Suppressed translation as a mechanism of initiation of CASP8 (caspase 8)-dependent apoptosis in autophagy-deficient NSCLC cells under nutrient limitation

ABSTRACT

Macroautophagy/autophagy inhibition under stress conditions is often associated with increased cell death. We found that under nutrient limitation, activation of CASP8/caspase-8 was significantly increased in autophagy-deficient lung cancer cells, which precedes mitochondria outer membrane permeabilization (MOMP), CYCS/cytochrome c release, and activation of CASP9/caspase-9, indicating that under such conditions the activation of CASP8 is a primary event in the initiation of apoptosis as well as essential to reduce clonogenic survival of autophagy-deficient cells. Starvation leads to suppression of CFLAR proteosynthesis and accumulation of CASP8 in SQSTM1 puncta. Overexpression of CFLARs reduces CASP8 activation and apoptosis during starvation, while its silencing promotes efficient activation of CASP8 and apoptosis in autophagy-deficient U1810 lung cancer cells even under nutrient-rich conditions. Similar to starvation, inhibition of protein translation leads to efficient activation of CASP8 and cell death in autophagy-deficient lung cancer cells. Thus, here for the first time we report that suppressed translation leads to activation of CASP8-dependent apoptosis in autophagy-deficient NSCLC cells under conditions of nutrient limitation. Our data suggest that targeting translational machinery can be beneficial for elimination of autophagy-deficient cells via the CASP8-dependent apoptotic pathway.

Introduction

Autophagy is a physiologically conserved mechanism essential for the degradation and recycling of intracellular constituents in lysosomes. This protective mechanism is activated in cells under stress conditions and also can be aberrantly controlled in some pathological conditions.¹ It is suggested that in tumors, autophagy is activated in cells distal from the blood supply (nutrient restriction) or as a response to therapy.² Several pathways are involved in the regulation of autophagy under starvation or nutrient limitation conditions. Thus, MTOR protein kinase, a regulator of cap-dependent protein translation, is a key player in the autophagy pathway. A lack of amino acids affects MTOR complex activity, leading to dephosphorylation of ATG13 and activation of the ULK1/2 autophagy initiator complex under nutrient-deprivation conditions.^3-5 Furthermore, a drop of ATP during starvation leads to activation of AMPK kinase, which either directly phosphorylates and controls activities of the autophagy proteins ULK1 and ATG13 or regulates ULK complex activity via inhibition of the MTOR complex.⁶

In most cases, upregulation of autophagy under starvation conditions preserves survival of cells and mice, and inhibition of autophagy under such conditions is often associated with increased cell death^7,8 however, activation of a type of autophagy-dependent cell death has also been suggested under some stress conditions.^9,10 Previously, several players involved in the regulation of both the autophagy and apoptosis pathways have been described. Thus, some transcriptional factors, such as TP53, activate expression of genes that are involved in both autophagy and apoptosis; BCL2 family members control apoptotic responses but also have a role in the regulation of autophagy by sequestering BECN1.^11-13 Increased levels of ROS can trigger permeabilization of the mitochondria membrane and initiate apoptosis but can also activate autophagy.¹⁴ Furthermore, several key autophagy proteins or their cleaved products may participate in the execution of an apoptotic program, and some apoptotic proteases inhibit autophagy by cleaving ATG proteins.¹⁵

Accumulating evidence suggests that one of the main mechanisms for activation of apoptosis in autophagy-deficient cells under stress conditions is accumulation of damaged mitochondria that triggers apoptosis via the CASP9/caspase-9-dependent pathway.¹⁶ In the present study, we demonstrate that under conditions of amino acid and growth factor deprivation, autophagy-deficient lung cancer cells efficiently die by caspase-dependent apoptosis, and activation of CASP8/caspase-8 is required for initiation of an apoptotic cascade in these cells. We show that due to nutrient limitation protein translation is suppressed, leading to downregulation of CFLARs and activation of CASP8 under such conditions. Similar to starvation, inhibition of protein translation with cycloheximide leads to efficient CASP8 activation and apoptosis in cells with silenced ATG13, suggesting that protein translation inhibition is one of the key mechanisms of increased apoptotic cell death in these cells under starvation conditions, and such activation of CASP8 takes place upstream of mitochondria outer membrane permeabilization (MOMP). Since CASP8 expression is upregulated in some lung adenocarcinomas, these results suggest that targeting protein translation can be a promising strategy to promote an efficient apoptotic response in autophagy-deficient lung cancer cells via the CASP8-dependent pathway.

Results

Autophagy inhibition facilitates caspase-dependent apoptosis in non-small cell lung cancer cells under conditions of amino acid and growth factor deprivation

ULK complex is the upstream initiator complex in the autophagy pathway. ATG13 is a component of the ULK complex that is directly controlled by MTOR, and it is one of the most upstream proteins, which is activated within this complex in response to starvation. Using the CRISPR/Cas9 system, we designed 2 gRNAs targeting different sequences of the human ATG13 gene. The efficiency of ATG13 silencing and the suppression of basal autophagy in the U1810 lung adenocarcinoma cell line were confirmed by immunoblotting using ATG13 and SQSTM1 antibodies, respectively (Fig. 1A). To activate autophagy, cells were grown in amino acid and growth factor-free Hank's balanced salt solution (HBSS) medium as previously described.¹⁷ Autophagy activation under starvation was confirmed by staining of autophagosomes with antibodies to MAP1LC3 and lysosomes with anti-LAMP2 (Fig. 1B, Fig. S1A).¹⁸ Under starvation conditions, control (EV/empty vector-transduced) cells showed accumulation of autophagosomes and their colocalization with lysosomes, and this effect was inhibited in ATG13-knockout cells.

Figure 1.

Autophagy inhibition facilitates caspase-dependent apoptosis in non-small lung cancer cells under conditions of amino acid and growth factor deprivation. (A) The ATG13 gene was knocked out by targeting 2 different sequences of the gene (ATG13_KO1 and ATG13_KO2; EV corresponds to the empty vector control) by using the CRISPR/CAS9 system. The efficiency of gene silencing was checked by immunoblotting, and the efficiency of autophagy inhibition was assessed by detecting accumulation of SQSTM1. (B) Control (EV) and ATG13-knockout U1810 cells were starved (24 h) in HBSS medium, and autophagy activation was detected by microscopy. Autophagosomes were stained using MAP1LC3 antibody (green); lysosomes, LAMP2 (red); nuclei were counterstained with Hoechst 33342 (blue). (C) Detection of CASP3- and CASP7-like activities in control and ATG13-knockout U1810 cells grown in complete RPMI1640 medium or starved for 24 h in HBSS. (D) Cleavage of PARP1 increased in ATG13-knockout cells under starvation conditions. U1810 cells were starved in HBSS medium (24 h) and treated with either DMSO or the pan-caspase inhibitor z-VAD-fmk. (E) Increase of caspase-dependent apoptotic cell death in ATG13-knockout U1810 cells starved (24 h) in HBSS medium and its inhibition by z-VAD-fmk (20 µM). Cell death was assessed by flow cytometry using ANXA5/annexin V and a PI detection kit. (F) Upregulation of cell death in H1299 and H661 cells starved for 24 and 8 h in HBSS, respectively. Cell death was measured by detecting ANXA5 exposure and membrane permeability with PI by flow cytometry. * P ≤0.05; ns, nonsignificant.

To assess the effect of autophagy inhibition on the activation of apoptosis, U1810 cells were starved for 24 h and CASP3- and CASP7-like activities were measured. Under starvation conditions, ATG13-deficient cells showed a dramatic increase in CASP3- and CASP7-like activities (Fig. 1C). Increased caspase-like activities were accompanied by accumulation of their cleaved PARP1 substrate, which was blocked by the pan-caspase inhibitor z-VAD-fmk (Fig. 1D). Increased caspase-like activities led to significant amplification of apoptotic cell death in several ATG13-deficient clones during nutrient limitation (Fig. S1B and S1C). Inhibition of caspase-like activities significantly reduced cell death, suggesting that the increased cell death response in autophagy-deficient U1810 cells under starvation was a caspase-dependent process (Fig. 1E; Fig. S1D). Furthermore, a similar increase in cell death was observed in other autophagy-deficient lung cancer cell lines (Fig. 1F).

Apoptosis in autophagy-deficient cells under starvation conditions is dependent on CASP8

To further explore which initiator caspase was activated the most upstream in the apoptotic pathway of ATG13-deficient cells, 4 lung cancer cell lines deficient in ATG13 were established using the CRISPR/Cas9 system, and activation of caspases was tested. Cells were starved in HBSS medium for the indicated period of time, and processed (active) fragments of CASP8 and CASP9 were detected by immunoblotting (Fig. 2A; Fig. S1E). The levels of SQSTM1 and MAP1LC3 proteins were measured to prove the efficiency of autophagy activation in control cells and its inhibition in ATG13-knockout cells. Both initiator CASP9 and CASP8 were efficiently activated in autophagy-deficient lung cancer cells. A similar effect on activation of CASP8 and CASP9 was observed in another cell type, HeLa (Fig. 2B). Activation of caspases under nutrient limitation conditions was accompanied by significant downregulation of anti-apoptotic BCL2 family proteins that are typically downregulated under this condition (Fig. 2C).

Figure 2.

Enhanced activation of caspase signaling in ATG13-deficient cells. (A) Control (EV) and ATG13-deleted U1810, H1299, and H661 cells were starved in HBSS medium for 24 h (U1810 and H1299 cells) or 8 h (H661 cells), and the expressions of cleaved forms of CASP8, CASP9, and PARP1 were used to detect caspase activation by immunoblotting. Detecting degradation of SQSTM1 and MAP1LC3 proteins proved autophagy activation under starvation conditions. GAPDH was used as a loading control. (B) ATG13 was silenced in HeLa cells using siRNA. Forty-eight h after transfection cells were starved in HBSS medium (16 h) and the expressions of cleaved forms of CASP8, CASP9, and cleaved PARP1 were used to detect caspase activation by immunoblotting. (C) Activation of caspase signaling and cleavage of PARP1 is associated with downregulation of anti-apoptotic proteins under conditions of nutrient limitation. Control or ATG13-deficient U1810 cells were starved in HBSS (24 h) and the expression of proteins was detected by immunoblotting.

To reveal the role of a specific initiator caspase in the apoptotic response of these cells to starvation, initiator CASP9, CASP8, and CASP2 were silenced in control and autophagy-deficient U1810 lung cancer cells using siRNAs (Fig. 3A; Fig. S2). These cells were starved, and cell death was measured using ANXA5/annexin V and propidium iodide (PI) staining. In contrast to CASP9 and CASP2, silencing of CASP8 significantly reduced cell death under starvation conditions (Fig. 3B). These data were further confirmed by using another siRNA, where only CASP8 silencing dramatically reduced effector caspase activities measured by cleavage of the caspase substrate LMNA (lamin A/C) (Fig. 3C). Moreover, silencing of CASP8 had a significant effect on long-term clonogenic survival in response to nutrient limitation (Fig. 3D). Similar effect on amplification of CASP8 apoptotic signaling under nutrient limitation conditions was confirmed by using ATG7 knockout cells, suggesting the importance of CASP8 in the apoptotic response of cells with inhibited autophagy (Fig. 3E). Moreover, the importance of CASP8 was further confirmed by using other lung cancer cell lines, where silencing of CASP8 led to an inhibition of CASP9 activation and PARP1 cleavage (Fig. 3F) and dramatically reduced cell death during starvation (Fig. 3G).

Figure 3.

ATG13-knockout lung cancer cells die by CASP8-dependent apoptosis under conditions of nutrient limitation. (A) Silencing of CASP2, CASP8, and CASP9 in U1810 cells by using siRNAs. (B) Silencing of CASP8 but not CASP9 or CASP2 significantly reduced apoptosis in U1810 cells starved in HBSS (24 h). (C) Silencing of CASP8, but not CASP9 significantly reduced effector caspase activity. To prove specificity of the effect, another sequence of siRNA targeting CASP8 and CASP9 were used. Expression of CASP8 and CASP9, and cleavage of LMNA was detected by immunoblotting. (D) 24 h after transfection with scramble or CASP8 siRNAs, cells were counted and seeded in 12-well plates, grown either in control medium or starved in HBSS (2 d). Two d after the seeding control cells were fixed with paraformaldehyde and stained with crystal violet, while HBSS was replaced with complete medium. Then, cells were grown for an additional 4 d, fixed with paraformaldehyde and stained with crystal violet. (E) CASP8 was silenced in U1810 cells with knockout of ATG7. The cells were starved for 24 h in HBSS and CASP8 processing and PARP1 cleavage were detected by immunoblotting. (F) Silencing of CASP8 in H661 and H1299 lung cancer cells significantly reduced activation of CASP9 in cells under starvation conditions. H661 cells were starved for 8 h and H1299 cells for 24 h, and processed forms of CASP9, CASP8 and the cleaved form of PARP1, were detected by immunoblotting. (G) Cell death in H1299 and H661 cells treated as in (F) and detected by using the ANXA5 assay. t-test, *P<0.05.

Caspases are enzymes that are activated when they are brought into close proximity with each other. Some data indicate that the autophagy receptor protein SQSTM1/p62 promotes aggregation of CASP8 within SQSTM1-dependent foci.¹⁹ Therefore, we decided to test whether such activation of CASP8 might take place under nutrient limitation conditions. SQSTM1-containing aggregates are insoluble in Triton X-100.²⁰ Thus, we performed fractionation and found that under starvation conditions SQSTM1 was dramatically reduced in EV cells in 2 fractions; however, its level was maintained in ATG13-deficient cells in both digitonin-extracted and Triton X-100 insoluble fractions (Fig. 4A). Moreover, we found that both full-length and active forms of CASP8 were accumulated in the Triton X-100-insoluble fraction of autophagy-deficient cells containing SQSTM1 under starvation conditions (Fig. 4A). Next, using immunocytochemistry we detected strong colocalization of active CASP8 with the aggregates containing SQSTM1, suggesting this platform for efficient activation of CASP8 in autophagy-deficient cells (Fig. 4B).

Figure 4.

Association of CASP8 with a fraction containing SQSTM1. (A) Antibodies for cleaved and full-length CASP8 and SQSTM1 were used to show their association within different fractions. CYCS (cytochrome c, somatic) and GAPDH antibodies, and Ponceau S staining were used to show the efficiency of cell fractionation and protein loading (B) Control (EV) and ATG13-knockout U1810 cells were starved (24 h) in HBSS medium, and localization of the cleaved form of CASP8 (green) and SQSTM1 (red) were detected by microscopy. Nuclei were counterstained with Hoechst 33342 (blue).

Mitochondria permeabilization in autophagy-deficient cells is downstream of CASP8

Mitochondria play a critical role in the initiation of the intrinsic apoptotic pathway, and it has been suggested that accumulation of damaged mitochondria in autophagy-deficient cells might lead to activation of an apoptotic response in these cells under different stress conditions, including starvation. We assessed the mitochondria functions by measuring the oxygen consumption rate (OCR) in control and ATG13-deleted U1810 cells. Wild-type and autophagy-deficient U1810 cells were characterized by comparable rates of basal and uncoupled (CCCP-stimulated) mitochondria activities (Fig. 5A). Furthermore, OCRs were dramatically reduced in both control and autophagy-deficient cells starved in HBSS medium, indicating that functions of mitochondria were significantly suppressed in both cell types during starvation. Moreover, under starvation conditions the OCR stimulation by CCCP was negligible, suggesting the deficiency of substrates for mitochondria respiration under such conditions. This was further confirmed by measuring the OCR rates of mitochondria after cell permeabilization with digitonin and supplementation of incubation medium with substrates for mitochondrial complexes I (pyruvate and malonate) and II (succinate) (Fig. S3), where both wild-type and ATG13-deficient cells actively respired in the buffer supplemented with these substrates.

Figure 5.

Mitochondria permeabilization in autophagy-deficient cells is downstream of CASP8. (A) U1810 cells were grown in complete medium or starved in HBSS (6 h). Basal and CCCP-uncoupled respiration were measured using the oxygraph system. The data are presented in the graph as the rate of oxygen consumption in nmols per min by 1 million cells (OCR). (B) Control (EV) or ATG13-knockout cells were grown in complete medium or starved (4 h) in HBSS. Treatments with 2-deoxyglucose (2DG; 10 mM) and antimycin A (Am A; 5 µM) were applied, and ATP content was measured according to the protocol described in the Materials and Methods section. (C) Release of CYCS/cytochrome c from mitochondria under starvation conditions detected by immunoblotting. U1810 and H661 control (EV) or ATG13-knockout (KO) cells were starved in HBSS medium for 16 or 8 h, respectively, and fractionation was performed according to the protocol described in the Materials and Methods section. GAPDH and SDHA antibodies were used as markers for cytosolic (Cyt) and mitochondrial (Membr) fractions, respectively. (D) Empty vector-transduced (EV) or ATG13-knockout H661 cells were untreated or treated with z-VAD-fmk (10 µM) and starved in HBSS (8 h). The number of cells with a drop of mitochondrial membrane potential (MMP) was assessed by flow cytometry. (E) Silencing of CASP8 (shCASP8) in control (EV) and ATG13-knockout (KO) U1810 and H661 cells using shRNA. (F) Control (EV) or ATG13-knockout U1810 and H661 cells with silenced CASP8 were starved for 16 or 8 h, respectively, and the number of cells with a drop of MMP was detected by flow cytometry using staining with tetramethylrhodamine ethyl ester perchlorate. (G) Silencing of CASP8 inhibited the release of CYCS from mitochondria of cells starved in HBSS. U1810 cells were starved in HBSS medium (16 h), and cell fractionation was performed as described in the Materials and Methods section. t-test, *P<0.05.

To assess the type of metabolic signature in wild-type and ATG13-deficient cells grown in complete and starvation medium, we measured ATP content by inhibiting glycolysis with 2-deoxyglucose, and mitochondria respiration with antimycin A. Thus, both wild-type and ATG13-deficient cells were more dependent on glycolysis under fed conditions. Inhibition of either glycolysis or mitochondria activities reduced the level of ATP during starvation, but without significant differences between control and autophagy-deficient cells (Fig. 5B). As expected, simultaneous inhibition of glycolysis and mitochondria respiration caused a rapid and dramatic drop of ATP (Fig. 5B) and cell death (Fig. S4) under both experimental conditions.

To assess the effect of autophagy inhibition on the permeabilization of the mitochondria membrane in wild-type and ATG13-knockout cells, U1810 and H661 cells were starved and cell fractionation was performed. Both U1810 and H661 autophagy-deficient cells were characterized by increased release of CYCS (cytochrome c, somatic) under starvation (Fig. 5C). Because CASP8 silencing blocked activation of CASP9, to prove the hypothesis that permeabilization of the mitochondria membrane under starvation was downstream of caspases, we inhibited caspases using the pan-caspase inhibitor z-VAD-fmk and starved those cells in HBSS medium. Indeed, inhibition of caspases significantly reduced the amount of cells with a complete drop of mitochondria membrane potential (MMP) (Fig. 5D).

Next, we silenced CASP8 in control and ATG13-knockout H661 and U1810 cells by using shRNA to prove its specific effect on mitochondria (Fig. 5E). Cells were starved for the indicated time, and the effect on MMP was measured by flow cytometry. Silencing of CASP8 significantly reduced the amount of cells with MMP drop, indicating that such an effect on mitochondria was mediated by CASP8 (Fig. 5F; Fig. S5). Furthermore, to prove that CASP8 also affected the permeabilization of the mitochondria membrane, U1810 cells were starved and fractionation was performed (Fig. 5G). Indeed, silencing of CASP8 blocked CYCS release in ATG13-knockout cells during starvation, further demonstrating that the permeabilization of the mitochondria membrane in autophagy-deficient cells was downstream of CASP8 activation.

Because autophagy-deficient cells are dependent on glutamine,²¹ we tested whether supplementation with glutamine inhibits cell death in ATG13-knockout cells. Indeed, supplementation of HBSS with glutamine was capable of rescuing all studied lung cancer cell lines under starvation conditions (Fig. 6A). Moreover, such inhibition of cell death was associated with inhibition of CASP8 and CASP9 activation and stabilization of CASP8 inhibitory CFLAR protein (Fig. 6B). Finally, we demonstrated that supplementation of HBSS with glutamine was sufficient to block the drop of MMP (Fig. 6C), suggesting that glutamine supplementation prevented mitochondria membrane permeabilization and cell death via inhibition of CASP8 activation.

Figure 6.

Glutamine inhibits CASP8 activation and cell death under conditions of nutrient limitation. (A) Supplementation of HBSS medium with glutamine was sufficient to rescue NSCLC cells from cell death. H661 or H1299 cells were starved for 8 h or U1810 starved for 24 h in HBSS medium with or without supplementation with glutamine. Cell death was measured using ANXA5-PI staining in flow cytometry. (B) Supplementation with glutamine inhibited CASP8 processing under starvation conditions in HBSS medium. H661 cells were starved (8 h) in HBSS with or without supplementation with glutamine or pretreated (30 min) with the pan-caspase inhibitor z-VAD-fmk (10 µM). Processed forms of CASP9 and CASP8, cleavage of PARP1, and expression of CFLAR and MAP1LC3 were detected by immunoblotting. GAPDH was used as a loading control. (C) H661 cells were starved in HBSS (8 h) with or without supplementation with glutamine, and MMP was measured as described in the Materials and Methods section. t-test, *P<0.05; ns, nonsignificant.

Suppressed translation downregulates CFLARs and activates CASP8 during starvation

Recently, we have reported that prolonged starvation conditions are associated with significant suppression of general protein translation.¹⁷ Indeed, starvation of U1810 or H661 cells in HBSS led to a dramatic reduction of protein translation, but such an effect was weaker in comparison to cell treatments with cycloheximide, an inhibitor of protein translation (Fig. 7A). Thus, both control and ATG13-knockout cells showed a dramatic reduction of protein translation in HBSS medium. It is known that CFLAR proteins have a short lifetime and, therefore, are sensitive to inhibition of protein translation. In both H661 and U1810 cell lines the levels of CFLARs were reduced already within 4 h of starvation (Fig. 7B). To reveal their potential contribution to the survival of these cells, we silenced CFLAR isoforms by using siRNAs targeting either a single CFLAR isoform (long, CFLAR-L, or short, CFLAR-S) or simultaneously both CFLAR isoforms (total, CFLARs). Silencing of CFLAR-L was sufficient to promote activation of CASP8 in ATG13-deleted cells even under nutrient-rich conditions, and this effect was significantly increased when both CFLAR isoforms were silenced, suggesting that conditions/treatments that downregulate this CASP8 inhibitory protein promote efficient activation of CASP8 in autophagy-deficient cells (Fig. 7C). Further studies showed that activation of CASP8 in U1810 cells with silenced CFLARs strongly correlated with an increase of apoptosis in cells with deletion of ATG13 (Fig. 7D). Moreover, specific inhibition of protein translation with cycloheximide caused rapid and efficient downregulation of both CFLAR isoforms and facilitated activation of CASP8 and cell death in ATG13-knockout cells (Fig. 7E and F).

Figure 7.

Suppressed translation is associated with downregulation of CFLARs and activation of CASP8 under starvation conditions. (A) H661 and U1810 cells were untreated, treated with cycloheximide (CHX; 10 µg/ml), or starved in HBSS medium for 8 h. Cell translation was measured by using puromycin incorporation as described in the Materials and Methods section. Staining with Ponceau S was used as a loading control. (B) U1810 or H661 cells were starved for different times in HBSS medium, and the levels of CFLARs were detected by immunoblotting. (C) Silencing of CFLARs promoted activation of CASP8 in U1810 lung cancer cells. Cells were transfected with scrambled siRNA or siRNAs targeting CFLAR-S (FS), CFLAR-L (FL), or both CFLAR isoforms (FT). Twenty-four h after transfection, the samples were collected for immunoblotting. Antibodies for cleaved CASP8 were used to detect processing of CASP8 and for cleaved PARP1 to assess the effector caspase activities. (D) Detection of cell death in control (EV) or ATG13_KO U1810 cells with silenced CFLARs. Cells were transfected with respective CFLAR siRNAs, and cell death was measured by flow cytometry 24 h after transfection by using an ANXA5 detection kit. (E) Inhibition of protein translation with cycloheximide (10 µg/ml) promoted activation of CASP8 in H661 and U1810 cells. Antibodies for cleaved CASP8, cleaved PARP1, and CFLARs were used to detect the apoptotic response. (F) Treatment with cycloheximide (10 µg/ml, 8 h) induced rapid cell death in U1810 cells with deleted ATG13. t-test, *P<0.05; ns, nonsignificant.

To prove the importance of CFLAR downregulation under starvation conditions, we established stable U1810 cells with overexpression of CFLAR-S or CFLAR-L isoforms. Autophagy in these cells was inhibited by silencing ATG13, and then these cells were starved in HBSS medium. As expected, the levels of both CFLAR isoforms were dramatically reduced in cells with overexpressed CFLAR during starvation; however, the remaining expressions of CFLARs were sufficient to significantly reduce processing of CASP8, cleavage of PARP1, and cell death under starvation conditions as well as affect clonogenic survival (Fig. 8A and B; Fig. S6). Moreover, similar to starvation, the effect of cycloheximide on CASP8 activation and cell death was reduced in cells with overexpressed CFLARs (Fig. 8C and D). The inhibitory effect of CFLAR-S was significant but less pronounced due to its very low stability under conditions of inhibited translation. Because it is known that CFLARs are degraded in proteasomes,^22,23 we measured their synthesis by blocking this degradation pathway. Measurement of mRNA expression of CFLAR isoforms showed that in contrast to protein, the mRNA levels were upregulated in starved cells, suggesting activation of prosurvival pathways to upregulate expression of CFLAR mRNAs under such conditions, and the potential accumulation of CFLAR protein in case of functional translation (Fig. 8E). Thus, to measure CFLAR proteosynthesis, cells were either grown in complete medium or starved in HBSS with or without treatments with the inhibitor of proteasomal degradation MG132 (5 µM) (Fig. 9 A and B). Densitometry analysis revealed that inhibition of the CFLARs degradation pathway led to their accumulation in cells grown in complete medium but not under starvation conditions, demonstrating the inhibition of CFLAR synthesis during starvation.

Figure 8.

Translation of CFLAR prevents cell death under starvation conditions. (A) Overexpression of CFLAR isoforms reduced CASP8 activation in U1810 cells during starvation. U1810 cells were transduced with retroviruses overexpressing CFLAR-S (FS) or CFLAR-L (FL) isoforms, and cells with stable CFLAR expression were established as described in the Materials and Methods section. To inhibit autophagy, cells overexpressing different CFLAR isoforms were transfected with scrambled siRNA or siRNA targeting ATG13 and then starved for 16 h in HBSS medium. The efficiency of CFLAR overexpression, ATG13 silencing, and autophagy inhibition were confirmed by using CFLAR, ATG13, and MAP1LC3 antibodies, respectively. Activation of CASP8 and cleavage of PARP1 were detected by immunoblotting. Equal loading was verified by using GAPDH antibody. (B) U1810 cells with stable expressions of CFLAR isoforms were transfected with scrambled or ATG13 siRNAs and starved as described in (A). Cell death was measured using an ANXA5 detection assay. (C) Assessment of cell death using ANXA5 and PI staining in U1810 cells with silenced ATG13 untreated or treated with cycloheximide (10 µg/ml, 4 h). (D) U1810 cells with overexpressions of CFLAR isoforms were treated with cycloheximide (10 µg/ml, 4 h), and expression of CFLAR and processing of CASP8 were detected by immunoblotting. (E) Expression of CFLARs mRNAs in wild-type and ATG13-deficient cells grown in complete or starvation medium (8 h). t-test, *P<0.05.

Figure 9.

Protein synthesis of CFLAR is significantly reduced under starvation conditions. (A) Expression of CFLAR isoforms in cells overexpressing EV, CFLAR-L, or CFLAR-S and grown in complete or starvation medium (8 h) were detected by immunoblotting. To check the proteosynthesis of CFLAR isoforms, protein degradation was inhibited by treating cells with the inhibitor of proteasomal degradation MG132 (5 µM) and (B) densitometry analysis was performed using ImageJ software. t-test, *P<0.05. (C) A proposed scheme for CASP8 activation and apoptosis in autophagy-deficient NSCLC cells under conditions of nutrient limitation.

Overall, our data suggest that mitochondria may function more as an amplification loop rather than initiators of apoptotic signaling in autophagy-deficient cells during starvation. Proteosynthesis of CFLARs is significantly reduced under starvation conditions, leading to efficient activation of CASP8 and apoptosis in autophagy-deficient lung cancer cells experiencing starvation.

Discussion

Autophagy inhibition under starvation is often associated with increased cell death responses. Because the level of autophagy could be differently modulated in tumors, it is important to understand how cell death programs are activated in cells under stress conditions when autophagy is being either activated or suppressed. Thus, accumulation of damaged mitochondria is suggested as the main mechanism of apoptosis activation in the response of cells with inhibited autophagy to various stress stimuli, and such an apoptotic response is associated with the formation of apoptosomes and activation of the CASP9-dependent pathway.¹⁶ In the present study, we show that the CASP8-dependent pathway mediates the apoptotic response in non-small cell lung cancer cells under starvation conditions.

Previously, we and others reported upregulation of CASP8 and CFLARs expression in human lung adenocarcinomas;^24,25 this observation has led to a potential interest in targeting/inhibiting autophagy in order to facilitate activation of CASP8-mediated apoptosis in these tumors. CASP8 is activated in cells as a response to death ligands, where it is recruited and autocatalytically processed at the DISC complex.²⁶ Interestingly, it has previously been reported that activation of CASP8 can be mediated even without treatment of cells with death ligands; just silencing the inhibitory CFLAR proteins can lead to activation of CASP8 and apoptosis in some non-small cell lung cancer cell lines.²⁵ Thus, the results obtained in our current study suggest that during starvation, suppression of CFLARs synthesis is involved in initiation of an apoptotic response. Our data demonstrate that some autophagy-deficient NSCLC cells efficiently activate CASP8 when CFLARs are silenced.

There are 2 possible mechanisms of upregulation of CASP8 activity in autophagy-deficient cells—an increase of its activation per se or inhibition of its degradation. Thus, activation of CASP8 after downregulation of CFLARs can be facilitated in autophagy-deficient cells via accumulation of the autophagy receptor and ubiquitin-binding protein SQSTM1, which promotes aggregation of CUL3-modified CASP8 within SQSTM1-dependent foci, and leads to its full activation in response to death ligands.¹⁹ In our current work we demonstrated that similar to activation of extrinsic apoptotic stimuli, under conditions of nutrient limitation CASP8 is translocated and activated within SQSTM1-containing foci. Additionally, active CASP8 can be eliminated by autophagy,²⁷ suggesting that downregulation of CFLARs under starvation will also lead to more accumulation of the active form of CASP8 in autophagy-deficient cells and an efficient apoptotic response. Interestingly, it was also suggested that in addition to the well-established role of CFLARs in apoptosis, viral CFLARs suppress autophagy by preventing ATG3 from binding and processing MAP1LC3. Thus, based on this report, one could suggest that downregulation of CFLARs under starvation conditions may facilitate an autophagy response.²⁸

It was previously reported that CFLAR is transcriptionally regulated by several growth and survival-promoting signaling pathways, including NFKB/NF-κB, AKT, and MAPK/ERK.^29-31 Activation of some of these pathways under stress conditions, for instance via upregulation of ROS (that also takes place during starvation), could potentially upregulate the mRNA levels of CFLARs. As we reported here, under starvation conditions the protein levels of CFLARs were downregulated. It is known that CFLARs are degraded in proteasomes, and their expression is very sensitive to inhibition of protein translation.³² Our data demonstrate that although the levels of CFLAR mRNA were increased during starvation, they were not sufficient to upregulate their protein levels even when the degradation process was blocked, further suggesting an important contribution of the processes downstream of transcription but upstream of degradation. Our data revealed that under starvation conditions, suppressed translation promotes downregulation of CFLAR proteins, which induces efficient activation of CASP8 and an apoptotic response in autophagy-deficient cells. Such activation of CASP8 is required for an apoptotic response in NSCLC cells with inhibited autophagy under starvation conditions. Our previous study proposed a model suggesting that inhibition of protein translation contributes to the limitation of autophagy under prolonged starvation conditions.¹⁷ In the present study, we showed that such inhibition of protein translation under starvation leads to amplification of apoptosis in autophagy-deficient cells, suggesting a novel targeting pathway for efficient elimination of autophagy-deficient cells.

Materials and methods

Reagents and antibodies

Mitochondrial inhibitor, uncoupler and substrates: Antimycin A (Sigma, A8674), carbonyl cyanide 3-chlorophenylhydrazone (CCCP; Sigma-Aldrich, C2759), sodium pyruvate (Sigma-Aldrich, P2256), succinate disodium salt (Sigma-Aldrich, S2378), malic acid (Sigma-Aldrich, M1000). Proteasomal inhibitor: MG132 (Enzo Life Sciences, BML-PI102). Proteosynthesis inhibitor: cycloheximide (Sigma, c6255). Caspase inhibitor: z-VAD-fmk (Peptide Institute Inc., 3188-v). Inhibitor of glycolysis: 2-deoxyglucose (Sigma, D8375). Antibodies: anti-ATG13 (Cell Signaling Technology, 6940), anti-MAP1LC3 (MBL, PM036), anti-GAPDH (Trevigen, 2275-PC-100), anti-ACTB/actin (Sigma, A2066), anti-TUBB (Sigma, T5168) anti-LAMP2 (Santa Cruz Biotechnology, sc-18822), anti-puromycin (Millipore, MABE343), anti-CFLAR and anti-CASP8 (kindly provided by profs P Krammer and I. Lavrik), anti-cleaved PARP1 (Cell Signaling Technology, 9546), anti-CASP2 (BD Biosciences, 611023), anti-PARP1 (BD Biosciences, 556494), anti-cleaved CASP3 (Cell Signaling Technology, 9661), anti-cleaved CASP9 (Cell Signaling Technology, 9505), anti-CASP9 (Cell Signaling Technology, 9508), anti-BCL2L1 (Santa Cruz Biotechnology, sc-634), anti-MCL1 (Sigma, M8434), anti-BCL2 (Santa Cruz Biotechnology, sc-492) anti-cleaved CASP8 (Cell Signaling Technology, 9496), anti-CYCS (Cell Signaling Technology, 556433), anti-SDHA (Cell SignalingTechnology, 5839), anti-SQSTM1 (Santa Cruz, 28359). Secondary antibodies: anti-mouse IgG (Pierce, 31430), anti-rabbit IgG (Pierce, 31460).

Cell culture

Human lung carcinoma cell lines H1299 (ATCC, CRL-5803), H661 (ATCC, HTB-183), H125 (ATCC, CRL-5801) and U1810 lung carcinoma (from collection at Uppsala University, Sweden) were cultured in RPMI 1640 medium (Gibco, 52400-041) or human cervical carcinoma cell line HeLa (ATCC, CCL-2) in DMEM medium (Gibco, 41965-039) both supplemented with 10% (v:v) fetal bovine serum (Gibco, 10270), 100 U/ml penicillin and 100 mg/ml streptomycin (Gibco, P0781) and maintained at 37°C with 5% CO₂ in air atmosphere. Starvation conditions were achieved by washing cells 3 times with phosphate-buffered saline (PBS; Sigma, D8537) and culturing them in HBSS medium (Gibco, 24020–133) for the indicated hours.

Clonogenic assay

Cells were seeded and grown in 12-well plates either in control (complete) medium or starved in HBSS as described in the figure legends. The cells were fixed in 4% paraformaldehyde (15 min at room temperature [RT]), washed with PBS and then stained with crystal violet.

LentiCRISPR ATG13 plasmids and cell transduction

ATG13 knockout cell lines were generated by adapting a CRISPR-Cas9 system using lentiCRISPR v1 vector (Addgene, 49535; deposited by Feng Zhang) according to the Dr. Feng Zhang Laboratory protocol³³ and as it was previously described.¹⁷ The following sequences of the primers were used (ΔATG13_#1 FW: CACCGGTGATTGTCCAGGCTCGGCT; RW: AAACAGCCGAGCCTGGACAATCACC; ΔATG13_#2 FW CACCGGATTTCACTTAAGACTTCTG; RW: AAACCAGAAGTCTTAAGTGAAATCC). Cells were transduced with lentiviruses and selected with 1 μg/mL puromycin.

Generation of stable CFLAR-long and CFLAR-short-overexpressing cells

Empty vector (EV), CFLAR-long- and CFLAR-short-containing pBABE puro vectors (kindly provided by Dr. D. Longley, Centre for Cancer Research and Cell Biology, Queen's University Belfast, Belfast, UK) were transfected into the HEK293T Phoenix™ Ampho packaging cell line using Lipofectamine LTX & PLUS™ Reagent transfection reagent (Invitrogen, 15338-100) according to the protocol. Following a 48-h transfection, the medium was filtered and polybrene (Sigma, 107689) was added (1:1000) to U1810 cells for 24 h. Transduced cells were selected with 1 μg/mL puromycin. Overexpression of CFLARs was confirmed by immunoblotting.

Lentiviral shRNA constructs and cell transduction

Plasmid expressing shRNA targeting CASP8 was obtained from Sigma (SHCLNG-NM_033356, clone NM_033356.3-415s21c1). To produce viruses, HEK293 cells were transfected with delta 8.9, VSV-G and either empty pLKO.1-vector (Sigma, SCC001) or shCASP8-pLKO1 plasmids (Sigma, TRCN00000 12246) using Lipofectamine LTX & PLUS™ Reagent transfection reagent according to the manufacturer's protocol. Following a 48-h transfection, the medium was filtered and polybrene added to the cells for 24 h. Expression of CASP8 was measured 3 d after viral infection. Silencing efficiency was confirmed by immunoblotting.

siRNA transfections

Cells were seeded in 6- or 12-well plates and in 24 h transfected with INTERFERin siRNA Transfection Reagent (Polyplus-transfection; 409–10). For each transfection, siRNAs (non-targeting pool (D-001810-10-05), si_CASP8 (L-003466-00-0005), si_CASP2 (L-003465-00-0005), si_CASP9 (L-003309-00-0005), si_ATG13 (M-020765-01-0005) all SMARTpool from Dharmacon; si_CASP9_2 (s2429; CAGAGGUUCUCAGACCGGAtt) and si_CASP8_2 (s2427; GAUACUGUCUGAUCAUCAAtt) all Silencer Select from Ambion (Life Technologies); si_CFLAR-long form AAGGAACAGCTTGGCGCTCAA and si_CFLAR-short form AACATGGAACTGCCTCTACTT (synthesized by Qiagen according to Galligan et al.³⁴), were mixed with INTERFERin in OPTI-MEM medium (Gibco, 51985-026). After 10 min incubation at RT the complexes were added to the cells. The final concentration of siRNA in the medium was 20 nM. Medium was replaced and treatments were administered 24–48 h after transfection.

Caspase activity assay

Cells were washed with ice-cold PBS, resuspended in 25 μl of PBS and after lysis in liquid nitrogen were loaded onto a microtiter plate. CASP3 substrate DEVD-AMC (50 μM; Peptide Institute, 409–10) was added and fluorescence was detected in a Fluoroscan II plate reader (Labsystems) using 355-nm excitation and 460-nm emission wavelengths. Fluorescent units were converted to pmols of released AMC and, subsequently, related to the amount of protein in each sample. Caspase activity was expressed as a fold-increase compared with the appropriate control.

Measurement of intracellular ATP

The cellular ATP content was determined using a Bioluminescent Somatic Cell Assay kit according to the manufacturer's instructions (Sigma-Aldrich, FLASC catalog number). Cells were grown in 12-well plates and ATP was extracted from cells by adding 80 µl of ATP releasing reagent directly to the wells. The ATP content was measured with the luciferin/luciferase method by using a Tecan Microplate Reader (Tecan Trading AG, Switzerland).

Measurement of mitochondrial oxygen consumption

Oxygen consumption rate (OCR) was monitored with an oxygen electrode (Hansatech Instruments, Norfolk, UK) and analyzed with the OxygraphPlus software (Hansatech Instruments, Norfolk, UK). To assess the maximum capacity of the respiratory chain, mitochondria were uncoupled with 5 μM carbonyl cyanide 3-chlorophenylhydrazone (CCCP). OCR was measured on intact or permeabilized (digitonin, 100 µg/ml; Calbiochem, 300410) cells. KCl buffer (140 mM KCl, 1 mM MgCl2, 5 mM KH2PO4, 5 mM Tris, pH 7.4) was used for studies of OCR in digitonin-permeabilized cells. Treatment with antimycin A (5 μM) was used at the end of each measurement to subtract non-mitochondrial oxygen consumption.

Immunoblotting

Cells were collected using trypsin (Sigma, 52400-041), washed with PBS and lysed using a complete Lysis-M buffer (Roche, 04719956001), supplemented with protease inhibitor (Roche, 05892970001) and phosphatase inhibitor (Roche, 04906837001) cocktails. Protein concentrations were determined using the BCA protein assay (Pierce, 23228) according to the manufacturer's instructions. Then, samples were mixed with Laemmli buffer and boiled for 5 min at 98°C. Equal amounts of proteins were separated by SDS-PAGE and blotted onto nitrocellulose membranes (Bio-Rad, 1620115). Membranes, blocked for 1 h with 5% skimmed milk in PBS, were incubated overnight with primary antibodies diluted in PBS (Sigma, D8537) containing 2% BSA (Sigma, A4503) and 0.05% Tween-20 (Sigma, P1379). The recognized proteins were detected using horseradish peroxidase-labeled secondary antibodies anti-mouse IgG and anti-rabbit IgG, and the enhanced chemiluminescence Clarity Western ECL Substrate (Bio-Rad, 1705061).

For preparation of digitonin extracts, cells were incubated for 5 min in a buffer containing 100 μg/ml digitonin (Calbiochem, 300410) in PBS (GIBCO, 20012), and then pelleted for 5 min at 7000 × g and the supernatant was collected. The pellet was washed in the same buffer and resuspended in Triton X-100 buffer (50 mM Tris-HCl, pH 7.5, 0.5% Triton X-100 (Sigma, X100), 137.5 mM NaCl, 10% glycerol (Sigma, G5516) to generate the Triton X-100-soluble fraction. The pellet (Triton X-100-insoluble fraction) was resuspended in complete lysis buffer, mixed with Laemmli buffer and boiled as described above.

Real-time quantitative PCR

RNA was extracted using an RNeasy mini kit (Qiagen, 74106) and 1 µg of total RNA extract was reverse-transcribed using superscript II reverse transcriptase (Invitrogen, 18064-014) according to the manufacturer's instructions. Gene expression levels were assessed in a 7500 Real-Time PCR System (Applied Biosystems, Stockholm, Sweden) on 10 ng of cDNA mixed with SYBR Select Master Mix (Applied Biosystems, 4472908) and 200 nM of primers. The reaction mixtures were subjected to an initial denaturation step, at 95°C for 10 min, and 40 cycles of amplification, each cycle consisting of a denaturation step at 95°C for 15 sec and an annealing/extension step at 60°C for 1 min. A melting curve analysis was used to confirm primer specificity and to ensure the absence of primer-dimer formation. The relative expression levels of each gene are presented as the fold increase relative to untreated cells after normalization against HPRT1. The following sets of primers from Life Technologies were used: CFLAR-short (FW: GGGCCGAGGCAAGATAAGCAAGG and RW: TCAGGACAATGGGCATAGGGTGT); CFLAR-long (FW: AGACACGCGAGTGGCCCTGT, RW: ACCCTCGCCGGACAAGCTCA); HPRT1 (FW: AATTATGGACAGGACTGAACGTCTTGCT, RW: TCCAGCAGGTCAGCAAAGAATTTATAGC).

Measurement of protein translation

A nonradioactive method based on the incorporation of puromycin was used to monitor protein synthesis.³⁵ Briefly, puromycin (1 μg/ml; Sigma, P8833) was added for 10 min to the medium of cultured cells. Then, cells were washed twice with PBS and collected with trypsin. The proteosynthesis level was assessed by immunoblot using anti-puromycin antibody.

Immunocytochemistry

Cells were seeded and grown on coverslips in 6-well plates. After desired treatments were applied, cells were fixed in 4% paraformaldehyde (15 min, RT). Afterwards, coverslips were handled as reported previously.³⁶ Briefly, cells were permeabilized with digitonin (100 µg/µl) and stained with anti-MAP1LC3 and anti-LAMP2 antibodies diluted in PBS-BSA (2%). The next day, the slides were incubated (1 h, RT) with Alexa Fluor 488-conjugated donkey-anti-rabbit (Molecular Probes, A21206; 1:500) or with Alexa Fluor 594-conjugated donkey-anti-mouse (Molecular Probes, A21203; 1:500) secondary antibodies. For SQSTM1 and cleaved CASP8 staining, cells were permeabilized with 0.1% Triton X-100 diluted in PBS. Nuclei were counterstained with DAPI (10 μg/ml; Molecular Probes, D1306) or Hoechst 33342 and the slides were mounted in Vectashield mounting medium (Vector Laboratories, H-1000) and examined under a Zeiss LSM 510 META confocal laser-scanning microscope (Carl Zeiss, Jena, Germany).

Detection of cell death using ANXA5-propidium iodide staining

ANXA5-propidium iodide double staining was carried out using an Annexin-V-FLUOS Staining Kit (Roche Applied Science, 11988549001) according to the manufacturer's protocol. The cells were analyzed by flow cytometry (BD Accuri, Becton Dickinson) and the data were evaluated using BD CSampler software.

Assessment of mitochondria membrane potential (MMP)

Cells were washed in PBS, incubated for 20 min at 37°C with 25 nM of tetramethylrhodamine ethyl ester perchlorate (Molecular Probes, T-669) in PBS. The cells were analyzed by flow cytometry (BD Accuri, Becton Dickinson) and the data were evaluated using BD CSampler software. Populations of cells with complete loss of MMP are presented in the graphs.

Statistical analysis

All data are expressed as means ± s.d. unless stated otherwise. The 2-tailed Student t-test was used to compare difference between groups, in which *P<0.05 was considered statistically significant.

Funding Statement

This work was supported by the Swedish Cancer Society (150371); The Stockholm Cancer Society (141382); Russian Science Foundaton (14-25-00056); The Swedish Childhood Cancer Foundation (PR2016-0090); The Swedish Research Council (521-2014-2258); and The World Wide Cancer Research Association (15-002).

Disclosure of Potential Conflicts of Interest

The authors declare that they have no competing interests.

Acknowledgements

The authors thank professors I. Lavrik (Institute of Experimental Internal Medicine, Magdeburg, Germany) and P. Krammer (German Cancer Research Center, Heidelberg, Germany) for providing CASP8 and CFLAR antibodies, prof. D. Longley (Queen's University Belfast, Belfast, UK) for providing CFLAR-long and CFLAR-short-expressing plasmids. The authors also thank Belen Espinosa for technical assistance. This project was supported by grants from the Swedish and Stockholm Cancer Societies, the Swedish Research Council, the Swedish Childhood Cancer Foundation, Karolinska Institutets Forskningsstiftelser and the Elsa Goljes Foundation. VG and BZ (assessment of respiration and mitochondria permeabilization experiments) were supported by the Russian Science Foundation (14-25-00056). GA and GB were supported by the Erasmus traineeship program and KJ was supported by EuroScholars program. FC was supported by the World Wide Cancer Research Association (15-002).