PMCCPMCCPMCC

Search tips
Search criteria 

Advanced

 
Logo of gthreeG3 Genes | Genomoes | GeneticsInformation for AuthorsEditorial BoardSubmit a Manuscript
 
G3 (Bethesda). 2017 March; 7(3): 871–880.
Published online 2017 January 23. doi:  10.1534/g3.116.037275
PMCID: PMC5345718

De Novo Transcriptome Characterization of a Sterilizing Trematode Parasite (Microphallus sp.) from Two Species of New Zealand Snails

Abstract

Snail-borne trematodes represent a large, diverse, and evolutionarily, ecologically, and medically important group of parasites, often imposing strong selection on their hosts and causing host morbidity and mortality. Even so, there are very few genomic and transcriptomic resources available for this important animal group. We help to fill this gap by providing transcriptome resources from trematode metacercariae infecting two congeneric snail species, Potamopyrgus antipodarum and P. estuarinus. This genus of New Zealand snails has gained prominence in large part through the development of P. antipodarum and its sterilizing trematode parasite Microphallus livelyi into a textbook model for host–parasite coevolutionary interactions in nature. By contrast, the interactions between Microphallus trematodes and P. estuarinus, an estuary-inhabiting species closely related to the freshwater P. antipodarum, are relatively unstudied. Here, we provide the first annotated transcriptome assemblies from Microphallus isolated from P. antipodarum and P. estuarinus. We also use these transcriptomes to produce genomic resources that will be broadly useful to those interested in host–parasite coevolution, local adaption, and molecular evolution and phylogenetics of this and other snail–trematode systems. Analyses of the two Microphallus transcriptomes revealed that the two trematode types are more genetically differentiated from one another than are the M. livelyi infecting different populations of P. antipodarum, suggesting that the Microphallus infecting P. estuarinus represent a distinct lineage. We also provide a promising set of candidate genes likely involved in parasitic infection and response to salinity stress.

Keywords: transcriptome, host-parasite interactions, microsatellites, coevolution, trematode

Snail-borne trematodes are important sources of morbidity and mortality in animals, including humans (Hotez 2013; Jurberg and Brindley 2015). For example, infections by trematodes in the genus Schistosoma affect >200 million people in sub-Saharan Africa (Rollinson et al. 2013). Trematodes and their mollusk intermediate hosts are also important models for the study of host–parasite coevolutionary interactions (e.g., Lively and Jokela 1996; King et al. 2009). While digenetic trematodes (parasitic flatworms) are one of the largest groups of metazoan parasites, with ~18,000 species (Cribb et al. 2001), the genomic and transcriptomic resources needed to understand trematode evolution, host–parasite coevolution, and the genomic basis of trematode infection are only available for 13 of these species (>50% members of the Schistosoma genus; NCBI, January 2017). Further development of such resources for this medically and scientifically important group of parasites is a necessary component of characterizing the molecular and genomic underpinnings of parasitic infection and host–parasite coevolutionary interactions.

New Zealand snails in the genus Potamopyrgus are often the intermediate host of sterilizing trematode parasites in the genus Microphallus (Winterbourn 1973). Here, we focus on Microphalluslivelyi’ (Hechinger 2012; hereafter “M. livelyi”), a species of digenetic trematode that uses Potamopyrgus antipodarum as an intermediate host and dabbling ducks (Anatinae) as the final host (Dybdahl and Lively 1996; Lively and Jokela 1996; King et al. 2009; Osnas and Lively 2011). Dabbling ducks are infected by M. livelyi while foraging, typically consuming infected P. antipodarum from submerged rocks and aquatic vegetation. The trematodes lay sexually produced eggs within the ducks, which are excreted within duck feces (Osnas and Lively 2011). The next round of snail infection begins after detritus-feeding P. antipodarum consume M. livelyi eggs while foraging along lake bottoms and in aquatic vegetation (Lively and Jokela 1996). Following snail consumption, the M. livelyi eggs clonally develop into thousands of encysted larvae (metacercariae) that rapidly sterilize the snail (Winterbourn 1973; King et al. 2009). M. livelyi individuals that cannot infect the first snail by which they are ingested will die before completing their lifecycle, and successful infection means that the M. livelyi metacercariae sterilize the snail, leading to strong reciprocal antagonistic selection pressures between these two species (King et al. 2011). These metacercariae are also the M. lively stage that is transmissible to the duck final host (Levri and Lively 1996) and thus represent an especially important point of no return in the life history of Microphallus: if the snail-inhabiting metacercariae are not ingested by the duck final host, the parasite will be unable to complete its life cycle (Lively and Jokela 1996; King et al. 2011). The importance of this metacercariae life stage is emphasized by new RNA sequencing (RNA-Seq) evidence that the presence of M. livelyi metacercariae influences patterns of gene expression in P. antipodarum (Bankers et al. 2016). By this logic and informed by these data, we chose to focus on the metacercariae stage for our analyses.

How interactions between M. livelyi and P. antipodarum play out in Microphallus are not well characterized, particularly at the genetic and molecular levels. The initial insights we do have into the mechanisms underlying Microphallus infectivity come from local adaptation and reciprocal infection experiments, providing preliminary lines of evidence in support of the likelihood that coevolutionary interactions between P. antipodarum and M. livelyi fit a “matching alleles” infection genetics model (Dybdahl and Krist 2004; Lively et al. 2004; King et al. 2009). With a matching alleles system, parasite infection success is thought to be determined by whether the parasite genotype at the loci involved in mediating infection match the corresponding loci in the host, enabling successful parasites to evade detection by host defenses. While matching alleles systems for infection have been demonstrated in several host–parasite interactions, the specific genetic or molecular mechanisms underlying such interactions remain unclear in P. antipodarum and in every other system (Carmona et al. 2015; Routtu and Ebert 2015).

The transcriptomic resources described here, the first genomic resources for the Microphallus genus of which we are aware, will provide a valuable starting point to characterize the molecular underpinnings of M. livelyi infection and the coevolutionary interactions between M. livelyi and P. antipodarum. We present annotated transcriptomes assemblies from M. livelyi metacercariae isolated from P. antipodarum, as well as from Microphallus of unknown species isolated from P. estuarinus, a closely related species native to New Zealand estuaries. While the coevolutionary interactions between M. livelyi and P. antipodarum are well characterized, much less is known about the Microphallus that infect P. estuarinus (Winterbourn 1973; Hechinger 2012). Expanding the genomic resources available for these two closely related types of Microphallus will allow for the characterization of genes that contribute broadly to infection processes, as well as genes that play species-specific roles in host–parasite interactions.

We identified and annotated 7584 pairs of one-to-one orthologous transcripts between the two transcriptomes, analyzed levels of genetic variation within and genetic differentiation between the two types of Microphallus at these genes, and performed gene ontology (GO)-based functional comparisons between these two samples. We also provide a broad set of genomic resources that set the stage for future studies aimed at characterizing the molecular underpinnings of the coevolutionary interactions between Microphallus and Potamopyrgus.

Materials and Methods

Sample collection and RNA-Seq

We isolated M. livelyi metacercariae from four individual adult female P. antipodarum (PA-Microphallus) collected in January of 2013 in shallow regions (<1 m lake depth) of each of the two New Zealand lakes Alexandrina and Kaniere, for a total of eight PA-Microphallus samples. We chose these lakes because their P. antipodarum populations contain relatively high frequencies of M. livelyi infection (10–20%; Vergara et al. 2013). We also isolated Microphallus metacercariae from a single adult female P. estuarinus (PE-Microphallus) collected from the Ashley River estuary (near Christchurch, New Zealand) in January of 2015. Metacercariae were collected from stage five infections (http://www.indiana.edu/~curtweb/trematodes/DATA_KEY.HTM), in which the entire snail body cavity is filled with clonally produced metacercariae cysts. We chose the metacercariae life stage both because it represents a critical time point in the life cycle of Microphallus and because this life stage is easily identifiable and allows for larger volumes of tissue to be collected than earlier stages.

We dissected individual Potamopyrgus snails to determine infection status and saved parasite tissue from snails containing stage five Microphallus infections. We separated parasite from snail tissue with a forceps and used a micropipette to isolate metacercariae cysts. We immediately placed the metacercariae in RNAlater Solution (Life Technologies Corporation) at 4° for 24 hr, and subsequently stored the RNAlater-submerged metacercariae at −80° (according to the manufacturer’s protocol) until RNA extraction. The eight PA-Microphallus metacercariae samples were pooled in a single tube. We had to use this pooling strategy for the PA-Microphallus because at this time (early 2013), pooling of cysts from multiple snails was necessary to obtain a sufficient amount of RNA to perform RNA-Seq. Technological advances in library preparation and RNA-Seq between 2013 and 2015 meant that we were able to obtain a sufficient amount of RNA from the metacercariae isolated from a single P. estuarinus individual for RNA-Seq.

We extracted RNA from metacercariae following the TRIzol protocol (Chomczynski and Sacchi 1987; Chomczynski 1993). We assessed RNA quantity and quality using a Bio-Rad Experion Automated Electrophoresis Station and Experion RNA analysis kit, requiring an RQI ≥ 8 and minimum of 2 µg of total RNA (following the manufacturer’s protocol). We prepared cDNA libraries following the Illumina Truseq LS protocol (Illumina, San Diego, CA) for the PA-Microphallus and PE-Microphallus samples. We barcoded the PA-Microphallus library and pooled the barcoded library with 11 other barcoded samples that were part of a different project in one Illumina HiSeq 2000 lane in 2013. For the PE-Microphallus library, we barcoded the library and pooled the barcoded library with five other samples that were part of a different project in one Illumina HiSeq 2000 lane in 2015. We performed 2 × 100 bp paired-end RNA-Seq for both Microphallus libraries. We obtained 31,635,450 paired-end reads (mean read length = 101 bp) for PA-Microphallus and 4,16,15,863 paired-end reads for PE-Microphallus (mean read length = 126 bp).

Transcriptome assembly and annotation

We used FASTQC (Andrews 2010) to assess RNA-Seq quality and the FASTX Toolkit (Gordon and Hannon 2010) to trim adapter sequences and remove poor quality reads from raw RNA-Seq data, requiring a mean Phred score ≥ 20. After quality filtering, 31,096,527 paired-end reads remained for PA-Microphallus and 40,413,790 paired-end reads remained for PE-Microphallus. We used these filtered reads to generate de novo transcriptome assemblies for each of the two Microphallus samples.

We performed the Trinity in silico normalization, requiring a minimum Kmer coverage of 2× and maximum Kmer coverage of 100× (following Haas et al. 2013), and otherwise default parameters (Grabherr et al. 2011). We then generated preliminary de novo transcriptome assemblies for both types of Microphallus using Trinity software with default parameters (Haas et al. 2013). Preliminary assemblies contained 65,665 contigs and 2,26,896 contigs for PA-Microphallus and PE-Microphallus, respectively. We used the Trinity plugin TransDecoder with default parameters to annotate likely protein-coding regions, identify long open reading frames of putative genes, and filter miscalled isoforms (Haas et al. 2013). Next, we used hierarchical clustering based on sequence identity to further reduce redundancy in the transcriptome assemblies, as implemented by CD-HIT-EST (Huang et al. 2010). To reduce the likelihood of collapsing truly nonredundant transcripts, we used a sequence identity of 0.95 and word size of 8 nt (e.g., Li et al. 2002). These steps resulted in assemblies containing 15,435 contigs for PA-Microphallus and 29,564 contigs for PE-Microphallus. We mapped reads back to their respective filtered transcriptome assemblies using TopHat2 and then used Cufflinks (Trapnell et al. 2013) to estimate mean FPKM for each transcript in each transcriptome. Mean FPKM (±SD) for the transcriptome assemblies were 71.6 (±274.8) for PA-Microphallus and 46.8 (±284.3) for PE-Microphallus. The large SD in FPKM for both types of Microphallus is due to the large ranges in FPKM (~0 to 10,866 for PA-Microphallus; ~0 to 19,279 for PE-Microphallus). The nearly twofold higher number of transcripts for PE-Microphallus relative to PA-Microphallus was likely a combination of advances in sequencing chemistry and technology between 2013 and 2015, higher sequencing depth for PE-Microphallus, and/or differences in heterozygosity of the sequenced individuals and/or samples. We annotated the transcriptomes using blastx with an E-value cut-off of 1e−6, followed by Blast2GO (using default parameters) to assign GO terms to blastx-annotated transcripts (Conesa et al. 2005). For PA-Microphallus, we obtained both blastx and GO annotations for 9272 transcripts and only blastx annotations for an additional 3814 transcripts. For PE-Microphallus we obtained both blastx and GO annotations for 13,778 transcripts and only blastx annotations for an additional 8822 transcripts. This analysis revealed 143 transcripts involved in photosynthesis in the PE-Microphallus assembly (none in PA-Microphallus), representing potential contaminants in the snail gut from which the PE-Microphallus was isolated. These 143 transcripts were filtered out of subsequent analyses. Finally, we used BUSCO (Simão et al. 2015) to assess the completeness of our transcriptome assemblies. This program uses tblastn to identify conserved single-copy orthologs among groups of organisms. We used the BUSCO OrthoDB Metazoan database to search our assemblies for the presence, absence, duplication, and/or fragmentation of 843 conserved single-copy Metazoan orthologs.

Identification of orthologous transcripts

We used “reciprocalblast_allsteps.py” (Warren et al. 2014) to identify putative orthologous transcripts between the two Microphallus transcriptomes based on one-to-one reciprocal best blastn hits. We performed reciprocal blastn searches between the two transcriptomes with an E-value cutoff of 1e−20 and identified pairs of transcripts that were reciprocally one another’s top blastn hit and that occurred once and only once in each transcriptome query. We identified 11,160 of such transcript pairs, which we will hereafter refer to as “one-to-one orthologs.” Using a custom python script (github.com/jsharbrough/grabContigs), we extracted the 11,160 orthologous sequences from each Microphallus transcriptome to generate putative ortholog transcriptomes for each sample. To improve the accuracy of our ortholog predictions, we then used blastx against the NCBI nonredundant dataset with an E-value cutoff of 1e−6 (Camacho et al. 2009) to annotate the two transcriptomes and filter out putative orthologs that received different blastx annotations. This step resulted in 7935 remaining putative one-to-one orthologs. To further ensure that we accurately predicted orthologs, we also performed reciprocal shortest molecular distance comparisons between the 7935 pairs of transcripts using “reciprocal_smallest_distance.py” (Wall et al. 2003). Here, we inferred orthologs using a combination of global sequence alignments between the two transcriptomes and maximum likelihood-based evolutionary pairwise distances, and identified orthologous pairs of transcripts as those sets of transcripts with the shortest pairwise molecular distance between them relative to their pairwise distances from any of the other 7934 transcripts in the dataset (Wall et al. 2003). This filtering step left 7584 remaining pairs of one-to-one orthologous transcripts that met all three of our criteria: (1) reciprocal best blastn hits, (2) the same blastx annotation, and (3) shared the shortest pairwise evolutionary distance relative to all other transcripts. There were 7851 and 21,980 transcripts for PA-Microphallus and PE-Microphallus, respectively, that did not meet these three criteria (hereafter, “nonone-to-one orthologs”). We extracted the 7584 orthologous transcripts from each reference transcriptome to generate ortholog transcriptomes for each of the two Microphallus datasets. We estimated mean FPKM for each transcript in the ortholog transcriptomes as described above. Mean FPKM (±SD) for the transcriptome assemblies were 114.3 (±403.2) for PA-Microphallus and 111.7 (±569.2) for PE-Microphallus.

Comparisons of genetic variation and genetic differentiation

We used Genome Tools to split both of the ortholog transcriptomes into individual fasta files for each transcript (Gremme et al. 2013). We then merged pairs of orthologous sequences into multifasta files and used Clustal Omega to align and compute Jukes–Cantor-corrected pairwise molecular distances for each pair of orthologous transcripts (Sievers et al. 2011). Finally, we used the pairwise molecular distance values between each pair of orthologous transcripts obtained from Clustal Omega to estimate mean and median pairwise molecular distance between the two transcriptomes. We used SPSS Statistics v.23 to generate a boxplot of pairwise molecular distance that enabled us to visualize the distribution of these values (see Supplemental Material, Figure S1).

We used the PoPoolation pipeline to assess allelic diversity (Watterson’s θ) within and relative levels of genetic differentiation (FST) between the two Microphallus samples (Kofler et al. 2011a,b). Watterson’s θ is an estimator of nucleotide diversity based on the number of segregating sites in a sample (here, SNPs) (Watterson 1975). FST is a descriptive statistic that estimates relative levels of genetic differentiation among populations based on variance in allele frequencies (Holsinger and Weir 2009). Analyses of levels of nucleotide variation and genetic differentiation from pooled RNA-Seq data with programs like PoPoolation and PoPoolation2 (Kofler et al. 2011a,b), which were developed to perform variant calling and calculate FST from pooled sequencing data, can effectively identity SNPs and candidate genes and perform FST-based measures of genetic differentiation (e.g., Fischer et al. 2013; Konczal et al. 2014; Li et al. 2014; Thavamanikumar et al. 2014; Ellison et al. 2016). In order to lower the amount of memory needed for analysis and to reduce biases linked to variation in gene expression among loci, we required a minimum coverage of 10× and a maximum coverage of 200× for both the θ and FST analyses described below. We also required that each SNP was sequenced at least four times and filtered out SNPs with minor allele frequencies <5% (e.g., Fischer et al. 2013; Konczal et al. 2014).

First, we used TopHat2 (Trapnell et al. 2013) to map Microphallus reads to their respective ortholog transcriptomes. For the Watterson’s θ estimates, this process included mapping the PA-Microphallus reads to the PA-Microphallus ortholog transcriptome and mapping the PE-Microphallus reads to the PE-Microphallus ortholog transcriptome. To estimate FST between the two types of Microphallus, PoPoolation2 requires the mapping of the two samples (PA-Microphallus and PE-Microphallus) to a single reference. In order to account for the ortholog transcriptome to which reads were mapped, we performed this analysis twice, first by mapping both sets of Microphallus reads to the PA-Microphallus ortholog transcriptome, and second by mapping both sets of Microphallus reads to the PE-Microphallus ortholog transcriptome. After mapping, we used Picard Tools v.1.119 (http://broadinstitute.github.io/picard/) to process and filter the bam files. This step included the addition of read groups and removal of duplicate reads from each bam file. We then used GATK (McKenna et al. 2010; DePristo et al. 2011; Van der Auwera et al. 2013) to realign reads around indels and to reassign mapping quality. Finally, we used SAMtools (Li et al. 2012) to sort each bam file by coordinates relative to the ortholog transcriptome to which they were mapped and to generate mpileups.

We generated separate mpileup files for each bam file and used PoPoolation (Kofler et al. 2011a) for further analyses. PoPoolation is a suite of programs designed to handle pooled sequencing data and is able to account for the number of individuals sequenced in each pool. First, we used PoPoolation’s Variation-at-position.pl script (which is not available in newer program versions) to calculate Watterson’s θ for each orthologous transcript for the two Microphallus types. We compared the medians and distributions of θ for each transcriptome using Bonferroni-corrected Mann–Whitney U tests and Kolmogorov–Smirnov tests, respectively, as implemented in SPSS Statistics v.23. We also used SPSS Statistics v.23 to generate a boxplot for Watterson’s θ and FST per SNP in order to visualize the distribution of values for each statistic (see Figure S1).

We then used the most up-to-date version of PoPoolation (PoPoolation2) (Kofler et al. 2011b) to generate synchronized mpileups (mpileup2sync.pl) for each bam file associated with the ortholog transcriptomes (i.e., PA-Microphallus and PE-Microphallus mapped to the PA-Microphallus ortholog transcriptome, and PA-Microphallus and PE-Microphallus mapped to the PE-Microphallus ortholog transcriptome). Next, we used the PoPoolation2 fst-sliding.pl to calculate FST per SNP to assess levels of relative genetic differentiation between the two Microphallus types. We used these FST values to determine mean and median FST per SNP between PA-Microphallus and PE-Microphallus relative to both of the ortholog transcriptomes. We applied outlier analyses within SPSS Statistics v.23 to identify transcripts containing FST outlier SNPs (i.e., SNPs with FST values outside of the 95% C.I. around the mean), which can be interpreted as representing significant genetic differentiation for these genes between these two types of Microphallus. Finally, we used blastx and Blast2GO to annotate FST outlier-containing transcripts and determine putative functions.

GO and KEGG-based functional comparisons of transcriptomes and orthologs

We used blastx (Camacho et al. 2009) and Blast2GO (Conesa et al. 2005) to annotate the orthologous sequences for both Microphallus types. We also annotated the transcripts from each transcriptome that did not fall into orthologous gene sets (nonone-to-one orthologs). We then performed GO functional enrichment analyses, using Fisher’s Exact tests as implemented by Blast2GO (Conesa et al. 2005), to identify overrepresented vs. underrepresented functional groups in the set of orthologous transcripts relative to the rest of the transcriptome, and for the set of nonone-to-one orthologous transcripts relative to the rest of the transcriptome. All functional enrichment analyses were performed on both Microphallus samples. This analysis approach allowed us to compare overrepresented vs. underrepresented functional groups for orthologous and nonorthologous transcripts between the two types of Microphallus in order to identify which functional groups were expressed in both Microphallus samples vs. those functional groups that were uniquely enriched in each Microphallus type. We also predicted KEGG (Kyoto Encyclopedia of Genes and Genomes database) pathways for each transcriptome as implemented by Blast2GO. This analysis allowed us to (1) identify potential among-gene interactions, (2) identify sets of interacting genes of relevance to the Microphallus life cycle stage that we sequenced, and (3) assess whether different pathways are being expressed between the two samples. Finally, we used Blast+ to generate local BLAST databases for both annotated transcriptomes.

Microsatellite loci and primer prediction

We used PrimerPro v.1.0 (https://webdocs.cs.ualberta.ca/~yifeng/primerpro/) to identify potential microsatellite loci in the two Microphallus transcriptomes. We limited these analyses to the ortholog transcriptomes to ensure the transcripts in which we identified microsatellites were distinct loci and to allow future direct comparisons among different types of Microphallus. The PrimerPro v.1.0 pipeline first predicts microsatellite loci by identifying simple sequence repeats (SSRs) using MISA (MIcroSAtellite Identification Tool; http://pgrc.ipk-gatersleben.de/misa/misa.html) and then uses Primer3 (Untergasser et al. 2012) and BLAST (Camacho et al. 2009) to generate primers isolating these loci. After these steps, PrimerPro v.1.0 outputs the putative microsatellites, the primer sequences, their transcriptomic locations, and the primer properties. Following Qiu et al. (2016), we allowed for a minimum of 10 repeats for mononucleotide repeats, six for dinucleotide repeats, five for trinucleotide repeats, four for tetranucleotide repeats, three for pentanucleotide repeats, and three for hexanucleotide repeats, and identified compound microsatellites as instances where there were more than one microsatellite separated by ≤150 nt.

Data availability

The raw RNA-Seq reads are deposited at NCBI Sequence Read Archive under the accessions SRR5170514 and SRR5170515 for PA-Microphallus and PE-Microphallus, respectively. The Transcriptome Shotgun Assembly projects have been deposited at DDBJ/EMBL/GenBank under the accessions GFFK00000000 and GFFL00000000, for PA-Microphallus and PE-Microphallus, respectively. The versions described in this paper are the first versions, GFFK01000000 and GFFL01000000. BLAST databases for each transcriptome are available at http://bioweb.biology.uiowa.edu/neiman/Potamomics.php. SNP and molecular distance data, microsatellite loci with predicted primer details, and transcriptome annotations are deposited at Dryad (doi: 10.5061/dryad.3ft30). Transcriptome annotation statistics and annotation details (see Table S1 and Table S4), a list of orthologous transcript IDs (see Table S2), boxplots to visualize distributions of θ, FST, and pairwise distances (see Figure S1), GO functional enrichment and KEGG analyses of orthologous transcripts (see Table S6 and Table S7), annotations of FST outlier-containing transcripts (see Table S3 and Table S5), and information about predicted microsatellite loci (see Figure S2, Table S8, and Table S9) are provided in the online supplemental information. The custom python program is available at github.com/jsharbrough/grabContigs.

Results and Discussion

Transcriptome assembly and annotation

We generated de novo transcriptome assemblies for stage five Microphallus metacercariae isolated from field-collected PA-Microphallus and PE-Microphallus. Our final transcriptome assemblies contained 15,435 and 29,564 transcripts for PA-Microphallus and PE-Microphallus, respectively. The assembly for PE-Microphallus contained approximately twice as many transcripts and was 33% longer than the assembly for PA-Microphallus. The PA-Microphallus assembly had, on average, 22% longer transcripts and 23% larger N50 than the PE-Microphallus assembly (Table 1). The marked differences in these assembly statistics are, at least in part, likely linked to differences in sequencing depth and technological advances between the two rounds of sequencing but could also reflect, for example, biological differences between Microphallus found in freshwater (inhabiting P. antipodarum) vs. estuarine (inhabiting P. estuarinus) habitats and/or differences in genetic variation (e.g., heterozygosity) of the sequenced samples.

Table 1
Assembly statistics for whole transcriptome assemblies and ortholog transcriptome assemblies for PA-Microphallus and PE-Microphallus (present study) as well as for two previously published Platyhelminthes transcriptomes: the trematode Schistosoma mansoni ...

We obtained both blastx and GO annotations for 60.1 and 46.6% of transcripts and only blastx annotations (including those with GO mapping, but not GO annotation) for an additional 24.6 and 29.8% of transcripts for PA-Microphallus and PE-Microphallus, respectively. There was a mean of ~3.4 GO annotations per transcripts per transcriptome. The remaining 15.2% of PA-Microphallus transcripts and 23.6% of PE-Microphallus transcripts were unable to be annotated, which is typical for de novo assemblies of nonmodel taxa (e.g., Guo et al. 2015; Theissinger et al. 2016). The species that were most frequently the top BLAST hit for a given transcript were most often other trematodes (10,053 transcripts, 77.4% for PA-Microphallus; 12,342 transcripts, 54.9% for PE-Microphallus), or nontrematode members of Platyhelminthes (an additional 332 transcripts, 2.6% for PA-Microphallus; 382 transcripts, 1.7% for PE-Microphallus) (see Table S1).

We used BUSCO to estimate the completeness of our transcriptome assemblies based on the presence, absence, duplication, and/or fragmentation of conserved Metazoan genes. Both transcriptomes contain complete transcripts for ~68% of the 843 conserved single-copy Metazoan genes in the BUSCO database. Our transcriptome assemblies contain gene fragments for an additional 7.8% (PA-Microphallus) and 14.5% (PE-Microphallus) of the conserved single-copy Metazoan genes, for a total of 76.5% (PA-Microphallus) and 82.4% (PE-Microphallus) of the 843 BUSCO genes represented in the transcriptomes (Table 2). Considered together with our assembly statistics (Table 1), these results indicate that our transcriptomes are of good quality, are reasonably complete, and are qualitatively similar to other recently published de novo transcriptome assemblies (e.g., Guo et al. 2015; Hara et al. 2015; Tassone et al. 2016; Theissinger et al. 2016), despite the differences in transcript number and N50 between the two transcriptome assemblies. Because we analyzed transcriptomes generated from only one stage of infection from a parasite with a complex life cycle, at least some of the missing BUSCO genes likely represent genes that are not expressed in metacercariae rather than due to sequencing or assembly limitations. Regardless, because stage five metacercariae are ready for transmission to the final host (Levri and Lively 1996), these data provide an excellent first step toward identifying the genes involved in a critical stage of parasite development.

Table 2
BUSCO assessment for transcriptome completeness

We generated local BLAST databases for both annotated Microphallus transcriptomes that allow for simple queries of nucleotide and protein sequences (http://bioweb.biology.uiowa.edu/neiman/Potamomics.php). This resource will be broadly useful for future studies geared toward identifying candidate genes from Microphallus as well as other related ecologically and/or medically important parasites by providing an easy way to search for genes in a well-characterized host–parasite system. These databases will also be an asset for phylogenetic or molecular evolution-focused studies that would benefit from the addition of data from Trematoda.

Ortholog identification and levels of genetic variation within and genetic differentiation between Microphallus samples

We identified 7584 pairs of one-to-one orthologous transcripts between the two transcriptomes (see Table S2) and 7851 and 21,980 nonone-to-one orthologous transcripts for PA-Microphallus and PE-Microphallus, respectively. The two one-to-one ortholog transcriptomes contain higher mean transcript lengths (18.6% longer for PA-Microphallus; 38.7% longer for PE-Microphallus) and larger N50 values (11.7% greater N50 for PA-Microphallus; 33.9% greater N50 for PE-Microphallus) than their corresponding reference transcriptomes (Table 1). The longer transcripts and higher N50 values in the one-to-one ortholog sets relative to the reference transcriptomes indicate that the one-to-one orthologs represent a set of approximately full-length high quality genes that occur once and only once in both transcriptome assemblies. More than half (PA-Microphallus: 51.7%; PE-Microphallus: 51.2%) of the BUSCO genes are retained in this reduced gene set, further indicating that our filtering steps likely removed poor quality transcripts from our ortholog transcriptomes (Table 2). The annotation statistics for the ortholog transcriptomes are broadly similar to the annotation statistics for the reference transcriptomes (see Table S1), suggesting the ortholog transcriptomes are likely representative of the reference transcriptomes. We also obtained transcriptome assemblies for two previously published Platyhelminthes transcriptomes that were readily available, well-characterized, and among the top-hit species in our transcriptome annotations (http://www.sanger.ac.uk/resources/downloads/helminths/), the trematode Schistosoma mansoni (Protasio et al. 2012) and the cestode Echinococcus granulosus (Tsai et al. 2013). We used these transcriptomes to further assess the quality of our transcriptomes relative to previously published assemblies (Table 1 and Table S1); this comparison reinforces our conclusion that our transcriptomes are of high quality (e.g., higher N50, mean transcript length).

Of the 7584 transcripts in the ortholog transcriptomes, a distinct majority received both blastx and GO annotation for PA-Microphallus (5746 transcripts, 75.8%) and PE-Microphallus (5793 transcripts, 76.4%). An additional 15.9% (1211 transcripts) and 15.8% (1203 transcripts) of these transcripts received only blastx annotations for PA-Microphallus and PE-Microphallus, respectively. The species that were top BLAST hits most often and the species that were hit most frequently in blastx searches include other trematodes (five species), as well as nontrematode Platyhelminthes (two species) and other metazoans with well-annotated genomic and/or transcriptomic resources (see Table S1). Species within Platyhelminthes were the top blastx hit for 90.6% (6337 transcripts) and 90.3% (6356 transcripts) of the orthologous transcripts that received blastx hits for PA-Microphallus and PE-Microphallus, respectively. The remaining 663 annotated orthologs for PA-Microphallus received top BLAST hits from mollusks (393 transcripts, 5.6%), other metazoans (253 transcripts, 3.6%), or nonmetazoan taxa (17 transcripts, 0.2%), and the remaining 682 annotated PE-Microphallus transcripts received top BLAST hits from mollusks (418 transcripts, 5.9%), other metazoans (250 transcripts, 3.6%), or nonmetazoan taxa (14 transcripts, 0.2%). The relatively small proportion of orthologous transcripts with a mollusk as their top hit could reflect low levels of contamination during the process of isolating metacercariae from snail tissue, but is likely also linked in part to availability of well-annotated genomic and/or transcriptomic resources for the specific species that were hit (see Table S1).

We used PoPoolation to calculate allelic diversity using SNPs (Watterson’s θ) for each orthologous transcript by mapping PA-Microphallus and PE-Microphallus reads to their own ortholog transcriptomes. Mean (±SD) θ per transcript for PA-Microphallus mapped to the PA-Microphallus orthologs was 0.0044 (±0.0039) and median θ was 0.0036. Mean (±SD) θ per transcript for PE-Microphallus mapped to the PE-Microphallus orthologs was 0.0051 (±0.0044) and median θ was 0.0046 (see Figure S1A). These estimates of θ are broadly consistent with observations from a previous allozyme-based estimate of θ (0.001–0.021) in several populations of P. antipodarum-infecting Microphallus (Dybdahl and Lively 1996). Comparisons of medians (Mann–Whitney U test) and distributions of θ (Kolmogorov–Smirnov test) for each transcriptome to one another revealed that θ values are significantly different between the two types of Microphallus (P < 0.001 for both analyses), with θ of PE-Microphallus being on average ~0.9 times higher than PA-Microphallus. This result thus provides an initial line of evidence for higher genetic variation in PE-Microphallus relative to PA-Microphallus, though one obvious caveat to definitive interpretation of these results is the differences in sampling between the two types of Microphallus in our study. In particular, because the PA-Microphallus sample was produced from pooled RNA from multiple individuals, we cannot tie sequence differences back to individual worms. On the other hand, because we only sequenced a single PE-Microphallus individual, we can definitively ascribe within-sample variation to individual variation, but are not able to evaluate across-individual polymorphism.

Next, we used PoPoolation2 to estimate FST at each SNP between PA-Microphallus and PE-Microphallus. This analysis allowed us to estimate relative levels of genetic differentiation among populations based on variance in allele frequencies between the two Microphallus samples (Holsinger and Weir 2009). In order to account for the ortholog transcriptome to which reads were mapped, we calculated FST per SNP between PA-Microphallus and PE-Microphallus mapped to the PA-Microphallus ortholog transcriptome and calculated FST per SNP between PA-Microphallus and PE-Microphallus mapped to the PE-Microphallus ortholog transcriptome. We found a mean (±SD) FST per SNP between the two types of Microphallus mapped to the PA-Microphallus ortholog transcriptome of 0.129 (±0.129); median FST per SNP was 0.085. Mean (±SD) FST per SNP between the two types of Microphallus mapped to the PE-Microphallus ortholog transcriptome was 0.148 (±0.138), and median FST per SNP was 0.102 (see Figure S1B). We supplemented these population genetic-based estimates of genetic differentiation with a phylogeny-based divergence estimation approach, using Clustal Omega to generate multiple sequence alignments and to calculate Jukes–Cantor-corrected pairwise molecular distance for each pair of orthologous transcripts. The mean pairwise molecular distance (±SD) between the two transcriptomes was 0.034 (±0.062) and the median was 0.019 (see Figure S1C).

These comparisons provide the first report of which we are aware suggesting that the Microphallus that infect P. antipodarum and P. estuarinus represent distinct genetic lineages. In particular, our FST-based estimates of divergence between these two Microphallus types are an order of magnitude higher than the FST values reported in an early allozyme-based study comparing FST across populations of Microphallus infecting P. antipodarum (Dybdahl and Lively 1996). This result thus provides an initial line of evidence for marked genetic differentiation between these two Microphallus types relative to the across-population differentiation observed for Microphallus infecting P. antipodarum. Our pairwise molecular distance analyses further suggest that these two types of Microphallus are genetically distinct and are consistent with estimates of genetic variation from prior studies from across the genus. O’Dwyer et al. (2014) estimated pairwise molecular distance (uncorrected p-distance) at cox1 to be ~0.002 for lineages of Microphallus that were isolated from the gastropod mollusk Austrolittorina cincta, collected in Otago Harbor, New Zealand. By contrast, pairwise distances between these sequences and previously published cox1 sequences of Microphallus infecting the mud snail Zeacumantus subcarinatus were orders of magnitude higher (0.21–0.25) than within the A. cinta-infecting Microphallus. Another relevant recent example comes from Galaktionov et al. (2012), who measured absolute pairwise differences and p-distances of ITS1 and ITS2 between six Microphallus species that infect gastropods from the genus Littorina from across the Holarctic, and found that pairwise distances within the Microphallus genus infecting congeneric gastropods (Littorina) ranged from 0 to 0.16 at these genes. Our results for mean pairwise molecular distance between PA-Microphallus and PE-Microphallus (0.034) are thus broadly consistent with other reports of variation within and among other Microphallus taxa. These new data also permit some useful comparisons: these pairwise distances are an order of magnitude higher than pairwise distances between Microphallus lineages collected from a single molluscan species (O’Dwyer et al. 2014), an order of magnitude lower than pairwise distances between Microphallus lineages collected from different genera of mollusks (O’Dwyer et al. 2014), and are within the range of previously reported pairwise distances for Microphallus lineages infecting congeneric gastropods (Galaktionov et al. 2012). Taken together, the outcomes of these analyses are consistent with a scenario where PA-Microphallus and PE-Microphallus represent distinct genetic lineages relative to Microphallus collected from different populations of P. antipodarum.

Orthologous transcripts containing FST outlier SNPs

Using the FST analyses described above, we identified 36 FST outlier-containing transcripts between PA-Microphallus and PE-Microphallus when mapping to the PA-Microphallus ortholog transcriptome. Of these 36 transcripts, 31 received blastx and GO annotation, one received only blastx annotation, and four were unable to be annotated (see Table S3). A total of 31 of the annotated transcripts had top blastx hit species within Trematoda (see Table S4). Based on Blast2GO annotations, these FST outlier-containing transcripts appear to be involved in cellular processes, metabolism, biological regulation, and localization (see Table S5). When mapping to the PE-Microphallus ortholog transcriptome, we identified 30 FST outlier-containing transcripts. Of these transcripts, 22 received both blastx and GO annotations, three received only blastx annotation, and three were unable to be annotated (see Table S3). All 25 of the annotated transcripts received top blastx hit species within Platyhelminthes, 24 of which were within Trematoda (see Table S4). The annotated FST outlier-containing transcripts identified when mapping to the PE-Microphallus orthologs appear to be involved in similar biological processes as those identified by mapping to PA-Microphallus (see Table S5).

Only two of the transcripts were identified as FST outliers in both comparisons. Both of these transcripts could represent genes that are evolving relatively rapidly compared to their respective ortholog transcriptome and/or may be potential targets of selection in one or both Microphallus types. These candidate genes are N-α-acetyltransferase, which is involved in the N-terminal acetyltransferase C (NatC) complex and acetylation of amino acids, and permease 1 heavy chain, which is involved in maltose metabolic processes (see Table S3). Genes involved in the NatC pathway have been implicated in modulating stress tolerance in Caenorhabditis elegans (Warnhoff and Kornfeld 2015). In schistosomes (Trematoda), permease 1 heavy chain is characterized as playing an important role in nutrient uptake from hosts and is expressed during all major schistosome life stages (Krautz-Peterson et al. 2007). This information from a related trematode group with a broadly similar life cycle indicates that permease 1 heavy chain might also play a role in nutrient acquisition in Microphallus. More directed study of patterns of molecular evolution and the roles that these two genes play in response to physiological stresses associated with freshwater vs. estuarine environments (N-α-acetyltransferase), as well as nutrient uptake by parasites from hosts (permease 1 heavy chain), in Microphallus will provide more definitive insights into their functions.

GO functional enrichment and KEGG pathway identification

We used Blast2GO to perform functional enrichment analyses comparing our ortholog transcriptomes against their associated reference transcriptomes, with the goal of identifying overrepresented vs. underrepresented functional groups that are shared between the two types of Microphallus. We also used functional enrichment analyses of our nonone-to-one orthologs against their associated reference transcriptomes to assess the types of genes that are overrepresented vs. underrepresented among the transcripts that did not represent one-to-one orthologs between PA-Microphallus and PE-Microphallus. These analyses revealed seven GO terms that were significantly underrepresented among orthologous transcripts in PA-Microphallus relative to the PA-Microphallus reference transcriptome. These same seven functional groups are significantly overrepresented among the nonone-to-one orthologs for PA-Microphallus relative to the PA-Microphallus reference transcriptome (see Table S6). For the PE-Microphallus orthologs relative to the PE-Microphallus reference, we identified 20 significantly underrepresented and six significantly overrepresented functional groups. There were five underrepresented and 14 overrepresented functional groups among the nonone-to-one PE-Microphallus orthologs relative to the PE-Microphallus transcriptome. All but four of the functional groups among the PE-Microphallus transcripts were only significantly overrepresented or underrepresented among either the one-to-one or nonone-to-one orthologs but were never significantly overrepresented or underrepresented in both transcript sets (see Table S6).

There were only two GO categories that were that were overrepresented or underrepresented among orthologs from both types of Microphallus. First, genes with functions related to hydrolase activity are underrepresented among one-to-one orthologs and overrepresented among nonone-to-one orthologs for both PA-Microphallus and PE-Microphallus. The presence of genes involved in hydrolase activity among genes that were significantly functionally enriched is interesting in light of evidence that hydrolases are involved in damaging epithelial lining of the digestive tracts of hosts by the parasitic nematode Contracaeum rudolphii (Dziekońska-Rynko and Rokicki 2005). Second, genes involved in carbohydrate metabolic processes are underrepresented among one-to-one orthologs for both types of Microphallus relative to the reference transcriptomes. Evidence that schistosome parasites dynamically shift carbohydrate metabolic processes throughout the course of their life cycle (Horemans et al. 1992) hint that these genes could also be of relevance to Microphallus. Altogether, both of these functional categories of genes provide a promising set of candidates for assessment of the changes in gene expression and physiological responses associated with the infection process in Microphallus. The marked differences in functional groups that are significantly overrepresented vs. underrepresented in the ortholog transcriptomes relative to the reference transcriptomes between the two Microphallus types suggest that the types of genes present in the two reference transcriptomes are often different. Whether this result is more a consequence of biological differences between PA-Microphallus and PE-Microphallus or differences in sequencing depth cannot be determined at this time but warrants future study.

Finally, we performed KEGG analyses as implemented in Blast2GO to identify the types of enzyme pathways that were being expressed in each transcriptome. This analysis identified 123 KEGG pathways in the PA-Microphallus transcriptome and 128 KEGG pathways in the PE-Microphallus transcriptome (see Table S7). All but 11 of the KEGG pathways that we identified were expressed in both transcriptomes, three were expressed in PA-Microphallus only, and eight were expressed in PE-Microphallus only. Most of the 120 pathways present in both Microphallus transcriptomes are involved in various metabolic processes (49 pathways, 40.2%) or biosynthetic processes (41 pathways, 33.6%). Of the 11 pathways only identified in a single transcriptome, the three pathways identified in only PA-Microphallus include the riboflavin metabolic pathway, the penicillin and cephalosporin biosynthesis pathway, and the biosynthesis of siderophore group nonribosomal peptides. The eight pathways that were identified in only PE-Microphallus include a variety of biosynthesis and metabolic processes (see Table S7). One such pathway, glycosphingolipid biosynthesis, has been implicated in response to salinity stress in Litopenaeus vannamei, the Pacific white shrimp (Chen et al. 2015). This result is suggestive of a scenario where these genes contribute to the ability of PE-Microphallus to tolerate salinity fluctuations associated with estuarine habitats. This important habitat difference between PA-Microphallus and PE-Microphallus is likely to differentially influence the selective pressures experienced by these two parasites and thus has the potential to influence patterns and rates of molecular evolution (e.g., Mitterboeck et al. 2016), making genes involved in salinity tolerance and/or stress particularly interesting candidates for future study.

Predictions of microsatellite loci and primers

We used MISA software to identify SSRs within the ortholog transcriptomes and used Primer3 and BLAST to predict primers that could be used for future microsatellite-based analyses of Microphallus. We identified 109 SSRs in 100 transcripts for PA-Microphallus and 117 SSRs in 107 transcripts for PE-Microphallus. We identified all six types of microsatellites for which we searched, with trinucleotide sequences with at least five repeats representing the most common microsatellite type (62.3% of SSRs identified in PA-Microphallus and 60.7% of SSRs identified in PE-Microphallus) (Table 3; see Figure S2). Eight and four SSRs, respectively, were defined as compound for PA-Microphallus and PE-Microphallus (Table 3). We also used perl scripts available with MISA (http://pgrc.ipk-gatersleben.de/misa/) to compile specific details about the primer sets (e.g., location, size of product, and melting temperature). We generated predicted primers for 84 of the PA-Microphallus SSRs from 100 sequences and for 83 of the PE-Microphallus SSRs from 113 sequences (see Table S8 and Table S9).

Table 3
Summary of potential microsatellite loci for the PA-Microphallus and PE-Microphallus one-to-one ortholog transcriptome assemblies

Future studies will be able to use these markers to characterize, for example, population genetic structure within and genetic differentiation between these two types of Microphallus isolated from two ecologically distinct snail taxa. Comparing patterns of molecular evolution among these orthologous genes from more Microphallus individuals isolated from P. antipodarum and P. estuarinus will allow for research directed at characterizing the strength and efficacy of selection and the importance of population structure in the context of a prominent model for host–parasite interactions and coevolution. Researchers can also use these resources to perform cost-effective analyses incorporating many Microphallus individuals from multiple populations to address key questions regarding the evolutionary connections between these two types of Microphallus (e.g., whether there is gene flow between the Microphallus infecting P. antipodarum and P. estuarinus and whether there is similar evidence for local adaptation and coevolution between P. estuarinus and Microphallus as exists for P. antipodarum and M. livelyi).

Caveats and conclusions

Multiple recent studies have shown that pooled RNA-Seq data can be used to effectively identify SNPs and candidate genes and to perform FST-based measures of genetic differentiation and other population genetic measures (e.g., Fischer et al. 2013; Konczal et al. 2014; Li et al. 2014; Thavamanikumar et al. 2014; Ellison et al. 2016). We used analysis approaches [PoPoolation (used to estimate θ), and PoPoolation2 (used to estimate FST)] that are designed to handle pooled sequencing data and explicitly incorporate number of individuals in the sequencing pools as a parameter in estimations of nucleotide variation and genetic differentiation (Kofler et al. 2011a,b). For these reasons, we believe that we applied the most appropriate means available to analyze our data.

Even so, it is important to acknowledge the potential caveats and limitations associated with pooled RNA-Seq approaches (Konczal et al. 2014). First, in order to reduce the likelihood of including artefactual SNPs, we required that each SNP to be sequenced at least four times and filtered out SNPs with minor allele frequencies <5%, at the risk of missing alleles at low frequencies. Second, differences in gene expression among individuals and/or loci can complicate comparisons among samples. We tried to minimize the effects of these potential expression differences by requiring a minimum of 10× coverage and a maximum of 200× coverage for both the θ and FST estimates. Another concern is that allele frequency estimates may be less accurate in regions of low expression (Konczal et al. 2014). As such, requiring a minimum of 10× coverage should reduce this effect, but does generate potential to miss some lowly expressed SNPs. The existence of allele-specific expression may be problematic in leading to underestimates of the number of heterozygous sites and overestimates of FST in pooled sequencing data. These concerns are at least partially obviated by results of simulations presented in Konczal et al. (2014), who found that allele-specific expression only influences allele frequency estimation in a small fraction (~1%) of genes (in a data set).

In conclusion, our study provides a set of widely useful transcriptomic and genomic resources for an evolutionarily and ecologically important group of parasites. P. antipodarum and M. livelyi are a textbook system for the study of host–parasite coevolutionary interactions, and the data generated herein represent the first large-scale genome-based resources for Microphallus. The annotated transcriptome assemblies, orthologous gene sets, BLAST databases, SNPs, and microsatellite loci and primers will facilitate research for scientists interested in host–parasite coevolution, local adaption, and/or patterns of molecular evolution and phylogenetics of these and other trematode parasites. Our FST, functional enrichment, and KEGG pathway analyses provide a promising set of candidate genes and pathways for future study and further characterization for potential roles in physiological and stress responses relevant to organisms inhabiting freshwater vs. estuarine environments, as well as genes involved in parasitic infection of host taxa.

Acknowledgments

We thank Gery Hehman for RNA sequencing assistance; Curt Lively, Daniela Vergara, Katelyn Larkin, and Mike Winterbourn for field collections; and Joel Sharbrough for the custom python program (github.com/jsharbrough/grabContigs). We are grateful to several anonymous reviewers for constructive feedback on earlier versions of this manuscript. This project was funded by Sigma-Xi (grant number 18911100 BR01 to L.B.), the Iowa Science Foundation (grant number 14-03 to M.N. and L.B.), the National Geographic Society (grant number 9595-14 to K. Larkin), and the National Science Foundation (National Science Foundation-Division of Molecular and Cellular Biosciences grant number 1122176 to M.N.).

Footnotes

Communicating editor: M. Walhout

Literature Cited

  • Andrews, S., 2010 FastQC: a quality control tool for high throughput sequence data. Available at: http://www.bioinformatics.babraham.ac.uk?/projects/fastqc/. Accessed: November 4, 2015.
  • Bankers L., Fields P., McElroy K. E., Boore J., Logsdon J. M., et al. , 2016.  Genomic evidence for population-specific responses to coevolving parasite in a New Zealand freshwater snail. bioRxiv DOI: .10.1101/045674 [Cross Ref]
  • Camacho C., Coulouris G., Avagyan V., Ma N., Papadopoulos J., et al. , 2009.  BLAST+: architecture and applications. BMC Bioinformatics 10: 421. [PMC free article] [PubMed]
  • Carmona D., Connor F. P., Johnson M. T., 2015.  Fifty years of co-evolution and beyond: integrating co-evolution from molecules to species. Mol. Ecol. 2: 5315–5329. [PubMed]
  • Chen K., et al. , 2015.  Transcriptome and molecular pathway analysis of the hepatopancreas in the Pacific white shrimp Litopenaeus vannamei under chronic low-salinity stress. PLoS One 10: e0131503. [PMC free article] [PubMed]
  • Chomczynski P., 1993.  A reagent for the single-step simultaneous isolation of RNA, DNA, and proteins from cell and tissue samples. Biotechniques 15: 532–537. [PubMed]
  • Chomczynski P., Sacchi N., 1987.  Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162: 156–159. [PubMed]
  • Conesa A., Götz S., García-Gómez J. M., Terol J., Talón M., et al. , 2005.  Blast2GO: a universal tool for annotation, visualization and analysis in functional genomics research. Bioinformatics 21: 3674–3676. [PubMed]
  • Cribb T. H., Bray T. A., Littlewood D. T., Pichelin S. P., Herniou E. A., 2001.  The digenea, Chapter 16 pp. 168–173 in Interrelationships of the Platyhelminthes, Vol. 1, Ser. 60. Taylor and Francis, New York, NY.
  • DePristo M., Banks E., Poplin R., Garimella K. V., Maguire J. R., et al. , 2011.  A framework for variation discovery and genotyping using next-generation DNA sequencing data. Nat. Genet. 43: 491–498. [PMC free article] [PubMed]
  • Dybdahl M. F., Lively C. M., 1996.  The geography of coevolution: comparative population structures for a snail and its trematode parasite. Evolution 50: 2264–2275.
  • Dybdahl M. F., Krist A. C., 2004.  Genotypic vs. condition effects on parasite-driven rare advantage. J. Evol. Biol. 17: 967–973. [PubMed]
  • Dziekońska-Rynko J., Rokicki J., 2005.  Activity of selected hydrolases in excretion-secretion products and extracts of adult Contracaecum rudolphii. Wiad. Parazytol. 51: 227–231. [PubMed]
  • Ellison A. R., DiRenzo G. V., McDonald C. A., Lips K. R., Zamudio K. R., 2017.  First in vivo Batrachochytrium dendrobatidis transcriptomes reveal mechanisms of host exploitation, host-specific gene expression, and expressed genotype shifts. G3 7: 269−278. [PMC free article] [PubMed]
  • Fischer M. C., Rellstab C., Tedder A., Zoller S., Gugerli F., et al. , 2013.  Population genomic footprints of selection and associations with climate in natural populations of Arabidopsis halleri from the Alps. Mol. Ecol. 22: 5594–5607. [PMC free article] [PubMed]
  • Galaktionov K. V., Blasco-Costa I., Olson P. D., 2012.  Life cycle, molecular phylogeny and historical biogeography of the ‘pygmaeus’ microphallids (Digenea: Microphallidae): widespread parasites of marine and coastal birds in the Holarctic. Parasitology 139: 1346–1360. [PubMed]
  • Grabherr M. G., Haas B. J., Yassour M., Levin J. Z., Thompson D. A., et al. , 2011.  Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nat. Biotechnol. 29: 644–652. [PMC free article] [PubMed]
  • Gremme G., Steinbiss S., Kurtz S., 2013.  GenomeTools: a comprehensive software library for efficient processing of structured genome annotations. IEEE/ACM Trans. Comput. Biol. Bioinform. 10: 645–656. [PubMed]
  • Gordon A., Hannon G., 2010.  Fastx-toolkit. FASTQ/A short-read pre-processing tools. http://hannonlab.cshl.edu/fastx_toolkit
  • Guo Y., Wiegert-Rininger K. E., Vallejo V. A., Barry C. S., Warner R. M., 2015.  Transcriptome-enabled marker discovery and mapping of plastochron-related genes in Petunia spp. BMC Genomics 16: 726. [PMC free article] [PubMed]
  • Haas B. J., Papanicolaou A., Yassour M., Grabherr M., Blood P. D., et al. , 2013.  De novo transcript sequence reconstruction from RNA-seq, using the Trinity platform for reference generation and analysis. Nat. Protoc. 8: 1494–1512. [PMC free article] [PubMed]
  • Hara Y., Tatsumi K., Yoshida M., Kajikawa E., Kiyonari H., et al. , 2015.  Optimizing and benchmarking de novo transcriptome sequencing: from library preparation to assembly evaluation. BMC Genomics 16: 977. [PMC free article] [PubMed]
  • Hechinger R. F., 2012.  Faunal survey and identification key for the trematodes (Platyhelminthes: Digenea) infecting Potamopyrgus antipodarum (Gastropoda: Hydrobiidae) as first intermediate host. Zootaxa 3418: 1–27.
  • Holsinger K. E., Weir B. S., 2009.  Genetics in geographically structures populations: defining, estimating and interpreting FST. Nat. Rev. Genet. 10: 639–650. [PMC free article] [PubMed]
  • Horemans A. M. C., Tielens A. G. M., van den Bergh S. G., 1992.  The reversible effect of glucose on the energy metabolism of Schistosoma mansoni cercariae and schistosomula. Mol. Biochem. Parasitol. 51: 73–79. [PubMed]
  • Hotez P. J., 2013.  Pediatric tropical diseases and the world’s children living in extreme poverty. J. Appl. Res. Child. 4: 10.
  • Huang Y., Niu B., Gao Y., Fu L., Li W., 2010.  CD-HIT Suite: a web server for clustering and comparing biological sequences. Bioinformatics 26: 680–682. [PMC free article] [PubMed]
  • Jurberg A. D., Brindley P. J., 2015.  Gene function in schistosomes: recent advances toward a cure. Front. Genet. 6: 144. [PMC free article] [PubMed]
  • King K. C., Delph L. F., Jokela J., Lively C. M., 2009.  The geographic mosaic of sex and the Red Queen. Curr. Biol. 19: 1438–1441. [PubMed]
  • King K. C., Jokela J., Lively C. M., 2011.  Trematode parasites infect or die in snail hosts. Biol. Lett. 7: 265–268. [PMC free article] [PubMed]
  • Kofler R., Orozco-terWengel P., De Maio N., Pandey R. V., Nolte V., et al. , 2011. a Popoolation: a toolbox for population genetic analysis of next generation sequencing data from pooled individuals. PLoS One 6: e15925. [PMC free article] [PubMed]
  • Kofler R., Pandey R. V., Schlötterer C., 2011. b Popoolation2: identifying differentiation between populations using sequencing of pooled DNA samples (Pool-Seq). Bioinformatics 27: 3435–3436. [PMC free article] [PubMed]
  • Konczal M., Koteja P., Stuglik M., Radwan J., Babik W., 2014.  Accuracy of allele frequency estimation using pooled RNA-Seq. Mol. Ecol. Resour. 14: 381–392. [PubMed]
  • Krautz-Peterson G., Camargo S., Huggel K., Verrey F., Shoemaker C. B., et al. , 2007.  Amino acid transport in schistosomes: characterization of the permease heavy chain SPRM1hc. J. Biol. Chem. 282: 21767–21775. [PubMed]
  • Levri E. P., Lively C. M., 1996.  The effect of size, reproductive condition, and parasitism on foraging behavior in a freshwater snail, Potamopyrgus antipodarum. Anim. Behav. 51: 891–901.
  • Li W., Jaroszewski L., Godzik A., 2002.  Tolerating some redundancy significantly speeds up clustering of large protein databases. Bioinformatics 18: 77–82. [PubMed]
  • Li J., Witten D. M., Johnstone I. M., Tibshirani R., 2012.  Normalization, testing, and false discovery rate estimation for RNA-sequencing data. Biostatistics 13: 523–538. [PMC free article] [PubMed]
  • Li C., Beck B. H., Fuller S. A., Peatman E., 2014.  Transcriptome annotation and marker discovery in white bass (Morone chrysops) and striped bass (Morone saxatilis). Anim. Genet. 45: 885–887. [PubMed]
  • Lively C. M., Jokela J., 1996.  Clinal variation for local adaptation in a host-parasite interaction. Proc. Biol. Sci. 263: 891–897.
  • Lively C. M., Dybdahl M. F., Jokela J., Osnas E. E., Delph L. F., 2004.  Host sex and local adaptation by parasites in a snail-trematode interaction. Am. Nat. 164: S6–S18. [PubMed]
  • McKenna A., Hanna M., Banks E., Sivachenko A., Cibulskis K., et al. , 2010.  The Genome Analysis Toolkit: a MapReduce framework for analyzing next-generation DNA sequencing data. Genome Res. 20: 1297–1303. [PubMed]
  • Mitterboeck T. F., Chen A. Y., Zaheer O. A., Ma Y. E. T., Adamowicz S. J., 2016.  Do saline taxa evolve faster? Comparing relative rates of molecular evolution between freshwater and marine eukaryotes. Evolution 70: 1960–1978. [PubMed]
  • O’Dwyer K., Blasco-Costa I., Poulin R., Faltýnková A., 2014.  Four marine digenean parasites of Austrolittorina spp. (Gastropoda: Littorinidae) in New Zealand: morphological and molecular data. Syst. Parasitol. 89: 133–152. [PubMed]
  • Osnas E. E., Lively C. M., 2011.  Using definitive host feces to infect experimental intermediate host populations: waterfowl hosts for New Zealand trematodes. N.Z. J. Zool. 38: 83–90.
  • 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. [PMC free article] [PubMed]
  • Qiu Z., Liu F., Lu H., Huang Y., 2016.  Characterization and analysis of a de novo transcriptome from the pygmy grasshopper Tetrix japonica. Mol. Ecol. Resour. DOI: .10.1111/1755-0998.12553 [PubMed] [Cross Ref]
  • Rollinson D., Knopp S., Levitz S., Stothard J. R., Tchuem Tchuenté L. A., et al. , 2013.  Time to set the agenda for schistosomiasis elimination. Acta Trop. 128: 423–440. [PubMed]
  • Routtu J., Ebert D., 2015.  Genetic architecture of resistance in Daphnia hosts against two species of host-specific parasite. Heredity 114: 241–248. [PMC free article] [PubMed]
  • Sievers F., Wilm A., Dineen D., Gibson T. J., Karplus K., et al. , 2011.  Fast, scalable generation of high-qualiy protein multiple sequence alignment using Clustal Omega. Mol. Syst. Biol. 7: 539. [PMC free article] [PubMed]
  • Simão F. A., Waterhouse R. M., Ioannidis P., Kriventseva E. V., Zdobnov E. M., 2015.  BUSCO: assessing genome assembly and annotation completeness with single-copy orthologs. Bioinformatics 31: 3210–3212. [PubMed]
  • Tassone E. E., Geib S. M., Hall B., Fabrick J. A., Brent C. S., et al. , 2016.  De novo construction of an expanded transcriptome assembly for the western tarnished plant bug, Lygus hesperus. BMC GigaScience 5: 6. [PMC free article] [PubMed]
  • Thavamanikumar S., Southerton S., Thumma B., 2014.  RNA-Seq using two populations reveals genes and alleles controlling wood traits and growth in Eucalyptus nitens. PLoS One 9: e101104. [PMC free article] [PubMed]
  • Theissinger K., Falckenhayn C., Blande D., Toljamo A., Gutekunst J., et al. , 2016.  De novo assembly and annotation of the freshwater crayfish Astacus astacus transcriptome. Mar. Genomics 28: 7–10. [PubMed]
  • Trapnell C., Hendrickson D. G., Sauvageau M., Goff L., Rinn J. L., et al. , 2013.  Differential analysis of gene regulation at transcript resolution with RNA-seq. Nat. Biotechnol. 31: 46–53. [PMC free article] [PubMed]
  • Tsai I. J., Zarowiecki M., Holroyd N., Garciarrubio A., Sanchez-Flores A., et al. , 2013.  The genomes of four tapeworm species reveal adaptation to parasitism. Nature 496: 57–63. [PMC free article] [PubMed]
  • Untergasser A., Cutcutache I., Koressaar T., Ye J., Faircloth B. C., et al. , 2012.  Primer3 – new capabilities and interfaces. Nucleic Acids Res. 40: e115. [PMC free article] [PubMed]
  • Van der Auwera G. A., Carneiro M. O., Hartl C., Poplin R., Del Angel G., et al. , 2013.  From FastQ data to high-confidence variant calls: the Genome Analysis Toolkit best practices pipeline. Curr. Protoc. Bioinformatics 43: 11.10.1–11.10.33. [PMC free article] [PubMed]
  • Vergara D., Lively C. M., King K. C., Jokela J., 2013.  The geographic mosaic of sex and infection in lake populations of a New Zealand snail at multiple spatial scales. Am. Nat. 182: 484–493. [PubMed]
  • Wall D. P., Fraser H. B., Hirsh A. E., 2003.  Detecting putative orthologs. Bioinformatics 19: 1710–1711. [PubMed]
  • Warnhoff K., Kornfeld K., 2015.  New links between protein N-terminal acetylation, dauer diapause, and the insulin/IGF-1 signaling pathway in Caenorhabditis elegans. Worm 4: e1023498. [PMC free article] [PubMed]
  • Warren I. A., Ciborowski K. L., Casadei E., Hazlerigg D. G., Martin S., et al. , 2014.  Extensive local gene duplication and functional divergence among paralogs in Atlantic salmon. Genome Biol. Evol. 6: 1790–1805. [PMC free article] [PubMed]
  • Watterson G. A., 1975.  On the number of segregating sites in genetical models without recombination. Theor. Popul. Biol. 7: 256–276. [PubMed]
  • Winterbourn M. L., 1973.  Larval trematode parasitising the New Zealand species of Potamopyrgus (Gastropoda: Hydrobiidae). Mauri Ora 2: 17–30.

Articles from G3: Genes|Genomes|Genetics are provided here courtesy of Genetics Society of America