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RNA editing

RNA editing is a co- or post-transcriptional modification process in which the nucleotide sequence of an RNA molecule is altered from that encoded by the corresponding DNA template, resulting in changes to the RNA's base composition and potential functional outcomes.^[1] The two predominant types of RNA editing in eukaryotes are adenosine-to-inosine (A-to-I) editing, which converts adenosine (A) to inosine (I) via hydrolytic deamination and is recognized as guanosine (G) during translation, and cytidine-to-uridine (C-to-U) editing, which deaminates cytosine (C) to uridine (U).^[1] These modifications are catalyzed by specific enzyme families: the adenosine deaminase acting on RNA (ADAR) proteins and ADATs for A-to-I events, primarily on double-stranded RNA or transfer RNA (tRNA), and the apolipoprotein B mRNA editing enzyme catalytic polypeptide-like (APOBEC) family for C-to-U events, often on single-stranded RNA substrates.^[1] A-to-I editing is the most widespread form in metazoans, with ADAR enzymes (including ADAR1, ADAR2, and the catalytically inactive ADAR3) targeting structured RNA regions to generate transcriptomic diversity, recode amino acids in proteins, and modulate RNA stability or splicing.^[2] For instance, ADAR1 plays a critical role in innate immunity by editing self double-stranded RNAs to prevent their recognition by sensors like MDA5, thus averting autoimmune responses, while ADAR2 facilitates essential recoding events in neuronal transcripts such as the glutamate receptor subunit GRIA2, changing a glutamine to arginine codon for proper channel function.^[2] In contrast, C-to-U editing is less prevalent but vital in specific contexts, such as APOBEC1-mediated editing of apolipoprotein B mRNA in the intestine to produce a truncated protein isoform, or APOBEC3 family members' roles in antiviral defense by hypermutating viral genomes.^[1] Beyond these core mechanisms, RNA editing expands the proteome without altering the genome, influences evolutionary adaptation through conserved recoding sites under positive selection, and has implications for disease when dysregulated, including neurological disorders, cancer, and immune pathologies.^[2] Recent advances in sequencing and bioinformatics have revealed millions of editing sites across transcriptomes, predominantly in non-coding regions like Alu elements, highlighting its role in fine-tuning gene regulation and inspiring therapeutic applications, such as site-directed editing tools for correcting pathogenic mutations.^[1]

Definition and Biological Significance

Definition of RNA Editing

RNA editing refers to a set of enzymatic processes that modify the nucleotide sequence of primary RNA transcripts after transcription, resulting in RNA molecules that encode proteins or perform functions distinct from those directly specified by the genomic DNA. These modifications can introduce changes that expand transcriptomic and proteomic diversity without altering the underlying DNA sequence. Unlike standard RNA processing events such as splicing, capping, or polyadenylation, RNA editing specifically involves targeted alterations to the RNA sequence itself.^[3] The phenomenon was first discovered in 1986 in the mitochondria of trypanosomes, where extensive insertion of uridine (U) residues into mitochondrial mRNAs was found to be necessary for proper coding of proteins, resolving apparent discrepancies between gene sequences and translated products. This unexpected finding challenged the central dogma of molecular biology and introduced the concept of RNA as a malleable information carrier. In the late 1980s, RNA editing was expanded to mammalian systems with the identification of cytidine-to-uridine (C-to-U) editing in apolipoprotein B (apoB) mRNA in the intestine, which generates a truncated protein isoform essential for lipid metabolism. Adenosine-to-inosine (A-to-I) editing in mammals was subsequently recognized in the early 1990s through studies of brain transcripts, particularly in ion channel mRNAs like those encoding glutamate receptor subunits, where it modulates neuronal excitability. The scope of RNA editing encompasses several types of sequence alterations, primarily base substitutions such as A-to-I and C-to-U, as well as insertion and deletion of nucleotides, particularly U residues in kinetoplastid protists. These processes occur across diverse RNA species, including mRNAs, non-coding RNAs, and viral RNAs, but are distinct from routine posttranscriptional modifications that do not change the sequence information.^[3] Major enzymes driving RNA editing include the adenosine deaminase acting on RNA (ADAR) family, which catalyzes A-to-I substitutions by deaminating adenosine to inosine in double-stranded RNA regions, and the apolipoprotein B mRNA editing enzyme catalytic polypeptide-like (APOBEC) family, particularly APOBEC1, which performs C-to-U editing via cytidine deamination. These enzymes are highly conserved and play pivotal roles in regulating gene expression across eukaryotes.

Role in Gene Expression and Diversity

RNA editing plays a pivotal role in expanding proteomic diversity by enabling post-transcriptional modifications that alter the coding potential of transcripts beyond the constraints of the genomic sequence. Through recoding events, such as codon changes that substitute amino acids in proteins, RNA editing generates protein isoforms with distinct functions, thereby increasing the complexity of the proteome without requiring genomic mutations.^[4] Additionally, editing influences gene expression by regulating alternative splicing, mRNA stability, and translational efficiency; for instance, adenosine-to-inosine (A-to-I) modifications can introduce or remove splice sites, leading to diverse transcript variants, while edited sequences may affect miRNA binding and mRNA decay rates.^[5] In humans, these processes contribute to editing in approximately 1-3% of transcripts, primarily through A-to-I changes mediated by ADAR enzymes.^[6] Biologically, RNA editing exhibits tissue-specific patterns that fine-tune cellular functions, particularly in dynamic environments like the nervous system. In the brain, elevated editing levels support synaptic plasticity by modulating neurotransmitter receptor properties and neuronal excitability, allowing adaptive responses to neural activity.^[7] Editing also facilitates rapid adaptation to environmental stresses, such as oxidative or hypoxic conditions, by altering transcripts involved in metabolic pathways or stress response genes, thereby enhancing cellular resilience without permanent genetic alterations.^[8] In organelles like plant mitochondria and chloroplasts, editing serves as an error-correction mechanism, converting mismatched nucleotides (e.g., C-to-U) to restore functional protein sequences that would otherwise be defective due to mutational biases in organellar genomes.^[9] A prominent example is the Q/R site editing in the glutamate receptor subunit GluA2 (GRIA2), where A-to-I deamination changes a glutamine codon to arginine, rendering AMPA receptors impermeable to calcium ions and preventing excitotoxicity in neurons.^[10] This nearly complete editing (>99%) in mature neurons exemplifies how precise modifications safeguard synaptic function. Genome-wide, ADAR-mediated A-to-I editing targets millions of sites in humans, with over 100 million potential sites in Alu repetitive elements alone, with editing occurring in nearly all adenosines within these editable Alu repeats, though often at low levels (<1%), and affecting a majority of genes containing such elements (67.4% of RefSeq genes).^[11] From an evolutionary perspective, RNA editing confers adaptive advantages by providing a flexible layer of transcriptome plasticity, enabling organisms to respond to changing environments or developmental cues without the fixation of potentially deleterious mutations in the genome. This mechanism is particularly evident in species facing variable conditions, where edited transcripts support physiological acclimation and increased genetic diversity at the RNA level.^[12]

Detection Methods

Next-Generation Sequencing

Next-generation sequencing (NGS) enables the genome-wide identification and quantification of RNA editing sites by aligning RNA-seq reads to a reference genome or matched genomic DNA sequence, detecting nucleotide mismatches that indicate editing events. For A-to-I editing, inosine is reverse-transcribed and sequenced as guanosine, appearing as A-to-G mismatches, while C-to-U editing manifests as C-to-T mismatches. This comparative approach distinguishes editing from genomic variants by requiring RNA-DNA discordance in the same sample. However, C-to-U editing detection faces additional challenges due to its lower prevalence and potential overlap with common SNPs, often requiring higher stringency filters.^[13] Standard protocols involve performing RNA-seq on poly(A)-selected or total RNA libraries with deep sequencing coverage, typically exceeding 100x per site to ensure sufficient read depth for reliable variant calling and to minimize stochastic errors. Specialized bioinformatics tools, such as REDItools and GIREMI, process aligned BAM files to annotate candidate sites, apply filters for strand bias, mapping quality, and allelic fraction thresholds (e.g., >0.01 for minor alleles), and output editing events while excluding repetitive regions or low-complexity sequences. These tools integrate statistical tests, like mutual information in GIREMI, to prioritize true positives without requiring multiple replicates initially.^[14]^[15] NGS offers high-throughput analysis, allowing cost-effective discovery of thousands to millions of editing sites across transcriptomes in a single experiment, and supports single-cell resolution through scRNA-seq to reveal cell-type-specific editing patterns. However, challenges include distinguishing true editing from single nucleotide polymorphisms (SNPs), sequencing artifacts, or alignment errors, particularly in polymorphic or repetitive genomic regions. Solutions involve using matched DNA-seq for validation, biological replicates to assess reproducibility, and statistical models—such as those estimating false discovery rates or employing generalized linear models—to correct for biases and filter false positives.^[14]^[16] The widespread adoption of NGS for RNA editing detection accelerated post-2010, driven by Illumina platforms that enabled scalable RNA-seq experiments, leading to the identification of over 2 million human A-to-I sites by 2013 through refined pipelines. NGS is often complemented by mass spectrometry for orthogonal validation.^[15]^[17]

Mass Spectrometry

Mass spectrometry (MS) serves as a powerful orthogonal method for validating RNA editing events by directly analyzing either modified nucleosides in RNA or the resulting altered amino acids in proteins derived from edited transcripts. At the nucleotide level, the principle involves enzymatic digestion of RNA into individual nucleosides, followed by liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS), where inosine (arising from A-to-I editing) is identified and quantified based on its unique mass-to-charge ratio (m/z) and chromatographic retention time relative to standard curves.^[18] This approach distinguishes inosine from adenosine, enabling precise measurement of editing efficiency in specific RNA populations, such as mRNA or viral RNA.^[19] At the protein level, MS targets tryptic peptides from proteomes, detecting recoding events like the Q/R site in AMPA receptor subunit GluA2, where A-to-I editing converts a glutamine codon to arginine, producing a detectable mass shift in the peptide spectrum.^[20] Key techniques include nanoLC-MS/MS for proteome-wide scans, which employs high-resolution instruments like the Orbitrap to sequence peptides from complex samples, often using tandem mass tag (TMT) labeling for multiplexed quantification across multiple samples.^[20] For inosine-specific quantification, methods like nucleic acid isotope labeling coupled mass spectrometry (NAIL-MS) incorporate stable isotopes during cell culture to track modification dynamics, allowing differentiation of newly synthesized versus pre-existing edited RNAs with temporal resolution.^[21] These approaches have been refined post-2015 for higher throughput; for instance, custom proteogenomic databases integrate RNA editing variants to match MS spectra, identifying edited peptides in human brain tissue from over 170 subjects.^[20] Similarly, advanced LC-MS/MS protocols with chemical derivatization have enabled detection of low-abundance inosine in single cells or yeast mRNA.^[22] The primary advantages of MS-based validation lie in its ability to confirm the functional consequences of RNA editing, such as amino acid substitutions that alter protein properties (e.g., calcium permeability in GluA2-edited AMPA receptors), providing direct evidence beyond nucleotide-level observations.^[20] It complements next-generation sequencing by verifying editing at the translational output, with high specificity for modified species.^[18] However, limitations include low sensitivity for rare editing events, often necessitating enrichment strategies like immunoprecipitation of specific RNAs or peptides, as seen in the sparse detection of only 10 out of 294 predicted recoding events in brain proteomes.^[20] Sample preparation challenges, such as RNA instability and ion suppression in complex matrices, further require rigorous controls.^[19] Post-2015 multiplexing innovations, including TMT and improved ionization, have enhanced scalability for validating editing impacts in aging or disease contexts, such as Alzheimer's brain tissue.^[20]

Emerging Techniques

Recent advances in RNA editing detection have introduced chemically-assisted sequencing methods that enable enzyme-free detection of A-to-I edits through bisulfite-like assays, which chemically modify inosine bases to distinguish them from adenosine without relying on enzymatic treatments that can introduce biases.^[18] These approaches, detailed in a 2025 review, improve specificity for non-canonical edits by leveraging reactivity differences, allowing for more accurate mapping in complex transcriptomes.^[18] Building on foundations from next-generation sequencing and mass spectrometry, such methods address longstanding gaps in sensitivity for low-frequency events.^[23] Nanopore direct RNA sequencing has emerged as a key tool for real-time detection of inosine reads, preserving native RNA structure to identify A-to-I editing sites without reverse transcription artifacts.^[24] The Dinopore algorithm, developed for nanopore platforms, uses signal patterns to pinpoint inosine with high precision across organisms, facilitating the study of editing dynamics in real time.^[25] A 2025 analysis highlights how these advancements in direct RNA nanopore sequencing enhance the detection of RNA modifications, including editing events, by integrating machine learning to classify base-level signals.^[26] In 2024, developments in AI-driven prediction tools have advanced site annotation for RNA editing, with models like those using fine-tuned GPT architectures predicting edit sites by analyzing sequence context and secondary structure features.^[27] These tools, building on convolutional neural networks such as EditPredict, achieve superior annotation accuracy for potential editing hotspots, aiding in the prioritization of candidates from large datasets.^[28] Such predictive frameworks are particularly valuable for identifying non-model organism edits where experimental validation is resource-intensive.^[29] Complementing this, 2025 multi-omics integration efforts have enhanced understanding of RNA editing by combining RNA-seq with epigenomic and proteomic data, revealing roles in tumor evolution.^[30] These emerging techniques offer substantial benefits, including enhanced accuracy for detecting low-abundance edits in cancer and brain tissues, where traditional methods often struggle with noise.^[24] By minimizing artifacts, they have been shown to substantially reduce false positives, enabling more reliable profiling of editing landscapes in disease states.^[31]

Core Mechanisms of RNA Editing

Deamination-Based Substitution

Deamination-based substitution represents the predominant mechanism of RNA editing in metazoans, involving the enzymatic removal of an amino group from specific nucleotide bases, which alters their base-pairing properties and can change codon meaning during translation.^[32] This process primarily targets cytidine (C) or adenosine (A) residues, converting them to uridine (U) or inosine (I), respectively, through hydrolytic deamination catalyzed by members of the metazoan-specific APOBEC family for C-to-U editing and the ADAR family for A-to-I editing.^[1] These modifications expand the proteome diversity without altering the genomic sequence, influencing gene expression, protein function, and cellular responses.^[33] The core reaction proceeds via hydrolysis, where water acts as the nucleophile to deaminate the base: for example, adenosine is converted to inosine plus ammonia (Adenosine + H₂O → Inosine + NH₃).^[32] This enzymatic activity requires double-stranded RNA (dsRNA) substrates, as the editing enzymes recognize structured RNA regions to ensure specificity and avoid off-target effects.^[34] ADAR enzymes, in particular, feature double-stranded RNA-binding domains (dsRBDs) that facilitate substrate recognition, followed by a catalytic deaminase domain that executes the deamination through a two-step mechanism involving a tetrahedral intermediate.^[32] APOBEC enzymes similarly rely on RNA structure for positioning but often require auxiliary factors for efficient activity in vivo.^[33] Deamination-based editing accounts for the vast majority of known RNA editing events in eukaryotic transcriptomes, far outnumbering other substitution or modification types.^[33] Editing efficiency and site selection are tightly regulated by factors such as the enzyme's subcellular localization—ADARs are primarily nuclear, though ADAR1 has a cytoplasmic isoform, while certain APOBECs exhibit variable nuclear or cytoplasmic distribution—and the local RNA secondary structure, which can enhance or inhibit access to target sites.^[35] These regulatory elements ensure that editing occurs in a controlled manner, often in response to cellular stress or developmental cues.^[1] Specific instances of C-to-U and A-to-I editing, such as those mediated by APOBEC1 in apolipoprotein B mRNA or ADAR2 in glutamate receptor transcripts, exemplify this mechanism's role in fine-tuning protein isoforms.^[33]

Insertion and Deletion Editing

Insertion and deletion editing represents a form of RNA modification that alters the length of the transcript by adding or removing uridine (U) residues, in contrast to substitution editing that changes individual bases without affecting length. This process is primarily observed in the mitochondria of kinetoplastid protists, such as trypanosomes, where it is directed by trans-acting guide RNAs (gRNAs).^[36]^[37] The mechanism involves the formation of a chimeric duplex between the pre-mRNA and a gRNA, which base-pairs with the target mRNA to specify editing sites through complementary regions. An endonuclease cleaves the mRNA at mismatched sites within this duplex, creating 5' and 3' fragments. For U insertion, the RET1 enzyme (a 3' terminal uridylyl transferase, or TUTase) adds U residues to the 3' end of the upstream fragment in a non-templated manner, with the number of Us determined by the gRNA anchor sequence to restore complementarity; the fragments are then joined by RNA ligase REL2. In U deletion, mismatched Us are removed from the 3' fragment by the 3'-5' exonuclease activity associated with the REH1 helicase, followed by re-ligation via REL1. This cycle progresses in a 3' to 5' direction along the mRNA, often requiring multiple gRNAs for extensive editing, and can involve the addition of poly-U tails during insertion steps that are subsequently trimmed to match the gRNA template. Up to over 200 Us may be inserted in a single transcript, such as in the cytochrome c oxidase subunit III (COIII) mRNA of Trypanosoma brucei, to generate functional open reading frames from cryptic pre-mRNAs.^[38]^[39]^[40] This type of editing is rare among eukaryotes and is essential for the expression of most mitochondrial genes in trypanosomes, where it corrects frameshifts and restores translatable sequences in up to 12 maxicircle-encoded mRNAs. No direct homologs of the editing machinery exist in mammals or other higher eukaryotes. Exceptions include limited U insertion events reported in dinoflagellate mitochondria, which primarily feature substitutional editing but occasionally add internal Us to refine transcripts. Additionally, some cases of physical editing occur via trans-splicing in certain protists, where RNA fragments are joined to effectively insert sequences, though this differs from the gRNA-templated U modifications in kinetoplastids.^[40]^[41]00468-0)

Deamination-Based Editing

C-to-U Editing

C-to-U RNA editing is a post-transcriptional modification in which cytidine residues in RNA transcripts are deaminated to uridine, altering the genetic code without changing the underlying DNA sequence. This process occurs predominantly in nuclear-encoded messenger RNAs in animals and in organellar transcripts in plants, contributing to protein diversity and functional optimization. The reaction is catalyzed by cytidine deaminases and is highly site-specific, often guided by auxiliary factors that recognize particular RNA motifs.^[42] In mammalian systems, the primary enzyme responsible for C-to-U editing is APOBEC1, a cytoplasmic cytidine deaminase that requires the cofactor APOBEC1 complementation factor (ACF) to form an active editing complex. APOBEC1 deaminates specific cytidines in target RNAs, with editing efficiency reaching over 90% in tissues like the small intestine. Site specificity is achieved through an 11-nucleotide mooring sequence located 3-5 nucleotides downstream of the edited cytidine, typically featuring a core motif of 5'-UUUN-3' that positions the enzyme correctly. A canonical example is the editing of apolipoprotein B (APOB) mRNA, where deamination of cytidine 6666 (C6666) to uridine creates a premature stop codon (UAA), truncating the full-length ApoB100 protein into the shorter ApoB48 isoform essential for lipid transport in the intestine.^[43]^[44]^[45] In plant mitochondria and plastids, C-to-U editing is mediated by pentatricopeptide repeat (PPR) proteins of the PLS subclass, which contain a C-terminal DYW deaminase domain responsible for the deamination activity. These DYW-type enzymes recognize cis-elements in organellar RNAs via the PPR array and catalyze editing at hundreds of sites, often restoring evolutionarily conserved amino acids or creating functional codons. For instance, in plastids, editing converts an ACG initiation codon to AUG in the psbL mRNA of tobacco, enabling translation of the photosystem II subunit PsbL. In mitochondria, similar editing events optimize codons for protein function, with over 400 sites identified in species like Arabidopsis thaliana.^[46]^[47]^[48] C-to-U editing is not typically observed in animal mitochondria, including humans, unlike the extensive editing in plant mitochondria and plastids. Recent research (as of 2025) has revealed cancer-specific hyper-editing patterns driven by APOBEC family members, such as elevated C-to-U modifications in hematologic malignancies that promote tumor progression through altered protein isoforms and increased mutational load. As of 2025, APOBEC3A has been shown to catalyze site-specific C-to-U editing of transfer RNAs (tRNAs), with implications for cancer and immune responses.^[8]^[49]^[50]

A-to-I Editing

A-to-I RNA editing, the conversion of adenosine (A) to inosine (I) in RNA, is the most prevalent form of RNA editing in animals and is catalyzed by the adenosine deaminase acting on RNA (ADAR) family of enzymes. These enzymes include ADAR1, ADAR2, and ADAR3, each with distinct expression patterns and functions. ADAR1 exists in two isoforms: the constitutively expressed nuclear p110 form and the interferon-inducible cytoplasmic p150 form, which is upregulated during immune responses. ADAR2 is primarily nuclear and responsible for constitutive editing events in specific transcripts. In contrast, ADAR3 lacks deaminase activity and acts as an inhibitor of editing by competing for double-stranded RNA (dsRNA) substrates.^[51] The editing process involves hydrolytic deamination of adenosine within dsRNA structures, where inosine is subsequently recognized as guanosine (G) during translation and splicing, potentially altering codon meaning or RNA stability. For instance, in messenger RNAs (mRNAs), this can lead to amino acid recoding; a prominent example is the Q/R site in the GRIA2 transcript encoding the AMPA receptor subunit GluA2, where editing changes CAG (glutamine) to CIG (read as CGG, arginine), modulating calcium permeability in neuronal synapses. Humans harbor over 100 confirmed recoding sites in mRNAs, many affecting neurotransmitter receptors and ion channels critical for brain function. In non-coding RNAs, particularly those with Alu repetitive elements, ADARs perform hyper-editing, extensively modifying adenosines to stabilize structures or prevent immune sensing of dsRNA.^[52] Functionally, A-to-I editing diversifies the proteome and regulates innate immunity; ADAR1 p150 edits viral dsRNA to evade interferon responses, thereby inhibiting excessive inflammation.^[53] In cancer, recent transcriptome-wide mapping has revealed dysregulated editing patterns, such as elevated ADAR1 activity promoting tumor immune evasion and progression in hematologic malignancies, highlighting therapeutic potential through ADAR modulation. As of 2024, advances show A-to-I editing's role in oncogenesis, including non-synonymous mutations in tumor progression.^[54]

Editing in Specific RNA Types

Messenger RNA Editing

Messenger RNA (mRNA) editing, predominantly mediated by deamination-based mechanisms such as A-to-I conversion, enables post-transcriptional diversification of the proteome by altering codon sequences in protein-coding transcripts. This process occurs in a majority of human protein-coding genes, with editing events being particularly prevalent in the brain, where they contribute to neuronal plasticity and function.^[11] For instance, the serotonin receptor 2C (HTR2C) transcript undergoes extensive A-to-I editing at five sites within its coding region, generating up to 24 distinct isoforms that modulate receptor signaling and G-protein coupling efficiency.^[55]^[56]^[57] The primary functions of mRNA editing include recoding amino acids to produce protein isoforms with modified properties, such as altered ion selectivity or channel kinetics in voltage-gated ion channels. For example, editing in subunits of potassium channels like KCNA1 changes an isoleucine to valine, altering recovery from inactivation and fine-tuning neuronal excitability.^[58] Additionally, exonic edits can regulate alternative splicing by introducing or disrupting exonic splicing enhancer motifs, thereby influencing isoform production and gene expression patterns.^[59]^[60]^[61] A canonical example is the glutamine/arginine (Q/R) site in the GluA2 subunit of AMPA receptors, where nearly complete A-to-I editing converts a CAG codon to CIG (read as CGG), replacing glutamine with arginine in the channel pore and preventing calcium influx, which is essential for synaptic transmission and neuroprotection. In diseases like amyotrophic lateral sclerosis (ALS), reduced ADAR2 activity leads to incomplete editing at this site, resulting in calcium-permeable AMPA receptors that promote motor neuron degeneration through excitotoxicity.^[10]^[62]

Transfer RNA Editing

Transfer RNA (tRNA) editing primarily involves post-transcriptional modifications that ensure accurate translation by refining anticodon functionality and wobble base pairing. One key type is C-to-U editing in anticodon loops, as observed in mitochondrial tRNAs where it alters codon recognition; for instance, in marsupial mitochondria, C-to-U editing at position 35 of the anticodon converts a tRNA to decode aspartate codons instead of a mismatched sequence.^[63] Another prominent type is A-to-I editing at the wobble position (position 34), which expands codon recognition by allowing inosine to pair with U, C, or A in the third codon position, thereby enabling a single tRNA to decode multiple synonymous codons.^[64] This A-to-I process is analogous to deamination-based editing in messenger RNA but is specialized for tRNA anticodon integrity.^[65] The enzymes mediating these edits are highly specific. For A-to-I editing, the ADAT complex—comprising ADAT1 (homodimeric, akin to yeast Tad1), ADAT2, and ADAT3 (forming a heterodimer)—catalyzes deamination at wobble and adjacent positions in various tRNAs, with ADAT2/3 targeting up to eight tRNA species in eukaryotes.^[66] These enzymes act independently or in complexes, recognizing tRNA structural motifs without requiring guide RNAs.^[67] tRNA editing serves critical functions, including correction of genomic encoding errors in tRNA genes and facilitation of the wobble hypothesis for degenerate codon decoding. By introducing inosine at the wobble position, editing compensates for limitations in tRNA gene diversity, allowing efficient translation of the full codon repertoire without requiring additional tRNA isoacceptors.^[68] This process is essential for viability in yeast, where Tad1/2/3 mutants exhibit severe growth defects due to impaired tRNA function, and in mammals, where ADAT disruptions lead to translational inefficiencies.^[69] Illustrative examples highlight tRNA editing's biological impact. In Escherichia coli, the TadA enzyme (a bacterial ADAT homolog) performs A-to-I editing at the wobble position of tRNA^Arg(ACG), forming inosine-34 essential for arginine codon decoding and cell viability.^[70] In humans, defects in mitochondrial tRNA modification due to mutations in tRNA genes, such as impaired wobble base modification in tRNA^Lys and tRNA^Leu, underlie diseases like myoclonic epilepsy with ragged-red fibers (MERRF) and mitochondrial encephalomyopathy, lactic acidosis, and stroke-like episodes (MELAS), where disruptions affect anticodon integrity and oxidative phosphorylation.^[71]^[72] These cases underscore tRNA editing's and modifications' role in preventing translational errors that propagate to cellular dysfunction.

Ribosomal RNA Editing

In eukaryotes, sequence-altering RNA editing in ribosomal RNA (rRNA) is rare compared to the abundant post-transcriptional chemical modifications that support ribosome biogenesis and function. While modifications such as pseudouridylation (isomerization of uridine to pseudouridine) and 2'-O-methylation enhance rRNA stability, folding, and translational efficiency, they do not change the nucleotide sequence and are distinct from editing.^[73] Rare instances of A-to-I deamination have been suggested in rRNA expansion segments, but this is not a primary mechanism and lacks widespread confirmation in cytoplasmic rRNA. Sequence editing is more prominent in organellar rRNAs, such as C-to-U changes in plant mitochondria and plastids (covered in the organelle editing section).^[2] The enzymes for rRNA modifications, though not editing, are organized into small nucleolar ribonucleoprotein (snoRNP) complexes. Box H/ACA snoRNPs guide pseudouridylation via the dyskerin (Cbf5 in yeast) synthase, and box C/D snoRNPs direct 2'-O-methylation by fibrillarin (Nop1 in yeast), using complementary snoRNA sequences for site specificity. Defects in these modification processes are linked to ribosomopathies, such as dyskeratosis congenita from DKC1 mutations impairing pseudouridylation, leading to telomere shortening, bone marrow failure, and premature aging.^[74]^[75]

Editing in Organelles and Viruses

Plant Mitochondrial and Plastid Editing

In plant mitochondria and plastids, RNA editing primarily involves cytidine-to-uridine (C-to-U) deamination, serving as a post-transcriptional mechanism to compensate for mutations in organellar genomes. In Arabidopsis thaliana, over 400 such sites have been identified in mitochondrial transcripts, with 456 C-to-U conversions reported exclusively in mRNAs, including 441 within open reading frames (ORFs), eight in introns, and seven in untranslated regions. Chloroplasts exhibit far fewer sites, with approximately 41 C-to-U edits detected in A. thaliana transcripts. While C-to-U editing predominates, rare U-to-C reversals occur in some land plants, though none were found in A. thaliana organelles. These edits are highly specific, occurring almost exclusively at conserved positions to restore evolutionarily conserved protein sequences. The editing machinery relies on pentatricopeptide repeat (PPR) proteins, particularly those of the PLS subclass containing DYW domains, which act as site-specific trans-factors by binding target RNAs via their RNA-recognition motifs. These PPR-DYW proteins recruit cytidine deaminases to catalyze the C-to-U conversion, often in complex with cofactors such as multiple organellar RNA-editing factors (MORFs) and RNA-editing interacting proteins (RIPs), which enhance editing efficiency through protein-protein interactions. For instance, MORF proteins stabilize PPR-RNA complexes and facilitate deaminase activity, while RIPs may bridge interactions between editing factors. This coordinated system ensures precise editing without off-target effects, reflecting the nuclear-encoded control over organellar RNA processing. Functionally, these edits correct mutations accumulated in organellar genomes over evolution, often restoring canonical amino acids essential for protein function; notable examples include the creation of AUG initiation codons or elimination of premature stop codons (UAG to UGG, leucine). Editing is also tissue-specific, with varying efficiencies observed across developmental stages or organs, potentially fine-tuning gene expression under environmental cues. In maize (Zea mays) chloroplasts, RNA editing is integral to the trans-splicing of the rps12 pre-mRNA, where PPR protein ZmPPR4 facilitates intron assembly and subsequent editing to produce functional ribosomal protein S12. The loss of editing sites in carnivorous plants, such as Drosera rotundifolia and Nepenthes ventricosa, correlates with organellar genome restructuring, including intron losses and reduced editing to only six sites in plastids, suggesting compensatory genomic changes in nutrient-stressed lineages.

Viral RNA Editing

Viral RNA editing primarily involves modifications to the viral genome or transcripts by host cellular enzymes, enabling viruses to adapt during replication and interact with host defenses. One key type is A-to-I editing mediated by host adenosine deaminases acting on RNA (ADAR), particularly ADAR1, which targets double-stranded regions in viral RNAs. This editing introduces inosine, read as guanosine during translation, potentially altering viral protein sequences and reducing the immunogenicity of viral double-stranded RNA intermediates. In contrast, C-to-U editing in coronaviruses like SARS-CoV-2 is driven by host APOBEC family deaminases, such as APOBEC3A, which convert cytosine to uracil in single-stranded viral RNA, leading to hypermutation patterns observed in emerging variants.^[76]^[77]^[78] These editing events serve critical functions in viral pathogenesis, including the generation of protein isoforms that enhance replication or evade immunity. A-to-I editing can disrupt viral gene expression to limit over-replication or create variants with altered antigenicity, while C-to-U hypermutation promotes genetic diversity akin to antigenic variation in other RNA viruses. Both types contribute to immune evasion by attenuating the host interferon response; for instance, ADAR1-mediated editing of viral RNAs prevents excessive activation of pattern recognition receptors like MDA5, thereby dampening type I interferon production and allowing persistent infection. In some cases, viruses exploit this to reduce antiviral signaling, as seen in hyper-edited defective interfering RNAs that sequester host sensors without productive replication.^[79]^[80]^[77] Notable examples illustrate host-virus dynamics in RNA editing. In measles virus, persistent infections in the brain feature extensive A-to-I hyper-editing by ADAR1, which suppresses viral replication and cytotoxicity but can be hijacked to facilitate long-term persistence by editing non-structural proteins. Similarly, in HIV-1, ADAR1 edits adenosines in the 5' untranslated region, as well as the Tat and Rev coding sequences, influencing nuclear export of unspliced viral transcripts and promoting virion production through modified protein isoforms. For SARS-CoV-2, 2024 analyses of variants revealed recurrent C-to-U editing sites biased toward single-stranded regions, driven by host APOBEC3A, which accelerates mutational instability and contributes to the evolution of immune-escape variants like Omicron sublineages. This interplay highlights ADAR1's dual role: it restricts viral spread by editing to inactivate genomes but enables evasion when viruses induce or tolerate editing, underscoring editing as a battleground in host-virus conflict.^[76]^[81]^[82]^[83]^[84]

Evolutionary Origins and Implications

Proposed Origins

One prominent hypothesis posits that RNA editing represents a vestige of the ancient RNA world, where self-replicating RNA molecules required post-transcriptional correction mechanisms to mitigate errors from inherently error-prone replication processes. In this view, enzymes like adenosine deaminases acting on RNA (ADARs) repurposed ancient RNA-modifying activities—originally involved in RNA catalysis or repair—for modern editing functions, allowing persistence of these "old players" in eukaryotic genomes. This theory aligns with the broader RNA world paradigm, suggesting that editing evolved as a legacy of primordial nucleic acid fidelity challenges before the emergence of DNA-based genomes. A complementary proposal frames RNA editing as a compensatory mechanism to alleviate mutational load in organelles, particularly mitochondria and plastids, which exhibit high mutation rates due to limited recombination and exposure to reactive oxygen species. In plant organelles, C-to-U editing frequently restores conserved amino acids at critical codon positions, counteracting deleterious genomic changes and maintaining protein functionality, such as hydrophobicity in membrane-bound complexes. This adaptive role is supported by the selective fixation of editing sites over evolutionary time, driven by functional constraints rather than neutral drift.^[85]^[86] Evidence for the deep antiquity of RNA editing comes from its presence in kinetoplastids, a basal eukaryotic lineage, where U-insertion/deletion editing likely originated in a common ancestor following the divergence from euglenoids over a billion years ago. Comparative analyses reveal that guide RNA-encoding minicircles and productively edited transcripts, such as those for ND8 and cytochrome oxidase subunits, are conserved across kinetoplastid subgroups, indicating an early innovation predating more specialized forms like extensive pan-editing. In plants, C-to-U editing shows signs of convergent evolution, independently arising in mitochondrial and plastid genomes to address similar mutational pressures, despite phylogenetic distance from kinetoplastid systems.^[41]^[87] Key proposals from the 1990s further suggest that A-to-I editing by ADARs arose via a "mistaken identity" mechanism, where deamination enzymes, evolved for antiviral defense against double-stranded RNA, inadvertently targeted cellular transcripts, leading to functional recoding events. The antiquity of RNA editing is underscored by conserved sites across eukaryotic clades, with estimates placing the last eukaryotic common ancestor approximately 1.8–2 billion years ago.^[88]

Evolutionary Conservation and Diversity

RNA editing mechanisms exhibit varying degrees of evolutionary conservation across eukaryotic lineages, reflecting their ancient origins while adapting to lineage-specific pressures. The core adenosine deaminase acting on RNA (ADAR) enzymes, responsible for A-to-I editing, are highly conserved among metazoans, with structural and functional similarities preserved from invertebrates to vertebrates, underscoring their essential role in neural function and development.^[89] In contrast, pentatricopeptide repeat (PPR) protein-mediated C-to-U editing in plant organelles shows significant loss in certain angiosperm lineages, where editing sites have diminished over time, accompanied by a reduction in PPR gene numbers, suggesting relaxed selective constraints in these advanced flowering plants.^[90] Diversity in RNA editing is evident in unique forms restricted to specific eukaryotic groups, highlighting independent evolutionary innovations. U-insertion/deletion editing, which extensively modifies mitochondrial mRNAs by adding or removing uridines, is characteristic of kinetoplastids within the excavate protists, enabling the restoration of functional coding sequences from highly derived genomes and distinguishing this mechanism from point-mutation editing prevalent elsewhere.^[91] Recent discoveries have revealed non-canonical A-to-I editing in fungi, independent of metazoan ADARs and mediated by tRNA-specific adenosine deaminases during sexual reproduction, allowing for adaptive responses such as antiviral defense through modulation of nearby gene expression.^[92] Evolutionary drivers of RNA editing vary by context and genomic region, balancing adaptive benefits with stochastic processes. In brain-expressed genes, nonsynonymous A-to-I editing sites show signatures of positive selection, particularly in Drosophila, where they enhance protein diversity and neural adaptability, contributing to lineage-specific traits.^[93] Conversely, the abundance of editing in non-coding regions, such as introns and repetitive elements, is often attributed to neutral drift, where off-target deaminase activity accumulates without strong purifying selection, potentially serving as a reservoir for future functional innovations.^[94] This duality may facilitate speciation, as editing-generated transcriptomic variability can promote reproductive isolation and ecological divergence, exemplified by elevated editing rates in species with recent adaptive radiations.^[95] Comparative genomic analyses further illuminate how editing sites correlate with mobile genetic elements, influencing their distribution and evolution. In primates, a substantial proportion of A-to-I editing events occur within Alu transposon insertions, where inverted Alu pairs create double-stranded RNA substrates that attract ADAR enzymes, leading to hyper-editing that shapes transcriptome diversity and potentially drives primate-specific adaptations.^[96]

Therapeutic Applications

Enzyme-Based RNA Editing Therapies

Enzyme-based RNA editing therapies harness endogenous cellular enzymes, such as ADAR for adenosine-to-inosine (A-to-I) editing and APOBEC for cytidine-to-uridine (C-to-U) editing, by delivering synthetic guide RNAs or engineered recruiters to direct site-specific modifications in target mRNAs. These approaches avoid direct genome alteration, focusing instead on transient RNA-level corrections that can restore protein function in genetic disorders. For A-to-I editing, platforms recruit the cell's own ADAR enzymes using partially double-stranded guide RNAs that anchor to the target transcript, enabling precise deamination without introducing foreign proteins. Wave Life Sciences' PRISM platform exemplifies this, utilizing chemically optimized oligonucleotides delivered subcutaneously to activate endogenous ADAR for editing the SERPINA1 mRNA in alpha-1 antitrypsin deficiency (AATD), a condition causing lung and liver damage due to misfolded protein accumulation.^[97]^[98] Therapeutic targets include monogenic diseases amenable to single-nucleotide corrections or exon modulation. In AATD, Wave's WVE-006 achieved the first demonstration of therapeutic RNA editing in humans during the phase 1b/2a RestorAATion-2 trial, where a 200 mg multidose regimen resulted in 7.2 μM wild-type M-AAT (64.4% of total AAT) with 60.3% reduction in mutant Z-AAT, sustained for at least 2 months; a single 400 mg dose yielded 5.3 μM M-AAT (47.2% of total AAT) with 49% Z-AAT reduction. As of September 2025, the therapy was well-tolerated with no serious adverse events; multidose expansion data expected in Q1 2026, and as of Q3 2025, the trial supports monthly or less frequent subcutaneous dosing with a favorable safety profile.^[99]^[100] For Duchenne muscular dystrophy (DMD), preclinical ADAR recruitment has corrected point mutations in the DMD gene, such as the missense 1682G>A variant, restoring dystrophin expression in mouse models via A-to-I editing to revert the codon, potentially addressing cases caused by such mutations.^[101] These therapies offer key advantages, including reversibility—edits last only as long as the RNA persists, typically days to weeks—and avoidance of DNA off-target effects, minimizing risks of permanent mutations or insertional mutagenesis seen in DNA editing. Delivery systems like LNPs for systemic administration or adeno-associated viruses (AAV) for tissue-specific targeting enhance accessibility, with LNPs enabling non-invasive subcutaneous dosing as in WVE-006. However, challenges persist, notably variable editing efficiencies in vivo due to suboptimal ADAR recruitment and guide RNA stability, though recent trials show levels up to 64%. Immune responses to synthetic guides, including innate activation via Toll-like receptors, can reduce efficacy and cause inflammation, as observed in early oligonucleotide trials. Specificity remains a hurdle, with off-target A-to-I edits potentially altering unintended transcripts, necessitating advanced chemical modifications like 2'-O-methyl substitutions to improve precision. Ongoing optimizations, such as AI-guided guide design, aim to address these to advance clinical viability.^[102]^[103]^[97]^[98]

CRISPR-Based RNA Editing Tools

CRISPR-based RNA editing tools leverage the programmable RNA-targeting capabilities of Cas13 enzymes, which are fused to deaminase domains to enable precise base conversions without altering the genomic DNA. These systems emerged as a safer alternative to DNA editing by allowing transient, reversible modifications at the RNA level, building briefly on native enzyme mechanisms like ADAR for adenine deamination. Unlike traditional CRISPR-Cas9, which cleaves DNA, Cas13 binds and edits single-stranded RNA transcripts guided by CRISPR RNAs (crRNAs), minimizing off-target effects and immunogenicity risks.^[104] Key systems include the RNA Editing for Programmable A to I Replacement (REPAIR), which uses a catalytically dead Cas13b (dCas13b) fused to the deaminase domain of human ADAR2 to achieve adenosine-to-inosine (A-to-I) editing. This fusion protein, guided by crRNA, recruits ADAR2 to target sites without sequence constraints beyond the protospacer flanking sequence, enabling correction of disease-associated mutations in transcripts like those for β-thalassemia.^[104] For combined knockdown and editing, Cas13d variants, such as those in the CasRx system, provide robust RNA cleavage alongside deaminase fusions; Cas13d's smaller size (~775 amino acids) facilitates delivery, and it has been adapted for isoform-specific targeting in mammalian cells, achieving up to 90% knockdown efficiency in primary T cells. Systems like LEAPER advance A-to-I editing using RNA-templated recruitment of ADAR without Cas13 dependency; for U-to-C, RESCUE uses Cas13b with engineered APOBEC1, demonstrating high specificity in vivo.^[105]^[106] Recent developments focus on engineering more compact and versatile Cas13 orthologs to improve delivery and multiplexing. Evolved variants like Cas13X and Cas13Y, discovered from uncultivated microbes, are the smallest in the family at 775–800 amino acids, reducing payload size for viral vectors and enabling RNA interference or base editing in hard-to-transfect cells. These compact enzymes support multiplex editing in vivo, as shown in a 2024 Cas13d platform that simultaneously regulates dozens of transcripts in mouse models via arrayed crRNAs, achieving coordinated knockdown of immune checkpoints without genomic integration.^[107] Such advancements postdate earlier Cas7-11 explorations, filling gaps in scalable, tissue-specific applications. In therapeutic contexts, these tools show promise in cancer immunotherapy by tuning immune responses; for instance, Cas13-mediated editing of PD-1 transcripts in T cells reduces inhibitory signaling, enhancing antitumor activity in preclinical models.^[108] A 2024 Stanford-developed Cas13d platform enables metabolic tuning in immune cells by regulating glycolytic enzyme transcripts, boosting CAR-T cell persistence and efficacy against solid tumors without permanent DNA changes.^[109] For neurological applications, similar Cas13 tools target metabolic pathways in neurons, with efficiencies reaching 50% in mouse brain tissues for correcting metabolic disorders like those in ALS models.^[110] The 2025 SPRING platform represents a precision breakthrough, integrating a hairpin-structured guide RNA with ADAR2 deaminase to displace non-target strands, achieving up to 67% editing efficiency and minimal bystander edits in human cells. This system enhances CRISPR-Cas13 fusions for therapeutic precision, particularly in vivo, by reducing off-target rates to below 1% in multiplex settings.^[111]

Comparison to DNA Editing

RNA editing therapies offer a transient modification to RNA transcripts, typically lasting hours to days, in stark contrast to the permanent genomic changes induced by DNA editing techniques such as CRISPR-Cas9.^[112] This reversibility stems from RNA's short lifespan and natural turnover, allowing effects to dissipate without lasting genomic impact, whereas DNA edits persist across cell divisions and generations.^[113] Furthermore, RNA editing circumvents the need for double-strand breaks (DSBs) required in many DNA editing methods, thereby minimizing risks of off-target mutations, insertions, or deletions that could lead to unintended genomic instability.^[114] These approaches are especially advantageous for non-dividing or post-mitotic cells, like neurons or mature muscle cells, where DNA editing efficiency is limited due to poor access or integration challenges.^[115] A key benefit of RNA editing lies in its capacity to address splicing defects—common in diseases like spinal muscular atrophy—by precisely altering pre-mRNA sequences to restore functional isoforms, all without modifying the underlying DNA.^[116] This preserves the genome's integrity, reducing long-term risks such as oncogenesis associated with DNA alterations.^[113] Ethically, RNA editing avoids germline transmission of changes, making it preferable for applications where heritable modifications raise concerns, unlike DNA therapies that could affect future generations.^[112] Despite these advantages, RNA editing requires repeated administrations to maintain therapeutic effects, given the ephemeral nature of RNA modifications, which contrasts with the one-time dosing potential of DNA edits.^[117] Delivery remains a hurdle, mirroring challenges in mRNA-based vaccines, including stability, immunogenicity, and tissue-specific targeting via lipid nanoparticles or viral vectors.^[118] Illustrative examples highlight these distinctions: CRISPR-Cas9 DNA editing has yielded approved therapies like Casgevy for sickle cell disease (SCD), involving ex vivo editing of hematopoietic stem cells to permanently reactivate fetal hemoglobin production.^[119] Conversely, ADAR-based RNA editing platforms are advancing for SCD and β-thalassemia, enabling reversible correction of aberrant transcripts in blood cells without genomic cuts.^[120] Looking ahead, the RNA editing technologies market is projected to expand from USD 262 million in 2024 to USD 397 million by 2030, driven by growing clinical pipelines.^[121]

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U-insertion/deletion mRNA editing holoenzyme: definition in sight
Abstract. RNA editing is a process that alters DNA-encoded sequence and is distinct from splicing, 5′ capping and 3′ additions.
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Adaptation of A-to-I RNA editing in Drosophila | PLOS Genetics
We detected strong signatures of positive selection on the nonsynonymous editing sites in Drosophila brains, and the beneficial editing sites were ...
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The preponderance of nonsynonymous A-to-I RNA editing ... - Nature
Nov 27, 2019 · For instance, in humans, the fraction of sites subject to nonsynonymous editing is lower than that subject to synonymous editing and the editing ...
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RNA editing: a driving force for adaptive evolution? - PMC
RNA editing changes a codon, which in turn leads to a protein with amino acid substitution and altered functional properties. Examples: neurotransmitter ...
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Adenosine-to-inosine RNA editing shapes transcriptome diversity in ...
Adenosine-to-inosine (A-to-I) RNA editing is a widespread modification of the transcriptome. ... GM Borchert, et al., Adenosine deamination in human transcripts ...Adenosine-To-Inosine Rna... · Results · Rna Editing Level In...<|control11|><|separator|>
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Wave Life Sciences Announces First-Ever Therapeutic RNA Editing ...
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Emerging clinical applications of ADAR based RNA editing - PMC
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RNA Editing Therapeutics: Advances, Challenges and Perspectives ...
Oct 14, 2022 · In this review, we discuss the main changes that RNA editing affects in cardiovascular gene expression and biology, as well as we introduce the ...
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Site-directed RNA editing: recent advances and open challenges
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RNA editing with CRISPR-Cas13 - Science
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Cas13d-mediated isoform-specific RNA knockdown with a unified ...
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A versatile CRISPR-Cas13d platform for multiplexed transcriptomic ...
Feb 29, 2024 · MEGA enables quantitative, reversible, and massively multiplexed gene knockdown in primary human T cells without targeting or cutting genomic DNA.
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Leveraging CRISPR gene editing technology to optimize the efficacy ...
Oct 25, 2024 · In this review we summarize potential uses of the CRISPR system to improve results of CAR T-cells therapy including optimizing efficacy and safety.
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A new RNA editing tool could enhance cancer treatment
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New CRISPR technology could help repair damaged neurons
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The Past, Present, and Future of RNA Editing Therapeutics
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Assessing and advancing the safety of CRISPR-Cas tools: from DNA ...
Jan 13, 2023 · Unlike DNA editing, RNA editing offers an alternative to genome editing in certain therapeutic applications in a reversible and tunable ...
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No Cuts, Just Edits: Therapeutic A-to-I RNA Editing as a Safer ...
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Gene editing: DNA versus RNA | Drug Discovery News
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RNA editing set to take off: could it outperform gene editing?
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RNA-editing drugs advance into clinical trials - Nature
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RNA editing: Expanding the potential of RNA therapeutics - PMC
ADAR-based RNA editing has emerged as a powerful tool to engineer RNAs, enable correction of disease-causing mutations, and modulate protein functions.
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FDA Approves First Gene Therapies to Treat Patients with Sickle ...
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Gene and RNA Editing: Methods, Enabling Technologies ... - arXiv
The purpose of gene and RNA editing is to modulate the root of disease by fixing the gene's DNA mutation [6] . Report issue for preceding element.
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Global RNA Editing Technologies Market By Share, Size and ...
The Global RNA Editing Technologies Market, valued at USD 262.28 Million in 2024, is projected to experience a CAGR of 7.16% to reach USD 397.16 Million by 2030 ...