Dataset Information

Improved CRISPR/Cas9 gene editing in primary human myoblasts using low confluency cultures on Matrigel.

ABSTRACT:

Background

CRISPR/Cas9 is an invaluable tool for studying cell biology and the development of molecular therapies. However, delivery of CRISPR/Cas9 components into some cell types remains a major hurdle. Primary human myoblasts are a valuable cell model for muscle studies, but are notoriously difficult to transfect. There are currently no commercial lipofection protocols tailored for primary myoblasts, and most generic guidelines simply recommend transfecting healthy cells at high confluency. This study aimed to maximize CRISPR/Cas9 transfection and editing in primary human myoblasts.

Methods

Since increased cell proliferation is associated with increased transfection efficiency, we investigated two factors known to influence myoblast proliferation: cell confluency, and a basement membrane matrix, Matrigel. CRISPR/Cas9 editing was performed by delivering Cas9 ribonucleoprotein complexes via lipofection into primary human myoblasts, cultured in wells with or without a Matrigel coating, at low (~ 40%) or high (~ 80%) confluency.

Results

Cells transfected at low confluency on Matrigel-coated wells had the highest levels of transfection, and were most effectively edited across three different target loci, achieving a maximum editing efficiency of 93.8%. On average, editing under these conditions was >4-fold higher compared to commercial recommendations (high confluency, uncoated wells).

Conclusion

This study presents a simple, effective and economical method of maximizing CRISPR/Cas9-mediated gene editing in primary human myoblasts. This protocol could be a valuable tool for improving the genetic manipulation of cultured human skeletal muscle cells, and potentially be adapted for use in other cell types.

SUBMITTER: Goullee H

PROVIDER: S-EPMC8456651 | biostudies-literature |

REPOSITORIES: biostudies-literature

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Project description:BackgroundCRISPR-Cas9-based technologies have revolutionized experimental manipulation of mammalian genomes. None-the-less, limitations of the delivery and efficacy of these technologies restrict their application in primary cells.AimsTo create an optimized protocol for penetrant, reproducible, and fast targeted genome editing in cell cultures derived from primary cells, using patient-derived glioblastoma stem-like cells (GSCs) and human neural stem/progenitor cells (NSCs) for proof-of-concept experiments.Methods and resultsWe employed transient nucleofection of Cas9:sgRNA ribonucleoprotein complexes composed of chemically synthesized 2'-O-methyl 3'phosphorothioate-modified sgRNAs and purified Cas9 protein. Insertion-deletion mutation (indel) frequency and size distribution were measured via computational deconvolution of Sanger sequencing trace data. We found that this optimized technique routinely allows for >90% indel formation in only 3 days, without the need to create clonal lines for simple loss-of-function experiments. Using Western blotting, we observed near-total protein loss of target genes in cell pools. Additionally, we found that this approach allows for the creation of targeted genomic deletions. Furthermore, by using RNA-seq in edited NSCs to assess gene expression changes resulting from knockout of tumor suppressors commonly altered in glioblastoma, we also demonstrated the utility of this method for quickly creating a series of gene knockouts that allow for the study of oncogenic activities.ConclusionOur data suggest that this relatively simple method can be used for highly efficient and fast gene knockout, as well as for targeted genomic deletions, even in hyperdiploid cells (such as GSCs). This represents an extremely useful tool for the cancer research community when wishing to inactivate not only coding genes, but also non-coding RNAs, UTRs, enhancers, and promoters. This method can be readily applied to diverse cell types by varying the nucleofection conditions.