Electroporation of Brain Endothelial Cells on Chip toward ...Article Electroporation of Brain Endothelial Cells on Chip toward Permeabilizing the Blood-Brain Barrier Mohammad Bonakdar,1,*

Biophysical Journal Volume 110 January 2016 503–513 503

Article

Electroporation of Brain Endothelial Cells on Chip toward Permeabilizingthe Blood-Brain Barrier

Mohammad Bonakdar,1,* Elisa M. Wasson,1 Yong W. Lee,2,3 and Rafael V. Davalos1,2,31Department of Mechanical Engineering and 2Department of Biomedical Engineering and Mechanics, Virginia Tech, Blacksburg, Virginia; and3School of Biomedical Engineering and Sciences, Virginia Tech – Wake Forest University, Virginia Tech, Blacksburg, Virginia

ABSTRACT The blood-brain barrier, mainly composed of brain microvascular endothelial cells, poses an obstacle to drug de-livery to the brain. Controlled permeabilization of the constituent brain endothelial cells can result in overcoming this barrier andincreasing transcellular transport across it. Electroporation is a biophysical phenomenon that has shown potential in permeabi-lizing and overcoming this barrier. In this study we developed a microengineered in vitro model to characterize the permeabili-zation of adhered brain endothelial cells to large molecules in response to applied pulsed electric fields. We found the distributionof affected cells by reversible and irreversible electroporation, and quantified the uptaken amount of naturally impermeable mol-ecules into the cells as a result of applied pulse magnitude and number of pulses. We achieved 81 5 1.7% (N ¼ 6) electropo-rated cells with 175 8% (N¼ 5) cell death using an electric-field magnitude of ~580 V/cm and 10 pulses. Our results provide theproper range for applied electric-field intensity and number of pulses for safe permeabilization without significantly compromisingcell viability. Our results demonstrate that it is possible to permeabilize the endothelial cells of the BBB in a controlled manner,therefore lending to the feasibility of using pulsed electric fields to increase drug transport across the BBB through the transcel-lular pathway.

INTRODUCTION

Many potential therapeutics for disorders of the central ner-vous system (CNS), which are proven to be effectivein vitro, have limited to no effect in vivo. This is due to theirlarge molecular weight, water solubility, and charge, whichrender it difficult for them to successfully cross the blood-brain barrier (BBB). Only small and lipid-soluble moleculesare able to passively cross the BBB, therefore protecting thebrain from harmful invading pathogens. Despite its protect-ing nature, most drugs used to treat CNS disorders are largewater-soluble molecules, therefore making the BBB a sig-nificant hindrance in treating patients. In the United Statesalone, 80,000,000 people are suffering from CNS disorderssuch as brain cancer, Alzheimer’s, and Parkinson’s dis-ease (1). Despite the high need for development of effectiveCNS drugs, only a small portion of newly discovered thera-peutics have been found to be clinically relevant and havethe ability to overcome the BBB.

The BBB is comprised of a layer of highly restrictive cellsthat line the walls of brain microvessels. Different vascularand parenchymal cell types contribute to the formation ofthis barrier including endothelial cells, astrocytes (2), peri-cytes (3), neurons, the extracellular matrix, and the base-ment membrane (4,5). Despite containing various celltypes, 75–80% of the contribution to barrier function is

Submitted August 14, 2015, and accepted for publication November 17,

2015.

*Correspondence: [email protected]

Editor: Jennifer Curtis.

� 2016 by the Biophysical Society

0006-3495/16/01/0503/11

made by the brain capillary endothelial cells liningthe luminal surface of the microvessels (6). These endothe-lial cells form intercellular tight junctions (TJs), whichlimit the transport of hydrophilic molecules larger than500 Da (7). Therefore, the biggest obstacle in deliveringan effective treatment to brain tissue is penetrating thisendothelial cell layer. Normal and disrupted transport acrossthis layer may occur through two pathways: the transcellularpathway, which is through the cell membrane and cytosol;and the paracellular pathway, which is located at thecell-cell TJs.

Many researchers have proposed different techniques totemporarily bypass or permeabilize the BBB through thesepathways. These include focused ultrasound (8,9), osmoticdisruption (10), drug delivery vehicles (11,12), and pulsedelectric fields (PEFs) (13,14) including electroporation(15,16). All of these techniques have inherent advantagesand drawbacks. Despite showing the capability to noninva-sively and reversibly disrupt the BBB, focused ultrasound islimited by the small coverage area the focal beam can target,leading to extended treatment times for large tissue volumes(17). Osmotic and pharmacological disruption using agentssuch as mannitol or bradykinin have also been studied as away to transiently open the BBB, but have been found to doso in a systemic manner that may lead to adverse effectssuch as headache, nausea, and abnormal neuron function(9,18). In addition, researchers have shown immunolipo-somes, endogenous peptides, or other vehicles successfullycarry therapeutic drugs across the BBB (19,20). Once there,

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504 Bonakdar et al.

many are unable to reach the concentration necessary foreffective treatment.

Alternatively, PEFs have shown promise in treatingneurological and psychiatric disorders. PEFs are short,intense electrical pulses that have been used in deep brainstimulation (21,22), electrochemotherapy (23,24), and irre-versible electroporation (IRE) (25–27) for tumor ablation.Electroporation is a biophysical phenomenon in whichPEFs applied across a cell cause the transmembrane poten-tial to exceed a certain threshold value, thus creating poresin the membrane (28). Electroporation can be either revers-ible (where pores reseal over time), or irreversible (wherepores grow too large to recover, according to the pulseparameters applied (29,30)). Irreversible electroporationresults in cell death due to the induced permanent defectsin the cell membrane. However, even when the cell mem-brane eventually recovers, the phenomenon may apoptoti-cally result in cell death due to the induced chemicalimbalance in the cell during the electroporation period(31). While irreversible electroporation may nonthermallyablate tissues (25), reversible electroporation may be usedin a controlled manner to temporarily disrupt the cell mem-brane for gene or drug delivery (32). In the paracellularpathway, TJs significantly restrict transport in between cellsdue to their secure connection to the cytoskeleton. BecausePEFs disrupt the cell membrane, and therefore the cytoskel-eton, they have the potential to disrupt the TJs as well (13).This indicates that PEFs have the potential to enhancepermeability of either the transcellular or the paracellularpathways of the BBB. In brain capillary cells, transcytosisof macromolecules is also extremely limited due to thelow amount of caveolae (33). Caveolae are lipid rafts thatselectively internalize molecules at the plasma membrane.According to the literature, one form of transcellular trans-port is passive diffusion through the lipid bilayer and thecytoplasm from the luminal to the abluminal side of theBBB (7). Because electroporation disrupts the lipid bilayer,it is concluded that it could potentially reduce the inherentresistance to passive diffusion across the transcellularpathway and increase the transcellular permeability of theBBB. To our knowledge, the distinction between permeabi-lization of different pathways across the BBB has rarelybeen studied in the literature.

Several studies have explored the effects of PEFs on theintegrity of the BBB. Lopez-Quintero et al. (13) probeddeep-brain-stimulation-relevant waveforms and intensitieson the permeability of the BBB using an in vitro setup.They found that the water flux across the endothelial celllayer is increased by applying high frequency PEFs of upto 185 Hz and 2.50 V/cm magnitude. Garcia et al. (15)investigated the effects of low intensity EF on BBB trans-port in an in vivo study. They found that the threshold forBBB disruption is between 400 and 600 V/cm for ninety50-ms pulses delivered at 1 Hz. In another in vivo study,Arena et al. (14) studied the effect of bipolar submicrosec-

Biophysical Journal 110(2) 503–513

ond pulses on BBB permeability and found that BBBdisruption occurs at much lower thresholds of 250 V/cmwithout any muscle contraction. Hjouj et al. (16) studiedreversible and irreversible electroporation thresholds ofthe BBB in rats and monitored the disruption by magneticresonance imaging. They found that reversible dis-ruption occurred between EF intensities of 330 V/cm and500 V/cm using a total number of 50–70 ms pulses at a fre-quency of 4 Hz, resulting in BBB disruption volumes largerthan tissue damage volumes. Other experimental studieshave also investigated the effect of electroporation on thepermeability of endothelial cells in vitro (34) and in vivo(35), with a focus on the paracellular pathway.

In vivo experiments on BBB permeabilization, such asthe ones mentioned above, provide mostly qualitative re-sults, therefore making in vitro models necessary for ob-taining more quantitative analysis; however, theirphysiological relevance is always a matter of debate.Several in vitro models have been previously developedto simulate the integrity and permeability of the BBB todifferent substances (36,37) as well as investigate the effectof different stimuli such as chemicals (38), pressure shock(39), radiation (40), and electromagnetic fields (41–43).Microfluidics studies provide an alternative platform tostandard in vitro systems for analyzing cells and tissuesin a more physiologically relevant environment (44,45).Organs-on-chip platforms utilize the advantages of micro-fluidics to recreate in vivo conditions of the body in aneasily testable, in vitro setting. Several investigators havedeveloped dynamic models of the BBB by culturing brainendothelial cells inside microfluidic channels (46–49) andinvestigating permeability of the BBB to differenttherapeutics.

While these in vivo and in vitro studies have providedinsight into the permeability of the BBB in response toPEFs and other therapeutics, no distinction has been madebetween the paracellular and transcellular pathways.Although both of these pathways are essential in transport-ing molecules across the BBB, each of them allow for quitespecific, distinct molecules to cross depending on their sizeand hydrophilic or hydrophobic nature. Therefore the abilityto target and controllably permeabilize each independentlymay allow for better delivery of drugs.

As previously mentioned, the transcellular pathway in-cludes the cell membrane, the cytoplasm, and again thecell membrane. For transcellular transport to occur, mole-cules are first uptaken through the luminal side of the cellmembrane into the cytoplasm, and then pass through tothe abluminal side of the membrane. In this work we studiedthe diffusion process by tracking the molecules during halfof their complete transcellular transport, which is from theluminal side to the cytosol of the endothelial cells. Moni-toring cellular uptake can give insight into possibleenhancement of transcellular transport of differentsubstances.

Electrode

Inlet

1mm 0.3mmA

B

BBB Electroporation on Chip 505

In this work we expanded on previous in vitro electropo-ration studies, which were mostly conducted on suspendedcells, by quantifying the uptaken amount of molecules inresponse to applied PEFs. The results from varying pulsestrength and number of applied pulses may provide informa-tion that can be used to find the proper parameters for thepossible enhancement of transcellular transport across theBBB using PEFs without causing any permanent damage.

Channel

FIGURE 1 (A) Tapered channel. (B) Microfluidic device comprising

three separate dual tapered channels.

MATERIALS AND METHODS

Device design and fabrication

The microfluidic channel used in these studies consisted of a nonlinearly

tapered geometry, 30 mm long and varying in width, ranging from 1000

mm at the ends to 300 mm at the center. Applying an electric potential to

the ends of the channel created a gradient of electric-field (EF) magnitude

inside the channel and over the cells. According to Ohm’s law, the EF

magnitude inside the channel is governed by

EðxÞ ¼ 1

sJðxÞ ¼ I

sAðxÞ; (1)

where s is the conductivity of the medium, J is the current density, x is the

length along the channel, and A is the cross-sectional area of the channel. It

was desired to have a linear gradient of EF along the length of the channel to

easily correlate these values with the results obtained from experiments.

According to Eq. 1 the magnitude of the EF and the cross section of the

channel are inversely proportional. Hence, for E(x) to be linear, the cross-

sectional area of the channel needed to be inversely proportional to the

length along the channel, x. That requires the channel-width profile to be

a section of the curve defined by the wðxÞ ¼ ðC=xÞ curve, where C is a con-

stant and the endpoints of the section are the values of the desired channel

widths, which are 1000 and 300. These endpoints and the constant C are

determined by satisfying the boundary conditions

wðxÞ ¼ C

x(wðaÞ ¼ 1000

wðaþ 15; 000Þ ¼ 300;

(2)

where a is the x axis value of the curve that gives us a width of 1000. Solv-

ing the above equations simultaneously gives the numeric values for a and

C. To have a symmetric channel geometry, two of the above sections were

attached end-to-end, as shown in Fig. 1 A.

Due to this geometry, the cells in themiddle of the channel experienced an

EF magnitude 3.3 times greater than the cells at the ends. Designing the de-

vice in thismanner ensured that each part of the channel showed cell behavior

in response to a specific EF magnitude. This gradient design allowed for

testing multiple conditions in one experiment. The symmetry of the dual

tapered channel also transitioned the high EF region from the channel end

(found in a single tapered channel) to itsmiddle,where, due to the ports, there

is no disruption of the induced EF. Moreover, the symmetry of the channel

mandated a symmetric response from the cells in the channel after exposure

to PEFs, which served as a verification tool for each experiment. Finite

element analysis was used to verify the magnitude of the EF at each point

along the channel. The analysis was performed in COMSOL Multiphysics

(Stockholm, Sweden) 4.4 in a two-dimensional model using the AC/DC

module. Boundary conditions were defined by a ground potential of 0 V

applied to one end of the channel (inlet) and a charged potential correspond-

ing to the pulse magnitude applied to the other end (outlet).

Standard photolithography was used to create a silicon mold and subse-

quently replication molding was used to fabricate the microfluidic channel

in PDMS (polydimethylsiloxane) (50) (Sylgard 184; Dow Corning, Wal-

tham, MA). PDMS was prepared by mixing the prepolymer and curing

agent with a 10:1 weight ratio. The mixture was then degassed and poured

onto the Si wafer. The PDMS was cured at 100�C for 1 h. After curing, the

inlets and outlets were punched and the PDMS device was bonded to a glass

slide using air plasma (Harrick Plasma, Ithaca, NY). Stainless-steel needles

were placed at the inlet and outlet to deliver the pulses. Fig. 1 B shows the

fabricated device.

Cell culture

Previous in vitro models of the BBB have been developed from a variety of

different primary cells and immortalized cell lines (51). In this studywe used

the mouse brain endothelial cell line, bEnd.3 (ATCC,Manassas, VA), which

has been shown to adequately represent the BBB (6). The bEnd.3 cells were

cultured in T-75 flasks at 37�C and 5% CO2 and maintained in complete

growth media consisting of DMEM (ATCC) supplemented with 10% (v/v)

fetal bovine serum (Atlanta Biologicals, Flowery Branch, GA) and

1% (v/v) penicillin-streptomycin (Life Technologies, Thermo Fisher Scien-

tific, Waltham, MA). The cells were routinely passaged at 70–90% conflu-

ence. To prepare the microfluidic device for cell seeding, the PDMS

channelswere first sterilizedwith ethanol. To promote cell adhesion and pro-

liferation, the channel was treated with 50 mg/mL human fibronectin (Trevi-

gen, Gaithersburg, MD) for 1 h in an incubator. Complete growth media was

then introduced into the device and incubated for 2 h. Endothelial cells were

collected by washing and detaching with trypsin. The trypsin was then

neutralized by the addition of media and centrifuged for 5 min at 120 � g.

The trypsin solutionwas removed and freshmediawas added to obtain a con-

centration of ~40,000,000 cells/mL. The cells were then introduced into the

device throughmanual injection using a syringe and tubing until an even dis-

tribution was achieved in the channel. The device was incubated at 37�C for

2 h, allowing the cells to fully attach. Then, complete media was provided to

the channel using media-filled pipet tips at the inlet and outlet. The devices

were incubated for two days at 37�C and 5% CO2 until the cells were

confluent in the channel.

Electroporation

A BTX pulse generator (Harvard Apparatus, Holliston, MA) was used to

apply PEFs to the microchannels. The pulse width and frequency were


200

300

400

500

600

700

800

-16 -14 -12 -10 -8 -6 -4 -2 0 2 4 6 8 10 12 14 16

EF N

orm

(V/c

m)

Channel Length (mm)

A

B

100μs1s

1500V

V/cm700

600

500

400

300

200

100

FIGURE 2 EF distribution in the channel as a result of applied potential

at the channel ends. (A) EF contours in the channel and (B) EF magnitude

along the channel centerline. To see this figure in color, go online.

506 Bonakdar et al.

set at 100 ms and 1 Hz, respectively, which is consistent with clinical elec-

troporation procedures (26). Pulse magnitude was set to 1500 V across the

channel for all experiments, which provides relevant EF magnitudes inside

the channel ranging from 214 to 714 V/cm. The number of pulses was var-

ied at 10, 30, and 90 to represent both standard electrochemotherapy and

IRE procedures and conditions between the two. To find the reversible

and irreversible electroporation thresholds, the cellular uptake of propidium

iodide (PI) (Life Technologies) both during and after application of PEFs

was monitored. PI has been previously used as a permeabilization tracer

during electroporation (52). In this study the choice of PI was based on

its molecular mass of 668 Da, which is comparable to several CNS drugs

listed in the Results and Discussion. To determine the threshold for the

onset of cellular uptake during electroporation, a solution of PI in PBS

was used as the electroporation buffer and injected into the channel imme-

diately before pulsing. While naturally impermeable, PI stains the cell

nuclei when the membrane is compromised. Therefore after pulsing and

disrupting the cell membrane, the PI molecules were able to enter the

cell and bind to DNA, allowing visualization of the affected cells. To deter-

mine the IRE threshold, the cells were pulsed and then allowed to recover

for 1 h. Small pores usually recover within a fraction of a second; however,

full recovery of larger pores may take longer (53). After recovery, the cells

were exposed to the PI solution. In this case, only the permanently damaged

cells were affected by PI, allowing visualization and quantification of the

IRE phenomenon. In both cases, NucBlue (Life Technologies) was used

to counterstain the nuclei and count the total number of cells. The uptake

of 4 kDa FITC-dextran (Sigma-Aldrich, St. Louis, MO) as a result of

applying PEFs was also investigated. In these experiments, the channels

were filled with a solution of the substance (10 mg/mL in PBS) before puls-

ing. After the pulsing is completed, the channel is rinsed with media for five

minutes to remove the FITC-dextran from the channel. That would restrict

the fluorescence to the cells and eliminate background light. The cells are

then visualized with fluorescent microscopy to quantify uptake.

Fluorescent microscopy and image processing

Fluorescent images were taken of the cells in the channel using an inverted

microscope (Carl Zeiss, Jena, Germany) equipped with a CoolSNAP HQ2

CCD camera (Photometrics, Tucson, AZ) and automated stage. The micro-

scope was controlled using the ZEN Pro 2012 software (Carl Zeiss). The

tiling feature of the software was used to image the entire length of the

channel. To enhance the contrast and brightness of the images and to re-

move background noise, images were processed in ImageJ (NIH, Bethesda,

MD). The intensity was then acquired along the channel and normalized to

the width of the channel. The normalized fluorescence intensity was used as

a quantitative measure of uptaken molecules as explained in the following

sections. To calculate the percentage of dead cells, the fluorescent light in-

tensities from PI (red) and NucBlue (blue) were adjusted in ImageJ to

restrict the light to the cell nuclei. This causes the light intensity from

each portion of the channel to be proportional to the number of cells.

The percentage of dead cells was obtained by dividing the intensity of

the dead cells (red) to that of the total cells (redþblue) in each 3-mm section

along the channel.

Confocal microscopy

Confocal microscopy was performed to obtain the average height of the cell

monolayer to develop the calibration curve used to quantify cellular uptake

of dextran. Before imaging, the cells were fixed with 4% paraformaldehyde

(Boston BioProducts, Ashland, MA) and stained with rhodamine phalloidin

and DAPI (Invitrogen, Carlsbad, CA), used to stain F-actin filaments and

nuclei, respectively. After staining, the cells in the microchannel were

imaged using a LSM880 confocal microscope (Carl Zeiss) with a 40�objective. The pinhole aperture for confocal fluorescence was adjusted

to produce a Z resolution of ~1 mm. Images were collected in steps of


0.1 mm in the Z direction. ZEN Black software (Carl Zeiss) was used to

analyze the sections and build the three-dimensional image.

Fluorescent calibration

To relate the fluorescence intensity of the images to the concentration of the

uptaken dextran molecules, a calibration curve was developed. This was

done by using microfluidic channels of different heights to simulate the

height of the adhered cells found using confocal microscopy. The channels

were filled with solutions of different concentrations of 4 kDa FITC-dextran

in PBS and imaged. The fluorescence intensity was then obtained using

ImageJ (National Institutes of Health).

Statistical analysis

Statistical analyses were performed using JMP Pro, ver. 11.0 (SAS Institute,

Cary, NC) with a confidence level of a ¼ 0.05. Two-way analysis of vari-

ance (ANOVA) was used to test for differences in the number of dead cells

after IRE treatment (N ¼ 5) as well as the cellular uptake of PI (N ¼ 6) and

dextran (N ¼ 3). When results of ANOVAwere significant, Tukey post hoc

comparisons were used to examine differences among treatment groups.

Data are presented as arithmetic mean 5 SD.

RESULTS AND DISCUSSION

EF distribution inside the channel using finiteelement analysis

Fig. 2 shows the EF distribution in the channel and the EFmagnitude along the centerline of the channel for an appliedpotential of 1500 V. As shown in the figure, the design of thechannel provided a linear variation in the EF magnitudefrom 213 V/cm at the ends to 714 V/cm at the center ofthe channel. It should also be noted that the EF magnitudedid not change significantly along the width of the channel.

Calibration of fluorescent intensity for differentheights in terms of the concentration offluorescent molecules

Fig. 3 shows the fluorescent intensity as a function ofthe concentration of 4 kDa FITC-dextran inside channelsof different heights. Our observations using confocal

0

50

100

150

200

250

0 5 10 15 20 25

Fluo

resc

ent I

nten

sity

Channel Height (μm)

60 µg/ml

300 µg/ml

600 µg/ml

B

0

50

100

150

200

250

0 100 200 300 400 500 600

Fluo

resc

ent I

nten

sity

Dextran Concentra�on (μg/ml)

4.78

8.8

15.3521.23

Aµm

µmµm

µm

FIGURE 3 Calibration curves for fluorescent intensity of 4 kDa FITC

dextran in the microchannel: (A) for different heights; (B) for different con-

centrations.


microscopy showed that after adhesion and spreading, thethickness of the cell monolayer varied between 2 mm (atthe cell edges) to ~7 mm (above the nuclei). Hence, fourdifferent heights around this range were selected for obtain-ing the calibration curves. It was found that the intensity var-ied linearly with respect to both the concentration and theheight. Linear regression was used to find an empirical rela-tion giving the fluorescence intensity as a function of FITC-dextran concentration and height. The linear dependence ofintensity on channel height was used in the following sec-tion to obtain the profile of adhered endothelial cells accord-ing to the fluorescence of absorbed dextran.

Characterization of the cell monolayer heightinside the microchannel

Fig. 4 shows cross-sectional images of the cell monolayer atdifferent heights from the channel base, as well as severalside views of the cells along different transverse sections.The maximum height of the cells was located above thecell nuclei with an average value of 7.0 5 1.4 mm. Therest of the cell body possessed a maximum height of<4 mm. The staining technique clearly visualized theboundary of the nucleus, but not the boundary of the cellbody, as shown in the side views of Fig. 4. Despite a sharply

contrasted nucleus, the rest of the cell body was blurred.Hence another technique using the fluorescent intensity ofuptaken FITC-dextran was used to find the outer profile ofadhered cells. Based on the size of the cells and the diffu-sivity of the dextran molecules inside the cytosol, it ispossible to assume a uniform distribution of absorbeddextran in the cell body after a few minutes. Therefore,the fluorescent intensity at each point becomes linearly pro-portional to the height of the adhered cell, as shown in thecalibration section (Fig. 3 B). We used this fact in additionto the absolute height of the cells above the nuclei—accord-ing to the confocal imaging—to find the height distributionof the adhered cells.

Fig. 5 shows the adhered bEnd.3 cells in the microchanneland the outer profile along their longitudinal directions. Us-ing this method, the average height of the endothelial cellmonolayer was calculated to be 3.15 mm (N ¼ 30).

Determination of the reversible and irreversibleelectroporation thresholds

When using PEFs to permeabilize the endothelial cells ofthe BBB for drug transport, it is desirable to avoid irrevers-ible effects on the cell membrane, which can result in per-manent damage and leakage of the blood vessels. Hence,it is necessary to know the IRE threshold and the marginfor reversible electroporation. The IRE threshold is a func-tion of an applied number of pulses. A treatment comprisingmore pulses needs a lower EF magnitude to induce IRE.Fig. 6 shows the distribution of dead cells inside the channelfor different numbers of pulses. (See Fig. S1 in the Support-ing Material for a high-resolution image of the electropo-rated cells in the channel.) As shown, by increasing thenumber of pulses, more cells in the lower EF region (widersection of the channel) were irreversibly electroporated. Ex-panding upon this, Fig. 7 shows the percentage of dead cellsalong the channel for different numbers of pulses and pulsemagnitudes. It should be noted that no chemical reaction orbubbling was observed around the pulsing electrodes insidethe channel, which is mainly attributed to the small pulsewidth (100 ms) and low current (~2 mA).

Factors such as cell orientation (54), cell size, and distri-bution density (55) cause individual cells to be affecteddifferently by the pulses of the same magnitude. Our exper-iments showed that the cells that are oriented parallel to theelectric field get electroporated at a lower electric field thanthose that are perpendicular to the field. (See Movie S1 fortime-lapse videos of cells during electroporation.) Previousmodeling for spheroidal cells has proven the same behavior(54). This results in a distribution of live/dead cells along thechannel instead of a sharp delineation between live and deadcells. The percentage of dead cells differed significantlyamong the examined treatment parameters (ANOVA, p <0.0001). Post hoc comparisons indicated that the dead cellpercentage for 10 pulses was significantly different from


FIGURE 4 Confocal images of the endothelial

cell monolayer. (Top) Cross-sectional images of

the bEnd.3 cell monolayer in the microchannel

taken at different heights. (Bottom) Side views

show the relative height of the nuclei (blue)

compared to the rest of the cell body (red). To see

this figure in color, go online.

508 Bonakdar et al.

30 and 90 pulses at higher EF magnitudes whereas lower EFmagnitudes (245–314 V/cm) did not show a significant dif-ference. These results can be seen in Fig. 7, which showsthat for a wide range of EF magnitude, the induced celldeath for 10 pulses (0–40%) is much lower than 30 and90 pulses (5–100%). This finding is consistent with thefact that usually 8–10 pulses are applied for drug deliveryduring electrochemotherapy. For 10 pulses, almost no celldeath is observed for EF <400 V/cm. It can also beseen that 30 and 90 pulses at higher EF magnitudes(646–714 V/cm) and low EF magnitudes (245 V/cm) didnot result in a significantly different percentage of cell deathwhereas an EF magnitude of 314 V/cm for 30 and 90 pulsesresulted in significantly different percentages. Similarin vitro experiments have been conducted by Dermol et al.(56), where pulse number and electric field were varied todetermine the influence these parameters have on cellsurvival and also to evaluate mathematical models of cellsurvival. The authors tested Chinese hamster ovary cellsin suspension with pulse numbers of 8, 30, and 90. Eachof these pulses were 100 ms in duration, with the electricfield ranging from 500 to 4000 V/cm. It was found that

1

2

3A

100 μm

0

4

8

0 20 40 60 80 100 120 140 160Cell

Heig

ht (μ

m)

Cell axis (μm)

sec�on 1 sec�on 2 sec�on 3B

FIGURE 5 Finding the height profile of cells from fluorescent imaging.

(A) Fluorescence image of adhered bEnd3 cells with absorbed FITC-

dextran. (B) Height profile of the adhered cells.


the majority of cells survived treatments with an electricfield of 1000 V/cm with little distinction between differentpulse numbers. A difference among 8, 30, and 90 pulseswas not seen until the applied electric field reached2000 V/cm. Therefore the threshold for irreversible electro-poration was found to be higher than what was found in ourexperiments, where close to 100% of cells became irrevers-ibly electroporated at 714 V/cm (Fig. 7). This may bebecause we are using a different cell type that containsstrong intercellular junctions. These junctions as well ascell geometry have been shown to influence transmembranepotential (57), causing results to be different from cellstested in suspension. Previous in vivo studies also showthat an EF magnitude of 500 V/cm and 90 pulses induceIRE (16). Our results show that the same pulsing, resultsin 85% cell death.

Fig. 8 shows the distribution of electroporated cells visual-ized by uptake of PI. The percentage of electroporated cellsdiffered significantly among the examined treatment param-eters (ANOVA, p< 0.0001). Post hoc comparisons indicatedthat the electroporated cell percentage for 10 pulses wassignificantly different from 30 and 90 pulses at mid-rangeEF magnitudes (314–513 V/cm), whereas low and high EFmagnitudes (245, 646–714 V/cm) did not show a significantdifference. Differences between 30 and 90 pulses were notstatistically significant for equivalent EF, which indicatesthat 30 pulses deliver an amount of PI into the cell comparableto 90 pulses. The electroporated cells spanned a wider rangeof EF compared to the irreversibly electroporated cells. Thisis because the cells were in contact with the PImolecules dur-ing treatment; as soon as the pores opened, the moleculespermeated the cells. However, in the case of IRE, only thepores that were not recovered after 1 h allowed the transportof PI molecules into the cell. (Refer to Movie S1 for videosshowing the uptake of PI during electroporation.) Dermoland Miklav�ci�c (58) also conducted electroporation experi-ments on adhered cells after delivering only 1 pulse of1 ms, and found that the higher the cell density, the lowerthe percentage of cells that are electroporated. It is difficultto compare our results directly with this study due to the dif-ference in the applied number of pulses. However, our resultsshow that the percentage of electroporated cells is higher than

90 Pulses

30 Pulses

10 Pulses

714

646

581

513

447

379

3 14

245

646

581

513

447

379

314

245

V/cm

FIGURE 6 Distribution of IRE-treated cells for

different numbers of pulses. (See Fig. S1 for high-

resolution color image.)


what is found in Dermol and Miklav�ci�c (58). This is becausethe bEnd3 cells spread out and become very long in the chan-nel, leading to a lower density of cells in the channel thanwhat would be found with Chinese hamster ovary cells.This lower density in turn lowers the electroporationthreshold. As previously mentioned, it is also possible thatsome of the bEnd3 cells were aligned parallel to the electricfield, therefore causing them to be electroporated at a lowerthreshold.

By comparing Figs. 7 and 8, one can find the proper num-ber and magnitude of pulses for cell electroporation withminimal cell death. This comparison is made in Fig. 9 forthe specific case of 10 and 90 pulses, which are typicallyused for electrochemotherapy and tumor ablation treatments,respectively. This comparison demonstrates how differentEF magnitudes and pulse numbers can electroporate apopulation of cells while keeping them viable. It is shownthat for 10 pulses, there is a drastic difference between theonset of electroporation and the onset of IRE. For example,10 pulses at 513V/cm electroporated ~60%of the cells whilecausing only 10% cell death. On the other hand for the case of

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FIGURE 7 Distribution of dead cell population after electroporation. (A)

Dead cell population along the channel for different numbers of pulses. (B)

Dead cell population as a function of PEF magnitude for different numbers

of pulses.

90 pulses, <20% of the cells that were electroporated re-mained alive. These results explain the reason eight pulsesare typically delivered during electrochemotherapy treat-ments (59). It should be noted that PI becomes fluorescentonce it binds to DNA after entering the cell. Because PI isnot intrinsically fluorescent, the emitted light is restrictedto the cell nuclei and does not reflect the quantitative amountof PI molecules present in the cell body. To study this uptakephenomenon in a more quantitative manner, the cells wereelectroporated in the presence of FITC-dextran.

Uptake of large impermeable molecules dueto PEFs

Four kiloDaltons of FITC-dextran is intrinsically fluorescentand naturally impermeable to the cells due to its large sizeand hydrophilic nature. However, this substance can pene-trate the cell membrane through induced pores formed byelectroporation. Fig. 10 shows the uptake of 4 kDa dextranafter electroporation. (For a high-resolution image, refer toFig. S2.) The percentage of affected cells differed signifi-cantly among the examined treatment parameters (ANOVA,p< 0.0001). Post hoc comparisons indicated that the uptakeof FITC-dextran into cells for 10 pulses was significantlydifferent from 30 and 90 pulses at high EF magnitudes(646–714 V/cm) whereas low EF magnitudes for 30pulses (245–314 V/cm) and midrange EF for 90 pulses(447–513 V/cm) did not show a significant difference.Differences between 30 and 90 pulses were statistically sig-nificant for midrange EF magnitudes (447–581 V/cm).Fig. 11 indicates that uptake of dextran into the cells wasmore effective for lower EF magnitudes for 30 and 90pulses, which initially may seem counterintuitive. For cellsthat were reversibly electroporated, the molecules becametrapped inside the cells after the pores were recovered. Forthe cells that were irreversibly electroporated, it is possiblethat the membrane remained leaky and could not retain theabsorbed molecules for the duration of the experiment.Therefore, we hypothesize that dextran molecules escapedthe cells during the 5-min washing period, reducing theemitted fluorescent light from those regions of the channel.This behavior was not observed for the uptake of PI (Fig. 8),because PI binds to the DNA as soon as uptake occurs andcannot exit the cell. Fig. 11 A shows the fluorescent intensityof absorbed 4 kDa dextran along the channel for differentnumbers of pulses. Knowing the calibration curve for fluo-rescent intensity and the cell monolayer thickness, it was


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FIGURE 8 Distribution of PI-permeated cells after electroporation. (A)

PI-permeated cell population along the channel for different numbers of

pulses. (B) PI-permeated cell population as a function of PEF magnitude.

510 Bonakdar et al.

possible to find the approximate concentration of the accu-mulated dextran inside the cells. Fig. 11 B shows dextranconcentration versus EF magnitude for different numbersof pulses. Depending on the applied number of pulses,different ranges of EF magnitude gave maximum uptakeof the dextran molecules. For the case of 10 pulses, gener-ally the higher the EF magnitude, the higher the amountof uptake. This is due to the dominant occurrence of revers-ible electroporation instead of IRE even at the highest EFmagnitude of 714 V/cm. In other words, despite some cellsdying by increasing the EF magnitude, other cells uptakeenough molecules that the overall uptake by the monolayeris seen as increasing. However, that was not the case whenthe number of applied pulses was increased. Increasingthe number of pulses resulted in more cell death at the

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higher EF zone. The maximum uptake for 10 pulses (at714 V/cm) is higher than the maximum uptake for 30 and90 pulses (at 447 and 379 V/cm, respectively), because thereis a low percentage of dead cells at 10 pulses compared to 30and 90, as seen in Fig. 7.

It should be noted that in the case of passive diffusionacross the intact cell membrane, the transcellular pathwayis restricted to lipophilic molecules. However, electropora-tion can induce hydrophilic pores into the membrane, whichmay facilitate the transport of hydrophilic molecules such asdextran in this case. In the above experiments, the rationalefor choosing PI (668 Da) and FITC-dextran (4 kDa) as thetarget molecules was their resemblance in size to severaldrugs, which are currently being administered to patientswith brain diseases and tumors for which penetration ofthe BBB remains a challenge. These drugs include Bleomy-cin, 1415 Da; Doxorubicin, 543 Da; Amphotericin B,923 Da; and Paclitaxel, 853 Da. Being slightly larger thanthese drugs, the results for the uptake of 4 kDa FITC-dextraninto brain endothelial cells gives an upper limit for the up-take of these drugs into the BBB by applying the properPEFs without causing any permanent damage to the BBB.

In addition to reversible electroporation that opens thetranscellular pathway for transfer of substances, PEFsmay also disrupt the TJs between the adjacent cells thatopen the paracellular pathway (60). This study was specif-ically aimed at finding the thresholds for reversibleelectroporation and cellular uptake of different-sized mol-ecules, which could yield relevant information about thepossibility of transcellular transport across the BBB.Studying the actual permeability coefficients for paracellu-lar and transcellular pathways requires access to bothluminal and abluminal sides of the BBB, which is leftfor future studies.

CONCLUSION

In this study we quantified the uptaken amount of moleculesinto adhered brain endothelial cells as a function of 1) EFmagnitude and 2) the number of pulses by using emittedfluorescent light from the electroporated cells. To the bestof our knowledge, it is the first time that such analysishas been performed for electroporated cells. We imple-mented a tapered microfluidic channel to apply a gradientof EF on adhered brain endothelial cells and visualizedthe electroporation phenomenon by tracking the uptake ofdifferent naturally impermeable molecules into the cells.The tapered design allowed for testing of multiple condi-tions in one experiment, therefore making it a useful plat-form to test drug delivery using PEFs. Using thisplatform, we were able to show the difference betweenreversible and irreversible electroporation for different pulsenumbers and a wide range of EF magnitudes. The majorityof the cells that were electroporated with 10 pulses recov-ered; however, electroporation with 30 and 90 pulses was

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FIGURE 10 Uptake of 4 kDa FITC-dextran for

different numbers of pulses. (See Fig. S2 for a

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mostly irreversible. Results for 30 and 90 pulses weresimilar for the entire range of EF magnitude, although 90pulses caused more cell death. These results provide theproper range of applied EF magnitude and number of pulsesfor safe permeabilization without significantly compro-mising cell viability. Our results demonstrate that it ispossible to permeabilize the endothelial cells of the BBBin a controlled manner, adding to the feasibility of usingPEFs to increase drug transport across the BBB via thetranscellular pathway.

SUPPORTING MATERIAL

Two figures and one movie are available at http://www.biophysj.org/

biophysj/supplemental/S0006-3495(15)04701-3.

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FIGURE 11 Dextran uptake during electroporation. (A) Fluorescent in-

tensity along the channel for different numbers of pulses. (B) Dextran con-

centration inside cell after pulsing versus EF for different numbers of

pulses. The dextran concentration in the pulsing medium was 10 mg/mL.

(Refer to the SupportingMaterial for a high-resolution image of dextran up-

take into the cells within the channel.)

AUTHOR CONTRIBUTIONS

M.B. designed and fabricated the device, ran the experiments, analyzed the

results, and wrote the article; E.M.W. analyzed the data, ran the statistics,

and contributed in the experiments and writing of the article; R.V.D. de-

signed the experiments and device, edited the manuscript, and the work

was done under his supervision; and Y.W.L. contributed in writing the

article. All authors have read and approved the article.

ACKNOWLEDGMENTS

This study was supported in part by the National Science Foundation under

NSF CAREER award Nos. CBET-1055913 and NSF EFRI-0938047 and by

the National Institutes of Health under NIH award No. 5R21 CA173092-01.

The authors also acknowledge support from Institute for Critical Technology

and Applied Science of Virginia Tech for general support of this research.

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Electroporation of Brain Endothelial Cells on Chip toward ...Article Electroporation of Brain Endothelial Cells on Chip toward Permeabilizing the Blood-Brain Barrier Mohammad Bonakdar,1,*