CRISPR-Cas9-mediated functional dissection of 3'-UTRs.

Many studies using reporter assays have demonstrated that 3' untranslated regions (3'-UTRs) regulate gene expression by controlling mRNA stability and translation. Due to intrinsic limitations of heterologous reporter assays, we sought to develop a gene editing approach to investigate the regulatory activity of 3'-UTRs in their native context. We initially used dual-CRISPR (clustered, regularly interspaced, short palindromic repeats)-Cas9 targeting to delete DNA regions corresponding to nine chemokine 3'-UTRs that destabilized mRNA in a reporter assay. Targeting six chemokine 3'-UTRs increased chemokine mRNA levels as expected. However, targeting CXCL1, CXCL6 and CXCL8 3'-UTRs unexpectedly led to substantial mRNA decreases. Metabolic labeling assays showed that targeting these three 3'-UTRs increased mRNA stability, as predicted by the reporter assay, while also markedly decreasing transcription, demonstrating an unexpected role for 3'-UTR sequences in transcriptional regulation. We further show that CRISPR-Cas9 targeting of specific 3'-UTR elements can be used for modulating gene expression and for highly parallel localization of active 3'-UTR elements in the native context. Our work demonstrates the duality and complexity of 3'-UTR sequences in regulation of gene expression and provides a useful approach for modulating gene expression and for functional annotation of 3'-UTRs in the native context.