Different types of genome-editing nucleases: advantages and disadvantages
Gene editing is a type of genetic engineering that allows the introduction of permanent and locus-specific DNA modifications in the genome. Four types of gene-editing nucleases have been used so far in research: meganucleases (MNs), zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs) and clustered regularly interspaced short palindromic repeats (CRISPR)-associated endonucleases (Cas) [
5,
6].
To guide the nuclease to the target site, MNs, ZFNs, and TALENs use a protein-DNA interaction while CRISPR-Cas systems are guided by RNA–DNA interactions. MNs are highly specific endonucleases, recognizing target sequences of about 14–40 base pairs (bp) [
7]. A drawback in using MNs is the limited number of target sites that they recognize and therefore the extreme difficulty to use these endonucleases in clinical settings, where a higher level of flexibility is desirable [
8]. The creation of new MNs is a laborious process requiring complex protein engineering procedures because the DNA binding and cleavage domains are difficult to separate [
7,
9]. Such laborious engineering required for MNs has constrained their widescale use [
10].
The main characteristics of genome-editing nucleases are listed in Table
2. Both ZFNs and TALENs utilize the nuclease FokI as a cleavage domain. Each ZFN module is composed of about 18–36 amino acids and recognizes a specific 3 bp sequence [
11,
12]. Therefore, several zinc finger modules need to be engineered to recognize the target sequence: each module will bind 3 bp in the target sequence and the FokI nuclease will be coupled to the DNA-binding modules [
13‐
15]. TALENs follow the same principle, but the DNA-binding module recognizes one single nucleotide, instead of 3 [
16]. TALENs modules are composed of 30–40 amino acids, resulting in a protein with higher specificity but larger than ZFNs [
17]. Additionally, for both ZFNs and TALENs, it is necessary to engineer two different enzymes for each target, one upstream and one downstream of the cut site. This is necessary because FokI dimerization is required for completion of the double-stranded break. Overall, the engineering of ZFNs and TALENs are technically challenging, and time consuming compared to the engineering of CRISPR-Cas nucleases [
8,
18]. Furthermore, the target sequence requirements for ZFNs render the selection of an appropriate and specific target difficult [
6,
19,
20].
Table 2
Main features of the different genome-editing nucleases
Phylogenetic origin | artificial restriction enzyme [ 252, 253] | Xanthomonas bacteria [ 254] | Streptococcus pyogenes [ 255] |
DNA binding domain | zinc finger protein [ 253, 256] | | |
DNA cleavage | | | |
DNA recognition range | 18–36 bp (3 bp per module) [ 253] | | 22 bp (DNA-RNA base pairing) [ 261] |
DNA cut | | | dsDNA complex protein-gRNA [ 259] |
Recognition sequence | | sequence with 5'-T and A-3' [ 16, 254, 263] | sequence immediately followed by 5'-NGG-3' [ 259, 266] |
Advantages | Small protein size (< 1 Kb), sequence-based module engineering [ 267] | High specificity, easy selection of target region [ 268] | Easy to multiplex, simple synthesis of gRNA, easy selection of target region [ 269] |
Disadvantages | Difficult sequence selection and protein engineering,, expensive and time consuming [ 267] | Large protein size (> 3 Kb), expensive and time-consuming [ 269, 270] | Large protein size (> 4 Kb) [ 269] |
Safety concerns | off-target effects: genome mutagenesis and GCRs [ 271] | off-target effects: genome mutagenesis and GCRs [ 270] | off-target effects: genome mutagenesis and GCRs [ 271] |
CRISPR-Cas technology
CRISPR-Cas nucleases have crucial advantages compared with ZFNs and TALENs, including the simplicity of the guide RNA (gRNA) design (Table
2). Such nucleases use a 22 bp gRNA to bind a complementary target sequence, which is subsequently cut by the Cas itself [
8,
21,
22]. By designing specific gRNAs, CRISPR-Cas systems could theoretically target any sequence in the genome. Indeed, the ease of target selection and the possibility of multiplexing the gRNAs while maintaining high specificity and efficiency led to the rapid development of CRISPR-Cas methods for clinical purposes [
18]. CRISPR-Cas systems allow to rapidly screen a large number of gRNAs and the scalability of this platform permits an accurate optimization of the study system [
23‐
25].
Stable integration of CRISPR-Cas is not necessary to provide a therapeutic effect and long-term expression is usually considered a disadvantage, as it can lead to off-target cleavage. However, the persistence of CRISPR-Cas in the cell must be sufficient to perform the editing function [
26].
The delivery of a native Cas protein in complex with a gRNA bypasses the requirement for transcription and translation. It introduces genome editing approximately 3 h after delivery and is degraded after 24–48 h [
26]. Circumventing transcription and translation is useful in post-mitotic or hard-to-transfect cells. Such transient functionality allows for rapid editing and reduced off-target effects. However, obtaining pure active protein is a difficult process, and the risk of endotoxin contamination remains of concern [
27]. The delivery of Cas proteins offers an improved dose-control compared to mRNA and DNA but, in order to produce a therapeutic effect, a significant amount of protein must be successfully delivered. This is due to the lack of amplification signal which normally occurs with mRNA and DNA. Moreover, the CRISPR-Cas protein is large, which may present a challenge for intracellular delivery.
Another strategy to minimize off-target editing events is through the delivery of Cas mRNA. This process results in rapid genome editing (5–7 h after transfection) and avoids the step of nuclear entry [
28]. The mRNA that codes for the Cas protein is immediately translated in the cytosol and the complex Cas-gRNA subsequently enters the nucleus. The transient expression of Cas proteins reduces off-target effects and risk of integration, but the dose and timing of mRNA delivery have to be carefully titrated [
29,
30].
Finally, the delivery of genome editing nucleases through plasmids is an easy procedure. Due to the necessity of nuclear entry, subsequent transcription, and translation into protein, the genome editing efficacy is significantly delayed. Plasmids are very stable molecules, therefore Cas protein expression may last several days, leading to a high risk of off-target effects and safety concerns [
31,
32]. Plasmid delivery may also trigger cytosolic DNA toxicity [
33]. Despite the fact that each of the above-mentioned nucleases present specific advantages and disadvantages, CRISPR-Cas technology has been widely adopted and improved in the last few years and it remains a promising route for preclinical and clinical investigations [
8]. Depending on the cell type and experimental conditions, the knockout efficiency for CRISPR-Cas9 varies between 40% (induced pluripotent stem cells, iPSCs) and 99.4% (cortex, hippocampus and spinal cord) [
34‐
37].
CRISPR-Cas9 can also be used for gene or targeted nucleotide knock-in experiments. Such manipulations are usually more challenging to perform and require accurate optimization, including the addition of an extra component in the form of a DNA donor template [
38]. Different studies reported a wide range of knock-in efficiencies depending on the method and the cell type used. Liu et al. [
39] compared the efficiencies of CRISPR-Cas9 versus ZFN and TALEN, performing knock-ins in fetal fibroblasts [
39]. They found that CRISPR-Cas9-mediated gene knock-in (70–80% efficiency) was 5.6 times more efficient than ZFN and around 3 times more efficient than TALEN. Nevertheless, other studies found lower knock-in efficiencies for CRISPR-Cas9, for example ∼20% in human primary T cells [
40]. In conclusion, the overall efficiency of knock-in seems to remain lower than the knockout efficiency, using CRISPR-Cas9. Moreover, delivery of CRISPR-Cas9 through viral vectors requires in vitro T cell activation and culture [
41,
42]. However, the use of electroporation (EP) methods to deliver Cas9-gRNA protein complex in knockout studies demonstrated the potential to overcome this issue and to achieve gene editing without in vitro T cell activation [
42,
43]. Human primary T cells are difficult to manipulate and chemically modified gRNAs were also tested to enhance genome editing efficiency [
44].
In the last decade many tools have been developed and optimized to investigate genome functional complexity based on Cas proteins. Among those, a Cas-based tool for epigenome editing (non-gene editing) called “nuclease dead Cas” (dCas) was developed by creating a mutant form of Cas which lacks endonuclease activity. This enzyme still retains the capability to bind the gRNA and it can target Cas-coupled effector proteins to a specific locus of the genome [
45]. Coupling dCas with VPR activator (CRISPRa) or KRAB repressor (CRISPRi) of transcription creates a powerful tool for precise epigenetic editing. For example, Schmidt et al. [
46] developed a CRISPRa and CRISPRi platform to perform genome-wide screens for functional regulators of cytokine production in response to T cell stimulation [
46]. Yang et al. [
47] developed a CAR-T cell product called RB-340–1, which was engineered through a CRISPRi circuit to prevent Programmed cell Death protein 1 (PD-1) expression upon antigen-encounter [
47]. RB-340–1 is the first application of CRISPRi toward a clinically relevant product and allows the conditional and reversible suppression of PD-1. The reversible nature of this editing also allows fine tuning of the degree of PD-1 expression. RB-340–1 demonstrated resilience to checkpoint inhibition and increased persistence and effectiveness against HER2-expressing cancer xenografts [
47].
Safety of genome editing-based techniques
A drawback that should be considered when performing genome editing is the immunogenicity of the nucleases – ZFNs, TALENs and CRISPR-Cas are exogenous proteins and may trigger an immune response in the patient. Regarding SpCas9, in vivo delivery has been found to elicit both antibody and T cell responses in immunocompetent mice [
48‐
51]. Cell therapy which employs products transiently edited ex vivo through plasmids, mRNA or protein, is expected to be safe as the Cas9 is diluted during cell proliferation [
52]. Early reports from clinical trials revealed persistence of T cells edited ex vivo through SpCas9, in cancer patients [
53‐
55]. However, in all these reports Cas9-directed immune responses were not directly evaluated [
56] and the available safety data derived from patients that had a compromised immune system. Despite the encouraging results, thorough investigations with ex vivo engineered T cell products may be needed to assess humoral and cellular immune response, after infusion.
Off-target genotoxicity together with the risk of creating translocations when multiplex genome editing is performed are a major drawback of genome editing nucleases [
57]. Indeed, chromosomal translocations are natural byproducts of inducing simultaneous genomic breaks [
58,
59]. Different nuclease combinations or the presence of a homologous single-stranded donor have been suggested as approaches to reduce chromosomal translocations in multiplex editing [
58]. For example, Bothmer et al. [
58], performed knockout at the TRAC and B2M loci in human T cells, including a single-stranded repair template in the reaction. The repair template presented 70 bp of homology on either side of the double-strand break (DSB) with a 10 bp stop cassette, to achieve functional knockout. With this strategy, the DSB repair mechanism was shifted from Non-Homologous End Joining (NHEJ) which can cause translocations, to single-stranded template repair (SSTR) [
58].
Clinical trials
Over the past few years, genome editing nucleases made their appearance in clinical trials although so far, no U.S. Food and Drug Administration (FDA) approved treatment based on this technology has been commercialized and no late-stage clinical trial has been approved.
In gene therapy, ZFN and CRISPR-Cas9 are currently being investigated in clinical trials to treat genetic diseases such as Mucopolysaccharidosis (NCT03041324, NCT04628871, NCT02702115), Hemophilia B (NCT02695160), β-Thalassemia (NCT03432364, NCT03728322, NCT03655678), Neurofibromatosis type 1 (NCT03332030), Sickle cell disease (NCT03745287) and LCA10 (NCT03872479). All these clinical trials are at an early stage: phase I or I/II and no clinical trials have been approved for TALEN so far.
The situation is similar for cell therapy: early-stage clinical trials are currently evaluating products to treat both hematological malignancies and solid tumors. Genome editing nucleases are employed to knock out target genes such as IL13Ralpha2, PD-1, CISH, TRAC and B2M. For example, TALEN is currently being tested in a clinical trial to knockout TRAC and CD52 in allogeneic CAR-T cells (NCT02808442). This is to create an off the shelf CAR product for a specific patient population [
60]. ZFN has been used to permanently disrupt the glucocorticoid receptor GRm13Z40-2 in anti-IL13Ralpha2 allogeneic CD8 + T cells, used to treat patients with recurrent/refractory malignant glioma (NCT01082926). Products engineered through CRISPR-Cas9 technology have been more frequently adopted in clinical trials. For solid tumors, PDC1 and TRAC knockouts have been tested (NCT02793856, NCT03081715, NCT03044743, NCT03545815, NCT03747965) aiming to decrease CAR-T cells exhaustion. For hematological malignancies CRISPR-Cas9 target genes include TRAC, B2M, CD7, CD28, CD19, CD20 and CD22 (NCT03190278, NCT03166878, NCT03398967, NCT03690011), depending on the cancer characteristics.