
==== Front
J Biol Chem
J Biol Chem
The Journal of Biological Chemistry
0021-9258
1083-351X
American Society for Biochemistry and Molecular Biology

S0021-9258(24)02177-X
10.1016/j.jbc.2024.107676
107676
Research Article
Classical apoptotic stimulus, staurosporine, induces lytic inflammatory cell death, PANoptosis
Sarkar Roman ‡
Choudhury Sk Mohiuddin ‡
Kanneganti Thirumala-Devi Thirumala-Devi.Kanneganti@StJude.org
∗
Department of Immunology, St. Jude Children's Research Hospital, Memphis, Tennessee, USA
∗ For correspondence: Thirumala-Devi Kanneganti Thirumala-Devi.Kanneganti@StJude.org
‡ These authors contributed equally to this work.

14 8 2024
9 2024
14 8 2024
300 9 1076769 3 2024
27 7 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Innate immunity is the body’s first line of defense against disease, and regulated cell death is a central component of this response that balances pathogen clearance and inflammation. Cell death pathways are generally categorized as non-lytic and lytic. While non-lytic apoptosis has been extensively studied in health and disease, lytic cell death pathways are also increasingly implicated in infectious and inflammatory diseases and cancers. Staurosporine (STS) is a well-known inducer of non-lytic apoptosis. However, in this study, we observed that STS also induces lytic cell death at later timepoints. Using biochemical assessments with genetic knockouts, pharmacological inhibitors, and gene silencing, we identified that STS triggered PANoptosis via the caspase-8/RIPK3 axis, which was mediated by RIPK1. PANoptosis is a lytic, innate immune cell death pathway initiated by innate immune sensors and driven by caspases and RIPKs through PANoptosome complexes. Deletion of caspase-8 and RIPK3, core components of the PANoptosome complex, protected against STS-induced lytic cell death. Overall, our study identifies STS as a time-dependent inducer of lytic cell death, PANoptosis. These findings emphasize the importance of understanding trigger- and time-specific activation of distinct cell death pathways to advance our understanding of the molecular mechanisms of innate immunity and cell death for clinical translation.

Keywords

staurosporine
cell death
innate immunity
pyroptosis
apoptosis
necroptosis
PANoptosis
PANoptosome
caspase
inflammasome
RIPK3
RIPK1
MLKL
gasdermin
macrophages
Abbreviation

STS staurosporine

Reviewed by members of the JBC Editorial Board. Edited by Craig Cameron
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pmcRegulated cell death is a tightly controlled biological process essential for tissue development and homeostasis. In addition, cell death is a critical component of the host innate immune response to clear intracellular pathogens and damaged cells (1, 2, 3). Pattern recognition receptors (PRRs) in the innate immune system recognize pathogens, pathogen- or damage-associated molecular patterns (PAMPs or DAMPs), or homeostatic alterations to induce immune responses, including innate immune cell death (1, 3, 4). The classical cell death pathways include non-lytic and lytic pathways (3). Apoptosis is the canonical non-lytic pathway, characterized by cellular blebbing, nuclear fragmentation, and the formation of apoptotic bodies, which are then phagocytosed (5), preventing the release of inflammatory cytokines (6). Apoptosis is mediated by initiator caspases, including caspases-8 and -9, that activate the executioner caspases-3, -6, and -7 (7, 8, 9, 10, 11, 12, 13, 14, 15, 16). In contrast to non-lytic apoptosis, caspases are also involved in controlling lytic cell death pathways, such as the molecularly, biochemically, and genetically characterized pathways pyroptosis, necroptosis, and PANoptosis. Pyroptosis is a form of lytic cell death activated by inflammatory caspases, such as caspases-1 and -11 in mice and caspases-1, -4, and -5 in humans, and executed by gasdermin D (GSDMD) (17, 18, 19, 20, 21, 22, 23). Pyroptosis generally involves the formation of inflammasomes, which are molecular platforms for the activation of caspase-1 (24). In contrast, necroptosis is activated in the absence or inhibition of caspase-8 via RIPK1/3, and it is executed by MLKL (25, 26, 27, 28, 29, 30). Distinct from these cell death pathways, PANoptosis is a lytic, inflammatory innate immune cell death pathway initiated by innate immune sensors and driven by caspases and RIPKs through PANoptosome complexes (3, 31, 32, 33, 34, 35, 36, 37, 38, 39, 40). This form of cell death is executed by gasdermins, MLKL, and potentially other yet to be identified molecules cleaved by caspases. Both non-lytic and lytic cell death pathways have been extensively implicated in several infections, inflammatory diseases, and cancer (3, 22, 40, 41, 42, 43), highlighting the importance of cell death pathways across the disease spectrum. However, our understanding of the time-, trigger-, and dose-dependent activation of cell death pathways remains incomplete, and defining these molecular mechanisms and the genetic basis for these pathways is essential for successful therapeutic targeting.

In this study, we unexpectedly discovered that the well-recognized apoptotic trigger staurosporine (STS) induced delayed lytic cell death, PANoptosis, via the RIPK1-dependent caspase-8/RIPK3 axis in a time- and dose-dependent manner. Moreover, the deletion of pyroptotic components, caspase-1 or GSDMD, and necroptotic components, RIPK3 or MLKL, did not significantly protect the cells from STS-induced cell death; however, deletion of the core PANoptosome components caspase-8 and RIPK3 inhibited the lytic cell death. Overall, our study defined key trigger- and time-specific mechanisms for lytic cell death in response to the canonical apoptosis activator STS. Given the roles of non-lytic and lytic cell death pathways in health and disease, improved understanding of these molecular mechanisms can shape therapeutic strategies.

Results

Staurosporine induces lytic cell death in a dose- and time-dependent manner

Non-lytic and lytic cell death pathways have been implicated in diverse diseases. The classical non-lytic pathway, apoptosis, can be triggered by a variety of stimuli, including cytotoxic drugs (3, 5, 6, 7, 44). STS, a broad-spectrum protein kinase inhibitor used in clinical settings to treat cancer and other conditions (45, 46, 47), is one such cytotoxic drug that has long been used to induce apoptosis (45, 47, 48, 49, 50, 51, 52). However, the kinetics of this process and its involvement beyond apoptosis in other cell death pathways, including lytic pathways, remains unknown. Therefore, to understand the time-dependent cell death kinetics of STS, we treated mouse BMDMs with different doses of STS (1 μM, 5 μM, and 20 μM) in the presence of a membrane impermeable lytic cell death marker, propidium iodide (PI) (Fig. 1, A and B). We observed that STS induced lytic cell death in BMDMs in a dose- and time-dependent manner. While almost no lytic cell death was induced at the lowest dose (1 μM), lytic cell death was induced with higher doses of STS (5 μM and 20 μM) (Figs. 1, A and B; S1A).Figure 1 Staurosporine induces lytic cell death in a dose- and time-dependent manner. Bone marrow-derived macrophages (BMDMs) were treated with the indicated doses of staurosporine (STS), and cell death analysis was performed using IncuCyte live cell imaging. A, real-time quantification and (B) representative images of lytic cell death following STS treatment, measured using propidium iodide (PI), a membrane-impermeable dye that is a marker of lytic cell death. C, BMDMs were treated with STS and stained for activated caspases-3 and -7 (activated CASP3/7; classical markers of non-lytic cell death; red dye) and Sytox green (a membrane-impermeable dye that is a marker of lytic cell death). The yellow dots in the merged images denote cells that are positive for both activated CASP3/7 and Sytox green. D–F, real-time quantification of the indicated markers of cell death. The scale bar measures 25 μm (B, C). The data are shown as the mean ± SD (A, D–F) and are representative of at least three independent biological replicates (A–F). Two-way ANOVA with Sidak’s or Tukey’s multiple comparisons test was used to determine the statistical significance. Specific p values are denoted alongside each representative plot for comparisons between the untreated and STS-treated groups. The representative data points were only included once the findings of each experiment were confirmed to be consistent across the biological replicates.

To further understand the time-dependent activation of lytic cell death, we used a cell-permeable cleaved caspase-3/7 red dye and a cell impermeable DNA-binding Sytox green stain (which enters through damaged plasma membranes and is impermeable to intact membranes). We then monitored cell death kinetics in response to STS in BMDMs using live cell imaging. Caspases-3 and -7 were activated by early time points (2 h), while there was limited Sytox green uptake at this time (Fig. 1, C–E). In contrast, at a later timepoint (8 h) post-STS treatment, the cells were positive for both Sytox green and the cleaved caspase-3/7 red dyes (Fig. 1, C–F). Using another marker of non-lytic cell death, Annexin V, we observed similar results. Annexin V uptake occurred by 2 h post-STS treatment, followed by delayed PI or Sytox green uptake at later time points (Fig. S1, B and C). Together, these results suggest that STS induces the activation of caspases at early timepoints, leading to lytic cell death at later timepoints.

Staurosporine induces time-dependent lytic cell death in multiple cell types

To determine whether the STS-induced lytic cell death was unique to mouse macrophages or was a more conserved mechanism, we treated mouse fibroblasts (L929 cells), human monocyte-derived macrophages, and a human keratinocyte cell line (HaCaT cells) with STS. We observed that STS induced lytic cell death in all three cell types in a time-dependent manner (Fig. 2, A–C). To determine whether the induction of delayed lytic cell death represented an STS-specific cell death mechanism, we treated murine BMDMs with different doses of other apoptotic triggers, including rotenone and etoposide. In contrast to our observations with STS, both rotenone and etoposide only induced low levels (<20%) of lytic cell death at the highest tested concentrations, and almost no cell death at lower concentrations (Fig. 2, D and E). Overall, our data suggest that STS has a unique ability to activate lytic cell death at later timepoints across different mouse and human cell types.Figure 2 Staurosporine induces time-dependent lytic cell death in multiple cell types.A–C, real-time quantification of lytic cell death and the corresponding representative images of cell death in (A) L929 mouse fibroblasts, (B) human macrophages, and (C) HaCaT human keratinocyte cells following treatment with 5 μM staurosporine (STS). D and E, the indicated doses of apoptosis triggers, rotenone (D) and etoposide (E), were used, and lytic cell death was assessed in mouse bone marrow-derived macrophages (BMDMs). Cell death analysis was performed using IncuCyte live cell imaging and staining with propidium iodide (PI). The scale bar measures 25 μm (A–C). The data are shown as the mean ± SD and are representative of at least three independent biological replicates (A–E). Two-way ANOVA with Sidak’s or Tukey’s multiple comparisons test was used to determine the statistical significance. Specific p values are denoted alongside each representative plot for comparisons between the untreated and treated (STS, rotenone, or etoposide) groups. The representative data points were only included once the findings of each experiment were confirmed to be consistent across the biological replicates.

Staurosporine induces time-dependent activation of lytic cell death molecules in immune and non-immune cells

To characterize the nature of STS-induced cell death, we investigated the activation of multiple cell death molecules at early and late time points across different cell types, including murine BMDMs, L929 cells, human macrophages, and HaCaT cells. In BMDMs, we observed robust time-dependent activation of cell death molecules, with caspases-8, -3, and -7 showing the earliest activation, followed by activation of molecules more traditionally associated with lytic cell death (Fig. 3A). Moreover, STS induced a time-dependent increase in the release of DAMPs associated with lytic cell death, such as LDH and HMGB1, in BMDMs (Fig. 3A). Similarly, we observed comparable activation of these cell death components in L929 cells, human macrophages, and HaCaT cells, though there were differences in the specific kinetics (Figure 3, Figure 4, A and B). Overall, the activation of these cell death components further suggests that STS treatment induces lytic cell death in a time-dependent manner.Figure 3 Staurosporine induces time-dependent activation of cell death molecules in murine immune and non-immune cells.A and B, immunoblot analysis for the activation of cell death molecules in mouse bone marrow-derived macrophages (BMDMs) (A) and mouse fibroblast L929 cells (B) treated with 5 μM staurosporine (STS) for the indicated time. Immunoblots for pro- (p45) and activated (p20) caspase-1 (CASP1), pro- (p53) and activated (p30) gasdermin D (GSDMD), pro- (p53) and activated (p34) gasdermin E (GSDME), pro- (p55) and cleaved (p45, p18) caspase-8 (CASP8), pro- (p35) and activated (p17) caspase-3 (CASP3), pro- (p35) and activated (p20) caspase-7 (CASP7), pro- (p47) and activated (p35) caspase-9 (CASP9), phosphorylated and total MLKL (pMLKL and tMLKL), and lytic cell death markers lactate dehydrogenase (LDH) and high mobility group box 1 (HMGB1) are shown. The data are representative of at least three independent biological replicates, with the most representative blot for each protein being selected from across the experiments. Comparable protein loading was verified between experiments by comparing β-actin as the internal control.

Figure 4 Staurosporine induces time-dependent activation of cell death molecules in human immune and non-immune cells.A and B, immunoblot analysis for the activation of cell death molecules in human monocyte-derived macrophages (A) and human keratinocyte (HaCaT) cells (B) treated with 5 μM staurosporine (STS) for the indicated time. Immunoblots for pro- (p45) and activated (p22, p20) caspase-1 (CASP1), pro- (p53) and activated (p30) gasdermin D (GSDMD), pro- (p53) and activated (p34) gasdermin E (GSDME), pro- (p57) and cleaved (p45, p18) caspase-8 (CASP8), pro- (p35) and activated (p19, p17, p15) caspase-3 (CASP3), pro- (p35) and activated (p20) caspase-7 (CASP7), pro- (p47) and activated (p35) caspase-9 (CASP9), phosphorylated and total MLKL (pMLKL and tMLKL), and lytic cell death markers lactate dehydrogenase (LDH) and high mobility group box 1 (HMGB1) are shown. The data are representative of at least three independent biological replicates, with the most representative blot for each protein being selected from across the experiments. Comparable protein loading was verified between experiments by comparing β-actin as the internal control. Red asterisk (∗) denotes a non-specific band.

Staurosporine-induced lytic cell death is PANoptosis, mediated by the caspase-8/RIPK3 axis

To further understand the molecular mechanisms of STS-induced lytic cell death, BMDMs derived from wild-type (WT) and different genetic knockout mice were treated with STS, and the cell death dynamics were examined. We first determined the role of caspases, which are known to be critical regulators and effectors across several cell death mechanisms (Fig. 5, A–F). We observed a small, but statistically significant, reduction in cell death in Casp1−/− and Casp7−/− BMDMs, as well as in BMDMs from mice with a catalytically inactive form of caspase-8 (Casp8DA/DA mice) at the 8 h timepoint (Fig. 5, A, E, and F); however, there is likely limited functional significance for this slight reduction in cell death. Furthermore, there was no significant reduction in cell death in Casp1−/−Casp11−/−, Casp6−/−, and Casp3−/− BMDMs (Fig. 5, B–D).Figure 5 Staurosporine-induced lytic cell death is PANoptosis, mediated by the caspase-8/RIPK3 axis.A–L, real-time quantification of cell death in untreated and 20 μM staurosporine (STS)-treated wild type (WT) and Casp1−/− (A), Casp1−/−Casp11−/− (B), Casp3−/− (C), Casp6−/− (D), Casp7−/− (E), Casp8DA/DA (F), Ripk3−/− (G), Casp7−/−Ripk3−/− (H), Casp8−/−Ripk3−/− (I), Casp8−/−Mlkl−/− (J), Casp1−/−Casp8−/−Ripk3−/− (K), and Casp1−/−Casp8−/−Mlkl−/− (L) bone marrow-derived macrophages (BMDMs) using IncuCyte live cell imaging and propidium iodide (PI) staining, a membrane impermeable dye that is taken up as a marker of lytic cell death. In some cases, a single WT was used to compare against multiple knockout lines inoculated and stimulated at the same time in the same experiment. The data are plotted separately in the figures for optimal visualization; therefore, the same WT cell death curve appears in multiple panels. The data are shown as the mean ± SD and are representative of at least three independent biological replicates (A–L). Two-way ANOVA with Tukey’s multiple comparisons test was used to determine the statistical significance. Specific p values are denoted alongside each representative plot for comparisons between the WT and knockout STS-treated groups. The representative data points were only included once the findings of each experiment were confirmed to be consistent across the biological replicates.

Given the limited protection against STS-induced lytic cell death that we observed in response to the deletion of individual caspases, we next considered other critical PANoptosis molecules. The PANoptosome cell death complex that drives PANoptosis contains caspases and RIPKs, and complex formation induces the activation of executioners and pore-forming molecules that trigger membrane damage and lytic cell death in response to various triggers (3, 32, 34, 38, 39, 40, 41, 53, 54). There is often functional redundancy between the molecules, and the combined loss or inhibition of more than one PANoptosome component or downstream effector is needed to protect cells from undergoing PANoptosis (31, 33, 39, 41). Therefore, we went on to genetically assess the involvement of key PANoptosis regulators in STS-induced lytic cell death. We observed that Ripk3−/− BMDMs did not show a significant reduction in STS-induced lytic cell death (Fig. 5G). However, the combined deletion of caspase-7 and RIPK3 demonstrated partial protection against STS-induced lytic cell death compared with the cell death in WT BMDMs (Fig. 5H), similar to what we observed in Casp7−/− BMDMs (Fig. 5E). In contrast, Casp8−/−Ripk3−/− BMDMs were almost completely protected from STS-induced lytic cell death (Fig. 5I). Similarly, BMDMs from other genetic crosses containing Casp8−/− and deletion of RIPK3 or its downstream executioner MLKL, including Casp8−/−Mlkl−/−, Casp1−/−Casp8−/−Ripk3−/−, and Casp1−/−Casp8−/−Mlkl−/−, were also protected from STS-induced lytic cell death (Fig. 5, J–L). These findings suggest that in response to the canonical apoptosis trigger STS, cells undergo delayed inflammatory, lytic PANoptotic cell death through the caspase-8/RIPK3 axis.

Executioner-deficient cells are partially protected against staurosporine-induced lytic cell death

Since pore-forming cell death executioners, such as gasdermins and MLKL, are downstream effectors for different modes of lytic cell death (3, 22), we next determined their roles in STS-induced inflammatory lytic cell death. Despite the roles of GSDMD and MLKL as the key executioners of the lytic cell death pathways pyroptosis and necroptosis, respectively, and the known role of GSDME in lytic cell death, we observed that Gsdmd−/− and Mlkl−/− BMDMs did not show significant protection from STS-induced lytic cell death even at later timepoints (Fig. 6, A and B), and Gsdme−/− BMDMs were marginally, but statistically significantly, protected (Fig. 6C).Figure 6 Executioner-deficient cells are partially protected against staurosporine-induced lytic cell death.A–F, real-time quantification of cell death in untreated and 20 μM staurosporine (STS)-treated wild type (WT) and Gsdmd−/− (A), Mlkl−/− (B), Gsdme−/− (C), Gsdmd−/−Gsdme−/− (D), Gsdmd−/−Mlkl−/− (E), and Gsdmd−/−Gsdme−/−Mlkl−/− (F) bone marrow-derived macrophages (BMDMs) using IncuCyte live cell imaging and propidium iodide (PI) staining, a membrane impermeable dye that is taken up as a marker of lytic cell death. In some cases, a single WT was used to compare against multiple knockout lines inoculated and stimulated at the same time in the same experiment. The data are plotted separately in the figures for optimal visualization; therefore, the same WT cell death curve appears in multiple panels. The data are shown as the mean ± SD and are representative of at least three independent biological replicates (A–F). Two-way ANOVA with Tukey’s multiple comparisons test was used to determine the statistical significance. Specific p values are denoted alongside each representative plot for comparisons between the WT and knockout STS-treated groups. The representative data points were only included once the findings of each experiment were confirmed to be consistent across the biological replicates.

Cell death executioners can often play functionally redundant roles, which may explain the limited protection from cell death observed upon deletion of a single executioner. Therefore, we also assessed STS-induced cell death upon deletion of multiple cell death executioners. We observed that Gsdmd−/−Gsdme−/− and Gsdmd−/−Mlkl−/− BMDMs were also marginally, but statistically significantly, protected against STS-induced lytic cell death (Fig. 6, D and E). However, the combined deletion of Gsdmd, Gsdme, and Mlkl provided greater protection from STS-induced cell death, though cell death was reduced by only ∼50% (Fig. 6F). Overall, these findings show that loss of the canonical cell death executioners provides only partial protection following STS treatment in a time-dependent manner, suggesting that additional executioners may be involved in driving the membrane damage.

Caspases-1, caspase-8, and RIPK3 drive staurosporine-induced inflammatory lytic cell death, PANoptosis

Given the clear role of STS in mediating lytic inflammatory cell death at later time points, we further used genetic knockouts to understand how disrupting different cell death pathways affected the molecular activation of STS-induced PANoptosis. We used cells with combined deletion of caspase-1 and caspase-11, which would inhibit pyroptosis but not PANoptosis; cells with deletion of caspase-7 alone, which would have reduced apoptosis but not PANoptosis; cells with deletion of RIPK3 alone, which would inhibit necroptosis but not PANoptosis; and cells with combined deletion of caspase-8 and RIPK3 or combined deletion of caspase-1, caspase-8, and RIPK3, which would inhibit PANoptosis. At the later time point (8 h post-STS treatment), we observed that combined deletion of caspase-1 and caspase-11 resulted in a substantial reduction in the cleavage of GSDMD, but not GSDME, when compared with WT BMDMs (Fig. 7). Additionally, cleaved GSDMD and GSDME were absent or markedly reduced in Casp8−/−Ripk3−/− and Casp1−/−Casp8−/−Ripk3−/− BMDMs following STS treatment (Fig. 7). In contrast, Ripk3−/− BMDMs had a partial reduction in GSDMD and GSDME cleavage post-STS treatment. Caspases-3 and -8 were activated at levels similar to those in WT BMDMs in Casp1−/−Casp11−/− and Casp7−/− BMDMs, but not in Casp8−/−Ripk3−/− or Casp1−/−Casp8−/−Ripk3−/− BMDMs (Fig. 7). Moreover, phosphorylation of MLKL following STS treatment was observed in WT, Casp1−/−Casp11−/−, and Casp7−/− BMDMs but was absent in Ripk3−/−, Casp8−/−Ripk3−/−, and Casp1−/−Casp8−/−Ripk3−/− BMDMs (Fig. 7). Additionally, LDH and HMGB1 were released from WT, Casp1−/−Casp11−/−, Casp7−/−, and Ripk3−/− BMDMs, but there was a decrease in their release from Casp8−/−Ripk3−/− and Casp1−/−Casp8−/−Ripk3−/− BMDMs (Fig. 7). Overall, these findings suggest that the caspase-8/RIPK3 axis, which is a crucial component of PANoptosis, is required for the biochemical activation of STS-induced lytic cell death.Figure 7 Caspase-1, caspase-8, and RIPK3 drive staurosporine-inducedlytic cell death, PANoptosis. Immunoblot analysis of pro- (p45) and activated (p20) caspase-1 (CASP1), pro- (p53) and activated (p30) gasdermin D (GSDMD), pro- (p53) and activated (p34) gasdermin E (GSDME), cleaved (p45, p18) caspase-8 (CASP8), pro- (p35) and activated (p17) caspase-3 (CASP3), pro- (p35) and activated (p20) caspase-7 (CASP7), pro- (p47) and activated (p35) caspase-9 (CASP9), phosphorylated and total MLKL (pMLKL and tMLKL), and lytic cell death markers lactate dehydrogenase (LDH) and high mobility group box 1 (HMGB1) are shown from untreated (UT) and 20 μM staurosporine (STS)-treated wild type (WT) and Casp1−/−Casp11−/− (deficient in pyroptosis), Casp7−/− (reduced apoptosis), Ripk3−/− (deficient in necroptosis), Casp8−/−Ripk3−/−, and Casp1−/−Casp8−/−Ripk3−/− (deficient in PANoptosis) bone marrow-derived macrophages (BMDMs) at the 8 h timepoint. The data are representative of at least three independent biological replicates, with the most representative blot for each protein being selected from across the experiments. Comparable protein loading was verified between experiments by comparing β-actin as the internal control. Red asterisk (∗) denotes a non-specific band.

RIPK1 regulates staurosporine-induced PANoptosis

To gain additional insight into the molecular regulation of STS-induced lytic cell death mechanisms, we examined potential upstream regulatory mechanisms. To date, multiple key PANoptosome-inducing molecules have been identified, namely, ZBP1, AIM2, NLRC5 and NLRP12, and RIPK1 (34, 36, 37, 38, 39, 55, 56, 57). Among these, PANoptosis is induced by ZBP1 in response to nucleic acids (36, 37), AIM2 in response to HSV1 or Francisella infections (39), NLRC5 and NLRP12 in response to heme-containing triggers (34, 57), and RIPK1 in the context of TAK1 kinase inhibition (38, 55, 56). Given the role of STS as a broad-spectrum kinase inhibitor, we hypothesized that STS could have an impact on the TAK1-mediated PANoptosis pathway regulated by RIPK1 and TNF signaling. Indeed, we observed that BMDMs deficient in TNF signaling components, such as TNF or TNFR1/2, were significantly protected from cell death in response to STS compared with WT BMDMs (Fig. 8A). We therefore sought to determine the role of RIPK1 as a regulator of STS-mediated PANoptosis. Due to the embryonic lethality of Ripk1 deletion, we evaluated cell death in BMDMs derived from mice with a kinase-dead mutant of RIPK1 (Ripk1KD/KD) following STS treatment. Ripk1KD/KD BMDMs demonstrated a significant reduction in cell death compared with WT BMDMs in response to STS (Fig. 8B). To complement this genetic approach, we also pharmacologically inhibited RIPK1 by treating with Nec1, and we observed that Nec1-treated BMDMs were significantly protected from STS-induced cell death (Fig. 8C). Combined treatment with Nec1 and the pan-caspase inhibitor zVAD did not further reduce cell death (Fig. 8C), suggesting that RIPK1 is the key upstream regulator of this process. Moreover, silencing Ripk1 provided significant protection against STS-induced cell death (Fig. 8D), whereas the combined silencing of Ripk1 and Casp8 did not provide any significant reduction in cell death beyond that achieved by silencing Ripk1 alone (Fig. 8D). Together, these findings provide multiple lines of evidence to suggest that the observed lytic cell death in response to STS is RIPK1-mediated PANoptosis.Figure 8 RIPK1 regulates staurosporine-induced PANoptosis.A–D, real-time quantification of cell death in untreated and 20 μM staurosporine (STS)-treated wild type (WT) bone marrow-derived macrophages (BMDMs) compared with (A) Tnf−/− and Tnfr1−/−Tnfr2−/− STS-treated BMDMs, (B) Ripk1KD/KD STS-treated BMDMs, (C) inhibitor treatments with the RIPK1 inhibitor (Nec1) and/or pan-caspase inhibitor (zVAD), and (D) WT BMDMs transfected with siRipk1, siCasp8, or both. The data are shown as the mean ± SD and are representative of at least two independent biological replicates. Two-way ANOVA with Tukey’s multiple comparisons test was used to determine the statistical significance. Specific p values are denoted alongside each representative plot for comparisons between the WT versus knockout STS-treated groups (A and B), or STS-treated versus STS + inhibitor-treated groups (C), or siControl versus siRipk1 and/or siCasp8 STS-treated groups (D). The representative data points were only included once the findings of each experiment were confirmed to be consistent across the biological replicates.

Discussion

Cell death is known to play an important role in disease progression and therapeutic outcomes (1, 2, 3, 58). Furthermore, both non-lytic (apoptotic) and lytic (pyroptosis, necroptosis, and PANoptosis) cell death pathways that involve caspases can be targeted through treatment strategies to improve patient outcomes. Apoptosis is a form of cell death that does not compromise cell membrane integrity or release cellular DAMPs, thereby preventing downstream inflammatory responses (5, 6, 44). STS, an ATP-competitive kinase inhibitor (59), has a high affinity for a variety of kinases, but it is particularly effective against STE20 family kinases, which include those in the MST, GCK, and PAK subfamilies (45, 48, 59, 60, 61). While STS is often utilized to trigger rapid, non-lytic apoptotic cell death and is employed in clinical settings to treat cancer and other disorders, its specific targets to activate cell death and its role in other lytic cell death pathways are unknown.

While non-lytic apoptosis is marked by the activation of executioner caspases-3 and -7 (7, 12, 13, 14, 15, 16), lytic cell death is characterized by membrane damage and the activation of various cell death molecules beyond caspases (3). Here, we reported that, in addition to its known ability to induce non-lytic apoptosis, STS can also induce RIPK1-mediated lytic cell death, PANoptosis, in a time- and dose-dependent manner through the caspase-8/RIPK3 axis. Moreover, cells deficient in components of pyroptosis, apoptosis, or necroptosis alone were not protected from the biochemical activation of cell death or the execution of cell death, and the deletion of PANoptotic components was required for rescue. Together, these findings provide multiple lines of evidence to show that the observed lytic cell death was not pyroptosis, necroptosis, or secondary necrosis but was instead the genetically controlled activation of PANoptosis. Additionally, our findings highlight the molecular crosstalk that occurs during PANoptosis. For instance, in response to STS, the activation of caspase-1 was dependent on caspase-8, and the activation of caspase-3 and GSDME was decreased in RIPK3-deficient cells. Our results also suggest a critical role for the TNF signaling axis. Innate immune activation through Toll-like receptors can induce the production of TNF, and TNF in combination with PAMPs or DAMPs is known to induce PANoptosis through multiple innate immune sensors (34, 38, 55, 56, 57). Our data suggest a specific role for RIPK1 in mediating STS-induced PANoptosis, though the specific upstream mechanisms through which STS can induce this lytic cell death require further investigation. Additionally, the combined deletion of GSDMD, GSDME, and MLKL, the most well-characterized pore-forming molecules, only provided partial protection from STS-mediated lytic cell death at later timepoints. Functional redundancy has increasingly been identified among these pore-forming molecules (58, 62), and the lack of full protection in Gsdmd−/−Gsdme−/−Mlkl−/− cells suggests that additional gasdermins or other pore-forming molecules also play a role in executing PANoptosis in response to STS (63).

While preclinical research investigating STS as an anti-cancer treatment has shown that it is generally ineffective due to its poor target specificity, the literature suggests that the effects of low-dose STS in various diseases are driven by apoptosis (64, 65, 66). Additionally, STS and its analogs have been used in clinical settings to treat non-small cell lung cancer and other advanced solid malignancies (67, 68, 69). However, their effectiveness is limited by a lack of specificity and the administration of low doses during treatment. Therefore, our findings that STS can also induce lytic cell death through PANoptosis suggest that it may be beneficial in scenarios where STS has previously not been considered, or in situations where low dose STS proved ineffective. Furthermore, in the cellular microenvironment, the local concentration of STS may be higher than the clinical dose administered, and PANoptosis could play a key role in that context. Additionally, several STS derivatives are also under preclinical and clinical investigation (64, 67, 68, 69, 70, 71, 72, 73) and may also have utility beyond the induction of apoptosis, which warrants further investigation.

Overall, our findings characterized the time-dependent induction of lytic cell death in response to the classical apoptotic trigger, STS. Leveraging the ability of STS to drive both non-lytic and lytic forms of cell death may have implications for inflammatory signaling and trigger a broader immune response. This immune activation is critical for pathogen or aberrant cell clearance but must be tightly regulated to prevent pathophysiology. Therefore, the time-dependent activation of lytic cell death, PANoptosis, by STS suggests new avenues for targeting PANoptosis components in treating diseases.

Experimental procedures

Mice

Wild-type (WT) C57/Bl6, Casp1−/− (74), Casp1−/−Casp11−/− (17), Casp3−/− (75), Casp6−/− (Jackson Laboratory, Cat# 006236), Casp7−/− (76), Casp8DA/DA (77), Ripk3−/− (78), Casp7−/−Ripk3−/− (63), Casp8−/−Ripk3−/− (79), Casp1−/−Casp8−/−Ripk3−/− (34), Gsdmd−/− (80), Gsdme−/− (81), Mlkl−/− (82), Gsdmd−/−Gsdme−/− (54), Gsdmd−/−Mlkl−/− (37), Tnf−/− (83), Tnfr1−/−Tnfr2−/− (84), Ripk1KD/KD (56), and Gsdmd−/−Gsdme−/−Mlkl−/− (41) mice used in the study have been described previously. Casp8−/−Mlkl−/− mice were generated by crossing Casp8−/−Ripk3−/− (79) and Mlkl−/− (82) mice and selecting for Casp8−/−Mlkl−/−Ripk3+/+ mice. Casp1−/−Casp8−/−Mlkl−/− mice were generated by crossing Casp8−/−Mlkl−/− mice with Casp1−/− (74) mice. All mice were housed and bred at the St Jude Children’s Research Hospital animal facility under specific pathogen-free conditions with aseptic handling techniques. Animal studies were conducted under protocols approved by St Jude Children’s Research Hospital’s committee on the use and care of animals.

Mouse bone marrow-derived macrophage and human blood monocyte-derived macrophage differentiation and culture

Primary bone marrow-derived macrophages (BMDMs) were cultured for 6 days in Iscove’s modified Dulbecco’s medium (Thermo Fisher Scientific, 12440-053) supplemented with 10% heat-inactivated fetal bovine serum (HI-FBS) (Biowest, S1620), 30% L929-conditioned medium, 1% non-essential amino acids (Thermo Fisher Scientific, 11140-050), and 1% penicillin and streptomycin (Thermo Fisher Scientific, 15070-063). The BMDMs were counted and seeded into 12-well plates at a density of 1 × 106 cells/well or 24-well plates at a density of 0.5 × 106 cells/well and incubated in a humidified incubator supplied with 5% CO2 overnight before use.

For human macrophages, fresh human blood was collected from the apheresis rings of anonymous, healthy blood donors from the blood bank at St Jude Children’s Research Hospital obtained through approval of the St Jude Institutional Review Board (IRB) in protocols consistent with the Declaration of Helsinki. Peripheral blood mononuclear cells (PBMCs) were isolated from the freshly collected blood using a lymphoprep solution (Stemcell Technologies, cat. #07801/07811). Naive monocytes were purified from the PBMCs using a monocyte isolation kit (EasySep direct human monocyte isolation kit, Stemcell Technologies, cat. #19669) strictly in accordance with the manufacturer’s protocol. These purified monocytes were further differentiated into monocyte-derived macrophages by culturing them in RPMI media supplemented with 10% HI-FBS, 1% penicillin-streptomycin, and 25 ng/ml human M-CSF (Peprotech, 300-25) for 6 days in a humidified incubator supplied with 5% CO2. On days 2 and 4, an additional 8 to 10 ml of similar media containing 25 ng/ml human M-CSF was added to the cells. On day 6, all the loosely attached and suspension cells were harvested and washed three times with PBS. The cells were resuspended in complete RPMI media at a cell density of 1 × 106 cells/ml containing 25 ng/ml human M-CSF. These cells were further used for downstream experiments.

Mouse fibroblast and human keratinocyte culture

Mouse fibroblast (L929 cells) and human keratinocyte (HaCaT cells) cell lines were originally obtained from ATCC and cultured and maintained in DMEM and RPMI, respectively, and supplemented with 10% HI-FBS and 1% penicillin-streptomycin in a humidified incubator supplied with 5% CO2. Cells were monitored for morphology, and protein expression was assessed by western blotting; no additional authentication was performed.

siRNA transfection in BMDMs

For siRNA transfection, BMDMs were washed and resuspended at a cell density of 1 × 107 cells/ml. Five pmoles of non-targeting control siRNA (D-001206-14-20, Horizon Discovery), mouse-specific Ripk1 siRNA (M-040150-00-0005, Horizon Discovery), and/or mouse-specific Casp8 siRNA (M-043044-01-0005, Horizon Discovery) per million cells were used for transfection by electroporation (Neon Transfection System kit, Thermo Fisher Scientific, cat. #MPK5000). The transfected cells were then seeded in 24-well tissue culture plates at a cell density of 0.5 × 106 cells/ml in Iscove’s modified Dulbecco’s medium supplemented with 10% HI-FBS, 30% L929-conditioned medium, 1% non-essential amino acids, and 1% penicillin and streptomycin. After overnight culture, the medium was replaced with fresh media. After 48 h of rest, the seeded transfected cells were treated with 20 μM of STS and analyzed for real-time cell death.

IncuCyte cell death analysis

The kinetics of cell death were measured using the IncuCyte S3 or SX5 imaging system (Sartorius). BMDMs and human macrophages were seeded in 12-well (1 × 106 cells/well) or 24-well (0.5 × 106 cells/well) cell culture plates. L929 cells and HaCaT cells were seeded at a density of 0.3 × 106 cells/well in 12-well plates or 0.15 × 106 cells/well in 24-well plates. The cells were treated with the indicated apoptosis triggers, including STS (Sigma Aldrich, S5921), etoposide (Sigma Aldrich, E1383), or rotenone (Sigma Aldrich, R8875), in the presence of the cell-impermeable DNA-binding fluorescent dye Sytox green (Life Technologies, S7020) or propidium iodide (PI, Life Technologies, P3566) solution. For the pharmacological inhibition of proteins, BMDMs were pre-treated for 1 h with the RIPK1 inhibitor necrostatin 1 (Nec1; Selleckchem, S8641), and/or the caspase inhibitor zVAD-FMK (zVAD; Apexbio, A1902) prior to STS stimulation in the presence of cell impermeable DNA-binding dye, PI. Cell permeable activated caspase-3/7 red dye (Sartorius, 4704, 1:1000) was used to detect and quantify cells with activated caspase-3/7 in real time. A series of images were acquired at 1 h time intervals for up to 8 h post-treatment with a 20 × objective and analyzed using the IncuCyte S3 or SX5 software, which enables precise analysis of the number of dye-positive cells present in each image. A minimum of four images per well were captured for the analysis of each timepoint. To assess Annexin V binding, BMDMs were seeded as previously described. The BMDMs were treated with IncuCyte Annexin V NIR Dye (Sartorius, 4768) along with cell impermeable DNA binding dyes, Sytox green or PI, in the presence of 5 mM calcium chloride (Sigma-Aldrich, C-34006) in complete DMEM media supplemented with 10% HI-FBS and 1% penicillin-streptomycin. A series of images were acquired for analysis.

Immunoblotting

For the immunoblotting analysis of caspases and other cell death markers, BMDMs and human macrophages were seeded a day before stimulation at a density of 1 × 106 cells/well, and L929 cells and HaCaT cells were seeded at a density of 0.3 × 106 cells/well in 12-well plates. For caspase immunoblotting analysis, the proteins from the indicated cell types were collected by combining cell lysates with culture supernatants in caspase lysis buffer (with 1 × protease inhibitor, 1 × phosphatase inhibitor, 10% NP-40, and 25 mM DTT) and 4 × sample loading buffer (containing SDS and 2-mercaptoethanol). For the immunoblotting of all other signaling proteins, the supernatant was collected and centrifuged to pellet cellular debris. The remaining adherent cells in the plate and cellular debris were lysed separately in RIPA buffer supplemented with protease inhibitors (Thermo Scientific, A32965) and phosphoStop (Roche, PHOSS-RO), according to the manufacturer’s instructions. These lysates were then combined with the sample loading buffer. To analyze LDH and HMGB1 release, the supernatants alone were combined with 4 × sample loading buffer. All samples were boiled at 100 °C for 10 min prior to loading onto gels and separated using SDS-PAGE, followed by transfer onto Amersham Hybond P polyvinylidene difluoride membranes (GE Healthcare Life Sciences, 10600023). All primary antibodies used in this study are commercially available and have been validated by the company or in house using the appropriate knockout controls. Antibodies were used for anti-caspase-1 (AdipoGen, AG-20B-0042, 1:2000), human anti-caspase-1 (Cell Signaling Technologies (CST), 3866, 1:2000), human anti-cleaved caspase-1 (CST, 4199, 1:2000), anti-caspase-3 (CST, 9662, 1:1000), anti-cleaved caspase-3 (CST, 9661, 1:1000), anti-caspase-7 (CST, 9492, 1:1000), anti-cleaved caspase-7 (CST, 9491, 1:1000), anti-caspase-8 (CST, 4927, 1:1000), anti-cleaved caspase-8 (CST, 8592, 1:1000), human anti-caspase-8 (Enzo Life Science, ALX-804-242, 1:1000), anti-caspase-9 (CST, 9504, 1:1000), anti-pMLKL (CST, 37333, 1:1000), human anti-pMLKL (Abcam, ab187091 (S-358), 1:1000), anti-GSDMD (Abcam, ab209845, 1:1000), human anti-GSDMD (Novus, NBP2-33422, 1:2000), anti-GSDME (Abcam, ab215191, 1:1000), anti-MLKL (Abgent, AP14272b, 1:1000), human anti-MLKL (Sigma, M6697, 1:1000), anti-LDHA (Proteintech, 19987-1-AP, 1:1000), anti-HMGB1 (Abcam, ab18256, 1:1000), anti-β-actin (Proteintech, 66009–1-IG, 1:5000), and HRP-conjugated secondary antibodies (Jackson ImmunoResearch Laboratories, anti-rabbit (111-035-047), 1:5000; and anti-mouse (315-035-047), 1:5000).

Statistical analysis

The GraphPad Prism software (v10) was used for the data analysis. Data are shown as the mean ± SD. Statistical significance was determined by a Two-way ANOVA analysis and Šidák or Tukey’s multiple comparisons test for all the cell death kinetics results. The number of experimental replicates is indicated in the corresponding figure legends. Specific p values are denoted alongside each representative plot in the figures.

Data availability

All datasets generated and analyzed in this study are provided within the manuscript and the accompanying figures.

Supporting information

This article contains supporting information.

Conflict of interests

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests

T.-D. K. was a consultant for Pfizer.

Supporting information

Supplemental legend

Supplemental Figure

Acknowledgments

We thank all the members of the Kanneganti laboratory for their scientific input, comments, and suggestions during the development of the manuscript. We thank Anu Sharma, PhD, and Rebecca Tweedell, PhD, for scientific editing and writing support, and Katie Thorne and Lauren Kneeland for mouse colony support.

Author contributions

R. S. and T.-D. K. conceptualization; R. S. and S. M. C. methodology; R. S. and S. M. C. investigation; R. S. and S. M. C. formal analysis; R. S. writing–original draft; R. S., S. M. C., and T-D. K. writing–review & editing; T-D. K. funding acquisition; T-D. K. supervision.

Funding and additional information

Work from our laboratory is supported by the 10.13039/100000002 US National Institutes of Health (AI101935 , AI124346 , AI160179 , AR056296 , and CA253095 to T.-D.K.) and the American Lebanese Syrian Associated Charities (to T.-D.K.). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.
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References

1 Kanneganti T.D. Intracellular innate immune receptors: life inside the cell Immunol. Rev. 297 2020 5 12 32856334
2 Christgen S. Kanneganti T.D. Inflammasomes and the fine line between defense and disease Curr. Opin. Immunol. 62 2020 39 44 31837596
3 Chen W. Gullett J.M. Tweedell R.E. Kanneganti T.D. Innate immune inflammatory cell death: PANoptosis and PANoptosomes in host defense and disease Eur. J. Immunol. 53 2023 e2250235
4 Kumar H. Kawai T. Akira S. Pathogen recognition by the innate immune system Int. Rev. Immunol. 30 2011 16 34 21235323
5 Kerr J.F. Wyllie A.H. Currie A.R. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics Br. J. Cancer 26 1972 239 257 4561027
6 Wyllie A.H. Kerr J.F. Currie A.R. Cell death: the significance of apoptosis Int. Rev. Cytol. 68 1980 251 306 7014501
7 Schwartzman R.A. Cidlowski J.A. Apoptosis: the biochemistry and molecular biology of programmed cell death Endocr. Rev. 14 1993 133 151 8325248
8 Pan G. O'Rourke K. Dixit V.M. Caspase-9, Bcl-XL, and Apaf-1 form a ternary complex J. Biol. Chem. 273 1998 5841 5845 9488720
9 Alnemri E.S. Livingston D.J. Nicholson D.W. Salvesen G. Thornberry N.A. Wong W.W. Human ICE/CED-3 protease nomenclature Cell 87 1996 171 8861900
10 Li P. Nijhawan D. Budihardjo I. Srinivasula S.M. Ahmad M. Alnemri E.S. Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade Cell 91 1997 479 489 9390557
11 Varfolomeev E.E. Schuchmann M. Luria V. Chiannilkulchai N. Beckmann J.S. Mett I.L. Targeted disruption of the mouse Caspase 8 gene ablates cell death induction by the TNF receptors, Fas/Apo1, and DR3 and is lethal prenatally Immunity 9 1998 267 276 9729047
12 Enari M. Talanian R.V. Wong W.W. Nagata S. Sequential activation of ICE-like and CPP32-like proteases during Fas-mediated apoptosis Nature 380 1996 723 726 8614469
13 Hasegawa J. Kamada S. Kamiike W. Shimizu S. Imazu T. Matsuda H. Involvement of CPP32/Yama(-like) proteases in Fas-mediated apoptosis Cancer Res. 56 1996 1713 1718 8620480
14 Schlegel J. Peters I. Orrenius S. Miller D.K. Thornberry N.A. Yamin T.T. CPP32/apopain is a key interleukin 1 beta converting enzyme-like protease involved in Fas-mediated apoptosis J. Biol. Chem. 271 1996 1841 1844 8567626
15 Fernandes-Alnemri T. Takahashi A. Armstrong R. Krebs J. Fritz L. Tomaselli K.J. Mch3, a novel human apoptotic cysteine protease highly related to CPP32 Cancer Res. 55 1995 6045 6052 8521391
16 Kesavardhana S. Malireddi R.K.S. Kanneganti T.D. Caspases in cell death, inflammation, and pyroptosis Annu. Rev. Immunol. 38 2020 567 595 32017655
17 Kayagaki N. Warming S. Lamkanfi M. Vande Walle L. Louie S. Dong J. Non-canonical inflammasome activation targets caspase-11 Nature 479 2011 117 121 22002608
18 Shi J. Zhao Y. Wang K. Shi X. Wang Y. Huang H. Cleavage of GSDMD by inflammatory caspases determines pyroptotic cell death Nature 526 2015 660 665 26375003
19 He W.T. Wan H. Hu L. Chen P. Wang X. Huang Z. Gasdermin D is an executor of pyroptosis and required for interleukin-1beta secretion Cell Res. 25 2015 1285 1298 26611636
20 Aglietti R.A. Estevez A. Gupta A. Ramirez M.G. Liu P.S. Kayagaki N. GsdmD p30 elicited by caspase-11 during pyroptosis forms pores in membranes Proc. Natl. Acad. Sci. U. S. A. 113 2016 7858 7863 27339137
21 Sborgi L. Ruhl S. Mulvihill E. Pipercevic J. Heilig R. Stahlberg H. GSDMD membrane pore formation constitutes the mechanism of pyroptotic cell death EMBO J. 35 2016 1766 1778 27418190
22 Man S.M. Karki R. Kanneganti T.D. Molecular mechanisms and functions of pyroptosis, inflammatory caspases and inflammasomes in infectious diseases Immunol. Rev. 277 2017 61 75 28462526
23 Cookson B.T. Brennan M.A. Pro-inflammatory programmed cell death Trends Microbiol. 9 2001 113 114 11303500
24 Martinon F. Burns K. Tschopp J. The inflammasome: a molecular platform triggering activation of inflammatory caspases and processing of proIL-beta Mol. Cell 10 2002 417 426 12191486
25 Degterev A. Huang Z. Boyce M. Li Y. Jagtap P. Mizushima N. Chemical inhibitor of nonapoptotic cell death with therapeutic potential for ischemic brain injury Nat. Chem. Biol. 1 2005 112 119 16408008
26 Hitomi J. Christofferson D.E. Ng A. Yao J. Degterev A. Xavier R.J. Identification of a molecular signaling network that regulates a cellular necrotic cell death pathway Cell 135 2008 1311 1323 19109899
27 Zhao J. Jitkaew S. Cai Z. Choksi S. Li Q. Luo J. Mixed lineage kinase domain-like is a key receptor interacting protein 3 downstream component of TNF-induced necrosis Proc. Natl. Acad. Sci. U. S. A. 109 2012 5322 5327 22421439
28 Sun L. Wang H. Wang Z. He S. Chen S. Liao D. Mixed lineage kinase domain-like protein mediates necrosis signaling downstream of RIP3 kinase Cell 148 2012 213 227 22265413
29 Galluzzi L. Kepp O. Chan F.K. Kroemer G. Necroptosis: mechanisms and relevance to disease Annu. Rev. Pathol. 12 2017 103 130 27959630
30 Dhuriya Y.K. Sharma D. Necroptosis: a regulated inflammatory mode of cell death J. Neuroinflammation 15 2018 199 29980212
31 Lukens J.R. Gurung P. Vogel P. Johnson G.R. Carter R.A. McGoldrick D.J. Dietary modulation of the microbiome affects autoinflammatory disease Nature 516 2014 246 249 25274309
32 Karki R. Lee S. Mall R. Pandian N. Wang Y. Sharma B.R. ZBP1-dependent inflammatory cell death, PANoptosis, and cytokine storm disrupt IFN therapeutic efficacy during coronavirus infection Sci. Immunol. 7 2022 eabo6294
33 Gurung P. Burton A. Kanneganti T.D. NLRP3 inflammasome plays a redundant role with caspase 8 to promote IL-1beta-mediated osteomyelitis Proc. Natl. Acad. Sci. U. S. A. 113 2016 4452 4457 27071119
34 Sundaram B. Pandian N. Mall R. Wang Y. Sarkar R. Kim H.J. NLRP12-PANoptosome activates PANoptosis and pathology in response to heme and PAMPs Cell 186 2023 2783 2801 37267949
35 Messaoud-Nacer Y. Culerier E. Rose S. Maillet I. Rouxel N. Briault S. STING agonist diABZI induces PANoptosis and DNA mediated acute respiratory distress syndrome (ARDS) Cell Death Dis. 13 2022 269 35338116
36 Kuriakose T. Man S.M. Malireddi R.K. Karki R. Kesavardhana S. Place D.E. ZBP1/DAI is an innate sensor of influenza virus triggering the NLRP3 inflammasome and programmed cell death pathways Sci. Immunol. 1 2016 aag2045
37 Christgen S. Zheng M. Kesavardhana S. Karki R. Malireddi R.K.S. Banoth B. Identification of the PANoptosome: a molecular platform triggering pyroptosis, apoptosis, and necroptosis (PANoptosis) Front. Cell Infect. Microbiol. 10 2020 237 32547960
38 Malireddi R.K.S. Kesavardhana S. Karki R. Kancharana B. Burton A.R. Kanneganti T.D. RIPK1 distinctly regulates yersinia-induced inflammatory cell death, PANoptosis Immunohorizons 4 2020 789 796 33310881
39 Lee S. Karki R. Wang Y. Nguyen L.N. Kalathur R.C. Kanneganti T.D. AIM2 forms a complex with pyrin and ZBP1 to drive PANoptosis and host defence Nature 597 2021 415 419 34471287
40 Malireddi R.K.S. Karki R. Sundaram B. Kancharana B. Lee S. Samir P. Inflammatory cell death, PANoptosis, mediated by cytokines in diverse cancer lineages inhibits tumor growth Immunohorizons 5 2021 568 580 34290111
41 Karki R. Sharma B.R. Tuladhar S. Williams E.P. Zalduondo L. Samir P. Synergism of TNF-alpha and IFN-gamma triggers inflammatory cell death, tissue damage, and mortality in SARS-CoV-2 infection and cytokine shock syndromes Cell 184 2021 149 168 33278357
42 Vitale I. Pietrocola F. Guilbaud E. Aaronson S.A. Abrams J.M. Adam D. Apoptotic cell death in disease-Current understanding of the NCCD 2023 Cell Death Differ. 30 2023 1097 1154 37100955
43 Mangalmurti A. Lukens J.R. How neurons die in Alzheimer's disease: Implications for neuroinflammation Curr. Opin. Neurobiol. 75 2022 102575
44 Fink S.L. Cookson B.T. Apoptosis, pyroptosis, and necrosis: mechanistic description of dead and dying eukaryotic cells Infect Immun. 73 2005 1907 1916 15784530
45 Yadav S.S. Prasad C.B. Prasad S.B. Pandey L.K. Singh S. Pradhan S. Anti-tumor activity of staurosporine in the tumor microenvironment of cervical cancer: an in vitro study Life Sci. 133 2015 21 28 26006035
46 Zhang J. Kinoh H. Hespel L. Liu X. Quader S. Martin J. Effective treatment of drug resistant recurrent breast tumors harboring cancer stem-like cells by staurosporine/epirubicin co-loaded polymeric micelles J. Control Release 264 2017 127 135 28842317
47 Zhou N. Wang R. Zhang Y. Lei Z. Zhang X. Hu R. Staurosporine induced apoptosis may activate cancer stem-like cells (CD44(+)/CD24(-)) in MCF-7 by upregulating Mucin1 and EpCAM J. Cancer 6 2015 1049 1057 26366219
48 Emanuelsson I. Norlin M. Protective effects of 27- and 24-hydroxycholesterol against staurosporine-induced cell death in undifferentiated neuroblastoma SH-SY5Y cells Neurosci. Lett. 525 2012 44 48 22884615
49 Moela P. Choene M.M. Motadi L.R. Silencing RBBP6 (Retinoblastoma Binding Protein 6) sensitises breast cancer cells MCF7 to staurosporine and camptothecin-induced cell death Immunobiology 219 2014 593 601 24703106
50 Krumschnabel G. Maehr T. Nawaz M. Schwarzbaum P.J. Manzl C. Staurosporine-induced cell death in salmonid cells: the role of apoptotic volume decrease, ion fluxes and MAP kinase signaling Apoptosis 12 2007 1755 1768 17624593
51 Yin J. Howe J. Tan K.S.W. Staurosporine-induced programmed cell death in Blastocystis occurs independently of caspases and cathepsins and is augmented by calpain inhibition Microbiology (Reading) 156 2010 1284 1293 20056704
52 Choudhury S.M. Sarkar R. Karki R. Kanneganti T.D. A comparative study of apoptosis, pyroptosis, necroptosis, and PANoptosis components in mouse and human cells PLoS One 19 2024 e0299577
53 Zheng M.K.,R. Vogel P. Kanneganti T.D. Caspase-6 is a key regulator of innate immunity, inflammasome activation and host defense Cell 181 2020 674 687.e613 32298652
54 Karki R. Sundaram B. Sharma B.R. Lee S. Malireddi R.K.S. Nguyen L.N. ADAR1 restricts ZBP1-mediated immune response and PANoptosis to promote tumorigenesis Cell Rep. 37 2021 109858
55 Malireddi R.K.S. Gurung P. Mavuluri J. Dasari T.K. Klco J.M. Chi H. TAK1 restricts spontaneous NLRP3 activation and cell death to control myeloid proliferation J. Exp. Med. 215 2018 1023 1034 29500178
56 Malireddi R.K.S. Gurung P. Kesavardhana S. Samir P. Burton A. Mummareddy H. Innate immune priming in the absence of TAK1 drives RIPK1 kinase activity-independent pyroptosis, apoptosis, necroptosis, and inflammatory disease J. Exp. Med. 217 2020 jem.20191644
57 Sundaram B. Pandian N. Kim H.J. Abdelaal H.M. Mall R. Indari O. NLRC5 senses NAD(+) depletion, forming a PANoptosome and driving PANoptosis and inflammation Cell 187 2024 4061 4077 38878777
58 Pandeya A. Kanneganti T.D. Therapeutic potential of PANoptosis: innate sensors, inflammasomes, and RIPKs in PANoptosomes Trends Mol. Med. 30 2024 74 88 37977994
59 Tanramluk D. Schreyer A. Pitt W.R. Blundell T.L. On the origins of enzyme inhibitor selectivity and promiscuity: a case study of protein kinase binding to staurosporine Chem. Biol. Drug Des. 74 2009 16 24 19519740
60 Seo S.R. Seo J.T. Calcium overload is essential for the acceleration of staurosporine-induced cell death following neuronal differentiation in PC12 cells Exp. Mol. Med. 41 2009 269 276 19299916
61 Rudolph J. Crawford J.J. Hoeflich K.P. Chernoff J. Chapter seven - p21-activated kinase inhibitors Tamanoi F. Der C.J. The Enzymes 2013 Academic Press Amsterdam, The Netherlands 157 180
62 Malireddi R.K.S. Sharma B.R. Bynigeri R.R. Wang Y. Lu J. Kanneganti T.D. ZBP1 drives IAV-induced NLRP3 inflammasome activation and lytic cell death, PANoptosis, independent of the necroptosis executioner MLKL Viruses 15 2023 2141 38005819
63 Han J.H. Karki R. Malireddi R.K.S. Mall R. Sarkar R. Sharma B.R. NINJ1 mediates inflammatory cell death, PANoptosis, and lethality during infection conditions and heat stress Nat. Commun. 15 2024 1739 38409108
64 Malsy M. Bitzinger D. Graf B. Bundscherer A. Staurosporine induces apoptosis in pancreatic carcinoma cells PaTu 8988t and Panc-1 via the intrinsic signaling pathway Eur. J. Med. Res. 24 2019 5 30686270
65 Ma D. Wang P. Fang Q. Yu Z. Zhou Z. He Z. Low-dose staurosporine selectively reverses BCR-ABL-independent IM resistance through PKC-alpha-mediated G2/M phase arrest in chronic myeloid leukaemia Artif. Cells Nanomed. Biotechnol. 46 2018 S208 S216 30618318
66 Bernard B. Fest T. Pretet J.L. Mougin C. Staurosporine-induced apoptosis of HPV positive and negative human cervical cancer cells from different points in the cell cycle Cell Death Differ. 8 2001 234 244 11319606
67 Monnerat C. Henriksson R. Le Chevalier T. Novello S. Berthaud P. Faivre S. Phase I study of PKC412 (N-benzoyl-staurosporine), a novel oral protein kinase C inhibitor, combined with gemcitabine and cisplatin in patients with non-small-cell lung cancer Ann. Oncol. 15 2004 316 323 14760128
68 Eder J.P. Jr. Garcia-Carbonero R. Clark J.W. Supko J.G. Puchalski T.A. Ryan D.P. A phase I trial of daily oral 4'- N -benzoyl-staurosporine in combination with protracted continuous infusion 5-fluorouracil in patients with advanced solid malignancies Invest. New Drugs 22 2004 139 150 14739662
69 Sparreboom A. Chen H. Acharya M.R. Senderowicz A.M. Messmann R.A. Kuwabara T. Effects of alpha1-acid glycoprotein on the clinical pharmacokinetics of 7-hydroxystaurosporine Clin. Cancer Res. 10 2004 6840 6846 15501960
70 Zambrano J.N. Williams C.J. Williams C.B. Hedgepeth L. Burger P. Dilday T. Staurosporine, an inhibitor of hormonally up-regulated neu-associated kinase Oncotarget 9 2018 35962 35973 30542510
71 Sausville E.A. Lush R.D. Headlee D. Smith A.C. Figg W.D. Arbuck S.G. Clinical pharmacology of UCN-01: initial observations and comparison to preclinical models Cancer Chemother. Pharmacol. 42 Suppl 1998 S54 S59 9750030
72 Propper D.J. McDonald A.C. Man A. Thavasu P. Balkwill F. Braybrooke J.P. Phase I and pharmacokinetic study of PKC412, an inhibitor of protein kinase C J. Clin. Oncol. 19 2001 1485 1492 11230495
73 Omura S. Asami Y. Crump A. Staurosporine: new lease of life for parent compound of today's novel and highly successful anti-cancer drugs J. Antibiot. 71 2018 688 701
74 Man S.M. Karki R. Sasai M. Place D.E. Kesavardhana S. Temirov J. IRGB10 Liberates bacterial Ligands for sensing by the AIM2 and caspase-11-NLRP3 inflammasomes Cell 167 2016 382 396 27693356
75 Zheng T.S. Hunot S. Kuida K. Momoi T. Srinivasan A. Nicholson D.W. Deficiency in caspase-9 or caspase-3 induces compensatory caspase activation Nat. Med. 6 2000 1241 1247 11062535
76 Lakhani S.A. Masud A. Kuida K. Porter G.A. Jr. Booth C.J. Mehal W.Z. Caspases 3 and 7: key mediators of mitochondrial events of apoptosis Science 311 2006 847 851 16469926
77 Wang Y. Karki R. Zheng M. Kancharana B. Lee S. Kesavardhana S. Cutting Edge: caspase-8 is a Linchpin in caspase-3 and gasdermin D activation to control cell death, cytokine release, and host defense during influenza A virus infection J. Immunol. 207 2021 2411 2416 34663620
78 Newton K. Sun X. Dixit V.M. Kinase RIP3 is dispensable for normal NF-kappa Bs, signaling by the B-cell and T-cell receptors, tumor necrosis factor receptor 1, and Toll-like receptors 2 and 4 Mol. Cell Biol. 24 2004 1464 1469 14749364
79 Oberst A. Dillon C.P. Weinlich R. McCormick L.L. Fitzgerald P. Pop C. Catalytic activity of the caspase-8-FLIP(L) complex inhibits RIPK3-dependent necrosis Nature 471 2011 363 367 21368763
80 Karki R. Lee E. Place D. Samir P. Mavuluri J. Sharma B.R. IRF8 regulates Transcription of Naips for NLRC4 inflammasome activation Cell 173 2018 920 933 29576451
81 Skarnes W.C. Rosen B. West A.P. Koutsourakis M. Bushell W. Iyer V. A conditional knockout resource for the genome-wide study of mouse gene function Nature 474 2011 337 342 21677750
82 Murphy J.M. Czabotar P.E. Hildebrand J.M. Lucet I.S. Zhang J.G. Alvarez-Diaz S. The pseudokinase MLKL mediates necroptosis via a molecular switch mechanism Immunity 39 2013 443 453 24012422
83 Pasparakis M. Alexopoulou L. Episkopou V. Kollias G. Immune and inflammatory responses in TNF alpha-deficient mice: a critical requirement for TNF alpha in the formation of primary B cell follicles, follicular dendritic cell networks and germinal centers, and in the maturation of the humoral immune response J. Exp. Med. 184 1996 1397 1411 8879212
84 Peschon J.J. Torrance D.S. Stocking K.L. Glaccum M.B. Otten C. Willis C.R. TNF receptor-deficient mice reveal divergent roles for p55 and p75 in several models of inflammation J. Immunol. 160 1998 943 952 9551933
