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RNA Modification

Yuru Wang, … Peter A. Beal, in The Enzymes, 2017

3.3 ADARs in Different Organisms

The ADAR family of proteins is found in most of metazoans[138–140]. Most species have more than one type of ADARprotein. If classified based on domain makeup, ADARs can be placed into three categories: ADAR1-like, ADAR2-like, and ADAD-like (ADAR-related proteins) [141]. There is at most one ADAR1-like ADAR present in most species. However, there are up to three versions of ADAR2-like proteins and two versions of ADAD-like proteins found in one species (Mnemiopsis leidyi)[141]. ADARs are not found in several nonmetazoaneukaryotes, such as yeast, fungi, plants, and choanoflagellates.

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RNA Editing

Liam P. Keegan, … Mary A. O'Connell, in Methods in Enzymology, 2007


ADAR editing enzymes are found in all multicellular animals and are conserved in sequence and protein organization. The number of ADAR genes differs between animals, ranging from three in mammals to one in Drosophila. ADAR is also alternatively spliced to generate isoforms that can differ significantly in enzymatic activity. Therefore, to study the enzyme in vitro, it is essential to have an easy and reliable method of expressing and purifying recombinant ADAR protein. To add to the complexity of RNA editing, the number of transcripts that are edited by ADARs differs in different organisms. In humans there is extensive editing of Alu sequences, whereas in invertebrates transcripts expressed in the central nervous system are edited and this editing increases during development. It is possible to quantify site‐specific RNA editing by sequencing of clones derived from RT‐PCRproducts. However, for routine assaying of an edited position within a particular transcript, this is both expensive and time consuming. Therefore, a nonradioactive method based on poison primer extension assay is an ideal alternative.

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miRNA and Cancer

Anjan K. Pradhan, … Paul B. Fisher, in Advances in Cancer Research, 2017

6.7 ADARs

ADAR or adenosine deaminase acting on RNA catalyzes adenine-to-inosine conversion in dsRNA. The inosine can then be converted to guanosine, which can pair to cytosine. Several primary and pre-miRNAs are substrates of ADARs. miR-142 is edited at the primary level, which results in its suppressed processing by Drosha (Yang et al., 2006). This unedited pri-miRNA is rapidly degraded by the ribonuclease Tudor-SN. miR-151 is another miR that is edited by ADAR and prevents processing by DICER, resulting in the accumulation of pre-miRNAs. This kind of modification can alter the expression level of mature miRNAs, which can change the level of a number of downstream mRNAs.

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RNA Editing by Mammalian ADARs

Marion Hogg, … Mary A. O'Connell, in Advances in Genetics, 2011

V RNA Binding by ADARs

ADAR belongs to a diverse group of proteins that have one or more copies of a dsRBD of approximately 70 amino acids (reviewed in Fierro-Monti and Mathews, 2000). Despite only a few residues being conserved, all dsRBDs analyzed fold into the same secondary structure of α-helices and β-sheets, organized as α1-β1-β2-β3-α2, where the α-helices make contact with the same face of the RNA. Other proteins with dsRBDs members include Staufen a protein involved in mRNA transport; PKR, an interferon-inducible, RNA-dependent protein kinase; and many proteins involved in the RNA interference pathway which have domain arrangements resembling ribonuclease III, an RNAnuclease.

The dsRBD-containing proteins do not exhibit sequence specificity, it is the secondary structure of the A-form RNA that they recognize. A-form nucleic acid differs from B-form in that the major groove is narrow and the minor groove is wide and shallow. Highly sequence-specific interaction between proteins and DNA occurs on the major grove; however, on A-form nucleic acid this is not possible so the protein–RNA interactions occur with the 2-hydroxyl group of the ribosesugar in the minor and are therefore not sequence specific.

However, the dsRBDs of ADAR2 do display a binding selectivity to the transcripts they edit. Experiments on a short RNA encompassing the GluR-B Q/R site demonstrated that ADAR2 dsRBDs exhibit selective RNA binding to the GluR-B Q/R site and this was distinct from the binding site of a dsRBD from PKR (Stephens et al., 2004). The dsRBD occupies approximately 16 bp of dsRNA but many can accommodate or prefer binding to substrates that contain bulges or loops within the dsRNA. The individual dsRBDs of ADAR2 exhibit different binding specificities when analyzed separately (Poulsen et al., 2006; Stefl et al., 2006). dsRBD1 preferentially binds to perfect duplex dsRNA located in the stem–loop region, whereas dsRBD2 shows preference for dsRNA containing an A–C mismatch which is usually located near to the editing site. Both dsRBDs are required for efficient editing by ADAR2 (Stefl et al., 2006).

In mammals, ADAR1 contains three dsRBDs, whereas ADAR2 and ADAR3 both contain two dsRBDs. Recently, an ADAR was cloned from squid that is an ortholog of ADAR2 (Palavicini et al., 2009). It encodes three dsRBD; however, the first dsRBD is in an alternatively spliced exon at the amino terminus of the protein. When both isoforms were expressed and assayed in vitro the isoform with three dsRBDs had increased enzymatic activity.

To investigate what was responsible for the specificity of ADAR1 and ADAR2, chimeric proteins were generated with the dsRBDs from one ADAR and the deaminase domain from another (Wong et al., 2001). This revealed that the editing specificity in the chimeric proteins was provided by the deaminase domain. However, when the dsRBDs of ADAR1 and PKR were exchanged, this resulted in a reduction or a complete loss of enzymatic activity on edited transcript; however, the chimeric protein was still active on dsRNA. Therefore, specificity is also provided by the dsRBDs (Liu et al., 2000). ADAR3 is the only member of the ADAR family known to bind to ssRNA, and experiments have shown this interaction is mediated through the arginine- and lysine-rich R domain in the amino terminus of the protein which is not found in other ADAR proteins (Chen et al., 2000; Fig. 3.1). ADAR3 is also capable of binding dsRNA through its canonical dsRBDs but it is unable to edit either known substrates or dsRNA.

The dsRBDs have additional roles other than binding dsRNA such as nucleocytoplasmic shuttling of ADAR1 as previously described. The deletion of dsRBD1 of ADAR2 reduced editing efficiency, whereas loss of dsRBD2 abolished editing altogether (Poulsen et al., 2006). The N-terminal region of ADAR2 had an autoinhibitory effect on catalytic activity as a truncated ADAR2 protein containing a deaminase domain and dsRB2 was capable of editing a 15-bp substrate, whereas the full-length protein did not. However, the addition of the N-terminal region of ADAR2 to the truncated protein in trans reduced editing efficiency on shorter substrates, suggesting that the RNA substrate has to be long enough for binding of both dsRBDs to relieve the autoinhibition for efficient editing (Macbeth et al., 2004).

The dsRBDs have also been implicated in dimer formation as binding to RNA has been shown to be a prerequisite for dimer formation. However, there have been conflicting reports and the issue of whether dimer formation requires RNA binding for its formation has not been resolved. Studies have shown that the formation of homodimers is required for ADAR activity (Cho et al., 2003; Gallo et al., 2003; Jaikaran et al., 2002; Poulsen et al., 2006). A ternary complex can be observed when increasing amounts of ADAR are added to substrate RNA, such that one monomer binds and then another, indicating dimerization is RNA-dependent (Jaikaran et al., 2002). Analysis of RNA editing in vitro using one wild-type monomer and one catalytically inactive monomer showed that both monomers contribute to hyperediting of dsRNA and site-specific editing of substrates (Cho et al., 2003). However, fluorescence energy resonance transfer (FRET) experiments indicate that ADAR1 and ADAR2 form homodimers in an RNA-independent manner, and are capable of forming heterodimers in vivo (Chilibeck et al., 2006).

To assess the role of RNA binding in dimerization (Valente and Nishikura, 2007), mutations were made in three conserved lysine residues (KKxxK → EAxxA) within each of the dsRBDs of ADAR1 and ADAR2. The mutations introduced were based on data from alanine-scanning mutagenesis of the DrosophilaRNA-binding protein Staufen, which demonstrated that mutation of exposed lysine residues within the conserved dsRBD eliminated RNA binding without disrupting structure (Ramos et al., 2000). Sequential purification of protein complexes containing both wild-type and mutant ADAR proteins demonstrated that homodimerization of ADAR proteins occurs independently of RNA binding (Valente and Nishikura, 2007). However, the dimeric ADAR proteins containing one mutant and one wild-type monomer behaved in a dominant negative manner in both an RNA-binding assay and an in vitro editing assay indicating that RNA binding of both monomers is required for deamination to occur.

The conflicting reports likely arise from the differentexperimental models, whether the experiments were performedin vitro or in vivo in cell culture. Also, posttranslational modification could be involved as some proteins were expressed in yeast and others in Sf9 insect cells. Furthermore, proteins from different species such as Drosophila and humanADARs were used.

In yeast, ADAT2 and ADAT3 have been shown to function as a heterodimer where ADAT2 provides the catalytic function and ADAT3 provides substrate specificity (Gerber and Keller, 1999). This raised the possibility that ADARs may function as heterodimers. Heterodimerization between ADAR proteins may reveal a role for ADAR3, where ADAR1 or ADAR2 provides the catalytic activity and ADAR3 affects the substrate specificity. FRET analysis indicates that heterodimers between ADAR1 and ADAR2 monomers form in vivo (Chilibeck et al., 2006) and heterodimers between ADAR1 and ADAR2 were co-immunoprecipitated from astrocytoma cell lines, where it was demonstrated that increased levels of ADAR1 can inhibit editing of substrates by ADAR2 (Cenci et al., 2008). It remains to be seen whether overexpression of ADAR3 can have the same effect. Dimers formed between the two isoforms of ADAR1 (p110 and p150) are readily detectable indicating that the Z-DNA binding domains absent in the p110 isoform are not required for dimerization (Cho et al., 2003).

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RNA Editing in Animals

J. Scott, in Encyclopedia of Genetics, 2001

Mechanism of A to I RNA Editing

ADARs edit A to I nonselectively in extended perfect double-stranded RNA duplexes. This activity is highly conserved and ancient in origin. The biological role for this promiscuous editing is not known. It has variously been suggested to be involved in gene regulationviral life cycle or defence against viruses. The A to I editing enzymes also have a site specific role in covering A to I in premessenger RNA. The targets for A to I editing are mainly found in the nervous system of vertebratesand invertebrates. They include transcripts for ligand or voltage gated ion channels and G protein-coupled receptors. A to I editing exists in a variety of other tissues, such as the heart, where I has been detected in mRNA.

The prototype of site selective A to I premessenger RNA editingis in the glutamate receptor sub-unit genes. At one site in the gluRB mRNA (the glutamine (Q)/arginine (R) site) undergoes editing from a genomically encoded Q codon (CAG) to an R codon (CGG). ADAR2 is the editing enzyme for this Q/R site. The functional consequence of this editing is a marked change in calcium permeability of the gluR channels. Site specific A to I editing is dependent on the formation of double stranded RNA between the editing site and a complementary downstream intron (Figure 4). The most plausible mechanism of action for this enzyme is suggested by the structural relationship of the ADARs to Hhal DNA methyl transferase. Thus ADAR would flip out the A to be modified to bring it into the active site of the enzyme. Other examples of A to I editing in mammals include other gluR editing sites, a variety of sites in serotonin receptor messenger RNA and in the ADAR2 gene premessenger RNA itself (Figure 4). The Q/R site is the most highly edited of all the A to I editing sites. More than 99% of transcripts are edited. Other examples of A to I editing have considerably lower frequencies.

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Nucleic Acid Sensing and Immunity, Part A

Takumi Kawasaki, Taro Kawai, in International Review of Cell and Molecular Biology, 2019

5.2 ADAR

ADARs (adenosine deaminases acting on RNA) catalyze the conversion of adenosine (A) to inosine (I) in double-stranded RNA (dsRNA) substrates. Such “A-to-I editing” alters the coding of RNA, as inosine is read as guanosine (G) instead of adenosine by ribosomes during translation, leading to the production of non-functional proteins. In humans, two autoimmune disorders, Dyschromatosis Symmetrica Hereditaria (DSH) and Aicardi-Goutières Syndrome (AGS), are attributed to specific mutations that have been mapped to theAdar1 gene (Rice et al., 2012). Furthermore, Adar1 inducible knockout (KO) mice exhibit a high level of IFN production in neuronal tissues, a key characteristic of AGS. In healthy cells, ADAR1-mediated nucleotide conversions are believed to be important for the suppression of viral replication and, more importantly, endogenous RNA recognition by the RNA sensor,MDA5. Correspondingly, mutations in either MDA5 or MAVS rescue the embryonic lethal phenotype of Adar1 KO mice (Liddicoat et al., 2015; Mannion et al., 2014; Pestal et al., 2015).

Approximately half of the mammalian genome is composed of non-coding retrotransposons, such as SINEs (short interspersed nuclear elements) and Alu elements, which typically form dsRNA duplexes (Lander et al., 2001; Waterston et al., 2002). Retrotransposons are subjected to extensive A-to-IRNA editing by ADAR (Ramaswami et al., 2012). It is possible that the location of repetitive elements determines theirimmunogenicity. Retrotransposons located within introns do not persist in the cytosol and therefore are unable to activate MDA5. Repetitive elements in 3′ UTRs, although rare, can be retained and form duplexes to activates MDA5. Editing of self-dsRNA by ADAR1 generates multiple I-U mismatches to avoid MDA5 recognition (Berke et al., 2012; Wu et al., 2013). In the absence of ADAR1 activity, long dsRNA stem loops can activate MDA5. However, A-to-I editing also alters the RNA secondary structure, which may lead to recognition by innate immune receptors (Liddicoat et al., 2015; Pestal et al., 2015). Recent data have shown that ADAR1 predominantly edits Alu elements in RNA polymerase II (pol II)-transcribed mRNAs, but not those transcribed by pol III in human cells (Chung et al., 2018), and A-to-I conversion in Alus suppresses MDA5 activation in vitro(Ahmad et al., 2018).

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Eukaryotic RNA Processing*

In Cell Biology (Third Edition), 2017

A-to-I Editing

The enzyme ADAR (adenosine deaminase acting on RNA) can convert adenine residues to inosine by deamination of the base (Fig. 11.7). Inosine acts like guanosine and base-pairs withcytosine rather than uracil, potentially altering the protein encoded by the mRNA. Most of the transcripts edited by ADARencode receptors of the central nervous system, and RNA editing is required to create the full receptor repertoire. Theamino acid substitutions that result from editing of the mRNAs can greatly alter the properties of ion channels, and aberrant editing occurs in various disorders ranging from epilepsy to malignant brain gliomas. ADAR binds as a dimer to imperfectdouble-stranded RNA duplexes, which are formed between the target site and sequences in a flanking intron. Editing is generally not 100% efficient, so heterogeneous populations of proteins are generated.

In addition to specific editing of individual nucleotides, ADARs can hyperedit long double-stranded RNAs (dsRNAs). In mammals, dsRNAs elicit a strong antiviral response from theinnate immune system and hyperediting is important to avoid inappropriate recognition of endogenous dsRNAs.

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RNA Editing

Mark R. Macbeth, Brenda L. Bass, in Methods in Enzymology, 2007


Various ADAR proteins purified using this scheme have been used for kinetic analyses, RNA binding assays, and X-ray crystallography (Haudenschild et al., 2004; Macbeth et al., 2004, 2005). Here we present an analysis of theoligomerization state of hADAR2, as purified from S. cerevisiae,using analytical gel filtration and equilibrium sedimentation.

3.1 Analytical gel filtration

For an analytical gel filtration analysis, 50 μl of purified, concentrated (6.6 mg/ml) hADAR2 is injected onto a Superdex 200 10/300 gel filtration column using an AKTA FPLC system (GE Healthcare). The protein was eluted with 20 mM Tris–HCl (pH 8.0), 200 mM NaCl, 5% glycerol, and 1 mM 2-mercaptoethanol, and the elution profile was analyzed with the PrimeView software package (GE Healthcare, Fig. 15.3B). The elution volume of the hADAR2 peak was determined to be 14.4 ml. Relative to the elution of standard molecular weight markers, hADAR2 has an observed molecular weight of 98.1 kDa and appears to be a monomer under these conditions (Fig. 15.3B).

3.2 Equilibrium sedimentation

To observe the oligomeric state of hADAR2, equilibrium sedimentation is performed as it allows an accurate determination of the molecular weight of a macromoleculethat is independent from its shape. Purified hADAR2 was dialyzed against a buffer containing 20 mM Tris–HCl (pH 8.0), 200 mM NaCl, and 1 mM 2-mercaptoethanol and diluted to 1.3, 3.3, and 6.6 μM. The protein was centrifuged in a Beckman XLA analytical ultracentrifuge, equipped with the AN-60 Ti rotor, at 16,000 and 18,000 rpm. Figure 15.3C shows data, fits, and residuals for hADAR2 spun at the three concentrations. The best overall fit of the data was to a single species molecule with an observed molecular weight of 75,320 Da. The residuals are relatively small and randomly distributed, indicating a good fit of the data. The MWobs/MWcalc is 0.98, suggesting hADAR2 exists as a monomer under these concentrations.

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Fidelity and Quality Control in Gene Expression

Stefan Maas, in Advances in Protein Chemistry and Structural Biology, 2012

C Enzyme-Substrate Recognition

Little is known about how ADARs interact with their specific substrates and how the RNA binding domains and the catalytic domain work together during the editing reaction. In contrast to the C-to-U RNA editing machinery that employs a primarysequence motif close to the editing site as critical determinant for substrate recognition (Niswender, 1998), the ADARenzymes lack any apparent intrinsic sequence specificity. In fact, when presented with completely double-stranded RNA molecules, ADAR1 as well as ADAR2 will deaminate almost promiscuously up to 50–60% of all adenosines in the sequence (Nishikura, 2010). However, the presence of loops, mismatches, and bulges within an RNA substrate structure increases site-selectivity and, as seen for some physiologicalADAR targets, the intricate RNA fold of such a substrate seems to restrict access for binding on the RNA and productive catalysis often to a single adenosine (Nishikura, 2010). Therefore, most of the target specificity of A-to-I editing may be residing within the three-dimensional fold of the substrate. However, recent NMR-based structural studies of ADAR protein RNA binding domains in complex with minimal RNA substrates indicate that individual dsRNA binding domains may also be involved in sequence-specific contacts that could contribute to selectivity for certain RNA targets over others (Stefl et al., 2006, 2011).

Another property of ADARs that may have implications for substrate recognition and interaction is the realization that for productive and high activity deamination, the ADAR proteins are forming a dimer on the substrate (Jaikaran et al., 2002; Cho et al., 2003; Gallo et al., 2003; Poulsen et al., 2006). Potentially, homo- as well as heterodimers between ADAR proteins may form (Chilibeck et al., 2006) and therefore represent another layer of regulation influencing substrate-specific recognition as well as RNA editing activity. Since to date there is no known editing target for ADAR3 and this protein does not display anyenzymatic activity in standard deaminase assays (Melcher et al., 1996a; Chen et al., 2000), its role has been suggested to be of regulatory nature through heterodimerization with the other ADARs.

Related to the formation of ADAR dimers for general editing activity is the observation that certain RNA editing events occurring within the same substrate exhibit positive or negative coupling. For example, adenosines directly neighboring each other or those that are located on the same side of the RNA duplex structure (about 11 nt distance within A-form RNA) often show coupling in editing by ADARs probably due to their proximity to the catalytic site of the enzyme in space (Koeris et al., 2005; Enstero et al., 2009).

Despite these insights, it is still not possible currently to predict an RNA editing site based on RNA primary sequence. This is mostly due to the complex nature of RNA tertiary structures and their dynamic behavior. Even when considering that dsRNA binding domains may be able to discriminate certain primary sequence motifs over others, small alterations in the surrounding primary sequence or secondary and tertiary structure may modulate or even override the sequence-specific contacts. As a result, attempts to devise search algorithms combining several of the molecular features (including base-pairing likelihood, sequence environment, sequence conservation) that are known to influence editing likelihood or activity in order to predict potential editing sites have resulted in long lists of candidate positions in mRNAs but few novel, site-selective and high-level modification sites have been validated as bona fide in vivo targets (Clutterbuck et al., 2005; Levanon et al., 2005; Gommans et al., 2008; Sie and Maas, 2009). The dichotomy between the detection of thousands of likely editing sites based on molecular features in pre-mRNAs and the paucity of high-level recoding sites suggests that either the prediction algorithms suffer from a high false positive rate and most of these candidate positions are not edited by ADARs, or the experimental detection of RNA editing is the problem and for many real editing sites the wrong tissues are tested (cell-type specific editing) or they are tested at the wrong time point (regulated editing). Alternatively, the in vivo editing levels of most of the sites are so small that current detection methods are not sensitive enough to prove or disprove editing.

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Biology and Mechanisms of Short RNAs in Caenorhabditis elegans

Alla Grishok, in Advances in Genetics, 2013

3.2.4 Competition between ADARs and endogenous RNAi

There are two C. elegans genes encoding ADARsadr-1 andadr-2 (Tonkin et al., 2002). Although only ADR-2 contains acatalytic domain and active editing function, ADR-1 modulates the activity of ADR-2 in vivo (Knight & Bass, 2002; Tonkin et al., 2002). It was shown earlier that somatic expression of repetitivetransgenes was silenced in ADAR mutants in an rde-1- and rde-4-dependent manner (Knight & Bass, 2002). Therefore, a similar competition between endogenous RNAi and ADARs for dsRNA substrates can be expected. Indeed, deep sequencinganalyses identified a number of low-to-moderate copy inverted repeat regions with a dramatic increase in short RNA reads in ADAR mutants; consistently, transcripts from such loci were found to be multiply edited (Wu, Lamm, & Fire, 2011). Interestingly, although a corresponding decrease in mRNA levels was often observed, histone messages remained unchanged in ADAR(−) animals despite dramatic increases in short RNAs corresponding to some histone loci (Wu et al., 2011).

The effect of ADARs on the biogenesis of short RNAs was examined in another study (Warf et al., 2012). Although the levels of many miRNAs were increased in the absence of ADARs, most of these effects were found to be indirect, likely due to decreased sequestering of pri-miRNAs by ADARs fromDrosha processing; only a couple miRNAs were found to be edited by ADARs (Warf et al., 2012). Surprisingly, generation of endo-siRNAs, which were selected by a 5′monophosphate-dependent sequencing protocol, was predominantly suppressed in the ADAR mutants, with ~40% of annotated loci producing fewer antisense siRNAs (Warf et al., 2012). These results are more consistent with the indirect effects of ADAR loss. Conversely, production of many Dicer-dependent 26G-antisense RNAs and their complementary 19 nt passenger strands was increased in the ADAR mutants, consistent with the competition between Dicer and ADARs for the dsRNA precursors of 26G-RNAs (Warf et al., 2012). As ADARs localize to the nucleus (Hundley, Krauchuk, & Bass, 2008), it is most plausible that the RRF-3-dependent synthesis of 26G-RNA precursors takes place there (Warf et al., 2012).

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