The type I IFNs (IFN-α and -β) are important for host defense against viral infections. In contrast, their role in defense against nonviral pathogens is more ambiguous. In this article, we report that IFN-β signaling in murine bone marrow–derived macrophages has a cell-intrinsic protective capacity against Mycobacterium tuberculosis via the increased production of NO. The antimycobacterial effects of type I IFNs were mediated by direct signaling through the IFN-α/β–receptor (IFNAR), as Ab-mediated blocking of IFNAR1 prevented the production of NO. Furthermore, M. tuberculosis is able to inhibit IFNAR-mediated cell signaling and the subsequent transcription of 309 IFN-β–stimulated genes in a dose-dependent way. The molecular mechanism of inhibition by M. tuberculosis involves reduced phosphorylation of the IFNAR-associated protein kinases JAK1 and TYK2, leading to reduced phosphorylation of the downstream targets STAT1 and STAT2. Transwell experiments demonstrated that the M. tuberculosis–mediated inhibition of type I IFN signaling was restricted to infected cells. Overall, our study supports the novel concept that M. tuberculosis evolved to inhibit autocrine type I IFN signaling to evade host defense mechanisms.

Type I IFNs are innate cytokines that are best known for their ability to induce an antiviral state in cells (1, 2). Upon binding to their shared receptor, IFN-α/β–receptor (IFNAR), a heterodimer composed of IFNAR1 and IFNAR2 transmembrane proteins, the receptor-associated tyrosine kinases JAK1 and TYK2 are activated, and this leads to the phosphorylation and activation of STAT1 and STAT2. Activated STAT1 can homodimerize, translocate to the nucleus, and bind to IFN-γ–activated sites to promote gene transcription of IFN-stimulated genes (ISGs). Alternatively, STAT1 will associate with STAT2 and IRF9 to form the transcription factor ISG factor 3 (ISGF3), which then translocates to the nucleus to bind to IFN-stimulated response elements of ISGs and induce their expression (3, 4).

Although type I IFNs clearly have a protective function during viral infection, the role of these cytokines during bacterial or protozoan infections is more ambiguous (2, 46). IFN-β is detrimental to the host during Mycobacterium tuberculosis infections (716). Despite the various outcomes of the type I IFN response to infection, it is well-documented that many intracellular, nonviral pathogens elicit a host response that leads to the increase in IFN-β production (2, 4, 5). Multiple cell-surface (TLR) and intracellular (e.g., retinoic acid inducible gene I) receptors recognize microbial products and initiate signaling pathways that activate IRF3, IRF7, or AP1 to induce transcription of type I IFN genes (2, 4, 5). In particular, M. tuberculosis gains access to the host cell cytosol via their ESX-1 type VII secretion system, in which secreted extracellular M. tuberculosis DNA (eDNA) binds to the cyclic GMP–AMP synthase (cGAS) that subsequently activates the STING/TBK1/IRF3 pathway, leading to the increased transcription of type I IFN genes (1721). The secretion of bacterial cyclic-di-AMP (c-di-AMP) can also mediate the cGAS-independent activation of the STING pathway (22, 23). Finally, M. tuberculosis can induce IFN-β production through mitochondrial stress and subsequent release of mitochondrial DNA (mtDNA), which activates the STING pathway (24).

The potential of nonviral pathogens to inhibit cell signaling via the IFNAR has not been studied in great detail. One reason for this is probably that the infected host cell detects the pathogen and responds by increased synthesis of IFN-β, which confounds the analysis. To overcome this problem, we used bone marrow–derived macrophages (BMDMs) from Ifn-β−/− knockout mice and investigated the effect of IFN-β on survival of M. tuberculosis and the capacity of M. tuberculosis to inhibit IFNAR-mediated cell signaling.

Ifn-β−/− mice were originally obtained by Dr. E. Fish (University of Toronto) and are described in (25). Ifnar1−/− mice were obtained from Dr. K. D. Mayer-Barber (National Institutes of Health [NIH]). C57BL/6J and NO synthase 2 (Nos2)−/− mice were obtained from The Jackson Laboratory. All animal studies were approved by the Institutional Animal Care and Use Committee and were conducted in accordance with the NIH. BMDMs were prepared from bone marrow cells flushed from the femurs and tibia of mice that were cultured in DMEM supplemented with 10% heat-inactivated FCS, 1% penicillin/streptomycin, and 20% L929 supernatant for a period of 6 d prior to infection. The Raw264.7-derived, Irf3-deficient, and IFNAR-signaling reporter cell line (RAW-Lucia ISG-KO-IRF3) is commercially available, and measurement of reporter activity was performed according to the manufacturer’s protocol (InvivoGen).

All animals were handled in accordance with the NIH guidelines for the housing and care of laboratory animals, and the studies were approved by the Institutional Animal Care and Use Committee at the University of Maryland (RJAN1702).

M. smegmatis (mc2155), M. bovis BCG-Pasteur, and M. tuberculosis H37Rv (25618; American Type Culture Collection) strains were obtained from Dr. W. R. Jacobs Jr. (Albert Einstein College of Medicine). M. kansasii strain Hauduroy (12478) was obtained from the American Type Culture Collection. Bacterial strains were grown in 7H9 media supplemented with 10% Middlebrook ADC growth supplement, 0.5% glycerol, and 0.05% Tween 80. Hygromycin (50 μg/ml) and kanamycin (40 μg/ml) were added to the mutant and complemented strain cultures, respectively.

Bacterial infections of BMDMs were performed as previously described (26). Postinfection, cells were incubated in media containing 100 μg/ml gentamicin in the absence or presence of IFN-β or IFN-γ (PeproTech). For assays using the RAW-Lucia ISG-KO-IRF3 reporter cell line, cells were infected at multiplicity of infection (MOI) 10 and stimulated with the indicated amount of IFN-β. For measurement of bacterial survival in macrophages, a total of 0.5 million Ifn-β−/−, Nos2−/−, Ifnar1−/−, or C57BL/6J BMDMs were seeded in 24-well plates and infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h. Selected BMDMs were then treated with 1000 pg/ml IFN-β twice a day for a total of 4 d. Cells were lysed at indicated time points with 0.1% Triton X-100 in PBS, and serial dilutions were plated on 7H11 agar plates (Difco). CFUs were counted after 15–20 d of incubation at 37°C.

Six-well transwells with a 0.4-μM membrane (Corning) were allowed to equilibrate in medium overnight before seeding cells. A total of 3 × 106Ifn-β−/− BMDMs were seeded into the upper transwell, and 3 × 106 RAW-Lucia ISG-KO-IRF3 cells were seeded in the lower transwell. Infections were performed as described earlier in the upper transwell. Selected conditions were then treated with 200 pg/ml IFN-β in the upper and lower transwells for the indicated time points.

Experiments with neutralizing Abs were performed using InVivoPlus anti-mouse IFNAR1 and InVivoMAB mouse IgG1 isotype control (Bio X Cell). Abs were added to cells at a final concentration of 20 μg/ml once, immediately after the 4-h infection period.

Cells were preincubated with DMSO as a solvent control or increasing amounts of InSolution JAK inhibitor I (EMD Millipore) for 1 h. Following preincubation with the inhibitor, cells were treated with 200 pg/ml IFN-β for 4 h, and whole-cell lysates were collected.

Whole-cell lysates were obtained by lysing cells with RIPA buffer containing protease and phosphatase inhibitor mixtures (Roche). Protein concentration was measured using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific), and proteins were subjected to SDS-PAGE followed by immunoblotting as described (26). Abs were detected using SuperSignal West Femto chemiluminescent substrate (34095; Thermo Fisher Scientific), and images were acquired using the LAS-300 imaging system (Fuji). All Western blots were performed at least three times, and the image of one representative result is shown. ImageJ software (NIH) was used for densitometry quantification as described in the figure legends for each Western blot.

Postinfection, BMDMs were blocked with 5% FCS and rat anti-mouse CD16/CD32 Fc Block (553141; BD Biosciences) for 15 min followed by incubation with either PE-conjugated mouse anti-IFNAR1 (127311; BioLegend) or PE-conjugated goat anti-IFNAR2 (FAB1083P; R&D Systems) for 30 min on ice. PE-conjugated mouse IgG1 (400111; BioLegend) and goat IgG1 (IC108P; R&D Systems) were used as isotype controls. Protein levels were quantified by acquiring 25,000 cells using the Accuri C6 flow cytometer and software (BD Biosciences). Histograms were processed using FlowJo software version 10 (BD Biosciences).

NO production was quantified in cell culture supernatants by the Griess reagent kit, which measures the NO derivate nitrite (G7921; Thermo Fisher Scientific) at the indicated time points. Absorbance was measured at 548 nm using a microplate reader (BioTek Instruments). Nitrite concentration was determined using a sodium nitrite standard curve (0–100 μM). Culture supernatants were pooled from three replicate wells per experiment for all infections. All samples were assayed in technical duplicates, and three independent experiments were performed.

Ifn-β−/− BMDMs were infected or not as indicated previously with M. tuberculosis H37Rv at an MOI of 3 for 4 h. At 4 h postinfection (hpi), cells were lysed with 1 ml of TRIzol (Ambion). RNA was extracted with chloroform, precipitated with 100% isopropanol, and washed with 70% ethanol. Purified RNA was treated with Turbo DNAse (Ambion) for 1 h.

RNA sequencing (RNAseq) libraries were prepared using the Illumina ScriptSeq v2 Library Preparation Kit according to the manufacturer’s protocol. Library quality was assayed by bioanalyzer (Agilent) and quantified by quantitative PCR (KAPA Biosystems). Sequencing was performed on an Illumina HiSeq 1500 generating 100-bp paired-end reads. RNAseq read quality was assessed using FastQC (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/), and low-quality bp were removed using Trimmomatic (27). The Ensembl Mus musculus GRCm38 reference genome (version 76) was downloaded from the Ensembl Web site (28), and reads were mapped to the genome using TopHat2 (29). HTSeq (30) was used to quantify expression at the gene level. Count tables were loaded into R/Bioconductor (31), log2-transformed, counts-per-million– and quantile-normalized (31), and variance bias was corrected for using Voom (32). Next, batch adjustment was performed using ComBat (33). We performed Pearson correlation, Euclidean distance, and principal component analyses, which revealed the presence of a single outlier M. tuberculosis sample (HPGL0627), which was removed from subsequent analyses (Supplemental Table I). The differential gene expression was measured for each of several contrasts: 1) uninfected (UI) versus UI plus IFN-β and 2) M. tuberculosis plus IFN-β versus UI plus IFN-β (Supplemental Table I). To determine which IFN-β–stimulated genes were specifically deregulated during infection with M. tuberculosis, the intersection of the set of genes found to be differentially expressed in both the UI versus UI plus IFN-β and M. tuberculosis plus IFN-β versus UI plus IFN-β contrasts was taken (Supplemental Table I). All raw RNAseq data were submitted to the Sequence Read Archive and can found at https://www.ncbi.nlm.nih.gov/sra/SRP130272.

Ifn-β−/− BMDMs were infected as described previously and treated with 50 pg/ml IFN-β. Additional 50 pg/ml IFN-β was added at 3 hpi, and cell culture supernatants were collected at 6 hpi. Concentrations of selected cytokines were determined using a custom ProcartaPlex magnetic bead–based multiplex assay (Thermo Fisher Scientific) on the Luminex MAGPIX platform according to the manufacturer’s instructions. IFN-α (PBL Assay Science), IFN-β (BioLegend), CCL12 (R&D Systems), and CCL3 (R&D Systems) protein levels were measured using ELISAs.

Densitometric analysis of Western blots was performed using ImageJ. Bands were normalized to the signal of the loading control (β-actin or GAPDH). Densitometric ratios are expressed relative to the indicated experimental group in each figure.

Statistical analysis was performed on at least three independent experiments using GraphPad Prism 7.0 software and one-way ANOVA with Tukey posttest or Student t test, and representative results of triplicate values are shown with SD unless otherwise noted in the legends. The range of p values is indicated as follows: *0.01 < p < 0.05, **0.001 < p < 0.01, ***0.0001 < p < 0.001, and ****p < 0.0001.

The analysis of the importance of IFN-β for host defense during infection with nonviral pathogens has proven to be complex because, during in vivo infections, host genetic components, tissue environment, and actual doses of IFN-β can all affect outcomes (2, 4, 6). To determine the differences in type I IFNs between nonvirulent and virulent mycobacterial species, we infected C57BL/6 BMDMs with M. smegmatis, M. tuberculosis H37Rv, or M. tuberculosis CDC1551 and measured levels of IFN-α and IFN-β. M. smegmatis produced a significant amount of IFN-α early in infection, and this increased production was sustained at later time points. In contrast, infection with both virulent M. tuberculosis strains produced much less IFN-α (Fig. 1A). Similarly, infection with M. smegmatis induced a robust production of IFN-β early during infection, and this was sustained at a later time point. Both virulent strains, in contrast, produced ∼10-fold less IFN-β than M. smegmatis (Fig. 1B). Thus, nonvirulent mycobacteria induce a stronger type I IFN response than their virulent counterparts. To investigate the importance of this finding, as well as the contribution of high levels of IFNAR signaling, we used a reductionist approach by eliminating bystander cell, host tissue effects, and the host cell production of IFN-β in response to infection using BMDMs from Ifn-β−/− mice (25). We infected Ifn-β−/− BMDMs with M. tuberculosis in the presence or absence of 1 ng/ml IFN-β, corresponding to the amount of IFN-β produced by infection with M. smegmatis. The bacterial burden was measured every 24 h for a total of 96 h by counting CFUs. During M. tuberculosis infection, we found that reduction in bacterial burden in IFN-β–treated cells was minimal at 24 and 48 hpi but steadily increased between 48 and 96 hpi. At 96 hpi, IFN-β treatment reduced bacterial burden by ∼50% (Fig. 1C). We also determined that IFN-β–treated M. tuberculosis–infected cells exhibited less necrosis compared with untreated infected cells because an increased release in gentamicin–containing medium could have otherwise accounted for the reduction of CFUs (Fig. 1D). These results demonstrate that IFN-β treatment promotes host defense against intracellular microbes during infection of primary macrophages. This study demonstrated the direct antimicrobial activity of IFNAR signaling on M. tuberculosis viability in a system in which the infection itself does not produce IFN-β.

FIGURE 1.

IFN-β treatment has antimicrobial activity against M. tuberculosis. (A and B) C57BL/6 BMDMs were infected with M. smegmatis, M. tuberculosis H37Rv, or M. tuberculosis CDC1551 at MOI 10:1. Supernatants were collected, and protein levels of IFN-α (A) or IFN-β (B) were measured using ELISA. (C) Ifn-β−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h in the presence or absence of 1000 pg/ml IFN-β. Cells were lysed, and serial dilutions were plated to determine CFUs. (D) Cell necrosis was assayed at each indicated time point using an adenylate kinase release assay and is represented as fold change over untreated infected cells. (E) Whole-cell lysates were collected at the indicated time points and immunoblotted for NOS2. Band densities were normalized to GAPDH. Densitometry ratios are shown relative to the UI condition. (F) Nitrite levels from culture supernatants were determined using the Griess reagent. (G and H) WT or Nos2−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h in the presence or absence of 1000 pg/ml IFN-β. Bacterial burden was determined as in Fig. 1C. All data shown are presented as mean ± SEM of at least three independent experiments. *0.01 < p < 0.05, **0.001 < p < 0.01, ***0.0001 < p < 0.001, and ****p < 0.0001.

FIGURE 1.

IFN-β treatment has antimicrobial activity against M. tuberculosis. (A and B) C57BL/6 BMDMs were infected with M. smegmatis, M. tuberculosis H37Rv, or M. tuberculosis CDC1551 at MOI 10:1. Supernatants were collected, and protein levels of IFN-α (A) or IFN-β (B) were measured using ELISA. (C) Ifn-β−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h in the presence or absence of 1000 pg/ml IFN-β. Cells were lysed, and serial dilutions were plated to determine CFUs. (D) Cell necrosis was assayed at each indicated time point using an adenylate kinase release assay and is represented as fold change over untreated infected cells. (E) Whole-cell lysates were collected at the indicated time points and immunoblotted for NOS2. Band densities were normalized to GAPDH. Densitometry ratios are shown relative to the UI condition. (F) Nitrite levels from culture supernatants were determined using the Griess reagent. (G and H) WT or Nos2−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h in the presence or absence of 1000 pg/ml IFN-β. Bacterial burden was determined as in Fig. 1C. All data shown are presented as mean ± SEM of at least three independent experiments. *0.01 < p < 0.05, **0.001 < p < 0.01, ***0.0001 < p < 0.001, and ****p < 0.0001.

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It was previously shown that type I IFN signaling may lead to the induction of Nos2 gene expression and subsequent NO production in infected BMDMs (34). To investigate whether NO production plays a role in IFN-β–mediated clearance ex vivo, we determined if NOS2 was expressed at time points correlating with the reduction in bacterial burden during M. tuberculosis infection. At 72 hpi, NOS2 was upregulated in M. tuberculosis–infected cells, and NOS2 levels were further increased in M. tuberculosis–infected cells stimulated with IFN-β (Fig. 1E). At 96 hpi, the NOS2 protein levels were decreased compared with 72 hpi; however, there was still a significant upregulation in M. tuberculosis–infected cells stimulated with IFN-β compared with UI cells (Fig. 1E). Consequently, we measured NO levels in the different experimental groups to assess the effect of IFN-β and infection on NO production. In M. tuberculosis–infected cells treated with 1 ng/ml IFN-β, we noticed a sharp increase in nitrite levels, which is consistent with the observed increase in NOS2 levels (Fig. 1F). We then investigated whether IFN-β could promote host resistance in Nos2−/− BMDMs. We infected wild-type (WT) or Nos2−/− BMDMs with M. tuberculosis in the presence or absence of 1 ng/ml IFN-β. Infected BMDMs showed a significant decrease in CFUs at 96 hpi in IFN-β–treated cells (Fig. 1G, 1H). In contrast, we measured no significant decrease in IFN-β–treated Nos2−/− BMDMs at either 72 or 96 hpi (Fig. 1G, 1H). These results demonstrate that IFN-β treatment promotes host resistance to M. tuberculosis via the production of NO at later time points during ex vivo infection.

Next, we sought to determine the contribution of IFNAR signaling to NO production and subsequent M. tuberculosis killing (716). To determine whether IFNAR1 neutralization could be sustained throughout our experiment, we used a RAW264.7 cell line with luciferase expression driven by an ISG reporter. Cells were treated with a single dose of isotype or anti-IFNAR1 Ab and then treated daily with 1 ng IFN-β. Secreted luciferase was then measured every 24 h (Fig. 2A). Cells treated with an isotype Ab had robust production of luciferase in the presence of IFN-β, and luciferase levels increased throughout the course of the experiment (Fig. 2B). In contrast, cells treated with an anti-IFNAR1 Ab had suppressed signaling in the presence of IFN-β, comparable to unstimulated cells (Fig. 2B). These data show that we are effectively able to neutralize IFNAR1 signaling throughout the course of the experiment.

FIGURE 2.

IFNAR1 is required for IFN-β–induced NO production. (A) Schematic representation of the experiment. (B) IRF3-deficient RAW264.7 macrophages transfected with an IFN-β–responsive luciferase gene (InvivoGen) were treated with 20 μg/ml of the indicated neutralizing Abs. One nanogram per milliliter IFN-β was added to the cells every 24 h, and the amount of secreted luciferase was quantified at the indicated time points and plotted as relative light units (RLU). (C and D) Ifn-β−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h and treated with IFN-β as in Fig. 1. (C) CFUs were enumerated at the indicated time points. (D) Nitrite levels from culture supernatants were determined using the Griess reagent. (E and F) C57BL6/6 or Ifnar1−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h and treated with IFN-β as in Fig. 1. (E) CFUs were enumerated at the indicated time points. (F) Nitrite levels from culture supernatants were determined using the Griess reagent. All data shown are presented as mean ± SEM of at least three independent experiments. *0.01 < p < 0.05, **0.001 < p < 0.01, and ****p < 0.0001.

FIGURE 2.

IFNAR1 is required for IFN-β–induced NO production. (A) Schematic representation of the experiment. (B) IRF3-deficient RAW264.7 macrophages transfected with an IFN-β–responsive luciferase gene (InvivoGen) were treated with 20 μg/ml of the indicated neutralizing Abs. One nanogram per milliliter IFN-β was added to the cells every 24 h, and the amount of secreted luciferase was quantified at the indicated time points and plotted as relative light units (RLU). (C and D) Ifn-β−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h and treated with IFN-β as in Fig. 1. (C) CFUs were enumerated at the indicated time points. (D) Nitrite levels from culture supernatants were determined using the Griess reagent. (E and F) C57BL6/6 or Ifnar1−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h and treated with IFN-β as in Fig. 1. (E) CFUs were enumerated at the indicated time points. (F) Nitrite levels from culture supernatants were determined using the Griess reagent. All data shown are presented as mean ± SEM of at least three independent experiments. *0.01 < p < 0.05, **0.001 < p < 0.01, and ****p < 0.0001.

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To show the role of IFNAR1 signaling without the confounding factor of host-produced IFN-β, we infected Ifn-β−/− BMDMs with M. tuberculosis and then treated cells with isotype or anti-IFNAR1 Ab immediately after the 4-h infection period. Cells were then treated with IFN-β as in Fig. 1D–H. At 96 hpi, cells treated with the isotype Ab saw a significant reduction in CFUs when treated with IFN-β (Fig. 2C). Conversely, cells treated with anti-IFNAR1 Ab had no significant difference in CFUs when treated with IFN-β (Fig. 2C). Importantly, there was no difference in CFUs between unstimulated cells treated with isotype or anti-IFNAR1 Ab. We also measured nitrite levels to determine if IFNAR neutralization abrogated NO production. In the absence of exogenous IFN-β stimulation, cells treated with isotype or anti-IFNAR1 had similar levels of NO production during M. tuberculosis infection (Fig. 2D). However, when stimulated with IFN-β, cells treated with anti-IFNAR1 Ab failed to reach NO levels comparable to those of isotype-treated cells (Fig. 2D). These data suggest that signaling through IFNAR1 is responsible for the host-protective effects we see upon addition of exogenous IFN-β.

Although these data support a protective role for IFNAR1 in the absence of endogenous IFN-β production, it is uncertain whether there will be a protective role for exogenous IFN-β in the context of host cells that are able to produce IFN-β. To address this question, we examined the role of IFN-β in M. tuberculosis killing in C57BL/6 (WT) or Ifnar1−/− BMDMs. At 96 hpi, Ifnar1−/− BMDMs had fewer CFUs than WT BMDMs infected with M. tuberculosis (Fig. 2E). However, Ifnar1−/− BMDMs treated with IFN-β did not see a reduction in CFUs, similar to the results seen with Ab neutralization (Fig. 2C, 2E). Interestingly, WT BMDMs treated with IFN-β had fewer CFUs than both treated and untreated Ifnar1−/− BMDMs. In line with this, Ifnar1−/− BMDMs failed to produce an amount of NO comparable to that produced by WT BMDMs (Fig. 2F). Taken together, these data demonstrate that there is a context-dependent role for IFN-β and IFNAR signaling during M. tuberculosis infection.

Viruses are well known to evade host-protective IFN-β responses by suppressing signaling via IFNAR (1, 2). Our results on the host-protective effect of IFN-β on M. tuberculosis infection prompted us to investigate the potential of M. tuberculosis to inhibit IFNAR signaling. To this purpose, we used a reporter RAW264.7 cell line (InvivoGen) that is deficient in Irf3 and has an ISG promoter in front of a reporter gene for easy quantification of IFNAR signaling. We first determined the capacity of these pathogens to induce activation of the reporter in the absence of an external IFN-β stimulus. M. smegmatis, M. tuberculosis H37Rv, and M. tuberculosis CDC1551 did not induce activation of the ISG reporter in the absence of external stimulation (data not shown). We also determined that the infection with the different mycobacterial species does not induce the production of IFN-β via ELISA, because these cells are deficient in IRF3 (data not shown). Next, we added increasing amounts of IFN-β to infected or UI cells and normalized the response to the response obtained in UI cells (Fig. 3A). Overall, both virulent M. tuberculosis strains consistently showed ∼50–60% of inhibition, whereas infection with M. smegmatis had only a minor effect (10–15%) at higher doses of IFN-β (Fig. 3A). Notably, this effect was reversed at 1000 pg/ml, suggesting there is a limit to the amount of signaling M. tuberculosis can inhibit. To our best knowledge, the described experiments are the first to demonstrate the capacity of M. tuberculosis to inhibit IFNAR signaling. The lack of investigation into this capacity of the pathogens might be explained by the proposed overall role of type I IFNs in the exacerbation of disease outcomes (2, 46).

FIGURE 3.

M. tuberculosis inhibits type I IFN signaling. (A) IRF3-deficient RAW264.7 macrophages transfected with an IFN-β–responsive luciferase gene were infected and treated with the indicated concentrations of IFN-β. After 20 hpi, the amount of secreted luciferase was quantified. ISG induction is represented as the fold change in relative light units (RLUs) compared with UI cells. (BE) Ifn-β−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h, washed, and treated with 50 pg/ml IFN-β for an additional 4 h before either total RNA or proteins were harvested. (B) Heatmap of downregulated genes selected for follow-up studies. Log-transformed expression ratios, as determined via RNAseq for M. tuberculosis plus IFNβ/UI plus IFN-β, are plotted for each gene. (C) Whole-cell lysates were collected and immunoblotted for IFI204, IFIT1, Rnf144A, and Rnf144B. Band density was normalized to β-actin. Densitometric ratios are relative to the UI plus IFN-β condition. (D) Cell lysates were collected and immunoblotted for either MX1 or IIGP1 and normalized to GAPDH. Densitometric ratios are relative to the UI plus IFN-β condition. (E) Supernatants were analyzed for levels of CCL12 using ELISA. All data shown are presented as mean ± SEM of at least three independent experiments. **0.001 < p < 0.01 and ****p < 0.0001.

FIGURE 3.

M. tuberculosis inhibits type I IFN signaling. (A) IRF3-deficient RAW264.7 macrophages transfected with an IFN-β–responsive luciferase gene were infected and treated with the indicated concentrations of IFN-β. After 20 hpi, the amount of secreted luciferase was quantified. ISG induction is represented as the fold change in relative light units (RLUs) compared with UI cells. (BE) Ifn-β−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h, washed, and treated with 50 pg/ml IFN-β for an additional 4 h before either total RNA or proteins were harvested. (B) Heatmap of downregulated genes selected for follow-up studies. Log-transformed expression ratios, as determined via RNAseq for M. tuberculosis plus IFNβ/UI plus IFN-β, are plotted for each gene. (C) Whole-cell lysates were collected and immunoblotted for IFI204, IFIT1, Rnf144A, and Rnf144B. Band density was normalized to β-actin. Densitometric ratios are relative to the UI plus IFN-β condition. (D) Cell lysates were collected and immunoblotted for either MX1 or IIGP1 and normalized to GAPDH. Densitometric ratios are relative to the UI plus IFN-β condition. (E) Supernatants were analyzed for levels of CCL12 using ELISA. All data shown are presented as mean ± SEM of at least three independent experiments. **0.001 < p < 0.01 and ****p < 0.0001.

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Signaling via the IFNAR induces the formation of the ISGF3 complex and/or STAT1:STAT1 homodimerization, which both translocate into the nucleus and bind to IFN-stimulated response elements or IFN-γ–activated sites, respectively, leading to the transcription of hundreds of genes involved in a variety of immunological functions (3). Consequently, we hypothesized that by inhibiting type I IFN signaling, M. tuberculosis can manipulate the expression of host genes to promote its intracellular survival. To investigate this, we used RNAseq technology to identify IFN-β–regulated genes that are up- or downregulated during M. tuberculosis infection in Ifn-β−/−–derived BMDMs. We analyzed the following experimental groups: 1) UI and untreated, 2) UI IFN-β–treated, and 3) M. tuberculosis H37Rv–infected and treated BMDMs (Supplemental Table I). The RNAseq data were reproducible, as indicated by principal component analysis, hierarchal clustering, and Pearson correlation (Supplemental Fig. 1). All three analyses depicted a satisfactory degree of clustering between biological replicates, with the exception of one outlier (HPGL0627) that was excluded from further analysis (Supplemental Fig. 1, Supplemental Table I). To identify IFN-β–stimulated genes that exhibited differential expression in M. tuberculosis–infected BMDMs, we first characterized all of the IFN-β–regulated genes in our experimental system. To that purpose, we compared the gene expression of UI, untreated to UI, IFN-β–treated conditions. We identified a total of 1144 IFN-β–stimulated genes and 956 genes with reduced expression (>2-fold change) (Supplemental Fig. 2A, Supplemental Table I). Next, to identify all of the genes that are impacted by M. tuberculosis infection, we compared gene expression levels between the UI plus IFN-β condition and the M. tuberculosis H37Rv–infected and treated condition and found 1296 upregulated and 1294 downregulated (>2-fold change and an adjusted p value <0.05) genes (Supplemental Fig. 2B, Supplemental Table I). Finally, M. tuberculosis infection causes the deregulation of many genes that are not regulated by IFN-β but will be included in the UI, IFN-β–treated versus M. tuberculosis H37Rv–infected and treated contrast. The overlap between these two gene sets was determined to be 309 genes with reduced expression and 170 with increased expression for a total of 479 deregulated genes (Supplemental Fig. 2C, Supplemental Table I).

To extend the results of our RNAseq analysis, we determined protein expression and secretion levels of selected targets in Ifn-β−/− BMDMs infected with M. tuberculosis in the presence or absence of IFN-β (Fig. 3B). The selection of candidate gene products for follow-up studies was based on the magnitude of deregulation and the availability of Abs (see list of 479 genes in Supplemental Table I). Rnf144A and Rnf144B proteins were not significantly upregulated upon treatment with IFN-β (Fig. 3C), which may reflect a low fold-increase at the RNA level as illustrated in the heatmap (Fig. 3B); however, IFI204, IFIT1, MX1, and IIGP1 were strongly induced after treatment with IFN-β (Fig. 3C, 3D). For all targets, we observed some degree of downregulation in IFN-β–treated cells infected with M. tuberculosis. In addition, we used ELISA to demonstrate the strong inhibition of IFN-β–driven secretion of CCL12 by M. tuberculosis infection (Fig. 3E). In conclusion, these data show a strong correlation of the RNAseq analysis with protein data for the subset of 309 IFN-β–regulated genes that are repressed by M. tuberculosis infection (Fig. 3B, Supplemental Table I). We additionally investigated some of the 170 genes that were strongly upregulated in the 479-gene set (Supplemental Table I), characterized several cytokine/chemokine targets using a combination of both multiplexed and regular ELISA, and determined that M. tuberculosis infection of BMDMs without the addition of IFN-β upregulated secretion levels of all assayed proteins (Supplemental Fig. 3). For IL-1β, TNF-α, and CXCL1, there seemed to be additive effects in M. tuberculosis plus IFN-β–treated cells when compared with UI plus IFN-β alone and M. tuberculosis minus IFN-β–mediated cytokine induction, whereas there was an antagonistic effect of the M. tuberculosis infection for IL-27, IL-1α, CCL5, and CCL3, suggesting that the induction of these cytokines in M. tuberculosis–infected cells is independent of IFNAR signaling and that their upregulation caused by the addition of IFN-β can be inhibited by M. tuberculosis (Supplemental Fig. 1). Overall, this analysis shows that this subset of 170 of M. tuberculosis–deregulated genes after IFN-β addition that is upregulated provides fewer insights because many of the genes seem to be upregulated by M. tuberculosis infection alone.

We discovered that M. tuberculosis inhibits signaling via IFNAR, and then we investigated further at what level of the signaling cascade the inhibition occurs. Several viral pathogens have evolved mechanisms to evade the IFN-β response by promoting the degradation of IFNAR (35, 36). We found using flow cytometry that surface expression levels of IFNAR1 and IFNAR2 remained unchanged by M. tuberculosis infection (Fig. 4A). After stimulation of IFNAR1/R2 by IFN-β, the cytosolic protein tyrosine kinases JAK1 and TYK2 are recruited and phosphorylated. We showed that M. tuberculosis inhibited tyrosine phosphorylation of both TYK2 and JAK1 as early as 20 min postinfection (Fig. 4B, 4C). Because JAK1 is involved in both type I and type II IFN signaling, we also analyzed phosphorylation levels of JAK1 and JAK2, another IFN-γ–activated protein kinase. In IFN-γ–treated and M. tuberculosis–infected cells, there was no significant change in JAK1 or JAK2 phosphorylation levels, showing that the inhibition of JAK1 phosphorylation is specific to type I IFN signaling (Fig. 4D, 4E). To show that inhibition of JAK1 is important for downstream signaling, we determined whether inhibition of JAK1 disrupts downstream signaling. BMDMs treated with increasing amounts of a JAK1 inhibitor show decreased levels of STAT1 phosphorylation compared with cells treated with DMSO (Fig. 4F).

FIGURE 4.

M. tuberculosis inhibits type I but not type II IFN–mediated activation of TYK2 and JAK1. (A) Ifn-β−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h, and flow cytometry was conducted at 4 hpi to measure surface receptor expression levels of IFNAR1 and IFNAR2. (B and C) Ifn-β−/− BMDMs were infected as described in the presence of 300 pg/ml IFN-β. Cell lysates were collected at 20 min postinfection and immunoblotted for p-JAK1 (Y1022/1023), total JAK1, p-TYK2 (Y1054/1055), and total TYK2. (D and E) Ifn-β−/− BMDMs were infected as described in the presence of 300 pg/ml IFN-γ. Cell lysates were collected at 20 min postinfection and immunoblotted for p-JAK1 (Y1022/1023), total JAK1, p-TYK2 (Y1054/1055), and total TYK2. (F) Ifn-β−/− BMDMs were treated with DMSO or increasing amounts of a JAK inhibitor for 1 h and then stimulated with 200 pg/ml IFN-β for 4 h. Cell lysates were then collected and immunoblotted for p-STAT1 (Y701), total STAT1, and β-actin. Densitometry was performed using ImageJ software, and phosphorylated protein bands were normalized to loading control for each condition. Densitometric ratios are relative to the UI plus IFN-β or the UI plus IFN-γ conditions. Data and densities shown represent one representative experiment out of three.

FIGURE 4.

M. tuberculosis inhibits type I but not type II IFN–mediated activation of TYK2 and JAK1. (A) Ifn-β−/− BMDMs were infected with M. tuberculosis H37Rv at an MOI of 3 for 4 h, and flow cytometry was conducted at 4 hpi to measure surface receptor expression levels of IFNAR1 and IFNAR2. (B and C) Ifn-β−/− BMDMs were infected as described in the presence of 300 pg/ml IFN-β. Cell lysates were collected at 20 min postinfection and immunoblotted for p-JAK1 (Y1022/1023), total JAK1, p-TYK2 (Y1054/1055), and total TYK2. (D and E) Ifn-β−/− BMDMs were infected as described in the presence of 300 pg/ml IFN-γ. Cell lysates were collected at 20 min postinfection and immunoblotted for p-JAK1 (Y1022/1023), total JAK1, p-TYK2 (Y1054/1055), and total TYK2. (F) Ifn-β−/− BMDMs were treated with DMSO or increasing amounts of a JAK inhibitor for 1 h and then stimulated with 200 pg/ml IFN-β for 4 h. Cell lysates were then collected and immunoblotted for p-STAT1 (Y701), total STAT1, and β-actin. Densitometry was performed using ImageJ software, and phosphorylated protein bands were normalized to loading control for each condition. Densitometric ratios are relative to the UI plus IFN-β or the UI plus IFN-γ conditions. Data and densities shown represent one representative experiment out of three.

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Type I and type II IFNs induce transcription of different subsets of genes, but STAT1 phosphorylation occurs in both signaling pathways (3). The canonical signaling pathways are defined as follows: type I IFNs induce the heterodimerization of STAT1 and STAT2, while type II IFNs (IFN-γ) induce homodimerization of STAT1, although type I IFNs can also induce STAT1 homodimerization (3). We performed an infection comparing STAT1 tyrosine phosphorylation levels in IFN-β–treated Ifn-β−/− BMDMs infected with virulent M. tuberculosis strains H37Rv or CDC1551 and observed that both could similarly inhibit STAT1 phosphorylation, suggesting that this mechanism of host cell manipulation is shared among virulent M. tuberculosis strains (Fig. 5A). We also showed that M. tuberculosis did not inhibit IFN-γ–dependent phosphorylation of STAT1 (Fig. 5B), which is consistent with previously published results (37) and suggests a separate molecular pathway engaged by M. tuberculosis for the inhibition of IFNAR signaling.

FIGURE 5.

M. tuberculosis inhibits phosphorylation of STAT1 and STAT2. (A and B) Ifn-β−/− BMDMs were infected with either M. tuberculosis strain H37Rv or CDC1551 at an MOI of 3 for 4 h and treated with 50 pg/ml IFN-β (A) or 50 pg/ml IFN-γ (B). Whole-cell lysates were collected at 4 hpi and immunoblotted for p-STAT1 (Y701) and total STAT1. (C) Ifn-β−/− BMDMs were infected with M. tuberculosis at an MOI of 3 for 4 h in the presence or absence of 300 pg/ml IFN-β. Whole-cell lysates were collected at 20 min postinfection and immunoblotted for p-STAT2 (Y690), total STAT2, p-STAT3 (Y705), or total STAT3 as indicated. (D) Cells were infected with the indicated Mycobacteria strains at an MOI of 3 for 4 h and treated with 50 pg/ml IFN-β. Whole-cell lysates were collected at 4 hpi and immunoblotted for p-STAT1 and total STAT1. (E and F) Ifn-β−/− BMDMs were infected with M. tuberculosis at an MOI of 3 for 4 h and treated with 1 ng/ml IFN-β. Whole-cell lysates were collected at 4 hpi and immunoblotted for p-STAT1, total STAT1, p-STAT2, or total STAT2 as indicated. Densitometric ratios are normalized to the loading control and are shown relative to the UI plus IFN-β or the UI plus IFN-γ conditions.

FIGURE 5.

M. tuberculosis inhibits phosphorylation of STAT1 and STAT2. (A and B) Ifn-β−/− BMDMs were infected with either M. tuberculosis strain H37Rv or CDC1551 at an MOI of 3 for 4 h and treated with 50 pg/ml IFN-β (A) or 50 pg/ml IFN-γ (B). Whole-cell lysates were collected at 4 hpi and immunoblotted for p-STAT1 (Y701) and total STAT1. (C) Ifn-β−/− BMDMs were infected with M. tuberculosis at an MOI of 3 for 4 h in the presence or absence of 300 pg/ml IFN-β. Whole-cell lysates were collected at 20 min postinfection and immunoblotted for p-STAT2 (Y690), total STAT2, p-STAT3 (Y705), or total STAT3 as indicated. (D) Cells were infected with the indicated Mycobacteria strains at an MOI of 3 for 4 h and treated with 50 pg/ml IFN-β. Whole-cell lysates were collected at 4 hpi and immunoblotted for p-STAT1 and total STAT1. (E and F) Ifn-β−/− BMDMs were infected with M. tuberculosis at an MOI of 3 for 4 h and treated with 1 ng/ml IFN-β. Whole-cell lysates were collected at 4 hpi and immunoblotted for p-STAT1, total STAT1, p-STAT2, or total STAT2 as indicated. Densitometric ratios are normalized to the loading control and are shown relative to the UI plus IFN-β or the UI plus IFN-γ conditions.

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Besides STAT1, other STAT isoforms may be stimulated by type I IFN signaling. STAT2 becomes phosphorylated and heterodimerizes with p-STAT1 and IRF9 to form the ISGF3 complex, which translocates into the nucleus to induce transcription of genes containing IFN-stimulated regulatory elements (3). In addition, it has also been shown that STAT3 can be induced by type I IFNs to regulate transcription of different gene subsets (38). We discovered that M. tuberculosis also inhibited the tyrosine phosphorylation of STAT2 (Fig. 5C). However, M. tuberculosis actually induced phosphorylation of STAT3 (Fig. 5C). STAT3 phosphorylation can inhibit type I IFN–mediated signaling, although this occurs at the level of nuclear translocation (39, 40). Thus, STAT3 activation is most likely not involved in our observed inhibition of TYK2/JAK1 phosphorylation by M. tuberculosis.

The levels of IFN-β production by mycobacteria-infected cells vary depending on the mycobacterial species and, in the context of M. tuberculosis, the specific strain that is used for the infection (24, 41, 42). We sought to test if the difference in type I IFN production observed for the different mycobacterial species correlated with their variable capacity to inhibit IFNAR-mediated cell signaling. Here we observed a more modest decrease in the relative STAT1 phosphorylation in cells infected with the vaccine strains M. bovis BCG and M. kansasii but only a minor reduction upon infection with M. smegmatis (Fig. 5D). Notably, the capacity of M. tuberculosis to inhibit STAT1 and STAT2 phosphorylation was also reversed upon addition of a high dose of IFN-β, supporting the data using our reporter cell line (Fig. 5E, 5F).

To determine whether inhibition of IFN-β signaling is specific to M. tuberculosis–infected cells and not due to the secretion of a soluble host factor, we sought to determine if there was an effect in bystander cells. To address this, we used a transwell system in which the upper transwell was seeded with Ifn-β−/− BMDMs and the lower transwell was seeded with our ISG reporter cell line (Fig. 6A). To confirm whether our experimental setup could transmit soluble factors that could result in a bystander effect, we measured IFN-β translocation from the upper transwell. The upper transwells were seeded with C57BL/6 BMDMs, with no cells in the lower transwell, and infected with M. smegmatis or treated with 1.5 ng of IFN-β. At 20 h poststimulation, supernatants from the upper and lower transwells were collected, and IFN-β levels were measured by ELISA. In both conditions, IFN-β levels equalized between the upper and lower transwells, showing that our experimental setup allowed for transmition of host-produced factors that could induce bystander effects (Fig. 6B). To determine if there was an effect on bystander cells, the upper transwell was then infected or not with M. tuberculosis, and both wells were stimulated or not with IFN-β as earlier (Fig. 6A). In this system, a reduction of secreted luciferase from the cells in the lower transwell would suggest that there is indeed a bystander effect. In our hands, we saw equal levels of reporter activity in bystander cells exposed to both UI and infected Ifn-β−/− BMDMs, suggesting that the inhibition is specific to infected cells (Fig. 6C). Western blots of Ifn-β−/− BMDM lysates (Fig. 6D) and the ISG reporter cell line (Fig. 6E) confirm that we do still see inhibition of STAT1 phosphorylation in infected Ifn-β−/− cells; however, STAT1 phosphorylation of bystander cells is not affected by the infection status of the Ifn-β−/− BMDMs (Fig. 7).

FIGURE 6.

M. tuberculosis does not inhibit type I IFN signaling in bystander cells. (A) Schematic of transwell experiments. (B) C57BL/6 BMDMs were seeded into the upper transwell, with no cells in the lower transwell. Upper transwells were then infected with M. smegmatis at an MOI of 3 for 4 h or treated with 1.5 ng of IFN-β. Twenty hours poststimulation, supernatants were harvested from the upper and lower transwells, and IFN-β protein levels were determined by ELISA. (C) Upper transwells were infected or not with M. tuberculosis at an MOI of 3 for 4 h, and then the upper and lower transwells were treated with 200 pg/ml IFN-β for 4 h. Cells in the lower transwell were lysed by addition of Triton X-100, and the amount of luciferase was quantified. (D) Whole-cell lysates from the upper transwell of Ifn-β−/− BMDMs were collected at 4 hpi and immunoblotted for p-STAT1, total STAT1, or β-actin as indicated. (E) Whole-cell lysates from the lower transwell were collected at 4 hpi and immunoblotted for p-STAT1, total STAT1, or β-actin as indicated. UI or M. tuberculosis refers to the infection condition of the upper transwell. Densitometric ratios are normalized to the loading control and are shown relative to the UI plus IFN-β condition.

FIGURE 6.

M. tuberculosis does not inhibit type I IFN signaling in bystander cells. (A) Schematic of transwell experiments. (B) C57BL/6 BMDMs were seeded into the upper transwell, with no cells in the lower transwell. Upper transwells were then infected with M. smegmatis at an MOI of 3 for 4 h or treated with 1.5 ng of IFN-β. Twenty hours poststimulation, supernatants were harvested from the upper and lower transwells, and IFN-β protein levels were determined by ELISA. (C) Upper transwells were infected or not with M. tuberculosis at an MOI of 3 for 4 h, and then the upper and lower transwells were treated with 200 pg/ml IFN-β for 4 h. Cells in the lower transwell were lysed by addition of Triton X-100, and the amount of luciferase was quantified. (D) Whole-cell lysates from the upper transwell of Ifn-β−/− BMDMs were collected at 4 hpi and immunoblotted for p-STAT1, total STAT1, or β-actin as indicated. (E) Whole-cell lysates from the lower transwell were collected at 4 hpi and immunoblotted for p-STAT1, total STAT1, or β-actin as indicated. UI or M. tuberculosis refers to the infection condition of the upper transwell. Densitometric ratios are normalized to the loading control and are shown relative to the UI plus IFN-β condition.

Close modal
FIGURE 7.

Overview of type I IFN signaling and production in M. tuberculosis–infected cells. M. tuberculosis induces the production of IFN-β in infected cells via release of bacterial DNA (17, 1922) or damage to host cell mitochondria followed by increased mtDNA in the cytosol (24). All these factors initiate the STING/TBK1 signaling pathway that leads to activation of the transcription factor IRF3 and increased IFN-β production. The secreted IFN-β acts on bystander cells to increase secretion of IL-10 and IL-1Ra, which leads to increased host cell necrosis and tissue damage, thus exacerbating the disease outcome (8). In this study, we show that M. tuberculosis inhibits autocrine IFNAR signaling, which limits not only the production of IFN-β but also the expression of genes with host cell defense properties such as Nos2.

FIGURE 7.

Overview of type I IFN signaling and production in M. tuberculosis–infected cells. M. tuberculosis induces the production of IFN-β in infected cells via release of bacterial DNA (17, 1922) or damage to host cell mitochondria followed by increased mtDNA in the cytosol (24). All these factors initiate the STING/TBK1 signaling pathway that leads to activation of the transcription factor IRF3 and increased IFN-β production. The secreted IFN-β acts on bystander cells to increase secretion of IL-10 and IL-1Ra, which leads to increased host cell necrosis and tissue damage, thus exacerbating the disease outcome (8). In this study, we show that M. tuberculosis inhibits autocrine IFNAR signaling, which limits not only the production of IFN-β but also the expression of genes with host cell defense properties such as Nos2.

Close modal

Although type I IFNs are largely considered to be beneficial in the context of viral infections, their role during bacterial infections is not completely understood and may vary depending on the bacterial pathogen and the site of infection. In the context of M. tuberculosis infections, type I IFNs are considered to be detrimental to the host, and numerous recent studies have worked toward better understanding why. Surprisingly, we discovered an antimicrobial effect of type I IFNs during M. tuberculosis infection in macrophages via the production of NO. The role of NO in host resistance to tuberculosis has been extensively investigated, and its bactericidal activity is attributed to the reactive nitrogen intermediates (RNI) such as NO2, NO3, and peroxynitrite (ONOO) (43, 44). Activation of macrophages with IFN-γ was especially powerful in augmenting RNI-mediated killing of M. tuberculosis (4547). Two recent studies demonstrate that the major host-protective role of NO during an in vivo infection with M. tuberculosis may not be its bactericidal activity but its immunosuppressive activity, leading to reduced host tissue pathology (48, 49). NO is generated in the cell cytosol by NOS2, and it diffuses rapidly and freely at an estimated 5–10 cell lengths/s (50). This means that during an in vivo infection, dilution is an important factor, as within 1 s the NO concentration has been diluted over 200 times in the NO-generating cell (50). In addition, the proteasome of M. tuberculosis has a role in resistance of the bacteria to RNI stress (51). It is, thus, possible that NO levels in the infected cells that actually generate the NO fail to accumulate to bactericidal threshold levels in vivo because of diffusion and dilution as compared with ex vivo infection experiments, which are in a closed system and contain mostly infected cells. In any case, the potential of M. tuberculosis to inhibit IFN-β–mediated NO production will be advantageous for the pathogen. We do not believe that the recently reported direct bactericidal effect of IFN-β (52) plays a role in our observed bactericidal effects because otherwise the Nos2−/− cells should not be different from WT BMDMs (Fig. 1G).

It seems confounding that M. tuberculosis would want to both induce IFN-β production and simultaneously inhibit its signaling. Considering that M. tuberculosis is a facultative intracellular pathogen, we hypothesize that inhibiting type I IFN signaling allows M. tuberculosis to mitigate the antibacterial effects of IFN-β–induced autocrine signaling (Fig. 1). The transcriptome analysis of IFN-β–regulated genes and their deregulation by M. tuberculosis infection suggests that there are some type I IFN responses that are detrimental to the bacterium during infection. CCL12/MCP-5 is one of the IFN-β–regulated cytokines that is strongly inhibited by M. tuberculosis infection at the mRNA and protein level (Fig. 3B, 3E). It is a chemoattractant for monocytes and an agonist of CCR2 (53). Its expression on macrophages is induced by LPS and IFN-γ (53). Interestingly, another chemoattractant for monocytes, CCL5, is also upregulated via IFN-β signaling, and its expression is inhibited by M. tuberculosis (Supplemental Fig. 3). Consequently, the repression of these chemokines could lead to a reduction of monocyte recruitment within the M. tuberculosis–infected lungs. The type I IFN–driven expression of chemokines has been shown to be an important signal for recruitment of bone marrow monocytes to the site of infection with Listeria monocytogenes (54). Among other top downregulated genes for which we also have confirmed reduced protein levels are the immunity-related GTPases Ifi204, Ifit1, and Iigp1 (Fig. 3) (55). IIGP1 is involved in host defense against Chlamydia trachomatis and Toxoplasma gondii (56, 57). It is unlikely that this protein, at least in vivo, is involved in host defense against M. tuberculosis because Iigp1−/− mice do not have a phenotype (58). IFI204 is a cytosolic DNA sensor that binds to eDNA and induces the cytosolic surveillance pathway via STING/TBK1/IRF3 signaling (17). Consequently, reduction of IFI204 expression by M. tuberculosis will reduce the activation of IRF3 and its regulon, which includes Ifn-β transcription. This could, thus, represent an additional mechanism, in addition to the expression of the phosphodiesterase CdnP (23), by which M. tuberculosis regulates IFN-β production. Interestingly, M. bovis does not inhibit IFI204 expression, and, hence, this host cell protein has an important role in host IFN-β production for this mycobacterial species (59). It is important to highlight, however, that the main cytosolic DNA sensor involved in recognition of M. tuberculosis DNA is cGAS (1921). IFIT1 has well established antiviral activity and is most strongly induced by IFN-β (60). Its activity is dependent upon selective binding of 5′-terminal regions of cap0-, cap1-, and 5′ppp-mRNAs (61), and, hence, IFIT1 is an unlikely candidate for functioning in host cell resistance against M. tuberculosis infection. The expression of the Nos2 gene is not affected by M. tuberculosis, which we think is due to the early time point (4 hpi) selected for the RNAseq analysis. Nos2 can be induced by signaling via many TLRs and cytokine receptors (43, 62). In our system, M. tuberculosis infection causes a strong induction of TNF secretion, which is amplified by addition of IFN-β (Supplemental Fig. 3). Consequently, we believe that TNF is the most likely cause for the observed late induction of NOS2, especially because it has been noted before that IFN-β and TNF synergistically mediate the induction of Nos2 gene transcription (43, 6264).

The capacity of M. tuberculosis to induce activation of STAT3 was recently reported (65), and the importance of STAT3 activation for the virulence of M. tuberculosis was recently demonstrated by showing that the deletion of STAT3 in myeloid cells increased resistance of mice to M. tuberculosis infection (66). STAT3 phosphorylation can inhibit type I IFN–mediated signaling, although this does not seem to occur at the level of STAT1/STAT2 phosphorylation but rather at the level of nuclear translocation (39, 40). Thus, STAT3 activation is likely not involved in our observed inhibition of TYK2/JAK1 phosphorylation by M. tuberculosis. However, we cannot exclude the possibility that M. tuberculosis exerts multiple strategies that synergize to prevent transcription of IFN-β–regulated genes and that STAT3-mediated inhibition may play a role during later time points to sustain the initial signaling inhibition. The capacity of STAT3 to inhibit Nos2 gene expression may explain why NOS2 is only detected after 72–96 h (Fig. 1E). SOCS1, SOCS3, and USP18 are negative regulators of the IFNAR signaling that are typically induced at a later time point during IFN-β stimulation and serve as negative feedback regulators (3). Considering that our observed inhibition occurs already as early as 5 min postinfection, albeit an infection period of 4 h, it is highly unlikely that the molecular mechanism of at least the early inhibition is dependent on these common negative regulators. It is known that M. tuberculosis possesses several phosphatases that have been shown to affect the host immune response by interfering with several signal transduction pathways. SapM is a phosphoinositide phosphatase that is essential in arresting phagosomal maturation by inhibiting phosphatidylinositol 3-phosphate phosphorylation (67). The tyrosine phosphatase PtpA is also involved in inhibiting phagosomal maturation through inhibition of V-ATPase, and PtpB has been shown to inhibit ERK 1/2 and p38 signaling cascades (6769). Although beyond the scope of this study, it would be interesting to investigate whether these proteins also regulate IFN-β–mediated signaling by directly dephosphorylating TYK2 or JAK1.

In this study, we show that M. tuberculosis is susceptible to host cell IFNAR signaling and has evolved to suppress it. In particular, M. tuberculosis inhibits IFNAR-mediated signaling at the level of the receptor-associated tyrosine kinases JAK1 and TYK2 (Fig. 4). Nevertheless, this inhibition can be overcome by high concentrations of extracellular IFN-β, leading to reduced viability of intracellular M. tuberculosis. It is well established that M. tuberculosis induces the STING/TBK1/IRF3 signaling axis via eDNA (17, 1921), damage to host cell mitochondria leading to an increase in cytosolic mtDNA (24) or secretion of c-di-AMP (22, 23) (Fig. 7). The capacity to produce IFN-β is associated with in vivo virulence of M. tuberculosis infections in mouse models and human studies (7, 12, 13, 15, 16). The levels of IFN-β production by mycobacteria-infected cells vary depending on the mycobacterial species and, in the context of M. tuberculosis, the specific strain that is used for the infection (24, 41, 42) (Fig. 1A, 1B). High levels of IFN-β drive production of IL-10 and IL-1Ra (Fig. 7), which antagonize the host-protective activity of IL-1β and ultimately lead to increased host tissue destruction, which establishes a replicative niche for M. tuberculosis (8). In contrast to the current dogma that M. tuberculosis induces production of type I IFNs to support its virulence, multiple studies have provided evidence that, in some settings, type I IFNs may have detrimental effects on M. tuberculosis. For example, Isg15 is one of the most highly upregulated genes after type I IFN stimulation of cells, and it has a host-protective role during M. tuberculosis infection (70). Furthermore, in the absence of IFN-γ, type I IFNs can promote the activation of macrophages for improved innate host response to M. tuberculosis (34). The strongest evidence for a host-protective element in the IFN-β response was produced by showing that Ifngr−/−Ifnar−/− double-knockout mice are more susceptible compared with Ifn-γ−/− mice (6, 34, 71). Additionally, macrophages derived from cGAS−/− and STING−/− knockout mice produced less IFN-β mRNA and had an increased bacterial burden compared with WT BMDMs (21). Nevertheless, in vivo there were no differences in bacterial burden (21). Importantly, cGAS−/− mice succumbed earlier to M. tuberculosis infection than WT mice during survival studies, and although most inflammatory cytokines were produced similarly, there was a 2-fold decrease in IFN-β serum levels compared with knockout mice; however, the absence of phenotype in STING−/− mice is confounding (21). IFN-β is protective during mouse infections against two nontuberculous mycobacterial species (M. smegmatis and M. avium ssp. paratuberculosis) (72). In this context, it is less surprising that M. tuberculosis has also evolved mechanisms to limit production of type I IFNs, possibly to achieve the “Goldilocks principle.” Overall, M. tuberculosis infection causes less production of type I IFNs in ex vivo–infected macrophages compared with nontuberculous mycobacteria (41) (Fig. 1A, 1B). Indeed, we have shown previously that M. tuberculosis can inhibit M. smegmatis–induced IFN-β production in an ESX-1–dependent manner in bone marrow–derived dendritic cells (41). There are potentially several mechanisms by which this inhibition occurs. The M. tuberculosis–mediated stimulation of TLR2 inhibits the induction of type I IFNs via TLR7/9 activation (73). In addition, the M. tuberculosis phosphodiesterase CdnP can reduce type I IFN production by hydrolyzing bacteria-derived c-di-AMP and host-derived cyclic GMP–AMP, thereby limiting activation of the STING pathway (23). Importantly, this inhibition is relevant for full virulence of M. tuberculosis because a CdnP transposon mutant of M. tuberculosis is attenuated in mice (23).

We propose that M. tuberculosis has evolved to inhibit autocrine IFN-β signaling and its host-protective effects to still take advantage of the benefits of paracrine IFN-β signaling on UI bystander cells, which is not inhibited by the M. tuberculosis–infected cells (Fig. 7). This model might explain why, in some settings, IFN-β may be beneficial for the host (6); for example, in the case of nontuberculous mycobacteria, which cannot inhibit IFNAR-mediated signaling (e.g., M. smegmatis) and are susceptible to a host type I IFN response (72).

We thank the University of Maryland Genomics Core facility for excellent technical assistance and Dr. Sophie Helaine (Imperial College London, London, U.K.) for advice and editing of the manuscript.

This work was supported by National Institutes of Health Grants R01AI139492 and R21AI107377 (to D.A.B., S.E.A., and V.B.), R01AI094773 (to N.M.E.-S. and V.K.H.), and R01AI125215 (to S.N.V.).

The sequences presented in this article have been submitted to the Sequence Read Archive under accession number SRP130272.

The online version of this article contains supplemental material.

Abbreviations used in this article:

BMDM

bone marrow–derived macrophage

c-di-AMP

cyclic-di-AMP

cGAS

cyclic GMP–AMP synthase

eDNA

extracellular M. tuberculosis DNA

hpi

hour postinfection

IFNAR

IFN-α/β–receptor

ISG

IFN-stimulated gene

ISGF3

ISG factor 3

MOI

multiplicity of infection

mtDNA

mitochondrial DNA

NIH

National Institutes of Health

Nos2

NO synthase 2

RNAseq

RNA sequencing

RNI

reactive nitrogen intermediate

UI

uninfected

WT

wild-type.

1
Diamond
,
M. S.
,
M.
Farzan
.
2013
.
The broad-spectrum antiviral functions of IFIT and IFITM proteins.
Nat. Rev. Immunol.
13
:
46
57
.
2
McNab
,
F.
,
K.
Mayer-Barber
,
A.
Sher
,
A.
Wack
,
A.
O’Garra
.
2015
.
Type I interferons in infectious disease.
Nat. Rev. Immunol.
15
:
87
103
.
3
Ivashkiv
,
L. B.
,
L. T.
Donlin
.
2014
.
Regulation of type I interferon responses.
Nat. Rev. Immunol.
14
:
36
49
.
4
Boxx
,
G. M.
,
G.
Cheng
.
2016
.
The roles of type I interferon in bacterial infection.
Cell Host Microbe
19
:
760
769
.
5
Kovarik
,
P.
,
V.
Castiglia
,
M.
Ivin
,
F.
Ebner
.
2016
.
Type I interferons in bacterial infections: a balancing act.
Front. Immunol.
7
:
652
.
6
Moreira-Teixeira
,
L.
,
K.
Mayer-Barber
,
A.
Sher
,
A.
O’Garra
.
2018
.
Type I interferons in tuberculosis: foe and occasionally friend.
J. Exp. Med.
215
:
1273
1285
.
7
Dorhoi
,
A.
,
V.
Yeremeev
,
G.
Nouailles
,
J.
Weiner
III
,
S.
Jörg
,
E.
Heinemann
,
D.
Oberbeck-Müller
,
J. K.
Knaul
,
A.
Vogelzang
,
S. T.
Reece
, et al
.
2014
.
Type I IFN signaling triggers immunopathology in tuberculosis-susceptible mice by modulating lung phagocyte dynamics.
Eur. J. Immunol.
44
:
2380
2393
.
8
Mayer-Barber
,
K. D.
,
B. B.
Andrade
,
S. D.
Oland
,
E. P.
Amaral
,
D. L.
Barber
,
J.
Gonzales
,
S. C.
Derrick
,
R.
Shi
,
N. P.
Kumar
,
W.
Wei
, et al
.
2014
.
Host-directed therapy of tuberculosis based on interleukin-1 and type I interferon crosstalk.
Nature
511
:
99
103
.
9
Mayer-Barber
,
K. D.
,
B. B.
Andrade
,
D. L.
Barber
,
S.
Hieny
,
C. G.
Feng
,
P.
Caspar
,
S.
Oland
,
S.
Gordon
,
A.
Sher
.
2011
.
Innate and adaptive interferons suppress IL-1α and IL-1β production by distinct pulmonary myeloid subsets during Mycobacterium tuberculosis infection.
Immunity
35
:
1023
1034
.
10
Novikov
,
A.
,
M.
Cardone
,
R.
Thompson
,
K.
Shenderov
,
K. D.
Kirschman
,
K. D.
Mayer-Barber
,
T. G.
Myers
,
R. L.
Rabin
,
G.
Trinchieri
,
A.
Sher
,
C. G.
Feng
.
2011
.
Mycobacterium tuberculosis triggers host type I IFN signaling to regulate IL-1β production in human macrophages.
J. Immunol.
187
:
2540
2547
.
11
Maertzdorf
,
J.
,
D.
Repsilber
,
S. K.
Parida
,
K.
Stanley
,
T.
Roberts
,
G.
Black
,
G.
Walzl
,
S. H. E.
Kaufmann
.
2011
.
Human gene expression profiles of susceptibility and resistance in tuberculosis.
Genes Immun.
12
:
15
22
.
12
Berry
,
M. P. R.
,
C. M.
Graham
,
F. W.
McNab
,
Z.
Xu
,
S. A. A.
Bloch
,
T.
Oni
,
K. A.
Wilkinson
,
R.
Banchereau
,
J.
Skinner
,
R. J.
Wilkinson
, et al
.
2010
.
An interferon-inducible neutrophil-driven blood transcriptional signature in human tuberculosis.
Nature
466
:
973
977
.
13
Antonelli
,
L. R. V.
,
A.
Gigliotti Rothfuchs
,
R.
Gonçalves
,
E.
Roffê
,
A. W.
Cheever
,
A.
Bafica
,
A. M.
Salazar
,
C. G.
Feng
,
A.
Sher
.
2010
.
Intranasal Poly-IC treatment exacerbates tuberculosis in mice through the pulmonary recruitment of a pathogen-permissive monocyte/macrophage population.
J. Clin. Invest.
120
:
1674
1682
.
14
Ordway
,
D.
,
M.
Henao-Tamayo
,
M.
Harton
,
G.
Palanisamy
,
J.
Troudt
,
C.
Shanley
,
R. J.
Basaraba
,
I. M.
Orme
.
2007
.
The hypervirulent Mycobacterium tuberculosis strain HN878 induces a potent TH1 response followed by rapid down-regulation.
J. Immunol.
179
:
522
531
.
15
Stanley
,
S. A.
,
J. E.
Johndrow
,
P.
Manzanillo
,
J. S.
Cox
.
2007
.
The Type I IFN response to infection with Mycobacterium tuberculosis requires ESX-1-mediated secretion and contributes to pathogenesis.
J. Immunol.
178
:
3143
3152
.
16
Manca
,
C.
,
L.
Tsenova
,
A.
Bergtold
,
S.
Freeman
,
M.
Tovey
,
J. M.
Musser
,
C. E.
Barry
III
,
V. H.
Freedman
,
G.
Kaplan
.
2001
.
Virulence of a Mycobacterium tuberculosis clinical isolate in mice is determined by failure to induce Th1 type immunity and is associated with induction of IFN-alpha/beta.
Proc. Natl. Acad. Sci. USA
98
:
5752
5757
.
17
Manzanillo
,
P. S.
,
M. U.
Shiloh
,
D. A.
Portnoy
,
J. S.
Cox
.
2012
.
Mycobacterium tuberculosis activates the DNA-dependent cytosolic surveillance pathway within macrophages.
Cell Host Microbe
11
:
469
480
.
18
Watson
,
R. O.
,
P. S.
Manzanillo
,
J. S.
Cox
.
2012
.
Extracellular M. tuberculosis DNA targets bacteria for autophagy by activating the host DNA-sensing pathway.
Cell
150
:
803
815
.
19
Wassermann
,
R.
,
M. F.
Gulen
,
C.
Sala
,
S. G.
Perin
,
Y.
Lou
,
J.
Rybniker
,
J. L.
Schmid-Burgk
,
T.
Schmidt
,
V.
Hornung
,
S. T.
Cole
,
A.
Ablasser
.
2015
.
Mycobacterium tuberculosis differentially activates cGAS- and inflammasome-dependent intracellular immune responses through ESX-1.
Cell Host Microbe
17
:
799
810
.
20
Watson
,
R. O.
,
S. L.
Bell
,
D. A.
MacDuff
,
J. M.
Kimmey
,
E. J.
Diner
,
J.
Olivas
,
R. E.
Vance
,
C. L.
Stallings
,
H. W.
Virgin
,
J. S.
Cox
.
2015
.
The cytosolic sensor cGAS detects Mycobacterium tuberculosis DNA to induce type I interferons and activate autophagy.
Cell Host Microbe
17
:
811
819
.
21
Collins
,
A. C.
,
H.
Cai
,
T.
Li
,
L. H.
Franco
,
X.-D.
Li
,
V. R.
Nair
,
C. R.
Scharn
,
C. E.
Stamm
,
B.
Levine
,
Z. J.
Chen
,
M. U.
Shiloh
.
2015
.
Cyclic GMP-AMP synthase is an innate immune DNA sensor for Mycobacterium tuberculosis.
Cell Host Microbe
17
:
820
828
.
22
Dey
,
B.
,
R. J.
Dey
,
L. S.
Cheung
,
S.
Pokkali
,
H.
Guo
,
J.-H.
Lee
,
W. R.
Bishai
.
2015
.
A bacterial cyclic dinucleotide activates the cytosolic surveillance pathway and mediates innate resistance to tuberculosis.
Nat. Med.
21
:
401
406
.
23
Dey
,
R. J.
,
B.
Dey
,
Y.
Zheng
,
L. S.
Cheung
,
J.
Zhou
,
D.
Sayre
,
P.
Kumar
,
H.
Guo
,
G.
Lamichhane
,
H. O.
Sintim
,
W. R.
Bishai
.
2017
.
Inhibition of innate immune cytosolic surveillance by an M. tuberculosis phosphodiesterase.
Nat. Chem. Biol.
13
:
210
217
.
24
Wiens
,
K. E.
,
J. D.
Ernst
.
2016
.
The mechanism for type I interferon induction by Mycobacterium tuberculosis is bacterial strain-dependent.
PLoS Pathog.
12
:
e1005809
.
25
Deonarain
,
R.
,
A.
Alcamí
,
M.
Alexiou
,
M. J.
Dallman
,
D. R.
Gewert
,
A. C. G.
Porter
.
2000
.
Impaired antiviral response and alpha/beta interferon induction in mice lacking beta interferon.
J. Virol.
74
:
3404
3409
.
26
Srinivasan
,
L.
,
S. A.
Gurses
,
B. E.
Hurley
,
J. L.
Miller
,
P. C.
Karakousis
,
V.
Briken
.
2016
.
Identification of a transcription factor that regulates host cell exit and virulence of Mycobacterium tuberculosis.
PLoS Pathog.
12
:
e1005652
.
27
Bolger
,
A. M.
,
M.
Lohse
,
B.
Usadel
.
2014
.
Trimmomatic: a flexible trimmer for Illumina sequence data.
Bioinformatics
30
:
2114
2120
.
28
Yates
,
A.
,
W.
Akanni
,
M. R.
Amode
,
D.
Barrell
,
K.
Billis
,
D.
Carvalho-Silva
,
C.
Cummins
,
P.
Clapham
,
S.
Fitzgerald
,
L.
Gil
, et al
.
2016
.
Ensembl 2016.
Nucleic Acids Res.
44
(
D1
):
D710
D716
.
29
Kim
,
D.
,
G.
Pertea
,
C.
Trapnell
,
H.
Pimentel
,
R.
Kelley
,
S. L.
Salzberg
.
2013
.
TopHat2: accurate alignment of transcriptomes in the presence of insertions, deletions and gene fusions.
Genome Biol.
14
:
R36
.
30
Anders
,
S.
,
P. T.
Pyl
,
W.
Huber
.
2015
.
HTSeq--a Python framework to work with high-throughput sequencing data.
Bioinformatics
31
:
166
169
.
31
Gentleman
,
R. C.
,
V. J.
Carey
,
D. M.
Bates
,
B.
Bolstad
,
M.
Dettling
,
S.
Dudoit
,
B.
Ellis
,
L.
Gautier
,
Y.
Ge
,
J.
Gentry
, et al
.
2004
.
Bioconductor: open software development for computational biology and bioinformatics.
Genome Biol.
5
:
R80
.
32
Law
,
C. W.
,
Y.
Chen
,
W.
Shi
,
G. K.
Smyth
.
2014
.
voom: precision weights unlock linear model analysis tools for RNA-seq read counts.
Genome Biol.
15
:
R29
.
33
Leek
,
J. T.
,
W. E.
Johnson
,
H. S.
Parker
,
A. E.
Jaffe
,
J. D.
Storey
.
2012
.
The sva package for removing batch effects and other unwanted variation in high-throughput experiments.
Bioinformatics
28
:
882
883
.
34
Moreira-Teixeira
,
L.
,
J.
Sousa
,
F. W.
McNab
,
E.
Torrado
,
F.
Cardoso
,
H.
Machado
,
F.
Castro
,
V.
Cardoso
,
J.
Gaifem
,
X.
Wu
, et al
.
2016
.
Type I IFN inhibits alternative macrophage activation during Mycobacterium tuberculosis infection and leads to enhanced protection in the absence of IFN-γ signaling.
J. Immunol.
197
:
4714
4726
.
35
Evans
,
J. D.
,
R. A.
Crown
,
J. A.
Sohn
,
C.
Seeger
.
2011
.
West Nile virus infection induces depletion of IFNAR1 protein levels.
Viral Immunol.
24
:
253
263
.
36
Xia
,
C.
,
M.
Vijayan
,
C. J.
Pritzl
,
S. Y.
Fuchs
,
A. B.
McDermott
,
B.
Hahm
.
2015
.
Hemagglutinin of influenza a virus antagonizes type I interferon (IFN) responses by inducing degradation of type I IFN receptor 1.
J. Virol.
90
:
2403
2417
.
37
Ting
,
L. M.
,
A. C.
Kim
,
A.
Cattamanchi
,
J. D.
Ernst
.
1999
.
Mycobacterium tuberculosis inhibits IFN-gamma transcriptional responses without inhibiting activation of STAT1.
J. Immunol.
163
:
3898
3906
.
38
Villarino
,
A. V.
,
Y.
Kanno
,
J. J.
O’Shea
.
2017
.
Mechanisms and consequences of Jak-STAT signaling in the immune system.
Nat. Immunol.
18
:
374
384
.
39
Ho
,
H. H.
,
L. B.
Ivashkiv
.
2006
.
Role of STAT3 in type I interferon responses. Negative regulation of STAT1-dependent inflammatory gene activation.
J. Biol. Chem.
281
:
14111
14118
.
40
Wang
,
W.-B.
,
D. E.
Levy
,
C.-K.
Lee
.
2011
.
STAT3 negatively regulates type I IFN-mediated antiviral response. [Published erratum appears in 2011 J. Immunol. 187: 6583.]
J. Immunol.
187
:
2578
2585
.
41
Shah
,
S.
,
A.
Bohsali
,
S. E.
Ahlbrand
,
L.
Srinivasan
,
V. A. K.
Rathinam
,
S. N.
Vogel
,
K. A.
Fitzgerald
,
F. S.
Sutterwala
,
V.
Briken
.
2013
.
Cutting edge: Mycobacterium tuberculosis but not nonvirulent mycobacteria inhibits IFN-β and AIM2 inflammasome-dependent IL-1β production via its ESX-1 secretion system.
J. Immunol.
191
:
3514
3518
.
42
Manca
,
C.
,
L.
Tsenova
,
S.
Freeman
,
A. K.
Barczak
,
M.
Tovey
,
P. J.
Murray
,
C.
Barry
III
,
G.
Kaplan
.
2005
.
Hypervirulent M. tuberculosis W/Beijing strains upregulate type I IFNs and increase expression of negative regulators of the Jak-Stat pathway.
J. Interferon Cytokine Res.
25
:
694
701
.
43
MacMicking
,
J.
,
Q. W.
Xie
,
C.
Nathan
.
1997
.
Nitric oxide and macrophage function.
Annu. Rev. Immunol.
15
:
323
350
.
44
Jamaati
,
H.
,
E.
Mortaz
,
Z.
Pajouhi
,
G.
Folkerts
,
M.
Movassaghi
,
M.
Moloudizargari
,
I. M.
Adcock
,
J.
Garssen
.
2017
.
Nitric oxide in the pathogenesis and treatment of tuberculosis.
Front. Microbiol.
8
:
2008
.
45
Flesch
,
I. E.
,
S. H.
Kaufmann
.
1991
.
Mechanisms involved in mycobacterial growth inhibition by gamma interferon-activated bone marrow macrophages: role of reactive nitrogen intermediates.
Infect. Immun.
59
:
3213
3218
.
46
Denis
,
M.
1991
.
Interferon-gamma-treated murine macrophages inhibit growth of tubercle bacilli via the generation of reactive nitrogen intermediates.
Cell. Immunol.
132
:
150
157
.
47
Chan
,
J.
,
Y.
Xing
,
R. S.
Magliozzo
,
B. R.
Bloom
.
1992
.
Killing of virulent Mycobacterium tuberculosis by reactive nitrogen intermediates produced by activated murine macrophages.
J. Exp. Med.
175
:
1111
1122
.
48
Mishra
,
B. B.
,
R. R.
Lovewell
,
A. J.
Olive
,
G.
Zhang
,
W.
Wang
,
E.
Eugenin
,
C. M.
Smith
,
J. Y.
Phuah
,
J. E.
Long
,
M. L.
Dubuke
, et al
.
2017
.
Nitric oxide prevents a pathogen-permissive granulocytic inflammation during tuberculosis.
Nat. Microbiol.
2
:
17072
.
49
Mishra
,
B. B.
,
V. A. K.
Rathinam
,
G. W.
Martens
,
A. J.
Martinot
,
H.
Kornfeld
,
K. A.
Fitzgerald
,
C. M.
Sassetti
.
2013
.
Nitric oxide controls the immunopathology of tuberculosis by inhibiting NLRP3 inflammasome-dependent processing of IL-1β.
Nat. Immunol.
14
:
52
60
.
50
Thomas
,
D. D.
,
L. A.
Ridnour
,
J. S.
Isenberg
,
W.
Flores-Santana
,
C. H.
Switzer
,
S.
Donzelli
,
P.
Hussain
,
C.
Vecoli
,
N.
Paolocci
,
S.
Ambs
, et al
.
2008
.
The chemical biology of nitric oxide: implications in cellular signaling.
Free Radic. Biol. Med.
45
:
18
31
.
51
Darwin
,
K. H.
,
S.
Ehrt
,
J.-C.
Gutierrez-Ramos
,
N.
Weich
,
C. F.
Nathan
.
2003
.
The proteasome of Mycobacterium tuberculosis is required for resistance to nitric oxide.
Science
302
:
1963
1966
.
52
Kaplan
,
A.
,
M. W.
Lee
,
A. J.
Wolf
,
J. J.
Limon
,
C. A.
Becker
,
M.
Ding
,
R.
Murali
,
E. Y.
Lee
,
G. Y.
Liu
,
G. C. L.
Wong
,
D. M.
Underhill
.
2017
.
Direct antimicrobial activity of IFN-β.
J. Immunol.
198
:
4036
4045
.
53
Sarafi
,
M. N.
,
E. A.
Garcia-Zepeda
,
J. A.
MacLean
,
I. F.
Charo
,
A. D.
Luster
.
1997
.
Murine monocyte chemoattractant protein (MCP)-5: a novel CC chemokine that is a structural and functional homologue of human MCP-1.
J. Exp. Med.
185
:
99
109
.
54
Jia
,
T.
,
I.
Leiner
,
G.
Dorothee
,
K.
Brandl
,
E. G.
Pamer
.
2009
.
MyD88 and Type I interferon receptor-mediated chemokine induction and monocyte recruitment during Listeria monocytogenes infection.
J. Immunol.
183
:
1271
1278
.
55
Kim
,
B.-H.
,
J. D.
Chee
,
C. J.
Bradfield
,
E.-S.
Park
,
P.
Kumar
,
J. D.
MacMicking
.
2016
.
Interferon-induced guanylate-binding proteins in inflammasome activation and host defense.
Nat. Immunol.
17
:
481
489
.
56
Martens
,
S.
,
I.
Parvanova
,
J.
Zerrahn
,
G.
Griffiths
,
G.
Schell
,
G.
Reichmann
,
J. C.
Howard
.
2005
.
Disruption of Toxoplasma gondii parasitophorous vacuoles by the mouse p47-resistance GTPases.
PLoS Pathog.
1
:
e24
.
57
Al-Zeer
,
M. A.
,
H. M.
Al-Younes
,
P. R.
Braun
,
J.
Zerrahn
,
T. F.
Meyer
.
2009
.
IFN-γ-inducible Irga6 mediates host resistance against Chlamydia trachomatis via autophagy.
PLoS One
4
:
e4588
.
58
Liesenfeld
,
O.
,
I.
Parvanova
,
J.
Zerrahn
,
S.-J.
Han
,
F.
Heinrich
,
M.
Muñoz
,
F.
Kaiser
,
T.
Aebischer
,
T.
Buch
,
A.
Waisman
, et al
.
2011
.
The IFN-γ-inducible GTPase, Irga6, protects mice against Toxoplasma gondii but not against Plasmodium berghei and some other intracellular pathogens.
PLoS One
6
:
e20568
.
59
Chunfa
,
L.
,
S.
Xin
,
L.
Qiang
,
S.
Sreevatsan
,
L.
Yang
,
D.
Zhao
,
X.
Zhou
.
2017
.
The central role of IFI204 in IFN-β release and autophagy activation during Mycobacterium bovis infection.
Front. Cell. Infect. Microbiol.
7
:
169
.
60
Fensterl
,
V.
,
G. C.
Sen
.
2015
.
Interferon-induced Ifit proteins: their role in viral pathogenesis.
J. Virol.
89
:
2462
2468
.
61
Kumar
,
P.
,
T. R.
Sweeney
,
M. A.
Skabkin
,
O. V.
Skabkina
,
C. U. T.
Hellen
,
T. V.
Pestova
.
2014
.
Inhibition of translation by IFIT family members is determined by their ability to interact selectively with the 5′-terminal regions of cap0-, cap1- and 5'ppp- mRNAs.
Nucleic Acids Res.
42
:
3228
3245
.
62
Bogdan
,
C.
2001
.
Nitric oxide and the immune response.
Nat. Immunol.
2
:
907
916
.
63
Farlik
,
M.
,
B.
Reutterer
,
C.
Schindler
,
F.
Greten
,
C.
Vogl
,
M.
Müller
,
T.
Decker
.
2010
.
Nonconventional initiation complex assembly by STAT and NF-kappaB transcription factors regulates nitric oxide synthase expression.
Immunity
33
:
25
34
.
64
Bachmann
,
M.
,
Z.
Waibler
,
T.
Pleli
,
J.
Pfeilschifter
,
H.
Mühl
.
2017
.
Type I interferon supports inducible nitric oxide synthase in murine hepatoma cells and hepatocytes and during experimental acetaminophen-induced liver damage.
Front. Immunol.
8
:
890
.
65
Queval
,
C. J.
,
O.-R.
Song
,
N.
Deboosère
,
V.
Delorme
,
A.-S.
Debrie
,
R.
Iantomasi
,
R.
Veyron-Churlet
,
S.
Jouny
,
K.
Redhage
,
G.
Deloison
, et al
.
2016
.
STAT3 represses nitric oxide synthesis in human macrophages upon Mycobacterium tuberculosis infection.
Sci. Rep.
6
:
29297
.
66
Gao
,
Y.
,
J. I.
Basile
,
C.
Classon
,
D.
Gavier-Widen
,
A.
Yoshimura
,
B.
Carow
,
M. E.
Rottenberg
.
2018
.
STAT3 expression by myeloid cells is detrimental for the T- cell-mediated control of infection with Mycobacterium tuberculosis.
PLoS Pathog.
14
:
e1006809
.
67
Wong
,
D.
,
J. D.
Chao
,
Y.
Av-Gay
.
2013
.
Mycobacterium tuberculosis-secreted phosphatases: from pathogenesis to targets for TB drug development.
Trends Microbiol.
21
:
100
109
.
68
Zhou
,
B.
,
Y.
He
,
X.
Zhang
,
J.
Xu
,
Y.
Luo
,
Y.
Wang
,
S. G.
Franzblau
,
Z.
Yang
,
R. J.
Chan
,
Y.
Liu
, et al
.
2010
.
Targeting mycobacterium protein tyrosine phosphatase B for antituberculosis agents.
Proc. Natl. Acad. Sci. USA
107
:
4573
4578
.
69
Wong
,
D.
,
H.
Bach
,
J.
Sun
,
Z.
Hmama
,
Y.
Av-Gay
.
2011
.
Mycobacterium tuberculosis protein tyrosine phosphatase (PtpA) excludes host vacuolar-H+-ATPase to inhibit phagosome acidification.
Proc. Natl. Acad. Sci. USA
108
:
19371
19376
.
70
Kimmey
,
J. M.
,
J. A.
Campbell
,
L. A.
Weiss
,
K. J.
Monte
,
D. J.
Lenschow
,
C. L.
Stallings
.
2017
.
The impact of ISGylation during Mycobacterium tuberculosis infection in mice.
Microbes Infect.
19
:
249
258
.
71
Desvignes
,
L.
,
A. J.
Wolf
,
J. D.
Ernst
.
2012
.
Dynamic roles of type I and type II IFNs in early infection with Mycobacterium tuberculosis.
J. Immunol.
188
:
6205
6215
.
72
Ruangkiattikul
,
N.
,
A.
Nerlich
,
K.
Abdissa
,
S.
Lienenklaus
,
A.
Suwandi
,
N.
Janze
,
K.
Laarmann
,
J.
Spanier
,
U.
Kalinke
,
S.
Weiss
,
R.
Goethe
.
2017
.
cGAS-STING-TBK1-IRF3/7 induced interferon-β contributes to the clearing of non tuberculous mycobacterial infection in mice.
Virulence
8
:
1303
1315
.
73
Liu
,
Y. C.
,
D. P.
Simmons
,
X.
Li
,
D. W.
Abbott
,
W. H.
Boom
,
C. V.
Harding
.
2012
.
TLR2 signaling depletes IRAK1 and inhibits induction of type I IFN by TLR7/9.
J. Immunol.
188
:
1019
1026
.

The authors have no financial conflicts of interest.

Supplementary data