Skip to main content


RNA editing in the forefront of epitranscriptomics and human health

Article metrics

  • 468 Accesses


Post-transcriptional modifications have been recently expanded with the addition of RNA editing, which is predominantly mediated by adenosine and cytidine deaminases acting on DNA and RNA. Here, we review the full spectrum of physiological processes in which these modifiers are implicated, among different organisms. Adenosine to inosine (A-to-I) editors, members of the ADAR and ADAT protein families are important regulators of alternative splicing and transcriptional control. On the other hand, cytidine to uridine (C-to-U) editors, members of the AID/APOBEC family, are heavily implicated in innate and adaptive immunity with important roles in antibody diversification and antiviral response. Physiologically, these enzymes are present in the nucleus and/or the cytoplasm, where they modify various RNA molecules, including miRNAs, tRNAs apart from mRNAs, whereas DNA editing is also possible by some of them. The expansion of next generation sequencing technologies provided a wealth of data regarding such modifications. RNA editing has been implicated in various disorders including cancer, and neurological diseases of the brain or the central nervous system. It is also related to cancer heterogeneity and the onset of carcinogenesis. Response to treatment can also be affected by the RNA editing status where drug efficacy is significantly compromised. Studying RNA editing events can pave the way to the identification of new disease biomarkers, and provide a more personalised therapy to various diseases.


The discovery of RNA editing and the field of Epitranscriptomics

RNA modifications refer to alterations in the chemical structure of RNA molecules occurring after DNA transcription and synthesis by the RNA polymerase enzyme. They were first described in 1968 with the discovery of RNA methylation in Hela cells [1]. Since then, modifications have been observed across many RNA types (miRNA, mRNA, rRNA, etc.) and detected in all domains of life, including archaea, prokaryotes and eukaryotes. So far, 112 nucleotide modifications have been observed, with the potential to affect the function and stability of the RNA molecule [2].

A unique type of RNA modification in trypanosome mitochondrial mRNA was discovered ~ 30 years ago [3]. The highly conserved mitochondrial cytochrome c oxidase subunit II (COX-2) gene mRNA was found to have four extra uridine (U) nucleotides, which could restore the reading frameshift to a functioning gene transcript. This post-transcriptional modification which edits the RNA transcript sequence, differentiating it from its corresponding DNA sequence, was named RNA editing. A year later, evidence of tissue-specific RNA editing was discovered in mammals [4]. The production of apolipoprotein-B48 in the intestine was observed to occur after a post-transcriptional cytidine to uridine (C-to-U) mRNA change in the gene’s transcript, which is responsible for the production of the hepatic apolipoprotein-B100. This change creates a translational stop codon and the functionally truncated intestinal protein (Fig. 1a).

Fig. 1

Cytidine and adenosine deaminases are critical RNA editors that play important functions in physiological events. a The vital role of APOBEC1 editing can be observed in the production of apolipoprotein B in the gut. The C-to-U editing at residue 2153 of hepatic Apo-B100 transforms the glutamate to a stop codon and produces a truncated protein Apo-B48 in intestinal cells [4]. b In neurons, mRNA editing of the glutamate receptor 2 (GluR2) at position 607 by ADAR2 results in an adenosine to inosine chance. This transforms the CAG codon for glutamine (Q) to CIG for arginine (R) as (CGG), since ribosomes read inosine (I) as guanosine (G). This neutralizes the diffusion of divalent cations and makes the receptor impermeable to calcium [112]

Early characterisation of RNA was time consuming and required substantial sample for sequencing because of its fragile state. In the beginning, researchers combined the knowledge of RNase enzymes to cut the RNA at specific sites and fragments, leading to the first complete tRNA sequence of Saccharomyces cerevisiae alanine tRNA [5, 6]. In 1977, alternative splicing was observed in adenoviruses demonstrating the capability of post-transcriptional modifications [7, 8]. Continued technological innovation led to the emergence of next-generation sequencing (NGS) technologies and made possible the high-throughput sequencing and identification of numerous RNA editing sites [9]. Furthermore, the programming of powerful computational methodologies enabled the study and prediction of RNA editing to become much more feasible [10].

The field of post-transcriptional RNA modifications is expanding beyond the common view of adjusting the structures and functions of mature RNA. Named RNA Epigenetics, or Epitranscriptomics, is an increasing group of RNA modifications classified in 4 groups: (1) the isomerization of uridine to pseudouridine, (2) alterations in the bases, including methylation and deamination, (3) methylation of the ribose 2′-hydroxyl (-OH) group and (4) complex and multiple modifications or hypermodifications [11]. Epitranscriptomics is progressively associated with many biological functions, from brain development and neuronal regulation to antibody diversification and immune defence [12, 13].

Conservation among species and the “Constructive Neutral Evolution” proposal

The identification of an increasing number of RNA edited sites in many organisms signifies the importance of this phenomenon in the evolution of species. The presence of RNA editing has been well observed in plants and animals [14, 15]. Phylogenetic analysis of the deaminase enzymes responsible for RNA editing, suggests that adenosine and cytidine deaminases emerged early in the metazoan radiation [15,16,17]. It is believed that they arose from an early transfer of an ancestral deaminase from bacteria toxin systems [18, 19]. Supporting the above hypothesis, recent evidence shows that RNA editing contributes in central processes in bacteria, thus, regulating evolutionarily conserved toxin-antitoxin systems [19]. From fruit flies to humans, RNA editing affects multiple cellular processes and is highly conserved [15, 20, 21].

The central dogma of molecular biology states that information is transferred from DNA to RNA to protein. Why RNA editing has emerged is a question that puzzled scientists, and is still under debate [22]. Simple answers are to repair genomic mutations or to provide another level of protein abundance. A three step model has been proposed for the development of RNA editing, accounting for the emergence of its activity, the mutation of editable nucleotide positions and fixation by drift, leading to its maintenance by natural selection [23]. This model has been expanded further, to describe how RNA editing could have evolved in the absence of positive selection [24]. Neutral theory suggests that neutral mutations and genetic drift account for the evolutionary changes at the molecular level [25]. Constructive neutral evolution (CNE) proposes that RNA editing systems emerged in existing proteins with metabolic activity where they remain active due to their ability to fix deleterious mutations in the RNA level. This can lead to an accumulation of gene mutations, since functional constrains are suppressed. However, RNA editing can be lost if these gene mutations are reversed; or if they accumulate, it becomes essential for the flow of genetic information [22, 26].

Deaminases acting as editors

Adenosine to inosine (A-to-I) editors

The unwinding of double stranded RNA (dsRNA) in oocytes of Xenopus laevis was the first evidence of adenosine to inosine (A-to-I) RNA editing [27]. The number of adenosine deaminases acting on RNA (ADARs) has increased, since then [28]. A-to-I conversion is the most common type of RNA editing [29]. In mammals, three ADAR genes (ADAR1-3) produce four isoforms, ADAR1p150, ADAR1p110, ADAR2 and ADAR3 [30, 31]. ADAR1 and ADAR2 arose from gene duplication around 700 million years ago in the early metazoan evolution [32]. ADAR3 most likely arose within the vertebrate linage from ADAR2 gene duplication [32]. The genome of Caenorhabditis elegans harbours two ADAR genes, adr-1 and adr-2 [33], while the genome of Drosophila melanogaster has only one, dADAR, that is under strict developmental control [34]. Two more ADAR-like genes (renamed to ADAD) are found in vertebrates; ADAD1 (or testes nuclear RNA-binding protein, TENR) which is required for spermatogenesis, and ADAD2, which is expressed in the brain [35, 36]. TENR-like genes have also been observed in the genomes of Danio rerio and Takifugu rubripes [32].

Adenosine deaminases that act on tRNAs (ADATs) form another class of A-to-I enzymes targeting tRNA molecules, and are believed to be evolutionary ancestors of ADARs [37]. In bacteria, TadA was the first prokaryotic RNA editing enzyme characterised in Escherichia coli [38]. ADATs have also been observed beyond metazoan, and are found in many prokaryotic and eukaryotic organisms [16]. In most eukaryotes, including humans, three ADAT enzymes (ADAT1-3) have been identified [39].

Cytidine to uridine (C-to-U) editors

Cytidine deaminases within the activation induced cytidine deaminase/apolipoprotein B editing complex (AID/APOBEC) family are responsible for the C-to-U mRNA editing, but also for DNA editing of deoxycytidines to deoxyuridines (dC-to-dU) [40, 41]. It was previously thought that the AID/APOBEC family of deaminases (AADs) was restricted to vertebrates. Nevertheless, further evidence showed that APOBEC-like proteins are also present across diverse metazoan dictyostelid, and algal lineages [17, 42].

In humans, 11 AADs are expressed, including AICDA (AID), APOBEC1, APOBEC2, APOBEC3A, APOBEC3B, APOBEC3C, APOBEC3D, APOBEC3F, APOBEC3G, APOBEC3H and APOBEC4. Phylogenetic analysis predicts that AID and APOBEC2 emerged in jawless fish (agnatha) ~ 500 million years ago, whereas APOBEC1 emerged in birds and reptiles ~ 300 million years ago. Gene duplication and divergence gave rise to the APOBEC3 subgroup in mammals ~ 100 million years ago, while APOBEC4 seems to have appeared ~ 20 million years ago [43]. Evolutionary analysis however, suggests that AADs emerged from bacterial lateral transfer; and are divided into secreted deaminases (SNADs) and classical AADs, which diversified and evolved rapidly with a widespread distribution across the tree of life [42, 44]. A unique example of cytidine deaminase acting on tRNA base C8 (CDAT8) enzyme was found in archaea, in Methanopyrus kandleri [45].

Alternative U-to-C and G-to-A editing

RNA editing can also occur in the form of U-to-C and G-to-A, called “alternative mRNA editing”. U-to-C was initially identified in the mRNA of Wilm’s tumor 1 (WT1) human transcript [46]. G-to-A editing is another alternative editing process, detected in the heterogeneous nuclear ribonucleoprotein K (hnRNP K) protein of colorectal cancer and surrounding tissues [47]. The precise mechanism of these RNA editing types is still under investigation. However, recent evidence suggests that APOBEC3A is implicated in G-to-A editing in WT1 transcripts, opening the door for unravelling these alternative processes [48].

Cellular localisation and tissue specificity of RNA editing

From the very beginning of the discovery of RNA editing, it was evident that this modification process can be tissue-specific. The hepatic apolipoprotein-B100 was found to be truncated in enterocytes through a C-to-U change and production of the intestinal apolipoprotein-B48 [4]. In cockroaches, U-to-C and A-to-I editing events of the sodium channel gene (BgNav) were observed in a tissue/cell-specific (ovary, gut, leg and nerve cord) and development-specific manner, thus, generating functional variants of sodium channels [49]. In plants, > 100 C-to-U edits were found in grape mitochondria, whereas 28% of them are significantly tissue-specific [50].

The regulation of RNA editing and tissue specificity can be closely observed during development [51]. For instance, ADAR2 deficient mice exhibit reduced RNA editing activity and are prone to seizures and early mortality [52]. In addition, Adar5G1 null mutant flies lack editing events in hundreds of central nervous system (CNS) transcripts and have defective locomotion, neurodegeneration and reduced survival [53]. In Drosophila, mice and human studies, there were significantly more A-to-I edits found in the brain compared to other tissues [54,55,56]. In the global scale, ADAR1 is the primary editor of repetitive sites (i.e., Alu repeats), ADAR2 is the primary editor of non-repetitive coding sites, while ADAR3 mainly inhibits editing [57]. On the other hand, ADATs seem to be expressed ubiquitously in human tissues [58].

In the cell, pre-mRNA editing is often constrained in the nucleus, hence the localisation of most ADAR enzymes. ADAR1 and ADAR2 are in steady motion all through the nucleolus and are recruited at regions of editing substrate accumulation like in the nucleoplasm [59]. ADAR1p150 is the most common ADAR1 isoform to be found outside the nucleus, trafficking between nucleus and cytoplasm [60]. It can bind to Exportin-1 and translocates in the cytoplasm where it edits (A-to-I) the 3′UTR of dsRNAs [61]. The ADAR1p110 isoform can also shuttle between the nucleus and cytoplasm, but it is predominantly nuclear and constitutively expressed [62]. ADAR2 is considered to be nuclear, since it is rapidly degraded by E3 ubiquitin ligase WWP2 in the cytoplasm, while ADAR3 appears to transiently translocate from the cytoplasm to the nucleus upon neuronal activation [63, 64].

The flux of AADs from the nucleus to the cytoplasm and reverse, is characterised by substantial tissue specificity [41]. AID is expressed in pluripotent tissues like embryonic germ and stem cells and oocytes [65]. APOBEC1 is primarily observed in the intestine of most mammals [66]. APOBEC2 is expressed in the skeletal muscle and cardiac tissue where it is essential for the development of muscles [67]. The subgroup of APOBEC3A-G proteins are heavily implicated in antiviral innate immunity; hence, they are found in immune cell populations [68]. Each family member has different subcellular localisations; namely APOBEC3A, APOBEC3C and APOBEC3H can be either nuclear or cytoplasmic, APOBEC3D, APOBEC3F and APOBEC3G are cytoplasmic, while APOBEC3B is mainly nuclear [69]. APOBEC4 is probably expressed in testes [70].

Cytidine/adenosine deaminase structural features and RNA targets

All AADs have a distinct zinc-dependent cytidine (C) or deoxycytidine (dC) deaminase domain (ZDD) (Fig. 2a) [41]. Most of them (AID, APOBEC1, APOBEC2, APOBEC3A, APOBEC3C, APOBEC3H and APOBEC4) have one ZDD while the others (APOBEC3B, APOBEC3D, APOBEC3F and APOBEC3G) have two ZDD domains, in tandem. APOBEC proteins with two ZDDs have a catalytically active C-terminal domain, and an inactive N-terminal domain [71]. APOBEC1 requires a trans-acting element, RNA binding protein cofactor (A1CF) or RNA-binding protein RBM47, and a cis-acting motif composed of 11 nucleotides, termed the mooring sequence [72,73,74].

Fig. 2

Human cytidine and adenosine deaminase family members. a The cytidine deaminase AID/APOBECs family is shown. Activation-induced cytidine deaminase (AID or AICDA) and all apolipoprotein B mRNA editing enzyme-catalytic polypeptide-like (APOBECs) have one catalytically active cytidine or deoxycytidine deaminase domain (ZDD). APOBEC3 diversifies in 7 submembers (APOBEC3A, APOBEC3B, APOBEC3C, APOBEC3D, APOBEC3F, APOBEC3G, APOBEC3H) whereas some have dual deaminase domain structures but the one in the N terminus is catalytically inactive. b The adenosine deaminase ADARs, ADATs and ADADs families are shown. Three members of the adenosine deaminase acting on RNAs (ADAR1, ADAR2, ADAR3). Two isoforms are known for ADAR1, ADAR1-p150 and ADAR1-p110 and harbour Z-DNA-binding domains. ADAR3 has a unique arginine-rich R domain. Three members of the adenosine deaminase acting on tRNAs (ADAT1, ADAT2 ADAT3). Up to three repeats of the dsRNA binding domain (dsRBD) and a catalytic deaminase domain are present in adenosine deaminases. Two adenosine deaminase domain-containing proteins (ADAD1, ADAD2) are also known as TENR and TENRL respectively. Amino acid length and motifs retrieved from UniProt database [198]. Length is not drawn to scale

In humans, three ADAR enzymes are expressed (Fig. 2b). ADAR1 has two splice variants, ADAR1p110 and ADAR1p150. Three dsRNA binding domains (dsRBD) are found in ADAR1, and two in ADAR2 and ADAR3, respectively [75]. The C-terminal regions are catalytically active but a homodimerization is needed for A-to-I activity [76]. ADAR1 also contains two Z-DNA binding domains, which are required for its localisation to stress granules [77]. An arginine-rich single-stranded RNA (ssRNA) binding domain (R domain) is uniquely present in the N-terminal region of ADAR3 [78].

RNA editing occurs primarily within noncoding regions, and only a small percentage takes place in coding regions resulting in amino acid change. In humans, A-to-I editing mostly occurs in introns and untranslated regions (UTRs) of protein coding genes [79]. A-to-I editing sites seem to happen more frequently in 3′ UTRs than in 5′-UTRs [80, 81]. Alu repeats, a repetitive short interspersed element (SINE), is the most favorable target of RNA editing. Up to 700,000 Alu elements are present in humans and can harbor at least 100 million A-to-I editing sites [82].

MicroRNAs (miRNAs) are also processed by ADARs (ADAR1 and ADAR2) for A-to-I editing in order to control miRNA biogenesis [83, 84]. In addition, evidence of RNA editing competition with RNA interference (RNAi) dsRNA substrates, suggests that ADARs act as modulators of the RNAi machinery [85]. Furthermore, tRNA can be edited by ADATs, as mentioned above. In humans, A-to-I edits at positions 34 and 37 of tRNAAla have been reported [86]. Even though widespread in plants, rRNA editing has not yet been observed in humans [87].

APOBEC1 has a preference for AU-rich sequences within mRNA 3′ UTRs [88]. APOBEC3 proteins are significantly active against endogenous retroelements and retroviruses, where they can target SINE RNA but also long interspersed nuclear elements (LINEs) [89, 90]. APOBEC2 and APOBEC4 are still under investigation since little evidence have emerged for their activities [70, 91, 92].

Target detection by next-generation sequencing (NGS) and computational processing

Classical molecular biology and sequencing techniques have contributed significantly in the discovery and early detection of RNA editing. The emergence NGS technologies in combination with accurate bioinformatic pipelines made possible the detection of thousands of new RNA editing sites [81, 93]. Matched DNA and RNA sequencing in single samples identifies RNA–DNA differences that can provide a plethora of possible edits [79, 94].

High-throughput sequencing methods like RNA-seq have now become readily available in the scientific community, leading to a wealth of accumulating transcriptomic data of various tissues and cell populations [95]. This extensive repertoire of information and the above techniques demand extensive computational tools. Besides software packages that identify RNA editing sites from matched sequencing samples, increasing programming is given to prediction models. Candidate RNA editing sites can be predicted by software packages like GIREMI (Genome-independent Identification of RNA Editing by Mutual Information,, RNAEditor ( and DeepRed ( from RNA-seq data in the absence of relevant genomic data [10, 80, 96].

RNA editing databases

Furthermore, freely available databases are now available, where one can explore RNA editing collections, in humans and other model organisms. The RNA Editing ATLAS ( is the first human inosinome atlas, comprising > 3.0 million A-to-I events identified in six tissues from three healthy individuals. Matched directional total-RNA-Seq and whole genome sequence datasets were generated and analysed within a dedicated computational framework, also capable of detecting hyper-edited reads. Inosinome profiles within the RNA Editing ATLAS are tissue specific and edited gene sets, consistently show enrichment of genes involved in neurological disorders and cancer. The RNA Editing ATLAS reports that overall, the frequency of editing varies, but is strongly correlated with the expression levels of ADAR [97].

Moreover, the RADAR database ( presents a comprehensive collection of A-to-I editing sites in human, mouse, and fly transcripts [98].

dbRES is another RNA editing database that contains only experimentally validated data that were manually collected from literature reporting related experiment results or from GeneBank [99].

REDIportal ( is another freely available database collecting > 4.5 millions of A-to-I editing events in 55 body sites of 150 healthy individuals from the GTEx project. In REDIportal, RNA editing sites can be searched by genomic region, gene name and other relevant features as the tissue of origin. Recently, REDIportal started collecting A-to-I events in non-human organisms [100].

REDIdb in another RNA editing database, where one can assess general information, editing features and the nucleotide genomic sequence and the corresponding transcript of the entry, annotated as cDNA [101].

Furthermore, LNCediting ( provides a comprehensive resource for the functional prediction of RNA editing in long noncoding RNAs (lncRNAs) in Homo sapiens, Macaca mulatta, Mus musculus and Drosophila melanogaster [102].

Role in health

Modulators of alternative splicing and transcriptional control

RNA editing can result in a number of functional effects (Fig. 3). A-to-I editing can produce or delete splice sites, regulating the production of different protein isoforms with varying traits [103,104,105]. For instance, in neurons RNA editing regulates synaptic transmission through editing and splicing of glutamate receptor pre-mRNA [103, 104]. Sequencing the RNA of different subcellular fraction showed that most (> 95%) of the A-to-I edits happen during mRNA maturation with ~ 500 editing sites in the 3′ acceptor sequences [106]. Modifications in these splice sites can easily result in alternative splicing of the associated exons. Interestingly, ADAR itself is regulated by RNA editing in order to produce the Drosophila dADAR variants [34]. In rats, ADAR2 edits twice its own pre-mRNA to produce four isoforms in order to control its own expression [105]. Moreover, ADAR1 and ADAR2 are related to spliceosomal proteins [107]. A nice example of RNA editing-mediated splice variant is the nuclear prelamin A recognition factor (NARF) exon 8 evolution in primates [108].

Fig. 3

Functional roles of RNA editing. Adenosine and cytidine deaminases target RNAs molecules and modify their sequence affecting multiple processes. A-to-I or C-to-U modifications in RNA transcripts are reflected in the folding of the RNA structure influencing its structural stability and binding accessibility for further processing [199]. mRNA abundance and gene expression is regulated by miRNA or miRNA target editing, influencing gene silencing by RNA degradation [84, 156]. Protein diversity is another outcome of RNA editing since it can create or abolish splicing sites regulating alternative splicing [54]. Moreover, editing in coding regions can recode amino acids and create an alternative protein with distinct functionalities [4]. Recoding protein example was built with SWISS-MODEL workspace [200]

RNA editing regulates neuronal dynamics

Studies in animal models (Drosophila and mice) implicated ADARs and A-to-I editing post-transcriptionally regulates circadian rhythm and sleep [109, 110]. ADAR1 deficiency in Drosophila led to constitutive release of neurotransmitter in glutamatergic neurons promoting sleep [111]. RNA editing association with neuronal activity is supported by the fact that ADAR2 and ADAR3 are generally highly expressed in the brain and CNS [64, 78]. During development and normal function, RNA editing acts as a regulator of neurotransmission and signal transduction by editing AMPA and kainate glutamate receptor subunits at the Q/R site making them impermeable to Ca2+, whilst regulating Ca2+ influx; a prerequisite for normal neuronal function (Fig. 1b) [112]. Additional properties (like targeting miRNA and circular RNA) have been attributed to RNA editing strengthening its role as a powerful and dynamic regulator of neuronal function, brain development and protection [113,114,115].

Fundamental players in innate and adaptive immunity

B lymphocytes produce a repertoire of antibodies in order to protect the organism from foreign and invading pathogens. This process is facilitated by genetic mechanisms such as gene rearrangement and conversion, somatic hypermutation (SHM) and class switch recombination (CSR). AID plays an important role in antibody diversity through both SHM and CSR [116]. Two models have been proposed for AID activity in immunoglobulins, DNA or RNA editing [117]. Most evidence however, support the notion that AID targets DNA hotspots; like the WRC motif of the variable region (where W = A or T and R = A or G) or the DGYW/WRCH motif (where G:C is the mutable position; D = A/G/T, H = T/C/A) of switch (S) region sequence causing U:G mismatches [118, 119]. These, either become point mutations and SHM or trigger double strand breaks (DSB) and CSR, respectively [119, 120].

APOBECs are heavily implicated in viral immune defense. APOBEC3G is the first family member that was identified as an antiviral factor and the most well studied [121, 122]. In the human immunodeficiency virus (HIV-1), APOBEC3G deaminates C-to-U in the reverse transcribed viral cDNA minus strand, causing G-to-A hypermutation on the plus strand that can lead to cDNA instability and inhibition of viral replication [123]. APOBEC3D, APOBEC3F and APOBEC3G in humans, and and APOBEC3H in the rhesus macaque are also shown to inhibit HIV-1 [124]. However, the HIV-1 virion infectivity factor (Vif) can ubiquinate APOBEC3 proteins targeting them for proteasomal degradation [123, 124].

Different levels of antiretroviral activity among APOBECs have also been demonstrated against Simian immunodeficiency virus (SIV), adeno-associated virus type 2 (AAV-2), murine leukemia virus (MLV), equine infectious anemia virus (EIAV), and foamy viruses (FVs) [89, 125,126,127,128]. Furthermore, hepatitis B virus (HBV) replication can be hampered by APOBEC3G in a deamination-dependent and -independent manner [129, 130].

Apart from viral C-to-U DNA editing, APOBEC3-mediated C-to-U RNA editing is also observed in HIV genomic RNA [131]. APOBECs also target RNA viruses such as members of the paramyxoviruses, measles and mumps, but also respiratory syncytial viruses (RSV) by C-to-U hypermutations [132].

ADARs (especially ADAR1 isoforms) target RNA viruses like measle virus, infuenza A virus, Rift Valley fever virus (RVFV), lymphocytic choriomeningitis virus (LCMV) and hepatitis C virus (HCV) by A-to-I hypermutation [133,134,135,136,137]. However, they also exhibit a significant degree of proviral effects (reviewed in [138]).

AID/APOBEC proteins have the capacity to restrict LTR and non-LTR retrotransposons [90, 139]. Retrotransposons are genetic elements able to transport themselves and multiply in the genome. LTR retrotransposons are mainly inhibited by APOBEC3 proteins through C-to-U DNA hyperediting [140]. Long interspersed nuclear element 1 (LINE-1) is the only functional family of transposable elements in humans [141]. Accumulating evidence supports that LINE-1 retrotransposon inhibition by AID/APOBECs and ADARs is editing-independent (reviewed in [142, 143]). Non-autonomous SINE-1 is another member of the non-LTR retrotransposons. The SINE Alu repeats exhibit strong A-to-I editability by ADARs and account for the majority of all editing sites [82, 144].

Role in disease

Aberrant involvement in human diseases

The contribution of RNA editing in the brain and CNS is not always beneficial. Evidence have emerged implicating RNA editing in the pathogenesis of neurodegenerative diseases like amyotrophic lateral sclerosis (ALS), Parkinson and Alzheimer diseases [145]. In ALS, mutation of the optineurin (OPTN) gene was found to be triggered by recombination between Alu elements, a favourable target of A-to-I editing [146]. In addition, down-regulation of ADAR2 is detrimental towards the physiological editing of glutamate receptor Q/R site and calcium regulation, leading to neuronal hyper-excitability and autophagy, which can contribute to the death of motor neurons in ALS [147, 148]. Reduction of glutamate receptor editing was also observed in Alzheimer disease [149]. A number of studies investigated editing patterns of the serotonin 2C receptor in patients with various psychological disorders like schizophrenia, depression, bipolar disorder, drug addiction and even in individuals who committed suicide (reviewed in [150]). Despite significant associations, results are conflicting and inconclusive requiring further investigation. ADAR mutations and altered editing have also been linked to autoimmune diseases like Aicardi-Goutières syndrome and systemic lupus erythematosus, respectively [151, 152].

RNA editing in cancer

Cancer pathogenesis is primarily attributed to genetic mutations in proto-oncogenes and tumour suppressor genes, which transform a normal cell into a malignant cancer cell. The rapid progress of NGS technologies and the readily available transcriptomic data have revealed a significant contribution of RNA editing in the pathogenesis and progression of cancer (Table 1) [153, 154]. Transcriptome analysis of various tumours showed deferential RNA editing levels depending on the cancer type. Decreased A-to-I editing patterns have been identified in brain, kidney, lung, prostate and testis tumours, with significant global hypo-editing of Alu elements [155]. On the other hand, recent studies on multiple cancer tissues found elevated editing levels in intergenic, intronic and 3′ UTR regions of most cancer types especially in thyroid, head and neck, breast and lung cancer tissues compared to their matched controls, which in most cases is associated with worst patient survival [153, 154]. These data clearly indicate that the editing level, high or low, can have different roles in the pathogenesis of cancer and different clinical outcomes in the progression of the disease.

Table 1 RNA editing events in cancer

As mentioned above, editing sites fall within noncoding regions of the genome, and this is also true for cancer genomes. MiRNAs are also important editing targets in cancer [156]. In glioblastoma, the anti-tumour function of ADAR2 was revealed, as it can regulate a large number of miRNAs, including miR-21 and miR-222/-221 precursors and reduce the expression of their mature oncogenic miRNAs [157]. Edited miR-376a was also found to exhibit anti-tumour effects in glioblastoma by targeting the RAP2A oncogene (a member of RAS oncogene family), while its unedited form targets the autocrine motility factor receptor (AMFR) receptor, thus promoting invasiveness [158]. In melanoma, unedited miR-455-5p promotes cancer growth and metastasis by inhibiting the tumour suppressor gene cytoplasmic polyadenylation element binding protein 1 (CPEB1) [159]. Overexpression of ADAR1 in lung cancer has been associated with poor outcome as it enhances editing of miR-381-associated stemness and chemoresistance, in addition to editing NEIL1, a DNA repair gene [160]. Besides unique miRNA editing events in individual cancer types, a recent study investigated 20 different cancers from The Cancer Genome Atlas (TCGA) and identified 19 RNA editing hotspots [161]. Among them, miR-200b, a tumour suppressor, was found to promote invasion and migration when edited by suppressing LIFR, a characteristic metastasis suppressor; while been associated with worst patient survival. Furthermore, it has been shown that editing of 3′ UTR can regulate miRNA binding sites affecting post-transcriptional regulation of tumour suppressor genes and oncogenes like MDM2 which promoted carcinogenesis [162]. Another study identified the secretion levels of miR-200 family members to be regulated by ADAR2 and protein kinase Cζ (PKCζ) axis in promoting liver metastasis in colorectal cancer [163].

Editing in the protein coding region is far less frequent, but it has major consequences in the regulation and function of the affected gene. Sixty recoding events have been identified in a large scale study that are associated with tumours [153]. Other recoding events include the RNA editing of GABRA3 and COPA in breast cancer, which suppresses AKT-mediated metastasis or promotes proliferation, invasion and metastasis, respectively [164, 165]. Gastric cancers display profound misediting of RNA since they exhibit significant genomic gain of ADAR1 and loss of ADAR2 genes acting as oncogenic and tumour suppressive mediators; hence the failure of ADAR2 recoding of PODXL gene allows its tumorigenic potential [166]. Other RNA editing events on coding sequences include the RHOQ gene in colorectal cancer, IGFBP7 and SLC22A3 in esophageal cancer and CDC14B in glioblastoma [167,168,169,170]. The most well characterised recoding event is the editing of the coding sequence of AZIN1 mRNA in cancers [162, 171,172,173]. In liver cancer, an ADAR1 A-to-I editing of AZIN1 transcripts results in a serine to glycine substitution at residue 367. This affects protein conformation and induces a nuclear translocation leading to tumour initiation potential and progression [171]. In colorectal cancer, AZIN1 RNA editing exhibits cancer stemness and augments oncogenic potential; while in esophageal squamous cell carcinoma it is associated with aggressive tumour behaviour [172, 173].

Cytidine deaminase mutations have also been observed in an array of human cancers [183]. APOBEC misregulation in cancer, contributes to genetic instability and affects prognosis depending on the type of cancer [181, 184]. APOBEC3A and APOBEC3B overexpression has been associated with localised C-to-T and/or C-to-G hypermutations termed “kataegis” in a number of cancers, suggesting that these signatures are important in cancer mutagenesis [183, 185, 186]. In breast cancer, APOBEC3B overexpression has been linked to DNA C-to-U editing and TP53 inactivation while it is correlated with poor treatment response and poor outcome for estrogen receptor-positive (ER+) tumours [180, 181, 187]. Furthermore, increased expression of APOBEC3G in colorectal tumours has been found to promote hepatic metastasis through inhibition of miR-29 mediated suppression of matrix metalloproteinase 2 (MMP2) [182]. However, in bladder cancer, APOBEC-low expressing tumours often present mutations in FGFR3 and RAS family of oncogenes; whereas APOBEC-high expressing tumours usually have mutations in DNA damage response genes and chromatin regulatory genes, an enhanced immune response and better overall survival [184]. C-to-U RNA editing by APOBEC1 is also observed in cancers and is especially enriched in lung tumorigenesis and hepatocellular carcinoma [188, 189].

Another important role of RNA editing in cancer is its involvement in the ability of tumours to evade immune responses. APOBEC overexpression and kataegis has been associated with programmed cell-death receptor-1 (PD-1) overexpression, an immune checkpoint molecule, leading to immune tolerance and exhaustion [190]. Conversely, RNA editing can elicit immune responses in tumours through increased levels of edited peptides that act as antigens, stimulating T cell responses [191].

In cancers, RNA editing not only changes the sequence of RNAs and their expression but also contributes to proteomic diversity [165, 192]. Combining the TCGA genomic data and the Clinical Proteomic Tumor Analysis Consortium (CPTAC) proteomic data (, Peng and colleagues present evidence that the A-to-I RNA editing events in cancer are manifested in protein diversity of cancer cells through changes in amino acid sequences. These intriguing observations suggest that RNA editing is a novel source of cancer protein heterogeneity.

Future perspectives in diagnosis and treatment

Increasing evidence proclaims that the levels of RNA editing, along with the expression of adenosine and cytidine deaminases and specifically edited genes (especially tumour suppressors and oncogenes), could all be used as important prognostic biomarkers in the pathogenesis and progression of cancer [165, 166, 171]. Deregulated expression patterns of ADARs and APOBECs observed between tumour and normal tissues, as well as within cancer types, revealed a promising scheme of clinical value towards a better understanding of cancer development and its corresponding treatment [154, 180, 183]. Distinctly edited genes like the ones discussed above, play a significant role in tumour pathophysiology [166, 172, 173]. Treatment strategies have also been challenged by nonsynonymous RNA editing events and expression levels, since they display considerable effects on drug sensitivity, as tamoxifen resistance in ER2+ breast cancers [154, 187]. Interestingly these processes also provide new therapeutic targets. ADAR inhibitors is a novel treatment strategy against ADAR-overexpressing tumours, such as in breast and lung, with positive results [193]. In addition, APOBEC inhibitors are still in the early stages of development, due to their recent involvement in cancer, but increasing interest is been directed towards this aspect, as well [194].

Overall, the modulation and application of RNA editing is an area of great potential. Besides traditional deaminase inhibitors to control expression, molecular tools such as antisense oligonucleotides are potent and selective inhibitors of RNA editing on targeted RNAs [195]. Engineered RNA editing-guided activity is a technique particularly useful for hypo-edited-related diseases, such as in prostate and brain cancers, but also in correcting disease-promoting genetic mutations [155, 196, 197].


In this review, we have discussed the diverse aspects of RNA editing, from its discovery to physiological function and involvement in human diseases. The road ahead still seems bright for RNA editing with new exciting findings from modern transcriptomics and new therapeutic developments in associated disorders.

Availability of data and materials

Not applicable.



cytochrome c oxidase subunit II


cytidine to uridine


adenosine to inosine


next-generation sequencing


constructive neutral evolution


double stranded RNA


adenosine deaminase acting on RNA


ADAR-like deaminase


testes nuclear RNA-binding protein


adenosine deaminases that act on tRNAs


tRNA adenosine deaminase


activation-induced cytidine deaminase


apolipoprotein B mRNA editing enzyme


AID/APOBEC family of deaminases


secreted deaminases


cytidine deaminase acting on tRNA base C8


Wilm’s tumor 1 human transcript

hnRNP K:

heterogeneous nuclear ribonucleoprotein K


zinc-dependent cytidine (C) or deoxycytidine (dC) deaminase domain


RNA binding protein cofactor


dsRNA binding domain

R domain:

single-stranded RNA binding domain


short interspersed element


long interspersed nuclear element


Genome-independent Identification of RNA Editing by Mutual Information


long noncoding RNAs




nuclear prelamin A recognition factor


somatic hypermutation


class switch recombination


virion infectivity factor


Simian immunodeficiency virus


adeno-associated virus type 2


murine leukemia virus


equine infectious anemia virus


foamy virus


Rift Valley fever virus


lymphocytic choriomeningitis virus


hepatitis C virus


amyotrophic lateral sclerosis




RAS related protein 2a (a member of the RAS oncogene family)


autocrine motility factor receptor


cytoplasmic polyadenylation element binding protein 1


The Cancer Genome Atlas


transformed mouse 3T3 cell double minute 2/MDM2 proto-oncogene


protein kinase Cζ


antizyme inhibitor 1


estrogen receptor


matrix metalloproteinase 2


fibroblast growth factor receptor 3


Clinical Proteomic Tumor Analysis Consortium


  1. 1.

    Iwanami Y, Brown GM. Methylated bases of ribosomal ribonucleic acid from HeLa cells. Arch Biochem Biophys. 1968;126:8–15.

  2. 2.

    Cantara WA, Crain PF, Rozenski J, McCloskey JA, Harris KA, Zhang X, et al. The RNA modification database, RNAMDB: 2011 update. Nucleic Acids Res. 2011;39:D195–201.

  3. 3.

    Benne R, Van den Burg J, Brakenhoff JP, Sloof P, Van Boom JH, Tromp MC. Major transcript of the frameshifted coxII gene from trypanosome mitochondria contains four nucleotides that are not encoded in the DNA. Cell. 1986;46:819–26.

  4. 4.

    Powell LM, Wallis SC, Pease RJ, Edwards YH, Knott TJ, Scott J. A novel form of tissue-specific RNA processing produces apolipoprotein-B48 in intestine. Cell. 1987;50:831–40.

  5. 5.

    Holley RW, Madison JT, Zamir A. A new method for sequence determination of large oligonucleotides. Biochem Biophys Res Commun. 1964;17:389–94.

  6. 6.

    Holley RW, Apgar J, Everett GA, Madison JT, Marquisee M, Merrill SH, et al. Structure of a ribonucleic acid. Science. 1965;147:1462–5.

  7. 7.

    Chow LT, Gelinas RE, Broker TR, Roberts RJ. An amazing sequence arrangement at the 5′ ends of adenovirus 2 messenger RNA. Cell. 1977;12:1–8.

  8. 8.

    Berget SM, Moore C, Sharp PA. Spliced segments at the 5′ terminus of adenovirus 2 late mRNA. Proc Natl Acad Sci USA. 1977;74:3171–5.

  9. 9.

    Ramaswami G, Li JB. Identification of human RNA editing sites: a historical perspective. Methods. 2016;107:42–7.

  10. 10.

    Ouyang Z, Liu F, Zhao C, Ren C, An G, Mei C, et al. Accurate identification of RNA editing sites from primitive sequence with deep neural networks. Sci Rep. 2018;8:6005.

  11. 11.

    Chow CS, Lamichhane TN, Mahto SK. Expanding the nucleotide repertoire of the ribosome with post-transcriptional modifications. ACS Chem Biol. 2007;2:610–9.

  12. 12.

    Noack F, Calegari F. Epitranscriptomics: a new regulatory mechanism of brain development and function. Front Neurosci. 2018;12:85.

  13. 13.

    O’Connell MA, Mannion NM, Keegan LP. The epitranscriptome and innate immunity. PLoS Genet. 2015;11:e1005687.

  14. 14.

    Freyer R, Kiefer-Meyer MC, Kossel H. Occurrence of plastid RNA editing in all major lineages of land plants. Proc Natl Acad Sci USA. 1997;94:6285–90.

  15. 15.

    Grice LF, Degnan BM. The origin of the ADAR gene family and animal RNA editing. BMC Evol Biol. 2015;15:4.

  16. 16.

    Gerber AP, Keller W. RNA editing by base deamination: more enzymes, more targets, new mysteries. Trends Biochem Sci. 2001;26:376–84.

  17. 17.

    Conticello SG, Thomas CJ, Petersen-Mahrt SK, Neuberger MS. Evolution of the AID/APOBEC family of polynucleotide (deoxy)cytidine deaminases. Mol Biol Evol. 2005;22:367–77.

  18. 18.

    Navaratnam N, Fujino T, Bayliss J, Jarmuz A, How A, Richardson N, et al. Escherichia coli cytidine deaminase provides a molecular model for ApoB RNA editing and a mechanism for RNA substrate recognition. J Mol Biol. 1998;275:695–714.

  19. 19.

    Bar-Yaacov D, Mordret E, Towers R, Biniashvili T, Soyris C, Schwartz S, et al. RNA editing in bacteria recodes multiple proteins and regulates an evolutionarily conserved toxin-antitoxin system. Genome Res. 2017;27:1696–703.

  20. 20.

    Slavov D, Crnogorac-Jurcevic T, Clark M, Gardiner K. Comparative analysis of the DRADA A-to-I RNA editing gene from mammals, pufferfish and zebrafish. Gene. 2000;250:53–60.

  21. 21.

    Keegan LP, McGurk L, Palavicini JP, Brindle J, Paro S, Li X, et al. Functional conservation in human and drosophila of metazoan ADAR2 involved in RNA editing: loss of ADAR1 in insects. Nucleic Acids Res. 2011;39:7249–62.

  22. 22.

    Gray MW. Evolutionary origin of RNA editing. Biochemistry. 2012;51:5235–42.

  23. 23.

    Covello PS, Gray MW. On the evolution of RNA editing. Trends Genet. 1993;9:265–8.

  24. 24.

    Stoltzfus A. On the possibility of constructive neutral evolution. J Mol Evol. 1999;49:169–81.

  25. 25.

    Kimura M. Evolutionary rate at the molecular level. Nature. 1968;217:624–6.

  26. 26.

    Stoltzfus A. Constructive neutral evolution: exploring evolutionary theory’s curious disconnect. Biol Direct. 2012;7:35.

  27. 27.

    Bass BL, Weintraub H. A developmentally regulated activity that unwinds RNA duplexes. Cell. 1987;48:607–13.

  28. 28.

    Nishikura K. A-to-I editing of coding and non-coding RNAs by ADARs. Nat Rev Mol Cell Biol. 2016;17:83–96.

  29. 29.

    Bazak L, Haviv A, Barak M, Jacob-Hirsch J, Deng P, Zhang R, et al. A-to-I RNA editing occurs at over a hundred million genomic sites, located in a majority of human genes. Genome Res. 2014;24:365–76.

  30. 30.

    Melcher T, Maas S, Herb A, Sprengel R, Higuchi M, Seeburg PH. RED2, a brain-specific member of the RNA-specific adenosine deaminase family. J Biol Chem. 1996;271:31795–8.

  31. 31.

    Melcher T, Maas S, Herb A, Sprengel R, Seeburg PH, Higuchi M. A mammalian RNA editing enzyme. Nature. 1996;379:460–4.

  32. 32.

    Jin Y, Zhang W, Li Q. Origins and evolution of ADAR-mediated RNA editing. IUBMB Life. 2009;61:572–8.

  33. 33.

    Tonkin LA, Saccomanno L, Morse DP, Brodigan T, Krause M, Bass BL. RNA editing by ADARs is important for normal behavior in caenorhabditis elegans. EMBO J. 2002;21:6025–35.

  34. 34.

    Palladino MJ, Keegan LP, O’Connell MA, Reenan RA. dADAR, a drosophila double-stranded RNA-specific adenosine deaminase is highly developmentally regulated and is itself a target for RNA editing. RNA. 2000;6:1004–18.

  35. 35.

    Schumacher JM, Lee K, Edelhoff S, Braun RE. Distribution of tenr, an RNA-binding protein, in a lattice-like network within the spermatid nucleus in the mouse. Biol Reprod. 1995;52:1274–83.

  36. 36.

    Hough RF, Bass BL. Analysis of xenopus dsRNA adenosine deaminase cDNAs reveals similarities to DNA methyltransferases. RNA. 1997;3:356–70.

  37. 37.

    Savva YA, Rieder LE, Reenan RA. The ADAR protein family. Genome Biol. 2012;13:252.

  38. 38.

    Wolf J, Gerber AP, Keller W. tadA, an essential tRNA-specific adenosine deaminase from escherichia coli. EMBO J. 2002;21:3841–51.

  39. 39.

    Torres AG, Pineyro D, Filonava L, Stracker TH, Batlle E, Ribas de Pouplana L. A-to-I editing on tRNAs: biochemical, biological and evolutionary implications. FEBS Lett. 2014;588:4279–86.

  40. 40.

    Knisbacher BA, Gerber D, Levanon EY. DNA editing by APOBECs: a genomic preserver and transformer. Trends Genet. 2016;32:16–28.

  41. 41.

    Smith HC, Bennett RP, Kizilyer A, McDougall WM, Prohaska KM. Functions and regulation of the APOBEC family of proteins. Semin Cell Dev Biol. 2012;23:258–68.

  42. 42.

    Krishnan A, Iyer LM, Holland SJ, Boehm T, Aravind L. Diversification of AID/APOBEC-like deaminases in metazoa: multiplicity of clades and widespread roles in immunity. Proc Natl Acad Sci USA. 2018;115:E3201–10.

  43. 43.

    Salter JD, Bennett RP, Smith HC. The APOBEC protein family: united by structure, divergent in function. Trends Biochem Sci. 2016;41:578–94.

  44. 44.

    Iyer LM, Zhang D, Rogozin IB, Aravind L. Evolution of the deaminase fold and multiple origins of eukaryotic editing and mutagenic nucleic acid deaminases from bacterial toxin systems. Nucleic Acids Res. 2011;39:9473–97.

  45. 45.

    Randau L, Stanley BJ, Kohlway A, Mechta S, Xiong Y, Soll D. A cytidine deaminase edits C to U in transfer RNAs in archaea. Science. 2009;324:657–9.

  46. 46.

    Sharma PM, Bowman M, Madden SL, Rauscher FJ 3rd, Sukumar S. RNA editing in the wilms’ tumor susceptibility gene, WT1. Genes Dev. 1994;8:720–31.

  47. 47.

    Klimek-Tomczak K, Mikula M, Dzwonek A, Paziewska A, Karczmarski J, Hennig E, et al. Editing of hnRNP K protein mRNA in colorectal adenocarcinoma and surrounding mucosa. Br J Cancer. 2006;94:586–92.

  48. 48.

    Niavarani A, Currie E, Reyal Y, Anjos-Afonso F, Horswell S, Griessinger E, et al. APOBEC3A is implicated in a novel class of G-to-A mRNA editing in WT1 transcripts. PLoS ONE. 2015;10:e0120089.

  49. 49.

    Song W, Liu Z, Tan J, Nomura Y, Dong K. RNA editing generates tissue-specific sodium channels with distinct gating properties. J Biol Chem. 2004;279:32554–61.

  50. 50.

    Picardi E, Horner DS, Chiara M, Schiavon R, Valle G, Pesole G. Large-scale detection and analysis of RNA editing in grape mtDNA by RNA deep-sequencing. Nucleic Acids Res. 2010;38:4755–67.

  51. 51.

    Wahlstedt H, Daniel C, Enstero M, Ohman M. Large-scale mRNA sequencing determines global regulation of RNA editing during brain development. Genome Res. 2009;19:978–86.

  52. 52.

    Higuchi M, Maas S, Single FN, Hartner J, Rozov A, Burnashev N, et al. Point mutation in an AMPA receptor gene rescues lethality in mice deficient in the RNA-editing enzyme ADAR2. Nature. 2000;406:78–81.

  53. 53.

    Li X, Overton IM, Baines RA, Keegan LP, O’Connell MA. The ADAR RNA editing enzyme controls neuronal excitability in drosophila melanogaster. Nucleic Acids Res. 2014;42:1139–51.

  54. 54.

    Mazloomian A, Meyer IM. Genome-wide identification and characterization of tissue-specific RNA editing events in D. melanogaster and their potential role in regulating alternative splicing. RNA Biol. 2015;12:1391–401.

  55. 55.

    Hwang T, Park CK, Leung AK, Gao Y, Hyde TM, Kleinman JE, et al. Dynamic regulation of RNA editing in human brain development and disease. Nat Neurosci. 2016;19:1093–9.

  56. 56.

    Huntley MA, Lou M, Goldstein LD, Lawrence M, Dijkgraaf GJ, Kaminker JS, et al. Complex regulation of ADAR-mediated RNA-editing across tissues. BMC Genomics. 2016;17:61.

  57. 57.

    Tan MH, Li Q, Shanmugam R, Piskol R, Kohler J, Young AN, et al. Dynamic landscape and regulation of RNA editing in mammals. Nature. 2017;550:249–54.

  58. 58.

    Maas S, Gerber AP, Rich A. Identification and characterization of a human tRNA-specific adenosine deaminase related to the ADAR family of pre-mRNA editing enzymes. Proc Natl Acad Sci USA. 1999;96:8895–900.

  59. 59.

    Desterro JM, Keegan LP, Lafarga M, Berciano MT, O’Connell M, Carmo-Fonseca M. Dynamic association of RNA-editing enzymes with the nucleolus. J Cell Sci. 2003;116:1805–18.

  60. 60.

    Fritz J, Strehblow A, Taschner A, Schopoff S, Pasierbek P, Jantsch MF. RNA-regulated interaction of transportin-1 and exportin-5 with the double-stranded RNA-binding domain regulates nucleocytoplasmic shuttling of ADAR1. Mol Cell Biol. 2009;29:1487–97.

  61. 61.

    Poulsen H, Nilsson J, Damgaard CK, Egebjerg J, Kjems J. CRM1 mediates the export of ADAR1 through a nuclear export signal within the Z-DNA binding domain. Mol Cell Biol. 2001;21:7862–71.

  62. 62.

    Strehblow A, Hallegger M, Jantsch MF. Nucleocytoplasmic distribution of human RNA-editing enzyme ADAR1 is modulated by double-stranded RNA-binding domains, a leucine-rich export signal, and a putative dimerization domain. Mol Biol Cell. 2002;13:3822–35.

  63. 63.

    Marcucci R, Brindle J, Paro S, Casadio A, Hempel S, Morrice N, et al. Pin1 and WWP2 regulate GluR2 Q/R site RNA editing by ADAR2 with opposing effects. EMBO J. 2011;30:4211–22.

  64. 64.

    Mladenova D, Barry G, Konen LM, Pineda SS, Guennewig B, Avesson L, et al. Adar3 is involved in learning and memory in mice. Front Neurosci. 2018;12:243.

  65. 65.

    Morgan HD, Dean W, Coker HA, Reik W, Petersen-Mahrt SK. Activation-induced cytidine deaminase deaminates 5-methylcytosine in DNA and is expressed in pluripotent tissues: implications for epigenetic reprogramming. J Biol Chem. 2004;279:52353–60.

  66. 66.

    Hadjiagapiou C, Giannoni F, Funahashi T, Skarosi SF, Davidson NO. Molecular cloning of a human small intestinal apolipoprotein B mRNA editing protein. Nucleic Acids Res. 1994;22:1874–9.

  67. 67.

    Liao W, Hong SH, Chan BH, Rudolph FB, Clark SC, Chan L. APOBEC-2, a cardiac- and skeletal muscle-specific member of the cytidine deaminase supergene family. Biochem Biophys Res Commun. 1999;260:398–404.

  68. 68.

    Refsland EW, Stenglein MD, Shindo K, Albin JS, Brown WL, Harris RS. Quantitative profiling of the full APOBEC3 mRNA repertoire in lymphocytes and tissues: implications for HIV-1 restriction. Nucleic Acids Res. 2010;38:4274–84.

  69. 69.

    Vieira VC, Soares MA. The role of cytidine deaminases on innate immune responses against human viral infections. Biomed Res Int. 2013;2013:683095.

  70. 70.

    Rogozin IB, Basu MK, Jordan IK, Pavlov YI, Koonin EV. APOBEC4, a new member of the AID/APOBEC family of polynucleotide (deoxy)cytidine deaminases predicted by computational analysis. Cell Cycle. 2005;4:1281–5.

  71. 71.

    Salter JD, Smith HC. Modeling the embrace of a mutator: APOBEC selection of nucleic acid ligands. Trends Biochem Sci. 2018;43:606–22.

  72. 72.

    Shah RR, Knott TJ, Legros JE, Navaratnam N, Greeve JC, Scott J. Sequence requirements for the editing of apolipoprotein B mRNA. J Biol Chem. 1991;266:16301–4.

  73. 73.

    Sowden MP, Ballatori N, Jensen KL, Reed LH, Smith HC. The editosome for cytidine to uridine mRNA editing has a native complexity of 27S: identification of intracellular domains containing active and inactive editing factors. J Cell Sci. 2002;115:1027–39.

  74. 74.

    Fossat N, Tourle K, Radziewic T, Barratt K, Liebhold D, Studdert JB, et al. C to U RNA editing mediated by APOBEC1 requires RNA-binding protein RBM47. EMBO Rep. 2014;15:903–10.

  75. 75.

    Nishikura K. Functions and regulation of RNA editing by ADAR deaminases. Annu Rev Biochem. 2010;79:321–49.

  76. 76.

    Cho DS, Yang W, Lee JT, Shiekhattar R, Murray JM, Nishikura K. Requirement of dimerization for RNA editing activity of adenosine deaminases acting on RNA. J Biol Chem. 2003;278:17093–102.

  77. 77.

    Ng SK, Weissbach R, Ronson GE, Scadden AD. Proteins that contain a functional Z-DNA-binding domain localize to cytoplasmic stress granules. Nucleic Acids Res. 2013;41:9786–99.

  78. 78.

    Chen CX, Cho DS, Wang Q, Lai F, Carter KC, Nishikura K. A third member of the RNA-specific adenosine deaminase gene family, ADAR3, contains both single- and double-stranded RNA binding domains. RNA. 2000;6:755–67.

  79. 79.

    Park E, Williams B, Wold BJ, Mortazavi A. RNA editing in the human ENCODE RNA-seq data. Genome Res. 2012;22:1626–33.

  80. 80.

    Zhang Q, Xiao X. Genome sequence-independent identification of RNA editing sites. Nat Methods. 2015;12:347–50.

  81. 81.

    Peng Z, Cheng Y, Tan BC, Kang L, Tian Z, Zhu Y, et al. Comprehensive analysis of RNA-seq data reveals extensive RNA editing in a human transcriptome. Nat Biotechnol. 2012;30:253–60.

  82. 82.

    Daniel C, Lagergren J, Ohman M. RNA editing of non-coding RNA and its role in gene regulation. Biochimie. 2015;117:22–7.

  83. 83.

    Luciano DJ, Mirsky H, Vendetti NJ, Maas S. RNA editing of a miRNA precursor. RNA. 2004;10:1174–7.

  84. 84.

    Yang W, Chendrimada TP, Wang Q, Higuchi M, Seeburg PH, Shiekhattar R, et al. Modulation of microRNA processing and expression through RNA editing by ADAR deaminases. Nat Struct Mol Biol. 2006;13:13–21.

  85. 85.

    Scadden AD, Smith CW. RNAi is antagonized by A–>I hyper-editing. EMBO Rep. 2001;2:1107–11.

  86. 86.

    Becker HF, Corda Y, Mathews MB, Fourrey JL, Grosjean H. Inosine and N1-methylinosine within a synthetic oligomer mimicking the anticodon loop of human tRNA(ala) are major epitopes for anti-PL-12 myositis autoantibodies. RNA. 1999;5:865–75.

  87. 87.

    Mahendran R, Spottswood MS, Ghate A, Ling ML, Jeng K, Miller DL. Editing of the mitochondrial small subunit rRNA in physarum polycephalum. EMBO J. 1994;13:232–40.

  88. 88.

    Anant S, Davidson NO. An AU-rich sequence element (UUUN[A/U]U) downstream of the edited C in apolipoprotein B mRNA is a high-affinity binding site for apobec-1: binding of apobec-1 to this motif in the 3′ untranslated region of c-myc increases mRNA stability. Mol Cell Biol. 2000;20:1982–92.

  89. 89.

    Yu Q, Chen D, Konig R, Mariani R, Unutmaz D, Landau NR. APOBEC3B and APOBEC3C are potent inhibitors of simian immunodeficiency virus replication. J Biol Chem. 2004;279:53379–86.

  90. 90.

    Kinomoto M, Kanno T, Shimura M, Ishizaka Y, Kojima A, Kurata T, et al. All APOBEC3 family proteins differentially inhibit LINE-1 retrotransposition. Nucleic Acids Res. 2007;35:2955–64.

  91. 91.

    Sato Y, Probst HC, Tatsumi R, Ikeuchi Y, Neuberger MS, Rada C. Deficiency in APOBEC2 leads to a shift in muscle fiber type, diminished body mass, and myopathy. J Biol Chem. 2010;285:7111–8.

  92. 92.

    Okuyama S, Marusawa H, Matsumoto T, Ueda Y, Matsumoto Y, Endo Y, et al. Excessive activity of apolipoprotein B mRNA editing enzyme catalytic polypeptide 2 (APOBEC2) contributes to liver and lung tumorigenesis. Int J Cancer. 2012;130:1294–301.

  93. 93.

    Leong WM, Ripen AM, Mirsafian H, Mohamad SB, Merican AF. Transcriptogenomics identification and characterization of RNA editing sites in human primary monocytes using high-depth next generation sequencing data. Genomics. 2018;111:899–905.

  94. 94.

    Bahn JH, Lee JH, Li G, Greer C, Peng G, Xiao X. Accurate identification of A-to-I RNA editing in human by transcriptome sequencing. Genome Res. 2012;22:142–50.

  95. 95.

    Lachmann A, Torre D, Keenan AB, Jagodnik KM, Lee HJ, Wang L, et al. Massive mining of publicly available RNA-seq data from human and mouse. Nat Commun. 2018;9:1366.

  96. 96.

    John D, Weirick T, Dimmeler S, Uchida S. RNAEditor: easy detection of RNA editing events and the introduction of editing islands. Brief Bioinform. 2017;18:993–1001.

  97. 97.

    Picardi E, Manzari C, Mastropasqua F, Aiello I, D’Erchia AM, Pesole G. Profiling RNA editing in human tissues: towards the inosinome atlas. Sci Rep. 2015;5:14941.

  98. 98.

    Ramaswami G, Li JB. RADAR: a rigorously annotated database of A-to-I RNA editing. Nucleic Acids Res. 2014;42:D109–13.

  99. 99.

    He T, Du P, Li Y. dbRES: a web-oriented database for annotated RNA editing sites. Nucleic Acids Res. 2007;35:D141–4.

  100. 100.

    Picardi E, D’Erchia AM, Lo Giudice C, Pesole G. REDIportal: a comprehensive database of A-to-I RNA editing events in humans. Nucleic Acids Res. 2017;45:D750–7.

  101. 101.

    Picardi E, Regina TM, Brennicke A, Quagliariello C. REDIdb: the RNA editing database. Nucleic Acids Res. 2007;35:D173–7.

  102. 102.

    Gong J, Liu C, Liu W, Xiang Y, Diao L, Guo AY, et al. LNCediting: a database for functional effects of RNA editing in lncRNAs. Nucleic Acids Res. 2017;45:D79–84.

  103. 103.

    Lai F, Chen CX, Carter KC, Nishikura K. Editing of glutamate receptor B subunit ion channel RNAs by four alternatively spliced DRADA2 double-stranded RNA adenosine deaminases. Mol Cell Biol. 1997;17:2413–24.

  104. 104.

    Liu Y, Samuel CE. Editing of glutamate receptor subunit B pre-mRNA by splice-site variants of interferon-inducible double-stranded RNA-specific adenosine deaminase ADAR1. J Biol Chem. 1999;274:5070–7.

  105. 105.

    Rueter SM, Dawson TR, Emeson RB. Regulation of alternative splicing by RNA editing. Nature. 1999;399:75–80.

  106. 106.

    Hsiao YE, Bahn JH, Yang Y, Lin X, Tran S, Yang EW, et al. RNA editing in nascent RNA affects pre-mRNA splicing. Genome Res. 2018;28:812–23.

  107. 107.

    Raitskin O, Cho DS, Sperling J, Nishikura K, Sperling R. RNA editing activity is associated with splicing factors in lnRNP particles: the nuclear pre-mRNA processing machinery. Proc Natl Acad Sci USA. 2001;98:6571–6.

  108. 108.

    Lev-Maor G, Sorek R, Levanon EY, Paz N, Eisenberg E, Ast G. RNA-editing-mediated exon evolution. Genome Biol. 2007;8:R29.

  109. 109.

    Terajima H, Yoshitane H, Ozaki H, Suzuki Y, Shimba S, Kuroda S, et al. ADARB1 catalyzes circadian A-to-I editing and regulates RNA rhythm. Nat Genet. 2017;49:146–51.

  110. 110.

    Sinigaglia K, Wiatrek D, Khan A, Michalik D, Sambrani N, Sedmik J, et al. ADAR RNA editing in innate immune response phasing, in circadian clocks and in sleep. Biochim Biophys Acta Gene Regul Mech. 2018;1862:356–69.

  111. 111.

    Robinson JE, Paluch J, Dickman DK, Joiner WJ. ADAR-mediated RNA editing suppresses sleep by acting as a brake on glutamatergic synaptic plasticity. Nat Commun. 2016;7:10512.

  112. 112.

    Sommer B, Kohler M, Sprengel R, Seeburg PH. RNA editing in brain controls a determinant of ion flow in glutamate-gated channels. Cell. 1991;67:11–9.

  113. 113.

    Jeck WR, Sorrentino JA, Wang K, Slevin MK, Burd CE, Liu J, et al. Circular RNAs are abundant, conserved, and associated with ALU repeats. RNA. 2013;19:141–57.

  114. 114.

    Ekdahl Y, Farahani HS, Behm M, Lagergren J, Ohman M. A-to-I editing of microRNAs in the mammalian brain increases during development. Genome Res. 2012;22:1477–87.

  115. 115.

    Riemondy KA, Gillen AE, White EA, Bogren LK, Hesselberth JR, Martin SL. Dynamic temperature-sensitive A-to-I RNA editing in the brain of a heterothermic mammal during hibernation. RNA. 2018;24:1481–95.

  116. 116.

    Muramatsu M, Kinoshita K, Fagarasan S, Yamada S, Shinkai Y, Honjo T. Class switch recombination and hypermutation require activation-induced cytidine deaminase (AID), a potential RNA editing enzyme. Cell. 2000;102:553–63.

  117. 117.

    Honjo T, Muramatsu M, Fagarasan S. AID: how does it aid antibody diversity? Immunity. 2004;20:659–68.

  118. 118.

    Shapiro GS, Aviszus K, Murphy J, Wysocki LJ. Evolution of ig DNA sequence to target specific base positions within codons for somatic hypermutation. J Immunol. 2002;168:2302–6.

  119. 119.

    Rogozin IB, Diaz M. Cutting edge: DGYW/WRCH is a better predictor of mutability at G: C bases in ig hypermutation than the widely accepted RGYW/WRCY motif and probably reflects a two-step activation-induced cytidine deaminase-triggered process. J Immunol. 2004;172:3382–4.

  120. 120.

    Chiarle R, Zhang Y, Frock RL, Lewis SM, Molinie B, Ho YJ, et al. Genome-wide translocation sequencing reveals mechanisms of chromosome breaks and rearrangements in B cells. Cell. 2011;147:107–19.

  121. 121.

    Madani N, Kabat D. An endogenous inhibitor of human immunodeficiency virus in human lymphocytes is overcome by the viral vif protein. J Virol. 1998;72:10251–5.

  122. 122.

    Sheehy AM, Gaddis NC, Choi JD, Malim MH. Isolation of a human gene that inhibits HIV-1 infection and is suppressed by the viral vif protein. Nature. 2002;418:646–50.

  123. 123.

    Yu X, Yu Y, Liu B, Luo K, Kong W, Mao P, et al. Induction of APOBEC3G ubiquitination and degradation by an HIV-1 vif-Cul5-SCF complex. Science. 2003;302:1056–60.

  124. 124.

    Hultquist JF, Lengyel JA, Refsland EW, LaRue RS, Lackey L, Brown WL, et al. Human and rhesus APOBEC3D, APOBEC3F, APOBEC3G, and APOBEC3H demonstrate a conserved capacity to restrict vif-deficient HIV-1. J Virol. 2011;85:11220–34.

  125. 125.

    Chen H, Lilley CE, Yu Q, Lee DV, Chou J, Narvaiza I, et al. APOBEC3A is a potent inhibitor of adeno-associated virus and retrotransposons. Curr Biol. 2006;16:480–5.

  126. 126.

    Delebecque F, Suspene R, Calattini S, Casartelli N, Saib A, Froment A, et al. Restriction of foamy viruses by APOBEC cytidine deaminases. J Virol. 2006;80:605–14.

  127. 127.

    Zielonka J, Bravo IG, Marino D, Conrad E, Perkovic M, Battenberg M, et al. Restriction of equine infectious anemia virus by equine APOBEC3 cytidine deaminases. J Virol. 2009;83:7547–59.

  128. 128.

    Harris RS, Bishop KN, Sheehy AM, Craig HM, Petersen-Mahrt SK, Watt IN, et al. DNA deamination mediates innate immunity to retroviral infection. Cell. 2003;113:803–9.

  129. 129.

    Turelli P, Mangeat B, Jost S, Vianin S, Trono D. Inhibition of hepatitis B virus replication by APOBEC3G. Science. 2004;303:1829.

  130. 130.

    Nguyen DH, Gummuluru S, Hu J. Deamination-independent inhibition of hepatitis B virus reverse transcription by APOBEC3G. J Virol. 2007;81:4465–72.

  131. 131.

    Bishop KN, Holmes RK, Sheehy AM, Malim MH. APOBEC-mediated editing of viral RNA. Science. 2004;305:645.

  132. 132.

    Fehrholz M, Kendl S, Prifert C, Weissbrich B, Lemon K, Rennick L, et al. The innate antiviral factor APOBEC3G targets replication of measles, mumps and respiratory syncytial viruses. J Gen Virol. 2012;93:565–76.

  133. 133.

    Ward SV, George CX, Welch MJ, Liou LY, Hahm B, Lewicki H, et al. RNA editing enzyme adenosine deaminase is a restriction factor for controlling measles virus replication that also is required for embryogenesis. Proc Natl Acad Sci USA. 2011;108:331–6.

  134. 134.

    Taylor DR, Puig M, Darnell ME, Mihalik K, Feinstone SM. New antiviral pathway that mediates hepatitis C virus replicon interferon sensitivity through ADAR1. J Virol. 2005;79:6291–8.

  135. 135.

    Suspene R, Renard M, Henry M, Guetard D, Puyraimond-Zemmour D, Billecocq A, et al. Inversing the natural hydrogen bonding rule to selectively amplify GC-rich ADAR-edited RNAs. Nucleic Acids Res. 2008;36:e72.

  136. 136.

    Samuel CE. ADARs: viruses and innate immunity. Curr Top Microbiol Immunol. 2012;353:163–95.

  137. 137.

    Zahn RC, Schelp I, Utermohlen O, von Laer D. A-to-G hypermutation in the genome of lymphocytic choriomeningitis virus. J Virol. 2007;81:457–64.

  138. 138.

    Samuel CE. Adenosine deaminases acting on RNA (ADARs) are both antiviral and proviral. Virology. 2011;411:180–93.

  139. 139.

    Chiu YL, Greene WC. The APOBEC3 cytidine deaminases: an innate defensive network opposing exogenous retroviruses and endogenous retroelements. Annu Rev Immunol. 2008;26:317–53.

  140. 140.

    Esnault C, Heidmann O, Delebecque F, Dewannieux M, Ribet D, Hance AJ, et al. APOBEC3G cytidine deaminase inhibits retrotransposition of endogenous retroviruses. Nature. 2005;433:430–3.

  141. 141.

    Richardson SR, Doucet AJ, Kopera HC, Moldovan JB, Garcia-Perez JL, Moran JV. The influence of LINE-1 and SINE retrotransposons on mammalian genomes. Microbiol Spectr. 2015;3:MDNA3-0061-2014.

  142. 142.

    Pizarro JG, Cristofari G. Post-transcriptional control of LINE-1 retrotransposition by cellular host factors in somatic cells. Front Cell Dev Biol. 2016;4:14.

  143. 143.

    Orecchini E, Frassinelli L, Michienzi A. Restricting retrotransposons: ADAR1 is another guardian of the human genome. RNA Biol. 2017;14:1485–91.

  144. 144.

    Chen LL, DeCerbo JN, Carmichael GG. Alu element-mediated gene silencing. EMBO J. 2008;27:1694–705.

  145. 145.

    Krestel H, Meier JC. RNA editing and retrotransposons in neurology. Front Mol Neurosci. 2018;11:163.

  146. 146.

    Maruyama H, Morino H, Ito H, Izumi Y, Kato H, Watanabe Y, et al. Mutations of optineurin in amyotrophic lateral sclerosis. Nature. 2010;465:223–6.

  147. 147.

    Kwak S, Kawahara Y. Deficient RNA editing of GluR2 and neuronal death in amyotropic lateral sclerosis. J Mol Med (Berl). 2005;83:110–20.

  148. 148.

    Sasaki S, Yamashita T, Kwak S. Autophagy in spinal motor neurons of conditional ADAR2-knockout mice: an implication for a role of calcium in increased autophagy flux in ALS. Neurosci Lett. 2015;598:79–84.

  149. 149.

    Akbarian S, Smith MA, Jones EG. Editing for an AMPA receptor subunit RNA in prefrontal cortex and striatum in alzheimer’s disease, huntington’s disease and schizophrenia. Brain Res. 1995;699:297–304.

  150. 150.

    Slotkin W, Nishikura K. Adenosine-to-inosine RNA editing and human disease. Genome Med. 2013;5:105.

  151. 151.

    Orlowski RJ, O’Rourke KS, Olorenshaw I, Hawkins GA, Maas S, Laxminarayana D. Altered editing in cyclic nucleotide phosphodiesterase 8A1 gene transcripts of systemic lupus erythematosus T lymphocytes. Immunology. 2008;125:408–19.

  152. 152.

    Rice GI, Kasher PR, Forte GM, Mannion NM, Greenwood SM, Szynkiewicz M, et al. Mutations in ADAR1 cause aicardi-goutieres syndrome associated with a type I interferon signature. Nat Genet. 2012;44:1243–8.

  153. 153.

    Paz-Yaacov N, Bazak L, Buchumenski I, Porath HT, Danan-Gotthold M, Knisbacher BA, et al. Elevated RNA editing activity is a major contributor to transcriptomic diversity in tumors. Cell Rep. 2015;13:267–76.

  154. 154.

    Han L, Diao L, Yu S, Xu X, Li J, Zhang R, et al. The genomic landscape and clinical relevance of A-to-I RNA editing in human cancers. Cancer Cell. 2015;28:515–28.

  155. 155.

    Paz N, Levanon EY, Amariglio N, Heimberger AB, Ram Z, Constantini S, et al. Altered adenosine-to-inosine RNA editing in human cancer. Genome Res. 2007;17:1586–95.

  156. 156.

    Kawahara Y, Zinshteyn B, Sethupathy P, Iizasa H, Hatzigeorgiou AG, Nishikura K. Redirection of silencing targets by adenosine-to-inosine editing of miRNAs. Science. 2007;315:1137–40.

  157. 157.

    Tomaselli S, Galeano F, Alon S, Raho S, Galardi S, Polito VA, et al. Modulation of microRNA editing, expression and processing by ADAR2 deaminase in glioblastoma. Genome Biol. 2015;16:5.

  158. 158.

    Choudhury Y, Tay FC, Lam DH, Sandanaraj E, Tang C, Ang BT, et al. Attenuated adenosine-to-inosine editing of microRNA-376a* promotes invasiveness of glioblastoma cells. J Clin Invest. 2012;122:4059–76.

  159. 159.

    Shoshan E, Mobley AK, Braeuer RR, Kamiya T, Huang L, Vasquez ME, et al. Reduced adenosine-to-inosine miR-455-5p editing promotes melanoma growth and metastasis. Nat Cell Biol. 2015;17:311–21.

  160. 160.

    Anadon C, Guil S, Simo-Riudalbas L, Moutinho C, Setien F, Martinez-Cardus A, et al. Gene amplification-associated overexpression of the RNA editing enzyme ADAR1 enhances human lung tumorigenesis. Oncogene. 2016;35:4407–13.

  161. 161.

    Wang Y, Xu X, Yu S, Jeong KJ, Zhou Z, Han L, et al. Systematic characterization of A-to-I RNA editing hotspots in microRNAs across human cancers. Genome Res. 2017;27:1112–25.

  162. 162.

    Zhang L, Yang CS, Varelas X, Monti S. Altered RNA editing in 3′ UTR perturbs microRNA-mediated regulation of oncogenes and tumor-suppressors. Sci Rep. 2016;6:23226.

  163. 163.

    Shelton PM, Duran A, Nakanishi Y, Reina-Campos M, Kasashima H, Llado V, et al. The secretion of miR-200s by a PKCzeta/ADAR2 signaling axis promotes liver metastasis in colorectal cancer. Cell Rep. 2018;23:1178–91.

  164. 164.

    Gumireddy K, Li A, Kossenkov AV, Sakurai M, Yan J, Li Y, et al. The mRNA-edited form of GABRA3 suppresses GABRA3-mediated akt activation and breast cancer metastasis. Nat Commun. 2016;7:10715.

  165. 165.

    Peng X, Xu X, Wang Y, Hawke DH, Yu S, Han L, et al. A-to-I RNA editing contributes to proteomic diversity in cancer. Cancer Cell. 2018;33(817–828):e7.

  166. 166.

    Chan TH, Qamra A, Tan KT, Guo J, Yang H, Qi L, et al. ADAR-mediated RNA editing predicts progression and prognosis of gastric cancer. Gastroenterology. 2016;151(637–650):e10.

  167. 167.

    Galeano F, Rossetti C, Tomaselli S, Cifaldi L, Lezzerini M, Pezzullo M, et al. ADAR2-editing activity inhibits glioblastoma growth through the modulation of the CDC14B/Skp2/p21/p27 axis. Oncogene. 2013;32:998–1009.

  168. 168.

    Han SW, Kim HP, Shin JY, Jeong EG, Lee WC, Kim KY, et al. RNA editing in RHOQ promotes invasion potential in colorectal cancer. J Exp Med. 2014;211:613–21.

  169. 169.

    Chen YB, Liao XY, Zhang JB, Wang F, Qin HD, Zhang L, et al. ADAR2 functions as a tumor suppressor via editing IGFBP7 in esophageal squamous cell carcinoma. Int J Oncol. 2017;50:622–30.

  170. 170.

    Fu L, Qin YR, Ming XY, Zuo XB, Diao YW, Zhang LY, et al. RNA editing of SLC22A3 drives early tumor invasion and metastasis in familial esophageal cancer. Proc Natl Acad Sci USA. 2017;114:E4631–40.

  171. 171.

    Chen L, Li Y, Lin CH, Chan TH, Chow RK, Song Y, et al. Recoding RNA editing of AZIN1 predisposes to hepatocellular carcinoma. Nat Med. 2013;19:209–16.

  172. 172.

    Qin YR, Qiao JJ, Chan TH, Zhu YH, Li FF, Liu H, et al. Adenosine-to-inosine RNA editing mediated by ADARs in esophageal squamous cell carcinoma. Cancer Res. 2014;74:840–51.

  173. 173.

    Shigeyasu K, Okugawa Y, Toden S, Miyoshi J, Toiyama Y, Nagasaka T, et al. AZIN1 RNA editing confers cancer stemness and enhances oncogenic potential in colorectal cancer. JCI Insight. 2018.

  174. 174.

    Chen W, He W, Cai H, Hu B, Zheng C, Ke X, et al. A-to-I RNA editing of BLCAP lost the inhibition to STAT3 activation in cervical cancer. Oncotarget. 2017;8:39417–29.

  175. 175.

    Takizawa M, Tolarova H, Li Z, Dubois W, Lim S, Callen E, et al. AID expression levels determine the extent of cMyc oncogenic translocations and the incidence of B cell tumor development. J Exp Med. 2008;205:1949–57.

  176. 176.

    Okazaki IM, Hiai H, Kakazu N, Yamada S, Muramatsu M, Kinoshita K, et al. Constitutive expression of AID leads to tumorigenesis. J Exp Med. 2003;197:1173–81.

  177. 177.

    Komeno Y, Kitaura J, Watanabe-Okochi N, Kato N, Oki T, Nakahara F, et al. AID-induced T-lymphoma or B-leukemia/lymphoma in a mouse BMT model. Leukemia. 2010;24:1018–24.

  178. 178.

    Matsumoto Y, Marusawa H, Kinoshita K, Niwa Y, Sakai Y, Chiba T. Up-regulation of activation-induced cytidine deaminase causes genetic aberrations at the CDKN2b–CDKN2a in gastric cancer. Gastroenterology. 2010;139:1984–94.

  179. 179.

    Saraconi G, Severi F, Sala C, Mattiuz G, Conticello SG. The RNA editing enzyme APOBEC1 induces somatic mutations and a compatible mutational signature is present in esophageal adenocarcinomas. Genome Biol. 2014;15:417.

  180. 180.

    Sieuwerts AM, Willis S, Burns MB, Look MP, Meijer-Van Gelder ME, Schlicker A, et al. Elevated APOBEC3B correlates with poor outcomes for estrogen-receptor-positive breast cancers. Horm Cancer. 2014;5:405–13.

  181. 181.

    Burns MB, Lackey L, Carpenter MA, Rathore A, Land AM, Leonard B, et al. APOBEC3B is an enzymatic source of mutation in breast cancer. Nature. 2013;494:366–70.

  182. 182.

    Ding Q, Chang CJ, Xie X, Xia W, Yang JY, Wang SC, et al. APOBEC3G promotes liver metastasis in an orthotopic mouse model of colorectal cancer and predicts human hepatic metastasis. J Clin Invest. 2011;121:4526–36.

  183. 183.

    Roberts SA, Lawrence MS, Klimczak LJ, Grimm SA, Fargo D, Stojanov P, et al. An APOBEC cytidine deaminase mutagenesis pattern is widespread in human cancers. Nat Genet. 2013;45:970–6.

  184. 184.

    Glaser AP, Fantini D, Wang Y, Yu Y, Rimar KJ, Podojil JR, et al. APOBEC-mediated mutagenesis in urothelial carcinoma is associated with improved survival, mutations in DNA damage response genes, and immune response. Oncotarget. 2017;9:4537–48.

  185. 185.

    Nik-Zainal S, Alexandrov LB, Wedge DC, Van Loo P, Greenman CD, Raine K, et al. Mutational processes molding the genomes of 21 breast cancers. Cell. 2012;149:979–93.

  186. 186.

    Alexandrov LB, Nik-Zainal S, Wedge DC, Aparicio SA, Behjati S, Biankin AV, et al. Signatures of mutational processes in human cancer. Nature. 2013;500:415–21.

  187. 187.

    Law EK, Sieuwerts AM, LaPara K, Leonard B, Starrett GJ, Molan AM, et al. The DNA cytosine deaminase APOBEC3B promotes tamoxifen resistance in ER-positive breast cancer. Sci Adv. 2016;2:e1601737.

  188. 188.

    Yamanaka S, Balestra ME, Ferrell LD, Fan J, Arnold KS, Taylor S, et al. Apolipoprotein B mRNA-editing protein induces hepatocellular carcinoma and dysplasia in transgenic animals. Proc Natl Acad Sci USA. 1995;92:8483–7.

  189. 189.

    Valdmanis PN, Roy-Chaudhuri B, Kim HK, Sayles LC, Zheng Y, Chuang CH, et al. Upregulation of the microRNA cluster at the Dlk1-Dio3 locus in lung adenocarcinoma. Oncogene. 2015;34:94–103.

  190. 190.

    Boichard A, Tsigelny IF, Kurzrock R. High expression of PD-1 ligands is associated with kataegis mutational signature and APOBEC3 alterations. Oncoimmunology. 2017;6:e1284719.

  191. 191.

    Zhang M, Fritsche J, Roszik J, Williams LJ, Peng X, Chiu Y, et al. RNA editing derived epitopes function as cancer antigens to elicit immune responses. Nat Commun. 2018;9:3919.

  192. 192.

    Ben-Aroya S, Levanon EY. A-to-I RNA editing: an overlooked source of cancer mutations. Cancer Cell. 2018;33:789–90.

  193. 193.

    Fritzell K, Xu LD, Otrocka M, Andreasson C, Ohman M. Sensitive ADAR editing reporter in cancer cells enables high-throughput screening of small molecule libraries. Nucleic Acids Res. 2018;47:e22.

  194. 194.

    Olson ME, Harris RS, Harki DA. APOBEC enzymes as targets for virus and cancer therapy. Cell Chem Biol. 2018;25:36–49.

  195. 195.

    Mizrahi RA, Schirle NT, Beal PA. Potent and selective inhibition of A-to-I RNA editing with 2′-O-methyl/locked nucleic acid-containing antisense oligoribonucleotides. ACS Chem Biol. 2013;8:832–9.

  196. 196.

    Montiel-Gonzalez MF, Vallecillo-Viejo I, Yudowski GA, Rosenthal JJ. Correction of mutations within the cystic fibrosis transmembrane conductance regulator by site-directed RNA editing. Proc Natl Acad Sci U S A. 2013;110:18285–90.

  197. 197.

    Hanswillemenke A, Kuzdere T, Vogel P, Jekely G, Stafforst T. Site-directed RNA editing in vivo can be triggered by the light-driven assembly of an artificial riboprotein. J Am Chem Soc. 2015;137:15875–81.

  198. 198.

    UniProt Consortium. UniProt: a worldwide hub of protein knowledge. Nucleic Acids Res. 2019;47:D506–15.

  199. 199.

    Wang IX, So E, Devlin JL, Zhao Y, Wu M, Cheung VG. ADAR regulates RNA editing, transcript stability, and gene expression. Cell Rep. 2013;5:849–60.

  200. 200.

    Waterhouse A, Bertoni M, Bienert S, Studer G, Tauriello G, Gumienny R, et al. SWISS-MODEL: homology modelling of protein structures and complexes. Nucleic Acids Res. 2018;46:W296–303.

Download references


Not applicable.



Author information

TC wrote the manuscript. AZ organized and supervised the study, critically read the manuscript. All authors read and approved the final manuscript.

Correspondence to Apostolos Zaravinos.

Ethics declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark


  • RNA editing
  • ADAR
  • AID
  • Cytidine/adenosine deaminases
  • Epitranscriptomics
  • Cancer
  • Post-transcriptional modifications