Apoptotic Caspases Suppress mtDNA-Induced STING-Mediated Type I IFN Production

HSC Dysfunction in Caspase-9-Deficient Mice Is Cell Extrinsic

To establish whether LSK expansion in Casp9^−/− mice was intrinsic to the LSK population itself, we generated mixed bone marrow chimeras by transplanting WT or Casp9^−/− E13.5 FLCs with WT filler FLCs into lethally irradiated recipients. Consistent with our previous observations, 12–16 weeks later, we observed an expansion of Casp9^−/− LSKs in the WT:Casp9^−/− chimeras. Strikingly, WT LSK numbers were also significantly increased relative to those in WT:WT chimeras (Figure 2A). Their Sca1 expression profile resembled that of Casp9^−/− Lin⁻ Kit⁺ cells (Figure 2B). Thus, WT LSKs expand in the presence of caspase-9-deficient hematopoietic cells, suggesting that HSC expansion in Casp9^−/− mice is driven by cell-extrinsic factors.

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Figure 2

Loss of Caspase-9 Results in Elevated Type I Interferon In Vivo

(A) FACS analysis of mixed bone marrow chimeras 12–16 weeks posttransplant of WT (CD45.2⁺) or Casp9^−/− (CD45.2⁺) with WT “bystander” (CD45.1 ⁺CD45.2⁺) E13.5 FLCs into lethally irradiated CD45.1⁺ recipients. Number of donor-derived bone marrow LSK cells from both fractions is displayed (n = 8 mixed bone marrow chimeras per genotype from 3–4 fetal livers per genotype). Means of WT (CD45.1 ⁺CD45.2⁺) “bystander” cells were compared using a two-tailed t test.

(B) Sca1 expression on Lin⁻Kit⁺ bone marrow cells from mixed bone marrow chimeras.

(C) Scatterplot of differentially expressed probes in microarray analysis of WT, Casp9^−/−, and Bak^−/− Bax^−/− Lin⁻Kit⁺CD45.2⁺ bone marrow cells. See also Table S1.

(D) Top ten gene sets (ranked by p value) from Gene Set Enrichment Analysis (GSEA) of Casp9^−/− in (C) (gray indicates type I IFN signatures).

(E) Top ten gene sets (ranked by p value) from GSEA of Bak^−/− Bax^−/− in (C).

(F and G) Real-time qPCR analysis of type I ISGs in fetal liver (F) and bone marrow cells (G) (n = 3–4 E13.5 fetal livers and 3–4 bone marrow chimeras per genotype). (H) IFN-β protein in serum of WT (n = 6), Casp9^−/− (n = 6), and Bak^−/− Bax^−/− (n = 8) bone marrow chimeras.

Unless indicated, means were compared to WT using a one-way ANOVA with Bonferroni correction. Data represent the mean ± SEM. *p ≤ 0.05, **p ≤ 0.01, and ***p ≤ 0.005.

Caspase-9-Deficient HSPCs Exhibit a Type I Interferon Response Signature

We therefore analyzed the gene expression profile of Lineage⁻Kit⁺ CD45.2⁺ hematopoietic stem and progenitor cells (HSPCs) harvested primary bone marrow chimeras. 495 genes (corresponding to 602 probes) were significantly upregulated in Casp9^−/− HSPCs (Figure 2C and Table S1 available online), with 275 genes (corresponding to 346 probes) differentially expressed (DE) in Bak^−/− Bax^−/− HSPCs. In Casp9^−/− HSPCs, gene set enrichment analysis using Camera identified enrichment for multiple type I IFN response signatures that were not apparent in Bak^−/− Bax^−/− HSPCs (Figures 2D and 2E). The top ten upregulated genes in Casp9^−/− HSPCs by fold change were all type I IFN targets (Table S1). Quantitative PCR (qPCR) analysis of canonical type I IFN-stimulated genes (ISGs) confirmed their upregulation relative to WT in Casp9^−/− FLCs and bone marrow cells (Figures 2F and 2G). We therefore examined levels of the type I IFNs, IFNα and IFN-β, in the serum of primary transplant recipients. Whereas IFNα was undetectable in all genotypic classes, we observed significantly elevated IFN-β in mice reconstituted with Casp9^−/−, but not Bak^−/− Bax^−/−, cells (Figure 2H).

HSC Expansion in Caspase-9-Deficient Mice Is Caused by Type I Interferons

Type I IFNs have been shown to induce HSC proliferation, leading to functional exhaustion in vivo (Essers et al., 2009; Sato et al., 2009). We therefore generated mice doubly deficient for caspase-9 and the type I IFN receptor (Ifnar1). Although loss of Ifnar1 did not rescue postnatal lethality of Casp9^−/− mice (data not shown), deletion of Ifnar1 prevented LSK expansion in Casp9^−/− fetal livers (Figures 3A and 3B). Next, bone marrow chimeras were generated by reconstituting recipients with Ifnar1^−/− Casp9^+/+, Ifnar1^+/+ Casp9^−/−, or Ifnar1^−/− Casp9^−/− FLCs. 16 weeks posttransplantation, recipients of Ifnar1^+/+ Casp9^−/− cells exhibited a 5-fold increase in LSKs relative to mice that received Ifnar1^−/− Casp9^+/+ cells (Figures 3C and 3D). In contrast, LSK numbers in Ifnar1^−/− Casp9^−/− chimeras were normal. Furthermore, deletion of Ifnar1 restored the ability of Casp9^−/− HSCs to engraft the host and contribute to all major lineages upon secondary transplantation (Figures 3E and 3F). Collectively, our data indicate that loss of caspase-9 in vivo leads to production of IFN-β, which feeds back to the HSC compartment, resulting in loss of self-renewal capacity.

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Figure 3

Type I Interferon Mediates the Hematopoietic Stem Cell Dysfunction Associated with Caspase-9 Loss

(A) Representative plots of LSK cell frequency in E13.5 fetal livers (gates display percentage of Lin⁻).

(B) FACS analysis of LSKs in Ifnar1^−/− Casp9^+/+ (n = 6), Ifnar1^+/+Casp9^−/− (n = 4), and Ifnar1^−/− Casp9^−/− (n = 6) E13.5 fetal liver.

(C) Bone marrow cellularity from Ifnar1^−/− Casp9^+/+ (n = 8), Ifnar1^+/+Casp9^−/− (n = 4), and Ifnar1^−/− Casp9^−/− (n = 5) bone marrow chimeras, 16 weeks posttransplant.

(D) Number of donor-derived LSK cells from Ifnar1^−/− Casp9^+/+ (n = 7), Ifnar1^+/+Casp9^−/− (n = 4), and Ifnar1^−/− Casp9^−/− (n = 5) bone marrow chimeras, 16 weeks posttransplant.

(E) Donor-CD45.2⁺ contribution to the peripheral blood B lymphocyte, T lymphocyte, myeloid cells, and bone marrow LSK cells of 1° and 2° recipients at 16 weeks posttransplant. Ifnar1^−/− Casp9^+/+ (n = 4), Ifnar1^+/+Casp9^−/− (n = 3), and Ifnar1^−/− Casp9^−/− (n = 4) donor fetal livers per genotype and three recipients per donor bone marrow.

(F) Plots of representative analysis of donor contribution to 2° recipient lymphoid lineages in (E).

Means were compared to WT using a one-way ANOVA with Bonferroni correction. Data represent the mean ± SEM. *p ≤ 0.05, **p ≤ 0.01, and ***p ≤ 0.005.

Apoptotic Mouse and Human Cells Produce Type I IFN When Caspases Are Inhibited

Activation of the apoptotic caspase cascade is impaired in the absence of caspase-9 (Li et al., 1997). We hypothesized that hematopoietic cells undergoing caspase-inhibited apoptosis might be the source of IFN-β. To test this, we treated WT murine splenocytes with the proapoptotic BH3 mimetic drug ABT-737, which targets the prosurvival proteins Bcl-x_L and Bcl-2 (Oltersdorf et al., 2005). ABT-737 induced caspase activation and cell death in splenocytes (Figures 4A and 4B). No IFN-β was detected in culture media of cells treated with ABT-737 alone. In contrast, when apoptosis was triggered in the presence of the pan-caspase inhibitor Q-VD-Oph, IFN-β was produced (Figure 4C). To test whether this mechanism is conserved between mice and humans, peripheral blood mononuclear cells (PBMCs) were isolated from the blood of five healthy adult donors and treated with ABT-737. Caspase-3/7 activation and loss of cell viability was observed over 24 hr (Figures 4D and 4E). Upon co-incubation with ABT-737 and Q-VD-Oph, IFN-β secretion was observed in all five human PBMC samples (Figure 4F). These data demonstrated that, in both human and murine hematopoietic cells, caspase-inhibited apoptosis results in the production of IFN-β.

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Figure 4

Apoptotic Hematopoietic Cells Produce Type I Interferon When Caspases Are Inhibited

(A) Viability of murine splenocytes treated with ABT-737 ± 20–30 µM Q-VD-OPh (QVD) for 24 hr.

(B and C) (B) Caspase activity and (C) IFN-β in culture supernatant after 24 hr of treatment (n = 4 mice). Means were compared using a two-tailed t test. Data represent the mean ± SEM. *p ≤ 0.05, **p ≤ 0.01, and ***p ≤ 0.005.

(D) Percentage of viable human PBMCs as quantitated by ATP levels (Cell Titer Glo) after 24 hr of treatment with ABT-737 and coincubation with 20–30 µM Q-VD-OPh (QVD).

(E) Bar graphs of caspase activity after 6 hr and (F) IFN-β in culture supernatant after 24 hr(n=5 healthy donor blood samples). Representative of two independent experiments.

Apoptotic MEFs Produce Type IIFN When Caspases Are Inhibited

To better examine the role of the intrinsic apoptosis pathway, we utilized immortalized mouse embryonic fibroblasts (MEFs). WT MEFs are dependent on the prosurvival proteins Mcl-1 and Bcl-x_L for survival (Figure 5A). MEFs lacking Mcl-1 undergo Bak- and Bax-mediated apoptosis in response to ABT-737 (van Delft et al., 2006). We therefore treated multiple Mcl1^−/− MEF lines with increasing concentrations of ABT-737 and coin-cubated them with either Q-VD-Oph or another pancaspase inhibitor, z-VAD.fmk. ABT-737 treatment of Mcl1^−/− MEFs induced caspase-3/7 activity and cell death (Figures 5B and 5C). Co-incubation with either z-VAD.fmk or Q-VD-Oph blocked caspase activation and prevented loss of viability. Although undetectable in supernatant from Mcl1^−/− MEFs treated with ABT-737 or caspase inhibitor alone, IFN-β was induced when ABT-737 (or other apoptotic stimuli) (Figure S2) was combined with z-VAD.fmk or Q-VD-Oph (Figure 5D). Upregulation of Ifnb1, the gene encoding IFN-β, was evident 4 hr posttreatment (Figure 5E). These data demonstrate that nonhematopoietic cells also produce IFN-β when undergoing caspase-inhibited apoptosis and implicate Bak and Bax as the initiators of the signal that triggers IFN-β production.

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Figure 5

Disabling Caspases Downstream of Bak and Bax Triggers Type IIFN Production

(A) Schematic diagram of the manipulation of intrinsic apoptosis in MEFs.

(B–D) (B) Viability of MEFs treated with ABT-737 ± 20–30 µM of Q-VD-Oph (QVD) or z-VAD.fmk (zVAD) for 24 hr, (C) bar graphs of caspase activity after 6 hr, and IFN-β in supernatant after 24 hr (n = 9 independent MEF lines). Means were compared using a two-tailed t test. See also Figure S2.

(E) Real-time qPCR analysis of Ifnb1 induction in Mcl1^−/− MEFs (n = 3 independent MEF lines). Means were compared using a two-tailed t test. (F and G) (F) Viability of MEFs expressing Bim_s2A or Bim_s4E and treated with ABT-737 for 20–24 hr and (G) caspase activity after 6 hr.

(H and I) (H) Immunoblot of lysates of MEFs treated with ABT-737 (1 µM) for 4 hr and (I) bar graph of IFN-β in supernatant after 20–24 hr (n = 3 independent MEF lines per genotype). See also Figures S2 and S3.

(J) Bone marrow cellularity from WT (n = 22), Bak^−/− Bax^−/− (n = 8), Apaf1^−/− (n = 5), Casp9^−/− (n = 11), Casp3^−/− (n = 13), Casp3^−/− Casp7^−/− (n = 7), and Bak^−/− Bax^−/− Casp9^−/− (n = 9) bone marrow chimeras.

(K) Number of donor-derived LSK cells from WT (n = 22),Bak^−/− Bax^−/−(n = 8), Apaf1^−/− (n = 5), Casp9^−/−(n = 11), Casp3^−/−(n = 13), Casp3^−/− Casp7^−/− (n = 7), and Bak^−/− Bax^−/− Casp9^−/− (n = 9) bone marrow chimeras.

(L) IFN-β in serum of WT (n = 10), Casp9^−/− (n = 5), Casp3^−/− (n = 5), Casp3^−/− Casp7^−/− (n = 4), and Bak^−/− Bax^−/− Casp9^−/− (n = 7) bone marrow chimeras. Not done, N.D.

(M) Real-time qPCR analysis of type I ISGs in bone marrow cells (n = 3–4 bone marrow chimeras per genotype). Means were compared using a two-tailed t test.

(N) Donor-CD45.2⁺ contribution to the peripheral blood B lymphocyte, T lymphocyte, myeloid cells, and bone marrow LSK cells of 1° and 2° recipients 16 weeks posttransplant (n = 3 donor fetal livers per genotype and 3 recipients per donor bone marrow).

Unless otherwise indicated, means were compared to WT using a one-way ANOVA with Bonferroni correction. Data represent the mean ± SEM. *p ≤ 0.05, **p ≤ 0.01, and ***p ≤ 0.005.

The Apoptotic Caspase Cascade Suppresses Bak/Bax-Mediated Type I IFN Production

To confirm whether Bak and Bax activation induces IFN-β production, we triggered mitochondrial apoptosis in WT, Casp9^−/−, and Bak^−/− Bax^−/− Casp9^−/− MEF lines by combining ABT-737 treatment with expression of Bim_S2A, which targets Mcl-1 (Lee et al., 2008). Expression of Bim_S4E, an inert form of Bim, served as a negative control. Casp3^−/− Casp7^−/− MEF lines were included to determine whether deletion of the effector caspases causes IFN production. In WT cells expressing Bim_S2A, ABT-737 treatment induced a 3-fold increase in caspase activity and an ~75% loss of viability (Figures 5F-5H). In contrast, both Casp9^−/− and Bak^−/− Bax^−/− Casp9^−/− cells were resistant to Bim_S2A + ABT-737. Analysis of the supernatant demonstrated abundant IFN-β secretion by Casp9^−/− and Casp3^−/− Casp7^−/− cells treated with Bim_S2A + ABT-737, but not WT or Bak^−/− Bax^−/− Casp9^−/− counterparts (Figures 5I and Figure S3). These data demonstrated that, in vitro, induction of Bak- and Bax-mediated apoptosis stimulates type I IFN production when either the apoptotic initiator (caspase-9) or effector (caspase-3/7) caspases are inactivated.

We reasoned that eliminating Bak and Bax would remove the stimulus for IFN-β production in caspase-deficient mice. To test this, we generated Casp9^−/−, Apaf1^−/−, Casp3^−/−, Casp3^−/− Casp7^−/−, and Bak^−/− Bax^−/− Casp9^−/− bone marrow chimeras. 16 weeks posttransplantation, LSKs were increased in recipients of Apaf1^−/−, Casp9^−/− or Casp3^−/− Casp7^−/− cells, demonstrating that inhibition of the apoptotic caspase cascade at either the apoptosome or effector stage triggers LSK expansion (Figures 5J and 5K). In contrast, the LSK profile in Bak^−/− Bax^−/− Casp9^−/− chimeras was unchanged. Thus, in the absence of Bak- and Bax-mediated apoptosis, deletion of caspases does not cause LSK expansion. Accordingly, IFN-β levels in Bak^−/− Bax^−/− Casp9^−/− chimeras were equivalent to WT, and no upregulation of type I IFN response genes was apparent (Figures 5L and 5M). Secondary bone marrow transplants confirmed that loss of Bak/Bax-mediated apoptosis also rescued HSC function, with Bak^−/− Bax^−/− Casp9^−/− bone marrow efficiently engrafting the host and contributing to the LSK compartment and all major lineages (Figure 5N). Thus, deletion of Bak and Bax prevents IFN-β production and HSC dysfunction in mice lacking a functional apoptotic caspase cascade.

Dying Cells Upregulate Type IIFN Production in a Cell-Intrinsic Manner

We next examined whether apoptotic cells secrete IFN-β in a cell-intrinsic manner. Ifnar1^−/− Casp9^−/− MEFs were transduced with Bim_S2A–GFP or Bim_S4E–GFP and co-plated with unmanipulated Ifnar1^−/− MEFs (Figures 6A and 6B). Ifnar1-deficient cells were utilized to eliminate paracrine/autocrine effects of IFN-β. Cocultures were treated with ABT-737, which induced apoptosis in the Ifnar1^−/− Casp9^−/− Bim_S2A–GFP cells, but not the other three cell populations. Subsequently, GFP⁺ (apoptotic or nonapoptotic) and GFP⁻ (bystander) cells were sorted, and expression of Ifnb1 was analyzed by qPCR. Relative to healthy bystanders, a significant upregulation of Ifnb1 mRNA was observed in apoptotic, but not nonapoptotic, cells (Figures 6B and 6C). When using WT (rather than Ifnar1^−/−) bystanders, type I IFN response genes were strongly induced in bystanders cocultured with apoptotic cells (Figure 6D), indicating that cells undergoing caspase-inhibited apoptosis actively transcribe Ifnb1 and secrete bioactive IFN-β.

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Figure 6

mtDNA Triggers Type IIFN Production during Caspase-Inhibited Apoptosis

(A) Representative plots of infection efficiency of the MEFs used in (B–D).

(B and C) Real-time qPCR analysis of nonapoptotic (expressing Bim_s4E, B) and apoptotic (expressing Bim_s2A, C) MEFs and their respective Ifnar1^−/− bystander from cocultures after treatment with ABT-737 (500 nM) for 18–20 hr (data combined from three experiments, with two independent MEF lines per genotype). mRNA expression is shown relative to two independent housekeeping genes, tbp and gapdh.

(D) Real-time qPCR analysis of WT bystanders from cocultures after treatment with ABT-737 for 18–20 hr (data combined from two experiments, with two independent MEF lines/genotype).

(E) Real-time qPCR analysis of mtDNA content from Mcl1^−/− MEFs cultured in ethidium bromide to generate mtDNA-depleted (ρ⁰) MEFs (used in F-J). Representative image of MEFs stained with PicoGreen nucleic acid stain. Arrows indicate mtDNA. Scale bars, 10 um. See also Figure S4.

(F) Scatterplot of differentially expressed probes from microarray analysis of Mcl1^−/−ρ⁰ MEFs compared to their respective parental Mcl1^−/− MEF. (n = 3 independent MEF lines). Chromosome M, ChrM. See also Table S2.

(G) Table of a selected set of type I IFN response genes from analysis in (F).

(H) Bar graph of IFN-β in the supernatant of ρ⁰ and parental MEFs transfected with Poly(I:C)(HMW) (n = 3 independent MEF lines).

(I–K) (I) Bar graphs of the viability of ρ⁰ and parental MEFs treated with ABT-737 ± 20–30 µM of Q-VD-Oph (QVD) for 24 hr, (J) caspase activity after 6 hr, and (K) IFN-β in supernatant after 24 hr (n = 4 independent MEF lines).

Means were compared using a two-tailed t test. Data represent the mean ± SEM. *p ≤ 0.05, **p ≤ 0.01, and ***p ≤ 0.005.

Apoptotic IFN Production Is Driven by Mitochondrial DNA

Considering the mechanism by which Bax and Bax stimulate a cell to produce type I IFN, we reasoned that release of a mitochondrial factor into the cytoplasm could be the initiating event. In the context of microbial invasion, type I IFNs are induced by a range of pathogen-associated molecular patterns (PAMPs). Viral nucleic acids are an important, and potent, example. Their presence is detected by cytosolic receptors that activate signaling cascades leading to the upregulation of IFN transcription (Paludan and Bowie, 2013). We hypothesized that Bak- and Bax-mediated damage to mitochondria may cause the release of mtDNA and that the latter would act as a DAMP capable of recognition by the dying cell’s innate nucleic acid sensors. We therefore generated mtDNA-depleted cells (so-called “ρ⁰” cells) by culturing Mcl1^−/− MEFs in ethidium bromide (King and Attardi, 1989). qPCR analysis revealed a near-complete absence of mtDNA after three passages (Figures 6E and Figure S4). Expression profiling of ρ⁰ cells demonstrated that 28 genes (corresponding to 42 probes) were downregulated and 61 genes (corresponding to 82 probes) were upregulated (Figure 6F and Table S2). 18 probes corresponding to 9 mitochondrially encoded genes were present on the array. These were the most downregulated genes in ρ⁰ cells (Camera p value = 0.0095). Gene set analysis using Camera detected no significant enrichment for any of the c2 expression signatures (data not shown). In addition, a subset of known type IIFN response genes was not significantly downregulated in ρ⁰ cells (Figure 6G, Camera p value for downregulation = 0.45). Consistent with there being no functional impairment of IFN response pathways, ρ⁰ cells responded normally when transfected with poly(I:C) (Figure 6H). Treatment of Mcl1^−/− ρ⁰ cells with ABT-737 resulted in caspase activation and a loss of viability similar in magnitude to that exhibited by the parental lines (Figures 6I and 6J). In the presence of Q-VD-Oph, IFN-β production was observed in the parental, but not ρ⁰, cells (Figure 6K). These data indicate that mtDNA is the trigger for IFN production downstream of Bak- and Bax-mediated mitochondrial damage.

Apoptotic versus Nonapoptotic Roles for Apoptotic Caspases

Apoptotic caspases have been ascribed a number of functions beyond the internal demolition of dying cells. Some of these (e.g., prostaglandin-induced tumor cell repopulation [Huang et al., 2011] and AMPA receptor internalization [Li et al., 2010b]) appear to be a by product of apoptotic cell death. Others (e.g., iPS cell reprogramming [Li et al., 2010a] and microglia activation [Burguillos et al., 2011]) are thought to represent “nonapoptotic” roles for caspases. Our genetic experiments, both in vitro and in vivo, demonstrate that, in suppressing IFN production, the apoptotic caspases play an apoptotic role, i.e., function downstream of Bak and Bax. Unless Bak and Bax are activated, mtDNA is not released, and caspases are not required to attenuate mtDNA-induced DAMP signaling. This has two important implications. First, blocking the apoptotic caspase cascade during apoptosis triggers the production of IFN-β, a potentially significant confounding experimental factor. This should be considered whenever the apoptotic caspase cascade is genetically or pharmacologically manipulated. Perturbations in type I IFN signaling may explain some of the published biological roles for caspases. Second, the fact that caspase-deficient mice present with a dramatic increase in LSKs, whereas Bak^−/− Bax^−/− animals do not, initially suggested that caspases play a “nonapoptotic” role in HSCs. However, the subsequent experiments with Bak^−/− Bax^−/− Casp9^−/− cells and bone marrow chimeras demonstrated that the phenotype is dependent on Bak/Bax activation. This highlights the importance of establishing whether apoptotic caspase activation in a given setting is the result of upstream mitochondrial damage or an alternative signaling mechanism.

mtDNA Activates the STING-Dependent Cytosolic DNA Sensing Pathway

Our data indicate that Bak/Bax-dependent mtDNA release triggers the cGAS/STING-dependent cytosolic DNA sensing pathway. mtDNA released during cell death has been previously reported to provide a second signal that cooperates with signal 1 (e.g., LPS) to activate the NLRP3 inflammasome and induce IL-1β production (Shimada et al., 2012). Whether mtDNA released via Bak/Bax also activates the inflammasome is unclear. However, other than IFN-β, mice with a caspase-9-deficient hematopoietic system did not exhibit elevated proinflammatory cytokine serum levels or any signs of systemic autoinflammatory disease when aged to 12 months (Figure S6). This suggests that, at least in the context of steady-state hematopoietic cell turnover, Bak/ Bax-mediated mtDNA release does not result in unbridled inflammasome activity when the caspase cascade is inactivated. Whether the elevations in IFN-β caused by loss of the caspase cascade lead to the development of autoimmune disease over the longer term and whether perturbations in caspase activity might contribute to human autoinflammatory/autoimmune disease remain to be established.

Mechanisms of Caspase-Mediated Suppression of Type IIFN

During apoptosis, caspases orchestrate a global program of cellular demolition, targeting ~1,000 protein substrates (Crawford and Wells, 2011). They cause DNA damage, suppress transcription, shut down protein translation, and disable a host of other essential cellular processes. Unlike genomic damage, which is triggered by cleavage of iCAD, the inhibitor of cas-pase-activated DNase (CAD) (Enari et al., 1998), and PS exposure, which is facilitated by cleavage of Xkr8 (Suzuki et al., 2013), it seems likely that caspase-mediated suppression of IFN-β production (and DAMP signaling generally) is the result of multiple redundant processes. First, the fact that apoptotic CAD^{CRISPR−/−} MEFs secrete detectable amounts of IFN-β suggests that CAD-mediated genomic damage contributes to the attenuation of gene expression (Figure S7). Second, caspase-3/7 might cleave and inactivate a component or components of the type I IFN production pathway. There is evidence that IFN signaling intermediates, including Irf3, can be targeted by caspases (Crawford et al., 2013). Third, caspase-3/7 could mediate the degradation of mtDNA, thereby preventing its interaction with cGAS. This might occur via CAD or perhaps lysosomal deoxyribonuclease (DNase) II, which can digest the genomic DNA of engulfed cells. Although the precise contribution these and other mechanisms make to suppressing IFN-β production in apoptotic cells remains to be established, it seems unlikely that any one of them is solely responsible. We suggest that caspase-mediated acceleration of cell death and suppression of DAMP signaling are inextricably linked. Both outcomes are achieved via global cellular demolition.

Potential Crosstalk between Apoptosis and Necroptosis

Necroptosis is a regulated form of necrotic cell death mediated by receptor-interacting protein kinase 3 (RIPK3) and the pseudokinase Mlkl (Linkermann and Green, 2014). It is triggered by prodeath ligands like TNFα and requires the inhibition or loss of caspase-8. In vitro, necroptosis is typically induced by TNFα and a caspase inhibitor such as z-VAD.fmk or Q-VD-Oph. The latter are both broad-spectrum inhibitors that target caspase-8, caspase-9, and caspase-3/7 (Chauvier et al., 2007). If necroptotic stimuli induce mitochondrial damage and mtDNA release, our results would predict that pan-caspase inhibition would result in IFN production by dying cells. Previous studies suggest that necroptosis triggered by TNFα, z-VAD.fmk, and cycloheximide causes Bak/Bax-dependent mitochondrial damage (Irrinki et al., 2011) and that Bak^−/− Bax^−/− MEFs are partially protected from necroptotic death induced by TNFα, Q-VD-Oph, and a Smac mimetic (TSQ) (Moujalled et al., 2013). Given that, like TNFα, type I IFNs, in combination with caspase inhibition, can induce necroptosis (Dillon et al., 2014; Thapa et al., 2013), our data suggest the potential for feed-forward effects driven by mtDNA-induced IFN production during caspase-inhibited necroptosis.

Pharmacological Inhibition of Apoptotic Caspases

Several caspase inhibitors have undergone clinical trials. They include the caspase-1 inhibitors VX-740 and VX-765 (Belnacasan) and the pan-caspase inhibitors IDN-6556 (Emricasan) and GS-9450 (O’Brien and Dixit, 2009). To date, none have received approval, although IDN-6556 is in ongoing trials for a number of indications, including alcoholic hepatitis, hepatic impairment, and islet transplantation. Evidence that VX-765 can block the pyroptotic death of HIV-infected CD4 T cells (Doitsh et al., 2014) has added to the hope that caspase inhibitors may yet find clinical application. That study highlighted their potential as a new class of antiviral drugs that target the host, not the virus. Our findings support this notion from an entirely different angle, suggesting that apoptotic caspase inhibition might be an effective means of amplifying endogenous IFN production. It will be interesting to see whether patients treated with a pancaspase inhibitor exhibit elevations in IFN-β levels.

Cell Death Assays

Cell death was induced by exposure to ABT-737 (AbbVie), Dexamethasone (Sigma), WEHI-539 (MedChemExpress), or Etoposide (Hospira). Where indicated, cells were preincubated for 1 hr with MRT-67307 (Sigma) and for 15–30 min with ABT-737, followed by continuous exposure to Q-VD-Oph (SM Biochemicals) or zVAD.fmk (R&D Systems). Cell viability was quantified by CellTiterGlo (Promega) or flow cytometric analysis of cells excluding 5 µg/ml propidium iodide (PI) (Sigma) and, where indicated, cells also negative for AnnexinV-FITC (InvivoGen) binding using a FACSCailbur (BD) or LSRII (BD). Caspase activity was assayed by the addition of caspase3/7Glo (Promega) or by immunoblotting as described in the Extended Experimental Procedures.

Measurement of IFN-β

IFN-β protein was measured using the VeriKine-HS Human or Mouse Interferon Beta ELISA (PBL Assay Science).

Generation of mtDNA-Depleted ρ⁰ Cells

MEFs were cultured in Dulbecco’s modified Eagle’s medium (DMEM)(GIBCO) supplemented with 4 mM L-glutamine, 4.5 g/l glucose, 10% FCS, 100 µg/ml sodium pyruvate, and 50 µg/ml uridine as described (Hashiguchi and Zhang-Akiyama, 2009). 100 ng/ml ethidium bromide was added to the medium for 6-0 days. qPCR evaluation of mtDNA content by qPCR, transfection with Pol-y(I:C)(HMW)/LyoVec (InvivoGen), and expression profiling are described in Extended Experimental Procedures.

Immunoprecipitation/PCR

Mcl1^−/− MEFs transduced with an expression plasmid (pMSCV-IRES-Hygro) encoding Flag-tagged mouse cGAS were treated with ABT-737 and Q-VD-Oph or left untreated. Following crosslinking of DNA and associated proteins, immunoprecipitation was performed with anti-FLAG M2 (Sigma) or mouse-IgG (BD) and coprecipitated DNA examined by real-time qPCR. Oligonucleotide sequences are listed in Extended Experimental Procedures.

The authors thank J. Corbin and L Di Rago for excellent technical assistance; K. Franks, E. Lanera, K. McGregor, S. Ross, K. Stoev, L. Wilkins, and E. Kryan for outstanding animal husbandry; S. Monard for expert flow cytometry assistance; T. Mak for Apaf1^+/− mice; R. Flavell for Casp3^+/− Casp7^+/− mice; C. Thompson for Bak^+/− and Bax^+/− mice; P. Crack for Ifnar1^+/− mice; D. Fairlie and E. Lee for Bim variants; K. Aumann and C. De Nardo for manuscript assistance; and A. Strasser, J. Choi, S. Best, A. Rongvaux, and R. Flavell for insightful discussions. This work was supported by Project Grants (516725 and 575535), Program Grants (461219, 461221, and 1016647), Fellowships (D.C.S.H. and B.T.K.), and an Independent Research Institutes Infrastructure Support Scheme Grant (361646) from the Australian National Health and Medical Research Council; a scholarship from the Leukemia Foundation of Australia (M.J.W.), fellowships from the Cancer Council of Victoria (M.J.W. and D.M.); an Australian Postgraduate Award scholarship (K.M.); a fellowship from the Sylvia and Charles Viertel Foundation (B.T.K.); the Australian Cancer Research Fund; and a Victorian State Government Operational Infrastructure Support Grant.