- Open Access
Genomics and transcriptomics of epizoic Seisonidea (Rotifera, syn. Syndermata) reveal strain formation and gradual gene loss with growing ties to the host
BMC Genomics volume 22, Article number: 604 (2021)
Seisonidea (also Seisonacea or Seisonidae) is a group of small animals living on marine crustaceans (Nebalia spec.) with only four species described so far. Its monophyletic origin with mostly free-living wheel animals (Monogononta, Bdelloidea) and endoparasitic thorny-headed worms (Acanthocephala) is widely accepted. However, the phylogenetic relationships inside the Rotifera-Acanthocephala clade (Rotifera sensu lato or Syndermata) are subject to ongoing debate, with consequences for our understanding of how genomes and lifestyles might have evolved. To gain new insights, we analyzed first drafts of the genome and transcriptome of the key taxon Seisonidea.
Analyses of gDNA-Seq and mRNA-Seq data uncovered two genetically distinct lineages in Seison nebaliae Grube, 1861 off the French Channel coast. Their mitochondrial haplotypes shared only 82% sequence identity despite identical gene order. In the nuclear genome, distinct linages were reflected in different gene compactness, GC content and codon usage. The haploid nuclear genome spans ca. 46 Mb, of which 96% were reconstructed. According to ~ 23,000 SuperTranscripts, gene number in S. nebaliae should be within the range published for other members of Rotifera-Acanthocephala. Consistent with this, numbers of metazoan core orthologues and ANTP-type transcriptional regulatory genes in the S. nebaliae genome assembly were between the corresponding numbers in the other assemblies analyzed. We additionally provide evidence that a basal branching of Seisonidea within Rotifera-Acanthocephala could reflect attraction to the outgroup. Accordingly, rooting via a reconstructed ancestral sequence led to monophyletic Pararotatoria (Seisonidea+Acanthocephala) within Hemirotifera (Bdelloidea+Pararotatoria).
Matching genome/transcriptome metrics with the above phylogenetic hypothesis suggests that a haploid nuclear genome of about 50 Mb represents the plesiomorphic state for Rotifera-Acanthocephala. Smaller genome size in S. nebaliae probably results from subsequent reduction. In contrast, genome size should have increased independently in monogononts as well as bdelloid and acanthocephalan stem lines. The present data additionally indicate a decrease in gene repertoire from free-living to epizoic and endoparasitic lifestyles. Potentially, this reflects corresponding steps from the root of Rotifera-Acanthocephala via the last common ancestors of Hemirotifera and Pararotatoria to the one of Acanthocephala. Lastly, rooting via a reconstructed ancestral sequence may prove useful in phylogenetic analyses of other deep splits.
Seisonidea (also Seisonacea or Seisonidae) includes worm-like creatures up to 2.5 mm in size that live in marine environment on opossum shrimps (Nebalia spec., Crustacea, Leptostraca) [1,2,3,4]. Traditionally, Seisonidea is subsumed under Rotifera or wheel animals, along with Monogononta and Bdelloidea the species of which are mostly free-living . However, as already noticed by Grube in 1861 , there are considerable differences between seisonids and other wheel animals regarding their morphology. Perhaps the most striking one is that seisonids have small ciliary brushes on each side of the mouth opening, but lack a rotatory or wheel organ (also corona), which in monogononts and bdelloids enables swimming and swirling food to the mouth [1,2,3, 6]. Seisonids are further distinguished by the continuous occurrence of well-developed males in addition to females and strictly sexual reproduction [1, 2, 6]. In contrast, males are absent in bdelloids and the sporadically occurring males in monogononts are dwarfed and morphologically simplified [3, 5, 7]. Nevertheless, Monogononta and Bdelloidea clearly belong to the closer phylogenetic relatives of Seisonidea – and they are joined by thorny-headed worms (Acanthocephala), i.e., endoparasites of mandibulate arthropods and gnathostome vertebrates [8, 9]. A monophyletic origin of these four taxa was probably proposed earliest in the middle of the previous century  and gained further impetus from ultrastructural studies in the 1990s [11,12,13]. The members of the group share a syncytial organization of the tegument (epidermis). Referring to this evolutionary novelty, the taxon is designated as Syndermata in part of the scientific literature (e.g., [14, 15]), while other authors prefer to subsume Acanthocephala under broader Rotifera (e.g., ). The monophyly of the group, which we here follow others in calling Rotifera-Acanthocephala , is in any case confirmed by numerous molecular analyses (e.g., [14, 16, 18, 19]).
With the growing number of genome assemblies, it has become increasingly clear that genome and transcriptome metrics vary considerably within Rotifera-Acanthocephala. Examples include haploid genome size estimates of 55 or 211 Mb in monogononts and of 122 Mb or larger in bdelloids [20,21,22]. The latter value is close to approximately 260 Mb of the first acanthocephalan draft genome published . However, while bdelloid genomes carry signatures of genome duplication (e.g., ), the organization of the acanthocephalan nuclear genome primarily testifies to expansion of the repetitive portion . Similarly, the proliferation of transposable elements was considered to be causative for the larger genomes inside Monogononta . This complexity impairs deriving a scenario of genome evolution within Rotifera-Acanthocephala. Further hampering the issue is the lack of comprehensive comparative data on a key taxon, the Seisonidea. In fact, for seisonids, solely mitochondrial gene sequences [14, 16, 18], a set of expressed sequence tags  and one mitochondrial genome  were available previous to this study. Reconstruction of first seisonid genome and transcriptome drafts would therefore significantly broaden the data basis for evolutionary analyses of Rotifera-Acanthocephala.
A broader molecular data basis might also shed new light on diversity within seisonids, for which four species have been described so far [1, 2, 25, 26]. In comparison, numbers of described species are many times higher for Monogononta (1570 species ), Bdelloidea (461 ) and Acanthocephala (ca. 1200 ). And these counts are growing, which partially reflects the discovery or resurrection of morphologically distinguished species (e.g., [29, 30]). For example, there is increasing evidence that the acanthocephalan Pomphorhynchus laevis might be a collective species containing several morphologically distinct lineages [23, 31]. In addition, molecular analyses have uncovered more and more cryptic species in monogononts, bdelloids and acanthocephalans [32,33,34]. Only for seisonids, cryptic speciation has not yet been reported although it might occur in this group as well. Grube, for example, distinguished four seisonid species in the Gulf of Naples, Italy, alone . And while this view has not gained acceptance, it is clear that different seisonid species can coexist on single opossum shrimps. This illustrates that co-existence of genetically distinct lineages is possible in seisonids .
Comprehensive genomic data may further elucidate the phylogenetic position of seisonids within the Rotifera-Acanthocephala clade. In fact, seisonids appeared as the acanthocephalan sister-group in part of the sequence-based analyses, especially when implementing measures for mitigating potential long branch attraction (LBA) [18, 23]. Such position of seisonids would be in accordance with mitochondrial gene order and ultrastructural data [11, 12, 24, 36] and also with combined analysis of morphological and molecular data . A monophylum containing Seisonidea and Acanthocephala may be referred to as Pararotatoria Sudzuki, 1964, a taxon name originally applied to Seisonacea or Seisonidea alone . Pararotatoria, in turn, could be the sister-group of Bdelloidea, which together would establish Hemirotifera (Bdelloidea+Pararotatoria) [19, 24, 37]. Yet, seisonids were basally branching in other molecular analyses, thus being sister to a clade comprised of monogononts, bdelloids and acanthocephalans [14, 16, 38]. In the respective trees, bdelloids grouped together with acanthocephalans, implying monophyly of Lemniscea as suggested by Lorenzen . Correspondingly, the phylogeny inside the Rotifera-Acanthocephala clade has repeatedly been regarded as unsettled [4, 5]. Moving further to unravel the phylogenetic relationships of monogononts, bdelloids, seisonids and acanthocephalans would thus be welcome and all the more so, as it should lead to a better understanding of the evolution of genomes and lifestyles inside the entire clade.
The present study aims at shedding light on the aforementioned issues. For achieving this, we present the first nuclear genome and transcriptome drafts for Seisonidea, as generated from next generation sequencing (NGS) data. We additionally screened for potential hidden diversity within specimens morphologically determined as S. nebaliae Grube, 1861 . Furthermore, we compared genome and transcriptome metrics within Rotifera-Acanthocephala. Finally, we re-evaluate the phylogenetic position of Seisonidea under particular consideration of potential LBA. Based on the results of tree reconstructions, we infer scenarios of how genomes and lifestyles might have evolved inside the Rotifera-Acanthocephala clade.
Two mitochondrial genomes in pooled S. nebaliae sample
MEGAHIT assembled two mitochondrial sequences based on DNA from 594 specimens determined morphologically to belong to S. nebaliae. With 15,114 bp, one of the two sequences was almost identical in length to the 15,120 bp long mitochondrial genome previously published for S. nebaliae (KP742964.1). As identity was high between these two sequences (99.9%), we collectively denote these as haplotype A and refer to Sielaff et al.  for a detailed description. According to the best hit from BLASTN (for references to programs see Methods), the second mitogenome sequence belonged to S. nebaliae too. However, with ca. 82.0% this consensus sequence showed considerably less nucleotide identity with the previously published reference (based on an alignment of 14,993 bp as compiled with MAFFT and Gblocks). In the following, we regard the corresponding mitogenome as haplotype B. Both haplotypes contained the 12 protein-coding genes, 2 rRNA genes and 22 tRNA genes to be expected for Gnathifera. Thus, tRNAs for lysine and serine were represented by two genes, each, while atp8 was lacking (e.g., [40, 41]). Haplotypes A and B also shared an identical gene order and all genes were encoded by the heavy strand in both of them (Fig. 1). Furthermore, RNA-Seq reads (enriched for coding sequences) mapped to all rRNA- and protein-coding genes in both mitogenomes (Fig. 2), thus indicating conserved transcription. On the other hand, with 16,246 bp, haplotype B exceeded haplotype A by about 1130 bp, due to length polymorphism of the non-coding regions (NCRs). Across the entire sequence of haplotype B, average gDNA read depth was 9111 (Bowtie 2). With 648, read depth was approximately 14-fold lower in haplotype A. In turn, RNA reads mapped with lower multiplicity to haplotype B (average 4598) than haplotype A (average 14,012).
Subsequently, we verified that the new haplotype B is attributable to S. nebaliae by phylogenetic analyses. Due to limited representation of orthologues in our WGA-based NGS dataset for Paraseison annulatus Claus, 1876 the approach was confined to cox1 and nd1 sequences. In unrooted maximum likelihood (ML) and Bayesian inference (BI) trees, none of the sequences from this study clustered with orthologues of crustaceans, thereunder a Nebalia species (see Supplementary Table S1, Additional file 1). In particular, all Seison sequences persistently formed a cluster, as exemplified by the BI tree in Fig. 3. Thus, the new mitogenome does not result from contamination of the sequenced sample with DNA of its host, Nebalia bipes, or the second seisonid species living on that crustacean, P. annulatus. Mitogenome-based tree reconstruction hence suggests variation within S. nebaliae. However, the extent of variation is so strong, given the 82% sequence identity mentioned above, that we consider the haplotypes found here as a first indication of distinct lineages within S. nebaliae.
Metrics of genome assemblies provide further evidence for distinct lineages in S. nebaliae
The appearance of two different mitochondrial genomes led us to take into consideration that two distinct lineages (A and B) of S. nebaliae could be included in the data. To further examine this working hypothesis we applied two alternative strategies for assembling the nuclear genome with MEGAHIT. Nuclear genome assembly 1 (GA1) was reconstructed without stringent filtering against less abundant reads. The assembly should thus represent nuclear genomes A and B, besides potential foreign DNA. In contrast, genome assembly 2 (GA2) was intended to be built from genome B reads only. For achieving this, we applied a multiplicity threshold of 20. The threshold was a slight enhancement compared to the multiplicity ratio of mitochondrial haplotypes as derived by mapping of gDNA reads.
Genome assembly 2 was more coherent compared to GA1 (Table 1). Specifically, GA2 had almost twice the N50 value of GA1 (ca. 45.7 kb in GA2, ca. 26.0 kb in GA1). Furthermore, GA2 contained disproportionally less contigs (3301 in GA2 versus 14,609 in GA1). Complementary search for BUSCO genes underlined better quality of GA2 relative to GA1 in fewer contigs of bacterial origin in the first relative to the latter (Table 1). This was additionally reflected in a lower error rate for GA2 relative to GA1, while the inverse relation held for the consensus quality value (Table 2). Nevertheless, Merqury gave the accuracy in consensus base calling with 99.9% (Q30) for GA1 and 99.99% (Q40) for GA2 (Table 2). Thus, the sequence information per se was of high quality in both assemblies.
With about 43.9 Mb, the length of GA2 undershot the GenomeScope estimate by only 1.8 Mb. Correspondingly, GA2 should cover approximately 96% of the haploid nuclear genome (B) of S. nebaliae. Moreover, GA2 spanned less than half of the 99 Mb in GA1 (Table 1). This is consistent with the expectation for GA2 representing genome B mainly and GA1 including genomes A and B in addition to a smaller fraction of contigs assembled from foreign DNA. Near-to completeness of GA1 and GA2 was underlined by additional Merqury estimates. In particular, k-mer completeness was given with about 93% for GA1 and about 92% for GA2 (Table 2). The difference between both estimates was in line with the fact that k-mers originating from foreign DNA were less contained in GA2 than GA1. These were included in k-mer distributions in low-multiplicity areas (Figs. 4, 5 and 6A, C). Thus, the application of a multiplicity threshold (20 x) effectively tailored GA2 to contain genome B contigs.
Distinct lineages in S. nebaliae were further reflected in a higher GC content of GA1 (34.0%) than GA2 (31.9%) (Fig. 7) and the rates at which mRNA-Seq reads mapped to both genome assemblies. In fact, also the RNA used for NGS was extracted from a pool of S. nebaliae specimens and hence should represent both lineages, although not necessarily at the same ratio as the DNA pool. In line with this, the mapping rate in the case of GA2 (32.8%) was clearly smaller than in GA1 (91.8%). In contrast, back-mapping rates of gDNA reads prior and after application of a multiplicity cutoff (Bowtie 2) were very similar in both genome assemblies (Table 1).
Representation of two nuclear genomes in GA1 additionally emerged from a high proportion of duplicated BUSCO Metazoa genes in this assembly (Table 1). In fact, 26.9% of the corresponding genes in GA1 were reported to be duplicated, whereas only 0.3% were considered duplicated in GA2. Specifically, out of 257 BUSCO Metazoa genes that occurred multiple times in GA1, 256 were duplicated. Of these, 252 occurred once each in GA2. In GA1, the corresponding gene pairs resided on contigs with markedly different multiplicities, whereby the median for high-multiplicity contigs (132 x) was about nine times higher than that for the low-multiplicity contigs (15 x). We observed the same ratio in all high and low multiplicity contigs of GA1 (see below). Furthermore, the direction of the ratio was the same in mitochondrial haplotypes B and A (see above). Thus, S. nebaliae specimens carrying mitochondrial haplotypes A and B were most probably identical with carriers of nuclear genomes A and B, respectively.
Results of read depth analysis is consistent with lineage formation in S. nebaliae
Analysis of k-mers provided further evidence for distinct lineages in S. nebaliae. In particular, k-mers that GenomeScope v. 1.0 classified as putative errors (1.3%) should partially have originated from less abundant carriers of nuclear genome A (Fig. 4: light blue fill to the left of the red curve). In turn, k-mers to the right of the red curve in Fig. 4 should primarily reflect more abundant DNA of S. nebaliae specimens that carried nuclear genome B. Based on B k-mers, GenomeScope estimated haploid genome size to be in the range of 45.7 Mb, with non-repetitive (unique) and duplicated portions of 81.3 and 2.1%, respectively. Probably due to genetic variation within B carriers, heterozygous k-mers peaked at higher multiplicity (coverage in GenomeScope terminology) than in the model graph for a diploid organism. In contrast, the larger peaks giving homozygous portions were at very similar multiplicity values in k-mers representing genome B and the model curve (for interpretation, see [42, 43]). The smaller peak of the blue-fill area in a range of ca. 200–250 x most likely indicated the duplicated portion in the nuclear genome of S. nebaliae.
Back-mapping of the gDNA reads used for assembling the contigs confirmed the intended pattern (Fig. 5). Thus, the graph for GA1 (dashed line) showed peaks in the low-multiplicity range (7 x and 15 x) which most likely corresponded to the heterozygous and homozygous portions in nuclear genome A in addition to foreign DNA. Two additional peaks at higher multiplicity values (81 x and 131 x) had smaller frequencies and probably represented genome B contigs. The corresponding distribution in GA2 (solid line) was confined to two stronger peaks in a higher multiplicity range. These should give the heterozygous and homozygous portions in nuclear genome B once more. Very similar multiplicities of these two peaks (79 x and 132 x) and their counterparts in GA1 (see above) reinforced that GA1 contained genome B contigs in addition to genome A contigs. Taking the homozygous peaks for both S. nebaliae genomes in GA1 as a reference, nuclear genome B (131 x) should have been nine times as abundant as genome A (15 x) in the in the DNA pool used for library preparation.
The previous pattern was reproduced in Merqury  analysis, where k-mers were mapped to the genome assemblies. Thus, GA1 and G2 shared k-mers peaking at higher multiplicity values of 72 x and 118 x (Fig. 6). These peaks corresponded to the ones at 77 x and 131 x in GenomeScope analysis (Fig. 4), thus representing heterozygous and homozygous proportions in the haploid nuclear genome of S. nebaliae genome B [42,43,44]. Slightly deviant multiplicity values for the two peaks in Merqury analysis in relation to GenomeScope analysis probably reflected differences in the settings. In particular, Merqury was run with a k of 18 whereas k was 21 in GenomeScope analysis. Either way, as in GenomeScope analysis (Fig. 4), the distribution for GA2 k-mers exhibited an additional smaller elevation at a multiplicity of 224 x, which should have represented the duplicated portion in the haploid nuclear genome of S. nebaliae (Fig. 6B). In contrast, the distribution for GA1 displayed a prominent k-mer peak at lower multiplicity (8 x), which in GenomeScope analysis was contained in the area left of the red line (Fig. 4).
Small repetitive portion and only few SINEs in the nuclear genome of S. nebaliae
According to Repeatmasker, long interspersed nuclear elements (LINEs) made up the largest share of the repetitive portion in GA2 (4.5%), followed by unclassified repeats (4.3%), DNA elements (2.4%), long terminal repeats (LTR: 2.2%) and simple repeats (2.2%). Yet, only a few short interspersed nuclear elements (SINEs) were annotated (0.03%). Altogether, the repetitive portion in GA2 amounted to 16.2% or 7.1 Mb according to Repeatmasker (Additional file 1). Thus, ca. 83.8% of the S. nebaliae nuclear genome should be repeat-free. This is in accordance with the range of the repeat-free complement as estimated by GenomeScope (36.7–37.2 Mb), thus reinforcing the quality of GA2.
Transcriptome metrics are in accordance with lineage formation and gene loss in S. nebaliae
Transcriptome assemblies 1 and 2 (TA1 and TA2) were highly congruent with the respective genome assemblies used for their guided reconstruction with Trinity . For example, only five of the 34,859 TA2 transcripts could not be mapped with GMAP  to GA2. The median score of all mappings was 98 (out of 100), the average score was 93. In 96% of the cases, the mapping score was 80 or higher (Additional file 2). More importantly, different GC contents of both draft transcriptomes (TA1: 36.0%, TA2: 34.0%) were consistent with distinct S. nebaliae lineages again, as were additional metrics of both transcriptome assemblies (Table 3). In particular, TA2 contained about half as many transcripts of 200 bp or more (~ 35,000) than TA1 (~ 72,000). These were condensed to about half as many SuperTranscripts in TA2 (~ 23,000) than TA1 (~ 47,000). In addition, SuperTranscripts spanned about 27 Mb in TA2 and about double as much in TA1 (56 Mb). According to quotients of transcriptome and genome spans (TA1/GA1, TA2/GA2), 56.7 and 62.1% of GA1 and GA2, respectively, should be occupied by coding sequences. As to be expected for coding sequences, their GC contents were overall increased in relation to the genome assemblies they were residing in (Tables 1, 3). Not least, both transcriptome assemblies testified to marked gene loss in S. nebaliae. In fact, 34.3 and 35.2% of the screened BUSCO Metazoa genes were classified as missing in TA1 and TA2, respectively (Table 3). This was close to corresponding percentages in GA1 (35.4%) and GA2 (36.5%) as reported in Table 1.
Similar to GA2 (0.3%), the proportion of duplicated BUSCO Metazoa genes was low in TA2 (1.8%) (Tables 1, 3). Recalling above evidence from frequency distributions, much higher proportions of “duplicated” genes in GA1 (26.9%) and TA1 (41.1%) presumably reflected the representation of gene pairs in nuclear genomes A and B. We took advantage of this fact and extracted cDNAs of the presumed gene pairs of genomes A and B, which BUSCO had classified as duplicated, from GA1. Of these, we retained 100 gene pairs residing on contigs with read depth differences in the range of 8–14 x, which corresponded to slightly relaxed demands when compared to the multiplicity ratios in mitochondrial haplotypes and genomic contigs (see above). As another precondition for further consideration of a gene pair, orthologues had to occur in all the species used for downstream comparisons. Plotting GC contents of the S. nebaliae contigs, which fulfilled these criteria, against their individual multiplicity values supported the genetic distinctness of genomes A and B. As in the aforementioned genome and transcriptome comparisons, the GC content was higher in the selection of contigs carrying BUSCO Metazoa genes representing genome A (mean = 36.2%, median = 36.1%) than in their counterparts representing genome B (mean = 31.5%, median = 31.2%) (Fig. 7). The difference was significant according to Mann-Whitney U (MWU) test (FDR = 0.000). In addition, the coding regions of the 100 BUSCO Metazoa genes analyzed exhibited differential codon usage at the third position. In particular, the skew toward AU into prospective S. nebaliae genome B was much more prominent than in presumed genome A (Fig. 8).
Taking the different lines of evidence (mitochondrial haplotypes and nuclear genomes with equidirectional ratios of abundancies, differential GC content, percentages of duplicated BUSCO genes, differential codon usage etc.), we consider the variation observed as indicative of strain formation in S. nebaliae. Thus, we will collectively refer to S. nebaliae specimens carrying mitochondrial haplotype A and nuclear genome A as to strain A. Carriers of mitochondrial haplotype B and nuclear genome B will be designated strain B henceforth.
Divergent gene compactness within Rotifera-Acanthocephala
We compared the aforementioned 100 BUSCO Metazoa genes of S. nebaliae strains A and B with their orthologues in the monogonont Brachionus plicatilis, bdelloid A. vaga, and acanthocephalan P. laevis. In support of the orthology of the proteins encoded, levels of protein length did not differ significantly across the five taxa studied according to Kruskal-Wallis (KW) test (FDR > 0.05; Fig. 9). In contrast, levels of gene length as derived from BUSCO annotations were significantly different across the five taxa (FDR = 0.000; KW test). Thereby, length distributions were most similar in both S. nebaliae strains and A. vaga, whereas gene annotations had overall larger spans in B. plicatilis and P. laevis (Fig. 10). Recalling similar levels of protein length, differences in gene length must have reflected divergent intronic spans. Thus, genes were the least compact in P. laevis, followed by B. plicatilis, while gene compactness was highest in A. vaga and both S. nebaliae strains.
Gene repertoire size in epizoic S. nebaliae ranges between corresponding numbers for (other) wheel animals and the acanthocephalan
Genome assembly 2 contained 606 BUSCO Metazoa genes classified as single copy, duplicated and fragmented. This was less than identified in the draft genomes for the monogonont B. plicatilis (863) and the bdelloid A. vaga (855). Still, the number of recognized genes in S. nebaliae was higher than in the acanthocephalan P. laevis (440). Missing BUSCO Metazoa genes showed the expected inverse order, with 91 in B. plicatilis, 99 in A. vaga, 348 in S. nebaliae strain B and 514 in P. laevis. In addition, the degenerate tetraploid A. vaga  had the fewest complete single copy (178) and by far the most duplicated BUSCO Metazoa genes (662). On the contrary, there were almost exclusively single copy genes in the other three genome assemblies. For S. nebaliae, this pattern was consistent with above results from k-mer analyses, according to which the duplicated portion of GA2 was small (Figs. 4, 6B).
Since genes classified as missing might actually have been overlooked due high divergence, we repeated the search for BUSCO Metazoa genes with the training option of the algorithm enabled. These additional analyses were confined to S. nebaliae (GA2) and P. laevis, since numbers of BUSCO Metazoa genes for the monogonont and the bdelloid already exceeded 90% without training. However, the second run did not result in a substantial increase in the number of BUSCO Metazoa genes detected. In fact, the training resulted in very similar percentages of missing genes (S. nebaliae: 35.5%, P. laevis: 52.3%) when compared to the values reported in Table 4 (S. nebaliae: 36.5%, P. laevis: 53.9%). This prompted us to additionally screen the S. nebaliae and P. laevis draft genomes for BUSCO Eukaryota genes, which are commonly regarded as being more conserved than metazoan ones. Although this approach uncovered relatively more genes, the general pattern was reproduced. Thus, the percentage of missing genes remained considerable in the S. nebaliae draft genome (GA2: 21.1%) and still was increased in the P. laevis draft genome (31.5%).
In a complementary approach, we checked for the presence of genes coding for transcription factors with ANTP-type homeodomains (Fig. 11). Corresponding screens failed to determine any member of five ANTP-type gene families (ro, nedX, xlox, hlx and abox) in the monogonont B. plicatilis, the bdelloid A. vaga, the seisonid S. nebaliae strain B (GA2) and the acanthocephalan P. laevis. The remaining 35 gene families searched occurred in at least one of the four draft genomes analyzed. Their presence-absence patterns yielded additional hints for an intermediate comprehensiveness of the seisonid gene repertoire, as observed before in BUSCO Metazoa genes. Thus, out of the aforementioned 35 ANTP-type gene families, B. plicatilis and A. vaga lacked four and two, respectively. With 18, more than half of the 35 ANTP-type gene families appeared to be absent in the draft genome of the acanthocephalan P. laevis. With eight ANTP-type gene families, for which no member was found, GA2 occupied the anticipated intermediate position once more. Furthermore, the draft genomes of S. nebaliae and P. laevis consistently lacked evidence of hox 5, mnx/hb9, gsx and nk4 (see black x’s in Fig. 11). In turn, there was no exclusive absence of any ANTP-type gene family in the bdelloid and acanthocephalan draft genomes.
Phylogenomics: rooting with the LCA places Seisonidea as sister to Acanthocephala
Seison nebaliae clustered with the platyhelminth Schmidtea mediterranea in unrooted BI and ML trees inferred from an alignment containing translated sequences of the above 100 BUSCO Metazoa genes (Concat100-6OTUs; Fig. 12A). In contrast, S. nebaliae grouped together with the acanthocephalan P. laevis in additional BI and ML trees, which we had derived from edited alignments. In particular, the alternative topology was reflected in unrooted ML and BI trees built from an alignment devoid of singleton positions and lacking the platyhelminth (Concat100-noSing-5OTUs; Fig. 12B). However, the branch to S. nebaliae was disproportionately long because amino acid states unique to S. nebaliae but shared by both strains were not recognized as singletons by the script used for editing the alignment. To account for this limitation, we additionally generated alignments, which were pruned from singleton positions and lacked either S. nebaliae strain A or B in addition to the platyhelminth (Concat100-noSing-4OTUs-A; Concat100-noSing-4OTUS-B). This enabled that character states unique to S. nebaliae were recognized as such, followed by removal of the corresponding alignment positions. Notably, branch lengths were homogenized in the resulting trees and, regardless of the strain kept, S. nebaliae clustered with P. laevis (Fig. 12C; Additional file 1). Support for the internal branch separating the seisonid-acanthocephalan cluster and the monogonont-bdelloid cluster reached the maximum value in all corresponding BI and ML trees (for a tabular summary, see Table S3 in Additional file 1). Collectively, these results suggested that the long seisonid branch in the tree derived from the unedited Concat100 alignment was drawn to the long outgroup branch (Fig. 12A). To avoid this, we inferred the Concat100 sequence for the last common ancestor (LCA) of Rotifera-Acanthocephala. This was done with an intree assuming a star-like topology for Rotifera-Acanthocephala. Thus, no particular phylogenetic hypothesis was implied. Prior to subsequent tree reconstruction, we replaced the platyhelminth by the LCA sequence (Concat100-6OTUs-LCA). Corresponding BI and ML trees consistently yielded maximum support for monophyletic Pararotatoria (Seisonidea+Acanthocephala) within maximally supported Hemirotifera (Bdelloidea+Pararotatoria) (Fig. 12D). The tree additionally implied paraphyly of Eurotatoria (Monogononta+Bdelloidea) and Lemniscea (Bdelloidea+Acanthocephala).
Two strains of S. nebaliae in the tidal flats off Roscoff
Based on DNA-Seq data for which DNA had been extracted from a pool of S. nebaliae specimens, we reconstructed two mitochondrial genome sequences (Fig. 1). One of the two was almost identical with a previously published mitogenome sequence of S. nebaliae , while the second one was novel. Both sequences exhibited considerably low sequence identity (82.5%) and markedly different lengths (strain A: 15.1 kb; strain B: 16.2 kb). Nevertheless, their sizes remained within the range of 13.6–16.8 kb reported for other members of the Rotifera-Acanthocephala clade [40, 47, 48] and phylogenetic analyses confirmed their origin in S. nebaliae (Fig. 3; Additional file 1). In addition, both mitogenomes showed the same gene order, which is unlikely to be coincidental . As transcriptional analyses were supportive of their functionality (Fig. 2), we refer to these mitochondial genomes in S. nebaliae as to haplotypes A and B. Since the almost complete assembly of the haploid nuclear genome of S. nebaliae (GA2: ca. 44 Mb) would fit approximately twice into a second assembly reconstructed under relaxed conditions (GA1: ca. 99 Mb), a total of two nuclear genomes should have been represented in the sequenced DNA(s). In line with this, we found evidence of variation between the presumed nuclear genomes in S. nebaliae, in terms of GC content, codon usage, and gene compactness (Figs. 7, 8, 9 and 10). Distributions of k-mer frequency and multiplicity provided additional hints for the presence of genome variants in S. nebaliae (Figs. 4, 5 and 6). Thereby, a common thread was that both the mitogenome and the nuclear genome were represented in a clearly more (B) and a less abundant variant (A). This consistency is hardly compatible with the occurrence of both mitochondrial haplotypes in single individuals of S. nebaliae, as reported for other Metazoa (e.g., ). Rather, the carriers of mitochondrial haplotypes A and B should be identical with specimens having genome variants A and B, respectively. Thus, there should be genetically more or less separated lineages within S. nebaliae in the tidal flats off Roscoff, France.
The degree of mitochondrial sequence dissimilarity (17.5%) would also justify the postulation of separate species, as done in other cases based on smaller values (e.g., ). Cryptic speciation in seisonids would fit well into the general picture of Rotifera-Acanthocephala, considering corresponding reports for Monogononta, Bdelloidea and Acanthocephala [32,33,34]. On the other hand, molecular features such as low mtDNA identity alone are not necessarily sufficient for species delimitation [52, 53] and revision of the Seison taxonomy seems premature to us as long as potential morphological or ecological specificities of both strains/species have not been checked for. For example, the lineages might differ in their spatial and nutritional niches on the common host N. bipes as it is commonly assumed for S. nebaliae and P. annulatus . Both S. nebaliae lineages might also show differential population dynamics in dependence of seasonal temperatures as described for sessile rotifers before . As these questions have not yet been adressed, we leave it at the distinction of S. nebaliae strains in the present study.
Consistent genome and transcriptome metrics in S. nebaliae and other members of Rotifera-Acanthocephala
The GC content found for S. nebaliae was within the previously known boundaries for Rotifera-Acanthocephala. However, a closer look shows that the GC content in the seisonid reference genome (GA2: 31.9%) was first of all very similar to identical to draft genomes of an acanthocephalan (P. laevis: 32.9%) and several bdelloids (A. vaga: 31.2%, A. ricciae: 35.6%, R. magnacalcarata: 31.9%, R. macrura: 32.6%) [21, 23]. Overall, their values were markedly higher than for the draft genomes of the monogononts B. calyciflorus (24.2%) and B. plicatilis (26.4%) [55, 56]. In light of the tree shown in Fig. 12D, the nuclear genome of the LCA of Hemirotifera might thus have had a GC content of ca. 30% or higher. In addition, the total number of nuclear genes in seisonids should be within the range of ca. 11,000 to ca. 69,000 as reported for other members of the Rotifera-Acanthocephala clade [21, 23, 55,56,57,58]. The number of SuperTranscripts inferred here for S. nebaliae (ca. 23,000) at least does not contradict such a vague estimation (Tables 1, 3).
Measures counteracting attraction to the root place the seisonid next to the acanthocephalan branch
The question remains why Seisonidea clustered with Acanthocephala in part of the previous molecular analyses (e.g., [18, 19]; see also Supplementary Fig. S1 to ), but otherwise branched basally within Rotifera-Acanthocephala. Present ML and BI based tree reconstructions add to previous hints that misleading signal could be the reason for a basal branching of Seisonidea . Indeed, the seisonid branch was the longest in trees in which the lineage diverged basally [38, 59]. However, it is getting increasingly clear that attraction of a long ingroup branch to the outgroup or root leads to systematic error in phylogenetic analyses (e.g., ). We found a strong indication of this phenomenon in the unrooted tree derived from 100 concatenated BUSCO Metazoa genes (Fig. 12A): Here, S. nebaliae grouped together with the outgroup representative S. mediterranea. Rooting the tree with the latter would hence result in a basal branching of S. nebaliae. But after implementation of measures preventing LBA between the platyhelminth and S. nebaliae, the latter clustered with P. laevis (Fig. 12B,C). A requirement for the rearrangement of the tree topology in analyses of the edited datasets was the combined removal of the platyhelminth and of singletons (see Additional file 1 for a detailed discussion). At this point, it remains to be noted that editing of the initial dataset provided clues according to which artificial signal might be the reason for a basal branching of S. nebaliae. A significant part of the problem is likely to be attraction to the outgroup.
Given hundreds of millions of years that will have elapsed elapsed since the split of Rotifera-Acanthocephala and any other lineage with extant species (reviewed in ), outgroups will always be highly divergent. Thus, attraction of a particular ingroup branch to the root would almost certainly persist in a wide range of outgroup species. In fact, corresponding results of other studies can be interpreted in this sense [38, 59]. Against this background, we consider rooting via a reconstructed ancestral sequence as a reasonable way to address the problematic resolution of deep splits. In the present study, the approach stably led to monophyletic Pararotatoria (Seisonidea+Acanthocephala) within monophyletic Hemirotifera (Bdelloidea+Pararotatoria) (Fig. 12D; Additional file 1 [19, 24, 37]). This adds to previous findings that Eurotatoria (Monogononta+Bdelloidea) is probably a paraphyletic assemblage [14, 38, 61,62,63]. Furthermore, Lemniscea would be paraphyletic according to the BI tree topology in Fig. 12D and the corresponding ML tree in Additional file 1. Actually, it would not even be possible to root the BI trees in present Fig. 12B, C and their ML counterparts in a way that monophyletic Lemniscea emerge. There also seems to be no morphological feature in support of monophyletic Lemniscea. In particular, the eponymous lemniscs, neck-based invaginations of the syncytial tegument in Acanthocephala, do not correspond to cellular hypodermal cushions supporting the wheel organ in Bdelloidea . In contrast, particular filament bundles in the tegument and two rows of electron-dense bodies alongside the sperm flagellum could be evolutionary novelties shared by Seisonidea and Acanthocephala [4, 11,12,13, 36]. Not least, mitochondrial gene order was shown to be in line with monophyly of Pararotatoria and Hemirotifera .
Genome evolution within Rotifera-Acanthocephala
The likely phylogenetic relationships depicted in Fig. 12D additionally implicate that the LCA of Rotifera-Acanthocephala may have passed a genome of about 50 Mb, as occurring inside Monogononta , to the descendant lineages of Hemirotifera and Pararotatoria. Thus, a smaller genome like in S. nebaliae (GA2: ca. 46 Mb) would have resulted from reduction as it appears to be reflected in the gene repertoire as well (see below). In turn, size increases should have independently occurred in the evolution of Bdelloidea, Acanthocephala and some Monogononta. Thus, within Monogononta, larger genome size has been associated with the spread of transposable elements . In Bdelloidea, genome duplication laid the basis for greater genomes although subsequent losses of genome parts obscure the original pattern of tetraploidy [20, 21, 65,66,67]. In line with this, we found the percentage of duplicated BUSCO Metazoa genes increased in the bdelloid A. vaga, whereas duplicated BUSCO genes were rare in the drafts for acanthocephalan, monogonont and seisonid genomes (Table 4). Lastly, genome size as reported for the acanthocephalan P. laevis is likely to result from multiplication of a wide variety of repeat types  in combination with a decrease in gene compactness (present study). But even if genome size increases reflect the spread of transposable elements, SINEs should have been of minor importance in the different lineages of Rotifera-Acanthocephala. In fact, SINEs appear generally to be little represented in rotiferan and acanthocephalan draft genomes available so far [21,22,23, 55].
Increasing gene loss with growing ties to the host
We cannot exclude that individual genes were missing in the draft genomes, although they are present in a given species. On the other hand, the possibility that missing BUSCO Metazoa genes are partially located in the unassembled regions should not be limited to S. nebaliae and P. laevis. But if incompleteness of the assemblies played a role at all, such an effect should not be substantial. Thus, the total size of the P. laevis haploid genome draft of approximately 260 Mb was very close to the GenomeScope prediction of 265–281 Mb . Similarly, GenomeScope estimated the size of the nuclear genome of S. nebaliae to be approximately 46 Mb, of which 44 Mb could be delivered in assembly GA2. Consistent with this, Merqury for GA2 calculated a k-mer completeness of approximately 92% (Table 2). However, a maximum of 8% of the genomes awaiting reconstruction in each of the two species is unlikely to explain the failure to determine substantial proportions of BUSCO Metazoa genes as reported in current Table 4. It remains possible that genes have not been detected because the genomes are more derived, especially in S. nebaliae and P. laevis. In agreement with this, we determined BUSCO Eukaryota genes, which are usually more conserved than BUSCO Metazoa genes, to larger proportions in the draft genomes of S. nebaliae and P. laevis. Nevertheless, the overall picture of more missing genes in the P. laevis than S. nebaliae draft genomes remained unchanged in BUSCO Eukaryota genes. Ultimately, the same picture seems to emerge in presence-absence analyses of ANTP-type gene families (Fig. 11). Still, it should be emphasized that we do not claim that the numbers of missing genes as reported by us are defintive. However, we consider valid the basic pattern that, under the same settings, the repertoires of discovered genes decrease from free-living rotifers via epizoic seisonids to endoparasitic acanthocephalans. This may be due to the increasing outsourcing of functions with growing ties to a host.
Outsourcing of functions and associated gene loss is well known for parasites (e.g., ). However, the close correspondence with the probable phylogenetic relationships within Rotifera-Acanthocephala is, in our view, striking. Thus, seisonids use leptostracan crustaceans as a substrate, probably also benefiting from their food as commensals. They may even feed on hemolymph depending on the seisonid species [1,2,3, 26]. Yet, living on a host should require less functional autonomy than living without host association such as in the monogonont B. plicatilis and the bdelloid A. vaga (compare ). An epizoic lifestyle as in extant seisonids is also probable for the LCA of Pararotatoria [18, 24].
Despite the preliminary nature of the results, it seems worthwhile mentioning that the ANTP-type gene families, for which no member was determined in the draft genomes for S. nebaliae and P. laevis (hox5, mnx/hb9, gsx and nk4), steer nervous system development in other taxa [69,70,71,72]. This does not have to be their only function, as the hox gene cluster has primarily been implicated in anterioposterior patterning [73, 74]. Regardless, the absence of all or part of the four gene families could indicate a simplified development of the nervous system in the pararotatorian LCA compared to free-living monogononts and bdelloids. Extending this argument, the low number of ANTP-type and other genes in P. laevis (Fig. 11) could indicate further reduced functional autonomy in endoparasitic acanthocephalans. In fact, thorny-headed worms are commonly considered to be simplified metazoans . Acanthocephalans such as P. laevis, for example, lack excretory organs and none of the extant species has a digestive tract (e.g., [9, 76]). At least it is not contradictory that two genes, hox1 and hox2, which are thought to play a role in the development of the stomatogastric nervous system [73, 74], could not be determined in the draft genome of P. laevis .
Present analyses suggest co-existence of two S. nebaliae strains on Nebalia bipes off the French Channel coast. Their mitogenomes (Fig. 1) differed considerably in terms of (NCR) length and sequence identity (ca. 82%). As shown by phylogenetic analyses, both mitochondrial genomes represented S. nebaliae. Thus, none of the mitochondrial sequences was derived from the crustacean host or from P. annulatus, the other seisonid species living on opossum shrimps (Fig. 3). Additional validation steps corroborated transcription and hence functionality for both S. nebaliae mitogenomes. Hence, none of these should represent a nuclear mitochondrial integration (Fig. 2). Therefore, we consider it appropriate to refer to the reconstructed mitochondrial genomes as to haplotypes (A and B) within S. nebaliae. We also found evidence for two nuclear genomes (A and B) that differed in gene compactness, GC content and codon usage (Figs. 7, 8, 9 and 10). Since the mitochondiral and nuclear genomes exhibited equidirectional abundance ratios, it is reasonable to assume that genetically distinct strains (A and B) exist in S. nebaliae. In both strains, haploid nuclear genome size should be ca. 46 Mb, of which ca. 44 Mb were contained in assembly GA2. Thus, the seisonid nuclear genome could be the smallest within Rotifera-Acanthocephala. Its size might reflect a reduction, provided haploid genome size was about 50 Mb in the LCA of Rotifera-Acanthocephala. Under the same premise, larger genomes should have evolved independently within Monogononta and in the stem lines of Bdelloidea and Acanthocephala [20, 22, 23]. Additional genome and transcriptome metrics for S. nebaliae were within the ranges in other members of Rotifera-Acanthocephala. In particular, the number of BUSCO Metazoa genes determined in the draft genome of S. nebaliae was intermediate between corresponding counts for free-living monogonont and bdelloid wheel animals and an endoparasitic acanthocephalan (Fig. 11; Table 4).
According to phylogenomic analyses of 100 proteins encoded by concatenated BUSCO Metazoa genes, a basal branching of Seisonidea within the Rotifera-Acanthocephala clade is primarily due to LBA to the outgroup or root. Indeed, measures avoiding such an effect led to a clustering of S. nebaliae with the acanthocephalan representative. In particular, rooting the tree via an ancestral sequence resulted in maximum support for monophyletic Pararotatoria (Seisonidea+Acanthocephala) within monophyletic Hemirotifera (Bdelloidea+Pararotatoria) (Fig. 12). Missing evidence can always be due to non-finding. Nevertheless, it is worthwhile noting that the exclusive non-determination of members of four ANTP-type gene families in the draft genomes of S. nebaliae and the acanthocephalan P. laevis at least does not contradict the Pararotatoria hypothesis. In contrast, none of the ANTP-type gene families searched for lacked members in draft genomes of the bdelloid A. vaga and P. laevis only (Fig. 11). Thus, Lemniscea (Bdelloidea+Acanthocephala)  gained no support from presence-absence analysis of ANTP-type gene families.
Mapping lifestyles on the tree in Fig. 12D, a parsimonious scenario would be that seisonids have retained an epizoic lifestyle on jawed arthropods (Mandibulata) from their last common ancestor with acanthocephalans. Early acanthocephalans would then have entered the mandibulate host, followed by upward inclusion of jawed vertebrates (Gnathostomata) into the lifecycle (e.g. [9, 24]). When combined with results of BUSCO analyses, the corresponding tree suggests gradually increasing gene loss from the LCAs of Rotifera-Acanthocephala and Hemirotifera, both of which should have been free-living, to the probably epizoic LCA of Pararotatoria to the LCA of Acanthocephala, for which an endoparasitic lifestyle is almost certain (for presumed lifestyles in LCAs, see, e.g., [8, 9]). Ultimately, however, the finding that growing ties to the host associate with increasing gene loss inside Rotifera-Acanthocephala should hold true regardless of the phylogenetic hypothesis.
In addition to the above phylogenetic and evolutionary implications, the S. nebalia draft genome might proof useful for the development of a specific drug for the control of acanthocephalans, which turned out to be a pest in fish aquaculture (e.g., [77, 78]). Thus, it might be possible now to identify acanthocephalan genes or proteins as potential targets for a highly specific control of the parasites. Follow-up studies might further uncover subtle differences in population dynamics and/or morphology between the S. nebaliae strains determined herein (compare ). Future analyses might additionally clarify if S. nebaliae is a complex of cryptic species. Finally, the path shown here of rooting a tree via a reconstructed ancestral sequence instead of the sequence of an extant outgroup representative may generally prove useful in phylogenetic analyses of deeply splitting taxa.
Permit for collecting and processing opossum shrimps (Crustacea: Leptostraca: N. bipes) and their epifauna was given by European Marine Biological Resource Centre (EMBRC). Collections of opossum shrimps were carried out in spring 2016 and late summer 2019 at low tide from rock pools in the tidal flats off Roscoff (Brittany, France). The crustaceans were kept in a Dewar vessel and brought to the laboratory at the Station Biologique Roscoff, an institution of the EMBRC. Under the stereomicroscope, specimens of S. nebaliae and P. annulatus (Rotifera: Seisonidea, each) were separated from the shrimps and stored at − 80 °C. Isolation of nucleic acids followed in Mainz upon transportation of the samples on dry-ice.
DNA sequencing and preliminary assembling
A pool of 594 S. nebaliae specimens (males and females) was digested using proteinase K, followed by DNA extraction with phenol-chloroform-isoamyl alcohol and precipitation in ethanol. The library prepared from this DNA was sequenced on four lanes on an Illumina NextSeq 500 platform (150 bp, paired-end mode, 74,690,183 pairs total). Quality of raw NGS data was checked with FastQC v. 0.11.9 (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/), followed by trimming of Illumina adaptor sequences and quality trimming (HEADCROP:6 SLIDINGWINDOW:4:15 LEADING:3 TRAILING:3 MINLEN:20) with Trimmomatic v. 0.39 . Processed reads were mapped against a masked human reference genome (HG19) with BBmap (https://sourceforge.net/projects/bbmap/), upon which sequences of potential human (Homo sapiens) origin were removed. In addition, DNA was isolated from a pool of 10 individuals of P. annulatus applying the same protocol. Following whole genome amplification (illustra GenomiPhi™ V2 DNA Amplification Kit) DNA was sequenced on one lane on an Illumina NextSeq 500 platform (150 bp, paired-end mode, 87,887,226 pairs total). Data quality assessment and trimming used the same programs as detailed above. Raw reads of both sequencing runs can be retrieved from EMBL Nucleotide Sequence Database (ENA) under the accession number PRJEB43415.
Mitochondrial genome assembly and analysis
We derived mitochondrial genome sequences of S. nebaliae from trimmed and filtered Illumina reads. First, we ran MEGAHIT v. 1.2.9  with standard settings and identified putative mitochondrial sequences by BLASTN searches (e-value cutoff ≤1e-05 ) against NCBI’s nt database (12-04-2020). The resulting assembly contained two mitogenomes, one of which prooved to be almost identical with the mitochondrial genome previously published for S. nebaliae (KP742964.1). We denoted this variant as haplotype A and kept it as is, as it was already validated in the reference study . Yet, the novel mitochondrial sequence from the MEGAHIT assembly was used as a seed for subsequent reconstructions with MITObim v. 1.9.1  (settings: --mismatch 0, −quick Megahit-seed) and NOVOPlasty v. 4.2  (settings: genome range = 14,500–17,000, K-mer = 61, Use Quality Scores = yes, Reference sequence = SeisonMT-Haplotype). NOVOPlasty was able to reconstruct a circular sequence despite two repetitive areas in the non-coding region (NCR). Finally, all three assemblies were aligned with MAFFT v. 7 (https://mafft.cbrc.jp/alignment/server/; ) for manual derivation of a consensus sequence, which was verified as being a Seison mitogenome by BLASTN against NCBI’s nt database. We refer to this novel mitogenome as to haplotype B. Downstream gene prediction by MITOS  was followed by validation of protein-coding genes with NCBI ORF finder (https://www.ncbi.nlm.nih.gov/orffinder/) and subsequent BLASTP against NCBI’s non-redundant protein database. Gene boundaries were expanded with preference given to non-overlapping genes. Annotation of tRNA genes was verified with ARWEN v. 1.2  and tRNAScan-SE v. 2.0 .
For inference of multiplicity values, we mapped the processed DNA and RNA reads (S. nebaliae, see RNA sequencing (RNA-seq) and de novo assembly of transcriptomes) with Bowtie 2 (http://usegalaxy.eu [88, 89]) onto both S. nebaliae mitogenomes. Sequence identity between the two haplotypes was derived by BioEdit v. 7.2.5 , after aligning with MAFFT v. 7 and revising with Gblocks v. 0.91b  in default settings.
A seisonid origin of both mitochondrial variants was verified by phylogenetic analyses of alignments including sequences of potential contaminants such as crustaceans or co-occuring P. annulatus. Limiting factor in alignment generation was the application of the same assembly pipeline as described above to trimmed P. annulatus reads, which resulted in a fragmented mitogenome assembly covering genes for cytochrome oxidase subunit I (cox1) and NADH dehydrogenase subunits 1 (nd1) only (see Additional file 1 for more details). In addition to the sequences of P. annulatus and S. nebaliae haplotypes A and B, the cox1 alignment contained an additional S. nebaliae sequence (DQ297765.1), which had been reconstructed by another working group . Due to a lack of respective sequences for N. bipes in GenBank, altogether three other crustacean species were taken as substitutes: N. pseudotroncosoi (Leptostraca; cox1: JX442539.1), Pacifastacus leniusculus (Astacoidea; nd1: NC_033509.1), P. (Litopenaeus) vannamei (Penaeoidea; cox1 and nd1: EF584003.1). We additionally included orthologues of the nematode C. elegans (cox1 and nd1: NC_001328.1).
Sequences were aligned with MAFFT v. 7 and pruned from uncertain alignment sections with Gblocks v. 0.91b . The latter was run with default settings, except for a lowered minimum block length (5 aa). For ML tree construction with PhyML v. 3.3 , the best-fit substitution model (GTR + G) was determined with Smart Model Selection (SMS) v. 1.8.4 , applying the Akaike Information Criterion (AIC). Branch support values in ML trees give results from approximate SH-like aLRT. For BI with MrBayes v. 3.2.7a [94, 95], trees were reconstructed from posterior distributions containing every 100th generation (the first 25% were discarded as burn-in) as drawn from two runs with four chains of 1 Mio generations, each. Trees were visualized iTOL v. 5 .
De novo assembly of the nuclear genome
Mitochondrial reads were excluded from reconstruction of the nuclear genome with the k-mer filtering function of BBDuk from the BBTools suite v. 38.73 (https://sourceforge.net/projects/bbmap/). Subsequent profiling with GenomeScope 1.0  used Jellyfish v. 2.2.8  k-mer counts (k = 21). For complementary Merqury analysis, we first determined the best k-mer size for both nuclear genome assemblies of S. nebalia (GA1: 99 Mb; GA2: 44 Mb) using the script best_k.sh . Subsequently, we created a meryl database for the best k-mer size (k = 18) from trimmed and filtered Illumina gDNA reads. With this database, we evaluated the k-mer frequency distribution in our two nuclear genome assemblies reconstructed with MEGAHIT .
In a first approach (GA1), we ran MEGAHIT with default settings, apart from a stricter error control (prune-level 3). We expected GA1 to contain two nuclear genomes (A and B) of carriers of mitochondrial haplotypes A and B. We aimed to reconstruct a second assembly (GA2) that represented a single nuclear genome (B) of carriers of mitochondrial haplotype B only. To achieve this, we implemented stricter settings in error control (prune-level 3, −-no-mercy, −-disconnect-ratio 0.2, −-low-local-ratio 0.4), higher minimum k-mer size (k-min 29) and a higher minimum k-mer count and average depth (−-prune-depth 20, −-min-count 4). The implementation of a multiplicity cut-off (≥ 20 x) to GA2 implicated a filtering against less abundant k-mers. Furthermore, we scrutinized GA2 contigs for BLASTN hits with E-values ≤1e-03 and IDs ≥ 80%, and deleted manually validated hits to N. bipes (PRJNA67309), Genostoma (Platyhelminthes) and bacteria. In addition, we filtered both nuclear genome assemblies, GA1 and GA2, for a minimal contig length of 1000 bp. We used Bowtie 2 to map the filtered DNA reads to both assemblies (GA1, GA2) and derived multiplicity values for each contig. Finally, we used BUSCO v. 4.0.6  to assess the level of completeness of genome assemblies GA1 and GA2. For doing so, the BUSCO pipeline was run with the metazoan database (odb10) and standard settings, except for the application of Schistosoma mansoni (Platyhelminthes, Trematoda) gene models. One hundred BUSCO Metazoa genes that had been classified as duplicated in GA1 were retained for further analyses, provided that the abundances of the corresponding contigs differed by 8–14-fold and orthologues were available for the other species included in downstream phylogenetic analyses. In case of the genome assemblies for S. nebaliae (GA2) and P. laevis sensu lato , we additionally searched for Metazoa and Eukaryota genes with the self-training function in Augustus being activated.
Annotation of the repetitive DNA complement
For de novo generation of a custom database of S. nebaliae repeats, we ran RepeatModeler v. 2.0.1  on the unfiltered draft genome GA2 and dnaPipeTE  and RepARK v. 1.3.0  on all trimmed and filtered Illumina DNA reads. As recommended , dnaPipeTE was started with alternative read depths to find best assembly conditions. Finally, we carried out 50 runs, with a multiplicity of 0.001 (for further details on de novo repeat generation see Additional file 1). The results of all these runs were collected into a single database. Since neither RepARK nor dnaPipeTE have an integrated annotation function, repeats were annotated with TE class . To avoid the masking of duplicated protein-coding genes, which were not derived from transposable elements (TEs), corresponding candidates were removed from the repeat database. For this purpose, we excluded contigs with BLASTX hits (E-value: 1e-05) to the Swiss-Prot database (release 2020_03), as long as the corresponding Swiss-Prot sequences lacked significant matches in RepBaseRepeatMaskerEdition-20,181,026 (TBLASTN; E-value: 1e-05). The repeats from all three annotations were merged into a single file. Finally, we added repeats from the RepBaseRepeatMaskerEdition-20,181,026 classified as root, Metazoa, Protostomia or Rotifera to the custom repeat database. We annotated the repeats in draft genome GA2 using RepeatMasker v. open-4.0.7, applying the more sensitive “slow search”.
RNA sequencing (RNA-seq) and de novo assembly of transcriptomes
RNA was extracted from 120 S. nebaliae specimens with the TRI Reagent™ (Invitrogen™) applying the manufacturers protocol. Pelleted RNA was resolved in HPLC-grade H2O. Following directional library preparation, NGS used one lane of an Illumina NextSeq 500 platform (150 bp, paired-end mode, 22,878,639 pairs total). Raw reads have been deposited under ENA accession number PRJEB43415. After having verified quality of raw data with FastQC, reads were processed with Trimmomatic (see above, for versions and references; Illumina adaptor sequence trimming, LEADING:3 TRAILING:3 SLIDINGWINDOW:4:15 AVGQUAL:20 MINLEN:40). For subsequent transcriptome reconstruction, we solely used mRNA reads, which STAR v. 2.7.6a  had mapped to GA1 and GA2. Genome-guided transcriptome reconstruction with Trinity then ran with the aid of STAR bam files. Reconstructions additionally used the jaccard_clip flag to avoid artificial fusion of transcripts. Using these data, Trinity v. 2.11.0  assembled two transcriptomes, whereby the first one should represent S. nebaliae strains A and B (TA1) and the second one strain B only (TA2). Running a script implemented in Trinity, we derived SuperTranscripts , which should combine all isoforms of individual genes into single sequences. BSUCO v. 4.0.6  was used in transcript mode with the metazoan database (odb10) and S. mansoni (Platyhelminthes, Trematoda) gene models, to assess the completeness of both SuperTranscript sets. To achieve an approximate map of the coding sequences, TA2 transcripts were mapped to the draft genome GA2 with GMAP v. 2021-05-27 .
Comparative analysis of conserved metazoan genes
Comparisons between the two S. nebaliae strains focused on GC content and nucleotide usage at third codon position as inferred by MEGA X  from 100 codon-based alignments of BUSCO metazoan genes generated with pal2nal v. 14.0  and the contigs they were residing on. Pairwise comparisons of nucleotide pair frequencies and codon usage were then conducted with the MWU test implemented in SPSS v. 23 (IBM). The genes considered had to reside on contigs with abundancies differing by a factor of 8–14, which roughly corresponded to the range observed in our mitochondrial and genome assemblies. As a second criterion, the genes selected had to have orthologues in additional species included in downstream comparisons: S. mediterranea (Platyhelminthes; GCA_002600895.1), B. plicatilis (Monogononta; GCA_010279815.1), A. vaga (Bdelloidea; GCA_000513175.1) and P. laevis (Acanthocephala; GCA_012934845.1). If a gene was annotated several times as in the degenerate tetraploid A. vaga , we accepted the copy with maximum BUSCO score. Based on the BUSCO annotations for the metazoan genes kept we compared protein and gene lengths, applying the KW test in SPSS. Alpha-error rates (p values) from MWU and KW tests were transformed into FDRs .
For phylogenetic analyses, amino acid sequences were aligned separately for each of the aforementioned 100 BUSCO metazoan genes (MAFFT v. 7), followed by curation with Gblocks 0.91b (minimum block length 5). The resulting alignments were concatenated to a single dataset of 21,042 aa with six operational taxonomic units (OTUs) (Concat100-6OTUs). Applying an in-house script , we built three additional datasets. In one of these datasets, positions in which at least one sequence had a private character (referred to as singletons) and the outgroup representative were removed. The resulting alignment contained five OTUs and had 13,268 aa (Concat100-noSing-5OTUs). We generated further alignments which lacked the outgroup representative and one of the S. nebaliae strains in addition to singletons. The dataset containing strain A had 9550 aa positions (Concat100-noSing-4OTUs-A), while the one with strain B extended over 9579 aa (Concat100-noSing-4OTUs-B). Another alignment contained an ancestral sequence which we had reconstructed for the Rotifera-Acanthocephala LCA whereas the platyhelminth was deleted (Concat100-6OTUs-LCA). Reconstruction of the LCA sequence used the ML framework of CodeML implemented in PAML package v. 4.9j . CodeML was run with lg.dat matrix and REVaa (189) + G + I model and the following intree: ((P. laevis, (S. nebaliae strain A, S. nebaliae strain B), A. vaga, B. plicatilis), S. mediterranea). Note that the intree was star-like for Rotifera-Acanthocephala. By doing so, reconstruction of the LCA sequence was unbiased in respect to the actual phylogenetic relationships within the clade. Based on AIC, PhyML determined LG + G + I + F as the best-fit model (SMS) for all five alignments. Tree reconstruction relied on PhyML v. 3.3 and MrBayes v. 3.2.7a once more. From two runs of four chains each was collected every 500th of a total of 100,000 generations.
We additionally compared complements of ANTP-type gene families between the monogonont B. plicatilis (GCA_010279815.1), A. vaga (GCA_000513175.1), S. nebaliae (present study), and P. laevis (GCA_012934845.1). For this purpose, we screened (TBLASTN) the (latest) genome assemblies specified in the previous sentence for aa sequences of homeodomains of various ANTP class gene homologues in species of Lophotrochozoa, Ecdysozoa and Vertebrata. Scaffolds with hits yielding incomplete homeodomains due to splice sites were analyzed with GENSCAN (Hollywood.mit.edu/GENSCAN.html) to recover complete homeodomains. First orthology assignments were made based on according BLASTX hits to NCBI’s GenBank. Results were verified by generating an alignment of the 60 aa homeodomains including sequences of homologous genes of several other species and subsequent phylogenetic analysis by FastML v. 3.1  and FastTreeMP v. 2.1.10  (data available upon request). Trees were visualized by FigTree v. 1.4.4 (https://github.com/rambaut/figtree/releases).
Availability of data and materials
The datasets analyzed in the current study are available in the EMBL Nucleotide Sequence Database (ENA) repository under the accession number PRJEB43415 [https://www.ebi.ac.uk/ena/browser/view/ERA3506149].
Grube AE. Ein Ausflug nach Triest und dem Quarnero, Beiträge zur Kenntnis der Thierwelt dieses Gebietes. Berlin: Nicolai'sche Verlagshandlung; 1861.
Claus C. Über die Organisation und systematische Stellung der Gattung Seison Gr. Festschrift zur Feier des Fünfundzwanzigjährigen Bestehens der K.K. Zoologisch-Botanischen Gesellschaft in Wien. Vienna: K.K. Zoologisch-Botanische Gesellschaft; 1876. p. 77–88.
de Beauchamp PM. Recherches sur les rotifères: les formations tégumentaires et l'appareil digestif. Arch Zool exp, Paris, ser 4. 1909;10:1–410.
Ahlrichs WH, Riemann O. Seisonidae. In: Schmidt-Rhaesa A, editor. Miscellaneous invertebrates. Berlin: De Gruyter; 2018. p. 55–86.
Fontaneto D, de Smet W. Rotifera. In: Schmidt-Rhaesa A, editor. Handbook of zoology, Gastrotricha and Gnathifera. Berlin: De Gruyter; 2015. p. 216–300.
Ricci C, Melone G, Sotgia C. Old and new data on Seisonidea (Rotifera). Hydrobiologia. 1993;255–256:495–511.
Wallace RL. Rotifers: exquisite Metazoans. Integr Comp Biol. 2002;42(3):660–7. https://doi.org/10.1093/icb/42.3.660.
Near TJ. Acanthocephalan phylogeny and the evolution of parasitism. Integr Comp Biol. 2002;41:668–77.
Herlyn H. Thorny-headed worms (Acanthocephala): jaw-less members of jaw-bearing worms that parasitize jawed arthropods and jawed vertebrates. In: de Baets K, Huntley J, editors. The Evolution and Fossil Record of Parasites: Springer; 2021. p. 273–313.
von Haffner K. Organisation und systematische Stellung der Acanthocephalen. Zool Anz. 1950;145:243–74.
Ahlrichs WH. Epidermal ultrastructure of Seison nebaliae and Seison annulatus, and a comparison of epidermal structures within the Gnathifera. Zoomorphology. 1997;117(1):41–8. https://doi.org/10.1007/s004350050028.
Melone G, Ricci C, Segers H, Wallace RL. Phylogenetic relationships of phylum Rotifera with emphasis on the families of Bdelloidea. Hydrobiologia. 1998;387/388:101–7.
Ferraguti M, Melone G. Spermiogenesis in Seison nebaliae (Rotifera, Seisonidea): further evidence of a rotifer-acanthocephalan relationship. Tissue Cell. 1999;31(4):428–40. https://doi.org/10.1054/tice.1999.0012.
García-Varela M, Nadler SA. Phylogenetic relationships among Syndermata inferred from nuclear and mitochondrial gene sequences. Mol Phylogenet Evol. 2006;40(1):61–72. https://doi.org/10.1016/j.ympev.2006.02.010.
Laumer CE, Fernández R, Lemer S, Combosch D, Kocot KM, Riesgo A, et al. Revisiting metazoan phylogeny with genomic sampling of all phyla. Proc R Soc B Biol Sci. 2019;286(1906):20190831. https://doi.org/10.1098/rspb.2019.0831.
Mark Welch DB. Evidence from a protein-coding gene that acanthocephalans are rotifers. Invertebr Biol. 2000;119:17–26.
Rieger RM, Tyler S. Sister-group relationship of Gnathostomulida and Rotifera-Acanthocephala. Invertebr Biol. 1995;114(2):186–8. https://doi.org/10.2307/3226891.
Herlyn H, Piskurek O, Schmitz U, Ehlers U, Zischler H. The syndermatan phylogeny and the evolution of acanthocephalan endoparasitism as inferred from 18S rDNA sequences. Mol Phylogenet Evol. 2003;26(1):155–64. https://doi.org/10.1016/S1055-7903(02)00309-3.
Wey-Fabrizius AR, Herlyn H, Rieger B, Rosenkranz D, Witek A, Welch DBM, et al. Transcriptome data reveal syndermatan relationships and suggest the evolution of endoparasitism in Acanthocephala via an epizoic stage. PLoS One. 2014;9(2):e88618. https://doi.org/10.1371/journal.pone.0088618.
Flot J, Hespeels B, Li X, Noel B, Arkhipova I, Danchin EGJ, et al. Genomic evidence for ameiotic evolution in the bdelloid rotifer Adineta vaga. Nature. 2013;500(7463):453–7. https://doi.org/10.1038/nature12326.
Nowell RW, Almeida P, Wilson CG, Smith TP, Fontaneto D, Crisp A, et al. Comparative genomics of bdelloid rotifers: insights from desiccating and nondesiccating species. PLoS Biol. 2018;16(4):e2004830. https://doi.org/10.1371/journal.pbio.2004830.
Blommaert J, Riss S, Hecox-Lea B, Mark Welch DB, Stelzer CP. Small, but surprisingly repetitive genomes: transposon expansion and not polyploidy has driven a doubling in genome size in a metazoan species complex. BMC Genomics. 2019;20(1):466. https://doi.org/10.1186/s12864-019-5859-y.
Mauer K, Hellmann SL, Groth M, Fröbius AC, Zischler H, Hankeln T, et al. The genome, transcriptome, and proteome of the fish parasite Pomphorhynchus laevis (Acanthocephala). PLoS One. 2020;15(6):e0232973. https://doi.org/10.1371/journal.pone.0232973.
Sielaff M, Schmidt H, Struck TH, Rosenkranz D, Mark Welch DB, Hankeln T, et al. Phylogeny of Syndermata (syn. Rotifera): mitochondrial gene order verifies epizoic Seisonidea as sister to endoparasitic Acanthocephala within monophyletic Hemirotifera. Mol Phylogenet Evol. 2016;96:79–92. https://doi.org/10.1016/j.ympev.2015.11.017.
Sørensen MV, Segers H, Funch P. On a new Seison Grube, 1861 from coastal waters of Kenya, with a reappraisal of the classification of the Seisonida (Rotifera). Zool Stud. 2005;44:34–43.
Leasi F, Rouse GW, Sørensen MV. A new species of Paraseison (Rotifera: Seisonacea) from the coast of California, USA. J Mar Biol Assoc United Kingdom. 2012;92(5):959–65. https://doi.org/10.1017/S0025315411000129.
Segers H. Annotated checklist of the rotifers (phylum Rotifera), with notes on nomenclature, taxonomy and distribution. Zootaxa. 2007;1564(1):1–104. https://doi.org/10.11646/zootaxa.1564.1.1.
Gibson DI, Bray RA, Hunt D, Georgiev BB, Scholz T, Harris PD, et al. Fauna europaea: helminths (animal parasitic). Biodivers Data J. 2014;2:e1060. https://doi.org/10.3897/BDJ.2.e1060.
Song MO, Min G-S. A new species and ten new records of bdelloid rotifers from Korea. Zootaxa. 2015;3964(2):211–27. https://doi.org/10.11646/zootaxa.3964.2.3.
Amin OM, Heckmann RA, Fiser Z, Zaksek V, Herlyn H, Kostanjsek R. Description of Acanthocephalus anguillae balkanicus subsp. n. (Acanthocephala: Echinorhynchidae) from Proteus anguinus Laurenti (Amphibia: Proteidae) and the cave ecomorph of Asellus aquaticus (Crustacea: Asellidae) in Slovenia. Folia Parasitol. 2019;66:1–13.
Reier S, Sattmann H, Schwaha T, Harl J, Konecny R, Haring E. An integrative taxonomic approach to reveal the status of the genus Pomphorhynchus Monticelli, 1905 (Acanthocephala: Pomphorhynchidae) in Austria. Int J Parasitol Parasites Wildl. 2019;8:145–55. https://doi.org/10.1016/j.ijppaw.2019.01.009.
Gómez A, Serra M, Carvalho GR, Lunt DH. Speciation in ancient cryptic species complexes: evidence from the molecular phylogeny of Brachionus plicatilis (Rotifera). Evolution. 2002;56(7):1431–44. https://doi.org/10.1111/j.0014-3820.2002.tb01455.x.
Fontaneto D, Iakovenko N, Eyres I, Kaya M, Wyman M, Barraclough TG. Cryptic diversity in the genus Adineta Hudson & Gosse, 1886 (Rotifera: Bdelloidea: Adinetidae): a DNA taxonomy approach. Hydrobiologia. 2011;662(1):27–33. https://doi.org/10.1007/s10750-010-0481-7.
Pinacho-Pinacho CD, García-Varela M, Sereno-Uribe AL, Pérez-Ponce de León G. A hyper-diverse genus of acanthocephalans revealed by tree-based and non-tree-based species delimitation methods: ten cryptic species of Neoechinorhynchus in Middle American freshwater fishes. Mol Phylogenet Evol. 2018;127:30–45. https://doi.org/10.1016/j.ympev.2018.05.023.
Plate L. Über einige ectoparasitische Rotatorien des Golfes von Neapel. Mitt Zool Stat Neapel. 1887;7:234–62.
Herlyn H, Ehlers U. Ultrastructure and function of the pharynx of Gnathostomula paradoxa (Gnathostomulida). Zoomorphology. 1997;117(3):135–45. https://doi.org/10.1007/s004350050038.
Sørensen MV, Giribet G. A modern approach to rotiferan phylogeny: combining morphological and molecular data. Mol Phylogenet Evol. 2006;40(2):585–608. https://doi.org/10.1016/j.ympev.2006.04.001.
Struck TH, Wey-Fabrizius AR, Golombek A, Hering L, Weigert A, Bleidorn C, et al. Platyzoan paraphyly based on phylogenomic data supports a noncoelomate ancestry of Spiralia. Mol Biol Evol. 2014;31(7):1833–49. https://doi.org/10.1093/molbev/msu143.
Lorenzen S. Phylogenetic aspects of pseudocoelomate evolution. In: Conway-Morris S, George JD, Gibson R, Platt HM, editors. The origins and relationships of lower invertebrates. Oxford: Clarendon Press; 1985. p. 210–23.
Wey-Fabrizius AR, Podsiadlowski L, Herlyn H, Hankeln T. Platyzoan mitochondrial genomes. Mol Phylogenet Evol. 2013;69(2):365–75. https://doi.org/10.1016/j.ympev.2012.12.015.
Gazi M, Kim J, García-Varela M, Park C, Littlewood DTJ, Park J-K. Mitogenomic phylogeny of Acanthocephala reveals novel class relationships. Zool Scr. 2016;45(4):437–54. https://doi.org/10.1111/zsc.12160.
Vurture GW, Sedlazeck FJ, Nattestad M, Underwood CJ, Fang H, Gurtowski J, et al. GenomeScope: fast reference-free genome profiling from short reads. Bioinformatics. 2017;33(14):2202–4. https://doi.org/10.1093/bioinformatics/btx153.
Kajitani R, Toshimoto K, Noguchi H, Toyoda A, Ogura Y, Okuno M, et al. Efficient de novo assembly of highly heterozygous genomes from whole-genome shotgun short reads. Genome Res. 2014;24(8):1384–95. https://doi.org/10.1101/gr.170720.113.
Rhie A, Walenz BP, Koren S, Phillippy AM. Merqury: reference-free quality, completeness, and phasing assessment for genome assemblies. Genome Biol. 2020;21:1–27.
Grabherr MG, Haas BJ, Yassour M, Levin JZ, Thompson DA, Amit I, et al. Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nat Biotechnol. 2011;29(7):644–52. https://doi.org/10.1038/nbt.1883.
Wu TD, Watanabe CK. GMAP : a genomic mapping and alignment program for mRNA and EST sequences. Bioinformatics. 2005;21(9):1859–75. https://doi.org/10.1093/bioinformatics/bti310.
Choi BS, Lee YH, Lee JS, Yamade T, Hagiwara A, Lee JS. Complete mitochondrial genome of the marine monogonont rotifer Proales similis (Rotifera, Proalidae). Mitochondrial DNA B Resour. 2020;5(2):1151–2. https://doi.org/10.1080/23802359.2020.1730265.
Muhammad N, Khan MS, Li L, Zhao Q, Ullah H, Zhu XQ, et al. Characterization of the complete mitogenome of Centrorhynchus clitorideus (Meyer, 1931) (Palaeacanthocephala: Centrorhynchidae), the largest mitochondrial genome in Acanthocephala, and its phylogenetic implications. Mol Biochem Parasitol. 2020;237:111274. https://doi.org/10.1016/j.molbiopara.2020.111274.
Boore JL, Brown WM. Big trees from little genomes: mitochondrial gene order as a phylogenetic tool. Curr Opin Genet Dev. 1998;8(6):668–74. https://doi.org/10.1016/S0959-437X(98)80035-X.
Ladoukakis ED, Saavedra C, Magoulas A, Zouros E. Mitochondrial DNA variation in a species with two mitochondrial genomes: the case of Mytilus galloprovincialis from the Atlantic, the Mediterranean and the Black Sea. Mol Ecol. 2002;11(4):755–69. https://doi.org/10.1046/j.1365-294X.2002.01473.x.
Söller R, Warnke K, Saint-Paul U, Blohm D. Sequence divergence of mitochondrial DNA indicates cryptic biodiversity in Octopus vulgaris and supports the taxonomic distinctiveness of Octopus mimus (Cephalopoda: Octopodidae). Mar Biol. 2000;136:29–35.
Thielsch A, Knell A, Mohammadyari A, Petrusek A, Schwenk K. Divergent clades or cryptic species? Mitonuclear discordance in a Daphnia species complex. BMC Evol Biol. 2017;17:1–9.
Vyskočilová S, Tay WT, van Brunschot S, Seal S, Colvin J. An integrative approach to discovering cryptic species within the Bemisia tabaci whitefly species complex. Sci Rep. 2018;8:1–13.
Jiménez-Santos MA, Sarma SSS, Nandini S. Temperature-dependent demographic differences in sessile rotifers of the genus Limnias (Rotifera: Gnesiotrocha). J Environ Biol. 2019;40(4):711–8. https://doi.org/10.22438/jeb/40/4/MRN-1046.
Kim H-S, Lee B-Y, Han J, Jeong C-B, Hwang D-S, Lee M-C, et al. The genome of the freshwater monogonont rotifer Brachionus calyciflorus. Mol Ecol Resour. 2018;18(3):646–55. https://doi.org/10.1111/1755-0998.12768.
Franch-Gras L, Hahn C. García- Roger EM, Carmona MJ, Serra M, Gómez a. genomic signatures of local adaptation to the degree of environmental predictability in rotifers. Sci Rep. 2018;8:1–14.
Han J, Park JC, Choi B-S, Kim M-S, Kim H-S, Hagiwara A, et al. The genome of the marine monogonont rotifer Brachionus plicatilis: genome-wide expression profiles of 28 cytochrome P450 genes in response to chlorpyrifos and 2-ethyl-phenanthrene. Aquat Toxicol. 2019;214:105230. https://doi.org/10.1016/j.aquatox.2019.105230.
Park JC, Choi B-S, Kim M-S, Shi H, Zhou B, Park HG, et al. The genome of the marine rotifer Brachionus koreanus sheds light on the antioxidative defense system in response to 2-ethyl-phenanthrene and piperonyl butoxide. Aquat Toxicol. 2020;221:105443. https://doi.org/10.1016/j.aquatox.2020.105443.
Laumer CE, Bekkouche N, Kerbl A, Goetz F, Neves RC, Sørensen MV, et al. Spiralian phylogeny informs the evolution of microscopic lineages. Curr Biol. 2015;25(15):2000–6. https://doi.org/10.1016/j.cub.2015.06.068.
Philippe H, Poustka AJ, Chiodin M, Hoff KJ, Dessimoz C, Tomiczek B, et al. Mitigating anticipated effects of systematic errors supports sister-group relationship between Xenacoelomorpha and Ambulacraria. Curr Biol. 2019;29(11):1818–26. https://doi.org/10.1016/j.cub.2019.04.009.
Witek A, Herlyn H, Meyer A, Boell L, Bucher G, Hankeln T. EST based phylogenomics of Syndermata questions monophyly of Eurotatoria. BMC Evol Biol. 2008;8(1):345. https://doi.org/10.1186/1471-2148-8-345.
Witek A, Herlyn H, Ebersberger I, Mark Welch DB, Hankeln T. Support for the monophyletic origin of Gnathifera from phylogenomics. Mol Phylogenet Evol. 2009;53(3):1037–41. https://doi.org/10.1016/j.ympev.2009.07.031.
Min GS, Park JK. Eurotatorian paraphyly: revisiting phylogenetic relationships based on the complete mitochondrial genome sequence of Rotaria rotatoria (Bdelloidea: Rotifera: Syndermata). BMC Genomics. 2009;10(1):533. https://doi.org/10.1186/1471-2164-10-533.
Ricci C. Are lemnisci and proboscis present in the Bdelloidea? Hydrobiologia. 1998;387/388:93–6.
Hur JH, Van Doninck K, Mandigo ML, Meselson M. Degenerate tetraploidy was established before bdelloid rotifer families diverged. Mol Biol Evol. 2008;26:375–83.
Welch DBM, Welch JLM, Meselson M. Evidence for degenerate tetraploidy in bdelloid rotifers. Proc Natl Acad Sci. 2008;105(13):5145–9. https://doi.org/10.1073/pnas.0800972105.
Nowell RW, Wilson CG, Almeida P, Schiffer PH, Fontaneto D, Becks L, et al. Evolutionary dynamics of transposable elements in bdelloid rotifers. Elife. 2021;10:e63194. https://doi.org/10.7554/eLife.63194.
Bushell E, Gomes AR, Sanderson T, Anar B, Girling G, Herd C, et al. Functional profiling of a Plasmodium genome reveals an abundance of essential genes. Cell. 2017;170(2):260–72. https://doi.org/10.1016/j.cell.2017.06.030.
Ferrier DE, Brooke NM, Panopoulou G, Holland PW. The Mnx homeobox gene class defined by HB9, MNR2 and amphioxus AmphiMnx. Dev Genes Evol. 2001;211(2):103–7. https://doi.org/10.1007/s004270000124.
Winterbottom EF, Illes JC, Faas L, Isaacs HV. Conserved and novel roles for the Gsh2 transcription factor in primary neurogenesis. Development. 2010;137(16):2623–31. https://doi.org/10.1242/dev.047159.
Lizen B, Moens C, Mouheiche J, Sacré T, Ahn MT, Jeannotte L, et al. Conditional loss of Hoxa5 function early after birth impacts on expression of genes with synaptic function. Front Mol Neurosci. 2017;10:369. https://doi.org/10.3389/fnmol.2017.00369.
Treffkorn S, Kahnke L, Hering L, Mayer G. Expression of NK cluster genes in the onychophoran Euperipatoides rowelli: implications for the evolution of NK family genes in nephrozoans. EvoDevo. 2018;9(1):17. https://doi.org/10.1186/s13227-018-0105-2.
Fröbius AC, Matus DQ, Seaver EC. Genomic organization and expression demonstrate spatial and temporal Hox gene colinearity in the lophotrochozoan Capitella sp. I PLoS ONE. 2008;3(12):e4004. https://doi.org/10.1371/journal.pone.0004004.
Fröbius AC, Funch P. Rotiferan Hox genes give new insights into the evolution of metazoan bodyplans. Nat Commun. 2017;8(1):9. https://doi.org/10.1038/s41467-017-00020-w.
Kennedy C. Ecology of the Acanthocephala. Cambridge: Cambridge University Press; 2006. p. 1–249. https://doi.org/10.1017/CBO9780511541902.
Meyer A. Acanthocephala. In: Bronn’s Klassen und Ordnungen des Tierreichs, vol. 4. Leipzig: Akademische Verlagsgesellschaft; 1932. p. 1–582.
Jerônimo GT, de Pádua SB, de Andrade Belo MA, Chagas EC, Taboga SR, Maciel PO, et al. Neoechinorhynchus buttnerae (Acanthocephala) infection in farmed Colossoma macropomum: a pathological approach. Aquaculture. 2017;469:124–7. https://doi.org/10.1016/j.aquaculture.2016.11.027.
Silva-Gomes AL, Gomes Coelho-Filho J, Viana-Silva W, Braga-Oliveira MI, Bernardino G, Ismael CJ. The impact of Neoechinorhynchus buttnerae (Golvan, 1956) (Eoacanthocephala: Neochinorhynchidae) outbreaks on productive and economic performance of the tambaqui Colossoma macropomum (Cuvier, 1818), reared in ponds. Lat Am J Aquat Res. 2017;45(2):496–500. https://doi.org/10.3856/vol45-issue2-fulltext-25.
Bolger AM, Lohse M, Usadel B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics. 2014;30(15):2114–20. https://doi.org/10.1093/bioinformatics/btu170.
Li D, Liu CM, Luo R, Sadakane K, Lam TW. MEGAHIT: an ultra-fast single-node solution for large and complex metagenomics assembly via succinct de Bruijn graph. Bioinformatics. 2015;31(10):1674–6. https://doi.org/10.1093/bioinformatics/btv033.
Camacho C, Coulouris G, Avagyan V, Ma N, Papadopoulos J, Bealer K, et al. BLAST+: architecture and applications. BMC Bioinformatics. 2009;10(1):421. https://doi.org/10.1186/1471-2105-10-421.
Hahn C, Bachmann L, Chevreux B. Reconstructing mitochondrial genomes directly from genomic next-generation sequencing reads- a baiting and iterative mapping approach. Nucleic Acids Res. 2013;41(13):e129. https://doi.org/10.1093/nar/gkt371.
Dierckxsens N, Mardulyn P, Smits G. NOVOPlasty: de novo assembly of organelle genomes from whole genome data. Nucleic Acids Res. 2016;45:e18.
Katoh K, Standley DM. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol Biol Evol. 2013;30(4):772–80. https://doi.org/10.1093/molbev/mst010.
Bernt M, Donath A, Jühling F, Externbrink F, Florentz C, Fritzsch G, et al. MITOS: improved de novo metazoan mitochondrial genome annotation. Mol Phylogenet Evol. 2013;69(2):313–9. https://doi.org/10.1016/j.ympev.2012.08.023.
Laslett D, Canback B. ARWEN: a program to detect tRNA genes in metazoan mitochondrial nucleotide sequences. Bioinformatics. 2008;24(2):172–5. https://doi.org/10.1093/bioinformatics/btm573.
Lowe TM, Chan PP. tRNAscan-SE on-line: integrating search and context for analysis of transfer RNA genes. Nucleic Acids Res. 2016;44(W1):W54–7. https://doi.org/10.1093/nar/gkw413.
Langmead B, Salzberg SL. Fast gapped-read alignment with bowtie 2. Nat Methods. 2012;9(4):357–9. https://doi.org/10.1038/nmeth.1923.
Afgan E, Baker D, Batut B, Van Den Beek M, Bouvier D, Čech M, et al. The galaxy platform for accessible, reproducible and collaborative biomedical analyses: 2018 update. Nucleic Acids Res. 2018;46(W1):W537–44. https://doi.org/10.1093/nar/gky379.
Hall TA. BioEdit: a user-friendly biological sequence alignment editor and analysis program for windows 95/98/NT. Nucleic Acids Symp Ser. 1999;41:95–8.
Talavera G, Castresana J. Improvement of phylogenies after removing divergent and ambiguously aligned blocks from protein sequence alignments. Syst Biol. 2007;56(4):564–77. https://doi.org/10.1080/10635150701472164.
Guindon S, Dufayard J-F, Lefort V, Anisimova M, Hordijk W, Gascuel O. New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Syst Biol. 2010;59(3):307–21. https://doi.org/10.1093/sysbio/syq010.
Lefort V, Longueville JE, Gascuel O. SMS: smart model selection in PhyML. Mol Biol Evol. 2017;34(9):2422–4. https://doi.org/10.1093/molbev/msx149.
Ronquist F, Teslenko M, van der Mark P, Ayres DL, Darling A, Hohna S, et al. MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Syst Biol. 2012;61(3):539–42. https://doi.org/10.1093/sysbio/sys029.
Huelsenbeck JP, Ronquist F. MRBAYES: Bayesian inference of phylogenetic trees. Bioinformatics. 2001;17(8):754–5. https://doi.org/10.1093/bioinformatics/17.8.754.
Letunic I, Bork P. Interactive tree of life (iTOL) v3: an online tool for the display and annotation of phylogenetic and other trees. Nucleic Acids Res. 2016;44(W1):W242–5. https://doi.org/10.1093/nar/gkw290.
Marçais G, Kingsford C. A fast, lock-free approach for efficient parallel counting of occurrences of k-mers. Bioinformatics. 2011;27(6):764–70. https://doi.org/10.1093/bioinformatics/btr011.
Simão FA, Waterhouse RM, Ioannidis P, Kriventseva EV, Zdobnov EM. BUSCO: assessing genome assembly and annotation completeness with single-copy orthologs. Bioinformatics. 2015;31(19):3210–2. https://doi.org/10.1093/bioinformatics/btv351.
Smit A, Hubley R. RepeatModeler Open-1.0 RepeatModeler website. Available at: http://www.repeatmasker.org/RepeatModeler/.
Goubert C, Modolo L, Vieira C, Moro CV, Mavingui P, Boulesteix M. De novo assembly and annotation of the Asian tiger mosquito (Aedes albopictus) repeatome with dnaPipeTE from raw genomic reads and comparative analysis with the yellow fever mosquito (Aedes aegypti). Genome Biol Evol. 2015;7(4):1192–205. https://doi.org/10.1093/gbe/evv050.
Koch P, Platzer M, Downie BR. RepARK - de novo creation of repeat libraries from whole-genome NGS reads. Nucleic Acids Res. 2014;42(9):e80. https://doi.org/10.1093/nar/gku210.
Schell T, Feldmeyer B, Schmidt H, Greshake B, Tills O, Truebano M, et al. An annotated draft genome for Radix auricularia (Gastropoda, Mollusca). Genome Biol Evol. 2017;9(3):585–92. https://doi.org/10.1093/gbe/evx032.
Abrusan G, Grundmann N, DeMester L, Makalowski W. TEclass - a tool for automated classification of unknown eukaryotic transposable elements. Bioinformatics. 2009;25(10):1329–30. https://doi.org/10.1093/bioinformatics/btp084.
Dobin A, Davis CA, Schlesinger F, Drenkow J, Zaleski C, Jha S, et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics. 2013;29(1):15–21. https://doi.org/10.1093/bioinformatics/bts635.
Davidson NM, Hawkins ADK, Oshlack A. SuperTranscripts: a data driven reference for analysis and visualisation of transcriptomes. Genome Biol. 2017;18(1):148. https://doi.org/10.1186/s13059-017-1284-1.
Kumar S, Stecher G, Li M, Knyaz C, Tamura K. MEGA X: molecular evolutionary genetics analysis across computing platforms. Mol Biol Evol. 2018;35(6):1547–9. https://doi.org/10.1093/molbev/msy096.
Suyama M, Torrents D, Bork P. PAL2NAL: robust conversion of protein sequence alignments into the corresponding codon alignments. Nucleic Acids Res. 2006;34(Web Server):W609–12. https://doi.org/10.1093/nar/gkl315.
Benjamini Y, Hochberg Y. Controlling the false discovery rate: a practical and powerful approach to multiple testing. J R Stat Soc Ser B Stat Methodol. 1995;57:289–300.
Yang Z. PAML 4: phylogenetic analysis by maximum likelihood. Mol Biol Evol. 2007;24(8):1586–91. https://doi.org/10.1093/molbev/msm088.
Ashkenazy H, Penn O, Doron-Faigenboim A, Cohen O, Cannarozzi G, Zomer O, et al. FastML: a web server for probabilistic reconstruction of ancestral sequences. Nucleic Acids Res. 2012;40(W1):W580–4. https://doi.org/10.1093/nar/gks498.
Price MN, Dehal PS, Arkin AP. FastTree: computing large minimum evolution trees with profiles instead of a distance matrix. Mol Biol Evol. 2009;26(7):1641–50. https://doi.org/10.1093/molbev/msp077.
We gratefully acknowledge excellent logistic support by Station Biologique Roscoff. We also thank two independent reviewers for constructive comments improving the manuscript. The freely distributable script used for alignment editing was thankfully written by Dr. David Rosenkranz, now at the Senckenberg Center for Human Genetics in Frankfurt, on the occasion of a previous study .
Funding was provided by Johannes Gutenberg University Mainz (Stufe 1) and Deutsche Forschungsgemeinschaft (HE 3487/5–1) to HH in collaboration with TH. Open Access funding enabled and organized by Projekt DEAL.
Ethics approval and consent to participate
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Additional file 1.
Additional file 2.
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.
About this article
Cite this article
Mauer, K.M., Schmidt, H., Dittrich, M. et al. Genomics and transcriptomics of epizoic Seisonidea (Rotifera, syn. Syndermata) reveal strain formation and gradual gene loss with growing ties to the host. BMC Genomics 22, 604 (2021). https://doi.org/10.1186/s12864-021-07857-y
- Cryptic speciation
- Evolution of parasitism
- Long branch attraction