Deep transcriptome profiling reveals limited conservation of A-to-I RNA editing in Xenopus.

BACKGROUND: Xenopus has served as a valuable model system for biomedical research over the past decades. Notably, ADAR was first detected in frog oocytes and embryos as an activity that unwinds RNA duplexes. However, the scope of A-to-I RNA editing by the ADAR enzymes in Xenopus remains underexplored. RESULTS: Here, we identify millions of editing events in Xenopus with high accuracy and systematically map the editome across developmental stages, adult organs, and species. We report diverse spatiotemporal patterns of editing with deamination activity highest in early embryogenesis before zygotic genome activation and in the ovary. Strikingly, editing events are poorly conserved across different Xenopus species. Even sites that are detected in both X. laevis and X. tropicalis show largely divergent editing levels or developmental profiles. In protein-coding regions, only a small subset of sites that are found mostly in the brain are well conserved between frogs and mammals. CONCLUSIONS: Collectively, our work provides fresh insights into ADAR activity in vertebrates and suggest that species-specific editing may play a role in each animal's unique physiology or environmental adaptation.

Identification of Bona Fide RNA Editing Sites: History, Challenges, and Opportunities.

Adenosine-to-inosine (A-to-I) RNA editing, catalyzed by the adenosine deaminase acting on the RNA (ADAR) family of enzymes of which there are three members (ADAR1, ADAR2, and ADAR3), is a major gene regulatory mechanism that diversifies the transcriptome. It is widespread in many metazoans, including humans. As inosine is interpreted by cellular machineries mainly as guanosine, A-to-I editing effectively gives A-to-G nucleotide changes. Depending on its location, an editing event can generate new protein isoforms or influence other RNA processing pathways. Researchers have found that ADAR-mediated editing performs diverse functions. For example, it enables living organisms such as cephalopods to adapt rapidly to fluctuating environmental conditions such as water temperature. In development, the loss of ADAR1 is embryonically lethal partly because endogenous double-stranded RNAs (dsRNAs) are no longer marked by inosines, which signal "self", and thus cause the melanoma differentiation-associated protein 5 (MDA5) sensor to trigger a deleterious interferon response. Hence, ADAR1 plays a key role in preventing aberrant activation of the innate immune system. Furthermore, ADAR enzymes have been implicated in myriad human diseases. Intriguingly, some cancer cells are known to exploit ADAR1 activity to dodge immune responses. However, the exact identities of immunogenic RNAs in different biological contexts have remained elusive. Consequently, there is tremendous interest in identifying inosine-containing RNAs in the cell.The identification of A-to-I RNA editing sites is dependent on the sequencing of nucleic acids. Technological and algorithmic advancements over the past decades have revolutionized the way editing events are detected. At the beginning, the discovery of editing sites relies on Sanger sequencing, a first-generation technology. Both RNA, which is reverse transcribed into complementary DNA (cDNA), and genomic DNA (gDNA) from the same source are analyzed. After sequence alignment, one would require an adenosine to be present in the genome but a guanosine to be detected in the RNA sample for a position to be declared as an editing site. However, an issue with Sanger sequencing is its low throughput. Subsequently, Illumina sequencing, a second-generation technology, was invented. By permitting the simultaneous interrogation of millions of molecules, it enables many editing sites to be identified rapidly. However, a key challenge is that the Illumina platform produces short sequencing reads that can be difficult to map accurately. To tackle the challenge, we and others developed computational workflows with a series of filters to discard sites that are likely to be false positives. When Illumina sequencing data sets are properly analyzed, A-to-G variants should emerge as the most dominant mismatch type. Moreover, the quantitative nature of the data allows us to build a comprehensive atlas of editing-level measurements across different biological contexts, providing deep insights into the spatiotemporal dynamics of RNA editing. However, difficulties remain in identifying true A-to-I editing sites in short protein-coding exons or in organisms and diseases where DNA mutations and genomic polymorphisms are prevalent and mostly unknown. Nanopore sequencing, a third-generation technology, promises to address the difficulties, as it allows native RNAs to be sequenced without conversion to cDNA, preserving base modifications that can be directly detected through machine learning. We recently demonstrated that nanopore sequencing could be used to identify A-to-I editing sites in native RNA directly. Although further work is needed to enhance the detection accuracy in single molecules from fewer cells, the nanopore technology holds the potential to revolutionize epitranscriptomic studies.

Direct identification of A-to-I editing sites with nanopore native RNA sequencing.

Inosine is a prevalent RNA modification in animals and is formed when an adenosine is deaminated by the ADAR family of enzymes. Traditionally, inosines are identified indirectly as variants from Illumina RNA-sequencing data because they are interpreted as guanosines by cellular machineries. However, this indirect method performs poorly in protein-coding regions where exons are typically short, in non-model organisms with sparsely annotated single-nucleotide polymorphisms, or in disease contexts where unknown DNA mutations are pervasive. Here, we show that Oxford Nanopore direct RNA sequencing can be used to identify inosine-containing sites in native transcriptomes with high accuracy. We trained convolutional neural network models to distinguish inosine from adenosine and guanosine, and to estimate the modification rate at each editing site. Furthermore, we demonstrated their utility on the transcriptomes of human, mouse and Xenopus. Our approach expands the toolkit for studying adenosine-to-inosine editing and can be further extended to investigate other RNA modifications.

Small-molecule enhancers of CRISPR-induced homology-directed repair in gene therapy: A medicinal chemist's perspective.

CRISPR technologies are increasingly being investigated and utilized for the treatment of human genetic diseases via genome editing. CRISPR-Cas9 first generates a targeted DNA double-stranded break, and a functional gene can then be introduced to replace the defective copy in a precise manner by templated repair via the homology-directed repair (HDR) pathway. However, this is challenging owing to the relatively low efficiency of the HDR pathway compared with a rival random repair pathway known as non-homologous end joining (NHEJ). Small molecules can be employed to increase the efficiency of HDR and decrease that of NHEJ to improve the efficiency of precise knock-in genome editing. This review discusses the potential usage of such small molecules in the context of gene therapy and their drug-likeness, from a medicinal chemist's perspective.

A Sensitive and Specific Fluorescent RT-LAMP Assay for SARS-CoV-2 Detection in Clinical Samples.

The raging COVID-19 pandemic has created an unprecedented demand for frequent and widespread testing to limit viral transmission. Reverse transcription loop-mediated isothermal amplification (RT-LAMP) has emerged as a promising diagnostic platform for rapid detection of SARS-CoV-2, in part because it can be performed with simple instrumentation. However, isothermal amplification methods frequently yield spurious amplicons even in the absence of a template. Consequently, RT-LAMP assays can produce false positive results when they are based on generic intercalating dyes or pH-sensitive indicators. Here, we report the development of a sensitive RT-LAMP assay that leverages on a novel sequence-specific probe to guard against spurious amplicons. We show that our optimized fluorescent assay, termed LANTERN, takes only 30 min to complete and can be applied directly on swab or saliva samples. Furthermore, utilizing clinical RNA samples from 52 patients with COVID-19 infection and 21 healthy individuals, we demonstrate that our diagnostic test exhibits a specificity and positive predictive value of 95% with a sensitivity of 8 copies per reaction. Hence, our new probe-based RT-LAMP assay can serve as an inexpensive method for point-of-need diagnosis of COVID-19 and other infectious diseases.

An engineered CRISPR-Cas12a variant and DNA-RNA hybrid guides enable robust and rapid COVID-19 testing.

Extensive testing is essential to break the transmission of SARS-CoV-2, which causes the ongoing COVID-19 pandemic. Here, we present a CRISPR-based diagnostic assay that is robust to viral genome mutations and temperature, produces results fast, can be applied directly on nasopharyngeal (NP) specimens without RNA purification, and incorporates a human internal control within the same reaction. Specifically, we show that the use of an engineered AsCas12a enzyme enables detection of wildtype and mutated SARS-CoV-2 and allows us to perform the detection step with loop-mediated isothermal amplification (LAMP) at 60-65 °C. We also find that the use of hybrid DNA-RNA guides increases the rate of reaction, enabling our test to be completed within 30 minutes. Utilizing clinical samples from 72 patients with COVID-19 infection and 57 healthy individuals, we demonstrate that our test exhibits a specificity and positive predictive value of 100% with a sensitivity of 50 and 1000 copies per reaction (or 2 and 40 copies per microliter) for purified RNA samples and unpurified NP specimens respectively.

RNA Editing in Human and Mouse Tissues.

Adenosine-to-inosine (A-to-I) RNA editing is a fundamental posttranscriptional mechanism that greatly diversifies the transcriptome in many living organisms, including mammals. Multiple studies have demonstrated the importance of this process not just in normal development and physiology but also in various human diseases. Importantly, the precise editing level of a site may have downstream consequences on cellular behavior. Hence, the editing levels should be quantified as accurately as possible. In this chapter, we describe how to examine RNA editing in human and mouse tissues. The rapid development of next-generation sequencing technologies is affording us an unprecedented ability to accurately measure the editing levels of numerous sites simultaneously. Our experimental workflow includes the harvesting of high-quality RNA samples and the construction of different high-throughput sequencing libraries. We also delineate the computational steps needed to analyze the sequencing data from an Illumina platform.

Determination of isoform-specific RNA structure with nanopore long reads.

Current methods for determining RNA structure with short-read sequencing cannot capture most differences between distinct transcript isoforms. Here we present RNA structure analysis using nanopore sequencing (PORE-cupine), which combines structure probing using chemical modifications with direct long-read RNA sequencing and machine learning to detect secondary structures in cellular RNAs. PORE-cupine also captures global structural features, such as RNA-binding-protein binding sites and reactivity differences at single-nucleotide variants. We show that shared sequences in different transcript isoforms of the same gene can fold into different structures, highlighting the importance of long-read sequencing for obtaining phase information. We also demonstrate that structural differences between transcript isoforms of the same gene lead to differences in translation efficiency. By revealing isoform-specific RNA structure, PORE-cupine will deepen understanding of the role of structures in controlling gene regulation.

A human expression system based on HEK293 for the stable production of recombinant erythropoietin.

Mammalian host cell lines are the preferred expression systems for the manufacture of complex therapeutics and recombinant proteins. However, the most utilized mammalian host systems, namely Chinese hamster ovary (CHO), Sp2/0 and NS0 mouse myeloma cells, can produce glycoproteins with non-human glycans that may potentially illicit immunogenic responses. Hence, we developed a fully human expression system based on HEK293 cells for the stable and high titer production of recombinant proteins by first knocking out GLUL (encoding glutamine synthetase) using CRISPR-Cas9 system. Expression vectors using human GLUL as selection marker were then generated, with recombinant human erythropoietin (EPO) as our model protein. Selection was performed using methionine sulfoximine (MSX) to select for high EPO expression cells. EPO production of up to 92700 U/mL of EPO as analyzed by ELISA or 696 mg/L by densitometry was demonstrated in a 2 L stirred-tank fed batch bioreactor. Mass spectrometry analysis revealed that N-glycosylation of the produced EPO was similar to endogenous human proteins and non-human glycan epitopes were not detected. Collectively, our results highlight the use of a human cellular expression system for the high titer and xenogeneic-free production of EPO and possibly other complex recombinant proteins.

Genome Editing in Mammalian Cell Lines using CRISPR-Cas.

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been successfully repurposed for genome engineering in many different living organisms. Most commonly, the wildtype CRISPR associated 9 (Cas9) or Cas12a endonuclease is used to cleave specific sites in the genome, after which the DNA double-stranded break is repaired via the non-homologous end joining (NHEJ) pathway or the homology-directed repair (HDR) pathway depending on whether a donor template is absent or present respectively. To date, CRISPR systems from different bacterial species have been shown to be capable of performing genome editing in mammalian cells. However, despite the apparent simplicity of the technology, multiple design parameters need to be considered, which often leave users perplexed about how best to carry out their genome editing experiments. Here, we describe a complete workflow from experimental design to identification of cell clones that carry desired DNA modifications, with the goal of facilitating successful execution of genome editing experiments in mammalian cell lines. We highlight key considerations for users to take note of, including the choice of CRISPR system, the spacer length, and the design of a single-stranded oligodeoxynucleotide (ssODN) donor template. We envision that this workflow will be useful for gene knockout studies, disease modeling efforts, or the generation of reporter cell lines.

An Expanded Synthetic Biology Toolkit for Gene Expression Control in Acetobacteraceae.

The availability of different host chassis will greatly expand the range of applications in synthetic biology. Members of the Acetobacteraceae family of Gram-negative bacteria form an attractive class of nonmodel microorganisms that can be exploited to produce industrial chemicals, food and beverage, and biomaterials. One such biomaterial is bacterial cellulose, which is a strong and ultrapure natural polymer used in tissue engineering scaffolds, wound dressings, electronics, food additives, and other products. However, despite the potential of Acetobacteraceae in biotechnology, there has been considerably little effort to fundamentally reprogram the bacteria for enhanced performance. One limiting factor is the lack of a well-characterized, comprehensive toolkit to control expression of genes in biosynthetic pathways and regulatory networks to optimize production and cell viability. Here, we address this shortcoming by building an expanded genetic toolkit for synthetic biology applications in Acetobacteraceae. We characterized the performance of multiple natural and synthetic promoters, ribosome binding sites, terminators, and degradation tags in three different strains, namely, Gluconacetobacter xylinus ATCC 700178, Gluconacetobacter hansenii ATCC 53582, and Komagataeibacter rhaeticus iGEM. Our quantitative data revealed strain-specific and common design rules for the precise control of gene expression in these industrially relevant bacterial species. We further applied our tools to synthesize a biodegradable cellulose-chitin copolymer, adjust the structure of the cellulose film produced, and implement CRISPR interference for ready down-regulation of gene expression. Collectively, our genetic parts will enable the efficient engineering of Acetobacteraceae bacteria for the biomanufacturing of cellulose-based materials and other commercially valuable products.

SRSF9 selectively represses ADAR2-mediated editing of brain-specific sites in primates.

Adenosine-to-inosine (A-to-I) RNA editing displays diverse spatial patterns across different tissues. However, the human genome encodes only two catalytically active editing enzymes (ADAR1 and ADAR2), suggesting that other regulatory factors help shape the editing landscape. Here, we show that the splicing factor SRSF9 selectively controls the editing of many brain-specific sites in primates. SRSF9 is more lowly expressed in the brain than in non-brain tissues. Gene perturbation experiments and minigene analysis of candidate sites demonstrated that SRSF9 could robustly repress A-to-I editing by ADAR2. We found that SRSF9 biochemically interacted with ADAR2 in the nucleus via its RRM2 domain. This interaction required the presence of the RNA substrate and disrupted the formation of ADAR2 dimers. Transcriptome-wide location analysis and RNA sequencing revealed 1328 editing sites that are controlled directly by SRSF9. This regulon is significantly enriched for brain-specific sites. We further uncovered a novel motif in the ADAR2-dependent SRSF9 binding sites and provided evidence that the splicing factor prevents loss of cell viability by inhibiting ADAR2-mediated editing of genes involved in proteostasis, energy metabolism, the cell cycle and DNA repair. Collectively, our results highlight the importance of SRSF9 as an editing regulator and suggest potential roles for other splicing factors.

Rapid Control of Genome Editing in Human Cells by Chemical-Inducible CRISPR-Cas Systems.

Genome editing using programmable DNA endonucleases enables the engineering of eukaryotic cells and living organisms with desirable properties or traits. Among the various molecular scissors that have been developed to date, the most versatile and easy-to-use family of nucleases derives from CRISPR-Cas, which exists naturally as an adaptive immune system in bacteria. Recent advances in the CRISPR-Cas technology have expanded our ability to manipulate complex genomes for myriad biomedical and biotechnological applications. Some of these applications are time-sensitive or demand high spatial precision. Here, we describe the use of an inducible CRISPR-Cas9 system, termed iCas, which we have developed to enable rapid and tight control of genome editing in mammalian cells. The iCas system can be switched on or off as desired through the introduction or removal of the small molecule tamoxifen or its related analogs such as 4-hydroxytamoxifen (4-HT).

Systematic evaluation of CRISPR-Cas systems reveals design principles for genome editing in human cells.

BACKGROUND: While CRISPR-Cas systems hold tremendous potential for engineering the human genome, it is unclear how well each system performs against one another in both non-homologous end joining (NHEJ)-mediated and homology-directed repair (HDR)-mediated genome editing. RESULTS: We systematically compare five different CRISPR-Cas systems in human cells by targeting 90 sites in genes with varying expression levels. For a fair comparison, we select sites that are either perfectly matched or have overlapping seed regions for Cas9 and Cpf1. Besides observing a trade-off between cleavage efficiency and target specificity for these natural endonucleases, we find that the editing activities of the smaller Cas9 enzymes from Staphylococcus aureus (SaCas9) and Neisseria meningitidis (NmCas9) are less affected by gene expression than the other larger Cas proteins. Notably, the Cpf1 nucleases from Acidaminococcus sp. BV3L6 and Lachnospiraceae bacterium ND2006 (AsCpf1 and LbCpf1, respectively) are able to perform precise gene targeting efficiently across multiple genomic loci using single-stranded oligodeoxynucleotide (ssODN) donor templates with homology arms as short as 17 nucleotides. Strikingly, the two Cpf1 nucleases exhibit a preference for ssODNs of the non-target strand sequence, while the popular Cas9 enzyme from Streptococcus pyogenes (SpCas9) exhibits a preference for ssODNs of the target strand sequence instead. Additionally, we find that the HDR efficiencies of Cpf1 and SpCas9 can be further improved by using asymmetric donors with longer arms 5' of the desired DNA changes. CONCLUSIONS: Our work delineates design parameters for each CRISPR-Cas system and will serve as a useful reference for future genome engineering studies.

Dynamic landscape and regulation of RNA editing in mammals.

Adenosine-to-inosine (A-to-I) RNA editing is a conserved post-transcriptional mechanism mediated by ADAR enzymes that diversifies the transcriptome by altering selected nucleotides in RNA molecules. Although many editing sites have recently been discovered, the extent to which most sites are edited and how the editing is regulated in different biological contexts are not fully understood. Here we report dynamic spatiotemporal patterns and new regulators of RNA editing, discovered through an extensive profiling of A-to-I RNA editing in 8,551 human samples (representing 53 body sites from 552 individuals) from the Genotype-Tissue Expression (GTEx) project and in hundreds of other primate and mouse samples. We show that editing levels in non-repetitive coding regions vary more between tissues than editing levels in repetitive regions. Globally, ADAR1 is the primary editor of repetitive sites and ADAR2 is the primary editor of non-repetitive coding sites, whereas the catalytically inactive ADAR3 predominantly acts as an inhibitor of editing. Cross-species analysis of RNA editing in several tissues revealed that species, rather than tissue type, is the primary determinant of editing levels, suggesting stronger cis-directed regulation of RNA editing for most sites, although the small set of conserved coding sites is under stronger trans-regulation. In addition, we curated an extensive set of ADAR1 and ADAR2 targets and showed that many editing sites display distinct tissue-specific regulation by the ADAR enzymes in vivo. Further analysis of the GTEx data revealed several potential regulators of editing, such as AIMP2, which reduces editing in muscles by enhancing the degradation of the ADAR proteins. Collectively, our work provides insights into the complex cis- and trans-regulation of A-to-I editing.

Discovery and engineering of a 1-butanol biosensor in Saccharomyces cerevisiae.

The present study aimed to develop a universal methodology for the discovery of biosensors sensitive to particular stresses or metabolites by using a transcriptome analysis, in order to address the need for in vivo biosensors to drive the engineering of microbial cell factories. The method was successfully applied to the discovery of 1-butanol sensors. In particular, the genome-wide transcriptome profiling of S. cerevisiae exposed to three similar short-chain alcohols, 1-butanol, 1-propanol, and ethanol, identified genes that were differentially expressed only under the treatment of 1-butanol. From these candidates, two promoters that responded specifically to 1-butanol were characterized in a dose-dependent manner and were used to distinguish differences in production levels among different 1-butanol producer strains. This strategy opens up new opportunities for the discovery of promoter-based biosensors and can potentially be used to identify biosensors for any metabolite that causes cellular transcriptomic changes.

Deficiency of microRNA miR-34a expands cell fate potential in pluripotent stem cells.

Embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs) efficiently generate all embryonic cell lineages but rarely generate extraembryonic cell types. We found that microRNA miR-34a deficiency expands the developmental potential of mouse pluripotent stem cells, yielding both embryonic and extraembryonic lineages and strongly inducing MuERV-L (MERVL) endogenous retroviruses, similar to what is seen with features of totipotent two-cell blastomeres. miR-34a restricts the acquisition of expanded cell fate potential in pluripotent stem cells, and it represses MERVL expression through transcriptional regulation, at least in part by targeting the transcription factor Gata2. Our studies reveal a complex molecular network that defines and restricts pluripotent developmental potential in cultured ESCs and iPSCs.

A chemical-inducible CRISPR-Cas9 system for rapid control of genome editing.

CRISPR-Cas9 has emerged as a powerful technology that enables ready modification of the mammalian genome. The ability to modulate Cas9 activity can reduce off-target cleavage and facilitate precise genome engineering. Here we report the development of a Cas9 variant whose activity can be switched on and off in human cells with 4-hydroxytamoxifen (4-HT) by fusing the Cas9 enzyme with the hormone-binding domain of the estrogen receptor (ERT2). The final optimized variant, termed iCas, showed low endonuclease activity without 4-HT but high editing efficiency at multiple loci with the chemical. We also tuned the duration and concentration of 4-HT treatment to reduce off-target genome modification. Additionally, we benchmarked iCas against other chemical-inducible methods and found that it had the fastest on rate and that its activity could be toggled on and off repeatedly. Collectively, these results highlight the utility of iCas for rapid and reversible control of genome-editing function.