAPI (blue) were detected by confocal microscopy. (C) Representative of images. Scale bar, 5 μm. (D) Statistical analysis of M. tuberculosis-ERFP and LAMP2 colocalization per cell containing 200 internalized mycobacteria. (E and F) BMDM were transduced with shTFEB-expressing lentivirus, infected with M. tuberculosis (MOI = 5, for 4 h), and then stimulated with GW7647 [(E); 2, 10, or 20 μM] or Wy14643 [(F); 20, 50, or 100 μM] for 3 d. Intracellular bacterial loads were determined by CFU assay. (G and H) BMDM were transduced with shTFEB-expressing lentivirus and infected with M. tuberculosis (G) or BCG [(H); MOI = 5] for 6 or 18 h. TNF-α and IL-6 levels in culture supernatants were determined by ELISA. Representative confocal microscopic images from three independent samples are shown (A and C). Data shown are mean ± SD of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, compared with the control condition (two-tailed Student t test). GW, GW7647; SC, solvent control (0.1% DMSO); Un, uninfected; Wy, Wy14643.

PPAR-α activation is required for lipid catabolism and increased mitochondrial respiratory function in macrophages infected with M. tuberculosis

The accumulation of lipid bodies is a hallmark of the foamy macrophages generated upon virulent mycobacterial infection (32). M. tuberculosis or BCG phagosomes merge with intracellular lipid bodies in foamy macrophages, a secure reservoir in which M. tuberculosis can hide and survive and from which they are released (32–34). Next, we investigated whether PPAR-α activation contributes to the reduction in fatty acid–rich lipid body formation in BMDM infected with M. tuberculosis or BCG. Bodipy 493/503 fluorescence staining revealed an increase in neutral lipid levels in the cytoplasm of M. tuberculosis–infected macrophages (Fig. 7A). Following treatment with PPAR-α agonists, the number and size of lipid bodies were greatly reduced in M. tuberculosis–infected macrophages (Fig. 7A–C). Infection of BMDM with BCG resulted in significantly reduced formation of lipid bodies in the cytosol compared with M. tuberculosis infection (Supplemental Fig. 3). Similar to M. tuberculosis infection, PPAR-α stimulation led to a significant reduction in lipid body number and size in BCG-infected BMDM (Supplemental Fig. 3). Importantly, the PPAR-α–mediated reduction in lipid body formation was absent in Ppara−/− BMDM during M. tuberculosis or BCG infection (Fig. 7A–C, Supplemental Fig. 3). We then quantified the cellular oxygen consumption rate as a measure of mitochondrial respiration and FAO in unstimulated and GW7647- or Wy14643-treated BMDM. PPAR-α agonist treatment increased the basal and maximum oxygen consumption rates, both of which were significantly decreased in Ppara−/− BMDM (Fig. 7D).

FIGURE 7.
FIGURE 7.

PPAR-α is required for lipid catabolism and mitochondrial respiratory function in macrophages during mycobacterial infection. (A–C) Ppara+/+ and Ppara−/− BMDM were infected with M. tuberculosis–ERFP (MOI = 5, for 4 h) then stimulated with GW7647 (10 μM) or Wy14643 (50 μM) for 24 h. M. tuberculosis–ERFP (red) and Bodipy 493/503 (green) were detected by confocal microscopy. (A) Representative images. Scale bar, 5 μm. (B) Representative three-dimensional reconstructed Z-stack immunofluorescence images of lipid droplets and M. tuberculosis–ERFP. Scale bar, 5 μm. (C) Statistical analysis of lipid droplets. (D) Ppara+/+ and Ppara−/− BMDM were stimulated with GW7647 or Wy14643 for 18 h. Real-time measurement of the oxygen consumption rate by sequential treatment with oligomycin, carbonyl cyanide 3-chlorophenylhydrazone (CCCP), and rotenone. (E) Ppara+/+ and Ppara−/− BMDM were infected with M. tuberculosis (MOI = 5, for 4 h) and stimulated with GW7647 or Wy14643 for 24 h. Lipa, Cd36, Cpt1a, and Acadl mRNA levels were determined by quantitative PCR analysis. Representative confocal microscopic images from four independent samples are shown, with each experiment including at least 50 cells scored from seven random fields (A and B). Data shown are mean ± SD of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, compared with the control condition (two-tailed Student t test). GW, GW7647; SC, solvent control (0.1% DMSO); Un, uninfected; Wy, Wy14643.

In addition, PPAR-α agonist treatment upregulated the expression of genes involved in lipid uptake, lipolysis, and FAO (Lipa, lysosomal acid lipase; Cd36, lipid uptake; Cpt1a and Acadl, lipid metabolism and FAO) in uninfected (Supplemental Fig. 4) or M. tuberculosis–infected Ppara+/+ BMDM (Fig. 7E). In addition, the transcriptional activation of these genes was markedly abrogated in Ppara−/− BMDM, when compared with that in Ppara+/+ BMDM (Fig. 7E). Collectively, these data indicate that PPAR-α activation is required for lipid catabolism, increased mitochondrial respiratory function, and upregulation of FAO in BMDM during mycobacterial infection.

Discussion

The findings of this work suggest that PPAR-α is required for innate host responses in macrophages to mycobacterial infection. They also suggest that PPAR-α is a critical and master regulator of host antimycobacterial responses through transcriptional autophagic activation, lysosomal biogenesis, promotion of phagosomal maturation, and induction of TFEB in macrophages. We discovered that TFEB mediates innate host defense against mycobacterial infection by enhancing Atg activation, lysosomal biogenesis, phagosomal maturation, and transcriptional activation of genes involved in FAO. Our studies have broad implications for molecular regulation of the autophagy pathway and lysosomal biogenesis by which PPAR-α activation reinforces antimicrobial host defense and prevents infectious diseases.

Autophagy is an intracellular degradative process with various homeostatic roles, including functions in innate immune regulation and cell-autonomous defense against intracellular microbes including M. tuberculosis (5). Atgs exert a concerted effect in autophagy, i.e., autophagosome initiation and elongation and lysosomal functions (28). Various Atgs play diverse roles in innate and inflammatory responses, such as the eradication of intracellular pathogens, murine gammaherpesvirus 68 reactivation from latency, Ag presentation, and innate immune signaling in a cell type–specific manner (28, 35, 36). Despite these findings, research on the molecular mechanisms of Atg regulation in macrophages during infection is in its infancy. Advances have recently been made toward our understanding of the transcriptional regulation of Atgs that function in autophagic processes and numerous physiological responses, including innate host defense against mycobacterial infection. As an example, recent studies have shown that activation of NR1D1 (an adopted orphan nuclear receptor subfamily 1, group D, member 1) enhances autophagy and the expression of LAMP1 and TFEB, to promote antimycobacterial effects in human macrophages (37). We reported previously that the AMPK-PGC1α pathway is essential for transcriptional activation of Atgs, including Atg5, Becn1, and Atg7, in macrophages via C/EBP-β signaling activation (18). The findings of the current study showed that PPAR-α activation led to activation of autophagy and promoted the expression of numerous Atgs, including Becn1, Atg7, Lc3b, Lamp1, Lamp2, Rab7, and Tfeb, in macrophages. We also found that PPAR-α deficiency led to excessive inflammatory responses in macrophages after M. tuberculosis infection. These data in part support previous reports that pharmacological activation of PPAR-α enhances the transcription of Atgs in the mouse liver (25), and that PPAR-α activation ameliorates hepatic injury and inflammatory responses through autophagic activation (27).

The roles of several nuclear receptors and their mechanisms in the regulation of antimycobacterial immunity have been investigated. Earlier studies revealed an essential role for vitamin d–receptor signaling in the activation of antimicrobial responses in human macrophages via induction of antimicrobial peptide cathelicidin LL-37, production of IL-1β, and enhancement of autophagy (38, 39). A recent study revealed that biallelic retinoic acid–related orphan receptor γ loss-of-function mutations in seven individuals lacking functional RAR-related orphan receptor γ and RAR-related orphan receptor γ T isoforms had a defect in IL-17A/F–producing T cells and impaired IFN-γ responses to mycobacterial infection (40). In addition, a macrophage transcriptomic profiling study showed that M. tuberculosis infection enhanced the levels of aryl hydrocarbon receptor (AHR) and its heterodimeric partners AHR nuclear translocator and RELB, and stimulated the expression of AHR target genes, including IL-1β (41). Another recent study reported that human xenobiotic nuclear receptor pregnane X receptor is essential for prolonging mycobacterial survival, which is mediated by induction of foamy macrophage formation and inhibition of phagolysosomal fusion (42). Importantly, each PPAR family member plays a distinct role in various physiological functions in a context-dependent manner, i.e., depending on ligand affinities, tissue, cell type, and so on (43). Our data are important in suggesting a novel function for PPAR-α in the promotion of antimicrobial responses through autophagic activation and induction of TFEB expression. However, PPAR-γ induced by BCG infection was involved in lipid body formation and PG E2 production in macrophages, thereby potentiating mycobacterial pathogenesis (33). In addition, the host lipid-sensing nuclear receptors PPAR-γ and testicular receptor 4 contributed to generation of foamy macrophages, induction of the anti-inflammatory factor IL-10, and enhancement of mycobacterial survival within macrophages (44). Thus, PPAR-γ and PPAR-α may play opposite roles in antimicrobial host defense against mycobacterial infection.

It was of interest that PPAR-α agonists play an essential role in anti-mycobacterial effects despite their involvement in the inhibition of proinflammatory cytokines such as TNF-α and IL-1β, which are critically involved in host protective immunity during mycobacterial infection (45, 46). Indeed, proinflammatory responses are regarded as a double-edged sword between protective immunity and detrimental inflammatory pathogenesis during mycobacterial infection. Higher IL-1β expression correlated with an increased accumulation of neutrophils in the lung, suggesting a driving cytokine for granulocytic inflammation and TB disease progression (47). In addition, defective recruitment of anti-inflammatory dendritic cells and regulatory T cells resulted in the enhanced susceptibility to Mtb infection and excessive lung inflammation (48). A selective inhibitor for soluble TNF receptors was beneficial for the treatment of mycobacterial infections through neutralization of excessive TNF (49). Extensive studies have reported the roles of PPAR-α agonists in the modulation of innate and adaptive immune responses (50–52). These data thus indicate that PPAR-α agonists may protect hosts against excessive inflammatory responses during mycobacterial infection to promote a balanced inflammatory response.

TFEB acts as a key coordinator of lysosomal function and biogenesis, cellular clearance, and autophagic processes by binding to the promoters of numerous genes involved in the Co-ordinated Lysosomal Expression and Regulation gene network (14, 15, 53). Metabolic signals such as mTORC1 inhibition drive dephosphorylation and nuclear translocation of TFEB, thus promoting the activation of genes involved in lysosomal function and autophagy (14, 53). PPAR-α agonists enhance the transcriptional activation of TFEB via direct binding to and regulation of its promoter regions in brain cells and fly models (54, 55). In addition, TFEB was activated in murine macrophages by Staphylococcus aureus infection and contributed to transcriptional induction of inflammatory cytokines and chemokines (56). Our data revealed a novel function of TFEB in the suppression of mycobacterial survival within macrophages through enhancement of phagosomal acidification. We found that TFEB plays a role in the expression of Atgs and lysosomal biogenesis, but inhibits inflammatory responses, in macrophages during M. tuberculosis or BCG infection. Thus, our findings suggest that TFEB is a key mediator of the anti-mycobacterial function and anti-inflammatory responses of PPAR-α.

Virulent M. tuberculosis infection modulates host lipid homeostasis and induces macrophage differentiation into foamy phenotypes that constitute a nutrient-rich niche for mycobacterial persistence and replication within tuberculous granuloma lesions (32, 34). In addition, lipid-laden mycobacteria exhibit a drug-resistance and dormancy-like phenotype, characteristics associated with latent infection (57). A recent study reported that M. tuberculosis–induced miR-33/33* inhibited FAO and enhanced lipid-body formation in macrophages (23). Importantly, we found that PPAR-α activation promoted lipid catabolism and upregulated the expression of genes involved in FAO in M. tuberculosis–infected macrophages. Moreover, silencing of TFEB downregulated the expression of genes associated with FAO in these macrophages (data not shown). Previous studies also reported that TFEB plays a crucial role in the link between lipophagy and FAO and in the transcriptional regulation of genes involved in several steps of lipid degradation (16). Moreover, silencing of miR-33/33* decreased lipid accumulation and induced lipolysis, which were correlated with increased xenophagy and antimicrobial responses against M. tuberculosis in macrophages (23). Thus, these data suggest that PPAR-α activation induces lipid catabolism to enhance destruction of the foamy refuges of mycobacteria in host cells.

In summary, the findings of this study demonstrate that PPAR-α activation results in the activation of autophagy through direct and indirect induction of Atgs via TFEB, which is essential for lysosomal biogenesis and enhancement of phagosomal maturation to promote host defense against M. tuberculosis infection. In addition, PPAR-α signaling played a major role in lipid catabolism in macrophages, also promoting host defense against mycobacteria. There is growing recognition of adjunctive host-directed therapy for human TB (58). A variety of PPAR-α–activating agents have antioxidant and anti-inflammatory properties, as well as protective effects against metabolic complications (such as diabetes and hyperlipidemia), cell death, and microvascular impairment (7). The potency of PPAR-α in infectious disease models presented in this study, including its mechanisms involving autophagic activation and lipid catabolism, might facilitate development of a novel therapeutic strategy against TB.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank Dr. Hyun-Woo Suh (Chungnam National University) for critical reading of manuscript, Dr. Chul-Ho Lee (Korea Research Institute of Bioscience and Biotechnology) and Yong Woo Back for excellent technical assistance, and Dr. Taeok Bae (University of Indiana) for generously providing reagents.

Footnotes

  • ↵1 Y.S.K. and H.-M.L. are cofirst authors.

  • This work was supported by the Korea Health Technology R&D Project through the Korea Health Industry Development Institute, Ministry of Health and Welfare, Republic of Korea (HI15C0395), by the National Research Foundation grant funded by the Korean government (MSIP) (NRF-2015M3C9A2054326) at Chungnam National University.

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    AHR
    aryl hydrocarbon receptor
    Atg
    autophagy-related gene
    BCG
    bacillus Calmette–Guérin
    BMDM
    bone marrow–derived macrophage
    COX-2
    cyclooxygenase 2
    ERFP
    enhanced red fluorescent protein
    FAO
    fatty acid β-oxidation
    IHC
    immunohistochemistry
    Lamp
    lysosomal-associated membrane protein
    mCherry-EGFP-LC3B
    tandem fluorescent-tagged LC3
    MOI
    multiplicity of infection
    OCR
    oxygen consumption rate
    PBST
    PBS plus 0.05% Tween-80
    PPAR-α
    Ppara, peroxisome proliferator–activated receptor α
    qPCR
    quantitative real-time PCR
    Rab
    Ras-related protein in brain
    RT
    room temperature
    shRNA
    short hairpin RNA
    shTFEB
    lentiviral shRNA specific to Tfeb
    TB
    tuberculosis
    TFEB
    transcription factor EB.

  • Received November 11, 2016.
  • Accepted February 3, 2017.

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