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. Author manuscript; available in PMC: 2020 Dec 1.
Published in final edited form as: Bioelectrochemistry. 2019 Jul 6;130:107328. doi: 10.1016/j.bioelechem.2019.107328

Electroporation-induced changes in tumor vasculature and microenvironment can promote the delivery and increase the efficacy of sorafenib nanoparticles

Hiroshi Kodama 1, Yosef Shamay 2, Yasushi Kimura 3, Janki Shah 4, Stephen B Solomon 1, Daniel Heller 4,**, Govindarajan Srimathveeravalli 3,5,*
PMCID: PMC6859646  NIHMSID: NIHMS1535260  PMID: 31306879

Abstract

Blood vessels, the extracellular space, and the cell membrane represent physiologic barriers to nanoparticle-based drug delivery for cancer therapy. We demonstrate that electroporation (EP) can assist in the delivery of dye stabilized sorafenib nanoparticles (SFB-IR783) by increasing the permeability of endothelial monolayers, improving diffusion through the extracellular space in tumorspheres, and by disrupting plasma membrane function in cancer cells. These changes occur in a dose-dependent fashion, increasing proportionally with electric field strength. Cell death from irreversible electroporation (IRE) was observed to contribute to the persistent transport of SFB-IR783 through these physiologic barriers. In a model of mice bearing bilateral xenograft HCT116 colorectal tumors, treatment with EP resulted in the immediate and increased uptake of SFB-IR783 when compared with the untreated contralateral tumor. The uptake of SFB-IR783 was independent of direct transfection of cells through EP and was mediated by changes in vascular permeability and extracellular diffusion. The combination of EP and SFB-IR783 was observed to result in 40% reduction in mean tumor diameter when compared with sham treatment (p < 0.05) at the time of sacrifice, which was not observed in cohorts treated with EP alone or SFB-IR783 alone. Treatment of tumor with EP can augment the uptake and increase the efficacy of nanoparticle therapy.

1. Introduction

Dysfunctional blood vessels and impaired lymphatic drainage in tumors aid the increased uptake of drugs that are encapsulated in or formulated as nanoparticles [1,2], forming the rationale for nanoparticle-based cancer therapy. Nevertheless, despite the propensity of nanoparticles to accumulate within tumors, microvascular permeability [3], the extracellular space [4], and the cell membrane [5] represent three physiologic barriers limiting intracellular drug concentration below cytotoxic levels. Therefore, active and passive modifications of nanoparticles as well as adjuvant techniques have been investigated to improve nanoparticle delivery into tumors [6-9]. Apart from functional and structural modifications, adjuvant techniques have also been identified and developed to enhance localized delivery of nanoparticles; for example, ionizing radiation [10] and non-ionizing energy in the form of heat [11-13] or light [14-15] can trigger biological changes in the tumor microenvironment that increase nanoparticle uptake. While nanoparticle modification and adjuvants targeting one or more physiologic barriers have been shown to increase treatment efficacy in the preclinical setting, we anticipate that a technique that can simultaneously modulate all three barriers is desirable and will have an important clinical impact.

The use of electric pulses to permeabilize the cell membrane is termed electroporation (EP), where permeabilization can be either transient (reversible electroporation, RE) or permanent (irreversible electroporation, IRE) [16]. While RE is widely used to facilitate the delivery of material that are cell impermeant (DNA plasmids) or material that diffuse poorly through an intact cell membrane (drugs such as bleomycin or cisplatin) [17], IRE is used to induce cell death through the loss of homeostasis [18] and has been developed for the ablation of tumors in the liver, lung, prostate, and pancreas. Besides membrane permeabilization, the electric pulses used for EP has also been reported to modulate vascular permeability. This was first demonstrated by Sersa et al. [19] in tumors, followed by Gehl et al. [20] in normal vasculature. In addition, EP has been reported to modulate extracellular diffusion properties under in vitro and in vivo conditions. This was demonstrated by Gibot et al. [21] who reported that RE increased the extent to which doxorubicin penetrates tumorspheres and by Zhang et al. [22,23] who reported that IRE altered tumor diffusion properties quantified with magnetic resonance imaging. Lastly, EP has been shown to modulate transfer into the cell membrane. Using a radiolabeled liposomal nanoparticle, Srimathveeravalli et al. [24] demonstrated that such secondary effects of RE significantly altered the pharmacokinetics of nanoparticle uptake in tumors, which was used to increase intratumoral uptake and concentration of liposomal doxorubicin.

As EP can be used to simultaneously modulate the barrier function of the cell membrane, tumor microvasculature, and the extracellular space, it is as an attractive candidate for promoting nanoparticle-based cancer therapy for locally advanced tumors. Our objective in this work was to study the effect of EP on endothelial monolayers and tumorspheres in vitro, with EP performed at varying energy doses to determine the optimal conditions for the treatment of tumors with near infrared (NIR) dye-stabilized sorafenib nanoparticles.

2. Materials and methods

2.1. Nanoparticle synthesis

Dye-stabilized sorafenib nanoparticles (SFB-IR783) were synthesized using a nano-precipitation method. Sorafenib dissolved in DMSO (10 mg/ml) was added drop-wise to a concentrated IR783 solution (1 mg/ml) in a sodium bicarbonate buffer (0.05 mM). The resultant mixture was centrifuged (at 20,000 g for 20 min) and resuspended in phosphate-buffered saline (PBS). Nanoparticles were characterized by Dynamic Light Scattering (DLS) and Atomic Force Microscopy (AFM). The content of the drug was measured by high performance liquid chromatography (HPLC) UV-VIS (Agilent) with a C18 column and gradient of Acetonitirle/Water (+Trifluoro acetic acid) and absorbance at 280 nm. The resultant SFB-IR783 nanoparticles had a zeta potential of -49 ± 5 mV and a size of 86 ± 3 nm (PDI = 0.09) (Supplemental Fig. 1).

2.2. Cell culture

Human endothelial (EA.hy926) and colorectal carcinoma (HCT116) cell lines were acquired from the American Type Culture Collection (Manassas, VA). All cell lines were grown in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum, 1% sodium pyruvate (100mM), 2% L-glutamine (200mM), penicillin, and streptomycin. Cells were cultured and maintained in an incubator at 37°C and 5% C02.

2.3. Endothelial monolayer assays

EA.hy926 endothelial cells were grown to confluence in either 24-well plates or in 6.5 mm transwell inserts with a polycarbonate membrane (3.0 μm pore size, Sigma-Aldrich, St. Louis, MO). The transwell inserts were coated with 200 μl of basement membrane gel (Matrigel, Sigma-Aldrich, St. Louis, MO). In both the well plates and transwell inserts, cells were seeded at 80% confluence and grown for 3–5 days prior to their use in experiments.

2.4. Creation of tumorspheres

HCT116 ce00lls were seeded (1.0 × 106 cells/ml) in ultra-low attachment cell culture flasks (Corning, Corning, NY) and grown for 2–5 days to allow for the aggregation of cells into tumorspheres. Tumorspheres that were approximately 200–300 μm in diameter were harvested for use in experiments. HCT116 cells were also grown as monolayers in 24-well plates to determine cell viability and proliferation following treatment with EP and nanoparticles.

2.5. In vitro EP

EP for all in vitro experiments was performed using an ECM 830 (BTX Technologies, Holliston, MA) square wave generator. The effect of EP was assessed at three energy settings (low: 500, intermediate: 1000, or high: 1500 V/cm) without varying the other pulse parameters (8 pulses, 100 μs pulse length, delivered at 1 Hz). Pulse delivery was performed using two stainless steel pin electrodes (25 mm length, 0.8 mm diameter, McMaster Carr, NJ) with 4 mm (for tumorspheres and transwell insert experiments) or 10 mm (for experiments in 24-well plates) spacing between electrodes, or using cuvettes (Micropulser Cuvette, BioRad, Hercules, CA) with a 4 mm electrode gap.

2.6. Endothelial monolayer morphology and permeability

Following EP, the nuclei and cell membrane of EA.hy926 cell monolayers in 24-well plates were stained with Hoechst 33258 and CellMask Green (Thermo fisher scientific, Waltham, MA both dyes at a concertation of 2 μl/ml), imaged at excitation/emission wavelengths of 460–480/500–550 nm (CellMask Green) and 340–380/420–480 nm (Hoechst 33258) at 1, 8, and 24 hours post EP with fluorescent microscopy (IX51, Olympus, Tokyo, Japan). Images were obtained at 20× magnification from at least 5 separate locations from each well and were evaluated for morphologic changes in the monolayer, resulting in loss of cell-cell contact and creation of intercellular “gaps”Image processing (ImageJ, National Institutes of Health, Bethesda, MD) was used to quantify the number, size, and distribution of such intercellular gaps. Pre- and post-EP comparison of nuclear staining were used identify gaps resulting from detached or dead cells. Confluent EA.hy926 cell monolayers in transwell inserts were treated with EP, and the transport of a small molecule dye (IR-783,MW 747 Sigma-Aldrich, St. Louis, MO) and SFB-IR783 nanoparticles was measured in the well underlying the insert at 1, 2, 4, 6, 8, 12, 24, 36, and 48 h post EP using a scanning spectrometer (Tecan, Mannedorf, Switzerland).

2.7. Tumorsphere morphology and permeability

Tumorspheres suspended in 300 μl of medium underwent EP in a cuvette and were subsequently seeded into 24-well plates. Changes in tumor morphology was imaged using a scanning electron microscope at 1-h post EP by collecting the tumorspheres in 1 mL tubes, allowing for gravity sedimentation, and placing them on poly-L-lysine-coated plastic coverslips (Thermonex) as follows. To assess changes in tumor permeability, the spheroids were then fixed in 2.5% paraformaldehyde (PFA) in 0.075 M cacodylate buffer for 1 h, rinsed in cacodylate buffer, and dehydrated in a graded series of alcohols: 50%, 75%, 95%, and absolute alcohol. The samples were then critical-point dried in a Denton Critical Point Dryer JCP-1. The coverslips were attached to SEM stubs and sputter-coated with gold/palladium in a Denton Vacuum Desk 1V. Images were obtained with a Zeiss Scanning Field Emission Supra 25 Scanning Electron Microscope. The penetration of a small molecule dye (Hoechst 33258) and SFB-IR783 nanoparticles into tumorspheres were imaged at excitation/emission wavelengths of 340-380/420-480 nm (Hoechst 33258) and at 690–730/750–800 nm (SFB-IR783) at 1 h post-EP with fluorescent microscopy. Images were taken with a fixed exposure time (200 μS). Changes in the mean intensity of fluorescence in individual tumorspheres from the presence of Hoechst 33258 or Sorafenib NP was quantified using ImageJ.

2.8. Viability and in vitro efficacy

Viability of EA.hy926 following EP and of HCT116 cell monolayers in 24-well plates when treated with EP, with or without SFB-IR783, was assessed using the MTT assay (Thermo Fisher Scientific, Waltham, MA), and values were normalized to sham controls. The viability of tumorspheres was determined by sequential measurement of size at 24 and 48 h post EP.

2.9. In vivo evaluation

Following techniques in a protocol approved by the Institutional Animal Care and Use Committee (IACUC), six-week-old female athymic NU/NU nude mice (Charles River Laboratories, Wilmington, MA) were implanted with HCT116 cells (1.0 × 106 cells in 100 μL of media) to generate unilateral or bilateral flank tumors. Tumors were allowed to grow until they reached 5–7 mm in size, after which the mice were randomly assigned to receive EP or sham treatment, with or without injection of SFB-IR783.

2.10. Imaging

Mice (n = 5) with bilateral flank tumors underwent treatment of one tumor with EP (1000 V/cm, 100 μs. 1 Hz, 8 pulses) while the contralateral side served as the control. EP was performed using Tweezertrodes (BTX Technologies, Holliston, MA) with eight 100 μs square pulses delivered at 1000 V/cm. SFB-IR783 (30mg/kg) was administered via tail vein injection within 3 min following pulse delivery. At 2 and 24 h after the treatment, animals underwent optical imaging (IVIS Spectrum, PerkinElmer Inc., Waltham, MA) and were then euthanized to determine the relative SFB-IR783 uptake in the tumor, lung, heart, liver, spleen, and kidney. Fluorescence intensity measurements (Living Image Software, PerkinElmer Inc., Waltham, MA) were compared between the EP-treated and the contralateral untreated tumor. Tumors extracted 24 h post EP underwent immunohistochemistry processing, with quantitative whole-slide evaluation for cell death (Cleaved Caspase-3) and proliferation (Ki-67).

2.11. Treatment Efficacy

Mice with unilateral flank subcutaneous tumor were randomized to the following cohorts: control, EP, SFB-IR783, and EP + SFB-IR783 (n = 5/cohort). EP and administration of SFB-IR783 were performed as per previous imaging experiments. Tumor growth was monitored with bi-weekly caliper measurements. Tumor volumes were calculated using the following formula: (length × width2) × (π/6). Tumor growth rate was computed as G.R. = In (V(t)V(i))∕t; where V(t) and V(i) represented the volume of the tumor at beginning and at the end of the measurement period denoted by t, respectively. The tumor doubling time was computed as T. D. = ln(2) /GR.

2.12. Statistical analysis

All in vitro experiments were performed in triplicate. All data was recorded as mean +/- standard deviation. Student’s t-test was used to determine the statistical significance of difference between cohorts when the data demonstrated normal distribution. The Mann-Whitney U test was used in datasets without normal distribution. A p value of less than 0.05 was considered indicative of significant difference between experimental conditions. All statistical calculations were performed using EZR software (Saitama Medical Center, Jichi Medical University, Saitama, Japan).

3. Results

3.1. Changes in cell diameter and cell death contribute to the formation of intercellular gaps in endothelial monolayers treated with EP

We assessed the effects of EP on EA.hy926 endothelial cells in response to increasing electric field strengths while other pulse parameters remained constant. Compared with controls (Fig 1 A)-(C), EP resulted in the rapid creation of intercellular gaps in confluent endothelial cell monolayers (Fig 1D-L) irrespective of the energy dose applied. At 1 h post EP, the appearance of intercellular gaps was associated with cell swelling (Fig 1 D) for all treatment groups, while cell shrinking (Fig 1 K,L), detachment (Fig 1 G-L), and cell death from IRE were observed only for the intermediate or high dose groups. Morphologically, the gaps were not evident at 8 h post EP in the low dose group (Fig 1 E), while they were persistent and obvious even at 24 h post EP in monolayers in the intermediate or high energy dose groups (Fig 1 I,L). The number of intercellular gaps per field of view (FOV) was highest at 1 h post EP for all treatment groups but rapidly declined to levels no different from the control group at 24 h (Fig 2A). The area of intercellular gaps as a percentage of FOV was also greatest at 1 h post EP for all treatment groups, though it remained at greater levels for only the intermediate and high dose groups when compared with controls (Fig 2B). Likewise, the average size of intercellular gaps was greatest at 1 h post EP for all treatment groups, with large intercellular gaps persisting for up to 24 h post-EP for only the intermediate and high dose groups (Fig 2C). EP resulted in the immediate reduction in cell viability in all treatment groups when compared to controls; with time, cell population in the low and intermediate treatment groups returned to baseline levels within 24 h while viability remained low in cells treated at the high dose treatment setting (Fig 2D).

Figure 1.

Figure 1.

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Evolution of intercellular gaps in endothelial cell monolayers following sham treatment (A-C) or electroporation at low (D-F), intermediate (G-I), or high energy (J-L). Cell membrane and nuclei were stained and imaging with fluorescent microscopy was performed at 1, 8 and 24 h after treatment. Compared with controls, electroporation resulted in immediate cell swelling at low and intermediate energy treatments (D,G) which was followed by the complete recovery of the endothelial layer integrity in the low energy group (E,F) but not in the other treatment groups. Cell shrinkage from possible pyknosis and cell death can be observed in the intermediate and high energy groups (H,K).

Figure 2.

Figure 2.

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Characteristics and evolution of intercellular gaps in endothelial monolayers after electroporation. (A) The number of intercellular gaps apparent in a field of view following treatment at different energy settings and the closing of gaps over 24 h. (B) The average size of the intercellular gaps and change in this area over time. (C) The average diameter of intercellular gaps and (D) cell proliferation in treatment groups as compared with the control group at different timepoints post treatment. (* p < 0.05). ■- 1 hour, ■ - 8 hours, □ - 24 hours.

Taken together, these results indicate that depending on the energy setting used to perform EP, changes in cell diameter due to cell swelling or shrinkage as well as loss of cells due to death can contribute to the formation of intercellular gaps in endothelial monolayers. At the low dose setting, which was expected to largely result in RE of cells, intercellular gap formation was driven by cell swelling and the resultant loss of cell-cell contact. Intercellular gap size and area of gaps as a proportion of monolayer area remained small in the low dose group, where the presence of intercellular gaps was no different from the control group as early as 8 h post EP. At the intermediate and high dose setting, changes in cell diameter from both cell swelling and shrinkage could be observed, with the latter occurring in dying cells.

3.2. EP increases the transport of nanoparticles and small molecules through endothelial monolayers

We sought to understand the impact of post-EP intercellular gap formation on endothelial barrier function. We performed EP on EA.hy926 monolayers grown in transwell inserts and studied the transport of small molecule dye or SFB-IR783 in this in vitro model. Despite the manifestation of intercellular gaps as early as 1 h post EP in endothelial monolayers, there was no apparent difference in dye or nanoparticle transport through this barrier till 8 h post EP (Fig 3), and the difference in transport was obvious in only the cells treated with a high energy dose. Regarding the transport of the small molecule dye, while increased dye transport could be observed in the intermediate and high dose group from 24 h onwards (Fig 3A), there was no obvious difference in dye transport between the low dose group and the control group at all timepoints. Increased nanoparticle transport was obvious only in the high dose group at early timepoints, while the intermediate dose group demonstrated greater nanoparticle accumulation at 48 hours (Fig 3B). The group treated with low dose EP was no different from the control group in the assessment of the barrier function of endothelial cells to the passage of nanoparticles.

Figure 3.

Figure 3.

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Transport of material through endothelial monolayers in a transwell insert following electroporation. Electroporation increases the transfer of both (A) small molecule dye and (B) SFB-IR783, with increased transport apparent after 24 hours following treatment, mostly in the intermediate and high dose settings. ■ - Low, ▴- Intermediat ▾- High, ●- Control.

3.3. EP promotes the penetration and accumulation of nanoparticles and small molecules into tumorspheres

We performed EP on tumorspheres to determine its effect on the diffusion of molecules and nanoparticles through the extracellular space. The SFB-IR783 or Hoechst 33258 dye was introduced to the media containing tumorspheres at 10 min post EP to avoid direct transfection of cells while the membranes were in a permeabilized state. When compared with sham-treated samples, EP resulted in the immediate increase in the diffusion of both the Hoechst 33258 dye and SFB-IR783 nanoparticles in tumorspheres (Fig 4). Regarding the Hoechst 33258 dye, penetration of the dye was observed throughout the tumorsphere in both the intermediate (Fig 4C) and high dose groups (Fig 4D), while the dye uptake pattern appeared similar in the low dose group (Fig 4B) as compared with the control group (Fig 4A). Increased dye uptake compared with the control group was observed only in tumorspheres treated with intermediate or high dose EP (Fig 4J). Regarding the SFB-IR783 nanoparticles, uptake in tumorspheres treated with EP seemed largely at the periphery in the low dose group (Fig 4F), while the penetration of SFB-IR783 deep within the tumorsphere could be observed in the intermediate (Fig 4G) and high dose (Fig 4H) groups; meanwhile, uptake of SFB-IR783 within control samples (Fig 4E) was unremarkable. SFB-IR783 uptake in all EP-treated samples was greater than in controls (Fig 4I).

Figure 4.

Figure 4.

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The effect of electroporation on the penetration of a small molecule dye (B–D) and SFB-IR783 (F–H) into the tumorspheres following treatment with EP, compared with control samples (A,E). Treatment with EP results in multifold increase in the penetration of the small molecule dye (J) and SFB-IR783 (I). (* p<0.05)

We also measured post-EP change in the diameter of tumorspheres to determine the possible cytotoxic effects of EP. Tumorspheres treated with low (Fig 5A) or intermediate (Fig 5B) dose did not demonstrate any perceivable morphologic changes, appearing similar to control samples when observed with SEM at 24 or 48 h post EP. Tumorspheres treated with high dose EP (Fig 5C) lost their spherical morphology and appeared to disintegrate by 48 h, possibly due to IRE-related cell death. When compared with control samples, changes in the diameter of tumorspheres from baseline measurement were significant only in samples treated with high dose EP (Fig 5D).

Figure 5.

Figure 5.

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The effect of EP on tumorsphere viability. EP does not affect tumorsphere viability at low energy (A) but results in substantial reduction in the integrity of the tumorsphere at higher energy settings (B–C). The effect of electroporation on tumorsphere viability was assessed by measuring change in diameter over 48 h (D). ■ - Low, ▴- Intermediat, ▾- High, ●- Control.

3.4. EP increases the uptake and efficacy of sorafenib nanoparticles in cancer cells

We performed in vitro studies using HCT116 monolayers to assess nanoparticle uptake by cancer cells during EP. To isolate the cytotoxic effects of IRE from cell death resulting from sorafenib, cells were treated with low dose EP which induces RE without concomitant cell death. Treatment with low dose EP resulted in an immediate and increased uptake (Fig 6A) of SFB-IR783 when compared with untreated cells incubated with SFB-IR783 nanoparticles (Fig. 6D). However, at 24 h, no difference was observed in the levels of intracellular levels of SFB-IR783 nanoparticles between EP-treated and untreated cells that had not been treated but incubated with EP (Fig 6B). This confirms that EP-mediated nanoparticle uptake is immediate, rapidly increasing the concentration of the potential drug payload comparable to uptake over 24 h in cells not treated but incubated with EP. When compared with low dose EP or SFB-IR783 alone, the combination of low dose EP and SFB-IR783 resulted in reduced cell proliferation (Fig 6C) at 24 h post treatment.

Figure 6.

Figure 6.

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EP-mediated nanoparticle uptake in cancer cells from the disruption of plasma membrane function. EP results in the immediate and increased uptake of SFB-IR783 in cancer cells treated with EP (A), but the difference in overall uptake is negligible when assessed at 24 hours post EP (B). EP at low dose has minimal cytotoxic effect on cancer cells, and cell viability is reduced when EP is performed at higher energies (C). The uptake of SFB-IR783 (red) immediately following EP can be observed in most cells (nuclei stained blue) (D) while uptake at the same timepoint in control cells is limited (E). (*p < 0.05)

3.5. Intermediate dose EP promotes nanoparticle delivery to tumors and can augment the therapeutic efficacy of sorafenib nanoparticles

We further assessed the utility of intermediate dose EP in promoting nanoparticle delivery in vivo with mice bearing HCT116 tumors. In mice bearing bilateral tumors (Fig. 7E), the tumors treated with intermediate dose EP demonstrated greater SFB-IR783 uptake at 2 h post EP compared with the contralateral untreated tumors (Fig. 7A,B,F p = 0.0203). The difference in uptake did not persist when the mice were imaged at 24 h (Fig. 7C,D,F p = 0.1202), mirroring our results from the in vitro experiments with HCT116 monolayers. The comparison of EP-treated and untreated contralateral tumors at the 24 h timepoint with immunohistochemistry did not reveal any difference in levels of cell death as measured by Cleaved Caspase-3 (Supplemental Fig. 2 A-D; levels: EP: 3.2%, EP + NP: 2.97%, NP: 3.15%, and Control: 2.005% ) and proliferation (Ki-67 positive nuclei, Supplemental Fig. 2 E-H; levels: levels: EP: 80.88%, EP + NP: 81.51%, NP: 82.8%, and Control: 82.78%). This finding suggests that IRE-related cell death was limited in our experiment model.

Figure 7.

Figure 7.

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Imaging of SFB-IR783 uptake in mice bearing bilateral tumors. Imaging at 2 and 24 h post EP/administration of SFB-IR783 revealed immediate and increased uptake in the EP-treated tumor (A,C) when compared with the untreated side (B,D). (E) Tumor on the right flank was treated with EP while the contralateral side served as a sham control. (F) The difference in SFB-IR783 uptake was significantly greater in EP-treated tumors at 2 h but the difference was not observed when measurements were performed at 24 h, except in select animals (G).

In mice bearing unilateral tumors that underwent EP, immediate uptake of SFB-IR783 nanoparticles was demonstrated based on imaging measurements and was elevated compared with untreated mice (Fig. 8 A,B). Tumor growth kinetics and tumor volume were similar in mice assigned to sham control, EP only, and SFB-IR783 only. Mice treated with combined EP and SFB-IR783 demonstrated reduced tumor growth rate resulting in smaller tumor volumes compared with mice from the other cohorts (Fig. 8C). The delay in tumor growth was similar or overlapping for control, EP only, and SFB-IR783 only cohorts while the delay in tumor growth was marked immediately after treatment but reduced at latter timepoints in the combined EP and SFB-IR783 cohort (Fig. 8D).

Figure 8.

Figure 8.

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Anti-tumor effect of combined therapy with EP and SFB-IR783. Treatment with EP increased the uptake of SFB-IR783 (A) when compared with mice given the nanoparticle without EP (B). The combined treatment with EP and SFB-IR783 retarded tumor volume in comparison with treatment with EP alone, treatment with SFB-IR783 alone, or control (C) ). ■ - EP+NP, ▴ - Control, ▾ - EP, ● - NP, with marked delay in tumor growth at early timepoints which did not persist at later assessment periods (D), ■ - EP, ● - Control.

4. Discussion

Our results demonstrate that EP performed at an intermediate energy dose can rapidly increase the intra-tumoral uptake and improve the efficacy of sorafenib nanoparticles in a subcutaneous tumor model. Our in vitro experiments demonstrate that the effects of EP performed at an intermediate energy dose increases the extravasation of nanoparticles through endothelial layers, improves their diffusion through the extracellular space, and augments transmembrane transport of the drug payload into cancer cells (Schematic. 1). EP can therefore serve as means for the modulation of all three physiologic barriers that impede the intratumoral penetration of nanoparticles.

Schematic 1.

Schematic 1.

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Treatment with EP disrupts three main physiologic barriers (endothelial layer, extracellular space, and cell membrane), thereby enhancing the uptake of nanoparticle drugs in the tumor microenvironment.

The effect of EP on blood vessels has been studied extensively. Sersa et al. [19] first reported EP-related disruption in blood flow within tumors []over two decades ago, showing that blood flow was increased at low electric field strengths (< 640 V) compared with higher electric field strengths. In subsequent work, Kanthou et al. [25] identified the disruption of the endothelial cytoskeleton due to cell swelling as one of the mechanisms contributing to increased permeability, in their in vitro study using transwell insert based assays. Markelc et al. [26] and Bellard et al. [27] investigated this phenomenon in vivo on healthy tissue using a window chamber model and reported that EP induces an initial vascular lock which is followed by a short period of increased vascular permeability lasting for approximately 30–60 min. More recent mechanistic work by Markelc et al. [28] indicates a loss of gap junction contact in endothelial cells in vitro and in vivo, with in vivo vascular permeability being mediated by complex biological effects such as platelet aggregation. Our in vitro and in vivo findings are consistent with these prior results. We also note that our choice of intermediate energy dose (1000 V/cm) mirrors the range of electric pulse settings evaluated in these (1000 or 1300 V/cm) publications.

Electrochemotherapy, where EP is performed in the presence of drugs such as bleomycin or cisplatin, has strong vascular effects marked by cell death [29,30], acute loss of perfusion [31], and sustained reduction in blood flow [32], In a side-to-side comparison of normal and tumor blood vessels after electrochemotherapy, Markelc et al. [33] reported that the effects of electrochemotherapy on the loss of blood vessels were more pronounced than that of EP, with the loss of blood vessels occurring around 24 h post treatment. Interestingly, some of these findings were recapitulated in our in vitro experiments with the high dose setting (1500 V/cm) where we observed increased cell death in endothelial cells, possibly due to IRE, which impeded the recovery of the endothelial barrier function in this setting.

Fluorescein isothiocyanate (FITC)-labeled dextran at different molecular weights (70 or 2000 kDA) has been used to study post-EP permeability in endothelial monolayers seeded on transwell inserts [25] and in blood vessels in vivo [26-28], The size of FITC-labeled dextran at 10 nm (70 kDA) and 50 nm (2000 kDA) is smaller than the SFB-IR783 nanoparticles used in our experiments (80 nm). Clinically-used nanoparticle formulations of liposomal doxorubicin, albumin-bound paclitaxel, and gemcitabine are of a similar size range (80–120 nm). Therefore, our findings are relevant to EP-mediated nanoparticle delivery in the clinical setting. In both in vivo and in vitro studies, the increased transport of 70 kDA FITC-labeled dextran has been shown to be apparent immediately after pulse application whereas in our in vitro experiments the transport of both small molecules and SFB-IR783 was delayed. While we do not know the mechanism underlying this difference, several factors could have contributed to this discrepant finding, including the difference in the size of the nanoparticle used in the experiments (~10 vs 80 nm), the presence of a basement membrane in our transwell assay, and the lack of positive blood pressure that could propel nanoparticles through the vessel wall in vivo. Further, cell death due to IRE was an important contributing factor in increasing the permeability of endothelial monolayers in our studies, the effects of which can take a few hours post treatment to fully manifest.

Both RE and IRE have both been shown to alter the kinetics of nanoparticle uptake in tumors. RE has been shown to increase the uptake of liposomal doxorubicin [24], superparamagnetic iron oxide nanoparticles [34-36], while IRE has been shown to increase the uptake of gold [37] and polystyrene [38] nanoparticles. Cell membrane permeabilization during EP was primarily thought to drive nanoparticle uptake in these studies, leading to the close timing of nanoparticle administration and pulse delivery to maximize uptake in cells that are in a permeabilized state. Srimathveeravalli et al. [24] used a Zirconium-labeled liposomal nanoparticle to demonstrate that EP-related vascular changes can greatly increase nanoparticle uptake in tumors, an effect which was found to be independent of cell membrane permeabilization, suggesting that EP-related vascular changes by itself can promote nanoparticle delivery to tumors. Using magnetic resonance imaging, Zhang et al. [22-23] identified that IRE induces significant changes to tumor diffusion; however, the implications of improved tumor diffusion properties on nanoparticle uptake was not evaluated. Our in vitro tumorsphere experiments provide evidence of another mechanism through which EP can promote the delivery of nanoparticles into tumors, namely that EP can alter the diffusion of nanoparticles through the extracellular space. This finding is supported by prior studies evaluating small molecule therapeutics such as doxorubicin, cisplatin and bleomycin in combination with EP in the tumorsphere model, but here we report the first such finding with nanoparticle therapeutics.

While a number of studies have evaluated the impact of RE or IRE on nanoparticle uptake in cells, the efficacy derived from the combination of EP and nanoparticle-based therapy has not been fully evaluated. Our work extends these results by reporting the different mechanisms by which EP treatment can facilitate nanoparticle delivery and also by demonstrating that combination of EP with non-cytotoxic drugs, such as sorafenib, in a nanoparticle formulation can be an effective anti-cancer treatment. EP is already in clinical use, where image guidance is used to place needle electrodes into deep-seated tumors during bleomycin electrochemotherapy [39]. Correlative imaging studies have shown that EP-driven nanoparticles are predictable and can be monitored with positron emission tomography [31,39,40] or magnetic resonance imaging [37,38], thereby improving the ability to plan nanoparticle delivery and predict treatment response.

Our experiments were limited to a single tyrosine kinase inhibitor (sorafenib), a simple nanoparticle formulation, and a single cancer type (xenograft colorectal cancer). Electroporation-mediated delivery of liposomal doxorubicin has been reported under different experimental conductions in preclinical models of pancreatic cancer [31] and in murine rabbit carcinoma [39,40]. However, further investigations are required to understand how the selection of electroporation parameters would influence tumor uptake when using nanoparticles of varying size and composition. EP relies on the use of high voltage electric pulses which can be a contraindication to its use in patients with cardiac comorbidities or when treating tumors adjacent to electrically active tissue such as skeletal muscle.

5. Conclusion

Intermediate dose EP, as demonstrated in our experiments, can be performed rapidly, where pulse application between electrode pairs can be completed in less than 10 seconds, allowing the treatment of large tumors with multiple electrode pairs within few minutes. Following established clinical evidence with electrochemotherapy with bleomycin and cisplatin [39], EP-assisted nanoparticle delivery is especially well suited to promote drug uptake in locally advanced cancer and can be immediately translated to enhance the treatment of diseases such as pancreatic cancer.

Supplementary Material

1

Highlights.

  • Electroporation (EP) increases nanoparticle (NP) deposition in tumors.

  • EP augments NP transport through vasculature, extracellular space and cell membrane.

  • EP mediated delivery of sorafenib nanoparticle potentiates anti-cancer effect.

Acknowledgements :

The authors acknowledge the support of NIH Cancer Center Support Grant (P30 CA008748) for core laboratory services that were used for the presented work. G.S. acknowledges grant support from the SIR Foundation and the National Cancer Institute of the National Institutes of Health under Award Number U54CA137788/U54CA132378. The authors thank the support of Dr. James Chambers, and the Light Microscopy core in the Institute for Applied Life Sciences in University of Massachusetts at Amherst for their assistance with imaging.

Source of funding:

The authors acknowledge the support of NIH Cancer Center Support Grant (P30 CA008748) for core laboratory services that were used for the presented work. G.S. acknowledges grant support from the SIR Foundation and the National Cancer Institute of the National Institutes of Health under Award Number U54CA137788/U54CA132378. The authors thank the support of Dr. James Chambers, and the Light Microscopy core in the Institute for Applied Life Sciences in University of Massachusetts at Amherst for their assistance with imaging.

Footnotes

Disclosures

The authors report no relevant disclosures related to the work presented here. S.B.S is a consultant to BTG, Johnson & Johnson, XACT, Adgero, Innobaltive, and Medtronic. S.B.S has funding support from GE Healthcare, J&J Ethicon, Elesta and Angiodynamics, and holds stock in Aperture Medical. G.S. holds stock options in Aperture Medical.

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References

  • 1.Maeda Hiroshi, et al. "Tumor vascular permeability and the EPR effect in macromolecular therapeutics: a review." Journal of controlled release 651-2 (2000): 271–284. [DOI] [PubMed] [Google Scholar]
  • 2.Fang Jun, Nakamura Hideaki, and Maeda Hiroshi. "The EPR effect: unique features of tumor blood vessels for drug delivery, factors involved, and limitations and augmentation of the effect." Advanced drug delivery reviews 633 (2011): 136–151. [DOI] [PubMed] [Google Scholar]
  • 3.Taurin Sebastien, Nehoff Hayley, and Greish Khaled. "Anticancer nanomedicine and tumor vascular permeability; where is the missing link?." Journal of Controlled Release 1643 (2012): 265–275. [DOI] [PubMed] [Google Scholar]
  • 4.Torchilin Vladimir. "Tumor delivery of macromolecular drugs based on the EPR effect." Advanced drug delivery reviews 633 (2011): 131–135. [DOI] [PubMed] [Google Scholar]
  • 5.Brigger Irene, Dubernet Catherine, and Couvreur Patrick. "Nanoparticles in cancer therapy and diagnosis." Advanced drug delivery reviews 64 (2012): 24–36. [DOI] [PubMed] [Google Scholar]
  • 6.Chithrani B. Devika, Ghazani Arezou A., and Chan Warren CW. "Determining the size and shape dependence of gold nanoparticle uptake into mammalian cells." Nano letters 64 (2006): 662–668. [DOI] [PubMed] [Google Scholar]
  • 7.Perrault Steven D., et al. "Mediating tumor targeting efficiency of nanoparticles through design." Nano letters 95 (2009): 1909–1915. [DOI] [PubMed] [Google Scholar]
  • 8.Li Shyh-Dar, and Huang Leaf. "Stealth nanoparticles: high density but sheddable PEG is a key for tumor targeting." Journal of controlled release: official journal of the Controlled Release Society 1453 (2010): 178. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Albanese Alexandre, Tang Peter S., and Chan Warren CW. "The effect of nanoparticle size, shape, and surface chemistry on biological systems." Annual review of biomedical engineering 14 (2012): 1–16. [DOI] [PubMed] [Google Scholar]
  • 10.Detappe A, Kunjachan S, et al. Advanced multimodal nanoparticles delay tumor progression with clinical radiation therapy. J Control Release. 2016. September 28;238:103–113. [DOI] [PubMed] [Google Scholar]
  • 11.Chilkoti Ashutosh, et al. "Targeted drug delivery by thermally responsive polymers." Advanced drug delivery reviews 545 (2002): 613–630. [DOI] [PubMed] [Google Scholar]
  • 12.Gasselhuber Astrid, et al. "Mathematical spatio-temporal model of drug delivery from low temperature sensitive liposomes during radiofrequency tumour ablation." International Journal of Hyperthermia 265 (2010): 499–513. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Ren Y, Wang R, Gao L, Li K, et al. Sequential co-delivery of miR-21 inhibitor followed by burst release doxorubicin using NIR-responsive hollow gold nanoparticle to enhance anticancer efficacy. J Control Release. 2016. April 28;28:74–86. [DOI] [PubMed] [Google Scholar]
  • 14.Sano Kohei, et al. "Markedly enhanced permeability and retention effects induced by photo-immunotherapy of tumors." ACS nano 71 (2012): 717–724.23214407 [Google Scholar]
  • 15.Shamay Yosi, et al. "Light induced drug delivery into cancer cells." Biomaterials 325 (2011): 1377–1386. [DOI] [PubMed] [Google Scholar]
  • 16.Yarmush ML, Golberg A, Serša G, Kotnik T, Miklavčič D. Electroporation-based technologies for medicine: principles, applications, and challenges. Annu Rev Biomed Eng. 2014. July 11; 16:295–320. [DOI] [PubMed] [Google Scholar]
  • 17.Rosazza C, Meglic SH, Zumbusch A, Rols MP, Miklavcic D. Gene Electrotransfer: A Mechanistic Perspective. Curr Gene Ther. 2016;16(2):98–129. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Rubinsky Boris, Onik Gary, and Mikus Paul. "Irreversible electroporation: a new ablation modality—clinical implications." Technology in cancer research & treatment 61 (2007): 37–48. [DOI] [PubMed] [Google Scholar]
  • 19.Sersa G, Cemazar M, Parkins CS, Chaplin DJ. Tumour blood flow changes induced by application of electric pulses. Eur J Cancer. 1999. April;35(4):672–7. [DOI] [PubMed] [Google Scholar]
  • 20.Gehl Julie, Skovsgaard Torben, and Mir Lluis M.. "Vascular reactions to in vivo electroporation: characterization and consequences for drug and gene delivery." Biochimica et Biophysica Acta (BBA)-General Subjects 15691–3 (2002): 51–58. [DOI] [PubMed] [Google Scholar]
  • 21.Gibot L, Wasungu L, Teissié J, Rols MP. Antitumor drug delivery in multicellular spheroids by electropermeabilization. J Control Release. 2013. April 28;167(2): 138–47. [DOI] [PubMed] [Google Scholar]
  • 22.Zhang Z, Li W, Procissi D, Tyler P, Omary RA, Larson AC. Rapid dramatic alterations to the tumor microstructure in pancreatic cancer following irreversible electroporation ablation. Nanomedicine (Lond). 2014;9(8): 1181–92. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Figini M, Wang X, Lyu T, Su Z, Wang B, Sun C, Shangguan J, Pan L, Zhou K, Ma Q, Yaghmai V, Procissi D, Larson AC, Zhang Z. Diffusion MRI biomarkers predict the outcome of irreversible electroporation in a pancreatic tumor mouse model. Am J Cancer Res. 2018. August 1;8(8): 1615–1623. [PMC free article] [PubMed] [Google Scholar]
  • 24.Srimathveeravalli Govindarajan, et al. "Reversible Electroporation-Mediated Liposomal Doxorubicin Delivery to Tumors Can Be Monitored With 89Zr-Labeled Reporter Nanoparticles." Molecular imaging 17 (2018): 1536012117749726. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Kanthou C, Kranjc S, Sersa G, Tozer G, Zupanic A, Cemazar M. The endothelial cytoskeleton as a target of electroporation-based therapies. Mol Cancer Ther. 2006. December;5(12):3145–52. [DOI] [PubMed] [Google Scholar]
  • 26.Markelc B, Bellard E, Sersa G, Pelofy S, Teissie J, Coer A, Golzio M, Cemazar M. In vivo molecular imaging and histological analysis of changes induced by electric pulses used for plasmid DNA electrotransfer to the skin: a study in a dorsal window chamber in mice. J Membr Biol. 2012. September;245(9):545–54. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Bellard Elisabeth, et al. "Intravital microscopy at the single vessel level brings new insights of vascular modification mechanisms induced by electropermeabilization." Journal of controlled release 1633 (2012): 396–403. [DOI] [PubMed] [Google Scholar]
  • 28.Markelc Bostjan, et al. "Increased permeability of blood vessels after reversible electroporation is facilitated by alterations in endothelial cell-to-cell junctions." Journal of Controlled Release 276 (2018): 30–41. [DOI] [PubMed] [Google Scholar]
  • 29.Cemazar M, Parkins CS, Holder AL, Chaplin DJ, Tozer GM, Sersa G. Electroporation of human microvascular endothelial cells: evidence for an anti-vascular mechanism of electrochemotherapy. Br J Cancer. 2001. February;84(4):565–70. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Meulenberg Cécil JW, Todorovic Vesna, and Cemazar Maja."Differential cellular effects of electroporation and electrochemotherapy in monolayers of human microvascular endothelial cells." PLoS One 712 (2012): e52713. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Sersa G, Krzic M, Sentjurc M, Ivanusa T, Beravs K, Kotnik V, Coer A, Swartz HM, Cemazar M. Reduced blood flow and oxygenation in SA-1 tumours after electrochemotherapy with cisplatin. Br J Cancer. 2002. October 21;87(9):1047–54. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Sersa G, Jarm T, Kotnik T, Coer A, Podkrajsek M, Sentjurc M, Miklavcic D, Kadivec M, Kranjc S, Secerov A, Cemazar M. Vascular disrupting action of electroporation and electrochemotherapy with bleomycin in murine sarcoma. Br J Cancer. 2008. January 29;98(2):388–98. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Markelc B, Sersa G, Cemazar M. Differential mechanisms associated with vascular disrupting action of electrochemotherapy: intravital microscopy on the level of single normal and tumor blood vessels. PLoS One. 2013;8(3):e59557. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.West DL, White SB, Zhang Z, Larson AC, Omary RA. Assessment and optimization of electroporation-assisted tumoral nanoparticle uptake in a nude mouse model of pancreatic ductal adenocarcinoma. Int J Nanomedicine. 2014. September 1;9:4169–76. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Image-guided local delivery strategies enhance therapeutic nanoparticle uptake in solid tumors. Mouli SK, Tyler P, McDevitt JL, Eifler AC, Guo Y, Nicolai J, Lewandowski RJ, Li W, Procissi D, Ryu RK, Wang YA, Salem R, Larson AC, Omary RA. ACS Nano. 2013. September 24;7(9):7724–33. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Melancon MP, Appleton Figueira T, Fuentes DT, Tian L, Qiao Y, Gu J, Gagea M, Ensor JE, Muñoz NM, Maldonado KL, Dixon K, McWatters A, Mitchell J, McArthur M, Gupta S, Tam AL. Development of an Electroporation and Nanoparticle-based Therapeutic Platform for Bone Metastases. Radiology. 2018. January;286(1):149–157. [DOI] [PubMed] [Google Scholar]
  • 37.Tam AL, Melancon MP, Abdelsalam M, Figueira TA, Dixon K, McWatters A, Zhou M, Huang Q, Mawlawi O, Dunner K Jr, Li C, Gupta S. Imaging Intratumoral Nanoparticle Uptake After Combining Nanoembolization with Various Ablative Therapies in Hepatic VX2 Rabbit Tumors. J Biomed Nanotechnol. 2016. February;12(2):296–307. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Bulvik BE, Rozenblum N, Gourevich S, Ahmed M, Andriyanov AV, Galun E, Goldberg SN. Irreversible Electroporation versus Radiofrequency Ablation: A Comparison of Local and Systemic Effects in a Small-Animal Model. Radiology. 2016. August;280(2):413–24. [DOI] [PubMed] [Google Scholar]
  • 39.Boc N, Edhemovic I, Kos B, Music MM, Brecelj E, Trotovsek B, Bosnjak M, Djokic M, Miklavcic D, Cemazar M, Sersa G. Ultrasonographic changes in the liver tumors as indicators of adequate tumor coverage with electric field for effective electrochemotherapy. Radiol Oncol. 2018. October 18;52(4):383–391. [DOI] [PMC free article] [PubMed] [Google Scholar]

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