PSC Resource Handbook – Genome editing

Broadly, gene engineering or genome editing involves changing an organism’s DNA through sequence disruption, replacement, or addition. While approaches for genetic manipulation of mouse ESCs have been widely used for decades in the generation of transgenic mouse models, recent advances in genome editing technologies now make this a tool that can readily be applied to hPSCs.

The capacity of hPSCs to self-renew and differentiate makes them ideally suited for generating both disease models and cells at the scale needed for drug development and cell therapy applications. The ability to genetically modify hPSCs further increases their usefulness for both research and clinical applications, enabling the generation of models for genetically complex disorders.

The dovetailing of iPSC and genome editing approaches supports a diverse range of applications (Figure 4.1), including:

• Generation of disease models by introducing known mutations in control iPSCs 
• Generation of isogenic controls by correcting mutations in patient-specific iPSCs 
• Testing the disease relevance of implicated genes by selectively knocking down or knocking out loci in wild type iPSCs
• Deconvoluting contributions of multiple loci in polygenic diseases by selectively correcting individual loci
• Generation of lineage-specific reporter lines
• Generation of gene-corrected disease-relevant cell types for cell replacement therapies 

To learn more about our genome editing products and services, go to thermofisher.com/genomeedit

Figure 4.1. Generation of disease-specific and isogenic control (wild type, WT) iPSCs and disease-relevant cell types using genome editing. Somatic cells such as fibroblasts or blood cells are isolated from healthy or patient donors (A) and reprogrammed to generate control and disease-specific iPSCs (B). Genome editing can be used to introduce disease-relevant mutations into control iPSCs to generate disease-specific iPSCs. Alternatively, gene correction can be used to generate isogenic controls from disease-specific iPSCs (C). Disease phenotypes can potentially be quantified by comparing the behavior of disease-relevant cell types, such as neurons or cardiomyocytes derived from control and patient-specific iPSCs, in functional assays (D). 
 

The design of a successful genome editing project will be impacted by several choices, starting with the choice of genome editing tool. Further factors that need to be considered include the delivery method, cell culture system, and editing validation technologies (Figure 4.2). Thermo Fisher Scientific provides a suite of tools optimized to help ensure a high level of success. These will be addressed in greater detail in following sections. 

Genome editing used to be a laborious and inefficient process, using either random mutagenesis or older technologies such as zinc finger nucleases that were difficult to design and target to specific sites in the genome. Recent advances in gene engineering tools now allow exquisite precision and control in a user-friendly workflow.

Genome editing is now routinely being achieved through the use of technology derived from clustered regularly interspaced short palindromic repeats (CRISPRs) and transcription activator–like (TAL) effectors. CRISPR gRNA and TAL effectors target nucleases to specific sites in the genome, creating double-stranded breaks at desired locations (Figure 4.3).

The natural repair mechanisms of the cell heal the break by either homologous recombination or nonhomologous end-joining (NHEJ). Homologous recombination is more precise because it requires a template for repair. By providing the cell with a synthetic template containing a sequence of interest, for example a disease-specific mutation, the researcher can introduce this sequence into the genome. In contrast, double-strand break repair by NHEJ is more error-prone, frequently introducing errors such as small insertions or deletions (indels). Since the resulting frameshift often leads to a nonfunctional gene, this approach can be harnessed to rapidly and efficiently generate specific gene knockouts.

Originally, Invitrogen CRISPR genome editing technologies were considered to be more efficient but also more prone to off-target effects when compared to TAL technologies. Recent advances in the tools and reagents available for both gene editing systems have negated some of these differences, and CRISPR and TAL technologies are now both widely used in a broad range of applications. A highlight of the benefits and limitations of the two technologies can be found in Table 4.1.

Figure 4.3. Genome editing technologies available.
 

Ease of design and construction

TAL target recognition occurs as a result of protein–DNA interaction between the TAL effector proteins and the DNA target sequence. Editing of a new site, therefore, requires the design and construction of a new protein specific to that sequence. Although the process of TAL effector protein cloning has been greatly streamlined through the availability of repeat combinations of modules, this can still present a significant pain point for users. On the other hand, CRISPR gene editing relies on the formation of a RNA–DNA complex. Targeting of a new site simply requires the design and generation of a 20 nt gRNA that provides specificity to the system. This can be achieved both readily and cost effectively, which is often particularly appealing to academic scientists.

Target site selection

While the nature of TAL targeting means that TAL pairs can be engineered to target virtually any position in the genome, CRISPR functionality requires that the gRNA target sequence immediately precedes a three-base PAM sequence, typically NGG. This is not typically an issue when designing knockouts, where there is greater flexibility in the exact gene region being targeted. However, this can severely limit the availability of sites that are amenable to homologous recombination using donor DNA, since the editing efficiency will depend on the distance from the PAM site. Therefore, when designing an elegant knock-in experiment, TAL effectors could be a better tool to use.

Specificity

TALs have traditionally been thought of as having fewer off-target effects due to the system’s longer DNA-binding sites, which reduce the likelihood of homology in other areas of the genome. The high degree of specificity inherent to TAL-based gene editing is particularly important for potential cell therapy applications, for example in the use of CAR T cells. In contrast, CRISPR tools have a comparatively high tolerance for mismatches between the gRNA and intended DNA target, which can negatively impact specificity. However, newer tools such as Invitrogen TrueCut Cas9 Protein v2, rather than DNA- or RNA-based systems, have a reduced half life and more acute effect once introduced into cells and this can greatly reduce off-target effects.

Efficiency

The high cleavage efficiencies achieved with CRISPR may be a deciding factor for many R&D applications. This feature also makes CRISPR suitable for high-throughput applications using lentiviral-based CRISPR libraries, as an alternative to RNAi-based screening, and further makes it amenable to simultaneous targeting of multiple sites. The efficiency of editing with TAL pairs can be impacted by the system’s sensitivity to CpG methylation. This can render some TAL pairs ineffective if they target areas with high levels of methylation.

Delivery

Both TAL and CRISPR can be introduced via lipid transfection and electroporation.

Licensing

For researchers pursuing commercial applications of their gene edited products, it is worth noting that TAL technologies provide a clear licensing path for this purpose. The intellectual property landscape relating to CRISPR editing has been less clear. It may still be some time before clear guidelines for the appropriate commercial use of CRISPR tools become available. For more information about licensing, contact us at outlicensing@thermofisher.com.

Genome editing uses engineered nucleases in conjunction with endogenous repair mechanisms to alter the DNA in a cell. The CRISPR-Cas9 system takes advantage of a short gRNA to target the bacterial Cas9 endonuclease to specific genomic loci. Because the gRNA supplies the specificity, changing the target only requires a change in the design of the sequence encoding the gRNA.

The CRISPR-Cas9 system is composed of a DNA endonuclease called Cas9 and a short, noncoding guide RNA (gRNA) that has two molecular components: a target-specific CRISPR RNA (crRNA) and an auxiliary trans-activating crRNA (tracrRNA). The gRNA guides the Cas9 protein to a specific genomic locus via base pairing between the crRNA sequence and the target sequence within the genomic locus (Figure 4.4).

With their highly flexible yet specific targeting, CRISPRCas9 systems can be manipulated and redirected to become powerful tools for genome editing. CRISPR-Cas9 technology permits targeted gene cleavage and gene editing in a variety of cells, and because the endonuclease cleavage specificity in CRISPR-Cas9 systems is guided by RNA sequences, editing can be directed to virtually any genomic locus by engineering the gRNA sequence and delivering it along with the Cas endonuclease to the target cell.

CRISPR-Cas9 delivery formats

Different formats of CRISPR tools are available for specific research needs including: CRISPR-Cas9 all-in-one expression plasmids, CRISPR-Cas9 mRNA and gRNA, Cas9 protein, Cas9 iPSC, and CRISPR libraries (Figure 4.5). PSCs can readily be edited using CRISPR plasmid vectors and mRNA; however, the highest cleavage efficiencies in hPSCs are observed using TrueCut Cas9 Protein v2.

Figure 4.5. Available CRISPR-Cas9 delivery formats.
 

Table 4.2. Comparison of Invitrogen CRISPR technologies.
 

TrueCut Cas9 Protein V2

The newly developed TrueCut Cas9 Protein v2 is a wild type Cas9 protein that has been designed to deliver consistently higher editing efficiency across a range of gene targets and cell types.

• High editing efficiency in all tested cell lines, including standard, immune, primary, and stem
• Up to 2x higher editing efficiency in difficult targets, compared to that obtained with other supplier’s kits 
• High quality—manufactured under strict ISO 13485 quality standards 
• Validated protocols—achieve success faster using protocols optimized for an extensive list of cell types

CRISPR gRNA

The design, production, and delivery of high-quality gRNAs are critical to achieving a successful result when using a CRISPR-Cas9 system for gene editing. CRISPR gRNAs are available in multiple formats including transfection-ready gRNAs and lentiviral delivery systems. Use our selection guide below to find the right gRNAs for your experiment (Table 4.3).

TrueGuide Synthetic gRNA

TrueGuide Synthetic gRNAs are ready-to-transfect synthetic gRNAs designed and validated to work with the Invitrogen suite of genome editing tools to provide consistent high-efficiency editing. Whether you need an economical solution for routine editing tasks or you want to drive maximum editing efficiency, particularly in primary or stem cells, TrueGuide Synthetic gRNAs offer the reagents required to introduce your specific edit in your cell line (Figure 4.6).

Figure 4.6. TrueGuide gRNAs perform equally to IVT gRNA in combination with TrueCut Cas9. Indel induction efficiency is shown for two targets, SNP induction for one (%HDR). tg: two-piece TrueGuide gRNA, sg: single-piece TrueGuide gRNA, IVT: IVT gRNA.
 

CRISPR gene editing workflow

Once a specific CRISPR format has been selected, it is introduced into the target cells via lipid-mediated transfection or electroporation. Cells are plated at low density to allow for expansion of clonal colonies. These are then selected and screened for gene editing events. A sample workflow is shown in Figure 4.7. 

Figure 4.7. Standard gene editing workflow using CRISPR-Cas9 technology. Following the expansion of hPSCs with the StemFlex media system, cells are singularized and electroporated using the Neon system to introduce precomplexed TrueCut Cas9 protein v2 and control TrueGuide Synthetic gRNA control. Cells recover in StemFlex medium in the presence or absence of RevitaCell supplement on either Geltrex substrate or rhLaminin-521. Following 48–72 hours of recovery, cleavage efficiency is assessed using the GeneArt Genomic Cleavage Detection Kit. Pending successful cleavage, cells are recovered and expanded for 2 passages prior to clonal expansion. At this time, viable PSCs are flow-sorted based on expression of Tra-1-60 and the absence of PI expression. Subsequently cells are plated, 1 cell/well in a 96-well plate in StemFlex Medium on rhLaminin-521–coated 96-well plates. Following 14 days of recovery, successful clonal expansion is determined, followed by confirmation of successful gene editing of clonally established lines through sequencing.
 

While gene editing in workhorse cell lines like 293Ts has become quite routine, successful editing of PSCs can still require significant optimization. In addition to optimizing the delivery of Cas9 and gRNA, the recovery following electroporation and later clonal expansion can be particularly challenging in PSCs. Below we outline our best practices that will maximize your chances of success.

1. Culture—While the maintenance of PSCs leading up to gene editing may seem like a routine step, having your cells adapted to a media that will support the survival of PSCs during the bottlenecks in the gene editing workflow is particularly critical. The StemFlex Medium has been specifically developed for optimal performance in these challenging applications. It is a feeder-free medium that eliminates the need for enzymatically or mechanically removing mouse embryonic fibroblasts (MEFs) that can otherwise lower transection efficiencies. Unlike other feeder-free culture systems, StemFlex Medium supports robust cell survival following singularization and transfection with a Cas9–gRNA complex, even in the absence of ROCK inhibitors. It also supports clonal expansion from a single cell and is compatible with every-other-day feed schedules, which not only reduces workload prior to gene editing but is also a critical step during the clonal expansion stage. 
2. Delivery—Delivery of Cas9 and the target gRNA can be achieved via either lipid transfection using reagents such as Invitrogen Lipofectamine Stem Transfection Reagent. However, the highest cleavage efficiencies are obtained using the TrueCut Cas9 Protein v2 and Invitrogen TrueGuide Synthetic gRNA RNP transfection via electroporation with the Invitrogen Neon system(Figures 4.8 and 4.9). Optimal Neon settings can vary depending on the culturing conditions, cell lines being used, and content being delivered. These may even differ between PSC lines. We therefore recommend that users test all 24 electroporation conditions using the optimization protocol and empirically determine the best program for their particular cell line, considering both editing efficiency and cell viability (Figure 4.10).
3. Recovery—For many scientists, post-electroporation recovery with Cas9 RNP presents the first significant bottleneck during gene editing of PSCs. Not only do cells undergo the stressful electroporation process, but the protocol necessitates the singularization of cells prior to electroporation. StemFlex Medium addresses this pain point by allowing for optimal recovery following electroporation with Cas9 RNP complex (Figure 4.11). 
4. Analysis—Prior to continuing with clonal expansion, it is recommended to first confirm successful cleavage at the target site. This allows for the identification of the condition with the best editing efficiency, which can then be used for further cell line generation. Please see Section 4.6. “Screening methods for TAL and CRISPR” for additional details.
5. Clonal expansion—The establishment of a successfully edited clonal PSC line is undoubtedly the most challenging step of the gene editing workflow. Cells can be plated at limiting dilution, however to reduce the number of wells screened, we recommend flow sorting and plating of viable, pluripotent cells (TRA-1-60+, PI– ) at 1 cell/well of a 96-well plate (Figure 4.12). StemFlex Medium is the only medium tested that allows for establishment of a clonal line from a single cell under these stressful conditions in the absence of ROCK inhibitor. Typically, 20–50% clonal recovery is achieved using this medium.
6. Sequencing—The correct editing of the desired target sequence in clonal populations can be confirmed via Sanger sequencing at the site of interest or via Ion PGM sequencing to simultaneously confirm the absence of off-target effects. Please see section 4.5, “Screening methods for TAL and CRISPR”, for additional details.

CRISPR gene editing of a disease-relevant target

In a proof-of-principle study, disease-causing mutations were introduced into a hiPSC line stable expressing Cas9. A number of cardiac and Parkinson’s disease–specific mutations were introduced through Neon electroporation delivery of gRNA and a single-stranded oligo donor carrying the SNP to be modified (Figure 4.13). Varying editing efficiencies were observed both for indel formation and homology-driven repair-mediated SNP introduction in pools generated for the mutations in the different genomic loci (Figure 4.13A). Isolation of single-cell clones via FACS yielded different distributions for the presence of wild type, indel, heterozygous, and homozygous edits for each of the targets, and in every case, a homozygous clone was identified regardless of the efficiency in the edited pools (Figure 4.13B). Through Ion PGM sequencing, clonality can then be verified quantitatively by calculating allele ratios from over 10,000 reads. Wild type or homozygote edits would have ~100% of one allele, and a heterozygote would have ~50% of one allele and ~50% of the other. These clonal lines can then be differentiated into desired cell types to model the disease of interest. 

Figure 4.13. Generation of disease models through the introduction of SNPs into a wild type iPSC overexpressing Cas9.(A) Indel and homology-driven repair efficiency for the indicated disease targets. (B) WT/indel/homozygous SNP/heterozygous SNP distribution of clonal lines isolated from genome edited pools via single-cell FACS sorting. (C) Next-generation sequencing analysis to confirm clonality of isolated hiPSC lines.

Transcription activator–like (TAL) effector proteins are plant pathogenic bacterial proteins that bind to specific DNA sequences and act as transcription factors during plant pathogenesis. The TAL DNA-binding domain contain highly conserved 32–34 amino acid repeat sequence except the amino acids in positions 12 and 13. These two amino acids, called the repeat variable di-residue or RVD, dictates specificity of each repeat to a single specific nucleotide within the target sequence. Because of the modular domain structure and well-defined amino acid–to-nucleotide code, fusion proteins containing TALs conjugated with various functional domains can be targeted to very specific loci within the genome.

The genome editing processes in products such as Invitrogen GeneArt PerfectMatch TALs use pairs of TALs that are fused to truncated FokI nuclease. FokI nuclease functions as a homodimer, and creates a double-stranded break in the DNA flanked by the TAL-binding sites. In the absence of DNA that shares homology across the region containing the break, the cell’s natural machinery will attempt to repair the break by NHEJ, which can lead to indels. In protein-coding regions, these indels can cause frameshift mutations that can result in a gene disruption (knockout). 

When this break is created in the presence of DNA that shares homology across the region, homology-directed repair can occur, which allows the added DNA to be incorporated at the site of the break. In this manner, specific bases or sequences can be introduced within user defined locations within the genome (Figure 4.14). A sample workflow for gene editing of iPSCs after culturing using TALs involves the following steps (Figure 4.15):

• Design and synthesis of TAL constructs
• Transfection and electroporation of iPSCs with TAL constructs in the presence or absence of donor DNA
• Clonal recovery of cells
• Picking and screening of colonies to determine successful cleavage and editing
• Expansion, characterization, and banking of successfully edited clones 

Figure 4.15. Workflow for genome editing of iPSCs using TAL technology. Feeder-free iPSCs cultured in StemFlex Medium and on Geltrex matrix– coated plates are treated with RevitaCell Supplement before dissociation with StemPro Accutase Cell Dissociation Reagent. Dissociated cells are electroporated with TAL constructs with or without donor DNA using the Neon Transfection System, and then plated at a low density onto Geltrex- or rhLaminin-521– coated plates for recovery. After 1–2 weeks, colonies are picked, screened, and selected based on results obtained with the GeneArt Genomic Cleavage Detection Kit, Applied Biosystems TaqMan SNP Genotyping Assay, or Ion PGM Sequencer. The final clones are expanded and characterized to confirm pluripotency and genomic integrity prior to banking.
 

TAL effector editing of a disease-relevant target

In a recent proof-of-concept study, Invitrogen GeneArt TALs were used to correct a LRRK2 G2019S mutation in iPSCs from a Parkinson’s disease patient. A review of this study is described below. Leucine-rich repeat kinase 2 (LRRK2) is a large multi-domain protein that contains protein–protein interaction domains flanking a catalytic core that harbors a GTPase and a kinase domain. Although the exact role of the LRRK2 gene in Parkinson’s disease is unknown, several mutations in LRRK2 have been linked to the disease, with G2019S being the most common one. Correcting the LRRK2 G2019S mutation back to wild type required editing via homologous recombination, which involved changing one nucleotide from an A back to the wild type G using a GeneArt TAL pair flanking the region along with a 1 kb donor DNA containing the desired correction (Figure 4.16A). In the initial screen for the LRRK2 correction, 2 out of 140 colonies (1.4%) were positive for editing in the assay using the Invitrogen GeneArt Genomic Cleavage Detection Kit(Figure 4.16B, colonies 26 and 27). The colonies were subcloned and rescreened with the TaqMan SNP Genotyping Assay(Figure 4.16C). Ion PGM sequencing was performed on positive colonies to confirm clonality of the population (Figure 4.16D).

To read the full study, go to thermofisher.com/diseasemodels

Figure 4.16. Generation of Parkinson’s disease donor iPSCs with LRRK2 G2019S corrected to wild type. (A) Sequence of LRRK2 G2019S region in the Parkinson’s disease line. The binding sites for the TAL pair are underlined in red. The TALs were electroporated into the cells along with a 1 kb purified PCR fragment containing the wild type sequence and 500 bp flanking sequences. (B) Colony screening by GeneArt Genomic Cleavage Detection Kit. Out of the 140 colonies screened, colonies 26 and 27 showed negligible cleavage product due to mismatch, indicating that the heterozygous mutation was mainly corrected to homozygous wild type. (C) A TaqMan SNP Genotyping Assay confirmed that clones 26 and 27, and their daughter colonies contain homozygous wild type allele, as these clones (red) were plotted in the same region as the wild type controls (from wild type donor plasmid or wild type template from HEK 293 cells) on the allelic discrimination plot. (D) Ion PGM sequencing showed progress from the heterozygous state of the parental line (53% G), to a predominantly edited form in colony 26 (93% G), and finally to a homozygous edited state (100% G) in each of three daughter colonies. 

When using genome editing tools such as CRISPRs or TAL effectors to obtain targeted mutations, it is recommended that you determine the efficiency with which these nucleases cleave the target sequence prior to continuing with labor-intensive and expensive clonal expansion steps. After gene editing, single-cell clones can be easily derived using single-cell sorting. Relying on StemFlex Medium, rhLaminin-521 substrate, and RevitaCell Supplement, single cells can be deposited and expanded in 96-well plates and expanded for ~2 weeks. Formed clones can then be consolidated and expanded for banking and screening for the occurrence of the genome editing event (Figure 4.17). This approach does not require a second round of clonal isolation and is therefore superior to low-density plating and manual picking.

Figure 4.17. Colony screening workflow. After genome editing of singularized hiPSCs, clonal lines can be generated through a FACS-based approach. Single viable pluripotent stem cells are deposited into 96-well plates and allowed to grow for 10–14 days in StemFlex Medium on rhLaminin-521 substrate. At that point, consolidated 96-well plates can be cryopreserved and screened for the genome edit with the indicated assays. 

A variety of tools and reagents, including the TaqMan SNP Genotyping Assay, Invitrogen GeneArt Genomic Cleavage Selection Kit, GeneArt Genomic Cleavage Detection Kit, and Ion PGM sequencing, can be used to quickly determine which cells have been successfully edited. A comparison of these technologies is presented in Table 4.4.

Table 4.4. Comparison of common genomic analysis methodologies.

Screening with the GeneArt Genomic Cleavage Selection Kit

The GeneArt Genomic Cleavage Selection Kit is a rapid and reliable tool for detecting functionality of engineered nucleases in transfected cells as well as enriching for modified cells (Figure 4.18). When using engineered nucleases to create double-stranded breaks in genomic DNA, it is necessary to know whether or not the designed nucleases are functional. Furthermore, to efficiently screen for modified cells, a way to enrich for the edited cells is also necessary, particularly if the engineered nuclease has low efficiency or the cell line used is difficult to transfect. The GeneArt Genomic Cleavage Selection Kit contains a vector with the Orange Fluorescent Protein (OFP) gene for a quick visual check of the functionality of the engineered nuclease. In addition, the reporter genes OFP and CD4 can be used to enrich for edited cells. It can be used in conjunction with genome editing tools such as ZFNs, TAL effector nucleases, and CRISPRs.

Screening with the GeneArt Genomic Cleavage Detection Kit

The GeneArt Genomic Cleavage Detection Kit provides a relatively quick, simple, and reliable assay that allows the assessment of the cleavage efficiency of genome editing tools at a given locus (Figure 4.19). A sample of the edited cell population is used as a direct PCR template with primers specific to the targeted region. The PCR product is then denatured and reannealed to produce heteroduplex mismatches where double-stranded breaks have occurred, resulting in indel introduction. The mismatches are recognized and cleaved by the detection enzyme. Using gel analysis, this cleavage is both easily detectable and quantifiable.

Find out more at thermofisher.com/genedetect

Figure 4.19. GeneArt Genomic Cleavage Detection Assay. To detect either an indel or a mutation within a specific sequence of DNA, the region is first amplified using primers specific for that region. A second nested PCR can be performed to increase sensitivity. After heating the sample and reannealing the PCR products, amplicons containing indels or other changes in sequence will result in the formation of heteroduplexes with amplicons containing unmodified sequences. When these heteroduplexes are treated with an endonuclease that only cleaves in the presence of a mismatch, two pieces of DNA of known size are generated, which can be detected by agarose gel electrophoresis.