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Biomedical Microdevices

, Volume 11, Issue 6, pp 1239–1250 | Cite as

Electrical characterization of a single cell electroporation biochip with the 2-D scanning vibrating electrode technology

  • Aeraj ul Haque
  • Mahvash Zuberi
  • Ruben E. Diaz-Rivera
  • D. Marshall Porterfield
Article

Abstract

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.

Keywords

Electroporation Biochip Micropore Current density Vibrating probe 

1 Introduction

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.

A tool with these capabilities does exist and has been employed by developmental biologists and electrophysiologists to study endogenous bioelectric fluxes across living cells and tissues for more than three decades. Originally developed by Jaffe and Nuccitelli (1974) the VP technology utilizes a 1–2 µm tip diameter insulated platinum (Pt) microelectrode with a 25–30 µm ball of mesoporous Pt black electroplated at its tip. This creates a low resistance, but highly capacitive tip which provides a high signal to noise ratio. The tip is vibrated in the extracellular ionic gradient of a cell or tissue while immersed in a medium of known conductivity. A lock-in amplifier drives the piezoelectric bender element inside the vibrating probe assembly with an AC signal that in turn vibrates the probe. The VP measures the voltage difference between its two extremes of vibration (usually 30 µm apart) which is recorded in the form of a sinusoidal output. Since the electric field E will be nearly constant over this small distance it is given by voltage difference ΔV divided by the excursion distance Δx. This electric field multiplied with the conductivity of the medium yields the current density traversing the center of vibration of the VP. Modern digital VP systems can measure two dimensional current density vectors and with nm resolution stepper motors are capable of achieving less then a micron spatial resolution. The working principle of the VP is shown in Fig. 1. The technology has been used extensively for measuring developmental currents across single cells (Nuccitelli and Jaffe 1974) injury and regeneration currents emanating from plant organs and mammalian tissues (Borgens et al. 1977, 1980; Weisenseel et al. 1992; Ried et al. 2007; Zuberi et al. 2008). It has also found a non-biological use in the corrosion industry and is used commonly in naval and automobile research (Aldykiewicz and Isaacs 1998; Krawiec et al. 2004).
Fig. 1

(a) Experimental setup for calibrating the VP. A 60 nA current passed through the calibration point source should report a current density of 21.2 μA/cm2 at a distance of 150 μm. This procedure was used to calibrate the VP. (Inset) 14x magnified image of the VP located at 150 µm from the point source while the vibration element is off. To observe the spatial current density profile of the point source, the VP was stepped back from a distance of 150 μm (b) to a distance of 600 μm (c) at fixed steps and the current density was measured at each point (d) Experimental setup for characterizing the microelectroporation biochip and measuring the pre/during/post electroporation current densities through a single cell (e) Working principle of the vibrating probe technology. The probe is oscillated in the extracellular bioelectric field of the sample at fixed frequency. The peak-to-peak voltage divided by the amplitude of vibration yields the electric field at the center of vibration. This electric field times the conductivity is equal to the current density

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

An electroporation system calibration measurement was performed before each characterization experiment by application of a square pulse on the electroporation chip. This was done to ensure that the pore was not clogged with any debris. Current, voltage, and resistance calibration measurements are shown in Fig. 2. For all measurements, the center of the pore was taken as origin (r = 0, z = 0). The systems output voltage represents an input voltage given by the equation
$$ U = \cos \theta \left( {L\rho J} \right) $$
(Jaffe and Nuccitelli 1974) where θ is the angle of the current vector with respect to the angle of vibration, L is the peak to peak amplitude of probe’s vibration, ρ is the resistivity of the medium and J is the magnitude of the current density in the probed region. From this equation it was immediately realized that the center of the pore will be the point where the probe measures zero current density in the r-direction and maximum current density in the z-direction. This principle was used to locate the origin. Electrical characterization of the biochip was performed by measuring the current densities for a 500 mV electroporation pulse at various distances in the r plane starting from origin at a fixed height (z) from the pore. The measurements were then repeated for increasing heights ranging from 40–150 µm. Measurements closer than 40 µm were not possible due to fear of crashing the probe with the surface of the biochip.
Fig. 2

Calibration measurements performed on the microelectroporation biochip. A 500 mV voltage pulse is applied for 2.5 s through the pore with DPBS as the conductive medium. Approximately 3.3 µA current passed through the pore at this voltage yielding a pore resistance equaling 150 kΩ

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

FEM simulations were performed in COMSOL Multiphysics software to model the expected current densities around a micropore subjected to an electroporation pulse. In this case, we were interested in modeling the current densities in a conductive media around a microscale pore excited by a DC static field. To achieve this, the static form of the continuity equation applied to Ohm’s law was used;
$$ \nabla \cdot J = - \nabla \cdot \left( {\sigma \nabla U} \right) $$
where J is the current density vector, σ is the conductivity of the medium (DPBS, 1.38 S/m at 22°C, Diaz-Rivera and Rubinsky 2006) and U is the voltage throughout the conductive medium. A 2-D model portraying half of the pore was used to exploit the symmetric nature of the pore in the angular (φ) direction and thus decrease the computation load. Boundary conditions were defined to be U = 500 mV at the top boundary (T), U = 0 at the bottom boundary (B), electrical insulation \( \left( {n \cdot J = 0} \right) \) at the LSN surface (L), and constant electrical potential in the angular direction with respect to the symmetry line (S). The final model after meshing and with boundary conditions explained is shown in Fig. 3. A finer mesh size was used near the pore and in the upper half well as these were the areas accessible by the VP, and the modeled data could be compared directly with the experimental data.
Fig. 3

Screen capture of the mesh used in the 2-D FEM model of the microelectroporation biochip generated in COMSOL Multiphysics. The boundary conditions were defined as top voltage U = 500 mV (T), bottom voltage at ground (B), insulated LSN membrane (L), and no angular variation in voltage with respect to the symmetry line (center of pore, S)

3 Results

3.1 Point source step back experiments

The patterns of electric fields and currents around a point source polarized at a fixed potential in a conductive medium is given by
$$ J = \frac{I}{{4\pi {r^2}}} $$
where I is the current at the source tip and J is the magnitude of the current density at a distance r from the tip. From this equation the current density measured at a distance of 150 µm from the source is 21.2 µA/cm2. The spatial current density profile of the point source was determined by the vibrating probe from this point backwards to a radial distance of 600 µm where only the background current is assumed to be present. The theoretically calculated data with the actual measured values from the VP are displayed in Fig. 4.
Fig. 4

Theoretical vs. measured spatial current density profile of the point source. Notice that the VP overestimates the current density profile at distances greater than 200 μm. This is however attributed to limitations in the theoretical model which does not account for diffusion related effects which might become dominant at larger distances. Therefore the VP measured data is considered to be more accurate

3.2 Finite element model

The finite element model simulated in COMSOL was used to visualize the current density profile near and around the pore. The current density vectors as displayed by COMSOL illustrate that a high current density can be observed near the pore which decreases at radial distances away from the pore. Figure 5 shows the current density vectors over the entire region and focused near the pore. The output from the model was used to plot the current density at various radial distances from the center of the pore at different heights to enable comparison with actual data collected from the VP which are shown in Fig. 6.
Fig. 5

(a) Modeled current density profile generated on the microelectroporation biochip on application of a 500 mV electroporation pulse displayed as current density vectors where the length of the arrows is proportional to the magnitude of the local current density. (b) Close up of the current density vectors near pore. Notice that the current densities are higher near the pore and drop rapidly away from it

Fig. 6

Modeled vs. measured spatial current density profiles generated on the application of a 500 mV electroporation pulse. Current densities were measured at various radial distances from the center of pore starting at 20 μm and ending at 150 μm where the current densities were indistinguishable from background noise. Similar measurements were made at various z planes. The measured values agree well with modeled values validating the accuracy of the model

3.3 Electrical characterization of pore with 2-D VP system

The initial goal of this research was to map the current density profile around a micropore subjected to an electroporation pulse. This would serve to elucidate the magnitude of current densities faced by a single cell under microelectroporation. Current density measurements were made with the 2-D VP at various positions from the pore in both the perpendicular (z) and parallel planes relative to the silicon chip surface, where the 2-D VP is vibrated in these planes. This should be representative of current densities in the entire 3-D plane because of pore symmetry. Measurements were made at 20–150 µm in the parallel plane and in the perpendicular planes ranging from 40–150 µm. Perpendicular plane measurements were limited to 40 µm or greater height from pore because of fear of damaging the probe. The outer extreme position away from the pore was where signals became indistinguishable from background. A representative VP data set is shown in Fig. 6 along with the modeled values which appear to follow each other very closely. A 4th order polynomial provided the best fit for the current density decay in the planes parallel to the micropore surface and is given by
$$ J\left( {r,z} \right) = a(z){r^4} + b(z){r^3} + c(z){r^2} + d(z)r + e(z) $$
([1])
where J is the current density, r is the distance from the center of the pore while a(z) to e(z) are quasi constants whose values depend on the perpendicular distance at which the parallel plane current densities were measured. Finally, the profile of the spatial current density measured by the VP in the perpendicular direction (z plane) from the center of the pore is shown in Fig. 7 and was found to fit a 3 parameter hyperbolic decay profile as
Fig. 7

Spatial current density profile for a 500 mV pulse at pore center as a function of perpendicular distance from the pore (distance in z). A three parameter hyperbolic decay model was found to best fit the data

$$ J\left( {r = 0,z} \right) = {y_o} + \frac{ab}{b + z} $$
([2])
where z is the perpendicular distance from the pore while y 0 , a, and b are constants whose values are y 0  = −26,455.94, a = 9,592,456.68, and b = 0.2654.

3.4 Single cell electroporation measurements

HepG2 cells ranging in diameter from 15–20 µm were selected as the model system for electroporation studies. Experimentation with different cell types revealed that any cell system can be used as long as their size in suspension is greater than 3 times the pore’s diameter (15–20 µm). Since all cells cannot bear the 2 kPa holding pressure, only those cells were used which could tolerate this pressure. For example 18–20 µm diameter P-19 embryonic stem cells would pass through the pore easily at the holding pressure while even 12 µm diameter HepG2 cells would stay in the pore. As such, HepG2 cells were selected for experimentation. Figure 8 shows a single HepG2 cell with the VP positioned near it. It was ensured that the rest of the cells settle atleast 150 µm away from the pore to prevent any interference form these cells. In practice, the VP was positioned on top of the cell at a height of 60 µm from the top of the pore with an approximate center of vibration 40 µm above the top pole of the cell. In the first set of experiments, the cells were subjected to three pulses; a 1 s 100 mV non-electroporative pre-pulse; a 100 ms 500 mV electroporative pulse followed by a 1 or 5 s 100 mV non-electroporative post-pulse. Current density traces from 7 cells and average pre/during/post electroporation current densities are shown in Fig. 9. The current observed during the pre-pulse is attributed to the leakage current passing around the cell, because the excitation voltage is below the threshold for electroporation. A higher current density is observed passing through the cell on application of the post electroporation pulse. For the seven cells a paired sample t-test reveals an average p-value of less then 1% implying that at a 5% significance level the pre and post electroporation current densities are significantly different. This clearly implies that post electroporation; there is a transient increase in membrane conductance of the cell which does not immediately recover to its initial level.
Fig. 8

Image of a HepG2 cell trapped in the micropore. Once trapped, the VP was positioned on top of the cell to take the measurements at a height of 40 μm from the top of the cell

Fig. 9

(a) Current density traces of 7 HepG2 cells subjected to a non-electroporative 100 mV pre-pulse, a 500 mV electroporation pulse and a 100 mV non-electroporative post-pulse. A higher current density was observed to traverse through the cells on application of an non-electroporative post pulse. (b) Average of pre/during/post electroporation current densities traversing through the 7 HepG2 cells. A statistically significant difference is observed between the pre and post-pulses which clearly shows that the cell membrane does not seal immediately post electroporation

A second set of experiments was performed to better understand the time scales involved in complete membrane recovery and are shown in Fig. 10 with the average linear leakage current subtracted from the current density traces. A cell was subjected to a three step pulse as before, with the only difference that the electroporation pulse was 2.5 s long. The cell was kept trapped in the pore with a 2 kPa negative pressure and then subjected to another 5 s 100 mV non-electroporative pulse 1 min after electroporation. The same process was repeated at 2.5 min after electroporation. We observed that even 2.5 min after electroporation a current density synonymous with that observed immediately post electroporation is recorded. This means that the membrane is still in the phase of electrophysiological recovery.
Fig. 10

Current densities through a HepG2 cell subjected to a 500 mV electroporation pulse and 100 mV pre and post non-electroporative pulses. A significant difference from pre-pulse current densities was observed even 2.5 min after electroporation clearly revealing that the cell takes a much longer time for complete structural and electrophysiological recovery under the current electroporative conditions. (Linear leak current is subtracted from the traces)

Finally to observe the effect of current density as function of distance from an electroporated cell, HepG2 cells were trapped in the micropore and subjected to subsequent electroporation pulses. The current density was simultaneously monitored at a height of 40 µm (z from pore ~55 µm with a 15 µm diameter cell) above the cell and at radial increments of 20 µm in the same z plane. The average current density spatial profile obtained from 5 cells is shown in Fig. 11. The highest current density is observed on top of the cell as expected. This is consistent with previous reports where the current density and thus the electric field is estimated to reach a local maximum in the centerline of the micropore (Huang and Rubinsky 2001; Diaz-Rivera and Rubinsky 2006; Lee et al. 2006). The magnitude of current density starts decreasing with radial distance from the top of the cell until it is indistinguishable from background at a radial distance of 100 µm.
Fig. 11

Spatial profile of electroporation current densities measured on 5 cells subjected to subsequent electroporation pulses. Highest current density is observed on top of the cell which decreases with radial distance away from the cell

4 Discussion

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.

5 Conclusion

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.

Notes

Acknowledgements

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.

Supplementary material

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References

  1. A.J. Aldykiewicz, H.S. Isaacs, Corros. Sci. 40, 1627 (1998)CrossRefGoogle Scholar
  2. F. Andre, L.M. Mir, Gene Ther. 11, S33 (2004)CrossRefGoogle Scholar
  3. R.B. Borgens, J.W. Vanable Jr., L.F. Jaffe, Proc. Nat. Acad. Sci. U.S.A. 74, 4528 (1977)CrossRefGoogle Scholar
  4. R.B. Borgens, L.F. Jaffe, M.J. Cohen, Proc. Nat. Acad. Sci. U.S.A. 77, 1208 (1980)Google Scholar
  5. D.C. Chang, Guide to electroporation and electrofusion (Academic Press, San Diego, 1992)Google Scholar
  6. A.-R. Denet, R. Vanbever, V. Preat, Adv. Drug Deliver. Rev. 56, 659 (2004)CrossRefGoogle Scholar
  7. R.E. Diaz-Rivera, B. Rubinsky, Biomed. Microdevices 8, 25 (2006)CrossRefGoogle Scholar
  8. K. Haas, W.C. Sin, A. Javaherian, Z. Li, H.T. Cline, Neuron 29, 583 (2001)CrossRefGoogle Scholar
  9. Y. Huang, B. Rubinsky, Biomed. Microdevices 2, 145 (1999)CrossRefGoogle Scholar
  10. Y. Huang, B. Rubinsky, Sensor. Actuat. A-Phys. 89, 242 (2001)CrossRefGoogle Scholar
  11. Y. Huang, B. Rubinsky, Sensor. Actuat. A-Phys. 104, 205 (2003)CrossRefGoogle Scholar
  12. L.F. Jaffe, R. Nuccitelli, J. Cell Biol. 63, (1974)Google Scholar
  13. M. Khine, A. Lau, C. Ionescu-Zanetti, J. Seo, L.P. Lee, Lab on a Chip. 5, 38 (2005)CrossRefGoogle Scholar
  14. H. Krawiec, V. Vignal, R. Oltra, Electrochem. Commun. 6, 655 (2004)CrossRefGoogle Scholar
  15. R.C. Lee, M.S. Kolodney, Plast. Reconstr. Surg. 80, 663 (1987)CrossRefGoogle Scholar
  16. R.C. Lee, D.J. Zhang, J. Hannig, Ann. Rev. Biomed. Eng. 2, 477 (2000)CrossRefGoogle Scholar
  17. E.S. Lee, D. Robinson, J.L. Rognlien, C.K. Harnett, B.S. Simmons, C.R.B. Ellis, R.V. Davalos, Bioelectrochem. 69, 117 (2006)CrossRefGoogle Scholar
  18. S. Lemaire, F. Van Bembeke, Mingeot-Leclercq M-P, P.M. Tulkins, Antimicrob. Agents Ch. 51, 2748 (2007)CrossRefGoogle Scholar
  19. J.A. Lundqvist, F. Sahlin, M.A.I. Aberg, A. Stromberg, P.S. Eriksson, O. Orwar, Proc. Nat. Acad. Sci. U.S.A. 95, 10356 (1998)CrossRefGoogle Scholar
  20. E. Neumann, A.E. Sowers, C.A. Jordan, Electroporation and electrofusion in cell biology (Plenum, New York, 1989)Google Scholar
  21. T. Niidome, L. Huang, Gene Ther. 9, 1647 (2002)CrossRefGoogle Scholar
  22. R. Nuccitelli, L.F. Jaffe, Proc. Nat. Acad. Sci. U.S.A. 71, 4955 (1974)CrossRefGoogle Scholar
  23. J.L. Rae, R.A. Levis, Pflug. Arch. Eur. J. Phy. 443, 664 (2002)CrossRefGoogle Scholar
  24. B. Ried, R. Nuccitelli, M. Zhao, Nat. Protoc. 2, 661 (2007)CrossRefGoogle Scholar
  25. F. Ryttsen, C. Farre, C. Brennan, S.G. Weber, K. Nolkrantz, K. Jardemark, D.T. Chiu, O. Orwar, Biophys. J. 79, 1993 (2000)CrossRefGoogle Scholar
  26. J. Teissie, M. Golzio, M.P. Rols, Biochim. Biophys. Acta. 1724, 270 (2005)Google Scholar
  27. T. Tryfona, M.T. Bustard, Biotechnol. Bioeng. 93, 413 (2005)CrossRefGoogle Scholar
  28. T.Y. Tsong, Biophys. J. 60, 297 (1991)CrossRefGoogle Scholar
  29. J.C. Weaver, IEEE T. Dielect. El. In. 10, 754 (2003)CrossRefMathSciNetGoogle Scholar
  30. M.H. Weisenseel, H.F. Becker, J.G. Ehlgotz, Plant Physiol. 100, 16 (1992)CrossRefGoogle Scholar
  31. M. Zuberi, P. Liu-Snyder, A. ul Haque, D.M. Porterfield, R.B. Borgens, J. Biol. Eng. 2, 17 (2008)CrossRefGoogle Scholar

Copyright information

© Springer Science+Business Media, LLC 2009

Authors and Affiliations

  • Aeraj ul Haque
    • 1
  • Mahvash Zuberi
    • 1
  • Ruben E. Diaz-Rivera
    • 2
    • 3
  • D. Marshall Porterfield
    • 4
    • 5
  1. 1.Department of Agricultural and Biological Engineering, Bindley Bioscience Center- Physiological Sensing FacilityPurdue UniversityWest LafayetteUSA
  2. 2.Department of Mechanical EngineeringUniversity of PuertoMayagüezPuerto Rico
  3. 3.Weldon School of Biomedical EngineeringPurdue UniversityWest LafayetteUSA
  4. 4.Department of Agricultural and Biological Engineering, Weldon School of Biomedical Engineering, Bindley Bioscience Center- Physiological Sensing FacilityPurdue UniversityWest LafayetteUSA
  5. 5.Department of Horticulture and Landscape Architecture, Weldon School of Biomedical Engineering, Bindley Bioscience Center- Physiological Sensing FacilityPurdue UniversityWest LafayetteUSA

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