Section 2 Technology

#### **Chapter 3**

## Maximizing the Efficacy of CRISPR/Cas Homology-Directed Repair Gene Targeting

*Terry S. Elton, Md. Ismail Hossain, Jessika Carvajal-Moreno, Xinyi Wang, Dalton J. Skaggs and Jack C. Yalowich*

#### **Abstract**

Clustered regularly interspaced short palindromic repeats/CRISPR-associated system (CRISPR/Cas) is a powerful gene editing tool that can introduce doublestrand breaks (DSBs) at precise target sites in genomic DNA. In mammalian cells, the CRISPR/Cas-generated DSBs can be repaired by either template-free error-prone end joining (e.g., non-homologous end joining/microhomology-mediated end joining [NHEJ]/[MMEJ]) or templated error-free homology-directed repair (HDR) pathways. CRISPR/Cas with NHEJ/MMEJ DNA repair results in various length insertions/ deletion mutations (indels), which can cause frameshift mutations leading to a stop codon and subsequent gene-specific knockout (i.e., loss of function). In contrast, CRISPR/Cas with HDR DNA repair, utilizing an exogenous repair template harboring specific nucleotide (nt) changes, can be employed to intentionally edit out or introduce mutations or insertions at specific genomic sites (i.e., targeted gene knock-in). This review provides an overview of HDR-based gene-targeting strategies to facilitate the knock-in process, including improving gRNA cleavage efficiency, optimizing HDR efficacy, decreasing off-target effects, suppressing NHEJ/MMEJ activity, and thus expediting the screening of CRISPR/Cas-edited clonal cells.

**Keywords:** CRISPR/Cas, homology-directed repair, gene editing, Cas9, Cas12, non-homologous end joining, microhomology-mediated end joining, knock-in

#### **1. Introduction**

Clustered regularly interspaced short palindromic repeats/CRISPR-associated system (CRISPR/Cas) technology has revolutionized biological research and holds great therapeutic potential, since it is remarkably flexible and reliable [1–3]. CRISPR/Cas genome editing (i.e., genetic engineering) is a programmable technology to introduce double-strand breaks (DSBs) at specific target sites in the genome of a living organism [1–3]. There are two major mechanisms by which Cas enzyme-mediated DSBs are subsequently repaired [4–6]. The first is by template-free end joining (e.g., nonhomologous end joining/microhomology-mediated end joining [NHEJ]/[MMEJ]), which introduces insertions/deletion mutations (indels) and can lead to targeted gene knock outs. The second mechanism is *via* the homology-directed repair (HDR) pathway, which produces a targeted gene knock-in or other specific mutations utilizing an exogenous donor template [4–6]. Given that DSBs generated in mammalian cells are predominantly repaired by NHEJ or MMEJ, the rate of precise editing through CRISPR/Cas/HDR with an exogenous repair template is significantly compromised/ reduced [4–6]. This review summarizes multiple strategies to enhance the efficacy of CRISPR/Cas/HDR as well as decrease off-target effects.

#### **2. CRISPR/Cas history**

CRISPR history began in 1987 when Ishino et al. [7–9] first observed five repetitive palindromic sequences of 29 nucleotides separated by random 32 nucleotides toward the end of the *E. coli* genome. Although Ishino et al. [7] did not decipher the biological significance of the puzzling repeat sequences, this report led to the discovery of similar patterns in other bacterial and archaea genomes [10–12]. Mojica et al. [13] then established that the unusual repetitive DNA sequences were functionally related. These curious sequences were later designated "CRISPR" by Jansen et al. [14] given that these loci harbor: 1) palindromic repeats with little sequence variation; 2) nonrepetitive spacer sequences between the repeats; and 3) a several hundred base pair (bp) common leader sequence on one side of the repeat cluster. The CRISPR locus is present in approximately 40% of the sequenced bacteria and 90% of the genomes of the different domains of archaea [15]. Finally, it was demonstrated that CRISPRassociated (Cas) genes (i.e., over 40), of which only a subset is found in any given prokaryote that harbor CRISPRs, are frequently located in close proximity to CRISPR loci [14, 16, 17]. The Cas genes were predicted to encode endo- and exonucleases, helicases, polymerases, and RNA-binding proteins [14, 15, 17].

Initially, CRISPR/Cas systems were expected to have a role in DNA repair or gene regulation due to their location near the DNA repair system in the bacterial genome [18]. However, in 2005, three seminal studies revealed that the CRISPR spacer sequences were homologous to bacteriophage, prophages, and conjugative plasmid sequences and suggested that they were the remnants of past invasions by extrachromosomal elements [19–21]. These investigators further speculated that there was a relationship between CRISPR and immunity against foreign DNAs by coding an anti-sense RNA [19–21]. In the following year, Makrova et al. [17] analyzed the link between the CRISPR and the Cas proteins and how this system is similar to the prokaryotic RNAi-mediated adaptive immune system, which led them to propose that the CRISPR/Cas system, with its "memory component," may function as inheritable adaptive immunity for bacteria.

Subsequently, Barrangou et al. [22] demonstrated that after a viral challenge, phage sequence was integrated into a CRISPR locus of *Streptococcus thermophilus* and provided immunity against the corresponding phage. When the protospacer sequence was deleted from the bacterial genome, they became sensitive to phage infection [22]. These investigators hypothesized that the nucleic acid based "immunity" system in prokaryotes was dictated by the CRISPR spacer sequence and that the Cas protein machinery mediated resistance against foreign DNAs [22].

In 2008, a pivotal study by Brouns et al. [23] established that the *E. coli* spacer sequences were transcribed into a precursor CRISPR RNA (pre-crRNA) that was matured to small crRNAs by a complex of Cas proteins. Additionally, it was demonstrated that mature crRNAs serves as a "guide" to a direct a protein to target viral

#### *Maximizing the Efficacy of CRISPR/Cas Homology-Directed Repair Gene Targeting DOI: http://dx.doi.org/10.5772/intechopen.109051*

nucleic acids, which results in an antiviral response in prokaryotes [23]. Subsequently, Mojica et al. [24] identified CRISPR-type-specific proto-spacer adjacent motifs (PAMs), which are important for discrimination between self and nonself sequences. Further, Garneau et al. [25] showed that CRISPR/Cas immunity resulted from the generation of DSBs at specific sites in bacteriophage and plasmid DNA. Finally, Sapranauskas et al. [26] demonstrated that the *S. thermophilus* CRISPR/Cas system could be transferred to *E. coli* and provide a Cas9-mediated immunity that required a PAM site. Their initial characterization of the Cas9 protein revealed that two domains are involved in the formation of DBSs [26]. The Cas9 McrA/HNH-like nuclease domain cleaves the DNA strand complementary to the guide RNA sequence (target strand), and the RuvC/RNaseH-like domain cleaves the noncomplementary strand (nontarget strand) (**Figure 1**) [26]. They also demonstrated that a 20-nucleotide crRNA, the trimmed version of the full-length crRNA, is sufficient for DNA target identification with efficient cleavage and that the target site of the Cas can be changed by changing the crRNA sequence (**Figure 1**) [26].

Next, Deltcheva et al. [27] discovered an additional small RNA designated the trans-activating CRISPR RNA (tracrRNA). This small RNA is transcribed from

#### **Figure 1.**

*Schematic representation of the CRISPR/Cas9-mediated DSB with a two-piece gRNA. The Cas9 gRNA is a two-piece RNA complex comprised of a crRNA required for DNA targeting (denoted in green) and the tracrRNA, which is necessary for nuclease activity (denoted in red) [27, 28]. The Cas9 protein (denoted in yellow) binds to the gRNA to form a RNP complex. The gRNA directs the Cas9 to a specific location in the genomic DNA (denoted in blue) through a user-defined 20 nt sequence at the 5' end of the crRNA, which is complementary to the target DNA (denoted by green and blue hash marks). If there is a PAM site (NGG) (denoted in pink) adjacent to the 3' end of the 20 nt sequence, then the Cas9 McrA/HNH-like nuclease domain (denoted in peach) cleaves the DNA strand complementary to the guide RNA sequence (target strand) and the RuvC/RNaseH-like domain (denoted in orange) cleaves the noncomplementary strand (nontarget strand) to introduce site-specific DSBs in the target DNA [26].*

sequence upstream of the CRISPR-Cas locus of *Streptococcus pyogenes* [27]. These investigators demonstrated that, upon maturation of both the tracrRNA and the crRNA, they form a duplex that has both single- and double-stranded regions (**Figure 1**) [27]. Furthermore, Jinek et al. [28] verified that the two-RNA complex, dual RNA (i.e., the crRNA [required for DNA targeting] and the tracrRNA [necessary for nuclease activity]; now designated as a guide-RNA [gRNA]) directs the Cas9 to introduce site-specific DSBs in the target DNA (**Figure 1**). They also demonstrated that Cas9 target recognition required complementary seed sequences between the crRNA and target DNA as well as a PAM sequence containing a GG dinucleotide adjacent to the crRNA-binding region in the DNA target (**Figure 1**) [28]. Moreover, Jinek et al. [28] established that the *S. pyogenes* Cas9 endonuclease could be programmed to target and cleave any dsDNA sequence, which harbors a NGG (N denotes any nt) PAM site, with an engineered gRNA which contains a 20-nucleotide crRNA sequence that is complementary to the target DNA (**Figure 1**). CRISPR-Cas technology is now widely adopted in the scientific community due to its simplicity and precision for gene editing, which has opened the possibility of numerous applications in the field of genetic engineering.

#### **3. DNA double-strand break repair**

Pathological DNA DSBs can arise from normal endogenous metabolic cellular processes (e.g., DNA replication and transcription) or from cellular exposure to exogenous sources (e.g., reactive oxygen species, ionizing radiation, radiomimetic chemicals, and anticancer chemotherapeutic drugs) [29–32]. However, physiologically important DNA DSBs are also required for several developmental and physiological cellular activities including chromosomal disjunction, meiosis, V(D)J, and immunoglobulin heavy chain (IgH) class switch recombination [29, 33]. Notably, both pathological and physiological DNA DSBs require efficient repair processes since these lesions can result in insertions, deletions, chromosomal translocations, and genomic instability, which can lead to numerous hereditary human diseases, including cancer, developmental disorders, and premature aging [29–31, 33]. Mammalian cells employ multiple DNA repair pathways to protect the integrity of their genomes. However, the two predominate DNA DSB repair pathways that are template-free NHEJ/MMEJ and templated HDR [32, 34, 35]. It is important to note that NHEJ and HDR are two competing pathways [4–6]. In mammalian cells, template-free NHEJ is favored over templated HDR since NHEJ is a rapid high-capacity pathway, which is active throughout the cell cycle and directly represses HDR [4–6]. In contrast, HDR is largely restricted to the S and G2 phases [4–6].

At the most basic level, the CRISPR/Cas genome editing technology is utilized to introduce a DSB at a specific target site in the genome and then relies upon the cellular machinery to repair this lesion by either the NHEJ/MMEJ or HDR repair pathways to yield the desired repair outcomes [32, 34, 35]. If the experimental goal is to knockout the function of a given gene of interest, then the error-prone NHEJ or MMEJ pathways would be utilized to repair DNA DSBs created by the Cas endonuclease at a programmed target site to introduce indels, which can shift the open reading frame (ORF) and result in targeted gene loss of function [32, 34, 35]. In contrast, if the experimental goal is to edit out or to introduce mutations at specific genomic sites (i.e., targeted gene knock-in), then the HDR pathway would be utilized to repair the Cas endonuclease-created DNA DSBs with an exogenous repair template harboring

specific nucleotide (nt) changes [32, 34, 35]. The cellular NHEJ/MMEJ and HDR repair pathways of endogenous and CRISPR/Cas-generated DSBs will be discussed in more detail below.

#### **3.1 Template-free error-prone end joining NHEJ/MMEJ pathways**

Non-homologous end joining (NHEJ) rejoins DNA DSBs as quickly as 30 minutes after break induction with minimal processing [4–6]. Briefly, after a DSB (i.e., DNA ends can be either blunt or possess a short 5′ overhang) has formed, the ring-shaped XRCC6 (X-ray repair cross complementing 6, also known as Ku70)/XRCC5 (X-ray repair cross complementing 5, also known as the Ku80) protein heterodimer quickly binds to the broken DNA ends [4–6]. This binding protects the DNA ends from further resection, preventing MMEJ and HDR pathway initiation [36, 37]. The XRCC6/XRCC5 heterodimer (Ku) then recruits and activates the catalytic subunit of the DNA-dependent protein kinase (DNA-PKcs) [38, 39]. The XRCC6/XRCC5 heterodimer subsequently recruits additional NHEJ factors including XRCC4 (X-ray repair cross complementing 4), NHEJ1 (non-homologous end joining factor 1, also known as XLF), and (DNA ligase IV) to the complex to ligate the DNA DSB ends [40]. Therefore, in the absence of DNA end processing, NHEJ-mediated repair is error-free [40]. In contrast, if the DSB ends are not ligatable due to nucleotide overhangs, DCLRE1C (DNA cross-link repair 1C, also known as Artemis), a singlestrand-specific 5′ → 3′ exonuclease, and specialized DNA polymerases POLL (DNA polymerase) and POLM (DNA polymerase μ) generate compatible DNA blunt ends, which can then be ligated by LIG4 [41]. Importantly, this process limits DNA end processing and minimizes mutagenesis (i.e., indels) [41].

#### **3.2 Microhomology-mediated end joining (MMEJ)**

Although it was originally thought that most CRISPR/Cas-generated DNA DSBs were repaired by the NHEJ pathway [4–6], it is now apparent that a significant number of these DSBs are also fixed by the MMEJ pathway (>50%) [42, 43]. MMEJ, like NHEJ, does not require a template for repairing DNA DSBs [4–6]. However, in contrast to NHEJ, MMEJ begins with a short-range resection of the DNA DSBs and functions independently of XRCC6/XRCC5 and LIG4 [44]. MMEJ resection is initiated by the MRN (i.e., MRE11 [MRE11 homolog double-strand break repair nuclease]-RAD50 [RAD50 double-strand break repair protein]-NBN [Nibrin, also known as NBS]) DNA DSB repair damage sensing complex with its stimulatory factor RBBP8 (RB-binding protein 8 endonuclease, also known as CTIP) [4–6, 44]. RBBP8 phosphorylation stimulates MRE11 endonuclease activity to create a nick at the 5′ strand near to the DSB, which promotes the removal of XRCC6/XRCC5 and DNA-PKcs, thus preventing NHEJ [45–47]. The resulting nick allows the MRE11 3′-to-5′ exonuclease to resect back toward the DNA DSB, which generates short 3′ overhangs, thus exposing potential single-strand DNA microhomologies (5–25 bps) on opposite strands, which allows the broken ends to realign and anneal [4–6, 44]. Any resulting heterologous 3′ single-stranded DNA (ssDNA) flaps must be removed by the ERCC1 (ERCC excision repair 1 endonuclease non-catalytic subunit)/ERCC4 (ERCC excision repair 4 endonuclease catalytic subunit, also known as XPF) endonuclease [48]. POLQ (DNA polymerase theta) is recruited to stabilize the annealed ssDNA and fills any gaps *via* template-directed DNA synthesis. LigI (DNA ligase 1) or LigIII (DNA ligase 3) subsequently seals the break [49]. Importantly, due to the resection

step, MMEJ is "error prone," and therefore, this repair mechanism can lead to indels, chromosomal translocations, and end-to-end chromosomal fusions [42, 43].

#### **3.3 CRISPR/Cas9-induced error-prone end joining DNA repair outcomes**

Then, the Cas9/gRNA complex binds to its target site, and the Cas9 HNH nuclease domain cleaves the target strand 3 bp upstream of the PAM site [50]. In contrast, the Cas9 RuvC-like nuclease domain cleaves the non-target strand 3, 4, or 5 bp upstream [50]. Therefore, Cas9-induced DSB ends are either blunt or have 1–2 bp 5′ overhangs [50]. As described above, the blunt ends can be directly ligated with the XRCC4/NHEJ1/LIG4 complex through NHEJ, without any further processing (i.e., "error-free" NHEJ) (**Figure 2**) [42]. Importantly however, even when DNA DSBs are repaired by nontemplated NHEJ, it has been established that the Cas9 cleavage cycle is repeated over and over until NHEJ mutagenic events prevent gRNA target recognition [51]. Thus, this repeated cleavage process enhances the number of non-templated indels (**Figure 2**) [51–55]. Likewise, Cas9-induced DNA DSB ends that have 1–2 bp 5′ overhangs are not ligatable and must be processed further by DCLRE1C, POLL, and POLM, which subsequently generates blunt ends followed by ligation through NHEJ [43]. Importantly, this process results in 1–2 bp indels (**Figure 2**) [55].

Similarly, Cas-induced DSBs repaired by the MMEJ pathway are also innately mutagenic due to the loss of sequence information when the extraneous heterologous 3′ ssDNA flaps are cleaved off [56, 57]. The frequency of deletions mediated by MMEJ is positively correlated with GC base content, and microhomology length, with deletions of two or more nucleotides occurring most often [50–55]. Interestingly, recent studies have established that MMEJ repair outcomes of Cas9-induced DSBs are not random and can be predicted [50–55].

Given the mutagenic nature of Cas9-induced DSBs repaired by NHEJ and MMEJ (i.e., the generation of non-templated indels), this type of end joining is leveraged frequently to silence gene expression (i.e., gene-specific knockout or loss of function)

#### **Figure 2.**

*Schematic representation of potential outcomes of error-prone NHJE/MMEJ repair of CRISPR/Cas9-mediated DSBs. The PAM site (denoted in pink) is shown relative to the DSB generated by CRISPR/Cas9 cleavage. Nontemplated DNA repair is mediated by the NHEJ/MMEJ pathway as described in the text. Three potential outcomes are shown. Nontemplated error-prone repair of CRISPR/Cas9-mediated DSBs can cause frameshift mutations leading to a stop codon and subsequent gene-specific knockout (i.e., loss of function).*

(**Figure 2**) [50–54]. Cas9-mediated error-prone NHEJ and MMEJ repair has been utilized to study the function of a wide variety of genes and noncoding elements in cellular and animal models [35, 37, 58]. Additionally, precise template-free endjoining-mediated genome editing through MMEJ has also been achieved [35, 37, 58].

### **4. Templated homology-directed repair**

HDR of endogenously generated DNA DSBs requires extensive DSB end resection and necessitates the physical base pairing interactions between the broken DNA strands and an identical sister chromatid, a homologous chromosome, or an ectopic site (i.e., a double-strand DNA [dsDNA] repair template) [4–6]. Therefore, HDR is most prominent during S and G2 cell cycle phases when an identical sister chromatid is available for recombination [59, 60]. Although HDR is typically an error-free process, indels, point mutations, genomic rearrangements, and subsequent genomic instabilities can result in a DNA donor-dependent or donor-independent manner [61].

#### **4.1 Rad51-dependent homology-directed repair**

The repair of DNA DSBs using an endogenous dsDNA repair template can occur through a RAD51 (RAD51 recombinase)-dependent mechanism [32, 34, 35, 62]. Initially, HDR, like MMEJ, begins with a short-range 3′-to-5′ resection (5–25 bps) of DNA ends mediated by the MRN/RBBP8 complex [32, 34, 35, 62]. The short-range resection is then followed by long-range 5′-to-3′ resection (>1000 bps) catalyzed by EXO1 (exonuclease 1) or DNA2 (DNA replication helicase/nuclease 2) with the assistance of BLM (BLM RecQ-like helicase) or WRN (WRN RecQ-like helicase) [32, 34, 35, 62]. The resected 3' ssDNA overhangs are subsequently stabilized by the binding of multiple RPA (heterotrimeric Replication Protein A) complexes [63]. RPA complexes are then replaced by the ATP-dependent nucleoprotein Rad51 (RAD51 recombinase) that forms long helical filaments on the resected 3′ ssDNA overhangs [64, 65]. RAD51 promotes the invasion of the overhangs (i.e., strand exchange), aligns, and pairs the ssDNA with a homologous sister chromatid sequence to form a displacement loop (D-loop) [32, 34, 35, 62]. The invading 3′ ssDNA overhang within the D-loop can then be extended by POLD1 (DNA polymerase delta 1) to synthesize sequences lost at the break site and by end resection using the homologous sister chromatid sequence as a template [66]. Finally, the resulting HDR intermediates can be resolved by multiple mechanisms, which include SDSA (synthesis-dependent strand annealing), crossover and non-crossover dHJ (double Holliday junction), and BIR (break-induced replication) [32, 34, 35, 62].

#### **4.2 Rad51-independent homology-directed repair**

Alternatively, endogenously generated DSBs cans also be repaired by a RAD51 independent HDR pathway designated single-strand annealing (SSA). Like Rad51- Dependent Homology-Directed Repair, SSA also requires long-range 5′-to-3′ resection (>1000 bps) catalyzed by EXO1 or DNA2/BLM [32, 34, 35, 62]. The resected 3' ssDNA overhangs are subsequently bound with RPA complexes; however, they are replaced by RAD52 (RAD52 homolog, DNA repair protein), which promotes the annealing of homologous sequences within the two DSB ends [67, 68]. The heterologous DNA flaps generated by SSA annealing are removed by the ERCC1 endonuclease complex, thus producing genomic deletions [48].

#### **4.3 CRISPR/Cas-induced homology-directed repair (HDR) DNA repair outcomes**

For HDR, subsequent to CRISPR/Cas-generated DSBs, an exogenous DNA template that shares homology to ends of the DSB and contains the desired genespecific nucleotide changes, mutations, or additions is required to incorporate these alterations intentionally and precisely *via* the HDR pathway (**Figure 3**) [32, 34, 35]. If a donor DNA template is not provided, then error-prone NHEJ/MMEJ will be the predominant mechanism utilized to repair the DSB and unwanted indels will occur [32, 34, 35].

If exogenous plasmids, PCR products, or chromatinized templates are utilized as dsDNA donor templates, then the Rad51-dependent HDR pathway described above is employed [32, 34, 35, 69]. In contrast, if single-strand oligodeoxynucleotides (ssODNs) are used as homologous donor templates to repair CRISPR/Cas-generated DSBs, then a RAD51-independent mechanism designated, single-stranded DNA donor-templated repair (SSTR) occurs through SSA and synthesis-dependent strand annealing (SDSA) [67, 68, 70, 71]. Like RAD51-dependent HDR, SSTR is initiated by resection of the DSB [67, 72–74] and like SSA, SSTR requires RAD52 to promote annealing of 3′ resected ssDNA tails with ssODN donor templates followed by DNAtemplated synthesis [68, 70, 74].

#### **Figure 3.**

*Schematic representation of precise gene modification mediated by HDR of CRISPR/Cas9-mediated DSBs. The PAM site (denoted in pink) is shown relative to the DSB generated by CRISPR/Cas9 cleavage. The ssODN donor template with symmetric 40-nt homology arms with the desired modifications placed in the middle of the template (denoted four red hash marks). Blocking PAM mutations (i.e., NGG → NCC) are also shown denoted with the pink and two red hash marks). After co-transfection of the Cas/RNP complex with the ssODN donor template, this template is utilized to repair the generated DSB by the HDR pathway. This allows for the precise knock-in of the sequence of interest.*

#### **5. Optimizing HDR efficiency**

#### **5.1 Allelic considerations**

Before initiating any CRISPR/Cas genome editing projects, regardless of whether knockout (i.e., NHEJ/MMEJ) or knock-in (i.e., HDR) experiments are planned, one must explore how many target gene alleles of interest are present in the cell line to be edited. This is a crucial consideration given that many cancer cell lines utilized for gene editing experiments often exhibit extensive somatic gene copy number variation (CNV) [75, 76]. Therefore, a chosen gene of interest could vary from a single copy (e.g., heterozygous deletion), two copies (e.g., normal), several copies (e.g., aneuploidy), or many copies (e.g., gene amplification) depending on which cell line is utilized for the CRISPR/Cas/NHEJ/MMEJ or CRISPR/Cas/HDR experiments.

For example, many CRISPR/Cas studies (i.e., from 2020 to 2022, greater than 100 published as per PubMed) have utilized K562 cells (an immortalized chronic myelogenous leukemia cell line) that are known to contain widespread aneuploidy and numerous obvious structural abnormalities [77–80]. Recently, Zhou et al. [81] published a comprehensive characterization of the K562 genome. This publication proved to be invaluable as our laboratory initiated CRISPR/Cas/HDR studies utilizing an anticancer drug (etoposide)-resistant K562 clonal subline, K/VP.5, previously generated by our laboratory [82, 83].

Briefly, our laboratory studies human DNA topoisomerase IIα (170 kDa, TOP2α/170), which generates transient double-strand DNA breaks to resolve nucleic acid topological entanglements [84, 85]. TOP2α/170 is an important target of anticancer drugs (such as etoposide), whose efficacy is often compromised due to decreased TOP2α/170 levels [86, 87] and resultant attenuation of cytotoxic druginduced TOP2α-DNA covalent complexes [84, 85]. Compared to parental K562 cells, etoposide-resistant K/VP.5 cells contain reduced TOP2α/170 levels and express high levels of a novel C-terminal truncated TOP2a isoform (90 kDa, TOP2α/90) [88, 89]. TOP2α/90 is the translation product of a short TOP2α mRNA that is generated from a cryptic poly(A) site harbored in intron 19 (i.e., I19 intronic polyadenylation; I19 IPA) [90, 91]. TOP2α/90 lacks the active site tyrosine 805 harbored in exon 20 of full-length TOP2α/170 necessary for TOP2α-mediated DNA strand breaks [88–91]. We hypothesized that, by utilizing CRISPR/Cas/HDR to enhance the TOP2α gene's suboptimal exon 19/intron 19 5´ SS (E19/I19 5´ SS), removal of intron 19 would be enhanced, which in turn would result in decreased TOP2α/90 mRNA/protein, increased TOP2α/170 mRNA/protein, and circumvention of etoposide resistance [92].

Since the human TOP2α gene is harbored on chromosome 17 (i.e., mapped to chromosome 17q21–22) [93], Zhou et al.'s [81] study was utilized to determine the number of TOP2α alleles (i.e., copy number) present in the K562/K/VP.5 cells before initiation of CRISPR/Cas/HDR experiments [92]. It was found that K562 and the isogenic-acquired resistant cell line, K/VP.5, contained three TOP2α alleles [81]. Therefore, our CRISPR/Cas9/HDR strategy was focused on editing all three TOP2α alleles in K/VP.5 cells at the E19/I19 5´ SS to maximize the desired phenotypic change (i.e., decreased TOP2α/90 mRNA/protein, and increased TOP2α/170 mRNA/protein levels) and to circumvent etoposide resistance [92]. qPCR and Sanger sequencing demonstrated that the ratio of wild-type to edited genomic sequence decreased by 1/3 with each allele edited [92]. TOP2α/90 progressively decreased and TOP2α/170 increased with each allele edited by CRISPR/Cas9/HDR. Etoposide resistance was completely reversed when all three TOP2α alleles were edited to enhance the E19/I19

5´ SS [92]. RNA seq confirmed that intron 19 was effectively spliced out in the three allele-edited clone [92].

#### **5.2 PAM site considerations**

Multiple studies have demonstrated that CRISPR/Cas-generated DSBs should be in close proximity to the edit site to achieve high HDR efficiencies [93–100]. These investigators established that if a Cas9 PAM site (i.e., NGG; N denotes any nt) was located more than 14 bp (on either DNA strand) from the desired gene-specific nt changes, mutations, or additions, then the efficiency of CRISPR/Cas9/HDR was dramatically reduced. However, Renaud et al. [96] observed that the 14-bp limitation may be pushed to 20 bp utilizing chemically modified ssODN donor templates (see "HDR Considerations" below). Paquet et al. [94] also demonstrated that it was easier to create homozygous gene edits when the PAM site was closer to the intended nucleotide changes and heterozygous gene editing by distance-dependent suboptimal mutation incorporation.

Importantly, Schubert et al. [100] indicated that although guide selection in close proximity with the required HDR changes is important, it was more significant that the gRNA utilized not only targeted Cas9 to the appropriate sequence but also activated Cas9 endonuclease activity. Therefore, since all gRNAs are not equally efficient in activating Cas9, it is essential that the cleavage efficiency for each gRNA utilized is calculated using a T7 endonuclease mismatch cleavage assay (i.e., measuring the extent of indel formation) before initiating HDR experiments [97, 100]. If several Cas9 PAM sites are identified within the 15 base HDR parameter, then the gRNA eliciting the highest cleavage efficiency should be utilized for CRISPR/Cas9/HDR experiments (see "gRNA Considerations" below).

Since the lack of gRNAs with appropriate cleavage efficiency and proximity to the desired HDR-mediated changes is a significant limitation for many CRISPR/Cas9/ HDR studies, Schubert et al. [100] also demonstrated that Cas9 D10A nickases (i.e., induce single DNA nicks) can be utilized for HDR mutation experiments if gRNAs target PAM sites on opposite strands of the genomic DNA to generate a staggered DSB provided that the desired mutation is placed between the two nick sites. Alternatively, the number of possible CRISPR/Cas/HDR editing sites can be expanded with the utilization of Cas12a (also known as Cpf1), which recognizes a unique PAM site (TTTV; V denotes an A, C, or G nt) [101].

Since Cas9/Cas12a PAM site recognition restricts targeting and affects CRISPR/Cas/ HDR editing efficiency and flexibility, there are efforts to genetically re-engineer CRISPR enzymes to target heretofore inaccessible PAMs [102–105]. For example, Kleinstiver et al. [106] have successfully altered *S. pyogenes* Cas9 (SpCas9) PAM specificity by utilizing bacterial selection-based directed evolution. Walton et al. [104] utilized structure-guided engineering to develop several "near-PAMless" SpCas9 variants capable of targeting NGN and NRN (R denotes an A or G), respectively. Finally, Kleinstiver et al. [105] have also utilized structure-guided protein engineering to improve the targeting range of *Acidaminococcus sp.* Cas12a. Together these studies suggest that the PAM site constraints that currently limit CRISPR/Cas/HDR editing will be circumvented in the future.

#### **5.3 gRNA considerations**

Most CRISPR/Cas genome editing experiments are now performed by delivering purified Cas9/Cas12 proteins and chemically synthesized gRNAs as a ribonucleoprotein (RNP) (i.e., Cas/RNP) complex to restrict their temporal activity, improve

*Maximizing the Efficacy of CRISPR/Cas Homology-Directed Repair Gene Targeting DOI: http://dx.doi.org/10.5772/intechopen.109051*

precision, decrease the immune response, and reduce off-target effects [106, 107]. Specifically, engineered gRNAs have been chemically modified to increase their stability and decrease off-target editing resulting in enhanced cleavage efficiency and improved HDR efficacy [108, 109].

gRNAs can be synthesized in two formats. First, like the endogenous Cas9 gRNA, the crRNA/tracrRNA is a two-piece gRNA where the crRNA (~36–42 nt) and tracrRNA (~67–89 nt) are synthesized as two independent oligonucleotides and are subsequently annealed together through a complimentary linker region to form a functional gRNA [28]. Second, a single guide (sgRNA, 100 nt) can be synthesized, which comprises both the crRNA and tracrRNA in a single oligonucleotide (no annealing is required) (**Figure 4**). It is important to note that the PAM sequence is not included in either gRNA format [28]. One advantage of the two-piece gRNAs is that the tracrRNA sequence is the same for all CRISPR/Cas9 experiments and only the crRNA sequence varies, based on the DNA site to be targeted [28]. Therefore, one chemically synthesized tracrRNA can be annealed to any chemically synthesized crRNA. Chemically synthesized chimeric sgRNAs have the advantage that they exhibit equivalent or greater efficiency compared to the native dual RNA system [109, 110]. We advocate for the use of sgRNAs since our laboratory tested several two-piece gRNAs that exhibited no activity [92] that when resynthesized as sgRNAs displayed high cleavage efficiency [111].

Since RNAs are inherently unstable and susceptible to endo- and exonucleases, considerable effort has been devoted to chemically modifying RNAs to improve their stability. Importantly, Hendel et al. [112] established that chemical modification of

#### **Figure 4.**

*Schematic representation of the CRISPR/Cas9-mediated DSB with a one-piece sgRNA. The CRISPR/Cas9 schematic is denoted as described in Figure 1, except Cas9 sgRNA is synthesized as a single molecule, which harbors both the crRNA and the tracrRNA (denoted in light blue). The DSB is created as described in Figure 1. sgRNAs have the advantage that they can exhibit greater efficiency compared to the native dual RNA system with no crRNA/tracrRNA annealing step required [109, 110].*

gRNAs protected them from degradation and enhanced genome editing efficiency. Specifically, these investigators demonstrated that when 2′-O-methyl 3′phosphorothioate (MS), or 2′-O-methyl 3′thioPACE (MSP) chemical modifications were incorporated at both the 5′ and 3′ three terminal nucleotides, and indel formation and HDR were significantly increased [112]. They concluded that chemically synthesized/ modified sgRNAs offer significant advantages over sgRNAs expressed by plasmids or by *in vitro* transcription, including 1) scalable and robust production for many applications; 2) greater sgRNA design flexibility; 3) lower toxicity; and 4) increased efficacy [112]. In conclusion, the studies reviewed in this section clearly suggest that the continued optimization of synthetic gRNAs will increase cleavage and on-target efficiency, which will help leading to future efficacious CRISPR-based therapies.

#### **5.4 HDR template considerations**

Regardless of whether dsDNA or ssODN donor templates are utilized by distinct HDR pathways to mend Cas RNP complex generated DSBs, the same precise, intentional repair outcomes can be achieved. However, ssODNs donor templates are most frequently used to introduce specific changes (e.g., introduce or correct mutations, and to create short insertions) into specific DNA sequences through HDR due to their superior efficiency, fidelity, and ease of synthesis (**Figure 3**) [71, 74, 95–97, 113]. Importantly, recent studies investigating 1) chemical modifications at the 5′ and 3′ ends of ssODNs donor templates; 2) optimal complementary length; 3) homology arm polarity and asymmetry; and 4) donor template design to prevent the re-cleavage of edited alleles have resulted in empirical rules to rationally design ssODN donor templates to maximize HDR efficiency and flexibility [94–100].

Renaud et al. [96] established that phosphorothioate (PS) chemical modifications at the two terminal nucleotides at both the 5′ and 3′ ends of ssODNs repair templates strongly enhanced genome editing efficiency in cultured cells. These investigators also demonstrated that PS-modified ssODN donor templates also permitted efficient insertion of over 100 nucleotides, while only limited integration was observed with nonchemically ssODNs [96]. Likewise, a higher frequency of insertions was attained in mice and rats using modified ssODNs [96]. The importance of utilizing PS-modified ssODN repair templates to enhance HDR editing efficiency was validated by Liang et al. [97].

Richardson et al. [95] demonstrated that although Cas9 dissociates slowly from dsDNA substrates, Cas9 releases the 3′ end of the cleaved nontarget strand (NT strand, the DNA strand that is not complementary to the gRNA and harbors the "NGG" PAM sequence) before complete Cas9 dissociation. They subsequently showed that ssODNs donor templates complementary to the NT strand increased HDR frequencies compared to donor templates complementary to the target strand (T strand; the DNA strand that is complementary to the gRNA and does not contain the "NGG" PAM sequence) [95]. Finally, these investigators established that ssODN donor templates asymmetrically oriented relative to the 5′- and 3′-side of the generated DSB and complementary to the NT strand also increased HDR rates [95]. In support of these results, Liang et al. [97] also showed that asymmetric ssODNs with 30-bp homology arms 3′ to the insertion and greater than 40 bp of homology at the 5′ end were preferred. This report indicated that the optimal amount of asymmetric ssODN was 10 pmol. However, in contrast to Richardson et al. [95], these investigators only observed a slight increase in HDR efficiency with NT strand compared with T strand ssODN repair templates [97]. Okamoto et al. [99] demonstrated that the optimal ssODN donor template should have a total length of ~75–85 nt with 30 to

#### *Maximizing the Efficacy of CRISPR/Cas Homology-Directed Repair Gene Targeting DOI: http://dx.doi.org/10.5772/intechopen.109051*

35 nt perfectly matched homology arms on the 5′ and 3′ ends and complementary to the gRNA strand (i.e., T strand). Recently, Schubert et al. [100] established further design parameters to improve HDR efficiencies by testing hundreds of genomic loci and multiple cell lines. First, they demonstrated that the ssODN donor template (i.e., NT or T strand) that leads to the highest HDR efficiencies varies greatly depending on the genomic locus and cell type utilized [100]. Second, they observed that the preferred strand (NT or T), relative to the gRNA, is dependent on where the desired HDR modification is located. For example, there is no repair strand preference when the HDR modification is placed precisely at the Cas9 cleavage site [100]. However, if the HDR modifications occur further from the Cas9 cleavage site, a NT strand ssODN donor template is preferred for PAM-distal mutations and a T strand ssODN donor template is ideal for PAM-proximal mutations [100]. Additionally, they showed that asymmetric homology arms did not improve HDR beyond symmetrical homology arms when arm length was ≥30-nt from both the mutation location and the Cas9 cleavage site [100]. These investigators advocated for ssODN donor templates with 40-nt homology arms with modifications placed in the middle (**Figure 3**) [100].

The CAS RNP complex can regenerate DSBs in alleles already appropriately edited, thereby lowering HDR efficiency [94, 99]. In another design innovation using ssODN donor templates, Paquet et al. [94] strategically prevented re-cutting of HDR-edited sites by introducing CRISPR/Cas-blocking PAM site mutations in their repair templates and observed increased HDR accuracy and effectiveness. Okamoto et al. [99] subsequently established that ssODN repair templates with a single mutation in the PAM site (i.e., NGG → NGC) showed the highest HDR efficiency. Their results clearly indicated that the re-cutting of edited alleles resulted in very low HDR efficiencies, and that introducing PAM site mutations within ssODN repair templates to prevent re-cutting is essential for efficient HDR knock-in [99]. Schubert et al. [100] also demonstrated that adding a blocking PAM mutation to the second or third base of the PAM (i.e., NGG → NCG or NGC) in ssODN repair templates resulted in greater HDR efficiency. Donor templates containing two blocking PAM mutations (i.e., NGG → NCC) resulted in the highest HDR efficiency (**Figure 3**). Finally, another important indication for blocking PAM mutations in HDR repair templates is to ensure that when multiple rounds CRISPR/Cas/HDR transfections are required to edit all gene-specific alleles in cell lines that exhibit aneuploidy; the previously edited alleles will not be re-cut in the subsequent rounds of transfection [91, 92].

#### **5.5 Pharmacological strategies to enhance HDR efficiency**

Most CRISPR/Cas/HDR genome editing experiments are now performed by transfecting Cas/RNP complexes and ssODN repair templates to restrict temporal activity, thereby reducing off target effects, decreasing immune responses, and increasing HDR efficiency [94–100, 106–110, 112, 113]. Discussion below of pharmacological and genetic strategies for gene editing by HDR will be limited to this experimental paradigm.

Since NHEJ/MMEJ are rapid high-capacity pathways which are active throughout the cell cycle (i.e., G1, S and G2 phases), while HDR is active only after DNA replication is completed and sister chromatids are available to serve as repair templates (i.e., late S and G2 phases) [60], one of the first attempts to pharmacologically enhance CRISPR/Cas/HDR efficiency was to time the delivery of Cas RNA complexes after synchronization of cells using aphidicolin or nocodazole [114]. Lin et al. [114] demonstrated that synchronization, with either aphidicolin or nocodzole, resulted in increased HDR rates (up to 38%) compared with unsynchronized cells.

Since DNA repair is also influenced by the accessibility of DNA binding factors, like Cas RNP complexes [115, 116]. Li et al. [117] hypothesized that CRISPR/Cas/ HDR efficiency would be enhanced with histone deacetylase (HDAC) inhibitors, trichostatin A (TSA) and PCI-24781, by promoting a more open chromatin structure [118]. These investigators established that HDR, single strand annealing and ssODN mediated HDR were all increased with HDAC inhibitor treatment [118]. Moreover, this study demonstrated that TSA and PCI-24781 usage also favored HDR by arresting the cell cycle in the G2/M phase [118].

Another pharmacological strategy to improve HDR efficiency is to target the competing NHEJ/MMEJ pathways. Riesenberg et al. [119], explored the efficacy of a wide range of small molecules reported to inhibit the NHEJ/MMEJ pathways or to activate/ increase the HDR protein components. These investigators determined that NU7026 (DNA-dependent protein kinase, DNA-PK inhibitor), TSA, MLN4924 (NEDD8 E1 Activating Enzyme Inhibitor), and NSC 15520 (replication protein A1, RPA1 inhibitor) increased HDR efficiencies in various genes and in specified cell lines when DSBs were generated by nickase Cas9n and Cas12/RNP complexes [119]. When Cas9/ RNP complexes were utilized to generate DSBs, only NU7026 significantly increased HDR efficacy [119]. NSC 19630 (WRN RecQ like helicase, WRN inhibitor), AICAR (protein kinase AMP-activated catalytic subunit alpha 1, PRKAA1 activator), RS-1 (RAD51 recombinase, RAD51 stimulator), Resveratrol (selective inhibitor of COX−1), SCR7 (DNA ligase IV inhibitor), and L755507 (potent β3-adrenergic receptor partial agonist), showed no clear effect on any Cas/HDR efficiency [119]. Finally, it was demonstrated that the combination of NU7026, TSA, MLN4924, and NSC 15520 resulted in the highest HDR levels observed with Cas9n and Cas12/RNP complexes [119].

In contrast to the results presented above, other investigators have successfully utilized SCR7 (DNA ligase IV inhibitor) to specifically impede the NHEJ pathway to increase HDR activity [120–124]. However, there are also conflicting reports on the ability of this compound to increase HDR [119, 125, 126]. The lack of consistency with this compound regarding HDR efficacy may have resulted from the use of different chemical derivatives of SCR7 [127]. Additionally, Greco et al. [127] demonstrated that SCR7 exhibited greater inhibitory activity against DNA ligases I and III than DNA ligase IV and therefore should target the MMEJ pathway (i.e., also involved in errorprone repair).

#### **6. Conclusion**

CRISPR/Cas/HDR is a robust gene editing methodology to purposefully edit out or introduce mutations or insertions at specific genomic sites (i.e., targeted gene knock-in) by creating a DSB along with the introduction of an exogenous template harboring the desired nt changes for DSB repair *via* the HDR pathway [32, 34, 35]. Since DSBs generated in mammalian cells are predominantly repaired by NHEJ/ MMEJ pathway [4–6], success of CRISPR/Cas/HDR gene editing will depend on maximizing overall HDR efficacy. We propose the following "workflow" strategies to facilitate the knock-in process (**Figure 5**). First, after developing an experimental CRISPR/Cas/HDR hypothesis, serious consideration must be given to the appropriate cellular system to utilize and the number of gene alleles of interest that may need be edited to obtain a resulting altered phenotype. If the cell line of choice is not well characterized, sequence analysis of CRISPR-edited cells may help determine the number of edited and non-edited alleles [91, 92, 111].

*Maximizing the Efficacy of CRISPR/Cas Homology-Directed Repair Gene Targeting DOI: http://dx.doi.org/10.5772/intechopen.109051*

#### **Figure 5.**

*Proposed experimental flow diagram to maximize CRISPR/Cas/HDR gene editing efficiency. An explanation of the flowchart is discussed in the conclusion section.*

Second, the location of the targeted gene knock-in must be analyzed to determine if a Cas9/12 PAM site(s) is/are harbored within 20 nt of the DNA sequence to be edited (both DNA strands should be analyzed) [93–100]. If more than one PAM site is identified that is in close proximity to sequence to be edited, we advocate that multiple chemically modified 5′ and 3′ terminal nucleotide sgRNAs be synthesized. Although chemically modified sgRNAs exhibit increased stability and decreased offtarget editing [108, 109], not all gRNAs are equally efficient in activating Cas enzymes [100]. Therefore, the cleavage efficiency for each sgRNA should be determined by employing a T7 endonuclease mismatch cleavage assay [97, 100] before proceeding with CRISPR/Cas/HDR editing experiments. The sgRNA with the highest cleavage efficiency should be utilized [100].

Third, for CRISPR/Cas/HDR, ssODN repair templates chemically modified at their 5′ and 3′ ends should be used to introduce specific changes (e.g., introduce or correct mutations, and to create short insertions) due to their superior efficiency, fidelity, and ease of synthesis [71, 74, 95–97, 113]. Although there are conflicting opinions regarding the optimal homology arm length, homology arm polarity, and asymmetry [94–100], it is now well established that HDR efficiencies varies greatly depending on the genomic locus and cell type utilized [94–100]. The most recent CRISPR/Cas/HDR editing data suggest that ssODN donor templates with symmetric 40-nt homology arms with the desired modifications placed in the middle of the template should be an appropriate standard approach (**Figure 3**) [100]. Regarding

the polarity of the ssODN donor templates with respect to the gRNA (see **Figure 1**), an NT strand ssODN donor template is preferred for PAM-distal mutations and a T strand ssODN donor template is ideal for PAM-proximal mutations [100]. Finally, blocking PAM mutations (i.e., NGG → NCC) should always be introduced in ssODN repair templates to further increase HDR efficiency [94, 99, 100] and to allow for potential repeated rounds of CRISPR/Cas/HDR transfections when editing multiple alleles in cell lines, which exhibits aneuploidy (**Figure 3**) [91, 92, 111].

Fourth, the chemically modified sgRNA with the highest cleavage efficiency should be incubated with high-quality, purified Cas9 or Cas12 enzymes that harbor a nuclear localization sequence (NLS) to form a Cas/RNP complex followed by co-transfection with an optimized ssODN donor template as above. Importantly, this procedure improves precision, restricts the temporal activity, decreases the immune response, and reduces the off-target effects of Cas proteins (**Figure 3**) [94–100, 106, 107].

Fifth, after transfection, small aliquots of cell suspension should be sampled to determine genomic cleavage efficiency (i.e., T7 endonuclease mismatch cleavage assay) [97, 100] to validate success in transfection, targeting, and DSB formation by the Cas/RNP complex at the appropriate genome location [91, 111]. Remaining cell suspensions should then be diluted (i.e., limiting dilution) and transferred to 96-well plates at a concentration less than one cell per well and allowed to grow until individual colonies are identifiable in some wells [128]. Individual clonal populations can be split into larger wells and then qPCR and/or Sanger sequencing [91, 92, 111] utilized to determine which clones contain the desired CRISPR/Cas/HDR editing. If electropherogram visualization of genomic sequence reveals only edited sequence, then this clone can be characterized for the hypothesized phenotypic changes. In contrast, if wild-type and edited genomic sequences are identified in the electropherogram, then the ratio of edited to wild-type to edited alleles can be determined. Editing all gene alleles of interest may be required to detect the hypothesized phenotypic change(s) [91, 92, 111]. If sequencing results reveal that a significant number of clonal cells have undergone NHEJ/MMEJ with no HDR editing, then pharmacological inhibitors (as described above) can be considered in an attempt to increase HDR efficiency.

The unique CRISPR/Cas/HDR gene editing experimental outline described in **Figure 5** incorporates the most comprehensive sgRNA and ssODN design considerations along with several important practical details that will help maximize the frequency of precise HDR. It is anticipated that, as strategies to enhance CRISPR/Cas/ HDR efficacy continue to advance, that this tractable experimental workflow will accelerate the development of therapeutic gene editing.

#### **Acknowledgements**

The work described was supported by grant CA226906-01A1 from the National Institutes of Health (J.C.Y. & T.S.E.).

*Maximizing the Efficacy of CRISPR/Cas Homology-Directed Repair Gene Targeting DOI: http://dx.doi.org/10.5772/intechopen.109051*

### **Author details**

Terry S. Elton\*, Md. Ismail Hossain, Jessika Carvajal-Moreno, Xinyi Wang, Dalton J. Skaggs and Jack C. Yalowich\* Division of Pharmaceutics and Pharmacology, College of Pharmacy, The Ohio State University, Columbus, Ohio, United States of America

\*Address all correspondence to: elton.8@osu.edu and yalowich.1@osu.edu

© 2022 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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#### **Chapter 4**

## The Prominent Characteristics of the Effective sgRNA for a Precise CRISPR Genome Editing

*Reza Mohammadhassan, Sara Tutunchi, Negar Nasehi, Fatemeh Goudarziasl and Lena Mahya*

#### **Abstract**

Clustered regularly interspaced short palindromic repeats (CRISPRs) technique is the most effective and novelist technique for genome editing. CRISPR mechanism has been widely developed for gene editing, gene silencing, high-specific regulation of the transcription, and reducing off-target effects through double-strand breaks (DSBs) in the genomic DNA and then modifying nucleotide sequences of the target gene in diverse plant and animal species. However, the application may be restricted by a high rate of off-target effects. So, there are many studies on designing precise single-guide RNAs (sgRNAs) to minimize off-target effects. Thus, the high-efficiency design of a specific sgRNA is critical. First, in the chapter, the sgRNA origin and different types of gRNA will be outlined. Then, the off-target effect will be described. Next, the remarkable characteristics of the sgRNA will be highlighted to improve precise gene editing. Finally, some popular *in silico* tools will be introduced for designing sgRNA.

**Keywords:** sgRNA, guide RNA, crRNA, tracrRNA, off-target effect, designing tools, CRISPR/Cas

#### **1. Introduction**

Clustered regularly interspaced short palindromic repeats (CRISPRs) and their CRISPR-associated (Cas) proteins system is an effective immune system among bacteria and archaea. This system was first discovered in the *E.coli* genome. CRISPR/ Cas is an acquired immunity mechanism in many bacteria and archaea against the genome of the infection factors such as viruses and plasmids [1]. CRISPR/Cas system is classified into three major groups (I, II, and III) with a specific functional mechanism and gene family encoding the specific Cas proteins. Types I and III apply several Cas proteins for endonuclease activity, while type II uses only one protein (Cas9) [2]. Evolutionary competition between the pathogens and host in the CRISPR/Cas system shows a very high variable rate in structures and functions. So, recent classification has stated that the CRISPR system has been categorized into two classes (I and II) and six types (I–VI) [3].

Most studies on genome engineering have been performed in system type II, derived from Streptococcus pyogenes (SpCas9). The advantage of system type II is that it needs only one protein (Cas9) for endonuclease activity. However, this system also needs the types of RNA, including CRISPR RNA (crRNA), which functions as a Cas protein guide for pairing with the target genome sequences, and trans-activating CRISPR RNA (tracrRNA), which play critical roles in crRNA maturation and directing Cas9 to the desired site [4, 5]. Merging of tracrRNA:crRNA sequences as a chimeric sequence known as single-guide RNA (sgRNA), which covers the features of both RNA types, makes it suitable and applicable for genome editing [6].

All CRISPR sites cover consecutive and spacer repetitions. These consecutive repetitions include identical sequences, while the spacer sequences originate from the genome of foreign factors [7, 8]. CRISPR sites and Cas proteins develop acquired immunity against the invading DNA. Suppose a microorganism survives the invasion of a pathogen. In that case, the integrated CRISPR system will be able to incorporate a piece of the pathogen DNA into its genome and then use it to fight against subsequent infections. This bacterial immune system degenerates the phage genome by integrating short fragments of the phage DNA in the spacer region of the CRISPR sites and transcribing the spacers (known as crRNAs) with associated Cas endonuclease in subsequent infections [9–11].

Briefly, the CRISPR system, as the RNA-mediated immune system in prokaryotes (bacteria and archaea), functions in four stages (**Figure 1**):


The classification represents the evolution of subtype-specific molecular defensive mechanisms for crRNA expression and maturation, as well as the inhibition of infectious factors [18]. The main known vital component of the CRISPR/Cas system is crRNA, which is common in types I and III. Pre-crRNAs are initially cleaved within the repeats by a Cas6 endoribonucleases family, and then intermediate crRNAs are further matured to generate shorter repeat-spacer crRNAs in type III. In both type I and type III, mature crRNAs direct a complex of multiple Cas proteins to the cognateinvading nucleic acids. Then, the target nucleic acids are cleaved by a Cas endonuclease of the ribonucleoprotein complex [19–21].

Pre-crRNA processing necessitates base-pairing of each pre-crRNA repeat with tracrRNA, a small noncoding RNA encoded near the *Cas* genes and spacer array [7]. *The Prominent Characteristics of the Effective sgRNA for a Precise CRISPR Genome Editing DOI: http://dx.doi.org/10.5772/intechopen.106711*

#### **Figure 1.**

*RNA-mediated CRISPR immune system in.*

The base-pairing drives cleavage and binding by RNase III and Cas9, respectively. Then, the crRNA:tracrRNA complex can direct Cas9 to bind target DNA sequences by matching PAM [16, 17]. In addition, type II CRISPR-Cas systems continuously utilize the crRNA:tracrRNA complex to identify and cleave double-stranded DNA (dsDNA). Recognition is driven by base-pairing between the guide sequence and the RNA in the crRNA:tracrRNA complex (**Figure 2**) [22, 23]. The tracrRNA is a common component of CRISPR-Cas systems among all three subtypes of type II systems and is required for crRNA biosynthesis [24]. These basic principles were discovered by following efforts to characterize tracrRNAs and crRNA biogenesis [25]. The biogenesis of crRNA, the structures of the crRNA:tracrRNA complex, and tracrRNA genomic location are variable among these subtypes [26]. The tracrRNA discovery as a key factor of crRNA biosynthesis allowed the sgRNA to be invented and Cas9 to be adopted as the core component of CRISPR technology [27].

**Figure 2.** *Schematic structure of the crRNA:tracrRNA complex.*

The major function of the sgRNA is efficiently the target region detection through the PAM sequence to edit a gene precisely. However, two vital challenges include efficacy and specificity for designing an effective sgRNA [28]. According to the significance of the sgRNA function, the role and designing tools of sgRNA will be outlined in the following.

#### **2. The role of sgRNA in CRISPR technology**

CRISPR/Cas genome editing technology can cause double-stranded DNA breaks (DSBs) in predefined genomic loci [29]. These DSBs are then repaired by the DNA repair systems of the target organism, which inherently can cause a mutation in the target gene. Despite the general processes driving all these genome editing systems being similar, CRISPR/Cas technology has emerged as the preferred technique due to its easy usage, low cost, outstanding adaptability, and ability to target several genes at once [30–32].

This technology consists of a Cas endonuclease, responsible for eliciting the DSB, and a short noncoding about sgRNA (20-nt), directing Cas to the correct genomic region for targeted genome editing. A chimeric gRNA (complementary to the target area) and trans-activating CRISPR-RNA are usually included in this sgRNA. Most Cas systems require the predesigned sgRNA to anneal immediately upstream of a PAM, which in the case of SpCas9 (the most extensively employed Cas protein for genome editing) is 5` NGG3` [4, 5, 33]. In these cases, the PAM is required to cleave target DNA around 3-nt upstream of this region. DSBs are repaired by either nonhomologous end joining (NHEJ) or homology-directed repair (HDR) as two central intrinsic DNA repair systems [34]. The error-prone nature of NHEJ is the main DNA repair route in species and the most common and straightforward pathway in genome

#### **Figure 3.**

*CRISPR/Cas9 technology for gene editing. The Cas9 DNA endonuclease is recruited by a single-guide RNA (sgRNA) that detects a genomic sequence followed by a 5*′*-NGG-3*′ *PAM motif.*

*The Prominent Characteristics of the Effective sgRNA for a Precise CRISPR Genome Editing DOI: http://dx.doi.org/10.5772/intechopen.106711*


#### **Table 1.**

*Comparison between genome editing systems.*

editing, causing small insertions or deletions (indels) to interrupt the target sequence (**Figure 3**) [35].

CRISPR/Cas technology relies on DNA-RNA interaction as well as simple design of RNA molecule for each specific sequence. However, protein-DNA interaction also depends on zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs). So there is a required reconstruction for each target DNA sequence. This is a significant benefit of CRISPR/Cas technology [36]. In addition, the technology has three other advantages over TALENs and ZFNs, including the following (**Table 1**):


#### **2.1 The role of PAM**

Although specific targeting significantly depends on the sgRNA sequence, PAM-specific sequence plays a crucial role in the effective enzymatic activity


#### **Table 2.**

*Different PAM sequences of some Cas endonucleases and their origins.*

of Cas endonuclease. In this system, Cas9 endonuclease can cleave any genome sequence immediately located on five nucleotides (nt) downstream of PAM sequence [25]. PAM sequences recognized by SpCas9 and StCas9 (from *Streptococcus thermophilus*) is, respectively, 3`-NGG-5` and 5`-NNAGAAW-3` (**Table 2**) [40, 41]. The SpCas9-sgRNA complex first seeks the complement sequence of PAM in the target genome and then the sgRNA base pairs to the target DNA, then the DNA is cleaved by SpCas9 to create DSB. Generally, the length of the DNA detection sequence in the crRNA region is 20 nt, though the more base pairs bind between RNA and DNA, the more the specificity of the sgRNA function can enhance. Hence, the 20 nt sequence of sgRNA and 3 nt in PAM play key roles in the specific targeting of CRISPR/Cas technology. However, there are some limitations in using the 3`-NGG-5` motif, particularly at high AT sequences of the target genome [42, 43].

#### **2.2 Variety of sgRNA types**

According to the development of the CRISPR system as technology and sgRNA invention, there are some improvements of sgRNA to enhance the efficiency, precision, and specificity of the genome editing technology [44]. The improvements include as follows:


*The Prominent Characteristics of the Effective sgRNA for a Precise CRISPR Genome Editing DOI: http://dx.doi.org/10.5772/intechopen.106711*

#### **2.3 Off-target effect**

Off-target mutation is a major challenge in CRISPR/Cas technology [48]. If the gRNA sequence contains less than three heterologous nucleotides to an off-target region, off-target effects will be observed [49]. Studies have indicated that mismatched pairs at the end of the 3` terminal of the target sequence are not tolerated (typically 8–14 nucleotides upstream of PAM sequence). In contrast, the mismatched pairs at the 5` terminal of the target sequence are more tolerable [50]. The sgRNA/ Cas9 also can influence the off-target effects [49].

Generally, although Cas9 protein can be differently used according to high endonuclease activity and the wide targeting range of the enzyme, the high molecular weight of the Cas9 endonuclease and off-target effects can restrict the popularity of the enzyme. Nevertheless, some variants of Cas9, such as SpCas9-HF and eSpCas9 [51, 52], can be mutated. So, the mutation can reduce the nonspecific interaction between the Cas9 protein and the target sequence. Digenome-seq, GUIDE-Seq (Genome-wide, Unbiased Identification of DSBs Enabled by Sequencing), and HTGTS (High-Throughput Genome-Wide Translocation Sequencing) can be employed for detecting off-target regions [53]. However, precisely designing sgRNA can significantly decrease the rate of the off-target effects [44].

#### **3. Characteristics of an effective sgRNA**

In addition to directing Cas endonuclease, sgRNA can stimulate Cas endonuclease activity. These functions of sgRNA can clarify how to tackle on-target effects [54]. It has been demonstrated that the proximal and distal ends of the PAM sequence are highly responsible for improving on-target effects. Besides, genomic frameworks of the target sequence, GC content, sgRNA length, and secondary structure play significant roles in enhancing the on-target rate [44]. Also, 5` terminal 20 nt of the sgRNA is highly efficient for on-target efficiency. However, there is insufficient information on the correlation between structure and sequence properties of sgRNA influencing the on-target effect [55]. At least, enhancement of the on-target rate can improve the efficiency of the CRISPR technology and facilitate the statistical interpretation of the edited genes rate [56].

#### **3.1 GC content of the sgRNA**

As mentioned above, the GC content of the sgRNA is closely related to the ontarget rate and the efficiency of the CRISPR gene editing. It has been indicated that too high or low GC content is unsuitable for achieving high rate of the on-target effects [57]. The knockout effectiveness of the CRISPR/Cas9 system was significantly improved by changing the sgRNA structure by expanding the duplex length (about 5 nt) and replacing the fourth T by C or G [58]. Many studies have reported that GC content plays a key role in improving the knockout efficiency of the CRISPR/Cas9 system. The effective rate of GC content is 40–60 percent. It is also recommended that sgRNA containing 50 percent GC content is efficient for CRISPR gene editing [59–61]. However, some studies suggest higher than 60 percent GC content for each organism, such as *Escherichia coli* (62.5%) and *Vitis vinifera* (65%) [62, 63]. In addition to GC content, purine residues and curvature in positions C3 and C16 of sgRNA could be effective in improving on-target activity [64]. It has also been reported that

sgRNAs containing 4 GC in the 6 nt close to the PAM sequence can effectively reduce off-target effects [65].

#### **3.2 The sgRNA length**

The most common sgRNA length is about 100 nt. Therefore, 5` terminal 20 nt of the sgRNA can be designed as the complement sequence of the target gene to direct precisely Cas endonuclease for achieving effective gene editing [66]. According to several studies, the less the sgRNA length is, the higher the rate of off-target effects increase [67–69]. However, as mentioned above, 17 nt length for tuRNA can be highly effective in reducing off-target effects. However, the length of the sgRNA recognition site is less than 15 nt, and Cas endonuclease will not show any activity [70, 71]. In addition to the sgRNA length, the efficiency and specificity of Cas cleavage activity in the targeting sequence are significantly influenced by the distance between the PAM site and the start codon [72].

#### **3.3 The sgRNA secondary structure**

The sgRNA secondary structure is highly responsible for effective Cas-target sequence binding [73]. There are also many reports to indicate that the presence of the quad stem-loop structure of sgRNA is a key factor in improving the efficiency of the riboprotein function. The repeat and anti-repeat region (stem-loop RAR, GAAA) can activate sgRNA processing before Cas-sgRNA binding. Besides, loops 2 (GAAA) and 3 (AGU) are demanded to create a stable riboprotein complex, but there is no report on the possible loop 1 role in sgRNA efficiency [74–77]. Besides, the hairpin structure of the sgRNA, particularly the inner side of the hairpin, plays a key role in cleaving target DNA by Cas9/sgRNA. In fact, the hairpin structure can provide a suitable conformation to bind Cas9 enzyme. If the loop-stem structure is elongated, the gene editing efficiency will be enhanced [78, 79]. It has been indicated that CRISPR efficiency can be improved when an engineered hairpin structure is inserted into the spacer region of sgRNA. The modified sgRNA can positively influence Cas-mediated transactivation and improve the function of the five different Cas9 and Cas12a variants. The evidence can demonstrate the effect of sgRNA secondary structure on the success rate of gene editing [80, 81]. There is also a correlation between the sgRNA secondary structure and the efficiency of the Cas9-mediated CRISPR [55]. It has been reported that sgRNA refolding can refine the destructive bonds of the deactivated sgRNAs. Also, heating or slowly chilling can thermodynamically activate these sgRNAs to improve Cas9 cleavage activity. At least, the sgRNA secondary structure can change the guide sequence activity and deactivated sgRNA can be recovered by refolding [82, 83].

#### **3.4 The sgRNA sequence**

In addition to these criteria, the sgRNA sequence features can show different efficiency levels. It has been reported that the functional sgRNA can be significantly accessed at certain nucleotide positions more than nonfunctional sgRNA. Particularly, 3` terminal nucleotides (positions 18–20) of the sgRNA can highly make a prominent difference in accessibility [84, 85]. The sgRNA 3` terminus, known as the seed region, is a key player in recognizing the target sequence. Therefore, accessibility of the final three bases of the seed region is a remarkable characteristic in

#### *The Prominent Characteristics of the Effective sgRNA for a Precise CRISPR Genome Editing DOI: http://dx.doi.org/10.5772/intechopen.106711*

distinguishing functional sgRNAs from nonfunctional ones based on structural analysis [86, 87]. Also, G at the 5` terminus of the spacer is demanded in the nonribosomal and ribosomal complexes. Besides, G is extremely proper at positions −1 and − 2, close to PAM associated with the sequence preference for loading Cas9 endonuclease [88]. According to the finding that multiple U in the spacer results in low sgRNA expression, T is not preferred at the four positions nearest to the PAM. The downstream nucleotides of the PAM cooperate with the efficiency of sgRNA, while upstream sequences of the spacer do not show significant effects. The early termination of sgRNA transcription is mostly responsible for the reduced sgRNA expression rates caused by the high frequency of nonconsecutive T clusters in the protospacer. Cytosine is favored at the −3 position as the cleavage site of the sgRNA/Cas9 complex.

Additionally, guanines are favored from positions −14 to −17, while adenines are favored positions −5 to −12 [89–93]. Most molecular characteristics that govern sgRNA stability, loading, and targeting *in vivo* are yet unknown. While variable Cas9 off-target binding, positioning of the nucleosome, and sgRNA loading are not key factors, adenine depletion and guanine enrichment improved sgRNA activity and stability. There is also a close correlation between sgRNA efficiency and guanine enrichment PAM-proximal site, supposedly caused by G-quadruple structure increasing sgRNA stability [94–97].

#### **4. Computational tools for designing sgRNA**

Generally, the potential off-target effect is still a vital concern for several applications of CRISPR technology. There are many strategies including Cas endonucleases engineering, transcriptome analysis, tunable systems (small-molecule induction of Cas9, light-activated and intein-inactivated Cas9), functional screening after dCas9 treatment, direct delivery (RNP complex), truncated sgRNAs (small-guide RNAs), and separate Cas9-binding approaches (paired Cas9 nickase) to reduce the off-target activity in CRISPR/Cas gene editing (**Table 3**) [105–107]. However, designing sgRNA can be the most simple, effective, low-cost, and time-saving approach [28].

There are some key factors for designing an efficient sgRNA for CRISPR editing and reducing off-target effects. First, the GC contact should be 40–80 per cent, although a higher percentage is more desirable and beneficial [28]. Second, the sgRNA length needs to be 17–20 nt, depending on the used Cas enzyme. The shorter the sequence is designed, the less the off-target effects are observed; however, too short sequences can increase the off-target effects [71]. As the third factor, off-target effects may result from mismatches between sgRNA and the target sequence, according to the mismatch numbers and positions [96]. ΔG calculation provides a significant benefit to assess sgRNA-DNA binding potential. The ΔG of a highly effective gRNA is from −64.53 to −47.09kcal/mol, but higher ΔG can increase mismatching rates, causing a high rate of the off-target effect [93]. Although the more the negative ΔG would be, the more stable the secondary structure of the sgRNA is observed, higher ΔG (~ −30 to −20 kcal/mol) can positively influence sgRNA transcription and practically make more effective RNA-RNA binding for a more functional secondary structure of the sgRNA [28].

According to all these criteria, designing sgRNA is a crucial concern in CRISPR technology [72]. As CRISPR/Cas contain two key players, including Cas endonuclease and sgRNA, to cooperate with genome editing, the development of each component could be beneficial to enhancing CRISPR/Cas editing. However, enzyme engineering


#### **Table 3.**

*Benefits and limitations of the strategies predicting off-target effects.*

is a costly, time-consuming, and complicated strategy. So, sgRNA designing could be more effective [108, 109]. Furthermore, an effective sgRNA should simultaneously show the highest on-target efficiency and the lowest off-target activity. So, several well-developed computational tools can be found to design sgRNA for high-efficiency genome editing [110]. In addition to the simplicity, high efficiency, and costeffectiveness, the *in silico* tool offers adaptability, automation, and fast processing to analyze many genes [111].

In the last decade, many *in silico* tools have been introduced to developing CRISPR technology because there has been an urgent demand to design an effective sgRNA to create precise mutations via CRISPR/Cas. Some tools have combined several scoring methods and/or algorithms to provide better design services [112, 113]. In addition to the different features of effectively designing sgRNA, these tools would be userfriendly [114]. The most popular computational sgRNA designing tools are outlined in the following and the other in silico tools are summarized in **Table 4**. The outlined tools are able to offer candidate sgRNAs and simultaneously score off-target activity. Besides, they are more user-friendly and fast-processing tools [28, 44].

#### **4.1 CHOPCHOP**

CHOPCHOP website, one of the most conventional *in silico* tools for detecting target sequences, includes a clear interface and complete functions. There are more than 200 reference genomes on the website so that the users can search for the target sequence, genomic coordinates, and name of the desired gene. The users are also able to choose two different methods to detect off-target and seven scoring approaches for on-target efficiency before sgRNA designing (**Table 5**) [117, 125, 126].

*The Prominent Characteristics of the Effective sgRNA for a Precise CRISPR Genome Editing DOI: http://dx.doi.org/10.5772/intechopen.106711*


#### **Table 4.**

*Common in silico tools for sgRNA designing in CRISPR technology.*

#### **4.2 CRISPR RGEN tools**

CRISPR/Cas-derived RNA-guided engineered nuclease (CRISPR RGEN) can provide several computational tools and sgRNA/Cas libraries, including nine tools such as Cas-OFFinder, Cas-Designer, and Digenome-Seq. Compared with other tools, Cas-Designer can rapidly detect potential off-target sites containing a DNA or RNA bulge. Besides, Cas-Designer can offer an out-of-frame score for each sgRNA to find the proper sites for the gene knockout [120, 127, 128]. Cas-OFFinder is also used to seek potential off-target positions of NmCas9 endonuclease (from *N.meningitides*), recognizing 5′-NNNNGMTT-3′ PAM sequence (M = A or C) as well as a 24-bp target sequence specific to the design sgRNA in the target genome. Also, mixed bases can be used by Cas-OFFinder to analyze the degeneracy of PAM sequences. At least, Cas-OFFinder can provide quick scanning for potential off-target positions in any sequenced genome, regardless of the mismatched nucleotides numbers or the PAM sequence limitation (**Table 5**) [120, 129, 130].

#### **4.3 CRISPOR**

Among these *in silico* tools for effectively designing sgRNA, CRISPOR includes various useful tools to design sgRNA, 417 genomes, and 19 PAM types. This *in silico* tool can receive genome coordinates and sequences with more than 2000 bp length as the inputs. After processing, comprehensive information is provided as the output. The result can be, by default, presented in two sections; first,


#### **Table 5.**

*Comparing three in silico tools.*

visualizing the PAM sites along the target sequence, available in different formats such as fasta, GenBank and SnapGene; second, providing a table containing all information such as 2 specificity scores and 10 efficiency scores for every predicted sgRNA (**Table 5**) [105, 124, 131, 132].

#### **4.4 Challenges and limitations of** *in silico* **tools**

Although the computational tools for sgRNA designing can facilitate on-target prediction and reduce off-target activity, some tools cannot cover all vital criteria. So, it is highly recommended to use different *in silico* tools. For example, some free, user-friendly, and reliable online tools include RNAfold and Mfold to predict sgRNA secondary structure [59, 79].

Moreover, all current prediction models struggle with four main challenges:


#### **5. Adenine base editors (ABEs)**

Base editors (BEs), as chimeric proteins, contain a catalytic domain and a DNA targeting modules which is able to deaminate adenines and cytosines. There is no need to make DSBs in DNA bases editing when BEs are used for base editing. So, these proteins can reduce the off-target effects and random indels at the on-target sequences. The BEs have been introduced as novel promising tools to make precise gene modification [6, 136, 137]. ABEs are the fused Cas9 nickase with a deaminase

#### *The Prominent Characteristics of the Effective sgRNA for a Precise CRISPR Genome Editing DOI: http://dx.doi.org/10.5772/intechopen.106711*

domain converting A-G and C-T (C > T) at the target sequence [138]. In fact, ABEs can effectively and precisely convert A-G and C-T base pairs at the target site within the editing frame while producing few by-products, consequently reducing off-target activity significantly. The ABE variants can improve the precision of adenine base editing by reducing the off-target activities of RNA and DNA [105, 139]. It has been reported that ABEs cooperate to induce free-sgRNA transcriptome editing. ABEs have also been discovered to display RNA off-target activity and the capacity to edit their own transcripts [140, 141]. ABEs can generally produce significant off-target singlenucleotide variations (SNVs) in RNA sequences. Therefore, deaminase engineering enables ABE variants to decrease off-target mutation of SNVs in RNA sequences while increasing on-target efficiency with DNA [105, 142].

#### **6. Conclusion**

Over the last 30 years, genome editing technology, particularly CRISPR/Cas, has promoted biosciences by editing and targeting the genomic DNAs of any species. CRISPR/Cas is the most precise, effective, and affordable among all these genome editing technologies. Although there are diverse types and classes of CRISPR/Cas systems, they are not all applicable due to the high rate of off-target effects. Different approaches have been developed to decrease the off-target effects for enhancing the precision and efficiency of the different CRISPR/Cas techniques. According to a refined reference genome, a well-designed sgRNA can support high on-target efficiency to create a precise and desirable mutation. Finally, it is highly recommended to consider the criteria as mentioned above, including GC content, length, secondary structure, and sequence, for designing an effective sgRNA to achieve high-precision CRISPR genome editing.

#### **Acknowledgements**

The chapter was funded by Amin Techno Gene Private Virtual Lab (NGO), Tehran, Iran.

### **Conflict of interest**

The authors declare no conflict of interest.

### **Author details**

Reza Mohammadhassan1 \*, Sara Tutunchi2 , Negar Nasehi3 , Fatemeh Goudarziasl4 and Lena Mahya5

1 Plant Sciences Department, Amin Techno Gene Private Virtual Lab (NGO), Tehran, Iran

2 Department of Medical Genetics, Shahid Sadoughi University of Medical Sciences, Yazd, Iran

3 Faculty of Biological Sciences, Department of Nanobiotechnology, Tarbiat Modares University, Tehran, Iran

4 Faculty of Biological Science, Department of Biochemistry, University of Mazandaran, Babolsar, Iran

5 Medical Sciences Department, Amino Techno Gene Private Virtual Lab (NGO), Tehran, Iran

\*Address all correspondence to: rezarmhreza22@gmail.com

© 2022 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

*The Prominent Characteristics of the Effective sgRNA for a Precise CRISPR Genome Editing DOI: http://dx.doi.org/10.5772/intechopen.106711*

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#### **Chapter 5**

## Recent Advances in *In Vivo* Genome Editing Targeting Mammalian Preimplantation Embryos

*Masahiro Sato, Masato Ohtsuka, Emi Inada, Shingo Nakamura, Issei Saitoh and Shuji Takabayashi*

#### **Abstract**

CRISPR-based genome engineering has been widely used for producing gene-modified animals such as mice and rats, to explore the function of a gene of interest and to create disease models. However, it always requires the *ex vivo* handling of preimplantation embryos, as exemplified by the microinjection of genome editing components into zygotes or *in vitro* electroporation of zygotes in the presence of genome editing components, and subsequent cultivation of the treated embryos prior to egg transfer to the recipient females. To avoid this *ex vivo* process, we have developed a novel method called genome-editing via oviductal nucleic acids delivery (GONAD) or improved GONAD (*i*-GONAD), which enables *in situ* genome editing of zygotes present in the oviductal lumen of a pregnant female. This technology does not require any *ex vivo* handling of preimplantation embryos or preparation of recipient females and vasectomized males, all of which are often laborious and time-consuming. In this chapter, recent advances in the development of GONAD/*i*-GONAD will be described.

**Keywords:** *in vivo* genome editing, GONAD, *i*-GONAD, preimplantation embryos, knock out, knock-in, *in vivo* electroporation, oviducts

#### **1. Introduction**

In mammals, embryogenesis begins when the oocytes (ovulated from an ovary of a female) fertilize with spermatozoa in the uterus of the female (**Figure 1**). The fertilized oocytes, called zygotes or 1-cell stage embryos, exist at the "ampulla," an area of the oviduct located near an ovary. In mice, early zygotes are surrounded by cumulus cells (also called follicular cells) and correspond to Day 0.4 of gestation (11:00 AM in the morning after mating with a male) (box in **Figure 1**). In this case, Day 0 of gestation is defined as the day when the copulation plug is recognized in the morning. Late zygotes corresponding to Day 0.7 of gestation (16:00 PM) exhibit dissociation of cumulus cells (**Figure 1**). Then, these zygotes develop into 2-cell (~Day 1.4 of gestation), 4-cell (~Day 1.8 of gestation), 8-cell (~Day 2.4 of gestation), 16-cell (also called morula; ~Day 2.7 of gestation), early blastocyst (~Day 3.4 of gestation), and

#### **Figure 1.**

*Schematic of preimplantation (Days 0.5 to 4.5; Day 0 of pregnancy is defined as the day a vaginal plug is found) and postimplantation (Days 5.5) development of mice. During preimplantation, early zygote (early 1-cell embryo) (at Day 0.4; see box), late zygote (late 1-cell embryo) (Day 0.7), 2-cell embryo (day 1.5), 8- to 16-cell embryo (Day 2.5), early blastocyst (Day 3.5), and late blastocyst (Day 4.5) float in the oviductal lumen or uterine horn. Embryos at days 0.5 to 4.5 have zona pellucida (ZP), but embryos at day 4.5 begin to escape from ZP, which is called "ZP hatching," and are ready to implant into the uterine epithelium. Note that at early zygote stage (see box), fertilized eggs are tightly surrounded by cumulus cells, but at late zygote stage, cumulus cells begin to detach from an embryo. This figure was drawn in-house and reproduced with permission from Sato et al., "Recent Advances and Future Perspectives of In Vivo Targeted Delivery of Genome-Editing Reagents to Germ cells, Embryos, and Fetuses in Mice"; published by MDPI, 2020.*

late blastocysts (~Day 4.4 of gestation) (at which stage implantation into the uterine epithelium starts). Notably, zygotes and 2-cell embryos exist in the ampulla, 4-cell to 8-cell embryos in the oviductal portion between the ampulla and isthmus, 16-cell embryos (morulae) in both the oviduct and uterus, and early- to late-blastocysts in the uterus (**Figure 1**).

For producing genetically engineered animals through pronuclear microinjection (MI) or viral infection, early zygotes are generally used [1]. To isolate early zygotes, oviducts dissected from pregnant female mice at ~Day 0.4 of pregnancy are teased using a needle at the ampulla under paraffin oil. The exposed cumulus-oocytecomplex is then transferred to a drop of hyaluronidase (HA)-containing medium. Brief incubation (~5 min at room temperature) relieves cumulus cells. The resulting "denuded" zygotes are always used for MI or viral infection.

Historically, the first attempt to obtain transgenic (Tg) mice was performed by microinjecting SV40 viral DNA into the blastocoel cavity of blastocysts [2]. The

resulting offspring had various levels of SV40 genome in their organs. In 1980, Gordon et al. [3] first produced germ-line Tg mice through MI of exogenous DNA. Since then, successful production of Tg rabbits, sheep, and pigs was reported by Hammer et al. [4]. This MI technique relies on the physical injection of any type of nucleic acids (NAs) (i.e., purified DNA fragments of 3–4 kb carrying an expression unit) using an expensive manipulator and requires personnel with specific skill. Furthermore, it generally takes 3–4 h to finish MI using ~100 zygotes per session (**Figure 2A**). Perry et al. [5] reported a novel technique to generate Tg mice through the intracytoplasmic injection of a sperm (which has been mixed with NAs) into a zygote. Later, this technique was recognized as a useful tool to introduce large sized DNA such as bacterial artificial chromosomes (BACs) and yeast artificial chromosomes (YACs) into the mammalian genome [6–8].

Mammalian zygotes, including those of mice, are surrounded by a translucent glycoprotein layer called zona pellucida (ZP), which exists as a barrier that protects the early embryo from environmental insults, including viral infections, and injury from chemical or physical substances [9, 10]. For example, ZP-enclosed embryos could not be transduced through simple incubation with solution containing lentivirus (LV), adenovirus (AV), or retrovirus (RV) [11–13]. However, injection of those viruses into the space between ZP and zygote (called "peri-vitelline space") results in successful transduction [14, 15]. Furthermore, transduction of mouse zygotes was possible when

#### **Figure 2.**

*Schematic of genome-edited mouse production through microinjection (MI) (A), in vitro electroporation (EP) (B), or genome-editing via oviductal nucleic acids delivery (GONAD) (or improved GONAD (i-GONAD)) (C). This figure was drawn in-house, and reproduced with permission from Sato et al., "Recent Advances in the Production of Genome-Edited Rats"; published by MDPI, 2022.*

they were subjected to laser perforation of ZP in the presence of LV vectors [16]. ZP can be removed by incubating ZP-enclosed embryos in the presence of proteolytic enzyme such as pronase, or under acidic conditions using acidic Tyrode's solution [1]; therefore, gene delivery can be accomplished by incubating the ZP-removed embryos in a solution containing liposome-complexed DNA [17] or the above-mentioned viruses [11, 18]. However, these treated embryos are often vulnerable, adhesive, and are easily damaged [19]. Furthermore, transferring the ZP-removed embryos (at least up to morulae) into the oviductal lumen of pseudopregnant recipient females cannot support their normal development, because the transferred embryos tend to adhere to the oviductal epithelium [20, 21]. To avoid this, the treated embryos have to be cultured at least up to blastocysts (showing reduced adhesive property), prior to uterine transfer. Notably, acquisition of Tg founders was reported using ZP-free embryos transfected liposomally [22] or when transduced with AV vectors [11, 18].

Electroporation (EP)-mediated gene delivery is also a method that efficiently introduces exogenous NAs into a cell or zygote through electric shock-induced, transient micropores in a cell membrane [23]. It requires an expensive electroporator, but does not require a skilled person, unlike in the case of MI (**Figure 2B**). In an early study regarding EP-based introduction of DNA into mouse preimplantation embryos, DNA was first injected into the peri-vitelline space of zygotes, and then the embryos were subjected to *in vitro* EP. Unfortunately, the transfection efficiency was very low [24]. In 2002, Grabarek et al. [25] first demonstrated that *in vitro* EP enabled incorporation of small-sized NAs (i.e., siRNA) into mouse zygotes. In this case, prior to EP, ZP has to be weakened by a brief treatment with acidic Tyrode's solution to facilitate transfer of NAs into an embryo and to protect the embryos from EP damage. Notably, *in vitro* EP was also successfully used to deliver double-stranded RNA (dsRNA) and morpholinos into mouse preimplantation embryos [26, 27].

To our knowledge, Peng et al. [27] first demonstrated that plasmid DNA can be effectively delivered into mouse preimplantation embryos when they were subjected to *in vitro* EP using optimal EP parameters (i.e., voltage, pulse duration, number of pulses, and repeats). Sato et al. [28] also demonstrated that plasmid DNA can be successfully introduced into early mouse embryos present within the oviductal lumen through *in vivo* EP. *In vitro* EP-based gene delivery is generally possible using over 100 zygotes per a trial and can be finished within 15–30 min. Thus, in terms of convenience, EP appears superior to the MI-based production of transgenics. However, this success appears largely to depend on the EP parameters and the type of electroporator used, as mentioned below.

Beside *in vitro* EP, MI, and viral transduction, substances capable of penetrating ZP (also called "ZP-penetrating reagents") can be used with NAs to perform gene delivery towards ZP-enclosed embryos. For examples, Ivanova et al. [29] employed a receptor-mediated gene transfer system, with insulin as the admission ligand in the DNA-carrying construct, because early embryonic cells are known to have internalizable insulin receptors on their surface [30]. They first made an insulin-polylysine conjugated with plasmid DNA. Next, this complex was mixed with a conjugate consisting of *streptavidin*-polylysine*-biotinylated* adenovirus. Short (3 h) incubation of ZP-enclosed mouse and rabbit preimplantation embryos with the resulting complex [called "(insulin-polylysine)-DNA and (insulinpolylysine)-DNA-(streptavidinpolylysine)-(biotinylated adenovirus)"] penetrated the constructs through ZP and accumulated in the peri-nuclear space of the embryos, leading to ligand/receptormediated transgenesis. Joo et al. [31] developed amphiphilic *chitosan*-based nanocarriers, called VisuFect. When murine zygotes were incubated with a solution containing

Cy5.5-labeled VisuFect conjugated with poly(A) oligonucleotides, the complex gradually penetrated the cytoplasm of the ZP-enclosed zygotes. This suggests that VisuFect could be used as a vehicle to deliver NAs to ZP-intact embryos. According to Joo et al. [31], VisuFect can deliver siRNA, but not large molecules such a plasmid DNA to embryos. Nanoparticles are also a promising tool to transfer exogenous NAs into ZP-enclosed embryos. Munk et al. [32] demonstrated that multiwall carbon nanotubes (MWNTs) can cross the ZP to help the delivery of plasmid DNA (carrying the green fluorescent protein (*GFP*) gene) into bovine embryos *in vitro*. According to Munk et al. [32], MWNTs themselves are non-harmful to embryos and do not affect their viability and gene expression. On the other hand, Jin et al. [33] first demonstrated that peptide nanoparticles can introduce siRNA into an intact mouse oocyte. When oocytes were incubated with peptide nanoparticle-complexed fluorescein isothiocyanate (FITC)-conjugated siRNA for 12–14 h, a cytoplasmic fluorescence of oocytes was observed together with a target gene knockdown. Jin et al. [33] suggested that peptide nanoparticle-mediated siRNA transfection was useful to explore the function of unknown genes in mouse oocytes. Unfortunately, except for the report by Ivanova et al. [29], these studies do not show germ-line transmission or chromosomal integration of transgenes.

As mentioned above, to obtain genetically-modified (GM) animals, isolation of zygotes from pregnant females or those obtained through *in vitro* fertilization (IVF), *in vitro* gene delivery towards the isolated embryos, transient cultivation of the treated embryos, and egg transfer (ET) of the cultivated embryos to the reproductive tracts of pseudopregnant recipient females to allow the GM embryos to develop *in vivo* further. These processes are called "*ex vivo* handling of embryos" (**Figure 2A** and **B**), and generally required for MI, *in vitro* EP, liposome- or viral transduction-based transgenesis. Notably, this *ex vivo* handling of embryos is costive, labor intensive, and laborious, because it requires preparation of sterile males called "vasectomized males" to create pseudopregnant females, timely supply of those pseudopregnant females, and ET technique, which is a more difficult task requiring people with specialized skill sets. To bypass this process, direct genetic manipulation must be performed in zygotes (or embryos at more advanced stages) existing within an oviductal lumen of a pregnant female. Relloso and Esponda [34, 35] first attempted to transfect epithelial cells lining oviductal lumen by injecting liposomally encapsulated DNA directly into the oviductal lumen of a female mouse. They found that 6% of oviductal epithelial cells were successfully transfected. Rios et al. [36] also demonstrated that intraoviductal injection of naked DNA or mRNA into the estradiol-treated female rats can help incorporate those substances into the oviductal cells. The introduced DNA or mRNA will then be translated into an active protein, possibly accelerating embryo transport.

Sato [37] employed *in vivo* EP to enhance transfection efficiency in the oviductal epithelium. *In vivo* EP is a method to transfect the tissue or organ *in situ* by injecting a fluid containing plasmid DNA into the target site, holding the injection site with tweezer-type electrodes, and subsequently giving an electric shock using an electroporator [38]. Using this system, several organs/tissues including kidney [39, 40], liver [41], brain [42], skin [43, 44], skeletal muscle [45], testis [46, 47], efferent duct [48], ovary [49], and fetuses [50–52], have been successfully transfected. According to Sato [37], ovary/oviduct/uterus were pulled out and exposed on the back of a female on Day 0.4 of pregnancy (~11:00 AM; corresponding to early zygotes). Then, a small amount (1–2 μL) of solution which contains an enhanced green fluorescent protein (eGFP) expression plasmid (0.5 μg/μL) and 0.2% (v/v) trypan blue (TB) (used as a marker for visualizing injected materials) was injected into the oviductal lumen using a mouthpiece-controlled glass pipette under a dissecting microscope. Immediately after injection, an entire oviduct was subjected to *in vivo* EP using tweezer-type electrodes in an attempt to transfect oviductal epithelium facing oviductal lumen and possibly zygotes (floating in the oviductal lumen) with the exogenous DNA. The EP condition was eight square-wave pulses with a pulse duration of 5 ms and an electric field intensity of 50 V, generated by a square-wave pulse generator (#T-820; BTX Genetronics Inc., San Diego, CA, USA). After EP, the treated oviduct was returned to the original position. One day after the surgery, oviducts were dissected from the female to isolate 2-cell embryos. When the eGFP-derived fluorescence in isolated embryos and oviducts was inspected, maximal 43% of oviductal epithelial cells facing oviductal lumen were fluorescent, while no fluorescence was discernible in the isolated embryos; only a cellular remnant probably derived from a part of zygotes was found to be fluorescent. Based on this finding, Sato [37] speculated that failure of gene delivery to zygotes may be due to the cumulus cells surrounding zygotes acting as a barrier. Because the oviductal epithelium can be efficiently transfected using *in vivo* EP method, Sato [37] named this technology "gene transfer via oviductal epithelium (GTOVE)."

Sato et al. [28] next attempted to transfect 2-cell embryos floating in the oviductal lumen using GTOVE, since those embryos are already free of cumulus cells. To address this issue, GTOVE was performed in pregnant females at Day 1.4 of pregnancy using the same conditions elaborated in the study by Sato [37]. One day after the GTOVE procedure, 8-cell embryos were collected from the GTOVE-treated females for checking eGFP-derived fluorescence. Of the 12 oviducts (6 females used) examined, 3 contained fluorescent 8-cell stage embryos (33%, 19/58 tested), but the intensity of fluorescence varied among the embryos. Unfortunately, gene expression was transient in this system, with no evidence for chromosomal integration of transgenes [28]. These results indicate that successful *in vivo* introduction of exogenous plasmid DNA into early mouse embryos is possible, as far as the T-820 electoporator is employed. However, the T-820 electoporator is currently unavailable. When we performed GTOVE to transfect 2-cell embryos using another electroporator (NEPA21; NEPA GENE Co., Chiba, Japan), the collected embryos failed to fluoresce [53]. This suggests a need to carefully examine the optimal EP condition enabling ZP penetration of larger sized molecules like plasmid DNA, as suggested by Peng et al. [27] and Hakim et al. [54].

#### **2. Development of genome editing technology**

Genome-editing technology includes zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (CRISPR/Cas9), all of which employ sequence-specific nucleases to induce modifications in a predefined region of the genome (reviewed by Harrison et al. [55]; Hsu et al. [56]). In the absence of the donor (or template) DNA [including longer genes (>1 kb), single-stranded (ss) sequences (>200 bp) or synthetic oligodeoxynucleotides (ODNs) 20–30 bp in size, all of which have sequences showing homology to the target sequence, these nucleases can induce double-strand breaks (DSBs) which are then repaired *via* nonhomologous end joining (NHEJ). This repair occurs through the cellular machinery, which frequently generates random insertions, deletions, or substitutions of nucleotides (called indels) at the break site. These indels often cause frameshift mutations, leading to the occasional

failure in protein expression. In the presence of the donor (or template) DNA, it will be introduced into the DSB site through homology-directed repair (HDR), and this event is called "knock-in (KI)." According to Yoshimi et al. [57], KI is more difficult to complete successfully than inducing NHEJ-mediated indels. Furthermore, NHEJ occurs in nondividing and dividing cells, but HDR occurs preferentially in dividing cells [58, 59].

Unlike ZFN and TALEN, the CRISPR/Cas9 gene editing system has several advantages, including easy design for any genomic targets, simplicity, and the ability to modify several target genes simultaneously (multiplexing). Owing to these properties, the CRISPR/Cas9 system is now widely used in various biological systems. It employs only two components: (1) a guide RNA (gRNA), comprised either of a duplex CRISPR RNA (crRNA)/trans-activating CRISPR RNA (tracrRNA) molecule or of single-guide RNA (sgRNA), a fusion between crRNA and tracrRNA, and (2) a Cas9 endonuclease (reviewed by Harrison et al. [55]; Hsu et al. [56]). The gRNA can bind to the specific DNA sequence together with Cas9. Once bound, the Cas9 cleaves dsDNA 3 bp upstream of the protospacer adjacent motif (PAM, 5′-NGG-3′), which is recognized by the Cas9 protein. The cleaved site is then repaired by various cellular machineries, such as NHEJ and HDR.

#### **3. Development of novel technologies enabling EP-based genome-editing**  *in vitro* **and** *in vivo*

As described in Section 2, in the late 2013, CRISPR/Cas9 system was recognized as a useful tool to manipulate a target gene in mammalian cells and embryos (reviewed by Harrison et al. [55]; Hsu et al. [56]). Since then, genome-edited animals from various species including mice, rats, pigs, bovines, and primates (monkeys) have been generated through MI [60–66]. Kaneko et al. [67] first demonstrated that *in vitro* EP of rat zygotes in the presence of genome editing reagents is a powerful tool to produce GM animals (**Figure 2B**). According to Kaneko et al. [67], intact rat zygotes were electroporated using an NEPA21 electroporator in a solution containing ZFN (40 μg/mL) mRNA [targeted to the rat interleukin 2 receptor subunit gamma gene (*Il2rg*)] under the EP condition of a poring pulse (Pp) (voltage: 225 V; pulse interval: 50 ms; pulse width: 1.5 and 2.5 ms; number of pulses: 4). The mechanism underlying gene delivery by this system is described in our previous paper [68]. As a result, they obtained genome-edited rat offspring with an efficiency of 73%, which is roughly 2-fold higher than that obtained through MI-mediated genome editing. Notably, they confirmed germ-line transmission of the genome-edited traits beyond next generation. This technology was thus named "technique for animal knockout system by electroporation (TAKE)." Since then, various genome-edited animals including mice [69–75], rats [76–78], and pigs [79–81] have been successfully generated using this technology. Furthermore, this technology has been applied to introduce nucleases into frozen embryos [82] and embryos derived from freeze-dried and frozen sperm [83].

Takahashi et al. [84] developed a novel *in vivo* EP-based method enabling *in situ* CRISPR/Cas9-based genome editing towards early mouse embryos (2-cell embryos) using a technique similar to GTOVE (**Figure 2C**). In this case, a solution (1–1.5 μL) containing *Cas9* mRNA (up to 1.1 μg/μL), sgRNA (0.6 μg/μL; targeted to *eGFP* cDNA), and 0.2% (v/v) TB was injected through the oviductal wall under observation into the oviductal lumen of pregnant non-Tg females at Day 1.4 of pregnancy (corresponding to the 2-cell stage) that had been mated with Tg males containing an *eGFP* expression cassette in a homozygous manner [85]. In this case, the pregnant

female should have zygotes, all of which are expected to have *eGFP* expression cassette in a heterozygous manner, and be fluorescent (**Figure 3**). If CRISPR/ Cas9-induced mutations (indels) occur in the genomes of those zygotes (*eGFP* cDNA), some of their offspring will most likely lose fluorescence from their entire body. Fluorescence inspection of mid-gestational fetuses dissected from the *in vivo* EP-treated females demonstrated that out of 6 fetuses obtained, two lost fluorescence completely, two exhibited weak fluorescence, and fluorescence in the remaining two fetuses remained unaltered. Molecular biological analysis revealed that the former fetuses comprised knock out (KO) cells, the middle fetuses contained a mixture of KO cells and intact cells (whose state is called "mosaic mutation"), and the latter fetuses had unedited intact cells. Based on these findings, the KO efficiency of these fetuses

#### **Figure 3.**

*Experimental flowchart for genome-editing via oviductal nucleic acids delivery (GONAD). Females (C57BL/6) are first mated to C57BL/6-Tg (CAG-eGFP) male mice that possess EGFP transgenes in a homozygous (Tg/Tg) state. All the fetuses are then expected to be eGFP-expressing fetuses carrying the transgenes in a heterozygous (Tg/+) state. Thus, successful genome editing targeted to eGFP at preimplantation stages are expected to reduce the levels of eGFP fluorescence in the mid-gestational fetuses, as a result of genome editing in the chromosomally integrated eGFP transgenes. When GONAD was performed using Cas9 mRNA and sgRNA towards pregnant females at Day 1.4 of pregnancy, there were three types of fetuses (showing complete loss of fluorescence, partial fluorescence, or no reduction in fluorescence) in view of fluorescence expression pattern.*

can be estimated to be approximately 29%. Based on these findings, Takahashi et al. [84] named this technology "genome-editing via oviductal nucleic acids delivery (GONAD)."

#### **4. Development of improved GONAD (***i***-GONAD)**

In 2018, Ohtsuka et al. [86] in the same group of Takahashi et al. [84] further elaborated on the GONAD technology. As was mentioned in Section 3, GONAD enables *in situ* genome editing in 2-cell embryos. In this case, only one blastomere among two blastomeres can be genome-edited, generating "mosaic" fetuses comprising edited and non-edited cells. To avoid this risk, genome editing at zygote (one-cell) stage is desirable. As was mentioned previously, early zygotes (corresponding to Day 0.4 of pregnancy; 11:00 AM) are tightly surrounded by cumulus cells. In our previous experience, GONAD at this stage failed, because almost all of the genome editing reagents introduced intaoviductally was trapped by the cumulus cells [86]. On the other hand, as mentioned previously, the detachment of cumulus cells from a zygote commences at late zygote stage (corresponding to Day 0.7 of pregnancy; 16:00 PM). Based on this finding, Ohtsuka et al. [86] first attempted to disrupt the forkhead box protein E3 (*Foxe3*) locus using ribonucleoprotein (RNP) (comprised of 1 μg/μL of Cas9 protein and 30 μM of crRNA/tracrRNA) using pregnant females at Day 0.7 of pregnancy. The advantage of using RNP is to induce genome editing more rapidly than using *Cas9* mRNA [87]. *In vivo* EP was performed using the NEPA21 apparatus under the following conditions: Pp: 50 V, 5-ms pulse, 50-ms pulse interval, three pulses, 10% decay (± pulse orientation); Tp: 10 V, 50 ms pulse, 50 ms pulse interval, three pulses, and 40% decay (± pulse orientation). This modification resulted in 97% of the embryos exhibiting indels in the target locus. They next attempted to perform KI of a sequence (coding for a gene of interest (GOI)) into the target locus using RNP containing 1–2 μg/μL of ssODN or 0.85–1.4 μg/μL of ssDNA (with ~925 bases in size) generated through a novel method, called Easi-CRISPR, a highly efficient KI technique using ssDNA as donor templates [88, 89]. As a result, ~50% and ~ 15% of embryos were found to have KI alleles for ssODN and longer ssDNA in their genome, respectively.

Ohtsuka et al. [86] also demonstrated that large deletion (LD) of a target sequence can be accomplished using this modified GONAD, which was re-named as "*i-*GONAD." It uses the Cas9 protein instead of the *Cas9* mRNA, and targets late zygotes. For example, they designed two gRNAs (16.2 kb distance apart), both of which recognize either the sites of the retrotransposon sequence in the intron 1 of *Agouti* locus in the C57BL/6 mouse genome. The *i-*GONAD-mediated deletion of the inserted sequence resulted in the generation of fetuses with agouti coat color with efficiencies of 50%. Molecular biological analysis of these rescued offspring revealed the evidence for LD in the *Agouti* locus containing retrotransposon of C57BL/6 mice.

Since the studies by Takahashi et al. [84] and Ohtsuka et al. [86], several reports have been provided using GONAD/*i*-GONAD technologies. In **Table 1**, past studies using those technologies are listed. Also, the detailed protocols for *i*-GONAD in mice have been provided by Gurumurthy et al. [116, 117] and Ohtsuka and Sato [118]. The GONAD/*i*-GONAD-based production of genome-edited rats is also possible using the same approach shown in mice. Notably, Namba et al. [119] demonstrated the protocols for GONAD/*i*-GONAD in rats.








*Abbreviations: AAV, adeno-associated virus; Agouti, Agouti-signaling protein (ASIP); AIMAs, anti-inhibin monoclonal antibodies; AMPAR, α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid receptor; ATF5, activating transcription factor 5; Axdnd1, axonemal dynein light chain domain containing 1; B6C3F1, a cross between female C57BL/6 and male C3H/He mice; B6D2F1, a cross between female C57BL/6 and male DBA/2 mice; BN, Brown Norway rat; Cas12a (AsCas12a), class 2 CRISPR-Cas endonuclease Cas12a (previously known as Cpf1); CCK, cholecystokinin; Cdkn1a, cyclin dependent kinase inhibitor 1A; Cdkn2a, cyclin dependent kinase inhibitor 2A; Clcf1, cardiotrophin-like cytokine factor 1; COL4A3, collagen type IV α3 chain; COL4A4, collagen type IV α4 chain; COL4A5, collagen type IV α5 chain; COL7A1, collagen type VII α1 chain; CRISPR/Cas9, clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9; DA, Dark Agouti rat; Dll1, Delta like canonical Notch ligand 1; eEF2, eukaryotic translation elongation factor 2; eGFP, enhanced green fluorescent protein; EP, electroporation; Fgf10, fibroblast growth factor 10; Fosl1, Fos-like antigen 1; Foxe3, forkhead box E3; Gbx2, gastrulation brain homeobox 2; GGTA1, α-1,3-galactosyltransferase; GONAD, genome-editing via oviductal nucleic acids delivery; Hprt, hypoxanthine guanine phosphoribosyl transferase; i-GONAD, improved GONAD; indels, insertions, deletions, or substitutions of nucleotides; KI, Knock-in; Kit, KIT proto-oncogene, receptor tyrosine kinase; Klrc2, killer cell lectin like receptor C2; KO, Knock out; LD, large deletion; LEW, Lewis rat; Mecp2, methyl-CpG binding protein 2; Mov10l1, Mov10 like RISC complex RNA helicase 1; Nrsn2, Neurensin 2; Pafah1b1, platelet activating factor acetylhydrolase 1b regulatory subunit 1; Pax6, paired box 6; Pitx3, paired like homeodomain 3; Plagl1, pleiomorphic adenoma gene-like 1; Qa-1b, a 48 kDa non-classical MHC class Ib molecule; rAAV, recombinant adeno-associated viruses; Rad51, RAD51 recombinase; RDEB, recessive dystrophic epidermolysis bullosa; SD, Sprague–Dawley rat; Serpina3n, serpin family A member 3; si-GONAD, sequential i-GONAD; spCas9, Streptococcus pyogenes-derived Cas9; ssODN, single-stranded oligodeoxynucleotide; Tis21, TPAinducible sequences 21; Tg, transgenics; Tprkb, TP53RK binding protein; Tyr, tyrosinase; WKY, Wistar Kyoto rat.*

#### **Table 1.**

*Summary of genome-edited animals produced using GONAD/i-GONAD between the years 2015–2020.*

#### **5. GONAD/***i***-GONAD in rats**

Rats (*Rattus norvegicus*) and mice (*Mus musculus*) are both classified into the same rodent family, and have been recognized as the most widely used models for biomedical research during the past four decades. However, these animals are different on many factors. For example, the rat is larger (roughly about eight- to ten-fold) in size than the mouse. Therefore, rats have been extensively used for pharmacological and surgical research, as exemplified by easier and more rapid microsurgery, multiple sampling of blood and tissues with relatively large amounts, and precise injection of substances into blood vessels (reviewed by Kjell and Olson [120]). Additionally, rats are considered as animals suitable for toxicological, neurobehavioral, and cardiovascular studies (reviewed by Jacob [121]). Since the first report in 1997 by Guerts et al. [122] on the production of genome-edited rats using ZFN technology, a total of 113 GM rats by MI (including pronuclear MI and cytoplasmic MI) or 9 rats by

*in vitro* EP method have been produced during the period 1997–2021 (reviewed by Sato et al. [68]). Notably, GM rats often exhibited disease phenotype similar to that observed in humans, comparable to those shown in GM mice. For example, Zhang et al. [123] produced KO rats through MI of CRISPR/Cas9 components to obtain rat models for hereditary tyrosinemia type I (HT1), a disease caused by a deficiency in fumarylacetoacetate hydrolase (FAH) enzyme. These *Fah* KO rats developed remarkable liver fibrosis and cirrhosis, which have not been observed in *Fah* mutant mice. Furthermore, dystrophin-coding gene (dystrophin) (*Dmd*) KO rats (but not mice) presented cardiovascular alterations close to those observed in humans, which are the main cause of death in patients [124, 125]. These findings encourage the speculation that rats may better mimic the human situation than mice.

In 2018, two groups in Japan succeeded in generating genome-edited rats using *i*-GONAD. Kobayashi et al. [90] examined optimal *in vivo* EP conditions, allowing successful *i*-GONAD using NEPA21 electroporator and red fluorescent dextran (RFD) (tetramethylrhodamine-labeled dextran 3 kDa) as a fluorescent dye to monitor the fate of injected materials. Wistar Kyoto (WKY) strain rats were first subjected to the intraoviductal injection of a solution containing RFD + TB into pregnant female rats at Day 0.7 of pregnancy and subsequent *in vivo* EP using the NEPA21 electroporator under the following EP conditions: Pp: 30, 40, or 50 V, 5-ms pulse, 50-ms pulse interval, 3 pulses, and 10% decay (± pulse orientation); Tp: 10 V, 50-ms pulse, 50-ms pulse, 6 pulses, and 40% decay (± pulse orientation). When two-cell embryos recovered one day after *i*-GONAD were inspected for RFD-derived fluorescence, EP with 50 V for Pp and 6 times for Tp yielded maximal fluorescence in those embryos, with 74% efficiency. Using these optimal EP conditions, they attempted to disrupt the endogenous tyrosinase (*Tyr*) gene, a gene coding for protein essential for eye pigmentation*,* in pigmented females through intraoviductal injection of RNP (1 μg/μL of Cas9 protein +30 μM of sgRNA targeted to *Tyr*). As a result, genome-edited pups with albino-colored coat were obtained with an efficiency of 42%, when a Pp of 50 V was employed. They also attempted to recover the coat-color mutation in WKY females using an ssODN-based KI approach. Consequently, the KI efficiency was 27% in the pups born. They named this rat-based *i*-GONAD as "rGONAD."

Takabayashi et al. [91] provided data similar to that of Kobayashi et al. [90], demonstrating that *i*-GONAD-based mutations (indels) resulted in the generation of fetuses (derived from pigmented Brown Norway (BN) × albino SD rat crosses) with non-pigmented eyes, with an efficiency of 56%. In this case, the *in vivo* EP condition used was almost the same as that used for *i*-GONAD in mice. They also tested the possibility of KI (targeted *Tyr* locus) using albino Lewis (LEW) rats and demonstrated that the *i-*GONAD-mediated KI efficiency was as low as ~5%, when the presence of fetuses with pigmented eyes was assessed. Takabayashi et al. [91] further attempted to disrupt another endogenous gene, paired box 6 (*Pax6*), an essential locus required for facial development, using *i*-GONAD. Out of 8 mid-gestational fetuses obtained, three had completely lacked eyes and lateral nasal prominence.

#### **6. GONAD/***i***-GONAD in hamsters**

The golden hamster (*Mesocricetus auratus*) is a small rodent that has been extensively used in biomedical research in fields including oncology, immunology, metabolic disease, cardiovascular disease, infectious disease, physiology, and behavioral and reproductive biology [126]. In 1976, hamster oocytes were first used for IVF assay to

test the fertilizing ability of human spermatozoa [127]. However, hamster embryos are highly vulnerable to *in vitro* conditions, hindering the generation of GM hamsters [128].

Hirose et al. [96] attempted to produce GM hamsters using *i-*GONAD, which allows embryo manipulation under the environment where the effects of handling embryos *in vitro* can be avoided as possible. Through intraoviductal injection of six sgRNAs (targeted *acrosin* gene) and the Cas9 protein, they produced KO hamsters lacking expression of acrosin, a protein thought to be essential for sperm penetration through ZP, to investigate how acrosin-KO hamster spermatozoa behaved both *in vivo* and *in vitro.* A total of 15 pups obtained, eight of which were weaned. Of these, five were found to have mutant alleles. Homozygous mutant males were completely sterile, as the mutant spermatozoa attached to ZP, but failed to penetrate it. This finding indicates that in hamsters, acrosin plays an indispensable role in allowing fertilizing spermatozoa to penetrate ZP.

#### **7. Application of GONAD/***i***-GONAD**

#### **7.1 Two-step** *i***-GONAD can introduce mutations at two sites located close to each other**

CRISPR/Cas9-based introduction of indels into two sites (which are closely located each other) through a single shot of transfection has been recognized difficult, because frequent deletion between these two sites occurs. However, sequential transfection of genome editing reagents can avoid such deletion between the two sites. To test the possibility, Sato et al. [97] tried to generate two types of indels at two target sites (that are located very close to each other; 44 bp apart) by performing *i*-GONAD sequentially (**Figure 4**). The two gRNAs were first designed to recognize the upper and lower portion of exon 4 of α-1,3-galactosyltransferase gene (*GGTA1*), coding for the protein essential for synthesizing the cell-surface α-Gal epitope [129]. For the 1st *i*-GONAD, a solution containing Cas9 protein, sgRNA (termed "A") recognizing the upper portion of exon 4, and dye (Fast Green FCF; for monitoring injection process) was injected intraoviductally at Day 0.7 of pregnancy (corresponding to late zygote stage). Next day, a solution containing Cas9 protein, sgRNA recognizing the lower portion of exon 4 (termed "B"), and dye was injected intraoviductally at Day 1.7–1.8 of pregnancy (corresponding to 2-cell stage). One day after the final surgery, morulae were isolated for single embryo-based analysis for possible indels at the target sites. As a result, the efficiency of successful generation of morulae with indels at both two sites was 18%. In contrast, *i*-GONAD using two sgRNAs (A and B) + Cas9 protein at Day 0.7 of pregnancy failed to generate morulae with mutations in both sites at exon 4 of *GGTA1*. Based on these findings, Sato et al. [97] named this approach "sequential *i*-GONAD (*si*-GONAD)."

#### **7.2 Preparation of floxed mice using** *i***-GONAD**

The Cre/*loxP* system is a useful tool for assessing *in vivo* gene function. Spatially and temporally-controlled expression of Cre recombinase enables precise deletion of *loxP*-floxed chromosomally integrated GOI. To realize it, two *loxP* sites must be simultaneously inserted in *cis* into the target locus. The resulting mice are called "conditional KO mice." Previously, this process for conditional KO mouse production was achieved by embryonic stem (ES) cell-based gene targeting and subsequent chimeric

#### **Figure 4.**

*Schematic of the detailed procedure of sequential improved genome-editing via oviductal nucleic acid delivery (si-GONAD). After first i-GONAD on Day 0.7 of pregnancy, second i-GONAD is performed as shown in the panel on Day 1.7 of pregnancy. This figure was drawn in-house, and reproduced with permission from Sato et al., "Sequential i-GONAD: An Improved In Vivo Technique for CRISPR/Cas9-Based Genetic Manipulations in Mice"; published by MDPI, 2020.*

mouse production, which is time-consuming and labor intensive [130]. To bypass this tedious process, attempts to produce mice carrying *loxP*-floxed GOI (which are generally called "floxed mice") have been made using genome editing technology. In the initial experiments, simultaneous injection of Cas9, two pairs of gRNAs, and two ssODNs containing *lox* sequences into mouse zygotes generates mice containing floxed alleles [61, 131–134]. This approach can generate floxed mice without using ES cells, since it does not require the construction of a KI vector, and production of floxed mice is finished in a short period of time (e.g., in a month). However, according to Horii et al. [130], the simultaneous introduction of two mutated *lox* sites

(to which Cre recombinase bind) at a target locus is difficult as it often causes LD of a sequence. To increase the possibility that the two *lox* sites are knocked-in, they used the sequential introduction approach to perform KI of mutated *lox* sites into the introns interposing exon 3 of methyl-CpG binding protein 2 gene (*Mecp2*) through *in vitro* EP. When the resulting embryos (blastocysts) were subjected to molecular analysis, 21% (33/155 tested) of the embryos had two floxed sites in the target *Mecp2* locus. Furthermore, the efficiency of generating LD was 36% (56/155 tested). Sato et al. [97] employed all the genome-editing components (gRNAs and ssODNs containing mutated *lox* sites as donor DNA) described by Horii et al. [130] for *si*-GONAD. Unfortunately, the generation of morulae with KI alleles (in which floxed sites had been knocked-in in both sides of the introns interposing exon 3 of *Mecp2*) failed; only a morula with one floxed site in the 5′ site of *Mecp2* was successfully generated.

Recently, Shang et al. [101] developed a new approach by integrating a unique design of asymmetric *loxP*-ssODN to create mouse conditional KO alleles in one step using the *i*-GONAD method. They injected a cocktail containing Cas9 protein, two gRNAs targeting the intron 2 and 3′ region of Fos-like antigen 1 gene (*Fosl1*), and two short ssODNs as HDR donors for *loxP* insertions. Each ssODN is 161 nucleotide (nt) long, composed of 91 nt of the 5′ homology arm from the PAM-proximal side, 34 nt of *loxP* sequence, and 36 nt of the 3′ homology arm from the PAM-distal side. Molecular biological analysis of the resulting pups demonstrated that out of 20 F0 mice obtained one mouse had the simultaneous 5′- and 3′-*loxP* insertions and 6 had either 5′- or 3′-*loxP* integrations. Similar experiments were also conducted to obtain floxed mice for genes coding for pleomorphic adenoma gene-like 1 (*Plagl1*), *Ak040954*, cardiotrophin-like cytokine factor 1 (*Clcf1*), and *Gm4438*. The overall targeting efficiency of producing floxed alleles by *i*-GONAD was 10% (8/76 tested).

#### **7.3 RFD is a useful reagent to master GONAD/***i***-GONAD in 2 days**

RFD has been recognized as a useful reagent to judge the success of gene delivery to early mammalian embryos after *in vitro* EP [67]. It has also proved useful for judging the success of GONAD/*i*-GONAD [53, 86, 90, 91, 97, 119]. For example, when we performed *i*-GONAD using a solution containing RFD and TB (or Fast Green FC) at Day 0.7 of pregnancy (corresponding to late zygote stage; 16:00 PM), the success of the approach can be judged by examining the RFD-derived red fluorescence in the isolated 2-cell embryos under fluorescence microscopic observation one day after *i*-GONAD. The presence of fluorescent embryos means successful *i*-GONAD, while the absence of fluorescent embryos indicates failure of *i*-GONAD. This short-term experience is especially beneficial for the beginners who want to master the technique, and is thus called "2-day protocol for mastering GONAD" [117]. Notably, Sato et al. [97] showed that FITC-labeled fluorescent dextran is also effective for reporting the success of gene delivery in mouse early embryos.

#### **7.4 Chromosomal engineering using** *i***-GONAD**

As shown in Section 7–1, CRISPR/Cas9-mediated genome editing using 2 types of gRNAs results in frequent generation of LD in the sequence flanked by the two sites recognized by these gRNAs. Iwata et al. [93] applied this technique to introduce chromosomal inversions of several megabases (Mb) in mice. When mouse zygotes were subjected to *in vitro* EP, a 7.67Mb inversion was successfully introduced, which is longer than any previously reported inversion produced using MI-based methods.

They confirmed that a similar event can be induced using *i*-GONAD. These findings suggest that CRISPR/Cas9 system *via in vitro* and *in vivo* EP is useful for examining genetic diseases with large-scale chromosomal rearrangements.

Notably, the same group [107] recently demonstrated that *i*-GONAD can be useful to maintain lethal mutant mice using an inversion balancer identified from the C3H/ HeJJcl strain. As a proof-of-principle, they created the Tp53rk binding protein gene (*Tprkb*) KO strain with an embryonic lethal mutation through *i*-GONAD in the presence of a non-targeted B6.C3H-In(6)1 J inversion. Iwata et al. [107] demonstrated that the edited lethal genes were stably maintained in heterozygotes, as recombination is strongly suppressed within this inversion interval. This strategy may facilitate further analysis of lethal mutants.

#### **7.5 Viral transduction using** *i***-GONAD**

Virus-based gene delivery approaches have been widely used in the biomedical sciences, especially for gene therapeutic purposes (reviewed by Sung and Kim [135]). The viral vectors widely used are RV, AV, LV, and adeno-associated viral vector (AAV). Each of these vectors have specific properties. For example, RV and LV can infect both dividing and senescent cells and enable chromosomal integration. AV can infect mainly dividing cells efficiently, but cannot integrate into host chromosomes. These viral vectors (but not AAV) can infect ZP-removed early embryos, but not ZP-enclosed (or intact) embryos [13]. Notably, the simple incubation of ZP-enclosed embryos with recombinant AAV (rAAV)-containing medium was recently shown to lead to the transduction of those embryos [13, 92]. Notably, there are over 10 different serotypes of AAVs, each of which exhibits different infectious ability depending on the type of cells [136]. Mizuno et al. [13] examined which serotype of AAV could effectively transduce ZP-intact mouse 2-cell embryos. The embryos were co-incubated for 16 h with several types of rAAVs carrying an eGFP expression unit and then transferred to normal medium; the morulae developing after co-incubation with AAV serotype 6 (which is hereinafter called rAAV-6) exhibited strong fluorescence [13]. The next vector showing relatively strong infectivity was rAAV-1. A similar observation was also made by Yoon et al. [92]. Importantly, rAAV-6 can transduce rat and bovine embryos [13], suggesting the multi-species infectivity of this vector.

However, genome editing could not be induced in early embryos through transduction with rAAV-6. The rAAV carrying Cas9 gene (~9 kb in size) and that carrying gRNA could not be co-delivered, because rAAV was unable to incorporate over 4.5-kb of an insert [137]. To perform successful CRISPR-based KI in mice, Mizuno et al. [13] employed a two-step gene delivery approach. Zygotes were first subjected to *in vitro* EP in the presence of RNP and then transduced with rAAV-6 carrying a 1.8-kb GFP expression cassette flanked by two 100-bp *Rosa26* homology arms. Molecular biological analysis of the newborn pups demonstrated that the KI efficiency in the *Rosa26* locus was 6%. Yoon et al. [92] performed intraoviductal injection of a solution containing rAAV6-Cas9 (carrying *spCas9* gene derived from *Streptococcus pyogenes*) and rAAV6-gTyr (carrying gRNA expression unit targeted *Tyr*) into the pregnant female mice at Day 0.5 of pregnancy, similar to GONAD/*i*-GONAD. Molecular biological analysis of the newborn pups demonstrated that the indel efficiency was 6%. All mutated founder (F0) mice generated albino offspring, indicating germ-line transmission; this suggested that AAV is a powerful tool for inducing genome editing in the ZP-enclosed early embryos *in vivo*. According to Sato et al. [138], this *in vivo* approach is referred to as "AAV-based GONAD."

As was mentioned earlier, AAV-based GONAD appears to be more convenient than GONAD/*i*-GONAD, since the former does not require any of the apparatus required for EP. Unfortunately, it still requires more detailed information concerning (1) which rAAV serotype is effective for *in vivo* transduction towards early mouse embryos, (2) the stage allowing maximal expression of GOI (included in rAAV) after infection at late zygote stage, and (3) whether the oviductal epithelial cells are infected through AAV-based GONAD. Sato et al. [139] first examined the above possibility using 4 types (1, 2, 5, and 6) of rAAVs carrying a unit for expression of eGFP (as GOI). When AAV-based GONAD was performed at Day 0.7 of pregnancy, and 2 days later the morulae were isolated to inspect eGFP fluorescence, rAAV-6 gave strongest fluorescence, though the fluorescence intensity varied among embryos. The fluorescence intensity provided by rAAV-1 was the next highest, but transduction with the other remaining serotypes (2 and 5) resulted in negative or faint fluorescence in the embryos. These results are consistent with the *in vitro* data from Mizuno et al. [13] and Yoon et al. [92]. A similar mode of transduction was also seen in the oviductal epithelial cells, suggesting the use of rAAV-6 (or possibly rAAV-1) for genome manipulation of those cells.

AAV-based GONAD using rAAV-6 was performed at Day 0.7 of pregnancy, and one day later, 2-cell embryos were isolated and cultured until the late blastocyst stage to monitor the eGFP fluorescence expression. Under the fluorescence microscope, fluorescence was first discernible at the 2-cell stage, attained at a maximal level at the morula stage, and declined towards late blastocyst stage [139]. These results suggest that one-day infection with rAAV-6 is enough to transduce ZP-enclosed zygotes floating in the oviductal lumen. Furthermore, the GOI expression was transient, peaking at the morula stage. These findings suggest a possibility that early mouse embryos from zygote to morula stages can be effectively transduced in a sequential manner, like *si*-GONAD.

#### **7.6 Effect of HA treatment on the efficiency of** *i***-GONAD-mediated genome editing**

As was mentioned in Section 4, *i*-GONAD at Day 0.4 of pregnancy (corresponding to early zygote) often failed to obtain genome-edited embryos/fetuses. This appears to be solely due to the presence of cumulus cells that tightly surround a zygote. One idea to overcome this problem may be that early zygotes are pretreated with HA, an enzyme capable of dispersing cumulus cells from a zygote [1], prior to *i*-GONAD. Kaneko and Tanaka [100] examined the possibility by injecting 1 μL of 0.1% HA into the ampulla of a female (ICR) at Day 0.4 of pregnancy (10:00–11:00) using a thin glass needle. As a control, the solvent (PBS) was similarly injected. Several minutes after the injection, a solution (1 μL) containing genome editing reagents (2 μg/μL Cas9 protein +60 mM dual gRNA (targeted fibroblast growth factor 10 gene (*Fgf10*) + 0.08% TB) was intraoviductally introduced and subsequently the entire oviducts were subjected to *in vivo* EP using tweezer-type electrodes. After that, the developing fetal offspring were isolated for examining the presence of possible genome editing in those samples. Consequently, the samples isolated from HA-treated group exhibited 2.5-fold higher genome editing (indels) efficiency than those isolated from the control group (68% *vs.* 27%). The *i*-GONAD on Day 0.7 of pregnancy (16:00–17:00; in which case no HA is used) yielded genome edited pups with an efficiency of 54%. These findings indicate that HA-mediated removal of cumulus cells at Day 0.4 of pregnancy is effective when *in situ* genome editing towards early

zygotes are intended. According to Kaneko and Tanaka [100], the operation time for introducing genome editing reagents into embryos in the oviducts can be adjusted by treatment with HA before EP. This improved protocol can also be used for efficient production of genome-edited mice and rats.

#### **7.7 Strain-difference can affect the efficiency of** *i***-GONAD-mediated genome editing**

According to Ohtsuka et al. [86], successful *i*-GONAD relies on the mouse strain used. For example, it worked successful under relatively stringent electrical conditions (40 V/100–200 Ω/~300 mA) when random-bred mice (such as MCH(ICR) and B6C3F1, a hybrid between C3H/He and C57BL/6), but not C57BL/6 strain, were used. Under less stringent conditions (40 V/350–400 Ω/~100 mA), *i*-GONAD was successful in the inbred C57BL/6 strain [86, 98, 117]. These findings suggest the importance of selecting the appropriate EP conditions, particularly when different mouse strains are used for *i*-GONAD experiment.

This is also true for *i*-GONAD using rats. For example, when a current of >500 mA was employed using the NEPA21 electroporator, albino SD and albino LEW rats were successfully genome-edited; however, no offspring were derived from pigmented BN rats (fetuses/newborns) [91]. In contrast, *i*-GONAD was performed under a current of 100–300 mA using the NEPA21 electroporator, leading to the production of genome-edited BN rats at efficiencies of 75–100% [99]. Similar success in producing GM BN rats was achieved with efficiencies of 24–55% when another electroporator CUY21EDIT II (BEX Co., Ltd., Tokyo, Japan) was employed under a current of 150–200 mA [99].

Notably, the most widely used electroporators (as exemplified by NEPA21) employ a constant voltage. Also, other electroporators (as exemplified by GEB15 (BEX Co., Ltd.)) employ a constant current. Kobayashi et al. [98] explored the conditions allowing the generation of a 100 mA current in C57BL/6 mice using two electroporators, NEPA21 and GEB15. As a result, *i-*GONAD performed under conditions of average resistance of 367 Ω and average voltage of 116 mA resulted in the production of genome-edited fetuses with efficiency of 39%.

#### **7.8 Attempt to increase the efficiency of KI using** *i***-GONAD**

In our previous study using *i*-GONAD to produce GM rats and KO/KI rats, the success rate of producing KI rats was lower than that of KI mice (5% *vs.* 60%, respectively) when ssODNs were used as KI donors [91, 117]. To improve the efficiency of *i*-GONAD in rats, Aoshima et al. [112] examined the effects of commercially available KI-enhancing drugs (including SCR7, L755,507, RAD51-stimulatory compound 1 (RS-1) and Alt-RⓇ HDR Enhancer (HDR enhancer)), some of which have been known to increase KI efficiency in culture cells and early embryos [140–143]. For example, *i*-GONAD was applied to SD female rats (albino) using a solution containing RNP complex (consisting of Cas9 protein and gRNA targeted *Tyr* locus), ssODN (used as a KI donor oligodeoxynucleotide), and various amounts (5 or 15 μM) of L755,507 on Day 0.7 of pregnancy. Inspection of mid-gestational fetuses revealed that 12% of fetuses obtained showed pigmented eyes when 5 μM L755,507 was used for *i*-GONAD, suggesting successful KI [112]. In addition to L755,507, some drugs (e.g., SCR7 and HDR enhancer) were found to be effective in *i*-GONAD in rats, but their effects were limited.

#### **Figure 5.**

*Schematic of knock-in (KI) experiment in rats towards the mutated tyrosinase gene (Tyr) locus performed by Aoshima et al. [112]. The target sequence (exon 2 of Tyr) recognized by crRNA1, 2, and 3 is shown in green. PAM sequences are underlined. Single-stranded oligodeoxynucleotide (ssODN) (containing wild-type nucleotide "G" that corresponds to mutated nucleotide "A") is shown in orange below the target sequence. The nucleotide "A/T" marked in red is the mutation causing the albino phenotype. This figure was drawn in-house, and reproduced with permission from Aoshima et al., "Modification of improved-genome editing via oviductal nucleic acids delivery (i-GONAD)-mediated knock-in in rats"; published by BioMed Central Ltd, 2021.*

In a study by Aoshima et al. [112], three gRNAs (called crRNA1, crRNA2, and crRNA3) were used. As shown in **Figure 5**, these gRNAs recognize different portions of the target locus, but also overlap each other in the target locus. Surprisingly, the KI efficiency in rat fetuses generated after *i*-GONAD with crRNA2 and ssODN was significantly higher (24%) than crRNA1 (5%) or crRNA3 (0%). The KI efficiency of *i*-GONAD with triple gRNAs was 11%. These findings demonstrated that the choice of gRNA is important for determining KI efficiency.

#### **7.9 Regulated timing of** *i***-GONAD by administration of gonadotrophins**

The *i*-GONAD experiment using C57BL/6 strain is always associated with the difficulty in consistently obtaining pregnant females, because estrous females are not always available. The administration of gonadotrophins has been frequently used for inducing superovulation in many mouse strains to obtain a number of early embryos [1]. This approach has an additional advantage in that it is capable of synchronizing the estrous cycle of females; thus, the estrous cycle need not be examined through smear testing or through visual inspection of the vagina.

Administration of higher dose (in this case, more than 5 international units (IU)) of gonadotrophins can induce superovulation, but often causes failure to deliver pups [144–146]. Administering low doses (less than 5 IU) of gonadotrophins facilitates ovulation of natural number of oocytes and successful delivery of pups [147, 148]. Notably, Sato et al. [53] reported that intraperitoneal (IP) administration of low-dose (2–0.5 IU) serum gonadotrophin (PMSG) from a pregnant mare on 11:00, followed by 5 IU of human chorionic gonadotrophin (hCG) 48 h later, is effective for inducing natural ovulation before *i*-GONAD. In case of administration of 5 IU PMSG, females having vaginal plug failed to deliver their pups. When females were inspected later, some had dead fetuses in their uterus. When 2 or 0.5 IU PMSG was administered, all females successfully delivered viable pups (average: 8 in each group). These findings suggest that *i-*GONAD can be performed on 11:00 at Day 0.7 of pregnancy when females were induced to ovulate by administering a low dose of PMSG. Indeed, Sato et al. [97] demonstrated that *i-*GONAD on 11:00 at Day 0.7 of pregnancy leads to generation of genome-edited morulae.

Unfortunately, the regime shown by Sato et al. [53] has only been used successfully on B6C3F1 hybrid mice. Kobayashi et al. [98] examined whether the administration of a single IP injection of low-dose PMSG **(**1.2 IU/10 g**)** is effective for synchronizing the estrous cycle in C57BL/6 females. Consequently, approximately 51% of C57BL/6 females had plugs upon mating with males 2 days after PMSG administration, which contrasts with the case (~26%) when C57BL/6 females were subjected to natural mating. Furthermore, 44% hormone-injected and plugged females delivered pups with an average litter size of six, which was comparable to the rate obtained from females that were not injected with hormones. These findings indicate that a single IP injection of low-dose PMSG increases the rate of acquiring plugged females before mating. This is particularly beneficial for *i*-GONAD which always requires desired number of plugged females obtained through a scheduled mating.

#### **7.10 Combinational use of** *i***-GONAD with anti-inhibin monoclonal antibodies (AIMAs) treatment to increase the number of GM mice**

Many attempts have been made to increase litter sizes (which is determined by the number of oocytes naturally ovulated) using conventional superovulation regimens (e.g., using PMSG/hCG), but had limited success because of unexpected decreases in the numbers of embryos surviving to term, as mentioned in the Section 7–9. Hasegawa et al. [115] attempted to overcome this problem using rat-derived AIMAs. They administrated progesterone (P4) once a day for 2 days (days 1 and 2) to synchronize the estrous cycle of female C57BL/6 mice, and AIMAs were injected into the same animals at Day 4 followed by mating with male C57BL/6 mice. When *i*-GONAD targeting *Tyr* was applied to the AIMA-treated C57BL/6 female mice on the day of vaginal plug formation during Days 6–8, a 1.5-fold increase in litter size was observed (7.3 *vs.* 4.8 for the untreated control). Notably, genome editing efficiency did not differ between these two groups. Therefore, AIMA treatment can reduce the number of females used for the *i*-GONAD experiment, which will fulfill the 3R principles of animal experimentation (i.e., Reduction, Replacement, and Refinement), and can be applied to other mouse strains and animals.

#### **7.11 GONAD/***i***-GONAD as a useful tool to check** *in vivo* **gene correction event**

As was shown previously, GONAD/*i*-GONAD enable gene delivery to early embryos present within the oviductal lumen and to the epithelial cells facing the lumen [37, 53, 139]. The success of gene delivery to the oviductal epithelial cells can be easily judged through direct observation of fluorescence under a fluorescence microscope [37, 53, 139]. Therefore, *in vivo* gene delivery approach targeting oviductal epithelial cells are excellent for testing the function of the GOI.

Miura et al. [109] attempted to examine whether *in vivo* CRISPR/Cas9-mediated gene correction is possible using GONAD/*i*-GONAD technologies. They first generated Δ*eGFP* KO mouse strain through MI of a solution containing *Cas9* mRNA and gRNA (targeted *eGFP* cDNA) into zygotes from Tg mice carrying *eGFP* cDNA [85]. The resulting Δ*eGFP* KO mice failed to exhibit systemic eGFP expression, due to frame-shift mutations in the coding region of *eGFP* cDNA that was chromosomally integrated in their genome. Next, they injected a solution containing sgRNA targeted to the mutation site, Cas9 protein, ssODN (as donor DNA sequence; in some cases), and Fast Green FCF into the oviductal lumen of female Δ*eGFP* KO mouse. Subsequently, the entire oviducts were electroporated, similar to that performed in

GONAD/*i*-GONAD. Three to 13 days later, the eGFP fluorescence was inspected in the oviducts dissected from the treated females. Consequently, fluorescence was detected in a portion of an oviduct, suggesting gene editing at the mutated site in the *eGFP* cDNA through HDR-mediated KI of ssODN or NHEJ-mediated indels. Molecular biological analysis of the oviduct confirmed the above events. Notably, in this system, editing of mutated site can be easily monitored by visually inspecting the gene-edited oviducts under UV illumination.

#### **7.12 GONAD/***i***-GONAD as a useful tool to generate mouse models for ovarian cancer**

As was mentioned previously, GONAD/*i*-GONAD could transfect both preimplantation embryos and the oviductal epithelium facing the oviductal lumen. Ovarian cancer is the most lethal gynecologic cancer to date. High-grade serous ovarian carcinoma (HGSOC) is the most common type of ovarian cancer and has the lowest rate of survival. Teng et al. [149] recreated the mutations found in ovarian cancer to generate somatic ovarian cancer mouse models, using an *in vivo* oviductal EP method similar to GONAD/*i*-GONAD. Using the CRISPR/Cas9 genome editing approach, they mutated the tumor suppressor genes (transformation related protein 53 (*Trp53*), breast cancer susceptibility gene I (*Brca1*), neurofibromin 1 (*Nf1*), and phosphatase and tensin homolog (*Pten*)) to study how these genes contribute to tumor development. When mutations were introduced in three of the four genes, namely *Trp53*, *Brca1*, and either *Nf1* or *Pten*, the sites transfected with the genome editing reagents displayed effects that were similar to human HGSOC and changes in chromosome number. Teng et al. [149] concluded that the *in vivo* oviductal EP method is highly useful for generating mouse models to advance the understanding and treatment of ovarian cancer.

#### **8. Challenges, limitations, concerns and future perspective**

#### **8.1 Challenges**

GONAD/*i*-GONAD is performed by injecting a small amount (1–2 μL) of a solution containing genome editing components into the specific site of an oviduct (called ampulla) of a pregnant female at Day 0.7 to 1.4 of pregnancy, using oral breath-controlled glass micropipette under a dissecting microscope. The genome editing components introduced within an oviductal lumen exist around ZP-enclosed early embryos (zygotes to 2-cell embryos), but are never incorporated into those embryos in an intact state. However, *in vivo* EP enables delivery of substances (present outside the embryos) into the internal portion of an embryo, leading to generation of genome-edited embryos. Another method to induce genome editing in early embryos *in situ* is viral transduction using rAAV. Unfortunately, EP requires expensive apparatus such as an electroporator, and rAAV transduction requires labor-intensive and time-consuming preparation of viral particles. In this context, the use of ZP-penetrating agents (e.g., MWNT and VisFect) would be ideal, because it does not require an electroporator or viral preparation. To date, there is no report on successful genome editing in early embryos using these ZP-penetrating agents. We expect that these agents could be useful for facilitating NA delivery to embryos and subsequent induction of genome editing.

Kaneko and Tanaka [100] demonstrated that pretreatment of early zygotes (tightly surrounded by cumulus cells) with HA led to increased efficiency of genome editing in those embryos after *i*-GONAD. This is based on the concept that cumulus cells surrounding a zygote hamper rapid transfer of genome editing reagents to zygotes [86]. In the previous approach using GONAD/*i*-GONAD, *in vivo* EP is applied immediately after intraoviductal injection of NAs [84, 86, 116]. In this case, it is highly likely that the reagents injected might not have been fully infiltrated between the intercellular space connecting cumulus cells. Waiting for several minutes after intraoviductal injection may permit sufficient infiltration of the reagents before *in vivo* EP, leading to increased efficiency of genome editing. This line of experiment has now been undertaken by Takabayashi and his colleagues.

#### **8.2 Limitations**

GONAD/*i*-GONAD can be applied to larger animals, as exemplified by its use in pigs; the demand for GM pigs has been rising due to the needs of the biomedical and agricultural fields [150, 151]. The current strategy for creating GM pigs is based on "*ex vivo* handling of embryos," where MI or EP is carried *in vitro* towards zygotes collected from individuals or produced through IVF, following which the treated embryos have to be subjected to ET to recipient females [138]. Similar to the case of MI or *in vitro* EP-mediated production of GM mice and rats, creation of GM pigs is highly costive, time-consuming, and labor-intensive. Additionally, it requires recipient female pigs, which are also expensive. In this context, GONAD/*i*-GONAD should be theoretically performed in more convenient and inexpensive manner, since it does not require "*ex vivo* handling of embryos." Notably, the porcine oviduct is generally ca. 100 mm in length and their form is linear, unlike the spiral form in rodents (mice and rats). Furthermore, it does not have an enlarged site called "ampulla" where rodent zygotes always stay. As a result, porcine zygotes will exist in a broad area throughout the oviduct. To perform GONAD/*i*-GONAD in pigs, researchers must inject a large amount of fluid (probably over 1 mL) and electroporate towards several sites whenever tweezer-type electrodes are used. It remains uncertain whether this is feasible. Therefore, GONAD/*i*-GONAD in pigs remains a challenge.

#### **8.3 Concerns**

As was mentioned in Section 1, in the initial step of development of GONAD/*i*-GONAD, gene delivery to early murine embryos using plasmid DNA were successful when the T-820 electroporator from BTX Co. was used [28]. However, it was impossible when the other electroporators such as NEPA21 and CUY21EDIT II were employed (Sato, Ohtsuka, unpublished). Notably, Hakim et al. [54] recently checked several *in vitro* EP parameters to seek optimal conditions enabling gene (plasmid DNA) delivery into mouse follicles, oocytes, and early embryos. When they were electroporated in the 1-mm gap cuvettes using Gene Pulser Mxcell System (Bio-rad Laboratories, Hercules, CA, USA), EP under 3 pulses of 30 V of 1 ms each at an interval of 10 s was ideal, with no need to weaken or loosen the ZP layer. This suggests that exploration of optimal EP condition using the above apparatuses (NEPA21 and CUY21EDIT II) may enable the transfection of ZP-intact embryos with plasmid DNA. If it is realized, transgenesis *via* introduction of plasmid-based transposons (e.g., *piggy*Bac (PB) transposon + PB transposase mRNA) may be possible.

#### **8.4 Future perspective**

Preimplantation embryos at zygote to morula stages, all of which exist within the oviductal lumen of a pregnant female, can be a target for gene delivery through GONAD/*i*-GONAD. As cleavage embryos (at 2-cell to early 8-cell stages) are comprised of blastomeres, each of which is facing the outside environment, genome editing at these stages may frequently result in the generation of mosaic offspring containing edited and unedited cells. This mosaic nature is tedious for the researchers who want to produce GM animals with high efficiency, but in turn beneficial for investigating the properties of embryonic lethal genes, because mosaic fetuses or pups produced through MI of genome editing components into one blastomere of two-cell embryos should be viable and carry heritable lethal mutations [152]. On the other hand, in the case of compacted 8-cell embryos and morulae, only blastomeres facing the external environment (but not the inner cells present inside an embryo) can be susceptible to genome editing. The outer blastomeres of morulae are thought to contribute to the formation of a trophectodermal cell, which is a cell involving implantation and placenta formation. Therefore, the GONAD/*i*-GONAD-mediated genome manipulation at these stages may be a novel tool to explore the molecular mechanisms underlying implantation and placenta formation.

#### **9. Conclusion**

Seven years have passed since the first report [84] on the development of GONAD using mice. During this period, successful genome editing was reported in other animals including rats and hamsters. The genes targeted by GONAD/*i*-GONAD technologies were *eGFP* cDNA (chromosomally integrated in the *eGFP* Tg mice) [84, 109] and endogenous genes such as *Acrosin*, *Agouti*, *Ak040954*, *ATF5*, *Axdnd 1*, *Cdkn1a*, *Cdkn2a*, *Clcf1*, *COL4A3*, *COL4A4*, *COL4A5*, *COL7A1*, *Dll1*, *eEF2*, *Fgf10*, *Fosl1*, *Foxe3*, *Gbx2*, *GGTA1*, *Gm30413*, *Gm44386*, *Hprt*, *Kit*, *Klrc2*, *Mecp2*, *Mov10l1*, *Nrsn2*, *Pafah1b1*, *Pax6*, *Pitx3*, *Plagl1*, *Serpina3n*, *Tis21*, *Tprkb*, and *Tyr* (**Table 1**). All these genes were disrupted by indel mutations (KO) or modified through HDRmediated KI using the CRISPR/Cas9 system. As for the components used for CRISPR/ Cas9-mediated genome editing, *Cas9* mRNA was initially employed, but later Cas9 protein was mainly used. For KI experiments, ssODN or *in vitro* synthesized long ssDNA was used as donor DNA. The dye used for monitoring successful injection process was also changed; while TB was initially employed, Fast Green FCF was used for later experiments. According to Ohtsuka et al. [86], TB (but not Fast Green FCF) often generates visible precipitates, when RNP is mixed with the dye. GONAD/*i*-GONAD technologies are very simple systems that only require the intraoviductal injection of NA-containing solution and subsequent *in vivo* EP. EP is now recognized as a powerful tool enabling efficient gene delivery, but often causes deleterious effects on cell/embryo survival, leading to reduction in litter size, as suggested by Kaneko and Tanaka [100]. Ohtsuka et al. [86] first demonstrated that the success of *in vivo* EP depends on the mouse strains used. This was also true for generating GM rats using *i*-GONAD [91]. Therefore, exploration of optimal EP conditions is required before GONAD/*i*-GONAD experiments start using new strains.

GONAD/*i*-GONAD requires an expensive electroporator. Notably, the intraoviductal injection of rAAV into a pregnant female mouse was also useful for the *in situ* transduction of zygotes [92, 139]. Furthermore, it has already been shown that

AAV-mediated genome editing at zygotes through AAV-based GONAD is powerful for the convenient acquisition of GM animals [92]. In this AAV-based GONAD system, the electroporator need not be used, although it is always associated with laborious and time-consuming tasks such as viral vector preparation. For developing systems that are simpler than the present GONAD/*i*-GONAD, the use of ZP-penetrating reagents will be highly desirable.

Recently, new genome editing systems known as prime editing [153–156] and base editing [157–159] were reported. These systems enable precise gene correction at a single nucleotide level. To date, these reagents have not been used for GONAD/*i*-GONAD-mediated production of GM animals. Future application of these new genome editing systems to GONAD/*i*-GONAD is highly expected.

#### **Acknowledgements**

We thank Kazusa Inada for her support for in-house drawing of the Figures, shown in **Figures 1–4**. This study was partly supported by a grant (no. 24580411 for Masahiro Sato; no. 21 K10165 for Emi Inada; no. 16H05049 for Shingo Nakamura; no. 22H03277 for Issei Saitoh; no. 16 K07087 for Shuji Takabayashi) from the Ministry of Education, Science, Sports, and Culture, Japan, and a fund for the Promotion of Joint International Research (Fostering Joint International Research) (no. 16KK0189 for Masato Ohtsuka) from JSPS, Japan.

#### **Conflict of interest**

The founding sponsors had no role in the design of the study, collection, analyses, or interpretation of data, writing of the manuscript, and decision to publish the results.

#### **Notes**

Masahiro Sato and Shuji Takabayashi designed the study and drafted the manuscript; Masato Ohtsuka, Emi Inada, Shingo Nakamura and Issei Saitoh critically revised the manuscript.

### **Author details**

Masahiro Sato1 \*, Masato Ohtsuka2,3,4, Emi Inada5 , Shingo Nakamura6 , Issei Saitoh7 and Shuji Takabayashi8

1 Department of Genome Medicine, National Center for Child Health and Development, Tokyo, Japan

2 Department of Molecular Life Science, Division of Basic Medical Science and Molecular Medicine, Tokai University School of Medicine, Isehara, Kanagawa, Japan

3 Center for Matrix Biology and Medicine, Graduate School of Medicine, Tokai University, Isehara, Kanagawa, Japan

4 The Institute of Medical Sciences, Tokai University, Isehara, Kanagawa, Japan

5 Department of Pediatric Dentistry, Graduate School of Medical and Dental Sciences, Kagoshima University, Kagoshima, Japan

6 Division of Biomedical Engineering, National Defense Medical College Research Institute, Saitama, Japan

7 Department of Pediatric Dentistry, Asahi University School of Dentistry, Mizuho, Japan

8 Laboratory Animal Facilities and Services, Preeminent Medical Photonics Education and Research Center, Hamamatsu University School of Medicine, Hamamatsu, Shizuoka, Japan

\*Address all correspondence to: sato-masa@ncchd.go.jp

© 2022 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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