Guidelines for evaluating myocardial cell death

Abstract

Cell death is a fundamental process in cardiac pathologies. Recent studies have revealed multiple forms of cell death, and several of them have been demonstrated to underlie adverse cardiac remodeling and heart failure. With the expansion in the area of myocardial cell death and increasing concerns over rigor and reproducibility, it is important and timely to set a guideline for the best practices of evaluating myocardial cell death. There are six major forms of regulated cell death observed in cardiac pathologies, namely apoptosis, necroptosis, mitochondrial-mediated necrosis, pyroptosis, ferroptosis, and autophagic cell death. In this article, we describe the best methods to identify, measure, and evaluate these modes of myocardial cell death. In addition, we discuss the limitations of currently practiced myocardial cell death mechanisms.

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INTRODUCTION

A cell succumbs to death either immediately after exposure to multiple stimuli extremely unfavorable to it, or it dies gradually in a regulated manner. In the pathological heart, myocardial cell death is controlled by several receptors and signaling. Based on intrinsic and extrinsic stimuli, myocardial cells adopt different mechanisms of cell death. A myocardial cell may have either an intact or disrupted plasma membrane during cell death. Death by apoptosis and autophagy maintains cell membrane integrity. On the other hand, pyroptosis and necroptosis lead to disruption of the cell membrane, resulting in the release of cytoplasmic contents, including inflammatory cytokines. Release of these cytokines from the dying cells provides stress signals that alert the neighboring cells and mobilize protective mechanisms in the organism. However, massive myocardial cell death is observed in ischemia-reperfusion injury, which is detrimental to the heart. Out of the multiple reported forms of cell death (73), the six major forms observed in the heart are apoptosis, necroptosis, mitochondrial-mediated necrosis, pyroptosis, ferroptosis, and autophagic cell death. We will describe the best practices to measure these six forms of myocardial cell death.

Programmed cell death is an essential process in the developmental stage and is required for the formation of different structures, such as separation of the digits in hands and legs. In the adult stage, cell death is an important process to prevent infection against viruses and microbes, because proliferation of viruses and microbes requires a host cell, and thus host cell death would prevent their replication (20). However, cell death in key organs such as the heart is detrimental, because myocardial cells are terminally differentiated, have limited capacity for division, and perform important functions. Death of myocardial cells such as cardiomyocytes compromises contractility of the heart muscle, which in turn instigates adverse remodeling that ultimately leads to cardiac dysfunction and subsequent heart failure. Thus, cell death that results in loss of myocardial cells is a critical step in cardiac pathology, and targeting cell death mechanisms is a promising therapeutic option to mitigate and/or revert cardiomyopathy.

The research area of cell demise has advanced rapidly in recent years and multiple forms of cell death mechanisms have been identified (73). However, there is a lack of standardization regarding the evaluation of myocardial cell death. In the heart, cell death is a critical step to initiate pathological remodeling. With the increased emphasis on the rigor and reproducibility and the importance of cell death in cardiac pathophysiology, it is necessary to set guidelines for the best practices to measure myocardial cell death. This guidelines article describes different methods to measure apoptosis, pyroptosis, necroptosis, mitochondrial-mediated necrosis, ferroptosis, and autophagic cell death in the heart, limitations of some of the methods, and recommendations for the best practices to assess cell death mechanisms in the heart.

APOPTOSIS

Apoptosis is the most studied form of cell death in the heart. Although low levels of myocyte apoptosis (80–250 myocytes/10⁵ nuclei) are observed in the failing human heart, the persistent dropout of myocytes is a causal mechanism of heart failure (270). This programmed cell death mechanism plays a crucial role in cardiovascular embryogenesis and cardiac pathology (109, 115, 257). Apoptosis is characterized by cell shrinkage, chromatin condensation, and systematic DNA cleavage (56) and may equally afflict cardiomyocytes, interstitial cells, and endothelial cells. This slow apoptotic death of cardiomyocytes in adult life is of particular importance, as it may contribute to the pathogenesis of numerous myocardial diseases (such as ischemia, inflammation, infection, and immunologically mediated insults), where it may precede occurrence of necrosis/necroptosis. Two central pathways mediate apoptosis, an extrinsic pathway involving cell surface death receptors and an intrinsic pathway initiated by mitochondria and the endoplasmic reticulum (Fig. 1) (41, 109). BH3-only proteins of the BCL2-family are initiators of the intrinsic pathway of apoptosis that exert pro-apoptotic functions through Bak/Bak (80, 83, 157, 269). Due to the fact that cardiomyocytes have little potential for division, prevention of cell death is essential in the treatment of cardiovascular disease, such as myocardial infarction and heart failure (109). Both the expansion of our knowledge on the role of apoptosis in the complex cardiovascular environment and the evaluation of the efficacy on new therapeutic interventions in treating cardiovascular disease depend strongly on proper measurements and analyses of myocardial cell death (257). Because myocardial apoptosis and necrosis may constitute a spectrum, attempts to retard any one of the processes may reduce the extent of cell death. Other forms of cell death, such as autophagy and necroptosis, may cross-talk within the spectrum and further justify the necessity of intervention.

Fig. 1.

Apoptosis pathway. There are 2 major pathways that lead to apoptosis: extrinsic and intrinsic. The extrinsic pathway is triggered when ligands bind to the death receptors (such as tumor necrosis factor receptor I and Fas) on the surface of the cell. The binding induces conformational changes of the receptors that, with the help of the adaptor protein Fas-associated protein with death domain (FADD), activate pro-caspase-8 to caspase-8. The activated caspase-8 then activates pro-caspases-3,-6, and -7, the activation of which eventually leads to apoptosis. The intrinsic pathway is mitochondria dependent and happens in response to insults such as DNA damage, oxidative stress, and high calcium concentration. Activation of pro-apoptotic proteins (such as BAX) neutralizes the effects of the antiapoptotic proteins of the Bcl-2 family and leads to the release of apoptogenic factor cytochrome c from the mitochondria. Cytochrome c binds to APAF1 (apoptosis protease activating factor-1), and together they bind to and activate pro-caspase-9. The complex of cytochrome c, APAF1, and activated caspase-9 forms apoptosome, which also activates pro-caspases-3, -6, and -7, and leads to apoptosis. The cross-talk between the extrinsic and intrinsic pathways is through BH3-interacting domain (BID). When BID is cleaved by activated caspase-8, it is activated to the truncated form of BID (tBID) and translocates to mitochondria, promoting cytochrome c release. See Refs. 91 and 210.

Biochemical Tests

Several common biochemical methods can be used to detect apoptosis (Table 1). Terminal transferase dUTP nick-end labeling (TUNEL) staining and DNA laddering assay can detect the presence of DNA fragmentation (108). Annexin V staining can label phosphatidylserine residues that become exposed to outer cell surface during apoptosis (108). Caspase isoform activation is detected by showing caspase cleavage at specific sites that generate processed (active) caspase fragments (108).

Table 1.

Evaluation of myocardial apoptosis

Method	Detection Target	Apoptotic Cell Presentation	Appropriate Studies	References
Biochemical
TUNEL	DNA fragmentation	TUNEL positive	In vitro	236
DNA laddering	DNA fragmentation	Appearance of nucleosomal DNA ladder in agarose	In vitro	92
Light and electron microscopy	Morphological changes	Rounded-up phenotype	In vitro	1, 246
Flow cytometry/confocal imaging using annexin V	Exteriorization of phospholipids (PE/PS)	Annexin V positive	In vitro	184
Activity assay/Western blot/immunohistochemistry for target proteins	Protease activation/mitochondrial dysfunction	Caspase-3/7 activity assay: increased fluorescence, Western blotting, and immunohistochemistry; detection of activated proteins involved in the signaling pathway, e.g., caspase-3/6/7/8/9	In vitro	196, 217
Bioimaging
Imaging using annexin V probes labeled with 99mTc	Exteriorization of phospholipids (PE/PS)	Annexin V positive	In vivo	113, 156
Cardiovascular magnetic resonance (CMR) imaging	Cytokinetics	Myocardial hypokinesis	In vivo	8, 172, 237

CMR, cardiovascular magnetic resonance; PE, phosphatidylethanolamine; PS, polyunsaturated ω-6 fatty acids TUNEL, terminal transferase dUTP nick end labeling.

TUNEL staining utilizes the activity of the terminal deoxynucleotidyl transferase enzyme to label the 3′ ends of DNA strand breaks, which are then identified in individual nuclei by microscopy (68). However, the specificity of TUNEL assay has been challenged, as TUNEL positive cells could be apoptotic, necrotic, or normal cells undergoing DNA repair (195). The same issue is shared by the DNA laddering assay (37). Therefore, it is recommended that when detecting the presence of apoptosis in the heart, one should perform TUNEL assays using triple staining- for the nuclei, TUNEL to detect DNA fragmentation in the apoptotic nuclei, and cardiomyocyte-specific markers (271) in the suspected ischemic, infarcted, and nonischemic areas. To further confirm the presence of apoptosis, it is also recommended that one or more additional indices of apoptosis [such as caspase activity, poly ADP ribose polymerase (PARP) cleavage, apoptosis inducing factor and cytochrome c translocation from the mitochondria] be measured (109).

Caspases are a group of cysteine proteases that play a crucial role in initiating and executing apoptosis. Initiator caspases, such as caspase-8 and -9, act upstream to initiate and regulate apoptosis, whereas the effector caspases, such as caspase-3, -6, and -7, act downstream of both the extrinsic and intrinsic pathways to carry out the final biochemical changes in apoptosis (109). The activity of specific caspases can be measured by Western blotting, immunohistochemical techniques, or caspase activity assays using specific fluoremetric substrate or antibodies against the cleaved forms of their target proteins, such as PARP (140, 219, 276).

Phospholipids, such as phosphatidylserine (PS) and phosphatidylethanolamine (PE), which normally reside on the inner leaflet, are exteriorized to the outer layer of the plasma membrane during apoptosis (257). Annexin V is a positively charged protein that binds to negatively charged PS on the cell surface of apoptotic cells (257). When annexin V is fluorescently conjugated, the translocation of phospholipids can be detected by flow cytometry (256). When used together with the membrane-impermeant dyes, apoptotic cells, displayed with only annexin V staining, can be distinguished from necrotic cells that display annexin V concurrently with vital dye uptake, because the externalization of PS is an early event of apoptosis that occurs with intact plasma membrane (68). Similar to annexin V, antibiotic duramycin is able to selectively target PE and can also be used for the detection of apoptosis (169, 229, 291). Therefore, proper controls and gating strategies need to be applied in this detection method. However, it should be noted that PS exposure before membrane compromise is also observed in oncotic cells (137). The identification of morphological changes using histological stains or fluorescent dyes and visualization under light or electron microscopy is very useful in the characterization of cell death (68).

Imaging In Vivo

Visualization of apoptosis through noninvasive imaging could be useful for understanding a disease, monitoring its course, prognostication, defining targeted therapy, and measuring the efficacy of therapeutic intervention (229). A number of targets, including cell surface markers, membrane permeability, intracellular apoptotic machinery activation (e.g., caspase), and mitochondrial imaging, can be investigated for noninvasive imaging of apoptotic cell death (229). Two kinds of noninvasive strategies have been employed for imaging of apoptosis: imaging of apoptosis-specific exteriorized targets and targeting of intracellular processes specific for apoptosis (Table 1).

Based on the affinity between annexin V and PS, or duramycin and PE, annexin V and duramycin tracers radiolabeled with ^99mTc can be used for imaging apoptotic cells in many cardiovascular disorders (229). The main disadvantage of the annexin V probe is the significant off-target radiation, leading to the slow adoption of annexin V into clinical practice (229). Radiolabeled duramycin may obviate this shortcoming of the radionuclide imaging. Magnetic resonance imaging has also been employed for annexin V targeting through late gadolinium enhancement (LGE) for T1 relaxation enhancement or iron oxide nanoparticles for T2 relaxation enhancement (229). Currently, nanoparticles are being developed to image apoptosis without radiation and enable better topographic localization by using an apoptosis-sensing ligand linked with an imaging molecule (229). One of the strengths of this approach is the ability to localize apoptosis, determine its extent, and correlate it with morphology and functional consequences (229). However, because nanoparticles can be taken up by macrophages, such techniques may be limited to cardiac disease without significant inflammation or macrophage infiltration (229).

Clinical apoptosis imaging in acute myocardial infarction or transplant rejection has mainly employed ^99mTc annexin V with single photon emission computed tomography (SPECT) (229). Although the imaging of nonexteriorized targets has not been attempted in cardiovascular diseases, data from the cancer imaging field suggest the possibility of developing radiolabeled substrates specific for executioner caspases; mitochondrial membrane potential imaging also seems feasible (229).

Recommendations to Evaluate Apoptosis

Taking together different characteristics that cardiac apoptosis possesses, we recommend that investigators should combine different biochemical methods in vitro and immunohistochemical methods (such as TUNEL or activated caspase-3 staining), or ultrastructural examination (such as immunogold staining for cytochrome c leakage from mitochondria in cardiomyocytes) in tissue specimens (Table 1). In addition, noninvasive imaging methods could offer great tools for in vivo detection of cardiac apoptosis and monitoring disease progression and therapeutic efficacy. Cleavage of caspase-8 should be measured for death receptor-mediated apoptosis, whereas cytochrome c release and caspase-9 cleavage should be measured to determine mitochondria-mediated apoptosis (Fig. 1).

In cell culture, apoptosis may often transition to necrosis over time with plasma membrane breakdown because there is no phagocytosis in cell culture. Thus, we recommend that apoptosis should be determined at early time points in cell culture or blocked by pan-caspase inhibitor such as zVAD-FMK.

NECROPTOSIS

The paradigm that cells with a ruptured plasma membrane die due to an accidental, chaotic, and unregulated cell death was discredited in 2005 with the recognition that tumor necrosis factor-α (TNFα), a cytokine known to trigger the canonical extrinsic apoptosis pathway, can cause cell loss manifesting with the necrotic morphotype (50). Regulated forms of necrosis include necroptosis, mitochondrial-mediated necrosis, pyroptosis, and ferroptosis. Necroptosis has been found to underlie pathomechanisms of inflammation (177, 208), malignancies (36, 102), and microbial and viral infections (102, 122, 197), as well as ischemic injury of the heart (3, 4, 159, 243, 286), brain (213, 279), retina (116), and kidneys (151). This form of necrotic cell death has also been identified in various types of human heart failure independently on the etiology (243) and has been shown to underlie, at least in part, some phenotypes of this cardiac damage (3, 4). In addition, several animal models of heart failure, such as drug/chemical-induced cardiomyopathy (286, 287) and after left anterior descending artery (LAD) ligation (77, 159, 194), have highlighted a role for necroptosis in adverse cardiac remodeling and worsening heart function. Although the precise mechanisms of necroptosis induction and execution under conditions of myocardial damage are still not fully known, inflammation and oxidative stress, both mediators of cardiac disease, have been associated with a pronecroptotic environment (159, 194). Very recently, it has been reported that NLRP3-associated inflammasome may play a role in both noninfarcted and infarcted areas of post-myocardial infarction, whereas a canonical necroptosis signaling has been detected in the latter one only (149). Thus, necroptosis exhibits not only an adaptive function upon failing of cellular response to stress, but also, it is involved in developmental safeguard programs and the innate immune response.

Necroptosis can be induced by several triggering molecules, mainly through the stimulation of death receptors [TNFR, FasR, TNF-related apoptosis-inducing ligand receptor (TRAIL-R)] by the TNF family cytokines, and Toll-like receptors (TLRs) (49, 50, 72, 181, 261). Likewise, type I interferons and certain pathogens as well genotoxic (67) and oxidative stresses (44, 59, 118) promote this type of regulated necrosis. Depending on the triggering stimulus, pathways that execute necroptotic cell loss are likely to be different, and the current understanding of its mechanisms is based largely on experiments with TNFR1 signaling, which follows the RIP1-RIP3-MLKL signaling cascade terminating with plasma membrane disruption and cell lysis. However, the death receptor and RIP1 requirement can be bypassed, and the core components of the necroptotic pathway are RIP3 and MLKL.

TNF-induced stimulation of TNFR1 leads to receptor trimerization and the recruitment of cytosolic adaptor proteins to form a membrane-bound complex known as complex I (175, 223). It is composed of TNFR-associated death domain (TRADD), TNFR-associated factors 2 (TRAF2) and 5 (TRAF5), receptor-interacting protein kinase 1 (RIP1), and cellular inhibitor of apoptosis 1/2 (cIAP1/2). Loss of cIAP1/2 or increased activity of its counterregulatory enzymes, such as cylindromatosis (CYLD) (181), causes deubiquitination of RIP1 and internalization of membrane-bound complex I, where it turns into a cell death-promoting complex by recruiting Fas-associated death domain (FADD) and caspase-8 (175, 223, 260). This complex, named complex II, determines cell mortality through either apoptosis (complex IIa) or necroptosis (complex IIb) (Fig. 2). In addition, the ripoptosome, a complex similar to complex II, is formed by genotoxic stress (loss of cIAP) or FasR stimulation and contains RIP1, FADD, cellular FLICE-like inhibitory protein (cFLIP) isoforms, and caspase-10 and caspase-8, can also act as a death-inducing complex (67, 193).

Fig. 2.

Necroptosis pathway. TNF receptors induce formation of complex I. If receptor-interacting protein kinase (RIP)1 remains ubiquitinated, the cell survives. Complex I forms complex II by deubiquitination of RIP1, and it is mediated by loss of cellular inhibitor of apoptosis 1/2 (cIAP1/2). Caspase 8 activation in complex II inhibits RIP3 and promotes apoptosis, whereas caspase inactivation promotes necroptosis by phosphorylating RIP1, which in turn phosphorylates RIP3. Phosphorylated RIP3 causes mixed-lineage kinase domain-like (MLKL) phosphorylation that triggers MLKL oligomerization and subsequently translocation to the cell membrane, where it makes pore and release damage-associated molecular patterns (DAMPs).

What particular cell death mode is promoted is controlled by the activity/presence of caspase-8. In fact, caspase-8 cleaves the kinase domain of RIP kinases, thereby making them inactive and inhibiting necroptosis signaling, but at the same time, it can promote apoptosis by the activation of apoptotic downstream signals (47, 66). In contrast, the insufficiently active caspase-8 or its genetic and pharmacological inhibition (106, 193) prevents RIP1 cleavage, thereby promoting its association with RIP3 to form an amyloid-like protein complex known as the necrosome (259). In addition, cFLIP isoforms, which are predominantly expressed as either long (cFLIPL) or short (cFLIPS) isoforms, serve as an intracellular regulator of caspase-8 activity. The former isoform blocks caspase-8 to a degree that is sufficient to cleave and thus inactivate RIP1 but is insufficient to promote the activation of procaspase-3 to caspase-3 (55, 67, 221, 239). On the other hand, cFLIPS completely blocks the proteolytic activity of caspase-8, therefore preventing RIP1 proteolysis and leaving it active for necroptosis signaling (67, 111) (Fig. 2).

The active RIP3 favors necroptosis by phosphorylation of other RIP1 molecules at Ser₂₂₇, which in turn phosphorylates the pseudokinase mixed-lineage kinase domain-like (MLKL) at Thr₃₅₇ and/or Ser³⁵⁸ (at Ser³⁵⁸ in mouse MLKL) (34, 240, 290). Phosphorylation of only one residue of MLKL is sufficient to cause a conformational change, which triggers the oligomerization of MLKL molecules and its translocation to membrane compartments (Fig. 2). Phosphorylated MLKL assembling into oligomers and their subsequent translocation and insertion into the plasma membrane have been suggested to be the key drivers of necroptosis. However, precise MLKL stoichiometry and mechanism of action are still only partially understood. Several partial events of MLKL being associated with necroptosis have been studied; nevertheless, it cannot be ruled out that they can occur concurrently. This includes the following: 1) MLKL forms homo-oligomers (tetramers and/or octamers) or hetero-oligomers with RIP3 (96, 243, 263, 268); 2) MLKL oligomerization and translocation to the plasma membrane occur due to a specific and nonredundant role of the heat shock protein 90-kDa α-family class A member-1 (99); 3) MLKL oligomerization induces phosphatidyl serine exposure, resulting in the phosphatidyl serine-exposing plasma membrane bubbles, which attract cell surface proteases of the a disintegrin and metalloprotease (ADAM) family inducing shedding of the diverse cell surface proteins, including receptors, adhesion molecules, growth factors, and cytokines (28); 4) MLKL oligomers bind to the plasma membrane due to electrostatic interactions between their conserved positively charged regions and negative headgroups of membrane phosphatidylinositol lipids and cardiolipin (214, 240, 263) [phosphatidylinositol transfer protein-α (PITPα) seems to facilitate this interaction (102)]; 5) MLKL oligomers in the plasma membrane directly form nonspecific pores (58) or modify sodium (27, 33), magnesium, and calcium influx through transient receptor potential cation channel, subfamily M member 7 (27); 6) upon the translocation to the plasma membrane, MLKL responds to RIP3-mediated phosphorylation by the activation of an oxidative burst produced by NADPH oxidase 1 (NOX1) (118, 120); 7) MLKL translocate to the nucleus (243) and nuclear p-MLKL associates wth p-RIP3 to form lower-order oligomers that further exported from the nucleus to contribute to the formation of cytoplasmic p-RIP3-p-MLKL higher order oligomers (268); and 8) MLKL induces NLRP3 inflammasome formation and caspase-1 activation, resulting in IL-1β cleavage before cell lysis, thereby suggesting that inflammation associated with necroptosis can occur independently of the release of proinflammatory cellular contents due to plasma membrane rupture (38).

As indicated above, necroptosis execution is consistently associated with plasma membrane mechanisms; nevertheless, mitochondria participation in this process has also been reported. RIP3-mediated phosphorylation of phosphatase phosphoglycerate mutase (PGAM-5) inducing the dephosphorylation of dynamin-related protein 1 (DRP1) with resultant mitochondrial fission and mitochondrial fragmentation has been suggested to mediate necroptosis (266). Two independent laboratories have indicated that this RIP3-PGAM5-Drp1 axis can promote necroptosis in response to ischemia-reperfusion (94, 228). On the other hand, Pgam5^−/− mouse embryonic fibroblasts have been reported to respond normally to multiple inducers of necroptosis, thereby suggesting a dispensable role of PGAM5 in necroptosis signaling, at least in some cell types (182). The significance of mitochondria in necroptosis remains controversial because nearly complete depletion of the mitochondria did not influence necroptotic cell loss (247).

From the aforementioned discussion, it is evident that TNFR-induced necroptosis requires both RIP1 and RIP3 kinase activities. However, the formation of a pronecroptotic protein complex, which is independent of RIP1 and occurs through the association of RIP3 with other RIP homotypic interaction motif-containing proteins, such as DAI (DNA-dependent activator of IFN regulatory factor), ICP6 (infected-cell protein 6), and TRIF (TIR-domain-containing adapter-inducing interferon-β), due to M45/vIRA mutant cytomegalovirus (MCMV) (255), human herpes simplex virus type 1 infection (HV-1) (264), and stimulation of TLR (89), has also been reported. Thus, RIP3, unlike RIP1, seems to be a central signal-transducing element in necroptosis signaling. It receives pronecroptotic upstream signals and transduces the signal by activating MLKL to execute cell death.

Although the TNFR1-RIP1 link can result in cell death, it is necessary to mention that this signaling can also lead to cell survival under conditions when RIP1 ubiquitination at Lys³⁷⁷ is catalyzed by ubiquitin ligases cIAP1/2 (181, 260). In this scenario, RIP1 serves as a scaffold protein for the recruitment of transforming growth receptor-β-activated kinase 1 (TAK1) and nuclear factor-κB essential modulator (NEMO), which are known to induce survival genes, including nuclear factor-κB (NF-κB) (61, 144, 192). Of note is that unlike its cell death promoting function, this scaffolding function of RIP1 is independent of the kinase activity.

Identification of Necroptosis

An integrated approach involving the assessment of morphological, biochemical, and molecular features of necroptosis is essential for its identification and discrimination from the other necrosis-like cell death modes described in this review, such as the late phase of apoptosis and/or passive necrosis, autophagy, pyroptosis, ferroptosis, and mitochondrial-mediated necrosis (39, 57, 73). This indicates that the identification of specific necroptosis biomarkers should be combined with the traditional monitoring of these cell death modes and live/dead cell exclusion analysis. It is also worthy to note that a real-time morphological analysis could be considered because cells can readily time-dependently shift from apoptosis to necroptosis (and vice versa), or secondary necrosis can occur in later phases of apoptosis. In this section, the identification of specific necroptosis biomarkers is primarily discussed, whereas other methods, which provide additional supportive information, are listed briefly, as they have been nicely reviewed previously (43, 129, 258) and are also referred to in other sections of this review.

Identification of Specific Necroptosis Markers

A general way to assess necroptosis signaling is to measure mRNA expression and/or protein levels of RIP3, MLKL, and, alternatively, RIP1 (78, 151, 159, 242, 245, 286). However, the increased levels of these proteins, and mainly RIP1 and RIP3, do not necessarily indicate necroptosis. In support, as aforementioned, these proteins possess a variety of pleiotropic effects. For instance, RIP1 serves as an essential platform for induction of NF-κB activation and the production of proinflammatory cytokines (TNF, IL-1) as well as proteins producing some prosurvival effects (antioxidant enzyme manganese superoxide dismutase and the anti-apoptotic protein BCL2) (31, 61, 79, 93, 192). RIP3 is necessary for NLRP3 inflammasome activation with resultant production of IL-1β and IL-8 (110). In addition, both RIP1 and RIP3 are also associated with the intrinsic apoptosis pathway (162, 284). Total levels of MLKL do not provide clear evidence of active necroptosis because assembly and function of the NLRP3 inflammasome have also been shown to be regulated by MLKL (110), although another study has indicated independent action of NLRP3 in the inflammatory response (136). Based on these effects of RIP kinases and MLKL, additional measurements, such as their phosphorylation state, as well as necrosome formation and MLKL localization within the plasma membrane, are essential for the robust identification of necroptosis (Table 2).

Table 2.

Evaluation of necroptosis

	Methods to Detect the Particular Necroptotic Signaling	Key Notes	References
Specific molecular events strongly suggesting necroptosis being activated and/or executed
Activation of canonical RIP1-RIP3-MLKL axis through phosporylation	1) Detection: immunostaining; 2) inhibitors of RIP1 (necrostatins, GSK2982772), RIP3 (GSK`872, GSK`843), and MLKL inhibitors (necrosulfonamide); 3) knockdown approaches	Phosphorylated RIP1 and RIP3 suggest necroptosis activation; however, phosphorylation of either residue of MLKL robustly identifies necroptotic environment	42, 49–51, 64, 102, 105, 114, 146, 159, 165, 191, 203, 242, 244, 263
MLKL phosphorylation and translocation to the plasma membrane	1) Detection: immunostaining, subcellular fractionation; 2) inhibitors: MLKL inhibitors (necrosulfonamide)	Evidence of MLKL in the membrane fraction indicates its prior activation by the phosphorylation and execution of necroptosis	27, 33, 59, 240, 242, 243, 263, 268
MLKL oligomerization in the nucleus and plasma membrane	Immunofluorescence, nonreducing SDS-PAGE, native page gels	Phosphorylation of MLKL is a prerequisite for homo/hetero-oligomerization and translocation to the membrane/nucleus	96, 242, 263
Necrosome formation: RIP1-RIP3 interaction, RIP3-MLKL interaction	Immunofluorescence, coimmunoprecipitation	Necrosome identification indicates necroptosis activation; however, end-stage effector of the necroptosis (p-MLKL) should be also provided	34, 139, 194, 268, 286
Other more nonspecific characteristics of necroptosis	Methods to detect nonspecific features of necroptotic cell death
Loss of cellular integrity, plasma membrane rupture	LDH, HMGB1 release, staining with impermeant dyes, annexin V/PI staining
Altered cell viability	XTT, MTT assay, etc.

HMGB1, high-mobility group box 1 protein; LDH, lactate dehydrogenase; MLKL, mixed-lineage kinase domain-like; p-MLKL, phosphorylated mixed-lineage kinase domain-like; RIP1 and -3, receptor-interacting protein kinase 1 and -3, respectively; XTT, 2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide.

Specific antibodies recognizing posttranslational modification via phosphorylation of particular residues have been developed, and thus phosphorylation of hRIP3 at Ser²²⁷ (Thr²³¹/Ser²³² in mRIP3) and hMLKL at Thr³⁵⁷ and Ser³⁵⁸ (at Ser³⁵⁸ in mMLKL) can be assessed by immunoblotting (102, 243, 245). Unlike in humans and mice, there is no commercial antibody recognizing necroptosis residues of rat MLKL, and therefore, Phos-tag Western blotting can be eligible for this purpose. It is important to note that while phosphorylation of MLKL is a strong indicator of necroptotic cell death, it can be questionable in some conditions (59, 243, 268). Necrosulfonamide, an MLKL inhibitor, was shown to affect neither phosphorylation nor oligomerization of MLKL, but it was capable of preventing MLKL membrane translocation and, consequently, membrane rupture. However, necrosulfonamide only appears to inhibit human MLKL but not mouse MLKL (240, 263). Given this fact, other approaches for necroptosis identification involving the detection of necrosome formation, and MLKL oligomers in nuclear and membrane fraction, might be more conclusive.

Necrosome formation can be detected by immunofluorescence or by coimmunoprecipitation as RIP1-RIP3 interaction (34, 139, 268, 286), and RIP3-MLKL interaction is also a powerful option (194, 268). The higher-molecular-weight bands running around 160–300 kDa representing MLKL tetramers and octamers can be used to identify MLKL oligomerization on a nonreducing SDS-PAGE or native PAGE gel (96, 242, 263). As the so-far accepted terminal event of the necroptosis pathway, membrane translocation of MLKL can be identified by cellular fractionation with subsequent Western blotting or by immunohistochemical/immunocytochemical staining (Table 2) (27, 33, 242, 263).

Additional Supportive Methods For Necroptosis Monitoring

As a form of necrosis, the main morphological hallmarks of necroptosis, monitored by a transmission electron microscope, time-lapse video microscopy with epifluorescence, or a phase-contrast inverted microscope, include an increase in cell size due to swelling, prominent nuclear swelling, swelling of other cytoplasmic organelles, and loss of mitochondrial and endoplasmic reticulum mass followed by rapid loss of plasma membrane integrity (19). Notably, optimizing the light exposure and fluorescent probe should be an important consideration to minimize technique-induced cytotoxicity. Likewise, depending on the necroptosis triggering stimuli and a particular cell line and thereby the kinetics of its cell cycle, the time points at which real-time morphology analysis is intended to be performed, should be individually optimized.

The conventional biochemical cytotoxicity assays for necroptosis identification are based on the evaluation of the leakage of intracellular molecules through the ruptured plasma membrane, such as lactate dehydrogenase (LDH), damage-associated molecular patterns (DAMPs) such as high-mobility group box 1 protein (HMGB1), IL-1α, and IL-33, and mitochondrial DNA (19). However, the presence of these molecules in in vivo experiments can also indicate a late phase of apoptosis/secondary necrosis or suggest macrophage activation. The loss of cellular integrity can be also identified by impermeant dyes [trypan blue, propidium iodide (PI), and 4′,6-diamidino2-phenylindole (DAPI)] that differentially enter the cytosol and/or are bound to DNA in dying cells, although they do not cross through the plasma membrane of the healthy cells. Similar to late apoptotic/secondary necrotic, necroptotic cells are annexin V and PI positive. To distinguish late apoptotic/secondary necrosis from necroptosis, image-based analysis can identify some specific morphological features. For the necroptotic cells, a diffuse PI staining and an intense fluorescence signal are characteristic, whereas late apoptotic/secondary necrotic cells, which are shrunken in size, with condensed chromatin, exhibit spots of PI staining and annexin V-positive blebs.

Another potential approach to discriminate the early stage of necroptosis from primary/secondary necrosis could be the assessment of mitochondrial membrane potential with fluorescent probes (JC-1, Rhodamine 123) that incorporate into mitochondria and fluoresce brightly, depending on the inner transmembrane potential (206). In cells with ruptured plasma membrane, which dissipates the potential, a decrease in fluorescence can be detected. This method is unlikely to be effective in distinguishing the later phase of necroptosis from necrosis, as both are known to exhibit the same loss of the plasma membrane integrity. In addition, as apoptotic cells can also exhibit a loss of mitochondrial membrane potential and thereby a decrease in fluorescent signal, additional analyses must be performed to avoid misinterpretation. However, in this regard, it should be noted that role of mitochondria in necroptosis is unclear, and although some authors have reported necroptosis execution due to mitochondrial fission associated with RIP3-PGAM5-Drp1 link (94, 228) and/or mitochondrial permeability transition (MPT) pore opening (150, 235) mediated by RIP3-CaMKII axis (287), others have provided contradictory data (186, 216). Thus, the above-mentioned fluorescent staining and other methods assessing dissipation of mitochondrial membrane potential, which are more indicative for mitochondrial-mediated necrosis, can be used as a supportive method to investigate mitochondrial involvement in necroptotic signaling.

Importantly, inhibitor and knockdown approaches, which allow rapid dissection of divergent cell death pathways, should be considered to test the involvement of necroptosis. Several pharmacological inhibitors of RIP1, RIP3, and MLKL have been synthetized. RIP1 inhibitors include a group of small-molecule chemicals named necrostatins, of which necrostatin-1 and its more selective analog necrostatin-1s have been extensively used in preclinical studies (49, 50, 117, 139, 152, 242, 244). They inhibit the death domain of RIP1, whereas other necrostatins target other steps in the necroptosis pathway through different mechanisms (49–51). In contrast to these molecules, another potent RIP1 inhibitor, GSK2982772, which binds in an allosteric pocket of the RIP1 kinase domain, possesses an exquisite kinase specificity and is a clinical candidate for treatment of inflammatory diseases, psoriasis, rheumatoid arthritis, and ulcerative colitis (86). RIP3 kinase inhibitors such as GSK`872, GSK`843, HS-1371, dabrafenib, and ponatinib (42, 64, 105, 146, 203) and necrosulfonamide, an MLKL inhibitor, which, however, blocks only human MLKL by targeting Cys⁸⁶ (240, 263), have also been appropriated for studying conditions exhibiting necroptosis. Nevertheless, some limitations such as protein selectivity and species specificity must be considered when selecting them for studies. Thus, additional knockdown strategies can be conducted to confirm data from pharmacological experiments. Nowadays, MLKL-deficient (273) and RIP3-deficient animals (159) are available. In contrast, as indicated above, RIP1 deficiency may not be a good choice for necroptosis study. RIP1^−/− mice have been found to display neonatal lethality (114), whereas mice with kinase-dead knock-in mutation of RIP1 are viable (191), supporting the evidence that this kinase has important pleotropic effects and that RIP1 with the dead kinase domain is a prosurvival factor affecting the downstream molecules (31, 79, 93).

Recommendations to Evaluate Necroptosis

Although the development of several accurate techniques for cell death detection has advanced, the precise determination of the extent of cell death due to necroptosis remains challenging. In addition, experimental conditions and material undergoing analysis (in vivo, in situ, in vitro, and ex vivo experiments), as well as the fact that some cells rapidly change phenotype, should be carefully considered when choosing particular methods of analysis. Regardless, the identification of necroptotic cell death is based on a set of analyses, with a cardinal focus on the monitoring of RIP3-MLKL necroptotic signaling-induced plasma membrane disruption (Table 2). Detailed analysis of necroptosis-associated relocalization of these proteins within individual cellular compartments as well as the formation of intracellular necroptotic complexes is recommended alongside indices of activation of the key necroptotic molecules (Table 2). Likewise, it is highly recommended that analyses of cell/tissue viability be provided in addition to conventional, nonspecific methods assessing cell membrane rupture. Importantly, pharmacological approaches and/or genetic engineering tools, such as ablation or transgenesis of particular necroptotic proteins, are needed to elucidate the exact role of this mode of cell death in certain cardiac pathologies. The specific molecular events strongly suggesting necroptosis, and their detection methods are described in Table 2.

MITOCHONDRIAL-MEDIATED NECROSIS

Just as mitochondria are key mediators of the intrinsic apoptotic pathway, they can also play an essential role in the execution of necrotic cell death, especially in cardiac myocytes. The archetypal mitochondrial-dependent necrotic pathway is that driven by the MPT pore. The MPT pore is a channel that spans the inner mitochondrial membrane (IMM) (25, 133, 211). Although not sufficient in size to enable the passage of proteins across the IMM, when open, the MPT pore allows the free movement of molecules <1.5 kD in size. Since this includes protons, pore opening will dissipate the mitochondrial membrane potential (ΔΨ_m) and, therefore, inhibit a variety of mitochondrial functions, most critical of which is ATP synthesis (25, 133, 211). In addition, water can enter the mitochondrial matrix down its osmotic gradient, causing the mitochondria to swell. This will disrupt cristae structure, further impairing oxidative phosphorylation (25, 133, 211). If mitochondral swelling is too excessive, the outer mitochondrial membrane can also rupture. Combined, these sequelae of pore opening ultimately cause the necrotic demise of the cell (Fig. 3).

Fig. 3.

Mitochondrial permeability transition (MPT)-mediated necrosis. Because of calcium overload and/or extremely high oxidative stress, the mitochondrial membrane becomes permeable and results in dissipation of mitochondrial membrane potential, leading to cessation of ATP synthesis. Increased permeability also causes influx of cytoplasmic water molecules, which causes matrix swelling that ultimately results in mitochondrial membrane rupture and release of apoptotic factors. In the presence of ATP, it leads to apoptosis. However, depletion of ATP causes MPT necrosis.

The primary inducers of MPT are Ca²⁺ and oxidants. In both cases, these stimulants act on the matrix side of the MPT pore (Fig. 3) (25, 85, 133, 211). Ca²⁺ needs to be taken up across the IMM through the mitochondrial Ca²⁺ uniporter (MCU) until a concentration of Ca²⁺ that is sufficient to induce MPT pore opening is reached. Oxidants react with vicinal thiols exposed to the matrix (85). However, which protein(s) constitutes the specific Ca²⁺ and/or redox sensor has yet to be established (see below). The induction of pore opening by these agents is further facilitated by any reductions in adenine nucleotides, which suppress MPT pore opening (85). Reactive oxygen species and/or Ca²⁺ are frequently elevated in multiple cardiac pathologies, and MPT has been reported to contribute to the cardiac myocyte necrosis underlying ischemia-reperfusion injury (9, 13, 35, 82, 188), mitochondrial, dystrophic, and myocarditic cardiomyopathies (166, 176, 183), and anthracycline cardiotoxicity (14, 165, 190).

The model of the proteins that make up the MPT pore is in constant flux with the voltage-dependent anion channel (VDAC), adenine nucleotide translocase (ANT), and phosphate carrier (PiC), and the ATP synthase itself being proposed as the channel-forming protein (25, 133, 211). However, genetic studies have questioned the identity of these proteins as the components of the MPT channel (14, 84, 87, 88, 125). As such, the only pore component that has stood up to genetic testing is the matrix isomerase cyclophilin-D (CypD) (13, 16, 188, 222). However, it should be remembered that although CypD is a critical positive regulator of the MPT pore, it is not the channel-forming protein itself, and its ablation can have several MPT-independent effects on mitochondrial metabolism and function (62, 211). We will describe the most commonly used techniques for measuring MPT in the following sections, but we would point out that additional methods can be employed (2, 76, 294).

Identification of MPT and MPT-Dependent Necrosis In Vitro

Several methods have been developed and established as indices of MPT pore opening in both isolated mitochondria and intact cells (Table 3). The oldest method is the spectrophotometric measurement of Ca²⁺ or oxidant-induced swelling in mitochondria (13, 14, 35). Here, mitochondria are isolated from cultured myocytes or excised hearts, typically by differential centrifugation in sucrose-based medium. Energized mitochondria are then challenged with a large bolus of Ca²⁺ in the micromolar range and swelling measured over time as a decrease in absorbance at 520–540 nM. The advantages of this system are its simplicity and reproducibility. However, it requires a relatively large number of mitochondria, which can be problematic when using cultured myocytes or mouse hearts such that often only a single concentration of Ca²⁺ can be tested when ideally a dose curve should be constructed.

Table 3.

Evaluation of mitochondrial permeability transition-dependent necrosis

Characteristics of MPT	Differences Between MPT-Dependent and Other Forms Of Cell Death	Methods to Detect Characteristics	References
Loss of ΔΨm	Opening of the MPT pore will dissipate the ΔΨm; however, other interventions can also induce loss of ΔΨm and therefore appropriate controls must be used.	1) Detection: assessment of using potentiometric fluorophores such as TMRE or JC-1 both in cultured myocytes and epicardially in isolate perfused hearts; 2) inhibitors: cyclosporine-A, bongkrekic acid, CypD depletion, positive controls: Ca2+, H2O2, ionomycin	9, 13, 35, 48, 82, 165, 166, 168, 176, 180, 183, 188, 190
Increased IMM permeability	Rapid increase in IMM permeability ≤1.5 kD, leading to swelling, separate from the increase in OMM permeability induced during intrinsic apoptosis.	1) Detection: measurement of swelling and CRC in isolated cardiac mitochondria and CRC and calcein CoCl2 in intact myocytes. Use of 2-DOG uptake or loss of mitochondrial NAD+ in vivo; 2) inhibitors: cyclosporine-A, bongkrekic acid, CypD depletion, positive controls: Ca2+, H2O2, ionomycin	9, 13, 35, 54, 81, 82, 165, 166, 176, 183, 188, 190, 207
Inhibition of mitochondrial ATP synthesis	By dissipating the ΔΨm, opening of the MPT pore inhibits ATP synthesis, leading to a depletion in adenine nucleotides.	1) Detection: measurement of mitochondrial ATP levels in vitro and in vivo using commercially available kits, measurement of ATP synthesis rates in isolated mitochondria; 2) inhibitors: cyclosporine-A, bongkrekic acid, CypD depletion, positive controls: Ca2+, H2O2, ionomycin	48, 53, 84, 227

CRC, Ca²⁺ retention capacity; CypD, cyclophilin-D; IMM, inner mitochondrial membrane; OMM, outer mitochondrial membrane; MPT, mitochondrial permeability transition; TMRE, tetramethylrhodamine ethyl ester.

An alternative method for measuring MPT is the Ca²⁺ retention capacity (CRC) assay (13, 87, 88, 188). This is an improvement on the swelling assay, as it can be done in intact (albeit permeabilized) cells, and the amount of starting material is considerably less, thereby making it more amenable to high throughput screens. In this assay, energized isolated mitochondria or permeabilized myocytes are incubated with the cell-impermeable Ca²⁺-sensitive dye Calcium Green-5N, which measures extramitochondrial Ca²⁺. Small boluses of Ca²⁺ are then added to the mix, which results in a spike in fluorescence as the ion binds the dye. The fluorescence then decays as the Ca²⁺ is taken up into the mitochondria via the MCU. This is repeated until the threshold required to open the MPT pore is reached. Opening of the pore allows the accumulated matrix Ca²⁺ to rapidly escape, resulting in a quick, large-amplitude increase in fluorescence. Because only small amounts of Ca²⁺ are added each time, a more precise calculation of the MPT pore’s Ca²⁺ sensitivity is obtained compared to that using the swelling assay. Moreover, depending on the fidelity of the detection system, it can also provide additional information regarding mitochondrial Ca²⁺ uptake kinetics.

Because both mitochondrial volume and CRC can be altered independently of direct changes in MPT, a role for the MPT pore should be verified using pharmacological inhibitors of the pore such as cyclosporine-A or bongkrekic acid, or RNAi-mediated depletion of CypD (Table 3). Pharmacological and/or genetic inhibition of MPT will attenuate swelling and greatly increase the amount of Ca²⁺ required to induce MPT in the CRC assay. However, it needs to be noted that the swelling and CRC assays measure only the sensitivity of the pore to Ca²⁺ and are typically done under “basal” conditions. Thus, although they can still provide useful information, they do not provide insight as to how MPT is occurring in response to a variety of noxious stimuli in intact cardiac myocytes.

In this regard, two commonly used methods for the assessment of MPT in intact myocytes are the measurement of ΔΨ_m and the calcein/Co²⁺ assay. ΔΨ_m can be measured using potentiometric dyes such as JC-1 or the rhodamine-based fluorophores such as tetramethylrhodamine ethyl ester (TMRE) (48). These dyes accumulate and fluoresce in the mitochondrial matrix due to the high negative charge. Opening of the MPT pore dissipates the ΔΨ_m, which reduces mitochondrial fluorescence. Typically, loading of the cell is done at the end of the stimulus incubation period, and microscopy, fluorimetry, or FACS can be used to measure differences between control and treated cells. Moreover, this assay is also amenable to measurement of ΔΨ_m over time once an intervention has been added. The major caveat is that although MPT pore opening will always reduce ΔΨ_m, ΔΨ_m can be reduced independently of MPT. Therefore, pharmacological and/or genetic inhibition of MPT, as described above, is required to ascribe any changes in ΔΨ_m to modulation of the MPT pore. H₂O₂ (oxidative stress) and ionomycin (Ca²⁺ overload), which induce MPT-dependent necrosis (13, 188), are also useful as positive controls.

To circumvent MPT-independent loss of mitochondrial membrane potential, Petronilli et al. (207) developed the calcein/CoCl₂ assay. In this method, cells are loaded with calcein-AM dye, which distributes between the cytosol and mitochondria. However, when co-loaded with CoCl₂, which cannot penetrate the mitochondria, the cytosolic signal is quenched so that only the mitochondria fluoresce. Opening of the MPT pore allows the equilibration of the mitochondrial calcein with the cytosolic CoCl₂ such that mitochondrial fluorescence is then reduced. Therefore, a reduction in calcein intensity is an index of pore opening. Like the ΔΨ_m-based assay, this can be analyzed by microscopy, fluorimetry, or FACS in real time. Again, established MPT inhibitors and activators should ideally be used as negative and positive controls, respectively. Unfortunately, the ΔΨ_m and calcein methods cannot be done simultaneously as TMRE quenches the calcein signal (207), and there is wavelength overlap between calcein and JC-1.

Although the assays described above can provide insight as to whether a compound/intervention sensitizes the MPT pore or induces its opening, they do not tell whether such changes in MPT are the causative driver of necrotic death in this context. There are no specific measures of MPT-driven necrosis. As described for the other pathways, general necrosis end point measurements such as vital dye exclusion or LDH and HMGB1 release are typically used in conjunction with genetic or pharmacological MPT inhibition.

Identification of MPT and MPT-Dependent Necrosis In Vivo

Although multiple indices have been established for assessing MPT in isolated mitochondria and cultured myocytes, reliable assays of MPT in vivo are lacking. Typically, mitochondria are isolated from control and affected hearts, and then MPT is assessed by swelling or CRC assays. But this does not indicate what is happening in situ. Epicardial fluorimetry of TMRE or JC-1 has been utilized in isolated perfused hearts as a real-time measure of MPT (168, 180) with the caveat of measuring surface mitochondria only in a limited area. Griffiths and Halestrap (81) utilized an isotope approach whereby rats were injected with ³H-2-deoxyglucose (DOG) and then subjected to cardiac ischemia/reperfusion injury. DOG is taken up into the myocyte but cannot enter mitochondria under normal circumstances. However, opening of MPT pore allows the uptake of the DOG into mitochondria. The authors isolated cardiac mitochondria following injury and found that mitochondrial ³H levels were increased upon reperfusion, but not during ischemia alone. However, the fact that DOG is an inhibitor of glycolysis and results in a tritiated animal greatly hampers the use of this model. Di Lisa et al. (54) took a reverse approach and measured the reduction in mitochondrial NAD⁺ due to its release through the MPT pore and subsequent hydrolysis in the cytosol after ischemia-reperfusion. While avoiding radioactive isotopes, cytosolic NAD⁺ can change independently of the release of mitochondrial pool. Moreover, both assays are really only amenable to interventions that induce massive MPT-driven necrosis, and pathologies where a more subtle loss of myocytes due to MPT occurs may not be detectable.

Thus, as with the cells, determination of MPT-dependent necrosis in vivo is reliant on manipulation of the MPT pore. As such chemical inhibitors of CypD such as cyclosporine-A, NIM-811, and sanglifehrin-A can be (and have been) used. Nevertheless, such drugs inhibit other cyclophilins, and cyclosporine-A inhibits calcineurin, which plays its own role in cardiac myocyte death and pathology (204). Consequently, any long-term use of these compounds as pore inhibitors can be problematic. Although the adenine nucleotide translocase (ANT) inhibitor bongkrekic acid inhibits the pore independently of CypD, its expense precludes its use in vivo, and long-term inhibition of ANT would be toxic. The still uncertain composition of the MPT pore also makes it hard to genetically inhibit in vivo. The primary approach has been to use mice lacking CypD, and useful data has been obtained with these animals. However, constitutive loss of CypD can also alter mitochondrial biology in ways that are likely MPT independent (62, 211), and care should be taken in data interpretation when using these mice. Fortunately, mice with a floxed CypD allele are available (134), and acute deletion of cardiac myocyte CypD using an inducible Cre may prove more informative as compensatory or non-pore-related changes may be circumvented, or at least reduced.

Recommendations for Evaluating Mitochondrial Permeability Transition-Dependent Necrosis

We recommend that at least two indices of MPT be measured in response to a drug or intervention (such as the swelling, CRC, ΔΨ_m, and/or calcein/CoCl₂ techniques described above), with or without MPT inhibition, in in vitro experiments. Measurement of ATP levels can also be used as a surrogate given that MPT inhibits ATP synthesis (53, 227). Similarly, pharmacological and/or genetic inhibition of the pore can provide insight for establishing MPT as a causative agent in increased cell necrosis. Given the intricacies and limitations of measuring MPT in vivo, we also recommend that two indices of MPT such as swelling or CRC be assessed ex vivo. As with in vitro studies, the role of MPT in myocyte injury and pathology would need to be substantiated by re-assessing the phenotype in the presence of MPT inhibition, either pharmacologically or using CypD-deficient mice. The characteristic features of MPT and detection methods of MPT-mediated necrosis are described in Table 3.

PYROPTOSIS

Pyroptosis is a lytic type of regulated cell death that is triggered by the formation of plasma membrane pores by members of the gasdermin (GSDM) family of proteins as a consequence of activation of inflammatory caspases, mainly caspase-1, but also murine caspase-11 and its human homologs caspase-4 and caspase-5 (70, 73, 230). Caspase-1 is activated by an intracellular multiprotein complex called the inflammasome that is formed in response to pathogen-associated molecular patterns (PAMPs) derived during infection and DAMPs derived during cellular stress and damage. Although several types of the inflammasome have been reported, the NLRP3 inflammasome is the best characterized and involves sterile inflammation in the pathophysiology of cardiovascular diseases, including myocardial infarction (248, 249, 251). The NLRP3 inflammasome is composed of NLRP3 [Nod-like receptor (NLR) family pyrin domain containing 3], the adaptor protein ASC (apoptosis-speck like protein containing a caspase recruiting domain, also known as PYCARD), and the cysteine protease caspase-1, and its formation induces caspase-1 activation (Fig. 4). Recent studies identified a pyroptosis executioner GSDMD that is a substrate of both caspase-1 and caspase-11-4/5 (127, 230). These caspases proteolytically cleave GSDMD into an NH₂-terminal domain (GSDMD-N) and COOH-terminal domain (GSDMD-C). Cleaved GSDMD-N binds to phosphoinositides in the plasma membrane (or cardiolipin of a bacterial membrane) and forms oligomeric death-inducing pores with an inner diameter of 10–20 nm. Activated caspase-1 can also process the pro-forms of inflammatory cytokines IL-1β and IL-18 into their mature forms (Fig. 4). Although IL-1β and IL-18 lack a secretion signal, the GSDMD-forming pore allows passage of these cytokines (∼5 nm) through the cell membrane, which probably explains the caspase-1-dependent secretory mechanism of IL-1β and IL-18.

Fig. 4.

The canonical NACHT, LRR, and PYD domains-containing protein 3 (NLRP3) inflammasome pathway of pyroptosis. A number of substances generated during ischemia-reperfusion (termed signal 2) trigger the assembly of the NLRP3 inflammasome, which results in activation of the cysteine-aspartic protease caspase-1. Simultaneous activation of Toll-like receptors (signal 1) increases the expression of pro-interleukins and NLRP3 via NF-κB. Caspase-1 activates pro-interleukins and releases the NH₂-terminal domain of gasdermin D, which makes large pores in the plasma membrane. Those pores allow the proinflammatory interleukins to be secreted, and the loss of membrane integrity also kills the cell (pyroptosis). Caspase 1 is also reported to attack many other proteins in the cell, including some vital enzymes like GAPDH, which could also contribute to its toxicity.

In contrast to caspase-1, caspase-11-4/5 directly binds cytosolic lipopolysaccharide (LPS) via its CARD (caspase activation and recruitment domain) domain, leading to the cleavage of GSDMD and subsequent pyroptosis. Although caspase-11-4/5 cannot process IL-1β and IL-18 directly, they induce GSDMD-mediated potassium efflux, which is sufficient to cause NLRP3 inflammasome activation and subsequent release of IL-1β and IL-18; this process is known as the noncanonical inflammasome pathway. Furthermore, recent studies indicated that pyroptosis can also be driven by caspase-3-induced cleavage of GSDME/DFNA5 (gasdermin E is also known as deafness associated tumor suppressor) (265). Because the pore-forming ability of GSDMD is considered to be conserved throughout GSDM family proteins (65), it is possible that other GSDM proteins are involved in the process of pyroptosis. The role of GSDMD, GSDME, and the noncanonical inflammasome pathway in myocardial ischemia-reperfusion injury is still poorly characterized.

Pyroptosis is characterized by loss of plasma membrane integrity, leading to cell swelling and rupture of the cell membrane, nuclear condensation, and release of inflammatory cytokines and intracellular DAMPs (Fig. 4). Some of these features are also observed in necrosis, apoptosis, and necroptosis. We herein propose several experimental models that are likely to involve pyroptosis and could be used to collect biochemical and pharmacological evidence that could either support or refute the presence and contributing role of pyroptosis on cardiac injury, with an emphasis on using multiple techniques (with assessment of expression and function) and models (both ex vivo and in vivo).

Canonical Mechanism of Caspase 1 Activation Within the Inflammasome

The inflammasome is a macromolecular structure that acts as a sensor for extracellular and intracellular injury, and as a mechanism for the initiation and amplification of the inflammatory response (90, 215). The inflammasome is formed by the aggregation of three main molecular entities. NLR or absent in melanoma 2-like receptors (ALR) constitute intracellular sensors of damage or danger signals that trigger the formation of the inflammasome by recruiting the adaptor protein ASC. The ASC scaffold allows for recruitment of caspase-1 and leads to oligomerization of pro-caspase-1 molecules and autocleavage into active caspase-1. The most widely studied sensor for the inflammasome is the NLRP3, also known as NALP3 or cryopyrin. A large number of unrelated stimuli activates NLRP3, including bacterial toxins, K⁺ efflux, and lysosomal failure (90).

An Alternative Mechanism of Caspase-1 Activation Independent of NLRP3 Inflammasome

The components of the NLRP3 inflammasome are not constitutively expressed in the heart. Formation of the NLRP3 inflammasome requires priming (e.g., during ischemia), followed by triggering by DAMPs released during ischemia (251–253). However, studies on LDH release with and without the use of a caspase-1/4 inhibitor, VRT-043198 (an active metabolite of VX-765), highlight a role for caspase-1 and -4 in membrane failure and cell death (Table 4). A benefit is seen with the introduction of a caspase-1/4 inhibitor given at the onset of reperfusion and assessment of infarct size measured after 2-h reperfusion (10). A smaller infarct size is seen after 24 h of reperfusion with a NLRP3 inhibitor, but the same benefit is not seen with an NLRP3 inhibitor given at reperfusion and infarct size is assessed only 3 h later (251). Therefore, the data suggest an NLRP3-independent yet rapid caspase-1/4 dependent membrane failure in this context.

Table 4.

Evaluation of pyroptosis

Characteristics of Pyroptosis	Differences Between Pyroptosis and Other Forms of Cell Death	Methods to Detect Characteristic Features	References
Human caspase-1/4/5 (or murine caspase-1/11)-mediated cell death by membrane failure that allows intracellular contents to be lost; pyroptosis is often associated with DNA fragmentation.	Pyroptosis is believed to result from caspase activation of GSDMD, which then forms large pores in the plasma membrane; proof of ischemia-triggered pyroptosis is complicated by the fact that loss of energetics alone during ischemia can cause membrane failure by necrosis, and inflammatory components may contribute to that energy deficit.	Cell death should be associated with caspase-1 activation and GSDMD cleavage; current evidence is that caspase-1 or inflammasome inhibition reduces ischemia-induced cell death; however, proof of a pyroptosis mechanism would also require a demonstration that the cell death was GSDMD dependent. Models to detect loss of membrane integrity include: • Isolated cells: trypan blue or propidium iodide exclusion assay. • Cells and heart tissue: release of cytosolic enzymes (CK-MB, troponin I or T, or LDH) • Whole heart: infarct size by pathology or by enzyme release	10, 112, 163, 167

GSDMD, gasdermin D; LDH, lactate dehydrogenase.

One possible explanation is that calpain activation upon reperfusion is releasing pro- caspase-1 sequestered in the actin cytoskeleton of the heart cells (288). Activation of calpain in macrophages by elevated Ca²⁺ digests a protein called flightless-1, which is a caspase-1 (and caspase-11)-binding protein and anchor to the cytoskeleton. Caspase-1 can be self-activated, but flightless-1 and other inhibitors function as caspase-1 pseudosubstrates preventing autocleavage. Calpain activation reduces flightless-1 levels, causing rapid self-activation of caspase-1 (145, 288). Whether calpain-dependent activation of caspase-1 also occurs in reperfused cardiac tissue is currently unknown.

Measuring Caspase-1 Activation in the Heart Tissue

Basal caspase-1 activity in the healthy heart is rather low; therefore, small changes in caspase-1 activation may be easily detected. Caspase-1 activation can be measured by detection of the appearance of the low-molecular weight active caspase-1 in a Western blot and expressed as its ratio to its heavier pro-caspase-1 precursor (22, 23). Caspase-1 activity can also be measured by the appearance of low-molecular weight fragments of endogenous caspase-1 substrates like IL-1, IL-18, GSDMD (Fig. 4), and others measured in the heart tissue or in the plasma (52), as well as by the cleavage of a fluorogenic substrate in which the heart tissue is incubated (22, 163, 164, 173). An intervention that reduces caspase-1 activity measured with one of the methods described and is associated with a reduction in infarct size may again be interpreted as evidence of possible pyroptosis-mediated cell death (Table 4). One limitation of this examination of biopsies in in vivo hearts is that it is not able to distinguish activation of caspase-1 in cardiomyocytes versus other resident or infiltrating cells.

Measuring Inflammasome Formation and Aggregation in the Heart Tissue

The components of the inflammasome are minimally expressed in the healthy heart, and the expression increases during insults like ischemia and infarction. The active inflammasome appears as a macromolecular structure or specks using fluorescence microscopy after staining for the scaffold protein ASC or for the mostly commonly involved effector of caspase-1, NLRP3 (18, 173, 254). Staining for one or more of these structures allows for the measurement of inflammasomal specks in the heart tissue, and using cell-specific costaining it allows for the determination of formation in cardiomyocytes versus other resident or nonresident cells. While the formation of the inflammasome specks can hypothetically occur without leading to activation of caspase-1, i.e., in presence of a powerful caspase-1 inhibitor, it is accepted that appearance of inflammasome specks can serve as a surrogate of caspase-1 activation and possible pyroptosis (173, 254). One of the limitations of this method is that it does not consider an inflammasome-independent mechanism of caspase-1 activation such as calpain activity, as discussed above. In rodent models of myocardial infarction, inflammasome specks have been detected in cardiomyocytes, cardiac fibroblasts, endothelial cells, and infiltrated leukocytes (173). Furthermore, activation of the NLRP3 inflammasome induces IL-1β release in cardiac fibroblasts and endothelial cells through pyroptosis, whereas in cardiomyocytes it only induces washout of cytosolic enzymes but not IL-1β (131, 132, 134). Therefore, pyroptosis may occur without secretion of IL-1β.

Supplementary Methods for Pyroptosis Monitoring

Triphenyl tetrazolium chloride (TTC) that stains viable myocardium red and nonviable myocardium unstained can be used to determine pyroptosis because it measures loss of dehydrogenase enzymes due to membrane failure. However, this method cannot distinguish pyroptosis from other forms of cell death where cell membrane integrity is compromised, such as necroptosis.

The release of the soluble cytosolic protein LDH is observed in pyroptosis. LDH release is a classical marker of necrosis, and therefore, it is not sufficient to identify, by itself, caspase-1 triggered pyroptosis. Exposing the isolated rat heart perfused with Krebs buffer to >20 min of global ischemia and 2–3 h of reperfusion will cause a large amount of the left ventricle to infarct (∼75% after 45 min of ischemia). Inhibiting caspase-1/4 with the highly selective inhibitor VRT-043198 will reduce that to ∼45% infarction, revealing a large component of caspase-1/4-dependent injury (10), which could reflect pyroptosis (Table 4). An advantage of the Krebs-perfused heart is that the venous effluent can be collected. The original demonstration of pyroptosis was the release of the soluble cytosolic protein LDH into the medium of cultured macrophages in which NLRP3 inflammasome formation had been stimulated (69). A reduction in LDH in the venous effluent with suppression of caspase-1/4 supports but still does not prove pyroptosis in this model, as caspase 1 is also known to digest important glycolytic enzymes that could also cause necrosis through loss of ATP required for cellular volume regulation (10, 226).

The time course of the membrane failure can be monitored in the isolated rat heart by measuring the rate of LDH washout during reperfusion with timed samples of effluent. By multiplying the volume collected for each sample by the LDH concentration and dividing by the sample duration, the rate of LDH release from the entire heart for each sample time can be calculated. This should then be normalized to the heart weight. Note that in hearts, which were reperfused with VRT-043198 containing buffer, the LDH begins to fall in the third minute of reperfusion, and the two curves continue to diverge until the end of the experiment (10). Thus, this model should be appropriate for studying mouse hearts that have been genetically manipulated to critically test the role of pyroptosis in this model.

Using inhibitors of caspase-1/4 or other components of the inflammasome followed by measurement of troponin T or I levels could also provide an indirect measure of pyroptosis.

Recommendations to Evaluate Pyroptosis

To date, the following parameters have been used to detect the presence of pyroptosis: 1) lytic cell death assessed by the release of cytoplasmic contents (e.g., LDH) and by the entry of cell-impermeable dyes [e.g., propidium iodide (PI)]; 2) activation of the inflammasome assessed by immunofluorescence (e.g., ASC speck) and activation of caspase-1 (and/or caspase-11–4/5) assessed by immunoblotting (e.g., processed fragments, p10/p33 of caspase-1) or assessment of enzymatic activity through cleavage of fluorogenic substrates; and 3) cleavage of GSDMD by immunoblotting (e.g., GSDMD-N). At present, however, a protocol to specifically differentiate pyroptosis from other forms of necrotic death is unavailable, especially considering a potential overlap between the different modes of necrosis. We recommend using multiple techniques described above to evaluate pyroptosis. The strongest evidence that pyroptosis is causing cell death would be a demonstration that the membrane failure is GSDMD dependent.

Additional Methods for Determining Necroptosis, Necrosis, and Pyroptosis

The measurement of viable and nonviable myocardium using TTC is the standard methodology to measure infarct size in laboratory animals (282). The infarct size is measured as the ratio of nonviable myocardium to the entire amount of myocardium that experienced ischemia (area of risk) or left ventricle (LV) by examining transverse sections of the heart from base to apex to evaluate infarct size (IS)/AR and IS/LV, respectively. TTC measures loss of dehydrogenase enzymes due to membrane failure and cannot distinguish between necrosis, necroptosis, pyroptosis, or any other modality of membrane failure.

Measuring LDH release from the isolated heart is a classical marker for necrosis and necroptosis in the damaged heart, for instance, in ischemia-reperfusion injury.

Detection of elevated troponin I or T levels is the mainstay of the diagnosis of acute myocardial infarction (AMI) in patients. As for other methods, plasma troponin levels provide a surrogate marker of infarction in in situ heart models of ischemia-reperfusion. The measurement of troponin levels at a given time point between 1 and 24 h after ischemia or, even better, measuring the area under the curve after three or more time points represents a good estimate of infarct size in the animal in vivo (163). Similarly to the other above-mentioned markers, increased levels of plasma troponin do not differentiate between necrosis, necroptosis, or pyroptosis.

FERROPTOSIS

Ferroptosis is a relatively new form of regulated cell death that was first described less than a decade ago (57). Since its discovery, ferroptosis has been shown to occur in several other disease models, including cancer (119), neurodegeneration (97), liver disease (289), kidney failure (5), and cardiovascular disease (63, 74, 148). The mechanisms underlying ferroptosis were discovered through small-molecule screens of anti-cancer drugs (57). Ferroptosis has characteristics that are distinct from other forms of regulated cell death: iron dependency, disturbances in redox balance mediated by glutathione (GSH) and glutathione peroxidase 4 (GPX4), and lipid peroxidation (Fig. 5) (29). The discovery of ferroptosis in cardiac tissue is even more recent, with proportionately few studies describing ferroptosis specifically in the context of the heart. Despite their limited number, those studies were able to repeat and demonstrate the same characteristic features of ferroptosis in cardiac tissue and successfully use methods of inducing and inhibiting ferroptosis in the heart (11, 12, 63, 74, 148, 153).

Fig. 5.

Signaling pathway of ferroptosis in cardiomyocytes. The defining characteristics of ferroptosis are 1) iron dependency 2), disturbances in glutathione (GSH) and glutathione peroxidase 4 (GPX4)-mediated redox balance, and 3) lipid peroxidation. Pointed arrowheads indicate transport, products, or activation. Blunt arrowheads indicate inhibition. Red arrows indicate ferroptotic activity or features that promote ferroptosis. Blue arrows indicate processes that inhibit ferroptosis. GSSG, glutathione disulfate; ROS, reactive oxygen species; RSL3, Ras-selective lethal 3.

Iron bestows ferroptosis with its namesake and is required for ferroptosis to occur. The importance of iron in ferroptosis was established by experiments that showed that increasing iron within the redox-active labile iron pool (LIP) made cells more prone to ferroptosis (74, 95). The necessity of iron in ferroptosis was established by experiments that showed that chelators and mechanisms that reduce iron in the LIP also inhibit ferroptosis (57, 132). Iron is an essential cofactor in several molecules and enzymes and is especially important in mitochondrial function (141, 238). The constant activity and contractions of cardiac tissue require a constant supply of energy from mitochondria, and thus it follows that the mitochondria compose a significant proportion of the cardiomyocyte volume, which further underscores the importance of iron in cardiac function (15). Iron deficiency impairs the normal functioning of the heart and is associated with cardiovascular disease (262). Conversely, excess iron is also harmful to normal cardiac function, and iron-overload diseases and ischemia-reperfusion injury can also cause accumulation of iron and impaired function in cardiac tissue (46, 71).

Cardiac tissue and cardiomyocytes can import and accumulate excess iron from iron-overload disease or cardiac pathologies (46, 71, 198). Soluble, redox-active free iron in the LIP is thought to be the catalyst that drives increased reactive oxygen species (ROS) production in ferroptosis (29, 250), thus making cardiomyocytes prone to oxidative stress in excess iron conditions (171). In addition to the LIP, excess iron is also absorbed into the mitochondria (63, 198). The mitochondria are already a hotbed of ROS production due to oxidative phosphorylation and H₂O₂ production within their matrices (141, 187), and therefore, excess iron will also cause mitochondrial ROS overproduction (75). Ferroptosis also results in altered mitochondrial morphology and fragmentation through an unknown mechanism (57). Increased ROS production is already associated with several cardiac pathologies, including ischemia-reperfusion injury, left ventricular remodeling, and cardiomyopathy (121, 185). Adding excess iron to the ROS generated by those pathologies increases ROS production beyond already elevated levels, thus pushing cardiomyocytes further toward oxidative stress and the ravaging of intracellular molecules by ROS. Membrane lipids are one of the targets of ROS, which leads to the production of lipid peroxides, a signature hallmark of ferroptosis (Fig. 5). Specifically, polyunsaturated fatty acids (PUFA) are targeted by lipoxygenases for oxidation into detectable byproducts (6, 135). Lipid peroxidation is also observed in various cardiac pathologies (170) and therefore represents another common intersection between ferroptosis and cardiovascular disease.

Redox homeostasis is crucial for the normal functioning of cardiac tissue and even more so for the protection of cardiomyocytes from the constant redox activity and ROS production in the mitochondria (187). In ferroptosis, GPX4 is an important antioxidant enzyme that uses GSH as a substrate to reduce and prevent the accumulation of ROS, thus protecting cells against oxidative stress and lipid peroxidation (40). Chemicals that cause ferroptosis, such as erastin and Ras-selective lethal 3 (RSL3), target either the production of GSH or GPX4 itself (29). Erastin is one of the first known ferroptosis-inducing agents and does so by targeting and inhibiting system Xc⁻ (cystine/glutamate antiporter) (57). System Xc⁻ imports cystine into the cell, where it is reduced into cysteine, one of the three amino acids needed to synthesize GSH. Erastin inhibits system Xc⁻, which deprives cells of cysteine and, consequently, GSH, thereby inhibiting the ability of GPX4 to reduce ROS in the cell (Fig. 5) (40). While system Xc⁻ is not expressed in the heart (142), another study showed that erastin is still able to induce ferroptosis in the heart (11). This suggests that despite the absence of system Xc⁻ in cardiac tissue, erastin may target another mechanism of glutathione synthesis or GPX4 activity in the heart. The mechanism of RSL3 is comparatively straightforward: RSL3 directly inhibits GPX4 activity, thus tipping the redox balance of the cell toward ROS accumulation, oxidative stress, and ferroptosis (Fig. 5) (29, 232).

A recent study investigated the mechanisms underlying doxorubicin cardiotoxicity and provided further evidence that ferroptosis is distinct from apoptosis and necrosis. The study used mice that had defective apoptotic or necroptotic signaling and showed that doxorubicin induced ferroptotic cell death in both types of knockout mice (63). This elegantly designed study demonstrated not only that doxorubicin induces ferroptosis but also that ferroptosis can occur in the absence of canonical apoptotic and necroptotic signaling proteins (63). However, although ferroptosis has distinct hallmarks that distinguish it from other forms of cell death in the heart, some cross-talk can occur. Ferroptosis-inducing agents also cause endoplasmic reticulum (ER) stress and the unfolded protein response, which in turn activate the PERK [protein kinase RNA (PKR)-like ER kinase]/eIF2α (eukaryotic initiation factor 2a)/ATF4 (activating transcription factor 4)/CHOP [C/EBP (CCAAT-enhancer-binding protein) homologous protein] pathway (138). The PERK/eIF2α/ATF4/CHOP pathway can also activate apoptotic and autophagic mechanisms and is observed during ischemia-reperfusion injury in the heart (283). Despite ferroptosis and apoptosis having a common intersection at the ER with the PERK/eIF2α/ATF4/CHOP pathway, ferroptosis does not necessarily cause apoptosis, as ferroptosis can occur without caspase activation, DNA laddering, or other classic apoptotic hallmarks (11, 57, 63).

Identification of Ferroptosis

The requirements, redox mechanisms, and byproducts of ferroptosis also serve as the basis for detecting ferroptosis (Table 5). In terms of mechanisms, disturbances in GSH and GPX4 activity are a unique feature of ferroptosis and are not a central pathway for any of the other types of cell death known to occur in the heart. Decreases in GSH and total glutathione can be measured using chromogenic assays that use 5,5′-dithiobis-2-nitrobenzoic acid (DTNB) as a detection reagent and oxidized glutathione (glutathione disulfate) as a standard (201). Total GPX enzymatic activity can be quantified by measuring the reduction rate of tert-butyl hydroperoxide catalyzed by GSH and nicotinamide adenine dinucleotide phosphate hydrogen (NADPH) (200). Measurements of GPX activity should also be supported with Western blot data to determine whether ferroptotic decreases in GPX4 activity are due to direct inhibition of enzymatic activity or downregulated expression of the GPX4 protein itself (Table 5) (277).

Table 5.

Evaluation of myocardial ferroptosis

Characteristics of Ferroptosis	Differences Between Ferroptosis and other Forms of Cell Death	Methods to Detect Characteristic Features	Limitations	References
GPX4 and GSH	GPX4 activity should be lowered only in cases of ferroptosis and should not be significantly altered in other forms of cell death like apoptosis; ferroptosis should also result in lowered GSH/total glutathione ratios, lower total glutathione, or both.	1) GPX4 activity can be assessed via colorimetric assays that measure GPX4-catalyzed consumption of GSH; 2) GPX4 itself can be detected by an antibody against GPX4.	1) GPX4 activity is often quick in nature, and thus experiments must be time sensitive; 2) antibody detection does not accurately depict the activity level of GPX4. High levels of GPX4 may still have reduced activity when inhibited.	1) 201; 2) 153
Lipid ROS	General ROS production is present in nonpathological conditions and can be found in other forms of cell death such as apoptosis and necrosis; lipid ROS is more specific to ferroptosis and accumulates in much higher amounts than other forms of cell death due to specific decreases in GPX4 activity.	1) Detection: Measurement of membrane-associated fluorescent probes (e.g., liperfluo) that fluoresce upon oxidation; lipid ROS downstream byproducts (e.g., 4-HNE protein adducts, MDA) can be quantified using ELISAs and TBARS assays; 2) inhibitors: Specific antioxidant ferroptosis inhibitors such as ferrostatin-1.	1) Downstream products such as 4-HNE often quickly dilute as time passes and thus create a time sensitive element, and thus fixation and assaying the cells should be done in a timely manner.	1) 104, 267, 281; 2) 11, 57, 63
Iron dependency	Ferroptosis requires iron but does not need abnormally high levels of iron; elevated levels of iron may be a sign that the cells are undergoing ferroptosis; if ferroptosis is occurring, chelators or other ways of reducing intracellular iron should significantly decrease cell death and lipid ROS and restore GPX4 and GSH; other non-iron-dependent forms of cell death should not be significantly affected if iron is removed via chelators or other techniques.	1) Iron can be detected via colorimetric assays that use compounds such as ferrozine, which produces a color when bound to iron; 2) chelators or genetic manipulation of proteins that affect iron concentrations (e.g., TfR, ferritin, FtMt) will also affect ferroptosis.	Iron levels may not indicate levels of ferroptosis and need to be paired with another assay.	1) 100, 218; 2) 63, 74
Supplementary features: mitochondrial morphology change	While mitochondrial fragmentation may not be distinct to ferroptosis; its pairing with high amounts of lipid ROS on the mitochondrial membrane sets it apart.	1) Smaller mitochondria smaller, with increased membrane densities and reduced cristal; 2) signs of cell fragmentation after a few hours of reducing GPX4 function (the morphology of the mitochondria can be assessed using a mitochondria marker like MitoTracker).	Morphology can vary, depending on the cells used, and must be analyzed using an imaging software for quantitative analysis.	57, 101

GPX4, glutathione peroxidase 4; 4-HNE, 4-hydroxynonenal; MDA, malondialdehyde; ROS, reactive oxygen species; TBARS, Thiobarbituric acid-reactive substance.

ROS production plays a significant role in ferroptosis, and ROS can be easily detected by using fluorescent probes such as 2′,7′-dichlorodihydrofluorescein diacetate (11) or more organelle-specific probes such as MitoSOX (184). However, simple ROS detection cannot be used alone to implicate ferroptosis in cardiac cell death, as ROS are also produced in necrosis and necroptosis. Therefore, more specific assays are required to distinguish ferroptotic ROS production from ROS produced by other forms of cell death in the heart.

Fortunately, there are assays that can specifically detect lipid peroxides, which are another distinct characteristic of ferroptosis and a downstream consequence of oxidative stress. Liperfluo (104) and C11-BODIPY (60) are both lipophilic fluorescent probes that associate with lipid bilayers and fluoresce when oxidized by lipid peroxides. Additionally, some specific lipid peroxides, such as 4-hydroxynonenal (HNE) and malondialdehyde (MDA), can be detected via various assays. 4-HNE is formed by the oxidation of polyunsaturated ω-6 fatty acids and can form adducts with membrane proteins (267, 281). 4-HNE production can be measured and visualized using various immunodetection methods, including ELISA (267) and immunohistochemistry (281). MDA is byproduct of further degradation of oxidized PUFAs and can be measured with a colorimetric assay using thiobarbituric acid as a detection reagent (Table 5).

Although the presence of redox-active iron is needed for ferroptosis to occur, three important caveats must be stated. 1) Ferroptosis can still occur at normal intracellular iron concentrations; 2) excess iron can induce ferroptosis, but is not necessary for ferroptosis; and 3) depriving cells of soluble, redox-active iron inhibits ferroptosis (11, 57, 63). Nevertheless, total iron can be detected in cell or tissue lysate samples to determine whether ferroptosis occurred due to an accumulation of excess iron. Ferrozine is an iron-detecting compound that forms a magenta-colored complex when bound to ferrous ions and can be used to detect and quantify total iron up to the nanomolar scale (100). Bathophenanthroline and Ferene-S can also detect total iron in samples using colorimetric assays, although Ferene-S is the more sensitive of the two and requires a smaller sample volume (209). For iron detection on an organismal level, T2 and T2* magnetic resonance imaging can be used to visualize iron deposition in the myocardium and other organs and tissues (26).

Several ferroptosis studies used chelators as a straightforward way of removing redox-active iron from cells without needing to alter the expression of genes and proteins that regulate iron homeostasis. In animal models, treatment with a chelator such as deferoxamine, dexrazoxane, or 2,2′-bipyridyl protects the heart against ferroptosis in ischemia-reperfusion injury and other models of cardiovascular disease (32, 63, 74). If ferroptosis is occurring as a result of experimental or pathological conditions, the addition of chelators to those conditions should inhibit cell death, as ferroptosis requires the presence of redox-active free iron (Table 5).

Recommendations and Strategies For Detecting Ferroptosis and Anti-Ferroptotic Effects

In addition to the ferroptosis detection methods and assays described above, anti-ferroptotic agents such as ferrostatin-1, liproxstatin-1, and vitamin E are also useful in determining whether ferroptosis is occurring (11, 63, 148, 293). All of those anti-ferroptotic agents are lipid-soluble antioxidants that scavenge free radicals, thereby reducing and decreasing ROS and lipid peroxides. If ferroptosis is suspected to be occurring in an experiment, the addition of ferrostatin-1 or a similar ferroptosis inhibitor should decrease the amount of lipid peroxidation and cell death.

Conversely, if a drug or intervention is thought to have anti-ferroptotic activity, it can be tested against treatments that induce ferroptosis, such as excess iron, erastin, or RSL3 (11, 12, 63). Excess iron causes ferroptosis due to an increase in overall ROS production, but treatment with erastin and RSL3 is a more targeted way of causing ferroptosis due to affecting glutathione synthesis and GPX4 activity, respectively. If a drug or intervention truly has anti-ferroptotic activity, it should decrease the amount of lipid peroxidation and cell death caused by ferroptotic agents such as erastin and RSL3. The detailed methods to evaluate ferroptosis are elaborated on in Table 5.

AUTOPHAGIC CELL DEATH

Autophagy is a cellular mechanism of bulk degradation of long-lived proteins and damaged organelles and recycling of the molecular components. It commences with nucleation and proceeds through sequestration of these materials in double-membraned structures termed autophagosomes, fusion of autophagosomes with lysosomes where enzymes catalyze the destruction of the cargo and release of the breakdown products into the cytosol (178). In contrast with other forms of cell death reviewed in this paper, autophagy as a process does not universally command the destruction of the cell. Earliest observations of autophagic structures were made in cells undergoing remodeling during differentiation or in response to stress (45). Whereas autophagic activity was reported in association with cellular demise as early as 1977 (17), autophagy often plays a role in a survival effort by supplying building blocks to facilitate cellular adaptation and nutrients for energy production. If these efforts are not successful, cell death may be observed to be concurrent with but not necessarily consequent to autophagy. Therefore, a distinction must be made between autophagy-associated and autophagy-mediated cell death, and experimental strategies should be designed accordingly.

Autophagy-Associated Cell Death

Autophagy-associated cell death is established by the detection of cell death, using standard approaches described herein, in cells in which autophagy is measured. Transmission electron microscopy (TEM) emerged as the earliest reliable method for visualizing autophagic compartments (241) and remains the tool with the highest resolution for characterizing their morphology. The presence of double membranes enclosing electron-dense cytoplasmic material or recognizable organelles is typical of autophagosomes and early autolysosomes (280). At later stages of degradation, definitive identification of these structures requires immuno-EM (electron microscopy) detection of lysosomal structural proteins or hydrolases. Furthermore, imaging of only a single section of the cardiomyocyte makes this identification more difficult due to significant limitations related to serial sections and representative sampling. For these reasons, collecting quantitative data such as autophagosome number and volume is challenging. Therefore, EM is better reserved for morphological characterization of autophagic vacuoles rather than monitoring changes in autophagic flux.

Alternative methods for microscopic visualization of autophagic machinery commonly involve tracking of autophagy-related proteins, such as microtubule-associated proteins 1A/1B light chain 3 (MAP1LC3; hereafter termed LC3). LC3, one of the mammalian homologs of Atg8, is a ubiquitin-like protein that is cleaved by Atg4 to produce LC3-I, which is then converted to LC3-II by Atg3- and Atg7-facilitated lipidation (103). LC3-II is incorporated into both sides of autophagosome membranes and can also be found on the outside of autolysosomes (274), making it a useful marker of these autophagic compartments (Fig. 6A). Given that an antibody capable of binding exclusively to the lipidated isoform does not exist, we rely on punctate localization of LC3-II versus diffuse distribution of LC3-I in the cytosol to differentiate between the two. For accurate interpretation of LC3 distribution, we recommend fixing samples with neutral-buffered formalin and preparing paraffin-embedded instead of frozen sections. Cryosections are not only inferior in morphology but also preserve lipid droplets, which have been shown to harbor LC3 on their surface (231). Because endogenous levels of LC3 are not robust in cardiomyocytes, permeabilization and antigen retrieval are also necessary. A gentle detergent such as digitonin or saponin can be used to avoid damaging membranes during permeabilization, as LC3-II is membrane-bound.

Fig. 6.

Autophagic cell death pathway and autosis. A: steps of adaptive autophagy/autophagic cell death and proteins involved in different steps. B: factors inducing autosis mediated by autophagy machinery and changes in the cell morphology during early and late phases of autosis. LC3, microtubule-associated proteins 1A/1B light chain 3; RER, rough endoplasmic reticulum.

Levels of LC3 expression may not always reach the threshold of detection even with careful tissue processing. To address this issue, transgenic animals expressing fluorescently tagged LC3 have been developed, starting with the ubiquitously expressed GFP-fused LC3 mouse engineered by Mizushima et al. (179) and Dr. Joseph Hill’s cardiomyocyte-specific, α-myosin heavy chain (α-MHC) promoter-driven GFP-LC3 mouse (292). These transgenic animals harbor higher levels of LC3 in the heart, which aids significantly in immunological detection but does not eliminate the need for it despite the fluorescent tag. While autophagosomes can be directly observed as green puncta in these mice, they can be difficult to distinguish from protein aggregates generated by GFP-LC3 overexpression or autofluorescent structures, such as lipofuscin. Additionally, autolysosomes cannot be visualized by the GFP signal alone, as GFP is quenched within their acidic environment. Use of fluorophore with lower pK_a, such as RFP, or mCherry, which is also more resistant to degradation, circumvents this issue (98).

Tandem reporter mice expressing two fluorophores with differing spectral properties have proven valuable in differentiating nucleation membranes and autophagosomes from autolysosomes (107, 147). Although the data remain only a snapshot of the number of autophagic components at a given time, quantification of components at earlier versus later stages of autophagy using these mice affords an estimate of flux through the system in what is otherwise a static method.

Further insight into the dynamic process of autophagy can be obtained by time-lapse imaging, which is possible by isolation of cardiomyocytes from these reporter animals or by labeling LC3 in in vitro cardiomyocyte models. Lipid or viral delivery of fluorescent-tagged LC3 has been successfully utilized for microscopy in HL-1 and H9c2 cell lines as well as isolated adult mouse cardiac myocytes and neonatal rat ventricular myocytes (NRVMs) (98, 124, 278, 285). Given that gene delivery to isolated cardiomyocytes in culture cannot be accomplished with high efficiency without the use of viral vectors (158), proper controls to account for cytotoxicity due to infection must be included. This involves optimization of multiplicity of infection to a level that maximizes the proportion of cells infected while maintaining cell viability. Because transfection alone can activate stress pathways and induce autophagy, deferring imaging for 24–48 h after infection can help decrease background. To fully account for this stress, we recommend the use of a control virus sharing the same sequence as the experimental vector, with the exclusion of LC3, for baseline comparison. Mutated versions of LC3 that are defective in lipidation or in interacting with ubiquitinated proteins can be substituted based on experimental goals (234). β-Gal substitution, with preservation of the chosen fluorescent protein(s) between the two viral vectors, is also feasible and commonly used (130). Because Atg4 cleaves the COOH-terminal region of LC3, care should be taken to attach the fluorescence tag to the NH₂ terminus. Once the in vitro model is selected, the optimal time frame for expression, adequate for detection with low basal levels of tagged LC3, will need to be determined empirically. Furthermore, to establish that the fluorescence signal truly reflects intrinsic changes in autophagy, it is necessary to evaluate the ratio of endogenous to exogenous LC3 and demonstrate a proportional response to known inhibitors and inducers of autophagic flux using an orthogonal method, such as Western blotting.

Once the in vitro model is established, fluorescence microscopy allows for quantification of autophagic structures at the cell and tissue levels at multiple time points (Table 6). These data can be expressed in many different ways, including but not limited to 1) number of puncta per cell, 2) mean signal intensity per cell, 3) mean puncta size/volume, and 4) total number of cells with puncta (123). We caution against the use of all but the number of puncta per cell divided by the cell area for accurate interpretation of results. Samples from transgenic animals and transfected cardiomyocytes often display fluorescently tagged LC3 puncta even under basal conditions due to high expression levels, meaning a punctum-free state may not be achievable. Furthermore, fluorescence intensity may not directly correlate with levels of autophagy due to the formation of protein aggregates, nonspecific binding of antibodies, and autofluorescence of other cellular structures. Therefore, it is preferable to quantify the number of autophagic puncta with fluorescence intensity over a threshold set to account for the sources of noise listed above. If a tandem reporter system utilizing fluorophores with different pK_a is used, colocalization analysis can be carried out to divide autophagic puncta into double-labeled autophagosome and single-labeled autolysosome pools. Although the results are less specific, colocalization analysis can also be performed on endogenous puncta by simultaneous labeling of autophagic and lysosomal structures via immunochemistry and/or commercially available dyes such as LysoTracker, acridine orange, or monodansylcadaverine (98, 174). Complementary data can be collected on the size, or volume in the case of confocal microscopy, of these puncta after verifying that lysosome fusion is intact, as failure in this process can also lead to enlarged puncta.

Table 6.

Evaluation of myocardial autophagy and features of autosis

Method for Autophagy	In vitro	In vivo	Live Imaging?	Quantitation	Specificity	Flux	Technical difficulty
EM	Yes	Yes	No	No	High	No	High
Fluorescence microscopy	Yes	Yes	Yes	Yes	Medium-High	Possible with additional tags/treatment	Medium
LC3 immunoblotting	Yes	Yes	No	Semi	Low	Possible with additional treatment	Low
Labeled substrate degradation	Yes	Yes	No	Yes	Low, requires proteasome inhibition	Yes	Medium

EM, electron microscropy; LC3, microtubule-associated proteins 1A/1B light chain 3. Features of autosis as follows. Early-stage autosis: 1) increased cell-substrate adherence, 2) nuclear membrane convolution and shrinkage, 3) dysmorphic endoplasmic reticulum. Late-stage autosis: 1) presence of a perinuclear space indicating separation of nuclear membranes focally; 2) ER disappearance; 3) swollen mitochondria.

Out of the four steps of autophagy, 1) nucleation, 2) autophagosome formation, 3) autophagosome to lysosome fusion, and 4) cargo degradation within the autolysosome (Fig. 6A), microscopy-based methods outlined above primarily evaluate steps 2 and 3. Appropriate interpretation of the differences in the number of autophagic compartments requires evaluation of passage of materials through the entire system, as an increase in their steady-state abundance can arise through increased formation (increased flux) and/or decreased clearance (decreased flux). One of the most widely used approaches to distinguish between these possibilities is the deliberate inhibition of flux at the distal end of the flux pathway. A large number of chemicals that inhibit autophagy at each step have been developed (205), but chloroquine, bafilomycin A1, and 3-methyladenine (3-MA) remain the most commonly used in cardiac research (174, 220, 285).

Unlike IHC/ICC, SDS-PAGE-based methods can differentiate between these two isoforms, because the PE moiety of LC3-II increases its hydrophobicity and, despite its larger size, allows it to migrate faster than LC3-I. Given that LC3-II is incorporated into autophagosomes, changes in LC3-II levels as quantified via densitometric analysis of blots are indicative of changes in the number of autophagosomes (Table 6). Alternatively, Western blotting can be used with inhibitors of autophagy to monitor the turnover of p62/SQSTM1, a receptor of polyubiquitinated proteins that interacts with LC3 and is incorporated in autophagosomes to be degraded alongside the protein cargo (202). It should be noted that, because autophagy is an established but not exclusive means of p62 degradation in cardiomyocytes, interpretation of results may be difficult. Furthermore, because expression of p62 can change in response to cellular stress, p62 mRNA levels should be measured in the same samples or synthesis blocked with cycloheximide.

Another approach to assessing autophagic flux involves measuring the breakdown of labeled substances (Table 6). Although diverse methods have been developed for various mammalian cell lines and yeast, the turnover of radioactively labeled long-lived proteins is the most commonly used method in cardiomyocytes (123). Radiolabeled amino acids such as [¹⁴C] valine or [³H]leucine are added to the cell culture medium during a pulse period to be incorporated into newly synthesized proteins, followed by a chase phase without radioactivity to allow for the degradation of short-lived proteins by the proteasome. Afterward, the release of radiolabeled amino acids from long-lived proteins being broken down by autophagy can be quantified as acid-soluble radioactivity at multiple time points (143). Further specificity can be obtained by addition of autophagy and proteasome inhibitors to delineate their relative contributions, but caution should be exercised, as suppression of one degradative pathway may activate the other. Radiolabeling of proteins can also be coupled with quantitative mass spectrometry to measure the degradation of thousands of proteins at once (30). To avoid technical difficulties associated with radioactivity, methods utilizing alternative means of labeling newly synthesized proteins are being developed; stable isotope labeling with amino acids in cell culture (SILAC) has been successfully utilized in cardiomyocytes in vitro and in vivo (224, 275). Work adapting SILAC-based labeling to measure autophagic protein degradation has been accomplished in other cell types (128) but remains to be adapted to cardiomyocytes. For further information on the application of proteomics to study autophagy, readers are directed to a comprehensive review by Wong et al. (272).

Autophagy-Mediated Cell Death

Autophagy-mediated cell death, also known as autophagy-dependent cell death, is the only form of autophagic death recognized by the Nomenclature Committee on Cell Death, emphasizing its distinction from cell death with adaptive autophagy (73). As part of the cellular response to stress, autophagy is commonly upregulated in cardiomyocytes that may later undergo death by necrosis, necroptosis, or apoptosis. Therefore, implicating autophagy as the proximal cause of death, as opposed to a compensatory response occurring in the setting of other death-triggering events, requires exclusion of cell death by alternate pathways. Furthermore, causality between autophagic machinery involvement and regulated cell death must be established, preferably by multiple complementary methods.

To determine conclusively that autophagy is the cause of cell death in a given cardiomyocyte, downregulation of autophagy should promote survival. Chemical interventions described above to block flux can be used for this purpose and cell survival evaluated both in vivo and in vitro (7, 98, 143). However, all well-characterized inhibitors of autophagy lack specificity, a fact that must be taken into consideration when designing and interpreting these studies. For instance, lysosomotropic alkaline chloroquine can inhibit apoptosis as well as autophagy, and 3-MA has autophagy-independent effects as a class III PI 3-kinase inhibitor (205). On the other hand, genetic approaches can be tailored to target specific steps in the autophagic process individually or simultaneously. Induction of autophagy and phagophore formation has been successfully downregulated in cardiomyocytes in vitro by adenovirus-delivered short-hairpin RNA targeting beclin1 (124). siRNA has been used to target autophagosome formation by knocking down atg7 (285) or atg5 (220), and lysosome function has been downregulated using siRNA against tfeb (174, 199) and adenovirus against lamp2 (160). Whenever possible, knockout/knockdown studies should be performed targeting more than one gene, as compensation is possible, and some atgs are involved in cellular processes other than autophagy, including apoptosis (161). Knockdown should be quantitatively confirmed both at the transcript level by qRT-PCR and at the protein level by Western blotting.

Because of the essential role of autophagy in cellular homeostasis, full-body knockout animals may either not survive or exhibit multi-organ dysfunction that contributes secondarily to a cardiac phenotype, as seen with atg5^−/− and atg7^−/− mice (126, 131, 212). Cardiomyocyte-specific phenotypes can be assessed by floxing the gene in question and crossing these mice with a Cre line driven by a cardiomyocyte-specific promoter, e.g., α-MyHC-Cre. Use of various Cre lines, such as MLC2v-Cre and α-MyHC-MerCreMer, also allows for evaluation of the role of a given gene at various developmental stages (189). For all knockout animal studies involving a lox-Cre system, controls should include Flox^+/+ Cre^−/− and Flox^−/− Cre^+/− littermates, as either of these genetic manipulations could elicit a baseline phenotype, depending on the lox insertion site. In addition, if an inducible system that is activated by drug delivery is used, experimental groups should include each genotype receiving treatment or the vehicle control, as α-MyHC-MerCreMer mice exposed to high doses of tamoxifen develop dilated cardiomyopathy (21). If knockout studies prove technically challenging, other strategies include knocking down the gene in question with lentiviral injection (285) or utilizing a haploinsufficiency model (292).

Conversely, if autophagy is truly responsible for cardiomyocyte death, its upregulation should hasten cell death. This can be achieved both in vivo and in vitro by starvation or chemical induction, most notably with rapamycin, the canonical inhibitor of mammalian target of rapamycin. Other agents known to activate autophagy in cardiomyocytes include AMPK activators, Sirt1 activators, and trehalose (225), all of which also elicit nonautophagic events. Similarly, hypoxia and exercise can upregulate autophagy, along with many other cellular processes, and therefore, they should be used with caution in studies testing for a causal role of autophagy in cell death. More targeted approaches with genetic overexpression of autophagic machinery elements include adenoviral overexpression of beclin1 (124) in NRVM and atg7 or beclin1 plasmid transfection in HL-1 cells (24, 285). In vivo techniques include overexpression by adeno-associated virus-9 (AAV9) delivery into the jugular vein of mice (220) and Beclin transgenic mice (292). Also, it should be noted that fluoro-tagged LC3 mouse models described above, such as CAG-RFP-GFP-LC3 transgenic mice, although initially designed for evaluating flux, also manifest inherently higher expression of these components and should be assumed to have increased autophagy at baseline unless it is shown otherwise experimentally. Finally, mice receiving an intraperitoneal injection of the Tat-Beclin-1 peptide, comprising HIV-1 Tat protein transduction domain linked to an 18-amino acid fragment of Beclin1, manifest elevated levels of autophagy in heart among other organs (233).

Autosis

Recently, studies employing Tat-Beclin have uncovered a previously unrecognized form of autophagy-dependent cell death termed autosis (155). In addition to the presence of numerous autophagosomes and autolysosomes indicative of autophagy, additional, distinct, morphological features mark autosis; cells in early stages of autosis are characterized by increased cell substrate adherence, nuclear membrane convolution and shrinkage, and dysmorphic endoplasmic reticulum, and at later stages a perinuclear space is observed, indicative of separation of the nuclear membranes focally (Table 6). These changes in nuclear membrane morphology may arise due to the involvement of the Na⁺,K⁺-ATPase (Fig. 6B), whose inhibition by cardiac glycosides or α₁-subunit knockdown prevent autosis (154). In addition to Tat-Beclin, starvation and hypoxia-ischemia have been shown to trigger autosis in the brain in vivo and in vitro. The role of autosis in cardiomyocyte death remains to be elucidated. Several methods to determine autophagy and autosis are described in Table 6.

Recommendation for Evaluating Autophagic Cell Death

We recommend that the autophagy process should be blocked to conclusively determine autophagic cell death or autosis. Different inhibitors are described above to inhibit autophagy. If blocking autophagy, both in vivo and in vitro, increases survival of myocardial cells, then it can be concluded that autophagic cell death/autosis is involved in the myocardial cell death. The characteristic features of autosis are described in Table 6.

OVERALL DISCUSSION AND CONCLUSIONS

Myocardial cell death is a critical step in the pathophysiology of cardiovascular disease. However, there is a gap in knowledge on how and why a particular form of cell death in preferred in a cardiac disease. Understanding the specific cell death mechanism in cardiomyopathies is important for increasing our fundamental knowledge on how myocardial cells die due to different insults and for developing therapeutic strategy by targeting specific cell death mechanism. One critical step in evaluating cell death in cardiomyopathies is correctly measuring different forms of cell death. It is important to notice limitations of methods during myocardial cell death assessment. Equally important is to pay attention to the common and the distinct/characteristic features of cell death when evaluating myocardial cell death because several forms of cell death can share common features (Fig. 8).

Fig. 8.

Evaluation of myocardial cell death. Different steps to distinguish and identify cell death mechanisms in the heart. GPX4, glutathione peroxidase 4; GSDMD, gasdermine D; LC3BII microtubule-associated proteins 1A/1B light chain 3BII; MLKL, mixed-lineage kinase domain-like; MTP, mitochondrial permeability transition pore; NLPR3, NACHT, LRR, and PYD domains-containing protein 3; PARP, poly-ADP ribose polymerase; RIP3, receptor-interacting protein kinase 3.

It is plausible that more than one form of cell death can occur in the heart during cardiomyopathies, and there can be cross-talk among these forms such as necroptosis can, which can induce pyroptosis via DAMPs (Fig. 7). Availability of ATP could be a deciding factor in initiating specific cell death signaling such as MPT necrosis in absence of ATP and apoptosis in presence of ATP (Fig. 7). One precursor can initiate more than one form of cell death, such as complex I can signal for both apoptosis and necrosis, and oxidative stress can trigger multiple forms of cell death, including apoptosis, MPT necrosis, pyroptosis, and ferroptosis (Fig. 7). Therefore, it is necessary to determine the specific forms of cell death by using its characteristic features and to understand its shared features with other cell death mechanisms. We have elaborated different approaches and techniques to measure characteristic and overlapping features of myocardial cell death mechanisms in Table 7.

Fig. 7.

Interactions between 6 forms of myocardial cell death. The nature of the signals in the myocardium determines the cell death pathway a cell undergoes. However, oxidative damage to mitochondria can trigger multiple cell death pathways. Different colors denote different forms of cell death. DAMPs, damage-associated molecular patterns; LC3, microtubule-associated proteins 1A/1B light chain 3; MPT, Mitochondrial permeability transition; ROS, reactive oxygen species.

Table 7.

Methods to evaluate myocardial cell death

Approach (Technique/Method)	Assessment	Cell Death Mechanism
Imaging
Observation of cell morphology under a microscope	Cell shrinkage, chromatin condensation	Apoptosis
TUNEL assay	DNA fragmentation	Apoptosis, necroptosis/necrosis, normal DNA repair
Annexin V staining with/without vital dye, duramycin with/without vital dye	Exteriorization of phosphatidylserine; exteriorization of phosphatidylethanolamine	No vital dye uptake apoptosis, vital dye uptake, necroptosis/necrosis, pyroptosis
Annexin V and PI staining	Diffused PI staining and intense Annexin V fluorescence; spots of PI with Annexin V-positive blebs	Necroptosis, late apoptosis/secondary necrosis
Immunofluorescence for necrosome formation	Co-localization of RIP3 and MLKL or RIP1-RIP3	Necroptosis
Immunohistochemical/immunocytochemical staining of MLKL	Membrane translocation of MLKL	Necroptosis
Decreased ΔΨm	Intensity of TMRE, JC-1, Rhodamine 123	Necroptosis, MPT-mediated necrosis
Decreased calcium retention capacity	Change in the intensity of calcium fluorescence	MPT-mediated necrosis
Observation of isolated cardiac mitochondria under a microscope	Mitochondrial swelling	MPT-mediated necrosis, autosis
Immunofluorescence of ASC, cleaved caspase-1	Measurement of fluorescence intensity	Pyroptosis
Immunofluorescence of mitoSOX	Measurement of fluorescence intensity	Ferroptosis, necroptosis/necrosis
Observing cell under microscope	Increased substrate adherence, ER fragmentation/loss, nuclear convolution and shrinkage, increased space between nuclear membranes, focal expansion of nuclear membrane	Autosis
Immunofluorescence of LC3-GFP	LC3B-GFP puncta intensity/cell area	Autophagy cell death, autosis
TTC staining	Viable myocardium red, unstained tissue non-viable	Necroptosis/necrosis, pyroptosis, other cell death types
Molecular/biochemical
LDH assay	Colorimetric analyses of releases of cytoplasmic protein LDH	Pyroptosis, necroptosis/necrosis
DNA laddering by agarose gel electrophoresis	DNA fragmentation	Apoptosis, necroptosis/necrosis, normal DNA repair
Western blotting for caspase-3 activity by	Cleaved to total caspase-3 protein	Apoptosis
Western blotting for PARP activity	Cleaved to total PARP protein	Apoptosis
Western blotting for BID	Expression of BID protein	Receptor-mediated apoptosis
Western blotting for RIP3 and MLKL	Phosphorylated to total protein	Necroptosis
Western blotting for MLKL oligomerization	Expression of 160- to 300-kDa proteins of MLKL tetramers and octamers	Necroptosis
Coimmunoprecipitation of RIP1-RIP3 and RIP3-MLKL	Protein-protein interactions	Necroptosis
Cellular fractionation and Western blotting of MLKL	MLKL translocation to membrane by measuring the levels of MLKL in the membrane and cytosolic fractions	Necroptosis
QPCR and Western blotting for NLRP3	mRNA and protein expression of NLRP3	Pyroptosis
Western blotting for caspase-1 activity	Cleaved to total caspase-1	Pyroptosis
Western blotting for IL-1β, IL-18, and GSDMD activity	Cleaved to total IL-1β, IL-18, and GSDMD	Pyroptosis
Western blotting for GSH, GPX4	Protein levels of GSH, GPX4	Ferroptosis
Western blotting for LC3-I and LC3-II	Protein levels of LC3-I and LC3-II	Autophagy

Because only 80–250 myocytes per 10⁵ nuclei undergo apoptosis in the failing human heart (270), it is likely that apoptosis may not be the predominant form of cell death in the failing human heart. There is a possibility of contribution of other forms of cell death, and thus understanding their roles in cardiomyopathies is valuable. The predominant cell death mechanism can vary in different types of cardiomyopathies, based on their microenvironment and the intrinsic factors. Thus, targeting the predominant form of cell death is crucial for developing an intervention approach for myocardial cell death in a specific cardiomyopathy.

To evaluate cell death, it is important to first assess whether the cells are dying by using a viability assay such as ATP/ADP ratio. Depending on cell membrane integrity, cell death mechanisms can be broadly categorized into two groups: 1) with intact cell membranes such as apoptosis, ferroptosis and autophagic cell death; and 2) necroptosis, mitochondrial permeability transition pore (MTP)-mediated necrosis, necroptosis, pyroptosis, and autosis with ruptured cell membranes. Thus, determining the cell membrane integrity could help only as an initial screen for cell death mechanisms. In each category, further assessment of the characteristic features will delineate the specific form of cell death (Fig. 8).

We have provided a brief characterization of six forms of myocardial cell death (Figs. 1–8) and elaborated different methods to evaluate key features of cell death (Tables 1–7). As indicated in several sections of this paper, cell death is a pivotal step in cardiac pathology, and developing guidelines to evaluate myocardial cell death is crucial for determining the cause of cardiomyopathies. Here, we have elaborated various approaches to measure apoptosis, necroptosis, MPT-mediated necrosis, pyroptosis, ferroptosis, and autophagic cell death/autosis. We have explained the concept of a particular method of myocardial cell death, different methods to assess myocardial cell death, limitations of these methods, and the best practices to evaluate a myocardial cell death mechanism. This article is intended to provide guidelines to evaluate myocardial cell death in cardiomyopathies using the best approaches, assessments of the data, and interpretations of the results, which will reduce the variability in results obtained from different laboratories in the world.

GRANTS

This work is supported by National Institutes of Health Grants HL-113281 and HL-116205 to P. K. Mishra; APVV-15-607, VEGA-1/0271/16, and 1/0016/20 to A. Adameova; HL-120732, HL-128215, and HL-126012 to J. A. Hill; HL-136816 to A. Abbate; and HL-094404 to CPB and P20GM113134 to T. Matsui.

DISCLOSURES

A. Abbate has served as a consultant to Astra-Zeneca, Janssen, Merck, Novartis, Olatec and Serpin pharma.

AUTHOR CONTRIBUTIONS

P.K.M. conceived and designed research; P.K.M. and S.K. prepared figures; P.K.M., A. Adameova, J.A.H., C.P.B., P.M.K., J.D., J.N., M.T., A. Abbate, H.C.P., S.S., J.K.H., N.K.K., and T.M. drafted manuscript; P.K.M., A. Adameova, J.A.H., C.P.B., P.M.K., J.D., J.N., M.T., A. Abbate, H.C.P., S.K., S.S., J.K.H., N.K.K., and T.M. edited and revised manuscript; P.K.M., A. Adameova, J.A.H., C.P.B., P.M.K., J.D., J.N., M.T., A. Abbate, H.C.P., S.K., S.S., J.K.H., N.K.K., and T.M. approved final version of manuscript.