Caspase dependent apoptosis

Caspase dependent apoptosis DEFAULT

Caspase-Dependent Apoptosis in Prostate Cancer Cells and Zebrafish by Corchorusoside C from Streptocaulon juventas

Abstract Image

Corchorusoside C (1), isolated from Streptocaulon juventas collected in Vietnam, was found to be nontoxic in a zebrafish (Danio rerio) model and to induce cytotoxicity in several cancer cell lines with notable selective activity against prostate DU-145 cancer cells (IC50 0.08 μM). Moreover, corchorusoside C induced DU-145 cell shrinkage and cell detachment. In CCD-112CoN colon normal cells, 1 showed significantly reduced cytotoxic activity (IC50 2.3 μM). A preliminary mechanistic study indicated that 1 inhibits activity and protein expression of NF-κB (p50 and p65), IKK (α and β), and ICAM-1 in DU-145 cells. ROS concentrations increased at 5 h post-treatment, and MTP decreased in a dose-dependent manner. Moreover, decreased protein expression of Bcl-2 and increased expression of PARP-1 was observed. Furthermore, corchorusoside C increased both the activity and protein levels of caspases 3 and 7. Additionally, 1 induced sub-G1 population increase of DU-145 cells and modulated caspases in zebrafish with nondifferential morphological effects. Therefore, corchorusoside C (1) induces apoptosis in DU-145 cells and targets the same pathways both in vitro and in vivo in zebrafish. Thus, the use of zebrafish assays seems worthy of wider application than is currently employed for the evaluation of potential anticancer agents of natural origin.

Sours: https://pubs.acs.org/doi/10.1021/acs.jnatprod.9b00140

Caspase-independent cell death: leaving the set without the final cut

Abstract

Apoptosis is dependent upon caspase activation leading to substrate cleavage and, ultimately, cell death. Although required for the apoptotic phenotype, it has become apparent that cells frequently die even when caspase function is blocked. This process, termed caspase-independent cell death (CICD), occurs in response to most intrinsic apoptotic cues, provided that mitochondrial outer membrane permeabilization has occurred. Death receptor ligation can also trigger a form of CICD termed necroptosis. In this review, we will examine the molecular mechanisms governing CICD, highlight recent findings demonstrating recovery from conditions of CICD and discuss potential pathophysiological functions of these processes.

Definition and characteristics of caspase-independent cell death

Apoptosis was originally defined by morphological criteria (Kerr et al., 1972). Countless studies have since shown that apoptosis is inextricably linked with caspase activity; apoptotic pathways induce and require caspase function to bring about the orderly demise of a cell (Taylor et al., 2008). We define caspase-independent cell death (CICD) as death that ensues when a signal that normally induces apoptosis fails to activate caspases. Even so, CICD often shares common characteristics with apoptotic cell death (Table 1). These include upstream signalling pathways that are critical for both forms of death such as mitochondrial outer membrane permeabilization (MOMP). Archetypal caspase-dependent events such as phosphatidylserine externalization and wide-scale chromatin condensation are notably absent during CICD. Cells undergoing CICD often display large-scale cytoplasmic vacuolization, autophagosome accumulation and peripheral nuclear condensation. In addition, CICD generally proceeds with much slower kinetics than apoptosis (Ekert et al., 2004). Furthermore, although apoptotic cells exhibit a relatively invariant phenotype, cells undergoing CICD may display widely varying characteristics dependent upon factors such as the initial stimulus, cell type and so on.

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Does caspase-independent cell death occur?

Our view is that the answer to this question is yes but with the proviso that it is not found in all animals. Genetic dissection of apoptosis signalling was begun using the nematode Caenorhabditis elegans as a model organism (Horvitz, 2003). In C. elegans, CED-3 represents the sole executioner caspase. No death is observed in cells destined to undergo apoptosis when CED-3 function is abrogated (Ellis and Horvitz, 1986). Therefore, CICD does not appear to occur in C. elegans. As far as it has been investigated, CICD rarely occurs in Drosophila melanogaster. Cells survive apoptosis induction in the absence of caspase activity in D. melanogaster (Fraser et al., 1997; Muro et al., 2006). Although recent studies have demonstrated alternative cell death pathways occurring in C. elegans and D. melanogaster, these do not fall under our definition of CICD as they manifest in the absence of caspase inhibition (Abraham et al., 2007; Berry and Baehrecke, 2007). The failure of cells, in both C. elegans and D. melanogaster, to undergo CICD is probably due to the absence of MOMP in these organisms (Varkey et al., 1999; Zimmermann et al., 2002).

In higher eukaryotes, the existence of CICD is supported by findings in both cell culture models and studies utilizing mice deficient in apoptotic signalling. Several studies using cell lines have shown that CICD occurs typically by inducing apoptosis in the presence of chemical caspase inhibitors such as zVAD-fmk (Hirsch et al., 1997; Ohta et al., 1997; Sarin et al., 1997). Alternatively, CICD can be observed following the induction of apoptosis in cells expressing genetically encoded caspase inhibitors such as XIAP, CrmA or p35 (Okuno et al., 1998; Denmeade et al., 1999; Wilkinson et al., 2004). With respect to pharmacological or genetic means of caspase inhibition, the argument has been made that perhaps the degree of caspase inhibition was insufficient or did not neutralize all caspases, thereby permitting a low level of caspase activation to drive cell death. Moreover, some commonly used caspase inhibitors such as zVAD-fmk can exhibit off-target effects (Rozman-Pungercar et al., 2003; Misaghi et al., 2006). Perhaps more definitively, CICD has been demonstrated in cells that lack components of the intrinsic apoptotic pathway such as Apaf-1, cytochrome c or caspase-9 (Hakem et al., 1998; Kuida et al., 1998; Li et al., 2000). Analagous to the lack of CICD in lower organisms, CICD in higher eukaryotes often requires MOMP. This is perhaps best illustrated in one study that compared CICD in cells overexpressing Bcl-2 (blocking MOMP) or lacking Apaf-1. CICD was only observed in cells that had undergone MOMP (lacking Apaf-1) (Haraguchi et al., 2000). However, as will be discussed further, there are examples of CICD that do not require MOMP.

Mice deficient in proteins required for caspase activation exhibit phenotypes consistent with the occurrence of CICD in vivo. Interdigital web loss is a characteristic apoptosis-dependent process that occurs during embryonic development. This process is signalled through the intrinsic apoptotic pathway requiring MOMP leading to apoptosome (Apaf-1/cytochrome c)-dependent activation of caspase-9. Importantly, genetic ablation of Apaf-1 slows but does not prevent interdigital web loss (Yoshida et al., 1998). It is intriguing to note that the phenotype of mice lacking Bax and Bak (therefore unable to undergo MOMP) is much more severe than those unable to activate caspases downstream of MOMP (Apaf-1/caspase-9 null mice) (Lindsten et al., 2000). This suggests that CICD can, at least partly, compensate for a lack of caspase-dependent apoptosis during development. Although supporting an in vivo role for CICD, mouse knockouts have also shown that CICD cannot fully compensate for a lack of apoptosis as mice lacking Apaf-1, caspase-9 or expressing non-apoptogenic cytochrome c frequently display pre- or perinatal lethality characterized by cleft palate and forebrain outgrowth (Cecconi et al., 1998; Hakem et al., 1998; Yoshida et al., 1998; Lindsten et al., 2000; Hao et al., 2005). These phenotypes are consistent with a defect in cell death that presumably cannot be rescued by CICD. One caveat of this interpretation is that there is no evidence that the extra neurons observed in the forebrain have previously received a signal to die (for example, did they undergo MOMP). Another hypothesis that may account for this phenotype is that slower cell death (under conditions of CICD) leads to an overabundance of survival or growth signalling resulting in aberrant proliferation of neighbouring cells (Chipuk and Green, 2005).

Collectively, these studies have convincingly demonstrated that diverse apoptotic stimuli can stimulate CICD. However, the question can be asked whether physiological CICD occurs. Some differentiated cell types, such as certain neuronal populations and cardiomyocytes, express low levels of Apaf-1 and might represent a physiological setting in which CICD occurs (Sanchis et al., 2003; Johnson et al., 2007). Supporting this, several studies have shown that CICD occurs in cardiomyocytes and neurons following apoptotic stimulation (Stefanis, 2005; Bahi et al., 2006). One study examining the removal of interdigital webs during limb bud development supports the physiological occurrence of CICD in vivo (Chautan et al., 1999). Using morphological criteria and Apaf-1 null mice as a reference, these authors concluded that up to 10% of dying cells in the developing limb bud undergo CICD. Clearly, the occurrence of CICD in vivo requires further investigation. Such in vivo studies are hampered (as are those in cell culture) by the lack of readily available tools to easily detect MOMP and CICD. It is likely that during CICD, besides the lack of caspase activity, MOMP will occur under most circumstances. Cells undergoing CICD fail to expose ‘eat-me’ signals such as phosphatidylserine found on apoptotic cells. This should allow cells undergoing CICD to persist longer in vivo thereby facilitating detection. Perhaps coupling fluorescent fusion proteins used to mark MOMP in cell culture such as cytochrome c-GFP with a means to rule out caspase activity will enable CICD detection in vivo (Goldstein et al., 2000).

Means to an end: how cells die in the absence of caspase activity

Death receptor-induced necroptosis

Apoptotic death receptor signalling is well elucidated. In short, following binding of their respective ligand, death receptors recruit intracellular adaptor molecules enabling dimerization and activation of caspase-8. Active caspase-8, in turn, drives the downstream apoptotic cascade (Lavrik et al., 2005). Intriguingly, several studies have shown cell death in the absence of caspase activation following death receptor ligation (Figure 1) (Jaattela and Tschopp, 2003). Here, we use the term necroptosis to describe CICD induced by death receptors to distinguish this from MOMP-induced CICD, as the mechanisms of death are almost certainly distinct (Degterev et al., 2005).

Death receptor-activated necroptosis. When caspase activity is blocked, death receptor activation can drive necroptosis through the upregulation of PLA2 activity that, in turn, increases oxidative stress. RIP-1 kinase is also activated triggering necroptosis by directly acting upon mitochondrial function or, perhaps, by affecting autophagy. PLA2, phospholipase A2; RIP-1, receptor-interacting protein-1.

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Tumour necrosis factor (TNF) stimulation can induce necroptosis. One study found that in vivo treatment of mice with TNF and the caspase inhibitor z-VAD-fmk led to rapid death within 4 h (compared with 24 h when TNF is administered alone) (Cauwels et al., 2003). Death was attributed to massive upregulation of reactive oxygen species as it was blocked by coadministration of antioxidants. Phospholipase A2 (PLA2) activation by TNF was a major contributor to reactive oxygen species production. PLA2 undergoes caspase-dependent cleavage during TNF-induced apoptosis. The authors’ proposed model is that PLA2 is activated following TNF treatment and under apoptotic conditions undergoes caspase cleavage, rendering PLA2 and the necroptotic pathway non-functional. When caspase activity is inhibited, reactive oxygen species generation by PLA2 contributes to necroptosis. One potential caveat of these studies is that zVAD-fmk can display some toxicity issues in vivo (Chauvier et al., 2007). However, another study using CrmA to inhibit caspase activity has also demonstrated the occurrence of necroptosis following TNF treatment (Vercammen et al., 1998).

Fas-induced apoptosis and necroptosis share some common molecular components. Fas-associated protein with death domain (FADD) acts as an essential adaptor molecule as it links death receptor oligomerization to caspase-8 activation. FADD is also required for necroptosis induced following Fas activation as loss of FADD expression blocks both Fas-induced apoptosis and necroptosis (Holler et al., 2000). Furthermore, FADD oligomerization itself is sufficient to induce necroptosis (Kawahara et al., 1998). However, FADD does not appear to be involved in TNF-induced necroptosis as no effect was observed following loss of FADD or overexpression of a dominant negative form (Holler et al., 2000).

Further dissection of death receptor-induced necroptosis has revealed a key function for receptor-interacting protein-1 (RIP-1). RIP-1, an intracellular adaptor molecule with kinase activity, has earlier been shown to be involved in recruiting the IκB kinase complex to the TNF receptor enabling nuclear factor-κB activation. Cells in which RIP-1 is lacking or in which RIP-1 kinase function is inhibited are refractory to necroptosis mediated by either TNF or Fas (Holler et al., 2000; Degterev et al., 2008). In contrast to its function in nuclear factor-κB signal transduction, several studies have shown a requirement for RIP-1 kinase function in signalling necroptosis (Kelliher et al., 1998). One possible mechanism by which RIP-1 can lead to necroptosis is by disrupting the interaction of the adenine nucleotide translocase with cyclophilin D. This, in turn, leads to mitochondrial dysfunction, although it remains unclear how RIP-1 signals to the mitochondrial inner membrane (Temkin et al., 2006).

An alternative means by which RIP-1 function could drive necroptosis has been suggested in a separate study (Yu et al., 2004). In this study, it was found that reduction of caspase-8 function through RNA interference or by using zVAD-fmk induces necroptosis. This suggests that a low level of caspase-8 activity is required for cell viability. Necroptosis induction was dependent upon RIP-1 and JNK function and appeared to occur through macroautophagy (hereafter termed autophagy). Autophagy is a homoeostatic cellular process regulating protein and organelle turnover by lysosomal destruction (Levine and Kroemer, 2008). Although inhibition of autophagy through RNAi inhibited immediate cell death, it was not examined whether these cells ultimately survived. Although there are putative links between death receptor signalling and autophagy, such as a molecular interaction between FADD and Atg5, it remains unclear as to whether an autophagic death pathway is driven by death receptors under conditions of necroptosis (Pyo et al., 2005). Furthermore, the idea that autophagy, per se, can promote cell death remains an open question.

Mitochondria and CICD

Death receptor-induced necroptosis may be restricted to certain cell types. In stark contrast, CICD, under conditions where MOMP has occurred, appears nearly universal. Numerous studies have demonstrated that various MOMP-inducing stimuli trigger CICD in a wide variety of cell types. Furthermore, direct induction of MOMP through ectopic Bax expression is sufficient to induce death in the presence of caspase inhibitors (Xiang et al., 1996). MOMP may contribute by one of several ways to bring about CICD. The main models by which MOMP can bring about CICD are either through a general decline in mitochondrial function and/or through the release of mitochondrial proteins that can actively induce CICD (Figure 2).

Mitochondrial roles in caspase-independent cell death. In the absence of caspase activity, following mitochondrial outer membrane permeabilization (MOMP), various mitochondrial intermembrane space proteins are released triggering loss of mitochondrial function and/or proactively contributing to cell death. Simultaneously, mitochondrial fission is activated and fusion inactivated thereby disrupting mitochondrial morphology. Collectively, these events contribute to caspase-independent cell death (CICD).

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Mitochondria have various critical cellular functions. Perhaps the most important function of mitochondria is the generation of ATP through the process of aerobic respiration. However, other mitochondrial functions such as the generation of acetyl CoA, nicotinamide adenine dinucleotide production and buffering intracellular calcium levels are also important. Under conditions of organelle damage, for example MOMP, some of these processes such as acetyl CoA generation for lipid biogenesis might be even more critical. Under apoptotic conditions, caspase-3 cleaves the p75 subunit of mitochondrial respiratory chain complex I. Cleavage of p75 leads to disruption of electron transport, loss of Δψm and reduction of ATP levels (Ricci et al., 2004). However, under conditions of CICD, mitochondria gradually lose Δψm even though caspase-dependent cleavage of p75 is blocked (Colell et al., 2007). Why this occurs is not clear but one reason may be a gradual release of respiratory chain components from the mitochondria following MOMP. Gradual loss of Δψm consequently leads to the loss of ATP generation and, ultimately, cell death independent of caspase activity. However, loss of ATP generation following MOMP most likely cannot account for CICD in transformed cells as they predominantly rely upon glycolysis for ATP generation. Besides ATP generation, maintenance of Δψm is required for mitochondrial protein import, loss of which is likely to impact on many different mitochondrial functions (Gasser et al., 1982). The relative contribution of other MOMP-induced mitochondrial defects to CICD remains an open question.

Concurrent with MOMP, the mitochondrial network undergoes dramatic fragmentation. This is due to the increased rates of mitochondrial fission coupled with decreased fusion (Arnoult, 2007). Mitochondrial fusion has been reported to have a protective function in cellular homoeostasis and enables cells to maintain a homogenous Δψm (Chen et al., 2003). It is thought that fusion may achieve this by promoting mitochondrial complementation thereby restoring function to damaged mitochondria. It is unknown whether long-term disruption of mitochondrial morphology also contributes to CICD.

Mitochondrial outer membrane permeabilization leads to the release of cytochrome c from the mitochondrial intermembrane space (IMS). Besides cytochrome c, numerous IMS proteins such as Smac/Diablo, HtrA2/Omi, Endonuclease G (Endo G) and apoptosis-inducing factor (AIF) are released (Susin et al., 1999; Du et al., 2000; Verhagen et al., 2000; Li et al., 2001; Suzuki et al., 2001). Various groups have proposed that these proteins can actively induce CICD following mitochondrial release.

Apoptosis-inducing factor, perhaps the best-studied example of a CICD mediator, is slowly released from mitochondria following MOMP in a caspase-independent manner although caspase activity may accelerate the release (Arnoult et al., 2003; Munoz-Pinedo et al., 2006). Following release, AIF translocates to the nucleus. Various in vitro studies have shown that nuclear AIF mediates chromatin condensation and CICD (Lorenzo et al., 1999). In vitro models of embryonic body cavitation suggested that AIF might have a function in cell death observed during cavitation; however, others have found no such effect in vivo (Joza et al., 2001; Brown et al., 2006). It has become clear that AIF has an important function in aerobic respiration. For example, Harlequin mice (which contain a proviral insertion in the AIF gene) display an 80% reduction in AIF levels. These mice exhibit a phenotype in line with defects in mitochondrial respiratory problems including progressive neuronal and retinal degeneration (Klein et al., 2002). Furthermore, cells lacking AIF are defective in electron transport chain complex I/III activity (Klein et al., 2002; Vahsen et al., 2004). From these studies, it is clear that mitochondrial release of AIF following MOMP would compromise mitochondrial function. This strongly suggests that, rather than acting as a positive mediator of CICD, AIF release promotes CICD by contributing to loss of mitochondrial function. However, a recent study addressed the two functions of AIF using mitochondrially tethered forms of AIF. These AIF proteins were not released from the mitochondria following MOMP. The authors found that these tethered proteins could not rescue enhanced CICD observed in Apaf-1 null relative to Apaf-1/AIF null neurons, suggesting an active role for AIF during CICD (Cheung et al., 2006). Ultimately, generation of knock-in mice using these or other approaches and crossing with mice deficient in intrinsic apoptosis signalling will reveal whether there is an in vivo role for nuclear AIF in mediating CICD.

Other mitochondrial IMS proteins that have been proposed to have a function in CICD include EndoG. EndoG is the predominant endonuclease of the mitochondrial IMS and is thought to be involved in regulating mitochondrial biogenesis, DNA synthesis and repair. Following MOMP, EndoG is released from the mitochondria whereupon it can translocate to the nucleus and, perhaps, induce DNA degradation (Li et al., 2001). It is unclear as to whether EndoG is involved in mediating CICD. Paradoxically, some studies have suggested that its release may require caspase activity (Arnoult et al., 2003). However, this may not always be the case. For example, a recent study in ischaemic cardiomyocytes demonstrated EndoG release following MOMP in a caspase-independent manner (Bahi et al., 2006). Revealingly, suppression of EndoG levels by RNAi had no effect on CICD.

HtrA2/Omi is a mammalian serine protease that resides in the mitochondrial IMS. Perhaps relevant to its primary function, HtrA2/Omi is evolutionarily conserved. Bacterial homologues exhibit chaperone activity (Vande Walle et al., 2008). Following MOMP, HtrA2/Omi is released into the cytoplasm whereupon it can interact with XIAP thereby promoting caspase activation during apoptosis (Suzuki et al., 2001). A more relevant function of HtrA2/Omi in CICD is that it exhibits serine protease activity. Ectopic cytoplasmic expression of HtrA2/Omi is sufficient to induce cell death that is not inhibited by zVAD-fmk. However, in vivo studies have convincingly demonstrated that, rather than being involved in cell death, HtrA2/Omi function is required for normal mitochondrial homoeostasis. HtrA2/Omi null mice exhibit muscle wasting and neurodegeneration (Jones et al., 2003). These are traits common to defects in mitochondrial physiology, in line with the function of HtrA2/Omi as a mitochondrial chaperone. Importantly, cells lacking HtrA2/Omi are more sensitive to cell death following stimulation with various stimuli arguing against a prodeath function for this protein.

Waking the dead: recovery from MOMP and resistance to CICD

Mitochondrial outer membrane permeabilization has often been viewed as a point of no return as it appears to commit a cell to death regardless of caspase activity. However, as we will discuss further, this is not always the case.

Sympathetic neurons deprived of nerve growth factor undergo MOMP and apoptosis that can be blocked by inhibiting caspase function (Deshmukh and Johnson, 1998; Martinou et al., 1999). Neurons typically express low levels of Apaf-1 protein and, as such, endogenous caspase inhibitors such as XIAP are at sufficient levels to block the ability of cytochrome c to induce apoptosis (Potts et al., 2003). Nerve growth factor deprivation induces a so-called competence to die likely due, in part, to antagonism of XIAP function by Smac/Diablo or HtrA2/Omi released following MOMP (Deshmukh et al., 2002). Importantly, re-addition of nerve growth factor under these circumstances effectively restores cell viability and therefore represents an example of recovery from CICD. It may be that similar recovery from CICD in vivo is responsible for the extra neurons and forebrain outgrowth observed in Apaf-1 and caspase-9 null mice. It is unclear as to the mechanism/s enabling neuronal recovery from CICD. Perhaps the increased glycolytic levels in neurons facilitate recovery. This would lessen the necessity for mitochondrial-dependent ATP production and mitigate the energy crisis that occurs following MOMP. However, this cannot be the sole factor determining recovery from CICD as many tumour cell lines that are highly glycolytic fail to recover from conditions of CICD.

In contrast to neurons, which are non-dividing, a recent study showed that proliferating cells can recover from conditions of CICD (Colell et al., 2007). In this study, the authors employed a retroviral screen to identify proteins that could promote recovery from CICD. Using this approach, the authors found that glyceraldehyde-3-phosphate dehydrogenase (GAPDH) could effectively protect cells from CICD downstream of MOMP. GAPDH was shown to promote recovery dependent upon two functions. First, the well-characterized function of GAPDH in glycolysis was shown to be necessary. A second novel function of GAPDH in promoting autophagy was also described and required for cellular recovery. Under conditions of CICD, GAPDH participates in transcriptional upregulation of ATG12 and enhances autophagy. Under these circumstances, as appears generally the case, autophagy acts primarily as a protective stress response. Upregulation of autophagy enabled mitochondrial clearance leading the authors to speculate that perhaps autophagy was required to clear the cell of damaged mitochondria. Alternatively or additionally, autophagy may also have facilitated energy production by catabolic breakdown of cellular components.

Many key questions remain as to how cells can recover from CICD. One critical issue is from where the ‘new’ mitochondria arise. Two equally exciting possibilities exist: either ‘MOMP-ed’ mitochondria can effectively heal and re-seal their outer membrane or some mitochondria may evade MOMP thereby providing a seed population to repopulate the cell. How exactly GAPDH stimulates autophagy and what functions autophagy has during cellular recovery remain to be deciphered. Moreover, how autophagy specifically targets damaged mitochondria, if indeed it does, is largely undefined (Yu et al., 2008). It is also likely that GAPDH represents only one of many mediators that enable recovery from CICD. Although technically challenging, means of assessing whether cells can recover from conditions of CICD in vivo should be sought.

CICD in disease and therapy

Aberrant levels of apoptosis have been linked with numerous diseases. For example, increased levels of apoptosis contribute to pathological damage observed following acute ischaemic damage in neuronal and cardiac tissues. Inhibition of apoptosis has multiple functions in tumorigenesis and in governing the sensitivity of tumour cells to conventional chemotherapies. Therefore, manipulating apoptosis obviously represents an attractive and much sought after therapeutic aim. It is also likely that pathways leading to CICD or recovery from CICD also represent therapeutic targets.

Cells are subject to ischaemia during neuronal stroke or cardiac infarction. Ischaemia, among other effects, typically induces an MOMP-dependent apoptotic programme leading to cell death (Chan, 2004; Gustafsson and Gottlieb, 2008). If MOMP is considered as a point of no return, then one would expect that inhibition of caspases would afford no protective effect. However, several studies have shown that administration of caspase inhibitors can provide significant clinical efficacy in murine models of stroke (Balsam et al., 2005; Braun et al., 2007). Taken at face value, this may indicate that cells destined to undergo apoptosis can effectively recover from CICD-inducing conditions. Perhaps an even better clinical outcome could be obtained through manipulation of metabolic pathways and/or enhancement of autophagy. However, as is the case for the extra neurons in the Apaf-1 or caspase-9 null mice, there is no proof that caspase inhibition during stroke enables cellular protection from CICD-inducing conditions. It remains possible that neurons dying under conditions of CICD perhaps release survival factors promoting survival in surrounding tissue. Furthermore, inhibition of other caspases that are primarily involved in inflammatory responses, such as caspase-1, may also provide a protective effect. A further caveat is that off-target inhibition of other proteases, such as calpains, may account for the ability of certain caspase inhibitors to provide a neuroprotective effect (Waterhouse et al., 1998).

Cells must evade apoptosis at multiple stages of the metastatic process for cancer to arise (Green and Evan, 2002). Apoptosis barriers to tumour formation include apoptotic signals driven by growth-promoting oncogenes, detachment from the extracellular matrix and apoptotic signalling induced by a hypoxic low-nutrient environment. As such, cancer cells have developed numerous ways to inhibit apoptosis. Perhaps the best described is upregulation of antiapoptotic Bcl-2 family members thereby inhibiting MOMP (Chipuk and Green, 2008). However, besides inhibition of MOMP, there is a growing body of evidence that caspase inhibition downstream of MOMP is a common event in multiple tumour types. This may support a role for recovery from CICD in promoting tumorigenesis. Moreover, if this is the case, it has implications for new therapies that solely rely on inducing MOMP as a means of killing tumour cells.

Several studies have shown loss of Apaf-1 expression in a variety of tumour cell lines. This was first described in metastatic melanoma in which Apaf-1 expression was lost due to the loss of promoter methylation (Soengas et al., 2001). Downregulation of Apaf-1 expression in metastatic melanoma correlates with poor prognosis (Fujimoto et al., 2004). Reduction of Apaf-1 expression has also been observed in other tumours such as acute myeloid leukaemias, cervical carcinoma and colorectal cancer, where low Apaf-1 levels correlate with disease progression (Jia et al., 2001; Leo et al., 2007; Zlobec et al., 2007). Besides loss of Apaf-1 expression, tumour cells can inhibit apoptosome-dependent caspase activity in other ways. For example, the oncogenic Bcr-Abl kinase, besides inhibiting MOMP, can also block apoptosome activity by inhibiting caspase-9/Apaf-1 binding (Deming et al., 2004). Other less well-defined means of inhibition of apoptosome activity have also been reported (Wolf et al., 2001). Importantly, apoptosome function can also be modulated by other proteins, intracellular ion and nucleotide levels (Schafer and Kornbluth, 2006). Under such conditions, levels of Apaf-1 or caspase-9 might be completely normal, whereas apoptosome function and caspase activity are compromised. Besides its well-characterized apoptotic function, a recent report demonstrated an apparent role for Apaf-1 in mediating DNA damage-induced cell cycle arrest (Zermati et al., 2007). Loss of this alternative Apaf-1 function would also be expected to impact on tumorigenesis. Various tumours have been shown to overexpress the endogenous caspase inhibitor, XIAP, which dependent on tumour cell type can correlate with a poor prognosis (Tamm et al., 2004). In addition, inhibition of XIAP function restores chemosensitivity in various tumour cell lines (Sasaki et al., 2000). Besides its ability to inhibit caspases, XIAP also has alternative functions that may be relevant for tumorigenesis such as its function in transforming growth factor-β and nuclear factor-κB signalling (Yamaguchi et al., 1999; Lu et al., 2007).

Collectively, these data indicate that tumour cells can inhibit caspase activation pathways downstream of MOMP and this might be physiologically relevant for both tumour progression and therapeutic sensitivity. As it appears that MOMP, in the absence of caspase activity, leads to CICD it is not unreasonable to predict that tumour cells can also recover from CICD. The discovery that GAPDH alone can promote recovery of proliferating cells under CICD-inducing conditions may be relevant for tumours (Colell et al., 2007). Metastatic cells typically display high levels of GAPDH, glycolytic activity and autophagy, all of which were deemed necessary for recovery from CICD (Revillion et al., 2000; Degenhardt et al., 2006; Deberardinis et al., 2008). Loss of Apaf-1 has been shown by some to promote transformation in vitro although this has been questioned by others (Soengas et al., 1999; Scott et al., 2004). Another study has shown that expression of a dominant negative version of caspase-9 efficiently promotes tumorigenesis in a myc-driven model of lymphoma (Schmitt et al., 2002). Finally, a recent study has shown that inhibition of clonogenic growth in a lung cancer cell line following chemotherapy could only be achieved by enhancing apoptosome activity. Evidently, these cells could survive and proliferate under conditions of CICD (Hoffarth et al., 2008).

If cancer cells can block the intrinsic apoptosis pathway and recover from CICD what is the best way to kill them? At least in some situations autophagy and glycolysis appear essential for recovery from CICD. It may be that inhibiting these processes using already available autophagy inhibitors such as chloroquine and/or glycolysis inhibitors, for example 2-deoxyglucose, will effectively block cellular recovery. Alternatively, attempts to restore apoptosome function under conditions of CICD might prove of therapeutic benefit.

Conclusion

Caspase-independent cell death is most likely engaged under physiological and pathological situations. However, understanding how cells actually die during CICD remains limited. In contrast to the study of apoptosis, there are limited means to detect CICD. More specifically, tools need to be developed to effectively discern CICD in vivo. The recent finding that cells can recover from conditions of CICD poses a number of key basic research questions. Moreover, this finding has clear implications for oncogenesis and the treatment of cancer. Accordingly, one might expect that apoptosis inducers coupled with drugs that block recovery from CICD might represent effective anti-cancer therapies.

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Sours: https://www.nature.com/articles/onc2008311
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Caspase-dependent cell death-associated release of nucleosome and damage-associated molecular patterns

Abstract

Apoptosis, which is anti-inflammatory, and necrosis, which is pro-inflammatory, represent the extremes of the cell death spectrum. Cell death is complex and both apoptosis and necrosis can be observed in the same cells or tissues. Here, we introduce a novel combined mode of cellular demise – caspase-dependent regulated necrosis. Most importantly, it is mainly characterized with release of marked amount of oligo- or poly-nucleosomes and their attached damage-associated molecular patterns (DAMPs) and initiated by caspase activation. Caspase-activated DNase has dual roles in nucleosomal release as it can degrade extracellularly released chromatin into poly- or oligo-nucleosomes although it prohibits release of nucleosomes. In addition, osmotically triggered water movement following Cl influx and subsequent Na+ influx appears to be the major driving force for nucleosomal and DAMPs release. Finally, Ca2+-activated cysteine protease, calpain, is an another essential factor in nucleosomal and DAMPs release because of complete reversion to apoptotic morphology from necrotic one and blockade of nucleosomal and DAMPs release by its inhibition.

Main

Apoptosis is characterized by membrane blebbing, cellular shrinkage, nuclear condensation, nuclear fragmentations, oligo-nucleosomal DNA fragmentation and formation of apoptotic bodies. These characteristics are attributed mainly to the caspase family of cysteine proteases.1,2 Necrosis is distinguished from apoptosis by cellular swelling, plasma membrane rupture, absence of oligo-nucleosomal degradation and, finally, rapid lysis of cells and cellular constituents including damage-associated molecular patterns (DAMPs) are massively exuded extracellularly to activate inflammatory and immune responses. 3, 4, 5

Calpains are a family of Ca2+-activated cysteine proteases consisting of 15 genes. Among them, μ-calpain (calpain I) and m-calpain (calpain II) are ubiquitously expressed in most cells as a heterodimer consisting of a large subunit (80 kDa; calpain 1 of μ-calpain and calpain 2 of m-calpain) and a common small subunit (29 kDa; calpain S1), which is processed into a smaller heterodimer (18–78 kDa) upon activation by Ca2+. Ubiquitous calpains are regulated by an endogenous inhibitor, calpastatin.6

It has long been observed that both apoptosis and necrosis can be simultaneously detected in tissues or cell culture. Therefore, apoptosis and necrosis have been assumed to be two extremes of the cell death spectrum capable of inter-conversion by key regulators.5,7 In this study, we introduce a novel mode of cell death involving the combination of apoptosis and necrosis, being a caspase-dependent process with necrotic morphology, involving the active release of DAMPs bound to nucleosomes.

Results

Release of nucleosomes and DAMPs from amino-acid-deprived HeLa cells

Amino-acid-deprived HeLa cells die and release of various pro-inflammatory mediators,8,9 with cellular morphology displaying detached and round plasma membranes, and diminished nuclei (Figure 1A). Surprisingly, when loaded with membrane-impermeable DNA dye, SYTOX Green, extracellular DNA release was revealed (Figure 1C). Amplification of genomic and mitochondrial gene sequences from the released DNA indicated the DNA was originated from nuclei and mitochondria (Figure 1B). Confocal microscopy revealed the released DNA co-stained with all major histones (Figure 1E), could be stained with SYTOX analogous with cellular bodies (Figure 1G), demonstrating the release of nucleosomes by dying cells. DNA also colocalized with interleukin 6 (IL6), high mobility group protein B1 (HMGB1), heat shock protein (Hsp) 90 and ERp57, a thiol oxidoreductase of the endoplasmic reticulum, which are characterized DAMPs (Figures 1D and F). The released DNAs and DAMPs was significantly increased by partial digestion with micrococcal nuclease (MNase) (Figures 1H and I), indicative of connection of released nucleosomes and DAMPs to cellular bodies. In live imaging, cells loaded with SYTOX Green and membrane-permeable DNA dye, DRAQ5, showed typical apoptotic morphologies, such as vigorous nuclear and cytoplasmic shrinkage, at early times, with abrupt plasma membrane swelling and the release of genomic DNA being evident after 5 h. DNA release continued slowly over the next 5 h together with slow contractions of swelled plasma membrane (Figures 1J and K,Supplementary Movie 1). Transmission electron microscopy (TEM) demonstrated swelled but intact nuclear membrane, partially condensed chromatin in the vicinity of nuclear membrane, sparse chromatin within the nucleoplasm and chromatin-like electron-dense material in the cytosol. Later examination of cells revealed disintegrated plasma membranes and small deflated nuclei with intact nuclear membranes (Figure 1L). Confocal microscopy revealed nuclei delineated by lamin A/C and nuclear pores appeared to have little DNA. In contrast, almost all DNA was located in the cytoplasm (Figure 1M), indicating that the cytosolic, chromatin-like, electron-dense material evident in TEM was definitely chromatin.

Release of nucleosomes and DAMPs from amino-acid-depleted HeLa cells. (A) An inverted microscopic image of HeLa cells in the condition of amino-acid depletion. Arrows designate dying HeLa cells. (B) Genomic sequences of glyceraldehyde-3-phosphate dehydrogenase (GAPDH), Fas, cytochrome oxidase subunit 1 (Co1) and ATP synthase subunit 6 (ATP6) were PCR amplified from extracellularly released DNA, genomic or mitochondrial DNAs. (C) Inverted and fluorescent microscopic images were taken from amino-acid-deprived HeLa cells in the presence of SYTOX, a membrane-impermeable DNA dye. HeLa cells deprived of amino acids for 15 h were fluorescence stained with histone H1 or IL6 antibodies (D), histone H2A, H2B, H3 or H4 antibodies (E) or HMGB1, Hsp90 or ERp57 antibodies (F) in combination with histone H1 antibody and 4',6-diamidino-2-phenylindole (DAPI). (G) Amino-acid-deprived HeLa cells were stained with SYTOX to determine viability, fixed and stained with DAPI and histone H1 antibodies. (H) Amino-acid-deprived HeLa cells were untreated or treated with MNase (500 mU/ml) for 10 min. Released DNA was quantitated at the indicated times. Data from triplicate samples are presented as mean±S.D. (I) Conditioned media from amino-acid-deprived HeLa cells treated or untreated with MNase were western blotted with histone H1, 2B, H3, H4, IL6, ERp57, HMGB1 or Hsp90 antibodies. (J) Images captured every hour from live imaging of amino-acid-deprived HeLa cells with SYTOX (green) and DRAQ5, membrane-permeable DNA dye (red). (K) SYTOX fluorescent intensities were measured from circularized areas of live imaging of amino-acid-deprived cells in 5-min intervals. (L) TEM images of control cells (La) and amino-acid-deprived HeLa cells (Lb–Ld). (M) Amino-acid-deprived HeLa cells were fluorescence stained with lamin and nuclear pore antibodies, or lamin antibody and wheat germ agglutinin (WGE)

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Apoptotic features of dying HeLa cells during amino-acid deprivation

During nucleosomal release (Supplementary Figure 1a, upper left), cell death measured by propidium iodide (PI) staining increased over time (Supplementary Figure 1a, upper middle), and mitochondrial membrane potential rapidly decreased (Supplementary Figure 1a, upper left). Interestingly, the increased activity of caspases 3 and 7 noted until 24 h showed similar time courses with DNA release (Supplementary Figure 1a, lower left). In contrast, cells did not show any Annexin V-positive populations (Supplementary Figure 1a, lower middle) and TUNEL-positive populations (Supplementary Figure 1a, lower right) in accordance with no subG1 population (Supplementary Figure 1b). The cells showed cleavage of effector caspases and degradation of caspase substrates (Supplementary Figure 1c). Collectively, cells showed several apoptotic features, albeit no DNA fragmentation and no exposure of phosphatidylserine.

Nucleosomal release is a common phenomenon provoked by various cytotoxic stimuli from various cells

Nucleosomal and DAMPs release is not a phenomenon confined to amino-acid deprivation. As well-known apoptosis inducers including an anticancer drug, VP16, tumor necrosis factor-alpha (TNFα) plus cycloheximide or staurosporine could promote the release of DNA in HeLa cells (Figure 2a). Several human cell lines showed comparable or lesser DNA release with HeLa when staurosporine treated until >50% of cellular viabilities reduced (Figure 2b) indicating that nucleosomal release is a phenomenon occurred by various cytotoxic stimuli formerly known to induce apoptosis, regardless of its cellular specificity. For example, U937 displayed activated caspases, cleavage of poly ADP ribose polymerase-1 (PARP-1) and lamin A/C (Figure 2c), but little or no release of nucleosomes and DAMPs (Figures 2b and d) with definitive DNA laddering and nuclear fragmentations, whereas HeLa did not show any DNA and nuclear fragmentation in both staurosporine treatment (Figures 2e and f) and amino-acid deprivation (Figure 1I and Supplementary Figure 1a, lower right and 1b). In confocal microscopy, staurosporine treatment showed nucleosomal release in HeLa, in contrast, nuclear fragmentations in U937 (Figures 1D–F and 2g), suggesting apoptotic cell death without fragmentation of intracellular DNA and nuclei would be a key figure of the cell death releasing nucleosomes and DAMPs.

Nucleosome release is a common phenomenon provoked by various cytotoxic stimuli from various cells. (a) Death of HeLa cells was caused by incubation with amino-acid-depleted medium (HBSS), VP16 (100 μM), TNF-α (50 ng/ml) plus cycloheximide (25 μg/ml), or staurosporine (1 μg/ml). Released DNA was measured with PicoGreen DNA dye. (b) Various human cell lines were incubated with staurosporine (1 μg/ml) to the time when >50% of cellular viability was reduced in a Calcein assay to detect extracellularly released DNA. (c) Staurosporine-treated HeLa cells and U937 cells were western blotted with caspases 9, 3, 6, 7, lamin A/C, PARP-1 and tubulin antibodies. Arrows and arrow heads designate parental proteins and cleaved fragments, respectively. (d) Total lysates (T) and conditioned medium (CM) from staurosporine-treated HeLa and U937 cells were western blotted with histone (H1, H2A, H2B, H3, H4), IL6, ERp57, HMGB1, Hsp60 and Hsp90 antibodies. (e) Genomic DNAs prepared from staurosporine-treated HeLa and U937 cells were separated by agarose gel electrophoresis. (f) Electron microscopy images were taken from HeLa and U937 cells treated with staurosporine for 8 h. (g) HeLa cells (upper panel) and U937 cells (lower panel) treated with staurosporine for 7 h, were fluorescence-stained with lamin A/C antibody, nuclear pore antibody and DAPI. Data from triplicate samples are presented as mean±S.D. (a and b)

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Release of nucleosomes and DAMPs is not associated with NETosis

Morphological features of dying HeLa appear to be very similar with NETosis, in that cells release DAMPs attached to nucleosomes instead of antimicrobial molecules. Regulatory factors of NETosis include Nox-derived reactive oxygen species (ROS), autophagy and histone citrullination by peptidylarginine deaminase (PAD).10 Autophagy, ROS and PAD did not confer any effect on nucleosomal and DAMPs release investigated through specific inhibitors and short hairpin RNA (shRNA) transfection (Supplementary Figures 2). Therefore, cell death releasing nucleosomes and NETosis are totally different from one another, although morphologically very similar.

Comparison between cell death involving release of nucleosomes and DAMPs, and primary necrosis

Next, we compared the cell death with primary necrosis. Primary necrosis induced by heating or H2O2 treatment showed earlier peak of DNA release than amino-acid deprivation or staurosporine treatment (Figure 3a). No fragmentation of genomic DNA was evident in all the conditions (Figure 3b, left panel). The released DNA from heated or H2O2-treated cells were completely degraded, whereas, surprisingly, the DNA released from staurosporine-treated and amino-acid-depleted cells showed classical apoptotic DNA cleavages into inter-nucleosomal fragments of roughly 180-base pairs and their multiples (Figure 3b, right panel). Heated cells released few histones and DAMPs. On the contrary, H2O2-treated cells released histones and DAMPs (Figure 3c). Total protein released from staurosporine-treated cells was analogous with that from H2O2-treated cells, but much more than from heated cells. The amount of released mitochondria, secretory pathway and cytosol proteins of staurosporine-treated cells was markedly higher than those of heated or H2O2-treated cells, investigated by GFP ELISA with conditioned media from cells expressing mitochondria-targeted GFP or GFP of secretory pathway, or LDH assay for detecting release of cytosolic components (Figures 3d and e). Furthermore, histone H1 and HMGB1 released from staurosporine-treated cells were bound to the released DNA, although those from heated or H2O2-treated cells were not, according to chromatin immunoprecipitation (ChIP) assay with conditioned medium (Figure 3f), implying that the cell death induced by staurosporine treatment releases proteins containing higher mitochondrial, cytosolic and secretory components, and DNAs as poly- or oligo-nucleosomes in comparison with primary necrosis in which totally degraded DNA and disjoined histones and DAMPs from DNA are released from the dying cells.

Comparison between cell death involving release of nucleosomes and DAMPs, and primary necrosis. HeLa cells were depleted of amino acids (HBSS), treated with staurosporine (1 μg/ml) (Stau) or H2O2 (32 mM) (H2O2), or heated at 56 °C for 1 h and incubated at 37 °C (Heat) during the indicated time periods. DNA release was measured with PicoGreen and data from triplicate samples are presented as mean±S.D. (a). Cellular genomic DNA (Pellet) and released DNA (SNT) were separated in agarose gels with molecular weight marker from the cells with amino-acid deprivation for 24 h, staurosporine treatment for 12 h, heating at 56 °C for 1 h and incubation at 37 °C for 3 h, or H2O2 treatment for 4 h (b). Released protein was western blotted for histones, IL6, ERp57, HMGB1, Hsp60 or Hsp90 (c). HeLa cells expressing green fluorescence protein (GFP) targeted to nucleus, mitochondria or secretory pathway were imaged via confocal microscopy with DAPI (d). Released GFP (Mito, Nuclear, Secretory) from HeLa cells was measured with GFP ELISA, total released protein (Total) was measured by the Bradford assay, and released lactate dehydrogenase (LDH; Cyto) was measured by a LDH assay (e). Data executed as triplicate are presented as mean values of released protein relative to released protein from staurosporine-treated HeLa cells±S.D. The concentrated conditioned medium from HeLa cells was immunoprecipitated with control IgG, anti-histone H1 antibody, or anti-HMGB1 antibody. The purified DNA from the antibody–protein–DNA complexes was PCR-amplified for GAPDH sequences (f)

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Release of nucleosomes and DAMPs in dying HeLa cells is dependent on caspase

To explore molecular mechanisms of cell death, the effects of cell death inhibitors were examined. There were no effects on release of nucleosome and DAMPs with co-treatment of Necrostatin-1, an inhibitor of necroptosis, or AG14699, an inhibitor of PARP-1-dependent cell death. Co-treatment with 3-methyl adenine (3MA), an inhibitor of autophagy, increased DNA release as well as protein release even further. On the contrary, nucleosomal and DAMPs releases were strikingly reduced by co-treatment with pan-caspase inhibitor zVAD-fmk and caspase 3 inhibitor zDEVD-fmk in staurosporine-treated and amino-acid-deprived cells (Figures 4a and c, and Supplementary Figures 5a and c). In addition, cellular viability was increased markedly using zVAD-fmk co-treatments with staurosporine, but not in the condition of amino-acid depletion (Figure 4b and Supplementary Figure 5b). In cells transfected with shRNA for caspases 1, 3, 6, 7 and 9 (Figure 4d), knock-down of caspases 3 or 9 significantly blocked DNA release (Figure 4e and Supplementary Figure 5d), which also showed notably decreased effector caspase activities (Figure 4f and Supplementary Figure 5e). Cells overexpressing caspase 3 displayed slight increase in DNA release than controls. Moreover, increased DNA and protein release were evident in cells overexpressing caspase 3 but knocked-down for caspase 9 when compared with cells knocked-down for caspase 9 (Figures 4g–i), indicating that caspases 3 and 9 are essential in release of nucleosomes and DAMPs.

Release of nucleosomes and DAMPs from dying HeLa cells is dependent on caspase. HeLa cells were incubated with staurosporine (1 μg/ml) in the presence of 3MA (10 mM), necrostatin-1 (Nec, 20 μM), zVAD-fmk (20 μM), zDEVD-fmk (20 μM) or AG014699 (10 μM) for 8 h. DNA release (a) and % viability (b) were measured by PicoGreen DNA dye method and Calcein assay, respectively. Released protein was western blotted for histones, IL6, ERp57, Hsp60 and Hsp90 (c). HeLa cells knocked-down by shRNA for caspases 1, 3, 6, 7 and 9 were confirmed on decreases of caspases by western blots (d), incubated with staurosporine, and DNA release at 8 h (e), and caspases 3 and 7 activity at 6 h (f) were monitored. HeLa cells knocked-down caspase 9 and/or overexpressing HA-tagged caspase 3 (g) were staurosporine-treated for 8 h, where released DNAs were measured by PicoGreen method (h) and released protein was detected by Western blot for histone H2B, histone H4, IL6, ERp57, HMGB1, Hsp60 or Hsp90 (i). Data from triplicate samples are presented as mean±S.D. (a, b, e, f and h). *P value<0.01

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Roles of caspase-activated DNase in nucleosomal release in dying HeLa cells

An intriguing finding was that apoptotic DNA laddering was observed only in the released DNA but not in nuclear DNA (Figure 3b). This prompted investigation of the role of caspase-activated DNase (DFF40/CAD) and its mitochondrial equivalent, endonuclease G (EndoG), which both function in DNA fragmentation during apoptosis. Addition of DNase I extracellularly during cell death decreased DNA release (Figure 5a) and release of histones (Figure 5b) in dose-dependent manners, implying that DNase activity impaired nucleosomal release. Moreover, knock-down of CAD increased DNA release and overexpression of CAD decreased release of DNA and histones, whereas there was no effect on release of DAMPs except for Hsp90, although knock-down or overexpression of EndoG had no effect on release of both nucleosomes and DAMPs (Figures 5c–e). Overexpression of CAD induced complete degradation of genomic and released DNAs (Figure 5f). These data are suggesting that CAD inhibits nucleosomal release through degrading nuclear DNA even if implicated in fragmentation of released nucleosomes, possibly in the extracellular space. Supporting this notion, released DNAs were additionally fragmentized by further in vitro incubation (Figure 5g), and CAD as well as inhibitor of CAD (ICAD) were released bound to extracellular released nucleosomes (Figures 5h and i).

Roles of CAD in release of nucleosomes and DAMPs from dying HeLa cells. Death of HeLa cells was induced by treatment of staurosporine (1 μg/ml) together with DNase I of the indicated concentrations, and released DNA (a), or histones and DAMPs (b) were identified by PicoGreen staining or western blot, respectively. In HeLa cells, either knocked-down or overexpressed for CAD or EndoG, confirmed by western blots (c), release of DNA (d) or histones and DAMPs (e) were measured by PicoGreen staining or western blots, respectively, after staurosporine treatments. (f) HeLa cells or cells transfected with control vector, CAD cDNA or EndoG cDNA were incubated with staurosporine for 10 h, from which genomic DNA (left panel) or released DNA (right panel) was separated by agarose gel electrophoresis. (g) Released DNA from HeLa cells treated with staurosporine for 10 h, was further incubated for the indicated times and analyzed by agarose gel electrophoresis. (h) CAD and ICAD were identified by western blot at the conditioned medium from staurosporine-treated HeLa cells (arrow: large isoform of ICAD; arrow head: small isoform of ICAD). (i) HeLa cells were incubated in staurosporine for 7 h and fluorescent-stained for CAD and histone H1. Data executed from triplicate samples are presented as mean±S.D. (a and d)

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Chloride channels are associated with release of nucleosome and DAMPs

Release of genomic DNA coincided with the beginning of cellular swelling (Figure 1j and Supplementary Movie 1), suggesting the possibility that osmotically triggered water movement and succeeding cellular swelling, caused by increased ion influx, may drive the release of nucleosomes and DAMPs. These were tested using blockers of various ion channels in the plasma membrane. Very interestingly, DNA release was obstructed nearly completely by the Cl channel inhibitors, 4,4′-Diisothiocyanatostilbene-2,2′-disulfonic acid (DIDS) and 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB), partially inhibited by the Na+ channel inhibitor, amiloride, and further prohibition of DNA release was observed by their combined use (Figure 6a), in accordance with decreased release of histones and DAMPs, whereas the inhibitors did not significantly affect cellular viability and death (Figures 6b–d). The effects were affirmed also in the condition of deprivation of amino acids (Supplementary Figures 6a–c). Requirement of Cl current in release of DNA was additionally validated by the cells in either Na+-free or Cl-free medium, and subsequently DNA release was completely inhibited in Cl-free medium (Figure 6e and Supplementary Figure 6d). Co-treatment of DIDS with staurosporine produced massive nuclear fragmentation (Figure 6f), increased TUNEL-positive population (Figure 6g) and inter-nucleosomal DNA fragmentation (Figure 6h). Therefore, hallmarks of apoptosis became apparent by inhibition of Cl channel. In fact, intracellular Cl concentration gradually increased with peak value at 10 h (Figure 6i) and Cl current also increased with a peak value 4 h after treatment of staurosporine (Figure 6j). Furthermore, DIDS partially suppressed Cl current, although amiloride produced little change (Figures 6k and l). However, the combined treatment of DIDS and amiloride completely blocked Cl current in staurosporine-treated cells (Figure 6m), indicating that the voltage gradient induced by Cl influx may have been neutralized by following Na+ influx, resulting in osmotically triggered movement of water and swelling. In addition, co-treatment of zVAD-fmk with staurosporine also inhibited Cl current (Figure 6n), indicating the probability that caspase activation may be associated with activation of Cl channels.

Chloride channels are associated with the release of nucleosomes and DAMPs in dying HeLa cells. (a) HeLa cells were incubated with staurosporine (1 μg/ml) in combination with various ion channel inhibitors for 12 h; an inhibitor of epithelial Na+ channel, amiloride hydrochloride hydrate; Cl channel inhibitors, DIDS (200 μM) and NPPB (200 μM); an inhibitor of Na+/Cl cotransporter, HCT (100 μM); an inhibitor of Na+ K+ ATPase, ouabain (100 μM); an inhibitor of non-selective cation channel, flufenamide (100 μM); an inhibitor of stretch-activated ion channel, Gad (III) chloride hexahydrate (40 μM); an inhibitor of K+ channel, tetraethyl ammonium chloride (TEA) (5 mM); an inhibitor of Na+/K+ cotransporter, bumetanide (100 μM). The released DNA was measured with PicoGreen dye (a) and the released protein from cells treated with staurosporine together with solvent, amiloride, DIDS or amiloride and DIDS was western blotted for histones and DAMPs (b). HeLa cells were treated with solvent or ion channel inhibitors with or without staurosporine for 8 h. Viability was measured with Calcein assay (c) and cell death was detected by SYTOX red staining (d). HeLa cells were incubated in MEM, Na+-deficient medium or Cl-deficient medium with staurosporine (1 μg/ml) for 10 h, and the released DNA was measured by PicoGreen (e). HeLa cells were incubated with staurosporine in combination with control solvent or DIDS. Nuclear fragmentation was detected with confocal microscopy (f), TUNEL staining (g) or agarose gel electrophoresis after genomic DNA preparation (h). Cellular chloride ion contents were measured by MQAE fluorescence in cells incubated with MEM containing either control solvent or staurosporine at the indicated time periods (i). Cellular Cl currents were measured in cells treated with staurosporine by halide-sensitive YFP quenching at the indicated times (j). HeLa cells treated with solvent or staurosporine in the presence or absence of DIDS, amiloride, DIDS and amiloride, or zVAD for 4 h and chloride ion currents were measured with halide-sensitive YFP quenching methods (kn). Data performed in triplicate are presented as mean±S.D. (a, c, d and e, right panel of g and i)

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Calpain activity is a prerequisite for nucleosome and DAMPs release

To evaluate the roles of intracellular free Ca2+, the effects of Ca2+ chelator, BAPTA-AM and the Ca2+ ionophore, A23187 were examined. BAPTA-AM reduced DNA release, whereas A23187 considerably increased DNA release in amino-acid-deprived or staurosporine-treated cells (Figure 7a). Furthermore, intracellular free Ca2+ was increased after staurosporine treatment (Figure 7b), implicating the presence of Ca2+-mediated effectors in nucleosomal release. Thus, we estimated the role of Ca2+-activated cysteine proteases, calpains because of their association with apoptotic and necrotic pathways.11,12 Calpain inhibitors ALLN or PD150606 partially blocked the release of nucleosomes and DAMPs except for Hsp90 (Figures 7c and d). Genomic DNA showed inter-nucleosomal fragmentation (Figure 7e), and nuclear condensation and fragmentation were produced upon co-treatment with PD150606 (Figure 7f) indicating that inhibition of calpain can convert necrotic cell death into apoptotic one. In contrast, there was no change in cellular viability (Figure 7g). Calpain activities were increased at 2 h, when DNA release and activities of caspases 3, 7 and 9 did not reach maximal levels, and decreased to the basal level by 4 h after staurosporine treatment (Figure 7h). Knock-down of calpastatin or overexpression of calpain 2 significantly increased DNA release, but knock-down of calpain S1 or overexpression of calpastatin considerably decreased DNA release (Figure 7i and Supplementary Figures 7a and b). In addition, calpain 2 began to show limited N-terminal degradation at 2 h after staurosporine treatment during the periods of increased calpain activities, whereas calpain 1 did not (Figure 7j), indicative of selective calpain II activation and its association with nucleosomal and DAMPs release. Regardless of previous reports revealing calpastatin degradation by caspases and subsequent calpain activation,13, 14, 15 calpastatin cleavage by caspases was not implicated in calpain activation, because DNA release from calpastatin mutant cells of caspases cleavage sites was not considerably different from that of calpastatin-overexpressing cells (Supplementary Figures 7c and 8a) and cleavage of calpastatin, although blocked by co-treatment of zVAD-fmk, only appeared in the beginning at 4 h at a time of no increased calpain activity (Supplementary Figure 8b). On the other hand, Cl currents were significantly inhibited by calpain inhibitors, indicating that one of possible roles of calpain could be activation of Cl channels (Figure 7k).

Calpain activity is a prerequisite for the release of nucleosomes and DAMPs in dying cells. HeLa cells were incubated in amino acid-depleted medium (HBSS) or staurosporine-containing medium (Stau) (1 μg/ml) for 24 or 8 h, respectively, in the presence of solvent control, BAPTA-AM (50 μM) or A23187 (1 μg/ml). Released DNA was quantified with PicoGreen (a). Amount of intracellular free Ca2+ was measured in cells treated with solvent control or staurosporine at the indicated time periods by eFluor 514 (b). Cells were treated with staurosporine for 8 h in the presence of control solvent, ALLN (20 μM), or PD150606 (20 μM), and released DNA or protein was detected by PicoGreen staining (c) or western blot for histones and DAMPs (d), respectively. Genomic DNA was separated in agarose gel electrophoresis from staurosporine-treated cells with or without PD150606 (e). The cells treated with staurosporine and PD150606 for 8 h, were stained for lamin A/C and histone H1, and examined by confocal microscopy; the arrows indicate nuclear fragmentations (f). Cells treated with control solvent, zVAD-fmk (20 μM), or PD150606 with or without staurosporine for 8 h were measured for viabilities with Calcein assay (g). Cells treated with staurosporine were measured for calpain activity, caspases 3 and 7 activity, or released DNAs at the indicated time periods (h). HeLa cells transfected with non-targeting shRNA (NT), calpastatin shRNA, calpain S1 shRNA, or overexpressing calpain 1, calpain 2, or calpastatin were incubated with staurosporine for 8 h and the released DNAs were measured by PicoGreen staining (i). Cells expressing calpain 1 or calpain 2 tagged with Flag at the N-terminus and HA at the C-terminus were treated with staurosporine and western-blotted with anti-Flag antibody or anti-HA antibody (j). HeLa cells expressing halide-sensitive YFP were treated with staurosporine and/or PD150606 for 4 h, and their chloride ion currents were measured by fluorometry by detecting quenching of YFP fluorescence (k). Data performed in triplicate are presented as mean±S.D. (ac and gi)

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Discussion

Recent advances in biology have differentiated regulated necrosis from the programmed cell death termed apoptosis.16, 17, 18, 19 Our data confirm that cell death involving the release of nucleosomes and DAMPs follows morphological necrosis (Figure 1), but is a caspase-dependent process (Figure 4). Therefore, the process should be classified as a kind of caspase-dependent regulated necrosis most importantly involving the conversion into apoptotic phenotypes by inhibition of the calpain pathway or Cl channel (Figures 6 and 7).

Necroptosis is a best understood caspase-independent regulatory necrosis, initiated by TNFα–TNFR1, FasL-Fas or Toll-like receptor pathway when apoptosis is blocked, requiring involvement of receptor interaction protein kinase 1 and 3 (RIPK1 and RIPK3).20, 21, 22 Execution phase of necroptosis has been largely unknown, however, implication of several elements including ROS, reactive nitrogen species, inhibition of mitochondrial adenine nucleotide translocase and phospholipase A2-lipoxigenase have been recently elucidated.22,23 Necroptosis and caspase-dependent regulated necrosis are presumed to be similar in morphological aspects, although both are different from each other in the requirements of caspases, RIPK1 or ROS (Figure 4 and Supplementary Figure 3). In addition, nucleosomal release in necroptosis is not reported, yet.

Apoptosis does not induce inflammation because its major regulators and processes have focused on eliminating pro-inflammatory materials.24,25 Contrastingly, the cell death shown here is completely at odds with this notion because caspases activation was indispensible for releasing the inflammatory materials. Although caspases function in initiating death processes and seem to be required for opening Cl channels, its precise molecular mechanisms remain unclear (Figures 4 and 6n).

DNA and nuclear DAMPs are reportedly known to be released in different types of cell death: late apoptosis or secondary necrosis, necrosis, or NETosis.26 Secondary necrosis is a necrotic change of terminal phase apoptotic cells when late apoptotic products are not adequately eliminated by nearby scavengers,27,28 from which extracellular nucleosome and DAMPs can be generated as membrane-bound vesicles after formation of apoptotic bodies,29 whereas DAMPs and nucleosomes are directly released in caspase-dependent necrosis as DAMPs bound poly-nucleosomes during relatively early phase when effector caspases are fully activated and mitochondrial membrane potential does not completely decline without apoptotic body formation and DNA fragmentation (Figure 1 and Supplementary Figure 1).

Major divergence of primary necrosis produced by H2O2 treatment or heating from caspase-dependent necrosis was a release of completely degraded DNA and detached DAMPs (Figures 3b and f). Moreover, some of DAMPs such as IL6 and ERp57 were not released in the primary necrosis (Figure 3c). Conclusively, both types of cell deaths are ultimately different from each other.

NETosis, neutrophil-specific form of generalized term ‘ETosis’ is a kind of regulated necrosis, releasing neutrophil extracellular trap (NET) composed of chromatin, and cytoplasmic and granular products,30 and coordinated by Nox-generated ROS, PAD4-mediated chromatin decondensation and autophagy.10,31 In spite of its morphological similarities just as rapid, explosive dislodge of chromatin and attached components, the regulatory factors of NETosis were not connected to release of nucleosome and DAMPs in the caspase-dependent necrosis (Supplementary Figure 2).

Nucleosomal release is associated with various cytotoxic stimuli (Figure 2a) well-known to promote apoptosis by different mechanisms but absolutely depending on activities of caspases 3 and 9 (Figure 4 and Supplementary Figure 5). But nucleosomes were not always equally released in all the cells. For examples, HeLa and A549 showed relatively very high amount of DNA release compared with other cells, on the contrary, BEAS-2B, Huh7 and U937 showed little release of DNA (Figure 2b), suggesting the probability that cell death processes accompanying caspase activation can be subdivided into necrotic (caspase-dependent regulated necrosis) or apoptotic (apoptosis) at the execution phase in cell type-specific manners.

One candidate modulator determining the cell fates seems plausible to be calpain systems as its prohibition converted necrotic status to be completely apoptotic (Figures 7e and f) notwithstanding no acquaintance with its exact molecular mechanism. Calpains have been known to have an important role in caspase-dependent demises.32, 33, 34, 35, 36, 37, 38, 39, 40, 41 According to the data in Figure 7, calpain II was instantaneously activated early and its inhibition strikingly reduced nucleosomal release with no influence on cellular viability, indicating that instant early calpain activation is not crucial for cell death but needed for the imperative process implicated in nucleosomal and DAMPs release, possibly by regulating Cl channel activity. Consistent with this, it has been previously reported that calpains contribute to necrotic deaths through increase of plasma membrane permeability to ions, progressive disruption of cytoskeleton and plasma membrane proteins, and mitochondrial dysfunction.42,43

One of hallmarks of apoptosis is oligo-nucleosomal fragmentation orchestrated by DFF40/CAD. Nevertheless, this has been hardly detected in some types of cells despite the presence of effector caspase activation.44, 45, 46, 47 Moreover, it may not be an essential factor for apoptosis.48, 49, 50, 51 In this regard, our results reveal the probable role of CAD to execute its extracellular function as a bound form to released chromatin, leading to digestion of chromatin, although overexpression of CAD reduces nucleosomal release (Figures 5g–i).

Apoptotic volume decrease (AVD) concurrent with cell shrinkage is induced by activation of K+ and Cl channels at early apoptosis before caspase activation,52 whereas necrotic volume increase is caused by activation of Na+ channels.53 Dying HeLa cells initially showed classical AVD but promptly began to release their DNA with cellular swelling (Figure 1j and Supplementary Movie 1), accompanying the increase of intracellular free Cl concentration and Cl influx (Figures 6i and j). Thus, we postulate that osmotically triggered water flow with Cl influx is a major driving force for the release of nucleosomes and DAMPs, although, hitherto, the specific Cl channels are not yet certain. Cl channels have been demonstrated to function mainly in cellular volume regulation and fluid secretion. Recent findings also have shown their close connection with various human diseases.54,55

In conclusion, we propose a novel entity of cellular demise, caspase-dependent regulated necrosis. It can be defined as follows (Figure 8): characterized by massive release of poly- and oligo-nucleosomes and their attached DAMPs and displaying necrotic morphology with no fragmentation of nuclei and DNA; subdivided from apoptosis at the step of caspases 9 and 3 activation, hence, the cell death can be blocked by zVAD-fmk; instant activation of calpains is needed for release of nucleosomes and DAMPs, subsequently, inhibition of calpain convert the necrotic cellular demise to the apoptotic one; osmotically triggered water movement following Cl influx and secondary Na+ influx is a critical factor, possibly through providing the driving force for nucleosomal and DAMPs release.

Schematic diagram showing the suggested molecular mechanisms of caspase-dependent regulated necrosis. Caspases 9 and 3 are sequentially activated by various cytotoxic stimuli (1). At the same time, focally released Ca2+ from either ER or mitochondria in the cellular stress activates calpain II (2), which opens Cl ion channels of plasma membrane cooperating together with caspase 3 by unknown mechanisms (3). Cl influx and secondary Na+ influx, mediated by voltage gradient produced by Cl influx, induce osmotically driven water movement into cytosol and subsequent cellular swelling (4). Hydrostatic pressure, generated by intracellular water increment, squeezes out nuclear chromatin through nuclear pore damaged by caspases to be released extracellularly by way of cytosol. During that time, nuclear and cytosolic DAMPs and discharged CAD from ICAD by caspase 3 are attached to the released chromatin (5, 6). Finally, the extracellular chromatin and its attached DAMPs are partially digested into poly- or oligo-nucleosomes with DAMPs by the CAD (7)

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Materials and Methods

Cells, antibodies and other reagents

HeLa cell line (human cervical cancer) was cultured in minimal essential media (MEM) supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin. HCT116 and HCT116 Bax (−/−) (human colon cancer), U87MG (human malignant glioma), U251MG (human neuronal glioblastoma), Hep3B, HepG2, and Huh7 (human hepatoma), DU145 and PC3 (human prostatic cancer), MDA-MB231 and MCF7 (human mammary cancer), HEK 293T (human kidney epithelium) cell lines were cultured in DMEM supplemented with 10% FBS, 2 mM L-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin. U937 (human histiocytic lymphoma) was cultured in RPMI1640 supplemented with 10% FBS, 2 mM L-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin. SKOV3 and SKOV3ip (human ovarian adenocarcinoma) was cultured in RPMI1640 supplemented with 20% FBS, 2 mM L-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin. Staurosporine and etoposide (VP16) were purchased from Cell Signaling Technology (Danvers, MA, USA). Cycloheximide was purchased from Calbiochem (Darmstadt, Germany). Human TNFα was purchased from R&D Systems (Minneapolis, MN, USA). Cl-amidine was purchased from Cayman Chemical (Ann Arbor, MI, USA). zVAD-fmk, zDEVD-fmk, Necrostatin-1, BAPTA-AM, diphenyliodonium, PD150606, ALLN and A23187 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). DNase I, N-acetyl cysteine (NAC), ascorbic acid, 3MA and AG14699 were from Sigma-Aldrich (Yongin, Korea). An inhibitor of epithelial Na+ channel, amiloride hydrochloride hydrate; Cl channel inhibitors, DIDS and NPPB; an inhibitor of Na+/Cl cotransporter, hydrochlorothiazide (HCT); an inhibitor of Na+ K+ ATPase, ouabain; an inhibitor of non-selective cation channel, flufenamide; an inhibitor of stretch-activated ion channel, Gad (III) chloride hexahydrate; an inhibitor of K+ channel, tetraethyl ammonium chloride (TEA); and an inhibitor of Na+/K+ cotransporter, bumetanide were purchased from Sigma-Aldrich. Na+-free media was made by substituting NaCl with N-methyl-D-glucamine (Sigma-Aldrich) from MEM media or Hank’s balanced salt solution (HBSS). Cl medium was made by substituting KCl, KH2PO4 and CaCl2 with sodium gluconate, potassium gluconate and calcium acetate hydrate (Sigma-Aldrich). Anti-histones (H1, H2A, H2B, H3 and H4), anti-ERp57, anti-IL6, anti-lamin A/C and anti-hemagglutinin tag (HA) antibodies were purchased from Santa Cruz Biotechnology. Anti-Hsp60, anti-Hsp90, anti-PARP, anti-caspase 3, anti-caspase 6, anti-caspase 7, anti-caspase 9, anti-ICAD, anti-calpastatin, anti-calpain 1 and anti-calpain 2 antibodies were purchased from Cell Signaling Technology; anti-HMGB1 antibody from Abcam (Cambridge, MA, USA); and anti-α-tubulin antibody from Calbiochem. Anti-CAD antibody was purchased from Novus Biologicals (Littleton, CO, USA). Anti-calpain S1 antibody was purchased from Thermo Scientific (Seoul, Korea). Anti-Flag tag and anti-HA tag antibodies were purchased from Sigma-Aldrich.

Expression constructs and lentiviral transfections

Lentiviral constructs expressing shRNAs for caspases 1, 3, 6, 7, 9, CAD, EndoG, calpastatin, calpain S1, PAD2, PAD4, Atg5, Vps34 and Beclin1 were purchased from Sigma-Aldrich. cDNAs of caspase 3, CAD, EndoG, calpain 1 and calpain 2 were purchased from OriGene (Rockville, MD, USA), calpastatin cDNA was from Shi-Yong Sun (Emory University School of Medicine), being subcloned into pCDH-EF2-MCS-T2A-Puro, a lentiviral vector for cDNA expression (System Biosciences, Mountain View, CA, USA). Mutation of calpastatin at cleavage sites of caspases was accomplished using a QuickChange site-directed mutagenesis kit (Stratagene, Santa Clara, CA, USA). All the lentiviral vectors were transfected to 293TN cells (System Biosciences) with Lipofectamine 2000 transfection reagent (Invitrogen, Seoul, Korea). Particles were collected 2 days after the transfection of lentiviral plasmids, and infected into the cells. Lentivirus-infected cells were puromycin-selected for 1 week.

Polymerase chain reaction (PCR)

The released DNAs and cellular genomic DNAs were PCR amplified using primers for genomic sequences; GAPDH (5′-IndexTermCCCCTTCATTGACCTCAACTAC-3′ and 5′-IndexTermGAGTCCTTCCACGATACCAAAG-3′), FAS (5′-IndexTermTCACCACTATTGCTGGAGTCAT-3′ and 5′-IndexTermTAAACATCCTTGGAGGCAGAAT-3′), and the released DNAs and mitochondrial DNAs were PCR-amplified using primers for mitochondrial genes; ATP synthase subunit 6 (ATP6) (5′-IndexTermATACACAACACTAAAGGACGAACCT-3′ and 5′-IndexTermGAGGCTTACTAGAAGTGTGAAAACG-3′), cytochrome oxidase c subunit 1 (CO1) (5′-IndexTermGGAGTCCTAGGCACAGCTCTAA-3′ and 5′-IndexTermGGAGGGTAGACTGTTCAACCTG-3′) for determining the presence of genomic and mitochondrial gene sequences in the released DNAs of dying cancer cells.56

Preparation of cell lysates and western blots

The conditioned medium from HBSS-incubated or staurosporine-treated cells were 100-fold concentrated with Amicon Ultra Centrifugal Filters (Millipore, Darmstadt, Germany; 3000 Da MW cut-off). For preparing total cell lysates, cells were lysed in high salt lysis buffer (50 mM HEPES (pH 7.5), 250 mM NaCl, 1% Triton X-100, 1 mM EDTA, 1 mM dithiothreitol, 1 mM Na3VO4, 1 mM NaF, 1 μg/ml pepstatin A, 10 μg/ml AEBSF, 2 μg/ml aprotinin and 1 μg/ml leupeptin), incubated on ice for 20 min and centrifuged for 20 min to remove cell debris. The concentrated conditioned media or total cell lysate was subjected to sodium dodecyl sulfate-polyacrylamide electrophoresis. The proteins were then electro-transferred to PVDF membranes and incubated overnight with antibodies at 4 °C. Subsequently, the membranes were incubated with peroxidase-conjugated secondary antibodies (Pierce, Rockford, IL, USA) for 1 h at room temperature, and the signal was detected using an enhanced chemiluminescence (ECL) detection kit (Amersham Biosciences, Seongnam, Korea).

Quantification of released DNAs

Cells were cultured in six-well plates for >24 h to 80% confluency and treated with various cytotoxic reagents for the indicated time periods. Released DNA from dying cells were digested with 500 mU/ml MNase (Thermo Scientific) for 5 min. Nuclease activity was stopped with 5 mM EDTA and the culture supernatants were collected and stored at −20 °C until quantification. Total genomic DNA from HeLa cells was extracted with 500 μl DNazol (Molecular Research Center, Cincinnati, OH, USA). Total DNA and released DNA were quantified using a PicoGreen dsDNA assay kit (Life Technologies, Seoul, Korea) according to the manufacturer’s instructions. Data are presented as % DNA release calculated as (released DNA/total genomic DNA) × 100.57

Live cell fluorescence imaging

Cells on Lab-Tek two-well glass chamber slide (NUNC, Rockford, IL, USA) were incubated in the presence of membrane-impermeable DNA dye, SYTOX Green (50 nM; Life Technologies), and membrane-permeable DNA dye DRAQ5 (5 μM; Cell Signaling Technology). The images were acquired every 5 min for 24 h via a 100 × objective on a Deltavision RT Deconcolution microscope (Applied Precision, Issaquah, WA, USA) using a Photometrics Cool SNAPHQ2 camera (Roper-Princeton Instruments, Trenton, NJ, USA) controlled by SoftWoRxTM Imaging workstation (Applied Precision).

Confocal microscopy

Cells grown on Lab-Tek four-well glass chamber slides (NUNC) were incubated in HBSS or medium containing appropriate reagents for the indicated times. Cells were fixed with 4% paraformaldehyde and permeabilized with 0.2% Triton X-100 for 5 min. They were washed with PBS and incubated with primary antibodies and subsequently with secondary antibody conjugates (Alexa Fluor 594 donkey anti-mouse IgG and/or Alexa Fluor 488 donkey anti-rabbit IgG; Invitrogen). Images were collected using a laser scanning confocal microscope LSM710 (Carl Zeiss, Oberkochen, Germany) equipped with argon (488 nm) and krypton (568 nm) lasers, using an x40 water immersion objective. Images were processed with ZEN 2009 light edition (Carl Zeiss).

Transmission electron microscopy

The cells were pelleted and washed twice with PBS. Fixation was performed with phosphate buffer pH 7.4 containing 2.5% glutaraldehyde for 30 min at 4 °C. The pellets were rinsed twice with cold PBS, post-fixed in buffered OsO4, dehydrated in graded acetone and embedded in Durcupan ACM resin (Fluka, Yongin, Korea). Ultrathin sections were obtained, mounted in copper grids and counterstained with uranyl acetate and lead citrate. The specimens were observed with a Hitachi H-7600 TEM (Schaumburg, IL, USA) at 80 kV.

Assays for cellular viability and cell death, TUNEL staining, Annexin V staining and JC-1 staining

Cellular viability was quantified with Calcein-AM (Invitrogen) as specified by the manufacturer. The cells were aliquoted into 96-black well plates, incubated with Calcein-AM (2.5 μM) for 15 min and fluorescent intensities were measured at excitation and emission wavelength of 490 nm and 520 nm, respectively. Data were the mean±S.D. of four independent measurements, and are presented as % viability ((fluorescence intensity of treated cells/fluorescent intensity of non-treated cells) × 100). Assays for determining cell death were done using cell-impermeable DNA dyes, PI (Calbiochem) or SYTOX Green (Life Technologies). Cells in triplicate were washed, stained briefly with PI (500 nM) or SYTOX Green (30 nM), and fluorescence intensities were analyzed by flow cytometry, and are presented as % cell death ((dead cellular counts/total cellular counts) × 100). Intracellular DNA fragmentation was detected using APO-BrdU TUNEL assay kit (Invitrogen) as specified by the manufacturer. Suspensions of dying cells were sequentially fixed by adding 1% paraformaldehyde in PBS for 15 min and 70% ethanol for 30 min in ice, washed twice, and DNA-labeled with TdT enzyme and BrdUTP for 60 min at 37 °C. The cells were stained with Alexa Fluor 488-conjugated anti-BrdU antibody for 30 min at room temperature and with PI/RNase A for 30 min at room temperature, and finally analyzed by flow cytometry. Mitochondrial membrane potential was measured by staining of a membrane-permeable JC-1 dye (MACS Miltenyi Biotech, Bergisch Gladbach, Germany) and analysis by flow cytometry. Annexin V staining was performed for detecting the exposure of phosphatidylserine to outer leaflet of plasma membrane that is a marker of early apoptosis using a Biotin Annexin V staining kit (BD Biosciences, San Jose, CA, USA) according to the manufacturer’s protocol. Annexin V-positive population was defined when Annexin V staining was positive and PI staining was negative in the population by flow cytometry.

ChIP assay

The assay was performed using EZ-Magna ChIP A/G assay kit (Millipore) according to the manufacturer’s instructions with minor modifications. Briefly, HeLa cells were treated with 1 μg/ml staurosporine for 10 h, with 32 mM H2O2 for 4 h, or heated at 65 °C for 1 h and incubated for 3 h. The conditioned media were fixed with 1% formaldehyde for 10 min, quenched with 1X glycine for 5 min and concentrated. The concentrated media were incubated overnight with goat polyclonal anti-histone H1 antibody (Santa Cruz Biotechnology), goat polyclonal anti-HMGB1 antibody (Santa Cruz Biotechnology) or normal goat serum. The antibody–protein–DNA complexes were captured by incubation with EZ-Magna ChIP A/G. Genomic DNA segments from complexes were eluted, digested with proteinase K digestion and purified using spin columns. Aliquots of the DNA were used as a template for PCR amplification using 35 cycles of 55 °C annealing temperature. The GAPDH primers for PCR amplification were 5′-IndexTermCCCCTTCATTGACCTCAACTAC-3′ and 5′-IndexTermGAGTCCTTCCACGATACCAAAG-3′.

Caspase activity assay

Activities of caspases 3/7 and 9 were measured by Caspase-Glo 3/7 Assay and Caspase-Glo 9 Assay (Promega, Seoul, Korea) according to manufacturer’s instructions.

Calpain assay

Calpain activity was measured using Calpain Activity Fluorometric Assay Kit (Biovision, San Francisco, CA, USA) according to the manufacturer’s instructions.

DNA laddering analysis

The cells were pelleted, washed twice and DNA extracted with DNazol (Molecular Research Center). DNA was precipitated with two volumes of ethanol and 0.1 volumes of 3 M sodium acetate, pH 5.2. DNA released from dying cells was concentrated, purified by phenol–chloroform extraction and precipitated with two volume of ethanol and 0.1 volumes of 3 M sodium acetate, pH 5.2. The precipitated DNA was resuspended in TE buffer pH 8.0 containing DNase-free RNase, and analyzed using 1.5% agarose gel in 1 × TBE buffer.

Measurements of intracellular chloride ion concentration

Fluorescence-based microplate assay applying intracellular quenching of N-(ethoxycarbonylmethyl)-6-methoxyquinolinium bromide (MQAE) fluorescence was used for quantifying intracellular Cl concentrations as described previously.58 HeLa cells loaded with 5 mM MQAE (Life Technologies) overnight were incubated with or without staurosporine (1 μg/ml) for the indicated times. One set of the cells was washed three times with Cl containing buffer (HBSS) and MQAE fluorescence was measured at excitation wavelength of 360 nm and an emission wavelength of 460 nm (Ft), and the other set of the cells were washed three times with Cl-free HBSS, suspended with Cl-free HBSS containing 10 μM tributyltin chloride (Sigma-Aldrich) and 5 μM nigericin sodium salt (Sigma-Aldrich). MQAE fluorescence was measured (F0). F0/Ft values were used as representative values for intracellular Cl concentrations according to Stern–Volmer equation (F0/Ft=1+KCl [Cl], where [Cl] is the intracellular Cl concentrations and KCl is the Stern–Volmer constant).

Measurements of chloride ion currents using halide-sensitive YFP

Cellular Cl currents were measured using Premo Halide Sensor (Invitrogen) as specified by the manufacturer. Briefly, HeLa cells infected with baculovirus encoding halide-sensitive yellow fluorescent protein (YFP) were incubated with various reagents for indicated times, mixed with an equal volume of 2X Premo halide stimulus buffer containing 150 mM NaI and the fluorescence intensities were monitored every 2 s for 60 s at an excitation wavelength of 480 nm and an emission wavelength of 560 nm to record YFP quenching evoked by inward currents of iodide ion.

Measurements of intracellular free calcium ion

Intracellular free Ca2+ was measured by Calcium Sensor Dye eFluor 514 (eBiosciences, San Diego, CA, USA). Briefly, cells were incubated for 30 min at 37 °C in medium containing 5 μM eFluor 514 and washed twice. Fluorescence was measured at an excitation wavelength of 490 nm and an emission wavelength of 514 nm with a FLUOstar Optima Microplate Fluorometer (BMG Labtech, Cary, NC, USA). Data were presented as relative fluorescence; MFI of treated cells/MFI of non-treated cells.

Real-time PCR

Total RNA was isolated using an RNeasy kit (Qiagen, Seoul, Korea). PrimeScript RT reagent Kit (TaKaRa, Seoul, Korea) was used to reverse transcribe mRNA into cDNA. PCR was then performed on an ABI PRISM 7000 machine (Applied Biosystems, Carlsbad, CA, USA) using SYBR Premix Ex Taq II (TaKaRa). The sequences of primers for PAD2 and PAD4 were: PAD2 (5′-IndexTermGTGACAACCCTCGGTGTGGA-3′ and 5′-IndexTermACATCAAGGTGGAAGCAGGAACTTA-3′), PAD4 (5′-IndexTermGAGGCTGTGGTGTTCCAAGA-3′ and 5′-IndexTermTCAGCTTGCACTTGGCTTTC-3′). Analysis of each sample was performed more than twice for each experiment, and data in the figures are reported as relative quantification: average values of 2−ΔΔCT±S.D.

Statistical analysis

All the values are presented as mean±S.E. A paired Student's t-test was used to identify significant differences in comparisons. A level of P<0.05 was considered statistically significant.

Abbreviations

3-methyl adenine

damage-associated molecular pattern

caspase-activated DNase

4,4′-Diisothiocyanatostilbene-2,2′-disulfonic acid

endonuclease G

high mobility group protein B1

heat shock protein 90

inhibitor of caspase-activated DNase

interleukin 6

minimal essential media

micrococcal nuclease

N-acetyl cysteine

2-(3-phenylpropylamino) benzoic acid

peptidylarginine deaminase

poly ADP ribose polymerase-1

short hairpin RNA

References

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Apoptosis caspase dependent

Apoptosis Dependent and Independent Functions of Caspases

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Introduction to Apoptosis - The Caspase Enzymes Part 1

Caspase

Family of cysteine proteases

Caspases (cysteine-aspartic proteases, cysteine aspartases or cysteine-dependent aspartate-directed proteases) are a family of protease enzymes playing essential roles in programmed cell death. They are named caspases due to their specific cysteine protease activity – a cysteine in its active site nucleophilically attacks and cleaves a target protein only after an aspartic acid residue. As of 2009, there are 12 confirmed caspases in humans[note 1] and 10 in mice, carrying out a variety of cellular functions.

The role of these enzymes in programmed cell death was first identified in 1993, with their functions in apoptosis well characterised. This is a form of programmed cell death, occurring widely during development, and throughout life to maintain cell homeostasis. Activation of caspases ensures that the cellular components are degraded in a controlled manner, carrying out cell death with minimal effect on surrounding tissues.[3]

Caspases have other identified roles in programmed cell death such as pyroptosis and necroptosis. These forms of cell death are important for protecting an organism from stress signals and pathogenic attack. Caspases also have a role in inflammation, whereby it directly processes pro-inflammatory cytokines such as pro-IL1β. These are signalling molecules that allow recruitment of immune cells to an infected cell or tissue. There are other identified roles of caspases such as cell proliferation, tumour suppression, cell differentiation, neural development and axon guidance and ageing.[4]

Caspase deficiency has been identified as a cause of tumour development. Tumour growth can occur by a combination of factors, including a mutation in a cell cycle gene which removes the restraints on cell growth, combined with mutations in apoptopic proteins such as caspases that would respond by inducing cell death in abnormally growing cells.[5] Conversely, over-activation of some caspases such as caspase-3 can lead to excessive programmed cell death. This is seen in several neurodegenerative diseases where neural cells are lost, such as Alzheimer's disease.[5] Caspases involved with processing inflammatory signals are also implicated in disease. Insufficient activation of these caspases can increase an organism's susceptibility to infection, as an appropriate immune response may not be activated.[5] The integral role caspases play in cell death and disease has led to research on using caspases as a drug target. For example, inflammatory caspase-1 has been implicated in causing autoimmune diseases; drugs blocking the activation of Caspase-1 have been used to improve the health of patients. Additionally, scientists have used caspases as cancer therapy to kill unwanted cells in tumours.[6]

Functional classification of caspases[edit]

Most caspases play a role in programmed cell death. These are summarized in the table below. The enzymes are sub classified into three types: Initiator, Executioner and Inflammatory.[7]

Note that in addition to apoptosis, caspase-8 is also required for the inhibition of another form of programmed cell death called necroptosis. Caspase-14 plays a role in epithelial cell keratinocyte differentiation and can form an epidermal barrier that protects against dehydration and UVB radiation.[11]

Activation of caspases[edit]

Caspases are synthesised as inactive zymogens (pro-caspases) that are only activated following an appropriate stimulus. This post-translational level of control allows rapid and tight regulation of the enzyme.

Activation involves dimerization and often oligomerisation of pro-caspases, followed by cleavage into a small subunit and large subunit. The large and small subunit associate with each other to form an active heterodimer caspase. The active enzyme often exists as a heterotetramer in the biological environment, where a pro-caspase dimer is cleaved together to form a heterotetramer.[12]

Dimerisation[edit]

The activation of initiator caspases and inflammatory caspases is initiated by dimerisation, which is facilitated by binding to adaptor proteins via protein–protein interaction motifs that are collectively referred to as death folds. The death folds are located in a structural domain of the caspases known as the pro-domain, which is larger in those caspases that contain death folds than in those that do not. The pro-domain of the intrinsic initiator caspases and the inflammatory caspases contains a single death fold known as caspase recruitment domain (CARD), while the pro-domain of the extrinsic initiator caspases contains two death folds known as death effector domains (DED).[13][14]

Multiprotein complexes often form during caspase activation.[12] Some activating multiprotein complexes includes:

Cleavage[edit]

Once appropriately dimerised, the Caspases cleave at inter domain linker regions, forming a large and small subunit. This cleavage allows the active-site loops to take up a conformation favourable for enzymatic activity.[15]

Cleavage of Initiator and Executioner caspases occur by different methods outlined in the table below.

  • Initiator caspases auto-proteolytically cleave whereas Executioner caspases are cleaved by initiator caspases. This hierarchy allows an amplifying chain reaction or cascade for degrading cellular components, during controlled cell death.
Initiator Caspase

Caspase-8

Inititator Pro-caspases have a prodomain that allows recruitment of other pro-caspases, which subsequently dimerise. Both pro-caspase moleuceules undergo cleavage by autocatalysis. This leads to removal of the prodomain and cleavage of the linker region between the large and small subunit. A heterotetramer is formed
PDB image of caspase 8 (3KJQ) in 'biological assembly'. Two shades of blue used to represent two small sunits, while two shades of purple represent two large subunits
Executioner

Caspase Caspase-3

Executioner caspase constitutively exist as homodimers. The red cuts represent regions where initiator caspases cleave the executioner caspases. The resulting small and large subunit of each Caspase-3 will associate, resulting in a heterotetramer.
[16]
PDB image of Caspase 3 (4QTX) in 'biological assembly'. Two shades of blue used to represent two small sunits, while two shades of purple represent two large subunits

Some roles of caspases[edit]

Apoptosis[edit]

Initiator caspases are activated by intrinsic and extrinsic apoptopic pathways. This leads to the activation of other caspases including executioner caspases that carry out apoptosis by cleaving cellular components.

Apoptosis is a form of programmed cell death where the cell undergoes morphological changes, to minimize its effect on surrounding cells to avoid inducing an immune response. The cell shrinks and condenses - the cytoskeleton will collapse, the nuclear envelope disassembles and the DNA fragments up. This results in the cell forming self-enclosed bodies called 'blebs', to avoid release of cellular components into the extracellular medium. Additionally, the cell membrane phospholipid content is altered, which makes the dying cell more susceptible to phagocytic attack and removal.[17]

Apoptopic caspases are subcategorised as:

  1. Initiator Caspases (Caspase 2, Caspase 8, Caspase 9, Caspase 10)
  2. Executioner Caspases (Caspase 3, Caspase 6 and Caspase 7)

Once initiator caspases are activated, they produce a chain reaction, activating several other executioner caspases. Executioner caspases degrade over 600 cellular components[18] in order to induce the morphological changes for apoptosis.

Examples of caspase cascade during apoptosis:

  1. Intrinsic apoptopic pathway: During times of cellular stress, mitochondrial cytochrome c is released into the cytosol. This molecule binds an adaptor protein (APAF-1), which recruits initiator Caspase-9 (via CARD-CARD interactions). This leads to the formation of a Caspase activating multiprotein complex called the Apoptosome. Once activated, initiator caspases such as Caspase 9 will cleave and activate other executioner caspases. This leads to degradation of cellular components for apoptosis.
  2. Extrinsic apoptopic pathway: The caspase cascade is also activated by extracellular ligands, via cell surface Death Receptors. This is done by the formation of a multiprotein Death Inducing Signalling Complex (DISC) that recruits and activates a pro-caspase. For example, the Fas Ligand binds the FasR receptor at the receptor's extracellular surface; this activates the death domains at the cytoplasmic tail of the receptor. The adaptor protein FADD will recruit (by a Death domain-Death domain interaction) pro-Caspase 8 via the DED domain. This FasR, FADD and pro-Caspase 8 form the Death Inducing Signalling Complex (DISC) where Caspase-8 is activated. This could lead to either downstream activation of the intrinsic pathway by inducing mitochondrial stress, or direct activation of Executioner Caspases (Caspase 3, Caspase 6 and Caspase 7) to degrade cellular components as shown in the adjacent diagram.[19]

Pyroptosis[edit]

Pyroptosis is a form of programmed cell death that inherently induces an immune response. It is morphologically distinct from other types of cell death – cells swell up, rupture and release pro-inflammatory cellular contents. This is done in response to a range of stimuli including microbial infections as well as heart attacks (myocardial infarctions).[20] Caspase-1, Caspase-4 and Caspase-5 in humans, and Caspase-1 and Caspase-11 in mice play important roles in inducing cell death by pyroptosis. This limits the life and proliferation time of intracellular and extracellular pathogens.[citation needed]

Pyroptosis by caspase-1[edit]

Caspase-1 activation is mediated by a repertoire of proteins, allowing detection of a range of pathogenic ligands. Some mediators of Caspase-1 activation are: NOD-like Leucine Rich Repeats (NLRs), AIM2-Like Receptors (ALRs), Pyrin and IFI16.[21]

These proteins allow caspase-1 activation by forming a multiprotein activating complex called Inflammasomes. For example, a NOD Like Leucine Rich Repeat NLRP3 will sense an efflux of potassium ions from the cell. This cellular ion imbalance leads to oligomerisation of NLRP3 molecules to form a multiprotein complex called the NLRP3 Inflammasome. The pro-caspase-1 is brought into close proximity with other pro-caspase molecule in order to dimerise and undergo auto-proteolytic cleavage.[21]

Some pathogenic signals that lead to Pyroptosis by Caspase-1 are listed below:

  • DNA in the host cytosol binds to AIM2-Like Receptors inducing Pyroptosis
  • Type III secretion system apparatus from bacteria bind NOD Like Leucine Rich Repeats receptors called NAIP's (1 in humans and 4 in mice)

Pyroptosis by Caspase-4 and Caspase-5 in humans and Caspase-11 in mice

These caspases have the ability to induce direct pyroptosis when lipopolysaccharide (LPS) molecules (found in the cell wall of gram negative bacteria) are found in the cytoplasm of the host cell. For example, Caspase 4 acts as a receptor and is proteolytically activated, without the need of an inflammasome complex or Caspase-1 activation.[21]

A crucial downstream substrate for pyroptopic caspases is Gasdermin D (GSDMD)[22]

Role in inflammation[edit]

Inflammation is a protective attempt by an organism to restore a homeostatic state, following disruption from harmful stimulus, such as tissue damage or bacterial infection.[18]

Caspase-1, Caspase-4, Caspase-5 and Caspase-11 are considered 'Inflammatory Caspases'.[7]

  • Caspase-1 is key in activating pro-inflammatory cytokines; these act as signals to immune cells and make the environment favourable for immune cell recruitment to the site of damage. Caspase-1 therefore plays a fundamental role in the innate immune system. The enzyme is responsible for processing cytokines such as pro-ILβ and pro-IL18, as well as secreting them.[21]
  • Caspase-4 and -5 in humans, and Caspase-11in mice have a unique role as a receptor, whereby it binds to LPS, a molecule abundant in gram negative bacteria. This can lead to the processing and secretion of IL-1β and IL-18 cytokines by activating Caspase-1; this downstream effect is the same as described above. It also leads to the secretion of another inflammatory cytokine that is not processed. This is called pro-IL1α.[21] There is also evidence of an inflammatory caspase, caspase-11 aiding cytokine secretion; this is done by inactivating a membrane channel that blocks IL-1β secretion[21]
  • Caspases can also induce an inflammatory response on a transcriptional level. There is evidence where it promotes transcription of nuclear factor-κB (NF-κB), a transcription factor that assists in transcribing inflammatory cytokines such as IFNs, TNF, IL-6 and IL-8. For example, Caspase-1 activates Caspase-7, which in turn cleaves the poly (ADP) ribose – this activates transcription of NF-κB controlled genes.[18]

Discovery of caspases[edit]

H. Robert Horvitz initially established the importance of caspases in apoptosis and found that the ced-3 gene is required for the cell death that took place during the development of the nematodeC. elegans. Horvitz and his colleague Junying Yuan found in 1993 that the protein encoded by the ced-3 gene is cysteine protease with similar properties to the mammalian interleukin-1-beta converting enzyme (ICE) (now known as caspase 1). At the time, ICE was the only known caspase.[23] Other mammalian caspases were subsequently identified, in addition to caspases in organisms such as fruit fly Drosophila melanogaster.

Researchers decided upon the nomenclature of the caspase in 1996. In many instances, a particular caspase had been identified simultaneously by more than one laboratory; each would then give the protein a different name. For example, caspase 3 was variously known as CPP32, apopain and Yama. Caspases, therefore, were numbered in the order in which they were identified.[24] ICE was, therefore, renamed as caspase 1. ICE was the first mammalian caspase to be characterised because of its similarity to the nematode death gene ced-3, but it appears that the principal role of this enzyme is to mediate inflammation rather than cell death.

Evolution[edit]

In animals apoptosis is induced by caspases and in fungi and plants, apoptosis is induced by arginine and lysine-specific caspase like proteases called metacaspases. Homology searches revealed a close homology between caspases and the caspase-like proteins of Reticulomyxa (a unicellular organism). The phylogenetic study indicates that divergence of caspase and metacaspase sequences occurred before the divergence of eukaryotes.[25]

See also[edit]

Notes[edit]

  1. ^ abFunctional CASP12 is only expressed in some individuals of African descent, while individuals of Asian or Caucasian descent express only a non-functional truncated form.[2]
  2. ^ abcCASP4 and CASP5 are considered to be the human orthologues of CASP11, which was found in mice and rats but not in humans.[9]

References[edit]

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  13. ^Lahm, Armin; Paradisi, Andrea; Green, Douglas R; Melino, Gerry (2003). "Death fold domain interaction in apoptosis". Cell Death and Differentiation. 10 (1): 10–2. doi:10.1038/sj.cdd.4401203. PMID 12655289. S2CID 32593733.
  14. ^Kumar, S (2006). "Caspase function in programmed cell death". Cell Death and Differentiation. 14 (1): 32–43. doi:10.1038/sj.cdd.4402060. PMID 17082813.
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  16. ^Lavrik, I.; Krueger, A.; Schmitz, I.; Baumann, S.; Weyd, H.; Krammer, P. H.; Kirchhoff, S. (2003-01-01). "The active caspase-8 heterotetramer is formed at the CD95 DISC". Cell Death & Differentiation. 10 (1): 144–145. doi:10.1038/sj.cdd.4401156. ISSN 1350-9047. PMID 12655304.
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  21. ^ abcdefEldridge, Matthew JG; Shenoy, Avinash R (2015). "Antimicrobial inflammasomes: unified signalling against diverse bacterial pathogens". Current Opinion in Microbiology. 23: 32–41. doi:10.1016/j.mib.2014.10.008. PMID 25461570.
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  25. ^Klim, Joanna; Gładki, Arkadiusz; Kucharczyk, Roza; Zielenkiewicz, Urszula; Kaczanowski, Szymon (2018-04-27). "Ancestral State Reconstruction of the Apoptosis Machinery in the Common Ancestor of Eukaryotes". G3: Genes, Genomes, Genetics. 8 (6): 2121–2134. doi:10.1534/g3.118.200295. ISSN 2160-1836. PMC 5982838. PMID 29703784.

External links[edit]

Sours: https://en.wikipedia.org/wiki/Caspase

You will also be interested:

Open Access

Peer-reviewed

  • Gang Liu ,
  • Hui Zou ,
  • Tongwang Luo ,
  • Mengfei Long,
  • Jianchun Bian,
  • Xuezhong Liu,
  • Jianhong Gu,
  • Yan Yuan,
  • Ruilong Song,
  • Yi Wang,
  • Jiaqiao Zhu ,
  • Zongping Liu
  • Gang Liu, 
  • Hui Zou, 
  • Tongwang Luo, 
  • Mengfei Long, 
  • Jianchun Bian, 
  • Xuezhong Liu, 
  • Jianhong Gu, 
  • Yan Yuan, 
  • Ruilong Song, 
  • Yi Wang
PLOS

x

Abstract

We designed this study to investigate whether cadmium induces caspase-independent apoptosis and to investigate the relationship between the caspase-dependent and caspase-independent apoptotic pathways. Cadmium (1.25–2.5 μM) induced oxidative stress in rat proximal tubular (rPT) cells, as seen in the reactive oxygen species levels; N-acetylcysteine prevented this. Cyclosporin A (CsA) prevented mitochondrial permeability transition pore opening and apoptosis; there was mitochondrial ultrastructural disruption, mitochondrial cytochrome c (cyt c) translocation to the cytoplasm, and subsequent caspase-9 and caspase-3 activation. Z-VAD-FMK prevented caspase-3 activation and apoptosis and decreased BNIP-3 (Bcl-2/adenovirus E1B 19-kDa interacting protein 3) expression levels and apoptosis-inducing factor/endonuclease G (AIF/Endo G) translocation. Simultaneously, cadmium induced prominent BNIP-3 expression in the mitochondria and cytoplasmic AIF/Endo G translocation to the nucleus. BNIP-3 silencing significantly prevented AIF and Endo G translocation and decreased the apoptosis rate, cyt c release, and caspase-9 and caspase-3 activation. These results suggest that BNIP-3 is involved in the caspase-independent apoptotic pathway and is located upstream of AIF/Endo G; both the caspase-dependent and caspase-independent pathways are involved in cadmium-induced rPT cell apoptosis and act synergistically.

Citation: Liu G, Zou H, Luo T, Long M, Bian J, Liu X, et al. (2016) Caspase-Dependent and Caspase-Independent Pathways Are Involved in Cadmium-Induced Apoptosis in Primary Rat Proximal Tubular Cell Culture. PLoS ONE 11(11): e0166823. https://doi.org/10.1371/journal.pone.0166823

Editor: Saeid Ghavami, University of Manitoba, CANADA

Received: July 26, 2016; Accepted: November 5, 2016; Published: November 18, 2016

This is an open access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.

Data Availability: All relevant data are within the paper and its Supporting Information files.

Funding: This work was supported by the National Natural Science Foundation of China (No. 31101866, No. 31372495, No. 31502128 and No. 31302058), Jiangsu Provincial Natural Science Foundation of China (BK20150447) and a Project Funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD). We state that the funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Abbreviations: rPT, rat proximal tubular; MPTP, mitochondrial potential translation pore; siRNA, small interfering RNA; BCA, bicinchonininc acid; PI, propidium iodide; ΔΨ, mitochondrial membrane potential

1. Introduction

Cadmium is gaining attention as a known occupational hazard and environmental pollutant that can cause a series of biochemical and physiological dysfunctions in humans. The exposure routes have principally been contact with batteries, paints, fertilizers, and automobiles. As with other complex organic pollutants, microorganisms cannot degrade cadmium. Cadmium accumulates in the ecosystem and enters the food chain through contaminated water and soil and has an extremely long biological half-life. As a multi-organ toxicant, cadmium exerts toxic effects on the brain, liver, kidney, heart, and bone [1]. The kidney is the primary site for the initial accumulation of cadmium, and the proximal tubule cells are sensitive to cadmium-induced damage [2].

The mitochondria play a central role in regulating apoptotic cell death. Numerous pro-apoptotic factors and damage pathways act on the mitochondria to induce oxidative stress, and reactive oxygen species (ROS) overproduction can directly result in mitochondrial permeability transition pore (MPTP) opening, followed by mitochondrial release of apoptogenic signaling molecules, such as procaspases, cytochrome c (cyt c), apoptosis-inducing factor (AIF), and endonuclease G (Endo G) [3, 4]. Cadmium-induced apoptosis occurs mostly via activation of the mitochondrial apoptotic pathways [5, 6].

The apoptogenic potential of cadmium on cells and primary rat kidney cell culture has been reported [7–10]. Previously, we showed that lead induces oxidative stress in rat proximal tubular (rPT) cells and resulted in apoptosis through MPTP opening [11]. ROS enhancement in murine splenocytes and thymocytes induces mitochondrial membrane depolarization, which leads to caspase-3 activation and DNA fragmentation [12, 13].

Many studies have also focused on the caspase-independent apoptotic pathway, known as the AIF/Endo G pathway. Caspase-independent apoptosis is activated by BNIP-3 (Bcl-2/adenovirus E1B 19-kDa interacting protein 3), which induces mitochondrial AIF release; Endo G acts as a modulator. Forced BNIP-3 expression by plasmid transfection results in mitochondrial Endo G release and nuclear translocation [14]. BH3 domain of BNIP-3 interacted with anti-apoptotic protein to form dimers, which was able to promote the apoptosis and the homodimerization of TM domain also promoted apoptosis. The investigation confirmed that homodimerization of BNIP-3’s TM domain involved in mitochondria apoptosis pathway [15]. While there was no evidence for homodimerization of TM domain involved in caspase-independent apoptosis pathway. Overexpression of BNIP-3, an upstream effector of AIF, induces MPT and cyt c release; BNIP-3 silencing by short hairpin RNA (shRNA) increases mitochondrial cyt c levels and blocks the caspase-dependent apoptotic pathway [16]. BNIP-3 located in different positions in cells. According to studies, BNIP3 was involved in promoting apoptosis mainly engaged in mitochondria, it could bind to mitochondria and make the mitochondrial dysfunction. While, BNIP3 bound to the promoter of the AIF gene and represses its expression when it translocated to nuclei. BNIP3-mediated reduction in AIF expression leads to decreased temozolomide-induced apoptosis in glioma cells and transcriptional repression function for BNIP3 causing reduced AIF expression and increased resistance to apoptosis [17]. BNIP-3 also involved in autophagy induction. BNIP-3's transmembrane domain that preserve mitochondrial localization, but disrupt dimerization fail to induce autophagy [18]. BNIP-3 dimerization is thought to free Beclin-1 from its interaction with anti-apoptotic Bcl-2 family proteins, then to cause autophagy [19]. Although the caspase-dependent and caspase-independent apoptotic pathways are separate, there is evidence of crosstalk between the two [20]. Furthermore, caspase inhibitors such as Z-VAD-FMK prevent mitochondrial AIF release [20–23].

We aimed to identify the role of the caspase-dependent and caspase-independent pathways in cadmium-induced apoptosis and the relationship between the two in rPT cells. We found that both pathways are involved in cadmium-induced rPT cell apoptosis and affect each other.

2. Materials and Methods

2.1. Animals and treatment

The Sprague-Dawley rats weighing between 180 g and 200 g were obtained from the Comparative Medicine Centre of Yangzhou University (Yangzhou, China). The animals were housed individually on a 12 h light/dark cycle with unlimited standard rat food and double distilled water (DDW). All experimental procedures were conducted in accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Research Council and were approved by the Animal Care and Use Committee of Yangzhou University (Approval ID: SYXK (Su) 2007–0005). All surgeries operations were performed under sodium pentobarbital anesthesia, and all efforts were made to minimize any suffering experienced by the animals used in this study.

2.2. Reagents

All chemicals were of the highest-grade purity available. Dulbecco’s modified Eagle’s medium (DMEM)-F12 (1:1), fetal bovine serum (FBS), trypsin-EDTA, and collagenase IV were from Gibco (Grand Island, NY, USA). Cadmium acetate, calcein acetoxymethyl ester (calcein-AM), cobalt chloride (CoCl2), 2’, 7’-dichlorofluorescein diacetate (DCFH-DA), and DAPI were from Sigma-Aldrich (St. Louis, MO, USA). Cell Counting Kit-8 (CCK-8) was from Dojindo Laboratories (Tokyo, Japan). The mitochondria isolation kit for cultured cells was from Pierce Biotechnology (Rockford, IL, USA).

The following primary antibodies were used: anti–cyt c (CST, #14940), anti–cyt c oxidase subunit IV (COX IV) (CST, #11967), anti–cleaved caspase-9 (CST, #9506), anti–cleaved caspase-3 (CST, #9664), anti–β-actin (CST, #4970S) were from Cell Signaling Technology (Boston, USA); anti–BNIP-3 (Abcam, ab 109362), anti-AIF (Abcam, ab1998), anti–Endo G (Abcam, ab9647), and anti–lamin B1 (Abcam, ab16048) were from Abcam (Cambridge, USA). All secondary antibodies were from Beijing Zhongshan Golden Bridge Biotechnology (Beijing, China). The PrimeScript RT reagent kit with gDNA Eraser and SYBR Premix Ex Taq RT-PCR kit were from Takara (Dalian, China). Accutase cell detachment solution and the annexin V–fluorescein isothiocyanate/propidium iodide (FITC/PI) apoptosis detection kit were from Becton-Dickinson (San Diego, CA, USA). All other chemicals were from Sigma-Aldrich.

2.3. Cell culture and cadmium exposure conditions

The rPT cells were obtained from the kidneys of Sprague-Dawley rats (from the Comparative Medicine Centre of Yangzhou University) with body weights between 180 g and 200 g. Intraperitoneal injection of sodium pentobarbital (2%, 0.31 ml/100 g) to anesthetize rats. Breaking the neck to death 5 min latter when the rats were in a deep coma. The rats were transferred to super-clean worktable after 75% alcohol soak for 2 minutes. Then, opened the abdominal cavity and removed kidneys of rats under aseptic conditions. rPT cell isolation, identification, and culture were performed as previously described [24]. Primary cells and subcultures were cultured in DMEM/F12 supplemented with 15% FBS, 0.25 g/L glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin at 37°C in 95% air and 5% CO2. rPT cell identity was confirmed by alkaline phosphatase antibody staining against specific proximal tubular antigens. The purity of the isolated primary rPT cells was >95%; the cells were subcultured using trypsin-EDTA digestion. Cells cultured for 12 h had the highest viability (according to the growth curve, data not shown). Based on the doses in a previous study [25], cells were treated with 1.25, 2.5, or 5.0 μM cadmium; the cadmium acetate stock solution was dissolved in sterile ultrapure water.

2.4. N-acetylcysteine (NAC) and Cyclosporin A (CsA) treatment

rPT cells were seeded in 6-well plates and pretreated 30 min with NAC or CsA before Cd treatment when the cell fusion rate was 60–70%. NAC or CsA were freshly prepared in deionized water and filter-sterilized before use, the pH of the NAC or CsA was adjusted to 7.0.

2.5. DAPI staining

Apoptotic morphological changes in the nuclei were detected by staining with DAPI (4',6-diamidino-2-phenylindole). rPT cells (2 × 105 cells per well) were seeded onto sterile cover slips in 24-well plates. After 12-h treatment with 0, 1.25, 2.5, or 5.0 μM cadmium, the medium was removed. The cells were washed with ice-cold phosphate-buffered saline (PBS), fixed with paraformaldehyde (4% w/v) for 10 min at room temperature, and incubated with DAPI staining solution (50 mM in PBS) for 10 min in the dark. After washing in PBS three times, the cells were viewed under a Leica inverted fluorescence microscope (Wetzlar, GER) at an excitation wavelength of 352 nm. To assess the extent of cadmium-induced apoptosis, 200 cells per experiment were randomly selected and the apoptotic cells therein were counted; each experiment was performed in triplicate.

2.6. Measurement of MPTP activity

MPTP opening in rPT cells was detected using calcein-AM and CoCl2 loading, resulting in mitochondrial localization of calcein fluorescence [26]; these reagents were used to monitor the MPTP activity. rPT cells (2 × 105 cells per well) were seeded onto sterile cover slips in 24-well plates, loaded for 30 min at 37°C with 2 μM calcein-AM, followed by 1-h incubation with 2 mM CoCl2 after 12-h incubation with 2.5 μM cadmium, and then washed twice with PBS. The cover slips were fixed with paraformaldehyde (4% w/v) for 10 min at room temperature and then imaged under a laser scanning confocal microscope (LSM 710; Zeiss, Jena, Germany). The change in fluorescence intensity was measured with excitation at 488 nm and emission at 525 nm.

2.7. Flow cytometry analysis

All subsequent assays were carried out on a Beckman Coulter fluorescence-activated cell sorter (CyAn ADP 7; Brea, CA, USA). rPT cells were seeded in 6-well plates and treated with 0, 1.25, 2.5, or 5.0 μM cadmium for 12 h when the cell fusion rate was 60–70%. Subsequently, the adherent cells were collected with the Accutase cell detachment solution by 5-min centrifugation at 1500 rpm. Each treatment group yielded at least 1.5 × 106 cells, which were washed twice with PBS and incubated with fluorescent dyes for the flow cytometric analysis.

2.8. Detection of apoptosis

Apoptotic cells were evaluated using annexin V–FITC/PI staining. The total apoptotic proportion is presented as the sum of early and late apoptotic cells, which was determined as the percentage of annexin V+/PI- and annexin V+/PI+ cells, respectively. After 12-h staining, the harvested cells were labeled with annexin V–FITC and PI according to the manufacturer’s protocol. FITC and PI fluorescence was characterized using an FL-1 filter (530 nm) and FL-2 filter (585 nm), respectively; 10,000 events were acquired.

2.9. ROS measurement

Intracellular ROS were determined using flow cytometry and DCFH-DA staining. DCFH-DA can be cleaved to form non-fluorescent dichlorofluorescein (DCFH) in the cells and is oxidized to fluorescent dichlorofluorescein (DCF) by ROS. Cells (1.5 × 106) were incubated with 100 μM DCFH-DA at 37°C for 30 min, washed twice with PBS, and the fluorescence intensity (FL-1, 530 nm) of 10,000 cells was measured using a flow cytometer.

2.10. Cell fraction preparation

After 12-h treatment with 0, 1.25, 2.5, or 5.0 μM cadmium, cells were harvested by Accutase™ Cell Detachment Solution and washed twice with PBS. To obtain the mitochondrial and cytosolic protein extracts, the harvested cells were subfractionized in homogenization buffer. The mitochondrial and cytosolic fractions were isolated with the method described by Jayanthi et al. [27]. The pellet and supernatant contained the mitochondrial fraction and cytosolic fraction, respectively.

2.11. Western blot analysis

After protein quantification with a bicinchoninic acid (BCA) protein assay kit (Beyotime, Shanghai, China), equal amounts of protein were separated by 8–15% sodium dodecyl sulfate–polyacrylamide gel electrophoresis and transferred to 0.22-μm or 0.45-μm polyvinylidene difluoride membranes followed by blocking in 5% skim milk for 1 h at room temperature. The membranes were incubated overnight at 4°C with the following primary antibodies: anti–cyt c (1:1000), anti–COX IV (1:1000), anti–cleaved caspase-9 (1:1000), anti–cleaved caspase-3 (1:1000), anti-AIF (1:1000), anti–BNIP-3 (1:1000), anti–Endo G (1:1000), and anti–β-actin (1:1000), and then with the appropriate secondary antibodies (1:5000) and enhanced chemiluminescence reagent. Each test was performed in three experiments with different batches of cells. Protein levels were determined by computer-assisted densitometric analysis (GS-800 densitometer, Quantity One; Bio-Rad). The band volumes were determined by standard scanning densitometry with normalization of densitometry measures to β-actin or COX IV.

2.12. Immunofluorescence assays

rPT cells (2 × 105 cells per well) were seeded onto sterile cover slips in 24-well plates and treated with 0, 1.25, 2.5, or 5.0 μM cadmium for 12 h; there were three replicates per group. Next, the cells were washed twice with PBS, and fixed on the coverslips with 4% paraformaldehyde. Then, the monolayer was permeated with 0.5% Triton X-100 and the cells were blocked with 5% bovine serum albumin (BSA). The cells were incubated with anti-AIF antibody (1:100) overnight at 4°C, washed with PBS, stained with Alexa Fluor 488–labeled goat anti-rabbit immunoglobulin G (IgG) (H+L) (1:500) for 1 h at room temperature, and the nuclei were stained with DAPI (5 μg/mL) in the dark. A Leica inverted phase contrast microscope (Wetzlar, GER) was used to examine and analyze the transfer rate of AIF into the nuclei.

2.13. Statistical Analysis

Data from the present study are presented as mean±SD from at least three independent experiments with different batches of cells, and each one was performed in duplicate or triplicate. Statistical comparisons were made using one-way analysis of variance (ANOVA) (Scheffe’s F test) after ascertaining the homogeneity of variance between the treatments. All statistical data were analyzed using SPSS 19.0 (SPSS, Chicago, IL, USA). The critical value for statistical significance was P<0.05.

3. Results

3.1. Cadmium induces mitochondrial oxidative stress and dysfunction in rPT cells

We detected intracellular superoxide accumulation in cadmium-treated rPT cells. The cells were stained with DCFH-DA, a superoxide-specific dye, after 12-h cadmium treatment (Fig 1A). Cadmium significantly increased the intracellular superoxide levels, which co-treatment with 100μM N-acetylcysteine (NAC) eliminated (Fig 1B). Next, we examined the changes in MPTP opening and mitochondrial morphology following cadmium treatment. Fig 1C and 1D shows that reduced mitochondrial calcein fluorescence represented MPTP opening and it was dose-dependent during cadmium exposure. The cadmium-induced changes in mitochondrial morphology were assessed by transmission electron microscopy. The morphological changes were typical of mitochondrial damage, i.e., swelling, rupture of the outer membrane, and distorted cristae (disruption or loss); the severity of the damage increased with the cadmium dose (Fig 1E).

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Fig 1.

(a) Cadmium induced ROS generation dose-dependently in rPT cells. (b) Intracellular ROS levels in rPT cells after 12-h cadmium (2.5 μM) treatment in the absence or presence of NAC (100 μM). DCF fluorescence was measured using a flow cytometer with FL-1 filter. Confocal microscopy of cadmium-induced MPTP opening. (c) Representative confocal images of cadmium-induced rPT cells. (d) Quantification of calcein fluorescence. Calcein fluorescence values were quantified relative to the control, where the fluorescence value was set at 100%. (e) Representative electron micrographs of rPT cell mitochondria following cadmium exposure (×6600 magnification). Figure shows membrane disruption (thin arrows), swelling, and damaged cristae (thick arrows). Fluorescence results are expressed as mean fluorescence intensity, and are the mean ± SD of three separate experiments, each performed in triplicate (n = 9). *P < 0.05, **P < 0.01 as compared to control.

https://doi.org/10.1371/journal.pone.0166823.g001

3.2. Effect of CsA on MPTP and apoptosis

Fig 2A and 2B show that co-incubation with CsA (an MPTP inhibitor) significantly reversed cadmium-mediated MPTP opening. The mitochondrial calcein fluorescence drastically increased from 35.7% (2.5 μM cadmium alone) to 83.6% (cadmium+CsA). However, CsA alone had no effect on MPTP opening. Annexin V/PI staining was used to determine the apoptotic cells after 12-h cadmium exposure. Fig 2C and 2D show that 2.5 μM cadmium significantly enhanced the number of apoptotic cells (early and late), being 2.84-fold that of the control. CsA co-treatment significantly prevented apoptosis in the treated cells, being 1.37-fold that of the control.

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Fig 2. Confocal microscopy of MPTP opening after 12-h cadmium (2.5 μM) treatment in the absence or presence of CsA (5 μM).

(a) Representative confocal images of cadmium-induced rPT cells. (b) Quantification of calcein fluorescence. The calcein fluorescence values were quantified relative to the control, where the fluorescence value was set at 100%. (c) Flow cytometry detection of the apoptosis rate after 12-h cadmium (2.5 μM) treatment in the absence or presence of CsA (5 μM) with annexin V–FITC/PI staining. (d) Percentage of apoptotic cells. Results are the mean ± SD of three separate experiments, each performed in triplicate (n = 9). **P < 0.01 as compared to control.

https://doi.org/10.1371/journal.pone.0166823.g002

3.3. Cyt c release and caspase-9 and caspase-3 activation as a measure of the caspase-dependent apoptotic pathway

Fig 3A show that immunoblotting indicated significant mitochondrial cyt c release to the cytoplasm after 12-h cadmium exposure. In addition, quantification (Fig 3B) demonstrated that cadmium induced cyt c release dose-dependently. Cyt c released into the cytoplasm activates caspase-9. Cadmium increased cleaved caspase-9 protein expression dose-dependently; Cleaved caspase-3 is an execution protein in apoptosis; cadmium also increased its expression dose-dependently (Fig 3C and 3D). These results confirm that the caspase-dependent pathway is involved in cadmium-induced apoptosis in rPT cells.

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Fig 3. Cadmium induced mitochondrial cyt c release to the cytoplasm and subsequent caspase-9 and caspase-3 activation in rPT cells.

(a, c) Representative western blots of cyt c, cleaved caspase-9, and cleaved caspase-3. (b, d) Quantitative analysis of cyt c, cleaved caspase-9, and cleaved caspase-3 western blots; grayscale of the control was set at 1. Quantitative analysis was performed with images from three independent experiments (mean ± SD, n = 3). *P < 0.05, **P < 0.01, and ##P < 0.01 as compared to control.

https://doi.org/10.1371/journal.pone.0166823.g003

3.4. BNIP-3 is involved in the caspase-independent apoptotic pathway and induces mitochondrial AIF and Endo G nuclear translocation

Immunoblotting demonstrated that BNIP-3 protein levels were increased, as was mitochondrial AIF and Endo G translocation to the nucleus, dose-dependently after 12-h cadmium exposure (Fig 4A–4F). Fig 4G shows that AIF staining exhibited a granular pattern in the cytosol of the control group and was restricted mainly to the nucleus, as indicated by colocalization with DAPI labeling after 12-h cadmium exposure. Immunoblotting showed that mitochondrial AIF and Endo G translocation to the nucleus was decreased after BNIP-3 knockdown (Fig 4H–4M). The results confirm that BNIP-3 is involved in the caspase-independent apoptotic pathway and causes AIF and Endo G nuclear translocation.

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Fig 4. Cadmium induced BNIP-3 expression and cytoplasmic AIF and Endo G translocation to the nucleus after 12-h cadmium treatment.

(a, c, e) Representative images of BNIP-3, AIF, and Endo G western blots. (b, d, f) Quantitative analysis of BNIP-3, AIF, and Endo G; grayscale of the control was set at 1. (g) Cadmium treatment (12 h) triggered AIF nuclear translocation dose-dependently. rPT cells were stained with anti-AIF antibodies and Alexa Fluor 488–labeled goat anti-rabbit IgG. AIF nuclear translocation was evaluated under fluorescence microscopy with DAPI staining. Scale bar: 50 μm. Effects of BNIP-3 silencing on changes in BNIP-3 expression and cytoplasmic AIF and Endo G translocation to the nucleus after 12-h cadmium treatment in the absence or presence of BNIP-3 small interfering RNA (siRNA). (h, j, l) Representative images of BNIP-3, AIF, and Endo G western blots. (i, k, m) Quantitative analysis of BNIP-3, AIF, and Endo G; grayscale of the control was set at 1. Results are from three independent experiments (mean ± SD, n = 3). **P < 0.01, #P < 0.05, and ##P < 0.01 as compared to control.

https://doi.org/10.1371/journal.pone.0166823.g004

3.5. Influence of caspase inhibitor on the caspase-independent apoptotic pathway

Cadmium (2.5 μM) and Z-VAD-FMK (20 μM, a caspase inhibitor) co-treatment decreased cleaved caspase-3 protein expression (Fig 5A and 5B). Flow cytometry revealed a significantly decreased apoptotic rate in the co-treatment group (Fig 5C and 5D). Immunoblotting showed that BNIP-3 protein levels (Fig 6A and 6B) and AIF transfer levels (Fig 6C) in the co-treatment group were significantly decreased compared with the cadmium-only group.

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Fig 5. Effects of Z-VAD-FMK on cadmium-induced rPT cell apoptosis.

(a) Representative images of cleaved caspase-3 western blot after 12-h cadmium treatment in the absence or presence of Z-VAD-FMK. (b) Quantitative analysis of cleaved caspase-3; grayscale of the control was set at 1. (c) Flow cytometry assessment of the rPT cell apoptosis rate after 12-h cadmium treatment with/without Z-VAD-FMK. (d) Percentage of apoptotic cells. Results are expressed as the mean ± SD of three separate experiments, each performed in triplicate (n = 9). **P < 0.01 as compared to control.

https://doi.org/10.1371/journal.pone.0166823.g005

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Fig 6. Effect of Z-VAD-FMK on BNIP-3 expression and cytoplasmic AIF translocation to the nucleus after 12-h cadmium treatment.

(a) Representative images of BNIP-3 western blot. (b) Quantitative analysis of BNIP-3; grayscale of the control was set at 1. (c) Twelve-hour cadmium treatment with/without Z-VAD-FMK triggered AIF nuclear translocation. rPT cells were stained with anti-AIF antibodies and Alexa Fluor 488–labeled goat anti-rabbit IgG. AIF nuclear translocation was evaluated under fluorescence microscopy with DAPI staining. Scale bar: 50 μm. Results are from three independent experiments (mean ± SD, n = 3). **P < 0.01 as compared to control.

https://doi.org/10.1371/journal.pone.0166823.g006

3.6. Influence of BNIP-3 silencing on the caspase-dependent apoptotic pathway

The earlier results confirmed that BNIP-3 is involved in the caspase-independent apoptotic pathway and induces mitochondrial AIF and Endo G translocation to the nucleus. BNIP-3 silencing blocked the caspase-independent pathway in cadmium-induced apoptosis, and clearly decreased the apoptotic rate (Fig 7A and 7B). Immunoblotting showed significantly decreased mitochondrial cyt c release to the cytoplasm after BNIP-3 knockdown (Fig 7C and 7D) and decreased cleaved caspase-9 and caspase-3 expression levels (Fig 7E and 7F).

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Fig 7. Effects of BNIP-3 silencing on cadmium-induced apoptosis, mitochondrial cyt c release to the cytoplasm, and changes in caspase-9 and caspase-3 expression.

(a) Flow cytometry assessment of the rPT cell apoptosis rate after 12-h cadmium and/or BNIP-3 siRNA treatment. (b) Percentage of apoptotic cells. (c, e) Representative images of cyt c, caspase-9, and caspase-3 western blots. (d, f) Quantitative analysis of cyt c, caspase-9, and caspase-3; grayscale of the control was set at 1. Results are from three independent experiments (mean ± SEM, n = 3). *P < 0.05, **P < 0.01, and ##P < 0.01 as compared to control.

https://doi.org/10.1371/journal.pone.0166823.g007

4. Discussion

Using rPT cells as an in vitro model, we demonstrate that cadmium preferentially induces mitochondrial oxidative stress, dysfunction, MPT, and apoptosis. Oxidative stress promotes apoptosis in primary rPT cell cultures exposed to cadmium [5, 25] and exploring the role of the mitochondrial apoptosis pathway, the caspase-dependent and caspase-independent apoptotic pathways, and the relationship between the two in rPT cell apoptosis, is necessary.

Cadmium induces rPT cell apoptosis, in which oxidative stress plays a pivotal role [25]. Our results showed that cadmium-induced preferential mitochondrial superoxide accumulation leads to mitochondrial dysfunction, which NAC prevented. ROS can directly result in MPTP opening, which facilitates MPT induction [3], the closing or opening state of MPTP, enabling tight regulation of mitochondria-mediated apoptosis [28]. Our results reveal that cadmium can lead to morphological changes typical of mitochondrial damage, including matrix swelling, outer membrane rupture, and distorted cristae. Cadmium also induced MPTP opening, triggering the release of apoptogenic proteins into the cytosol, which CsA prevented. In the cadmium and CsA co-treatment group, the rate of rPT cell apoptosis was prevented partially as compared to the cadmium-only group. Therefore, the cadmium-induced oxidative stress–induced MPTP opening, which triggers apoptogenic factor release, was halted.

The release of mitochondrial pro-apoptotic proteins such as cyt c (caspase-dependent), and AIF and Endo G (caspase-independent) strengthened the occurrence of multiple apoptotic pathways. Caspase-9 activation subsequently activates caspase-3, and requires cyt c for apoptosome formation [29, 30]. We found that oxidative stress induced cyt c release and caspase-9 and caspase-3 activation, and ultimately led to apoptosis, which is in accordance with previous research on different cell types [31–34]. A similar study indicated that the use of 10 μmol/L of Cd induces cyt c release after 24 hours of Cd treatment [35]. Compared with our results, they used a lower dose and shorter time that induced cyt c release may due to primarily cultured cells was more sensitive to Cd. Intracellular zinc (Zn) depletion also induce apoptosis and shown as loss of ΔΨ, release of cyt c and activation of caspase-9 and caspase-3 [36]. This similar phenomenon may due to Zn from the zinc enzyme could be replaced by Cd and loss of function. Meanwhile, Zn is a part of the antioxidant defence system. Zn depletion will lead to oxidative stress which also induced by Cd.

AIF is a mitochondrial protein that translocates to the cytosol and the nucleus, mediating caspase-independent apoptosis in a number of model systems [37–40]. Endo G participates in mitochondrial DNA copying, recombination, and repair [41]; induced by oxidative stress, it translocates from the mitochondria to the nucleus [42]. Endo G released from the mitochondria interacts with AIF in the nucleus and is involved in caspase-independent apoptosis in Caenorhabditis elegans [43, 44]; in neurodegenerative disease, both Endo G and AIF expression levels are decreased in the mitochondria but are increased in the nuclei [45]. AIF cannot cut DNA; it is possible that both Endo G and AIF are involved in nuclear DNA degradation [44, 46]. BNIP-3 is a BH3-only pro-apoptotic member of the Bcl-2 family: it mediates cell death via different pathways, including the mitochondrial pathway [47]. Endoplasmic reticulum–targeted BNIP-3 induces cell death that the anti-apoptotic protein Bcl-2 can block. Mitochondria-targeted BNIP-3 initiates apoptosis, inducing MPT and mitochondrial membrane potential dissipation [48], and Bcl-2 expression cannot prevent it [47]. BNIP-3 and AIF cooperate to induce apoptosis and cavitation in epithelial morphogenesis [16]. Meanwhile, silencing BNIP-3 prevents Endo G translocation and DNA degradation [49]. We found that BNIP-3 protein levels and AIF and Endo G translocation were increased during cadmium-induced rPT cell apoptosis, while BNIP-3 silencing decreased AIF and Endo G translocation. In short, BNIP-3 is involved in the caspase-independent apoptotic pathway and it is located upstream of AIF/Endo G.

Under different stimuli, PARP-1 [poly (ADP-ribose) polymerase-1] activation triggers mitochondrial AIF release and translocation to the nucleus [50–52]. PARP-1 activation produces PAR in the nucleus, which is released into the cytosol and colocalizes with the mitochondria to induce AIF release [53]. The mature AIF is loosely bound on the mitochondrial outer membrane [54], from which PAR can detach it [55]. Others have reported that without caspase activity, AIF/Endo G still translocates to the nucleus under cadmium induction [6, 56]. Consequently, the relationship between the caspase-dependent and caspase-independent apoptotic pathways is controversial due to the uncertainty as to whether there is a site upstream of AIF/Endo G. We show that BNIP-3 is involved in the caspase-independent apoptotic pathway during cadmium treatment and that it is located upstream of AIF/Endo G. BNIP-3 silencing inhibits mitochondrial cyt c release to the cytosol and caspase-dependent apoptosis in embryoid body differentiation [57]. BNIP-3 overexpression or recombinant BNIP-3 treatment of isolated mitochondria induce MPT and cyt c release in fibroblasts [48, 58]. We obtained similar results in cadmium-induced rPT cells. Here, the cadmium and Z-VAD-FMK co-treatment group had significantly decreased BNIP-3 protein levels, and apoptosis was significantly prevented, as expected. This proves that the caspase-dependent apoptotic pathway affects the caspase-independent apoptotic pathway. Z-VAD-FMK co-treatment revealed that the two pathways play a similar role (co-promotion or co-suppression), acting synergistically in cadmium-induced rPT cell apoptosis.

The part data was also observed in programmed necrosis induced by Cd. Therefore, we detected level of intracellular ATP and expression level of HMGB1 in the cytoplasm of Cd-treated rPT cells. Mitochondrial ATP production is essential for maintaining ΔΨ and preventing apoptosis [59]. In this study, decreased ATP level (S1 Fig) indicated that abnormal cellular energy metabolism promoted Cd-induced apoptosis in rPT cells; and it was not detected any HMGB1 in the cytoplasm (date not shown). Thus, rPT cells experienced apoptosis rather than programmed necrosis during Cd exposure.

In summary, BNIP-3 acts as an upstream factor in the caspase-independent apoptotic pathway to induce AIF/Endo G translocation. Cadmium activates both the caspase-dependent and caspase-independent apoptotic pathways in rPT cells, and inhibiting one restrains the other. That is, the caspase-dependent and caspase-independent apoptotic pathways are complementary in cadmium-induced rPT cell apoptosis.

Supporting Information

S1 Fig. Effect of Cd on intracellular ATP levels in rPT cells.

Cells were treated with Cd (0, 1.25, 2.5 and 5 μmol/L) for 12 h and then collected to measure the cellular ATP levels. Values represent mean ± SEM made in six different primary cultures (n = 6). **P < 0.01 as compared to control.

https://doi.org/10.1371/journal.pone.0166823.s001

(TIF)

Acknowledgments

The authors are thankful to Dr Maozhi Hu for technical assistance in flow cytometry analysis.

Author Contributions

  1. Conceptualization: GL.
  2. Data curation: ML.
  3. Formal analysis: TL.
  4. Investigation: GL.
  5. Methodology: GL HZ.
  6. Project administration: JB.
  7. Resources: ZL YW RS.
  8. Software: YY.
  9. Supervision: XL.
  10. Validation: JZ.
  11. Visualization: JG.
  12. Writing – original draft: GL.
  13. Writing – review & editing: JZ.

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  • Apoptosis 
  • Cadmium 
  • Mitochondria 
  • Cell staining 
  • DAPI staining 
  • Oxidative stress 
  • Cytoplasm 
  • Flow cytometry 
Sours: https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0166823


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