Genome editing tools such as the clustered regularly interspaced short palindromic repeat (CRISPR)-associated system (Cas) have been widely used to modify genes in model systems including animal zygotes and human cells, and hold tremendous promise for both basic research and clinical applications. To date, a serious knowledge gap remains in our understanding of DNA repair mechanisms in human early embryos, and in the efficiency and potential off-target effects of using technologies such as CRISPR/Cas9 in human pre-implantation embryos. In this report, we used tripronuclear (3PN) zygotes to further investigate CRISPR/Cas9-mediated gene editing in human cells. We found that CRISPR/Cas9 could effectively cleave the endogenous β-globin gene (HBB). However, the efficiency of homologous recombination directed repair (HDR) of HBB was low and the edited embryos were mosaic. Off-target cleavage was also apparent in these 3PN zygotes as revealed by the T7E1 assay and whole-exome sequencing. Furthermore, the endogenous delta-globin gene (HBD), which is homologous to HBB, competed with exogenous donor oligos to act as the repair template, leading to untoward mutations. Our data also indicated that repair of the HBB locus in these embryos occurred preferentially through the non-crossover HDR pathway. Taken together, our work highlights the pressing need to further improve the fidelity and specificity of the CRISPR/Cas9 platform, a prerequisite for any clinical applications of CRSIPR/Cas9-mediated editing.Electronic supplementary materialThe online version of this article (doi:10.1007/s13238-015-0153-5) contains supplementary material, which is available to authorized users.

β-Thalassemia is a global health issue, caused by mutations in the HBB gene. Among these mutations, HBB −28 (A>G) mutations is one of the three most common mutations in China and Southeast Asia patients with β-thalassemia. Correcting this mutation in human embryos may prevent the disease being passed onto future generations and cure anemia. Here we report the first study using base editor (BE) system to correct disease mutant in human embryos. Firstly, we produced a 293T cell line with an exogenous HBB −28 (A>G) mutant fragment for gRNAs and targeting efficiency evaluation. Then we collected primary skin fibroblast cells from a β-thalassemia patient with HBB −28 (A>G) homozygous mutation. Data showed that base editor could precisely correct HBB −28 (A>G) mutation in the patient’s primary cells. To model homozygous mutation disease embryos, we constructed nuclear transfer embryos by fusing the lymphocyte or skin fibroblast cells with enucleated in vitro matured (IVM) oocytes. Notably, the gene correction efficiency was over 23.0% in these embryos by base editor. Although these embryos were still mosaic, the percentage of repaired blastomeres was over 20.0%. In addition, we found that base editor variants, with narrowed deamination window, could promote G-to-A conversion at HBB −28 site precisely in human embryos. Collectively, this study demonstrated the feasibility of curing genetic disease in human somatic cells and embryos by base editor system.Electronic supplementary materialThe online version of this article (doi:10.1007/s13238-017-0475-6) contains supplementary material, which is available to authorized users.

Targeted point mutagenesis through homologous recombination has been widely used in genetic studies and holds considerable promise for repairing disease-causing mutations in patients. However, problems such as mosaicism and low mutagenesis efficiency continue to pose challenges to clinical application of such approaches. Recently, a base editor (BE) system built on cytidine (C) deaminase and CRISPR/Cas9 technology was developed as an alternative method for targeted point mutagenesis in plant, yeast, and human cells. Base editors convert C in the deamination window to thymidine (T) efficiently, however, it remains unclear whether targeted base editing in mouse embryos is feasible. In this report, we generated a modified high-fidelity version of base editor 2 (HF2-BE2), and investigated its base editing efficacy in mouse embryos. We found that HF2-BE2 could convert C to T efficiently, with up to 100% biallelic mutation efficiency in mouse embryos. Unlike BE3, HF2-BE2 could convert C to T on both the target and non-target strand, expanding the editing scope of base editors. Surprisingly, we found HF2-BE2 could also deaminate C that was proximal to the gRNA-binding region. Taken together, our work demonstrates the feasibility of generating point mutations in mouse by base editing, and underscores the need to carefully optimize base editing systems in order to eliminate proximal-site deamination.Electronic supplementary materialThe online version of this article (doi:10.1007/s13238-017-0418-2) contains supplementary material, which is available to authorized users.

To develop a noninvasive medium-based preimplantation genetic diagnosis (PGD) test for α-thalassemias-SEA.The embryos of α-thalassemia-SEA carriers undergoing in vitro fertilization (IVF) were cultured. Single cells were biopsied from blastomeres and subjected to fluorescent gap polymerase chain reaction (PCR) analysis; the spent culture media that contained embryo genomic DNA and corresponding blastocysts as verification were subjected to quantitative-PCR (Q-PCR) detection of α-thalassemia-SEA. The diagnosis efficiency and allele dropout (ADO) ratio were calculated, and the cell-free DNA concentration was quantitatively assessed in the culture medium.The diagnosis efficiency of medium-based α-thalassemias–SEA detection significantly increased compared with that of biopsy-based fluorescent gap PCR analysis (88.6% vs 82.1%, P < 0.05). There is no significant difference regarding ADO ratio between them. The optimal time for medium-based α-thalassemias–SEA detection is Day 5 (D5) following IVF.Medium-based α-thalassemias–SEA detection could represent a novel, quick, and noninvasive approach for carriers to undergo IVF and PGD.

Purpose To identify differentially expressed microRNAs (miRNAs) and expression patterns of specific miRNAs during meiosis in human oocytes. Materials and methods To identify differentially expressed miRNAs, GV oocytes and MII oocytes matured at conventional FSH levels (5.5 ng/ml) were analyzed by miRNA microarray. Real-time RT-PCR was used to confirm the changed miRNAs. To validate the dynamic changes of miRNAs from GV to MII stages, oocytes were divided into four groups (#1-4), corresponding to GVoocytes, MI oocytes, MII oocytes matured in conventional FSH level and MII oocytes matured in high FSH level (2,000 ng/ml) respectively. Results Compared with GVoocytes, MII oocytes exhibited up-regulation of 4 miRNAs (hsa-miR-193a-5p, hsa-miR-297, hsa-miR-625 and hsa-miR-602), and down-regulation of 11 miRNAs (hsa-miR-888*, hsa-miR-212, hsa-miR-662, hsa-miR-299-5p, hsa-miR-339-5p, hsa-miR-20a, hsa-miR-486-5p, hsa-miR-141*, hsa-miR-768-5p, hsa-miR-376a and hsa-miR-15a). RT-PCR analysis of hsa-miR-15a and hsamiR-20a expression revealed concordant dynamic changes in oocytes from group 1 to group 4. Conclusion(s) Specific miRNAs in human oocytes had dynamic changes during meiosis. High-concentration FSH in IVM medium led to reverse effect on the expression of hsa-miR-15a and hsa-miR-20a.Keywords miRNA . Oocytes . In vitro maturation . qRT-PCR Oocyte growth and early development requires large amounts of maternally-derived transcripts which are subjected to massive destruction as oocytes mature. Of the estimated 85 pg of polyadenylated mRNAs present in a germinal vesicle (GV)-stage mouse oocyte, 50 pg of mRNAs are degraded during oocyte maturation [1]. Furthermore, transcript degradation is highly selective, primarily affecting genes involved in processes associated with meiotic arrest at the GV-stage and progression of oocyte maturation, such as oxidative phosphorylation, energy production, protein synthesis and metabolism [2]. On the other hand, up-regulation of a number of transcripts during oocyte maturation was also observed in mice, cattle, and humans in recent years [3][4][5].miRNAs are a family of small non-coding RNAs that play important regulatory roles in gene expression. Specifically, miRNA-mediated translational regulation involves cleavage of messenger RNAs or repression of mRNA translation [6]. It has been estimated that 30% or more of human mRNAs are regulated by miRNAs [7]. Likewise, miRNAs may also play an important role in modulating gene expression in oocytes [8].Dynamic changes in miRNA expression during oogenesis were first revealed by a real-time PCR-based miRNA expression profiling method in single mouse oocytes [8]. A Yan-Wen Xu and Bin Wang contributed equally to this study. Capsule Differentially expressed miRNAs in the human oocytes between GV and MII stages were identified by miRNA microarrays. Dynamic changes of miR-15a, hsa-miR-20a, and miR-602 during meiosis were validated by qRT-PCR.

Abstractm5C is one of the longest-known RNA modifications, however, its developmental dynamics, functions, and evolution in mRNAs remain largely unknown. Here, we generate quantitative mRNA m5C maps at different stages of development in 6 vertebrate and invertebrate species and find convergent and unexpected massive methylation of maternal mRNAs mediated by NSUN2 and NSUN6. Using Drosophila as a model, we reveal that embryos lacking maternal mRNA m5C undergo cell cycle delays and fail to timely initiate maternal-to-zygotic transition, implying the functional importance of maternal mRNA m5C. From invertebrates to the lineage leading to humans, two waves of m5C regulatory innovations are observed: higher animals gain cis-directed NSUN2-mediated m5C sites at the 5' end of the mRNAs, accompanied by the emergence of more structured 5'UTR regions; humans gain thousands of trans-directed NSUN6-mediated m5C sites enriched in genes regulating the mitotic cell cycle. Collectively, our studies highlight the existence and regulatory innovations of a mechanism of early embryonic development and provide key resources for elucidating the role of mRNA m5C in biology and disease.

The application of a modified SCNT technique (OSM) followed by embryo culture in hamster embryo culture medium-10 (HECM-10) allows, for the first time, the routine production of SCNT blastocysts, most of which appear normal by immunochemical, cytochemical and in vitro developmental criteria. These embryos will provide a resource for isolating ES cells and for studies of nuclear reprogramming by monkey cytoplasts.

It is now well known that somatic cells can be efficiently reprogrammed into induced pluripotent stem cells (iPSCs) by forced expression of defined factors [1][2][3]. These cells, like embryonic stem cells (ESCs), have true pluripotency as shown by the live, fertile mice that can be generated through the tetraploid complementation assay using these iPSCs [4,5]. So far, iPSCs have been generated from many species including mice, primate, rat, as well as pigs [1,3,[6][7][8][9][10][11][12]; however, the latter failed to pass the gold standard test of pluripotency, that is, the tetraploid complementation assay, and iPSCs from some species have not even generated offspring with germline transmission. In addition, whether these iPSCs are capable of generating offspring through nuclear transfer remained to be determined.In this study, we first looked at three iPSC lines generated using the four Yamanaka factors (Oct3/4, Sox2, Klf4, and c-Myc) [1] and tagged with an Oct4-fused enhanced green fluorescent protein (GFP) [13]. These iPSC lines were originated from B6D2 F1 mouse embryonic fibroblasts (MEF) isolated from E13.5 fetuses, as we previously reported [4]. They express the correct pluripotency markers, and can form teratomas in severe combined immunodeficient mice. All three lines have the ability to produce chimeric mice, but only one (IP14D-1) can generate live mice through tetraploid complementation.To investigate the ability of different iPSC lines to generate cloned mice through somatic cell nuclear transfer (SCNT), and to compare with other nuclear donor cells, we first treated the three iPSC lines and their parental untransfected MEF cells, as well as two ESC lines (R1 and ESC2), with demecolcine to synchronize them at metaphase. Nuclei from each of the donor cell types were extracted and transferred to metaphase II (M II) oocytes of 8-week-old B6D2 F1 female mice using the Piezo-assisted one-step nuclear transfer method [14]. The reconstructed embryos were activated as previously described [15]. Embryos originated from iPSCs were selected by detection of fluorescence from Oct4-fused GFP expression. Interestingly, green fluorescence was observed starting from the eight-cell stage and was most apparent in the inner cell mass of the reconstructed embryos containing iPSC nuclei; this timing and localization coincide with normal Oct4 expression patterns. All of these embryos exhibited normal embryo morphology, indicating the likelihood of successful reprogramming of iPSCs by mouse M II ooplasm ( Figure 1A-1F).Preimplantation and postimplantation developmental efficiencies for the iPS-NT embryos were very similar to that of the ES-NT, and much higher than observed with MEF-NT embryos (Table 1). This difference is most exaggerated at the postimplantation stage. After transferring the nuclear transferred embryos back to the uterus of pseudopregnant white-coated CD-1 mice, 16 live cloned pups (1.0%) were born after 19.5 days of gestation, 3 of which were derived from the two iPSC lines that were not tetraploid comple...