FUNDC1 Protects against Doxorubicin-Induced Cardiomyocyte PANoptosis through Stabilizing mtDNA via Interaction with TUMF

doi:10.21203/rs.3.rs-1773973/v1

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FUNDC1 Protects against Doxorubicin-Induced Cardiomyocyte PANoptosis through Stabilizing mtDNA via Interaction with TUMF

https://doi.org/10.21203/rs.3.rs-1773973/v1

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published 05 Dec, 2022

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Doxorubicin (DOX) is an effective anti-cancer anthracyclines chemotherapy drug with its potentially life-threatening cardiotoxicity severely limiting its clinical application. Mitochondrial damage-induced cardiomyocyte death is considered an essential cue for DOX cardiotoxicity. FUN14 domain containing 1 (FUNDC1) is a mitochondrial membrane protein participating in the regulation of mitochondrial integrity in multiple diseases although its role in DOX cardiomyopathy remains elusive. Here we examined whether PANoptosis, a novel type of programmed cell death closely associated with mitochondrial damage, was involved in DOX-induced heart injury, and whether FUNDC1 regulated cardiomyocyte PANoptosis. FUNDC1 was downregulated in heart tissues in patients with dilated cardiomyopathy (DCM) and DOX-challenged mice. FUNDC1 deficiency aggravated DOX-induced cardiac dysfunction, mitochondrial injury and cardiomyocyte PANoptosis. Further examination revealed that FUNDC1 countered against cytoplasmic release of mitochondrial DNA (mtDNA) and activation of PANoptosome through interaction with mitochondrial Tu translation elongation factor (TUFM), a key factor in the translational expression and repair of mitochondrial DNA, via its 96–133 amino acid domain. TUFM intervention reversed FUNDC1-elicited protection against DOX-induced mtDNA cytosolic release and cardiomyocyte PANoptosis. These findings shed light towards the protective role of FUNDC1 against DOX cardiotoxicity and cardiomyocyte PANoptosis, which provides a potentially therapeutic approach for Dox induced cardiotoxicity.

doxorubicin cardiotoxicity

mitochondrial damage

cardiomyocyte death

FUNDC1

PANoptosis

Doxorubicin (DOX) is a widely employed potent chemotherapeutic agent for a variety of cancers, including leukemias and solid tumors^{1, 2}. Unfortunately, DOX treatment can also cause adverse drug reactions with cardiovascular toxicity as one of the most common and life threatening-side effect^{3, 4, 5}. DOX cardiotoxicity leads to dilated cardiomyopathy (DCM) and heart failure (HF), ultimately, limiting its clinical application ⁶. Although the underlying mechanisms behind DOX-driven cardiotoxicity have been unveiled to some extent, feasible targets for preventing or reversing DOX cardiotoxicity are still lacking.

Mitochondrial perturbations play a critical role in DOX cardiotoxicity ⁷. Upon exposure to DOX, mitochondria are certainly the most extensively impaired intracellular organelles, and mitochondrial injury is believed to represent one of the first events of cardiotoxicity ⁸. Mitochondrial dysfunction along with mitochondrial DNA (mtDNA) damage provoke multiple programmed cell death (PCD) patterns in cardiomyocytes in response to DOX insult, involving apoptosis^{9, 10}, ferroptosis^{11, 12}, as well as necroptosis^{13, 14}. The mammalian mitochondrial membrane protein FUN14 domain containing 1 (FUNDC1) is a mitophagy receptor that interacts with and recruits LC3 to mitochondria for mitophagy¹⁵. Several studies have shown that FUNDC1 also exerts vital roles in regulating mitochondrial dynamics¹⁶, maintaining mitochondrial calcium homeostasis¹⁷, and participating in cellular plasticity by sustaining oxidative bioenergetics, buffering ROS production, and supporting cell proliferation¹⁸, thus to prevent mitochondrial damage in multiple diseases^{17, 19, 20}. Nevertheless, the role of FUNDC1 in DOX-induced cardiotoxicity has not been clarified.

PANoptosis is not only a newly recognized proinflammatory programmed cell death pattern, but also an emerging concept that highlights the crosstalk and coordination occurs between the three cell death patterns, pyroptosis, apoptosis, and necroptosis^{21, 22}. PANoptosis is driven through a multiprotein complex named the PANoptosome and enables crosstalk and co-regulation among these processes^{23, 24}. Recent studies have showed that the PANoptosome contains molecules critical for pyroptosis, apoptosis, and necroptosis including gasdermin D (GSDMD), Caspase1, Caspase3, Caspase8, RIP kinase 1 (RIPK1) as well as RIP kinase 3 (RIPK3), and is able to activate all of the three pathways to execute cell death^{25, 26}. The conceptualized complex would form a flexible skeleton, whereby the core components of different cell death patterns may be recruited to execute cell death^{23, 25}. Studies focused on the mechanisms manipulating these cell death patterns have revealed that several molecules are capable of regulating pyroptosis, apoptosis, and necroptosis. Z-DNA-binding protein 1 (ZBP1) was found to be crucial for the activation of all the three pathways^{21, 25}. Additionally, cytosolic double-stranded DNA (dsDNA) senser AIM2 was identified to be critical for the activation of PANoptosis by forming AIM2-PANoptosome^{27, 28}. AIM2, inflammasome sensor pyrin, and ZBP1 were members of the large multi-protein complex PANoptosome, along with Caspase1, Caspase8, RIPK3 and RIPK1^{27, 28}. Except for sensing dsDNA of the invading virus, AIM2 was also demonstrated could be activated by mtDNA release in metabolic diseases²⁹. However, whether PANoptosis drives cardiovascular pathology especially DOX cardiotoxicity remains largely unknown.

In this study, we revealed induction of PANoptosis by DOX treatment in cardiomyocytes. We found that FUNDC1 was downregulated in heart tissues from patients with dilated cardiomyopathy (DCM) and mice with DOX challenge. While FUNDC1 deficiency in mice aggravated cardiac function, mitochondrial damage and cardiomyocyte PANoptosis induced by DOX. Our mechanistic approach revealed that FUNDC1 defended against mtDNA release to cytoplasm and resultant PANoptosome activation, through interaction with mitochondrial Tu translation elongation factor (TUFM), a key factor in the translational expression and repair of mitochondrial DNA^{30, 31}, via its 96–133 amino acid domain. TUFM ablation reversed the protective effect of FUNDC1 on mtDNA cytosolic release and cardiomyocyte PANoptosis induced by DOX. These findings illustrated the protective role of FUNDC1 in DOX-driven cardiotoxicity and cardiomyocyte PANoptosis, offering a potentially therapeutic approach for DOX cardiotoxicity.

FUNDC1 deficiency aggravated DOX-induced cardiac dysfunction in mice

To evaluate levels of FUNDC1 in DCM and DOX-challenged heart tissues, heart tissues from patients with DCM and healthy donors were evaluated. Levels of FUNDC1 were significantly downregulated in heart samples from DCM patients compared with healthy donors (Fig. 1a-1b). Next, a mouse model of DOX-induced cardiac injury was established using intraperitoneal injection with DOX (5 mg/kg, four doses, once per week) or an equal dose of saline prior to assessment of myocardial function at 1, 2, 3 and 4 W following completion of DOX challenge (for 4 weeks) (scheme shown in Fig. 1c). A downregulation of FUNDC1 was observed in heart tissues at 1, 2, 3 and 4 W following DOX exposure, with a most pronounced effect at 1 W (Fig. 1d-1e). Besides, echocardiographic analysis revealed that DOX impaired fractional shortening at 1, 2, 3 and 4 W following DOX exposure, with the most pronounced response at 1 W (Fig. 1f). Thus, 1 week following completion of the 4-week DOX treatment was used for further analysis (expect for Masson staining for fibrosis analysis). Besides, AC-16 cardiomyocytes were treated with DOX (1 µM, 24 hrs) to evaluate the expression of FUNDC1. DOX treatment downregulated FUNDC1 levels in AC-16 cardiomyocytes (Fig.S1a-S1b). These results denoted downregulated FUNDC1 in human DCM heart tissues and in DOX-challenged mouse heart tissues and human cardiomyocytes.

To discern the potential role of FUNDC1 in DOX-induced cardiac injury, FUNDC1 knockout transgenic (FUNDC1^−/−) and wild type (WT) mice were challenged with DOX. FUNDC1 knockout efficiency was verified using western blot (Fig. 1g-1h). DOX challenge overtly decreased mouse survival rate, with a more pronounced response in FUNDC1 ablation although FUNDC1 knockout itself did not affect mouse survival (Fig. 1i). Meanwhile, DOX challenge overtly decreased heart weight-to-tibial length (TL) ratio, the effect of which was exacerbated by FUNDC1 ablation with little effect from FUNDC1 deficiency itself (Fig. 1j). In addition, echocardiographic analysis exhibited that DOX insult decreased fractional shortening, ejection fraction (Fig. 1k-1m) (consistent with Fig. 1f), in conjunction with elevated left ventricular end-diastolic diameter (LVEDD), and left ventricular end-systolic dimension (LVESD) (Fig.S1c-S1d). Although FUNDC1 deficiency itself failed to affect echocardiographic indices, it aggravated DOX-induced changes in cardiac geometry and function (Fig. 1k-1m, Fig.S1c-S1d). Neither DOX challenge nor FUNDC1 deficiency (or both) overtly affected heart rate and left ventricular posterior wall diameter in diastole (LVPWD) in all groups (Fig. 1n and Fig.S1e). Moreover, Masson staining (performed at 4 W following the completion of DOX challenge) revealed an overtly increased fibrosis area in respond to DOX insult, the effect of which was exacerbated by FUNDC1 ablation with little effect from FUNDC1 deficiency itself (Fig. 1o-1p). These results suggested that FUNDC1 deficiency aggravated DOX-induced cardiac injury and cardiac dysfunction in mice.

FUNDC1 deficiency exacerbated DOX-induced cardiomyocyte contractile dysfunction in mice

Consistent with echocardiographic findings, adult cardiomyocytes (AMCMs) isolated from DOX-insulted mice exhibited overtly dampened contractile function, as manifested by decreased peak shortening (PS), maximal velocity of shortening/relengthening (± dL/dt), and prolonged time-to-90% relengthening (TR90), the effect of which was exacerbated by FUNDC1 ablation with little effect from FUNDC1 deficiency itself. Resting cell length and time-to-PS (TPS) remained unchanged in all experimental groups (Fig. 2a-2f).

Next, intracellular Ca²⁺ handling was evaluated using Fura-2 fluorescence microscopy. Adult cardiomyocytes from DOX-insulted mice displayed overtly decreased intracellular Ca²⁺ release in response to electrical stimuli (ΔFFI) and prolonged intracellular Ca²⁺ decay rate, the effect of which was aggravated by FUNDC1 ablation with little effect from FUNDC1 deficiency itself. Resting intracellular Ca²⁺ (Baseline FFI) kept unchanged in all groups (Fig. 2g-2i). These results indicated that FUNDC1 deficiency exacerbated DOX-induced cardiomyocyte contractile dysfunction.

FUNDC1 deficiency worsened DOX-induced cardiomyocyte mitochondrial injury in vivo and invitro

To evaluate the effect of DOX on cardiomyocyte mitochondrial function, transmission electron microscopy (TEM) was performed to evaluate myocardial ultrastructure. Our data revealed pronounced myocardial damage including disrupted mitochondria, distortion of sarcomeres and myofilaments following DOX challenge, the effect of which was exacerbated by FUNDC1 deletion (Fig. 3a). Moreover, DOX impaired O₂ consumption rate (OCR) in AMCMs (from DOX-treated mice) including ATP production and maximal respiration, the effect of which were accentuated by FUNDC1 ablation with little effect from FUNDC1 deficiency itself (Fig. 3b-3d). Next, cytosolic ROS level was evaluated using the DHE staining in vivo, and mitochondrial ROS production and mitochondrial membrane potential (MMP) were examined using the fluorescent dyes MitoSOX and TMRM, respectively, in AMCMs isolated from WT and FUNDC1 deficient mice. DOX treatment overtly elevated ROS levels in heart tissues (Fig. 3e-3f), promoted mitochondrial ROS production (Fig. 3g-3h), albeit declined MMP in AMCMs (Fig. 3i-3j), the effect of which was aggravated by FUNDC1 ablation with little effect from FUNDC1 deficiency itself (Fig. 3e-3j). These findings depicted that FUNDC1 deficiency worsened DOX-induced cardiomyocyte mitochondrial injury in vivo and in vitro.

FUNDC1 protected against DOX-induced cardiomyocyte mtDNA release, mitochondrial damage, and cardiomyocyte death

DOX is known to trigger mtDNA release into cytoplasm⁵. To evaluated mtDNA release in the face of DOX insult and FUNDC1 deficiency or overexpression, cytosolic and nuclear fractions were used to quantitate the level of DNA containing specific mitochondrial (mtCOI) and nuclear (18S rDNA) genes by qPCR (primer sequence listed in Table S1). AC-16 cells were silenced or transfected with FUNDC1 prior to DOX exposure. DOX treatment increased the mtDNA-to-nDNA ratio in AC-16 cardiomyocytes, suggesting that DOX provoked mtDNA release to cytoplasm, the effect of which was deteriorated by FUNDC1 ablation (using siRNA) while it mitigated by FUNDC1 overexpression (plasmid transfection) (Fig. 4a). Along the same line, immunofluorescence staining revealed that DOX challenge boosted the level of dsDNA not co-localized with MitoTracker or DAPI (denoting mtDNA release to cytoplasm), the effect of which was deteriorated by FUNDC1 ablation while it was mitigated by FUNDC1 overexpression (Fig. 4b-4c). Moreover, DOX insult led to cell death (shrunken and brightened cells), the effect of which was worsen by FUNDC1 ablation while alleviated by FUNDC1 overexpression (Fig. 4d). MitoSOX and TMRM staining revealed that FUNDC1 ablation aggravated DOX-induced mitochondrial ROS production and mitochondrial membrane potential collapse, while FUNDC1 overexpression exhibited opposite effect (Fig. 4e-4f, Fig.S2a-S2b). These data illustrated that FUNDC1 protected against DOX-induced cardiomyocyte mtDNA release and mitochondrial damage.

FUNDC1 deficiency promoted DOX-induced cardiomyocyte apoptosis, necroptosis, and pyroptosis (PANoptosis)

PANoptosis is a newly recognized programmed cell death pattern^{21, 22}, while whether PANoptosis participates in DOX-induced cardiotoxicity remains unknown. PANoptosis is driven through a multiprotein complex activated by DNA senser AIM2, ZBP1 and Pyrin to form a flexible skeleton to recruit Caspase1, Caspase3, Caspase8, RIPK1 as well as RIPK3, thus to execute cell pyroptosis, apoptosis and necroptosis concurrently (Fig. 5a). Hence, PANoptosis was evaluated in DCM patient heart tissues. Our result indicated that PANoptosis was activated in heart tissues of DCM patients manifested by the increased expression of AIM2, ZBP1, Pyrin (members of PANoptosome) (Fig.S3a-S3c), active forms of Caspase1 and GSDMD (pyroptosis markers) (Fig.S3d-S3e), active forms of Caspase3 and Caspase8 (apoptosis markers) (Fig.S3f-S3g), and phosphorylation of MLKL, RIPK1, RIPK3 (necroptosis markers) (Fig.S3h-S3j). Next, we examined PANoptosis in DOX-challenged heart tissues in mice. Consistent with the results in human heart tissues, DOX boosted PANoptosis in mouse hearts as evidenced by increased levels of AIM2, ZBP1, Pyrin (members of PANoptosome) (Fig. 5b-5d), active forms of Caspase1 and GSDMD (pyroptosis markers) (Fig. 5e-5f), active forms of Caspase3 and Caspase8 (apoptosis markers) (Fig. 5g-5h), and the phosphorylation of MLKL, RIPK1, RIPK3 (necroptosis markers) (Fig. 5i-5k), the effect of which was aggravated by FUNDC1 ablation with little effect from FUNDC1 deficiency itself (Fig. 5b-5k).

Co-immunoprecipitation (Co-IP) was performed to detect AIM2-PANoptosomes in AC-16 cardiomyocytes with AIM2 siRNA or scrambled siRNA transfection (Fig. 5l). ZBP1, AIM2, Pyrin, Caspase1, Caspase8, RIPK1 and RIPK3 were found in RIPK3 immune complexes, of which without IgG identification (Fig. 5l). Moreover, AIM2 knockdown using siRNA abolished elevated expression of ZBP1 and Pyrin following DOX challenge with or without FUNDC1 deficiency in AC-16 cardiomyocytes (Fig. 5m-5o). These findings manifested that FUNDC1 deficiency promoted DOX-induced cardiomyocyte PANoptosis.

FUNDC1 deficiency promoted Dox-induced cardiomyocyte PANoptosis in a mtDNA-dependent manner

To examine the role of mtDNA in the regulation of PANoptosis by FUNDC1 in DOX-induced cardiotoxicity, nucleoside analog 2'3'-dideoxycytidine (ddC), a selective inhibitor of the mitochondrial DNA polymerase γ ³² was added to AC-16 cells with scrambled or FUNDC1 siRNA transfection in the presence of DOX treatment. As precited, ddC overtly decreased mtDNA level in cytoplasm regardless of the presence of DOX or not (Fig. 6a). Besides, DOX treatment provoked PANoptosis in AC-16 cardiomyocytes as evidenced by upregulated levels of AIM2, ZBP1, Pyrin (members of PANoptosome) (Fig. 6b-6d), the elevated active forms of Caspase1 and GSDMD (pyroptosis markers) (Fig. 6e-6f), upregulated active forms of Caspase3 and Caspase8 (apoptosis markers) (Fig. 6g-6h), and higher phosphorylation of of MLKL, RIPK1, RIPK3 (necroptosis markers) (Fig. 6i-6k), the effect of which was aggravated by FUNDC1 ablation. Interestingly, ddC treatment reversed FUNDC1 deficiency-elicited response (Fig. 6b-6k). These results suggested that FUNDC1 deficiency promoted DOX-induced cardiomyocyte PANoptosis in a mtDNA-dependent manner.

FUNDC1 directly interacted with TUFM through its 96–133 amino acid fragment

To discern possible interacting partners of FUNDC1, Mass spectrometry was performed following immunoprecipitation (IP-MS) in AC-16 cells transfected with FUNDC1 plasmids with Flag tag (FUNDC1-Flag) prior to immunoprecipitation with an anti-flag antibody. We identified that TUFM ranked atop among possible FUNDC1 binding proteins associated with mitochondria (Table S2). PRISM tool analysis revealed that FUNDC1 and TUFM shared structural motif for direct binding. Prediction results from PRISM revealed that FUNDC1 and TUFM may form a protein interaction interface (Fig. 7a). To further explore specific interaction between FUNDC1 and TUFM, truncated mutants of FUNDC1 were constructed with deletion of 2–47 amino acid fragment (Delta-2-47 aa), 69–74 amino acid fragment (Delta-69-74 aa) or 96–133 amino acid fragment (Delta-96-133 aa) (Fig. 7b). WT and truncated variants of flag-tagged FUNDC1 plasmids were transfected with His-tagged TUFM plasmid in AC-16 cardiomyocytes. The result of Duolink proximity ligation assay (PLA) detection revealed that WT and truncated variants of FUNDC1 interacted with TUFM with the exception of mutation of Delta-96-133 aa (Fig. 7c-7d), suggesting an obligatory role for 96–133 aa domain of FUNDC1 in the direct interaction with TUFM. To explore spatial relationship between FUNDC1 and TUFM, AC-16 cardiomyocytes were transfected with Flag-tagged WT or truncated variants of FUNDC1 together with His-tagged TUFM. Consistent with Duolink PLA findings, immunofluorescence staining demonstrated that WT and truncated variants of FUNDC1 co-localized with TUFM with the exception of mutation of Delta-96-133 aa (Fig. 7e-7i). In addition, Co-IP assays also showed that WT and truncated variants of FUNDC1 interacted with TUFM, with the exception of mutation of Delta-96-133 aa (Fig. 7j). These findings suggested that FUNDC1 directly interacted with TUFM through its 96–133 amino acid fragment.

Expression of TUFM was evaluated in heart tissues from WT and FUNDC1^−/− mice with or without DOX insult. Our data revealed that DOX downregulated the expression of TUFM, the effect of which was unaffected by FUNDC1 deficiency (Fig.S4a). Next, to detect whether FUNDC1 affect the distribution of TUFM in the face of DOX treatment, we examined levels of TUFM in mitochondria and cytosol in AC-16 cardiomyocytes, the result indicated that FUNDC1 overexpression increased mitochondrial TUFM content (Fig. 7k, Fig.S4b). These data manifested FUNDC1 recruit TUFM to promote its mitochondrial translocation.

TUFM was responsible for FUNDC1-mediated mtDNA stabilization and inhibition of PANoptosis

To examine the role of TUFM in FUNDC1-elicited regulation of mtDNA and PANoptosis in the face of DOX-induced cardiotoxicity, AC-16 cardiomyocytes were transfected with FUNDC1 plasmid, either alone or with TUFM siRNA, prior to DOX insult. FUNDC1 overexpression was found to protect against DOX-induced mtDNA release, the effect of which was abolished by TUFM deficiency (Fig. 8a-8c). Besides, FUNDC1 overexpression defended against DOX-provoked PANoptosis in AC-16 cardiomyocytes as evidenced by reversed levels of AIM2, ZBP1, Pyrin (members of PANoptosome) (Fig. 8d-8f), active forms of Caspase1 and GSDMD (pyroptosis markers) (Fig. 8g, Fig.S5a), and active forms of Caspase3 and Caspase8 (apoptosis markers) (Fig. 8h, Fig.S5b), and the incremental phosphorylation forms of MLKL, RIPK1, RIPK3 (necroptosis markers) (Fig. 8i, Fig.S5c-S5d), the effect of which was abolished by TUMF ablation with little effect from TUFM deficiency itself (Fig. 8d-8i, Fig.S5a-S5d). In addition, we observed AC-16 cell state transfected with FUNDC1 plasmid, either alone or with TUFM siRNA, prior to DOX insult. We found that FUNDC1 overexpression protected against cell death (shrunken and brightened cells), the effect of which was abolished by TUFM ablation (Fig. 8j). These results suggested that TUFM was responsible for FUNDC1-mediated mtDNA stabilization and inhibition of PANoptosis in cardiomyocytes under DOX challenge.

Salient findings from our present study revealed that levels of FUNDC1 were significantly downregulated in heart tissues of patients with dilated cardiomyopathy (DCM), in heart tissues of mice with DOX insult, as well as in DOX-treated AC-16 cardiomyocytes. In DOX-induced murine model of cardiac injury, a downregulated expression of FUNDC1 was observed in heart tissues at 1, 2, 3 and 4-week following DOX-modeling completion, with the most pronounced response at 1 week. In line with that, echocardiographic analysis showed that DOX challenge impaired fractional shortening of left ventricle at 1, 2, 3 and 4-week following DOX-modeling completion, also with the most pronounced at 1 week, suggesting a connection of downregulation of FUNDC1 with cardiac dysfunction in the face of DOX. We demonstrated that FUNDC1 deficiency aggravated myocardial, cardiomyocyte contractile dysfunction, and cardiomyocyte mitochondrial injury in the face of DOX. Besides, using qPCR and immunofluorescence staining detections, FUNDC1 was found to protect against DOX-induced cardiomyocyte mtDNA release into cytoplasm, which then turn on DNA senser AIM2 to form AIM2-PANoptososme and boosted PANoptosis in cardiomyocytes. Further mechanistic study revealed that FUNDC1 directly interacted with TUFM through its 96–133 amino acid fragment as evidenced by Co-IP, Duolink PLA and co-localization of immunofluorescence staining assays. Moreover, our data suggested that TUFM was responsible for FUNDC1 mediated mtDNA stabilization and PANoptosis inhibition in cardiomyocytes. These results uncovered a protective role of FUNDC1 in DOX-induced cardiac injury through inhibition of mtDNA release and PANoptosis in cardiomyocytes.

PANoptosis, the newly proposed programmed cell death pattern, is believed to provoke severe damage to cells, tissues and organs^{33, 34}. Whereas, whether PANoptosis participates in cardiovascular diseases especially DOX cardiotoxicity remains unclear. Here, by detection of PANoptosis-related proteins and formation of PANoptosomes in heart tissues of patients with DCM and DOX-challenged mice, DOX was found to evoke PANoptosis in cardiomyocytes, which shed some lights towards a novel mechanism of DOX cardiotoxicity. Although studies have revealed that DOX challenge can lead to pyroptosis³⁵, apoptosis^{36, 37} and necroptosis³⁸ in cardiomyocytes, the crosstalk and coordination that occurs among the three cell death patterns have not been explicated. The new concept highlights that during the process of PANoptosis activation, a flexible skeleton formed in PANoptosome and core components of the three cell death patterns are recruited and activated to provoke pyroptosis, apoptosis and necroptosis concurrently and jointly to initiate PANoptosis^{23, 24}. In addition, previous studies focused on the resistant role of PANoptosis in immune response to virus infection³⁹, our study indicates that under pathological condition, mtDNA release is another trigger of AIM2-PANoptosome and PANoptosis, providing more insights into the activation of PANoptosis.

In the present study, TUFM was identified as an interacted partner of FUNDC1. TUFM is a nuclear gene-encoded mitochondrial protein widely expressed in different tissues including the heart⁴⁰. It is critical for mitochondrial respiratory function in addition to its role as a key factor for translation of mitochondrial genes in the governance of amino acid elongation of mtDNA-encoded peptides³⁰. Besides, TUFM also plays an essential role in nucleotide excision repair of mtDNA³¹. Moreover, as a mitochondrion-cytosol dual-localized protein, a majority of TUFM is transported to mitochondria following the completion of synthesis⁴¹. Interestingly, our data suggested that FUNDC1 did not bind TUFM through its inner-mitochondrial domain, but rather through its 96–133 amino acid fragment, an outer-mitochondria domain of FUNDC1, suggesting that FUNDC1 recruited TUFM to promote its mitochondrial translocation, supported by the increased mitochondrial content of TUFM.

FUNDC1 is a mitophagy receptor regulating multiple chronic disease progresses including various heart diseases^{17, 19, 42, 43}. Our results also suggested the role of FUNDC1 as a positive factor in DOX-induced heart injury through fending off mtDNA damage and release. A protective role of FUNDC1 for mtDNA has been previously proposed, mainly through its function of mediating mitophagy and mitochondrial dynamics^{44, 45, 46}. In our study, we unveiled that FUNDC1 helped to defend against mtDNA damage by recruiting a mtDNA-repair protein TUFM and promoting its mitochondrial translocation. These results provide new insights for FUNDC1 function in mitochondrial quality control beyond mitophagy.

In conclusion, findings from our current study have offered a new perspective that FUNDC1 protects against DOX-induced cardiomyocyte PANoptosis and heart injury. Mechanically, FUNDC1 restrains mtDNA release to cytoplasm by direct binding and recruiting TUFM to mitochondria to defend against mtDNA damage and cytosolic release in the face of DOX insult, thus preventing activation PANoptosome and PANoptosis in the face of DOX challenge. To this end, induction or stimulation of FUNDC1 should be considered as a novel and promising strategy in the therapeutics of DOX-induced cardiotoxicity.

Human samples

Human heart samples in the current study were obtained from patients who have undergone heart transplant surgery due to DCM or the doner heart failed with transplantation due to non-cardiac reasons (n = 6/group), from Sun Yat-Sen Memorial Hospital, Sun Yat-Sen University (Guangzhou, China). The study was approved by the institutional ethics committee of Sun Yat-Sen Memorial Hospital.

Experimental animals and Dox challenge

All animal procedures were approved by our institutional Animal Care and Use Committees at the Zhongshan Hospital Fudan University (Shanghai, China) and the University of Wyoming (Laramie, WY, USA). Global FUNDC1 knockout (FUNDC1^−/−) mice were generated as described in our earlier report¹⁷. Wild type (WT) and FUNDC1^−/− mice (6–8 weeks old) were subjected to Dox (5 mg/kg, i.p. four doses, once weekly).

Echocardiographic assessment

Following Dox treatment (for 4 weeks) completion, mice were anesthetized with isoflurane (1–2%) prior to the M-mode echocardiographic assessment. A two dimensional (2D) guided M-mode echocardiography was applied to measure mouse cardiac geometry and function (Vevo 2100, FUJIFILM Visualsonics, Toronto, ON, Canada) equipped with a 22 − 55 MHz transducer (MS550D, FUJIFILM VisualSonics) according to previously published procedures^{47, 48}.

Histological examination

Following anesthesia, hearts were arrested in 10% KCl solution, then exercised and placed in diastole with 10% neutral-buffered formalin for 24 hrs at room temperature. Specimens were embedded in paraffin, cut into 5-µm sections and stained with the Masson trichrome. Percentage of fibrosis was calculated using a digital microscope (× 400) and the Image J (version1.34S) software.

Adult mouse cardiomyocyte isolation, shortening/relengthening and intracellular Ca²⁺ recording

Langendorff perfusion system was employed to isolate cardiomyocytes from mouse hearts following previous protocols⁴⁷. Mechanical properties of cardiomyocytes were assessed using a Softedge MyoCam system (IonOptix Corporation, Milton, MA, USA) with an IX-70 Olympus inverted microscope. Contractile buffer containing NaCl 135 mM, KCl 4.0 mM, CaCl2 1.0 mM, MgCl 1.0 mM, glucose 10 mM and HEPES 10 mM was added and cardiomyocytes were electrically stimulated at 0.5 Hz. Cell shortening was assessed including peak shortening (PS), maximal velocity of shortening (+ dL/dt), maximal velocity of relengthening (-dL/dt), time-to-PS (TPS), and time-to-90% relengthening (TR₉₀). For intracellular Ca²⁺ recording, cardiomyocytes were loaded with Fura-2/AM (0.5 µM) for 10 min, and fluorescence measurements were recorted with a dual-excitation fluorescence photomultiplier system (IonOptix). To assess intracellular Ca²⁺ signaling, cells were exposed to light emitted by to light emitted by a 75-W lamp and passed through 360 nm or a 380 nm filter, while being stimulated to contract at 0.5Hz. Fluorescence emissions were detected between 480 and 520 nm and the alterations in fura-2 fluorescence intensity (FFI) were quantitated from the FFI ratio at 360 nm to 380 nm. Fluorescence decay time was assessed as an indicator of intracellular Ca²⁺ clearing^{47, 49}.

Transmission electron microscopy (TEM)

Cubic heart pieces were dissected and immersed with 2.5% glutaraldehyde in 0.1 M sodium phosphate (pH 7.4) for at least 24 hrs at 4°C. Tissues were dehydrated through graded alcohols and were embedded in Epon Araldite then fixed in 1% OsO₄ for 1 hr. Ultrathin sections (75–80 nm) were produced using an ultramicrotome (Leica, Wetzlar, Germany) equipped with a Diatome diamond knife, and were stained with uranyl acetate for 10 min and lead citrate for another 5-min. Specimens were observed under an 40–120 kV transmission electron microscope (Hitachi H600 Electron Microscope, Hitachi, Japan. Images were captured using the Ditital Micrograph software.

Cell isolation, culture and treatment

The protocol to isolate adult mouse cardiomyocytes (AMCMs) was described previously ^{50, 51}. In brief, adult male C57/BL6J mice (8 to 10 weeks old) were anesthetized, then opened the chest to fully exposed heart. Cut inferior vena cava and descending aorta, and inject EDTA buffer immediately into the right ventricle. Then tightly clamped ascending aorta and removed the heart. Next, inject EDTA buffer and perfusion buffer into left ventricles prior to perfusion of a collagenase buffer (type II and IV collagenase). When the tissues became slightly pale and flaccid, cut left ventricle into 1 mm³ pieces in stop buffe. Cell suspension underwent four sequential rounds of gravity settling, using three intermediate Ca²⁺ reintroduction buffers to gradually restore extracellular Ca²⁺ concentration to 1.2 mM. A yield of at least 80% rod- shaped CMs were deemed successful ^{50, 51}.

The human AC-16 cardiomyocytes were cultured in a Dulbecco’s Modified Eagle Medium (DMEM, Gibco, Waltham, Maine state, USA), containing 10% fetal bovine serum albumin (FBS, Gibco, Waltham, USA) at 37°C, 5% CO₂. AC-16 cardiomyocytes were treated with Dox (1 µM, 24 hrs, Sigma Aldrich, MO, USA). To determine involvement of mtDNA, a nucleoside analog 2'3'-dideoxycytidine (ddC, 40 µg/mL for 5 days, Selleck Chemicals, Houston, TX, USA,) was added to AC-16 cells when cells grow to a proper density for further experimentation.

Plasmid construction and transfection

Full-length, domain truncated FUNDC1 plasmid (FUNDC1-Delta-2-47aa, FUNDC1-Delta-69-74aa, and FUNDC1-Delta-96-133) tagged with Flag were cloned in pcDNA3.1+ (Dongxuan Genes, Kunshan, China). TUFM tagged with His and negative controls were cloned in pcDNA3.1+ (Dongxuan Genes, Kunshan, China). Plasmids were transfected into AC-16 cardiomyocytes for 48 hrs.

Mitochondrial respiration measurement

Mitochondrial respiration was measured by analyzing the mitochondrial oxygen consumption rates (OCR) using a Seahorse XFe96 analyzer (Seahorse Biosciences, North Billerica, MA, USA), as our previously described ⁵². In brief, cells were seeded at 40,000 cells/well on 96- well XFe96 cell culture microplates then treated as designed. For respiration assays, cells were incubated in a CO²-free environment for 1 hr, and OCR was measured every 3 min over 90 min. First, OCR was quantified in basal conditions (20 mmol/L glucose), next 1 µM oligomycin (ATP synthase inhibitor), then with 0.125 µM FCCP (mitochondrial respiration uncoupler), and finally with 1 µM rotenone/antimycin A (complex I and III inhibitors, respectively). Seahorse XF-24 software was used to calculate the OCR automatically.

Cytosolic mtDNA isolation

AC-16 cells were lysed in cell lysis buffer (Thermo Fisher Scientific, Foster, CA) and centrifuged (700 × g, for 10 minutes, at 4°C) to remove the nuclei. Then normalized the volume of the supernatant according to the protein concentration. Next, centrifuged the cell lysate (10,000 × g, for 30 minutes, at 4°C) again to isolate the cytosolic fraction, then the DNA in the cytosolic fraction was isolated. The mtDNA was detected using quantitative PCR with the gene sequences coding for mitochondrial cytochrome c oxidase 1 (mtCOI) as primers. The nuclear DNA was measured using sequences coding 18S ribosomal RNA as primers⁵³. The copy numbers of mtDNA were normalized by the copy numbers of nuclear DNA. The primers for human mtCOI and 18S rDNA were listed in Table S1 (Dongxuan Genes, Kunshan, China).

Realtime quantitative PCR

Trizol Reagent (Invitrogen, NY, Empire State, USA) was used to extract total RNA. Then the purity and concentration of RNA were determined using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, Maine state, USA). PrimeScriptTM RT Master Mix (Takara, Shiga, Japan) was used to conduct reverse-transcription for synthesis of cDNA. Realtime quantitative PCR was performed using FastStart Essential DNA Green Master (Roche, Shanghai, China). GAPDH was employed as the reference gene. Relative gene expression was calculated using 2-ddCt method. The primers were listed in Table S1 (Dongxuan Genes, Kunshan, China).

Isolation of mitochondria

Preparation of mitochondria was conducted using the Mitochondria/Cytosol Fractionation Kit (Abcam, #ab65320). Isolation of mitochondria was performed on ice to prepare the mitochondrial and cytosolic fractions used for Western blot analysis.

Western blot analysis

Heart tissues and cells were homogenized and sonicated in a RIPA lysis buffer (Beyotime, Shanghai, China) with protease inhibitor cocktail. Target proteins were separated by SDS/PAGE gel, then transferred to 0.22 µm PVDF membrane. After blocking with 5% bovine serum albumin (BSA), the PVDF membrane were incubated overnight with primary antibodies at 4°C. Protein samples were incubated with anti-Vinculin (Abcam, #ab219649), anti-FUNDC1(Abcam, #ab224722), anti-AIM2 (Abcam, #ab204995), anti-ZBP1 (Abcam, #ab290736), anti-Pyrin (Abcam, #ab195975), anti-Caspase1 (Abcam, #ab207802), anti-Caspase3 (Abcam, #ab32351), anti-Caspase8 (Proteintech, 13423-1-AP), anti-GSDMD (Abcam, #ab209845), anti-MLKL (Abcam, #ab184718), anti-p-MLKL (Abcam, #ab196436) anti-RIPK1 (Cell Signaling Technology, #73271), anti-p-RIPK1 (Cell Signaling Technology, #44590), anti-RIPK3 (Cell Signaling Technology, #10188), anti-p-RIPK3 (Cell Signaling Technology, #93654), anti-p-TUFM antibodies (Abcam, #ab173300). All antibodies were obtained from Cell Signaling Technology (Danvers, MA), Abcam (Cambs, UK) or Proteintech (Wuhan, China). Secondary antibodies were used for membrane incubation. Films were scanned and detected with a Bio-Rad calibrated densitometer.

Structure-based protein interaction interface analysis between FUNDC1 and TUFM

The protein structure of FUNDC1 was predicted by template-based homology structure modeling tool SWISS-MODEL (https://www.swissmodel.expasy.org), using PDB structure 3BK6, chain A (covering residues 74–256, sequence identity = 21.64%), and 2IP6, chain A (covering residues 82–131, sequence identity = 10.00%), as the template, respectively. Structure of TUFM was downloaded from the PDB database (PDB ID:7A5G). Prediction of potential interaction interface between TUFM and FUNDC1 was obtained from PRISM tool (http://cosbi.ku.edu.tr/prism). Prediction results were visualized using the PyMol tool (http://pymol.org).

Co-Immunoprecipitation (Co-IP)

Total protein was extracted from AC-16 cells transfected with plasmids as designed using NP-40 lysis buffer, 50 µL total protein was extracted as input, then protein samples was exposed to the primary antibody of His (Abcam, #ab18184) or isotype control immunoglobulin G (Cell Signaling Technology, USA, #3900S) at 4°C overnight. 20 mL protein A/G agarose beads (Sea Biotech, China, #P001-2) were added into the mixture, and incubated for 3 hrs at 4°C with rotation. Centrifuged the cell lysate 10,000 × g for 30 minutes at 4°C to collect protein A/G agarose beads and washed the beads 3–5 times with NP-40 lysis buffer. Finally, proteins were analyzed by Western blotting.

Immunoprecipitation assay and Mass spectrometry

AC-16 cells were transfected with FUNDC1-Flag plasmids for 48 hrs, and cell lysates were immunoprecipitated with the anti-Flag magnetic beads (Cell Signaling Technology, #82103). Precipitates were separated by SDS-PAGE and stained with coomassie blue subsequently. Then cut the protein bands into small pieces and subjected in-gel digestion. The extracted peptides from the gel pieces were prepared for proteomic data analysis by liquid chromatography tandem mass spectrometry (LC-MS/MS).

Immunofluorescence staining

The treated cells were fixed by 4% paraformaldehyde for 10 min, then permeabilized and blocked them for 1 hr prior to incubation with specified antibodies (dsDNA, Abcam, #ab27156, His, Abcam, #ab18184, and Flag, Abcam, #ab205606) overnight at 4°C, and followed by the incubation of the corresponding secondary antibodies conjugated with Alexa Fluor (Cell Signaling Technology, Boston, USA) for 2 hr in the dark. After staining, examined the immunofluorescence using a laser confocal microscope with a 630 × oil immersion objective with 488 and 561 nm laser excitation (Leica, Wetzlar, Germany). Results were analyzed using a coloc2 plug-in (Fiji, version 2.0, Rawak Software Inc., Stuttgart, Germany) of Image J software.

Measurement of mitochondrial ROS (mtROS)

Cells were incubated with the MitoSox Red fluorescent dye (5 µM, Thermo Fisher, Waltham, Maine state, USA) at 37°C for 30 min. Cells were washed with a warm phosphate buffered saline (PBS) buffer for 3 times prior to observation under a Leica confocal microscopy (Wetzlar, Germany).

Measurement of mitochondrial membrane potential (MMP)

Mitochondrial membrane potential (MMP) was evaluated using tetramethylrhodamine methyl Ester (TMRM) staining and mitochondrial morphology was assessed using MitoTracker staining. Cells were stained with TMRM (20 nM, Molecular Probes, Invitrogen, Invitrogen, Carlsbad, California, USA) and MitoTracker Red solution (20 nM, Molecular Probes, Invitrogen, Carlsbad, California, USA) for 30 min at 37°C, and were imaged using a fluorescence microscope. Image J was used to evaluate red fluorescence intensity.

Statistical analysis

All quantitative data were presented as mean ± standard error of mean (SEM). Results were analyzed using a Prism 8.0 software (GraphPad, San Diego, CA). Comparation between two groups were conducted using the Student’s t-test (two-tailed). Comparation among multiple groups were conducted using one-way ANOVA followed Tukey's test for post hoc analysis. Statistical significance was set at p < 0.05.

ACKNOWLEDGEMENT:

Drs. Jun Ren and Yingmei Zhang are the guarantors of this work and, as such, had full access to all data in the study and takes responsibility for the integrity of the data and the accuracy of data analysis.

CONFLICT OF INTEREST:

None.

AUTHORS CONTRIBUTION:

YB, HX, HZ, XW and JR involved in conception and design, performance of experiments, data analysis and interpretation, and manuscript preparation; JR and YZ involved in study supervision and revision of the manuscript.

FUNDING

This study was supported by grants from the Program of

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FUNDC1 Protects against Doxorubicin-Induced Cardiomyocyte PANoptosis through Stabilizing mtDNA via Interaction with TUMF

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Journal Publication

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Abstract

Figures

Introduction

Results

FUNDC1 deficiency aggravated DOX-induced cardiac dysfunction in mice

FUNDC1 deficiency exacerbated DOX-induced cardiomyocyte contractile dysfunction in mice

FUNDC1 deficiency worsened DOX-induced cardiomyocyte mitochondrial injury in vivo and invitro

FUNDC1 protected against DOX-induced cardiomyocyte mtDNA release, mitochondrial damage, and cardiomyocyte death

FUNDC1 deficiency promoted DOX-induced cardiomyocyte apoptosis, necroptosis, and pyroptosis (PANoptosis)

FUNDC1 deficiency promoted Dox-induced cardiomyocyte PANoptosis in a mtDNA-dependent manner

FUNDC1 directly interacted with TUFM through its 96–133 amino acid fragment

TUFM was responsible for FUNDC1-mediated mtDNA stabilization and inhibition of PANoptosis

Disscussion

Methods And Materials

Human samples

Experimental animals and Dox challenge

Echocardiographic assessment

Histological examination

Adult mouse cardiomyocyte isolation, shortening/relengthening and intracellular Ca2+ recording

Transmission electron microscopy (TEM)

Cell isolation, culture and treatment

Plasmid construction and transfection

Mitochondrial respiration measurement

Cytosolic mtDNA isolation

Realtime quantitative PCR

Isolation of mitochondria

Western blot analysis

Structure-based protein interaction interface analysis between FUNDC1 and TUFM

Co-Immunoprecipitation (Co-IP)

Immunoprecipitation assay and Mass spectrometry

Immunofluorescence staining

Measurement of mitochondrial ROS (mtROS)

Measurement of mitochondrial membrane potential (MMP)

Statistical analysis

Declarations

References

Additional Declarations

Supplementary Files

Status:

Journal Publication

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Adult mouse cardiomyocyte isolation, shortening/relengthening and intracellular Ca²⁺ recording