Syrian hamsters (Mesocricetus auratus) have long served as valuable model organisms in diverse research fields such as oncology, immunology, and physiology owing to their unique biological and pathological characteristics. Although embryo manipulation techniques such as embryo collection, pronuclear microinjection, and embryo transfer have been established, gene knock-in (KI) hamsters have not yet been reported. Here, we report the successful generation of gene KI Syrian hamsters by microinjecting CRISPR/Cas9 components and plasmid DNA into pronuclear-stage zygotes. Targeted insertion of a DNA cassette up to 8 kb was achieved at the ROSA26 orthologous locus and other genomic sites. Importantly, we confirmed functional expression of a reporter cassette inserted at the ROSA26 site, providing evidence of transcriptional activity at this locus in Syrian hamsters. Furthermore, we demonstrated that frozen-thawed KI embryos could give rise to live offspring using a simplified freezing and thawing protocol originally developed for mice and rats. These results confirm the feasibility and applicability of advanced genome editing technologies in Syrian hamsters. These technological advancements enable the development of versatile KI models for applications such as gene expression monitoring and conditional mutagenesis, thereby expanding the utility of Syrian hamsters as model organisms, comparable to mice and rats.

Cyp19a1-Cre-EGFP: A placenta-specific Cre transgenic mouse model for targeted gene recombination in trophoblast cells.

The phenomenal progress in biotechnology and genomics is both inspiring and overwhelming-a classic curse of choice, particularly when it comes to selecting methods for mapping transgene DNA integration sites. Transgene localization remains a crucial task for the validation of transgenic mouse or other animal models generated by pronuclear microinjection. Due to the inherently random nature of DNA integration, reliable characterization of the insertion site is essential. Over the years, a vast number of mapping methods have been developed, and new approaches continue to emerge, making the choice of the most suitable technique increasingly complex. Factors such as cost, required reagents, and the nature of the generated data require careful consideration. In this review, we provide a structured overview of current transgene mapping techniques, which we have broadly classified into three categories: classic PCR-based methods (such as inverse PCR and TAIL-PCR), next-generation sequencing with target enrichment, and long-read sequencing platforms (PacBio and Oxford Nanopore). To aid in decision-making, we include a comparative table summarizing approximate costs for the methods. While each approach has its own advantages and limitations, we highlight our top four recommended methods, which we believe offer the best balance of cost-effectiveness, reliability, and simplicity for identifying transgene integration sites.

The genetic modification of rats is a key technology for advancing biomedical research on human diseases. CRISPR/Cas9-mediated genome editing enables the generation of knockout rats in a single step, without the need for embryonic stem cells, by directly injecting genome editing components into zygotes. This simplifies the process, reduces costs, and accelerates gene function analysis in rats. However, the insertion of a gene cassette into a target site has remained inefficient, limiting the generation of knockin (KI) rats. To overcome this issue, we developed an optimized method that covers the entire process from zygote harvesting with superovulation to timed microinjection, ensuring the consistent generation of KI rats. We successfully generated four different fluorescent reporter lines at the ROSA26 locus in rats. Our study provides detailed, step-by-step protocols for donor vector design, zygote collection, microinjection, founder screening, and cryopreservation in rats.

In pronuclear microinjection, the Cas9 endonuclease is employed to introduce in vivo DNA double-strand breaks at the genomic target locus or within the donor vector, thereby enhancing transgene integration. The manner by which Cas9 interacts with DNA repair factors during transgene end processing and integration is a topic of considerable interest and debate. In a previous study, we developed a barcode-based genetic system for the analysis of transgene recombination following pronuclear microinjection in mice. In this approach, the plasmid library is linearized with a restriction enzyme or a Cas9 RNP complex at the site between a pair of barcodes. A pool of barcoded molecules is injected into the pronucleus, resulting in the generation of multicopy concatemers. In the present report, we compared the effects of in vivo Cas9 cleavage (RNP+ experiment) and in vitro production of Cas9- linearized transgenes (RNP- experiment) on concatenation. In the RNP+ experiment, two transgenic single-copy embryos were identified. In the RNP- experiment, six positive embryos were identified, four of which exhibited lowcopy concatemers. Next-generation sequencing (NGS) analysis of the barcodes revealed that 53 % of the barcoded ends had switched their initial library pairs, indicating the involvement of the homologous recombination pathway. Out of the 20 transgene-transgene junctions examined, 11 exhibited no mutations and were presumably generated through re-ligation of Cas9-induced blunt ends. The majority of mutated junctions harbored asymmetrical deletions of 2-4 nucleotides, which were attributed to Cas9 end trimming. These findings suggest that Cas9-bound DNA may present obstacles to concatenation. Conversely, clean DNA ends were observed to be joined in a manner similar to restriction-digested ends, albeit with distinctive asymmetry. Future experiments utilizing in vivo CRISPR/ Cas cleavage will facilitate a deeper understanding of how CRISPR-endonucleases influence DNA repair processes.

Mitochondria play a crucial role in sperm development; however, the mechanisms regulating their function in sperm remain poorly understood. Developing a method to regulate the expression of a target gene within the mitochondria of sperm is a vital step in this area of research. In this study, we aimed to create a system for expressing a transgene in the mitochondria of sperm. As a proof of concept, we generated transgenic mice that express green fluorescent protein (GFP) fused with a mitochondrial localization signal (MLS) driven by the phosphoglycerate kinase 2 (PGK2) promoter, which facilitates the transgene expression in the sperm. Although the PGK2 promoter has previously shown to drive gene expression in spermatocytes and spermatids, the novelty of our approach lies in the combination of PGK2-driven MLS-GFP expression to study mitochondria in vivo. We established two founder lines of transgenic mice through pronuclear microinjection, and MLS-GFP expression was confirmed in the mitochondria of sperm cells using fluorescence microscopy and flow cytometry. Consequently, we provide a novel platform for investigating mitochondrial function in sperm, where GFP can be substituted with other genes of interest to examine their effects on mitochondria. This system specifically targets sperm mitochondria, offering an innovative approach for studying mitochondrial function in vivo.

Gene-engineered animals created using gene-targeting technology have long been recognized as beneficial, valid, and valuable tools for exploring the function of a gene of interest, at least in early 2013. This approach, however, suffers from laborious and time-consuming tasks, such as the production of successfully targeted embryonic stem (ES) cells, their characterization, production of chimeric blastocysts carrying these gene-modified ES cells, and transplantation of those manipulated blastocysts to the recipient (pseudopregnant) females to deliver chimeric mice. Since the appearance of genome editing technology, which is now exemplified by the CRISPR/<italic>Cas9</italic> system, in late 2013, significant advances have been made in the generation of genome-edited animals through pronuclear microinjection (MI) of genome-editing components into fertilized eggs (zygotes) or electroporation (EP) of zygotes in the presence of these reagents. However, these procedures require the transfer of genome-edited embryos into the reproductive tracts of recipient females for further development. <underline>G</underline>enome editing via <underline>o</underline>viductal <underline>n</underline>ucleic <underline>a</underline>cids <underline>d</underline>elivery (GONAD) and its modified version, called "improved GONAD (<italic>i</italic>-GONAD)," were developed as an alternative to the MI- or EP-based genome-edited animal production and now recognized to be very convenient and straightforward as genome editing can only be performed <italic>in</italic> <italic>vivo</italic> (within the oviductal lumen where fertilized embryos exist). This system also enables the simultaneous transfection of epithelial cells <italic>lining the oviductal lumen</italic>. In this review, we summarize the recent advances in GONAD/<italic>i</italic>-GONAD and their derivatives and discuss the potential of these technologies to study various biological systems related to female reproduction.

To complete microinjection as quickly as possible, we have developed Vibratory Microinjection Systems (VMSs) that vibrate a micropipette in its longitudinal direction and can significantly reduce the time needed for pronuclear microinjection compared to ordinary (non-vibratory) microinjection. The longest breakdown of the time is the time required to pierce the cell membrane and the pronuclear membrane simultaneously. Because cytoplasmic microinjection, which pierces the cell membrane alone, is far more difficult and time-consuming than pronuclear microinjection, we next aimed to develop a VMS capable of penetrating the cell membrane instantly. In this new and latest version, two types of ultrasonic-wave vibrators were developed: the first for commercially available micropipettes (Femtotip) and the second for self-made micropipettes. The two vibrators differ only in their airtight structure, where the micropipettes connect to their respective vibrators: a female screw plus O-ring for the first vibrator (VMS6_1) and a silicone-rubber tube for the second (VMS6_2). The tube-type joint used in VMS6_2 only slightly damped or amplified vibrations from the vibrator to the micropipette tip, propagating them much more accurately than the screw-type joint in VMS6_1. In addition, VMS6_2 significantly shortened the time taken to pierce the cell membrane of a fertilized egg: an average of 1.52 s (N = 410) vs. 3.62 s (N = 65) in VMS6_1. The VMS6_2 group achieved a piercing time of zero in 86.1% of the allocated eggs, while only 10.8% of the VMS6_1 group did. In each vibrator, we also compared vibratory microinjection (VM; N = 475) and ordinary microinjection (OM; N = 457), which uses injection pressure in place of vibration. None of the eggs in the OM group achieved the zero-second piercing time. Compared to the OM, the VM group showed a significantly shorter piercing time, 1.80 vs. 10.69 s on average, and a significantly better survival rate, 90.3 vs. 81.8% on average. VMS6_2 not only improved on the already demonstrated superiority of VM to OM but also enabled instantaneous piercing of the cell membrane.

The first genetically modified large animals were developed in 1985 by microinjection to increase the growth of agricultural livestock such as pigs. Since then, it has been a difficult trail due to the lack of genetic tools. Although methods and technologies were developed quickly for the main experimental mammal, the mouse, e.g., efficient pronuclear microinjection, gene targeting in embryonic stem cells, and omics data, most of it was-and in part still is-lacking when it comes to livestock. Over the next few decades, progress in genetic engineering of large animals was driven less by research for agriculture but more for biomedical applications, such as the production of pharmaceutical proteins in the milk of sheep, goats, or cows, xeno-organ transplantation, and modeling human diseases. Available technologies determined if a desired animal model could be realized, and efficiencies were generally low. Presented here is a short review of how genome editing tools, specifically CRISPR/Cas, have impacted the large animal field in recent years. Although there will be a focus on genome engineering of pigs for biomedical applications, the general principles and experimental approaches also apply to other livestock species or applications.

β-catenin is a conserved molecule that plays an important role in hair follicle development. In this study, we generated skin-specific overexpression of ovine β-catenin in transgenic mice by pronuclear microinjection. Results of polymerase chain reaction (PCR) testing and Southern blot showed that the ovine β-catenin gene was successfully transferred into mice, and the exogenous β-catenin gene was passed down from the first to sixth generations. Furthermore, real-time fluorescent quantitative PCR (qRT-PCR) and western blot analysis showed that β-catenin mRNA was specifically expressed in the skin of transgenic mice. The analysis of F6 phenotypes showed that overexpression of β-catenin could increase hair follicle density by prematurely promoting the catagen-to-anagen transition. The results showed that ovine β-catenin could also promote hair follicle development in mice. We, therefore, demonstrate domestication traits in animals.

In the CRISPR/Cas9-mediated gene cassette knockin (KI) strategy, a gene cassette is integrated into a target locus through a proper DNA repair pathway after the Cas9-induced double-strand DNA breaks; the activation of the DNA repair pathway is known to be correlated with the cell cycle. Recently, we have reported a new KI approach named SPRINT (S-phase pronuclear injection for targeting)-CRISPR, focusing on the correlation between the cell cycle and the KI efficiency in the mouse zygote microinjection. Our results suggest that the CRISPR-mediated KI with a homologous recombination-based donor vector during S-phase enhances the KI efficiency. For SPRINT-CRISPR, the uniformity of the zygotes in the cell cycle is achieved by in vitro fertilization, and the zygotes are cryopreserved until use. These reproductive techniques are necessary for efficient KI. Furthermore, Piezo-assisted microinjection has been successful in improving the survival rate of the injected embryos. In this chapter, we describe the protocols that focus on the zygote preparation and Piezo-assisted microinjection of the SPRINT-CRISPR method.

Abstract Under the background of food security, using non‐grain feed instead of corn–soybean‐based feed is an effective measure to alleviate the food‐feed competition. While, non‐grain feeds are often rich in fiber, which cannot be digested by non‐ruminants. Producing heterologous enzymes in non‐ruminants to improve cellulose utilization rate is a new research strategy by transgenic technology. In this study, porcine transthyretin (TTR) promoter, signal peptide‐coding sequence (CDS), Saccharomycopsis fibuligera β‐glucosidase gene (BGL1)‐CDS, 6×His sequences fragments were fused into pGL3‐control vector to generate transgenic vector. Then, transgenic mice were generated by pronuclear microinjection of the linearized expression vectors. Transgenic mice and their offspring were examined by PCR‐based genotyping and copy number variation. Results showed that BGL1 was successfully integrated into the mouse genome and transmitted stably. Furthermore, reverse transcription‐polymerase chain reaction (RT‐PCR), Western blotting, and β‐glucosidase activity assay demonstrated that BGL1 was specifically expressed in the liver, and β‐glucosidase activity significantly increased. In addition, liver weight index, cellular morphology, and collagen fiber content of the liver showed that exogenous gene insertion did not cause any lesions to live. Taken together, our findings suggest that β‐glucosidase driven by TTR promoter was specifically expressed in the liver of transgenic mice.

One-step generation of a conditional allele in mice using a short artificial intron

Myostatin (Mstn) is a major negative regulator of skeletal muscle mass and initiates multiple metabolic changes. The deletion of the Mstn gene in mice leads to reduced mitochondrial functions. However, the underlying regulatory mechanisms remain unclear. In this study, we used CRISPR/Cas9 to generate myostatin-knockout (Mstn-KO) mice via pronuclear microinjection. Mstn-KO mice exhibited significantly larger skeletal muscles. Meanwhile, Mstn knockout regulated the organ weights of mice. Moreover, we found that Mstn knockout reduced the basal metabolic rate, muscle adenosine triphosphate (ATP) synthesis, activities of mitochondrial respiration chain complexes, tricarboxylic acid cycle (TCA) cycle, and thermogenesis. Mechanistically, expressions of silent information regulator 1 (SIRT1) and phosphorylated adenosine monophosphate-activated protein kinase (pAMPK) were down-regulated, while peroxisome proliferator-activated receptor γ coactivator-1α (PGC-1α) acetylation modification increased in the Mstn-KO mice. Skeletal muscle cells from Mstn-KO and WT were treated with AMPK activator 5-aminoimidazole-4-carboxamide riboside (AICAR), and the AMPK inhibitor Compound C, respectively. Compared with the wild-type (WT) group, Compound C treatment further down-regulated the expression or activity of pAMPK, SIRT1, citrate synthase (CS), isocitrate dehydrogenase (ICDHm), and α-ketoglutarate acid dehydrogenase (α-KGDH) in Mstn-KO mice, while Mstn knockout inhibited the AICAR activation effect. Therefore, Mstn knockout affects mitochondrial function by inhibiting the AMPK/SIRT1/PGC1α signaling pathway. The present study reveals a new mechanism for Mstn knockout in regulating energy homeostasis.

In this work, we set out to create mice susceptible to the SARS-CoV-2 coronavirus. To ensure the ubiquitous expression of the human ACE2 gene we used the human EF1a promoter. Using pronuclear microinjection of the transgene construct, we obtained six founders with the insertion of the EF1a-hACE2 transgene, from which four independent mouse lines were established. Unfortunately, only one line had low levels of hACE2 expression in some organs. In addition, we did not detect the hACE2 protein in primary lung fibroblasts from any of the transgenic lines. Bisulfite sequencing analysis revealed that the EF1a promoter was hypermethylated in the genomes of transgenic animals. Extensive analysis of published works about transgenic animals indicated that EF1a transgenic constructs are frequently inactive. Thus, our case cautions against using the EF1a promoter to generate transgenic animals, as it is prone to epigenetic silencing.Supplementary InformationThe online version contains supplementary material available at 10.1007/s11248-022-00319-5.

Transgenic mice encoding modern imaging probes: Properties and applications.

Embryonic morphogenesis is strictly dependent on tight spatiotemporal control of developmental gene expression, which is typically achieved through the concerted activity of multiple enhancers driving cell type-specific expression of a target gene. Mammalian genomes are organized in topologically associated domains, providing a preferred environment and framework for interactions between transcriptional enhancers and gene promoters. While epigenomic profiling and three-dimensional chromatin conformation capture have significantly increased the accuracy of identifying enhancers, assessment of subregional enhancer activities via transgenic reporter assays in mice remains the gold standard for assigning enhancer activity in vivo. Once this activity is defined, the ideal method to explore the functional necessity of a transcriptional enhancer and its contribution to target gene dosage and morphological or physiological processes is deletion of the enhancer sequence from the mouse genome. Here we present detailed protocols for efficient introduction of enhancer-reporter transgenes and CRISPR-mediated genomic deletions into the mouse genome, including a step-by-step guide for pronuclear microinjection of fertilized mouse eggs. We provide instructions for the assembly and genomic integration of enhancer-reporter cassettes that have been used for validation of thousands of putative enhancer sequences accessible through the VISTA enhancer browser, including a recently published method for robust site-directed transgenesis at the H11 safe-harbor locus. Together, these methods enable rapid and large-scale assessment of enhancer activities and sequence variants in mice, which is essential to understand mammalian genome function and genetic diseases.

Genetically modified mice are commonly used in biologic, medical, and drug discovery research, but conventional microinjection methods used for genetic modification require extensive training and practical experience. Here we present a fully automated system for microinjection into the pronucleus to facilitate genetic modification. We first developed software that automatically controls the microinjection system hardware. The software permits automatic rotation of the zygote to move the pronucleus to the injection pipette insertion position. We also developed software that recognizes the pronucleus in 3-dimensional coordinates so that the injection pipette can be automatically inserted into the pronucleus, and achieved a 94% insertion rate by linking the 2 pieces of software. Next, we determined the optimal solution injection conditions (30 hPa, 0.8–2.0 s) by examining the survival rate of injected zygotes. Finally, we produced transgenic (traditional DNA injection and piggyBac Transposon system) and knock-in (genomic editing) mice using our newly developed Integrated Automated Embryo Manipulation System (IAEMS). We propose that the IAEMS will simplify highly reproducible pronuclear stage zygote microinjection procedures.

To study the pathophysiology of human cardiac diseases and to develop novel treatment strategies, complex interactions of cardiac cells on cellular, tissue and on level of the whole heart need to be considered. As in vitro cell-based models do not depict the complexity of the human heart, animal models are used to obtain insights that can be translated to human diseases. Mice are the most commonly used animals in cardiac research. However, differences in electrophysiological and mechanical cardiac function and a different composition of electrical and contractile proteins limit the transferability of the knowledge gained. Moreover, the small heart size and fast heart rate are major disadvantages. In contrast to rodents, electrophysiological, mechanical and structural cardiac characteristics of rabbits resemble the human heart more closely, making them particularly suitable as an animal model for cardiac disease research. In this review, various methodological approaches for the generation of transgenic rabbits for cardiac disease research, such as pronuclear microinjection, the sleeping beauty transposon system and novel genome-editing methods (ZFN and CRISPR/Cas9)will be discussed. In the second section, we will introduce the different currently available transgenic rabbit models for monogenic cardiac diseases (such as long QT syndrome, short-QT syndrome and hypertrophic cardiomyopathy) in detail, especially in regard to their utility to increase the understanding of pathophysiological disease mechanisms and novel treatment options. LINKED ARTICLES: This article is part of a themed issue on Preclinical Models for Cardiovascular disease research (BJP 75th Anniversary). To view the other articles in this section visit http://onlinelibrary.wiley.com/doi/10.1111/bph.v179.5/issuetoc.

Genetic engineering (GE) of livestock initially has been accomplished primarily using pronuclear microinjection into zygotes (1985-1996). The applications of the technology were limited due to low integration efficiency, aberrant transgene expression resulting from random integration and the presence of genetic mosaicism in transgenic founder animals. Despite enormous efforts to established embryonic stem cells (ESCs) for domestic species, the ESC GE technology does not exist for livestock. Development of somatic cell nuclear transfer (SCNT) has bypassed the need in livestock ESCs and revolutionized the field of livestock transgenesis by offering the first cell-based platform for precise genetic manipulation in farm animals. For nearly two decades since the birth of Dolly (1996-2013), SCNT was the only method used for the generation of knockout and knockin livestock. Arrival of CRISPRS/Cas9 system, a new generation of gene-editing technology, gave us an ability to introduce precise genome modifications easily and efficiently. This technological advancement accelerated production of GE livestock by SCNT and reinstated zygote micromanipulation as an important GE approach. The primary advantage of the SCNT technology is the ability to confirm in vitro that the desired genetic modification is present in the somatic cells prior to animal production. The edited cells could also be tested for potential off-target mutations. Additionally, this method eliminates the risk of genetic mosaicism frequently observed following zygote micromanipulation. Despite its low efficiency, SCNT is a well-established procedure in numerous laboratories around the world and will continue to play an important role in the GE livestock field.