Section 2 Animal Research

**17**

this species.

**Chapter 2**

**Abstract**

Research

the future prospects.

**1. Introduction**

**technology**

Application and Development of

CRISPR/Cas9 Technology in Pig

Pigs provide valuable meat sources, disease models, and research materials for humans. However, traditional methods no longer meet the developing needs of pig production. More recently, advanced biotechnologies such as SCNT and genome editing are enabling researchers to manipulate genomic DNA molecules. Such methods have greatly promoted the advancement of pig research. Three gene editing platforms including ZFNs, TALENs, and CRISPR/Cas are becoming increasingly prevalent in life science research, with CRISPR/Cas9 now being the most widely used. CRISPR/Cas9, a part of the defense mechanism against viral infection, was discovered in prokaryotes and has now developed as a powerful and effective genome editing tool that can introduce and enhance modifications to the eukaryotic genomes in a range of animals including insects, amphibians, fish, and mammals in a predictable manner. Given its excellent characteristics that are superior to other tailored endonucleases systems, CRISPR/Cas9 is suitable for conducting pig-related studies. In this review, we briefly discuss the historical perspectives of CRISPR/Cas9 technology and highlight the applications and developments for using CRISPR/Cas9-based methods in pig research. We will also review the choices for delivering genome editing elements and the merits and drawbacks of utilizing the CRISPR/Cas9 technology for pig research, as well as

**Keywords:** applications, CRISPR/Cas9, delivery methods, gene editing, pig

**1.1 The status of pig production and current application of CRISPR/Cas9** 

Worldwide, pig (*Sus scrofa domestica*) production accounted for 42% of total livestock production in 2018, and this percentage is expected to go up by the year 2050 [1, 2]. Pork, which makes up nearly 40% of all meat consumed by the world population, is clearly an important meat source for humans [3]. These production and consumption data reveal the significant implications of pigs for humans. Indeed, pigs bring many benefits for the convenience and survival of human beings. In light of the importance and necessity for pig production, researchers all around the world are using various methods to actively investigate

*Huafeng Lin, Qiudi Deng, Lili Li and Lei Shi*

### **Chapter 2**

## Application and Development of CRISPR/Cas9 Technology in Pig Research

*Huafeng Lin, Qiudi Deng, Lili Li and Lei Shi* 

### **Abstract**

 Pigs provide valuable meat sources, disease models, and research materials for humans. However, traditional methods no longer meet the developing needs of pig production. More recently, advanced biotechnologies such as SCNT and genome editing are enabling researchers to manipulate genomic DNA molecules. Such methods have greatly promoted the advancement of pig research. Three gene editing platforms including ZFNs, TALENs, and CRISPR/Cas are becoming increasingly prevalent in life science research, with CRISPR/Cas9 now being the most widely used. CRISPR/Cas9, a part of the defense mechanism against viral infection, was discovered in prokaryotes and has now developed as a powerful and effective genome editing tool that can introduce and enhance modifications to the eukaryotic genomes in a range of animals including insects, amphibians, fish, and mammals in a predictable manner. Given its excellent characteristics that are superior to other tailored endonucleases systems, CRISPR/Cas9 is suitable for conducting pig-related studies. In this review, we briefly discuss the historical perspectives of CRISPR/Cas9 technology and highlight the applications and developments for using CRISPR/Cas9-based methods in pig research. We will also review the choices for delivering genome editing elements and the merits and drawbacks of utilizing the CRISPR/Cas9 technology for pig research, as well as the future prospects.

**Keywords:** applications, CRISPR/Cas9, delivery methods, gene editing, pig

### **1. Introduction**

### **1.1 The status of pig production and current application of CRISPR/Cas9 technology**

Worldwide, pig (*Sus scrofa domestica*) production accounted for 42% of total livestock production in 2018, and this percentage is expected to go up by the year 2050 [1, 2]. Pork, which makes up nearly 40% of all meat consumed by the world population, is clearly an important meat source for humans [3]. These production and consumption data reveal the significant implications of pigs for humans. Indeed, pigs bring many benefits for the convenience and survival of human beings. In light of the importance and necessity for pig production, researchers all around the world are using various methods to actively investigate this species.

 Benefitting from the rapid development of genome-editing technologies during the last decade, many laboratories have applied this tool to animals, plants, and microorganisms in order to obtain both higher yield and better quality varieties. With the advent of the CRISPR (clustered regularly interspaced short palindromic repeats)/Cas9 technique and the melioration of delivery methods, gene editing can be more successfully performed in livestock such as swine. In addition, evidence shows that, in addition to primates, pigs share many similar characteristics with humans such as organ size, genome length, blood glucose levels, and the complexity and composition of chromosomes [4, 5], as well as the early embryonic development trajectory [6]. Therefore, pigs are not only used as important domestic animals for food and pharmaceutical applications, but also served as ideal animal models for simulating various human diseases (e.g., diabetes, obesity, and cardiovascular disease). In this manuscript, we first introduce the historical perspectives of gene-editing technologies in pigs, review the latest advances in the utilization of CRISPR/Cas9 strategies for swine research, and then describe possible methods for delivering these genome-editing components, as well as the future perspective on pig studies by using this technology.

### **1.2 Historical background of gene editing in pigs**

 CRISPR, discovered in 1987, is a family of DNA sequences of short direct repeats interspaced with short sequences. Its mechanism of action has been confirmed to be related with acquired immunity of microbes [7–9]. By 2000, researchers had discovered that these specific sequences occurred in about 40% of bacteria and 90% of archaea [10, 11]. In 2002, this interesting architecture, initially named short regularly spaced repeats (SRSRs), was renamed as the clustered regularly interspaced short palindromic repeats (CRISPRs) [10, 12]. Between 2002 and 2009, a series of proteins associated with these palindromic sequences were identified as constituents of the complicated mechanism of microbial adaptive immunity [11]. In 2014, the X-ray crystal structure of *Streptococcus pyogenes* Cas9 (*Sp*Cas9) in complex with sgRNA was elucidated [13, 14]. Nowadays, *Sp*Cas9 endonuclease, which requires a protospacer adjacent motif (PAM) sequence (5'-NGG-3′), is routinely designed as a 'molecular scissor' guided by a single guide RNA (or dual-tracrRNA) due to simple structural characteristics, the advantages of easy operation, and high efficiency [11, 15]. Notably, the multiplex abilities of the Cas9-associated guided RNAs (gRNAs) and the diverse Cas9 orthologs (e.g*.*, *Sp*Cas9, *Sa*Cas9, *St*Cas9) as well as the diversified Cas9 variants (**Figure 1**) have enabled CRISPR/Cas9 systems to be used in a wide range of research applications [16, 17].

As early as 1985, the first transgenic pig was created by direct DNA microinjection of the metallothionein-I/human growth hormone (MT/hGH) fusion gene into a fertilized egg [18]. Further technical enhancements occurred during the next 20 years, until, in 2011, Whitworth and his co-workers were the first to successfully apply ZFN technology to generate cloned eGFP knockout pigs [19]. Similarly, Carlson et al. (2012) pioneered the application of TALENs in editing the porcine genome, and they produced low-density lipoprotein receptor (LDLR) knockout pigs [20]. By 2013, the groundbreaking work of genome engineering in mammalian cells based on the CRISPR/Cas9 system had been achieved [21]. The first examples of genome-modified pigs engineered using the CRISPR/Cas9 technique were reported almost simultaneously by Hai et al. (2014) [22] and Whitworth et al. (2014) [23]. From then on, rapid and efficient CRISPR/Cas9-mediated genome editing in pigs has opened up many more possibilities for applications in biology and biomedicine.

*Application and Development of CRISPR/Cas9 Technology in Pig Research DOI: http://dx.doi.org/10.5772/intechopen.85540* 

### **Figure 1.**

*Diagram illustrating different types of engineered CRISPR/Cas9 and its Cas9 variants. (A) The wild-type SpCas9 nuclease. (B) The wild-type SaCas9 nuclease. (C) The wild-type NmCas9 nuclease. (D) The wildtype StCas9 nuclease. (E) The dCas9 variant can bind DNA but cannot cut DNA strands. (F) The SpCas9 nickase that can only introduce a single strand break at the HNH nuclease domain. (G) The SpCas9 nickase that can only introduce a single strand break at the RuvC nuclease domain. (SpCas9, Streptococcus pyogenes Cas9; SaCas9, Staphylococcus aureus Cas9; NmCas9, Neisseria meningitides Cas9; StCas9, Streptococcus thermophilus Cas9; dCas9, catalytically inactive ("dead") Cas9; sgRNA, single-guide RNA; PAM, protospacer adjacent motif; W = A or T). Refer to [16].* 

### **2. Application and development**

### **2.1 Applications in the antimicrobial and antiviral fields**

Currently, the traditional methods for developing pig anti-viral vaccines are time-consuming and labor-intensive [24]. Cas9 endonucleases, as molecular DNA scissors guided by gRNA, are now used to target and cut exogenous DNA arising from virus or plasmids [25]. With the development of state-of-the-art biotechnologies, scientists now can utilize this revolutional tool to prevent domestic pigs from pathogenic bacterial and viral attack. In 2016, Liang and his colleagues developed a rapid vaccine development method based on the combination of CRISPR/Cas9 and the Cre/Lox system to fight against the re-emerging pseudorabies virus (PRV). The results demonstrated the protective efficacy of this candidate vaccine in swine and showed promise in controlling the outbreak of pseudorabies [26]. In another trial, Whitworth et al. (2015) employed the CRISPR/Cas9 system to directionally mutate the CD163 gene (cluster of differentiation 163 gene, a gate keeper gene associated with PRRSV) in order to create biallelic gene knockout pigs which had protective immunity against infection of porcine reproductive and respiratory syndrome virus (PRRSV) [27]. In 2018, Xie and his co-workers applied the combinational method of CRISPR/Cas9 and RNAi to generate anti-CSFV transgenic pigs and confirmed that these pigs could impede the multiplication of classical swine fever virus (CSFV). They further proved that the disease resistance traits presented in the

 transgenic sows could be stably transmitted to their F1-generation offspring. This study suggested that the use of such transgenic pigs would offer potential benefits over commercial vaccination, could substantially reduce CSFV-related economic losses, and would also improve the well-being of livestock [28]. Compared to CSFV, African swine fever virus (ASFV) is a very acute, lethal viral pathogen for both domestic and wild pigs, but unfortunately, a vaccine candidate that effectively prevents ASFV infection remains elusive. HüBner et al. (2018) applied the CRISPR/ Cas9 nuclease system to target the double-stranded DNA genome of ASFV. In vitro culture experiments showed that mediated targeting of the ASFV p30 gene using this system is a feasible strategy to fight against ASFV infection, and may also be applied to the natural animal host [29].

### **2.2 Applications to breeding and reproduction**

 Traditional breeding methods, which comprise selective breeding and crossbreeding, have clearly hit a bottleneck. Additionally, due to the long time, high cost, and high labor intensity of traditional breeding methods [30], researchers now hope to find other alternatives that are more convenient and efficient than previously. Genome-editing technology can help us to achieve a good result in a short time, and help better understand swine reproduction. Interestingly, many aspects of pig reproduction are suitable as translational models of reproduction in humans, including oocyte maturation, sperm-egg interaction mechanism, tubo-uterine contractility, early embryo development, pregnancy, fetal genome modification, and reproductive diseases [31]. Strategies that use the CRISPR/Cas9 technique to improve the reproduction in swine are becoming more prevalent. PRRSV, a virus associated with reproductive and respiratory disease, can cause severe unsuccessful reproductive outcomes in sows, decrease sperm quality in infected boars, and lower the birth rates of healthy piglets [32]. In 2016, Tao et al. generated efficient biallelic mutation in porcine parthenotes by cytoplasmic injection of Cas9/sgRNA mixtures. These data emphasize the function of parthenotes in revealing early embryonic development and assessing mutation efficiency [33]. In the same year, Whitworth et al. used CRISPR/Cas9 to generate CD163-knockout pigs to protect pig from PRRSV and reduce the incidence of reproductive disease, important for pig studies in both the fields of reproduction and anti-viruses [27]. In 2017, Park et al. utilized CRISPR/Cas9 technology to program the NANOS2 gene in domestic pigs to generate offspring with monoallelic and biallelic mutations. They found that NANOS2 knockout pigs presented the phenotype of male specific germ line ablation but other aspects of testicular development were normal. The exception was male pigs with one intact NANOS2 allele and female knockout pigs which both maintained good reproductive performance [34].

### **2.3 Applications in immunization and xenotransplantation**

Swines, having many highly similar anatomical and physiological features to humans, are considered to be the excellent donors for patients in the case of a shortage of human organs for allogenic transplantation [35, 36]. However, several issues still need to be addressed such as hyperacute rejection which can develop in recipients within several minutes after organ xenotransplantations [36, 37]. The advancement of the CRISPR/Cas9 technique has greatly strengthened the ability to effectively manipulate porcine genome in order to evaluate and generate porcine organs that can assist in xenotransplantation.

An early study, undertaken by Sato and his research team in 2013, used a modified CRISPR/Cas9 system to knockout the porcine GGTA1 gene, whose protein

### *Application and Development of CRISPR/Cas9 Technology in Pig Research DOI: http://dx.doi.org/10.5772/intechopen.85540*

product is responsible for the biosynthesis of the a-Gal epitope, which leads to hyperacute rejection upon pig-to-human xenotransplantation. This trial not only demonstrated that CRISPR/Cas9 is a promising tool for producing knockout cloned piglets, but also paved the way for pig-to-human xenotransplantation [38]. Piglets with biallelic knockouts of GGTA1 gene were eventually created by Petersen and his colleagues [39] using the combined technologies of CRISPR/Cas9 and somatic cell nuclear transfer (SCNT).

Swine could also serve as an ideal animal model for investigating viral immunity and immune rejection in xenotransplantation if they are deficient in class I MHC. Research published by Reyes et al. in 2014 utilized the Cas9 endonuclease with chimeric gRNAs to generate class I MHC knockout piglets as promising experimental animals for immunological research [40]. In 2015, Yang and coworkers designed two Cas9 gRNA molecules to inactivate 62 copies of the pol gene required for porcine endogenous retrovirus (PERV) activity. This study performed on porcine kidney epithelial cell lines demonstrated that the modifications could greatly reduce in vitro spreading of PERVs to human cells, raising the hope of the eradication of such viruses from pigs for heterograft donors [41]. One year later, Yang's research team (2017) made further progress in employing CRISPR/Cas9 technology to inactivate all the PERVs in a porcine primary cell line and produced PERV-eliminated pigs using the SCNT technique. The experimental results addressed the safety problem in clinical xenotransplantation due to the success of impeding interspecific transmission of viruses [42].

### **2.4 Disease models and translational medical research**

The CRISPR/Cas9 technology has both simplified and expedited biomedical modeling for some refractory human diseases. One way to combat human diseases is to create genetically modified animal models for investigating the mechanism of diseases enabling the development of safe and effective drugs. An effective animal disease model should appropriately simulate the *in vivo* environment under investigation and respond or react to stimuli in a similar manner to the human body [43–45]. Commonly used animal models in the laboratory include mice, rats, dogs, monkey, and swine. The pig models have been developed to faithfully mimic various human diseases including neurodegenerative diseases [46], cancers [45], and gastrointestinal (GI) tract diseases [47] as they share similar features to humans in terms of anatomy, physiology, and genetics [43]. Gene editing using CRISPR/Cas9 technology is proving an innovative and effective research tool, which is greatly revolutionizing our ability to manipulate the porcine genome to create appropriate disease models.

 As early as 2013, Tan et al. used two custom endonucleases (TALEN and CRISPR/Cas9 system) to edit azoospermia-like (DAZL) and adenomatous polyposis coli (APC) loci in the pig genome. The results suggested that gene editing could be incorporated into selection programs to accelerate genetic improvement, with applications in animal breeding and human personalized medicine [48]. In 2014, Zhou et al. were the first to report that zygote injection of a customized CRISPR/ Cas9 system could efficiently generate genome-modified pigs (biallelic knockout pigs) in one step, which provided an important animal model for the treatment of human type I and III *von* Willebrand disease [22]. At the end of 2015, Peng et al. adopted the CRISPR/Cas9 method to knockin human cDNA into the albumin gene locus in pig zygotes and successfully produced human albumin from porcine blood [49]. Additionally, Feng et al. (2015) reported the potential of using the combination of CRISPR/Cas9 and human pluripotent stem cells (PSCs) to harvest human organs from chimeric swine [50]. In 2016, Wang et al. performed a study in

which Cas9 mRNA and multiple single guide RNAs (sgRNAs), which respectively specifically target to parkin, DJ-1, and PINK1 gene loci, were coinjected into in vivo derived pronuclear embryos of Bama miniature pigs. There were only minor low off-target events. These results demonstrated the capability of using the CRISPR/ Cas9 system to trigger genetic modification of multiple sites in pigs, yielding positive results with high medical value [51]. In the same year, Lee and his team utilized genome-specific CRISPR/Cas9 systems to target runt-related transcription factor 3 (RUNX3, a known tumor suppressor gene) to generate a pig model that can recapitulate the pathogenesis of RUNX3-associated stomach cancer in humans. The results demonstrated that the CRISPR/Cas9 system was effective in inducing mutations on a specific locus of the pig genome, resulting in the generation of piglets lacking RUNX3 protein in their internal organs. This system brings useful resources (RUNX3 knockout pigs) for human cancer research and the development of novel cancer therapies [52]. In 2017, Zhang et al. designed an experiment that applied the CRISPR/Cas9 system and SCNT technology to generate complement protein C3 targeted piglets, which could be a valuable large animal model for elucidating the roles of C3, a protein of the immune system that plays a central role in the complement system and contributes to innate immunity [53]. By 2018, following many years' efforts, scientists have now made significant progress in using CRISPR/Cas9 mediated knockin techniques to produce a Huntington's disease (HD) pig model, which assists in the investigation of the pathogenesis of neurodegenerative diseases and the development of appropriate therapeutics [54]. Recently (2018), Cho and co-workers successfully used the CRISPR/Cas9 and SCNT technologies to generate INS knockout pigs (insulin-deficient pigs) and demonstrated the efficacy of the CRISPR/Cas9 system in producing pig models for use in diabetes research and pharmaceutical testing [55].

### **2.5 Improvement of meat quality and food safety**

 Pig meat quality is controlled by multiple factors. To some extent, genetics are considered as the dominating factor influencing pork quality in the pig industry, although environmental conditions can also potentially influence the porcine genetics in the long term. In addition, fat and lean meat contents are both important for the palatability of the pork [56, 57] and diet considerations. Consequently, scientists now propose to improve pork traits to cater for the taste of the general public by using gene-editing technology. In 2016, Bi et al. constructed isozygous, functional myostatin (MSTN) knockout cloned pigs without selectable marker gene (SMG) by combined use of CRISPR/Cas9 and Cre/LoxP. The results showed that compared to the control group, the skeleton muscles were more pronounced and the back fat thickness decreased slightly in such gene-edited pigs [58]. In 2017, Zheng et al. established a CRISPR/Cas9-mediated homologous recombination-independent approach to efficiently insert mouse adiponectin-UCP1 into the porcine endogenous uncoupling protein 1 (UCP1) locus. The resultant UCP1 knockin pigs showed an enhanced ability to control their body temperature during acute cold exposure, lower fat deposition, and increased carcass lean meat [59]. In 2018, Xiang et al. used CRISPR/Cas9 technology to effectively edit insulin-like growth factor 2 (IGF2) intron 3–3072 site as the method of choice for the improvement of meat production in Bama pigs. The result showed that it was the first time to demonstrate that editing a noncoding region can ameliorate economic traits in livestock [60].

CRISPR/Cas9 gene-editing technology has multiple benefits. In gene detection fields, Zhou et al. developed a CRISPR/Cas9-triggered nicking endonucleasemediated strand displacement amplification method (namely CRISDA) for amplifying and detecting double-stranded DNA [61]. CRISDA promises to be a


### *Application and Development of CRISPR/Cas9 Technology in Pig Research DOI: http://dx.doi.org/10.5772/intechopen.85540*



**Table 1.** *Examples for the applications of CRISPR/Cas9 technology in pigs.* 

*Application and Development of CRISPR/Cas9 Technology in Pig Research DOI: http://dx.doi.org/10.5772/intechopen.85540* 

powerful isothermal tool for ultrasensitive and specific detection of nucleic acids in pig pathogeny detection and food safety. Consequently, by making good use of this precision editing engineered technology in agriculture, a reliable avenue for elite swine production could be guaranteed, potential biological risks can be minimized, and a higher food safety can be protected.

### **2.6 Applications in transgenesis and beyond**

 Pig transgenesis is an important facet for functional investigation of biological pathways, as well as for biotechnology in animal husbandry. As a promising tool, CRISPR/Cas9 now has the ability to accelerate the process of pig transgenesis. Several studies have successfully constructed a CRISPR/Cas9 system for targeting the pig GGTA1 gene [38, 39, 62]. Ruan et al. (2015) inserted a gene fragment larger than 9 kb at the newly named pH 11 genomic locus using CRISPR/Cas9 technology and then confirmed that it was highly expressed in cells, embryos, and animals [63]. Similarly, Zhou et al. (2015) worked on CRISPR/Cas9-mediated gene targeting in porcine fetal fibroblasts (PFFs), in which TYR, PARK2, and PINK1 loci were effectively edited [64]. In 2016, Yang and colleagues edited the porcine INS (pINS) gene in fibroblasts by using TALENs or CRISPR/Cas9 [65], and in 2017, Zheng et al. inserted a mouse adiponectin-UCP1 gene efficiently into the porcine endogenous UCP1 locus by the utilization of a CRISPR/Cas9-mediated homologous recombination-independent approach [59]. In the same year, Wang et al. applied the combined system of Cre recombinase and Cas9/sgRNAs to simultaneously inactivate five tumor suppressor genes (TP53, PTEN, APC, BRCA1, and BRCA2) and activate one oncogene (KRAS) to develop a rapid lung tumor model in pigs [66]. By 2018, Whitworth et al. had developed a method that utilized the CRISPR/Cas9 technology to remove a loxP flanked neomycin cassette by direct zygote injection of RNA encoding Cre recombinase. This new technique can be used to efficiently remove selectable markers in genetically engineered animals without the need for long-term cell culture and subsequent somatic cell nuclear transfer (SCNT) [67]. Almost certainly, it has a very promising future for transgenic pigs with the advantages of enhancing body growth and minimizing environmental pollution that would be created by the CRISRP/Cas9 method. **Table 1** shows applications of CRISRP/Cas9 technology in transgenic pigs.

### **3. Delivery methods of CRISPR/Cas9**

### **3.1 The appropriate choices for delivery: viral systems or nonviral platforms?**

In order to introduce precise and efficient genome modification, the proper delivery modalities of CRISPR/Cas9 genome-editing materials are a crucial factor in the generation of genetically engineered pigs. A variety of strategies have been used for delivering the CRISPR/Cas9 system which can be mainly divided into viral and nonviral delivery methods (**Figure 2**) [82].

Viral systems are the traditional tools that have been widely used for delivering genome editing materials (DNA or mRNA). To-date, three viral vectors including lentivirus [83], adenovirus, and adeno-associated virus (AAV) have been used for delivery of CRISPR/Cas9 components in various biological studies [84, 85]. However, there are several limitations associated with viral vectors including immunogenicity, packaging capacity, broad tropism, and difficulty in production.

*Application and Development of CRISPR/Cas9 Technology in Pig Research DOI: http://dx.doi.org/10.5772/intechopen.85540* 

### **Figure 2.**

*Delivery techniques for the CRISPR/Cas9 system. (iTOP: induced transduction by osmocytosis and propanebetaine; AAV: adeno-associated virus).* 

Nonviral platforms for transferring the CRISPR/Cas9 components can be achieved by physical and chemical approaches. In contrast to viral vectors, nonviral vectors have lower immunogenicity, are not constrained by packaging sizes, are facile to synthesize, and are capable of carrying multiple sgRNAs simultaneously [86, 87]. In nonviral methods, genome editing reagents are delivered either as mRNA or as a combination of Cas9 endonuclease and sgRNA. To date, nonviral methods available include microinjection, electroporation [88], hydrodynamic injection, lipid particles, nanoclews, zwitterionic amino lipid (ZAL) nanoparticles, and iTOP as well as the combinations of viral and nonviral methods [82]. Herein, we compared the various methods for delivering the CRISRP/Cas9 system (**Table 2**).

 Delivery methods of gene modification in the field of pig research have even used sperms as vectors for foreign genes (*e.g.* sperm-mediated gene transfer (SMGT), and intracytoplasmic sperm injection (ICSI)-mediated gene transfer), and delivery strategies such as retroviruses and lentiviruses are still current [100]. Somatic cell nuclear transfer (SCNT), a technique that consists of taking an enucleated oocyte and then implanting a donor nucleus from a somatic cell, is a remarkable breakthrough in the history of swine genetic engineering [101, 102]. SCNT combined with the rapid development of gene editing technologies such as TALENs and CRISPR/Cas9 has excellent prospects.

### **3.2 Challenges for delivering the CRISPR/Cas9 systems**

 The CRISPR/Cas9 system has been applied to genome modification in a variety of microorganisms, plants, and animals (including pigs), but the efficient transfer of such system is still a challenge that affects the precise genome-editing activity [103]. If the CRISPR/Cas9 systems are to effectively function in the targeted cells or organisms, choosing a suitable delivery system is of critical importance. According to existing research, the CRISPR/Cas9 system can be broadly divided into three kinds of packaging formats: Cas9 protein and sgRNA, Cas9 mRNA and


### **Table 2.**

*Comparison of different delivery methods for CRISPR/Cas9 system.* 

sgRNA, and CRISPR/Cas9 plasmid. Different CRISPR/Cas9 formats cooperate with special transport vehicle to complete the transportation task for gene-editing elements. Some research studies indicate that CRISPR/Cas9 ribonucleoprotein (RNP) delivery seems to exceed gene delivery as it provides multiple function advantages: short-term delivery, no insertional mutagenesis, minimal immunogenicity, and low off-target effect [87]. As previously mentioned, viral vectors usually have their own limitations to be overcome compared to nonviral vectors. However, nonviral vectors are generally used for *in vitro* genome editing studies due to their biological incompatibility or cytotoxicity [95]. Recently, developing efficient and biocompatible nonviral vectors (e.g., liposome and nanocarrier) has just emerged, and achievements have been made. For example, a low cytotoxic cationic polymer has been proven to mediate efficient CRISPR/Cas9 plasmid delivery for genome editing [92]. In addition, a research article presented that lipid-based Cas9 mRNA delivery has lower off-target effects than lentiviruspackaged Cas9 mRNA transportation [104]. Generally speaking, the packaging modes and delivery tools are two biggest factors that affect efficiency of the CRISPR/Cas9 system apart from this system itself. In order to describe the possible challenges for delivering the CRISPR/Cas9 system and the strategies used to overcome these challenges, we form a table to illustrate in detail (**Table 3**) and further to promote much research applications appropriately.


*Application and Development of CRISPR/Cas9 Technology in Pig Research DOI: http://dx.doi.org/10.5772/intechopen.85540* 

### **Table 3.**

*Challenges for delivering the CRISPR/Cas9 system and the strategies that respond to these challenges.* 

### **4. Discussion**

CRISPR/Cas9 technology is not only simple and easy to perform, but also has significantly improved performances for mutational studies, which has accelerated the application of the CRISPR/Cas9 toolkit [68, 111]. However, there are still some limitations and difficulties in the use of the CRISPR/Cas9 system for pig research.


### **5. Conclusion**

Over the past few years, genome-editing technology clearly allows scientists to produce genetically engineered pigs that are healthier to consume and more resistant to diseases in an efficient way. Nowadays, the use of the CRISPR/Ca9 technique on pigs in immunity, autoimmunity, obesity, aging, etc. is increasingly expanding and showing advantages over the conventional methods. In addition, another version of CRISPR named CRISPR/Cpf1 was discovered in microbes, which further expanded the CISPR toolkit, and holds promise to be applied in pig research. CRISPR/Ca9-modified pigs are providing a better perspective for understanding various aspects of pig biology and are paving the way for advancing the fields of basic biology, translational medicine, biomedicine, and drug development.

### **Acknowledgements**

We would like to thank Professor XF Qi (Jinan University, China), Professor Edouard C. Nice (Monash University, Australia), and Professor Mark Baker (Macquarie University, Australia) for providing assistance and guidance in this project. We specially acknowledge Professor Edouard C. Nice for revising this article.

### **Conflict of interest**

None declared.

### **Fundings**

 This work was collectively supported by grants from the Guangdong Innovative and Entrepreneurial Research Team Program (2014ZT05S136), National Key Research and Development Plan (2016YFD0500600), Guangdong Provincial Science and Technology Plan Project (2017B020207004), and Fundamental Research Funds for the Central Universities (21618309).

### **Acronyms and abbreviations**


*Application and Development of CRISPR/Cas9 Technology in Pig Research DOI: http://dx.doi.org/10.5772/intechopen.85540* 


### **Author details**

Huafeng Lin1,2\* † , Qiudi Deng1†, Lili Li2 and Lei Shi2

1 Department of Biotechnology, College of Life Science and Technology, Jinan University, Guangzhou, Guangdong, PR China

2 Institute of Food Safety and Nutrition, Jinan University, Guangzhou, PR China

\*Address all correspondence to: 88983088@qq.com

† These authors contributed equally to this work and should be considered co-first authors

© 2019 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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**41**

**Chapter 3**

**Abstract**

sperm based on recent findings.

intratesticular injection

**1. Introduction**

Possible Production of

*Masahiro Sato and Shingo Nakamura*

Gene-Engineered Sperm

Genome-Edited Animals Using

CRISPR/Cas9 is widely used for genome editing in a variety of organisms, including mammals, fishes, and plants. In mammals, zygotes are considered an appropriate target for gene delivery of CRISPR/Cas9 components [Cas9 endonuclease and a single-guide (sgRNA)] via microinjection or *in vitro* electroporation. However, these approaches require *ex vivo* handling of zygotes, which is necessary for egg transfer to recipient females to allow the treated zygotes to develop fullterm. These procedures are often laborious, time-consuming, and use numerous mice. In our previous experiments, the plasmid DNA encapsulated by liposomal reagent introduced into the internal portion of a testis can be transferred to the mature sperm present in the epididymal ducts, and is finally transferred to oocytes via fertilization. Although it was not integrated into their genome, this approach would be useful for creating genome-edited animals, since CRISPR/Cas9 can be performed by transient interaction of Cas9 and sgRNA, whereby chromosomal integration of the CRISPR components is not a prerequisite. Here, we will review past achievements concerning in vivo transfection of immature/mature sperm and present experimental proposals for possible genome editing via gene-engineered

**Keywords:** sperm, CRISPR/Cas9, guide RNA, testis-mediated gene transfer,

*in vivo* transfection, genome editing, vas deferens, epididymis, artificial insemination,

Transgenesis is a method to induce genetic change in an organism by delivering exogenous DNA (also called transgenes) to early embryos (i.e., zygotes), and is now considered an important technique to examine gene function *in vivo* and for creating animal models of human disease [1, 2]. In 1980, Gordon et al. [3] first demonstrated that microinjection of purified DNA fragments into the pronuclei of zygotes led to the production of mice carrying the transgenes, which are generally referred to as transgenic (Tg) or genetically modified (GM) mice. When the injected transgenes are successfully integrated into the host chromosomes of the zygotes, they are transmitted to the next generation through mating with normal mice in a Mendelian ratio, and gene expression derived from the integrated transgenes will occur in the Tg offspring depending on the property of the promoter used. For the production of Tg animals through zygote microinjection, several steps

### **Chapter 3**

## Possible Production of Genome-Edited Animals Using Gene-Engineered Sperm

*Masahiro Sato and Shingo Nakamura* 

### **Abstract**

 CRISPR/Cas9 is widely used for genome editing in a variety of organisms, including mammals, fishes, and plants. In mammals, zygotes are considered an appropriate target for gene delivery of CRISPR/Cas9 components [Cas9 endonuclease and a single-guide (sgRNA)] via microinjection or *in vitro* electroporation. However, these approaches require *ex vivo* handling of zygotes, which is necessary for egg transfer to recipient females to allow the treated zygotes to develop fullterm. These procedures are often laborious, time-consuming, and use numerous mice. In our previous experiments, the plasmid DNA encapsulated by liposomal reagent introduced into the internal portion of a testis can be transferred to the mature sperm present in the epididymal ducts, and is finally transferred to oocytes via fertilization. Although it was not integrated into their genome, this approach would be useful for creating genome-edited animals, since CRISPR/Cas9 can be performed by transient interaction of Cas9 and sgRNA, whereby chromosomal integration of the CRISPR components is not a prerequisite. Here, we will review past achievements concerning in vivo transfection of immature/mature sperm and present experimental proposals for possible genome editing via gene-engineered sperm based on recent findings.

**Keywords:** sperm, CRISPR/Cas9, guide RNA, testis-mediated gene transfer, *in vivo* transfection, genome editing, vas deferens, epididymis, artificial insemination, intratesticular injection

### **1. Introduction**

Transgenesis is a method to induce genetic change in an organism by delivering exogenous DNA (also called transgenes) to early embryos (i.e., zygotes), and is now considered an important technique to examine gene function *in vivo* and for creating animal models of human disease [1, 2]. In 1980, Gordon et al. [3] first demonstrated that microinjection of purified DNA fragments into the pronuclei of zygotes led to the production of mice carrying the transgenes, which are generally referred to as transgenic (Tg) or genetically modified (GM) mice. When the injected transgenes are successfully integrated into the host chromosomes of the zygotes, they are transmitted to the next generation through mating with normal mice in a Mendelian ratio, and gene expression derived from the integrated transgenes will occur in the Tg offspring depending on the property of the promoter used. For the production of Tg animals through zygote microinjection, several steps are required for the "*ex vivo* handling of embryos," including: collection of zygotes, DNA microinjection using an expensive micromanipulator, temporal incubation of the injected zygotes, and egg transfer (ET) to the oviducts of the pseudo-pregnant females to allow full-term development of the injected eggs [4, 5]. Furthermore, all of this requires highly specialized and skilled personnel for the preparation of pseudo-pregnant females and vasectomized males, which is time-consuming and tedious, and requires a large number of mice.

 Since 1980, several methods for bypassing microinjection-based transgenesis have been provided, which include infection of zygotes with viral vectors like a retrovirus [6, 7], embryonic stem (ES) cell-based gene transfer [8–10], transgenesis via somatic cell nuclear transfer [11–13], and intracytoplasmic sperm injection (ICSI) using sperm associated with the transgene (TransICSI) [14, 15]. All of these methods deal with zygotes and require *ex vivo* handling of embryos, although a micromanipulator system is not used in almost the cases.

 Genome-editing via Oviductal Nucleic Acids Delivery (GONAD) is a recently developed method for creating GM mice and rats without *ex vivo* handling of embryos [16–21]. It can be simply performed by injecting a solution containing nucleic acids into the oviductal lumen of pregnant females at zygote to two-cell stages and subsequent *in vivo* electroporation (EP) to enhance DNA uptake by early embryos *in situ*. In this case, there is no need for the large number of mice that is normally required for the traditional microinjection-based transgenesis: only four to five pregnant females are required for modifying an endogenous gene [18]. Thus, creation of GM animals is simplified with the development of GONAD, but an expensive apparatus electroporator is still required for the technique.

In 1989, Lavitrano et al. [22] reported the simplest, convenient, and cost-effective method to create GM animals, which was called sperm-mediated gene transfer (SMGT), where isolated sperm were incubated in the presence of naked plasmid DNA for a short period and these DNA-associated sperm were subjected to *in vitro* fertilization (IVF) with normal oocytes. The resulting progeny are later judged as those carrying the exogenous DNA in their genome. Since the report, there has been controversy over its reproducibility among researchers [23–25]. However, several recent improvements were made on this SMGT system by several researchers who employed reagents capable of enhancing gene delivery towards isolated sperm [26–28]. For example, Shen et al. [29] incubated mouse sperm in a solution containing 3% dimethyl sulfoxide (DMSO) and plasmid DNA for 10–15 min at 4°C prior to IVF. Embryos (42%; 25/60) obtained from this experiment showed bright enhanced green fluorescent protein (EGFP)-derived fluorescence. Furthermore, Kim et al. [30] reported that nanoparticles, such as magnetic nanoparticles, can be a vehicle for delivering exogenous DNA to sperm from various animals such as boar. They incubated boar sperm in the presence of 0.5% (v/v) of magnetic nanoparticles (MNPs) and plasmid DNA coding for green fluorescent protein (GFP) on the magnetic field for 90 min, and the magnetofected sperm were subjected to IVF with normal oocytes. As a result, they obtained fertilized eggs expressing GFP. Unfortunately, for further development of IVF-derived embryos, it still required ET towards recipient females, which is laborious and requires specialized skill. Notably, it is possible to perform artificial insemination (AI) using *in vitro*-transfected sperm, which can be simply done by injecting those sperm into the uterine horn or lumen of the oviducts of recipient females showing oocyte ovulation. This method, called "SMGT-based AI" (SMGT-AI), has already been performed by several laboratories and will be discussed in more detail in the last part of this paper.

 Testis-mediated gene transfer (TMGT) is the *in vivo* version of SMGT, in which sperm is transfected *in vivo*. This technology was first developed by Sato et al. [31], who performed intratesticular injection of calcium phosphate-precipitated plasmid DNA using a glass micropipette in mice. The injected exogenous DNA is transferred

 to epididymal sperm or spermatogenic cells within a seminiferous tubule (ST) of the testis, and those transfected sperm will transmit it to an oocyte through fertilization (**Figure 1**). They could detect the injected DNA in isolates of sperm from the epididymis and from the uteri of females mated with the injected males, but the DNA could not be detected in embryos [31]. In contrast with SMGT, TMGT does not require *ex vivo* handling of embryos such as collection of oocytes, IVF and ET. In this context, TMGT appears to be more convenient and simpler than SMGT for the purpose of GM animal production. Since the report of Sato et al. [31], several *in vivo*  gene delivery approaches targeted to male reproductive systems have been reported: gene delivery to spermatogenic cells by intratubular injection of STs (ST-mediated gene transfer, STGT) (**Figure 2a**), to epididymal sperm present in the epididymal ducts (epididymis-mediated gene transfer, EpiGT) (**Figure 2b**), and to sperm present in the vas deferens (vas deferens-mediated gene transfer, VDGT) (**Figure 2c**).

 Based on this background, the previous terminology TMGT appears to be now recognized as "direct *in vivo* gene delivery approach towards male reproductive system." In this context, it may be better to re-name TMGT as "intratesticular injection-based gene transfer" (IIGT), which involves direct injection of genetic materials into the interstitial space of a testis. Thus, IIGT, STGT, EpiGT and VDGT can be considered as TMGT-related experiments. In **Table 1**, a summary of previous studies on TMGT-related experiments is listed. Furthermore, there are several excellent papers reviewing the SMGT/TMGT-related studies [28, 90–93], which provide a helpful survey of this field.

### **Figure 1.**

*IIGT in mice. To perform IIGT, at least three different ways to inject a DNA-containing solution into the testis have been employed. The first way (shown in a) is to perform IIGT towards a testis (that is exposed outside after surgery) under anesthesia [31, 32]. The second way (shown in b) is to perform IIGT through insertion of a needle via scrotum under anesthesia [33]. The third way (shown in c) is to insert a needle at three times to different sites [34]. In these latter two cases, no surgery is required. Three to five days after IIGT, the IIGTtreated males are subjected to mating with normal estrous females. The in vivo transfected sperm may fertilize ovulated oocytes, leading to creation of offspring carrying the introduced exogenous DNA.* 

### **Figure 2.**

*There are several routes for introducing exogenous DNA into the male reproductive system. The routes for DNA injection are as follows: (a) STGT via rete testis; (b) EpiGT by inserting a glass pipette into the proximal region of caput epididymis; (c) EpiGT by inserting a glass pipette into the proximal region of cauda epididymis; and (d) VDGT by inserting a glass pipette into the proximal region of vas deferens.* 









*1 TMGT can be defined as a method for in vivo gene delivery towards male reproductive systems (testis, epididymis, and vas deferens) and includes intratesticular injection-based gene transfer (IIGT), seminiferous tubule-mediated gene transfer (STGT), epididymis-mediated gene transfer (EpiGT), and vas deferens-mediated gene transfer (VDGT).* 

*Abbreviations: BMPs, bacterial magnetic particles; DMSO, dimethyl sulfoxide; DOTAP, N-[1-(2,3-Dioleoyloxy) propyl]-N,N,N-trimethylammonium methyl sulfate; DMA, N,N-dimethylacetamide; EP, electroporation; GFP, green fluorescent protein; IVF, in vitro fertilization; PEI, polyethylenimine; STs, seminiferous tubules; Tg, transgenic; TB, trypan blue.* 

### **Table 1.**

*Summary of testis-mediated gene transfer (TMGT)1 -related studies.* 

In the following sections, the TMGT-related experiments will be mentioned in more detail.

### **2. Historical background of TMGT-related experiments**

### **2.1 IIGT-related experiments**

 Between 1994 and 2006, over 20 reports on IIGT-related experiments were reported using various animal models such as mice, rats, hamsters, rabbits, boar, goats, chicks, fishes, and shellfishes [29, 31–41, 43–53, 68]. The DNA used were mainly plasmid DNA that had been mixed with calcium phosphate, liposomes/ lipids, polyethylenimine (PEI), or DMSO, all of which were intended to facilitate uptake of DNA by sperm or spermatogenic cells [29, 31–34, 37–40, 44, 47, 48, 51, 68]. The method to use *in vivo* EP towards the entire testis after IITG with naked plasmid DNA was also employed [36, 41, 45, 49]. Furthermore, adenoviral vectors were introduced by IIGT [35, 43, 46, 50].

Through these works, the following information became available:


In 2007 and onward, attempts to improve IIGT systems were made by several laboratories to enhance the gene delivery efficiency [54–67]. In the following sections, we will describe several examples [(v) to (x)] about the improvement of IIGT, *in vitro* assessment for gene expression after IIGT or possible mechanism underlying IIGT.

 v.**IIGT at a young stage:** In mammalian testis, spermatogenesis occurs in the STs of a testis (**Figure 3d**). In the ST, there are spermatogenic cells called spermatogonia (spermatogonial cells) that can be further matured into spermatocytes and spermatids. Spermatogonia are largely divided into two types, type A and type B cells. The former undergoes active mitosis and divide to produce type B cells. The type B cells divide to give rise to spermatocytes and spermatids, which move towards the lumen of the ST as they mature. According to Hui-ming et al. [59], type A spermatogonia first appear between 3 and 7 days postnatally in mice and are the only immortalized diploid cells. They considered that if these type A spermatogonia are stably transfected with the exogenous DNA, the transfected cells would be able to produce mature Tg

> sperm leading to production of Tg mice through natural fertilization. Based on this hypothesis, they performed IIGT using GFP-expressing plasmid DNA encapsulated by the ExGen500 transfection reagent on 7-day old male ICR mice. When these treated mice reached different stages of sexual maturity (6, 12, and 24 weeks of age), they were mated with normal females. The resulting pups were identified as Tg, with efficiencies of 11–14%. They observed GFP expression in sperm cells isolated from F0 and F1 pups. They referred to this technology as "type A spermatogonia-mediated gene transfer" (TASMGT).


### **Figure 3.**

*EP-based IIGT in mice. a-c. Schematic illustration for EP-based IIGT. After IIGT towards both testes (a), they are held by a pair of tweezer-type electrodes and then subjected to the first in vivo EP (b). The second EP was next performed by changing the direction of electric pulse (c). (d) Structure of a ST. The colored spermatogenic cells and Sertoli cells indicate cells successfully transfected by the exogenous DNA that have been instilled within a lumen of ST.* 

### **Figure 4.**

*In vitro EP-based IIGT in goats. The isolated goat testis is subjected to IIGT with a total of eight repeated injections from different directions (a). Then, the entire testis is hold by a pair of electrodes prior to EP (b). After that, the EP-treated STs are partially dissected and subjected to in vitro cultivation (c). By this, the transgene expression on the spermatogenic cells of STs can be monitored continuously (d).* 

 such as nucleosides inside the cell. The authors hypothesized that a hypotonicTris-HCl solution at a certain hypotonic concentration might allow the germ cells to internalize the surrounding solutes like DNA *in vivo* without being killed and the sperm produced from transfected germ cells may carry a desired DNA fragment (transgene) to generateTg animals. Usmani et al. [65] suspended the linearized plasmid DNA (transgenes) in hypotonic Tris-HCl solution (pH 7.0) and simply performed IIGT to internalize the injected transgenes in the genome of spermatogonia residing at basal compartment of tubules. As a result, such males successfully generated Tg progeny by natural mating. This technique is easy and simple and does not require expensive apparatuses like electroporators. Usmani et al. [65] proposed that such a procedure enables researchers to generate their

own Tg animals, instead of outsourcing, and would drastically minimize the time required for studies on functional genomics.


Only a minor portion of the solution injected into the interstitial space of a testis is transferred inside the STs. This may be elicited by mechanical shearing of STs upon insertion of a needle or glass capillary. To increase the transfection efficiency in the spermatogenic cells existing within STs, repeated needle insertions (over at least three times from different sites) have been employed by some research groups [34, 46, 53, 59, 63]. At present, it remains unknown how many STs are indeed transfected by this treatment. Usmani et al. [65] reported that spermatogenic cells, including spermatogonia present within STs, are transfected after IIGT and subsequent *in vivo* EP using fluorescent marker genes.

### **2.2 STGT-related experiments**

ST is a tubular structure packed in a testis, which contains spermatogenic cells, such as spermatogonia, a precursor for mature sperm, and maturing sperm cells

### **Figure 5.**

*Possible mechanism of how the exogenous DNA introduced into the interstitial space of a testis is transmitted to epididymal sperm or to spermatogenic cells within a ST.* 

 (spermatocytes and spermatids), and Sertoli cells, which support the proliferation and differentiation of spermatogonia (**Figure 3d**). If the exogenous DNA is introduced within a lumen of the ST, it is possible to transfect those spermatogenic cells, from which transfected spermatozoa are transported via epididymal ducts and finally ejaculated upon mating with estrous females. As a result, the introduced exogenous DNA will be transmitted to oocytes via fertilization with the transfected sperm.

To our knowledge, STGT was first performed in 1997 by two groups: Blanchard and Boekelheide [35], with rats using adenoviral vector, and Kim et al. [68], with mouse and pigs using liposomally encapsulated plasmid DNA. The aim of the former group was to study transgene expression in the adult rat testis *in vivo*. They demonstrated that there was transgene expression in Sertoli cells and principal cells of the epididymis, and expression persisted for at least 10 days. The latter group demonstrated that in mice, 8.0–14.8% of STs expressed the introduced transgene, as evaluated by histochemical staining for the lacZ gene, and 7–13% of epididymal sperm had the exogenous DNA, as evaluated by PCR analysis. In pigs, 15–25% of the STs contained lacZ-positive germ cells. They suggested that STGT can be used as a powerful tool for producing Tg livestock.

 In 1998, Yamazaki et al. [69] employed STGT for examining transcriptional regulatory elements of spermatogenic specific genes. After injecting DNA into STs, subsequent *in vivo* EP was used to enhance DNA uptake by spermatogenic cells. Based on these experiments, they suggested that spermatogenic stem cells and/or spermatogonia can incorporate foreign DNA, and that the transgene could be transmitted to the progenitor cells derived from a transfected proliferating germ cell. Later, the same group [70] examined the possibility to create Tg offspring using STGT coupled with *in vivo* EP (EP-based STGT). Although longlasting transgene expression could be detected in the spermatogenic cells even 2 months after EP, no Tg offspring were obtained after natural mating with normal adult females.

 Huang et al. [71] used the EP-based STGT towards an entire mouse testis after intratubular injection of plasmid DNA that coded for fluorescent genes. To trace the fate of transfected spermatogenic cells, they obtained fluorescent sperm by fluorescence activated cell sorting (FACS), and performed ICSI to obtain their offspring. Almost all the individuals produced from fluorescent sperm were Tg. The authors

claim that this is the first report of gene transfer into germ cells and subsequent production of Tg offspring.

 STGT appears to be the direct approach to transfect spermatogenic cells *in situ*, but most experiments [46, 49, 51, 72–84] have been confined to successful transfection of spermatogenic cells and Sertoli cells for rescuing damaged/inactive Sertoli cells, *in vivo* testing of efficiency of RNA interference (RNAi), or establishment of an *in vivo* assay system to evaluate the promoter activity of the gene of interest. There have been no trials to create Tg offspring through STGT. Only some groups have tried to test the possibility of creating Tg animals by STGT. For example, Celebi et al. [74] performed STGT using circular plasmid carrying the lacZ reporter gene mixed with noncommercial cationic lipids. These injected males were mated with wild-type females and the progeny were analyzed by PCR and Southern blot assay. They demonstrated that the transgenes were transmitted to the offspring, but remained episomal, since it was found in the tail of the young animals and was lost at adulthood. Therefore, the plasmid seemed to be lost during the numerous germ cell divisions. This plasmid stayed in some tissues, such as in the skeletal and cardiac muscles. No integrative forms have yet been found with the use of circular DNA. Kanatsu-Shinohara et al. [75] described a novel approach for producing Tg animals by transducing spermatogonial stem cells *in vivo* using a retroviral vector by STGT. When these injected males were mated with wild-type females, Tg offspring were obtained with an efficiency of 2.8%. The transgene was transmitted stably and expressed in the next generation. The authors, thus, concluded that this technique will be useful as an alternative to the pre-exiting microinjection-based transgenesis, as well as provide a means for analyzing the self-renewal and differentiation processes of spermatogonial stem cells *in vivo*. Sehgal et al. [83] described a technique for the generation of Tg mice by infection of spermatogonial stem cells with recombinant lentiviruses expressing EGFP with a high rate of success. When the infected males were mated to normal females, over 60% of the delivered pups were found to be Tg. Li et al. [84] employed methods similar to those of Kanatsu-Shinohara et al. [75] and Sehgal et al. [83] and reported that the Tg efficiency is around 2.4%, which is similar to the previous report of Kanatsu-Shinohara et al. [75].

### **2.3 EpiGT-related experiments**

 Epididymal sperm present on the ducts of caput and cauda epididymides and epididymal epithelial cells can be targeted for transfection by the exogenous DNA. Kirby et al. [85] performed intraluminal injections (2–5 μL) of plasmid DNA into the lumen of an initial segment tubule of caput epididymis (**Figure 2b**), and subsequent *in vivo* EP towards the injected portion to examine the function of epididymal epithelial cells, which are thought to play critical role in sperm maturation during transport through epididymides. They concluded that this procedure is useful for elucidating the activity of promoter elements included in the injected plasmid that may not be identified when traditional *in vitro* methods are used. Esponda and Carballada [86] injected plasmid DNA mixed with the lipid FuGENE6 into the lumen of mouse cauda epididymis (**Figure 2c**). Successful transfection was observed in about 40% of cells after 2 days and in about 32% after 7 days, and then diminished progressively over time. Gene expression continued up to 15 days after gene injection and occupied about 22% of the area of the tubules. They concluded that intraluminal injections of exogenous DNA are effective for the study of epididymal physiology or to change the fertilizing ability of sperm. These studies are not aimed to create Tg animals, but they hold a potential to transfect epididymal sperm as a useful Tg tool.

### **2.4 VDGT-related experiments**

 In 1998, the Esponda's group [87] first attempted to examine whether exogenous plasmid DNA introduced into the lumen of the proximal region of the vas deferens could be taken up by mouse and rat sperm (**Figure 2d**). They demonstrated that 60–70% of sperm recovered 6 h after DNA injection had positive signal for successful transfection in their sperm nucleus, which was not affected by DNase treatment. This was also confirmed by PCR and slot blot analyses. They concluded that sperm within the vas deferens had the ability to incorporate exogenous DNA, which can be transferred to their nuclei, and vas deferens secretions do not block these capacities. In 2000, the same group [88] showed that this VDGT is useful for production of Tg mice. They injected plasmid DNA encoding GFP into the lumen of mouse vas deferens. The night after injections, males were mated with normal estrous females, and the offspring were analyzed. About 8% (4/53) of the newborns delivered expressed the GFP gene. They concluded that VDGT is a simple alternative to the pre-existing microinjection-based production of Tg animals and can be used for species in which the microinjection procedure is not feasible. This technology was later found to also be useful for transfection of epithelial cells of the vas deferens using a direct injection of DNA-liposome complexes, which could modify vas fluid contents [89].

### **3. Historical background of SMGT-based AI**

 As mentioned previously, AI of transfected sperm with exogenous DNA through SMGT is a highly convenient route for producing Tg animals. To our knowledge, Sperandio et al. [98] was the first to demonstrate its usefulness in domestic animals, such as bovine and swine. They performed AI towards ten sows with boar sperm cells that had been preincubated with plasmid DNA and obtained 82 offspring. Southern blot analysis of the DNA extracted from the animal tails showed that five animals were Tg and contained sequences complementary to the exogenous plasmid DNA that appeared to be rearranged compared to the original plasmid. From this study, it was suggested that SMGT-AI can be successfully adapted for the generation of Tg livestock. Yonezawa et al. [99] tested whether liposome-peptide (derived from human protamine)-DNA complex (LPD), a new reagent known to stabilize transfection in cultured cells, was useful to increase the efficiency of SMGT. They performed AI using rat epididymal sperm that had been incubated in a solution containing GFP expressing plasmid DNA and LPD complex. Expression of GFP was detectable in the morulae isolated from the treated animals. Furthermore, the AI-treated animals produced pups carrying foreign DNA.

This SMGT-AI is applicable to avian species. Yang et al. [100] performed AI using freshly-ejaculated chicken sperm that had been incubated in the presence of plasmid DNA and liposome, and found that about 4% (2/53) newly hatched chicks was identified as Tg. Harel-Markowits et al. [101] employed restriction enzymemediated insertion (REMI) to increase the efficacy of the transfection towards the isolated chicken sperm. REMI was used to insert exogenous DNA linearized with a restriction enzyme that cuts the genomic DNA at sites that enable the exogenous DNA to integrate via its matching cohesive ends [102, 103]. Following insemination with sperm transfected with linearized DNA, restriction enzyme, and liposome, they obtained Tg offspring. Furthermore, when chicken sperm are incubated in a solution containing plasmid DNA and DMSO or N,N-dimethylacetamide (DMAc) and subsequently subjected to AI, the resultant newborn chicks have the transgene, with efficiencies of 38% (for the DMSO-treated group) and 19% (for the

DMAc-treated group) [104]. However, Chaparian et al. [105] recently reported that they were unable to create Tg chicks by SMGT-AI.

### **4. Exosomes as a possible carrier to deliver genetic materials to sperm**

 Exosomes, membrane-enclosed sub-cellular microvesicles shed from most cell types, are present in a wide variety of body fluids [106]. Recently, it was found that they can mediate various effects on the behavior of recipient cells, since they contain cytokines, growth factors, and membrane proteins [107]. Furthermore, they contain a substantial amount of small and functional RNA molecules, called microRNAs (<100 nucleotides in length) [108], which could potentially control gene expression of various endogenous genes. It has recently been shown that these exosomes are (1) found in human semen [109], (2) involved in sperm maturation process during the transit along the male epididymal tracts [110], (3) accumulated in mature spermatozoa nuclei [111], and (4) delivered to oocytes through fertilization [112].

Notably, there are some reports describing non-Mendelian germline-independent inheritance of phenotypes in the absence of any classically identifiable mutation or predisposing genetic lesion in the genome of individuals who develop the disease [113–115]. For example, Cossetti et al. [116] performed subcutaneous inoculation of EGFP-expressing human melanoma cells into an immunocompromised mouse, from which EGFP RNA was released from the grafted melanoma cells, delivered to the bloodstream, and finally brought to sperm. When epididymal sperm isolated from these tumor-bearing males were examined carefully, the EGFP RNA was found to be tightly associated with the extracellular fraction of these mature sperm. They termed this phenomenon "soma-to-germ line transmission of information," and thought that exosomes may be involved in this phenomenon as the carrier to deliver EGFP RNA. The findings of Cossetti et al. [116] appear to be well correlated with those obtained from the TMGT-related experiments done at earlier stages of IIGT development, which include (1) non-Mendelian transmission of the exogenous DNA in the offspring obtained [33], (2) extreme low copy number of the exogenous DNA (<1 copy per diploid cell) transmitted to these offspring [32], (3) mosaic expression of the exogenous DNA in the offspring (blastocysts) obtained [33, 40], and (4) reduction in the number of offspring carrying the exogenous DNA during development [40]. As mentioned in Section 2.1 (x), parts of a solution introduced into the interstitial space of a testis is transferred to the excurrent ducts of epididymides, and the exogenous DNA may be taken up by the extracellular fraction of epididymal sperm, possibly through exosomes. We detected the presence of exogenous DNA in the DNase I-treated epididymal sperm, which have been isolated from the IIGT-treated males [42]. This may be due to the fact that exosomes can protect its exogenous DNA against DNase I-mediated digestion.

Notably, in their review article, Jiang and Gao [117] demonstrated that exosomes can be used as naturally occurring cell-to-cell transporters or as novel biocarriers for gene and drug delivery. These exosomes are naturally secreted by the cells and pass through additional biological barriers. They are more biocompatible and biodegradable and can avoid immune response which is most likely due to the surface expression of the complement regulatory proteins, such as CD55 and CD59. For these natural characteristics, exosomes are being extensively explored as gene delivery vehicles. For example, in 2011, Alvarez-Erviti et al. [118] first demonstrated that exosomes are useful for delivering short interfering (si)RNA to the mouse brain. They engineered dendritic cells to express lysosome-associated membrane protein 2 (Lamp2) isoform (Lamp2b), an exosomal membrane protein fused to a neuron-specific peptide. Following that, exosomes were isolated from

the gene-engineered dendritic cells and loaded with siRNA using electroporation and were administered intravenously to mice. As a result, the targeting peptide was shown to be successfully delivered to the brain. The concomitantly delivered siRNA caused reduced expression of a target protein associated with the pathogenesis of Alzheimer's disease. Furthermore, Lin et al. [119] proposed that exosomes can be a good carrier to introduce various cargoes, including plasmid DNA, into a cell. They prepared a mixture composed of purified exosomes isolated from HEK293FT cell line, pEGFP-C1 plasmid DNA, and Lipofectamine 2000 liposomes *in vitro*. During the incubation at 37°C for 12 h, exomes and liposomes are fused together and the exogenous plasmid DNA becomes incorporated into exosome-liposome hybrid nanoparticles. Transfecting mesenchymal stem cells (MSCs), which cannot be transfected by the liposome alone, with this complex resulted in successful generation of fluorescent cells when evaluated by FACS. Now, an exosome-based transfection kit, possibly based on this principle, is commercially available: Exo-Fect Exosome Transfection Kit (System Biosciences). We confirmed the usefulness of this kit by *in vivo* transfecting oviductal epithelial cells through intraoviductal instillation of a solution prepared using this kit. Some oviductal epithelial cells were found to be fluorescent after transfection with a plasmid expressing GFP (unpublished results). Thus, it may be possible to transfect isolated sperm by incubating plasmid DNA and exosome/liposome hybrid vesicles provided from the Exo-Fect Exosome Transfection Kit, prior to AI as mentioned below.

### **5. Genome-editing sperm**

Gene modification based on recently developed techniques such as zinc-finger nucleases (ZFNs), TAL effector nucleases (TALENs), and clustered regularly interspersed short palindromic repeats/CRISPR-associated protein 9 (CRISPR/ Cas9) are now recognized as a revolutionary genetic engineering tool *in vitro* and *in vivo* [120–124]. Three types of endonucleases from ZFNs, TALENs, and CRISPR/ Cas9 have been developed to promote precise genome editing at a target gene. All these enzymes have a DNA-binding ability and an ability to elicit double-strand DNA break (DSB) at a target genomic locus. Subsequently, in the absence of a homologous template to repair, nonhomologous end joining (NHEJ) occurs and causes small insertions or deletions (termed "indels"). In the presence of a template donor DNA, site-specific recombination through homology-directed repair (HDR) occurs. Generally, the frequency of NHEJ is thought to be higher than that of HDR in most of the cell types [125]. **Table 2** shows comparison among these three technologies.

ZFNs are the first engineered endonucleases [127] that combine the DNA recognition ability of zinc-finger protein (called zinc-finger motifs) and restriction enzyme *Fok I* to introduce DSB [120–124]. In 2005, Urnov et al. [128] first demonstrated that ZFNs are effective as a genome editing system in the human cells.

TALENs are similar to ZFNs and require a string of TALEN motif (consisting of a series of 33–35 amino acid repeats) to bind to the specific sequence of a target gene and *Fok I* enzyme to introduce DSB [120–124]. TALENs provide more flexibility to the target sequences since ZFNs are known to be more active towards GC-rich region, whereas TALENs can be assembled to the target AT-rich regions [120–124].

Since both ZFNs and TALENs require assembling of an array to build each set, which is a complex and time-consuming process [124], CRISPR/Cas9 has become the favorite because of its easy application. CRISPR/Cas9 requires only two components,


### **Table 2.**

*Comparison of ZFN, TALEN, and CRISPR/Cas9-mediated genome editing systems1 .* 

 the Cas9 nuclease and a single-guide RNA (sgRNA), which is a short sequence to guide the Cas9 protein to a target site. More importantly, these events are performed by transient interaction of Cas9 and sgRNA, whereby chromosomal integration of the CRISPR components is not a prerequisite [129]. There is a concern of off-target cleavage activity from the endonuclease from CRISPR/Cas9 because the system requires recognition of only 20 bp target sequence and allows up to 5 bp mismatches for the formation of DSB [124]. Several strategies for minimizing the off-target cleavage have been employed including use of double nickase mutant form of Cas9, which induces a single-strand break instead of DSB [130]; use of Cas9-sgRNA ribonucleoprotein (RNP) complex, whose half-life is shorter than that the time in which plasmid or viral nucleic acid is transcribed [131]; or use of fusions of catalytically inactive Cas9 with *Fok* I nuclease domain (fCas9) to improve the DNA cleavage specificity [132].

 In the case of producing GM animals using SMGT or TMGT, it is better for the exogenous DNA (transgenes) to be integrated into the chromosomes of sperm. This event appears to occur more frequently in immature sperm cells present in the ST of a testis than in the mature epididymal sperm because the chromosomal DNA in the latter cells is tightly packed in the head region of a sperm. In this context, STGT is a preferable system to create GM animals because it is targeted to transfection of spermatogenic cells present within the STs. However, it takes about 4 weeks for mature sperm to reach the epididymal portion for fertilization. If a researcher wants to generate GM animals within a short period of time, direct transfection of mature sperm present in epididymides or vas deferens is recommended. In this case, as mentioned above, the introduced exogenous DNA may be associated to the extracellular fraction of a sperm, as episomal DNA. Notably, CRISPR/Cas9 based genome editing does not always require chromosomal integration of its components; it can be performed by transient expression of their components after transfection [129]. In this sense, an attempt to transfect mature sperm would be a useful alternative for GM animal production.

### **6. Proposal of new experimental systems for simple creation of genome-edited animals using** *in vivo* **or** *in vitro* **transfected sperm**

In the following section, we propose two experimental plans to create genomeedited animals using VDGT or AI-based systems, all of which are simpler and more convenient than the previously described systems.

### **6.1 VDGT-based genome editing**

As previously described in Section 2.4, VDGT enables transfer of exogenous DNA to oocytes via fertilization by mature sperm transfected within vas deferens [87, 88]. Injecting a solution containing genome editing components (e.g., sgRNA + DNA/ mRNA/protein for Cas9) into the lumen of vas deferens of anesthetized males and subsequent mating between the VDGT-treated males and normal estrous females the day (night) after the surgery may result in production of genome-edited offspring.

 In **Figure 6a**–**c**, experiments obtained after TB injection into the lumen of mouse vas deferens is shown (unpublished results). Under anesthesia, cauda epididymis and vas deferens were pulled out and a small slit was made at the proximal region of vas deferens using micro scissors (**Figure 6a**). Then, a glass micropipette containing TB was inserted into the lumen of vas deferens under observation using a dissecting microscope and about 15 μL of the solution is slowly injected (**Figure 6b**). It is easily discernible that the injected TB still remains within the proximal portion of vas deferens immediately after the injection (arrows in **Figure 6c**). However, the injected TB moved to the distal portion of vas deferens the next day (arrow in **Figure 6d**). Thus, to produce genome-edited animals by VDGT, a solution containing CRISPR/Cas9 components (sgRNA + DNA/mRNA/protein for Cas9), gene delivery enhancing reagents (such as DMSO, liposomes, microparticles, etc.) and fluorescent marker expression plasmid DNA has to be prepared first (**Figure 6e**).

### **Figure 6.**

*Procedure for VDGT-based genome editing. (a–d) Experimental procedure of VDGT when TB as a visible marker is injected into the lumen of vas deferens. Under anesthesia, a small slit is made at the proximal region of vas deferens using micro scissors (a). Then, a glass micropipette is inserted into the lumen of vas deferens under observation using a dissecting microscope and the solution is slowly injected (b). After TB injection, the injected TB still remains within the proximal portion of vas deferens (arrows in c). One day after VDGT, the injected TB moves to the distal portion of vas deferens (arrow in d), showing the flow of the injected substance. e. Experimental procedure of VDGT when genome-editing components are injected into the lumen of vas deferens. First, a solution containing CRISPR/Cas9 components, gene delivery enhancing reagents (such as DMSO, liposomes, microparticles, etc.), and fluorescent marker expression plasmid DNA is prepared in a tube. Then, about 15 μL of this solution is immediately injected into the lumen of vas deferens. On the night following the VDGT or the next day, the VDGT-treated males are mated to normal estrous females. Later, cleavage stage embryos are collected to examine the presence/expression of the transgene (plasmid), as well as possible mutations in a target locus.* 

 After short incubation period, this solution is injected into the lumen of vas deferens. Then, the VDGT-treated males are mated to normal estrous females on the day (night) or next day. Later, cleavage stage embryos were collected from the VDGTtreated females to examine the presence and expression of the transgene (plasmid) (**Figure 6e**, bottom) and occurrence of mutations in a target locus. In some cases, the SMGT-AI-treated females were allowed to deliver their pups to see whether genome editing is induced in their chromosomes.

### **6.2 SMGT-AI-based genome editing**

AI is one of the assisted reproduction technologies that is based on the introduction of isolated sperm into the female reproductive tracts, such as uterine horn or oviductal lumen, to *in vivo* fertilize ovulated oocytes. As previously described in Section 3, isolated sperm are incubated in a solution containing exogenous DNA and gene delivery enhancing reagents such as DMSO, liposomes, and microparticles, for a short period (SMGT), and then the transfected sperm are subjected to AI, called "SMGT-based AI" (SMGT-AI). During this process, the exogenous DNA should be transmitted to oocytes via fertilization resulting in Tg embryos.

We previously reported that transfer of sperm into a space near the infundibulum between the ovary and ovarian bursa enables *in vivo* fertilization of ovulated oocytes in the ampulla region of the oviduct [133, 134]. In more detail, 2 μL of fresh epididymal B6C3F1 (F1 hybrid mice between C57BL/6 and C3H) sperm (containing 2 x 105 spermatozoa) were intrabursally injected 7 h after human chorionic gonadotropin (hCG) administration to B6C3F1 females that had been administrated with pregnant mare serum gonadotropin (PMSG) 48 h before. At 1.7 days after AI, normal cleaving embryos were recovered at rates of 40–100%. We called this AI technology "intrabursal transfer of sperm" (ITS) [133]. In **Figure 7a**, the ITS procedure is schematically illustrated. In **Figure 7b**  and **c**, photographs before (b) and after (c) ITS are shown by using TB as a dye to visualize the process of AI. It is clear that the injected solution is present between the ovary and ovarian bursa (arrow in **Figure 7c**). In **Figure 7d**, an example for SGMT-AI-mediated genome editing in embryos is schematically shown. First, sperm isolated from the vas deferens are treated with CRISPR/Cas9 components (sgRNA + DNA/mRNA/protein for Cas9), gene delivery enhancing reagents (such as DMSO, liposomes, microparticles, etc.), and fluorescent marker expression plasmid DNA for a short period. Then, the solution containing the transfected sperm is subjected to AI towards females 7 h after hCG administration. The next day, 2-cell embryos are collected from the AI-treated females to examine the presence and expression of the transgene (plasmid) and occurrence of mutations in a target locus. In some cases, the SMGT-AI-treated females are allowed to deliver their pups to see whether genome editing is induced in their chromosomes. Notably, the selection of a successfully genome-edited sperm prior to AI may accelerate the production efficiency of genome edited offspring, although the practical approach for this remains unknown at present. Therefore, the success or failure of genome editing performed in this system may depend on the molecular analysis of the offspring (e.g. blastocysts or fetuses) generated after AI of the SMGT-treated sperm.

### **7. Conclusion**

TMGT, based on direct *in vivo* gene delivery towards interstitial space of a testis, ST within a testis, or excurrent ducts of epididymides and vas deferens, is less labor

### **Figure 7.**

*Procedure for SMGT-AI-based genome editing. (a) ITS procedure schematic. (b, c) Photographs during before (b) and after (c) ITS, which is shown by intrabursal injection of TB. Note the presence of TB between the ovary and ovarian bursa (arrow in c). (d) Experimental procedure of SMGT-AI when genome-editing components are injected between the ovary and ovarian bursa. First, sperm isolated from the vas deferens are incubated in a solution containing CRISPR/Cas9 components, gene delivery enhancing reagents (such as DMSO, liposomes, microparticles, etc.) and fluorescent marker expression plasmid DNA for a short period. Then, the solution containing the transfected sperm is subjected to AI towards females 7 h after hCG administration. Later, cleavage stage embryos are collected to examine the presence/expression of the transgene (plasmid), as well as possible mutations in a target locus.* 

 intensive and less time consuming for the production of GM animals. This testicular route is also ethically superior since fewer mice are required than existing alternative methods of transgenesis. The TMGT-treated males can be used to mate with estrous females, through which the exogenous genetic materials are transferred to oocytes at fertilization. During this process, there is no need for *ex vivo* handling of embryos, which is strictly required for zygote-based gene modification such as microinjection, EP, viral infection, and TransICSI. SMGT-AI, based on AI of sperm that have been transfected *in vitro* with the exogenous DNA, is also a convenient system for production of Tg animals, like TMGT. The CRISPR/Cas9 system, one of the recently developed genome editing technologies, is now recognized as a powerful and simple tool to create GM animals. More importantly, in this system, chromosomal integration of the genome editing components is not the prerequisite. Coupling this genome editing system with TMGT or SMGT-AI would accelerate creation of genome-edited animals in a more convenient manner. Furthermore, TMGT/SMGT-AI will be particularly useful for other animals that are difficult to manipulate as early embryos *in vitro*.

### **Acknowledgements**

This study was partly supported by a grant (no. 24580411 for M.S.; no. 16H05049 and 16 K15063 for S.N.) from the Ministry of Education, Science, Sports, and Culture, Japan. Author Contributions: Masahiro Sato designed the study and drafted the manuscript; Shingo Nakamura critically revised the manuscript.

### **Conflicts 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.

### **Author details**

Masahiro Sato1 \* and Shingo Nakamura<sup>2</sup>

1 Section of Gene Expression Regulation, Frontier Science Research Center, Kagoshima University, Kagoshima, Japan

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

\*Address all correspondence to: masasato@m.kufm.kagoshima-u.ac.jp

© 2019 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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Section 3

Genetically Modified

Organism (GMO)

## Section 3
