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RNA editing
RNA editing
RNA editing
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RNA editing (also RNA modification) is a molecular process through which some cells can make discrete changes to specific nucleotide sequences within an RNA molecule after it has been generated by RNA polymerase. It occurs in all living organisms and is one of the most evolutionarily conserved properties of RNAs.[1][2][3] RNA editing may include the insertion, deletion, and base substitution of nucleotides within the RNA molecule. RNA editing is relatively rare, with common forms of RNA processing (e.g. splicing, 5'-capping, and 3'-polyadenylation) not usually considered as editing. It can affect the activity, localization as well as stability of RNAs, and has been linked with human diseases.[1][2][3][4]

RNA editing has been observed in some tRNA, rRNA, mRNA, or miRNA molecules of eukaryotes and their viruses, archaea, and prokaryotes.[5] RNA editing occurs in the cell nucleus, as well as within mitochondria and plastids. In vertebrates, editing is rare and usually consists of a small number of changes to the sequence of the affected molecules. In other organisms, such as squids,[6] extensive editing (pan-editing) can occur; in some cases the majority of nucleotides in an mRNA sequence may result from editing. More than 160 types of RNA modifications have been described so far.[7]

RNA-editing processes show great molecular diversity, and some appear to be evolutionarily recent acquisitions that arose independently. The diversity of RNA editing phenomena includes nucleobase modifications such as cytidine (C) to uridine (U) and adenosine (A) to inosine (I) deaminations, as well as non-template nucleotide additions and insertions. RNA editing in mRNAs effectively alters the amino acid sequence of the encoded protein so that it differs from that predicted by the genomic DNA sequence.[8]

The editosome complex

Detection of RNA editing

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Next generation sequencing

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To identify diverse post-transcriptional modifications of RNA molecules and determine the transcriptome-wide landscape of RNA modifications by means of next generation RNA sequencing, recently many studies have developed conventional[9] or specialised sequencing methods.[1][2][3] Examples of specialised methods are MeRIP-seq,[10] m6A-seq,[11] PA-m5C-seq [12], methylation-iCLIP,[13] m6A-CLIP,[14] Pseudo-seq,[15] Ψ-seq,[16] CeU-seq,[17] Aza-IP[18] and RiboMeth-seq[19]). Many of these methods are based on specific capture of the RNA species containing the specific modification, for example through antibody binding coupled with sequencing of the captured reads. After the sequencing these reads are mapped against the whole transcriptome to see where they originate from.[20] Generally with this kind of approach it is possible to see the location of the modifications together with possible identification of some consensus sequences that might help identification and mapping further on. One example of the specialize methods is PA-m5C-seq. This method was further developed from PA-m6A-seq method to identify m5C modifications on mRNA instead of the original target N6-methyladenosine. The easy switch between different modifications as target is made possible with a simple change of the capturing antibody form m6A specific to m5C specific.[12] Application of these methods have identified various modifications (e.g. pseudouridine, m6A, m5C, 2′-O-Me) within coding genes and non-coding genes (e.g. tRNA, lncRNAs, microRNAs) at single nucleotide or very high resolution.[4]

Mass spectrometry

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Mass spectrometry is a way to quantify RNA modifications.[21] More often than not, modifications cause an increase in mass for a given nucleoside. This gives a characteristic readout for the nucleoside and the modified counterpart.[21] Moreover, mass spectrometry allows the investigation of modification dynamics by labelling RNA molecules with stable (non-radioactive) heavy isotopes in vivo. Due to the defined mass increase of heavy isotope labeled nucleosides they can be distinguished from their respective unlabelled isotopomeres by mass spectrometry. This method, called NAIL-MS (nucleic acid isotope labelling coupled mass spectrometry), enables a variety of approaches to investigate RNA modification dynamics.[22][23][24]

Types of RNA

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Messenger RNA modification

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Recently, functional experiments have revealed many novel functional roles of RNA modifications. Most of the RNA modifications are found on transfer-RNA and ribosomal-RNA, but also eukaryotic mRNA has been shown to be modified with multiple different modifications. 17 naturally occurring modifications on mRNA have been identified, from which the N6-methyladenosine is the most abundant and studied.[25] mRNA modifications are linked to many functions in the cell. They ensure the correct maturation and function of the mRNA, but also at the same time act as part of cell's immune system.[26] Certain modifications like 2'O-methylated nucleotides has been associated with cells ability to distinguish own mRNA from foreign RNA.[27] For example, m6A has been predicted to affect protein translation and localization,[1][2][3] mRNA stability,[28] alternative polyA choice [14] and stem cell pluripotency.[29] Pseudouridylation of nonsense codons suppresses translation termination both in vitro and in vivo, suggesting that RNA modification may provide a new way to expand the genetic code.[30] 5-methylcytosine on the other hand has been associated with mRNA transport from the nucleus to the cytoplasm and enhancement of translation. These functions of m5C are not fully known and proven but one strong argument towards these functions in the cell is the observed localization of m5C to translation initiation site.[31] Importantly, many modification enzymes are dysregulated and genetically mutated in many disease types.[1] For example, genetic mutations in pseudouridine synthases cause mitochondrial myopathy, sideroblastic anemia (MLASA) [32] and dyskeratosis congenital.[33]

Compared to the modifications identified from other RNA species like tRNA and rRNA, the amount of identified modifications on mRNA is very small. One of the biggest reasons why mRNA modifications are not so well known is missing research techniques. In addition to the lack of identified modifications, the knowledge of associated proteins is also behind other RNA species. Modifications are results of specific enzyme interactions with the RNA molecule.[25] Considering mRNA modifications most of the known related enzymes are the writer enzymes that add the modification on the mRNA. The additional groups of enzymes readers and erasers are for most of the modifications either poorly known of not known at all.[34] For these reasons there has been during the past decade huge interest in studying these modifications and their function.[20]

Transfer RNA modifications

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Transfer RNA or tRNA is the most abundantly modified type of RNA.[35] Modifications in tRNA play crucial roles in maintaining translation efficiency through supporting structure, anticodon-codon interactions, and interactions with enzymes.[36]

Anticodon modifications are important for proper decoding of mRNA. Since the genetic code is degenerate, anticodon modifications are necessary to properly decode mRNA. Particularly, the wobble position of the anticodon determines how the codons are read. For example, in eukaryotes an adenosine at position 34 of the anticodon can be converted to inosine. Inosine is a modification that is able to base-pair with cytosine, adenine, and uridine.[37]

Another commonly modified base in tRNA is the position adjacent to the anticodon. Position 37 is often hypermodified with bulky chemical modifications. These modifications prevent frameshifting and increase anticodon-codon binding stability through stacking interactions.[37]

Ribosomal RNA modification

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Ribosomal RNA (rRNA) is essential to the makeup of ribosomes and peptide transfer during translation processes.[38] Ribosomal RNA modifications are made throughout ribosome synthesis, and often occur during and/or after translation. Modifications primarily play a role in the structure of the rRNA in order to protect translational efficiency.[38] Chemical modification in rRNA consists of methylation of ribose sugars, isomerization of uridines, and methylation and acetylation of individual bases.[39]

Methylation

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Methylation of rRNA upholds structural rigidity by blocking base pair stacking and surrounds the 2'-OH group to block hydrolysis. It occurs at specific parts of eukaryotic rRNA. The template for methylation consists of 10-21 nucleotides.[38] 2'-O-methylation of the ribose sugar is one of the most common rRNA modifications.[40] Methylation is primarily introduced by small nucleolar RNA's, referred to as snoRNPs. There are two classes of snoRNPs that target methylation sites, and they are referred to box C/D and box H/ACA.[39][40] One type of methylation, 2′-O-methylation, contributes to helical stabilization.[38]

Isomerization

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The isomerization of uridine to pseudouridine is the second most common rRNA modification. These pseudouridines are also introduced by the same classes of snoRNPs that participate in methylation. Pseudouridine synthases are the major participating enzymes in the reaction.[41] The H/ACA box snoRNPs introduce guide sequences that are about 14-15 nucleotides long.[39] Pseudouridylation is triggered in numerous places of rRNAs at once to preserve the thermal stability of RNA.[39] Pseudouridine allows for increased hydrogen bonding and alters translation in rRNA and tRNA.[40][41] It alters translation by increasing the affinity of the ribosome subunit to specific mRNAs.[38]

Base Editing:

Base editing is the third major class of rRNA modification, specifically in eukaryotes. There are 8 categories of base edits that can occur at the gap between the small and large ribosomal subunits.[38] RNA methyltransferases are the enzymes that introduce base methylation.[38] Acetyltransferases are the enzymes responsible for acetylation of cytosine in rRNA. Base methylation plays a role in translation. These base modifications all work in conjunction with the two other main classes of modification to contribute to RNA structural stability. An example of this occurs in N7-methylation, which increases the nucleotide's charge to increase ionic interactions of proteins attaching to the RNA before translation.

Editing by insertion or deletion

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The effect of uracil insertion in pre-mRNA transcripts

RNA editing through the addition and deletion of uracil has been found in kinetoplasts from the mitochondria of Trypanosoma brucei.[42] Because this may involve a large fraction of the sites in a gene, it is sometimes called "pan-editing" to distinguish it from topical editing of one or a few sites.

Pan-editing starts with the base-pairing of the unedited primary transcript with a guide RNA (gRNA), which contains complementary sequences to the regions around the insertion/deletion points. The newly formed double-stranded region is then enveloped by an editosome, a large multi-protein complex that catalyzes the editing.[43][44] The editosome opens the transcript at the first mismatched nucleotide and starts inserting uridines. The inserted uridines will base-pair with the guide RNA, and insertion will continue as long as A or G is present in the guide RNA and will stop when a C or U is encountered.[45][46] The inserted nucleotides cause a frameshift, and result in a translated protein that differs from its gene.

The mechanism of the editosome involves an endonucleolytic cut at the mismatch point between the guide RNA and the unedited transcript. The next step is catalyzed by one of the enzymes in the complex, a terminal U-transferase, which adds Us from UTP at the 3' end of the mRNA.[47] The opened ends are held in place by other proteins in the complex. Another enzyme, a U-specific exoribonuclease, removes the unpaired Us. After editing has made mRNA complementary to gRNA, an RNA ligase rejoins the ends of the edited mRNA transcript.[48][49] As a consequence, the editosome can edit only in a 3' to 5' direction along the primary RNA transcript. The complex can act on only a single guide RNA at a time. Therefore, a RNA transcript requiring extensive editing will need more than one guide RNA and editosome complex.

Editing by deamination

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C-to-U editing

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The effect of C-to-U RNA editing on the human ApoB gene

The editing involves cytidine deaminase that deaminates a cytidine base into a uridine base. An example of C-to-U editing is with the apolipoprotein B gene in humans. Apo B100 is expressed in the liver and apo B48 is expressed in the intestines. In the intestines, the mRNA has a CAA sequence edited to be UAA, a stop codon, thus producing the shorter B48 form. C-to-U editing often occurs in the mitochondrial RNA of flowering plants. Different plants have different degrees of C-to-U editing; for example, eight editing events occur in mitochondria of the moss Funaria hygrometrica, whereas over 1,700 editing events occur in the lycophytes Isoetes engelmanii.[50] C-to-U editing is performed by members of the pentatricopeptide repeat (PPR) protein family. Angiosperms have large PPR families, acting as trans -factors for cis -elements lacking a consensus sequence; Arabidopsis has around 450 members in its PPR family. There have been a number of discoveries of PPR proteins in both plastids and mitochondria.[51]

A-to-I editing

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See also: Adenine deamination

Adenosine-to-inosine (A-to-I) modifications contribute to nearly 90% of all editing events in RNA. The deamination of adenosine is catalyzed by the double-stranded RNA-specific adenosine deaminase (ADAR), which typically acts on pre-mRNAs. The deamination of adenosine to inosine disrupts and destabilizes the dsRNA base pairing, therefore rendering that particular dsRNA less able to produce siRNA, which interferes with the RNAi pathway.

The wobble base pairing causes deaminated RNA to have a unique but different structure, which may be related to the inhibition of the initiation step of RNA translation. Studies have shown that I-RNA (RNA with many repeats of the I-U base pair) recruits methylases that are involved in the formation of heterochromatin and that this chemical modification heavily interferes with miRNA target sites.[52] There is active research into the importance of A-to-I modifications and their purpose in the novel concept of epitranscriptomics, in which modifications are made to RNA that alter their function.[53][54] A long established consequence of A-to-I in mRNA is the interpretation of I as a G, therefore leading to functional A-to-G substitution, e.g. in the interpretation of the genetic code by ribosomes. Newer studies, however, have weakened this correlation by showing that inosines can also be decoded by the ribosome (although in a lesser extent) as adenosines or uracils. Furthermore, it was shown that I's lead to the stalling of ribosomes on the I-rich mRNA.[55]

The development of high-throughput sequencing in recent years has allowed for the development of extensive databases for different modifications and edits of RNA. RADAR (Rigorously Annotated Database of A-to-I RNA editing) was developed in 2013 to catalog the vast variety of A-to-I sites and tissue-specific levels present in humans, mice, and flies. The addition of novel sites and overall edits to the database are ongoing.[56] The level of editing for specific editing sites, e.g. in the filamin A transcript, is tissue-specific.[57] The efficiency of mRNA-splicing is a major factor controlling the level of A-to-I RNA editing.[58][59] Interestingly, ADAR1 and ADAR2 also affect alternative splicing via both A-to-I editing ability and dsRNA binding ability.[60][61]

Alternative mRNA editing

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Alternative U-to-C mRNA editing was first reported in WT1 (Wilms Tumor-1) transcripts,[62] and non-classic G-A mRNA changes were first observed in HNRNPK (heterogeneous nuclear ribonucleoprotein K) transcripts in both malignant and normal colorectal samples.[63] The latter changes were also later seen alongside non-classic U-to-C alterations in brain cell TPH2 (tryptophan hydroxylase 2) transcripts.[64] Although the reverse amination might be the simplest explanation for U-to-C changes, transamination and transglycosylation mechanisms have been proposed for plant U-to-C editing events in mitochondrial transcripts.[65] A recent study reported novel G-to-A mRNA changes in WT1 transcripts at two hotspots, proposing the APOBEC3A (apolipoprotein B mRNA editing enzyme, catalytic polypeptide 3A) as the enzyme implicated in this class of alternative mRNA editing.[66] It was also shown that alternative mRNA changes were associated with canonical WT1 splicing variants, indicating their functional significance.

RNA editing in plant mitochondria and plastids

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It has been shown in previous studies that the only types of RNA editing seen in the plants' mitochondria and plastids are conversion of C-to-U and U-to-C (very rare).[67][68][69][70][71][72][73][74][75][76][77][78][79] RNA-editing sites are found mainly in the coding regions of mRNA, introns, and other non-translated regions.[69] In fact, RNA editing can restore the functionality of tRNA molecules.[71][72] The editing sites are found primarily upstream of mitochondrial or plastid RNAs. While the specific positions for C to U RNA editing events have been fairly well studied in both the mitochondrion and plastid,[80] the identity and organization of all proteins comprising the editosome have yet to be established. Members of the expansive PPR protein family have been shown to function as trans-acting factors for RNA sequence recognition.[81] Specific members of the MORF (Multiple Organellar RNA editing Factor) family are also required for proper editing at several sites. As some of these MORF proteins have been shown to interact with members of the PPR family, it is possible MORF proteins are components of the editosome complex.[82] An enzyme responsible for the trans- or deamination of the RNA transcript remains elusive, though it has been proposed that the PPR proteins may serve this function as well.

RNA editing is essential for the normal functioning of the plant's translation and respiration activity. Editing can restore the essential base-pairing sequences of tRNAs, restoring functionality.[83] It has also been linked to the production of RNA-edited proteins that are incorporated into the polypeptide complexes of the respiration pathway. Therefore, it is highly probable that polypeptides synthesized from unedited RNAs would not function properly and hinder the activity of both mitochondria and plastids.

C-to-U RNA editing can create start and stop codons, but it cannot destroy existing start and stop codons. A cryptic start codon is created when the codon ACG is edited to be AUG.

Summary of the Various Functions of RNA Editing

RNA editing in viruses

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Viruses (i.e., measles, mumps, or parainfluenza), especially viruses that have an RNA genome, have been shown to have evolved to utilize RNA modifications in many ways when taking over the host cell. Viruses are known to utilize the RNA modifications in different parts of their infection cycle from immune evasion to protein translation enhancement.[27] RNA editing is used for stability and generation of protein variants.[84][85] Viral RNAs are transcribed by a virus-encoded RNA-dependent RNA polymerase, which is prone to pausing and "stuttering" at certain nucleotide combinations. In addition, up to several hundred non-templated A's are added by the polymerase at the 3' end of nascent mRNA.[86] These As help stabilize the mRNA. Furthermore, the pausing and stuttering of the RNA polymerase allows the incorporation of one or two Gs or As upstream of the translational codon.[86] The addition of the non-templated nucleotides shifts the reading frame, which generates a different protein.

Additionally, the RNA modifications are shown to have both positive and negative effects on the replication and translation efficiency depending on the virus.  For example, Courtney et al.[12] showed that an RNA modification called 5-methylcytosine is added to the viral mRNA in infected host cells in order to enhance the protein translation of HIV-1 virus. The inhibition of the m5C modification on viral mRNA results in significant reduction in viral protein translation, but interestingly it has no effect on the expression of viral mRNAs in the cell. On the other hand, Lichinchi et al.[87] showed that the N6-methyladenosine modification on ZIKV mRNA inhibits the viral replication.

Origin and evolution of RNA editing

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The RNA-editing system seen in the animal may have evolved from mononucleotide deaminases, which have led to larger gene families that include the apobec-1 and adar genes. These genes share close identity with the bacterial deaminases involved in nucleotide metabolism. The adenosine deaminase of E. coli cannot deaminate a nucleoside in the RNA; the enzyme's reaction pocket is too small for the RNA strand to bind to. However, this active site is widened by amino acid changes in the corresponding human analog genes, APOBEC1 and ADAR, allowing deamination.[88][89] The gRNA-mediated pan-editing in trypanosome mitochondria, involving templated insertion of U residues, is an entirely different biochemical reaction. The enzymes involved have been shown in other studies to be recruited and adapted from different sources.[43][90] But the specificity of nucleotide insertion via the interaction between the gRNA and mRNA is similar to the tRNA editing processes in the animal and Acanthamoeba mitochondria.[91] Eukaryotic ribose methylation of rRNAs by guide RNA molecules is a similar form of modification.[92]

Thus, RNA editing evolved more than once. Several adaptive rationales for editing have been suggested.[93] Editing is often described as a mechanism of correction or repair to compensate for defects in gene sequences. However, in the case of gRNA-mediated editing, this explanation does not seem possible because if a defect happens first, there is no way to generate an error-free gRNA-encoding region, which presumably arises by duplication of the original gene region. A more plausible alternative for the evolutionary origins of this system is through constructive neutral evolution, where the order of steps is reversed, with the gratuitous capacity for editing preceding the "defect".[94]

Therapeutic mRNA editing

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See also: CRISPR § Cas13 (formerly C2c2)

Directing edits to correct mutated sequences was first proposed and demonstrated in 1995.[95] This initial work used synthetic RNA antisense oligonucleotides complementary to a pre-mature stop codon mutation in a dystrophin sequence to activate A-to-I editing of the stop codon to a read through codon in a model xenopus cell system.[95] While this also led to nearby inadvertent A-to-I transitions, A to I (read as G) transitions can correct all three stop codons, but cannot create a stop codon. Therefore, the changes led >25% correction of the targeted stop codon with read through to a downstream luciferase reporter sequence. Follow on work by Rosenthal achieved editing of mutated mRNA sequence in mammalian cell culture by directing an oligonucleotide linked to a cytidine deaminase to correct a mutated cystic fibrosis sequence.[96] More recently, CRISPR-Cas13 fused to deaminases has been employed to direct mRNA editing.[97]

In 2022, therapeutic RNA editing for Cas7-11 was reported.[98][99] It enables sufficiently targeted cuts and an early version of it was used for in vitro editing in 2021.[100]

Comparison to DNA editing

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Unlike DNA editing, which is permanent, the effects of RNA editing − including potential off-target mutations in RNA − are transient and are not inherited. RNA editing is therefore considered to be less risky. Furthermore, it may only require a guide RNA by using the ADAR protein already found in humans and many other eukaryotes' cells instead of needing to introduce a foreign protein into the body.[101]

See also

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References

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Revisions and contributorsEdit on WikipediaRead on Wikipedia

RNA editing

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from Grokipedia
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. 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). 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. A-to-I editing is the most widespread form in metazoans, with enzymes (including ADAR1, ADAR2, and the catalytically inactive ADAR3) targeting structured regions to generate transcriptomic diversity, recode in proteins, and modulate stability or splicing. For instance, ADAR1 plays a in innate immunity by editing self double-stranded RNAs to prevent their recognition by sensors like , thus averting autoimmune responses, while ADAR2 facilitates essential recoding events in neuronal transcripts such as the glutamate receptor subunit GRIA2, changing a to codon for proper channel function. 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 , or APOBEC3 family members' roles in antiviral defense by hypermutating viral genomes. Beyond these core mechanisms, RNA editing expands the without altering the , influences evolutionary adaptation through conserved recoding sites under positive selection, and has implications for when dysregulated, including neurological disorders, cancer, and immune pathologies. 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.

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. The phenomenon was first discovered in 1986 in the mitochondria of trypanosomes, where extensive insertion of (U) residues into mitochondrial mRNAs was found to be necessary for proper coding of proteins, resolving apparent discrepancies between sequences and translated products. This unexpected finding challenged the and introduced the concept of as a malleable information carrier. In the late , editing was expanded to mammalian systems with the identification of cytidine-to- (C-to-U) editing in (apoB) mRNA in the intestine, which generates a truncated essential for . Adenosine-to-inosine (A-to-I) editing in mammals was subsequently recognized in the early through studies of brain transcripts, particularly in mRNAs like those encoding 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. Major enzymes driving RNA editing include the acting on RNA () family, which catalyzes A-to-I substitutions by deaminating to in double-stranded RNA regions, and the mRNA editing enzyme catalytic polypeptide-like () family, particularly APOBEC1, which performs C-to-U editing via deamination. These enzymes are highly conserved and play pivotal roles in regulating 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 in proteins, RNA editing generates protein isoforms with distinct functions, thereby increasing the complexity of the without requiring genomic mutations. 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. In humans, these processes contribute to editing in approximately 1-3% of transcripts, primarily through A-to-I changes mediated by enzymes. Biologically, RNA editing exhibits tissue-specific patterns that fine-tune cellular functions, particularly in dynamic environments like the . In the , elevated editing levels support by modulating neurotransmitter receptor properties and neuronal excitability, allowing adaptive responses to neural activity. also facilitates rapid 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. In organelles like plant mitochondria and chloroplasts, editing serves as an error-correction mechanism, converting mismatched (e.g., C-to-U) to restore functional protein sequences that would otherwise be defective due to mutational biases in organellar genomes. 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. 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). From an evolutionary perspective, RNA editing confers adaptive advantages by providing a flexible layer of plasticity, enabling organisms to respond to changing environments or developmental cues without the fixation of potentially deleterious in the . This mechanism is particularly evident in species facing variable conditions, where edited transcripts support physiological acclimation and increased at the RNA level.

Detection Methods

Next-Generation Sequencing

Next-generation sequencing (NGS) enables the genome-wide identification and quantification of RNA editing sites by aligning reads to a or matched genomic DNA sequence, detecting mismatches that indicate editing events. For A-to-I editing, is reverse-transcribed and sequenced as , 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. Standard protocols involve performing on poly(A)-selected or total 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, aligned BAM files to annotate sites, apply filters for , 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 in GIREMI, to prioritize true positives without requiring multiple replicates initially. NGS offers high-throughput , 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 patterns. However, challenges include distinguishing true 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. 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.

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. This approach distinguishes inosine from adenosine, enabling precise measurement of editing efficiency in specific RNA populations, such as mRNA or viral RNA. 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. Key techniques include nanoLC-MS/MS for proteome-wide scans, which employs high-resolution instruments like the to sequence peptides from complex samples, often using tandem mass tag (TMT) labeling for multiplexed quantification across multiple samples. For inosine-specific quantification, methods like nucleic acid isotope labeling coupled mass spectrometry (NAIL-MS) incorporate stable isotopes during to track modification dynamics, allowing differentiation of newly synthesized versus pre-existing edited RNAs with . 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 tissue from over 170 subjects. Similarly, advanced LC-MS/MS protocols with chemical derivatization have enabled detection of low-abundance inosine in single cells or mRNA. The primary advantages of MS-based validation lie in its ability to confirm the functional consequences of RNA editing, such as substitutions that alter protein properties (e.g., calcium permeability in GluA2-edited receptors), providing direct evidence beyond nucleotide-level observations. It complements next-generation sequencing by verifying editing at the translational output, with high specificity for modified species. However, limitations include low sensitivity for rare editing events, often necessitating enrichment strategies like of specific RNAs or peptides, as seen in the sparse detection of only 10 out of 294 predicted recoding events in proteomes. Sample preparation challenges, such as RNA instability and ion suppression in complex matrices, further require rigorous controls. Post-2015 multiplexing innovations, including TMT and improved , have enhanced scalability for validating editing impacts in aging or disease contexts, such as Alzheimer's tissue.

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 bases to distinguish them from without relying on enzymatic treatments that can introduce biases. 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. Building on foundations from next-generation sequencing and , such methods address longstanding gaps in sensitivity for low-frequency events. Nanopore direct sequencing has emerged as a key tool for real-time detection of inosine reads, preserving native structure to identify A-to-I sites without reverse transcription artifacts. The Dinopore algorithm, developed for platforms, uses signal patterns to pinpoint with high precision across organisms, facilitating the study of dynamics in real time. A 2025 analysis highlights how these advancements in direct enhance the detection of modifications, including events, by integrating to classify base-level signals. In 2024, developments in AI-driven 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. 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. Such predictive frameworks are particularly valuable for identifying non-model organism edits where experimental validation is resource-intensive. Complementing this, 2025 multi-omics integration efforts have enhanced understanding of RNA editing by combining with epigenomic and proteomic data, revealing roles in tumor evolution. 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. By minimizing artifacts, they have been shown to substantially reduce false positives, enabling more reliable profiling of editing landscapes in disease states.

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. 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. These modifications expand the proteome diversity without altering the genomic sequence, influencing gene expression, protein function, and cellular responses. The core reaction proceeds via , where water acts as the to deaminate the base: for example, is converted to plus ( + H₂O → + NH₃). This enzymatic activity requires double-stranded (dsRNA) substrates, as the editing enzymes recognize structured regions to ensure specificity and avoid off-target effects. enzymes, in particular, feature double-stranded -binding domains (dsRBDs) that facilitate substrate recognition, followed by a catalytic deaminase domain that executes the through a two-step mechanism involving a tetrahedral intermediate. enzymes similarly rely on structure for positioning but often require auxiliary factors for efficient activity . Deamination-based editing accounts for the vast majority of known RNA editing events in eukaryotic transcriptomes, far outnumbering other substitution or modification types. 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. These regulatory elements ensure that editing occurs in a controlled manner, often in response to cellular stress or developmental cues. Specific instances of C-to-U and A-to-I editing, such as those mediated by APOBEC1 in mRNA or ADAR2 in transcripts, exemplify this mechanism's role in fine-tuning protein isoforms.

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 (U) residues, in contrast to substitution 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 guide RNAs (gRNAs). 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 , 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 REL2. In U deletion, mismatched Us are removed from the 3' fragment by the 3'-5' activity associated with the REH1 , 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 subunit III (COIII) mRNA of , to generate functional open reading frames from cryptic pre-mRNAs. 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 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.00468-0)

Deamination-Based Editing

C-to-U Editing

C-to-U RNA editing is a in which residues in transcripts are deaminated to , altering the without changing the underlying DNA sequence. This process occurs predominantly in nuclear-encoded messenger RNAs in animals and in organellar transcripts in , 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. In mammalian systems, the primary 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 s in target RNAs, with editing efficiency reaching over 90% in tissues like the . Site specificity is achieved through an 11-nucleotide mooring located 3-5 nucleotides downstream of the edited , typically featuring a core motif of 5'-UUUN-3' that positions the correctly. A canonical example is the editing of (APOB) mRNA, where of 6666 (C6666) to creates a premature (UAA), truncating the full-length ApoB100 protein into the shorter ApoB48 isoform essential for lipid transport in the intestine. 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 at hundreds of sites, often restoring evolutionarily conserved or creating functional codons. For instance, in plastids, editing converts an ACG initiation codon to AUG in the psbL mRNA of , enabling translation of the subunit PsbL. In mitochondria, similar editing events optimize codons for protein function, with over 400 sites identified in species like . 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- patterns driven by 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, has been shown to catalyze site-specific C-to-U editing of transfer RNAs (tRNAs), with implications for cancer and immune responses.

A-to-I Editing

A-to-I RNA editing, the conversion of (A) to (I) in , is the most prevalent form of editing in animals and is catalyzed by the acting on (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 (dsRNA) substrates. The editing process involves hydrolytic deamination of within dsRNA structures, where is subsequently recognized as (G) during 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 subunit GluA2, where editing changes CAG () to CIG (read as CGG, ), modulating calcium permeability in neuronal synapses. Humans harbor over 100 confirmed recoding sites in mRNAs, many affecting receptors and channels critical for 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. 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. 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.

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 by altering codon sequences in protein-coding transcripts. This process occurs in a majority of protein-coding genes, with editing events being particularly prevalent in the , where they contribute to neuronal plasticity and function. For instance, the serotonin receptor 2C (HTR2C) transcript undergoes extensive A-to-I editing at five sites within its , generating up to 24 distinct isoforms that modulate receptor signaling and G-protein coupling efficiency. The primary functions of mRNA editing include recoding to produce protein isoforms with modified properties, such as altered selectivity or channel kinetics in voltage-gated channels. For example, editing in subunits of channels like KCNA1 changes an to , altering recovery from inactivation and fine-tuning neuronal excitability. Additionally, exonic edits can regulate by introducing or disrupting exonic splicing enhancer motifs, thereby influencing isoform production and patterns. A canonical example is the / (Q/R) site in the GluA2 subunit of receptors, where nearly complete A-to-I editing converts a CAG codon to CIG (read as CGG), replacing with in the channel pore and preventing calcium influx, which is essential for synaptic transmission and . In diseases like (), reduced ADAR2 activity leads to incomplete editing at this site, resulting in calcium-permeable receptors that promote degeneration through .

Transfer RNA Editing

Transfer RNA (tRNA) editing primarily involves post-transcriptional modifications that ensure accurate 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 mitochondria, C-to-U editing at position 35 of the anticodon converts a tRNA to decode aspartate codons instead of a mismatched sequence. Another prominent type is A-to-I editing at the wobble position (position 34), which expands codon recognition by allowing to pair with U, C, or A in the third codon position, thereby enabling a single tRNA to decode multiple synonymous codons. This A-to-I process is analogous to deamination-based in but is specialized for tRNA anticodon integrity. The enzymes mediating these edits are highly specific. For A-to-I editing, the ADAT complex—comprising 1 (homodimeric, akin to Tad1), ADAT2, and ADAT3 (forming a heterodimer)—catalyzes at wobble and adjacent positions in various tRNAs, with ADAT2/3 targeting up to eight tRNA species in eukaryotes. These enzymes act independently or in complexes, recognizing tRNA structural motifs without requiring guide RNAs. tRNA editing serves critical functions, including correction of genomic encoding errors in tRNA and facilitation of the for degenerate codon decoding. By introducing at the wobble position, editing compensates for limitations in tRNA gene diversity, allowing efficient of the full codon repertoire without requiring additional tRNA isoacceptors. This process is essential for viability in , where Tad1/2/3 mutants exhibit severe growth defects due to impaired tRNA function, and in mammals, where disruptions lead to translational inefficiencies. Illustrative examples highlight tRNA editing's biological impact. In , the TadA enzyme (a bacterial homolog) performs A-to-I editing at the wobble position of tRNA^Arg(ACG), forming inosine-34 essential for codon decoding and cell viability. 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, , and stroke-like episodes (MELAS), where disruptions affect anticodon integrity and . 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 in (rRNA) is rare compared to the abundant post-transcriptional chemical modifications that support and function. While modifications such as pseudouridylation ( of to ) and 2'-O-methylation enhance rRNA stability, folding, and translational efficiency, they do not change the nucleotide sequence and are distinct from . Rare instances of A-to-I have been suggested in rRNA expansion segments, but this is not a primary mechanism and lacks widespread confirmation in cytoplasmic rRNA. Sequence is more prominent in organellar rRNAs, such as C-to-U changes in mitochondria and plastids (covered in the organelle editing section). 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 ) synthase, and box C/D snoRNPs direct 2'-O-methylation by fibrillarin (Nop1 in ), 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, failure, and premature aging.

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 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 accumulated in organellar genomes over , often restoring canonical essential for protein function; notable examples include the creation of AUG initiation codons or elimination of premature stop codons (UAG to UGG, ). Editing is also tissue-specific, with varying efficiencies observed across developmental stages or organs, potentially fine-tuning under environmental cues. In (Zea mays) chloroplasts, RNA 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 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 (), particularly ADAR1, which targets double-stranded regions in viral RNAs. This editing introduces , read as during , potentially altering sequences and reducing the of viral double-stranded RNA intermediates. In contrast, C-to-U editing in coronaviruses like is driven by host APOBEC family deaminases, such as APOBEC3A, which convert to uracil in single-stranded viral RNA, leading to hypermutation patterns observed in emerging variants. These editing events serve critical functions in , including the generation of protein isoforms that enhance replication or evade immunity. A-to-I editing can disrupt viral to limit over-replication or create variants with altered antigenicity, while C-to-U hypermutation promotes akin to antigenic variation in other RNA viruses. Both types contribute to immune evasion by attenuating the host response; for instance, ADAR1-mediated editing of viral RNAs prevents excessive activation of pattern recognition receptors like , thereby dampening type I 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. 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 and 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 , 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 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.

Evolutionary Origins and Implications

Proposed Origins

One prominent hypothesis posits that RNA editing represents a vestige of the ancient , 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 or repair—for modern editing functions, allowing persistence of these "old players" in eukaryotic genomes. This theory aligns with the broader paradigm, suggesting that editing evolved as a legacy of primordial 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 . 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. 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 oxidase subunits, are conserved across kinetoplastid subgroups, indicating an early innovation predating more specialized forms like extensive pan-editing. In , C-to-U editing shows signs of , independently arising in mitochondrial and genomes to address similar mutational pressures, despite phylogenetic distance from kinetoplastid systems. Key proposals from the further suggest that A-to-I editing by ADARs arose via a "mistaken identity" mechanism, where enzymes, evolved for antiviral defense against double-stranded , 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.

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 acting on RNA () enzymes, responsible for A-to-I editing, are highly conserved among metazoans, with structural and functional similarities preserved from to vertebrates, underscoring their essential role in neural function and development. 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 numbers, suggesting relaxed selective constraints in these advanced flowering . 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. Recent discoveries have revealed non-canonical A-to-I editing in fungi, independent of metazoan ADARs and mediated by tRNA-specific deaminases during , allowing for adaptive responses such as antiviral defense through modulation of nearby . 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 , where they enhance protein diversity and neural adaptability, contributing to lineage-specific traits. 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. This duality may facilitate , as editing-generated transcriptomic variability can promote and ecological divergence, exemplified by elevated editing rates in species with recent adaptive radiations. Comparative genomic analyses further illuminate how editing sites correlate with , influencing their distribution and evolution. In , 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 enzymes, leading to hyper-editing that shapes diversity and potentially drives primate-specific adaptations.

Therapeutic Applications

Enzyme-Based RNA Editing Therapies

Enzyme-based RNA editing therapies harness endogenous cellular enzymes, such as for adenosine-to-inosine (A-to-I) editing and 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 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 enzymes using partially double-stranded guide RNAs that anchor to the target transcript, enabling precise without introducing foreign proteins. Wave Life Sciences' platform exemplifies this, utilizing chemically optimized delivered subcutaneously to activate endogenous for editing the SERPINA1 mRNA in (AATD), a condition causing and liver due to misfolded protein accumulation. Therapeutic targets include monogenic diseases amenable to single-nucleotide corrections or modulation. In AATD, Wave's WVE-006 achieved the first demonstration of therapeutic RNA editing in humans during the phase 1b/2a RestorAATion-2 , 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. For (DMD), preclinical recruitment has corrected point mutations in the DMD gene, such as the missense 1682G>A , restoring expression in mouse models via A-to-I editing to revert the codon, potentially addressing cases caused by such mutations. 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 seen in DNA editing. Delivery systems like LNPs for 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 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 , 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.

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. 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 ADAR2 to achieve adenosine-to-inosine (A-to-I) editing. This , 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. 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 . 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. Such advancements postdate earlier Cas7-11 explorations, filling gaps in scalable, tissue-specific applications. In therapeutic contexts, these tools show promise in 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. 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. 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 models. The 2025 SPRING platform represents a precision breakthrough, integrating a hairpin-structured with ADAR2 deaminase to displace non-target strands, achieving up to 67% editing efficiency and minimal bystander edits in cells. This system enhances CRISPR-Cas13 fusions for therapeutic precision, particularly , by reducing off-target rates to below 1% in multiplex settings.

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. 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. 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. 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. 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. This preserves the genome's integrity, reducing long-term risks such as oncogenesis associated with DNA alterations. 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. 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. Delivery remains a hurdle, mirroring challenges in mRNA-based , including stability, , and tissue-specific targeting via nanoparticles or viral vectors. Illustrative examples highlight these distinctions: CRISPR-Cas9 DNA editing has yielded approved therapies like Casgevy for (SCD), involving editing of hematopoietic stem cells to permanently reactivate fetal hemoglobin production. 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. 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.

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