Immunoblotting was performed using antibodies against phospho-TBK1, TBK1, NLRP3, Caspase-1, phospho-p65, p65, phospho-IκB, IκB, phospho-S6K, S6K (Cell Signaling Technology, Danvers, MA, USA), Caspase-1 (Santa Cruz Biotechnology, Dallas, TX, USA), and α-tubulin (Developmental Studies Hybridoma Bank, Iowa City, IA, USA). LPS (Escherichia coli LPS, serotype 0127: B8), ATP, and rapamycin were purchased from Sigma-Aldrich (St. Louis, MO, USA). Amlexanox and Torin 1 were purchased from the Cayman Chemical Company, Ann Arbor, MI, USA. The sources of other reagents are indicated in the specified methods.

Female C57BL/6 mice (6–8-weeks-old) were purchased from Samtako (Osan, Korea). All mice were maintained under a controlled temperature (20–25°C) and humidity (50 ± 5%) with a 12 h light/dark cycle and free access to food and drinking water. Female mice were mated with male mice, and the presence of vaginal plugs the following day confirmed successful mating. The timed pregnant mice received an intraperitoneal injection of LPS (200 µg/kg) or saline on gestation day 17.5. In the amlexanox + LPS group, the pregnant mice were intraperitoneally injected with amlexanox (2.5 mg/kg) at 27 h and 3 h before the intraperitoneal administration of LPS. The pregnant mice were sacrificed at various time points (4, 6, 8, 10, or 12 h) after LPS treatment, and placental samples were collected for molecular analysis. All animal studies were conducted following the Guidelines for the Care and Use of Laboratory Animals of the National Institutes of Health and approved by the Animal Ethics Committee of Konyang University (P-21-19-A-01).

Human first-trimester trophoblast cell lines, HTR-8/SVneo (American Type Culture Collection, Rockville, MD, USA) and Sw.71 (gifted by Dr. Gil Mor, Yale University School of Medicine, New Haven, CT, USA), were used in our experiments. HTR-8/SVneo cells were cultured in RPMI 1640 medium (Welgene, Gyeongsan, Korea) supplemented with 10% fetal bovine serum (Welgene), 4500 mg/L D-glucose, 10 mM HEPES, 2mM L-glutamine, 1 mM sodium pyruvate, and 100 U/mL penicillin-streptomycin (Welgene). The Sw.71 cells were cultured in Dulbecco’s modified Eagle’s medium (Welgene) containing 10% fetal bovine serum, 4500 mg/L D-glucose with L-glutamine, and 100 U/mL penicillin-streptomycin. All cultures were maintained in a humidified 5% CO₂ atmosphere at 37°C. For LPS treatment, cells were incubated in the presence of LPS or treated with LPS and then treated with 5 mM ATP for 45 min, as described previously (17). Identical volumes of phosphate-buffered saline (PBS) were used as the vehicle controls. When indicated, the cells were incubated with the TBK1 inhibitor amlexanox.

The HTR-8/SVneo cell survival was measured using the WST-1 assay (EZ-CytoX Enhanced Cell Viability Assay Kit, Daeil Lab Service, Seoul, Korea) according to the manufacturer’s protocol (25). Briefly, the HTR-8/SVneo cells (1 × 10⁴ cells/well) were plated into 96-well plates and treated with the indicated concentrations of amlexanox for 24 h. Subsequently, 10 μL of WST-1 reagent was added to each well and incubated for 30 min at 37°C in a 5% CO₂ incubator (Thermo Scientific, Waltham, MA, USA). The absorbance was measured at 450 nm using an Epoch 2 microplate reader (Bio-Tek Instruments, Winooski, VT, USA), and the absorbance values of the treated cells were expressed as a percentage of the absorbance values of the control.

Using a polyethylenimine reagent, HEK293T cells were transfected with the following lentiviral constructs with the packaging plasmids: sh-Luciferase (sh-Luc) and sh-TBK1 constructs (obtained from the RNAi Consortium collection of the Broad Institute). Lentiviral supernatants were collected and filtered 48 and 72 h after transfection. HTR-8/SVneo cells and Sw.71 cells were incubated 2–3 d with the lentiviral medium in the presence of 4 μg/mL polybrene.

Total RNA was extracted from placental tissues and HTR-8/SVneo cells using TRIzol reagent (Takara, Shiga, Japan) according to the manufacturer’s instructions. Complementary DNA was synthesized from RNA using a cDNA synthesis kit (BioFact, Daejeon, Korea). Quantitative real-time reverse transcription PCR (qRT-PCR) was performed in triplicate with the SYBR Green Real-Time PCR Master Mix Reagent (BioFact) using the QuantStudio 3 Real-time PCR System (Life Technologies, Carlsbad, CA, USA). Relative mRNA expression was calculated using comparative threshold cycle (C_t) values normalized to those of mouse or human cyclophilin. The following primers were used: mouse Nlrp3: forward 5′- AGCCTTCCAGGATCCTCTTC-3′, reverse 5′-CTTGGGCAGCAGTTTCTTTC-3′; mouse Tnf-α: forward 5′-TCCCAGGTTCTCTTCAAGGGA-3′, reverse 5′-GGTGAGGAGCACGTAGTCGG-3′; mouse Tgf-β1: forward 5′-CTCCCGTGGCTTCTAGTGC-3′, reverse 5′-GCCTTAGTTTGGACAGGATCTG-3′; mouse Il-6: forward 5′- TAGTCCTTCCTACCCCAATTTCC-3′, reverse 5′-TTGGTCCTTAGCCACTCCTTC-3′; mouse Il-10: forward 5′-GCTCTTACTGACTGGCATGAG-3′, reverse 5′-CGCAGCTCTAGGAGCATGTG-3′; mouse Mcp-1: forward 5′-CATCCACGTGTTGGCTCA-3′, reverse 5′-GATCATCTTGCTGGTGAATGAGT-3′; mouse Il-1β: forward 5′-TCTTTGAAGTTGACGGACCC-3′, reverse 5′-TGAGTGATACTGCCTGCCTG-3′; mouse IFN-β: forward 5′-GCACTGGGTGGAATGAGACT-3′, reverse 5′-AGTGGAGAGCAGTTGAGGAC-3′; mouse Cyclophilin A: forward 5′-GAGCTGTTTGCAGACAAAGTTC-3′, reverse 5′-CCCTGGCACATGAATCCTGG-3′; human NLRP3: forward 5′-GATCTTCGCTGCGATCAACAG-3′, reverse 5′-CGTGCATTATCTGAACCCCAC-3′; human IL-1β: forward 5′-AAAGAGGCACTGGCAGAA-3′, reverse 5′-AGCTCTGGCTTGTTCCTCAC-3′; and human Cyclophilin A: forward 5′-GCAAAGTGAAAGAAGGCATGAA-3′, reverse 5′-CCATTCCTGGACCCAAAGC-3′.

Placental tissues, HTR-8/SVneo cells, and Sw.71 cells were lysed in an ice-cold radioimmunoprecipitation assay buffer containing cOmplete protease inhibitor cocktail (Roche, Basel, Switzerland). Lysates were incubated for 20 min on ice and centrifuged at 18,000 × g for 15 min at 4°C. Protein concentrations were measured using a bicinchoninic acid protein assay (Pierce, Rockford, IL, USA). Lysates were boiled in 1× sodium dodecyl sulfate (SDS) and Laemmli sample buffer for 5 min. Proteins were separated by SDS-polyacrylamide gel electrophoresis and transferred to polyvinylidene fluoride membranes (Millipore, Burlington, MA, USA), which were probed with primary antibodies against p-TBK1, TBK1, NLRP3, caspase-1, p-p65, p65, p-IκBα, IκBα, p-S6K, S6K, and α-tubulin. After incubation with secondary antibodies conjugated with horseradish peroxidase (Bio-Rad, Hercules, CA, USA), chemiluminescence was detected using a Fusion Solo System (Vilber Lourmat, Marne-la-Vallée, France). Densitometric analysis of the blots was performed using ImageJ (National Institutes of Health, Bethesda, MD, USA), in which the background was removed for each band.

Mouse placental tissues were fixed in 10% neutral buffered formalin for 24 h, dehydrated, and embedded in paraffin. Sections (5 μm) of the embedded tissue were prepared and stained with hematoxylin and eosin dyes or periodic acid-Schiff dyes. For immunohistochemistry, the paraffin-embedded sections were deparaffinized, rehydrated, and treated for antigen retrieval. Endogenous peroxidase was quenched using 3% hydrogen peroxide. After blocking the nonspecific antigens, the sections were incubated with anti-NLRP3 antibody or anti-IL-1β antibody overnight at 4°C, followed by incubation with biotinylated secondary antibody (Vector Laboratories, Burlingame, CA, USA). Antibodies were visualized with streptavidin-HRP (BD Pharmingen, San Diego, CA, USA) using diaminobenzidine (Sigma-Aldrich). Hematoxylin was used to visualize nuclei. The samples were analyzed under a light microscope (DM2500; Leica, Wetzlar, Germany).

Cell supernatants collected from the HTR-8/SVneo cells were assayed for the levels of the secreted inflammatory cytokine IL-1β using the DuoSet ELISA development kit (R&D Systems, Minneapolis, MN, USA) according to the manufacturer’s instructions. The optical density of the final colored reaction product was determined at 450 nm using an Epoch 2 microplate reader (Bio-Tek Instruments).

Results are presented as the mean ± standard error of the mean (SEM). Data presented in the figures are representative of at least three independent experiments unless stated otherwise. The significance of differences between the two experimental groups was determined using a two-tailed Student’s t-test. Multiple comparisons were conducted using one-way ANOVA followed by Tukey’s or Fisher’s least significant difference (LSD) post hoc test. Differences were considered statistically significant at p < 0.05, *p < 0.05, **p < 0.01, ***p < 0.001).

We evaluated the phosphorylation of TBK1 protein in the mouse placenta by LPS-induced maternal inflammatory response. Immunoblot analysis showed that the placental TBK1 phosphorylation was significantly increased at 4, 6, 8, and 10 h after LPS treatment ( Figures 1A, B ). To measure the phosphorylated levels of TBK1 in trophoblasts during maternal inflammation, human first-trimester extravillous trophoblasts (HTR-8/SVneo and Sw.71) were stimulated with various LPS concentrations at various time points. Immunoblot analysis showed that LPS increased TBK1 phosphorylation in HTR-8/SVneo after both short- ( Figures 1C, D ) and long-term treatments ( Figures 1E, F ) in a dose-dependent manner. The time course of TBK1 phosphorylation was further investigated by treating the cells with LPS (1 μg/mL), and the results showed that short- and long-term exposure to LPS significantly increased phosphorylated TBK1 levels in HTR-8/SVneo cells in a time-dependent manner ( Figures 1G–J ). Similar to that in HTR-8/SVneo cells, the expression of phosphorylated TBK1 was stimulated by LPS treatment in Sw.71 cells ( Supplemental Figure 1 ).

NLRP3 inflammasome activation in trophoblasts is implicated in the pathogenesis of placental inflammation (17, 26). To investigate the effects of LPS on placental NLRP3 protein expression, placentas from mice exposed to LPS were analyzed by immunohistochemistry. The results showed that NLRP3 protein expression was increased in trophoblasts in the junctional zone and the labyrinth of the placentas of the LPS-treated mice compared with that of the control placentas of saline-treated mice ( Supplemental Figure 2 ). Consistent with the above results, the NLRP3 inflammasome was activated in HTR-8/SVneo and Sw.71 cells upon stimulation with LPS and ATP ( Supplemental Figures 3A, B ). Furthermore, we investigated whether TBK1 is involved in LPS-induced NLRP3 inflammasome activation. Amlexanox, a TBK1 inhibitor, was used to explore the effect of TBK1 deficiency on LPS-induced NLRP3 inflammasome. We measured the cell viability of HTR-8/SVneo and Sw.71 cells treated with different concentrations of amlexanox. In vitro cytotoxicity assays revealed that amlexanox exhibited low cytotoxicity against cells up to 200 µM ( Supplemental Figures 4A, B ). Immunoblot analysis showed that amlexanox markedly decreased LPS-induced NLRP3 mRNA levels and NLRP3 and cleaved caspase-1 protein levels ( Figures 2A–C and Supplemental Figure 3C ). To confirm these results, IL-1β levels were analyzed using ELISA. Pretreatment of HTR-8/SVneo cells with amlexanox resulted in significantly lower levels of IL-1β compared to that in LPS-treated cells ( Figure 2D ).

To clarify the specific signaling mechanism by which amlexanox suppresses LPS-induced NLRP3 inflammasome activation in trophoblasts, lentiviral vectors expressing small hairpin RNAs (shRNAs) targeting human TBK1 were transduced to HTR-8/SVneo and Sw.71 cells. Knockdown of TBK1 was sufficient to block the upregulation of NLRP3 and cleaved caspase-1 in LPS-treated cells ( Figures 2E–G and Supplemental Figure 4D ), consistent with the inhibitory effect of amlexanox on NLRP3 inflammasome activation. ELISA showed that the lentiviral knockdown of TBK1 also decreased IL-1β production ( Figure 2H ), suggesting that it is an inhibitory effect of TBK1, not a nonspecific effect of amlexanox.

NF-κB signaling mediates the priming signal of NLRP3 inflammasome activation and upregulates the expression levels of NLRP3 and pro-IL-1β (27, 28). LPS increased the phosphorylation of NF-κB/p65 in a time-dependent manner, and the maximal effect was observed at 1 h following 1 μg/mL LPS treatment ( Supplemental Figure 5A ). To investigate the effect of amlexanox on the NF-κB pathway, we measured the phosphorylation levels of IκBα and NF-κB/p65 in HTR-8/SVneo and Sw.71 cells. Pretreatment with amlexanox decreased LPS-induced phosphorylation of IκBα and NF-κB/p65 ( Figures 3A, B , and Supplemental Figure 5B ). Further, we investigated the nuclear translocation of NF-κB/p65. LPS treatment resulted in a significant increase in NF-κB/p65 in the nucleus of HTR-8/SVneo cells, whereas the nuclear translocation of NF-κB/p65 was suppressed by amlexanox ( Figure 3C ).

We also examined whether the knockdown of TBK1 reversed the effect of LPS on the NF-κB/p65 pathway. Lentiviral knockdown of TBK1 reduced LPS-induced phosphorylation of IκBα and NF-κB/p65 ( Figures 3D, E , and Supplemental Figure 4C ). Immunofluorescence staining showed that the knockdown of TBK1 was sufficient to inhibit the nuclear translocation of NF-κB/p65 in LPS-treated cells ( Figure 3F ). These results collectively indicate that TBK1 deficiency prevents the LPS-induced nuclear translocation of NF-κB/p65 by phosphorylation of IκBα.

To evaluate whether maternal administration of amlexanox affects the fetal-placental development of LPS-treated mice, we first measured the fetal and placental weight and length in each group. Fetal and placental weight and length were significantly reduced in the LPS-treated mice compared to that in the control mice, whereas amlexanox treatment prevented the decrease in fetal and placental weight and length associated with LPS treatment ( Figures 4A–C ). We also evaluated whether amlexanox affects placental growth and morphology in LPS-treated mice. Periodic acid-Schiff staining revealed that LPS-exposed placenta had significantly reduced length of decidual layers and junctional zones; however, no appreciable change was observed in the length of the labyrinth layer. Interestingly, amlexanox treatment abrogated the LPS-induced placental morphological changes ( Figures 4D, E ). These findings suggest that maternal administration of amlexanox alleviates LPS-induced morphological changes in the placenta.

To further confirm whether maternal administration of amlexanox influences placental NLRP3 inflammasome activation in vivo, we measured the expression levels of NLRP3 inflammasome markers in the placentas of LPS-treated mice. Immunoblot analysis showed that NLRP3 and cleaved caspase-1 protein levels were elevated in response to LPS treatment; this increase was suppressed in amlexanox-treated mice exposed to LPS ( Figures 5A, B ). Correspondingly, amlexanox treatment decreased NLRP3 mRNA levels in the LPS-treated mouse placenta ( Figure 5C ). Immunohistochemical analysis showed that NLRP3 and IL-1β staining was significantly increased in the cytoplasm of trophoblasts in the junctional zone and labyrinth layer in LPS-treated mouse placenta; however, their expression was decreased with amlexanox treatment ( Figure 5D ).

To determine whether the effect of amlexanox on the NF-κB pathway could be observed in vivo, we measured the phosphorylation levels of NF-κB/p65 in mouse placental tissues. Immunoblot analysis revealed that phosphorylation of NF-κB/p65 was increased, whereas amlexanox treatment prevented the increase in phosphorylation levels of NF-κB/p65 ( Figures 6A, B ). We also evaluated whether maternal administration of amlexanox affected gene expression of inflammatory cytokines and chemokines using qRT-PCR. The expression of IL-1β, IL-6, IL-10, TNF-α, TGF-β1, MCP-1, and IFN-β genes was elevated in the placenta of LPS-treated mice as compared to that of control mice; this increase was largely attenuated in amlexanox-treated mice exposed to LPS ( Figure 6C ). Taken together, these data indicate that TBK1 inhibition by amlexanox alleviates placental inflammation and NLRP3 inflammasome activation.

In response to LPS, TBK1 can directly activate mTORC1, but this function of TBK1 has been controversial (29, 30). Therefore, we checked the expression of downstream target proteins of mTORC1 in the placental tissues of control, vehicle-, and amlexanox-treated LPS mice. Immunoblot analysis indicated that mice treated with amlexanox had lower levels of phosphorylated p70 ribosomal protein S6 kinase (p70S6K) compared to that in LPS-treated mice ( Figures 7A, B ). Further, we evaluated the effect of amlexanox on LPS-induced mTORC1 activation in HTR-8/SVneo cells and found a decrease in the phosphorylation of p70S6K after pretreatment with amlexanox ( Figures 7C, D ). Consistent with the inhibitory effect of amlexanox on mTORC1 signaling, lentiviral knockdown of TBK1 blocked LPS-induced phosphorylation of p70S6K in HTR-8/SVneo cells ( Figures 7E, F ), indicating that TBK1 is involved in LPS-mediated mTORC1 activation. To investigate whether mTORC1 inhibition influences LPS-induced NLRP3 inflammasome activation, we pretreated HTR-8/SVneo cells with mTOR inhibitors, rapamycin and Torin 1. Immunoblot analysis revealed suppression of LPS-induced increase in NLRP3 and cleaved caspase-1 protein levels after pretreatment with rapamycin or Torin 1 ( Figures 7G–J ). Thus, these data suggest that TBK1 deficiency alleviates LPS-induced NLRP3 inflammasome activation in an mTORC1-dependent manner.