Epidemiological studies suggest that chronic exposure to air pollution increases susceptibility to respiratory infections, including tuberculosis in humans. A possible link between particulate air pollutant exposure and antimycobacterial immunity has not been explored in human primary immune cells. We hypothesized that exposure to diesel exhaust particles (DEP), a major component of urban fine particulate matter, suppresses antimycobacterial human immune effector cell functions by modulating TLR-signaling pathways and NF-κB activation. We show that DEP and H37Ra, an avirulent laboratory strain of Mycobacterium tuberculosis, were both taken up by the same peripheral human blood monocytes. To examine the effects of DEP on M. tuberculosis-induced production of cytokines, PBMC were stimulated with DEP and M. tuberculosis or purified protein derivative. The production of M. tuberculosis and purified protein derivative-induced IFN-γ, TNF-α, IL-1β, and IL-6 was reduced in a DEP dose-dependent manner. In contrast, the production of anti-inflammatory IL-10 remained unchanged. Furthermore, DEP stimulation prior to M. tuberculosis infection altered the expression of TLR3, -4, -7, and -10 mRNAs and of a subset of M. tuberculosis-induced host genes including inhibition of expression of many NF-κB (e.g., CSF3, IFNG, IFNA, IFNB, IL1A, IL6, and NFKBIA) and IFN regulatory factor (e.g., IFNG, IFNA1, IFNB1, and CXCL10) pathway target genes. We propose that DEP downregulate M. tuberculosis-induced host gene expression via MyD88-dependent (IL6, IL1A, and PTGS2) as well as MyD88-independent (IFNA, IFNB) pathways. Prestimulation of PBMC with DEP suppressed the expression of proinflammatory mediators upon M. tuberculosis infection, inducing a hyporesponsive cellular state. Therefore, DEP alters crucial components of antimycobacterial host immune responses, providing a possible mechanism by which air pollutants alter antimicrobial immunity.

Air pollution and tuberculosis (TB) each contribute significantly to global burden of disease. The World Health Organization estimates that air pollution, which is linked to a variety of illnesses including cardiopulmonary diseases and cancer, causes ∼2 million premature deaths worldwide per year (1). According to the World Health Organization’s 2009 Global Health Risk report (2), indoor air pollution (from biomass fuel and coal combustion) and urban outdoor air pollution rank 10th and 14th, respectively, among 19 leading risk factors for mortality in low- and middle-income countries. Exposure to ambient-air fine particulate matter <2.5 μm in aerodynamic diameter (PM2.5) is estimated to cause ∼1% of mortality from acute respiratory infections in children >5 y worldwide (3).

TB was estimated to afflict ∼10 million people and cause 2 million deaths in 2010 alone (4). Epidemiologic associations have been established between the incidence of TB and, respectively, cigarette smoking (511), occupational exposure to aerosolized silica (1214), and indoor air pollution (10, 11, 15). Despite these associations, to the best of our knowledge, no studies have been conducted to assess the mechanisms that underlie associations between air pollution and TB development in primary human cells.

Human exposure to diesel exhaust particles (DEP), a major component of urban PM2.5 in most industrialized urban areas (16), typically occurs by inhalation. Due to their small size, DEP remain airborne for prolonged time periods and deposit in the lungs upon inhalation. Likewise, infection with Mycobacterium tuberculosis typically occurs by inhalation of aerosolized M. tuberculosis-containing droplet nuclei (1–5 μm) that are released from patients with active TB during respiratory maneuvers (1720). Thus, concurrent respiratory exposure to DEP and M. tuberculosis can be expected to occur under real-life conditions and may subsequently alter host immune responses.

Protective antimycobacterial human host immunity involves innate and adaptive immune mechanisms that are primarily cell-mediated, involving monocytes/macrophages, dendritic cells, CD4+ and CD8+ T cells, NK, and γδT cells. M. tuberculosis Ag-specific immunity is characterized predominantly by a Th1 response (IL-2, IFN-γ, and TNF-α) (21) that is strongly enhanced and compartmentalized at the infection sites, most commonly the lungs (2224). IFN-γ, secreted from activated T cells and NK cells, is known to activate macrophages and promote bacterial killing (25) by permitting phagosomal maturation and production of antimicrobial inducible NO synthase and reactive oxygen intermediates (26). IFN-γ has an essential role in the control of mycobacterial infections both in the murine model (27, 28) and in humans (2933). Likewise, TNF-α plays an important role in killing of intracellular M. tuberculosis (34, 35) and granuloma formation. As evidenced by the increased risk of reactivation TB during TNF-α inhibitor treatment (36, 37), TNF-α is required in the maintenance of latent M. tuberculosis infection. IL-6 and IL-10 are important regulatory cytokines during the early phase of the mycobacterial infection of macrophages and in the inflammatory responses to M. tuberculosis and the regulation of IFN-γ, respectively (22).

Innate host resistance against M. tuberculosis depends to a large extent on the engagement of TLRs (3843) and nucleotide oligomerization domain-like receptors (44). TLR2, TLR4, and TLR9 are activated by M. tuberculosis-derived ligands such as lipoarabinomannan and mycobacterial 19-kDa protein (39, 40, 45), M. tuberculosis heat shock proteins 65 and 71 (38), and mycobacterial DNA (42), respectively. Stimulation of these TLRs (38, 4143) on monocytes, alveolar macrophages, or dendritic cells (4649) activates signal transduction pathways that culminate in the activation of MAPK, transcription factor NF-κB, and the IFN regulatory factor (IRF) family (47), leading to the release of antimicrobial effector molecules (50), proinflammatory cytokines (5157), and chemokines (58).

DEP exposure has been shown to shift Th1 to Th2 cytokine production by T, monocyte-derived dendritic, and spleen cells (59, 60), decrease secretion of IFN-γ, and increase secretion of IL-10 in murine bone marrow-derived dendritic cells (61). A study of the direct effects of DEP on M. tuberculosis immune responses showed that DEP increases the pulmonary M. tuberculosis burden concomitant with decreased production of proinflammatory cytokines (e.g., IL-1β, IL-12p40, and IFN-γ) in experimentally infected mice (62). DEP exposure also decreases the phagocytosis and bactericidal activity of rat alveolar macrophages exposed to Listeria monocytogenes infection (63, 64).

The current study was undertaken to examine the effects of DEP on human host immune responses to M. tuberculosis. We show that DEP significantly alter live M. tuberculosis and purified protein derivative (PPD)-induced cytokine production in primary human PBMC. Furthermore, DEP modulate M. tuberculosis-induced host gene expression by inhibiting TLR-mediated signaling including NF-κB and IRF pathways crucial for M. tuberculosis-induced host immunity. Suppression of M. tuberculosis-induced activation of these two pathways by DEP provides insights into the possible mechanisms by which particulate air pollution may modify pathogen-induced human host immune responses.

Approval to perform this study, collect personal health information, and perform venipunctures was given by the Institutional Review Boards of the University of Medicine and Dentistry of New Jersey (UMDNJ) in Newark and New Brunswick (Institutional Review Board protocol number 0120060235).

A total of 20 healthy individuals (13 male, 7 female, mean age 33.5 y [range 20–52 y]) were recruited from students and staff of the UMDNJ-School of Public Health, Rutgers University, the Environmental and Occupational Health Sciences Institute, and volunteers at the Chandler Clinic in New Brunswick, NJ. Inclusion and exclusion criteria were as follows: inclusion criteria: healthy women and men, 18–65 y of age; exclusion criteria: concurrent infections, use of immunosuppressive medications (steroidal and/or nonsteroidal anti-inflammatory medications, TNF-α inhibitors, antineoplastic chemotherapy), illnesses affecting host immunity (diabetes, HIV-1 infection, chronic liver or kidney diseases, and malignancies), smoking, and illicit drug use. A total of 80 ml heparinized venous whole blood was obtained from each individual from a cubital vein by venipuncture. Not all subjects were examined in all experiments. Subject numbers for each experiment are shown in the figures or figure legends. All study subjects provided written informed consent before personal health information was acquired and whole heparinized peripheral blood obtained by venipuncture.

Reagents were obtained from the following sources: ELISPOT assays: capture and biotinylated detection Abs for IL-1β and IL-6 (Cell Sciences, Canton, MA), IFN-γ, TNF-α (Endogen Pierce, Rockford, IL), IL-10, IL-4 (BD Pharmingen, San Diego, CA), streptavidin-peroxidase (Sigma-Aldrich, St. Louis, MO), chromogen 1% 3-amino-9-ethylcarbazole (Pierce, Rockford, IL), and MultiscreenHTS high protein binding 96-well plates (Millipore, Bedford, MA); PBMC isolation, culture, and stimulation: PHA (Sigma-Aldrich), LPS (Escherichia coli 026:B6; Sigma-Aldrich), PPD (Statens Serum Institute, Kopenhagen, Denmark), RPMI 1640 (BioWhittaker, Walkersville, MD), l-glutamine (Cellgro, Manassas, VA), pooled human AB serum (Gemini Bioproducts, Woodland, CA), and Ficoll-Paque (GE Healthcare Biosciences, Pittsburgh, PA); RNA extraction and quantitative RT-PCR (qRT-PCR): RNeasy mini kit (Qiagen, Germantown, MD), RNase-free DNase set (Qiagen), RT2 First Strand kit and RT2 qPCR Master Mix (SuperArray Bioscience, Frederick, MD), Power SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA), TaqMan reverse transcription reagents (Applied Biosystems); pathway-specific gene expression arrays: human Th1-Th2-Th3 (PAHS-34E) and TLR (PAHS-18E; SuperArray Bioscience); M. tuberculosis culture: H37Ra (ATCC no. 25177; American Type Culture Collection, Manassas, VA), Middlebrook 7H9 broth medium (Difco Laboratories, Detroit, MI) supplemented with 10% albumin dextrose catalase (Difco Laboratories), 7H10 solid agar plates (BD Biosciences, San Jose, CA); flow cytometry: Annexin V/propidium iodide (PI) apoptosis detection assay (BD Biosciences), PE-conjugated CD14 (BioLegend, San Diego, CA), and allophycocyanin-conjugated CD3 (eBioscience, San Diego, CA) mAbs; and DEP and carbon black (CB): DEP was a gift from Dr. Sagai (Tokyo, Japan), generated by a diesel-powered automobile, collected in a condensation trap, and stored at −80°C (65), and CB (Printex 90, primary particle diameter 16 nm) was from Degussa (Frankfurt, Germany).

DEP and CB stock suspensions (10 mg/ml) were prepared by 15-min sonication in PBS containing 0.05% Tween, aliquoted, and kept frozen at −20°C until use. Prior to addition to cell cultures, frozen aliquots of DEP or CB were thawed and vortexed for 5 min (a period determined to provide reliable dispersion of DEP in preliminary studies). DEP were then diluted in complete cell-culture medium to generate desired final concentrations (0.1, 1, 10, 50, and 100 μg/ml).

Presence of any endotoxin in complete culture fluid with and without DEP (100 μg/ml) was assessed using a commercial endotoxin detection service (Lonza Walkersville, Walkersville, MD) that employed the quantitative kinetic chromogenic Limulus amebocyte lysate (LAL) method (KQCL). The assay met all validation requirements per U.S. Food and Drug Administration-established guidelines for LAL testing (“Guideline on Validation of the Limulus Amebocyte Lysate Test as an End-Product Endotoxin Test for Human and Animal Parenteral Drugs, Biological Products, and Medical Devices”). The assays were conducted in adherence to USP <85> Bacterial Endotoxin Test and Lonza’s Quality System requirements. Testing was performed according to standard operating procedure 162.6.

To rule out the possibility of bacterial or fungal contamination of DEP, complete culture media (RPMI 1640 + 10% pooled human serum + L-Gln) and complete culture media containing DEP (100 μg/ml) were examined on blood brain heart infusion and inhibitory mode agar plates as well as on sheep blood and chocolate agar plates (all from Voigt Global Distribution, Lawrence, KS) at the Department of Microbiology at Robert Wood Johnson University Hospital in New Brunswick, NJ.

PBMC were isolated from whole heparinized venous blood by Ficoll gradient centrifugation (66). Briefly, whole blood was diluted 1:1 with l-glutamine supplemented RPMI 1640 medium and subjected to gradient density centrifugation (1200 rpm, 45 min, 21°C) over Ficoll-Paque. Following removal from the interface, PBMC were washed three times in RPMI 1640, resuspended in complete culture medium, counted, and adjusted at required concentrations. Viability of PBMC was 98–100% by trypan blue exclusion in all experiments.

Suspensions of M. tuberculosis (avirulent M. tuberculosis strain H37Ra) were prepared in Middlebrook 7H9 broth medium supplemented with 10% albumin dextrose catalase and 0.2% glycerol. After a 21-d incubation period at 37°C on an orbital shaker, M. tuberculosis stock suspensions were harvested, aliquoted, and stored at −86°C until use. The concentrations of the M. tuberculosis stock suspensions were confirmed by assessing CFU numbers from serial culture dilutions after 21 d of incubation on 7H10 solid agar plates. For PBMC infection experiments, single-cell suspensions of M. tuberculosis were prepared as follows: frozen M. tuberculosis stock was thawed, centrifuged for 5 min at 6000 × g, and resuspended in complete culture medium. To generate single-bacterial-cell suspensions, M. tuberculosis stock suspensions were declumped by vortexing for 5 min in the presence of five sterile 3-mm glass beads. Remaining M. tuberculosis clumps were removed with additional centrifugation at 350 × g for 5 min. Appropriate volumes of the supernatant M. tuberculosis suspensions were used for in vitro infections to obtain multiplicities of infection (MOIs) of 1 and 10. MOIs represent the ratio of M. tuberculosis to monocytes (bacteria/monocytes), and monocytes were assumed to constitute 10% of PBMC. Concentrations of frozen M. tuberculosis stock suspensions were confirmed after thawing of aliquots in each experiment by CFU assays.

Enriched CD14+CD3 peripheral blood monocytes were generated by negative selection through immunomagnetic depletion of nonmonocytes using a mixture of magnetic bead-coupled biotin-conjugated mAbs against CD3, CD7, CD16, CD19, CD56, CD123, and Glycophorin A (Miltenyi Biotec, Auburn, CA) from whole heparinized blood of an adult male healthy donor. The purity of the enriched monocytes was assessed by flow cytometry using a Cytomics FC500 Cytometer (Beckman Coulter, Miami, FL) and CXP software (Beckman Coulter). Monocytes were incubated on ice for 30 min with CD14-PE and CD3- allophycocyanin mAbs, washed with PBS, and acquired immediately without fixation for analysis. Cells were then gated in a forward scatter and side scatter dot plot to exclude debris, an FL2 and FL4 dot plot was drawn from the monocyte gate, and 20,000 cells were acquired for analysis. The CD14+CD3 monocyte population had a purity of 83.5%, which matched the manufacturer-predicted range (Miltenyi Biotec). CD14+CD3 monocytes were subsequently stimulated with DEP (10 μg/ml) and M. tuberculosis (MOI 10) in vitro and then incubated at 37°C in 5% CO2 for 24 h in 15-ml round-bottom polypropylene tubes. This cell-culture incubation period reflected the duration of the IFN-γ, IL-6, and IL-1β ELISPOT assays and that of the gene expression studies and provided ample time for DEP and M. tuberculosis uptake. CD14+CD3 monocytes were used for cytospin preparations and transmission electron microscopy (TEM) studies.

Uptake of DEP and M. tuberculosis by CD14+CD3 blood monocytes was assessed by histochemistry on thin-layer cytospin preparations of 150,000 cells per glass slide (cytoslides; Thermo Shandon, Pittsburgh, PA) using a cytocentrifuge (Shandon CytoSpin 4; Thermo Fisher Scientific, Waltham, MA). Cytospins were stained with Kinyoun stain (Alpha-Tec Systems, Vancouver, WA) and methyl blue counterstain (Thermo Shandon) following the manufacturer’s protocol to visualize M. tuberculosis and DEP particles. Photomicrographs were taken at original magnification ×1000 (oil immersion) using a BX51 microscope equipped with UPlanSApo lenses and a DP71 camera with DP image manager software (both from Olympus, Tokyo, Japan).

DEP and M. tuberculosis uptake within CD14+CD3 monocytes was further corroborated by TEM using a Philips CM12 transmission electron microscope (Philips). Cells were fixed in 2.5% glutaraldehyde/4% paraformaldehyde in 0.1 M cacodylate buffer and then postfixed in buffered 1% osmium tetroxide. Samples were subsequently dehydrated in a graded series of acetone and embedded in EMbed 812 resin (Electron Microscopy Sciences, Hatfield, PA), and 90-nm thin sections were cut on a Leica EM UC6 ultramicrotome (Leica Microsystems). Sectioned grids were stained with a saturated solution of uranyl acetate and lead citrate. Images were captured with an Advanced Microscopy Techniques XR111 digital camera (Advanced Microscopy Techniques) at 80 Kv.

DEP effects on apoptosis and necrosis in PBMC were assessed by flow cytometry (FACSCalibur; BD Biosciences, San Jose, CA) using an Annexin V/PI apoptosis detection kit as described by the manufacturer (BD Biosciences). Proportions of mononuclear cells that were simultaneously undergoing apoptosis (Annexin V positive) and necrosis (PI positive) were determined. DEP were added to PBMC at final concentrations of 0.1, 1, 10, and 100 μg/ml for 2, 24, and 72 h.

ELISPOT assays are highly sensitive tools for quantitation of frequencies of cytokine-producing cells and the semiquantitative assessment of their cytokine output (spot size) (67). In the current study, ELISPOT assays were performed to enumerate frequencies of IL-1β–, IL-4–, IL-6–, IL-10–, IFN-γ–, and TNF-α–producing cells. PBMC were stimulated simultaneously with DEP (0, 1, 10, 50, and 100 μg/ml) and M. tuberculosis at MOI 1 or MOI 10, or PPD (10 μg/ml), or LPS (100 ng/ml), or complete culture medium (control) in duplicate wells and incubated at 37°C in 5% CO2. PBMC and stimuli were cultured in a total volume of 200 μl/well in high protein-binding 96-well plates coated with appropriate anti-human primary capture Abs. Optimal densities of PBMC per well for the detection of the various cytokines were as follows: 20,000 for IL-6 and TNF-α, 100,000 for IL-1β, and 200,000 for IL-4, IL-10, and IFN-γ. Optimal short-term cell culture incubation periods for the ELISPOT assays were 4 h for TNF-α, 20 h for IL-1β, 24 h for IL-6 and IFN-γ, and 48 h for IL-4 and IL-10. These incubation periods were chosen on the basis of published studies (22, 23, 67, 68) and the manufacturer’s recommendations. Plates were then washed three times with PBS and PBS-Tween 0.05% followed by the addition of appropriate detection Abs. Plates were incubated for 1 h at 37°C for IL-1β and overnight at 4°C for all other cytokines. Following washing, cytokine spots were visualized with peroxidase-conjugated streptavidin, HRP, and chromogen 1% 3-amino-9-ethylcarbazole. After washing, plates were dried and scanned, images acquired, and frequencies of cytokine-producing cells (cytokine spots, spot-forming units) enumerated with an automated ELISPOT reader (Immunospot series 5 analyzer, software version 5.0; Cellular Technology, Cleveland, OH). Frequencies were calculated by averaging spot numbers from duplicate wells per stimulant and exposure condition. Frequencies of cytokine-producing cells (y-axes) were plotted as a function of DEP concentration in micrograms per milliliter (x-axes, shown in the bottom IL-1β panel only).

PBMC cultures were incubated at 37°C in 5% CO2 and stimulated with DEP, M. tuberculosis, M. tuberculosis plus DEP simultaneously, M. tuberculosis following a 20-h DEP prestimulation, and M. tuberculosis following a 20-h LPS prestimulation or left untreated (control) in six-well plates (Falcon; BD Biosciences) at concentrations of 1.5 × 106 cells/ml for 24 h. Total RNA was extracted from unstimulated (in complete culture medium only) or stimulated PBMC using the RNeasy mini kit (Qiagen), and DNA was removed from RNA samples by RNase-free DNase treatment (Qiagen). Concentration and purity of RNA were determined using a Nanodrop spectrophotometer (Nanodrop Technologies, Wilmington, DE). Total RNA (400 ng) was reverse transcribed into cDNA with RT2 First Strand kit (SuperArray Bioscience) according to the manufacturer’s protocol.

Human Th1-Th2-Th3 and TLR pathway-specific qRT-PCR arrays were used to assess mRNA expression in stimulated and unstimulated PBMC samples. First-strand cDNA diluted in water (102 μl) was mixed with 2× RT2 qPCR master mix (550 μl; SuperArray Bioscience), and the total reaction volume was adjusted to 1100 μl with RNAse-free water. Sample cocktails (10 μl/well) were loaded onto 384-well plates. A two-step cycling program consisting of 1 cycle of 95°C for 10 min, 40 cycles of 95°C for 15 s, and 60°C for 1 min was performed with an ABI 7900HT (Applied Biosystems, Foster City, CA) instrument. A dissociation curve specific to this instrument was used to assure primer quality. Levels of cDNA were calculated by the relative quantitation method (ΔΔCt method) from the PCR array data using the analysis software accessed at http://sabiosciences.com/pcrarraydataanalysis.php. Statistical differences in fold mRNA expression levels between stimulated and unstimulated PBMC were calculated using the same software.

Gene expression data obtained by qRT-PCR arrays were validated by real-time PCR with gene-specific forward and reverse primer pairs. Forward and reverse primers were designed using Oligoperfect software (Invitrogen), and cDNA was generated from RNA from PBMC using TaqMan reverse transcription reagents (Applied Biosystems) according to the manufacturer’s protocols. Briefly, 400 ng total RNA was used for the generation of cDNA in a 25-μl reaction (1× reverse transcription buffer, 5.5 mM MgCl2, 500 mM 2′-deoxynucleoside 5′-triphosphate, 2.5 μM random hexamer, 0.4 U/μl RNAse inhibitor, 1.25 U/μl Multiscribe Reverse Transcriptase, 0.4 M forward primer, and 0.4 M reverse primer). Thermal cycling parameters of the reverse transcription reaction were 25°C for 10 min, 48°C for 10 min, and 95°C for 5 min. Real-time PCR was performed with Power SYBR Green PCR master mix in an ABI 7900HT (Applied Biosystems). A two-step cycling program followed by dissociation curve as described in the section above was used.

We performed statistical analyses to test the following two hypotheses: the first was whether there was an effect for DEP within each stimulant; the second was whether each DEP dose affected M. tuberculosis MOI 1, MOI 10, and PPD stimulation effects differentially. Histograms of the values were examined for normality and other model assumptions. Linear mixed-effects models were employed with cytokine counts measured as a response and DEP dose as the predictor. Because all DEP doses were applied to each PBMC sample, a random effect for subject was included to account for the repeated measures. Also, based on observed heterogeneity, the variance of the cytokine counts was allowed to vary by DEP dose. The analyses were then stratified by each stimulant (M. tuberculosis MOI 1, M. tuberculosis MOI 10, PPD, and LPS). All the statistical analyses were conducted using the SAS 9.1 for Windows (SAS Institute).

DEP used in this study consist of a carbonaceous core to which organic chemicals such as polycyclic aromatic hydrocarbons (PAH), nitro derivatives of PAH, oxygenated derivatives of PAH (ketones, quinines, and diones), heterocyclic compounds, aldehydes, and aliphatic hydrocarbons are adsorbed (65, 6971). To determine any potential endotoxin contamination of the DEP used in this study, a quantitative kinetic chromogenic LAL method (KQCL) was employed (see 1Materials and Methods). Endotoxin content of culture media without and with DEP (100 μg/ml) was <0.05 EU/ml, indicating that no additional endotoxin was introduced into the PBMC cultures as a consequence of stimulation with DEP. To further rule out microbial contaminants in the DEP, complete culture media without and with DEP (100 μg/ml) were cultured for bacterial and fungal growth. No bacterial or fungal growth was detected in either of the samples after 1 wk and 1 mo, respectively.

M. tuberculosis is an intracellular bacterium that is taken up in monocytes, macrophages, and dendritic cells by phagocytosis, a prerequisite for the initiation of innate and subsequent adaptive host immune responses. Because human mononuclear phagocytes have been demonstrated to take up DEP (72), uptake of DEP and M. tuberculosis was assessed in enriched peripheral CD14+CD3 monocytes. Cytospin analysis of monocytes stimulated with DEP and M. tuberculosis (Fig. 1) provided initial evidence for a concurrent cellular uptake of DEP and M. tuberculosis into the boundaries of the cells. Subsequent studies by TEM confirmed presence of DEP and M. tuberculosis within the same transect of the monocytes (Fig. 2), implying the possibility of functional interactions of DEP and M. tuberculosis within the same cells.

FIGURE 1.

DEP and M. tuberculosis uptake in human blood monocytes. Magnetic bead-enriched CD14+CD3peripheral blood monocytes were stained with Kinyoun acid-fast stain on cytospins following overnight incubation with DEP (10 μg/ml) and M. tuberculosis (MOI 10). Five randomly selected monocytes that have incorporated both DEP and M. tuberculosis are shown. Arrows indicate the internalized DEP and Kinyoun-stained M. tuberculosis visible as black spots and red rods, respectively, at original magnification ×1000. Scale bars, 10 μM.

FIGURE 1.

DEP and M. tuberculosis uptake in human blood monocytes. Magnetic bead-enriched CD14+CD3peripheral blood monocytes were stained with Kinyoun acid-fast stain on cytospins following overnight incubation with DEP (10 μg/ml) and M. tuberculosis (MOI 10). Five randomly selected monocytes that have incorporated both DEP and M. tuberculosis are shown. Arrows indicate the internalized DEP and Kinyoun-stained M. tuberculosis visible as black spots and red rods, respectively, at original magnification ×1000. Scale bars, 10 μM.

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FIGURE 2.

TEM of DEP and M. tuberculosis uptake in human peripheral blood monocytes. The micrographs [(A and B) overviews of two independent cells; (C) details of the area surrounded by rectangle in (A); (D) intracellular compartment containing both DEP and three M. tuberculosis] show uptake of DEP (asterisks) and M. tuberculosis (arrows) within the same transects of monocytes, providing evidence of presence of DEP and M. tuberculosis within the same cell. Enriched peripheral blood CD14+CD3 monocytes were cultured with DEP (10 μg/ml) and M. tuberculosis (MOI 10) for 24 h and processed as described in 1Materials and Methods. Scale bars and direct magnifications are included in the figure.

FIGURE 2.

TEM of DEP and M. tuberculosis uptake in human peripheral blood monocytes. The micrographs [(A and B) overviews of two independent cells; (C) details of the area surrounded by rectangle in (A); (D) intracellular compartment containing both DEP and three M. tuberculosis] show uptake of DEP (asterisks) and M. tuberculosis (arrows) within the same transects of monocytes, providing evidence of presence of DEP and M. tuberculosis within the same cell. Enriched peripheral blood CD14+CD3 monocytes were cultured with DEP (10 μg/ml) and M. tuberculosis (MOI 10) for 24 h and processed as described in 1Materials and Methods. Scale bars and direct magnifications are included in the figure.

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To examine the extent of DEP-induced toxicity in PBMC, we first assessed apoptosis and necrosis using a flow cytometry-based Annexin V/PI apoptosis detection assay. PBMC from healthy subjects (n = 5) were exposed to DEP for 2, 24, and 72 h at final concentrations of 0.1–100 μg/ml and proportions of mononuclear cells undergoing combined apoptosis (Annexin V positive) and necrosis (PI positive) determined. Stimulation of PBMC with DEP for 2 h did not result in significant toxicity (combined apoptosis and necrosis) at any of the concentrations of DEP (0–100 μg/ml) tested. Furthermore, no significant cell toxicity with DEP stimulation in the dose range from 0.1–10 μg/ml relative to unstimulated cells was observed at any of the time points examined. Combined apoptosis and necrosis levels were increased only by 20–25% above the spontaneous level in PBMC exposed to DEP at a concentration of 100 μg/ml (Table I). It is worth noting that even at the highest concentration of DEP examined, ∼90, 70, and 60% of cells were viable at 2, 24, and 72 h, respectively. Based on these observations, we chose the DEP concentrations and PBMC stimulation durations for the subsequent ELISPOT assay and mRNA expression studies.

Table I.
Induction of apoptosis and necrosis by DEP in PBMC
Annexin V FITC/Propidium Iodide Positive (Mean %)
DEP Concentration (μg/ml)2 h24 h72 h
3.2 6.3 23.6 
0.1 4.3 7.2 24.0 
4.9 6.1 22.4 
10 5.4 6.8 24.9 
100 8.5 31.4 40.6 
Annexin V FITC/Propidium Iodide Positive (Mean %)
DEP Concentration (μg/ml)2 h24 h72 h
3.2 6.3 23.6 
0.1 4.3 7.2 24.0 
4.9 6.1 22.4 
10 5.4 6.8 24.9 
100 8.5 31.4 40.6 

Mean proportions of PBMC from healthy subjects (n = 5) showing positivity for Annexin V and PI by flowcytometry.

ELISPOT assays were used to assess frequencies of cytokine-producing PBMC as a measure of cytokine protein production. Representative images of cytokine-producing PBMC in developed wells are shown in Supplemental Fig. 1. To assess whether DEP alter antimycobacterial host immune responses, PBMC were stimulated with DEP alone or M. tuberculosis MOI 1, MOI 10, and PPD in presence of DEP at 0 (no DEP control), 1, 10, 50, and 100 μg/ml.

Stimulation of PBMC with DEP alone induced negligible amounts of IFN-γ (Fig. 3, leftmost column). Frequencies of IFN-γ–producing cells in PBMC stimulated with M. tuberculosis MOI 1 and MOI 10 were reduced significantly at DEP concentrations of ≥10 μg/ml relative to DEP 0 μg/ml. As expected, frequencies of PBMC producing all cytokines tested (IFN-γ, TNF-α, IL-1β, IL-6, and IL-10) were greater upon stimulation with M. tuberculosis MOI 10 than with MOI 1 (DEP 0) (Fig. 3, second and third columns from the left). Following PPD stimulation, IFN-γ production was significantly decreased at DEP concentrations of ≥50 μg/ml (Fig. 3, IFN-γ). Interestingly, suppressive effects of DEP were also observed on the per-cell IFN-γ output, as the size of the IFN-γ spots (diameters) decreased with increasing DEP concentrations (data not shown).

FIGURE 3.

Simultaneous stimulation with DEP alters pathogen-induced cytokine production. Frequencies of cytokine-producing PBMC (IFN-γ [n = 20 subjects], TNF-α [n = 18 subjects], IL-6 [n = 15 subjects], IL-10 [n = 20 subjects, data for DEP 50 μg/ml from n = 18 only], and IL-1β [n = 5 subjects]) stimulated with DEP alone (0, 1, 10, 50, and 100 μg/ml) or the stimuli M. tuberculosis (M.tb) MOI 1, MOI 10, and PPD (10 μg/ml) in presence (1, 10, 50, and 100 μg/ml) or absence (0 μg/ml) of DEP, were measured by ELISPOT assay. Cytokine data are arranged horizontally for each of the stimuli, which are arranged vertically. Frequencies of cytokine-producing cells (y-axes) are plotted as a function of DEP concentration in micrograms per milliliter (x-axes shown below the IL-1β panel only). Data are presented in box plots showing from top to bottom: the maximum value (black dot), the 95th (whisker), 75th (top of box plot), 50th (median, dotted line), mean (solid line in middle of box plot), 25th (solid line bottom of box plot), and 5th (whisker) percentiles and the minimum value (black dot). The p values are shown on top of box plots in which differences in cytokine expression between no DEP and different doses of DEP were regarded to be statistically significant (p ≤ 0.05). Additional statistical comparisons are shown in Supplemental Table I.

FIGURE 3.

Simultaneous stimulation with DEP alters pathogen-induced cytokine production. Frequencies of cytokine-producing PBMC (IFN-γ [n = 20 subjects], TNF-α [n = 18 subjects], IL-6 [n = 15 subjects], IL-10 [n = 20 subjects, data for DEP 50 μg/ml from n = 18 only], and IL-1β [n = 5 subjects]) stimulated with DEP alone (0, 1, 10, 50, and 100 μg/ml) or the stimuli M. tuberculosis (M.tb) MOI 1, MOI 10, and PPD (10 μg/ml) in presence (1, 10, 50, and 100 μg/ml) or absence (0 μg/ml) of DEP, were measured by ELISPOT assay. Cytokine data are arranged horizontally for each of the stimuli, which are arranged vertically. Frequencies of cytokine-producing cells (y-axes) are plotted as a function of DEP concentration in micrograms per milliliter (x-axes shown below the IL-1β panel only). Data are presented in box plots showing from top to bottom: the maximum value (black dot), the 95th (whisker), 75th (top of box plot), 50th (median, dotted line), mean (solid line in middle of box plot), 25th (solid line bottom of box plot), and 5th (whisker) percentiles and the minimum value (black dot). The p values are shown on top of box plots in which differences in cytokine expression between no DEP and different doses of DEP were regarded to be statistically significant (p ≤ 0.05). Additional statistical comparisons are shown in Supplemental Table I.

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TNF-α production was barely detectable upon stimulation of PBMC with DEP alone (Fig. 3, leftmost column). TNF-α production by PBMC stimulated with DEP at a concentration of ≥50 μg/ml was significantly lower (p ≤ 0.05) compared with that of unstimulated PBMC (culture media). TNF-α production upon stimulation with M. tuberculosis (MOI 1 and MOI 10) and PPD was significantly decreased at DEP concentrations of ≥50 and ≥10 μg/ml, respectively (Fig. 3, TNF-α panel).

As shown for IFN-γ and TNF-α, M. tuberculosis and PPD-induced IL-1β and IL-6 production was significantly decreased by DEP in a dose-dependent manner (Fig. 3, IL-1β and IL-6 panels). Unlike IFN-γ and TNF-α, however, stimulation of PBMC with DEP alone induced significant levels of cytokine production at concentrations of 1 and 10 μg/ml for IL-6 and 10 μg/ml for IL-1β compared with PBMC in culture media (Fig. 3, leftmost column). It is worth noting that the DEP-mediated suppression of cytokine production was most striking for IFN-γ and IL-1β, both of which are crucial for M. tuberculosis-induced host immune responses (21, 2830, 73).

We also investigated the effect of DEP on the production of the immunoregulatory cytokines IL-4 and IL-10, which have suppressive effects on innate and acquired immune responses to intracellular pathogens including M. tuberculosis (7477). In contrast to the findings with IFN-γ, TNF-α, IL-1β, and IL-6, IL-10 production in response to M. tuberculosis or PPD stimulation remained unchanged in presence of DEP concentrations up to 50 μg/ml relative to unstimulated PBMC (DEP 0) (Fig. 3, IL-10). No significant IL-10 production was detected when PBMC were stimulated with DEP alone (Fig. 3, leftmost column). No IL-4 production by PBMC was detected following stimulation with DEP, M. tuberculosis, or PPD (data not shown).

To examine whether DEP effects on M. tuberculosis-induced cytokine expression were similar to those induced by LPS, a TLR4 agonist, we determined frequencies of IFN-γ–, TNF-α–, IL-1β–, IL-6–, and IL-10–producing cells in PBMC stimulated with LPS in presence and absence of DEP. Effects of DEP on frequencies of IFN-γ–, TNF-α–, IL-1β–, IL-6–, and IL-10–producing PBMC following stimulation with LPS were comparable to effects of DEP following stimulation with M. tuberculosis or PPD (Fig. 4). Production of all cytokines was completely abrogated at the DEP concentration 100 μg/ml (Figs. 3, 4). It is worth noting that DEP-mediated apoptosis and necrosis of PBMC cannot account for the observed inhibition of cytokine production because 60–90% of PBMC were viable even after 72 h of incubation with DEP (Table I).

FIGURE 4.

Simultaneous stimulation with DEP alters LPS-induced cytokine production. Frequencies of IFN-γ–, TNF-α–, IL-6–, IL-10–, and IL-1β–producing PBMC (y-axis) represent spot frequencies following stimulation with LPS in presence or absence of indicated amounts of DEP (0, 1, 10, 50, and 100 μg/ml) were measured by ELISPOT assay. Subject numbers, incubation time, and cell numbers for each cytokine were identical to that stated in legend to Fig. 3. Presentation of data in the box plots is described in Fig. 3. Additional statistical comparisons are shown in Supplemental Table I.

FIGURE 4.

Simultaneous stimulation with DEP alters LPS-induced cytokine production. Frequencies of IFN-γ–, TNF-α–, IL-6–, IL-10–, and IL-1β–producing PBMC (y-axis) represent spot frequencies following stimulation with LPS in presence or absence of indicated amounts of DEP (0, 1, 10, 50, and 100 μg/ml) were measured by ELISPOT assay. Subject numbers, incubation time, and cell numbers for each cytokine were identical to that stated in legend to Fig. 3. Presentation of data in the box plots is described in Fig. 3. Additional statistical comparisons are shown in Supplemental Table I.

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Taken together, these results indicate that stimulation of PBMC with DEP inhibited M. tuberculosis, PPD, and LPS-induced production of IFN-γ, TNF-α, IL-1β, and IL-6 in a concentration-dependent manner. On the contrary, IL-10 production remained unchanged at all but the highest concentration of DEP examined. The observed suppression of M. tuberculosis-induced production of proinflammatory cytokines together with the concomitantly unaltered IL-10 production strongly suggest that DEP interfere with protective host antimycobacterial immunity.

To determine whether the observed effects of DEP were due to its particle nature or the adsorbed surface chemical species (65, 6971), we assessed the effects of CB as a chemically inert particle control on M. tuberculosis-induced production of IFN-γ, TNF-α, and IL-10, in parallel with DEP. CB represents the chemically inert inner carbon core of DEP (78) and is devoid of surface chemical species. In contrast to the findings following DEP exposure, M. tuberculosis- or PPD-induced production of IFN-γ, TNF-α, and IL-10 did not change with increasing CB concentrations (0, 1, 10, and 50 μg/ml) (Fig. 5). Taken together, these results suggest that the alterations in cytokine production observed in PBMC stimulated with DEP (Figs. 3, 4) are in large part due to adsorbed surface chemical species (see 18Characterization of DEP) rather than the inert carbon core of DEP.

FIGURE 5.

Comparison of DEP and CB effects on M. tuberculosis- (M.tb) and PPD-induced cytokine production. PBMC from healthy subjects (n = 3) were stimulated with M. tuberculosis at MOI 10 and PPD (10 μg/ml) in presence of varying doses of DEP or CB. DEP or CB was added at final concentrations of 0 (DEP0, CB0, negative control), 1, 10, and 50 μg/ml. Numbers of IFN-γ–, IL-10–, and TNF-α–producing PBMC were enumerated by ELISPOT assay, and frequencies of cytokine-producing cells (y-axes) are plotted as a function of DEP concentration in micrograms per milliliter (x-axes shown below the TNF-α panel only). Mean frequencies ± SEM are shown.

FIGURE 5.

Comparison of DEP and CB effects on M. tuberculosis- (M.tb) and PPD-induced cytokine production. PBMC from healthy subjects (n = 3) were stimulated with M. tuberculosis at MOI 10 and PPD (10 μg/ml) in presence of varying doses of DEP or CB. DEP or CB was added at final concentrations of 0 (DEP0, CB0, negative control), 1, 10, and 50 μg/ml. Numbers of IFN-γ–, IL-10–, and TNF-α–producing PBMC were enumerated by ELISPOT assay, and frequencies of cytokine-producing cells (y-axes) are plotted as a function of DEP concentration in micrograms per milliliter (x-axes shown below the TNF-α panel only). Mean frequencies ± SEM are shown.

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A broad spectrum of gene targets was studied using Th1-Th2-Th3 and TLR pathway-specific gene expression qRT-PCR arrays to determine the effect of DEP on M. tuberculosis-induced host gene expression.

M. tuberculosis-induced mRNA expression.

In a first step, mRNA expression profiles of PBMC stimulated with M. tuberculosis MOI 10 were compared with that of unstimulated PBMC (cultured in complete medium). Levels of M. tuberculosis-induced mRNAs (increased or decreased by ≥2-fold; p ≤ 0.01) relative to unstimulated cells were evaluated with Th1-Th2-Th3 (Fig. 6A, 6B) and TLR pathway-specific (Fig. 6C, 6D) qRT-PCR arrays. Using human Th1-Th2-Th3 qRT-PCR arrays, a significant increase (100–1000-fold) of mRNAs encoding Th1 cytokines IFNG, IL12B, and CSF2 relative to untreated controls was observed (Fig. 6A). In addition, the level of mRNA encoding Th1 cytokines and related genes IL12RB2, IL2RA, IRF1, IL18R1, SOCS1, STAT4, TBX21, and TNFA was increased by 2–50-fold upon M. tuberculosis stimulation (Fig. 6A). The expression of a smaller number of Th2 cytokines and related genes (e.g., CCL7, IL10, IL13, IL5, ICOS, and NFATC2) was increased by 2–50-fold in M. tuberculosis-stimulated cells relative to unstimulated PBMC. Increased expression (2- to >2,000-fold) of genes encoding CD4+ T cell markers CD40LG, CTLA4, FASG, IL6, IL7, LAG3, TNFRSF8, TNFRSF9, and TNFRSF4 was observed in M. tuberculosis-stimulated PBMC (Fig. 6A), whereas IL18, TLR4, IL13RA1, IL1R2, CD4, and CD86 genes were downregulated by 2–30-fold following M. tuberculosis stimulation (Fig. 6B).

FIGURE 6.

M. tuberculosis (M.tb)-induced mRNA expression. Upregulation (A, C) and downregulation (B, D) of mRNA levels in M. tuberculosis MOI 10-stimulated PBMC relative to that of untreated PBMC from healthy subjects using Th1-Th2-Th3 (n = 11) and TLR signaling (n = 8) pathway-specific qRT-PCR arrays are shown. y-axes represent statistically significant (p ≤ 0.01) mean fold changes (≥2-fold) ± SEM in the mRNA expression level of M. tuberculosis-induced genes compared with that in untreated PBMC. No M. tuberculosis-induced fold change relative to untreated (defined by <2-fold; p > 0.01) was set as 1 in (A) and (C) and as 0 in (B) and (D), in which logarithmic and linear scales were used, respectively.

FIGURE 6.

M. tuberculosis (M.tb)-induced mRNA expression. Upregulation (A, C) and downregulation (B, D) of mRNA levels in M. tuberculosis MOI 10-stimulated PBMC relative to that of untreated PBMC from healthy subjects using Th1-Th2-Th3 (n = 11) and TLR signaling (n = 8) pathway-specific qRT-PCR arrays are shown. y-axes represent statistically significant (p ≤ 0.01) mean fold changes (≥2-fold) ± SEM in the mRNA expression level of M. tuberculosis-induced genes compared with that in untreated PBMC. No M. tuberculosis-induced fold change relative to untreated (defined by <2-fold; p > 0.01) was set as 1 in (A) and (C) and as 0 in (B) and (D), in which logarithmic and linear scales were used, respectively.

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Using human TLR signaling pathway-specific qRT-PCR arrays, upregulation of NF-κB target genes CCL2, CSF2, CSF3, IFNA1, IFNB1, IFNG, IL1A, IL1B, IL6, IL8, IL10, TNF, NFKB1, and NFKB1A and of genes downstream of IRF pathway, CXCL10, IFNA1, IFNB1, IFNG, and IRF1, were observed in M. tuberculosis-stimulated PBMC relative to unstimulated PBMC (Fig. 6C). In addition to M. tuberculosis-mediated upregulation (Fig. 6C), downregulation of several genes related to TLR-mediated signal transduction pathway by M. tuberculosis was observed (Fig. 6D).

DEP-mediated modification of M. tuberculosis-induced mRNA expression.

Engagement of TLRs by M. tuberculosis ligands initiates signaling pathways that ultimately activate transcription factor NF-κB, leading to subsequent release of cytokines and chemokines (38, 41, 42, 73). The effects of DEP were studied upon simultaneous stimulation of PBMC with DEP and M. tuberculosis and upon a 20-h prestimulation of PBMC with DEP followed by stimulation with M. tuberculosis. The prestimulation with DEP simulated the sequential exposures to DEP and M. tuberculosis that may occur in real life. Thus, PBMC were stimulated with DEP, M. tuberculosis plus DEP (simultaneous), M. tuberculosis following DEP (prestimulation), and M. tuberculosis alone or left unstimulated. The mRNA expression profile of PBMC that had been stimulated with M. tuberculosis following DEP pre-exposure was considerably different from that of PBMC stimulated with M. tuberculosis plus DEP simultaneously (compare Fig. 7B, 7C and Fig. 8B, 8C, Table II). The mRNA fold changes plotted on y-axes in Figs. 7B, 7C, 8B, and 8C represent a ratio of the mRNA levels of M. tuberculosis plus DEP and M. tuberculosis alone. Thus, all DEP-mediated mRNA expression changes are represented relative to the mRNA expression levels in PBMC stimulated with M. tuberculosis alone (set as baseline). IFNG, TLR4, and CXCL10 mRNAs decreased to comparable levels during both the simultaneous stimulation (Figs. 7B, 8B, Table II) and the 20-h prestimulation with DEP (Figs. 7C, 8C, Table II). DEP prestimulation followed by M. tuberculosis infection, however, led to the downregulation of a larger number of M. tuberculosis-induced genes with known involvement in antimycobacterial immunity such as costimulatory molecules, type I and type II IFNs, proinflammatory cytokines, chemokines, chemokine receptors, and TLRs 3, 4, 7, and 10. In addition to the downregulation of the genes mentioned above, stimulation of PBMC with DEP upregulated the expression of several M. tuberculosis-induced genes including CSF2, IL1R1 during simultaneous, and CD86, CEBPB, IGSF6, IL13RA1, and IL18 during prestimulation by 2–4-fold (p ≤ 0.01) compared with PBMC stimulated with M. tuberculosis alone (Figs. 7B, 7C, 8B, 8C, Table II). Thus, prestimulation with DEP followed by M. tuberculosis stimulation alters M. tuberculosis-induced host responses more profoundly than simultaneous stimulation with DEP and M. tuberculosis as evidenced by the alteration of 4 and 5 mRNAs in Figs. 7B and 8B, respectively, compared with 23 and 17 mRNAs in Figs. 7C and 8C, respectively. Stimulation of PBMC with DEP alone altered the expression of only a small number of mRNAs (Figs. 7A, 8A). The expression of proinflammatory mediators including IL1R1, IL8, PTGS2, and TNF was increased, whereas the expression of the receptors CD4 and CD80 (B7-1) was decreased (Figs. 7A, 8A).

FIGURE 7.

DEP effects on M. tuberculosis (M.tb)-induced Th1-Th2-Th3 pathway-specific mRNA expression. Depicted on y-axes are statistically significant (p ≤ 0.01) mean fold changes (≥2-fold) ± SEM in mRNA expression levels of in vitro-stimulated PBMC from healthy subjects (n = 11). (A) The DEP-mediated (10 μg/ml) mRNA expression in PBMC relative to mRNA expression levels of unstimulated PBMC in culture medium (set as 0-line). (B) mRNA expression levels from simultaneous DEP (10 μg/ml) and M. tuberculosis (MOI 10) stimulation of PBMC compared with mRNA expression levels from PBMC stimulated with M. tuberculosis alone (set as 0-line). (C) The effect of DEP prestimulation on M. tuberculosis-induced mRNA expression. PBMC were prestimulated with DEP (10 μg/ml) for 20 h and subsequently stimulated with M. tuberculosis at MOI 10 for an additional 24 h. The mRNA expression levels of DEP prestimulated and subsequently M. tuberculosis-stimulated PBMC were compared with that of PBMC stimulated with M. tuberculosis alone (set as 0-line).

FIGURE 7.

DEP effects on M. tuberculosis (M.tb)-induced Th1-Th2-Th3 pathway-specific mRNA expression. Depicted on y-axes are statistically significant (p ≤ 0.01) mean fold changes (≥2-fold) ± SEM in mRNA expression levels of in vitro-stimulated PBMC from healthy subjects (n = 11). (A) The DEP-mediated (10 μg/ml) mRNA expression in PBMC relative to mRNA expression levels of unstimulated PBMC in culture medium (set as 0-line). (B) mRNA expression levels from simultaneous DEP (10 μg/ml) and M. tuberculosis (MOI 10) stimulation of PBMC compared with mRNA expression levels from PBMC stimulated with M. tuberculosis alone (set as 0-line). (C) The effect of DEP prestimulation on M. tuberculosis-induced mRNA expression. PBMC were prestimulated with DEP (10 μg/ml) for 20 h and subsequently stimulated with M. tuberculosis at MOI 10 for an additional 24 h. The mRNA expression levels of DEP prestimulated and subsequently M. tuberculosis-stimulated PBMC were compared with that of PBMC stimulated with M. tuberculosis alone (set as 0-line).

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FIGURE 8.

DEP Effects on TLR pathway-specific mRNA expression. Depicted in y-axes are statistically significant (p ≤ 0.01) mean fold changes (≥2-fold) ± SEM in mRNA expression levels of in vitro-stimulated PBMC from healthy subjects (n = 8). (A) DEP-mediated alteration of mRNA expression in PBMC. DEP-induced mRNA expression levels were compared with that of unstimulated PBMC in culture medium (set as 0-line). (B) The effect of DEP (10 μg/ml final concentration) on M. tuberculosis (M.tb) MOI 10-induced mRNA expression during simultaneous combined stimulation compared with mRNA expression levels from PBMC that were stimulated with M. tuberculosis MOI 10 alone (set as 0-line). (C) The effect of DEP prestimulation on M. tuberculosis MOI 10-induced mRNA expression. PBMC were prestimulated with DEP (10 μg/ml) for 20 h and subsequently stimulated with M. tuberculosis at MOI 10 for an additional 24 h. The mRNA expression levels of DEP prestimulated and subsequently M. tuberculosis-stimulated PBMC were compared with that of PBMC stimulated with M. tuberculosis alone (set as 0-line).

FIGURE 8.

DEP Effects on TLR pathway-specific mRNA expression. Depicted in y-axes are statistically significant (p ≤ 0.01) mean fold changes (≥2-fold) ± SEM in mRNA expression levels of in vitro-stimulated PBMC from healthy subjects (n = 8). (A) DEP-mediated alteration of mRNA expression in PBMC. DEP-induced mRNA expression levels were compared with that of unstimulated PBMC in culture medium (set as 0-line). (B) The effect of DEP (10 μg/ml final concentration) on M. tuberculosis (M.tb) MOI 10-induced mRNA expression during simultaneous combined stimulation compared with mRNA expression levels from PBMC that were stimulated with M. tuberculosis MOI 10 alone (set as 0-line). (C) The effect of DEP prestimulation on M. tuberculosis MOI 10-induced mRNA expression. PBMC were prestimulated with DEP (10 μg/ml) for 20 h and subsequently stimulated with M. tuberculosis at MOI 10 for an additional 24 h. The mRNA expression levels of DEP prestimulated and subsequently M. tuberculosis-stimulated PBMC were compared with that of PBMC stimulated with M. tuberculosis alone (set as 0-line).

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Table II.
Effect of DEP exposure on M. tuberculosis-induced host gene expression
Functional GroupGenesDEP Stimulation ConditionFold Change Relative to M. tuberculosis
Cytokines and cytokine receptors IL5 Pre ↓ 9 
 IL6 Pre ↓ 8.9-9.5 
 IFNG Simultaneous and Pre ↓ 5–5.4 and 6.1–7.3 
 IFNA1 Pre ↓ 6.4 
 IFNB1 Pre ↓ 4.7 
 IL1A Pre ↓ 4 
 CSF3 Pre ↓ 3.5 
 IL18 Pre ↑ 3.2 
 CSF2 Simultaneous ↑ 2.4–2.7 
 IL7 Pre ↓ 2.2 
 IL2RA Pre ↓ 2.3 
 IL18R1 Pre ↓ 2.2 
 IL1R1 Simultaneous ↑ 2.2 
 IL13RA Pre ↑ 2.1 
Chemokines and receptors CXCL10 Simultaneous and Pre ↓ 3.48 and 5.3 
 CXCR3 Pre ↓ 3 
 CCL5 Pre ↓ 2.4 
 CCL7 Simultaneous ↓ 2.3 
 CCR4 Pre ↓ 2.1 
Transcription factors and inhibitors NFKB1A Pre ↓ 9.6 
 SOCS1 Pre ↓ 2.6 
 FOS Pre ↑ 2.6 
 GATA3 Pre ↓ 2 
 CEBPB Pre ↑ 2 
TLR TLR7 Pre ↓ 4.7 
 TLA4 Simultaneous and Pre ↓ 2.3 and 3.8 
 TLR3 Pre ↓ 3.1 
 TLR10 Pre ↓ 2.4 
 TLR5 Simultaneous ↑ 2.1 
Costimulatory molecules, T cell activation CD86 Pre ↑ 2.8–3.5 
 CD80 Pre ↓ 2.4–2.7 
 LAG3 Pre ↓ 2.7 
 CTLA4 Pre ↓ 2.6 
 ICOS Pre ↓ 2.3 
 TNFSF4 Pre ↓ 2.2 
IFN-responsive effector EIF2AK2 Pre ↓ 5.6 
 SPP1 Pre ↑ 2.1 
Others FASLG Pre ↓ 3.0 
 PTGS2 Pre ↓ 3.6 
 IGSF6 Pre ↑ 3.9 
Functional GroupGenesDEP Stimulation ConditionFold Change Relative to M. tuberculosis
Cytokines and cytokine receptors IL5 Pre ↓ 9 
 IL6 Pre ↓ 8.9-9.5 
 IFNG Simultaneous and Pre ↓ 5–5.4 and 6.1–7.3 
 IFNA1 Pre ↓ 6.4 
 IFNB1 Pre ↓ 4.7 
 IL1A Pre ↓ 4 
 CSF3 Pre ↓ 3.5 
 IL18 Pre ↑ 3.2 
 CSF2 Simultaneous ↑ 2.4–2.7 
 IL7 Pre ↓ 2.2 
 IL2RA Pre ↓ 2.3 
 IL18R1 Pre ↓ 2.2 
 IL1R1 Simultaneous ↑ 2.2 
 IL13RA Pre ↑ 2.1 
Chemokines and receptors CXCL10 Simultaneous and Pre ↓ 3.48 and 5.3 
 CXCR3 Pre ↓ 3 
 CCL5 Pre ↓ 2.4 
 CCL7 Simultaneous ↓ 2.3 
 CCR4 Pre ↓ 2.1 
Transcription factors and inhibitors NFKB1A Pre ↓ 9.6 
 SOCS1 Pre ↓ 2.6 
 FOS Pre ↑ 2.6 
 GATA3 Pre ↓ 2 
 CEBPB Pre ↑ 2 
TLR TLR7 Pre ↓ 4.7 
 TLA4 Simultaneous and Pre ↓ 2.3 and 3.8 
 TLR3 Pre ↓ 3.1 
 TLR10 Pre ↓ 2.4 
 TLR5 Simultaneous ↑ 2.1 
Costimulatory molecules, T cell activation CD86 Pre ↑ 2.8–3.5 
 CD80 Pre ↓ 2.4–2.7 
 LAG3 Pre ↓ 2.7 
 CTLA4 Pre ↓ 2.6 
 ICOS Pre ↓ 2.3 
 TNFSF4 Pre ↓ 2.2 
IFN-responsive effector EIF2AK2 Pre ↓ 5.6 
 SPP1 Pre ↑ 2.1 
Others FASLG Pre ↓ 3.0 
 PTGS2 Pre ↓ 3.6 
 IGSF6 Pre ↑ 3.9 

The mRNA levels of PBMC stimulated simultaneously with M. tuberculosis and DEP (simultaneous) and with M. tuberculosis following DEP prestimulation (Pre) were compared with that of PBMC stimulated with M. tuberculosis alone. mRNA expression levels that were increased or decreased by ≥2-fold (p ≤ 0.01) relative to the mRNA expression levels of PBMC stimulated with M. tuberculosis alone are shown.

The gene expression analyses with qRT-PCR arrays were validated by real-time PCR of cDNAs generated from a subset of RNA samples in the qRT-PCR arrays using primers listed in Fig. 9. The expression of eight genes altered by simultaneous and/or prestimulation with DEP is shown in Fig. 9. In general, real-time PCR data were consistent with the data obtained from the qRT-PCR arrays.

FIGURE 9.

Validation of qRT-PCR array results. Total RNA (400 ng) from PBMC of a subset of donors (n = 4–6) was reverse transcribed to generate cDNA. Real-time PCR was performed and mRNA fold change in DEP-prestimulated and M. tuberculosis (M.tb) MOI 10-exposed PBMC relative to M. tuberculosis was evaluated. Means ± SEM are shown. IFNG: forward primer, 5′-GCATCCAAAAGAGTGTGGAGAC-3′, reverse primer, 5′-TACTGGGATGCTCTTCGACCT-3′; CXCL10: forward primer, 5′-TGGCATTCAAGGAGTACCTCTC-3′, reverse primer, 5′-CTTGATGGCCTTCGATTCTG-3′; CD86: forward primer, 5′-GGTGCTGCTCCTCTGAAGATT-3′, reverse primer, 5′-TGA AGTCTCAGGGTCCAACTG-3′; IL13RA: forward primer, 5′-GTCCCAGTGTAGCACCAATGA-3′, reverse primer, 5′-CCAGGCTTCTGTGCCAATAGT-3′; IL6: forward primer, 5′-AGACAGCCACTCACCTCTTCA-3′, reverse primer, 5′-CACCAGGCAAGTCTCCTCATT-3′; CSF3: forward primer, 5′-ATAGCGGCCTTTTCCT CTACC-3′, reverse primer, 5′-GCCATTCCCAGTTCTTCCAT-3′; PTGS2: forward primer, 5′-GGTGATGAGCAGTTGTTCCAG-3′, reverse primer, 5′-GAAGGGGATGCCAGTGATAGA-3′; and LAG3: forward primer, 5′-TCCTGGTGACTGGAGACAATG-3′, reverse primer, 5′-TGGAGTCACCTCACAAAGCAG-3′.

FIGURE 9.

Validation of qRT-PCR array results. Total RNA (400 ng) from PBMC of a subset of donors (n = 4–6) was reverse transcribed to generate cDNA. Real-time PCR was performed and mRNA fold change in DEP-prestimulated and M. tuberculosis (M.tb) MOI 10-exposed PBMC relative to M. tuberculosis was evaluated. Means ± SEM are shown. IFNG: forward primer, 5′-GCATCCAAAAGAGTGTGGAGAC-3′, reverse primer, 5′-TACTGGGATGCTCTTCGACCT-3′; CXCL10: forward primer, 5′-TGGCATTCAAGGAGTACCTCTC-3′, reverse primer, 5′-CTTGATGGCCTTCGATTCTG-3′; CD86: forward primer, 5′-GGTGCTGCTCCTCTGAAGATT-3′, reverse primer, 5′-TGA AGTCTCAGGGTCCAACTG-3′; IL13RA: forward primer, 5′-GTCCCAGTGTAGCACCAATGA-3′, reverse primer, 5′-CCAGGCTTCTGTGCCAATAGT-3′; IL6: forward primer, 5′-AGACAGCCACTCACCTCTTCA-3′, reverse primer, 5′-CACCAGGCAAGTCTCCTCATT-3′; CSF3: forward primer, 5′-ATAGCGGCCTTTTCCT CTACC-3′, reverse primer, 5′-GCCATTCCCAGTTCTTCCAT-3′; PTGS2: forward primer, 5′-GGTGATGAGCAGTTGTTCCAG-3′, reverse primer, 5′-GAAGGGGATGCCAGTGATAGA-3′; and LAG3: forward primer, 5′-TCCTGGTGACTGGAGACAATG-3′, reverse primer, 5′-TGGAGTCACCTCACAAAGCAG-3′.

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Simultaneous and prestimulation with DEP were found to downregulate the expression of a large number of genes including that of TLRs (Fig. 8C). Considering the importance of TLR-mediated antimycobacterial immunity, DEP-induced suppression of TLR pathways could potentially inhibit robust host immune responses against microbial infection.

DEP prestimulation modified M. tuberculosis-induced cytokine responses in a manner comparable to that shown in other studies following repeated LPS stimulation of cells (40, 7981). Such LPS stimulation induces a state of cellular tolerance of monocytes/macrophages to subsequent secondary stimulation, which is characterized by impairment of IFN-γ, TNF-α, IL-1β, and IL-6 and preservation of anti-inflammatory IL-10 production (80). Stimulation with LPS, a potent ligand of TLR4 and CD14, leads to activation of the NF-ĸB pathway via MyD88-dependent and MyD88-independent signaling (47, 82). To examine whether LPS prestimulation suppresses M. tuberculosis-induced gene expression, the gene expression profile of PBMC infected with M. tuberculosis following prestimulation with LPS was compared with that of M. tuberculosis-infected PBMC without LPS prestimulation using Th1-Th2-Th3 (Fig. 10A) and TLR pathway-specific (Fig. 10B) qRT-PCR arrays. Prestimulation with LPS, like with DEP, suppressed several M. tuberculosis-induced genes encoding host immune effector proteins including IFN-γ and CXCL10, which are crucial for protective antimycobacterial immunity. However, unlike with DEP, LPS prestimulation inhibited the M. tuberculosis-induced expression of IL12B, IL12RB2, and IRF4 (Fig. 10, Table III). As seen in Table III, prestimulation with DEP suppressed a broader spectrum of M. tuberculosis-induced host immune genes in PBMC than prestimulation with LPS. This is not surprising considering that LPS prestimulation suppresses predominantly TLR4 signaling, whereas DEP may potentially suppress signaling through other TLRs involved in M. tuberculosis-induced host responses. Thus, prestimulation with DEP, like with LPS, induced a state of cellular hyporesponsiveness toward subsequent stimulation with M. tuberculosis.

FIGURE 10.

Effect of LPS prestimulation on M. tuberculosis (M.tb)-induced mRNA expression. Depicted in y-axes are statistically significant (p ≤ 0.01) mean fold changes (≥2-fold) ± SEM in mRNA expression levels of in vitro-stimulated PBMC from healthy subjects (n = 11 for Th1-Th2-Th3, n = 8 for TLR pathway-specific qRT-PCR arrays). PBMC were prestimulated with LPS (100 ng/ml) for 20 h and subsequently stimulated with M. tuberculosis at MOI 10 for an additional 24 h. Following RNA extraction and generation of cDNA, gene expression was assessed by qRT-PCR arrays. mRNA expression levels were compared with that from PBMC stimulated with M. tuberculosis alone (set as 0-line). (A) Effects of LPS prestimulation on M. tuberculosis-induced Th1-Th2-Th3 pathway-specific mRNA expression. (B) Effects of LPS prestimulation on M. tuberculosis-induced TLR pathway-specific mRNA expression.

FIGURE 10.

Effect of LPS prestimulation on M. tuberculosis (M.tb)-induced mRNA expression. Depicted in y-axes are statistically significant (p ≤ 0.01) mean fold changes (≥2-fold) ± SEM in mRNA expression levels of in vitro-stimulated PBMC from healthy subjects (n = 11 for Th1-Th2-Th3, n = 8 for TLR pathway-specific qRT-PCR arrays). PBMC were prestimulated with LPS (100 ng/ml) for 20 h and subsequently stimulated with M. tuberculosis at MOI 10 for an additional 24 h. Following RNA extraction and generation of cDNA, gene expression was assessed by qRT-PCR arrays. mRNA expression levels were compared with that from PBMC stimulated with M. tuberculosis alone (set as 0-line). (A) Effects of LPS prestimulation on M. tuberculosis-induced Th1-Th2-Th3 pathway-specific mRNA expression. (B) Effects of LPS prestimulation on M. tuberculosis-induced TLR pathway-specific mRNA expression.

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Table III.
Alteration of M. tuberculosis-induced genes by DEP and LPS
M. tuberculosis-Induced Genes Altered by DEP Prestimulation (Relative to M. tuberculosis)M. tuberculosis-Induced Genes Altered by LPS Prestimulation (Relative to M. tuberculosis)M. tuberculosis-Induced Genes Not Altered by DEP Prestimulation (Relative to M. tuberculosis)M. tuberculosis-Induced Genes Not Altered by LPS Prestimulation (Relative to M. tuberculosis)
CCL5 ↓ CCL7 ↓ CCL2 CCL2 
CCR4 ↓ CLEC4E ↑ CCL7 CCL5 
CD86 ↑ CSF2 ↓ CD180 CCR4 
CEBPB ↑ HAVCR2 ↓ CD40LG CD180 
CSF3 ↓ IL1R1 ↑ CLEC4E CD40LG 
CTLA4 ↓ IL12B ↓ CSF2 CD86 
CXCR3 ↓ IL12RB2 ↓ ELK1 CEBPB 
FOS ↑ INHBA ↓ HAVCR2 CSF3 
GATA3 ↓ IRF4 ↓ IL10 CTLA4 
ICOS ↓ TBX21 ↓ IL12RB2 CXCR3 
IFNA1 ↓ TNF ↓ IL13 ELK1 
IGSF6 ↑ TNFRSF9 ↓ IL1R2 FOS 
IL13RA ↑ CD80 IL23A GATA3 
IL18 ↑ CXCL10 IL8 ICOS 
IL1A ↓ EIF2AK2 INHBA IFNA1 
IL5 ↓ FASLG ↓ IRAK2 IGSF6 
IL6 ↓ IFNB1 IRF1 IL10 
IL7 ↓ IFNG IRF4 IL13 
NFkB1A ↓ IL18R1 JUN IL13RA 
PTGS2 ↓ IL2RA LTA IL18 
SOCS1 ↓ LAG3 ↓ MAP4K4 IL1A 
SSP1 ↑ TLR3 NFKB1 IL23A 
TLR10 ↓  RIPK2 IL6 
TLR4 ↓  SIGIRR IL7 
TLR7 ↓  SOCS2 IL8 
TNFSF4 ↓  STAT4 IRAK2 
CD80  TBX21 IRF1 
CXCL10  TLR1 JUN 
EIF2AK2  TLR5 LTA 
FASLG ↓  TLR8 MAP4K4 
IFNB1  IL12B NFKB1 
IFNG  TNF NFKB1A 
IL18R1  TNFRSF8 PTGS2 
IL2RA  TNFRSF9 RIPK2 
LAG3 ↓  UBE2V1 SIGIRR 
TLR3   SOCS1 
   SOCS2 
   SSP1 
   STAT4 
   TLR1 
   TLR10 
   TLR4 
   TLR5 
   TLR7 
   TLR8 
   IL5 
   TNFRSF8 
   TNFSF4 
   UBE2V1 
M. tuberculosis-Induced Genes Altered by DEP Prestimulation (Relative to M. tuberculosis)M. tuberculosis-Induced Genes Altered by LPS Prestimulation (Relative to M. tuberculosis)M. tuberculosis-Induced Genes Not Altered by DEP Prestimulation (Relative to M. tuberculosis)M. tuberculosis-Induced Genes Not Altered by LPS Prestimulation (Relative to M. tuberculosis)
CCL5 ↓ CCL7 ↓ CCL2 CCL2 
CCR4 ↓ CLEC4E ↑ CCL7 CCL5 
CD86 ↑ CSF2 ↓ CD180 CCR4 
CEBPB ↑ HAVCR2 ↓ CD40LG CD180 
CSF3 ↓ IL1R1 ↑ CLEC4E CD40LG 
CTLA4 ↓ IL12B ↓ CSF2 CD86 
CXCR3 ↓ IL12RB2 ↓ ELK1 CEBPB 
FOS ↑ INHBA ↓ HAVCR2 CSF3 
GATA3 ↓ IRF4 ↓ IL10 CTLA4 
ICOS ↓ TBX21 ↓ IL12RB2 CXCR3 
IFNA1 ↓ TNF ↓ IL13 ELK1 
IGSF6 ↑ TNFRSF9 ↓ IL1R2 FOS 
IL13RA ↑ CD80 IL23A GATA3 
IL18 ↑ CXCL10 IL8 ICOS 
IL1A ↓ EIF2AK2 INHBA IFNA1 
IL5 ↓ FASLG ↓ IRAK2 IGSF6 
IL6 ↓ IFNB1 IRF1 IL10 
IL7 ↓ IFNG IRF4 IL13 
NFkB1A ↓ IL18R1 JUN IL13RA 
PTGS2 ↓ IL2RA LTA IL18 
SOCS1 ↓ LAG3 ↓ MAP4K4 IL1A 
SSP1 ↑ TLR3 NFKB1 IL23A 
TLR10 ↓  RIPK2 IL6 
TLR4 ↓  SIGIRR IL7 
TLR7 ↓  SOCS2 IL8 
TNFSF4 ↓  STAT4 IRAK2 
CD80  TBX21 IRF1 
CXCL10  TLR1 JUN 
EIF2AK2  TLR5 LTA 
FASLG ↓  TLR8 MAP4K4 
IFNB1  IL12B NFKB1 
IFNG  TNF NFKB1A 
IL18R1  TNFRSF8 PTGS2 
IL2RA  TNFRSF9 RIPK2 
LAG3 ↓  UBE2V1 SIGIRR 
TLR3   SOCS1 
   SOCS2 
   SSP1 
   STAT4 
   TLR1 
   TLR10 
   TLR4 
   TLR5 
   TLR7 
   TLR8 
   IL5 
   TNFRSF8 
   TNFSF4 
   UBE2V1 

The mRNA levels of PBMC stimulated with M. tuberculosis following DEP or LPS prestimulation were compared with that of PBMC stimulated with M. tuberculosis alone. Increased or decreased (≥2-fold; p ≤ 0.01) mRNA expression levels relative to mRNA expression levels of M. tuberculosis-stimulated PBMC (set as 0-value) are shown. mRNAs altered by both DEP and LPS (columns 1 and 2) are in boldface.

It is worth noting that the mRNA expression profile of the M. tuberculosis-induced host immune response in presence of DEP stimulation compared with that in the presence of M. tuberculosis alone was consistent with the reduced IFN-γ, IL-1β, and IL-6 and increased or unchanged IL-10 protein expression in the ELISPOT assays (compare Figs. 7B, 7C, 8B, 8C with Fig. 3). DEP had no effect on M. tuberculosis-induced TNF-α mRNA expression (measured 24 h after stimulation), whereas DEP decreased M. tuberculosis-induced TNF-α protein expression (Fig. 3). This discrepancy in gene and protein expression is likely due to the transient and early expression of mRNA encoding TNF-α. Taken together, these cytokine production and mRNA expression data suggest that DEP exposure modulates and interferes (see hypothetical model in Fig. 11) with antimycobacterial immune responses.

TB remains a major global public health concern that is aggravated by the HIV pandemic and emergence of drug-resistant M. tuberculosis strains. Deteriorating air quality from rapid industrial growth and urban traffic coincide with high prevalence and incidence rates of endemic TB in densely populated regions of low- and middle-income countries. Although epidemiological evidence suggests that exposure to air pollutants (e.g., silica, indoor pollution from fossil fuel combustion, cigarette smoke) increase the incidence of TB, only a few studies have assessed mechanisms that may underlie the effects of urban air PM2.5 on innate and adaptive human immune responses to mycobacteria (62, 72, 83).

Because concomitant aerosol exposure to DEP and M. tuberculosis occur frequently in real-life situations, we hypothesized that DEP may alter human antimycobacterial host immunity. This study is the first, to our knowledge, to assess the effects of DEP, a major component of urban air pollution, on human primary immune cell responses to M. tuberculosis. We focused on the effects of DEP on M. tuberculosis-specific cytokine production and gene expression in PBMC, which, unlike studies in isolated primary cell populations or cell lines (61, 84,,,,,,91), permit evaluation of immunological interactions between APCs and lymphocytes in a quasi-physiological context. The current study demonstrates that DEP and M. tuberculosis localize within the same monocytes (Figs 1, 2), a finding that is consistent with a report of combined uptake of bacillus Calmette-Guérin (BCG) and DEP by human monocyte-derived macrophages (72). Because combined uptake of M. tuberculosis and DEP in host immune cells suggests the possibility of subsequent alterations of cellular functions, we studied the DEP effects on TLR-mediated proinflammatory immune responses and adaptive immunity (92).

Our studies show that DEP suppress M. tuberculosis, PPD, and LPS-induced IFN-γ, IL-6, IL-1β, and TNF-α protein production as determined by a dose-dependent reduction of the frequencies of cytokine-producing PBMC, whereas the frequencies of IL-10–producing PBMC remain unaltered except at the highest DEP dose (Figs. 3, 4). To gain further insights into the mechanisms of these immunosuppressive effects of DEP, we studied a broad spectrum of genes corresponding to TLRs and their associated pathways as well as Th1-Th2-Th3 cytokines, chemokines, and their receptors. The central finding of these studies is that DEP prestimulation impairs M. tuberculosis-induced effector mechanisms by suppressing NF-κB (IL6, IFNG, NFKNA1, IFNB1, IFNA1, IL1A, and CSF3) and IRF (IFNG, IFNA1, IFNB1, and CXCL10) pathway target genes (Fig. 8C, Table II), thus modulating the production of IFN-γ, IL-6, IL-1β, and TNF-α. It has been shown that M. tuberculosis-mediated TLR activation leads to the recruitment of various adaptor proteins and the initiation of TLR signaling, such as the activation of adaptor protein MyD88-dependent and -independent pathways and the activation of various transcription factors including NF-κB and IRFs (38, 4143, 46, 47). Our data suggest that DEP prestimulation suppresses these MyD88-dependent signaling pathways, as indicated by the downregulation of several genes such as IL1A, IL6, and PTGS2 (Fig. 8B). In addition, DEP may also affect MyD88-independent pathways (91), as indicated by the inhibition of the expression of M. tuberculosis-induced type I IFN mRNAs (IFNA1 and IFNB1) following DEP prestimulation. Curiously, the observed increase in expression of proinflammatory mediators IL8, PTGS2, and TNF and decrease in expression of costimulatory receptors CTLA4 and CD80 (B7-1) in PBMC stimulated with DEP alone (Fig. 8A) coincide with that in monocyte-derived immature dendritic cells following in vitro exposure to ambient PM (93). Further, our finding of suppressed expression of M. tuberculosis-induced TLR3, TLR4, TLR7, and TLR10 mRNA following DEP prestimulation (Fig. 8C) is consistent with the reported suppression of expression and function of TLRs (TLR2 and TLR4) in monocyte-derived dendritic cells exposed to ambient PM (93, 94). In addition, suppression of TLR signaling may also result from the reduced expression and function of signaling intermediates downstream of the TLRs, as indicated by recent studies (80, 95, 96).

Based on the presented findings and published evidence, we propose a hypothetical model of DEP-mediated suppression of M. tuberculosis-induced TLR signaling (Fig. 11) in which DEP prestimulation of PBMC suppresses both MyD88-dependent and -independent M. tuberculosis-induced signaling pathways, leading to a hyporesponsive state upon subsequent exposure to M. tuberculosis. Such a DEP-induced hyporesponsive state is reminiscent of the effects of tobacco smoke on alveolar macrophages, which have been shown to become refractory to challenge with TLR2 and TLR4 agonists compared with alveolar macrophages of nonsmokers (97). The DEP-induced hyporesponsive state toward M. tuberculosis-stimulation described in this study also shows similarities to LPS-induced tolerance (40, 7981), a state characterized by a transitory hyporesponsiveness to secondary stimulation following repeated preceding LPS stimulation. It is interesting in this context that in contrast to our DEP prestimulation experiments, simultaneous stimulation of PBMC with DEP and M. tuberculosis affected the expression of fewer M. tuberculosis-induced genes (Figs. 7B, 8B). DEP prestimulation thus appears to be a prerequisite for the observed induction of the cellular hyporesponsiveness to subsequent M. tuberculosis-induced cellular responses. These studies are of relevance, as periods of respiratory exposure to air pollutants may precede subsequent M. tuberculosis infection events frequently in real-life situations.

FIGURE 11.

A hypothetical representation of DEP-mediated suppression of M. tuberculosis-induced TLR signaling and effector functions. M. tuberculosis-derived ligands (lipoarabinomannan [LAM], 19-kDa protein, heat shock proteins 65 and 71, and M. tuberculosis DNA) bind to TLR2, -4, and -9 and recruit adaptor protein MyD88 to the TLR receptor complex, which associates with IL-1R–associated kinase (IRAK) 1 and 4. TNFR-associated factor 6 (TRAF6) is also recruited to the receptor complex following phosphorylation of IRAK1 by IRAK4. Association of TRAF6–IRAK complex with TGF-β–activated kinase (TAK) and TAK-binding protein (TAB) 1 and 2 induces activation of TAK1 that phosphorylates IκB-kinase (IKK) complex consisting of three subunits: IKKα, β, and γ. IKK complex phosphorylates IκB, leading to its dissociation from NF-κB complex followed by translocation of NF-κB into the nucleus, where NF-κB binds and activates target genes. In addition to MyD88, ligand binding to TLR4 recruits adaptor protein Toll/IL-1R domain-containing adaptor protein inducing IFN-β (TRIF), which associates with TANK-binding kinase 1 (TBK1), leading to the phosphorylation of IRF3. Phosphorylated IRF3 translocates to the nucleus and induces production of type I IFNs (4749). DEP suppress M. tuberculosis-induced expression of target genes IL6, IL1A, and PTGS2 via MyD88-dependent and type I IFNs via MyD88-independent pathways. DEP-mediated downregulation of expression of TLR4, TLR7, and NF-κB target genes and type I IFNs is shown by downward arrows.

FIGURE 11.

A hypothetical representation of DEP-mediated suppression of M. tuberculosis-induced TLR signaling and effector functions. M. tuberculosis-derived ligands (lipoarabinomannan [LAM], 19-kDa protein, heat shock proteins 65 and 71, and M. tuberculosis DNA) bind to TLR2, -4, and -9 and recruit adaptor protein MyD88 to the TLR receptor complex, which associates with IL-1R–associated kinase (IRAK) 1 and 4. TNFR-associated factor 6 (TRAF6) is also recruited to the receptor complex following phosphorylation of IRAK1 by IRAK4. Association of TRAF6–IRAK complex with TGF-β–activated kinase (TAK) and TAK-binding protein (TAB) 1 and 2 induces activation of TAK1 that phosphorylates IκB-kinase (IKK) complex consisting of three subunits: IKKα, β, and γ. IKK complex phosphorylates IκB, leading to its dissociation from NF-κB complex followed by translocation of NF-κB into the nucleus, where NF-κB binds and activates target genes. In addition to MyD88, ligand binding to TLR4 recruits adaptor protein Toll/IL-1R domain-containing adaptor protein inducing IFN-β (TRIF), which associates with TANK-binding kinase 1 (TBK1), leading to the phosphorylation of IRF3. Phosphorylated IRF3 translocates to the nucleus and induces production of type I IFNs (4749). DEP suppress M. tuberculosis-induced expression of target genes IL6, IL1A, and PTGS2 via MyD88-dependent and type I IFNs via MyD88-independent pathways. DEP-mediated downregulation of expression of TLR4, TLR7, and NF-κB target genes and type I IFNs is shown by downward arrows.

Close modal

DEP stimulation has also been linked to a potent Th2 adjuvant activity and induction of allergic conditions (98102) and increased Th2 cytokine responses (103). These observations are consistent with our findings of a DEP prestimulation-mediated downregulation of the expression of IFN-γ and type I IFNs (Figs. 7C, 8C), which promote Th1 differentiation, inhibit Th2 responses, and are required for robust Th1 responses.

The DEP-induced alterations described in this study suggest that exposure to DEP may promote M. tuberculosis survival and progressive M. tuberculosis pathogenesis. Indeed, intrapulmonary instillation of DEP in C57BL/J6 mice followed by infection with attenuated bovine M. tuberculosis strain BCG results in a 5-fold increased pulmonary BCG load compared with mice exposed to BCG without prior DEP exposure and decreases the responsiveness of the murine alveolar macrophages to IFN-γ, IFN-γ–induced NO production, and the recovery of IFN-γ–producing lung lymphocytes (83). Similarly, long-term respiratory exposure of mice to DEP has been shown to result in increased pulmonary burden of M. tuberculosis (Kurono strain) following experimental aerogenic infection (62) and decrease in IL-1β, IL-12p40, IFN-γ, and inducible NO synthase production (62). Thus, the net effects of DEP-induced alterations of host immunity in rodent studies translate into decreased clearance of bacteria (63), further promoting progressive M. tuberculosis pathogenesis. Functional studies assessing the impact of the observed DEP effects on M. tuberculosis growth control in vitro (24), and in vivo human DEP exposure effects on antimycobacterial immunity are under way in our laboratory.

Future studies will have to determine whether the observed decrease in IFN-γ, IL-1β, and IL-6 protein and CXCL10 and IFNB mRNA, the increased IL13RA mRNA, and the unchanged IL-10 protein expression is indicative of a phenotype switch from classically to alternatively activated macrophages (104, 105). Indeed, recent studies suggest that tolerance and alternative macrophage activation (M2 polarization) in human monocytes and monocyte-derived macrophages are related processes (106).

Previous studies have shown that suppressive effects of DEP on macrophages reside with polar aromatic hydrocarbons and resins containing fractions (107). Consistent with these observations, our study points into the same direction as CB, an inert control for particle effects, did not suppress cytokine responses at the same concentrations of DEP tested (Fig. 5). This observation is also consistent with findings in rats in which DEP, but not CB exposure, decreased the pulmonary clearance of L. monocytogenes following intratracheal infection, suggesting that the exposure to the condensed organic and inorganic species adsorbed to the carbon core in DEP were responsible for the failing antimicrobial activity (65).

In conclusion, this study suggests that the DEP-mediated alterations of M. tuberculosis-induced NF-κB and IRF pathway target gene expression and a concomitant decrease in Th1 immunity may favor the progression of M. tuberculosis infection. This DEP-induced dysregulation of host immunity may thus result in increased susceptibility to M. tuberculosis infection and/or facilitate reactivation of latent M. tuberculosis infection and increased incidence of TB. Given the wide geographical scales of both air pollution and M. tuberculosis infections, the mechanistic insights from this study may have significant public health implications, particularly in cities and regions with high levels of air pollution and high TB prevalence rates.

We thank the study volunteers for participation in this study. We also thank Drs. S. Levine and S. Marcella at the Chandler Clinic in New Brunswick and Kathie Kelly McNeill at the Environmental and Occupational Health Sciences Institute for invaluable help with the study subject recruitment and administrative work, Megan Rockafellow (UMDNJ-School of Public Health) for help with cytospin preparations, Raj Patel (Electron Microscopy Core Imaging Lab, Department of Pathology, UMDNJ) for expert TEM, and Drs. Martha Torres (National Institute for Respiratory Diseases, Mexico City, Mexico) and Andrew Gow (Rutgers University, Piscataway, NJ) for reviewing the manuscript.

This work was supported by National Institute of Environmental Health Sciences Grants 5R21ES16928-2 (to S. Schwander), U19ES019536 (to J.Z.), and K08 ES013520 (to R.J.L.) and National Institute of Environmental Health Sciences-sponsored University of Medicine and Dentistry of New Jersey Center for Environmental Exposures and Disease Grant P30ES005022.

The online version of this article contains supplemental material.

Abbreviations used in this article:

BCG

bacillus Calmette-Guérin

CB

carbon black

DEP

diesel exhaust particle

IRAK

IL-1R–associated kinase

IRF

IFN regulatory factor

LAL

Limulus amebocyte lysate

MOI

multiplicity of infection

PAH

polycyclic aromatic hydrocarbon

PI

propidium iodide

PM2.5

particulate matter <2.5 μm in aerodynamic diameter

PPD

purified protein derivative

qRT-PCR

quantitative RT-PCR

TB

tuberculosis

TEM

transmission electron microscopy

UMDNJ

University of Medicine and Dentistry of New Jersey.

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The authors have no financial conflicts of interest.