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Commun Biol
Commun Biol
Communications Biology
2399-3642
Nature Publishing Group UK London

39289441
6882
10.1038/s42003-024-06882-3
Article
Activation of the NLRP1B inflammasome by caspase-8
https://orcid.org/0000-0002-2215-5623
Meade Justin J. 1
Stuart Sarah 1
Neiman-Zenevich Jana 1
Krustev Christian 1
http://orcid.org/0000-0001-9208-8879
Girardin Stephen E. 12
http://orcid.org/0000-0003-3081-9247
Mogridge Jeremy jeremy.mogridge@utoronto.ca

1
1 https://ror.org/03dbr7087 grid.17063.33 0000 0001 2157 2938 Department of Laboratory Medicine and Pathobiology, University of Toronto, Toronto, On M5S 1A8 Canada
2 https://ror.org/03dbr7087 grid.17063.33 0000 0001 2157 2938 Department of Immunology, University of Toronto, Toronto, ON M5S 1A8 Canada
17 9 2024
17 9 2024
2024
7 116423 5 2023
12 9 2024
© The Author(s) 2024
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.
Cleavage of the innate immune receptor NLRP1B by various microbial proteases causes the proteasomal degradation of its N-terminal fragment and the subsequent release of a C-terminal fragment that forms an inflammasome. We reported previously that metabolic stress caused by intracellular bacteria triggers NLRP1B activation, but the mechanism by which this occurs was not elucidated. Here we demonstrate that TLR4 signaling in metabolically stressed macrophages promotes the formation of a TRIF/RIPK1/caspase-8 complex. Caspase-8 activity, induced downstream of this TLR4 pathway or through a distinct TNF receptor pathway, causes cleavage and activation of NLRP1B, which facilitates the maturation of both pro-caspase-1 and pro-caspase-8. Thus, our findings indicate that caspase-8 and NLRP1B generate a positive feedback loop that amplifies cell death processes and promotes a pro-inflammatory response through caspase-1. The ability of NLRP1B to detect caspase-8 activity suggests that this pattern recognition receptor may play a role in the defense against a variety of pathogens that induce apoptosis.

Metabolic stress is shown to impair IKKβ function, which leads to the activation of pro-caspase-8 downstream of TLR4 engagement. Caspase-8 cleaves and activates the NLRP1B inflammasome, thereby enhancing a cell death signal.

Subject terms

Cell death and immune response
Immune cell death
NOD-like receptors
https://doi.org/10.13039/501100000024 Gouvernement du Canada | Canadian Institutes of Health Research (Instituts de Recherche en Santé du Canada) issue-copyright-statement© Springer Nature Limited 2024
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pmcIntroduction

Detection of microbes by innate immune cells initiates processes that facilitate the elimination of pathogens and that generate signals to shape the immune response. Pattern recognition receptors are key drivers of pathogen detection and within this class of receptors are those that form inflammasomes1. Inflammasomes are protein complexes that consist of oligomerized pattern recognition receptors that bind and activate pro-caspase-1 and in some cases pro-caspase-82–5. Caspase-1 processes pro-IL-1β and pro-IL-18 as well as an executioner of cell death, Gasdermin D (GSDMD). Processed GSDMD forms pores in the plasma membrane that facilitate the release of IL-1β and IL-18 and that cause a type of pro-inflammatory cell death termed pyroptosis6. Pyroptotic cell death and the release of pro-inflammatory mediators are important contributors to mounting an inflammatory response against invading pathogens.

Human NLRP1 and murine NLRP1B belong to a family of inflammasome-forming nucleotide oligomerization domain-like receptors (NLRs)7. Unlike other members of this family, NLRP1 and NLRP1B contain a FIIND (function-to-find-domain) that auto-processes into a ZU5 domain and an UPA domain8–10. This auto-processing event divides the protein into a non-covalently associated N-terminal fragment (consisting of a PYD domain [in NLRP1, but not in NLRP1B], NACHT domain, LRR domain, and ZU5 domain) and a C-terminal fragment (consisting of an UPA domain and a CARD). The auto-processing of FIIND is not efficient, however, as both unprocessed and auto-processed NLRP1/NLRP1B are found in cells8–10. Prior to activation, the inhibited complex consists of full-length NLRP1/NLRP1B bound to an UPA-CARD fragment and to dipeptidyl proteases 8 and 9 (DPP8/9)11,12.

Activation of NLRP1 and NLRP1B occurs when the N-terminal fragment is ubiquitinated and then destroyed by the proteasome to release UPA-CARD fragments that self-associate and recruit ASC (human) and pro-caspase-113,14. Ubiquitination can occur following cleavage by anthrax lethal toxin13 or viral proteases15–17, phosphorylation by ZAKα18 or p3819, inhibition of DPP8/9 by Val-boroPro20, reductive stress21, binding of dsRNA22, or after recognition of NLRP1B by a bacterial ubiquitin ligase13. In contrast, Kaposi’s sarcoma associated herpesvirus ORF4523 activates NLRP1 by disrupting the auto-inhibited complex without causing its ubiquitination.

We previously reported that metabolic stress activates the NLRP1B inflammasome in LPS-stimulated macrophages24. This inflammasome is activated by Shigella flexneri, an invasive Gram-negative pathogen that induces metabolic stress in the host cell. It is also activated by the combination of LPS and the glycolysis inhibitor 2-deoxyglucose (2DG), which reduces cellular ATP levels24.

Here we delineate the pathway leading from the treatment of cells with LPS/2DG to the assembly of the NLRP1B inflammasome. Using CRISPR-Cas9 gene editing of RAW264.7 cells, we found that TLR4 was required for NLRP1B-dependent processing of pro-caspase-1, but this was not a result of de novo transcription because processing was not blocked by Actinomycin D. We found that signaling through TLR4 and its adaptor TRIF induced maturation of pro-caspase-8 when cells were treated with 2DG. 2DG prevented the LPS-dependent autoactivation of IKKβ, which allowed RIPK1 to recruit pro-caspase-8 to the TLR4/TRIF complex where it was matured. Caspase-8 caused the cleavage of NLRP1B between the LRR and FIIND domains to generate a ZU5-UPA-CARD fragment and induced NLRP1B-dependent processing of both pro-caspase-1 and pro-caspase-8. Thus, our work demonstrates that metabolic stress redirects signals downstream of TLR4 from a transcriptional program to a cell death program mediated by caspase-8. The activation of pro-caspase-8 by LPS/2DG is likely the only contributing factor caused by this treatment that promotes NLRP1B inflammasome formation because NLRP1B is also activated by treating cells with TNF and a TAK1 inhibitor, which initiates a mechanistically distinct process of activating pro-caspase-8.

Results

LPS and 2DG cause caspase-8-dependent, NLRP1B-independent, cell death

In our previous study, we found that treating RAW264.7 macrophages with a combination of LPS and 2DG led to NLRP1B-dependent processing of pro-caspase-124. To determine whether this treatment was sufficient to compromise membrane integrity, we monitored the cellular uptake of the membrane-impermeant dye propidium iodide (PI). Cells treated with either LPS or 2DG alone did not exhibit permeabilized membranes, whereas the combined treatment caused ~70% of cells to stain positively for PI (Fig. 1a). Surprisingly, NLRP1B was not required for the membrane damage (Fig. 1b). We confirmed that inflammasome activation was not necessary for cell death by using a Casp1-/- cell line generated by CRISPR-Cas9 gene editing (Fig. 1b, S1). Thus, although LPS/2DG activates NLRP1B24, cell death occurs through a different mechanism.Fig. 1 LPS/2DG-mediated cell death is dependent on caspase-8 and not on the NLRP1B inflammasome.

a, b RAW264.7 cells (wild-type, Nlrp1b–/–, or Casp1–/–) were treated for 3 h with LPS and 2DG. Cells were stained with propidium iodide (PI) and analyzed by flow cytometry. c RAW264.7 cells were treated with LPS/2DG, a pan-caspase inhibitor (Z-VAD) or a caspase-8 inhibitor (Z-IETD) for 3 h and were stained with PI and analyzed by flow cytometry. d RAW264.7 cells were treated with LPS/2DG, a pan-caspase inhibitor (Z-VAD) or a caspase-8 inhibitor (Z-IETD) as indicated for 2 h and cell lysates were immunoblotted for caspase-1, caspase-8, and β-actin. e RAW264.7 cells were treated with LPS/2DG for indicated times. Cell lysates were immunoblotted for caspase-1, caspase-8, and β-actin. f–i RAW264.7 cells (wild-type, Casp8–/– #1, Casp8–/– #2 pool, Nlrp1b–/–) were treated with LPS/2DG for 3 h (f) or 2 h (g–i). Cells were stained with PI and analyzed by flow cytometry or cell lysates were immunoblotted for caspase-1, caspase-8, and β-actin. j Schematic of NLRP1B activation by caspase-8. Made using Biorender.com. Blots are representative of three independent experiments. Error bars represent standard error of the mean of three independent experiments. Statistical significance was determined using a one-way (a, c) or (b, f) two-way ANOVA followed by a Sidak (a, b) or Tukey’s (c, f) post hoc test. *p < 0.05; **p < 0.01; ****p < 0.0001; NS nonsignificant.

We next sought to determine if caspase-8 is involved in LPS/2DG-mediated cell death because this extrinsic apoptotic initiator is activated by LPS under certain conditions25–27. Both a pan-caspase inhibitor and a caspase-8 inhibitor diminished PI uptake (Fig. 1c) and reduced the amount of processed caspase-8 p18 fragment in LPS/2DG-treated cells (Fig. 1d). Interestingly, NLRP1B-dependent processing of pro-caspase-1 was blocked by the caspase-8 inhibitor (Fig. 1d), suggesting that caspase-8 activity is required for triggering the NLRP1B inflammasome. Consistent with the notion that NLRP1B activation is downstream of caspase-8, processed caspase-8 was detected slightly earlier than the caspase-1 fragments (Fig. 1e). To confirm that caspase-8 is required for both cell death and NLRP1B activation, we generated a knock-out cell line from a single clone and found that LPS/2DG-treated Casp8–/– cells neither stained positively for PI, nor exhibited cleaved caspase-1 (Fig. 1f, g). We verified that caspase-8 is required for pro-caspase-1 processing using an unselected pool of Casp8–/– cells generated by a different gRNA (Fig. 1h).

Previously published work demonstrated that NLRP1B promotes pro-caspase-8 auto-processing28. Consistent with this study, we observed a reduced level of pro-caspase-8 processing in Nlrp1b–/– cells (Fig. 1i) and reduced PI uptake (Fig. 1b) indicating a positive feedback loop (Fig. 1j). Together these experiments reveal a pro-caspase-8 activation pathway induced by the combination of LPS and 2DG that signals the NLRP1B inflammasome.

TLR4 and TRIF are required for LPS/2DG-mediated cell death

We next wanted to address the role of LPS in this cell death pathway. LPS can influence cell death through transcriptionally dependent or independent mechanisms. To address whether LPS promotes the expression of genes involved in LPS/2DG-mediated cell death, we used the RNA polymerase inhibitor Actinomycin D. Actinomycin D did not prevent the LPS/2DG-mediated processing of pro-caspase-8 and pro-caspase-1 at a concentration that blocked transcription (Fig. 2a, b). That LPS is not required to induce gene expression for this cell death pathway is consistent with the rapid activation of pro-caspase-8 in LPS/2DG-treated cells (Fig. 1e).Fig. 2 TLR4 and TRIF are required for LPS/2DG-mediated cell death.

a RAW264.7 cells were treated with LPS and Actinomycin D (ActD) for 2 h and analyzed for Il1b transcript levels using quantitative real-time PCR. b RAW264.7 cells were treated with LPS, 2DG and ActD for 2 h. Cell lysates were immunoblotted for caspase-1, caspase-8 and β-actin. c, d RAW264.7 cells (wild-type, Tlr4–/–, Myd88–/–, Ticam1–/–) were treated with LPS/2DG for 3 h (c) or 2 h (d) (Ticam1 encodes TRIF). Cells were stained with propidium iodide (PI) and analyzed by flow cytometry; or cell lysates were immunoblotted for caspase-1, caspase-8, and β-actin. e RAW264.7 cells were treated with LPS, 2DG and dynasore for 2 h. Cell lysates were immunoblotted for caspase-1, caspase-8 and β-actin. f Bone marrow-derived macrophages (BMDMs) were treated with LPS, 2DG and dynasore for 2 h. Cell lysates were immunoblotted for caspase-1, caspase-8 and β-actin. Blots are representative of three independent experiments. Error bars represent standard error of the mean of three independent experiments. Statistical significance was determined by a one-way (a) or two-way ANOVA followed by a Tukey’s post hoc test. *p < 0.05; ****p < 0.0001; NS nonsignificant.

Signaling downstream of TLR4 is mediated through the adaptors MyD88 and TRIF29,30. MyD88 binds TLR4 indirectly through TIRAP and generates a signaling complex called the Myddosome. During endocytosis of TLR4, the Myddosome dissociates from TLR4 to allow binding of TLR4 to TRAM and TRIF. Using knock-out cell lines, we found that TLR4 and TRIF were required for pro-caspase-8 activation and cell death, whereas MyD88 was not (Fig. 2c,d, Fig. S2). Consistent with these observations, the endocytosis inhibitor dynasore reduced the processing of pro-caspase-8 and pro-caspase-1 in RAW264.7 cells and in bone marrow-derived macrophages (BMDMs) (Fig. 2e, f). Together, our data indicate that metabolically stressed macrophages induce pro-caspase-8 activation and cell death through a TLR4/TRIF-dependent but transcriptionally independent pathway.

Hypo-phosphorylated RIPK1 recruits pro-caspase-8 to TLR4/TRIF in LPS/2DG-treated cells

RIPK1 participates in cell death pathways and is regulated by post-translational modifications31. RIPK1 has a RHIM domain that binds the RHIM domains of TRIF and RIPK3. During TLR4-mediated cell death, RIPK1 binds TRIF and then forms large multimeric complexes with RIPK332. RIPK1 can also bind FADD, an adaptor protein that interacts with pro-caspase-8 and thereby facilitates the recruitment of pro-caspase-8 to the RIPK1/3 multimers to promote proximity-induced auto-processing and cell death33. Using co-immunoprecipitation assays, we detected RIPK1 complexed with pro-caspase-8 in untreated cells, and with pro-caspase-8 and caspase-8 p43 and p18 fragments in cells treated with the combination of LPS and 2DG (Fig. 3a). We also observed the TRIF-dependent recruitment of RIPK1 to TLR4 in LPS/2DG-treated cells (Fig. 3b, c). Together our experiments support a model in which 2DG promotes the recruitment of a cytosolic complex containing RIPK1 and pro-caspase-8 to an LPS-induced TLR4/TRIF complex29,34 where pro-caspase-8 undergoes proteolysis.Fig. 3 Hypo-phosphorylated RIPK recruits pro-caspase-8 to TLR4/TRIF in LPS/2DG-treated cells.

a RAW264.7 cells (wild-type or Casp8–/–) were treated for 1 h with LPS and 2DG. Endogenous RIPK1 was immunoprecipitated from cell lysates with an anti-RIPK1 antibody and then immunoblotted for pro-caspase-8 and RIPK1. Mouse IgG1 was used as a control. b, c RAW264.7 cells (wild-type or Ticam-/-) were treated for 30 min with LPS, 2DG and the TAK1 inhibitor 5z-7-Oxozeaenol (5z-7) as indicated. Endogenous RIPK1 was immunoprecipitated from cell lysates and immunoblotted for TLR4 and RIPK1. d RAW264.7 cells were treated for 30 min with LPS, 2DG and 5z-7 and cell lysates were immunoblotted for RIPK1. e RAW264.7 cells were treated for 30 min with LPS. Cell lysates were incubated in Alkaline Phosphatase (AP) for 30 min and then immunoblotted for RIPK1. f RAW264.7 cells were treated for 2 h with LPS, 2DG and 5z-7. Cell lysates were immunoblotted for pro-caspase-8. g C57BL/6 BMDMs were treated as in (d). h RAW264.7 and (i) C57BL/6 BMDMs were treated with LPS, 2DG, for 2 h and TNFα/SMAC mimetic/Z-VAD (T/S/Z) for 3 h and immunoblotted for pro-caspase-8, phospho-MLKL S345, MLKL, and β-actin. Blots are representative of three independent experiments.

We speculated that 2DG causes a reduction of RIPK1 phosphorylation to promote its binding to TRIF because phosphorylation of RIPK1 has been shown to prevent its involvement in cell death pathways35–38. It has also been shown that inhibition of TAK1, which directly phosphorylates RIPK135, causes activation of pro-caspase-8 in LPS-treated cells25–27. Consistent with this work, we observed that the TAK1 inhibitor 5z-7-oxozeaenol (5z-7) prevented the LPS-induced phosphorylation of RIPK1, detected as phosphatase-sensitive lower mobility species in immunoblots, and caused the processing of pro-caspase-8 (Fig. 3d–f). TAK1 inhibition also promoted the binding of RIPK1 to TLR4/TRIF (Fig. 3c). 2DG treatment recapitulated each of these events in RAW264.7 macrophages that were exposed to LPS (Fig. 3c–f) and prevented LPS-induced RIPK1 phosphorylation in BMDMs (Fig. 3g). Together, our data indicates that glycolytic inhibition interferes with kinase pathways that prevent cell death.

RIPK1 participates in cell death pathways through kinase-dependent and kinase-independent mechanisms39. To gain further insight into the role of RIPK1 in pro-caspase-8 activation, we treated cells with LPS/2DG and the RIPK1 kinase inhibitor Necrostatin 1S (Nec1s). We found that LPS/2DG-dependent pro-caspase-8 processing was not inhibited by Nec1s. Nec1s did, however, block necroptosis (induced by TNFα/SMAC mimetic/ZVAD) in RAW264.7 macrophages and BMDMs as determined by the inhibition of MLKL phosphorylation (Fig. 3h, i). Interestingly, 2DG also inhibited necroptosis (Fig. 3i), which suggests that glycolytic perturbation interferes with the necroptotic machinery. Together, our data support a model in which glycolytic inhibition promotes the assembly of a pro-caspase-8 activation complex involving TLR4, TRIF, and the scaffolding function of RIPK1.

2DG inhibits IKKβ kinase activity

Next, we wanted to identify the kinase(s) that 2DG inhibits to prevent the phosphorylation of RIPK1. TAK1, IKKβ40,41, p38/MK236,37,42 and TBK1/IKKε43,44 have each been shown to phosphorylate RIPK1. In LPS-stimulated RAW264.7 macrophages, the phosphorylation of RIPK1 was reduced by the TAK1 inhibitor 5z-7, the IKKβ inhibitor TPCA1, the p38/MK2 inhibitor SB203580, but not by the TBK1/IKKε inhibitor MRT67307 (Fig. 4a). Among the inhibitors that blocked RIPK1 phosphorylation, only 5z-7 and TPCA1 promoted caspase-8 activation in cells treated with LPS (Fig. 4b), suggesting that metabolic stress induced by 2DG may interfere with TAK1 and/or IKKβ activity. Consistent with these findings, we found that 2DG interfered with LPS-induced phosphorylation of the NF-κB p65 subunit (Fig. 4c), which is mediated through TAK1 and IKKβ45–48. 2DG did not, however, prevent the phosphorylation of the transcription factor IRF3 by the IKK-related kinases TBK1/IKKε (Fig. 4d), indicating that some kinases are more susceptible to inhibition by metabolic stress than others.Fig. 4 Inhibition of IKKβ by 2DG prevents LPS-dependent phosphorylation of RIPK1 to drive caspase-8 activation.

a RAW264.7 cells were stimulated for 30 min with LPS, 2DG, 5z7 (TAK1 inhibitor), TPCA1 (IKKβ inhibitor), SB203580 (p38i; p38 inhibitor), and MRT67307 (TBK1/IKKε inhibitor) as indicated. Cell lysates were immunoblotted for RIPK1 and tubulin. b RAW264.7 cells were treated for 2 h with LPS, SB203580, 5z7, TPCA1, and 2DG. Cell lysates were immunoblotted for caspase-8 and β-actin. c RAW264.7 cells were treated for 15 and 30 min with LPS, TPCA1, and 2DG. Cell lysates were immunoblotted for phospho-Serine-536-p65 NFκB, total p65-NFκB, and β-actin. d RAW264.7 cells were treated for 30 and 60 min with LPS, MRT67307 and 2DG. Cell lysates were immunoblotted for phospho-Serine 396-IRF3, total IRF3, and β-actin. e Schematic model of the two-step phosphorylation cascade that leads to IKKβ activation. RAW264.7 cells were stimulated for 15 min with LPS, 5z7, TPCA1, and 2DG. Cell lysates were immunoblotted for phospho-Serine 176/177 IKKα/β, phospho-Serine 176/177 and Serine 180/181 IKKα/β, total IKKβ, and β-actin. f HEK293T cells were transfected with the indicated plasmids for 22 h followed by a 2 h treatment with 2DG. Cell lysates were immunoblotted for phospho-Serine 536 p65 NFκB, total p65 NFκB, and β-actin. All blots are representative of three independent experiments. Asterisk (*) indicates non-specific band. Schematic made with Biorender.

We next monitored IKKβ phosphorylation to determine whether 2DG inhibits TAK1, IKKβ, or both of these enzymes. IKKβ is activated through a stepwise process in which TAK1 phosphorylates IKKβ at Serine 177, which then licenses IKKβ autophosphorylation at Serine 181 (Fig. 4e). Dual phosphorylated IKKβ-pS177/181 then phosphorylates its downstream substrates including NF-κB and RIPK1. We employed phospho-specific antibodies that recognize IKKβ-pS177 (TAK1-dependent phosphorylation) and IKKβ-pS177/181 (TAK1- and IKKβ-dependent phosphorylation) to assess the effects of metabolic stress on IKKβ activation. We observed both IKKβ-pS177 and IKKβ-pS177/181 in cells treated with LPS, but not in untreated cells (Fig. 4e). As expected, 5z-7 reduced the level of both IKKβ-pS177 and IKKβ-pS177/181, whereas TPCA1 only diminished the generation of IKKβ-pS177/181 (Fig. 4e). Similar to what was observed with TPCA1, 2DG reduced the level of IKKβ-pS177/181 and increased the level of IKKβ-pS177 (note that the antibody recognizes IKKβ-pS177 but not IKKβ-pS177/181, so preventing autophosphorylation increases the IKKβ-pS177 signal) indicating that metabolic stress interferes with IKKβ activity but not TAK1 activity. Next, we mutated Serine 177 and Serine 181 of IKKβ to glutamic acids which renders this protein constitutively active49,50. Expression of wild-type IKKβ and IKKβ-S177E/S181E in HEK293T cells resulted in the phosphorylation of p65 NF-κB, which was blocked by incubation with 2DG (Fig. 4f), indicating that metabolic stress prevents IKKβ activity irrespective of its phosphorylation at Serine 177 and 181. Together, these data support a model where metabolic stress negatively modulates the kinase activity of IKKβ, which is sufficient to prevent RIPK1 phosphorylation and drive pro-caspase-8 activation downstream of TLR4.

Shigella flexneri induces caspase-8-dependent activation of NLRP1B

We previously found that NLRP1B was activated in Shigella-infected RAW264.7 cells24, but it was unclear whether this activation resulted from metabolic stress or from another signal generated by the pathogen. We sought to determine, therefore, if Shigella triggers the inflammasome through the same caspase-8 pathway as LPS/2DG treatment does. We found that infection of cells with Shigella (wild-type strain M90T) caused processing of pro-caspase-8 and pro-caspase-1 (Fig. 5a). In contrast, the Shigella strain BS176 that lacks the ability to invade cells did not induce processing of these pro-caspases and did not reduce the level of cytosolic ATP (Fig. 5b, c). Importantly, the NLRP1B-dependent maturation of pro-caspase-1 was reduced in Tlr4–/– and Casp8–/– cell lines (Fig. 5d). We conclude, therefore, that Shigella infection and LPS/2DG treatment induce a common inflammasome activation pathway.Fig. 5 Shigella flexneri induces caspase-8-dependent inflammasome activation.

a RAW264.7 cells were incubated with Shigella flexneri at indicated multiplicities of infection (MOI) for 1 h. Cell lysates were immunoblotted for caspase-1, pro-caspase-8, cleaved caspase-8, and β-actin. b, c Shigella flexneri (wild-type, M90T; and mutant, BS176) were incubated with RAW264.7 cells at an MOI of 5 for 1 h. Intracellular ATP concentrations were measured and cell lysates were immunoblotted for caspase-1, pro-caspase-8, cleaved caspase-8 and β-actin. d RAW264.7 cells (wild-type, Nlrp1b–/–, Tlr4–/–, Casp8–/–) were incubated with Shigella flexneri at an MOI of 5 for 1 h. Cell lysates were immunoblotted for caspase-1, pro-caspase-8, cleaved caspase-8, and β-actin. e, f RAW264.7 cells were incubated with lethal toxin (LT) for 3 h, Shigella flexneri for 1 h, LPS/2DG, and MG132 for 2 h. Cell lysates were immunoblotted for caspase-1, GSDMD, and β-actin. Blots are representative of at least three independent experiments. Error bars represent standard error of the mean of three independent experiments. Statistical significance was determined using a one-way ANOVA followed by a Tukey’s post hoc test. ***p < 0.001; NS nonsignificant.

The Shigella effector IpaH7.8 has been shown to activate NLRP1B by ubiquitinating the N-terminal fragment of the protein. This fragment undergoes proteasome-mediated degradation, which allows the C-terminal fragment to assemble into an inflammasome complex—a process known as functional degradation13. That NLRP1B activation by Shigella flexneri strain M90T was dependent on TLR4 and caspase-8, whereas activation by strain 2457 T was dependent on the effector IpaH7.813 suggested different processes, so we sought to determine whether functional degradation occurred during M90T infection. We found that inhibition of the proteasome with MG-132 slightly increased the processing of pro-caspase-1 in Shigella-infected and LPS/2DG-treated cells (Fig. 5e, f). As expected, MG-132 decreased the amount of the caspase-1 p10 and GSDMD p30 generated in anthrax lethal toxin-treated cells (Fig. 5f). Consistent with another study51, inhibition of the proteasome slightly increased the amount of the 43 kDa caspase-8 fragment that correlated with the slight induction of the ~14 kDa caspase-1 fragment (Fig. 5f). Our results indicate that the M90T strain of Shigella flexneri (and LPS/2DG treatment) can induce NLRP1B through a mechanism dependent on caspase-8 but likely independent of the proteasome.

Caspase-3, but not membrane damage, facilitates inflammasome activation

Loss of membrane integrity has been associated with the activation of the NLRP3 inflammasome52,53, so we next sought to determine how membrane breach was occurring in LPS/2DG-treated cells and whether permeabilization of the plasma membrane triggers NLRP1B. Using quantitative real-time PCR, we found that RAW264.7 cells express Gsdmd and Gsdme, but none of the other Gasdermin family members (Table S1). GSDMD and GSDME were cleaved in both RAW264.7 macrophages and BMDMs treated with LPS/2DG: GSDMD was cleaved primarily into inactive 20 kDa and 43 kDa fragments, whereas GSDME was cleaved into an active ~36 kDa fragment (Fig. 6a, c). Using a Gsdmd-/-Gsdme-/- double knock-out cell line, we determined that these proteins were not, however, required for uptake of PI or for pro-caspase-1 processing (Fig. 6d, e).Fig. 6 Caspase-3, but not membrane damage, facilitates inflammasome activation.

a, b RAW264.7 cells and (c) C57BL/6 BMDMs were treated with LPS and 2DG for 2 h. Cell lysates were immunoblotted for GSDMD, GSDME, Pannexin-1 and β-actin. RAW264.7 cells (wild-type, Gsdmd–/–Gsdme–/–) were treated with LPS/2DG and trovafloxacin for 2 h (d) or 3 h (e). Cells were stained with PI and analyzed by flow cytometry, or cell lysates were immunoblotted for caspase-1 and β-actin. f, g RAW264.7 cells (wild-type, Casp3–/–) were treated with LPS/2DG for 2 h (f) or 3 h (g). Cells were stained with PI and analyzed by flow cytometry, or cell lysates were immunoblotted for caspase-1, GSDMD, GSDME and β-actin. Blots are representative of three independent experiments. Error bars represent standard error of the mean of three independent experiments. Statistical significance was determine using a two-way ANOVA followed by a Tukey’s (e) or Sidak (f) post hoc test. ****p < 0.0001; NS nonsignificant.

Previous work has shown that caspase-3 and caspase-7, executioner caspases downstream of caspase-8, cleave pannexin-1 to activate its channel activity54. Because we observed pannexin-1 cleavage in LPS/2DG-treated cells (Fig. 6b, c), we used the pannexin inhibitor trovafloxacin to determine whether pannexin channels contribute to membrane permeability55. We found that trovafloxacin diminished PI uptake in wild-type and Gsdmd-/-Gsdme-/- cells but did not reduce the maturation of pro-caspase-1 (Fig. 6d, e). These results suggest that pannexin-1 channels contribute to permeabilization of the plasma membrane, but that this may not be involved in the activation of NLRP1B.

Next, we assessed whether caspase-3 was involved in triggering the inflammasome. Treating Casp3-/- cells with LPS/2DG led to PI uptake, but not to processing of pro-caspase-1 (Figs. 5f, g, Fig. S3). We note that NLRP1B was functional in these cells as demonstrated by its activation by Val-boroPro (Fig. S4). Thus, these results indicate that in cells treated with LPS/2DG, caspase-3 facilitates the activation of NLRP1B and that permeabilization of the plasma membrane is not sufficient to activate NLRP1B.

Caspase-8 induces the cleavage of NLRP1B

Although we have shown that caspase-8 is required to activate NLRP1B in LPS/2DG-treated cells, it is possible that the treatment generates an additional signal that is required for inflammasome formation. To address this possibility, we sought to determine if different treatments that activate pro-caspase-8 also lead to NLRP1B inflammasome activation. TAK1 inhibition activates pro-caspase-8 downstream of either TLR4 or tumor necrosis factor receptor 1 (TNFR1) signaling25,26 (Fig. 7a). We found that treatment with LPS/5z-7 or TNFα/5z-7 led to the processing of pro-caspase-1 in wild-type cells, but not in Casp8-/- knock-out cells (Fig. 7a). When cells were treated with concentrations of LPS/5z-7, TNFα/5z-7, and LPS/2DG that led to low levels of caspase-8 p43 and p18, the processing of pro-caspase-1 was completely dependent on NLRP1B (Fig. 7b). Processing of pro-caspase-8 was also decreased in the Nlrp1b-/- cells, indicating that a NLRP1B-caspase-8 positive feedback loop is activated by each of these conditions (Fig. 7b). Cells treated with fivefold more 5z-7 in combination with LPS or TNFα exhibited higher levels of mature caspase-8 and caspase-1, but the processing of these caspases was only slightly reduced in Nlrp1b–/– knock-out cells (Fig. 7c). Therefore, these data indicate that a lower level of pro-caspase-8 activation causes NLRP1B-dependent processing of pro-caspase-1, whereas a higher level of pro-caspase-8 activation results in additional NLRP1B-independent processing of pro-caspase-1 (Fig. 7d, S5). Furthermore, these results suggest that the only functionally significant consequence of LPS/2DG treatment that causes NLRP1B activation is the generation of mature caspase-8.Fig. 7 Caspase-8 induces proteasome-independent cleavage of NLRP1B.

a–c RAW264.7 cells (wild-type, Casp8–/–, Nlrp1b–/–) were treated as indicated for 2 h. Cell lysates were immunoblotted for pro-caspase-8, pro-caspase-1 and β-actin. d Schematic of caspase-1 activation by weak and strong stimuli of caspase-8. Made using Biorender.com. e–g RAW264.7 cells (wild-type, Casp3–/–, Casp8–/– #1, Casp8–/– #2 pool, Nlrp1b–/–) were treated as indicated for 2 h. Cell lysates were immunoblotted for pro-caspase-8, NLRP1B, cleaved caspase-3, and β-actin. h, i BALB/c BMDMs were treated as indicated for 2 h. Cell lysates were immunoblotted for pro-caspase-8, NLRP1B, and β-actin. Blots are representative of three independent experiments except (h) where n = 2. Asterisk (*) indicates NLRP1B cleavage products.

As NLRP1B is cleaved and activated by microbial proteases13,17, we speculated that NLRP1B might be cleaved by caspase-8 or a downstream protease. Using an antibody generated against the CARD domain of NLRP1B, we observed the auto-processed UPA-CARD fragment and unprocessed NLRP1B in untreated cells. Interestingly, the combination of LPS and 2DG promoted the appearance of bands of ~37 kDa-45 kDa in size in wild-type RAW264.7 cells that were absent in Casp8-/- knock-out cells (Fig. 7e, f). Proteasomal inhibition did not block LPS/2DG-induced cleavage of NLRP1B; in fact, MG132 increased the level of caspase-8 and cleaved NLRP1B (Fig. 7f). The increased NLRP1B cleavage in the presence of MG-132 correlates with the MG-132-induced increase in NLRP1B-dependent pro-caspase-1 processing in cells treated with LPS/2DG or infected with Shigella (Fig. 5e, f). Cells treated with LPS/5z-7 also led to the processing of NLRP1B (Fig. 7g). Together, these data demonstrate that activation of caspase-8 leads to the cleavage of NLRP1B.

Caspase-3 facilitates the activation of NLRP1B (Fig. 6g), so we sought to determine if caspase-3 does so by cleaving NLRP1B. Although LPS/2DG did not induce NLRP1B cleavage in Casp3–/– knock-out cells, the processing of pro-caspase-8 was also impaired in these cells (Fig. 7g), likely because of a caspase-8/caspase-3 positive feedback loop. To overcome the need for the feedback loop, we increased the concentration of 5z-7 to induce caspase-3-independent processing of pro-caspase-8 and we observed cleavage of NLRP1B in these conditions (Fig. 7g). Thus, caspase-3 acts upstream of caspase-8 and may play a substantial role in inflammasome activation only in conditions that weakly activate pro-caspase-8.

We next asked if the cleavage of NLRP1B occurs in primary macrophages. There are five alleles of NLRP1B, so we generated BMDMs from BALB/c mice as these mice harbor allele 1 of NLRP1B, which is the allele found in RAW264.7 macrophages. BMDMs treated with LPS/2DG and LPS/5z-7 produced NLRP1B cleavage products that were diminished when the cells were treated with the pan-caspase-inhibitor Z-VAD (Fig. 7h, i).

Caspase-8-mediated cleavage of NLRP1B occurs between the LRR and FIIND domains

To observe NLRP1B cleavage in a reconstituted system, we expressed NLRP1B with either wild-type pro-caspase-8 or the catalytically inactive pro-caspase-8-C362A mutant in HEK293T cells. Expression of wild-type but not mutant pro-caspase-8 led to its auto-activation, as demonstrated by the appearance of the p18 cleavage product (Fig. 8a). Expression of NLRP1B with wild-type, but not mutant, pro-caspase-8 resulted in the cleavage of NLRP1B into products of a similar size as those observed in RAW264.7 macrophages and in BALB/c BMDMs (Fig. 8a).Fig. 8 Caspase-8-mediated cleavage of NLRP1B signals inflammasome formation.

a HEK293T cells were transfected with the indicated plasmids for 24 h. Cell lysates were immunoblotted for pro-caspase-8, NLRP1B, and beta-actin. (b, Top panel) Schematic of NLRP1B WT and the NLRP1BΔ749-870 mutant lacking the linker between the LRR and FIIND domain. (b, Bottom panel) HEK293T cells were transfected with the indicated plasmids for 24 h. Cell lysates were immunoblotted for pro-caspase-8, NLRP1B, and β-actin. (c, Top panel) Schematic of murine NLRP1B allele 1 with the amino acid sequence between residues 785 and 849 with red indicating residues mutated to alanine. (c, Bottom panel) d HEK293T cells were transfected with the indicated plasmids for 24 h. Cell lysates were immunoblotted for pro-caspase-8, NLRP1B and β-actin. e Activated caspase-8 cleaves NLRP1B to form an inflammasome that activates caspase-8 and caspase-1. Asterisk (*) indicates cleavage products of NLRP1B. All immunoblots are representative of three independent experiments. Schematics made using Biorender.

The size of the NLRP1B cleavage products suggested that proteolysis occurs in the linker region between the LRR and FIIND domains (Fig. 8b). To test this idea, we used a construct that lacks the linker region (NLRP1BΔ749-870)56. Co-expression of this mutant with pro-caspase-8 did not generate any fragments, suggesting that this linker is the segment that is cleaved (Fig. 8b). NLRP1BΔ749-870 did not generate an UPA-CARD fragment (Fig. 8b), suggesting that deletion of this linker region also prevented FIIND auto-processing. The reason why NLRP1BΔ749-870 does not undergo auto-processing is unclear.

Caspases recognize tetra-peptide motifs and cleave adjacent to a mandatory aspartic acid in position P157. Caspase-8 recognizes the motif XEXD (where “X” represents any amino acid residue)57. We found three sequences (785TEED788, 821DEED824, and 846TEED849) between the LRR and FIIND domains of NLRP1B that conform to the caspase-8 recognition motif and that would produce fragments in the expected size range when cleaved (Fig. 8c). We individually mutated these aspartic acid residues to alanine and determined whether the mutants were cleaved in HEK293T cells that co-expressed caspase-8 (Fig. 8c). We found that NLRP1B-D788A and NLRP1B-D849A were not cleaved into the larger and smaller cleavage products, respectively, whereas NLRP1B-D824A was cleaved into the wild-type products (Fig. 8c). Furthermore, the double mutant NLRP1B-D788A/D849A was not cleaved by caspase-8 (Fig. 8d). These data indicate, therefore, that caspase-8-mediated cleavage of NLRP1B occurs between the LRR and FIIND domains at two sites, D788 and D849, to produce 37 kDa and 45 kDa C-terminal fragments.

Discussion

Cells that detect infections can initiate protective processes or commit programmed cell death. Immunologically silent cell death, such as apoptosis, limits self-inflicted tissue damage whereas pro-inflammatory cell death may be better able to initiate a response that clears the infection. Elucidating how the signals that lead to these outcomes are detected, integrated and used by interconnected processes is key to our understanding of the immune system. Our work here has focused on how the combination of two signals generated by Gram-negative intracellular pathogens—LPS and metabolic stress—invoke a pro-inflammatory cell death response (Figs. S6, S7).

We found that metabolic stress regulates TLR4 signaling to promote the activation of pro-caspase-8. While TLR4 has long been known to trigger gene expression and cell death programs, the ways in which cells decide between these two programs is incompletely understood. RIPK1 is central to this decision as phosphorylated RIPK1 potentiates NF-κB function, whereas unphosphorylated RIPK1 assembles a pro-caspase-8 activation complex58. RIPK1 phosphorylation is inhibited by YopJ, a Yersinia pestis effector that acetylates an upstream kinase of RIPK1, TAK125,26,35,59,60. The authors of these studies suggest that the evolved response to having a virulence factor shut down a signaling pathway involved in defence is to commit to cell death. Similarly, our finding suggests that a metabolically impaired cell, which may be limited in its ability to initiate defensive measures such as cytokine production, initiates cell death as the ultimate response to infection.

We furthermore discovered that metabolic stress prevents RIPK1 phosphorylation by inhibiting IKKβ. A previous study that measured IKKβ enzyme kinetics demonstrated that the Ki for ADP, a product of the kinase reaction, is 0.77 μM61. This unusually low Ki suggested to the authors that ADP may play a physiological role in the regulation of IKKβ activity. This is consistent with our findings – metabolic stress caused by 2DG or bacterial invasion decreases cellular ATP (Fig. 5b)24, which would increase the ADP:ATP ratio and thereby reduce IKKβ activity. This low Ki value could also explain why metabolic stress inhibited IKKβ activity (Fig. 4e, f), but not the activities of TBK1/IKKε (Fig. 4d) or TAK1 (Fig. 4e). It is an intriguing idea that IKKβ might function as a sensor of metabolic integrity that regulates the response to infection.

We speculated previously that NLRP1B is a sensor of metabolic stress62, but our data now support the notion that it is caspase-8 and not specifically metabolic stress that activates NLRP1B. This conclusion is supported by the observations that LPS/2DG does not activate NLRP1B in Casp8-/- cells and that NLRP1B is activated in cells that are treated with either TNFα/5z-7 or LPS/5z-7, treatments that activate pro-caspase-8 without inducing metabolic stress. In light of work that demonstrates that NLRP1B is a target of microbial proteases13,15–17, it did not come as a surprise to find that NLRP1B is cleaved in cells that have active caspase-8 (Figs. 7e–i, 8a).

We mapped the cleavage of NLRP1B allele 1 to two sites that conform to the caspase-8 recognition motif. NLRP1B allele 1 has a repeated sequence as a result of an exon duplication, whereas alleles 2–5 have only one of the repeats and only one predicted caspase-8 recognition site (Fig. S8). Of the two human FIIND-containing proteins, NLRP1 does not have a consensus caspase-8 recognition site in this region, but CARD8 has a site that aligns near the second motif found in NLRP1B allele 1 (Fig. S8). It remains to be determined whether CARD8 can be activated by caspase-8. We have not conclusively determined that caspase-8 directly cleaves NLRP1B, although we have shown that NLRP1B cleavage occurs in LPS/5z7-treated Casp3-/- RAW264.7 macrophages and in HEK293T cells in which caspase-8 and NLRP1B are co-expressed.

Cleavage occurs in a linker region between the LRR and FIIND domains and although the resulting C-terminal fragments might be subjected to proteasomal degradation, which would fit the functional degradation model, we found that a proteasome inhibitor did not prevent caspase-8-mediated NLRP1B activation (Fig. 5f). It is possible, therefore, that cleavage of the linker may disrupt the NLRP1B auto-inhibited complex to release active FIIND-CARD and UPA-CARD fragments (Fig. 8e). We also note that the antibody used in this study binds the NLRP1B CARD domain, which would allow it to detect caspase-8-mediated cleavage of unprocessed NLRP1B, but not processed NLRP1B – whether caspase-8 cleaves processed NLRP1B is not known.

A shortcoming of this study is that we have not proven that it is the cleavage of the linker domain that activates NLRP1B. We were not able to reconstitute Nlrp1b-/- RAW264.7 cells with wild-type and caspase-8 resistant NLRP1B to test this because neither transfection nor transduction of the genes led to sufficient protein levels. We were also not able to test whether the linker mutant was resistant to caspase-8-mediated activation in HEK293T cells because overexpression of pro-caspase-8 in this system leads to NLRP1B-independent processing of pro-caspase-1.

While previous studies have described NLRP1B activators that initiate cell death, our work describes a mechanism in which NLRP1B amplifies cell death. Caspase-8 activates NLRP1B, which can serve as a scaffold to activate pro-caspase-828 and pro-caspase-1. A weak apoptotic signal can thereby be amplified to initiate pro-inflammatory pyroptotic death. Strong pro-caspase-8 activation, such as we observed in cells treated with LPS/5z-7, does not require NLRP1B for amplification and does not seem to require an inflammasome to activate pro-caspase-1 (Fig. 7c, d). Thus, apoptotic cells that express NLRP1B appear to be predisposed to pyroptotic cell death, which might be beneficial for pathogen clearance.

Methods

Antibodies

The following primary antibodies were used for Western blotting, immunoprecipitation, and immunofluorescence: mouse monoclonal anti-β-actin (Millipore-Sigma, #A5441; RRID:AB_476744), rabbit polyclonal anti-Caspase-1 p10 (Santa Cruz, #sc-514; RRID:AB_2068895), rabbit polyclonal anti-Caspase-3 (Cell Signaling Technology, #9662; RRID:AB_331439), rabbit polyclonal anti-Caspase-8 (Cell Signaling Technology, #4927; RRID:AB_2068301), rabbit monoclonal anti-Cleaved Caspase-8 (Asp387) (Cell Signaling Technology, #8592; RRID:AB_10891784), rabbit monoclonal anti-DFNA5/GSDME, N-terminal (Abcam, #ab215191; RRID:AB_2737000), rabbit monoclonal anti-GSDMD (Abcam, #ab209845; RRID:AB_2783550), mouse monoclonal anti-human HA, clone HA-7 (Millipore-Sigma, # H9658; RRID:AB_260092), mouse monoclonal anti-RIPK1 (Abcam, #ab72139; RRID:AB_2178115); rabbit monoclonal anti-RIPK1 (Cell Signaling Technology, #3493; RRID:AB_2305314); rabbit monoclonal anti-Toll-like Receptor 4 (Cell Signaling Technology, #14358; RRID:AB_2798460), mouse monoclonal anti-α-tubulin (Millipore-Sigma, #T9026; RRID:AB_477593), mouse monoclonal anti-NLRP1B (Adipogen, #AG-20B-0084-C100), rabbit monoclonal anti-MyD88 (Cell Signaling Technology, #4283; RRID:AB_10547882), rabbit monoclonal anti-Phospho-MLKL (Cell Signaling Technology, #37333; RRID:AB_2799112), rabbit monoclonal anti-MLKL (Cell Signaling Technology, #37705; RRID:AB_2799118) rabbit polyclonal anti-Phospho-NFκB-p65-Ser536 (Cell Signaling Technology, #3031), rabbit monoclonal anti-NFκB-p65 (Cell Signaling Technology, #8242), rabbit monoclonal anti-Phospho-IKKα/β-Ser176/177 (Cell Signaling Technology, #2078), rabbit monoclonal anti-Phospho-IKKα/β-Ser176/180 (Cell Signaling Technology, #2697), rabbit monoclonal anti-IKKα/β (Cell Signaling Technology, #8943) and, rabbit monoclonal anti-Phospho-IRF3-Ser396 (Cell Signaling Technology, #4947).

Secondary antibodies used for Western blotting were goat anti-mouse IgG (H + L) secondary antibody, HRP (Thermo Fisher, #31430; RRID:AB_228307) or goat anti-rabbit IgG (H + L) secondary antibody, HRP (Thermo Fisher, #31460; RRID:AB_228341). Secondary antibodies for immunofluorescence were goat anti-mouse IgG (H + L) highly cross-adsorbed secondary antibody, Alexa Fluor Plus 555 (Thermo Fisher, #A32727; RRID:AB_2633276) and goat anti-rabbit IgG (H + L) cross-adsorbed secondary antibody, Alexa Fluor 405 (Thermo Fisher, # A-31556; RRID:AB_221605).

For fluorescence-activated cell sorting, rat anti-mouse CD16/CD32 (Mouse BD Fc Block™) (BD Biosciences, #553141; RRID:AB_394656) and TLR4/MD-2 Complex Monoclonal Antibody (MTS510), APC, eBioscience™ (Thermo Fisher, #17-9924-82; RRID:AB_657858) were used.

Cell lines

RAW264.7 cells (sex: male) were purchased from ATCC (Cat.#TIB-71; RRID:CVCL_0493) and propagated in the lab; knockout cell lines were derived from parental RAW264.7 cells as described below. RAW264.7 cells were cultured in RPMI 1640 medium (Wisent, Cat.#350-000-CL) supplemented with 5% fetal bovine serum (FBS; Wisent, Cat.#091-150 or Cat.#080-150) in a humidified atmosphere at 37˚C, 5% CO2. HEK293T cells were purchased from ATCC (Cat.#CRL-3216) and propagated in the lab. HEK293T cells were cultured in DMEM medium (Wisent, Cat.#319-070-CL) supplemented with 10% fetal bovine serum (FBS; Wisent, Cat.#091-150).

Murine primary cell culture

Wild-type C57BL/6 and BALB/c mice were originally purchased from the Jackson Laboratory and were bred and housed under specific pathogen-free conditions at the Division of Comparative Medicine (DCM) of the University of Toronto. All experiments were performed according to DCM guidelines, using protocols approved by the Local Animal Ethics Review Committee (protocol 20012048 to S.E.G.). Primary bone marrow-derived macrophages (BMDMs) were derived from 7 to 30-week-old male and female mice as follows: the bone marrow was flushed from femurs and tibiae into RPMI 1640 and centrifuged at 3000 × g for 1 min. Cells were resuspended in RPMI 1640 containing L-glutamine and sodium pyruvate (Wisent, #350-007-CL) and supplemented with 20% conditioned medium from L929 cells, 10% FBS (Wisent, #080-150), 1% penicillin/streptomycin (Wisent, #450-201-EL), 1% non-essential amino acids (Thermo Fisher, #11140076), and 0.1% 2-mercaptoethanol (Thermo Fisher, #21985023). Cells were incubated in a humidified incubator at 37 ˚C, 5% CO2 for 7 days (replacing the medium on the 4th day) to differentiate the cells into macrophages. BMDMs were seeded at 2×106 cells/well in 24-well plates on the 7th day.

Microbial strains and cultures

Shigella flexneri wild-type strain M90T63 and BS176, a non-invasive, virulence plasmid-cured derivative of M90T64, were cultured at 37 ˚C in tryptic soy broth (TSB; Wisent, #800-056-CG) medium containing 100 µg/mL spectinomycin (Bioshop, #SPE201) or 50 µg/mL ampicillin (Millipore-Sigma, #A9518), respectively. TSB-agar plates were supplemented with 0.01% Congo Red (Millipore-Sigma, #C6277) and the appropriate antibiotic in order to select for virulent (red) or non-virulent (white) colonies.

E. coli strains XL1-blue and BL21 (DE3) were cultured at 37˚C in LB (Lennox) broth (Bioshop, #LBL405). Ampicillin (50 μg/mL) was added to maintain pET22b-PA, as well as pX330 and pHR constructs. Bacillus megaterium was cultured at 37˚C in LB broth; tetracycline (10 μg/mL; Millipore-Sigma, #T3258) was added to maintain pWH1520-LF.

Purification of PA and LF

Protective antigen was purified from E. coli and lethal factor was purified from B. megaterium65.

Generation of knockout cell lines

RAW264.7-Nlrp1b–/– was described previously24.

CRISPR guides targeting Caspase-1, Caspase-3, Caspase-8, Gsdmd, Gsdme, Myd88, Tlr4, and Ticam-1, were designed (Table S2) using the CRISPR design tool from MIT (http://crispr.mit.edu; no longer available), CRISPOR, or Synthego (https://www.synthego.com). Each pair of oligonucleotides was annealed and then ligated to BbsI-digested pX330-U6-Chimeric_BB-CBh-hSpCas9 (a gift from Feng Zhang; Addgene plasmid # 42230; http://n2t.net/addgene:42230; RRID:Addgene 42230) to generate pX330 constructs66.

Homologous regions flanking each guide RNA were amplified from RAW264.7 genomic DNA (QIAamp DNA Mini kit; QIAGEN, #51304) using Phusion high-fidelity DNA polymerase (Thermo Fisher, #F-530L). Primers containing restriction site overhangs were designed to amplify homologous arms of approximately 1 kb each that were ideally less than 10 nucleotides from the Cas9 cut site (Table S2). Each amplicon was digested with the appropriate restriction enzyme and ligated to MCS1-EF1α-GFP-T2A-Puro-pA-MCS2 (System Biosciences, #HR410PA-1) to generate pHR constructs.

RAW264.7-Tlr4-/- was generated by electroporating 2×106 low-passage RAW264.7 cells with 750 ng pX330-Tlr4 guide RNA using an Amaxa Nucleofector™ I and Amaxa™ Cell Line Nucleofector™ Kit V (Lonza, # VCA-1003), according to the manufacturer’s instructions. Cells were expanded, and then were selected for the absence of TLR4 using a fluorescently labeled TLR4 antibody. Briefly, one million RAW264.7-WT and RAW264.7-Tlr4-gRNA cells were washed twice with ice-cold PBS, centrifuged (100 g for 10 min) and resuspended in 100 μL of PBS containing 5% FBS. Cells were incubated at 4 ˚C for 20 min with 1 μL mouse Fc block™ (BD Biosciences, #553141; RRID:AB_394656), then incubated in the dark at 4 ˚C for 30 min with 2.5 μL APC-conjugated TLR4/MD2 antibody (Thermo Fisher, #17-9924-82; RRID:AB_657858). Cells were washed twice in ice-cold PBS, then resuspended in PBS containing 5 mM EDTA, 25 mM HEPES pH 7, and 1% FBS. A BD FACSAria™ IIIu cell sorter was used to select a population of RAW264.7-Tlr4-gRNA cells that showed little or no APC staining relative to RAW264.7-WT. This population was expanded, and single-cell clones were isolated by diluting into 96-well plates (~0.05 cells/well). Knockout clones were confirmed by Western blotting for TLR4.

For Caspase-1–/–, Caspase-3–/–,Caspase-8–/–, Myd88–/–, and Ticam-1–/– cell lines, low-passage RAW264.7 cells were electroporated as above with 375 ng of each pX330-guide RNA plasmid and its corresponding homologous region donor vector, which was linearized prior to electroporation. Surviving cells were allowed to recover, then were grown in complete medium containing 6 μg/ml puromycin (Millipore-Sigma, #P9620) to select for integration of the selection cassette. Cells were then seeded into 96-well plates (~0.05 cells/well) to isolate single-cell clones. Knockout was validated by Western blotting for Caspase-1, Caspase-3, Caspase-8, and MyD88. As TRIF (Accession # NP_778154.1) was undetectable by Western blotting, knockout was confirmed by sequencing of genomic DNA (QIAamp DNA Mini kit; QIAGEN, #51304). One mutation resulted in the introduction of the GFP/puromycin selection cassette after the cut site (nucleotide 283 of the coding region), while the second mutation was the insertion of a T. Both mutations resulted in a frameshift and a premature stop codon.

To generate the Gsdmd–/–Gsdme–/– double knockout cell line, the Gsdme-/- cell line was established first, as described above, and knockout was confirmed by immunoblotting for GSDME. Subsequently, Cre-Lox recombination was used to remove the selection cassette: Gsdme–/– cells were electroporated with 500 ng pCDH-CB-iCre (a gift from Kazuohiro Oka [Addgene plasmid # 72257; http://n2t.net/addgene:72257; RRID:Addgene 72257]) and GFP-negative cells were collected using a BD FACSAria IIIu™ cell sorter. The resulting Gsdme-/- GFP-negative cells were then electroporated with Gsdmd guide RNA and homologous region donor plasmids, and cells were selected for puromycin resistance, as above. Knockout was validated by immunoblotting for both GSDMD and GSDME.

Plasmid construction

To generate the caspase-8 expression plasmid, total RNA was isolated from approximately 1.5 million RAW264.7 cells using RNeasy mini kit (QIAGEN, #74106) and was treated with RNase-free DNase (QIAGEN, #79254). RNA was reverse transcribed with iScript™ Advanced cDNA Synthesis kit (Bio-Rad, #1725037). Primers containing the start and stop codon of caspase-8 were used to amplify caspase-8 from cDNA (Table S2). pLJM1-EGFP (a gift from David Sabatini (Addgene plasmid # 19319; http://n2t.net/addgene:19319 ; RRID:Addgene_19319)), and the caspase-8 PCR product were digested with AgeI and BstBI (NEB) and ligated using T4 DNA ligase (NEB). The resulting construct (pLJM1-caspase-8) was confirmed by sanger sequencing. QuikChange Site-Directed Mutagenesis Kit (Agilent, #200519) was used to generate pLJM1-Caspase-8-C362A, following the manufacturer’s protocol (Table S2).

To generate NLRP1B point mutants, nucleotides encoding Iso598 to Leu965 of murine NLRP1B allele 1 were ordered with Asp 788, 824 and 849 mutated to alanine. pNTAP-NLRP1B and NLRP1B598-965 were digested with EcoRV and XbaI and ligated together to generate the mutant constructs. Constructs were ordered using Invitrogen GeneArt Gene Synthesis services.

pCR-Flag-IKKβ was a gift from Hiroyasu Nakano (Addgene plasmid # 15465 ; http://n2t.net/addgene:15465 ; RRID:Addgene_15465). QuikChange Site-Directed Mutagenesis Kit (Agilent, #200519) was used to generate pCR-FLAG-IKKβSSEE following the manufacturers (Table S2).

Plasmids pNTAP-NLRP1B, pNTAP-NLRP1BΔ749-870, pcDNA3-caspase-1-FLAG and, pcDNA3-caspase-1-C284A-FLAG have been described previously10,56.

Transfection of HEK293T cells

HEK293T cells were seeded into a 6-well dish at 5 × 105 cells/well and incubated at 37°C, 5% CO2 for approximately 16 h. 4 µg of total plasmid DNA was mixed with 100 µL of serum-free DMEM and then 16 µL of 1 mg/mL PEI (Polyethyleneimine) was added and vortexed. The mixture was incubated for 20 minutes and room temperature. Cells were washed 3x in DMEM supplemented with 10% FBS and then 100 µL of the DNA:PEI mixture was added to each well and incubated for 24 h.

Western blotting

RAW264.7 cells (wild-type, Caspase-1–/–, Caspase-3–/–, Caspase-8–/–, Gsdmd–/–Gsdme–/–, MyD88–/–, Ticam1–/–, and Tlr4–/–) were seeded into 24-well plates at 2 × 106 cells/well and incubated at 37˚C, 5% CO2 for approximately 16 h. Cells were treated at 37 ˚C, 5% CO2 for 2 h, unless otherwise indicated, with LPS (1 μg/mL; Millipore-Sigma, #L4391), 2DG (200 mM; Millipore-Sigma, #D8375), Z-VAD-FMK (20 μM; Enzo Life Sciences, #ALX-260-020-M001), Z-IETD-FMK (50 μM; R&D Systems, #FMK007), actinomycin D (1 μg/mL; Millipore-Sigma, #A9415), dynasore (80 μM; Millipore-Sigma, #1202867-00-2), lethal toxin (10-8 M PA and 10-9 M LF), MRT67307 (10 μM; Millipore-Sigma, #SML0702), MG132 (10 μM; Millipore-Sigma #474790), Nec1s (50 μM, Cayman Chemical, #20924),  SM164 (1 μM; BioVision, #B1816),  SB202190 (10 μM; Millipore-Sigma, #S7067), TNFα (50 ng/mL; PeproTech, #315-01 A), TPCA (10 μM; Abcam, #145522), trovafloxacin (10 μM; Millipore-Sigma, #PZ0015), or (5Z)-7-oxozeaenol (0.2 μM or as indicated; Cayman Chemical, #17459), as indicated. Cells were pre-treated for 1 h with dynasore (or 0.5% DMSO); for 30 min with NSCI (or 0.3% DMSO); or for 20 min with trovafloxacin (or 0.1% DMSO).

BMDM cells were seeded into 24-well plates at 2 × 106 cells/well and incubated at 37 ˚C, 5% CO2 for approximately 16 h. Cells were pre-treated for 1 h at 37˚C, 5% CO2 with dynasore (248 µM or 1.6% DMSO), then were treated with LPS (200 ng/mL), 2DG (200 mM), or dynasore, as indicated, for an additional 2 h.

Cells were lysed in EBC buffer (50 mM Tris pH 8.0, 120 mM NaCl, 0.5% NP-40) containing 1 mM PMSF or 1X Halt™ Protease and Phosphatase Inhibitor Cocktail (Thermo Fisher, #78441) and were rotated for 1 h at 4 ˚C or sonicated 3 times for 5 s on ice. For NLRP1B blots, HEK293T and BMDMs were lysed in RIPA buffer followed by sonication 2x for 3 s. Lysates were clarified by centrifugation at 20,000 × g for 10 min at 4 ˚C and quantified using the Bio-Rad Protein Assay (Bio-Rad, #500-0006). Equivalent amounts of protein were boiled for 5 min (with the exception of lysates being probed for TLR4 or NLRP1B), then were resolved on SDS-PAGE under reducing conditions and transferred to a PVDF membrane (Bio-Rad, #162-0177) in 20% methanol, 22 mM Tris, 171 mM glycine and 0.01% SDS. Membranes were blocked for 1 h at room temperature in TBS-T (20 mM Tris pH 7.6, 137 mM NaCl and 0.05% Tween 20) containing 5–10% skim milk, then incubated overnight at 4 ˚C in primary antibody, diluted according to manufacturer’s instructions in TBS-T containing 5% BSA and 0.02% sodium azide. Membranes were washed in TBS-T and incubated at room temperature for 45 min in HRP-conjugated secondary antibody, diluted 1:30,000 in blocking solution. After the last wash, membranes were incubated with SuperSignal™ West Dura Extended Duration substrate (Thermo Fisher, #34076), Pierce™ ECL Western Blotting Substrate (Thermo Fisher, #32106), or SuperSignal™ West Femto Maximum Sensitivity substrate (Thermo Fisher, #34096). Proteins were visualized on a G:BOX Chemi XX6 imaging system (Syngene) using GeneSys imaging software (Syngene).

Propidium iodide uptake by flow cytometry

RAW264.7 cells (wild-type, Caspase-1–/–, Caspase-3–/–, Caspase-8–/–, Gsdmd–/–Gsdme–/–, MyD88–/–, Ticam1–/–, and Tlr4–/–) were seeded at 1.5 × 106 cells/well into 24-well plates and incubated at 37 ˚C, 5% CO2 for approximately 16 h. Cells were treated at 37˚C, 5% CO2 for 3 h with LPS (1 μg/mL), 2DG (200 mM), Z-VAD-FMK (20 μM), Z-IETD-FMK (50 μM), or trovafloxacin (10 μM), as indicated. Cells were pre-treated for 30 min with NSCI (or 0.3% DMSO), or for 20 min with trovafloxacin (or 0.1% DMSO). Membrane integrity was assessed by replacing treatment medium with 37˚C flow assay buffer (2% FBS and 5 mM EDTA in PBS) containing 1 µg/mL propidium iodide (Millipore Sigma, #P4864) and incubated in the dark for 30 min at room temperature. Approximately 1 × 106 cells (in 1 mL flow assay buffer) were strained through a 35 μm filter (Corning, #352235) to remove cell clumps and placed on ice until analysis by flow cytometry. For trovafloxacin experiments, TFX (or 0.1% DMSO) was added during propidium iodide incubation. Cells were analyzed using a BD™ LSRII flow cytometer (BD Biosciences) and BD FACSDiva™ software (BD Biosciences). FSCA/H characteristics of cells were used to exclude doublets from the analysis. Ten-thousand single cells were assayed for the emission of photons within the range of the B610/20 filter. The propidium iodide positive (PI + ) gate was set such that untreated RAW264.7 cells maintained a PI+ population of approximately 1–7%. Data analysis was performed on FlowJo software, and propidium iodide-positive values were plotted using GraphPad Prism.

Shigella infection

RAW264.7 cells (wild-type, Nlrp1b–/–, Tlr4–/–, and Caspase-8–/–) were seeded into 24-well plates at 2 × 106 cells/well and incubated overnight at 37 ˚C, 5% CO2. Shigella flexneri strain M90T was grown as described above. A single virulent colony was used to inoculate 1 ml TSB containing spectinomycin and was grown overnight at 37 ˚C with shaking. Shigella flexneri strain BS176 was grown as described above. A single non-virulent colony was used to inoculate 1 ml TSB-ampicillin and grown overnight at 37 ˚C with shaking. M90T cultures (diluted 1:100) and BS176 cultures (diluted 1:300) were grown to an OD600 of 0.5. Cultures were centrifuged at 1000 × g for 5 min, washed with PBS, and resuspended in serum-free RPMI.

RAW264.7 cells were washed once with PBS and were either uninfected or were infected with S. flexneri at the indicated multiplicity of infection (MOI). Plates containing RAW264.7 cells plus Shigella were centrifuged at 1000 × g for 13 min, and then incubated for 30 min at 37 ˚C, 5% CO2. The medium was aspirated; cells were washed once with PBS and treated with RPMI containing 5% FBS and 30 μg/ml gentamycin (Wisent, #450-134-XL) for 1 h at 37 ˚C, 5% CO2. Lysates were prepared as described above.

For MG132 assays, RAW264.7 cells were pre-treated for 30 min at 37˚C, 5% CO2 in complete medium containing 10 μM MG132 or 0.1% DMSO. Cells were washed once with PBS and infected with S. flexneri as above; MG132 (or 0.1% DMSO) was present in treatments throughout the duration of the experiment.

Measurement of ATP levels

Intracellular ATP levels were determined using the CellTiter-Glo® Luminescent Cell Viability Assay (Promega, #G7571), according to manufacturer’s instructions. Briefly, treated cells were equilibrated at room temperature for 30 min, then CellTiter-Glo® reagent was added and incubated with shaking for 2 min at room temperature. Samples were clarified by centrifugation, diluted 1:200 in water, and incubated in the dark at room temperature for 10 min. Luminescence was recorded using a Lumat LB 9507 Ultra Sensitive Tube Luminometer (Berthold Technologies) with an integration time of 0.3 seconds. Background luminescence was subtracted from sample values, which were then graphed using GraphPad Prism.

Real-time qPCR

Total RNA was isolated from approximately 3 million RAW264.7 cells using RNeasy mini kit (QIAGEN, #74106) and was treated with RNase-free DNase (QIAGEN, #79254). RNA (1 μg) was reverse transcribed with iScript™ Advanced cDNA Synthesis kit (Bio-Rad, #1725037). Genomic DNA was isolated from RAW264.7 cells (QIAamp DNA Mini kit; QIAGEN, #51304) and used to generate a standard curve. Three-fold dilutions of genomic DNA, from 8100 copies to 100 copies, or cDNA equivalent to 20 ng RNA, were added to the SsoAdvanced™ Universal SYBR Green Supermix (Bio-Rad, #1725270) and 800 nM each primer. All standards and cDNA samples were assayed in triplicate wells of a 384-well plate (Bio-Rad #HSP3805) in a reaction volume of 10 μL. qPCR was performed on the Bio-Rad CFX384™ Real-Time PCR Detection System using the following amplification conditions: 95 ˚C (3 min); 40 cycles of 95˚C (10 s), 60˚C (30 s); 95 ˚C (10 s), 65˚C (30 s); 65˚C (5 s), +0.5˚C/cycle (60 cycles). Primers used to determine mRNA levels are listed in Table S2. Il1b mRNA levels (expressed relative to untreated sample) were determined using relative quantification (EΔΔCt method) and were normalized to β-actin. Transcript levels of Gasdermin family members and β-actin were extrapolated from the standard curve. Values represent the means of three independent experiments ± SEM in which each sample was performed in triplicate.

Co-immunoprecipitation experiments

RAW264.7 cells were seeded into 10 cm dishes at a density of 4 × 107 cells/dish and incubated overnight at 37 ˚C, 5% CO2. For TLR4-RIPK1 interaction, duplicate dishes of RAW264.7 cells were treated for 30 min with LPS, 2DG, or a combination of LPS and 2DG (37˚C, 5% CO2). For RIPK1-caspase-8 interactions, single dishes of RAW264.7 cells were treated with LPS, 2DG, or a combination of LPS and 2DG, as indicated, for 1 h at 37˚C. Cells were scraped in PBS, centrifuged (220 g, 3 min), and lysed by rotating for 30 min at 4 ˚C in EBC lysis buffer containing 1X Halt™ Protease and Phosphatase Inhibitor Cocktail. Lysates were clarified by centrifugation (20,000 × g for 10 min at 4 ˚C) and were quantified using the Bio-Rad Protein Assay. Equivalent amounts of lysate were pre-cleared with BSA-blocked Protein A resin (Repligen, #10-2500-03) by rotating for 30 min at 4 ˚C, then centrifuged (10,000 × g for 5 min.). Supernatants were then incubated with 1 μg of mouse monoclonal RIPK1 antibody or 1 μg of irrelevant-antibody control (mouse anti-HA antibody), as indicated, and rotated for 2 h at 4˚C; BSA-blocked Protein A resin was added and samples were rotated for an additional 1 h at 4 ˚C. Beads were washed three times with EBC lysis buffer, then eluted by incubating for 10 min at room temperature with SDS sample buffer. Samples were centrifuged (16,000 × g, 2 min); eluates and input samples were heated to 100˚C for 5 min for caspase-8 blots but were not boiled for TLR4 blots. Samples were resolved by SDS-PAGE and blotted as described above. To detect TLR4 being immunoprecipitated by RIPK1, blots were incubated at 4 ˚C for 2–4 days in primary antibody and were developed using SuperSignal™ West Femto Maximum Sensitivity substrate.

RIPK1 phosphorylation/Alkaline phosphatase treatment

RAW264.7 cells were seeded into 24-well plates at a density of 2 × 106 cells/well and were incubated overnight at 37 ˚C, 5% CO2. Duplicate wells were treated with RPMI alone (untreated) or 1 μg/mL LPS for 30 min at 37 ˚C, then scraped in PBS. Cells were centrifuged (4500 × g, 3 min) and each treatment was divided into two samples. Cells to be treated with alkaline phosphatase (Millipore-Sigma, #P6774) were sonicated on ice (2 × 3 s) in EBC lysis buffer containing PMSF, while those that would not be treated with alkaline phosphatase were sonicated in EBC containing 1X Halt™ Protease and Phosphatase Inhibitor Cocktail. Lysates were clarified by centrifugation (16,000 × g, 10 min), and 20 μg were added to a reaction containing 0.1 mM DTT in a total volume of 40 μL EBC lysis buffer. Alkaline phosphatase (0.5 μL) was added to samples, as indicated, and tubes were incubated on ice for 20 min. Reactions were terminated by the addition of SDS sample buffer; samples were resolved by SDS-PAGE and subjected to Western blotting as described above.

Statistics and reproducibility

Statistic were performed on samples containing three or more replicates from independent experiments. We define an independent experiment/replicate as a procedure performed on different days, using the same cell line. The statistical test utilized for each experiment can be found within the figure caption. We declare that all experiments were reproducibile to a minimum of three independent experiments except for Fig. 7h which was reproduced twice.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Supplementary information

Supplementary Information

Description of Additional Supplementary File

Supplementary Data

reporting summary

Supplementary information

The online version contains supplementary material available at 10.1038/s42003-024-06882-3.

Acknowledgements

This work was supported by grants from the CIHR to S.E.G. and J.M. J.J.M received funding from the Ontario Graduate Scholarship program. We thank Keisuke Mori for generating a plasmid used in this study.

Author contributions

Conceptualization, J.J.M., S.S., and J.M.; Investigation, J.J.M., S.S., J.N.-Z., and K.C.; Writing-Original Draft, J.J.M., S.S., and J.M.; Supervision, S.E.G. and J.M.; Funding Acquisition, S.E.G., J.J.M., and J.M.

Peer review

Peer review information

Communications Biology thanks Seth Masters and the other, anonymous, reviewers for their contribution to the peer review of this work. Primary Handling Editors: Kaiwen Chen and Tobias Goris.

Data availability

All data supporting the findings of this study are available within the paper and its Supplementary Information. All source data and unedited/uncropped western blots can be found in the supplementary data.

Competing interests

The authors declare no competing interests.

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
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