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Characteristics of the nuclear (18S, 5.8S, 28S and 5S) and mitochondrial (12S and 16S) rRNA genes of Apis mellifera (Insecta: Hymenoptera): structure, organization, and retrotransposable elements

J. J. Gillespie, J. S. Johnston, J. J. Cannone and 1 more

Insect Molecular Biology · 2006

doi:10.1111/j.1365-2583.2006.00689.x

Abstract

As an accompanying manuscript to the release of the honey bee genome, we report the entire sequence of the nuclear (18S, 5.8S, 28S and 5S) and mitochondrial (12S and 16S) ribosomal RNA (rRNA)‐encoding gene sequences (rDNA) and related internally and externally transcribed spacer regions of Apis mellifera (Insecta: Hymenoptera: Apocrita). Additionally, we predict secondary structures for the mature rRNA molecules based on comparative sequence analyses with other arthropod taxa and reference to recently published crystal structures of the ribosome. In general, the structures of honey bee rRNAs are in agreement with previously predicted rRNA models from other arthropods in core regions of the rRNA, with little additional expansion in non‐conserved regions. Our multiple sequence alignments are made available on several public databases and provide a preliminary establishment of a global structural model of all rRNAs from the insects. Additionally, we provide conserved stretches of sequences flanking the rDNA cistrons that comprise the externally transcribed spacer regions (ETS) and part of the intergenic spacer region (IGS), including several repetitive motifs. Finally, we report the occurrence of retrotransposition in the nuclear large subunit rDNA, as R2 elements are present in the usual insertion points found in other arthropods. Interestingly, functional R1 elements usually present in the genomes of insects were not detected in the honey bee rRNA genes. The reverse transcriptase products of the R2 elements are deduced from their putative open reading frames and structurally aligned with those from another hymenopteran insect, the jewel wasp Nasonia (Pteromalidae). Stretches of conserved amino acids shared between Apis and Nasonia are illustrated and serve as potential sites for primer design, as target amplicons within these R2 elements may serve as novel phylogenetic markers for Hymenoptera. Given the impending completion of the sequencing of the Nasonia genome, we expect our report eventually to shed light on the evolution of the hymenopteran genome within higher insects, particularly regarding the relative maintenance of conserved rDNA genes, related variable spacer regions and retrotransposable elements.

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Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 20, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction

Ribosomal RNA (rRNA)-encoding genes (rDNA) and related genetic elements have been well studied for over six decades (see for a recent review), with interests ranging from pharmaceutical and biochemical investigations to comparative biological studies garnering a wealth of information on the structural, functional and evolutionary characteristics of these molecules. Phylogenetic studies, in particular, have propagated a large number of rRNA gene sequences on public genetic databases, as the organismal universality and typically high gene copy number per cell facilitate gene amplification and sequencing (e.g.;; ). Within rRNA genes, varying evolutionary rates of base substitution can be localized to conserved core rRNA sequences (e.g.;;;;;; ) as well as rapidly evolving variable regions and expansion segments (e.g.;,;;;; ). Rates of substitution are even faster in transcribed spacer regions and non-coding spacer regions, which can reveal high levels of divergence even when compared across closely related species (e.g.;; ), as well as within individual genomes (e.g.;;; ). These conserved and variable regions of rRNA genes have not only been useful for recovering phylogenetic relationships (e.g.;,), but also for understanding the structural and functional constraints of both the coding and the non-coding organizational sequences within rRNA genes (see ).

Recent crystal structures of the ribosome have verified many of the long-standing structures of the rRNAs that were predicted by comparative sequence analysis or substantiated with site-directed or random mutagenesis (e.g.,;; ). Still, these three-dimensional models have yielded further information on the organization of the small and large subunits (SSU and LSU, respectively) with the numerous ribosomal proteins comprising the mature ribosome. In particular, new information on enzymatic function (e.g. ), non-canonical base pairing (e.g.;; ), tertiary interactions via coaxial stacking (e.g.; ), RNA turn motifs (e.g.; ), rRNA–protein interactions (e.g.;;; ), metal ions and rRNA–rRNA packing interactions, lone pair triloops and dynamism has been elicited from observations of tertiary models not easily obtained from comparative sequence analysis. It is essential that these characteristics be incorporated into multiple sequence alignments to improve the assignment of structurally and functionally homologous bases, as well as to verify that generated sequences are indeed functional rRNA genes and not paralogues and/or pseudogenes (e.g.;;; ) or artefacts of the sequencing process (see;;; ).

In this study, we predict entire secondary and some tertiary structures of the nuclear (nl) and mitochondrial (mt) SSU and LSU rRNAs from the honey bee, Apis mellifera, as the recent compilation of its genome has provided sequences spanning the entire rDNA region. Wherever possible, we include information from published crystal structures of the ribosome and chemical probing studies to annotate and update diagrams of arthropod nl and mt rRNA structure. These diagrams (and related multiple sequence alignments) will be valuable for related studies on the phylogeny of arthropods, as well as the evolution of the structure and function of eukaryotic and organellar rRNAs.

With the completion of the honey bee genome comes the opportunity to begin analysing the intragenomic variation associated with repetitive sequence motifs, particularly those associated with multicopy genes. Here, we make an attempt to characterize the sequences associated with the externally transcribed spacer regions (ETS) and intergenic spacers (IGS) that are conserved across all rDNA cistrons in the honey bee genome. We describe several repeat regions that occur in the 5′- and 3′-ends of the IGS regions. Our preliminary mapping of these complete ETS sequences and partial IGS sequences will be useful for future studies that attempt to span the further repetitive motifs of the honey bee rRNA genes that occur in the central portion of the IGS, including promoter and enhancer regions.

As a result of the compilation of all rRNA genes in the honey bee genome, we were able to assess the degree of rRNA gene inactivation by the insertion of retrotransposable elements. Two types of retrotransposable elements, R1 and R2, can disrupt the complete transcription of arthropod rRNA genes by inserting into conserved sequences of the nl LSU rDNA (e.g.;;,;,; ). Our analysis of R1 and R2 element retrotransposition in the honey bee rRNA genes is somewhat consistent with other studies on arthropod retrotransposition, and we provide here the predicted open reading frame (ORF) of an R2 element retrotransposase (RT).

Predicted rRNA secondary structures

Our results are presented within the typical organization and pattern of transcription of eukaryotic rRNA genes. A schematic diagram illustrates the composition of typical arthropod rRNA genes within the nuclear and mitochondrial genomes (Fig. 1). Within the honey bee genome, the nl rRNA gene arrays have been identified as occurring on two different chromosomes subsequently localized to linkage groups 6 and 12 (Aquino-Perez, unpublished data), while the mt rRNA genes are organized in the ancestral arthropod position (see; ) within the mitochondrial genome. In insects, pioneering structural studies of rRNA molecules were conducted on flies (Diptera) and one beetle (Coleoptera). While these studies predicted structures mostly from comparisons with other early published complete rDNA sequences from bacteria, yeast, frog, rodent, cow, human and other mammals, they were lacking a larger collection of arthropod sequences to make thorough comparisons of most of the variable regions outside of the core rRNA [although see the early structural model of the mt LSU rRNA of the bird spider]. Thus, the following sections review subsequent structural studies that were performed with comparisons across several to many arthropod/insect rDNA sequences.

Nuclear SSU rRNA

This model (Fig. 2A) is in concordance with the conserved 16S (prokaryote) and 16S-like (eukaryote) structures of the SSU rRNA from the literature and is based on an updated model of Drosophila melanogaster. Aside from D. melanogaster, structural models of five insect taxa have been published, including Tenebrio molitor (Coleoptera), Acyrthosiphon pisum (Heteroptera), peloridiid Hemiptera, Loricera foveata (Coleoptera) and ichneumonoid Hymenoptera. Our model is based loosely on these earlier models, and relies more heavily on the structure predictions from three recent studies wherein the 18S rRNA secondary structure was predicted across most insect orders as well as evidence from SSU rRNA crystal structures. For referencing purposes, our new model follows the convention (variable region nomenclature, helix numbering, etc.) put forth by ). The SSU rRNA contains 65 helices that are conserved across arthropods (Fig. 2A).

The predicted structure of most regions of the 18S rRNA from honey bee are consistent with structures of the major lineages of Arthropoda, and differences between the nl SSU rRNA model of ) and that presented here are illustrated on the jRNA website (see Experimental procedures). Variable region four (V4) is shown with refinements to the double pseudoknot model of, such that their proposed base triple is removed in place of the recently predicted tertiary interaction between the hairpin loop of the second pseudoknot and an unpaired sequence in variable region 2 (V2). Our alignment across arthropod V4 sequences spanning most orders shows strong support for the internal seven base pairs of this helix (HV2–V4); however, base pairs 1 and 9 are minimally supported with covariation evidence across arthropods (CRW Site bp frequency tables; ).

Accurate structural models of arthropod V4 are of particular interest because this region of 18S rRNA is the most commonly sequenced marker for higher level arthropod phylogeny estimation (reviewed in ). Still, multiple structural predictions exist for this large variable region of SSU rRNA. For instance, proposed a secondary structure model for insect V4 based on the comparative analysis of 22 insect sequences. That same year, presented a refined model with the 3′-half of this variable region consisting of two pseudoknots. One of the pseudoknots was previously predicted, while the other, a rare type of pseudoknot similar to that found in plant viruses, was newly reported. Within these two contrasting models of the 3′-half of arthropod V4, 32 nucleotides form different base pairs (Fig. 3A). Six nucleotides form the same base pairs but within different structural contexts (grey dashed lines in Fig. 3A). Interestingly, across most eukaryotes this region alternates the number of base pairs within helices E23-13 and E23-14. Our analysis of honey bee (and most arthropod) 18S rRNA sequences is consistent with an expanded helix E23-13 (6–11 bp) and a contracted helix E23-14 (11–3 bp) (Fig. 3B). A base pair frequency table (available at the jRNA website) suggests that the model of, including HV2–V4, is more probable based on more covariation at the positions in the proposed base pairs; however, a minimal amount of compensatory base change evidence supports the model such that a dynamic nature of this region of nl SSU rRNA cannot be disproved. These findings are consistent with a recent comparison of the and, ) models in the V2 region across most insect lineages (; see table 3).

Of the remaining regions of insect 18S rRNA, domain I (including variable regions V1–V3) is sequenced less frequently than domain III (including variable regions V6–V9). Variable regions V1, V3, V6 and V8 (and V5 of domain II) are considerably smaller in size than V4 and yield little phylogenetic information. The V2 region is difficult to align with confidence due to length variation. Studies on arthropod V7 have divulged a region characteristic of extraordinary variation in the length of helices. Sequence alignments of V7 across divergent insect taxa contain many variable positions, yet structure is necessary to guide the alignment. This is because, while only one helix is present in nearly all non-holometabolous orders, as well as in Amphiesmenoptera (Lepidoptera + Trichoptera), the remaining seven (minus Thysanoptera) holometabolous orders contain a second helix within V7. Additionally, this smaller less-conserved helix, H1118b-2, can be present or absent within these orders, with its base pairs often containing minimal comparative support. Finally, sometimes it is difficult to predict base pairing in the V9 at the end of the penultimate helix because many of the published sequences are missing the extreme 3′-end of the 18S rRNA.

Nuclear LSU rRNA

This model is in concordance with the conserved 23S (prokaryote) and 23S-like (eukaryote) structures of the LSU rRNA from the literature. With existing predicted structure models for Aedes albopictus, D. melanogaster (; others therein), Acyrthosiphon pisum, Tenebrio sp. and ichneumonoid Hymenoptera, this is the sixth predicted structure of a complete or nearly complete insect nl LSU rRNA (Fig. 2B,C). To date, complete comparative models (alignments) for insect 28S rRNA have not been published, so we present our model following the nearly complete model of ichneumonoid Hymenoptera with adherence to LSU rRNA crystal structures. Eukaryotic nl LSU rRNA is interrupted by an internally transcribed spacer region (ITS-2) in the terminal loop of helix H150 that separates the LSU into 5.8S and 28S rRNAs. Flies (Diptera) have a second ITS in the nl LSU that occurs within the terminal loop of helix H131. The A. mellifera LSU rRNA (minus 5S rRNA) structure model contains 143 helices (133 in the 28S rRNA and five in the 5.8S rRNA, with five helices comprised of both 28S and 5.8S rRNAs) that are conserved across arthropods (Fig. 2B,C).

Thirteen regions previously described as extremely variable or expansion segments of the 28S rRNA sequence are difficult to align across the arthropods (e.g. ). These regions of honey bee rRNA have the typical variation seen across insect orders and families. Regarding Hymenoptera, the structures for two of the largest of these regions, expansion segments D2 and D3, were recently predicted in studies of 349 ichneumonoid wasps and are comparable to the model predicted across 527 chalcidoid wasps and 60 evanioid wasps. Two other studies have used secondary structure as an alignment guide for phylogeny estimation of hymenopteran D2 sequences but did not produce global structural models. While expansion segment D2 is phylogenetically informative within orders and families of arthropods (for a comparison of our model with Coleoptera see ), the alignment of these sequences across highly divergent taxa (i.e. class and phylum levels) is extraordinarily difficult (see ). However, the expansion segment D3 is conserved enough in its secondary structure to be of practical use in arthropod phylogeny estimation (e.g.;; ), although its phylogenetic signal is weak when analysed without other data partitions (see ).

As expansion segments D2 and D3 of the 28S rRNA can be amplified in one PCR reaction, and because they usually contain characteristic variation in sequence length and base composition that is informative across family, genus and species levels, this region of nl LSU rRNA has become one of the most common phylogenetic markers (e.g.; ). Thus, the majority of published insect nl LSU rRNA sequences are comprised of expansion segments D2 and D3 and related core sequences. Some insect phylogenetic studies have generated sequences for expansion segments D4 to D10 (e.g.;; ); however, in-depth structural predictions with reference to previous models were not provided. This lack of structural information from these studies, coupled with limited sequences covering the majority of the 3′-half of the 28S rRNA, makes it difficult to propose models that are thoroughly supported with base pair covariation. Therefore, our proposed structures for this region of the nl LSU rRNA, particularly expansion segments D7, D10, D12 and R2832, should not be considered well supported. Our labs are currently generating more sequences from the 3′-half of the 28S rRNA from a wide range of insect taxa that should improve these preliminary structure predications.

Sequences beyond the D3 in domain II include expansion segments D4–D6, which are relatively small for candidate phylogenetic markers (Fig. 2B). Earlier predicted structures for these regions in insects have recently been modified based on a more thorough taxon sampling. In domain IV, a notable expansion segment, D7a, has been identified as the region of metazoan nl LSU rRNA wherein an internally processed cleavage site termed the ‘hidden break’ occurs. As originally characterized in the dipteran Sciara coprophila, this cleavage site separates the 28S rRNA into α and β fragments of roughly equal size. Aside from pea aphid (Acyrthosiphon pisum) 28S rRNA; which does not separate into α and β fragments, most insects contain this hidden break (e.g.; ), which was speculated to occur near an unpaired 5′-UAAU-3′ sequence within expansion segment D7. Despite this prevalent pattern, the proposed cleavage signal is absent in many hymenopteran 28S rDNA sequences, including the honey bee. However, it is probable that this region of the D7a contains a cleavage site in the 28S rRNA, given that the loop formed by helix D7a-3 is extraordinarily variable in sequence length and base composition and contains no detectable conserved secondary structural elements across arthropods (e.g. ) or other metazoan taxa studied (e.g.; ). Also, some insect D7a sequences contain repeated microsatellite motifs that are indicative of slippage events during replication of the rRNA gene array, further suggesting that this portion of the 28S rRNA is non-functional and likely cleaved out of the mature rRNA molecule. This observation is consistent with site-directed mutagenesis studies on the yeast Saccharomyces cerevisiae, which show that the D7a variable region of the 26S LSU rRNA is dispensable. Recently, provided evidence for the possible involvement of a highly conserved eukaryotic processing machinery responsible for cleavage of the α and β fragments in LSU rRNA, as microinjected Sciara coprophila (fungus fly) rDNA into Xenopus laevis oocytes resulted in transcription and pre28S rRNA fragmentation. Still, without a characterized processing machinery, an autocatalytic nature of the cleavage of cytoplasmic LSU rRNA cannot be ruled out.

Nearly half of domain IV of arthropod 28S rRNA is comprised of expansion segment D8 (Fig. 2C). Site-directed mutagenesis studies have implicated the D8 region with ribosome function, and it is likely that small nucleolar RNA E2 interacts with D8 in the human LSU rRNA. ) have characterized the structure of expansion segment D8 across aculeate Hymenoptera, and our analyses of more diverse taxa support their predicted structure. Domain V of arthropod 28S rRNA contains expansion segments D9, which is small and highly conserved in many Hymenoptera, including the honey bee, and D10, which is larger and highly variable in sequence composition but conserved in secondary structure. As more sequences accumulate on genetic databases, it is likely that expansion segment D10 will become a more commonly used marker for phylogeny estimation, especially because it can be amplified in some arthropod taxa with all of expansion segment D12 (Gillespie, unpublished data).

Expansion segment D11 is positioned between domains V and VI in cytoplasmic LSU rRNA (Fig. 4C) and is typically short and without base pairing. The remaining variable regions of arthropod 28S rRNA are expansion segments D12 and R2832, for which little information on base pairing can be derived given the paucity of published arthropod sequences spanning these regions. We found minimal compensatory base change support for the four helices within expansion segment D12, and the remainder of the structured bases within this region should be considered preliminary at best. Similarly, despite having a complete 28S rRNA sequence for the honey bee, our prediction of variable region R2832 is supported minimally because most ‘complete’ published arthropod rDNA sequences are nearly complete, having the 3′-primer designed from the D. melanogaster sequence (it is no coincidence that the majority of published arthropod sequences containing complete or nearly complete R2832 sequences are flies). The difficulty in sequencing the entire 3′-half of the 28S rDNA is a consequence of the extreme variability within the IGS region that immediately flanks the last nucleotides of the 28S rDNA (see below).

The nuclear-encoded cytoplasmic large subunit of the ribosome contains a 5S rRNA in Archaea, Bacteria and Eukaryota, and while chloroplast and plant mitochondria also have a 5S rRNA in their large ribosomal subunit, the mitochondria in fungi, animals and many protists do not. Our predicted structure of the nuclear-encoded honey bee 5S rRNA (Fig. 2D) is similar to other published structures of arthropod 5S rRNA (e.g.; ), and we confirmed our model with a recent interpretation of the crystal structure of the LSU rRNA of the halophile archaea Haloarcula marismortui. We have no supporting evidence that this 5S rRNA, or an additional 5S rRNA encoded in the nucleus, is imported into mitochondria; however, evidence suggests that nuclear-encoded 5S rRNA is a constituent of mt ribosomes in several eukaryotes, including trypanostomatids, yeast and mammals.

Mitochondrial SSU rRNA

In most insects, the 12S rRNA gene is flanked on the 5′-side by the non-coding portion of the mitochondrial genome that contains the control region and on the 3′-side by the valine transfer RNA (tRNA-Val). The honey bee 12S rRNA gene follows this conserved arrangement of the rRNA genes, despite seven gene rearrangements within the mitochondrial genome. Our structural model (Fig. 4A) follows the Drosophila virilis model of with minor modifications made based on an alignment of 83 neopteran 12S rRNA sequences. Our numbering system follows the convention established at the CRW Site (see Experimental procedures). There are 30 helices within insect 12S rRNA that are supported by compensatory base change evidence across our alignment.

Other than Drosophila spp., the first structural models for insect 12S rRNA were based on comparisons with other mt SSU rRNA models, such as mouse, frog, primate and sea anemone. Initially used for large-scale phylogenetic analyses (e.g.; ), it soon after became clear that the phylogenetic utility of SSU mt rRNA genes was limited to lower level divergences. Thus, a plethora of lower level studies within Insecta utilizing 12S rDNA sequences emerged. Some sequences were collected in various structural compilations, and together with more in-depth models on birds, vertebrate classes, rodents and insects, presented a refined model of animal 12S rRNA that differed from previous models in that several variable regions were adjusted from the global SSU rRNA model to fit a custom animal model [results also reached by, ) in his analysis of amphibians]. Of particular note, the honey bee 12S rRNA sequence confounded the alignment of helix H1047 [helix 38 of ] due to its high AT bias. This resulted in the proposal for a radical structure for this helix in honey bee. Analysing the secondary structure of 225 insect 12S rRNA sequences using maximum weighted matching, predicted a structure for domain III of honey bee 12S rRNA that was in more agreement with earlier insect models (e.g. ). Our predicted model for domain III of honey bee 12S rRNA is more consistent with the model of than that of, and we have identified two regions within the alignment of that were misaligned, causing the majority of domain III to be predicted inaccurately (Fig. 4A, boxed sequences 1 and 2). The second misaligned sequence (5′-GAA-3′) was noted by as being misaligned, prompting a follow-up note about the dangers of relying on conserved motifs to anchor alignments. Our realignment of the first sequence (5′-AUUUAUGU-3′) further suggests that caution should be used when anchoring variable regions with thermodynamic evidence rather than support from crystal structures, as helix H1047 contains several non-canonical base pairs in its proximal region across many arthropod taxa. As with, we note that, despite its misalignment in, the honey bee 12S rDNA sequence likely did not affect the analysis by, wherein their structural alignment was used to evaluate the performance of a suite of automated alignment programs.

Despite the continued use of 12S rDNA sequences, few recent studies have presented structural predictions. The predicted structures for lice (Phthiraptera), Trirhabda leaf beetles (Coleoptera: Chrysomelidae), species in the taxon Arthropleona (Collembola) and burnet moths (Lepidoptera) all slightly differ from our model in that they are refined for the lower taxonomic levels analysed in each study. This latter study is interesting because, while nearly all previous studies of 12S rRNA structure and evolution have focused primarily on domain III, predict a structure of the entire 12S rRNA molecule, providing primers to amplify the less common sequence domains I and II. Hopefully more studies will utilize this information, as well as our 12S rRNA structural diagram and related alignments, to compile more sequences of domains I and II from diverse taxa to improve our knowledge of this seldom-sequenced region of arthropod 12S rRNA.

Mitochondrial LSU rRNA

In most insects, the 16S rRNA gene is flanked on the 5′-side by the tRNA-Val gene, and on the 3′-side by the tRNA-LeuCUN gene. Like the 12S rRNA gene, the honey bee 16S rRNA gene follows this conserved arrangement (Fig. 1). Our secondary structure model (Fig. 4B) follows the D. melanogaster model of with minor modifications made based on an alignment of 244 neopteran 16S rRNA sequences. Our numbering system follows the convention established at the CRW Site (see Experimental procedures). There are 49 helices within insect 16S rRNA that are supported by compensatory base changes across our alignment. Despite the presence of a complete sequence for nearly 20 years, we believe that this is the first published secondary structural model for honey bee mitochondrial 16S rRNA.

Since the earliest structural predictions for arthropod 16S rRNA, namely those from Drosophila yakuba, Locusta spp., D. melanogaster and gypsy moth, Lymantria dispar, more refined models have been proposed based on larger comparisons across divergent taxa, e.g. Deltocephalus-like leafhoppers, families of Hymenoptera, families of Dermaptera, families of Orthoptera and groups of spiders. These studies have undoubtedly prompted refinements to older structural diagrams present on the CRW Site and other rRNA secondary structure databases.

analysed 16S rRNA secondary structure from over 400 taxa representing 13 insect orders. We assume that honey bee was included in their study; however, no secondary structure diagram was included for honey bee, and no honey bee sequence was present in their condensed alignment. Aside from minor deviances, the authors noted several major differences between their predicted insect model and previous structures proposed by Gutell (unpublished then, now compiled on the CRW Site) and. Specifically, helices 84 and 91 (our H2347 and H2520) were proposed to form different base pairs within their models than in the older models, and these structures are drawn as alternatives in our 16S rRNA structural diagram (Fig. 4B). Our search across 244 neopteran 16S rDNA sequences did not yield a high amount of compensatory substitutions supporting these alternative structures (data not shown).

Consistent with most structural and phylogenetic studies on 12S rRNA (see above), nearly all studies on 16S rRNA have analysed only a portion of the 3′-half of the molecule, representing conserved domains IV and V (Fig. 4B). Recently, however, analysed domains I, II, IV and V of 16S rRNA across major groups of dragonflies and damselflies (Odonata), predicting a nearly complete model for this large insect group (domain III is absent in arthropods and domain VI is rarely sequenced due to the position of the universal 3′-primer). Much of the odonate model is consistent with our predicted 16S rRNA structure presented here, with the major differences resulting from additional base pairs proposed in regions left unpaired in our model, for example the region in domain II that interacts with ribosomal protein 11 (Fig. 4B; discussed below), and helices H1–H3 in domain I proposed by. Additionally, as with, helices H2347 and H2520 of our model are different in the model. Interestingly, many of the regions in the odonate model that differ from our model, likely by being more odonate-based, do not reflect phylogenetic information consistent with expected relationships within Odonata (; see ).

Mitochondrial rRNA structural conservation.

Arthropod mt rRNA has a characteristic structure (Fig. 4) that may be correlated with genome reduction in mitochondria (e.g. ) such that many features common to rRNA from all domains of life are absent in the organellar rRNA structure. A lack of recombination in the mitochondria (e.g. ) likely contributes to a high level of non-canonical base pairs or unstructured regions that are highly conserved in non-organellar rRNAs of other organisms. The highly oxidative nature of the mitochondria is probably responsible for mutational loads not seen in most regions of the nucleus (e.g. ), particularly in nuclear coding sequences. Rates of evolution within the mitochondrial genome are typically much faster than genes within the nucleus (e.g.;;;;; ), a possible consequence of the lack of recombination and proofreading, and high oxidative environment. Studies showing gene rearrangement in arthropod mtDNA are growing (e.g.;;,;;;;,;;; ), and it has recently been demonstrated that mt genes in arthropods show a positive correlation between high rates of nucleotide substitution and gene rearrangement. While once considered stable in their position within the arthropod mitochondrial genome, evidence for rearrangements of the rRNA genes is growing (e.g.;; ). It is interesting that, despite some of the highest levels of rates of substitution and AT bias, the honey bee mt rRNA genes are arranged in the conserved ancestral manner of most other arthropods (Fig. 1B).

The paucity of crystal structures of mt rRNAs, coupled with the difficulty of aligning their variable regions due to a high AT bias, does not permit us to evaluate accurately the regions of mt rRNAs that are truncated or deleted relative to the cytoplasmic rRNAs. Ideally, significant differences from our conserved structural model, i.e. the louse 12S rRNA model or the odonate 16S rRNA model, should be evaluated across all published arthropod mt rRNA sequences to determine the base pairings with the strongest positional covariation. Unfortunately, this difficult task is beyond the scope of our current investigation. However, we stress that regions of insect mt rRNA that differ across diverse taxa in both base composition and predicted secondary structures, may in fact be conserved in tertiary interactions within the ribosome. For example, the ribosomal protein 11 binding-domain within domain II of LSU rRNA (Fig. 4B) and associated ribosomal protein 11 (L11) is one of three structural domains that are proximal on the 50S subunit, forming the GTPase-associated centre. Recent evidence suggests that the structure of the LSU rRNA stabilized by the C-terminal domain of L11 is necessary to alleviate elongation factor G (EF-G) binding in the post-translocation state of the ribosome. In arthropod 16S rRNA sequences between the L11 binding-domain and helix H991, which are shorter and closer to the core rRNA due to the loss of pseudoknot H1005 and helices H1011 and H1030, nucleotides must remain unpaired and in a ‘stretched’ conformation in order for the L11–L11 binding domain to proximally interact with two other structural domains in the GTPase-associated centre of the LSU rRNA. Thus, base pairing in these regions, even if supported by compensatory base change evidence (which by chance alone would outperform unpaired sites; see mfold structure in Fig. 4B), would be deleterious to ribosome assembly and function, as recently suggested for mammalian 16S rRNA. Given this, we doubt the formation of secondary structure in this region of 16S rRNA, although, as in the case of V4 of 18S rRNA (Fig. 3), it cannot be ruled out that base pairing in these regions may be temporary, ultimately dissolving during the tertiary interactions formed before and during translation. However, unlike the V4 region of 18S rRNA, transitionally less optimal structures are so far not supported across all arthropod sequences, and hence cannot support a hypothetical dynamic nature of this region of LSU rRNA.

Taxon-specific structures in mt rRNA variable regions would seemingly imply changes to associated ribosomal proteins and other ribosome cofactors, such that all of these molecules would coevolve to maintain the structural integrity and function of the mt ribosome. This is unlikely as ribosomal proteins are usually highly conserved across deep levels of divergence (e.g.;; ). Studies characterizing the coevolution of mitochondrial ribosomal proteins and their variable rRNA counterparts are needed to determine, if any, the secondary structures that are homologous across insects.

Characterization of IGS- and ETS-rDNA boundaries

Despite the growing number of sequenced eukaryotic genomes, few studies have predicted full sequences spanning the entire IGS, especially in arthropods. Usually, only those studies that sequence clones report complete IGS sequences, as intergenomic variation compounds other simpler methods. For genome sequencing studies, two main problems confound the accurate assemblage of IGS sequences: variation in subrepeat number, and the presence of multiple subrepeats that are not contiguous (i.e. separated by conserved or other repetitive sequences). This second problem is critical because if the variation within subrepeats is high then multiple non-contiguous subrepeats can be spuriously assembled in the wrong locale of the IGS, ultimately compounding the rebuilding of the accurate IGS sequence. Methods for accommodating these problems are in great need, as IGS regions contain the rDNA promoter region, as well as enhancer sequences that facilitate transcription of arthropod rRNA genes (e.g. ). These regions are also useful for species identification in arthropods (e.g.;;;; ) and assessing the fitness and the evolutionary ecology of organisms (e.g.; ).

Relative to coding regions of the rRNA genes, complete IGS sequences from arthropods have only been characterized for a few taxa, and many of these are dipterans: D. melanogaster, D. orena, D. virilis, D. hydei, Aedes albopictus, A. aegypti, Anopheles sinensis, the tstetse fly, Glossina sp. (Glossinidae), the black fly, Simulium sanctipauli (Simuliidae) and several species of Chironomus midges (Chironomidae). Other published arthropod complete IGS sequences are from the German cockroach Blattella germanica (Blattaria: Blattellidae), the bulldog ant, Myrmecia croslandi (Hymenoptera: Formicidae) and the crustaceans Daphnia pulex (Cladocera) and Tigriopus californicus (Copepoda). Much of the IGS and ETS have also been characterized for the pea aphid, Acyrthosiphon pisum and the silkmoth, Bombyx mori. In agreement with complete IGS sequences from other eukaryotes, including the nematode Caenorhabditis elegans, several diverse plants and trypanostomatids, IGS sequences contain one to several ‘types’ of repeat regions that are variable within species and difficult to compare across even closely related species (e.g. ). Because these repetitive regions are associated with the rDNA transcription promoter, it is no surprise that RNA polymerase I complexes of one species fail to transcribe the rRNA genes of another (e.g.; ).

IGS

We predicted the sequence and structure of the conserved regions flanking the 3′-end of the 28S rDNA (473 nts of the IGS) and the 5′-end of the 18S rDNA (2042 nts of the IGS + ETS) mostly from the repeat reads of assembly 3 (Fig. 5). While a distinct boundary between the IGS and ETS was not determined, we treat the description of the ETS separately (next section). Immediately flanking the boundary of the 28S rDNA and the IGS we predict a helix of eight base pairs that can also form in the bulldog ant (S1 in Fig. 5). Interestingly, predicted a compound helical region in this same position in bulldog ant consisting of over 639 nts. While not experimentally proven, it is possible that this region of the IGS fosters a helical element; however, structural prediction in honey bee yielded nothing substantially significant other than helix S1 (data not shown). The remaining nts of the IGS flanking the 3′-end of the 28S rDNA contain six conserved repetitive (CRp) regions, three variable repetitive (VRp) regions, one A-rich region, one T-rich region and a subrepetitive (SRp) region (Fig. 5). This first SRp region, SRp1, is estimated to repeat from four to 11 times in the honey bee genome, but could likely exceed this range. Due to the variability in sequence length and base composition of SRp1, we could not accurately predict sequences flanking the 3′-end (dashed line in Fig. 5).

As with the region of the IGS flanking the 3′-end of the 28S rDNA, the region of the IGS adjacent to the ETS cannot be predicted beyond a SRp region (Fig. 5). This second SRp region, SRp2, is more complicated than SRp1 because it is repeated at least twice within conserved stretches of sequence. SRp2 is present from three to six times (and likely more), and is then interrupted by a conserved sequence containing two CRp regions and two VRp regions (Fig. 5). SRp2 then repeats again from two to four times before flanking a conserved stretch of 1113 nts that is likely comprised mostly of the ETS. It is difficult to predict the composition and structure of the internal region of the IGS flanked by SRp1 and SRp2 for two reasons. First, the number of SRp regions vary in arthropods, ranging from one in the pea aphid to three in D. melanogaster. Thus, although we identified two distinct SRp regions in honey bee IGS, a third (or more) SRp region may exist in the central portion of the spacer. It is interesting, however, that the bulldog ant has two SRp regions, and this may be a characteristic of hymenopteran IGS sequences. Second, the composition of the central portion of the IGS in arthropods is variable, in that it can be comprised entirely of repetitive regions (e.g. bulldog ant) or alternating conserved and repetitive regions (e.g. A. sinensis). While we have not reported a complete sequence for the IGS from honey bee, we predict that our characterization of the conserved sequences flanking two SRp regions will allow for primer design and sequencing of the remaining internal sequence.

ETS

Three CRp regions and one VRp region occur within a highly conserved stretch of nts that contains the rDNA gene promoter and probable transcription enhancer elements. Spacer and gene promoters have been determined for several arthropods (listed in ), with a consensus sequence of 5′-TA-TATANGRRRR-3′ accounting for the variation across diverse taxa. We did not find a sequence within the putative ETS region of honey bee that matched this motif; however, using the Neural Network Promoter Prediction tool at the Berkeley Drosophila Genome project (http://www.fruitfly.org/seq_tools/promoter.html), a promoter sequence 5′-ATATATATATATATATATAGTCGATCGGTGAGGGGGCACCGACGACGAAA-3′ was predicted with a score of 1.0 (Fig. 5). A second promoter sequence 5′-ATTATATATATATATATATATATATATAGTCGATCGGTGAGGGGGCACCG-3′, shifted nine nts upstream from the first predicted promoter, was predicted with a score of 0.93. The bold nucleotide in each sequence is the predicted transcription initiation site (TIS), and the predicted TIS in the more optimal promoter is shaded in Fig. 5. Despite high scores, these promoter sequences are likely too close to the start of the 18S rDNA, especially when compared to the location of promoters in other arthropod ETS sequences. These putative promoter sequences and TIS should be taken with caution, as it will be necessary to determine the true promoter and TIS with S1 nuclease mapping.

Despite the high level of variation across ETS regions from different taxa, some characteristics are conserved and may allude to functional constraints not easily identifiable from primary sequence comparison. For instance, the GC content within the putative ETS region of honey bee is 35.3% and similar to other holometabolous insects; e.g. B. mori (36.7%), D. melanogaster (24%) and D. orena (23.7%) and tsetse fly, Glossina morsitans (28%). The GC content of a non-holometabolous insect, the pea aphid, is 69%, indicating a possible correlation between base composition in the ETS and phylogeny. Helical elements have been predicted within the ETS region in other eukaryotes and likely have an important role in the regulation of transcription and processing of rRNA. Using secondary structure, these regions can be aligned across closely related species, providing information on putative rRNA processing sites. Within the honey bee ETS region we calculated two putative helices using mfold that warrant further validation from closely related hymenopteran taxa (Fig. 5). We were unable to identify these helices in the bulldog ant; however, given the variability present in other ETS helical elements, e.g. in legumes or trypanostomatids, the bulldog ant and honey bee may have very divergent structures involved in the processing of the ETS region.

R2-element insertion sites

We report the presence of three distinct R2 elements inserted into the 28S rDNA at the typical target sites described for other arthropods (e.g.;;,;,; ). In general, most arthropod R2 elements insert between nucleotides 1928 and 1929 of the lonepair triloop (LPTR) H1925, however, R2 elements have also been shown to insert in helices H1835a and H1906. It now appears that only the initial nick in the target 28S rDNA sequence is conserved across arthropods, with the cleavage of the second strand highly variable such that the 5′-junctions of the R2 element and 28S rDNA are highly variable in sequence length and base composition, even within species. Indeed, our findings suggest at least two types of R2 element insert within nucleotides 1928 and 1929 of LPTR H1925 (Fig. 6A, B). Type 1 R2 elements are more conserved at their 5′-junction with the 28S rDNA and have a putative 5′-untranslated region (5′-UTR) of approximately 335 nts (Fig. 6E). In contrast, type 2 R2 elements are extraordinarily variable in their junction with the 28S rDNA and have a putative 5′-UTR of approximately 1039 nts (Fig. 6D,E). The 3′-region of this larger 5′-UTR from type 2 R2 elements is nearly identical with the entire 5′-UTR of type 1 R2 elements (Fig. 6E), suggesting that the additional nucleotides in the 5′-half of the 5′-UTR of type 2 R2 elements are not likely contributing to the structure and function of the conserved 5′-UTR. The ultimate 3′-nucleotides of the 5′-UTR in all type 1 and type 2 R2 elements from honey bee flank a predicted start codon of the putative ORF of the RT protein (Figs 6E and 7A).

We also discovered a third type of R2 element in honey bee 28S rDNA. Type 3 R2 elements were found to insert in an unusual position in the 28S rDNA, helix H1935 (Fig. 6A,B), and did not reveal a typical ORF of other arthropod R2 elements (Fig. 6G). This, coupled with a lack of a 3′-junction with the 28S rDNA, and the fact that we only detected a few copies of type 3 R2 elements, led us to doubt that this position in honey bee 28S rDNA fosters the insertion of R elements. It could also be that this particular region of 28S rDNA was erroneously assembled in the building of the genome. Still, as the number of arthropod genome sequences continue to grow, it is likely that new insertion sites in 28S rDNA will be discovered, especially within this region of domain IV, given that it is one of the most interrupted regions of rDNA across all domains of life.

Predicted R2-element ORF and 3′-UTR

Given that ORFs for the RT of R2 elements have been predicted for a variety of arthropods (e.g.;,; ), we aligned and predicted the domain structures for honey bee R2-RT sequences by comparison with another hymenopteran insect, the jewel wasp Nasonia sp. (Chalcidoidea: Pteromalidae) (Fig. 7B,D,E). Nasonia sp. is remarkable because it contains two different types of R2 elements: one that is comparable to most other arthropod R2 elements (type A), and a second that differs mainly in the presence of two additional zinc-finger domains in the amino-terminal region (type B). Only one other arthropod, the horseshoe crab Limulus polyphemus, has a type B R2 element. Interestingly, not only does our analysis of the honey bee genome identify the presence of a type B R2 element similar to horseshoe crab and Nasonia sp. type B elements (a finding consistent with ) (Fig. 7B), it also suggests that the honey bee does not have the more common type A R2 element. Within a phylogenetic context of Hymenoptera (e.g. ), wherein Chalcidoidea is derived in regards to a more primitive Apidae, this implies that Nasonia sp. has retained both A and B type R2 elements, whereas honey bee has lost the more typical A type but retained the less frequent B type, which is likely a more primitive parasite of arthropod genomes. Two types of R2 elements have been detected in the cicada killer, Sphecius speciosus (Sphecoidea; Sphecidae); however, the limited portion of the published sequence cannot lead us to determine if this insect contains type A or B R2 elements (or both). In response to this, we have identified several regions within the ORF alignment of Nasonia sp. B and honey bee R2 elements that are highly conserved for the design of PCR and sequencing primers (Fig. 7). We are currently using primers from these regions to determine the presence or absence of type B R2 elements across hymenopteran families. Given that R2 elements typically evolve at rates similar to their rDNA hosts, we expect our investigation to yield a reasonable estimate of the evolution of R2 elements across Hymenoptera, and the degree of vertical and horizontal transmission of these mobile elements.

The remaining conserved domains of the honey bee R2 element ORF, namely the RT domain with fingers/palm and thumb motifs (Fig. 7D) and the carboxyl-terminal domains that include the CCHC DNA-binding motif and endonuclease domain (Fig. 7E), are highly conserved across honey bee and jewel wasp. The sequence between the ORF stop codon (UGA) and the 3′-junction of the 28S rDNA, or 3′-UTR, is more conserved than the 5′-UTR and comprises 504 nts. Thus, it is similar in size to the Nasonia spp. 3′-UTR, which is 594 nts. While predicted secondary structures exist for R2 element-3′-UTRs from several arthropods, including the lepidopterans Bombyx mori (silkmoth), Samia cynthia (Ailanthus silkmoth), Callosamia promethea (Promethea silkmoth), Coscinocera hercules (Hercules moth) and Saturnia pyri (great peacock moth), the earwig (Forficula auricularia) and several Drosophila spp., no secondary structures have been predicted for hymenopteran insects. This is likely due to the large size of the hymenopteran R2 element-3′-UTR as compared to other arthropods, but also due to a lack of more sequences to facilitate the comparative method for predicting structure. Even using the Nasonia spp. (A and B) for comparative analysis, our attempts at predicting a secondary structure for honey bee R2 element-3′-UTR were futile.

While non-LTR retrotransposable elements are likely highly abundant in arthropod genomes, most copies are likely defective elements that drift with their inactivated rRNA gene hosts (e.g.;;; ). Subject to the process of concerted evolution or biased gene conversion of rDNA arrays, non-functional R2 elements within inactivated rRNA genes are rapidly eliminated from arthropod genomes. Despite the likely non-functional putative type 3 R2 element described above, we found limited evidence for non-functional R2 elements within the rRNA genes of the honey bee. This is probably an artefact of our method for constructing the R2 element sequences from conserved flanking regions of 28S rDNA, as the majority of non-functional R2 elements would likely reside within heavily mutated 28S rDNA sequences, which would not have been sampled in our procedure. Of the conserved R2 elements with putatively functional ORFs, we identified four nucleotide positions that contain variation in excess of single nucleotide polymorphisms (SNPs). All four cases pertain to either first or third codon positions of predicted amino acids, with the state in less frequency resulting in a stop codon (Fig. 7C,D). These variable positions likely indicate the presence of some non-functional R2 elements that have yet to be pruned out of the genome by concerted evolution of rRNA genes.

Lack of functional R1 elements in the honey bee genome

R1 elements are found in arthropod 28S rRNA genes in a conserved insertion site 74 nts downstream of the R2 insertion site (Fig. 6A). Our search for R1 elements within the honey bee genome did not recover any functional or complete sequences. Only truncated copies of putative R1 elements were assembled at the 3′-junction of the 28S rDNA (Fig. 6C). It is surprising that the honey bee does not contain complete and functional R1 elements, given that other hymenopteran insects have been purported to harbour them. According to, Nasonia sp., S. speciosus and the carpenter bee, Xylocopa sp., all contain R1 elements, although ORFs have not been determined for any of these species, and only the 3′-junctions with the 28S rDNA were actually sequenced. Similarly, detected the presence of R1 elements in 12 Hymenoptera, including the honey bee; however, the distinction between functional and truncated copies could not be determined by Southern blot analysis. The results of their RFLP analysis of the R1 insertion site (Fig. 3, ) showed a small amount of DNA in honey bee as compared to other sampled Hymenoptera, a finding consistent with our detection of only truncated R1 elements. Indeed, 5′-truncated R1 elements of sizes 0.5 and 1 kb long are known from the 28S rRNA genes of D. melanogaster. In agreement, our truncated consensus R1 element is 577 nts long, and thus could reflect the remnant of once functional R1 elements in the genome of the honey bee that have since been pruned by rRNA gene homogenization. Alternatively, R1 elements could occur in such low frequency in the honey bee genome that no functional copies were included in its assembly.

Conclusion

Our analysis of the rRNA genes from the honey bee suggests that the functional rRNA-coding regions are structurally conserved and homogeneous throughout the nuclear and mitochondrial genomes. Some regions of mt rRNAs that are variable in sequence length and base composition do not contain secondary structures that are conserved across Insecta. Studies evaluating these variable regions within the tertiary structure of the ribsome, including rRNA-rRNA and rRNA–ribosomal protein interactions, are needed to determine their structural and functional significance. Our preliminary characterization of the IGS and ETS regions linking nl rRNA genes in the honey bee suggests that these highly variable sequences are relatively similar to other holometabolous insects in organization, repetitive nature and base composition. Like most other arthropods studied, honey bee rRNA genes are subject to parasitism by retrotransposable elements, although they lack both the most common type of R2 element and functional R1 elements. In Hymenoptera, it has been hypothesized that the intriguing haplo-diploid system, in which males come from parthenogenic eggs (n) and females come from fertilized eggs (2n), is correlated with a low level of genetic variability, relative to other arthropod genomes. The characteristics of the honey bee rRNA genes we present here cannot contest this claim.

rRNA secondary structure prediction

The assembled honey bee rDNA sequences were integrated into the arthropod rRNA models (nl 18S, 5.8S, 28S, 5S rRNAs; mt 12S and 16S rRNAs) predicted and compiled at the Comparative RNA website (CRW Site) (http://www.rna.icmb.utexas.edu) and the jRNA website (http://hymenoptera.tamu.edu/rna). Helix numbering follows the E. coli system available at the CRW Site. Information pertaining to the alignment of RNA sequences using secondary structure models, including covariation analysis, thermodynamic algorithms, and ambiguously aligned regions, is available at both websites. Secondary structure model diagrams were generated with the program XRNA (developed by B. Weiser and H. Noller, University of Santa Cruz, CA). Base pair frequency tables and structural diagrams are available at the CRW Site. Differences between our previous arthropod rRNA structures and those presented here are illustrated at the jRNA website.

IGS sequence comparison

Conserved rDNA sequences flanking the 3′-terminal end of the 28S rRNA and 5′-terminal end of the 18S rRNA were used for Blast searches at the Honey Bee Genome Database (http://racerx00.tamu.edu/bee_resources.html). All options in assembly 3 were explored, including unassigned groups (bin 0); however, almost without exception, all Blast results were within the repeat reads of assembly 3. Default Blast settings were used, except that we did not filter for low complexity. Results were viewed as master-slave with identities and displayed with 500 descriptions and alignments. We used master-slave to identify reads used in subsequent Blast searches to extend the sequence. Only sequence differences repeated four or more times were reported. Unique and rare SNP differences were not reported. Sequences were then aligned manually in SeAl v2.0a11. Only one alignment was made for the 5′-end of the IGS, while three alignments were made for the 3′-end of the IGS and ETS regions. Alignments are available at the jRNA website.

R1 and R2 element prediction rDNA sequences spanning the conserved insertion sites for R1 and R2 elements in arthropods were compiled using the Blast strategy discussed above. Results containing non-rDNA sequences inserted either at the 5′- or 3′-end of the rDNA insertion site were exported to SeAl for manual alignment. Further Blast searches were performed using conserved regions of aligned sequences, allowing us to ‘walk’ across the R elements from both the 5′- and 3′-ends. Upon completion of the R2 elements, we translated the consensus sequence in six frames to determine the ORF of RT protein. This allowed for the identification of putative start and stop codons. Nucleotide and amino acid sequences of honey bee R2 element were aligned with the R2-B element of the jewel wasp, Nasonia sp. (GenBank accession no. AF090145). Alignment was performed manually with reference to the published structure of RT proteins from arthropods.

Supplementary material

The following material is available for this article online:

S1. Base pair frequency table comparing the models of Wuyts et al. (2000) and Ouvrard et al. (2000) for variable region 4 (V4) of arthropod 18S rRNA.

S2. Alignments of the IGS and ETS regions of unassembled honey bee rDNA sequences.

S3. Alignments of R1 and R2 retrotransposable elements of the honey bee.

This material is available from the Comparative RNA website, http://www.rna.icmb.utexas.edu, and the jRNA website http://hymenoptera.tamu.edu.

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    Clary DO, Wolstenholme DR. The ribosomal RNA genes of Drosophila DNA. Nucleic Acids Res. 1985;13:4029-4045.

    Not in our library10.1093/nar/13.11.4029

  43. 43.

    Clary DO, Wolstenholme DR. Drosophila mitochondrial DNA: conserved sequences in the A+T-rich region and supporting evidence for a secondary structure model of the small ribosomal RNA. J Mol Evol. 1987;25:116-125.

    Not in our library10.1007/bf02101753

  44. 44.

    Cluster PD, Marinkovic D, Allard RW, Ayala FJ. Correlations between development rates, enzyme activities, ribosomal DNA spacer-length phenotypes, and adaptation in Drosophila melanogaster. Proc Natl Acad Sci USA. 1987;84:610-614.

    Not in our library10.1073/pnas.84.2.610

  45. 45.

    Cooper A. Molecular evolutionary studies of New Zealand birds. PhD Thesis. 1994.

    Not in our library

  46. 46.

    Covacin C, Shao R, Cameron S, Barker SC. Extraordinary number of gene rearrangements in the mitochondrial genomes of lice (Phthiraptera: Insecta). Insect Mol Biol. 2006;15:63–68.

    Not in our library10.1111/j.1365-2583.2005.00608.x

  47. 47.

    Crease TJ. Sequence of the intergenic spacer between the 28S and 18S rRNA-encoding genes of the crustacean, Daphnia pulex. Gene. 1993;134:245-249.

    Not in our library10.1016/0378-1119(93)90101-8

  48. 48.

    Crease TJ, Colbourne JK. The unusually long small-subunit ribosomal RNA of the crustacean, Daphnia pulex: sequence and predicted secondary structure. J Mol Evol. 1998;46:307-313.

    Not in our library10.1007/pl00006307

  49. 49.

    Crease TJ, Taylor DJ. The origin and evolution of variable-region helices in V4 and V7 of the small-subunit ribosomal RNA of branchiopod crustaceans. Mol Biol Evol. 1998;15:1430-1446.

    Not in our library10.1093/oxfordjournals.molbev.a025871

  50. 50.

    Cross NC, Dover GA. Tsetse fly rDNA: an analysis of structure and sequence. Nucleic Acids Res. 1987a;15:15-30.

    Not in our library10.1093/nar/15.1.15

  51. 51.

    Cross NC, Dover GA. A novel arrangement of sequence elements surrounding the rDNA promoter and its spacer duplications in tsetse species. J Mol Biol. 1987b;195:63-74.

    Not in our library10.1016/0022-2836(87)90327-5

  52. 52.

    Crozier RH, Crozier YC. The mitochondrial genome of the honeybee Apis mellifera: complete sequence and genome organization. Genetics. 1993;133:97-117.

    Not in our library10.1093/genetics/133.1.97

  53. 53.

    Crozier RH, Crozier YC, Mackinlay AG. The CO-I and CO-II region of honeybee mitochondrial DNA: evidence for variation in insect mitochondrial evolutionary rates. Mol Biol Evol. 1989;4:399-411.

    Not in our library10.1093/oxfordjournals.molbev.a040553

  54. 54.

    Dams E, Hendriks L, Van de Peer Y, Neefs J-M, Smits G, Vandenbempt I, De Wachter R. Compilation of small ribosomal subunit RNA sequences. Nucleic Acids Res. 1988;16(Suppl.):r87-173.

    Not in our library10.1093/nar/16.suppl.r87

  55. 55.

    Davis RE, Kelly TJ, Masler EP, Thyagaraja BS, Rote CA, Imberski RB. Complete base sequences for the mitochondrial large subunit ribosomal RNA of the gypsy moth Lymantria dispar (L.). Insect Mol Biol. 1994;3:219-228.

    Not in our library10.1111/j.1365-2583.1994.tb00170.x

  56. 56.

    De Rijk P, Neefs J-M, Van de Peer Y, De Wachter R. Compilation of small ribosomal subunit RNA sequences. Nucleic Acids Res. 1992;20(Suppl.):2075-2089.

    Not in our library10.1093/nar/20.suppl.2075

  57. 57.

    De Rijk P, Van de Peer Y, Chapelle S, De Wachter R. Database on the structure of large ribosomal subunit RNA. Nucleic Acids Res. 1994;22:3495-3501.

    Not in our library10.1093/nar/22.17.3495

  58. 58.

    De Rijk P, Van de Peer Y, De Wachter R. Database on the structure of the large ribosomal RNA. Nucleic Acids Res. 1997;25:117-123.

    Not in our library10.1093/nar/25.1.117

  59. 59.

    De Rijk P, Caers A, Van de Peer Y, De Wachter R. Database on the structure of small ribosomal subunit RNA. Nucleic Acids Res. 1998;26:183-186.

    Not in our library

  60. 60.

    De Rijk P, Wuyts J, Van de Peer Y, Winkelmans T, De Wachter R. The European large subunit ribosomal RNA database. Nucleic Acids Res. 2000;28:177-178.

    Not in our library10.1093/nar/28.1.177

  61. 61.

    Deans AR, Gillespie JJ, Yoder MJ. An evaluation of ensign wasp classification (Hymenoptera: Evaniidae) based on molecular data and insights from ribosomal RNA secondary structure. Syst Entomol. 2006;31:517-528.

    Not in our library

  62. 62.

    Degelmann A, Royer H-D, Hollenberg CP. The organization of the ribosomal RNA genes of Chironomus tentans and some closely related species. Chromosoma. 1979;71:263-281.

    Not in our library10.1007/bf00287136

  63. 63.

    Dover GA, Flavell RB. Molecular coevolution: DNA divergence and the maintenance of function. Cell. 1984;38:622-623.

    Not in our library10.1016/0092-8674(84)90255-1

  64. 64.

    Dover GA, Tautz D. Conservation and divergence in multigene families: alternatives to selection and drift. Philos Trans Roy Soc Lond B Biol Sci. 1986;312:275-289.

    Not in our library10.1098/rstb.1986.0007

  65. 65.

    Dowton M, Austin AD. Molecular phylogeny of the insect order Hymenoptera: Apocritan relationships. Proc Natl Acad Sci USA. 1994;91:9911-9915.

    Not in our library10.1073/pnas.91.21.9911

  66. 66.

    Dowton M, Austin AD. Increased genetic diversity in mitochondrial genes is correlated with the evolution of parasitism in the Hymenoptera. J Mol Evol. 1995;41:958-965.

    Not in our library10.1007/bf00173176

  67. 67.

    Dowton M, Austin AD. Evolutionary dynamics of a mitochondrial rearrangement ‘hotspot’ in the Hymenoptera. Mol Biol Evol. 1999;16:298-309.

    Not in our library10.1093/oxfordjournals.molbev.a026111

  68. 68.

    Dowton M, Campbell NJH. Intramitochondrial recombination: is it why some mitochondrial genes sleep around? Trends Ecol Evol. 2001;16:269-271.

    Not in our library10.1016/s0169-5347(01)02182-6

  69. 69.

    Dunon-Bluteau D, Brun G. The secondary structures of the Xenopus laevis and human mitochondrial small ribosomal subunit RNA are similar. FEBS Lett. 1986;198:333-337.

    Not in our library10.1016/0014-5793(86)80431-8

  70. 70.

    Eickbush TH, Craig NL, Craigie R, Gellert M, Lambowitz A. R2 and related site-specific non-long terminal repeat retrotransposons. Mobile DNA II. 2002;813-835.

    Not in our library

  71. 71.

    Eickbush DG, Eickbush TH. Vertical transmission of the retrotransposable elements R1 and R2 during the evolution of the Drosophila melanogaster species subgroup. Genetics. 1995;139:671-684.

    Not in our library10.1093/genetics/139.2.671

  72. 72.

    Eickbush DG, Lathe WD, Francino MP, Eickbush TH. R1 and R2 retrotransposable elements of Drosophila evolve at rates similar to those of nuclear genes. Genetics. 1995;139:685-695.

    Not in our library10.1093/genetics/139.2.685

  73. 73.

    Elgavish T, Cannone JJ, Lee JC, Harvey SC, Gutell RR. AA.AG@Helix.Ends: A:A and A:G base-pairs at the ends of 16S and 23S rRNA helices. J Mol Biol. 2001;310:735-753.

    Not in our library10.1006/jmbi.2001.4807

  74. 74.

    Ellis RE, Sulston JE, Coulson AR. The rDNA of C. elegans: sequence structure. Nucleic Acids Res. 1986;14:2345-2364.

    Not in our library10.1093/nar/14.5.2345

  75. 75.

    Entelis NS, Kolesnikova OA, Dogan S, Martin RP, Tarassov IA. 5S rRNA and tRNA import into human mitochondria: comparison of in vitro requirements. J Biol Chem. 2001;276:45642-45653.

    Not in our library10.1074/jbc.m103906200

  76. 76.

    Entelis N, Kolesnikova O, Kazakova H, Brandina I, Kamenski P, Martin RP, Tarassov I. Import of nuclear encoded RNAs into yeast and human mitochondria: experimental approaches and possible biomedical applications. Genet Eng. 2002;24:191-213.

    Not in our library10.1007/978-1-4615-0721-5_9

  77. 77.

    Fang Q, Black WCIV, Blocker HD, Whitcomb RF. A phylogeny of New World Deltocephalus-like leafhopper genera based on mitochondrial 16S ribosomal DNA sequences. Mol Phylogenet Evol. 1993;2:119-131.

    Not in our library10.1006/mpev.1993.1012

  78. 78.

    Fernández M, Polanco C, Ruiz ML, Pérez de la Vega M. A comparative study of the structure of the rDNA intergenic spacer of Lens culinaris Medik. & other legume species. Genome. 2000;43:597-603.

    Not in our library10.1139/g00-022

  79. 79.

    Field KG, Olsen GJ, Lane DJ, Giovannoni SJ, Ghiselin MT, Raff EC, Pace NR, Raff RA. Molecular phylogeny of the animal kingdom. Science. 1988;239:748-753.

    Not in our library10.1126/science.3277277

  80. 80.

    Flook PK, Rowell CHF. The effectiveness of mitochondrial rRNA gene sequences for the reconstruction of the phylogeny of an insect order, Orthoptera. Mol Phylogenet Evol. 1997;8:177-192.

    Not in our library10.1006/mpev.1997.0425

  81. 81.

    Friedrich M, Tautz D. An episodic change of rDNA nucleotide substitution rate has occurred at the time of the emergence of the insect order Diptera. Mol Biol Evol. 1997a;14:644-653.

    Not in our library10.1093/oxfordjournals.molbev.a025804

  82. 82.

    Friedrich M, Tautz D. Evolution and phylogeny of the Diptera: a molecular phylogenetic analysis using 28S rDNA sequences. Syst Biol. 1997b;46:674-698.

    Not in our library10.1093/sysbio/46.4.674

  83. 83.

    Fujiwara H, Ishikawa H. Structure of the Bombyx mori rDNA: Initiation site for its transcription. Nucleic Acids Res. 1987;15:1245-1258.

    Not in our library10.1093/nar/15.3.1245

  84. 84.

    Gerbi SA, MacIntyre RJ. Evolution of ribosomal DNA. Molecular Evolutionary Genetics. 1985;419-517.

    Not in our library

  85. 85.

    Gerbi SA, Zimmermann RA, Dahlberg AE. Expansion segments: Regions of variable size that interrupt the universal core secondary structure of ribosomal RNA. Ribosomal RNA: Structure, Evolution, Processing, and Function in Protein Synthesis. 1996;71-87.

    Not in our library

  86. 86.

    Gillespie JJ, Kjer KM, Duckett CN, Tallamy DW. Convergent evolution of cucurbitacin feeding in spatially isolated rootworm taxa (Coleoptera: Chrysomelidae; Galerucinae, Luperini). Mol Phylogenet Evol. 2003;29:161-175.

    Not in our library10.1016/s1055-7903(03)00256-2

  87. 87.

    Gillespie JJ, Cannone JJ, Gutell RR, Cognato AI. A secondary structural model of the 28S rRNA expansion segments D2 and D3 from rootworms and related leaf beetles (Coleoptera: Chrysomelidae; Galerucinae). Insect Mol Biol. 2004;13:495-518.

    Not in our library10.1111/j.0962-1075.2004.00509.x

  88. 88.

    Gillespie JJ, McKenna CH, Yoder MJ, Gutell RR, Johnston JS, Kathirithamby J, Cognato AI. Assessing the odd secondary structural properties of nuclear small subunit ribosomal RNA sequences (18S) of the twisted-wing parasites (Insecta: Strepsiptera). Insect Mol Biol. 2005a;14:625-643.

    Not in our library10.1111/j.1365-2583.2005.00591.x

  89. 89.

    Gillespie JJ, Munro JB, Heraty JM, Yoder MJ, Owen AK, Carmichael AE. A secondary structural model of the 28S rRNA expansion segments D2 and D3 for chalcidoid wasps (Hymenoptera: Chalcidoidea). Mol Biol Evol. 2005b;22:1593-1608.

    Not in our library10.1093/molbev/msi152

  90. 90.

    Gillespie JJ, Yoder MJ, Wharton RA. Predicted secondary structures for 28S and 18S rRNA from Ichneumonoidea (Insecta: Hymenoptera: Apocrita): impact on sequence alignment and phylogeny estimation. J Mol Evol. 2005c;61:114-137.

    Not in our library10.1007/s00239-004-0246-x

  91. 91.

    Glotz C, Brimacombe R. An experimentally-derived model for the secondary structure of the 16S ribosomal RNA from Escherichia coli. Nucleic Acids Res. 1980;8:2377-2395.

    Not in our library10.1093/nar/8.11.2377

  92. 92.

    Gorski JL, Gonzalez KL, Schmicuez RD. The secondary structure of human 28S rRNA: The structure and evolution of a mosaic rRNA gene. J Mol Evol. 1987;24:236-251.

    Not in our library10.1007/bf02111237

  93. 93.

    Grauer G. Gene diversity in Hymenoptera. Evolution. 1985;39:190-199.

    Not in our library10.1111/j.1558-5646.1985.tb04091.x

  94. 94.

    Graybeal A. Evaluating the phylogenetic utility of genes: a search for genes informative about deep divergences among vertebrates. Syst Biol. 1994;43:174-193.

    Not in our library

  95. 95.

    Green R, Ellington AD, Szostak JW. In vitro genetic analysis of the Tetrahymena self-splicing intron. Nature. 1990;347:406-408.

    Not in our library10.1038/347406a0

  96. 96.

    Greenberg JR. Synthesis and properties of ribosomal RNA in Drosophila. J Mol Biol. 1969;46:85-98.

    Not in our library10.1016/0022-2836(69)90059-x

  97. 97.

    Gruendler P, Unfried I, Pascher K, Schweizer D. rDNA intergenic region from Arabidopsis thaliana: structural analysis, infraspecific variation and functional implications. J Mol Biol. 1991;221:1209-1222.

    Not in our library10.1016/0022-2836(91)90929-z

  98. 98.

    Gutell RR. Collection of Small Subunit (16S- and 16S-like) ribosomal RNA structures. Nucleic Acids Res. 1993;21:3051-3054.

    Not in our library10.1093/nar/21.13.3051

  99. 99.

    Gutell RR. Collection of Small Subunit (16S- and 16S-like) ribosomal RNA structures: 1994. Nucleic Acids Res. 1994;22:3502-3507.

    Not in our library10.1093/nar/22.17.3502

  100. 100.

    Gutell RR, Dahlberg AE, Zimmerman RA. Comparative sequence analysis and the structure of 16S and 23S rRNA. Ribosomal RNA: Structure, Evolution, Processing and Function in Protein Synthesis. 1996;111-128.

    Not in our library

  101. 101.

    Gutell RR, Fox GE. A compilation of large subunit RNA sequences presented in a structural format. Nucleic Acids Res. 1988;16S:r175-r269.

    Not in our library10.1093/nar/16.suppl.r175

  102. 102.

    Gutell RR, Weiser B, Woese CR, Noller HF. Comparative anatomy of 16S-like ribosomal RNA. Prog Nucleic Acid Res Mol Biol. 1985;32:155-216.

    Not in our library10.1016/s0079-6603(08)60348-7

  103. 103.

    Gutell RR, Noller HF, Woese CR. Higher order structure in ribosomal RNA. EMBO J. 1986;5:1111-1113.

    Not in our library10.1002/j.1460-2075.1986.tb04330.x

  104. 104.

    Gutell RR, Schnare MN, Gray MW. A compilation of large subunit (23S-like) ribosomal RNA sequences presented in a secondary structure format. Nucleic Acids Res. 1990;18S:2319-2330.

    Not in our library10.1093/nar/18.suppl.2319

  105. 105.

    Gutell RR, Schnare MN, Gray MW. A compilation of large subunit (23S- and 23S-like) ribosomal RNA structures. Nucleic Acids Res. 1992;20S:2095-2109.

    Not in our library10.1093/nar/20.suppl.2095

  106. 106.

    Gutell RR, Gray MW, Schnare MN. A compilation of large subunit (23S- and 23S-like) ribosomal RNA structures: 1993. Nucleic Acids Res. 1993;21S:3055-3074.

    Not in our library10.1093/nar/21.13.3055

  107. 107.

    Gutell RR, Cannone JJ, Du Shang ZY, Serra MJ. A story: unpaired adenosine bases in ribosomal RNAs. J Mol Biol. 2000;304:335-354.

    Not in our library10.1006/jmbi.2000.4172

  108. 108.

    Hancock JM. The contribution of DNA slippage to eukaryotic nuclear 18S rRNA evolution. J Mol Evol. 1995;40:629-639.

    Not in our library10.1007/bf00160511

  109. 109.

    Hancock JM, Dover GA. Molecular coevolution among cryptically simple expansion segments of eukaryotic 26S/28S rRNAs. Mol Biol Evol. 1988;5:377-392.

    Not in our library10.1093/oxfordjournals.molbev.a040505

  110. 110.

    Hancock JM, Tautz D, Dover GA. Evolution of the secondary structures and compensatory mutations of the ribosomal RNAs of Drosophila melanogaster. Mol Biol Evol. 1988;5:393-414.

    Not in our library10.1093/oxfordjournals.molbev.a040501

  111. 111.

    Harms J, Schluenzen F, Zarivach R, Bashan A, et al. High resolution structure of the large ribosomal subunit from a mesophilic eubacterium. Cell. 2001;107:679-688.

    Not in our library10.1016/s0092-8674(01)00546-3

  112. 112.

    Hassouna N, Michot B, Bachellerie J-P. The complete nucleotide sequence of mouse 28S rRNA gene: Implications for the process of size increase of the large subunit rRNA in higher eukaryotes. Nucleic Acids Res. 1984;12:3563-3583.

    Not in our library10.1093/nar/12.8.3563

  113. 113.

    Hayward DC, Glover DM. Analysis of the Drosophila rDNA promoter by transient expression. Nucleic Acids Res. 1988;25:4253-4268.

    Not in our library10.1093/nar/16.10.4253

  114. 114.

    Hendriks L, de Baere R, van Broeckhoven C, De Wachter R. Primary and secondary structure of the 18S ribosomal RNA of the insect species Tenebrio molitor. FEBS Lett. 1988a;232:115-120.

    Not in our library10.1016/0014-5793(88)80398-3

  115. 115.

    Hendriks L, van Broeckhoven C, Vandenberghe A, Van de Peer Y, De Wachter R. Primary and secondary structure of the 18S ribosomal RNA of the bird spider Eurypelma californica and evolutionary relationships among eukaryotic phyla. Eur J Biochem. 1988b;177:15-20.

    Not in our library10.1111/j.1432-1033.1988.tb14339.x

  116. 116.

    Hickson RE. Evolutionary tails from the South Pacific. Being a wondrous and exciting account of investigations into the histories of New Zealand skinks, and in which it is shown that things are not what they seem. PhD Thesis. 1993.

    Not in our library

  117. 117.

    Hickson RE, Simon C, Cooper A, Spicer GS, Sullivan J, Penny D. Conserved sequence motifs, alignment, and secondary structure for the third domain of animal 12S rRNA. Mol Biol Evol. 1996;13:150-169.

    Not in our library10.1093/oxfordjournals.molbev.a025552

  118. 118.

    Hickson RE, Simon C, Perrey SW. The performance of several multiple-sequence alignment programs in relation to secondary-structure features for an rRNA sequence. Mol Biol Evol. 2000;17:530-539.

    Not in our library10.1093/oxfordjournals.molbev.a026333

  119. 119.

    Hillis DM, Dixon MT. Ribosomal DNA: molecular evolution and phylogenetic inference. Quart Rev Biol. 1991;66:411-453.

    Not in our library10.1086/417338

  120. 120.

    Hixson JE, Brown WM. A comparison of the small ribosomal RNA genes from the mitochondrial DNA of the great apes and humans: sequence, structure, evolution, and phylogenetic implications. Mol Biol Evol. 1986;3:1-18.

    Not in our library10.1093/oxfordjournals.molbev.a040379

  121. 121.

    Hoang L, Fredrick K, Noller HF. Creating ribosomes with an all-RNA 30S subunit P site. Proc Natl Acad Sci USA. 2004;101:12439-12443.

    Not in our library10.1073/pnas.0405227101

  122. 122.

    Hovmöller R, Pape T, Kallersjö M. The Paleoptera problem: basal pterygote phylogeny inferred from 18S and 28S rDNA sequences. Cladistics. 2002;18:313-323.

    Not in our library10.1111/j.1096-0031.2002.tb00153.x

  123. 123.

    Hsieh H-M, Hou R-J, Chen K-F, Tsai L-C, Liu S-W, Liu K-L, Linacre A, Lee JC-I. Establishing the rDNA IGS structure of Cannabis sativa. J Forensic Sci. 2004;49:1-4.

    Not in our library

  124. 124.

    Huysmans E, De Wachter R. Compilation of small ribosomal subunit RNA sequences. Nucleic Acids Res. 1986;14(Suppl.):r73-r118.

    Not in our library10.1093/nar/14.suppl.r73

  125. 125.

    Hwang UI, Kim W, Tautz D, Friedrich M. Molecular phylogenetics at the Felsenstein zone: approaching the Strepsiptera problem using 5.8S and 28S rDNA sequences. Mol Phylogenet Evol. 1998;9:470-480.

    Not in our library10.1006/mpev.1998.0518

  126. 126.

    Hypsa V. Parasite histories and novel phylogenetic tools: alternative approaches to inferring parasite evolution from molecular markers. Int J Parasitol. 2006;36:141-155.

    Not in our library10.1016/j.ijpara.2005.10.010

  127. 127.

    Ishikawa H. Evolution of ribosomal RNA. Comp Biochem Physiol B. 1977;58:1-7.

    Not in our library10.1016/0305-0491(77)90116-x

  128. 128.

    Ishikawa H, Newburgh RW. Studies of thermal conversion of 28S RNA of Galleria mellonella (L.) to an 18S product. J Mol Biol. 1972;64:135-144.

    Not in our library10.1016/0022-2836(72)90325-7

  129. 129.

    Israelewski N. Structure and function of an AT-rich, interspersed repetitive sequence from Chironomus thummi: solenoidal DNA, 142 bp palindrome-frame and homologies with the sequence for site-specific recombination of bacterial transposons. Nucleic Acids Res. 1983;11:6985-6996.

    Not in our library10.1093/nar/11.20.6985

  130. 130.

    Israelewski N, Schmidt ER. Spacer size heterogeneity in ribosomal DNA of Chironomus thummi is due to a 120 bp repeat homologous to a predominantly centromeric repeated sequence. Nucleic Acids Res. 1982;10:7689-7700.

    Not in our library10.1093/nar/10.23.7689

  131. 131.

    Jackson SA, Cannone JJ, Lee JC, Gutell RR, Woodson SA. Distribution of rRNA introns in the three-dimensional structure of the ribosome. J Mol Biol. 2002;323:35-52.

    Not in our library10.1016/s0022-2836(02)00895-1

  132. 132.

    Jakubczak JL, Xiong Y, Eickbush TH. Type I (R1) and type II(R2): ribosomal DNA insertions of Drosophila melanogaster are retrotransposable elements closely related to those of Bombyx mori. J Mol Biol. 1990;212:37-52.

    Not in our library10.1016/0022-2836(90)90303-4

  133. 133.

    Jakubczak JL, Burke WD, Eickbush TH. Retrotransposable elements R1 and R2 interrupt the rRNA genes of most insects. Proc Natl Acad Sci USA. 1991;88:3295-3299.

    Not in our library10.1073/pnas.88.8.3295

  134. 134.

    Jakubczak JL, Zenni MK, Woodruff RC, Eickbush TH. Turnover of R1 (type I) and R2 (type II) retrotransposable elements in the ribosomal DNA of Drosophila melanogaster. Genetics. 1992;131:129-142.

    Not in our library10.1093/genetics/131.1.129

  135. 135.

    Jamrich M, Miller OL. The rare transcripts of interrupted rRNA genes in Drosophila melanogaster are processed or degraded during synthesis. EMBO J. 1984;3:1541-1545.

    Not in our library10.1002/j.1460-2075.1984.tb02008.x

  136. 136.

    Kambhampati S, Kjer KM, Thorne BL. Phylogenetic relationships among termite families based on DNA sequence of mitochondrial LSU ribosomal RNA gene. Insect Mol Biol. 1996;5:229-238.

    Not in our library10.1111/j.1365-2583.1996.tb00097.x

  137. 137.

    van Keulen H, Mertz PM, LoVerde T, Shi H, Rekosh DM. Characterization of a 54-nucleotide gap region in the 28S rRNA gene of Schistosoma mansoni. Mol Biochem Parasitol. 1991;45:205-214.

    Not in our library10.1016/0166-6851(91)90087-m

  138. 138.

    Kjer KM. Use of rRNA secondary structure in phylogenetic studies to identify homologous positions: an example of alignment and data presentation from the frogs. Mol Phylogenet Evol. 1995;4:314-330.

    Not in our library10.1006/mpev.1995.1028

  139. 139.

    Kjer KM. An alignment template for amphibian 12S rRNA, domain III: conserved primary and secondary structural motifs. J Herpetol. 1997;31:599-604.

    Not in our library

  140. 140.

    Kjer KM. Aligned 18S and insect phylogeny. Syst Biol. 2004;53:506-514.

    Not in our library10.1080/10635150490445922

  141. 141.

    Kjer KM, Baldridge GD, Fallon AM. Mosquito large subunit ribosomal RNA: simultaneous alignment of primary and secondary structure. Biochim Biophys Acta. 1994;1217:147-155.

    Not in our library10.1016/0167-4781(94)90028-0

  142. 142.

    Klein DJ, Schmeing TM, Moore PB, Steitz TA. The kink-turn: a new RNA secondary structure motif. EMBO J. 2001;20:4214-4221.

    Not in our library10.1093/emboj/20.15.4214

  143. 143.

    Klein DJ, Moore PB, Steitz TA. The roles of ribosomal proteins in the structure assembly, and evolution of the large ribosomal subunit. J Mol Biol. 2004a;340:141-177.

    Not in our library10.1016/j.jmb.2004.03.076

  144. 144.

    Klein DJ, Moore PB, Steitz TA. The contribution of metal ions to the structural stability of the large ribosomal subunit. RNA. 2004b;10:1366-1379.

    Not in our library10.1261/rna.7390804

  145. 145.

    Kojima KK, Fujiwara H. Cross-genome screening of novel sequence-specific non-LTR retrotransposons: various multicopy RNA genes and microsatellites are selected as targets. Mol Biol Evol. 2004;21:207-217.

    Not in our library10.1093/molbev/msg235

  146. 146.

    Kojima KK, Fujiwara H. Long-term inheritance of the 28S rDNA-specific retrotransposon R2. Mol Biol Evol. 2005;22:2157-2165.

    Not in our library10.1093/molbev/msi210

  147. 147.

    Koonin EV, Fedorova ND, Jackson JD, Jacobs AR, et al. A comprehensive evolutionary classification of proteins encoded in complete eukaryotic genomes. Genome Biol. 2004;5:R7.1-R7.28.

    Not in our library10.1186/gb-2004-5-2-r7

  148. 148.

    Kwon O-Y, Ishikawa H. Unique structure in the intergenic and 5′ external transcribed spacer of the ribosomal RNA gene from the pea aphid Acyrthosiphon pisum. Eur J Biochem. 1992;206:935-940.

    Not in our library10.1111/j.1432-1033.1992.tb17003.x

  149. 149.

    Kwon OY, Ogino K, Ishikawa H. The longest 18S ribosomal RNA ever known. Eur J Biochem. 1991;202:827-833.

    Not in our library10.1111/j.1432-1033.1991.tb16439.x

  150. 150.

    Lakshmikumaran M, Negi MS. Structural analysis of two length variants of the rDNA intergenic spacer from Eruca sativa. Plant Mol Biol. 1994;24:915-927.

    Not in our library10.1007/bf00014445

  151. 151.

    Lanadu C, du Montcel HT, Jolivot MP, Glaszmann JC, de Leon DG. Variation of ribosomal gene spacer length among wild and cultivated banana. Hereditas. 1992;68:147-156.

    Not in our library10.1038/hdy.1992.23

  152. 152.

    Lang BF, Burger G, O’Kelly CJ, Cedergren R, et al. An ancestral mitochondrial DNA resembling a eubacterial genome in miniature. Nature. 1997;387:454-455.

    Not in our library10.1038/387493a0

  153. 153.

    de Lanversin G, Jacq B. Sequence and secondary structure of the central domain of Drosophila 26S rRNA: a universal model for the central domain of the large rRNA containing the region in which the central break may happen. J Mol Evol. 1989;28:403-417.

    Not in our library10.1007/bf02603076

  154. 154.

    Larsen N. Higher order interactions in 23S rRNA. Proc Natl Acad Sci USA. 1992;89:5044-5048.

    Not in our library10.1073/pnas.89.11.5044

  155. 155.

    Lee JC, Gutell RR. Diversity of base-pair conformations and their occurrence in rRNA structure and RNA structural motifs. J Mol Biol. 2004;344:1225-1249.

    Not in our library10.1016/j.jmb.2004.09.072

  156. 156.

    Lee JC, Cannone JJ, Gutell RR. The lonepair triloop: a new motif in RNA structure. J Mol Biol. 2003;325:65-83.

    Not in our library10.1016/s0022-2836(02)01106-3

  157. 157.

    Leontis NB, Westhof E. Analysis of RNA motifs. Curr Opin Struct Biol. 2003;13:300-308.

    Not in our library10.1016/s0959-440x(03)00076-9

  158. 158.

    Lescoute A, Leontis NB, Massire C, Westhof E. Recurrent structural RNA motifs, Isostericity Matrices and sequence alignments. Nucleic Acids Res. 2005;33:2395-2409.

    Not in our library10.1093/nar/gki535

  159. 159.

    Levinson G, Gutman GA. Slipped-strand mispairing: a major mechanism for DNA sequence evolution. Mol Biol Evol. 1987;4:203-221.

    Not in our library10.1093/oxfordjournals.molbev.a040442

  160. 160.

    Lin C-P, Danforth BN. How do insect nuclear and mitochondrial gene substitution patterns differ? Insights from Bayesian analyses of combined datasets. Mol Phylogenet Evol. 2004;30:686-702.

    Not in our library10.1016/s1055-7903(03)00241-0

  161. 161.

    Long EO, Dawid IB. Restriction analysis of spacers in ribosomal DNA of Drosophila melanogaster. Nucleic Acids Res. 1979;7:205-215.

    Not in our library10.1093/nar/7.1.205

  162. 162.

    Long EO, Dawid IB. Repeated genes in eukaryotes. Annu Rev Biochem. 1980;49:727-764.

    Not in our library10.1146/annurev.bi.49.070180.003455

  163. 163.

    Luan DD, Eickbush TH. Downstream 28S gene sequences on the RNA template affect the choice of primer and the accuracy of initiation by the R2 reverse transcriptase. Mol Cell Biol. 1996;16:4726-4734.

    Not in our library10.1128/mcb.16.9.4726

  164. 164.

    Lynch M. Mutation accumulation in transfer RNAs: molecular evidence for Muller's ratchet in mitochondrial genomes. Mol Biol Evol. 1995;13:209-220.

    Not in our library10.1093/oxfordjournals.molbev.a025557

  165. 165.

    Lynch M. Mutation accumulation in nuclear, organelle, and prokaryotic transfer RNA genes. Mol Biol Evol. 1997;14:914-925.

    Not in our library10.1093/oxfordjournals.molbev.a025834

  166. 166.

    Macas J, Navrátilová A, Mézáros T. Sequence subfamilies of satellite repeats related to rDNA intergenic spacer are differentially amplified on Vicia sativa chromosomes. Chromosoma. 2003;112:152-158.

    Not in our library10.1007/s00412-003-0255-3

  167. 167.

    Magalhães PJ, Andreu AL, Schon EA. Evidence for the presence of 5S rRNA in mammalian mitochondria. Mol Biol Cell. 1998;9:2375-2382.

    Not in our library10.1091/mbc.9.9.2375

  168. 168.

    Mahapatra S, Ghosh T, Adhya S. Import of small RNAs into Leishmania mitochondria in vitro. Nucleic Acids Res. 1994;22:3381-3386.

    Not in our library10.1093/nar/22.16.3381

  169. 169.

    Márquez LM, Miller DJ, MacKenzie JB, van Oppen MJH. Pseudogenes contribute to the extreme diversity of nuclear ribosomal DNA in the hard coral Acropora. Mol Biol Evol. 2003;20:1077-1086.

    Not in our library10.1093/molbev/msg122

  170. 170.

    Martin AP, Palumbi SR. Body size, metabolic rate, generation time, and the molecular clock. Proc Natl Acad Sci. 1993;90:4087-4091.

    Not in our library10.1073/pnas.90.9.4087

  171. 171.

    Masta SE. Mitochondrial sequence evolution in spiders: Intraspecific variation in tRNAs lacking the TΨC arm. Mol Biol Evol. 2000;17:1091-1100.

    Not in our library10.1093/oxfordjournals.molbev.a026390

  172. 172.

    Masta SE, Boore JL. The complete mitochondrial genome sequence of the spider Habronattus oregonensis reveals rearranged and extremely truncated tRNAs. Mol Biol Evol. 2004;21:893-902.

    Not in our library10.1093/molbev/msh096

  173. 173.

    Mathews DH, Banerjee AR, Luan DD, Eickbush TH, Turner DH. Secondary structure model of the RNA recognized by the reverse transcriptase from the R2 retrotransposable element. RNA. 1997;3:1-16.

    Not in our library

  174. 174.

    Mathews DH, Sabina J, Zuker M, Turner DH. Expanded sequence dependence of thermodynamic parameters improves prediction of RNA secondary structure. J Mol Biol. 1999;288:911-940.

    Not in our library10.1006/jmbi.1999.2700

  175. 175.

    McLain DK, Collins FH, Brandling-Bennet AD, Were JBO. Microgeographic variation in rDNA intergenic spacers of Anopheles gambiae in Western Kenya. Heredity. 1989;62:257-264.

    Not in our library10.1038/hdy.1989.36

  176. 176.

    McTaggart S, Crease T. Selection on the structural stability of a ribosomal RNA expansion segment in Daphnia obtusa. Mol Biol Evol. 2005;22:1309-1319.

    Not in our library10.1093/molbev/msi119

  177. 177.

    Mears JA, Sharma MR, Gutell RR, McCook AS, Richardson PE, Caulfield TR, Agrawal RK, Harvey SC. A structural model for the large subunit of the mammalian mitochondrial ribosome. J Mol Biol. 2006;358:193-212.

    Not in our library10.1016/j.jmb.2006.01.094

  178. 178.

    Michot B, Bachellerie J-P, Raynal F. Sequence and secondary structure of mouse 28S rRNA 5′-terminal domain: organisation 5 8S−28S rRNA complex. Nucleic Acids Res. 1982;10:5273-5283.

    Not in our library10.1093/nar/10.17.5273

  179. 179.

    Michot B, Hassouna N, Bachellerie J-P. Secondary structure of mouse 28S rRNA and general model for the folding of the large rRNA in eukaryotes. Nucleic Acids Res. 1984;12:4259-4279.

    Not in our library10.1093/nar/12.10.4259

  180. 180.

    Misof B, Fleck G. Comparative analysis of mt LSU rRNA secondary structures of Odonates: structural variability and phylogenetic signal. Insect Mol Biol. 2003;12:535-547.

    Not in our library10.1046/j.1365-2583.2003.00432.x

  181. 181.

    Misof B, Anderson CL, Buckley TR, Erpenbeck D, Rickert A, Misof KJ. An empirical analysis of mt 16S rRNA covarion-like evolution in insects: site-specific rate variation is clustered and frequently detected. J Mol Evol. 2002;554:460-469.

    Not in our library10.1007/s00239-002-2341-1

  182. 182.

    Morales-Hojas R, Post RJ, Wilson MD, Cheke RA. Completion of the sequence of the nuclear ribosomal DNA subunit of Simulium sanctipauli, with descriptions of the 18S, 28S genes and the IGS. Med Vet Entomol. 2002;16:386-394.

    Not in our library10.1046/j.1365-2915.2002.00392.x

  183. 183.

    Muir G, Fleming CC, Schlötterer C. Three divergent rDNA clusters predate the species divergence in Quercus petrea (Matt.) Liebl. & Quercus robur L. Mol Biol Evol. 2001;18:112-119.

    Not in our library10.1093/oxfordjournals.molbev.a003785

  184. 184.

    Mukha DV, Sidorenko AP, Lazebnaya IV, Wiegmann BM, Schal C. Analysis of intraspecies polymorphism in the ribosomal DNA cluster of the cockroach Blattella germanica. Insect Mol Biol. 2000;9:217-222.

    Not in our library10.1046/j.1365-2583.2000.00175.x

  185. 185.

    Mukha DV, Wiegmann BM, Schal C. Saltatory changes in the structure of the ribosomal DNA external transcribed spacer during the evolution of cockroaches of genus Blattella. Doklady Biol Sci. 2002;387:549-552.

    Not in our library10.1023/a:1021745709410

  186. 186.

    Murtif VL, Rae PM. In vivo transcription of rDNA spacers in Drosophila. Nucleic Acids Res. 1985;13:3221-3239.

    Not in our library10.1093/nar/13.9.3221

  187. 187.

    Musters W, Venema J, van der Linden G, van Heerikhuizen H, Klootwijk J, Planta RJ. A system for the analysis of yeast ribosomal DNA mutations. Mol Cell Biol. 1989;9:551-559.

    Not in our library10.1128/mcb.9.2.551-559.1989

  188. 188.

    Musters W, Goncalves PM, Boon K, Raué HA, van Heerikhuizen H, Planta RJ. The conserved GTPase center and variable region V9 from Saccharomyces cerevisae 26 rRNA can be replaced by their equivalents from other prokaryotes or eukaryotes without detectable loss of ribosomal function. Proc Natl Acad Sci USA. 1991;88:1469-1473.

    Not in our library10.1073/pnas.88.4.1469

  189. 189.

    Neefs J-M, De Wachter R. A proposal for the secondary structure of a variable area of eukaryotic small ribosomal subunit RNA involving the existence of a pseudoknot. Nucleic Acids Res. 1990;18:5695-5704.

    Not in our library10.1093/nar/18.19.5695

  190. 190.

    Neefs J-M, Van de Peer Y, Hendriks L, De Wachter R. Compilation of small ribosomal subunit RNA sequences. Nucleic Acids Res. 1990;18(Suppl.):2237-2317.

    Not in our library10.1093/nar/18.suppl.2237

  191. 191.

    Neefs J-M, Van de Peer Y, De Rijk P, Goris A, De Wachter R. Compilation of small ribosomal subunit RNA sequences. Nucleic Acids Res. 1991;19:1987-2015.

    Not in our library10.1093/nar/19.suppl.1987

  192. 192.

    Neefs J-M, Van de Peer Y, De Rijk P, Chapelle S, De Wachter R. Compilation of small ribosomal subunit RNA structures. Nucleic Acids Res. 1993;21:3025-3049.

    Not in our library10.1093/nar/21.13.3025

  193. 193.

    Niehuis O, Naumann CM, Misof B. Identification of evolutionary conserved structural elements in the mt SSU rRNA of Zygaenoidea (Lepidoptera): a comparative sequence analysis. Organ Division Evol. 2006;6:17-32.

    Not in our library

  194. 194.

    Nissen P, Hansen J, Ban N, Moore PB, Steitz TA. The structural basis of ribosome activity in peptide bond synthesis. Science. 2000;289:920-930.

    Not in our library10.1126/science.289.5481.920

  195. 195.

    Noller HF. The driving force for molecular evolution of translation. RNA. 2004;10:1833-1837.

    Not in our library10.1261/rna.7142404

  196. 196.

    Noller HF. RNA structure: reading the ribosome. Science. 2005;309:1508-1514.

    Not in our library10.1126/science.1111771

  197. 197.

    Noller HF, Baucom A. Structure of the 70 S ribosome: implications for movement. Biochem Soc Trans. 2001;30:1159-1161.

    Not in our library10.1042/bst0301159

  198. 198.

    Noller HF, Woese CR. Secondary structure of 16S ribosomal RNA. Science. 1981;212:403-411.

    Not in our library10.1126/science.6163215

  199. 199.

    Noller HF, Kop J, Wheaton V, Brosius J, et al. Secondary structure model for 23S ribosomal RNA. Nucleic Acids Res. 1981;9:6167-6189.

    Not in our library10.1093/nar/9.22.6167

  200. 200.

    Ogino K, Edafujiwara H, Fujiwara H, Ishikawa H. What causes the aphid 28S ribosomal RNA to lack the hidden break? J Mol Evol. 1990;30:509-513.

    Not in our library10.1007/bf02101106

  201. 201.

    Ogle JM, Brodersen DE, Clemons WM, Tarry MT, Carter AP, Ramakrishnan V. Recognition of cognate transfer RNA by the 30S ribosomal subunit. Science. 2001;292:897-902.

    Not in our library10.1126/science.1060612

  202. 202.

    Ohnishi H, Yamamoto M-T. The structure of a single unit of ribosomal RNA gene (rDNA) including intergenic subrepeats in the Australian bulldog ant Myrmecia croslandi (Hymenoptera: Formicidae). Zoo Sci. 2004;21:139-146.

    Not in our library10.2108/zsj.21.139

  203. 203.

    Orlando TC, Rubio MA.T, Sturm NR, Campbell DA, Floeter-Winter LM. Intergenic and external transcribed spacers of ribosomal RNA genes in lizard-infecting Leishmania: molecular structure and phylogenetic relationship to mammal-infecting Leishmania in the subgenus Leishmania (Leishmania). Mem Inst Oswaldo Cruz, Rio Janeiro. 2002;97:1-7.

    Not in our library10.1590/s0074-02762002000500020

  204. 204.

    Osorio CR, Collins MD, Romalde JL, Toranzo AE. Variation in 16S−23S rRNA intergenic spacer regions in Photobacterium damselae: a mosaic-like structure. Appl Environ Microbiol. 2005;71:636-645.

    Not in our library10.1128/aem.71.2.636-645.2005

  205. 205.

    Ouvrard D, Campbell BC, Bourgoin T, Chan KL. 18S rRNA secondary structure and phylogenetic position of Peloridiidae (Insecta, Hemiptera). Mol Phylogenet Evol. 2000;16:403-417.

    Not in our library10.1006/mpev.2000.0797

  206. 206.

    Pace NR, Olsen GJ, Woese CR. Ribosomal RNA phylogeny and the primary lines of evolutionary descent. Cell. 1986;45:325-326.

    Not in our library10.1016/0092-8674(86)90315-6

  207. 207.

    Page RDM. Comparative analysis of insect mitochondrial small subunit ribosomal RNA using maximum weighted matching. Nucleic Acids Res. 2000;28:3839-3845.

    Not in our library10.1093/nar/28.20.3839

  208. 208.

    Page RDM. On the dangers of aligning RNA sequences using ‘conserved’ motifsTechnical reports in taxonomy.

    Not in our library

  209. 209.

    Page RD, Lee PL, Becher SA, Griffiths R, Clayton DH. A different tempo of mitochondrial DNA evolution in birds and their parasitic lice. Mol Phylogenet Evol. 1998;9:276-293.

    Not in our library10.1006/mpev.1997.0458

  210. 210.

    Page RD.M, Cruickshank R, Johnson KP. Louse (Insecta: Phthiraptera) mitochondrial 12S rRNA secondary structure is highly variable. Insect Mol Biol. 2002;11:361-369.

    Not in our library10.1046/j.1365-2583.2002.00346.x

  211. 211.

    Paques F, Samson ML, Jordan P, Wegnez M. Structural evolution of the Drosophila 5S ribosomal genes. J Mol Evol. 1995;41:615-621.

    Not in our library10.1007/bf00175820

  212. 212.

    Park Y-J, Fallon AM. Mosquito ribosomal RNA genes: characterization of gene structure and evidence for changes in copy number during development. Insect Biochem. 1990;20:1-11.

    Not in our library

  213. 213.

    Pepera OP, Cockburn AF, Mitchell SE, Conn J, Seawright JA. Species-specific repeat units in the intergenic spacer of the ribosomal RNA cistron of Anopheles aquasalis Curry. Am J Trop Med Hyg. 1998;59:673-678.

    Not in our library10.4269/ajtmh.1998.59.673

  214. 214.

    Perez-Gonzalez CE, Eickbush TH. Dynamics of R1 and R2 elements in the rDNA locus of Drosophila. Genetics. 2001;158:1557-1567.

    Not in our library10.1093/genetics/158.4.1557

  215. 215.

    Perez-Gonzalez CE, Eickbush TH. Rates of R1 and R2 retrotransposition and elimination from the rDNA locus of Drosophila melanogaster. Genetics. 2002;162:799-811.

    Not in our library10.1093/genetics/162.2.799

  216. 216.

    Pont-Kingdon GA, Beagley CT, Okimoto R, Wolstenholme DR. Mitochondrial DNA of the sea anemone, Metridum senile (Cnidaria): Prokaryote-like genes for tRNA (f-Met) and small-subunit ribosomal RNA, and standard genetic code specificities for AGR and ATA codons. J Mol Evol. 1994;39:387-399.

    Not in our library10.1007/bf00160271

  217. 217.

    Rambaut A. Se-Al: Sequence Alignment Editor.

    Not in our library

  218. 218.

    Reese MG. Application of a time-delay neural network to promoter annotation in the Drosophila melanogaster genome. Comput Chem. 2001;26:51-56.

    Not in our library10.1016/s0097-8485(01)00099-7

  219. 219.

    Rimoldi OJ, Raghu B, Mag MK, Eliceiri GL. Three new small nucleolar RNAs that are psoralen cross-linked in vivo to unique regions of pre-rRNA. Mol Cell Biol. 1993;13:4382-4390.

    Not in our library10.1128/mcb.13.7.4382-4390.1993

  220. 220.

    Rocheford TR. Change in ribosomal DNA intergenic spacer-length composition in maize recurrent selection populations. 1. Analysis of BS13, BSSS, and BSCB1. Theor Appl Genet. 1994;88:541-547.

    Not in our library10.1007/bf01240916

  221. 221.

    Roiha H, Miller RM, Woods LC, Glover DM. Arrangements and rearrangements of sequences flanking the two types of rDNA insertion in D. melanogaster. Nature. 1981;290:749-753.

    Not in our library10.1038/290749a0

  222. 222.

    Ruschak AM, Mathews DH, Bibillo A, Spinelli SL, Childs JL, Eickbush TH, Turner DH. Secondary structure models of the 3′ untranslated regions of diverse R2 RNAs. RNA. 2004;10:978-987.

    Not in our library10.1261/rna.5216204

  223. 223.

    Sassaman DM, Dombroski BA, Moran JV, Kinberland ML, et al. Many human L1 elements are capable of retrotransposition. Nat Genet. 1997;16:37-43.

    Not in our library10.1038/ng0597-37

  224. 224.

    Schluenzen F, Tocilj A, Zarivach R, Harms J, et al. Structure of functionally activated small ribosomal subunit at 3.3 Å resolution. Cell. 2000;102:615-623.

    Not in our library10.1016/s0092-8674(00)00084-2

  225. 225.

    Schnare MN, Collings JC, Spencer DF, Gray MW. The 28S−18S rDNA intergenic spacer from Crithidia fasciculata: repeated sequences, length heterogeneity, putative processing sites and potential interactions between U3 and small nucleolar RNA and the ribosomal precursor. Nucleic Acids Res. 2000;28:3452-3461.

    Not in our library10.1093/nar/28.18.3452

  226. 226.

    Schnare MN, Damberger SH, Gray MW, Gutell RR. Comprehensive comparison of structural characteristics in eukaryotic cytoplasmic large subunit (23S-like) ribosomal RNA. J Mol Biol. 1996;256:701-719.

    Not in our library10.1006/jmbi.1996.0119

  227. 227.

    Schulmeister S. Simultaneous analysis of basal Hymenoptera (Insecta): Introducing robust-choice sensitivity analysis. Biol J Linn Soc. 2003;79:245-275.

    Not in our library

  228. 228.

    Shao R, Barker SC. The highly rearranged mitochondrial genome of the plague thrips, Thrips imaginis (Insecta: Thysanoptera): convergence of two novel gene boundaries and an extraordinary arrangement of rRNA genes. Mol Biol Evol. 2003;20:362-370.

    Not in our library10.1093/molbev/msg045

  229. 229.

    Shao R, Campbell NJH, Barker SC. Numerous gene rearrangements in the mitochondrial genome of the wallaby louse, Heterodoxus macropus (Phthiraptera). Mol Biol Evol. 2001a;18:858-865.

    Not in our library10.1093/oxfordjournals.molbev.a003867

  230. 230.

    Shao R, Campbell NJH, Schmidt ER, Barker SC. Increased rate of gene rearrangement in the mitochondrial genomes of three orders of hemipteroid insects. Mol Biol Evol. 2001b;18:1828-1832.

    Not in our library10.1093/oxfordjournals.molbev.a003970

  231. 231.

    Shao R, Dowton M, Murrell A, Barker SC. Rates of gene rearrangement and nucleotide substitution are correlated in the mitochondrial genomes of insects. Mol Biol Evol. 2003;20:1612-1619.

    Not in our library10.1093/molbev/msg176

  232. 232.

    Shine J, Dalgarno L. Occurrence of heat-dissociable ribosomal RNA in insects: presence of 3 polynucleotide chains in S26 RNA from cultured Aedes aegypti cells. J Mol Biol. 1973;75:57-72.

    Not in our library10.1016/0022-2836(73)90528-7

  233. 233.

    Simeone A, La Volpe A, Boncinelli E. Nucleotide sequence of a complete ribosomal spacer of D. melanogaster. Nucleic Acids Res. 1985;13:1089-1101.

    Not in our library10.1093/nar/13.4.1089

  234. 234.

    Simon C, Hewitt GM, Johnston AWB, Young JPW. Molecular systematics at the species boundary. Exploiting Conserved and Variable Regions of the Mitochondrial Genome of Animals Via Direct Sequencing from Amplified DNA. 1991;33-71.

    Not in our library

  235. 235.

    Simon C, Frati F, Beckenbach A, Crespi B, Liu H, Flook P. Evolution, weighting, and phylogenetic utility of mitochondrial gene sequences and a compilation of conserved polymerase chain reaction primers. Ann Entomol Soc Am. 1994;87:651-701.

    Not in our library

  236. 236.

    Simon C, Paabo S, Kocher T, Wilson AC, Cleggs M, O’brien S. Evolution of the mitochondrial ribosomal RNA in insects as shown by the polymerase chain reaction. Molecular Evolution. 1990;235-244.

    Not in our library

  237. 237.

    Smith SD, Bond JE. An analysis of the secondary structure of the mitochondrial large subunit rRNA gene (16S) in spiders and its implications for phylogenetic reconstruction. J Arach. 2003;31:44-54.

    Not in our library

  238. 238.

    Spahn CM, Beckmann R, Eswar N, Penczek PA, Sali A, Blobel G, Frank J. Structure of the 80S ribosome from Saccharomyces cerevisiae-tRNA-ribosome and subunit–subunit interactions. Cell. 2001;107:373-386.

    Not in our library10.1016/s0092-8674(01)00539-6

  239. 239.

    States DJ. Molecular sequence accuracy: analysing imperfect data. Trends Genet. 1992;8:52-55.

    Not in our library10.1016/0168-9525(92)90349-9

  240. 240.

    Stiegler P, Carbon P, Ebel JP, Ehresmann C. A general secondary-structure model for procaryotic and eucaryotic RNAs from the small ribosomal subunits. Eur J Biochem. 1981;120:487-495.

    Not in our library10.1111/j.1432-1033.1981.tb05727.x

  241. 241.

    Stuart GW, Berry MW. An SVD-based comparison of nine whole eukaryotic genomes supports a coelomate rather than ecdysozoan lineage. BMC Bioinformatics. 2004;5:204.

    Not in our library10.1186/1471-2105-5-204

  242. 242.

    Sullivan J, Holsinger KE, Simon C. Among-site rate variation and phylogenetic analysis of 12S rRNA in sigmodontine rodents. Mol Biol Evol. 1995;12:988-1001.

    Not in our library10.1093/oxfordjournals.molbev.a040292

  243. 243.

    Sweeney R, Yao M-C. Identifying functional regions of rRNA by insertion mutagenesis and complete gene replacement in Tetrahymena thermophila. EMBO J. 1989;8:933-938.

    Not in our library10.1002/j.1460-2075.1989.tb03454.x

  244. 244.

    Sweeney R, Chen L, Yao M-C. An rRNA variable region has an evolutionary conserved essential role despite sequence divergence. Mol Cell Biol. 1994;14:4203-4215.

    Not in our library10.1128/mcb.14.6.4203-4215.1994

  245. 245.

    Swigonova Z, Kjer KM. Phylogeny and host–plant association in the leaf beetle genus Trirhabda LeConte (Coleoptera: Chrysomelidae). Mol Phylogenet Evol. 2004;32:358-374.

    Not in our library10.1016/j.ympev.2004.02.010

  246. 246.

    Szymanski M, Barciszewska MZ, Erdmann VA, Szymanski JB. 5S ribosomal RNA database. Nucleic Acids Res. 2002;30:176-178.

    Not in our library10.1093/nar/30.1.176

  247. 247.

    Tan THP, Pach R, Crausaz A, Ivens A, Schneider A. tRNAs in Trypanosoma brucei: genomic organization, expression, and mitochondrial import. Mol Cell Biol. 2002;22:3707-3717.

    Not in our library10.1128/mcb.22.11.3707-3716.2002

  248. 248.

    Tautz D, Hancock JM, Webb DA, Tautz C, Dover GA. Complete sequences of the rRNA genes of Drosophila melanogaster. Mol Biol Evol. 1988;5:366-376.

    Not in our library10.1093/oxfordjournals.molbev.a040500

  249. 249.

    Tautz D, Tautz C, Webb D, Dover GA. Evolutionary divergence of promoters and spacers in the rDNA family of four Drosophila species. J Mol Biol. 1987;195:525-542.

    Not in our library10.1016/0022-2836(87)90181-1

  250. 250.

    Insights into social insects from the genome of the honeybee Apis mellifera. Nature. 2006.

    Not in our library10.1038/nature05260

  251. 251.

    Thomas WK, Maa J, Wilson AC. Shifting constraints on tRNA genes during mitochondrial DNA evolution in animals. New Biologist. 1989;1:93-100.

    Not in our library

  252. 252.

    Uhlenbusch I, McCracken A, Gellissen G. The gene for the large (16S) ribosomal RNA from the Locusta migratoria mitochondrial genome. Curr Genet. 1987;11:631-638.

    Not in our library10.1007/bf00393927

  253. 253.

    Van de Peer Y, Neefs J-M, De Rijk P, De Wachter R. Reconstructing evolution from eukaryotic small-ribosomal-subunit RNA sequences: calibration of the molecular clock. J Mol Evol. 1993;37:221-232.

    Not in our library10.1007/bf02407359

  254. 254.

    Van de Peer Y, van den Broeck, De Rijk P, De Wachter R. Database on the structure of small ribosomal subunit RNA. Nucleic Acids Res. 1994;22:3488-3392.

    Not in our library10.1093/nar/22.17.3488

  255. 255.

    Van de Peer Y, Nicolai S, De Wachter R. Database on the structure of small ribosomal subunit RNA. Nucleic Acids Res. 1996;24:86-91.

    Not in our library10.1093/nar/24.1.86

  256. 256.

    Van de Peer Y, Jansen J, De Rijk P, De Wachter R. Database on the structure of small ribosomal subunit RNA. Nucleic Acids Res. 1997;25:111-116.

    Not in our library10.1093/nar/25.1.111

  257. 257.

    Van de Peer Y, Caers A, De Rijk P, De Wachter R. Database on the structure of small ribosomal subunit RNA. Nucleic Acids Res. 1998;26:179-182.

    Not in our library

  258. 258.

    Van de Peer Y, Robbrecht E, de Hoog S, Caers A, De Rijk P, De Wachter R. Database on the structure of small subunit ribosomal RNA. Nucleic Acids Res. 1999;27:179-183.

    Not in our library10.1093/nar/27.1.179

  259. 259.

    Van de Peer Y, De Rijk P, Wuyts J, Winkelmans T, De Wachter R. The European Small Subunit Ribosomal RNA database. Nucleic Acids Res. 2000;28:175-176.

    Not in our library10.1093/nar/28.1.175

  260. 260.

    Varricchio P, Gargiulo G, Graziani F, Manzi A, Pennacchio F, Digilio M, Tremblay E, Malva C. Characterization of Aphidius ervi (Hymenoptera: Braconidae) ribosomal genes and identification of site-specific insertion elements belonging to the non-LTR retrotransposon family. Insect Biochem Mol Biol. 1994;25:603-612.

    Not in our library10.1016/0965-1748(94)00102-n

  261. 261.

    Vawter L, Brown WM. Rates and patterns of base change in the SSU ribosomal RNA gene. Genetics. 1993;134:597-608.

    Not in our library10.1093/genetics/134.2.597

  262. 262.

    Veldman GM, Klootwijk J, De Regt VCFH, Planta RJ, Branlant C, Krol A, Ebel J-P. The primary and secondary structure of yeast 26S rRNA. Nucleic Acids Res. 1981;9:6935-6952.

    Not in our library10.1093/nar/9.24.6935

  263. 263.

    Vlasak I, Burgschwaiger S, Kreil G. Nucleotide sequence of the large ribosomal RNA of honeybee mitochondria. Nucleic Acids Res. 1987;15:2388.

    Not in our library10.1093/nar/15.5.2388

  264. 264.

    Ware VC, Tague BW, Clark CG, Gourse RL, Brand RC, Gerbi SA. Sequence analysis of 28S ribosomal DNA from the amphibian Xenopus laevis. Nucleic Acids Res. 1983;11:7795-7817.

    Not in our library10.1093/nar/11.22.7795

  265. 265.

    Ware VC, Renkawitz R, Gerbi SA. rRNA processing: removal of only nineteen bases at the gap between 28S alpha and 28S beta rRNAs in Sciara coprophila. Nucleic Acids Res. 1985;13:3581-3597.

    Not in our library10.1093/nar/13.10.3581

  266. 266.

    Weider LJ, Elser JJ, Crease TJ, Mateos M, Cotner JB, Markow TA. The functional significance of ribosomal (r) DNA variation: Impacts on the evolutionary ecology of organisms. Annu Rev Ecol Evol Syst. 2005;36:219-242.

    Not in our library

  267. 267.

    Weiner AM, Deininger PL, Efstratiadis A. Nonviral retroposons: genes, pseudogenes, and transposable elements generated by the reverse flow of genetic information. Annu Rev Biochem. 1986;55:631-661.

    Not in our library10.1146/annurev.bi.55.070186.003215

  268. 268.

    Whang I-J, Jung J, Par J-K, Min G-S, Kim W. Intragenomic length variation of the ribosomal DNA intergenic spacer in a malaria vector, Anopheles sinensis. Mol Cells. 2002;14:158-162.

    Not in our library

  269. 269.

    Wharton RA, Yoder MJ, Gillespie JJ, Patton JC, Honeycutt RL. Relationships of Exodontiella, a non-alysiine, exodont member of the family Braconidae (Insecta, Hymenoptera). Zool Scripta. 2006;35:323-340.

    Not in our library

  270. 270.

    Wheeler WC, Whiting MF, Wheeler QD, Carpenter JC. The phylogeny of extant insect orders. Cladistics. 2001;17:113-169.

    Not in our library10.1111/j.1096-0031.2001.tb00115.x

  271. 271.

    Whiting MF, Carpenter JC, Wheeler QD, Wheeler WC. The Strepsiptera problem: phylogeny of the holometabolous insect orders inferred from 18S and 28S ribosomal DNA sequences and morphology. Syst Biol. 1997;46:1-68.

    Not in our library10.1093/sysbio/46.1.1

  272. 272.

    Wiegmann BM, Yeates DK, Thorne JL, Kishino H. Time flies, a new molecular time-scale for brachyceran fly evolution without a clock. Syst Biol. 2003;52:745-756.

    Not in our library

  273. 273.

    Wimberly BT, Guymon R, McCutcheon JP, White SW, Ramakrishnan V. A detailed view of a ribosomal active site: the structure of the L11-RNA complex. Cell. 1999;97:491-502.

    Not in our library10.1016/s0092-8674(00)80759-x

  274. 274.

    Wimberly BT, Brodersen DE, Clemons WM, Morgan-Warren RJ, et al. Structure of the 30S ribosomal subunit. Nature. 2000;407:327-339.

    Not in our library10.1038/35030006

  275. 275.

    Winker S, Woese CR. A definition of the domains Archaea, Bacteria and Eucarya, in terms of small subunit ribosomal RNA characteristics. Syst Appl Microbiol. 1991;14:305-310.

    Not in our library10.1016/s0723-2020(11)80303-6

  276. 276.

    Woese CR. Bacterial evolution. Microbiol Rev. 1987;51:221-271.

    Not in our library10.1128/mr.51.2.221-271.1987

  277. 277.

    Woese CR, Pace NR, Gesteland RF, Atkins JF. Probing RNA structure, function and history by comparative analysis. The RNA World. 1993;91-117.

    Not in our library

  278. 278.

    Woese CR, Magrum LJ, Gupta R, Siegel RB, Stahl DA, Kop J, Crawford N, Brosius J, Gutell RR, Hogan JJ, Noller HF. Secondary structure model for bacterial 16S ribosomal RNA: phylogenetic, enzymatic and chemical evidence. Nucleic Acids Res. 1980;8:2275-2293.

    Not in our library10.1093/nar/8.10.2275

  279. 279.

    Woese CR, Kandler O, Wheelis ML. Towards a natural system of organisms: Archaea, Bacteria, Eucarya. Proc Natl Acad Sci USA. 1990;87:4576-4579.

    Not in our library10.1073/pnas.87.12.4576

  280. 280.

    Woods PE, Guttman SI. Genetic variation in Neodiprion (Hymneoptera: Symphyta: Diprionnidae) sawflies and a comment on low levels of genetic diversity within Hymenoptera. Ann Entomol Soc Am. 1987;80:590-599.

    Not in our library

  281. 281.

    Wool IG, Hardesty J, Kramer G. Studies of the structure of eukaryotic (mammalian) ribosomes. Structure, Function and Genetics of Ribosomes. 1986;391-411.

    Not in our library

  282. 282.

    Wu CC, Fallon AM. Analysis of a ribosomal DNA intergenic spacer region from the yellow fever mosquito, Aedes aegypti. Insect Mol Biol. 1998;7:19-29.

    Not in our library10.1046/j.1365-2583.1998.71194.x

  283. 283.

    Wuyts J, De Rijk P, Van de Peer Y, Pison G, Rousseeuw P, De Wachter R. Comparative analysis of more than 3000 sequences reveals the existence of two pseudoknots in area V4 of eukaryotic small subunit ribosomal RNA. Nucleic Acids Res. 2000;28:4698-4708.

    Not in our library10.1093/nar/28.23.4698

  284. 284.

    Wuyts J, De Rijk P, Van de Peer Y, Winkelmans T, De Wachter R. The European large subunit ribosomal RNA database. Nucleic Acids Res. 2001;29:175-177.

    Not in our library10.1093/nar/29.1.175

  285. 285.

    Wuyts J, Van de Peer Y, Winkelmans T, De Wachter R. The European database on small subunit ribosomal RNA. Nucleic Acids Res. 2002;30:183-185.

    Not in our library10.1093/nar/30.1.183

  286. 286.

    Wuyts J, Perriere G, Van de Peer Y. The European ribosomal RNA database. Nucleic Acids Res. 2004;32:D101-D103.

    Not in our library10.1093/nar/gkh065

  287. 287.

    Xiong Y, Eickbush TH. Origin and evolution of retroelements based upon their reverse transcriptase sequences. EMBO J. 1990;9:3353-3362.

    Not in our library10.1002/j.1460-2075.1990.tb07536.x

  288. 288.

    Yoshionari S, Koike T, Yokogawa T, Nishikawa K, Ueda T, Miura K, Watanabe K. Existence of nuclear-encoded 5S rRNA in bovine mitochondria. FEBS Lett. 1994;338:137-142.

    Not in our library10.1016/0014-5793(94)80351-x

  289. 289.

    Yoshizawa K, Johnson KP. Aligned 18S for Zoraptera (Insecta): phylogenetic position and molecular evolution. Mol Phylogenet Evol. 2005;37:572-580.

    Not in our library10.1016/j.ympev.2005.05.008

  290. 290.

    Yusupov MM, Yusupova GZ, Baucom A, Lieberman K, Earnest TN, Cate JH, Noller HF. Crystal structure of the ribosome at 5.5 Å resolution. Science. 2001;292:883-896.

    Not in our library10.1126/science.1060089

  291. 291.

    Zarlenga DS, Dame JB. The identification and characterization of a break within the large subunit ribosomal RNA of Trichinella spiralis: comparison of gap sequences within the genus. Mol Biochem Parasitol. 1992;51:281-290.

    Not in our library10.1016/0166-6851(92)90078-x

  292. 292.

    Zuker M. Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res. 2003;31:3406-3415.

    Not in our library10.1093/nar/gkg595

  293. 293.

    Zwieb C, Glotz C, Brimacombe R. Secondary structure comparisons between SSU ribosomal RNA molecules from six different species. Nucleic Acids Res. 1981;9:3621-3640.

    Not in our library10.1093/nar/9.15.3621

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