Electrical characterization of a single cell electroporation biochip with the 2-D scanning vibrating electrode technology
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Advancements in microfabrication technology have lead to the development of planar micro-pore electroporation technology. This technology has been shown to provide greater control in single cell manipulation, and electroporation which is independent from cell size. In this work we report direct and spatially resolved characterization of electric currents within a planar micropore electroporation biochip to better understand this phenomenon at the cellular level. This work was performed using a two-dimensional (2-D) vibrating probe (VP). Analysis of the spatial patterns of current density yielded a 4th order polynomial profile in the planes parallel to the biochip’s surface and a three parameter hyperbolic decay profile in the planes perpendicular to the chip surface. A finite element model was developed which correlates with actual measurements on the micropore. Preliminary VP current density measurements of electroporated HepG2 cells revealed a significantly high current density minutes after electroporation even with non-electroporative pulses. These results indicate that cells take a considerable amount of time for complete electrophysiological recovery and indicate the use of the VP as a cell viability indicator for optimized electroporation.
KeywordsElectroporation Biochip Micropore Current density Vibrating probe
Every eukaryotic cell is enclosed inside a dynamic phospholipid bi-layer known as the cell membrane. The cell membrane serves to maintain the microenvironment of the cell, maintain the membrane potential, and also acts as a selectivity filter for controlled transport of biological species into and out of the cell. An uncontrollable breach in the cell membrane can cause severe damage to the cell leading to necrosis and apoptosis. Recent advances in medical sciences have offered new diagnostic and therapeutic paradigms including targeted drug delivery (Denet et al. 2004), gene therapy (Andre and Mir 2004), and intracellular organelle modulation (Lemaire et al. 2007). Successful implementation of these techniques requires controlled and reversible penetration through the cell membrane into the cytoplasm. This can be achieved through several mechanisms including: a) biological processes (endocytosis, membrane fusion, viral vectors); b) chemical processes (surfactants) and; c) physical methods (membrane abrasion, microneedle penetration, gene gun, ultrasound, hydrodynamic injection, electropermeabilization) (Tryfona and Bustard 2005; Niidome and Huang 2002). Among these, electropermeabilization has been the preferred method because of its ease of use and high turnaround time. Commonly known as electroporation, this involves the transient increase in membrane permeability through application of an appropriate electrical pulse (Diaz-Rivera and Rubinksy 2006). Application of a high intensity electric field pulse (~1 kV) for a short duration (10−6 to 10−1 s) is believed to cause localized rearrangement of the lipid bilayer. This results in the creation of water filled pores which provide a free path for ions or molecules to move into the cell (Neumann et al. 1989; Tsong 1991; Chang 1992; Weaver 2003).
Conventional electroporation is generally performed on cells suspended in a conductive medium between two electrodes. In spite of the ease of use this setup provides, there are certain drawbacks. The electroporation voltage required for optimum reversible electroporation is dependent on cell size. Hence, the application of a voltage specified for a given cell size on a batch of cells with size variations (generally the case), can result in no transfection (smaller cell size) or irreversible electroporation (larger cell size). In addition, the high voltages required in the electroporation process induces Joule heating of the medium, which causes cell injury (Lee and Kolodney 1987; Lee et al. 2000). Thus the low cell viability and low transfection rates are trademarks of conventional electroporation. To circumvent these issues, achieve a greater level of control and understand the fundamental mechanisms of electroporation, researchers have shifted their focus towards single cell electroporation through miniaturization of the electroporation technique. This provides several advantages such as a) use of low voltages thereby decreasing damage, b) site specific electroporation through concentration of electric field, c) independence from cell size, and d) enabling the study of biokinetics and biochemical mechanisms at the cellular level (Diaz-Rivera and Rubinsky 2006).
Single cell electroporation has been performed using carbon microelectrodes (Lundqvist et al. 1998) and glass micropipettes (Haas et al. 2001; Rae and Levis 2002). The technique which has received the most attention and is also the focus of our lab involves the use of micro apertures or pores on planar surfaces. Called microelectroporation, the technique uses micropores (1/3rd the size of the cell diameter) fabricated on a planar substrate in which a single cell can be trapped through application of slight negative pressure. Application of a small voltage pulse concentrates the electric field through the pore resulting in localized electroporation (Huang and Rubinsky 1999). Electrical parameters can be measured across the cell for real time feedback and control which are the hallmarks of this technology (Huang and Rubinsky 1999, 2001, 2003).
In order to optimize microelectroporation and understand its fundamental mechanistic properties it is necessary to elucidate the microscale physical and electrical properties of the systems involved in this process. An important parameter is the current density traversing through the conducting medium and converging on the pore. More important are the currents traversing through a cell before, during, and after electroporation. For instance, a single cell has a standing membrane potential and a dynamic equilibrium of cytoplasmic ion concentrations, and this fundamental electrophysiological equilibrium can be potentially abolished during electroporation. For successful reversible electroporation, the membrane has to seal back and the cell needs to return to its original electrophysiological equilibrium to survive and remain viable.
Ryttsen et al. (2000) adopted a unique approach to indirectly map the electroporation electric field spatial distribution of micro electrode based single cell electroporation. Two 5 µm diameter carbon fiber ultramicroelectrodes placed within 2–5 µm on either side of single cell created a focused electric field for single cell electroporation. Simultaneously a patch was drawn on the cell at an angle of 90º from the carbon electrodes with a patch-clamp pipette. Measurement of current responses of the cell in whole cell configuration during electroporation enabled determination of electropermeabilization potential. Maximum transmembrane current response was determined by withdrawing the patched cell in small steps away from the focus of the electric field. Electrophysiological response of the cell during and post electroporation was also measured through real time measurement of cell current response during and after electroporation pulse. A threshold voltage of 250 mV was estimated to cause electroporation.
The impressive technique used by Ryttsen however, poses some challenges especially for characterizing micropore based single cell electroporation. First, patch-clamping by nature is an invasive technique which disturbs the natural environment of the cell in addition to electroporation. Second, this technique cannot be used to directly measure the electric field or current density around the cell or micropore, which is essential for the validation of a computational model. Third, characterization of a micropore based electroporation biochip without a cell is not possible. A technology is required which can non-invasively characterize the spatial distribution of electric fields on and around the pore of a microelectroporation biochip with high resolution and do so in real time with and without a cell.
Here we report on the electrical characterization of a microelectroporation biochip using the 2-D VP technology. This is not only the first report on real time current density measurements around a microelectroporation pore but also, to our best knowledge, the first application of the VP technology to study a MEMS based device. Current density measurements were performed at various distances from a micropore subjected to an electroporation pulse, both in the perpendicular (the z plane) and parallel planes (the r plane) relative to the plane of the biochip surface. A Finite Element Method (FEM) simulation was also developed and the experimental measurements were found to strongly correlate with it. Extracellular current density measurements were also performed on Human hepatocellular carcinoma (HepG2) cells pre-, during-, and post-electroporation.
2 Materials and methods
2.1 Micropore fabrication
The micropore was fabricated in silicon nitride on an underlying silicon substrate with a process described previously (Diaz-Rivera and Rubinsky 2006). Briefly, a 1 µm thick LSN film was chemically grown on a <100> double sided polished silicon substrate through a Low Pressure Chemical Vapor Deposition (LPCVD) process. Low stress silicon nitride (LSN) is optically translucent making it compatible with upright or inverted microscope configurations. It is also electrically nonconductive which is necessary to ensure that the micropore provides the only passage for the ionic current. Positive photoresist was spun on the wafer and the micropore pattern was photolithographically defined in it. This served as a mask for the Reactive Ion Etching (RIE) process, which created the micropore in the silicon nitride film. The average diameter of the microfabricated pores throughout the wafer measured 5.5 ± 0.26 µm (n = 10). The top photoresist was then removed followed by a backside photolithography step which opened a square window for etching the silicon nitride. After RIE etching of the backside silicon nitride, the photoresist was removed and the wafer was subjected to an anisotropic KOH silicon etch (H2O:KOH = 2:1) at 80°C. KOH etches silicon at an angle of 57.4° but does not etch silicon nitride. Therefore, the silicon nitride acted as an etch mask (backside) as well as an etch stop (front side) for the KOH etch. This step yielded a 1 µm thick LSN membrane suspended on a silicon substrate with a 5.5 µm pore at its center. Finally, a 0.1 µm silicon oxide layer was thermally grown on the exposed silicon.
2.2 2-D vibrating probe system
Electrical characterization measurements were made with a 2-D vibrating probe system (Applicable Electronics, Forestdale, MA). The r/z vibrating probe, which vibrates the probe along the r and z planes, (with respect to the top surface of the biochip) was employed. The probe consists of a parylene insulated Pt/Ir microelectrode (Microprobe Inc., Gaithersburg, MD) with a tip diameter of 2-5 µm. The tip of the probe was electroplated in a 10% hexachloroplatinate solution with 1% lead acetate (Sigma-Aldrich, St. Louis, MO).This process electroplates a ball of mesoporous Pt on the electrode tip. This highly dense structure visually appears black. Therefore it is called Pt black. After plating, the capacitance of the probe was measured with a digital oscilloscope. Only probes with a capacitance >2 nF were used. The probe was mounted on the 2-D vibrating probe assembly which houses the piezoresistive vibration elements. The VP assembly was attached to a 3-D motion control system. Both, the VP assembly and the motion control system were set up on a vibration isolation table that was enclosed by a Faraday cage. An upright video zoom microscope with a maximum optical magnification of 14X was integrated with the system for probe and sample location. When the probe is vibrated in a bioelectric field while immersed in a conductive medium, it measures the voltages at the two extremes of vibration which are usually 20–30 µm apart. The electric field is now equal to the voltage differential between the two points divided by this distance (amplitude of vibration). The electric field multiplied by the conductivity of the medium yields the current density traversing the center point of probe vibration. The output of the probe is an AC voltage, which is fed into the Phase Sensitive Detection Amplifier (PSDA) or lock-in amplifier after pre-amplification. The PSDA served the triple function of modulating the driving frequencies of the vibrating probe, signal amplification and signal rectification to report a DC output. Thus, the data was collected only at the frequencies at which the probe was driven, effectively filtering out most of the external noise. The probe was always vibrated away from 50/60 Hz and its multiples to avoid power line noise. General vibrating frequencies were approximately 260 Hz for the r and 150 Hz for the z vibration axis. The amplitude of vibration was always kept to be one tip diameter for both r and z vibrations. All calibrations, measurements, imaging, and motion control were performed using the ASET software (Sciencewares Inc., Falmouth, MA). The stepper motors of this modern 2-D VP have a resolution in the 100’s of nm. This gives the VP a very high spatial resolution necessary for mapping current densities at various points around a cell.
2.3 Probe calibration
The 2-D VP was calibrated using a 1.5 mm outer diameter glass capillary pulled to a tip diameter of 2–5 µm on a vertical puller (David Kopf Instruments, Tujunga, CA) and filled with 3 M KCl to yield a point source. After taking a reference measurement, the VP was brought at a distance of 150 µm from the tip of the point source in the r-plane (z = 0) and was calibrated by the ASET software. The same process was repeated in the z-plane. For a 60 nA current emanating from the tip of the source in a medium of known resistivity, the current density at 150 µm from the source tip is 21.2 µA/cm2 (see section 3.1). A probe was considered to be calibrated when the measured current was within ±1 µA/cm2 of the theoretically expected value. Since the calibrations are dependent on the resistivity of the medium, all calibrations were performed in Dulbecco’s Phosphate Buffered Saline (DPBS) (Sigma-Aldrich, St. Louis, MO), which is also the medium in which characterization and electroporation studies were performed. Calibration was followed by step back experiments on the point source to determine the spatial profile of the point source and also to ensure the viability of the probe as a spatial current density profiling tool. Once calibrated, the probe frequency and amplitude were left undisturbed. Setup for calibration is shown in Fig. 1.
2.4 Experimental setup
For the experiments, fluid reservoirs needed to be constructed above and below the micropore. The top well contained the cells and all measurements would be performed in this half. The bottom chamber was connected to fluid inlet and outlets and was used to maintain the negative pressure required to hold a cell in the pore. A custom polycarbonate assembly was machined to achieve this goal. The top half of the assembly forms an open well around the pore and also has grooves to house the top silver/silver chloride (Ag/AgCl) electrode (In Vivo Metric Biomedical Products, Healdsburg, CA). This served as the counter/reference electrode. The bottom half forms a closed well and also houses the working Ag/AgCl electrode. The pressure inside the pore is controlled manually with a syringe and is read out using a differential pressure transducer (Omega, PX26-015DV, Stamford, CT). The micropore assembly was then mounted on the stage of the vibrating probe rig and purged with DPBS ensuring removal of all air bubbles. The micropore was further brought into view and the vibrating probe was positioned near the pore for taking measurements. Cell electroporation measurements were performed similarly with the probe brought into position after a cell had been trapped in the pore. A schematic of the experimental setup is shown in Fig. 1.
2.5 Cell preparation
HepG2 cells used in these experiments were obtained from the Purdue University Cytometry Laboratories (PUCL). Cells were grown in an incubator supplied with 5% CO2 at 37°C in Dulbecco’s Modified Eagle Medium (DMEM) (Invitrogen Coro., Carlsbad, CA) supplemented with 10% fetal bovine serum, 100 u/ml penicillin and 10 µg/ml streptomycin. Cells were passaged every 4 days with trypsin/EDTA which was utilized to detach the cells. Before the experiments began, the cells were re-suspended in DPBS.
2.6 Experimental procedure
For cell electroporation studies, a 5 ml solution of HepG2 cells suspended in DPBS was pipetted into the top well. A slight negative pressure of 2.0 ± 0.1 kPa was applied to trap and hold a cell in the micropore. Once the cell was trapped, the system was kept on hold for one minute to achieve stability. The cells were then subjected to a 2 s 100 mV non-electroporative pre-pulse, a 100 ms or 2.5 s 500 mV electroporation pulse (Khine et al. 2005), and finally another 10 s 100 mV non-electroporative post-pulse. Current density measurements were performed during this entire phase with the vibrating probe stationed at a height of 60 µm from the center of the pore, approximately 40 µm from the top poles of the 15–20 µm diameter cells.
2.7 Finite element model
3.1 Point source step back experiments
3.2 Finite element model
3.3 Electrical characterization of pore with 2-D VP system
3.4 Single cell electroporation measurements
The spatial profile of a point source was mapped by the vibrating probe and compared to a theoretical current density profile (Fig. 7). At distances less than 200 µm from the point source the VP agrees with the theoretical current density profile, but at distances beyond 200 μm the model values are constantly lower than the measured VP values. The reason the theoretical values do not match the measured values at these distances is that the simple theoretical model of the current density profile fails to account for diffusional parameters associated with the ionic components of the current. Since the point source current is actually an ionic current, these effects become more important at larger distances from the point source. Therefore, the VP profile is more accurate then the theoretical profile, thus illustrating the need to directly measure these profiles in microelectroporation systems.
The COMSOL Multiphysics software is becoming increasingly popular in the modeling sector, and thus it was selected to simulate the theoretical current density profiles. The simulated and VP measured current density profiles match each other closely, validating the model. We observe from both these data sets that the actual current density at the pore is extremely high, approximately 9566000 µA/cm2, with an electroporation voltage that is much lower than the one employed in traditional electroporation. The rapid drop in current density in the perpendicular and parallel planes is the key to the localized electroporation effect as well as minimum collateral damage observed in microelectroporation (Huang and Rubinsky 2003). This is attributed to the electric field concentration near the micropore, which strangulates the ionic current flow and works as an ideal target site. The drop in current density even at 20 µm from the micropore is 100 fold smaller which might not be enough to cause electroporation. This agrees with Ryttsen’s observation that for a 250 mV electroporation pulse, electroporation does not occur for a cell even at 27 µm from the focus point of a microelectroporation electric field. This is the first time we have been able to directly measure the electric field near the micropore and thus elucidate the reasons behind the single cell targeting attributes of the microelectroporation biochip. These are the hallmarks of microelectroporation which cannot be obtained by traditional means.
Single cell experiments on the combined microelectroporation and VP rig reveal a high current density passing through the cell during electroporation which is not the case with a small non-electroporating pre-pulse. However, a small current density is observed even with the non-electroporative pre-pulse, which is attributed to a leakage current. This is expected as it is not possible to attain a giga-ohm seal between the cell membrane and the nitride surface. We expect to use glass in future designs which is known to achieve better seals. An electroporated cell subjected to a non-electroporative pulse was found to pass a significantly higher current, clearly indicating that the cell membrane does not immediately recover. This phenomenon was observed even 2.5 min post electroporation. This agrees with reports that indicate that even though membrane resealing might take a few seconds, membrane structural and electrophysiological properties take a much longer time to recover (Teissie et al. 2005).
Finally, to demonstrate the spatial profiling capabilities of the 2-D VP, electroporation current densities traversing through a single cell which was subjected to subsequent electroporation pulses were measured at various radial distances from the top of the cell. Highest current density was observed on top of the cell which is the site of pore formation along with the membrane portion trapped in the pore. The electroporation current density magnitude decreases away from the cell. Our current limitations with the video microscope zoom and access limitations to the bottom portion of the cell membrane prevent us from mapping current densities all around the cell. This should be possible with a modified setup in the future where the cell can be trapped in to the pore from the bottom well.
These preliminary results support the application of the VP as a tool for measuring cell viability—the key to successful reversible microelectroporation. Where microelectroporation provides a platform to study the intricacies of membrane permeabilization and cell arrangement, the VP complements it to non-invasively characterize the electrical parameters of the cell and biochip. By selectively removing extracellular ions from the medium, contribution of each ion to the electrophysiological recovery current can be determined. Of special importance is Ca2+ which can trigger secondary signaling pathways including apoptosis even with nM increase in its cytosolic levels. We intend to study dynamic Ca2+ flux patterns on electroporated cells in the future using the self-referencing Ca2+ selective electrode in combination with the microelectroporation system. We also intend to study dynamic O2 flux on electroporated cells using the self referencing oxygen optrode, with the potential of using cell metabolism as a cell viability indicator.
With the rapid progress in micro and fabrication technologies and reduction in costs associated with them, the research community has steadily moved towards micro and nano tools to interface with organisms at the cellular level. The planar microelectroporation technology is one these examples. The ability to locally induce electroporation on single cells with minimum collateral damage has made this a very important tool for cell transfection and electroporation research. Tools are needed which can characterize these devices on the micro scale in order to optimize their operational parameters. The Vibrating Probe technology, a tool initially developed for developmental biology research, has been used to characterize the 3-D spatial microelectroporation current density profiles. The results compare well with those obtained from an FEM model and were found to follow a 4th order polynomial in the planes parallel to the surface of the biochip while a three parameter hyperbolic decay model approximates the current density decay in the perpendicular plane. Preliminary results on cells suggest that though membrane resealing might occur within seconds, complete electrophysiological recovery might take a much longer time. Our results thus emphasize the importance of the microelectroporation technology in focused localized electroporation as well the importance of the VP technology as a potential tool for measuring cell viability as well as a general tool for characterizing MEMS based devices. In the future, we expect to be able to elucidate the dynamic transport of physiologically relevant ions during and post electroporation and cellular metabolism through integration of the self referencing ion selective electrode and optrode technology with the microelectroporation system.
The authors would like to acknowledge members of the Purdue University Cytometry Laboratories (PUCL) for their support with microscopy and cell culture. We would also like to thank Eric McLamore for his assistance on the microprobe rigs and the staff of Bindley Bioscience Center. This work was partially funded by the Institute for Functional Nanomaterials and the Collaboration in Biomedical Engineering Research, a joint initiative between the University of Puerto Rico at Mayagüez and the Weldon School of Biomedical Engineering at Purdue University.
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