The authors have no acknowledgments.

The following protocol involves working with mammalian cells; familiarity with sterile technique/cell culture is expected.
1. Cell Line and Culture Conditions
<ol>
	<li>Make 500 mL of medium for the culture of HCT 116 cells: McCoy&#39;s 5A modified medium supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, and 1% penicillin (this is complete medium). 
	NOTE: Grow HCT 116-19 cells in a T-75 or T-175 flask prior to plating. When 90% confluent, each T-75 flask will yield 8.4 x 106 cells, roughly five 10-cm plates, and each T-175 will yield 18.4 x 106 cells, roughly fifteen 10-cm plates.</li>
</ol>
2. Harvesting Cells from the Flask
<ol>
	<li>Aspirate away the medium, wash with Dulbecco&#39;s Phosphate-Buffered Saline without calcium and magniesium (PBS) (10 mL for T-75 or 25 mL for T-175), and aspirate.</li>
	<li>Add trypsin dropwise to the flask using a 2-mL pipette (2 mL for T-175 or 1 mL for T-75). Place the flask in an incubator at 37 &#176;C and 5% CO2 for 5 min to allow the cells to detach.</li>
	<li>Tap the flask to make sure that all cells are dislodged and then quench with complete medium by dispersing it over the entire surface of the flask (8 mL for T-175 or 4 mL T-75).</li>
	<li>Pipette up and down multiple times to break up cell clumps and transfer the cells to a 15-mL conical tube</li>
	<li>Before spinning the cells down, take 10 &#181;L from the 15-mL conical and combine it with 10 &#181;L of trypan blue to count the cells. Pellet the cells by spinning for 5 min at 125 x g and 16 &#176;C.</li>
</ol>
3. Counting the Cells
<ol>
	<li>Transfer 10 &#181;L of the cells mixed with trypan blue to the hemocytometer. Count the 4 grids around the outside (each grid contains 16 squares).
	<ol>
		<li>Take the average cell count from each set of sixteen corner squares.</li>
		<li>Multiply by 10,000 (104).</li>
		<li>Multiply by the total volume of medium used to harvest the cells to correct for the dilution from the trypan blue addition. 
		NOTE: The equation format to calculate the volume to resuspend the cells follows: 
		<img alt="Equation 1" src="/files/ftp_upload/56195/56195eq1.jpg" /> 
		<img alt="Equation 2" src="/files/ftp_upload/56195/56195eq2.jpg" /></li>
	</ol>
	</li>
</ol>
4. Plating the Cells
<ol>
	<li>For each 10-cm plate of cells to be synchronized, add 5 mL of complete medium and 6 &#181;M of aphidicholin (12 &#181;L of a 2.5 mM stock in 200-proof ethanol).</li>
	<li>Transfer 100 &#181;L of re-suspended cell pellet, 2.5 x 106 cells, to each 10-cm plate and swirl gently to mix.</li>
	<li>Incubate the plates at 37 &#176;C and 5% CO2 for 16-24 h to synchronize cells at the G1/S border.</li>
</ol>
5. Releasing the Cells from Aphidicolin Synchronization
<ol>
	<li>4 h prior to targeting, aspirate the medium, wash with PBS, aspirate the PBS, and add 5 mL of complete medium</li>
	<li>Place it back in the incubator at 37 &#176;C and 5% CO2 for 4 h</li>
</ol>
6. RNA Complexing
<ol>
	<li>Enter the mutant eGFP gene sequence into the Zhang Lab&#39;s online generator25 (http:// crispr.mit.edu/) and choose the CRISPR guide sequences that bind with close proximity to the target site. Obtain the CRISPR guide sequences from a commercial source.</li>
	<li>Store the CRISPR RNA (crRNA), trans-activating crRNA (tracrRNA), and Cas9 protein at 20 &#176;C and use according to manufacturer suggestions.
	<ol>
		<li>Mix the RNA in equimolar concentrations to 45 &#181;M. Add 6.75 &#181;L of a 200 &#181;M stock of crRNA and 6.75 &#181;L of a 200 &#181;M stock of tracrRNA to a 1.5-mL centrifuge tube. Add 16.50 &#181;L of TE Buffer to make a final volume of 30 &#181;L.</li>
	</ol>
	</li>
	<li>Heat at 95 &#176;C for 5 min in a heat block or PCR machine. 
	Caution: Hot!</li>
	<li>Allow to cool to room temperature. 
	NOTE: If using a PCR machine, set the cooling to 0.2 &#176;C/s.</li>
	<li>Perform the following steps for each sample.
	<ol>
		<li>Dilute 2.22 &#181;L of crRNA:tracrRNA complex in 2.78 &#181;L of TE Buffer (10 mM Tris, pH 8.0 and 0.1 mM EDTA; pH 8.0) to a final volume of 5 &#181;L.</li>
		<li>Dilute 1.67 &#181;L of Cas9 Protein from a 60 &#181;M stock in 3.33 &#181;L of low-serum medium to final volume of 5 &#181;L.</li>
	</ol>
	</li>
	<li>Mix 5 &#181;L of Cas9 protein with 5 &#181;L of complexed RNA</li>
</ol>
7. Harvesting the Cells for Targeting
<ol>
	<li>Aspirate the medium, wash with 5 mL of PBS, aspirate the PBS, and add 1 mL of pre-warmed trypsin to each 10-cm plate. Put the plates in the incubator at 37 &#176;C and 5% CO2 for 5 min.</li>
	<li>Tap on the 10-cm plate to make sure all cells are dislodged and then quench with 4 mL of complete medium by dispersing it over the entire surface of the plate.</li>
	<li>Pipette up and down multiple times to break up cell clumps and transfer the cells to a 15-mL conical tube.</li>
	<li>Before spinning the cells down, take 10 &#181;L from the 15-mL conical tube and combine it with 10 &#181;L of trypan blue to count the cells. Pellet the cells by spinning for 5 min at 125 x g and room temperature.</li>
	<li>Aspirate the medium and wash with 5 mL of PBS. Pellet the cells by spinning at 125 x g for 5 min at room temperature.</li>
</ol>
8. Counting the Cells
<ol>
	<li>Transfer 10 &#181;L of the cells mixed with trypan blue to the hemocytometer. Count the 4 grids around the outside (each grid contains sixteen squares).
	<ol>
		<li>Take the average cell count from each set of sixteen corner squares.</li>
		<li>Multiply by 10,000 (104).</li>
		<li>Multiply by the total volume of medium used to harvest the cells to correct for the dilution from the trypan blue addition. 
		NOTE: Following is the equation format to calculate the volume to resuspend the cells. Re-suspend the required number of cells in serum-free McCoy&#39;s 5A modified medium. 
		<img alt="Equation 3" src="/files/ftp_upload/56195/56195eq3.jpg" /> 
		<img alt="Equation 4" src="/files/ftp_upload/56195/56195eq4.jpg" /></li>
	</ol>
	</li>
</ol>
9. Targeting Samples
<ol>
	<li>Transfer 100 &#181;L of cell suspension (5 x 105 cells) from step 8.1 to each electroporation 4-mm gap cuvette. Add 10 &#181;L of RNP complex from step 6.7 to 100 &#181;L of cells at a 5 x 105 cell density. Add ODN (2 &#181;M) to each sample. 
	NOTE: For a positive control, add 1 &#181;L of eGFP at 1 &#181;g/&#181;L expressing plasmid
	<ol>
		<li>Take the rack to an electroporation machine, lightly flick each sample, and place them in the chamber. Electroporate at 250 V, LV; 2 pulses, 1 s; 13 ms; unipolar pulse.</li>
	</ol>
	</li>
	<li>Transfer the rack back to the hood. Transfer each sample to a well containing 2 mL of complete medium in a 6-well plate. Incubate at 37 &#176;C and 5% CO2 for 72 h before checking for correction levels.</li>
</ol>
10. Analysis of Gene Edited Cells and Transfection Efficiency
<ol>
	<li>Aspirate the medium and wash the cells with 2 mL of PBS. Aspirate the PBS and add 500 &#181;L of pre-warmed trypsin to each well of the 6-well plate. Put the plates in the incubator at 37 &#176;C and 5% CO2 for 5 min.</li>
	<li>Tap the plate to make sure all cells are dislodged and then quench with 1 mL of complete medium by dispersing it over the entire surface of the well.</li>
	<li>Pass the cells into a 1.5-mL centrifuge tube and pellet at 5,000 x g for 5 min at room temperature.</li>
	<li>Aspirate the medium. Re-suspend the cell pellet in 500 &#181;L of FACS buffer (0.5% BSA, 2 mM EDTA, and 2 &#181;g/mL propidium iodide in PBS).</li>
	<li>Measure the cell fluorescence (eGFP+) by flow cytometry.
	<ol>
		<li>Calculate the correction efficiency as the percentage of the total live eGFP-positive cells over the total number of live cells in each sample, as described in Rivera Torres et al30.</li>
	</ol>
	</li>
</ol>
11. DNA Sequence Analysis
<ol>
	<li>Electroporate the synchronized and released HCT 116-19 cells at a concentration of 5 x 105&#8197;cells/100&#8197;&#181;L, with RNP complex at 100 pmols and 72NT ODN at 2.0 &#181;M.</li>
	<li>Transfer the cells to 6-well plates and allow them to recover for 72 h.</li>
	<li>Sort the cells individually into 96-well plates using a FACS sorter with a 488-nm (100 mw) laser for eGFP+/-, as described in Rivera-Torres et al30. 
	NOTE: Not all wells will successfully grow.</li>
	<li>Expand the cells over 6 weeks and harvest as described in step 7.</li>
	<li>From the wells that have growth, isolate cellular gDNA using a commercially available DNA isolation kit (see the Table of Materials) and amplify the region surrounding the target base via PCR (718 bp; forward primer 5&#39;-ATGGTGAGCAAGGGCGAGGA-3&#39; and reverse primer 5&#39;-ACTTGTACAGCTCGTCCATGC-3&#39;).</li>
	<li>Perform DNA sequencing analysis on the samples.</li>
</ol>

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X., Zhang, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rationally+engineered+Cas9+nucleases+with+improved+specificity.">Rationally engineered Cas9 nucleases with improved specificity.</a> Science. 351 (6268), 84-88 (2016).</li></ol>

The authors having nothing to disclose.

Gene editing has emerged as a mainstream scientific discipline primarily because of the emergence of the CRISPR/Cas9 system. This remarkable pathway, which facilitates adoptive immunity in bacterial cells, has been repurposed as a molecular tool to enable genomic alteration in human chromosomes. The natural function of CRISPR/Cas9 is to disable viral DNA by introducing double-stranded DNA breaks leading to fragmentation30,31. This activity results in the destruction of invading exogenous DNA and leads to prokaryotic immunity that suppresses subsequent infection by the same viral particle. Amazingly, this system exhibits pneumonic behavior, in that it can reactivate specific CRISPR gRNA created by previous infection events.
The efficiency and accuracy of gene disruption catalyzed by CRISPR/Cas9 has been used in numerous eukaryotic genetic systems where the objective was to disable a functioning gene32,33. When reduced to its basal level, the process consists of two fundamental steps. The first is a double-stranded break, while the second relies on the inherent process of non-homologous end joining (NHEJ) to complete the knockout. In most cases, the double-stranded break results in the creation of blunt-ended, disengaged chromosomal segments, and enzymes involved in NHEJ react to the chromosomal break and act to conjoin the broken ends. This process can sometimes involve the loss of individual nucleotides at the severed site. The loss of even a few bases can cause a frameshift, and functional expression ceases. The use of CRISPR/Cas9 to create genetic knockout by engaging the natural process of NHEJ has revolutionized eukaryotic genetics; the loss of DNA at the target site is both predictable and expected.
In contrast to gene knockout, a number of researchers have been engaged in attempting to redirect and repurpose CRISPR/Cas9 activity toward the process of homology-directed repair. The aim of the studies is gene correction. In this strategy, CRISPR/Cas9 is combined with a donor DNA template that provides the genetic information to repair an inborn error or to insert mutations into healthy genes. Early reports highlighting the remarkable capacity of CRISPR/Cas9 to catalyze homology-directed repair indicated (directly or indirectly) that the process occurred in a highly precise fashion24,34. Our laboratory has been studying homology-directed repair or gene correction using single-stranded oligonucleotides in an attempt to define the mechanism and regulatory circuitry surrounding it15,17,18,23. We have focused primarily on the gene editing single point mutations, since this is the most basic genetic mutation known to be responsible for many inherited disorders. Our fundamental knowledge of gene editing led us to question the precision of CRISPR/Cas9 activity in these reactions, since the CRISPR/Cas9 function in gene knockout results in the formation of indels. We used a well-defined model system, rather than a non-selectable yet clinically relevant gene, to study on-site mutagenesis in a decidedly reductionist fashion. By using a simple target gene, upon which gene correction can be measured at both the genotypic and phenotypic levels, we reasoned that heterogeneity occurring at the target site could be identified in a reliable and robust fashion.
Our data confirm that the well-established gene editing system, consisting of a mutant eGFP gene integrated into HCT116 cells, can provide foundational information with regard to the generation of genetic lesions and the process of on-site mutagenesis. CRISPR/Cas9 and single-stranded oligonucleotide donor DNA molecules working in tandem can lead to the precise repair of the point mutation in the eGFP gene. We propose a new model for the repair of point mutations, a molecular pathway in which the donor DNA acts as a replication template for the repair of the mutant base, a process we have termed ExACT30. The targeted population, not exhibiting the corrected phenotype, displays a variety of cells containing heterogeneous and widely ranging DNA indels surrounding the target site. In approximately half the clones isolated from the uncorrected population, deletion mutagenesis was observed at the target site. Since no previous report has indicated that single-stranded oligonucleotides acting as single-agent gene-editing tools can induce indels at the target site, we conclude that CRISPR/Cas9 activity is responsible for these mutations.
In this manuscript, we provide a detailed methodology so that genetic heterogeneity at the target site can be measured in a reliable and robust fashion. While an enormous amount of attention has been paid to the analysis and mapping of off-site mutagenesis, it is likely that a heterogeneous mutation created at the target site will have a greater effect on the success or failure of gene editing in the clinical arena. Additional technologies or modified Cas9 proteins may be required to improve the precision of the homology-directed repair of inborn errors in mammalian cells35. Some of these technologies include the use of auxiliary oligonucleotides to act as a bridge, holding the chromosomal ends together and avoiding the destructive action of NHEJ. Defining the degree of heterogeneity at the target site as a result of gene-editing activity is and must be an important part of any protocol designed for therapeutic intervention.

Pioneering studies by Mandecki (1986) demonstrated permanent changes in plasmid DNA using oligonucleotides transformed into bacteria1, while Walder and Walder (1986) carried out similar studies in yeast2. Shortly thereafter, Sherman and colleagues published a series of papers in which single-stranded oligonucleotides were introduced into yeast cells to make heritable changes in genes3. Building on the seminal work in microbial cells, Kmiec and colleagues began to develop unique single-agent oligonucleotides that directed single base repair in mammalian cells4. This concept was based on biochemical data that showed that molecules bearing RNA bind more tightly to the target site than those composed entirely of DNA bases. Although there were numerous reports of gene-editing success using chimeric oligonucleotides5,6,7, the editing levels remained highly variable between experiments and across different target/cell combinations.
Single-stranded donor DNA is preferred to double-stranded donor DNA because it is less likely to integrate at random sites within the genome8. However, exogenously introduced single-stranded oligonucleotides are prone to rapid degradation by cellular nucleases. Several strategies to protect the termini of oligonucleotides from degradation have been employed. An exonuclease-protected, single-stranded DNA containing the desired sequence was sufficient to obtain an editing efficiency in the 0.1-1% range6. Improved and more consistent transfection techniques, coupled with phenotypic readouts, allowed for more consistent editing by multiple research groups8,9,10,11,12,13.
Over the past 10 years, there has been a significant effort to make target cells more amenable to gene editing. One strategy employs the modulation of the cell cycle14,15,16. While editing frequencies under normal reaction conditions with single-stranded oligonucleotides (ssODNs) introduced into cultured mammalian cells hovered between 0.1% and 1%, the frequencies of gene editing increased 3- to 5-fold when the oligonucleotides were introduced into cells during their transition through the S phase. In a separate series of experiments, Brachman and Kmiec (2005) demonstrated that relatively high levels of precise gene editing (3-5%) were obtained when 2&#39;3&#39;dideoxycytosine (ddC) was incubated in cells for 24 h prior to the addition of the oligonucleotide17. ddC reduces the rate of DNA replication fork movement, suggesting that gene editing by ssODNs in replicating cells likely involves the incorporation of ODNs into regions of active replication11,18. Taken together, these studies led to the concept that donor DNA becomes incorporated into a growing replication fork as part of the true mechanism of action of gene editing, independent of whether a double-stranded break is present or not19. Thus, the correction of a point mutation or the replacement of a segment of DNA within the chromosome occurs via a DNA pairing and single-strand assimilation pathway.
Inducing random double-stranded DNA breaks in growing cells with small-molecule agents creates an environment in which DNA replication events are stalled as the cell attempts to repair the damage20,21. This transitory slowdown of the replication forks enables a more efficient penetration of the chromatin structure by single-stranded editing oligonucleotides, improving target accessibility15. The creation of double-stranded DNA breaks by drugs such as VP16, bleomycin, or camptothecin22,23, has been shown to stimulate gene editing levels by nearly 10-fold, up to 6-8%. However, random double-stranded DNA breaks are undesirable in a therapeutic setting.
Gene editing with programmable nucleases and oligonucleotides is also stimulated during cell division24,25. When nucleic acid-based delivery was used to carry out homology-directed repair in synchronized HCT 116 cells, Rivera-Torres and colleagues demonstrated the same increase in targeting activity when transcription activator-like effector nucleases (TALENs) were employed with single-stranded oligonucleotides, both introduced into a replicating cell population26,27 . More recently, Bialk and colleagues showed that the repair of single base mutations with single-stranded oligonucleotides and an array of clustered regularly interspaced short palindromic repeats Cas9 (CRISPR/Cas9) molecules takes place with higher efficacy when the cell population is traversing S phase28. The CRISPR molecule is comprised of RNA and functions to identify the region within the target DNA sequence that is designated for cleavage. Cas9 is a bacterial enzyme whose function is to cleave double-stranded DNA in a nucleolytic exchange reaction. Thus, CRISPR positions the complex on the genomic target, and Cas9 nuclease executes the double-stranded break. Lin et al. (2014) also demonstrated the importance of the cell cycle for achieving high frequencies of gene editing, using a modified CRISPR/Cas9 ribonulceoprotien (RNP) complex-mediated delivery system in primary neonatal fibroblasts, human embryonic stem cells, and other cell lines24. Thus, the relationship established between gene editing and cell cycle progression for single-agent gene editing is applicable to combinatorial gene editing using programmable nucleases and donor DNA templates. While the combinatorial approach to gene editing has brought together a variety of partners with the donor DNA template, most workers in the field use CRISPR/Cas9 to provide the double-stranded break function. This choice is predicated upon the ease-of-use of this particular genetic tool and the flexibility with which it can be employed to disable gene function or to introduce a foreign piece of DNA into a specific site. The generation of a knockout is technically easier compared to a gene replacement, where the incorporation of &#34;correct&#34; or normal copies of a gene into the disease site must be carried out with precision. A number of researchers are identifying and studying the use of specific drugs and reagents that enable the correction of mutant bases and the insertion of normal genetic sequences in the proper position at elevated frequencies29.
Recently, Rivera-Torres et al.30 used combinatorial gene editing, taking advantage of the double-stranded break activity of a specifically designed CRISPR/Cas9 system and the genetic information provided by a single-stranded oligonucleotide donor DNA template, to repair a point mutation in a single copy of the enhanced green florescent protein (eGFP) gene integrated into HCT116 cells. The authors took advantage of this well-characterized model cell line to evaluate the specificity of cleavage surrounding the target site. The data reveal that heterogeneity at the target site exists, especially in cells that do not contain a corrected point mutation. In this manuscript, we detail and focus upon the methodology used by these workers to examine on-site mutational heterogeneity created by CRISPR/Cas9 gene editing.

<table>
	<tbody>
		<tr>
			<td>McCoy&#8217;s 5A Modified medium</td>
			<td>ATCC</td>
			<td>30-2007</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>ATCC</td>
			<td>30-2020</td>
			<td></td>
		</tr>
		<tr>
			<td>L-glutamine</td>
			<td>ATCC</td>
			<td>30-2214</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin Solution</td>
			<td>ATCC</td>
			<td>30-2300</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Phosphate Buffered Saline</td>
			<td>ATCC</td>
			<td>30-2200</td>
			<td>No Calcium or magnesium</td>
		</tr>
		<tr>
			<td>Trypsin EDTA Solution</td>
			<td>ATCC</td>
			<td>30-2101</td>
			<td></td>
		</tr>
		<tr>
			<td>Aphidicolin 1mg</td>
			<td>Thermo Fisher</td>
			<td>AC611970010</td>
			<td></td>
		</tr>
		<tr>
			<td>Alt-R CRISPR crRNA, 10 nmol</td>
			<td>IDT</td>
			<td></td>
			<td>No catalog number since its made to your specific gene target.</td>
		</tr>
		<tr>
			<td>CRISPR-Cas9 tracrRNA, 20 nmol</td>
			<td>IDT</td>
			<td>1072533</td>
			<td></td>
		</tr>
		<tr>
			<td>S.p. Cas9 Nuclease 3NLS, 500 &#181;g</td>
			<td>IDT</td>
			<td>1074182</td>
			<td></td>
		</tr>
		<tr>
			<td>IDTE Buffer</td>
			<td>IDT</td>
			<td>11-01-03-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeus Electroporation Cuvettes 0.4 Cm</td>
			<td>VWR</td>
			<td>10497-474</td>
			<td></td>
		</tr>
		<tr>
			<td>Gene Pulser Xcell Electroporation Systems</td>
			<td>BioRad</td>
			<td>1652660</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin 
			lyophilized powder, crystallized, &#8805;98.0% (GE)</td>
			<td>Sigma</td>
			<td>05470-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA (0.5 M), pH 8.0</td>
			<td>ThermoFisher Scientific</td>
			<td>AM9260G</td>
			<td></td>
		</tr>
		<tr>
			<td>Propidium Iodide - 1.0 mg/mL Solution in Water</td>
			<td>ThermoFisher Scientific</td>
			<td>P3566</td>
			<td></td>
		</tr>
		<tr>
			<td>DNeasy Blood &#38; Tissue Kit (250)</td>
			<td>Qiagen</td>
			<td>69581</td>
			<td></td>
		</tr>
		<tr>
			<td>6-well Standard Line Multiwell Cell Culture Plates</td>
			<td>VWR</td>
			<td>10062-892</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri Dishes</td>
			<td>Fisher</td>
			<td>12-565-90</td>
			<td></td>
		</tr>
		<tr>
			<td>Conical Tubes (15 mL) (racked)</td>
			<td>Thermo Fisher</td>
			<td>AM12500</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue Solution, 0.4%</td>
			<td>Thermo Fisher</td>
			<td>15250061</td>
			<td></td>
		</tr>
		<tr>
			<td>Reduced Serum Medium</td>
			<td>Thermo Fisher</td>
			<td>31985062</td>
			<td></td>
		</tr>
	</tbody>
</table>

a standard methodology to examine on-site mutagenicity as a function of point mutation repair catalyzed by crispr/cas9 and ssodn in human cells

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.

We used a model system to study gene editing in mammalian cells, which relies upon the correction of a point mutation embedded within the eGFP gene integrated as a single copy in HCT116 cells. It is important to note that this is a single-copy gene; thus, a less complicated view of DNA alterations or mutagenesis can be made. Figure 1A displays the mutant eGFP gene sequence with the targeted base, the third base of the stop codon TAG, highlighted in red. The 72-base oligonucleotide, which is partially complementary to the non-transcribed strand of the eGFP gene (72NT) and is designed to induce the base exchange from a G to a C, is also illustrated. In addition, a CRISPR, designated as 2C, with the indicated protospacer sequence in a 5&#39; to 3&#39; orientation, is also depicted in Figure 1A. To carry out this gene-editing reaction, we used a ribonucleoprotein (RNP) consisting of the CRISPR (cr) RNA and the tracr (tr) RNA coupled to purified Cas9 protein (Figure 1B), instead of using a mammalian expression vector consisting of the Cas9 gene and the appropriate specific guide RNA sequence.
When the specifically designed RNP particle is delivered with the single-stranded oligonucleotide into HCT116 cells by electroporation, gene editing, evidenced by the repair of the single base mutation in eGFP, is observed after 72 h of incubation using a flow cytometer. Functional repair is observable by the emergence of green fluorescence in targeted cells, cells that can also be separated from the entire population by sorting because of this fluorescence. As shown in Figure 2, a gradual dose response can be seen as the coordinated levels of RNP and 72NT increase. The molecular ratio, picomoles of RNP, and micromolar concentration of the 72NT as displayed in the figure, are based on optimal dosages used in gene-editing reactions that are dependent on the introduction of Cas9 and specific guide RNA from transfected expression vectors. The inset in Figure 2 displays a gene-editing reaction carried out in the absence of the RNP particle. Here, approximately 1% of the targeted cells are corrected when a 10-fold higher concentration of the 72NT oligonucleotide is used in the single-agent gene-editing reaction.
In order to determine if genetic heterogeneity exists at the target site-the so-called on-site mutagenesis effect-we decided to examine the outcome of gene editing activity in individual cells. While much more laborious than examining the overall population, a true measure of genetic footprints or lesions can be ascertained when the genome of clonally expanded cells is examined. We repeated the experiment described in Figure 2, this time only using 100 pmol of RNP complex and 2.0 &#181;mol of the 72NT oligonucleotide. As above, HCT 116 cells were synchronized for 24 h with aphidicholine and arrested at the G1/S border. 4 h afterwards, the cells were released and the gene-editing tools were introduced by electroporation. 72 h later, the cells were analyzed using FACS and sorted individually into 96-well plates (the experimental process is illustrated in Figure 3). Cells displaying green fluorescence were sorted by flow cytometry into individual wells of a 96-well plate for clonal expansion. Importantly, cells lacking eGFP expression were also isolated and sorted in a similar fashion for expansion under the same conditions.
After 14 days of growth, most of the individual clones had expanded sufficiently to enable DNA isolation. As such, 16 clones of the eGFP-positive samples were selected, and the genetic integrity surrounding the target site was analyzed by DNA sequencing. Information surrounding the DNA sequence of alleles within the population was generated using Sanger sequencing, assembled using sequence visualization software to compare the sequence of a wildtype allele (Figure 4A). The cut site of the RNP complex is indicated by a black arrow, located on the green bar (2C crRNA). As also shown in Figure 4A, all 16 eGFP-positive cells contain the predicted nucleotide exchange at the target site. The converted C residue is highlighted in red, and the peak profile reflecting that precise change is provided underneath the eGFP-positive sequence. In a similar fashion, 15 non-green clonal isolates, sufficiently expanded to enable DNA extraction and sequencing, were analyzed for heterogeneity at the target site. As predicted, in approximately half the samples, no DNA base exchange was observed. This is reflected in the maintenance of the G residue at the target site, as shown in Figure 4B. The remainder of the clonal expansions examined in these experiments displayed a heterogeneous population of deletion mutations, therefore accounting for the lack of green fluorescence. The deletion size ranged from one base to 19 bases. It is important to note that we have only examined 15 samples of the eGFP-positive cells, and while we believe that this is quite representative of the type of genetic lesions left behind by CRISPR/Cas9 activity, there is a possibility that other types or forms of indels could be present in the targeted population.
Taken together, the results displayed in Figure 4 confirm the phenotypic readout in the eGFP targeting system. Conversion of the G to C nucleotide enables the emergence of green fluorescence in corrected HCT116 cells. No base substitution surrounding the target site has been observed in the clones isolated for this experiment or in previous experiments27,28. The data also demonstrate that cells failing to undergo gene editing via point-mutation repair remain uncorrected but, in some cases, not unaltered, with a range of genetic heterogeneity surrounding the target site.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56195/56195fig1.jpg" /> 
Figure 1. (A) Model system for the gene editing of the mutant eGFP gene. The appropriate segments of the wildtype and mutated eGFP gene with the targeted codon, located in the center of the sequence, are displayed in green and red, respectively. The nucleotide targeted for exchange is bolded and underlined. The highlighted bases in blue represent the 2C CRISPR protospacer sequence, and the orange bases highlight the PAM site. The oligonucleotide used in these experiments is 72 bases in length, bearing phosphorothioate modified linkages at the three terminal bases; the 72-mer targets the non-transcribed (NT) strand (72NT). (B) CRISPR/Cas9 ribonucleoprotein assembly reaction. crRNA provides target specificity (20 bases, red section) corresponding to the 2C protospacer sequence and an interaction domain (blue) with the tracrRNA (green). crRNA and tracrRNA are annealed in equimolar concentrations. Cas9 protein (gray) is added to complete the RNP assembly. Guide RNAs (gRNAs) direct and activate the Cas9 endonuclease, which then cleave the target DNA. The lower section of the figure shows the 2C seed sequence and the tracrRNA sequence. This figure was modified from Rivera-Torres, N. et al. (2017). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56195/56195fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56195/56195fig2.jpg" /> 
Figure 2. Gene editing is dose-dependent when directed by the RNP and the ssODN. Synchronized and released HCT 116-19 cells were electroporated with 24-120 pmol of CRISPR/Cas9 RNP and 0.6-3.0 &#181;M of 72mer. After a 72-h recovery period, gene-editing activity was measured using a flow cytometer. Gene editing is displayed as the correction efficiency (%), determined by the number of viable eGFP-positive cells divided by the total number of viable cells in the population. Error bars are produced from three sets of data points generated over three separate experiments using basic calculations of standard error. Inset: Single-agent gene editing. Gene-editing activity directed by the single-stranded oligonucleotide (72NT) in the absence of the RNP complex under identical conditions is presented as a function of increasing concentration. This figure was modified from Rivera-Torres, N. et al. (2017). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56195/56195fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56195/56195fig3.jpg" /> 
Figure 3. Experimental strategy for the isolation of single-cell clones. Cells exhibiting eGFP expression were scored as positive and sorted using a flow cytometer as single cells into individual wells for clonal expansion. Cells lacking eGFP expression were isolated and sorted in a similar fashion and expanded under the same conditions. The DNA was then isolated and the eGFP gene was amplified and subjected to Sanger sequencing to analyze the gene-editing activity surrounding the target site. This figure was modified from Rivera-Torres, N. et al. (2017). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56195/56195fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56195/56195fig4.jpg" /> 
Figure 4. (A) Allelic analysis of eGFP-positive cells expanded as a clonal population. Clonally isolated and expanded eGFP-positive samples (sixteen clones) were analyzed at the site surrounding the targeted base and DNA from each, harvested, purified, amplified, and sequenced. Allelic analysis was carried out using Sanger sequencing, assembled using sequence visualization software and compared to the sequence of a wildtype allele, which is illustrated at the top of the figure. The cut site of the RNP complex is indicated as a small black arrow located on the green bar (2C crRNA). (B) Allelic analysis of eGFP-negative cells expanded as a clonal population. Fifteen individual samples, expanded from clones originating from the uncorrected population, were randomly selected and analyzed for indel formation at the site surrounding the target nucleotide. As above, allelic analysis was carried out using Sanger sequencing and assembled using a sequence visualization software. Once again, the sequence of a wildtype allele at the top of the figure, along with the cut site of the RNP, are presented. This figure was modified from Rivera-Torres, N. et al. (2017). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56195/56195fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells at JoVE.com

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site&#8212;in essence, analyzing onsite mutagenesis.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point ...

a-standard-methodology-to-examine-on-site-mutagenicity-as-function

Nanyang Technological University

Research

JoVE Journal

Genetics

7.7K Views.  Christiana Care Health Services. This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site—in essence, analyzing onsite mutagenesis.

Video: A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Scientific knowledge is intrinsically linked to available technologies and methods. This article will present two methods that allowed for the identification and verification of a drug resistance gene in the Apicomplexan parasite Toxoplasma gondii, the method of Quantitative Trait Locus (QTL) mapping using a Whole Genome Sequence (WGS) -based genetic map and the method of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9 -based gene editing. The approach of QTL mapping allows one to test if there is a correlation between a genomic region(s) and a phenotype. Two datasets are required to run a QTL scan, a genetic map based on the progeny of a recombinant cross and a quantifiable phenotype assessed in each of the progeny of that cross. These datasets are then formatted to be compatible with R/qtl software that generates a QTL scan to identify significant loci correlated with the phenotype. Although this can greatly narrow the search window of possible candidates, QTLs span regions containing a number of genes from which the causal gene needs to be identified. Having WGS of the progeny was critical to identify the causal drug resistance mutation at the gene level. Once identified, the candidate mutation can be verified by genetic manipulation of drug sensitive parasites. The most facile and efficient method to genetically modify T. gondii is the CRISPR/Cas9 system. This system comprised of just 2 components both encoded on a single plasmid, a single guide RNA (gRNA) containing a 20 bp sequence complementary to the genomic target and the Cas9 endonuclease that generates a double-strand DNA break (DSB) at the target, repair of which allows for insertion or deletion of sequences around the break site. This article provides detailed protocols to use CRISPR/Cas9 based genome editing tools to verify the gene responsible for sinefungin resistance and to construct transgenic parasites.

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Details are presented on how QTL mapping with a whole genome sequence based genetic map can be used to identify a drug resistance gene in Toxoplasma gondii and how this can be verified with the CRISPR/Cas9 system that efficiently edits a genomic target, in this case the drug resistance gene.

Scientific knowledge is intrinsically linked to available technologies and methods. This article will present two methods that allowed for the ...

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system provides a promising tool for genetic engineering, and opens up the possibility of targeted integration of transgenes. We describe a homology-mediated end joining (HMEJ)-based strategy for efficient DNA targeted integration in vivo and targeted gene therapies using CRISPR/Cas9.

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration ...

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical regions, is caused by infection with Plasmodium parasites. The development of more effective drugs to combat malaria can be accelerated by improving our understanding of the biology of this complex parasite. Genetic manipulation of these parasites is key to understanding their biology; however, historically the genome of P. falciparum has been difficult to manipulate. Recently, CRISPR/Cas9 genome editing has been utilized in malaria parasites, allowing for easier protein tagging, generation of conditional protein knockdowns, and deletion of genes. CRISPR/Cas9 genome editing has proven to be a powerful tool for advancing the field of malaria research. Here, we describe a CRISPR/Cas9 method for generating glmS-based conditional knockdown mutants in P. falciparum. This method is highly adaptable to other types of genetic manipulations, including protein tagging and gene knockouts.

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

We describe a method for generating glmS-based conditional knockdown mutants in Plasmodium falciparum using CRISPR/Cas9 genome editing.

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical ...

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or C-terminal fluorescent tags. The prokaryotic CRISPR/Cas9 system (clustered regularly interspaced short palindromic repeats/CRISPR-associated 9) may be used to introduce large exogenous sequences into genomic loci via homology directed repair (HDR). To achieve the desired knock-in, this protocol employs the ribonucleoprotein (RNP)-based approach where wild type Streptococcus pyogenes Cas9 protein, synthetic 2-part guide RNA (gRNA), and a donor template plasmid are delivered to the cells via electroporation. Putatively edited cells expressing the fluorescently tagged proteins are enriched by fluorescence activated cell sorting (FACS). Clonal lines are then generated and can be analyzed for precise editing outcomes. By introducing the fluorescent tag at the genomic locus of the gene of interest, the resulting subcellular localization and dynamics of the fusion protein can be studied under endogenous regulatory control, a key improvement over conventional overexpression systems. The use of hiPSCs as a model system for gene tagging provides the opportunity to study the tagged proteins in diploid, nontransformed cells. Since hiPSCs can be differentiated into multiple cell types, this approach provides the opportunity to create and study tagged proteins in a variety of isogenic cellular contexts.

Described here is a protocol for tagging endogenously expressed proteins with fluorescent tags in human induced pluripotent stem cells using CRISPR/Cas9. Putatively edited cells are enriched by fluorescence activated cell sorting and clonal cell lines are generated.

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or ...

CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of CRISPR/Cas9 elements into the target cells by transfection is a prerequisite for efficient gene editing. This protocol demonstrates that tube electroporation (TE) machine-mediated delivery of CRISPR/Cas9 ribonucleoprotein (RNP), along with single-stranded oligodeoxynucleotide (ssODN) donor templates to different types of mammalian cells, leads to robust precise gene editing events. First, TE was applied to deliver CRISPR/Cas9 RNP and ssODNs to induce disease-causing mutations in the interleukin 2 receptor subunit gamma (IL2RG) gene and sepiapterin reductase (SPR) gene in rabbit fibroblast cells. Precise mutation rates of 3.57%-20% were achieved as determined by bacterial TA cloning sequencing. The same strategy was then used in human iPSCs on several clinically relevant genes including epidermal growth factor receptor (EGFR), myosin binding protein C, cardiac (Mybpc3), and hemoglobin subunit beta (HBB). Consistently, highly precise mutation rates were achieved (11.65%-37.92%) as determined by deep sequencing (DeepSeq). The present work demonstrates that tube electroporation of CRISPR/Cas9 RNP represents an efficient transfection protocol for gene editing in mammalian cells.

Presented here is a protocol for efficient CRISPR/Cas9 ribonucleoprotein-mediated gene editing in mammalian cells using tube electroporation.

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of ...

Generation of Defined Genomic Modifications Using CRISPR-CAS9 in Human Pluripotent Stem Cells

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise heterozygous genetic modifications is challenging due to CRISPR-CAS9 mediated indel formation in the second allele. Here, we demonstrate a protocol to help overcome this difficulty by using two repair templates in which only one expresses the desired sequence change, while both templates contain silent mutations to prevent re-cutting and indel formation. This methodology is most advantageous for gene editing coding regions of DNA to generate isogenic control and mutant human stem cell lines for studying human disease and biology. In addition, optimization of transfection and screening methodologies have been performed to reduce labor and cost of a gene editing experiment. Overall, this protocol is widely applicable to many genome editing projects utilizing the human pluripotent stem cell model.

This protocol provides a method to facilitate the generation of defined heterozygous or homozygous nucleotide changes using CRISPR-CAS9 in human pluripotent stem cells.

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise ...