Chinese hamster ovary (CHO) cells are the superior host cell culture models used for the bioproduction of therapeutic proteins. One of the prerequisites for bioproduction using CHO cell lines is the need to generate stable CHO cell lines with optimal expression output. Antibiotic selection is commonly employed to isolate and select CHO cell lines with stable expression, despite its potential negative impact on cellular metabolism and expression level. Herein, we present a novel proline-based selection system for the isolation of stable CHO cell lines. The system exploits a dysfunctional proline metabolism pathway in CHO cells by using a pyrroline-5-carboxylate synthase gene as a selection marker, enabling selection to be made using proline-free media. The selection system was demonstrated by expressing green fluorescent protein (GFP) and a monoclonal antibody. When GFP was expressed, more than 90% of stable transfectants were enriched within 2 weeks of the selection period. When a monoclonal antibody was expressed, we achieved comparable titers (3.35 ± 0.47 μg/mL) with G418 and Zeocin-based selections (1.65 ± 0.46 and 2.25 ± 0.07 μg/mL, respectively). We further developed a proline-based coselection by using S. cerevisiae PRO1 and PRO2 genes as markers, which enables the generation of 99.5% double-transgenic cells. The proline-based selection expands available selection tools and provides an alternative to antibiotic-based selections in CHO cell line development.

CRISPR/Cas9 is increasingly being used as a tool to prosecute functional genomic screens. However, it is not yet possible to apply the approach at scale across a full breadth of cell types and endpoints. In order to address this, we developed a novel and robust workflow for array-based lentiviral CRISPR/Cas9 screening. We utilized a β-lactamase reporter gene assay to investigate mediators of TNF-α-mediated NF-κB signaling. The system was adapted for CRISPR/Cas9 through the development of a cell line stably expressing Cas9 and application of a lentiviral gRNA library comprising mixtures of four gRNAs per gene. We screened a 743-gene kinome library whereupon hits were independently ranked by percent inhibition, Z′ score, strictly standardized mean difference, and T statistic. A consolidated and optimized ranking was generated using Borda-based methods. Screening data quality was above acceptable limits (Z′ ≥ 0.5). In order to determine the contribution of individual gRNAs and to better understand false positives and negatives, a subset of gRNAs, against 152 genes, were profiled in singlicate format. We highlight the use of known reference genes and high-throughput, next-generation amplicon and RNA sequencing to assess screen data quality. Screening with singlicate gRNAs was more successful than screening with mixtures at identifying genes with known regulatory roles in TNF-α-mediated NF-κB signaling and was found to be superior to previous RNAi-based methods. These results add to the available data on TNF-α-mediated NF-κB signaling and establish a high-throughput functional genomic screening approach, utilizing a vector-based arrayed gRNA library, applicable across a wide variety of endpoints and cell types at a genome-wide scale.

KEY MESSAGE:A novel and robust lipofection-mediated transfection approach for the use of DNA-free Cas9/gRNA RNP for gene editing has demonstrated efficacy in plant cells. Precise genome editing has been revolutionized by CRISPR/Cas9 systems. DNA-based delivery of CRISPR/Cas9 is widely used in various plant species. However, protein-based delivery of the in vitro translated Cas9/guide RNA (gRNA) ribonucleoprotein (RNP) complex into plant cells is still in its infancy even though protein delivery has several advantages. These advantages include DNA-free delivery, gene-edited host plants that are not transgenic, ease of use, low cost, relative ease to be adapted to high-throughput systems, and low off-target cleavage rates. Here, we show a novel lipofection-mediated transfection approach for protein delivery of the preassembled Cas9/gRNA RNP into plant cells for genome editing. Two lipofection reagents, Lipofectamine 3000 and RNAiMAX, were adapted for successful delivery into plant cells of Cas9/gRNA RNP. A green fluorescent protein (GFP) reporter was fused in-frame with the C-terminus of the Cas9 protein and the fusion protein was successfully delivered into non-transgenic tobacco cv. 'Bright Yellow-2' (BY2) protoplasts. The optimal efficiencies for Lipofectamine 3000- and RNAiMAX-mediated protein delivery were 66% and 48%, respectively. Furthermore, we developed a biolistic method for protein delivery based on the known proteolistics technique. A transgenic tobacco BY2 line expressing an orange fluorescence protein reporter pporRFP was targeted for knockout. We found that the targeted mutagenesis frequency for our Lipofectamine 3000-mediated protein delivery was 6%. Our results showed that the newly developed lipofection-mediated transfection approach is robust for the use of the DNA-free Cas9/gRNA technology for genome editing in plant cells.

GUIDE-seq was developed to detect CRISPR/Cas9 off-target. However, as originally reported, it was associated with a high level of nonspecific amplification. In an attempt to improve it, we developed target-enriched GUIDE-seq (TEG-seq). The sensitivity level reached 0.1–10 reads-per-million depending on the NGS platform used, which was equivalent to 0.0002–1% measured by Targeted Amplicon-seq. Application of TEG-seq was demonstrated for the evaluation of various Cas9/gRNA configurations, which suggests delivery of Cas9/gRNA ribonucleoprotein results in significantly fewer off-targets than Cas9/gRNA plasmid. TEG-seq was also applied to 22 gRNAs with relatively high in silico ranking score that targeted the biological relevant SNPs. The result indicated the initial selection of gRNAs with high score is important, although it cannot exclude the possibility of off-target.

Abstract Identifying and validating targets that underlie disease mechanisms and can be addressed to provide efficacious therapies remains a significant challenge in the drug discovery and development process. Use of siRNA and shRNA to knock-down RNA and suppress gene function, have provided insights into mechanism of action, but depending on the nature of the targets, cells, biology and end-point assays these approaches may suffer variously from their transient nature, design complexity, incomplete knock-down or off-target effects. The use of CRISPR (clustered regularly interspaced short palindromic repeat)-associated Cas9 nuclease and guide RNA (gRNA) provides a strong alternative that can produce long-lasting impact, straightforward design, knock-out of genes and increased specificity. A number of laboratories have already published reports demonstrating how pools of gRNA can be delivered to cells and “hits” can be established through enrichment or depletion of cells following a “survival” assay and identified by sequencing the introduced gRNAs in the remaining cell population. Here we demonstrate a knock-out screening approach that utilizes the Invitrogen™ LentiArray™ CRISPR library to interrogate the impact of individual gene knock-outs on the NFκB pathway as measured by a functional cell-based assay. We describe the library design concepts, the assay development, initial screening results and validation of specific identified hits. The gRNAs are designed to primarily 5’ coding exons of a target gene using our CRISPR design tool to maximize knock-out efficiency and minimize off-target effects. Each gRNA is delivered as a separate lentiviral particle including an antibiotic-resistant marker and each gene is targeted by 4 gRNAs per well, delivered in a 96-well plate. We tested the approach using a library that targets the human kinome and developed a loss-of-function assay using our CellSensor® NF-κB-bla ME180 cell line, which is based on the ratiometric blue/green reporter assay and easily enables identification of genomic targets associated with the NF-κB pathway. We elucidate the key factors in developing a robust assay including both transduction and assay optimization to achieve the highest levels of transduction efficiency and assay window. Using these optimized parameters, we screened the Invitrogen™ LentiArray™ CRISPR kinome library that targets >800 kinases and demonstrate how we followed-up on and validated a subset of the identified hits. We expect these approaches to be scalable to the entire human genome and portable to multiple cell types and end-point assays including both high-throughput plate-based assays and high-content imaging based assays. Citation Format: Chetana M. Revankar, Justin Wetter, Julia Braun, Natasha Roark, Veronica Magnon, LaiYee Wong, Yanfei Zou, Namritha Ravinder, Jian-Ping Yang, Jonathan Chesnut, David Piper. Functional genomics screening using LentiArray™ CRISPR libraries and CellSensor™ assays [abstract]. In: Proceedings of the American Association for Cancer Research Annual Meeting 2017; 2017 Apr 1-5; Washington, DC. Philadelphia (PA): AACR; Cancer Res 2017;77(13 Suppl):Abstract nr LB-116. doi:10.1158/1538-7445.AM2017-LB-116

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. (C) 2016 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY license

To 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.Using 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.Lipofectamine 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.

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.

Genetic Engineering & Biotechnology NewsVol. 35, No. 15 OMICS TutorialProofing Genome EditsDetecting and Analyzing Genomic Modifications Induced with Engineered NucleasesJian-Ping Yang, Natasha Roark, Pei-Zhong Tang, Veronica Blackston, and Jonathan D. ChesnutJian-Ping YangJian-Ping Yang, Ph.D. (E-mail Address: jianping.yang@thermofisher.com), is a staff scientist, Natasha Roark is a scientist II, Pei-Zhong Tang, Ph.D., is a senior staff scientist, Veronica Blackston is a scientist II, and Jonathan D. Chesnut, Ph.D., is a senior director in R&D at Thermo Fisher Scientific. Website: www.lifetechnologies.com.Search for more papers by this author, Natasha RoarkSearch for more papers by this author, Pei-Zhong TangJian-Ping Yang, Ph.D. (E-mail Address: jianping.yang@thermofisher.com), is a staff scientist, Natasha Roark is a scientist II, Pei-Zhong Tang, Ph.D., is a senior staff scientist, Veronica Blackston is a scientist II, and Jonathan D. Chesnut, Ph.D., is a senior director in R&D at Thermo Fisher Scientific. Website: www.lifetechnologies.com.Search for more papers by this author, Veronica BlackstonSearch for more papers by this author, and Jonathan D. ChesnutJian-Ping Yang, Ph.D. (E-mail Address: jianping.yang@thermofisher.com), is a staff scientist, Natasha Roark is a scientist II, Pei-Zhong Tang, Ph.D., is a senior staff scientist, Veronica Blackston is a scientist II, and Jonathan D. Chesnut, Ph.D., is a senior director in R&D at Thermo Fisher Scientific. Website: www.lifetechnologies.com.Search for more papers by this authorPublished Online:26 Aug 2015https://doi.org/10.1089/gen.35.15.11AboutSectionsView articleView Full TextPDF/EPUB Permissions & CitationsPermissionsDownload CitationsTrack CitationsAdd to favorites Back To Publication ShareShare onFacebookTwitterLinked InRedditEmail View articleFiguresReferencesRelatedDetails Volume 35Issue 15Sep 2015 Information© 2015 by GEN PublishingTo cite this article:Jian-Ping Yang, Natasha Roark, Pei-Zhong Tang, Veronica Blackston, and Jonathan D. Chesnut.Proofing Genome Edits.Genetic Engineering & Biotechnology News.Sep 2015.26-27.http://doi.org/10.1089/gen.35.15.11Published in Volume: 35 Issue 15: August 26, 2015PDF download

With the advent of synthetic biology and cell engineering, the demand for large synthetic DNA fragments has been steadily increasing. Consequently, a number of multi-fragment cloning technologies optimized for the assembly of sizable DNA constructs have been developed. Still, screening for the right clone can be tedious because the high incidence of illegitimate assembly results in a relatively large proportion of missing or shuffled DNA elements. To mitigate this risk, we have developed a strategy that reduces the rate of fragment mis-assembly and is compatible with a variety of cloning methodologies. The approach is based on the positive selection of truncated plasmid markers, which are rendered active by providing their missing sequences during the assembly process. The method has been successfully validated in the context of complex in vivo and in vitro homologous recombination workflows, but it could be readily adapted to other cloning strategies, including those based on restriction endonucleases.

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

Since the discovery of restriction enzymes and the generation of the first recombinant DNA molecule over 40 years ago, molecular biology has evolved into a multidisciplinary field that has democratized the conversion of a digitized DNA sequence stored in a computer into its biological counterpart, usually as a plasmid, stored in a living cell. In this article, we summarize the most relevant tools that allow the swift assembly of DNA sequences into useful plasmids for biotechnological purposes. We cover the main components and stages in a typical DNA assembly workflow, namely in silico design, de novo gene synthesis, and in vitro and in vivo sequence assembly methodologies.

Transcription activator-like effectors (TALEs), secreted by the pathogenic bacteria Xanthomonas, specifically activate expression of targeted genes in plants. Here, we designed synthetic TALEs that bind to the flanking regions of the TATA-box motif on the CaMV 35S promoter for the purpose of understanding the engineerable 'hot-spots' for increasing transgene expression. We demonstrated that transient expression of de novo-engineered TALEs using agroinfiltration could significantly increase reporter gene expression in stable transgenic tobacco expressing the orange fluorescent protein reporter gene pporRFP under the control of synthetic inducible, minimal or full-length 35S promoters. Moreover, the additive effects of a combination of two different synthetic TALEs could significantly enhance the activation effects of TALEs on reporter gene expression more than when each TALE was used individually. We also studied the effects of the C-terminal domain and the activation domain of synthetic TALEs, as well as the best 'hot-spots' on the 35S promoter on targeted transgene activation. Furthermore, TALE activation of the Arabidopsis MYB transcription factor AtPAP1 (PRODUCTION OF ANTHOCYANIN PIGMENT 1) in stable transgenic tobacco gave rise to a dark purple colour on infiltrated leaves when driven by four copies of cis-regulatory elements of pathogenesis-related gene (PR1) with enhancer motifs B and A1 from the 35S promoter. These results provide novel insights into the potential applications of synthetic TALEs for targeted gene activation of transgenes in plants.

Prokaryotic regulatory proteins respond to diverse signals and represent a rich resource for building synthetic sensors and circuits. The TetR family contains >10(5) members that use a simple mechanism to respond to stimuli and bind distinct DNA operators. We present a platform that enables the transfer of these regulators to mammalian cells, which is demonstrated using human embryonic kidney (HEK293) and Chinese hamster ovary (CHO) cells. The repressors are modified to include nuclear localization signals (NLS) and responsive promoters are built by incorporating multiple operators. Activators are also constructed by modifying the protein to include a VP16 domain. Together, this approach yields 15 new regulators that demonstrate 19- to 551-fold induction and retain both the low levels of crosstalk in DNA binding specificity observed between the parent regulators in Escherichia coli, as well as their dynamic range of activity. By taking advantage of the DAPG small molecule sensing mediated by the PhlF repressor, we introduce a new inducible system with 50-fold induction and a threshold of 0.9 μM DAPG, which is comparable to the classic Dox-induced TetR system. A set of NOT gates is constructed from the new repressors and their response function quantified. Finally, the Dox- and DAPG- inducible systems and two new activators are used to build a synthetic enhancer (fuzzy AND gate), requiring the coordination of 5 transcription factors organized into two layers. This work introduces a generic approach for the development of mammalian genetic sensors and circuits to populate a toolbox that can be applied to diverse applications from biomanufacturing to living therapeutics.

Genetic Engineering & Biotechnology NewsVol. 34, No. 5 Drug Discovery TutorialTAL Effectors for Genome EditingTALEs Enable Rapid and Easy Design of Effectors to Control Gene Expression in CellsJian-Ping Yang, Veronica Blackston, Swarup Dash, and Jonathan ChesnutJian-Ping YangJian-Ping Yang, Ph.D. (E-mail Address: jianping.yang@lifetech.com), is a staff scientist, Veronica Blackston is a scientist II, Swarup Dash was a scientist, and Jonathan Chesnut is a research fellow in R&D at Life Technologies. Website: www.lifetechnologies.com.Search for more papers by this author, Veronica BlackstonSearch for more papers by this author, Swarup DashSearch for more papers by this author, and Jonathan ChesnutSearch for more papers by this authorPublished Online:27 Feb 2014https://doi.org/10.1089/gen.34.05.09AboutSectionsView articleView Full TextPDF/EPUB Permissions & CitationsPermissionsDownload CitationsTrack CitationsAdd to favorites Back To Publication ShareShare onFacebookTwitterLinked InRedditEmail View articleFiguresReferencesRelatedDetails Volume 34Issue 5Mar 2014 Information© 2014 by GEN PublishingTo cite this article:Jian-Ping Yang, Veronica Blackston, Swarup Dash, and Jonathan Chesnut.TAL Effectors for Genome Editing.Genetic Engineering & Biotechnology News.Mar 2014.16-17.http://doi.org/10.1089/gen.34.05.09Published in Volume: 34 Issue 5: February 27, 2014PDF download