Exogenous enzymes upgrade transgenesis and genetic engineering of farm animals

Abstract

Transgenic farm animals are attractive alternative mammalian models to rodents for the study of developmental, genetic, reproductive and disease-related biological questions, as well for the production of recombinant proteins, or the assessment of xenotransplants for human patients. Until recently, the ability to generate transgenic farm animals relied on methods of passive transgenesis. In recent years, significant improvements have been made to introduce and apply active techniques of transgenesis and genetic engineering in these species. These new approaches dramatically enhance the ease and speed with which livestock species can be genetically modified, and allow to performing precise genetic modifications. This paper provides a synopsis of enzyme-mediated genetic engineering in livestock species covering the early attempts employing naturally occurring DNA-modifying proteins to recent approaches working with tailored enzymatic systems.

Introduction

Transgenic farm animals can serve as excellent models of human diseases, of biopharming, and of basic research [1–5]. During the past few years, transgenic farm animals have gained renewed popularity, because of the availability of annotated genome depositories (http://www.ensembl.org; http://www.ncbi.nlm.nih.gov/genome) and because of the introduction of active methods of transgenesis. Active transgenesis refers to the introduction of exogenously provided enzymes or nucleic acids encoding them [2, 6, 7], which catalyze specific gain-of-function or loss-of-function genetics in an unprecedented pace. The exogenous enzymes are only transiently present; however, by carefully selecting highly active or hyperactive variants [8–11], the desired genetic modification can be performed in individual cells, such as the mammalian zygote. Prominent examples are hyperactive transposon systems, such as Sleeping Beauty [9, 10], piggyBac [11], as well as designer nucleases, including zinc finger nucleases (ZFNs), transcription activator-like element nucleases (TALENs) and RNA-guided nucleases [1–3, 12]. In addition, Cre recombinase and ΦC31 integrase found some interest for farm animal transgenesis [13, 14]. Viral integrases (retrovirus, lentivirus, adeno-associated virus) apply similar mechanisms; however, viral transgenesis is already covered by a number of excellent reviews [15–18], and will not be discussed in this paper. The repertoire of molecular tools now allows the precise modification of large mammalian genomes at rapid pace and has led to a recent boost in this field [2, 19–22].

Brief time course of livestock transgenesis

Since the isolation of class II restriction enzymes (RE), the hypothesis that simultaneous delivery of a RE in combination with a transgene would increase the efficiency of foreign DNA incorporation was postulated. First evidence for this hypothesis came from a study in which illegitimate integration events of non-homologous DNA fragments into yeast genome was several fold enhanced when a standard RE was included in the transformation mixture [23]. The restriction enzyme-mediated integration (REMI) was subsequently used in unicellular organisms, fungi [24, 25] and xenopus [26]. The catalytic activity of RE in cultured mammalian cells [27, 28] prompted researchers to apply REMI in combination with pronuclear microinjection of mouse zygotes [29]. In the mouse model, the rate of transgenic embryos and live pups by PCR analysis was doubled (18 vs 9 %) in REMI versus standard pronuclear injection [29]. However, no information regarding number of copies, genomic sites of transgene incorporation, expression and transmission to progeny were given [29]. Further independent replications of this approach are warranted to unequivocally establish the usefulness of REMI for animal transgenesis.

It has been postulated that the co-delivery of transgene and a site-specific RE could increase efficiency of integration into the host genome by three non-excluding ways: (1) protecting the ends of the transgene constructs, (2) inducing DNA breaks, and (3) stimulating endogenous DNA repair mechanisms [30].

It has been well established that frequently cutting RE may pose a potential risk of causing genotoxic damage [31, 32]. In fact, introduction of a RE by electroporation into mammalian cells has been shown to induce genomic rearrangements such as deletions, duplications, and translocations [28, 33, 34]. To avoid these detrimental effects, REMI can be performed with rare-cutting meganucleases. Meganucleases or homing endonucleases (HE) are naturally occurring enzymes that recognize long consensus sequences spanning 12–40 bp [35]. One of the best characterized is I-SceI from Saccharomyces cerevisiae [36]. Albeit the consensus sequence spans 18 bp, I-SceI seems to allow some ambiguity in the recognition site. Co-injection of a transgene flanked by two I-SceI restriction sites with purified I-SceI into fertilized eggs of Oryzias latipes (medaka fish) and Xenopus tropicalis resulted in improved transgenic efficiencies [37, 38]. Preliminary studies assessed I-SceI for transgenesis in livestock [39]. I-SceI-injected bovine zygotes resulted in an increased proportion of embryos expressing the reporter gene and a reduced percentage of mosaic embryos. In a preliminary report [40], the I-SceI approach significantly increased ratio of transgenic bovine fibroblasts, suggesting that I-SceI can enhance transgene integration into the cattle genome. Recently, the first reporter transgenic pigs were generated by an I-SceI approach; however, no details about copy numbers, integration sites and transgene silencing have been reported [41]. Further studies are warranted to reveal the mechanistic role of I-SceI during mammalian transgenesis.

The first evidence for sperm-mediated transport of native simian virus 40 (SV40) DNA into rabbit oocytes came from a study by Brackett et al. [42]. The SV40 DNA encoded the complete genome of the virus, and infective SV40 virions could be recovered by coculture of fertilized embryos with a permissive kidney cell line from African green monkey. Eighteen years later Lavitrano et al. [43] used sperm mixed with plasmid DNA to produce transgenic mice. This report of sperm-mediated gene transfer (SMGT) was soon challenged by an independent study, which failed to replicate the experiment [44]. Since then, SMGT has been assessed for transgenesis in several invertebrates as well as vertebrates [45–47], including domestic species [48–50]. However, most of the studies in mammalian species provided poor evidences for transgene integration or recombinant protein expression.

In 1999, an alternate technique called intracytoplasmic sperm injection-mediated transgenesis (ICSI-Tr) was published [51]. Developed originally to produce transgenic mice, the method was later translated to other mammalian species [52–56] and birds [57]. In this methodology, double-stranded DNA molecules are complexed with membrane-damaged (dead) spermatozoa, which were subsequently microinjected into the cytoplasm of metaphase II oocytes. In this modification of SMGT, the physical or chemical disruption of sperm cell membranes is a prerequisite for successful gene transfer, which then requires the troublesome ICSI procedure. With ICSI-Tr, high percentages of transgenic offspring with low incidence of mosaicism have been reported [51]. Despite the success of this technique in terms of transgenic ratios, it does not escape from the numerous drawbacks of methodologies that rely on passive integration of transgenes, such as concatemeric transgene integration, silencing, and variegated transgene expression [58, 59].

To address some of these concerns, approaches to combine ICSI-Tr with the delivery of ectopic enzymes were assessed [60]. Initial experiments addressed the effect of a bacterial recombinase (RecA) [60] and Tn5 transposase [61] on mouse and livestock transgenesis [62, 63]. Both enzymes were able to increase the proportion of live transgenic animals compared to classic pronuclear microinjection and ICSI-Tr methods [64], but seemed to suffer from sub-optimal activities of the employed enzymes [29, 30].

The need for advanced transgenic methodologies that permit precise genetic and highly efficient modifications in preselected DNA sequences has driven research efforts to develop hyperactive and codon-optimized transposases (SB, PB, Tol2) [10, 11, 65–68], recombinases (Cre, flippase) [13, 69] and customized programmable nucleases, like zinc finger nucleases (ZFN), transcription activator-like element nucleases (TALEN), and RNA-guided nucleases [2, 19, 22, 70], which already initiated a revolution in the field of animal transgenesis.

Application of transposon systems for genetic engineering

Transposons or jumping genes belong to a diverse family of genetic elements that are able to move horizontally in genomes. Transposons were originally described in maize [71], but later identified as widespread components in the genomes of prokaryotes and eukaryotes [72]. Interestingly, transposable elements comprise high proportions of eukaryotic genomes (i.e., about 45 % of the human genome [73]) and the vast majority of them are inactive due to accumulated deleterious mutations [74].

Transposons are grouped in two distinct categories according to the mechanism used for mobilization (transposition). Class I transposons also called retrotransposons rely on a RNA intermediate, which is reverse-transcribed in a new genomic locus. As consequence the number of genomic units increases by a mechanism that can be characterized as “copy and paste” [75]. Retrotransposon mobilization is capable of inducing random mutations at high frequency, disrupting endogenous genes, and therefore it has been held responsible for causing several genetic disorders [75, 76].

Class II transposons or DNA transposons are mobilized by a process that operates by a “cut and paste” mechanism [10, 77]. A transposase specifically recognizes inverted terminal repeats (ITRs), and precisely removes and relocates the ITR-flanked DNA segment to a different genome position [10, 77] (Fig. 1).

Fig. 1.

General mechanism for transposase-mediated transgene integration into the host genome. Transposase (TPase) is commonly synthesized from an expression vector (helper plasmid), mRNA or it is (rarely) provided as protein. The gene of interest (GOI) flanked by ITRs is delivered on an independent vector (donor plasmid) (a). TPase recognizes/binds ITRs flanking the GOI and catalyzes staggered double-strand breaks at the ends of ITRs (b). The resulting transposon–transposase complex binds at specific target sites in the host genomic DNA (c), and integrates the ITR-flanked transposon (d–e)

DNA transposons have been manipulated as useful gene transfer vectors for germline transgenesis, insertional mutagenesis and somatic cell transgenesis (gene therapy) (reviewed in [78]). Bicomponent transgenic systems have been developed [79] in which the gene of interest (GOI) is flanked by ITRs, and the transposase is provided in trans as mRNA or as an expression plasmid (helper plasmid) (Fig. 1). The excised transposon plus transposase proteins binds to a target DNA, where the insertion takes place [80]. Most transposases catalyze integration at short consensus sequences, for example TC1/mariner transposases recognize TA dinucleotides, and PB transposase recognizes TTAA tetranucleotides (Fig. 1). Through this mechanism, one monomeric copy of a transposon is integrated in the genome, leaving the empty backbone of the donor plasmid, which is eventually degraded (Fig. 1) [77].

Due to the development of hyperactive transposase variants, two-component transposon system has been adopted as an improved tool for germline transgenesis in a broad range of invertebrate and vertebrate species (reviewed in [81]). Integration efficiencies reported for various transposon systems (Table 1) rival the high integration rates of viral-based methods. In contrast to viral methods, transposon systems are characterized by being safe and capable of delivering large cargos [68, 82–85]. An advantage of transposase-mediated transgenesis is the monomeric transgene integration, making these constructs less prone to silencing in transposon transgenic animals [65].

Table 1.

Transposon transgenesis approaches in livestock

Transposon	Species	Construct	Antibiotic selection	Overall efficiency*	Expression pattern	Unspecific integrations	Germline transmission generations	Method	References
Sleeping Beauty	Rabbit	CAGGS-Venus	AB-free	1.4 %	Ubiquitous	No	F0, F1, F2	PNI	[258]
	Pig	CAGGS-Venus	AB-free	6.8 %	Ubiquitous	~5 %	F0, F1, F2	CPI	[65]
	Pig	floxedUbi-GIN	G418	NA	Ubiquitous	~25 %	F0	SCNT (HMC)	[13]
	Pig	DIV PuroΔtk APOBEC3G	Puromycin	~3 %	Ubiquitous	No	F0	CT	[87]
	Pig	INV-hITGA2/NV-hITGB1	G418	NA	Keratinocyte	ND	F0.	SCNT (HMC)	[259]
	Pig	HCR-hAAT-D374Y-PCSk9	Puromycin	NA	Liver	ND	F0	SCNT (HMC)	[260]
PiggyBac	Chicken	CAGGS-EGFP-IRES-Puro	Puromycin	NA	Ubiquitous	ND	Prefounder, F1	PGC transfection in vitro	[261]
	Chicken	CMV-EGFP SV40-Neo	G418	49.6 %	Ubiquitous	ND	Prefounder, F1, F2	PGC transfection in vitro	[262]
	Chicken	IRES-LacZ-CAGGS-EGFP-PGK-Neo	G418	NA	Ubiquitous	ND	Prefounder	Embryo microinjection and electroporation	[263]
	Pig	CMV-Neo-EGFP	G418	1.3 %	Ubiquitous	ND	F0	SCNT	[111]
	Pig	CAA–tdTomato	AB-free	7.0 %	Eye lens	5 %	F0	CPI	Unpub. (KW)
	Pig	CAGGS-EGFP, SV40-Hygro	none	5.7 %	Ubiquitous	ND	F0	CPI	[106]
Tol2	Chicken	CAGGS-EGFP-IRES-Puro	Puromycin	NA	Ubiquitous	ND	Prefounder	PGC transfection in vitro	[261]
	Chicken	CAGGS-EGFP	AB-free	1.5 %	Ubiquitous	ND	Prefounder, F1	PGC transfection in vivo	[264]

AB-free antibiotic selection marker-free, NA Efficiencies as transgenic offspring per treated embryos are not applicable, ND not determined, CAGGS cytomegalovirus early enhancer/chicken beta-actin promoter, CMV cytomegalo virus (immediate early) promoter, Ubi ubiquitin C promoter, SV40 simian virus 40 promoter, DIV diverse promoters were tested, PGK phosphoglycerate kinase promoter, CAA crystallin Aα promoter, GIN EGFP-IRES-neomycin, APOBEC3G apolipoprotein B mRNA-editing enzyme, INV involucrin promoter, hITGB1 human beta1 integrin, hD374-PCSK9 D374Y gain-of-function mutation in the proprotein convertase subtilisin/kexin type 9, HCR-hAAT hepatocyte control region and human α1-antitrypsin promoter, hITGA2 human Integrin α2 (CD49b), HMC hand made cloning, PGC primordial germ cell

* Transgenic offspring per treated oocytes or embryos

The reported instability of transgene expression from sequences inserted by non-facilitating mechanisms has been linked to methylation of CpG-rich vector sequences [86] that flank the transgene and are co-inserted with the transgene. An added advantage of transposition transgenesis is that each event can be later segregated in the descendants [65, 79, 87]. Segregation of independently inserted sequences by transposition would maximize the overall efficiency of the methodology. The same segregation process can serve to recycle marker/antibiotic selection cassettes to comply with current regulatory guidelines regarding transgenic animals.

Under certain scenarios, intentional removal of the stably inserted sequences is required to turn on or off transgene expression, being the conditional transgenesis an illustrative example of such applications [88]. Since transposition does not change the ITRs [80], the transposon is susceptible to be remobilized, and eventually removed, if the transposase is reintroduced in the system. This can be exploited to excise unwanted genomic DNA sequences flanked by transposon ITRs. Proof of principle for this potentially useful strategy has come from experiments with induced pluripotent stem (iPS) cells [89, 90]. The recent development of transposase variants, which are excision competent, but integration deficient will facilitate the seamless removal of transposons [91]. Thus, transposon systems combine high delivery rates of transgenes and the possibility of seamless transposon removal.

The use of DNA transposons to engineer vertebrate genomes began in 1997, when an active transposase, SB, was reconstructed from non-functional transposon sequences isolated from several salmonid species [79]. It was demonstrated that the original SB variant can transpose DNA sequences in a broad range of vertebrate species [65, 92–95] with moderate activity [9, 96–98]. Using an in vitro evolutionary approach, Mátés et al. [10] finally came up with a hyperactive version: SB100X. Since then, this hyperactive transposase has become the gold standard for transposition approaches in animals.

Successful implementation of SB-mediated integration for germline transgenesis in small animal models was followed by translational research aimed to produce transgenic livestock animals (Table 2). There are two established methodologies to generate transgenic large animals, DNA microinjection of zygotes or somatic cell nuclear transfer (SCNT) (Fig. 2).

Table 2.

Recombinase superfamily divisions (adapted from [126])

Superfamily	Family	Subdivisions	Recognition sites	Activity	Representative members
Site-specific recombinases	Tyrosine recombinases	Bidirectional	Identical	Reversible	Cre
				Inversion, excision and integration	FLP
					R
		Unidirectional	Non-identical	Irreversible	Lambda
				Inversion, excision and integration	HK101
					pSAM2
	Serine recombinases	Small	Identical	Irreversible	Beta-six
				Excision	CinH
					ParA
		Large	Non-identical	Irreversible	Bxb1
				Inversion, excision and integration	ΦC31
					TP901

Fig. 2.

Genome engineering via in vivo and in vitro approaches. Injection of nucleic acids and/or protein into the pronucleus of a zygote (a) or into the cytoplasm (b). Genetic modification in primary cells, which are subsequently used in SCNT (c)

Microinjection of pronuclear stage embryos, developed by Gordon et al. [99], became a routine technique to produce transgenic mice. Later, pronuclear microinjection (PNI) was adapted to livestock zygotes [100, 101]. Unlike mouse oocytes, the porcine and bovine counterparts are darkened by lipid droplets precluding the visualization of pronuclei. Therefore, high-speed centrifugation of zygotes is mandatory to visualize the pronuclei [100, 101]. PNI is a technically demanding methodology, which has been characterized by low efficiency in terms of number of transgenic offerings per injected embryo, and variable and instable expression of the recombinant protein in the transgenic animals. The cytoplasmic injection (CPI) of plasmids into the cytoplasm of one-cell embryos [102] represents a simplified alternative, making it suitable for species with opaque zygotes (Fig. 2). Both PNI and the CPI methods were successfully employed with SB, PB and Tol2 transposon components for germline transgenesis in fish [103], frogs [92], mice [10, 104], rats [105] and domestic pigs [65, 87]. A significant increase in the ratio of transgenic animals per microinjected zygotes has been consistently reported. The feasibility and efficiency of transposon-mediated transgene integration into the pig genome are supported by the high proportion of born animals carrying at least one copy of the transgene (>40 %) [65, 66, 106]. Reported overall efficiency was also very impressive reaching 5.7 and 6.8 % of transgenic live pigs per microinjected zygote for PB and SB transposon systems, respectively [65, 106] (Table 1). Moreover, most transposon integrations corresponded to monomeric integrations and very low incidence of passive incorporation of vector backbone or SB transposase vector sequences (Table 1). Interestingly, all transgenic pigs stably expressed the transgene in a promoter-dependent manner in SB transgenic animals and only one case of variegated reporter expression was observed in the PB transgenic group [106]. This can be interpreted as transposase prefers safe harbor loci for integration.

Transposition transgenesis is also compatible with SCNT [87, 107]. SCNT involves the introduction of a somatic cell into an enucleated metaphase II-arrested oocyte, followed by activation by chemical or electric stimulation, and subsequent transfer to synchronized surrogate females for development to term [108]. Since the first report of the successful cloning of sheep from cultured cells [109], SCNT has become a major method to produce transgenic livestock. Advantages of SCNT are: (1) the high rate of transgenic animals per born animals, which often reaches 100 % (but low overall efficiency), and (2) the possibility to characterize the genotype of the somatic cells before use as nuclear donor [110]. High likelihood of obtaining a transgenic animal by SCNT with known genetic makeup would reduce costs associated with producing a transgenic animal. This is particularly relevant for monotocous species with long-generation interval like cattle, in which husbandry expenditures of surrogate females negatively impact the sustainability of transgenic endeavors.

Donor cells can be transfected with linear or circular DNA transgenes for passive integration or with the components of an active system, such as SB or PB (Fig. 2) [107, 111]. Although donor cells are not considered a limiting resource for SCNT, the use of transposons is associated with significantly enhanced proportion of stably transfected cells. Delivery of pmGENIE-3, a helper independent PB transposon, to bovine primary fibroblast cells in culture caused an impressive 42-fold increase in the number of resistant cell colonies over controls [112]; similar results were reported for an established immortalized porcine cell line, as well as in primary porcine cells transfected with the SB, PB, Tol2 or Passport transposon systems [67, 68, 87, 95]. A disadvantage of the SCNT approach is that an antibiotic selection cassette is usually needed for isolation of transgenic cell clones and therefore, it is carried over into SCNT transgenic animals. This drawback, which is strongly discouraged by current regulatory guidelines, can be overcome if the antibiotic gene is supplied on a separate vector and consequently is genomically integrated independently from the GOI. Under this circumstance, the antibiotic selection cassette may be removed by segregation of the GOI from unwanted sequences by an additional round of breeding [87]. Alternatively, unwanted sequences can be removed by Cre or Flp recombinase systems as addressed in depth in the next section.

Clean and stable genomic insertion events mediated by transposases are the most striking features of these systems, making transposons first choice when gene addition for gain of function or loss of function by expression of a dominant negative allele or RNA interference is sought. The availability of different transposon systems with distinct characteristics regarding their recognition sites and/or biased genome distribution confer versatility to the system by offering the possibility of choosing a specific transposon according to the application or particular goal. For instance, PB transposase has a slight tendency to land in or close to transcriptionally active regions of the chromatin [113], so it may be more appropriate for insertional mutagenesis studies. On the other hand, SB transposase shows no predilection for transcription units, it rather prefers intergenic chromosomal regions [114, 115], which makes it the system of choice for safe delivery of transgenes. As new transposable elements are discovered and recombinant transposases with optimum enzymatic activity in the mammalian environment and improved targeting activity are developed, it is expected that transposon-based systems will gain ground in the field of large animal transgenesis.

Application of site-specific recombinases

Site-specific recombinase systems occur naturally in prokaryotes and fungi, where they perform several biological functions such as bacterial phase variation, plasmid copy number regulation, bacteriophage integration/excision from bacterial genome and amplification of yeast plasmids [116–118]. Site-specific recombinases have in common the capacity to bring together two DNA partners, catalyze double-strand cleavage at specific sites, and rejoin reciprocal strands (Fig. 3).

Fig. 3.

Recombination reactions catalyzed by Cre and Flp. The outcome of the recombination reaction is determined by the relative orientation of target sites. Inverted target sites in the same linear molecule dictate inversion of intervening DNA (a). Recombination between a target site located in a genomic address and an identical target site present in a circular DNA molecule results in insertion of circular DNA (b; thin arrow). The intramolecular recombination (thick arrow) is favored over the intermolecular reaction (thin arrow) (b). RMCE involves recombinase-mediated insertion and excision reactions, which lead to mutual exchange of the DNA between target sites (c). The inclusion of mutated target sites makes the recombination reaction irreversible

Importantly, site-specific recombination can proceed in heterologous environments, opening new avenues to engineer genomes in a predictable manner [119, 120], overcoming many problems associated with traditional methods, namely silencing or unpredictable expression of transgenes [121, 122] and unwanted remnant sequences left behind after genome manipulations.

Two basic elements comprise a site-specific recombinase system: two short consensus sequences and an enzyme that specifically recognizes those motifs and mediates strand exchange between the two DNA partner molecules [123, 124]. This process may lead to insertion, inversion, deletion or translocation of a DNA fragment in a reversible or irreversible manner [123, 125].

Members of the recombinase superfamily can be grouped according to the active amino acid present within the catalytic site in tyrosine or serine recombinases [126]. The mechanism of strand breakage, exchange and reunion markedly differs for each family (for details see [123]). The former group is further classified according to the mechanism of action in bidirectional or unidirectional [126] (Table 2).

From the tyrosine recombinase family, the Cre-loxP and Flp-FRT systems are by far the most extensively characterized members [127]. The minimum requirements for the recombination process to take place are two specific 34 base pair recognition sites and the recombinase [123]. In addition, based on the length of the recognition sites (34 bp) the probability that an identical sequence occurs by chance is extremely low (p ~ 10⁻²¹), and it is conveniently short enough as to be normally neutral toward gene expression when positioned in the genome.

Each loxP or FRT site comprises two inverted 13-bp symmetry elements, which serve as recombinase binding motif, flanking an 8-bp non-palindromic core element where strand recombination is catalyzed. The core nucleotide sequence asymmetry gives directionality to the reaction and therefore determines the type of modification [128]. Recombination of two identical target sites located on a circular and a linear DNA molecule each one will cause integration into the linear molecule. Inverted target sites in the same linear molecule dictates inversion of the DNA between recognition target sites (Fig. 3). If each identical target site is located on different linear DNA molecules, Cre or Flp recombination causes mutual exchange of sequences distal to the two recognition sites. Interestingly, it is known that recombination reaction occurs with the same efficiency regardless of the DNA topology (supercoiled or relaxed), and circular or linear molecules [128]. This versatility has positioned these members of tyrosine recombinase family at the vanguard of tools for genetic studies as well as biotechnological developments [125].

Based on the general properties of site-specific recombinase systems, different strategies for genome engineering have been developed to tailor diverse objectives, initially applied to classic model mammals and later to large domestic species. Early reports on the successful use of Flp and Cre recombinase systems for mammalian genome modifications employed recombinase-mediated DNA insertion (RMDI) [119, 120]. RMDI relies on recombination between two identical target sites, one inserted in a genomic address and the other one provided in a donor plasmid carrying the sequence to be inserted [129, 130] (Fig. 3). Upon recombination, the newly integrated DNA sequence is flanked by tandem-oriented target sites; therefore, it is prone to be excised by another round of recombination, explaining the low efficiency of this transgenic approach [119, 120]. One alternative is the use of heteromeric target sites, which recombine into inactive double mutant site and wild-type site, thus precluding another round of recombination [131–134].

Subsequently, an alternative methodology that addressed some pitfalls of RMDI, known as recombinase-mediated cassette exchange (RMCE), was introduced [135]. RMCE requires the host genome being previously tagged with compatible docking sites [132–134]. Once the appropriate tagged cell line is obtained and characterized, a site-specific recombinase mediates the exchange of the genomic-tagged cassette with the sequence of interest (Fig. 3). Recombinase may be provided as protein [136, 137], mRNA [138] or most commonly as an expression plasmid [129]. The GOI is delivered in a donor vector flanked by target sites that are homologous to the ones that were previously inserted in the genome. In summary, RMCE offers the precise engineering of animal genomes.

Inclusion of a positive selectable marker is normally required to enrich cell populations in which the desire transgenic event has occurred. However, retention of strong promoter and enhancer sequences associated with the selectable marker may have unpredictable effects on expression of linked genes [139–142], even those located at long distances from the inserted cassette [143]. From the perspective of the future introduction of transgenic animal products or derivatives into the food chain, production of selectable marker-free animals will be a mandatory condition to comply with regulatory agency guidelines raising concern regarding the possibility that antibiotic resistance genes being transferred to intestinal or environmental bacteria [144]. Therefore, deletion of selectable genes after in vitro selection of clonal cell lines is of upmost importance for both research and commercial application of transgenesis.

Removal of selectable marker genes introduced as part of the transgenic strategy can be accomplished by homologous recombination (HR), using the so-called “hit-and-run” or “tag-and-exchange” approaches [145–147]. Although successful, these methods subject the cell line to a second cycle of selection, which is not only time consuming but also may compromise the proliferative capacity of primary cell cultures. Therefore, the use of floxed or flirted selection cassettes provides the opportunity of marker deletion or replacement by site-specific recombination. Several reports have provided proof of concept for the potential of site-specific recombination technology for livestock genome engineering [65, 148–150]. Generation of marker-free cattle [14, 151] and goats [152] originated from cells that were selected and subsequently subjected to recombinase-mediated marker removal has been documented (Table 3).

Table 3.

Recombinase and integrase approaches in livestock

Enzyme	Species	Recombination site	Targeted modification	Selection	Recombinase activity	Comments	Method	Germline transmission, generation	References
Cre	Goat	LoxP	Deletion of Neo and tk	G418	Excision		SCNT	Yes	[152]
	Pig	LoxP/loxP257	Venus >mCherry	FACS	RMCE		SCNT	Yes, F1	[65]
	Pig	LoxP/LoxP?	GFP >PSEN1M146I	G418/Puromycin	RMCE		SCNT	Yes	[13]
	Cattle	LoxP	Deletion of Neo	FACS	Excision		SCNT	Preimplantation embryos	[151]
Flp	Goat	FRT	Puromycin >GFP	Puromycin/hygromycin	Gene replacement		SCNT	35-day fetus	[150]
ΦC31/Cre	Cattle	AttB-pseudo-attP/loxP	Artificial locus integration/deletion of Neo and DsRed	G418/FACS	Excision	Ambiguity	SCNT	ND	[14]
ΦC31/Cre/Dre	Cattle	AttB-pseudo-attP/loxP/rox	Deletion of Neo, tk, EGFP and plasmid backbone	G418/GCV/FACS	Excision		SCNT	Preimplantation embryos	[149]

RMCE recombinase-mediated cassette exchange, ND not determined, FACS fluorescence-activated cell sorting, GCV ganciclovir, PSEN1M146I mutated presenilin-1 gene (Alzheimer’s disease-causing gene), Neo neomycin gene, tk thymidine kinase

The potential of site-specific recombinase technology is not limited to deletion of resistance marker sequences from the manipulated genome. In mice, strategies that combine marker gene removal with complex targeted sequence modifications have been developed [125]. These include large deletions [153], non-selectable subtle mutations [154], large-scale chromosomal rearrangements (translocation, duplication, inversion, deletion, or chromosomal gain or loss) [155] and swapping gene endogenous sequences for heterologous sequences [156]. Another application of site-specific recombinases that has revolutionized mouse genetic studies is the so-called conditional gene targeting. With the conditional gene targeting methodology the specific genetic modification is triggered in a specific cell type (tissue-specific) [157] or at a particular stage of development (temporal-specific) [158, 159].

Applying the basic principles of site-specific recombination and conditional gene targeting system, a myriad of novel strategies for mouse genomic manipulation have been developed, which are revolutionizing genetic research in the post-genomic era (reviewed in [159]). Equivalent conditional methods for genetic engineering of livestock are not yet available. Albeit, some steps toward establishing conditional gene targeting methods in large animals, like generation of pigs with Cre-induced expression [160, 161] have been undertaken, the complete conditional system has not been validated in domestic species.

Further flexibility to site-specific recombination applications has come with the introduction of the large family of serine recombinases. For example, the integrase ΦC31 [162, 163] induces recombination between two different target sites known as attP (39 bp minimal size) and attB (43 bp minimum size) [162]. Upon recombination, it originates two sequence hybrid sites, attL and attR, making the reaction unidirectional [164]. Depending on the configuration, ΦC31 can induce inversion, excision or integration of DNA sequences in heterologous genomes [165, 166]. For ΦC31-based strategies, an attP-tagged genome has to be generated by random integration or HR. Once the genomic single-copy tagging is achieved, unidirectional recombination between the genomic attP site and a vector attB site is catalyzed by ΦC31 [163].

An alternative approach to accomplish chromosomal targeting with ΦC31 integrase involves recombination at cryptic endogenous genomic recognition sites, also known as pseudo-attP sites. Recombination occurs at these pseudo-sites, because of their similarity in nucleotide sequence with the wild-type attP [167]. Pseudo-attP sites have been reported to be present not only in invertebrates [168, 169], lower vertebrates [170], but also in mouse [163], human [167, 171], cattle [172, 173], sheep [174], goat [175] and pig [176] genomes. Accumulating experimental evidence indicates that these pseudo-target sites reside in genomic locations that conform to the definition of “safe harbors” [177, 178]. Further improvement to this transgenic method was achieved by the introduction of evolved and mutated ΦC31 integrases, showing enhanced sequence specificity and integration frequency at preintegrated and pseudo-attP sites [179, 180].

Although ΦC31 has been the integrase that has received most attention, new members of the large serine subfamily [181] are constantly discovered. Such novel recombinase systems include: R4 [182], TP901-1 [183, 184], and Bxb1 [181, 185]. The wide spectrum of site-specific recombinases identified so far offers a set of tools to tailor controlled and sophisticated genome modifications for basic and applied research endeavors.

Application designer nucleases

Designer nucleases, also known as programmable nucleases, are regarded as a new generation of transgenic tools characterized by being efficient, customizable and capable of precise targeted genome modifications for a broad spectrum of applications [186–191]. Programmable nucleases include ZFNs, TALENs and a RNA-guided genome modification system termed CRISPR/Cas9 [192]. The CRISPR/Cas9 has recently emerged as a powerful and facile alternative to ZFNs and TALENs for inducing targeted genetic alterations in cells and embryos [2, 20, 22]. Generically, these chimeric proteins harbor a domain (protein or RNA) that recognizes and interacts with a specific genomic sequence and an associated catalytic module that induces site-specific DNA single- or double-strand breaks (DSBs). Enzyme-catalyzed DNA cleavage in turn activates host repair mechanisms through error-prone non-homologous end joining (NHEJ) and/or homology-directed repair (HDR) [188, 189, 193], which are ultimately responsible for the targeted genome modification. Depending on the system configuration, programmable nucleases can predictably alter nucleotide sequences to achieve gene knockout, gene insertion, gene correction or point mutations at predefined endogenous loci [189–191]. Moreover, long-range chromosomal rearrangements, including deletions, inversions and translocations can be accomplished by nuclease-induced DSBs.

The introduction of SCNT opened the possibility of conventional HR-based gene targeting in somatic cells of livestock [194, 195], an approach that has been inapplicable before because of the lack of germline competent stem cells in these species [196–199]. However, the extremely low rate of HR in somatic cells (10- to 100-fold lower than that in murine ES cells) [200–202], along with the inherent inefficiency of the SCNT technique [203] made this approach cumbersome and tedious. In consequence, only few loci were knocked out by conventional HR in livestock species since the establishment of SCNT in 1997.

Designer nucleases rapidly changed the scene in livestock transgenesis, evidenced by a burst in the number of published reports since their recent introduction (summarized in Table 4). The most appealing features of programmable nucleases are that their DNA-binding domain can be engineered to target almost any predefined DNA sequence in a particular genome [12]. During NHEJ, the break ends are ligated and small base pair deletions or insertions (indels) are commonly introduced at the site of breakage. Indels in coding exons frequently result in reading frameshift mutations and inactivation of the allele.

Table 4.

Selected designer nuclease approaches in livestock and monkeys

Designer nuclease	Species	Target gene/gene KO	Deletion	Generation	Overall efficiency*	Method	Off-target effects/comments	Selection of somatic cells	References
ZFN	Rabbit	IgM	NHEJ	F0, F1	3.1 %	CPI	Mosaic F0; 14–22 % without mod.	NA	[265]
	Pig	EGFP	NHEJ, m	F0	Not given	SCNT	80 % without mod.	FACS	[209]
	Pig	PPAR-γ	NHEJ, m	F0	0.15 %	SCNT	10 % cotransfection	G418	[210]
	Pig	GGTA1	NHEJ, b	F0	1.4 %	SCNT	Resorbed fetuses and neonatal mortality	α-gal− cells counter selection	[186]
	Pig	GGTA1	NHEJ, b	F0	1.6 %	SCNT	Resorbed fetuses	α-gal− cells counter selection	[266]
	Pig	GGTA/CMAH	NHEJ, b	F0	0.8 %	SCNT	Resorbed fetuses	Clonal exp.	[267]
	Pig	CMAH	NHEJ, b	F0	2.6 %	SCNT		Clonal exp.	[268]
	Pig	CMAH	HDR, b, m	F0	0.4 %	SCNT	No integration of ZFN constructs	G418	[268]
	Pig	RELA	NHEJ, b	F0	11 %	CPI		NA	[269]
	Pig	IL2rg	NHEJ, b, m	Fetuses	2 %	SCNT	High neonatal mortality		[270]
	Cattle	BLG	NHEJ, b	F0	0.8 %	SCNT	Weak off-target effect	Clonal exp.	[211]
	Cattle	Beta-casein	Nick + HR	F0	<0.5 %	SCNT	High levels of off-target mutations	Clonal exp.	[223]
	Rabbit	APOE	NHEJ, b	F1	25 %	PNI		No selection	[271]
TALEN	Cattle	Several	NHEJ, m, b	Preimplant	50 %	CPI	High deletion/insertion	NA	[187]
	Cattle	MSTN	NHEJ	F0	15 %	CPI	Mosaicism	NA	[233]
	Sheep	MSTN	NHEJ	F0	3.8 %	CPI		NA	[233]
	Pig	RELA	NHEJ, m, b	Preimplant	29 %	CPI	High deletion/insertion	NA	[187]
	Pig	LDLR	NHEJ, m, b	F0	Not given	CT		Puromycin	[187]
	Pig	RELA	NHEJ, m, b	F0	21 %	CPI		NA	[269]
	Mini-pig	GGTA	NHEJ, b	F0	0.16 %	SCNT	High deletion	G418	[272]
	Pig,	DAZL, APC	HDR, NHEJ	F0	Not given	CT		No selection	[19]
	Monkey	MECP2	NHEJ	F0	1.2 %	CPI	High male fetal mortality, chimeric F0	NA	[273]
	Chicken	OV	NHEJ	Founder, F1	8 %	Microinjection of PGC	No off-target effects	FACS	[274]
CRISPR/Cas9	Pig	vWF	NHEJ, b, m	F0	13.2 %	CPI	Small effect on embryo development	NA	[248]
	Pig	RELA	HDR	F0	Not given	CT	Off-target cleavage	No selection	[19]
	Pig	CD163, CD1D	NHEJ	F0	1.2–2.1 %	SCNT	Very low HDR	No selection or G418	[246]
	Pig	CD163, CD1D	NHEJ	F0	4.2–3.6 %	CPI		NA	[246]
	Pig	GGTA1, CMAH, iGb3S	NHEJ	F0	3.9 %	SCNT		α-gal− cells counter selection	[245]
	Mini-pig	TYR	NHEJ, b	F0	0.9 %	SCNT	No off-target effects	G418?	[244]
	Mini-pig	PARK2, PINK1	NHEJ, b	F0	1.2 %	SCNT	No off-target effects	G418?	[244]
	Goat	Several	NHEJ, b	F0	1.1 %	SCNT	No off-target effects	No selection	[247]
	Sheep	MSTN	NHEJ	F0	5.7 %	CPI	No off-target effects	NA	[249]
	Monkey	PPAR-γ, RAG1	NHEJ	F0	Not given	CPI	No off-target effects	NA	[275]
	Rabbit	TYR	NHEJ	F0, F1	3 %	PNI	No off-target effects	No selection	[276]
	Rabbit	IL2rg; Tiki1; IL2rg + RAG1 a	NHEJ b	F0	7.5–16.6 %	CPI	Low off-target effects	No selection	[238]

PPAR-γ peroxisome proliferator-activated receptor gamma, GGTA1 α1,3-galactosyltransferase gene, CMAH cytidine monophosphate-N-acetylneuraminic acid hydroxylase, PRNP prion protein, RELA v-rel reticuloendotheliosis viral oncogene homolog A (avian), LDLR LDL receptor, IL2rg interleukin 2 receptor gamma gene, BLG beta-lactoglobulin, MECP2 methyl-CpG-binding protein, vWF Von Willebrand disease gene, MSTN myostatin gene, RAG1 recombination-activating gene 1, DAZL deleted in azoospermia-like, APC adenomatous polyposis coli, CD163 porcine reproductive and respiratory syndrome virus (PRRSV) receptor, CD1D non-classical major histocompatibility complex protein, iGb3S iGb3 synthase candidate glycosyltransferase, TYR tyrosinase gene, PARK2 parkin, RBR E3 ubiquitin protein ligase, PINK1 PTEN-induced putative kinase, OV ovalbumin gene, m mono-allelic, b bi-allelic, NA not applicable

* Transgenic offspring per treated oocytes or embryos

^aMultiplex gene targeting

The other cell pathway triggered by specific nuclease cleavage of the genome is HDR. The likelihood of a HR increases several orders of magnitude in the vicinity of a DSB [204, 205]. Therefore designer nuclease-mediated DNA scission will favor precise modification of the target sequence by HR between the endogenous sequence and the provided donor template [206, 207]. Genome editing through HR is highly versatile allowing for targeted introduction of large genetic segments to precise single-base mutations.

Such improved efficiencies associated with programmed nucleases make it feasible to target both alleles of a gene simultaneously, to perform one-step multiplex gene targeting and to omit selectable markers [19, 186]. With this emerging transgenic technology it is now possible to achieve biallelic targeting in livestock [187, 208]. Perhaps one of the major advantages of engineered nuclease-mediated gene editing for commercial purposes is that beneficial traits or mutations can be introduced in livestock genomes without inserting surplus genetic material, which is one of the concerns associated with genetically modified organisms [144].

The first designer nuclease platform to be used in livestock was the ZFN [186, 209–211], followed by TALEN [187, 190, 212] and more recently by CRISPR/Cas9 [19, 213]. ZFNs utilize zinc finger motifs tethered to a non-specific nuclease (Fok1) to bind chromosomal DNA and perform a double cut [214]. This system is comprised of an array of zinc fingers (which recognizes 12 nucleotides in the target sequence; Fig. 4). As FokI nuclease requires dimerization to cleave DNA, two ZFN monomers are necessary to interact with specific sequences in opposite DNA strands to form an active nuclease dimer. This configuration doubles the length of the recognition site, which substantially increases the specificity of ZFNs. Despite the length of the recognition site, off-target cleavages may still occur [215, 216].

Fig. 4.

Schematic representation of components of ZFNs, TALENs and CRISPR/Cas9 systems. a Structure of ZFNs. A ZFN enzyme comprise a DNA-binding domain formed by zinc finger modules (ZF), each recognizing a unique 3-base pair sequence on the target DNA, and a DNA-cleaving domain composed of FokI. Two ZFN are designed to recognize DNA sequences that flank the desired cleavage site. In the example, each ZFN comprises four ZF which recognize 12 bp on opposite strands. Upon interaction of ZF with the target site, a FokI dimer catalyzes a targeted double-strand break. b Model of TALEN system. Targeted DNA cleavage is achieved by a pair of TALEN molecules. Each TALEN is comprised by a TALE and a FokI catalytic domain. TALE-targeting domain comprise a variable number of tandem arrays of repeats of typically 34 amino acids each (shown as colored boxes in b). The RVD is responsible for the repeat specificity to associate with a particular base pair on the target DNA. c Model of CRISPR/Cas9. Cas9 nuclease is directed to a specific sequence in the genomic DNA by the first 20 nucleotides of the sgRNA, which hybridizes with the target genomic DNA, which has to be followed by a mandatory protospacer adjacent motif (PAM; 5′-NGG for Cas9 system derived from Streptococcus pyogenes). Cas9 catalyzes a DSB upstream of the PAM (red arrowheads)

The first report on ZFN-mediated gene knockout in porcine somatic cells came from the Nagashima laboratory in 2010 [217], followed by the ZFN-mediated knockout of a single-copy GFP transgene in cloned pigs [218]. These initial studies were rapidly extended by the ZFN-mediated mono-allelic disruption of an endogenous gene (PPARγ) in cloned piglets [210] and the biallelic disruption of the α1,3-galactosyltransferase (GGTA1) gene in pig fibroblasts, subsequently employed in SCNT to produce knockout pigs (Table 4; [186, 219]). In cattle, ZFN-mediated disruption of the beta-lactoglobulin (BLG) gene in bovine fibroblasts and production of cloned cows have been recently reported (Table 4, [211]). Variants of ZFNs are the zinc finger nickases (ZFNickases), which induce site-specific single-strand breaks in genomic DNA. This modified version of ZFNs can be engineered by mutating the FokI catalytic domain in one of the ZFN monomers [220–222]. ZFNickases, and other programmable nucleases with single-strain cleavage capability, are better suited for situations in which HDR-based genome edition is sought. Single-strand break stimulates resolution by HDR; therefore the faulty prone NHEJ pathway is avoided [220, 221]. A ZFNickase was used to stimulate gene addition (lysostaphin) by HDR into the endogenous beta-casein (CSN2) locus of bovine fibroblasts. Treated cells were subsequently used to generate cloned cows that produced the antimicrobial transgene product in milk [223].

Although ZFNs have been exceptionally effective for knocking out genes in farm animal genomes [186], the lack of proprietary algorithms to predict active ZFN molecules has restricted their use [190, 224].

TALENs are fusion proteins that comprise an assembled DNA-targeting domain coupled to a DNA cleavage motif. The DNA-binding domain is derived from proteins secreted by a plant pathogen belonging to the genus Xanthomonas. The DNA-binding domain is tethered to a catalytic domain of the non-specific restriction endonuclease FokI. DNA recognition and binding is mediated by tandem repeats of typically 34 amino acids, except for the last module, called half-repeat, which comprises 20 amino acids [212]. Thirty-two of the amino acids that comprise the repeat are highly conserved, whereas variable residues at positions 12 and 13, repeat variable di-residues (RVDs), dictate the binding specificity to a single nucleotide [225, 226]. Based on this code, arrays of tandem repeats can be assembled to target almost any DNA sequence of choice. Similar to ZFNs, TALEN-mediated cleavage depends on the dimerization of a pair of TALEN monomers binding to opposite DNA strands, which activates the FokI nuclease domains. Typical a TALEN monomer contains up to twenty tandem repeats, such that upon dimerization a 40-bp target sequence is recognized. Despite this theoretical high specificity, there exists evidence that TALEN can bind to degenerate sequences, and induce DNA cleavage at off-target sites [227, 228].

Several particular features of TALENs make them easier to develop and use than ZFNs [229]. Tan et al. [19] assessed the potential of TALENs (and CRISPR/Cas9 nucleases, see below) to edit the genome of many commercially important species. The authors demonstrated that TALENs can efficiently target a variety of alleles involved in food production, reproductive efficiency and external traits (hornlessness) [230] in the genomes from different livestock species. In the same study, TALENs were used to induce NHEJ- or HDR-directed edits at specific loci to mimic mutations that are known to be associated with genetic diseases in humans. From the edited cells, live pigs carrying the induced mutations were generated by chromatin transfer (CT) and these animals promise to become valuable large mammalian models in translational medicine. The targeting efficiency ranged from approximately 10 % for single-nucleotide polymorphisms to >50 % for some larger alterations. According to recent data, the TALEN system is also functional and efficient in the preimplantation embryo context, since microinjection of TALEN mRNA directed to the GDF-8 gene [231, 232] into bovine and ovine zygotes resulted in correctly edited cattle and sheep [233]. In light of these encouraging findings, it is conceivable that genome edition by designer nucleases will become a practical strategy to introduce or suppress genetic characteristics in livestock populations to accelerate the genetic progress in harmony with classic breeding strategies.

The RNA-guided CRISPR/Cas9 system [234, 235] was recently discovered in bacteria and archaea, in which the RNA-guided foreign-DNA cleavage process provides adaptive immunity against invading phages or plasmids [236, 237]. The CRISPR/Cas9 sequence specificity is determined by Watson–Crick base complementarity with a single-guide RNA (sgRNA). The induced DNA damage is repaired by either HR or error-prone NHEJ that normally causes indels at the cleavage site. Design and generation of the synthetic sgRNA is markedly easier compared with the cumbersome protein engineering required to produce ZFNs and TALENs. By changing the nucleotide sequence of the sgRNA it is possible to target almost any site in the genome. The activity of the CRISPR/Cas9 system allows the implementation of high-throughput methodologies and multiplex editing of genomic loci in preimplantation mammalian embryos [238].

From 2013, the CRISPR/Cas9 system bursts into the genome engineering scenario through several independent reports providing encouraging evidence for simplicity and effectiveness of this system to engineer large animal genomes [2, 239–243]. Tan et al. [19] were the first researchers to apply CRISPR/Cas9 technology to target endogenous genes (P65 and APC) in primary pig cells in culture. Despite the demonstration that CRSPR/Cas9 works in livestock cells, recovery of CRISPR mutant cell clones was much lower than that with TALENs, suggesting that CRISPR/Cas9 system needs further optimization to achieve targeting efficiencies comparable to TALENs. Follow-up studies have demonstrated that CRISPR/Cas9 genome-edited cells can support development to term when used as nuclear donor in SCNT in pigs [244–246] and goats [247]. An appealing and straightforward alternative to SCNT to generate genome-edited animals is the injection of the CRISPR/Cas9 components in livestock one-cell embryos produced in vivo or in vitro. Using the CPI approach, Hai et al. [248] managed to obtain live pigs with mono- and biallelic mutations in the vWF gene to generate a relevant large animal model for hemophilia. The reported efficiency is quite impressive for a zygote microinjection-based method; with 10 out of 16 born piglets (~63 %) carrying one or both vWF alleles mutated. Another study with sheep zygotes microinjected with sgRNA/Cas9 mRNA [249] produced modest results in terms of the number of mutated offspring to born animals (2/32); however, no off-target mutations were detected. A recurrent problem of zygote microinjections is the high incidence of mosaic animals [227, 246, 249, 250], which is believed to originate when the nuclease remains active beyond the first embryo cleavage. Another potential problem associated with the use CRISPR/Cas9 editing system is the introduction of undesired mutations at off-target genomic sites [251, 252]. Based on the relatively short CRISPR/Cas9 recognition site (20 nt) and the known mismatch tolerance, especially at the 5′ region of the target sequence, occurrence of off-target mutations should not be disregarded [253]. A strategy to minimize off-target activity is to use a mutated version of Cas9 (D10A mutation [192, 254]) with nickase activity. DNA single-strand break stimulates HDR with negligible NHEJ-mediated mutations. Moreover, when required, a DSB can be simultaneously induced at the target site using a pair of appropriately spaced and oriented sgRNAs along with Cas9 nickase, that enhances genome editing specificity [255]. More research is warranted to ascertain if off-target DNA cleavage rates induced by CRISPR/Cas9 are a concern in the context of genome edition in large animals.

Conclusions

Transgenic methodologies are constantly evolving, providing researchers and biotechnologists with advanced tools for efficient and controlled genome modifications. Initial transgenic interventions in livestock were confined to simple gene insertion at random places in the genome. Thanks to constant advances in the area of genetic engineering, today it is possible to achieve precise genome modifications by inserting, replacing or removing predefined DNA sequences. In this regard, introduction of methodologies that enable enzymatic manipulation of animal genomes have opened new possibilities to create genetically modified animals for agriculture or biomedicine. Although the transgenic toolbox for large animals is currently equipped with powerful methodologies there are many aspects to improve in the associated reproductive technologies required to generate a transgenic animal. Low success rates of SCNT and zygote microinjection, two of the most commonly used methods to generate transgenic large animals, still represent a bottleneck. Efficiency of SCNT has remained low in spite of considerable efforts invested in developing more successful protocols, while embryo microinjection has been invariably associated with undesired chimerism. Thus, further improvements in surpassing the limitations of these techniques may impact favorably on the overall efficiency of transgenic methods.

Transposon-based systems are a straightforward alternative to achieve transgene integrations with persistent transgene expression and germline transmission. These characteristics along with increased transgenic efficiencies will certainly reduce costs and contribute to animal welfare by reducing the number of animal required to produce the desired genotype and by avoiding unwanted phenotypes. Transposon-based methods alone or combined with site-directed recombinases will simplify the production of marker-free animals to comply with regulatory guidelines for animal transgenesis.

The ground-breaking feature of designer nucleases is that they brought the possibility of purposely directing the genomic modification to a specific and unique chromosomal locus. Among the members of the engineered nucleases, RNA-guided nucleases are the ones that promise to change the paradigm of genome editing in large animals. The CRISPR/Cas9 system combines facile design and construction with high specificity, effectivity and real possibility of multiplex gene edition. However, there is still room for improvement in particular areas like minimizing off-target effects of designer nucleases, enhancement of nuclease activity, and development of methods to enrich cell population with targeted genome edits. Another avenue to improve engineered nuclease-based methods is through the genetic or pharmacologic manipulation of the DSB repair pathway. For instance, for many applications, enhancement of low-frequency HDR over the NHEJ would be convenient.

The launching of high-throughput genome sequencing at accessible prices will make it possible to improve the quality of current genome data in farm animals and it will become a valuable tool to verify transgenic lines at genome scale. It is anticipated that new generation transgenic tools in concert with updated genomic data will facilitate the production of large animal models for translational medicine. These large animal models will be instrumental for understanding disease pathogenesis and development of better therapeutic approaches of severe human pathologic conditions.

It is foreseen that similar opportunities will arise in agricultural applications of transgenic livestock. Genome sequencing and phenotyping will provide unprecedented opportunities for the identification of molecular markers that affect livestock performance, which can be readily addressed and manipulated at will by site-directed nucleases to improve productive traits. Experimental evidence has provided proof of principle that non-meiotic introgression of natural or novel genetic variants in livestock genomes is attainable using designer nucleases. Numerous reports cited in this review strongly indicate that designer nucleases have earned enough merit as genome engineering tools as to be considered in the near future in selection programs to advance genetic improvement when selective breeding is impracticable or inefficient. Importantly, the toolbox for genome engineering is still expanding, as new enzymatic systems are constantly discovered. One recent example are the bacterial casposons, which seem to combine the features of CRISPR/Cas and transposons [256, 257], suggesting that more sophisticated options for genome engineering will become feasible in the near feature.

Abbreviations

Cas9

CRISPR-associated protein 9

CPI

Cytoplasmic injection

Ct

Chromatin transfer

CRISPR

Clustered regularly interspaced short palindromic repeats

DSB

Double-strand break

GOI

Gene of interest

HR

Homologous recombination

HDR

Homology-directed repair

ICSI

Intracytoplasmic sperm injection

ICSI-Tr

Intracytoplasmic sperm injection-mediated transgenesis

iPS

Induced pluripotent stem (cell)

I-SceI

Homing endonuclease

ITR

Inverted terminal repeat

KO

Knockout

NHEJ

Non-homologous end joining

PB

piggyBac transposon system

PNI

Pronuclear injection

RE

Restriction enzyme

RecA

Recombinase A

REMI

Restriction enzyme-mediated integration

RMCE

Recombinase-mediated cassette exchange

RMDI

Recombinase-mediated DNA insertion

SB

Sleeping Beauty transposon system

SCNT

Somatic cell nuclear transfer

sgRNA

Single-guide RNA

SMGT

Sperm-mediated gene transfer

SV40

Simian virus 40

TALEN

Transcription activator-like element nuclease

Tol2

Tol2 transposon system

ZFN

Zinc finger nuclease