DED or alive: assembly and regulation of the death effector domain complexes

Abstract

Death effector domains (DEDs) are protein–protein interaction domains initially identified in proteins such as FADD, FLIP and caspase-8 involved in regulating apoptosis. Subsequently, these proteins have been shown to have important roles in regulating other forms of cell death, including necroptosis, and in regulating other important cellular processes, including autophagy and inflammation. Moreover, these proteins also have prominent roles in innate and adaptive immunity and during embryonic development. In this article, we review the various roles of DED-containing proteins and discuss recent developments in our understanding of DED complex formation and regulation. We also briefly discuss opportunities to therapeutically target DED complex formation in diseases such as cancer.

Facts

• FADD, FLIP, procaspase-8 and procaspase-10 all contain death effector domains (DEDs).

• The DED is a conserved protein sub-domain that mediates important protein–protein interactions.

• DED-containing proteins form a variety of complexes that regulate key cellular processes, most notably apoptosis, necroptosis and autophagy.

• Recent reports also highlight the critical role of DED proteins in other key processes linked to development and inflammation.

Open Questions

• Does caspase-10 (absent in mice) have overlap** functions with caspase-8, or is it functionally distinct?

• Under what physiologically relevant conditions does necroptosis occur rather than apoptosis?

• In which cellular contexts are FADD, FLIP, (pro)caspase-8 and (pro)caspase-10 critical for regulating autophagy?

• What are the best ways of targeting DED-containing proteins to therapeutically activate cell death (e.g., in cancers) or prevent cell death (e.g., in neurodegenerative diseases)?

• Are DED-containing proteins potential therapeutic targets for inflammatory diseases?

Cell death is critical for maintaining homeostasis in multicellular organisms; too much can result in pathologies such as neurodegeneration, whereas too little can lead to the accumulation of malignant cancerous cells. Cell death can be either active, where the cell participates in its own destruction or passive, for example, when a cell undergoes irreparable physical damage.¹ The most biochemically well-characterised form of cell death is apoptosis, an active process in which cysteine-dependent aspartate-directed proteases (caspases) are activated in response to extracellular stimuli or internal damage culminating in a form of cell death defined by distinct molecular events and characteristic changes in the morphology of the dying cell. Recently, a number of actively regulated non-apoptotic mechanisms of cell death have emerged, including necroptosis, pyroptosis and ferroptosis, which have been comprehensively reviewed elsewhere.^{2, 3} Here, we focus on those mechanisms of cell death arising following stimulation of death receptors, broadly termed the ‘extrinsic pathway’. For authoritative reviews on mitochondrial-mediated ‘intrinsic’ cell death, we direct the Reader elsewhere.^{4, 5} Central to receptor-mediated cell death pathways are proteins containing ‘death–fold superfamily’ interaction motifs such as the death domain (DD), caspase activation and recruitment domain (CARD), pyrin domain and the death effector domain (DED).

The DED-containing proteins, which are key decision makers in determining the life and death of cells, are the primary focus of this review. We will first introduce the main members of the DED protein family and discuss advances in the understanding of the assembly and stoichiometry of death receptor complexes. We will then summarise the recent literature surrounding the regulation of these complexes and consider the role of these proteins in disease.

The DED Proteins

The death-fold motif is characterised by its globular structure containing six amphipathic α-helices that run anti-parallel in α-helical bundles.⁶ When folded, a conserved hydrophobic core forms, although differences in helical length and residue distribution give rise to significant variations between the different sub-families.^{6, 7, 8, 9} The DED death-fold sub-family consists of procaspases-8 and -10, FLIP, FADD, DEDD, DEDD2, and PEA-15 (Figure 1). FADD, DEDD, DEDD2 and PEA-15 contain a single DED, whereas FLIP, procaspase-8 and procaspase-10 each have tandem DEDs. Procaspases-8 and -10 have catalytically active domains in the regions C-terminal to their tandem DEDs; whereas the long splice form of FLIP, FLIP_L, has a pseudo-caspase domain C- terminal to its tandem DEDs, in which the cysteine residue critical for enzymatic activity is absent. Shorter splice forms of FLIP (FLIP short, FLIP_S, and FLIP Raji, FLIP_R) arising through alternative mRNA splicing lack the pseudo-caspase domain, but contain the tandem DEDs.¹⁰

Figure 1

Schematic representations of the structures of DED proteins (ribbon format with transparent solvent-accessible surface area) along with their complete linear domain organisation. Protein structures: (a) Homology model of human FLIP DEDs⁴⁸ (b) NMR solution structure of FADD DED (PDB ID: 2GF5) ⁷ (c) NMR solution structure of PEA-15 DED (PDB ID: 2LS7)¹⁹¹ (d) Homology model of procaspase-8 DEDs⁴⁸ (e) Homology model of procaspase-10 DEDs generated using the I-TASSER web server¹⁹²

Procaspase-8 is a highly conserved protease, displaying ~20% sequence similarity to its Caenorhabditis elegans homolog CED-3.¹¹ Eight splice forms of procaspase-8 have been identified at the mRNA level, although only two of these, procaspases-8A and 8B are expressed as functional proteases.¹² Additionally, a long splice form (procaspase-8L), which contains a 136-bp insert between exons 8 and 9 encoding an early stop codon, contains both DEDs but lacks a functional catalytic domain; it is found in undifferentiated cells and neoplasms and has been reported to act in a dominant negative manner to inhibit apoptosis.^{13, 14, 15}

Procaspase-10 is also expressed as multiple splice forms: procaspases-10A, B, D and G. All contain tandem DEDs and proteolytic domains except G, which is truncated and only consists of the DEDs.^{16, 17, 18} In vitro studies have shown that procaspase-10 is activated by induced proximity in a manner similar to procaspase-8.¹⁹ Despite the similarity between procaspase-10 and procaspase-8, whether procaspase-10 can initiate death receptor-mediated apoptosis in the absence of procaspase-8 remains controversial,^{20, 21, 22} although they share common substrates, notably BID and RIPK1.^{23, 24} However, no ortholog of the gene encoding procaspase-10 (Casp10) is present in the mouse genome,²⁵ suggesting that procaspase-10 is not required for activation of the extrinsic apoptotic pathway. Despite this, expression of both procaspase-8 and procaspase-10 is frequently downregulated in cancer.^{26, 27, 28} Interestingly, the genes encoding procaspase-8 (Casp8), procaspase-10 and FLIP (CFLAR) are present at the same loci (2q33-q34) and clearly evolved via gene duplication events. Evolutionary studies have identified the predecessors of Casp8, Casp10 and CFLAR in fish.²⁹

FADD is an adaptor protein containing a DD which allows it to associate with the DDs of TRAIL- R1, TRAIL-R2, CD95 and TRADD, and a DED, which enables it to recruit other DED-containing proteins, namely procaspase-8, procaspase-10 or FLIP. As with most other components of the extracellular apoptosis signalling pathways, FADD is highly evolutionarily conserved.³⁰ As a protein linking death receptors to death initiators, it is not only a key player in cell death, but also has reported roles in non-apoptotic processes. For example, FADD has been identified in the nucleus and has been postulated to have functions in regulating cell cycle progression,^{31, 32} NF-κB activity,³³ autophagy,³⁴ cytokine signalling^{35, 36} and T-cell activation.³⁷ Indeed, roles beyond core apoptosis signalling have also been identified for FLIP, procaspase-8 and procaspase-10, and frequently these roles involve complex interactions between these proteins and FADD.

Assembly of the Death-inducing Signalling Complex (DISC)

The death receptors TRAIL-R1 (DR4), TRAIL-R2 (DR5) and CD95 (Fas) are specialised members of the TNF receptor superfamily and are key mediators of apoptosis triggered by ligands expressed by cells of the immune system, namely TRAIL (TNF-related apoptosis-inducing ligand), which activates TRAIL-R1 and TRAIL-R2, and CD95L (FasL), which activates CD95. Following extracellular ligand binding, pre-associated TRAIL-R1, TRAIL-R2 and CD95 trimers, cluster through interactions between their intracellular DDs.^{38, 39} The DD of FADD can then interact with the cytoplasmic DDs of the death receptors, after which its DED becomes available for protein–protein interactions with other DED proteins, thereby creating a platform for assembly of the DISC.

Death domain interactions

The interactions between the CD95 and FADD DDs have been described (Figure 2). Scott et al.⁴⁰ reported a 2.7 Å resolution co-crystal structure, which suggests that CD95 and FADD bind in dimeric units, that is 2 × FADD-DD interacting with 2 × CD95-DD. These units are then proposed to further associate into tetrameric structures (4 × FADD-DD to 4 × CD95-DD), although in vivo, the dimeric form is favoured. They report that, following CD95 receptor activation, CD95 undergoes a conformational change, exposing its hydrophobic core and revealing a multitude of interaction surfaces capable of binding the DD of FADD. This represents a possible safety mechanism whereby the apoptotic cascade only proceeds when sufficient CD95 DDs cluster. However, the FADD interaction sites in the CD95 DD predicted by this model do not correlate with mutations observed in patients with autoimmune lymphoproliferative syndrome, a disease defined by mutations in CD95 which prevent DISC formation.⁴¹ In a different study, Wang et al.⁴² used electron microscopy to visualise CD95 DD–FADD DD interactions and observed that they bore a striking resemblance to the PIDD DD–RAIDD DD complex, being principally composed of 5 × CD95 DDs and 5 × FADD DDs layered together. This stoichiometry is in agreement with data by Esposito et al.,⁴³ who also reported a ratio of 5 × CD95:5 × FADD together with some 6 × CD95:5 × FADD and 7 × CD95:5 × FADD ratios, but not the 4 × CD95:4 × FADD suggested by Scott et al. Disparities between these models could be explained by the different conditions used for protein crystallisation; however, the models proposed by Wang et al. and Esposito et al. are supported by the fact that they account for mutations frequently seen in autoimmune lymphoproliferative syndrome. Most disease-causing mutations present in autoimmune lymphoproliferative syndrome patients reside in the DD of CD95,⁴¹ resulting in an inability to bind FADD and form a DISC. Map** these mutated residues onto the structure proposed by Wang et al. reveals that they reside on the exposed surface of the DD and would be likely to prevent the binding of FADD.⁴²

Figure 2

Crystal structure of the human CD95–FADD DD heterodimer. CD95 DD (blue ribbon) binds to FADD DD (green ribbon) via several key interacting residues (highlighted as grey sticks) (PDB ID: 3EZQ).⁴⁰ Hydrogen bonds are depicted as black dotted lines. These heterodimers oligomerise into higher order structures

DED interactions

In addition to death receptor:FADD DD interactions, FADD has been reported to self-associate through its DED, which stabilises its association with the death receptor. Sandu et al.⁴⁴ identified a ‘hydrophobic patch’ (F25, L28 and K33) as the critical surface for FADD-FADD interactions and not an RxDL motif as had previously been reported. Such interactions between FADD molecules generates higher order complexes of FADD and death receptors that may be the basis of the SPOTS (signaling protein oligomerisation transduction structures), which have been reported to form soon after CD95 receptor activation.⁴⁵ The RxDL motif is found in both DEDs of viral FLIP MC159 and is critical for its ability to be recruited to the DISC and inhibit apoptosis.^{9, 46, 47} However, this appears not to be the case for murine FLIP, for which the hydrophobic patch was instead found to be indispensable for DISC recruitment and apoptosis inhibition.⁴⁷ MC159 was also reported to interact with the RxDL motif of FADD, blocking FADD self-association and preventing the formation of a competent caspase-recruiting platform.⁸ The role of the RxDL motif may differ for cellular and viral forms of FLIP. Our recent data suggest that this motif is important for human FLIP’s anti-apoptotic function; however, not because of its direct involvement in mediating inter- and intra-molecular interactions, but rather because it controls the spatial orientation of the hydrophobic patch defined by the α2 and α5 helices of FLIP’s DEDs (Figure 3); our data and those of others indicate that it is this hydrophobic patch that mediates intra-molecular interactions between FLIP’s tandem DEDs and inter-molecular interactions between FLIP and FADD and procaspase-8 (Figure 4).⁴⁸

Figure 3

Conserved features of the DED protein family. (a) Homology model of the tandem DEDs of human FLIP (orange ribbon) depicting the main conserved residues part of the hydrophobic patch (highlighted in grey), which are essential for inter- and intra-DED–DED interactions. (b) Sequence alignment of residues that form part of the hydrophobic patches, which are highly conserved amongst DED proteins. (c) Homology model of human FLIP DEDs (orange ribbon) with a close-up view of the two E/D-RxDL motifs. Only residues that form the charged triad motifs (highlighted in cyan) are shown. Hydrogen bonds are highlighted in black dotted lines. (d) Sequence alignment of the E/D-RxDL motifs which are conserved amongst DED proteins

Figure 4

Binding modes of FLIP–FADD and procaspase 8–FADD interactions. FLIP binds to the DED of FADD using its DED2, whereas procaspase-8 binds to FADD using its DED1. The main residues that are important for the interactions are shown in grey: FLIP uses its F114 residue on the α2 helix to bind into a groove between α1/α4 helices in the DED of FADD with a reciprocal interaction from FADD H9 into the α2/α5 hydrophobic patch of FLIP; procaspase-8 uses the Y8 residue on its α1 helix to bind into the hydrophobic patch between the α2/α5 helices in FADD, with a reciprocal interaction from FADD F25 into the α1/α4 groove in procaspase-8

Emerging models of DISC assembly at the level of DED interactions

Two independent studies proposed a novel model of DISC assembly after finding that FADD is sub-stoichiometric at the DISC compared with death receptors and caspase-8, with three to five receptors and as many as nine caspase-8 molecules for every FADD molecule recruited to the complex.^{49, 50} DED-containing proteins interact with themselves and one another in a homotypic manner through their DEDs, so both groups proposed that one FADD molecule could recruit multiple DED-only proteins (procaspase-8, procaspase-10 or FLIP) as ‘chains’. In support of this model, formation of caspase-8 chains was observed in single cell studies using fluorescently tagged caspase-8. Such DED ‘filaments’ have been described before for caspase-8 and FADD;^{46, 51} however, their physiological relevance is questionable as they are generated in cells expressing supra-physiological levels of each DED protein.

By creating models of FLIP and procaspase-8 DEDs based on the published structure of vFLIP MC159,⁸ we used the NMR structure of FADD⁷ to perform docking experiments between the DEDs of the three proteins. These modelling experiments suggested that each protein pair could potentially interact in two distinct orientations, which involved mainly hydrophobic interactions between the α2/α5 surface (the aforementioned hydrophobic patch) in one DED and α1/α4 surface of the adjacent DED.⁴⁸ Subsequent mutagenesis studies revealed that FLIP and procaspase-8 have differential affinities for the two available interaction surfaces of the FADD DED. FLIP preferentially binds to the α1/α4 surface of FADD’s DED, whereas procaspase-8 binds to FADD’s α2/α5 surface (Figure 4). Our analysis of the stoichiometry of the TRAIL-R2 DISC was not in agreement with the caspase chain models described above: in our study, sub-apoptotic DISC stimulation resulted in an approximate 1:1:1 ratio of FADD:caspase-8:FLIP; while at higher levels of DISC stimulation, there was more caspase-8 than FADD or FLIP, although there remained approximately one FADD molecule for every two molecules of FLIP/caspase-8.

Activation of caspase-8

The current model of procaspase-8 activation is that 53/55 kDa procaspase-8 zymogens are recruited to FADD as monomers via their DEDs leading to dimerisation of the procaspases, initially via their DEDs (Figure 5).^{52, 53, 54, 55, 56} Dimerisation of the caspase domains then occurs and results in conformational changes that reveal the enzymatic activity necessary for intra-molecular cleavage of the C-terminal portion of the caspase, liberating a p12 subunit (subsequently processed to the small p10 catalytic subunit) and simultaneously stabilising the dimer. Next, the 41/43 kDa caspase-8 intermediates in the dimer cleave one another in a trans-catalytic manner in the region between their DEDs and the large p18 catalytic subunit. The two molecules of p18-caspase-8 that are subsequently released associate with the two p10 subunits to form the active protease.⁵⁷ These two steps are critical, as cleavage in the absence of dimerisation does not result in an active protease.⁵⁸

Figure 5

Processing of procaspase-8. When two procaspase-8 molecules are co-recruited to the DISC via their DEDs, their caspase domains undergo conformational changes that exposes the enzymatic activity necessary for cleavage of the C-terminal portion of the caspase, liberating a p12 subunit, which is subsequently processed to the small p10 catalytic subunit. This initial processing step may occur in an inter-dimer manner between adjacent procaspase-8 dimers rather than the intra-dimer manner depicted.⁶³ The 41/43 kDa caspase-8 intermediates cleave one another in a trans-catalytic manner in the region between their DEDs and the large p18 catalytic subunit. The two molecules of pro-caspase-8 that are subsequently released associate with the two p10 subunits to form the active protease.⁵⁷ At lower levels of DISC stimulation or when FLIP is highly expressed, FLIP/caspase-8 heterodimers assemble at the DISC via interactions between their DEDs and those of FADD.⁴⁸ The pseudo-caspase domain of FLIP_L is able to induce the conformational change in procaspase-8’s caspase domain that is necessary to create its active site.⁵⁹ The FLIP_L:caspase-8 heterodimer is processed between the p18 and p12 subunits of both proteins, but is unable to be further processed owing to FLIP_L’s lack of enzymatic activity, and this heterodimer is unable to activate apoptosis. In the case of FLIP_S, heterodimerisation fails to activate procaspase-8 as the initial conformational change cannot take place in procaspase-8’s caspase domain

Procaspase-8 can also heterodimerise with FLIP at the DISC. In the case of FLIP_S/R, heterodimerisation fails to activate procaspase-8 as the initial conformational change cannot take place in procaspase-8’s caspase domain;⁵⁹ thus, FLIP_S/R effectively acts in a dominant negative manner (Figure 5). For FLIP_L, heterodimerisation results in an active enzyme, as the pseudo-catalytic domain of FLIP_L is able to induce the conformational change in procaspase-8’s caspase domain that is necessary to create the active site (Figure 5).⁵⁹ Indeed, it appears that FLIP_L’s pseudo-caspase domain is more efficient at inducing the conformational change in the dimer than the caspase domain of another molecule of procaspase-8.⁶⁰ The FLIP:caspase-8 heterodimer remains tethered to the DISC because the second step of activation, cleavage between the DEDs and p18-subunits cannot occur because of FLIP_L’s lack of enzymatic activity, and the heterodimer is unable to activate the apoptotic cascade. However, the FLIP_L:caspase-8 heterodimer’s enzymatic activity can cleave local substrates, most notably RIPK1, an important regulator of necroptosis (see below).⁶¹

Procaspase-8 dimerisation is required for formation of the active site.^{55, 56} However, as well as intra-dimer cleavage, procaspase-8 can be cleaved in an inter-dimeric manner (i.e., dimers acting in a trans manner).^{54, 115} a protein known to bind to ubiquitin, that moves caspase-8 into ubiquitin-rich foci.¹¹⁴ However, it is unknown whether translocation into these foci is necessary for caspase-8′s activation, and the implications this has for the observation that caspase-8 and p62 are both found in intracellular DISCs and autophagosomal membranes is unclear.^{103, 104} In subsequent work, the same group found that cytosolic p43 and p18 fragments of caspase-8 are degraded in a proteasome-dependent manner. In this latter study, caspase-8 was reported to be conjugated by degradative K48-linked ubiquitin chains on its p18 fragment by another E3 ligase, TRAF2.¹¹⁶ Rather than increasing its activation as is the case for CUL3-mediated K63-ubiquitination, TRAF2 acts to degrade the pool of activated caspase-8, decreasing the propensity of the cell to commit to apoptosis. Caspase-8-processed FLIP_L has also been reported to interact with TRAF2, promoting activation of the NF-κB transcriptional pathway, but it is not yet clear whether these observations are related.¹¹⁷

Regulation of FLIP by the UPS

An additional layer of control of death receptor-mediated apoptosis is achieved by the ubiquitination of FLIP. Similar to the anti-apoptotic BCL-2 family member MCL-1, FLIP_S is an extremely short-lived protein that is rapidly turned over through the ubiquitin-proteasome system.¹¹⁸ FLIP_S is ubiquitinated on K192 and K195 in DED2 and, between these two lysine residues at position 193 is a serine residue which, when phosphorylated, inhibits the ubiquitination of the adjacent lysines.^{118, 119} Notably, mutational studies showed that ubiquitin-deficient mutants of FLIP_S had increased half-lives (as expected) but were still recruited to the DISC, retaining their anti-apoptotic ability. In further work, the same lab identified PKCα/β as the key mediators of FLIP_S phosphorylation on S193.¹¹⁹ FLIP_L is less labile than FLIP_S, although it too is turned over relatively rapidly, with a typical half-life of 2–3 h.¹¹⁸ A similar interplay between phosphorylation and ubiquitination is true for FLIP_L, where ROS production induces the phosphorylation and subsequent ubiquitination and degradation of FLIP_L.¹²⁰ Moreover, K195 is a site of ubiquitin conjugation on FLIP_L in response to hyperthermia.¹²¹ Our group has identified a role for the DNA repair protein Ku70 in regulation of FLIP ubiquitination. Ku70 forms a complex with FLIP protecting it from ubiquitination and subsequent degradation. This complex is regulated by the acetylation of Ku70 and thus can be manipulated pharmacologically by histone deacetylase inhibitors leading to rapid degradation of FLIP.¹²² Chang et al.¹²³ reported that following TNFα stimulation, JNK is activated which in turn activates the E3 Ubiquitin ligase Itch, resulting in FLIP_L ubiquitination and subsequent proteasomal degradation. This apparent link between FLIP and Itch was partly confirmed by other studies, including Panner et al.,^{124, 125} who identified a PTEN-Akt-Itch pathway controlling FLIP_S stability and degradation.

DED Complexes in Mammalian Host Defence

Antiviral immunity

DED-containing protein complexes are critical for innate immune reactions and can assemble to induce apoptosis in response to viral infection, generally in a mitochondrial antiviral signalling adaptor (MAVS)-dependent manner. Typically, cytosolic viral RNA is recognised by one of the three RIG-1-like receptors (RLRs) retinoic acid-inducible gene I (RIG-1), melanoma differentiation-associated gene 5 (MDA5) or laboratory of genetics and physiology 2 (LGP2). RIG-1 and MDA5 undergo conformational changes upon binding to viral RNA, exposing an N-terminal CARD that interacts with and activates MAVS,^{126, 127} which in turn activates a type I interferon response.¹²⁶

FADD and RIPK1 can interact with MAVS (Figure 7),¹²⁸ and FADD- and RIPK1-deficient cells are hypersensitive to viral infection owing to an inability to induce the transcription of key antiviral genes.³⁵ Tschopp and co-workers elucidated the mechanistic basis of these findings and found that the antiviral RIG-1 signalling pathway bears a striking resemblance to the TNFR1 pathway.^{64, 129} They propose that following viral infection, a complex termed the ‘TRADDosome’ (composed of TRADD, RIPK1 and FADD) forms on the mitochondrial membrane via MAVS. In this MAVS-located TRADDosome, RIPK1 can be K63 ubiquitinated by TRAF2/3 and recruit NEMO resulting in activation of IKKα and IKKβ and subsequent phosphorylation and degradation of IκB leading to NF-κB activation. Finally, via FADD, the TRADDosome can recruit procaspases-8 and -10 and FLIP, potentially inducing cell death.¹²⁹

Figure 7

DED proteins in MAVS signaling. Following viral infection of cells, viral RNA is detected by CARD-containing RIG-1-like receptors, for example, RIG-1 and MDA5. RIG-1 binds the dsRNA, exposing its normally hidden CARD domains. K63 ubiquitin chains are conjugated to the CARDs, facilitating the assembly of a complex composed of four polyubiquitin chains and four RIG-1 molecules (not shown).^{200, 201} This in turn induces the formation of prion-like aggregates of MAVS, which strongly activate IRF3.^{202, 203} These MAVS aggregates form a platform which can recruit TRAF2, TRAF3 and TRAF6.²⁰³ TRADD also binds MAVS followed by TANK and TBK2, activating antiviral IRF3.^{129, 203} However, TRADD can also recruit RIPK1, FADD and caspase-8, a complex dubbed the ‘TRADDosome’. Caspase-8 cleaves RIPK1 and the resulting RIPK1 fragment can inhibit IRF3, ceasing the antiviral response.²⁰⁴ RIPK1 is conjugated by K63 ubiquitin chains inducing two distinct signaling pathways from the TRADDosome: firstly, NF-κB signaling through NEMO, IKKα and IKKβ, and secondly NEMO can interact with NAP1, TBK1 and IKKɛ to activate IRF3 or IRF7.^{205, 206, 207, 208} In addition to caspase-8, the TRADDosome can also recruit FLIP (not shown) and may under certain conditions, trigger cell death

DED proteins and the inflammasome

The inflammasome is a multi-protein oligomeric structure formed in macrophages and monocytes in response to inflammatory stimuli (Figure 8).¹³⁰ Inflammasomes are comprised of a stimulus-specific sensor protein belonging to either the NLR, AIM2 or ALR family, the adaptor protein ASC and the inactive zymogen procaspase-1.¹³¹ Formation of the inflammasome leads to activation of caspase-1, which processes pro-IL-1β (and IL-18) to its mature form.^{132, 133} Different inflammasomes assemble in response to distinct stimuli, for example, the NLRP3 inflammasome forms in response to a plethora of pathogens, including influenza A,^{134, 135} Klebsiella pneumoniae and Staphylococcus aureus, in addition to endogenous danger signals such as ATP, uric acid crystals, nigericin and hyaluronan.¹³¹ The NLRC4 inflammasome reacts to bacterial flagellin and PrgJ, and the AIM2 inflammasome detects foreign dsDNA.^{136, 137} RIG-1 senses RNA viruses and forms a signaling complex with ASC, activating an inflammasome.¹³⁸ Toll-like receptors (TLRs) detect pathogens by recognising pathogen membrane proteins (TLR4) or cytoplasmic nucleotides (TLR3). Full activation of inflammasomes requires two distinct stages: a ‘priming’ signal 1, for example, from either a TLR agonist or a pro-inflammatory cytokine, that activates NF-κB and upregulates pro-IL-1β expression,^{139, 140} followed by stimulus-specific inflammasome activation and processing of pro-IL-1β.^{141, 142}

Figure 8

Involvement of DED proteins in the inflammasome. (a) Full activation of the inflammasomes requires a two-step process. Firstly, a priming signal is induced by a PAMP (such as LPS) activating a transcriptional cascade resulting in the de novo synthesis of inflammasome components such as NLRP3 and pro-IL-1β.¹³⁹ Of note, subsequent work has shown that this is not always the case, as TLR-mediated priming of the NLRP3 inflammasome does not always require transcriptional upregulation of NLRP3.^{209, 210} (b) Assembly of the inflammasome components NLRP3, ASC and pro-caspase-1 occurs following a second signal. This can be any one of numerous different stimuli, for example, ATP activation of the P2X7 receptor, bacterial toxins, nigericin and silica.^{211, 212, 213} A subsequent efflux of potassium from the cell permits the components to form a functional canonical NLRP3 inflammasome where pro-caspase-1 is cleaved into its active form.²¹⁴ Catalytically active caspase-1 cleaves pro-IL-1β and pro-IL-18 into their mature forms, which are then released from the cell.²¹¹ (c) Fungi and mycobacterium activate the β-glucan receptor dectin-1 resulting in the phosphorylation of cytoplasmic domain allowing the recruitment of SYK kinase.²¹⁵ This elicits the formation of the non-canonical caspase-8 inflammasome, consisting of CARD9, BCL-10, MALT1, ASC and caspase-8. In this complex, active caspase-8 cleaves pro-IL-1β into its mature form which is released from the cell.²¹⁶ (d) IL-1β and IL-18 can be processed directly by caspase-8 in an ASC-independent manner following ligation of members of the TNF receptor family, such as CD95, although the mechanism for this remains unclear.¹⁴⁶ (e) The Ripoptosome forms upon loss of IAPs and can lead to activation of caspase-8, which can potentially cleave IL-1β directly or indirectly via caspase-1.^{150, 217}

DED-containing proteins are emerging as players in inflammasome signalling. Caspase-8-dependent TLR4 signalling is critical for inflammasome assembly and IL-1β processing in glaucoma,¹⁴³ and engagement of TLR3 or TLR4 can result in processing of IL-1β by caspase-8, independent of NLRP3 and caspase-1.¹⁴⁴ Studies by Kanneganti and colleagues¹⁴⁵ revealed that FADD and caspase-8 are obligatory for the correct priming and activation of both the canonical and non-canonical NLRP3 inflammasome, and that CD95 can induce IL-1β and IL-18 maturation in a caspase-8-dependent, but RIPK3-independent manner.¹⁴⁶ However, work from other groups contest this: Allam et al.¹⁴⁷ and Weng et al.¹⁴⁸ show that caspase-8 is required only for TLR-induced inflammasome priming. A number of other studies show normal canonical NLRP3 inflammasome activation in caspase-8-deficient cells, refuting an essential role for caspase-8 in NLRP3 inflammasome activation.^{149, 150, 151} Together, these data suggest that caspase-8 may promote caspase-1 activity under certain conditions, but is not absolutely required for NLRP3 inflammasome activation.

FLIP_L has been shown to be involved in activation of the NLRP3 and AIM2 inflammasome and directly interacts with NLRP3, AIM2 and procaspase-1.¹⁵² In contrast, FLIP decreased IL-1β generation in response to SMAC mimetics and CD95 receptor activation, indicating that its role in regulating the inflammasome is context-dependent.¹⁵²

DED proteins and the adaptive immune system

It has long been known that caspase-8- or FADD-deficient T cells do not proliferate in response to T-cell receptor activation;^{37, 153} it had previously been thought that this was due to a defective ability to activate NF-κB, but recent work has shown that it is due to induction of necroptosis.^{154, 155} Rescue of T cells deficient in either caspase-8 or FADD can be achieved by simultaneously deleting RIPK1¹⁵⁶ or by inhibiting RIPK1 pharmacologically with necrostatin-1¹⁰⁸; furthermore, FADD^−/−RIPK1^−/− and Casp8^−/−RIPK3^−/− T cells undergo normal rates of clonal expansion following viral stimulation.^{97, 156, 157} Stimulation of the T-cell receptor by antigens induces the formation of the CARMA1-BCL-10-MALT1 complex which activates NF-κB.^{158, 159, 160, 161} Downstream of this, a complex composed of FADD, caspase-8 and FLIP_L forms, which presumably prevents aberrant activation of RIPK1, blocking necroptosis and promoting cell survival and proliferation.¹⁶² TIPE2 (tumor necrosis factor-α-induced protein-8, TNFAIP8) is the newest member of the DED-containing protein family, identified as highly expressed in a murine model of spine inflammation.^{163, 164, 165} TIPE2^−/− mice develop spontaneous fatal inflammatory disease with concomitant elevated production of pro-inflammatory cytokines, suggesting a role for TIPE2 in the immune system and, more specifically, T cells.¹⁶⁴ Upon infection or immunisation, TIPE2^−/− mice exhibit increased levels of CD8+ T cells and inflammatory cytokine production, implying that TIPE2 is a negative regulator of T-cell-mediated immunity by impeding the NF-κB and AP1 transcriptional pathways and TLR signalling in macrophages.¹⁶⁴ Interestingly, Sun et al.¹⁶⁵ also found that TIPE2 interacts with caspase-8, but not FLIP, in macrophages, and blockade of caspase-8 function in TIPE2^−/− cells rescues the hypersensitive phenotype; however, subsequent papers have disputed this.

Therapeutically Exploiting DED Complexes

Evasion of apoptosis is a hallmark of cancer,¹⁶⁶ but aberrant cell death is also a feature of other human pathologies such as inflammation and neurodegenerative diseases. The DED family of proteins constitute key decision makers in these processes, with the ability to switch outcomes from life to death, or to different modes of death. As such, they represent an attractive set of proteins to target therapeutically.

Death receptors such as TRAIL-R1 and TRAIL-R2 are overexpressed in many types of cancer,¹⁶⁷ and there has been much effort to develop agents (recombinant forms of TRAIL and antibodies) that activate these receptors, particularly as they appear to selectively target malignant tissue while sparing normal cells.¹⁶⁸ Although pre-clinical data for TRAIL receptor-targeted therapeutics were promising and these agents were well tolerated in phase I trials, they showed limited anti-cancer effects in patients when used alone or in combination with chemotherapy or proteasome inhibition (reviewed in Lemke et al.¹⁶⁹). However, a major shortcoming of these clinical studies was that they failed to learn from the experiences with other molecularly targeted agents and were conducted in unselected patient populations.^{170, 171} Another limitation of first generation TRAIL receptor agonists may have been insufficient levels of receptor super-clustering; a number of second generation TRAIL agonists are now in development with novel mechanisms of action that overcome this limitation, for example, MedImmune’s multivalent ‘superagonist’, which efficiently engages and clusters TRAIL-R2.¹⁷² However, increased valency may increase the toxicity of second generation TRAIL-R agonists, and a recent phase I clinical trial with a tetravalent agonistic Nanobody targeting TRAIL-R2, TAS266, had to be halted at the lowest dose owing to hepatotoxicity. ¹⁷³ The opposite approach is required for the TNFα pathway, where biologics have been developed to block TNFα itself or TNFR1, preventing downstream activation of the NF-κB pathway and/or apoptosis induction and providing effective treatment for a number of inflammatory diseases, including SLE, rheumatoid arthritis and septic shock.^{174, 175, 176} TNFα-induced necroptosis has been implicated in a number of pathophysiological conditions such as Crohn’s disease⁹⁹ (reviewed by Linkermann and Green¹⁷⁷); thus, TNFα blockade may prove to be therapeutically beneficial in these situations.

A number of IAP antagonists (SMAC mimetics) are currently in clinical development and have shown potential as anti-cancer agents (reviewed by Fulda¹⁷⁸). They promote caspase activation and elicit an apoptotic response by binding to and inhibiting IAPs, which are overexpressed in many types of cancer.^{70, 71, 179, 180} Additionally, they activate the non-canonical NF-κB pathway through the accumulation of NIK, which is normally degraded by cIAP1.^{181, 182} This results in an upregulation of NF-κB target genes, including TNFα. TNFα can signal in an autocrine or paracrine manner, stimulating the assembly of TNFR1 Complex II (Figure 6) and activation of cell death via apoptosis and/or necroptosis.^{86, 92}

As it is a potent anti-apoptotic molecule, lowering FLIP expression in malignancies could lower the threshold for cell death. We and others have reported that histone deacetylase inhibitors such as vorinostat trigger the rapid ubiquitination and degradation of FLIP, thus sensitising cells to TRAIL or chemotherapy.^{122, 170} Additionally, a number of chemotherapeutics and other anti-cancer agents downregulate FLIP expression via multiple mechanisms (reviewed by Safa¹⁸³). As previously mentioned, the gene encoding procaspase-8 (Casp8) is silenced by methylation in several cancers, such as small cell lung cancer and neuroblastomas;¹⁸⁴ treatment with another class of epigenetic drugs, the demethylating agents such as 5-azacytidine can reverse this effect, thereby enhancing the potential for caspase-8-mediated apoptosis.²⁷

Conclusion and Perspective

As a result of their key roles in determining life and death outcomes, much work has focussed on the complexes formed by DED proteins. From this work, FLIP in particular has emerged as a master regulator of the signalling outputs from DED-containing complexes. It is probably for this reason that FLIP expression is regulated at multiple levels: by numerous transcription factors (such as NF-κB,¹⁸⁵ NFAT,¹⁸⁶ AP-1¹⁸⁷ and c-Myc¹⁸⁸); alternative splicing;¹⁸⁹ mRNA translation¹⁹⁰ and by posttranslational modifications, including its rapid turnover via the ubiquitin-proteasome system.^{118, 119} This exquisite level of regulation may have evolved to allow swift responses to various cellular stresses, for example, to safeguard against inappropriate activation of cell death or enhance cell death, depending on the cellular context. Biochemical and structural studies have demonstrated that DED-containing complexes are highly intricate with ubiqutination playing a key role. These complexes are also more numerous than previously appreciated, with the discovery of complexes such as the intracellular DISC and Ripoptosome and involvement of DED proteins in complexes such as the inflammasome. It is anticipated that future studies will reveal novel ways of therapeutically targeting DED protein complexes that could find clinical applications in cancers, inflammatory diseases and neurodegenerative diseases.