Proteolytic degradation of the Yap1 transcription factor is regulated by subcellular localization and the E3 ubiquitin ligase Not4.

Saccharomyces cerevisiae Yap1 is a transcriptional regulatory protein that serves as a central determinant of oxidative stress tolerance. Activity of this factor is regulated in large part by control of its subcellular location. In the absence of oxidants, Yap1 is primarily located in the cytoplasm. Upon oxidant challenge, Yap1 accumulates rapidly in the nucleus where it activates expression of genes required for oxidative stress tolerance such as the thioredoxin TRX2. Here, we demonstrate that Yap1 degradation is accelerated in response to oxidative stress. Yap1 is folded differently depending on the oxidant used to induce its nuclear localization but is degraded similarly, irrespective of its folded status. Mutant forms of Yap1 that are constitutively trapped in the nucleus are degraded in the absence of an oxidant signal. Degradation requires the ability of the protein to bind DNA and a domain in the amino-terminal region of the factor. Inhibition of the proteasome prevents Yap1 turnover. Screening a variety of mutants involved in ubiquitin-mediated proteolysis demonstrated an important role for the nuclear ubiquitin ligase Not4 in Yap1 degradation. Not4 was found to bind to Yap1 in an oxidant-stimulated fashion. The Candida albicans Yap1 homologue (Cap1) also was degraded after oxidant challenge. These data uncover a new, conserved pathway for regulation of the oxidative stress response that serves to temporally limit the duration of Yap1-dependent transcriptional activation.

Shuttle peptide delivers base editor RNPs to rhesus monkey airway epithelial cells in vivo.

Gene editing strategies for cystic fibrosis are challenged by the complex barrier properties of airway epithelia. We previously reported that the amphiphilic S10 shuttle peptide non-covalently combined with CRISPR-associated (Cas) ribonucleoprotein (RNP) enabled editing of human and mouse airway epithelial cells. Here, we derive the S315 peptide as an improvement over S10 in delivering base editor RNP. Following intratracheal aerosol delivery of Cy5-labeled peptide in rhesus macaques, we confirm delivery throughout the respiratory tract. Subsequently, we target CCR5 with co-administration of ABE8e-Cas9 RNP and S315. We achieve editing efficiencies of up-to 5.3% in rhesus airway epithelia. Moreover, we document persistence of edited epithelia for up to 12 months in mice. Finally, delivery of ABE8e-Cas9 targeting the CFTR R553X mutation restores anion channel function in cultured human airway epithelia. These results demonstrate the therapeutic potential of base editor delivery with S315 to functionally correct the CFTR R553X mutation in respiratory epithelia.

Tumor Apolipoprotein E is a key checkpoint blocking anti-tumor immunity in mouse melanoma.

Immunotherapy is a key modality in the treatment of cancer, but many tumors remain immune resistant. The classic mouse model of B16-F10 melanoma is immune resistant even in the face of checkpoint inhibition. Apolipoprotein E (apoE), a known immune suppressant is strikingly elevated in many human tumors, but its role in cancer immunology is not defined. We investigated the role of apoE in the immune micro-environment using a mouse melanoma model. We demonstrate that ApoE is -highly expressed in wild-type B16-F10 melanoma and serum levels progressively increase as tumors grow. The conditioned media from wild type ApoE secreting melanoma cells suppress T-cell activation in vitro while this suppressive effect is absent in conditioned media from ApoE knock out tumor cells. Mechanistically, apoE induces IL-10 secreting dendritic cells and stimulates T-cell apoptosis and arrest partially via the lrp8 receptor. Ablating ApoE in mice inoculated with tumor cells enabled tumor cell rejection and was associated with induction of immune pathway activation and immune cell infiltration. Tumor secreted apoE appears to be a potent immune cell checkpoint and targeting apoE is associated with enhanced tumor immunity in the mouse melanoma model.

Pharmacological interventions enhance virus-free generation of TRAC-replaced CAR T cells.

Chimeric antigen receptor (CAR) redirected T cells are potent therapeutic options against hematological malignancies. The current dominant manufacturing approach for CAR T cells depends on retroviral transduction. With the advent of gene editing, insertion of a CD19-CAR into the T cell receptor (TCR) alpha constant (TRAC) locus using adeno-associated viruses for gene transfer was demonstrated, and these CD19-CAR T cells showed improved functionality over their retrovirally transduced counterparts. However, clinical-grade production of viruses is complex and associated with extensive costs. Here, we optimized a virus-free genome-editing method for efficient CAR insertion into the TRAC locus of primary human T cells via nuclease-assisted homology-directed repair (HDR) using CRISPR-Cas and double-stranded template DNA (dsDNA). We evaluated DNA-sensor inhibition and HDR enhancement as two pharmacological interventions to improve cell viability and relative CAR knockin rates, respectively. While the toxicity of transfected dsDNA was not fully prevented, the combination of both interventions significantly increased CAR knockin rates and CAR T cell yield. Resulting TRAC-replaced CD19-CAR T cells showed antigen-specific cytotoxicity and cytokine production in vitro and slowed leukemia progression in a xenograft mouse model. Amplicon sequencing did not reveal significant indel formation at potential off-target sites with or without exposure to DNA-repair-modulating small molecules. With TRAC-integrated CAR+ T cell frequencies exceeding 50%, this study opens new perspectives to exploit pharmacological interventions to improve non-viral gene editing in T cells.

Optimized design parameters for CRISPR Cas9 and Cas12a homology-directed repair.

CRISPR-Cas proteins are RNA-guided nucleases used to introduce double-stranded breaks (DSBs) at targeted genomic loci. DSBs are repaired by endogenous cellular pathways such as non-homologous end joining (NHEJ) and homology-directed repair (HDR). Providing an exogenous DNA template during repair allows for the intentional, precise incorporation of a desired mutation via the HDR pathway. However, rates of repair by HDR are often slow compared to the more rapid but less accurate NHEJ-mediated repair. Here, we describe comprehensive design considerations and optimized methods for highly efficient HDR using single-stranded oligodeoxynucleotide (ssODN) donor templates for several CRISPR-Cas systems including S.p. Cas9, S.p. Cas9 D10A nickase, and A.s. Cas12a delivered as ribonucleoprotein (RNP) complexes. Features relating to guide RNA selection, donor strand preference, and incorporation of blocking mutations in the donor template to prevent re-cleavage were investigated and were implemented in a novel online tool for HDR donor template design. These findings allow for high frequencies of precise repair utilizing HDR in multiple mammalian cell lines. Tool availability: https://www.idtdna.com/HDR.

AsCas12a ultra nuclease facilitates the rapid generation of therapeutic cell medicines.

Though AsCas12a fills a crucial gap in the current genome editing toolbox, it exhibits relatively poor editing efficiency, restricting its overall utility. Here we isolate an engineered variant, "AsCas12a Ultra", that increased editing efficiency to nearly 100% at all sites examined in HSPCs, iPSCs, T cells, and NK cells. We show that AsCas12a Ultra maintains high on-target specificity thereby mitigating the risk for off-target editing and making it ideal for complex therapeutic genome editing applications. We achieved simultaneous targeting of three clinically relevant genes in T cells at >90% efficiency and demonstrated transgene knock-in efficiencies of up to 60%. We demonstrate site-specific knock-in of a CAR in NK cells, which afforded enhanced anti-tumor NK cell recognition, potentially enabling the next generation of allogeneic cell-based therapies in oncology. AsCas12a Ultra is an advanced CRISPR nuclease with significant advantages in basic research and in the production of gene edited cell medicines.