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Biophysical studies of nanosecond pulsed electric field induced cell membrane permeabilization
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Biophysical studies of nanosecond pulsed electric field induced cell membrane permeabilization
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Content
Biophysical Studies of Nanosecond Pulsed Electric Field
Induced Cell Membrane Permeabilization
by
Yu-Hsuan Wu
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(Materials Science)
May 2014
Copyright
2014
Yu-‐Hsuan
Wu
i
Dedication
This thesis is dedicated to my parents, sisters and brother
for their endless support and encouragement.
ii
Acknowledgements
I would like to gratefully thank my advisor, Prof. P. Thomas Vernier. He
guided me to this exciting field and taught me how to think and behave as a
good researcher. I appreciate all his encouragement and ideas to let me
accomplish my Ph.D. researches at USC. I would like to thank the enthusiastic
supervision of Prof. Martin Gundersen for his guidance and encouragement in
my research and study. It has been an honor to be a Ph.D. student in his research
group.
I would also like to thank my colleagues, Dr. Andras Kuthi, Dr. Chunqi
Jiang, Dr. Jason M. Sanders, Dr. Zach Levine, Yung-Hsu Lin, Ming-Chak Ho.
Without their good advices and collaboration, I could not have done all this
work here.
Finally, I would like to thank our collaborator, Prof. Delia Arnaud-
Cormos, Dr. Stefania Romeo, Dr. Tina Batista Napotnik and Maura Casciola. It
is grateful to work with them.
iii
Table of Contents
Dedication i
Acknowledgements ii
List of Figures vi
Abbreviations xiii
Abstract xiv
Chapter 1: Introduction 1
1.1 The Action of Nanosecond Pulsed Electric Field 1
1.2 nsPEF and Apoptosis 3
1.3 nsPEF and Plasma Membrane Permeability 6
1.4 Parameters that Affect nsPEF-Induced Responses 10
1.5 nsPEF Delivery Systems 13
Chapter 2: Lipid Peroxidation in Living Cells Promotes Plasma Membrane
Electropermeabilization 17
2.1 Abstract 17
2.2 Introduction 18
2.3 Materials and Methods 20
iv
2.4 Lipid Peroxidation Enhances Membrane Electropermeabilization 25
2.5 MD Simulations Show Evidences of Electropermeabilization Enhanced by
Oxidized Membrane Lipids 29
2.6 Conclusion 32
Chapter 3: The Effects of Nanosecond Pulsed Electric Field on Mitochondrial
Membrane 34
3.1 Abstract 34
3.2 Introduction 35
3.3 Materials and Methods 37
3.4 Localization of Mitochondria 42
3.5 nsPEF Cause a Loss of Mitochondrial Potential 44
3.6 Conclusion 53
Chapter 4: Cell Swelling and Membrane Permeabilization after Nanosecond
Pulsed Electric Field 55
4.1 Abstract 55
4.2 Introduction 56
4.3 Materials and Methods 58
4.4 Electric Pulse-Induced Cell Swelling 61
4.5 Effects of Pulse Repetition Rate on nsPEF-Induced Cell Swelling 67
4.6 A Scaling Law for nsPEF-Induced Cell Swelling 71
4.7 nsPEF-Induced Ion Flux 74
4.8 MD Simulation of Water Permeation of Intact Lipid Bilayer 78
4.9 Conclusion 81
Chapter 5: Moveable Wire Electrode Microchamber for Nanosecond Pulsed
Electric Field Delivery 82
v
5.1 Abstract 82
5.2 Introduction 82
5.3 Setup of nsPEF Exposure System with Tungsten-Wire Electrodes (TWE) 85
5.4 Electromagnetic Analysis of TWE Delivery System 96
5.5 Biological Experiments with 2.5 and 5.0 ns, 10 MV/m nsPEFs 105
5.6 Conclusion 107
Chapter 6: Determination of Nanoelectropulse-Induced Pore Size by Blocking
Osmotic Swelling 108
6.1 Abstract 108
6.2 Introduction 109
6.3 Materials and Methods 111
6.4 Cell Volume Changes Induced by nsPEFs 115
6.5 Impermeant Solute Inhibits Cell Swelling (PEG1000) 118
6.6 Cell Swelling Reduced by Inositol, Blocked by Sucrose 120
6.7 Pulse Dose and Pore Diameter 122
6.8 Conclusion 126
Chapter 7: Conclusion 128
References 131
vi
List of Figures
Figure 1.1 A map of applications and effects of pulsed electric field in pulse
duration-strength space. Both values of duration and field strength are
shown in log. The figure was published by Weaver et al. [33]. 5
Figure 1.2: Increases in intracellular fluorescence intensity resulting from YO-
PRO-1 influx, indicating electropermeabilization, are greater for 4 ns pulses
than for 3 ns pulses at 10 MV/m. Doubling the field from 10 MV/m to 20
MV/m increases permeabilization ten-fold. Data represents the ratio of YO-
PRO-1 fluorescence emission 3 minutes after and immediately before pulse
exposure from the integrated intensity of at least 10 cells for each condition
from each of 3 independent experiments. YO-PRO-1 (5 µM): Invitrogen;
λex: 491 nm, λem: 509 nm; Filter set: Emission filter: 480/30 nm, Dichroic
mirror: 505D, Excitation filter: 535/40. 11
Figure 1.3: Constructed pulse generator with a cuvette load by Tang et al. in
USC [66]. 14
Figure 1.4: Four-channel electrode microchamber on a standard microscope
glass slide. Top: ideal electrode chamber with straight sidewalls and
appropriate dimensions. Bottom: top view of the electrode microchamber
[68]. 15
Figure 2.1: Membrane lipid peroxidation visualized with C11-BODIPY
581/591
.
(a): Fluorescence images of Jurkat cells showing C11-BODIPY
581/591
in
control and peroxidized cells. (b): Integrated ratio C11-BODIPY
581/591
fluorescence intensity from more than 300 individual cells from three
independent experiments for each condition. Error bars are standard error of
the mean. 26
Figure 2.2: Electropermeabilization enhancement by treatment with
peroxidation agents. (A). Fluorescence images of Jurkat cells showing
pulse-induced (30 ns, 3 MV/m, 50 Hz) YO-PRO-1 influx into control and
peroxidized cells after 10 min exposure to 500 µM H
2
O
2
+1000 µM FeSO
4
.
(B). Integrated YO-PRO-1 fluorescence intensity from more than 300
individual cells from three independent experiments for each condition.
Error bars are standard error of the mean. 28
Figure 2.3: Molecular structures of PLPC and two oxidized variants, oxPLPC
(12-al) and oxPLPC (13-tc). 29
vii
Figure 2.4: Consecutive snapshots of a system with an 11 % concentration of
12-al oxPLPC. In the first snapshot all lipids are displayed whereas in all
subsequent snapshots, only the oxidized sn-2 tails are shown. Carbon is
colored cyan and oxygen is colored in red. The enlarged red spheres
represent aldehyde oxygens which appear to attract water deep into the
bilayer interior, thus helping mediate the process of pore formation. 31
Figure 2.5: Large quilted system which contains PLPC lipids (light gray) and
selectively placed oxPLPC lipids (dark blue) which were inserted by
replacing PLPC lipids with oxPLPC lipids in opposing quadrants. The
concentration of oxPLPC in an oxidized patch is 50 %. After an electric
field is applied, it appears that there is a clear correlation between pore
location and local oxidation sites. 32
Figure 3.1: Experimental setup for pulse exposure. Cells were loaded in the
homemade michcrochamber lay on a specially designed board with voltage
divider on it. The board was connected to the pulse generator and
oscilloscope, and located on the fluorescence microscope beforehand. This
setup made the real-time observation of pulsing experiment happen. 39
Figure 3.2: Localization of mitochondria in Jurkat cells. Cells were treated with
cobalt-quenched calcein assay (A), Rhodamine 123 (B) or TMRE (C), and
incubated with either MitoTracker Orange (A, B) or MitoTracker Green (C)
together. A1-C1: permeabilization indicator images. A2-C2: Mitotracker
stained images. A3-C3: superimposed images. 43
Figure 3.3: A schematic diagram shows how Rhodamine 123 (R123) works.
R123 is a lipophilic cationic fluorescent dye that is accumulated within
active mitochondria because of large negative potential across the
mitochondrial membrane. The decrease of mitochondria potential will
decrease R123 concentration in mitochondria and cause the drop of its
fluorescent intensity. TMRE works in a similar way. 45
Figure 3.4: Dose-dependent R123 fluorescence intensity before and after pulsing
(mean ± SEM). CCCP was used as positive control. Significant differences
from zero-pulse controls are designated by asterisks (*P < 0.05).
#Statistically different from previous column of same incubation time (P <
0.05). There are at least 60 cells for each data point and each condition was
repeated at least three times.. 46
Figure 3.5: Dose-dependent TMRE fluorescence intensity before and after
pulsing (mean ± SEM). CCCP was used as positive control. Significant
differences from zero-pulse controls are designated by asterisks (*P <
viii
0.05). #Statistically different from previous column of same incubation
time (P < 0.05). There are at least 124 cells for each data point and each
condition was repeated at least three times. 47
Figure 3.6: Non-fluorescent calcein-AM and Co
2+
enter the cell. AM groups are
cleaved from calcein via non-specific esterase activity in the cytosol and
mitochondria. Co
2+
quenches the cytosolic calcein signal. Co
2+
cannot
readily enter healthy mitochondria. Co
2+
influx after pore opening quenches
the mitochondrial calcein fluorescence. 49
Figure 3.7: Dose-dependent calcein fluorescence without (A) or with (B) Co
2+
in
the external medium. Data show the relative fluorescence intensity before
and 3 min after pulsing (mean ± SEM). Significant differences from zero-
pulse controls are designated by asterisks (*P < 0.05). #Statistically
different from previous column (P < 0.05). There are at least 38 cells for
each data point and each condition was repeated at least three times. 50
Figure 3.8: Dose-dependent YO-PRO-1uptake.Fluorescence images were
recorded 3 min after pulse exposure. Data show YO-PRO-1 fluorescence 3
min after pulsing (mean ± SEM) in arbitrary units. Significant differences
from zero-pulse controls are designated by asterisks (*P < 0.05). There are
at least 36 cells for each data point and each condition was repeated at least
three times. 51
Figure 3.9: Dose-dependent propidium uptake. Fluorescence images were
recorded 3 min after pulse exposure. Data show PI fluorescence 3 min after
pulsing (mean ± SEM) in arbitrary units. There are at least 32 cells for each
data point and each condition was repeated at least three times. 52
Figure 4.1: Osmotic swelling after nanoelectropulse exposure. (A) Jurkat cells
before pulse exposure. (B) Same cells 60 s after exposure to 30, 5 ns, 10
MV/m pulses at 1 kHz. (C), (D). Cells in (A) and (B) are outlined for area
extraction with ImageJ. Note pulse-induced swelling, blebbing, and
intracellular granulation and vesicle expansion, a result of the osmotic
imbalance caused by electropermeabilization of the cell membrane. 62
Figure 4.2: Representative images of morphological changes in Jurkat cells
before (0 s) and after (10 s, 20 s, 30 s, 40 s, 60 s, 120 s, 180 s, 300 s and
420 s) exposure to 30, 5 ns, 10 MV/m pulses at 1 kHz. Even at 420 s, the
longest monitoring time permitted by our apparatus, cells were not
observed to recover their initial volume. 63
ix
Figure 4.3: Dose-dependent area increase of Jurkat cells after exposure to 0, 10,
20, 30 and 50 5 ns, 10 MV/m electric pulses delivered at 1 kHz. Note initial
rapid swelling, which declines over about 1 min to a slower, but non-zero
rate. Results are presented as mean ± standard error for at least 30 cells,
from at least 3 independent experiments, for each pulsing condition. 64
Figure 4.4: Cell swelling can be reliably detected after even a single 5 ns, 10
MV/m pulse. Results are presented as mean ± standard error for at least 30
cells, from at least 3 independent experiments, for each pulsing condition. 65
Figure 4.5: Frequency-dependent YO-PRO-1 influx after exposure of Jurkat
cells to 30 4 ns, 10 MV/m electric pulses delivered at different repetition
rates. Fluorescence microscopic images of Jurkat cells in growth medium
containing YO-PRO-1 (5.0 µM) were captured immediately before and 3
min after exposure. Fluorescent intensity of YO-PRO-1 was integrated
from the captured images. Results are presented as mean ± standard error
for at least 30 cells, from at least 3 independent experiments, for each
pulsing condition. 68
Figure 4.6: Rate and extent of pulse-induced swelling is greater at higher pulse
repetition rates. Results are presented as mean ± standard error for at least
30 cells, from at least 3 independent experiments for each pulsing condition.
69
Figure 4.7: Electric pulse-induced area increase dose-dependency. (a) The
phenomenon is described for times up to about 60 s after exposure by the
electrical impact dose factor, the product of the electric field, the pulse
duration, and the square root of the number of pulses. The impact dose
variable for these experiments is the pulse number N, which has the
following values: 0, 1, 3, 5, 10, 20, 30, and 50. All pulses were 5 ns, 10
MV/m, delivered at 1 kHz. Straight gray lines are a linear fit to the swelling
data at 10 s and 60 s; (b) fitting of the cell swelling data with Eq. (2); (c)
fitting parameters over time. 73
Figure 4.8: Random permeation of an intact POPC bilayer. A single H
2
O
molecule crosses the phospholipid bilayer interface. Applied electric field is
340 MV/m downward in the diagram. (A) 0 ps, (B) 100 ps, (C) 200 ps, (D)
300 ps. Only acyl oxygens and water molecules in the interface are shown.
The crossing water molecule is enlarged and colored yellow. Light blue
spheres are acyl oxygens; red and white atoms are water; the empty space
in the middle of figures between two water phases represents the
hydrocarbon region of the lipid bilayer. From a system containing 128
POPC, 4480 H
2
O, at 310 K, 1 bar. 80
x
Figure 5.1: General experimental delivery system set-up composed of the nsPEF
generator, the measurement devices, the TWE delivery system, the
micromanipulator and the microscope stage. 85
Figure 5.2: The power measurement device with 100:1 attenuator was designed
and assembled in USC. Three transmission lines connected to the pulse
generator, oscilloscope and TWE separately. 87
Figure 5.3: Sketch shows the design of TWE. The parallel TWE connected to
nsPEF generator is able to work on a microscope stage. A 3-axis
micromanipulator can control the movement of TWE precisely. 89
Figure 5.4: Pictures show the fabrication process of PDMS mold. To making the
PDMS mold, a thick layer of SU8 was coated on a silicon wafer followed
by post soft baking. The baked wafer was covered by the mask designed for
wire alignment, and exposed to UV light. After UV exposure, a hard bake
was applied. Finally, a thick layer of PDMS was spread on the wafer to
finish the fabrication. 90
Figure 5.5: Tungsten wire electrodes delivery system. A) Overview of the TWE
components, materials and dimensions. B) Zoom on the TWE containing
the cells under nsPEF exposure. All dimensions are in mm. 91
Figure 5.6: TWE delivery system frequency characterization obtained by FDTD
simulations and measurements with vector network analyzer. (A)
Reflection coefficient (S
11
). (B) Real and imaginary parts of the impedance. 99
Figure 5.7: Time domain measurements with both generators. A) Applied
Pulses. B) Normalized Power spectra in dB. 101
Figure 5.8: Electric field distributions in the TWE channel. A) TWE without
biological medium placed in the coverglass well. B) Same as previous but
with biological medium. C) Same as previous but without the coverglass
chamber. 104
Figure 5.9: Dose-dependent area increase of Jurkat cells after exposure to 0, 5
and 10 electric pulses delivered with 10 MV/m field strength and a
frequency repetition of 1 kHz. Results are presented as mean ± standard
error for at least 70 cells, from at least 3 independent experiments, for each
pulsing condition. (A) 5 ns duration pulses delivered by nsPEF1. (B) 2.5 ns
duration pulses delivered by nsPEF2. 106
xi
Figure 6.1: The molecular structures of tested sugars were drawn by VMD
software. The minimum cross-sectional areas were shown and the smallest
diameters measured from the cross-sectional areas were listed in the table. 113
Figure 6.2: Time series of DIC images showing a Jurkat cell swelling after
nsPEF exposure (10 5 ns, 10 MV/m at 1 kHz) in RPMI 1640. Scale bar is 5
µm. 116
Figure 6.3: Dose-dependent area increase of Jurkat cells after nsPEF exposure to
0, 5, 10, 20 and 30 5 ns electric pulses delivered with 10 MV/m field
strength and a repetition rate of 1 kHz in RPMI 1640. Swelling ratio
depends on the applied pulse intensity. With increasing pulse counts there
was an increase in the volume changes. The results are presented as
mean±SE for at least 60 cells from at least three independent experiments
for each pulsing condition at 25 °C 117
Figure 6.4: Isoosmotic substitution of PRMI 1640 with PEG 1000 with different
concentration. Jurkat cells were exposed to 30 5 ns pulses delivered with 10
MV/m field strength and 1 kHz repetition rate. Increasing of PEG 1000
concentration inhibited the nsPEF-induced cell swelling. The results are
presented as mean±SE for at least 60 cells from at least three independent
experiments for each pulsing condition at 25 °C. 119
Figure 6.5: : Protective action of inositol and sucrose against nsPEF-induced cell
swelling. Jurkat cells were exposed to 0, 5, 10, 20 and 30 5 ns pulses
delivered with 10 MV/m field strength and 1 kHz repetition rate in
solutions isoosmotically replaced 116 mOsm RPMI1640 with inositol or
sucrose. The extent of swelling in solution containing inositol and the
inhibition of swelling in solution with sucrose indicated that inositol can
pass through the 5 ns electric pulses created pores but sucrose can not. The
results are presented as mean±SE for at least 60 cells from at least three
independent experiments for each pulsing condition at 25 °C. 121
Figure 6.6: Cell shrinking after exposed to 30 and 100 5 ns pulses delivered with
10 MV/m field strength and 1 kHz repetition rate in solutions
issosmotically replaced RPMI 1640 with higher concentration of sucrose.
The continuous shrinking in the observation time course demonstrated that
even 100 5 ns electric pulses created membrane pores smaller than sucrose
molecules. The results are presented as mean±SE for at least 60 cells from
at least three independent experiments for each pulsing condition at 25 °C. 123
Figure 6.7: The effect of extremely large number of 5 ns pulses on membrane
pore size in solutions isoosmotically replaced RPMI 1640 with PEG 1000
xii
with different concentration. Large number of 5 ns pulses did not create
pores larger the PEG 1000. However, the data in 50 and 116 mOsm PEG
1000 shows that the intracellular solutes that were impermeable to cell
plasma membrane at 30 pulses started to be able to penetrate the membrane
while the pulse number over 100. The results are presented as mean±SE for
at least 60 cells from at least three independent experiments for each
pulsing condition at 25 °C. 125
xiii
Abbreviations
CCCP: carbonyl cyanide 3-chlorophenylhydrazone
DIC: differential interference contrast
DNA: deoxyribonucleic acid
ECT: electrochemotherapy
EGT: electro-gene therapy
IRE: irreversible electroporation
mPTP: mitochondira permeability transition pore
nsPEF: nanosecond pulsed electric field
PI: propidium iodide
PS: phosphatidylserine
R123: rhodamine 123
TMRE: tetramethyl rhodamine ethyl ester
xiv
Abstract
Nanosecond megavolts-per-meter pulsed electric field (nsPEF) offers a non-
invasive manipulation of intracellular organelles and functions of biological cells.
Accordingly, nsPEF is a potential technique for biophysical research and cancer
therapy, and is of growing interest. Although, the application of nsPEF has shown
electroperturbation on cell plasma membranes and intracellular membranes as well,
the mechanisms underlying the electropermeabilization are still not clear. In this
thesis, we systematically study nsPEFs (5 and 30 ns) induced membrane
permeability change in biological cell in-vitro with different pulse parameters. In
Chapter 3, we investigate the nsPEF-induced intracellular membrane
permeabilization of mitochondria which play key roles in activating apoptosis in
mammalian cells. The results show the evidences of nsPEF-induced membrane
permeability increase in mitochondria, and suggest that nsPEF is a potential
technology for cancer cell ablation without delivery of drug or gene into cells.
In Chapter 2, 4 and 6, we study the properties of nsPEF-induced plasma
membrane permeabilization. In the beginning, the change of plasma membrane
permeability is studied by uptake of YO-PRO-1 and propidium iodide, fluorescent
dyes specifically used as indicators of plasma membrane permeabilization.
However, the detection is limited by the fluorescent emission efficiency and detector
capability. To increase the detection sensitivity, we later develop a method based on
cell volume change due to regulation of osmotic balance that causes water and small
xv
ions transport through plasma membrane. We find that even a single 10 MV/m pulse
of 5 ns duration produces measureable cell swelling. The results demonstrate that
cell swelling is susceptible to nsPEF and can detect membrane permeabilization
more easily and precisely than fluorescent dyes. We compare the effects of different
pulse parameters (pulse duration, pulse number, electric field amplitude and pulse
repetition rate) on electropermeabilization. The effects of chemical agents that either
promote (H
2
O
2
) or inhibit (lanthanide ions and Hg
2+
) electropermeabilization are
also studied. To characterize the population of pores created by nsPEFs, we
isoosmotically substitute different size of neutral molecules in the pulsing medium,
and estimate pore size by analyzing cell volume changes that result from the
permeation of these substituted molecules through the plasma membrane of Jurkat T
lymphoblasts. The basis of this method is regulation of osmotic balance across the
plasma membrane as well. We find that most pores opened by 5-100 5 ns pulses in
plasma memebrane of Jurkat T lymphoblasts have diameter between 0.7-0.9 nm.
In Chapter 5, we report the design and construction of a delivery system for
nsPEF. We integrate a pair of delicately fabricated tungsten wire electrodes spaced
100 µm, a solid-state high-voltage nanosecond pulse generator and a fluorescent
microscope coupling with a fast and sensitive digital recording camera. This system
enables real-time biophotonic investigations of the nsPEF-induced biological
responses of living mammalian cells in-vitro.
1
Chapter 1: Introduction
1.1 The Action of Nanosecond Pulsed Electric Field
Intense pulsed electric fields are capable to alter the barrier function of
biological cell membrane by rapidly raising the transmembrane voltage to a
threshold value (at least 0.2 V and usually 0.5 to 1 V), causing membrane
breakdown and thus increasing the membrane conductivity [1, 2]. Neumann et al.
first reported the evidence in 1972 showing that the pulsed electric fields can
transiently increase cell plasma membrane permeability that may further lead to
release of biogenic amines [3]. This transient effect induced by electric pulses is
often attributed to a burst of pore formation and known as electroporation or
electropermeabilization. It has been proved to permit the transfer of ions and
molecules, which are normally excluded by cell plasma membrane, into cytoplasm
[4-6]. In conventional electroporation, pulses in the range from milliseconds with
several tens of kV/m to a few microseconds with several hundred kV/m have been
wildly used to facilitate the transport of pharmacological and genetic materials into
cell interior for laboratory and clinical applications [7-13].
Electric pulse with nanoseconds duration and MV/m field intensity (nsPEF)
was available since mid-90s. The equivalent circuit modeling and molecular
dynamic simulation suggested that nsPEFs create much smaller pores at higher pore
2
density than conventional electroporation (diameter less than 1 nm). Moreover,
contrary to conventional electroporation, nsPEFs with the increase in field strength
can penetrate exterior plasma membrane, produce significant voltage across
intracellular organelle membranes and thus porate these small organelles [14-21].
The idea makes nsPEF a potential tool to target and to manipulate intracellular
organelles without direct lethal effects. The applications of nsPEF induced
intracellular effects include activation of platelets and release of growth factors for
accelerated wound healing [22], stimulation of neuromuscular responses [23-25],
control of apoptosis [26-29], and shrinkage or elimination of melanoma tumors [30,
31].
Joule heating due to energy dissipation in the applications of pulsed electric
field is unavoidable. The magnitude of temperature increase is related to the pulse
duration, applied filed strength and the electric conductivity of sample. Large and
long pulses can create significant temperature rises which lead to thermal damages
(ex: burns) [32]. This undesirable side effect can be reduced by delivering short and
low field pulses in low repetition rates. Esser et al. [19] and Weaver et al. [33] used
Pennes bioheat equation to estimate the temperature increase by electric pulse. A
single 100 µs, 2 kV/m electric pulse increases ~ 1 °C in in-vitro environment [19]. A
single 20 ns, 6 MV/m electric pulse increases ~ 0.15 °C in multicellular tissue
model. Moreover, Chen et al. used resistance temperature detector (RTD) for direct
measurement and reported that 15000 15 ns pulses of maximum 10 MV/m field
3
intensity, 50 Hz repetition rate (pulses delivered by needles or plate electrode)
increases less than 1 °C in in-vitro environment [34]. These studies suggest that
temperature change due to ultra short pulses (ns) is small and might be negligible
while treatment protocols are taken carefully. However, the estimations and
measurements above are happened in simulated model and in-vitro experiments. The
contribution of Joule heating should be always taken into account in in-vivo
application where higher conductivity pathway is expected [35].
1.2 nsPEF and Apoptosis
Cell death is important in regulating organismal development, tissue
homeostasis and interconnects with cell survival and proliferation [36]. During
tumor development, uncontrolled cell proliferation is aided by the disablement of
cell death responses triggered by specific oncogenes [37]. Apoptosis is one form of
cell death mechanisms. It is a consequence of a series of precisely regulated events.
In contrast to necrosis, another form of cell death mechanisms that is usually
characterized by rupture of the plasma membrane with a consequent localized
inflammatory response and damage to surrounding cells and tissues, apoptosis is
associated with the rapid engulfment and removal of cell debris by phagocytes that
recognize “eat-me” signals displayed on the outer surface of the apoptotic cell [38].
4
The difference between these two mechanisms of cell death makes apoptosis more
desirable for the induction of cell death in cancer cells than necrosis.
Conventional electroporation using electric pulses with duration from few µs
to ms has been used in tumor treatment for a period of time. These include
electrochemotherapy (ECT), electro-gene therapy (EGT) and irreversible
electroporation (IRE) [33, 39]. ECT uses pulsed electric field in the regime that can
cause reversible electroporation in tumor cell plasma membrane. After poration,
anticancer drugs such as bleomycin which usually poorly permeable to plasma
membrane can be delivered into cells through the electropores [40, 41]. EGT uses
the modality similar to ECT but deliver genes that have anticancer activity [39,40].
IRE causes irreversible electroporation by increasing electric field intensity and
induces cell death mainly by necrosis [42, 43]. In contrast, nsPEF with higher field
intensity and much shorter pulse durations than conventional electroporation can
permeabilize not only cell plasma membrane but also the intracellular membrane.
These unique features are proposed to be responsible for apoptosis induction. Based
on here, nsPEF is used as a potential tool for cancer treatment without delivering
either drugs or genes into cells [26, 44]. Figure 1.1 shows a map guiding the
approximate locations of effects and applications due to pulsed electric field in
strength-duration space by James C. Weaver et al. [33].
5
Figure 1.1 A map of applications and effects of pulsed electric field in pulse
duration-strength space. Both values of duration and field strength are shown in
log. The figure was published by Weaver et al. [33].
The presence of apoptotic responses, including DNA fragmentation [44-46],
changes in cell morphology, phosphatidylserine externalization [47], caspase
activation [29, 48] and increase of intracellular calcium (Ca
2+
) concentration [49,
50], has been demonstrated in a variety of malignant cells in-vitro. The actual
nsPEF-induced apoptosis pathways are still not clear yet. One possible hypothesis is
the nsPEF-induced regulation of mitochondria involving apoptosis pathway. It is
known that nsPEFs can cause the disruption of mitochondrial membrane potential.
The effects on the change of potential are due to the open of mitochondrial
6
permeability transition pore complex (mPTP) [51] and/or mitochondrial membrane
voltage-dependent anion channels [52]. Thus, nsPEF can induce the release of
cytochrome c, proteins normally exiting in the inner mitochondrial membrane,
through these pores, activate caspase family, and finally lead cells to apoptosis [53,
54]. In Chapter 3, we demonstrate that multiple nsPEFs (5 or more 4 ns 10 MV/m
pulses delivered at 1 kHz repetition rate) can target intracellular mitochondrion,
increase the inner mitochondrial membrane permeability and disrupt the
mitochondrial membrane potential. At the same time, the plasma membrane
permeability was detected by fluorescent dyes with different size (YO-PRO-1 and
propidium iodide).
1.3 nsPEF and Plasma Membrane Permeability
Previously, in contrast to conventional electroporation, researchers paid
more attention on nsPEF-induced effects on intracellular organelles than the effects
on cell plasma membrane. Early papers published using nsPEFs on various types of
cells reported that biological responses occurred without permeabilizing cell plasma
membrane. The conclusion is supported by the lack of triggering an uptake of
membrane integrity markers, such as propidium iodide and Trypan blue [26, 27, 49,
55]. However, the modeling results reveal that nsPEFs form high density of pores of
nanometer scale in plasma membrane [14-16]. Collaborating with the observations
7
of nsPEF-induced phosphatidylserine externalization from the internal plasma
membrane in both simulations and living cells [20, 55-57], these results suggest that
(1) nsPEFs do affect plasma membrane and create nanopores in plasma membrane,
(2) the nsPEFs-induced nanopores are too small for fluorescent markers that are
traditionally used to detect membrane permeability, ex: PI (~1.5 nm in diameter).
Indeed, opening of small plasma membrane pores in nsPEFs-treated cells is
demonstrated by detection of YO-PRO-1 uptake and thallium (with thallium-
sensitive fluorophore) uptake before responding to PI [58, 59]. Van der Waals
diameter for YO-PRO-1 is about 1 nm and for thallium is about 0.39 nm. These
experimental data show that the nsPEF-created plasma membrane pores are smaller
than 1.0-1.5 nm. However, the sensitivity of fluorescent, membrane impermeant
dyes for the study of nsPEF-induced electropermeabilization is limited by the
number and size of pores created. By decreasing the intensity of nsPEF exposure
(either electric field strength or pulse number), only small amount of dye that can
enter into cells and the fluorescence emission from them is difficult to distinguish
from background with affordable and flexible imaging and detection systems.
Recently, a phenomenon of cell volume change (typically, cell swelling)
following nsPEF exposure was reported [60, 61]. Presumably, the increase of cell
volume is due to nsPEF-initiated water uptake triggered by osmotic imbalance
across cell plasma membrane [60, 62, 63]. It has been demonstrated for nsPEF that
the process of pore creation dominates that of pore expansion, so the transport of
8
even relatively small dye molecules through the nanometer-sized pores produced by
nsPEF exposure is much less than the flux of smaller species like calcium and
monovalent ions which cause osmotic imbalance [18]. Thus, the analysis of cell
swelling can be a more sensitive method for study of nsPEF-induced plasma
membrane permeabilization than fluorescent membrane integrity indicators. In
Chapter 4, we illustrate the mechanism of nsPEF-induced osmotic swelling in detail.
Jurkat T lymphoblasts were exposed to 5 ns, 10 MV/m pulsed electric field with
different pulse number (from 1 to 50) and pulse repetition rate (1 Hz or 1 kHz).
Standard white light microscopic images were captured by DIC channel
immediately before and after pulse exposure in different time intervals. By
systematically analyzing these images, we exhibit the dependence of nsPEF-induced
cell swelling on pulse number and repetition rate. We also observed that even a
singe pulse induces cell swelling which demonstrates that cell swelling can be an
extremely sensitive indicator for nsPEF-induced electropermeabilization.
The phenomenon of cell volume change is further used for the study of pore
populations created by nsPEFs. In brief, the size of nsPEFs-created pores can be
estimated by isosmotic replacement of pulsing solution with small molecules that
can not permeate through intact cell plasma membrane. If the size of test molecule
small enough to pass the nsPEF-created pores, cell swelling will take place. In
contrast, if the size of test molecule prohibits it from going through the nsPEF-
created pores, cell swelling will be replaced with shrinking. Therefore, the pore size
9
can be estimated from the dimension of test molecules [64]. In Chapter 6, we use
neutral sugars with different sizes and PEG 1000 to estimate the diameter of pores
created in Jurkat T lymphoblast cell plasma membranes by nsPEF exposure. Various
numbers of pulses of 5 ns duration, 10 MV/m field strength were delivered at 1 kHz.
The dependence of pore size on the pulse number is also analyzed and discussed.
10
1.4 Parameters that Affect nsPEF-Induced Responses
The physiological response of biological cells to nsPEFs depends on specific
pulse parameters, which include pulse shape, pulse duration, electric field
amplitude, pulse repetition rate, pulse number and electrode arrangement. For
example, Figure 1.2 shows pulse numbers and field amplitude affect the amount of
nsPEFs-induced YO-PRO-1 uptake by Jurkat T lymphoblasts. In addition to pulse
parameters, the efficiency of nsPEF is also related to cell type, cell concentration
and even the ambient temperature. Therefore, to optimize protocols for nsPEF
exposure, experimental studies that provide database with different variables are
important. From the established information, researchers can predict the biological
effects caused by nsPEFs and design the pulsing protocol without guess and time
consuming. We presented most of our works here with different pulse parameters,
especially in different pulse numbers and repetition rates. As well known, multiple
pulsing can not be considered as causing a simple additive effects. In Chapter 4, we
extract the swelling data and built an empirical scaling law to describe the nsPEFs-
caused plasma membrane permeabilization.
11
Figure 1.2: Increases in intracellular fluorescence intensity resulting from YO-
PRO-1 influx, indicating electropermeabilization, are greater for 4 ns pulses
than for 3 ns pulses at 10 MV/m. Doubling the field from 10 MV/m to 20 MV/m
increases permeabilization ten-fold. Data represents the ratio of YO-PRO-1
fluorescence emission 3 minutes after and immediately before pulse exposure
from the integrated intensity of at least 10 cells for each condition from each of
3 independent experiments. YO-PRO-1 (5 µM): Invitrogen; λex: 491 nm, λem:
509 nm; Filter set: Emission filter: 480/30 nm, Dichroic mirror: 505D,
Excitation filter: 535/40.
The possibility of chemical and physical agents that affect nsPEF-induced
electropermeabilization is also discussed. In Chapter 1, we investigate the role of
oxidative stress in plasma membrane because oxidative stress is commonly
encountered in cultured cells and in whole organisms under a variety of conditions.
The results demonstrate that peroxidative cells are more susceptible to nsPEFs-
12
induced electropermeabilization. This suggests that an appropriately controlled
peroxidizing regimen may enhance the efficiency of the applications requiring
membrane electropermeabilization, such as electrotransfection of genetic materials,
without significantly decreasing the cell viability.
In Chapter 4, we investigate the effects of lanthanide ions (Gd
3+
and La
3+
)
and mercury ions on nsPEF-induced permeabilization. Lanthanide ions are known
as the membrane ion channel blockers and mercury ion is the membrane aquaporin
channel blocker. Lanthanide ions have been found to inhibit the nsPEF-induced
membrane permeabilization, and also prevent swelling and blebbing in severely
exposed cells [55]. Andre et al. reported that Gd
3+
, even in sub-micromolar
concentrations, can weaken or revert electropermebilization of the plasma
membrane by 60 and 600 ns pulsed electric fields, and increase cell viability after
pulse exposure [65]. However, we did not observe the membrane protection effects
by lanthanide ions in our work with 5 ns pulses. These results suggest that longer
(60 and 600 ns) and shorter (5ns) pulses may affect membrane structure in different
ways.
13
1.5 nsPEF Delivery Systems
Design of a delivery system for nanosecond, megavolts per meter electric
pulses is a challenge. The delivery device must be compatible with the biological
sample and does not allowed the distortion of pulse shape and amplitude.
The most common available delivery device is commercialized
electroporation cuvette. The cuvette chamber consists of two stainless steels or
aluminum electrodes with 1 mm separation in molded plastic. This design is suitable
to expose suspension of large number of cells. By coupling with appropriately
designed pulse generator for cuvette (10 ohm load), the cuvette allows the delivery
of tens nanosecond electric pulses with field intensity up to 5 MV/m [66]. Figure
1.3 shows the delivery system with cuvette load that were designed and constructed
in USC. This system is also used in the work presented in Chapter 2 in this thesis.
14
Figure 1.3: Constructed pulse generator with a cuvette load by Tang et al. in
USC [66].
However, the cuvette does not allow real time observation of cells during
and immediately after nsPEF exposure. Also, Kenaan et al. reported that, at
frequencies higher than 100 MHz, the cuvette presents not negligible mismatching
the pulse exposure which suggests that cuvette is suitable for pulses longer than 10
ns [67].
15
To enable real time imaging and investigation of cells exposed to pulses
shorter than 10 ns, Sun et al. designed and fabricated a microchamber utilizing
photolithographic and microelectronic methods [68]. SU-8 photoresist was patterned
on a transparent glass slide to form straight sidewalls and further deposited with
gold film for conductive, nonreactive electrodes and a uniform electric field. The
dimension of microchamber is 100 µm wide, 30 µm deep, and 15 mm long, as
shown in Figure 1.4.
Figure 1.4: Four-channel electrode microchamber on a standard microscope
glass slide. Top: ideal electrode chamber with straight sidewalls and appropriate
dimensions. Bottom: top view of the electrode microchamber [68].
16
We have performed a lot of experiments with the microchamber as presented
in Chapter 3 and 4. However, there are some drawbacks of this device. Cells are
easily floating inside the chamber which makes observation and image recording
difficult. Moreover, microchamber does not permit long-term observation, ex: 24
hours post-exposure. In Chapter 5, we report the development of a new delivery
system that integrated fluorescent microscope, nanosecond pulse generator and
delivery device based on tungsten-wire electrodes. This delivery system solves the
problems existing in microchamber and improves the experimental flexibility (ex:
easy to locate cells, flexible observation duration).
17
Chapter 2: Lipid Peroxidation in Living
Cells Promotes Plasma
Membrane
Electropermeabilization
2.1 Abstract
Pulsed electric field technology is widely used to facilitate the delivery of
genetic material and pharmaceutical agents into living cells. To optimize pulsing
protocols, a better understanding of the factors affecting the susceptibility of cells to
electric pulse exposure is needed. In this chapter, the effects of membrane oxidation
on electropermeabilization sensitivity are investigated by molecular dynamics (MD)
simulations and in-vitro cell experiments. Jurkat T lymphoblast cells were gently
peroxidized and then exposed to 30 ns, 3 MV/m pulses containing YO-PRO-1, a
membrane-impermeant dye that fluoresces significantly only when cell membranes
become permeabilized. The results reveal that peroxidation significantly increases
nanoelectropulse-induced influx of YO-PRO-1 into cells. In MD simulations we
applied electric fields to phospholipid bilayers containing varying concentrations of
oxidized lipid species. Systems with higher concentrations of oxidized lipids form
hydrophilic electropores in significantly shorter times and at lower electric fields
than do systems with lower oxidized lipid concentrations. Sites of water defect
18
formation and electroporation appear to coincide with the clustering of oxidized
lipids in the bilayer. In large-area simulations containing localized highly oxidized
lipid concentrations, pores formed preferentially in these oxidized regions.
2.2 Introduction
Recently, pulse electric field induced electropermeabilization technology is
reported as an useful tool for applications in laboratories and clinics, such as cell
biology, genetic engineering, and cancer therapy [31, 69-74]. To optimize pulsing
protocols for specific applications, a better understanding of the mechanisms of
electropermeabilization and of the factors affecting the susceptibility of cells to
electric pulse exposure is needed.
In addition to knowledge of the effects of varying the relevant physical
parameters (electric field strength and duration, temperature, cell concentration,
composition of the medium, electrode arrangement) [75-77], optimizing the
efficiency of electropermeabilization methods requires an understanding of the
relation between the physiological state of cells and their susceptibility to
electropermeabilization. For example, it may be that starved cells can be most
effectively treated under conditions that are very different from those that are
optimal for actively respiring cells.
Oxidative stress is readily imposed and commonly encountered in cultured
19
cells and in whole organisms under a variety of conditions. It not only impacts
cellular activities across the metabolic spectrum but also directly affects the physical
properties of the cell membrane [78-80]. Because of these reasons, we speculated
that this might be an important factor in the effectiveness of electroporation
methods. Studies of the peroxidation of membrane lipids after electroporation have
been reported [81-84]. However, the effects of pre-treatment oxidative stress have
received little experimental attention. Molecular dynamics simulations have recently
shown that incorporating oxidized lipids into phospholipid bilayers increases the
water permeability of these membranes [85], suggesting that bilayers containing
oxidized lipids will also electroporate more readily (the formation of membrane-
spanning water defects is one of the initial steps in molecular dynamics
representations of electroporation). In this work, we investigated the effect of
membrane oxidative damage on the sensitivity of cells by exposing Jurkat cells to
30 ns 30 ns, 3 MV/m electric pulses. In parallel to experiments of living cells,
molecular dynamics simulations of lipid bilayers using varying concentrations of
oxidized lipids with were perform to validate the hypotheses [86].
20
2.3 Materials and Methods
2.3.1 Cell Experiments
Cell Line and Cell Culture
Jurkat T lymphoblasts (ATCC TIB-152) were grown in RPMI 1640 medium
(Mediatech) containing 10 % heat-inactivated fetal bovine serum (FBS; Mediatech),
2 mM L-glutamine (Invitrogen), 50 units/mL penicillin (Gibco), and 50 µg/mL
streptomycin (Gibco) at 37 °C in a humidified, 5 % carbon dioxide atmosphere.
Cell Preparation
For membrane peroxidation, Jurkat cells were exposed to peroxidizing
conditions — H
2
O
2
(200 and 500 µM) and FeSO
4
(400 and 1000 µM) in RPMI
1640 — for 15 minutes before pulse exposure. After peroxidation, 5 µM YO-PRO-1
(Molecular Probes, Invitrogen) was added as a permeabilization indicator, and the
cells were immediately pulsed.
Pulse Generator and Pulse Exposure
30 ns, 3 MV/m pulses were delivered at a 50 Hz repetition rate to cell
suspensions in commercial electroporation cuvettes (VWR) with 1 mm electrode
spacing from a USC pulse generator based on a magnetic compression, diode
opening-switch architecture [66].
21
Fluorescence Microscopy
Images were captured and analyzed with a Zeiss AxioCam MRm and
AxioVision 3.1 software (Carl Zeiss Goettingen, Germany) on a Zeiss Axiovert
200M epifluorescence microscope. Intracellular YO-PRO-1 (λex = 480 nm, λem =
535 nm) fluorescence was used as a sensitive indicator of membrane
permeabilization [58]. Membrane lipid peroxidation was monitored qualitatively
with the fluorescent dye C11-BODIPY
581/591
[87]. Upon oxidation, the dye shifts
from a red-emitting from (595 nm) to a green-emitting form (520 nm), which
increases the green:red emission ratio. Cells were incubated in RPMI 1640 medium
containing 4 µM C11-BODIPY
581/591
for 30 minutes at 37 °C before exposure to
peroxidizing reagents.
2.3.2 Simulations
Molecular Dynamics Simulations
All simulations were performed using the GROMACS set of programs
version 3.3.1 [88] on the University of Southern California High Performance
Computing and Communications Linux cluster (http://www.usc.edu/hpcc/). Lipid
parameters were derived from OPLS, united-atom parameters [89] modified for
PLPC and oxPLPC [90]. We used the Simple Point Charge (SPC) model for water.
Systems were coupled to a temperature bath at 310K with a relaxation time of 0.1 ps
and a pressure bath at 1 bar with a relaxation time of 1 ps, each using a weak
22
coupling algorithm [91]. Pressure was scaled semi-isotropically with a
compressibility of 4.5×10
-5
bar
-1
in the plane of the membrane and 4.5×10
-5
bar
-1
perpendicular to the membrane. Bond lengths were constrained using LINCS [92]
for lipids and SETTLE [93] for water. Short-range electrostatics and Lennard-Jones
interactions were cut off at 1.0 nm. Long-range electrostatics were calculated by the
PME algorithm [94] using fast Fourier transforms and conductive boundary
conditions. Reciprocal-space interactions were evaluated on a 0.12 nm grid with
fourth order B-spline interpolation. The parameter ewald_rtol, which controls the
relative error for the Ewald sum in the direct and reciprocal space, was set to 10
-5
.
Periodic boundary conditions were employed to mitigate system size effects.
23
Electroporation Simulations
To determine a baseline porating electric field [95], simulations of
equilibrated (constant area per lipid), fully hydrated (40 water/lipid) PLPC bilayer
systems containing 72 lipid molecules and 2880 water molecules [85] were run with
applied electric fields ranging from 300 to 500 mV/nm. The lowest field which
results in pore formation within 25 ns in at least one of three parallel PLPC
simulations is 360 mV/nm, and that value was used for comparisons of pore
formation times in oxidized (PLPC:oxPLPC) and non-oxidized (PLPC) systems.
Oxidized lipid systems included 8 (11.1%), 18 (25%), or 36 (50%) randomly
distributed molecules of oxPLPC, either 12-al or 13-tc [85].
Electroporation times were calculated by measuring at each time step the
number of phosphorus atom groups in a system, where a group is a cluster of
phosphorus atoms each no more than 1.2 nm from another phosphorus atom [96].
The two phosphorus atom groups in an intact bilayer (one in each of the two leaflets)
merge when the bilayer interior is bridged by a hydrophilic pore. We call the time at
which this occurs the electroporation time. In some simulations the merged groups
split again, in each case in less than 400 ps after merger. This was not considered
pore formation.
Assembly of Larger Systems
Systems with four times the area were created by doubling pure PLPC
systems in x and y using the GROMACS function genconf. Using custom code,
24
oxidized groups were inserted on the sn-2 linoleate tails in two opposite quadrants to
create quilted systems where two quadrants contained pure PLPC and two quadrants
contained 50 % oxPLPC, either 12-al or 13-tc. For 12-al an aldehyde group was
added at C12. For 13-tc, a hydroperoxy group was added at C13, and the double
bond at C12 was shifted to C11. The system was energy-minimized for 1 ps using
the GROMACS steepest descent method to test for bad contacts each time a new
oxidized group was inserted. As with the smaller systems, the oxPLPC lipids were
distributed equally between the two leaflets of the bilayer, and the water-to-lipid
ratio was 40. Each system was then equilibrated long enough (11.5 ns for 12-al, 32.5
ns for 13-tc) to bring the area per lipid within 2 % of its final equilibrated mean
value while maintaining the four-quadrant distribution of the lipids.
Images
Molecular graphics images were generated with Visual Molecular Dynamics
(VMD) [97].
25
2.4 Lipid Peroxidation Enhances Membrane
Electropermeabilization
To peroxidize membrane lipids, Human Jurkat T lymphoblasts were treated
with sublethal doses of peroxidizing agents, ferrous sulfate
(0, 400, 1000 µM) and
hydrogen peroxide (0, 200, 500 µM), in culture medium (RPMI 1640) for 15
minutes at 37 °C. In addition, cells were also treated with hydrogen peroxide and
ferrous sulfate alone to examine the effects of the individual reagents. Conditions
for the peroxidation treatment were developed by maximizing the green:red
fluorescence emission ratio from the membrane-staining fluorescent dye C11-
BODIPY
581/591
[87] while at the same time minimizing changes in cell morphology
and membrane integrity. Figure 2.1a displays the representative images of cells
stained with C11-BODIPY
581/591
and treated with different doses of H
2
O
2
and FeSO
4
.
Figure 2.1b shows the integrated ratio C11-BODIPY
581/591
fluorescence intensity
(green/red) of cells treated with the peroxidizing reagents as the dosage used in
nsPEF exposures in this work.
26
Figure 2.1: Membrane lipid peroxidation visualized with C11-BODIPY
581/591
.
(a): Fluorescence images of Jurkat cells showing C11-BODIPY
581/591
in control
and peroxidized cells. (b): Integrated ratio C11-BODIPY
581/591
fluorescence
intensity from more than 300 individual cells from three independent
experiments for each condition. Error bars are standard error of the mean.
27
Cells were then exposed to 30 ns, 5 MV/m electric pulses after lipid
peroxidation and the membrane permeability was monitored by the influx of the
fluorescent dye YO-PRO-1, a sensitive indicator of membrane permeabilization [58].
Results are shown in Figure 2.2. Fluorescence imaging analysis reveals that pulse-
induced YO-PRO-1 uptake in peroxidized Jurkat cells is significantly higher than in
non-oxidized cells.
The results demonstrate that the lipid peroxidation enhances the
electropermeabilization. On the other hand, reducing the oxidative stress in
environment might be expected to protect cells against electropermeabilization. This
hypothesis suggests possible applications by appropriately controlling peroxidizing
regimen. For example, by employing dose which increases permeabilization without
significantly affecting viability, we might increase efficiency of electrotransfection
protocols either indirectly by enabling the use of lower porating voltages (higher
voltages are associated with lower cell survival rates) or directly by increasing the
amount of genetic material that enters the cells for a given series of electrical pulses.
On the contrary, electrochemotherapy and direct ablation and killing of tumor cells
using electrical pulse therapy might be enhanced by procedures which promote
oxidative stress in the tumor tissue before pulse delivery [98]. In any mixed
population of cells, the different native sensitivities of various cell types to
permeabilizing electric fields [29, 99, 100] and oxidative stress might be exploited
by adjusting electrical, physical, and chemical parameters to selectively transfect
28
subsets of cells, in-vitro and in-vivo.
Figure 2.2: Electropermeabilization enhancement by treatment with
peroxidation agents. (A). Fluorescence images of Jurkat cells showing pulse-
induced (30 ns, 3 MV/m, 50 Hz) YO-PRO-1 influx into control and peroxidized
cells after 10 min exposure to 500 µM H
2
O
2
+1000 µM FeSO
4
. (B). Integrated
YO-PRO-1 fluorescence intensity from more than 300 individual cells from
three independent experiments for each condition. Error bars are standard error
of the mean.
electroporation may lead to a more sophisticated approach to
controlling electrotransfection yield, including both the efficiency
of the incorporation of the genetic material and the subsequent
viability of the cells, which may be decreased by pulse-induced
apoptosis. Similar considerations apply to potential improvements
in electroporation protocols. Thus the knowledge gained from an
exploration of the interplay between nanosecond membrane
permeabilization [17,58] and conventional electroporation [59]
in the presence of excess reactive oxygen species and the initiation
of apoptosis may result in improved electroporation and electro-
transfection procedures, with benefits from cell science to cancer
therapeutics.
Permeabilizing Defects and Membrane Boundaries
Facilitationofelectricfield-driven waterdefectformationbythe
aldehyde and hydroperoxy oxygens of the oxPLPC species
reported here may in fact be a relatively simple example of a
more general tendency for water intrusion, permeabilization, and
othermembranerestructuring eventsto occuratmembranephase
or domain boundaries, especially where these discontinuities have
charged or hydrophilic components that extend into the
membrane interior [44,60–62]. In this context it will no doubt
prove instructive to examine the combined effects of lipid
peroxidation and cholesterol on membrane electropermeabiliza-
tion. Adding cholesterol to a bilayer is likely to render the
membrane more difficult to electropermeabilize [63], but the
consequences of incorporating peroxidized lipids, which promote
the formation of cholesterol domains and lipid rafts [64,65], into
the system are difficult to predict. Increasing computationalpower
makes it possible now to address these more complex (and more
realistic) systems with an approach that ties together atomically
detailed simulations and experimental cell biology.
Materials and Methods
Cell Lines and Culture Conditions
Jurkat T lymphoblasts (ATCC TIB-152) were grown in RPMI
1640 medium (Mediatech) containing 10% heat-inactivated fetal
bovine serum (FBS; Mediatech), 2 mM L-glutamine (Invitrogen),
50 units/mL penicillin (Gibco), and 50 mg/mL streptomycin
(Gibco) at 37uC in a humidified, 5% carbon dioxide atmosphere.
DC-3F adherent cells (Chinese hamster lung fibroblast cells) were
grown in Minimum Essential Medium (Invitrogen, France)
containing 10% heat-inactivated FBS (Invitrogen), 500 U/ml
penicillin, 500 mg/ml streptomycin (Invitrogen) at 37uC in a
humidified, 5% carbon dioxide atmosphere.
Cell Preparation
Jurkat cells were exposed to peroxidizing conditions — H
2
O
2
(200 and 500 mM) and FeSO
4
(400 and 1000 mM) in RPMI 1640
—for15 minutesbeforepulseexposure.Afterperoxidation,5 mM
YO-PRO-1 (Molecular Probes, Invitrogen) was added as a
permeabilization indicator, and the cells were immediately pulsed.
DC-3F cells were incubated with 1 mM calcein-AM (Sigma-
Aldrich, France) in culture medium for 1 hour at 37uC, then
rinsed with PBS, trypsinized, and centrifuged at 1000 rpm for
10 minutes. For peroxidation, DC-3F cells were incubated with
H
2
O
2
and FeSO
4
(each 1000 mM for long pulse experiments,
1500 mM for short pulse experiments) for 1 hour at 37uC, rinsed
withPBSand centrifuged at1000 rpmfor 10 minutes.Thepellets
were suspended in sucrose buffer (250 mM sucrose, 10 mM Tris,
1 mM MgCl
2
— Sigma), pH 7, containing 2% low-melting
agarose (Tebu-Bio, France) at about 50 000 cells/mL. Cell
suspensions were kept at 37uC until electric pulse exposure.
Pulse Generator and Pulse Exposures
For the Jurkat cell experiments 30 ns, 3 MV/m pulses were
delivered at a 50 Hz repetition rate to cell suspensions in
commercial electroporation cuvettes (VWR) with a 1 mm
electrode spacing in ambient atmosphere at room temperature
from a USC pulse generator based on a magnetic compression,
diode opening-switch architecture [66]. For the DC-3F experi-
ments the cell suspension was placed within the 2 mm gap
between two copper electrodes fixed to a microscope slide. Cells
wereexposedto1longpulse(100 ms,50or60 kV/m)deliveredby
a micropulse generator (Cliniporator, IGEA, Italy) or to 1000
Figure 4. Electropermeabilization enhancement by treatment
with peroxidation agents. (A). Fluorescence images of Jurkat cells
showing pulse-induced (30 ns, 3 MV/m, 50 Hz) YO-PRO-1 influx into
control and peroxidized cells after 10 min exposure to 500 mM
H
2
O
2
+1000 mM FeSO
4
. (B). Integrated YO-PRO-1 fluorescence intensity
frommorethan300individualcellsfromthreeindependentexperiments
for each condition. Error bars are standard error of the mean.
doi:10.1371/journal.pone.0007966.g004
Porating Oxidized Membranes
PLoS ONE | www.plosone.org 5 November 2009 | Volume 4 | Issue 11 | e7966
29
2.5 MD Simulations Show Evidences of
Electropermeabilization Enhanced by Oxidized
Membrane Lipids
In addition to experimental observations, we applied electric fields
during molecular dynamics simulations of phospholipid bilayers (PLPC)
containing varying concentrations of oxidized lipid species, which are in part
comprised of peroxidized linoleic acid derivatives containing either an aldehyde
group (12-al oxPLPC) or a hydroperoxide group (13-tc oxPLPC), to validate the
effects of oxidative stress in electropermeabilization. The structures of
individual lipid molecules are shown in Figure 2.3.
Figure 2.3: Molecular structures of PLPC and two oxidized variants, oxPLPC
(12-al) and oxPLPC (13-tc).
30
The stabilization associated with hydration of the tail oxygens in the
oxidized lipids results in a tendency for the oxidized tails to spend more time
near the aqueous interface than the hydrocarbon tails of PLPC. Thus, Increasing
the oxidized lipid content decreases the bilayer thickness, increases the average
area per lipid, and reduces the time to poration in an external electric field [88].
Moreover, the observations from MD simulation suggested that the site
of pore initiation in MD simulations is specifically associated with single or
aggregated oxPLPC molecules, as shown in Figure 2.4. A more clear view of
this phenomenon is shown in Figure 2.5 with large quilted systems containing
50 % oxPLPC or only PLPC.
31
Figure 2.4: Consecutive snapshots of a system with an 11 % concentration of
12-al oxPLPC. In the first snapshot all lipids are displayed whereas in all
subsequent snapshots, only the oxidized sn-2 tails are shown. Carbon is colored
cyan and oxygen is colored in red. The enlarged red spheres represent aldehyde
oxygens which appear to attract water deep into the bilayer interior, thus helping
mediate the process of pore formation.
32
Figure 2.5: Large quilted system which contains PLPC lipids (light gray) and
selectively placed oxPLPC lipids (dark blue) which were inserted by replacing
PLPC lipids with oxPLPC lipids in opposing quadrants. The concentration of
oxPLPC in an oxidized patch is 50 %. After an electric field is applied, it
appears that there is a clear correlation between pore location and local
oxidation sites.
2.6 Conclusion
In conclusion, we demonstrate oxidized lipid membrane enhances the
nsPEF-induced membrane permeability by in-vitro cell experiments and MD
simulations. Cells pre-treated with peroxidizing reagents (H
2
O
2
+ FeSO
4
) show
significant increase in membrane permeabilization after exposure to 30 ns, 5
MV/m electric pulses. The result is consistent with MD simulations in which
PLPC bilayers containing higher concentration of oxidized lipids form
33
hydrophilic electropores in significantly shorter times than do those with lower
oxidized lipid concentrations. Sites of water defect formation and subsequent
electroporation appear to coincide with local clustering of oxidized lipids in the
bilayer. The presence of aldehyde and hydroperoxy oxygens on an otherwise
nonpolar lipid tail appears to facilitate the penetration of water into the bilayer
interior.
34
Chapter 3: The Effects of Nanosecond
Pulsed Electric Field on
Mitochondrial Membrane
3.1 Abstract
Nanosecond, high-voltage pulsed electric fields (nsPEFs) induce
permeabilization of the plasma membrane and the membranes of cell organelles,
leading to various responses in cells including cytochrome c release from
mitochondria and caspase activation associated with apoptosis. In this chapter, we
provide the evidence of nsPEFs-induced permeabilization of mitochondrial
membranes in Jurkat T lymphoblasts by employing three different methods with
fluorescence indicators—rhodamine 123 (R123), tetramethyl rhodamine ethyl ester
(TMRE), and cobalt-quenched calcein. The results demonstrate that multiple
nsPEFs (5 or more 4 ns 10 MV/m pulses delivered at 1 kHz repetition rate)
increased the inner mitochondrial membrane permeability and an associated loss of
mitochondrial membrane potential. In addition to mitochondrial membrane
permeabilization, nsPEFs-induced plasma membrane permeabilization was detected
by the influx of YO-PRO-1.
35
3.2 Introduction
Cell plasma membrane permeabilization (electroporation) by intense pulsed
electric fields has been studied for several decades and widely used in research,
biotechnology and medicine for delivering of foreign molecules into cells. Recently,
the ability of reducing the pulse duration less than the membrane charging time
constant (human lymphocytes in growth medium the charging time constant for the
plasma membrane lies in the range of 50 to 100 ns) showed that high intensity,
nanosecond, pulsed electric fields (nsPEFs) are possible to target intracellular
organelles and induce subcellular effects.
Previous studies of nsPEFs-induced subcellular effects demonstrate that
nsPEFs with durations less than 100 ns and electric field strengths over 1 MV/m can
permeabilize intracellular granules and large endocytosed vacuoles [101-103],
trigger calcium ion release [49, 104], induce externalization of phosphatidylserine
[57], damage cell nuclei and DNA [46, 105], and cause platelet activation [106].
Moreover, they can also provoke apoptosis [29, 44] and are considered as a
promising new tool for cancer therapy [31, 107, 108].
Apoptosis is an evolutionarily conserved type of programmed cell death
essential for development, homeostasis, and self-defense against infection.
Mitochondria play a crucial role in apoptosis due to the participation of the intrinsic
pathways of apoptosis [109]. Mitochondria regulate the release of several apoptosis-
36
inducing proteins from the space between the inner and outer mitochondrial
membranes to the cytosol, such as cytochrome c [110], second mitochondria-derived
activator of caspase/direct IAP-binding protein with low pI (Smac/Diablo) [111],
and AIF (apoptosis-inducing factor) [112]. The release of these proteins is through
the mitochondria membrane permeabilization (MMP) controlled by B-cell
lymphoma 2 (Bcl-2) family and/or disruption of the outer mitochondrial membrane,
following the mitochondrial permeability transition [109, 113]. When cytochrome c
is released from mitochondria, it binds to apoptotic peptidase activating factor 1
(Apaf-1). Activated Apaf-1 then forms complexes with pro-Caspase 9. This results
in the activation of caspase cascade and induction of apoptosis [114]. Release of
cytochrome c from mitochondria after applying nsPEFs to cells has been reported
[115, 116], and mitochondrial membrane potential was decreased in a portion of
cells after exposure to 300 ns pulses [53], even in the absence of cytochrome c
release [117].
Theoretical evaluation predicts that mitochondria could be targeted by
nsPEFs [118] and suggests that a mitochondria-dependent apoptosis could be due to
electroporation of these organelles. Therefore, we investigated the effects of nsPEF
(4 ns, 10 MV/m) on mitochondrial membrane permeability and mitochondrial
membrane potential. A decrease in mitochondrial membrane potential indicates a
change in the ability of the cell to maintain proton and other ion concentration
gradients across the inner mitochondrial membrane [109]. We monitored
37
mitochondrial membrane potential with rhodamine 123 (R123) and tetramethyl
rhodamine ethyl ester (TMRE), fluorescent lipophilic cationic dyes that accumulate
in mitochondria in a potential-dependent manner [119, 120]. We also employed a
cobalt-quenched calcein method for detecting mitochondrial membrane
permeabilization [121]. In addition, we monitored plasma membrane
permeabilization by looking for influx of the fluorescent dyes YO-PRO-1 and
propidium iodide (PI). Our results are consistent with pulse-induced
permeabilization of mitochondrial membranes with an associated loss of
mitochondrial membrane potential [122].
3.3 Materials and Methods
3.3.1 Cell Line and Cell Culture
Human Jurkat T lymphoblasts (obtained from American Type Culture
Collection (ATCC), Manassas, VA; ATCC cell number: TIB-152) were grown in
RPMI-1640 medium (Mediatech, Manassas, VA) containing 10% heat-inactivated
fetal bovine serum (Gibco/Invitrogen, Carlsbad, CA), 2 mM L-glutamine
(Gibco/Invitrogen), 50 units/ml penicillin (Gibco/Invitrogen), and 50 mg/ml
streptomycin (Gibco/Invitrogen). Cells were cultured at 37 °C in a humidified, 5 %
carbon dioxide atmosphere and concentrated to 2x10
7
cells/ml for pulse treatment.
38
3.3.2 Pulse Generator and Pulse Exposure
For microscopic observation, cells were placed in a microchamber 100 mm
wide, 30 mm deep, and 15 mm long with platinum electrode walls, on a glass
microscope slide. A resonant-charged, solid state Marx bank-driven, hybrid-core
compression, diode-opening switch pulse generator, designed and assembled at the
University of Southern California (Los Angeles, CA) [123], delivered 4 ns, 10
MV/m electrical pulses at a 1 kHz repetition rate to the microchamber electrodes
mounted on the microscope stage in ambient atmosphere at room temperature. The
impedance of the microchamber load, when it was loaded with a cell suspension in
our standard media, was about 350 Ω. Setup of pulsing experiments described above
is shown in Figure 3.1.
39
Figure 3.1: Experimental setup for pulse exposure. Cells were loaded in the
homemade michcrochamber lay on a specially designed board with voltage
divider on it. The board was connected to the pulse generator and oscilloscope,
and located on the fluorescence microscope beforehand. This setup made the
real-time observation of pulsing experiment happen.
3.3.3 Fluorescence Microscopy and Fluorescent Molecular Probes
Observations of live cells during pulse exposures were made using a Zeiss
Axiovert 200 epifluorescence microscope (Zeiss, Go¨ttingen, Germany) with a 63x
water immersion objective and a Hamamatsu ImageEM EM-CCD camera
(Hamamatsu Photonics KK, Hamamatsu City, Japan). Captured images were
analyzed with Hamamatsu SimplePCI software. Cells were stained with the
fluorescent probes for mitochondria (cobalt-quenched calcein, R123, TMRE, and
MitoTrackers) and for plasma membrane permeability (PI and YO-PRO-1).
40
3.3.4 Fluorescent Molecular Probes
Cobalt-quenched calcein. Calcein-AM (Molecular Probes/Invitrogen,
Carlsbad, CA) is an anionic fluorochrome that enters cells freely and labels
cytoplasmic as well as mitochondrial regions following removal of the
acetoxymethyl (AM) group by intracellular esterases. Because cobalt ions are taken
up by cells but do not readily pass through the mitochondrial membrane,
mitochondria can be identified by the cobalt quenching of cytoplasmic, but not
mitochondrial, calcein fluorescence [121, 124]. Cells were loaded with 500 nM
calcein-AM in the presence of 1 mM CoCl
2
at 37 °C in 5 % CO
2
atmosphere for 20
min. Before pulse treatment, cells were washed and resuspended in fresh RPMI-
1640 medium with or without 1 mM Co
2+
. Fluorescent images were captured before
pulsing and 3 min after pulsing, using the appropriate filter (Ex 482/35, Em 536/40,
and D 506DCLP). For quantification, the cells in the images were encircled and the
fluorescence of cells was measured. The background fluorescence was subtracted.
The results were expressed relative to cells before pulsing.
Mitochondrial membrane potential. Rhodamine 123 (R123) and TMRE
were used to assess changes in mitochondrial membrane potential [119, 125, 126].
Cells were incubated with mM R123 (Molecular Probes/Invitrogen) for 45 min, or
50 nM TMRE (Molecular Probes/Invitrogen) for 20 min. After incubation, cells
were centrifuged and resuspended in fresh RPMI-1640 medium for pulse exposure.
41
Fluorescent images were captured before pulsing and 30 s and 3 min after pulsing,
using the appropriate filters (R123: Ex 482/35, Em 536/40, D 506DCLP; TMRE: Ex
543/22, Em 593/40, D 562DCLP). For quantification, the cells in the images were
encircled and the fluorescence of cells was measured. The background fluorescence
was subtracted.
For a positive control, the proton ionophore uncoupler of oxidative
respiration carbonyl cyanide 3-chlorophenylhydrazone (CCCP) was added to the
R123- or TMRE-stained cells in suspension at 20 mM [127]. Images were captured
and quantified the same way as pulsed cells, only the results were expressed relative
to cells that were not treated with CCCP.
MitoTrackers. MitoTracker Green and MitoTracker Orange (Molecular
Probes/Invitrogen) were used to confirm the labeling of mitochondria by double
labeling (MitoTracker Orange with cobalt-quenched calcein and R123, and
MitoTracker Green with TMRE). Cell loading was performed according to the
manufacturer’s protocol (100 nM MitoTracker Orange and 50 nM MitoTracker
Green). Images of double-stained cells were captured by using appropriate filters
(MitoTracker Orange: Ex 543/22, Em 593/40, D 562DCLP; MitoTracker Green: Ex
482/35, Em 536/40, D 506DCLP).
42
Plasma membrane permeability. Intracellular YO-PRO-1 (Molecular
Probes/Invitrogen) and PI (Molecular Probes/Invitrogen) fluorescence were used as
indicators of plasma membrane permeabilization as described previously [58]. Cells
were suspended in RPMI-1640 medium with 10 µM YO-PRO-1 or 7.5 µM PI just
before pulsing. Fluorescent images were captured before pulsing and 3 min after
pulsing, using the appropriate filters (YO-PRO-1: Ex 472/30, Em 520/35, D
495DCLP; PI: Ex 562/40, Em 624/40, D 593DCLP). For quantification, cells in the
images were encircled and the fluorescence of cells was measured. The background
fluorescence was subtracted. The results were expressed in arbitrary units.
3.4 Localization of Mitochondria
In order to localize the position of mitochondria in Jurkat T lymphoblasts for
nsPEFs experiments, we employed three fluorescent labeling methods: (1) cobalt-
quenched calcein, (2) rhodamine 123 (R123), and (3) TMRE. For all of them,
mitochondria in cells are seen as bright spots in the cytoplasm because of the
accumulation of fluorophores inside mitochondria. In the meantime, we stained cells
with widely used mitochondria trackers, MitoTracker Orange or MitoTracker Green,
to confirm the efficacy of labeling methods used in pulsing experiments. As shown
in Figure 3.2, the images revealed almost complete overlap of the MitoTracker dyes
with the structures stained with R123 and TMRE. The bright particles in calcein-
43
stained cells are not dense enough to merge over the entire intracellular area labeled
by the MitoTracker dyes, but the calcein-labeled regions are coincident with the
MitoTracker-labeled regions.
Figure 3.2: Localization of mitochondria in Jurkat cells. Cells were treated with
cobalt-quenched calcein assay (A), Rhodamine 123 (B) or TMRE (C), and
incubated with either MitoTracker Orange (A, B) or MitoTracker Green (C)
together. A1-C1: permeabilization indicator images. A2-C2: Mitotracker stained
images. A3-C3: superimposed images.
44
3.5 nsPEF Cause a Loss of Mitochondrial Potential
R123 and TMRE are lipophilic cationic dyes that accumulate in active
mitochondria because of the large negative potential that appears across normal
mitochondrial membranes. When the mitochondrial membrane potential decreases,
the concentration of these dyes in the mitochondria decreases with an associated
decrease in R123 and TMRE fluorescence. The idea of employing R123 and TMRE
as a detector for nsPEFs-induced mitochondrial membrane potential change is
described in Figure 3.3. Our results show that nsPEFs cause a significantly reduced
fluorescence of both dyes, indicating a loss of mitochondrial membrane potential
(Figures 3.4 and 3.5). The extent of pulse-induced mitochondrial membrane
permeabilization depends on the number of pulses—more pulses cause a greater
reduction in R123 and TMRE fluorescence. At 10 pulses (4 ns, 10 MV/m, 1 kHz
repetition frequency), R123 and TMRE fluorescence decreased to 76% and 78% of
pre-pulse fluorescence, respectively. At 50 pulses, R123 and TMRE fluorescence
decreased to 63% and 30% of the pre-pulse fluorescence emission, respectively. In
our data, TMRE exhibited a more rapid decrease in fluorescence emission compared
to R123. This phenomenon might be explained by the natural of R123 and TMRE.
R123 and TMRE both are rapidly and reversibly equilibrated across membranes in a
voltage-dependent manner. However, TMRE shows less specific binding and more
membrane permeable, therefore more suitable for dynamic measurements [128,
45
129]. CCCP, a proton ionophore uncoupler that causes a loss of mitochondrial
membrane potential [127], was used as a positive control. After exposing the cells to
CCCP (Figure 3.4 and 3.5), fluorescence decreased to 61% and 12% of control cells
for R123 and TMRE, respectively. Comparing to R123 and TMRE fluorescence
emission intensities at 30 seconds after pulsing, decreases in the fluorescence
emissions after 3 minutes revealed that the nsPEFs-induced mitochondrial
membrane potential drop is a progressive event and lasts for several minutes after
pulsing.
Figure 3.3: A schematic diagram shows how Rhodamine 123 (R123) works.
R123 is a lipophilic cationic fluorescent dye that is accumulated within active
mitochondria because of large negative potential across the mitochondrial
membrane. The decrease of mitochondria potential will decrease R123
concentration in mitochondria and cause the drop of its fluorescent intensity.
TMRE works in a similar way.
46
Figure 3.4: Dose-dependent R123 fluorescence intensity before and after pulsing
(mean ± SEM). CCCP was used as positive control. Significant differences from
zero-pulse controls are designated by asterisks (*P < 0.05). #Statistically
different from previous column of same incubation time (P < 0.05). There are at
least 60 cells for each data point and each condition was repeated at least three
times.
47
Figure 3.5: Dose-dependent TMRE fluorescence intensity before and after
pulsing (mean ± SEM). CCCP was used as positive control. Significant
differences from zero-pulse controls are designated by asterisks (*P < 0.05).
#Statistically different from previous column of same incubation time (P <
0.05). There are at least 124 cells for each data point and each condition was
repeated at least three times.
Mitochondrial Membrane Permeabilization After nsPEF Exposure
In addition to conventional mitochondrial membrane potential detectors
(R123 and TMRE), we used cobalt-quenched calcein method for probing
mitochondrial membrane permeability change. The mechanism of this method is
described in Figure 3.6. We first labeled mitochondria with cobalt-quenched calcein
according to the protocol described in Materials and Methods (Chapter 3.3.4), and
then re-suspended these cells in the fresh RPMI 1640 medium without or with
cobalt ions. Three minutes after treating calcein-stained, cobalt-quenched cells with
nsPEFs in the medium without cobalt ions in the external medium (Figure 3.7A), we
48
observed calcein fluorescence emission decreased in cells exposed to 5, 10, and 20
pulses as an evidence for mitochondrial membrane permeabilization. However, we
did not observe fluorescence decrease after exposure to 30 pulses or more under the
same buffer conditions. These results suggest that plasma membrane
permeabilization at these higher pulse numbers allows cobalt ions to leak out of the
cells, dequenching the calcein fluorescence; hence, the fluorescence emission of
calcein in cells was high.
On the other hand, when calcein-stained, cobalt-quenched cells were treated
in medium with cobalt ions (Figure 3.7B), the calcein fluorescence decreased
significantly after the application of 5 pulses and decreased further at higher pulse
numbers (30–100). The extent of mitochondrial permeabilization depends on the
number of pulses applied since the fluorescence is lower with higher pulse numbers;
it decreased to 73% and 51% of the fluorescence before pulsing with 30 and 100
pulses, respectively. These results suggest that cobalt ions in the external medium
allow sufficient calcein quenching even after permeabilization of the plasma
membrane since the Co
2+
concentration gradient favors an inward flux of cobalt ions
rather than a decrease in cytoplasmic Co
2+
.
49
Figure 3.6: Non-fluorescent calcein-AM and Co
2+
enter the cell. AM groups are
cleaved from calcein via non-specific esterase activity in the cytosol and
mitochondria. Co
2+
quenches the cytosolic calcein signal. Co
2+
cannot readily
enter healthy mitochondria. Co
2+
influx after pore opening quenches the
mitochondrial calcein fluorescence.
50
Figure 3.7: Dose-dependent calcein fluorescence without (A) or with (B) Co
2+
in
the external medium. Data show the relative fluorescence intensity before and 3
min after pulsing (mean ± SEM). Significant differences from zero-pulse
controls are designated by asterisks (*P < 0.05). #Statistically different from
previous column (P < 0.05). There are at least 38 cells for each data point and
each condition was repeated at least three times.
51
Plasma Membrane Permeabilization After nsPEF Exposure
In addition, plasma membrane permeabilization was monitored with YO-
PRO-1 and PI uptake. YOPRO-1 influx is significantly increased at five or more
pulses (Figure 3.8). Fluorescence intensity increased 8, 12, and 19 times higher than
in untreated control cells after applying 20, 30, and 100 pulses, respectively.
Although PI is a less sensitive detector of plasma membrane permeabilization than
YO-PRO-1 [58], a measurable influx of PI was detected after applying 20 pulses or
more (Figure 3.9). PI fluorescence intensity increased three and four times higher
than in untreated control cells after applying 20 and 30 pulses, respectively.
Figure 3.8: Dose-dependent YO-PRO-1uptake.Fluorescence images were
recorded 3 min after pulse exposure. Data show YO-PRO-1 fluorescence 3 min
after pulsing (mean ± SEM) in arbitrary units. Significant differences from zero-
pulse controls are designated by asterisks (*P < 0.05). There are at least 36 cells
for each data point and each condition was repeated at least three times.
52
Figure 3.9: Dose-dependent propidium uptake. Fluorescence images were
recorded 3 min after pulse exposure. Data show PI fluorescence 3 min after
pulsing (mean ± SEM) in arbitrary units. There are at least 32 cells for each data
point and each condition was repeated at least three times.
Taking the results from three different indicators of mitochondrial membrane
integrity in living cells together, we have shown that high-voltage, nanosecond
electric pulses (4 ns, 10 MV/m, 1 kHz repetition) cause an increase in mitochondrial
membrane permeability and an associated decrease in mitochondrial membrane
potential. Moreover, the data of YO-PRO-1 and PI shows that with similar pulse
parameters, exposure to fewer pulses (<20) permeabilize the mitochondrial
membrane while leaving the plasma membrane relatively impermeable to propidium
yet permeable to small but significant amounts of YO-PRO-1 (after five pulses,
fluorescence reached 1.8 times the value of control cells). Higher pulse numbers
!50,000.0&
0.0&
50,000.0&
100,000.0&
150,000.0&
200,000.0&
250,000.0&
300,000.0&
0&p& 1&p& 5&p& 10&p& 20&p& 30&p&
Rela/ve&Fluorescence&Intensity&&
Number&of&Pulses&
53
(≥20), however, produce significant plasma and mitochondrial membrane
permeabilization. These relative permeabilization sensitivities are consistent with
our observations of intracellular cobalt-quenched calcein fluorescence with and
without cobalt in the external medium. However, we cannot conclude with certainty
from the results reported here whether the nsPEF-induced decrease in the
mitochondrial membrane potential results from the direct electroporation of the
inner mitochondrial membrane or the formation of mitochondrial membrane
permeability transition pores, or from some other permeabilizing mechanism. Since
nsPEF also causes plasma membrane permeabilization (as observed by cobalt-
quenched calcein and YO-PRO-1 and PI influx), other molecules and ions including
calcium ions could enter the cell and trigger the mitochondrial permeability
transition [130] as a secondary effect of plasma membrane permeabilization.
3.6 Conclusion
In conclusion, using three different methods with fluorescence indicators—
rhodamine 123, TMRE, and cobalt-quenched calcein — we show that nsPEFs (4 ns
duration, 10 MV/m, 1 kHz repetition rate) cause a loss of the mitochondrial
membrane potential. The most likely explanation for this observation is nsPEFs-
induced permeabilization of the inner mitochondrial membrane, although it is
possible that nsPEF exposure triggers the mitochondrial permeability transition
54
through some mechanism still to be identified. At the same time, we also detected
plasma membrane permeabilization, indicated by YO-PRO-1 and propidium iodide
influx and the responses of cobalt-quenched calcein labeled cells.
55
Chapter 4: Cell Swelling and Membrane
Permeabilization after
Nanosecond Pulsed Electric
Field
4.1 Abstract
In this chapter, cell plasma membrane electropermeabilization is
characterized by exposing Human Jurkat T lymphoblasts to 5 ns pulsed electric
fields (including single pulse exposures), and by monitoring the resulting
osmotically driven cell swelling as a function of pulse number and pulse repetition
rate. In addition, molecular dynamic (MD) simulations are established to exhibit
how water molecules permeate a lipid bilayer under fixed electric field. We find that
even a single 10 MV/m pulse of 5 ns duration produces measurable swelling of
Jurkat T lymphoblasts in RPMI 1640 growth medium, and the degree of swelling
depends on the pulse number and repetition rate. From the measured and simulated
values, we estimate from the swelling kinetics the ion and water flux that follows
the electropermeabilization of the membrane.
56
4.2 Introduction
High-intensity, nanosecond pulsed electric fields (nsPEFs) induced cell
plasma membrane electropermeabilization is believed to results from formation of
long-lived hydrophilic pores in lipid bilayers through a mechanism that is strongly
dependent on the applied field intensity and on the pulse duration [20, 131].
Although the exact nature of such pores remains poorly understood,
electropermeabilization is proved to facilitate the delivery of various substances,
such as drugs, oligonucleotides, antibodies, genes and plasmids, as well as for
cancer therapy, genetic engineering and wound healing [71, 132-135].
Membrane permeabilization is traditionally detected by measuring the
uptake of fluorescent dyes like propidium iodide or YO-PRO-1 that cannot pass
through an intact membrane, and this method has been widely used in studies of
permeabilization by nanosecond pulses [58, 59, 122]. However, the quantification of
membrane permeabilization based on influx of fluorescent dyes is limited by the
number and size of the pores created. Below certain dose, the fluorescence emission
from the small amount of dye that enters the cell is difficult to distinguish from
background with affordable and flexible imaging and other detection systems.
Recently, measurement of cell volume changes arising from colloidal
osmotic imbalance has been introduced as a more sensitive method for
characterizing nsPEFs-induced plasma membrane permeabilization [62, 65, 136-
57
138]. When the cell membrane is permeabilized, small intracellular and extracellular
solutes can pass freely through the membrane, causing a loss of the concentration
gradients maintained by active living cells and an equilibration of the intracellular
and extracellular concentrations of these species. Larger intracellular solutes cannot
pass through the permeabilized membrane, creating an osmotic imbalance. In the
general case the osmotic pressure resulted from the trapped intracellular colloidal
solutes leads to water influx from the extracellular medium and an increase in cell
volume (cell swelling). The advantage of cell swelling analysis is that it is not
dependent on the dye selection and on the sensitivity of fluorescence detection
methods. Thus, cell swelling can be used for the study of plasma membrane
electropermeabilization even in response to mild stresses (single, nanosecond-
duration pulses), and for following the time dynamic of the phenomenon [64].
In this chapter, we exposed Jurkat T lymphoblasts to 5 ns, 10 MV/m pulsed
electric fields. Instead of relying on the fluorescent dye uptake, we tracked cell
volume change following plasma membrane permeabilization by using standard
white light microscopic imaging [61]. Systematical analysis of the osmotically
driven cell swelling show that nsPEF-induced cell volume increase is a function of
pulse number (from 1-50) and pulse repetition rate (1 Hz or 1 kHz). Cell swelling
caused by a single pulse exposure observed in this chapter helps to eliminate the
cascading cellular responses to a train of pulses and reduce the complexity of data
interpretation. From the rate and magnitude of the cellular response, we characterize
58
the dynamics of electropermeabilization. Moreover, we show how water transport
through intact lipid bilayers by building molecular dynamic (MD) simulations of
lipid bilayers with no membrane proteins. By correlating the experimental results
with the simulated results, we improve the accuracy of electropermeabilization
mechanism and applicability of current model. Finally, we examine the participation
of membrane protein channels in nsPEF-induced cell swelling by ion channel
blocker (Gd
3+
and La
3+
) and aquaporin channel blocker (Hg
2+
).
4.3 Materials and Methods
4.3.1 Cell Line and Cell Culture
Jurkat T lymphoblasts (ATCC TIB-152, Manassas, VA) were grown in
RPMI 1640 (Mediatech,Manassas, VA) containing 10% heat-inactivated fetal
bovine serum (Gibco, Carlsbad, CA), 2 mM L-glutamine (Gibco), 50 U/mL
penicillin (Gibco), and 50 µg/mL streptomycin (Gibco). Cells were maintained in
exponential growth at 37 °C in a humidified, 5 % CO
2
atmosphere.
4.3.2 Cell Preparation
For pulse treatment, cells were concentrated to 2 × 10
7
cells/mL and in some
cases incubated for 15 min with 0.5 µM calcein-AM (acetoxymethyl ester,Molecular
Probes, Eugene, OR), which enhances visualization of the cell outline using
59
fluorescence microscopy. After loading of the dye, cells were centrifuged and
resuspended in fresh RPMI 1640 (or in some cases 150 mM NaCl) at 2 × 10
7
cells/mL. For exposure in presence of lanthanide or mercuric ions, GdCl
3
or LaCl
3
(Aldrich Chem. Co, Milwaukee, WI), 100 µM or 1 mM, or HgCl
2
, 50 µM, was
added to the cell suspension 5 min before pulse treatment.
4.3.3 Pulsed Electric Field Exposures
Setup of pulse exposure for life-time microscopic observation is same as
description in Chapter 3.3.2. A resonant-charged, solid-state Marx bank-driven,
hybrid-core compression, diode-opening switch pulse generator designed and
assembled at the University of Southern California [123] delivered 5 ns, 10 MV/m
electrical pulses at a 1 kHz or 1 Hz repetition rate to the microchamber electrodes
mounted on the microscope stage in ambient atmosphere at room temperature.
4.3.4 DIC and Fluorescence Microscopy
Observations of live cells during and after pulse exposure were made with a
Zeiss (Göttingen, Germany) Axiovert 200 epifluorescence microscope with 63×
water immersion objective and Hamamatsu (Higashi-ku, Hamamatsu City, Japan)
ImageEM EM-CCD camera. Captured images were analyzed with Hamamatsu
SimplePCI and ImageJ (http://imagej.nih.gov/ij/) software. The cell perimeter was
tracked using a freehand selection function, and then the area defined by the drawn
perimeter was measured. To reduce variability, cells in the center of the exposure
60
chamber, not adjacent to the electrode surfaces, were selected in the images captured
immediately before pulsing, and then the same cells were analyzed in the post-pulse
images. For each pulsing condition, at least 3 experiments were carried out, and a
total of at least 30 cells were analyzed.
4.3.5 Molecular Dynamics Simulations
Simulations were performed using the GROMACS set of programs version
4.0.5 [139] on the University of Southern California High Performance Computing
and Communications Linux cluster (http://www.usc.edu/hpcc/). Lipid topologies
derived from OPLS united-atom parameters [89] were obtained from Peter Tieleman
(http://moose.bio.ucalgary.ca). The Simple Point Charge (SPC) water model [140]
was used, and all simulations were coupled to a temperature bath at 310 K with a
relaxation time of 0.1 ps and a pressure bath at 1 bar with a relaxation time of 1 ps,
each using a weak coupling algorithm [91]. Pressure was coupled semi-isotropically
(using a compressibility of 4.5 × 10
−5
bar
−1
) normal to and in the plane of the
membrane (NpT). Bond lengths were constrained using the LINCS algorithm [92]
for lipids and SETTLE [93] for water. Short-range electrostatic and Lennard–Jones
interactions were cut off at 1.0 nm. Long-range electrostatics was calculated with
the PME algorithm [94] using fast Fourier transforms and conductive boundary
conditions. Reciprocal-space interactions were evaluated on a 0.12 nm grid with
fourth order B-spline interpolation. The parameter ewald_rtol, which controls the
relative error for the Ewald sum in the direct and reciprocal space, was set to 10
−5
.
61
Periodic boundary conditions were employed to mitigate system size effects.
Diffusion of water was calculated from mean square displacements over 50 ps using
the ‘g_msd’ tool in GROMACS.
All phospholipid systems contain 128 1-palmitoyl-2-oleoyl-sn-glycero-3-
phosphatidylcholine (POPC) lipids and 8960 water molecules (70 waters/lipid),
which results in a system box size of approximately 7 nm × 7 nm × 10 nm. To
ensure that replicated simulations are independent, each atom is assigned a
randomized velocity from a Maxwell distribution at the beginning of a simulation (a
built-in function of GROMACS). POPC systems are equilibrated before
electroporation by waiting until they reach a constant area per lipid — 0.66 nm
2
.
Molecular graphic images were generated with Visual Molecular Dynamics
(VMD) [97].
4.4 Electric Pulse-Induced Cell Swelling
In order to qualitatively study the effect of nsPEF on cell swelling, we
exposed Jurkat T lymphoblasts to 5 ns, 10 MV/m electric pulses in
physiological medium (RPMI 1640) with different pulse numbers and repetition
rates. Cell response was detected by DIC channel of microscope and recorded
on a fast camera at different time intervals before and after pulse exposure. As
shown in Figure 4.1, Jurkat cells exhibit an increase in cross-sectional area 60 s
62
after being exposed to 30 5 ns, 10 MV/m electric pulses at 1 kHz repetition rate.
Cell cross-sectional area was measured by freehand selection function of Image
J (Figure 4.1C and 4.1D)
Figure 4.1: Osmotic swelling after nanoelectropulse exposure. (A) Jurkat cells
before pulse exposure. (B) Same cells 60 s after exposure to 30, 5 ns, 10 MV/m
pulses at 1 kHz. (C), (D). Cells in (A) and (B) are outlined for area extraction
with ImageJ. Note pulse-induced swelling, blebbing, and intracellular
granulation and vesicle expansion, a result of the osmotic imbalance caused by
electropermeabilization of the cell membrane.
Cells do not respond uniformly, as shown in Figure 4.1 and Figure 4.2. The
responses of individual cells vary not only in the magnitude of the volume increase,
but also in the visual appearance of the cells during the first minute after pulsing.
63
Some cells expand more or less uniformly, maintaining their roughly spherical
shape. Others initially form one or more blebs of various sizes, which are subsumed
as the cell continues to swell and recover its spheroidal geometry. Figure 4.2 shows
a series of images that exhibit the morphological changes in Jurkat cells over 7
minutes post pulse exposure.
Figure 4.2: Representative images of morphological changes in Jurkat cells
before (0 s) and after (10 s, 20 s, 30 s, 40 s, 60 s, 120 s, 180 s, 300 s and 420 s)
exposure to 30, 5 ns, 10 MV/m pulses at 1 kHz. Even at 420 s, the longest
monitoring time permitted by our apparatus, cells were not observed to recover
their initial volume.
64
Dose-dependent cell area increase of Jurkat cells after exposure to 0, 10, 20,
30 and 50 5 ns, 10 MV/m electric pulses at 1 kHz repetition rate in RPMI 1640 is
shown in Figure 4.3. The rate of swelling is greatest immediately following pulse
exposure, declining exponentially over tens of seconds, but not falling to zero after 3
min. The ratio of swelling is proportional to the number of pulses applied. Moreover,
cells were not observed to recover their initial volume, under the conditions in these
experiments in this chapter, even after 7 min, the longest monitoring time permitted
by our apparatus (Figure 4.2).
Figure 4.3: Dose-dependent area increase of Jurkat cells after exposure to 0, 10,
20, 30 and 50 5 ns, 10 MV/m electric pulses delivered at 1 kHz. Note initial
rapid swelling, which declines over about 1 min to a slower, but non-zero rate.
Results are presented as mean ± standard error for at least 30 cells, from at least
3 independent experiments, for each pulsing condition.
65
An increase in cell cross-sectional area was detected after a single 5 ns, 10
MV/m pulse, as shown in Figure 4.4. Furthermore, almost 10 % cross-sectional area
increase was observed 3 minutes after pulsing. Comparing to pervious data of
nsPEF-induced YO-PRO-1 influx with similar pulsing conditions (Figure 1.2), a
single pulse results shown here suggests that cell measurement of swelling could be
a more sensitive way for detection of cell plasma membrane permeability.
Figure 4.4: Cell swelling can be reliably detected after even a single 5 ns, 10
MV/m pulse. Results are presented as mean ± standard error for at least 30 cells,
from at least 3 independent experiments, for each pulsing condition.
66
This is the first systematic study of cell swelling following exposure to
pulsed electric fields shorter than 10 ns. However, the observed phenomenon seems
qualitatively similar to that reported for much longer pulses in previous studies with
cells exposed to 60- and 600 ns electric pulses [65, 138]. It is likely that the primary
mechanisms are the same for short and long pulses. The phenomenon of pulse-
induced osmotic imbalance and the resulting cell swelling are consistent with the
mechanistic hypothesis that nanosecond pulses open a large number of small pores
on the plasma membrane, with a pore creation rate larger than the pore expansion
one. Inorganic ions and small molecules, but not larger solutes, pass through these
defects [141].
From the presented results, and building on conclusions supported by
previous studies [62, 136-138, 142, 143], we propose the following hypothetical
sequence of events in Jurkat cells responding to 5 ns, 10 MV/m electric pulses
delivered at 1 kHz. All of the observations we describe below are consistent with
this scheme.
1. Pulse exposure results in the immediate formation of a dose-dependent number
of lipid nanopores of a dose-dependent total area, large enough to allow passage
of small inorganic ions [138]. These pores are formed during the 5 ns duration of
a pulse.
2. Equilibration of intracellular and extracellular [Na
+
], [K
+
], and [Cl
−
] by
diffusive ion transport through the permeabilized membrane (presumably
67
through the lipid nanopores but perhaps also through other structures) proceeds
for several minutes [143-145].
3. During [Na
+
], [K
+
], and [Cl
−
] equilibration, water flux (bidirectional under
balanced conditions) becomes net positive inward, driven by the osmotic
gradient caused by the reduction of the mobile ion concentration gradients and
the remaining intracellular colloidal anions and other large molecules. This
osmotically driven water influx is the primary cause of pulse-induced cell
swelling [64].
4. When the [Na
+
], [K
+
], and [Cl
−
] concentration gradients across the membrane
reach zero (or some minimum value), water influx continues at a continually
decreasing rate, as the internal osmolytes are diluted, until the ion-permeant
structures (pores) close, allowing re-establishment of small ion concentration
gradients, or until mechanical constraints become significant, or until the cell
bursts.
4.5 Effects of Pulse Repetition Rate on nsPEF-
Induced Cell Swelling
Repetition rate is an important parameter that impacts the efficacy of nsPEFs
on cell plasma membrane. Previous study by Vernier et al. reported that the influx of
the normally impermeant fluorescent dye YO-PRO-1 into Jurkat cells after nsPEF
68
exposure (30 4 ns, 8 MV pulses delivered at 10, 100, 1k and 10 KHz) is dependent
on the pulse repetition rate [58]. This result in fact corresponds to our observation of
nsPEF-induced YO-PRO-1 influx into Jurkat cells after exposure to 30 4 ns, 10 MV
electric pulses, as shown in Figure 4.5. For a given number of pulses, the amount of
YO-PRO-1 that enters the cell is considerably greater for higher pulse repetition rate
than it is for lower pulse repetition rates.
Figure 4.5: Frequency-dependent YO-PRO-1 influx after exposure of Jurkat
cells to 30 4 ns, 10 MV/m electric pulses delivered at different repetition rates.
Fluorescence microscopic images of Jurkat cells in growth medium containing
YO-PRO-1 (5.0 µM) were captured immediately before and 3 min after
exposure. Fluorescent intensity of YO-PRO-1 was integrated from the captured
images. Results are presented as mean ± standard error for at least 30 cells, from
at least 3 independent experiments, for each pulsing condition.
69
In the experiments of swelling, we found a similar result that the amount of
nsPEF-induced volume increase is dependent on the pulse repetition rate, as shown
in Figure 4.6. The results of YO-PRO-1 and cell swelling demonstrate that
nanosecond pulsed electric field induced cell membrane permeabilization is directly
proportional to the repetition rate. However, both of them are in contrast to results
reported for longer but still sub-microsecond pulses [146]. These differences in
repetition rate dependencies might suggest that longer pulses (≥60 ns) produce a
qualitatively different restructuring of the membrane than is produced by 5 ns pulses.
Figure 4.6: Rate and extent of pulse-induced swelling is greater at higher pulse
repetition rates. Results are presented as mean ± standard error for at least 30
cells, from at least 3 independent experiments for each pulsing condition.
70
Considering the 5 ns pulses used in the work reported here, we can
hypothesize that high-repetition-rate (1 kHz) pulses either expand the area of the
pores formed by the first pulse or they create new pores or both. If high-repetition-
rate pulses only expand the pores formed by the first pulse, then the expansion must
produce more pore area per pulse than multiple pulses widely separated in time. To
distinguish among these possibilities we will need to learn more about the pore
density and the population distribution of the pore area.
One parameter that might explain the influence of pulse repetition rate on
nsPEF-induced membrane permeability change is the resealing process of
membrane pores. We assume that repair process of some membrane pores created
by nsPEF take longer than 1 ms. When a second pulse is delivered in 1 ms or less
after the first pulse, the permeabilizing damage has not yet been repaired, or has
been only partially repaired, and so the damage is compounded. In other words,
more damages (either larger pores or more number of pores) will be created on
plasma membrane by higher repetition rate. In contrast, when pulses are delivered at
lower repetition rate and more than 1 ms passes between pulses, there is sufficient
time for this repair process to complete, and the damage is merely additive.
However, the actual mechanism is still unclear. To resolve these possibilities,
further observations and more accurate modeling are needed.
71
4.6 A Scaling Law for nsPEF-Induced Cell Swelling
Electrical impact dose is a value to describe and quantify the pulsed field
stimulated bioelectric effects. Schoenbach et al. first proposed this idea and he
generalized an empirical equation to draw it [147]:
𝑆= 𝑆(𝐸𝜏𝑁
!/!
) (1)
S is electrical impact dose which describes the intensity of the observed
bioelectric effect. E is the field amplitude in V/m, τ is the pulse duration in s, and N
is the number of pulses. The equation is so called scaling law and holds for
secondary bioelectric effects, which are caused by permeability change in the
plasma membrane or subcellular membrane. We present our swelling data by using
equation (1), as shown in Figure 4.7a. At least for the first 60 s, the relative cell area
scales with the square of the pulse count, probably a reflection of the stochastic
character of the electropore formation [16, 90]. To approach a better fitting, equation
(2) was modified from equation (1) by adding two fitting parameters, x and k:
𝐴
!
(𝑡)= 𝐸𝜏𝑁
!
+𝑘 (2)
A
R
(t) is the pulse-number-dependent cross-sectional area of the cells,
evaluated at each sampled time point. Figure 4.7b shows the new data fitting with
equation (2) at 0, 10, 60 and 180 after pulsing. The behavior of fitting parameters x
72
and k over the considered time interval is shown in Figure 4.7c. Different from the
definition of equation (1), the parameter x which determines the contribution of
pulse number N to the scaling law is not exactly 0.5 but slowly increases from 0.3 to
0.6. Besides, the parameter k that does not present in equation (1) can be interpreted
as the relative cell area when the pulse number is N=0. However, the number of k is
not exactly 1 in our case. The deviation of k might come from the uncertainty of our
measurement of cell area.
The mismatch between our data and equation (1) may be explained through
the definition of the model built by Schoenbach et al. This model bases on the
hypothesis of “random rotation” of cells. The validity of random rotation requires
that recovery time of permeabilized cell membrane is longer than the time between
pulses. It is also requires that the rotation of cells, with respect to the electric field
direction, is completely random between pulses. If there is only minor cell rotation
between pulses, the scaling law will approach a liner dependence on N.
Consequently, for high repletion rate pulsing events where the position of cells will
not change much between pulses, the observed effects will scale with N
x
, where x is
expected to exceeding 0.5 and might even reach values larger than 1, in case of
nonlinear membrane effects. In our pulsing condition with 1 kHz, the motion of
cells is likely not respected the random rotation concept, but our observation
displays non-liner dependence on N.
73
(b) (c)
Figure 4.7: Electric pulse-induced area increase dose-dependency. (a) The
phenomenon is described for times up to about 60 s after exposure by the
electrical impact dose factor, the product of the electric field, the pulse duration,
and the square root of the number of pulses. The impact dose variable for these
experiments is the pulse number N, which has the following values: 0, 1, 3, 5,
10, 20, 30, and 50. All pulses were 5 ns, 10 MV/m, delivered at 1 kHz. Straight
gray lines are a linear fit to the swelling data at 10 s and 60 s; (b) fitting of the
cell swelling data with Eq. (2); (c) fitting parameters over time.
(a)
74
4.7 nsPEF-Induced Ion Flux
Recently, both MD simulation and experimental studies found that nsPEF
increase membrane permeability by open long-lasting (minutes) nanometer-size
electropores in the cell plasma membran [56, 58, 65, 138, 148]. For healthy cells
with intact plasma membrane, membrane barrier function permits the establishment
of the Na
+
and K
+
concentration gradients that result primarily from the operation of
Na
+
/K
+
-ATPase pump and K
+
leak channels, and which are an essential component
in the maintenance of physiological osmotic balance. On the contrary, cells exposed
to nsPEF form a number of nanopores in plasms membrane and these nanopores
provide alternative pathways for Na
+
and K
+
transport that are not under the control
of cells. Without regulation, the nsPEF-induced ion flux will lead to an osmotic
imbalance which the cells can not quickly restore, and finally result in colloidal
osmotic swelling.
Nonetheless, we can not rule out the possible participation of membrane
protein channels in nsPEF-induced swelling from our data. To clarify the role of
these ion channels in nsPEF-induced swelling process, we set two observations with
different types of channel blocker, lanthanide ions (Gd
3+
and La
3+
) and Hg
2+
.
Lanthanides are know to block multiple types of ion channels, including
voltage-gated calcium channels, stretch-activated channels, several ionotropic and
metabotropic ligand-gated channels; they can also inhibit Ca
2+
- and Mg
2+
-ATPases
75
and Na
+
/Ca
2+
exchanger [149-156]. The mechanism of the action of lanthanide ions
is not completely understood, but it appears to be related to the similarity of their
cationic radii to the size of Ca
2+
ions [149]. We exposed Jukart cells to 0-50 5 ns, 10
MV/m electric pulses at 1 kHz in RPMI1640 growth medium containing 0.1 mM or
1 mM Gd
3+
/La
3+
. No significant inhibition of nanoelectropulses-indeuced swelling
by Gd
3+
and La
3+
was observed (Table 4.1). However, it is important to note that
lanthanide ions promote the precipitation of lanthanide phosphates in cell culture
media depending on time and concentration. Even though we carried out the
experiment with lanthanide ions in growth medium and did not observe precipitation
during the image capture time, we set another experiment with cells in NaCl as a
check on the possibility of lanthanide chelation and complex formation by
phosphates and other constituents of the complex growth medium. The result does
not show significant difference to data with lanthanide ions (Table 4.1) which, not
exactly, but may partially exclude the effects from formation of soluble complexes.
Aquaporins are a large family of specific protein channels that facilitate the
passive permeation of water across biological membranes in response to osmotic
pressure [157, 158]. To see the possibility of voltage-regulated aquaporins and the
effects of opening of aquaporin channels on nsPEF-induced swelling response, we
carried out pulse exposures in the presence of mercury ions (Hg
2+
), a channel
blocker that inhibits most aquaporins by interacting with cysteine residues [159,
76
160]. The result is similar to those obtained with lanthanide ions. Again, we did not
observe significant inhibition of nsPEF-induced swelling by Hg
2+
(Table 4.1).
77
0 s 10 s 60s
0 p 10 p 30 p 50 p 0 p 10 p 30 p 50 p 0 p 10 p 30 p 50 p
RPMI
RPMI
alone
1.0 1.0 1.0 1.0 1.0
1.07±
0.002
1.11±
0.003
1.15±
0.006
1.0
1.21±
0.009
1.30±
0.003
1.39±
0.007
Gd
3+
0.1 mM
1.0 1.0 1.0 1.0 1.0
1.01±
0.006
1.09±
0.002
1.18±
0.003
1.0
1.13±
0.002
1.32±
0.004
1.34±
0.008
Gd
3+
1 mM
1.0 1.0 1.0 1.0 1.0
1.04±
0.01
1.12±
0.006
1.11±
0.004
1.0
1.15±
0.01
1.32±
0.012
1.35±
0.011
La
3+
0.1 mM
1.0 1.0 1.0 1.0 1.0
0.99±
0.008
1.08±
0.039
1.10±
0.021
1.0
1.05±
0.017
1.24±
0.038
1.33±
0.028
La
3+
1 mM
1.0 1.0 1.0 1.0 1.0
0.98±
0.015
1.08±
0.018
1.08±
0.021
1.0
1.04±
0.013
1.36±
0.039
1.33±
0.036
Hg
2+
50 µM
1.0 1.0 1.0 1.0 1.0
1.03±
0.012
1.09±
0.017
1.11±
0.011
1.0
1.16±
0.015
1.37±
0.035
1.35±
0.017
NaCl
NaCl
alone
1.0 1.0 1.0 1.0 1.0
1.02±
0.001
1.10±
0.001
1.13±
0.010
1.0
1.12±
0.007
1.34±
0.005
1.33±
0.001
Gd
3+
0.1 mM
1.0 1.0 1.0 1.0 1.0
1.04±
0.003
1.04±
0.008
1.09±
0.007
1.0
1.14±
0.008
1.30±
0.006
1.32±
0.006
La
3+
0.1 mM
1.0 1.0 1.0 1.0 1.0
1.00±
0.006
1.04±
0.008
1.08±
0.009
1.0
1.10±
0.013
1.25±
0.016
1.34±
0.020
Table 4.1: Pulse-induced Jurkat cell swelling in presence of lanthanides or mercury
ions. Relative cell area increase over time in Jurkat cells exposed to 0, 10, 30 or 50
pulses in RPMI/150 mM NaCl alone or in presence of GdCl
3
or LaCl
3
(0.1 and 1
mM) or HgCl
2
(50 µM). Similar results were. Data are presented as mean ± standard
error over three independent experiments (at least 30 cells for each condition).
120 s 180 s
0 p 10 p 30 p 50 p 0 p 10 p 30 p 50 p
RPMI
RPMI
alone
1.0
1.27±
0.007
1.38±
0.005
1.47±
0.019
1.0
1.31±
0.012
1.50±
0.014
1.52±
0.008
Gd
3+
0.1 mM
1.0
1.09±
0.007
1.42±
0.005
1.42±
0.019
1.0
1.16±
0.007
1.47±
0.024
1.54±
0.010
Gd
3+
1 mM
1.0
1.23±
0.01
1.37±
0.014
1.43±
0.019
1.0
1.25±
0.008
1.44±
0.009
1.51±
0.018
La
3+
0.1 mM
1.0
1.11±
0.025
1.36±
0.044
1.0
1.15±
0.028
1.43±
0.054
La
3+
1 mM
1.0
1.08±
0.014
1.44±
0.031
1.43±
0.049
1.0
1.12±
0.016
1.46±
0.051
1.51±
0.048
Hg
2+
50 µM
1.0
1.25±
0.016
1.55±
0.048
1.52±
0.030
1.0
1.31±
0.022
1.68±
0.066
1.59±
0.035
NaCl
NaCl
alone
1.0
1.20±
0.010
1.44±
0.006
1.48±
0.011
1.0
1.26±
0.012
1.50±
0.020
1.57±
0.017
Gd
3+
0.1 mM
1.0
1.20±
0.014
1.42±
0.002
1.53±
0.016
1.0
1.27±
0.023
1.51±
0.005
1.53±
0.008
La
3+
0.1 mM
1.0
1.16±
0.015
1.38±
0.023
1.46±
0.028
1.0
1.26±
0.019
1.47±
0.031
1.53±
0.032
78
Similar to the repetition rate dependent swelling response, the result we
observed with lanthanide ions is again in contrast to the results reposted for longer
pulses [65]. By comparison, our pulses are 5 ns, 10 MV/m; the pulses in both
Pakhomov’s work of repetition rate dependencies for the magnitude [146] and
Andre’s work of Gd
3+
sensitivity of the swelling response [65] are 60 ns and 600 ns,
1.2 MV/m. These differences in the results suggest that longer pulses (≥60 ns) might
produce a qualitatively different restructuring of the membrane than is produced by
5 ns pulses.
4.8 MD Simulation of Water Permeation of Intact
Lipid Bilayer
Molecular simulations reveal how water permeates a lipid bilayer without
pore formation [161, 162]. About once per nanosecond in simulations of 50 nm
2
POPC bilayers at 310 K [163], an isolated water molecule crosses from one leaflet
of an intact bilayer to the other (Figure 4.8). No pore is involved, and application of
a porating electric field has little effect on either the frequency or the direction of
these random, diffusive crossing events. In simple bilayer systems the primary
determinant of the rate of crossing from one side of the membrane to the other
(expressed experimentally as the permeability coefficient) is the area per lipid [164].
Systems with a smaller area per lipid have smaller permeability coefficients [165].
79
Although detailed calculations would be premature at the present level of accuracy
and understanding of the correspondence between MD simulations of lipid bilayers
and living cell membranes, we can translate the random and bidirectional 1 H
2
O per
nanosecond per 50 nm
2
that we observe in our POPC simulations to an osmotically
driven (initially 100 mOsm) net transport of about 10
4
H
2
O per nanosecond per cell,
declining exponentially as the ion concentration imbalance is reduced.
80
Figure 4.8: Random permeation of an intact POPC bilayer. A single H
2
O
molecule crosses the phospholipid bilayer interface. Applied electric field is 340
MV/m downward in the diagram. (A) 0 ps, (B) 100 ps, (C) 200 ps, (D) 300 ps.
Only acyl oxygens and water molecules in the interface are shown. The crossing
water molecule is enlarged and colored yellow. Light blue spheres are acyl
oxygens; red and white atoms are water; the empty space in the middle of
figures between two water phases represents the hydrocarbon region of the lipid
bilayer. From a system containing 128 POPC, 4480 H
2
O, at 310 K, 1 bar.
81
4.9 Conclusion
In this work, we have demonstrated that osmotically driven cell swelling is a
sensitive method for studying plasma membrane electropermeabilization induced by
very short and very intense pulsed electric fields. Cell swelling has been observed
even after a single 5 ns, 10 MV/m electric pulse exposure. Moreover we have
characterized the swelling response as a function of pulse number and repetition rate,
demonstrating that it increases with the number of pulses delivered, is frequency-
dependent (increasing when pulses are delivered at higher repetition rate). The
participation of membrane protein channels has been checked by ion channel
blockers, lanthanide ions, and aquaporin channel inhibitor, Hg
2+
. In our
experimental conditions, it is not sensitive to the presence of either the lanthanide
ions (Gd
3+
and La
3+
) or mercuric ions, (Hg
2+
) in the external medium. Our results
are consistent with the idea that the primary permeabilizing event leading to cell
swelling is the formation of lipid pores in the plasma membrane. The swelling
responses can be described with a empirical scaling equation. The scaling equation
summarizes the effects of pulsing parameters (pulse duration, electric field
amplitude and pulse number) on nsPEF-induced primary cell responses. The
approach allows people to predict the events triggered by nsPEF, although further
investigations are needed to complete the scaling law and clarify the details of the
mechanisms underlying nsPEF-induced responses.
82
Chapter 5: Moveable Wire Electrode
Microchamber for Nanosecond
Pulsed Electric-Field Delivery
5.1 Abstract
Constructing a delivery system for megavolt-per meter nanoelectropulses is
a bioelectrical engineering challenge. In this chapter, we report fabrication of a
parallel tungsten wire electrode assembly with sub-millimeter spacing, integrated
with nanosecond pulse generators. Coupling of the delivery system and a fluorescent
microscope enables real-time biophotonic investigations of the electroperturbation
of living cells. We report also electrical characterization of the system through
experimental measurements and numerical simulations to evaluate the actual field
delivered to the cells. The frequency and time domain analyses demonstrate the
utility of the proposed assembly for delivering pulses as short as 2.5 ns.
5.2 Introduction
In recent years, the biological effects from nanosecond, high-voltage electric
pulses with electric field strengths of megavolts per meter have been widely
reported. These cellular and subcellular responses of nanosecond pulsed electric
83
fields (nsPEFs) show important applications in cancer therapy, genetic engineering,
and cell biology [31, 44, 108]. The effectiveness of nsPEFs depends on the electric
field actually applied to the cell. To achieve nsPEFs with MV/m field strength,
appropriate exposure systems, i.e., pulse generators, delivery systems and
measurement devices must be used [123, 166, 167]. However, constructing an
exposure system for successfully delivering nsPEFs to cells is still a challenge.
For conventional electroporation applications of pulse duration in
microsecond range, the development of pulse generators and electrodes-based
delivery systems have benefitted from the very low output impedance (close to 0 Ω).
Those generators enable one to fix electrical parameters of microsecond pulses
regardless of the impedance of biological samples [168-170]. On the contrary, for
the applications of pulse duration in nanosecond range (nsPEFs) with a given output
impedance, special care must be taken to design devices compatible in terms of
frequency bandwidth and impedance matching (typically 50Ω). An impedance
mismatch between the generator and the biological load will impact on pulse shape
and magnitude and, thus, on biological results. Moreover, applications of pulses in
nanosecond range need much higher field intensity, and thus delivery systems with
reduced distances- in the micrometer range - are preferred.
To increase the efficacy of nsPEFs experiments, we developed a new
delivery system based on tungsten wire electrodes (TWE) for nsPEFs delivery.
Combined with nanosecond pulse generator and automated micromanipulator, the
84
TWE based nsPEFs delivery system provides real-time and in-situ observation for
in-vitro cell experiments involving in nsPEFs exposure [171]. In this chapter, we
describe the design and assembly of the new tungsten wire electrodes. Simulation
results using custom finite-difference time-domain (FDTD)-based code are
presented to evaluate the actual electric field delivered by tungsten wire electrodes.
We also report a detailed characterization of the electromagnetic properties of the
delivery system in terms of frequency bandwidth. Finally, we present the results of
in-vitro cell experiments to validate the delivery system.
85
5.3 Setup of nsPEF Exposure System with TWE
5.3.1 System Setup
Figure 5.1 shows the setup of nsPEF exposure system with TWE for in-vitro
cell experiments. This system integrates nanosecond pulse generator, TWE-base
delivery system, microscope, and oscilloscope as a pulse signal monitor. The
delivery system can deliver nanosecond electric pulse without distorting the pulse
shape losing the pulse amplitude, and allows real-time and in-situ observations.
Figure 5.1: General experimental delivery system set-up composed of the nsPEF
generator, the measurement devices, the TWE delivery system, the
micromanipulator and the microscope stage.
nsPEF
Oscilloscope
100-to-1
Attenuator
Tungsten wire
electrodes
50
Motorized
command unit
3-axis
micromanipulator
Lens
Coverglass
chamber
tt
86
5.3.1.1 Pulse Generators
Two different nanosecond pulse generators were used for nsPEF exposure in
our system. One is a resonant-charged, solid-state Marx bank-driven, hybrid-core
compression, diode-opening switch pulse generator (nsPEF1) designed and
assembled at the University of Southern California (USC). It delivered 5 ns, l kV
electric pulses to TWE [123]. The other pulse generator (nsPEF2) using a laser-
triggered photoconductive semiconductor switches (PCSSs) and on a microchip
laser developed by Horus–Laser HT (Limoges, France) [167, 172] delivered 2.5 ns,
1 kV electric pulses to TWE. Both pulse generators used in this study present 50 Ω
output impedance.
5.3.1.2 Pulse Signal Measurement
The delivered electric pulses were monitored on a digital storage
oscilloscope (DSO, 725ZI, LeCroy, NY). A power measurement device (Figure 5.2)
with a 100:1 attenuator designed and assembled at USC was inserted in the setup
allowing voltage measurement on the DSO [173]. Three transmission lines (RG 58
C/U, Pasternack Enterprises, Irvine, CA) served to connect: the generator to the
power measurement device, the power measurement device to the TWE and to the
DSO.
The measurement devices have BNC connectors (Pasternack Enterprises,
Irvine, CA).
87
Figure 5.2: The power measurement device with 100:1 attenuator was designed
and assembled in USC. Three transmission lines connected to the pulse
generator, oscilloscope and TWE separately.
5.3.1.3 Tungsten Wire Electrode (TWE)
In order to perform real-time and in-situ observation, the TWE was designed
to be able to work on microscope stage and tiny enough to locate targeted cells. The
idea was shown in Figure 5.3. 100 µm in diameter, gold plated tungsten wires
(45086, Alfa Aesar, MA) were selected to fabricating the electrodes because
tungsten is rigid and not easy to be deformed and the gold plated surface ensure
biocompatibility with the cells. In order to achieve parallel alignment between two
wires, 3 mm long, 100 µm wide and 50 µm deep micro-grooves separated 100 µm
Pulse&
generator&
Oscilloscope&
TWE&
Power&
Measurement&
device&with&100:1&
a=enuator&&
88
from each other were fabricated on a polydimethylsiloxane (PDMS) as a mold to
support the alignment (Figure 5.4). Two wires were placed in two adjacent grooves
individually and then fixed both of the grooves ends by epoxy adhesive (5 min
epoxy, Pacer Technology, CA) so that wires can maintain in parallel position once
removed from the PDMS support. After removing the wires from the PDMS, the
wires were carefully bent at a right angle by using flat-tip forceps to facilitate the
placement of the electrodes in a coverglass chamber. The vertical part of the wires
was covered with silicone rubber to give stability to the structure (see Figure 5.5).
Compared to the previously designed systems [74, 146, 174, 175], the parallel
electrode configuration in our design permits the inclusion of many more cells
within the region of uniform field between the electrodes. The wires were soldered
to a BNC connector (Amphenol rfx, Danbury, CT) with a 50-Ω resistor (RR5025,
Vishay, PA). The resistor, placed in parallel to the TWE, allows impedance
matching with the transmission lines and the generator. While working with cells,
the TWE was inserted in an 8-well coverglass chamber (Lab-Tek II, Nalge Nunc
International, Penfield, New York) with RPMI 1640 biological culture medium (200
µL) containing the cells.
89
Figure 5.3: Sketch shows the design of TWE. The parallel TWE connected to
nsPEF generator is able to work on a microscope stage. A 3-axis
micromanipulator can control the movement of TWE precisely.
90
Figure 5.4: Pictures show the fabrication process of PDMS mold. To making the
PDMS mold, a thick layer of SU8 was coated on a silicon wafer followed by
post soft baking. The baked wafer was covered by the mask designed for wire
alignment, and exposed to UV light. After UV exposure, a hard bake was
applied. Finally, a thick layer of PDMS was spread on the wafer to finish the
fabrication.
91
Figure 5.5: Tungsten wire electrodes delivery system. A) Overview of the TWE
components, materials and dimensions. B) Zoom on the TWE containing the
cells under nsPEF exposure. All dimensions are in mm.
Tungsten wire
electrodes
Front view
3.15% 4.68% 5.81%
15%
3.00%
Top view
Side view
50 ohms
resistor
Gold-plated
Tungsten wires
Silicon
Epoxy
adhesive
Top view
Front view
3.00$
0.10$
0.10$ 0.10$ 0.10$
Tungsten wires
electrodes
RPMI 1640 solution containing cells
Coverglass
well
!
A
B
92
5.3.1.4 Micromanipulator and Microscope
A three-axis, motorized micromanipulator (MP 225/M-20139, Sutter
Instruments, CA, USA) was attached to the transmission line leading to the TWE to
enable precise positioning. The micromanipulator permits controlled, programmable
access to the cells in the coverglass chambers on the microscope stage, quickly
moving from one set of cells to another within a chamber, or from one chamber to
the next. A Zeiss Axiovert 200 epifluorescence microscope and a Hamamatsu
ImageEM CCD camera were used for cell imaging before, during, and after pulse
exposure.
5.3.2 Measurement Protocol
Coverglass chambers loaded with cells were placed on the microscope stage
in ambient atmosphere at room temperature allowing real-time microscopic
observation under nsPEF exposure. The cells were exposed to 2.5 and 5.0 ns pulses,
with varying pulse counts (0, 3, 5, 10) and 1 kHz repetition rates.
5.3.2.1 Electromagnetic Measurements
A vector network analyzer (VNA, HP 8720ET, Palo Alto, CA) with a 20-
GHz bandwidth allowed the measurement of the TWE delivery system reflection
coefficient S
11
. The measurements were carried out at the input of the BNC
connector. The relative permittivity ε
r
and the electrical conductivity σ of the
93
biological medium (RPMI 1640) were measured with a dielectric probe (85070E
Dielectric probe kit, Agilent, Santa Clara, CA).
5.3.2.2 Cell Culture
Human Jurkat T lymphoblasts (ATCC TIB- 152) were grown inRPMI 1640
containing 10% heat-inactivated fetal bovine serum, 2 mM L-glutamine, 50-
units/mL penicillin, and 50 µg/mL streptomycin. Cells were cultured at 37 °C in a
humidified, 5% carbon dioxide atmosphere and concentrated to 1 × 10
6
cells/mL for
nsPEF treatment.
5.3.2.3 Cell Imaging and Image Analysis
nsPEF-induced cell swelling was introduced as a result of cell plasma
membrane permeabilization. In this chapter, cell swelling was used to validate the
TWE delivery system. In order to enhance visualization of the cell outline, cells
were incubated with 0.5 µM calcein-AM (acetoxymethyl ester) for 15 min. After
incubation, cells were centrifuged and resuspended in fresh RPMI 1640 for pulse
exposure. Differential interference contrast (DIC) and fluorescence images were
obtained with a 63X, water-immersion objective. The images were taken as a time
series, starting immediately before the pulse exposure, and at 10, 20, 30, 40, 60, 120,
and 180, 240, 300 s after the exposure. Cell swelling was assessed by comparing the
sizes of cells in DIC-fluorescence merged images. Image processing was carried out
with Image J 1.43u (Wayne Rasband, National Institute of Health, Bethesda, MD;
94
http://rsb.info.nih.gov/ij). The cross-sectional area of individual cells was measured,
and the swelling was characterized by comparing area changes before and at
different time intervals after pulse exposure.
5.3.2.4 Electromagnetic Numerical Modeling
The electromagnetic characterization of the TWE delivery system was
accomplished also through numerical modeling using custom finite-difference time-
domain (FDTD)-based code. The FDTD method is a 3-D full-wave numerical
method which allows the rigorous analysis of complex and inhomogeneous
structures. For the structure to be simulated with the FDTD method, a geometrical
model with spatial meshing is required. The reflection coefficient S
11
, the input
impedance, and the electric field between the electrodes can be extracted from the
FDTD simulations and correlated with the measurements. The TWE delivery system
was modeled with the geometry presented in Figure 5.5, i.e., with a BNC connector,
a 50-Ω resistor, the vertical tungsten wires covered with silicone rubber, the TWE,
and the epoxy adhesive delimiting the TWE channel. In addition, the coverglass
chamber containing the biological medium (RPMI 1640) was also simulated. The
BNC inner and outer metallic parts and the gold-plated tungsten wires were
considered as perfect conductors. Since the dimensions of the primary area of
interest for our study, i.e., the channel formed by the TWE where the cells are
exposed to nsPEF, are small compared to rest of the structure, the TWE delivery
95
system was meshed with a non-uniform mesh. The grids imposed were 50×50×50
µm
3
for the TWE channel and 200 × 200 × 200 µm
3
otherwise. All the dielectric
materials have been taken into account in the simulations through their
electromagnetic macroscopic properties, i.e., relative permittivity and electrical
conductivity. The relative permittivities are summarized in Table 5.1. The electrical
conductivity of the biological medium is 1.5 S/m. The electromagnetic feed is
placed at the input of the BNC connector as a 50 Ω localized source, and classical
perfectly matched layers boundary conditions envelop the computational volume
during the simulations [176].
Table 5.1: The relative permittivities of the dielectric materials accounted in
simulations.
96
5.4 Electromagnetic Analysis of TWE Delivery
System
The electromagnetic analysis of the versatile TWE-based delivery system
carried out with numerical modeling was confronted to experimental measurement
results. Impedance matching of pulse generators and delivery systems is important
because it determines the energy transferred to the biological cells. To address this
point the following studies were conducted: TWE frequency characterization,
electric field distribution in the TWE channel, and time domain measurements of the
applied pulses.
5.4.1 Frequency Characterization
To determine the TWE frequency performance, its impedance and reflection
coefficient were studied (Figure 5.6). For both parameters, the simulated results
were consistent with the experimental ones. For frequencies up to about 750 MHz, a
particularly good agreement can be observed. The discrepancy above can be
explained by the differences between the modeled structure and the manufactured
one. As illustrated by Figure 5.6A, the |S
11
| < -10 dB for frequencies lower than
about 300 MHz. The latter value can be seen as an upper boundary defining the
frequency bandwidth of the TWE delivery system. The phenomenon observed after
1 GHz can be related to the TWE delivery system geometrical resonance (i.e., the
97
structure could be seen as a matched antenna).
Figure 5.6B shows the TWE impedance as a function of frequency. The
effective impedance of the structure becomes complicated at high frequencies, but it
can most simply be thought of as a 50 Ω surface mount resistor in parallel with the
effective impedance of the electrode structure. The rigid part of the structure that is
held in place by silicone has an impedance that depends on the geometry as well as
the permittivity of the silicone and epoxy adhesive. Similarly, the TWE impedance
depends on its geometry and the biological medium’s electrical parameters, such as
conductivity and permittivity. All of these elements contribute to the complex
evolution of the equivalent impedance as a function of the frequency as illustrated
Figure 5.6B. Using approximate analytical formulas [167, 177], an impedance of a
few hundred ohms can be calculated for the TWE. Thus, a 50 Ω resistor is placed in
parallel with the tungsten wires to ensure matching with the generator impedance.
For frequencies lower than about 300 MHz, the imaginary part has a low value. The
real part of the impedance does not drift too much from 50 Ω and it can be assumed
constant (to approximately 40 Ω). As the frequency increases, the real part of the
impedance progressively shifts to higher values thus increasing the impedance
mismatch with the generator.
The impedance of the entire electrode assembly with biological medium
between the electrodes is 40 Ω. The impedance of the tungsten wires only, with
biological medium between them, is about 250 Ω. When manufacturing the
98
electrode structure, the tungsten wires are placed as precisely as possible in a PDMS
support to obtain reproducible alignment. Manufacturing tolerances of ±150 µm on
the microchamber length and ±10 µm on the microchamber width induce about
±10% variation on the impedance of the electrode structure and less than ±2%
variation on the global TWE impedance, at low frequencies.
99
Figure 5.6: TWE delivery system frequency characterization obtained by FDTD
simulations and measurements with vector network analyzer. (A) Reflection
coefficient (S
11
). (B) Real and imaginary parts of the impedance.
0 0.5 1 1.5 2
−20
−15
−10
−5
0
Frequency (GHz)
|S
11
| (dB)
FDTD simulation
Measured with the NVA
0 0.5 1 1.5
−50
0
50
100
150
200
Frequency (GHz)
Impedance (Ω)
R(Ze) Simulation
I(Ze) Simulation
R(Ze) Measured
I(Ze) Measured
A
B
100
5.4.2 Time-Domain Characterization
Figure 5.7A shows the measured applied pulses obtained by adding the
incident and the reflected pulses using the method described in [167, 178]. For
nsPEF1, the pulse characteristics are 5.0 ns full width at half of the maximum value
(FWHM), 1.2 kV maximum amplitude and 6.0 ns rise time. For nsPEF2, the pulse
characteristics are 2.5 ns FWHM duration, 1.1 kV maximum amplitude, and 1.7 ns
rise time. The rise times of the pulses delivered by the generators are 5.0 and 1.5 ns
for nsPEF1 and nsPEF2, respectively. The slight distortions on the rise times are due
to the high-frequency mismatch between the generator and the delivery system.
The power spectra corresponding to the applied pulses are presented in
Figure 5.7B. The spectra are normalized with respect to the maximum value of each
pulse. The frequency band edge at −10 dB can be defined as a reasonable limit of
power spectra containing the main pulse energy. This band edge is around 75 and
220 MHz for nsPEF1 and nsPEF2, respectively. The -10 dB band edge of the
applied pulses power spectra is of course related to the frequency bandwidth of the
TWE delivery system. Indeed, the shorter the pulses, the broader their frequency
distribution, and the stronger the impedance mismatch with the delivery system.
Since the TWE impedance bandwidth is about 300 MHz, this proposed assembly is
adapted for delivering pulses as short as 2.5 ns.
101
Figure 5.7: Time domain measurements with both generators. A) Applied
Pulses. B) Normalized Power spectra in dB.
10 15 20 25 30
0
0.2
0.4
0.6
0.8
1
Time (ns)
Voltage (kV)
nsPEF1
nsPEF2
1 10 100 1000
−30
−25
−20
−15
−10
−5
0
Frequency (MHz)
Normalized |TF(v
app
(t)|, (dB)
nsPEF1
nsPEF2
A
B
102
5.4.3 Electric Field Distribution
Figure 5.8 shows the distribution of the electric field between the electrodes
as determined by numerical simulations. The voltage applied between the electrodes
was 1.2 kV. We have used a 3-D-FDTD with 50 µm voxels for the tungsten
electrode modeling. This allowed obtaining the impedance and also the electric
field, but with low resolution compared to the size of the electrodes (only four
voxels between the electrodes). This 50 µm resolution does not allow a good
meshing for the space between the electrodes. To determine the electric field with a
resolution comparable to the dimensions of biological cells, a quasi-static approach
was taken, with a resolution of 1 µm, which produces a more realistic and smooth
electric field distribution. The center-to-center distance between the electrodes is
meshed using 200 voxels. To determine the influence of the biological medium and
the coverglass chamber, three configurations were studied: the TWE placed in the
coverglass chamber without biological medium, the TWE placed in the coverglass
chamber with biological medium (actual experimental conditions) and for the last
configuration the coverglass chamber was removed.
Figure 5.8A and 5.8B indicate that the electric field is less homogenous
when the TWE channel is filled with biological medium. However, a good
homogeneity is obtained in a central region around the half-distance mark between
the two electrodes. The homogeneous region near the surface of coverglass chamber
is around 80 µm in width. This region should then be preferred when conducting
103
biological experiments with this delivery system. The electric field value in the
bottom of that central region is about 10 MV/m (Figure 5.8B). This field
corresponds to the actual electric field applied on the cells and this value is
consistent with the applied voltage of ∼1.2 kV per 100 µm gap between circular
electrodes. Figure 8B and Figure 8C indicate that electric field at the position of the
cells is more homogenous without the coverglass chamber. However, its value has
decreased to about 8 MV/m. Consequently, the coverglass chamber increases the
electric-field value, particularly for cells placed in the central region.
104
Figure 5.8: Electric field distributions in the TWE channel. A) TWE without
biological medium placed in the coverglass well. B) Same as previous but with
biological medium. C) Same as previous but without the coverglass chamber.
(A)
(B)
(C)
105
5.5 Biological Experiments With 2.5 and 5.0 ns, 10
MV/m nsPEFs
To demonstrate the biological effectiveness of this exposure system, we
conducted experiments with living cells. As shown in Figure 5.9, changes in cell
volume that occur as a result of membrane permeabilization induced osmotic
imbalance are used for estimating the effects of nsPEF on the cell membrane. Jurkat
T lymphoblasts were exposed to 0, 5, and 10, 5.0 ns and 2.5 ns, 10 MV/m electric
pulses with 1 kHz repetition rate. Under these experimental conditions, the cells
swelled and were not observed to recover after 5 min. The results confirmed that
nsPEFs were successfully delivered to cells by the TEW delivery system.
106
Figure 5.9: Dose-dependent area increase of Jurkat cells after exposure to 0, 5
and 10 electric pulses delivered with 10 MV/m field strength and a frequency
repetition of 1 kHz. Results are presented as mean ± standard error for at least
70 cells, from at least 3 independent experiments, for each pulsing condition.
(A) 5 ns duration pulses delivered by nsPEF1. (B) 2.5 ns duration pulses
delivered by nsPEF2.
A
B
107
5.6 Conclusion
In conclusion, we designed and assembled the tungsten wire electrodes
based exposure system for nsPEF delivery. This system enables real-time and in-situ
cell observations. To ensure the matching with the generator output impedance, a 50
Ω resistor was placed in parallel with the TWE ensures matching with the generator
output impedance. The electromagnetic behavior of the TWE delivery system was
studied. The matching bandwidth for a reflection coefficient ≤ -10 dB of the TWE
delivery system containing a biological medium is around 300 MHz. We also
performed time-domain measurements of two pulse generators, nsPEF1 and
nsPEF2, with different technologies. Pulse durations of them were 5 ns and 2.5 ns
respectively, and the maximum pulse amplitudes were around 1.2 kV. The electric
field distribution simulations were provided by FDTD. The results suggest that the
actual electric field delivered to cells was 10 MV/m with the maximum 1.2 kV
applied voltage. In-vitro cell experiments studying nsPEF-induced cell membrane
permeabilization were taken by using this delivery system. Dose-dependent area
increase of Jurkat cell was observed after exposure to pulses as short as 2.5 ns.
Moreover, this broadband TWE delivery system could also be used for classical cell
electroporation with µs and ms pulses.
108
Chapter 6: Determination of
Nanoelectropulse-induced Pore
Size by Blocking Osmotic
Swelling
6.1 Abstract
Pulsed power technology with high intensity nanosecond pulsed electric
fields (nsPEFs) are used to permeablilize cell plasma membranes by creating stable
pores in biotechnology and biomedical applications. Although the mechanism of
nsPEFs-induced membrane poration has been reported in many studies, the
dependence of pore size and distribution on nsPEF pulse duration, strength and
pulse counts are still not clear. In this paper, we characterized the size of nsPEFs-
induced membrane pores by isoosmotically replacing the pulsing buffers with
polyethylene glycols and sugars before exposing Jurkat T lymphoblasts to 5 ns, 10
MV/m electric pulses with different pulse number. To this end, the pore size was
evaluated by analyzing cell volume changes that result from the permeation of
substituted extracellular molecules through the plasma membrane of Jurkat T
lymphoblasts. We find that most pores opened by 5-100 5 ns pulses have diameter
between 0.7-0.9 nm. Either the diameter of pores or the number of pores, or the
combination of both phenomena increased with increasing the pulse counts. The
109
prevention of cell swelling by PEG 1000 after 2000 5 ns pulses suggests that 5 ns
pulses can only opened membrane pores with diameter smaller than 1.9 nm even
after such extreme exposure.
6.2 Introduction
High-intensity, nanosecond pulsed electric fields (nsPEFs) have been
extensively reported to be versatile nondestructive tools capable of producing a
variety of specific cellular responses such as membrane permeabilization [58],
intracellular calcium release [49], cellular apoptosis for tumor shrinking [31, 48, 73,
74], temporary blockage of action potential propagation in nerves [179],
catecholamine secretion [180], activation of platelets, and release of growth factors
for accelerated wound healing [181]. In particular, cell membrane permeabilization
caused by nsPEFs is believed to begin with the formation of hydrophilic pores in
lipid bilayers through a mechanism that is strongly dependent on the applied field
intensity and on the pulse duration [131, 182]. Until the nature of the pore structure
and the details of the mechanisms that govern pore size and lifetime are better
understood, the practical utilization of nsPEF-induced electropermeabilization in
biomedical and biotechnological applications, with the potential for greater energy
efficiency and less invasive procedures, will be ad hoc and empirical.
Membrane permeabilization is traditionally detected by measuring the
110
uptake of fluorescent dyes like propidium iodide, YO-PRO-1 and thallium ions that
cannot pass through an intact membrane, and this method has been used in studies
of permeabilization by nanosecond pulses [58, 59, 122]. The results of those studies
are consistent with models that predict membrane electropore diameters of 1 to 1.5
nm [59, 183, 184]. Quantification of membrane permeabilization based on influx of
fluorescent dyes, however, is limited by dye emission and excitation efficiency and
detector sensitivity. Recently, measurement of cell volume changes arising from
colloidal osmotic imbalance has been introduced as a more sensitive method for
characterizing nsPEFs-induced plasma membrane permeabilization [62, 65, 136-
138]. Chapter 4 describes the mechanism of the cell swelling based on colloidal
osmotic imbalance in detail.
Multiple studies have shown that cell swelling can be suppressed or reversed
(cell shrinking) if the extracellular medium contains appropriate concentrations of
large solutes which cannot pass through the permeabilized membrane, thus
counterbalancing the intracellular colloid osmotic pressure. Pore size can be
estimated based on this method. by isosmotically replacing larger extracellular
solutes, such as polyethylene glycols (PEG) and neutral sugars of various sizes [62,
185-187]. Nesin et al. has reported that, for nsPEF of 60 ns and 300 ns durations, the
pore estimation method can be used to determine the size of nanopores created on
the cell plasma membrane [64].
In the present work we employed PEG and two sugars of different sizes to
111
estimate the diameter of pores created in Jurkat T lymphoblast cell membranes by
nsPEF exposure. The pulse duration in this study is 5 ns, considerably shorter than
the pulses investigated in other studies. In addition we examined the dependence of
pore size on pulse parameters, especially the number of pulses.
6.3 Materials and Methods
6.3.1 Cell Line and Cell Culture.
Jurkat T lymphoblasts (ATCC TIB-152, Manassas, VA) were grown in
RPMI 1640 (Mediatech, Manassas, VA) containing 10% heat-inactivated fetal
bovine serum (Gibco, Carlsbad, CA), 2 mM L-glutamine (Gibco), 50 units/mL
penicillin (Gibco), and 50 µg/mL streptomycin (Gibco). Cells were maintained in
exponential growth at 37 °C in a humidified, 5% CO
2
atmosphere.
6.3.2 Cell Preparation
For pulse treatment, cells were concentrated to 1 x 10
6
cells/mL and
incubated for 15 minutes with 0.5 µM calcein-AM (acetoxymethyl ester, Molecular
Probes, Eugene, OR), which enhances visualization of the cell outline using
fluorescence microscopy. After loading of the dye, cells were centrifuged and
resuspended in different bath buffers at 1 x 10
6
cells/mL as needed.
112
6.3.3 Chemicals and Buffers
A reference buffer for measuring nanosecond pulsed electric field effect on
cell cross-sectional area was RPMI 1640 (Mediatech, Manassas, VA) containing
10% heat-inactivated fetal bovine serum (Gibco, Carlsbad, CA), 2 mM L-glutamine
(Gibco), 50 units/mL penicillin (Gibco), and 50 µg/mL streptomycin (Gibco).
The osmolality of pure RPMI 1640 was ~ 280 mOsm. In experiments with
different sugars, sugars were first dissolved in distilled water and mixed with RPMI
1640. 116 mM of RPMI 1640 was isoosmotically replaced with inositol (VWR).
116 mM, 150 mM and 200 mM of RPMI 1640 was isoosmotically replaced with
sucrose (EM Science). In experiments with different concentration of PEGs, buffers
were formulated similarly. Because the osmotic property of large PEGs depends on
their molecular size and concentration, the isosmotic PEG 1000 (Polysciences, Inc.)
concentrations have to be found by trial and error [185, 186]. As a result, 50
mOsm/kg, 116 mOsm/kg, 150 mOsm/kg and 200 mOsm/kg PEG1000
isoosmotically replaced RPMI 1640. The osmolality (mOsm/kg, denoted here after
as mOsm) of each solution was measured by vapor pressure osmometer (Wescor
Inc., Utah). All solutions were between 280 and 300 mOsm.
The molecular structures of tested sugars were drawn and determined using
the software package VMD (Visual Molecular Dynamics). Van der Waal radii were
presented in the plotted geometries in Figure 61. The minimum diameters of sugars
were measured from their structure planes with the smallest cross sections. The
113
hydrodynamic radius of a PEG random coil depends on its molecular weight and the
viscosity of solution through the Einstein viscosity relation [188, 189]. The
hydrodynamic diameter of PEG 1000 presented in this study was 1.9 nm that
employed the value estimated from viscosity measurement of Ringer solution in
previous work [185].
Figure 6.1: The molecular structures of tested sugars were drawn by VMD
software. The minimum cross-sectional areas were shown and the smallest
diameters measured from the cross-sectional areas were listed in the table.
6.3.4 Nanosecond Pulsed Electric Field Exposures
Tungsten wire electrodes system was designed for nanosecond pulse electric
field exposure. The detail of set up was described in Chapter 5 [171]. A resonant-
charged, solid-state Marx bank-driven, hybrid-core compression, diode-opening
114
switch pulse generator designed and assembled at the University of Southern
California [123] delivered 5 ns, 10 MV/m electrical pulses at 1 kHz repetition rate to
the electrodes in ambient atmosphere at room temperature.
Before nsPEF exposure, 8-well coverglass chamber was placed on
microscope stage at proper position, and 200 µL of 1x10
6
cells/mL was injected in
to the chamber followed by quickly and precisely inserting tungsten wire electrodes
into the same chamber with the help of micromanipulator (MP-225, Sutter). Cells
were allowed to settle and adhere to the chamber floor for 10-15 minutes before
exposed to nsPEF.
6.3.5 DIC/ Fluorescence Microscopy and Data Analysis
Observations of live cells were made with a Zeiss (Göttingen, Germany)
Axiovert 200 epifluorescence microscope with 63X water immersion objective and
ImageEM EM-CCD camera (Hamamatsu, Japan). Images of cells were taken right
before and at various time intervals of 10 s up to 5 minutes after nsPEF exposure.
Captured images were analyzed with Hamamatsu SimplePCI and ImageJ
(http://imagej.nih.gov/ij/) software. To reduce variability, cells in the center of the
exposure chamber, not adjacent to the electrode surfaces, were selected in the
images captured before pulsing, and the identical cells were selected in the post-
pulse images, too. The cell perimeter was tracked using a freehand selection
function, and then the cell cross-section area defined by the drawn perimeter was
measured. Cell cross-section area (A) of different time intervals was normalized to
115
the area before pulsing (A
0
) as: A
normalied
=A/A
0
, and plot against time for data
presentation. For each pulsing condition, at least 3 experiments (n) were carried out,
and a total of at least 30 cells were analyzed. Pooled data were given as mean ± SE
of experiments. If the SE value was smaller than the size of symbol, the error bar
may not be seen in the graph.
6.4 Cell Volume Changes Induced by nsPEFs
Cell swelling after nsPEF treatment has been shown to indicate plasma
membrane permeabilization for pulses lasting 5 ns or longer [61, 64, 65]. Figure 6.2
shows a representative series of images of a single Jurkat cell before and at different
times after exposure to 10, 5 ns, 10 MV/m pulses delivered at 1 kHz in RPMI 1640
medium. The cross-sectional areas of this cell extracted from serial images are listed
in Table 6.1.
116
Figure 6.2: Time series of DIC images showing a Jurkat cell swelling after
nsPEF exposure (10 5 ns, 10 MV/m at 1 kHz) in RPMI 1640. Scale bar is 5 µm.
Table 6.1: Cross-sectional area of a representative Jurkat cell after nsPEF
exposure (10 pulses, 5 ns, 10 MV/m at 1 kHz). Values were extracted from the
images in Figure 6.2.
Time
[s]
0 10 20 30 40 60 120 180 240 300
Area
[µm
2
]
72.4 74.3 78.4 78.6 84.1 85.6 94.4 97.8 108 108
117
The extent of swelling is a function of the number of pulses delivered. The
change of cross-sectional area (from which the volume change can be readily
extracted) after nsPEF treatment increases with increasing pulse count (Figure 6.3).
Five minutes after exposure to 30 pulses, the area has increased by a factor of 1.6
(two-fold volume increase, assuming spherical geometry). Previously we have
shown influx of normally impermeant fluorescent dye after similar pulse doses
[190]. These images and data are consistent then with the interpretation that cell
swelling after 5 ns pulses is a dose-dependent indicator of pulse-induced plasma
membrane permeabilization.
Figure 6.3: Dose-dependent area increase of Jurkat cells after nsPEF exposure to
0, 5, 10, 20 and 30 5 ns electric pulses delivered with 10 MV/m field strength
and a repetition rate of 1 kHz in RPMI 1640. Swelling ratio depends on the
applied pulse intensity. With increasing pulse counts there was an increase in
the volume changes. The results are presented as mean±SE for at least 60 cells
from at least three independent experiments for each pulsing condition at 25 °C.
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
1.4$
1.5$
1.6$
1.7$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela1ve$Area$Changes$$
Time$(second)$
RPMI$1640$
118
6.5 Impermeant Solute Inhibits Cell Swelling
(PEG1000)
To verify that the colloidal osmotic swelling mechanism is operating in
cells permeabilized with 5 ns pulses, we compared cell volume changes in cells
suspended in isosmotic solutions containing the large molecule PEG 1000 at
several concentrations. Figure 6.4 shows the swelling response of Jurkat cells
exposed to 30 pulses, 10 MV/m at 1 kHz in isosmotic RPMI1640 solutions
containing 50–200 mOsm PEG 1000. Swelling induced by 5 ns pulse exposure
is inhibited by 116 mOsm PEG 1000. At higher concentrations of PEG 1000 the
cells shrink after pulse treatment, consistent with observations reported for
significantly longer pulses [64]. In sham exposures, cell volume did not change
during the five-minute observation period, demonstrating that osmoregulatory
mechanisms are not affected by PEG 1000 in the medium over this time period.
These results suggest that PEG 1000 does not penetrate into cells
exposed to the 5 ns pulse doses used in these experiments and that the
concentration of the large intracellular solutes that contribute to colloidal
osmotic swelling is less than 150 mOsm. In subsequent experiments we used
116 mOsm for the external solute concentration, which seems to provide an
approximately osmotically balanced condition (no swelling or shrinking with
PEG 1000) in suspensions of 5 ns pulse-permeabilized Jurkat cells.
119
Figure 6.4: Isoosmotic substitution of PRMI 1640 with PEG 1000 with different
concentration. Jurkat cells were exposed to 30 5 ns pulses delivered with 10
MV/m field strength and 1 kHz repetition rate. Increasing of PEG 1000
concentration inhibited the nsPEF-induced cell swelling. The results are
presented as mean±SE for at least 60 cells from at least three independent
experiments for each pulsing condition at 25 °C.
120
6.6 Cell Swelling Reduced by Inositol, Blocked by
Sucrose
To compare the effects of solutes smaller than PEG 1000 on cell volume
changes triggered by 5 ns, 10 MV/m pulses, we treated cells in RPMI containing
either inositol or sucrose, isosmotically added to a final concentration of 116
mM (Figure 6.5). In contrast to the significant volume increases observed in
standard RPMI, cells in medium containing inositol showed reduced swelling,
and sucrose blocked swelling completely.
Assuming that colloidal osmotic swelling is the mechanism underlying
the observed volume increase, we can extract from these results an approximate
size for the permeabilizing structures produced by 5 ns electric pulse exposure.
For simplicity we call these structures pores, although we emphasize that their
precise nature has not been definitively established. When a molecule (inositol)
is small enough to pass through the pores, the swelling will not be stopped,
because these molecules cannot offset the osmolality of the large impermeant
molecules inside the cell. On the other hand, if the molecule (sucrose) is too
large to pass through the pores and the concentration is high enough, swelling
will not occur because there is an osmotic balance. Although we do not have
direct evidence for the size of membrane electropores, we can estimate an upper
limit on pore size from the diameter of the molecules.
121
The minimum molecular cross sectional diameters of inositol and
sucrose are 0.7 nm and 0.9 nm, respectively. As shown in Figure 6.5, Pores
created by 5 pulse exposures must have an effective diameter less than 0.7 nm,
since neither inositol nor sucrose blocks swelling at the doses used in this study.
For 10–30 pulses, cell swelling was blocked by sucrose but not by inositol
(Figure 6.5), indicating that a pulse dose in this range produces pores with an
effective diameter greater than 0.7 nm but less than 0.9 nm.
Figure 6.5: : Protective action of inositol and sucrose against nsPEF-induced cell
swelling. Jurkat cells were exposed to 0, 5, 10, 20 and 30 5 ns pulses delivered
with 10 MV/m field strength and 1 kHz repetition rate in solutions
isoosmotically replaced 116 mOsm RPMI1640 with inositol or sucrose. The
extent of swelling in solution containing inositol and the inhibition of swelling
in solution with sucrose indicated that inositol can pass through the 5 ns electric
pulses created pores but sucrose can not. The results are presented as mean±SE
for at least 60 cells from at least three independent experiments for each pulsing
condition at 25 °C.
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
1.4$
1.5$
1.6$
1.7$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela1ve$Area$Changes$$
Time$(second)$
Inositol$
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
1.4$
1.5$
1.6$
1.7$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela1ve$Area$Changes$$
Time$(second)$
Sucrose$
122
6.7 Pulse Dose and Pore Diameter
The results summarized in Figure 6.5 show that with isosmotic inositol
in the medium the swelling rate increases with the number of pulses delivered.
This could indicate either that the number of inositol-permeable pores increases
with the number of pulses, or that the mean diameter of a population of initially
inositol-impermeant pores is increased by the application of multiple pulses.
However, sucrose remains impermeant under these conditions.
To investigate whether increasing the number of pulses delivered
increases the diameter of the permeabilizing structures (pores), we applied
higher pulse dose and increased the sucrose concentration to 150 mOsm and 200
mOsm, so that if sucrose molecules cannot pass through the pores, the cells will
shrink, as we observe with PEG 1000 (Figure 6.4).
As seen in Figure 6.6, cells in the suspensions with higher sucrose
concentrations shrink after 30 or even 100 5 ns pulses, with no volume recovery.
These results imply that sucrose does not penetrate into cells exposed to a large
number of 5 ns pulses, or that the influx is minimal and thus does not produce a
detectable volume change. The result suggests that that multiple pulses do not
increase the diameter of the permeabilizing structures beyond the 0.9 nm
produced at lower pulse counts. ). If pores larger than 0.9 nm are produced, we
can conclude from these data that they are a small fraction of the pore
123
population, since their contribution to the volume change cannot be detected in
our measurements.
Figure 6.6: Cell shrinking after exposed to 30 and 100 5 ns pulses delivered with
10 MV/m field strength and 1 kHz repetition rate in solutions issosmotically
replaced RPMI 1640 with higher concentration of sucrose. The continuous
shrinking in the observation time course demonstrated that even 100 5 ns
electric pulses created membrane pores smaller than sucrose molecules. The
results are presented as mean±SE for at least 60 cells from at least three
independent experiments for each pulsing condition at 25 °C.
To explore further the possibility that pore diameter is increased by
multiple pulses, we applied a much larger dose — 2000 pulses — to cells in
isosmotic solutions containing 50 and 116 mOsm PEG 1000.
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
1.4$
1.5$
1.6$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela0ve$Area$Changes$$
Time$(second)$
200$mOsm$Sucrose$$
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
1.4$
1.5$
1.6$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela0ve$Area$Changes$$
Time$(second)$
150$mOsm$Sucrose$$
124
As expected, since the PEG 1000 molecule is about twice the size of
sucrose (the hydrodynamic diameter of PEG 1000 is about 1.9 nm [23]), cell
shrinking is observed after 100 5 ns pulses at PEG 1000 concentrations above
116 mOsm (150 and 200 mOsm; Figure 6.7), as it was for 30 pulses (Figure 6.4).
However, at a lower PEG 1000 concentration, 50 mOsm, where we
expect to see swelling, since the extracellular concentration of PEG 1000 is less
than the intracellular concentration of large, impermeant solutes, we observe for
a 100-pulse exposure instead of simple swelling an initial increase in cell
volume followed by shrinking (Figure 6.7). And when we increase the pulse
count further, to 2000, we see no swelling at all, only a reduction in cell volume
(Figure 6.7).
These results could be explained if some of the large intracellular solutes,
which remain impermeant after a dose of 30 pulses, are able to pass through the
membrane after a 100-pulse exposure, even though the larger PEG 1000
molecules in the extracellular medium are still blocked. In that case one might
expect to see first a swelling, resulting from the more rapid diffusion of water
and small molecules, followed by an osmotic equilibration resulting from the
100-pulse induced large-molecule leakage (associated with the slower diffusion
of the larger molecules), and subsequent shrinkage, since the PEG 1000
molecules still cannot diffuse into the cell. When 2000 pulses are delivered
instead of 100, the greater extent of the permeabilization, and the consequently
125
more rapid outward diffusion of intracellular solutes, eliminates the initial
swelling observed with the lower pulse dose.
Figure 6.7: The effect of extremely large number of 5 ns pulses on membrane
pore size in solutions isoosmotically replaced RPMI 1640 with PEG 1000 with
different concentration. Large number of 5 ns pulses did not create pores larger
the PEG 1000. However, the data in 50 and 116 mOsm PEG 1000 shows that
the intracellular solutes that were impermeable to cell plasma membrane at 30
pulses started to be able to penetrate the membrane while the pulse number over
100. The results are presented as mean±SE for at least 60 cells from at least
three independent experiments for each pulsing condition at 25 °C.
0.6$
0.7$
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela0ve$Area$Changes$$
Time$(second)$
116$mOsm$PEG1000$
0.6$
0.7$
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela0ve$Area$Changes$$
Time$(second)$
50$mOsm$PEG1000$
0.6$
0.7$
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela0ve$Area$Changes$$
Time$(second)$
200$mOsm$PEG1000$
0.6$
0.7$
0.8$
0.9$
1.0$
1.1$
1.2$
1.3$
0$ 50$ 100$ 150$ 200$ 250$ 300$
Rela0ve$Area$Changes$$
Time$(second)$
150$mOsm$PEG1000$
126
With a smaller osmotic differential at a PEG 1000 concentration of 116
mOsm, the initial pulse-induced swelling does not occur with either 100 or 2000
pulses. As with 50 mOsm PEG 1000, as the large intracellular solutes leak out
of the cell, decreasing the intracellular osmolality, water diffuses outward as
well, and the cell shrinks. In all of these cases the membrane remains
impermeant to PEG 1000, but the evidence can be interpreted to indicate that at
larger pulse exposures, some permeabilizing structures that may or may not be
pores, permit intracellular solutes larger than sucrose, but not as large as PEG
1000, to pass through the membrane,
The result shows that PEG1000 was impermeable to cell membrane even
after exposed to 2000 5 ns pulses. In other words, the size of pores created by
2000 5 ns pulses was still smaller than the size of PEG1000 molecules.
6.8 Conclusion
In conclusion, we have used inositol, sucrose, and PEG 1000 to estimate
the size of cell membrane pores created by 5 ns, 10 MV/m electric pulses. Pores
created by 5–100 pulses have a diameter between 0.7 nm and 0.9 nm. Cell
swelling is prevented by isosmotic PEG 1000 in the external medium with pulse
127
counts up to 2000, indicating that the pore diameter even after this extreme
exposure is still smaller than the size of PEG 1000 (about 1.9 nm, assuming that
the hydrodynamic radius of PEG 1000 is equivalent to the Einstein-Stokes
radius, which is 0.95 nm). It is not possible to determine with the method used
in this work the distribution of sizes in the population of pores created by the
electric pulse exposures, and for this reason it cannot be excluded that many
smaller pores are created along with pores in the 0.7 nm – 0.9 nm range. That
determination will require a careful calibration of transport rates for the various
solutes as a function of pore cross section and then an accurate measurement of
the flux observed experimentally.
128
Chapter 7: Conclusion
We have demonstrated the nsPEF-induced permeability change in
plasma membrane and intracellular membrane of malignant mammalian cells in-
vitro. The dependency of degree of electropermeabilization and pulse
parameters (pulse duration, pulse number, electric field amplitude and pulse
repetition rate) are studied and described in each chapter with different
experimental conditions. In addition to using fluorescent dyes for detection of
membrane integrity, we explore a new method based on osmotically driven cell
swelling in Chapter 4 and demonstrate that cell swelling can be used as a
sensitive indicator for studying plasma membrane electropermeabilization
induced by very short and very intense pulsed electric fields. In general,
increasing the pulse intensity, which including longer pulse duration, higher
electric field, faster pulse repetition rate and larger pulse number, will also
increase the degree of nsPEF-induced plasma membrane
electropermeabilization. Moreover, we characterize the population of plasma
membrane pores created by nsPEFs in Chapter 6. We have used small molecules
with different diameters, inositol, sucrose, and PEG 1000, to estimate the size of
cell membrane pores created by 5 ns, 10 MV/m electric pulses. The results show
that most pores created by 5–100 pulses have a diameter between 0.7 nm and
0.9 nm.
129
In Chapter 1, we demonstrated that cells pretreated with peroxidation
reagents (H
2
O
2
+FeSO
4
) show significant increase in membrane
permeabilization after exposure to 30 ns, 5 MV/m electric field. In Chapter 4,
we add membrane channel blockers, lanthanide ions (Gd
3+
and La
3+
) which
show protective effect of plasma membrane in pervious nsPEF study with
longer pulse (60 and 600 ns), or mercuric ions, (Hg
2+
) in the external medium.
After exposing to 5 ns and 10 MV/m electric pulses, we do not observe
significant change in plasma membrane permeability. Our results suggests that
the primary permeabilizing event triggered by our pulsing condition is formation
of lipid pores in the plasma membrane but not opening the membrane protein
channels. In conclusion, the studies of chemical reagents which can promote or
inhibit plasma membrane electropermeabilization can help one modulates the
pulsing protocols for different applications.
The evidence of permeabilizing mitochondrial membrane by nsPEF (4
ns duration, 10 MV/m, 1 kHz repetition rate) is shown in Chapter 3 by using
three different methods with fluorescence indicators—rhodamine 123, TMRE,
and cobalt-quenched calcein. The nsPEF may induce mitochondrial membrane
permebilization in a direct or indirect manner. Although we can not identify the
actual mechanism in our studies, these results still contribute to the
understanding of the nsPEF-induced intracellular effects in biological cells, and
130
demonstrating the potential applications of nsPEF to ablation of cancer cells
without drug and gene delivery.
In Chapter 6, we report a new delivery device for nsPEF. A pair of gold
plated tungsten wires is carefully aligned in parallel and spaced with 100 um in
between. We show that, by coupling the wire electrodes with nanosecond pulse
generator, fluorescent microscope and sensitive CCD camera, we can perform a
biocompatible, flexible and real-time observation of biological cells in-vitro.
131
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Abstract (if available)
Abstract
Nanosecond megavolts‐per‐meter pulsed electric field (nsPEF) offers a noninvasive manipulation of intracellular organelles and functions of biological cells. Accordingly, nsPEF is a potential technique for biophysical research and cancer therapy, and is of growing interest. Although, the application of nsPEF has shown electroperturbation on cell plasma membranes and intracellular membranes as well, the mechanisms underlying the electropermeabilization are still not clear. In this thesis, we systematically study nsPEFs (5 and 30 ns) induced membrane permeability change in biological cell in‐vitro with different pulse parameters. In Chapter 3, we investigate the nsPEF‐induced intracellular membrane permeabilization of mitochondria which play key roles in activating apoptosis in mammalian cells. The results show the evidences of nsPEF‐induced membrane permeability increase in mitochondria, and suggest that nsPEF is a potential technology for cancer cell ablation without delivery of drug or gene into cells. ❧ In Chapter 2, 4 and 6, we study the properties of nsPEF‐induced plasma membrane permeabilization. In the beginning, the change of plasma membrane permeability is studied by uptake of YO‐PRO‐1 and propidium iodide, fluorescent dyes specifically used as indicators of plasma membrane permeabilization. However, the detection is limited by the fluorescent emission efficiency and detector capability. To increase the detection sensitivity, we later develop a method based on cell volume change due to regulation of osmotic balance that causes water and small ions transport through plasma membrane. We find that even a single 10 MV/m pulse of 5 ns duration produces measureable cell swelling. The results demonstrate that cell swelling is susceptible to nsPEF and can detect membrane permeabilization more easily and precisely than fluorescent dyes. We compare the effects of different pulse parameters (pulse duration, pulse number, electric field amplitude and pulse repetition rate) on electropermeabilization. The effects of chemical agents that either promote (H₂O₂) or inhibit (lanthanide ions and Hg²⁺) electropermeabilization are also studied. To characterize the population of pores created by nsPEFs, we isoosmotically substitute different size of neutral molecules in the pulsing medium, and estimate pore size by analyzing cell volume changes that result from the permeation of these substituted molecules through the plasma membrane of Jurkat T lymphoblasts. The basis of this method is regulation of osmotic balance across the plasma membrane as well. We find that most pores opened by 5-100 5 ns pulses in plasma memebrane of Jurkat T lymphoblasts have diameter between 0.7-0.9 nm. ❧ In Chapter 5, we report the design and construction of a delivery system for nsPEF. We integrate a pair of delicately fabricated tungsten wire electrodes spaced 100 μm, a solid‐state high‐voltage nanosecond pulse generator and a fluorescent microscope coupling with a fast and sensitive digital recording camera. This system enables real‐time biophotonic investigations of the nsPEF‐induced biological responses of living mammalian cells in-vitro.
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Asset Metadata
Creator
Wu, Yu-Hsuan
(author)
Core Title
Biophysical studies of nanosecond pulsed electric field induced cell membrane permeabilization
School
Viterbi School of Engineering
Degree
Doctor of Philosophy
Degree Program
Materials Science
Publication Date
02/24/2014
Defense Date
01/15/2014
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cell membrane permeabilization,nanosecond pulsed electric field,OAI-PMH Harvest
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English
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Gundersen, Martin A. (
committee chair
), Goo, Edward K. (
committee member
), Wang, Pin (
committee member
), Zhou, Chongwu (
committee member
)
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sharonyhw@gmail.com,yuhsuanw@usc.edu
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cell membrane permeabilization
nanosecond pulsed electric field