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. 2014 Jun 13;197(4):1201–1211. doi: 10.1534/genetics.114.162875

Operons Are a Conserved Feature of Nematode Genomes

Jonathan Pettitt 1,1, Lucas Philippe 1, Debjani Sarkar 1, Christopher Johnston 1, Henrike Johanna Gothe 1, Diane Massie 1, Bernadette Connolly 1, Berndt Müller 1
PMCID: PMC4125394  PMID: 24931407

Abstract

The organization of genes into operons, clusters of genes that are co-transcribed to produce polycistronic pre-mRNAs, is a trait found in a wide range of eukaryotic groups, including multiple animal phyla. Operons are present in the class Chromadorea, one of the two main nematode classes, but their distribution in the other class, the Enoplea, is not known. We have surveyed the genomes of Trichinella spiralis, Trichuris muris, and Romanomermis culicivorax and identified the first putative operons in members of the Enoplea. Consistent with the mechanism of polycistronic RNA resolution in other nematodes, the mRNAs produced by genes downstream of the first gene in the T. spiralis and T. muris operons are trans-spliced to spliced leader RNAs, and we are able to detect polycistronic RNAs derived from these operons. Importantly, a putative intercistronic region from one of these potential enoplean operons confers polycistronic processing activity when expressed as part of a chimeric operon in Caenorhabditis elegans. We find that T. spiralis genes located in operons have an increased likelihood of having operonic C. elegans homologs. However, operon structure in terms of synteny and gene content is not tightly conserved between the two taxa, consistent with models of operon evolution. We have nevertheless identified putative operons conserved between Enoplea and Chromadorea. Our data suggest that operons and “spliced leader” (SL) trans-splicing predate the radiation of the nematode phylum, an inference which is supported by the phylogenetic profile of proteins known to be involved in nematode SL trans-splicing.

Keywords: operons, SL trans-splicing, nematode genome

THE organization of open-reading frames into operons, such that multiple, distinct gene products are produced from a single, polycistronic transcript, is commonplace in prokaryote genomes (Jacob et al. 1960). Operons are also found in eukaryotes, although their distribution is sporadic and it does not seem likely that they represent an ancestral eukaryotic trait (Lawrence 1999; Hastings 2005). In prokaryotes, translation of multiple open reading frames in a polycistronic RNA occurs through multiple independent translation initiations. In eukaryotes, the polycistronic RNAs must first be processed into individual mRNAs before being translated. This creates a problem in that the processed, downstream mRNAs would lack a cap structure necessary for RNA stability and translation. A number of eukaryotes are able to circumvent this problem through the trans-splicing of a short “spliced leader” (SL) RNA onto the 5′ end of the mRNA. Because the precursor SL RNAs that donate the SL are trimethylguanosine-capped, the trans-splicing event provides the cap structure for the mRNA. Thus by providing a mechanism that allows the formation of monocistronic, capped mRNAs from polycistronic RNA, SL trans-splicing enables the organization of eukaryotic genes into operons. It is striking that, at least to date, all eukaryotes in which operon usage is widespread also undergo SL trans-splicing (Johnson et al. 1987; Spieth et al. 1993; Davis and Hodgson 1997; Blumenthal et al. 2002; Ganot et al. 2004; Guiliano and Blaxter 2006; Satou et al. 2008; Marlétaz et al. 2008; Dana et al. 2012; Protasio et al. 2012; Tsai et al. 2013), suggesting that the resolution of polycistronic RNA is dependent upon SL trans-splicing.

Although operon organization is widespread in numerous eukaryotic taxa, the evolutionary mechanisms that have resulted in this form of gene organization are not well understood. The most detailed analysis of the problem has come from studies in Caenorhabditis elegans, which led to the hypothesis that operon organization allows the marshalling of multiple genes under the control of a single promoter. This makes cells better able to cope with situations when transcription factors are present in limiting concentrations, such as recovery from growth arrest (Zaslaver et al. 2011). However, it is far from clear whether this is the only mechanism responsible for the evolution of operon organization, and the general applicability of this hypothesis to other members of the nematode phylum is not known.

To better understand the relationship between operon evolution and SL trans-splicing, it is necessary to determine the distribution of operon organization across the nematode phylum. Nematodes can be divided into two major classes: Enoplea and Chromadorea (Holterman et al. 2006; Meldal et al. 2007), with the latter class being much better characterized in terms of gene expression mechanisms, largely because it contains C. elegans. Both SL trans-splicing and operons have been identified in multiple nematodes within the Chromadorea (Evans et al. 1997; Lee and Sommer 2003; Guiliano and Blaxter 2006). However, the presence of operons has not been reported in nematodes from the other taxon.

We have previously identified SL trans-splicing in the enoplean nematodes Trichinella spiralis and Prionchulus punctatus (Pettitt et al. 2008; Harrison et al. 2010), suggesting that they may also possess operons. The draft genome of T. spiralis should be a useful resource for identifying operons in this nematode (Mitreva et al. 2011). However, identification of operons is not straightforward. The original discovery of operons in C. elegans was dependent upon the discovery of a specific spliced leader, SL2, which is trans-spliced to most mRNAs derived from genes downstream of the first gene in operons (Spieth et al. 1993), but not all nematodes use a specialized SL RNA to resolve polycistronic RNAs (Guiliano and Blaxter 2006). Thus, this feature cannot be considered diagnostic for mRNAs derived from nematode operons. The other feature common to operonic genes is that, at least in C. elegans, the distance between genes in an operon (the intercistronic region, ICR) is unusually short, with a mean ICR size of 126 bp (Blumenthal et al. 2002). Again, this trait is not definitive: the ICR size can be considerably larger in the operons of other nematodes (Guiliano and Blaxter 2006; Ghedin et al. 2007) and even in C. elegans operons exist with large ICR distances (Morton and Blumenthal 2011).

Previous approaches to identify operons in T. spiralis (Mitreva et al. 2011) looked for pairs of T. spiralis genes whose homologs were in the same operon in C. elegans. This resulted in a limited set of 16 neighboring pairs of genes that potentially correspond to T. spiralis operons; however, further characterization of these candidate operons was not undertaken. We have used conserved synteny, coupled with the fact that mRNAs derived from downstream genes in operons are dependent on SL trans-splicing to elucidate a set of putative T. spiralis operons. Detailed analysis of two of these putative operons indicates that they display all the molecular characteristics expected of loci that generate polycistronic RNA. Taken together our data indicate that the organization of genes into operons was present in the last common ancestor of the Chromadorea and Enoplea.

Materials and Methods

Bioinformatic identification of T. muris SL RNA genes

Trichuris muris SL RNA genes Tmu-SL1, Tmu-SL2, Tmu-SL3, and Tmu-SL9 were identified by searching the T. muris genome dataset with T. spiralis SL sequences using the BLASTN tool. Hits were considered if two of the three following criteria were met: a candidate Sm protein binding site was detected (AATTTTTG), the 5′ splice site sequence was conserved (AGGT), and a run of at least three Ts was found located ∼100 bp from the end of the putative SL sequence. T. muris SL RNA genes Tmu-SL4, Tmu-SL5, Tmu-SL6, Tmu-SL7, Tmu-SL8, and Tmu-SL10 were identified by searching the T. muris genome dataset with the Tmu-SL1, Tmu-SL2, Tmu-SL3, or Tmu-SL9 sequences using the BLASTN tool and fulfilling the same criteria as above. Genes for Tmu-SL1, Tmu-SL2, Tmu-SL6, Tmu-SL8, Tmu-SL9, Tmu-SL10, and Tmu-SL11 were also identified with a PERL script (Pettitt et al. 2008) used previously to identify T. spiralis SL genes, except that the parameters for the Sm binding site were changed to AATTTTTG/TG.

Analysis of T. spiralis SL containing ESTs and Identification of Putative Conserved Operons

The ESTs were identified earlier (Pettitt et al. 2008). To identify the corresponding gene from which each EST was derived, the EST sequences were mapped onto the T. spiralis draft genome sequencing using BLASTN. The corresponding gene was annotated as being in a putative operon if its upstream or downstream neighbor genes were on the same DNA strand with an intercistronic distance of ≤1 kb. If the neighbors were on the same strand, but between 1 kb and 5 kb away, they were recorded as ambiguous. Otherwise the genes were annotated as nonoperonic. A minority of the ESTs matched to more than one predicted T. spiralis gene.

To identify the operonic status of the C. elegans homologs of each SL trans-spliced T. spiralis EST, BLASTX searches were carried out. We used an E-value cutoff of 10−5 to determine homology. In addition, if we obtained similar E-values for multiple C. elegans genes, we excluded that EST from the analysis.

T. spiralis and Romanomermis culicivorax homologs of operonic gene pairs conserved between C. elegans and Brugia malayi (Ghedin et al. 2007) were identified using BLASTP searches with the C. elegans upstream homologs from each pair as a query in searches against the respective gene predictions. The predicted coding region of the T. spiralis/R. culicivorax gene immediately downstream of the gene identified by this search was then used as a query sequence in a “reciprocal” BLASTP search against the C. elegans gene predictions. Since it was apparent that T. spiralis genes, which are separated by unusually short intergenic distance (such as might be expected in genes organized into operons), are prone to misannotation and conflation into a single gene prediction, we also carried out an additional step using both C. elegans gene pairs as query sequences in BLASTP searches against the T. spiralis gene predictions, looking for cases where both C. elegans genes return matches to the same T. spiralis gene. Manual examination of such putative gene prediction errors, guided by the results of the sequence similarity searches, was then used to identify the intercistronic regions in each case. In all BLASTP searches, we used an E-value cutoff of 10−5 for the establishment of homology.

Phylogenetic profiling of SL trans-splicing snRNPs

Homologs of C. elegans sna-1, sut-1, and sna-2 were identified by carrying out BLAST searches against the National Center for Biotechnology Information (NCBI) nonredundant database, except in the case of R. culicivorax and T. muris, where BLAST searches were carried out against datasets downloaded from http://www.nematodes.org/genomes/romanomermis_culicivorax/ and http://www.sanger.ac.uk/resources/downloads/helminths/trichuris-muris.html, respectively. Phylogenetic tree construction was carried out with the online implementation of PhyML (Dereeper et al. 2008) using default settings.

Nematode isolation and RNA preparation

T. spiralis RNA was produced as described (Pettitt et al. 2008). T. muris RNA was a generous gift from Allison Bancroft and Richard Grencis (University of Manchester).

Analysis of RNA 5′ ends

The 5′ ends of cDNAs were obtained through 5′ RACE using the GeneRacer kit (Invitrogen), according to the manufacturer’s instructions. Gene-specific primers used are given in Supporting Information, Table S3, and the cDNAs, amplified by PCR, using either GoTaq polymerase (Promega) or Expand High Fidelity polymerase (Roche), were cloned into pGEM T-Easy (Promega). The resulting plasmid inserts were sequenced by the University of Dundee Sequencing Service.

Detection of processing intermediates of polycistronic transcripts

RNA was reverse transcribed using SuperScript III Reverse Transcriptase (Invitrogen) and random primers according to the instructions of the manufacturer. In control reactions (“−RT”) all reagents were included except the reverse transcriptase. Processing intermediates were normally amplified by two rounds of PCR with nested primer pairs (Table S4) and either GoTaq polymerase (Promega) or Expand High Fidelity polymerase (Roche) and visualized by agarose gel electrophoresis. The identity of the PCR products was determined by cloning into pGEM-T Easy (Promega) and sequencing of plasmid inserts.

Identification of SL RNA 3′ ends

SL RNA 3′ ends were determined essentially as described previously (Pettitt et al. 2008). T. muris total RNA (∼5 µg) was poly(A) tailed using yeast poly(A) polymerase, reverse transcribed using an oligo-dT-anchor primer (GCGAGCTCCGCGGCCGCGTTTTTTTTTTTTTTT) and then PCR amplified using an SL-specific primers (GGTTAATTACCCAATTTAAAAG) and an anchor primer (GCGAGCTCCGCGGCCGCG). PCR fragments were inserted into pGEM-T Easy (Promega), and inserts were sequenced at the University of Dundee DNA Sequencing Facility.

SL RNA secondary structure prediction:

Secondary structure prediction of T. muris SL RNA was performed using MFOLD Version 2.3 (Zuker 2003) using the default folding conditions (1 M NaCl, 37°) and with the constraint that the Sm-binding site (5′-AAUUUUUUG-3′) was required to be single stranded.

Generation and analysis of synthetic operon constructs

The GFP coding region was amplified from pTG96 using the primers 5′-CAATACAGACTTCCCGGGATTGGCCAAAGGACCCAAA-3′ and 5′-GCTCACCATGCTAGCCTATTTGTATAGTTCATCCATGC-3′. The mCherry coding region, coupled to the unc-54 3′-UTR, was amplified from pPD95.75Cherry (a derivative of pPD95.75 in which the GFP coding region was replaced by mCherry) using the primers 5′-ACAAATAGGCTAGCATGGTGAGCAAGGGCGAG-3′ and 5′-CGCGCGAGACGAAAGGGCCCAGGAAACAGTTATGTTTGGTAT-3′. The primers were designed so that they had overlapping complementary 5′ extensions that introduced an NheI restriction site. The two amplicons were purified and fused using a PCR fusion strategy (Hobert 2002). The resulting amplicon consisting of the GFP and mCherry coding regions flanking an NheI site was cloned into SmaI–ApaI cut pTG96 using In-Fusion HD (Clontech Laboratories) to generate pTG96-Op. The ICRs were cloned from PCR products amplified from genomic DNA. The Tsp-cpt-2∼nuaf-3 ICR was amplified from T. spiralis genomic DNA using primers 5′-ATACAAATAGGCTAGCACGAATTATCACTTTTATAAC-3′ and 5′-TGCTCACCATGCTAGCTTACGCCAAACTAGGAAATTATTGA-3′, and the Cel-cpt-2∼prx-14 ICR was amplified from C. elegans genomic DNA using primers 5′-ATACAAATAGGCTAGCTTGTTTGATGACATTTATGTATTTAT-3′ and 5′-TGCTCACCATGCTAGCTTTCAACCTGAAGCTTTAAAAT-3′. The resulting PCR products were cloned into NheI cut pTG96-Op using In-Fusion HD (Clontech Laboratories) and the resultant plasmids, pPE#LP1 (Tsp-cpt-2∼nuaf-3 ICR clone) and pPE#LP2 (Cel-cpt-2∼prx-14 ICR clone) were sequenced to confirm the integrity of the cloning process. To generate transgenic C. elegans strains, the plasmids were co-injected (100 ng/μl) with Pmyo-2::dTomato (10 ng/μl) into Bristol (N2) wild-type hermaphrodites. For each construct, several lines were obtained, each of which gave identical expression patterns. Single lines for each construct were selected for the experiments reported here: PE612, feEx304 [sur-5::gfp::ICRTsp-cpt-2∼nuaf-3::mCherry Pmyo-2::dTomato] and PE613, feEx305 [sur-5::gfp::ICRCel-cpt2∼prx-14::mCherry Pmyo-2::dTomato]. Trans-splicing of reporter gene transcripts was analyzed as described previously (Harrison et al. 2010). Briefly, total RNA was reverse transcribed and trans-spliced transcripts were PCR amplified using C. elegans SL2-specific (5′-GGTTTTAACCCAGTTACTCAAG-3′) and mCherry-specific (5′-CCGTCCTCGAAGTTCATCAC-3′) primers. Primers derived from gpd-1 (5′-CCAACTGTCTGGCACCACT-3′ and 5′-GTCTTCTGGGTTGCGGTTAC-3′) were used to normalize the reactions. cDNA fragments were cloned into pGEM-T Easy (Promega), and inserts were sequenced at the University of Dundee DNA Sequencing Facility.

Results

A putative enoplean operon

As part of the analysis of the transcriptome of the free-living enoplean, Prionchulus punctatus, we identified an EST corresponding to an SL trans-spliced mRNA. Sequence similarity searches using this sequence identified a single predicted T. spiralis gene, Tsp_06075. However, subsequent sequence analysis of Tsp_06075 showed that it corresponds to an erroneous gene prediction, which conflates three genes that are the orthologs of the C. elegans genes zgpa-1 (C33H5.17), dif-1, and aph-1, respectively. That Tsp_06075 is actually three separate genes was confirmed by sequence analysis of 5′ RACE products. It seems likely that the unusually short intergenic regions that exist between these three T. spiralis genes caused the gene annotation error (Figure 1). Such short intergenic distances are characteristic of nematode genes that are arranged into operons (Blumenthal et al. 2002; Guiliano and Blaxter 2006; Ghedin et al. 2007), and we thus decided to investigate the possibility that Tsp-zgpa-1, Tsp-dif-1, and Tsp-aph-1 constitute an operon. In parallel, we also analyzed the homologs of these three genes in the closely related enoplean, T. muris, which show the same syntenic arrangement, although the intergenic distance between Tmu-dif-1 and Tmu-aph-1 is much larger than expected for an ICR. The three C. elegans homologs although organized into operons, are not found in the same operon. However, in a close relative of C. elegans, Pristionchus pacificus, zgpa-1 and dif-1 could potentially constitute a single operon, but again the intergenic space between the two genes is also relatively large compared to the average size of ICRs in C. elegans operons.

Figure 1.

Figure 1

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Evidence for the existence of an enoplean operon. (A) Schematic showing the genomic organization of zgpa-1, dif-1, and aph-1 in selected nematodes mapped onto their phylogenetic relationships. Arrows represent genes, and the gray lines represent the intercistronic regions (ICRs). Numbers above the ICRs represent the distances, in base pairs, between the stop and start codons of the upstream and downstream genes, respectively. The C. elegans operon numbers are given where appropriate. Fractions below the T. spiralis and T. muris genes represent the proportion of cDNAs derived from those genes that begins with a spliced leader sequence (see also Table S2). In C. elegans, the three genes are part of different operons. * indicates the distance between genes on chromosome IV. (B) Detecting polycistronic RNAs derived from the zgpa-1∼dif-1∼aph-1 operon in enoplean nematodes. The exon–intron structures of the amplicons used to identify polycistronic RNAs are shown, with exons represented by boxes (shaded to identify the genes from which they are derived using the same color coding that was used in A. The intercistronic regions are represented by cream-colored boxes. The positions of the SL trans-spliced 3′ splice sites are indicated. The length of each cDNA is indicated.

We determined the overall pattern of SL trans-splicing of the mRNAs derived from the putative operons in both T. spiralis and T. muris by analyzing the 5′ ends of zgpa-1, dif-1, and aph-1 mRNAs using 5′ RACE (Figure 1A; Table S2). The analysis of Tsp-zgpa-1 and Tmu-zgpa-1 transcripts mapped the mRNA 5′ ends to a region 200–250 bp upstream of the start codon, and we failed to detect any SL trans-spliced transcripts derived from this gene in either nematode. In contrast, all dif-1 transcripts analyzed were subject to SL trans-splicing in both organisms (Table S2). Tsp-dif-1 transcripts were trans-spliced to Tsp-SL10 [note this SL was previously given the designation TSL-10 (Pettitt et al. 2008), but we have renamed it to conform to accepted nematode gene nomenclature rules, which employ a species-specific prefix (Beech et al. 2010)] and Tmu-dif-1 transcripts were trans-spliced to the newly identified Tmu-SL1, Tmu-SL4, and Tmu-SL12 (Table S2). Analysis of aph-1 transcripts showed that in some cases the transcripts are SL trans-spliced, but we were also able to detect transcripts that initiated ∼200–300 bp upstream of the start codon, indicating that they were not subject to SL trans-splicing (Figure 1A; Table S2). It is notable that the distance between Tmu-aph-1 and Tmu-dif-1 is relatively large, suggesting the possibility that there are promoter elements immediately upstream of Tmu-aph-1 that would allow the production of transcripts without the need for SL trans-splicing. Such “hybrid operons” have been described in C. elegans (Huang et al. 2007).

As part of this analysis, we identified spliced leaders in T. muris, leading to the discovery of 13 Tmu-SLs (Figure 2; Table S2). Previous studies have shown that the primary sequences of spliced leaders in T. spiralis are much more variable than those found in the Chromadorea (Pettitt et al. 2008), and many lack the conserved motifs that characterize spliced leaders from these latter nematodes. In contrast, those of P. punctatus do not show the same diversity and display a greater degree of sequence similarity to the Chromadorid spliced leaders (Harrison et al. 2010). Analysis of the 13 distinct T. muris spliced leaders, designated Tmu-SL1–13, support this view, since the T. muris spliced leaders possess the same 5′ GGUWW and central CCC motifs that are highly conserved in the P. punctatus spliced leaders and Chromadorid SL1 and SL2 families, but missing in most of the T. spiralis spliced leaders. The presence of canonical nematode spliced leaders in T. muris and P. punctatus, despite the fact that the former nematode is more closely related to T. spiralis, supports the inference that the T. spiralis spliced leaders are derived features.

Figure 2.

Figure 2

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T. muris SL sequences and SL RNA structure. (A) Tmu-SL1–13 genes were identified using a combination of cDNA sequencing and bioinformatics tools as described in Materials and Methods. Tmu-SL12 was found by 5′ RACE trans-spliced to nuaf-3 mRNA, and Tmu-SL13 was found trans-spliced to aph-1 mRNA. In the alignment, only the SL sequences are shown. T. muris SL sequences were manually aligned and conserved groups are countershaded. C. elegans SL1 and SL2 and the previously identified P. punctatus SL sequences were included for comparison. (B) The intron of Tmu-SL2 was experimentally identified and also found in the genome sequence. The proposed secondary structure was produced using M-fold (Zuker 2003). The SL sequence is shown in outline font and the putative Sm sequence motif is countershaded.

If zgpa-1, dif-1, and aph-1 are components of a bona fide operon in the two enoplean nematodes, we would expect to be able to detect the polycistronic RNA from which their mRNAs are derived. Although not a definitive criterion, the presence of polycistronic, partially processed pre-mRNAs is a predicted property of operon usage. We tested for the presence of such RNA molecules in both T. spiralis and T. muris (Figure 1B; Figure S1) by reverse transcription of total RNA followed by PCR with gene-specific primers. PCR products were then analyzed by agarose gel electrophoresis. In T. spiralis, we detected RNA species connecting the open reading frames of Tsp-zgpa-1 with Tsp-dif-1 (Tsp-zgpa-1∼dif-1) and Tsp-dif-1 with Tsp-aph-1 (Tsp-dif-1∼aph-1) (Figure 1B). As we failed to amplify any products in control reactions performed in parallel with RNA subjected to mock reactions without reverse transcriptase (Figure S1), these products represent processing intermediates of polycistronic transcripts.

The Tsp-zgpa-1∼dif-1 intermediates contained the intercistronic region, and two of the intermediates lacked introns. The Tsp-dif-1∼aph-1 processing intermediates detected were all subject to cis-splicing of dif-1 introns, but we failed to detect an intermediate containing the complete ICR. Instead, the ICR was removed by cis-splicing of a cryptic splice donor site located in exon 7 of the dif-1 gene to the SL splice acceptor site of aph-1 (Figure 1B). Such cis-splicing events have also been detected in putative polycistronic RNAs discovered in tapeworm genomes (Tsai et al. 2013). Moreover, this demonstrates that Tsp-dif-1 and aph-1 are transcribed as a single transcript. In T. muris we also detected processing intermediates corresponding to Tmu-zgpa-1∼dif-1 and Tmu-dif-1∼aph-1 transcripts (Figure 1B; Table S2). The latter observation is significant, since the ICR between Tmu-dif-1 and Tmu-aph-1 is predicted to be 3033 nt long, a distance substantially longer than the length of an average ICR in C. elegans, although ICRs of similar length are also present in some C. elegans operons (Morton and Blumenthal 2011).

Identification of additional putative enoplean operons

To more systematically identify enoplean operons, we adopted two approaches. First, we used a set of EST sequences derived from SL trans-spliced T. spiralis mRNAs (Pettitt et al. 2008) to identify their corresponding genes via sequence similarity searches (we also identified two T. muris mRNAs via the same approach: FF145866 and CB277782). For each gene, we then looked for neighboring genes predicted to be transcribed in the same orientation and that lay within 1 kb. Using this approach, we were able to identify multiple potential operons in the T. spiralis genome (Table 1; Table S1). We further analyzed this set of genes by identifying the C. elegans orthologs of each T. spiralis gene and determining whether these correspond to genes within operons. Our analysis revealed that at least 75 of the C. elegans orthologs are arranged in operons. This represents 44% of the C. elegans genes identified as orthologs of our T. spiralis SL trans-spliced EST set. Since only 15% of C. elegans genes are organized into operons (Allen et al. 2011), we would expect only 15% of T. spiralis genes in our dataset to match C. elegans operonic homologs if they were selected at random. It is difficult to determine the reason for the increased likelihood of matches to C. elegans operonic genes among the T. spiralis SL trans-spliced ESTs; it may be that the corresponding T. spiralis gene set is biased for highly expressed genes, for instance. However, it is consistent with the possibility that this dataset is enriched for transcripts derived from operonic genes.

Table 1. Operonic status of genes that match ESTs derived from T. spiralis SL trans-spliced transcripts.

Species Location in operon Nonoperonic
Upstream Downstream Ambiguous
T. spiralis 30 35 40 54
C. elegans 29 46 5a 89
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The status of each T. spiralis gene was determined using criteria given in Materials and Methods. The status of C. elegans genes was obtained from WormBase (Release WS237). Eleven EST matches were absent from the T. spiralis data relative to the C. elegans data as the corresponding T. spiralis gene could not be identified (see Materials and Methods). Full details of the individual EST sequence matches are given in Table S1.

a

Genes not annotated as operons, but having intergenic spacing with respect to their neighbors, that suggests they may be organized in an operon.

As an alternative approach, which would potentially identify operons that have been conserved since the separation of the Enoplea and Chromadorea, we looked for T. spiralis homologs of a set of putative operons conserved between C. elegans and B. malayi (Ghedin et al. 2007). Of the 107 operonic gene pairs screened, we identified 12 T. spiralis gene pairs that displayed conserved synteny, and whose component genes were separated by an average intergenic distance of 607 bp (Table 2). To determine whether any of these operons are also conserved in another enoplean species, we examined the organization of the corresponding homologous genes in the genome of R. culicivorax (Schiffer et al. 2013). This analysis revealed that 4 of the 12 gene pairs were arranged in putative operons (assuming a maximum ICR distance of up to 1 kb) in this nematode (Table 2).

Table 2. Putative conserved nematode operons.

T. spiralis R. culicivorax C. elegans B. malayi
Upstream gene Downstream gene Upstream gene Downstream gene Upstream gene Downstream gene Operon no. Upstream gene Downstream gene
Tsp_00685 a Nonoperonic mrps-17 C05D11.9 3372 Bm1_13520 Bm1_13525
Tsp_03140 Tsp_03139 t32947 t32944 T26E3.4 par-6 1672 Bm1_48785 Bm1_48780
Tsp_05540 Tsp_05541 Nonoperonic K11B4.1 K11B4.2 1764 Bm1_55805 Bm1_55810
Tsp_06077 Tsp_06076 Nonoperonic Y62E10A.2 Y62E10A.6 4540 Bm1_54855 Bm1_54850
Tsp_06996 a t05598/9 t05596 sel-1 mrps-5 5365 Bm1_45745 Bm1_45750
Tsp_09103 Tsp_09102 t35569 t35568 snu-23 ZK686.3 3452 Bm1_13735 Bm1_13740
Tsp_09506 a Nonoperonic H20J04.6 mog-2 2124 Bm1_15855 Bm1_15860
Tsp_09539 a Nonoperonic E02H1.5 E02H1.6 2436 Bm1_24720 Bm1_24715
Tsp_10673 Tsp_10674 t34344.1 t34344.2 B0491.1 B0491.7 2532 Bm1_10780 Bm1_10785
Tsp_10698 Tsp_10702 Nonoperonic trpp-8 vha-10 1264 Bm1_12140 Bm1_12135
Tsp_10959 a Nonoperonic ubxn-2 Y94H6A.8 4665 Bm1_36515 Bm1_36520
Tsp_11898 a Nonoperonic lst-6 sqv-7 2276 Bm1_24075 Bm1_24080
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a

Single gene annotation matches to both C. elegans genes in the operon pair consistent with annotation error caused by short intergenic spacing.

Taken together, our analysis indicates that there are multiple T. spiralis gene pairs whose genomic arrangement is consistent with their corresponding to operons. Moreover, it is possible to identify gene pairs conserved between T. spiralis, R. culicivorax, B. malayi, and C. elegans, suggesting that these represent operons that were present in the last common ancestor of the three species.

Characterization of a conserved nematode operon

The analysis of one of the two SL trans-spliced T. muris mRNAs (GenBank accession no. FF145866) led to the identification of a putative operon conserved between multiple nematodes (Figure 3A). The two genes contained in these putative operons have C. elegans homologs, cpt-2 and nuaf-3, respectively, which are in the same operon (CEOP4424) (Figure 3A), although there is an additional gene, prx-14, located between these genes in CEOP4424 that is not present in the putative homologous T. spiralis operon. Examination of the genomic organization of the homologous genes in a selection of nematode species confirmed the evolutionary conservation of the synteny of the cpt-2 and nuaf-3 homologs (Figure 3A). This analysis also showed that insertion of prx-14 into the cpt-2∼nuaf-3 operon was a relatively recent event, since it is present only in C. elegans and other closely related Caenorhabditis species. We also find, based on the head-to-tail organization and spacing between coding regions, that there is variation in the composition of both operons in different species, and these genes in R. culicivorax are not in operons.

Figure 3.

Figure 3

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Evidence for an evolutionarily conserved nematode operon. (A) The structure of the cpt-2-nuaf-3 genomic regions from a range of nematode species mapped onto the nematode phylogeny. Genes are represented by arrows, and the gray lines represent the intercistronic regions (ICRs). Numbers above the ICRs represent the distances, in base pairs, between the stop and start codons of the upstream and downstream genes, respectively. The C. elegans operon numbers are given where appropriate. Fractions below the T. spiralis and T. muris genes represent the proportion of cDNAs derived from those genes that begins with a spliced leader sequence. (B) Detecting polycistronic RNAs derived from the cpt-2∼nuaf-3 operon in T. spiralis. The exon–intron structures of the amplicons used to identify polycistronic RNAs are shown, with exons represented by boxes (shaded to identify the genes from which they are derived using the same color coding that was used in A. The region removed during operon processing is represented by cream-colored boxes. The positions of the SL trans-splice 3′ spliced sites are indicated.

We further focused on the cpt-2∼nuaf-3 operon in T. spiralis and T. muris to determine the pattern of SL trans-splicing exhibited by the mRNAs derived from this operon and to verify that we were able to detect cDNAs consistent with the production of polycistronic RNAs.

Determination of the 5′ ends of cpt-2 and nuaf-3 transcripts by 5′ RACE revealed that nuaf-3 mRNA is subject to SL trans-splicing (Table S2) in both nematodes. However, we failed to detect any SL trans-spliced cpt-2 transcripts, similar to the situation with zgpa-1 transcripts in the zgpa-1∼dif-1∼aph-1 operon.

We were also able to detect processing intermediates derived from the putative polycistronic T. spiralis cpt-2∼nuaf-3 transcripts. As for the zgpa-1∼dif-1∼aph-1 operon, in addition to unprocessed, polycistronic transcripts, we detected cis-spliced intermediates lacking cpt-2 and nuaf-3 introns and a transcript in which the ICR was removed by splicing from a cryptic 3′ splice site in cpt-2 to the nuaf-3 SL splice acceptor site (Figure 3B).

The T. spiralis cpt-2∼nuaf-3 ICR can mediate polycistronic RNA processing in C. elegans

Analysis of the intercistronic region between Tsp-cpt-2 and nuaf-3 downstream of the polyadenylation signal of Tsp-cpt-2 revealed a clear Ur element and there are several U-rich regions, characteristics of the ICRs in C. elegans operons (Graber et al. 2007). To investigate the possibility that this region is able to function in polycistronic RNA processing, we determined whether the ICR from it could be recognized and processed if heterologously expressed in C. elegans. We generated an artificial operon consisting of sur-5::gfp (Gu et al. 1998) and mCherry genes flanking the ICR from Tsp-cpt-2∼nuaf-3. Transgenic animals carrying this construct expressed nuclear GFP and cytoplasmic mCherry, consistent with the processing of the two coding regions under the direction of the Tsp-cpt-2∼nuaf-3 ICR. We confirmed that this involved trans-splicing to SL2, as expected for polycistronic RNA processing in C. elegans, by showing that we could detect SL2 trans-splicing to the mCherry mRNA in RNA derived from transgenic animals (Figure 4). Thus, the predicted ICR between Tsp-cpt-2 and nuaf-3 is recognized and used as a substrate for polycistronic RNA processing in C. elegans.

Figure 4.

Figure 4

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Processing of a synthetic operon containing the T. spiralis cpt-2∼nuaf-3 intercistronic region in C. elegans. (A) Schematic of the structure of sur-5::gfp∼mCherry synthetic operon construct containing the Tsp-cpt-2∼nuaf-3 intercistronic region. The sequence immediately downstream from the Tsp-cpt-2 3′-UTR is shown, illustrating the presence of the Ur motif and the trans-splice acceptor site. Strain PE613 contains an identical construct, but with the ICR replaced with that from between Cel-cpt-2 and prx-14. (B) Detection of operon transcript processing by SL2 trans-splicing. mCherry transcripts trans-spliced to SL2 were detected by reverse transcription of RNA prepared from either PE612 or PE613 animals followed by PCR with an SL2 primer and a primer located in the mCherry coding region (+RT). Primers amplifying gpd-1 were included to control for sample variation. gpd-1 genomic DNA was detected in the −RT control reaction (*) and SL2-ZK1236.7a is a minor product detected in the +RT reactions. Reactions with RNA isolated from N2 wild-type animals were included as control. M is a DNA size standard. (C) Alignment of transgene sequences and SL2-mCherry transcripts confirming correct splice site usage. The beginning of the mCherry open reading frame is countershaded black, the NheI cloning site is countershaded gray, and the SL2 sequences are underlined.

Conservation of SL trans-splicing snRNPs between Enoplea and Chromadorea

Our data show that both SL trans-splicing and operons were likely present in the last common ancestor of the nematode phylum. This suggests that the processing machinery necessary for the coordination of these processes was already in place prior to radiation of the nematode phylum. To address this, we sought to determine the conservation of known protein components that are specifically involved in SL trans-splicing. Previous studies have shown the existence of two interacting proteins, conserved between Ascaris suum and C. elegans, that are components of the SL small nuclear ribonucleic particle (snRNP) (Denker et al. 2002; MacMorris et al. 2007). The two proteins, termed SNA-1 and SNA-2 in C. elegans, form a complex with SL RNA. In addition, a paralog of SNA-1, SUT-1, forms novel snRNPs containing SNA-2 and a family of nematode-specific RNAs, designated Sm Y (MacMorris et al. 2007; Jones et al. 2009). The function of Sm Y RNAs is not known, but they are associated with SL trans-splicing (Maroney et al. 1996), and a role in the recycling of Sm proteins following SL trans-splicing has been proposed (MacMorris et al. 2007).

We were able to identify credible homologs of sna-2 in the genomes of all nematodes in which searches were carried out, including T. spiralis and R. culicovorax, indicating that this gene encodes a nematode-wide SL trans-splicing component (Figure S2). Searches of the same datasets identified clear sna-1 and sut-1 orthologs in the chromadorean nematodes (Figure 5), extending the previously reported phylogenetic distribution of sut-1 (MacMorris et al. 2007). In contrast, the only enoplean genome in which we were able to identify a sut-1 homolog was R. culicivorax (Figure 5). We could not detect sna-1 homologs in any of the enoplean genomes that we assayed. These data suggest that the gene duplication event that gave rise to sna-1 and sut-1 occurred after the separation of the two major nematode taxa; however, we cannot rule out that the possibility that our failure to detect sna-1 homologs is due to the incomplete state of the enoplean genome drafts.

Figure 5.

Figure 5

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Evolutionary relationship between snRNPs associated with SL trans-splicing. Unrooted PhyML tree showing the relationship between sna-1 and sut-1 homologs identified in selected nematodes. Genes were named on the basis of their C. elegans homologs. The following species-specific prefixes were used: Aca, Angiostrongylus cantonensis; Asu, Ascaris suum; Bma, Brugia malayi; Cel, C. elegans; Cbr, C. briggsae; Cre, C. remanei; Hco, Haemochus contortus; Llo, Loa loa; Nam, Necator americanus; Rcu, Romanomermis culicivorax; and Wba, Wucheria bancrofti. The numbers at each node are approximate likelihood ratio test statistics.

Discussion

The incidence of operons as a means to coordinate gene expression has been investigated in only one of the two main nematode taxa, leaving unanswered the question about when this mechanism first occurred during nematode evolution (Guiliano and Blaxter 2006; Ghedin et al. 2007; Liu et al. 2010). The work presented here provides strong support for the existence of operons in the Enoplea, and that operons were likely present in the ancestor of the Chromadorea and the Enoplea. The unequivocal identification of operons is not straightforward in those nematodes that do not utilize a specialized spliced leader to resolve polycistronic RNAs; the use of SL2 in C. elegans and its close relatives has greatly facilitated the identification of operons in these species. In contrast, in enoplean species we can only infer the presence of operons through multiple lines of evidence.

Our work demonstrates the existence of clusters of genes ordered in a head-to-tail arrangement with short, less than 1 kb, intergenic distances, consistent with their being organized into operons. The fact that we can identify homologous pairs of closely spaced genes conserved between enoplean and chromadorean nematodes suggests that at least in these cases there has been selective pressure to retain short intergenic distances, consistent with what would be expected if they were part of operons. Analysis of the transcripts produced by these putative operons shows that, as expected, genes predicted to be downstream in operons produce mRNAs that are SL trans-spliced. Further supporting evidence comes from the fact that we can detect unprocessed, polycistronic pre-mRNAs derived from these putative operons. Thus these genes exhibit the molecular properties expected of operons. Most significantly, the intergenic region between Tsp-cpt-2 and nuaf-3 acts as a substrate for the polycistronic RNA processing machinery of C. elegans, providing the strongest evidence that it is part of an operon in T. spiralis.

The identification of SLs from T. muris has extended our understanding of nematode SL evolution. Previous studies have shown that T. spiralis and T. pseudospiralis have unusually diverse SLs that do not readily correspond to those SLs found within Chromadorea (Pettitt et al. 2008, 2010). Another enoplean, P. punctatus, possesses SLs that resemble the specialized SL2 associated with trans-splicing and polycistronic RNA resolution found in C. elegans and other closely related nematodes. At least one of these is specifically recognized by the SL2 trans-splicing machinery in C. elegans when expressed heterogeneously (Harrison et al. 2010). Rather surprisingly, since it is more closely related to Trichinella species than P. punctatus, the T. muris SLs are more similar in terms of sequence composition to those of the latter nematode (the most extreme example of this being Ppu-SL3 and Tmu-SL13, which possess >95% sequence identity). Thus, the diverse SL complement of the Trichinella species is likely to be a derived rather than ancestral trait.

In addition to elucidating the extent of nematode SL RNA conservation, we have determined the phylogenetic distribution of the protein components of the SL snRNP. This reveals that one of these components, SNA-2, is conserved throughout the phylum, consistent with its essential role in SL trans-splicing in both A. suum and C. elegans (Denker et al. 2002; MacMorris et al. 2007). Our analysis of SNA-1/SUT-1 has revealed that these two paralogs are distributed throughout the Chromadorea. These data, together with the presence of a single clear SUT-1 homolog in R. culicivorax (and absence of a SNA-1 homolog) is consistent with an evolutionary scenario whereby the ancestral sut-1 gene was duplicated in the ancestor of the Chromadorea, with one paralog, sut-1, retaining the ancestral function, while the other, sna-1, evolved a derived function. However, the fact that sna-1 and sut-1 show a synthetic loss of function phenotype in C. elegans (MacMorris et al. 2007) indicates that their functions are related and possibly overlapping. The puzzling absence of SUT-1 homologs in T. spiralis and T. muris means that such an interpretation must be provisional, pending broader sampling of genome sequences from other enopleans. If these nematodes actually lack these proteins this would be surprising, given that loss of both sna-1 and sut-1 function in C. elegans results in embryonic lethality (MacMorris et al. 2007) and SNA-1 is required for SL trans-splicing in A. suum splicing extracts (Denker et al. 2002).

An important question that remains to be answered is whether there is any evidence for functional specialization of the enoplean SLs with regard to polycistronic RNA resolution. In C. elegans, SL2-type SLs are clearly specific for this latter process (Blumenthal et al. 2002), with SL1 trans-splicing almost exclusively employed in mRNAs derived from nonoperonic genes, or the first gene in an operon. Other nematodes within the Rhabditina also show the same specialization (Evans et al. 1997; Lee and Sommer 2003), but other taxa within the Chromadorea appear to use the same set of SLs for all SL trans-spliced mRNAs (Guiliano and Blaxter 2006). Within the Enoplea we clearly see multiple different SLs used in the trans-splicing of mRNAs derived from downstream genes in operons, but our data are not comprehensive enough to determine whether some SLs are preferentially used to process the transcripts arising from such genes.

We have identified multiple examples of gene pair synteny conserved between T. spiralis, R. culicivorax, B. malayi, and C. elegans, suggesting these correspond to conserved operons present in the last common ancestor of the Enoplea and Chromadorea. Nevertheless, the majority of the putative operons that we have identified in T. spiralis are not syntenic with C. elegans operons, although in many cases we see that the component genes of these putative T. spiralis operons have C. elegans orthologs that are found in operons. An example of such an operon is that comprising Tsp-zgpa-1, Tsp-dif-1, and Tsp-aph-1, whose C. elegans orthologs are located in different operons. We also observed putative T. spiralis operon genes whose C. elegans orthologs are not part of operons. The changes in the operon complements between the two nematodes could arise from the lineage-specific rearrangements of an ancestral set of nematode operons, but could also be accounted for by differential de novo operon generation in the two clades, or a mixture of the two processes. A key question is whether the synteny shown between the putative enoplean operons and their chromadorean orthologs is significant, i.e., are the genes that comprise these operons more constrained to be located in the same operon than other operonic genes, or is the conservation of synteny merely random chance? The availability of tools to engineer the C. elegans genome (Golic 2013) might allow this question to be addressed by assaying the function of selected operons compared with their individual component genes each expressed under their own promoters.

Finally, it is clear from this work and previous studies that SL trans-splicing and operon organization arose prior to the divergence of the Enoplea and Chromadorea. An important question is whether “nematode” SL trans-splicing and operons predate the foundation of the phylum. It will thus be important to establish whether they are also present in the other, so far uncharacterized, phyla that are closely related to the nematodes.

Supplementary Material

Supporting Information
supp_197_4_1201__index.html (1.8KB, html)

Acknowledgments

We thank Allison Bancroft and Richard Grencis for providing T. muris RNA. The Bristol (N2) C. elegans N2 strain was provided by the Caenorhabditis Genetics Center, which is funded by National Institutes of Health Office of Research Infrastructure Programs (P40 OD010440). This work was funded by a Biotechnology and Biological Sciences Research Council project grant BB/J007137/1.

Footnotes

Communicating editor: M. P. Colaiácovo

Literature Cited

  1. Allen M. A., Hillier L. W., Waterston R. H., Blumenthal T., 2011.  A global analysis of C. elegans trans-splicing. Genome Res. 21: 255–264. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Beech R. N., Wolstenholme A. J., Neveu C., Dent J. A., 2010.  Nematode parasite genes: what’s in a name? Trends Parasitol. 26: 334–340. [DOI] [PubMed] [Google Scholar]
  3. Blumenthal T., Evans D., Link C. D., Guffanti A., Lawson D., et al. , 2002.  A global analysis of Caenorhabditis elegans operons. Nature 417: 851–854. [DOI] [PubMed] [Google Scholar]
  4. Dana C. E., Glauber K. M., Chan T. A., Bridge D. M., Steele R. E., 2012.  Incorporation of a horizontally transferred gene into an operon during cnidarian evolution. PLoS ONE 7: e31643. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Davis R. E., Hodgson S., 1997.  Gene linkage and steady state RNAs suggest trans-splicing may be associated with a polycistronic transcript in Schistosoma mansoni. Mol. Biochem. Parasitol. 89: 25–39. [DOI] [PubMed] [Google Scholar]
  6. Denker J. A., Zuckerman D. M., Maroney P. A., Nilsen T. W., 2002.  New components of the spliced leader RNP required for nematode trans-splicing. Nature 417: 667–670. [DOI] [PubMed] [Google Scholar]
  7. Dereeper A., Guignon V., Blanc G., Audic S., Buffet S., et al. , 2008.  Phylogeny.fr: robust phylogenetic analysis for the non-specialist. Nucleic Acids Res. 36: W465–W469. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Evans D., Zorio D., MacMorris M., Winter C. E., Lea K., et al. , 1997.  Operons and SL2 trans-splicing exist in nematodes outside the genus Caenorhabditis. Proc. Natl. Acad. Sci. USA 94: 9751–9756. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Ganot P., Kallesøe T., Reinhardt R., Chourrout D., Thompson E. M., 2004.  Spliced-leader RNA trans splicing in a chordate, Oikopleura dioica, with a compact genome. Mol. Cell. Biol. 24: 7795–7805. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Ghedin E., Wang S., Spiro D., Caler E., Zhao Q., et al. , 2007.  Draft genome of the filarial nematode parasite Brugia malayi. Science 317: 1756–1760. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Golic K. G., 2013.  RNA-guided nucleases: a new era for engineering the genomes of model and nonmodel organisms. Genetics 195: 303–308. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Graber J. H., Salisbury J., Hutchins L. N., Blumenthal T., 2007.  C. elegans sequences that control trans-splicing and operon pre-mRNA processing. RNA 13: 1409–1426. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Gu T., S. Orita, M. Han, 1998.  Caenorhabditis elegans SUR-5, a novel but conserved protein, negatively regulates LET-60 Ras activity during vulval induction. Mol. Cell. Biol. 18: 4556–4564. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Guiliano D. B., Blaxter M. L., 2006.  Operon conservation and the evolution of trans-splicing in the phylum Nematoda. PLoS Genet. 2: e198. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Harrison N., Kalbfleisch A., Connolly B., Pettitt J., Müller B., 2010.  SL2-like spliced leader RNAs in the basal nematode Prionchulus punctatus: New insight into the evolution of nematode SL2 RNAs. RNA 16: 1500–1507. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Hastings K. E. M., 2005.  SL trans-splicing: easy come or easy go? Trends Genet. 21: 240–247. [DOI] [PubMed] [Google Scholar]
  17. Hobert O., 2002.  PCR fusion-based approach to create reporter gene constructs for expression analysis in transgenic C. elegans. Biotechniques 32: 728–730. [DOI] [PubMed] [Google Scholar]
  18. Holterman M., van der Wurff A., van den Elsen S., van Megen H., Bongers T., et al. , 2006.  Phylum-wide analysis of SSU rDNA reveals deep phylogenetic relationships among nematodes and accelerated evolution toward crown Clades. Mol. Biol. Evol. 23: 1792–1800. [DOI] [PubMed] [Google Scholar]
  19. Huang P., Pleasance E. D., Maydan J. S., Hunt-Newbury R., O’Neil N. J., et al. , 2007.  Identification and analysis of internal promoters in Caenorhabditis elegans operons. Genome Res. 17: 1478–1485. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Jacob F., Perrin D., Sánchez C., Monod J., Edelstein S., 1960.  The operon: a group of genes with expression coordinated by an operator. C. R. Acad. Sci. Paris 250: 1727–1729.14406329 [Google Scholar]
  21. Johnson P. J., Kooter J. M., Borst P., 1987.  Inactivation of transcription by UV irradiation of T. brucei provides evidence for a multicistronic transcription unit including a VSG gene. Cell 51: 273–281. [DOI] [PubMed] [Google Scholar]
  22. Jones T., Otto W., Marz M., Eddy S., Stadler P., 2009.  A survey of nematode SmY RNAs. RNA Biol. 6: 5– 8. [DOI] [PubMed] [Google Scholar]
  23. Lawrence J., 1999.  Selfish operons: the evolutionary impact of gene clustering in prokaryotes and eukaryotes. Curr. Opin. Genet. Dev. 9: 642–648. [DOI] [PubMed] [Google Scholar]
  24. Lee K.-Z., Sommer R. J., 2003.  Operon structure and trans-splicing in the nematode Pristionchus pacificus. Mol. Biol. Evol. 20: 2097–2103. [DOI] [PubMed] [Google Scholar]
  25. Liu C., Oliveira A., Chauhan C., Ghedin E., Unnasch T. R., 2010.  Functional analysis of putative operons in Brugia malayi. Int. J. Parasitol. 40: 63–71. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. MacMorris M., Kumar M., Lasda E., Larsen A., Kraemer B., et al. , 2007.  A novel family of C. elegans snRNPs contains proteins associated with trans-splicing. RNA 13: 511–520. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Marlétaz F., Gilles A., Caubit X., Perez Y., Dossat C., et al. , 2008.  Chaetognath transcriptome reveals ancestral and unique features among bilaterians. Genome Biol. 9: R94. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Maroney P. A., Yu Y. T., Jankowska M., Nilsen T. W., 1996.  Direct analysis of nematode cis- and trans-spliceosomes: a functional role for U5 snRNA in spliced leader addition trans-splicing and the identification of novel Sm snRNPs. RNA 2: 735–745. [PMC free article] [PubMed] [Google Scholar]
  29. Meldal B. H. M., Debenham N. J., De Ley P., De Ley I. T., Vanfleteren J. R., et al. , 2007.  An improved molecular phylogeny of the Nematoda with special emphasis on marine taxa. Mol. Phylogenet. Evol. 42: 622–636. [DOI] [PubMed] [Google Scholar]
  30. Mitreva M., Jasmer D. P., Zarlenga D. S., Wang Z., Abubucker S., et al. , 2011.  The draft genome of the parasitic nematode Trichinella spiralis. Nat. Genet. 43: 228–235. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Morton J. J., Blumenthal T., 2011.  Identification of transcription start sites of trans-spliced genes: uncovering unusual operon arrangements. RNA 17: 327–337. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Pettitt J., Harrison N., Stansfield I., Connolly B., Müller B., 2010.  The evolution of spliced leader trans-splicing in nematodes. Biochem. Soc. Trans. 38: 1125–1130. [DOI] [PubMed] [Google Scholar]
  33. Pettitt J., Müller B., Stansfield I., Connolly B., 2008.  Spliced leader trans-splicing in the nematode Trichinella spiralis uses highly polymorphic, noncanonical spliced leaders. RNA 14: 760–770. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Protasio A. V., Tsai I. J., Babbage A., Nichol S., Hunt M., et al. , 2012.  A systematically improved high quality genome and transcriptome of the human blood fluke Schistosoma mansoni. PLoS Negl. Trop. Dis. 6: e1455. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Satou Y., Mineta K., Ogasawara M., Sasakura Y., Shoguchi E., et al. , 2008.  Improved genome assembly and evidence-based global gene model set for the chordate Ciona intestinalis: new insight into intron and operon populations. Genome Biol. 9: R152. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Schiffer P. H., Kroiher M., Kraus C., Koutsovoulos G. D., Kumar S., et al. , 2013.  The genome of Romanomermis culicivorax: revealing fundamental changes in the core developmental genetic toolkit in Nematoda. BMC Genomics 14: 923. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Spieth J., Brooke G., Kuersten S., Lea K., Blumenthal T., 1993.  Operons in C. elegans: polycistronic mRNA precursors are processed by trans-splicing of SL2 to downstream coding regions. Cell 73: 521–532. [DOI] [PubMed] [Google Scholar]
  38. Tsai I. J., Zarowiecki M., Holroyd N., Garciarrubio A., Sanchez-Flores A., et al. , 2013.  The genomes of four tapeworm species reveal adaptations to parasitism. Nature 496: 57–63. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Zaslaver A., Baugh L. R., Sternberg P. W., 2011.  Metazoan operons accelerate recovery from growth-arrested states. Cell 145: 981–992. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Zuker M., 2003.  Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res. 31: 3406–3415. [DOI] [PMC free article] [PubMed] [Google Scholar]

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