Acute ATP Loss During Irreversible Electroporation Mediates Caspase Independent Cell Death

Abstract

Irreversible electroporation (IRE) has been reported to variably cause apoptosis, necrosis, oncosis or pyroptosis. Intracellular ATP is a key substrate for apoptosis which is rapidly depleted during IRE, we sought to understand whether intracellular ATP levels is a determinant of the mode of cell death following IRE. A mouse bladder cancer cell line (MB49) was treated with electric fields while increasing the number of pulses at a fixed electric field strength, and pulse width. Cell proliferation and viability and ATP levels were measured at different timepoints post-treatment. Cell death was quantified with Annexin-V/Propidium Iodide staining. Caspase activity was measure with a fluorometric kit and western blotting. A pan-caspase (Z-VAD-FMK) inhibitor was used to assess the impact of signal inhibition. We found cell death following IRE was insensitive to caspase inhibition and was correlated with ATP loss. These findings were confirmed by cell death assays and measurement of changes in caspase expression on immunoblotting. This effect could not be rescued by ATP supplementation. Rapid and acute ATP loss during IRE interferes with caspase signaling, promoting necrosis. Cell necrosis from IRE is expected to be immunostimulatory and may be effective in cancer cells that carry mutated or defective apoptosis genes.

1. Introduction

The application of high-intensity pulsed electric fields cells induces the molecular rearrangement of the cell membrane, leading to the formation of aqueous pathways or “pores” that can be temporary (Reversible Electroporation; RE) or long-lasting (Irreversible Electroporation; IRE) [1]. Following RE, the integrity of the cell membrane recovers, allowing most cells to survive. During IRE the majority of cells are unable to recover homeostasis, leading to cell death [2]. IRE is currently used for the treatment of tumors involving sensitive structures such as the bile duct, large blood vessels or the urinary tract, allowing therapy of patients who are unable to undergo external beam radiation or conventional thermal ablation using radiofrequency, microwave or cryoablation technique [3]. IRE has been associated with immune priming; with increased infiltration of T cells and macrophages in the treatment region and enhanced anti-cancer immune cell activity, generating interest in using it as an adjuvant to improve the efficacy of immune checkpoint inhibition therapy [4–6]. Likewise, evasion or emergence of resistance to apoptosis is a hallmark of cancer cells, and it is clinically important to ensure that IRE can effectively treat tumors with such characteristics [7].

Cell death during IRE and RE has been reported to occur through programmed and unprogrammed pathways, namely apoptosis and necrosis respectively. Literature suggests the variable presence of both pathways in cells dying following exposure to electric pulses, with occasional involvement of other non-canonical forms of cell death such as pyroptosis, necroptosis and oncosis [8–11]. Apoptosis is characterized by numerous morphological changes in cell structure paired with caspase-dependent biochemical processes leading to the clearance of cells from the body with minimal damage to the surrounding tissues. Apoptosis occurs physiologically at a pace of millions of events per second in a healthy human and is broadly considered non-immunogenic. In contrast, necrosis is a sudden and uncontrolled process that follows an acute physiologic insult, resulting in the release of cellular content into the surrounding tissues and damage to the cellular microenvironment [12]. Necrosis creates a pro-inflammatory environment inducing the secretion of Th1 cytokines, attraction of antigen-presenting cells dendritic cells, and cross-priming of potent T cell response against the cancer cells. In addition, damage-associated molecular pattern molecules (DAMPs) are actively or passively exposed or released to extracellular space leading to immune stimulation [2,13]. As the clinical application of IRE to augment cancer immunotherapy is dependent on inducing necrosis, we seek to understand whether this is indeed a dominant mode of cell death following IRE and identify the factors mediate the process.

Permeabilization of the cell membrane during IRE is the preliminary injury that initiates cell death. Multiple secondary events such as the of loss homeostasis, increased intracellular calcium levels, and damage to the cell membrane and organelles by Reactive Oxygen Species (ROS) are reported to be potential pathways mediating cell death following IRE [14,15]. However, the specific mechanisms that lead to apoptosis or necrosis have not been clearly delineated and it is unknown whether a desired form of cell death can be obtained in a selective fashion. ATP leakage and depletion are hallmarks of both RE and IRE. Studies investigating the interaction between calcium, RE and ATP have shown that cellular ATP levels drop following electroporation [2,16]. This was associated with the increased activity of Ca²⁺ pumps, a disturbance in mitochondrial metabolism and ATP leakage through the permeabilized cell membrane, leading to different types of cell death varying based on pulse parameters used, cell and tissue type being treated, and other factors [9,16]. ATP loss may be a key determinant of the mode of cell death following IRE as considerable intracellular ATP is consumed during caspase signaling, and therefore essential for the completion of the apoptosis cascade. Several studies have reported induction of necrosis in cells treated with drugs and electroporation, under conditions of depleted intracellular ATP levels [2,17,18].

The objective of our study was to explore the link between intracellular ATP levels and to necrosis following IRE, and to understand whether IRE can be effective in killing cells under conditions of caspase signaling inhibition. For the same, we have used a cell permeable, pan-caspase inhibitor (Z-VAD-FMK) to study post-IRE cell death signaling [19]. The central hypothesis of our experiments was that rapid depletion of intracellular ATP during IRE renders necrosis as the dominant mode of cell death.

2. Materials and methods

Cell culture

Experiments were performed using a mouse bladder cancer cell line (MB49, kind gift from the Coleman Lab, Memorial Sloan Kettering Cancer Center, NYC, USA). The cell line was screened for pathogens and grown in DMEM with antibiotic-antimycotic: Penicillin G (50–100 U/ml), Streptomycin Sulfate (100 U/ml), Amphotericin B (0.25–2.5 μg/ml) (Gibco), supplemented with 10 % fetal bovine serum (One Shot, Gibco). The cells were maintained in an incubator at 37 °C with 5% CO2, and all experiments were performed on cells between passage numbers 5 and 15. Schematic of the experimental design is described in Schematic.1

Schematic 1.

Description of experimental conditions, assessment timepoints and assays.

Reagents

Benzoyloxycarbonyl-Val-Ala-Asp-fluoromethylk-etone (Z-VAD-FMK) was purchased from R&D Systems (USA). It was reconstituted using 107 μL of DMSO to yield a 20 mM stock solution, that was then diluted in culture-media to derive a final concentration of 50 μM in solution. Tris buffered Adenosine 5’-triphosphate (ATP) was purchased from Thermo Scientific (USA). The initial 100 mM aqueous solution in a 0.025 ml initial volume was prepared with serial tenfold dilutions to derive a final concentration of 1.5 μM in solution.

Treatment parameters

The cell line was trypsinized, centrifuged, suspended in DMEM, seeded at a cell density of 10⁵ cells in a 24-well plate and incubated for 24 hours at 37°C and 5% CO₂ prior to treatment with or without the addition of Z-VAD-FMK or exogeneous ATP (both added 1 hour before pulse delivery). The cells were removed from the incubator, transferred to a 24-well plate treated by electroporation using pin electrodes (Fig. S1, see supplementary material). A square wave pulse generator (BTX ECM 830, Harvard Apparatus, Holliston, MA, USA) was used with the following electroporation parameters: 0, 10, 30, 50, 90 pulses of 100 μs width, 1 Hz frequency, and 1500 V/cm field strength. After treatment, all cells were returned to the incubator at 37°C and 5% CO₂ until the desired endpoint for appropriate assay.

Cell metabolism and cell survival Assays

Cell Counting Kit-8 (CCK-8) assay (Dojindo Laboratories, Kumamoto, Japan) was used to measure the cell proliferation at 4- and 24-hours following treatment. CCK-8 solution (50 μL) was added to each well of the 24-well plate, followed by incubation for 4 hours at 37°C. The absorbance at 450 nm was determined by a multiplate reader (SpectraMax, Molecular Device). Cell proliferation was expressed as a percentage of the control group (untreated cells). Relative cell viability was assessed by Trypan blue staining, following by automatic image acquisition and analysis using the ImageXpress Pico Automated Imaging System (Molecular Device). Relative survival and proliferation rate was expressed as percentage of the sham control group.

Cell death analysis by Annexin-V/PI Assay

Cell death was measured at the 4 hours post-electroporation using Annexin/Propidium Iodide double staining. Cell pellets were extracted from the 24 well plates and 100 μL of Annexin V Binding Buffer 1X (Sigma-Aldrich), 10 μL of PI solution (Sigma-Aldrich) and 5 μL of APC Annexin V (Fisher scientific) were added. Stained cells were seeded into 96 well plates and the whole plate was imaged and automatically quantified using ImageXpress Pico software.

ATP assay

ATP levels in culture was measured using Cell-Titer-Glo^® Luminescent Cell viability Assay (Promega) at serial timepoints (30 minutes, 2 hours and 24 hours). Measurements were performed separately to quantify levels of ATP in the supernatant and the cell pellet. The absorbance at 578 nm was determined by a multiplate reader (SpectraMax, Molecular Device). Concentration of ATP was calculated using standard curves.

Caspase 3/7 activity

Caspase activity was measured with the use of CellEvent Caspase-3/7 Green Detection Reagent (ThermoFisher, USA), imaged with a ImageXpress Pico Automated Imaging System. This reagent consists of a four amino acid peptide (DEVD) conjugated to a nucleic acid binding dye. After activation of caspase-3 or caspase-7 in apoptotic cells, the DEVD peptide is cleaved, enabling the dye to bind to DNA and produce a bright, fluorogenic response with an absorption/emission maximum of 502/530 nm. Caspase activity was measured at 4 hours following pulse application with or without the presence of Z-VAD-FMK. Image acquisition was performed using the ImageXpress Pico system to detect FITC and Alexa FluorTM dye at the green fluorescence wavelength of 502/530 nm.

Western blot analysis

MB49 cells were treated with reversible electroporation settings (10 pulses) or irreversible electroporation settings (50 pulses) with or without Z-VAD-FMK (50 μM) and ATP (1.5 μM). At 4 hours following treatment, cells were harvested, lysed with M-PER (Thermo Scientific) plus HALT protease inhibitor cocktail (Thermo Scientific) and frozen at −80°C. Equal amount (10μg) of proteins from total cell lysate were mixed with Laemmli sample buffer at 95°C for 5min. Sample solutions were loaded to electrophoresis gels and transferred onto a PVDF membrane (Bio-Rad) after electrophoresis. After transfer, the membrane was blocked by 5% non-fat dry milk in tris-buffered saline with 0.1% Tween 20 detergent (TBS-T). The protein was probed with primary antibody (caspase 8 #4790, pro-caspase 3 #9664, Beta Actin #4970, Cell Signaling; 1:1000) diluted into 5% BSA in TBS-T for 1 hour and then with secondary antibody (anti-Rabbit immunoglobulin G (IgG); 1:5000) conjugated to HRP diluted into 5% non-fat dry milk in TBS-T for 1 hour. The HRP signal was developed using enhanced chemiluminescence (ECL) reagent. Blots were imaged and densitometry values obtained using the ChemiDOC western blot imaging system.

Statistical analysis

Statistical analyses were performed using the GraphPad Prism 8 software. Normality of the data was verified using Shapiro-Wilk test. Values were presented as a mean ± SD (Gaussian distribution) or median (5^th-95^th) (non-Gaussian distribution); Data were compared using unpaired Student’s t-test was used to determine the statistical significance of difference between cohorts when the data demonstrated normal distribution. The Mann–Whitney U test was used in datasets without normal distribution. Values are deemed to be statistically significant for p values of * p<0.05.

3. Results

3.1. Cell death during IRE but not RE occurs independent of caspase activity.

We performed a dose escalation study by increasing the number of pulses applied with or without Z-VAD-FMK to understand the dependency of cell death on caspase mediated apoptosis. Increasing the number of electric pulses applied at a fixed electric field strength (1500V/cm) resulted in the progressive reduction of proliferation and cell viability at both 4 and 24 hours after treatment (Fig. 1). Cells incubated with Z-VAD-FMK prior to the pulse application did not demonstrate differences in cell viability or proliferation at the assessed timepoints when treated with higher number of pulses (30 and 90 pulses). In contrast, sham control cells demonstrated increased proliferation at both timepoints (Fig. 1a,b) without change in viability (Fig. 1c,d), while cells treated with 10 pulses and Z-VAD-FMK had greater proliferation at 24 hours when compared to the no Z-VAD-FMK group (Fig. 1b). Despite this effect, no difference was found in relative cell viability at the same pulse number with or without the pan-caspase inhibitor (Fig. 1c,d). These results suggest that cell death under RE-like conditions (10 pulses or fewer) may involve caspase dependent mechanisms, while these are largely absent during IRE-like treatment conditions (30 pulses or greater).

Fig 1.

Cell proliferation and viability in response to dose escalation by increasing pulse numbers. Cell proliferation and viability relative to sham control was assessed at 4 (a,c) and 24 (b,d) hours following electroporation. Caspase inhibition was performed by incubating cells in Z-VAD-FMK for 1 hour prior to pulse application (white bar) or without (black bar). Data presented as mean and standard deviations. (* p < .05)

We further tested the mode of cell death under RE (10 pulses) or IRE (50 pulses) conditions with or without the pan-caspase inhibitor by differential staining using Annexin (apoptosis or caspase mediated cell death), PI staining (necrosis) or dual Annexin + PI staining (both apoptosis and necrosis) (Fig. 2). Both RE and IRE groups demonstrated positive staining for Annexin or Annexin+PI, but limited staining for PI alone (Fig. 2b,c). The IRE group had greater number of cells positive for both stains, but with substantially greater number of cells staining for Annexin+PI (Fig. 2d). These numbers were statistically significant when compared to control or solely Z-VAD-FMK condition. The addition of the pan-caspase inhibitor slightly reduced RE treated cells staining for Annexin only or Annexin+PI, but these results were not statistically significant (Fig. 2d). Similar changes were observed in IRE treated cells, where the addition of Z-VAD-FMK resulted in a slight in reduction in cells staining for Annexin+PI, but these results were also not statistically significant (Fig. 2d).

Fig 2.

Cell death assessment by AnnexinV-APC (green) and Propidium iodide (PI, red) staining in MB49 cells. Cellular response following sham control (a), RE (10 pulses, b) and IRE (50 pulses, c) treatment with or without pre-treatment with Z-VAD-FMK. Samples were analyzed at 4 hours after treatment. Representative pictures bright field (left) and fluorescence images (right) at 10x magnification (White scale bar = 50 μm). Results are quantified with percentage of stained cells for each dye or combination (d) (* p < .05)

To determine whether caspase pathways were involved in the induction of cell death following electroporation (EP), we performed an assay for Caspase 3/7 at 4 and 24 hours after treatment under RE or IRE conditions (Fig. 3). Following pulse delivery, no change in caspase signaling was observed at 4 hours post-treatment. At 24 hours post-treatment, a statistically significant increase of caspase 3/7 activity was seen in the RE group, which was reversed using Z-VADFMK (Fig. 3b,e). For the IRE group, a significant increase in caspase 3/7 activity compared to sham group appeared only at 24 hours after treatment, with no observable effect from treatment with Z-VAD-FMK (Fig. 3c,e). These findings suggest the predominance of caspase mediated apoptosis during RE which can be rescued by the addition of the pan-caspase inhibitor, while post-IRE cell death seemed to progress independent of caspase signaling.

Fig 3.

Caspase 3/7 activity measured with a fluorescent assay. Representative images for sham (a), RE (10 pulses, b) and IRE (50 pulses, c) conditions with or without pre-treatment with Z-VAD-FMK. Samples were analyzed at 4 hours and 24 hours after treatment. Representative pictures bright field and fluorescence images at 10x magnification 24 hours after treatment (White Scale bar = 100 μm). Results are quantified for percentage cells staining positive at 4 (d) and 24 (e) hour timepoint. (* p < .05)

3.2. Post-EP ATP loss determines mode of cell death

As the caspase signaling cascade is an ATP dependent process, we sought to understand the interaction between EP related ATP loss and the associated mode of cell death. RE or IRE was performed in cells with or without the presence of Z-VAD-FMK, followed by assessment of intracellular and extracellular ATP at 30 minutes, 2- or 4-hours endpoint after pulse application (Fig. 4). Compared to sham controls, RE and IRE induced a significant decrease in intracellular ATP content at all timepoints (30 minutes, 2 hours, 4 hours) (Fig. 4a,b,c) with a moderate release of ATP into the supernatant at 30 minutes post-pulse application (Fig. 4d). The loss of intracellular ATP was greater following IRE when compared to RE, where the ATP levels in the latter group recovered towards baseline within 4 hours post-treatment (Fig. 4c). Addition of Z-VAD-FMK did not alter intracellular ATP levels in the IRE group, but a sharp reduction could be observed at both 30 minutes and 2 hours timepoints in the RE group (Fig. 4a,b). Likewise, Z-VAD-FMK significantly reduced the amount of extracellular ATP in the RE group at the acute timepoint (30 minutes) (Fig. 4d). These findings were absent in the IRE group.

Fig 4.

Kinetics of intracellular and extracellular ATP levels following electroporation. Intracellular and extracellular ATP levels following RE or IRE in cells with (white box) or without (black box) Z-VAD-FMK pre-treatment at (30 minutes (a,d), 2 hours (b,e) and 4 hours (c,f) following pulse application. (* p<.05)

To further understand the interaction of ATP and post-EP cell death, we studied the effect of exogenous ATP supplementation to overcome EP related loss (Fig. 5). The addition of exogenous ATP did not alter early apoptotic and late apoptotic/necrotic cells ratio for all groups at 4-hours post-treatment (Fig. 5a,b,c). ATP supplementation and Z-VAD-FMK reduced the caspase activity in the absence of electric pulse application (Fig. 5d). ATP supplementation did not alter the caspase 3/7 activity for IRE in contrast to RE (Fig. 5f). Likewise, for RE, combination ATP supplementation and the pan-caspase inhibitor was associated with significantly decreased caspase activity with a potentiator effect (Fig. 5e). The addition of the pan-caspase inhibitor was able to reverse the RE condition related reduction of cell proliferation without altering overall viability (Fig. 6). Neither Z-VAD-FMK nor ATP-supplementation was able to reverse reduction in proliferation or viability in the IRE group (Fig. 6)

Fig 5.

Impact of exogenous ATP on electroporation related cell death. The effect of exogenous ATP (1.5 μM) given to MB49 cells after no pulse, RE or IRE with or without Z-VAD-FMK pre-treatment at 4 hours post-pulse application. Annexin-V-APC and PI double staining (a,b,c) and Caspase 3/7 assay (d,e,f) were performed. Data show as mean +/− standard deviation. (*p<.05)

Fig 6.

Cell viability and proliferation when performing RE or IRE in cells treated with exogenous ATP or Z-VADFMK. Cell viability (a) and proliferation (b) relative to sham control were assessed at 4 hours post pulse application. (* p < 0.05)

Caspase 8 and caspase 3 play initiator and executioner roles respectively in the apoptotic cascade. Reduction in caspase 8 levels is associated with the start of the apoptosis process which ends with decreased levels of pro-caspase 3 due to proteolytic cleavage. We measured relative levels of these proteins following EP with or without ATP supplementation or Z-VAD-FMK (Fig. 7). In the control condition, treatment with the pan-caspase inhibitor decreased the levels of pro-caspase 3 without change in caspase 8 levels. Following RE, ATP supplementation or presence of Z-VAD-FMK increased pro-caspase 3 and caspase 8 levels, consistent with interruption of the apoptotic cascade. Similar findings were observed in the IRE treated cells, but changes in the levels of pro-caspase 3 and 8 did not correlate with the extent of cell death in these cohorts.

Fig 7.

Immunoblotting of caspase 8 and pro-caspase 3 in MB49 cells following RE or IRE, with or without exogenous ATP and Z-VAD-FMK.

4. Discussion

Our results demonstrate that cell death following IRE but not RE progresses unhindered even under conditions of caspase inhibition, suggesting independence from this classical apoptosis pathway. Cell death markers for both apoptosis and necrosis manifest following IRE, indicating the initiation but not the completion of the latter signaling cascade. We demonstrate that necrosis emerges as the dominant cell death mode following IRE due to these conditions, and that ATP loss was potentially contributing to this effect, where supplementation with exogenous ATP did not rescue the cells. These findings contribute to our understanding on factors that can improve cell viability following RE for gene transfection applications. Further establishing necrosis as the dominant cell death mode solidifies the utility of IRE as an inflammatory adjuvant therapy for combination with immunotherapy applications.

The effect of the IRE on cancer cell lines has been studied extensively and its ability to induce cell death is widely recognized by the scientific community but the specific mode by which it occurs has not been clearly established [20]. One study using high-frequency irreversible electroporation in 4T1 cells showed a necrosis predominant mode of cell death [21]. At the same time, another study using BAX and TUNEL assays reported that IRE-induced mode of cell death was by apoptosis [22]. Apoptosis through the activation of caspase-3 or -7 was previously considered as the primary mode of cell death following IRE [23–26]. However, there is growing evidence that IRE induces necrosis, possibly involving programmed cell death pathway as necroptosis or pyroptosis [27]. Our results demonstrate that 4 hours after treatment, there is increased levels of early apoptotic patterns proportional to the number of pulses applied, and at later timepoints dual apoptotic/necrotic staining emerges. Our findings are in agreement with the work of Fernandes et al. showing pancreatic cancer cells treated with electroporation stain positive for both Annexin and PI and that electrochemotherapy increased staining by AnnexinV/PI without a coincident increase of AnnexinV population, suggesting a predominance of necrotic cell death in the latter condition. Taken together, it seems that IRE may produce markers for both apoptosis and necrosis with time dependent kinetics where cell death eventually occurs via the latter process. In contrast, there is considerable evidence that apoptosis may be the dominant form of cell death following RE [28,29]. Our results are consistent with the literature where there is a significant increase of pro-caspase 3 cleavage and caspase-3/7 activity following treatment under RE-like conditions, where such activity is curtailed by treatment with Z-VAD-FMK [30,31].

Electropermeabilization of the cell membrane is well known to elicit the release of ATP, which has been used as a permeability marker for RE [2,17,32]. Simultaneously, ATP is required for caspase activation, serving as a key substrate for the signaling cascade. The lack of intracellular ATP can impede the cell’s ability to undergo apoptosis [33]. This provides a potential rationale for the progression of cell death even when blocking caspase signaling during IRE in our experiments. Our results are consistent with previous studies showing acute intracellular ATP depletion in proportion to the number of pulses applied [2,16]. During IRE, the greater magnitude of intracellular ATP loss may impede the activation and progression of caspase signaling, and therefore limiting the execution of programmed cell death. We tested whether exogenous supplementation of ATP can influence or alter the mode of cell death. Previously, Leist et al. showed that intracellular ATP depletion > 50–70% was the trigger level to switch from an apoptotic process to a necrotic process in Jurkat cells and higher ATP concentration favored the ordered continuation of apoptotic programs [34]. We found that such supplementation did not impact cell death during IRE, suggesting a deeper alteration in the cell metabolism preventing induction of apoptotic pathways. It is well known that mitochondria are essential for ATP production, and for cellular functions such as maintenance of Ca ²⁺ homeostasis and the elimination of reactive oxygen reactive species (ROS) [35]. Esser et al. suggested that upon membrane permeabilization induced by electroporation, voltage-sensitive organelles such as mitochondria could be affected by external electric field [36]. Reynaud et al showed that severe damages to mitochondria could be observed when electroporation is performed with field strength greater than 1kV/cm [37]. Mitochondria are also known to be essential for apoptosis by releasing proteins for the activation of caspase pathways. This is accomplished by cytochrome C release into the cytosol which after binding with APAF-1 results in the formation of a scaffold necessary for caspase activation. This process is energy expensive, requiring considerable levels of ATP to complete. Therefore, in addition to acute ATP depletion by leakage, injury to the mitochondria may be another factor that contributes to cell death following IRE under our experimental conditions. The mechanistic details, as well as interaction with Ca2+ can form basis of future, follow-up studies in the topic.

To further understand the interaction of caspase signaling cascade and IRE, we studied the levels of caspase 8 and pro-caspase 3 with or without the presence of Z-VAD-FMK and exogenous ATP. The induction of the apoptosis was marked by a decrease in pro-caspase 3 by proteolytic cleavage, and the consumption of caspase 8 which acts as an initiator for the process. In the sham treatment condition, treatment with Z-VAD-FMK increased pro-caspase 3 levels with minimal impact on the levels of caspase 8, consistent with increased cell viability. This observation was similar to what has been reported in literature, showing caspase blockade after caspase 3 cleavage and a direct inhibition of the initiator caspase 8 [19,38–40]. RE or IRE treatment resulted in an immediate decrease in caspase 8 and pro-caspase 3 levels when compared to sham treatment, suggesting initiation of the apoptotic processes. These levels were substantially increased by treatment with the pan-caspase inhibitor or by exogenous ATP in the RE condition but was impacted only by the addition of ATP for the IRE condition. These results are suggestive of a protective effect of ATP. However, increased expression of the caspase proteins did not impact cell viability in cells treated with IRE. It is also possible that the presence of exogenous ATP may hinder apoptosis by serving as a survival factor. During periods of stress, ATP can sequester cytochrome C leaked by the mitochondria, protecting the cells from accidental apoptosis [41]. However, due to the severity of insult to the cell by IRE this alone may be insufficient to abrogate cell death as the cell still dies by other caspase-independent necrotic pathways [42].

Our experiments were limited by our experimental conditions where we used a single caspase inhibitor (Z-VAD-FMK) and a single murine cancer cell line (the MB49 bladder cancer cell line). Our findings have to be further validated in cells of different lineages and cancer types. Despite its broad activity, the caspase inhibitor we tested does not abrogate all apoptotic signaling, such as what may be initiated by TRAIL ligands or Cytochrome C pathways [43,44]. Thus, the contribution of these other cell death pathways need to studied to truly ensure necrosis as the outcome of IRE. We tried to offset some of these weaknesses by multi-assay investigation of caspase signaling using colorimetric assays, western blotting and molecular biology techniques. In addition, the use of a necrosis inhibitor as necrostatine-1 to trigger the necrotic pathway could further confirm the full role of this mode of cell death following IRE.

Conclusion

Our experiments suggest that the progression of caspase signaling during IRE can be impeded by acute ATP loss, resulting in abrupt cell death by necrosis. We observed that this effect could not be altered by exogenous ATP supplementation and was unaffected by pre-treatment with caspase-inhibitors such as Z-VAD-FMK. As necrosis is considered an inflammatory form of cell death, tumor ablation with IRE is anticipated to induce robust stimulation of the immune system even cancer cells with deficient apoptotic pathways. Taken in context with existing body of literature, we anticipate this process may have interactions with mitochondrial injury and dysfunction following IRE. There may also exist other dependencies with cell metabolic status and activity, which need to be uncovered by experiments using cells of different lineage and cancer types. Our work addressed the impact of caspase inhibition but interactions with other ligands and pathways that can trigger apoptosis, such as Fas ligand, gasdermin or TRAIL merit investigation. It would be interesting to study cell fate in the presence of both apoptosis and necrosis inhibitors to ascertain whether cell death is from abrupt necrosis or if programmed pathways are involved.

Highlights.

• Rapid and acute ATP loss during IRE promotes necrotic cell death.

• Cell death from IRE progresses even during caspase signaling inhibition.

• IRE can be immunostimulatory and effective in cancers with mutant apoptosis genes

Abbreviations

IRE

Irreversible electroporation

IR

Interventional radiologist

ATP

Adenosine triphosphate

EP

Electroporation

z-VAD-fmk

Benzoyloxycarbonyl-Val-Ala-Asp-fluoromethylk-etone

DAMPS

Damage-associated molecular pattern molecules (DAMPs)

DMEM

Dulbecco modified Eagle’s minimal essential medium

DMSO

Dimethyl sulfoxide

MMC

Mitomycin C

PBS

Phosphate buffered saline

CCK8

Cell Counting Kit-8

EDTA

Ethylenediaminetetraacetic acid

PI

Propidium Iodide

FITC

Fluorescein isothiocyanate

ROS

Oxygen Reactive species