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Molecular determinants and guided evolution of species-specific RNA editing.
Author: Robert A. Reenan
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".............................................................. Molecular determinants and guided evolution of species-specific RNA editing Robert A. Reenan Department of Genetics and Developmental Biology, University of Connecticut Health Center, Farmington, Connecticut 06030, USA ............................................................................................................................................................................. Most RNA editing systems are mechanistically diverse, informa- tionally restorative, and scattershot in eukaryotic lineages 1 .In contrast, genetic recoding by adenosine-to-inosine RNA editing seems common in animals; usually, altering highly conserved or invariant coding positions in proteins 2?4 . Here I report striking variation between species in the recoding of synaptotagmin I (sytI). Fruitflies, mosquitoes and butterflies possess shared and species-specific sytIediting sites, all within a single exon. Honey- bees, beetles and roaches do not edit sytI. The editing machinery is usually directed to modify particular adenosines by infor- mation stored in intron-mediated RNA structures 5?7 . Combining comparative genomics of 34 species with mutational analysis revealsthat complex,multi-domain, pre-mRNA structures solely determine species-appropriate RNA editing. One of these is a previously unreported long-range pseudoknot. I show that small changes to intronic sequences, far removed from an editing site, can transfer the species specificity of editing between RNA substrates. Taken together, these data support a phylogeny of sytI gene editing spanning more than 250 million years of hexapod evolution. The results also provide models for the genesis of RNA editing sites through the stepwise addition of structural domains, or by short walks through sequence space from ancestral structures. RNA editing systems programmatically alter messenger RNA sequences after transcription from genomic templates and are found enigmatically scattered among phyla. One example, base modification through the hydrolytic deamination of adenosine to inosine (A-to-I) by ADARs (for ?adenosine deaminases acting on RNAs?), can result in informational recoding: the ribosome inter- prets inosine as guanosine 8 . Curiously, this recoding occurs almost exclusively in gene products whose primary function is fast neuronal signalling 3 , in keeping with the observed neurological phenotypes of ADAR-deficient Caenorhabditis elegans, Drosophila and mice 9?11 . Further, human neurological disease has been associ- ated with altered gene recoding 12,13 . The biological consequence of ADAR action at a particular site can vary markedly between genes and between species. Certain mammalian ionotropic glutamate receptor (iGluR) genes recode a functionally critical glutamine (Q) codon to that of arginine (R). Studies involving animals genetically altered in this recoding event are revealing. For instance, the GluR-2 (Q/R) site must be edited at nearly 100% or mice developepilepsy and die postnatally 14 , whereaslackof editing at the GluR-6 (Q/R) site (normally about 75%) is relatively benign 15 , resulting in modest changes in synaptic plasticity and seizure vulnerability. C. elegans iGluR genesdo not editthe same conserved amino acid; however, when an R-encoding version is introduced into worms, neurotoxicity and lethality ensue 16 . Previous reports indicate that A-to-I RNA editing varies between arthropod species 3,17,18 . Although the mechanism of gene recoding frequently involves imperfect base pairing of exonic and intronic sequences 5?7,19 , the molecular basis of species-specific editing is unknown. Drosophila synaptotagmin I (dsytI), the Ca 2� sensor for synchronous neurotransmitter release 20 , is a target of A-to-I RNA editing 3 . One exon, whose boundaries are conserved in all synapto- tagmins 21 , was shown to possess four editing sites (A to D) that recode highly conserved positions. To investigate whether editing varied between species, I performed reverse-transcriptase polymer- ase chain reaction (RT?PCR) to obtain sytI complementary DNAs from Anopheles gambiae (malaria mosquito), Manduca sexta (tobacco hawkmoth), Apis mellifera (honeybee), Tribolium casta- neum (red flour beetle) and Blattella germanica (German cock- roach). Direct sequenceanalysis ofRT?PCR amplification products was performed to identify RNA editing sites, as described pre- viously 3 . Editing was not detectable in A. mellifera, T. castaneum or B. germanica sytI genes. The remaining arthropods edited sytI, but no twospecies possessed the same set ofediting sites (Fig.1a, b). All Figure 1 Species-specific genetic recoding of synaptotagmin I. a, Left: ribbon structure of the synaptotagmin I C 2 B domain. Yellow corresponds to the conserved exon found in all sytI orthologues. Side chains of D. melanogaster sytI editing sites A?D (red), mosquito- specific site N (blue) and lepidopteran-specific site L (green) are indicated. Right: electrostatic surface potential showing surface-projecting residues of editing sites B, C, N and L (colours as in a). b, Cladogram of taxa in this study and editing sites present: Diptera (Drosophilidae (Drosophila melanogaster), Culicidae (Anopheles gambiae)), Lepidoptera (Manduca sexta), Hymenoptera (Apis mellifera), Coleoptera (Tribolium castaneum) and Blattaria (Blattella germanica). Dashes indicate no detectable editing. c, Protein sequence alignment of synaptotagmin I editing exon orthologues of the animal kingdom. Shaded invariant (red) and conserved (blue) amino acid residues are shown. Editing sites A?D, L and N result in the following recoding events (single-letter amino acid codes): site A, I ! V; site B, K ! R; site C, I ! V; site D, I ! M; site L, T ! A; site N, K ! R. letters to nature NATURE|VOL 434|17 MARCH 2005|www.nature.com/nature 409 � 2005 Nature Publishing Group had a common editing site (D). In addition, D. melanogaster and A. gambiae shared a site (C). Finally, each species was found to possess species-specific editing sites (A, B, N and L). That species- specificity of RNA editing was a conserved feature among closely related species was confirmed byanalysing sytI editing in additional representatives within each taxonomic group (Supplementary Table S1). These editing sites in insects alter invariant or highly conserved residues within the SytI C2B domain (Fig. 1c), a Ca 2� -dependent phospholipid-binding machine essential for the rapid and synchro- nousreleaseofneurotransmitter.Threespecies-specificeditingsites (B, N and L) as well as shared editing site C recode amino acids positioned on an interaction surface that is crucial for proper SytI function 22?25 (Fig. 1a). Aplysia californica (sea hare) employs alternative splicing of the same exon, generating functionally distinct SytI isoforms 26 . A single amino acid difference accounted for differential function of these isoforms, corresponding to editing site L, and resulting in nearly the same amino acid change (Thr! Gly by alternative splicing, versus Thr ! Ala by RNA editing). Conservation of specific ADAR modification between related species can result in the simultaneous conservation of cis elements that direct RNA structure formation 17,27?29 . To test this rule for sytI, genomicsequenceswerenextclonedandsequencedforthegenomic region spanning the editing sites from ten members of the family Drosophilidae with estimated divergence times ranging from 15 million to 80 million years ago. Sequence alignment revealed two invariant intronic elements, E1 (33 nucleotides) and E2 (48 nucleotides) (Fig. 2a, Supplementary Fig. S1). The remaining intron sequences were highly divergent, although interelement spacing was a conserved feature. To determine whether the E1/E2-containing intron directs the RNA editing of Drosophila sytI, the sequences spanning the edited exon, the intron and the downstream exon were expressed as a minigene in Drosophila S2 cells together with Drosophila ADAR (dADAR).Efficientandspecificeditingwasobservedwiththeuseof a restriction-enzyme assay for sites C and D (see Methods), demonstrating the sufficiency of these sequences to direct ADAR modification (Fig. 2c, WT). Noediting wasobserved at inappropri- ate adenosines or in the absenceof co-transfected dADAR (data not shown). RNA structural computations with MFOLD consistently predicted two mutually exclusive lowest-energy structures: one pairing E1 with the upstream exonic region of editing sites B and C (domain I), and one pairing the downstream distal E2 with the exonic region of editing site D (domain II). The assumption of a pseudoknot structure reconciled these two duplexes in one struc- ture (Fig. 2b). To probe this hypothetical structure, potentially disruptive mutations were introduced into domains I and II (Fig. 2b, c). Mutations M1 and M2 singly abolished editing at site D, whereas editing at site C occurred normally. Likewise, mutations M3 and M4 singlyabolished editing at site C, whereas editing at site D was unaffected. Structurally compensatory double mutations M12 and M34 each restored editing in their respective domains, thus validating numerous predicted base-pair interactions. Because Figure 3 Structure of lepidopteran sytI editing site and heterologous editing by dADAR. a, Predicted structure of lepidopteran sytI pre-RNA. Exon (black), intron (blue) and variant positions (red) are indicated (see Supplementary Fig. S3). Variation in loop sequence length is indicated (mean ^ s.d). Intronic mutations are indicated (LM1 and LM2). b, Electropherograms of M. sexta sytI sites L (left) and D (right). Sequences were generated from genomic DNA PCR products (top), RT?PCR products from M. sexta brain RNA (middle) and RT?PCR products from M. sexta minigene expressed in Drosophila S2 cells with dADAR (bottom). c, Mutations LM1 and LM2 (as in a) were introduced into the M. sexta minigene and expressed in S2 cells with dADAR. Values are means ^ s.d. Figure 2 Drosophila sytI pre-mRNA forms a pseudoknot. a, Genomic organization of Drosophilid sytI gene editing sites. Exons (blue boxes), conserved elements E1 and E2 (yellow) and intron (line) shown with spacings as indicated (means ^ s.d.). b, Predicted pseudoknot domain structure of dsytI pre-mRNA. Exon (black), intron (blue), and editing site C (red) and D (green) sequences are indicated. Mutations introduced into domains I and II are indicated above or below mutated sequences (M1?M4). c, Effects of mutations on the editing of sites C (red) and D (green) are indicated for disruptive single mutations (M1?M4) and compensatory double mutations (M12 and M34). WT, wild type. Values are means ^ s.d. letters to nature NATURE|VOL 434|17 MARCH 2005|www.nature.com/nature410 � 2005 Nature Publishing Group editing in each domain of the dsytI structure can be disrupted independently,itseemsunlikely that dsytI editing proceeds through a stepwise mechanism invoking sequential editing. Rather, both duplex regions probably exist in the pre-mRNA, forming a long- range pseudoknot containing two domains of ADAR action. Lepidopterans modify sytI at site D, as well as a Lepidoptera- specificsite(L).GenomicDNAsequencesspanningthesameregion studied in Drosophila species wereobtainedfor tenspecies of moths and butterflies. Comparison revealed highly conserved intronic sequences downstream of the editing sites, comprising a single extended region with limited sequence variation (Supplementary Fig.S2). Totest whether these sequencesdirectediting, M. sexta sytI genomic sequences encompassing the edited exon, downstream intron and downstream exon were expressed as a minigene in Drosophila S2 cells along with dADAR. Efficient and specific editing wasobserved at M. sexta sites Dand L(Fig.3b).The M. sexta intron is clearly sufficient to direct species-specific editing with a heter- ologous editing enzyme. Thus, little of the species-specificityof syt I editing must be due to differences in ADAR enzymes between species. Structural predictions for the M. sexta sytI pre-mRNA consist- ently paired conservedintronic sequenceswiththe region of editing sites D and L (Fig. 3a). Although the intronic conserved elements varied in sequence between moth and butterfly species, none of the differences altered the predicted secondary structure of the RNA (Fig. 3, Supplementary Fig. S3). Like dsytI, the lepidopteran substrate contains two duplex domains, each with a site of adeno- sine modification. Because dADAR efficiently and accurately edits the M. sexta minigene in Drosophila cells, the predicted structure was tested by mutation (Fig. 3a, c). Two mutations were engineered into intronic conserved sequences (LM1 and LM2). The LM1 mutation decreased editing at site L but editing at site D was relatively unaffected. Mutation LM2 disrupted editing at nearby site D but editing at site L occurred at nearly wild-type levels (Fig.3c).Thus, sytI RNAeditinginLepidopteraoccursthroughtwo mutationally separable domains of ADAR action whose overall structural arrangement is substantially different from that seen in Drosophilidae. Anopheles gambiae and Aedes aegypti sytI share editing sites with Drosophila species (C and D), whereas mosquito-specific site N occurs between sites C and D (Fig. 1c). Because the editing of Drosophila sytI at sites C and D is directed by a conserved pseudoknot structure, a comparative sequence analysis of five mosquito species was performed to identify conserved intronic sequences (Supplementary Fig. S4). Like Drosophila, mosquitoes possess distinct E1-like and E2-like elements that are predicted to form a homologous pseudoknot structure, with some alterations. Many of these differences are structurally silent; however, key changesareobservedinmosquitodomainIinthevicinityofediting site C, including the extensive disruption of base pairing near editing site B, which is not edited in mosquitoes (Fig. 4a and Supplementary Fig. S5). Expression of the Anopheles gambiae sytI Figure 4 Guided evolution of species-specific RNA editing. a, Structure of pseudoknot domain I showing exon (black) and intron (blue) sequences for Drosophila (D.m., left), Anopheles (A.g., middle) and MAG4 mutation (D.m. MAG4, right). Differences of Anopheles domain I from Drosophila are indicated in orange (middle). The MAG4 mutations (right) are also indicated in orange. Editing status is shown in electropherograms below structures. b, Phylogeny of sytI RNA editing. Extant synaptotagmin I pre-mRNA structures and proposed ancestor molecules are shown associated with a cladogram of ordinal relationships of taxa in this study 30 . Nodes denoting ancestral origins of particular editing events are indicated by circles. Unedited sytI mRNAs are depicted as unstructured (black). letters to nature NATURE|VOL 434|17 MARCH 2005|www.nature.com/nature 411 � 2005 Nature Publishing Group substrate in Drosophila cells along with dADAR resulted in the Anopheles pattern of editing, indicating that dADAR recognized the mosquito-specific editing site N (data not shown). To determine whether the editing of site N results from structural differences in domain I, two nucleotide changes were made in the Drosophila sytI expression construct in element E1, guided by the A. gambiae domainIstructure(MAG4;Fig.4a).The intronic MAG4mutations are located more than 1,200 nucleotides from editing site C. Whereas no editing of site N occurs in the wild-type dsytI construct, the MAG4 mutations confer efficient modification of the mosquito site N adenosine on the Drosophila minigene RNA. Thus, small non-coding changes, far removed from a potential target adenosine, are capable of inducing sufficient structural change to directly evolve a site of gene recoding by ADAR modification. The editing sites described here recode invariant or nearly invariant positions, a phenomenon seen in other targets of A-to-I editinginarthropods,molluscsandvertebrates.Together,thesedata imply a selective advantage for RNA editing by allowing protein sequences access to a mutational forbidden zone wherein histori- cally invariant amino acids can be altered bydegrees, in mRNA, but not through discrete genetic change in coding sequence. Access to this normally unattainable realm of protein space is mediated through complex RNA secondary structures under intense purify- ing selection. I suggest that the data presented here comprise a credible phylogeny of RNA editing for a gene, graphically illustrat- ing descent with modification (Fig. 4b). RNA editing in insect sytI first seems to have evolved in the common ancestor of dipteran and lepidopteran lineages, beginning with site D. Beetles, roaches and reported vertebrate sytI genes are genomically incapable of evolving Ile ! Met editing at the same location, for lack of a third-position adenosine in their isoleucine codons (ATTorATC). The ancestral editing site D structure probably generated new editing sites through two methods. One, global intronic variation, led to the synthesis of entirely new, add-on oligonucleotide domains, such as those that direct editing sites C and L in dipterans and lepidopter- ans,respectively(Figs2and3).Alternatively,siteswendedtheir way through sequence space, generating nascent editing sites by means of a small number of changes to ancestral structures, as shown by thenatureofsitesA,BandNandtheabilitytotransfereditingfrom the mosquito to fly pre-mRNA substrate readily by simple mutation.Ofcourse,species-specific sytI RNAeditingsites,directed by different structures from those presented here, might exist in other animals. The evolutionary methods ofcreatingediting sites proposedhere have probably shaped most targets of ADAR-mediated recoding. Further challenges posed by this study lie in determining what advantage particular editing sites confer on species, the extent of standing variation of RNA editing, and the role, if any, of RNA recoding in the process of speciation. A Methods Multiple sequence alignment and structural predictions Sequences were aligned with the GCG software package. Protein alignment shown in Fig. 1c was performed with the Pileup program using default settings. Sequences are abbreviated as follows: Mmu (Mus musculus), Hro (Halocynthia roretzi), Aga (Anopheles gambiae), Dme (Drosophila melanogaster), Ame (Apis mellifera), Mse (Manduca sexta), Tca (Tribolium castaneum), Dja (Dugesia japonica) and Lst (Lymnaea stagnalis). For Drosophila genomic DNA sequence alignments (Supplementary Fig. S1, Supplementary Table S1), alignments wereperformed on sequences spanning the editing exonthrough to element E2 because of large discrepancies in intron size downstream of E2. No significant conservedsequenceswereobtainedinalignmentsdownstreamofE2(datanotshown).All DNA sequence alignments were performed with the gap creation and gap extension penalties set to a value of 1. Structural predictions were performed using the MFOLD program of the Macfarlane Burnet Centre MFOLD server (http://mfold.burnet.edu.au/), setting the folding temperature to 258C. For Drosophila substrates, only the sequences from the editing exon as far as 100 nucleotides downstream of E2 were used. RNA editing analysis of endogenous synaptotagmins For flies, mosquitoes and beetles, whole-organism RNAs were isolated with TRI Reagent (Molecular Research Center). For moths, butterflies, bees and roaches, whole-head RNA was isolated. Species-specific primers were used for first-strand cDNA synthesis. PCR was then performed with species-specific sytI primers. Amplification products of the correct size were gel-purified and sequenced. The presence or absence of editing was assessed by the presenceofmixedA/Gsignalsintheelectropherograms andsingletAsignalfromPCR products from genomic DNA for each species. Schneider cell RNA-editing system and substrate mutagenesis RNA-editingreporterminigeneconstructsweregeneratedbycloningthe sytI editingexon, downstream intron and downstream exon into the pMT-V5/His vector (Invitrogen). dADARexpressionconstructwasgeneratedbycloningthe�1,23aalternativespliceform of dADAR into pMT-V5/His. Schneider S2 cells were transfected with various editing reporter constructs (about 200ng) with or without 2mg of dADAR expression construct. Cultures were transfected with DNAusing GeneJuice (Novagen) transfection reagent and induced with copper sulphate 5h after transfection. Cell were harvested 3 days after transfection and total RNAwas prepared with TRI Reagent. In all cases the RNA samples were treated with DNase (DNA-free; Ambion) to remove contaminating input DNA. SytI minigene transcripts were amplified by RT?PCR from S2-cell total mRNA with gene-specific and vector-specific primers. RNA editing was quantified as follows. RNA editing of dsytI creates a PshAI restriction enzyme cutting site at site C or a HpyCH4V cuttingsiteatsiteD. M. sexta RNAeditingcreatesa BcgI cleavagesiteateditingsiteLanda HpyCH4V cuttingsiteatsiteD.RNAsampleswerepreparedfromtwotofourindependent transfections for each construct. For each RNA sample quantified, three independent RT?PCRs wereperformed with different primer sets; the resulting products were digested with either PshAI, BcgI or HpyCH4V and subjected to electrophoresis on an agarose gel. The intensities of bands corresponding to edited and unedited products were quantified; band intensities were corrected for band size. Editing frequencies reported are means ^ s.d. All images were obtained on a Kodak Gel Logic 100 system and were taken under subsaturation conditions. Data were quantified with 1D v.3.6 image analysis software (Kodak). Mutations indicated in Figs 2?4 were introduced with PAGE-purified mutagenic primers (about 50 nucleotides in length) (IDT) using the QuikChange II XL Site-directed Mutagenesis Kit (Stratagene). Mutations were induced on the full-length editing constructs in pMT-V5/His. All mutagenized templates were subject to 12?14 rounds of amplification, and the resultant transformed mutants were subjected to sequence analysis of the entire insert to confirm the mutations and lack of secondary mutations. Received 25 October 2004; accepted 14 January 2005; doi:10.1038/nature03364. 1. Bass, B. L. (ed.) RNA Editing (Oxford University Press, Oxford, 2001). 2. Seeburg, P. & Hartner, J. Regulation of ion channel/neurotransmitter receptor function by RNA editing. Curr. Opin. Neurobiol. 13, 279?283 (2003). 3. Hoopengardner, B., Bhalla, T., Staber, C. & Reenan, R. Nervous system targets of RNA editing identified by comparative genomics. Science 301, 832?836 (2003). 4. Bhalla, T., Rosenthal, J. J., Holmgren, M. & Reenan, R. Control of human potassium channel inactivation by editing of a small mRNA hairpin. Nature Struct. Mol. Biol. 11, 950?956 (2004). 5. Herb, A., Higuchi, M., Sprengel, R. & Seeburg, P. Q/R site editing in kainate receptor GluR5 and GluR6pre-mRNAsrequiresdistantintronicsequences. Proc. Natl Acad. Sci. USA 93,1875?1880(1996). 6. Dawson, T., Sansam, C. & Emeson, R. Structure and sequence determinants required for the RNA editing of ADAR2 substrates. J. Biol. Chem. 279, 4941?4951 (2004). 7. Higuchi, M. et al. RNA editing of AMPA receptor subunit GluR-B: a base-paired intron-exon structure determines position and efficiency. Cell 75, 1361?1370 (1993). 8. Bass, B. L. RNA editing by adenosine deaminases that act on RNA. Annu. Rev. Biochem. 71, 817?846 (2002). 9. Tonkin,L.A. et al. RNAeditingbyADARsisimportantfornormalbehaviorin Caenorhabditis elegans. EMBO J. 21, 6025?6035 (2002). 10. Palladino, M. J., Keegan, L. P., O?Connell, M. A. & Reenan, R. A. A-to-I pre-mRNA editing in Drosophila is primarily involved in adult nervous system function and integrity. Cell 102, 437?449 (2000). 11. Higuchi, M. et al. Point mutation in an AMPA receptor gene rescues lethality in mice deficient in the RNA-editing enzyme ADAR2. Nature 406, 78?81 (2000). 12. Gurevich, I. et al. Altered editing of serotonin 2C receptor pre-mRNA in the prefrontal cortex of depressed suicide victims. Neuron 34, 349?356 (2002). 13. Kawahara, Y. et al. Glutamate receptors: RNA editing and death of motor neurons. Nature 427, 801 (2004). 14. Brusa,R. et al. Early-onsetepilepsyandpostnatallethalityassociatedwithanediting-deficientGluR-B allele in mice. Science 270, 1677?1680 (1995). 15. Vissel,B. et al. The roleof RNAeditingof kainatereceptorsin synapticplasticityand seizures. Neuron 29, 217?227 (2001). 16. Aronoff, R., Mellem, J. E., Maricq, A. V., Sprengel, R. & Seeburg, P. H. Neuronal toxicity in Caenorhabditis elegans from an editing site mutant in glutamate receptor channels. J. Neurosci. 24, 8135?8140 (2004). 17. Hanrahan,C.J.,Palladino,M.J.,Ganetzky,B.&Reenan,R.A.RNAeditingofthe Drosophila paraNa � channel transcript. Evolutionary conservation and developmental regulation. Genetics 155, 1149?1160 (2000). 18. Grauso, M., Reenan, R. A., Culetto, E. & Sattelle, D. B.Novelputative nicotinic acetylcholinereceptor subunit genes, Da5, Da6andDa7, in Drosophila melanogaster identify a new and highly conserved target of adenosine deaminase acting on RNA-mediated A-to-I pre-mRNA editing. Genetics 160, 1519?1533 (2002). 19. Lehmann, K. A. & Bass, B. L. The importance of internal loops within RNA substrates of ADAR1. J. Mol. Biol. 291, 1?13 (1999). 20. Yoshihara, M. & Littleton, J. T. Synaptotagmin I functions as a calcium sensor to synchronize neurotransmitter release. Neuron 36, 897?908 (2002). letters to nature NATURE|VOL 434|17 MARCH 2005|www.nature.com/nature412 � 2005 Nature Publishing Group 21. Craxton, M. Genomic analysis of synaptotagmin genes. Genomics 77, 43?49 (2001). 22. Mackler, J. M. & Reist, N. E. Mutations in the second C2 domain of synaptotagmin disrupt synaptic transmission at Drosophila neuromuscular junctions. J. Comp. Neurol. 436, 4?16 (2001). 23. Chapman, E. R., Desai, R. C., Davis, A. F. & Tornehl, C. K. Delineation of the oligomerization, AP-2 binding, and synprint binding region of the C2B domain of synaptotagmin. J. Biol. Chem. 273, 32966?32972 (1998). 24. Rickman, C. et al. Synaptotagmin interaction with the syntaxin/SNAP-25 dimer is mediated by an evolutionarily conserved motif and is sensitive to inositol hexakisphosphate. J. Biol. Chem. 279, 12574?12579 (2004). 25. Grass, I., Thiel, S., Honing, S. & Haucke, V. Recognition of a basic AP-2 binding motif within the C2B domain of synaptotagmin is dependent on multimerization. J. Biol. Chem. 279, 54872?54880 (2004). 26. Nakhost, A., Houeland, G., Blandford, V. E., Castellucci, V. F. & Sossin, W. S. Identification and characterization of a novel C2B splice variant of synaptotagmin I. J. Neurochem. 89, 354?363 (2004). 27. Reenan,R.,Hanrahan,C.&Ganetzky,B.Themle(napts)RNAhelicasemutationin Drosophila results in a splicing catastrophe of the para Na � channel transcript in a region of RNA editing. Neuron 25, 139?149 (2000). 28. Aruscavage, P. & Bass, B. A phylogenetic analysis reveals an unusual sequence conservation within introns involved in RNA editing. RNA 6, 257?269 (2000). 29. Kung,S.S.,Chen,Y.C.,Lin,W.H.,Chen,C.C.&Chow,W.Y.Q/RRNAeditingoftheAMPAreceptor subunit 2 (GRIA2) transcript evolves no later than the appearance of cartilaginous fishes. FEBS Lett. 509, 277?281 (2001). 30. Wheeler,W.C.,Whiting,M.,Wheeler,Q.D.&Carpernter,J.M. Thephylogenyof theextanthexapod orders. Cladistics 17, 113?169 (2001). Supplementary Information accompanies the paper on www.nature.com/nature. Acknowledgements I thank L. Reenan for discussions; B. Hoopengardner, T. Bhalla and A. Das forcommentsonthemanuscript;B.Hoopengardner forsharingcertaingenomicDNAtemplates andforassistancewithS2cellculture;UCHCMolecularCoreFacilitystafffordiligentsequencing efforts; and M. Lalande for his encouragement. This work was supported by grants from the National Science Foundation and National Institutes of Health (R.A.R.). Competing interests statement The author declares that he has no competing financial interests. Correspondence and requests for materials should be addressed to R.A.R. (rreenan@neuron.uchc.edu). ............................................................................................................................... erratum Pleistocene to Holocene extinction dynamics in giant deer and woolly mammoth A. J. Stuart, P. A. Kosintsev, T. F. G. Higham & A. M. Lister Nature 431, 684?689 (2004). ................................................................................................................................................................................................................................................................................................................................................................... In Fig. 4 of this Letter, some of the data were not properly aligned with their location labels. The corrected figure is shown here. A letters to nature NATURE|VOL 434|17 MARCH 2005|www.nature.com/nature 413 � 2005 Nature Publishing Group "
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