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DddA-derived cytosine base editors (DdCBEs) enable the targeted introduction of C•G-to-T•A conversions in mitochondrial DNA (mtDNA). DdCBEs work in pairs, with each arm composed of a transcription activator-like effector (TALE), a split double-stranded DNA deaminase half, and a uracil glycosylase inhibitor. This pioneering technology has helped improve our understanding of cellular processes involving mtDNA and has paved the way for the development of models and therapies for genetic disorders caused by pathogenic mtDNA variants. Nonetheless, given the intrinsic properties of TALE proteins, several target sites in human mtDNA are predicted to remain out of reach to DdCBEs and other TALE-based technologies. Specifically, due to the conventional requirement for a thymine immediately upstream of the TALE target sequences ( i.e. , the 5'-T constraint), over 150 loci in the human mitochondrial genome are presumed to be inaccessible to DdCBEs. Previous attempts at circumventing this requirement, either by developing monomeric DdCBEs or utilizing DNA-binding domains alternative to TALEs, have resulted in suboptimal specificity profiles with reduced therapeutic potential. Here, aiming to challenge and elucidate the relevance of the 5'-T constraint in the context of DdCBE-mediated mtDNA editing, and to expand the range of motifs that are editable by this technology, we generated DdCBEs containing TALE proteins engineered to recognize all 5' bases. These modified DdCBEs are herein referred to as αDdCBEs. Notably, 5'-T-noncompliant canonical DdCBEs efficiently edited mtDNA at diverse loci. However, they were frequently outperformed by αDdCBEs, which exhibited significant improvements in activity and specificity, regardless of the most 5' bases of their TALE binding sites. Furthermore, we showed that αDdCBEs are compatible with the enhanced DddA tox variants DddA6 and DddA11, and we validated TALE shifting with αDdCBEs as an effective approach to optimize base editing outcomes. Overall, αDdCBEs enable efficient, specific, and unconstrained mitochondrial base editing.

DddA-derived cytosine base editors (DdCBEs) enable the targeted introduction of C•G-to-T•A conversions in mitochondrial DNA (mtDNA). DdCBEs are often deployed as pairs, with each arm comprised of a transcription activator-like effector (TALE), a split double-stranded DNA deaminase half, and a uracil glycosylase inhibitor. This pioneering technology has helped improve our understanding of cellular processes involving mtDNA and has paved the way for the development of models and therapies for genetic disorders caused by pathogenic mtDNA variants. Nonetheless, given the intrinsic properties of TALE proteins, several target sites in human mtDNA remain out of reach to DdCBEs and other TALE-based technologies. Specifically, due to the conventional requirement for a thymine immediately upstream of the TALE target sequences (i.e., the 5'-T constraint), over 150 loci in the human mitochondrial genome are presumed to be inaccessible to DdCBEs. Previous attempts at circumventing this constraint, either by developing monomeric DdCBEs or utilizing DNA-binding domains alternative to TALEs, have resulted in suboptimal specificity profiles with reduced therapeutic potential. Here, aiming to challenge and elucidate the relevance of the 5'-T constraint in the context of DdCBE-mediated mtDNA editing, and to expand the range of motifs that are editable by this technology, we generated αDdCBEs that contain modified TALE proteins engineered to recognize all 5' bases. Notably, 5'-T-noncompliant, canonical DdCBEs efficiently edited mtDNA at diverse loci. However, DdCBEs were frequently outperformed by αDdCBEs, which consistently displayed significant improvements in activity and specificity, regardless of the 5'-most bases of their TALE binding sites. Furthermore, we showed that αDdCBEs are compatible with DddA tox and its derivatives DddA6, and DddA11, and we validated TALE shifting with αDdCBEs as an effective approach to optimize base editing outcomes at a single target site. Overall, αDdCBEs enable efficient, specific, and unconstrained mitochondrial base editing.

Background & Aims: Although depletion of neuronal nitric oxide synthase (NOS1)-expressing neurons contributes to gastroparesis, stimulating nitrergic signaling is not an effective therapy. We investigated whether hypoxia-inducible factor 1α (HIF1A), which is activated by high O 2 consumption in central neurons, is a Nos1 transcription factor in enteric neurons and whether stabilizing HIF1A reverses gastroparesis., Methods: Mice with streptozotocin-induced diabetes, human and mouse tissues, NOS1 + mouse neuroblastoma cells, and isolated nitrergic neurons were studied. Gastric emptying of solids and volumes were determined by breath test and single-photon emission computed tomography, respectively. Gene expression was analyzed by RNA-sequencing, microarrays, immunoblotting, and immunofluorescence. Epigenetic assays included chromatin immunoprecipitation sequencing (13 targets), chromosome conformation capture sequencing, and reporter assays. Mechanistic studies used Cre-mediated recombination, RNA interference, and clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9)-mediated epigenome editing., Results: HIF1A signaling from physiological intracellular hypoxia was active in mouse and human NOS1 + myenteric neurons but reduced in diabetes. Deleting Hif1a in Nos1-expressing neurons reduced NOS1 protein by 50% to 92% and delayed gastric emptying of solids in female but not male mice. Stabilizing HIF1A with roxadustat (FG-4592), which is approved for human use, restored NOS1 and reversed gastroparesis in female diabetic mice. In nitrergic neurons, HIF1A up-regulated Nos1 transcription by binding and activating proximal and distal cis-regulatory elements, including newly discovered super-enhancers, facilitating RNA polymerase loading and pause-release, and by recruiting cohesin to loop anchors to alter chromosome topology., Conclusions: Pharmacologic HIF1A stabilization is a novel, translatable approach to restoring nitrergic signaling and treating diabetic gastroparesis. The newly recognized effects of HIF1A on chromosome topology may provide insights into physioxia- and ischemia-related organ function., (Copyright © 2023 AGA Institute. Published by Elsevier Inc. All rights reserved.)

CD8 + T cell exhaustion (Tex) limits disease control during chronic viral infections and cancer. Here, we investigated the epigenetic factors mediating major chromatin-remodeling events in Tex-cell development. A protein-domain-focused in vivo CRISPR screen identified distinct functions for two versions of the SWI/SNF chromatin-remodeling complex in Tex-cell differentiation. Depletion of the canonical SWI/SNF form, BAF, impaired initial CD8 + T cell responses in acute and chronic infection. In contrast, disruption of PBAF enhanced Tex-cell proliferation and survival. Mechanistically, PBAF regulated the epigenetic and transcriptional transition from TCF-1 + progenitor Tex cells to more differentiated TCF-1 - Tex subsets. Whereas PBAF acted to preserve Tex progenitor biology, BAF was required to generate effector-like Tex cells, suggesting that the balance of these factors coordinates Tex-cell subset differentiation. Targeting PBAF improved tumor control both alone and in combination with anti-PD-L1 immunotherapy. Thus, PBAF may present a therapeutic target in cancer immunotherapy., Competing Interests: Declaration of interests E.J.W. is a member of the Parker Institute for Cancer Immunotherapy, which supports research in the Wherry laboratory. E.J.W. is an advisor for Danger Bio, Janssen, New Limit, Marengo, Pluto Immunotherapeutics Related Sciences, Santa Ana Bio, Synthekine, and Surface Oncology. E.J.W. is a founder of and holds stock in Surface Oncology, Danger Bio, and Arsenal Biosciences., (Copyright © 2023 Elsevier Inc. All rights reserved.)

The FusX TALE Based Editor (FusXTBE) is a programmable base editing platform that can introduce specific TC-to-TT variations in the mitochondrial DNA (mtDNA). Here, we provide a protocol describing the synthesis and testing of the FusXTBE plasmids in cultured human cell lines. This tool is designed to be easily modified to work in diverse applications where editing of mitochondrial DNA is desired. For complete details on the use and execution of this protocol, please refer to Sabharwal et al. (2021) and Ma et al. (2016)., Competing Interests: Mayo Clinic has patent filings that cover the FusX TALE assembly kit and applications. K.J.C. and S.C.E. are shareholders in LifEngine Technologies Inc., which has a license to use the FusX TALE assembly kit., (© 2022 The Author(s).)

Nearly 90% of human pathogenic mutations are caused by small genetic variations, and methods to correct these errors efficiently are critically important. One way to make small DNA changes is providing a single-stranded oligo deoxynucleotide (ssODN) containing an alteration coupled with a targeted double-strand break (DSB) at the target locus in the genome. Coupling an ssODN donor with a CRISPR-Cas9-mediated DSB is one of the most streamlined approaches to introduce small changes. However, in many systems, this approach is inefficient and introduces imprecise repair at the genetic junctions. We herein report a technology that uses spatiotemporal localization of an ssODN with CRISPR-Cas9 to improve gene alteration. We show that by fusing an ssODN template to the trans-activating RNA (tracrRNA), we recover precise genetic alterations, with increased integration and precision in vitro and in vivo . Finally, we show that this technology can be used to enhance gene conversion with other gene editing tools such as transcription activator like effector nucleases.

The development of CRISPR-associated proteins, such as Cas9, has led to increased accessibility and ease of use in genome editing. However, additional tools are needed to quantify and identify successful genome editing events in living animals. We developed a method to rapidly quantify and monitor gene editing activity non-invasively in living animals that also facilitates confocal microscopy and nucleotide level analyses. Here we report a new CRISPR "fingerprinting" approach to activating luciferase and fluorescent proteins in mice as a function of gene editing. This system is based on experience with our prior cre recombinase (cre)-detector system and is designed for Cas editors able to target lox P including gRNAs for SaCas9 and ErCas12a. These CRISPRs cut specifically within lox P, an approach that is a departure from previous gene editing in vivo activity detection techniques that targeted adjacent stop sequences. In this sensor paradigm, CRISPR activity was monitored non-invasively in living cre reporter mice (FVB.129S6(B6)-Gt(ROSA)26Sortm1(Luc)Kael/J and Gt(ROSA)26Sortm4(ACTB-tdTomato,-EGFP)Luo/J, which will be referred to as LSL-luciferase and mT/mG throughout the paper) after intramuscular or intravenous hydrodynamic plasmid injections, demonstrating utility in two diverse organ systems. The same genome-editing event was examined at the cellular level in specific tissues by confocal microscopy to determine the identity and frequency of successfully genome-edited cells. Further, SaCas9 induced targeted editing at efficiencies that were comparable to cre, demonstrating high effective delivery and activity in a whole animal. This work establishes genome editing tools and models to track CRISPR editing in vivo non-invasively and to fingerprint the identity of targeted cells. This approach also enables similar utility for any of the thousands of previously generated lox P animal models.

CRISPR and CRISPR-Cas effector proteins enable the targeting of DNA double-strand breaks to defined loci based on a variable length RNA guide specific to each effector. The guide RNAs are generally similar in size and form, consisting of a ∼20 nucleotide sequence complementary to the DNA target and an RNA secondary structure recognized by the effector. However, the effector proteins vary in protospacer adjacent motif requirements, nuclease activities, and DNA binding kinetics. Recently, ErCas12a, a new member of the Cas12a family, was identified in Eubacterium rectale . Here, we report the first characterization of ErCas12a activity in zebrafish and expand on previously reported activity in human cells. Using a fluorescent reporter system, we show that CRISPR-ErCas12a elicits strand annealing mediated DNA repair more efficiently than CRISPR-Cas9. Further, using our previously reported gene targeting method that utilizes short homology, GeneWeld, we demonstrate the use of CRISPR-ErCas12a to integrate reporter alleles into the genomes of both zebrafish and human cells. Together, this work provides methods for deploying an additional CRISPR-Cas system, thus increasing the flexibility researchers have in applying genome engineering technologies.

The rapid growth of the field of gene editing can largely be attributed to the discovery and optimization of designer endonucleases. These include zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and clustered regular interspersed short palindromic repeat (CRISPR) systems including Cas9, Cas12a, and structure-guided nucleases. Zebrafish (Danio rerio) have proven to be a powerful model system for genome engineering testing and applications due to their external development, high fecundity, and ease of housing. As the zebrafish gene editing toolkit continues to grow, it is becoming increasingly important to understand when and how to utilize which of these technologies for maximum efficacy in a particular project. While CRISPR-Cas9 has brought broad attention to the field of genome engineering in recent years, designer endonucleases have been utilized in genome engineering for more than two decades. This chapter provides a brief overview of designer endonuclease and other gene editing technologies in zebrafish as well as some of their known functional benefits and limitations depending on specific project goals. Finally, selected prospects for additional gene editing tools are presented, promising additional options for directed genomic programming of this versatile animal model system., (Copyright © 2018 The Authors. Published by Elsevier Inc. All rights reserved.)