A novel rapid and reproducible flow cytometric method for optimization of transfection efficiency in cells

Transfection is one of the most frequently used techniques in molecular biology that is also applicable for gene therapy studies in humans. One of the biggest challenges to investigate the protein function and interaction in gene therapy studies is to have reliable monospecific detection reagents, particularly antibodies, for all human gene products. Thus, a reliable method that can optimize transfection efficiency based on not only expression of the target protein of interest but also the uptake of the nucleic acid plasmid, can be an important tool in molecular biology. Here, we present a simple, rapid and robust flow cytometric method that can be used as a tool to optimize transfection efficiency at the single cell level while overcoming limitations of prior established methods that quantify transfection efficiency. By using optimized ratios of transfection reagent and a nucleic acid (DNA or RNA) vector directly labeled with a fluorochrome, this method can be used as a tool to simultaneously quantify cellular toxicity of different transfection reagents, the amount of nucleic acid plasmid that cells have taken up during transfection as well as the amount of the encoded expressed protein. Finally, we demonstrate that this method is reproducible, can be standardized and can reliably and rapidly quantify transfection efficiency, reducing assay costs and increasing throughput while increasing data robustness.


Introduction
Transfection is one of the most common used techniques in molecular biology [1,2]. Transfection is the process of introducing plasmid nucleic acid (DNA that carries a gene of interest or mRNA) into target cells that then eventually express the desired nucleic acid or protein. There are a number of strategies for introducing nucleic acids into cells that use various biological, chemical, and physical methods [1][2][3]. However, there is a wide variation with respect to transfection efficiency, cell toxicity, the level of gene expression, etc. To determine how these factors influence transfection, a sensitive and robust detection assay is required to quantify and  [19] were cultured in RPMI 1640 medium (Invitrogen) supplemented with 10% fetal bovine serum (FBS), L-glutamine, penicillin, and streptomycin. All cells were incubated at 37˚C and 5% CO 2 .

Nucleic acid labeling of plasmids
Plasmid DNA for transfection was extracted from Stbl3™ (Thermo Fisher, Waltham, MA) or Stellar (Clontech, Mountain View, CA) bacteria using the PureLink HiPure Midiprep Kit (Thermo Fisher, Waltham, MA). The concentration was determined using spectrophotometry and the purity was between 1.8-2.0 (260/280 ratio). Plasmid DNA was labeled with DNA LabelIT Tracker (Mirus, Madison, WI) the day before transfection using 0.5 ul FITC/1 μg DNA according to the manufacture's protocol. Unreacted LabelIT Tracker reagent was removed using ethanol precipitation. According to the manufacturer 1 ul of labeling dye per 1 μg plasmid DNA will yield labeling efficiencies of approximately one Label molecule every 40 base pairs (on average) of double-stranded DNA. The purity and concentration of labeled DNA was determined by spectrophotometry.

Transfection reagents
The following commercially available transfection reagents were purchased: TransIT-X2 (Mirus, Madison, WI), Jet Prime (Polyplus, France), Lipofectamine 2000 (Thermo Fisher, Waltham, MA) and Fugene HD (Promega, Madison, WI). All reagents were used at recommended reagent to DNA ratio to transfect a constant amount of 1 μg DNA for comparison.

Chemical-based transfection conditions
For transfection experiments, 3x10 5 293T cells passaged less than 20 times were plated into 12 well plates the day before transfection. At this density, cells were attached to the well bottom after 24 h at a confluency of 70%-80%. Twenty-four hours after seeding, cells were transfected with FITC-labeled and unlabeled pNL4-3 or pUltraHot encoding mCherry. The transfection was performed using 1 μg plasmid DNA and 3 μl TransIT X2 in serum free media per well, according to the manufactures'protocol. Cells were cultured in complete media. Transfection complexes were removed after 6 hours by replacing all of the media in the well. For time course experiments, the cells were transfected at the same time and harvested at 0, 6, 12, 24, 36, 48 and 72hrs. For experiments comparing different transfection reagents, the cells were transfected with 1 μg DNA/well with varying amounts of transfection reagents to facilitate the recommended ratio of transfection reagent to DNA for each reagent: 3 μl TransIT X2, 4 μl Lipofectamine 2000, 2 μl Jet Prime and 3 μl Fugene HD. Cells were harvested at 24 hrs posttransfection and stained with a viability dye and for intracellular target protein. All experiments were performed in triplicates.
transferred into 4mm cuvettes (Gene Pulser cuvettes, Biorad, Hercules, CA) and 4 μg of labeled or unlabeled pUltraHot plasmid were added to the suspension. Jurkat E6-1 cells were electroporated with a BioRad Gene Pulser Xcell electroporation system using the exponential protocol with 250 volts, 350uF capacitance, 1000 ohm Resistance. Cells were transferred immediately into pre-warmed media and incubated for 24 hours before staining for viability and flow cytometric analysis.

Flow cytometry reagents
Anti-HIV-1 p24 monoclonal antibody (71-31) was obtained through the NIH AIDS reagent program #530 and was conjugated with a CF647 dye (Mix-n-Stain Kit from Biotium, Hayward, CA). The matching human IgG1 control was purchased from Biolegend (San Diego, CA) and conjugated with a CF647 dye (Mix-n-Stain Kit from Biotium, Hayward, CA). Amine reactive (Ghost 780 Live/dead dye) and nucleic acid dye (7-AAD: 7-Aminoactinomycin D) were obtained from Tonbo Biosciences (San Diego, CA).   kits for A488 and A647 (Bangs Laboratories, Fishers, IN) as previously described [21]. The fluorescent MESF (Molecules of Equivalent Soluble Fluorochrome) beads contain five different populations of beads, each labelled with a known number of fluorochrome molecules [21]. When this bead mixture is analyzed, the mean fluorescence intensity (MFI) values for each bead peak corresponds to the approximate number of fluorescence molecules associated with a cell in the same MFI range [21]. The FITC and APC channel can be calibrated with these MESF beads on a logarithmic scale, giving a standard curve for MESF values, and by extension FITC and APC molar concentration. Determining the MFI value for each bead peak in the FITC and APC channel allows for the determination of the equivalent MESF values associated with each sample. With the amount of FITC conjugated to the plasmid being a known value, the amount of DNA plasmid can then be calculated at the molar level.

Statistical analysis
Experiments were performed in three independent cell culture preparations (e.g., biological replicates) unless stated otherwise in the figures. Differences of MFI between different transfection variations were analyzed by two-way analysis of variance (ANOVA) or by Student's t-test, or non-parametric equivalent as appropriate. Correlations were assessed with the Spearman rank correlation coefficients. Statistical analyses were performed using GraphPad Prism. Significant differences were accepted when p 0.05.

Simultaneous quantification of nucleic acid uptake and protein expression by flow cytometry
To develop a robust flow cytometric method that can be used to reliably quantify transfection efficiency at the single cell level and to avoid limitations of GFP-based flow cytometric methods that can compromise accuracy of determination of transfection efficiency [6,[9][10][11][12][13][14][15], we developed a flow cytometric method that is based on two independent readouts of transfection efficiency and on direct labeling of the transfected plasmid DNA. The FITC fluorochrome is used to detect intracellular levels of the transfected plasmid that has been labeled with FITC (Label IT tracker) (Fig 1A-1E). The second fluorochrome is used to quantify expression of the target protein [by directly measuring fluorescence of the expressed protein if the target protein is fluorescent e.g. mCherry ( Fig 1C) or by using a fluorescent-labeled antibody against the target protein ( Fig 1D)]. Fig 1 shows the gating strategy of the flow cytometric method. To develop and optimize this method, we focused on the 293T cell line that is known to be an excellent cell line for quantification of transfection efficiency of various plasmids. Of note the labeling of the transfected nucleic acids did not affect their function, i.e. transcription since similar amounts of target protein were detected with use of labelled or unlabeled (standard method) plasmids (Figs 1 and 2). However, our novel flow cytometric method can also be used as a tool to optimize transfection efficiency in difficult to transfect human cells without the need to co-transfect a GFP reporter plasmid that can further compromise transfection efficiency ( Fig 1D). We validated our method using two independent labeled plasmids, the pNL4-3 plasmid (14 kB) that encodes the entire HIV genome and the smaller (8.3 kB) pUltraHot plasmid that encodes for the mCherry fluorescent protein. Using this method both plasmid DNA and protein can be quantified simultaneously in the same tube at the single cell level, while gating in viable cells. The confounding effect of cellular toxicity on determination of transfection efficiency can be minimized with the novel flow cytometric method Toxicity of DNA and the transfection method per se should also be taken into consideration during transfection [22] but most standard methods that are used to quantify transfection efficiency (such as the luciferase method) do not take this important variable into consideration. A third fluorochrome can be used to quantify cellular toxicity (death) as a direct result of the transfection (Fig 3A and 3D). The direct toxicity as a result of the DNA plasmid and transfection reagent can be quantified by the percentage of cells expressing the FITC labeled DNA plasmid that are also positive for the death dye detected in an appropriate channel (Fig 3A and  3D). Alternatively, the confounding effect of cellular toxicity can also be minimized without Overall cellular toxicity was similar (10-40%) among the 3 methods [FSC/SSC (Fig 1), nucleic acid and amine reactive death dyes] (Fig 4). The mean viability of the untransfected 293T cells was >85%. In addition, determination of the transfection efficiency based on two independent readouts (DNA plasmid uptake and protein expression) was similar by gating on viable cells based on either FSC/SSC (B, E) or viability dye (C, F). use of a death dye since quantification of cell death by forward and side scatter parameters, and doublet discrimination gave similar results compared to quantification of cell death by nucleic acid (7-AAD) or amine viability dye (Figs 3 and 4). In addition, determination of the transfection efficiency was similar by gating on viable cells based on either FSC/SSC or viability dye (Figs 2 and 3). Since viability dyes cannot be used in certain experiments such as electroporation, gating on viable cells based on FSC/SSC is a reasonable approach. Thus, the novel flow cytometric method can be used to limit analysis of transfection efficiency to viable cells and can minimize the effects of direct cellular toxicity of transfection on the determination of transfection efficiency.
The novel flow cytometric method that quantifies transfection efficiency is reproducible Using the approach described in Fig 1 and standardized fluorescence units (MESF) beads, the transfection efficiency for a specific target can be standardized using two independent readouts (amount of DNA uptake and detected expressed protein) (Fig 5). Overall, the MESF of FITC (labeled DNA) in viable cells was the most reproducible readout of transfection efficiency (mean interassay coefficient of variation 8.56%, range 4.19 to 12.66%) (Fig 5). Thus, the standardized fluorescence per cell provides a more reproducible measure of transfection efficiency compared to the percentage of transfected cells in a cell population.
Optimization of timing to determine transfection efficiency using the novel flow cytometric method There is limited published evidence regarding the optimal timing after transfection for determination of transfection efficiency [9,10,22]. Using the methodology described in Fig 1 we found that the peak uptake of two independent labeled plasmids occurred after 6 hours after transfection and that the peak protein expression was evident within 48 hours (Fig 6). For both plasmids, the peak uptake of plasmid DNA (FITC) was seen at 12 hours. A linear reduction in the DNA FITC signal with a simultaneous increase in the protein signal was seen after 12 hours. Similar optimization of timing to determine transfection efficiency can be established with various cells and transfection methods (e.g. Jurkat E6 cells electroporation with FITClabeled DNA mCherry plasmid). Thus, the current method can be used as a tool to optimize the timing of determination of transfection efficiency, but specific time-course experiments need to be performed for specific plasmid and transfection reagents/methods.
The novel flow cytometric method can be used as a tool to optimize transfection by selecting the best for a specific type and amount of plasmid transfection reagent To further optimize transfection efficiency, we directly compared four major commercially available nonviral reagents of chemical transfection using the same approach as in Fig 1 and the best time point (24 hours as determined in Fig 6) for the use of both independent readouts (labeled plasmid and expressed protein) of transfection efficiency. For each transfection reagent, we chose the optimal ratio of transfection reagent to the amount of DNA plasmid that produced the highest FITC signal based on recommendations from the manufacturer. Using two independent plasmids, TransIT-X2 and Jet Prime gave the best (and comparable) transfection efficiencies, whereas the efficiency was reduced with Lipofectamine 2000 and Fugene HD (Figs 7 and 8). We also determined the viability of the cells during the chemical transfection with different reagents. Using the approach described in Figs 1 and 2 and death dye, all transfection reagents had similar cellular toxicity at 24 hours after transfection (overall cell death 8.2-12.6%). Thus, using our flow cytometric method for a specific amount of a certain plasmid, different transfection reagents can be directly compared with regards to efficiency and cellular toxicity and thus the efficiency of chemical transfection can be further optimized.
Using the novel flow cytometric method, transfection efficiency can be reliably quantified as early as 6 hours after transfection The MESF readouts (at 24 hours after transfection) for labeled plasmid and expressed protein, correlated better compared to the % positivity for labeled plasmid and expressed protein (Fig  9). In addition an early (6 hours) readout from labeled DNA (FITC) correlated significantly (r = 0.65-0.8, p<0.05) with the maximum (48 hours) median fluorescence per cell that corresponded to expression of protein per cell. Thus, our method is rapid and can reliably be used to quantify transfection efficiency as early as 6 hours after transfection.

Discussion
Here we provide a relatively simple, rapid and robust flow cytometric method that can be used as a tool to optimize transfection efficiency at the single-cell level. This method takes into consideration cell toxicity as a direct result of the transfection and the nucleic acid per se and uses two independent readouts of transfection efficiency: a) the amount of plasmid nucleic acid that cells have taken up during transfection b) the amount of the encoded expressed protein.
It is important to emphasize that any nucleic acid plasmid can be labeled with the LabelIT Tracker, making this method very versatile and applicable to any transfection method. We observed a higher transfection efficiency based on DNA FITC signal and protein expression with a smaller plasmid, which is in line with the nature of large plasmid requiring more transfection reagent to complex the DNA and being more difficult to be transported across the plasma membrane than smaller constructs. We here also discuss important considerations required to successfully implement this method that can be used to optimize several variables that can affect transfection efficiency such as cellular toxicity related to the transfection method, the time of exposure of a standard cell line to specific nucleic acid plasmid and the type of transfection reagent. This method overcomes limitations of prior established methods such as dual reporter systems [6,[9][10][11][12][13][14][15] and measures reliably the efficiency of transfection into human cells and cell lines.
Transfection is an important tool in molecular biology that is applicable for gene therapy studies in humans using methods such as LTR-Specific Tre-Recombinase [23], the Sleeping Beauty (SB) transposon system [24] and the clustered, regularly interspaced, short palindromic repeat (CRISPR) technology [25]. One of the biggest challenges to investigate the protein function and interaction in gene therapy studies is to have reliable monospecific detection reagents, particularly antibodies, for all human gene products [26,27]. Transfection efficiency can be affected by multiple experimental parameters such as the choice of the transfection method, health, and viability of the cell line, degree of confluency, quality and quantity of the nucleic acid used [1,2]. The ideal approach should be selected depending on specific cell type and experimental needs, and should have high transfection efficiency, low cell toxicity, minimal effects on normal physiology, and be easy to use and reproducible [1,2]. Thus, a rapid, robust Comparison of major commercially available transfection reagents using described flow cytometric method that quantifies transfection efficiency. 293T cells underwent chemical transfection using labeled or standard transfected nucleic acids (pNL4-3 or mCherry plasmids) and different commercially available transfection reagents as described in Methods. Gating on viable cells was performed as shown in Fig 1. Each experiment was performed in triplicates and six independent times (6 per plasmid and 12 in total) with specific (un)labelled nucleic acids. Data in each experiment were normalized by the average of the and reliable method that can be used to efficiently quantify transfection efficiency based on not only expression of the target protein of interest but also the uptake of the nucleic acid plasmid, can be an important tool in molecular biology.
The measure of transfection efficiency, the percentage of transfected cells in a cell population, is a subjective measure prone to many variable factors, such as cell cycle progression, the circadian rhythm of gene expression activity, promoter activity, and general activity of a given experimental control (standard transfected nucleic acids) in each experiment and were then pooled together. This approach increases statistical power while taking into consideration the inherent differences in transfection efficiency among different transfection methods (different chemical transfection reagents and plasmids). The non-parametric statistical Mann-Whitney test was used for comparisons between groups. Median and interquartile range (IQR) are shown. The transfection efficiency was assessed by both amount of labelled DNA (A) and protein (B). The use of TransIT-X2 and Jet Prime gave the best (and comparable) transfection efficiencies, whereas the efficiency was reduced with Lipofectamine 2000 and Fugene HD. There was a major decrease in the amount of detectable labelled DNA with Lipofectamine 2000 and Fugene HD.
https://doi.org/10.1371/journal.pone.0182941.g008 Flow cytometry and transfection efficiency cell type. All of these factors can inhibit a cell from actually expressing the transfected protein.
Commonly used methods to quantify transfection efficiency such as the luciferase assay, require cell lysis, do not quantify overall cell toxicity, and thus the determination of the percentage of transfected cells in a cell population may not be accurate [4]. Indeed, in prior work using luciferase reporter constructs to compare different transfection reagents, arbitrary units per protein content were used as readout instead of a percentage of transfected cells [28]. On the other hand, flow cytometric methods to quantify transfection efficiency in cells based on transfection of GFP-fusion proteins or co-transfection of GFP plasmids have limitations including competition in expression of the two different plasmids [9] and artifacts of GFP fluorescence during processing of cells or tissues [6,[9][10][11][12][13][14][15]. Maeβ et al reported fluorescence labelling of siRNA in the setting of transfection of macrophages but did not clarify details of the labelling method of the nucleic acid (siRNA), the method was not standardized and there was no simultaneous measurement of target protein as a tool to quantify transfection efficiency [29]. Herein, we standardized median fluorescence intensity of two independent readouts of transfection efficiency (amount of DNA that cells have taken up during transfection as well as the amount of the expressed target protein) using MESF beads. This approach provides an objective and more reproducible (compared to the percentage of transfected cells in a cell population) measure of transfection efficiency.
We also used this method to demonstrate that it can be used as a tool to detect transfected DNA in difficult to transfect human cells without the need to co-transfect plasmids for reporters that can further compromise transfection efficiency. The use of plasmid vectors for the ex vivo genetic modification of human tissue/cell types for therapeutic purposes has been limited by the low efficiency of currently available plasmid transfection systems and their cell toxicity [30][31][32]. Since lymphocytes are notoriously refractory to most kinds of nonviral DNA delivery methods that have significant toxicity, robust methods to quantify transfection efficiency as well as transfection-induced cellular toxicity are needed. Thus, our method can directly detect labeled transfected nucleic acid in viable cells after electroporation at the single cell without the need to use co-transfection of two different plasmids involving a reporter gene.
One of the limitations of nonviral gene delivery systems is also their toxicity that can significantly compromise transfection efficiency. Other common methods that quantify transfection efficiency such as luciferase assay do not quantify cell toxicity of the transfection method [4]. Our assay can quantify not only the transfection efficiency with 2 different readouts (DNA-FITC and protein) but also the cellular toxicity. Quantification of transfection efficiency at the single cell level is based on viable target cells and this approach minimizes confounders such as the effect of cellular toxicity on overall transfection efficiency and the effect of detachment of adherent cells on cell death rate. Thus depending on the application and the cell type this information can be used to further optimize transfection efficiency.
Incubation time for optimal protein detection after transfection varies (3-48 hours) between different plasmids and transfection methods depending on the protein of interest and its post-translational modifications, location and turnaround time [10]. Due to aforementioned limitations of prior methods to quantify transfection efficiency, there is limited published evidence regarding the optimal timing after transfection for determination of transfection efficiency [9,22]. Interestingly, we found that the DNA uptake peaked between 6-12 hours after transfection. The low capacity of DNA to escape from endosomes is regarded a major limitation of their transfection efficiency [33]. Thus carriers (such as polymers, peptides and lipids) based on the exploitation of the imidazole ring as an endosome destabilization device after often used with plasmids to enhance nucleic acid delivery in the cytosol [33]. DNA plasmids are also often condensed by protamine to enhance cytosolic delivery [34][35][36]. Cellular processes related to DNA are governed by the intricate interplay between different states of DNA compaction that often have surprising and poorly described properties [37]. Thus, the DNA plasmid may open later after initial cellular uptake and an unexpected peak in its detection may be seen hours after initial uptake. Our method based on fluorescent labeling of DNA plasmid can be further optimized with the use of novel methods such as ImageStream flow to study in detail cytoplasmic and nuclear transfer process [38]. Thus, with our flow cytometric method cells can be harvested at multiple times throughout an experiment to generate a time-course experiment and optimize the best time to determine peak transfection efficiency.
There is limited published evidence regarding direct comparison of the major commercially available transfection reagents (for the same plasmid and cell type) [22]. With our proposed flow cytometric method, it is possible to compare several different transfection products simultaneously and thereby reduce result variability. Out of the numerous commercially available transfection reagents, we chose four nonviral transfection reagents which can be used in serum-containing media. The first one, Lipofectamine™ 2000 (Invitrogen) transfection reagent, is based on the lipofection method [39]. Lipofectamine reagents are often considered the "gold-standard" of transfection reagents and are used as a basis of comparison for efficiencies of other transfection methods [40]. The second transfection reagent evaluated in this study was JetPrime™ (Polyplus transfection, France), which is a nonliposomal, polymer-based transfection reagent. Both lipofection and polymer transfections, like JetPEI [41] and JetPrime, involve extensive membrane interaction in the form of liposome fusion or charge binding and subsequent endocytosis, respectively. The third transfection reagent was TransIT-X2 1 Dynamic Delivery System an advanced non-liposomal system of proprietary polymers that aid in nucleic acid complexation, uptake and endosomal release thereby enabling superior transfection of plasmid DNA and smaller nucleic acids. Finally, the fourth transfection reagent was FuGENE 1 HD Transfection Reagent a nonliposomal formulation designed to transfect DNA into a wide variety of cell lines with high efficiency and low toxicity. While these four different transfection reagents only represent a small fraction of commercially available products, our flow cytometric assay is a valuable tool for future evaluation of any transfection reagent in specific cell types.
Interestingly, a drastic reduction in the detection of the FITC labeled DNA was seen when Lipofectamine 2000 and Fugene HD were used for chemical transfection. It is possible that quenching of FITC in endosomes/lysosomes may be an explanation for our findings given the nature of the transfection reagents and that FITC is a pH-sensitive fluorescent dye [42,43]. It is also possible that fluorescence labeling inhibits complex formation/transfection with Lipo/ Fugene. In addition, we found that addition of amine reactive death dyes interfered with the FITC readout and gave variable and inconsistent results. Different fluorochromes (CX-Rhodamine, Cy 1 3, Cy 1 5, TM-Rhodamine) that are not as pH-sensitive dyes as FITC can be used to label plasmid nucleic acid (RNA or DNA) [44][45][46]. The precise mechanisms of how the plasmid DNA interacts with different cellular compartments are not well characterized. Based on our results, we suggest that chemicals (such as TransIT-X2 1 ) that are optimized to allow escape of the nucleic acid plasmid from endosomes/lysosomes should be preferred when our flow cytometric method is used to quantify efficiency of chemical transfection. Finally, possible biochemical interactions with individual fluorochromes and the DNA LabelIT Tracker need to be individually tested for each transfection method.
Any nucleic acid (DNA or RNA) plasmid can be labeled with the LabelIT Tracker, making this method very versatile and applicable to any transfection method. This method is applicable to transfection methods that have revolutionized the field of gene therapy such as the bacterial CRISPR/Cas9 system. CRISPR consists of two components: a "guide" RNA (gRNA) and a non-specific CRISPR-associated endonuclease (Cas9) [47][48][49][50]. The genomic target of Cas9 can be changed by simply changing the targeting sequence present in the gRNA. Once expressed, the Cas9 protein and the gRNA form a riboprotein complex [47][48][49][50]. For some CRISP libraries, Cas9 (or Cas9 derivative) is included on the gRNA-containing plasmid; for others, they must be delivered to the cells separately [47][48][49][50]. Mammalian expression vector containing promoters for Cas9 and gRNA, lentiviral transfer vector containing Cas9 and gRNA, RNA plasmids containing gRNA and Cas9 mRNA and purified Cas9 protein-gRNA riboprotein complexes have all been used as major expression systems and variables for using CRISPR in mammalian cells [25,[51][52][53]. Although genomic cleavage PCR detection methods and sequencing of genomic DNA are ultimately needed to quantify efficiency and accuracy of the CRISP system for a specific application, these methods are not efficient tools to optimize all pertinent experimental variables related to the transfection per se [53][54][55]. Reporter gene (e.g. GFP) to identify and enrich positive cells, or selection marker to generate stable cell lines have been used but reporter systems have several aforementioned limitations. The approach presented here allows labelling of any nucleic acid and simultaneous detection of target protein and could be adapted to optimize delivery of Cas9/gRNA cassettes for genome editing in a variety of systems. Commercially available monoclonal antibodies targeting the Cas protein (clone 7A9-3A3) are available (Cell Signalling Technology) and can be used for flow cytometry. In addition, different fluorochromes (FITC, CX-Rhodamine, Cy 1 3, Cy 1 5, TM-Rhodamine) [44][45][46] can be used to simultaneously label both the gRNA and the Cas9 nucleic acid (mRNA or pDNA).

Conclusion
Taken together, the described relatively simple, rapid and robust flow cytometric method can be used as a tool to optimize transfection efficiency at the single-cell level. This method overcomes limitations of prior established methods (such as dual reporter systems) to quantify transfection efficiency. Any DNA or RNA template can be labelled and is suitable for a wide range of applications. The one-step chemical method is easy and allows consistent control of the labeling reactions and high sensitivity of detection of plasmid DNA can be achieved with optimally labeled DNA and RNA. In addition, the fluorescent labels do not impact hybridization performance and the covalent mechanism allows permanent, non-destructive modification of nucleic acid residues that is ideal for many diverse applications. Given the importance of transfection in molecular biology that has set the basis for gene therapy studies in humans, this novel flow cytometric method can be an important tool in molecular biology and gene therapy studies using methods such as the Sleeping Beauty (SB) transposon system, the CRISPR technology and in cases where reliable monospecific detection methods for human gene products (protein) do not exist.