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.

Chinese hamster ovary (CHO) cell engineering based on CRISPR/Cas9 knockout (KO) technology requires the delivery of guide RNA (gRNA) and Cas9 enzyme for efficient gene targeting. With an ever-increasing list of promising gene targets, developing, and optimizing a multiplex gene KO protocol is crucial for rapid CHO cell engineering. Here, we describe a method that can support efficient targeting and KO of up to 10 genes through sequential transfections. This method utilizes Cas9 protein to first screen multiple synthetic gRNAs per gene, followed by Sanger sequencing indel analysis, to identify effective gRNA sequences. Using sequential transfections of these potent gRNAs led to the isolation of single cell clones with the targeted deletion of all 10 genes (as confirmed by Sanger sequencing at the DNA level and mass spectrometry at the protein level). Screening 704 single cell clones yielded 6 clones in which all 10 genes were deleted through sequential transfections, demonstrating the success of this decaplex gene editing strategy. This pragmatic approach substantially reduces the time and effort required to generate multiple gene knockouts in CHO cells.

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The successes of chimeric antigen receptor (CAR) T cells in treating blood cancers have highlighted the cell therapy era. However, the difficulty of delivering molecules into immune cells has been an obstacle to more rapid advancement. Here we present an innovative large-scale Lentivirus (LV) production system as a solution to lower the cost and time of viral production. On the other hand, the next generation cell therapy will rely heavily on gene editing, especially in a safer non-viral integration manner. We have demonstrated that our novel non-viral all-in-one electroporation method provides high efficiency of gene knock-in in primary T cells. The new LV production system was developed for the clinical grade production of lentiviral vectors (LVVs) on a large-scale serum-free suspension platform. This technology employs a newly developed propriety set of GMP reagents comprising of culture media, suspension cells, transfection reagent and boosting enhancers. The system is able to deliver greater than 1 × 108 (TU/mL) functional titer with un-concentrated LVVs. For CRISPR/Cas9 gene editing in primary T cells, we were able to reach more than 90% knockout efficiency for most genes we tested, including T cell receptor (TCR), with Cas9 RNP electroporation using Neon Transfection System. More importantly, gene knock-in efficiency can be reached to greater than 30% with all-in-one electroporation, which delivers Cas9 RNP and donor DNA in one reaction using our newly developed electroporation buffer. Moreover, TCR knockout in addition to a knock-in at another locus can also be done in a single electroporation using our all-in-one method, which is ideal for developing next generation CAR and TCR-T cells.