Abstract
Eiger (Egr), the homolog of the mammalian tumor-necrosis factor (TNF), is the ligand of the c-Jun N-terminal kinase (JNK) stress response signaling pathway in Drosophila. Although expression of Egr frequently leads to apoptosis, it has also been implicated in activation of non-apoptotic cell death. However, it is not yet clear how Egr can induce both apoptosis and non-apoptotic cell death, and if so, how such processes are coordinated. Here, we show that expression of Egr in the developing Drosophila eye induces apoptosis and non-apoptotic developmental defects, both of which are JNK-dependent. Intriguingly, when apoptotic effector caspases DrICE and Dcp-1 are defective or inhibited, expression of Egr induces necrosis characterized by loss of cell membrane integrity, translucent cytoplasm and aggregation of cellular organelles. Surprisingly, the induction of necrosis depends on the catalytic activity of the initiator caspase Dronc and the input from JNK signaling independently of their roles in apoptosis. Therefore, similar to the mammalian caspase-8, caspases in Drosophila also have dual roles in promoting TNF-mediated apoptosis and inhibiting necrosis.
Introduction
Apoptosis, a major form of programmed cell death, plays crucial roles in elimination of both developmentally unwanted cells and stress-induced damaged cells.1 The key factors driving apoptosis are caspases, a family of cysteine proteases. These apoptotic caspases can be further grouped into initiator (or apical) versus effector (or executioner) caspases based on the structure of their N-terminal prodomains and their cleavage substrates.2 The initiator caspases have long N-terminal prodomains. Once activated, they further cleave and activate the effector caspases. In contrast, the effector caspases have short prodomains. Once activated, effector caspases cleave a broad spectrum of intracellular substrates to execute cell death. In mammals, caspases can be activated by either intrinsic or extrinsic apoptotic machineries.3 Mitochondria are key components in the intrinsic pathway. In response to apoptotic stimuli, cytochrome c is released from mitochondria and binds to the adaptor protein Apaf-1 forming the apoptosome. Caspase-9, an initiator caspase, then interacts with the apoptosome and becomes activated. Activated caspase-9 further cleaves and activates effector caspases such as caspase-3 and –7 leading to apoptosis. Activities of caspase-9, −3 and −7 are also subject to repression by the inhibitor of apoptosis proteins (IAPs). This inhibition can be removed by pro-apoptotic proteins (or IAP antagonists) such as Smac/Diablo and HtrA2/Omi which are also released from mitochondria.4
In contrast to the intrinsic pathway, the extrinsic apoptosis pathway is initiated by activation of cell surface death receptors.3 Examples of death ligands that can activate death receptors are the tumor necrosis factor (TNF) superfamily, a group of cytokines which was initially discovered because of its anti-tumor activity.5 Binding of TNF ligands to their receptors (TNF receptors, TNFRs) promotes TNFR-mediated recruitment of various death-inducing protein complexes depending on the context.6 A key component common in these complexes is caspase-8, an initiator caspase. Once activated, caspase-8 can further cleave and activate effector caspases, such as caspase-3 and –7, triggering apoptosis or activate the intrinsic pathway to further enhance the apoptotic response.7 Importantly, caspase-8 also cleaves and inactivates two receptor-interacting serine/threonine-protein kinases (RIPKs), RIPK1 and RIPK3, thus preventing activation of necroptosis, another type of programmed cell death (programmed necrosis) which is morphologically distinct from apoptosis.6, 8 Therefore, when caspase-8 is deficient or inhibited, activation of TNF and TNFR can lead to necroptosis through RIPK1 and PIPK3. In addition to cell death, functions of TNF family members have also been revealed in immunity, inflammation, cell survival and proliferation.6 Many of these functions are mediated by activation of the nuclear factor-κB (NF-κB), a transcription factor controlling multiple target gene expression in a context-dependent manner.9 In addition to NF-κB, the TNF signaling can also activate the c-Jun N-terminal kinase (JNK) and the p38 mitogen-activated protein kinase (p38-MAPK) pathways. Both of these pathways are pleiotropic in regulating a variety of cellular processes including cell death, proliferation and differentiation.10 It is therefore not surprising that dysregulated TNF-TNFR signaling is associated with numerous pathological situations including cancer and inflammatory diseases.11
The core components in the apoptosis pathway are evolutionarily conserved. In Drosophila, apoptotic stimuli induce expression of pro-apoptotic proteins Head involution defective (Hid), Reaper (Rpr) and Grim.12 These pro-apoptotic proteins act as antagonists to Diap1, the major Drosophila IAP, which inhibits activities of both initiator caspases, e.g. Dronc13, 14, and effector caspases, e.g. DrICE15 and Dcp-116. Once Dronc is released from the inhibition of Diap1, it induces formation of the apoptosome through its interaction with Dark, the Drosophila homolog of Apaf-1.17 Dronc is then activated. It further cleaves and activates effector caspases such as DrICE and Dcp-1 to induce apoptosis. Compared to mammals, mitochondria are less important in apoptosis in Drosophila. However, the pro-apoptotic proteins Hid, Rpr and Grim need to localize to mitochondria to exert their apoptotic functions.18–20 Although it is still a subject of debate that the extrinsic apoptosis pathway exists in Drosophila, one TNF homolog, Eiger (Egr), has been identified.21, 22 Two TNFRs Wengen (Wgn)23 and, more recently, Grindelwald (Grnd)24 have also been reported. Similar to mammalian TNFs, Egr plays multiple roles in regulating cell death, host defense, pain sensation, nutrient response, tissue growth and regeneration in a context-dependent manner.25–27 However, Egr exerts its functions mainly through activation of the JNK pathway in Drosophila. For example, expression of Egr under the control of an eye-specific driver GMR (GMR>egr) activates JNK and cell death resulting in small adult eyes.21, 22, 28 This cell death may not be apoptotic because it cannot be suppressed by P35 (ref.21), an inhibitor of the effector caspases DrICE and Dcp-1 (ref.29). Paradoxically, GMR>egr-induced eye ablation phenotype can be partially rescued by defective apoptosis through reduction of pro-apoptotic gene expression or inhibition of the initiator caspase Dronc.22, 28 It is therefore not yet clear whether and how Egr can induce apoptosis and non-apoptotic cell death in parallel.
The core components in the apoptosis pathway are evolutionarily conserved. In Drosophila, apoptotic stimuli induce expression of pro-apoptotic proteins Head involution defective (Hid), Reaper (Rpr) and Grim.12 These pro-apoptotic proteins act as antagonists to Diap1, the major Drosophila IAP, which inhibits activities of both initiator caspases, e.g. Dronc13, 14, and effector caspases, e.g. DrICE15 and Dcp-116. Once Dronc is released from the inhibition of Diap1, it induces formation of the apoptosome through its interaction with Dark, the Drosophila homolog of Apaf-1.17 Dronc is then activated. It further cleaves and activates effector caspases such as DrICE and Dcp-1 to induce apoptosis. Compared to mammals, mitochondria are less important in apoptosis in Drosophila. However, the pro-apoptotic proteins Hid, Rpr and Grim need to localize to mitochondria to exert their apoptotic functions.18–20 Although it is still a subject of debate that the extrinsic apoptosis pathway exists in Drosophila, one TNF homolog, Eiger (Egr), has been identified.21, 22 Two TNFRs Wengen (Wgn)23 and, more recently, Grindelwald (Grnd)24 have also been reported. Similar to mammalian TNFs, Egr plays multiple roles in regulating cell death, host defense, pain sensation, nutrient response, tissue growth and regeneration in a context-dependent manner.25–27 However, Egr exerts its functions mainly through activation of the JNK pathway in Drosophila. For example, expression of Egr under the control of an eye-specific driver GMR (GMR>egr) activates JNK and cell death resulting in small adult eyes.21, 22, 28 This cell death may not be apoptotic because it cannot be suppressed by P35 (ref.21), an inhibitor of the effector caspases DrICE and Dcp-1 (ref.29). Paradoxically, GMR>egr-induced eye ablation phenotype can be partially rescued by defective apoptosis through reduction of pro-apoptotic gene expression or inhibition of the initiator caspase Dronc.22, 28 It is therefore not yet clear whether and how Egr can induce apoptosis and non-apoptotic cell death in parallel.
Here, we report that GMR>egr primarily induces JNK-dependent apoptosis as well as non-apoptotic developmental defects in the Drosophila eye. However, when apoptosis is blocked through inhibition of effector caspases DrICE and Dcp-1, necrosis is induced instead. Intriguingly, loss of one copy of the gene encoding the initiator caspase Dronc suppresses necrosis but not Egr-induced apoptosis. Moreover, activation of regulated necrosis requires the catalytic activity of Dronc as well as the additional input from the JNK signaling pathway. Therefore, caspases are crucial in regulating TNF-triggered necrosis in Drosophila which is analogous to its mammalian counterparts.
Results
The cleaved Dcp-1 antibody is a specific marker for activated effector caspases DrICE and Dcp-1 in Drosophila
To determine whether GMR>egr induces apoptosis and/or non-apoptotic cell death, we first sought to identify a marker that specifically recognizes activated effector caspases, e.g. cleaved DrICE and Dcp-1, in Drosophila because antibodies recognizing the cleaved human caspase-3 are not specific to cleaved DrICE and Dcp-1.30 The recently developed cleaved Dcp-1 (Asp216) antibody (referred to as cDcp1) from Cell Signaling Technology is a polyclonal antibody raised against the large 22 kDa fragment of cleaved Dcp-1. Although this antibody has been increasingly used to label apoptosis in Drosophila 31, a detailed characterization of its specificity has not been carried out. We therefore addressed this by using GMR-hid, a transgene leading to induction of apoptosis specifically in the developing Drosophila eye.32 In late 3rd instar eye discs, compared to wild type where apoptosis occurs at a very low level (Fig.2e), GMR-hid induces two waves of apoptotic cells as indicated by TUNEL, an assay detecting DNA fragmentation a hallmark of apoptotic cells (Fig.1a).33 cDcp1 antibodies also recognize these two apoptotic waves in GMR-hid as expected (Fig1b). To determine whether cDcp1 only detects the cleaved Dcp-1, we analyzed its staining in dcp-1 null mutants which does not obviously affect GMR-hid-induced apoptosis (Fig. 1c). Intriguingly, cDcp1-labeling of two apoptotic waves persists (Fig.1d) suggesting at least one other apoptotic protein is recognized by cDcp1. In addition to Dcp-1, DrICE is the second major effector caspase mediating apoptosis in somatic tissues including eye discs.34, 35 Moreover, DrICE and Dcp-1 share the same cleavage site for their activation.30, 36 It is therefore possible that cDcp-1 also recognizes the cleaved DrICE. To examine this, we labeled GMR-hid discs with cDcp1 in drICE null mutants. Although apoptosis is almost completely lost in drICE mutants as indicated by TUNEL (Fig.1e), cDcp1 detects a relatively low level of signals in the GMR domain (Fig.lf) which are presumably the cleaved Dcp-1 proteins. Importantly, cDcpl-and TUNEL-labeling is lost in dcp-1; drICE double null mutants (Fig.1g,h). Therefore, cDcpl recognizes both cleaved Dcp-1 and cleaved DrICE.
Expression of Eiger induces strong apoptosis through the canonical apoptosis pathway and the pro-apoptotic gene hid
GMR>egr induces a strong eye ablation phenotype (compare 2b to 2a).21, 22 As controls, we validated that GMR>egr-induced small eyes are completely suppressed by knocking down egr itself through RNAi (Fig.2c) or expressing pucker (puc) (Fig.2d), a negative regulator of JNK.37, 38. Therefore, GMR>egr-induced eye ablation is indeed due to expression of egr and activation of JNK. To assess whether GMR>egr induces apoptosis, we used cDcp1 to label 3rd instar larval eye discs in which the GMR promoter is expressed. Compared to wild type (Fig.2e), a strong wave of cDcp1-labeling was observed in GMR>egr discs (Fig.2f). To further confirm that apoptosis is induced by GMR>egr genetic analyses on key components of the canonical apoptosis pathway were conducted. Loss of Dronc, the major initiator caspase mediating apoptosis in Drosophila 39, 40, or expression of P35, an inhibitor of activated DrICE and Dcp-1 (ref.29), completely blocks cDcp1 signals induced by GMR>egr (Fig.2g,h). Consistently, although dronc null mutants are mostly pharate adult lethal, they show suppressed GMR>egr eyes (compare 2m to 2b) when they are dissected out of the pupal cases. Furthermore, expression of a RING domain-deleted (therefore stabilized) form of Diap1 (BIR), the apoptosis inhibitor upstream of Dronc 41, also strongly suppresses GMR>egr-induced small eyes (compare Fig.2l to 2b). Therefore, GMR>egr induces massive apoptosis in the developing Drosophila eye.
To identify which pro-apoptotic genes mediate GMR>egr-induced apoptosis, we first examined expression of hid and rpr by using their reporters.42 Compared to the control, expression of hid, but not rpr, shows significant increase (Supplementary Fig.Sl) in response to egr expression. This is consistent with a previous report.28 To determine whether hid, but not rpr, is required for GMR>egr-induced apoptosis, mutant analyses were performed. While loss of rpr by using a combination of a deletion 43 and a null mutant of rpr (ref.44), rpr87/XR38, does not significantly affect the level of GMR>egr-induced apoptosis (Fig.2j), GMR>egr-induced apoptosis is lost in hid mutant clones (Fig.2i-i’’). Consistently, GMR>egr-induced small eyes are suppressed by hid mosaics (Fig.2k), but not by rpr mutants (Fig.2n). Taken together, expression of Egr activates apoptosis through the pro-apoptotic gene hid in the developing Drosophila eye.
Expression of Eiger also induces JNK-dependent non-apoptotic defects
Intriguingly, unlike expression of puc (Fig.2d), loss-of-dronc or expression of Diapl (GMR>BIR) does not completely restore GMR>egr eyes back to normal indicated by their glassy appearance suggesting irregular ommatidial patterning (Fig.2i,m). Thus expression of Egr in the eye can induce defects other than apoptosis. These defects appear to depend on JNK activity. Similar to expression of puc, expression of a dominant negative form of bsk (bskDN, bsk=Drosophila JNK) or hemizygous mutants of Takl (Tak12527, null mutant 45), an upstream kinase of JNK, almost completely rescue GMR>egr-induced adult eye defects (Fig.3a,b). Therefore, in addition to apoptosis, GMR>egr can also induce JNK-dependent non-apoptotic defects (Fig.3c).
To further characterize GMR>egr-induced non-apoptotic defects at the cellular level, we examined ommatidial organization in pupal eye discs which is a sensitive readout of defects in eye development. Developing pupal eye discs are composed of well-patterned ommatidia (Fig.3d,d’’, ELAV labelling) and interommatidial cells (Fig.3d,d’). At 25°C, developmental apoptosis occurs at around 28h after puparium formation (APF28h) to remove extra interommatidial cells in eye discs (Supplemental Fig.S2a-a’’).46, 47 However, no apoptosis is observed at APF22h (Fig.4a,a’). We therefore performed our analysis of GMR>egr at APF22h to avoid developmental apoptosis. Compared to wild type eye discs where ommatidia and interommatidial cells are well-organized (Fig.3d-d’’), GMR>egr leads to irregular organization of ommatidia and interommatidial cells (Fig.3e-e’’) with both increased interommatidial spacing (Fig.3e’’, arrows) and ommatidial fusion (Fig.3e’’, arrowheads) observed. These GMR>egr-induced defects persist when apoptosis is inhibited in dronc null mutants (Fig.3g-g’’) or by expression of Diap1 (Fig.3i-i’’). As controls, at APF22h, ommatidial organization is not affected by loss-of-dronc (Fig.3f-f”) or expression of Diap1 (Fig.3h-h’’) alone. In contrast, GMR>egr-induced ommatidial organization defects are strongly suppressed by expression of bskDN (Fig.3j-j’’) or in hemizygous mutants of Tak1(Fig.3k-k’’). Altogether, these data indicate that GMR>egr induces both apoptosis and non-apoptotic, but JNK-dependent, defects in the developing Drosophila eye (Fig.3c).
Inhibition of effector caspases in GMR>egr induces non-apoptotic cell death
Another intriguing observation that has been reported previously is that, unlike dronc mutants or expression of Diap1, expression of P35, an inhibitor of effector caspases, cannot or only slightly suppress GMR>egr-induced eye ablation phenotype (compare Fig.4d to Fig.2b, quantified in Fig.5i). 21,22,48 One possibility is that expression of P35 may not be sufficient to block GMR>egr-induced apoptosis because P35 is a pseudo-substrate of effector caspases.29, 49 We looked into this possibility by using cDcp1 to label apoptotic cells. As a control, cDcp1 detects developmental apoptosis in pupal discs at APF28h (Supplementary Fig.S2). We therefore examined eye discs at APF22h to avoid developmental apoptosis. Compared to the wild type discs where no apoptosis is detected at APF22h (Fig.4a,a’), GMR>egr induces massive apoptosis and defective ommatidial organization (Fig.4b,b’). Notably, GMR>egr-induced apoptosis, but not ommatidial mis-organization, is almost completely blocked by expression of P35 (Fig.4c,c’). However, unlike dronc mutants or expression of Diap1, expression of P35 in GMR>egr still results in small eyes although it suppresses GMR>egr-induced apoptosis (compare Fig.4d to Fig.2l,m). Importantly, this is not due to any unspecific effects induced by P35 itself because double RNAi knockdown of drICE and dcp-1 or using drICE mutants does not suppress, or even enhance, GMR>egr- induced eye ablation phenotype (Fig.4e,f). Both knockdown of drICE and dcp-1 and loss-of-drICE can suppress GMR-hid-induced apoptosis confirming these RNAi lines and mutants are functional (Fig.4g-i). Taken together, these data suggest that inhibition of effector caspases, in particular DrICE, may lead to non-apoptotic cell death, while it suppresses apoptosis in GMR>egr.
The initiator caspase Dronc mediates Eiger-induced necrosis when apoptosis is blocked
To further characterize the non-apoptotic cell death induced by GMR>egr when effector caspases are inhibited, we performed labeling with propidium iodide (PI), a dye that enters the cells and binds to their nucleic acids when the cell membrane integrity is disrupted. Therefore, PI has been used to label necrotic cells, characterized by membrane rupture, in both Drosophila eye and wing tissues (Supplementary Fig.S3a,b)50, 51. Interestingly, no PIpositive cells were detected in late 3rd instar larval GMR>egr discs with or without expression of P35 (Supplementary Fig.S3c,d) suggesting the non-apoptotic cell death may not occur immediately following apoptosis inhibition. We then examined eye discs in the pupal stage of APF22h to allow more time for non-apoptotic cell death to occur. In GMR>egr a very low level of PI-labeling was observed (Fig.5a,a’). Notably, most of these PI signals do not co-localize with a DNA marker Hoechst (Fig.5a,a’’, arrowheads) suggesting that such a low level of PI-labeling may be unspecific. This is further supported by a comparable low level of PI signals observed in wild type discs (quantified in Fig.5c) under the same experimental condition. Therefore, only a low level of, probably unspecific, PI-labeling can be detected in wild type or GMR>egr pupal discs. In contrast, expression of P35 strongly induces PI-labeling in GMR>egr pupal discs (Fig.5b,b’). Importantly, the majority of these PI signals co-localizes with Hoechst-positive nuclei (Fig.5b,b’’, arrowheads) suggesting that they specifically label membrane-compromised cells. As a control, expression of P35 alone does not induce PI-labeling (Fig.5c). Altogether, these data suggest that a factor(s) upstream of effector caspases may trigger necrosis when apoptosis is blocked by P35. We therefore examined PI-labeling in GMR>egr discs in a background of dronc mutants or with expression of the stabilized Diapl (GMR-BIR) as they both suppresses GMR>egr-induced apoptosis upstream of effector caspases (see Fig.2l,m). Only background levels of PI-labeling were detected in these conditions (Fig.5c). Strikingly, loss of one copy of dronc, although it only weakly affects GMR>egr-induced apoptosis (Fig.5d,d’) and eye ablation phenotype (Fig.5e,i), it strongly suppresses PI-labeling induced in GMR>egr/GMR-p35 (Fig.5c). Consistently, dronc heterozygous mutants together with expression of P35 strongly suppress GMR>egr-induced eye ablation phenotype (Fig.5f,i). Therefore, Dronc is a key component mediating PI-positive cell death when GMR>egr-induced apoptosis is blocked by expression of P35.
It is known that Dronc, the initiator caspase, cleaves its downstream effector caspases DrICE and Dcp-1 to activate apoptosis in Drosophila.13, 36 We thus examined whether the catalytic activity of Dronc is required for non-apoptotic cell death induced in GMR>egr/GMR-p35 eye discs. To do this, either a wild type Dronc (droncwt) or a catalytic site-mutated form of Dronc (droncC>A) transgene was used.13 As expected, expression of the wild type Dronc restores PI-positive cell death (Fig.5c), and therefore eye ablation (Fig.5g,i), in GMR>egr/GMR-p35; dronc+/− animals. However, expression of droncC>A in the same background does not have these effects (Fig.5c,h,i). This suggests that the catalytic activity of Dronc is required for Egr-induced non-apoptotic cell death when apoptosis is blocked.
To further determine the type of non-apoptotic cell death induced in GMR>egr/GMR-p35, we performed the Transmission Electron Microscopy (TEM) analysis on APF22h pupal eye discs. Compared to the wild type control (Fig.6a), cells with typical apoptotic features 8 such as high-electron-density chromatin condensation and apoptotic bodies were frequently observed in GMR>egr discs (Fig.6b,b’). In contrast, cells with typical necrotic features were observed in GMR>egr/GMR-p35 eye discs (Fig.6c-c’’). These cells often have translucent cytoplasm, mal-shaped or unidentifiable nuclei, and aggregation of endoplasmic reticulum and other cellular organelles.8, 52 Therefore, it appears that necrosis is induced in GMR>egr/GMR-p35 pupal eye discs.
JNK signaling is required for Eiger-induced necrosis when apoptosis is blocked
Unlike GMR>egr, hid-induced apoptosis e.g. GMR-hid can be suppressed by expression of P35 (ref.32) or reduction of DrICE and Dcp-1 (Fig.4g-i). Therefore, factors other than inhibitors of apoptosis are also required for induction of necrosis in GMR>egr/GMR-p35 discs. Because Egr activates JNK signaling upstream of hid-mediated apoptosis, we examined whether there is non-apoptotic input from the JNK signaling contributing to induction of necrosis. Hypomorphic mutants of bsk, hep, MKK4 and Tak1,genes encoding various kinases in the JNK pathway 53, were used. We observed that heterozygosity of these mutants can only weakly or moderately suppress GMR>egr-induced eye ablation phenotype (compare Fig.7a-d to Fig.2b). Intriguingly, except hep1 (Fig.7f), heterozygosity of other mutants including bsk1, MKK4G680 and Tak12 can strongly suppress GMR>egr/GMR-p35-induced small eyes (compare Fig.7e,g,h to Fig.4d). This suggests that, in addition to activate apoptosis through hid, JNK signaling also contributes to induction of necrosis when apoptosis in blocked (Fig.7i).
Discussion
Our findings in this study suggest an analogy between Drosophila and mammals in regulation of TNF-induced necrosis (Fig.7i). Caspases play critical roles in these processes. In mammals, caspase-8 exerts a permissive role on apoptosis but an inhibitory role on necroptosis, a form of regulated necrosis. Here, we report that, in Drosophila, regulated necrosis can also occur when apoptosis is blocked. Interestingly, both apoptosis and necrosis induced by Egr, the Drosophila TNF, depend on the initiator caspase Dronc. However, loss of one copy of dronc blocks necrosis, but not apoptosis, suggesting that different levels of Dronc are required for apoptosis versus necrosis. In contrast to Dronc, effector caspases DrICE and Dcp-1, in particular DrICE, appear to inhibit Egr-induced necrosis. This inhibition can be released either by expression of the inhibitor P35 or reduction of DrICE and Dcp-1 by RNAi or mutants. However, inhibition of DrICE and Dcp-1 alone following apoptotic stresses is not sufficient to induce necrosis. For example, no necrosis is evident when apoptosis induced by expression of hid is inhibited (Fig.4g-i).29 This is further supported by our observation that heterozygous mutants of bsk, MKK4 and Tak1, genes encoding various kinases in the JNK pathway, suppress eye ablation in GMR>egr/GMR-p35 much better than in GMR>egr. Therefore, an additional input from Egr-induced JNK signaling is also required for induction of necrosis (Fig.7i).
How do DrICE and Dcp-1 inhibit Egr-induced necrosis? How does Dronc promote it when DrICE and Dcp-1 are inhibited? What factors coordinate the inputs from Dronc and JNK in this process? These are intriguing questions remained open (Fig.7i, question marks in red). In mammals, caspase-8 cleaves and inactivates RIPK1 and PIPK3 therefore inhibits TNF-induced necroptosis (Fig.7i). While pyroptosis, another type of regulated necrosis induced by pathogens, is activated by several inflammatory (non-apoptotic) caspases, such as caspase-1, –11 in mice and caspase-1, –4, –5 in humans, through their cleavage of gasdermin-D (GSDMD).54 Interestingly, capase-1, –4, –5 and –11 are structurally similar to the apoptotic initiator caspases. However, no homologs of RIPK1, RIPK3 and GSDMD have been reported in Drosophila. Our study suggests that the catalytic activity of Dronc is required for Egr-induced necrosis because only expression of the wild type Dronc, but not the catalytic site-mutated form of Dronc, can restore such necrosis in dronc mutants. Therefore, the key to understanding Egr-induced necrosis in Drosophila is to identify the non-apoptotic substrates of Dronc, DrICE and Dcp-1 that mediate the process. Interestingly, roles of necrosis-like cell death in Drosophila have also been reported in stress-induced loss of neural stem cells in the larval brain52, the development of male testes55 and female ovaries56. In these processes, the catalytic activity of Dronc or even Dronc itself doesn’t seem to be required. Hence, further studies are needed to understand how these different types of necrosis are regulated at the molecular level. Notably, a series of intracellular events specific to necrosis including mitochondrial dysfunction, ATP depletion, increased cytosolic Ca2+and organelle clustering appear to be conserved in multiple organisms in which necrosis has been studied.57 Are there conserved mechanisms regulating these necrotic cellular events throughout evolution despite death factors executing necrosis might be different across eukaryotes? Future genetic studies on various necrosis models in Drosophila will likely provide some insights into this question.
In addition to cell death, mammalian TNFs, as a family of pleiotropic cytokines, regulate a variety of cellular processes through activating multiple signaling pathways such as the NF-κB, JNK and p38-MAPK pathways (Fig.7i).6, 10 In Drosophila, although Egr exerts its functions mainly through the JNK pathway, context-dependent roles of the Egr-JNK signaling have also been reported in various cellular processes.25 Consistent with previous reports 21, 22, 28, we found that GMR>egr induces not only apoptotic but also non-apoptotic effects downstream of JNK activation in the developing Drosophila eye. The apoptotic effects are mediated by the pro-apoptotic gene hid and the canonical apoptosis pathway.
While the non-apoptotic effects are characterized by irregular ommatidial fusion and interommatidial spacing which may lead to the glassy appearance of the adult eyes. Recent studies suggest that these Egr-induced non-apoptotic effects might be mediated by the metabolic energy production pathways and the Toll/NF-κB pathway, a pathway regulates both immunity and development in Drosophila.48, 58 Further studies are required to elucidate how these pathways are coordinated to regulate eye development in response to Egr-induced activation of JNK signaling. It is also interesting to investigate whether these pathways contribute to Egr-induced necrosis when apoptosis is blocked.
Material and Methods
Drosophila genetics
Genetic crosses for all experiments were reared at 25°C. GMR-GAL4 UAS-egr (GMR>egr) 22, GMR-hid10 (ref.32), dcp-1Prev1 (ref.59), drICEΔ1 (ref.34), dronc129 (ref.39), rpr87(ref.44), XR38 (ref.43), hid05014 (ref.32), GMR-BIR41, GMR-p35 (ref.29), hid20-10-lacZ42, rprXRE- lacZ42, sev>GluLc50, MKK4G680 (ref.60), UAS-droncwt and UAS-droncC>A 13 were as described. bsk1, hep1, Tak12527, Tak12 and UAS-bskDN were obtained from the Bloomington stock center. UAS-drICERNAi and UAS-dcp-1RNAi were obtained from the NIG-Fly stock center. UAS-egrRANi (108814) was obtained from the VDRC stock center. For mosaic analysis with hid mutant clones, larvae of the following genotype were analyzed at the late 3rd instar larval stage: ey-FLP/+; GMR>egr/+; FRT80B /hid05014 FRT80B.
Immunohistochemistry
Pupal or larval discs were dissected, fixed (with 4% paraformaldehyde for 30min at room temperature), and then labeled with antibodies using standard protocols. Primary antibodies used are rabbit anti-cDcp1 (the cleaved Dcp-1 antibody, 1:500, Cell Signaling), rat anti-ELAV, mouse anti-Dlg and mouse anti-β-Gal (all 1:50, DHSB). Secondary antibodies were goat Fab fragments conjugated to Alex488, 555, or 647 (all 1:1000) from Molecular Probes.
Propidium Iodidum (PI), Hoechst labeling and TUNEL
For PI and Hoechst double labelling on pupal discs, freshly dissected discs were incubated in dark with 4μM PI (Sigma-Aldrich) and 16μM Hoechst (ThermoFisher) in Schneider’s media for 1h at room temperature as previously described.51 The discs were then fixed with 4% paraformaldehyde for 20min at room temperature followed by gentle washes. For PI labeling alone, larval or pupal eye discs were incubated with 15 μI for 10min at room temperature followed by fixation (with 4% paraformaldehyde for 20min at room temperature) and gentle washes as previously reported.50 PI labeling can also be observed and scored without fixation. For TUNEL (terminal deoxynucleotide transferase-mediated dUTP end labelling), dissected and fixed (with 4% paraformaldehyde for 20min at room temperature) discs were incubated in 100 mM Na-citrate with 0.1% Triton X-100 for 30 min at 65°C, followed by detection of dying cells using an in situ cell death detection kit (Roche).
Imaging and statistical analysis
Fluorescent eye disc images were taken with either a Zeiss or Leica confocal microscope. Adult fly eye images were taken using a Zeiss stereomicroscope equipped with an AxioCam ICC1 camera. For statistical analysis of PI-labeling (Fig.5c), at least 20 pupal discs from each genotype were used for counting the number of PI-positive cells. For quantification of adult eye size (Fig.5d), the ‘‘histogram’’ function in Adobe Photoshop CS was used to measure at least 10 representative adult eyes of each genotype. The statistical significance was evaluated through a one-way ANOVA with Bonferroni multiple comparison test using GraphPad Prism.
Transmission electron microscopy (TEM)
Freshly dissected pupal eye-brain complexes were fixed in 2.5% glutaraldehyde in 0.1M sodium cacodylate buffer (pH 7.4) for 45 min followed by a secondary fixation of 1% Osmium Tetroxide for 1 hour at room temperature. The samples were then washed with the buffer (10×5min) and dehydrated in ascending concentrations of ethanol before they were embedded in epoxy resin. Sections (90nm) were prepared and stained with uranyl acetate and lead citrate followed by examination using a JEOL 1200EX electron microscope fitted with a tungsten filament. Images were acquired through a GATAN MultiScan camera.
Conflict of Interest
The authors declare that there is no conflict of interest.
Acknowledgement
We apologize to colleagues whose work was not cited due to space limitations. We would like to thank Paul Stanley and Theresa Morris at the Centre for Electron Microscopy in the University of Birmingham for their assistance on TEM. We thank Kai Liu for advice on PI-labeling. We are grateful to Konrad Basler, Andreas Bergmann, Lei Liu, Pascal Meier, the Bloomington Stock Center, the NIG-Fly stock center in Japan, the VDRC stock center in Vienna and the Developmental Studies Hybridoma Bank (DSHB) in Iowa for fly stocks and reagents. ML is supported by the China Scholarship Council (CSC)-Birmingham joint PhD program. YF is supported by the Birmingham Fellowship, Grant BB/M010880/1 from the Biotechnology and Biological Sciences Research Council (BBSRC) UK and the Marie Curie Career Integration Grant (CIG) 630846 from the European Union’s Seventh Framework Programme (FP7).
References
Supplemental References
- S1.