TLR2 or TLR4 Stimulation Induces Transmembrane TNF-Driven Priming in Macrophages Which Results in Improved Clearance of a Subsequent Staphylococcus aureus Infection Open Access

Macrophage recognition of microbes or their associated products, termed pathogen-associated molecular patterns (PAMPs) occurs via ligation to host pattern recognition receptors (PRRs) [1, 2]. One class of PRRs that recognizes PAMPs are toll-like receptors (TLRs). TLRs encompass a group of evolutionarily conserved transmembrane receptors capable of recognizing PAMPs both extracellularly and intracellularly [3, 4]. Two particularly important TLRs for extracellular PAMP recognition are TLR2 and TLR4, which are known to induce inflammation in response to a wide range of bacteria, including gram-negative and gram-positive microbes as well as mycobacteria [3‒5]. PAMP ligation to TLR2 or TLR4 on immune cells is critical for initiating intracellular signaling cascades which promote host defense via inflammation [6‒8]. During this inflammatory response, tumor necrosis factor alpha (TNF) is one of the first and most robustly produced cytokines, and it plays a major role in perpetuating the inflammatory host response [9, 10].

TNF is a pleiotropic cytokine that exists in two bioactive forms. The 26 kDa tmTNF is the precursor protein anchored to the cellular membrane. Due to this anchoring, tmTNF requires cell-to-cell contact to signal effectively. This limits tmTNF signaling outcomes to nearby cells [11, 12]. The metalloprotease ADAM Metallopeptidase Domain 17 (ADAM17) cleaves tmTNF to produce the mature soluble form of TNF (sTNF), a 17 kDa protein that is able to exert distal systemic effects [11, 12]. Both forms of TNF are capable of binding either of the two TNF receptors (TNFR1 or TNFR2). However, the inability of sTNF to efficiently signal through TNFR2 makes TNFR1 the primary receptor for sTNF [13, 14]. Broadly, ligation of TNFR1 by sTNF leads to classical pro-inflammatory responses via activation of canonical NFκB or MAPK signaling pathways [12, 15‒17]. Conversely, TNFR2 is thought to be the primary receptor for tmTNF. Although tmTNF is also capable of signal transduction via TNFR1, tmTNF displays higher binding avidity for TNFR2 [12‒14, 18]. Ligation of TNFR2 leads to processes involved in cellular proliferation via activation of noncanonical NFκB or PI3K/AKT [19, 20]. Several studies suggest that differences in cellular expression between TNFR1 and TNFR2 could contribute to distinct signaling effects by sTNF and tmTNF [21‒23]. Unlike TNFR1, which is ubiquitously expressed across cell types, TNFR2 expression is primarily found on immune cells [19, 22, 24, 25]. Furthermore, TNFR2 is a strictly inducible receptor, whereas TNFR1 is constitutively expressed [19, 22, 24, 25]. Together, the different affinities of TNF isoforms to TNFR1 and TNFR2 and the differential TNFR expression patterns allow for functional pleiotropy of TNF-induced inflammation.

Here, we hypothesize that tmTNF expressed as a result of TLR stimulation may be critical for macrophage priming, which leads to increased clearance of a subsequent Staphylococcus aureus infection by primed macrophages. The primary goals of this study were to (1) characterize both forms of TNF within our cell culture system and (2) determine whether/which bioactive form of TNF is required for macrophage priming that leads to improved clearance of S. aureus.

Although effects of signaling by either bioactive form of TNF induced during infection have been characterized, not much is known regarding whether and how sTNF and tmTNF induced prior to infection contribute to innate priming. A better understanding of the function of TNF following TLR stimulation (PAMP recognition) in macrophages could provide novel insights into the early key signaling events that could improve macrophage response to infection.

Six-to-eight-week-old female wild type (C57BL/6) or TNF−/− mice were purchased from Jackson Laboratory. Mice were maintained at the Montana State University (MSU) Animal Resource Center. All care and procedures were in accordance with the recommendations of the National Institutes of Health, the US Department of Agriculture, and the Guide for the Care and Use of Laboratory Animals (8th ed) [26]. Animal protocols were reviewed and approved by the MSU Institutional Animal Care and Use Committee (IACUC). For RNA sequencing experiments, wild-type (WT) mice were intratracheally inoculated with either 10 μg of PAM2CSK4 or PAM3CSK4 (Invivogen) in 100 μL volume, equal volume sterile PBS, or were left untreated (naïve). At 6 h posttreatment (0 h for naïve), mice were euthanized, the lungs were extracted, cut into small pieces (<0.5 cm) and were immediately stored in RNA-later (Thermofisher). RNA was extracted from the lungs by homogenizing the lung pieces with a TissueRuptor II (Qiagen) in 3 mL of TRIzol Reagent (Invitrogen) with TissueRuptor disposable probes. Following homogenization, RNA was extracted from half of the sample (other half flash frozen). Briefly, 0.3 mL chloroform was added to the 1.5 mL of TRIzol homogenate, vortexed, incubated 3 min, and the aqueous phase was collected into a new tube following a 15-min centrifugation at 12,000 g at 4°C. Equal volume of 70% ethanol was added to the aqueous layer, mixed, and added to a RNeasy spin column. Following the Qiagen RNeasy protocol, the column was spun, flow through discarded, washed 2× with RPE Buffers. The column was dried with a brief centrifugation and the RNA was eluted. The quality of RNA was measured for each sample using a Bioanalyzer (Agilent) and only RNA samples with a RIN number >9.0 and OD260/280 >2.0 were sent off for sequencing. Processing of RNA for library generation and sequencing on an Illumina Platform PE150 instrument was done by NovoGene Corporation (Sacramento, CA, USA).

Downstream analysis was performed by NovoGene Corporation and their protocol is as described below. A combination of programs including STAR (v2.6.1), HTseq (v0.6.1), Cufflink and NovaGene Corporation wrapped scripts were utilized. Alignments were parsed using STAR program and differential expressions were determined through DESeq2 (v2_1.6.3)/edgeR (v3.16.5). Gene fusion and difference of alternative splicing event were detected by Star-fusion and rMATS software. Reference genome and gene model annotation files were downloaded from genome website browser (NCBI/UCSC/Ensembl) directly (GRCm38 Mus Musculus genome). Indexes of the reference genome was built using STAR and paired-end clean reads were aligned to the reference genome using STAR (v2.5). STAR used the method of Maximal Mappable Prefix which can generate a precise mapping result for junction reads. STAR will count number reads per gene while mapping. The counts coincide with those produced by htseq-count with default parameters. Fragments per kilobase of exon per million mapped fragments of each gene were calculated based on the length of the gene and reads count mapped to this gene. Fragments per kilobase of exon per million mapped fragments considers the effect of sequencing depth and gene length for the reads count at the same time, and is currently the most commonly used method for estimating gene expression levels [27]. For DESeq2 with biological replicates, differential expression analysis between two conditions/groups (two biological replicates per condition) was performed using the DESeq2 R package (1.14.1). DESeq2 provide statistical routines for determining differential expression in digital gene expression data using a model based on the negative binomial distribution. The resulting p values were adjusted using the Benjamini and Hochberg’s approach for controlling the False Discovery Rate (FDR). Genes with an adjusted p value <0.05 found by DESeq2 were assigned as differentially expressed.

For edgeR without biological replicates, prior to differential gene expression analysis, for each sequenced library, the read counts were adjusted by edgeR program package through one scaling normalized factor. Differential expression analysis of two conditions was performed using the edgeR R package (3.16.5). The p values were adjusted using the Benjamini & Hochberg method. Corrected p value of 0.05 and absolute fold change of 1 were set as the threshold for significantly differential expression.

RNA sequencing data received from NovoGene Corporation analysis was organized and plotted using Linux and R provided by the National Center of Genomics Research (NCGR; Santa Fe, NM, USA). Pathway analysis performed using Cytoscape and ClueGo software (access provided by NM-INBRE and the National Center of Genomics Research).

Isolation of primary BMDMs was collected from WT C57BL6 or TNF−/− mice purchased from Jackson Laboratory. Mice were euthanized according to IACUC protocol and both pairs of femurs, tibias, and fibulas were collected per mouse and bones were soaked in 70% ethanol for 5 min before bone marrow was flushed from the bones with 1X Roswell Park Memorial Institute 1640 (RPMI; Cytiva, Marlborough, MA, USA) supplemented with 10% heat-inactivated fetal bovine serum (FBS; Atlas Biologicals, Fort Collins, CO, USA), 0.1 m HEPES (Fisher Scientific, Waltham, MA, USA), 10,000 U/mL penicillin-streptomycin (Fisher Scientific, Waltham, MA, USA), 4 mg/mL glutamine (Gibco, Grand Island, NY, USA), and 1.792 × 10⁻³ mM 2-Mercaptoethanol (Fisher Scientific, Waltham, MA, USA). Bone marrow was centrifuged at 408 g for 5 min at 4°C and subsequently incubated with ACK lysis buffer (Fisher Scientific, Waltham, MA, USA) for 4 min at room temperature. Bone marrow cells were centrifuged, resuspended, and differentiated into macrophages in 1X RPMI1640 supplemented with 10% heat-inactivated FBS, 0.1 m HEPES, 10,000 U/mL penicillin-streptomycin, 4 mg/mL glutamine, 1.792 × 10⁻³ mM 2-Mercaptoethanol, and L929 conditioned media for 6 days at 37°C with 5% CO₂.

Immortalized bone marrow macrophages derived from WT, TLR2^−/−, and TLR6^−/− mice on a C57BL/6 background (NR-9456, NR-9457, and NR-19972) were obtained from BEI Resources, NIAID, NIH. Cells were maintained in Dulbecco’s modified Eagles medium (Cytiva, Marlborough, MA, USA) supplemented with 10% heat-inactivated fetal bovine serum (FBS; Atlas Biologicals, Fort Collins, CO, USA), 2 mm glutamine (Gibco, Grand Island, NY, USA), 1 mm sodium pyruvate (Gibco, Grand Island, NY, USA), and 10 µg/mL ciprofloxacin (Bioworld, Dublin, OH, USA) and cultured at 37°C with 5% CO₂ until 90% confluency was reached.

S. aureus (USA300 LAC) was a kind gift from Dr. Jovanka Voyich, Department of Microbiology and Cell Biology at MSU. BBL Trypticase soy broth (TSB; BD, Franklin Lakes, NJ, USA) was inoculated with frozen S. aureus and grown overnight at 37°C with 250 rpm shaking. The following morning, fresh TSB was inoculated with the overnight culture 1:100 and cultured for 2 h 15 min at 37°C with 250 rpm shaking to achieve a day culture of S. aureus. One mL of day culture or uninfected TSB (blank) was transferred to a cuvette and OD₆₀₀ was determined. Bacteria were then diluted to appropriate concentration based on growth curve.

Murine immortalized macrophage cell line NR-9456, NR-9457, or NR-19972 (BEI Resources; Manassas, VA, USA) were grown and maintained as previously described until 90% confluency was achieved. Cells were lifted with a sterile-filtered solution of 3 mm EDTA (MilliporeSigma; Burlington, MA, USA) dissolved in dPBS (Cytiva, Marlborough, MA, USA) for 10 min at 37°C with 5% CO₂. Cells were washed once in sterile dPBS, resuspended in complete media, and counted before being plated in 24-well plates at a cell density of 5 × 10⁵ cells/well and allowed to adhere for 12 h at 37°C with 5% CO₂. Cells were then treated for 12 h with 500 µL complete media containing either equal volume PBS, 1 µg/mL Pam2CSK4 (PAM2; InvivoGen, San Diego, CA, USA), 1 μg/mL Pam3csk4 (PAM3; InvivoGen, San Diego, CA, USA), or 1 μg/mL ultrapure lipopolysaccharide (LPS; InvivoGen, San Diego, CA, USA) with or without 100 μg/mL of p75TNFR:Fc, a dimeric fusion protein composed of two extracellular ligand binding domains of the 75 kDa receptor for TNF linked to the Fc fragment of murine IgG1, a kind gift from Amgen Inc., 1 μm TMI-005 (Cayman Chemical, Ann Arbor, MI, USA), or 1 μg anti-mouse TNFR2 antibody (BioXcell, Lebanon, NH, USA). After treatment, cells were washed once with dPBS and incubated with S. aureus at an MOI of 2:1 for 5 h at 37°C with 5% CO₂. Cell lysis was performed with 1% saponin (MilliporeSigma; Burlington, MA, USA), and CFUs were determined based on plating serial dilutions using the drop plate method onto tryptic soy agar [28] (TSA; Fisher Scientific, Waltham, MA, USA). Cellular supernatants before treatment, after treatment, and after S. aureus were collected into sterile Eppendorf tubes and frozen at −80°C for cytokine analysis.

Cells were seeded into 6 well plates at 2 × 10⁶ cells/well and allowed to adhere for 12 h at 37°C with 5% CO₂. Cells were treated for 12 h with 1 mL complete media containing either equal volume PBS, 0.5 μg/mL PAM2, 0.5 μg/mL PAM3, or 0.5 μg/mL LPS. Subsequently, media was removed, cells were washed once with ice cold PBS and lysed with 500 μL RIPA buffer (ThermoFisher Scientific; Waltham, MA, USA) supplemented with protease inhibitor cocktail (MilliporeSigma; Burlington, MA, USA) at 1:100 dilution and phosphatase inhibitor cocktail 2 (MilliporeSigma; Burlington, MA, USA) at 1:500 dilution. Cells were rocked at 4°C for 15 min before being transferred to Eppendorf tubes and centrifuged at top speed for 10 min at 4°C. Supernatant was collected and BCA (ThermoFisher Scientific; Waltham, MA, USA) was performed to assess total protein concentration. Total protein was separated on 10% SDS-PAGE gels at constant amperage (30 mA) and transferred to nitrocellulose membranes (MilliporeSigma; Burlington, MA, USA) for 1 h at constant amperage (300 mA). Membranes were blocked in 3% (w/v) bovine serum albumin (BSA; ThermoFisher Scientific; Waltham, MA, USA) for 1 h with rocking at room temperature and subsequently incubated overnight with rocking at 4°C with the following primary antibodies (1:1000 dilution): anti-β-actin, anti-GAPDH (Cell Signaling Technologies, Danvers, MA; USA), or anti-TNF (clone EPR19147, Abcam, Cambridge, MA, USA). Membranes were then incubated 1:2000 in secondary anti-rabbit IgG HRP-linked antibody (Cell Signaling Technologies, Danvers, MA; USA) for 45 min with rocking at room temperature before detection with ECL substrate and imaging (BioRad, Hercules, CA, USA).

Cells were seeded into 24-well plates at 5 × 10⁵ cells/well and allowed to adhere for 24 h at 37°C with 5% CO₂. Cells were then treated for 3 h with 500 μL complete media containing either equal volume PBS, 1 μg/mL Pam2CSK4, 1 μg/mL Pam3CSK4, or 1 μg/mL ultrapure LPS (PAM2; PAM3; LPS; all from InvivoGen, San Diego, CA, USA). Treatments were removed, cells were washed with sterile dPBS, and lysed with 400 μL TRIzol Reagent (Invitrogen, Waltham, MA, USA). RNA was extracted using chloroform (ThermoFisher Scientific; Waltham, MA, USA), 70% ethanol (MSU chemistry store), Buffer RPE (Qiagen, USA), RNase-free water (Invitrogen, Waltham, MA, USA), and RNA mini spin columns (Epoch Life Science Inc., Missouri City, TX, USA) before quantification of RNA via nanodrop. All RNA samples were normalized to a concentration of 20 ng/μL prior to cDNA library synthesis. The cDNA library was created using high-capacity cDNA Reverse Transcription kit (Applied Biosystems, Waltham, MA, USA) according to manufacturers’ recommendation with a total input of 100 ng RNA. qPCR was performed in triplicate using SsoAdvanced Universal SYBR Green Supermix (BioRad, Hercules, CA, USA) according to manufacturers’ recommendation with TNF, β-actin, LTα, and GAPDH primers (all from IDT, Coralville, IA, USA). All quantitative reverse transcription-PCRs were run on QuantStudio 5 Real-Time PCR system (ThermoFisher Scientific; Waltham, MA, USA). A total of forty cycles were run for each qRT-PCR and a CT of ≤25 cycles was used as a cutoff for positive values. Relative fold change was calculated using delta CT method and data were normalized to β-actin.

TNF sequence primer 1: 5′AGA​CCC​TCA​CAC​TCA​GAT​CA-3′

TNF sequence primer 2: 5′ TCT​TTG​AGA​TCC​ATG​CCG​TTG-3′

β-actin sequence primer 1: 5′ GAT​TAC​TGC​TCT​GGC​TCC​TAG-3′

β-actin sequence primer 2: 5′ GAC​TCA​TCG​TAC​TCC​TGC​TTG-3′

GAPDH sequence primer 1: 5′-AAT​GGT​GAA​GGT​CGG​TGT​G-3′

GAPDH sequence primer 2: 5′-GTG​GAG​TCA​TAC​TGG​AAC​ATG​TAG-3′

Cellular supernatants were collected from bacterial clearance assays before treatment, after treatment, and after S. aureus infection and frozen back at −80°C for later cytokine analysis. Cellular supernatants collected post S. aureus infection were centrifuged at 835 g for 15 min before aliquoting and storage at −80°C. ELISAs were performed with ELISA MAX Deluxe Set Mouse TNF-α kit according to manufacturers’ recommendations (BioLegend, San Diego, CA, USA).

Cells were seeded into 24-well plates containing sterile collagen coated coverslips (Neuvitro Corporation, Vancouver, WA, USA) at 5 × 10⁵ cells/well and allowed to adhere for 12 h at 37°C with 5% CO₂. Cells were then treated for 12 h with 500 μL complete media containing either equal volume PBS, 1 μg/ml Pam2CSK4 (PAM2; InvivoGen, San Diego, CA, USA), 1 μg/mL Pam3CSK4 (PAM3; InvivoGen, San Diego, CA, USA), or 1 μg/mL ultrapure LPS (InvivoGen, San Diego, CA, USA). Coverslips were washed twice with ice cold dPBS, fixed with 4% PFA (ThermoFisher Scientific; Waltham, MA, USA) for 15 min at room temperature, and blocked/permeabilized in 400 μL 2% BSA (ThermoFisher Scientific; Waltham, MA, USA) with 0.1% Triton X-100 (MilliporeSigma; Burlington, MA, USA) for 45 min on ice. Coverslips were then blocked with 1× avidin/biotin blocking solution for 15 min at room temperature with washing steps in-between each blocking step (ThermoFisher Scientific; Waltham, MA, USA). Primary rat anti-mouse TNF antibody conjugated to biotin (clone MP6-XT3; a kind gift from Dr. Mark Jutila at MSU) was diluted 1:250 in 2% BSA and incubated with coverslips for 1 h at room temperature protected from light. Coverslips were washed with dPBS and incubated with secondary avidin-HRP antibody (BD Pharmingen Inc., Franklin Lakes, New Jersey, USA) at 1:1,000 dilution in 2% BSA for 1 h at room temperature protected from light. Subsequently, coverslips were washed with dPBS and incubated with 3,3-diaminobenzidine chromogen (ImmPACTDAB; Vector Labs, Burlingame CA, USA) for 10 min at room temperature. Nuclear staining was performed with methyl green counterstain (Vector Labs, Burlingame CA, USA) for 5 min at 60°C. Visualization was performed with standard compound light microscope.

Murine immortalized macrophages (cell line NR-9456, BEI Resources; Manassas, VA, USA) were grown and maintained as previously described until 90% confluency was achieved. Cells were lifted with a sterile-filtered solution of 3 mm EDTA dissolved in dPBS for 10 min at 37°C with 5% CO₂. Cells were washed once in sterile dPBS, resuspended in complete media, and counted before being plated in 24-well plates at a cell density of 5 × 10⁵ cells/well and allowed to adhere for 6 h at 37°C with 5% CO₂. Cells were then treated for 12 h with 500 µL complete media containing either equal volume dPBS or 1 µg/mL Pam2CSK4. Cells were additionally treated with or without 1 or 100 μg/mL of p75TNFR:Fc (Amgen Inc.) or 1 or 100 µm TMI-005 (Cayman Chemical, Ann Arbor, MI, USA.

After treatment, cells were washed once with dPBS. Cells were lifted from wells with 3 mm EDTA dissolved in dPBS for 10 min at 37°C with 5% CO₂. Cells were transferred to 5 mL round-bottom polystyrene FACS tubes (Falcon Plastics, Brookings, SD, USA) and rinsed once with ice cold dPBS. A 1:500 LIVE/DEAD Fixable Dead Cell Yellow staining solution (Invitrogen, Waltham, MA, USA) was prepared in ice cold dPBS. Cells were resuspended in 1 mL staining solution and incubated on ice protected from light for 20 min. Cells were washed once and then incubated in FACS blocking buffer (5% sterile filtered mouse serum, Equitech Bio, Kerrville, TX, USA, and 2% FBS, SeraPrime, Fort Collins, CO, USA, dissolved in ice cold dPBS) on ice for 20 min to block unspecific binding. Cells were pelleted and resuspended in 200 µL FACS blocking buffer. Here, 100 µL of each cell suspension was transferred to a new FACS tube to receive either isotype control or fluorescence minus one (FMO) staining. Cells were incubated with 5 µL mouse anti-mouse CD45.2 IgG2a, clone 104, conjugated to FITC (BioLegend) and 5 µL rat anti-mouse TNF IgG1, clone MP6-XT22, conjugated to APC (BioLegend). Live/dead stained isotype control cells were treated with 5 µL nonspecific mouse anti-mouse IgG2a FITC and 2 µL rat anti-mouse IgG1 APC. Alternatively, live/dead stained FMO control cells were stained with 5 µL mouse anti-mouse CD45.2 IgG2a, clone 104, conjugated to FITC (BioLegend). Cells were incubated for 30 min on ice protected from light, then washed twice with FACS blocking buffer. Cells were resuspended in 300 µL Cytofix (BD Biosciences, Franklin Lakes, NJ, USA) and immediately run on a BioRad ZE5 Cell Analyzer. Cells for analysis on the ImageStream X Mark II imaging flow cytometer (Amnis, Seattle, WA, USA) were cultured were resuspended in 50 µL Cytofix.

To account for the variance within and between individual experiments, bacterial clearance assay dilutions were plated onto two agar plates. On each plate every dilution was in duplicate and each duplicate included 5 drops. The values of CFU counts for all 5 drops were averaged to obtain one value per dilution and the same was done for the duplicate dilution on each plate. The two values were averaged to obtain a replicate. Shown is the average of two replicates per experiment. Results from at least two independent experiments are shown for bacterial clearance assays. Western blots were performed a minimum of three times independently. Data from ELISA assays included three technical replicates and were performed a minimum of two times independently. Western blot data were analyzed using AzureSpot Pro software and plotted with GraphPad Prism software version 10. Quantitative RT-PCR was performed a minimum of 2 times independently, with each experiment containing three technical replicates. Relative fold change was calculated using delta CT method and data was normalized to β-actin. All other data were analyzed and plotted using either GraphPad Prism or R Studio software. With an exception of flow cytometry data in Figure 4 and Suppl Figure 9 (for all online suppl. material, see https://doi.org/10.1159/000546011), data were expressed as geometric mean with geometric standard deviation. Geometric means were calculated by taking the log means of experimental values and calculating antilog. Geometric standard deviations were calculated from the variance of log experimental values. Statistical significance was determined using either one-way analysis of variance (ANOVA), two-way ANOVA, or Student’s t test, and only statistically significant comparisons are shown in Figure (p ≤ 0.05).

Flow cytometry output was analyzed using FlowJo V10.10.0 (BD Biosciences, Ashland, OR, USA). Statistics were performed using GraphPad Prism software, V10.4.1. Fold change for tmTNF expression was calculated by averaging the geometric mean of compensated APC signal area across technical replicates, then normalizing the geometric mean of each treatment condition to the PBS control values. Plots contain 3 experimental replicates.

The statistical test used to compare treatment groups in flow cytometry analyses was a one-way ANOVA using Dunnett’s multiple comparisons test. ImageStream output was analyzed on the IDEAS analysis platform V. 6.2.

We found that prior stimulation of immortalized WT bone marrow-derived macrophages (BMDMs) with TLR2/6 agonist (PAM2), TLR4 agonist (LPS), or TLR2/1 agonist (PAM3) reduced bacterial burden of a subsequent S. aureus infection by ∼40% when compared to mock stimulated WT BMDMs (Fig. 1a and online suppl. S1A). Consistent with the findings of others [29‒32], BMDMs stimulated with these agonists required their cognate TLR2 or TLR4 receptors to productively clear bacterial infection (Figures 1b, online suppl. S1b, S1c). Together, this indicated that signaling through either TLR2 or TLR4 was critical for improving subsequent BMDM clearance of S. aureus infection. Here, we sought to investigate the intracellular signaling cascade initiated by the detection of PAMPs (through TLR2 or TLR4) that provided BMDMs with improved capacity for clearance of a subsequent bacterial infection.

Stimulation of innate immune cells with PAMPs often results in rapid secretion of pro-inflammatory cytokines such as IL-1β, IL-6, and TNF [33]. Thus, we analyzed transcript expression in whole lung homogenates of WT mice intratracheally inoculated with either PAM2 (TLR2/6) or PAM3 (TLR2/1) agonist for 6 h. We found that TNF was among one of the most strongly upregulated genes in response to TLR2 stimulation (Fig. 1c). Additionally, previously published literature has shown that TNF is one of the most strongly upregulated genes in response to TLR4 stimulation via LPS [34]. As such, we focused our subsequent investigation on macrophages, which are the primary producers of TNF [35, 36] and one of the first innate immune cells to respond to microbial invasion [37]. Dependent on the isoform (tmTNF or sTNF), TNF protein signals through TNFR1 or TNFR2. Pathway analysis of RNA seq data within the Reactome database identified that the tumor necrosis factor receptor type II (TNFR2) and noncanonical NFkB pathway were upregulated among mice that received either PAM2 or PAM3 (online suppl. Fig. S2). TNFR2 has two naturally occurring ligands, TNF and lymphotoxin α (LTα) [14, 38] both of which were identified to be upregulated in the RNA sequencing dataset. However, LTα upregulation (≤1.1 log fold) was mild in comparison to the robust upregulation of TNF (≥3.8 log fold) and as such, we chose to focus our analysis on TNF.

Next, we quantified TNF gene expression in immortalized WT BMDMs stimulated with either TLR2 or TLR4 agonists. Indeed, we found that at 3 h following WT BMDM stimulation, TNF gene expression was increased at least 5-fold relative to PBS (Fig. 1d, online suppl. S3a).

To address whether the observed increase in TNF expression in WT BMDMs was indeed dependent on the TLR2 or TLR4 agonists binding their cognate receptors. We stimulated immortalized murine TLR2−/− or TLR4−/− with their respective ligands for 3 h before quantifying TNF transcript via qRT-PCR. In TLR2−/− BMDMs stimulated with either PAM2 or PAM3, the absence of TLR2 receptor subunit resulted in deficiency of TNF gene expression (Fig. 1e, online suppl. S3b). To the contrary, even in the absence of the TLR2 receptor subunit, BMDMs upregulated TNF gene expression in response to LPS stimulation (Fig. 1e). In TLR4−/− BMDMs, LPS stimulation did not induce TNF gene expression. However, TNF gene was expressed in TLR4−/− BMDMs following either PAM2 or PAM3 stimulation (Fig. 1f, online suppl. S3c).

Next, we asked whether TNF mediated priming of BMDMs was necessary for improved clearance of subsequent bacterial infection. Unlike WT BMDMs (Fig. 1a), primary TNF−/− BMDMs stimulated with either TLR2 or TLR4 agonists did not improve their clearance of a subsequent S. aureus infection when compared to PBS treated primary TNF−/− BMDMs (Fig. 1g, online suppl. S4). This suggested the TNF produced as a direct result of TLR2 or TLR4 stimulation played a central role in priming BMDMs prior to S. aureus infection. Based on these data, we sought to further elucidate the mechanism of TLR-induced TNF mediated priming which resulted in improved clearance of a subsequent bacterial infection.

TNF exists in two bioactive forms, soluble/secreted (sTNF) and tmTNF, both of which bind distinct receptors to elicit differential signaling outcomes [12, 39‒41]. Cellular supernatants collected from WT BMDMs after 12-h stimulation with either TLR2 or TLR4 agonist exhibited a significant increase in sTNF protein production compared to PBS treated BMDMs (Fig. 2a). Immunocytochemistry and Western blot analyses following 12-h stimulation with either TLR2 or TLR4 agonists also demonstrated an increase in tmTNF protein production compared to mock treated BMDMs (Fig. 2b, c).

To address the possibility that this early sTNF and tmTNF could improve subsequent bacterial clearance by simply enhancing TNF production, we quantified TNF in supernatants from TLR2 or TLR4 stimulated WT BMDMs compared to PBS stimulated WT BMDMs after a 5-h infection period with S. aureus. PBS, PAM2, PAM3, and LPS stimulated BMDMs all exhibited comparable concentrations of sTNF after S. aureus infection (∼7,000 pg/mL) (Fig. 2d, online suppl. S5). This suggested the presence of TNF following TLR2 or TLR4 stimulation was not influencing the subsequent production of TNF during active S. aureus infection.

In Figure 1g, we demonstrated that PAM2 or LPS stimulated BMDMs deficient in TNF were not able to improve their ability to clear a subsequent S. aureus infection. Because either TLR2 or TLR4 stimulation of WT BMDMs resulted in abundant production of both sTNF and tmTNF, we sought to investigate if both bioactive forms of TNF equally participated in BMDM priming that granted BMDMs improved clearance of a subsequent S. aureus infection.

We first tested whether sTNF is sufficient to mediate the priming event. PAM2 or PBS stimulated TNF−/− BMDMs were supplemented with 2,000 pg/mL of exogenous murine recombinant sTNF or equal concentration of irrelevant protein control (OVA). Our exogenous sTNF dosage choice was guided by results in Figure 2a, which demonstrated that WT BMDMs stimulated with TLR2 or TLR4 agonist induced a minimum of 2,000 pg/mL sTNF. The validation of TNF−/− BMDMs prior to exogenous sTNF supplementation confirmed deficiency of sTNF production both after 12-h stimulation with TLR2 or TLR4 stimulation and after S. aureus infection (online suppl. Fig. S6).

Consistent with results in Figure 1g, we found that TLR2 stimulation in the absence of TNF was insufficient for BMDM priming (Fig. 3a). More importantly, we found that addition of exogenous sTNF to TLR2 stimulated TNF−/− BMDMs was also not sufficient to prime BMDMs prior to bacterial infection. This was evident by no change in S. aureus clearance between treatment groups that lacked exogenous sTNF and treatment groups that were supplemented with exogenous sTNF (Fig. 3a).

Next, we employed a reductionist approach to complement experiments which supplemented sTNF. The tmTNF is the bioactive precursor to mature sTNF and as such, the extracellular domain of tmTNF and sTNF have nearly homologous sequences [42, 43]. This posed an obstacle for selective TNF inhibition. To address this issue, we employed two approaches which target TNF protein regulation at different cellular levels. To ensure that our approaches yielded depletion to baseline concentrations, we first quantified sTNF concentrations in PBS treated BMDMs to establish a baseline. We determined that in the absence of TLR stimulation, sTNF concentrations in these cells was on average ∼100 pg/mL (online suppl. Fig. S7).

In our first approach, we leveraged the critical function of tumor necrosis factor-α-converting enzyme (TACE; ADAM17) for regulating TNF. ADAM17 is the enzyme responsible for cleavage of the precursor form of tmTNF to its secreted sTNF form [42, 43]. Therefore, we postulated the use of ADAM17 small molecule inhibitor, TMI-005 (Apratastat) would prevent cleavage of tmTNF, thereby reducing sTNF protein levels. Indeed, inhibition of ADAM17 allowed us to enrich cell surface associated tmTNF protein while simultaneously reducing concentrations of sTNF protein by ∼75%–∼500 pg/mL (Fig. 3b). Despite this reduction of sTNF protein, WT BMDMs stimulated with TLR2 agonist retained their improved ability to clear subsequent S. aureus infection (Fig. 3c). While a ∼75% decrease in sTNF was a substantial reduction, 500 pg/mL was still representative of a 3-fold increase of sTNF compared to PBS treated BMDMs. Thus, in our second approach we employed a murine biologic, p75TNFR:Fc, which acts as a soluble decoy receptor for sTNF. Similar to TMI-005 treatment, cellular supernatants collected from WT BMDMs treated with increasing concentrations of murine p75TNFR:Fc in the presence of TLR2 agonist showed a dose dependent reduction of sTNF. The optimal dosage of p75TNFR:Fc was determined to be 100 μg/mL, which reduced sTNF by >90% from ∼2,000 pg/mL to ∼125 pg/mL (online suppl. Fig. S8). Treatment of WT BMDMs with 100 μg/mL p75TNFR:Fc for 12 h concurrent with TLR2 stimulation effectively reduced sTNF protein concentrations to near baseline concentrations, achieving a >90% reduction of sTNF to ∼171 pg/mL (Fig. 3d). Importantly, reduction of sTNF to near baseline levels (prior to bacterial infection) did not abrogate improved clearance of S. aureus following TLR2 stimulation (Fig. 3e). Cumulatively, our results to this point indicate that the presence of sTNF following TLR2 stimulation was not required for BMDM priming which led to improved clearance of a subsequent S. aureus infection. This suggested that tmTNF signaling in response to TLR2 stimulation may be the critical signal that primes BMDMs.

Despite TNF expression being necessary for TLR-induced BMDM priming, sTNF protein was neither sufficient nor required for improved S. aureus clearance by primed BMDMs. Thus, we next sought to examine the contribution of tmTNF to TLR2-induced priming of BMDMs. The TMI-005 and the p75TNFR:Fc reduced sTNF either by inhibition of tmTNF cleavage (TMI-005) or by acting as a soluble decoy receptor for TNF protein (p75TNFR:Fc), and neither of these treatments affected viability of BMDMs (online suppl. Fig. S9). Using flow cytometry, we next investigated whether these reagents affected expression of tmTNF on the surface of WT BMDMs. In agreement with Western blot analysis in Figure 2c flow cytometric analysis found that PAM2 treatment of BMDMs led to a consistent, albeit non-significant increase in the expression of surface TNF (Fig. 4a, b). Consistent with its role in inhibiting TNF cleavage by ADAM17, both concentrations of TMI-005 enriched for tmTNF expression when compared to PBS-treated BMDMs (Fig. 4a, b). In contrast, treatment with the decoy receptor p75TNFR:Fc did not result in enhanced tmTNF expression by flow cytometry. Interestingly, treatment with either TMI-005 or p75TNFR:Fc increased background fluorescence. These results were confirmed using imaging cytometry, which revealed co-localization of TNF with the surface marker CD45 in a proportion of BMDMs treated with PAM2 or with PAM2 and TMI-005 (Fig. 4c).

To investigate whether tmTNF contributed to BMDM priming following TLR stimulation, we leveraged the physical proximity constraint inherent to tmTNF signaling. Because tmTNF is anchored to the cell membrane [41, 43], it requires cell-to-cell contact for ligand/receptor binding and signaling to occur. We capitalized on this cell-to-cell contact requirement to assess tmTNF involvement in BMDM priming prior to infection with S. aureus. Specifically, we performed bacterial clearance assays in which WT BMDMs were plated at various cell densities prior to TLR2 stimulation. The MOI was scaled for each cell density group, such that MOI was 2:1 for all conditions. At low cell density with limited cell-to-cell contact, bacterial burden remained the same across treatment groups, indicating that BMDMs were deficient in clearing S. aureus (Fig. 5a). As cell density and cell-to-cell contact increased, bacterial burden decreased in TLR2 stimulated WT BMDMs. At our final concentration of 2 million cells/well (4 × 10⁶ cells/ml) no TLR2-induced priming advantage in bacterial clearance was observed (Fig. 5a). This suggested that cell-to-cell contact may be important for TLR-induced BMDM priming that leads to improved S. aureus clearance.

To more directly address a possible role of tmTNF in BMDM priming following TLR stimulation, we targeted TNFR2 by antibody-based blocking. TNFR2 is the primary receptor for tmTNF, whereas sTNF cannot signal through TNFR2 [13, 14]. WT BMDMs stimulated with TLR2 agonist in the presence of TNFR2 blocking antibody prior to infection lost their ability to efficiently clear subsequent S. aureus infection. In fact, the bacterial burden of TLR2 stimulated BMDMs in the presence of TNFR2 blocking antibody resembled that of PBS treated BMDMs (Fig. 5b). This suggested that signaling through tmTNF may be important for priming BMDMs following TLR2 stimulation. Interestingly, WT BMDMs stimulated with PBS in the presence of TNFR2 blocking antibody showed a subsequent improved ability for S. aureus clearance. Collectively, our results provide evidence that tmTNF, but not sTNF, induced in response to TLR stimulation contributes to BMDM priming that enables these cells to improve their ability to clear a subsequent S. aureus infection.

Microbial structures or products are collectively known as PAMPs and upon invasion bind to host membrane PRRs including TLRs. TLR binding of PAMPs triggers a myriad of intracellular signaling events which orchestrate a complex network of cellular activation [1, 44]. Although TLRs are among one of the most well studied family of PRRs [45], there is still much of the TLR signaling cascade that needs to be unraveled. A deeper understanding of the TLR signaling cascade will provide insight into the individual signaling components that comprise cellular activation post PAMP ligation by TLRs.

This study demonstrated that treatment of macrophages with TLR2 or TLR4 agonists prior to infection improved their capacity to clear S. aureus infection due to TNF mediated priming. We found that stimulation of WT BMDMs with TLR2 or TLR4 agonists significantly upregulated both bioactive protein forms of TNF (tmTNF; sTNF). Interestingly, however, we found that sTNF was not required for BMDM priming following TLR stimulation. Depletion of sTNF in WT BMDMs prior to S. aureus infection did not compromise improved bacterial clearance. Supplementation of exogenous sTNF to TNF−/− BMDMs in the presence of TLR stimulation was not sufficient restore improved clearance capacity. While we did not explicitly determine that tmTNF is required for macrophage priming, we did confirm that both cell-to-cell contact and TNFR2 signaling, which are critical for tmTNF but not sTNF signaling, are required to improve bacterial clearance capacity of primed BMDMs.

TLR2 stimulation in the absence of TNF (TNF−/− BMDMs) was insufficient to establish signaling which resulted in BMDM priming. This was reflected by the loss of improved S. aureus clearance by TLR2 stimulated TNF−/− BMDMs. TLR2 stimulation followed by exogenous sTNF addition prior to bacterial infection also was not sufficient to rescue the signaling that occurs in TLR-stimulated WT BMDMs prior to S. aureus infection. Altogether, these data support our hypothesis that tmTNF signaling resulting from TLR stimulation prior to S. aureus infection could prime BMDMs by enhancing their capacity for bacterial clearance.

To assess whether tmTNF was required for BMDM priming, we leveraged the physical constraint inherent to membrane-bound proteins such as tmTNF. These experiments revealed an association between cell-to-cell contact and improved BMDM S. aureus clearance ability. Because MOI was scaled and consistent between treatment groups, the possibility of S. aureus overgrowth and overwhelming BMDMs was limited. This suggested that lack of improved S. aureus clearance by TLR2 stimulated BMDMs at lower cell densities could be due to limited cell-to-cell contact that is required for tmTNF signaling.

To more directly examine tmTNF involvement in BMDM priming, we blocked TNFR2 to prevent signaling following TLR2 stimulation. In TLR2 stimulated BMDMs, blocking tmTNF/TNFR2 signaling led to deficiencies in S. aureus clearance compared to TLR2 stimulated BMDMs capable of signaling through TNFR2. This provided evidence that tmTNF signaling following TLR stimulation could be a key signaling event involved in priming BMDMs prior to infection which leads to improved S. aureus clearance.

While blocking TNFR2 in TLR2 stimulated BMDMs reduced responsiveness of these cells to a subsequent infection, we also found that blocking TNFR2 in PBS treated BMDMs had an opposite effect. Not much is known about TNFR2 in regards to macrophage priming, nonetheless this result was unexpected. There are several important differences between PBS- and TLR2-stimulated macrophages that could help explain this phenomenon. BMDMs stimulated with PBS did not produce TNF, therefore for this treatment condition, TNF was only produced in response to S. aureus infection. Blocking TNFR2 in PBS stimulated BMDMs likely directed TNF produced during S. aureus infection to bind and signal through TNFR1. Because signaling through TNFR1 is known to initiate and perpetuate inflammatory host responses [46, 47], it is possible that funneling TNF through TNFR1 in the presence of an active infection bolstered macrophage inflammatory responses and aided in the improved clearance of S. aureus. TLR2 stimulated BMDMs produced both sTNF and tmTNF prior to infection. Thus, blocking TNFR2 in TLR2-stimulated BMDMs likely directed TNF to bind and signal through TNFR1 prior to S. aureus infection. As a result, tmTNF may have been blocked from signaling through its cognate receptor which could be important for priming BMDMs to respond to S. aureus infection.

Studies highlighting the importance of TLR activation and subsequent sTNF production in host defense against infection have prompted interest in therapeutic manipulation of TLRs or sTNF, especially in the context of antibiotic resistance [48‒53]. However, immunotherapies which directly target TLRs or sTNF can result in numerous global off-target effects which are disadvantageous for the host. Several studies suggest that prolonged activation of TLRs and subsequent inflammation can contribute to a hyperinflammatory host environment which damages native host cellular structures and is associated with auto-immune disease [46, 47, 54‒56]. On the other hand, studies suggest that sustained suppression of sTNF signaling is associated with increased host susceptibility to infection [50, 57‒59].

In the present study, we found that a critical signaling event for activation of macrophage microbial defense via TLR2 ligation may rely on tmTNF signaling, which is the bioactive precursor form of sTNF. Interestingly, we also found that the mature secreted protein form of TNF, sTNF, was not necessary for this priming event. This finding was unexpected because the majority of the existing literature which characterizes the importance of TNF in resolving infection has historically focused on the role of sTNF, while the role of tmTNF remains poorly defined [60‒62]. Nonetheless, more recent reports suggest that tmTNF, independent of sTNF, may be sufficient for protection against mycobacterium and listeria infection [48, 63]. These studies found that the presence of tmTNF alone was sufficient to either partially or fully rescue mice from succumbing to mycobacterial or listeria infection as compared to mice globally deficient in TNF, which exhibited 100% mortality [48, 63]. The findings from these reports attribute the protective function of tmTNF to tmTNF’s retained ability to functionally induce signaling [48, 63]. Thus, these studies highlight the overlooked importance and contribution that tmTNF could have on infection resolution. The function of tmTNF is beginning to be elucidated, there are still many gaps in our understanding of tmTNF signaling and how it may influence resolution of bacterial infections, specifically in the context of macrophages. While our study did not evaluate whether and how tmTNF is directly involved in resolution of bacterial infection, we provided evidence of tmTNF contribution to macrophage priming. To our knowledge this is the first report of tmTNF conferring an early advantage which improved subsequent S. aureus clearance.

In conclusion, we have shown that TLR2 or TLR4 activation of BMDMs results in signaling which leads to increased BMDM ability to clear S. aureus. We sought to understand what specific signaling events are critical in promoting BMDM priming following early TLR stimulation (PAMP detection) that led to this increased ability of S. aureus clearance. We found that sTNF signaling was not required to prime BMDMs, however, tmTNF signaling contributed to priming of BMDMs allowing them to efficiently respond to a subsequent S. aureus infection. These findings will provide insight into the TLR-based downstream signaling mechanisms which drive host-pathogen resolution mechanisms.

We would like to thank Dr. Fermin Guerra for his valuable discussion, suggestions, and guidance.

This work was approved by Montana State University’s Institutional Biosafety Committee (IBC). Protocol numbers 2022-127-IBC and 2023-64-IBC. Animal protocols were reviewed and approved by Montana State University’s Institutional Animal Care and Use Committee (IACUC). Protocol number 2022-49-IA.

The authors have no conflicts of interest to declare.

Research reported in this publication was supported by the National Institutes of Health under awards number TL1TR002318 (Luu) and R21AI171903 (Bimczok), institutional support start-up funds from Montana State University (Rynda-Apple), an NSF Research Traineeship (NRT) award (Gattiker), the Montana State University Agricultural Experiment Station, and the M.J. Murdock Charitable Trust. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Conceptualization: A.M.L., A.A.H., and A.R.-A.; methodology: A.M.L., A.A.H., J.G., K.M.S., D.B., M.N.H., and E.B.; data curation: A.M.L., A.A.H., J.G., and D.B.; project administration/oversight: A.M.L. and A.R.-A., writing original draft: A.M.L., writing, revisions, and editing: A.M.L., A.A.H., M.N.H., K.M.S., E.B., A.R.-A., J.G., D.B.; funding: A.M.L., J.G., D.B., and A.R.-A.

All data generated or analyzed during this study are included in this article and its supplementary material files. Further enquiries can be directed to the corresponding author.