The TLR2 agonist BLP, a synthetic bacterial lipopeptide (Pam3Cys-Ser-Lys4-OH), and the TLR4 agonist ultrapure LPS from E. coli serotype O55:B5 were purchased from EMC Microcollections (Tubingen, Germany) and InvivoGen (San Diego, CA), respectively. The NOD1 agonist L-Ala-γ-D-Glu-mDAP (Tri-DAP) and NOD2 agonist MDP were obtained from InvivoGen. Antibodies (Abs) that recognize TLR4, NOD1, NOR2, myeloid differentiation factor 88 (MyD88), IL-1 receptor-associated kinase-1 (IRAK-1), CARD9, and RIP2 were purchased from Cell Signaling Technology (Beverly, MA), Santa Cruz Biotechnology (Santa Cruz, CA), and Abcam (Cambridge, MA), respectively. Abs that recognize NF-κB p65, phosphor-p65 at Ser586, the inhibitor of κBα (IκBα), phosphor-IκBα at Ser32/36, MAPK p38, and phospho-p38 at Th180/Try182 were purchased from Cell Signaling Technology. All culture medium and reagents for cell cultures were obtained from Invitrogen Life Technologies (Paisley, Scotland, U.K.). All other chemicals, unless indicated, were purchased from Sigma-Aldrich (St. Louis, MO).

Pyrogen-free, 8- to 10-week-old C3H/HeN mice, TLR2- and TLR4-deficient mice on the C3H background, C57BL/6 mice, NOD1- and NOD2-deficient mice on the C57BL/6 background were obtained from Harlan (Oxon, UK), Jackson Laboratories (Bar Harbor, ME), and Carsten J. Kirschning at Technische Universitat Munchen, Munich, Germany. Mice were housed in barrier cages under controlled environmental conditions (12/12 h light/dark cycle, 55 ± 5% humidity, 23°C) and had free access to standard laboratory chow and water. All animal studies were conducted with the ethical approval granted from the Institutional Animal Care and Use Committee of Soochow University and the Ethics Committee of University College Cork, and complied with the animal welfare act. The methods applied in the present study were performed in accordance with the approved guidelines.

Peritoneal macrophages were collected from wild-type, TLR4- and TLR2-deficient, and NOD1- and NOD2-deficient mice by peritoneal lavage and incubated with DMEM containing 10% heat-inactivated fetal calf serum (FCS) in 24-well plates (Falcon, Lincoln Park, NJ) for 90 min to remove non-adherent cells as previously described (36, 37). Bone marrow-derived macrophages (BMMs) were isolated from the femurs and tibias of wild-type, TLR4- and TLR2-deficient, and NOD1- and NOD2-deficient mice, and cultured in DMEM containing 20% heat-inactivated FCS, penicillin (100 units/ml), streptomycin sulfate (100 μg/ml), and supplemented with 10 ng/ml of recombinant mouse macrophage colony-stimulating factor (CSF) (R&D Systems, Minneapolis, MN) for 7 days at 37°C in a humidified 5% CO2 atmosphere as previously described (36, 37). The purity of both peritoneal macrophages and BMMs was >95%, as confirmed by FACScan analysis of the positive F4/80 antigen (Ag) staining with a rat anti-mouse F4/80 Ab (Serotec, Oxford, U.K.).

Isolated peritoneal macrophages or BMMs were incubated with PBS, LPS (10 ng/ml), BLP (10 ng/ml), Tri-DAP (5 μg/ml), MDP (5 μg/ml), LPS + Tri-DAP (10 ng/ml + 5 μg/ml), LPS + MDP (10 ng/ml + 5 μg/ml), BLP + Tri-DAP (10 ng/ml + 5 μg/ml), and BLP + MDP (10 ng/ml + 5 μg/ml) for 6 h, and further challenged with gram-positive or gram-negative bacteria to assess their ability in bacterial phagocytosis, killing, and phagosome maturation.

Isolated peritoneal macrophages or BMMs were plated onto 96-well plates (Falcon) at 2 × 10⁴ cells/well and incubated with PBS as the control or stimulated with LPS (10 ng/ml), BLP (10 ng/ml), Tri-DAP (5 μg/ml), MDP (5 μg/ml), and their combinations for 12 h. Cell-free supernatants were collected and stored at −80°C until analysis. Concentrations of inflammatory cytokines TNF-α, IL-6, IL-12p70, and chemokine CXCL2 in the supernatant were assessed by cytometric bead array (BD Biosciences, San Joes, CA) and ELISA (R&D Systems), respectively.

Isolated peritoneal macrophages were incubated with PBS, LPS (10 ng/ml), BLP (10 ng/ml), Tri-DAP (5 μg/ml), MDP (5 μg/ml), and their combinations for 2 h, and stained with anti-complement receptor type 3 (CR3) (BD PharMingen, San Diego, CA) and anti-FcγIII/II receptor (FcγR) (BD PharMingen) monoclonal antibodies (mAbs) conjugated with PE or FITC. PE- or FITC-conjugated isotype-matched mAbs (BD PharMingen) were used as the control. FACScan analysis was performed from at least 10,000 events for detecting the surface expression of CR3 and FcγR on macrophages using CellQuest software (BD Biosciences).

Gram-positive Staphylococcus aureus (S. aureus) and gram-negative Salmonella typhimurium (S. typhimurium) were obtained from American Type Culture Collection (ATCC, Manassas, VA) and the National University of Ireland Culture Collection, respectively. Bacteria were cultured at 37°C in trypticase soy broth (Merck, Darmstadt, Germany), harvested at the mid-logarithmic growth phase, washed twice, and resuspended in PBS for in vitro use. The concentration of resuspended bacteria was determined and adjusted spectrophotometrically at 550 nm.

Bacterial uptake, phagocytosis, and intracellular bacterial killing were determined as previously described (38, 39). Briefly, S. aureus and S. typhimurium were heat-killed at 95°C for 20 min and labeled with 0.1% FITC (Sigma-Aldrich). After stimulation with LPS, BLP, Tri-DAP, MDP, and their combinations for 6 h as mentioned above, isolated peritoneal macrophages or BMMs were incubated with heat-killed, FITC-labeled S. aureus or S. typhimurium at a ratio of 1/20 (macrophage/bacteria) at 37°C for 30 min. Bacterial uptake was assessed by FACScan analysis and bacterial ingestion was further determined after the external fluorescence of the bound, but non-ingested, bacteria was quenched with 0.025% crystal violet (Sigma-Aldrich). Intracellular bacterial killing was assessed by incubation of macrophages with live S. aureus or S. typhimurium (macrophage/bacteria = 1/20) at 37°C for 60 min in the presence or absence of cytochalasin B (5 μg/ml) (Sigma-Aldrich). After macrophages were lysed, total, and extracellular bacterial killing were determined by incubation of serial 10-fold dilutions of the lysates on tryptone soy agar (Merck) plates at 37°C for 24 h. Intracellular bacterial killing was calculated according to the total and extracellular bacterial killing.

Phagosome luminal pH was assessed as previously described (40, 41). Briefly, heat-killed S. aureus and S. typhimurium were doubly labeled with 5 μg/ml carboxyfluorescein-SE (a pH-sensitive fluorescent probe) (Molecular Probes, Eugene, OR) and 10 μg/ml carboxytetramethylrhodamine-SE (a pH-insensitive fluorescent probe) (Molecular Probes). After stimulation with LPS, BLP, Tri-DAP, MDP, and their combinations for 6 h, isolated peritoneal macrophages or BMMs were pulsed with the labeled bacteria (macrophage/bacteria = 1/20) for 20 min, and then chased at 37°C for the indicated time periods. Macrophage-based mean fluorescence intensity (MFI) of fluorescein on FL1 and rhodamine on FL2 were simultaneously detected by FACScan analysis using CellQuest software (BD Biosciences). Phagosomal pH was calculated according to the ratio of fluorescein/rhodamine fluorescence using a calibration curve.

After stimulation with LPS, BLP, Tri-DAP, MDP, and their combinations for 6 h, isolated peritoneal macrophages or BMMs were labeled with a red fluorescent cell membrane linker PKH26 (20 μM) (Sigma-Aldrich) for subsequent phagosome recognition, as previously described (40, 42). PKH26-labeled macrophages were pulsed and chased with heat-killed S. aureus or S. typhimurium (macrophage/bacteria = 1/20) at 37°C for the indicated time periods. Cells were lysed in a hypotonic buffer, and phagosomes were prepared by centrifugation. The isolated phagosomes were permeabilised with 0.2% saponin (Sigma-Aldrich) and stained with FITC-conjugated anti-LAMP-1 mAb (Abcam) that specially recognizes late endosomes/lysosomes, or FITC-conjugated isotype-matched mAb (Abcam) as the control. The green MFI of LAMP-1 on the positive red fluorescent events (phagosomes that have ingested bacteria), representing phagolysosome fusion and/or phagosome maturation, was quantitatively assessed by FACScan analysis using CellQuest software (BD Biosciences).

Isolated BMMs were incubated with PBS, BLP (10 ng/ml), Tri-DAP (5 μg/ml), MDP (5 μg/ml), and their combinations for various time periods. For determination of intracellular NF-κB p65 and MAPK p38 phosphorylation, BMMs were fixed and permeabilized simultaneously with Phosflow fix and perm buffers (BD Biosciences) for 30 min on ice. Cells were then stained with anti-phospho p65 and anti-phospho p38 mAbs conjugated with PE or Alexa Fluor 488 (Cell Signaling Technology). PE or Alexa Fluor 488-conjugated isotype-matched mAbs (Cell Signaling Technology) were used as the control. FACScan analysis was performed from at least 10,000 events for detecting the intracellular staining of phosphorylated NF-κB p65 and MAPK p38 in BMMs using CellQuest software (BD Biosciences).

Following stimulation of isolated BMMs with LPS (10 ng/ml), Tri-DAP (5 μg/ml), MDP (5 μg/ml), and their combinations for the indicated time periods, cells were collected, washed with ice-cold PBS, and lysed on ice in cell lysis buffer (Cell Signaling Technology), supplemented with 1 mM phenylmethylsulfonyl fluoride and protease inhibitor cocktail (Roche Life Science, Indianapolis, IN). The resultant lysates were centrifuged and supernatants containing the cytoplasmic proteins were collected. Protein concentrations were determined using a micro bicinchoninic acid (BCA) protein assay (Pierce, Rockford, IL). Equal amounts of protein extracts were separated on SDS-polyacrylamide gels and trans-blotted onto polyvinylidene difluoride (PVDF) membranes (Schleicher and Schuell, Dassel, Germany). The membrane was blocked for 1 h at room temperature with PBS containing 0.05% Tween-20 and 5% nonfat milk, and probed overnight at 4°C with the respected primary Abs. Blots were then incubated with appropriate horseradish peroxidase-conjugated secondary Abs (Dako, Cambridge, U.K.) at room temperature for 1 h, developed with SuperSignal chemiluminescent substrate (Pierce), and captured with LAS-3000 imaging system (Fujifilm, Tokyo, Japan).

ChIP assay was performed using the ChIP-IT Express kit (Active Motif, Carlsbad, CA) according to the manufacturer's instructions. Briefly, isolated BMMs were stimulated with LPS (10 ng/ml), BLP (10 ng/ml), Tri-DAP (5 μg/ml), MDP (5 μg/ml), and their combinations for 1 h, and then washed with PBS and fixed with 1% formaldehyde at room temperature for 10 min. The cells were lysed in ice-cold lysis buffer and sheared by sonication to generate 200–1,000 bp DNA fragments. Immunoprecipitation was carried out by incubation of the diluted sonicates with protein G magnetic beads and a specific anti-NF-κB p65 Ab (Santa Cruz Biotechnology) with rotation overnight. Protein G magnetic beads were collected with magnetic stand (Active Motif) and washed extensively. Protein–DNA complexes were eluted, the cross-link was reversed, and proteins were digested with proteinase K. Immunoprecipitated DNA and nonimmunoprecipitated DNA (input control) were amplified by quantitative PCR (qPCR) using the following promoter-specific primers: mouse TNF-α (sense-5′-TCCTTGATGCCTGGGTGTCCC-3′ and antisense-5′-GCAGACGGCCGCCTTTATAGC-3′) and mouse IL-6 (sense-5′-TCCAATCAGCCCCACCCACTC-3′ and antisense-5′-GGTGGGCTCCAGAGCAGAATG-3′).

Pyrogen-free, 8- to 10-week-old male C3H/HeN mice received intraperitoneal injection of 200 μl PBS, LPS (1 mg/kg), BLP (1 mg/kg), Tri-DAP (15 mg/kg), LPS + Tri-DAP (1 mg/kg + 15 mg/kg), and BLP + Tri-DAP (1 mg/kg + 15 mg/kg), respectively, 6 h before septic challenge. Polymicrobial sepsis was induced using a CLP method as previously described (40, 43). Briefly, mice were anesthetized by intramuscular injection of 150 μl of a ketamine/xylazine admixture (150 μl ketamine + 150 μl xylazine made up to 1 ml with 0.9% saline). A midline laparotomy was performed at which the cecum was delivered, ligated at the base, just distal to the ileocaecal juncture with a 2/0 mersilk tie. A single through puncture was then made distal to the ligature with a 17G needle. The cecum was returned to the peritoneal cavity and the abdomen was closed with 6/0 prolene sutures. Survival rates were recorded and monitored for at least 7 days. Blood samples were collected at 2 and 6 h post CLP, and serum TNF-α and IL-6 were assessed by cytometric bead array (BD Biosciences).

Bacterial counts were determined as previously described (39, 40). Briefly, mice were culled at 12 and 24 h post CLP-induced polymicrobial sepsis. Blood samples were obtained by retinal artery puncture, and the dissected liver and spleen were homogenized in sterile PBS. Serial 10-fold dilutions of heparinized whole blood and organ homogenates in sterile water containing 0.5% Triton X-100 (Sigma-Aldrich) were plated on brain heart infusion agar (BD Biosciences) and incubated for 24 h at 37°C for determination of bacterial colony-forming unit (CFU).

All data are expressed as the mean ± SD. Statistical analysis was performed using the log rank test for survival, and the ANOVA or Mann-Whitney U test for all others with GraphPad software version 5.01 (Prism, La Jolla, CA). Differences were judged to be statistically significant when the p-value was less than 0.05.

We first stimulated macrophages isolated from wild-type mice with the TLR4 agonist LPS, NOD1 agonist Tri-DAP, NOD2 agonist MDP, and their combinations for 12 h to assess the production of inflammatory cytokines and chemokines. As shown in Figure 1A, stimulation of macrophages with LPS led to an increased release of inflammatory cytokines TNF-α, IL-6, IL-12p70, and chemokine CXCL2 (p < 0.01 vs. PBS-treated macrophages), whereas Tri-DAP or MDP stimulation caused moderate but significant increases in TNF-α, IL-6, IL-12p70, and CXCL2 release (p < 0.05 vs. PBS-treated macrophages). Of note, a combined stimulation of LPS with Tri-DAP or MDP maximized the inflammatory response with substantially augmented release of TNF-α, IL-6, IL-12p70, and CXCL2 when compared to the response observed with LPS, Tri-DAP, or MDP alone (p < 0.05, p < 0.01). Consistent with the finding from the TLR4 agonist LPS stimulation, stimulation of macrophages with the TLR2 agonist BLP also induced a markedly increased release of TNF-α, IL-6, IL-12p70, and CXCL2 (p < 0.01 vs. PBS-treated macrophages), which was further augmented by a combination of BLP plus Tri-DAP or MDP (p < 0.05, p < 0.01 vs. macrophages stimulated with BLP, Tri-DAP, or MDP alone) (Figure 1B). These results indicate that activation of both TLR and NOD signaling in macrophages leads to an augmented inflammatory response.

To examine whether TLR and NOD signaling are both critical for the observed optimal inflammatory response, we further stimulated macrophages isolated from TLR4- and TLR2-deficient, and NOD1- and NOD2-deficient mice with LPS, BLP, Tri-DAP, MDP, and their combinations. LPS stimulation failed to induce TNF-α release and was unable to augment Tri-DAP- or MDP-induced TNF-α release in TLR4-deficinet macrophages (Figure 2A). Moreover, the amplified TNF-α release induced by a combined stimulation of LPS with the NOD1 agonist Tri-DAP or NOD2 agonist MDP observed in wild-type macrophages was receded in NOD1- and NOD2-deficient macrophages, respectively (Figure 2A). Similarly, BLP also lost its stimulatory ability in TLR2-deficient macrophages, and failed to augment Tri-DAP- or MDP-induced TNF-α release in NOD1- or NOD2-deficient macrophages (Figure 2B). These results suggest that the augmented inflammatory response by a combined stimulation of TLR and NOD agonists is entirely dependent on intact TLR and NOD signaling.

To examine whether the crosstalk between TLR and NOD signaling results in an enhanced activation of intracellular signal transduction pathways, thus causing an augmented inflammatory response, we assessed TLR- and NOD-mediated upstream and downstream pathways in macrophages stimulated with TLR and NOD agonists. Stimulation of BMMs with LPS, Tri-DAP, MDP, and their combinations did not affect TLR- and NOD-mediated upstream pathways including the expression of TLR4, NOD1, NOD2, MyD88, IRAK1, RIP2, and CARD9 (Figure 3A). However, stimulation of BMMs with a combination of LPS plus Tri-DAP resulted in a vigorous activation in the downstream NF-κB pathway with considerably increased phosphorylation of NF-κB p65 at Ser586 and IκBα at Ser32/36 (p < 0.05, p < 0.01), but had no augmentative effect on phosphorylation of MAPK p38 at Th180/Try182, when compared to BMMs stimulated with LPS or Tri-DAP alone (Figures 3B,D). A markedly enhanced expression of phosphorylated NF-κB p65 at Ser586 and IκBα at Ser32/36 was also observed in BMMs stimulated by LPS in combination with MDP (p < 0.05, p < 0.01 vs. BMMs stimulated with LPS or MDP alone) (Figures 3C,E). We also stimulated BMMs with BLP, Tri-DAP, MDP, and their combinations, and observed a substantially augmented activation of NF-κB p65 in BMMs stimulated with a combination of BLP plus Tri-DAP (p < 0.01) (Figure S1A) or MDP (p < 0.05) (Figure S1B) compared with BMMs stimulated with BLP, Tri-DAP, or MDP alone. Again, stimulation of BMMs by BLP in combination with Tri-DAP (Figure S1C) or MDP (Figure S1D) did not induce a further activation of MAPK p38 compared with BMMs stimulated with BLP, Tri-DAP, or MDP alone. Consistent with the amplified downstream NF-κB activation by a combined stimulation of TLR and NOD agonists, stimulation of BMMs with a combination of LPS plus Tri-DAP or MDP strongly augmented the nuclear transactivation of NF-κB p65 at both TNF-α and IL-6 promoters (p < 0.05, p < 0.01 vs. BMMs stimulated with LPS, Tri-DAP, or MDP alone) (Figure 3F). A substantially enhanced recruitment of NF-κB p65 to either the TNF-α promoter or IL-6 promoter was also observed in BMMs stimulated by BLP in combination with Tri-DAP or MDP (p < 0.05, p < 0.01 vs. BMMs stimulated with BLP, Tri-DAP, or MDP alone) (Figure 3G). These results indicate that the crosstalk between TLR and NOD signaling by a co-stimulation with TLR and NOD agonists triggers an amplified downstream activation of the NF-κB pathway with subsequently augmented nuclear transactivation of p65 at both TNF-α and IL-6 promoters.

We stimulated macrophages isolated from wild-type mice with LPS, Tri-DAP, MDP, and their combinations for 6 h, and further challenged these macrophages with gram-negative S. typhimurium to assess bacterial uptake, phagocytosis, and intracellular killing. Stimulation with LPS, Tri-DAP, or MDP alone did not affect macrophage-associated bactericidal activity; however, a combined stimulation of LPS with Tri-DAP or MDP significantly enhanced uptake and phagocytosis of S. typhimurium with substantially increased intracellular killing of the engulfed S. typhimurium (p < 0.05 vs. macrophages stimulated with LPS, Tri-DAP, or MDP alone) (Figure 4A). BLP stimulation in combination with Tri-DAP or MDP also resulted in an augmented bactericidal activity against gram-positive S. aureus, as represented by significantly enhanced uptake, phagocytosis, and intracellular killing of S. aureus (p < 0.01 vs. macrophages stimulated with BLP, Tri-DAP, or MDP alone), whereas stimulation with BLP, Tri-DAP, or MDP alone had no such effect (Figure 4B). Nevertheless, a combined stimulation of LPS or BLP with either Tri-DAP or MDP significantly augmented macrophage-associated bactericidal activity against gram-positive S. aureus (p < 0.05 vs. macrophages stimulated with LPS, Tri-DAP, or MDP alone) (Figure S2A) and gram-negative S. typhimurium (p < 0.01 vs. macrophages stimulated with BLP, Tri-DAP, or MDP alone) (Figure S2B). These results indicate that activation of both TLR and NOD signaling in macrophages is required to induce an enhanced antimicrobial response.

We next stimulated macrophages isolated from TLR4- and TLR2-deficient, and NOD1- and NOD2-deficient mice with LPS, BLP, Tri-DAP, MDP, and their combinations for 6 h, and further challenged these macrophages with live bacteria to assess the intracellular bacterial killing. The augmented bactericidal activity observed in wild-type macrophages following a combined stimulation of LPS with Tri-DAP or MDP was totally abolished in TLR4-deficient macrophages and selectively lost in NOD1- and NOD2-deficient macrophages upon live gram-negative S. typhimurium (Figure 5A) or gram-positive S. aureus (Figure S3A) infection. An enhanced intracellular killing of the engulfed S. aureus (Figure 5B) or S. typhimurium (Figure S3B) seen in wild-type macrophages after stimulation with a combination of BLP plus Tri-DAP or MDP was also absent in TLR2-deficient macrophages as well as in NOD1- or NOD2-deficient macrophages. These results suggest that TLR and NOD signaling are both critical for an efficient phagocyte-associated bactericidal activity.

We first examined whether co-stimulation of macrophages with TLR and NOD agonists induces an enhanced expression of phagocytic receptors. Stimulation with LPS, but not Tri-DAP or MDP, increased surface expression of CR3 and FcγR on macrophages, and a combined stimulation of LPS with Tri-DAP or MDP led to a further upregulation of CR3 and FcγR expression (Figure S4A). An enhanced expression of CR3 and FcγR was also observed in macrophages stimulated with BLP alone, which was further augmented in macrophages stimulated by a combination of BLP plus Tri-DAP or MDP (Figure S4B). All phagocytic processes including the engulfment of microbial pathogens within the phagocyte are primarily driven by a tightly controlled rearrangement of the actin cytoskeleton or actin polymerization (44), we next assessed actin polymerization in macrophages stimulated with TLR and NOD agonists. Significantly enhanced actin polymerizations as represented by the F-actin ratio were observed in macrophages stimulated by LPS in combination with Tri-DAP or MDP, but not by LPS, Tri-DAP, or MDP alone, upon gram-positive S. aureus or gram-negative S. typhimurium infection (p < 0.05, p < 0.01 vs. macrophages stimulated with LPS, Tri-DAP, or MDP alone) (Figure S5A). A combined stimulation of BLP with Tri-DAP or MDP also caused markedly increases in actin polymerization in response to S. aureus or S. typhimurium infection (p < 0.01 vs. macrophages stimulated with BLP, Tri-DAP, or MDP alone) (Figure S5B).

We further determined whether activation of TLR and NOD signaling by their specific agonists results in an accelerated phagosome maturation. Stimulation of macrophages with LPS, Tri-DAP, or MDP alone had no effect on phagosomal acidification; however, a combined stimulation of LPS with Tri-DAP or MDP substantially accelerated phagosomal acidification after engulfment of gram-negative S. typhimurium (p < 0.01 vs. macrophages stimulated with LPS, Tri-DAP, or MDP alone) (Figures 6A,C). A similar acceleration in phagosomal acidification was also seen in macrophages stimulated by a combination of BLP plus Tri-DAP or MDP after ingestion of gram-positive S. aureus (p < 0.01 vs. macrophages stimulated with BLP, Tri-DAP, or MDP alone) (Figures 6B,D). Consistent with an accelerated phagosomal acidification, macrophages stimulated by a combination of LPS or BLP plus either Tri-DAP or MDP displayed significantly increased phagolysosome fusion at 30, 60, and 90 min after ingestion of S. typhimurium (p < 0.05 vs. macrophages stimulated with LPS, Tri-DAP, or MDP alone) (Figures 6E,G) and S. aureus (p < 0.05 vs. macrophages stimulated with BLP, Tri-DAP, or MDP alone) (Figures 6F,H), as determined in a cell-free organelle system.

Upregulation and activation of membrane-trafficking regulators and lysosomal enzymes in macrophages during the process of bacterial phagocytosis are critical events for subsequent phagolysosome fusion and efficient killing of the ingested microbial pathogens (45, 46), we next examined whether co-stimulation of macrophages with TLR and NOD agonists upregulates membrane-trafficking regulator and lysosomal enzyme expression. Stimulation of macrophages with LPS, but not Tri-DAP or MDP, led to increased expression of membrane-trafficking regulators Rab10 and Stx1A as well as lysosomal enzymes Camp and Acp5 (p < 0.05 vs. PBS-treated macrophages); however, a combined stimulation of LPS with Tri-DAP or MDP maximized mRNA expressing levels of Rab10, Stx1A, Camp, and Acp5 when compared with macrophages stimulated with LPS, Tri-DAP, or MDP alone (p < 0.05, p < 0.01) (Figure S6A). A similar augmented expression of Rab10, Stx1A, Camp, and Acp5 was also observed in macrophages stimulated by BLP in combination with Tri-DAP or MDP (p < 0.01 vs. macrophages stimulated with LPS, Tri-DAP, or MDP alone) (Figure S6B). These results indicate that activation of both TLR and NOD signaling in macrophages results in upregulated phagocytic receptor expression, enhanced actin polymerization, accelerated phagosome maturation, and increased membrane-trafficking regulators and lysosomal enzymes in response to microbial infection.

To further clarify whether activation of both TLR and NOD signaling in vivo affords protection against microbial infection, wild-type mice were pretreated with PBS, LPS, Tri-DAP, or LPS plus Tri-DAP for 6 h, and further challenged with CLP-induced polymicrobial sepsis. Survival rates were recorded and monitored for at least 7 days. All mice receiving PBS succumbed within 60 h of septic challenge, while mice receiving LPS or Tri-DAP alone had a similar mortality rate at 95%, respectively, upon septic challenge (Figure 7A). However, mice that received a combination of LPS plus Tri-DAP were shown to be more resistant to polymicrobial sepsis, with an overall survival of 34% compared with the survival rate of 0% in mice that received PBS (p = 0.0112), 5% in mice that received LPS alone (p = 0.0275), and 5% in mice that received Tri-DAP alone (p = 0.0375) (Figure 7A). Consistent with a substantial survival advantage, mice treated with LPS plus Tri-DAP displayed moderate but significantly increased serum peak levels of TNF-α at 2 h and IL-6 at 6 h post septic challenge (p < 0.05 vs. mice treated with LPS or Tri-DAP alone) (Figure 7B). Furthermore, substantially reduced bacterial counts in the blood, spleen, and liver were observed at 12 and 24 h post septic challenge in mice treated with LPS plus Tri-DAP (p < 0.05, p < 0.01 vs. mice treated with LPS or Tri-DAP alone) (Figure 7C), indicating an accelerated bacterial clearance in these mice. Stimulation with a combination of BLP plus Tri-DAP also protected mice against polymicrobial sepsis-associated lethality, with a significant reduction in mortality from 100% in mice receiving PBS (p = 0.0091), 100% in mice receiving BLP alone (p = 0.0085), and 95% in mice receiving Tri-DAP alone (p = 0.0298) to 61% in mice receiving BLP plus Tri-DAP (Figure 7D). Consistent with the findings in mice treated with LPS plus Tri-DAP, mice treated with BLP plus Tri-DAP showed increased serum TNF-α and IL-6 (p < 0.05 vs. mice treated with BLP or Tri-DAP alone) (Figure 7E), and reduced bacterial load in the circulation and visceral organs (p < 0.05 vs. mice treated with BLP or Tri-DAP alone) (Figure 7F) post septic challenge. These results demonstrate that activation of both TLR and NOD signaling by the combined TLR and NOD agonists protects mice against microbial sepsis-associated lethality, which is associated with simultaneously augmented both inflammatory and antimicrobial responses.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.