Name: the fabrication and operation of a continuous flow, micro-electroporation system with permeabilization detection
Uploaded: 2022-01-07T13:00:00.000+00:00
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Description: Watch this Scientific Journal Video about The Fabrication and Operation of a Continuous Flow, Micro-Electroporation System with Permeabilization Detection at JoVE.com

The authors would like to acknowledge financial support by the National Science Foundation (NSF CBET 0967598, DBI IDBR 1353918) and the U.S. Department of Education&#39;s Graduate Training in Emerging Areas of Precision and Personalized Medicine (P200A150131) for funding&#160;graduate student J.J.S. on fellowship.

<ol>
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The authors have nothing to disclose.

The methodology presented within this protocol primarily focuses on the microfabrication of a microfluidic device that is then integrated into a specialized electroporation experimental setup. The term &#39;recipe&#39;, which is often used when describing the specifics of the microfabrication process, hints at the importance of following/optimizing each step to successfully fabricate a functioning device. However, certain critical steps within the process, when not optimized, such as UV exposure time/energy, PVD sputtering rates/durations, and oxygen plasma generator settings, can be problematic to both the fabrication process as well as the successful execution of the electroporation experiments. Troubleshooting the fabrication process is primarily done via trial and error or a more controlled Design of Experiments experimental design. Additionally, there are alternative microfabrication techniques, such as Deep Reactive Ion Etching (DRIE), that can be substituted to perform the different steps within the protocol (i.e., using a DRIE etched molding structure to perform the soft lithography process). Furthermore, optimizing recipes and designing/fabricating devices can be time-consuming for novices in the field. However, once the microfabrication process has been successfully developed, the engineer/scientist has the freedom to design a device that is suitable to their specific needs.
To that end, the device described within this protocol was developed to expand upon our previous work32. This entailed the utilization of the single-cell membrane permeabilization electrical detection but in a higher-throughput manner. The experimental setup described within requires the need for specialized equipment, i.e., lock-in amplifier, that may be uncommon to the standard research lab and thus limiting the potential outreach and adaptability of this technique. However, a &#39;bare-bones&#39; microfluidic electroporation device can be implemented following this protocol, requiring only a function generator and possibly a voltage amplifier to generate the electroporation pulses.
Nevertheless, this micro-electroporation platform distinguishes itself from other single-cell electroporation technologies. The ability to both electrically detect and optimize electroporation parameters on a single-cell suspension in a continuous-flow environment is truly innovative. Future work involves optimizing the other important experimental parameters related to successful electroporation outcomes (see Figure 1) to further improve the overall effectiveness of this platform. Additional viability and metabolic assays will be developed and implemented to assess any potential negative downstream effects associated with the micro-electroporation platform. Furthermore, the microfluidic design can continue to be improved upon to achieve higher cellular throughput, as has been demonstrated by other groups40. Upon addressing these concerns, this technology has the potential to be adopted into the cell therapy manufacturing process to perform gene delivery and/or gene editing, as this methodology is highly amenable to both a closed and automated process.

Current therapeutic innovations in biomedical research, such as CAR-T (Chimeric Antigen Receptor Engineered T cell) cell therapy and genetic editing using CRISPR (clustered regularly interspaced short palindromic repeat DNA sequences)/Cas9, heavily rely on the ability to deliver exogenous material both successfully and efficiently into the intracellular space1. In CAR-T therapy, the gold standard to perform the gene delivery step in cell therapy manufacturing is using viral vectors2. Though viral-mediated gene delivery is an efficient delivery modality, it also has several drawbacks. These include manufacturing costs, cytotoxicity, immunogenicity, mutagenesis/tumorigenesis potential, and size limitations on the gene(s) to be delivered3. These limitations have led to the research and development of alternative, non-viral delivery technologies.
Electroporation, an alternative to viral-mediated gene delivery, relies on the application of an optimal electrical pulse waveform to perform DNA, RNA, and protein transfections of cells. Following the application of an external electric field, the cell membrane is briefly compromised, making the cell susceptible to the intracellular delivery of otherwise impermeable exogenous materials4. Compared to viral-mediated delivery, electroporation is advantageous as it is generally safe, easy to operate, and has low operating costs. Electroporation can deliver both small and large molecular cargo and can be efficient in transfecting cells regardless of lineage5. To achieve desirable outcomes following electroporation, i.e., good viability and good electro-transfection efficiency, a variety of experimental parameters need to be co-optimized. These include cell type6, cell density, molecule concentration7, electroporation buffer properties (e.g., molecular composition, conductivity, and osmolarity)8, electrode size/geometry9, and electrical pulse waveform (shape, polarity, number of pulses)10 (refer to Figure 1 for an illustration). Although each of these parameters can have a significant effect on the outcomes of electroporation experiments, pulse waveform has been especially studied in great detail, as the electrical energy of the applied pulse(s) is the root of the intrinsic trade-off between the resulting cell viability and electro-transfection efficiency8.
Typically, electroporation experiments are performed on the macro-scale, where cells are suspended in 100s of microliters of buffer between a set of large, parallel-plate electrodes within an electroporation cuvette. The electrodes are commonly manufactured out of aluminum with an electrode distance of 1-4 mm. Once the cells are manually loaded via pipette, the cuvette is electrically connected to a bulky, electrical pulse generator where the user can set and apply the pulse waveform parameters to electroporate the cell suspension. Although macro-scale or bulk electroporation can process cell densities &#62;106 cells/mL, this feature can be wasteful when optimizing the electrical pulse waveform settings. This is particularly of concern when electroporating primary cell types where the cell population numbers can be limited. Additionally, due to the large distance between the electrodes, the pulse generator must be able to supply large voltages to achieve electric field strengths &#62;1kV/cm11. These high voltages cause resistive power dissipation through the electrolyte buffer resulting in Joule heating, which can be detrimental to the resulting cell viability12. Lastly, performing electroporation on a dense suspension of cells will consistently be burdened with an innate variability in the resulting electro-transfection efficiency and cell viability. Each cell in suspension could experience a different electric field strength due to the surrounding cells. Depending on whether the experienced electric field strength is either increased or decreased, the resulting cell viability or electro-transfection efficiency may each be negatively impacted11. These downsides to macro-scale electroporation have led to the pursuit and development of alternative technologies that operate on the micro-scale and allow for better control at the single-cell level.
The field of BioMEMS, or biomedical micro-electro-mechanical systems, stems from the technological advancements made in the microelectronics industry. Specifically, utilizing microfabrication processes to develop micro-devices for the advancement of biomedical research. These advancements include the development of micro-electrode arrays for in vivo electrical monitoring13, capacitive micro-electrodes for in situ electroporation14, miniaturized organ-on-a-chip devices15, microfluidic point-of-care diagnostics16, biosensors17, and drug delivery systems18, including nano- and micro-electroporation devices19,20,21. Due to the ability to design and manufacture devices at the same size scale as biological cells, nano- and micro-electroporation technologies are advantageous when compared to their macro-scale counterpart22,23. These electroporation devices eliminate the requirement of high voltage pulse applications, as electrode sets with spacings of 10s to 100s of micrometers are typically integrated. This feature drastically reduces the current through the electrolyte, which in turn reduces the accumulation of toxic electrolysis products and the effects of Joule heating in these systems. The micro-scale channels also ensure that a much more uniform electric field is reliably applied to the cells during pulse application, resulting in more consistent outcomes24. In addition, it is also commonplace for micro-electroporation devices to be integrated into a microfluidic platform which lends itself for future integration into a fully automated technology, a highly desirable capability in cell therapy manufacturing25. Lastly, micro-scale electroporation allows for the electrical interrogation of electroporation events. For example, the degree of cell membrane permeabilization can be monitored in real-time at a single cell level26,27. The purpose of this method is to describe the microfabrication, system operation, and analysis of a microfluidic, single-cell micro-electroporation device capable of measuring the degree of cell membrane permeabilization for optimizing electroporation protocols, yet increasing throughput over the previous state-of-the-art.
Performing single-cell level electroporation is no longer a novel technique, as it was first demonstrated by Rubinsky et al. in 2001 with the development of a static cell electroporation technology28. Their micro-device was innovative as they were the first to demonstrate the ability to electrically monitor the event of electroporation. This has further led to the development of static, single-cell electroporation technologies capable of electrically detecting the degree of cell membrane permeabilization in a parallelized manner to increase the throughputs of the devices. However, even with parallelization and batch processing, these devices severely lack the total number of cells they can process per unit time29,30. This limitation has led to the development of flow-through devices capable of performing single-cell level micro-electroporation at much greater throughputs31. This device transition, from static to flow-through environment, limits the capability of electrically monitoring the degree of cell membrane permeabilization following the application of the electroporation pulse. The method described in this work bridges the gap between these two technologies, a micro-electroporation technology capable of electrically detecting, pulsing, and monitoring the degree of cell membrane permeabilization of individual cells, in a continuous-flow, serial fashion.
This technology was recently described in Zheng et al. In that work, the capabilities of this technology were introduced with the completion of a parametric study, where both the amplitude and duration of the electroporation pulse were varied, and the ensuing electrical signal, indicative of cell membrane permeabilization, was explored32. The results showed that an increase in the intensity of the electroporation pulse (i.e., increase in applied electric field or increase in pulse duration) caused an increase in the measured cell membrane permeabilization. To further validate the system, a common fluorescent indicator of successful electroporation, propidium iodide33, was added to the cell suspension, and a fluorescence image was captured immediately following the application of the electrical pulse. The optical signal, i.e., the fluorescence intensity of propidium iodide inside the cell, was strongly correlated with the electrical measurement of the degree of cell membrane permeabilization, verifying the reliability of this electrical measurement. However, this work only considered the delivery of the small molecule propidium iodide, which has little to no translatable significance.
In this work, a new application of this technology is introduced to improve upon the throughput of the system while delivering a biologically active plasmid DNA (pDNA) vector and assessing the electro-transfection efficiency of cells replated and cultured following electroporation. Though the previous work outperforms existing micro-electroporation technologies that are capable of electrically measuring the event of electroporation, the current state of the device still requires long cell transit times between the electrode set (~250 ms) to perform the cell detection, pulse application, and the cell membrane permeabilization measurement. With a single channel, this limits the throughput to 4 cells/s. To combat this limitation, a new concept of cell-population-based feedback-controlled electroporation is introduced to perform pDNA electro-transfection. By using a hypo-physiologic conductivity electroporation buffer, this system allows for the electrical interrogation of single cells across a multitude of electroporation pulse applications. Based on the electrical response, an &#39;optimal&#39; electroporation pulse is then determined. A &#39;high-throughput&#39; mode is then implemented where the cell membrane permeabilization determination is nullified, the flow rate is increased, and the electroporation pulse duty cycle is matched to the cell transit time to ensure one pulse per cell in transit between the electrodes. This work will provide extensive details into the microfabrication steps for the manufacturing of the micro-device, the material/equipment and their setup required to perform the experimentation, and the operation/analysis of the device and its electro-transfection efficiency (eTE).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63103/63103fig01.jpg" /> 
Figure 1: Experimental factors affecting electroporation outcomes. (Left) Cell Suspension-Important factors to consider prior to the onset of electroporation include: Payload (in this case, pDNA), concentration, cell density, and electroporation buffer properties. Electroporation buffer properties to consider are conductivity, osmolarity, and the exact molecular composition contributing to these values. (Middle) Pulse Application-The exact pulse-type (square wave vs. exponential decay) and pulse waveform (single pulse vs. pulse train) must be optimized to maximize both the resulting cell viability and electro-transfection efficiency. Common pulse trains implemented in electroporation processes are typically composed of a series of High Voltage (HV) pulses or series of pulses rotating between HV and Low Voltage (LV) pulse magnitudes. (Right) Cell Recovery-Down-stream processing steps, in particular, the recovery cell culture media that cells are transferred to, should be optimized. Not featured (Far Left), additional upstream cell processing steps can be implemented for overall electroporation process optimization. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63103/63103fig1v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>150-mm diameter petri dishes</td><td>VWR</td><td>25384-326</td><td>step 6.1.1 to secure wafer</td></tr><tr><td>24-well tissue culture plates</td><td>VWR</td><td>10062-896</td><td>step 10.3.6 to plate electroporated cells</td></tr><tr><td>33220A Waveform/Function generator</td><td>Agilent</td><td></td><td>step 9.2.3 electroporation pulse generator</td></tr><tr><td>4&#039;&#039; Si-wafers</td><td>University Wafer</td><td></td><td>subsection 2.1 for microfluidic channel fabrication</td></tr><tr><td>6-well tissue culture plates</td><td>VWR</td><td>10062-892</td><td>step 8.1.8 to plate cells</td></tr><tr><td>Acetone</td><td>Fisher Scientific</td><td>A18-4</td><td>step 2.1.2 for cleaning and&amp;nbsp; step 5.1 photoresist lift-off</td></tr><tr><td>Allegra X-22R Centrifuge</td><td>Beckman Coulter</td><td></td><td>steps 8.1.4 , 8.3.2. and 8.3.3. to spin down cells</td></tr><tr><td>AutoCAD 2018</td><td>Autodesk</td><td></td><td>subsection 1.1. to design transparency masks</td></tr><tr><td>Buffered oxide etchant 10:1</td><td>VWR</td><td>901621-1L</td><td>subsection 3.1 for HF etching</td></tr><tr><td>CCD Monochrome microscope camera</td><td>Hamamatsu</td><td>Orca 285 C4742-96-12G04</td><td>step 11.2.3. for imaging</td></tr><tr><td>CMOS camera- Sensicam QE 1.4MP</td><td>PCO</td><td></td><td>subsection 9.3 part of the experimental setup</td></tr><tr><td>Conductive Epoxy</td><td>CircuitWorks</td><td>CW2400</td><td>subsection 7.6. for wire attachement</td></tr><tr><td>Conical Centrifuge Tubes, 15 mL</td><td>Fisher Scientific</td><td>14-959-70C</td><td>step 8.1.4. for cell centrifuging</td></tr><tr><td>Dektak 3ST Surface Profilometer</td><td>Veeco (Sloan/Dektak)</td><td></td><td>step 2.1.15 and 5.4 for surface profilometry</td></tr><tr><td>Disposable biopsy punch, 0.75 mm</td><td>Robbins Instruments</td><td>RBP075</td><td>step 6.2.3 for inlet access</td></tr><tr><td>Disposable biopsy punch, 3 mm</td><td>Robbins Instruments</td><td>RBP30P</td><td>step 6.2.3 for outlet access</td></tr><tr><td>DRAQ5</td><td>abcam</td><td>ab108410</td><td>step 11.2.2. for live cell staining</td></tr><tr><td>Dulbecco&amp;rsquo;s Modified Eagle&amp;rsquo;s Medium</td><td>ThermoFisher Scientific</td><td>11885084</td><td>step 8.1.2. part of media composition</td></tr><tr><td>E3631A Bipolar Triple DC power supply</td><td>Agilent</td><td></td><td>step 9.2.1.-9.2.2.part of the experimental setup</td></tr><tr><td>Eclipse TE2000-U Inverted&amp;nbsp; Microscope</td><td>Nikon</td><td></td><td>&amp;nbsp;subsection 9.3. part of the experimental setup</td></tr><tr><td>EVG620 UV Lithography System</td><td>EVG</td><td></td><td>&amp;nbsp;step 2.1.9. and 2.2.7. for UV Exposure</td></tr><tr><td>Fetal Bovine Serum</td><td>Neuromics</td><td>FBS001</td><td>step 8.1.2. part of media composition</td></tr><tr><td>FS20 Ultrasonic Cleaner</td><td>Fisher Scientific</td><td></td><td>subsection 5.1. for photoresist lift-off</td></tr><tr><td>Glass Media Bottle with Cap, 100mL</td><td>Fisher Scientific</td><td>FB800100</td><td>step 8.2.1. for buffer storage</td></tr><tr><td>Glass Media Bottle with Cap, 500mL</td><td>Fisher Scientific</td><td>FB800500</td><td>step 8.1.2.for media storage</td></tr><tr><td>HEK-293 cell line</td><td>ATCC</td><td>CRL-1573</td><td>subsection 8.1 for cell culturing</td></tr><tr><td>HEPES buffer solution</td><td>Sigma Aldrich</td><td>83264-100ML-F</td><td>step 8.2.1 part of electroporation buffer composition</td></tr><tr><td>Hexamethyldisilazane</td><td>Sigma Aldrich</td><td>379212-25ML</td><td>step 2.2.3 adhesion promoter</td></tr><tr><td>HF2LI Lock-in Amplifier</td><td>Zurich Instruments</td><td></td><td>subsection 9.2 part of the experimental setup</td></tr><tr><td>HF2TA Current amplifier</td><td>Zurich Instruments</td><td></td><td>subsection 9.2 part of the experimental setup</td></tr><tr><td>Isopropyl Alcohol</td><td>Fisher Scientific</td><td>A459-1</td><td>step 2.1.2 for cleaning, step 2.1.14 for rinsing wafer following SU-8 development, and step 6.3.1 for cleaning PDMS</td></tr><tr><td>IX81 fluorescence microscope</td><td>Olympus</td><td></td><td>step 11.2.3 for imaging</td></tr><tr><td>L-Glutamine Solution</td><td>Sigma Aldrich</td><td>G7513-20ML</td><td>step 8.1.2. part of media composition</td></tr><tr><td>M16878/1BFA 22 gauge wire</td><td>AWC</td><td>B22-1</td><td>subsection 7.5 for device fabrication</td></tr><tr><td>Magnesium chloride</td><td>Sigma Aldrich</td><td>208337-100G</td><td>step 8.1.2 part of electroporation buffer composition</td></tr><tr><td>MF 319 Developer</td><td>Kayaku Advanced Materials</td><td>10018042</td><td>step 2.2.9. photoresist developer</td></tr><tr><td>Microposit S1818 photoresist</td><td>Kayaku Advanced Materials</td><td>1136925</td><td>step 2.2.4 positive photoresist for electrode patterning</td></tr><tr><td>Microscope slides, 75 x 25 mm</td><td>VWR</td><td>16004-422</td><td>step 2.2.1 electrode soda lime glass substrate</td></tr><tr><td>Model 2350 High voltage amplifier</td><td>TEGAM</td><td>2350</td><td>step 9.2.5. part of the experimental setup</td></tr><tr><td>National Instruments LabVIEW</td><td>National Instruments</td><td></td><td>data acquisition</td></tr><tr><td>Needle, 30G x 1 in</td><td>BD Scientific</td><td>305128</td><td>step 10.1.1. part of the system priming</td></tr><tr><td>PA90 IC OPAMP Power circuit</td><td>Digi-key</td><td>598-1330-ND</td><td>Part of the custom circuit</td></tr><tr><td>Penicillin-Streptomycin</td><td>Sigma Aldrich</td><td>P4458-20ML</td><td>step 8.1.2. part of media composition</td></tr><tr><td>Plasmid pMAX-GFP</td><td>Lonza</td><td>VCA-1003</td><td>step 8.3.4. for intracellular delivery</td></tr><tr><td>Plastic tubing, 0.010&#039;&#039; x 0.030&quot;</td><td>VWR</td><td>89404-300</td><td>step 10.1.2. for system priming</td></tr><tr><td>Platinum targets</td><td>Kurt J. Lesker</td><td></td><td>subsection 4.2. for physical vapor deposition</td></tr><tr><td>Potassium chloride</td><td>Sigma Aldrich</td><td>P9333-500G</td><td>step 8.2.1. part of electroporation buffer composition</td></tr><tr><td>Pump 11 PicoPlus microfluidic syringe pump</td><td>Harvard Apparatus</td><td>MA1 70-2213</td><td>step 10.1.4. for system priming</td></tr><tr><td>PVD75 Physical vapor deposition system</td><td>Kurt J. Lesker</td><td></td><td>subsection 4.1. for physical vapor deposition</td></tr><tr><td>PWM32 Spinner System</td><td>Headway Research</td><td></td><td>steps 2.1.6 and 2.2.2. for substrate coating with photoresist</td></tr><tr><td>PX-250 Plasma treatment system</td><td>March Instruments</td><td></td><td>subsection 7.2 for PDMS and glass substrate bonding</td></tr><tr><td>SDG1025 Function/Waveform generator</td><td>Siglent</td><td></td><td>step 9.2.2. part of the experimental setup</td></tr><tr><td>Sodium hydroxide</td><td>Sigma Aldrich</td><td>S8045-500G</td><td>step 8.2.1. part of electroporation buffer composition</td></tr><tr><td>SU-8 2010 negative photoresist</td><td>Kayaku Advanced Materials</td><td>Y111053</td><td>step 2.1.7. for microfluidic channel patterning</td></tr><tr><td>SU-8 developer</td><td>Microchem</td><td>Y010200</td><td>step 2.1.12. for photoresist developing</td></tr><tr><td>Sucrose</td><td>Sigma Aldrich</td><td>S7903-1KG</td><td>step 8.2.1. part of electroporation buffer composition</td></tr><tr><td>Sylgard 184 elastomer kit</td><td>Dow Corning</td><td>3097358-1004</td><td>step 6.2.1&amp;nbsp; 10 : 1 mixture of PDMS polymer and hardening agent</td></tr><tr><td>Syringe, 1 ml</td><td>BD Scientific</td><td>309628</td><td>step 8.3.4. part of system priming</td></tr><tr><td>SZ61 Stereomicroscope System</td><td>Olympus</td><td></td><td>subsection 7.3. for channel and electrode alignment</td></tr><tr><td>Tissue Culture Treated T25 Flasks</td><td>Falcon</td><td>353108</td><td>step 8.1.2 for cell culturing</td></tr><tr><td>Titanium targets</td><td>Kurt J. Lesker</td><td></td><td>subsection 4.2. for physical vapor deposition</td></tr><tr><td>Transparency masks</td><td>CAD/ART Services</td><td></td><td>steps 2.1.9. and 2.2.7. for photolithography</td></tr><tr><td>Trichloro(1H,1H,2H,2H-perfluorooctyl)silane</td><td>Sigma Aldrich</td><td>448931-10G</td><td>step 6.1.2. for wafer silanization</td></tr><tr><td>Trypsin-EDTA solution</td><td>Sigma Aldrich</td><td>T4049-100ML</td><td>steps 8.1.3. and 8.3.1. for cell harvesting</td></tr></tbody></table>

the fabrication and operation of a continuous flow, micro-electroporation system with permeabilization detection

Current therapeutic innovations, such as CAR-T cell therapy, are heavily reliant on viral-mediated gene delivery. Although efficient, this technique is accompanied by high manufacturing costs, which has brought about an interest in using alternative methods for gene delivery. Electroporation is an electro-physical, non-viral approach for the intracellular delivery of genes and other exogenous materials. Upon the application of an electric field, the cell membrane temporarily allows molecular delivery into the cell. Typically, electroporation is performed on the macroscale to process large numbers of cells. However, this approach requires extensive empirical protocol development, which is costly when working with primary and difficult-to-transfect cell types. Lengthy protocol development, coupled with the requirement of large voltages to achieve sufficient electric-field strengths to permeabilize the cells, has led to the development of micro-scale electroporation devices. These micro-electroporation devices are manufactured using common microfabrication techniques and allow for greater experimental control with the potential to maintain high throughput capabilities. This work builds off a microfluidic-electroporation technology capable of detecting the level of cell membrane permeabilization at a single-cell level under continuous flow. However, this technology was limited to 4 cells processed per second, and thus a new approach for increasing the system throughput is proposed and presented here. This new technique, denoted as cell-population-based feedback control, considers the cell permeabilization response to a variety of electroporation pulsing conditions and determines the best-suited electroporation pulse conditions for the cell type under test. A higher-throughput mode is then used, where this 'optimal' pulse is applied to the cell suspension in transit. The steps for fabricating the device, setting up and running the microfluidic experiments, and analyzing the results are presented in detail. Finally, this micro-electroporation technology is demonstrated by delivering a DNA plasmid encoding for green fluorescent protein (GFP) into HEK293 cells.

Figure 4 highlights the operating principles behind the single-cell-level membrane permeabilization detection for a single pulse amplitude. Following the initiation of the electroporation experiment, the cell detection algorithm determines an optimal threshold for cell detection via a point-by-point, slope-based detection method. The system then continuously monitors (1) for a significant negative change in the measured electrical current, which is indicative of the entry of a cell. This is due to the insulative nature of the biological cell membrane, such that when the cell traverses through the electrode set, there is an instantaneous increase in impedance, resulting in a sharp, decrease in the measured current, allowing for consistent cell detection (2), which ultimately triggers the switch to the computer-controlled pulse application (4). The insulated cell displaces a volume of electrolyte between the electrodes, resulting in a drop in current that is proportional to the size of the cell. This change in current is denoted as &#916;IC (3). Immediately following &#916;IC calculation, the pre-determined, electrical pulse is administered (4) to the cell in transit. This instantaneous influx of energy introduces a brief sensing artifact into the system (grey box). Upon re-locking onto the signal, i.e., switching back to cell monitoring, (5) it is evident that the electroporation pulse permeabilized the cell membrane as the magnitude of current change due to the cell&#39;s presence between the electrode set drops upon exit (6). The difference in the two drops in current due to the cell&#39;s impedance magnitude pre/post electroporation pulse application is termed the permeabilization current and is denoted as &#916;IP. Once the cell exits the volume between the electrodes, the baseline stabilizes, and the system returns to cell detection mode (1). After a pre-determined number of cells are electroporated, the next highest energy electroporation pulse is tested (Refer to Table 1 for pulse settings). For each electroporation pulse tested, an average &#39;degree of membrane permeabilization&#39; is determined. This value is calculated as &#916;IP/&#916;IC. Once each pre-determined electroporation pulse is tested, the &#916;IP/&#916;IC&#160;is plotted against the applied electrical energy density (&#963;&#160;x E2 x t), where &#963; is the solution conductivity (S/cm), E is the electric field strength (kV/cm), and t is the pulse duration (ms). Refer to Figure 5 for the cell membrane permeabilization map for HEK293 cells used in this example.
Table 1: Electroporation pulse parameters. For this study, electroporation pulses were chosen such that the charge flux (&#963;&#215;E&#215;t) remains constant, where &#963; is the solution conductivity (S/cm), E is the electric field strength (kV/cm) and t is the pulse duration (ms). The result is a spectrum of the applied pulse electrical energy. Examples of the required duty cycle (d.c.) to achieve the specified pulse parameters are provided for both a 5&#215; and 10&#215; increase in the initial (single-cell-detection) flow rate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63103/Table 1.xlsx" target="_blank">Please click here to download this Table.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63103/63103fig04.jpg" /> 
Figure 4: Single cell membrane permeabilization - Algorithm operation. (Top) Electrical recording of a series of single-cell detections / pulse applications (indicated by the sharp spikes in current). (Bottom) System operation for the detection and pulsing of a single cell. (1) System is continuously sensing for a change in the current, via a point-by-point slope calculation. (2) A sharp decrease in the slope is detected, indicative of the entry of a cell between the electrodes and triggers the computer-controlled pulse application. (3) A current drop (&#916;IC) is determined and is proportional to the size of the cell. (4) The electroporation pulse is applied to the cell in transit, causing a sensing artifact in the electrical signal (grey box). (5) The lock-in amplifier switches back to cell monitoring as it re-locks into the cell in transit. (6) The cell exits the electrode set, causing another, smaller magnitude spike in current (&#916;IC &#62; (I6 - I5)). The difference in the impedance measurements is due to pore formation through the insulated cell membrane. This change in current is termed the permeabilization current (&#916;IP). The degree of cell membrane permeabilization is calculated (&#916;IP/&#916;IC). The baseline stabilizes and the system returns to detection mode (1). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63103/63103fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
A distinct correlation is observed between the applied electrical energy and the degree of cell membrane permeabilization (Figure 5), with the existence of a transition region where a substantial increase in the degree of cell membrane permeabilization occurs. To that end, a pulse with electrical energy that surpasses this transition region is selected for the high-throughput phase of the micro-electroporation process (Figure 6). In this experiment, the 1.8 kV/cm: 670 &#181;s pulse was determined as &#39;optimal&#39;. As was described in detail in subsection 10.3 of the protocol, the system flow rate is increased, and the function generator set to continuously output a pulse with a set pulse and duty cycle (refer to Table 1 for pulse settings for 1.5 &#181;L/min and 3.0 &#181;L/min flow rates) to ensure 1 pulse is applied to each cell in transit. In this study, the flow rate was increased by 5x, thus the pulse width was set to 50 ms (matching the cell transit time) at a duty cycle (d.c.) of 2.7%.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63103/63103fig05.jpg" /> 
Figure 5: HEK293 cell membrane permeabilization mapping -&#916;Ip/ &#916;Ic versus electrical energy. The electrical data (&#916;IP/ &#916;IC) is represented as the Mean &#177; SEM. Pulsing conditions (left to right)- 0.4 kV/cm : 3 ms, 0.8 kV/cm : 1.5 ms, 1.0 kV/cm : 1.2 ms, 1.2 kV/cm : 1 ms, 1.8 kV/cm : 0.67 ms, 2.4 kV/cm : 0.5 ms. A clear correlation is observed between the degree of cell membrane permeabilization and the electrical energy density of the applied pulse. For this round of experimentation, the 1.8 kV/cm: 0.67 ms pulsing condition was selected as the &#39;optimal&#39; electroporation pulse for the high-throughput module. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63103/63103fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63103/63103fig06.jpg" /> 
Figure 6: Cell-population-based feedback-controlled electroporation-Process workflow. To start, an initial flow rate is programmed (Q0) to allow for single-cell-level electrical interrogation. A programmable number of cells is pulsed at each pre-determined electroporation pulsing conditions (E0/t0 to EN/tN), with the applied electrical energy increasing with each iteration of electroporation pulse applications. Following the completion of the highest electrical energy pulse included in the study, EN/tN, the cell membrane permeabilization curve is plotted, and the optimal electroporation pulse is determined for the cell population under test. The system proceeds to high-throughput mode, where the flow rate is increased to Qthroughput, and the rate-limiting single-cell interrogation steps are omitted. The optimal pulse train will be continuously applied Eopt / topt at d.c.opt such that each cell in transit will receive a single electroporation pulse based on the cell transit time and the pulse width duty cycle (d.c.). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63103/63103fig6v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Following 24 h of post-electroporation recovery, the cells were imaged to determine the electro-transfection efficiency (eTE). As described in subsection 11.2 of the protocol, the eTE was determined as the total number of cells expressing GFP normalized to the total number of cells stained with DRAQ5. The eTE for the 1.8 kV/cm : 670 &#181;s pulse was determined to be ~70% (Figure 7A). To highlight the importance of the system to accurately map out the degree of cell membrane permeabilization and select a sufficiently high electroporation pulse energy when transitioning to the high-throughput mode, the 0.4 kV/cm: 3 ms pulse condition was also explored in terms of eTE (Figure 7B). In this case, the resulting eTE at 24 hours was less than 5%.
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/63103/63103fig07.jpg" /> 
Figure 7: electro-Transfection Efficiency-GFP expression at 24 h. HEK293 cells were incubated at 37 &#176;C for 24 hours following micro-electroporation experiments. All cells were stained with DRAQ5 (red), and the electro-transfection efficiency (eTE) was determined based on the ratio of cells expressing GFP (green) to the total cell number (red). Cell viability was not assessed as an outcome metric in this study. (A) Representative, stacked 4&#215; fluorescence image of HEK293 cells successfully transfected via a 1.8 kV/cm: 670 &#181;s pulse showing eTE of approximately 70%. (B)&#160;Representative, stacked 4&#215; fluorescence image of HEK293 cells unsuccessfully transfected via a 0.4 kV/cm : 3 ms pulse showing eTE &#60;&#60; 5%. Scale bar: 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63103/63103fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: 2-Dimensional CAD schematic. The micro-electroporation device consists of a straight, 100 &#181;m wide micro-channel with a 1 mm diameter inlet and a 3 mm diameter outlet. Each electrode trace is 100 &#181;m wide and the electrode set encompasses the electroporation region of the device, which is 300 &#181;m long. The 3-dimensional height of the micro-channel is controlled by the thickness of the photoresist. In this work, the height of the device was 20 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63103/Supplementary File 1-R2.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about The Fabrication and Operation of a Continuous Flow, Micro-Electroporation System with Permeabilization Detection at JoVE.com

The Fabrication and Operation of a Continuous Flow, Micro-Electroporation System with Permeabilization Detection

This protocol describes the microfabrication techniques required to build a lab-on-a-chip, microfluidic electroporation device. The experimental setup performs controlled, single-cell-level transfections in a continuous flow and can be extended to higher throughputs with population-based control. An analysis is provided showcasing the ability to electrically monitor the degree of cell membrane permeabilization in real-time.

Current therapeutic innovations, such as CAR-T cell therapy, are heavily reliant on viral-mediated gene delivery. Although efficient, this technique is ...

the-fabrication-operation-continuous-flow-micro-electroporation

Nanyang Technological University

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Bioengineering

2.7K Views.  Rutgers, The State University of New Jersey. Deze methode stelt onderzoekers in staat om optimale elektroporatiepulsingsomstandigheden voor het gewenste celtype te bepalen, terwijl de behoefte aan de traditionele empirische experimentele benadering wordt geminimaliseerd. Deze methode maakt gebruik van micro-elektroporatietechnieken om een schaalbaar microfluïdisch actuatieapparaat te vervaardigen dat in staat is om de mate van celmembraanpermeabilisatie tijdens het elektroporatieproces elektrisch te bewaken. Kishankumar Busha, een mast...

Video: The Fabrication and Operation of a Continuous Flow, Micro-Electroporation System with Permeabilization Detection

Microfabricated Platforms for Mechanically Dynamic Cell Culture

The ability to systematically probe in vitro cellular response to combinations of mechanobiological stimuli for tissue engineering, drug discovery or fundamental cell biology studies is limited by current bioreactor technologies, which cannot simultaneously apply a variety of mechanical stimuli to cultured cells. In order to address this issue, we have developed a series of microfabricated platforms designed to screen for the effects of mechanical stimuli in a high-throughput format. In this protocol, we demonstrate the fabrication of a microactuator array of vertically displaced posts on which the technology is based, and further demonstrate how this base technology can be modified to conduct high-throughput mechanically dynamic cell culture in both two-dimensional and three-dimensional culture paradigms.

In this protocol, we demonstrate the fabrication of a microactuator array of vertically displaced posts on which the technology is based, and how this base technology can be modified to conduct high-throughput mechanically dynamic cell culture in both two-dimensional and three-dimensional culture paradigms.

The ability to systematically probe in vitro cellular response to combinations of mechanobiological stimuli for tissue engineering, drug discovery or ...

Fabrication of Electrochemical-DNA Biosensors for the Reagentless Detection of Nucleic Acids, Proteins and Small Molecules

As medicine is currently practiced, doctors send specimens to a central laboratory for testing and thus must wait hours or days to 
receive the results. Many patients would be better served by rapid, bedside tests. To this end our laboratory and others have developed a versatile, reagentless 
biosensor platform that supports the quantitative, reagentless, electrochemical detection of nucleic acids (DNA, RNA), proteins (including antibodies) and small 
molecules analytes directly in unprocessed clinical and environmental samples. In this video, we demonstrate the preparation and use of several biosensors in this 
"E-DNA" class. In particular, we fabricate and demonstrate sensors for the detection of a target DNA sequence in a polymerase chain reaction mixture, an HIV-specific antibody and the drug cocaine. The preparation procedure requires only three hours of hands-on effort followed by an overnight incubation, and their use requires only minutes.

"E-DNA" sensors, reagentless, electrochemical biosensors that perform well even when challenged directly in blood and other complex matrices, have been adapted to the detection of a wide range of nucleic acid, protein and small molecule analytes. Here we present a general procedure for the fabrication and use of such sensors.

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Parallel-plate Flow Chamber and Continuous Flow Circuit to Evaluate Endothelial Progenitor Cells under Laminar Flow Shear Stress

The overall goal of this method is to describe a technique to subject adherent cells to laminar flow conditions and evaluate their response to well quantifiable fluid shear stresses1.
Our flow chamber design and flow circuit (Fig. 1) contains a transparent viewing region that enables testing of cell adhesion and imaging of cell morphology immediately before flow (Fig. 11A, B), at various time points during flow (Fig. 11C), and after flow (Fig. 11D). These experiments are illustrated with human umbilical cord blood-derived endothelial progenitor cells (EPCs) and porcine EPCs2,3.
This method is also applicable to other adherent cell types, e.g. smooth muscle cells (SMCs) or fibroblasts.
The chamber and all parts of the circuit are easily sterilized with steam autoclaving. In contrast to other chambers, e.g. microfluidic chambers, large numbers of cells (&gt; 1 million depending on cell size) can be recovered after the flow experiment under sterile conditions for cell culture or other experiments, e.g. DNA or RNA extraction, or immunohistochemistry (Fig. 11E), or scanning electron microscopy5. The shear stress can be adjusted by varying the flow rate of the perfusate, the fluid viscosity, or the channel height and width. The latter can reduce fluid volume or cell needs while ensuring that one-dimensional flow is maintained. It is not necessary to measure chamber height between experiments, since the chamber height does not depend on the use of gaskets, which greatly increases the ease of multiple experiments. Furthermore, the circuit design easily enables the collection of perfusate samples for analysis and/or quantification of metabolites secreted by cells under fluid shear stress exposure, e.g. nitric oxide (Fig. 12)6.

We are describing a method to subject adherent cells to laminar flow shear stress in a sterile continuous flow circuit. The cells' adhesion, morphology can be studied through the transparent chamber, samples obtained from the circuit for metabolite analysis and cells harvested after shear exposure for future experiments or culture.

The overall goal of this method is to describe a technique to subject adherent cells to laminar flow conditions and evaluate their response to well ...

Bacterial Detection &amp; Identification Using Electrochemical Sensors

Electrochemical sensors are widely used for rapid and accurate measurement of blood glucose and can be adapted for detection of a wide variety of analytes. Electrochemical sensors operate by transducing a biological recognition event into a useful electrical signal. Signal transduction occurs by coupling the activity of a redox enzyme to an amperometric electrode. Sensor specificity is either an inherent characteristic of the enzyme, glucose oxidase in the case of a glucose sensor, or a product of linkage between the enzyme and an antibody or probe.
Here, we describe an electrochemical sensor assay method to directly detect and identify bacteria. In every case, the probes described here are DNA oligonucleotides. This method is based on sandwich hybridization of capture and detector probes with target ribosomal RNA (rRNA). The capture probe is anchored to the sensor surface, while the detector probe is linked to horseradish peroxidase (HRP). When a substrate such as 3,3',5,5'-tetramethylbenzidine (TMB) is added to an electrode with capture-target-detector complexes bound to its surface, the substrate is oxidized by HRP and reduced by the working electrode. This redox cycle results in shuttling of electrons by the substrate from the electrode to HRP, producing current flow in the electrode.

Bacterial Detection & Identification Using Electrochemical Sensors

We describe an electrochemical sensor assay method for rapid bacterial detection and identification. The assay involves a sensor array functionalized with DNA oligonucleotide capture probes for ribosomal RNA (rRNA) species-specific sequences. Sandwich hybridization of target rRNA with the capture probe and a horseradish peroxidase-linked DNA oligonucleotide detector probe produces a measurable amperometric current.

Electrochemical  sensors are widely used for rapid and accurate measurement of blood glucose and  can be adapted for detection of a wide variety of ...

Microscale Vortex-assisted Electroporator for Sequential Molecular Delivery

Electroporation has received increasing attention in the past years, because it is a very powerful technique for physically introducing non-permeant exogenous molecular probes into cells. This work reports a microfluidic electroporation platform capable of performing multiple molecule delivery to mammalian cells with precise and molecular-dependent parameter control. The system&#8217;s ability to isolate cells with uniform size distribution allows for less variation in electroporation efficiency per given electric field strength; hence enhanced sample viability. Moreover, its process visualization feature allows for observation of the fluorescent molecular uptake process in real-time, which permits prompt molecular delivery parameter adjustments in situ for efficiency enhancement. To show the vast capabilities of the reported platform, macromolecules with different sizes and electrical charges (e.g., Dextran with MW of 3,000 and 70,000 Da) were delivered to metastatic breast cancer cells with high delivery efficiencies (&#62;70%) for all tested molecules. The developed platform has proven its potential for use in the expansion of research fields where on-chip electroporation techniques can be beneficial.

A microfluidic vortex assisted electroporation platform was developed for sequential delivery of multiple molecules into identical cell populations with precise and independent dosage control. The system&#8217;s size based target cell purification step preceding electroporation aided to enhance molecular delivery efficiency and processed cell viability.

Electroporation has received increasing attention in the past years, because it is a very powerful technique for physically introducing non-permeant ...

Label-free Isolation and Enrichment of Cells Through Contactless Dielectrophoresis

Dielectrophoresis (DEP) is the phenomenon by which polarized particles in a non-uniform electric field undergo translational motion, and can be used to direct the motion of microparticles in a surface marker-independent manner. Traditionally, DEP devices include planar metallic electrodes patterned in the sample channel. This approach can be expensive and requires a specialized cleanroom environment. Recently, a contact-free approach called contactless dielectrophoresis (cDEP) has been developed. This method utilizes the classic principle of DEP while avoiding direct contact between electrodes and sample by patterning fluidic electrodes and a sample channel from a single polydimethylsiloxane (PDMS) substrate, and has application as a rapid microfluidic strategy designed to sort and enrich microparticles. Unique to this method is that the electric field is generated via fluidic electrode channels containing a highly conductive fluid, which are separated from the sample channel by a thin insulating barrier. Because metal electrodes do not directly contact the sample, electrolysis, electrode delamination, and sample contamination are avoided. Additionally, this enables an inexpensive and simple fabrication process.
cDEP is thus well-suited for manipulating sensitive biological particles. The dielectrophoretic force acting upon the particles depends not only upon spatial gradients of the electric field generated by customizable design of the device geometry, but the intrinsic biophysical properties of the cell. As such, cDEP is a label-free technique that avoids depending upon surface-expressed molecular biomarkers that may be variably expressed within a population, while still allowing characterization, enrichment, and sorting of bioparticles.
Here, we demonstrate the basics of fabrication and experimentation using cDEP. We explain the simple preparation of a cDEP chip using soft lithography techniques. We discuss the experimental procedure for characterizing crossover frequency of a particle or cell, the frequency at which the dielectrophoretic force is zero. Finally, we demonstrate the use of this technique for sorting a mixture of ovarian cancer cells and fluorescing microspheres (beads).

Contactless dielectrophoresis (cDEP) achieves sorting and enrichment of particles via their intrinsic dielectric properties. Fluidic electrode channels replace metallic electrodes traditional to DEP, suiting cDEP to non-damaging sterile characterization and sorting of biological particles. We demonstrate how to prepare a cDEP microdevice and conduct cell characterization and sorting experiments.

Dielectrophoresis (DEP) is the phenomenon by which polarized particles in a non-uniform electric field undergo translational motion, and can be used to ...

Medicine

Fast Enzymatic Processing of Proteins for MS Detection with a Flow-through Microreactor

The vast majority of mass spectrometry (MS)-based protein analysis methods involve an enzymatic digestion step prior to detection, typically with trypsin. This step is necessary for the generation of small molecular weight peptides, generally with MW &#60; 3,000-4,000 Da, that fall within the effective scan range of mass spectrometry instrumentation. Conventional protocols involve O/N enzymatic digestion at 37 &#186;C. Recent advances have led to the development of a variety of strategies, typically involving the use of a microreactor with immobilized enzymes or of a range of complementary physical processes that reduce the time necessary for proteolytic digestion to a few minutes (e.g., microwave or high-pressure). In this work, we describe a simple and cost-effective approach that can be implemented in any laboratory for achieving fast enzymatic digestion of a protein. The protein (or protein mixture) is adsorbed on C18-bonded reversed-phase high performance liquid chromatography (HPLC) silica particles preloaded in a capillary column, and trypsin in aqueous buffer is infused over the particles for a short period of time. To enable on-line MS detection, the tryptic peptides are eluted with a solvent system with increased organic content directly in the MS ion source. This approach avoids the use of high-priced immobilized enzyme particles and does not necessitate any aid for completing the process. Protein digestion and complete sample analysis can be accomplished in less than ~3 min and ~30 min, respectively.

A quick protocol for proteolytic digestion with an in-house built flow-through tryptic microreactor coupled to an electrospray ionization (ESI) mass spectrometer is presented. The fabrication of the microreactor, the experimental setup and the data acquisition process are described.

The vast majority of mass spectrometry (MS)-based protein analysis methods involve an enzymatic digestion step prior to detection, typically with trypsin. ...

Dry Film Photoresist-based Electrochemical Microfluidic Biosensor Platform: Device Fabrication, On-chip Assay Preparation, and System Operation

In recent years, biomarker diagnostics became an indispensable tool for the diagnosis of human disease, especially for the point-of-care diagnostics. An easy-to-use and low-cost sensor platform is highly desired to measure various types of analytes (e.g., biomarkers, hormones, and drugs) quantitatively and specifically. For this reason, dry film photoresist technology - enabling cheap, facile, and high-throughput fabrication - was used to manufacture the microfluidic biosensor presented here. Depending on the bioassay used afterwards, the versatile platform is capable of detecting various types of biomolecules. For the fabrication of the device, platinum electrodes are structured on a flexible polyimide (PI) foil in the only clean-room process step. The PI foil serves as a substrate for the electrodes, which are insulated with an epoxy-based photoresist. The microfluidic channel is subsequently generated by the development and lamination of dry film photoresist (DFR) foils onto the PI wafer. By using a hydrophobic stopping barrier in the channel, the channel is separated into two specific areas: an immobilization section for the enzyme-linked assay and an electrochemical measurement cell for the amperometric signal readout.
The on-chip bioassay immobilization is performed by the adsorption of the biomolecules to the channel surface. The glucose oxidase enzyme is used as a transducer for electrochemical signal generation. In the presence of the substrate, glucose, hydrogen peroxide is produced, which is detected at the platinum working electrode. The stop-flow technique is applied to obtain signal amplification along with rapid detection. Different biomolecules can quantitatively be measured by means of the introduced microfluidic system, giving an indication of different types of diseases, or, in regard to therapeutic drug monitoring, facilitating a personalized therapy.

A microfluidic biosensor platform was designed and fabricated using low-cost dry film photoresist technology for the rapid and sensitive quantification of various analytes. This single-use system allows for the electrochemical readout of on-chip-immobilized enzyme-linked assays by means of the stop-flow technique.

In recent years, biomarker diagnostics became an indispensable tool for the diagnosis of human disease, especially for the point-of-care diagnostics. An ...

Amplification of Escherichia coli in a Continuous-Flow-PCR Microfluidic Chip and Its Detection with a Capillary Electrophoresis System

Polymerase chain reaction (PCR) is a traditional method employed for the amplification of a target gene that has played an important role in biomolecular diagnostics. However, traditional PCR is very time-consuming because of the low-temperature variation efficiency. This work proposes a continuous-flow-PCR (CF-PCR) system based on a microfluidic chip. The amplification time can be greatly reduced by running the PCR solution into a microchannel placed on heaters set at different temperatures. Moreover, as capillary electrophoresis (CE) is an ideal way to differentiate positive and false-positive PCR products, a CE system was built to achieve efficient separation of the DNA fragments. This paper describes the process of amplification of Escherichia coli (E. coli) by the CF-PCR system built in-house and the detection of the PCR products by CE. The results demonstrate that the target gene of E. coli was successfully amplified within 10 min, indicating that these two systems can be used for the rapid amplification and detection of nucleic acids.

Amplification of Escherichia coli in a Continuous-Flow-PCR Microfluidic Chip and Its Detection with a Capillary Electrophoresis System

This protocol describes how to build a continuous-flow-polymerase chain system based on a microfluidic chip and how to build a capillary electrophoresis system in the lab. It presents a simple method for the analysis of nucleic acids in the lab.

Polymerase chain reaction (PCR) is a traditional method employed for the amplification of a target gene that has played an important role in biomolecular ...

TEEI Method for Electroporation Analysis and Delivery Prediction in A431 Cells

Porous Substrate-Based Electroporation with Transepithelial Electrical Impedance Monitoring

Porous substrate electroporation (PSEP) is an emerging method of electroporation that provides high throughput and consistent delivery. Like many other types of intracellular delivery, PSEP relies heavily on fluorescent markers and fluorescent microscopy to determine successful delivery. To gain insight into the intermediate steps of the electroporation process, a PSEP platform with integrated transepithelial electrical impedance (TEEI) monitoring was developed. Cells are cultured in commercially available inserts with porous membranes. After a 12 h incubation period to allow for the formation of a fully confluent cell monolayer, the inserts are placed in transfection media located in the wells of the PSEP device. The cell monolayers are then subjected to a user-defined waveform, and delivery efficiency is confirmed through fluorescent microscopy. This workflow can be significantly enhanced with TEEI measurements between pulsing and fluorescent microscopy to collect additional data on the PSEP process, and this additional TEEI data is correlated with delivery metrics such as delivery efficiency and viability. This article describes a protocol for performing PSEP with TEEI measurements.

Author Spotlight: Optimizing Porous Substrate Electroporation Through Micro and Nanochannels for Enhanced Monitoring and Intermediate Stage Characterization

Porous substrate electroporation (PSEP) pairs consistent, high throughput delivery with high cell viability. Introduction of transepithelial electrical impedance (TEEI) measurements provides insight into the intermediate processes of PSEP and allows for label-free delivery. This article discusses a method for performing PSEP delivery experiments and TEEI measurement analysis simultaneously.

Porous substrate electroporation (PSEP) is an emerging method of electroporation that provides high throughput and consistent delivery. Like many other ...

The Fabrication and Operation of a Continuous Flow, Micro-Electroporation System with Permeabilization Detection

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