Although electric field–induced cell membrane permeabilization (electroporation) is used in a wide range of clinical applications from cancer therapy to cardiac ablation, the cellular- and molecular-level details of the processes that determine the success or failure of these treatments are poorly understood. Nanosecond pulsed electric field (nsPEF)–based tumor therapies are known to have an immune component, but whether and how immune cells sense the electroporative damage and respond to it have not been demonstrated. Damage- and pathogen-associated stresses drive inflammation via activation of cytosolic multiprotein platforms known as inflammasomes. The assembly of inflammasome complexes triggers caspase-1–dependent secretion of IL-1β and in many settings a form of cell death called pyroptosis. In this study we tested the hypothesis that the nsPEF damage is sensed intracellularly by the NLRP3 inflammasome. We found that 200-ns PEFs induced aggregation of the inflammasome adaptor protein ASC, activation of caspase-1, and triggered IL-1β release in multiple innate immune cell types (J774A.1 macrophages, bone marrow–derived macrophages, and dendritic cells) and in vivo in mouse skin. Efflux of potassium from the permeabilized cell plasma membrane was partially responsible for nsPEF-induced inflammasome activation. Based on results from experiments using both the NRLP3-specific inhibitor MCC950 and NLRP3 knockout cells, we propose that the damage created by nsPEFs generates a set of stimuli for the inflammasome and that more than one sensor can drive IL-1β release in response to electrical pulse stimulation. This study shows, to our knowledge, for the first time, that PEFs activate the inflammasome, suggesting that this pathway alarms the immune system after treatment.

Treatments with intense pulsed electric fields (PEFs) are central for many existing and emerging medical applications, including tumor treatment (1–7), vaccination (8–11), cardiac ablation (12, 13), and gene therapy (14–16). PEF technologies include electrochemotherapy (ECT), gene electrotransfer (GET), irreversible electroporation (IRE) and nanosecond PEFs (nsPEFs) (17). In these methods the delivery of electrical energy causes cell membrane permeabilization in the treated area; in most protocols the rate of energy deposition is controlled so that the concurrent Joule heating does not cause thermal damage (3, 17–20). Depending on the application, this disruption of the membrane barrier function, called electroporation (or electropermeabilization), can be either reversible or irreversible. Protocols causing reversible electroporation, in which permeabilizing structures are transient and membrane integrity is quickly recovered, are used in ECT and GET to introduce into cells substances that are otherwise impermeant, such as drugs, proteins, and nucleic acids (21, 22). Conversely, IRE creates damage that exceeds the cell’s membrane repair capacity, leading to Ca2+ overload, efflux of ATP and other metabolites, and disturbances in transmembrane ion gradients (Na+, K+, Cl) required for maintenance of membrane resting potential and for osmotic and cell volume regulation (17, 23–30). Lethal PEF applications such as IRE and nsPEFs cause both immediate and delayed cell death by multiple mechanisms, still incompletely defined (1, 3, 23, 31, 32).

PEF protocols currently in use in the clinic were developed to maximize tumor tissue ablation (IRE and nsPEFs) or to achieve the highest drug uptake or the most persistent gene expression (ECT and GET). Recent research in both healthy and tumor tissue indicates that PEFs have a potent immune stimulatory effect (5, 7, 33–36), a novel dimension that merits consideration during treatment planning. Activation of a robust immune response against cancerous cells or a pathogen Ag can be a significant advantage for any anticancer therapeutic modality or vaccine. Conversely, PEF-induced immune activation can be harmful for other applications such as cardiac ablation and gene therapy. Understanding the mechanisms responsible for PEF-induced immune stimulation, and their dependence on pulse parameters, is essential for the improvement of existing PEF applications and the development of new ones.

The immune system detects threats such as tissue damage, infections, and metabolic stress through pattern recognition receptors. The nucleotide-binding domain, leucine-rich repeat–containing receptors (NLRs) are pattern recognition receptors that initiate inflammatory responses (37). Upon activation NLRs trigger the assembly of multiprotein complexes known as inflammasomes. Inflammasomes are typically formed by three components: a sensor, an adaptor, and an effector. Activation of the inflammasome promotes the secretion of proinflammatory cytokines and induces pyroptosis. Among the multiple NLRs, NLR family, pyrin domain containing 3 (NLRP3) has been the most extensively studied (38, 39). The NLRP3 inflammasome is regulated by a two-step process: a “priming” stimulus is required to initiate expression of key inflammasome components, followed by a secondary “activating” stimulus that results in assembly of the inflammasome complex (38, 39). This process involves the oligomerization of NLRP3 proteins, which then recruit the adaptor apoptosis-associated speck-like protein containing a CARD (ASC) and caspase-1. Autocatalytic activation and cleavage of caspase-1 enables cleavage of proinflammatory cytokines IL-1β and IL-18, as well as the pore-forming protein gasdermin D (GSDMD). IL-1β is released through GSDMD pores, and in larger amounts during pyroptosis, the lytic cell death that often follows GSDMD pore formation (38, 39). IL-1β and IL-18 are involved in the innate immune response to infection and trauma, creating a generalized proinflammatory environment. Detection of active caspase-1 and mature IL-1β and IL-18 are commonly used in research as indicators of NLRP3 activation (40).

The NLRP3 inflammasome sensor is activated by several chemically and structurally diverse triggers, including markers of cell damage (e.g., ATP), environmental pollutants such as silica, and pore-forming toxins (38, 39). Because a direct interaction with each activator is unlikely, it is assumed that the NLRP3 inflammasome either senses a common secondary activator downstream of these stimuli or responds to cellular stress signals associated with infection or damage. For instance, a feature common to all NLRP3 stimuli is K+ efflux, an indicator of cell membrane permeabilization. Other signals proposed to be critical for NLRP3 activation include reactive oxygen species (ROS), cell volume changes, elevation of intracellular calcium levels, mitochondria destabilization, and endoplasmic reticulum (ER) stress (38, 39, 41, 42). Interestingly, several of these mechanisms mirror PEF bioeffects, especially responses to nanosecond pulses used in nsPEFs. Unlike the microsecond and millisecond PEFs, nanosecond pulses permeabilize not only the outer (plasma) membrane of the cell, but also intracellular membranes of organelles such as the ER and the mitochondria, and these “ultrashort” pulses deposit proportionally more energy in intracellular membranes than in plasma membranes (28, 29, 43–46). Membrane permeabilization by nsPEFs causes K+ efflux (47), intracellular Ca2+ mobilization (28, 29, 48, 49), and cell swelling (45). Moreover, cell damage by nsPEFs has also been found to increase ROS production and trigger ER stress (7, 50, 51).

In this study we investigated whether the damage created by nsPEFs is sensed intracellularly by the NLRP3 inflammasome. We show that nsPEFs trigger caspase-1 activation and IL-1β release in innate immune cells in vitro and in vivo in mouse skin. We also found that nsPEF-induced inflammasome activation is partially dependent on K+ efflux and that longer, microsecond pulses are less effective. Finally, whereas IL-1β release in response to nsPEFs was blocked in cells treated with the NLRP3 inhibitor MCC950, its release was not impaired in bone marrow–derived macrophages (BMDMs) from NLRP3 knockout (NLRP3-KO) mice. These results suggest that nsPEFs generate a set of stimuli for the inflammasome that are sensed not only by the NLRP3 platform but also by other, not yet identified sensors.

Murine macrophage J774A.1 cells (TIB-67) and human embryonic kidney (HEK)293T cells (CRL-3216) were obtained from American Type Culture Collection (Manassas, VA). Cells were cultured in high-glucose DMEM (Corning, New York, NY) supplemented with l-glutamine (American Type Culture Collection), 10% FBS (Atlanta Biologicals, Norcross, GA), 100 U/ml penicillin, and 0.1 mg/ml streptomycin (Life Technologies, Gaithersburg, MD) at 37°C and 5% CO2.

Wild-type, NLRP3-KO BMDMs and dendritic cells (DCs) were differentiated as previously described (52, 53). Briefly, female BALB/c or C57BL/6 (NLRP3-KO) mice (The Jackson Laboratory, Bar Harbor, ME) were euthanized and contents of femurs and tibiae were flushed with cold medium using a 5-ml syringe and a 25G needle. For BMDMs, cells were plated at 6–8 × 106 cells/plate in 7 ml of DMEM with 20% FBS, 100 U/ml penicillin, 0.1 mg/ml streptomycin, 25 mM HEPES, and 30 ng/ml recombinant murine M-CSF (PeproTech, Cranbury, NJ). To differentiate DCs, 6–8 × 106 cells/plate were cultured in RPMI 1640 with 10% FBS, 50 µM 2-ME (Thermo Fisher Scientific, Waltham, MA), and penicillin/streptomycin and supplemented with 10 ng/ml recombinant murine GM-CSF (PeproTech) and 20 ng/ml murine IL-4 (PeproTech). On day 3, 3 ml of the same medium was added to both BMDM and DC cultures. Cells were used for experiments at day 7. Both BMDM and DC phenotypes were confirmed by flow cytometry analysis using specific macrophage (CD11b and F4/80) and DC (CD11c and MHC class II) markers (data not shown).

J774A.1 cells overexpressing ASC fused to GFP (J774A.1 ASC-GFP) were generated by lentiviral transduction using a pLEX-MCS-ASC-GFP construct (Addgene, Watertown, MA). Viral particles were produced in HEK293T cells using a Lenti-X packaging single shot kit (Takara Bio, San Jose, CA) according to the manufacturer’s instructions. J774A.1 cells were transduced with 2 ml of HEK293T viral particle-containing supernatant in the presence of 5 µg/ml Polybrene (Sigma-Aldrich, St. Louis, MO) for 18 h. After 72 h, cells expressing ASC-GFP were selected with 2 µg/ml puromycin (Sigma-Aldrich).

All cell types were primed with 1 μg/ml LPS from Escherichia coli 0111:B4 (Sigma-Aldrich) for 4 h to induce the expression of pro–IL-1β and NLRP3 inflammasome precursors. Because intracellular delivery of LPS by electroporation is a widely used method for noncanonical activation of the NLRP3 inflammasome (54, 55), the extracellular LPS was removed by three washes with PBS. Cells were then detached, centrifuged, and resuspended in growth medium for PEF exposure. ATP (5 mM; Sigma-Aldrich) for either 30 or 60 min was used as positive control for caspase-1 activation or IL-1β release, respectively. MCC950 (75 nM; InvivoGen, San Diego, CA) was used to inhibit the NLRP3 inflammasome. The incubation with the inhibitor started 30 min before stimulation of the inflammasome.

In experiments in vitro PEFs were delivered to cells either attached to coverglasses or in suspension in electroporation cuvettes. Both methods have been previously described (7, 56).

Trapezoidal 200-ns pulses were produced by either a custom pulse generation system with an output impedance of 100 Ω, adjustable pulse amplitude (up to 15 kV), duration of 200–1000 ns, and frequency of 1–100 Hz (Pulse Biosciences, Hayward, CA) (57) or from an Avtech AVOZ-D2-B-ODA generator (Avtech Electrosystems, Ottawa, ON, Canada). The 200-ns pulse waveforms from each generator are shown in Fig. 1A. To deliver trains of 100-μs pulses (rectangular), we used an ECM 830 square wave electroporation system (BTX/Harvard Apparatus, Holliston, MA).

FIGURE 1.

nsPEF exposure setups. (A) Representative 200-ns waveforms generated either by the Pulse Biosciences generator used for experiments in cuvette (left graph) or by the Avtech AVOZ-D2-B-ODA used in time-lapse microscopy experiments (right graph). (B) Setup for nsPEF exposure in time-lapse microscopy experiments. (Bi) Parallel tungsten wire electrodes mounted on a pipette tip. (Bii) Electrode system positioned at the bottom of the cover glass chamber on the microscope stage. (Biii) Position of the nsPEF-delivering electrodes (E1 and E2) relative to the exposed cells (red rectangle) and sham control cells (white rectangle). (C) Setup for the in vivo nsPEF stimulation. (Ci) Mouse skin trapped between the silicon support and the ring through which the electrode array was inserted into the skin. (Cii) Needle electrode array. (Ciii) Two-dimensional simulation of the electric field in the skin when 15 kV are applied across the electrode array.

FIGURE 1.

nsPEF exposure setups. (A) Representative 200-ns waveforms generated either by the Pulse Biosciences generator used for experiments in cuvette (left graph) or by the Avtech AVOZ-D2-B-ODA used in time-lapse microscopy experiments (right graph). (B) Setup for nsPEF exposure in time-lapse microscopy experiments. (Bi) Parallel tungsten wire electrodes mounted on a pipette tip. (Bii) Electrode system positioned at the bottom of the cover glass chamber on the microscope stage. (Biii) Position of the nsPEF-delivering electrodes (E1 and E2) relative to the exposed cells (red rectangle) and sham control cells (white rectangle). (C) Setup for the in vivo nsPEF stimulation. (Ci) Mouse skin trapped between the silicon support and the ring through which the electrode array was inserted into the skin. (Cii) Needle electrode array. (Ciii) Two-dimensional simulation of the electric field in the skin when 15 kV are applied across the electrode array.

Close modal

In time-lapse microscopy experiments, cells were maintained at laboratory room temperature (RT; 22 ± 2°C) on the stage of a Leica TCS SP8 laser scanning confocal microscope (Leica, Buffalo Grove, IL) in cell culture medium. nsPEFs (0–100, 200-ns pulses, 10 Hz, 0.9 MV/m) were delivered to cells by placing a pair of parallel tungsten wire electrodes with a 60-μm interelectrode gap at the bottom of the well (Fig. 1Bi, ii) (30, 58). Biological responses in unexposed (sham) and exposed cells were measured in cells outside and within the nsPEF-delivering electrodes, respectively (Fig. 1Biii).

For exposure of cells in cuvettes, samples were resuspended at 1.2–2 × 106 cells/ml in the growth medium, loaded in 1-mm gap electroporation cuvettes (BioSmith, San Diego, CA), and subjected to either nsPEFs (0–300 pulses, 200 ns, 10 Hz, 0.9 MV/m) or sham exposure at RT.

To treat the skin in vivo, mice were anesthetized by inhalation of 3% isoflurane in oxygen (Patterson Veterinary, Devens, MA). Pulses of 200 ns (50–200, 15 kV, 4 Hz) were applied by sandwiching the skin of the mouse between a flat round silicon stage and a plastic ring into which a needle electrode array made of two parallel rows of six needles, 4.5 mm apart, was inserted through the skin into the underneath silicon support (Fig. 1Ci, ii). To ensure an efficient electrical continuity, electrodes were covered with Vaseline. The electric field was calculated by two-dimensional numerical simulations using a finite element analysis software COMSOL Multiphysics, release 5.0 (COMSOL, Stockholm, Sweden). The electric field distribution was modeled under static conditions and the electrical conductivity was assigned to 0.148 S/m (https://itis.swiss/virtual-population/tissue-properties/database/low-frequency-conductivity/). Fig. 1Ciii shows that the electric field is homogeneous in the area bounded by the two rows of electrodes and its value is 3 MV/m for 15 kV applied. Animals in the sham control group underwent anesthesia and the probe insertion procedure but no nsPEF delivery.

The pulse amplitude and shape were monitored in all experiments using a WaveSurfer 3034z oscilloscope (Teledyne Lecroy, Chestnut Ridge, NY).

Immediately after PEF application, cell samples were plated in triplicate at 0.03 × 106 cells/well in black wall 96-well plates (Thermo Fisher Scientific), and viability was measured by a Presto Blue assay after 24 h (Thermo Fisher Scientific). For the ELISA assays, cell samples were seeded at 0.6 × 106 cells/well in 48-well plates immediately after PEFs. In 1 h, the supernatant was harvested, centrifuged at 200 × g for 5 min at 4°C, and IL-1β was measured using a mouse-specific ELISA kit according to the manufacturer’s instructions (88-7013-88; Thermo Fisher Scientific) (59–62). In experiments using high K+ solutions, cells were resuspended in high K+ Tyrode’s solution (140 mM KCl, 2 mM CaCl2, 1.5 mM MgCl2, 10 mM glucose, 25 mM HEPES) or, as control, regular Tyrode’s solution (140 mM NaCl, 5.4 mM KCl, 2 mM CaCl2, 1.5 mM MgCl2, 10 mM glucose, 25 mM HEPES), and supernatant was assessed for IL-1β concentration after 1 h as above described. TNF-α was also measured 1 h after treatment using a mouse-specific ELISA kit (887324-22; Thermo Fisher Scientific) (61, 62).

The Western blot procedure was described previously (7, 63). Either 30 min (for caspase-1 and caspase-11) or 1 h (GSDMD and caspase-3) after nsPEF treatment, cells were washed twice with ice-cold PBS and lysed for 30 min in a buffer containing 20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1 mM Na2EDTA, 1 mM EGTA, 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM β-glycerophosphate, 1 mM Na3VO4, 1 µg/ml leupeptin (Cell Signaling Technology, Danvers, MA) and 1 mM PMSF (Sigma-Aldrich) added immediately before use. For IL-1β detection both supernatants and cell lysates were collected at 1 h after nsPEFs. Supernatants were concentrated 10 times using 10-kDa centrifugal filter devices (Thermo Fisher Scientific). Protein concentration was measured by a bicinchoninic acid assay (Thermo Fisher Scientific), and 20 μg of lysate (or supernatant) per sample was separated by SDS-PAGE and transferred onto polyvinylidene difluoride membranes. Membranes were bathed in 5% BSA TBS with 0.1% Tween (TBST) solution for 1 h at RT and then incubated overnight at 4°C and for 1 h at RT with primary and secondary Abs, respectively. Anti–caspase-1 (p20) (AG-20B-0042-C100; Adipogen, San Diego, CA) (64–66) was used at 1:1000 and 1:2500 dilutions for BMDM and skin lysates, respectively. Anti-GSDMD (ab209845; Abcam, Cambridge, MA) (64), anti–IL-1β (AF-401-NA; R&D Systems, Minneapolis, MN) (59, 64, 65, 67, 68), anti–caspase-3 (9662; Cell Signaling Technology) (64), anti–caspase-11 (NB120-10454; Novus Biologicals, Englewood, CO) (69), and anti-vinculin primary Abs (ab129002; Abcam) were used at 1:1000 dilution. A rabbit anti-mouse IgG (ab6728; Abcam) was used to detect caspase-1, and a goat anti-rabbit IgG (7074, Cell Signaling Technology) was used for GSDMD, caspase-3, and vinculin detection. IL-1β and caspase-11 were detected using a rabbit anti-goat IgG (HAF017; Novus Biologicals) and goat anti-rat IgG (7077; Cell Signaling Technology) Ab, respectively. Images were captured with a ChemiDoc MP imaging system (Bio-Rad, Hercules, CA).

Quantification of band intensities was conducted in ImageJ (ImageJ-win64). The fraction of the cleaved protein (IL-1β, caspase-1, and GSDMD) was calculated as K = 100 × S/(S + L), where L and S are the intensities of the full-length protein and of the cleaved fragment, respectively.

For K+ efflux measurements, cells were loaded with ion potassium green-2 (IPG-2) AM (ION Biosciences, San Marcos, TX). IPG-2 AM is a membrane-permeant form of the K+-sensitive fluorescent dye IPG-2 (IPG-2 fluorescence increases in the presence of K+ ions, proportional to the K+ concentration). J774A.1 cells were plated the night before the experiment at 0.1 × 106 cells/well in a cover glass chamber whereas BMDMs were detached and seeded at 0.5 × 106 cells/well on day 7 of differentiation. Cells were loaded for 10 min at 37°C in cell culture medium containing 1 µM IPG-2 AM, then washed and placed under the microscope for the recording. Recording started 25 s before nsPEF exposure (0–100 pulses, 200 ns, 0.9 MV/m, 10 Hz). Images were captured every 5 s for 10 min (125 frames). IPG-2 AM was excited with a blue 488-nm laser through a ×63, water-immersion, epifluorescence objective, and emission of the dye was detected between 525 and 575 nm. To report as accurately as possible the changes in intracellular concentrations of K+, we evaluated volume changes resulting from PEF-induced swelling (Vt/V0, where Vt is the volume at any time t, and V0 is the volume at the beginning of the recording). For a perfectly spherical cell that swells or shrinks symmetrically in all directions, the volume change is equal to the area change in the mid-cell slice raised to the 1.5 power, that is Vt/V0 = (At,mid-cell/A0,mid-cell)1.5, where At,mid-cell is the area of the mid-cell slice at time t, and A0,mid-cell is the initial area of the mid-cell slice.

The actual volume/area relationship for both J774A.1 cells and BMDMs was extracted from measurements on pulse-exposed cells in high K+ (140 mM) buffer solution. Matching (approximately) intracellular and extracellular K+ concentrations minimizes the K+ diffusion from the cell that would otherwise occur down the intracellular–extracellular K+ concentration gradient, so that fluorescence changes are only due to volume changes (dilution resulting from swelling). By monitoring the area and volume (from z stacks) over time, we were able to extract the relationship between volume and area: Vt/V0 = (At,mid-cell/A0,mid-cell)x.

For J744A.1 cells, which were exposed to PEF while attached to the cover glass chamber, for 25 and 50 pulses, x = 1.5 and 1.1, respectively. For BMDMs, which were exposed in suspension, for both 25 and 100 pulses, x = 1.1.

To correct fluorescence intensities for volume changes, we used those expressions to convert area change ratio measurements to volume change ratios, so that a volume change is not interpreted as a change in concentration of the indicator material. This procedure allows quantitative accounting for total material transport across the cell membrane.

To convert IPG-2 fluorescence to intracellular K+ concentration, we generated two fluorescence calibration curves (Supplemental Fig. 1) from solutions of the tetramethylammonium salt of IPG-2 (IPG-2 TMA+ salt, ION Biosciences), one for IPG-2 fluorescence as a function of [IPG-2] (0–40 µM) with constant [K+] (140 mM), the other for IPG-2 fluorescence as a function of [K+] (0–300 mM) with constant [IPG-2] (5 µM). The slope of the curve in Supplemental Fig. 1A is slope = α([K+]/Kd), where [K+] is the K+ concentration (millimolar), Kd is 18 mM, and α (0.75 arbitrary U/M) is constant at fixed K+ concentration. From this relationship we calculated the intracellular concentration of IPG-2, at rest condition after 10 min of incubation, using the following formula: [IPG-2] = (F/α) × (Kd/[K+] ≅ 9 μM.

Then, to determine the K+ concentration corresponding to any measured IPG-2 fluorescence, a fitting curve was constructed in MATLAB (Supplemental Fig. 1B). In this case, α is not constant but depends on the K+ concentration. Moreover, through experiments in high K+ solution (not shown) we ensured that under these conditions the dye is not leaking out of the cell. In this case [IPG-2] can be considered constant inside the cells in all the experiments. The equation for the curve in Supplemental Fig. 1B is
F=(c1e[K+]k)[IPG2]Kd[K+]+c2.

By solving the equation, we obtained c1 = 0.19, c2 = 25, and k = 60 mM. [K+] at any given time can then be calculated from the fluorescence.

Four- to 6-wk-old BALB/c female mice (The Jackson Laboratory) were housed in groups of five animals in individually ventilated cages under pathogen-free conditions. The flank skin was treated with nsPEFs as described above. As a positive control for the activation of the inflammasome in vivo, animals were injected s.c. with 10 mg/kg LPS. After 1 h, mice were humanly euthanized, and skin samples were collected and snap-frozen in liquid nitrogen. In 24 h, the tissue was weighed and sonicated in lysis buffer to measure IL-1β and TNF-α release by ELISA (Thermo Fisher Scientific) and caspase-1 activation by Western blot analysis as above described. Three independent experiments were conducted with one mouse per each treatment condition at a time.

Data are presented as mean ± SE for n independent experiments with overlap of all data points. Statistical analyses were performed using either a two-tailed t test or a one-way ANOVA, where p < 0.05 was considered statistically significant. Statistical calculations, including data fits and data plotting were carried out in Grapher 11 (Golden Software, Golden, CO) and Prism (GraphPad Software, San Diego, CA).

The effect of PEFs on inflammasome activation was measured using 200-ns pulses in innate immune cells that express the NLRP3 inflammasome, namely macrophages and DCs (Fig. 1). Previous studies reported marked differences in sensitivity of various cell types exposed to nsPEFs (70). Therefore, to identify electric pulse doses causing comparable damage across multiple cell types, we investigated the sensitivity to nsPEFs of J774A.1 macrophages, BMDMs, and DCs (Fig. 2, top panels). We identified 25 and 50 pulses for J774A.1 (Fig. 2A), 25 and 100 pulses for BMDMs (Fig. 2B), and 25 and 200 pulses for DCs (Fig. 2C) as isoeffective doses for 200-ns pulses (0.9 MV/m, 10 Hz) causing 0–10% (low dose) and 35–45% (high dose) cell death at 24 h, respectively. For each cell type the above-indicated pulse numbers are referred to as high dose and low dose throughout this study. Inflammasome activation was monitored by measuring the secretion of IL-1β in cells primed with LPS. For each cell type the higher pulse dose triggered statistically significant IL-1β release as measured by ELISA at 1 h (Fig 2, bottom panels). We further investigated the role of the pulse dose in BMDMs and found that nsPEF doses causing 80% cell death at 24 h, namely 150 pulses (200 ns, 0.9 MV/m, 10 Hz), did not cause IL-1β release (Supplemental Fig. 2A). These results can be explained by the fact that incrementing the number of pulses beyond a certain limit increases the percentage of cells dying immediately of primary necrosis due to irreversible permeabilization of the cell plasma membrane. In this scenario most cells do not have time to activate the inflammasome, whereas at lower doses a higher percentage of cells can repair the electroporation damage and cope with it by activating stress responses, including inflammasome activation. We further confirmed the presence of the active form of IL-1β (p17) in the supernatant of treated cells by Western blot analysis (Supplemental Fig. 2B). Moreover, the release of the inflammasome-independent marker TNF-α was not affected, suggesting that nsPEFs preferentially stimulate IL-1β release (Supplemental Fig. 2C).

FIGURE 2.

nsPEFs trigger IL-1β release in J774A.1 cells, BMDMs, and DCs. (AC) Results are shown for J774A.1 cells (A), BMDMs (B), and DCs (C). Cells were primed by incubation with LPS (1 µg/ml). After 4 h, samples were washed with PBS three times, detached, resuspended in growth medium, and either treated with the indicated numbers of 200-ns pulses (0.9 MV/m, 10 Hz) or subject to a sham exposure. Viability was measured at 24 h after nsPEFs by a Presto Blue assay and is reported as the percentage compared to a sham-exposed parallel control (top graphs). Filled circles identify isoeffective pulse doses causing 0–10% (low dose; 25 pulses) and 35–45% (high dose; 50, 100 and 200 pulses for J774A.1 cells, BMDMs, and DCs, respectively) cell death at 24 h. Bottom graphs report IL-1β measured in the cell supernatant by ELISA 1 h after treatment with the above-indicated isoeffective nsPEF doses. As the positive control for inflammasome induction, cells were treated with 5 mM ATP for 1 h. Means ± SEM are shown; n = 5–16. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 2.

nsPEFs trigger IL-1β release in J774A.1 cells, BMDMs, and DCs. (AC) Results are shown for J774A.1 cells (A), BMDMs (B), and DCs (C). Cells were primed by incubation with LPS (1 µg/ml). After 4 h, samples were washed with PBS three times, detached, resuspended in growth medium, and either treated with the indicated numbers of 200-ns pulses (0.9 MV/m, 10 Hz) or subject to a sham exposure. Viability was measured at 24 h after nsPEFs by a Presto Blue assay and is reported as the percentage compared to a sham-exposed parallel control (top graphs). Filled circles identify isoeffective pulse doses causing 0–10% (low dose; 25 pulses) and 35–45% (high dose; 50, 100 and 200 pulses for J774A.1 cells, BMDMs, and DCs, respectively) cell death at 24 h. Bottom graphs report IL-1β measured in the cell supernatant by ELISA 1 h after treatment with the above-indicated isoeffective nsPEF doses. As the positive control for inflammasome induction, cells were treated with 5 mM ATP for 1 h. Means ± SEM are shown; n = 5–16. *p < 0.05, **p < 0.01, ***p < 0.001.

Close modal

Most inflammasomes comprise members of the NLR family of intracellular pathogen receptors that associate with the ASC adaptor protein to recruit caspase-1. ASC-dependent inflammasome activation is accompanied by rapid relocation of NLRs and ASC into a singular, perinuclear, punctate “speck” structure of ∼1 µm that can be observed with fluorescence microscopy (71). To visualize the formation of ASC specks in response to inflammasome activation, we generated a J774A.1 cell line stably expressing ASC fused to GFP (J774A.1 ASC-GFP). To test the functionality of this cell line, ASC speck formation was monitored in response to extracellular ATP. As expected, ATP triggered the aggregation of ASC in LPS-treated cells whereas it failed to do so in cells that were not primed (Fig. 3A). Notably, ASC specks formed also in response to 200-ns pulses (50 pulses, 0.9 MV/m, 10 Hz) (Fig. 3A). In nsPEF-treated cells, clustering of the ASC adaptor correlated with the activation caspase-1 (Fig. 3B) whereas caspase-11 (Supplemental Fig. 3A) was not affected, suggesting that these brief electrical stimuli activate the canonical inflammasome.

FIGURE 3.

nsPEFs trigger ASC specks formation and caspase-1 activation but not GSDMD cleavage. (A) Representative fluorescence images of LPS-primed J774A.1 cells expressing ASC-GFP and treated with either 5 mM ATP or 50 (200-ns) pulses (0.9 MV/m, 10 Hz). As negative controls, cells were either treated with ATP without priming with LPS or with a sham exposure. Characteristic fluorescence changes from cytosolic diffuse to granular specks are indicated by the white arrows. Scale bar, 100 µm. (B and C) Representative Western blot images and quantifications for caspase-1 and GSDMD cleavage measured in J774A.1 cells at 30 min and 1 h after nsPEFs, respectively. Low dose and high dose correspond to 25 and 50 pulses (200 ns, 0.9 MV/m, 10 Hz) causing 0–10% and 35–45% cell death (Fig. 2A), respectively. Vinculin was used as the housekeeping control. Means ± SEM are shown; n = 3. *p < 0.05, **p < 0.01.

FIGURE 3.

nsPEFs trigger ASC specks formation and caspase-1 activation but not GSDMD cleavage. (A) Representative fluorescence images of LPS-primed J774A.1 cells expressing ASC-GFP and treated with either 5 mM ATP or 50 (200-ns) pulses (0.9 MV/m, 10 Hz). As negative controls, cells were either treated with ATP without priming with LPS or with a sham exposure. Characteristic fluorescence changes from cytosolic diffuse to granular specks are indicated by the white arrows. Scale bar, 100 µm. (B and C) Representative Western blot images and quantifications for caspase-1 and GSDMD cleavage measured in J774A.1 cells at 30 min and 1 h after nsPEFs, respectively. Low dose and high dose correspond to 25 and 50 pulses (200 ns, 0.9 MV/m, 10 Hz) causing 0–10% and 35–45% cell death (Fig. 2A), respectively. Vinculin was used as the housekeeping control. Means ± SEM are shown; n = 3. *p < 0.05, **p < 0.01.

Close modal

Although secretion of IL-1β typically requires formation of membrane pores by GSDMD, we did not measure cleavage of GSDMD in response to nsPEFs (Fig. 3C). Our results are consistent with previous reports of IL-1β secretion through GSDMD-independent pathways controlled by caspase-1, such as exocytosis of secretory lysosomes and microvesicle shedding (72–75).

Altogether, our data show that nanosecond pulses induce ASC speck formation, caspase-1 activation, and unconventional secretion of IL-1β.

Next, we asked whether the damage created by longer pulses such as the 100-μs pulses used in IRE and ECT was also sensed by the inflammasome. For direct comparison with nsPEF experiments, we measured IL-1β release in cells treated with 100-μs pulse doses (0.1 MV/m, 5 Hz) causing either 0–10% (low dose) or 35–45% (high dose) cell death, namely 2 and 15 pulses and 2 and 7 pulses for BMDMs and J774A.1, respectively (Fig. 4, top panels). Compared to the 200-ns pulses used in our nsPEF experiments, 100-μs pulses failed to trigger the release of IL-1β in BMDMs and had a weaker effect in J774A.1 cells (Fig. 4, bottom panels).

FIGURE 4.

One hundred–microsecond pulses are weaker activators of the inflammasome. (A and B) Primed BMDMs (A) and J774A.1 cells (B) were treated with the indicated numbers of 100-μs pulses (0.1 MV/m, 5 Hz) or subject to a sham exposure. Top panels show viability measured by a Presto Blue assay at 24 h posttreatment. Viability is reported as percentage compared to a sham-exposed parallel control; filled circles identify isoeffective 100-μs pulse doses causing 0–10% (low dose, 2 pulses for both BMDMs and J774A.1 cells) and 35–45% (high dose; 15 and 7 pulses for BMDMs and J774A.1 cells, respectively) cell death. Bottom graphs report IL-1β measured in the cell supernatant by ELISA 1 h after treatment with the above-indicated isoeffective low and high pulse doses. Means ± SEM are shown; n = 6–10. *p < 0.05, **p < 0.01, ***p < 0.001. n.s., not significant.

FIGURE 4.

One hundred–microsecond pulses are weaker activators of the inflammasome. (A and B) Primed BMDMs (A) and J774A.1 cells (B) were treated with the indicated numbers of 100-μs pulses (0.1 MV/m, 5 Hz) or subject to a sham exposure. Top panels show viability measured by a Presto Blue assay at 24 h posttreatment. Viability is reported as percentage compared to a sham-exposed parallel control; filled circles identify isoeffective 100-μs pulse doses causing 0–10% (low dose, 2 pulses for both BMDMs and J774A.1 cells) and 35–45% (high dose; 15 and 7 pulses for BMDMs and J774A.1 cells, respectively) cell death. Bottom graphs report IL-1β measured in the cell supernatant by ELISA 1 h after treatment with the above-indicated isoeffective low and high pulse doses. Means ± SEM are shown; n = 6–10. *p < 0.05, **p < 0.01, ***p < 0.001. n.s., not significant.

Close modal

It has been demonstrated in several systems that K+ efflux is both necessary and sufficient for NLRP3 inflammasome activation (76–78). For example, the NLRP3 inflammasome can be activated by incubation of primed macrophages in a K+-free medium, and activation can be blocked by high concentrations of extracellular K+ (77, 79). Inflammasome activation by nsPEFs could result from K+ efflux caused by permeabilization of the cell plasma membrane.

To investigate the effect of nsPEFs on K+ efflux, both J774A.1 cells and BMDMs were loaded with the K+ indicator IPG-2 AM, treated with either a low- or a high-pulse dose, and intracellular fluorescence changes were measured over time. Fig. 5A shows that nsPEF exposure causes an immediate decrease in IPG-2 intracellular fluorescence. IPG-2·K+ fluorescence measurements were corrected for both photobleaching and PEF-induced cell swelling, as described in Materials and Methods, so this reduction in IPG-2 fluorescence can be attributed to K+ leakage from the cells due to nsPEF exposure. Interestingly, except for J774A.1 cells treated with 50 pulses, all cell samples recovered the initial (resting) intracellular K+ concentration within a few minutes, suggesting that the membrane damage created by nsPEFs was repaired.

FIGURE 5.

Blocking nsPEF-induced K+ efflux reduces but does not block IL-1β release. (A) BMDMs (left panel) and J744A.1 (right panel) loaded with the K+ indicator IPG2-AM were treated with nsPEFs (200 ns, 0.9 MV/m, 10 HZ) and fluorescence changes were monitored for 10 min. Cells were treated with isoeffective pulse doses causing either 0–10% (25 pulses for both BMDMs and J774A.1 cells) or 35–45% (100 and 50 pulses for BMDMs and J774A.1 cells, respectively) cell death at 24 h. The pulse treatment was delivered 25 s into the recording (dashed line). Each point represents the mean ± SEM; n > 25 cells (three independent experiments). (B) J774A.1 cells were resuspended in Tyrode’s solution containing 5 mM KCl (white bars) or in high K+ Tyrode’s solution containing 140 mM KCl (gray bars). Samples were treated with either low (25 pulses) or high (50 pulses) 200-ns pulse doses (0.9 MV/m, 10 Hz) and IL-1β was measured by ELISA at 1 h posttreatment. Means ± SEM are shown; n = 3. *p < 0.05, **p < 0.01.

FIGURE 5.

Blocking nsPEF-induced K+ efflux reduces but does not block IL-1β release. (A) BMDMs (left panel) and J744A.1 (right panel) loaded with the K+ indicator IPG2-AM were treated with nsPEFs (200 ns, 0.9 MV/m, 10 HZ) and fluorescence changes were monitored for 10 min. Cells were treated with isoeffective pulse doses causing either 0–10% (25 pulses for both BMDMs and J774A.1 cells) or 35–45% (100 and 50 pulses for BMDMs and J774A.1 cells, respectively) cell death at 24 h. The pulse treatment was delivered 25 s into the recording (dashed line). Each point represents the mean ± SEM; n > 25 cells (three independent experiments). (B) J774A.1 cells were resuspended in Tyrode’s solution containing 5 mM KCl (white bars) or in high K+ Tyrode’s solution containing 140 mM KCl (gray bars). Samples were treated with either low (25 pulses) or high (50 pulses) 200-ns pulse doses (0.9 MV/m, 10 Hz) and IL-1β was measured by ELISA at 1 h posttreatment. Means ± SEM are shown; n = 3. *p < 0.05, **p < 0.01.

Close modal

Next, we asked whether blocking K+ efflux impairs inflammasome activation by nsPEFs. J774A.1 cells were exposed to nsPEFs (25 or 50 pulses, 200 ns, 0.9 MV/m, 10 Hz) in either regular Tyrode’s or high K+ Tyrode’s solution, and IL-1β release was measured after 1 h. Fig. 5B shows a statistically significant reduction in IL-1β release from nsPEF-treated cells in high extracellular [K+]. As expected, the high K+ extracellular solution completely blocked IL-1β secretion in response to ATP.

To assess the relevance of the NLRP3 inflammasome sensor, we initially tested whether IL-1β release in response to nsPEFs was affected by treatment with the NLRP3 inhibitor MCC950 (68). MCC950 (75 nM) blocked IL-1β release in both LPS-primed BMDMs and J774A.1 cells (Fig. 6A). In addition, MCC950 rescued 200-ns-treated J774A.1 cells from death (Fig. 6B), whereas it did not protect 100-μs-treated cells (Fig. 6C). These results suggested that the release of IL-1β and cell death in response to nsPEFs were both dependent on the activation of the NLRP3 inflammasome. Recently, Schneider et al. (72) reported that the NLRP3 inflammasome can drive GSDMD-independent secondary pyroptosis through activation of apoptotic caspases. However, we did not measure caspase-3 activation (Supplemental Fig. 3B), suggesting that this cell death pathway is not engaged by nsPEFs in J774A.1 cells.

FIGURE 6.

MCC950 blocks IL-1β release and rescued cells treated with nsPEFs from death. (A) BMDMs (left panel) and J774A.1 cells (right panel) were primed by incubation with LPS (1 µg/ml) for 4 h. Treatment with the NLRP3 inflammasome inhibitor MCC950 (75 nM) started 30 min before the indicated nsPEF treatments (200 ns, 0.9 MV/m, 10 Hz) and IL-1β was measured in 1 h. (B and C) J774A.1 cells were primed and then incubated with MCC950 (75 nM) for 30 min before nsPEFs (200 ns, 0.9 MV/m, 10 Hz) (B) or 100-μs (0.1 MV/m, 5 Hz) (C) pulse treatments. Viability was measured in 24 h. Means ± SEM are shown; n = 3–8 (A), 8 (B), and 6 (C). *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 6.

MCC950 blocks IL-1β release and rescued cells treated with nsPEFs from death. (A) BMDMs (left panel) and J774A.1 cells (right panel) were primed by incubation with LPS (1 µg/ml) for 4 h. Treatment with the NLRP3 inflammasome inhibitor MCC950 (75 nM) started 30 min before the indicated nsPEF treatments (200 ns, 0.9 MV/m, 10 Hz) and IL-1β was measured in 1 h. (B and C) J774A.1 cells were primed and then incubated with MCC950 (75 nM) for 30 min before nsPEFs (200 ns, 0.9 MV/m, 10 Hz) (B) or 100-μs (0.1 MV/m, 5 Hz) (C) pulse treatments. Viability was measured in 24 h. Means ± SEM are shown; n = 3–8 (A), 8 (B), and 6 (C). *p < 0.05, **p < 0.01, ***p < 0.001.

Close modal

To further investigate the role of the NLRP3 inflammasome sensor, we used BMDMs from NLRP3-KO mice. As expected, the lack of NLRP3 expression completely blocked IL-1β release in response to ATP (Fig. 7). However, contrary to our results with the MCC950 inhibitor, the lack of NLRP3 expression did not impair IL-1β release in response to nsPEFs. These results suggested that either nsPEFs trigger another inflammasome sensor in NLRP3-KO cells or MCC950 has an off-target effect that impacts both IL-1β release and viability. To distinguish between these two scenarios, we measured the effect of MCC950 on nsPEF-treated NLRP3-KO cells. Notably, MCC950 did not block IL-1β release in NLRP3-deficient cells, ruling out the possibility of an off-target effect (Fig. 7).

FIGURE 7.

Relevance of NLRP3 expression for nsPEF-induced IL-1β release. NLRP3-KO BMDMs were primed by incubation with LPS (1 µg/ml). After 4 h, cells were treated with a high 200-ns pulse dose (100 pulses, 0.9 MV/m, 10 Hz) and IL-1β was measured by ELISA in 1 h. In selected experiments, treatment with the NLRP3 inflammasome inhibitor MCC950 (75 nM) started 30 min before nsPEF treatments (filled bars). As the control for NLRP3 inflammasome induction, cells were treated with 5 mM ATP for 1 h. Means ± SEM are shown; n = 3–8. **p < 0.01.

FIGURE 7.

Relevance of NLRP3 expression for nsPEF-induced IL-1β release. NLRP3-KO BMDMs were primed by incubation with LPS (1 µg/ml). After 4 h, cells were treated with a high 200-ns pulse dose (100 pulses, 0.9 MV/m, 10 Hz) and IL-1β was measured by ELISA in 1 h. In selected experiments, treatment with the NLRP3 inflammasome inhibitor MCC950 (75 nM) started 30 min before nsPEF treatments (filled bars). As the control for NLRP3 inflammasome induction, cells were treated with 5 mM ATP for 1 h. Means ± SEM are shown; n = 3–8. **p < 0.01.

Close modal

Taken together, our results show that the nsPEF damage is sensed intracellularly by the NLRP3 inflammasome and that other sensors triggering IL-1β release can be activated when the NLRP3 is not expressed.

The activation of the inflammasome in vivo in response to nsPEFs was monitored in the skin of mice, which is easy to access and highly infiltrated by innate immune cells. Electric pulses were delivered to the mouse’s flank skin using a needle electrode array as described in Materials and Methods. To address the inherent uncertainty of the extrapolation of nsPEF efficacy from in vitro to in vivo, we tested three pulse number doses delivered at 3 MV/m, an electric field that is widely used in preclinical and clinical settings (80–83). At 1 h after nsPEFs, all tested pulse doses (50, 100, and 200 pulses, 200 ns, 3 MV/m, 4 Hz) activated the inflammasome as measured by caspase-1 activation (Fig. 8A) and increased the concentration of IL-1β in skin tissue lysates (Fig. 8B). Notably TNF-α, another proinflammatory cytokine produced by macrophages, was not affected by the pulse treatment, suggesting that nsPEFs selectively induce the release of IL-1β.

FIGURE 8.

Two hundred–nanosecond pulses activate the inflammasome in mouse skin. The skin of mice (three animals per group) was either treated with nsPEFs (50, 100, and 200 pulses, 200 ns, 3 MV/m, 4 Hz) or left untreated. As a positive control for inflammasome activation, animals were injected intradermally with LPS (10 mg/kg). At 1 h posttreatment, all mice were humanely euthanized to collect the skin. (A and B) Tissue lysates were analyzed for caspase-1 activation (A) and IL-1β and TNF-α concentration (B). Symbols identify animal per condition done in parallel. Means ± SEM are shown; n = 3. *p < 0.05, **p < 0.01.

FIGURE 8.

Two hundred–nanosecond pulses activate the inflammasome in mouse skin. The skin of mice (three animals per group) was either treated with nsPEFs (50, 100, and 200 pulses, 200 ns, 3 MV/m, 4 Hz) or left untreated. As a positive control for inflammasome activation, animals were injected intradermally with LPS (10 mg/kg). At 1 h posttreatment, all mice were humanely euthanized to collect the skin. (A and B) Tissue lysates were analyzed for caspase-1 activation (A) and IL-1β and TNF-α concentration (B). Symbols identify animal per condition done in parallel. Means ± SEM are shown; n = 3. *p < 0.05, **p < 0.01.

Close modal

We have presented evidence in the current study that the damage created by PEFs activates the NLRP3 inflammasome, initiating IL-1β release. These effects are better seen when using nanosecond pulses compared with equivalent treatments with longer (100-µs) electric pulses. Experiments using the NLRP3 inhibitor MCC950 also highlight the relevance of the pulse duration. Preincubation with the inhibitor protects J774A.1 cells from 200-ns pulses but not from 100-µs pulses (Fig. 7B, 7C). Notably, the effect of the inhibitor is best seen at doses that also cause IL-1β secretion.

Why are shorter pulses more effective at activating the inflammasome? Unlike electroporative pulses in the microsecond and millisecond range, which produce voltages across the cytoplasmic membrane with no direct effects on the cell interior, pulses with rise and fall times faster than the membrane charging time constant (<1 μs for tightly packed cells in a tissue [84]) generate an electric field in the cytoplasm and across the intracellular membranes of the nucleus, mitochondria, and other organelles (85, 86). For nanosecond pulses the cytoplasmic membrane appears electrically transparent, and, when the pulse amplitude is large enough, depolarizing and porating electric fields can appear across internal organelles and dissipate before charges redistribute on the plasma membrane to shield the cell interior from the external electric field. Multiple studies have shown that nsPEFs permeabilize intracellular structures (28, 29, 43, 44, 46, 86), including mitochondria (43, 87), ER (28, 29, 44–46, 88), and nuclei (89).

This organelle-penetrating property of nsPEFs may explain why nsPEF damage is sensed by inflammasomes, the guardians of cytosolic integrity. Mitochondria contain several damage-associated molecular patterns for inflammasome activation (90). For instance, oxidized mitochondrial DNA released into the cytosol upon mitochondrial dysfunction is a potent NLRP3 inflammasome activator (91–93). Mitochondrial DNA also activates inflammasomes that use absent in melanoma 2 (AIM2) as a sensing component (94). Notably, AIM2 can also sense alteration of nuclear envelope integrity by detecting nuclear DNA in the cytosol (95). Our results showing that the lack of NLRP3 expression did not impair nsPEF-induced IL-1β release (Fig. 8) indicate that nsPEFs generate a set of stimuli that can trigger more than one inflammasome sensor. Although it has been reported that nsPEFs cause cytochrome c release (44, 96, 97) and reduction of mitochondrial membrane potential (43), no one yet has looked for evidence of mitochondrial DNA in the cytoplasm. More research is needed to define the minimal requirements needed to trigger the inflammasome with nsPEFs. These short pulses initiate multiple interrelated cellular events, including membrane permeabilization, mitochondrial damage, and ROS production, complicating the distinction between bystander and causative events.

Unexpectedly, we found that GSDMD is not activated by nsPEFs, at least not at 1 h posttreatment, where we measured significant IL-1β release in the supernatant of treated cells (Fig. 3C). IL-1β lacks a secretory signal and is synthetized as a precursor, pro–IL-1β, which requires cleavage for activation. Pro–IL-1β is cleaved by caspase-1, which enables its secretion (38, 39). Caspase-1 also cleaves GSDMD, triggering the formation of GSDMD pores in the plasma membrane and pyroptosis (38, 39). Cell lysis and active IL-1β secretion are temporally associated (98–100), and therefore IL-1β is often proposed to be passively released during cell rupture (101). However, a growing number of studies report IL-1β release from living cells (59, 102, 103), and various cell lysis–independent secretion mechanisms have been reported, including through GSDMD pores, secretory lysosomes, and microvesicle shedding (59, 72–75).

Lipid electropores (nanopores) are too small to conduct IL-1β (45, 104). However, Pakhomova et al. (105) have reported that an initial pulse-induced opening of nanopores minimally permeable to propidium is followed by an abrupt propidium inflow that can occur 10s of minutes after the exposure. This increase in permeability was shown to be due to a sudden pore dilation or perhaps de novo opening of larger pores (105). These events are followed by characteristic and irreversible changes in cell appearance (granulation, pyknosis, loss of differential interference contrast, and volume changes), and could be regarded as a sign of cell death. IL-1β could then be released through either dilated nanopores or during cell lysis.

One more intriguing hypothesis is that the release of IL-1β, which localizes to the plasma membrane after maturation (73) and can also be packed into lysosomes (75), occurs in connection with the active membrane repair initiated to restore membrane integrity after electroporation. Indeed, our results suggest that cells repair the plasma membrane damage, as in most cases cell samples recovered the resting intracellular K+ concentration within a few minutes. Membrane repair mechanisms involve outward vesiculation or shedding of damaged membranes (106, 107), and exocytosis of lysosomes (108) and inflammasome activity correlate with enhanced secretion of extracellular vesicles containing IL-1β (74). Notably, the repair by lysosomal exocytosis was demonstrated in epithelial cells and fibroblasts treated by millisecond and microsecond pulses (109), while we found that annexin V, a protein that self-assembles into lattices and is involved in patch resealing through vesicle fusion, is activated by nsPEFs and contributes to the membrane resealing (110).

Another key question is whether inflammasome activation and cell death are linked or independent processes in nsPEF-treated cells. The lack of GSDMD cleavage suggests that inflammasome activation is not triggering bona fide pyroptosis, but then why is cell death blocked by the NLRP3 inflammasome inhibitor MCC950? Several studies have shown that GSDMD-deficient cells are still susceptible to inflammasome-driven cell death via activation of apoptotic caspases (72, 111–113). However, we did not measure caspase-3 activation (Supplemental Fig. 3B), suggesting that nsPEFs engage an alternative still unknown lytic cell death pathway. Further experiments are required to determine whether nsPEF-induced cell death is dependent on caspase-1 activation and on the expression of other GSDMs such as GSDME.

Finally, what are the implications of inflammasome activation for PEF applications?

Inflammasome-mediated inflammation can be beneficial for PEF applications such as tumor ablation. Our results, showing higher efficiency of short pulses at inducing the inflammasome, suggest that the immunogenicity of the PEF treatment may be increased by shortening the pulse duration. For instance the release of inflammatory cytokines such as IL-1β and damage-associated molecular patterns can create an immune-adjuvanted environment within which tumor Ags are released. Although longer pulses of 100 μs are routinely used in the clinic for cancer applications, including IRE and ECT, treatment with nsPEFs is a newer technology that has been investigated mostly in preclinical settings using pulses ranging from 100 to 300 ns (114). Multiple studies, including ours, have shown that nsPEFs induce an immunogenic form of cell death that assists tumor eradication and prevents the formation of new tumors (5, 7, 34, 35, 82, 115–120). Although the inflammasome has been investigated most often in the context of inflammation and innate immunity, there is growing evidence that inflammasome products, particularly IL-1 family cytokines, have an essential role in stimulating adaptive immune responses, including those involved in anticancer immunosurveillance (121). For instance, Ghiringhelli et al. (122) reported that the activation of the NLRP3 inflammasome in DCs is critical for the priming of IFN-γ–producing, tumor Ag–specific CD8 T cells.

Conversely, an enhanced inflammatory response may be harmful to other PEF applications such as cardiac ablation for the treatment of atrial fibrillation (12, 13). Cardiomyocytes, cardiac fibroblasts, and cardiac macrophages express inflammasome sensors including NLRP3 (123), AIM2 (124), and NLRC4 (124), suggesting that ablation of cardiac tissue may potentially trigger inflammation via inflammasome activation. Notably, an increase in inflammatory markers such as high-sensitivity C-reactive protein (hs-CRP), troponin-T, and fibrinogen are predictive of early recurrence of atrial arrhythmias after radiofrequency ablation (125). Hence, anti-inflammatory agents such as corticosteroids and colchicine are currently being tested for their efficacy at preventing atrial fibrillation recurrence (126–129).

Current research is focused on identifying the stimuli that trigger the inflammasome as well as the executors of cell death in nsPEF-treated cells. A better understanding of these responses will facilitate optimization of the treatment conditions to achieve the desired outcome, whether it is exploiting the immune adjuvant effects of PEFs or avoiding them.

The authors have no financial conflicts of interest.

This work was supported by a grant from Pulse Biosciences, Inc. (to P.T.V. and C.M.) and by the Biomedical Sciences Program, Old Dominion University.

The online version of this article contains supplemental material.

AIM2

absent in melanoma 2

ASC

apoptosis-associated speck-like protein containing a CARD

BMDM

bone marrow–derived macrophage

DC

dendritic cell

ECT

electrochemotherapy

ER

endoplasmic reticulum

GET

gene electrotransfer

GSDMD

gasdermin D

HEK

human embryonic kidney

IPG-2

ion potassium green-2

IRE

irreversible electroporation

KO

knockout

NLR

nucleotide-binding domain, leucine-rich repeat–containing receptor

NLRP3

NLR family, pyrin domain containing 3

nsPEF

nanosecond PEF

PEF

pulsed electric field

ROS

reactive oxygen species

RT

room temperature

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