Two Modes of Cell Death Caused by Exposure to
Nanosecond Pulsed Electric Field
Olga N. Pakhomova*, Betsy W. Gregory, Iurii Semenov, Andrei G. Pakhomov
Frank Reidy Research Center for Bioelectrics, Old Dominion University, Norfolk, Virginia, United States of America
High-amplitude electric pulses of nanosecond duration, also known as nanosecond pulsed electric field (nsPEF), are a novel
modality with promising applications for cell stimulation and tissue ablation. However, key mechanisms responsible for the
cytotoxicity of nsPEF have not been established. We show that the principal cause of cell death induced by 60- or 300-ns
pulses in U937 cells is the loss of the plasma membrane integrity (‘‘nanoelectroporation’’), leading to water uptake, cell
swelling, and eventual membrane rupture. Most of this early necrotic death occurs within 1–2 hr after nsPEF exposure. The
uptake of water is driven by the presence of pore-impermeable solutes inside the cell, and can be counterbalanced by the
presence of a pore-impermeable solute such as sucrose in the medium. Sucrose blocks swelling and prevents the early
necrotic death; however the long-term cell survival (24 and 48 hr) does not significantly change. Cells protected with
sucrose demonstrate higher incidence of the delayed death (6–24 hr post nsPEF). These cells are more often positive for the
uptake of an early apoptotic marker dye YO-PRO-1 while remaining impermeable to propidium iodide. Instead of swelling,
these cells often develop apoptotic fragmentation of the cytoplasm. Caspase 3/7 activity increases already in 1 hr after
nsPEF and poly-ADP ribose polymerase (PARP) cleavage is detected in 2 hr. Staurosporin-treated positive control cells
develop these apoptotic signs only in 3 and 4 hr, respectively. We conclude that nsPEF exposure triggers both necrotic and
apoptotic pathways. The early necrotic death prevails under standard cell culture conditions, but cells rescued from the
necrosis nonetheless die later on by apoptosis. The balance between the two modes of cell death can be controlled by
enabling or blocking cell swelling.
Citation: Pakhomova ON, Gregory BW, Semenov I, Pakhomov AG (2013) Two Modes of Cell Death Caused by Exposure to Nanosecond Pulsed Electric Field. PLoS
ONE 8(7): e70278. doi:10.1371/journal.pone.0070278
Editor: Maria Rosaria Scarfi, National Research Council, Italy
Received April 12, 2013; Accepted June 19, 2013; Published July 23, 2013
Copyright: ? 2013 Pakhomova et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: The study was supported by RO1CA125482 from the National Cancer Institute and RO1GM088303 from the National Institute of General Medical
Sciences. 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.
* E-mail: firstname.lastname@example.org
Cell death induction by nsPEF has recently been proposed as a
new therapeutic modality to ablate cancer. Cytotoxic efficiency of
nsPEF against multiple cancer types has been demonstrated both
in vitro [1–7] and in vivo [1,5,6,8–10]. Interestingly, cancer cells
reportedly were more vulnerable than matching normal cell lines
. Contemplated advantages of nsPEF over other ablation
methods include higher probability of complete elimination of
cancer cells; reduced collateral damage to healthy tissues and
extracellular matrix; relative simplicity of the treatment; inhibition
of angiogenesis; minimal systemic side effects; and fast recovery.
The exact mechanisms responsible for nsPEF cytotoxicity have
been a subject of numerous studies [1–4,11–17], but nonetheless
remain elusive. Early studies in this area have noted fast and
massive externalization of phosphatidylserine in nsPEF-treated
cells, which was interpreted as a sign of apoptosis and a proof that
apoptosis is the prevailing or even the sole mode of cell death after
nsPEF [4,18,19]. As a result, the vast majority of studies into
nsPEF-induced cell death focused solely on the apoptotic death
pathway. Indeed, various types of cells exposed at lethal nsPEF
doses expressed such manifestations of apoptosis as caspase
activation, poly-ADP ribose polymerase (PARP) cleavage, cyto-
chrome C release into the cytoplasm, and internucleosomal DNA
fragmentation [4,6,12,14]. The only type of necrosis considered in
these studies was the so-called ‘‘secondary necrosis’’ (a final cell
destruction following the apoptotic process in vitro).
However, the validity of PS externalization as a sign of apoptosis
has been challenged with understanding that nsPEF opens pores in
the cell plasma membrane. These pores could provide passage for
calcium ions into the cell, causing scramblase activation and fast
PS externalization [20,21]. In addition, the pores can serve as a
lipid-water interface pathway from the cell inside to the outside,
allowing for calcium-independent lipid scrambling by a hypothet-
ical ‘‘lateral drift’’ mechanism [22,23]. In either case, the fast onset
of PS externalization (,1 sec after nsPEF) suggested that this effect
is not necessarily a step in the organized apoptotic process. These
findings suggested that the conclusion about apoptosis prevalence
after nsPEF (which was based primarily on the PS externalization
data) may need to be revisited and revised.
Concurrently, several groups reported that nsPEF-treated cells
typically swell [17,24–27], which is a morphological hallmark of
necrosis. Permeabilization of the cell plasma membrane was
identified as the principal cause of the necrotic cell transformation
[17,24]. Recently we reported that a significant fraction of nsPEF-
treated Jurkat and U937 cells died at intervals much shorter than
what it typically takes to complete the apoptotic process . The
number of ‘‘live’’ cells (impermeable to Trypan blue) decreased
almost twofold already at 2 hr after the nsPEF exposure (600
pulses, 10 ns, 100 kV/cm), but the internucleosomal DNA
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fragmentation developed only in 3 hr; hence, a large fraction of
cells died before reaching this apoptotic step. Significant cell death
could be observed in the absence of PARP cleavage, suggesting a
caspase-independent mechanism . In agreement with the
above findings, Yin et al.  observed destroyed cells and cell
fragments after intense nsPEF treatments in vitro, and interpreted it
as a necrotic effect of exposure. Several studies have reported both
apoptotic and necrotic cell death after nsPEF treatments of tumors
in vivo [9,29].
In this study, we show that cell swelling and membrane rupture
are the predominant mechanisms of the early cell death following
nsPEF exposure. The prevalence of the early necrotic death was
characteristic for nsPEF treatments with either ‘‘long’’ 300-ns
pulses or ‘‘short’’ 60-ns pulses, within a wide range of doses, and
for diverse pulse delivery protocols. This primary necrotic death
prevented the development and observation of apoptosis in nsPEF-
treated cells. However, the inhibition of the primary necrosis led to
a much higher incidence of delayed cell death by apoptosis.
Materials and Methods
Cells and Media
Experiments were performed in a suspension cell line U937
(human monocytes). The cells were obtained from ATCC
(Manassas, VA) and propagated at 37uC with 5% CO2in air in
RPMI-1640 medium supplemented with 10% fetal bovine serum,
2 mM L-glutamine, 100 IU/ml penicillin, and 0.1 mg/ml strep-
tomycin. The media and its components were purchased from
Mediatech Cellgro (Herdon, VA) except for serum (Atlanta
Biologicals, Norcross, GA). Other chemicals used for this study
were from Sigma–Aldrich (St. Louis, MO) unless noted otherwise.
Modifications of the Growth Medium to Inhibit nsPEF-
induced Cell Swelling
The nsPEF-induced water uptake is driven by the colloid-
osmotic mechanism and can be blocked by the presence of a
nanopore-impermeable solute such as sucrose . Importantly,
this effect is achieved without changing the integral osmolality of
the extracellular medium.
In this study, the RPMI medium containing sucrose (hereinaf-
ter, ‘‘RPMI+sucrose’’) was produced by mixing RPMI (containing
cells and all supplements listed above) with an isoosmotic
(290 mOsm/kg) water solution of sucrose. Mixing was performed
at the proportion of 4:1 or 7:3, yielding fractional osmolalities due
to the sucrose of 58 or 87 mOsm/kg, respectively. As found in
preliminary experiments (data not shown), such fractions of
sucrose provided an accurate colloid-osmotic balance to the
cytosol, thereby preventing any volume changes in cells permea-
bilized by nsPEF.
An unavoidable side effect of the isoosmotic mixing of RPMI
with sucrose was the dilution of nutrients, salts, serum, and other
ingredients of the medium. This dilution was well tolerated by
cells, although could cause a minor slowdown of the propagation
rate. Nonetheless, in order to match this dilution, the parallel
control samples were diluted by an isoosmotic NaCl solution at the
same proportions (‘‘RPMI+NaCl’’). Na+and Cl-ions are small
solutes capable of passing the nanopores and therefore do not
prevent the water uptake [25,26]. As shown below, the growth rate
of control U937 cells (not exposed to nsPEF) in RPMI+sucrose was
the same as in RPMI+NaCl.
In several sets of experiments, the parallel control samples were
diluted with fresh RPMI instead of NaCl (‘‘RPMI+RPMI’’). In
such cases, cells were left in RPMI+sucrose and in RPMI+RPMI
only for a brief time interval (e.g., 30 min). Afterwards, all samples
were diluted 10x with fresh RPMI, thereby canceling out any
differences in the medium composition.
The exact protocols that were employed for each specific
experiment are described in the Results section and in figure
captions. As shown below, the RPMI+sucrose medium always
modified the effects of nsPEF in a similar way, irrespective of the
specific protocol employed.
nsPEF Exposure Methods and Protocols
In most experiments, we used trapezoidal pulses of 300 ns
duration from an AVTECH AVOZ-D2-B-ODA generator
(AVTECH Electrosystems, Ottawa, Ontario, Canada). Pulse
trains of needed duration at a selected repetition rate of 200 Hz
were triggered externally from a model S8800 stimulator (Grass
Instruments Co., Quincy, MA). Pulses were delivered to an
electroporation cuvette with cells using a 50- to 10-Ohm transition
module (AVOZ-D2-T, AVTECH Electrosystems) modified into a
cuvette holder. The pulse amplitude and shape were monitored
using a Tektronix TDS 3052B oscilloscope. More details of this
exposure procedure were reported earlier .
Main findings of this study were replicated using 60-ns pulses
from a high-voltage home-made pulse generator described
previously [4,31]. The goal for testing 60-ns pulses and different
exposure protocols was to demonstrate that the established
mechanisms of cell death hold true for diverse nsPEF treatments
rather than just for a specific, randomly chosen type of treatment.
The 60-ns pulse generator utilizes the pulse forming network
technology and a simple spark gap in the atmospheric air works as
a switch. With this device, the pulse repetition rate can only be
approximately controlled by the rate of network charging. We
utilized the pulse rate of about 1 Hz, and the number of pulses
delivered to the sample was controlled manually. Because of
multiple differences in the pulse delivery protocols for 300- and 60-
ns pulses, any quantitative comparison between these treatments
was not intended.
For nsPEF exposure, cells were harvested during the logarith-
mic growth phase, pelleted by centrifugation, and resuspended in a
fresh growth medium. As found in preliminary experiments, the
cell density at the time of exposure within the range from 0.5 to
86106cells/ml did not affect the nsPEF efficiency (data not
shown). For cell survival studies, the cell density at the time of
exposure was 0.6 or 1.26106
measurements which required larger cell quantities, the density
was increased to 76107cells/ml.
Immediately following nsPEF exposure, cells were diluted to
0.2–0.76106cells/ml into RPMI+RPMI, RPMI+sucrose, or
RPMI+NaCl medium and stored in the incubator until further
measurements or manipulations.
In several early series of experiments, cells were placed in the
modified medium prior to nsPEF exposure. Although slightly
lower electrical conductance of RPMI+sucrose compared to other
media could affect the efficiency of nsPEF, we observed no
differences compared to post-exposure dilutions.
In any series of experiments, samples in different media and/or
exposed to different nsPEF parameters were handled in exactly the
same manner, and different treatments were applied in a random
sequence. All series were also accompanied by parallel sham-
NsPEF exposures were performed at a room temperature of 22–
24uC. Heating of cell samples by nsPEF did not exceed 7uC, as
measured with a fiber optic ReFlex-4 thermometer (Nortech
Fibronic, Quebec City, Canada).
cells/ml; for Western blot
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Cell survival was measured at different times after nsPEF
exposure using either MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5-
diphenyltetrazolium bromide) assay or a fluorescent dye exclu-
sion/quenching method (AO/PI assay). Both assays were
described in detail previously .
In brief, for the MTT assay (BioAssay Systems, Hayward, CA)
we used a 96-well format and a Synergy 2 microplate reader
(BioTEK, Winooski, VT). 10 ml of the MTT reagent were added
to 100 ml of cell suspension and incubated for 4 hr until adding the
solubilization buffer. The plates were left on an orbital shaker
overnight and the absorbance was read at 570 nm.
The AO/PI assay utilized a mixture of a membrane-permeable
dye acridine orange (AO) and a membrane-impermeable dye
propidium iodide (PI). This method detected only massive PI
uptake characteristic for dead cells with fully ruptured plasma
membrane. Immediately prior to measurements, a 20 ml aliquot of
the cell suspension was mixed with the equal volume of staining
solution (0.5 mg/ml AO and 100 mg/ml PI in a phosphate-
buffered saline, PBS). The sample was loaded into a counting
chamber of the automated cell counter Cellometer Vision with
two-channel cell fluorescence detection (Nexcelom Bioscience
LLC, Lawrence, MA). Live cells were distinguished by bright AO
fluorescence (exc./em. 475/535 nm). In cells with compromised
membrane AO emission was quenched by PI uptake. Combined
fluorescence of either AO or PI (exc./em. 525/595 nm) was used
to determine the total (live+dead) cell counts.
Cell Diameter Measurement
Cells in the counting chamber of the Cellometer were imaged in
bright field, automatically de-clustered and distinguished from
debris. The automated recognition of cells in the sample was
verified manually and corrected if needed. The diameters of 400–
600 cells per sample were automatically measured from the image
and logged using Cellometer software.
Cell images were taken using an Olympus IX71 inverted
microscope equipped with a Retiga 2000R Fast 1394 CCD
camera (QImaging, Surrey, BC, Canada). We used 5 mg/ml PI
and 1 mM YO-PRO-1 dye (Life Technologies, Grand Island, NY)
as fluorescent markers of membrane permeabilization. The dyes
were added about 10 min prior to scheduled measurements, and
cells were allowed to settle down. Images taken with a 20x,
0.40 NA dry objective were captured and processed with
MetaMorph 7.5.2 software (Molecular Devices, Foster City, CA).
For a more detailed but mostly qualitative analysis of
morphological effects of nsPEF, we used an Olympus FluoView
1000 confocal scanning system. It utilized an IX81 microscope
equipped with differential-interference contrast (DIC) optics. 10-ml
aliquots of cells with the dyes already added were placed on a
coverslip, allowed to settle down for 5–10 min, and then examined
with a 40x, 0.95 NA objective. Due to the small size of the sample
and varied times for cell settling, the images were not used for cell
Caspase 3/7 Activity
We utilized a Caspase- GloH3/7 Assay from Promega
Corporation (Madison, WI) according to manufacturer’s instruc-
tions. Briefly, cells were exposed at 76106cells/ml and diluted
tenfold into RPMI+sucrose or RPMI+NaCl. The cells were
incubated at 37uC in 5% CO2humidified air. In 1, 2, 6, and 24 hr
after nsPEF, cells were aliquoted in triplicate at 50 ml/well into a
96-well plate; 10 ml of Caspase- GloH3/7 reagent were added to
each well, and the plate was briefly mixed on an orbital shaker.
After 40 min of incubation at room temperature, the level of
luminescence was measured by the Synergy 2 reader.
U937 cells incubated with 10 mM staurosporin were used as a
positive control for the induction of apoptosis.
Immunoblot Analysis and Quantitation of Poly-ADP
Ribose Polymerase (PARP) Cleavage
Specific PARP cleavage is an established hallmark of apoptosis
[32,33]. Both the full-length 116 kDa PARP and its 89 kDa
fragment can be detected together by immunoblotting. Quantita-
tion of the apoptotic fraction of cells from the relative amounts of
intact and cleaved PARP is intrinsically ratiometric and therefore
more quantitative than most of comparable assays.
At 1, 2, 6, or 9 hrs after nsPEF exposure, samples containing
approximately 46105cells were chilled on ice and pelleted by
centrifugation. The pellet was washed twice with ice-cold PBS and
lysed in 30 ml of a buffer containing 20 mM HEPES (pH 7.5),
200 mM NaCl, 10 mM EDTA, 1% Triton X-100, and freshly
added 1 mM DTT (dithiothreitol), 10 mM Leupeptin, 1 mM
PMSF (phenylmethanesulfonyl fluoride), and 0.2 mg/ml Benza-
midin. The samples were vortexed and centrifuged at 15,000 g at
4uC for 10 min. The supernatant containing PARP was stored at
Proteins were separated by electrophoresis on a NuPAGE 4–
12% Bis-Tris SDS-polyacrylamide gel (Life Technologies) and
then transferred to Immun-Blot Low Fluorescence PVDF mem-
brane (Bio-Rad Laboratories, Hercules, CA) by wet electroblotting
at 30 volts for 1 hr. Odyssey marker IRDye 680/800 was added as
a molecular weight standard (LI-COR Biosciences, Lincoln, NE).
The blots were blocked by incubation overnight at 4uC in the
Odyssey blocking buffer (LI-COR Biosciences).
Rabbit anti-PARP primary polyclonal antibodies (Roche
Diagnostics GmbH, Mannheim, Germany) were diluted 1:2,000
in the Odyssey blocker with 0.2% Tween-20. Donkey anti-rabbit
IgG(H+L) secondary antibodies conjugated with an infra-red
fluorophore IRDye-680LT (LI-COR Biosciences) were diluted
1:20,000 in the same buffer. The blots were treated with the
primary antibodies for 1 hr at room temperature, washed 4 times
(5 min each) in PBS with 0.1% Tween-20, treated with secondary
antibodies for 1 hr, and washed again.
The membranes were imaged using Odyssey 9120 Infrared
Imaging System (LI-COR Biosciences) in the 700 nm channel.
The images were quantified using MetaMorph software (Molec-
The fraction of the cleaved PARP (K, %) was measured as:
intensities of the 116 kDa full-length PARP and of the 89 kDa
PARP fragment, respectively. The coefficient 1.3 was used for S
mass correction. The quantitative data from 4–5 independent
experiments were processed for each timepoint and for each type
of nsPEF treatment. Staurosporin-induced apoptosis was used as a
Þ where L and S are the fluorescence
General Protocols and Statistics
All of experiments were designed to minimize potential biases
and to ensure the accuracy and reproducibility of results. All
experiments included a sham-exposed parallel control group,
which was subjected to all the same manipulations and procedures
as the nsPEF-exposed samples, excluding only the nsEP exposure
itself. Various regimens of the nsPEF treatment and parallel
control experiments alternated in a random manner, and no
‘‘historical’’ controls were accepted. Diverse buffer conditions were
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also tested in parallel. When measurements were made in triplicate
(e.g., cell viability using MTT assay), the mean of the three values
was counted as a single experiment. To achieve statistical
significance, we usually ran 4–6 independent experiments per
each group (a minimum of 3). Student’s t-test with Dunnet’s
correction when applicable [34,35] was employed to analyze the
significance of differences. The data were presented in the graphs
as mean values +/2 s.e. The difference at p,0.05 level (2-tailed)
was regarded as statistically significant. Due to multiple statistical
comparisons made (exposures versus controls; different buffers;
different timepoints; etc) we chose to let the error bars speak for
the statistical difference with minimum use of special symbols. For
clarity, the special symbols were only used for the RPMI+sucrose
groups, to indicate the significant difference from the RPMI+NaCl
group (*) and from the sham-exposed control (#). In fact, most
effects reported below were quite robust and statistically significant
at p,0.01 or better at least for several timepoints.
Sucrose Inhibits nsPEF-induced Cell Swelling and
Prevents Membrane Rupture
In a recent study , we showed that 60- as well as 600-ns
pulses cause water uptake and cell swelling due to the colloid-
osmotic imbalance [25,36–38]. In brief, the water uptake results
from the fact that small solutes can diffuse through membrane
pores towards the concentration equilibrium, whereas the larger
solutes cannot. Hence, the larger solutes remain trapped inside the
cell, thereby creating an osmotic gradient to attract water. This
gradient can be counterbalanced by replacing small solutes (such
as Na+and Cl-) in the bath buffer with larger, pore-impermeable
solutes such as sucrose. Such replacement prevents cell swelling
even though the osmolality of the extracellular buffer remains
Fig. 1 shows a typical time dynamics of cell volume changes
following nsPEF treatment. All cell samples were exposed to 600
pulses (300-ns pulse duration, 7 kV/cm, 200 Hz) in a standard
RPMI medium. Immediately after the exposure, the samples were
mixed 7:3 with isoosmotic NaCl or sucrose as described above.
Sham-exposed cell samples that served as controls were diluted the
The diameter of control cells did not depend on the time of
incubation after nsPEF or on whether the sucrose or NaCl was
added to the medium. The distribution of cell diameters was bell-
shaped, with the peak at 16–18 mm. NsPEF exposure caused rapid
swelling in the RPMI+NaCl group, eventually followed by
membrane rupture and cell destruction. The destroyed cells
shrunk abruptly, almost to the size of the nucleus, so the cell death
was manifested as an increased fraction of smaller cells. This mode
of cell death was essentially identical to the classic scenario of
hemolysis caused by the electroporation of erythrocytes .
Consistent with the earlier observations , the dilution of
RPMI with sucrose fully prevented cell swelling (Fig. 1, right
column). Consequently, sucrose also prevented the secondary cell
shrinkage due to the membrane rupture.
Dual Effect of Sucrose on Cell Survival
While it was most logical to expect that the inhibition of cell
swelling and prevention of membrane rupture by sucrose should
improve cell survival, the experiments showed that it was not the
case. At 24 hr after nsPEF exposure, and for a wide range of
exposure intensities, the cell survival stayed remarkably the same
in the presence or absence of sucrose (Fig. 2).
This unexpected finding has stimulated the analyses of the time
course of nsPEF-induced cell death under diverse conditions
(Fig. 3). Panels A, B, and C represent three independent series of
experiments. In all these series, cell survival was monitored by the
AO/PI assay for up to 24 hr (A) or 48 hr (B and C) after the
nsPEF exposure. For experiments in panel A, cells in RPMI were
diluted with either sucrose or fresh RPMI prior to nsPEF
treatment. For experiments in panels B and C, the dilution with
either sucrose or NaCl was performed immediately after the
exposure. The nsPEF exposure was either 600 pulses, 300 ns,
200 Hz at 7 kV/cm (panels A and B), or 50 pulses, 60 ns,
approximately 1 Hz at 40 kV/cm (panel C).
Irrespective of the methodological differences, the effects
observed in these experiments were similar. In sham-exposed
cells, modifications of the growth medium had little or no effect:
cells grew similarly in RPMI+sucrose and in RPMI+NaCl (panels
B and C), and perhaps slightly faster in the RPMI (panel A). At the
same time, the presence of sucrose profoundly improved the
survival of nsPEF-exposed cells at early time intervals (1–8 hr).
However, at the later time intervals the cells protected by sucrose
continued to die, whereas those without sucrose protection already
started to recover. Eventually, the percent of viable cells became
the same, and the protective effect of sucrose was nullified.
This phenomenon can also be illustrated by measuring the
fraction of dead (PI-positive) cells at different times after nsPEF
exposure (Fig. 4). Without the sucrose protection, the fraction of
dead cells increased rapidly to its maximum at 8–10 hr, and
gradually decreased afterwards. With the sucrose protection, the
fraction of PI-positive cells increased after a delay of several hours,
but showed no reduction within the period of observation.
To summarize, the presence of sucrose efficiently inhibited the
early cell death (just as expected from the blockage of cell swelling),
but the rescued cells nonetheless died later on because of some
other reason. As a result, the cell survival at intervals of 24 and
48 hr was not significantly improved by the inhibition of cell
swelling (Figs. 2 and 3).
Blockage of Cell Swelling Switches the Cell Death
Pathway from Necrosis to Apoptosis
The cell survival data reported above were consistent with
gradual changes in the cell appearance and the uptake of
membrane-impermeable dyes with time after nsPEF (Fig. 5).
At 0.5–1 hr after the exposure, cells left in the RPMI were
swollen and developed necrotic-type blebs (also sometimes called
‘‘blisters’’). In the presence of sucrose, nsPEF-exposed cells
displayed few if any morphological changes. In both these cell
populations, the plasma membrane was partially compromised,
allowing the uptake of YO-PRO-1 but not of PI (propidium ion is
larger than YO-PRO-1 and has low permeability through nsPEF-
opened pores [25,26,39]). Later on, membrane rupture in swollen
cells resulted in massive PI uptake and dual staining of dead cells
by both Yo-PRO-1 and PI. However, cells that did not rupture
became impermeable to YO-PRO-1 and regained the normal
appearance. At 4 and 6 hr after nsPEF, most of cells in RPMI
were either dead (double-stained) or normal (no staining). In the
presence of sucrose, many cells developed cytoplasm fractionation
(apoptotic blebbing) and remained permeable to YO-PRO-1.
These manifestations suggested that the delayed death in sucrose-
rescued cells could be a result of apoptosis.
Indeed, protection of nsPEF-treated cells by sucrose caused
profound activation of caspase 3/7 already at 1 hr after the
exposure, reaching maximum at 3–6 hr (Fig. 6). The cells left in
RPMI+NaCl after nsPEF exposure showed just minor caspase
activation. These findings were corroborated by another hallmark
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of apoptosis, namely the markedly increased cleavage of PARP
(Fig. 7). The data looked similar for exposures to 300- and 60-ns
pulses; taken together with the cell swelling and survival
measurements, the data indicated that the cell damage and cell
death mechanisms caused by 300- and 60-ns pulses were similar.
Figs. 6 and 7 also include the data for a chemically-induced
apoptosis as a positive control. We used staurosporin, a well-
established agent which, at the tested concentration, caused
apoptotic cell death in almost 100% of U937 cells (data not
shown). Interestingly, caspase 3/7 activation and PARP cleavage
in the staurosporin-induced apoptosis developed as much as 2–
3 hr later as compared to the nsPEF-induced apoptosis. One
possible interpretation of this observation is that nsPEF just
‘‘bypassed’’ the initial steps of the staurosporin-induced apoptotic
cascade . At the same time, we showed earlier that the
internucleosomal DNA fragmentation in nsPEF-exposed cells
developed later than in heat-shocked cells . Overall, the time
course of the nsPEF-induced apoptosis appeared within the time
limits reported for other apoptotic factors.
The Balance between the Two Modes of Cell Death
The proportion of necrotic, apoptotic, and non-apoptotic live
cells, as determined by different approaches, is presented in Figs. 8
In Fig. 8, the fraction of PI-positive (dead) cells was determined
by the AO/PI assay. The remaining fraction of live cells was split
into the ‘‘apoptotic’’ and ‘‘non-apoptotic’’ subpopulations based
on the relative amounts of intact and cleaved PARP measured in
the same sample. With time after nsPEF exposure, non-apoptotic
cells could shift into either ‘‘apoptotic’’ or ‘‘dead’’ categories, and
the ‘‘apoptotic’’ cells could also become ‘‘dead’’.
After exposure to 600 pulses (300 ns, 7 kV/cm), 60% of cells
were already dead at 1 hr if kept in the RPMI+NaCl medium.
Taking into account the early occurrence of the cell death,
morphological signs (cell swelling and membrane rupture), and the
lack of concurrent caspase 3/7 activation or PARP cleavage, the
early cell death can be categorized as a primary necrosis. Later on,
a fraction of cells kept in RPMI+NaCl entered the apoptotic
pathway; however, even assuming that the entire cell loss after
2 hr was due to the apoptosis only, the cumulative fraction of
Figure 1. Sucrose inhibits cell swelling and membrane rupture caused by nsPEF. The bar charts show the frequency distribution of cell
diameter values at the indicated time intervals after nsPEF exposure and in sham-exposed controls. Cells were exposed in the RPMI medium and
placed immediately afterwards into either RPMI+NaCl or RPMI+sucrose (87 mOsm/kg due to sucrose); see text for more details. 400–600 cells per
group were measured at each timepoint. Note fast cell swelling followed by membrane rupture and apparent shrinkage in RPMI+NaCl, but not in the
RPMI+sucrose. Representative cell images in the differential interference contrast (DIC) and propidium iodide (PI) fluorescence channels illustrate
swelling and eventual membrane rupture in the RPMI+NaCl medium.
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apoptotic cells was just 16%. In contrast, the same calculation for
sucrose-protected cells yields over 50% of apoptotic cells. Thus,
the primary necrosis was the predominant cell death pathway
unless cells were protected with sucrose. For the data in Fig. 8, the
primary necrosis was responsible for about 87% of the cell loss,
versus 43% in the presence of sucrose.
Fig. 9 shows the time dynamics of cell subpopulations
permeable to either YO-PRO-1, or both PI and YO-PRO-1, or
not permeable to any of the dyes. Notably, YO-PRO-1 is both a
sensitive indicator of membrane nanoelectroporation [25,39,41]
and a marker of selective membrane permeabilization early in the
course of apoptosis . In contrast, the uptake of PI through
nanopores is minimal; with the employed method of PI detection,
its uptake manifests the irreversible cell destruction, by either
primary or secondary necrosis.
Nanopores created by nsPEF still remained permeable to YO-
PRO-1 at 20 min after the exposure, as seen by YO-PRO-1
uptake by most cells. In the RPMI+NaCl medium, these cells
swelled and got destroyed (became PI-permeable) already within
an hour. In the RPMI+sucrose group, many cells remained
permeable just to YO-PRO-1 for several hours after the nsPEF
exposure. The fraction of PI-positive cells was much smaller and
stable at 1–4 hr after the exposure, followed by a delayed increase
by 6 hr.
Figure 2. Lack of the effect of sucrose on the 24-hr survival of
nsPEF-treated cells. Cells in RPMI were mixed with sucrose
(RPMI+sucrose; 60 mOsm/kg due to sucrose) or fresh RPMI (RPMI+RPMI)
before nsPEF exposure (600 pulses, 300-ns). At 30 min after the
exposure, all samples were diluted tenfold with fresh RPMI and
incubated until measuring cell survival by the MTT assay at 24 hr (mean
values +/2 s.e. for 6 independent experiments).
Figure 3. Inhibition of swelling improves the short-term but not the long-term survival after nsPEF exposure. Panels A, B, and C
represent the data from three independent sets of experiments performed under different exposure conditions and using different protocols. For
panel A, cells in RPMI were mixed with sucrose (RPMI+sucrose; 60 mOsm/kg due to sucrose) or fresh RPMI (RPMI+RPMI) before nsPEF exposure (the
same protocol as in Fig. 2). For panels B and C, cells were exposed in the RPMI and placed immediately afterwards into either RPMI+NaCl or
RPMI+sucrose (87 mOsm/kg due to sucrose), same as in Fig. 2. See graph legends and text for more details. Cell survival was measured by the AO/PI
assay and normalized to the pre-exposure value (mean+/2 s.e., n=4–6). Cell survival in sham-exposed controls is shown by dashed lines and open
symbols. * p,0.05 for the difference of RPMI+sucrose from RPMI+NaCl (or RPMI+RPMI); # p,0.05 for the difference of RPMI+sucrose from the
respective sham-exposed control. Other significant differences are not shown for clarity.
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While it is widely accepted that nsPEF-exposed cells die by
apoptosis, our results demonstrate for the first time that the
primary necrosis was the predominant cell death mode. We also
established that necrosis results from plasma membrane permea-
bilization, followed by water uptake, cell swelling, and eventual
membrane rupture. This necrotic pathway is similar to what is
seen with ‘‘classic’’ electroporation pulses or when applying
various other necrotic factors.
This result may appear contradictory to the prevalence of
nsPEF-induced apoptosis as reported by multiple other studies
Figure 4. Inhibition of swelling blocks the early cell death after nsPEF. Dead cells were identified by the AO/PI assay. The total number of
cells counted at each timepoint was taken as 100% (mean+/2 s.e., n=4–6). The data in panels A and B are from the same experiments as in Fig. 3, B
and C. See text and Fig. 3 for details. PI uptake in sham-exposed controls is shown by dashed lines and open symbols.
Figure 5. Effects of sucrose on cell swelling and membrane permeability. DIC and fluorescence images of nsPEF-exposed cells incubated in
either RPMI+RPMI or RPMI+sucrose. Green: YO-PRO-1; red: PI; yellow: both dyes overlapped. Parameters of exposure and times after it are given in the
legend. Cells were handled the same way as in Fig. 2 but without additional dilution at 30 min. The dyes were added 5–10 min prior to taking an
image. Note early cell swelling and rupture in the RPMI+RPMI but not in the RPMI+sucrose medium. The survivors show no YO-PRO-1 uptake in the
RPMI+RPMI, but remain permeable to the dye in the RPMI+sucrose group. An arrow points to a group of cells that display the apoptotic blebbing and
Cell Death Triggered by Nanosecond Electric Pulses
PLOS ONE | www.plosone.org7 July 2013 | Volume 8 | Issue 7 | e70278
[1,3,4,10,14,43]. However, these reports were based primarily on
the flow cytometry counts of cells that display PS externalization;
as discussed above, the relevance of this parameter to apoptosis in
nsPEF-treated cells is questionable.
Figure 6. Inhibition of swelling in nsPEF-exposed cells
facilitates caspase 3/7 activation. The exposure parameters and
media are identified in the legend. Growth media were changed the
same way as in Fig. 1. Caspase-3/7 was measured by a luminescence
assay. For a positive control, apoptosis was induced by 10 mM of
staurosporin. Mean values +/2 s.e. for n=3. See text and Fig. 3 for
Figure 7. PARP cleavage in nsPEF-exposed cells. A: The fraction of cleaved PARP is increased when nsPEF-exposed cells are protected with
sucrose. Mean values +/2 s.e. for n=4–5. Growth media were changed the same way as in Fig. 1. NsPEF and media conditions are specified in the
legend. The numbers in parentheses correspond to the lanes in panel B, which shows representative Western blots for intact and cleaved PARP (116
and 89 kDa, respectively). See text and Fig. 3 for details.
Figure 8. The structure of nsPEF-exposed cell populations with
and without blockage of cell swelling with sucrose. Bars show
relative fractions of non-apoptotic, apoptotic, and dead cells at different
timepoints after nsPEF (600 pulses, 300 ns, 7 kV/cm). Growth media
were changed the same way as in Fig. 1. Dead and live cells were
counted by the AO/PI assay. The fraction of apoptotic cells among live
cells was considered proportional to the fraction of cleaved PARP. The
data were averaged from 4–5 experiments; the error bars are omitted
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Although a number of studies reported evidence for necrotic cell
death after nsPEF [4,6,9,17,28], this pathway has received little
attention. Most of research focused on cellular mechanisms of
nsPEF-induced apoptosis, whereas necrosis was viewed as a less
common and less important event. In this study, for the first time
we report that under standard cell culture conditions necrosis can
be a prevalent mode of cell death. This finding holds true for
rather diverse nsPEF exposure conditions, including different pulse
durations, E-field values, and pulse delivery protocols.
With that said, the balance between apoptosis and necrosis can
be profoundly dependent on the cell type and on the cell
environment. For example, cells that do not have a large ‘‘stock’’
of spare membrane to swell will have less time for membrane
repair after nsPEF, and are more likely to die from the membrane
rupture. This fact may explain why smaller Jurkat cells were more
vulnerable than larger U937 . Cells within tissues in vivo may
have limited room for swelling. Instead of the presence of sucrose,
swelling can potentially be limited by the space constraints,
thereby shifting the in vivo cell death towards apoptosis.
The profound increase of apoptosis in nsPEF-treated cells in the
presence of sucrose raises a question if sucrose just ‘‘unmasked’’
the latent apoptosis or also facilitated the apoptotic cell death. For
example, in Fig. 9 (right panel, RPMI+sucrose), the pool of YO-
PRO-1 positive cells remained large for several hours after the
exposure. This pool concurrently shrunk due to both resealing of
nanopores and cell death, and expanded due to the development
of apoptosis. One may speculate that the presence of sucrose could
somehow inhibit the cell membrane repair, thereby leaving it
permeable to YO-PRO-1 for longer time. Such long-lasting
membrane disruption due to the impaired repair would be a
plausible explanation for the onset of apoptosis in sucrose-
protected cells; however, this mechanism does not appear to be
supported by the data. Indeed, a large increase in the fraction of
cells that did not uptake any of the dyes (between 0.3 and 2 hr)
argued for the successful pore resealing in the RPMI+sucrose
group. Therefore the development of apoptosis was not a side
effect of the sucrose; instead, it was an effect of nsPEF exposure
itself, which was masked by the faster necrotic process under the
normal cell culture conditions.
The fact that nsPEF triggers both necrotic and apoptotic death
mechanisms makes it an attractive modality for cancer ablation.
First, the concurrent induction of both cell death modes reduces
the chance for malignant cells to escape the death sentence,
despite the sophisticated death evasion mechanisms developed by
various cancers. Second, varying the nsPEF exposure parameters
and combining nsPEF with sucrose injection or a similar treatment
is an approach to deliberately induce either the apoptotic or
necrotic death, or a combination thereof. For an in vivo cancer
treatment, a carefully tuned combination of necrotic and apoptotic
cell death may be an optimal solution for tumor elimination
without excessive pain and inflammation while stimulating the
immunogenicity of tumor cells [2,44].
Conceived and designed the experiments: ONP BWG IS AGP. Performed
the experiments: ONP BWG. Analyzed the data: ONP BWG IS AGP.
Wrote the paper: ONP AGP.
Figure 9. The effect of sucrose on Yo-PRO-1 and PI uptake by nsPEF-exposed cells. Growth media were changed after nsPEF exposure (600
pulses, 300 ns, 7 kV/cm) the same way as in Fig. 1. The number of cells displaying no dye uptake, YO-PRO-1 uptake, and both YO-PRO-1 and PI uptake
were automatically counted in microscope images as described in Methods. The total number of cells counted in each sample was taken as 100%.
The PI-positive cells were presumed dead. YO-PRO-1-positive cells could be either apoptotic or just transiently permeabilized to this dye by nsPEF.
Cells negative for either dye were regarded as live, non-apoptotic. The data were averaged from 3 experiments; error bars are omitted for clarity.
Cell Death Triggered by Nanosecond Electric Pulses
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