TLR-4 Agonist Induces IFN-γ Production Selectively in Proinflammatory Human M1 Macrophages through the PI3K-mTOR– and JNK-MAPK–Activated p70S6K Pathway

Macrophages, a heterogeneous population of myeloid cells, are present in almost all organ systems of the body as tissue-resident macrophages (1–3). They play a key role in innate and adaptive immunity and in the pathogenesis of various chronic and inflammatory autoimmune diseases, cancer, and infection (1–3). These cells are long-lived with a life span extending from months to years (4, 5). Based on the cytokine milieu, the macrophage populations are believed to polarize into two major phenotypes: the activated M1 macrophages that help to control infection and promote inflammation; and the anti-inflammatory M2 macrophages that play a key role in the resolution of inflammation, tissue remodeling, and tissue repair (3, 6). M1 macrophages are derived in vitro following stimulation of monocyte-derived macrophages (MDMs) with IFN-γ in the presence or absence of LPS (3, 7–9). M2 macrophages are further subdivided into M2a, M2b, and M2c subsets, based on their exposure to distinct cytokines and/or bacterial products, their state of alternate activation and regulatory functions, cytokine and chemokine profiles, and relative expression of surface markers (7, 10, 11). M2a macrophages are induced following stimulation with IL-4 or IL-13, whereas M2b macrophages are generated by stimulation with immune complexes, LPS or IL-1 receptor ligands (3, 11). M2c macrophages are generally induced by glucocorticoids and/or IL-10 (11, 12). M2a and M2c subsets are anti-inflammatory whereas the M2b subset secretes both pro- and anti-inflammatory cytokines (7, 11–13). There is evidence that an M1 polarization signature is associated with bacterial infections including typhoid fever, tuberculoid leprosy, active tuberculosis, and Helicobacter pylori gastritis (8, 14–17), and with chronic autoimmune and inflammatory diseases such as rheumatoid arthritis, atherosclerosis, and type-2 diabetes (18, 19). Conversely, M2 polarization program is observed in lepromatous leprosy, Whipple’s disease, and chronic rhinosinusitis (14, 20, 21).

IFN-γ plays a key role in immunity against intracellular pathogens and cancer (22). Mice genetically deficient in IFN-γ or its receptor are highly susceptible to several microbial pathogens (23). It activates phagocytes by upregulating their lysosomal enzymatic activity and increasing the level of reactive oxygen species (24). IFN-γ is primarily produced by activated CD4⁺ and CD8⁺ T cells and NK and NKT cells (25). There is evidence to suggest that macrophages in murine models produce IFN-γ under certain conditions (26–29). Peritoneal- and bone marrow–derived murine macrophages were shown to produce IFN-γ following stimulation with either IL-12 or IL-12 and IL-18 (26, 28) and stimulation of macrophages with LPS increased steady-state levels of IFN-γ mRNA (27). Further, in vivo and in vitro models of pulmonary mycobacterial infection studies have shown that murine lung macrophages produce IFN-γ (29). In humans, there are two reports which show that macrophages produce IFN-γ in disease conditions such as pulmonary sarcoidosis (30), and in vitro infection of alveolar macrophages with Mycobacterium tuberculosis (31). Recently, we have also shown that macrophages derived from hepatitis C patients’ with stage F3-4 produced high levels of IFN- γ (32). Whether human macrophages or any of their subsets produce IFN-γ in health remains unknown.

The TLRs, expressed on macrophages and dendritic cells, recognize structurally conserved molecules found on microbes leading to macrophage activation and development of immune responses (33). Whether microbial products such as TLR-2, TLR-3, and TLR-4 agonists induce IFN-γ production in human macrophages or any of their subsets is not known. We report that human M1 macrophages, unlike the M2 subset, are a significant source of IFN-γ. Moreover, stimulation of M1 macrophages with TLR-2, TLR-3, or TLR-4 agonists, unlike the M2 subset, further enhanced IFN-γ production. We also investigated the signaling pathways underlying IFN-γ production by human M1 macrophages following stimulation with LPS, the TLR-4 agonist. In addition to the Jak/STAT-mediated differentiation of M1 cells, we identified, to our knowledge for the first time, a dependence on the S6 kinase activated by multiple upstream signaling pathways including PI3K, the mammalian target of rapamycin complex 1/2 (mTORC1/2), p38 MAPK, and the JNK MAPK in the regulation of LPS-induced IFN-γ production.

Blood was collected from healthy donors and COVID-19–infected patients admitted into the Ottawa Hospital as per the protocol approved by the Ottawa Health Sciences Network Research Ethics Board. Informed consent was obtained from the donors in a written form. PBMCs were isolated by collecting the buffy coat generated after Ficoll-Paque (GE Healthcare, Buckinghamshire, UK) density centrifugation. Briefly, PBMCs were resuspended (4 × 10⁶ cells/ml) in Iscove’s Modified DMEM 1× medium (Sigma-Aldrich, Oakville, Ontario, Canada) and seeded into 12-well polystyrene plates (Thermo Fisher Scientific, Rochester, NY) to isolate monocytes by adherence. Monocytes were allowed to adhere at 37°C, 5% CO₂/air mixture for 3 h. Nonadherent cells were washed off using Iscove’s Modified DMEM 1× medium. The adherent monocytes were cultured for 6 d in Iscove’s Modified DMEM 1× medium supplemented with 10% FBS (GE Healthcare, Canada), 10 U/ml penicillin/gentamicin (Sigma-Aldrich), and 10 ng/ml recombinant macrophage-CSF (R&D Systems, Minneapolis, MN) to generate MDMs. The medium was changed every second day. The MDMs were polarized using appropriate stimuli for 2 d: IFN-γ (20 ng/ml) (Thermo Fisher Scientific, Rochester, NY) for M1 macrophages, IL-4 (20 ng/ml) (R&D Systems) for M2a macrophages, LPS (1 μg/ml) (Sigma-Aldrich) and IL-1β (10 ng/ml) (R&D Systems) for M2b macrophages, and IL-10 (10 ng/ml) (R&D Systems) for M2c macrophages, as described previously (11). The polarized MDMs thus generated were washed thoroughly three times to remove the polarizing stimuli followed by stimulation with either LPS (1 µg/ml), lipoteichoic acid (LTA) (5 µg/ml), or polyinosinic-polycytidylic acid (poly(I:C)) (50 µg/ml) for 24 h or IL-12 (10 ng/ml) and IL-18 (10 ng/ml) (both from R&D Systems) for 48 h.

The following reagents were purchased: ruxolitinib (RUX, Jak inhibitor), rapamycin (mTORC inhibitor), PF-4708671 (S6K inhibitor), and PD-0325901 (Erk inhibitor) from Sigma-Aldrich (St. Louis, MO); SP-600125 (JNK inhibitor), SB-203580 (p38 inhibitor), and LY294002 (PI3K inhibitor) from Calbiochem (La Jolla, CA); MK-2206 (Akt inhibitor) from Cayman Chemicals (Ann Arbor, MI); and Torin (mTORC1/2 inhibitor) from Tocris Bioscience (Oakville, Ontario, Canada). DMSO was used as a control vehicle as this was used for dissolving all the above inhibitors used in the study.

Cell surface staining was performed based on a previously described protocol (11). Briefly, the unpolarized and polarized macrophages were washed twice with PBS. The cells were dislodged using 500 μl of Accutase (Stemcell Technologies) for 45 min at 37°C. Cells were then collected and centrifuged at 2000 rpm for 5 min, followed by two times wash with PBS. The cells were incubated with human FcR blocking buffer (Miltenyi Biotec, Auburn, CA) for 20 min at 4°C followed by staining using conjugated Abs for CD14, CD80, CD86, CD163, CD200R, and HLA-DR (BD Biosciences, San Jose, CA) for 1 h at 4°C in the dark. The cells were washed with PBS. PBS (200 μl) was added to each tube and the fluorescence was measured using PE (CD14, CD200R, and CD80), FITC (CD86 and CD163), and PE/Cy7 (HLA-DR) channels in the flow cytometer BD LSRFortessa cell analyzer (BD Life Sciences, Mississauga, Ontario, Canada). All samples tested were compared with M0 samples. The histograms and mean fluorescence intensity (MFI) were analyzed and plotted using FlowJo software version 10.0.7 (BD Biosciences, San Jose, CA) and GraphPad Prism 5.0 software (San Diego, CA). The cell surface marker expression between the samples was quantified by gating the untreated macrophages and applying the same gate to all the treated samples followed by calculation and quantification of MFI.

Briefly, MDMs and M1-polarized macrophages were pretreated with anti-IFN-γ receptor Abs or isotype control Abs (40 μg/ml each, BioLegend, San Diego, CA) for 2 h to block IFN-γ binding to its cognate receptor. The cells were then stimulated with the collected supernatants for 24 h. These blocked MDMs were treated with the previously collected supernatants for 24 h to induce the expression of TLR-4. The cells were labeled with fluorescently conjugated Abs specific for TLR-4 and fluorescence was detected by flow cytometry using a FACSCanto flow cytometer and FACSDiva software (BD Biosciences, Franklin Lakes, NJ). Histograms were plotted using FlowJo software version 10.0.7 (BD Biosciences, San Jose, CA) as described previously (11, 34, 35).

The human IFN-γ Duoset (R&D Systems) ELISA was used to measure IFN-γ levels in macrophage supernatants as described previously (11). The plates were coated overnight at 4°C with 100 μl/well of capture Ab (4 μg/ml) in PBS. The plates were blocked with 300 μl/well of 1% BSA in PBS for 2 h. Following blocking, the plates were incubated with 100 μl/well of detection Ab (200 ng/ml) in 1% BSA in PBS and 2% heat-inactivated normal goat serum. Streptavidin-HRP (100 μl/well in 1% BSA in PBS) was added for 30 min followed by the addition of 100 μl/well of substrate (BioFX Laboratories, Owing Mills, MD). The reaction was stopped by adding 50 μl/well of Stop Solution (BioFX Laboratories, Owing Mills, MD). The intensity of light in each well was measured using iMark Microplate reader at 490 nm (Bio-Rad Laboratories, Mississauga, Ontario, Canada) and data were processed using Micro Plate Manager 6 software.

Intracellular IFN-γ was analyzed as described previously (32, 36). Macrophages were transferred to polypropylene tubes, washed, and resuspended in 100 µl of media containing 10 µl of dilute Golgi transport inhibitor (containing 15 μg/ml brefeldin A, MilliporeSigma, Oakville, Ontario, Canada) for the last 6 h of cell culture to prevent the release of IFN-γ. Cells were then washed with PBS and fixed using 4% paraformaldehyde in the dark for 15 min at room temperature. Cells were washed again and permeabilized using 100 µl of permeabilization buffer containing saponin (MilliporeSigma) in 10% human AB serum (Valley Biomedical, Winchester, VA) + anti-human IFN-γ-FITC Abs (clone 4S.B3, BioLegend) or IgG1κ-FITC isotype control (clone MOPC-21, BioLegend). Cells were incubated for 30 min in the dark at 37°C. Cells were then washed and resuspended in 300 µl of PBS. To minimize nonspecific binding, human AB serum (10%) was included in all buffers. Samples were analyzed by flow cytometry within 1 h and data were analyzed using FCS Express Research Edition 4.0 (De Novo Software, Los Angeles, CA).

An RNeasy Plus Mini Kit (Qiagen, Hilden, Germany) was used to extract total RNA from cells, according to the manufacturer’s instructions. A master mix was then prepared as follows using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Carlsbad, CA): 7.5 μl of 10 × reverse transcriptase buffer, 3 μl of 2′-deoxynucleoside 5′-triphosphate, 7.5 μl of random primers, 3.75 μl of reverse transcriptase, and 53.25 μl of ddH2O. The master mix (25 μl) was added to the isolated RNA samples and the GeneAmp PCR System 2700 amplifier (Applied Biosystems, Carlsbad, CA) was used to carry out reverse transcription for 2 h to yield cDNA. For real-time PCR, the generated cDNA was mixed with 12.5 μl of Taqman DNA polymerase, 1.25 μl of primer pairs for either IFN-γ (Hs00989291_m1) or the internal PCR control β-actin (ACTB Hs99999903_m1), and 8.75 μl of ddH2O. Using the 7500 Real Time PCR System (Bio-Rad Laboratories), a quantitative PCR program was carried out as follows: an initial incubation for 2 min at 50°C and 10 min at 95°C, followed by 40–50 cycles of 15 s at 95°C to denature DNA and 2 min at 60°C to allow for annealing and elongation. Cycle threshold values in amplification plots (Applied Biosystems) were used to calculate mRNA transcript levels normalized to the mRNA transcript expression of β-actin.

Western immunoblotting was performed as described previously (37). Briefly, total cell proteins were subjected to SDS-PAGE and transferred onto a polyvinylidene difluoride membrane (Bio-Rad Laboratories). Membranes were probed with the following primary Abs: phospho-p70 S6 kinase (-p70S6K) (Thr389), phospho-Akt (Thr308), phospho-p38 MAPK (Thr180/Tyr182), phospho-SAPK/JNK (Thr183/Tyr185), phospho-p44/42 MAPK (Erk1/2) (Thr202/Tyr204), and phospho-Jak1 (Tyr1034/1035); all were purchased from Cell Signaling Technology (Danvers, MA). Membranes were then washed and probed with goat secondary Abs conjugated to HRP for 1 h (Bio-Rad Laboratories). For total loading proteins, membranes were restriped and probed with either anti-β-actin (#8457) or anti-GAPDH (#2118) Abs (Cell Signaling Technology). Protein bands were visualized using the Amersham ECL Western blotting detection system (GE Healthcare, Buckinghamshire, UK) and GeneSnap software.

GraphPad Prism 5 software was used to generate graphs and to conduct statistical analyses. Means between the two groups were compared using the Student t test. The results are expressed as a mean ± SD of the mean from at least three experiments, unless otherwise stated.

We first phenotypically characterized human M0, M1, M2a, M2b, and M2c macrophages by flow cytometry using a nonexhaustive panel of well-established cell surface markers, namely CD14, CD80, CD86, CD200R, CD163, and HLA-DR as described previously (11, 32). M0 macrophages expressed all the above surface markers. However, these surface markers were differentially expressed in the remaining macrophage subsets. The surface markers expressed at substantially high levels at ∼1.5–2-fold in MFI compared with the M0 macrophages were designated as “high” and chosen as characteristic for that macrophage subset. As shown in (Fig. 1 and Table I, M1 macrophages differentially expressed very high levels of CD80, CD86, and HLA-DR compared with the M0 macrophages and were designated as CD80^high, CD86^high, and HLA-DR^high. Similarly, M2a macrophages were defined as CD86^high, CD200R^high, and HLA-DR^high; M2b macrophages were defined as CD14^high, CD80^high, and CD163^high; and M2c were characterized as CD14^high and CD163^high.

IL-12, produced by macrophages upon pathogen recognition during the innate immune response, acts on NK cells and T cells to promote IFN-γ secretion which in turn activates macrophages (38). Similarly, IL-18 induces IFN-γ in murine and human T cells (39). IL-12 and IL-18 also synergize to produce IFN-γ in murine T cells, NK cells (39, 40), and macrophages (28). To determine whether IL-12/IL-18 induce IFN-γ production in human macrophages, we show that M0 macrophages treated with IL-12 and IL-18 produced significantly high levels of IFN-γ compared with the untreated or LPS-, LTA-, or poly(I:C)-treated M0 macrophages (Fig. 2A).

Stimulation of pattern recognition receptors like TLRs leads to the activation of macrophages and results in cytokine production and eventual clearance of the pathogen (33). Therefore, it was of interest to examine IFN-γ production in macrophages following activation of TLRs known to be stimulated by bacterial and viral pathogens. Ligands against two cell surface–bound TLRs and one against an intracellular membrane–bound TLR were used: LPS (TLR-4), LTA (TLR-2), and poly(I:C) (TLR-3), respectively (33). IL-12/IL-18–primed macrophages were treated with LPS, LTA, or poly(I:C) for 24 h and IFN-γ production was measured. Interestingly, IL-12/IL-18–primed macrophages exhibited significantly enhanced production of IFN-γ following TLR-2, TLR-3, and TLR-4 stimulation compared with the IL-12/IL-18–primed macrophages alone (Fig. 2B).

Subsequently, we determined if M1, M2a, M2b, and M2c macrophages, generated following stimulation with IFN-γ, IL-4, LPS + IL-1β, and IL-10, respectively, would produce IFN-γ on their own and without any TLR stimulation. For this, M1, M2a, M2b, and M2c subsets were generated as above. The polarizing stimulus was then removed after 48 h, and the cells were washed thrice and cultured with fresh medium for an additional 24 h followed by quantification of IFN-γ in the culture supernatants. The results show that M1 macrophages produced significantly high levels of IFN-γ compared with the M2a, M2b, and M2c macrophages, which produced basal levels of IFN-γ (Fig. 2C). IFN-γ production by M1 macrophages is not just leftover IFN-γ from the initial stimulation as cells were washed thoroughly prior to IFN-γ quantification.

Because M1 macrophages are generated following IFN-γ stimulation and IL-12/IL-18 priming induces IFN-γ production in human macrophages, we determined whether M1, M2a, M2b, and M2c macrophages produce IFN-γ following stimulation with TLR-2, TLR-3, or TLR-4 agonists. M1 macrophages exhibited significantly enhanced IFN-γ production following stimulation with TLR-2, TLR-3, or TLR-4 agonists (Fig. 2D). In contrast, M2a, M2b, and M2c subsets did not produce significant levels of IFN-γ following stimulation with the TLR agonists tested (Fig. 2E–G). To determine if M1 macrophages from patients with inflammatory diseases such as COVID-19 produce IFN-γ similar to the normal healthy M1 macrophages, we show that M1 macrophages generated from COVID-19 patients produced significantly high levels of IFN-γ compared with the M0 or LPS-stimulated M0 macrophages. Moreover, LPS stimulation of COVID-19 M1 macrophages led to significantly high IFN-γ production compared with the COVID-19 M1 macrophages alone (Fig. 2H). We also validated the ability of M1 macrophages to induce IFN-γ mRNA transcription by quantitative real-time PCR in contrast to M2a macrophages stimulated with TLR-2, TLR-3, and TLR-4 ligands (Fig. 3).

Because LPS stimulation of M1 macrophages induced high levels of IFN-γ production, we focused on LPS-induced IFN-γ production by M1 macrophages in subsequent studies. The kinetics of IFN-γ production by M1 macrophages after LPS stimulation revealed that enhanced production of IFN-γ is highest at 18 and 24 h after LPS stimulation (Fig. 4A). Intracellular staining for IFN-γ also revealed similar results with significant high levels of IFN-γ expression in M1 cells compared with M0 macrophages (8% versus 25%). Moreover, LPS-stimulated M1 macrophages expressed a significantly higher percentage of IFN-γ–expressing cells compared with the M1 macrophages alone (80% versus 25%) (Fig. 4B). A representative histogram is shown in (Fig. 4C. These results suggest that IL-12/IL-18–primed macrophages produce IFN-γ and IFN-γ induces its own production in M1 macrophages. Moreover, IL-12/IL-18–primed M0 macrophages and M1 macrophages, unlike M2a, M2b, and M2c macrophages, exhibit significantly enhanced production of IFN-γ following stimulation with TLR-2, TLR-3, and TLR-4 ligands.

Because mutations in N-terminal amino acid residues 1–10 and 14 aa at the carboxyl-terminal residues of the mature IFN-γ protein are critically involved in its biological activity (41, 42), we examined whether IFN-γ derived from M1 macrophages is biologically active. IFN-γ is known to upregulate the surface expression of TLR-4 in macrophages (43). Hence, we determined whether IFN-γ produced by M1 macrophages after LPS stimulation is capable of inducing TLR-4 expression in macrophages. For this, supernatants were collected from M1 macrophages left unstimulated or stimulated with LPS for 24 h. Newly untreated MDMs were treated for 2 h with anti-IFN-γR Ab or with isotype control IgG (40 μg/ml each) to block the functional activity of the endogenously produced IFN-γ. Macrophages were subsequently treated with culture supernatants collected previously from unstimulated or LPS-stimulated M1 macrophages followed by analysis of TLR-4 expression by flow cytometry. The results show that treatment with anti-IFN-γR Ab of macrophages with supernatants from unstimulated M1 macrophages (M1 media + anti-IFN-γR) significantly reduced TLR-4 expression compared with the IgG-treated macrophages stimulated with media from unstimulated M1 macrophages (M1 media + IgG; (Fig. 5, upper panels). Similarly, treatment of macrophages with culture supernatants from LPS-stimulated M1 macrophages (M1 LPS) or LPS-stimulated M1 macrophages along with control IgG showed similar levels of TLR-4 expression (Fig. 5, lower panels). However, treatment with anti-IFN-γR Ab of macrophages with supernatants from LPS-stimulated M1 macrophages (M1 LPS + anti-IFN-γR) significantly reduced TLR-4 expression compared with the untreated or IgG-treated macrophages stimulated with media from LPS-stimulated macrophages (Fig. 5B, lower panel). The representative histograms are shown in (Fig. 5A (lower panels).

To determine whether inhibition of M0 to M1 differentiation will inhibit LPS-induced IFN-γ production by M1 macrophages, M0 macrophages were pretreated with Jak-1 inhibitor, RUX, followed by IFN-γ stimulation to generate M1 macrophages. The biological activity of RUX was confirmed by demonstrating IFN-γ–induced Jak-1 phosphorylation in M0 macrophages that was inhibited by RUX (Fig. 6A). M0 macrophages were treated with RUX for 2 h prior to IFN-γ stimulation to inhibit M1 differentiation. After 48 h, macrophages were stimulated with LPS, followed by measurement of IFN-γ production. M1 macrophages stimulated with LPS secreted high concentrations of IFN-γ that was significantly inhibited by RUX (Fig. 6B). This reduction in IFN-γ concentration became more pronounced with increasing concentrations of the Jak-1 inhibitor.

We then investigated the signaling pathways involved in the LPS-induced IFN-γ production by M1 macrophages. The role of Erk, JNK, and p38 MAPK pathways was determined by employing their specific inhibitors PD-98059, SP-600125, and SB-203580, respectively (44). IFN-γ–differentiated M1 macrophages were treated with these inhibitors for 2 h prior to stimulation with LPS followed by IFN-γ measurement. Interestingly, p38 and JNK inhibitors significantly and completely inhibited LPS-induced IFN-γ production in M1 macrophages whereas Erk inhibitor PD-98059 did not have a significant effect (Fig. 7, left panels). The biological activity of these inhibitors was confirmed by inhibition of LPS-induced Erk, p38, and JNK phosphorylation in M1 macrophages (Fig. 7, right panels). We also show that LPS stimulation of M2c macrophages induced p38, Erk, and JNK phosphorylation that was inhibited by their respective inhibitors (Fig. 7, right panels).

The PI3K-Akt cascade was investigated by employing PI3K and Akt inhibitors, LY294002 and MK-2206, respectively. M1 macrophages were treated with LY294002 or MK-2206 for 2 h prior to stimulation with LPS for 24 h followed by IFN-γ measurement. The results show that LPS-induced IFN-γ production was significantly reduced by both inhibitors to the levels observed in M1 macrophages and in a dose-dependent manner (Fig. 8A, 8B). The activity of LY294002 was determined by showing inhibition of LPS-induced Akt activation in M1 macrophages (Fig. 8C). Furthermore, LPS stimulation of M2c macrophages induced Akt phosphorylation that was inhibited by PI3K inhibitor (Fig. 8D).

The mTOR protein complex is a serine/threonine kinase that acts downstream of phosphoinositide-dependent kinase-1 in the PI3K pathway and functions as a nutrient/energy/redox sensor and controls protein synthesis (45). mTOR links with other proteins and serves as a core component of two distinct protein complexes, mTORC1 and mTORC2, which regulate different cellular processes (45, 46). Rapamycin inhibits mTORC1 whereas torin inhibits both complexes (46, 47). To determine the involvement of mTORC1/2 pathway, M1 macrophages were treated with rapamycin or torin for 2 h prior to stimulation with LPS for 24 h followed by IFN-γ measurement as above. Rapamycin significantly reduced LPS-induced IFN-γ production in a dose-dependent manner (Fig. 9A, left panel) whereas torin significantly and completely inhibited LPS-induced IFN-γ production (Fig. 9B, left panel) by M1 macrophages. The biological activity of torin and rapamycin was confirmed by the Western blot analysis showing that M1 macrophages treated with rapamycin and torin exhibited reduced levels of LPS-induced p70S6K proteins compared with the LPS-stimulated macrophages in the absence of the inhibitor (Fig. 9A, 9B, right panels).

Because p70S6K is a downstream target of mTOR (48), we investigated the role of p70S6K in the LPS-induced IFN-γ production by M1 macrophages. As above, M1 macrophages were treated with the p70S6K inhibitor (PF-4708671) prior to LPS stimulation followed by IFN-γ measurement. M1 macrophages treated with the p70S6K inhibitor significantly inhibited LPS-induced IFN-γ production compared with the M1 macrophages stimulated with LPS alone (Fig. 9C, left panel). The biological activity of the p70S6K inhibitor was confirmed by the Western blot analysis showing inhibition of the p70S6K activity compared with the macrophages stimulated with LPS alone (Fig. 9C, right panel). Moreover, LPS stimulation of M2c macrophages induced p70S6K activity that was inhibited by PF-4708671, the p70S6K inhibitor (Fig. 9C).

The above results suggest that LPS-induced IFN-γ production by M1 macrophages is regulated by multiple pathways, including p38 MAPK and JNK MAPK, PI3K, mTOR, and S6K. To determine whether these pathways are interconnected and merge at the p70S6K enzymatic junction or run parallel to each other, M1 macrophages were pretreated with 25 and 50 µM of the PI3K inhibitor (LY294002) or the JNK inhibitor (SP-600125), followed by stimulation with LPS. The results show that LY294002 and SP-600125 inhibited the phosphorylation of p70S6K compared with the controls (Fig. 10) suggesting that LPS-induced IFN-γ production is regulated by the p70S6K pathway through the activation of JNK MAPK and the PI3K/Akt pathway.

Because our results indicated multiple signaling pathways intersecting at the S6K junction, we then inhibited the activation of S6K to assess if this downstream protein could cross-regulate the upstream p38, JNK, and Jak/STAT pathways. M0 macrophages were treated with p70S6K inhibitor for 2 h followed by IFN-γ stimulation for 30 min. As expected, the levels of p70S6K decreased with increasing concentrations of S6K inhibitor compared with the controls (Fig. 11A). However, the p70S6K inhibitor did not decrease the expression of phosphorylated Jak-1 (Fig. 11A). Similarly, M1 macrophages were pretreated with p70S6K inhibitor for 2 h followed by LPS stimulation for 30 min. The p70S6K inhibitor did not decrease the expression of phosphorylated p38 MAPK or JNK MAPK (Fig. 11B). Overall, although inhibiting upstream cell signaling proteins resulted in a decreased level of p70S6K, inhibiting p70S6K did not feed back to inhibit the activation of the upstream proteins, suggesting that LPS-induced IFN-γ production by M1 macrophages is regulated by the p70S6K through the activation of JNK MAPK and PI3K/Akt pathways (Fig. 12).

In this study, we show that human macrophages primed by inflammatory mediators IL-12/IL-18 and M1 macrophages generated from healthy individuals and from patients with inflammatory COVID-19 disease secrete IFN-γ spontaneously as well as following activation with TLR-2, TLR-3, and TLR-4 ligands. This reflects a broad range of bacterial/viral products, which significantly enhanced IFN-γ production in M1 compared with the M0, M2a, M2b, and M2c macrophages activated with the same ligands. Our studies regarding the signaling pathway(s) responsible for LPS-induced IFN-γ production in M1 macrophages revealed that LPS-induced IFN-γ production is regulated by the p70S6K pathway through the activation of multiple signaling pathways including PI3K-mTORC1/2 and JNK MAPKs. These findings suggest that IFN-γ produced by M1 macrophages and IL-12/IL-18–primed M0 macrophages may contribute to inflammation that may be further augmented following bacterial/viral infections, an increasingly emerging theme in the field with clinical relevance to acute and chronic inflammatory diseases. Moreover, targeting of mTOR, PI3K, and JNK MAPKs may be of potential translational significance to prevent the macrophage-associated inflammatory disorders.

Phenotypic characterization of macrophage subsets generated from human MDMs revealed distinct expression of a set of cell surface markers. The proinflammatory M1 macrophages expressed higher levels of well-established CD80, CD86, and HLA-DR compared with the undifferentiated M0 macrophages (49). CD80 and CD86 play a role in the differentiation of Th1 and Th2 cells and generation of immune responses (50). Moreover, M1 macrophages in contrast to M2a, M2b, and M2c subsets were distinguishable by secretion of IFN-γ spontaneously as well as following stimulation with TLR ligands. In contrast, anti-inflammatory M2a and M2c macrophages exhibited a distinctly higher CD200R expression. CD200–CD200R interactions deliver negative regulatory signals in endothelial cells and can inhibit macrophage activation (11). Moreover, CD200 homologs such as herpesvirus 8K14 protein (51), CMV e127 (52) protein, and myxoma virus M141R (53) in the Herpesviridae and Poxviridae families (54) can inhibit macrophage activation resulting in the dampening of immune responses and viral pathogenesis. Moreover, M2a and M2c macrophages could be distinguished from M1 macrophages by their lower expression of CD80. Similar to CD200R, M2b and M2c macrophages characteristically exhibited low levels of HLA-DR that may also dampen the immune response observed in stress, such as trauma, burns, surgery, and sepsis (55). In addition, M2c macrophages express high levels of CD163 that highlights their role in preventing tissue inflammation (56, 57).

In innate or adaptive immunity, differentiated hematopoietic cells limit tissue damage caused by inflammation and infection. This is accomplished by fine tuning the immune response influenced by the cytokine milieu to achieve a polarized state of macrophages that can effectively contribute to the re-establishment of homeostasis (58). Macrophage polarization into proinflammatory M1 and anti-inflammatory M2 is acquired following a change in type, timing, or amount of microenvironmental signals. These states are flexible and develop quickly to respond to environmental changes (9, 59). Moreover, macrophages provide a rapid immune response by recognizing TLRs present on invading pathogens (2, 60). Herein, we stimulated polarized macrophages with agonists against two cell surface–bound TLRs (TLR-2 and TLR-4) and against one intracellular membrane–bound TLR (TLR-3) that reflect the broad range of microbial ligands and thus were used to measure IFN-γ production.

IFN-γ plays a profound role in determining the effectiveness of the innate and adaptive immunity (25, 61). It is primarily produced by CD4⁺ and CD8⁺ T cells, γ/δ T cells, NK cells, and NKT cells spontaneously, following receptor-mediated and cytokine-mediated pathways (62). There are reports showing IFN-γ production by murine macrophages following stimulation with IL-12 and IL-18, and LPS-stimulated peritoneal macrophages (26–29). Although IFN-γ production by human alveolar macrophages with pulmonary sarcoidosis and subsequent in vitro infection of macrophages with M. tuberculosis has been shown (30, 31), it is not clear whether human macrophages or their subsets under normal healthy conditions produce IFN-γ. In this study, to our knowledge for the first time, we show that IL-12/IL-18–primed human M0 macrophages and the M1 macrophages produce IFN-γ spontaneously. IL-12 and IL-18 synergize to promote IFN-γ production in T cells and NK cells (39, 40). We observed that stimulation of IL-12/IL-18–primed human M0 and M1 macrophages with TLR-2, TLR-3, and TLR-4 ligands further increased IFN-γ production in contrast to the similarly stimulated M0, M2a, M2b, and M2c macrophages. This suggests that IL-12 and IL-18 produced during the innate immune response can lead to IFN-γ production by macrophages and promote their differentiation into the M1 state in an autocrine manner, further contributing to their activation. In addition, interaction of IL-12/IL-18–primed macrophages or M1 macrophages with bacterial/viral products following infection may result in severe inflammation. It is notable that IFN-γ produced under our experimental conditions was entirely derived from macrophages as MDMs generated by macrophage-CSF followed by stimulation with either IL-12/IL-18 or IFN-γ were 100% pure without any contamination with T cells, B cells, or NK cells (data not shown). This was also confirmed by showing intracellular IFN-γ expression by flow cytometry in M1 macrophages. In addition, IFN-γ production by M1 macrophages, confirmed as inhibition of M1 differentiation by Jak-1 inhibitors, inhibited LPS-induced IFN-γ production. Because mutations in N-terminal amino acid residues 1–10 and 14 aa at the carboxyl-terminal residues (41, 42) of the mature IFN-γ protein are critically involved in its biological activity, we demonstrated that IFN-γ produced by macrophages is biologically active.

IFN-γ expression is regulated epigenetically, transcriptionally, and posttranscriptionally (63–68). The regulation of IFN-γ has been extensively studied in murine and human NK cells and T cells, both of which share the regulatory signaling pathways and the transcription factors involved in IFN-γ synthesis (65–68). NK cells and T cells can be stimulated to secrete IFN-γ through both receptor (TCR)- and cytokine (IL-12 and IL-18)-mediated pathways (62, 69, 70). The receptor-activated signaling is mediated through immunoreceptor tyrosine-based activating motifs, which phosphorylate members of the Src family of protein tyrosine kinases, the Ras/MAPK, PI3K/Akt, and phospholipase-C-γ (71, 72). Src activation then leads to the activation of MAPKs such as Erk and p38 MAPKs, which in turn activate transcription factors such as Fos and Jun, leading to IFN-γ transcription (63, 73). Cytokine-mediated activation is regulated by IL-12 produced by macrophages and other APCs following infection (74). IL-12 interacts with its receptors on these cells leading to the activation of Jak-STAT-4 pathway and eventual activation of NF-κB (65, 66). IFN-γ is also regulated by noncoding RNAs including several microRNAs (75, 76). IFN-γ is also known to be regulated posttranscriptionally. For example, several cytokines including IL-12 stabilize IFN-γ transcripts in T cells through a p38 MAPK–dependent pathway (65, 66).

The molecular mechanism by which IL-12 and IL-18 induce IFN-γ expression in murine macrophages or IFN-γ induces its own expression in human macrophages is not known. Herein, we investigated the mechanism regulating IFN-γ expression in M1 macrophages following LPS stimulation. We show for the first time, to our knowledge, that LPS-induced IFN-γ production by M1 macrophages is regulated by multiple signaling pathways including p38 MAPK and JNK MAPK, mTOR-PI3K/Akt, and S6K. Furthermore, LPS-induced IFN-γ production in M1 macrophages is regulated by p70S6K through the activation of JNK MAPKs and mTOR-Akt/PI3K pathways, and is independent of the Jak/STAT pathway responsible for the differentiation of M0 macrophages into M1 macrophages. Interestingly, although M2c macrophages stimulated with LPS induced the activation of the same signaling pathways including p38 MAPK, JNK MAPKs, PI3K/Akt, and p70S6K, they did not lead to IFN-γ production following LPS stimulation. These results suggest that M1 macrophages acquire a unique IFN-γ–activated signaling phenotype during M1 differentiation responsible for IFN-γ production spontaneously as well as following LPS stimulation. However, the mechanism by which IFN-γ is regulated in M1 macrophages epigenetically, transcriptionally, or posttranslationally remains unknown. The promoter and gene sequences for IFN-γ have been characterized in T cells and NK cells. Binding sites for many transcription factors have been located in these regulatory elements, including Fos, Jun, GATA3, NFAT, NF-κB, and T-bet (65, 66, 77–79). Whether the same transcription factors regulate IFN-γ gene expression in LPS-activated M1 macrophages requires investigation.

Although recent studies involving genome-wide gene expression profiling suggest a multidimensional model of macrophage activation rather than the M1/M2 model (80–83), our novel findings show that IFN-γ–differentiated M1 macrophages are a significant source of IFN-γ that may be further enhanced following bacterial/viral infections, setting it apart from the M2 counterparts. This may be exemplified in several human diseases wherein a shift toward the proinflammatory M1 environment is associated with immune-mediated tissue damage and inflammation (8, 14–16, 18, 19, 82, 84). We show that M1 macrophages generated from COVID-19 patients produced significantly higher levels of IFN-γ compared with the M0 or LPS-stimulated M0 macrophages from the same patients. Moreover, LPS stimulation of M1 macrophages from COVID-19 patients significantly enhanced their IFN-γ production. Although blood monocytes from COVID-19 patients are exposed to a variety of pro- and anti-inflammatory cytokines, interestingly M1 macrophages generated from COVID-19 patients produced high levels of IFN-γ spontaneously as well as following LPS stimulation. In addition, IFN-γ production by human alveolar macrophages with pulmonary sarcoidosis and following in vitro infection of macrophages with M. tuberculosis has been shown (30, 31). We have also shown that macrophages derived from hepatitis C patients with stage F3-4 produced high levels of IFN- γ (32). Thus, targeting M1 cells via the p70S6K pathway through kinases such as PI3K, mTOR, p38 MAPK, or JNK MAPK and more specifically the expression of immune-modulating IFN-γ may help mitigate inflammation and reduce the burden of disease.

We thank the nurses for generous help in collecting blood samples. We also thank the healthy donors for providing their blood samples.

The authors have no financial conflicts of interest.