Heme Oxygenase-1 Regulates Inflammation and Mycobacterial Survival in Human Macrophages during Mycobacterium tuberculosis Infection

Caitlyn R. Scharn, Angela C. Collins, Vidhya R. Nair, Chelsea E. Stamm, Denise K. Marciano, Edward A. Graviss and Michael U. Shiloh
J Immunol June 1, 2016, 196 (11) 4641-4649; DOI: https://doi.org/10.4049/jimmunol.1500434

Abstract

Mycobacterium tuberculosis, the causative agent of tuberculosis, is responsible for 1.5 million deaths annually. We previously showed that M. tuberculosis infection in mice induces expression of the CO-producing enzyme heme oxygenase (HO1) and that CO is sensed by M. tuberculosis to initiate a dormancy program. Further, mice deficient in HO1 succumb to M. tuberculosis infection more readily than do wild-type mice. Although mouse macrophages control intracellular M. tuberculosis infection through several mechanisms, such as NO synthase, the respiratory burst, acidification, and autophagy, how human macrophages control M. tuberculosis infection remains less well understood. In this article, we show that M. tuberculosis induces and colocalizes with HO1 in both mouse and human tuberculosis lesions in vivo, and that M. tuberculosis induces and colocalizes with HO1 during primary human macrophage infection in vitro. Surprisingly, we find that chemical inhibition of HO1 both reduces inflammatory cytokine production by human macrophages and restricts intracellular growth of mycobacteria. Thus, induction of HO1 by M. tuberculosis infection may be a mycobacterial virulence mechanism to enhance inflammation and bacterial growth.

Introduction

One-third of the world’s population is latently infected with Mycobacterium tuberculosis, leading to 9 million new cases of tuberculosis every year and another 1–2 million deaths (1). M. tuberculosis is spread from person to person through aerosolized droplets (2). Following infection, the bacteria are taken up by alveolar macrophages found in the terminal alveoli of the lungs (3, 4). Cytokines produced by these macrophages recruit a large number of immune cells to the lung, including dendritic cells, B cells, T cells, and additional monocytes. In the majority of cases, bacterial replication is restricted, leading to latent infection (5, 6). However, in ∼5% of early cases, bacterial replication is not controlled, which leads to active disease. Importantly, how the human immune system responds to M. tuberculosis infection remains poorly understood.

We (7) and others (8) have shown that M. tuberculosis infection in mice induces heme oxygenase (HO1), and M. tuberculosis senses CO, a downstream product of HO1, to alter its transcriptional program (7, 9). HO1 catalyzes the breakdown of heme into carbon monoxide, biliverdin, and iron (7, 10). We found that M. tuberculosis infection of mouse macrophages in vitro results in production of HO1, and HO1 accumulation is observed within macrophages in M. tuberculosis–infected mice (7). In addition, HO1 knockout mice show increased susceptibility to M. bovis bacillus Calmette-Guérin and M. avium infection (11, 12), likely owing to the cytotoxic effects of free heme accumulation (12). Although HO1 is important for controlling murine M. tuberculosis infection, whether HO1 plays a similar role in human infection is not known. Based on previous work, we hypothesized that HO1 would be produced by human macrophages in response to M. tuberculosis infection and that HO1 expression would have an impact on tuberculosis pathogenesis.

In this article, we report that HO1 was robustly expressed both in human tuberculosis lesions and in human macrophages in response to M. tuberculosis infection, and HO1 directly colocalized with M. tuberculosis within the phagosomes of infected human macrophages. Moreover, HO1 regulated inflammatory cytokine production such that significant reductions in proinflammatory cytokines were observed when HO1 was inhibited in both macrophage cell lines and primary human macrophages. Surprisingly, when HO1 was inhibited in primary human macrophages infected with M. tuberculosis, intracellular growth of M. tuberculosis was diminished coincident with reduced autocrine exposure to proinflammatory cytokines. Together, our results suggest a vital role for HO1 during human tuberculosis by mediating inflammatory cytokine production and facilitating intracellular growth.

Materials and Methods

Strains and media

The Erdman wild-type strain of M. tuberculosis was grown in Middlebrook 7H9 medium or on Middlebrook 7H11 agar plates supplemented with 10% oleic acid–albumin–dextrose–catalase (Thermo Fisher). Tween 80 (Fisher) was added to liquid medium to a final concentration of 0.05%. M. tuberculosis Erdman expressing GFP or mCherry was used for immunofluorescence experiments and was grown in the same 7H9 broth medium as above.

Cell maintenance and macrophage differentiation

THP1 and U937 cells were grown in suspension in RPMI 1640 media supplemented with 10% FBS, 2.5% D-glucose, 2 mM L-glutamine, ± penicillin/streptomycin (100 U/ml and 100 μg/ml, respectively), 10 mM HEPES, and 1 mM sodium pyruvate. To differentiate THP1 and U937 cells into macrophages, cells were treated with PMA (100 ng/ml; 16561-29-8, Fisher) and plated at a density of 5 × 105 cells per well of a 24-well plate. Cells were kept at 37°C, 5% CO2 for 2 d, at which time the media were changed to antibiotic-free media. Cells were kept at 37°C, 5% CO2 for an additional 2 d prior to M. tuberculosis infection.

Primary human macrophages were collected from 30 ml citrate-treated blood samples and prepared according to standard techniques (13). Briefly, blood was separated on a Ficoll-Paque PLUS gradient by centrifugation at 750 × g for 20 min without braking. Autologous serum was collected and frozen for M. tuberculosis infection. The lymphocyte/monocyte layer was transferred to a 50-ml centrifuge tube, diluted to 50 ml with PBS, and centrifuged at 350 × g for 20 min at 4°C without braking. Monocytes were washed three more times in PBS at 4°C. Finally, monocytes were resuspended in 5 ml RPMI 1640 + 10% autologous human serum. To differentiate into macrophages, monocytes were diluted with RPMI 1640 + 10% human serum + 100 ng/ml M-CSF (216-MC-025, R&D Systems) and plated in 24-well Primaria Plates (353847, Corning). Cells were allowed to attach at 37°C 5% CO2 for 4 h before being rinsed with PBS and adding fresh RPMI 1640 + 10% human serum + M-CSF. Cells were differentiated for 7 d, and medium was changed every other day before infection.

Western blotting

THP-1 cells were infected at a multiplicity of infection (MOI) of 10. At 24 h postinfection, cells were lysed in 0.5% Triton X-100 (Sigma-Aldrich) plus protease inhibitors (Roche), and samples were filtered twice with a 0.22-μm microcentrifuge filter (Millipore). Blotting was performed with rabbit anti-HO1 polyclonal Ab (1:1000; ADI-SPA-896-F, Enzo Life Sciences) or mouse anti–β-actin Ab (1:1000; sc-47778, Santa Cruz) and donkey anti-rabbit–HRP conjugate secondary Ab (1:5000; sc-2305, Santa Cruz) or goat anti-mouse–HRP conjugate (1:5000; sc-2005, Santa Cruz), respectively. Supersignal West Femto Chemiluminescent Substrate (Pierce) was used for signal detection.

HO1 ELISA

The HO1 ELISA was performed according to the manufacturer’s instructions (HO1 ELISA Kit, ADI-EKS-800, Enzo). Clinical samples of serum from individuals with active tuberculosis were obtained from the Houston Tuberculosis Initiative (14–17). Briefly, primary human macrophages were lysed in extraction buffer at various time points after infection or hemin treatment. Primary human macrophage lysates were diluted to 20 ng/ml total protein with kit-provided sample diluent, and human M. tuberculosis patient serum was diluted 1:10 in sample diluent. A total of 100 uL of each diluted sample and standard was added in duplicate to a 96-well plate and plate incubated at room temperature for 30 min. Excess sample was flicked off the plate, and wells were washed six times with 200 μl kit-provided wash buffer. Each well was treated with 100 μl kit-provided rabbit anti-human HO1 Ab for 1 h at room temperature. After washing and drying, 100 μl kit-provided anti-rabbit IgG:HRP conjugate was added to each well for 30 min at room temperature. The plate was washed again, and 100 μl kit-provided TMB Substrate was added to each well for 15 min at room temperature, followed directly by 100 μl kit-provided Stop Solution. The plate was read immediately on a Synergy HT plate reader (BioTek Instruments) at 450 nm.

Quantitative PCR

THP-1 cells were infected at an MOI of 10. At 24 h, infected cells were lysed using TRIzol and the RNA was purified. cDNA was prepared using the iScript cDNA synthesis kit (Bio-Rad). For quantitative PCR (qPCR) analysis, Fast Sybr Green (Life Technologies) was used. Primers were from Sigma-Aldrich. The qPCR was performed on an Applied Biosystems Vii7 using the following primers: HO-1 forward (5′-CAACAAAGTGCAAGATTCTG-3′), HO-1 reverse (5′-TGCATTCACATGGCATAAAG-3′), IFN-β forward (5′-CGCTGCGTTCCTGCTGTGCTT-3′), IFN-β reverse (5′-AGGTGAGGTTGATCTTTCCATTCA-3′), β-actin forward (5′-GGTGTGATGGTGGGAATGG-3′), and β-actin reverse (5′-GCCTCGTCACCCACATAGGA-3′). All fold change calculations for gene expression were made using the ΔΔCt method (18).

Mouse infections

BALBc mice (The Jackson Laboratory) were infected using a Glas-Col aerosol-exposure chamber to deliver ∼200 bacilli per mouse, as previously described (19, 20). Prior to aerosolization, bacteria were washed repeatedly and sonicated to generate a single-cell suspension. At day 0, we plated total organ homogenates from both lungs (five mice per group) to determine the initial inoculum. At subsequent time points, lungs from infected animals were fixed in 10% neutral buffered formalin for 24 h and paraffin embedded. Animal experiments were reviewed and approved by the Institutional Animal Care and Use Committee of the University of Texas (UT) Southwestern.

Immunohistochemistry and immunofluorescence of tissue specimens

Sections from paraffin-embedded mouse and human lung were deparaffinized in xylene and subjected to heat-mediated Ag retrieval in 1 mM EDTA with 0.05% Tween 20 (pH 8.0); endogenous peroxide activity was quenched in 2.5% hydrogen peroxide solution in methanol; and slides were permeabilized in 0.1% Triton-100 (Sigma-Aldrich) and blocked in SuperBlock Blocking Buffer (Thermo Scientific). For immunohistochemistry, sections were stained with rabbit anti-HO1 (1:100; ADI-SPA-896-F, Enzo Life Sciences), rabbit anti-HA (1:50; sc-805, Santa Cruz), or mouse anti-CD68 (1:100; ab955, Abcam). HRP-conjugated secondary Abs were obtained from Jackson Immunochemicals and used at 1:250. Staining was amplified with AB reagent (VECTASTAIN) and detected using diaminobenzidine (DAB) reagent (Thermo Scientific). Images were acquired using a Zeiss Axioplan 2 microscope. For immunofluorescence, HO1 was identified using rabbit anti-HO1 (1:100) and a donkey anti-rabbit–HRP conjugate secondary Ab (1:500; Santa Cruz) followed by amplification with cyanine 3 tyramide (1:100; PerkinElmer). M. tuberculosis was identified using guinea pig anti–M. tuberculosis (1:25; NR-13818, NR-13823, BEI Resources) and an Alexa 488–conjugated donkey anti–guinea pig secondary Ab (1:100; 706-545-148, Jackson Immunochemicals). All commercial Abs were prepared without Freund’s adjuvant. Images were acquired using a Leica TCS SP5 confocal microscope.

Quantification of immunohistochemistry in mouse lungs

DAB-stained sections were imaged as above and analyzed with ImageJ using the color deconvolution plug-in (http://wiki.imagej.net/Colour_Deconvolution), as described (21). Briefly, images were opened in ImageJ and then processed using the color deconvolution tool to separate brown and purple images. The area of brown staining was then quantified and divided by the total (purple) area to yield a percentage of staining area. Three mice were analyzed per time point.

Mycobacterial preparation for macrophage infections

Bacteria for infection were prepared as described previously (7, 22). Briefly, M. tuberculosis were grown to late-log phase and washed repeatedly with PBS. After the final wash, bacterial clumps were removed by slow-speed centrifugation (300 × g), and the resulting supernatant was then sonicated to break up any remaining clumps and generate a single-cell suspension. After sonication, the bacterial OD600 was recorded and the bacteria were diluted into macrophage-infection media (RPMI 1640 plus 10% horse serum [THP1 and U937 cells] or 10% autologous human serum [primary cells]).

Immunofluorescence of macrophages infected in vitro

U937 or THP-1 cells were differentiated as described above. Cells were infected at an MOI of 5, and 24 h postinfection, cells were washed with PBS and fixed in 4% paraformaldehyde (Alfa Aesar). Cells were permeabilized with 0.1% Triton X-100 and blocked with SuperBlock Blocking Buffer (Thermo Scientific). HO1 was identified using rabbit anti-HO1 (1:100) and an HRP-conjugated donkey anti-rabbit secondary (1:500; Santa Cruz), followed by amplification with cyanine 3 tyramide (1:100; PerkinElmer). M. tuberculosis expressing GFP were used for infections in THP1 and U937 cells. Images were acquired as z-stacks using a Zeiss Axioplan 2 microscope and were deconvoluted using Imaris and Autoquant softwares. Additional Abs used to identify the location of HO1-positive bacteria were mouse anti-LAMP1 (lysosome, 1:100; sc-20011, Santa Cruz), mouse anti-Rab7 (late endosome, 1:100; ab50533, Abcam), mouse anti-SQSTM1 (for p62/sequestosome, 1:2000; H00008878-MO1, Abnova), and mouse anti-Sec22B (endoplasmic reticulum–Golgi intermediate compartment, 1:50; sc-101267, Santa Cruz).

Immunofluorescence of mycobacteria

M. tuberculosis expressing mCherry were grown in 7H9 media to log phase and then fixed in 4% paraformaldehyde for 30 min. Fixed M. tuberculosis were then incubated with primary Abs against M. tuberculosis (rabbit anti–M. tuberculosis), HO1 (rabbit anti-HO1; Enzo Life Sciences), or rabbit IgG, all at a 1:100 dilution. Then, an Alexa 488–conjugated donkey anti-rabbit secondary Ab (1:500; A21206, Life Technologies) was added. After incubation, M. tuberculosis were immobilized on glass slides and imaged using a Zeiss Axioplan 2 fluorescence microscope.

Magpix cytokine analysis

U937 cells were infected at an MOI of 10 for 2 h. Cells were washed with PBS, and fresh medium ± 0.1 M NaOH (vehicle control) or tin protoporphyrin (SnPP) (50 μM final concentration) was added. SnPP blocks the active site of HO1, leading to competitive inhibition with a nanomolar Ki (23). At 24 h, conditioned medium was collected and filtered twice through a 0.22-μm filter. Cytokine levels were measured using a 29-plex Magpix cytokine assay (HCYTMAG-60K-PX29; Millipore).

Primary human cells were infected at an MOI of 10 for 2 h. Cells were washed with PBS, and fresh medium ± SnPP (50 μM) or cobalt protoporphyrin (CoPP) (10 μM final concentration) was added (24). At 24 h, conditioned medium was collected and filtered twice through a 0.22-μm filter. IL-1β, IL-8, TNF-α, IL-6, and GM-CSF levels were measured using the Human Inflammatory Magnetic 5-Plex Panel Assay (LHC0003M; Thermo Fisher). SnPP and CoPP were both from Fisher Scientific. Both were dissolved in 0.1 M NaOH and sterile filtered prior to use.

Survival in macrophages

Primary human macrophages were infected at an MOI of 0.1 for 2 h with a single-cell suspension of bacteria prepared as described above, using autologous serum for each donor during the initial infection period. Macrophages were washed, and macrophage medium without antibiotics (±50 μM SnPP or 10 μM CoPP) was then replaced. To avoid the need for antibiotics, cells were gently washed with PBS daily and fresh medium ± SnPP or CoPP was added. On days 0, 3, and 6, cells were lysed using 0.5% Triton X-100 (Sigma-Aldrich) and serial dilutions were plated to 7H11 agar without antibiotics to determine CFU values.

Human studies

The Institutional Review Boards of the University of Texas Southwestern and Methodist Hospital approved all research using human specimens.

Statistical analysis

The qPCR was analyzed by unpaired Student t test. Cytokine data from conditioned media were first analyzed by two-way ANOVA, followed by head-to-head comparisons with Tukey’s correction for multiple comparisons. Colocalization of M. tuberculosis with markers was analyzed by unpaired Student t test. HO1 cytokine data in serum were analyzed by unpaired Student t test. All analyses were performed using Prism 6 (GraphPad Software).

Results

HO1 colocalizes with M. tuberculosis during chronic aerosol infection

Previously we showed that HO1 is induced 10 d after i.v. infection of mice with M. tuberculosis (7). To determine if HO1 expression is induced during a more physiologically relevant M. tuberculosis infection, we used a mouse aerosol infection model. We performed an aerosol infection with a low dose of M. tuberculosis Erdman (200 CFU per mouse); harvested lungs from BALB/c mice 21, 42, 77, and 112 d postinfection; and assessed HO1 and M. tuberculosis expression by immunohistochemistry and immunofluorescence microscopy. By immunohistochemistry we found that HO1 was modestly expressed at 21 d and then gradually increased over time, so that at 112 d it was robustly expressed within characteristic foamy macrophages (Fig. 1A, 1B). To show that HO1 and M. tuberculosis exist within the same cellular organelles, we used confocal microscopy and found that within infected cells, HO1 and M. tuberculosis colocalized, with HO1 expression often surrounding individual bacteria (Fig. 1C). Thus, HO1 expression appeared to increase over the course of infection, and HO1 and M. tuberculosis colocalized to the same cells and organelles.

FIGURE 1.
FIGURE 1.

HO1 colocalizes to M. tuberculosis (Mtb)–infected macrophages in an aerosol model of infection. (A) BALB/c mice were infected with M. tuberculosis via a low-dose aerosol infection, and lungs were harvested at days 21, 42, 77, and 112. Shown is immunohistochemistry of paraffin sections with anti-HO1 (brown) and counterstained with hematoxylin (purple). Scale bars, 500 μm for 4× images and 100 μm for 20× images. (B) The extent of DAB positivity was quantified using ImageJ and graphed over time as percentage of positive area (three mice per time point). *p < 0.05 by Kruskall–Wallis test. (C) Immunofluorescence of day 112 paraffin sections with rabbit anti-HO1 and guinea pig anti–M. tuberculosis. Scale bars, 10 μm.

HO1 is expressed in CD68+ cells within tissue from M. tuberculosis–infected individuals

To determine the role of HO1 in human tuberculosis, we first examined HO1 expression by immunohistochemistry and immunofluorescence microscopy in lung biopsy specimens from 10 M. tuberculosis–infected individuals. These individuals and their specimens were randomly selected from a pool of individuals identified by the pathology department at Parkland Hospital (Dallas, TX) as being positive for M. tuberculosis by acid-fast bacillus staining and culture. A representative patient sample demonstrated an accumulation of HO1 in infected tissues, compared with results from staining with secondary Ab alone or with an isotype Ab control, both of which showed minimal DAB staining (Fig. 2A). HO1 expression was found in 10 of 10 tested individuals to varying degrees, whereas HO1 expression was only rarely observed in 4 of 4 autopsy specimens from uninfected individuals (Fig. 2A, Supplemental Fig. 1, and 5 other M. tuberculosis–infected individuals not shown). Occasionally, HO1 staining was found in areas outside traditional granuloma, including in epithelial cells and cells with a monocytic appearance, although the majority of HO1 was found within granuloma. To exclude that the HO1 Ab was detecting M. tuberculosis Ags, we stained fluorescent M. tuberculosis with a rabbit anti–M. tuberculosis Ab, rabbit polyclonal Ig, or rabbit anti-HO1 Ab and found that the HO1 Ab did not cross-react with M. tuberculosis alone (Supplemental Fig. 2). To determine if HO1 expression was increased in M. tuberculosis–infected monocytes and macrophages, we stained serial sections for HO1, M. tuberculosis, or the macrophage/monocyte marker CD68. We found that HO1 and M. tuberculosis–positive cells were also CD68 positive (Fig. 2B). By deconvolution of z-stacked immunofluorescence microscopy images, we found that, similar to our observations of M. tuberculosis–infected mouse tissue (Fig. 1), HO1 colocalized to M. tuberculosis–infected cells in lungs from individuals with active tuberculosis (Fig. 2C). Furthermore, HO1 appeared to envelop discrete bacteria (Fig. 2C) in the same staining pattern as M. tuberculosis–infected mouse lungs (Fig. 1). Thus HO1 was expressed in human lung specimens from M. tuberculosis–infected individuals within monocytes and macrophages.

FIGURE 2.
FIGURE 2.

HO1 is expressed in human tuberculosis specimens. (A) Immunohistochemistry was performed on paraffin-embedded lung sections from M. tuberculosis (Mtb)–infected individuals using anti-HO1 Ab and DAB detection (brown). Sections were counterstained with hematoxylin (purple). Control specimens were treated with either rabbit polyclonal serum (control Ab) or secondary Ab alone. Scale bars, 100 μm. (B) Serial sections were stained with anti-HO1, anti-CD68, or anti–M. tuberculosis, as in (A). Scale bars, 10 μm. (C) Immunofluorescence was performed on paraffin sections with anti-HO1 (green) and anti–M. tuberculosis (red).

M. tuberculosis infection is sufficient to induce HO1 expression in human macrophages

To determine whether M. tuberculosis infection alone is sufficient to induce HO1 in human macrophages, we measured the regulation of HO1 by M. tuberculosis infection in the human cell lines U937 and THP1, and in primary human macrophages differentiated from healthy donor monocytes. In THP1 cells we observed that M. tuberculosis infection was sufficient to induce HO1 transcription (Fig. 3A) and accumulation of HO1 protein by Western blotting (Fig. 3B). We also quantified HO1 using a specific HO1 ELISA in primary human macrophages and noted a 3-fold increase in HO1 after 8 h of infection (Fig. 3C). Furthermore, we confirmed a prior report (25) that individuals with active tuberculosis have increased serum HO1 by comparing serum HO1 from healthy donors with those from individuals with active tuberculosis (Fig. 3D). We next infected macrophages with fluorescent M. tuberculosis and found by deconvolution of z-stacked microscopy images that M. tuberculosis–infected THP1, U937, and primary human macrophages induced HO1 expression in a very focal distribution compared with uninfected cells (Fig. 3E , 3G, 3I, Supplemental Fig. 3, Supplemental Video 1). Whereas uninfected cells demonstrated faint and diffuse HO1 staining, during M. tuberculosis infection HO1 frequently colocalized with M. tuberculosis (Fig. 3E, 3G, 3I). In particular, HO1 staining surrounded individual bacteria in infected macrophages (Fig. 3E, 3G, 3I and insets, Supplemental Fig. 3, Supplemental Video 1), with ∼40–50% of bacteria colocalizing with HO1 (Fig. 3F, 3H, 3J).

FIGURE 3.
FIGURE 3.

M. tuberculosis (Mtb) infection is sufficient to induce HO1 expression in human macrophages. (A) qPCR analysis of HO1 gene expression in THP1 cells following M. tuberculosis infection (MOI 10) for 24 h. *p < 0.05 by Student t test. (B) Western blot of lysates from THP1 cells mock-infected or infected with M. tuberculosis (MOI 10) for 24 h and probed with anti-HO1 or anti-actin Ab. (C) HO1 ELISA from macrophages derived from a healthy donor and then infected with M. tuberculosis for 8 or 24 h. *p < 0.05 by Student t test. (D) HO1 concentration in the serum of M. tuberculosis–infected versus healthy control individuals was determined by ELISA. (E–J) Confocal microscopy and quantification showing HO1 localization in THP1 (E and F), U937 (G and H), and primary human macrophages (I and J) following M. tuberculosis infection at an MOI of 5 and stained 24 h postinfection. (E, G, and I) HO1 (green) was detected using anti-HO1 Ab, and M. tuberculosis (shown in red) was detected by its GFP fluorescence and false colored to red in ImageJ. Scale bars, 10 μm. (F, H, and J) Quantification of HO1 and M. tuberculosis colocalization in macrophages. A total of 100 bacteria were counted in triplicate slides, and colocalization was assessed by deconvolution of z-stacked images. Percentage of colocalization reflects the number of HO1-colocalizing bacteria per 100 total bacteria counted. Data are representative of at least two similar experiments. ****p < 0.0001 by Student t test.

To further determine the M. tuberculosis–containing organelle to which HO1 was recruited, we stained M. tuberculosis–infected THP1 macrophages with markers for the endoplasmic reticulum–Golgi intermediate compartment [also known as ERGIC (Sec22B)], late endosome (Rab7), autophagosome (p62), and lysosome (LAMP1) (Fig. 4, Supplemental Figs. 3, 4). In these experiments, we consistently observed 40% of M. tuberculosis colocalizing with HO1. Of the HO1-positive, M. tuberculosis–containing organelles, ∼40% also colocalized with LAMP1 (Fig. 4A–D, Supplemental Fig. 3, Supplemental Video 2). Thus, a total of ∼20% of total intracellular M. tuberculosis were HO1+/Lamp1+. In addition, we observed that although M. tuberculosis–containing organelles also colocalized with Rab7 (∼20% colocalization, Fig. 4E–H) and with p62 (–20% colocalization, Supplemental Fig. 4), these M. tuberculosis–containing organelles were not HO1 positive. Neither HO1 nor M. tuberculosis colocalized with the ERGIC marker Sec22B (Supplemental Fig. 4). Thus, M. tuberculosis infection of human macrophages in vitro upregulated HO1 expression, and HO1 was targeted to M. tuberculosis–containing organelles, some of which appeared to be lysosomes. In addition, some bacteria that were not HO1 positive also localized to either late endosomes or autophagosomes.

FIGURE 4.
FIGURE 4.

HO1 colocalizes with M. tuberculosis (Mtb) in LAMP1-positive organelles in human macrophages. Confocal microscopy and quantification showing HO1 colocalization with LAMP1 (A–D) or Rab7 (E–H) in THP1 cells following M. tuberculosis infection at an MOI of 5 and stained 24 h postinfection. HO1 (red) was detected using anti-HO1 Ab, LAMP1 or Rab7 (green) was detected with specific Abs, and M. tuberculosis (blue) was detected by its GFP fluorescence and false colored to blue in ImageJ. Nuclei (cyan) were detected using DAPI. Scale bars, 10 μm. For quantification of HO1, M. tuberculosis, or LAMP1/Rab7 colocalization in macrophages, 100 bacteria were counted on triplicate slides, and colocalization was assessed by deconvolution. Percentage of colocalization reflects the number of bacteria colocalizing with HO1 per 100 total bacteria counted (B and F), LAMP1 or Rab7 with M. tuberculosis (C and G), or HO1+ M. tuberculosis with LAMP1/Rab7 (D and H). Data are representative of two similar experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by Student t test.

HO1 inhibition prevents inflammatory cytokine production by M. tuberculosis–infected macrophages

To explore the functional significance of HO1 activity during M. tuberculosis infection, we infected U937 macrophages in the presence or absence of the HO1 inhibitor SnPP and determined the effect on inflammatory cytokine production. As has been previously observed, M. tuberculosis infection of human macrophages induced a number of cytokines, including IL-1β, TNF-α, and IL-6 (Fig. 5A). Surprisingly, inhibition of HO1 activity significantly decreased the accumulation of IL-1β, TNF-α, and IL-6 in the conditioned media (Fig. 5A). Inhibition of HO1 did not uniformly affect all cytokines, as another M. tuberculosis–induced cytokine, MIP-1β, was unaffected by treatment with SnPP (Fig. 5A). We also measured the induction of IFN-β by qPCR and found that HO1 inhibition significantly reduced its transcription in M. tuberculosis–infected macrophages (Fig. 5B). To confirm these results, we tested the role of HO1 in cytokine induction in primary human macrophages derived from healthy human donors. Similar to the results with the U937 cell line, primary human macrophages also induced IL-1β, TNF-α, IL-6, and GM-CSF when infected with M. tuberculosis, and this M. tuberculosis–dependent induction was partially blunted by SnPP treatment (Fig. 5C). In contrast, treatment with the HO1 inducer CoPP (24) did not have a statistically significant effect on cytokine production. Thus, we conclude that HO1 activity is important for the production of some, but not all, inflammatory cytokines following M. tuberculosis infection.

FIGURE 5.
FIGURE 5.

HO1 inhibition reduces inflammatory cytokine production during M. tuberculosis (Mtb) infection. (A and B) U937 cells were infected with M. tuberculosis or treated with PBS in the presence of SnPP or NaOH control for 24 h. (A) Supernatants were collected and assayed for IL-1β, IL-6, TNF-α, and MIP-1β, using Magpix multiplexing technology. Data are mean ±SD of samples in triplicate and are representative of two similar experiments. **p < 0.01, ***p < 0.001 by Student t test. (B) U937 cells were infected as described above with and without the presence of SnPP for 24 h. Macrophage RNA was collected, and qPCR was performed using IFN-β–specific primers with actin as the internal control. ***p < 0.001 by Student t test. (C) Primary human macrophages were infected with M. tuberculosis or treated with PBS in the presence of SnPP, CoPP, or control for 24 h. Supernatants were collected and assayed for IL-1β, IL-6, TNF-α, and GM-CSF, using Magpix multiplexing technology. Data are mean ±SD of samples in triplicate and are representative of two similar experiments. *p < 0.05, **p < 0.01, ***p < 0.001 by Student t test.

HO1 activity is permissive for M. tuberculosis growth within primary human macrophages

Because HO1-deficient mice are hypersusceptible to mycobacterial infection (11, 12), we hypothesized that HO1 induction in human macrophages might have antimicrobial activity. We first confirmed that SnPP or CoPP alone had no direct effect on M. tuberculosis by growing the bacteria with and without SnPP or CoPP added to the media and performing CFU assays over the course of 6 d. No significant growth difference was seen between bacteria treated with SnPP or CoPP and untreated controls (not shown). We next infected primary human macrophages with M. tuberculosis in the presence or absence of SnPP or CoPP and measured CFU from infected macrophages 3 and 6 d postinfection. Control, SnPP-treated, and CoPP-treated macrophages all demonstrated an initial inhibition of M. tuberculosis growth (Fig. 6A–C). However, by 6 d postinfection, M. tuberculosis growth was restricted in SnPP-treated macrophages compared with CoPP-treated and control macrophages (Fig. 6A, 6B). The effect of SnPP treatment was consistent across six independent human donors, where SnPP-treated macrophages had, on average, a 4- to 5-fold lower M. tuberculosis CFU 6 d postinfection (Fig. 6C). All of the donors induced HO1 in response to both M. tuberculosis and hemin (Fig. 6D). Thus, we conclude that HO1 activity facilitates growth of M. tuberculosis within primary human macrophages.

FIGURE 6.
FIGURE 6.

HO1 inhibition reduces bacterial growth in primary human macrophages. (A and B) Primary human macrophages from two independent donors were infected in quadruplicate with M. tuberculosis (Mtb) in the presence of NaOH, SnPP, or CoPP and CFU enumerated at 0, 3, and 6 d postinfection. Data are mean ±SD. ***p < 0.001 by ANOVA. (C) Primary human macrophages from six donors were infected in quadruplicate with M. tuberculosis in the presence of NaOH or SnPP and CFU enumerated at 0, 3, and 6 d postinfection. Fold CFU was calculated by dividing the mean CFU at each time point by the day 0 CFU for each donor. Data include donor 1 and 2 from (A) and (B). ***p < 0.001 by ANOVA. (D) HO1 ELISA was performed on lysates from donor macrophages infected with M. tuberculosis or treated with hemin. ****p < 0.001 by Student t test.

Discussion

We and others have shown previously that HO1 is induced shortly after M. tuberculosis infection in mice (7, 8). Compared with wild-type mice, HO1-deficient mice are less able to control mycobacterial infection, as indicated by increased bacterial numbers and mortality following infection (11, 12). In this article, we demonstrate that HO1 accumulated throughout the course of a chronic low-dose mouse infection. Modest HO1 expression was observed at day 21 and increased over time, so that by day 112, robust levels of HO1 were observed. In addition, HO1 colocalized with M. tuberculosis within mouse macrophages, appearing to surround the bacteria. Taken together, these data further support an important role for HO1 in murine M. tuberculosis infection.

Human tuberculosis and experimental murine tuberculosis infection differ in many ways, including how infected macrophages kill M. tuberculosis (26, 27), what cell types are found in experimental infections (28), how M. tuberculosis Ags are presented to the adaptive immune system (29, 30), and the architecture of infected tissue (31). The experimental models used in mice, such as i.v. infection, could potentially explain some of these differences. However, even the aerosol mouse model, although more physiological, still differs significantly from human infection (32, 33). Thus, it is important to test whether the functions of host genes identified in mouse models are shared in humans.

To determine the role of HO1 in human tuberculosis, we examined tissue samples from individuals infected with M. tuberculosis and identified that HO1 is expressed in human tuberculosis lesions. Similar to its localization in the lungs of mice infected with tuberculosis, HO1 colocalized with M. tuberculosis in human granulomas at both the cellular and subcellular level. The observation that HO1 is expressed within infected human lungs is consistent with the recent finding that plasma HO1 levels are elevated during active tuberculosis in humans, and that HO1 levels return to baseline following successful treatment (25). Because HO1 is known to be upregulated by a variety of stimuli, we wanted to determine if M. tuberculosis infection alone is sufficient to induce HO1. We found that in human macrophage–like cell lines and in primary human macrophages derived from healthy donors, HO1 was increased following infection and colocalized with M. tuberculosis, commonly colocalizing with a lysosomal marker. However, ∼50% of the HO1 and M. tuberculosis colocalization is in organelles not marked by p62, Sec22B, Rab7, or Lamp1, and some of the HO1 colocalization may be with intracytoplasmic M. tuberculosis (34).

Previously, HO1 was found to regulate expression of multiple cytokines (10). In purely inflammatory disease models, HO1 is anti-inflammatory (35, 36). Splenocytes from HO1 knockout mice secrete increased levels of proinflammatory cytokines following mitogenic stimulation with LPS or anti-CD3/anti-CD28 Abs (37). In addition, HO1 transgenic mice that constitutively express HO1 in the lung show a significant reduction in the production of proinflammatory cytokines and chemokines in response to hypoxia (38). However, in microbial infectious disease models, recent studies suggest HO1 may be involved in the induction of inflammatory cytokines. In mice, conditional deletion of HO1 from myeloid cells results in impaired IFN-β production as well as decreased production of IRF3-dependent target genes such as RANTES, IP-10, and MCP-1 in peritoneal macrophages in the setting of both viral and bacterial infection (39). Further, HIV infection of LPS-activated primary human monocyte–derived macrophages leads to increased HO1 expression and increased levels of the inflammatory cytokines MIP-1α and MIP-1β, which are attenuated following treatment with the HO1 inhibitor SnPP (40). Finally, it was recently demonstrated that CO derived from HO1 causes bacteria to release ATP, thus triggering P2X7 receptors to activate the NALP3 inflammasome and release the proinflammatory cytokine IL-1β (41). We found that inhibition of HO1 concurrent with M. tuberculosis infection in both macrophage cell lines and primary human macrophages suppressed the accumulation of multiple inflammatory cytokines, including IL-1β. Taking a human host–centric view, HO1 might therefore act as an important early component of the innate immune response leading to the production of proinflammatory cytokines such as TNF-α and IL-1β. Conversely, taking a mycobacteria-centric view, induction of HO1 may be a virulence mechanism. Studies in both zebrafish and mice indicate that granuloma formation in the context of M. marinum infection requires an appropriate level of TNF-α (42, 43), which can be induced by bacterial factors (44). Indeed, it was recently shown that HO1 could be induced in macrophages via the region of difference 1 (RD1) locus and its effector ESAT-6 (45).

The cytokines produced by M. tuberculosis–infected macrophages are IFN-β, IL-1β, IL-6, and TNF-α (46). A number of genetic determinants for M. tuberculosis susceptibility have been described (47, 48), including polymorphisms in the genes for TNF-α (49, 50), IL-1β (51), and IL-6 (52). Furthermore, TNF-α blockade is an established risk factor for developing active tuberculosis (53). Although we have demonstrated that secretion of these cytokines was significantly reduced following HO1 inhibition, what remains unclear is the mechanism behind this inhibition. One possibility is that reduced intracellular mycobacterial growth broadly diminishes the signals for cytokine production. Another possibility is that HO1-dependent production of proinflammatory cytokines is mediated through one of the products of HO1, such as iron, biliverdin, or CO, as has been reported for Enterococcus faecalis–infected macrophages (41). Alternatively, cytokine induction may be due to direct interaction of HO1 with other host molecules. For example, M. tuberculosis can directly activate the cytoplasmic DNA surveillance pathway through cGAS, resulting in IRF-3 phosphorylation and IFN-β production by infected macrophages (19, 54, 55). Of interest, HO1 directly interacts with IRF3, and HO1-deficient macrophages show a reduction in the expression of IFN-β and other IRF3 target genes (39), and we also observed diminished IFN-β transcription in infected macrophages. Whether inhibition of HO1 activity would also alter binding to, and nuclear translocation of, IRF-3 is unknown.

HO1 is important in controlling a variety of infections in mice, including M. avium (12), Listeria monocytogenes (56), Plasmodium falciparum (57), and Toxoplasma gondii (58). In human cells, HO1 can inhibit the replication of viruses such as HIV (40), Ebola (59), and hepatitis C virus (60). However, a direct role for HO1 in human macrophages infected with M. tuberculosis has not previously been established. We found that, in contrast to HO1−/− mice infected with mycobacteria, inhibition of HO1 in primary human macrophages was more restrictive to M. tuberculosis growth. This unexpected result is consistent with a recent report demonstrating reduced M. abscessus growth in THP-1 cells when HO-1 was inhibited by SnPP treatment or small interfering RNA–mediated silencing of HO1 (24). For M. abscessus–infected macrophages, the reduced growth was attributed to enhanced phagolysosomal fusion and increased reactive oxygen intermediate production (24). Another explanation for the enhanced growth of M. tuberculosis in human cells in an HO1-dependent manner is that HO1 may increase the availability of intracellular and intraphagosomal iron through heme catabolism. Because iron acquisition is essential for M. tuberculosis survival in the host (61, 62), increasing the free iron concentration could increase the ability of M. tuberculosis to replicate (63, 64). Indeed, for a number of intracellular pathogens, including Salmonella typhimurium, Chlamydia species, and Legionella pneumophila, increased intracellular iron enhances replication (65, 66). Thus, the cell-autonomous effect of HO1 in the context of M. tuberculosis infection of macrophages could be beneficial for M. tuberculosis growth.

In conclusion, we show that HO1 has an important dual role in human tuberculosis infection. HO1 was induced by aerosol M. tuberculosis infection in mice and in human macrophages, where it colocalized within infected cells. HO1 also mediated proinflammatory cytokine production in a human macrophage cell line and primary human macrophages while at the same time creating a permissive environment for intracellular bacterial replication. Future studies will be necessary to determine whether HO1 is beneficial or detrimental during human tuberculosis infection.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank D. Cavuoti for help with pathology specimens; B. Greenberg, D. Greenberg, S. Daly, and S. Hughes for help with human blood donors; and members of the Shiloh laboratory for helpful discussions.

Footnotes

  • C.R.S., A.C.C., and M.U.S. conceived and designed the study; C.R.S., C.E.S., A.C.C., and V.R.N. performed all of the experiments; C.R.S., A.C.C., V.R.N., and D.K.M. obtained microscopy images; E.A.G. provided human serum samples from the Houston Tuberculosis Initiative; C.R.S., A.C.C., and M.U.S. drafted the manuscript; and all authors edited and approved the final manuscript.

  • This work was supported by National Institutes of Health Grants RO1 AI099439, R21 AI111023, and U19 AI109725 (to M.U.S.), T32 AI5284 (to C.R.S.), T32 AI7520 (to C.E.S.), R01 DK099478 (to D.K.M.), and R01 DA09238 (to E.A.G.). Clinical specimens used for this project were partially supported by the National Institutes of Health (National Institute of Allergy and Infectious Diseases) under Contract N01-AO02738. M.U.S. acknowledges support from the Disease Oriented Clinical Scholars Program at University of Texas Southwestern.

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    CoPP
    cobalt protoporphyrin
    DAB
    diaminobenzidine
    HO1
    heme oxygenase
    MOI
    multiplicity of infection
    qPCR
    quantitative PCR
    SnPP
    tin protoporphyrin.

  • Received February 20, 2015.
  • Accepted March 22, 2016.

References

  1. ↵
    1. WHO
    . 2011. Global Tuberculosis Control. WHO Press, World Health Organization
  2. ↵
    1. Fennelly K. P.,
    2. J. W. Martyny,
    3. K. E. Fulton,
    4. I. M. Orme,
    5. D. M. Cave,
    6. L. B. Heifets
    . 2004. Cough-generated aerosols of Mycobacterium tuberculosis: a new method to study infectiousness. Am. J. Respir. Crit. Care Med. 169: 604–609.
    OpenUrlCrossRefPubMed
  3. ↵
    1. Smith D. W.,
    2. E. Wiegeshaus,
    3. R. Navalkar,
    4. A. A. Grover
    . 1966. Host-parasite relationships in experimental airborne tuberculosis. I. Preliminary studies in BCG-vaccinated and nonvaccinated animals. J. Bacteriol. 91: 718–724.
    OpenUrlAbstract/FREE Full Text
  4. ↵
    1. Dannenberg A. M., Jr..
    1982. Pathogenesis of pulmonary tuberculosis. Am. Rev. Respir. Dis. 125: 25–29.
    OpenUrlPubMed
  5. ↵
    1. Silva Miranda M.,
    2. A. Breiman,
    3. S. Allain,
    4. F. Deknuydt,
    5. F. Altare
    . 2012. The tuberculous granuloma: an unsuccessful host defence mechanism providing a safety shelter for the bacteria? Clin. Dev. Immunol. 2012: 139127.
    OpenUrlCrossRefPubMed
  6. ↵
    1. Modlin R. L.,
    2. B. R. Bloom
    . 2013. TB or not TB: that is no longer the question. Sci. Transl. Med. 5: 213sr6.
    OpenUrlFREE Full Text
  7. ↵
    1. Shiloh M. U.,
    2. P. Manzanillo,
    3. J. S. Cox
    . 2008. Mycobacterium tuberculosis senses host-derived carbon monoxide during macrophage infection. Cell Host Microbe 3: 323–330.
    OpenUrlCrossRefPubMed
  8. ↵
    1. Kumar A.,
    2. J. S. Deshane,
    3. D. K. Crossman,
    4. S. Bolisetty,
    5. B. S. Yan,
    6. I. Kramnik,
    7. A. Agarwal,
    8. A. J. Steyn
    . 2008. Heme oxygenase-1-derived carbon monoxide induces the Mycobacterium tuberculosis dormancy regulon. J. Biol. Chem. 283: 18032–18039.
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Zacharia V. M.,
    2. M. U. Shiloh
    . 2012. Effect of carbon monoxide on Mycobacterium tuberculosis pathogenesis. Med. Gas Res. 2: 30.
    OpenUrlCrossRefPubMed
  10. ↵
    1. Grochot-Przeczek A.,
    2. J. Dulak,
    3. A. Jozkowicz
    . 2012. Haem oxygenase-1: non-canonical roles in physiology and pathology. Clin. Sci. 122: 93–103.
    OpenUrlCrossRefPubMed
  11. ↵
    1. Regev D.,
    2. R. Surolia,
    3. S. Karki,
    4. J. Zolak,
    5. A. Montes-Worboys,
    6. O. Oliva,
    7. P. Guroji,
    8. V. Saini,
    9. A. J. Steyn,
    10. A. Agarwal,
    11. V. B. Antony
    . 2012. Heme oxygenase-1 promotes granuloma development and protects against dissemination of mycobacteria. Lab. Invest. 92: 1541–1552.
    OpenUrlCrossRefPubMed
  12. ↵
    1. Silva-Gomes S.,
    2. R. Appelberg,
    3. R. Larsen,
    4. M. P. Soares,
    5. M. S. Gomes
    . 2013. Heme catabolism by heme oxygenase-1 confers host resistance to Mycobacterium infection. Infect Immun. 81: 2536–2541.
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Erbel C.,
    2. G. Rupp,
    3. C. M. Helmes,
    4. M. Tyka,
    5. F. Linden,
    6. A. O. Doesch,
    7. H. A. Katus,
    8. C. A. Gleissner
    . 2013. An in vitro model to study heterogeneity of human macrophage differentiation and polarization. J. Vis. Exp. (72): e50332.
  14. ↵
    1. Ma X.,
    2. S. Dou,
    3. J. A. Wright,
    4. R. A. Reich,
    5. L. D. Teeter,
    6. H. M. El Sahly,
    7. R. J. Awe,
    8. J. M. Musser,
    9. E. A. Graviss
    . 2002. 5′ Dinucleotide repeat polymorphism of NRAMP1 and susceptibility to tuberculosis among Caucasian patients in Houston, Texas. Int. J. Tuberc. Lung Dis. 6: 818–823.
    OpenUrlPubMed
    1. Feske M. L.,
    2. L. D. Teeter,
    3. J. M. Musser,
    4. E. A. Graviss
    . 2011. Giving TB wheels: public transportation as a risk factor for tuberculosis transmission. Tuberculosis (Edinb.) 91(Suppl 1): S16–S23.
    OpenUrlCrossRef
    1. Marquez L.,
    2. M. L. Feske,
    3. L. D. Teeter,
    4. J. M. Musser,
    5. E. A. Graviss
    . 2012. Pediatric tuberculosis: the litmus test for tuberculosis control. Pediatr. Infect. Dis. J. 31: 1144–1147.
    OpenUrlCrossRefPubMed
  17. ↵
    1. Teeter L. D.,
    2. N. P. Ha,
    3. X. Ma,
    4. J. Wenger,
    5. W. A. Cronin,
    6. J. M. Musser,
    7. E. A. Graviss
    . 2013. Evaluation of large genotypic Mycobacterium tuberculosis clusters: contributions from remote and recent transmission. Tuberculosis (Edinb.) 93(Suppl): S38–S46.
    OpenUrlCrossRefPubMed
  18. ↵
    1. Livak K. J.,
    2. T. D. Schmittgen
    . 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods 25: 402–408.
    OpenUrlCrossRefPubMed
  19. ↵
    1. Collins A. C.,
    2. H. Cai,
    3. T. Li,
    4. L. H. Franco,
    5. X. D. Li,
    6. V. R. Nair,
    7. C. R. Scharn,
    8. C. E. Stamm,
    9. B. Levine,
    10. Z. J. Chen,
    11. M. U. Shiloh
    . 2015. Cyclic GMP-AMP synthase is an innate immune DNA sensor for Mycobacterium tuberculosis. Cell Host Microbe 17: 820–828.
    OpenUrlCrossRefPubMed
  20. ↵
    1. Zacharia V. M.,
    2. P. S. Manzanillo,
    3. V. R. Nair,
    4. D. K. Marciano,
    5. L. N. Kinch,
    6. N. V. Grishin,
    7. J. S. Cox,
    8. M. U. Shiloh
    . 2013. cor, a novel carbon monoxide resistance gene, is essential for Mycobacterium tuberculosis pathogenesis. MBio 4: e00721–e13.
    OpenUrlCrossRefPubMed
  21. ↵
    1. Ruifrok A. C.,
    2. D. A. Johnston
    . 2001. Quantification of histochemical staining by color deconvolution. Anal. Quant. Cytol. Histol. 23: 291–299.
    OpenUrlPubMed
  22. ↵
    1. Stanley S. A.,
    2. S. Raghavan,
    3. W. W. Hwang,
    4. J. S. Cox
    . 2003. Acute infection and macrophage subversion by Mycobacterium tuberculosis require a specialized secretion system. Proc. Natl. Acad. Sci. USA 100: 13001–13006.
    OpenUrlAbstract/FREE Full Text
  23. ↵
    1. Drummond G. S.,
    2. A. Kappas
    . 1981. Prevention of neonatal hyperbilirubinemia by tin protoporphyrin IX, a potent competitive inhibitor of heme oxidation. Proc. Natl. Acad. Sci. USA 78: 6466–6470.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Abdalla M. Y.,
    2. I. M. Ahmad,
    3. B. Switzer,
    4. B. E. Britigan
    . 2015. Induction of heme oxygenase-1 contributes to survival of Mycobacterium abscessus in human macrophages-like THP-1 cells. Redox Biol. 4: 328–339.
    OpenUrlCrossRefPubMed
  25. ↵
    1. Andrade B. B.,
    2. N. Pavan Kumar,
    3. K. D. Mayer-Barber,
    4. D. L. Barber,
    5. R. Sridhar,
    6. V. V. Rekha,
    7. M. S. Jawahar,
    8. T. B. Nutman,
    9. A. Sher,
    10. S. Babu
    . 2013. Plasma heme oxygenase-1 levels distinguish latent or successfully treated human tuberculosis from active disease. PLoS One 8: e62618.
    OpenUrlCrossRefPubMed
  26. ↵
    1. Philips J. A.,
    2. J. D. Ernst
    . 2012. Tuberculosis pathogenesis and immunity. Annu. Rev. Pathol. 7: 353–384.
    OpenUrlCrossRefPubMed
  27. ↵
    1. Stanley S. A.,
    2. J. S. Cox
    . 2013. Host-pathogen interactions during Mycobacterium tuberculosis infections. Curr. Top. Microbiol. Immunol. 374: 211–241.
    OpenUrlPubMed
  28. ↵
    1. Huynh K. K.,
    2. S. A. Joshi,
    3. E. J. Brown
    . 2011. A delicate dance: host response to mycobacteria. Curr. Opin. Immunol. 23: 464–472.
    OpenUrlCrossRefPubMed
  29. ↵
    1. Abebe F.,
    2. G. Bjune
    . 2009. The protective role of antibody responses during Mycobacterium tuberculosis infection. Clin. Exp. Immunol. 157: 235–243.
    OpenUrlCrossRefPubMed
  30. ↵
    1. Behar S. M.
    2013. Antigen-specific CD8(+) T cells and protective immunity to tuberculosis. Adv. Exp. Med. Biol. 783: 141–163.
    OpenUrlCrossRefPubMed
  31. ↵
    1. Tsai M. C.,
    2. S. Chakravarty,
    3. G. Zhu,
    4. J. Xu,
    5. K. Tanaka,
    6. C. Koch,
    7. J. Tufariello,
    8. J. Flynn,
    9. J. Chan
    . 2006. Characterization of the tuberculous granuloma in murine and human lungs: cellular composition and relative tissue oxygen tension. Cell. Microbiol. 8: 218–232.
    OpenUrlCrossRefPubMed
  32. ↵
    1. Via L. E.,
    2. P. L. Lin,
    3. S. M. Ray,
    4. J. Carrillo,
    5. S. S. Allen,
    6. S. Y. Eum,
    7. K. Taylor,
    8. E. Klein,
    9. U. Manjunatha,
    10. J. Gonzales,
    11. et al
    . 2008. Tuberculous granulomas are hypoxic in guinea pigs, rabbits, and nonhuman primates. Infect. Immun. 76: 2333–2340.
    OpenUrlAbstract/FREE Full Text
  33. ↵
    1. Nuermberger E.
    2008. Using animal models to develop new treatments for tuberculosis. Semin. Respir. Crit. Care Med. 29: 542–551.
    OpenUrlCrossRefPubMed
  34. ↵
    1. van der Wel N.,
    2. D. Hava,
    3. D. Houben,
    4. D. Fluitsma,
    5. M. van Zon,
    6. J. Pierson,
    7. M. Brenner,
    8. P. J. Peters
    . 2007. M. tuberculosis and M. leprae translocate from the phagolysosome to the cytosol in myeloid cells. Cell 129: 1287–1298.
    OpenUrlCrossRefPubMed
  35. ↵
    1. Willis D.,
    2. A. R. Moore,
    3. R. Frederick,
    4. D. A. Willoughby
    . 1996. Heme oxygenase: a novel target for the modulation of the inflammatory response. Nat. Med. 2: 87–90.
    OpenUrlCrossRefPubMed
  36. ↵
    1. Paine A.,
    2. B. Eiz-Vesper,
    3. R. Blasczyk,
    4. S. Immenschuh
    . 2010. Signaling to heme oxygenase-1 and its anti-inflammatory therapeutic potential. Biochem. Pharmacol. 80: 1895–1903.
    OpenUrlCrossRefPubMed
  37. ↵
    1. Kapturczak M. H.,
    2. C. Wasserfall,
    3. T. Brusko,
    4. M. Campbell-Thompson,
    5. T. M. Ellis,
    6. M. A. Atkinson,
    7. A. Agarwal
    . 2004. Heme oxygenase-1 modulates early inflammatory responses: evidence from the heme oxygenase-1-deficient mouse. Am. J. Pathol. 165: 1045–1053.
    OpenUrlCrossRefPubMed
  38. ↵
    1. Minamino T.,
    2. H. Christou,
    3. C. M. Hsieh,
    4. Y. Liu,
    5. V. Dhawan,
    6. N. G. Abraham,
    7. M. A. Perrella,
    8. S. A. Mitsialis,
    9. S. Kourembanas
    . 2001. Targeted expression of heme oxygenase-1 prevents the pulmonary inflammatory and vascular responses to hypoxia. Proc. Natl. Acad. Sci. USA 98: 8798–8803.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    1. Tzima S.,
    2. P. Victoratos,
    3. K. Kranidioti,
    4. M. Alexiou,
    5. G. Kollias
    . 2009. Myeloid heme oxygenase-1 regulates innate immunity and autoimmunity by modulating IFN-beta production. J. Exp. Med. 206: 1167–1179.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    1. Zhou Z. H.,
    2. N. Kumari,
    3. S. Nekhai,
    4. K. A. Clouse,
    5. L. M. Wahl,
    6. K. M. Yamada,
    7. S. Dhawan
    . 2013. Heme oxygenase-1 induction alters chemokine regulation and ameliorates human immunodeficiency virus-type-1 infection in lipopolysaccharide-stimulated macrophages. Biochem. Biophys. Res. Commun. 435: 373–377.
    OpenUrlCrossRefPubMed
  41. ↵
    1. Wegiel B.,
    2. R. Larsen,
    3. D. Gallo,
    4. B. Y. Chin,
    5. C. Harris,
    6. P. Mannam,
    7. E. Kaczmarek,
    8. P. J. Lee,
    9. B. S. Zuckerbraun,
    10. R. Flavell,
    11. et al
    . 2014. Macrophages sense and kill bacteria through carbon monoxide-dependent inflammasome activation. J. Clin. Invest. 124: 4926–4940.
    OpenUrlCrossRefPubMed
  42. ↵
    1. Tobin D. M.,
    2. J. C. Vary Jr..,
    3. J. P. Ray,
    4. G. S. Walsh,
    5. S. J. Dunstan,
    6. N. D. Bang,
    7. D. A. Hagge,
    8. S. Khadge,
    9. M. C. King,
    10. T. R. Hawn,
    11. et al
    . 2010. The lta4h locus modulates susceptibility to mycobacterial infection in zebrafish and humans. Cell 140: 717–730.
    OpenUrlCrossRefPubMed
  43. ↵
    1. Tobin D. M.,
    2. F. J. Roca,
    3. S. F. Oh,
    4. R. McFarland,
    5. T. W. Vickery,
    6. J. P. Ray,
    7. D. C. Ko,
    8. Y. Zou,
    9. N. D. Bang,
    10. T. T. Chau,
    11. et al
    . 2012. Host genotype-specific therapies can optimize the inflammatory response to mycobacterial infections. Cell 148: 434–446.
    OpenUrlCrossRefPubMed
  44. ↵
    1. Watkins B. Y.,
    2. S. A. Joshi,
    3. D. A. Ball,
    4. H. Leggett,
    5. S. Park,
    6. J. Kim,
    7. C. D. Austin,
    8. A. Paler-Martinez,
    9. M. Xu,
    10. K. H. Downing,
    11. E. J. Brown
    . 2012. Mycobacterium marinum SecA2 promotes stable granulomas and induces tumor necrosis factor alpha in vivo. Infect. Immun. 80: 3512–3520.
    OpenUrlAbstract/FREE Full Text
  45. ↵
    1. Andrade B. B.,
    2. N. Pavan Kumar,
    3. E. P. Amaral,
    4. N. Riteau,
    5. K. D. Mayer-Barber,
    6. K. W. Tosh,
    7. N. Maier,
    8. E. L. Conceição,
    9. A. Kubler,
    10. R. Sridhar,
    11. et al
    . 2015. Heme oxygenase-1 regulation of matrix metalloproteinase-1 expression underlies distinct disease profiles in tuberculosis. J. Immunol. 195: 2763–2773.
    OpenUrlAbstract/FREE Full Text
  46. ↵
    1. Torrado E.,
    2. A. M. Cooper
    . 2013. Cytokines in the balance of protection and pathology during mycobacterial infections. Adv. Exp. Med. Biol. 783: 121–140.
    OpenUrlCrossRefPubMed
  47. ↵
    1. Azad A. K.,
    2. W. Sadee,
    3. L. S. Schlesinger
    . 2012. Innate immune gene polymorphisms in tuberculosis. Infect. Immun. 80: 3343–3359.
    OpenUrlAbstract/FREE Full Text
  48. ↵
    1. Sivangala R.,
    2. M. Ponnana,
    3. S. Thada,
    4. L. Joshi,
    5. S. Ansari,
    6. H. Hussain,
    7. V. Valluri,
    8. S. Gaddam
    . 2014. Association of cytokine gene polymorphisms in tuberculosis patients and their household contacts. Scand. J. Immunol. 79: 197–205.
    OpenUrlCrossRefPubMed
  49. ↵
    1. Correa P. A.,
    2. L. M. Gomez,
    3. J. Cadena,
    4. J. M. Anaya
    . 2005. Autoimmunity and tuberculosis. Opposite association with TNF polymorphism. J. Rheumatol. 32: 219–224.
    OpenUrlAbstract/FREE Full Text
  50. ↵
    1. Merza M.,
    2. P. Farnia,
    3. S. Anoosheh,
    4. M. Varahram,
    5. M. Kazampour,
    6. O. Pajand,
    7. S. Saeif,
    8. M. Mirsaeidi,
    9. M. R. Masjedi,
    10. A. A. Velayati,
    11. S. Hoffner
    . 2009. The NRAMPI, VDR and TNF-alpha gene polymorphisms in Iranian tuberculosis patients: the study on host susceptibility. Braz. J. Infect. Dis. 13: 252–256.
    OpenUrlPubMed
  51. ↵
    1. Awomoyi A. A.,
    2. M. Charurat,
    3. A. Marchant,
    4. E. N. Miller,
    5. J. M. Blackwell,
    6. K. P. McAdam,
    7. M. J. Newport
    . 2005. Polymorphism in IL1B: IL1B-511 association with tuberculosis and decreased lipopolysaccharide-induced IL-1beta in IFN-gamma primed ex-vivo whole blood assay. J. Endotoxin Res. 11: 281–286.
    OpenUrlPubMed
  52. ↵
    1. Amirzargar A. A.,
    2. N. Rezaei,
    3. H. Jabbari,
    4. A. A. Danesh,
    5. F. Khosravi,
    6. M. Hajabdolbaghi,
    7. A. Yalda,
    8. B. Nikbin
    . 2006. Cytokine single nucleotide polymorphisms in Iranian patients with pulmonary tuberculosis. Eur. Cytokine Netw. 17: 84–89.
    OpenUrlPubMed
  53. ↵
    1. O’Garra A.,
    2. P. S. Redford,
    3. F. W. McNab,
    4. C. I. Bloom,
    5. R. J. Wilkinson,
    6. M. P. Berry
    . 2013. The immune response in tuberculosis. Annu. Rev. Immunol. 31: 475–527.
    OpenUrlCrossRefPubMed
  54. ↵
    1. Manzanillo P. S.,
    2. M. U. Shiloh,
    3. D. A. Portnoy,
    4. J. S. Cox
    . 2012. Mycobacterium tuberculosis activates the DNA-dependent cytosolic surveillance pathway within macrophages. Cell Host Microbe 11: 469–480.
    OpenUrlCrossRefPubMed
  55. ↵
    1. Watson R. O.,
    2. S. L. Bell,
    3. D. A. MacDuff,
    4. J. M. Kimmey,
    5. E. J. Diner,
    6. J. Olivas,
    7. R. E. Vance,
    8. C. L. Stallings,
    9. H. W. Virgin,
    10. 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.
    OpenUrlCrossRefPubMed
  56. ↵
    1. Tachibana M.,
    2. M. Hashino,
    3. T. Nishida,
    4. T. Shimizu,
    5. M. Watarai
    . 2011. Protective role of heme oxygenase-1 in Listeria monocytogenes-induced abortion. PLoS One 6: e25046.
    OpenUrlCrossRefPubMed
  57. ↵
    1. Seixas E.,
    2. R. Gozzelino,
    3. A. Chora,
    4. A. Ferreira,
    5. G. Silva,
    6. R. Larsen,
    7. S. Rebelo,
    8. C. Penido,
    9. N. R. Smith,
    10. A. Coutinho,
    11. M. P. Soares
    . 2009. Heme oxygenase-1 affords protection against noncerebral forms of severe malaria. Proc. Natl. Acad. Sci. USA 106: 15837–15842.
    OpenUrlAbstract/FREE Full Text
  58. ↵
    1. Araujo E. C.,
    2. B. F. Barbosa,
    3. L. B. Coutinho,
    4. P. V. Barenco,
    5. L. A. Sousa,
    6. C. M. Milanezi,
    7. G. Bonfá,
    8. W. R. Pavanelli,
    9. J. S. Silva,
    10. E. A. Ferro,
    11. et al
    . 2013. Heme oxygenase-1 activity is involved in the control of Toxoplasma gondii infection in the lung of BALB/c and C57BL/6 and in the small intestine of C57BL/6 mice. Vet. Res. (Faisalabad) 44: 89.
    OpenUrlCrossRef
  59. ↵
    1. Hill-Batorski L.,
    2. P. Halfmann,
    3. G. Neumann,
    4. Y. Kawaoka
    . 2013. The cytoprotective enzyme heme oxygenase-1 suppresses Ebola virus replication. J. Virol. 87: 13795–13802.
    OpenUrlAbstract/FREE Full Text
  60. ↵
    1. Chen M. H.,
    2. M. Y. Lee,
    3. J. J. Chuang,
    4. Y. Z. Li,
    5. S. T. Ning,
    6. J. C. Chen,
    7. Y. W. Liu
    . 2012. Curcumin inhibits HCV replication by induction of heme oxygenase-1 and suppression of AKT. Int. J. Mol. Med. 30: 1021–1028.
    OpenUrlPubMed
  61. ↵
    1. Rodriguez G. M.
    2006. Control of iron metabolism in Mycobacterium tuberculosis. Trends Microbiol. 14: 320–327.
    OpenUrlCrossRefPubMed
  62. ↵
    1. Rodriguez G. M.,
    2. I. Smith
    . 2006. Identification of an ABC transporter required for iron acquisition and virulence in Mycobacterium tuberculosis. J. Bacteriol. 188: 424–430.
    OpenUrlAbstract/FREE Full Text
  63. ↵
    1. Ganz T.
    2009. Iron in innate immunity: starve the invaders. Curr. Opin. Immunol. 21: 63–67.
    OpenUrlCrossRefPubMed
  64. ↵
    1. Soares M. P.,
    2. G. Weiss
    . 2015. The iron age of host-microbe interactions. EMBO Rep. 16: 1482–1500.
    OpenUrlAbstract/FREE Full Text
  65. ↵
    1. Nairz M.,
    2. I. Theurl,
    3. S. Ludwiczek,
    4. M. Theurl,
    5. S. M. Mair,
    6. G. Fritsche,
    7. G. Weiss
    . 2007. The co-ordinated regulation of iron homeostasis in murine macrophages limits the availability of iron for intracellular Salmonella typhimurium. Cell. Microbiol. 9: 2126–2140.
    OpenUrlCrossRefPubMed
  66. ↵
    1. Paradkar P. N.,
    2. I. De Domenico,
    3. N. Durchfort,
    4. I. Zohn,
    5. J. Kaplan,
    6. D. M. Ward
    . 2008. Iron depletion limits intracellular bacterial growth in macrophages. Blood 112: 866–874.
    OpenUrlAbstract/FREE Full Text
Back to top

In this issue

Print
Download PDF
Article Alerts
Email Article
Citation Tools

Jump to section