Combined Stimulation of Toll-Like Receptor 5 and NOD1 Strongly Potentiates Activity of NF-κB, Resulting in Enhanced Innate Immune Reactions and Resistance to Salmonella enterica Serovar Typhimurium Infection

Inbred BALB/c female mice weighing 18 to 20 g purchased from the Pushchino nursery (Institute of Bioorganic Chemistry of the Russian Academy of Sciences, Pushchino, Russia) were used for cytokine and survival assays. Salmonella infection of female BALB/c mice was done at the Gamaleya Institute (Moscow, Russia) under biosafety level 2 (BSL-2)-equivalent conditions using NIH-approved ethical standards. For in vivo detection of NF-κB activity, 6- to 8-week-old BALB/c-Tg(IκBα-luc)Xen mice (Jackson Laboratory, Bar Harbor, ME) carrying an NF-κB-inducible luciferase reporter gene (Xenogen, Alameda, CA) were used at Roswell Park Cancer Institute (Buffalo, NY, USA) following protocols approved by the Roswell Park Cancer Institute IACUC. The mice were fed a complete pelleted laboratory chow and had access to food and tap water ad libitum.

The following PRR ligands were used: synthetic TLR2 and TLR5 agonists CBLB612 and CBLB502, respectively (Cleveland BioLabs, Inc. Buffalo, NY, USA), and NOD1 and NOD2 agonists C12-iE-DAP and L18-MDP, respectively (InvivoGen, San Diego, CA, USA). Lipopolysaccharide (LPS) purified by gel filtration chromatography was purchased from Sigma-Aldrich (USA).

A Bradford protein assay kit was purchased from Bio-Rad (USA), complete protease inhibitor cocktail tablets were from Roche Diagnostics (Deutschland GmbH, Mannheim, Germany), and tissue protein extraction reagent (T-PER) was purchased from Pierce (Rockford, IL, USA). Inhibitors celastrol, triptolide, gefitinib, and dexamethasone (DEX) were obtained from (InvivoGen, San Diego, CA, USA).

THP1-XBlue-CD14 cells (InvivoGen, USA) derived from THP-1 human acute monocytic leukemia cells were maintained at approximately 1 × 10⁶ cells/ml in RPMI 1640 (Gibco, USA) medium, supplemented with 10% fetal calf serum (Thermo scientific, USA), 50 U/ml penicillin, 50 μg/ml streptomycin, 2 mM l-glutamine, 0.1 M NaHCO₃ (all from PanEco, Russia), 200 μg/ml Zeocin (a formulation containing phleomycin D1), and 250 μg/ml G418 (both from InvivoGen, USA) at 37°C with 5% CO₂.

Salmonella enterica serovar Typhimurium IE147 strain was the kind gift of L. N. Nesterenko and Y. M. Romanova (N. F. Gamaleya Research Institute for Epidemiology and Microbiology, Moscow, Russia). Bacteria were cultured overnight in LB broth at 37°C with shaking at 300 rpm. An overnight culture of S. Typhimurium was washed with phosphate-buffered saline (PBS) and adjusted to 2.5 × 10⁷ CFU/ml. The number of CFU was determined on the next day by counting colonies that grew on Salmonella-Shigella (SS) agar (Himedia, India). Female BALB/c mice were injected subcutaneously (s.c.) with PBS or with CBLB502 (1 μg/mouse), C12-iE-DAP (200 μg/mouse), or their combination 9 h before oral administration of 5 × 10⁶ CFU (0.2 ml) of S. Typhimurium. The time period between PRR agonist injection and S. Typhimurium infection was selected based on previous experiments for induction of maximum protective effect (data not shown). For the survival experiment, mice (10 mice per group) were monitored for 35 days. For determination of bacterial load, spleens were isolated from mice (10 mice per group) at 3, 6, and 9 days after bacterial infection and homogenized in 0.5 ml of PBS using a FastPrep 24 device (MP Biomedicals, USA). Aliquots (100 μl) of diluted or undiluted homogenates were plated in duplicate on SS agar (Himedia, India). After overnight incubation at 37°C, colonies were counted manually.

Female BALB/c-Tg(IκBα-luc)Xen reporter mice were injected s.c. with PBS vehicle or with CBLB502 (1 μg/mouse) or C12-iE-DAP (200 μg/mouse) alone or in combination. At 2, 4, 6, 8, or 10 h after PRR agonist injection, mice were injected intraperitoneally (i.p.) with d-luciferin (3 mg/mouse; Promega, USA) and anesthetized using 2.5% isoflurane. Luminescence images were captured 2 min after d-luciferin injection using an IVIS Imaging System, 100 series (Caliper Life Sciences, USA), with an integration time of 10 s. In parallel, luciferase activity was also measured in tissue (liver, spleen, kidney, lung, and small and large intestine) homogenates prepared at the same time points following PRR agonist treatment as described above. Small and large intestine sections (approximately 3 cm long) were dissected 2 cm below the stomach (referred to as duodenum) and 5 cm below the cecum (referred to as colon), correspondingly. Sections were surgically isolated from omentum and feces and washed in ice-cold PBS. Organ homogenates were prepared at +4°C in 1× Reporter Lysis Buffer (Promega, USA) supplemented with protease inhibitor cocktail (Sigma-Aldrich, USA) using a FastPrep 24 device with Lysing Matrix A (MP Biomedicals, USA). All homogenates were normalized (10 mg) by protein concentration using Bradford reagent (Sigma-Aldrich, USA). To detect luciferase activity, aliquots of homogenates (50 μl) were mixed with 50 μl of Bright-Glo Luciferase Assay Buffer containing luciferin substrate in a 96-well plate (all from Promega, USA). Plates were briefly vortexed and then immediately read using a Wallac 1420 plate reader (PerkinElmer, USA).

Briefly, secreted embryonic alkaline phosphatase (SEAP) activity was determined as follows. Aliquots of culture medium were clarified by centrifugation at 14,000 × g for 2 min, heated at 65°C for 5 min to inhibit endogenous phosphatase activities, adjusted to 1× SEAP assay buffer (0.5 M carbonate, pH 9.8, 0.5 mM MgCl₂), and then incubated at 37°C for 10 min in a 96-well culture dish. Fifty microliters of 60 μM p-nitrophenylphosphate (Sigma-Aldrich, USA) dissolved in SEAP assay buffer (prewarmed to 37°C) was added to the mixture (to a final volume of 200 μl). The A₄₀₅ of the reaction mixture was read in a Wallac 1420 plate reader (PerkinElmer, USA). SEAP activity is given in milliunits (mU) per ml. One milliunit is defined as the amount of phosphatase that hydrolyzes 1.0 pmol of p-nitrophenylphosphate per min, and this corresponds to an increase of 0.04 mU per min.

For evaluation of p65 nuclear translocation, THP1-XBlue-CD14 cells were seeded in duplicate in 6-cm dishes at a density of 1 × 10⁶ cells/ml. The next day, cells were treated with NOD1 and TLR5 agonists, alone or in combination, or left untreated. Thirty minutes after addition of agonists, cells were harvested, and nuclear and cytoplasmic cell extracts were prepared using an NE-PER kit (Thermo Fisher Scientific, USA) according to the manufacturer's instructions. For detection of PRR expression levels, total protein extracts from THP-1 cells were prepared using radioimmunoprecipitation assay (RIPA) buffer supplemented with protease inhibitor cocktail (all, Sigma-Aldrich, USA). For in vivo detection of antimicrobial peptides, small intestine homogenates were prepared 16 h after s.c. administration of PRR agonists as described below (see the paragraph “Ex vivo cytokine/chemokine assay”). All samples were normalized based on total protein concentrations measured using Bradford reagent (Sigma-Aldrich, USA). Samples were mixed with 2× Laemmli sample buffer (Sigma-Aldrich, USA) and run under denaturation conditions on 15% polyacrylamide gels. Proteins were transferred to nitrocellulose Hybond C membranes (GE Healthcare, USA) using a semidry Trans-Blot Transfer Cell (Bio-Rad, USA). Primary antibodies were anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH), anti-p65 (Santa Cruz Biotechnology, CA, USA), anti-TLR5, anti-NOD1, and anti-beta-defensin-3 (Santa Cruz Biotechnology, USA). Horseradish peroxidase (HRP)-conjugated secondary anti-rabbit and anti-mouse antibodies were from GE Healthcare (GE Healthcare, Germany). Antibody-bound protein bands were visualized using an enhanced chemiluminescence (ECL) detection kit and cassette on Hyperfilm membrane (GE Healthcare, Germany).

For ex vivo cytokine/chemokine assays, BALB/c mice were injected s.c. with PBS or with CBLB502 (1 μg/mouse), C12-iE-DAP (200 μg/mouse), or their combination. Two hours later, samples of small intestine, large intestine, and peripheral blood from tail vein were collected. Tissue samples were placed immediately into ice-cold T-PER extraction buffer containing complete protease inhibitor (Sigma-Aldrich, USA) and homogenized using a FastPrep 24 device with Lysing Matrix A (MP Biomedicals, USA). The homogenates were centrifuged at 12,000 rpm for 12 min at +4°C. All supernatant samples were normalized (to 10 mg/ml) by total protein concentration as measured using Bradford reagent (Sigma-Aldrich, USA). Peripheral blood samples were incubated for 20 min at +37°C for clot formation. Serum samples were obtained using subsequent centrifugation at 1,000 rpm for 10 min. The levels of 20 cytokines and chemokines (IL-1α, -2, -4, -5, -6, -10, -13, -17, -21, -22, -27, gamma interferon [IFN-γ], TNF-α, CXCL1/keratinocyte-derived chemokine [KC], monocyte chemotactic protein 3 [MCP-3], MCP-1, macrophage inflammatory protein 1α [MIP-1α], MIP-1β, RANTES, and granulocyte-macrophage colony-stimulating factor [GM-CSF]) were measured in the prepared mouse serum and small and large intestine homogenate samples using mouse Th1/Th2 and chemokine bead-based FlowCytomix kits (Bender MedSystems GmbH, Austria) according to the manufacturer's instructions.

THP1-XBlue-CD14 cells carrying an NF-κB-dependent SEAP reporter gene were seeded in 96-well plates at 10⁵ cells per well in complete RPMI 1640 medium. Eighteen hours after cells were treated with PRR ligands, plates were centrifuged at 1,000 rpm for 10 min, and culture supernatants were collected. Levels of 19 cytokines and chemokines (IL-1β, -2, -4, -5, -6, -8, -9, -10, -12p70, -13, -17A, -22, IFN-γ, TNF-α, MCP-1, MIP-1α, MIP-1β, G-CSF, and monokine induced by IFN-γ [MIG]) were measured in the prepared supernatants in triplicate using human Th1/Th2/Th9/Th17/Th22 and chemokine bead-based FlowCytomix kits (Bender MedSystems GmbH, Austria) according to the manufacturer's instructions.

The data shown are representative results. Experimental values are given as the means ± standard deviations (SD) of triplicate assays. Groups were compared using Student's t test. In the mouse infection study Kaplan-Meier survival curves were compared using a log rank test. P values of less than 0.05 were considered statistically significant.

Previous work showed that costimulation of NODs (NOD1 and NOD2) and TLRs (TLR2, TLR4, and TLR9) in various cell types resulted in enhanced production of several cytokines, such as interleukin-1β (IL-1β), IL-6, IL-8, and tumor necrosis factor alpha (TNF-α) compared to stimulation of a single type of PRR (9, 10, 12–14). However, the molecular mechanism(s) underlying these enhanced responses were not defined. Since NF-κB controls expression of many cytokines and is activated by both TLR5 and NOD1 (11), we hypothesized that synergistic activation of NF-κB was responsible for the enhanced effect of costimulation with TLR5 and NOD1 (TLR5+NOD1) on cytokine production. We tested this hypothesis in vitro using THP1-XBlue-CD14 cells expressing endogenous NOD1 and TLR5 receptors (Fig. 1A) along with an introduced secreted embryonic alkaline phosphatase (SEAP) reporter gene controlled by an NF-κB-dependent promoter. These reporter cells were left untreated or treated with NOD1 and TLR5 agonists alone or in combination. For stimulation of NOD1, we used C12-iE-DAP, an acylated derivative of iE-DAP (d-glutamyl-meso-diaminopimelic acid, the moiety in bacterial molecules recognized by NOD1) (7). For activation of TLR5, we used CBLB502, a pharmacologically optimized derivative of the natural TLR5 agonist, bacterial flagellin (15). Based on SEAP expression in cells treated with C12-iE-DAP or CBLB502 as a single agent over a wide range of concentrations, the NOD1 agonist was a much weaker NF-κB activator than the TLR5 agonist (Fig. 1B)]. Interestingly, however, when used in combination with the TLR5 agonist, concentrations of NOD1 ligand that did not produce substantial NF-κB-dependent responses by themselves markedly increased (up to 3.88-fold) the already strong NF-κB-dependent response generated by TLR5 stimulation. Similarly, addition of CBLB502 at suboptimal concentrations significantly potentiated the NF-κB response after NOD1 stimulation (up to 6.76-fold). For many tested doses (from 10 pg/ml up to 100 μg/ml), the level of NF-κB-dependent reporter expression detected after combined stimulation of TLR5 and NOD1 was greater than the sum of the levels seen in cells with stimulation of only one receptor type. It is important to note that detected levels of NF-κB activity after combined stimulation of TLR5 and NOD1 could be reached using up to 100-fold higher doses of isolated agonist or in some cases could not be reached at all. These results indicate that costimulation of TLR5 and NOD1 receptors has a synergistic, rather than additive, effect on activation of NF-κB.

The results obtained in the SEAP reporter system were confirmed using a second, non-reporter-based method to assess NF-κB activation: Western blot analysis of nuclear translocation of the p65 subunit of NF-κB (Fig. 1C). In this experiment, suboptimal concentrations of TLR5 (10 ng/ml) and NOD1 (100 ng/ml) agonists applied to THP-1 cell cultures as single agents had no effect or a minimal effect on p65 nuclear accumulation, whereas their combination induced substantial p65 translocation.

It is noteworthy that combined stimulation of other TLRs (TLR2, TLR4, and TLR5) with either NOD1 or NOD2 produced similar synergistic effects on NF-κB activation (see Fig. S1 in the supplemental material). Therefore, positive cross talk between members of the TLR and NOD receptor families appears to be a general phenomenon that is not restricted to TLR5 and NOD1.

It was previously shown that combined stimulation of TLR2, TLR4, or TLR9 with NOD1 or NOD2 significantly enhanced production of IL-8 in cultured THP-1 cells (13). Here, we examined whether combined NOD1 and TLR5 stimulation enhanced cytokine production in THP-1 cells compared to stimulation of only NOD1 or TLR5. Levels of 19 cytokines (see Materials and Methods) were measured in culture supernatants collected 18 h after addition of C12-iE-DAP and/or CBLB502 to cell cultures using bead-based FlowCytomix kits. For isolated stimulation of NOD1 and TLR5 receptors, we used C12-iE-DAP and CBLB502, respectively, in final concentrations ranging from 10 pg/ml to 100 μg/ml. For combined stimulation, we used a fixed suboptimal C12-iE-DAP concentration (1 μg/ml) previously determined to provide maximal enhancement of the NF-κB response when used in combination with different concentrations of CBLB502. As seen for NF-κB activation, combined stimulation of NOD1 and TLR5 receptors resulted in significant enhancement of production of 5 of the 19 analyzed cytokines compared to stimulation of either single receptor type. Cytokines that showed a synergistic (greater than additive) response to combined TLR5+NOD1 stimulation included IL-1β (up to a 4.4-fold increase), IL-8 (up to a 7.8-fold increase), MIP-1α (up to a 13.2-fold increase), MIP-1β (up to a 6.2-fold increase), and TNF-α (up to a 4.1-fold increase) in comparison to isolated stimulation (Fig. 2).

Having demonstrated that costimulation of TLR5 and NOD1 leads to both potentiation of NF-κB activity and production of cytokines, we next tested whether NF-κB was required for the effect on cytokines. This was accomplished by using several inhibitors for blocking the RIP2-TAK1-IKK-NF-κB pathway: (i) gefitinib for inhibition of the upstream common signal kinase of TLR and NOD receptors (RIP2) (16), (ii) celastrol for inhibition of the RIP2-activated TAK1 signal kinase (17), (iii) triptolide for inhibition of NF-κB transcriptional activation (18), and (iv) synthetic glucocorticoid dexamethasone (DEX) to inhibit the activity of both the NF-κB and mitogen-activated protein kinase (MAPK)–AP-1 pathways (19). Celastrol (5 μM), triptolide (10 nM), gefitinib (10 μM), and DEX (100 μg/ml) were added in final concentrations to cultures of THP-1 cells 1 h prior to addition of PRR agonists (C12-iE-DAP at 1 μg/ml and/or CBLB502 at 1 μg/ml). Eighteen hours later, levels of NF-κB activity (NF-κB-dependent SEAP reporter expression) and cytokine production were assessed (Fig. 3A and B). IL-1β and TNF-α were measured since their production was found to be strongly enhanced by combined stimulation of NOD1 and TLR5. The results of this experiment showed that nontoxic concentrations of NF-κB inhibitors abrogated not only potentiation of the NF-κB response but also production of IL-1β and TNF-α. These data show the important role of enhancement of NF-κB activity in potentiation of cytokine production levels after combined TLR5 plus NOD1 stimulation. An inhibitor of both AP-1 and NF-κB pathways, DEX, was slightly more efficient in reducing production of IL-1β and TNF-α, reflecting a possible role of the MAPK–AP-1 pathway in the potentiation effect of cytokine production (9).

To determine whether the synergistic activation of NF-κB by combined NOD1 and TLR5 stimulation that was observed in vitro also occurs in vivo, we used BALB/c-Tg(IκBα-luc)Xen reporter mice carrying an NF-κB-dependent luciferase reporter gene. NF-κB activity in these mice can be detected by live imaging of bioluminescence in intact sedated animals after administration of NF-κB-activating agents (PRR ligands, UV radiation, etc.) and luciferin (20).

NF-κB-Luc transgenic mice were injected s.c. with 100 μl of C12-iE-DAP (200 μg/ml) or CBLB502 (1 μg/ml) alone or in combination. Doses of individual agonists resulting in suboptimal activation of NF-κB activity were determined in preliminary experiments (data not shown). Control mice were injected s.c. with 100 μl of PBS. NF-κB activity was detected by live imaging of mice at 2, 4, 6, 8, or 10 h after administration of PRR agonists by i.p. injection of d-luciferin 2 min before imaging. The obtained data showed that the NOD1 agonist C12-iE-DAP induced significantly lower levels of NF-κB activity than the TLR5 agonist CBLB502 (Fig. 4A). When used as single agents, both PRR agonists caused maximal reporter induction at 2 to 4 h postinjection. The combination of C12-iE-DAP and CBLB502 also induced maximal NF-κB reporter activity at 2 to 4 h after administration. However, compared to single-agonist treatments, combined-agonist treatment produced more sustained NF-κB activation, which was strong even at 10 h after agonist injection, when the signals induced by either agent alone were practically undetectable. These imaging data also revealed strong enhancement of NF-κB activity in the abdominal region of mice that received the combination of agonists.

To obtain more precise quantitative results, we measured luciferase activity in tissue lysates of livers, spleens, kidneys, lungs, and small and large intestines isolated from NF-κB-Luc transgenic mice at different times after administration of PRR agonists (Fig. 4B). This showed that the degree of enhancement of reporter activation induced by combined stimulation of NOD1 and TLR5 compared to stimulation of either receptor type alone varied greatly between different organs. There was no significant enhancement detected in the liver and spleen with administration of combined agonists compared to administration of CBLB502 alone. In contrast, the luciferase signal in small intestines, large intestines, lung, and kidney was significantly higher in mice treated with combined NOD1 and TLR5 agonists than in mice treated with either agonist alone. Maximal levels of enhancement of NF-κB activation by the combination of agonists was detected in the small intestines. In this tissue, NF-κB showed synergistic activation at 4, 6, 8, and 10 h following combined PRR agonist treatment. The greatest effect was seen at the 10-h time point, with NF-κB showing 2.76-fold, 8.24-fold, and 60-fold activation (relative to PBS treatment) in samples from CBLB502-treated, C12-iE-DAP-treated, and CBLB502- and C12-iE-DAP-treated mice, respectively. Thus, combined TLR5+NOD1 stimulation led to induction of NF-κB that was 7.5-fold stronger than the sum of the induction produced by treatment with CBLB502 and C12-iE-DAP separately. It is notable that the maximal synergistic effect in the small intestine was observed at 10 h after agonist administration; at this late time point, NF-κB activity had significantly declined in all other tissues. Synergistic activation of NF-κB was also observed in the large intestine, kidney, and lung; however, its scale and duration were lower than in the small intestine. To determine whether the high level of NF-κB activity reached after combined stimulation of NOD1 and TLR5 could be achieved by using higher doses of TLR5 agonist as a single agent, we treated reporter mice with a range of ascending CBLB502 doses (from 0.04 μg to 25 μg/mouse in 5-fold increments) for 4 h (Fig. 5). In most of the studied organs, including kidney, spleen, large intestine, and liver, the levels of reporter activation reached by combined stimulation of NOD1 and TLR5 could be achieved by using higher doses of CBLB502 as a single agent. Significant enhancement of NF-κB activity in lungs after combined stimulation of NOD1 and TLR5 was likely due to strong NF-κB activation in response to C12-iE-DAP administration (the effect of C12-iE-DAP alone at 4 h is shown in Fig. 4B). In the small intestine, however, the enhanced level of NF-κB activity induced by combined stimulation of NOD1 and TLR5 could not be reached by much higher doses (25-fold increment) of CBLB502 administered as a single agent and could not be explained by an additive response to C12-iE-DAP (Fig. 4B).

Having demonstrated that combined NOD1 and TLR5 stimulation leads to synergistic activation of NF-κB in mouse small intestines, we examined its biological outcome in terms of cytokine production in vivo. For this purpose, BALB/c mice were injected s.c. with 100 μl of C12-iE-DAP (200 μg/ml) or CBLB502 (1 μg/ml) alone or in combination. Control mice were injected s.c. with 100 μl of PBS. Two hours after administration of PRR ligands, mice were euthanized, and small intestine, liver, and blood samples were isolated for measurement of cytokine levels in tissue homogenates and blood serum using bead-based immunoassays as described above. In mouse serum, production of 3 of the 20 measured cytokines (IL-6, IL-22, and TNF-α) was enhanced following combined TLR5+NOD1 stimulation compared to stimulation of either receptor alone (Fig. 6A). In mouse small intestine samples, 6 of the 20 measured cytokines (IL-5, -6, -13, -21, -22, and TNF-α) showed enhanced production following combined TLR5+NOD1 stimulation (Fig. 6B). In both serum and small intestine samples, combined TLR5+NOD1 stimulation had a synergistic effect (greater than the additive effect of single-receptor stimulation) on production of at least some of the induced cytokines, including IL-6 in serum and IL-5, IL-13, and TNF-α in small intestines.

In contrast to what was observed in serum and small intestine samples, none of the analyzed cytokines showed enhanced production in liver samples from mice treated with combined TLR5+NOD1 agonists compared to each agonist alone (data not shown). These data demonstrate that combined stimulation of NOD1 and TLR5 receptors has both local, organ-specific (particularly in the small intestine) effects as well as systemic (blood serum) effects involving synergistic cytokine production. The observed correlation between enhancement of cytokine production and enhancement of NF-κB activation (see above) is consistent with potentiation of the cytokine response being NF-κB dependent.

As an additional indicator of the effect of combined TLR5+NOD1 stimulation on NF-κB-dependent immune responses, we evaluated production of beta-defensin-3, which was previously shown to be induced during bacterial infection (21). Using Western blot analysis of small intestine homogenates, we found that stimulation of TLR5 with CBLB502 induced production of beta-defensin-3. There was no production of beta-defensin-3 following stimulation of NOD1 with C12-iE-DAP; however, combined stimulation of NOD1 and TLR5 led to increased production of these peptides relative to that observed with TLR5 stimulation alone (Fig. 6C). Taken together, these results clearly indicate that combined stimulation of TLR5 and NOD1 receptors potentiates NF-κB activation and downstream immune responses not only in vitro but also in vivo.

The results described above suggested that combined TLR5 and NOD1 stimulation might lead to more effective antimicrobial defense responses than stimulation of either receptor alone, particularly in the small intestine. To test this possibility in vivo, we used an enteropathogenic bacterial strain, Salmonella enterica serovar Typhimurium. BALB/c mice were injected s.c. with PBS or with C12-iE-DAP (200 μg/ml), CBLB502 (1 μg/ml), or their combination 9 h before oral infection with 5 × 10⁶ CFU of S. Typhimurium in a volume of 0.5 ml. Bacterial loads were monitored in spleens isolated 3, 6, and 9 days after infection (Fig. 7A). The bacterial loads in spleens of mice treated with C12-iE-DAP or CBLB502 alone were not significantly different from those in spleens from PBS-treated mice. In contrast, spleens from mice treated with a combination of C12-iE-DAP and CBLB502 to stimulate both NOD1 and TLR5 had 10-fold fewer bacterial CFU. This reduced bacterial load resulted in significantly prolonged mouse survival: on day 35 after infection, 80% of mice treated with both C12-iE-DAP and CBLB502 were alive, compared to only 20% of mice treated with PBS or either PRR agonist alone (Fig. 7B). Therefore, in this animal model of enteropathogenic bacterial infection, combined stimulation of NOD1 and TLR5 receptors is critically important for efficient bacterial clearance and improved mouse survival. These results illustrate the key role that synergistic activation of immune responses stemming from stimulation of multiple PRRs likely plays in a broad spectrum of scenarios of in vivo pathogen exposure.