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Microfluidic electro-sonoporation: a multi-modal cell poration methodology through simultaneous application of electric field and ultrasonic wave3 Whitney Longsine-Parker,{a Han Wang,{a Chiwan Koo,b Jeongyun Kim,c Beomjoon Kim,d Arul Jayaramane and Arum Han*ab A microfluidic device that simultaneously applies the conditions required for microelectroporation and microsonoporation in a flow-through scheme toward high-efficiency and high-throughput molecular delivery into mammalian cells is presented. This multi-modal poration microdevice using simultaneous application of electric field and ultrasonic wave was realized by a three-dimensional (3D) microelectrode scheme where the electrodes function as both electroporation electrodes and cell flow channel so that acoustic wave can be applied perpendicular to the electric field simultaneously to cells flowing through the microfluidic channel. This 3D microelectrode configuration also allows a uniform electric field to be applied while making the device compatible with fluorescent microscopy. It is hypothesized that the simultaneous application of two different fields (electric field and acoustic wave) in perpendicular directions allows formation of transient pores along two axes of the cell membrane at reduced poration intensities, hence maximizing the delivery efficiency while minimizing cell death. The microfluidic electrosonoporation system was characterized by delivering small molecules into mammalian cells, and showed average poration efficiency of 95.6% and cell viability of 97.3%. This proof of concept result shows that by combining electroporation and sonoporation together, significant improvement in molecule delivery efficiency could be achieved while maintaining high cell viability compared to electroporation or

Received 2nd August 2012, Accepted 8th April 2013

sonoporation alone. The microfluidic electro-sonoporation device presented here is, to the best of our knowledge, the first multi-modal cell poration device using simultaneous application of electric field and ultrasonic wave. This new multi-modal cell poration strategy and system is expected to have broad

DOI: 10.1039/c3lc40877a www.rsc.org/loc

applications in delivery of small molecule therapeutics and ultimately in large molecule delivery such as gene transfection applications where high delivery efficiency and high viability are crucial.

1. Introduction Delivering genes and molecules such as probes and therapeutics into cells has immense applications in molecular biotechnology and gene therapy, though cell membranes act as an effective barrier between the cytoplasm and the extracellular environment.1,2 Various methods have been

a

Department of Electrical and Computer Engineering, Texas A&M University, College Station, TX, 77843, USA. E-mail: [email protected]; Fax: +1 979-845-6259; Tel: +1 979-845-9686 b Department of Biomedical Engineering, Texas A&M University, College Station, TX 77843, USA c Department of Nanobiomedical Science and WCU Research Center, Dankook University Graduate School, South Korea d CIRMM, Institute of Industrial Science, The University of Tokyo, Tokyo, Japan e Department of Chemical Engineering, Texas A&M University, College Station, TX 77843, USA 3 Electronic supplementary information (ESI) available. See DOI: 10.1039/ c3lc40877a { These authors contributed equally to the paper.

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developed to make the membrane temporarily permeable for the introduction of foreign substances into cells. The introduction of genetic materials and constructs, a process commonly referred to as transfection, is of particular interest for applications in genetic engineering and gene therapy.2,3 Examples of transfection methods include viral vector methods, chemical-based methods, and physical methods such as particle bombardment (gene gun), microneedle injection, as well as methods that create transient pores in the cell membrane, such as electroporation and sonoporation.4 Viral vector methods are widely used due to their specific delivery capabilities and high efficiency, but there are potential complications such as mutagenesis and inherent risks that may incite an immune response.5 Chemical-based methods have limited efficiency in general, and certain physical methods like direct bombardment and injection are not selective and may permanently disrupt cell function.4 Among the physical methods, both electroporation and sonoporation methods are widely used to allow foreign materials such as

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Lab on a Chip dyes, drugs, and genetic materials to transverse into the cells. Electroporation is the use of an externally applied electric field to form transient pores in the cell membrane to deliver genes and drugs into cells, both in vitro and in vivo.4,6–8 Rols et al. demonstrated that the creation of transient pores in cell membrane requires two criteria: the applied electrical field intensity needs to be greater than a threshold to induce transient permeated structures, and the number and duration of the electrical pulses need to be sufficient for expansion of these structures.9 Short pulses (ms–ms) of high electric fields (1–10 kV cm21) are typically used to break down the cell membrane and form transmembrane pores.1,4,10,11 Permeabilization is reversible up to a certain limit, beyond which the electric field exposure compromises the viability of the cells.12 Bench-top electroporators can process large numbers of cells rapidly, however drawbacks include the large amount of reagent consumed, rising ambient temperature over time that reduces cell viability, and limited transfection efficiency caused by cells being exposed to non-uniform electric field.3,4,7,13,14 In the past decade, extensive efforts have been made in developing various forms of microchip electroporation devices to overcome these limitations.7,15,16 Using microfluidic channels and integrated microelectrodes, high electric fields sufficient for electroporation could be achieved with a supply voltage of only 0.1–20 V.11,15,16 The low voltage and rapid heat dissipation in microchips with continuous flow also helped diminish heating issues, which improved the survival rate after treatment.4,11 Overall, significant improvement in efficiency and cell viability have been achieved through microchip electroporators. Single-cell electroporation methods have been also extensively investigated to achieve very high poration rates, with the capability to directly monitor the poration procedures.1,2,13,17–19 Sonoporation, another physical method, is the use of ultrasonic wave to create transient permeability in cell membranes and, like electroporation, is effective both in vitro and in vivo.20 Acoustic cavitation is believed to be the primary mechanism for cell membrane disruption, and relies on dissolved air and/or solution impurities as cavitation nuclei.20,21 Cavitation can be enhanced by the introduction of microbubbles known as ultrasound contrast agents (UCA), which greatly reduce the cavitation threshold.21 However, UCA has also been found to compromise cell viability in sonoporation at high excitation.22,23 Similar to electroporation, the membrane permeabilization is not permanent, however increased exposure dose to improve efficiency also leads to lower cell viability.20,21,24 Cell damage and death is typically far less extensive than with electroporation, however the overall transfection efficiency in vitro is lower than electroporation, with efficiencies and viabilities varying widely in different reports.20,25–29 This is likely due to the non-uniform acoustic field that cells are exposed to in a cuvette-like suspension. If cells can be exposed to a uniform acoustic field by being positioned in the same focal plane, significant improvement in both efficiency and viability may be achieved.


Paper Microfluidic chambers combined with laser-induced cavitation have been utilized for controlled cavitation and improved efficiency and viability in sonoporation.30 However the laser setup is costly and difficult to integrate with other postexposure applications. Rodamporn et al. demonstrated UCAfree sonoporation in a microfluidic chamber with a single piezoelectric ceramic, which utilized a microfluidic platform to greatly reduce transducer-to-reflector spacing and thus generate a highly uniform acoustic field at resonance frequency.31 However such static chamber configuration limits the throughput of cell transfection. Recently Carugo et al. developed a flow-through type UCA-free ultrasound-microfluidic chip for continuous single-cell sonoporation using ultrasonic standing wave.23 Although many of these electroporation and sonoporation methods are currently widely used, there is a trade-off between membrane permeabilization efficiency and cell viability. Methods that can achieve high efficiency transfection with extremely low cell death will have significant impact to this field. Based on the extensive success in both the fields of electroporation and sonoporation, investigation of the combined effect of these methods is a natural progression, however, very little has been reported and no microfluidicsbased devices exist. Yamashita et al. presented in vivo transfection into the quadriceps of mice via a plasmid DNA injection followed by sonoporation for 2.5 min and simultaneous electro-sonoporation for another 2.5 min, increasing luciferase activity in the muscle two-fold compared to electroporation alone.32 Escoffre et al. presented a sequential electro-sonoporation method in vitro, first electroporating CHO cells using a conventional-type setup with plasmid DNA encoding enhanced green fluorescent protein (EGFP) and then sonoporating the cells in the presence of microbubbles.22 In this sequential treatment of electroporation and sonoporation, optimized conditions for each modality were based on maximum transfection efficiency with minimum fatality separately, resulting in increased overall transfection efficiency. However the exposure of cells to two fields at high strength probably resulted in low cell viability. Also, the combined effect may have been limited by the long time interval between the two modalities since relaxation of the transient pores is spontaneous.9 Our hypothesis is that simultaneously applying electric field and acoustic wave in perpendicular directions to cells will result in highly efficient delivery of molecules by maximizing total pore formation while minimizing cell death by limiting the degree of pore formation (i.e. overexposure) for a given cell membrane area. The objective of our presented work is to demonstrate the proof of concept that such a multi-modal electro-sonoporation scheme can be advantageous compared to both microchip-based electroporation and sonoporation methods.

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Fig. 1 Schematic showing the concept of the microfluidic electro-sonoporation device. (a) Cells flowing through a microfluidic channel made of two opposing 3D microelectrodes are exposed to electric field generated between the microelectrode pair. An ultrasonic transducer placed on top of the channel couples ultrasonic wave through the PDMS layer to cells flowing through the microfluidic channel. (b) A single cell being exposed simultaneously to electric field and ultrasonic wave while flowing through the microfluidic channel.

2. Materials and methods 2.1. Design of the microfluidic 3D electrode electrosonoporation device The microfluidic system developed here enables flow-through microchip electroporation utilizing a novel 3D microelectrode configuration where the microelectrode serves both as the electric field generator as well as the microchannel itself, and flow-through sonoporation, simultaneously. It is to be noted that we have purposely selected a simple straight-channel electroporation microchip design without using more elaborate microfluidic electroporation device designs demonstrated by others for higher efficiency,33–36 so that the comparison between the proposed electro-sonoporation method to electroporation or sonoporation only is easier and fair. Fig. 1 shows the overall design concept and device configuration. The device consists of two layers, the 3D electrode/channel layer and the poly(dimethylsiloxane) (PDMS) polymer channel/cover layer (Fig. 2). The electrode layer on a glass substrate consists of two parallel 3D electrodes

Lab on a Chip 30 mm high and separated by a 50 mm gap. These two electroporation electrodes also function as cell flow microchannels, where cells can be guided to flow between the two electrodes while being exposed to a uniform electric field generated between the electrodes. The inlet area of the 3D electrode/channel structure was shaped as a funnel so that cells can be more easily guided into the microchannel. Typically, 2D planar electrode pairs are not efficient for electroporation due to the non-uniformity and shallow penetration depth of electric field into a microfluidic channel. A top-bottom electrode pair scheme provides uniform electric field exposure, however requires a transparent electrode material such as indium-tin-oxide (ITO) to be compatible with optical microscopy for real-time monitoring. More importantly, such design is not compatible with combined sonoporation due to obstruction of ultrasonic wave coupling into the microfluidic channel as required here. The channel structure in the polymer layer was generally shaped as an inverse replica of the 3D electrode/channel structure, but with slightly larger dimension so that once flipped and assembled on top of the 3D electrode/channel layer, it could incorporate the entire 3D electrode/channel structure. The layer-to-layer assembly of the 3D electrode design relied primarily on the strong bonding between PDMSto-glass, rather than bonding the PDMS layer on top of the electroplated 3D electrode, where surface roughness can result in fluidic leakage. The PDMS layer had three inlets, a center cell inlet and two side sheath flow inlets, so that cell-flow can be focused into the inlet of the 3D electrode/channel structure with high efficiency utilizing microfluidic flow focusing. Three outlets were incorporated, a center cell collection outlet that collects cells undergoing electro-sonoporation, and two side waste outlets collecting buffer and any unfocused, hence untreated, cells. The bottom glass substrate made the device compatible with light/fluorescent microscopy. The top polymer layer allowed easy coupling of ultrasonic waves from the acoustic transducer placed directly on top of the PDMS channel layer, so that electroporation and sonoporation conditions could be simultaneously applied to cells guided and flowing between the 3D electrodes. The electric field and acoustic field inside the microfluidic channel structure were simulated using finite element method (FEM)-based models from a commercially available FEM software (COMSOL Multiphysics1, COMSOL Inc., Los Angeles, CA, USA). Electric field and acoustic pressure field at the center part of the microfluidic channel where cells are

Fig. 2 Schematic and fabrication steps of the two-layer microfluidic electro-sonoporation device. (a) The glass/electrode layer serves as optically transparent substrate and channel floor, while electroplated electrodes act as channel sidewalls and electroporation actuators. (b) The PDMS layer contains a chamber serving as channel ceiling and outer boundaries of cell flow; optical transparency allows viewing of cell flow in the chip. Ultrasonic wave from a piezoelectric transducer (not shown in this image) is directly coupled from the top side through the PDMS channel layer. (c) Assembled device showing cell flow. (d) Fabricated device.

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2.2. Fabrication The 3D electrodes were patterned on glass slides (5.08 cm 6 7.62 cm) by electroplating. Starting from an evaporated seed layer of Au/Cr (tAU = 2000 Å, tCR = 200 Å), the electroplating molds for the 3D electrodes were defined by photolithography (Futurrex NR2-20000P, Futurrex, Inc., Franklin, NJ, USA), and then nickel electrodes were electroplated (Techni Nickel Sulfamate RTU, Technic Inc., Cranston, RI, USA) to a thickness of 30 mm. The photoresist masking layer and the Au/Cr seed layer were removed following electroplating. The PDMS layer (Sylgard1 184, Dow Corning Corp., Midland, MI, USA) was fabricated with soft lithography using a patterned SU-8 master mold. Since the bonding between PDMS and nickel electrodes is poor due to the surface roughness of electroplated nickel, the PDMS channel depth was intentionally fabricated to be 5 mm lower than the electrode height to achieve tight sealing. Fluidic inlet and outlet holes, as well as holes to expose the electrical contact pads, were formed using punch tools. The PDMS slab was then bonded to the glass substrate patterned with nickel electrodes by O2 plasma treatment (Expanded Plasma Cleaner, Harrick Plasma, Ithaca, NY, USA). Electrical connections were provided via wires soldered to the contact pads built into the electrode design and uncured PDMS was used to both secure the contact wires in place and to provide a fluidic seal to the outside edges of the electroplated electrodes. 2.3. Device preparation for electro-sonoporation experiment Prior to each cell experiment, the microfluidic channel was treated with a series of surface treatment steps to avoid cell adhesion to the channel surfaces. First, deionized (DI) water was flowed at a rate of 0.2 ml h21 for 10 min to rinse the microfluidic channel. The channel was then coated with a bovine serum albumin (BSA) solution to prevent cell adsorption to the Ni plated sidewalls. BSA was dissolved in phosphate buffered saline (PBS) to a 5% concentration and flowed in the channel for 10 min at a rate of 0.2 ml h21 and then left to sit for 1 h. Fresh BSA solution was injected for an additional 10 min and then PBS was used for a final rinsing step for 10–20 min at 0.2 ml h21. 2.4. Cell preparation Human cervical cancer (HeLa) cells (ATCC; American Type Culture Collection, Manassas, VA, USA) were grown in Dulbecco’s Modified Eagle Medium (DMEM, Gibco1, Invitrogen, Carlsbad, CA, USA) supplemented with 10% bovine serum (BS, Invitrogen, Carlsbad, CA, USA) to 90% confluency in a 37 uC, 5% CO2 incubator. Prior to electro-sonoporation experiments, cells were harvested from the culture flask using trypsin-EDTA (0.25% with EDTA 4Na, Invitrogen, Carlsbad, CA, USA), centrifuged, and resuspended in 2 ml of serum-free DMEM medium. A hemacytometer was used to verify the target cell density of approximately 2 6 106 cells ml21.


Paper 2.5. Cell poration and staining In this study, propidium iodide (PI, 668.4 Da) was used to simulate small molecule delivery into cells due to its wide acceptance as a marker for cell membrane permeability, as well as easy visualization for evaluating poration efficiency.12,37,38 PI has similar molecular weight compared to many small molecule therapeutic agents such as doxorubicin (543.52 Da) and apigenin (270.24 Da).23 Permeabilization of porated cells was evaluated by the penetration of PI and cell viability was evaluated by Calcein AM staining post treatment, a standard procedure in poration characterization.6,34 Viable cells with intact membranes exclude PI, whereas membranes of dead and damaged cells as well as porated membranes are permeable to PI. On the contrary, Calcein AM is membrane permeable, but only live cells can provide active esterases to cleave acetomethoxy for Calcein AM to be fluorescent. Therefore, only cells that are viable and porated will show fluorescence of both PI and Calcein AM. PI (Invitrogen, Carlsbad, CA, USA) was added to resuspended cells in serum-free DMEM at 1 : 20 (vDYE : vMED) ratio prior to treatment, for a final concentration of 50 mg ml21. Calcein AM (Invitrogen, Carlsbad, CA, USA) staining on the other hand, was performed at 1 : 1000 (vDYE : vMED) ratio 20 min post-treatment, for a final concentration of 5 mM, followed by incubation at 37 uC, 5% CO2 for 30 min. It has been reported that the reversible transient pores in cell membrane would reseal within 20 min after poration,12,18,39 therefore the added PI would only present in porated or non-viable cells. 2.6. Microchip electroporation protocol In all experiments, microelectroporation, microsonoporation, and combined microelectro-sonoporation were performed in series for a total of 5 repetitions. Individual devices could be used for multiple experiments after rinsing with isopropyl alcohol and DI water. To achieve the desired E-field range for HeLa cell electroporation (0.3–2.3 kV cm21)37,40,41 and help prevent cell clogging in the channel, the distance between the electrodes was designed to be 50 mm. A function generator (33220A 20 MHz, Agilent Technologies, Santa Clara, CA, USA) was used to apply an AC square wave with a peak-to-peak amplitude of 5 V to 8 V and a frequency of 1 kHz or 100 kHz. The duty cycle was 50% in all experiments. These conditions were selected based on the required electroporation conditions and the electrolysis threshold of PBS buffer (16, pH = 7.4, Invitrogen, Carlsbad, CA, USA). The cell solution was loaded into a 1 ml syringe and driven by a syringe pump (KDS100, KD Scientific Inc., Holliston, MA, USA) at a flow rate of 0.05–0.1 ml h21. At this flow rate range, cells were exposed to electric field for 1.46–0.73 s. Microfluidic flow focusing using sheath flow was utilized to focus the cell flow to the middle of the channel so that all cells are exposed to the same electric field. Cells exposed to the electric field were collected at the outlet. The collection time required for each sample of 10 ml varied with the selected flow rate, but was typically within 5 min.

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2.7. Microchip sonoporation protocol A 1 MHz piezoelectric immersion transducer (TSI–I–#300, Techno Scientific Inc., Concord, ON, CA) was driven by a function generator via a power amplifier. An immersion style transducer was chosen because the acoustic impedance of PDMS (1–1.9 MRayl) is close to that of water (y1.5 MRayl), and also close to that of biological tissue (1.3–1.7 MRayl).42,43 An impedance matching gel (Aquasonic 100 Ultrasound Transmission Gel, Parker Laboratories, Inc., Fairfield, NJ, USA) was applied at the interface of the transducer and the PDMS device. The function generator (33220A 20 MHz, Agilent Technologies, Santa Clara, CA, USA) supplied 1 MHz sine wave with a peak-to-peak amplitude of 100 mVpp. The signal from the function generator was amplified by a 50 dB power amplifier (E&I 2100L RF Amplifier, Electronics & Innovation Ltd, Rochester, NY, USA). The maximum supply power was limited to 100 mVpp from function generator to avoid overheating of the transducer and thus to improve cell survival rates. Microsonoporation was conducted in the same device after microelectroporation. With a cell flow rate of 0.05 ml h21 (with 0.1 ml h21 total sheath flow), the ultrasound wave was applied through the impedance matching gel continuously for 5 min to collect 10 ml of sample. The transducer area covers the entire flow channel and funnel area, resulting in exposure time of 1.5 s for an individual cell. As mentioned in the introduction, reported sonoporation data often involves the incorporation of microbubbles or UCA to enhance cavitation and in turn, improve transfection efficiency. However, Yamashita et al. reported that in conventional electro-sonoporation using microbubbles, significant increase in the resistance to electric pulses was observed, thus limiting electroporation.32 Carugo et al. also reported significantly reduced cell viability due to UCA usage, and demonstrated that it is also possible to achieve high transfection efficiency without UCA.23 Therefore, microbubbles or UCA were not used in our experiments. 2.8. Combined microelectro-sonoporation Microelectro-sonoporation was performed at the same flow conditions (0.05 ml h21) and by applying both electric field and acoustic wave simultaneously using the same parameters as described above. Besides the three treatment groups, a negative control group without any microelectroporation nor microsonoporation was tested by following the same cell preparation and flow procedures but without any electric field or ultrasonic wave. 2.9. Cell imaging and analysis Treated cells, following incubation, were diluted and plated onto microslides for imaging with an inverted fluorescent microscope (Eclipse TS100, Nikon Instruments Inc., Melville, NY, USA) to analyze successful poration through PI incorporation into cells (red fluorescence) and cell viability through Calcein AM staining (green fluorescence). Thus, cells that exhibit both red and green fluorescence are deemed to be successfully porated and viable. Cells that are only fluorescing green indicate that these cells are alive but without successful

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Lab on a Chip poration, whereas cells that are only fluorescing red indicate that these cells are either dead from the beginning or died during the poration process. Small cell debris also show red fluorescence but is not counted. The criteria in determining a cell being fluorescent are 1) at least 50% of the cell area needs to show fluorescence (see Supplementary Fig. S2(a), ESI3 where the marked cell is NOT counted as fluorescence) and 2) any cell that is showing significantly weaker fluorescence (less than 40% mean intensity) is NOT counted as fluorescence (see Supplementary Fig. S2(b), ESI3). The number of transfected cells were counted and compared against the total number of cells to determine the efficiency of the poration methods.

3. Results & discussion 3.1. 3D electrode device testing A photograph of a successfully fabricated electro-sonoporation microchip, with all electrical and fluidic connection ports, is shown in Fig. 2(d). The funnel-shaped channel inlet and laminar flow based flow-focusing scheme employed here successfully guided the majority of incoming cells between the 3D electrode-formed channels. This scheme was critical to prevent incoming cells from partially going into the two side waste channels. The simulated electric field of the 3D electrodes as shown in Supplementary Fig. S1, ESI3 showed uniform field intensity along the microchannel, which guaranteed uniform and continuous exposure of cells to the electric field. The device was first tested to determine the maximum electric pulse condition applicable without bubble formation due to electrolysis of the buffer. At a frequency of 1 kHz, 5.2 Vpp was determined to be the upper limit. When the frequency was increased to 100 kHz, bubble generation was not observed with an applied voltage as high as 8 Vpp, consistent with previous reports. For example, Ziv et al. demonstrated that bubble generation, or electrolysis, can be reduced by using an AC source and is frequency dependent (higher frequencies are preferred).10 Fox et al. reported that electrolysis can be further reduced by applying bipolar pulses, making the time-averaged current zero.11 Therefore, the applied voltage was kept at 8 Vpp at 100 kHz in all microelectroporation and microelectrosonoporation experiments, which translates to 1600 V cm21. This value is well within the range of previously reported HeLa cell electroporation conditions.37,38,40,41,44,45 3.2. Microelectroporation Fig. 3 shows example images of HeLa cells that were collected at the outlet of the device without electroporation or sonoporation (Fig. 3(a), negative control), with electroporation only (Fig. 3(b)), with sonoporation only (Fig. 3(c)), and with electro-sonoporation (Fig. 3(d)). The experimental results shown in Fig. 3(a–d) are from a single batch of cells and were performed in immediate succession of one another. Note that the images here show the collected cells diluted to low concentration for easy single-cell visualization and analysis on a glass cover slide. Additional cell images, including higher


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Paper cence. For microelectroporation, the average poration efficiency through five different experiments (total of 1687 cells analyzed) was 77.8% ¡ 3.9% (nEXP = 5), and the average viability of these experiments was 89.3% ¡ 4.3% (nEXP = 5). 3.3. Microsonoporation


Microsonoporation experiments were performed in the same devices right after the microelectroporation experiment. Fig. 3(c) shows example images of a successful sonoporation experiment, where cells showing up as both green and red indicate successfully porated cells while still being viable. The average poration efficiency of microsonoporation through five different experiments (total of 1710 cells analyzed) was 84.9% ¡ 4.1% (nEXP = 5), and the average viability of these experiments was 97.1% ¡ 2.3% (nEXP = 5). The higher poration efficiency of microsonoporation compared to microelectroporation can be attributed to the higher cell viability of microsonoporation, since the efficiency is calculated as percentage of viable transfected cells out of total number of cells. In fact, the ratio of transfected cells to live cells in microsonoporation (87.4%) was almost equal to that of microelectroporation (87.1%). 3.4. Combined microelectro-sonoporation

Fig. 3 (a)–(d) Images of cells from a single batch of experiment. Left: brightfield images, middle: fluorescent images of live cells stained by Calcein AM, right: fluorescent images of porated or non-viable cells stained by PI. (a) Negative control, nCELLS = 599. (b) Microelectroporation: Efficiency = 77.8%, Viability = 89.3%, nCELLS = 1687. (c) Microsonoporation: Efficiency = 84.9%, Viability = 97.1%, nCELLS = 1710. (d) Microelectro-sonoporation: Efficiency = 95.6%, Viability = 97.3%, nCELLS = 1249. (e) Comparison of average poration efficiency (red) and cell viability (blue). Error bar indicates standard deviation. *p , 0.05, nEXP = 5. Scale bar = 200 mm.

magnification images, from three different repetitions for each condition are included as Supplementary Fig. S3., ESI3. In the particular batch shown in Fig. 3(b), 100% of cells were viable and 96.8% of cells were both viable and successfully porated, evident from cells showing up as both green (Calcein AM staining) and red (propidium iodide staining) fluores-


Microchip electro-sonoporation experiments were performed in the same devices right after the microsonoporation experiment. Fig. 3(d) shows example images of a successful electro-sonoporation experiment. The average poration efficiency of microchip electro-sonoporation through five different experiments (total of 1249 cells analyzed) was 95.6% ¡ 2.1% (nEXP = 5), and the average viability of microchip electrosonoporation was 97.3% ¡ 1.8% (nEXP = 5). The efficiency of electro-sonoporation was 11% greater (or 13% improvement) than that of sonoporation alone (95.6% vs. 84.9%) and 18% greater (or 23% improvement) than that of electroporation alone (95.6% vs. 77.8%). Furthermore, the cell viability after electro-sonoporation was higher compared to electroporation (97.3% vs. 89.3%) and same as sonoporation (97.3% vs. 97.1%), showing that combining two different physical transfection modalities could result in increased transfection efficiency while showing slightly higher or similar cell viability level compared to using only one modality. Fig. 3(e) shows average and standard deviation of poration efficiency and viability from the five batches of experiments conducted. Student’s T-test result shows that combined microelectro-sonoporation shows statistically higher (defined as p , 0.05) efficiency compared to microelectroporation and microsonoporation. When comparing cell viability, microelectro-sonoporation showed statistically significant improvement compared to microelectroporation (p , 0.05) but insignificant change compared to microsonoporation (p = 0.8938). Cells showing PI incorporation in negative control was 1.7% with cell viability of 98.5% (nCELLS = 599). This indicates that spontaneous incorporation of PI is negligible, thus PI can be used as an indicator for molecular delivery into cells. Additional images showing cells in negative control as well as with electroporation, sonoporation, and combined electrosonoporation at 106 and 206 magnifications are shown in Supplementary Fig. S3, ESI3. The throughput of the device was

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Table 1 HeLa cell electroporation efficiencies and viabilities (%)

Methods

Researcher

Efficiency

Viability

Electroporation

Glahder et al.b 38 Kim et al.b 45 Rodamporn et al.c He et al.c 46 Kim et al.b 45 This Workc This Workc

40%a 40%a 48.74% 75%a 85%a 77.8% 95.6%

25%a 67%a — 80%a 77%a 89.3% 97.3%

Microelectroporation Electro-sonoporation

40


a

Percentages not mentioned explicitly in text, values est. from graphs. b EGFP as transfection indicator. c PI as permeabilization indicator.

2 6 105 cells h21, demonstrating that this highly efficient and highly viable method can be achieved in a high-throughput manner.

4. Discussion In order to effectively compare our microfluidic electrosonoporation system to other previously reported methods, existing literature was reviewed for reported HeLa cell transfection efficiencies and viabilities, and a detailed discussion is given in the Supplementary Material.3 In fact, a primary reason for selecting HeLa cells for our experiments was the availability of several reported electroporation and sonoporation results to compare our findings against. A summary of the transfection efficiency and cell viability data for HeLa cell electroporation and sonoporation is shown in Table 1 and Table 2, respectively. Our microelectroporation result shows similar transfection efficiency while maintaining higher cell viability compared to other microchip electroporation results, which can be attributed to the flow focusing of cells and 3D electrodes construction so that all cells are exposed to a uniform electric field and maximum heat dissipation due to the continuous flow. Our microsonoporation result shows much higher efficiency and viability compared to others. This can be attributed to the fact that all cells are on the same plane and thus exposed to a uniform acoustic field (Supplementary Fig. S1, ESI3). This result is similar in concept with the recent

Table 2 HeLa cell sonoporation efficiencies and viabilities (%)

Methods

Researcher

Efficiency

Viability

Sonoporation

Feril et al.b 47 Chen et al.b 48 Lai et al.c 49 Rodamporn et al.b This Workc This Workc

16.1% 26% 35%a 68.9% 87.9% 95.6%

80%a 50–60% 45%a 77% 97.1% 97.3%

Microsonoporation Electro-sonoporation a

31

Percentages not mentioned explicitly in text, values est. from graphs. b EGFP as transfection indicator. c PI as permeabilization indicator.

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finding by Rodamporn et al.31 and Carugo et al.23 that higher cell viability and efficiency can be achieved using ultrasonic standing wave (USW) that can better control the acoustic radiation strength applied to cells. The combined simultaneous microelectro-sonoporation result from our device clearly shows that combining two modalities simultaneously can result in higher efficiency while maintaining high viability. The underlying mechanism of this synergistic behavior needs further investigation, however we hypothesize that applying relatively low stimulation conditions for electroporation and sonoporation which does not irreversibly damage cells (i.e. high cell viability) can still result in high transfection efficiency due to the larger degree of pore formation per cell. Though various models have been proposed for electroporation, it is now widely accepted that the transient pore formation can be divided into two phases: (1) induction of transient permeated structures when electric field is stronger than a threshold, and (2) expansion of such permeated structure to form transient pores. The expansion step depends on pulse number and duration, and occurs in regions which satisfies E?cos(h) . ES

(1)

where E is the electric field intensity, ES is the threshold value to induce cell membrane permeability, and h is the polar angle in reference to the applied electric field.8,9,50,51 Kim et al. recently developed a microchannel single cell electroporation device for direct visualization of molecular uptake direction using fluorescent dyes.52 This work showed in real time that molecular uptake by electroporation occurs in a cone shaped region in reference to the applied electric field, and the polarity depends on the charge property of delivered molecules. To the best of our knowledge, there is no literature on the directionality of transient pore formation in sonoporation. However we assume the pore formation in sonoporation follows the same pattern as in electroporation where pores are mostly formed along the acoustic field direction. Thus by combing the two modalities simultaneously in perpendicular directions, transient pore formation can be found in directions of both electric field and acoustic field, which can significantly increase the degree of pore formation (e.g. pore numbers) per cell under the same intensities than only one modality applied. However since the degree of pore formation per cell surface area is still sub-threshold due to pores from electroporation and pores from sonoporation forming on different parts of the cells, this will result in high efficiency while maintaining high cell viability. In this study we have demonstrated the capability of the developed microchip electro-sonoporation system in successfully delivering small molecules. Future studies will be to test this multimodal poration strategy for delivery of larger genetic molecule such as siRNA (y6 kDa) or even DNA-plasmid (y20 kDa) into mammalian cells. These results, especially in terms of cell viability, clearly show that combining electroporation and sonoporation in a


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microscale flow-through configuration could be a new modality for high efficiency, high viability transfection of cells. We expect that further optimization of experimental protocols, utilizing some of the latest microelectroporation configurations that show higher efficiency compared to a straight-channel design as used here, further characterization beyond cellular uptake of dyes such as gene expression, and long-term viability studies will make this new multi-modal method a next-generation tool in molecular biotechnology.

5. Conclusion We present a microfluidic device for combined microelectroporation and microsonoporation to improve overall transfection efficiency for mammalian cells with high cell viability. The flow-through type microfluidic platform utilizes lateral 3D electrodes that function as both the electric field applying electrodes and as a cell-flowing channel structure. This novel design enables uniform electric field application to cells flowing through the metal channels as well as direct coupling of ultrasonic waves generated from a piezoelectric transducer applied at the topside through an acoustic impedance matching layer. With this device, cells flowing through the microchannel can be simultaneously exposed to electric field and ultrasonic wave in perpendicular directions for combined microscale electro-sonoporation. Results from our microfluidic electro-sonoporation showed higher transfection efficiency compared to microelectroporation alone or microsonoporation alone, with cell viability higher than 90%. While others have reported electro-sonoporation protocols, to the best of our knowledge, we are the first to incorporate both electroporation and sonoporation on a chip format and to perform simultaneous poration. Overall, we have presented a proof-ofconcept device to combine microelectroporation and microsonoporation in a microfluidic flow-through format that, in its current form, meets or outperforms the transfection efficiency and cell viability standards set by other reported electroporation and sonoporation methods.

Acknowledgements AH would like to thank the Institute of Industrial Science (IIS) of the University of Tokyo for their support through the visiting scholar program. This project was partially funded by the Korean Ministry of Knowledge Economy grant #10039890 to AH.

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