All blood samples were collected per approved institutional review board protocols. CD14 magnetic beads (Miltenyi Inc., USA) were used to purify monocytes from PBMCs that were plated in 6 or 24 tissue culture well plates at a density of 5x10⁶ and 1x10⁶ cells per well, respectively. Eight-well slide chambers or coverslips received 10⁴ cells per chamber for confocal/immunofluorescent imaging studies. CD14 bead purified monocytes were grown in Iscove’s medium (IMDM) with 10% fetal bovine serum (FBS) and 10 μg/mL penicillin and gentamycin for 24 hours and then plated in antibiotics-free medium until 7 days for before differentiation into macrophages.

PBMCs were collected from deidentified, known TB patients and their healthy contacts under the approved IRB protocols of Dr. Restrepo from Reynosa, Mexico, under a collaboration. Approved IRB protocol: HSC-SPH-12-0037.

Cell supernatants were tested for cytokines using sandwich ELISA kits (Biolegend and R&D systems, USA) for various cytokines (TNF-α, IL-1β, IL-4, IL-6, IL-10, IL-12, and IL-13) secreted by GM-CSF competent and GM-CSF depleted macrophages after infection with M. tuberculosis. For the quantification of Cathilicidin through ELISA (Novus Biologicals), cell lysates of GM-CSF competent and GM-CSF depleted macrophages after infection with M. tuberculosis were used.

Macrophages were lysed with 0.05% SDS at different time points post-M. tuberculosis infection with or without hGM-CSF supplementation, α-hGM-CSF antibodies, or other pharmacological agents/inhibitors. Cell culture media along with hGM-CSF and α-hGM-CSF antibodies were replaced every alternate day during these experiments and assays. At different time points, lysates were plated at serial 10-fold dilutions in PBS using 7H11 Middlebrook agar plates (Difco Laboratories, Surrey, UK). The plates were incubated at 37°C for 3 weeks before counting CFUs. Data were expressed as log10- CFUs per million macrophages. Alamar Blue cell viability reagent (Life Technologies, DAL1025) was used to assess cell viability by adding the ready-to-use 1X solution to uninfected or M. tuberculosis-infected macrophages at various time points, followed by fluorescence measurement through fluorimeter per the manufacturer’s protocol. A fluorescence-based apoptosis/necrosis detection kit (Abcam #ab176749) that can simultaneously monitor apoptotic, necrotic, and healthy cells was used to measure and quantify apoptosis and necrosis in M. tuberculosis infected, uninfected, differentiated, and undifferentiated macrophages using a fluorescence microscope.

We have previously described this method in detail, and the original method described by the Harding lab has been extensively used by us and others for in vitro antigen presentation by macrophages (18). Briefly, M. tuberculosis-infected macrophages were washed after a 4-hour infection and overlaid with the F9A6-CD4 T cell hybridoma (gift from Dr. David Canaday lab) that recognizes an Ag85B epitope in the context of human HLA-DR1. IL-2 secreted from hybridoma T cells or other cytokines secreted from M. tuberculosis-infected macrophages were determined using a sandwich ELISA kit (Biolegend).

Six well tissue culture plates were seeded with macrophages. They were infected with M. tuberculosis for 4 hours at MOI of 1 and then washed three times with PBS and re-plated in the medium. At different time points, MΦs were washed three times with 1x PBS, and 50 µL RIPA buffer containing APM (anti-protease mix) was added to each well and incubated for 15 minutes. Lysates were then collected, and protein quantification (Bradford assay, Pierce Coomassie Plus cat no. 23238; Thermo Fisher Scientific) was performed. The quantitative Wes capillary immunoassay was used for the Western blot, in which the lysates were separated and detected using Wes separation capillary cartridge 12-230 kDa along with Wes Anti-Rabbit Detection Module (Simple Western system and Compass Software, Protein Simple). In brief, glass microcapillaries were loaded with stacking and separation matrices followed by sample loading. During capillary electrophoresis, proteins were separated by size and then immobilized to the capillary wall (19). Samples were loaded at 1 mg/ml dilution and the primary rabbit antibody ATG 5 and 7 was used at a dilution of 1:50 (ATG5: cat no.2360;cell signaling and ATG7: cat no.8558) and β-actin (Rabbit monoclonal, Sigma-Aldrich # SAB5600204) were used at 1:50 dilution. Data were analyzed with the Compass software (version 2.6.7). The area under the curve (AUC), which represents the signal intensity of the chemiluminescent reaction was analyzed for all the antibodies and β-actin. Values given for protein expression were normalized to β-actin. Quantitation of protein levels (area under each peak; arbitrary units [A.U.]) were performed using the Compass software (version 2.6.7).

GM-CSF competent and depleted macrophages were plated in 8-well chamber slides at a density of 10⁴ cells per chamber and infected with red fluorescent protein (RFP) expressing M. tuberculosis (rfp M. tuberculosis) at an MOI of 1. Per established procedures, autophagy was evaluated using at least three criteria. Fixed macrophages were permeabilized and stained using specific antibodies against LC3 followed by secondary staining with Alexfluor488 (Jackson Immunoresearch # 109-545-003) as described previously (19). Percent rfp M. tuberculosis phagosomes colocalizing with antibodies was conducted as described previously (20).

Total RNA from macrophages was extracted using RNAeasy mini kit (Qiagen, Germany). RNA concentration and purity ratios (OD260/280, OD260/230) were measured using the NanoDrop ND-1000 spectrophotometer (Thermo Fisher Scientific, USA). cDNA synthesis was performed on a CFX96 Real-Time PCR System (Biorad, USA) using the 2X OneStep qRT-PCR Mastermix Kit (Applied Biosystems, USA) according to manufacturer’s instruction. Quantitative PCR (qPCR) was performed using SYBR green probe and gene specific primers ( Supplementary Table S2 ). Threshold cycle numbers were transformed to ΔΔCt values, and the results were expressed relative to the reference gene, GAPDH. Gene expression data was performed using GraphPad Prism ver. 6.0 suite (GraphPad Software). Student’s t-test was used for mean comparison between GM-CSF competent and depleted macrophages. Significance was set at the 0.05 level.

Macrophages were plated in 96-well plates at a density of 1x10⁴ cells per well in triplicate and infected with M. tuberculosis (MOI=1). Cells were then treated with fluorescent probes for the quantification of NO using diaminofluorescein diacetate (DAF-2 DA), per manufacturer’s instructions (Enzo Life Sciences, USA); DCFDA was used to detect ROS. Quantification of NO/ROS release was determined by calculating the fluorescence emitted over time using an ^ex485 nm/^em515 nm and plotting AFUs (± SD) against time using Ascent fluoroscan software version 2.6.

All experiments were performed at least 3 times containing triplicate wells per experimental group or combination. Statistical parameters including the definition of central value and spread (mean ± SD) of macrophages per group are annotated in the corresponding figure legends. Statistical analysis was performed with GraphPad Prism version by using, unless otherwise stated, unpaired t tests with 95% confidence intervals. P values for CFU count in macrophages were determined using 1- or 2-way ANOVA with Tukey’s posthoc test (GraphPad PRISM software).

We compared the levels of GM-CSF produced by monocytes isolated from the peripheral blood of various healthy subjects (HS) (n=26) and analyzed their bacterial load and survival during M. tuberculosis infection in vitro. Prior to M. tuberculosis infection, we allowed their differentiation into macrophages and measured GM-CSF production. CD14+ classic monocytes, isolated from the peripheral blood of different HS were differentiated via adherence on cell culture plates. Quantifiable levels of GM-CSF in cell culture supernatants were not detectable before 48 h using the Sandwich ELISA method ( Supplementary Figure S1 ). After seven days of differentiation, adherent, fully differentiated macrophages were isolated and infected with M. tuberculosis at a low multiplicity of infection (MOI) of one. GM-CSF levels were immediately detectable post-infection in these differentiated cells, though they varied significantly over time between donors ( Figure 1A ). We also measured GM-CSF levels in an additional, uninfected set of differentiated macrophages for comparison with infected cells from the same donors set. Relative to uninfected cells, infected cells produced significantly more GM-CSF early after infection (1-3 days) in most of the HS. However, GM-CSF levels became more similar between these two groups at three days post infection. We compared intracellular colony-forming units (CFUs) of M. tuberculosis, macrophage viability, and GM-CSF levels over time after infection and found that intracellular CFUs declined in most individuals, though three individuals (11.5%) showed an increase in intracellular M. tuberculosis burden ( Figure 1B ). Similarly, M. tuberculosis-infected macrophage viability declined slowly but remained over 80% seven days post-infection in a majority of individuals (77%) ( Figure 1C , Supplementary Table 1 ). Macrophages from six individuals (23%) lost over 20% cell viability by seven days post-infection and exhibited accelerated cell death following infection ( Figure 1C ). Increased production of GM-CSF directly correlated with prolonged survival and improved intracellular bacterial control of macrophages ( Figures 1D, E ). GM-CSF levels and bacterial burden were inversely correlated in macrophages ( Figure 1D ). On the other hand, there was a strong positive correlation between GM-CSF levels and macrophage viability after infection ( Figure 1E ) that was also observed in uninfected cells ( Supplementary Figure S2 ). Thus, GM-CSF may contribute to long-term survival in macrophages while preventing M. tuberculosis-induced cell death. To investigate the association between macrophage GM-CSF production, cell survival, and M. tuberculosis infection control in undifferentiated cells, we infected CD14+ monocytes from different donors prior to differentiation ( Figures 1F, G ). Undifferentiated monocytes did not produce detectable levels of GM-CSF following infection with M. tuberculosis (data not shown), and an increase in bacterial burden was observed in these cells ( Figure 1F ). Furthermore, viability of undifferentiated monocytes decreased rapidly relative to differentiated macrophages over the seven-day infection period ( Figure 1G ). These data suggest that undifferentiated monocytes are more permissive for M. tuberculosis growth and infection-induced cell death as compared to differentiated macrophages.

To determine whether macrophage permissiveness for M. tuberculosis growth and infection-induced cell death were dependent on GM-CSF, we assessed M. tuberculosis proliferation in healthy and infected macrophages in the presence and absence of GM-CSF. We used an anti-human GM-CSF antibody (α-hGM-CSF) to neutralize GM-CSF effect on macrophages. We added α-hGM-CSF antibodies (2 μg/1 million cells) to monocytes before or after differentiation, as earlier described. M. tuberculosis-infected undifferentiated monocytes treated with α-hGM-CSF showed increased bacterial burden and decreased survival over a period of seven days relative to those treated with IgG isotype ( Figures 2A, B ). Uninfected and undifferentiated monocytes treated with α-hGM-CSF antibody had significantly reduced survival over a period of seven days relative to untreated controls ( Figure 2C ). Microscopic examination revealed inhibition of cell differentiation in uninfected and infected monocytes treated with α-hGM-CSF ( Figure 2D ). Similar results were also observed when hGM-CSF receptor (CSF2RA) was blocked through anti-CD116 antibody (Data not shown).

We next assessed whether exogenous hGM-CSF enhanced monocyte survival during differentiation in the presence and absence of M. tuberculosis infection. We added a range of hGM-CSF doses (1-10 ng/5 million cells) to undifferentiated monocytes isolated from three human subjects. Survival in infected and uninfected controls was assessed in cells from the same donor in the presence or absence of exogenous hGM-CSF. hGM-CSF (2 ng/1 million cells) enhanced cell survival by over 30% by day 28 post-M. tuberculosis infection relative to untreated infected macrophages ( Figure 2E ). Doses exceeding 2 ng hGM-CSF/1 million cells enhanced cell survival but also induced cell proliferation and inhibited differentiation (data not shown). Uninfected monocytes treated with 2 ng hGM-CSF/1 million monocytes survived significantly longer relative to untreated monocytes ( Figure 2F ). These data suggest that a limited supplementation of GM-CSF could further extend the survival of monocyte during M. tuberculosis infection.

Exogenous hGM-CSF also increased the formation of granulomatous structures in peripheral blood mononuclear cells (PBMC)-derived monocytes ( Figure 3A ), a previously documented phenomenon (21). While M. tuberculosis infection itself induced granuloma-like structures in macrophage culture, exogenous hGM-CSF further increased the size and number of granuloma-like centers ( Figure 3A ). Conversely, treatment with α-hGM-CSF reduced the granulomatous response in macrophages and significantly reduced the number and the size of granulomatous structures. These granuloma-like structures were not seen in uninfected macrophages (data not shown). Granuloma-like structures began to disappear 14 days post-infection/exogenous hGM-CSF treatment, but the number of M. tuberculosis CFUs remained significantly low relative to untreated macrophages ( Figures 3B, C ). With increasing exogenous hGM-CSF doses, a time-dependent decrease in intracellular CFUs was observed, indicating that GM-CSF may restrict M. tuberculosis by promoting granulomatous response in cells ( Figure 3C ). Relative to untreated cells, treatment with 2 ng of hGM-CSF/1 million macrophages reduced bacterial burden by over 2.5 logs by day 28 post-infection. Notably, granuloma-like structures were also seen early during infection (5-10 days) in untreated macrophages, and though these cells could prevent intracellular proliferation of M. tuberculosis bacilli, they could not reduce bacterial burden with time ( Figures 3A–C ). We next examined if intrinsic levels of GM-CSF produced by macrophages could prevent intracellular proliferation of M. tuberculosis. The endocrine effect of GM-CSF in macrophages was blocked using α-hGM-CSF, and intra-macrophage CFUs were examined up to 28 days post-infection ( Figure 3D ). A marked increase in M. tuberculosis CFUs was observed in the first 14 days relative to untreated macrophages. After 21 days 100% cell death was observed in α-hGM-CSF treated macrophages whereas untreated macrophages maintained more than 50% viability at this time point ( Figure 3E ). Taken together, these results clearly indicated that self-survival of macrophages as well as their ability to form granuloma and contain M. tuberculosis infection is dependent on GM-CSF.

The molecular mechanism through which GM-CSF may regulate host cell survival and bacterial containment is unknown. We investigated the downstream effects of GM-CSF supplementation/depletion on cell death and survival pathways in uninfected and M. tuberculosis-infected cells. Specifically, we assessed transcriptional expression of cell death/survival related genes HSP27, BCL-xL, BCL-2, BAX/BAK, and MCL1 and the mode of cell death during exogenous hGM-CSF supplementation/antibody-mediated depletion. M. tuberculosis-infected macrophages supplemented with exogenous hGM-CSF (2 ng/1 million cells) showed significantly less apoptotic cell death over 14 days relative to untreated macrophages ( Figure 4A ). Moreover, apoptotic cell death was more common than necrotic cell death in infected macrophages. GM-CSF supplementation resulted in significantly lower levels of apoptosis even in uninfected macrophages ( Supplementary Figure S3 ), indicating a homeostatic role for GM-CSF in preventing apoptotic cell death. Anti-apoptotic gene BCL-2 was upregulated ~2-fold in hGM-CSF-treated macrophages in infected and uninfected groups at day 7 ( Figure 4B ). However, BCL-xL, another anti-apoptotic gene, did not vary between groups. Pro-apoptotic gene BAX was not significantly up or downregulated in hGM-CSF-treated macrophages relative to untreated controls. Apoptosis as well as necrosis significantly increased in infected and uninfected macrophages treated with α-hGM-CSF antibodies, and we observed a corresponding increase in pro-apoptotic BAX expression ( Figure 4C ). Cell survival-associated gene HSP27 was downregulated significantly in M. tuberculosis-infected macrophages treated with α−hGM-CSF antibody; uninfected macrophages showed a similar profile with comparatively lower magnitude of difference.

Together, these data indicated a pattern of apoptosis and necrosis inhibition in macrophages with higher GM-CSF levels that were most prominent in infected macrophages. To determine whether this GM-CSF effect was mediated by apoptosis inhibition or other antimicrobial host defense pathways, we assessed antimicrobial effector mechanisms, including reactive oxygen species (ROS)/nitric oxide (NO) generation, antimicrobial peptide production, autophagic flux in GM-CSF-competent and GM-CSF-depleted macrophages during M. tuberculosis infection. GM-CSF-competent macrophages did not show any significant differences in ROS production levels compared to GM-CSF-depleted macrophages ( Figure 4D ). NO levels in GM-CSF-competent macrophages were significantly higher compared to GM-CSF-depleted macrophages, indicating that NO may be an antimicrobial effector mechanism induced by GM-CSF signaling. To determine if an NO-mediated effector mechanism was associated with macrophage survival, we measured host and bacterial cell survival in GM-CSF-competent and GM-CSF-depleted macrophages treated with a pharmacological inhibitor of NO; NG-Methyl-L-arginine acetate salt (L-NMMA) ( Figures 4E, F ). In GM-CSF-competent macrophages, L-NMMA treatment resulted in increased bacterial burden but not significant cell death. In GM-CSF-depleted macrophages, untreated and L-NMMA-treated cells showed increased bacterial burden and host cell death. These data suggest that NO contributes to GM-CSF-mediated antimicrobial effector mechanisms but not host cell survival.

Since NO is known to induce production of antimicrobial peptides and activate autophagy, which could also contribute to boosting intracellular host defenses, antimicrobial peptide cathilicidin (LL-37) and autophagic flux in GM-CSF competent and depleted macrophages were further examined. LL-37 levels were similar between GM-CSF-competent and GM-CSF-depleted macrophages, indicating that GM-CSF-mediated restriction of M. tuberculosis bacilli in macrophages is not dependent on LL-37 ( Figure 4G ). Strikingly, increased autophagy flux (as examined through the LC3B positive puncta) in GM-CSF competent macrophages was observed as compared to GM-CSF depleted macrophages ( Figure 4H ). Increased LC3 B gene expression at the transcription level was also observed in hGM-CSF-treated infected and uninfected macrophages relative to untreated macrophages ( Supplementary Figure S4 ). Interestingly, blockade of NO via L-NMMA did not affect autophagic flux in GM-CSF competent/depleted macrophages, indicating that GM-CSF-mediated induction of autophagy is NO-independent ( Figure 4H ). To determine whether GM-CSF-mediated autophagy affects host and bacterial cell survival, we induced and blocked autophagy using rapamycin and 3-MA, respectively, in GM-CSF-competent macrophages during M. tuberculosis infection. Low-dose rapamycin resulted in decreased bacterial burden, but higher doses reversed this effect. Blockade of autophagy through 3-MA resulted in increased bacterial burden and decreased cell survival ( Figures 4I, J ). Thus, GM-CSF-dependent host cell survival and antimicrobial effector mechanisms may be partially autophagy-dependent; however, the data also indicated that precise regulation of autophagy may be equally important for pathogen restriction and host cell survival.

We evaluated a panel of pro- and anti-inflammatory cytokines in GM-CSF-competent/deficient macrophages three days post-M. tuberculosis infection ( Figure 5A ). GM-CSF-depleted macrophages secreted significantly lower levels of IL-1β, IL-12, and IL-10 and significantly higher levels of TNF-α and IL-6 compared to GM-CSF-competent macrophages. No significant difference was observed in IL-4 and IL-13 levels. Thus, GM-CSF signaling may simultaneously be inducing pro-and anti-inflammatory immune responses and a dual GM-CSF response may be considered more of a regulatory immune response than a pro-inflammatory response. Interestingly, IL-12 levels from GM-CSF-competent macrophages were high relative to GM-CSF-deficient macrophages, suggesting that GM-CSF may induce macrophage maturation (22).

We next evaluated whether GM-CSF-competent/deficient macrophages had different T cell priming capabilities during M. tuberculosis infection. We used an in vitro antigen presentation assay using a CD4 hybridoma T cell recognizing a specific epitope on Ag85B to assess T cell priming by GM-CSF-competent/deficient macrophages. IL-2 levels secreted from hybridoma T cells were monitored over time with overlay to M. tuberculosis-infected macrophages ( Figure 5B ). GM-CSF-competent macrophages induced higher levels of antigen presentation after two days of infection, though the difference was not significant relative to GM-CSF-depleted macrophages. However, GM-CSF competent macrophages were able to sustain a much higher levels of antigen presentation beyond day 3 post infection as compared to GM-CSF deficient macrophages, hence further suggesting the critical role of GM-CSF signaling in sustenance of antigen presentation and efficient processing and presentation of antigens to CD4T cells. it is also possible that low number of viable cells in GM-CSF deficient macrophages could have also contributed to the reduction in overall levels of antigen presentation. However, the magnitude of difference in IL-2 levels secreted by untreated vs anti-GM-CSF antibody treated macrophages is much higher at day 7 post infection (almost 10 fold: 30 picograms vs 300 picograms) ( Figure 5B ) as compared to the magnitude of difference in cell viability of untreated vs anti-GM-CSF antibody treated macrophages (less than 2 fold) ( Figure 3E ) at day 7 post infection. We thus believe that during the late phase of infection, GM-CSF is affecting the antigen presentation capability of macrophages.

We further compared phagosome maturation events in GM-CSF-competent/deficient macrophages after M. tuberculosis infection. At three days post-infection, over 50% of M. tuberculosis-containing phagosomes colocalized with LC3B positive endosomes in GM-CSF-competent macrophages, with GM-CSF deficient macrophages exhibiting less than 20% co-localization ( Figure 5C ). Notably, this difference was not significant during the first 24 h of infection (data not shown). Thus, increased fusion of LC3B containing endosomes with M. tuberculosis bacilli indicated an autophagy dependent maturation of phagosomes. Autophagosome-associated LC3B and other autophagy-related genes, including LC3B, Atg5 and Atg7, also demonstrated increased expression at the transcription/translational level in hGM-CSF-treated macrophages and decreased expression in α-GM-CSF-treated macrophages ( Figures 5C, D ), further indicating the role of GM-CSF in macrophage autophagy regulation. Thus, increased maturation of M. tuberculosis-containing phagosomes via GM-CSF-mediated autophagy may degrade M. tuberculosis bacilli and process its antigens, resulting in improved and sustained antigen presentation in macrophages.

As our results indicated that natural variations in GM-CSF production/signaling could be a factor in determining an individual’s vulnerability to TB, we conducted parallel studies to compare patterns of GM-CSF production in macrophages from individuals with active TB to healthy controls. Monocytes from healthy controls and newly diagnosed TB patients were isolated and compared for their ability to produce GM-CSF, control host cell survival, and restrict M. tuberculosis growth. M. tuberculosis infection increased macrophage GM-CSF production in both populations after seven days of infection ( Figure 6A ), though macrophages from active TB patients (n=4) produced significantly less than those from healthy controls (n=5). These levels remained low throughout the 14-day infection period ( Supplementary Figure S5 ) and translated to increased bacterial burden and host cell death relative to healthy controls ( Figures 6B, C ).

Because some of our active TB population also had preexisting diabetic conditions, which can cause a metabolic reprogramming of immune cells that leads to defective immune response against TB, we examined whether diabetes-induced metabolic changes were associated with macrophage GM-CSF production. We first characterized the central metabolic state of macrophages from active TB patients and healthy controls by examining the expression of genes related to glycolysis and oxidative phosphorylation. Macrophages from active TB patients exhibited significant downregulation (1.5- 20 fold) of genes encoding glucose transporters 1 and 6 (GLUT1 and 6), glycolytic enzymes/isozymes hexokinases (HK2 and HK3), phosphofructokinase 1 (PFK1), and lactate dehydrogenase A (LDHA) relative to healthy controls ( Figure 6D ). Further, decreased glycolytic flux also resulted in concurrent downregulation of lipid metabolism-related gene Peroxisome Proliferator-Activated Receptor Gamma (PPAR-γ) in macrophages from active TB patients relative to controls.

We next evaluated whether metabolic dysregulation was causing reduced GM-CSF production in macrophages. We induced metabolic dysregulation in macrophages by treating cells with glucose, insulin, and palmitate (23) and quantified GM-CSF production in these metabolically dysregulated macrophages (MME macrophages) relative to a control group of untreated cells from the same donor. MME macrophages produced significantly reduced levels of GM-CSF, and exhibited significantly reduced survival following M. tuberculosis infection relative to untreated control macrophages ( Figures 6E, F ). We also observed reduced expression of glycolysis-related genes Protein glycosylation K (PGLK), Monocarboxylate transporter 4 (MCT4), LDHA, and HK3, and lipid metabolism-related gene PPAR-γ in MME macrophages relative to control macrophages ( Figure 6G ). MME macrophages also showed a significantly higher intracellular M. tuberculosis burden relative to untreated macrophages ( Figure 6H ). Thus, metabolic dysregulation may also cause a reduction in macrophage GM-CSF production that results in their reduced survival and increased permissiveness to M. tuberculosis growth.