Ferroptosis in heart failure

Xinquan Yang; Nicholas K Kawasaki; Junxia Min; Takashi Matsui; Fudi Wang

doi:10.1016/j.yjmcc.2022.10.004

. Author manuscript; available in PMC: 2024 Jul 5.

Published in final edited form as: J Mol Cell Cardiol. 2022 Oct 20;173:141–153. doi: 10.1016/j.yjmcc.2022.10.004

Ferroptosis in heart failure

Xinquan Yang ^1,^#, Nicholas K Kawasaki ^2,^#, Junxia Min ^1,^*, Takashi Matsui ^2,^*, Fudi Wang ^1,^*

PMCID: PMC11225968 NIHMSID: NIHMS2005881 PMID: 36273661

Abstract

With its complicated pathobiology and pathophysiology, heart failure (HF) remains an increasingly prevalent epidemic that threatens global human health. Ferroptosis is a form of regulated cell death characterized by the iron-dependent lethal accumulation of lipid peroxides in the membrane system and is different from other types of cell death such as apoptosis and necrosis. Mounting evidence supports the claim that ferroptosis is mainly regulated by several biological pathways including iron handling, redox homeostasis, and lipid metabolism. Recently, ferroptosis has been identified to play an important role in HF induced by different stimuli such as myocardial infarction, myocardial ischemia reperfusion, chemotherapy, and others. Thus, it is of great significance to deeply explore the role of ferroptosis in HF, which might be a prerequisite to precise drug targets and novel therapeutic strategies based on ferroptosis-related medicine. Here, we review current knowledge on the link between ferroptosis and HF, followed by critical perspectives on the development and progression of ferroptotic signals and cardiac remodeling in HF.

Keywords: Ferroptosis, Heart failure, Iron metabolism, Lipid metabolism, Glutathione homeostasis

1. Introduction

Ferroptosis, a new form of regulated cell death, is characterized by an iron-dependent lethal accumulation of lipid peroxides in the membrane system [1]. The key metabolic factors of ferroptosis include iron, glutathione, and lipid. In general, the disturbance of iron metabolism is critical in the production and activation of lipid peroxides [2]. The latter increases plasma cytomembrane permeability, affects membrane fluidity or ionic function, and promotes the formation of non-range pores, leading to cell death and ferropotic signal propagation in various manners [3–5]. A growing number of studies have identified system Xc⁻-GPX4, GCH1/BH4, FSP1/CoQ10, FSP1/Vitamin K, mitochondria DHODH/CoQH2, and other pathways as mechanisms which suppress ferroptosis through scavenging excess toxic lipid peroxides and ferrous iron [6, 7].

Heart failure (HF) is an increasingly prevalent global epidemic that threatens human health [8]. Although effective treatments are constantly being innovated and improved, patients with HF still have a poor prognosis [9]. Ferroptosis has been identified to play an important pathophysiological role in the development and progression of HF in various settings such as myocardial infarction and cardiomyopathy [10]. Thus, comprehensive understanding of the mechanism underlying ferroptosis is a prerequisite to explore promising targets for improving the prognosis of HF. In this review, we summarize the latest findings in different aspects of ferroptosis and discuss the research literatures of ferroptosis in HF.

2. Signaling of ferroptosis

2.1. Iron homeostasis

Dietary iron including heme and non-heme iron is absorbed by intestinal epithelial cells through two different mechanisms. Heme iron is directly absorbed at the apical site of cells through heme carrier protein1 (HCP1) and then catalyzed by hemoxygenase-1 (HO-1) [11]. The non-heme iron, mainly composed of trivalent iron, transported by divalent metal transporter1 protein (DMT1) [12]. Ferrous iron (Fe²⁺) can be stored in ferritin [13] or released to circulation by ferroportin (FPN) which is the only known protein that exports non-heme iron outside of the cell [14–19]. Circulating transferrin (TF) is responsible for iron transport and binds to transferrin receptor 1 (TfR1) on the cell membrane. Iron is distributed to peripheral tissues and moved into the cell through the internalization of the TF-TfR1 protein complex and subsequent release of iron [20].

Iron dyshomeostasis occurs during iron import, storage, export, and transport, which affects the susceptibility to ferroptosis (Figure 1) [6, 21]. Due to the changing levels of intracellular free Fe²⁺, the Haber–Weiss reaction and Fenton reaction occur dynamically [22]. A state of iron in excess within cells initiates and develops ferroptosis by Fenton reaction-mediated overproduction of reactive oxygen species (ROS) and then rapid amplification of hydroperoxy-phospholipids (PLOOHs) in the membrane, especially through inhibition of antioxidant glutathione peroxidase 4 (GPX4) [23]. Iron can also serve as a co-factor for many enzymes such as lipoxygenases, a family of iron-containing enzymes that catalyze deoxygenation of polyunsaturated fatty acids (PUFA) to produce PUFA hydroperoxides in a stereospecific manner [24]. In addition, phosphorylase kinase G2 (PHKG2) has been reported to increase iron availability to lipoxygenase enzymes, which subsequently promotes ferroptosis via oxidation of PUFAs at bis-allylic positions [25]. For example, 15-lipoxygenases catalyze the formation of pro-ferroptotic hydroperoxyl lipids, such as 15-OOH-eicosatetraenoic (HpETE) [26]. Phosphatidylethanolamine-binding protein 1 (PEBP1) forms a complex with 15-lipoxygenases to reduce HpETE by specifically affecting their substrate competence [26]. Interestingly, arachidonic acid (AA) lipoxygenase12 (ALOX12) has also been functionally linked to p53-mediated ferroptosis, which is independent of acyl-coenzyme A (CoA) synthetase long chain family member 4 (ACSL4) [27].

Heme is important for the function of the P450 system in the endoplasmic reticulum, whereas cytochrome P450 oxidoreductase (POR) has been shown to contribute to the initiation of PL peroxidation during ferroptosis [28]. These findings highlight cellular iron homeostasis and its related redox-based metabolic processes are under exquisite control through both enzymatic and non-enzymatic pathways. Conceivably, any condition that raises intracellular labile iron levels increases the sensitivity of ferroptosis. For example, hepatocyte-specific transferrin knockout mice exhibit significant ferroptosis-mediated liver fibrosis in response to a high-iron diet [20]. At the cellular level, deficiency of TfR1 can be effective in dealing with ferroptosis inducers [29]. In fact, anti-TfR1 and anti-malondialdehyde (MDA), one of the lipid peroxides, adduct antibodies, can serve as effective markers for evaluating the state of ferroptosis [30]. Regulatory mechanisms that promote intracellular iron export lay the foundation of ferroptosis resistance. In contrast, loss of FPN induces ferroptosis-mediated memory impairment and promotes disease development [31]. These also indicate that hepcidin, RNF217, and others regulating the expression of FPN might be regarded as potential targets of ferroptosis [16, 32, 33]. In addition, labile iron pool (LIP) alterations in the cytoplasm or organelles (peroxisome, mitochondria, and lysosome) have also been implicated in ferroptosis [34–36]. Ferritinophagy, the autophagy-mediated degradation of ferritin, induces ferroptosis by increasing LIP levels [37, 38]. In general, mitochondrial Fe²⁺ is used for storage in mitochondrial ferritin or to synthesize heme or Fe-S clusters [39, 40]. Under oxidative stress, heme oxygenase 1 (HMOX1)-mediated degradation of heme, abnormal assembly of iron-sulfur clusters (e.g. suppression of NFS1-mediated activation of IREB2), or irregular mitochondrial iron metabolism (e.g. mitochondria iron transporters SLC25A37 and SLC25A28, outer membrane of mitochondria anchored proteins CISD1 and CISD2, and mitochondria iron storage protein mFTH) can sensitize cellular responses to ferroptosis [41]. Although CD44 has been reported to mediate lysosomal iron uptake through the endocytic vesicle system, increasing sensitivity to ferroptosis, its exact regulatory mechanisms are still unknown [42, 43]. Additionally, understanding the mechanism of iron metabolism and pro-ferroptotic signal propagation between organelles (e.g. ER, Golgi apparatus, lipid droplets, and nucleus) may help to develop novel insights.

2.2. The reprogramming of lipid and cholesterol pathways in ferroptosis

Lipids are essential components of the cellular membrane system and drive cellular functions such as energy supply and signal transmission [68]. Lipid composition creates the structural and functional foundation of diversity in organelles and some subdomains. Among cellular lipids, PLs are an important determinant of triggering ferroptosis. Alkeny chain at the sn-1 position is preferentially occupied by 16:0 or 18:0 saturated fatty acids (SFA) or 18:1 monounsaturated fatty acids (MUFA). The sn2 position-linked chain is predominantly PUFA such as AA (20:4) or docosahexaenoic acid (22:6) [69].

ACSL4 and lysophosphatidylcholine acyltransferase 3 (LPCAT3) are important regulators in synthesizing PUFA-containing phospholipids (PUFA-PLs) [70]. The Fenton reaction-dependent peroxidation of PUFA-PLs, especially AA, leads to a lethal accumulation of PLOOHs in the membrane, overcoming the compensatory capacity of antioxidants and eventually triggering ferroptosis [71]. Peroxidation of PUFAs-PLs requires initiation, propagation, and termination processes, which are dependent on both enzymatic and non-enzymatic pathways. These pathways produce products possessing marked differences due to variations in molecular positional isomerism and stereoisomerism [2, 72]. Lipid peroxide adducts, such as 4-hydroxynonenal (4-HNE) and MDA, or oxidized proteins eventually lead to functional failure of organelles and cell death through formation of ferroptotic-like nano pores [3]. AA and eicosapentanoic acid (20:5) are regarded as primary substrates of ACSL4, which enrich membranes with long chain PUFAs through coenzyme A in an ATP-dependent manner [73]. ALOX15 induction ignites PUFA-PL oxidization into ferroptotic signals, with PEs being the most predominant class of oxidized phospholipids [74]. Indeed, oxidized PEs are specific products of ferroptosis and accumulate in the mitochondria, lysosome, and endoplasmic reticulum, highlighting the role of subcellular organelles in PL peroxidation as well as activating ferroptotic signaling [75, 76]. Docosahexaenoic acid (DHA) containing PUFA with three biallylic groups appears to be more oxidizable than AA. ACSL1 remodels both conjugated and non-conjugated fatty acids of membrane lipids to adjust vulnerability to ferroptosis by promoting the incorporation of conjugated linoleates, including α-eleostearic acid, into specific neutral lipids (e.g. diacylglycerols and triacylglycerols) [77]. The unrestrained accumulation of PLOOHs is widely accepted as the most critical step of ferroptosis, yet it is noted that not all damages caused by lipid peroxidation or general accumulation of ROS could lead to ferroptosis. There is no doubt that the peroxidation of specific lipids in various contexts is involved in the process of ferroptosis.

Exogenous non-oxidizable MUFAs, such as oleic acid and palmitoleic acid, reduce the amount of oxidizable plasma membrane lipids and thereby increase ferroptotic resistance through displacing oxidizable PUFAs in an ACSL3-dependent manner [78]. MUFAs preponderantly compete with PUFAs for the biological reaction of PLs or the chemical/structural characteristics of MUFAs can reprogram lipid metabolism within cellular membranes. Similarly, stearoyl-coA desaturase 1 (SCD1) catalyzes the desaturation of SFAs to MUFAs [e.g. stearic acid (18:0) and palmitic acid (16:0) to oleic acid (18:1) and palmitoleic acid (16:1)] facilitating cell growth and anti-ferroptosis properties [79]. Polyunsaturated ether phospholipids (PUFA-ePLs) act as crucial substrates for lipid peroxidation. Fatty acyl-CoA reductase1 (FAR1) is necessary for catalyzing SFAs to fatty alcohols [80]. The FAR1-ether lipids-TMEM189 axis has been identified as a novel ferroptosis regulatory pathway. Due to its potential antioxidative function, plasmalogens containing vinyl ether moiety linked fatty alcohol at the sn-1 position may be important [81].

Cholesterol participates in the modulation of membrane trafficking, signal transduction, and other biological activities [82]. In the cholesterol synthesis pathway, downregulation of squalene monooxygenase catalyzing the oxidation of squalene to 2,3-oxisdosqualene accounts for ferroptosis resistance in specific cancer cells [83]. Inhibition of HMGCR catalyzing HMG-CoA into mevalonate or high-density lipoprotein receptor has been proposed as a cholesterol depleting therapeutic strategy for ferroptosis-related nanomedicine [84, 85]. Isopentenyl pyrophosphate, an intermediate of cholesterol synthesis, regulates selenocysteine tRNA and therefore affects the synthesis of GPX4 [86]. Decreased CoQ10 synthesis by inhibition of the upstream mevalonate pathway may affect ferroptotic signals, suggesting that cholesterol, sterol intermediates of the mevalonate pathway, and cholesterol derivatives, are important for future research. Loss of 15-LOXs, reduces the total amount of cholesterol, oxysterols in IL-4-stimulated human macrophages, and cholesterol intermediates through sterol regulatory element binding transcription factor2 (SREBP2)-inhibition [87]. Cells chronically treated with 27-hydroxycholesterol (27HC), an abundant circulating cholesterol metabolite, are resistant to ferroptosis with substantially increased tumorigenic and metastatic capacity. This is possibly through accommodating cellular metabolic stress associated with its activity through lipid uptake and cumulating in upregulation of GPX4 expression [88]. However, the precise role of 27HC in ferroptosis remains undefined. CD36-mediated uptake of cholesterol or fatty acids leads to the exhaustion of CD8+ T cells through uncontrollable lipid peroxidation and ferroptosis [89, 90]. CD8+ T cell-derived interferon altering lipid patterning through activation of ACSL4 in tumor cell fatty acids, mostly palmitoleic acid and oleic acid, promotes tumor ferroptosis [71]. Moreover, dietary n-3 LC-PUFAs exhibits satisfactory anti-tumor activity when combined with diacylglycerol acyltransferase inhibitor (DGATi) or other ferroptosis inducers [91]. n-3 and n-6 PUFAs tend to accumulate in lipid droplets and then induce ferroptosis in tumor cells under ambient acidosis upon exceeding the capacity of lipid droplet triglyceride storage.

2.3. Ferroptosis defense system

The cellular antioxidant system directly neutralizing lipid peroxidation is primarily regarded as a ferroptosis defense mechanism [92]. Based on the distinctive characteristic of subcellular localizations, several ferroptosis defense mechanisms have been reported.

2.3.1. The system XC (−)/GPX4 axis

SLC7A11 and SLC3A2 are referred to as system Xc⁻ amino acid antiporter and mediate the exchange of glutamate and cystine across the plasma membrane to synthesize glutathione (GSH) after series of enzymic reactions (Figure 1). Functional failure of Xc− antiporter caused by loss of expression of transporters or pharmacological inhibitors such as erastin or its analog imidazole ketone erastin (IKE) triggers ferroptosis due to decreased intracellular cysteine and subsequent reduction of GSH concentration [93]. Moreover, methionine, a sulfur donor, synthesizes cysteines via the trans-sulfuration pathway and may act as a compensatory source of cysteine in the absence of Xc− antiporter [94]. It is noted that cystine starvation triggers more severe ferroptosis than the loss of GSH, indicating the presence of additional mechanisms [95]. Increasing enzymatic activity of glutamate-cysteine ligase catalytic subunit alleviates intracellular glutamate toxicity through catalyzing excess glutamate to γ-glutamy-l peptide and thereby protecting against ferroptosis via a mechanism involving non-canonical regulation of glutathione [96]. Cystine starvation has been identified to impair GPX4 protein synthesis at least partly through inhibition of the mTORC1/4E-BP1 axis-mediated protein translation [97]. A screening study using erastin and cystine deprivation identified sideroflexin 1 (SFXN1), a mitochondrial serine transporter, as a top scoring gene [98]. Enhanced levels of two sulfur-containing cysteine-derived metabolites, which may act as antioxidants in the mitochondria upon SFXN1 loss, provide a survival advantage upon cysteine deprivation [99]. In addition, NFS1 is the iron–sulfur cluster biosynthetic enzyme that harvests sulfur from cysteine. Considering the role of NFS1 and CoQ10 in regulating cellular redox homeostasis [35, 46, 100], loss of cystine may form a complex metabolic-gene-protein network to drive pro-ferroptotic signal transduction in different intracellular compartmentalization. Except for cysteine itself, intracellular accumulation of glutamate mediating the susceptibility of ferroptosis needs to be further elucidated.

GPX4, a selenocysteine-containing enzyme, catalyzes PL hydroperoxides to PL alcohols in the presence of reduced GSH [101]. GSH is regenerated by glutathione reductase mediated reduction of GSSG by using reduced NADPH for the electronic transmission. In contrast to other GPXs, GPX4 is the only member capable of reducing oxidized lipids in free form or in complex with lipids and proteins [102]. Based on genetic loss or inactive function of GPX4 leads to embryonic lethality in a lipid peroxidation-dependent manner, highlighting the importance of GPX4 for early development [103, 104]. Studies focusing on identification of its cell-protective functions were performed with heterozygous or conditional knockout mice [105–107], indicating sufficiency of one allele and tissue-specific effect of GPX4 on increased susceptibility to oxidative injury. GPX4 comprises three isoforms encoded by the same gene with different subcellular localizations: cytosolic, mitochondrial and nuclear GPX4 in mammalian cells. Mitochondrial GPX4 has a mitochondrial signal at the N terminus and the signal peptide can be cleaved when it crosses inside the mitochondria [108]. Due to a lack of such signal peptide, cytosolic GPX4 synthesized by using the second translation start codon can be found in subcellular locations such as the cytoplasm, nucleus, and microsome [108]. Nuclear GPX4 encoded from an alternative first exon called exon Ib is believed to be expressed mainly in sperm nuclei [109], however specific disruption in mice does not result in male infertility [109]. Nevertheless, cytosolic GPX4 is the predominant isoform found in somatic tissues and can be targeted correctly to mitochondria through crossing the mitochondria outer membrane and then accumulating in the intermembrane space, which is essential for embryonic survival [110]. In contrast, mitochondrial GPX4 mainly confers normal sperm function and male fertility during spermatogenesis and is dispensable for embryonic survival since such form may not be expressed endogenously in the majority of cell types [110]. Overexpression of cytosolic GPX4 but not the other forms could completely prevent the death of Gpx4-deleted mouse embryonic fibroblasts, suggesting an essential role for cytosolic GPX4 in somatic and germinal cells likely due to its scavenging of lipid peroxides [111]. Interestingly, p53 plays a paradoxical role in ferroptosis [112]. On the one hand, p53 triggers ferroptosis by downregulation of SLC7A11 expression [113]. Unlike GPX4, loss of phospholipase 2β (iPLA2β) does not affect normal development or cell viability in normal tissues, but regulates ferroptosis upon ROS-induced stress, suggesting the suppressive function of iPLA2β for controlling p53-driven ferroptosis in a GPX4-independent manner [114]. GPX4 can convert oxidized PLs into non-toxic lipid alcohols, while iPLA2β cleaves peroxidized lipids, indicating that distinct lipid repair mechanisms determine the different physiological function and regulation of ferroptosis. Activation of iPLA2β is mediated by p53 under low levels of stress and can be diminished under high levels of stress, suggesting high levels of ferroptotic signals facilitate the scavenging of dead cells, but the precise role of the p53-iPLA2β axis in ferroptosis and cancer development needs to be further studied. In addition, intestinal ischemia/reperfusion-induced acute lung injury can be improved by inhibition of apoptosis-stimulating protein of p53 induced ferroptosis in part through Nrf2/HIF-1/TF signaling pathway [115]. ALOX12 is also reported as a critical gene for p53-mediated ferroptosis, but ACSL4 is dispensable for p53-mediated ferroptosis upon GPX4 inhibition [27]. However, in response to cystine deprivation, p53 also inhibits ferroptosis through regulating cyclin-dependent kinase inhibitor 1A (encoding p21) through conservating intracellular glutathione.[116]. Delaying onset of ferroptosis by activation p53-p21 axis might provide novel insights into understanding how cancer cells survive transient exposure to harsh environments, however, how p21 potentially enhances GSH retention in certain context remain to be further investigated. In addition, activating transcription factor 3 (ATF3) is reported to promote erastin-induced ferroptosis by directly binding to the SLC7A11 promoter and then repressing its expression in a p53-independent manner [117]. ATF3 might be required for slight induction of p53 as current findings support the interaction between ATF3 and p53 [118]. However, one important unaddressed question is whether ATF3, p53 and SLC7A11 coordinately mediate cellular ferroptotic state. Therefore, a greater understanding of how p53 regulates ferroptosis in both a canonical and non-canonical manner is required for clinical translation of p53-modulated ferroptosis.

2.3.2. The GPX4-independent anti-ferroptosis mechanisms

GPX4-independent ferroptosis defense systems have been identified in cancer cells under epithelial state exhibiting ferroptotic resistance upon loss of GPX4 function. Ferroptosis-suppressor-protein 1 (FSP1), also known as apoptosis-inducing factor mitochondria-associated 2 (AIFM2), complements GPX4 loss by using ubiquinone (CoQ10) to protect against ferroptosis elicited by GPX4 deletion and was identified using unbiased cloning approaches. FSP1 is predominantly cytosolic and localized on the cell membrane, especially the cytosolic surface of the mitochondrial outer membrane [35]. FSP1 acts as NAD(P)H-dependent oxidoreductase to catalyze ubiquinone to its reduced form ubiquinol thereby trapping lipid peroxyl radicals and suppressing ferroptosis [35, 119]. Recently, FSP1 is reported to efficiently reduce vitamin K to hydroquinone which acts as potent lipophilic radical-trapping antioxidants and protect the cell against lethal lipid peroxidation and ferroptosis [7]. Subsequent serial studies have identified several upstream regulatory signals in response to ferroptosis such as the MDM2/MDMX complex [120], miR-4443-mediated FSP1 m6A modification [121], or miR-672-3p-mediated transcriptional control of FSP1 [122].

Beyond ubiquinol metabolism, FSP1-dependent endosomal sorting complexes are necessary for transporting (ESCRT)-III recruitment in the plasma membrane and activating repair mechanisms that regulate membrane budding and fission, contributing to ferroptosis resistance [123]. FSP1 induces caspase-independent apoptosis in humans and mice [124, 125]. Its expression is regulated by p53, which binds to responsive elements in the FSP1 promoter region. Upon DNA damage, increasing expression of p53 acts in apoptosis and senescence. Unlike apoptosis, its activation alone is not sufficient to induce ferroptosis. Instead, p53 is a key regulator of ferroptotic signals in the presence of ferroptosis inducers through regulating key metabolic targets involved in ferroptosis [112]. This finding suggests that the p53-FSP1 axis may exert multiple functions in maintaining cellular homeostasis by regulating the transformation among ferroptosis, apoptosis, and other types of cell death, especially by coordinating the mitochondria and nucleus.

Dihydroorotate dehydrogenase (DHODH) in the mitochondrial inner membrane has been identified to inhibit ferroptosis through reducing ubiquinone to ubiquinol through conversion of dihydroorotate to orotate [46]. This occurs in parallel to mitochondrial GPX4, but independently of cytosolic GPX4 or FSP1. The DHODH inhibitor brequinar effectively suppresses both the growth of GPX4-low tumor cells, in combination with sulfasalazine, for the growth of GPX4-high tumor cell types by inducing ferroptosis, highlighting the antitumor activity of DHODH inhibitors. DHODH and mitochondrial GPX4 act as defensive compensatory systems to inhibit ferroptosis in the mitochondria; with a loss of the two arms triggering ferroptosis.

GTP cyclohydrolase 1(GCH1) has been identified as a rate-limiting enzyme for catalyzing tetrahydrobiopterin (BH4) synthesis and as a potent antagonist of ferroptosis [4]. BH4 is an important cofactor for producing aromatic amino acids, neurotransmitters, nitric oxide, and others in the presence of iron in enzyme catalytic sites and is capable of trapping lipid peroxyl radicals. GCH1-overexpressing cells exhibit potent anti-ferroptotic effects through BH4/BH2 mediated protective functions of phosphatidylcholine phospholipids with PUFAs. In addition, BH4 can be secreted and appears to protect neighboring cells from ferroptosis, suggesting that GCH1-high tumor cells may have a wider range of protection against radiation [4]. Dihydrofolate reductase regenerates oxidized BH4 and blocks its function through genetic or pharmacological inhibition, which synergizes with GPX4 inhibition to induce ferroptosis [98]. Inhibition of GCH1 can enhance erastin-induced ferroptosis by promoting ferritinophagy, suggesting GCH1/BH4 metabolism might serve as a novel ferroptosis defensive mechanism and a therapeutic target[126]. The subcellular compartment in interaction between anti-ferroptosis systems wherein the GCH1–BH4 axis operates remains to be understood.

3. The ferroptosis signaling pathway in cardiomyopathy

3.1. Chemotherapy-induced cardiomyopathy

Doxorubicin (DOX) is commonly used in chemotherapy, but it also has severe cardiac side-effects, such as cardiomyopathy. Previously, DOX has been reported to induce ferroptosis as evidenced by iron chelators and ferroptosis inhibitors uniquely suppressing cardiac remodeling after induction with DOX [57]. Tadokoro et al. have shown that GPX4 overexpression inhibited the progression of DOX-induced cardiomyopathy (DIC), but GPX4 heterodeletion exacerbated cardiac ferroptosis [127]. The finding highlights the interconnection between GPX4-mediated ferroptosis and mitochondrial dysfunction in DIC. Trastuzumab, a monoclonal antibody against HER2 receptor that is a target for breast cancer, induces a significant reduction of cell viability in H9c2 cells whilst increasing intracellular and mitochondrial ROS levels in a dose- and time-dependent manner. Given that one third of intracellular iron in cardiomyocytes is located in the mitochondria [128], mitochondrial iron is considered as a pro-oxidative core. Cardiac ferroptosis may be confined to the subcellular localization of free iron release and lipid peroxidation, with ferroptosis’ uniqueness caused by these different levels in context. Acyl-CoA thioesterase 1, an important enzyme in fatty acid metabolism catalyzing the reaction of fatty acyl-CoA to CoA-SH and free fatty acids [129], exerts a protective effect in DOX-induced cardiac ferroptosis through reshaping bio-membrane free fatty acids composition and desensitizing cardiomyocytes to ferroptosis induction [130]. Mitochondrial aldehyde dehydrogenase counters ACSL4-mediated lipid peroxidation and ferroptosis, which may also occur in the maintenance of cardiac homeostasis in a DOX-induced setting [131]. Additional studies have reported cardiac TRIM21-P62-Keap1-Nrf2 axis [132], SIRT1/Nrf2 [133], MITOL/MARCH5 [134], METTL14-KCNQ1OT1-miR-7-5p-TFR1 [135], NLRP3-MyD88 [136] and AMPK-dependent signaling pathways [137, 138] may also be considered as therapeutic approaches to control DIC.

3.2. Hypertrophic Cardiomyopathy

While compensated hypertrophy in the myocardium is initially observed under hemodynamic stress, the continued pathological stress causes adverse LV remodeling and finally leads to HF [139]. In animal models of cardiac hypertrophy, pressure-overload [140–143], infusion of angiotensin II (Ang II) [144, 145] or isoproterenol [146] cause HF, accompanied by excess lipid peroxidation and decreased levels of GPX4 and ferritin heavy chain 1(FTH1). In mice, pressure overload induced cardiomyopathy increased MDA and 4-HNE. Deletion of Ncoa4 reduced both the ratio of Fe²⁺ to FTH1, representing a reduction in free iron, and Ptgs2. Administration of Fer-1 improved cardiac function and reduced MDA, Ptgs2, and fibrotic scarring [38]. Puerarin, a phytoestrogen with antioxidant properties, prevents HF and exhibits stereotypical anti-ferroptotic signals through significant inhibition of ROS [140]. Pressure overload promotes higher levels of Mixed Lineage Kinase 3(MLK3) resulting in a decrease in GPX4 and xCT, culminating in ferroptosis [143]. In response to pressure overload, Mlk3 knockout mice exhibited lower ROS levels, reduced fibrotic scaring, higher GPX4/xCT levels, and higher cardiac functionality compared to controls [143]. Ang II induced hypertrophy initially downregulated xCT, but by week two xCT was upregulated. xCT inhibition increased expression of hypertrophic markers (BNP and Myh7) and reduced left ventricle ejection fraction (LVEF), while treatment with Fer-1 reversed hypertrophic markers and LVEF levels to near original [144]. The expression of elabela (ELA), a second endogenous ligand for the apelin receptor, reduces hypertension induced by Ang II infusion [145]. Treatment with ELA ameliorates cardiac ferroptosis, remodeling, and dysfunction in Ang II-induced cardiomyopathy via suppressing the IL-6/STAT3 signaling pathway [145]. When given a high-iron diet, mice lacking ferritin exhibit severe cardiac injury and hypertrophic cardiomyopathy, with marked molecular features of ferroptosis, however, overexpressing Slc7a11 in these mice prevented cardiac ferroptosis and remodeling [147]. Hypertrophic cardiomyopathy results from various sources, yet ferroptosis and its inhibition has been shown to play a key role in these animal disease models.

3.3. Diabetic Cardiomyopathy

Diabetic cardiomyopathy has been shown to increase MDA, a typical marker associated with ferroptosis due to it being a consistent product of the lipid peroxidation of PUFAs [148] and Ptgs2, a gene that is often upregulated when GPX4 is inhibited and as such is another marker for ferroptosis [149]. Additionally, diabetic cardiomyopathy decreases SLC7A11 and GSH, suggesting the role of ferroptosis in diabetic cardiomyopathy. Treatment with Liproxstatin-1 (Lip-1), which inhibits ferroptosis alongside Fer-1 by stopping PUFA-PL peroxidation [150], improved diastolic function in diabetic mouse models. Sulforaphane improved cardiac function and inhibited ferroptosis through an AMPK-NRF2 axis dependent manner. [60]. Inducing an I/R injury with a cofactor of diabetes increased both overall levels of ACSL4 and specific localized expression in the infarct zone, while inhibition with Fer-1 reduced the amount of ACSL4 and injury size [151]. The role of ferroptosis in diabetic cardiomyopathy has been recently reported, but further studies are needed to fully understand its role.

3.4. Other Cardiomyopathies

In response to sepsis, nuclear receptor coactivator 4 (NCOA4)-mediated degradation of ferritin leads to the accumulation of cytoplasmic ferrous iron and subsequently activates the expression of SFXN1 on the mitochondrial membrane, transporting cytoplasmic ferrous iron into the mitochondria and giving rise to mitochondrial ferroptosis, indicating ferritinophagy-mediated ferroptosis as critical mechanism in sepsis-induced cardiomyopathy [152]. AMPK or islet cell autoantigen 69-STING-mediated ferroptosis signaling have also been implicated in this setting [153, 154]. In alcohol cardiomyopathy, which frequently exhibits atrial fibrillation (AF), ferroptosis plays an essential role in its pathogenesis [155]. Recently, SARS-CoV-2 infection-associated ferroptosis in cardiac SARS-CoV-2 has been discovered and two drug candidates including deferoxamine and imatinib have been identified as specific ferroptosis inhibitors, suggesting that ferroptosis may be a potential mechanism for causing arrhythmias-related cardiomyopathy in patients with COVID-19 [156].

4. Ferroptosis in Myocardial Ischemia-Reperfusion (I/R) Injury

Although myocardial reperfusion therapies, such as primary percutaneous coronary intervention (PCI), reduce infarct size in patients with acute myocardial infarction, reperfusion induces further injury in the myocardium, known as reperfusion injury [157, 158]. Previous studies suggest that lethal reperfusion injury accounts for up to 50% of the final myocardial infarct size [158].

The pathophysiology of myocardial I/R injury is not well understood but is believed to have numerous factors such as calcium, inflammation, and ROS [159]. Ferroptosis has become an important area to further understand the pathophysiology behind I/R injury given their shared connection to iron metabolism [160]. The role of iron in the Fenton reaction and subsequent accumulation of lipid peroxidation necessitates examining the relationship of iron and I/R injury as it pertains to ferroptosis.

Ferritin, specifically FTH1, helps regulate stored iron by converting it between Fe²⁺ and Fe³⁺ [161]. Iron metabolism regulation is important for maintaining the heart’s homeostatic functions. Yet, iron accumulation, resulting from MI, plays a key role in adverse left ventricle (LV) remodeling and is important to track in relation to I/R injury [160]. T2* magnetic resonance imaging (MRI) can be used to detect intra-myocardial hemorrhaging and is a predictor for adverse LV remodeling [162]. Furthermore, using T2* MRI mapping, a larger amount of residual iron was localized to the infarct zone of patients who suffered from STEMI, suggesting a correlation between residual iron and adverse LV remodeling [163]. With iron being a key factor in LV remodeling, it is then necessary to understand the regulation of iron after I/R injury and its overarching effects.

Regulation of the LIP is a key factor in reducing cell death after an I/R injury. BTB domain and CNC homolog 1 (BACH1) is key in iron metabolism with knockout of BACH1 leading to an increase in FTH1 and a reduction in both labile iron and the overall infarct size after I/R injury, showcasing a relationship between labile iron and infarct size [5]. In mouse models, ligation of the left anterior descending artery (LAD) inducing I/R injury leads to reduced GPX4 activity alongside higher levels of iron and embryonic lethal-abnormal vision like protein 1 (ELAVL1), which directly binds to and stabilizes Beclin-1 mRNA. Inhibition of ELAVL1 with siRNA reduces intracellular iron levels, infarct size, and lipid hydroperoxide levels [164]. Other groups have shown MIs increase DMT1 expression, while inhibition of DMT1 leads to lower Fe²⁺ and reduced infarct size; suggesting a relationship between I/R injury and iron regulation [165].

Cardioprotective proteins, like mTOR, confer resistance against iron-induced cell death. Transgenic overexpression of mTOR reduced cell death in isolated cardiomyocytes exposed to excess Fe³⁺ [166]. Cyanidin-3-glucoside (C3G), a flavonoid, has been shown to reduce FTH1 and TfR1, thereby reducing available Fe²⁺. Administration of C3G reduces both infarct size in I/R injury and cell death of H9c2 cells exposed to hypoxia/reperfusion (H/R) [167]. While the presence and regulation of iron can be localized to the infarct region and correlated to its size; the role of the antioxidant system is equally important in understanding ferroptosis during I/R injury.

In terms of lipid peroxidation and I/R injury, multiple studies have found an increase in MDA in infarct regions, emphasizing the role of ferroptosis [165]. After ligation of the LAD, GPX4 is downregulated during the first one to seven days after ligation, but is upregulated by the thirty-day mark. Uniquely, GPX4 is the only member of the glutathione peroxidase family to be downregulated after an ischemic event. Additionally, H9c2 cells exposed to H/R conditions exhibited a similar reduction in GPX4 [168]. Due to a reduction in GPX4, groups have found markers of ferroptosis such as elevated oxidized phosphatidylcholines (OxPCs) in rats after coronary ligation. OxPCs applied to cardiomyocytes in vitro causes elevated cell death, while inhibition of OxPCs reduces both cell death in vitro and the infarct size of rat hearts suffering from I/R injury in vivo [169]. OxPCs have a negative feedback effect on GPX4 as application of them to isolated cardiomyocytes results in a decrease in GPX4 activity and increases cell death. Ferroptosis inhibitors, such as Fer-1, have been shown to have an ameliorating effect on cell death caused by I/R injury. Fer-1 applied during reperfusion improved contractile function after I/R injury and through nuclear magnetic resonance is shown binding to OxPCs [170]. Subsequently, Fer-1 has been shown to decrease both creatine kinase and arachidonic acid metabolites, hydroxyeicosatetraenoic acid and epoxyeicosatrienoic acid, after I/R injury, highlighting the role of ferroptosis in I/R injury. Additionally, administration of Fer-1 reduced cell death in the LV when applied during heart transplant in mouse models [171]. Groups have reported that GPX4 activity reduction is specific to the reperfusion stage and is reduced after one hour of reperfusion following ischemic injury. While GPX4 reduction remains constant throughout reperfusion, ACSL4 activity increases throughout the entirety of reperfusion. DFO, an iron chelator, applied during reperfusion, restored GPX4 and reduced infarct size [172]. Lip-1, a ferroptosis inhibitor, has also been shown to reduce infarct size after I/R injury by around 20%. Lip-1 reduces voltage-dependent anion channel 1 (VDAC1) and increases GPX4. While ROS levels were lowered with Lip-1, the calcium required to open pores remained unchanged [172]. Even without the presence of a ferroptosis inhibitor, transgenic upregulation of GPX4 is sufficient to improve contractile function after I/R injury [173]. Regulation of iron or traditional anti-oxidation through GPX4 reduces cell death and lowers infarct size after I/R injury.

Non-traditional ferroptotic inhibitors also reduce I/R injury infarct size through antioxidation. C3G, while an iron regulator, has been shown to, either by proxy or directly, reduce ROS and MDA resulting in a smaller infarct size [167]. Baicalin, another flavonoid, reduces ROS levels and infarct size, but had a more compensatory interaction with ACSL4 [174]. USP22, a deubiquitinase, reduces I/R injury through upregulating the antioxidant system by increasing expression of SLC7A11. Additionally, USP22 downregulates p53 and acts as a counterbalance against ferroptosis [175]. Similarly, USP1 inhibits TfR1 and p53, but USP7 is upregulated in I/R injury. Inhibition of USP7 has been shown to decrease infarct size, creatine kinase levels, and lipid peroxide levels [176]. Propofol, common in general anesthesia, protects the heart against I/R injury by inhibiting the p53 pathway, which is regulated by Akt, and restoring GPX4 [177]. Importantly, reduction of ferroptosis generated ROS is beneficial in reducing cellular damage after an I/R injury with both traditional and non-traditional ferroptosis inhibitors.

5. HF with preserved ejection fraction (HFpEF) and ferroptosis

Clinical studies suggest that HF with preserved ejection fraction (HFpEF) is more dominant, compared to HF with reduced ejection fraction (HFrEF) as HFpEF now comprises more than 50% of all the patients with HF [178]. A recent clinical study showed that SGLT2 inhibition led to a lower risk of hospitalization for patients with HFpEF [179]. However, mechanisms underlying HFpEF remain undefined. In rats, cardiac function in HFpEF was preserved by an SGLT2 inhibitor, along with the suppression of ferroptosis [180], suggesting that ferroptosis could play an integral role in furthering understanding of HFpEF. One of the established models of HFpEF is created by a combination of high fat diet (HFD) and hypertension induced by L-NAME (constitutive nitric oxide synthase inhibitor) [181]. A study using HFpEF mice created by HFD and L-NAME demonstrated that increased inducible nitrous oxide (iNOS) activity is a major pathogenesis of HFpEF by inactivating the inositol-requiring protein 1α (IRE1α)-X-box binding protein 1 (Xbp1s) axis, a key signal in the unfolded protein response [182]. In another study, HFD and L-NAME induced HFpEF resulted in lower levels of GPX4. Treatment with imeglimin, an anti-diabetic medication, restored GPX4 and Xbp1s levels [183]. New models of HFpEF start to elucidate a relationship between HFpEF and ferroptosis, but further research is necessary to understand the role of GPX4 and more broadly ferroptosis in HFpEF. While the mechanism of HFpEF still remains unclarified, further study of HFpEF focused on ferroptosis will provide a potential target for both HFrEF and HFpEF therapies (Figure 2).

Figure 2. — Multiple physiological events ranging from I/R injury to diabetes can cause an oxidative dysfunctional state resulting in ferroptosis. Cell death resulting from ferroptosis in the heart plays a key role in the onset of cardiomyopathy and heart failure. ROS, reactive oxygen species; PLOOH, hydroperoxy-phospholipid; MDA, malondialdehyde; GPX4, Glutathione Peroxidase 4; FSP1, ferroptosis-suppressor-protein 1; GCH1, GTP Cyclohydrolase 1; BH4, tetrahydrobiopterin; Fer-1, Ferrostatin-1; Lip-1, Liproxstatin-1; DFO, deferoxamine; HFrEF, heart failure with reduced ejection fraction; HFpEF, heart failure with preserved ejection fraction.

6. Ferroptosis in non-cardiomyocytes

Fibrosis is thought as an important and intractable factor in the progression of HF, especially in HFpEF [184]. Accumulating evidence demonstrates that iron homeostasis and non-cardiomyocyte function including fibroblasts, endothelial cells and other immune cells are closely related to cardiac inflammation and remodeling and affect HF development [185]. Excess ROS-induced release of non-transferrin bond iron induces endothelial dysfunction and inflammation in vascular endothelial cells, which is strongly improved by low-iron diet and iron chelation therapy [186]. Dysfunction of iron uptake and storage as shown by aberrant expression of TfR1, FTL and FTH1 induces ferroptosis in endothelial cells, which however can be ameliorated by ferroptosis inhibitors [187]. These studies suggest that dysregulation of iron metabolism and ferroptosis can be considered as a major driver in endocardial remodeling and immunologic derangement. A previous study has shown that inhibition of glutaminolysis, an important process of ferroptosis, reduces cardiac injury and dysfunction triggered by I/R [188]. Recent findings highlight the essential role of glutaminolysis for myofibroblast persistence in chronic HF and can be attributable to carbon derived from the glutamine for maintaining the fibrotic phenotype though regulating bioenergetics, anabolism, collagen biosynthesis, and even epigenetics [189]. Nevertheless, there is a need for more direct evidence demonstrating that ferroptosis is involved in the macrophages and other immune cells in the context of HF, thus, additional studies are needed to identify the ferroptosis-related roles of non-cardiomyocytes in the pathological development of HF.

7. Conclusion

Ferroptosis is an ever-growing field that is interconnected with many fields of study due to the ubiquitous nature of lipid oxidation. Novel therapeutic strategies based on ferroptosis-related medicine such as elucidation of the structure of erastin-bound xCT-4F2hc complex [190] are integral to understanding the role of ferroptosis in HF and will be important in developing new potential therapies, both preventative and diagnostic. Subsequent preclinical studies should be based on established phenotypes including pathological markers and other observations to comprehensively explore the potential role of ferroptosis in the different stages of HF and discuss the in-depth molecular interaction and mechanism of different cardiac cells though deep multiomic profiling and functional screening.

Table 1.

Ferroptosis in in vivo animal models of cardiovascular disease

Disease models	Species	Mechanisms in ferroptotic cell death.
Dox-induced cardiomyopathy	Mouse	Nrf2-HMOX1 [57], GPX4 [127], Acot1[130], TRIM21-P62-Keap1-Nrf2 [132], MITOL/MARCH5 [134], METTL14-KCNQ1OT1-miR-7–5p-TFR1 [135], NLRP3-MyD88 [136], AMPK-dependent signaling pathways [137, 138]
Dox-induced cardiomyopathy	Rat	Nrf2 [191], SIRT1[113], HMGB1 [192]
Hypertrophic cardiomyopathy (Pressure-overload models)	Mouse	FTH1 [140], MLK3 [143]
Hypertrophic cardiomyopathy (Pressure-overload models)	Rat	TLR4-NOX4 [141] IRF3-SLC7A11-ALOX12 [142]
Hypertrophic cardiomyopathy (infusion of Ang II)	Mouse	SLC7A11/xCT [144] IL-6, STAT3, GPX4 [145]
Hypertrophic cardiomyopathy (infusion of isoproterenol)	Mouse	FTH1 [146]
Sepsis induced Cardiomyopathy	Mouse	NCOA4 [152]
Diabetic Cardiomyopathy	Mouse	AMPKα2-Nrf2 [60], FTH1 [193]
Diabetic Cardiomyopathy	Rat	AMPK-Nox2 [194]
I/R Injury	Mouse	Bach1[5], ELAVL1 [164], DMT1 [165], OxPCs [169], Cyanidin-3-glucoside [167], ACSL4 [174], Akt [177], USP22 [175]
MI (no reperfusion)	Mouse	GPX4 [168]
I/R Injury	Rat	USP22 [175], Beclin-1 [195], ACSL4 [172], USP7-p53-TFR1 [176]
I/R injury in diabetes	Rat	NOX2-AMPK [194], ER stress [151]

Open in a new tab

Acot1, Acyl-CoA thioesterase 1; ACSL4, Acyl-coenzyme A synthetase long chain family member 4; AKT, Protein Kinase B; ALOX12, arachidonate 12-lipoxygenase; AMPK; AMP-activated protein kinase; AMPKα2, AMP-activated apha 2 catalytic subunit; Bach1, BTB domain and CNC homolog 1; DMT1, divalent metal transporter1; ELAVL1, ELAV like RNA binding protein 1; FTH1, ferritin heavy chain 1; GPX4, glutathione peroxidase 4; HMGB1, hih mobility group box 1; HMOX1, heme oxygenase 1; IRF3, interferon regulatory factor 3; KCNQ1OT1, KCNQ1 overlapping transcript 1; Keap1, Kelch like ECH associated protein 1; IL-6, Interleukin 6; MARCH5, membrane associated ring-CH-type finger 5; METTL14, methyltransferase-like 14; miR-7-5p, microRNA-7-5p; MITOL, mitochondrial ubiquitin ligase; MyD88, myeloid differentiation primary response 88; MLK3, mixed lineage kinase 3; NCOA4, nuclear receptor coactivator 4; NLRP3, NOD-,LRR-and pyrin domain-containing protein 3; NOX4 and NOX2, NADPH oxidase 4 and 2; Nrf2, Nuclear factor erythroid 2-related factor 2; OxPc, oxidized phosphatidylcholine; SIRT1; sirtuin 1; SLC7A11 and SLC3A2, Solute carrier family 7 member 11; STAT3, signal transducer and activator of transcription 3; TFR1, transferrin receptor1; TRIM21, Tripartite motif-containing 21; TLR4, toll-like receptor 4; USP22 and USP7; ubiquitin specific peptidase 22 and 7

Acknowledgements:

The authors thank Kyoko Komai, Wang & Min laboratory members for a critical discussion for the manuscript, and apologize to colleagues whose work cannot be cited in this manuscript due to space limitations. Figure 2 was created with Biorender.com.

Funding:

This work was supported by the National Natural Science Foundation of China (31970689 to J.M., 31930057 to F.W); the National Key R&D Program (2018YFA0507801 to J.M., 2018YFA0507802 to F.W.); the Hawai‘i Community Foundation (MedRes-2022-771 to TM); the NIH grant (P20GM113134 to TM).

Abbreviation

ACSL4: Acyl-coenzyme A synthetase long chain family member 4
ALOX: arachidonate lipoxygenase
DHODH: dihydroorotate dehydrogenase
DMT1: divalent metal transporter1
DOX: doxorubicin
FAR1: fatty acyl-coA reductase1
FPN: ferroportin
FSP1: ferroptosis-suppressor-protein 1
FTH1: ferritin heavy chain 1
GPX4: glutathione peroxidase 4
GSH: glutathione
H/R: hypoxia/reperfusion
HCP1: heme carrier protein1
HF: heart failure
HFpEF: HF with preserved ejection fraction
HfrEF: HF with of reduced ejection fraction
HMOX1: heme oxygenase 1
HMGCR: 3-hydroxy-3-methylglutaryl-coA reductase
HO-1: hemoxygenase-1
I/R: ischemic reperfusion
LPCAT3: lysophosphatidylcholine acyltransferase 3
MI: myocardial infarction
mTOR: mechanistic target of rapamycin
mTORC1: mechanistic target of rapamycin complex 1
NCOA4: nuclear receptor coactivator 4
POR: cytochrome P450 oxidoreductase
ROS: reactive oxygen species
SCD1: stearoyl-coA desaturase 1
SLC7A11 and SLC3A2: Solute carrier family 7 member 11 and 3 member 2
STEMI: ST-elevation myocardial infarction
TF: transferrin
TfR1: transferrin receptor1

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PERMALINK

Ferroptosis in heart failure

Xinquan Yang

Nicholas K Kawasaki

Junxia Min

Takashi Matsui

Fudi Wang

Abstract

1. Introduction

2. Signaling of ferroptosis

2.1. Iron homeostasis

Figure 1. Mechanisms of ferroptosis.

2.2. The reprogramming of lipid and cholesterol pathways in ferroptosis

2.3. Ferroptosis defense system

2.3.1. The system XC (−)/GPX4 axis

2.3.2. The GPX4-independent anti-ferroptosis mechanisms

3. The ferroptosis signaling pathway in cardiomyopathy

3.1. Chemotherapy-induced cardiomyopathy

3.2. Hypertrophic Cardiomyopathy

3.3. Diabetic Cardiomyopathy

3.4. Other Cardiomyopathies

4. Ferroptosis in Myocardial Ischemia-Reperfusion (I/R) Injury

5. HF with preserved ejection fraction (HFpEF) and ferroptosis

Figure 2. The role of ferroptosis in the pathophysiological development of heart failure.

6. Ferroptosis in non-cardiomyocytes

7. Conclusion

Table 1.

Acknowledgements:

Funding:

Abbreviation

References

ACTIONS

PERMALINK

RESOURCES

Cite

Add to Collections

PERMALINK

Ferroptosis in heart failure

Xinquan Yang

Nicholas K Kawasaki

Junxia Min

Takashi Matsui

Fudi Wang

Abstract

1. Introduction

2. Signaling of ferroptosis

2.1. Iron homeostasis

Figure 1. Mechanisms of ferroptosis.

2.2. The reprogramming of lipid and cholesterol pathways in ferroptosis

2.3. Ferroptosis defense system

2.3.1. The system XC (−)/GPX4 axis

2.3.2. The GPX4-independent anti-ferroptosis mechanisms

3. The ferroptosis signaling pathway in cardiomyopathy

3.1. Chemotherapy-induced cardiomyopathy

3.2. Hypertrophic Cardiomyopathy

3.3. Diabetic Cardiomyopathy

3.4. Other Cardiomyopathies

4. Ferroptosis in Myocardial Ischemia-Reperfusion (I/R) Injury

5. HF with preserved ejection fraction (HFpEF) and ferroptosis

Figure 2. The role of ferroptosis in the pathophysiological development of heart failure.

6. Ferroptosis in non-cardiomyocytes

7. Conclusion

Table 1.

Acknowledgements:

Funding:

Abbreviation

References

ACTIONS

PERMALINK

RESOURCES

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