research paper on genome editing

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• Cold Spring Harb Perspect Biol
• v.8(12); 2016 Dec

Genome-Editing Technologies: Principles and Applications

1 Department of Bioengineering, University of California, Berkeley, California 94720

Shannon J. Sirk

2 Department of Chemical Engineering, Stanford University, Stanford, California 94305

Sai-lan Shui

3 Shanghai Institute for Advanced Immunochemical Studies, ShanghaiTech University, Shanghai, China

Targeted nucleases have provided researchers with the ability to manipulate virtually any genomic sequence, enabling the facile creation of isogenic cell lines and animal models for the study of human disease, and promoting exciting new possibilities for human gene therapy. Here we review three foundational technologies—clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9), transcription activator-like effector nucleases (TALENs), and zinc-finger nucleases (ZFNs). We discuss the engineering advances that facilitated their development and highlight several achievements in genome engineering that were made possible by these tools. We also consider artificial transcription factors, illustrating how this technology can complement targeted nucleases for synthetic biology and gene therapy.

Three technologies—CRISPR-Cas9, TALE nucleases, and zinc-finger nucleases—have facilitated a genome-editing revolution. But several challenges (e.g., effectively treating human diseases) remain.

In recent years, the emergence of highly versatile genome-editing technologies has provided investigators with the ability to rapidly and economically introduce sequence-specific modifications into the genomes of a broad spectrum of cell types and organisms. The core technologies now most commonly used to facilitate genome editing, shown in Figure 1 , are (1) clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9), (2) transcription activator-like effector nucleases (TALENs), (3) zinc-finger nucleases (ZFNs), and (4) homing endonucleases or meganucleases.

Genome-editing technologies. Cartoons illustrating the mechanisms of targeted nucleases. From top to bottom : homing endonucleases, zinc-finger nucleases (ZFNs), transcription activator-like effector (TALE) nucleases (TALENs), and clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9). Homing endonucleases generally cleave their DNA substrates as dimers, and do not have distinct binding and cleavage domains. ZFNs recognize target sites that consist of two zinc-finger binding sites that flank a 5- to 7-base pair (bp) spacer sequence recognized by the FokI cleavage domain. TALENs recognize target sites that consist of two TALE DNA-binding sites that flank a 12- to 20-bp spacer sequence recognized by the FokI cleavage domain. The Cas9 nuclease is targeted to DNA sequences complementary to the targeting sequence within the single guide RNA (gRNA) located immediately upstream of a compatible protospacer adjacent motif (PAM). DNA and protein are not drawn to scale.

In particular, the ease with which CRISPR-Cas9 and TALENs can be configured to recognize new genomic sequences has driven a revolution in genome editing that has accelerated scientific breakthroughs and discoveries in disciplines as diverse as synthetic biology, human gene therapy, disease modeling, drug discovery, neuroscience, and the agricultural sciences.

The diverse array of genetic outcomes made possible by these technologies is the result, in large part, of their ability to efficiently induce targeted DNA double-strand breaks (DSBs). These DNA breaks then drive activation of cellular DNA repair pathways and facilitate the introduction of site-specific genomic modifications ( Rouet et al. 1994 ; Choulika et al. 1995 ). This process is most often used to achieve gene knockout via random base insertions and/or deletions that can be introduced by nonhomologous end joining (NHEJ) (Fig. 2A) ( Bibikova et al. 2002 ). Alternatively, in the presence of a donor template with homology to the targeted chromosomal site, gene integration, or base correction via homology-directed repair (HDR) can occur (HDR) (Fig. 2B) (see Fig. 2 for an overview of other possible genome-editing outcomes) ( Bibikova et al. 2001 , 2003 ; Porteus and Baltimore 2003 ; Urnov et al. 2005 ). Indeed, the broad versatility of these genome-modifying enzymes is evidenced by the fact that they also serve as the foundation for artificial transcription factors, a class of tools capable of modulating the expression of nearly any gene within a genome.

Genome-editing outcomes. Targeted nucleases induce DNA double-strand breaks (DSBs) that are repaired by nonhomologous end joining (NHEJ) or, in the presence of donor template, homology-directed repair (HDR). ( A ) In the absence of a donor template, NHEJ introduces small base insertions or deletions that can result in gene disruption. When two DSBs are induced simultaneously, the intervening genomic sequence can be deleted or inverted. ( B ) In the presence of donor DNA (plasmid or single-stranded oligonucleotide), recombination between homologous DNA sequences present on the donor template and a specific chromosomal site can facilitate targeted integration. Lightning bolts indicate DSBs.

Here we review key principles of genome editing, emphasizing many of the engineering advances that have laid the groundwork for the creation, refinement, and implementation of the current suite of genome-modifying tools. We also provide an overview of the achievements made possible by genome editing, illustrating how this technology can enable advances throughout the life sciences.

TARGETED NUCLEASES

Zinc-finger nucleases.

ZFNs, which are fusions between a custom-designed Cys 2 -His 2 zinc-finger protein and the cleavage domain of the FokI restriction endonuclease ( Kim et al. 1996 ), were the first targeted nuclease to achieve widespread use ( Porteus and Carroll 2005 ; Urnov et al. 2010 ). ZFNs function as dimers, with each monomer recognizing a specific “half site” sequence—typically nine to 18 base pairs (bps) of DNA—via the zinc-finger DNA-binding domain ( Fig. 1 ). Dimerization of the ZFN proteins is mediated by the FokI cleavage domain, which cuts DNA within a five- to seven-bp spacer sequence that separates two flanking zinc-finger binding sites ( Smith et al. 2000 ). Each ZFN is typically composed of three or four zinc-finger domains, with each individual domain composed of ∼30 amino acid residues that are organized in a ββα motif ( Pavletich and Pabo 1991 ). The residues that facilitate DNA recognition are located within the α-helical domain and typically interact with three bps of DNA, with occasional overlap from an adjacent domain ( Wolfe et al. 2000 ). Using methods such as phage display ( Choo and Klug 1994 ; Jamieson et al. 1994 ; Wu et al. 1995 ), a large number of zinc-finger domains recognizing distinct DNA triplets have been identified ( Segal et al. 1999 ; Dreier et al. 2001 , 2005 ; Bae et al. 2003 ). These domains can be fused together in tandem using a canonical linker peptide ( Liu et al. 1997 ) to generate polydactyl zinc-finger proteins that can target a wide range of possible DNA sequences ( Beerli et al. 1998 , 2000a ; Kim et al. 2009 ). In addition to this “modular assembly” approach to zinc-finger construction, selection-based methods for constructing zinc-finger proteins have also been reported ( Greisman and Pabo 1997 ; Isalan et al. 2001 ; Hurt et al. 2003 ; Magnenat et al. 2004 ), including those that consider context-dependent interactions between adjacent zinc-finger domains, such as oligomerized pool engineering (OPEN) ( Maeder et al. 2008 ). In addition, specialized sets of validated two-finger, zinc-finger modules have been used to assemble zinc-finger arrays ( Kim et al. 2009 ; Bhakta et al. 2013 ), including those that take context-dependent effects into account ( Sander et al. 2011b ; Gupta et al. 2012 ).

One major concern associated with the use of ZFNs for genome editing (in addition to all targeted nucleases) is off-target mutations ( Gabriel et al. 2011 ; Pattanayak et al. 2011 ). As a result, several approaches have been undertaken to enhance their specificity. Among the most successful of these has been the creation of obligate heterodimeric ZFN architectures that rely on charge–charge repulsion to prevent unwanted homodimerization of the FokI cleavage domain ( Miller et al. 2007 ; Doyon et al. 2011 ), thereby minimizing the potential for ZFNs to dimerize at off-target sites. Additionally, protein-engineering methods have been used to enhance the cleavage efficiency of the FokI cleavage domain ( Guo et al. 2010 ). One particularly promising approach for improving ZFN specificity is to deliver them into cells as protein. Because of the intrinsic cell-penetrating activity of zinc-finger domains ( Gaj et al. 2014a ), ZFN proteins themselves are inherently cell-permeable and can facilitate gene editing with fewer off-target effects when applied directly onto cells as purified protein compared to when expressed within cells from nucleic acids ( Gaj et al. 2012 ). Modified ZFN proteins endowed with improved cell-penetrating activity have since been described ( Liu et al. 2015a ). ZFNickases can also facilitate gene correction in the absence of a DSB ( Kim et al. 2012 ; Ramirez et al. 2012 ; Wang et al. 2012 ). These enzymes, which consist of one catalytically inactivated ZFN monomer in combination with a second native ZFN monomer, can stimulate HDR by nicking or cleaving one strand of DNA and are derived from a concept first illustrated by Stoddard and colleagues using homing endonucleases ( McConnell Smith et al. 2009 ).

Unlike TALENs and CRISPR-Cas9, the difficulty associated with constructing zinc-finger arrays has hindered their widespread adoption in unspecialized laboratories. In particular, it remains challenging to create zinc-finger domains that can effectively recognize all DNA triplets, especially those of the 5′-CNN-3′ and 5′-TNN-3′ variety. As a result, ZFNs lack the target flexibility inherent to more recent genome-editing platforms. Nevertheless, the potential for ZFNs to mediate specific and efficient genome editing is evidenced by ongoing clinical trials based on ZFN-mediated knockout of the human immunodeficiency virus (HIV)-1 coreceptor CCR5 for treatment of HIV/acquired immune deficiency syndrome (AIDS) ( Tebas et al. 2014 ) and a planned clinical trial based on site-specific integration of the factor IX gene into the albumin locus to treat hemophilia B (Clinical Trial ID: {"type":"clinical-trial","attrs":{"text":"NCT02695160","term_id":"NCT02695160"}} NCT02695160 ) ( Sharma et al. 2015 ).

TALE Nucleases

TALE proteins are bacterial effectors. In 2009, the code used by TALE proteins to recognize DNA was uncovered ( Boch et al. 2009 ; Moscou and Bogdanove 2009 ). In a matter of months, this discovery enabled the creation of custom TALENs capable of modifying nearly any gene. Like ZFNs, TALENs are modular in form and function, comprised of an amino-terminal TALE DNA-binding domain fused to a carboxy-terminal FokI cleavage domain ( Christian et al. 2010 ; Miller et al. 2011 ). Also like ZFNs, dimerization of TALEN proteins is mediated by the FokI cleavage domain, which cuts within a 12- to 19-bp spacer sequence that separates each TALE binding site ( Fig. 1 ) ( Miller et al. 2011 ). TALEs are typically assembled to recognize between 12- to 20-bps of DNA, with more bases typically leading to higher genome-editing specificity ( Guilinger et al. 2014a ). The TALE-binding domain consists of a series of repeat domains, each ∼34 residues in length. Each repeat contacts DNA via the amino acid residues at positions 12 and 13, known as the repeat variable diresidues (RVDs) ( Boch et al. 2009 ; Moscou and Bogdanove 2009 ). Unlike zinc fingers, which recognize DNA triplets, each TALE repeat recognizes only a single bp, with little to no target site overlap from adjacent domains ( Deng et al. 2012 ; Mak et al. 2012 ). The most commonly used RVDs for assembling synthetic TALE arrays are: NI for adenine, HD for cytosine, NG for thymine, and NN or HN for guanine or adenine ( Boch et al. 2009 ; Moscou and Bogdanove 2009 ; Cong et al. 2012 ; Streubel et al. 2012 ). TALE DNA-binding domains can be constructed using a variety of methods, with the most straightforward approach being Golden Gate assembly ( Cermak et al. 2011 ). However, high-throughput TALE assembly methods have also been developed, including FLASH assembly ( Reyon et al. 2012 ), iterative capped assembly ( Briggs et al. 2012 ), and ligation independent cloning ( Schmid-Burgk et al. 2013 ), among others. More recent advances in TALEN assembly, though, have focused on the development of methods that can enhance their performance, including specificity profiling to uncover nonconventional RVDs that improve TALEN activity ( Guilinger et al. 2014a ; Yang et al. 2014 ; Juillerat et al. 2015 ; Miller et al. 2015 ), directed evolution as means to refine TALE specificity ( Hubbard et al. 2015 ), and even fusing TALE domains to homing endonuclease variants to generate chimeric nucleases with extended targeting specificity (discussed in more detail below) ( Boissel et al. 2014 ).

Compared to ZFNs, TALENs offer two distinct advantages for genome editing. First, no selection or directed evolution is necessary to engineer TALE arrays, dramatically reducing the amount of time and experience needed to assemble a functional nuclease. Second, TALENs have been reported to show improved specificity and reduced toxicity compared to some ZFNs ( Mussolino et al. 2014 ), potentially because of their increased affinity for target DNA ( Meckler et al. 2013 ) or perhaps a greater energetic penalty for associating with base mismatches. However, TALENs are substantially larger than ZFNs, and have a highly repetitive structure, making their efficient delivery into cells through the use of lentivirus ( Holkers et al. 2013 ) or a single adeno-associated virus (AAV) particle challenging. Methods for overcoming these limitations have emerged as TALENs can be readily delivered into cells as mRNA ( Mahiny et al. 2015 ; Mock et al. 2015 ) and even protein ( Cai et al. 2014 ; Liu et al. 2014a ), although alternative codon usage and amino acid degeneracy can also be leveraged to express RVD arrays that might be less susceptible to recombination ( Kim et al. 2013a ). In addition, adenoviral vectors have also proven particularly useful for mediating TALEN delivery to hard-to-transfect cell types ( Holkers et al. 2014 ; Maggio et al. 2016 ).

CRISPR-Cas9

The CRISPR-Cas9 system, which has a role in adaptive immunity in bacteria ( Horvath and Barrangou 2010 ; Marraffini and Sontheimer 2010 ), is the most recent addition to the genome-editing toolbox. In bacteria, the type-II CRISPR system provides protection against DNA from invading viruses and plasmids via RNA-guided DNA cleavage by Cas proteins ( Wiedenheft et al. 2012 ; Sorek et al. 2013 ). Short segments of foreign DNA are integrated within the CRISPR locus and transcribed into CRISPR RNA (crRNA), which then anneal to trans -activating crRNA (tracrRNA) to direct sequence-specific degradation of pathogenic DNA by the Cas9 protein ( Jinek et al. 2012 ). In 2012, Charpentier, Doudna, and co-workers reported that target recognition by the Cas9 protein only requires a seed sequence within the crRNA and a conserved protospacer-adjacent motif (PAM) upstream of the crRNA binding site ( Jinek et al. 2012 ). This system has since been simplified for genome engineering ( Cho et al. 2013 ; Cong et al. 2013 ; Jinek et al. 2013 ; Mali et al. 2013b ) and now consists of only the Cas9 nuclease and a single guide RNA (gRNA) containing the essential crRNA and tracrRNA elements ( Fig. 1 ). Because target site recognition is mediated entirely by the gRNA, CRISPR-Cas9 has emerged as the most flexible and user-friendly platform for genome editing, eliminating the need for engineering new proteins to recognize each new target site. The only major restriction for Cas9 target site recognition is that the PAM motif—which is recognized by the Cas9 nuclease and is essential for DNA cleavage—be located immediately downstream of the gRNA target site. The PAM sequence for the Streptococcus pyogenes Cas9, for example, is 5′-NGG-3′ (although in some cases 5′-NAG-3′ can be tolerated) ( Hsu et al. 2013 ; Jiang et al. 2013 ; Mali et al. 2013a ). Several studies have now shed light on the structural basis of DNA recognition by Cas9, revealing that the heteroduplex formed by the gRNA and its complementary strand of DNA is housed in a positively charged groove between the two nuclease domains (RuvC and HNH) within the Cas9 protein ( Nishimasu et al. 2014 ), and that PAM recognition is mediated by an arginine-rich motif present in Cas9 ( Anders et al. 2014 ). Doudna and colleagues have since proposed that DNA strand displacement induces a structural rearrangement within the Cas9 protein that directs the nontarget DNA strand into the RuvC active site, which then positions the HNH domain near target DNA ( Jiang et al. 2016 ), enabling Cas9-mediated cleavage of both DNA strands.

The Cas9 nuclease and its gRNA can be delivered into cells for genome editing on the same or separate plasmids, and numerous resources have been developed to facilitate target site selection and gRNA construction, including E-CRISP ( Heigwer et al. 2014 ), among others. Although Cas9 boasts the highest ease of use among the targeted nuclease platforms, several reports have indicated that it could be prone to inducing off-target mutations ( Cradick et al. 2013 ; Fu et al. 2013 ). To this end, considerable effort has been devoted to improving the specificity of this system, including using paired Cas9 nickases ( Mali et al. 2013a ; Ran et al. 2013 ), which increase gene-editing specificity by requiring the induction of two sequential and adjacent nicking events for DSB formation, or truncated gRNA that are more sensitive to mismatches at the genomic target site than a full-length gRNA ( Fu et al. 2014 ). Off-target cleavage has also been reduced by controlling the dosage of either the Cas9 protein or gRNA within the cell ( Hsu et al. 2013 ), or even by using Cas9 variants configured to enable conditional genome editing, such as a rapamycin-inducible split-Cas9 architecture ( Zetsche et al. 2015b ) or a Cas9 variant that contains a strategically placed small-molecule-responsive intein domain ( Davis et al. 2015 ). Nucleofection ( Kim et al. 2014 ) or transient transfection ( Zuris et al. 2015 ) of a preformed Cas9 ribonucleoprotein complex has also been shown to reduce off-target effects, enabling DNA-free gene editing in primary human T cells ( Schumann et al. 2015 ), embryonic stem cells ( Liu et al. 2015b ), Caenorhabditis elegans gonads ( Paix et al. 2015 ), mouse ( Menoret et al. 2015 ; Wang et al. 2015a ) and zebrafish embryos ( Sung et al. 2014 ), and even plant protoplasts ( Woo et al. 2015 ). The incorporation of specific chemical modifications known to protect RNA from nuclease degradation and stabilize secondary structure can further enhance Cas9 ribonucleoprotein activity ( Hendel et al. 2015 ; Rahdar et al. 2015 ). In a clever marriage of genome-editing platforms, the FokI cleavage domain has even been fused to an inactivated Cas9 variant to generate hybrid nucleases that require protein dimerization for DNA cleavage ( Guilinger et al. 2014b ; Tsai et al. 2014 ), theoretically increasing CRISPR-Cas9 specificity. Similarly, fusing Cas9 to DNA-binding domains has also proven effective at improving its specificity ( Bolukbasi et al. 2015 ). Finally, several studies have recently showed that protein engineering can broadly enhance Cas9 specificity ( Kleinstiver et al. 2016 ; Slaymaker et al. 2016 ) and even alter its PAM requirements ( Kleinstiver et al. 2015 ), the latter having the potential to enable creation of customized variants of Cas9 for allele-specific gene editing, although Cas9 orthologs ( Cong et al. 2013 ; Esvelt et al. 2013 ; Hou et al. 2013 ; Ran et al. 2015 ) or alternative CRISPR systems ( Zetsche et al. 2015a ) with unique PAM specificities have been uncovered in nature.

Homing Endonucleases

Homing endonucleases, also known as meganucleases, represent the final member of the targeted nuclease family. These enzymes have been reviewed at length elsewhere ( Silva et al. 2011 ; Stoddard 2014 ) but, briefly, members of the LAGLIDADG family of endonucleases—so named for the conserved amino acid motif present within these enzymes that interacts with DNA—are a collection of naturally occurring enzymes that recognize and cleave long DNA sequences (14–40 bps) ( Fig. 1 ). These enzymes make extensive sequence-specific contacts with their DNA substrate ( Stoddard 2011 ), and thus typically show exquisite specificity. However, unlike ZFNs and TALENs, the binding and cleavage domains in homing endonucleases are not modular. This overlap in form and function make their repurposing challenging, and limits their utility for more routine applications of genome editing. More recently megaTALs—fusions of a rare-cleaving homing endonuclease to a TALE-binding domain—have been reported to induce highly specific gene modifications ( Boissel et al. 2014 ; Lin et al. 2015a ). These enzymes have enabled integration of antitumor and anti-HIV factors into the human CCR5 gene in both primary T cells and hematopoietic stem/progenitor cells ( Sather et al. 2015 ), as well as disruption of endogenous T-cell receptor elements in T cells ( Osborn et al. 2016 ), indicating their potential for enabling and enhancing immunotherapies.

GENOME-EDITING APPLICATIONS

Engineering cell lines and organisms.

Before the emergence of engineered nucleases, genetically modifying mammalian cell lines was labor intensive, costly, and often times limited to laboratories with specialized expertise. However, with the advent of cost-effective and user-friendly gene-editing technologies, custom cell lines carrying nearly any genomic modification can now be generated in simply a matter of weeks. Examples of some of the outcomes that have become routine because of the emergence of targeted nucleases include gene knockout ( Santiago et al. 2008 ; Mali et al. 2013b ), gene deletion ( Lee et al. 2010 ), gene inversion ( Xiao et al. 2013 ), gene correction ( Urnov et al. 2005 ; Ran et al. 2013 ), gene addition ( Moehle et al. 2007 ; Hockemeyer et al. 2011 ; Hou et al. 2013 ), and even chromosomal translocation ( Fig. 2 ) ( Torres et al. 2014 ). In addition to cell line engineering, targeted nucleases have also expedited the generation of genetically modified organisms, facilitating the rapid creation of transgenic zebrafish ( Doyon et al. 2008 ; Sander et al. 2011a ; Hwang et al. 2013 ), mice ( Cui et al. 2011 ; Wang et al. 2013 ; Wu et al. 2013 ), rats ( Geurts et al. 2009 ; Tesson et al. 2011 ; Li et al. 2013 ), monkeys ( Liu et al. 2014c ), and livestock ( Hauschild et al. 2011 ; Carlson et al. 2012 ), which together have the capacity to accelerate human disease modeling and the discovery of new therapeutics.

Targeted nucleases have also emerged as powerful tools for plant engineering ( Baltes and Voytas 2015 ). Both TALENs and CRISPR-Cas9 have been used to modify multiple alleles within hexaploid bread wheat to confer heritable resistance to powdery mildew ( Wang et al. 2014b ). In another study, TALENs were used to knock out nonessential genes in the fatty acid metabolic pathway in soybean to generate a simplified plant cell with reduced metabolic components ( Haun et al. 2014 ). Of special note, two recent reports showed that purified nuclease proteins can be introduced directly into plant protoplasts, enabling the introduction of germline-transmissible modifications that are virtually indistinguishable from naturally occurring ( Luo et al. 2015 ; Woo et al. 2015 ). This technical advance could help to overcome certain regulatory hurdles associated with the use of transgenic crops. Finally, targeted nucleases have also been used to inactivate pathogenic genes to prevent viral ( Lin et al. 2014 ) or parasitic ( Ghorbal et al. 2014 ) infection, as well as to introduce knockin-specific factors capable of imparting pathogen resistance ( Wu et al. 2015 ).

Intriguingly, targeted nucleases could also serve as conduits for curbing mosquito- or insect-borne diseases through a technique known as gene drive ( Burt 2003 ; Sinkins and Gould 2006 ), which harnesses genome editing to facilitate the introduction of a specific gene or mutation that can then confer a particular phenotype into a host and also be transmitted to its progeny ( Windbichler et al. 2011 ). Gene drives have now been tested in the malaria vector mosquitos Anopheles stephensi ( Gantz et al. 2015 ) and Anopheles gambiae ( Hammond et al. 2016 ) as a means for achieving population control and to prevent disease transmission, respectively. However, owing to the ease with which CRISPR-Cas9 can be programmed ( Gantz and Bier 2015 ), debate has ignited on the potential societal and environmental impact of this technology ( Esvelt et al. 2014 ; Akbari et al. 2015 ), spurring the development of safeguard elements that could help to minimize the risk of gene-edited organisms escaping from the laboratory ( DiCarlo et al. 2015 ).

Synthetic Biology and Genome-Scale Engineering

Targeted nucleases also offer a facile means for generating modified bacterial and yeast strains for synthetic biology, including metabolic pathway engineering. Bacterial species of the order Actinomycetales , for instance, are one of the most important sources of industrially relevant secondary metabolites. However, many Actinomycetales species are recalcitrant to genetic manipulation, a fact that has severely hampered their use for metabolic engineering. CRISPR-Cas9 has been used to inactivate multiple genes in actinomycetes ( Tong et al. 2015 ), indicating its ability to enable the creation of designer bacterial strains with enhanced metabolite production capabilities. CRISPR has also facilitated multiplexed metabolic pathway engineering in yeast at high efficiencies ( Jakociunas et al. 2015a , b ), as well as random mutagenesis of yeast chromosomal DNA for phenotypic screening of desired mutants ( Ryan et al. 2014 ). Indeed, genome-wide CRISPR-based knockout screens hold tremendous potential for functional genomics ( Hilton and Gersbach 2015 ), having facilitated the discovery of genomic loci that confer drug resistance to cells ( Koike-Yusa et al. 2014 ; Shalem et al. 2014 ; Wang et al. 2014a ; Zhou et al. 2014 ), uncovered how cells can control induction of the host immune response ( Parnas et al. 2015 ), provided new insights into the genetic basis of cellular fitness ( Hart et al. 2015 ; Wang et al. 2015b ), and even shed light on how certain viruses induce cell death ( Ma et al. 2015 ). Genome-wide CRISPR screens can also facilitate the discovery of functional noncoding elements ( Kim et al. 2013b ; Korkmaz et al. 2016 ), and provide a means for studying the structure and evolution of the human genome. In a remarkable example of the latter, Shendure and colleagues used Cas9 to mediate integration of short randomized DNA sequences into the BRCA1 and DBR1 genes ( Findlay et al. 2014 ). They then measured the functional consequences of these mutations on fitness, achieving an unprecedented look at some of the factors driving genome and disease evolution. Finally, CRISPR screens have even proven effective in vivo, enabling the identification of factors involved in zebrafish development ( Shah et al. 2015 ) and disease progression in mice ( Chen et al. 2015 ).

Therapeutic Genome Editing

Genome editing itself also holds tremendous potential for treating the underlying genetic causes of certain diseases ( Cox et al. 2015 ; Porteus 2015 ; Maeder and Gersbach 2016 ). In one of the most successful examples of this to date, ZFN-mediated disruption of the HIV coreceptor CCR5 was used to engineer HIV resistance into both CD4 + T cells ( Perez et al. 2008 ) and CD34 + hematopoietic stem/progenitor cells (HSPCs) ( Holt et al. 2010 ), proving safe and well-tolerated in a phase I clinical trial that infused these gene-modified T cells into individuals with HIV/AIDS ( Tebas et al. 2014 ). In addition to enabling the introduction of gene modification that can enhance autologous cell therapies, targeted nucleases can also be combined with viral vectors—including AAV—to mediate genome editing in situ ( Gaj et al. 2016 ). For instance, delivery of an AAV vector encoding a ZFN pair designed to target a defective copy of the factor IX gene, along with its repair template, led to efficient gene correction in mouse liver, increasing factor IX protein production in both neonatal ( Li et al. 2011 ) and adult ( Anguela et al. 2013 ) models of the disease. In vivo genome editing also recently enabled the restoration of dystrophin gene expression and the rescue of muscle function in mouse models of Duchenne muscular dystrophy ( Long et al. 2015 ; Nelson et al. 2015 ; Tabebordbar et al. 2015 ). Therapeutic gene editing in a mouse model of human hereditary tyrosinemia has also been reported using both hydrodynamic injection of plasmid DNA encoding CRISPR-Cas9 ( Yin et al. 2014 ) and by combining nanoparticle-mediated delivery of Cas9-encoding mRNA with AAV-mediated delivery of the DNA template for gene correction ( Yin et al. 2016 ). More recently, a dual particle AAV system, wherein one AAV vector carried the Cas9 nuclease and a second harbored the gRNA and donor repair template, was able to mediate correction of a disease-causing mutation in the ornithine transcarbamylase gene in the liver of a neonatal model of the disease ( Yang et al. 2016 ). This work, in particular, showed that therapeutic levels of gene correction could be achieved in a regenerating tissue even when using multiple AAV particles. Although highly promising, numerous hurdles still need to be overcome for in vivo applications of genome editing to reach its full potential. Chief among these are methods that can facilitate nuclease delivery or expression to only diseased cells or tissues, and the development of new strategies that can enhance HDR in disease-associated postmitotic cells in vivo.

TARGETED TRANSCRIPTION FACTORS

Tools for modulating gene expression.

The modular qualities of zinc-finger and TALE proteins, in addition to the highly flexible DNA recognition ability of CRISPR-Cas9, also provide investigators with the ability to modulate the expression of nearly any gene from its promoter or enhancer sequences via their fusion to transcriptional activator and repressor protein domains. Among the first fully synthetic transcriptional effector proteins to be generated ( Beerli et al. 1998 ) were those based on the fusion of engineered zinc-finger proteins with either the Herpes simplex–derived transactivation domain ( Sadowski et al. 1988 ) or the Krüppel-associated box (KRAB) repression protein ( Margolin et al. 1994 ). Over the course of the next 15 years, zinc-finger-based transcriptional modulators were expanded and featured several other types of effector domains ( Beerli and Barbas 2002 ), including, for example, the Dnmt3a methyltransferase domain ( Rivenbark et al. 2012 ; Siddique et al. 2013 ) and the ten-eleven translocation methylcytosine dioxygenase 1 (TET1) ( Chen et al. 2014 ), which can modulate transcription via targeted methylation or demethylation, respectively. As a natural extension of zinc-finger transcription factors, and further drawing on the parallels with zinc-finger proteins, TALE transcription factors have also emerged as an especially effective platform for achieving targeted transcriptional modulation ( Miller et al. 2011 ; Zhang et al. 2011 ). Effector domains are generally fused to the carboxyl terminus of the synthetic TALE array and, contrary to the longer sequence typically required for efficient modulation by zinc-finger transcription factors, TALEs have been reported to regulate gene expression with as few as 10.5 repeats ( Boch et al. 2009 ). Like zinc fingers, TALEs are also compatible with numerous epigenetic modifiers, including the TET1 hydroxylase catalytic domain ( Maeder et al. 2013b ) and the lysine-specific histone demethylase 1 (LSD1) ( Mendenhall et al. 2013 ) domains, which have been used for targeted CpG demethylation and histone demethylation, respectively. In particular, the ease with which a large number of TALEs can be constructed has enabled the discovery that tiling a promoter sequence with combinations of synthetic transcription factors can lead to a synergistic increase in gene expression ( Maeder et al. 2013b ; Perez-Pinera et al. 2013 ). And, like zinc fingers ( Beerli et al. 2000b ; Pollock et al. 2002 ; Magnenat et al. 2008 ; Polstein and Gersbach 2012 ), TALE activators have also been successfully engineered to regulate gene expression in response to external ( Mercer et al. 2014 ) or endogenous ( Li et al. 2012 ) chemical stimuli, optical signals ( Konermann et al. 2013 ), and even proteolytic cues ( Copeland et al. 2016 ; Lonzaric et al. 2016 ).

Because of the exquisite ease with which it can be programmed, the CRISPR-Cas9 system has also been adapted for transcriptional modulation through fusion of specific effector domains to a catalytically inactivated variant of the Cas9 protein ( Qi et al. 2013 ). Deactivation is achieved by introducing two amino acid substitutions, D10A and H840A, into the RuvC and NHN endonuclease domains of Cas9, respectively. Although unable to cleave DNA, this mutant, referred to as dCas9, retains its ability to bind DNA in an RNA-directed manner. Effector domains are fused to the carboxyl terminus of the dCas9 protein and can modulate gene expression from either strand of the targeted DNA sequence ( Farzadfard et al. 2013 ; Maeder et al. 2013a ; Perez-Pinera et al. 2013 ). Genome-scale activation studies have indicated that the most robust levels of activation are generally observed when dCas9 activators are targeted to -400 to -50 bp upstream from the transcriptional start site ( Gilbert et al. 2014 ; Hu et al. 2014 ). Additionally, dCas9 can inhibit gene expression by simply blocking transcriptional initiation or elongation through a process known as CRISPR interference ( Qi et al. 2013 ), although fusing dCas9 to transcriptional repressor domains can also lead to efficient silencing from the promoter ( Gilbert et al. 2013 ; Zalatan et al. 2015 ). Much like zinc fingers and TALEs, methods for achieving conditional gene modulation using dCas9 have also been reported, including the fusion of a dihydrofolate reductase destabilization domain to dCas9, which can provide chemical control over activation, enabling cellular reprogramming or differentiation ( Balboa et al. 2015 ). Light-inducible dCas9-based systems capable of providing optical control of gene expression provide another means for achieving conditional control of gene expression ( Nihongaki et al. 2015 ; Polstein and Gersbach 2015 ).

Although flexible, first-generation dCas9 activators were routinely found to display suboptimal levels of activation. As a result, the development of second-generation CRISPR activators quickly emerged as a highly active area of research. One particularly elegant approach for overcoming the low activation thresholds inherent within first-generation systems was by strategically inserting an RNA aptamer within a functionally inert region of the gRNA. This aptamer recruits specific activation helper proteins that work in concert with a dCas9 activator to enhance transcription ( Konermann et al. 2015 ; Zalatan et al. 2015 ). Other strategies based on directly fusing additional helper activation domains to dCas9 have also been shown to enhance transcription ( Chavez et al. 2015 ). Targeted acetylation of histone proteins within a promoter or enhancer sequence via epigenome editing using the catalytic core of the human acetyltransferase p300 fused to dCas9 can also lead to robust levels of gene activation ( Hilton et al. 2015 ). Similarly, dCas9 repressor proteins targeted to distal regulatory elements have been found to facilitate chromatin remodeling and gene repression via epigenomic modification ( Thakore et al. 2015 ). Finally, by simply reducing the length of the gRNA, catalytically active variants of Cas9 can stimulate transcription without inducing DNA breaks ( Dahlman et al. 2015 ; Kiani et al. 2015 ), enabling orthogonal gene knockout and activation with the same Cas9 variant in a single cell.

Applications of Targeted Transcriptional Regulation

Early work on the use of engineered zinc-finger transcription factors revealed that synthetic transcriptional modulators are effective tools for a broad range of applications, enabling such tasks as inhibiting viral replication ( Papworth et al. 2003 ; Reynolds et al. 2003 ; Segal et al. 2004 ; Eberhardy et al. 2006 ), modulating the expression of disease-associated loci ( Graslund et al. 2005 ; Wilber et al. 2010 ), inducing angiogenesis for accelerated wound healing ( Rebar et al. 2002 ), and genomic screening of cellular targets for cancer progression and drug resistance ( Park et al. 2003 ; Blancafort et al. 2005 , 2008 ). Facilitated by many of the insights gained from zinc-finger transcription factor technology, both TALEs and CRISPR-Cas9 have now further expanded the possibilities of engineered transcriptional activators and repressors. For example, TALEs and CRISPR-Cas9 have enabled rapid construction of custom genetic circuits and logic gates ( Gaber et al. 2014 ; Lebar et al. 2014 ; Liu et al. 2014b ), complex gene regulation networks ( Nielsen and Voigt 2014 ; Nissim et al. 2014 ), and even facilitated cellular reprogramming ( Gao et al. 2013 ) and the differentiation of mouse embryonic fibroblasts to skeletal myocytes ( Chakraborty et al. 2014 ). dCas9 transcriptional effectors have even been used to efficiently mediate repression and activation of endogenous genes in Drosophila ( Lin et al. 2015b ) and in plant cells ( Piatek et al. 2015 ). Both TALE and Cas9 activators have also been configured to stimulate transcription of latent HIV ( Zhang et al. 2015 ; Ji et al. 2016 ; Limsirichai et al. 2016 ; Perdigao et al. 2016 ; Saayman et al. 2016 ), indicating their potential to work in concert with antiretroviral therapy for eradicating HIV infection. Importantly, because of the ease with which the CRISPR-Cas9 system can be used, genome-wide screens using Cas9 transcriptional activators ( Gilbert et al. 2014 ; Konermann et al. 2015 ) and repressors ( Gilbert et al. 2014 ) can be easily implemented to discover genes involved in a number of diverse processes, including drug resistance and cancer metastasis. In particular, CRISPR-based genome-scale screening methods have the potential to overcome many of the technical hurdles associated with other contemporary screening technologies, such as cDNA libraries and RNAi, indicating its potential for facilitating drug discovery and basic biological research.

CONCLUSIONS

Despite the successes already achieved, many challenges remain before the full potential of genome editing can be realized. First and foremost are the development of new tools capable of introducing genomic modifications in the absence of DNA breaks. Targeted recombinases ( Akopian et al. 2003 ; Mercer et al. 2012 ), which can be programmed to recognize specific DNA sequences ( Gaj et al. 2013 ; Sirk et al. 2014 ; Wallen et al. 2015 ) and even integrate therapeutic factors into the human genome ( Gaj et al. 2014b ), are one such option. More recent work has indicated that single-base editing without DNA breaks can be achieved using an engineered Cas9 nickase complex ( Komor et al. 2016 ), although it remains unknown how effective this technology is in therapeutically relevant settings. By linking genomic modifications induced by targeted nucleases to their own self-degradation, self-inactivating vectors are also poised to improve the specificity of genome editing, especially because the frequency of off-target modifications can be directly proportional to the duration of cellular exposure to a nuclease ( Pruett-Miller et al. 2009 ). In addition, much of the knowledge behind genome engineering has been obtained in immortalized cell lines. However, in the case of regenerative medicine, it is highly desirable to genetically manipulate progenitor or stem-cell populations, both of which can differ markedly from transformed cell lines with respect to their epigenome or three-dimensional organization of their genomic DNA. These differences can have profound effects on the ability of genome-editing tools to either modify a specific sequence or regulate gene expression. Although the union between genome engineering and regenerative medicine is still in its infancy, realizing the full potential of these technologies in stem/progenitor cells requires that their functional landscape be fully explored in these genetic backgrounds. Only then will genome editing technologies truly be able to reprogram cell fate and behavior for the next generation of advances in synthetic biology and gene therapy.

ACKNOWLEDGMENTS

We gratefully acknowledge the support and mentorship of the late Carlos F. Barbas, III (1964–2014). This work is supported by the National Institutes of Health (F32GM113446 to T.G.) and ShanghaiTech University (to J.L.). We apologize to those whose important contributions were not cited because of space constraints.

Editors: Daniel G. Gibson, Clyde A. Hutchison III, Hamilton O. Smith, and J. Craig Venter

Additional Perspectives on Synthetic Biology available at www.cshperspectives.org

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• 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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• CRISPR-Cas system
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Recent advances in CRISPR-based genome editing technology and its applications in cardiovascular research

• Zhen-Hua Li 1   na1 ,
• Jun Wang 1   na1 ,
• Jing-Ping Xu 1 , 2 ,
• Jian Wang 1 &
• Xiao Yang 1  

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The rapid development of genome editing technology has brought major breakthroughs in the fields of life science and medicine. In recent years, the clustered regularly interspaced short palindromic repeats (CRISPR)-based genome editing toolbox has been greatly expanded, not only with emerging CRISPR-associated protein (Cas) nucleases, but also novel applications through combination with diverse effectors. Recently, transposon-associated programmable RNA-guided genome editing systems have been uncovered, adding myriads of potential new tools to the genome editing toolbox. CRISPR-based genome editing technology has also revolutionized cardiovascular research. Here we first summarize the advances involving newly identified Cas orthologs, engineered variants and novel genome editing systems, and then discuss the applications of the CRISPR-Cas systems in precise genome editing, such as base editing and prime editing. We also highlight recent progress in cardiovascular research using CRISPR-based genome editing technologies, including the generation of genetically modified in vitro and animal models of cardiovascular diseases (CVD) as well as the applications in treating different types of CVD. Finally, the current limitations and future prospects of genome editing technologies are discussed.

Genome editing technology refers to a series of technologies capable of manipulating cellular DNA sequences at desired genomic sites by generating altered DNA sequences through nuclease-mediated site-specific DNA breaks that are resolved through DNA repair pathways [ 1 , 2 , 3 ]. Among genome editing-associated nucleases, clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein (Cas) nucleases are convenient, efficient, and precise, and are currently the most widely used [ 4 , 5 , 6 , 7 , 8 ]. After the CRISPR-Cas9 system was characterized and programmed to perform RNA-guided DNA cleavage at specific sites in prokaryotes, it was immediately proven to be an efficient tool for editing eukaryotic genomes [ 9 ]. Since then, CRISPR-based genome editing technology has drawn a worldwide attention and initiated extensive development. Emerging CRISPR-based tools with broadened targeting ranges, improved editing specificity and efficiency, and other distinct capabilities have facilitated eukaryotic genome editing by selecting optimal CRISPR-Cas tools. In addition to expanding the CRISPR-Cas nuclease arsenal, this system has also been applied to transcriptional regulation, epigenetic modification, and live-cell imaging by incorporation with other effector proteins [ 7 , 10 , 11 ].

The exponential development of genome editing technology has dramatically changed the landscape of biological and medical research, heralding a new era of precision medicine based on genome editing [ 12 ]. CRISPR-based nucleases are able to cut target DNA and generate double-strand breaks (DSBs), followed by the introduction of random mutations mediated by non-homologous end-joining (NHEJ), or make precise editing through homology-directed repair (HDR) [ 13 ]. The therapeutic potential of CRISPR-based tools has been investigated using the mouse models of various human diseases [ 7 , 14 ]. However, precise gene correction for in vivo therapeutic utility remains challenging, which is partially due to the low efficiency of HDR-mediated DNA replacement. This strategy is usually not applicable to post-mitotic cells, as HDR occurs mainly in the S/G 2 phase during cell division [ 15 ]. Precise genome editing tools have been developed and continuously optimized by fusing activity-impaired Cas nucleases with deaminases, called base editors, or with reverse transcriptases, called prime editors (PEs) [ 16 , 17 , 18 ]. Despite not achieving the goal of arbitrarily introducing any genetic substitutions at any targeted genomic site, we are now closer to this aspiration than ever.

Cardiovascular disease (CVD) is a group of disorders of the heart and blood vessels that has consistently been ranked as the leading threat to human health worldwide. Many gene mutations have been linked to CVD, and the number is still increasing [ 19 , 20 ]. Loss-of-function studies in animals are required to address the causal relationship between these mutations and cardiovascular pathologies. With the help of the CRISPR-based toolbox, creating animal models of human diseases has become much easier, faster, and more flexible than ever before; these models will greatly advance our understanding of cardiovascular pathogenesis and the development of therapeutic strategies [ 14 ]. Furthermore, CRISPR-based genome editing technology holds promise for treating inherited CVD caused by rare mutations.

In this review, we highlight the recent advances in CRISPR-based genome editing technology, mainly in the past three years, and discuss the tremendous innovation this epoch-making technology has brought to the field of cardiovascular research.

Novel Cas orthologs and engineered variants

Natural CRISPR-Cas systems are originally identified as adaptive immune systems in bacteria and archaea, and can be divided into two classes based on their composition and mechanisms. These systems are further divided into six types (I–VI) and dozens of subtypes based on the characteristics and accessory genes flanking the CRISPR array [ 21 ]. The most widely used class 2 CRISPRs are characterized by their single effector proteins, including type II Cas9 and type V-A Cas12a. Although class 2 natural Cas nucleases have long been used for efficient genome editing, their applications are limited because of the requirement of specific protospacer adjacent motif (PAM) sequences, off-target DNA cleavage, and occasionally, large sizes. Class 1 CRISPR systems possess multiple effector molecules that have unique features, such as distinct PAM preferences, higher on-target specificity through longer target recognition, and production of long-range genomic deletion [ 22 , 23 , 24 , 25 ]. However, the requirement of multiple effectors and the relatively low editing efficiency must be improved before their widespread application. Continuous efforts have been made to characterize novel Cas orthologs and engineered Cas variants to improve genome editing efficiency and broaden compatibility (Fig.  1 ).

Characteristics of novel Cas orthologs and engineered variants. a Representative type I Cas orthologs capable of large-range deletions. b Representative Cas orthologs of miniature sizes. c Engineered Cas variants with diverse protospacer adjacent motif recognition capabilities. d Structure-guided strategies for improving DNA specificity without affecting the on-target cleavage efficiency

Characterizing novel Cas orthologs with distinctive features

The class 1 type I CRISPR system is the most prevalent CRISPR system, in which the multi-subunit CRISPR-associated complex for antiviral defense (Cascade) identifies DNA targets, and the helicase-nuclease enzyme Cas3 degrades DNA [ 26 ] (Fig.  1 a). Several type I CRISPR systems have been characterized and applied to mammalian genome editing. Type I-E and type I-D systems have been used to induce unidirectional and bidirectional long-range deletions in human cells [ 22 , 23 , 24 ]. Recently, supplying Cas11 was shown to enable divergent I-C, I-D, and I-B CRISPR-Cas3 editors for eukaryotic applications, and efficiently produced large unidirectional deletions [ 25 ]. Therefore, type I CRISPR systems can greatly expand the genome editing toolbox owing to their unique mechanisms and advantages in deleting full-length genes, gene clusters, and non-coding sequences.

Recently, the IS200/605 transposon family encoded RNA-guided nucleases have been identified as ancestors of CRISPR-Cas nucleases [ 27 , 28 ]. Cas9 endonucleases could likely have evolved from ancestral IscB proteins, whereas Cas12 endonucleases descended from TnpB proteins [ 27 ]. These transposon-encoded nucleases, together with the IsrB proteins, which are shorter IscB homologs also encoded in IS200/605 superfamily transposons, are called the obligate mobile element-guided activity (OMEGA) system [ 27 ]. IscB and TnpB are guided by non-coding RNAs called ωRNAs, which are derived from the left- or right-end elements of a transposon and combine the functions of CRISPR RNA (crRNA) and trans-activating CRISPR RNA (tracrRNA) [ 27 , 28 , 29 ]. Being only two-fifths the size of Cas9, IscB and TnpB can mediate double-strand DNA cleavage at the target sites with a 3′ or 5′ transposon-associated motif (TAM), and both have been adopted for genome editing in human cells [ 27 , 28 ].

The in vivo application of most CRISPR systems is challenging because of their large size, especially when delivered by the widely used adeno-associated virus (AAV). As a result, the exploitation of miniature Cas proteins with high efficiency is in sustained demand (Fig.  1 b). For instance, SaCas9 (1053 aa), CjCas9 (984 aa), and Nme1Cas9 (1082 aa) have been validated as mammalian genomic editors [ 30 , 31 , 32 ]. Recently, a compact Nme2Cas9 (1082 aa) recognizing an N 4 CC PAM was described with an identical target density as SpCas9 and few off-target effects [ 33 ]. In addition to the Cas9 orthologs mentioned above, several Cas12 nucleases with smaller sizes, such as Cas12e (or CasX, 986 aa), and Cas12f (400–700 aa), have also been identified as genome editing tools in mammalian cells [ 34 , 35 , 36 , 37 ]. The Un1Cas12f1 (522 aa) system was optimized to enable efficient genome editing in human cells [ 35 ]. Notably, with only a size of 422 aa, AsCas12f1 is currently the smallest RNA-guided Cas nuclease, and has been shown to be an effective programmed genome editing tool in both bacterial and human cells [ 36 ]. With approximately equal to or less than half the size of the widely used SpCas9, these miniature CRISPR tools facilitate the AAV-mediated all-in-one delivery of CRISPR components or catalytically inactive Cas variants fused with other functional proteins.

Very recently, CRISPR-Cas systems were found to be widely encoded in the genomes of diverse bacteriophages, where they are involved in competition with other viruses [ 38 , 39 ]. The bacteriophage-encoded Cas proteins contain all known types of CRISPR-Cas systems, but have phage-specific properties. These Cas proteins, such as CasΦ [ 38 ] and Casλ [ 39 ], tend to have remarkably small sizes due to the compact viral genome. These hypercompact systems have been shown to edit the genomes of human and plant cells, indicating that viral Cas nucleases could serve as a new source of genome editing tools.

Expanding the range of genomic targets

Genomic targeting by Cas nucleases requires a PAM sequence near the site where the Cas nuclease binds DNA sequences complementary to the single guide RNA (sgRNA). The requirement of PAM is the gatekeeper for CRISPR-Cas mediated genome targeting, as whether a genomic sequence possesses a PAM for a certain Cas nuclease determines whether the site can be targeted and edited by CRISPR. Efforts have been made to develop Cas nucleases with broader PAM compatibility to pursue true PAM-free nucleases. However, PAM-free nucleases might have potential drawbacks, such as self-targeting of gRNA-expressing DNA constructs and reduced efficiency as more time is required for interrogating the whole genome. Therefore, it is better to develop an arsenal of divergent PAM-dependent Cas nucleases that collectively cover all genomic sequences [ 40 ] (Fig.  1 c).

Using phage-assisted non-continuous and continuous evolution strategies, three new SpCas9 variants (SpCas9-NRRH, SpCas9-NRCH, and SpCas9-NRTH) were characterized to recognize most NR PAM sequences, together with SpCas9-NG (N = A/T/C/G, R = A/G, H = A/C/T) [ 41 ]. To relax the PAM preference of SpCas9, two SpCas9 variants, SpG (targeting NGN) and SpRY (targeting NYN), have been generated by structure-guided substitutions in several residues, making most of the genome targetable (Y = C/T) [ 42 ]. Structure-motivated engineering has also been used to expand targeting range of LbCas12a and AsCas12a [ 43 , 44 ]. Chimeric Cas proteins created by exchanging PAM-interacting domains between naturally occurring Cas orthologs have also been applied to expand PAM recognition. Substituting the loop sequence of Cas9 from Streptococcus anginosus , together with the T1227K mutation, into the open reading frame (ORF) of ScCas9 generates ScCas9++ with NNG PAM compatibility [ 45 ]. A similar strategy was used to generate a variant iSpyMac by grafting the PAM-interacting domain of SmacCas9 into SpyCas9, which recognizes all adenine dinucleotide PAM (NAAN PAM) sequences [ 46 ]. The chimera generation approach has also been applied to replace PAM-interacting domains in SaCas9 [ 47 ] and Cas12a [ 48 ].

Improving the DNA specificity without affecting on-target cleavage efficiency

Off-target activity is a major challenge when using CRISPR tools for disease-related gene therapy. Over the years, continuous efforts have been made to construct high-fidelity Cas9 variants including eSpCas9(1.1) [ 49 ], SpCas9-HF1 [ 50 ], HypaCas9 [ 51 ], evoCas9 [ 52 ], HiFi Cas9 [ 53 ] and Sniper-Cas9 [ 54 ]. In addition, high-fidelity SaCas9s have been identified either by rational engineering [ 55 ] or directional screening [ 56 ]. The achievement of enhanced discrimination between on-target and off-target binding of these variants relies mainly on the energetic destabilization of the Cas9:sgRNA:DNA complex at off-target sites [ 57 ]. However, the improvement of these high-fidelity Cas9 variants seems to occur at the cost of decreased on-target efficiency [ 58 , 59 ]. Recently, kinetics-guided cryo-electron microscopy was used to show that mismatches distal to PAM can be stabilized by a loop in the RuvC domain, allowing Cas9 activation [ 60 ]. Based on this observation, they designed a high-fidelity variant with mutations in the RuvC domain, named SuperFi-Cas9, which displayed significantly improved mismatch discrimination without compromising on-target DNA cleavage efficiency [ 60 ].

DpbCas12e has been validated as a naturally occurring high-fidelity Cas nuclease with striking avoidance of off-target activity [ 61 ]. In a recent cryo-EM-based structural engineering study, unique nucleotide-binding loops within Cas12e were found to be important for DNA cleavage efficacy. Based on this finding, newly designed chimeric Cas12e proteins (DpbCasX-R3 and PlmCasX-R1) and sgRNA (sgRNAv2) exhibited substantially improved DNA editing efficiency in mammalian cells [ 62 ]. Structure-guided protein engineering has also been used to improve the performance of AsCas12a [ 43 ]. E174R/S542R/K548R substitutions were introduced into AsCas12a to construct a variant called enAsCas12a that possesses an expanded targeting range and increased cleavage activity. A high-fidelity version of enAsCas12a (enAsCas12a-HF1) with an additional N282A substitution was also engineered to reduce off-target effects [ 43 ] (Fig.  1 d).

Advances in precise genome editing

Precise genome editing is essential for preclinical research and clinical gene therapy, and HDR-mediated gene editing has long been the only option. Efforts have been made to improve HDR efficiency, such as the use of rationally designed single-stranded oligodeoxynucleotide (ssODN) templates instead of double-stranded DNA [ 63 ] or the addition of NHEJ chemical inhibitors [ 64 ]. The delivery of Cas9 and HDR templates by AAVs has accomplished precise genome editing in post-mitotic hippocampal neurons and cardiomyocytes in mice [ 65 , 66 , 67 ]. However, the efficiency of HDR-mediated editing is still relatively low compared to that of the predominant NHEJ repair pathway, and DSBs made by this conventional method may introduce undesired damage to the genome [ 68 , 69 , 70 ]. Its clinical application is hampered by the need for additional DNA templates and HDR-promoting chemical agents with potential cytotoxicity, such as SCR7 [ 64 ], azidothymidine, trifluridine [ 71 ], NU7026, and NU7441 [ 72 ]. Motivated by these problems, novel precise genome editing tools that do not require DSBs or exogenous DNA templates have been developed (Fig.  2 ).

CRISPR-Cas-based DNA base editing tools. a-c Schematic diagrams of CBE ( a ), ABE ( b ), and CGBE ( c ). d Schematic of PE. UGI uracil-DNA glycosylase inhibitor, AID activation-induced cytidine deaminase, UNG uracil-DNA glycosylase, TadA deoxyadenosine deaminases, RT reverse transcriptase, PBS primer binding site, RTT RT template, CBE cytosine base editor, ABE adenine base editor, CGBE C-to-G base editor, PE Prime editor

Base editors

Base editors can make precise base substitutions without requiring DSBs or donor DNA templates, and are independent of HDR, providing a promising therapeutic tool for human genetic diseases in which the most relevant variants are single nucleotide mutations [ 73 , 74 ]. Current base editors are constructed by fusing DNA deaminase enzymes to catalytically impaired Cas nucleases, which can precisely change a single base in a targeted sequence [ 75 ]. Base editing jointly harnesses the genome-targeting function of a Cas protein and the DNA base modification role of a deaminase, and sometimes additional regulatory elements are also required to achieve the desired performance. To date, base editors can be used not only for precise genome editing at specific single loci, but also for large-scale functional screening of genetic variants or key amino acid residues [ 76 , 77 , 78 ].

The cytosine base editor (CBE) and adenine base editor (ABE) are widely used base editors that enable the editing of all four types of base transitions (C-to-T, A-to-G, T-to-C, and G-to-A) [ 16 , 17 ] (Fig.  2 a, b). Since these two editors were developed in 2016 and 2017, subsequent efforts have significantly expanded their genome-targeting range and improved their efficiency and product purity. The use of Cas nickase, fusion of a second uracil-DNA glycosylase inhibitor (UGI) domain, the addition of a nuclear localization sequence, and linker and codon optimization have greatly increased the editing efficiencies of base editors [ 79 , 80 , 81 ]. To maximize the editing scope of base editors, diverse base editors with natural, engineered, or evolved Cas variants that recognize alternative PAMs and various deaminases have been created [ 82 , 83 , 84 , 85 , 86 , 87 , 88 ].

Programmable C-to-G base editors (CGBEs) that can achieve targeted C-to-G and G-to-C base transversions have recently been developed [ 89 , 90 ]. CGBEs originate from CBE by replacing the UGI with a uracil-DNA glycosylase (UNG), which excises the U base generated by cytosine deaminase, resulting in an abasic site followed by the preferential installation of a G base through the DNA repair mechanism (Fig.  2 c). Although CGBEs can provide efficient C-to-G and G-to-C editing, very few sites are suitable for CGBE editing. Several studies have used machine learning to optimize CGBEs to improve editing efficiency and product purity by changing the species origin, modifying the relative positions of UNG and deaminase, and optimizing codons [ 91 , 92 ].

The therapeutic applications of base editors have been hampered by their genome-wide off-target effects. Recent studies have shown that cytidine deaminases used in CBE induce genome-wide off-target editing independently of sgRNA or Cas9 [ 93 , 94 ]. In addition, both ABE and CBE can cause transcriptome-wide mutations [ 95 , 96 ]. Continuous efforts have been made to reduce off-target effects by engineering the DNA- [ 97 , 98 ] or RNA-binding domain [ 95 , 96 , 99 ], thereby bringing base editors closer to clinical applications.

Prime editing is a newly developed precise genome editing technology that enables all types of base conversion, small deletions, and insertions, as desired. PEs consist of a prime editor protein and prime editing gRNA (pegRNA). The PE protein is constructed by fusing an engineered Cas9 nickase (H840A) with reverse transcriptase, which can be targeted to the genomic locus by pegRNA [ 18 ]. The pegRNA combines a gRNA recognizing the target genomic sequence, a reverse transcriptase template encoding the desired edits, and a primer binding site to initiate reverse transcription [ 18 ] (Fig.  2 d). The newly synthesized edited DNA strand is incorporated into the target locus to generate heteroduplex DNA, in which the non-edited strand is eventually replaced by an edited strand through DNA repair. Compared to base editing, which often introduces bystander editing of extra bases in an activity window, prime editing is more versatile and precise.

A series of PE systems, namely PE2, PE3b, PE4, and PE5b, have been developed and are most widely used. All these systems share a common PE2 protein with an engineered Moloney murine leukemia virus (M-MLV) reverse transcriptase instead of the wild-type M-MLV reverse transcriptase in PE1 to increase editing efficiency [ 18 , 100 ]. The PE3 system contains an additional sgRNA that targets the non-edited strand to increase the editing efficiency [ 18 ]. The PE2 and PE3 systems were further optimized by introducing a DNA mismatch repair-inhibiting domain MLH1dn to generate PE4 and PE5 systems, respectively [ 18 , 100 ]. Systems ending in “b”, namely PE3b and PE5b, use an edit-specific nicking sgRNA to reduce indel levels [ 18 , 100 ]. Constant efforts are being devoted to optimizing PEs, with a primary focus on improving their editing efficiency. Optimization of PE2 protein architecture by codon optimization, SpCas9 mutation, and alterations of the nuclear localization signal and peptide linker sequence results in PEmax protein architecture, which greatly enhances editing efficiency [ 100 ]. They also constructed two types of engineered pegRNAs (epegRNAs) by incorporating 3′ structural motifs, which stabilize pegRNA and increase prime editing efficiency [ 101 ]. Many other groups have adopted similar strategies by optimizing either PE proteins [ 102 , 103 , 104 ] or pegRNAs [ 105 ].

In addition, a dual pegRNA strategy has been used to improve editing efficiency, which can also achieve programmable insertion, deletion, and replacement of large genomic sequences at specific genomic sites [ 106 , 107 , 108 , 109 , 110 ]. Using a pair of pegRNAs, each of which targets a different DNA strand and template the synthesis of complementary DNA flaps, endogenous targeted DNA sequence between the PE-induced nick sites is successfully replaced. This strategy also achieves targeted insertion of gene-sized DNA plasmids (> 5 kb) and targeted inversions of 40 kb in human cells when co-expressing a site-specific serine recombinase, Bxb1 integrase [ 108 ]. The dual pegRNA strategy with expanded capabilities of precision genome editing provides new possibilities for treating genetic disorders caused by large DNA deletions or complex structural mutations.

CRISPR-associated transposon (CAST) systems for large DNA insertion

CAST systems, consisting of transposase subunits and CRISPR effectors, facilitate the RNA-guided transposition of mobile genetic elements, making it a promising system for targeted, precise, and efficient insertion of large DNA segments. Most identified CASTs are derived from Tn7-like transposons that retain the core genes of the transposition machinery, but have no genes for target selection [ 111 , 112 ]. Instead, CASTs co-opt nuclease-deficient CRISPR-Cas proteins to induce RNA-guided transposition [ 111 , 112 ]. Several CAST systems have been experimentally or bioinformatically characterized, including type I-B, type I-C, type I-F, type IV, and type V-K CAST systems [ 111 , 113 , 114 , 115 ]. Bioinformatic analysis of the metagenomic database also revealed a non-Tn7 CAST system that co-opts a nuclease-inactive Cas12 and type I-E cascade [ 111 ].

Type I-F and type V-K CAST systems have been successfully reconstituted to achieve the integration of donor DNA into specific bacterial genome sites [ 113 , 114 ] (Fig.  3 a). An improved version of the type I-F CAST system enables highly specific and effective integration of up to 10 kb DNA fragments in the bacterial genome [ 116 ]. However, the application of these two systems in mammalian cells has not been reported. A very recent study developed an artificial transposon-associated CRISPR-Cas system named find and cut-and-transfer (FiCAT) system by coupling a SpCas9 protein with an engineered piggyBac (PB) transposase (Fig.  3 b), which achieved the targeted integration of multi-kilobase DNA fragments into the genomes of mammalian cell lines and mouse liver [ 117 ]. The discovery of CAST systems has expanded the genome editing toolkit, although CAST systems still require extensive modification and optimization until they can be conveniently and effectively applied to biomedical research.

CRISPR-Cas-based transposon systems. Schematic of CRISPR-based transposon systems, CAST system ( a ) and FiCAT system ( b ), which mediate site-specific DNA integration. Tns Tn7-like transposases, PB piggyBac transposase, LE transposon left end sequences, RE transposon right end sequences, CAST CRISPR-associated transposon, FiCAT find and cut-and-transfer

Delivery systems for CRISPRs

The safe, effective, and tissue-specific delivery of CRISPR-Cas tools in vivo determines whether CRISPR-based gene therapy can be used for this tissue. Therapeutic in vivo delivery systems for CRISPR-Cas have recently been discussed [ 118 , 119 ]. CRISPR-Cas tools can be delivered in the form of DNA, mRNA, or ribonucleoprotein complexes (RNP) through ex vivo or in vivo approaches. Various robust methods have been established to deliver genome editing reagents ex vivo, some of which have been used in multiple clinical trials involving different types of diseases [ 120 , 121 , 122 ]. The most efficient method of in vivo delivery of editors reported so far is the use of AAV, which can deliver editor-encoding DNA to target tissues and has been applied in clinical trials [ 118 , 123 , 124 ]. However, AAV-based delivery of DNA-encoding editing agents has a number of disadvantages, such as the possibility of viral vector integration into the transduced cell genome and increased frequency of off-target editing due to prolonged expression [ 75 , 119 , 125 , 126 ], which limits its clinical application. Therefore, safer alternative strategies for in vivo delivery of genome editors must be developed.

As a gene therapy delivery system approved by the Food and Drug Administration (FDA), lipid nanoparticles (LNP) have been demonstrated to safely deliver therapeutic small molecules and nucleic acid drugs to hepatocytes and antigen-presenting cells via systemic administration or intramuscular injection. The LNP system was used to deliver gene editing tools in the first clinical trial involving human gene editing in vivo [ 123 ]. However, because intravenously delivered LNP showed liver tropism, delivering editors to non-hepatocytes has been a huge challenge. Recent studies have shown that high-throughput screening identifies nanoparticles targeting non-hepatocytes, including endothelial cells (ECs) and spleen immune cells [ 127 , 128 ]. In addition, cell-type specificity of LNP-mediated Cas9 therapies could be modified by reducing Cas9-mediated insertions and deletions in hepatocytes using inhibitory oligonucleotides and siRNAs [ 129 ].

Virus-like particle (VLP) systems, which combine the advantages of viral and non-viral delivery systems, are another promising in vivo gene editing delivery vehicle [ 126 ]. VLPs can package genome editing agents in the forms of mRNA or RNP. The short cellular lifespan of RNPs effectively restricts off-target editing. Almost all current VLPs are derived from retroviruses and contain most viral components but no viral genome [ 118 , 130 , 131 , 132 , 133 ]. Very recently, fourth-generation engineered VLPs (eVLPs) based on M-MLV have been developed to deliver Cas9 or base editor RNPs both in vitro and in vivo [ 126 ]. A single intravenous injection of eVLPs carrying a base editor targeting proprotein convertase subtilisin/kexin type 9 ( Pcsk9 ) can achieve base editing in multiple tissues, reduce serum PCSK9 levels by 78%, and partially restore visual function when designed for retinal editing in a mouse model of blindness [ 126 ]. The mammalian endogenous retrovirus-like protein PEG10 has also been programmed as a VLP system called selective endogenous encapsidation for cellular delivery (SEND) platform, which can package and deliver mRNA encoding Cas9 in vivo [ 134 ]. Based on endogenous mammalian proteins, the SEND system may be less immunogenic than bona fide retrovirus-based VLP systems.

Applications of genome editing in modeling and treating CVD

CVD, including heart and vascular diseases, are leading causes of morbidity and mortality at different ages [ 135 , 136 ]. In recent years, a tremendous amount of new genetic information related to CVD has been identified using next-generation sequencing technologies [ 19 , 20 ]. Owing to the advent and development of the CRISPR-Cas system, we can now handle this information and determine CVD-related functions much more easily than ever. The CRISPR-Cas system also provides more possibilities for treating inherited CVD by correcting disease-causing mutations in the patient genome. As the most commonly used Cas proteins, SpCas9 and SaCas9 have been broadly applied in CVD-related modeling and therapeutic purposes, both in vitro [ 137 , 138 , 139 , 140 ] and in vivo [ 141 , 142 , 143 , 144 ]. Newly developed base editing and prime editing systems have also been used [ 145 , 146 ]. Additionally, delivering CRISPR-Cas components to the cardiovascular system remains challenging, and AAV-based systems are currently the most widely used methods [ 147 , 148 , 149 , 150 ].

Modeling CVD using CRISPR

Genetic studies have identified various pathogenic genetic variants associated with the occurrence of CVD [ 151 , 152 ]. Revealing the consequences of specific mutations in CVD-related genes is important for CVD genetic diagnosis and precise medicine. CVD models have played a critical role in establishing causal links between genetic variants and CVD, dissecting the molecular mechanisms underlying CVD, validating therapeutic targets, and preclinical evaluation of therapeutic agents. At present, multiple gene editing tools have been applied to create in vitro and in vivo models of CVD [ 151 ].

In vitro models of CVD

Human induced pluripotent stem cells (hiPSCs) are promising for modeling human cardiomyopathies in vitro because they can differentiate into cardiomyocytes [ 153 ]. Through genome editing of hiPSCs followed by their differentiation into cardiomyocytes (hiPSC-CMs), isogenic hiPSC-CMs have been broadly used to verify causative genes or mutations in cardiomyopathies. Gene disruption induced by CRISPR-Cas9 in hiPSC-CMs is straightforward and suitable for determining the role of a gene in CVD. For example, DNA methyltransferase 3A ( DNMT3A ) gene-deleted hiPSC-CMs generated using CRISPR-Cas9 gene editing showed altered contraction kinetics and impaired glucose/lipid metabolism, suggesting an important role of DNA methylation in cardiac diseases [ 137 ]. The homozygous SCN10A gene (encoding Na V 1.8) knockout hiPSC-CMs help demonstrate that the voltage-gated sodium channel Nav1.8 contributes to late Na + current (I NaL ) formation and displays a harmful proarrhythmogenic function [ 138 ].

The precise introduction of point mutations into hiPSC-CMs facilitates the determination of the causal relationship between genetic mutations and heart diseases. Striated muscle-enriched protein kinase (SPEG) E1680K homozygous mutant hiPSC-CMs recapitulate the hallmarks of dilated cardiomyopathy (DCM), confirming that SPEG E1680K is a novel DCM-causing mutation [ 139 ]. Genome editing of hiPSCs has also been used to identify several causative mutations of arrhythmias. hiPSC-CMs expressing an R211H substitution in the Ras-related associated with diabetes ( RRAD ) gene mimic the single-cell electrophysiological characteristics of Brugada syndrome, a disorder predisposing the patient to ventricular arrhythmias, indicating that RRAD is possibly a novel susceptibility gene for Brugada syndrome [ 154 ]. CRISPR-Cas9-engineered hiPSC-CMs carrying three different mutations of ryanodine receptor 2 (RyR2), R420Q, Q4201R, or F2483I, exhibit various pathological features of catecholaminergic polymorphic ventricular tachycardia 1 (CPVT1)-associated arrhythmia, suggesting that different RyR2 mutations cause varied Ca 2+ signaling consequences and drug sensitivities [ 155 ]. Genome-edited hiPSC-CMs can also be used as high-throughput platforms for scalable functional validation of the pathogenicity and pathophysiology of genetic variants identified in the human population. To determine the functional significance of cardiac troponin T ( TNNT2 ) variants, the endogenous TNNT2 gene was knocked out in hiPSCs using CRISPR-Cas9, and 51 different TNNT2 variants were expressed using lentivirus in differentiated TNNT2 knockout hiPSC-CMs. The results revealed that various TNNT2 variants exhibit different pathogenic mechanisms, greatly expanding the knowledge of which and how TNNT2 variants cause hypertrophic cardiomyopathy (HCM) and DCM [ 156 ].

Genome editing has been used to correct mutations to generate optimal isogenic controls for patient-derived iPSC-CMs, enabling the determination of genotype–phenotype relationships more precisely. Genome editing has been performed to correct the missense mutation T618I in the potassium channel gene KCNH2 in short QT syndrome patience-specific hiPSC-CMs to elucidate the single-cell phenotype of short QT syndrome [ 157 ]. Using iPSC-CMs derived from doxorubicin-treated pediatric patients, cytosine base editing has helped identify the single nucleotide polymorphism rs11140490 in the SLC28A3 locus, which is a novel protector against doxorubicin-induced cardiotoxicity [ 145 ]. Isogenic hiPSC-CM controls generated by CRISPR-based gene correction have also been used as platforms to evaluate other therapeutic methods. Type 1 long QT syndrome is caused by loss-of-function variants in the KCNQ1-encoded Kv7.1 potassium channel α-subunit. iPSC-CMs generated from patients with KCNQ1-V254M and -A344A/spl mutations have recently been corrected using CRISPR-Cas9 to act as isogenic controls, which have been used to evaluate a dual-component suppression-and-replacement gene therapy method [ 158 ]. Base editing and prime editing could possibly be widely used in establishing hiPSC-CM-based CVD models in the near future.

Animal models of CVD

CRISPR-based germline genome editing tools have revolutionized the generation of genetically modified animal models of CVD. Compared to conventional gene targeting technologies using embryonic stem cells, CRISPR-based gene editing technologies are easier to operate, faster, and applicable to most species. One strategy for generating animal models of CVD is to introduce targeted point mutations, insertions, or deletions using HDR-mediated germline genome editing. A mouse model of HCM with a Myh6 R404Q mutation was generated using SpCas9/ssODN-mediated directed genomic DNA editing, and heterozygous mice developed a typical HCM phenotype [ 141 ]. A similar approach was also utilized to insert an additional adenine nucleotide into the lysosomal acid alpha-glucosidase ( Gaa ) gene at the c.1826 locus and generate a novel mouse model of infantile-onset Pompe disease (IOPD), which recapitulates HCM and the skeletal muscle weakness of human IOPD [ 142 ]. A CRISPR-Cas9-generated rat model, with a 9 bp deletion within the hotspot analogous to the novel mutation of the human PDE3A gene, recapitulates arterial hypertension with brachydactyly, demonstrating that mutant PDE3A causes arterial hypertension [ 143 ]. Recently, a 94 bp out of frame deletion was generated in exon 1 of Kcnk3 using SpCas9/ssODN-mediated genome editing, creating a novel rat model of pulmonary arterial hypertension [ 159 ]. Another strategy is to delete exon(s) using two sgRNAs flanking specific exon(s). Exon deletion mutations in the dystrophin are among the most common causes of Duchenne muscular dystrophy (DMD). Several mouse models of DMD have been generated using CRISPR-Cas9 genome editing [ 144 , 160 ], which are discussed further in the next section. CRISPR-Cas9 mediated mosaic inactivation of zebrafish ccm2 led to a lethal multi-cavernous lesion that histologically mimics the typical human hemorrhagic cerebral cavernous malformation [ 161 ].

Compared with germline genome editing, somatic genome editing is a more flexible method for obtaining CVD models, which overcomes the challenges of germline modification, such as embryonic lethality and the cost and time required to establish, reproduce, and maintain these models. It is also suitable for rapid and relatively high-throughput studies on the functions of CVD-related genes. As early as 2016, a cardiomyocyte-specific SpCas9 transgenic mouse model was successfully generated to achieve somatic editing in the heart [ 162 ]. Following this study, intraperitoneal injection of AAV9 encoding sgRNA against three genes critical for the heart, Myh6 , Sav1 , and Tbx20 , in postnatal cardiomyocyte-Cas9 transgenic mice caused a similar degree of DNA disruption and subsequent mRNA downregulation, but only Myh6 disruption induced HCM and heart failure, suggesting that the effect of postnatal cardiac genome editing is target-dependent [ 147 ]. Mouse models can also be generated by activating endogenous gene expression through CRISPR-mediated genome editing in the postnatal heart [ 163 ]. CRISPR-mediated endogenous activation of myocyte enhancer factor 2D ( Mef2d ) leads to cardiac hypertrophy in mice, indicating that CRISPR-mediated genome editing can be used to generate CVD mouse models by controlling transcription in the postnatal heart [ 163 ]. Several recent studies have shown that CRISPR-Cas9 can be used to edit endothelial genes in vivo to obtain vascular disease models and enable reverse genetic studies of gene function in the mammalian vascular endothelium. Co-injection of an adenovirus harboring sgRNAs targeting the Alk1 gene and AAV1-VEGF successfully induced mutations in Alk1 in brain ECs and generated brain arteriovenous malformations in adult mice [ 164 ]. We recently generated a blood–brain barrier (BBB) breakdown mouse model by AAV-BR1-CRISPR mediated somatic genome editing. A single intravenous administration of brain microvascular EC targeting AAV-BR1 encoding sgRNA against the β-catenin ( Ctnnb1 ) gene resulted in a mutation of 36.1% of the Ctnnb1 alleles and dramatically decreased levels of CTNNB1 in brain ECs, leading to BBB breakdown in EC-restricted Tie2 Cas9 mice [ 148 ]. The AAV-BR1-CRISPR system established in this study allowed for the rapid construction of BBB perturbation models in vivo and may be helpful for developing drug delivery systems in the central nervous system. Recently, the nanoparticle-mediated delivery of CRISPR plasmid DNA expressing Cas9 under the control of the Cdh5 promoter resulted in efficient genome editing in the ECs of the peripheral vasculature in adult mice, which provides a powerful tool to construct animal models of peripheral vascular diseases [ 165 ].

Genome editing in CVD treatment

Therapeutic genome editing can be used to treat monogenic CVD, and the technology could permanently correct mutations and eventually eradicate specific CVD. Programmed edits were introduced into the human germline genome [ 166 , 167 , 168 , 169 ]. However, human germline genome editing faces significant ethical concerns and is prohibited in most countries [ 170 ]. Somatic editing is a promising technology for editing CVD-causing mutations without the risk of passing genomic changes to the offspring. Table 1 summarizes the latest applications of genome editing in treating different types of CVD.

DMD is an X-linked disorder characterized by proximal muscle weakness and cardiomyopathy caused by mutations in the largest human gene, dystrophin ( DMD ) [ 182 ]. A variety of mutations exist throughout the DMD gene, most of which are located in the regions crossing exons 43 to 53 and disrupt the ORF, resulting in non-functional truncated proteins.

A single-cut genome editing strategy was applied in both iPSC-CMs and mouse models of DMD bearing an exon 44 deletion mutation (Δ44), one of the most common causative mutations of DMD. The ORF can be restored by disrupting the exon splice site to skip the adjacent exon, inserting one nucleotide, or deleting two nucleotides in exon 44 [ 144 ]. Systemic delivery of gene editing components by a single dose of AAV9 restores ~ 90% dystrophin protein expression in the hearts of Δ44 mice within 4 weeks [ 144 ]. This approach also helps to correct DMD models bearing deletions of exons 43, 45, and 52 (Δ43, Δ45, and Δ52) both in vitro and in vivo [ 171 ]. A dual-AAV system was used to deliver SpCas9 and a single sgRNA targeting the splice donor site of exon 44 (for Δ43 or Δ45 mice) or splice acceptor site of exon 53 (for Δ52 mice) in vivo to restore dystrophin expression. Both exon skipping and reframing were induced in Δ45 and Δ52 mice, and the efficacy of dystrophin in these two models was higher than that in Δ43 mice, in which only exon skipping was generated [ 171 ]. Restoration of dystrophin has also been achieved in hiPSC-CMs from these DMD models [ 171 ]. However, this study did not specify whether exon skipping and/or reframing could subsequently rescue the cardiac phenotypes of DMD models [ 171 ]. Another strategy is to delete exon(s) using two sgRNAs flanking on either side, thus restoring the ORF of the DMD gene. The systemic application of AAV9 carrying an intein-split SpCas9 and a pair of sgRNAs targeting sequences flanking exon 51 in a pig model of DMD lacking exon 52 induced dystrophin expression in the heart and reduced arrhythmogenic vulnerability [ 172 ]. The long-term efficacy and safety of therapeutic editing for DMD have also been studied [ 149 , 150 ]. AAV vectors carrying SaCas9 and a pair of sgRNAs targeting exon 23 or exons 21–23 were administrated for one year or 19 months, respectively, to mdx mice. Cardiac functions were improved without serious adverse effects, indicating that in vivo CRISPR genome editing may be a safe therapeutic strategy for DMD [ 149 , 150 ].

Base editing and prime editing show great promise for treating DMD. Both ABE and PE can restore dystrophin protein expression by inducing exon skipping or exon reframing to correct the Dmd exon 51 deletion mutation in iPSC-CMs, and intramuscular delivery of AAV9 encoding ABE components amends the mutation in ∆E51 DMD mice [ 146 ]. CBE has been shown to rescue dystrophic cardiomyopathy in Dmd E4* mice, which harbor a 4 bp deletion in exon 4 of the Dmd gene and recapitulate many characteristics of human DMD [ 160 ]. A single-dose administration of AAV9-eTAM encoding a fused nuclease-defective SaCas9 (KKH) with activation-induced cytidine deaminase (AID) and UGI, together with AAV9-sgRNA, efficiently induced splice site mutation and exon 4 skipping of the Dmd gene and restored up to 90% of dystrophin proteins in the heart of Dmd E4* mice, resulting in improved cardiac function and an increased life span [ 160 ]. Alternatively, a dual AAV-mediated protein trans-splicing approach was used to deliver a modified ABE-NG to an mdx 4cv mouse model carrying a premature stop codon (CAA-to-TAA) in exon 53 of the Dmd gene. After 10 months of treatment, a near-complete rescue of dystrophin was found in the hearts of mdx 4cv mice without obvious toxicity [ 173 ].

HCM and DCM

Inherited cardiomyopathies, including HCM and DCM, are candidate genetic disorders that are suitable for genome editing-related treatment. ABEmax-NG has been shown to correct a pathogenic R404Q/+ mutation in embryos of the HCM mouse model [ 141 ]. Administration of ABEmax-NG mRNA to Myh6 R404Q/+ embryos corrects the mutant allele at a rate of 62.5% to 70.8%, abolishing the HCM phenotype in postnatal mice and their progeny. Moreover, in utero delivery of intein-split ABEmax-NG induced a high correction rate without introducing indels or off-target editing in Myh6 R404Q/+ fetuses [ 141 ]. Intronic CRISPR repair has been demonstrated as efficient in a preclinical iPSC-CM model of Noonan syndrome-associated HCM [ 174 ]. CRISPR-Cas9-mediated destruction of the mutation-induced additional intronic donor splice site can reverse the hypertrophic phenotypes in Noonan syndrome patient-derived iPSC-CMs carrying biallelic mutations in intron 16 of the leucine zipper-like transcription regulator 1 ( LZTR1 ) gene, indicating new possibilities for personalized therapeutic genome editing in HCM patients [ 174 ]. Notably, CRISPR-based genome editing has been shown to have potential to correct a well-documented heterozygous dominant 4 bp deletion in exon 16 of MYBPC3 , which causes familial HCM, in human embryos [ 175 ]. Co-injection of Cas9 proteins, mutation-specific sgRNAs, and mutant sperm into healthy metaphase II oocytes corrected the deletion by wild-type maternal allele-mediated HDR, resulting in a high yield of homozygous embryos carrying the wild-type MYBPC3 gene without mosaicism or off-target mutations [ 175 ].

Genome editing has also been used to correct DCM-causing mutations. In addition to previous studies using hiPSC-CMs showing that truncated titin (TTNtv) mutations are the most common causes of DCM [ 183 , 184 , 185 ], recently, the pathological mechanisms of TTNtv-associated DCM have been highlighted and a new genome editing strategy has been developed to treat TTNtv-associated DCM. iPSC-CMs with patient-derived or CRISPR-Cas9-generated TTN mutations were corrected using SpCas9/ssODN. Engineered heart muscle generated from corrected hiPSC-CMs shows normalized titin protein levels and contractile function [ 140 ]. Genome editing using SpCas9 and A-band TTNtv -specific sgRNA was also shown to restore the reading frame of TTN protein in hiPSC-CMs, leading to increased full-length TTN protein levels and normalized sarcomere function [ 176 ]. More recently, the application of precise genome editing technology for treating DCM caused by mutations in RBM20 has been reported [ 177 ]. The RBM20 R634Q and RBM20 R636S mutant iPSCs were corrected by ABE and PE, with the efficiency of 92% and 40%, respectively. In addition, AAV9-mediated systemic delivery of ABE components corrected 66% of the RBM20 transcripts expressed in cardiomyocytes of postnatal RBM R636Q/R636Q mice. The corrected mice showed restored cardiac size and function, and prolonged life span [ 177 ].

Cardiac arrhythmia

Cardiac arrhythmia caused by autosomal-dominant mutations can be treated with CRISPR-mediated specific disruption of the mutant allele, which has been validated in several mouse models [ 186 , 187 ]. Recently, humanized mice expressing a human mutant PLN (hPLN-R14del) demonstrated bi-ventricular dilation and a higher propensity for sustained ventricular tachycardia. Disruption of the hPLN-R14del allele by AAV9-CRISPR-Cas9 improved cardiac function and reduced sustained ventricular tachycardia susceptibility in young adult humanized PLN-R14del mice, providing a potential therapeutic strategy for the arrhythmogenic phenotype in human patients with the PLN-R14del mutation [ 178 ].

Atherosclerosis

Atherosclerosis is a chronic disease that refers to the formation of fibrofatty lesions in the arterial wall, and causes ischemic stroke, ischemic cardiomyopathy, myocardial infarction, and peripheral arterial disease. Blood concentration of low-density lipoprotein cholesterol (LDL-C) is one of the best-established causal risk factors for atherosclerosis.

The lipid metabolism-related gene, Pcsk9 , is specifically expressed in the liver and functions primarily as an antagonist to the LDL receptor. Disruption of PCSK9 activity can reduce circulating LDL-C levels, thereby lowering the risk of atherosclerosis [ 188 ]. Several clinical trials have investigated monoclonal antibodies targeting PCSK9. However, even if these antibody-based drugs are effective, their effect on LDL-C is short-lived. Genome editing using CRISPR systems provides an alternative method for reducing PCSK9 levels. A single administration of adenovirus co-expressing SpCas9 and sgRNA targeting exon 1 of the mouse Pcsk9 gene can efficiently introduce loss-of-function mutations into endogenous Pcsk9 genes in vivo and chronically decrease plasma cholesterol levels in the blood [ 179 ]. The AAV-SaCas9 system has also been proven to be effective in editing the Pcsk9 gene in vivo, leading to significantly decreased serum PCSK9 and cholesterol levels [ 30 ]. Single injections of engineered DNA-free VLPs targeting the Pcsk9 gene into adult mice demonstrated 63% base editing in the liver, resulting in 78% reduction in serum PCSK9 levels [ 126 ]. In addition to these studies carried out in rodents, somatic Pcsk9 gene editing has also been validated in nonhuman primates [ 180 , 181 ]. CRISPR base editors delivered using LNPs proved highly effective in editing the Pcsk9 gene in the liver of macaques and cynomolgus monkeys. A single-dose treatment of LNPs carrying CRISPR base editors leads to stable Pcsk9 knockdown in the liver and a 60% reduction in blood LDL-C for at least 8 months [ 180 ]. To ensure the safety of gene editing in comparatively more proliferative organs, such as the liver, the proportion of edited cells that remain stable over time must be investigated. All these genome editing approaches offer the potential for once-and-done therapies for the lifelong treatment of atherosclerosis-associated CVD.

Perspectives

CRISPR-based genome editing technology has been rapidly applied in almost all fields, from basic biology to translational medicine. The development of novel systems and tools for more accurate, efficient, and faster genome editing and tighter control of the duration, efficiency, and specificity of genome editors will further benefit their translational applications. Newly uncovered thousands of phage-encoded CRISPR systems provide a valuable resource for searching novel miniature single-effector CRISPR-Cas systems [ 39 ]. In addition, newly developed Cas13a-based RNA editing tools can achieve RNA knockdown and precise base editing of mammalian transcripts without causing DNA damage, providing a promising potential therapeutic strategy in translational cardiovascular medicine [ 11 , 189 ]. Notably, type III-E CRISPR-Cas7-11 effector has recently been shown to cleavage protein under target RNA guidance [ 190 , 191 ], bringing new potential CRISPR tools for CVD diagnosis and treatment.

Genome editing technologies have been successfully translated into human clinical trials for enhanced chimeric antigen receptor (CAR) T-cell therapy, cell-based regenerative medicine, and treatment of monogenic diseases, such as transfusion-dependent beta thalassemia (TDT) and sickle cell disease (SCD) [ 192 , 193 ]. Researchers have used in vivo genome editing to target the transthyretin (TTR) gene to treat transthyretin amyloidosis and have achieved very encouraging results in phase 1 clinical trials, taking the most critical step towards applying CRISPR-based genome editing technology to treat human genetic diseases [ 123 ]. Taking the most optimistic view, CVD with known causal genes can theoretically be treated with CRISPR technology. However, there are still several important challenges. Recently, CRISPR-based genome editing in human embryos was shown to cause unpredictable genomic alterations, including DNA rearrangements, large deletions, and even loss of allele-specific chromosomes [ 168 , 194 , 195 ]. Therefore, potential technical safety concerns, including mosaicism, off-target effects, and long-term risks caused by genome editing, need to be addressed before the therapeutic applications of CRISPR technology in treating CVD. Perhaps striking a balance between the efficiency and safety of genome editing is crucial. At present, efficient delivery of CRISPR-Cas systems to human cardiovascular system remains a challenge. In addition, the efficacy and safety of each therapeutic gene editing strategy for each CVD need to be confirmed by clinical trials. Although this paper uses the CVD as example to illustrate the progress of CRISPR-based genome editing in modeling and treating diseases, the same strategies could also be used for numerous diseases in other tissues.

Like any cutting-edge technology, gene editing technology could be a double-edged sword. Genome editing has been listed as a potential weapon of mass destruction in the 2016 annual worldwide threat assessment report of the U.S. intelligence community, indicating a high risk of extreme misuse. Recent rapid advances have made genome editing technologies more accessible and difficult to control, which may further lower the threshold for genome editing misuse and increase biosecurity threats. The possible misuse of genome editing technology and biosecurity risks may include, but are not limited to creating (1) pathogens with increased virulence, (2) new pathogens and biotoxins, and (3) gene-driven animals that may have irreversible effects on specific populations and the environment. Regulations and guidelines should be developed after extensive consultation to ensure that the development of gene editing technologies will not harm living organisms, including humans, or the environment.

Conclusions

The emerging novel Cas nucleases and their extended applications have greatly expanded the CRISPR-based genome editing toolbox and promoted the development of life science and medicine. CRISPR-based genome editing technology has also revolutionized cardiovascular research, accelerating the generation of genetically modified models of CVD and its application in the treatment of different types of CVD. However, this technology may also bring huge potential biological threats, which should be strictly controlled to prevent its abuse.

Availability of data and materials

Not applicable.

Abbreviations

Adeno-associated virus

Adenine base editor

Blood–brain barrier

Clustered regularly interspaced short palindromic repeats

CRISPR-associated protein

• Cardiovascular disease

CRISPR-associated complex for antiviral defense

Cytosine base editor

C-to-G base editor

CRISPR-associated transposon

Chimeric antigen receptor

Dilated cardiomyopathy

Duchenne muscular dystrophy

Double-strand break

Endothelial cell

Find and cut-and-transfer

Human induced pluripotent stem cell

Hypertrophic cardiomyopathy

Homology-directed repair

Lipid nanoparticle

Low-density lipoprotein cholesterol

Moloney murine leukemia virus

Non-homologous end-joining

Open reading frame

Obligate mobile element-guided activity

Protospacer adjacent motif

Prime editor

Prime editing gRNA

Ribonucleoprotein complexes

Single guide RNA

Single-stranded oligodeoxynucleotide

Sickle cell disease

Trans-activating CRISPR RNA

Transposon-associated motif

Transfusion-dependent beta thalassemia

Uracil-DNA glycosylase inhibitor

Uracil-DNA glycosylase

Virus-like particle

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This work was supported by the National Natural Science Foundation of China (82270355, 82270354, 81970134, 82030011, 31630093), and the National Key Research and Development Program of China (2019YFA0801601, 2021YFA1101801).

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State Key Laboratory of Proteomics, Beijing Proteome Research Center, National Center for Protein Sciences, Beijing Institute of Lifeomics, Beijing, 100071, China

Zhen-Hua Li, Jun Wang, Jing-Ping Xu, Jian Wang & Xiao Yang

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ZHL, JW, JW and XY drafted the manuscript. ZHL and JPX prepared figures. JW and XY conceived and supervised the study. All authors have read and approved the final manuscript.

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Li, ZH., Wang, J., Xu, JP. et al. Recent advances in CRISPR-based genome editing technology and its applications in cardiovascular research. Military Med Res 10 , 12 (2023). https://doi.org/10.1186/s40779-023-00447-x

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Ethics and Genomic Editing Using the Crispr-Cas9 Technique: Challenges and Conflicts

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The field of genetics has seen major advances in recent decades, particularly in research, prevention and diagnosis. One of the most recent developments, the genomic editing technique Clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9, has opened the possibility for genetic therapies through genome modification. The technique marks an improvement on previous procedures but poses some serious ethical conflicts. Bioethics is the discipline geared at finding answers to ethical challenges posed by progress in medicine and biology and examining their repercussions for society. It can also offer a conceptualization of these ethical dilemmas. The aim of this paper is to offer a map of the ethical dilemmas associated with this technique by way of a critical analysis of current literature. The main issues can be grouped in four areas: efficacy and security; the types of cells which can be targeted by the technique (somatic, embryonic and gametes); the goal of the therapy; and accessibility and justice.

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Introduction

The field of genetics has undergone remarkable development in recent years, and promising advances are constantly being made. One example of this is the recent development of the Clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 genome editing technique, which has made the modification of the human genome a distinct possibility. This is reminiscent of the news of the birth of two babies genetically modified in 2018 by the Chinese scientist He Jiankui.

These new possibilities raise ethical questions about this technique that have important implications for individuals, society and indeed the whole human species. Literature on the technical characteristics of CRISPR and the ethical challenges that it poses is already abundant, but the rapid pace of progress makes it difficult to establish a clear and precise view of the challenges that it poses.

Bioethics is the discipline geared at finding answers to ethical challenges posed by progress in medicine and biology. Its aim is to discuss the problems posed by medical and biological advances and the impact that they have on society and its value systems Abel [ 1 ].

As Bauman [ 2 ] observed: ‘Ethics…must deal with what-has-not-happened-yet, with a future that is endemically the realm of uncertainty and the playfield of conflicting scenarios. Visualization can never pretend to offer the kind of certainty which experts with their scientific knowledge and with greater or lesser credibility claim to offer. The duty to visualize the future impact of action (undertaken or not undertaken) means acting under the pressure of acute uncertainty. The moral stance consists precisely in seeing to it that this uncertainty is neither dismissed not suppressed, but consciously embraced’. This is not about preventing progress, but rather about being able to visualize the impact of our actions and thereby to minimize any possible negative effects.

This article aims to present a map of all the ethical issues raised by the CRISPR technique and to succinctly reflect upon them. The literature on the ethical challenges posed by CRISPR is already plentiful, but most of the articles published to date have tended to focus on only a limited number of these issues (and especially on those related to technical questions). We aim to present a global overview of all of the issues that arise (including technical, anthropological and social considerations). It should be noted that some of these arise in almost all genome editing techniques and are not exclusive to CRISPR; even so, CRISPR may cause them to surface in an easier, quicker or more intense way [ 3 ].

This work is the product of a project on the ethical implications of using the CRISPR technique that was developed by a multi-disciplinary group of researchers working at our institute. It is based on previous work, published in Medicina Clínica [ 4 ], whose objective was limited to making a brief enumeration of these issues (but without examining them in depth). Their mapping was, however, able to identify and define a series of potential conflicts and made it possible to analyse and provide a response to each of them, either in the form of regulation or a moratorium.

In presenting this global overview, the article addresses the following topics: the characteristics of the technique and its application; related ethical issues; the uses and purposes of the technique; and, finally, the question of social justice.

Characteristics of the Technique

The CRISPR technique is a genetic editing procedure that was first used in 2012 by a group of researchers from the University of Berkeley. Footnote 1 There are also other, similar, editing techniques (based on recombinant DNA), such as TALEN (transcription activator-like effector nucleases) and ZFN (zinc-finger nucleases), but CRISPR provides a better combination of three key factors: precision, accessibility and price. It is also easier to use than the alternatives, as the other techniques require more time and more specialised personnel.

It is a procedure that starts with the DNA of bacteria that provide an immune mechanism against viruses. These sequences are able to recognize viruses that enter bacteria and which ‘activate’ an enzyme that is able to break them down. In doing this, the enzyme makes use of the resulting fragments to immunise the bacteria against the virus. This process is made possible by CRISPR (the NRA molecule which transmits the biological information contained in the DNA for protein synthesis) using Cas9, which is a specific enzyme belonging to bacteria that can repair fragments pf DNA with great precision [ 5 , 6 ].

Application of the Technique

Genetic editing (and, in particular, the CRISPR technique) can be used in people, animals and plants [ 7 , 8 ].

In animals, genetic editing [ 9 ] can be used in food production (to increase muscle mass, improve nutritional content and breed more manageable animals) and to avoid, or prevent, diseases that could affect humans (for example, genetically modifying vectors to eradicate disease, as in the case of the Aedes aegypti mosquito, which transmits dengue fever, or in certain subspecies of the Anopheles mosquito genus, which carry the malaria parasite). Another hypothetical, although as-of-yet unfeasible, use of the technique would be to obtain organs to transplant into humans. In plants, genetic editing has also been used to improve food destined for human or animal consumption.

Some of the ethical problems associated with using this technique in animals and plants involve: potentially causing significant transformations of insect or plant species that could alter important ecological balances; producing ‘off-target’ Footnote 2 effects that may not be possible to control; having effects on animals and people that consume genetically modified animals; and the risk of unnecessarily and irresponsibly reducing the level of biodiversity [ 9 , 10 ].

Genetic Editing in Humans

The application of genetic editing techniques to humans is undoubtedly one of the issues that causes most debate and ethical interest in genetic engineering [ 3 ]. It has, however, been presented as one of the best tools for potentially avoiding, or preventing, diseases, as well as for genetically modifying an organism.

For example, certain types of cancer are currently treated using gene editing in somatic cells and there are on-going trials to treat Cooley's anaemia (β-thalassemia), sickle cell anaemia, mucopolysaccharidosis (types I and II) and haemophilia B, amongst other pathologies. For example, in 2017, S. Mitalipov and his team used the CRISPR technique to ‘repair’ a mutation associated with hypertrophic cardiomyopathy in an ovule, just prior to fertilisation [ 11 ]. The mutation was not ‘inherited’ by the resulting zygote and no mosaicisms Footnote 3 or ‘off-target’ effects were observed [ 12 , 13 , 14 , 15 ].

There are three different types of cells which can be modified by genetic editing, with different repercussions for the subjects:

Somatic cells: Their genetic modification only affects the individual, not their offspring.

Embryonic (pluripotential) cells: Their genetic modification only tends to affect the individual (although, in some cases, it can also affect their offspring).

Gametes: Their genetic modification affects the individual and is also transmitted to their offspring [ 10 ].

When possible, germ-line modification (which alters the genetic inheritance) can be achieved in two ways: by modifying germ cells (gametes: sperm cells or oocytes /ovules) and by modifying the zygote, or embryo, at an early stage in its development: before the formation of its reproductive organs [ 12 ].

Map of Ethical Conflicts in the Genetic Editing Technique

The European Society of Human Reproduction and Embryology (ESHRE) and the European Society of Human Genetics (ESHG) issued a document early in 2018, which was subsequently followed by another, setting out their rationale and outlining a series of practical recommendations. In this way, they officially stated their position regarding the ethical questions posed by the new technique [ 12 , 13 ]. Both documents particularly focused on issues related to the genetic editing of germ cells.

Amongst the various problems, or ethical issues, raised by genetic editing using the CRISPR technique, it is possible to distinguish four different groups. The first is associated with the technique itself and its effectiveness and safety; the second is related to the type of cells to which the technique is applied; the third refers to the purpose for which it is applied; and the fourth is related to its accessibility. Although these different groups of issues are closely related, they are not really the same in nature.

Ethical Issues Related to the Technique

The most important ethical problems presented by this technique are surely those related to its safety and efficacy [ 15 ]. With respect to precision, it should be underlined that this approach is not as accurate as might be expected [ 10 , 15 , 16 ]. Guttinger [ 6 ] states that total accuracy is not possible due to the complexity of DNA sequencing and its relationship with the RNA molecule.

Another difficult issue is related to the efficiency of the technique and the difficulty involved in controlling and determining its off-target effects. If its accuracy were total, and if it were possible to always intervene on the desired gene, the next problem would be how to determine whether the effects of the intervention were, in fact, only the ones sought [ 6 , 10 , 15 , 16 ]. CRISPR is the gene editing technique that has the most off-target effects (compared to TALEN and ZFN) [ 15 ], although efforts are currently being made to reduce this unpredictability [ 10 ].

Such problems of safety and efficacy could also be present in He Jiankui’s experiments (He [ 17 ], according to two articles published in December 2019, in the MIT Technology Review [ 18 , 19 ]). Regalado [ 19 ] has explained the trajectory of the publication of He’s work ‘Birth of Twins After Genome Editing for HIV Resistance’ in several scientific journals and highlighted some serious methodological and ethical irregularities. Having reviewed He Jiankui’s work, Musurunu (2019) stated that evidence of mosaicism was present in both twin embryos, so they could still have been vulnerable to HIV, and that the possible presence of off-target mutations could not be completely ruled out. The negative impact of He Jiankui’s experiments may not only have caused damage to those directly affected, but it could also have slowed down research into genetic editing and in similar fields [ 20 , 21 ].

As a result, more knowledge is required before this technique should be applied to humans. For genetic editing to be successful, it is necessary to know how to determine the impact of small changes in DNA (or its ‘packaging’) on the chemical components and physical properties of cells. It is, therefore, important to improve our existing knowledge of genetic and epigenetic effects in order to subsequently determine, and predict, the phenotypic effects of genetic editing [ 15 ].

Bearing in mind these shortcomings, O’Keefe, Perrault and other researchers have asked whether it would not be a good idea to change the language used (and the metaphors reflected within it) when describing, or discussing, the CRISPR technique, and especially when referring to it in the press and other non-specialised types of literature. In fact, in the video produced by He Jiankui [ 17 ], the most repeated words are ‘safely’ and ‘healthily’. The verbs most frequently used to describe the process (‘edit’, ‘cut’, ‘erase’ and ‘repair’) suggest a degree of accuracy and security that, in practice, do not exist,or, at least, not to such a high degree. This is why authors often advocate using terms like ‘modify’, ‘change’ and ‘alter’, which are more realistic and have fewer potentially misleading connotations [ 22 ].

Ethical Issues Related to Its Use

As previously mentioned, there are three types of cells that can be the object of genetic editing: somatic cells, embryonic (pluripotential) cells and germ cells (gametes). Somatic cells, and most embryonic cells, present the fewest ethical problems since the intervention upon them only affects the individual, but not their offspring. In these cases, the main ethical criteria to consider are non-maleficence, the ratio between risk and benefit, and consent.

In the case of interventions on germline cells, any modifications should (when technically possible) be carried out in one of two ways: by modifying the germ cells (gametes: sperm cells or oocytes) or by modifying the zygote, or embryo, at an early stage [ 15 ].

Interventions involving germline cells could potentially affect offspring. Any modification, where feasible, could, however, be carried out in one of two ways: by modifying the germ cells (gametes: sperm or oocytes) or by modifying the zygote or the embryo at an early stage [ 15 ].

Liang, Xu, Zhang, Ding and several other Chinese scientists first applied the CRISPR technique to embryos in 2015. A year later, another group of Chinese researchers (Kang, He, Huang and others) repeated the process [ 22 ]. Both experiments involved applying the technique to in vitro fertilized zygotes that were defective and unviable for reproductive implantation (because they were triploid). Neither of the projects was successful, and they were characterised by imprecision, mosaicism and numerous ‘off-target’ mutations [ 8 , 12 ].

Both projects highlighted the technical and ethical problems of genetic editing in the germline. From a technical point of view, there are basically three types of embryos that can be investigated: (1) those that are not viable or which are unsuitable for fertilization treatments; (2) viable/adequate but leftover embryos; and (3) embryos created specifically for research [ 10 ].

Genetic research is currently performed on embryos of the second and third types: embryos with a high probability of mosaicism. The specific creation of embryos for research purposes is not legally permitted in most countries [ 12 ].

However, even if access to optimal zygotes and embryos were possible, there are still certain technical obstacles that would have to be overcome. The possibility of analysing the genetically modified embryo is limited, since this analysis would have to be carried out during the subsequent in vitro culture period; this would impose limitations in terms of both technique and time. In addition, reducing the usable embryos to those obtained from in vitro fertilization processes, as opposed to being able to specifically create embryos for research, would reduce the quantity and quality of the embryos available Footnote 4 [ 23 ].

These types of technical limitations involve primary ethical issues. As previously stated, as of today, a completely safe and precise genetic alteration of the germline is effectively impossible. To achieve this, better knowledge and better technology are required, and this would call for more research to be carried out. Secondary ethical issues relate to the very use of embryos in scientific research.

There are some scientists who believe that research with human embryos is not ethically acceptable because it involves dealing with organisms, or entities, with human status or which have dignity [ 5 , 10 ]. Others, including members of the ESHRE and ESHG, argue that the embryo has a lower moral ‘status’ than the foetus which, in turn, has a lower moral ‘status’ than a child or adult. The ESHRE and ESHG have no objections to using discarded, or leftover, embryos for research [ 12 ]. Along the same lines, Savulescu et al. [ 24 ] argue that the main consideration concerning genetic experimentation on embryos (and, more specifically, the use of the CRISPR technique) is that they would not be useful in any other way.

Similarly, the ESHRE and ESHG see no problems with and have no moral objections to the creation of embryos explicitly and directly for research purposes, since their moral status is the same as that of embryos that have been left over from, or discarded during, in vitro fertilization processes [ 12 ].

There are some researchers, however, who point out that any use of embryos for research, regardless of their origin, runs the risk of falling upon a slippery slope: Accepting certain practices will easily open the door to others that are regarded as unacceptable. It should be noted that the ethical problem is not the ‘slippery slope’ itself, but rather these potentially unacceptable practices. Along these lines, several authors have suggested that the following limitations should be accepted by the entire scientific community (as has already occurred in the USA, China, the UK and Sweden): no investigating with embryos that are more than 14 days old and no implantations for reproduction (in any species) of any embryos that have already been used in research [ 10 ].

The genetic modification of germ cells consists of the application of genetic editing techniques in the process of gametogenesis: their application to original, or primordial, cells that will engender sperm and the mature ovum [ 25 ]. The female germ cell is more accessible, and susceptible, to genetic editing, but it is not yet possible to act upon the previous, or precursor, cell of the oocyte (the immature precursor of the ovum), and there is controversy in this regard [ 12 ].

Another option would be to produce in vitro gametes from pluripotential stem cells, but this technique has yet to be fully developed in animals [ 12 ]. As a result, the genetic editing of this type of cell is currently impossible in humans, even using the CRISPR technique. This should be mentioned as a research channel, however, because it is considered a possible future option.

Having already mentioned the use of embryos in experimentation, it is necessary to underline some other ethical issues that may help to configure the map of ethical conflicts associated with the genetic modification of the germline.

Although this article analyses the issue from an ethical point of view, to fully address this question, it is also important to make reference to the legal framework. From the legal perspective, at least in Europe, this matter is not subject to discussion and there is no leeway for different possibilities of implementation. This is because, on the one hand, the European Convention on Human Rights and Biomedicine (Convention of Oviedo; Article 13 (1997)) strictly prohibits any modification of the human genome that could affect their offspring and, on the other, the European regulation governing clinical trials (EU Regulation No. 536/2014) does not allow any type of genetic experimentation that could alter the subject’s germ line genetic identity [ 12 ]. However, not all European countries have ratified the Oviedo Convention [ 26 ] and, moreover, within the field of ethics, there is no general agreement amongst the scientific community as to the use that should be made of the CRISPR technique in genetic experimentation [ 27 ]. Footnote 5

Both the ESHRE and ESHG consider that no theoretical ethical objections should prevent accepting germline gene modification. They do, however, recognize the previously mentioned technical objections related to safety and precision, avoiding off-target effects and anticipating the consequences of genetic modifications on subsequent generations. Overcoming these limitations would require more research, and in the opinion of these institutions, it would be ethically unacceptable to attempt any modifications in germline genetics until this has been completed [ 12 , 27 ].

Other authors do not consider the unpredictable impact that (at least some of) these consequences could have on future generations to be sufficient, in themselves, to warrant a solid moral objection. This is, for example, a thesis supported by Sugarman and Savulescu, who argue that it is not, in fact, possible to do research into science and technology while knowing, or being able to anticipate, all of the effects that a given discovery, or technique, may have on future generations [ 24 , 28 ].

At the same time, several authors have stated that it would raise a fundamental moral problem if any alteration to the germline were to imply converting future generations into effective research participants. This would make it necessary to monitor, or even control, its development. The ‘participants’ in that control group would always need to give their informed consent which, for obvious reasons, could not possibly be obtained at the time of starting the trial. Any genetic modification to the germline that could cause effects that would be transmissible to offspring would therefore require monitoring during subsequent generations [ 29 ]. However, the reply to that objection is that such a limitation would, in fact, effectively constitute a limitation to any type of research that could directly, or significantly, affect members of future generations. It should be added that it is never possible to request the consent of all the individuals who are directly involved in research initiated by a previous generation [ 13 , 24 , 28 ]. However, in this particular case, the involvement of subjects without their consent would be direct.

Mintz [ 30 ] argued that although the embryo does not have the capacity for autonomous decision-making at the time of the germline engineering, the decision taken by the parents could affect its future autonomy. Citing Feinberg [ 31 ], he argued that to protect the autonomy of future children, parents should be helped to make ethical decisions that would give their children an open future.

The ethical debate could be summarized in relation to the four questions, or criteria, put forward by Lehmann [ 14 ] for morally assessing any genetic modification (whether in the germ line, or not). The author called these criteria the ‘4-S’: (1) S afety, (2) the S ignificance of the harm to be avoided or averted, (3) S uccessive generations and (4) S ocial consequences.

Hildt [ 23 ] argued that any attempt to genetically modify the germline should meet three conditions that do not tend to be met at present:

Previous solid and safe experience in gene therapy with somatic cells

Tests on animals that would guarantee safety and reproducibility and suggest that any subsequent human interventions would be successful

Public approval (social, political…) of the technique

The criteria mentioned in the last two points have several social implications that must be considered when evaluating the moral permissibility of genetic research. In particular, these are relevant to actions that involve changes in the germline. These social consequences are not usually included amongst the criteria considered in the literature, but we think that they have a specific weight in the moral debate.

Ethical Issues Related to Its Purpose

To expand the map of ethical issues related to genetic editing and the use of the CRISPR technique, one fundamental question within the debate is its purpose. There are, or can be, two main reasons for genetic editing:

The first is therapy: the treatment or prevention of a given disease

The second is improvement: modifying a characteristic (or several characteristics) of an individual (or species) that is (or are) not a disease

The debate and questions about these two points could be expressed in terms of the following questions: (1) Is a genetic modification ethically acceptable when performed for therapeutic purposes: to cure or prevent disease? And (2) Is it ethically acceptable to produce a change in a patient when what is modified (or improved) does not constitute or involve a disease.

Lehmann has posed the following questions related to this debate: (1) Should humans use genetic editing not only to prevent disease, but also to improve a given individual, or indeed, the whole species (for example by helping them to adapt to climate change)? (2) If this is the case, who should have the power to control, or decide, this and also how to distribute the available resources? (3) Should parents be able to decide whether to genetically modify their children (at the embryonic stage) and to take decisions that would affect them (and future generations) for as long as they live? [ 14 ].

Agar [ 32 ] suggests that we should distinguish between ‘morally wrong’ practices, which should be condemned, and ‘morally problematic’ practices, which call for ‘solutions’. According to Agar, genetic editing in order to make improvements would fall into the latter ‘problematic’ category, with the issue being complex, but not necessarily ‘wrong’.

Broadly speaking (and without entering into intermediate nuances), there are two opposing positions here: Some argue that genetic editing is only ethically acceptable to cure, or prevent, disease, while others argue that, in some cases, gene editing for human enhancement is not only acceptable, but even a moral duty.

The argument put forward by advocates of improvement is that medicine is already used to 'improve' individuals and species (for example, through vaccines and surgery). They also argue that there could even be a moral obligation to improve the human body and health, if the means are available to do so [ 14 , 24 ]. In contrast, others hold that the use of genetics or genomic modification to improve non-pathological features of individual humans (and the species as a whole) could easily lead to eugenic practices and to the value of the individual being given less importance than certain bodily and cognitive characteristics [ 5 ].

This reflection on human improvement poses important questions whose scope goes beyond genetics and the CRISPR technique but which could have a powerful influence upon them, both now and in the future. Check raises several of these questions: (1) Which diseases should be eradicated or prevented? (2) Should all disabilities be eradicated? (3) In fact, what is a disease? (4) Is it ethically and socially good to eradicate everything that is judged to be ‘defective’ (or any feature considered to be so)? [ 33 ].

Ethical Issues Related to Justice

In health care and scientific research, one of the main principles, or fundamental ethical criteria, is that of justice. This principle establishes, amongst other duties, the obligation to always consider the social consequences of technological and scientific ‘advances’. Paradoxically, these social consequences are a factor that the scientific literature does not usually consider. In the case of genetic editing techniques, in general, and the CRISPR technique, in particular, these relate to questions concerning accessibility, equality and representativeness.

The financial cost of therapy is a fundamental factor when considering the question of justice. Marketing an expensive therapy could effectively exacerbate existing inequalities in health provision, in both poor and rich countries (an example is the USA, with its marked inequalities in the reception of health) [ 34 , 35 , 36 ]. On the other hand, in the public health systems of Western countries, the problem would be the impact of its cost for the health system, which should not threaten the sustainability of the whole system. Such inequalities could, as Bellver [ 5 ] has pointed out, cause a genetic alteration of the germline that would potentially generate excessive inequalities between different individuals, communities and societies, giving rise to what has been referred to as aristocracy genetics.

Likewise, the inclusion and representativeness of all sectors of the population is also important, as in the case of minorities (such as different ethnic groups) [ 35 ]. When engaging in debate and considering the scope of such hypothetical genetic alterations, the diversity of the genetic sequencing found in certain individuals, or groups, within the human population should not be placed in jeopardy.

The CRISPR genetic editing technique is a technological procedure that, while improving on previous approaches, is not, in itself, exempt from certain ethical problems (some of which are technical). The scientific literature has already focused on some of these, but there is still a relative lack of publications highlighting the full range of this problem. It can, however, be concluded that the map of ethical problems associated with this technique should consider the following points:

Efficiency and safety: Efficiency and safety are two considerations that have not, to date, received as much attention as might be expected.

Cell type: The types of cells that are, or may be used, in this technique, and the applications that it may be given, are the subject of a moral debate. The ethical questions related to somatic cell interventions are not the same as those associated with germ cell interventions. The use of embryos should also be debated in a similar way (whether it is ethical for them to be subject to research, or the application of the technique, and which types of embryos can be used, etc.).

Purpose: From the perspective of morality, the implications are not the same when the purpose of using the technique is associated with therapy to treat certain diseases, as when the objective is to improve certain aspects, or traits, that are not considered ‘pathological’ (whether these apply to individuals, or to the human species as a whole).

Justice: It is also important to consider the social impact that extending the application of the technique could have regarding accessibility, costs, possible inequalities and the position of ethnic or social minorities.

Rosenbaum outlined the need to establish a social consensus [ 37 ] and to perhaps declare a moratorium. The conceptualization of a map of ethical conflicts, of the type presented in this article, clearly defines the different problem areas, helps to define a reference framework for debate and establishes positions and a base for subsequent legal developments.

It should be pointed out that the technique had already been successfully used, but that had been for editing DNA [ 2 ].

An ‘off-target’ effect is an unforeseen mutation that may modify the phenotype [ 9 ]. This is the consequence of the non-specific activity of the Cas nuclease at a non-targeted location ‘in’ the genome [ 27 ].

Mosaicism is a genetic alteration, whereby cells with different genotypes coexist in the same organism. In embryos, they arise due to a fault in the nuclease ruptures. They are the result of either the imprecise reparation of DNA before the embryo has begun its cell-division stage [ 12 ] or the onset of cell-division preceding any form of genetic editing [ 27 ].

For example, Savulescu points out that hundreds, or even thousands, of embryos are needed to run the experiments required for research into polygenic diseases [ 6 ].

Hence, in July 2017, Chneiweiss, Hirsch, Montoliu and other researchers (representing over 20 institutions) proposed creating a European Steering Committee to analyse and evaluate the advantages and dangers of the new technique [ 23 ].

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Grup Investigació en Bioètica (GIB): David Lorenzo 1,4 , Montse Esquerda 1,5 , Margarita Bofarull 1 , Helena Roig 1 , Victoria Cusí 1 , Francisco J. Cambra 1,3 , Joan Carrera 1 , Francesc Palau 2,3

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Lorenzo, D., Esquerda, M., Palau, F. et al. Ethics and Genomic Editing Using the Crispr-Cas9 Technique: Challenges and Conflicts. Nanoethics 16 , 313–321 (2022). https://doi.org/10.1007/s11569-022-00425-y

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Targeted genome-modification tools and their advanced applications in crop breeding

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• 1 New Cornerstone Science Laboratory, Center for Genome Editing, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, Beijing, China.
• 2 College of Advanced Agricultural Sciences, University of Chinese Academy of Sciences, Beijing, China.
• 3 Hainan Yazhou Bay Seed Laboratory, Sanya, China.
• 4 State Key Laboratory of Plant Genomics, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, Beijing, China.
• 5 New Cornerstone Science Laboratory, Center for Genome Editing, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, Beijing, China. [email protected].
• 6 College of Advanced Agricultural Sciences, University of Chinese Academy of Sciences, Beijing, China. [email protected].
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• DOI: 10.1038/s41576-024-00720-2

Crop improvement by genome editing involves the targeted alteration of genes to improve plant traits, such as stress tolerance, disease resistance or nutritional content. Techniques for the targeted modification of genomes have evolved from generating random mutations to precise base substitutions, followed by insertions, substitutions and deletions of small DNA fragments, and are finally starting to achieve precision manipulation of large DNA segments. Recent developments in base editing, prime editing and other CRISPR-associated systems have laid a solid technological foundation to enable plant basic research and precise molecular breeding. In this Review, we systematically outline the technological principles underlying precise and targeted genome-modification methods. We also review methods for the delivery of genome-editing reagents in plants and outline emerging crop-breeding strategies based on targeted genome modification. Finally, we consider potential future developments in precise genome-editing technologies, delivery methods and crop-breeding approaches, as well as regulatory policies for genome-editing products.

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A small factor makes a big impact on genome editing

by Caitlin Sedwick, Princeton University

Through years of engineering gene-editing systems, researchers have developed a suite of tools that enable the modification of genomes in living cells, akin to "genome surgery." These tools, including ones based on a natural system known as CRISPR/Cas9, offer enormous potential for addressing unmet clinical needs, underscored by the recent FDA approval of the first CRISPR/Cas9-based therapy.

A relatively new approach called "prime editing" enables gene-editing with exceptional accuracy and high versatility, but has a critical tradeoff: variable and often low efficiency of edit installation. In other words, while prime edits can be made with high precision and few unwanted byproducts, the approach also often fails to make those edits at reasonable frequencies.

In a paper that appeared in print in the journal Nature on April 18, 2024 , Princeton scientists Jun Yan and Britt Adamson, along with several colleagues, describe a more efficient prime editor.

Prime editing systems minimally consist of two components: a modified version of the protein element of CRISPR/Cas9 and a ribonucleic acid (RNA) molecule called a pegRNA. These components work together in several coordinated steps: First, the pegRNA binds the protein and guides the resulting complex to a desired location in the genome.

There, the protein nicks the DNA and, using a template sequence encoded on the pegRNA, "reverse transcribes" an edit into the genome nearby. In this way, prime editors "write" exact sequences into targeted DNA.

"Prime editing is such an incredibly powerful genome editing tool because it gives us more control over exactly how genomic sequences are changed," Adamson said.

At the outset of their study, Adamson and Yan, a graduate student in Adamson's research group and the Department of Molecular Biology, reasoned that unknown cellular processes may aid or hinder prime editing. To identify such processes, Yan laid out a conceptually simple plan: First, he would engineer a cell line that would emit green fluorescence when certain prime edits were installed. Then, he would systematically block expression of proteins normally expressed within those cells and measure editing-induced fluorescence to determine which of those proteins impact prime editing.

By executing this plan, the team identified 36 cellular determinants of prime editing, only one of which—the small RNA-binding protein La—promoted editing.

"Although promoting prime editing is obviously not a normal function of the La protein, our experiments showed that it can strongly facilitate the process," Yan said.

Within cells, La is known to bind specific sequences often found at the ends of nascent small RNA molecules and it protects those RNAs from degradation. The Princeton team recognized right away that the pegRNAs deployed in Yan's first experiments likely contained those exact sequences, called polyuridine tracts, as they are a typical but often overlooked byproduct of pegRNA expression in cells. Subsequent experiments suggested that such pegRNAs inadvertently harness La's end-binding activity for protection and to promote prime editing.

Motivated by their results, the team asked if fusing the part of La that binds polyuridine tracts to a standard prime editing protein could boost prime editing efficiencies. They were thrilled to find that the resulting protein , which they call PE7, substantially enhanced intended prime editing efficiencies across conditions and, when using some prime editing systems, left the frequencies of unwanted byproducts very low.

Their results quickly drew the attention of colleagues interested in using prime editing in primary human cells, including Daniel Bauer at Boston Children's Hospital and Harvard Medical School and Alexander Marson at the University of California, San Francisco. Together with scientists from these labs, the team of researchers went on to demonstrate that PE7 can also enhance prime editing efficiencies in therapeutically relevant cell types, offering expanded promise for future clinical applications.

"This work is a beautiful example of how deeply probing the inner workings of cells can lead to unexpected insights that may yield near-term biomedical impact," Bauer noted.

Journal information: Nature

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Generative A.I. Arrives in the Gene Editing World of CRISPR

Much as ChatGPT generates poetry, a new A.I. system devises blueprints for microscopic mechanisms that can edit your DNA.

The physical structure of OpenCRISPR-1, a gene editor created by A.I. technology from Profluent. Credit... Video by Profluent Bio

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By Cade Metz

Has reported on the intersection of A.I. and health care for a decade.

• April 22, 2024

Generative A.I. technologies can write poetry and computer programs or create images of teddy bears and videos of cartoon characters that look like something from a Hollywood movie.

Now, new A.I. technology is generating blueprints for microscopic biological mechanisms that can edit your DNA, pointing to a future when scientists can battle illness and diseases with even greater precision and speed than they can today.

Described in a research paper published on Monday by a Berkeley, Calif., startup called Profluent, the technology is based on the same methods that drive ChatGPT, the online chatbot that launched the A.I. boom after its release in 2022 . The company is expected to present the paper next month at the annual meeting of the American Society of Gene and Cell Therapy.

Much as ChatGPT learns to generate language by analyzing Wikipedia articles, books and chat logs, Profluent’s technology creates new gene editors after analyzing enormous amounts of biological data, including microscopic mechanisms that scientists already use to edit human DNA.

These gene editors are based on Nobel Prize-winning methods involving biological mechanisms called CRISPR. Technology based on CRISPR is already changing how scientists study and fight illness and disease , providing a way of altering genes that cause hereditary conditions, such as sickle cell anemia and blindness.

Previously, CRISPR methods used mechanisms found in nature — biological material gleaned from bacteria that allows these microscopic organisms to fight off germs.

“They have never existed on Earth,” said James Fraser, a professor and chair of the department of bioengineering and therapeutic sciences at the University of California, San Francisco, who has read Profluent’s research paper. “The system has learned from nature to create them, but they are new.”

The hope is that the technology will eventually produce gene editors that are more nimble and more powerful than those that have been honed over billions of years of evolution.

On Monday, Profluent also said that it had used one of these A.I.-generated gene editors to edit human DNA and that it was “open sourcing” this editor, called OpenCRISPR-1. That means it is allowing individuals, academic labs and companies to experiment with the technology for free.

A.I. researchers often open source the underlying software that drives their A.I. systems , because it allows others to build on their work and accelerate the development of new technologies. But it is less common for biological labs and pharmaceutical companies to open source inventions like OpenCRISPR-1.

Though Profluent is open sourcing the gene editors generated by its A.I. technology, it is not open sourcing the A.I. technology itself.

The project is part of a wider effort to build A.I. technologies that can improve medical care. Scientists at the University of Washington, for instance, are using the methods behind chatbots like OpenAI’s ChatGPT and image generators like Midjourney to create entirely new proteins — the microscopic molecules that drive all human life — as they work to accelerate the development of new vaccines and medicines.

(The New York Times has sued OpenAI and its partner, Microsoft, on claims of copyright infringement involving artificial intelligence systems that generate text.)

Generative A.I. technologies are driven by what scientists call a neural network , a mathematical system that learns skills by analyzing vast amounts of data. The image creator Midjourney, for example, is underpinned by a neural network that has analyzed millions of digital images and the captions that describe each of those images. The system learned to recognize the links between the images and the words. So when you ask it for an image of a rhinoceros leaping off the Golden Gate Bridge, it knows what to do.

Profluent’s technology is driven by a similar A.I. model that learns from sequences of amino acids and nucleic acids — the chemical compounds that define the microscopic biological mechanisms that scientists use to edit genes. Essentially, it analyzes the behavior of CRISPR gene editors pulled from nature and learns how to generate entirely new gene editors.

“These A.I. models learn from sequences — whether those are sequences of characters or words or computer code or amino acids,” said Profluent’s chief executive, Ali Madani, a researcher who previously worked in the A.I. lab at the software giant Salesforce.

Profluent has not yet put these synthetic gene editors through clinical trials, so it is not clear if they can match or exceed the performance of CRISPR. But this proof of concept shows that A.I. models can produce something capable of editing the human genome.

Still, it is unlikely to affect health care in the short term. Fyodor Urnov, a gene editing pioneer and scientific director at the Innovative Genomics Institute at the University of California, Berkeley, said scientists had no shortage of naturally occurring gene editors that they could use to fight illness and disease. The bottleneck, he said, is the cost of pushing these editors through preclinical studies, such as safety, manufacturing and regulatory reviews, before they can be used on patients.

But generative A.I. systems often hold enormous potential because they tend to improve quickly as they learn from increasingly large amounts of data. If technology like Profluent’s continues to improve, it could eventually allow scientists to edit genes in far more precise ways. The hope, Dr. Urnov said, is that this could, in the long term, lead to a world where medicines and treatments are quickly tailored to individual people even faster than we can do today.

“I dream of a world where we have CRISPR on demand within weeks,” he said.

Scientists have long cautioned against using CRISPR for human enhancement because it is a relatively new technology that could potentially have undesired side effects, such as triggering cancer, and have warned against unethical uses, such as genetically modifying human embryos.

This is also a concern with synthetic gene editors. But scientists already have access to everything they need to edit embryos.

“A bad actor, someone who is unethical, is not worried about whether they use an A.I.-created editor or not,” Dr. Fraser said. “They are just going to go ahead and use what’s available.”

Cade Metz writes about artificial intelligence, driverless cars, robotics, virtual reality and other emerging areas of technology. More about Cade Metz

Explore Our Coverage of Artificial Intelligence

News  and Analysis

Saudi Arabia is plowing money into glitzy events, computing power and artificial intelligence research, putting it in the middle of an escalating  U.S.-China struggle for technological influence.

Microsoft gave more signs that its hefty investments in A.I.  were beginning to bear fruit, as it reported a 17 percent jump in revenue and a 20 percent increase in profit for the first three months of the year.

Meta projected that revenue for the current quarter  would be lower than what Wall Street anticipated and said it would spend billions of dollars more on its artificial intelligence efforts, even as it reported robust revenue and profits for the first three months of the year.

The Age of A.I.

A new category of apps promises to relieve parents of drudgery, with an assist from A.I . But a family’s grunt work is more human, and valuable, than it seems.

Despite Mark Zuckerberg’s hope for Meta’s A.I. assistant to be the smartest , it struggles with facts, numbers and web search.

Much as ChatGPT generates poetry, a new A.I. system devises blueprints for microscopic mechanisms  that can edit your DNA.

Could A.I. change India’s elections? Avatars are addressing voters by name, in whichever of India’s many languages they speak. Experts see potential for misuse  in a country already rife with disinformation.

Which A.I. system writes the best computer code or generates the most realistic image? Right now, there’s no easy way to answer those questions, our technology columnist writes .

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Targeted genome-modification tools and their advanced applications in crop breeding

• Boshu Li 1 , 2   na1 ,
• Chao Sun 1 , 2   na1 ,
• Jiayang Li   ORCID: orcid.org/0000-0002-0487-6574 3 , 4 &
• Caixia Gao   ORCID: orcid.org/0000-0003-3169-8248 1 , 2  

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Crop improvement by genome editing involves the targeted alteration of genes to improve plant traits, such as stress tolerance, disease resistance or nutritional content. Techniques for the targeted modification of genomes have evolved from generating random mutations to precise base substitutions, followed by insertions, substitutions and deletions of small DNA fragments, and are finally starting to achieve precision manipulation of large DNA segments. Recent developments in base editing, prime editing and other CRISPR-associated systems have laid a solid technological foundation to enable plant basic research and precise molecular breeding. In this Review, we systematically outline the technological principles underlying precise and targeted genome-modification methods. We also review methods for the delivery of genome-editing reagents in plants and outline emerging crop-breeding strategies based on targeted genome modification. Finally, we consider potential future developments in precise genome-editing technologies, delivery methods and crop-breeding approaches, as well as regulatory policies for genome-editing products.

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Applications of CRISPR–Cas in agriculture and plant biotechnology

Genome editing for horticultural crop improvement

Recent advances in crop transformation technologies

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Acknowledgements

This work was supported by the National Natural Science Foundation of China (32388201), the National Key Research and Development Program (2022YFF1002802), the Ministry of Agriculture and Rural Affairs of China, the Strategic Priority Research Program of the Chinese Academy of Sciences (Precision Seed Design and Breeding, XDA24020102), and the New Cornerstone Science Foundation. The authors thank K. T. Zhao, C. Xue, R. Liang, G. Liu, J. Hu, H. Li, Y. Li, F. Qiu, S. Li, Y. Lei and X. Jiang for their insightful comments on the manuscript.

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New Cornerstone Science Laboratory, Center for Genome Editing, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, Beijing, China

Boshu Li, Chao Sun & Caixia Gao

College of Advanced Agricultural Sciences, University of Chinese Academy of Sciences, Beijing, China

Hainan Yazhou Bay Seed Laboratory, Sanya, China

State Key Laboratory of Plant Genomics, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, Beijing, China

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B.L., C.S. and C.G. researched the literature. All authors substantially contributed to discussions of the content and wrote the article. J.L. and C.G. reviewed and/or edited the manuscript before submission.

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A bacterium used for plant delivery. It can induce the formation of hairy roots in the infection site. It contains a root-inducing plasmid that carries a T-DNA segment capable of integrating into the plant genome. Typically, this T-DNA harbours the desired sequences intended for transfer into the plant genome.

A bacterium used for plant delivery. It contains a modified tumour-inducing plasmid that carries a T-DNA segment capable of integrating into the plant genome. Typically, this T-DNA harbours the desired sequences intended for transfer into the plant genome, as well as marker genes for selecting positive events.

(CRISPRi). CRISPR interference utilizes dCas9 either alone or with a transcription repressor to inhibit gene expression by targeting specific DNA sequences without altering the genetic code, offering precise control for studying gene functions and regulatory processes within cells.

(gRNA). An RNA molecule used to direct Cas9 or similar enzymes to a specific DNA or RNA sequence for precise modification.

A phenomenon in which the offspring of two different inbred lines or varieties exhibit improved traits compared to their parents, such as increased yield, growth or biotic or abiotic resistance. Also known as heterosis.

The DNA strand that is not complementary to the guide RNA sequence. DNA nicking by PE2 and base deamination by base editors occur on the non-targeted DNA strand.

A genetic transformation technique, also known as gene gun or biolistic delivery, that involves loading exogenous DNA onto microscopic metal particles that are accelerated and propelled into plant cells or other target cells by compressed gas or physical force.

(PAM). A short DNA sequence immediately adjacent to the target site that is essential for the recognition and binding of Cas protein to the target DNA.

A specific structure consisting of one DNA strand, its complementary DNA strand and an RNA strand located between them.

The DNA strand that is complementary to the guide RNA sequence. DNA nicking by base editors occurs on the targeted DNA strand.

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Li, B., Sun, C., Li, J. et al. Targeted genome-modification tools and their advanced applications in crop breeding. Nat Rev Genet (2024). https://doi.org/10.1038/s41576-024-00720-2

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• Study from Phoenix Children’s Research Institute Reveals New Way to Prevent Lung Cancer from Spreading

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PHOENIX [April 25, 2024] – Research conducted at the Phoenix Children’s Research Institute at the University of Arizona College of Medicine — Phoenix shows normalizing cancer tumor vessels and alleviating low oxygen levels in the tumor microenvironment can improve the effectiveness of chemotherapy treatment in lung cancer, according to a paper published in EMBO Molecular Medicine .

This study, authored by Tanya Kalin, MD, PhD, vice chair of translational research for the Phoenix Children’s Center for Cancer and Blood Disorders and professor of Child Health at the University of Arizona College of Medicine — Phoenix, ”FOXF1 Promotes Vessel Normalization and Prevents Lung Cancer Progression Through FZD4, is a complete pivot from how researchers have historically studied cancer treatment by providing a solution that strengthens blood vessels feeding cancer tumors.

“Cancer research is often focused on ways to deplete cancer tumors of blood vessels and oxygen by basically starving the cancer cells to prevent tumor growth and metastasis, but it was shown this approach can make the tumor cells become even more aggressive and metastatic,” said Dr. Kalin. “Our study takes a completely different approach and instead of obliterating the tumor-associated blood vessels, we normalize and repair the tumor vessels by increasing the FOXF1 protein in endothelial cells, which, in turn, prevents lung cancer progression.”

Dr. Kalin, a renowned scientist focused on developing effective treatments for pediatric cancers, has spent the last few decades researching how to create a non-toxic small molecule inhibitor compound to kill cancer cells and developing tumor-cell-specific nanoparticles that deliver the inhibitor compound directly to cancer cells, blocking specific targeted proteins. This lung cancer study builds upon her years of research to develop a more targeted approach to cancer treatment. The insertion of the FOXF1 protein increases tumor vessel stability and stimulates nanoparticle delivery into the cancer-causing cells which destroys them from the inside with minimal chemotherapy.

“This is a promising study for future therapies in non-small cell lung cancer and other types of cancer and will hopefully change how we look at cancer treatment moving forward,” said Dr. Kalin.

Lung cancer is the leading cause of cancer-related mortality worldwide. Current treatment strategies include chemotherapy and/or radiotherapy, and surgery in the case of patients diagnosed with early-stage lung cancer. The 5-year survival rate of patients with advanced Non-Small Cell Lung Cancer (NSCLC) remains less than 20 percent, emphasizing a need to develop better treatment strategies. Interactions between the tumor-microenvironment and tumor cells play a crucial role in tumor progression and to this point traditional therapies have failed to improve overall survival of lung cancer patients, suggesting a deeper understanding of tumor-associated vascular biology is required.

“Every breakthrough in pediatric cancer research is cause for celebration, especially studies like Dr. Kalin’s that shed light on tumor microenvironments and stopping the spread of cancer,” said Stewart Goldman, MD, senior vice president of research for Phoenix Children’s and Sybil B. Harrington endowed chair and professor of Child Health at University of Arizona College of Medicine — Phoenix. “Dr. Kalin’s remarkable findings will have implications for non-small cell lung cancer, as well as many childhood cancers, hopefully changing cancer therapies in the near term and I believe this research will be one of the most cited cancer research studies for years to come.”

In March, Phoenix Children’s released another study, “CRISPR/Cas9 Genome Editing Allows Generation of the Mouse Lung in a Rat,” that focuses on finding an innovative solution for babies born with chronic lung diseases caused by either prematurity or severe genetic conditions.

The Phoenix Children's Research Institute at the University of Arizona College of Medicine — Phoenix launched in May 2023, formalizing a longstanding research collaboration between the health system and the University of Arizona College of Medicine — Phoenix. The Research Institute includes more than 700 active studies, 640 research investigators and 90 research staff members including research scientists, associates, biostatisticians, pharmacists, nurses and coordinators. Scientists engage in research across multiple clinical disciplines including cancer , neurology , cardiology , pulmonology and more.

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