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Principles of Genetic Engineering

Thomas m. lanigan.

1 Biomedical Research Core Facilities, Vector Core, University of Michigan, Ann Arbor, MI 48109, USA; ude.hcimu@tnaginal (T.M.L.); ude.hcimu@hgnohc (H.C.K.)

2 Department of Internal Medicine, Division of Rheumatology, University of Michigan, Ann Arbor, MI 48109, USA

Huira C. Kopera

3 Department of Human Genetics, University of Michigan, Ann Arbor, MI 48109, USA

Thomas L. Saunders

4 Biomedical Research Core Facilities, Transgenic Animal Model Core, University of Michigan, Ann Arbor, MI 48109, USA

5 Department of Internal Medicine, Division of Genetic Medicine, University of Michigan, Ann Arbor, MI 48109, USA

Genetic engineering is the use of molecular biology technology to modify DNA sequence(s) in genomes, using a variety of approaches. For example, homologous recombination can be used to target specific sequences in mouse embryonic stem (ES) cell genomes or other cultured cells, but it is cumbersome, poorly efficient, and relies on drug positive/negative selection in cell culture for success. Other routinely applied methods include random integration of DNA after direct transfection (microinjection), transposon-mediated DNA insertion, or DNA insertion mediated by viral vectors for the production of transgenic mice and rats. Random integration of DNA occurs more frequently than homologous recombination, but has numerous drawbacks, despite its efficiency. The most elegant and effective method is technology based on guided endonucleases, because these can target specific DNA sequences. Since the advent of clustered regularly interspaced short palindromic repeats or CRISPR/Cas9 technology, endonuclease-mediated gene targeting has become the most widely applied method to engineer genomes, supplanting the use of zinc finger nucleases, transcription activator-like effector nucleases, and meganucleases. Future improvements in CRISPR/Cas9 gene editing may be achieved by increasing the efficiency of homology-directed repair. Here, we describe principles of genetic engineering and detail: (1) how common elements of current technologies include the need for a chromosome break to occur, (2) the use of specific and sensitive genotyping assays to detect altered genomes, and (3) delivery modalities that impact characterization of gene modifications. In summary, while some principles of genetic engineering remain steadfast, others change as technologies are ever-evolving and continue to revolutionize research in many fields.

1. Introduction

Since the identification of DNA as the unit of heredity and the basis for the central dogma of molecular biology [ 1 ] that DNA makes RNA and RNA makes proteins, scientists have pursued experiments and methods to understand how DNA controls heredity. With the discovery of molecular biology tools such as restriction enzymes, DNA sequencing, and DNA cloning, scientists quickly turned to experiments to change chromosomal DNA in cells and animals. In that regard, initial experiments that involved the co-incubation of viral DNA with cultured cell lines progressed to the use of selectable markers in plasmids. Delivery methods for random DNA integration have progressed from transfection by physical co-incubation of DNA with cultured cells, to electroporation and microinjection of cultured cells [ 2 , 3 , 4 ]. Moreover, the use of viruses to deliver DNA to cultured cells has progressed in tandem with physical methods of supplying DNA to cells [ 5 , 6 , 7 ]. Homologous recombination in animal cells [ 8 ] was rapidly exploited by the mouse genetics research community for the production of gene-modified mouse ES cells, and thus gene-modified whole animals [ 9 , 10 ].

This impetus to understand gene function in intact animals was ultimately manifested in the international knockout mouse project, the purpose of which was to knock out every gene in the mouse genome, such that researchers could choose to make knockout mouse models from a library of gene-targeted knockout ES cells [ 11 , 12 , 13 ]. Thousands of mouse models have resulted from that effort and have been used to better understand gene function and the bases of human genetic diseases [ 14 ]. This project required high-throughput pipelines for the construction of vectors, including bacterial artificial chromosome (BAC) recombineering technology [ 13 , 15 , 16 , 17 ]. BACs contain long segments of cloned genomic DNA. For example, the C57BL/6J mouse BAC library, RPCI-23, has an average insert size of 197 kb of genomic DNA per clone [ 18 ]. Because of their size, BACs often carry all of the genetic regulatory elements to faithfully recapitulate the expression of genes contained in them, and thus can be used to generate BAC transgenic mice [ 19 , 20 ]. Recombineering can be used to insert reporters in BACs that are then used to generate transgenic mice to accurately label cells and tissues according to the genes in the BACs [ 21 , 22 , 23 , 24 , 25 , 26 ]. A panoply of approaches to genetic engineering are available for researchers to manipulate the genome. ES cell and BAC transgene engineering have given way to directly editing genes in zygotes, consequently avoiding the need for ES cell or BAC intermediates on the way to an animal model.

Prior to the adaptation of Streptococcus pyogenes Cas9 protein to cause chromosome breaks, three other endonuclease systems were used: (1) rare-cutting meganucleases, (2) zinc finger nucleases (ZFNs), and (3) transcription activator-like effector (TALE) nucleases (TALENs) [ 27 ]. The I-CreI meganuclease recognizes a 22 bp DNA sequence [ 28 , 29 ]. Proof-of-concept experiments demonstrated that the engineered homing endonuclease I-CreI can be used to generate transgenic mice and transgenic rats [ 30 ]. I-CreI specificity can be adjusted to target specific sequences in DNA by protein engineering methodology, although this limits its widespread application to genetic engineering [ 31 ]. Subsequently, ZFN technology was developed to cause chromosome breaks [ 32 ]. A single zinc finger is made up of 30 amino acids that bind three base pairs. Thus, three zinc fingers can be combined to specifically recognize nine base pairs on one DNA strand and a triplet of zinc fingers is made to bind nine base pairs on the opposite strand. Each zinc finger is fused to the DNA-cutting domain of the FokI restriction endonuclease. Because FokI domains only cut DNA when they are present as dimers, a ZFN monomer binding to a chromosome cannot induce a DNA break [ 32 ], instead requiring ZFN heterodimers for sequence-specific chromosome breaks. It is estimated that 1 in every 500 genomic base pairs can be cleaved by ZNFs [ 33 ]. Compared with meganucleases, ZFNs are easier to construct because of publicly available resources [ 34 ]. Additionally, the value of ZFNs in mouse and rat genome engineering was demonstrated in several studies that produced knockout, knockin, and floxed (described below) animal models [ 35 , 36 , 37 ]. The development of transcription activator-like effector nucleases (TALENs) followed after ZFN technology [ 38 ]. TALENs are made up of tandem repeats of 34 amino acids. The central amino acids at positions 12 and 13, named repeat variable di-residues (NVDs), determine the base to which the repeat will bind [ 38 ]. To achieve a specific chromosomal break, 15 TALE repeats assembled and fused to the FokI endonuclease domain (TALEN monomer) are required. Thus, one TALEN monomer binds to 15 base pairs on one DNA strand, and a second TALEN monomer binds to bases on the opposite strand [ 38 ]. When the FokI endonuclease domains are brought together, a double-stranded DNA break occurs. In this way, a TALEN heterodimer can be used to cause a sequence-specific chromosome break. It has been estimated that, within the entire genome, TALENs have potential target cleavage sites every 35 bp [ 39 ]. Compared with ZFNs, TALENs are easier to construct with publicly available resources [ 40 , 41 ], and TALENs have been adopted for use in mouse and rat genome engineering in several laboratories that have produced knockout and knockin animal models [ 42 , 43 , 44 , 45 , 46 ].

The efficiencies of producing specific double-strand chromosome breaks, using prior technologies such as meganucleases, ZFNs, and TALENs [ 28 , 32 , 38 ], were surpassed when CRISPR/Cas9 technology was shown to be effective in mammalian cells [ 47 , 48 , 49 ]. The essential feature that all of these technologies have in common is the production of a chromosome break at a specific location to facilitate genetic modifications [ 50 ]. In particular, the discovery of bacterial CRISPR-mediated adaptive immunity, and its application to genetic modification of human and mouse cells in 2013 [ 47 , 48 , 49 ], was a watershed event to modern science. Moreover, the introduction of CRISPR/Cas9 methodology has revolutionized transgenic mouse generation. This paradigm shift can be seen by changes in demand for nucleic acid microinjections into zygotes, and ES cell microinjections into blastocysts at the University of Michigan Transgenic Core ( Figure 1 ). While previously established principles of genetic engineering using mouse ES cell technology [ 51 , 52 , 53 ] remain applicable, CRISPR/Cas9 methodologies have made it much easier to produce genetically engineered model organisms in mice, rats, and other species [ 54 , 55 ]. Herein, we discuss principles in genetic engineering for the design and characterization of targeted alleles in mouse and rat zygotes, or in cultured cell lines, for the production of animal and cell culture models for biomedical research.

Recent trends in nucleic acid microinjection in zygotes, and embryonic stem (ES) cell microinjections into blastocysts, for the production of genetically engineered mice at the University of Michigan Transgenic Core. As shown, prior to the introduction of CRISPR/Cas9, the majority of injections were of ES cells, to produce gene-targeted mice, and DNA transgenes, to produce transgenic mice. After CRISPR/Cas9 became available, adoption was slow until 2014, when it was enthusiastically embraced, and the new technology corresponded to a reduced demand for ES cell and DNA microinjections.

2. Principles of Genetic Engineering

2.1. types of genetic modifications.

There are many types of genetic modifications that can be made to the genome. The ability to specifically target locations in the genome has expanded our ability to make changes that include knockouts (DNA sequence deletions), knockins (DNA sequence insertions), and replacements (replacement of DNA sequences with exogenous sequences). Deletions in the genome can be used to knockout gene expression [ 56 , 57 ]. Short deletions in the genome can be used to remove regulatory elements that knockout gene expression [ 58 ], activate gene expression [ 59 ], or change protein structure/function by changing coding sequences [ 60 ].

Insertion of new genomic information can be used to knock in a variety of genetic elements. Knockins are also powerful approaches for modifying genes. Just as genomic deletions can be used to change gene function, knockins can be used to block gene function by inserting fluorescent reporter genes such as eGFP or mCherry, in such a way as to knock out the gene at the insertion point [ 61 , 62 ]. It is also possible to knock in fluorescent protein reporter genes, without knocking out the targeted gene [ 63 , 64 ]. Just as fluorescent proteins can be used to label proteins and cells, short knockins of epitope tags in proteins can be used to label proteins for detection with antibodies [ 64 , 65 ].

Replacement of DNA sequences in the genome can be used to achieve two purposes at the same time, such as blocking gene function, while activating the function of a new gene such as the lacZ reporter [ 66 ]. Large-scale sequence replacements are possible with mouse ES cell technology, such as the replacement of the mouse immunoglobulin locus with the human immunoglobulin locus to produce a “humanized” mouse [ 67 ]. Furthermore, very small replacements of single nucleotides can be used to model point mutations that are suspected of causing human disease [ 68 , 69 , 70 ].

A special type of DNA sequence replacement is the conditional allele. Conditional alleles permit normal gene expression until the site-specific Cre recombinase removes a loxP-flanked critical exon to produce a “floxed” (flanked by loxP) exon. Cre recombinase recognizes 34 bp loxP (locus of recombination) elements, and catalyzes recombination between the two loxP sites [ 71 , 72 ]. Therefore, deletion of the critical exon causes a premature termination codon to occur in the mRNA transcript, triggering its nonsense-mediated decay and failure to make a protein [ 13 , 73 ]. Engineering conditional alleles was the approach used by the international knockout mouse project [ 13 ]. Mice with cell- and tissue-specific Cre recombinase expression are an important resource for the research community [ 74 ].

Other site-specific recombinases, such as FLP, Dre, and Vika, that work on the same principle have also been applied to mouse models [ 75 , 76 , 77 , 78 , 79 , 80 ]. Recombinase knockins can be designed to knock out the endogenous gene or preserve its function [ 81 , 82 ]. A variation in the conditional allele is the inducible allele, which is silent until its expression is activated by Cre recombinase [ 79 ]. For example, reporter models can activate the expression of a fluorescent protein [ 83 ], change fluorescent reporter protein colors from red to green [ 84 ], or use a combinatorial approach to produce up to 90 fluorescent colors [ 85 ]. Another type of inducible allele is the FLEX allele. FLEX genes are Cre-dependent gene switches based on the use of heterotypic loxP sites [ 86 ]. In one application that combined Cre and FLP recombinases, it was demonstrated that a gene inactivated in ES cells by a gene trap could be switched back on and then switched off again [ 87 ]. In another application of heterotypic loxP sites in mouse ES cells, it was demonstrated that genes could be made conditional by inversion (COIN) [ 88 ]. This application has been used to produce mice with conditional genes for point mutations [ 89 ] and has been applied to produce conditional single exon genes that lack critical exons by definition [ 90 ].

2.2. Genetic Engineering with CRISPR/Cas9

The central principle of gene targeting with CRISPR/Cas9, or other directed DNA endonucleases, is that a double-strand DNA break is generated in the cell of interest. Following a chromosomal break, the principal outcomes of interest are nonhomologous end joining (NHEJ) repair [ 91 ] or homology-directed repair (HDR) [ 92 ]. When the break is directed to a coding exon in a gene, the outcome of NHEJ is usually a small insertion or deletion of DNA sequence at the break (indel), causing frame shifts in mRNA transcripts that lead to premature termination codons, causing nonsense-mediated mRNA decay and loss of protein expression [ 73 ]. The HDR pathway copies a template during DNA repair, and thus the insertion of modified genetic sequences in the form of a DNA donor. This DNA donor can introduce new information into the genome flanked by homology arms on either side of the chromosome break. Typical applications of HDR include the use of genetic engineering to abrogate gene expression (gene knockouts), to modify amino acid codons (i.e.; point mutations), to replace genes with new genes (e.g.; knockins of fluorescent reporters, Cre recombinase, cDNA coding sequences), to produce conditional genes (floxed genes that are normally expressed until they are inactivated by Cre recombinase), to produce Cre-inducible genes (genes that are only expressed after Cre recombinase activates them), and to delete DNA from chromosomes (e.g.; delete regulatory elements that control gene expression, delete entire genes, or delete up to a megabase of chromosome segments). The simplest of these modifications is abrogation of gene expression. Multifunctional alleles, such as FLEX alleles, require the cloning or synthesis of multi-element plasmid DNA donors for HDR.

The processes of CRISPR/Cas9-mediated modifications of genes (gene editing) to produce a new cell line or animal model have in common a series of steps to achieve the final product. First, a gene of interest is identified and the final desired allele is specified. The next step is to identify single guide RNA(s) (gRNAs) that will be used to target a chromosomal break in one or more places. There are numerous online websites that can be used for this purpose [ 93 ]. One of the most up-to-date and versatile sites is CRISPOR ( http://crispor.tefor.net ) [ 94 ]. Interestingly, the authors provide evidence that the predictive powers of algorithms vary depending on whether they were based on the analysis of gRNAs delivered as RNA molecules, versus gRNAs delivered as U6-transcribed DNA molecules [ 94 ]. In any event, the selection of a gRNA target (20 nucleotides), adjacent to a protospacer-adjacent motif (PAM; NGG motif), should not be done without the aid of a computer algorithm that minimizes the possibility of off-target hits. After a gRNA target is identified, a decision is made to obtain gRNAs. While it is possible to produce in vitro-transcribed gRNAs, this may be inadvisable in so much as in vitro-transcribed RNAs can trigger innate immune responses and cause cytotoxicity in cells [ 95 ]. Chemically synthesized gRNAs using phosphorothioate modifications that improve gRNA stability may be preferable alternatives to in vitro-transcribed molecules [ 96 , 97 ]. With a gRNA in hand, a Cas9 protein is then selected. There are numerous forms of Cas9 that can be used for different purposes [ 98 ]. For practical purposes, we limit our discussion to Cas9 varieties that are on the market. A number of commercial entities sell wild-type Cas9 protein. When wild type Cas9 is used to target the genome with nonspecific guides, the frequency of off-target genomic hits, besides the desired Cas9 target, is very likely to increase [ 94 , 99 ]. Alternatives to the wild-type protein include enhanced specificity Cas9 from Sigma-Aldrich [ 100 ], and high-fidelity Cas9 from Integrated DNA Technologies [ 101 ]. In addition, there are other versions such as HF1 Cas9 [ 102 ], hyperaccurate Cas9 [ 103 ], and evolved Cas9 [ 104 ], all available in plasmid format from Addgene.org. As may be inferred from the names of these engineered Cas9 versions, they are designed to be more specific than wild type Cas9. Once the gRNAs and Cas9 protein are on hand, then it is a “simple” matter to combine them and deliver them to the target cell to produce a chromosome break and achieve a gene knockout by introducing premature termination codons or DNA sequence deletion of regulatory regions or entire genes.

2.3. Locus-Specific Genetic Engineering Vectors in Mouse and Rat Zygotes

The most challenging type of genetic engineering is the insertion (i.e.; knockin) of a long coding sequence to express a fluorescent reporter protein, Cre recombinase, or conditional allele (floxed gene). In addition to these genetic modifications, numerous other types of specialized reporters can be introduced, each designed to achieve a different purpose. There is great interest in achieving rapid and efficient gene insertions of reporters in animal models with CRISPR/Cas9 technology. It is generally recognized that, the longer the insertion, the less efficient it is to produce a knockin animal. Additional challenges are allele-specific differences that affect efficiency. For example, it is fairly efficient to produce knockins into the genomic ROSA26 locus in mice, while other loci are targeted less efficiently, and thus refractory to knockins. This accessibility to CRISPR/Cas9 complexes mirrors observations in mouse ES cell gene targeting technology, in which it was reported that some genes are not as efficiently targeted as others [ 105 ].

When the purpose of the experiment is to specifically modify the DNA sequence by changing amino acid codons, or introducing new genetic information, then a DNA donor must be delivered to the cells with Cas9 reagents. After the selected gRNAs and Cas9 proteins are demonstrated to produce the desired chromosome break, the DNA donor is designed and procured. The donor should be designed to insert into the genome such that it will not be cleaved by Cas9, usually by mutating the PAM site. The DNA donor may take the form of short oligonucleotides (<200 nt) [ 106 , 107 ], long single-stranded DNA molecules (>200 nt) [ 108 ], or double-stranded linear or circular DNA molecules of varying lengths [ 109 , 110 ].

DNA donor design principles should include the following: (1) nucleotide changes that prevent CRISPR/Cas9 cleavage of the chromosome, after introduction of the DNA donor; (2) insertion of restriction enzyme sites unique to the donor, to simplify downstream genotyping; (3) insertions of reporters or coding sequences, at least 1.5 kb in length, that can be introduced as long single-stranded DNA templates with short 100 base pair arms of homology [ 111 ], or as circular double-stranded DNA plasmids with longer (1.5 or 2 kb) arms of homology [ 63 , 110 ]; and (4) insertions of longer coding sequences, such as Cas9, that use circular double-stranded DNA donors with longer arms of homology [ 63 , 112 ]. It is also possible to use linear DNA fragments as donors [ 63 , 110 , 113 ], although random integration of linear DNA molecules is much higher than those of circular donors, thus requiring careful quality control.

The establishment of genetically modified mouse and rat models can be divided into three phases, after potential founder animals are born from CRISPR/Cas9-treated zygotes. In the first phase, animals with genetic modifications are identified. The first phase requires a sensitive and specific genotyping assay to identify cells or animals harboring the desired knockin. Genotyping potential founder mice for knockins typically begins with a PCR assay using a primer that recognizes the exogenous DNA sequence and a primer in genomic DNA outside of the homology arm in the targeting vector. Accordingly, PCR assays are designed to specifically detect the upstream and downstream junctions of the inserted DNA in genomic DNA. Subsequent assays may be used to confirm that the entire exogenous sequence is intact. Conditional genes represent a special case of insertion, as PCR assays designed to detect correct insertion of loxP-flanked exons will also detect genomic DNA [ 108 ]. In the second phase, founders are mated and G1 pups are identified that inherited the desired mutation [ 114 ]. In the third phase, it is essential to sequence additional genomic regions upstream and downstream of the inserted targeting vector DNA, because Cas9 is very efficient at inducing chromosomal breaks, but has no repair function. Thus, it is not unusual to identify deletions/insertions that flank the immediate vicinity of the Cas9 cut site or inserted targeting vector DNA sequences [ 115 , 116 ]. If such deletions affect nearby exons, gene expression can be disrupted, and confounding phenotypes may arise.

For gene knockouts, PCR amplicons from primers that span the chromosome break site are analyzed by DNA sequencing. Any animals that are wild-type at the allele are not further characterized or used, so as to prevent any off-target hits from entering the animal colony or confounding phenotypes. Animals that show disrupted DNA sequences at the Cas9 cut site are mated with wild-type animals for the transmission of mutant alleles that produce premature termination codons, for gene knockout models [ 57 , 73 ]. As founders from Cas9-treated zygotes are genetic mosaics [ 55 , 115 ], it is essential to mate them to wild-type breeding partners, such that obligate heterozygotes are produced. In the heterozygotes, the wild-type sequence and the mutant sequence can be precisely identified by techniques such as TOPO TA cloning (Invitrogen, CA, USA) or next-generation sequencing (NGS) methods [ 117 , 118 , 119 , 120 ]. Animals carrying a defined indel, with the desired properties, are then used to establish lines for phenotyping. The identical approach is used when short DNA sequences are deleted by two guide RNAs [ 58 ]. Intercrossing mosaic founders will produce offspring carrying two different mutations with different effects on gene expression. These animals are not suitable for line establishment.

2.4. Gene Editing in Immortalized Cell Lines

CRISPR/Cas9 gene editing in immortalized cell lines presents a set of challenges unique from those used in the generation of transgenic animals. Cell lines encompass a wide range of characteristics, resulting in each line being handled differently. Some of these characteristics include phenotype heterogeneity, aberrant chromosome ploidy, varying growth rates, DNA damage response efficiency, transfection efficiency, and clonability. While the principles of CRISPR/Cas9 experimental design, as stated above, remain the same, three major considerations must be taken into account when using cell lines: (1) copy number variation, or the number of alleles of the gene of interest; (2) transfection efficiency of the cell line; and (3) clonal isolation of the modified cell line. In cell lines, all alleles need to be modified in the generation of a null phenotype, or in the creation of a homozygous genotype. Unlike transgenic animals, where single allele gene edits can be bred to homozygosity, CRISPR/Cas9-edited cells must be screened for homozygous gene edits. Copy number variations within the cell line can decrease the efficiency and add labor and time (i.e.; editing 3 or 4 copies versus editing 1 or 2). Furthermore, an aberrant number of chromosomes, deletions, duplications, pseudogenes, and repetitive regions complicate genetic backgrounds for PCR analysis of the CRISPR edits. To help with some of these issues, one common approach is to use NGS on all the clonal isolates for a complete understanding of copy number variations for each clonal cell line generated, and the exact sequence for each allele.

As all cell types are not the same, different CRISPR/Cas9 delivery techniques may need to be tested to identify which method works best. One approach is to use viruses or transposons to deliver CRISPR/Cas9 reagents (detailed below). However, the viruses and transposons themselves will integrate into the genome, as well as allowing long-term expression of CRISPR/Cas9 in the cell. This prolonged expression of gRNAs and Cas9 protein may lead to off-target effects. Moreover, transfection and electroporation can have varying efficiencies, depending on the cell lines and the form of CRISPR/Cas9 reagents (e.g.; DNA plasmids or ribonucleoprotein particles (RNPs)).

Following delivery, clonal isolation is required to identify the edited cell line, and at times, can result in the isolation of a cell phenotype different than that expected, arising from events apart from the desired gene edit. While flow cytometry can aid in isolating individual cells, specific flow conditions, such as pressure, may require adjustment to ensure cell viability. Furthermore, one clonal isolate from a cell line may possess a different number of alleles for the targeted gene than another clonal isolate. Additionally, not all cell lines will grow from a single cell, thus complicating isolation. Growth conditions and cell viability can also change when isolating single cells.

Despite these challenges, new advances in CRISPR technology can likely alleviate some of these difficulties when editing cell lines. For example, fluorescently tagged Cas9 and RNAs help to isolate only transfected cells, which helps to eliminate time wasted on screening untransfected cells. Cas9-variants that harbor mutations that only create single-strand nicks (Cas9-nickases) complexed with two different, but proximal gRNAs can increase HDR-mediated knockin [ 48 , 121 ]. Similarly, fusing Cas9 with base-editing enzymes can also increase the efficiency of editing, without causing double-strand breaks [ 121 ].

2.5. Viruses and Transposons as Genetic Engineering Vectors

Viral and transposon vectors have been engineered to be safe, efficient delivery systems of exogenous genetic material into cells. The natural lifecycle of some viruses and transposons includes the stable integration into the host genome. In the field of genome engineering, these vectors can be used to modify the genome in a non-directed fashion, by inserting cassettes expressing any cDNA, shRNA, miRNA, or any non-coding RNA. The most widely used vectors capable of integrating ectopic genetic material into cells are retroviruses, lentiviruses, and adeno-associated virus (AAV). These viruses are flanked by terminal repeats that mark the boundaries of the integration. In engineering these viruses into recombinant vector systems, all the viral genes are removed from the flanking terminal repeats and supplied in trans for the recombinant virus to be packaged. These “gutted”, nonreplicable viral vectors allow for the packaging, delivery, integration, and expression of cDNAs of interest, shRNAs, and CRISPR/Cas9, without viral replication in various biological targets.

Similar to recombinant viruses, transposon vectors are also “gutted”, separating the transposase from the terminal repeat-flanked genetic material to be inserted into the genome. DNA transposons are mobile elements (“jumping genes”) that integrate into the host genome through a cut-and-paste mechanism [ 122 ]. Transposons, much like viral vectors, are flanked by repeats that mark the region to be transposed [ 123 ]. The enzyme transposase binds the flanking DNA repeats and mediates the excision and integration into the genome. Unlike viral vectors, transposons are not packaged into viral particles, but form a DNA-protein complex that stays in the host cell. Thus, the transgene to be integrated can be much larger than the packaging limits of some viruses.

Two transposons, Sleeping Beauty (SB) and piggybac (PB), have been engineered and optimized for high activity for generating transgenic mammalian cell lines [ 124 , 125 , 126 ]. Sleeping Beauty is a transposable element resurrected from fish genomes. The SB system has been used to generate transgenic HeLa cell lines, T-cells expressing chimeric antigen receptors that recognize tumor-specific antigens, and transgenic primary human stem cells [ 127 , 128 , 129 ]. The insect-derived PB system also has been used to generate transgenic cell lines [ 126 , 130 , 131 ]. The PB system was used to generate induced pluripotent stem cells (iPSCs) from mouse embryonic fibroblasts, by linking four or five cDNAs of the reprogramming (Yamanaka) factors [ 132 ] with intervening peptide self-cleavage (P2A) sites, thus delivering all of the factors in one vector [ 130 ]. Furthermore, once reprogrammed, the transgene may be removed by another round of PB transposase activity, leaving no genetic trace of integration or excision (i.e.; transgene-free iPSCs). Following PB transposase activity, epigenetic differences remaining at the endogenous promoters of the reprogramming factor genes result in sustained expression and pluripotency, despite transgene removal.

Aside from transgene insertion, Sleeping Beauty (SB) and piggyback (PB) have both been engineered to deliver CRISPR/Cas9 reagents into cells [ 133 , 134 , 135 ]. Similar to lentivirus, the stable integration of CRISPR/Cas9 by transposons could increase the efficacy of targeting and modifying multiple alleles. SB and PB have been used to deliver multiple gRNAs to target multiple genes (instead of just one), aiding in high-throughput screening. Furthermore, owing to the nature of PB excision stated above, the integrated CRISPR/Cas9 can be removed once a clonal cell line is established, to limit off-target effects. However, engineered transposons must be transfected into cells. As stated above, efficiencies vary between different cell lines and transfection methods. One potential solution to overcome this challenge is to merge technologies. For example, instead of transfecting cells with a plasmid harboring a gRNA flanked by SB terminal repeats (SB-CRISPR), the SB-CRISPR may be flanked by recombinant AAV (rAAV) terminal repeats (AAV-SB-CRISPR), allowing for packaging into rAAV. To that end, rAAV-SB-CRISPR has been used to infect primary murine T-cells, and deliver the SB-CRISPR construct [ 136 ].

2.6. Genetic Engineering Using Retroviruses

Retroviruses are RNA viruses that replicate through a DNA intermediate [ 137 ]. They belong to a large family of viruses including both onco-retroviruses, such as the Moloney murine leukemia virus (MMLV) (simply referred to as retrovirus), and lentiviruses, including human immunodeficiency virus (HIV). In all retroviruses, the RNA genome is flanked on both sides by long terminal repeats (LTRs); packaged with viral reverse transcriptase, integrase, and protease, surrounded by a protein capsid; and then enveloped into a lipid-based particle [ 138 ]. Envelope proteins interact with specific host cell surface receptors to mediate entry into host cells through membrane fusion. Then, the RNA genome is reverse-transcribed by the associated viral reverse transcriptase. The proviral DNA is then transported into the nucleus, along with viral integrase, resulting in integration into the host cell genome [ 139 ]. By contrast, the retroviral MMLV pre-integration complex is incapable of crossing the nuclear membrane, thus requiring the cell to undergo mitosis to gain access to chromatin [ 139 ], while lentiviral pre-integration complexes can cross nuclear membrane pores, allowing genome integration in both dividing and non-dividing cells.

Large-scale assessments of genomic material composition have uncovered features associated with retroviral insertion into mammalian genomes [ 140 ]. Although determination of integration target sites remains ill-defined, it does depend on both cellular and viral factors. For retroviruses such as MMLV, integration is preferentially targeted to promoter and regulatory regions [ 140 , 141 , 142 ]. Such preferences can be genotoxic owing to insertional activation of proto-oncogenes in patients undergoing gene therapy treatments for X-linked severe combined immunodeficiency [ 143 , 144 ], Wiskott–Aldrich syndrome [ 143 ], and chronic granulomatous disease [ 145 ]. Likewise, retroviral integration can generate chimeric and read-through transcripts driven by strong retroviral LTR promoters, post-transcriptional deregulation of endogenous gene expression by introducing retroviral splice sites (leading to aberrant splicing), and retroviral polyadenylation signals that lead to premature termination of endogenous transcripts [ 142 , 146 , 147 ].

Unlike retroviruses, lentiviruses prefer to integrate into transcribed portions of expressed genes in gene-rich regions, distanced from promoters and regulatory elements [ 140 , 142 , 148 ]. The cellular protein LEDGF/p75 aids in the target site selection by binding directly to both the active gene and the viral integrase within the HIV pre-integration complex [ 149 ]. Although the propensity of lentivirus to integrate into the body of expressed genes should increase the incidence of post-transcriptional deregulation, deletion of promoter elements from the lentiviral LTR (self-inactivating (SIN) vectors) has been reported to decrease transcriptional termination, but increase the generation of chimeric transcripts [ 149 ]. Overall, it appears that lentiviral SIN vectors are less likely to cause tumors than retroviral vectors with an active LTR promoter [ 148 , 150 , 151 , 152 ].

The 7.5–10 kb packaging limit of lentiviruses can accommodate the packaging, delivery, and stable integration of Cas9 cDNA, gRNAs, or Cas9 and gRNAs (all-in-one) to cells [ 153 , 154 ]. Often, a selectable marker, such as drug resistance, can also be included to isolate transduced cells. The high transduction efficiency of lentivirus can result in an abundance of CRISPR/Cas9-expressing cells to screen, compared with more traditional transfection methods. Stable and prolonged expression of CRISPR/Cas9 can facilitate targeting of multiple alleles of the gene of interest, resulting in more cells harboring homozygous gene modifications. Conversely, stable integration of CRISPR/Cas9 increases potential off-target effects. Moreover, lentiviral integration itself is a factor that may confound cellular phenotypes and should be considered when characterizing CRISPR-edited cell lines.

2.7. Gene Targeting Using Adeno-Associated Virus

Adeno-associated virus (AAV) is a human parvovirus with a single-stranded DNA genome of 4.7 kb, which was originally identified as a contaminant of adenoviral preparations [ 155 ]. The genome is flanked on both sides by inverted terminal repeats (ITR) and contains two genes, rep and cap [ 156 , 157 ]. Different capsid proteins confer serotype and tissue-specific targeting of distinct AAVs, in vivo. AAV cannot replicate on its own, and requires a helper virus, such as adenovirus or herpes simplex virus (HSV), to provide essential proteins in trans. AAV is the only known virus to integrate into the human genome in a site-specific manner at the AAVS1 site on chromosome 19q13.3-qter [ 158 , 159 , 160 ]. Although the precise mechanism is not well understood, the Rep protein functions to tether the virus to the host genome through direct binding of the AAV ITR and the AAVS1 site [ 158 , 160 , 161 ]. In the recombinant AAV (rAAV) vector system, the rep and cap genes are removed from the packaged virus, resulting in the loss of site-specific integration into the AAVS1 site. Despite removal of Rep, it has been shown that rAAV can still integrate, albeit randomly, into the host genome, via nonhomologous recombination, at low frequencies [ 162 , 163 , 164 ]. Furthermore, numerous clinical trials, to date, have shown that rAAV integration is safe and has no genotoxicity [ 165 , 166 , 167 ]. However, this “safety” is controversial, owing to preclinical studies suggesting genotoxicity in mouse models [ 168 , 169 , 170 , 171 ]. More studies are needed to understand the cellular impact of rAAV integration.

rAAVs have been used to deliver one or two CRISPR guide RNAs (gRNAs), in cells and model animals, by taking advantage of different rAAV serotypes to target specific cells or tissue types. Owing to the packaging capacity of rAAV, SpCas9 must be delivered as a separate virus, unlike lentivirus, which can be delivered as an “all-in-one” CRISPR/Cas9 vector. However, alternate, smaller Cas9s can be packaged into rAAVs [ 172 ]. Furthermore, rAAVs can be used to deliver repair templates or single-stranded donor oligonucleotides (ssODNs) for homology-directed repair (HDR), relying on the single-stranded nature of the AAV genome [ 173 , 174 ]. It has also been observed that rAAVs can integrate into the genome at CRISPR/Cas9-induced breaks in various cultured mouse tissue types, including neurons and muscle [ 175 ]. This observation goes against the notion of rAAVs integrating only at the AAVS1 locus, and should be considered when analyzing and characterizing rAAV-mediated CRISPR-edited cells.

3. Conclusions

There are many approaches to inserting new genetic information into chromosomes in cells and animals. At this time, the most appealing method is single copy gene insertion at a defined locus. This approach has numerous advantages, with respect to reproducible transgene expression. Random insertion transgenesis has been effectively used to probe gene function in mouse models [ 176 ]. It is generally accepted that this requires a spontaneous chromosome break [ 176 ]. Recent NGS data suggest that the repair mechanism resembles chromothripsis [ 118 , 177 ]. In addition to unintended gene disruptions owing to chromosome damage, the random insertion of transgenes exposes them to “position effects” in which their expression is controlled by neighboring genes [ 118 , 178 ]. Ideally, the insertion of reporter cDNAs in the genome results in single copy transgene insertions in defined loci in such a way that endogenous genes are not disrupted, and reporters are placed under the control of specific endogenous promoters [ 179 ]. The application of CRISPR/Cas9 technology to address this problem shows it can be used to achieve these goals [ 63 , 82 , 180 ]. The development of CRISPR/Cas9 base editing technology shows that it is possible to make single-nucleotide changes in the genome [ 181 , 182 , 183 , 184 ]. Base editors have the advantage that double-strand chromosome breaks are not produced, thus lessening the chances of undesirable mutations in the genome. A novel approach to small insertions in the genome by the use of a RNA donor sequence fused to the sgRNA in combination with a reverse transcriptase fused to dead Cas9 also avoids the need to produce double-strand breaks on chromosomes. This approach is referred to as “prime editing” [ 185 ]. CRISPR technology that avoids chromosome breaks, while making changes to the genome, is extremely important in clinical applications where unintended changes can adversely affect patients. These advanced versions of CRISPR technology will be important for future research.

The desire to apply CRISPR/Cas9 for the targeted insertion of transgenes is reflected in the profusion of methods directed towards this purpose [ 63 , 108 , 110 , 112 , 186 , 187 ]. Each method was successfully used to engineer mouse and rat genomes ( Table 1 ). Each method was shown to be more cost-effective and rapid than the application of mouse or rat ES cell technology. For the practitioner of the art, the question remains: which method is most efficient? That is to say, which method minimizes the number of animals needed for zygote production and maximizes the number of gene-targeted founders? One approach to this question is to compare the transgenic efficiency of each method [ 188 ]. The results in Table 1 show that the highest efficiency experiments were obtained when long single-stranded DNA donors and Cas9 ribonucleoproteins were used to produce genetically engineered mice. All methods are very effective compared with traditional methods of gene targeting in zygotes. Perhaps future avenues to even more efficient gene targeting lie in the application of small molecule activators for HDR [ 189 , 190 , 191 ].

Analysis of targeting vector knockin by CRISPR/Cas9 in mouse and rat zygotes.

1 Conditional: A critical exon was flanked by loxP sites, so as to produce a Cre-dependent knockout allele. Reporter: an exogenous coding sequence, such as for a fluorescent protein, was inserted. 2 RNP: ribonucleoprotein; Cas9 protein was complexed with guide RNA. Cas9 mRNA: in vitro transcribed mRNA from a plasmid containing Cas9 mixed with guide RNA. Cas9-mSa: in vitro transcribed mRNA from a plasmid containing Cas9 fused to monomeric streptavidin. 3 ssDNA: single-stranded DNA repair template. BioPCR: PCR was used to prepare biotinylated PCR amplicons. dsDNA: circular double-stranded DNA repair template. HMEJ: homology-mediated end joining; circular double-stranded DNA repair template incorporating sgRNA targets that flank homology arms. Tild: linear double-stranded DNA repair template. AAV: an adeno-associated vector donor was cultured with zygotes loaded with Cas9 RNP, by electroporation. 4 Efficiency, as calculated as the number of genetically engineered mice or rats produced per 100 zygotes treated with CRISPR/Cas9 reagents and transferred to pseudopregnant females.

Author Contributions

Conceptualization, T.L.S. Writing—review and editing, T.M.L.; H.C.K.; and, T.L.S. All authors have read and agreed to the published version of the manuscript.

This research was supported by Institutional Funds from the University of Michigan Biomedical Research Core Facilities.

Conflicts of Interest

The authors declare no conflict of interest.

• Open access
• Published: 29 October 2020

The genome editing revolution: review

• Ahmad M. Khalil   ORCID: orcid.org/0000-0002-1081-7300 1  

Journal of Genetic Engineering and Biotechnology volume  18 , Article number:  68 ( 2020 ) Cite this article

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Development of efficient strategies has always been one of the great perspectives for biotechnologists. During the last decade, genome editing of different organisms has been a fast advancing field and therefore has received a lot of attention from various researchers comprehensively reviewing latest achievements and offering opinions on future directions. This review presents a brief history, basic principles, advantages and disadvantages, as well as various aspects of each genome editing technology including the modes, applications, and challenges that face delivery of gene editing components.

Genetic modification techniques cover a wide range of studies, including the generation of transgenic animals, functional analysis of genes, model development for diseases, or drug development. The delivery of certain proteins such as monoclonal antibodies, enzymes, and growth hormones has been suffering from several obstacles because of their large size. These difficulties encouraged scientists to explore alternative approaches, leading to the progress in gene editing. The distinguished efforts and enormous experimentation have now been able to introduce methodologies that can change the genetic constitution of the living cell. The genome editing strategies have evolved during the last three decades, and nowadays, four types of “programmable” nucleases are available in this field: meganucleases, zinc finger nucleases, transcription activator-like effector nucleases, and the clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR associated protein 9 (Cas9) (CRISPR/Cas-9) system. Each group has its own characteristics necessary for researchers to select the most suitable method for gene editing tool for a range of applications. Genome engineering/editing technology will revolutionize the creation of precisely manipulated genomes of cells or organisms in order to modify a specific characteristic. Of the potential applications are those in human health and agriculture. Introducing constructs into target cells or organisms is the key step in genome engineering.

Conclusions

Despite the success already achieved, the genome editing techniques are still suffering certain difficulties. Challenges must be overcome before the full potential of genome editing can be realized.

In classical genetics, the gene-modifying activities were carried out selecting genetic sites related to the breeder’s goal. Subsequently, scientists used radiation and chemical mutagens to increase the probability of genetic mutations in experimental organisms. Although these methods were useful, they were time-consuming and expensive. Contrary to this, reverse genetics goes in the opposite direction of the so-called forward genetic screens of classical genetics. Reverse genetics is a method in molecular genetics that is used to help understanding the function of a gene by analyzing the phenotypic effects of specific engineered gene sequences. Robb et al. [ 68 ] defined and compared the three terms: “genome engineering”, “genome editing”, and “gene editing”. Genome engineering is the field in which the sequence of genomic DNA is designed and modified. Genome editing and gene editing are techniques for genome engineering that incorporate site-specific modifications into genomic DNA using DNA repair mechanisms. Gene editing differs from genome editing by dealing with only one gene.

This review briefly presents the evolution of genome editing technology over the past three decades using PubMed searches with each keyword of genome-editing techniques regarding the brief history, basic principles, advantages and disadvantages, as well as various aspects of each genome editing technology including the modes, future perspective, applications, and challenges.

Genome-wide editing is not a new field, and in fact, research in this field has been active since the 1970s. The real history of this technology started with pioneers in genome engineering [ 36 , 59 ]. The first important step in gene editing was achieved when researchers demonstrated that when a segment of DNA including homologous arms at both ends is introduced into the cell, it can be integrated into the host genome through homologous recombination (HR) and can dictate wanted changes in the cell [ 10 ]. Employing HR alone in genetic modification posed many problems and limitations including inefficient integration of external DNA and random incorporation in undesired genomic location. Consequently, the number of cells with modified genome was low and uneasy to locate among millions of cells. Evidently, it was necessary to develop a procedure by which scientists can promote output. Out of these limitations, a breakthrough came when it was figured out that, in eukaryotic cells, more efficient and accurate gene targeting mechanisms could be attained by the induction of a double stranded break (DSB) at a specified genomic target [ 70 ].

Furthermore, scientists found that if an artificial DNA restriction enzyme is inserted into the cell, it cuts the DNA at specific recognition sites of double-stranded DNA (dsDNA) sequences. Thus, both the HR and non-homologous end joining (NHEJ) repair can be enhanced [ 14 ]. Various gene editing techniques have focused on the development and the use of different endonuclease-based mechanisms to create these breaks with high precision procedures [ 53 , 78 ] (Fig. 1 ). The mode of action of what is known as site-directed nucleases is based on the site-specific cleavage of the DNA by means of nuclease and the triggering of the cell’s DNA repair mechanisms: HR and NHEJ.

Genome editing outcomes. Genome editing nucleases induce double-strand breaks (DSBs). The breaks are repaired through two ways: by non-homologous end joining (NHEJ) in the absence of a donor template or via homologous recombination (HR) in the presence of a donor template. The NHEJ creates few base insertions or deletion, resulting in an indel, or in frameshift that causes gene disruption. In the HR pathway, a donor DNA (a plasmid or single-stranded oligonucleotide) can be integrated to the target site to modify the gene, introducing the nucleotides and leading to insertion of cDNA or frameshifts induction. (Adapted from [ 78 ])

One of the limitations in this procedure is that it has to be activated only in proliferating cells, adding that the level of activity depends on cell type and target gene locus [ 72 ]. Tailoring of repair templates for correction or insertion steps will be affected by these differences. Several investigations have determined ideal homology-directed repair (HDR) donor configurations for specific applications in specific models systems [ 67 ]. The differences in the activities of the DNA repair mechanisms will also influence the efficiency of causing indel mutations through NHEJ or the classical microhomology-mediated end joining (c-MMEJ) pathway, and even the survival of the targeted cells. The production of such repair in the cell is a sign of a characteristic that errors may occur during splicing the ends and cause the insertion or deletion of a short chain. Simply speaking, gene editing tools involve programmed insertion, deletion, or replacement of a specific segment of in the genome of a living cell. Potential targets of gene editing include repair of mutated gene, replacement of missing gene, interference with gene expression, or overexpression of a normal gene.

The human genome developments paved the way to more extensive use of the reverse genetic analysis technique. Nowadays, two methods of gene editing exist: one is called “targeted gene replacement” to produce a local change in an existing gene sequence, usually without causing mutations. The other one involves more extensive changes in the natural genome of species in a subtler way.

In the field of targeted nucleases and their potential application to model and non-model organisms, there are four major mechanisms of site-specific genome editing that have paved the way for new medical and agricultural breakthroughs. In particular, meganucleases (MegNs), zinc finger nucleases (ZFNs), transcription activator-like effector nuclease (TALENs), and clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) (CRISPR/Cas-9) (Fig. 2 ).

Schematic diagram of the four endonucleases used in gene editing technologies. a Meganuclease (MegN) that generally cleaves its DNA substrate as a homodimer. b Zinc finger nuclease (ZFN) recognizes its target sites which is composed of two zinc finger monomers that flank a short spacer sequence recognized by the FokI cleavage domain. c Transcription activator-like effector nuclease (TALEN) consists of two monomers; TALEN recognizes target sites which flank a fok1 nuclease domain to cut the DNA. d CRISPR/Cas9 system is made of a Cas9 protein with two nuclease domains: human umbilical vein endothelium cells (HuvC) split nuclease and the HNH, an endonuclease domain named for the characteristic histidine and asparagine residue, as well as a single guide RNA (sgRNA). (Adapted from [ 1 , 51 ]; Gaj et al., 2016 [ 53 ];)

Meganucleases (MegNs)

Meganucleases (MegNs) are naturally occurring endodeoxyribonucleases found within all forms of microbial life as well as in eukaryotic mitochondria and chloroplasts. The genes that encode MegNs are often embedded within self-splicing elements. The combination of molecular functions is mutually advantageous: the endonuclease activity allows surrounding introns and inteins to act as invasive DNA elements, while the splicing activity allows the endonuclease gene to invade a coding sequence without disrupting its product. The high specificity of these enzymes is based on their ability to cleave dsDNA at specific recognition sites comprising 14–40 bp (Fig. 2 a). Unlike restriction enzymes, which provide defenses to bacteria against invading DNA, MegNs facilitate lateral mobility of genetic elements within an organism. This process is referred to as “homing” and gives the name homing endonucleases to these enzymes. The high DNA specificity of MegNs makes them a powerful protein scaffold to engineer enzymes for genome manipulation. A deep understanding of their molecular recognition of DNA is an important prerequisite to generate engineered enzymes able to cleave DNA in specific desired genome sites. Crystallographic analyses of representatives from all known MegNs families have illustrated both their mechanisms of action and their evolutionary relationships to a wide range of host proteins. The functional capabilities of these enzymes in DNA recognition vary widely across the families of MegNs. In each case, these capabilities, however, make a balance between what is called orthogonal requirements of (i) recognizing a target of adequate length to avoid overt toxicity in the host, while (ii) accommodating at least a small amount of sequence drift within that target. Indirect readout in protein-DNA recognition is the mechanism by which the protein achieves partial sequence specificity by detecting structural features on the DNA.

Several homing endonucleases have been used as templates to engineer tools that cleave DNA sequences other than their original wild-type targets.

Meganucleases can be divided into five families based on sequence and structure motifs: LAGLIDADG, GIY-YIG, HNH, His-Cys box, and PD-(D/E) XK [ 74 ]. I-CreI is a homodimeric member of MegNs family, which recognizes and cleaves a 22-bp pseudo-palindromic target (5′-CAAAACGTCGTGAGACAGTTTG-3′). The important role of indirect readout in the central region of the target DNA of these enzymes I-CreI suggested that indirect readout may play a key role in the redesign of protein-DNA interactions. The sequences of the I-CreI central substrate region, four bp (± 1 and ± 2) called 2NN, along with the adjacent box called 5NNN, are key for substrate cleavage [ 64 ]. Changes in 2NN significantly affect substrate binding and cleavage because this region affects the active site rearrangement, the proper protein-DNA complex binding, and the catalytic ion positioning to lead the cleavage.

An exhaustive review of each MegN can be found in Stoddard [ 75 ] as well as in Petersen and Niemann [ 63 ]. Several MegNs have been used as templates to engineer tools that cleave DNA sequences other than their original wild-type targets. This technology have advantages of high specificity of MegNs to target DNA because of their very long recognition sites, ease in delivery due to relatively small size, and giving rise to more recombinant DNA (i.e., more recombinogenic for HDR) due to production of a 3′ overhang after DNA cleavage. This lowers the potential cytotoxicity [ 53 , 78 ].

Meganucleases have several promising applications; they are more specific than other genetic editing tools for the development of therapies for a wide range of inherited diseases resulting from nonsense codons or frameshift mutations. However, an obvious drawback to the use of natural MegNs lies in the need to first introduce a known cleavage site into the region of interest. Additionally, it is not easy to separate the two domains of MegNs: the DNA-binding and the DNA-cleavage domains, which present a challenge in its engineering. Another drawback of MegNs is that the design of sequence-specific enzymes for all possible sequences is time-consuming and expensive. Therefore, each new genome engineering target requires an initial protein engineering step to produce a custom MegN. Thus, in spite of the so many available MegNs, the probability of finding an enzyme that targets a desired locus is very small and the production of customized MegNs remains really complex and highly inefficient. Therefore, routine applications of MegNs in genome editing is limited and proved technically challenging to work with [ 24 ].

Zinc finger nucleases (ZFNs)

The origin of genome editing technology began with the introduction of zinc finger nucleases (ZFNs). Zinc finger nucleases are artificially engineered restriction enzymes for custom site-specific genome editing. Zinc fingers themselves are transcription factors, where each finger recognizes 3–4 bases. Zinc finger nucleases are hybrid heterodimeric proteins, where each subunit contains several zinc finger domains and a Fok1 endonuclease domain to induce DSB formation. The first is zinc finger, which is one of the DNA binding motifs found in the DNA binding domain of many eukaryotic transcription factors responsible for DNA identification. The second domain is a nuclease (often from the bacterial restriction enzyme FokI) [ 6 ]. When the DNA-binding and the DNA-cleaving domains are fused together, a highly specific pair of “genomic scissors” is created (Fig. 2b ). In principle, any gene in any organism can be targeted with a properly designed pair of ZFNs. Zinc finger recognition depends only on a match to DNA sequence, and mechanisms of DNA repair, both HR and NHEJ, are shared by essentially all species. Several studies have reported that ZFNs with a higher number of zinc fingers (4, 5, and 6 finger pairs) have increased the specificity and efficiency and improved targeting such as using modular assembly of pre-characterized ZFs utilizing standard recombinant DNA technology.

Since they were first reported [ 41 ], ZFN was appealing and showed considerable promise and they were used in several living organisms or cultured cells [ 11 ]. The discovery of ZFNs overcame some of the problems associated with MegNs applications. They facilitated targeted editing of the gene by inducing DSBs in DNA at specific sites. One major advantage of ZFNs is that they are easy to design, using combinatorial assembly of preexisting zinc fingers with known recognition patterns. This approach, however, suffered from drawbacks for routine applications. One of the major disadvantages of the ZFN is what is called “context-dependent specificity” (how well they cleave target sequence). Therefore, these specificities can depend on the context in the adjacent zinc fingers and DNA. In other terms, their specificity does not only depend on the target sequence itself, but also on adjacent sequences in the genome. This issue may cause genome fragmentation and instability when many non-specific cleavages occur. It only targets a single site at a time and as stated above. Although the low number of loci does not usually make a problem for knocking-out editing, it poses limitation for knocking in manipulation [ 32 ]. In addition, ZFNs cause overt toxicity to cells because of the off-target cleavages. The off-target effect is the probability of inaccurate cut of target DNA due to single nucleotide substitutions or inappropriate interaction between domains.

Transcription activator-like effector nucleases (TALENs)

The limitations mentioned in the previous section paved the way for the development of a new series of nucleases: transcription activator-like effector nucleases (TALENs), which were cheaper, safer, more efficient, and capable of targeting a specified region in the genome [ 13 ].

In principle, the TALENs are similar to ZFNs and MegNs in that the proteins must be re-engineered for each targeted DNA sequence. The ZFNs and TALENs are both modular and have natural DNA-binding specificities. The TALEN is similar to ZFN in that it is an artificial chimeric protein that result from fusing a non-specific FokI restriction endonuclease domain to a DNA-binding domain recognizing an arbitrary base sequence (Fig. 2c ). This DNA-binding domain consists of highly conserved repeats derived from transcription activator-like effectors (TALE). When genome editing is planned, a pair of TALEN is used like ZFNs. The TALE protein made of three domains: an amino-terminal domain having a transport signal, a DNA-binding domain which is made of repeating sequences of 34 amino acids arranged in tandem, and a carboxyl-terminal domain having a nuclear localization signal and a transcription activation domain. Of the 34 amino acids, there is a variable region of two amino acid residues located at positions 12 and 13 called repeat variable di-residues (RVD). This region has the ability to confer specificity to one of the any four nucleotide bps [ 15 ].

Unlike ZFNs, TALENs had advantages in that one module recognizes just one nucleotide in its DNA-binding domain, as compared with 3 bps recognized by the first single zinc finger domains [ 39 ]. So, interference of the recognition sequence does not occur even when several modules are joined. In theory, because cleavage of the target sequence is more specific than ZFN, it became possible to target any DNA sequence of any organism genome. This difference facilitates creation of TALEN systems which recognize more target sequences. Another benefit of the TALEN system over ZFN’s for genome editing is that the system is more efficient in producing DSBs in both somatic cells and pluripotent stem cells [ 35 ]. In addition, TALENs exhibit less toxicity in human cell lines due to off-target breaks that result in unwanted changes and toxicity in the genome. Another advantage of TALENs is a higher percentage of success in genome editing through cytoplasmic injection of TALEN mRNA in livestock embryos than observed with ZFN induction [ 39 ]. In addition, TALENs have been more successfully used in plant genome engineering [ 88 ]. It is hoped that TALENs will be applied in the generation of genetically modified laboratory animals, which may be utilized as a model for human disease research [ 24 , 39 ].

The TALEN-like directed development of DNA binding proteins was employed to improve TALEN specificity by phage-assisted continuous evolution (PACE). The improved version was used to create genetically modified organisms [ 34 ]. Nucleases which contain designable DNA-binding sequences can modify the genomes and have the promise for therapeutic applications. DNA-binding PACE is a general strategy for the laboratory evolution of DNA-binding activity and specificity. This system can be used to generate TALEN with highly improved DNA cutting specificity, establishing DB-PACE as a diverse approach for improving the accuracy of genome editing tools. Thus, similar to ZFN, TALEN is used for DSBs as well as for knocking in/knocking out. In comparison with the ZFN, two important advantages for this editing technique have been reported: first, the simple design, and second, the low number of off-target breaks [ 35 ].

In spite of the improvement and simplification of the TALEN method, it is complicated for whom not familiar with molecular biological experiments. Moreover, it is confronted with some limitations, such as their large size (impeding delivery) in comparison to ZFN [ 24 , 39 ]. The superiority of TALEN relative to ZFN could be attributed to the fact that in the TALEN each domain recognizes only one nucleotide, instead of recognizing DNA triplets in the case of ZEF. The design of TALEN is commonly more obvious than ZNF. This results in less intricate interactions between the TALEN-derived DNA-binding domains and their target nucleotides than those among ZNF and their target trinucleotides [ 35 , 39 ].

Clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9)

The CRISPR/Cas system is the most recent platform in the field of genome editing. The system was developed in 2013 and is known as the third generation genomic editing tools. The clustered regularly interspaced short palindromic repeats, which are sometimes named “short regularly spaced repeats” were discovered in the 1980s. Computational analysis of these elements showed they were found in more than 40% of sequenced bacteria and 90% of archaea [ 37 , 56 ]. The acronym CRISPR was suggested, and a group of genes adjacent to the CRISPR locus, which was termed “CRISPR-associated system”, or Cas was established [ 37 ]. Cas proteins coded by these genes carry functional domains similar to endonucleases, helicases, polymerases, and nucleotide-binding proteins. In addition, the role of CRISPRs as bacterial and archaeal adaptive immunity system against invading bacteriophages and other and in DNA repair was realized [ 17 , 77 ].

Unlike the two previous technologies (ZFN and TALEN), in which the recognition of the DNA site was based on the sequence recognition by artificial proteins requiring interaction between protein and DNA, the DNA recognition of the CRISPR/Cas system is based on RNA-DNA interactions. This offers several advantages over ZFNs and TALENs. These include easy design for any genomic targets, easy prediction regarding off-target sites, and the probability of modifying several genomic sites simultaneously (multiplexing). CRISPR-Cas systems are diverse and have been classified thus far into two classes, six types, and over 20 subtypes based on locus arrangement and signature cas genes [ 33 , 44 , 51 ]. Types I, III, and IV, with multiprotein crRNA-effector complexes, are class 1 systems; types II, V, and VI, with a single protein-crRNA effector complex, are class 2. All CRISPR-Cas systems require Cas proteins and crRNAs for function, and CRISPR- cas expression is a prerequisite to acquire new spacers, process pre-crRNA, and assemble ribonucleoprotein crRNA interference complexes for target degradation. Herein, we will focus on the CRISPR-Cas9 technology, the reader should keep in mind other available variants of the system such as CRISPR-Cas6 [ 5 ], CRISPR-Cas12a, -Cas12b [ 42 ], as well as the most recently discovered c2c2 (Cas13a) and c2c6 (Cas13b [ 19 , 69 ]. The CRISPR/Cas9 system is made of Cas9 nuclease and single-guide RNA (sgRNA). The sgRNA is an engineered single RNA molecule containing crispr RNA and tracr RNA parts. The sgRNA recognizes the target sequence by standard Watson-Crick base pairing. It has to be followed by a DNA motif called a protospacer adjacent motif (PAM). The commonly used wild-type Streptococcus pyogenes Cas (SpCas9) protein has a specific PAM sequence, 5’-NGG-3’, where “N” can be any nucleotide base followed by two guanine (“G”) nucleobases. This sequence is located directly downstream of the target sequence in the genomic DNA, on the non-target strand. Targeting is constrained to every 14 bp (12 bp from the seed sequence and 2 bp from PAM) [ 15 ]. SpCas9 variants may increase the specificity of genome modifications at DNA targets adjacent to NGG PAM sequences when used in place of wild-type SpCas9.

DNA cleavage is performed by Cas9 nuclease and can result in DSB in the the case of a wild-type enzyme, or in a SSB when using mutant Cas9 variants called nickases (Fig. 2d ). It should be emphasized that the utilization of this approach in editing eukaryotes’ genome only needs the manipulation of a short sequence of RNA, and there is no need for complicated manipulations in the protein domain. This enables a faster and more cost-effective design of the DNA recognition moiety compared with ZFN and TALEN technologies. Applications of CRISPR-Cas9 systems are variable like those for ZFNs, TALENs, and MegNs. But, because of the relative simplicity of this system, its great efficiency and high tendency for multiple functions and library construction, it can be applied to different species and cell types [ 35 ].

As shown in Fig. 3 , in all CRISPR/Cas systems, immunity occurs in three distinct stages [ 77 , 81 ]: (1) adaptation or new spacer acquisition, (2) CRISPR transcription and processing (crRNA generation), and (3) interference or silencing. The advantages of the CRISPR/Cas system superseded those of both of the TALEN and ZFN tools, the ZFN in particular. This is due to its target design simplicity since the target specificity depends on ribonucleotide complex formation and non-protein/DNA recognition. In addition, the CRISPR/Cas approach is more efficient because changes can be introduced directly by injecting RNAs that encode the Cas protein and gRNA into developing embryos. Moreover, multigene mutations can be induced simultaneously by injecting them with multiple gRNAs. This is an example that explains the rapid spread of CRISPR/Cas 9 application in various fields. Still, the system has certain drawbacks. Although the CRISPR/Cas9 is much less complicated than TALEN, in terms of execution and construction, the off-target effect in CRISPR/Cas9 is higher than TALEN. Since the DSB results only after accurate binding of a pair of TALEN to the target sequence, the off-target effect problem is considered to be low. These two are different in restriction of target sequence. CRISPR/Cas9 is much more efficient than TALEN in multiple simultaneous modification. Table 1 compares the three main systems of site-directed synthetic nuclease employed in genome editing: ZFN, TALEN, and CRISPR/Cas9.

Schematic representation of CRISPR loci and targeting of DNA sequence, which include Cas genes, a leader sequence, and several spacer sequences derived from engineered or foreign DNA that are separated by short direct repeat sequences. The three major steps of CRISPR-Cas immune systems. In the adaptation phase, Cas proteins excise specific fragments from foreign DNA and integrate it into the repeat sequence neighboring the leader at the CRISPR locus. Then, CRISPR arrays are transcribed and processed into multiple crRNAs, each carrying a single spacer sequence and part of the adjoining repeat sequence. Finally, at the interference phase, the crRNAs are assembled into different classes of protein targeting complexes (cascades) that anneal to, and cleave, spacer matching sequences on either invading element or their transcripts and thus destroy them. (Adapted from [ 3 , 53 , 78 ])

The off-target effect is an essential subject for future studies if CRISPR/Cas9 is to achieve its promises as a powerful method for genome editing. Non-specific and unintended genetic modifications (off-target effect) can result from the use of CRISPR/Cas9 system which is one of the drawbacks of this tool. Therefore, this point should be considered for use in researches. One strategy to reduce the off-target activity is to replace the Streptococcus pyogenes Cas9 enzyme (SpyCas9) for a mutant Cas9 nickase (nSpyCas9; ncas9), which cleaves a single strand through the inactivation of a nuclease domain Ruvc or HNH [ 9 ]. Our understanding of off-target effects remains fragmentary. A deeper understanding of this phenomenon is needed. Several approaches that could be followed to characterize the binding domains and consequently Cas9 targeting specificity have been reviewed and summarized [ 83 ].

It has previously been stated that CRISPR/Cas9 system needs both gRNA and PAM to detect its target sequence of interest by integration of a gRNA component that binds to complementary double-stranded DNA sequences. Cell culture studies have shown that off-target effects may be due to the incorrect detection of genomic sequences by sgRNA. This, in turn, affects cleavage when the mismatch is in the vicinity of the PAM (up to 8 bases), but if the PAM is too far apart, these effects will be small [ 4 ], even a slight mismatch between sgRNA and target sequences can lead to a failure. Dependence of this method on specific PAM sequences to act functionally limits the number of target loci, and it can reduce off-target breaks [ 86 ]. For this goal, another type of specific PAM-containing nucleases has been prepared to compensate for this limitation. Genetic engineering and enzyme changing have also been able to overcome the limitation [ 42 ]. For a sgRNA, many similar sequences depending on the genome size of the species may exist [ 86 ]. Interestingly, the initial targeting scrutiny of the CRISPR/Cas9-sgRNA complex showed that not every nucleotide base in the gRNA is necessary to be complementary to the target DNA sequence to effect Cas9 nuclease activity. Regarding that where the similar sequences are found in the genome, their breaks could lead to malignancies or even death [ 86 ]. Various methods have been proposed to prevent off-target breaks, among which the double nicking method, the FokI-dCas9 fusion protein method, and the truncated sgRNA method [ 76 ] (Fig. 4 ).

a Summary of the Cas9 nickases methods in efficient genome editing. Two gRNAs target opposite strands of DNA. These double nicks create a DSB that is repaired using non-homologous end joining (NHEJ) or edits via homology-directed repair (HDR) (adapted from www.addgene.org/crispr/nick ). b FokI-dCas 9 fusion protein method. Two FokI-dCas9 fusion proteins are used to adjacent target sites by two different sgRNAs to facilitate FokI dimerization and DNA cleavage. These fusions would have enhanced specificity compared to the standard monomeric Cas9 nucleases and the paired nickase system because they should require two sgRNAs for activity. c Truncated sgRNA method. Cas9 interacting with either a full-length sgRNA (20 nucleotide sequence complementary to target site) or truncated gRNA (less than 15 nucleotide sequence complementary to target site). (Retrieved from blog.addgene.org )

To overcome these problems, researchers explored another generation of base editing technologies, which combine CRISPR and cytidine deaminase (Fig. 5 ). This is a diverse method called CRISPR-SKIP (Fig. 6 ) which uses cytidine deaminase single-base editors to program exon skipping by mutating target DNA bases within splice acceptor sites [ 25 ]. Given its simplicity and precision, CRISPR-SKIP will be widely applicable in gene therapy. Base editing utilizes Cas9 D10A nickases fused to engineered base deaminase enzymes to make single base changes in the DNA sequence without the need of DNA DSB. Also, base editing does not require an external repair template. The Cas9 nickase part of the base editor protein plays a dual function. The first is to target the deaminase activity to the wanted region and the second is to localize the enzyme to certain regions of double-stranded RNA. The deaminase domains in base editors (BEs) occur in two versions: either adenosine deaminase or cytosine deaminase, which catalyze only base transitions (C to T and A to G) and cannot produce base transversions [ 26 , 68 ]. In these base editing tools, the targeted activity of adenosine deaminase can result in an A:T to G:C sequence alteration in a very similar way [ 26 , 68 ].This approach avoided the requirement of breaking DNA to induce an oligonucleotide. In addition, compared to knocking system, it exerted a higher output with lower off-targets [ 40 , 43 ]. Adenosine is deaminated to inosine (I) that is subsequently utilized to repair the nicked strand with a cytosine, and the I:C base pair is resolved to G:C [ 26 ]. More recently, new genome editing technologies have been developed: glycosylase base editors (GBEs), which consist of a Cas9 nickase, a cytidine deaminase, and a uracil-DNA glycosylase (Ung), are capable of transversion mutations by changing C to A in bacterial cells and from C to G in mammalian cells [ 45 , 89 ]. The new BEs can also be designed to minimize unwanted (“off-target”) mutations that could potentially cause undesirable side effects. The novel BE platform may help researchers understand and correct genetic diseases by selective editing of single DNA “alphabets” across nucleobase classes. However, the technique with this new class of transversion BEs is still at an early stage and requires additional optimization, so it would be premature to say this is ready for the clinic applications.

Base editing uses engineered Cas9 variants to induce base changes in a target sequence. Cas9 nickase is fused to a base deaminase domain. The deaminase domain works on a targeted region within the R-loop after target binding and R-loop formation. Simultaneously, the target strand is nicked. DNA repair is started in response to the nick using the strand which contains the deaminated base as a repair template. Repair leads to a transition mutations: C:G to T:A and A:T to G:C for cytosine and adenosine base editors, respectively [ 68 ]

Essential steps in CRISPR-SKIP targeting approach: a Nearly every intron ends with a guanosine (asterisked G). It is hypothesized that mutations that disrupt this highly conserved G within the splice acceptor of any given exon in genomic DNA would lead to exon skipping by preventing incorporation of the exon into mature transcripts base. b In the presence of an appropriate PAM sequence, this G can be effectively mutated by converting the complementary cytidine to thymidine using CRISPR-Cas9 C>T single-base editors. (From [ 25 ])

Gene delivery

From biotechnology’s point of view, the main obstacle that is facing molecular technology is to select the right method that is simple but effective to transfer the gene to the host cell. The components of gene editing have to be transferred to the cell/nucleus of interest using in vivo, ex vivo, or in vitro route. In this regard, several concerns must be considered including physical barriers (cell membranes, nuclear membranes) as well as digestion by proteases or nucleases of the host. Another important issue is the possible rejection by the immune system of the host if the components are delivered in vivo. In general, the gene delivery routes can be categorized in three classes of physical delivery, viral vectors, and non-viral agents. Although the direct delivery of construct plasmids may sound easy and more efficient and specific than the physical and the chemical methods, it proves to be an inappropriate choice because the successful gene delivery system requires the foreign genetic molecule to remain stable within the host cells [ 52 ]. The other possible procedure is to use viruses. However, because plant cells have thick walls, the gene transfer systems for plants involve transient and stable transformation using protoplast-plasmid in vitro [ 54 ]: agrobacterium-mediated transformation, gene gun and viral vectors (transient expression by protoplast transformation), and agro-infiltration [ 1 ]. Viruses may present a suitable vehicle to transfer genome engineering components to all plant parts because they do not require transformation and/or tissue culture for delivering and mutated seeds could easily recovered. For many years, scientists employed different species of Agrobacterium to systematically infect a large number of plant species and generate transgenic plants. These bacterial species have small genome size and this facilitates cloning and agroinfections, and the virus genome does not integrate into plant genomes [ 1 ].

Of the challenges and approaches of delivering CRISPR, it was pointed out [ 18 , 51 ] that although the present genome engineering is in favor of CRISPR tools, TALENs may still be of a primary choice in certain experimental species. For example, TALENs have been utilized in targeted genomic editing in Xenopus tropicalis by knocking-out Klf4 [ 49 , 50 ] or thyroid hormone receptor α [ 23 ]. In addition, TALENs have been utilized to modify genome of human stem cells [ 47 ]. Also TALEN approach has been applied to create amniotic mesenchymal stem cells overexpressing anti-fibrotic interleukin-10 [ 12 ]. Lately, a geminivirus genome has been prepared to deliver various nucleases platforms (including ZFN, TALENs, and the CRISPR/Cas system) and repair template for HR of DSBs [ 62 ].

To deliver the carrying DNA sequence to target cells, non-viral techniques such as electroporation, lipofection, and microinjection can also be used [ 18 ]. In addition, these techniques also reduce off-target cleavages problems. Gene transfer via microinjection is considered the gold standard procedure since its efficiency is approximately 100% [ 85 ]. The advantage of this approach is its high efficacy and less constrains on the size of the delivery. A disadvantage is that it can be employed only in in vitro or ex vivo cargo. Recently, small RNAs, including small interfering RNA (siRNA) and microRNA (miRNA), have been widely adopted in research to replace laboratory animals and cell lines. Development of innovative nanoparticle-based transfer systems that deliver CRISPR/Cas9 constructs and maximize their effectiveness has been tested in the last few years [ 29 , 58 ].

Applications of gene technology

The ability of the abovementioned gene delivery systems to target and manipulate the genome of living organisms has been attractive to many researchers worldwide. Despite all limitations, the interest in this technology has developed its capabilities and enhanced its scope of applications. Genome/gene engineering technology is relatively applicable and has potential to effectively and rapidly revolutionize genome surgery and will soon transform agriculture, nutrition, and medicine. Some of the most important applications are briefly described below.

Plant-based genome editing

The appearance of genome editing has been appealing especially to agricultural experts. One of the major goals for utilizing genome editing tools in plants is to generate improved crop varieties with higher yields and clear-cut addition of valuable traits such as high nutritional value, extended shelf life, stress tolerance, disease and pest resistance, or removal of undesirable traits [ 1 ]. However, several obstacles related to the precision of the genetic manipulations and the incompatibility of the host species have hampered the development of crop improvements [ 2 ]. The use of site-specific nucleases is one of the important promising techniques of gene editing that helped overcome certain limitations by specifically targeting a suitable site in a gene/genome. The employment of the gene editing technologies, including those discussed in this review, seems to be endless ever since their emergence, and several improvements in original tools have further brought accuracy and precision in these methods [ 78 ].

Animal-based genome editing

Recent genome editing techniques has been extensively applied in many organisms, such as bacteria, yeast, and mouse [ 53 , 73 ]. Genetic manipulation tools cover a wide range of fields, including the generation of transgenic animals using embryonic stem cells (ESC), functional analysis of genes, model development for diseases, or drug development. Genome editing techniques have been used in many various organisms. Among the livestock and aquatic species, ZFN is only used for zebrafish, but two other technologies, TALEN and CRISPR, have been used at the cell level in chicken, sheep, pig, and cattle. Engineered endonucleases or RNA-guided endonucleases (RGENs) mediated gene targeting has been applied directly in a great number of animal organisms including nematodes and zebrafish [ 20 , 57 ], as well as pigs [ 71 , 85 ]. Since the first permission to use CRISPR/Cas9 in human embryos and in vivo genome editing via homology-independent targeted integration (HITI), an increasing number of studies have identified striking differences between mouse and human pre-implantation development and pluripotency [ 66 ], highlighting the need for focused studies in human embryos. Therefore, more specific criteria and widely accepted standards for clinical research have to be met before human germline editing would be deemed permissible [ 31 ]. In this regard, results of some research on the human genome editing have been questioned. The “He Jiankui experiments at the beginning of 2019”, which claimed to have created the world’s first genetically edited babies, is simply the most recent example. He Jiankui said he edited the babies’ genes at conception by selecting CRISPR/cas9 to edit the chemokine receptor type 5 (CCR5) gene in cd4+ cells in hopes of making children resistant to the AIDS virus, as their father was HIV-positive. Researchers said He’s actions exposed the twins to unknown health risks, possibly including a higher susceptibility to viral illnesses. For more information on the scientific reactions around the world, the reader may find helpful several excellent sources of information [ 38 , 49 , 79 , 84 ].

• Gene therapy

The original principles of gene therapy arose during the 1960s and early 1970s when restriction enzymes were utilized to manipulate DNA [ 22 ]. Since then, researchers have done great efforts to treat genetic diseases but treatment for multiple mutations is difficult. Different clinical therapy applications have been attempted to overcome these problems. Much of the interest in CRISPR and other gene editing methods revolves around their potential to cure human diseases. It is hoped that eradication of human diseases is not too far to achieve via the CRISPR system because it was employed in other fields of biological sciences such as genetic improvement and gene therapy. It is important to mention that the therapeutic efficiency of gene editing depends on several factors, such as editing efficacy, which varies widely depending on the cell type, senescence status, and cell cycle status of the target [ 69 ]. Other factors that also influence therapeutic effectiveness include cell aptitude, which refers to the feasibility of accomplishing a therapeutic modification threshold, and the efficient transfer of programmable nuclease system to the target tissue, which is only considered to be effective if the engineered nuclease system reaches safely and efficiently to the nucleus of the target cell. Finally, the precision of the editing procedure is another important aspect, which refers to only editing the target DNA without affecting any other genes [ 80 ].

The genome editing tools have enabled scientists to utilize genetically programmed animals to understand the cause of various diseases and to understand molecular mechanisms that can be explored for better therapeutic strategies (Fig. 7 ). Genome editing gives the basis of the treatment of many kinds of diseases. In preliminary experiments, the knocking-in procedure was used to reach this goal. There are examples of gene editing techniques applied in different genetic diseases in cell lines, disease models, and human [ 48 , 53 , 82 ]. These encouraging results suggest the therapeutic capability of these gene editing strategies to treat human genetic diseases including Duchenne muscular dystrophy [ 8 , 28 , 55 ], cystic fibrosis [ 21 ], sickle cell anemia [ 62 ], and Down syndrome [ 7 ]. In addition, this technology has been employed in curing Fanconi anemia by correcting point mutation in patient-derived fibroblasts [ 60 ], as well as in hemophilia for the restoration of factor VIII deficiency in mice [ 61 , 87 ]. The CRISPR tools have also demonstrated promising results in diagnosis and curing fatal diseases such as AIDS and cancer [ 16 , 30 , 84 ].

Outline of the ex vivo and in vivo genome editing procedures for clinical therapy. Top: In the ex vivo editing therapy, cells are removed from a patient to be treated, corrected by gene editing and then re-engrafted back to the patient. To achieve therapeutic success, the target cells must be capable of surviving in vitro and autologous transplantation of the corrected cells. Below: In the in vivo editing therapy, designed nucleases are administered using viral or non-viral techniques and directly injected locally to the affected tissue, such as the eye, brain, or muscle. (Adapted from [ 48 ])

Other applications

The applications mentioned above were more about knock out or modification of genes Gapinske et al. [ 25 ]. However due to inactivate nuclease activity nature of the dCas9, CRISPR can be used in other applications as well. By selecting the target sequence, gene expression can be controlled by inhibiting the transcription rate of RNA polymerase II (polII) or inhibiting the transcription factor binding [ 65 ]. Additionally, combining gene expression inhibitors such as Krüppel-associated box with the inactivated Cas9 has led to generate a special kind of gene inhibitors, which are called CRISPR interference (CRISPRi), and downregulate gene expression [ 46 ]. It is also possible to control gene expression by fusing transcription-activating molecule, the transcription-repressing molecule, or the genome-modifying molecule to dCas9 [ 27 ].

Genome editing is a fast-growing field. Editing nucleases have revolutionized genomic engineering, allowing easy editing of the mammalian genome. Much progress has been accomplished in the improvement of gene editing technologies since their discovery. Of the four major nucleases used to cut and edit the genome, each has its own advantages and disadvantages, and the choice of which gene editing method depends on the specific situation. The current genome editing techniques are still buckling up with problems, and it is difficult to perform genome editing in cells with low transfection efficiency or in some cultured cells such as primary cultured cells. Genotoxicity is an inherent problem of enzymes that act on nucleic acids, though one can expect that highly specific endonucleases would reduce or abolish this issue. Exceptional efforts are needed in future to complement and offer something novel approaches in addition to the already existing ones. It is anticipated that research in gene editing is going to continue and tremendously advance. With the development of next-generation sequencing technology, new extremely important clinical applications, such as manufacturing engineered medical products, eradication of human genetic diseases, treatment of AIDS and cancers, as well as improvement of crop and food, will be introduced. Combination of genomic modifications induced by targeted nucleases to their own self-degradation, self-inactivating vectors may help overcoming confronting limitations discussed above to improve the specificity of genome editing, especially because the frequency of off-target modifications. Our understanding of off-target effects remains poor. This is a vital area for continued study if CRISPR/Cas9 is to realize its promise. Regarding gene cargo delivery systems, this remains the greatest obstacle for CRISPR/Cas9 use, and an all-purpose delivery method has yet to emerge. The union between genome engineering and regenerative medicine is still in its infancy; realizing the full potential of these technologies in reprograming the fate of stem/progenitor cells requires that their functional landscape be fully explored in these genetic backgrounds. Humankind can only wait to see what the potential of these technologies will be. One major question is whether or not the body’s immune response will accept or reject the foreign genetic elements within the cells. Another important concern is that along with the revolutionary advances of this biotechnology and related sciences, bioethical concerns and legal problems related to this issue are still increasing in view of the possibility of human genetic manipulation and the unsafety of procedures involved [ 49 , 50 , 66 ]. The enforcement of technical and ethical guidelines, and legislations should be considered and need serious attention as soon as possible.

Availability of data and materials

Not applicable

Abbreviations

CRISPR-associated protein 9

Clustered regularly interspaced short palindromic repeats

Double-stranded break

Embryonic stem cells

Homology-directed repair

Homology-independent targeted integration

Homologous recombination

Human umbilical vein endothelium cells

Intron-encoded endonuclease

• Meganucleases

Microhomology-mediated end joining

Non-homologous end joining

Phage-assisted continuous evolution

Protospacer adjacent motifs

RNA-guided endonucleases

Repeat variable di-residues

Single guide RNA

Streptococcus pyogenes Cas9

Single-strand break

Transcription activator-like effector nuclease

Zinc finger nucleases

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In this chapter I will apply the PP to ethical and policy issues related to genetic engineering of microbes, plants, animals, and human beings. I will argue that the PP can provide some useful insights into these issues, due to the scientific and morally uncertainty surrounding the consequences of genetic engineering for public health, the environment, society, and patients.

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By “genetic engineering” I mean technologies that involve direct modification or alteration of the genomes of cells or organisms. Changes brought about by genetic engineering might or might not be inheritable, depending on the type of change and the organism. Modification of the genomes of somatic cells in humans (discussed below) does not normally result in inheritable genetic changes, but modification of human germ cells, sperm, eggs, or embryos does (Resnik et al. 1999 ). Modification of bacterial genomes always results in inheritable genetic changes because bacteria are unicellular organisms. Ooplasm transfer, nuclear transfer, and reproductive cloning in human beings raise important ethical and social issues, but these procedures are not genetic engineering, according to my definition, because their purposes is not modify genomes, even though they involve the manipulation of genetic material. Synthetic biology uses genetic engineering methods to design cells, organisms, and biological system that do not already exist in the natural world (Biotechnology Innovation Organization 2020b ).

Some viruses encode their genetic information in RNA (ribonucleic acid).

A polymer is a large molecule.

James Watson (1928–) and Francis Crick (1916–2004) won the Nobel Prize in Physiology of Medicine in 1962 for discovering the structure of DNA. Their model was confirmed by Rosalind Franklin’s x-ray crystallography data, Watson and Crick did not name Franklin as an author on the paper that described their model of the structure of DNA. Franklin (1920–1958) was also not awarded the Nobel Prize for her contribution, because she died of ovarian cancer in 1958, and the Nobel Prize is not awarded posthumously (Maddox 2003 ).

Because mitochondria have their own DNA, scientists have speculated that mitochondria were at one time independent organisms that became incorporated into primordial, unicellular organisms (Alberts et al. 2015 ).

Prokaryotes are single-celled organisms with no distinct cell nucleus or organelles.

Mitochondria replicate independently of the cell.

Most higher life forms, including most plants, mammals, and human beings, are diploid (Alberts et al. 2015 ).

Many species of plants and animals that reproduce sexually can also propagate asexually. Growing a new plant from a cutting is a form of asexual propagation.

Plant stem cells can also generate different tissue types.

Berg, Gilbert, and Sanger won the Nobel Prize in chemistry in 1980 for their development of recombinant DNA techniques (Nobel Prize.org 2021 ).

Doudna and Charpentier won the Nobel Prize in Chemistry in 2000 for the discovery of CRISPR (Ledford and Callaway 2020 ).

Laboratory animals are used to produce monoclonal antibodies. An antigen is introduced into the animal, which produces antibodies in its lymphocyte cells. These cells are cultured and then antibodies are isolated. Since these antibodies would be rejected by the human immune system, the cells are genetically modified so that they produce antibodies with a human protein component, or humanized antibodies. The genetically modified cells are then cultured and humanized antibodies are isolated for production (GenScript 2020 ).

Somatic cells are cells other than the reproductive or germ cells, such as skin, nerve, muscle, liver or bone marrow cells.

Monsanto has developed GM crops (known as Bt crops) that produce Bacillus thuringiensis toxins, which are deadly to insects. Farmers were already using these toxins as pesticides were Bt crops were developed (Resnik 2012 ).

Monsanto has developed GM crops (known as “Roundup Ready” crops) that are immune to the effects of glyphosate, the active ingredient in the widely-used herbicide Roundup ™. Farmers can control weeds with damaging their crops by spraying their crops with Roundup (Resnik 2012 ).

Golden rice, for example, contains more beta carotene than normal rice (McDivitt 2019 ).

In 2018, 228 million people worldwide contracted malaria and 405,000 people died from the disease (World Health Organization 2020a ). About 390 million people contract the dengue virus each year and about 4000 die from the disease (World Health Organization 2020b ).

Oxitec has also genetically engineered diamondback moths (Plutella xylostella) to control these populations. Diamondback moths are a destructive pests that feed on cauliflower, cabbage, broccoli and canola (Campbell 2020a ).

E.g. Bt crops. See Footnote 12.

These are the sorts of problems encountered by the natural law approaches to morality, discussed in Chapter 3 .

Most defenders of the slippery slope argument in genetic only apply it to using genome editing in humans, but it could be applied to other applications of genetic engineering.

I am assuming that GM microbes will not be intentionally released into the environment, which would create risks not discussed here. Scientists have developed GM microbes to clean up oil spills but have not deployed them yet, mostly due to regulatory issues. In nature, microbes already play an important role in cleaning up oil spills (Ezezika and Singer 2010 ).

The reproduction rate is how many people infected persons infect. R 0  = 1 means that an infected person infects one more person on average; R 0  = 2 means an infected person infects two people on average.

It is worth noting, however, that a voluntary moratorium was a reasonable option when this technology was emerging in the 1970s.

As noted in Chapter 6 , a black market for alcohol emerged during Prohibition era in the US (1919–1933). The desire to avoid creating a black market for any product is an relevant to regulatory actions that involve prohibitions.

As a side note, members of Greenpeace broke into a research farm in Australia in 2011 and destroyed an entire crop of GM wheat. Members of another environmental damaged a crop of golden rice in the Philippines (Zhang et al. 2016 ).

To date, 156 Nobelists have signed the petition (Nobel Prize Winners 2016 ).

For a review of the GM food safety literature, also see Domingo ( 2016 ).

It is worth noting the long-term animal studies pose some scientific and technical challenges because most of the rodent species used in these types of experiments have a lifespan of about three years and normally develop tumors and other health problems as they age. So, it can be difficult to determine whether an adverse effect in a laboratory animal is due to an exposure to a GM food or the natural aging process. A two-year study published by Séralini et al. ( 2012 ) claiming that mice fed a diet of Roundup Ready GM corn had more tumors than mice fed the normal diet (the control group) was later retracted by the journal due to serious methodological flaws that undermined the validity of the data (Resnik 2015a ).

See Footnote 12.

Davidson ( 2001 ) defends a principle of charity for interpreting language. The basic idea here is that one should interpret a speaker’s statements as being rational, other things being equal. Interpreting disagreements about GM foods/crops as based on differing value priorities portrays these disagreements as rational, rather than based on irrational fear or ignorance.

It is also worth noting that bans on GM plants can create black markets because of the high demand for these products.

As of the writing of this book, Kenya is currently rethinking its ban on GM crops (Meeme 2019 ).

Most of the debate about chimeras so far has focused on inserting human cells into early animal embryos (or blastocysts), not on inserting human genes into animals.

It is also worth noting that a ban would probably create a black market because demand for GM animals and animal products it high.

There is a potential regulatory gap in the genetic engineering of animals for meat or animal products. Although regulations and ethical guidelines require IACUCs to review and oversee genetic engineering of animals for research conducted at academic institutions, there are no such requirements for genetic engineering of animals for non-research purposes, such as meat production. One could argue that companies that genetically engineer animals for non-research purposes should form ethics committees similar to IACUCs to oversee these activities.

Anderson led the research team that conducted the world’s first human gene therapy clinical trial. The experiment used an adenovirus vector to insert the adenosine deaminase gene into the T-cells of two young children with combined immunodeficiency. The trial showed that the procedure was safe and effective even if did not cure the patients (Blaese et al. 1995 ). In 2006, Anderson was convicted of molesting and sexually abusing a girl over a four-year period, beginning when she was 10 years old, and he served 12 years in prison. Anderson maintains that he is innocent and that his conviction was based on falsified evidence (Begley 2018 ).

See Footnote 29.

An example of somatic genetic enhancement would be a transferring a gene to an adult male to stimulate production of testosterone to enhance athletic and sexual performance.

It is worth noting that not everyone regards genetic enhancement immoral or morally questionable. The transhumanist movement embraces various forms of enhancement to benefit mankind and allow people to express creative freedom (Harris 2007 ; Bostrom 2008 , 2010 ; More and Vita-More 2013 ; Porter 2017 ; Rana and Samples 2019 ).

Some have attempted to define health in terms of a normal range of variation for an organism. In medicine, a normal physiological trait is a trait that falls within a range of variation for healthy functioning of the organism (Boorse 1977 ; Schaffner 1993 ). For example, normal fasting blood sugar levels range from 60 mg/dL to 100 mg/dL (WebMD 2020 ). Fasting blood sugar levels that are too high cause diabetes and levels that are too low cause hypoglycemia, both of which are unhealthy conditions. However, normality cannot be equated with the statistical norm for a population, since the statistical norm might be unhealthy. If most people in a population have a fasting blood sugar greater than 100 mg/dL, we would not say that a fasting blood sugar greater than 100 mg/dL is normal, even though it would be the statistical norm for that population. Thus, the concept of a normal range of variation cannot be defined statistically and depends on a broader concept of health, which may be influenced by moral, social, and cultural factors.

Some argue that “gene therapy” is a misleading term because it implies that the genetic interventions are likely to benefit the patient or human subject, when often they do not (Henderson et al. 2006 ).

See Resnik ( 2018a ) for discussion of additional safety protections for subjects enrolled in clinical research.

In 1996, the US Congress passed a ban, known as the Dickey-Wicker amendment, on the use of federal funds to create human embryos for research (Green 2001 ). Though the ban has been interpreted differently by different administrations, it is still in effect.

For further discussion of creating embryos for research, see Green ( 2001 ).

I will assume that parents who are willing to use medical technology to prevent the birth of children with genetic diseases view abortion as morally acceptable, at least for this purpose.

Prenatal genetic testing can also be used to avoid giving birth to children with chromosomal abnormalities, such as Trisomy 21 (Down Syndrome).

Embryos that are not implanted would be destroyed. I am assuming that parents would view this as morally acceptable.

See Resnik et al. ( 1999 ) and National Academies of Sciences, Engineering, and Medicine ( 2017 ) for additional examples of monogenic disorders that GGE might be used to prevent.

The concept of a parent can be confusing here, because people who related to the child genetically might not be related socially. The concept of a parent can be even more confusing when surrogate pregnancy is used to produce children, since woman who gestates and gives birth to the child might not be genetically related to the child, if she is carrying a fetus created by another couple in vitro.

This is one of the themes of the science fiction movie GATTACA.

This cost estimate is based on dividing the total cost of the Human Genome Project--$3 billion—by three. The Human Genome Project was a US-funded research project that took place from 1990 to 2003. Although sequencing the human genome was the primary goal of the project, it also included other activities, such as studies of human diseases, model organisms, genetic technologies, computational methods, and ethical issues (Human Genome Project 2020 ).

Interestingly, two of the scientists who called for the moratorium, David Baltimore and Paul Berg, participated in the Asilomar conference on recombinant DNA (discussed earlier).

These studies could include the creation of human embryos to study the safety and efficacy of GGE methods and techniques (Liang et al. 2015 ).

This is an example of the problem of incoherence discussed in Chapter 4 .

Alopecia areata is a condition that leads to hair loss. It is thought to have a genetic basis (McIntosh 2017 ).

The moratorium would not apply to GGE for research purposes.

The moratorium would not apply to research on embryos created by GGE, which would be necessary to obtain the knowledge needed to better understand the safety and efficacy of using GGE to produce children (Liang et al. 2015 ; Baltimore et al. 2015 ).

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Resnik, D.B. (2021). Genetic Engineering. In: Precautionary Reasoning in Environmental and Public Health Policy. The International Library of Bioethics, vol 86. Springer, Cham. https://doi.org/10.1007/978-3-030-70791-0_7

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