The Importance of Internal Loops within RNA Substrates of ... · The Importance of Internal Loops within RNA Substrates of ADAR1 Katrina A. Lehmann and Brenda L. Bass* Department

Article No. jmbi.1999.2914 available online at http://www.idealibrary.com on J. Mol. Biol. (1999) 291, 1±13

The Importance of Internal Loops within RNASubstrates of ADAR1

Katrina A. Lehmann and Brenda L. Bass*

Department of Biochemistryand HHMI, University ofUtah, 50 N. Medical DriveSalt Lake City, UT 84132, USA

E-mail address of the [email protected]

Abbreviations used: ADARs, adethat act on RNA; dsRNA, double-stRNA-binding proteins; dsRBM, dsRss, single-stranded.

0022-2836/99/310001±13 $30.00/0

Adenosine deaminases that act on RNA (ADARs) are a family of RNAediting enzymes that convert adenosines to inosines within double-stranded RNA (dsRNA). Although ADARs deaminate perfectly base-paired dsRNA promiscuously, deamination is limited to a few, selectedadenosines within dsRNA containing mismatches, bulges and internalloops. As a ®rst step in understanding how RNA structural featurespromote selectivity, we investigated the role of internal loops withinADAR substrates. We observed that a dsRNA helix is deaminated at thesame sites whether it exists as a free molecule or is ¯anked by internalloops. Thus, internal loops delineate helix ends for ADAR1. SinceADAR1 deaminates short RNAs at fewer adenosines than long RNAs,loops decrease the number of deaminations within an RNA by dividinga long RNA into shorter substrates. For a series of symmetric internalloops related in sequence, larger loops (5six nucleotides) acted as helixends, whereas smaller loops (4four nucleotides) did not. Our workprovides the ®rst information about how secondary structure withinADAR substrates dictates selectivity, and suggests a rational approachfor delineating minimal substrates for RNAs deaminated by ADARsin vivo.

# 1999 Academic Press

Keywords: deaminase; double-stranded RNA; inosine; RNA editing
*Corresponding author
Introduction

Adenosine deaminases that act on RNA(ADARs) convert adenosines to inosine residueswithin cellular and viral RNAs (see reviews byBass, 1997; Maas et al., 1997; O'Connell, 1997). Theenzyme activity has been detected in manymetazoa, usually in the nucleus, and cDNAs for atleast two distinct ADARs (ADAR1 and ADAR2)have been cloned. Like RNA splicing, adenosinedeamination is a way to diversify the informationencoded by a gene. For example, ADARs candeaminate adenosines within codons so thatmultiple protein isoforms can be synthesized froma single encoded mRNA. ADARs are involved inproducing isoforms of mammalian serotoninreceptors (Burns et al., 1997), several mammalianglutamate receptor subunits (Egebjerg &Heinemann, 1993; Higuchi et al., 1993; Lomeli et al.,

ing author:

nosine deaminasesranded RNA; RBP,NA binding motif;

1994), and the virally encoded hepatitis delta anti-gen (Polson et al., 1996). ADARs may haveadditional functions as well; in fact, other cellularand viral substrates have been identi®ed for whichthe function of deamination is unclear (Bass, 1997).

ADAR substrates fall into two general categoriesbased on how many of their adenosines aredeaminated, i.e. how selectively they are deami-nated (Bass, 1997). (The term selectivity has ade®ned meaning in the ADAR ®eld and refers tothe percentage of the total number of adenosines ina base-paired region that are deaminated atcomplete reaction). Selectively modi®ed substratesare deaminated at less than 10 % of theiradenosines, while substrates that are promiscu-ously, or non-selectively, modi®ed are deaminatedat 50-60 % of their adenosines. Obviously,promiscuous deamination is ill-suited for generat-ing protein isoforms of precise function, and inmRNAs where ADARs produce a functionallyimportant codon change, deamination occurs selec-tively. Although it was initially speculated thatADARs would require accessory factors for selec-tive deamination, many studies show that ADARsalone are capable of very selective deamination(Hurst et al., 1995; Dabiri et al., 1996; Melcher et al.,

# 1999 Academic Press

2 Internal Loops Promote Selective ADAR Deamination

1996; Polson et al., 1996). It is now generallyaccepted that features of the RNA substrate dictateselectivity.

The speci®c features of ADAR substrates thatdetermine selectivity are unclear. In vitro studiesshow that ADAR1 prefers to deaminate adenosineswith a 50 neighbor of A, U, or C, over those with a50 neighbor of G. In addition, adenosines very closeto the 30 end of a strand are deaminatedinfrequently. These preferences are observed inboth selectively and non-selectively deaminatedsubstrates, but are not suf®cient to promote theexquisite selectivity observed within some biolo-gical substrates (see Polson et al., 1996). Althoughfew studies have been performed, ADAR2 appearsto have overlapping, but slightly different,speci®cities (Melcher et al., 1996).

It has been proposed that the structure of theRNA substrate, rather than its sequence, mightdetermine selectivity (Bass, 1997). ADARs are cate-gorized as double-stranded RNA-binding proteins(dsRBPs) and like other dsRBPs (e.g. RNase III andPKR) require substrates that are completely, orlargely, double-stranded. A survey of characterizedADAR substrates shows that RNAs that are com-pletely double-stranded are deaminated promiscu-ously (non-selectively), while RNAs whose base-paired structures are periodically interrupted bymismatches, bulges and loops are deaminated

Figure 1 (Legen

selectively. Although mutagenesis studies clearlyshow that base-pairs surrounding a given editingsite are important for the precise and ef®cient dea-mination of that site (Higuchi et al., 1993; Lomeliet al., 1994; Yang et al., 1995; Polson et al., 1996), theeffect of loops, mismatches and bulges on the totalnumber of deamination sites within a given helix,i.e. selectivity, has not been investigated.

As a ®rst step in understanding how RNA struc-ture allows selective deamination, we have focusedon the role of internal loops within ADAR sub-strates. Previous studies show that short duplexesare deaminated more selectively than longduplexes (Nishikura et al., 1991; Polson & Bass,1994). Thus, we reasoned that internal loops withinan ADAR substrate could uncouple adjacenthelices to convert a long, promiscuously deami-nated substrate into a series of short, selectivitydeaminated substrates. Our experiments supportthis hypothesis, thus providing the ®rst infor-mation in regard to how RNA structure determinesADAR selectivity.

Results

We examined the role of internal loops in pro-moting ADAR1 selectivity using six RNA mol-ecules (Figure 1(a)). One molecule consisted of acompletely base-paired dsRNA of 36 bp (36mer),

d opposite)

Figure 1. Nucleotide sequence and RNase T2 structure mapping of RNAs. (a) The same numbering scheme is usedfor the 36mer sequence of each RNA (blue), and purine (R) and pyrimidine (Y) rich strands are indicated. Internalloops, GC-rich helices, and the central helix of barbell molecules are labeled. (b) L0, L4, L6 and L12 moleculescontaining an R strand labeled at its 50 end with 32P were incubated in the presence or absence of RNase T2, underthe same native conditions used in reactions with ADAR1 (see Materials and Methods). In separate reactions, RNAwas digested with RNase T1 under denaturing conditions for sequence orientation purposes (data not shown). Incontrast to a single R strand (ss), the barbells were protected from RNase T2 cleavage in regions predicted to be base-paired. Consistent with the predicted structures, the left loop, right loop and 30 overhang were digested with RNaseT2. The L12, L6 and L4 loops show the expected number of RNase T2-accessible nucleotides, and one or two accessi-ble nucleotides 30 of the predicted loop closure; moderate increases in solvent accessibility of nucleotides 30 of internalloops has been observed previously (Weeks & Crothers, 1993). The L6 and L4 loops show a degree of T2 accessibilityintermediate to that of the L12 loops and the corresponding region of L0. All four RNAs show similar RNase T2

cleavages in the 30 overhang. The secondary structure of L8 was not determined, but subsequent studies showed thatL8 was deaminated by ADAR1 with the same selectivity as barbell RNAs with larger (L12) and smaller (L6) loops(see Figure 3(a) and (b)) whose structures were con®rmed. OH- indicates alkaline hydrolysis lanes. Numbering isshown to the right of the gel, along with parentheses indicating the central helix, left loop, right loop, and30 overhang.

Internal Loops Promote Selective ADAR Deamination 3

formed between a purine (R) and a pyrimidine (Y)rich strand; previous studies show that this mol-ecule is deaminated by ADAR1 and has six deami-nation sites at complete reaction (Polson & Bass,1994). Four of the molecules were ``barbell'' mol-ecules that contained the 36mer sequence boundedon each side by symmetric internal loops of 12(L12), eight (L8), six (L6), or four (L4) nucleotides;

loops were closed with GC-rich helices at eachterminus. Barbell molecules of decreasing loopsizes (L8, L6, L4) were created by changing thesequence of the L12 Y strand so as to extend thelength of the GC-rich helix and decrease the loopsize; thus, the R and Y strands had identicallengths from one barbell to the next. Importantly,the external GC-rich helices of all molecules were


predicted to be too short to serve as ADAR sub-strates if separated from the rest of the molecule.In the sixth molecule, L0, the GC-rich helices wereextended completely to form a continuous helixwith the 36mer sequence; this molecule was calledL0 to convey the fact that it contained loops ofzero nucleotides. All molecules except the 36mercontained a 12 nt 30 overhang on each strand tofacilitate primer binding during primer extensionanalyses.

The loops of the barbells consisted of adenosinesand cytidines and were designed to minimizepairing and stacking interactions. We con®rmedthat these sequences were unpaired internal loopsby treating the molecules with ribonuclease T2

(RNase T2; Figure 1(b)), which cleaves 30 ofunpaired nucleotides. Consistent with thepredicted structures, pronounced T2 cleavageoccurred in all loops, as well as the 30 overhangs,but not within base-paired regions. As expected,only the 30 overhang was cleaved in the L0molecule.

As mentioned, our starting hypothesis was thatinternal loops, by uncoupling adjacent helices,could convert one long ADAR substrate into aseries of shorter substrates. If this hypothesis wascorrect, and each RNA helix was indeed uncoupledfrom adjacent helices by the loops, ADAR1 shouldtreat internal loops as helix termini. That is, thedeamination pattern within a particular sequenceshould be identical whether the sequence existedas a free dsRNA or was bounded by internalloops.

To test this idea we reacted 36mer and L12 withADAR1 and compared the reaction products.RNAs were internally labeled with [32P]ATPduring transcription, and adenosine deaminationwas measured by determining the amount of[32P]AMP converted to [32P]IMP at various timepoints (Figure 2(a)). Previous studies show thatdsRNAs greater than �50 bp are deaminated pro-miscuously by ADAR1, showing 50-60 % deamina-tion at reaction completion (Polson & Bass, 1994).Since the L12 molecule contained a total of 54canonical base-pairs, we expected at least 50 % ofthe adenosines would be deaminated at completereaction if the internal loops were not acting ashelix termini. However, we found that 36mer andL12 were deaminated to the same extent through-out the course of the experiment (zero to fourhours, see the legend to Figure 2). A 5 fmol sampleof each substrate was reacted, and both moleculesshowed between 21-23 fmol of inosine at completereaction. These data showed that ADAR1 deami-nated the 36mer and L12 with the same selectivity,which supported our idea that the L12 loops wereacting to uncouple the 36 bp helix from its adjacentexternal helices.

We next determined which adenosines weredeaminated in each molecule using a primerextension assay (Figure 2(b)). Since inosine prefersto pair with cytidine, in this assay, inosines areevident by the loss of a ddTTP stop and the corre-

sponding appearance of a ddCTP stop. Again, theresults supported the idea that ADAR1 recognizesinternal loops as helix termini. At complete reac-tion with ADAR1, 36mer and L12 were deami-nated at the same six primary deamination sites(A5, A6, A9, A10, A14, A15; Figure 2(c)). The onlydifference between the two molecules occurred atA2, which was deaminated to a very minor extentin the L12 molecule (see Discussion). Thus, for themost part, ADAR1 appeared to recognize the L12loops as helix ends, and the selectivity observed inthe free 36mer molecule was maintained. The factthat the deamination sites were almost identicalin the two molecules further emphasized that theloops were functioning as effective helixends, since ADAR1 disfavors adenosines close to30 termini. If the loops were not acting as effectivehelix ends, we would expect deamination ofadditional adenosines closer to the loop, such asthose between A16 and A34.

We next wanted to determine how short aninternal loop could be and still function as a helixterminus. We extended our analysis to include bar-bells L8, L6, and L4, which have loops of eight, sixand four nucleotides, respectively (Figure 1(a)). Wealso analyzed L0 (Figure 1(a)), the molecule inwhich the barbell loops had been closed to form acompletely base-paired molecule of 66 bp. Wereasoned that barbells with loops that coulduncouple helices would be deaminated similarly to36mer, while those with loops that could notuncouple adjacent helices would be deaminatedpromiscuously, and show a deamination patternsimilar to that of L0. We performed a time-courseexperiment like that described in the legend toFigure 2(a), and analyzed the accumulation of ino-sine within all six RNAs over time (Figure 3(a)). Ateach time point, the amount of inosine observedfor L12, L8, and L6 was almost identical with thatof the 36mer, suggesting the loops in these mol-ecules could uncouple adjacent helices. However,throughout the time-course the L4 barbell con-tained much more inosine, approaching thatobserved in L0; this suggested that, at least for thesequences we tested, loops 5six nucleotides, but

Figure 2. Comparison of the reaction of 36mer and L12 RNAs. (a) RNAs were incubated with ADAR1, and at var-ious times, aliquots were removed, digested to 50 nucleotide monophosphates (50NMPs), and analyzed by TLC. Thegraph shows the amount of IMP (fmol) produced during the incubation of 36mer (squares) and L12 (circles) withADAR1. Each point is the average of two to four experiments, with standard deviation bars shown. The amount ofIMP observed at four hours (data not shown) veri®ed that the reaction was complete at two hours, and control exper-iments con®rmed that the enzyme was active throughout the incubation. (b) The sites of deamination within 36merand L12 were mapped by primer extension sequencing using reverse transcriptase (RT). RNAs were incubated fortwo hours with (�) or without (ÿ) ADAR1 and subsequently reverse-transcribed in the absence of dideoxynucleo-tides (lanes 2-3), or in the presence of ddGTP (lane 4), ddATP (lane 5), ddTTP (lanes 6-7) or ddCTP (lanes 8-9). Lane1 was loaded with the primer only. Numbers to the right of each gel show adenosine number, and parentheses indi-cate the left and right loops of L12. (c) The diagram shows the location of inosines as determined in (b), and wascon®rmed by mapping with a mutant (E46Q) RNase T1 (data not shown; see Materials and Methods). Based oncomparisons with the RNase T1 mapping data, deaminations evident by the loss of a ddTTP stop, as well as thecorresponding gain of a ddCTP stop, were considered to occur in the majority of the population of molecules(I), whereas deaminations evident only by the gain of a ddCTP stop were considered to be minor deaminations thatexisted in a minor fraction of the population (i). Italicized adenosines within L12 mark a region of possible locaconformational heterogeneity. Deamination sites were not observed within the L12 loops.


l

Figure 3. Determination of the minimal loop lengthrequired to uncouple helices in the barbell molecules. (a)All RNAs shown in Figure 1(a) were incubated withADAR1 and the amount of IMP produced over timewas determined as described in the legend to Figure 2.For each RNA, the reaction was complete at two hours,as veri®ed by analyses of four hour time points. Thesymbols used in the graphs are correlated with theappropriate RNAs to the right of the plot. The two hourpoints are the average of four (36mer, L12 and L8), ®ve(L6), or six (L4 and L0) experiments, whereas shortertime points are the average of two experiments;standard deviation (s.d.) bars are shown. (b) The TLCplate shows relative amounts of AMP and IMP at arepresentative two hour time point used for the graphshown in (a). The positions of AMP, IMP, and the origin(ori) are labeled to the right of the plate. RNAs wereincubated with (�) or without (ÿ) ADAR1 prior todigestion to 50NMPs. The average amount of IMPobserved for all two hour time points is given below thecorresponding lane, with s.d.


not those 4four nucleotides, were equivalent tohelix termini.

Figure 3(b) shows the primary data for the twohour time point (when the reaction is complete), aswell as the amount of inosine observed. Reactionscontaining 5 fmol of 36mer, L12, L8 or L6contained 21-23.5 fmol of inosine at reaction

completion. In contrast, much more inosine wasobserved after the complete reaction of 5 fmol ofL4 (�47 fmol) or L0 (�58 fmol). Although therange of values observed for the L4 and L0samples were slightly overlapping at the two hourtime point (see error bars, Figure 3(a)), within agiven experiment the amount of inosine observedin L0 was consistently 20 % higher than thatobserved in L4. The overall time-course alsoemphasizes that L4 and L0 react with ADAR1slightly differently. While the loops of four nucleo-tides are not acting as internal loops to uncoupleadjacent helices, they are probably affecting thesubstrate in other important ways (see Discussion).

We next mapped the inosines within L6 and L4by primer extension (Figure 4(a)). L4 containedmany more deamination sites than L6, and thelocation of each deamination site is shown inFigure 4(b). L6 showed exactly the same deamina-tion pattern as 36mer, indicating that its loopswere acting as helix termini. In contrast, L4 con-tained more than twice the number of deaminationsites as L6, and had a pattern of deamination verysimilar to that seen in L0. Consistent with the ideathat the external GC-rich helices are too short toact as ADAR substrates by themselves, none of theadenosines in the external helices of L6 were dea-minated. In contrast, in L4 two adenosines in oneof the external helices were deaminated (see smalli's in Figure 4(b)); these two adenosines were alsodeaminated in L0, emphasizing that ADAR1 isrecognizing L4 and L0 as similar molecules.

Each time we decreased the length of a loop tomake a different barbell RNA, we concurrentlyincreased the length of the adjacent GC-rich helix.Thus, it was possible that the lower selectivityobserved in L4 compared to L6 was due to theincrease in the length of the external helices (andthus an increase in the overall stability of theduplex) rather than its smaller loops. To test thispossibility, we created a chimeric barbell RNA thatcontained the L6 loops adjacent to L4 externalhelices (L6/L4 chimera). The L6/L4 chimerashowed the same selectivity as L6 (25.9 (�2.9) fmolinosine, n � 5; data not shown) but half the totalinosines as L4. Thus, the ability of the L6 barbell tobe deaminated with the same selectivity andpreferences as 36mer is dependent on its internalloops and not its external helices.

Discussion

Here we show that internal loops play animportant role in determining the number of dea-mination events that occur within an ADAR sub-strate. In particular, we show that internal loopsare equivalent to helix termini for ADAR1. Thus, asingle, long dsRNA substrate can be converted intoa series of short, independent substrates by theinsertion of internal loops. Since ADARs deaminateshort dsRNAs more selectively than long dsRNAs,internal loops can dramatically decrease the num-


ber of deamination events that occur within anRNA. The restriction of adenosine deamination toa select number of sites is essential when ADARsact on mRNAs to alter speci®c codons. Our workprovides the ®rst information on how secondarystructure within ADAR substrates contributes toselective deamination and also adds to the generalunderstanding of how internal loops functionwithin RNAs.

A selectivity model

In our favorite model, all aspects of ADAR selec-tivity can be understood with respect to its nucleicacid binding properties. Simply put, ADARs bindto double-stranded RNA but not single-strandedRNA (Bass & Weintraub, 1987; Wagner &Nishikura, 1988). Since ADARs are not sequencespeci®c (Polson & Bass, 1994), an increase in thelength of a dsRNA, or the number of base-pairs itcontains, should increase the number of ADARbinding sites; similar arguments have been madewith regard to sequence-independent DNA-bind-ing proteins (McGhee & von Hippel, 1974).According to our model, a long dsRNA would bedeaminated at more adenosines because it hasmore binding sites. For example, if we assume thata single molecule of ADAR1 interacts with 20 bp,we would predict the 36mer has 17 differentADAR binding sites (36 ÿ 20 � 1), while the L0molecule, with 66 bp, would have 47 differentbinding sites.

For the barbell molecules analyzed, we observedthat loops 5six nucleotides uncoupled adjacenthelices and increased selectivity, while loops4four nucleotides did not (Figures 3(b) and 4(b)).According to our model, and as shown in Figure 5,this difference exists because the internal loops ofL6 decrease the number of binding sites comparedto L4. To understand why loops must be a certainlength to uncouple adjacent helices, it is importantto remember that a particular sequence of RNA isnever completely paired or unpaired, but at equili-brium favors one state over the other. Since theunfavorable entropy associated with incorporatingloop nucleotides into a helix increases with looplength, the population of L6 molecules will havefewer paired loops at equilibrium than the popu-lation of L4 molecules. Further, the addition of adsRBP such as ADAR1 would shift the equilibriumtoward the paired state. Although the bindingenergy incurred when an ADAR interacts with asubstrate might be able to compensate for disrup-tion of a helix by a single mismatch, successiveunpairing of adjacent base-pairs will eventuallycreate an internal loop whose free energy favorsthe unpaired state. From these considerations weexpect that the minimal length necessary for a loopto function as a helix end will vary with thesequence of the loop and adjacent helices, andfurther, that certain asymmetric internal loops willalso act as helix termini. Our future studies will beaimed at exploring these issues.

Our experiments with the L12 barbell suggestthat some loops will be too large to maintain the®delity of deamination within an adjacent helix. Incontrast to the other barbell molecules studied, thedeamination sites within L12 were not identicalwith those within the 36mer. In particular, oneadditional, very minor, deamination site occurredat A2 of L12 (Figure 2(c)). We suspect that thelarge loops of L12 are more destabilizing than theloops of other barbells, and may be exploringalternative conformers that involve nearbysequences. Possibly, an alternate pairing betweenthe loops and bases within the helix promotes dea-mination at A2. Based on our observations withL12 and the other barbell molecules, we predictthat loops that can both de®ne helix termini forADARs and maintain the ®delity of deamination,will have an optimal size range, being neither toosmall to uncouple helices (like L4), nor too large tosample competing conformations with nearbysequences.

Given that ADAR1 prefers to bind dsRNA overssRNA, anything that shifts the equilibriumbetween the paired and unpaired state shouldeffect ADAR binding. Thus, in addition to helicallength, alterations of the thermodynamic stabilityof a duplex, such as the insertion of mismatches,should alter its selectivity. We propose that L4 isdeaminated a bit more selectively than L0, becauseits internal loops, although too short to act as helixtermini, increase the single-stranded character ofthe RNA. These ideas are related to those raisedduring early studies of ADAR to explain why dea-mination of completely base-paired RNAs stoppedwhen 50-60 % of the adenosines were deaminated.Since modi®cation of AU base-pairs in dsRNAcreates the considerably less stable IU mismatch(Bass & Weintraub, 1988; Stroebel et al., 1994), itwas proposed that the reaction stopped when themolecule no longer had enough double-strandedcharacter to be bound and deaminated by anADAR (Bass & Weintraub, 1987; Polson & Bass,1994).

What structural features allow internal loops tofunction as helix ends?

The RNase T2 experiments were performedunder the same conditions used for ADAR1 reac-tions, and indicate that under these conditions, thestructures of the barbells are largely as wedesigned them. That is, sequences designed to existas unpaired loops were digested with the single-strand-speci®c ribonuclease, while regionsdesigned to exist as double-stranded helices wereresistant. However, the RNase T2 data do not givespeci®c details about possible non-canonical fea-tures of the loops. Further, since RNase T2 is asingle-stranded RNA binding protein, the additionof RNase T2 to the RNA solution could shift theequilibrium between the paired and unpairedstates from that normally encountered by ADAR1.

Figure 4 (Legend shown opposite)


Figure 5. A model to explain why internal loops promote selectivity. According to our model, internal loops pro-mote selective deamination by decreasing the number of ADAR1 binding sites within a substrate. ADAR1 molecules(green) are shown interacting with a completely base-paired dsRNA (c), or the same molecule with internal loopslarge enough to uncouple helices (a) or too short to uncouple helices (b). Since ADARs are not sequence speci®c,binding sites occur along the entire length of a completely base-paired dsRNA, and these molecules are deaminatedpromiscuously (c). Internal loops like those in (a) decrease the number of binding sites and allow more selectivedeamination; these loops may create a physical end to the helix by thermodynamically uncoupling adjacent helices.As depicted for the external helices of the molecule in (a), internal loops may sometimes increase selectivity by creat-ing helices that are too short to support ADAR binding. The internal loops in the molecule shown in (b), while tooshort to uncouple helices, decrease the overall stability of the molecule. As previously proposed (Polson & Bass, 1994;Bass, 1997), mismatches and small internal loops may decrease the number of deamination events by shifting theconformational equilibrium towards the single-stranded state, thus decreasing the number of ADAR binding sites inthe population. As shown for each example, the IU mismatches (red) that accumulate during the reaction ultimatelylead to a molecule that is too unstable to be further modi®ed, and at this point the reaction terminates. See the textfor further discussion. ADAR1 molecules are depicted according to previous observations regarding deaminationpreferences (Polson & Bass, 1994).


Previous studies show that internal loops withinan RNA molecule can increase or decrease thermo-dynamic stability (Peritz et al., 1991; SantaLuciaet al., 1991; Xia et al., 1997), promote conformation-

Figure 4. Location of inosines within L6 and L4. (a) Thewere mapped by primer extension sequencing as describedwithin the central helix is shown to the left of the gel. The pwell as whether the RNA was incubated in the presence (�autoradiogram. RT sequencing products (cDNAs) producedhave a faster gel mobility due to their altered nucleotide cunmodi®ed samples (ÿADAR1; lanes 1, 3, 5, 7). Further, sinpopulations of molecules, bands in these lanes are less sharthe same method (data not shown). In some instances, seunclear, possibly due to abortive RT elongation of the primwithin 36mer, L6, L4 and L0 RNAs. The continuous blackloops are long enough to uncouple helices (5six nucleotides)(4four nucleotides). Inosines were not observed within theinosines, since previous studies (Polson & Bass, 1994) shownot deaminated. While L0 may contain deaminated adenosiduplex, the possible presence of such deaminations does not

al ¯exibility such as bending (Zacharias &Hagerman, 1996), and widen the major groove ofhelical regions (Weeks & Crothers, 1993). Structuralanalyses show that the nucleotides of some loops

locations of inosines in L4 (lanes 1-4) and L6 (lanes 5-8)in the legend to Figure 2(b). Numbering of adenosines

articular dideoxynucleotide included in each reaction, as) or absence (ÿ) of ADAR1 is indicated at the top of thefrom deaminated RNAs (� ADAR1; lanes 2, 4, 6, 8-10)

omposition (cytidines instead of thymidines) relative toce cDNAs derived from deaminated RNAs are complex

p in appearance. Inosines within L0 were mapped usingquence information near the primer annealing site was

er. (b) The diagram compares the location of inosinesline indicates the boundary between molecules whoseand those whose loops are too short to uncouple helices

L6 and L4 loops. Barbell Y strands were not mapped forthat the three adenosines within the 36mer Y strand arenes on its Y strand due to the increased stability of thisaffect the conclusions drawn in this study.


incorporate into adjacent helices (Holbrook et al.,1991; Jang et al., 1998), while those of others dis-rupt the structure of an otherwise continuousA-form helix (Wimberly et al., 1993; Cai & Tinoco,1996; Correll et al., 1997). At present we do notknow which of these structural features, if any, areimportant for interactions of loops with ADARs.However, as suggested by the selectivity modelproposed above, we prefer the idea that internalloops act as helix ends not because ADARs recog-nize and interact with their particular structuralfeatures, but because their particular structural fea-tures cannot be recognized as an RNA helix. Thisidea is consistent with the fact that editing sites inRNAs whose structures are proven occur in cano-nical base-pairs or single base mismatches. How-ever, it is important to note that there is one reportof deamination sites occurring within internalloops (Herb et al., 1996). At present it is not clear ifthis is a true anomaly or that the predicted struc-ture is incorrect.

Our work ®ts nicely in the context of previousstudies that demonstrate the ability of internalloops to thermodynamically uncouple RNA helices(Weeks & Crothers, 1993). These previous studiesindicate that internal loops are able to uncouplehelices by decreasing enthalpically favorable stack-ing interactions so as to create an effective helicalend similar to a blunt end. Although future studieswill be required to understand the physical basisof our observations, recognition of RNAs byADARs may provide an example of a protein:RNAinteraction that makes use of the ability of internalloops to thermodynamically uncouple adjacenthelices.

Relationship of ADAR1 to other dsRBPs

Interestingly, like ADAR1, other dsRNA-bindingproteins have been observed to distinguishbetween perfect and irregular RNA duplexes,suggesting that internal loops may serve a similarfunction in these proteins. RNase III acts promiscu-ously on long, perfectly base-paired dsRNA, cleav-ing the RNA at numerous sites to yield 12-15 bpfragments (Robertson, 1982; Nicholson, 1996). Incontrast, imperfect duplexes are cleaved at onlyone to two sites, and internal loops can furtherrestrict cleavage to a single site in some substrates(Chelladurai et al., 1993, and see reviews byRobertson, 1982; Court, 1993; Nicholson, 1996).Likewise, the kinase activity of PKR is activated by33-80 bp perfect dsRNAs, but not by irregular orshorter dsRNAs (see reviews by Mathews &Shenk, 1991; Clemens & Elia, 1997).

All of these proteins (ADAR1, RNase III andPKR) bind dsRNA using an amino acid sequenceknown as the dsRNA binding motif (dsRBM),suggesting it is this shared sequence that mediatesthe discrimination between a perfectly base-pairedduplex and one containing mismatches, bulges andloops. Consistent with our selectivity model, whichproposes differences in secondary structures alter

selectivity by affecting protein binding, bothRNase III and PKR bind perfect dsRNAs withhigher af®nity than irregular dsRNAs (Chelladuraiet al., 1993; Bevilacqua & Cech, 1996). Studies ofthe effects of internal loops within substrates ofother ADARs, such as ADAR2, have not yet beenperformed. However, the fact that all ADARs havedsRBMs, show promiscuous deamination oncompletely base-paired dsRNA and selectivedeamination on dsRNA containing periodicstructural disruptions, suggests internal loops func-tion to uncouple helices in substrates of allADARs. Of course, future studies will need to beperformed to con®rm this.

Implications for biological substrates

Our results help to explain the exquisite selectiv-ity observed in some natural ADAR substrates. Forinstance, the production of the two isoforms ofhepatitis delta antigen relies on highly selectivedeamination of the HDV antigenomic RNA(Polson et al., 1996). In this case, less than 1 % ofthe �340 adenosines of the antigenomic RNA aredeaminated in vivo, or in vitro with puri®edADAR1. The HDV antigenome, while largely base-paired, is predicted to contain numerous internalloops. Based on our studies, we propose that theseinternal loops allow ADAR1 to recognize the long,�1700 nt HDV antigenome as a series of short seg-ments. Due to the frequency of loops within theHDV antigenome, we predict that most of the heli-cal segments are too short to act as ADAR sub-strates, thus limiting deamination to a selectnumber of sites.

Since ADAR1 deaminates helices ¯anked byappropriate internal loops with the same selectivityand preferences as those with blunt termini, it fol-lows that a helical segment between adjacent loopscould be removed from a biological substrate with-out loss of deamination speci®city. Thus, our stu-dies suggest the ®rst rational approach fordelineating minimal substrates for the RNAs thatare deaminated by ADARs in vivo.

Materials and Methods

Construction of transcription vectors

Plasmids pKL36A and pKL36S were used as templatesfor transcription of the Y and R strands of L12, respect-ively, and were constructed from plasmid pGEM-9zf(Promega) after excision of the multiple-cloning regionand phage T7 and SP6 promoter sequences (NotI to S®I).Excised sequences were replaced by an insert thatincluded an optimal T7 promoter sequence (Milliganet al., 1987) and the template to transcribe one RNAstrand of L12. Inserts were created by annealing com-plementary DNA oligos to create dsDNA inserts withNotI and S®I compatible ends, and are shown 50 to 30below (36Acpy and 36Atpl anneal to create the pKL36Ainsert; 36Scpy and 36Stpl anneal to make the pKL36Sinsert): 36Acpy, CGGCCGTCGACTAGTAATACGACTCACTATAGGGCCCGGGAACACACCTGGTCCCTGTC


CTTGTTATTTTCCTTGGTTAATTACACAAGGCGCGCCCACACCACGGATCCATGGGCATGC; 36Atpl, GGCCGCATGCCCATGGATCCGTGGTGTGGGCGCGCCTTGTGTAATTAACCAAGGAAAATAACAAGGACAGGGACCAGGTGTGTTCCCGGGCCCTATAGTGAGTCGTATTACTAGTCGACGGCCGACT; 36Scpy, CGGCCGTCGACTAGTAATACGACTCACTATAGGGCGCGCCAACACAAATTAACCAAGGAAAATAACAAGGACAGGGACCAGGACACAACCCGGGCCCACGCACCAGATCTAGAGCATGC; and 36Stpl, GGCCGCATGCTCTAGATCTGGTGCGTGGGCCCGGGTTGTGTCCTGGTCCCTGTCCTTGTTATTTTCCTTGGTTAATTTGTGTTGGCGCGCCCTATAGTGAGTCGTATTACTAGTCGACGGCCGACT.

Gel-puri®ed oligo nucleotides were phosphorylatedwith phage T4 polynucleotide kinase (T4 PNK) and ATPprior to annealing in 10 mM Tris (pH 8.0), 1 mM EDTA(1 � TE), and ligating with the 2818 bp fragment of thepGEM-9zf vector.

Nucleic acid preparation

Transcription vectors pKL36A and pKL36S were lin-earized at the BamH1 and BglII (NEB) sites, respect-ively, within the inserted sequence for use in run-offtranscription. Transcription reactions (100 ml) contained1 unit/ml phage T7 RNA polymerase (Life Technol-ogies) and the manufacturer's buffer, and included500 mM NTPs, 1.8 units/ml rRNasin (Promega),100 mM DTT, 5 mg linearized plasmid. Other ssRNAs(Y and R strands of 36mer, Y and R strands of theL6/L4 chimera, and Y strands of L8, L6, L4 and L0)were synthesized from partially ssDNA templates thatincluded an optimal T7 promoter sequence (Milliganet al., 1987) using the T7 MegaShortScript in vitro tran-scription kit (Ambion).

RNAs were transcribed in the presence of either0.2 mCi 3000 Ci/mmol [a-32P]ATP for TLC applications(in which case the non-radioactive ATP concentrationwas lowered to 250mM), or 0.1 mCi [5,6-3H]UTP toenable quanti®cation of the transcript. For RNase T2 andRNase T1 mapping applications, the RNA strand to beanalyzed was gel puri®ed, then labeled at its 50 endusing 6000 Ci/mmol [g-32P]ATP as described (Polson &Bass, 1994).

The L12 R strand was annealed with other Y strandsto create all the barbell RNAs shown in Figure 1(a). Gel-puri®ed Y and R strand RNAs were combined in equalmolar amounts in a 15 ml volume in 10 mM Tris (pH 8.0)and annealed by heating to 95 �C for three minutes, fol-lowed by slow cooling to room temperature for �45minutes. Annealed RNAs were incubated brie¯y atÿ20 �C before the addition of 15 ml native gel buffer(2 � � 20 % (w/v) sucrose, 1 � TBE, 0.5 % (w/v) SDS,0.01 % (w/v) xylene cyanol and 0.01 % (w/v) bromophe-nol blue), and electrophoresis at 100 V for 12 hours on a10 % (w/v) polyacrylamide native gel. dsRNAs wereprocessed as described (Polson & Bass, 1994) prior toquanti®cation in triplicate by diethylaminoethyl cellulose(DEAE, Whatman) ®lter paper binding.

Deamination reactions

Adenosine deamination reactions (50 ml) contained5 ml Xenopus laevis xADAR1 AF-Blue-650 M pool (Hough& Bass, 1994) and 5 fmol dsRNA in the standard assaybuffer (1 � AB) described by Hough & Bass (1994),except additional glycerol was excluded and EDTA was

5.1 mM. Both ADAR1 and dsRNA were pre-incubatedseparately in 1 � AB for ten minutes on ice or at 25 �C,respectively, prior to mixing of these components. RNAswere reacted to completion with ADAR1 (120 minutes,25 �C) unless otherwise noted. Reactions were stopped,and RNAs puri®ed as described (Polson & Bass, 1994).

Thin-layer chromatography

For TLC, RNAs were digested to 50NMPs as described(Saccomanno & Bass, 1994), except nuclease P1 reactionswere at 50 �C in 40 ml of 1 � TE. Puri®ed 50NMPs werelyophilized, Cerenkov counted, and resuspended inwater to the same cpm/ml.

PEI-cellulose plates (Bodman) were pre-run in waterand dried as described (Paul & Bass, 1998), prior to spot-ting nuclease P1 reaction products and 50AMP and 50IMPmarkers (prepared as described by Bass & Weintraub(1988)). Spotted plates were dried, then soaked in anhy-drous methanol as described (Paul & Bass, 1998), priorto one-dimensional chromatography in a standard sol-vent (Saccomanno & Bass, 1994). Percentage deaminationwas calculated as described (Saccomanno & Bass, 1994).The molar amount of inosine was determined as the pro-duct of the percentage of deamination, the number ofadenosines per molecule of unreacted duplex, and theamount of dsRNA reacted (5 fmol).

Primer extension

Oligonucleotides used as primers for primer extensionwere gel puri®ed, then 50-end-labeled with phage T4PNK (USB) and 6000 Ci/mmol [g-32P]ATP. Oligo-1(50 CCTGGTCCCTGT 30) was used for primer extension of36mer and L12, and SR14D (50 TCTGGTGCGTGGGC 30)was used for primer extension of barbell RNAs. Oligo-1hybridizes to the last 12 nt of the R strand of the 36mersequence. Mapping of inosines using a mutant (E46Q)RNase T1 (Steyaert et al., 1991) as described (Polson &Bass, 1994) demonstrated that the oligo-1 primer did notobscure deaminated adenosines (data not shown). SR14Dhybridizes to nucleotides in the right GC-rich helix and30 overhang of the R strand.

Inosines were mapped by primer extension asdescribed (Inoue & Cech, 1985) with the following modi-®cations. RNA (50 fmol) was combined with 50 end-labeled primer in 1 � hybridization buffer, incubated at90 �C for three minutes, and placed on dry ice to facili-tate annealing of primer to the RNA. Reverse transcrip-tion reactions contained 2 ml of the above RNA-primermixture, and were done as described (Inoue & Cech,1985), except reactions were incubated at 42 �C for 30minutes with 1.5 units AMV-RT (Boehringer Mannheim).Reactions were performed as full extensions to verifyRNA integrity, or with the addition of 40 mM ddNTP toobtain sequence information. An equal volume of2 � formamide loading buffer (Polson & Bass, 1994) wasadded prior to heating cDNAs (®ve minutes, 90 �C).Electrophoresis (50 W, three hours) was on a 12 % (w/v,19:1) polyacrylamide, 1 � TBE, 8 M urea gel (43 cmlong). Gels were dried and subjected to autoradiographyand Phosphorimager analysis.

RNase T2 structure mapping

Approximately 30,000 cpm of (32P-50)-end-labeledRNA was incubated in a 10 ml reaction containing1 � AB (see above), 10 mg Torula yeast RNA and RNase


T2 (Life Technologies). RNase T2 activity was inef®cientunder our reaction conditions at 25 �C, and so was usedwithout dilution (23 units). Prior to addition of RNaseT2, RNAs were incubated in 1 � AB (ten minutes, 25 �C).RNase T2 reactions were incubated for 20 minutes, thenstopped by adding 10 ml of 2 � formamide loading buf-fer and placing tubes on dry ice. Alkaline hydrolysis of45,000 cpm RNA and 10 mg Torula yeast RNA wasperformed as described (Polson & Bass, 1994). Sampleswere heated (90 �C, three minutes), then loaded onto a12 % (w/v) polyacrylamide, 8 M urea, 1 � TBE gel(43 cm long) prior to electrophoresis (50 W, one hour 45minutes). Gels were dried and subjected to autoradiog-raphy and Phosphorimager analysis.

Acknowledgments

We thank members of our research group for helpfuladvice and discussions, and Mark Macbeth and BrianWimberly for critically reading this manuscript. Wethank Ron Hough for providing puri®ed xADAR1, andEd Meenan for the synthesis of DNA oligos. Oligonu-cleotides were synthesized by the Howard HughesMedical Institute oligonucleotide synthesis facility at theUniversity of Utah supported by the Department ofEnergy (Grant # DE-FG03-94ER61817). This work wassupported by funds to B.L.B from the National Instituteof General Medical Sciences (GM44073). B.L.B. is anHHMI Associate Investigator.

References

Bass, B. L. (1997). RNA editing and hypermutation byadenosine deamination. Trends Biochem. Sci. 22, 157-162.

Bass, B. L. & Weintraub, H. (1987). A developmentallyregulated activity that unwinds RNA duplexes. Cell,48, 607-613.

Bass, B. L. & Weintraub, H. (1988). An unwindingactivity that covalently modi®es its double-strandedRNA substrate. Cell, 55, 1089-1098.

Bevilacqua, P. C. & Cech, T. R. (1996). Minor-groove rec-ognition of double-stranded RNA by the double-stranded RNA-binding domain from the RNA-acti-vated protein kinase PKR. Biochemistry, 35, 9983-9994.

Burns, C. M., Chu, H., Rueter, S. M., Hutchinson, L. K.,Canton, H., Sanders-Bush, E. & Emeson, R. B.(1997). Regulation of serotonin-2C receptor G-pro-tein coupling by RNA editing. Nature, 387, 303-308.

Cai, Z. & Tinoco, I., Jr (1996). Solution structure of loopA from the hairpin ribozyme from tobacco ringspotvirus satellite. Biochemistry, 35, 6026-6036.

Chelladurai, B., Li, H., Zhang, K. & Nicholson, A. W.(1993). Mutational analysis of a ribonuclease III pro-cessing signal. Biochemistry, 32, 7549-7558.

Clemens, M. J. & Elia, A. (1997). The double-strandedRNA-dependent protein kinase PKR: structure andfunction. J. Interferon Cytokine Res. 17, 503-524.

Correll, C. C., Freeborn, B., Moore, P. B. & Steitz, T. A.(1997). Metals, motifs, and recognition in the crystalstructure of a 5S rRNA domain. Cell, 91, 705-712.

Court, D. (1993). RNA processing and degradation byRNase III. In Control of Messenger RNA Stability(Belasco, J. G. & Braverman, G., eds), pp. 71-116,Academic Press, San Diego.

Dabiri, G. A., Lai, F., Drakas, R. A. & Nishikura, K.(1996). Editing of the GluR-B ion channel RNAin vitro by recombinant double-stranded RNA ade-nosine deaminase. EMBO J. 15, 34-45.

Egebjerg, J. & Heinemann, S. F. (1993). Ca2� per-meability of unedited and edited versions of thekainate selective glutamate receptor GluR6. Proc.Natl Acad. Sci. USA, 90, 755-759.

Herb, A., Higuchi, M., Sprengel, R. & Seeburg, P. H.(1996). Q/R site editing in kainate receptor GluR5and GluR6 pre-mRNAs requires distant intronicsequences. Proc. Natl Acad. Sci. USA, 93, 1875-1880.

Higuchi, M., Single, F. N., Kohler, M., Sommer, B.,Sprengel, R. & Seeburg, P. H. (1993). RNA editingof AMPA receptor subunit GluR-B: a base-pairedintron-exon structure determines position and ef®-ciency. Cell, 75, 1361-1370.

Holbrook, S. R., Cheong, C., Tinoco, I., Jr & Kim, S. H.(1991). Crystal structure of an RNA double helixincorporating a track of non-Watson-Crick basepairs. Nature, 353, 579-581.

Hough, R. F. & Bass, B. L. (1994). Puri®cation of theXenopus laevis double-stranded RNA adenosine dea-minase. J. Biol. Chem. 269, 9933-9939.

Hurst, S. R., Hough, R. F., Aruscavage, P. J. & Bass, B. L.(1995). Deamination of mammalian glutamatereceptor RNA by Xenopus dsRNA adenosine deami-nase: similarities to in vivo RNA editing. RNA, 1,1051-1060.

Inoue, T. & Cech, T. R. (1985). Secondary structure ofthe circular form of the Tetrahymena rRNA inter-vening sequence: a technique for RNA structureanalysis using chemical probes and reverse tran-scriptase. Biochemistry, 82, 648-652.

Jang, S. B., Hung, L. W., Chi, Y. I., Holbrook, E. L.,Carter, R. J. & Holbrook, S. R. (1998). Structure ofan RNA internal loop consisting of tandem C-A�base pairs. Biochemistry, 37, 11726-11731.

Lomeli, H., Mosbacher, J., Melcher, T., Hoger, T.,Geiger, J. R., Kuner, T., Monyer, H., Higuchi, M.,Bach, A. & Seeburg, P. H. (1994). Control of kineticproperties of AMPA receptor channels by nuclearRNA editing. Science, 266, 1709-1713.

Maas, S., Melcher, T. & Seeburg, P. H. (1997). Mamma-lian RNA-dependent deaminases and editedmRNAs. Curr. Opin. Cell Biol. 9, 343-349.

Mathews, M. B. & Shenk, T. (1991). Adenovirus virus-associated RNA and translation control. J. Virol. 65,5657-5662.

McGhee, J. D. & von Hippel, P. H. (1974). Theoreticalaspects of DNA-protein interactions: co-operativeand non- co-operative binding of large ligands to aone-dimensional homogeneous lattice. J. Mol. Biol.86, 469-489.

Melcher, T., Maas, S., Herb, A., Sprengel, R., Seeburg,P. H. & Higuchi, M. (1996). A mammalian RNAediting enzyme. Nature, 379, 460-464.

Milligan, J. F., Groebe, D. R., Witherell, G. W. &Uhlenbeck, O. C. (1987). Oligoribonucleotide syn-thesis using T7 RNA polymerase and syntheticDNA templates. Nucl. Acids Res. 15, 8783-8798.

Nicholson, A. W. (1996). Structure, reactivity, andbiology of double-stranded RNA. Prog. Nucl. AcidRes. Mol. Biol. 52, 1-65.

Nishikura, K., Yoo, C., Kim, U., Murray, J. M., Estes,P. A., Cash, F. E. & Liebhaber, S. A. (1991). Sub-strate speci®city of the dsRNA unwinding/modify-ing activity. EMBO J. 10, 3523-3532.


O'Connell, M. A. (1997). RNA editing: rewriting recep-tors. Curr. Biol. 7, R437-439.

Paul, M. S. & Bass, B. L. (1998). Inosine exists in mRNAat tissue-speci®c levels and is most abundant inbrain mRNA. EMBO J. 17, 1120-1127.

Peritz, A. E., Kierzek, R., Sugimoto, N. & Turner, D. H.(1991). Thermodynamic study of internal loops inoligoribonucleotides: symmetric loops are morestable than asymmetric loops. Biochemistry, 30, 6428-6436.

Polson, A. G. & Bass, B. L. (1994). Preferential selectionof adenosines for modi®cation by double-strandedRNA adenosine deaminase. EMBO J. 13, 5701-5711.

Polson, A. G., Bass, B. L. & Casey, J. L. (1996). RNAediting of hepatitis delta virus antigenome bydsRNA-adenosine deaminase. Nature, 380, 454-456.

Robertson, H. D. (1982). Escherichia coli ribonuclease IIIcleavage sites. Cell, 30, 669-672.

Saccomanno, L. & Bass, B. L. (1994). The cytoplasm ofXenopus oocytes contains a factor that protectsdouble-stranded RNA from adenosine-to-inosinemodi®cation. Mol. Cell. Biol. 14, 5425-5432.

SantaLucia, J., Jr, Kierzek, R. & Turner, D. H. (1991).Stabilities of consecutive A. C, C. C, G. G, U. C,and U. U mismatches in RNA internal loops: evi-dence for stable hydrogen-bonded U.U and C.C.�pairs. Biochemistry, 30, 8242-8251.

Steyaert, J., Opsomer, C., Wyns, L. & Stanssens, P.(1991). Quantitative analysis of the contribution ofGlu46 and Asn98 to the guanosine speci®city ofribonuclease T1. Biochemistry, 30, 494-499.

Stroebel, S. A., Cech, T. R., Usman, N. & Beigelman, L.(1994). The 2,6-diaminopurine riboside-5-methyliso-cytidine wobble base pair: an isoenergetic substi-tution for the study of G-U pairs in RNA.Biochemistry, 33, 13824-13835.

Wagner, R. W. & Nishikura, K. (1988). Cell cycleexpression of RNA duplex unwindase activity inmammalian cells. Mol. Cell. Biol. 8, 770-777.

Weeks, K. M. & Crothers, D. M. (1993). Major grooveaccessibility of RNA. Science, 261, 1574-1577.

Wimberly, B., Varani, G. & Tinoco, I., Jr (1993). The con-formation of loop E of eukaryotic 5S ribosomalRNA. Biochemistry, 32, 1078-1087.

Xia, T., McDowell, J. A. & Turner, D. H. (1997). Thermo-dynamics of nonsymmetric tandem mismatchesadjacent to G.C base pairs in RNA. Biochemistry, 36,12486-12497.

Yang, J.-H., Sklar, P., Axel, R. & Maniatis, T. (1995).Editing of glutamate receptor subunit B pre-mRNAin vitro by site-speci®c deamination of adenosine.Nature 374, 77-81.

Zacharias, M. & Hagerman, P. J. (1996). The in¯uence ofsymmetric internal loops on the ¯exibility of RNA.J. Mol. Biol. 257, 276-289.

Edited by D. E. Draper

(Received 12 March 1999; received in revised form 25 May 1999; accepted 28 May 1999)

Documents

The Importance of Internal Loops within RNA Substrates of ... · The Importance of Internal Loops within RNA Substrates of ADAR1 Katrina A. Lehmann and Brenda L. Bass* Department