Comparison of DNA Delivery Methods

Research will be limited to:
- Application: DNA delivery
- Cell Type: Mammalian cells
- Method: Lipofectamine transfection vs Electroporation
- Evaluation Criteria: Efficiency, Toxicity, Cost
- Scope: Protocol comparison

Strategy: I will conduct a comparative analysis focusing on the quantitative performance metrics of Lipofectamine-mediated chemical transfection versus physical electroporation for DNA delivery in mammalian systems. The search will prioritize head-to-head studies that evaluate transfection efficiency (percentage of positive cells), cell viability (toxicity), and operational costs. I will also look for protocol-specific nuances, such as the impact of cell type (adherent vs. suspension) and DNA concentration on these outcomes.
Queries:
- comparison Lipofectamine vs electroporation DNA delivery mammalian cells efficiency toxicity
- cost-benefit analysis chemical transfection vs electroporation mammalian cell culture
- Lipofectamine 3000 vs Neon electroporation system DNA transfection efficiency toxicity
- protocol optimization DNA delivery mammalian cells Lipofectamine vs electroporation
- quantitative comparison DNA delivery methods mammalian cells viability cost efficiency

Executing 5 concurrent searches:
- comparison Lipofectamine vs electroporation DNA delivery mammalian cells efficiency toxicity
- cost-benefit analysis chemical transfection vs electroporation mammalian cell culture
- Lipofectamine 3000 vs Neon electroporation system DNA transfection efficiency toxicity
- protocol optimization DNA delivery mammalian cells Lipofectamine vs electroporation
- quantitative comparison DNA delivery methods mammalian cells viability cost efficiency

Found 8 papers. Key excerpts:

Comparative Efficiency for in vitro Transfection of Goat Undifferentiated Spermatogonia Using Lipofectamine Reagents and Electroporation. — Wilkister Nabulindo Nakami et al., 2022

Spermatogonial stem cells are the precursor cells that produce spermatozoa. Their ability to proliferate in vitro and reestablish spermatogenesis in recipient testes make them ideal for genetic manipulation. 1 Utilization of SSC in the production of genome-edited animals is increasingly becoming an area of research interest. 2 , 3 Intra-testicular transplantation of SSC that carries desired transgenes, can result in the production of donor genotype-derived transgenic spermatozoa. 4 , 5 Transfection of genes into the germline stem cells shortens the generational interval for the production of transgenic animals. 6
Previously, viral vectors and carrier molecules transfection techniques have been deployed for gene transfer. 4 Despite successful reports of efficient viral transduction, the method risks viral gene transcription and insertion into the host genome. 6 , 7 Non-viral methods, use of liposome carriers -lipofection-and electroporation have been used to transfer the foreign DNA into testicular cells and SSC cells in sheep and bovine. 6 , 8 , 9
Lipofection has been used to transfer foreign DNA into spermatogonial stem cells. 8 The DNA-lipid complex is introduced into the cell culture wells, and the liposome transports the foreign DNA into the cells. 10 Bovine SSC have successfully been transfected with eGFP plasmid lipofectamine reagent. 8
The electroporation process involves the exposure of cells to high-intensity electric field pulses. This creates temporary pores in the cell membrane through which exogenous DNA diffuse into the cytoplasm, or nucleus. 6 , 11 Electroporation of sheep testicular cells 6 and porcine spermatogonial stem cells has been reported. 9
There is paucity of data on the comparative efficiency of various transfection methods in goat SSC. Here, we compare the efficiency of eGFP gene transfer through lipofection and electroporation methods.
Ten prepubertal Galla bucks aged 3–6 months were obtained from the International Livestock Research Institute (ILRI) farm. Prepubertal bucks are ideal for spermatogonial stem cells isolation. 12 The study was approved by the Institutional Animal Care and Use Committees (IACUC Ref no: 2018–15) of the International Livestock Research Institute

(ILRI) in Kenya and the University of Nairobi (Ref: FVM BAUEC/2019/243). The handling of the animals adhered strictly to animal welfare considerations by using local anaesthesia recommended for castration and pain management post-surgery.
The prepubertal goats were castrated, and the testes were transported on ice to ILRI laboratory within three hours in Hanks Balanced Salt Solution (HBSS, Gibco TM Grand Island, New York, USA). The transport media was supplemented with 100 IUmL −1 penicillin (Sigma-Aldrich St Louis, Missouri USA) and 100 ug/mL streptomycin (Sigma-Aldrich St Louis, Missouri USA). A testicular cell isolation procedure was performed as described by Oatley et al with minor adaptation. 13 Briefly, testes were washed in HBSS and gently disentangled to expose seminiferous tubules. In a water bath at 37°C, 150–200 mg of testicular tissue was digested in 0.25mg/mL of collagenase type IV enzyme (Gibco TM Grand Island, New York, USA) and 7 mg/mL DNase 1(ROCHE Sandhoferstrasse,Mannheim Germany) in HBSS for 5–7 minutes. To eliminate interstitial cells, gravity sedimentation of seminiferous tubules was performed on ice and the supernatant discarded. The sedimentation on ice and washing process was repeated five times.
The seminiferous tubules were incubated in 0.25% Trypsin/0.04 EDTA (Gibco TM Grand Island, New York, USA) and DNase 7 mg/mL in a 37°C water bath for 30 minutes. The trypsin reaction was terminated by adding Foetal Bovine Serum (FBS, Gibco TM Brazil). The cell suspension was passed through a 40μm cell-strainer and washed twice in HBSS through centrifugation. Somatic cells were removed through differential plating on gelatin-coated plates. The somatic cells are attached to the plate bottom. The floating SSC-rich population was collected and washed through centrifugation. The cell pellet was resuspended in Stempro serum-free medium (Gibco™ Grand Island, New York, USA) containing bovine serum

albumin with Stempro nutrient supplement (Gibco™ Grand Island, New York, USA) and seeded on 96-well laminin-coated cell culture plates for four days. The medium was refreshed on alternate days. Trypan blue exclusion staining was performed to evaluate the viability of primary isolated cells. The total number of cells present in the suspension was determined using a hemocytometer observed under a light microscope.
Lipofectamine 2000 (Invitrogen by Life Technologies, Renfrew, United Kingdom) and Lipofectamine Stem (Invitrogen by Life Technologies, Renfrew, UK) reagents were used for transfection. Commercially available Enhanced Green Fluorescent Protein (eGFP) plasmid DNA (NepaGene, Clontech, Japan) was transferred to the cells. The protocol described by Tajik et al was adopted with minor modifications. Briefly, the cells were recovered from the bottom of the culture wells, and clumps were broken up by pipetting gently. Cells were washed and resuspended in Dulbecco's Modified Eagle Medium (DMEM, Sigma-Aldrich St Louis, Missouri USA) serum and antibiotic-free medium, then plated into 24-well cell culture plates (Corning ® , Glendale, Arizona USA).
There were four experimental groups, treated as follows ( Table 1 ). Table 1 Description of Culture Experimental Groups Group Treatment 1 Culture of SSC with DNA + Lipofectamine 2000 complex 2 Culture of SSC with DNA + lipofectamine Stem complex 3 Culture of SSC with DNA only 4 Culture of SSC without any other inclusion
Description of Culture Experimental Groups
To transfer eGFP into goat SSC, lipofectamine 2000 DNA transfection reagent and lipofectamine stem reagent liposomal carriers were used. After the four-day culture, the SSC clumps were recovered and plated at a concentration of 0.5–2×10 5 cells in 500μL of serum-free and antibiotic-free growth medium 24 hours prior to transfection. On the day of transfection, the cells were detached from the bottom of the plate by gently pipetting and washing through centrifugation at 600xg for 7 minutes at 4°C. The cells were then re-suspended in an antibiotic-free, serum-free DMEM medium and

Improved delivery of Cas9 protein/gRNA complexes using lipofectamine CRISPRMAX — Xin Yu et al., 2016

The use of cell-penetrating peptides (CPPs) facilitates the delivery of active proteins into cells (Copolovici et al.
Systematic design of experiments used for screening transfection reagents The screening of transfection reagents was conducted in a 96-well format. 1 day prior to transfection, six commonly used cell lines, A549, HEK293, HeLa, HepG2, MCF-7, and U2OS, were seeded in 96-well plates at 10,000–20,000 cells per well. On the day of transfection, a master mix of Cas9 protein and HPRT1 gRNA was prepared in Opti-MEM medium and incubated for 5 min at 25 °C to form the Cas9 RNPs. The amount of Cas9 RNPs was held constant at 40 ng GeneArt Platinum Cas9 nuclease and 8.5 ng gRNA per well. On the other hand, the amount of each transfection reagent, which was also prepared in Opti-MEM medium, varied from 0.1, 0.2, 0.4 and 0.6 µl per well. Lipofectamine 3000 and Lipofectamine RNAiMAX served as controls. The Cas9 RNPs in Opti-MEM medium were added to the transfection reagents diluted in Opti-MEM medium. The mixture was incubated at 25 °C for 10–15 min to form the Cas9 RNPs and transfection reagent complexes, followed by addition to the cells. After incubation for 48 h, the cells were lysed and percentage of Indel (insertion and deletion) was measured by GeneArt Genomic Cleavage Detection Kit. The experimental data was then analyzed using JMP, Version 11. SAS Institute Inc. (Cary, NC, USA). Cell transfection in a 24-well plate using CRISPRMAX One day prior to transfection, adherent cells were plated onto 24-well plates at 0.4 to 1.5 × 10 For transfection of human iPSC, the cells were treated with TrypLE and plated onto Geltrex-coated 24-well plates at 40,000 cells per well, leading to approx. 30–40 % confluence at the

time of transfection. One µg GeneArt Platinum Cas9 nuclease, 250 ng gRNA and 6 µl Cas9 Plus reagent were used to prepare the Cas9 RNPs instead, while the amount of Lipofectamine CRISPRMAX reagent remained constant at 1.5 µl. At around 6 h post-transfection, the media containing the transfection reagent was removed and replaced with fresh Essential 8 Medium. The cells were analyzed at 48 h post-transfection. A ‘reverse’ transfection protocol was used to transfect MCF-7 and HepG2 cells. In this case, the Cas9 RNPs and Lipofectamine CRISPRMAX reagent were prepared in two separate tubes as described above with 500 ng GeneArt Platinum Cas9 nuclease, 125 ng gRNA, and 1 µl Cas9 Plus reagent in Tube-1, and 1.5 µl Lipofectamine CRISPRMAX reagent in Tube-2, respectively. The Cas9 RNPs solution was then transferred to the Lipofectamine CRISPRMAX solution. Upon vortexing, the mixture was incubated at 25 °C for 10–15 min to form the Cas9 RNPs and Lipofectamine CRISPRMAX complexes. Meanwhile, MCF-7 and HepG2 cells were detached with TrypLE and counted, followed by seeding at 2 × 10 For Neon electroporation, adherent cells were detached from culture dishes and counted. In general, 10 Genomic cleavage assay The genomic modification efficiency was determined by GeneArt Genomic Cleavage Detection kit as described in the manual. The targeting gRNA sequence: 5′-gcatttctcagtcctaaaca Generation of a disrupted EmGFP stable cell line for homologous recombination assay GripTite HEK293 stable cells expressing EmGFP were prepared via the Jump-In system as described in the manual (Thermo Fisher Scientific). To generate a disrupted EmGFP mutant stable cell line, 1.5 µg of GeneArt Platinum Cas9 nuclease was associated with 300 ng gRNA targeting the 5′-ctcgtgaccaccttcaccta To create homologous recombination assays, a gRNA targeting the 5′-

gaagcactgcacgccgtaggtgg-3′sequence within the disrupted EmGFP reporter gene (T2) was designed and synthesized. This gRNA only recognized the disrupted EmGFP gene but not the wild type EmGFP gene. One day prior to transfection, the cells were seeded on a 24 well plate at 10
Identification of Lipofectamine CRISPRMAX To identify the best transfection reagents for delivery of Cas9 RNPs, initially we compared several commercially available protein transfection reagents, including Lipofectamine 2000, Lipofectamine 3000, Lipofectamine RNAiMAX, Lipofectamine MeassengerMax, TurboFect, and Xfect Protein transfection reagent. Cas9 protein and HPRT1 gRNA were transfected into HEK293 and HCT116 cell lines according to manufacturer’s protocol. Upon 48 h post-transfection, the cells were harvested to analyze the genome modification efficiencies. As depicted in Fig.  As shown in Fig.  Time course of Cas9 RNPs activity Upon identification of Lipofectamine CRISPRMAX as the best transfection reagent, we determined the functional activity of Cas9 RNPs by examining the times required for complexation of Cas9 protein with gRNA (Cas9 RNPs) and Cas9 RNPs with Lipofectamine CRISPRMAX in A549, HEK293, and HeLa cells. As shown in Fig.  Main factors regulating transfection efficiency Next, we examined the key factors that governed the transfection efficiency by varying the dose of transfection reagent, the amount of Cas9 RNPs, and cell density. Six commonly used cell lines were transfected with increasing amount of Cas9 RNPs, followed by genome cleavage assay. As shown in Fig.  Low cell toxicity of Lipofectamine CRISPRMAX We then scaled up to 24 wells to test a set of 23 cell lines, including a variety of adherent and suspension cells from different species. The morphologies of more than a dozen adherent cell lines were recorded prior to transfection and at 48 h post-transfection (Supplementary Fig. 1). Most of the cells looked healthy under the microscope with examples shown in Fig.  Comparison of Lipofectamine CRISPRMAX to electroporation Suspension

In situ electroporation of mammalian cells through SiO 2 thin film capacitive microelectrodes. — M Maschietto et al., 2021

A variety of methods are routinely employed to transfect mammalian cells, i.e. to introduce DNA molecules across the bi-lipid plasma membrane that physiologically separates the intracellular cytoplasm from the extracellular fluid. Among physical methods, electroporation is the most used 1 . Traditionally, it is performed in a cuvette on a population of cells in suspension between two facing metal electrodes (e.g. platinum). The field applied between the two electrodes has the purpose to cause the voltage across the cell membrane to change abruptly reaching a critical threshold (i.e., between 250 and 500 mV), thus leading to the opening of transient pores 2 , 3 and providing a pathway to exogenous molecules to enter the cytoplasm 4 – 6 . In alternative, there is a growing interest for methods of in-situ electroporation, which avoid the potentially detrimental enzymatic and mechanical treatments to detach cells growing in adhesion on a solid substrate. In particular, recent developments include miniaturized and on-chip integrated microsystems that selectively operate in-situ electroporation on subpopulations of cells in adhesion 7 – 9 . With respect to bulk electroporation, these devices enable to operate spatially confined transfections of preselected subpopulations of cells 10 , 11 or single cells 12 , 13 . In one approach cells are grown on top of microelectrode arrays integrated in the solid culture substrate. Selectivity is achieved because cells are targeted by stimulating the corresponding microelectrodes, which also enable serial transfection by repeating the electroporation procedure on the same target cells 13 . In an alternative configuration, the array of microelectrodes is approached from the top of the cell culture 14 , 15 . In general, in-situ electroporation allows for proper tuning of electroporation parameters, favouring transfection efficiency mainly thanks to the reduced intensity of the electric fields and their homogeneity across cells with respect to suspensions 16 – 19 . Moreover, microfabrication offers additional opportunities, e.g. to modulate the DNA concentration in the proximity of target cells by microfluidics 20 , by electrophoresis 21 , 22 or by coating the surface with cationic polymers 23 , 24 . Noteworthy, with respect to electrodes design, 3D hollow nanoelectrodes were recently demonstrated to further reduce electroporation voltages 25 , 26 . In

conclusion, on-chip microdevices based on integrated active sites are promising tools for selective in-situ electroporation and transfection of adherent cells, and has been proven with a variety of cell types including primary cultures 27 – 29 and transfection molecules comprising DNA for exogenous gene expression or short oligonucleotides for RNA interference 27 , 30 – 33 . On the other hand, improvements are still necessary in terms of control over the field applied to the cells across a culture to optimize transfection reliability and efficiency and in terms of microelectrode stability for serial transfections or re-use. Metal-electrolyte interfaces, for instance, can suffer from non-linearities in the current–voltage relationship due to mixed faradaic and non-faradaic behaviour when subjected to large voltages (e.g., 1 V or above) 34 . This can be the case during field application in routinely used electroporation protocols. In this work, we demonstrate that in-situ electroporation of mammalian cells can be achieved efficiently through a SiO 2 thin film that, similarly to other oxide films, is known to largely suppress faradaic currents 35 – 40 . We validate the method by demonstrating high transfection efficiency and cell selectivity for exogenous gene expression and by performing double serial transfections of adherent mammalian cells, including primary neurons.
The basic structure and operation of the electroporation chip with integrated an array of 32 × 2 SiO 2 thin film capacitive microelectrodes is described in Fig. 1 (see also ‘The equivalent circuit’ in the Methods section) 41 . Each stimulation site consists of an octagonal microelectrode formed by highly conductive p-type silicon coated with a 15 nm thick layer of SiO 2 (Fig. 1 a–d). The electrolyte/SiO 2 /Si stack forms a capacitor, that is charged by applying voltage transients to the p-doped Si line realized on the substrate. The associated ionic current flowing into the cell-substrate cleft region causes a transient, non-null and localized extracellular field to which cells growing on top of the microelectrode are exposed (Fig. 1 e–f). Each microelectrode is individually addressable. Figure 1 Cells on SiO 2 thin-film capacitive microelectrodes . ( a ) Top

view of the electroporation microchip including the cell culture plastic chamber glued on a ceramic package providing mechanical support and electrical contacts to the voltage generator. Scale bar: 1 cm. ( b ) Magnification of three octagonal microelectrodes out of sixty-four (organized in two linear arrays as shown in Fig. 2 ) and with CHO-K1 cells growing in adhesion on top of them. Scale bar: 10 μm. ( c , e ) Drawing of the cell-chip interface and microelectrode structure (cross-section). The cell grows in adhesion on top of the SiO 2 thin film from which it is separated by a narrow (i.e., in the range of a few tens of nanometers) cleft occupied by electrolyte. The highly conductive p-type silicon forms a capacitor together with the electrolyte and the oxide dielectric film separating the two. Insulation between microelectrodes is ensured by SiO 2 . ( d ) Equivalent electrical circuit of the cell-capacitive microelectrode (CME) system. The cell is described by two compartments of lipid membrane, one for the region of adhesion (with resistance R AM and capacitance C AM ), the other one for the free portion exposed to the bulk electrolyte (R FM and C FM ). R cleft represents the resistance sealing the cleft from the bulk electrolyte. C Ox is the capacitance of the oxide and V S is the voltage applied to one face of the dielectric through the p-type Si. Accordingly, upon a change of V s two potentials, Δ V F and Δ V A , develop across the two compartments of the cell membrane, eventually causing electroporation. V B : is the bias voltage applied to the bulk n-type silicon. ( f ) Stimulation settings. The bath electrolyte is kept at ground potential through a Ag/AgCl electrode and the bulk n-type Si at a bias potential V B . A voltage source, V S , modulates the p-type Si. A typical stimulation waveform consisting of a repetition of sawtooth waves is sketched at the bottom.
Cells on SiO 2 thin-film capacitive microelectrodes . ( a ) Top view of the electroporation microchip including the cell culture plastic chamber glued on a ceramic package providing mechanical support and electrical contacts to the voltage generator.

Transfection by Electroporation — Huntington Potter et al., 2017

INTRODUCTION

Electroporation-the use of high-voltage electric shocks to introduce DNA into cells-is a procedure that is gaining in popularity for standard gene transfer and that also allows the generation of genetically modified mice (Potter, 1988). It can be used with most cell types, yields a high frequency of both stable transformation and transient gene expression, and, because it requires fewer steps, can be easier than alternative techniques.
Basic Protocol 1 describes the electroporation of mammalian cells, including ES cells for the generation of transgenic and knock-out/knock-in mice. The in vivo protocol (Basic Protocol 2) describes the use of electroporation to deliver plasmid DNA to muscle and skin.

Harvest cells by centrifuging 5 min as in step 2.

1. Resuspend cells at 1 x 10 7 /ml in electroporation buffer at 0degC for permanent transfection.
   Higher concentrations of cells (up to 8 x 10 7 ) may be used for transient expression.
2. Transfer 0.5-ml aliquots of the cell suspension into the desired number of electroporation cuvettes set on ice.
   Add DNA and electroporate the cells 7. Add DNA to the cell suspension in the cuvettes on ice.
   For stable transformation, DNA should be linearized by cleavage with a restriction enzyme that cuts in a nonessential region, and then purified by phenol extraction and ethanol precipitation (UNIT 10.1;Moore, 1993 8. Mix DNA/cell suspension by holding the cuvette on the two "window sides" and flicking the bottom. Incubate 5 min on ice.
3. Place cuvette in the holder in the electroporation apparatus (at room temperature) and shock one or more times at the desired voltage and capacitance settings.
   The number of shocks and the voltage and capacitance settings will vary depending on the cell type and should be optimized (see Critical Parameters).
4. After electroporation, return cuvette containing cells and DNA to ice for 10 min.
   Culture and harvest the transfected cells 11. Dilute transfected cells 20-fold in nonselective complete medium and rinse cuvette with this same medium to remove all transfected cells.
   12a. For stable transformation: Grow cells 48 hr (about two generations) in nonselective medium,

then transfer to antibiotic-containing medium.
Selection conditions will vary with cell type. For example, neo selection generally requires ss400 ug/ml G418 in the medium. XGPRT selection requires 1 ug/ml mycophenolic acid, 250 ug/ml xanthine, and 15 ug/ml hypoxanthine in the medium (see UNIT 10.17A;Kingston, 1994).
It is often convenient to plate adherent cells at limiting dilution (see UNIT 3.14;Kruisbeek, 1997) immediately following the shock, or suspension cells at the time of antibiotic addition.
12b. For transient expression: Incubate cells 50 to 60 hr, then harvest cells for transient expression assays.
Transfected cells can be visualized by standard transient expression assays (Brasier and Fortin, 1995;Kingston et al., 1995;Kain and Ganguly, 1998).

BASIC PROTOCOL 2

ELECTROPORATION INTO MUSCLE OR SKIN

Electroporation has been used successfully to deliver plasmid DNA to a variety of tissues in vivo ). Because of its physical nature, electroporation can be applied to practically any cell or tissue. Plasmid DNA in the appropriate diluent is injected into the tissue. Electrodes are then placed around the injection site and the cells within the tissue are subjected to a high-voltage electrical pulse of defined magnitude and length. The animals are then allowed to recover and the tissue is evaluated at specified time points following delivery. Factors that can be varied to optimize electroporation effectiveness are pulse width, number, amplitude, and electrode configuration.

Materials

Linear or supercoiled, purified DNA preparation (see annotation to step 1) DNA amplification kit (e.g., Qiagen, cat. no. 12991) Animals to undergo procedure Anesthetic: 2% to 4% isoflurane in O 2
Electric razor, disposable razor, or hair removal product Anesthesia apparatus 1-cc syringe 25-to 30-G needle Electrodes for administering the pulses (available from multiple sources, e.g., Harvard Apparatus has both plate and needle electrodes) Electroporation power source Additional reagents and equipment for harvesting tissue or evaluating expression levels and efficiency (Lucas and Heller, 2001;)
Prepare

DNA for procedure 1. Amplify DNA.
For in vivo procedures, DNA used will typically be supercoiled. The plasmid to be delivered will need to be amplified, and stock solution should be at a concentration of 2 to 5 ug/ul.
There are several commercially available kits for performing amplification, as well as commercial entities that will prepare the plasmid at the appropriate concentration and quantity. DNA should be prepared with low endotoxin levels. While tissue-specific promoters can be used, for both muscle and skin, plasmids containing the CMV promoter are often used and are very effective.
2. Suspend DNA at appropriate concentration.
Typically for muscle this will be 0.5 to 1.0 ug/ul, and for skin 1.0 to 2.0 ug/ul. The diluent is typically sterile 0.9% saline. Both sterile phosphate-buffered saline and sterile water have also been used successfully. DNA should be stored cold.
In vivo delivery procedure 3. Remove hair from area (skin or skin above muscle) to be transfected, e.g., using an electric razor, disposable razor, or hair removal product.

Anesthetize the animals.

Using an induction chamber, animals can be anesthetized in 2% to 4% isoflurane in oxygen. Once animal is anesthetized, fit with an appropriate mask and keep under general anesthesia (2% to 3% isoflurane in oxygen) for the entire procedure.
5. Inject DNA into tissue.

A standard injection volume is 50 ul, although volumes between 10 and 100 ul have been used. For muscle, the concentration of DNA should be between 0.5 and 1.0 ug/ul, and for skin the concentration should be between 1.0 and 2.0 ug/ul.

Several muscles have been used successfully with in vivo electroporation including tibialis anterior, gastrocnemius, and rectus femoris. Muscle chosen will be dependent on

animal species utilized and specific application. For skin, delivery is via an intradermal injection and is typically on the flank, abdomen, or base of tail. The concentration and injection volume will also be dependent on the specific application. For vaccines and immunotherapy, the volumes and concentrations may be lower than for protein replacement therapies.
6

An individually addressable suspended-drop electroporation system for high-throughput cell transfection. — Youchun Xu et al., 2014

Introduction

Cell transfection is a basic and important technology in most cell biological studies. Traditional chemical transfection methods 1,2 are toxic and inefficient at delivering exogenous molecules into cells, especially into primary cells and stem cells. Although viral transfections 3,4 display a higher efficiency, such procedures are experimentally difficult, and these approaches are not suitable for protein or drug transfection. As an alternative technology, electroporation is a non-toxic, economical, efficient, and convenient approach to cell transfection via the application of electric pulses. However, current commercial electroporation systems are of limited use due to several technical problems. For instance, the most widely used electroporation platforms are cuvette-type electroporators and they are not suitable for high-throughput experimentation because only one sample can be electroporated at a time, and a large volume of sample is required. Although some multi-well format electroporation devices exist, the manual steps for sample loading, transferring, and device cleaning are tedious and time-consuming, which limits the use of these devices for high-throughput experimentation. 9 To achieve high-throughput transfection, various techniques have been developed.
The reverse transfection method was developed for parallel chemical transfection, wherein plasmid DNA or siRNA are pre-spotted on the surface of a slide to locally reversetransfect into the cells cultured on the slide. This method has the shortcomings of the chemical approaches, precluding its use for hard-to-transfect cell lines and primary cells. Additionally, cells easily migrated away from the transfection area, raising the issue of cross-contamination of different spots. To overcome these challenges, a microwell array-based electroporation device was established to transfer DNA or siRNA into mammalian cells confined to the bottom of microwells by energizing the top and bottom electrodes. 16,17 This method is feasible for high-throughput transfection but not suitable for the transfection of various cell types in parallel. More importantly, in most cell biological research, cell transfection is only an initial step; many following operations must be performed. Yet, in these studies, 16 types of transfected cells. To provide an electroporation method that is compatible with standard biological operations, a suspended-drop electroporation system consisting of two machined gold-coated copper pieces

was designed to create an array of 96 pairs of vertical electrodes aligned on a standard 96-well plate to facilitate cell transfer. 9 However, this device cannot provide customizable electroporation parameters for different chambers.
Here, we present an individually addressable suspendeddrop electroporation system on a printed circuit board (PCB) for selectable high-throughput cell transfection. This PCBbased suspended-drop electroporation (PSE) device has 96 individually addressable through-holes that function as electroporation chambers. The metal inside each through-hole was broken into an electrode pair by milling the metal on the two vertexes of the through-hole. The samples can be toploaded into the through-holes on the PCB, electroporated by the electrode pairs inside the through-holes, and finally flushed into the cell culture multi-well plate by direct addition of the cell culture medium. Compared to a previous report, 9 our device relies on standard PCB technology, which endows this method with cost and manufacturing cycle advantages. In addition, the individually addressable feature makes it possible to achieve customizable electroporation of different cell types with varied biomolecules. Using this PSE device, we effectively introduced plasmid and synthetic siRNA into cultured and primary cells with high cell viability and transfection efficiency.

Methods

Preparation of cells

HeLa, MCF7, LO2, HepG2, and 3T3-L1 cells were obtained from the American Type Culture Collection (ATCC, Manassas, VA), and SMMC-7721 cells were obtained from Peking Union Medical College (PUMC, Beijing, China). SH-SY5Y cells were provided by Dr Kai Gong from Tsinghua University. Goat fibroblast cells were kindly provided by Dr Yanping Xing from Inner Mongolia Agricultural University. The HeLa cell line that expresses enhanced green fluorescent protein (GFP) was established in our laboratory. All of these cells were maintained in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (Gibco BRL Inc, Grand Island, NY) and cultured in a standard cell culture incubator (37 degC, 100% humidity, and 5% CO 2 ). For cell seeding, a monodisperse cell suspension was prepared using standard tissue culture techniques with 0.25% trypsin containing

0.53 mM EDTA.

Transfection of the plasmid and siRNA

For plasmid transfection assays, cell suspensions were centrifuged and re-suspended to a density of 1-4 x 10 6 cells mL -1 in a hypoosmolar electroporation buffer containing 40 mg mL -1 of the pEGFP-N1 plasmid. The electroporation buffer contained 25 mM KCl, 0.3 mM KH 2 PO 4 , 0.85 mM K 2 HPO 4 , and 90 mM myo-inositol (pH adjusted to 7.4 at 25 degC). After electroporation, the cells were transferred to 96-well plates for cell culture. Twenty-four hours later, the transfection efficiency was analyzed with a conventional flow cytometer (BD FACSCalibur, Beijing, China) by calculating the fraction (%) of the total cells which are GFP-positive cells, while the viability ratio was obtained by comparing the number of living cells in the electroporation well to the number of living cells in an untreated group. All the experiments were performed at least in triplicate. All experimental data represent the mean and standard deviation.
For RNA interference assays, the siRNA-GAPDH and siRNA-GFP were dissolved in an electroporation buffer (4 mM final concentration) and electroporated into HeLa and GFPexpressing HeLa cells, respectively. In contrast, conventional electroporation based on cuvette-type chambers using the BTX ECM830 system was also performed with the same electric field strength, duration and pulse number. Moreover, the chemical transfection of siRNA-GFP and siRNA-GAPDH (100 nM final concentration) was also performed as a control using Lipofectamine 2000. To exclude artifacts due to this change in the transfection procedure, control experiments with negative control siRNA were performed in exactly the same way. The sequence of siRNA-GFP and siRNA-GAPDH are presented in Table S1. +

Results and discussion

PSE system setup

The core of the PSE system used in our experiments is a PCB that was designed using commercial electric circuit layout software (Altium Designer Winter 09, Altium Corp., Sydney, Australia) and fabricated with modified PCB technology. The PCB included 3-

An electroporation protocol for efficient DNA transfection in PC12 cells — Giuseppina Covello et al., 2014

PC12 cells are a cell line originating from pheochromocytoma in the rat adrenal medulla (Schaefer et al. Various transfection methods have been attempted to transfect this cell line. In general, cells can be gene-modified in vitro and in vivo using chemical or physical methods (Azzam and Domb In the particular case of PC12 cells, cationic lipids formulations have been employed to increase transfection efficiency. Using Lipofectamine 2000 (Invitrogen) the transfection efficiency was about 14 % and was similar to the efficiency obtained with polyethyleneimine (PEI) (15 %) (Lee et al. Physical methods, including electroporation, biolistics and injection, are used with varying success and are cell cycle-independent but may be more toxic for some cell types and usually require cell suspensions in vitro and specialized equipment (Villemejane and Mir The aim of this work was to find a protocol ensuring high transfection efficiency in PC12 cells, while retaining viability and ability to differentiate. We compared two electroporation systems (Neon transfection and Gene Pulser Xcell) and three chemical transfection methods (Lipofectamine 2000, Lipofectamine LTX, TransIT-LT1).
Plasmid DNA plasmid used for transfection or electroporation was pEGFP-C1 (BD Biosciences Clontech, Palo Alto, CA, USA) driving the expression of an enhanced green fluorescent protein (EGFP) under the control of the CMV promoter. Plasmid was amplified in Culture of PC12 cells Rat PC12 cells (ATCC entry CRL-1721) were grown at 37 °C (5 % CO Viable cells counts A Trypan Blue Stain exclusion test (Invitrogen, Carlsbad, CA, USA) was used to distinguish viable from nonviable cells. The suspended cells (9 μl) were mixed with 1 μl of 0.4 % Trypan Blue Stain and analyzed in the Countess™ Automated Cell Counter (Invitrogen) chamber slide. The percentage of viable cells was calculated as follows: (number of viable cells/total number of cells) × 100. Lipid-mediated DNA transfection The transfection reagents used in this

study were: TransIT-LT1 Transfection Reagent (Mirus, Madison, WI, USA), Lipofectamine 2000 and Lipofectamine LTX (Invitrogen). 4 × 10 Transfection by electroporation DNA electroporation was performed with the Neon For the electroporation with the Neon System, PC12 cells were grown to 70 % confluence in a poly- For the electroporation with the Gene Pulser Xcell System, PC12 cells were grown to 70 % confluence in a poly- High-content image acquisition and analysis Upon chemical transfection or electroporation of pEGFP-C1 vector, EGFP expression and cell nuclei were visualized using Operetta High Content Imaging System (PERKIN ELMER, Monza, Italy). To count total cell numbers, nuclei were counterstained with Hoechst 33342 (Invitrogen). Forty-eight hours after transfection, cells were washed once with PBS and incubated in 0.5 ml of supplemented DMEM containing 1 mg/ml of Hoechst 33342, for 20 min at 37 °C and 5 % CO Cell differentiation Electroporated PC12 cells were plated at a density of 1 × 10 Neurite analysis Neuronal differentiation was estimated every day, for 7 days, after exposure to NGF 2.5S, by measurement of morphological parameters. Images were acquired using a fluorescence microscope (DFC420C, Leica, Milan, Italy) and a 20X objective (magnification). Filter A (Exciter BP340–380 nm; Dichromatic Mirror 400 nm) and I3 (Exciter BP450–490 nm; Dichromatic Mirror 510 nm) were used and the images were analysed with ImageJ software ( Statistical analysis All statistical analyses were performed using Graphpad Prism software package (GraphPad Software, San Diego, CA, USA). Student’s
Comparison of transfection chemical methods in PC12 cells We compared transfection efficiencies obtained in PC12 cells with the lipopolyplex transfection reagent TransIT-LT1 (Mirus) and the cationic lipids Lipofectamine 2000 and LTX (Invitrogen). Cells were grown in conditions promoting proliferation including use of an enriched medium, but the

transfections were carried out in a serum- and antibiotic- free environment. We performed these experiments with different amounts of pEGFP-C1 DNA (Clontech), a plasmid driving the expression of an enhanced green fluorescent protein (EGFP) under the control of the CMV promoter. The ratio DNA (μg): transfection reagent (μl) was 1:3 and we transfected increasing amounts of plasmid (0.25, 0.5, 0.75 and 1 μg). Forty-eight hours after transfection, nuclei were stained with the viable Hoechst 33342 fluorescent dye and images were acquired on an Operetta System, which combines fluorescence microscopy in a multi-well format with automated image acquisition and quantitative analysis (Fig.  As shown in Fig.  On the contrary, when we performed the transfection experiments by using Lipofectamine LTX and Lipofectamine 2000 we managed to transfect DNA into PC12 cells, with efficiencies that ranged from 7 and 15 % respectively, with 0.25 μg DNA, to 35 and 46 % respectively, when 1 μg of DNA was used. As expected, transfection efficiency for both reagents correlates with the amounts of DNA used. However, in comparing the transfection efficiency of the two cationic lipids, Lipofectamine LTX seems to perform better than Lipofectamine 2000 at low DNA amounts, while Lipofectamine 2000 outperforms Lipofectamine LTX at higher DNA amounts. In particular, comparing our results with those by Lee and colleagues (2008), with Lipofectamine 2000 we reached 21 % transfection efficiency (with 0.5 μg of DNA in a 24-well plate) while Lee and collaborators reported 14 % efficiency in similar conditions (1 μg of DNA in a 12-well plate). Cell viability was measured by Trypan Blue Staining 48 h after transfection (Fig.  DNA electroporation in PC12 cells As the transfection efficiency obtained with Lipofectamine 2000 was not sufficient for our purposes, we investigated if we could obtain a higher percentage of transfected PC12 cells with an electroporation method (Neon Transfection System, Invitrogen). To optimize conditions, 0.5 μg of plasmid

Comparative Evaluation of Lipofectamine and Dendrimer for Transfection of Short RNA Into Human T47D and MCF-10A Cell Lines. — Zohreh Jahanafrooz et al., 2023

Short RNAs can influence different hallmarks of breast cancer such as proliferation, apoptosis resistance, invasion/migration, and angiogenesis/new vessel formation. Efficacy and feasibility of short RNA therapies has been confirmed in preclinical attempts. 1 - 3 On the other hand, the onset of viral diseases and emerging nucleic acid-based drugs, which usually have better effects than protein-based drugs, has led pharmacists and biotechnologists to produce more efficient and safer nucleic acid carriers into the cells. Because of the negative charge of the plasma membrane and the negative charge of nucleic acids, the delivery of nucleic acid-based drugs and vaccines is inefficient. Recently, new methods for short RNA transfection offer ways for inactivation of target genes and also analyses of gene function. 4 - 7 Production of genetically modified cells through delivery of foreign nucleic acids into the cells is called transfection, which includes biologically, physically, and chemically mediated methods with various benefits for particular applications. 8 Due to the complications of the traditional delivery methods, researchers are looking for new methods and systems to more efficient delivery. Several factors such as kind of the nucleic acid, cell types and medium conditions have an influence on transfection efficiency. An appropriate transfection method has both high transfection efficiency and low cytotoxicity. 9 , 10 Virus-mediated transfection or transduction is commonly utilized in clinical researches. Although viral vector is an efficient and promising delivery method, it has some disadvantages, including viral recombination, limited size of DNA, host immunological response, and some other possible undesired effects such as off-target or oncogenic effect. 10 , 11 Physical transfection methods, including electroporation, biolistic particle delivery or micro-projectile bombardment, microinjection and laser-based transfection, require various physical tools. 12 Lipidoids, cationic polymers, carbon nanotubes, cell-penetrating peptides and cationic protein–antibody fusions are used in chemical transfection methods. 8 , 13
Liposomes are one of the most notable candidates for drug delivery. Liposomes are spherical vesicles that have an aqueous nucleus surrounded by one or more phospholipid layers and are usually divided according to size (small, large and very large), the number of bilayers (monolayer and multilayer) and

the charge of phospholipids (neutral, anionic or cationic). Cationic liposomes are much more effective and safer, and previous studies have shown that they improve the efficiency of gene transfer compared to other techniques. Cationic lipids apply chemical vectors to condense nucleic acid through ionic interaction. Lipid subunits of Lipofectamine reagent can form liposomes in an aqueous environment and entrap oligonucleotide. 14 Lipofectamine 2000 is a cationic liposome composed of both poly-cationic and natural lipids. Lipofectamine 2000 compared to the old prototype of Lipofectamine series such as Lipofectin is less toxic, and can transfect a wide variety of both adherent and suspend cells in the absence or presence of FBS. Lipofectamine 3000 is even more improved prototype of Lipofectamine. 15 In one recent study, five commonly used transfection reagents, including Lipofectamine 3000, Lipofectamine 2000, Fugene, RNAiMAX and Lipofectin, were comprehensively analyzed in ten cell lines. According their results, Lipofectamine 3000, Fugene and RNAiMAX showed high transfection efficacy with lower toxicity compared to Lipofectamine 2000. The mentioned study also showed that both transfection efficacy and toxicity of the transfection reagents are cell type dependent. 16
PAMAM G5 (polyamidoamine), as a cationic dendrimer, has homogenous and nanometric size and globular shape with various changeable surface functional groups and wide molecular weight range for drug and gene delivery. Over the last decades, there have been many promising results for applications of dendrimers in biology or medicine. PAMAM dendrimer suggested as a promising nonviral gene carrier in cancer therapy. PAMAM dendrimer are stable and hardly oxidized compared to cationic liposomes. 17 , 18 The unique characteristic of dendrimer makes the dendrimer more favorable, in which drugs have been attached or encapsulated to the peripheral active amine groups of dendrimer that modulate its solubility and cytotoxicity. 19 , 20 These highly branched macromolecules are effective carriers for drug and gene delivery in cancer therapy. In a recent study, PAMAM G3 was cross

-linked with 4,4′-dithiodibutryic acid (DA) to form nanoclusters (NCs). The synthesized G3-DA NCs increased 2.3 and 2.1 times gene transfection to cancer cells compared to the PAMAM G3 and PAMAM G5, respectively, under the same conditions. 21
Hydrocortisone is a glucocorticoid hormone, which stimulates cellular growth in cell culture. 22 Hydrocortisone is one of the ingredients in MCF-10A cell culture. Since hydrocortisone can interact with Lipofectamine and interfere in transfection process, 23 the T47D cells were also cultured with hydrocortisone, to evaluate the possible effect of hydrocortisone on transfection efficiency. Our team reported the transfection efficiencies and toxicity of DOTAP (a liposome) and PAMAM G5 in stem cells previously. 4 , 14 In the present study we evaluated transfection efficiency and toxicity of PAMAM G5 dendrimer and Lipofectamine in T47D (breast cancer cell line) and MCF-10A (non-malignant breast cell line) cells. For this purpose, scrambled FITC-conjugated RNA as a type of short RNA was transfected into mentioned cell lines with both transfection reagents. In addition, the necrosis of both cells was evaluated to determine the susceptibility of cells to mentioned reagents.
T47D cells (human ductal breast epithelial tumor cell line) were cultured in RPMI-1640 containing 10% fetal bovine serum (FBS) and 1% Penicillin/streptomycin antibiotics (Gibco, UK) incubated at 37°C in 5% CO2 under 95% humidity. MCF-10A cells (non-cancerous human breast epithelial cell line) were cultured in DMEM/F12 (Invitrogen, USA), supplemented with 15% FBS, 10 μg/ml insulin (Sigma-Aldrich, USA), 100 μg/mL hydrocortisone (Sigma-Aldrich Chemie GmbH, Germany), and 1% pen/strep (Gibco, UK) incubated at 37°C in 5% CO2 under 95% humidity.

A cost-effective approach to microporate mammalian cells with the Neon Transfection System. — Chantal Brees et al., 2014

a b s t r a c t

Electroporation is one of the most efficient nonviral methods for transferring exogenous DNA into mammalian cells. However, the relatively high costs of electroporation kits and reagents temper the routine use of this fast and easy to perform technique in many laboratories. Several years ago, a new flexible and easy to operate electroporation device was launched under the name Neon Transfection System. This device uses specialized pipette tips containing gold-plated electrodes as electroporation chamber. Here we report a protocol to regenerate these expensive tips as well as some other Neon kit accessories, thereby reducing the cost of electroporation at least 10-fold.
O 2014 Elsevier Inc. All rights reserved.
Many cell biology experiments require the introduction of genetic material into cells. A variety of viral and nonviral delivery systems have been developed, but electroporation is often the selected method of choice . The underlying reason is that this method, which employs electrical pulses to create transient pores in cell membranes, reproducibly yields high transfection efficiencies in combination with low cell mortality in a wide range of cell lines . However, a major drawback limiting its broad use, especially in less well-financed research groups, is the high and ongoing cost associated with the purchase of the electroporation kits. Here we present a simple and cost-saving approach to electroporate mammalian cells with the Neon Transfection System (Invitrogen), a device that is increasingly gaining in popularity.
The Neon system employs specialized consumable pipette tips containing gold-plated electrodes as electroporation chamber. In general, the use of such a capillary electroporation system (which is often called a microporation device) has important advantages compared with conventional cuvette-based electroporation chambers: (i) the method is rapid, reproducible, and free of cell loss; (ii) the overall cell viability is higher; and (iii) the transfection efficiencies are superior even for hard-to-transfect cells . However, to ensure repeatability and eliminate variation of the transfection conditions, the supplier strongly advises users not to employ Neon pipette tips more than two times. In addition, the supplier does not provide a protocol to regenerate these tips or any other component of the electroporation kit (e.g., the Neon tubes that hold the electrolytic buffer during electroporation) so that they

can be reused later on to perform high-quality transfections with different plasmids. To overcome this problem, we have developed a fast, reliable, and inexpensive protocol to regenerate Neon pipette tips and tubes for multiple reuses. We found that the Neon pipette tips and tubes can be recycled at least 10 times without significant loss in transfection efficiency (Fig. 1A). In addition, no cross-contamination from the previous plasmid preparation could be detected in transfected cells (Fig. 1B).
Here we provide a protocol for recycling Neon pipette tips and tubes. Note that Neon kits are available for electroporation of 10-or 100-ll samples and that our protocol has been developed with the 10-ll tips. However, by adjusting the volume amounts, the approach should also work with the 100-ll tips. After electroporation, the used tips and tubes are collected and stored at room temperature until regeneration (normally once a week). To regenerate the tips, residual plasmid DNA is first removed by pipetting they are first thoroughly washed with distilled water, thereafter rinsed with 70% (v/v) ethanol, and finally also air-dried in a sterile laminar flow hood. The recycled materials can subsequently be used for electroporation according to the procedures recommended by the manufacturer. Importantly, the limited amounts of the patented resuspension and electrolytic buffers that are provided with the electroporation kit, and of which the composition is not specified, can be easily replaced by a filter-sterilized sucrose-based buffer (SBB: 250 mM sucrose and 1 mM MgCl 2 in DPBS [Dulbecco's phosphate-buffered saline, Gibco cat. no. 14190]) without effecting transfection efficiency (data not shown; the results shown in Fig. 1A were obtained with homemade SBB). Based on our experience, the Neon tips and tubes can be regenerated at least 10 times with little or no deterioration in performance (Fig. 1A). In addition, it is worthwhile to note that the described procedure has already been routinely carried out in our laboratory for several years and that other laboratories have expressed their interest in using this easy and effective Neon kit component recycling protocol to perform microporation experiments at affordable cost. To assess potential crosscontamination in sequential experiments, the two plasmids were used in alternate order

. After electroporation, the cells were cultured in standard growth medium for 2 days, fixed, counterstained with DAPI (4 0 ,6-diamidino-2-phenylindole) to visualize nuclei, and processed for fluorescence microscopy as described elsewhere .