CRISPR-Cas9 systems provide a platform for high efficiency genome editing that are enabling innovative applications of mammalian cell engineering. However, the delivery of Cas9 and synthesis of guide RNA (gRNA) remain as steps that can limit overall efficiency and ease of use. Here we describe methods for rapid synthesis of gRNA and for delivery of Cas9 protein/gRNA ribonucleoprotein complexes (Cas9 RNPs) into a variety of mammalian cells through liposome-mediated transfection or electroporation. Using these methods, we report nuclease-mediated indel rates of up to 94% in Jurkat T cells and 87% in induced pluripotent stem cells (iPSC) for a single target. When we used this approach for multigene targeting in Jurkat cells we found that two-locus and three-locus indels were achieved in approximately 93% and 65% of the resulting isolated cell lines, respectively. Further, we found that the off-target cleavage rate is reduced using Cas9 protein when compared to plasmid DNA transfection. Taken together, we present a streamlined cell engineering workflow that enables gRNA design to analysis of edited cells in as little as four days and results in highly efficient genome modulation in hard-to-transfect cells. The reagent preparation and delivery to cells is amenable to high throughput, multiplexed genome-wide cell engineering.

While CRISPR-based gene knock out in mammalian cells has proven to be very efficient, precise insertion of genetic elements via the cellular homology directed repair (HDR) pathway remains a rate-limiting step to seamless genome editing. Under the conditions described here, we achieved up to 56% targeted integration efficiency with up to a six-nucleotide insertion in HEK293 cells. In induced pluripotent stem cells (iPSCs), we achieved precise genome editing rates of up to 45% by co-delivering the Cas9 RNP and donor DNA. In addition, the use of a short double stranded DNA oligonucleotide with 3' overhangs allowed integration of a longer FLAG epitope tag along with a restriction site at rates of up to 50%. We propose a model that favors the design of donor DNAs with the change as close to the cleavage site as possible. For small changes such as SNPs or short insertions, asymmetric single stranded donor molecules with 30 base homology arms 3' to the insertion/repair cassette and greater than 40 bases of homology on the 5' end seems to be favored. For larger insertions such as an epitope tag, a dsDNA donor with protruding 3' homology arms of 30 bases is favored. In both cases, protecting the ends of the donor DNA with phosphorothioate modifications improves the editing efficiency.

ObjectivesTo identify the best lipid nanoparticles for delivery of purified Cas9 protein and gRNA complexes (Cas9 RNPs) into mammalian cells and to establish the optimal conditions for transfection.ResultsUsing a systematic approach, we screened 60 transfection reagents using six commonly-used mammalian cell lines and identified a novel transfection reagent (named Lipofectamine CRISPRMAX). Based on statistical analysis, the genome modification efficiencies in Lipofectamine CRISPRMAX-transfected cell lines were 40 or 15 % higher than those in Lipofectamine 3000 or RNAiMAX-transfected cell lines, respectively. Upon optimization of transfection conditions, we observed 85, 75 or 55 % genome editing efficiencies in HEK293FT cells, mouse ES cells, or human iPSCs, respectively. Furthermore, we were able to co-deliver donor DNA with Cas9 RNPs into a disrupted EmGFP stable cell line, resulting in the generation of up to 17 % EmGFP-positive cells.ConclusionLipofectamine CRISPRMAX was characterized as the best lipid nanoparticles for the delivery of Cas9 RNPs into a variety of mammalian cell lines, including mouse ES cells and iPSCs.Electronic supplementary materialThe online version of this article (doi:10.1007/s10529-016-2064-9) contains supplementary material, which is available to authorized users.

Alexander disease (AxD) is a fatal neurodegenerative disorder caused by mutations in glial fibrillary acidic protein (GFAP), which supports the structural integrity of astrocytes. Over 70 GFAP missense mutations cause AxD, but the mechanism linking different mutations to disease-relevant phenotypes remains unknown. We used AxD patient brain tissue and induced pluripotent stem cell (iPSC)-derived astrocytes to investigate the hypothesis that AxD-causing mutations perturb key post-translational modifications (PTMs) on GFAP. Our findings reveal selective phosphorylation of GFAP-Ser13 in patients who died young, independently of the mutation they carried. AxD iPSC-astrocytes accumulated pSer13-GFAP in cytoplasmic aggregates within deep nuclear invaginations, resembling the hallmark Rosenthal fibers observed in vivo. Ser13 phosphorylation facilitated GFAP aggregation and was associated with increased GFAP proteolysis by caspase-6. Furthermore, caspase-6 was selectively expressed in young AxD patients, and correlated with the presence of cleaved GFAP. We reveal a novel PTM signature linking different GFAP mutations in infantile AxD.

The generation of induced pluripotent stem cells (iPSCs) from somatic cells has enabled the possibility of providing unprecedented access to patient-specific iPSC cells for drug screening, disease modeling, and cell therapy applications. However, a major obstacle to the use of iPSC for therapeutic applications is the potential of genomic modifications caused by insertion of viral transgenes in the cellular genome. A second concern is that reprogramming often requires the use of animal feeder layers and reagents that contain animal origin products, which hinder the generation of clinical-grade iPSCs. Here, we report the generation of iPSCs by an RNA Sendai virus vector that does not integrate into the cells genome, providing transgene-free iPSC line. In addition, reprogramming can be performed in feeder-free condition with StemPro hESC SFM medium and in xeno-free (XF) conditions. Generation of an integrant-free iPSCs generated in xeno-free media should facilitate the safe downstream applications of iPSC-based cell therapies.

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