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.
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.
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 2'3'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 "correct" 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.
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.
The authors have nothing to disclose.
The authors have no acknowledgments.
McCoy’s 5A Modified medium | ATCC | 30-2007 | |
Fetal Bovine Serum (FBS) | ATCC | 30-2020 | |
L-glutamine | ATCC | 30-2214 | |
Penicillin-Streptomycin Solution | ATCC | 30-2300 | |
Dulbecco's Phosphate Buffered Saline | ATCC | 30-2200 | No Calcium or magnesium |
Trypsin EDTA Solution | ATCC | 30-2101 | |
Aphidicolin 1mg | Thermo Fisher | AC611970010 | |
Alt-R CRISPR crRNA, 10 nmol | IDT | No catalog number since its made to your specific gene target. | |
CRISPR-Cas9 tracrRNA, 20 nmol | IDT | 1072533 | |
S.p. Cas9 Nuclease 3NLS, 500 µg | IDT | 1074182 | |
IDTE Buffer | IDT | 11-01-03-01 | |
Zeus Electroporation Cuvettes 0.4 Cm | VWR | 10497-474 | |
Gene Pulser Xcell Electroporation Systems | BioRad | 1652660 | |
Bovine serum albumin lyophilized powder, crystallized, ≥98.0% (GE) |
Sigma | 05470-1G | |
EDTA (0.5 M), pH 8.0 | ThermoFisher Scientific | AM9260G | |
Propidium Iodide – 1.0 mg/mL Solution in Water | ThermoFisher Scientific | P3566 | |
DNeasy Blood & Tissue Kit (250) | Qiagen | 69581 | |
6-well Standard Line Multiwell Cell Culture Plates | VWR | 10062-892 | |
Petri Dishes | Fisher | 12-565-90 | |
Conical Tubes (15 mL) (racked) | Thermo Fisher | AM12500 | |
Trypan Blue Solution, 0.4% | Thermo Fisher | 15250061 | |
Reduced Serum Medium | Thermo Fisher | 31985062 |