RIPK3 Promotes Mefv Expression and Pyrin Inflammasome Activation via Modulation of mTOR Signaling

Mutations in the pyrin-encoding gene MEFV are associated with familial Mediterranean fever, an autoinflammatory disorder (1, 2). Pyrin has been recognized as an innate sensor that assembles an inflammasome complex with apoptosis-associated speck-like protein containing a caspase recruitment domain and caspase-1 in response to Rho modifications induced by bacterial toxins (3–5). Rho modification is a common mechanism employed by bacteria to hijack the host cytoskeleton and subvert effector responses (6). The ability of the host cell to identify this subversion is essential for an effective innate immune response.

Clostridium difficile is an enteric pathogen whose prevalence has been increasing because of microbial dysbiosis induced by prolonged antibiotic use and compromised immune status (7, 8). C. difficile produces two large exotoxins, toxin A (TcdA) and toxin B (TcdB), that glycosylate and inactivate Rho, thereby instigating the antipathogen host response (9). Toxin production by C. difficile is an integral part of its pathogenesis, and several treatment regimens target toxin neutralization or absorption to alleviate the clinical symptoms (8–11). C. difficile mutants that lack Rho glycosylating toxins have significantly reduced pathology, and challenge with toxin alone is sufficient to promote pathology in various animal models (9, 12, 13). These findings highlight the importance of the toxin-induced immune response in C. difficile pathogenesis and the need to investigate the underpinnings of toxin-mediated pathology.

TcdB-mediated Rho inactivation was recently shown to activate the pyrin inflammasome. This makes pyrin a unique innate sensor because it recognizes a bacteria-induced host modification instead of a molecular pattern associated with pathogens (3). Pyrin activation has diverse roles in host defense, and its dysregulation can be detrimental during infectious or sterile insults. Pyrin inflammasome activation contributes to inflammation and pathology in a murine model of familial Mediterranean fever (14, 15), promotes host defense in response to Burkholderia cenocepacia infection (5, 16), and is required for maintenance of epithelial barrier integrity during mucosal injury (17). Furthermore, inflammasome activation in general has been shown to be detrimental to the host during C. difficile infection and can modulate various aspects of the resulting inflammation (18). Although Rho modifications play a key role in activating the pyrin inflammasome, little is known about the details of the molecular pathways involved in this process.

To fill this gap in knowledge, we have investigated the pathways involved in pyrin inflammasome activation and previously identified a role for TNF signaling (19). TNF signaling promotes major cellular pathways, including inflammation, cell survival, and cell death (20). Receptor-interacting protein kinase (RIPK) 1 and RIPK3 are critical components of a signaling network downstream of the TNF receptor. The fate of a cell in response to a TNF stimulus varies between NF-κB–induced cell survival, Fas-associated death domain/caspase-8–induced apoptotic cell death, and mixed lineage kinase domain-like pseudokinase (MLKL)–induced necroptotic cell death (21). We sought to investigate the impact of pathways downstream of TNF signaling on pyrin inflammasome activation.

In this study, we demonstrate that RIPK3 is involved in pyrin inflammasome activation, and this inflammasome activation is independent of the necroptotic and apoptotic cell death pathways. We further demonstrate that RIPK3 modulates the mechanistic target of rapamycin (mTOR) pathway to regulate Mefv expression and pyrin inflammasome activation. This study highlights the impact of RIPK3 and mTOR modulation on pyrin inflammasome activation.

Pyrin^−/− (22), Casp1^−/− (23), Tnf^−/− (24), Nlrp3^−/− (25), Ripk3^−/− (26), Mlkl^−/− (27), Casp3^−/− (28), Casp7^−/− (29), Myd88^−/− (30), and Trif^−/− (31) mice have been previously described. Mice were maintained in a specific pathogen-free facility, and animal studies were approved by St. Jude Children’s Research Hospital Committee on the Use and Care of Animals.

C. difficile r20291 AB⁻ and AB⁺ strains were provided by Dr. N. Minton, and the toxin was prepared as described previously (9). Briefly, C. difficile strain r20291 (AB⁻ and AB⁺) was cultured in tryptone–yeast extract media for 24 h in an anaerobic chamber at 37°C. Cultures were diluted to an OD of 1 (corresponding to 2 × 10⁷ CFU/ml) and spun down, and the supernatant was sterilized using 0.22-μM filters. The supernatant prepared from the toxin-positive strain was used to stimulate bone marrow–derived macrophages (BMDMs) at a 1:5 dilution. This stimulation is referred to as “C. difficile toxin” stimulation. The supernatant prepared from the toxin-negative strain is referred to as “control” stimulation and was included in all experiments with crude C. difficile toxin as the stimulus.

BMDMs were generated as previously described (32). For pyrin inflammasome activation, cells were resuspended in Opti-MEM (Life Technologies) and stimulated with C. difficile supernatant (AB⁻ [control] or AB⁺) for 12–16 h. Alternately, cells were stimulated with 0.2 μg/ml TcdB (List Biological Laboratories) or medium for 6–8 h. For mTOR inhibition, 1 μM rapamycin (InvivoGen), 200 nM torin 1 (Selleck Chemicals), or 200 nM PP242 (Selleck Chemicals) was added to cells 30 min prior to stimulation with toxins.

BMDMs differentiated from specific mouse strains were seeded in 12-well cell culture plates and treated with the indicated stimuli for predetermined amounts of time. Image-based light microscopy data were collected using an Olympus CKX41 microscope with a ×40 objective lens. The acquired data were digitally analyzed using the INFINITY ANALYZE Software (Lumenera).

Cell death was analyzed using a two-color Incucyte ZOOM in an incubator imaging system (Essen BioScience). BMDMs were subcultured at 1.0 × 10⁶ cells/well in treated 12-well cell culture plates and were treated with different experimental conditions inducing cell death in the presence of 100 nM of the cell-impermeable DNA binding fluorescent dye SYTOX Green (S7020; Life Technologies). Loss of membrane integrity in the dying cells results in uptake and positive staining for the SYTOX Green dye. Several images at different timepoints were collected using a ×20 objective and analyzed using the Incucyte S3 software, which allows precise quantification of the number of SYOTX Green–positive dead cells present in each image. The number of dead cells for each of the stimulations and mouse strains was exported for quantitative analysis of the total cell death.

BMDM cell lysates and supernatants were combined in caspase lysis buffer (protease inhibitors, phosphatase inhibitors, 10% NP40, and 25 mM DTT) and boiled in NaDodSO₄ (SDS) sample buffer for Western blot analysis. These lysates were used for analysis of caspase processing. For signaling analysis, BMDMs were lysed in complete RIPA buffer (containing protease inhibitors and phosphatase inhibitors [Calbiochem]) and boiled in SDS sample buffer for Western blot analysis. Pyrin phosphorylation status was assessed as previously described (33) using Mn²⁺-based Phos-tag gel analysis (FUJIFILM Wako Chemicals). Proteins were separated by electrophoresis using 6–12% polyacrylamide gels. Following electrophoresis, proteins were transferred to PVDF membranes (MilliporeSigma), blocked in 5% skim milk to reduce nonspecific binding, and probed with primary Abs. Membranes were then washed and incubated with appropriate horseradish peroxide–conjugated secondary Ab (1:5000; Jackson ImmunoResearch Laboratories). Proteins were visualized using the Luminata Forte Western horseradish peroxide substrate (MilliporeSigma). The primary Ab used for caspase processing analysis was anti–caspase-1 (AG-20B-0042-C100, 1:3000; AdipoGen). Abs used for signaling immunoblotting were anti–p-ERK1/2 (no. 9101, 1:1000; Cell Signaling Technology), anti–total-ERK1/2 (no. 9102, 1:1000; Cell Signaling Technology), anti–p-IκBα (no. 2859, 1:1000; Cell Signaling Technology), anti–total-IκBα (no. 9242, 1:1000; Cell Signaling Technology), anti–p-p38 (no. 9211, 1:1000; Cell Signaling Technology), anti–p-JNK (no. 9251, 1:1000; Cell Signaling Technology), anti–p-pyrin S241 (ab200420, 1:1000; Abcam), anti-pyrin (ab195975, 1:1000; Abcam), anti–14-3-3 (no. 8312, 1:1000; Cell Signaling Technology), anti–p-mTOR S2481 (no. 2974, 1:1000; Cell Signaling Technology), anti–p-mTOR S2448 (no. 2971, 1:1000; Cell Signaling Technology), anti–p-GSK3β (no. 5558, 1:1000; Cell Signaling Technology), anti–p-S6K (no. 9205, 1:1000; Cell Signaling Technology), and anti–p-S6 (no. 4856, 1:1000; Cell Signaling Technology). Anti-GAPDH (no. 5174, 1:1000; Cell Signaling Technology) was used as a control.

RNA was extracted using TRIzol (Thermo Fisher Scientific) according to the manufacturer’s instructions. The isolated RNA was reverse transcribed using the First-Strand cDNA Synthesis Kit (Applied Biosystems), and real-time quantitative PCR was performed using 2× SYBR Green (Applied Biosystems) using appropriate primers on Applied Biosystems 7500 RT-PCR instrument. RT-PCR primer sequences are TNF: forward 5′-CATCTTCTCAAAATTCGAGTGACAA-3′, reverse 5′-TGGGAGTAGACAAGGTACAACCC-3′; pyrin: forward 5′-TCATCTGCTAAACACCCTGGA-3′, reverse 5′-GGGATCTTAGAGTGGCCCTTC-3′, forward primer 2 5′-AGGCTTCAAGGACTTTACAACAA-3′, reverse primer 2 5′-TCATGCGAATGAGACTCCCA-3′; and GAPDH: forward 5′-CGTCCCGTAGACAAAATGGT-3′, reverse 5′-TTGATGGCAACAATCTCCAC-3′.

Lactate dehydrogenase (LDH) assay (Promega, Madison, WI) was carried out as per the manufacturer’s instructions. Briefly, cell culture supernatants were incubated with the substrate at 37°C for 15 min, and the end point colorimetric assay was read at 450 nM. A standard curve was generated using the cellular lysate of known density and percentage of cell death extrapolated using the curve.

Toxin was prepared as described above, and 1 ml (control or toxin) was injected i.p. into gender-matched mice at 8–10 wk of age. Four hours postinjection, blood was harvested through cardiac puncture, and serum was isolated for cytokine analysis.

Cytokines in the serum and cell culture supernatants were measured by ELISA, according to the manufacturers’ instructions. The IL-18 and multiplex ELISA kits were obtained from eBioscience and MilliporeSigma, respectively.

All statistical analysis was performed using Prism v6.0 software. Student t test, one-way ANOVA followed by Fisher least significant difference (LSD) or Kruskal–Wallis followed by Dunn posttest was used for statistical analysis as indicated. The p values <0.05 were considered significant.

The pyrin inflammasome is activated in response to purified C. difficile toxins TcdA and TcdB in BMDMs (3, 33), and we have previously shown that loss of TNF leads to reduced caspase-1 cleavage, IL-18 release, and pyroptotic cell death in response to pyrin inflammasome activation by C. difficile toxin (19). In this study, we went on to test whether the loss of major components of the TNF signaling pathway affected pyrin inflammasome activation. Wild type (WT) BMDMs stimulated with C. difficile toxin underwent pyrin inflammasome activation, indicated by pyrin-dependent caspase-1 cleavage and IL-18 maturation (Fig. 1A, 1B), and pyroptotic cell death, indicated by LDH release and membrane permeability (Fig. 1C–E). Loss of RIPK3 resulted in a reduction in pyrin inflammasome activation (Fig. 1A–E), demonstrating that RIPK3 promotes pyrin inflammasome activation in response to C. difficile toxin.

RIPK3 has important roles in regulating inflammatory signaling and promoting necroptosis (34). We therefore tested whether disrupting necroptosis impacted pyrin inflammasome activation. Loss of MLKL, the necroptosis executioner downstream of RIPK3, did not affect pyrin inflammasome activation as levels of caspase-1 cleavage, IL-18 release, and pyroptotic cell death were similar between WT and Mlkl^−/− BMDMs treated with C. difficile toxin (Fig. 2A–D). This suggests that RIPK3 promotes pyrin inflammasome activation independent of its role in necroptosis. Similarly, the loss of apoptosis executioners, caspase-3 and -7, also did not affect levels of caspase-1 cleavage, IL-18 release, and pyroptotic cell death in response to treatment with C. difficile toxin (Fig. 2E–H). These data demonstrate that the function of RIPK3 in promoting pyrin inflammasome activation is independent of its role in necroptosis and independent of apoptosis.

Supernatants obtained from C. difficile cultures contain various pathogen-associated molecular patterns that can modulate inflammatory signaling (35, 36). The pyrin inflammasome has been previously shown to be primed by TLR signaling (22). We assessed the role of RIPK3 in the activation of inflammatory signaling pathways downstream of TLRs including MAPK and NF-κB. C. difficile toxin stimuli promoted the phosphorylation of ERK, p38, JNK, and IκBα, demonstrating that these pathways were activated in response to the bacterial components (including toxin) (Fig. 3A). The loss of RIPK3 did not affect the activation of MAPK and NF-κB pathways or the expression of 14-3-3 (Fig. 3A). Pyrin expression, in contrast, was notably reduced in the absence of RIPK3 (Fig. 3A). To test whether differential pyrin expression was caused by transcriptional regulation, we assessed the level of Mefv transcript in response to the stimuli. The Mefv transcript levels mirrored the protein expression pattern and were reduced in Ripk3^−/− cells compared with WT cells, both under basal and stimulated conditions, whereas Tnf transcript levels were similar between Ripk3^−/− and WT cells (Fig. 3B).

Given the relevance of the TNF–TNFR axis in pyrin activation (19) and the role of RIPK3 in TNFR signaling, we assessed the role of TLR signaling in TNF induction. TLR signaling relies on the adaptor proteins MyD88 and TRIF; we therefore tested pyrin inflammasome activation in macrophages deficient in MyD88 or TRIF. MyD88 was required for caspase-1 cleavage (Supplemental Fig. 1A), IL-18 release (Supplemental Fig. 1B), and cell death induced by C. difficile toxin (Supplemental Fig. 1C, 1D), whereas TRIF was dispensable (Supplemental Fig. 1A, 1B). We further found that MyD88, and not TRIF, was required for TNF production and Mefv expression and pyrin production in response to C. difficile toxin (Supplemental Fig. 1B, 1E, 1F). These data demonstrate that MyD88 is required for the TNF production that engages the RIPK3 pathway to promote pyrin activation in response to C. difficile toxin stimuli.

To specifically test whether pyrin inflammasome activation is independent of the signaling engaged by other pathogen-associated molecular patterns in the C. difficile toxin stimuli, we stimulated BMDMs with purified TcdB. Similar to the response to C. difficile toxin stimuli, pyrin inflammasome activation in response to purified TcdB was significantly reduced in Ripk3^−/− BMDMs compared with WT BMDMs as indicated by reduced caspase-1 cleavage, IL-18 release, and cell death (Fig. 3C, 3D, Supplemental Fig. 2).

Recent studies have identified pyrin dephosphorylation as a critical step upstream of pyrin inflammasome activation (33, 37, 38). We therefore assessed whether pyrin dephosphorylation was affected in the absence of RIPK3. We found that the loss of RIPK3 reduced pyrin expression but did not affect pyrin dephosphorylation (Fig. 3E). Overall, these data suggest that the modulation of pyrin inflammasome activation by RIPK3 occurs at the level of Mefv transcript and protein expression and is independent of MAPK and NF-κB signaling and pyrin dephosphorylation.

In addition to MAPK and NF-κB signaling, Mefv expression is regulated by the mTOR (39) and PI3K pathways (40). We assessed mTOR activation in BMDMs following C. difficile toxin stimuli exposure. We observed increased phosphorylation of mTOR, GSK3β, and downstream targets S6K and S6 in the absence of RIPK3 (Fig. 4A). These data demonstrate that RIPK3 restricts mTOR activation following C. difficile toxin stimuli. To assess whether mTOR activation modulates pyrin inflammasome activation, we inhibited mTOR activation using rapamycin treatment during stimulation with C. difficile toxin (Fig. 4B). Treatment with rapamycin increased the amount of caspase-1 cleavage and IL-18 released in response to the C. difficile toxin stimuli (Fig. 4C, 4D). Additionally, rapamycin treatment increased Mefv transcript levels independent of MAPK or NF-κB modulation (Supplemental Fig. 3A, 3B). To confirm that mTOR inhibition specifically led to pyrin inflammasome activation in response to C. difficile toxin stimuli, we tested the effect of rapamycin on C. difficile toxin–induced inflammasome activation in Nlrp3^−/− BMDMs (41). We observed that rapamycin promoted C. difficile toxin–induced inflammasome activation that was independent of NLRP3 and dependent on pyrin (Supplemental Fig. 3C, 3D).

To confirm the role of mTOR signaling in pyrin inflammasome activation, we also tested the effect of mTOR inhibitors PP242 and torin 1 on pyrin inflammasome activation. Similar to the effects of rapamycin treatment, treatment with PP242 and torin 1 promoted pyrin inflammasome activation in response to C. difficile toxin stimuli (Fig. 4E, 4F). In line with the observed pyrin inflammasome activation, mTOR inhibition also promoted IL-1β release, but it did not consistently affect release of the noninflammasome cytokines TNF and IL-6 (Supplemental Fig. 3E). Pyrin inflammasome activation was promoted in response to mTOR inhibition even upon stimulation with the purified toxin, TcdB (Supplemental Fig. 3F). Overall, these data establish an inverse correlation between the mTOR and pyrin inflammasome activation, demonstrating that negative regulation of mTOR signaling by RIPK3 can regulate Mefv expression and pyrin inflammasome activation.

To test the in vivo relevance of these findings, we injected supernatant derived from C. difficile cultures (toxin-negative C. difficile and toxin-positive C. difficile) into the mouse peritoneum and assessed inflammasome activation. Peritonitis as a model allows us to specifically address the toxin-induced innate inflammasome response independent of alterations observed in the adaptive immune system (42–44). Similar to our in vitro data, C. difficile toxin promoted pyrin- and caspase-1–mediated IL-18 release, suggesting that the toxin activates the pyrin inflammasome and promotes systemic IL-18 release in the peritonitis model (Fig. 5A). C. difficile toxin stimuli (irrespective of toxin status) also induced production of the inflammatory mediators TNF and KC that, as expected, did not rely on inflammasome activation (Fig. 5A). Genetic deletion of TNF or RIPK3 reduced IL-18 release in response to the toxin stimuli, highlighting the significance of this signaling axis in pyrin inflammasome activation (Fig. 5B). These data demonstrate that RIPK3 plays an important role in promoting activation of the pyrin inflammasome.

Our observations demonstrate a critical role for RIPK3 in Mefv expression and pyrin inflammasome activation. RIPK3 promotes Mefv expression downstream of TNF production. Neither RIPK3- and MLKL-mediated necroptosis nor caspase-3– or -7–mediated apoptosis was involved in the regulation of the pyrin inflammasome. RIPK3 was instead involved in transcriptional regulation of Mefv via negative control of mTOR signaling, and inhibition of mTOR activity is sufficient to upregulate Mefv expression and pyrin inflammasome activation. The regulation of Mefv expression by the RIPK3–mTOR axis could involve epigenetic control, mRNA stability, or microRNA-mediated control; these are interesting topics for further research. Overall, these data identify a novel regulatory mechanism for pyrin inflammasome activation.

Pyrin inflammasome activation is modulated through Rho-mediated pyrin phosphorylation and binding to 14-3-3 proteins (16, 33, 37, 38). Levels of 14-3-3 expression, Rho-mediated pyrin phosphorylation, and toxin-induced dephosphorylation were similar between WT and Ripk3^−/− cells, suggesting that these factors do not contribute to the differential pyrin inflammasome activation observed in RIPK3-deficient cells. Mefv expression, both basal and stimulation induced, was modulated by RIPK3 and inversely correlated with the status of mTOR activity. This observation is similar to the increased pyrin expression in cells lacking PI3K catalytic p110δ subunit (40). Our findings demonstrate that RIPK3 promotes Mefv expression and inflammasome activation in response to C. difficile toxin.

RIPK3 promotes inflammation independent of MLKL-mediated necroptosis in various settings (45–48). We similarly observed that the role of RIPK3 in the regulation of the pyrin inflammasome was independent of its role in necroptosis or inflammatory signaling and instead relied on regulation of the mTOR pathway. RIPK3-mediated necroptosis is mediated by mTOR–AKT activation (49), suggesting that RIPK3 promotes mTOR activation in response to necroptotic stimuli. However, we observed increased mTOR activation in Ripk3^−/− BMDMs both at baseline and in response to C. difficile toxin.

In addition to the impact of mTOR on pyroptosis and the pyrin inflammasome shown in this study, mTOR can also influence other forms of cell death, including apoptosis (50, 51) and necroptosis (49, 52). Additionally, mTORC1 activation promotes NLRP3 inflammasome activation (53), and rapamycin limits NLRP3 inflammasome activation in response to alum and viral infections (54). This is in contrast to the role of mTOR signaling in pyrin inflammasome activation. This contrasting regulatory function of mTOR on the two inflammasomes might provide the host with a competitive advantage in cases in which bacteria or other pathogenic agents actively subvert the mTOR pathway.

It was recently shown that intestinal cell death early during C. difficile infection is protective during self-limiting pseudomembranous colitis and proceeds independent of the pyrin inflammasome (55). However, inflammasome activation and inflammasome-dependent IL-1 signaling have been shown to be deleterious in toxin-mediated intestinal pathology (18). Thus, pyrin activation and downstream IL-1 signaling could, under certain conditions, impact C. difficile–induced pathology.

Overall, our data identified RIPK3 as a critical regulator of Mefv expression and the pyrin inflammasome via modulation of the mTOR pathway. The regulation of the pyrin inflammasome by molecules involved in cellular functions of cell death and metabolism highlights the significance of the interplay between these pathways during inflammation. Dysregulation of the pyrin inflammasome is associated with autoinflammatory disorders and impaired host defense following infectious or sterile insults. Defining the molecular mechanisms involved in pyrin inflammasome activation is therefore critical to understanding the etiology of, and identifying therapeutic targets for, the treatment of associated inflammatory diseases.

We thank Dr. Nigel Minton (University of Nottingham) and Dr. William A. Petri, Jr., (University of Virginia) for providing the C. difficile strains. We also thank Amanda Burton for technical assistance and Drs. Teneema Kuriakose, Parimal Samir, and Rebecca Tweedell for valuable input in editing the manuscript.

The authors have no financial conflicts of interest.