The second mitochondrial activator of caspases (SMAC) regulates growth, inflammation and mitochondrial integrity in cancer cells

Amer, Tarek N.; Haimovici, Aladin; Kirschnek, Susanne; Vier, Juliane; Moeed, Abdul; Ukachukwu, Uzochukwu; Neugebauer, Daniela; Neubert, Philip; Helmstädter, Martin; Kayser, Severine; Öllinger, Rupert; Rad, Roland; Groß, Olaf; Holzschuh, Jochen; Driever, Wolfgang; Weber, Arnim; Badr, Mohamed Tarek; Häcker, Georg

doi:10.1038/s41418-026-01667-z

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Published: 03 February 2026

The second mitochondrial activator of caspases (SMAC) regulates growth, inflammation and mitochondrial integrity in cancer cells

Cell Death & Differentiation (2026)Cite this article

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Abstract

SMAC is a mitochondrial intermembrane space protein, which is released during apoptosis and whose known function is antagonism of inhibitor of apoptosis proteins in the cytosol, to facilitate caspase activation. Recent data suggest that SMAC can also be released by sub-lethal signals in the apoptosis pathway, in the absence of cell death. We here explored potential functions of SMAC in non-apoptotic cells. We found that a portion of SMAC is spontaneously released into the cytosol in the absence of apoptosis, regulated by the BCL-2-family proteins BAX and BAK and the fission GTPase DRP1. In cancer cell lines, SMAC was required for the activation of caspases in lethal and non-lethal conditions, while this contribution to caspase-activation was much smaller in non-malignant fibroblast lines. In cells with high levels of cytosolic SMAC, SMAC deficiency reduced in vitro migration, invasion and anchorage-independent growth as well as metastasis in a xenograft model in zebrafish. SMAC-deficient cells further showed a reduced activity in interferon signaling, associated with reduced cytosolic presence of mitochondrial DNA and activation of the stimulator of interferon genes (STING), and SMAC expression levels correlated with interferon-induced genes in cancer data sets. We further found that SMAC can regulate mitochondrial morphology and integrity. Finally, high gene-expression of SMAC was associated with poor prognosis in patients of several cancer types. These results identify SMAC as a regulator of inflammation and growth behavior of cancer cells. They further report a mitochondrial function of SMAC and demonstrate a role of SMAC in human cancer biology across several cancer entities.

Introduction

One of the functions of mitochondria is the regulation of apoptosis [1]. Following the integration of upstream signals, BCL-2-family proteins can permeabilize the outer mitochondrial membrane and release intermembrane space proteins that trigger the cytosolic activation of caspases [2]. Outer membrane permeabilization eventually kills the cell; caspase-activation causes the apoptotic morphology and counteracts the inflammatory activity of mitochondrial contents such as mitochondrial (mt) DNA [3].

The release of cytochrome c is indispensable for caspase-activation and apoptosis [4]. SMAC, also known as Direct IAP Binding protein with Low PI (DIABLO), was identified as an inhibitor of IAPs (inhibitor of apoptosis proteins) in the cytosol [5, 6]. SMAC translocates to the cytosol alongside cytochrome c during apoptosis [7] and in the cytosol releases the inhibition of XIAP on caspase-9 [8]. In a number of experimental models, for instance in TRAIL signaling, this XIAP inhibition has been found to be required for apoptosis [9], and the role of the SMAC-XIAP interaction for apoptosis has been modeled mathematically [10, 11]. However, SMAC appears to be variably relevant for apoptosis depending on cell type. Analysis of SMAC-deficient mice suggests no or little importance of SMAC in thymocyte, lymphocyte, embryonic stem cell or fibroblast apoptosis [12]. On the other hand, SMAC was essential for the activation of caspases upon mitochondrial permeabilization in human cell lines [13]. Small molecules have been developed that mimic the molecular function of SMAC [14, 15]. These molecules have identified a potential pro-inflammatory role of SMAC by antagonism of cIAP1/2 [16].

SMAC is a mitochondrial protein: like most mitochondrial proteins it is encoded in the nuclear genome, synthesized in the cytosol and imported into mitochondria. During import, its N-terminus is cleaved by the protease PARL [13]. SMAC has also been proposed to regulate phospholipid synthesis during tumorigenesis [17], and A549 lung cancer cells deficient in SMAC showed reduced growth and tumorigenesis [18]. SMAC may therefore have functions in non-apoptotic cells [19] but it is unclear how it achieves these effects.

During apoptosis, the release of cytochrome c into the cytosol is rapid and complete [20], and the kinetics of SMAC release follow a similar pattern [7]. Recent results show that the release of mitochondrial cytochrome c can be partial when cells are treated with a sub-lethal dose of BCL-2 inhibitor. In this situation, only a small portion of mitochondria permeabilize, and the level of caspase-activation elicited is too low to kill the cell (minority MOMP) [21, 22]. Such sub-lethal signaling has been found in cellular infection with numerous pathogens [23], and during infection with the bacterium Helicobacter pylori, SMAC can be released in the absence of cell death but in sufficient amounts to cause NF-κB-activation [24]. SMAC may therefore have a function in sub-lethal signaling following its release into the cytosol. It is further surprising that SMAC is a mitochondrial protein but its reported molecular functions are exclusively cytosolic. Lastly, an unexpected correlation of SMAC-expression and tumor progression has been reported. Given its function as a pro-apoptotic protein and the general role of apoptosis as a cancer-preventing mechanism, a correlation of high levels of SMAC and a good prognosis in cancer may be expected. However, the opposite was the case in metastatic melanoma: high SMAC-levels were linked to poor survival [25].

We here tested whether non-lethal release of SMAC into the cytosol may be relevant for its biology. We observed spontaneous cytosolic presence of SMAC in human cell lines, which was relevant for the regulation of growth behavior. SMAC levels further regulated the interferon (IFN) system in cancer cells, associated with the cytosolic presence of mtDNA. SMAC-deficient cells showed changes in their mitochondrial morphology and integrity.

Results

Requirement for SMAC in lethal and sub-lethal caspase-activation

As discussed above, the dependency of cells on SMAC for caspase-activation following mitochondrial permeabilization appears to be variable. We deleted SMAC in three human cancer cells lines (HeLa cervical carcinoma, MDA-MB231 breast cancer and 1205Lu melanoma cells) and tested for the requirement for SMAC in caspase-activation (gene deletions for all cell lines used here are documented in Fig. S1). The activation of caspase-3 during induction of apoptosis with the combination of the BCL-2/BCL-X_L-inhibitor ABT-737 and the MCL-1-inhibitor S63845 was almost completely SMAC-dependent in these cell lines (Fig. 1A), expanding earlier results [13]. This clear SMAC dependency is in contrast to the findings in primary cells from SMAC-deficient mice where no apoptosis defect had been observed [12]. We therefore tested non-transformed cells from mouse and human origin. SMAC-deficient human WI-38 fibroblasts and two Diablo-deficient mouse embryonic fibroblast lines established from gene deficient mice did show some reduction in apoptosis sensitivity to BH3-mimetics but this effect was relatively subtle (Fig. 1B). There were some differences in XIAP expression, albeit not statistically significant, that may explain this difference (Fig. S2B). With this sample of cells, it appears that cancer cell lines show a substantially higher dependency than non-malignant cells on SMAC for caspase-activation downstream of cytochrome c-release.

**Fig. 1: SMAC is required for lethal and sub-lethal mitochondrial signaling in cancer cell lines.**

We established conditions where no or very little cell death was induced (“sub-lethal conditions”) using low concentrations of either BH3-mimetic (Fig. 1C, S2A). We measured sub-lethal signals by the caspase-activated DNase (CAD)-dependent appearance of the DNA-damage response signal γH2AX [25]. Again, the signal required SMAC in MDA-MB-231 (Fig. 1D, E), HeLa (Fig. 1F) and 1205Lu cells (Fig. 1G). The required activity of SMAC was the inactivation of IAPs, presumably XIAP, because the addition of the small molecule SMAC mimetic/IAP inhibitor Birinapant reversed the effect of SMAC-loss and permitted caspase-activation, as measured by the γH2AX-signal in MDA-MD-231 (Fig. 1H) and HeLa cells (Fig. 1I).

We have previously reported spontaneous activity of the mitochondrial apoptosis pathway in proliferating cells in culture [25]. Spontaneous caspase-3-activation also depended on SMAC: Western blotting detected cleaved caspase-3 in HeLa and MDA-MB-231 cells, and this required the genomic presence of the functional DIABLO gene (Fig. 2A). Effector caspase (DEVD-cleaving) activity was also detectable, and this activity was substantially reduced in the absence of SMAC in either cell line (Fig. 2B). Active caspase-3 could further be precipitated from control but not from SMAC- or caspase-9-deficient HeLa or MDA-MB231 cells when a biotinylated caspase-substrate was added to the cells prior to cell lysis and processing, avoiding potential artificial activation during preparation of cell lysates (Fig. 2C). The results identify the requirement for SMAC in the activation of caspases in the mitochondrial pathway during lethal and sub-lethal signaling. Intriguingly, such requirement was even seen in the absence of any pro-apoptotic stimulus. This suggests that some SMAC is constantly released into the cytosol in steady state to trigger caspase-activation.

**Fig. 2: A stable cytosolic fraction of SMAC drives constitutive activation of caspase-3.**

A portion of SMAC is found in the cytosol in non-apoptotic cells

Fractionation of cells into cytosolic and mitochondrial fractions showed that a portion of SMAC, and also of cytochrome c, is detectable in the cytosol in unstimulated cells. A signal for SMAC was detected in the cytosol, and this signal varied greatly between cell lines, from a very strong signal in two cell lines from the gastrointestinal tract (AGS, gastric, and CaCo2, colon) to no detectable signal in the metastatic melanoma cell line 1205Lu (Fig. 2D–F, S2C). A cytosolic signal for SMAC was also detected using confocal imaging (Fig. S2D). The cytosolic presence of SMAC was reduced but not abolished by the deletion of BAX and BAK in AGS gastric carcinoma cells (Fig. 2D). Individual deletion of either BAX or BAK had a relatively small effect in AGS (Fig. 2G), and a more pronounced effect in MDA-MB-231 cells (Fig. 2H), suggesting that either protein can contribute to spontaneous release of these proteins into the cytosol. The term “minority MOMP” has been coined to describe the release of cytochrome c from only a minor share of mitochondria [21]. The Induction of minority MOMP by ABT-737 was blocked by forced mitochondrial fusion [22]. We therefore deleted the GTPase DRP1, which is required for mitochondrial fission. Loss of DRP1 appeared to block the spontaneous release of SMAC and cytochrome c completely (Fig. 2I), indicating that both fission and the BCL-2-family can regulate the release. A previous study has suggested that the apoptotic release of cytochrome c is more strongly affected by RNAi-mediated knock-down of DRP1 than the release of SMAC [26]. In MDA-MB231 and HeLa cells, the inhibition of the release of both proteins was similar following DRP1 deletion (Fig. 2J, K).

SMAC can regulate growth in cell lines and fibroblasts

Spontaneous activity of the mitochondrial apoptosis pathway can affect growth behavior of human and mouse cells [25]. Because this spontaneous activity depended on SMAC, we tested the effect of SMAC loss on aspects of growth behavior. SMAC-deficient MDA-MB231 cells showed reduced migration, reduced invasion through matrix and reduced colony formation in soft agar (Fig. 3A). To test whether this phenotype is downstream SMAC’s antagonistic function against XIAP, we also tested XIAP-, and XIAP/caspase-9 deficient cells for colony formation in soft agar. Deletion of XIAP or XIAP/caspase-9 in SMAC-deficient cells did not rescue the defect in colony formation in MDA-MB-231 cells (Fig. 3A), indicating that the SMAC effect was independent of its “canonical” role in antagonizing XIAP. MDA-MB-231 have easily detectable SMAC levels in the cytosol. In HeLa cells, where the cytosolic SMAC levels were lower, there was no significant difference in wound healing or invasion by SMAC deletion but a small decrease in proliferation (Fig. 3B). No cytosolic SMAC was detectable in 1205Lu melanoma cells, and SMAC-deficient 1205Lu cells showed no defect in colony formation in soft agar (Fig. 3C). In AGS cells, which again have high levels of cytosolic SMAC, SMAC deficiency was also associated with a reduction of colony growth in soft agar, which was rescued upon re-expression of SMAC in SMAC-deficient cells (Fig. 3D). The small effect or lack of effect of SMAC deletion in HeLa and 1205Lu cells is most likely linked to the small amount of cytosolic SMAC in these cells (see Fig. 2E). The result in the two Diablo-deficient MEF lines was heterogeneous, with one but not the other showing a clear growth deficiency (Fig. 3E). The data suggest that SMAC, which is spontaneously released from mitochondria into the cytosol, affects growth behavior of cancer cell lines. For in vivo validation of this growth phenotype, we used the zebrafish xenograft model of metastasis. By injection of cells into zebrafish larvae and monitoring metastases in the tail fin, the metastatic activity of human cell lines can be estimated [27]. We co-injected control and SMAC-deficient MDA-MB231 cells into the larvae and found that SMAC-deficient MDA-MB-231 cells formed fewer metastases 48 h after injection in zebrafish (Fig. 3F). This confirms the conclusion that SMAC can drive growth behavior of human cancer cells.

**Fig. 3: SMAC can regulate cellular growth behavior.**

SMAC controls interferon response and mtDNA release

The results of growth and metastatic behavior suggested that SMAC has cytosolic functions beyond the regulation of caspases in apoptosis. We conducted RNA-sequencing in control and SMAC deficient MDA-MB-231 cells. To control for effects due to caspase-activation by SMAC, we included caspase-9-deficient cells. Principal component analysis showed a clear separation of SMAC-deficient cells from control cells while caspase-9 deficiency had only a very small effect (Fig. 4A). Comparing control to SMAC-deficient cells, we identified a set of 1454 differentially regulated genes, while caspase-9-deficient cells showed a difference from control cells only in 13 genes (10 of which were also differentially regulated in SMAC-deficient cells) (Fig. 4B, Table S1). Gene enrichment analysis showed a downregulation of genes known from the IFN response in SMAC-deficient cells compared to control cells (Fig. 4C and S3A).

**Fig. 4: SMAC drives an interferon response characterized by STING phosphorylation and an increase in cytosolic mtDNA in MDA-MB-231 cells.**

An IFN response is often driven by the stimulator of interferon genes (STING), which signals upon recognition of cytosolic DNA by the receptor of dsDNA, cGAS. We identified easily detectable levels of phosphorylation of STING and of the key transcription factor of the IFN pathway, IRF3, in MDA-MB-231 control cells, which was substantially reduced in SMAC-deficient cells. The levels of IRF3 phosphorylation were comparable in SMAC- and in STING-deficient cells, compatible with the interpretation that SMAC-dependent STING activation accounted for a large part of IRF3 phosphorylation (Fig. 4D). A well-characterized ligand of cGAS is mitochondrial DNA (mtDNA), which can be released into the cytosol [28]. We found that SMAC-deficient cells had less mtDNA in their cytosol compared to control cells, indicated by reduced cytosolic levels of the mitochondrial transcription factor A (TFAM), reduced cytosolic staining for cytosolic dsDNA and reduced mtDNA in cytosolic fractions of SMAC-deficient cells measured by quantitative PCR (Fig. 4E, F, G). The total number (mitochondria and cytosol) of dsDNA spots detected in SMAC-deficient MDA-MB-231 was similar to that of CTRL cells (Fig. S3B). No differences were observed in the levels of mtDNA detected in the cytosolic fraction of MDA-MB-231 CTRL and caspase-9 deficient cells (Fig. S3C). Surprisingly, using mitoSIRF [29] we found that SMAC and mtDNA exist in close proximity in the mitochondria (Fig. 4H), potentially suggesting an interaction between both. SMAC-deficient MDA-MB-231 cells further secreted reduced amounts of IL-6, a cytokine that is at least partly regulated by STING [30] (Fig. 4I). To test for this SMAC-dependent regulation of the IFN response in cancer, we analyzed gene sets of the TCGA database. In a large number of cancer entities, we detected a significant positive correlation of SMAC expression and the expression of IFN-regulated genes (Table S2); Fig. 4J shows the data for invasive carcinoma of the breast. These results suggest the surprising conclusion that the loss of SMAC leads to a reduction of spontaneous release of mitochondrial DNA into the cytosol, and this may account for the correlation of SMAC expression and the IFN response in human cancer.

SMAC loss affects mitochondrial characteristics

SMAC therefore appears to have control over the release of mtDNA. mtDNA can be released during apoptosis [31, 32] but various forms of stress can also increase cytosolic levels of mtDNA [33]. The architecture of mitochondrial cristae, the invaginations of the inner membrane into the matrix, appears to be a major regulator of mtDNA release [34]. Because SMAC expression correlates with cytosolic mtDNA presence, SMAC may have a function in determining mitochondrial structure, which will affect spontaneous release of mtDNA. We found that SMAC-deficient MDA-MB-231 cells and Diablo-deficient MEFs had a different morphology of the mitochondrial network, with an increase in longer mitochondria and ellipticity index compared to control cells (Fig. 5A, B). MDA-MB-231 cells showed unaltered mitochondrial mass (Fig. S4A) but the mitochondrial membrane potential was reduced in SMAC-deficient cells (Fig. S4B). The oxygen consumption rate (OCR) of SMAC-deficient MDA-MB-231 cells was unchanged (Fig. S4C). There was no interaction between SMAC and OPA1 or the MICOS complex protein MIC10 detectable by immunoprecipitation (Fig. S4D).

**Fig. 5: SMAC can regulate mitochondrial dynamics and architecture.**

When we visualized mitochondrial structure by electron microscopy, it became apparent that the cristae structure was different in SMAC-deficient cells. These cells had a significantly reduced cristae density (number of cristae along the mitochondria) and showed a pronounced increase in the number of “looped” cristae in their mitochondria (Fig. 5C). The results suggest that SMAC deficiency leads to a structural defect in mitochondria, which is associated with a reduced spontaneous release of mtDNA. We tested whether SMAC deficiency also affected the apoptotic release of cytochrome c. We found reduced release of cytochrome c in SMAC deficient MDA-MB-231, HeLa and HCT116 cells upon treatment with BH3 mimetics or with raptinal, which permeabilizes the outer mitochondrial membrane independent of BAX/BAK [35]. As expected, the sensitivity of the cell lines to these treatments varied. However, the reduction of cytochrome c release was observed in all of them (Fig. 6A–C).

**Fig. 6: SMAC can regulate mitochondrial integrity.**

SMAC expression correlates with survival in cancer patients

Our results show regulation of sub-lethal caspase activation and of in vitro growth characteristics by SMAC in cancer cell lines. We have reported earlier that sub-lethal signals in the apoptosis pathway can drive metastasis in xenograft models and are associated with patient survival [25]. We analyzed the TCGA database for the association of DIABLO mRNA expression and patient survival. As shown in Fig. 7, we found a significant association of high levels of DIABLO mRNA expression and shorter patient survival in a number of cancer entities. The association was significant for lung adenocarcinoma and renal clear cell carcinoma across all stages, and mostly for the later tumor stages in the other tumor entities. An association of enhanced SMAC protein expression and poor prognosis has been reported before in patients with metastatic melanoma [36], and a manuscript available on bioRxiv provides the analysis of a large set of human colorectal cancer patients. In that study, higher protein levels of SMAC were also associated with poor clinical outcome [37]. Together with the functional data shown above, it seems most likely that the activation of sub-lethal signals downstream of mitochondria by SMAC translates into aggressive growth of cancer cells and thereby a poor prognosis of cancer patients.

**Fig. 7: association of DIABLO gene expression with prognosis in cancer patients.**

Discussion

Our results indicate non-apoptotic functions of SMAC in both cytosol and mitochondria. In the cytosol, SMAC can regulate not only lethal but also non-lethal caspase activity. SMAC also regulated mitochondrial morphology and integrity. The results extend the correlation of SMAC expression and tumor progression. One intriguing association is the level of SMAC with IFN signaling in cancer. Although molecular details remain to be clarified, we believe that it is relevant to describe these findings all focusing on the role of SMAC as one study. It will still have to be worked what exactly the molecular function of SMAC in mitochondria is. It seems, however, important to describe the function in regulating mitochondrial morphology and the release of mitochondrial molecules, even though underlying mechanisms remain unresolved at this stage.

It was surprising to see the substantial difference regarding the requirement for SMAC between the tested cells. The observations are however in line with the results that have been reported for individual cell types. Notably, cells from Diablo gene deficient mice showed no apoptosis defect; mostly analyzed were lymphocytes and thymocytes, which may be principally different, but embryonic stem cells and embryonic fibroblasts were also tested [12]. On the other hand, human tumor cell lines near-completely depended on SMAC for caspase-activation following mitochondrial permeabilization [13]. Because this could also be a mouse-human difference, we included a non-malignant human fibroblast line, which turned out also to have a reduced requirement for SMAC for caspase activation. We have no explanation of this differential response in molecular terms. The only obvious possibility might have been a difference in XIAP levels between the cells, which we didn’t find. It therefore appears on the basis of these data that SMAC dependency is found in transformed cells but not in non-malignant cells. The reliance was not complete, and ~10% of cell death was still seen. Because we don’t know what the molecular process is that requires SMAC, it is difficult to know why some cells still die. It seems likely that in some way SMAC is required for caspase-activation in cancer cells. Perhaps levels or preformed complexes of APAF-1 and caspases may regulate this dependency. The analysis is of course limited to a relatively small number of cells and cell lines, and the results will have to be tested in further cells.

SMAC is one of several apoptosis signaling proteins whose expression levels are linked to cancer prognosis. It is clear that apoptosis is an anti-cancer mechanism: to become malignant, cells have to develop apoptosis resistance. The relationship is different however in established cancer. Here, high levels of pro-apoptotic proteins can be associated with poor prognosis – in addition to SMAC, this has been noted for BAX and BAK in melanoma – and high levels of BCL-2 are clearly associated with good prognosis in numerous cancer types [38]. Because low activity in the apoptosis pathway will not only reduce apoptosis but also sub-lethal signals, these correlations may be the result of non-lethal activity in the apoptosis pathway. Sub-lethal signals can drive tumor growth [39], aggressive growth in cancer cell lines and metastasis in xenograft models [25]. The results reported here show that SMAC can regulate the growth behavior of cancer cells, including metastasis in zebrafish. Part of this effect appears to be due to DNA damage and the DNA damage response, as a consequence of sub-lethal activity of the apoptosis pathway.

We have reported spontaneous activity of the mitochondrial apoptosis pathway in cancer cells and in mouse primary intestinal organoids before [25]. The results shown here add the requirement for SMAC, presumably upon its release into the cytosol. The earlier results had suggested that mitochondrial ‘leakage’ occurs in unstimulated cells. Here we confirm this directly. The mechanism is not clear. Earlier studies of sub-lethal signaling have reported that individual fissioned-off mitochondria were particularly sensitive to BH3-mimetic but their permeabilization still required this stimulus [22]. Both BAX and BAK appeared to be able to release SMAC and cytochrome c in healthy cells in culture. Double-deletion of BAX and BAK reduced but did not abolish the release. Loss of DRP1, on the other hand, appeared to block it completely; this may mean that indeed release occurs from mitochondria that have fissioned off from the network. Although the link between fission and apoptosis signaling is still incompletely understood, a role for DRP1 in mitochondrial permeabilization has been suggested to occur through its interaction with BAX [40, 41]. What is still unclear, however, is how DRP1 could regulate mitochondrial permeabilization in the absence of BAX and BAK, as it is suggested by the stronger phenotype of the DRP1-deficient cells (see above). Perhaps mitochondrial fission releases small amounts of intermembrane space content by simple leakage. It will be important to understand the importance of this release for mitochondrial function. During apoptosis, mitochondria are completely permeabilized, the cristae structure is destroyed [42] and at least in some cells the inner membrane ruptures [31, 32]. During minority MOMP, it has been found that individual mitochondria rupture [21, 22] and presumably are removed by mitophagy [43]. We think that the possibility could also be considered that there is leakage of intermembrane space proteins in the absence of ‘lethal’ damage to mitochondria but there is no evidence for that as yet.

Our results suggest a role of SMAC in mitochondrial structure and integrity. SMAC deficiency changed the morphology of mitochondria and reduced the release of cytochrome c upon MOMP. Most cytochrome c is not actually localized in the intermembrane space but within the cristae, which themselves are blocked off from the intermembrane space through cristae junctions [42]. SMAC-dependent structural changes may further shift this balance and thereby reduce the cytochrome c available for release. Similarly, deletion of SMAC reduced the release of mtDNA into the cytosol. Because this release is also under the control of the cristae architecture [34], a regulatory effect of SMAC on cristae structure would have the same effect. The release was independent of caspase-9 and therefore very likely not a feedback from caspases. Spontaneous release of mtDNA has been found to occur through VDAC pores [44]. It may therefore be a function of SMAC to regulate VDAC, and its loss may interfere with that function. Release of cytochrome c was also affected by the loss of SMAC. Because the greater part of cytochrome c (85% in one estimate [42]) is localized to the cristae, cristae structure may also affect the release of cytochrome c. The structural consequences of SMAC deficiency may therefore be the basis for changed mitochondrial integrity. The actual function of SMAC in mitochondria remains unresolved. Surprisingly, we found evidence of co-localization of mtDNA with SMAC. Whether this is direct interaction or an interaction through proteins is unclear. It is also unclear at which site in the mitochondria exactly this interaction takes place. It does, however, open the possibility that the release of the two molecules is somehow linked.

SMAC-deficient MDA-MB-231 cells showed a discrete reduction in mitochondrial membrane potential but no reduction in OCR. These findings are not contradictory but rather reflect the complex regulation of mitochondrial bioenergetics. Depending on the precise conditions, reduced membrane potential can be accompanied by increased, decreased, or unaltered OCR. The outcome depends on what is cause and what is effect, the degree of the perturbation, and the contribution of compensatory mechanisms. For example, although reduced electron transport/oxygen consumption and thereby reduced proton pumping will reduce the membrane potential, changes to complex V activity can compensate for this. Reversely, although strong uncoupling (directly reducing the membrane potential) will lead to increased OCR, mild mitochondrial uncoupling (such as facilitated by endogenous uncoupling proteins), while reducing the membrane potential, does not affect OCR. Our OCR data suggest that the reduced mitochondrial membrane potential in the absence of SMAC is not a consequence of reduced electron transport.

Cytosolic mtDNA is known to drive the IFN response through cGAS/STING stimulation [45]. It seems a likely possibility that this SMAC-dependent regulation of the cytosolic levels of mtDNA is the basis for the correlation of IFN-induced genes and SMAC expression levels in cancer cells. cGAS stimulation in cancer cells affects the immune response to cancer through STING stimulation in infiltrating dendritic cells. In this way, SMAC is likely to be a co-determinant of the anti-cancer immune response, and this mechanism may be relevant to cancer progression and patient prognosis.

Materials and methods

Cell lines and culture conditions

HeLa229 (HeLa) cells (ATCC Cat# CCL-2.1) and MDA-MB-231 cells (ATCC Cat# HTB-26) were cultured in RPMI 1640 medium (Thermo Fisher Scientific, Gibco) with 10% FCS (Sigma–Aldrich, #F7524). Mouse embryonic fibroblasts (MEFs) were generated from wild type and Diablo-deficient embryos and cultured in DMEM (Thermo Fisher Scientific, 41965062) with 10% FCS (Anprotec) and 50 µM 2-mercaptoethanol. WI-38 fibroblasts, a gift from Clemens Schmitt, Berlin, were cultured in DMEM supplemented with 10% FCS (Anprotec), 1% sodium pyruvate, 1% glutamax and 1% (v/v) penicillin/streptomycin (Gibco). The metastatic melanoma cell line 1205Lu (Meenhard Herlyn, Wistar Institute, Philadelphia) was cultured in TU2% melanoma medium containing 80% (v/v) MCDB153 (Sigma–Aldrich, #M7403), 20% (v/v) Leibovitz’s L-15, 2% (v/v) FCS (Thermo Fisher, Gibco), 5 µg/ml insulin (bovine, Sigma–Aldrich, #I4011), and 1.68 mM CaCl2. AGS cells (ATCC CRL-1739; obtained from ECACC, Sigma-Aldrich) were maintained in Ham’s F-12 K medium (Gibco) with 10% FCS. HCT116 (from Christoph Borner), were cultured in DMEM with 10% FCS. Caco-2 cells (Tilman Brummer), were cultured in DMEM with 10% FCS. Jurkat cells (Georg Bornkamm, Munich) were cultured in DMEM with 10% FCS. All cells were cultured at 37 °C with 5% CO₂. Gene-deficient cells were generated by CRISPR/Cas9 genome editing by transducing the cells with the lentiviral vector lentiCRISPR v2 (Addgene; Sanjana et al., [46]) and selection with puromycin (Invivogen). Guide RNAs (gRNA) used for CTRL CAD, SMAC, BAX, BAK and Caspase-9 cells as well as HeLa and AGS cells created with those gRNAs have been described [23,24,25, 47]. For generating Drp1-deficient cells, the gRNA AAATAGCTACGGTGAACCCG was used.

Antibodies

The following primary antibodies were used: anti-γH2AX (#9718S, Cell Signaling), anti-GAPDH (MAB374, Millipore), anti-cleaved caspase-3 (#9661, Cell Signaling Technology), anti-caspase-3 (#9662, Cell Signaling Technology), anti-SMAC (#2954 and #15108, Cell Signaling Technology), anti-cytochrome c (#11940 and #12963, Cell Signaling Technology), anti-phospho-STING (#19781S, Cell Signaling Technology), anti-phospho-IRF3 (#ab76493, Abcam), anti-TFAM (#8076S, Cell Signaling Technology), anti-dsDNA (#ab27156, Abcam) anti-biotin (#SAB4200680-100UL, Sigma-Aldrich) anti-LAMP2 (#49067T, Cell Signaling Technology), anti-BiP (#3177T, Cell Signaling Technology), anti-COX IV-Alexa Flour 488 conjugate (#4853S, Cell Signaling Technology), anti-OPA1 (#67589S, Cell Signaling Technology), anti MIC10 (#31561-1-AP, Proteintech) and anti-TOM20 (#42406, Cell Signaling Technology). The following secondary antibodies were used: Alexa Fluor 488 donkey anti-rabbit (#711-545-152, Dianova), Alexa Fluor 488 donkey anti-mouse antibody (#115-545-062,Dianova), Alexa Fluor 647 donkey anti-rabbit (#711-605-152, Dianova), Cy5 donkey anti-mouse (#715-175-151, Dianova), goat IgG anti-mouse IgG (H + L)-HRP (#115-035-166, Dianova) and goat IgG anti-rabbit IgG (H + L)-HRP (#A6667, Sigma-Aldrich).

Flow cytometry for cleaved caspase-3

Cells were trypsinized, combined with supernatants, centrifuged, and washed twice with DPBS. They were re-suspended in DPBS and fixed in 2% paraformaldehyde (PFA; Morphisto, catalog no. 11762.00500) for 15 min at room temperature. After washing with DPBS containing 0.5% bovine serum albumin (BSA; PAN-Biotech, catalog no. P06-1391500), cells were further washed with DPBS containing 0.5% BSA and 0.5% saponin (Roth, catalog no. 4185.1), stained with an antibody against cleaved caspase-3 and analyzed using a FACSCalibur (Becton, Dickinson).

AnnexinV/PI assay

Cells were harvested, washed and resuspended on ice in 1x AnnexinV Binding Buffer (#556454, BD Pharmingen) then incubated with rh AnnexinV/FITC Recombinant Protein (#BMS306FI-300, Invitrogen) for 15 min. Before measuring cells, propidium iodide was added to a concentration of 5 µg/ml, followed by measuring cells using flow cytometry.

Western blot

Cells were lysed with RIPA buffer in the wells. Samples were mixed with Laemmli loading buffer and heated to 95 °C before loading to SDS PAGE. Nitrocellulose membranes were blocked with 5% milk. Proteins were detected with ECL substrate.

Assay for anchorage-independent growth

Cells were plated in triplicate for each experiment. A 2% agarose solution was heated to 95 °C and combined with medium to prepare a 0.6% agarose basal layer. The cells were then mixed with medium and agarose to achieve a final concentration of 0.3% agarose in 6-well plates. The plates were kept at 37 °C, and 100 µl of cell culture medium was carefully added to each well twice a week. After an incubation period of 2–3 weeks, colonies were stained with a 0.005% crystal violet solution. Images of the plates were captured, and the stained colonies were quantified using ImageJ software.

Migration assay

For HeLa Cells, 5 × 10⁵ cells were plated in a 6-well plate. Upon confluency, a 200 µl pipette tip was used to create a scratch in the monolayer. The wells were rinsed three times with DPBS to remove any non-adherent cells. Medium containing 2% serum was then added, and images were captured immediately after scratching and twice daily in the same region of each well until the scratches had fully closed. The scratch areas were measured using ImageJ software and normalized to the scratch area of the control cell line at 0 h. For MDA-MB-231 cells, the kit Radius™ Cell Migration Assay (#CBA-126, Cell BioLabs) was used according to manufacturer’s instructions. Briefly, 2.5 × 10⁴ cells were plated in the provided 96-well plate. After attachment, the Radius™ Gel is removed and cells are allowed to migrate for 24 h.

Invasion assay

The Falcon® Permeable Support for 24-well plates was used following the manufacturer’s instructions. Individual permeable supports (#353097, Corning) were placed into Falcon cell culture permeable support companion plates (#353504, Corning) and coated with 250 µg/ml Corning Matrigel basement membrane matrix (#354234, Corning) for 2 h. Cells were seeded at a density of 5 × 10⁴ per condition and allowed to adhere overnight. After attachment, serum-free medium was added to the cells, while medium containing 10% FCS was added to the lower chamber of the well. The cells were incubated for 24 h to allow invasion. Non-invasive cells were carefully removed using a wet cotton swab, and the membranes were fixed with absolute methanol before being stained with 1% crystal violet. Once dried, five representative images were captured for each permeable support at 10× magnification. The invaded cells were manually counted using ImageJ software.

Caspase-3 activity assay

Cells from duplicates were pooled and lysed using buffer (#9803, Cell Signaling) supplemented with protease inhibitors (#04693132001, Roche). Ten microliters of the lysate were combined with reaction buffer (90 µl, MDB buffer) containing 11 µM Ac-DEVD-AMC (#I-1660.0005, Bachem), 100 µg/ml BSA, and 0.1% CHAPS, and the reaction was performed in triplicates. Analyses were conducted using a Spark 10 M plate reader (Tecan). To precipitate active caspases, biotinylated VAD-fmk (#sc-311290A, Santa Cruz) was added 3 h prior to cell harvesting. Cells were lysed with RIPA buffer, and aliquots of the lysate were heated to 95 °C in Laemmli buffer. Supernatants were incubated overnight with neutravidin beads (#29204, Thermo Fisher) at 4 °C. The beads were washed with RIPA buffer, re-suspended in Laemmli buffer, boiled, and analyzed via Western blotting to detect active caspase-3.

RNA-sequencing

Library preparation for bulk-sequencing of poly(A)-RNA was done as described previously [25]. Barcoded cDNA of each sample was generated with a Maxima RT polymerase (Thermo Fisher) using oligo-dT primer containing barcodes, unique molecular identifiers (UMIs) and an adaptor. 5′-Ends of the cDNAs were extended by a template switch oligo (TSO) and full-length cDNA was amplified with primers binding to the TSO-site and the adaptor. NEB UltraII FS kit was used to fragment cDNA. After end repair and A-tailing a TruSeq adapter was ligated and 3′-end-fragments were finally amplified using primers with Illumina P5 and P7 overhangs. The library was sequenced on a NextSeq 500 (Illumina) with 65 cycles for the cDNA in sread1 and 19 cycles for the barcodes and UMIs in read2. Data was processed using the published Drop-seq pipeline (v1.0) to generate sample- and gene-wise UMI tables [48]. Reference genome (GRCh38) was used for alignment. Transcript and gene definitions were used according to GENCODE v38. Downstream analysis of feature counts was performed with R Studio. First, genes expressed below 10 read counts were excluded. Then, differentially expressed genes were determined using the R Bioconductor package DeSeq2 (version 1.30.1). Next, differentially expressed genes were filtered according to their adjusted p value (<0.05) and absolute value of log2 fold change (>0.5). Heatmaps were generated using the R package pheatmap (version 1.0.12). Gene set enrichment analysis was performed using the R Bioconductor package fgsea (version 1.16.0) [1] together with the Hallmark pathways from the MSigDb. As suggested by fgsea, all differentially expressed genes were used for the analysis. Ggplot2 (version 3.3.5) package was used to create dot plots of pathway analysis. Data was deposited at Mendeley Data under doi: 10.17632/bk9m6kf8s5.1

Immunofluorescence

Cells were incubated on IBIDI slides, fixed with 4% PFA for 10 min, followed by a 10 min permeabilization with PBS containing 0.2% Triton-X. Samples were blocked with PBS 5% BSA 0.2% Triton-X 0.1% Tween for 30 min. For mitochondrial DNA detection, samples were incubated with primary antibody anti-dsDNA 1:500 and anti-TOM20 1:400 in the blocking solution for an hour. Samples were washed then incubated with Alexa Flour 488 anti-mouse and Alexa Flour 647 anti-rabbit, both 1:500 in the blocking solution for an hour. For mitochondrial dynamics, samples were incubated with anti-TOM20 1:400 for an hour. Samples were washed and incubated with Alexa Flour 488 anti-rabbit for an hour. For SMAC detection, cells were incubated with anti-SMAC 1:300 followed by Alexa Flour 488 anti-rabbit 1:500 Samples were washed afterwards and stained with DAPI for 10 min. Samples were washed and covered with IBIDI mounting medium (#50001-4, IBIDI). Images were acquired using Zeiss LSM 880 with AiryScan. Analysis was done using ImageJ.

Subcellular fractionation and mitochondrial DNA extraction

Cells were detached and re-suspended in DPBS. For detection of mitochondrial DNA in cytosol using qPCR, each sample was divided into two tubes. To extract the cytosol, cells were centrifuged and re-suspended in a solution of 70 mM Tris and 250 mM sucrose at pH 7. On ice, digitonin was added to 100 μg/ml for a minute followed by centrifugation at 920 × g for 3 min. The supernatant was collected as cytosol and the pellet was solubilized in RIPA buffer for western blotting. To isolate mitochondrial DNA from the cytosolic fraction, QIAquick Nucleotide Removal Kit (#28306, QIAGEN) was used according the manufacturer’s protocol. Total DNA was isolated from whole cells with the DNeasy Blood & Tissue Kit (#69506, QIAGEN) according the manufacturer’s protocol. Equal volumes were used for the elution of DNA. Alternatively, cytosolic fraction was isolated by mechanically disrupting the cells using 27G syringe followed by ultracentrifugation as described previously [49].

qPCR

Equal volume of DNA extracted from cytosol or whole cell was mixed with SYBR Green master mix (#4472918, Applied Biosystems) and primers described previously [50, 51], followed by detection using QuantStudio 5 Real-Time PCR detection system (Applied Biosystems). Results were analyzed using QuantStudio Design and Analysis software v.1.5.2. The comparative CT method (ΔΔCT method) was used to quantify cytosolic mitochondrial D-loop region (Primers forward CATAAAGCCTAAATAGCCCACACG, reverse CCGTGAAGTGTTTAAGAGGTTGATA) relative to KCNJ gene (Primers forward GGCGAAAGGCTCCTCTCATT, reverse CCCTTCCTGTTTGTGGTGGG) in the whole cell extract.

Survival and correlation analysis

TCGA data were used for survival analysis assessing expression levels of the gene DIABLO and the pathological stage of tumors. Survival analysis of patients within highest and lowest quartiles of expression was done either with all tumor stages combined or with only late stages using the Xena platform [52]. For correlation analysis, significantly downregulated IFN-regulated genes were identified using all available resources (in vitro/in vivo data of Homo sapiens) on INTERFEROME v2.0 database [53]. The expression of significantly downregulated IFN-regulated genes was compared with expression levels of DIABLO in different tumors for significant correlation using Gene Expression Profiling Interactive Analysis (GEPIA) [2, 54].

Electron microscopy

For transmission electron microscopy (TEM), samples were fixed using 2.5% glutaraldehyde (GA) in 0.1 M sodium cacodylate buffer (pH 7.4) at room temperature for 1 h, followed by an overnight incubation at 4 °C. After fixation, the samples were stored in 0.1 M sodium cacodylate buffer at 4 °C until further processing. Contrasting was done with a single wash in 0.1 M sodium cacodylate buffer, followed by primary contrasting using a solution containing 1% osmium tetroxide (OsO₄) and 1.5% potassium ferrocyanide in the same buffer (1 h on ice). The samples were washed twice with deionized water. A secondary contrasting step involved a 1% osmium tetroxide solution and was carried out on ice for 1 h. Whole-mount staining was performed using a 1% uranyl acetate solution. For embedding, samples were dehydrated followed by a resin infiltration protocol using increasing concentrations of Durcupan in acetone (25%, 50%, 75%). Pure Durcupan infiltration followed (30 min at room temperature and an overnight incubation at 4 °C). Final sample embedding was performed in Durcupan filled PCR tubes by polymerization for 2 days at 61 °C in an oven. Ultrathin sections of 70 nm were cut and briefly stained with lead citrate for 1 min. The imaging was done using a Talos L120C transmission electron microscope operated at 120 kV, with 10 cells imaged at a magnification of 13,500×.

Xenotransplantations in zebrafish

Embryos were obtained from wild-type AB/TL zebrafish (Danio rerio) breeding pairs. 50 embryos per Petri dish were maintained in a solution of egg water (60 µg/mL sea salts) at a temperature of 28 °C. Subsequent to gastrulation, the embryos were transferred to an E3 medium containing 200 µM of 1-phenyl-2-thiourea (PTU), to prevent pigmentation. Thereafter, the embryos were subjected to further incubation at a temperature of 28 °C. For xenotransplantation 48 h post-fertilization (hpf) embryos were dechorinated with fine tweezers and anesthetized with tricaine (150 µg/ml, Sigma-Aldrich). Then, 15 embryos were picked randomly and mounted in 1% low melting agarose in E3 on a glass microscope slide. Approximately 20 nl cells suspension of a mix of 300 MDA-MB-231 CTRL cells labeled with DiD (#V22887, Thermofisher) and 300 SMAC KO cells labeled with DiO (#V22886, Thermofisher) was transferred with glass capillary needles in the perivitelline space between the yolk sac and the periderm of each zebrafish embryo. Following a 48 h period post-transplantation, embryos exhibiting abnormal morphology or exhibiting cardiac and/or yolk edema were excluded from further analysis. From the remaining embryos, only those that exhibited fluorescent metastases in the trunk or tail under a fluorescence binocular microscope (Leica MZ 16F, GFP-Filter) were taken for further analysis. Experiment was not blinded. Quantification was automated.

Mitochondrial mass and membrane potential

Cells were stained with MitoTracker™ green for mitochondria mass and red CMXROS for membrane potential (#M46750 and #M46752 respectively, Thermofisher) according to the manufacturer’s instructions, then analyzed using flow cytometry.

Seahorse analysis

OCR was assessed using extracellular flux analysis on a Seahorse XFe96 Analyzer (Agilent). HeLa and MDA-MB-231 cells were seeded in XFe96 cell culture microplates at a density of 5 × 10⁴ cells per well. On the day of measurement, the culture medium was replaced with XF RPMI medium (pH 7.4) containing 1 mM pyruvate, 2 mM L-glutamine, and 10 mM glucose. Cells were then incubated for 1 h at 37 °C in a CO₂-free environment. Mitochondrial respiration modulators were prepared in Seahorse base medium and injected as specified during the assay. Data were analyzed using Wave software (Agilent, version 2.6.1).

mitoSIRF assay

Proximity of SMAC and nascent mtDNA was measured as described by Lozen et al. [29]. Newly-synthesized mtDNA was labeled using Click-iT™ EdU Alexa Fluor™ 647 Imaging Kit (#C10340, Fisher Scientific). The fluorescence stain was omitted. Instead, the mtDNA was labeled with Biotin-azide (#B10184, Thermo Scientific), which was detected using mouse anti-biotin. SMAC was stained as described in immunofluorescence, omitting the secondary antibody. The proximity was detected using Duolink® Proximity Ligation Assay (#DUO82049-4L, #DUO92013-100RXN, #DUO92005-100RXN and #DUO92001-100RXN, Sigma-Aldrich) according to manufacturer’s instructions. Signal was detected using Zeiss LSM 880 Airyscan.

Quantification and statistical analysis

Experiments were done in three independent biological replicates, unless otherwise indicated, to allow meaningful variance calculation. Reported is mean with standard error of mean (SEM) unless otherwise indicated. For xenotransplantation, we used 14 animals after excluding those in which the injection failed to account for biological variability. For Statistical analysis was done using the software Graphpad prism 8. Data was tested for normal distribution then statistical significance was tested using unpaired t-test with Welch’s correction, unless otherwise indicated, with * for p < 0.05, ** for p < 0.01 *** for p < 0.001. All experiments were included in quantification. For xenotransplantation in zebrafish, one animal was excluded due to injection failure. Experiments were not blinded, but quantification of experiments was automated. For image analysis, the software Fiji was used. For each experiment, an automated macro was developed to eliminate bias.

Data availability

RNA-sequencing data were deposited at Mendeley Data under https://doi.org/10.17632/bk9m6kf8s5.1.

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Acknowledgements

We are grateful for support by the Core Facility for Electron Microscopy (EMcore) at the University Freiburg Medical Center—IMITATE is registered with the DFG (German Research Foundation) under the reference number RI_00555. We thank the staff of the Life Imaging Center (LIC) in the Hilde Mangold Haus (HMH), Albert-Ludwigs-University Freiburg for providing microscopy resources and support in image analysis. (LIC is funded by the Deutsche Forschungsgemeinschaft (DFG)—DFG 91b; Prof. Dr. G. Walz and Dr. R Nitschke, DFG project No. 317784314).

Funding

This work was supported by a grant from the Wilhelm Sander-Stiftung to GH (2019.019.1). Open Access funding enabled and organized by Projekt DEAL.

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Authors and Affiliations

Institute of Medical Microbiology and Hygiene, Medical Center, Faculty of Medicine, University of Freiburg, Freiburg, Germany
Tarek N. Amer, Aladin Haimovici, Susanne Kirschnek, Juliane Vier, Abdul Moeed, Uzochukwu Ukachukwu, Daniela Neugebauer, Philip Neubert, Arnim Weber, Mohamed Tarek Badr & Georg Häcker
EMcore, Internal Medicine IV, Faculty of Medicine, Medical Center, University of Freiburg, Freiburg, Germany
Martin Helmstädter & Severine Kayser
Institute of Molecular Oncology and Functional Genomics, Department of Medicine II and TranslaTUM Cancer Center; TUM School of Medicine, Technical University of Munich, Munich, Germany
Rupert Öllinger & Roland Rad
Institute of Neuropathology, Faculty of Medicine, Medical Center, University of Freiburg, Freiburg, Germany
Olaf Groß
Signaling research centres BIOSS and CIBSS, University of Freiburg, Freiburg, Germany
Olaf Groß, Wolfgang Driever & Georg Häcker
Developmental Biology, Institute Biology 1, Faculty of Biology, University of Freiburg, Freiburg im Breisgau, Germany
Jochen Holzschuh & Wolfgang Driever

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Tarek N. Amer
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Contributions

TNA, AH, SKi, JV, UU, DN, PN, JH and AW performed the experiments. TNA, AM, PN, MH, SKa, RÖ, RR, OG, JH and WD acquired and analyzed data. TNA, AH and GH designed experiments. AH and GH supervised the work. TNA and GH wrote the manuscript. All authors contributed to the discussion of content.

Corresponding author

Correspondence to Tarek N. Amer.

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Competing interests

The authors declare no competing interests.

Ethics

The study used established human cell lines from qualified repositories. The isolation of MEFs from SMAC-deficient mice and control mice was done under the permission granted by the local authority (Regierungspräsidium Freiburg, registration number X-23/03 K). Breeding and holding of zebrafish were approved by Regierungspraesidium Freiburg permit 35-9185.64/1.1, and experiments with embryos were carried out in accordance with the German Animal Welfare Act. No human material was used.

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Supplementary information

Figure S1: validation of gene deficiency in cell lines

41418_2026_1667_MOESM2_ESM.pdf

Figure S2: validation of sublethal doses of BH3 mimetics, the levels of XIAP in human cancer cells and primary cells and cytosolic presence of SMAC

Figure S3: Pathway enrichment analysis in MDA-MB-231 SMAC KO and qPCR for cytosolic mtDNA in caspase-9 deficient cells

Figure S4: Mitochondrial changes in SMAC-deficient cells

Table S1: Differentially regulated genes in MDA-MB-231 cells deficient in SMAC or caspase-9

41418_2026_1667_MOESM6_ESM.xlsx

Table S2: Expression of DIABLO is positively correlated with the expression of interferon-regulated genes in multiple cancer

Supplementary Figure Legends

Uncropped western blots

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Amer, T.N., Haimovici, A., Kirschnek, S. et al. The second mitochondrial activator of caspases (SMAC) regulates growth, inflammation and mitochondrial integrity in cancer cells. Cell Death Differ (2026). https://doi.org/10.1038/s41418-026-01667-z

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Received: 04 April 2025
Revised: 11 December 2025
Accepted: 21 January 2026
Published: 03 February 2026
Version of record: 03 February 2026
DOI: https://doi.org/10.1038/s41418-026-01667-z