IL-1β Promotes Antimicrobial Immunity in Macrophages by Regulating TNFR Signaling and Caspase-3 Activation

Pushpa Jayaraman, Isabel Sada-Ovalle, Tomoyasu Nishimura, Ana C. Anderson, Vijay K. Kuchroo, Heinz G. Remold and Samuel M. Behar
J Immunol April 15, 2013, 190 (8) 4196-4204; DOI: https://doi.org/10.4049/jimmunol.1202688

Abstract

In vivo control of Mycobacterium tuberculosis reflects the balance between host immunity and bacterial evasion strategies. Effector Th1 cells that mediate protective immunity by depriving the bacterium of its intracellular niche are regulated to prevent overexuberant inflammation. One key immunoregulatory molecule is Tim3. Although Tim3 is generally recognized to downregulate Th1 responses, we recently described that its interaction with Galectin-9 expressed by M. tuberculosis–infected macrophages stimulates IL-1β secretion, which is essential for survival in the mouse model. Why IL-1β is required for host resistance to M. tuberculosis infection is unknown. In this article, we show that IL-1β directly kills M. tuberculosis in murine and human macrophages and does so through the recruitment of other antimicrobial effector molecules. IL-1β directly augments TNF signaling in macrophages through the upregulation of TNF secretion and TNFR1 cell surface expression, and results in activation of caspase-3. Thus, IL-1β and downstream TNF production lead to caspase-dependent restriction of intracellular M. tuberculosis growth.

Introduction

Host resistance to Mycobacterium tuberculosis relies on the cooperation between innate and adaptive immunity. The factors that drive this cooperation involve cytokines secreted by Th1 cells through cell contact–dependent signals and myeloid cells that are activated by Th1 cells to produce antimicrobial effector molecules. Of particular note, IFN-γ and TNF are produced by M. tuberculosis–specific Th1 cells and activate infected macrophages (Mφ) to induce intracellular mediators such as NO and promote changes in intracellular physiology, including phagolysosomal fusion (1, 2). Both IFN-γ−/− (−/−, knockout) and nitric oxide synthase-2 (NOS2−/−) mice are extremely susceptible to M. tuberculosis, which indicates the crucial role of IFN-γ and NO in immunity against tuberculosis (3–5). TNF plays a key role in granuloma formation, thereby molding the extracellular milieu in which M. tuberculosis–infected Mφ interact with M. tuberculosis–specific T cells. TNF blockade in M. tuberculosis–infected wild-type (WT) mice or latently infected humans exacerbates disease (6, 7). Together, IFN-γ and TNF play an important part in shaping the unique microenvironment in lung granulomas and differentially modulate effector T cell immune reactivity.

Following resolution and clearance of infection, effector T cells are deleted, which prevents excessive tissue inflammation and development of immunopathology. The expression of cell surface inhibitory receptors, such as T cell–Ig and mucin-domain–containing molecule-3 (Tim3), negatively regulates effector Th1 cells (8). In addition to its role in T cell exhaustion, we previously described that Tim3 expressed by T cells interacts with Gal9 expressed by infected Mφ to promote a program of cellular activation indicated by cytokine secretion and increased antimycobacterial activity (9). Cytokine secretion induced by Tim3–Gal9 interaction relied on caspase-1–dependent IL-1β secretion, suggesting that autocrine feedback by IL-1β further promotes Mφ activation and antimicrobial activity (9). Of interest, both IFN-γ and IL-1β induce Galectin-9 (Gal9), the ligand for Tim3 that upon binding to Tim3 transduces a signal to the T cells that triggers apoptosis, resulting in clonal contraction and/or deletion of effector Th1 cells (10–13). Thus, Tim3 and Gal9 define a bidirectional regulatory pathway that results in two distinct cellular outcomes—activation of Mφ and deactivation of T cells. Although such a mechanism may be appropriate for acute infection, it appears to be detrimental in the case of persistent pathogens such as HIV, hepatitis C virus, and M. tuberculosis.

As the antibacterial activity induced by Tim3 is mediated by IL-1β, we became interested in how IL-1β promotes intracellular control of M. tuberculosis replication. IL-1αβ−/− and IL-1R−/− mice are extremely susceptible to low-dose aerosol M. tuberculosis infection and die nearly as rapidly as IFN-γ−/−, IFN-γR−/−, and TNF−/− mice, despite elevated levels of IFN-γ and TNF in their lungs (14–18). These compelling data highlight the important contribution of IL-1β in defense against tuberculosis. The biological activity of IL-1β is tightly regulated (19). Regulation occurs at the level of 1) gene expression, 2) posttranscriptional activation of an inactive proform by proteolytic cleavage, and 3) competition with decoy receptors and soluble (s) IL-1R antagonists (19). Although production of IL-1β by Mφ in vitro generally requires both TLR signaling and inflammasome/caspase-1, IL-1β production during the early host response to M. tuberculosis infection appears to be independent of these two factors (16, 19, 20).

Despite the abundance of data on the importance of IL-1β in defense against tuberculosis, the molecular mechanism by which IL-1β enhances host resistance is unknown. In our low multiplicity of infection (MOI) model, <10% of Mφ are infected and IL-1β secretion is not detected (9). We evaluated how IL-1β restricts M. tuberculosis replication under conditions that induce IL-1β (e.g., Tim3) or by directly treating infected Mφ with recombinant IL-1β. We report that IL-1β activates M. tuberculosis–infected Mφ to restrict intracellular bacterial replication. The molecular basis for IL-1β–mediated M. tuberculosis control requires recruitment of TNF, upregulation of cell surface TNFR expression, and caspase-3 activation. These data support a model by which IL-1β promotes Mφ apoptosis, which inhibits the growth of intracellular M. tuberculosis.

Materials and Methods

Ethics statement

All animal work has been conducted according to relevant U.S. guidelines. The animals were housed in Association for Assessment and Accreditation of Laboratory Animal Care–approved animal facilities and meets National Institutes of Health standards as set forth in the Guide for the Care and Use of Laboratory Animals (Revised, 2010). The Institutional Animal Care and Use Committee of the Dana-Farber Cancer Research Institute (Animal Welfare Assurance A3431-01) approved all experimental procedures. The institution also accepts as mandatory the Public Health Service Policy on Humane Care and Use of Laboratory Animals by Awardee Institutions and National Institutes of Health Principles for the Utilization and Care of Vertebrate Animals Testing, Research, and Training. Human blood collected from healthy donors was purchased from Research Blood Components (Boston, MA), and its use was approved by the Institutional Review Board of Brigham and Women’s Hospital (Human Subjects Assurance FWA00000484). Written informed consent was obtained from healthy volunteers recruited at the National Institute of Respiratory Diseases, Mexico City, Mexico.

Materials

Reagents were as follows: anti-mouse CD120a (TNFR1/p55; clone 55R-286; BioLegend), anti-mouse CD120b (TNFR2/p75; TR75-89; BioLegend), anti-mouse CD11b (M1/70; BioLegend), anti-mouse F4/80 (BM8; BioLegend), rat anti-mouse CD16/CD32 (Fc-Block; BioLegend), CD11b microbeads (Miltenyi Biotec), human IgG1κ (I5154; Sigma-Aldrich), human serum (Gemini Bioproducts), Caspase-3 Inhibitor II (Z-DEVD-Fmk; 264155; Calbiochem), Caspase-8 Inhibitor II (Z-IETD-Fmk; 218759; Calbiochem), Caspase-9 Inhibitor I (Z-LEHD-Fmk; 218761; Calbiochem), Caspase inhibitor negative control (Z-FA-Fmk; 342000; Calbiochem), Green FLICA Caspases 3, 7, 8, and 9 Assay Kits (Immunochemistry Technologies), recombinant mouse IL-1α (rIL-1α; R&D Systems), recombinant mouse IL-1β (rIL-1β; R&D Systems), recombinant mouse TNF-α (rTNF-α; R&D Systems), recombinant human IL-1β (rIL-1β; R&D Systems), recombinant human TNF-α (rTNF; R&D Systems), anti-mouse IL-1β Ab (B122; eBioscience), anti-human IL-1β Ab (H1b-27; BioLegend), anti-mouse TNF-α Ab (MP6-XT22; BioLegend), anti-human TNF-α Ab (MAb1; BioLegend), Armenian hamster IgG (eBio299Arm; eBioscience), Rat IgG1 (RTK2071, BioLegend), Cytotoxicity Detection Kit PLUS (LDH; 04744926001, Roche), full-length (FL) and soluble forms of Tim3–Ig fusion protein (Tim3–Ig; V. Kuchroo), anti-Tim4 Ab (anti-phosphatidyl serine receptor; clone 4G3; V. Kuchroo), mouse TNF-α ELISA MAX Standard (430901; BioLegend), mouse IL-1β ELISA Max Standard (432602; BioLegend), mouse IL-1α ELISA Max Standard (433402; BioLegend), mouse IL-6 ELISA Max Standard (431302; BioLegend), and Ficoll-Paque PLUS (GE Healthcare Life Sciences).

Mice

C57BL/6J, IL-1R−/−, TNF−/−, TNFR1−/−, and TNFR2−/− mice 6–10 wk old were obtained from The Jackson Laboratory; MyD88−/− mice were obtained from K. Kobayashi, Harvard Medical School (Boston, MA) (21).

Human peripheral blood

Human blood collected from healthy donors was purchased from Research Blood Components, Boston, MA. Research Blood Components tests each donor's blood for standard blood-borne pathogens prior to the initial donation, and at the time of each subsequent donation. Only blood samples negative for HIV, human T cell leukemia virus, hepatitis C virus, and hepatitis B virus were used. Healthy donors were also recruited at the National Institute of Respiratory Diseases, Mexico City, Mexico. All healthy donors from Mexico City had received bacillus Calmette-Guérin vaccination at birth. Written informed consent was obtained from all participants.

Bacteria, cells, and culture

Virulent M. tuberculosis (H37Rv) was grown to midlog phase in Middlebrook 7H9 broth (BD Biosciences) with BBL Middlebrook OADC Enrichment (Becton Dickinson) and 0.05% Tween 80 (Difco). Aggregation was prevented by sonication for 10 s. CD11b+ peritoneal Mφ were harvested after being elicited with 3% thioglycolate, followed by CD11b+ selection using MACS columns. Purified cells were >95% F4/80+ CD11b+ as determined by flow cytometry. Mφ (1 × 105 cells per well) were seeded in a 96-well culture plate in complete RPMI 1640 medium (Invitrogen Life Technologies) supplemented with 10% FBS (Hyclone), 1 mM pyruvate, 1% nonessential amino acids, 1% minimal essential amino acids, 2 mM l-glutamine, 7 mM NaOH, and 20 mM HEPES (all from Life Technologies). Cells were allowed to adhere for 2–24 h prior to in vitro infection with M. tuberculosis.

In vitro infections and cocultures

Peritoneal Mφ were infected with H37Rv at MOI = 10, as described previously (22). In brief, M. tuberculosis were opsonized for 5 min using RPMI 1640 medium supplemented with 2% human serum/10% FBS/0.05% Tween 80, washed twice with complete medium without antibiotics. Bacteria were passed through a 5-μm syringe filter (Millipore), counted in a Petroff–Hausser chamber, and added to Mφ at the MOI indicated. The length of infection was 2 h for all experiments (unless otherwise indicated). Infected Mφ were cultured overnight before the addition of cytokines, Tim3–Ig, and other conditions. At days 1 and 5 post infection, cells were lysed with 1% Triton X-100 for 5 min, and mycobacteria were enumerated by plating serial dilutions of cell lysates on Middlebrook 7H10 agar plates and culture at 37°C. Colonies were counted after 21 d. In all the culture conditions mentioned below, CFU were enumerated at day 1 (unless otherwise indicated) in untreated infected Mφ to determine initial inoculum and at day 5 post infection to determine growth of intracellular M. tuberculosis in untreated infected Mφ and relative suppression or killing mediated by various treatments. The “experimental” MOI is calculated by dividing the CFU recovered on day 1 by the number of Mφ per well. For example, recovery of 20,000 CFU in a well containing 100,000 Mφ indicates an experimental MOI of 0.2 (Fig. 3D). The % growth inhibition of M. tuberculosis was calculated as follows: 100 × (CFUd5; conditions – CFUd1; media)/(CFUd5; media – CFUd1; media).

Human monocyte-derived Mφ isolation and infection

Human peripheral monocyte-derived Mφ (MDMs) were from heparinized blood from healthy donors by centrifugation using Ficoll-Paque PLUS gradient. Monocytes from PBMCs were enriched through CD14+ selection using MACS columns. Purified cells were >90% CD14+, as determined by flow cytometry. CD14+ monocytes were plated at 1 × 105 cells per well in 96-well plates supplemented with RPMI 1640 medium (Invitrogen Life Technologies) supplemented with 2 mM l-glutamine, streptomycin, penicillin (all from Life Technologies), and 10% human serum. After 7-d incubation, nonadherent cells were washed off with sterile PBS, and the medium was replenished; viable cells were considered monocyte-derived Mφ (MDMs) based on their adherence to tissue culture plates and expression profile of CD14 and CD68. Human monocyte-derived Mφ (HMDMs) were infected with H37Rv according to the methods mentioned above for peritoneal Mφ.

Tim3–Ig, IL-1β, TNF treatment and TNF, IL-1β, and caspase inhibition studies

Tim3–Ig, constructed as human IgG1 Fc tail fusion protein, is available as FLTim3–Ig or sTim3–Ig based on the domains included in the fusion protein construct (23, 24). Fusion proteins contained <0.1 EU/μg LPS (Chimerigen Laboratories, Allston, MA). In all in vitro infections, both FLTim3–Ig and sTim3–Ig were used, and data obtained were identical. However, for simplicity, data from either fusion protein are included in the figures. FLTim3–Ig, sTim3–Ig, and human IgG1 [human IgG (Hu IgG); control] were added to M. tuberculosis–infected Mφ at concentrations indicated in the figure legends. Following 20-min incubation, goat F(ab′)2 anti-human Ig at a final concentration of 2.5 μg/ml was added to cross-link the fusion proteins or Hu IgG. To test control of M. tuberculosis by IL-1α, IL-1β in murine peritoneal Mφ, cytokines at final concentrations of 10, 1, and 0.1 ng/ml were added directly to media containing infected Mφ. In HMDMs, IL-1β at final concentrations of 10, 5, 2.5, 1.25, and 0.625 ng/ml were added directly to media containing infected Mφ. For all cytokine blockade studies using cytokine-neutralizing Abs (anti–IL-1β and anti-TNF), 25 μg/ml neutralizing Ab was added either separately to infected Mφ or along with treatment (Tim3–Ig, TNF, or IL-1β), as indicated in the figure legend. For example, to determine whether TNF was required for Tim3–Ig–mediated M. tuberculosis killing, anti-TNF (25 μg/ml; final concentration) was added separately to infected Mφ along with 10 μg/ml Tim3–Ig. To account for any cells that died following Tim3–Ig treatment and and released mycobacteria into cell culture supernatant, leading to underestimation of total M. tuberculosis in infected Mφ, we measured CFU following 1) removal of supernatant, lysis of Mφ in 1% Triton X-100, and plating the supernatant and the Mφ lysate; or 2) lysing Mφ without the removal of cell culture supernatant by adding 10% Triton X-100 at 1/10 the cell culture volume. Under the standard in vitro infection conditions mentioned above, we detect < 10% of the total CFU in the cell culture supernatant. The CFU present in the supernatant was not statistically significant between Mφ treated with media, Tim3–Ig, Hu IgG, IL-1β, or TNF. To determine whether caspase-3, -8, and -9 were involved in Tim3–Ig– and IL-1β–mediated control, caspase-3 (Z-DEVD-Fmk), caspase-8 (Z-IETD-Fmk), or caspase-9 (Z-LEHD-Fmk) peptide inhibitors at 10 μM final concentration were added to the culture media 30 min prior to Tim3–Ig or Il-1β treatment. Negative control peptide (Z-FA-Fmk) was added at 10 μM final concentration. As a control for cytotoxicity, we treated Mφ with caspase-3, -8, or -9 inhibitors and the negative control peptide in the absence of Tim3–Ig or IL-1β. Our ability to recover similar levels of intracellular mycobacteria in caspase-3, -8, -9, or negative peptide inhibitor–treated Mφ and untreated Mφ indicated that minimal cytotoxicity was associated with these inhibitors at the concentrations used.

Cytokine detection

Culture supernatants from uninfected and infected Mφ following IL-1β treatment were filtered through a 0.2-μM filter to remove any bacteria. Supernatants were assayed for IL-1α, IL-1β, IL-6, and TNF by sandwich ELISA performed in accordance with the manufacturer’s instructions (BioLegend), and absorbance was recorded at 405 nm on SoftMax Pro ELISA analysis software (Molecular Devices).

Flow cytometry

Uninfected and M. tuberculosis–infected Mφ following 24-h Tim3–Ig, IL-1β, or TNF treatment were stained for 30 min at 4°C with 25 μg/ml anti-CD120b-PE, anti-CD120a-APC, anti-CD11b-APC-Cy7, and anti-F4/80-Pacific Blue Abs. To inhibit nonspecific staining, murine Fc receptors were blocked with 25 μg/ml Fc-Block for 20 min at 4°C prior to staining with fluorochrome-conjugated Abs and appropriate isotype controls. To determine apoptosis following 24-h Tim3–Ig or IL-1β treatment, uninfected and M. tuberculosis–infected Mφ were stained for activated caspase-3, using 1) FLICA reagent FAM-DEVD-FMK, 1 h at 37°C (Immunochemistry Technologies; 1:10; optimal excitation range = 490–495 nm, and emission range = 515–525 nm); and 2) Alexa Fluor 647 rabbit anti-active caspase-3, 30 min at 4°C (BD560626; 1:25). Activated caspase-8 and -9 were also detected using FLICA reagents FAM-LETD-FMK and FAM-LEHD-FMK, respectively. FLICA reagent FAM-DEVD-FMK, FAM-LETD-FMK, and FAM-LEHD-FMK are a cell-permeable nontoxic fluorescently conjugated caspase-3, -8, and -9 peptide that irreversibly binds to activated caspases with a preference for its target peptide sequence (FAM-DEVD-Fmk, FAM-LETD-FMK, and FAM-LEHD-FMK highly specific for caspase-3/7, -8, and -9 respectively). Because the FLICA reagent becomes covalently coupled to the active caspase enzymes, it is retained within the cell during wash steps, whereas any unbound FLICA reagent diffuses out of the cell and is washed away. The remaining green fluorescent signal is a direct measure of the amount of caspase-3, -8, or -9 activity present in the cell at the time the reagent was added. Alexa Fluor 647 rabbit anti-active caspase-3 preferentially binds activated caspase-3 and was added to Mφ following fixation and permeabilization performed according to the manufacturer’s instructions (BD Biosciences). Both staining strategies (caspase-3 FLICA for 1 h and Alexa Fluor 647 rabbit anti-active caspase-3 for 30 min) yielded similar results. The cells were costained with LIVE/DEAD Viability dye (Life Technologies). Data were collected using a FACSCanto II (BD Biosciences) and analyzed with FlowJo (Tree Star).

In vitro assays of necrosis and apoptosis

Necrosis/pyroptosis in Mφ was evaluated through the release of the intracellular enzyme LDH in cell culture supernatants. The in vitro infections performed were done at a low (actual) MOI, resulting in infection of 1–2% of cells. Under these standard in vitro infection conditions, we did not detect LDH release. To improve the sensitivity of the LDH assay, we sought to increase the frequency of infected cells by infecting the Mφ overnight as opposed to 2 h. At the times indicated post infection, the LDH activity of the culture supernatants of infected cells was measured using a cytotoxicity detection kit according to the manufacturer’s protocol. The percentage of LDH release was calculated according to the following formula: (LDH activitytest sample, SN – LDH activityuntreated uninfected Mφ, SN)/(Maximal releasable LDHSN+lys − LDH activityuntreated uninfected Mφ, SN), where SN is the amount in the supernatant and Lys is the amount in the lysates. Apoptosis and necrosis were measured by cell ELISA (Cell Death Detection ELISA PLUS; 11 920685 001; Roche Applied Science) for quantification of cytoplasmic (apoptosis) and extracellular (necrosis) histone-associated DNA fragments according to the specifications of the manufacturer. The relative amount of necrosis or apoptosis was calculated as a ratio of the absorbance of infected Mφ to that of uninfected control Mφ.

Statistical analysis

One-way ANOVA was used to analyze the in vitro Mφ infections, and Dunnett’s posttest was used to compare appropriate control conditions, as indicated in the figure legends. Analysis was performed using Prism 5.0 software (GraphPad Software, San Diego, CA).

Results

IL-1 restricts intracellular bacterial replication in M. tuberculosis–infected murine Mφ

We previously showed that Tim3 binding to Galectin-9 expressed by M. tuberculosis–infected Mφ led to Mφ activation and IL-1β–dependent M. tuberculosis control in both murine peritoneal and alveolar Mφ (9). To determine the full potential of IL-1β to mediate antimicrobial activity in murine Mφ, we treated M. tuberculosis–infected Mφ with recombinant IL-1β. IL-1β inhibited M. tuberculosis replication in thioglycolate-elicited peritoneal Mφ in a dose-dependent manner (Fig. 1A). At a dose of 10 ng/ml, bacterial replication was inhibited by 80–90% (p < 0.05, one-way ANOVA). In addition to IL-1β, IL-1α also binds specifically to the IL-1R. However, IL-1α suppressed bacterial growth less potently (Fig. 1A). The greater activity of IL-1β is consistent with its greater affinity for the IL-1R. Suppression of M. tuberculosis replication was a specific action of IL-1β, because neither Tim3–Ig nor IL-1β was able to inhibit bacterial growth in infected Mφ lacking IL-1R (Fig. 1B). Finally, the antimicrobial function of both Tim3–Ig and IL-1β was dependent on MyD88, as neither molecule was able to control M. tuberculosis growth in MyD88−/− Mφ (Fig. 1B). These data show that the antimicrobial action of IL-1β requires signaling via the IL-1R and the adaptor molecule MyD88.

FIGURE 1.
FIGURE 1.

IL-1 restricts intracellular bacterial replication in M. tuberculosis–infected murine Mφ. (A) M. tuberculosis–infected thioglycolate-elicited peritoneal Mφ were cultured alone or in the presence of increasing amounts of IL-1α or IL-1β. Bacterial growth was determined on day 4. Percent M. tuberculosis inhibition was calculated as described in Materials and Methods. (B) Percent inhibition of M. tuberculosis growth in WT, IL-1R−/−, and MyD88−/− Mφ cultured in the absence or presence of Tim3–Ig or Hu IgG (10 μg/ml), or 10 ng/ml of IL-1β. Data are representative of four independent experiments in (A), or pooled data from three independent experiments are shown in (B). Data points in each graph represent mean ± SEM. *p < 0.05, ***p < 0.001, one-way ANOVA compared with control conditions: (A) untreated Mφ alone (0 ng/ml); (B) WT Mφ.

IL-1β suppresses intracellular mycobacterial growth in human Mφ

An essential question is whether IL-1β induced similar antimycobacterial activity in human Mφ. Peripheral monocytes were isolated from healthy donors and allowed to differentiate into Mφ (MDMs). Treatment with recombinant human IL-1β led to a significant reduction in intracellular mycobacterial growth in HMDMs (Fig. 2A). MDMs from six anonymous donors from the United States and Mexico were infected in vitro with M. tuberculosis and treated with human IL-1β. IL-1β inhibited intracellular growth of M. tuberculosis in all donors. A significant CFU reduction was seen at a dose of 1.25 ng/ml human IL-1β, and 10 ng/ml completely suppressed M. tuberculosis replication (Fig. 2B). These results show that IL-1β signaling activates human Mφ to suppress intracellular growth of mycobacteria.

FIGURE 2.
FIGURE 2.

IL-1β restricts intracellular bacterial replication in M. tuberculosis–infected human Mφ. (A) Peripheral blood HMDMs from a single donor were infected in vitro with M. tuberculosis and were cultured alone or in the presence of 10 ng/ml IL-1β. Six replicate cultures per condition were performed. (B) M. tuberculosis–infected MDMs from six healthy donors were cultured alone or in the presence of increasing concentrations of IL-1β. The effect of IL-1β on bacterial growth was calculated as percent inhibition, as described in Materials and Methods. *p < 0.05, **p < 0.01, ***p < 0.001, one-way ANOVA compared with control conditions: (A) day 6 media; (B) untreated Mφ alone (0 ng/ml).

Tim3/Gal9- and IL-1β–induced M. tuberculosis control in Mφ requires TNF

We previously observed that Tim3–Ig treatment of M. tuberculosis–infected Mφ not only induced IL-1α and IL-1β but also stimulated significant production of IL-6, TNF, MIP-1α, MIP-1β, and G-CSF, compared to untreated infected or Tim3–Ig-treated uninfected Mφ (9). Our previous observation that induction of these cytokines by Tim3–Ig was dependent on caspase-1 suggested that Tim3-induced IL-1β acts in an autocrine manner to stimulate the production of additional cytokines. Given the importance of TNF in antimicrobial immunity, we hypothesized that IL-1β induction of TNF is important in restricting bacterial growth. Indeed, neutralizing Abs to TNF inhibited Tim3–Ig–mediated suppression of M. tuberculosis growth (Fig. 3A), and Tim3–Ig could not induce antimicrobial activity in TNF-deficient Mφ (Fig. 3B). Addition of TNF to infected Mφ also suppressed M. tuberculosis growth (Fig. 3A).

FIGURE 3.
FIGURE 3.

Tim3- and IL-1β–mediated control of M. tuberculosis replication requires TNF. (A) M. tuberculosis–infected WT Mφ were cultured either alone or in the presence of 10 μg/ml Tim3–Ig with and without 25 μg/ml anti-TNF neutralizing Ab (αTNF). (B) WT and TNF−/− Mφ were infected with M. tuberculosis in parallel. On day 1, 10 μg/ml of Tim3–Ig or Hu IgG (control) was added to the Mφ. (C) M. tuberculosis–infected WT Mφ were cultured alone or with 10 ng/ml of IL-1β. At 24 h, culture supernatants from triplicate wells were assayed for TNF. Open bars indicate uninfected Mφ, and closed bars indicate M. tuberculosis–infected Mφ. (D) M. tuberculosis (Mtb)–infected murine peritoneal Mφ were cultured either alone or in the presence of 10 ng/ml IL-1β or 10 ng/ml TNF with and without 25 μg/ml anti-murine TNF (αTNF) or anti-murine IL-1β (αIL-1β) neutralizing Ab. (E) Percent inhibition in CFU in WT, IL-1R−/−, and MyD88−/− Mφ treated with and without 10 ng/ml of IL-1β or TNF. (F) M. tuberculosis–infected HMDMs were cultured either alone or in the presence of 10 ng/ml IL-1β with and without 25 μg/ml anti-murine TNF neutralizing Ab (αTNF). CFUs in (A), (B), (D), and (F) (left panel) were determined on day 1 and day 4 or day 5 post infection. Percent inhibition in (E) and (F) (right panel) is CFU growth in test conditions normalized to untreated Mφ alone. Data are representative of two (A, B), three (C, F, left panel), or four (D) independent experiments. Pooled data from three independent experiments are shown in (E) and (F), right panel. Bars represent mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001, one-way ANOVA compared with control conditions: (A) isotype control; (B) Hu IgG; (D) isotype control; (E) WT Mφ; (F) isotype control. IC, Isotype control.

Because both IL-1β and TNF were essential for Tim3–Ig–mediated control, we wished to determine whether IL-1β and TNF act in parallel or in series, and possibly regulate each other. IL-1β treatment of uninfected and M. tuberculosis–infected Mφ increased TNF transcription (data not shown) and TNF secretion (Fig. 3C). In contrast, TNF did not induce IL-1β transcription or secretion (Supplemental Fig. 1). Furthermore, TNF did not induce the secretion of other NF-κB–dependent cytokines, such as IL-1α or IL-6 (Supplemental Fig. 1). These data indicate that IL-1β induces TNF production by M. tuberculosis–infected Mφ. To determine whether TNF contributed to the antimicrobial activity of IL-1β, M. tuberculosis—infected Mφ were treated with IL-1β in the presence of neutralizing Ab to TNF. Treatment with anti-TNF, but not an isotype control, abrogated the antimicrobial effect of IL-1β (Fig. 3D). In contrast, the antimicrobial effect of TNF was not dependent on IL-1β, because anti–IL-1β neutralizing Ab had no intrinsic effect on TNF-mediated M. tuberculosis control (Fig. 3D). Neither anti-TNF nor anti–IL-1β mAb by itself has any effect on M. tuberculosis growth. As expected, the antimicrobial effect of TNF was independent of IL-1R and MyD88 (Fig. 3E).

To determine whether IL-1β–stimulated TNF production led to control of intracellular M. tuberculosis growth in human Mφ, infected MDMs were treated with IL-1β in the presence of TNF blocking Abs. IL-1β efficiently suppressed M. tuberculosis growth by ∼90%. As predicted on the basis of our murine data, addition of anti-human TNF mAbs abrogated the antimicrobial activity of IL-1β (Fig. 3F). Collectively, these data show that IL-1β signaling induces TNF, which in turn activates downstream antimicrobial activity in both murine and human Mφ.

IL-1β modulates TNFR1 expression to mediate M. tuberculosis control

Soluble and membrane-bound TNF signals via two distinct receptors, TNFR1 and TNFR2 (25). On average, 80–95% of thioglycolate-elicited peritoneal Mφ express cell surface TNFR2, but only a small fraction (4–5%) of these Mφ express cell surface TNFR1 (Fig. 4A). The expression of TNFR1 and TNFR2 was not significantly altered following M. tuberculosis infection. However, IL-1β treatment of M. tuberculosis–infected Mφ, but not uninfected Mφ, upregulated cell surface levels of TNFR1 but had little effect on TNFR2. A similar effect was observed when the Mφ were activated by TNF. In the absence of IL-1R, IL-1β had no effect on TNFR1 expression in M. tuberculosis–infected Mφ (Fig. 4B). To determine which TNFR was required for Tim-3– and IL-1β–mediated M. tuberculosis control, we treated TNFR1−/− and TNFR2−/− Mφ with Tim3–Ig, IL-1β, and TNF. TNFR1 was essential, whereas TNFR2 was dispensable for sTNF-mediated M. tuberculosis control, confirming previously established TNF signaling pathways for TNF receptors. Tim3–Ig– and IL-1β–mediated M. tuberculosis control was dependent on TNFR1 and largely independent of TNFR2 signaling. (Fig. 4C). These data indicate that TNFR1 is required for antimicrobial activities of Tim3–Ig and IL-1β.

FIGURE 4.
FIGURE 4.

Regulation of TNF receptor expression and its requirement for IL-1β–mediated M. tuberculosis control. (A) Uninfected and M. tuberculosis–infected (MOI 10) peritoneal Mφ were cultured alone or in the presence of IL-1β or TNF (10 ng/ml). Cell surface expression of TNFR1 and TNFR2 of CD11b+ F4/80+ Mφ was assessed by flow cytometry after 24 h. (B) Cell surface expression of TNFR1 is shown for uninfected and M. tuberculosis–infected (MOI 10) WT and IL-1R−/− Mφ. (C) M. tuberculosis–infected WT, TNFR1−/−, and TNFR2−/− Mφ were treated with 10 μg/ml of Tim3–Ig or Hu IgG (control), or 10 ng/ml IL-1β or TNF. Data are representative of two (A, B) independent experiments. Pooled data from two independent experiments are shown in (C). **p < 0.01, ***p < 0.001, one-way ANOVA compared with control conditions: (C) WT Mφ.

Tim3/Gal9 binding and IL-1β both induce caspase-3 activation in M. tuberculosis–infected Mφ

As Tim3–Ig and IL-1β treatment promotes control of intracellular bacterial replication by infected Mφ in a TNF-dependent manner, we hypothesized that IL-1β was modulating the death modality of infected Mφ via the extrinsic death pathway. We measured the release of the intracellular enzyme LDH into cell culture supernatant, as a measure of plasma cell membrane disruption. According to our standard infection protocol, no necrosis was observed. Because the actual multiplicity of infection (MOI) is very low, with <10% of Mφ being infected, we increased the length of infection from 2 h to overnight and varied the MOI to increase the percentage of Mφ that were infected. Under these conditions, necrosis was detected 3 d post infection and correlated with increasing MOI; however, neither IL-1β nor Tim3–Ig caused an increase in LDH release (data not shown). These data are consistent with our prior data that Tim3–Ig does not induce necrosis (9).

We next asked whether Tim3–Ig or IL-1β induced apoptosis of M. tuberculosis–infected Mφ. Using an ELISA that detects intracellular and extracellular DNA–histone complexes as a measure of apoptosis and necrosis, respectively, we previously did not find any evidence that Tim-Ig induces apoptosis. However, the ELISA is of unknown sensitivity, and because only 1–10% of the Mφ are infected, we sought to use an assay that can detect apoptosis at the single-cell level. Mφ infected with MCherry–M. tuberculosis were treated with Tim3–Ig or IL-1β, and after fixation, a TUNEL assay was performed. Despite trying multiple MOIs and time points, a consistent increase in TUNEL+ cells was not detected after Tim3–Ig or IL-1β treatment (data not shown). In parallel, to determine whether Tim3–Ig or IL-1β leads to the activation of caspase-3, the executioner caspase that has a pivotal role in apoptosis, we measured caspase-3 activation in M. tuberculosis–infected Mφ, using the fluorescently conjugated caspase-3 substrate Z-DEVD-Fmk. Caspase-3 is activated by both extrinsic (death ligand) and intrinsic (mitochondrial) pathways via caspase-8 and capase-9 initiator caspases. On average, 5–8% of uninfected thioglycolate-elicited peritoneal Mφ express active caspase-3. The number of Mφ expressing active caspase-3 increased 2-fold following M. tuberculosis infection (Fig. 5A). Tim3–Ig–treated M. tuberculosis–infected Mφ, but not uninfected Mφ or control treated M. tuberculosis–infected Mφ, had increased caspase-3 activity (Fig. 5A, 5B). A similar effect was observed when murine Mφ were treated with IL-1β (Fig. 5C). The action of IL-1β was specific because IL-1β treatment of M. tuberculosis–infected IL-1R−/− Mφ did not increase active caspase-3 (Fig. 5D). As predicted on the basis of our murine data, IL-1β also induced caspase-3 activation in M. tuberculosis–infected human Mφ (Fig. 5E). To further confirm TNF was responsible for this caspase-3 activation, infected MDMs were treated with IL-1β in the presence of TNF blocking Abs. The addition of anti-human TNF mAb diminished the increased frequency of active caspase-3 in HMDMs to baseline levels in untreated media–alone conditions (Fig. 5E, 5F). Similar results were obtained when intracellular staining with an mAb specific for active caspase 3 was used (data not shown). In addition, IL-1β treatment of M. tuberculosis–infected HMDMs led to TNF-dependent activation of caspase-8 and -9 (see Supplemental Fig. 2). We conclude that in M. tuberculosis–infected Mφ, stimulation of IL-1β by Tim3/Gal9 binding, or addition of exogenous IL-1β, leads to activation of caspase-3, the key executioner caspase that mediates apoptosis.

FIGURE 5.
FIGURE 5.

Tim3 and IL-1β induce caspase-3 activation. (A) M. tuberculosis–infected and uninfected thioglycolate-elicited Mφ were cultured either alone or in the presence of 10 μg/ml Tim3–Ig or Hu IgG (control). At 24 h, intracellular expression of active caspase-3 (aCaspase-3) in CD11b+ F4/80+ Mφ was assessed by flow cytometry. (B) Fold induction of active caspase-3 compared with untreated uninfected Mφ is graphically plotted. Stauro, staurosporine [1 μM], positive control. (C) M. tuberculosis–infected and uninfected thioglycolate-elicited Mφ were cultured either alone or in the presence of 10 ng/ml IL-1β. At 24 h, intracellular expression of active caspase-3 in CD11b+ F4/80+ Mφ was assessed by flow cytometry. (D) Fold induction of active caspase-3 over untreated uninfected Mφ is graphically plotted for uninfected and M. tuberculosis–infected WT and IL-1R−/− Mφ. (E) M. tuberculosis–infected and uninfected HMDMs were cultured either alone or in the presence of 10 ng/ml IL-1β with and without 25 μg/ml anti-TNF neutralizing Ab (αTNF). At 24 h, intracellular expression of active caspase-3 in CD14+ Mφ was assessed by flow cytometry. (F) Fold induction of active caspase-3 compared with untreated uninfected Mφ is graphically plotted. Data are representative of two (A, B, E, F) and three (C) independent experiments. Pooled data from three independent experiments are shown in (D). Bars indicate mean ± SEM from three replicate cultures. *p < 0.05, **p < 0.01, ***p < 0.001, one-way ANOVA compared with conditions: (B) Tim3–Ig and staurosporine; (D) IL-1β–treated WT Mφ; or (F) IL-1β–treated HMDMs.

The antimicrobial effect of IL-1β requires efferocytosis

We next evaluated whether active caspase-3, -8, and -9 were required for Tim3–Ig– and IL-1β–mediated M. tuberculosis control in murine peritoneal Mφ and HMDMs. M. tuberculosis–infected Mφ were treated with Tim3–Ig or IL-1β in the presence of Z-DEVD-Fmk, Z-IETD-Fmk, and Z-LEHD-Fmk, peptide inhibitors of caspase-3, -8 and -9 activity, respectively. Z-DEVD-Fmk, Z-IETD-Fmk, and Z-LEHD-Fmk abrogated Tim3–Ig–and IL-1β–mediated control of M. tuberculosis infection (Fig. 6A, 6B), showing that caspase-3, -8, and -9 are required for Tim3–Ig– and IL-1β–mediated M. tuberculosis control in murine and human Mφ. In the absence of Tim3–Ig or IL-1β, Z-DEVD-Fmk, Z-IETD-Fmk, and Z-LEHD-Fmk had no effect on bacterial replication (data not shown).To rule out any possible toxic effects of caspase inhibitors, we monitored the secretion of TNF, IL-1α, and IL-6 in the presence of Tim3–Ig and IL-1β treatment (Supplemental Fig. 3). Although we saw reduction in the levels of IL-6 following addition of peptide inhibitors, we did not observe any reduction in the levels of sTNF and IL-1α following addition of caspase-3 peptide inhibitor, suggesting that loss of CFU control in IL-1β treatment conditions was not due to caspase-3–mediated inhibition of TNF secretion.

FIGURE 6.
FIGURE 6.

Active caspase-3 and efferocytosis are required for M. tuberculosis control. (A) H37Rv-infected WT Mφ were cocultured with Tim3–Ig or IL-1β in the presence or absence of Z-DEVD-Fmk, Z-IETD-Fmk, and Z-LEHD-Fmk, peptide inhibitors of caspase-3, -8, and -9 activity, respectively. (B) H37Rv-infected HMDMs were cocultured with 10 ng/ml IL-1β in the presence or absence of Z-DEVD-Fmk, Z-IETD-Fmk, and Z-LEHD-Fmk, peptide inhibitors of caspase-3, -8, and -9 activity, respectively. Control peptide is Z-FA-Fmk. (C) H37Rv-infected WT Mφ were cultured either alone or in the presence of 10 ng/ml IL-1β, with and without 25 μg/ml blocking Ab to PS receptor (αPS-R). Data are representative of four (A, B) or two (C) independent experiments. Bars indicate mean ± SEM from three replicate cultures. *p < 0.05, **p < 0.01, ***p < 0.001, one-way ANOVA compared with conditions: (A, B) Tim3–Ig– or IL-1β–treated WT Mφ; (C) isotype control. IC, Isotype control.

We have recently shown that the viability of M. tuberculosis is reduced when M. tuberculosis sequestered within apoptotic vesicles is rapidly taken up by uninfected bystander Mφ by a constitutive process called efferocytosis (26). In contrast, necrosis of M. tuberculosis–infected Mφ leads to bacterial dispersal and ongoing growth. Uninfected bystander Mφ recognize dying apoptotic cells through “find me” and “eat me” signals such as phosphatidylserine (PS). To determine whether an IL-1β–mediated program of antimicrobial activity requires efferocytic uptake of apoptotic Mφ, M. tuberculosis–infected Mφ were treated with blocking Ab to the PS receptor (Tim-4) in the presence of IL-1β (27, 28). Blocking Ab to the PS receptor, but not the control Ab, completely abolished IL-1β–mediated M. tuberculosis control (Fig. 6C). We found that IL-1β did not enhance efferocytosis of apoptotic cells (data not shown). These data support the hypothesis that IL-1β induces apoptosis in M. tuberculosis–infected Mφ that then are engulfed by the process of efferocytosis, which is required to restrict M. tuberculosis replication.

Discussion

The inflammasome complex, IL-1β, and downstream innate adaptor MyD88 are all important in modulating host susceptibility to M. tuberculosis (16). Mice deficient in either IL-1R or IL-1β are acutely susceptible to M. tuberculosis (14, 16). Although it has been recently described that inflammatory monocytes and dendritic cells, distinguished by their cell surface expression of CD11c and Ly6C, are the primary sources of IL-1β production in the lungs of M. tuberculosis–infected mice, why IL-1β is essential for host resistance to M. tuberculosis is unknown (29). In this article, we provide the first evidence, to our knowledge, that IL-1β directly activates innate antimicrobial activity in murine Mφ and HMDMs leading to restriction of M. tuberculosis replication. Using in vitro models of M. tuberculosis infection in Mφ, we were able to ascertain the antimicrobial molecules activated following IL-1R signaling that contribute to IL-1β–mediated M. tuberculosis control.

We previously reported that the caspase-1–processed cytokine IL-1β promotes innate antimicrobial immunity in M. tuberculosis–infected Mφ following interaction between Tim3 expressed by T cells and Gal9 expressed by Mφ (9). We found that in M. tuberculosis–infected Mφ, Tim3 activates a program of antimicrobial immunity that involves the initial secretion of IL-1β followed by IL-1β–mediated recruitment of other antimicrobial effectors, including IL-6, TNF, MIP-1α, MIP-1β, and G-CSF. Our previous finding that genetic ablation of caspase-1 in M. tuberculosis–infected Mφ suppressed TNF production strongly indicated that IL-1β was acting in an autocrine manner to promote production of additional antimicrobial effectors (9). In this article, we show that IL-1β directly increased TNF transcription and secretion, an activity that was augmented in M. tuberculosis–infected Mφ. We do not believe that autocrine action of IL-1β on Mφ is the sole source of TNF in vivo. Indeed, IL-1R−/− and IL-1β−/− exhibit higher levels of TNF in lungs of infected mice than in those of WT controls (16, 29). This finding indicates that mechanisms other than IL-1β activation of Mφ are important for TNF production during M. tuberculosis infection. Finally, we found that IL-1β potentially augments TNF signal transduction by upregulating TNFR1 cell surface expression on infected Mφ. The antimicrobial actions of both Tim3–Ig and IL-1β were largely dependent on TNFR1 and partially dependent on TNFR2. TNFR1 can be activated by both sTNF and membrane TNF, whereas TNFR2 has some exclusivity for membrane TNF. The nature of the TNF ligand produced following IL-1β signaling is unknown. However, one explanation for our finding is the “ligand-passing” mechanism by which TNFR2 holds ligand (TNF) and enhances the local TNF concentration in the vicinity of TNFR1, which in turn accepts TNF from TNFR2 leading to TNFR1 signaling (25). Whereas TNFR1 is indispensable for IL-1β–mediated M. tuberculosis control, TNFR2 appears to play a minor role. Thus, the ability of Tim3–Ig and IL-1β to stimulate TNF production by M. tuberculosis–infected Mφ is required for their antimicrobial activity.

The ability of the Tim3/Gal9 pathway to restrict intracellular M. tuberculosis replication is independent of IFN-γ and NO (9), two of the chief mediators that lead to bacterial control. Therefore, we considered whether Tim3/Gal9 signaling altered the death modality of infected Mφ. Induction of Mφ necrosis, which is the cellular outcome more frequently associated with infection by virulent M. tuberculosis, leads to enhanced bacterial growth in vitro. In contrast, apoptosis of M. tuberculosis–infected Mφ is associated with the control of intracellular bacterial growth (30). Apoptosis, a tightly regulated complex process, is initiated by extrinsic (caspase-8 dependent) or intrinsic (caspase-9 dependent) signals and culminates in the activation of executioner caspase-3 or caspase-7. TNF is a pleiotropic molecule that plays crucial roles in cellular stress, inflammation during infection, tissue damage, and tuberculosis granuloma formation in the lung (6, 7, 31–33). TNF signaling via its receptors TNFR1 and TNFR2 influences cell fate and can lead to three distinct outcomes: 1) necrosis, 2) apoptosis through the death domain TRADD, and 3) cellular survival via NF-κB activation (25). M. tuberculosis can evade TNF-mediated apoptosis by stimulating human Mφ to secrete sTNFR2, which serves as an antagonist of TNFR1 signaling (34). Virulent mycobacteria can also evade TNF-mediated apoptosis by inducing necrosis (30, 35, 36).

Tim3–Ig did not induce more necrosis or apoptosis of infected Mφ (9). However, the characterization of cell death during M. tuberculosis infection suffers from difficulties, including differing death modalities, lack of synchronization, secondary necrosis, varying kinetics, and poor performance of some experimental assays following fixation. In particular, we were concerned that the DNA–histone ELISA may lack the sensitivity to detect rare events, particularly in our low MOI culture system. We next turned to a microscopy-based assay using MCherry-expressing H37Rv and TUNEL. Although sensitive, this assay gave inconsistent results. However, other data suggested that Tim3–Ig was modulating cell death. After treatment with Tim3–Ig, more M. tuberculosis–infected Mφ expressed activated capsase-3. Although the frequency of caspase-3+ cells was higher than expected based on our TUNEL staining, these results were confirmed by showing that CFU control was dependent on caspase-3, suggesting that Tim3–Ig induced greater apoptosis. In parallel, CFU control was inhibited by caspase-8 and -9 inhibitors. Such codependence on both the intrinsic and the extrinsic pathways is observed when the initial precursor pool of caspase-8 is not sufficient to drive subsequent caspase-3 activation. In certain cell types (M2 Mφ) the initial precursor pool of active caspase-8 leads to truncation of Bid, which translocates to the mitochondria that serve as a platform for caspase-9 activation and subsequent caspase-3 activation and apoptosis (37). We cannot completely rule out that the effect of these peptide inhibitors is due to inhibition of serine proteases other than caspase-3, -8, and -9. However, the known enzymes that are cross inhibited are other caspases or cathepsins, which also have a role in cell death.

We have recently described that Mφ engulfment of apoptotic infected cells leads to destruction of intracellular M. tuberculosis (26). This process, commonly referred to as efferocytosis, occurs when Mφ recognize and engulf apoptotic cells. Apoptosis is not inherently bactericidal; rather, the efferocytic uptake of dying apoptotic Mφ by uninfected bystander Mφ restricts M. tuberculosis viability. We have shown that intracellular bacterial control occurs when M. tuberculosis sequestered within an apoptotic macrophage is further compartmentalized within the efferocytic phagosome, which is subsequently delivered along with the apoptotic cell debris to the lysosomal compartment for degradation. (26). The recognition of PS by a variety of PS receptors is critical for efferocytosis. Although several PS receptors have been identified, the dominant one used by peritoneal Mφ is Tim4. Our data that anti-Tim4 mAb blocks bacterial control induced by IL-1β and Tim3–Ig implicate efferocytosis of apoptotic cells as part of the mechanism.

Our finding that IL-1β requires active caspase-8, -9, and -3 for its antimicrobial activity suggests that IL-1R signaling leads to TNF/TNFR1-mediated activation of the extrinsic caspase-8 pathway that feeds into the intrinsic caspase-9 pathway, resulting in caspase-3–mediated apoptosis of M. tuberculosis–infected Mφ and control of intracellular bacterial replication. Phagocytic cells clear apoptotic cells by efferocytosis, a highly conserved process that is important in preventing autoimmunity and inappropriate inflammation. Defective efferocytosis often seen in atherosclerotic plaques is a failure to clear lipid-laden apoptotic Mφ (also called foamy Mφ) that leads to postapoptotic necrosis (secondary necrosis) (38). Foamy Mφ are also characteristic of caseating necrotic lesions in tuberculous granuloma (39). Could the presence of foamy Mφ in TB granulomas therefore be indicative of defective efferocytosis whereby lipid-laden Mφ are not efficiently cleared by bystander Mφ? We find that IL-1β 1) induces apoptosis in M. tuberculosis–infected Mφ, 2) requires efferocytosis for M. tuberculosis control, but 3) does not by itself regulate efferocytosis. These data suggest a model in which IL-1β production by infected Mφ or neighboring cells stimulates TNF production by M. tuberculosis–infected Mφ, leading to their apoptotic cell death. These dying cells are engulfed by recruited Mφ (efferocytosis), which we believe represents a common final pathway that restricts M. tuberculosis replication.

The extensive pulmonary necrosis seen in infected M. tuberculosis IL-1R−/− mice may reflect the role that IL-1β plays in modulating cell death and host resistance to M. tuberculosis (16). Given the high levels of IL-1R antagonist in lungs of infected WT mice that can block IL-1R signaling, we speculate that inhibited IL-1R signaling could impair control of bacterial growth and contribute to recrudescence and lung disease (caseating necrosis) in M. tuberculosis–infected WT mice (16). The functional overlap between IL-1R signaling and TNFR1/TNFR2 signaling has been previously illustrated in lung models of airway inflammation. IL-1β augments TNF-mediated secretion of chemokines such as MIP-2 and KC, key chemoattractants for neutrophils and Mφ, thereby altering the microenvironment of the lung (40–42). Although the outcome of enhanced Mφ and neutrophil recruitment on host resistance is unpredictable, it is likely that tight regulation of the IL-1β/TNF signaling axis prevents overexuberant inflammation and tissue damage, possibly at the cost of optimal bacterial control (43).

In summary, our work provides evidence that IL-1β directly activates antimicrobial effector pathways in M. tuberculosis–infected human and murine Mφ. We find that IL-1β acts in an autocrine fashion to modulate TNFR signaling through increased TNF and TNFR1 expression in Mφ. This activity leads to caspase-3 activation and apoptosis, which restricts M. tuberculosis replication by efferocytosis.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank K. Kobayashi (Harvard Medical School, Boston, MA) for MyD88−/− mice.

Footnotes

  • P.J. and S.M.B. conceived of and designed the experiments, analyzed the data, and wrote the paper; P.J., I.S.-O., and T.N. performed the experiments; A.C.A., V.K.K., and H.G.R. provided reagents and intellectual input.

  • This work was supported by National Institutes of Health Grants R01 AI085669 and AI098637 (to S.M.B.), R01 AI072143 and R01 AI 073774 (to H.G.R.), and T Cell-Immunoglobulin and Mucin-Domain-Containing Molecule program project grant P01 AI 073748 (to V.K.K.); and the American Lung Association postdoctoral research training fellowship (RT-123085-N) and Harvard University Center for AIDS Research Scholar Award (to P.J.). The Harvard University Center for AIDS Research Scholar Award (to P.J.) is a National Institutes of Health–funded program (P30 AI060354), which is supported by the following National Institutes of Health Co-funding and Participating Institutes and Centers: National Institute of Allergy and Infectious Diseases; National Cancer Institute; National Institute of Child Health and Human Development; National Heart, Lung, and Blood Institute; National Institute on Drug Abuse; National Institute of Mental Health; National Institute on Aging; National Center for Complementary and Alternative Medicine; Fogarty International Center; and Office of AIDS Research.

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    −/−
    knockout
    FL
    full-length
    Gal9
    Galectin-9
    HMDM
    human monocyte-derived Mφ
    Hu IgG
    human IgG
    LDH
    lactate dehydrogenase
    Mφ
    macrophage
    MDM
    monocyte-derived Mφ
    MOI
    multiplicity of infection
    PS
    phosphatidylserine
    s
    soluble
    Tim3
    T cell–Ig and mucin-domain–containing molecule-3
    Tim3–Ig
    Tim3–Ig fusion protein
    WT
    wild-type.

  • Received September 25, 2012.
  • Accepted February 13, 2013.

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