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Challenging the Boundaries Between Treatment, Prevention, and Enhancement in Human Genome Editing.
Traditional distinctions between treatment and enhancement goals for human genome editing (HGE) have animated oversight considerations, yet these categories have been complicated by the addition of prevention as a possible target for HGE applications. To assess the role these three categories might play in continued HGE governance efforts, we report on interviews with genome editing scientists and governance group members. While some accepted traditional distinctions between treatment and enhancement and rejected the latter as unacceptable, others argued that the concept of enhancement is largely irrelevant or not as morally problematic as suggested. Others described how preventive goals for HGE create gray zones where prevention and enhancement may be difficult to distinguish, which may stymie uses of HGE. We conclude by discussing the governance implications of these various understandings of treatment, prevention, and enhancement as HGE research moves beyond the treatment of serious disease to embrace longer range preventive goals.
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Differential expression of genes in pre-blastoderm embryos of oriental fruit fly, Bactrocera dorsalis (Hendel) (Diptera: Tephritidae) microinjected with white locus CRISPR/Cas9 ribo nucleo protein (RNP) complex.
CRISPR/Cas9-based genome editing is currently revolutionizing applications in agriculture, human health, and other domains. Targeted gene editing employing CRISPR/Cas9 is achieved by delivering the various components viz. guide RNA, traCrRNA, and the Cas9 protein in DNA or RNA or RNP format. RNP enables fast, DNA-free editing without genomic integration. While innate immunity to guide RNA is studied in humans and plants, similar research in insects remains unexplored. In the present study, the RNP complex (sgRNA+Cas9) was administered via microinjection of embryos (eggs) in the pre-blastoderm (G0) stage. Transcriptome sequencing was performed using the Illumina Novoseq 6000 platform to understand the prevalence of innate immunity against the delivered guide RNA for the white locus of Bactrocera dorsalis. 33 up- and 67 down-regulated genes were found in the differential gene expression analysis from the RNA-Seq data. The correlation between RT-qPCR and RNA-Seq gene expression levels showed a strong coefficient of determination (R2 = 0.984). The genes related to stress, intron removal, and effector recognition were overexpressed. However, the genes responsible for growth and development were significantly downregulated. Thus, the present study identified the possible reasons that could play a vital role in influencing the editing outcomes employed through the RNP complex.
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Rapid Generation of Recombinant Poxviruses Using CRISPR/Cas9 Gene Editing.
The low-frequency natural recombination that is detected in poxvirus-infected cells has long been used to genetically modify poxviruses. Such recombinant poxviruses have found many applications as vaccines for preventing infectious diseases and as experimental cancer therapeutics. Unfortunately, these methods are time consuming, can leave behind "scars" or selectable markers, and many months of work may be required to generate plaque-purified recombinants bearing multiple virus gene substitutions, deletions, and/or inserted transgenes. Over the last decade, several reports have described how CRISPR/Cas9 technologies can be used to better facilitate genetic manipulation of vaccinia virus (VACV). These protocols use Cas9/gRNA complexes to introduce double-stranded breaks into specific sites in virus genomic DNA either in vivo or in vitro. Recombination-repair reactions are then employed to repair the breaks using transfected DNAs encoding the required homologies and desired mutation(s). Here we describe a method where we combine CRISPR/Cas9 genome editing in vitro, followed by Leporipoxvirus-catalyzed repair and reactivation of the cut VACV DNA using repair fragments provided in trans. This method optimizes several steps in the preparation of the CRISPR/Cas9-cut VACV DNA and can be used to introduce mutations at multiple sites without requiring selectable markers. It also provides some guidance regarding how the position of the CRISPR/Cas9-cuts can affect co-conversion of flanking markers embedded in the repair fragment. The method allows researchers to quickly generate recombinant VACV bearing multiple genetic alterations and using only a single round of reactivation and plating.
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Analysis of Oestrus and Hormonal Changes in the Progeny of 2 Genotypes of FecB Gene-Edited Sheep.
The FecB gene, the first gene identified for multiple births, can improve reproductive performance in sheep. We used the CRISPR/Cas9 editing system to obtain the FecB gene in Kazakh sheep and mated with unedited Kazakh sheep. After the genotypic screening of the progeny, four B+ and ++ Kazakh ewes, each 2 weeks old and healthy, and close to the same body weight, were selected to study oestrus and hormonal changes in the progeny population of FecB gene-edited Kazakh sheep. Our results showed that the FecB gene was moderately polymorphic (0.25 < PIC < 0.5) in the progeny population, was able to be stably inherited, and showed increased lambing numbers. The number of follicles and tiny follicles differed significantly (p < 0.05) between the 2 genotypes of ewes in the progeny population. Physiologically, FecB gene-edited sheep did not differ from wild-type sheep. Throughout the oestrous cycle, FSH, LH, and E2 in serum hormones of both sheep genotypes showed variability (p < 0.05) between 72-360 h, 360-528 h, and 12-72 h, respectively, compared to other periods. The concentration of FSH and PROG in follicular fluid was much higher than that of LH and E2 (p < 0.05). Overall, it is possible to make Kazakh sheep carry the FecB gene by CRISPR/Cas9 editing and stably inherit it into the progeny population. The enhancement of reproductive hormones by the FecB gene is beneficial to follicular development and ovulation and improves the reproductive efficiency of sheep.
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Isolation and Analysis of a Genome-Edited Single-Hepatocyte from a Cas9 Transgenic Mouse Line.
The primary cells isolated from the freshly dissected organ are thought to be different from those cultured for a long time in vitro. For instance, hepatocytes isolated in situ from the liver, display the ability to produce albumin, cultured for about a week often tend to cease production of albumin, including loss of proliferation capability. Thus, it is difficult to perform genome editing (i.e., production of genome-edited hepatocytes by in vitro gene delivery) in such cultured cells. Furthermore, hepatic cell lines available so far do not produce albumin and they would also have lost several characteristics of native liver cells. This poses a serious disadvantage when researchers want to study gene expression profiles under specific experimental settings, for example before and after genome editing. However, this demerit can be overcome if genome-editing is performed in situ in liver and single hepatocytes (both genome-edited and wild-type) can be isolated for analysis immediately following transient gene editing. Previously, we demonstrated successful isolation of genome-edited single hepatocytes, using mice expressing systemic Cas9 transgene (called "sCAT" mouse) and by tail-vein-mediated hydrodynamics-based gene delivery of gRNA targeted to Albumin gene (Sakurai et al., Sci Rep 6:20011, 2016). Here, we describe the detailed protocols for collection and analysis of single genome-edited hepatocytes, which will be useful for many types of hepatocyte functional studies.
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Generation of heritable germline mutations in the jewel wasp Nasonia vitripennis using CRISPR/Cas9.
The revolutionary RNA-guided endonuclease CRISPR/Cas9 system has proven to be a powerful tool for gene editing in a plethora of organisms. Here, utilizing this system we developed an efficient protocol for the generation of heritable germline mutations in the parasitoid jewel wasp, Nasonia vitripennis, a rising insect model organism for the study of evolution, development of axis pattern formation, venom production, haplo-diploid sex determination, and host-symbiont interactions. To establish CRISPR-directed gene editing in N. vitripennis, we targeted a conserved eye pigmentation gene cinnabar, generating several independent heritable germline mutations in this gene. Briefly, to generate these mutants, we developed a protocol to efficiently collect N. vitripennis eggs from a parasitized flesh fly pupa, Sarcophaga bullata, inject these eggs with Cas9/guide RNA mixtures, and transfer injected eggs back into the host to continue development. We also describe a flow for screening mutants and establishing stable mutant strains through genetic crosses. Overall, our results demonstrate that the CRISPR/Cas9 system is a powerful tool for genome manipulation in N. vitripennis, with strong potential for expansion to target critical genes, thus allowing for the investigation of several important biological phenomena in this organism.
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CRISPR takes on cancer.
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Considerations in engineering viral vectors for genome editing in plants.
Plant viruses have been engineered to express proteins and induce gene silencing for decades. Recently, plant viruses have also been used to deliver components into plant cells for genome editing, a technique called virus-induced genome editing (VIGE). Although more than a dozen plant viruses have been engineered into VIGE vectors and VIGE has been successfully accomplished in some plant species, application of VIGE to crops that are difficult to tissue culture and/or have low regeneration efficiency is still tough. This paper discusses factors to consider for an ideal VIGE vector, including insertion capacity for foreign DNA, vertical transmission ability, expression level of the target gene, stability of foreign DNA insertion, and biosafety. We also proposed a step-by-step schedule for excavating the suitable viral vector for VIGE.
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CRISPR/Cas9-Mediated CCR5 Ablation in Human Hematopoietic Stem/Progenitor Cells Confers HIV-1 Resistance In Vivo.
Transplantation of hematopoietic stem cells (HSCs) with a naturally occurring CCR5 mutation confers a loss of detectable HIV-1 in the patient, making ablation of the CCR5 gene in HSCs an ideal therapy for an HIV-1 cure. Although CCR5 disruption has been attempted in CD4+ T cells and hematopoietic stem/progenitor cells (HSPCs), efficient gene editing with high specificity and long-term therapeutic potential remains a major challenge for clinical translation. Here, we established a CRISPR/Cas9 gene editing system in human CD34+ HSPCs and achieved efficient CCR5 ablation evaluated in long-term reconstituted NOD/Prkdcscid/IL-2Rγnull mice. The CCR5 disruption efficiency in our system remained robust in secondary transplanted repopulating hematopoietic cells. More importantly, an HIV-1 resistance effect was observed as indicated by significant reduction of virus titration and enrichment of human CD4+ T cells. Hence, we successfully established a CRISPR/Cas9 mediated CCR5 ablating system in long-term HSCs, which confers HIV-1 resistance in vivo. Our study provides evidence for translating CCR5 gene-edited HSC transplantation for an HIV cure to the clinic.
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Correction of muscular dystrophies by CRISPR gene editing.
Muscular dystrophies are debilitating disorders that result in progressive weakness and degeneration of skeletal muscle. Although the genetic mutations and clinical abnormalities of a variety of neuromuscular diseases are well known, no curative therapies have been developed to date. The advent of genome editing technology provides new opportunities to correct the underlying mutations responsible for many monogenic neuromuscular diseases. For example, Duchenne muscular dystrophy, which is caused by mutations in the dystrophin gene, has been successfully corrected in mice, dogs, and human cells through CRISPR/Cas9 editing. In this Review, we focus on the potential for, and challenges of, correcting muscular dystrophies by editing disease-causing mutations at the genomic level. Ideally, because muscle tissues are extremely long-lived, CRISPR technology could offer a one-time treatment for muscular dystrophies by correcting the culprit genomic mutations and enabling normal expression of the repaired gene.
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