Research Article

Functional degradation: A mechanism of NLRP1 inflammasome activation by diverse pathogen enzymes

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Science  05 Apr 2019:
Vol. 364, Issue 6435, eaau1330
DOI: 10.1126/science.aau1330

Degradation triggers the alarm

Inflammasomes are multiprotein complexes that orchestrate proinflammatory cytokine secretion and cell death. Proteases such as anthrax lethal factor can activate an inflammasome known as NLRP1B, but the mechanism for this activation has been unclear. Chui et al. used genome-wide knockout screens to show that proteolysis of NLRP1B by lethal factor induces proteasomal degradation of the amino-terminal domains of NLRP1B and eventual cell death. Sandstrom et al. found that degradation of the amino-terminal domains of NLRP1B resulted in the release of a carboxyl-terminal fragment that activates caspase-1. This process, called “functional degradation,” allows the immune system to detect pathogen-associated activities, much as it recognizes pathogen-associated antigens.

Science, this issue p. 82, p. eaau1330

Structured Abstract

INTRODUCTION

Detection of pathogens by the innate immune system is an essential first step in successful host defense against infection. Pathogens are typically detected by germline-encoded innate immune receptors that bind directly to pathogen-derived ligands. Here, we describe a distinct mechanism of pathogen-sensing mediated by an immune sensor protein called NLRP1B. NLRP1B is not known to bind directly to a pathogen ligand but instead appears to detect the enzymatic activity of lethal factor (LF), a protease toxin produced by the anthrax bacterium, Bacillus anthracis. LF was previously shown to cleave NLRP1B at its N terminus, leading to activation of a multiprotein complex called an inflammasome. Inflammasomes initiate immune responses by activating a proinflammatory protease called caspase-1. However, the mechanism by which cleavage of NLRP1 results in inflammasome assembly was unknown.

RATIONALE

NLRP1B is a member of the nucleotide-binding domain leucine-rich repeat (NLR) protein superfamily. Unlike other NLRs, NLRP1B encodes a C-terminal function-to-find domain (FIIND) followed by a caspase activation and recruitment domain (CARD). The FIIND undergoes constitutive autoproteolytic processing, which cleaves NLRP1B into two separate polypeptides, which nevertheless remain noncovalently associated. The functional importance of this distinctive protein architecture has been a long-standing mystery. Also unexplained is the prior observation that NLRP1B activation requires the activity of the proteasome. We therefore sought a mechanism that explains how N-terminal cleavage by LF, FIIND autoprocessing, and the proteasome cooperate to initiate NLRP1B inflammasome formation.

RESULTS

We found that N-terminal cleavage of NLRP1B by LF protease results in destabilization of NLRP1B and its degradation by the proteasome. Paradoxically, NLRP1B inflammasome activation inversely correlated with the stability of the NLRP1B protein after protease cleavage: Proteasome inhibitors stabilized cleaved NLRP1B and prevented NLRP1B inflammasome activation. Indeed, we found that targeted degradation of NLRP1B induced inflammasome activation, even in the absence of protease cleavage, indicating that proteasomal degradation is not only necessary but also sufficient for NLRP1B inflammasome activation. FIIND autoprocessing was required for NLRP1B activation regardless of whether activation occurred via LF cleavage or targeted degradation. To explain these observations, we hypothesized that the function of the proteasome during NLRP1B activation is to degrade the N-terminal domains of NLRP1B, leading to release of a bioactive C-terminal CARD-containing fragment. Consistent with this model, we found that the C-terminal CARD-containing fragment of NLRP1B is sufficient to self-assemble, recruit caspase-1, and form a functional inflammasome. LF protease treatment resulted in specific degradation of the N-terminal domains of NLRP1B, whereas the liberated C-terminal domain associated with inflammasome puncta in cells. We refer to this proteasome-dependent mechanism of NLRP1B inflammasome activation as functional degradation. Similar conclusions were reached independently by Bachovchin and colleagues (this issue).

The functional degradation model raised the possibility that NLRP1B could sense not only proteases but also any pathogen effector that induced proteasomal degradation of NLRP1B. In accord with this prediction, we identified IpaH7.8, an E3 ubiquitin ligase secreted by the pathogen Shigella flexneri, as another activator of NLRP1B. IpaH7.8 directly and specifically ubiquitylates NLRP1B, leading to NLRP1B degradation and inflammasome activation in Shigella-infected macrophages.

CONCLUSION

We speculate that functional degradation may also explain the activation of other FIIND-containing proteins, such as CARD8 and PIDD1. More broadly, our results reveal a mechanism, distinct from recognition of pathogen-associated ligands, by which hosts can initiate protective immune responses via detection of diverse pathogen-associated activities.

Diverse pathogen enzymes activate the NLRP1B inflammasome by functional degradation.

NLRP1B autoprocesses within the FIIND to generate two noncovalently associated fragments. Cleavage of NLRP1B by anthrax LF, or ubiquitylation by Shigella IpaH7.8, results in proteasome-mediated degradation of NLRP1B, leading to release of the bioactive C-terminal fragment and inflammasome assembly. LRR, leucine-rich repeat; NBD, nucleotide binding domain; Nt, N terminus; Ub, ubiquitylation.

Abstract

Inflammasomes are multiprotein platforms that initiate innate immunity by recruitment and activation of caspase-1. The NLRP1B inflammasome is activated upon direct cleavage by the anthrax lethal toxin protease. However, the mechanism by which cleavage results in NLRP1B activation is unknown. In this study, we find that cleavage results in proteasome-mediated degradation of the amino-terminal domains of NLRP1B, liberating a carboxyl-terminal fragment that is a potent caspase-1 activator. Proteasome-mediated degradation of NLRP1B is both necessary and sufficient for NLRP1B activation. Consistent with our functional degradation model, we identify IpaH7.8, a Shigella flexneri ubiquitin ligase secreted effector, as an enzyme that induces NLRP1B degradation and activation. Our results provide a unified mechanism for NLRP1B activation by diverse pathogen-encoded enzymatic activities.

In animals, pathogens are generally recognized by germline-encoded innate immune receptors that bind directly to conserved pathogen-associated molecular patterns (PAMPs) such as bacterial lipopolysaccharide or flagellin (1). The recognition of PAMPs permits robust self-nonself discrimination, but PAMP receptors cannot readily distinguish pathogens from nonpathogens because PAMPs are found on innocuous microbes as well as pathogens. Plants also use germline-encoded receptors to detect PAMPs but additionally respond to infection by indirect detection of secreted pathogen enzymes called effectors (2). In this mode of recognition, called effector-triggered immunity, intracellular proteins of the nucleotide-binding domain leucine-rich repeat (NLR) superfamily sense effector-induced perturbation of host signaling pathways. Because innocuous microbes do not deliver effectors into host cells, effector-triggered immunity is inherently pathogen specific. Animals may also detect pathogen-encoded activities (310), but there have been relatively few examples of this mode of pathogen recognition understood in molecular detail.

We sought to determine how the mammalian NLR protein NLRP1 senses pathogen-encoded activities. NLRP1 is the foundational member of a class of proteins that form inflammasomes (11), multiprotein platforms that initiate immune responses by recruiting and activating proinflammatory proteases, including caspase-1 (CASP1) (1214). CASP1 cleaves and activates specific cytokines [interleukin (IL)–1β and IL-18] and the pore-forming protein gasdermin D, resulting in a type of host cell death called pyroptosis. In certain murine strains, a NLRP1 paralog, NLRP1B, is activated via direct proteolysis of its N terminus by the lethal factor (LF) protease secreted by Bacillus anthracis (1518). Previous studies demonstrated that N-terminal proteolysis is sufficient to initiate NLRP1B inflammasome activation (16), but the molecular mechanisms by which proteolysis activates NLRP1B have been elusive.

Like other NLRs, NLRP1B contains a nucleotide-binding domain (NBD) and leucine-rich repeats (LRRs) (Fig. 1A). However, NLRP1B also exhibits several distinctive features. First, the NLRP1B caspase activation and recruitment domain (CARD) is C-terminal instead of N-terminal. Second, NLRP1B is the only NLR that contains a function-to-find domain (FIIND). The FIIND constitutively undergoes an autoproteolytic event, resulting in two separate NLRP1B polypeptides that remain noncovalently associated (1921). Mutations that abolish FIIND autoprocessing block inflammasome activation (20, 21), but it remains unclear why FIIND autoprocessing is essential for NLRP1B function. Last, proteasome inhibitors specifically block NLRP1B inflammasome activation but do not affect other inflammasomes or inhibit LF protease, suggesting that the proteasome is specifically required for activation of NLRP1B itself (2226).

Fig. 1 The N-terminal domain of NLRP1B does not mediate autoinhibition.

(A) Schematic of mouse NLRP1B domain architecture. Nt, N terminus. FIIND autoprocessing (white triangle) is not complete and thus NLRP1B appears as a doublet [(B to D), upper blot]. Orange triangle, LF cleavage site. (B and C) The indicated N-terminal amino acids of NLRP1 were mutated to alanine [(B), AAA] or glycine-serine-glycine [(C), GGSGG], and inflammasome activation was induced by coexpression of the TEV protease, which cleaves a TEV site engineered into the NLRP1B N terminus. Activation was monitored by immunoblot (IB) for p17, generated upon CASP1 processing of pro-IL-1β. MBP, maltose binding protein tag. (D) The N terminus of NLRP1B was replaced with a heterologous sequence from flagellin (FlaNt), and inflammasome activation was assessed as in (B). (E to G) A TEV site was positioned at the indicated positions along the NLRP1B Nt, and inflammasome activation was assessed as in (B). Cleaved NLRP1B was detected by IB for a C-terminal hemagglutinin (HA) tag [(E), upper blot]. The relative intensity of cleaved IL-1β p17 was plotted relative to the position of the TEV site (F) or the relative level of cleaved NLRP1B protein (G). Gel images are representative of a single experiment performed once [(B), (C), and (E)] or three times (D). a.u., arbitrary units; aa, amino acid.

Degradation of NLRP1B correlates with its activation

Cleavage of NLRP1B by LF results in a loss of 44 amino acids from the N terminus of NLRP1B, an event that correlates with its activation (1518). We and others have previously proposed an autoinhibition model to explain NLRP1B activation (13, 27). In this model, the N terminus of NLRP1B functions as an autoinhibitory domain that is lost after cleavage by LF. The NLRP1B N terminus may mediate autoinhibition either through direct engagement with other NLRP1B domains or by binding to an inhibitory cofactor. A clear prediction of the autoinhibition model is that sequences within the N terminus should be required to prevent spontaneous inflammasome activation. To identify such sequences, we systematically mutated the NLRP1B N terminus by replacing groups of three consecutive amino acids with alanine residues (Fig. 1B) or by replacing groups of five sequential amino acids with a flexible GGSGG (G, Gly; S, Ser) motif (Fig. 1C). Each mutant was also engineered to contain an N-terminal TEV (tobacco etch virus) protease site to enable inducible NLRP1B cleavage and activation (16, 28). Inflammasome activity was monitored by CASP1-dependent processing of pro-IL-1β to p17 in a reconstituted inflammasome system in transfected 293T cells (16, 28). To our surprise, none of the mutants were autoactive, even though all showed full activity upon cleavage. To test if any N-terminal sequence could mediate autoinhibition, we replaced the entire N terminus with a heterologous α-helical domain from bacterial flagellin. Again, surprisingly, the hybrid Fla-NLRP1B protein was not autoactive but was still functional after N-terminal cleavage (Fig. 1D). Together with prior results (20, 28, 29), these data led us to reconsider a model in which specific N-terminal sequences of NLRP1B mediate autoinhibition.

We next asked whether the precise site of N-terminal proteolysis is a major determinant of NLRP1B activation. We generated a series of NLRP1B variants in which a TEV protease cleavage site was positioned at regular intervals from the N terminus. We found that cleavage of as few as 10 amino acids from the N terminus was sufficient to activate NLRP1B. Furthermore, there was no substantial correlation between the position of TEV cleavage and NLRP1B activity (Fig. 1, E and F). By contrast, there was a marked negative correlation between the amount of TEV-cleaved NLRP1B protein and inflammasome activation (Fig. 1, E and G). This correlation, as well as prior evidence that proteasome inhibitors block NLRP1B activity (2226), led us to hypothesize that proteasome-mediated degradation of NLRP1B is an important step in its activation. The proteasome inhibitors MG132 and bortezomib not only abrogated NLRP1B activation in our reconstituted 293T system but also prevented loss of the cleaved NLRP1B protein after LF treatment (Fig. 2A). By contrast, the inhibition of p97/VCP by NMS-873 had no effect on NLRP1B activation.

Fig. 2 Degradation of NLRP1B is necessary and sufficient for NLRP1B inflammasome activation.

(A) 293T cells transfected with constructs encoding NLRP1B, CASP1, and pro-IL-1β were assayed for inflammasome activation, as in Fig. 1, in the presence (+) or absence (–) of proteasome inhibitors [MG132 (10 μM) and bortezomib (1 μM)] or p97/VCP inhibitor [NMS-873 (0.5 μM)]. (B) Immortalized 129 BMDMs were treated with LF to activate the NLRP1B inflammasome ± MG132. Endogenous NLRP1B was detected by IB with 2A12. Relative band intensities are indicated. (C) The plant AID was fused to the N terminus of the indicated GFP-NLRP1B variants. Specific degradation was induced with IAA in TIR1-expressing 293T cells. Inflammasome activation was assessed by IB for IL-1β p17. Relative band intensities are indicated. Gel images are representative of experiments performed at least three times. S984A, Ser984→Ala.

Proteasomal degradation is necessary and sufficient for NLRP1B activation

To determine whether endogenous NLRP1 is also lost from cells after cleavage, we derived a monoclonal antibody (2A12) against the C-terminal CARD of NLRP1B (fig. S1). Using this antibody, we could track endogenous NLRP1B in 129S1/SvimJ (129) bone marrow–derived macrophages (BMDMs) after treatment with LF (Fig. 2B). As in reconstituted 293T cells, LF treatment led to a loss of full-length NLRP1B, which was at least partially reversed by MG132 treatment. By contrast, proteasome inhibitors have no effect on NLRP3 (23, 26) or NAIP5/NLRC4 inflammasome activation (fig. S2), consistent with the hypothesis that the proteasome functions distinctively in NLRP1B inflammasome activation.

A limitation of our study is that the precise mechanism by which NLRP1B is targeted to the proteasome after LF cleavage remains unknown. However, a protein quality control pathway known as the N-end rule pathway ubiquitylates cleaved proteins (30, 31), resulting in their proteasomal degradation, and N-end rule inhibitors block NLRP1B activation (25). Therefore, although our assays were insufficiently sensitive to detect NLRP1B ubiquitylation after LF cleavage, we propose that cleavage of NLRP1B reveals a destabilizing neo-N terminus, which targets NLRP1B for ubiquitylation by N-end rule E3 ubiquitin ligases. Consistent with this hypothesis, in independent and parallel work, the Bachovchin laboratory has identified N-end rule pathway components that mediate NLRP1B ubiquitylation and are critical for LF-mediated NLRP1B activation (32). We note that the TEV-cleavable NLRP1B variants that we examined generate a glycine residue at the neo-N terminus after TEV cleavage. According to the N-end rule, glycine is a stabilizing N-terminal amino acid (33), a prediction apparently at odds with our observation that TEV cleavage results in NLRP1B destabilization (Fig. 1). However, aminopeptidase inhibitors block the LF-mediated killing of RAW264.7 cells (25). Thus, it appears likely that the neo-N terminus generated by primary cleavage is further processed by aminopeptidases, resulting in the exposure of otherwise internal amino acids to N-end rule recognition. When we swapped the P2′ residues between two differentially activated TEV-cleavable NLRP1B variants, we found that the P2′ residue could also modulate activity (fig. S3). Thus, the stability and activity of cleaved NLRP1B depends on more than just the identity of the neo-N-terminal amino acid, consistent with a growing body of evidence that multiple determinants underlie N-end rule degradation (34).

In the above experiments, the inhibition of NLRP1B activation by proteasome inhibitors may have been due to stabilization of a negative regulator of NLRP1B rather than to stabilization of NLRP1B itself. Therefore, we next asked whether specific degradation of NLRP1B is sufficient to induce its activation. To achieve the selective degradation of NLRP1B, the auxin-inducible degron (AID) (35, 36) was fused to the N terminus of NLRP1B. Upon addition of the auxin hormone indole-3-acetic acid (IAA), AID recruits a coexpressed TIR1 E3 ligase that specifically ubiquitylates AID-fusion proteins, targeting them to the proteasome. IAA induced the rapid degradation of the AID-NLRP1B fusion protein and stimulated robust IL-1β processing (Fig. 2C). Notably, FIIND autoprocessing was also required for IAA-induced activation of AID-NLRP1B (Fig. 2C). Thus, the proteasomal degradation of NLRP1B itself is both necessary and sufficient for NLRP1B inflammasome activation.

Functional degradation of NLRP1B liberates an active C-terminal fragment

To explain the seemingly paradoxical observation that NLRP1B degradation leads to its activation, we propose the following functional degradation model (Fig. 3A). This proposal relies on the prior observation that FIIND autoprocessing is required for activation of NLRP1B. The FIIND comprises two separate subdomains, termed ZU5 and UPA, with autoprocessing occurring near the C-terminal end of the ZU5 domain (at Phe983|Ser984) (1921). After autoprocessing, the C-terminal fragment of NLRP1B consists of an UPA domain fused to the CARD that is required for CASP1 recruitment and activation. Before activation, the FIIND(UPA)-CARD fragment is noncovalently associated with the rest of NLRP1B. After cleavage by LF, NLRP1B is targeted to the proteasome, a processive protease that degrades polypeptides by feeding them through a central barrel (37). Critically, however, directional (N to C terminus) and processive degradation of NLRP1B by the proteasome would be interrupted by the covalent break within the autoprocessed FIIND. Thus, we propose that the C-terminal FIIND(UPA)-CARD fragment is released and can seed inflammasome assembly (Fig. 3A).

Fig. 3 NLRP1B is activated by functional degradation.

(A) Model for NLRP1B activation via functional degradation. (i) Constitutive autoprocessing of the NLRP1B FIIND results in two noncovalently associated polypeptides: NBD-LRR-FIIND(ZU5) and FIIND(UPA)-CARD. (ii) LF protease cleavage of the NLRP1B Nt exposes a neo-Nt. (iii) N-end rule factor recognition of the neo-Nt results in ubiquitylation (Ub) of NLRP1B. (iv) NLRP1B is degraded by the proteasome, resulting in (v) release of the FIIND(UPA)-CARD fragment. (vi) The FIIND(UPA)-CARD fragment self-assembles into a high–molecular weight oligomer that (vii) serves as a platform for CASP1 maturation and downstream inflammasome signaling. (B) The ability of the indicated C-terminal HA-tagged expression constructs to induce inflammasome activity was tested in 293T cells, as in Fig. 1. (C) 293T cells were transfected with the indicated amounts (nanograms) of plasmid encoding HA-tagged FIIND(UPA)-CARD fragment (left) or full-length (FL) NLRP1B (± cotransfected TEV protease), and inflammasome activation was measured as in Fig. 1. L, ladder of protein molecular weight standards. (D) 293T cells were transfected with CASP1 and HA-tagged constructs as depicted in (B). Lysates were immunoprecipitated (IP) with anti-CASP1 and immunoblotted (IB) as indicated. (E) 293T cells were transfected with CASP1 and HA-tagged constructs as depicted in (B) and analyzed by native PAGE and IB with anti-NLRP1B antibody (2A12). Proteins were native or denatured with sodium dodecyl sulfate (SDS) as indicated. MW, molecular weight. (F) 293T cells were transfected with an expression construct encoding AID-FIIND(ZU5+UPA)-CARD and treated with IAA or MG132 as indicated. Relative band intensities are shown. Gel images are representative of experiments performed at least three times [(B) and (D) to (F)] or once (C). (G) 293T cells were transfected with expression constructs for ASC (blue) and TEV-cleavable NLRP1B encoding a C-terminal FLAG (green) tag and HA (magenta) tag (inserted after the TEV site, as shown). The number of ASC specks per field (± SD) was quantified for TEV-treated samples and compared with that of cells expressing a S984A FIIND autoprocessing NLRP1B mutant. Representative images depict cytosolic FLAG and HA signal in untreated samples, with FLAG colocalization with ASC specks (white arrowheads) and concomitant loss of the HA signal in TEV-expressing cells. The total number of ASC specks or ASC specks positive for both FLAG and HA (n = 7) or only FLAG (n = 32) or HA (n = 0) in TEV-treated samples is quantified from 12 total fields representative of three independent experiments. Scale bars, 10 μm. Significance was determined by Student’s t test; ***P < 0.001.

The functional degradation model of NLRP1B inflammasome activation has several merits. First, it explains how N-terminal cleavage results in proteasome-dependent NLRP1B activation without a requirement for specific N-terminal autoinhibitory sequences. Second, the model accounts for why the NLRP1B CARD is C-terminal, rather than N-terminal, as only the C terminus of NLRP1B remains after proteasome-mediated degradation. Last, the model explains why FIIND autoprocessing is required for NLRP1B activity: An unprocessed FIIND mutant would be fully degraded and would not release a C-terminal CARD-containing fragment.

A strong prediction of the functional degradation model is that the C-terminal FIIND(UPA)-CARD fragment possesses inflammasome activity. Consistent with prior work (21, 38), we determined that the FIIND(UPA)-CARD fragment was indeed sufficient to promote robust CASP1 activity in our 293T reconstituted inflammasome assay, whereas the full-length FIIND(ZU5+UPA)-CARD appeared inactive despite autoprocessing (Fig. 3B). Likewise, the isolated CARD (lacking any portion of the FIIND) also appeared inactive, implying that the FIIND(UPA) contributes to inflammasome formation. This pattern was observed across a range of expression levels (fig. S4). Notably, the FIIND(UPA)-CARD fragment appears to be a highly potent activator of inflammatory signaling. By titrating the amount of expression construct, we found that the FIIND(UPA)-CARD fragment appeared to be up to ~150 times as potent as TEV-cleavable full-length NLRP1B (Fig. 3C). These results imply that only a small fraction of the total NLRP1B in a cell may need to be degraded to liberate sufficient amounts of the FIIND(UPA)-CARD fragment for robust inflammasome activation. Thus, even if NLRP1B is only degraded a fraction of the time in the “productive” N-to-C-terminal direction, this would still likely be sufficient for robust inflammasome activation.

The functional degradation model predicts that the mature (i.e., assembled) NLRP1B inflammasome consists of the FIIND(UPA)-CARD fragment and CASP1. In support of this hypothesis, only the FIIND(UPA)-CARD fragment coimmunoprecipitated robustly with CASP1 (Fig. 3D) and assembled into higher-order oligomers when assessed via nondenaturing polyacrylamide gel electrophoresis (PAGE) (Fig. 3E). These results indicate that the NBD and LRR domain of NLRP1B are dispensable for inflammasome activation. Indeed, IAA-induced degradation of an AID-FIIND(ZU5+UPA)-CARD fragment is sufficient to induce CASP1 activity (Fig. 3F). Thus, the FIIND-CARD module appears to be sufficient to impart both autoinhibition and proteasome-induced activation to NLRP1B. Prior studies found that NBD and LRR mutants of NLRP1B are constitutively active (27, 28, 38, 39), a result that could be explained by functional degradation if the mutations destabilize NLRP1B. Moreover, although the role of the NBD and LRRs of NLRP1B remains to be determined, we hypothesize that these domains may contribute to recognition of pathogen-encoded effectors.

To visualize the simultaneous degradation of the N-terminal domains and the release of the FIIND(UPA)-CARD fragment upon NLRP1B activation, we engineered a variant of NLRP1B with a C-terminal FLAG tag and an HA tag following the N-terminal TEV cleavage site. In unstimulated cells expressing this variant, FLAG and HA costained within the cytosol (Fig. 3G). By contrast, cells cotransfected with TEV protease showed inflammasome activation, as indicated by formation of “specks” containing the ASC adaptor protein. Notably, we found that the FLAG-tagged UPA-CARD fragment, but not the HA-tagged N-terminal domains, robustly formed puncta colocalized at the ASC speck (Fig. 3G). Moreover, we also observed a near-complete loss of the HA signal, consistent with N-terminal degradation upon TEV cleavage. The FLAG signal was also lost upon TEV cleavage when FIIND processing was disrupted (fig. S5). Thus, the C-terminal UPA-CARD appears to be released to seed inflammasome formation upon proteolytic cleavage and the subsequent proteasomal degradation of the NLRP1B N-terminal domains.

The Shigella effector IpaH7.8 degrades and activates NLRP1B

A major implication of our functional degradation model is that NLRP1B can potentially sense any enzymatic activity that results in NLRP1B degradation in addition to detecting pathogen-encoded proteases such as LF. Several pathogens encode E3 ubiquitin ligases that promote virulence through degradation of host target proteins (40). We therefore tested whether the type III secretion system IpaH family of E3 ubiquitin ligases, encoded by the intracellular bacterial pathogen Shigella flexneri (4143), is detected by NLRP1B. Using our reconstituted 293T cell system, we found that IpaH7.8, but not IpaH1.4, -4.5 or -9.8, markedly reduced NLRP1B protein levels and induced IL-1β processing in an NLRP1B-dependent manner (Fig. 4A). IpaH4.5 also reduced IL-1β levels in cells, but we did not pursue this observation. IpaH7.8 selectively activated the 129 allele but not the C57BL/6 (B6) allele of NLRP1B (Fig. 4B). As expected, FIIND autoprocessing was required for IpaH7.8-induced NLRP1B activation (Fig. 4C). Truncation of either the IpaH7.8 LRR or E3 domains, as well as mutation of the catalytic cysteine residue (CA) required for E3 ligase activity, also abolished IpaH7.8-mediated inflammasome activation (Fig. 4D). We observed that IpaH7.8, but not IpaH9.8 or an IpaH7.8 catalytic mutant (IpaH7.8CA), was able to directly ubiquitylate the 129 allele but not the B6 allele of NLRP1B (Fig. 4E) in a reconstituted ubiquitylation assay. Thus, IpaH7.8/ubiquitylation-dependent degradation of NLRP1B can result in its activation.

Fig. 4 The secreted S. flexneri IpaH7.8 E3 ubiquitin ligase activates NLRP1B.

(A and B) 293T cells were transfected with expression constructs for the 129 [(A) and (B)] or B6 (B) alleles of NLRP1B plus CASP1; pro-IL-1β; and GFP-tagged IpaH1.4, 4.5, 7.8, or 9.8. NLRP1B expression and inflammasome activation was assessed as in Fig. 1. (C) As in (A), but cells were transfected with a Ser984→Ala mutant of NLRP1B. (D) As in (A), but cells were transfected with expression constructs for mutant IpaH7.8. CA, catalytic mutant; ∆E3, deletion of Ub ligase domain; ∆LRR, deletion of LRR. (E) An in vitro ubiquitylation assay (see Materials and methods) was used to assess the ability of IpaH7.8, IpaH7.8 catalytic mutant (7.8CA), or IpaH9.8 to ubiquitylate the 129 or B6 alleles of NLRP1B-FLAG. Reactions were immunoprecipitated with anti-FLAG before IB with anti-ubiquitin or with anti-NLRP1B (2A12). Images are representative of at least three independent experiments [(C) to (E)], except for those depicted in (A) and (B), which were performed twice. (F) WT, Casp1–/–, Nlrp1b–/–, or Nlrc4–/– RAW264.7 cells were infected (MOI 30) with WT S. flexneri strain 2457T (black circle), BS103 (virulence plasmid-cured, white box), Δ7.8 (ipaH7.8 deletion, blue triangle), p7.87.8 strain complemented with pCMD136 ipaH7.8, green inverted triangle), or vec (Δ7.8 strain complemented with pCMD136 empty vector, red diamond). Inflammasome-induced pyroptotic cell death was monitored by assaying for LDH activity in culture supernatants 30 min postinfection (± SD). (G) Immortalized 129 (i129) BMDMs were infected with S. flexneri strains as in (F). Cell lysates were IB with anti-NLRP1B (2A12) or anti-mouse CASP1, or cell death was measured by LDH, as in (F), 2 hours postinfection (± SD). Data in (F) and (G) are representative of at least three independent experiments. Data sets [(F) and (G)] were analyzed using one-way ANOVA (analysis of variance). P values were determined by Dunnet’s multiple comparison post hoc test. *P < 0.05; **P < 0.01; ***P < 0.001.

S. flexneri robustly activates multiple inflammasomes (44) and can cause macrophage cell death in an IpaH7.8- and NLRP1B-dependent manner (4547). However, a connection between IpaH7.8 and NLRP1B has not been established. Consistent with earlier studies (45, 47), wild-type (WT) S. flexneri induced robust lactate dehydrogenase (LDH) release from infected RAW264.7 macrophages, which was reduced in infections with a ΔipaH7.8 mutant (Fig. 4F). Cell killing by the ΔipaH7.8 strain was complemented with a plasmid expressing ipaH7.8. IpaH7.8-dependent cell death was markedly reduced in cells lacking CASP1 or NLRP1B (Fig. 4F) (48). The NLRC4 inflammasome also recognizes S. flexneri (4951). In Nlrc4–/– RAW cells, inflammasome activation was almost entirely IpaH7.8 dependent (Fig. 4F). Although the N-end rule ubiquitin ligase Ubr2 is required for LF-mediated NLRP1B activation (32, 52), as expected, this requirement was circumvented by the direct ubiquitylation of NLRP1B by S. flexneri (fig. S6). Immortalized 129 macrophages were also sensitive to IpaH7.8-dependent killing, which correlated with decreased levels of endogenous NLRP1B and the induction of CASP1 maturation (Fig. 4G). S. flexneri is not a natural pathogen of mice; this may be partly due to species-specific NLRP1B effector recognition, as human NLRP1 does not appear to detect IpaH7.8 (fig. S6D). Thus, a mechanistic understanding of NLRP1B has led us to identify ubiquitin ligases as an additional category of pathogen-encoded enzymes that activate NLRP1B.

Discussion

Prior work in Arabidopsis has shown that an NLR called RPS5 detects proteolytic cleavage of the host PBS1 kinase by the translocated Pseudomonas syringae effector AvrPphB (53). In this system, RPS5 appears to detect the cleavage products of PBS1 (54). Although NLRP1B also appears to detect a pathogen-encoded protease, our results suggest that the underlying mechanism is very different from that of RPS5. Instead of proteolysis generating a specific ligand, it appears that NLRP1B is itself the target of proteolysis, leading to its proteasomal degradation and release of a functional inflammasome fragment. Thus, NLRP1B is in essence a sensor of its own stability, permitting detection of diverse pathogen-encoded enzymes, potentially including those of viruses or parasites (5557). Although IpaH7.8 and LF protease both activate NLRP1B, we favor a scenario in which the intended targets of these pathogen-encoded enzymes are other host proteins and NLRP1B has evolved as a decoy target of these proteins. Although decoy sensors are widely deployed in plant immunity (58), evidence for such sensors in animals has been scant.

NLRP1 was the first protein shown to form an inflammasome (11). Our results provide a possible mechanism explaining how NLRP1 is activated. Our proposed mechanism may also apply to other FIIND-death domain fold containing proteins, including PIDD1 and CARD8 (19). In addition to explaining how NLRP1 senses pathogens, functional degradation likely provides an explanation for why naturally occurring mutations that destabilize the N-terminal pyrin domain in human NLRP1 (27) also result in its activation. It has previously been suggested that NLRP1B is activated upon adenosine triphosphate (ATP) depletion in cells (39, 46). Our results are not incongruous with this model; indeed, ATP depletion may also indirectly affect NLRP1B stability. Thus, our results may provide a foundation for identifying pathogen-encoded activators of human NLRP1 and offer a conceptual basis for designing therapeutic interventions targeting NLRP1.

Materials and methods

Plasmids and constructs

The coding sequence of Nlrp1b allele 1: 129S1/SvimJ (129) Nlrp1b (DQ117584.1), allele 2: C57BL/6J (B6) Nlrp1b (BC141354) or allele 3: AKR/J, and variants thereof, were cloned into either pCMSCV-IRES-GFP or pAcSG2, with an N-terminal maltose binding protein (MBP) tag followed by a 3C protease cleavage site and a C-terminal HA or FLAG tag, respectively, or into pQCXIP with N-terminal green fluorescent protein (GFP) and C-terminal HA tags. The Fla-NLRP1B hybrid was constructed by replacing the first 45 N-terminal amino acids of NLRP1B with residues 431 to 475 of Legionella pneumophila flagellin (ANN95373) followed by the TEV cleavage sequence. CASP1, IL-1β, TEV, and LF producing constructs were described previously (16). The Nlrp1b coding sequence was subcloned in-frame with AID-GFP (Addgene 80076). GFP-fused IpaH producing plasmids were constructed as follows: the ipaH coding sequences from the S. flexneri 2a str. 2457T virulence plasmid were transferred using the Gateway vector conversion system (ThermoFisher) from Gateway entry clones (59) into the SmaI restriction site of the Gateway-compatible destination vector pC1-eGFP (Clontech) via LR reactions. The ipaH7.8 coding sequence was also subcloned into pQCXIP with an N-terminal mCherry tag. For protein expression in Escherichia coli, the ipaH coding sequences were subcloned into pET28a with a C-terminal 6X-HIS tag. Mutations were engineered by overlapping PCR.

Cell culture

293T and RAW264.7 cells were grown in DMEM supplemented with 10% FBS, 100 U/ml penicillin, 100 mg/ml streptomycin, and 2 mM l-glutamine. Primary BMDMs were cultured in RPMI supplemented with 5% FBS, 5% mCSF, 100 U/ml penicillin, 100 mg/ml streptomycin, and 2 mM l-glutamine. BMDM immortalization was performed as previously described (60).

Bacterial strains and infections

2457T S. flexneri–derived ipaH deletion strains were constructed using the λ red recombinase–mediated recombination system (61), as previously described (62). To construct complemented strains, the coding sequence of ipaH7.8 and 407 base pairs upstream, representing the endogenous promoter, were Gateway cloned into the pCMD136 plasmid and transformed into the ΔipaH7.8 mutant strain. S. flexneri was grown at 37°C on tryptic soy agar plates containing 0.01% Congo red, supplemented with 100 μg/ml spectinomycin for growth of complemented strains. For infections, 5 ml of tryptic soy broth (TSB) was inoculated with a single Congo red-positive colony and grown overnight shaking at 37°C. Saturated cultures were back-diluted 1:100 in 5 ml of fresh TSB and incubated for 2 to 3 hours shaking at 37°C. Bacteria were washed in cell culture medium and spun onto cells for 10 min at 300×g. Infected cells were incubated at 37°C for 20 min and then washed twice with cell culture medium containing 25 μg/ml gentamicin, then returned to 37°C for further incubation (30 min to 2 hours). Cells were infected at an MOI of 30 unless otherwise specified. Cell death was assessed by LDH activity in clarified culture supernatants as previously described (63). Protein in supernatants was TCA precipitated for anti-CASP1 immunoblotting.

Reconstituted NLRP1B activity assays

To reconstitute inflammasome activity in 293T cells, constructs producing NLRP1B (or mutants), CASP1, and IL-1β were cotransfected with constructs producing TEV, LF, IpaHs, or empty vector (MSCV2.2 or pcDNA3) using Lipofectamine 2000 (Invitrogen) following the manufacturer’s protocol. For experiments using recombinant proteins, fresh media containing 10 μg/ml PA and 2.5 μg/ml LF, supplemented with or without 10 μM MG132, 1 μM bortezomib, or 0.5 μM NMS-873, was added to cells for 2 to 4 hours. For auxin-inducible degradation, AID-NLRP1B and TIR1-producing constructs (TIR1, Addgene 80073) were cotransfected and treated with 500 μM indole-3-acetic acid sodium salt (IAA) (Sigma) for 3 to 6 hours in the presence or absence of 10 μM MG132. In all experiments, cells were lysed in RIPA buffer with protease inhibitor cocktail (Roche) 24 hours posttransfection.

Endogenous NLRP1B activity assays

Immortalized 129 (i129) BMDMs (2.5 × 106) were plated in six-well plates. Two hours before challenge, cells were primed with 1.0 μg/ml Pam3CSK4 (Invivogen). Cells were washed with PBS and media was replaced with 0.5 ml of Opti-MEM (Gibco) with or without 20 μg/ml PA, 10 μg/ml LF, and/or 10 μM MG132. Cells and media were lysed by addition of 120 μl of 10X RIPA buffer with protease inhibitor cocktail 2.5 hours posttreatment.

Immunoblotting and antibodies

Lysates were clarified by spinning at ~16,000×g for 10 min at 4°C. Clarified lysates were denatured in SDS loading buffer. Samples were separated on NuPAGE Bis–Tris 4% to 12% gradient gels (ThermoFisher) following the manufacturer’s protocol. Gels were transferred onto Immobilon-FL PVDF membranes at 35 V for 90 min and blocked with Odyssey blocking buffer (Li-Cor). Proteins were detected on a Li-Cor Odyssey Blot Imager using the following primary and secondary antibodies: 100 ng/ml anti-HA clone 3F10 (Sigma), 200 ng/ml anti-IL-1β (R&D systems, AF-401-NA), 1 μg/ml anti-GFP (Clontech, JL8), 2 μg/ml anti-mCherry (ThermoFisher, 16D7), 1 μg/ml anti-CASP1 (Adipogen, AG-20B-0042-C100). Anti-MBP (NEB, E8032S) and anti-ubiquitin (Cell Signaling, P4D1) antibodies were used at 1:1,000 dilution of manufacturer’s stock. Alexa Fluor 680-conjugated secondary antibodies (Invitrogen) were used at 0.4 μg/ml. Band intensities were quantified with Image Studio Lite software v5.2.5. The complete gel images are shown in fig. S7.

To produce the 2A12 mouse anti-NLRP1B monoclonal antibody, the pAcSG2-Nlrp1b construct was cotransfected with BestBac linearized baculovirus DNA (Expression Systems) into SF9 cells following the manufacturer’s protocol to generate NLRP1B expressing baculovirus. Primary virus was amplified in SF9 cells. NLRP1B was produced by infecting 4 liters of High Five cells with 1 ml of amplified virus per 1 liter of cells. Cells were harvested 48 hours after infection by centrifugation at 300×g for 15 min. Cell pellets were resuspended in lysis buffer (50 mM Hepes pH 7.5, 150 mM NaCl, 1% NP-40, and 5% glycerol) and lysed on ice using a dounce homogenizer. Homogenized samples were clarified at 24,000×g for 30 min, and supernatants were batch bound to 1 ml of amylose resin for 2 hours at 4°C. Samples were column purified by gravity. Resin was washed with 50 ml of wash buffer (20 mM Hepes pH 7.4, 150 mM NaCl, 0.02% NP-50, 5% glycerol). Samples were eluted with 1 ml of elution buffer (20 mM Hepes pH 7.4, 150 mM NaCl, 0.02% NP-50, 5% glycerol, 20 mM maltose) fractions. Peak elutions were pooled and MBP was cleaved by treatment overnight with 3C protease. Free MBP was removed by passing the sample over amylose resin. BALB/c mice were immunized with 10 μg of NLRP1B in 100 μl of Sigma adjuvant on days 0, 21, and 42, and with 10 μg of NLRP1B without adjuvant on day 60. Mice were sacrificed on day 63. Splenocytes were fused the with the P3X63-Ag8.653 parental line. Clones were screened via ELISA against recombinant NLRP1B protein or recombinant FLAG-tagged MBP protein to identify clones specifically reactive to NLRP1B. Clarified supernatant from the hybridoma clone 2A12 was used for immunoblotting.

In vitro ubiquitylation assay

Recombinant 129 or B6 NLRP1B was produced in insect cells and purified as described above before 3C treatment. Recombinant IpaH7.8, IpaH7.8 C357A (catalytic mutant), and IpaH9.8 were expressed in BL21 E. coli. Cells (1 liter) were grown to ~0.7 OD600 and induced with 1 M IPTG (Sigma) for 4 hours at 37°C. Pellets were resuspended in 50 mM Tris pH 7.4, 150 mM NaCl, 1% NP-40 and sonicated to lyse. Samples were clarified at 24,000×g for 30 min. The NaCl concentration of the supernatants was increased to 400 mM and 20 mM imidazole pH8.0 was then added to samples. Supernatants were batch bound to 1 ml of Ni resin (Qiagen) at 4°C for 2 hours. Samples were purified by gravity, washed with 50 ml of 20 mM Tris pH 7.4, 400 mM NaCl, and 20 mM imidazole pH 8.0. Protein was eluted in 1-ml fractions of 20 mM Tris pH 7.4, 150 mM NaCl, and 250 mM imidazole pH 8.0. Elution peaks were pooled and desalted into 20 mM HEPES pH 7.4, 150 mM NaCl, and 2 mM DTT.

In vitro ubiquitylation assays were performed in 25 mM Tris pH 7.4, 50 mM NaCl, 10 mM MgCl2, 5 mM ATP, 0.1 mM DTT with 60 nM Ubiquitin E1 (Boston Biochemistry), 200 nM UbcH5c (Boston Biochemistry), 10 μM Ubiquitin (Boston Biochemistry), 300 nM IpaH, and 270 nM NLRP1B. The reaction was run for 1 hour at 37°C. Solutions were then batch bound to anti-FLAG M2 agarose gel (Sigma) at 4°C for 2 hours. Bound samples were column-purified and washed with 5 ml of HBS (20 mM Hepes, 150 mM NaCl). Final samples were eluted in 150 μl HBS+150 μg/ml of FLAG peptide.

Native gel oligomerization assay

Samples were transfected into 293T cells with constructs as above in a six-well plate. After 24 hours, samples were harvested by removing media and washing cells off plate with cold PBS. Harvested cells were centrifuged at 300×g for 10 min at 4°C. Cells were lysed in lysis buffer (50 mM Hepes pH 7.5, 150 mM NaCl, 1% NP-40, 5% glycerol), and samples were clarified by spinning at 16,000×g for 10 min at 4°C. Samples were run on NativePAGE Bis–Tris gels (ThermoFisher) according to manufacturer’s protocols.

Detection of UPA-CARD and ASC speck formation by immunofluorescence

293T cells were grown on fibronectin-coated coverslips. Constructs producing NLRP1B and ASC were cotransfected with constructs producing TEV or an empty vector. The NLRP1B construct was designed with a C-terminal FLAG and N-terminal HA tag, where the HA sequence was inserted following the P1′ position of the TEV cleavage site. Twenty hours posttransfection, cells were fixed with 4% PFA/PBS (20 min) and permeabilized in 0.5% saponin in PBS (5 min). Blocking and antibody staining was performed at room temperature in 5% BSA/0.1% saponin in PBS. Primary antibodies: 0.5 μg/ml of rabbit anti-ASC (Santa Cruz, N-15), 0.5 μg/ml of rat anti-HA (Roche, 3F10), and 1 μg/ml of mouse anti-FLAG (Sigma, M2). Secondary antibodies: 3 μg/ml of AMCA-labeled goat anti-rabbit IgG (Jackson Laboratories), 4 μg/ml of Alexa Fluor 647-labeled goat anti-rat IgG, and 4 μg/ml of Alexa Fluor 555-labeled goat anti-mouse (Molecular Probes). Coverslips were mounted onto slides using Vectashield medium (Vector Laboratories, Inc., H-1000), and imaged on a ZEISS LSM 710 with a W Plan-Apochromat 40×/1.0 DIC oil immersion objective. Fluorophores were excitated at 405, 543, and 633 nM. To quantify the presence of FLAG or HA per ASC speck, the Imaris imaging software (Bitplane) was used to first identify ASC specks as objects, then to count FLAG- and/or HA-positive objects based on a fluorescence intensity threshold set manually for each channel. Thresholds were determined manually using a training set of samples and controls (i.e., no primary antibody) and were applied in batch to all samples.

Supplementary Materials

References and Notes

Acknowledgments: We are grateful to J. Chavarría-Smith for discussions and for laying the experimental foundations for this work. We thank P. R. Beatty and UC Berkeley undergraduates in the MCB150L course for help with generating the 2A12 monoclonal antibody; the Rape laboratory for guidance with ubiquitylation assays; J. Mogridge for the AKR/J Nlrp1b allele construct (21); A. Holland for the AID and TIR1 constructs; the Bachovchin laboratory for the HEK and RAW cell lines and for sharing results before submission; G. Barton, J. Chavarría-Smith, H. Darwin, M. Dorrington, and J. Tenthorey for comments on the manuscript; and members of the Vance and Barton laboratories for discussions. Funding: R.E.V. is an HHMI Investigator and is supported by NIH AI075039 and AI063302. P.S.M. is supported by a Jane Coffin Childs Memorial Fund postdoctoral fellowship. C.F.L. is a Brit d’Arbeloff MGH Research Scholar and is supported by NIH AI064285. Author contributions: Conceptualization, A.S., P.S.M., and R.E.V.; Methodology, A.S., P.S.M., and R.E.V.; Investigation, A.S., P.S.M., L.G., and E.W.M.; Resources, A.S., P.S.M., L.G., C.F.L., and R.E.V; Writing – Original Draft, A.S., P.S.M., and R.E.V.; Writing – Review & Editing, A.S., P.S.M., C.F.L., and R.E.V.; Visualization, A.S. and P.S.M.; Supervision, A.S., P.S.M., C.F.L., and R.E.V. Competing interests: A patent related to this work has been submitted by R.E.V., A.S., and P.S.M. R.E.V. is a scientific advisory board member for Metchnikoff Therapeutics, Inc. Data and materials availability: All data are available in the main text or the supplementary materials.
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