Expression of NK cluster genes in the onychophoran Euperipatoides rowelli: implications for the evolution of NK family genes in nephrozoans

Treffkorn, Sandra; Kahnke, Laura; Hering, Lars; Mayer, Georg

doi:10.1186/s13227-018-0105-2

Research
Open Access
Published: 18 July 2018

Expression of NK cluster genes in the onychophoran Euperipatoides rowelli: implications for the evolution of NK family genes in nephrozoans

Sandra Treffkorn ORCID: orcid.org/0000-0002-9160-6862¹,
Laura Kahnke¹,
Lars Hering¹ &
Georg Mayer¹

EvoDevo volume 9, Article number: 17 (2018) Cite this article

2166 Accesses
11 Citations
15 Altmetric
Metrics details

Abstract

Background

Understanding the evolution and development of morphological traits of the last common bilaterian ancestor is a major goal of the evo-devo discipline. The reconstruction of this “urbilaterian” is mainly based on comparative studies of common molecular patterning mechanisms in recent model organisms. The NK homeobox genes are key players in many of these molecular pathways, including processes regulating mesoderm, heart and neural development. Shared features seen in the expression patterns of NK genes have been used to determine the ancestral bilaterian characters. However, the commonly used model organisms provide only a limited view on the evolution of these molecular pathways. To further investigate the ancestral roles of NK cluster genes, we analyzed their expression patterns in the onychophoran Euperipatoides rowelli.

Results

We identified nine transcripts of NK cluster genes in E. rowelli, including single copies of NK1, NK3, NK4, NK5, Msx, Lbx and Tlx, and two copies of NK6. All of these genes except for NK6.1 and NK6.2 are expressed in different mesodermal organs and tissues in embryos of E. rowelli, including the anlagen of somatic musculature and the heart. Furthermore, we found distinct expression patterns of NK3, NK5, NK6, Lbx and Msx in the developing nervous system. The same holds true for the NKL gene NK2.2, which does not belong to the NK cluster but is a related gene playing a role in neural patterning. Surprisingly, NK1, Msx and Lbx are additionally expressed in a segment polarity-like pattern early in development—a feature that has been otherwise reported only from annelids.

Conclusion

Our results indicate that the NK cluster genes were involved in mesoderm and neural development in the last common ancestor of bilaterians or at least nephrozoans (i.e., bilaterians to the exclusion of xenacoelomorphs). By comparing our data from an onychophoran to those from other bilaterians, we critically review the hypothesis of a complex “urbilaterian” with a segmented body, a pulsatile organ or heart, and a condensed mediolaterally patterned nerve cord.

Background

Understanding morphological and major developmental features of the metazoan stem species is one of the major challenges of comparative developmental biology [1, 2]. In particular, the “urbilaterian” has drawn a great deal of attention and resolving the putative morphology of this hypothetical triploblastic animal has proven to be particularly challenging [1,2,3,4,5]. The reconstruction of the “urbilaterian” is primarily based on striking similarities of molecular patterning mechanisms and developmental pathways between extant bilaterian taxa, including the most popular model organisms like the fruit fly Drosophila melanogaster, the nematode Caenorhabditis elegans, the annelid Platynereis dumerilii, and the mouse Mus musculus [1]. These molecular pathways include the anterior–posterior patterning system involving the Hox genes [2, 6, 7], the dorsoventral patterning system using the antagonistic interaction of sog/chordin and dpp/BMP2/4 signaling [8, 9], the pax6/eyeless patterning system for eye development [1, 10, 11], body segmentation processes [4, 12], the anterior patterning system involving empty spiracles and orthodenticle [13], and the posterior patterning system involving even skipped and caudal [14, 15].

Due to these genetic similarities, a complex “urbilaterian” has been proposed, which might have possessed a complex morphology including, e.g., body segmentation, a centralized nervous system, a differentiated somatic and visceral musculature, and a circulatory system with a pulsatile vessel or heart [1, 2]. However, recent phylogenetic analyses have shown that the morphologically simpler organized Xenacoelomorpha (Acoela + Nematodermatida + Xenoturbella) might be the sister group of all remaining bilaterians (Protostomia + Deuterostomia, or “Nephrozoa”), leading some authors to suggest a rather simple “urbilaterian” [16]. Hence, studies increasingly focus on revealing the morphological traits of the “urnephrozoan” rather than the “urbilaterian” [16, 17].

In addition to the conserved developmental genes mentioned above, the NK homeobox genes have been identified as key players in many molecular pathways that have been used to reconstruct the morphological characteristics of the “urbilaterian” or “urnephrozoan” [7, 17,18,19]. The NK genes were first identified in Drosophila melanogaster in 1989 by Kim and Niremberg [20], who established the term “NK genes” using their own initials. These genes belong to the Antennapedia class (ANTP) of homeobox genes, which is one of the largest homeobox gene classes in bilaterians, comprising the Hox, ParaHox, NK, Mega-homeobox and SuperHox gene families [7, 21]. Like the Hox and ParaHox genes, the NK genes are typically arranged in one or several clusters. However, the NK cluster shows a much higher degree of rearrangement due to extensive breakup, inversion and reunion events in different taxa [22, 23]. Based on the composition of the NK cluster in insects, combined with the information from chordates, it has been deduced that the ancestral bilaterian NK cluster comprised nine genes, namely NK1, NK3, NK4, NK5, NK6, NK7, Msx, Lbx and Tlx [7, 21,22,23,24]. In addition to the NK cluster genes, there is a variety of other related genes—the so-called NK-linked (NKL) genes [7]. These genes are defined as NKL genes because they show more sequence similarities to the NK cluster genes than to any other genes of the ANTP class, although they are located outside the NK cluster [7]. However, their evolutionary origin is not fully understood [7, 21].

Comparative genomic analyses and gene expression studies revealed that the NK cluster genes are expressed in mesodermal derivatives across bilaterians, including the dorsal pulsatile vessel or heart, the visceral mesoderm, and various somatic muscles [6, 23, 25,26,27]. For example, NK4 is involved in the development of pulsatile organs in all bilaterians studied thus far [18, 19, 28,29,30]. There has been a long debate on whether or not the simple contractile vessels of invertebrates are homologous to the complex vertebrate heart. This is reflected by the existence of two different definitions of what a “heart” is [31,32,33]. While some authors regard only the chambered circular pumps of vertebrates as a heart, implying an anatomical homology, others use a functional definition, regarding any organ that propels fluid through a circulatory system as a heart [31, 32]. In this study, we apply the latter, more general and common definition [18, 33,34,35,36]. The striking similarities in the NK4 expression patterns across bilaterians have led to the hypothesis that the invertebrate pulsatile vessels and the complex vertebrate heart might indeed be homologous structures [35]. Other NK cluster genes, such as Msx and Lbx, are expressed in specific sets of muscles, suggesting that they specify identity and position for the somatic musculature among bilaterians [18, 19, 37], whereas NK3 is mainly expressed in the developing visceral mesoderm in most species studied [19, 28, 38,39,40,41]. These similarities have been used to propose a complex “urbilaterian,” which is believed to have possessed a differentiated somatic and visceral musculature as well as a pulsatile dorsal vessel [1, 19].

In addition to a function in mesoderm patterning, some NK cluster and NKL genes also show conserved expression patterns in the developing nervous system in both vertebrates and invertebrates [17,18,19, 42, 43]. In particular, Msx, NK6 and NK2.2 are involved in the mediolateral patterning (“dorsoventral patterning” sensu Martín-Durán et al. [17]; we favor the term “mediolateral” to avoid confusion with the dorsoventral patterning regulated by the sog/dpp system) of the trunk nervous system in both protostomes [18, 19, 44] and deuterostomes [42, 43, 45, 46]. Together with other transcription factors, these genes are expressed in a staggered mediolateral pattern in the neuroectoderm [17, 42,43,44,45,46]. These similarities have been used as a key argument for proposing a condensed medial ventral nerve cord for the bilaterian ancestor [43, 44]. However, this hypothesis has been challenged by a recent, more detailed analysis of the mediolateral patterning system in representatives of Xenacoelomorpha and Spiralia, which revealed that this pattern is missing in many of these taxa, indicating convergent evolution of the bilaterian nerve cords [17].

Interestingly, a study of NK genes in the annelid Platynereis dumerilii even revealed that some NK genes are expressed in a segment polarity pattern early in development, which has lead the authors to assume that NK genes might have been involved in segment formation in the last common bilaterian ancestor [19]. Hence, the question arises of whether or not this pattern is also found in other bilaterians.

In order to further explore the putative ancestral roles of NK cluster genes among nephrozoans, we investigated their expression patterns in embryos of the onychophoran Euperipatoides rowelli. Together with tardigrades (water bears) and arthropods (spiders, insects and allies), onychophorans or “velvet worms” comprise the Panarthropoda, which, together with the Cycloneuralia (nematodes, priapulids and allies), form the Ecdysozoa—the clade of molting animals ([47, 48], but see Ref. [49] for a critical review of ecdysozoan phylogeny with respect to the uncertain phylogenetic position of tardigrades and a putative paraphyly of cycloneuralians). We first screened different transcriptomic libraries of E. rowelli as well as the publicly available genomes of the tardigrades Hypsibius exemplaris and Ramazzottius varieornatus [50,51,52] for the putative NK and NKL homologs. We used the identified sequences for a phylogenetic analysis to clarify gene orthology within the entire NK family. We then generated specific probes and performed in situ hybridization on embryos of E. rowelli to clarify the expression patterns of NK cluster genes and the NKL gene NK2.2 in Onychophora. Our findings provide insights into the evolutionary history and putative ancestral roles of NK cluster genes in nephrozoans.

Results

Identification and analysis of NK and NKL homologs in onychophorans and tardigrades

NK/NKL repertoire in Onychophora

Screening of the transcriptome libraries of the onychophoran E. rowelli revealed 17 putative NK and NKL homologs (Fig. 1A, B). Reciprocal BLAST searches against the nr database of GenBank retrieved only single copies of most NK homologs, except for NK6, which is represented by two copies, NK6.1 and NK6.2, in the transcriptome of E. rowelli. To clarify the orthology of the identified homologs, we conducted a phylogenetic analysis of NK and NKL genes across bilaterians (Fig. 1A). Using the ANTP-class member distal-less as an outgroup, the NK and NKL homeobox genes form a monophyletic clade. Within this clade, the individual NK and NKL genes also mostly comprise monophyletic groups with the exception of the vax/Noto assemblage, which includes the monophyletic Emx clade. Taken together, the NK/NKL repertoire of E. rowelli consists of 17 genes, including nine NK cluster genes (NK1, NK3, NK4, NK5, Msx, Lbx, Tlx, NK6.1, and NK6.2) and eight NKL genes (Nedx, vax, Emx, Bari, BarH, Hhex, NK2.1, and NK2.2). The NK cluster gene NK7 and the NKL genes Abox, Ro, Noto, Hlx, Dbx, Bsx, Barx, Nanog and Ventx are missing from the transcriptome of E. rowelli.

NK/NKL repertoire in Tardigrada

Screening of our own transcriptomic libraries of H. exemplaris as well as the publicly available genomes of the two tardigrade species H. exemplaris and R. varieornatus and subsequent phylogenetic analyses revealed single copies of the eight NK cluster genes NK1, NK3, NK4, NK5, NK6, Msx, Lbx and Tlx in both species (Fig. 1A, B). In contrast to this set of NK cluster genes, which is identical in H. exemplaris and R. varieornatus, the complement of NKL genes differs between the two eutardigrade species (Fig. 1A, B). In H. exemplaris, we identified single copies of Abox, Ro, Nedx, Emx, Hhex, two copies of Barx and three copies of BarH, while NK2.1, NK2.2, vax, Noto, Bari, Hlx, Dbx, Bsx, Nanog and Ventx are missing. In R. varieornatus, we identified single copies of NK2.1, NK2.2, Abox, Ro, Nedx, Emx, Hhex and two copies of BarH, while vax, Noto, Bari, Hlx, Dbx, Bsx, Nanog and Ventx are missing. We further analyzed the genomic location of the identified NK cluster genes in both tardigrade species (Fig. 2; Table 1). In the genome of H. exemplaris, we identified two scaffolds that carry an NK gene pair each. Tlx and NK6 are located on the forward strand of scaffold0001, 340,377 bp apart from each other and with a total of 59 non-NK genes between them. Msx and NK3 are located on the reverse strand of scaffold0004, separated by 79,490 bp with a total of 17 non-NK genes (Fig. 2). The analysis of the R. varieornatus genome revealed that five NK genes, NK4, Tlx, NK6, Msx and NK3 are located on scaffold002, spanning a region of more than 2 Mb of the scaffold with distances between the individual genes ranging from 100 kb to 1 Mb, and a total of 518 non-NK genes located in between (Table 1; Fig. 2). NK4, NK6 and NK3 are located on the forward strand, while Tlx and Msx occur on the reverse strand. Another gene pair is found on scaffold005, with Lbx being located on the reverse strand and NK1 on the forward strand. Both genes are separated by ~ 700 kb and a total of 194 non-NK genes (Table 1; Fig. 2).

Table 1 Genomic location and orientation of NK cluster genes in the tardigrades H. exemplaris and R. varieornatus. Genes that are located on the same scaffolds are highlighted in dark blue and light blue for H. exemplaris and dark purple and light purple for R. varieornatus

Full size table

Expression of NK cluster genes in embryos of the onychophoran E. rowelli

Transcription levels of NK cluster genes and the NKL gene \(\text{NK2.2}\)

To assess the abundance of the NK/NKL mRNA sequences and their transcription levels in individual developmental stages of E. rowelli, we performed a short read mapping using the segemehl v0.1.7 software and the ribosomal gene RPL31 as a reference (Fig. 3; Additional file 2). Msx shows the highest (> 12% of the RPL31 level) and NK3 and NK4 the lowest expression levels (< 1% of the RPL31 level) throughout development, whereas NK1, NK5, Lbx, Tlx, NK6.1, NK6.2 and NK2.2 are expressed at levels ranging from 1 to 7%. Interestingly, the transcription levels of the two copies of NK6 are strikingly similar throughout development (Fig. 3).

Expression of \(\text{NK1}\)

This gene is first expressed in the mesoderm of stage I embryos in weak stripes in the anterior half of the jaw, slime papilla, and subsequent five leg-bearing segments, that are delineated by transverse segmental furrows, but not in the antennal segment (Fig. 4A, A′; note that we use the term “segmental furrows” for the morphological indentations of the body surface that are clearly seen in the embryo and that have also been referred to as “furrows,” “grooves” or “segment boundaries” in other studies [53,54,55,56], irrespective of whether or not they are homologous to the segmental/parasegmental grooves of arthropods [57]). As more segments are added posteriorly, the expression extends and the intensity of the signal increases in the anterior segments, while it is still weak in the posterior segments (Fig. 4B, C). At this stage of development, the expression is restricted to the anterior half of the mesoderm of each segment (Fig. 4C, D). As development proceeds and the limb buds start to protrude distally, a mesodermal U-shaped signal appears in each limb bud, which is larger and more prominent in the anterior limbs (Fig. 4E–H). In addition to this U-shaped domain, a weak stripe appears along the dorsal midline of the fourth and fifth legs (Fig. 4F). In later developmental stages, this expression is also visible in the limb buds posterior to the fourth and fifth leg-bearing segments but never anterior to them (Fig. 4H–J).

Expression of \(\text{NK3}\)

NK3 is expressed weakly throughout development, as indicated by the short read mapping (Fig. 3). Its expression is first detectable in stage IV embryos in two individual mesodermal stripes in the anterior developing limbs with one of these stripes being larger and located posteriorly and the other one being smaller and located anterior to the larger stripe (Fig. 5A, D, D′). As development proceeds, the expression in the developing limbs disappears. Additionally, a diffuse NK3 signal appears in the developing central brain neuropil and in the anlagen of the ventral nerve cords (Fig. 5B, C).

Expression of \(\text{NK4}\)

Similar to NK3, the short read mapping revealed a relatively low abundance of NK4 transcripts in the transcriptomes, indicating that its expression is weak throughout development (Fig. 3). The signal developed only after 2 days of staining and the embryos show a high level of unspecific background (Fig. 5E–G). However, at stage IV, two continuous stripes of expression appear on the dorsal side of the embryo, along the dorsal rim of the paired lateral germ bands (Fig. 5E, E′, F, F′). Stripes of expression also occur in the mesoderm of the posterior-most developing limbs (Fig. 5G). Expression of NK4 was undetectable in other developmental stages.

Expression of \(\text{NK5}\)

NK5 expression is first detectable in stage II embryos in two distinct domains in the cephalic lobes that are clearly separated from each other. The anterior domain is thin and curved, while the posterior one is larger and more roundish (Fig. 6A, A′). This signal persists and intensifies in stage III embryos (Fig. 6B). As development proceeds, NK5 expression occurs in a diffuse domain in the developing brain cortex as well as in the developing ventral nerve cords (Fig. 6C–E, J, K). Additional domains appear later in the mesoderm of the posterior developing legs from the fourth leg pair onward as well as in the distal mesoderm of the antennae (Fig. 6F–I).

Expression of \(\text{NK6.1}\) and \(\text{NK6.2}\)

The two copies of NK6 do not only show similar transcription levels throughout development (Fig. 3), but they also exhibit the same expression patterns in all developmental stages studied (Figs. 7A–H, 8A, B; Additional file 3). NK6.1 and NK6.2 are initially expressed at an early stage of development, when the blastoporal slit closes. At that stage, weak expression is seen around the closing blastoporal slit (Fig. 7A, B). Additionally, a diffuse and continuous band of expression is seen along the germ bands, beginning in the jaw segment and ending in a ring-shaped domain around the proctodeum (dotted lines with arrows in Fig. 7A; Additional file 3). This expression is condensed into a thin, continuous stripe in the ventrolateral ectoderm at around the stage when the limb buds start to protrude distally (Fig. 7C–G; Additional file 3). Additionally, diffuse dot-like thickenings—a small anterior and a larger posterior one—are detected next to this band of expression in each segment, decreasing in size and intensity toward the posterior end (Fig. 7C, G; Additional file 3). As development proceeds, these diffuse dots become more defined and oval shaped in the anterior segments, while they still appear as diffuse spots in the posterior segments (Figs. 7G, H, 8A; Additional file 3). Cross sections revealed that these oval-shaped domains are located in the ventromedial ectoderm (Fig. 8C, C′), whereas the longitudinal bands of expression are confined to the developing nerve cords (Fig. 8D, D′). In later developmental stages, the expression of NK6.1 and NK6.2 persists in each ventral nerve cord and expands anteriorly into the jaw segment, while the oval-shaped ectodermal domains disappear (Fig. 8A, B; Additional file 3). An additional domain appears in the distal mesoderm of the developing antennae (Fig. 8B).

Expression of \(\text{Msx}\)

This gene is initially expressed in stage I embryos in each segment except for the antennal segment (Fig. 9A, B). While the expression pattern is still diffuse and continuous in the posterior-most part of the lateral germ bands, i.e., it is not subdivided into segmental domains, the domains are clearly separate from each other in the anterior-most segments, where they occupy the posterior two-thirds of each segment (Fig. 9A–E). In stage III embryos, expression occurs in the mesoderm of the developing limbs, including the antennae (Fig. 9F–H). Additionally, similar to NK6.1 and NK6.2, paired dot-like Msx domains—a small anterior and a larger posterior one—appear in the ventrolateral ectoderm (Figs. 9F, G, 10A, B). As the limb buds grow distally, the expression in the developing limbs becomes restricted to the proximal portion and is absent from the distal portion of each limb (Fig. 9I, J). Furthermore, a continuous band of expression appears in the nerve cords lateral to the paired dot-like domains. Cross sections revealed that the paired dot-like domains are restricted to the center of the ventral ectoderm, while the longitudinal bands of expression are located in the developing nerve cords (Figs. 9K, 10A, B). In later developmental stages, Msx expression disappears from the developing limbs apart from a distinct domain in the distal mesoderm of the developing antennae (Fig. 10C, D). At this stage of development, Msx expression is initiated in the developing brain, including the central brain neuropil, the antennal tracts, the brain cortex, and the connecting pieces that link the brain to the ventral nerve cords (Fig. 10E–H). Furthermore, Msx expression persists in the ventral nerve cords with a decreasing intensity from anterior to posterior, while the paired dot-like domains in the ventral ectoderm have disappeared (Fig. 10C). Later in development, the intensity of expression in the developing brain and ventral nerve cords increases, while the signal in the distal mesoderm of the developing antennae disappears (Fig. 10D, F).

Expression of \(\text{Lbx}\)

Initially, Lbx is expressed in the posterior half of the mesoderm of the jaw segment, slime papilla segment and the first two leg-bearing segments (Fig. 11A, A′). As more segments are added, the expression occurs in all segments that are delineated by transverse furrows (Fig. 11B–F). These domains are restricted to the posterior mesoderm of each segment (Fig. 11B′, E, F). Additionally, a weak signal is detected in the anlagen of the antennae (Fig. 11C, D). Later in development, when the limb buds start to grow distally, the expression in the limb buds becomes restricted to the distal mesoderm (Fig. 11G, H). At around stage IV, these domains expand toward the proximal portion of the anterior limbs, while they are still restricted only to the distal portion of the posterior limbs (Fig. 11I, K). Additionally, a ring-like expression appears in the anterior portion of the cephalic lobes at this stage of development (Fig. 11J).

Expression of \(\text{Tlx}\)

The initial Tlx expression appears as two domains in the anterior mesoderm and one domain in the distal ectoderm of the anterior limb anlagen of the early stage III embryo (Fig. 12A, B). The two mesodermal domains show a stronger signal than the ectodermal domain at stage III (Fig. 12B). However, the signal in the ectoderm increases later in development and reaches a similar level to that in the mesoderm in stage IV and V embryos (Fig. 12D, E). As development proceeds, additional limb buds begin to express Tlx in two mesodermal domains and one distal ectodermal domain (Fig. 12C, F, F′). At this stage of development, the distal mesodermal domain of the anterior limb buds extends proximally, while the corresponding domains in the posterior limbs are still restricted to the tip (Fig. 12D, E, F, F′). The ectodermal expression in the jaw anlagen is delayed with respect to the subsequent segments, as it is first detectable at stage IV (Fig. 12C, D). Moreover, the corresponding domain occurs posterior rather than distally in the anlagen of the jaws (Fig. 12D).

Expression of \(\text{NK2.2}\)

The expression of NK2.2 is initiated in the developing ventral nerve cords of stage II embryos, beginning in the jaw segment. At this stage, a single continuous stripe of expression is seen in most segments, with additional thickenings in the jaw and slime papilla segments (Fig. 13A, C). As development proceeds, this stripe of NK2.2 expression is subdivided into a stronger medial and a weaker lateral domain (Fig. 13B, D–F). Cross-sectioned embryos show that the medial domain is restricted to the medial ectoderm in early and late developmental stages, whereas the lateral domain corresponds to the expression pattern in the lateral ectoderm as well as in the mediolateral portion of the nerve cord in later developmental stages (Fig. 13G–J). This paired pattern persists until late in development (Fig. 13D, F). Additionally, a bilateral pattern of expression is seen in the walls of the stomodeum at stage IV (Fig. 13K), whereas the expression occurs in the walls of the entire mouth cavity at stage VI (Fig. 13L).

Discussion

NK and NKL gene repertoires in Onychophora and Tardigrada

Our searches and phylogenetic analyses revealed a total of 17 NK and NKL genes in the onychophoran E. rowelli and the tardigrade R. varieornatus, whereas the other analyzed tardigrade species, H. exemplaris, possesses 18 genes. We identified nine NK cluster genes, including single copies of NK1, NK3, NK4, NK5, Msx, Lbx, Tlx and two different transcripts of NK6, in the transcriptome of E. rowelli. The genomes of the two tardigrade species and the transcriptome of H. exemplaris show identical sets of eight NK cluster genes, which comprise single copies of NK1, NK3, NK4, NK5, NK6, Msx, Lbx and Tlx. These results show that, apart from the duplication of NK6 in onychophorans (or the onychophoran subgroup containing E. rowelli), onychophorans and tardigrades possess an identical set of eight NK cluster genes, whereas NK7 is missing in both taxa. However, the presence of NK7 in the genomes of several arthropods [Additional file 1; 23, 58] as well as other bilaterians [Additional file 1; 24, 59] suggests that a complete “protoNK cluster” [7, 23, 60, 61] consisting of nine genes was present in the last common ancestor of Panarthropoda and that NK7 was most likely lost in the onychophoran and tardigrade lineages.

Interestingly, localization of NK cluster genes in the genomes of the two tardigrade species revealed that most of these genes are located on different scaffolds. While the NK cluster genes are encoded on six scaffolds in H. exemplaris (with four scaffolds containing one and two scaffolds including two NK genes each), they are encoded on three scaffolds in R. varieornatus (with each scaffold containing one, two and five NK genes, respectively). The gene pairs NK3/NK4 and NK6/Tlx occur in both tardigrade species, suggesting that these pairs were present in the genome of the last common ancestor of the two species. However, even those NK genes that are located on the same scaffold do not neighbor each other but lie at a distance ranging from several kb to up to ~ 1.5 Mb. Moreover, large numbers of non-NK genes are interspersed between the individual NK genes. For comparison, the NK cluster of the fruit fly D. melanogaster spans only about 180 kb and contains only four non-NK genes [18, 23]. We therefore conclude that the NK cluster might have been fragmented in tardigrades. Similar breakups have been reported for the tardigrade Hox cluster (see supplementary figure S3 in Ref. [62]) as well as for the Hox and ParaHox clusters in other bilaterians, including the fruit fly D. melanogaster, the nematode C. elegans and the tunicate Ciona intestinalis [63,64,65]. Comparative analyses of the genomes of H. exemplaris and R. varieornatus generally revealed a low level of synteny, which might be due to extensive intrachromosomal rearrangements similar to those in C. elegans [66]. It has been hypothesized that such breakups of gene clusters are correlated with rapid modes of embryogenesis, possibly due to the loss of temporal collinearity and corresponding regulatory mechanisms in these taxa [63]. Since tardigrades also show rapid embryonic development (4–5 days in H. exemplaris [67]), this might be indeed due to the disintegrated homeobox clusters in these animals.

In contrast to the two tardigrade species, well-assembled genomic data are currently unavailable for Onychophora. Since neither the location nor the orientation of NK genes are known in E. rowelli, it remains unclear whether or not the ancestral NK cluster has been retained in the onychophoran genome.

In contrast to the almost identical sets of NK cluster genes in onychophorans and tardigrades, the NKL gene complement is variable in the three species studied. While E. rowelli shows eight NKL genes (NK2.1, NK2.2, Nedx, vax, Emx, Bari, BarH and Hhex), we found ten NKL genes in H. exemplaris (Abox, Ro, Nedx, Emx, Hhex, plus two copies of Barx and three copies of Barh) and nine in R. varieornatus (NK2.1, NK2.2, Abox, Ro, Nedx, Emx, Hhex, plus two copies of Barh). These results, together with the NKL complements of various arthropods, indicate that the last common ancestor of Panarthropoda might have possessed single copies of each NKL gene except for Nanog and Ventx, which were missing, and that several gene losses and gene duplication events occurred in different lineages (Fig. 1B; Additional file 1). The high variation of the NKL gene repertoire in tardigrades, onychophorans and arthropods suggests that the NKL genes are more prone to evolutionary changes than the NK cluster genes.

Segment polarity-like expression patterns of NK1, Lbx and Msx in the onychophoran E. rowelli

Our data revealed that NK1, Lbx and Msx are expressed early in development and show a stereotypic pattern in the mesoderm (somites) of the onychophoran embryo. Each somite of E. rowelli exhibits a wide anterior NK1 domain, followed by a diffuse medial Msx domain and a posterior Lbx domain (Fig. 14A). This pattern strongly resembles the expression of the so-called segment polarity genes (SPG), which are also expressed in segmentally reiterated domains early in development that are aligned in defined anteroposterior positions within each segment [55, 68,69,70]. To our knowledge, a similar expression pattern of these three NK genes has not been reported from any arthropod species studied thus far, except for an early Lbx expression in D. melanogaster that does show an SPG-like pattern in both ectoderm and mesoderm [71]. Interestingly, NK1, Lbx and Msx are also expressed in an SPG-like pattern in the annelid Platynereis dumerilii, which shares with E. rowelli the formation of somites during its larval development [19]. Based on the distinct complementary expression of these genes early in development of P. dumerilii, the authors [19] have concluded that the role of the NK genes in anterior–posterior segment patterning might be conserved in protostomes. If so, one would expect these genes to be expressed in a similar anterior–posterior alignment in onychophorans.

However, besides NK1, Lbx and Msx at least two other NK cluster genes, namely NK4 and Tlx, are expressed in an SPG-like pattern in P. dumerilii (Fig. 14B). Furthermore, despite the SPG-like expression patterns of NK1, Lbx and Msx, the relative positions and dimensions of individual domains within each somite clearly differ between E. rowelli and P. dumerilii. For example, while NK1 and Lbx are expressed in the anterior and posterior halves of each somite in E. rowelli, respectively, both genes are co-expressed in the anterior portion of each somite in P. dumerilii (Fig. 14A, B). Likewise, Msx is expressed in the posterior two-thirds of each somite in E. rowelli, while it occupies the anterior portion of each somite in P. dumerilii. Most importantly, the expression of Lbx and Msx is restricted to the mesoderm in E. rowelli, whereas these genes show additional ectodermal domains in P. dumerilii (Fig. 14A, B). These major positional discrepancies suggest considerable differences in segment patterning mechanisms in these two taxa.

As another line of evidence, Saudemont et al. [19] state that the expression of Lbx is co-localized with the expression of wingless in P. dumerilii in a domain anterior to engrailed, which is similar to the situation in D. melanogaster, where Lbx expression is co-localized with and dependent on wingless expression. Based on these similarities, they consider it as unlikely that the complementary expression of Lbx, wingless and engrailed might have been recruited independently in annelids and arthropods [19]. In contrast to this, Lbx is not expressed in the ectoderm of E. rowelli at any developmental stage and, thus, is not co-localized with the exclusively ectodermal wingless expression [55]. Instead, the Lbx domain might partially overlap with the mesodermal engrailed domain in onychophorans, which is devoid of Lbx expression in D. melanogaster and P. dumerilii. Consequently, one would have to assume that this pattern has been modified to a large extent in onychophorans, while it was retained in annelids and arthropods. However, the argumentation of Saudemont et al. [19] is based on the expression of only a single NK gene in two distantly related species. Further data from other protostome taxa, including other annelids [72, 73] and arthropods [74,75,76,77,78], so far have not revealed any evidence for an involvement of NK genes in segment formation, which would indicate several independent losses of this pattern in many protostome taxa.

In summary, if the NK genes were involved in the anterior–posterior regionalization of segments in the protostome ancestor, as proposed by Saudemont et al. [19], one would expect a similar set of NK genes to be expressed in defined anteroposterior positions in most protostome taxa, similar to what has been described from the segment polarity genes engrailed, hedgehog, wingless and cubitus interruptus [55, 57, 68]. However, the considerable differences and the lack of specific similarities in the expression patterns observed in P. dumerilii and E. rowelli, as well as the absence of similar patterns in almost all protostomes studied thus far rather indicate that NK1, Msx and Lbx might have been recruited independently to fulfill similar functions in the regionalization of segments in annelids and onychophorans. On the other hand, even though these genes show a segmentally reiterated pattern in both species, which resembles an SPG-like expression, so far there is no convincing evidence for their potential role in the anterior–posterior regionalization of segments.

Conserved expression patterns of NK cluster genes in the mesoderm of E. rowelli

One of the most intriguing features of the NK cluster genes is their seemingly conserved expression in mesodermal derivatives across bilaterians, including various somatic muscles [6, 7, 18]. For example, NK1, Msx and Lbx are expressed in non-overlapping patterns in different sets of longitudinal and parapodial muscles in the annelid P. dumerilii, suggesting that their transcripts might provide identity information for the differentiation of these muscles [19]. In the fruit fly D. melanogaster, NK1, Msx and the two Lbx orthologs (lbl and lbe) are mainly expressed in subsets of muscle founder cells, including the developing dorsal body wall muscles, lateral and segmental muscles [18, 37, 71, 77, 79]. In vertebrate embryos, including the mouse Mus musculus and the chicken Gallus gallus, the Msx and Lbx orthologs are expressed in specific sets of muscle precursors that will give rise to the limb musculature [38,39,40,41, 80,81,82].

Similarly, our data revealed that NK1, NK3, NK4, NK5, Lbx and Tlx are expressed in distinct, mesodermal domains in the developing limbs in embryos of E. rowelli. NK1, NK4 and Lbx show a similar pattern in each developing leg, which follows an anterior-to-posterior progression in development. Similar Tlx domains appear in all developing limbs, although they differ in size and shape in the developing jaws and slime papillae, which are modified cephalic appendages and show a derived muscle arrangement ([56, 83]; see also Fig. 9A–C in Ref. [84]).

These findings indicate that the mesodermal domains of NK1, NK4, Lbx and Tlx might correspond in position to the individual developing limb muscles, and that these genes might provide positional identity information for the limb muscles in onychophorans. However, since development of the somatic musculature of onychophorans has not been studied and since the exact number and arrangement of individual leg muscles has not yet been clarified [85,86,87,88,89,90], the relation of these expression patterns to the development of specific sets of muscles in the onychophoran limbs remains unclear. Nevertheless, our results indicate that the involvement of NK genes in the development and differentiation of somatic musculature might be conserved in Bilateria [19], or at least Nephrozoa, since comparative NK gene expression data are missing from Xenacoelomorpha, the sister group of Nephrozoa [16].

In contrast to the mostly similar mesodermal expression patterns of NK1, NK4, Lbx and Tlx, the posterior domains of NK3 and NK5 are elongated in the fourth and fifth developing legs with respect to their corresponding domains in the remaining limbs. Compared to the jaws and slime papillae, however, these limbs do not show a derived muscle arrangement that would explain these differences [56]. Interestingly, these patterns resemble the expression of odd-skipped, pox-neuro and pax3/7, which also show enlarged domains in the fourth and fifth legs [11, 68]. This pattern might correspond to the anlagen of specific types of nephridia along the onychophoran body: The small NK3 domains in the first three leg-bearing segments might be localized in the tiny nephridial anlagen of these segments, while the elongated NK3 and NK5 domains in the fourth and fifth leg-baring segments might be associated with the large, specialized “labyrinth organs” developing in these segments ([91,92,93]; see also Fig. 10A in [94]). This striking pattern indicates that these genes might be involved in the development of nephridia and their derivatives in Onychophora. To our knowledge, a comparable role of NK genes in nephridiogenesis has only been reported from the annelid P. dumerilii thus far [19].

Apart from the expression of NK genes in the developing limb muscles, we observed peculiar mesodermal stripes of NK4 expression along the dorsal rim of the lateral blastoderm bands. The position of this expression might correspond to mesenchymal cells of the dorsal coelomic linings that move into the space above the midgut to form the future heart [56, 95,96,97]. This pattern strongly resembles the previously described expression of the T-box gene H15 in the dorsal tube of the onychophoran Euperipatoides kanangrensis [98]. This, in turn, is reminiscent of the overlapping expression of NK4 and H15 in the beetle Tribolium castaneum, the fly D. melanogaster and the spider Cupiennius salei along the dorsal rim of the lateral germ bands in cells that have been identified as heart precursors in these animals, suggesting that this pattern might be conserved in onychophorans and arthropods [18, 28,29,30, 35]. These similarities support the hypothesis that the panarthropod hearts are homologous. In contrast to onychophorans and arthropods, however, a vascular system with a pulsatile organ is absent from the third major panarthropod taxon, the tardigrades. This absence might be a secondary loss due to the miniaturized body of tardigrades [99]. Depending on the phylogenetic position of tardigrades, our data suggest that the last common ancestor of either panarthropods or onychophorans plus arthropods possessed a pulsatile dorsal vessel that expressed NK4 during development (Fig. 15A, B).

Interestingly, studies of NK4 expression in other bilaterians, including annelids and chordates, revealed similar patterns in the pulsatile dorsal vessel or heart, which has led to the hypothesis that a role in the formation of the pulsatile dorsal vessel or heart might be conserved among bilaterians, or at least nephrozoans, since xenacoelomorphs do not possess a circulatory system [19, 100,101,102]. Thus, the “urnephrozoan” might have already possessed a pulsatile dorsal vessel. A possible conservation of the NK4 expression in heart precursor cells and a common origin of the heart in panarthropods or onychophorans plus arthropods might support this hypothesis. However, many protostome taxa, including cycloneuralians and numerous spiralian/lophotrochozoan taxa, do not possess heart-like organs or pulsatile vessels. If a heart-like structure indeed evolved only once in the nephrozoan ancestor, one would have to assume multiple independent losses of pulsatile organs in these taxa. Like tardigrades, many of these taxa show extensive miniaturization [103]. Since it has been shown that miniaturization often results in the reduction or complete loss of organs or entire organ systems, multiple independent losses of the vascular system including heart-like structures seem possible [103]. In contrast to this, NK4 expression data from the cephalopod mollusk Sepia officinalis revealed that NK4 is not involved in the formation of the heart but rather in somatic muscle development in this species [104]. Consequently, if one assumes a single origin of the heart in the nephrozoan ancestor, mollusks would have lost NK4 expression in cardiac tissue but retained the heart itself (Fig. 15A).

Interestingly, the example of S. officinalis shows that NK4 is not essential for proper heart development and that the formation of pulsatile tissue can be mediated by other molecular mechanisms [104]. The expression of NK4 in the somatic musculature in S. officinalis rather indicates an ancestral role of this gene in somatic muscle development. Thus, an alternative scenario of heart evolution in nephrozoans is conceivable (Fig. 15B). According to this scenario, a heart was absent in the last common ancestor of nephrozoans and pulsatile organs evolved four times independently in deuterostomes, panarthropods (or onychophorans plus arthropods), annelids, and mollusks. According to this scenario, NK4 might have been involved in somatic muscle development in the nephrozoan ancestor and was recruited independently to fulfill major regulatory functions during heart development in annelids, panarthropods and deuterostomes but not in mollusks.

Non-regionalized neural expression of NK cluster genes and the NKL gene NK2.2 in E. rowelli supports convergent evolution of bilaterian nerve cords

In addition to mesodermal domains, we observed an expression of NK3, NK5, NK6.1, NK6.2, Lbx, Msx and NK2.2 in the developing nervous system of the onychophoran embryo. While NK3, NK5, Msx and NK2.2 are expressed in the anlagen of both the brain and the ventral nerve cords, the expression of the two NK6 copies is restricted to the ventral nervous system, and Lbx is confined to the developing brain. Interestingly, an involvement of NK5, NK6, Msx, Lbx, Tlx and NK2.2 in neural development has also been reported from other bilaterians, including arthropods, annelids and vertebrates [18, 19, 44, 105]. This supports the assumption that these genes were involved in neural development in the last common bilaterian ancestor [18, 19, 44].

In the trunk of E. rowelli, NK5, NK6.1, NK6.2, Msx, Lbx, Tlx and NK2.2 are expressed in uniform domains along the body. However, NK2.2, Msx and both copies of NK6 show peculiar mediolaterally regionalized patterns (Fig. 16A). While NK2.2 is expressed in parallel, continuous, medial and lateral stripes along the body, Msx and both copies of NK6 show both, continuous bands of expression within each nerve cord as well as segmentally reiterated domains in the medial ectoderm (Fig. 16A). Interestingly, Msx, NK6 and NK2.2 have been reported to be involved in the mediolateral regionalization of the nervous system in vertebrates, the fly D. melanogaster and the annelid P. dumerilii [19, 44, 46, 105]. This regionalization is mediated by the staggered expression of the transcription factors NK6, Msx, NK2.2, Pax6 and Pax3/7 in the neuroectoderm of these animals (Fig. 16B), thus defining a specific mediolateral arrangement of neurogenic domains and neuron types [19, 44, 105]. While the medial NK2.2⁺/NK6⁺ domain gives rise to a medial column of serotonergic neurons, the adjacent NK6⁺/pax6⁺ area forms an intermediate column of cholinergic motor neurons and the lateral pax6⁺/pax3/7⁺ and pax3/7⁺/msx⁺ domains establish a lateral column of interneurons and lateral sensory trunk neurons [17, 44]. This seemingly conserved mediolateral patterning has been used as a major argument for proposing an ancestral condensed, mediolaterally patterned ventral nerve cord in the “urbilaterian” [17, 44, 105]. The absence of a mediolateral patterning of these genes in hemichordates, nematodes and planarians has been interpreted as independent losses in these lineages [44].

However, recent studies of representatives of Xenacoelomorpha revealed that NK6, Msx, NK2.2, pax6 and pax3/7 expression is unrelated to the trunk neuroanatomy in these taxa, suggesting that the mediolateral patterning evolved after the Xenacoelomorpha–Nephrozoa split [17]. Interestingly, investigation of these genes in additional nephrozoan taxa, including Rotifera, Nemertea, Brachiopoda and Enteropneusta, revealed that a mediolateral regionalization is also absent or only partially present in these taxa [17, 42]. Moreover, despite the presence of mediolateral regionalization in the annelid P. dumerilii (Errantia, Phyllodocida), there is no evidence for such a pattern in the annelid Owenia fusiformis (Sedentaria, Sabellida), although its trunk neuroanatomy largely resembles that of P. dumerilii, D. melanogaster and vertebrates [17]. Thus, two possible scenarios on the evolution of condensed medial nerve cords among bilaterians have been proposed ([17] but see an opposing view in Ref. [106]). According to the first scenario, the mediolateral patterning of the central nervous system in vertebrates, D. melanogaster and P. dumerilii is homologous, thus reflecting the ancestral bilaterian or nephrozoan state, which would imply multiple independent losses or modifications. Alternatively, the absence of mediolateral regionalization of the central nervous system in xenacoelomorphs, many spiralians and some annelids might indicate a convergent evolution of this patterning system in vertebrates, arthropods and some annelids.

Our results show that NK2.2, NK6.1, NK6.2 and Msx are expressed in dynamic patterns early and late in development of E. rowelli. At the onset of neurogenesis, when neural precursors segregate from the ventral ectoderm [56], expression of these genes is mainly restricted to the ventral ectoderm (Fig. 16A). NK2.2 and Msx are expressed in non-overlapping medial and lateral domains, respectively, that resemble the NK2.2 and Msx domains of other bilaterians. In contrast to this, NK6 is expressed in a lateral domain, overlapping with Msx, but not with NK2.2 (Fig. 16A). This is different from what has been reported from arthropods, annelids and vertebrates, where NK6 is expressed in a medial domain, overlapping with NK2.2 but not with Msx (Fig. 16B).

Later in development, after most neural precursors have been segregated and the developing nerve cords have delaminated from the ventral ectoderm, these genes are expressed in both the ventral ectoderm and the nerve cords where they show different patterns (Fig. 16A). In the ventral ectoderm, NK2.2 and NK6 are expressed in medially restricted, largely overlapping domains, while Msx expression is confined to the central region, partially overlapping with the medial NK2.2 and NK6 domains. NK2.2 shows an additional domain in the lateral ectoderm, which partially overlaps with the Msx domain, but not with the medial NK6 domain. In the nerve cords, NK6 and Msx are expressed in broad, largely overlapping domains that are confined to the medial part, while NK2.2 is restricted to a spot-like expression in the lateral nerve cord, overlapping with the NK6 and Msx domains (Fig. 16A).

Similarly, previous gene expression data on pax6 and pax3/7 did not provide any evidence for regionalized mediolateral patterning in Onychophora [11, 107]. Instead, pax6 is expressed in a broad domain, which covers the entire width of the nerve cord in late developmental stages (see Fig. 9D in [11]), thus overlapping with the NK2.2, NK6 and Msx domains and extending further laterally (Fig. 16A). In contrast, pax3/7 is not expressed in the developing nerve cords at any developmental stage (see Fig. 8A–D in [11] and Fig. 5A–C in [68]). Thus, irrespective of the developmental stage studied, neither our results nor previously published data on pax6 and pax3/7 expression provide any evidence for the presence of adjacent NK2.2⁺/NK6⁺, NK6⁺/pax6⁺, pax6⁺/pax3/7⁺ and pax3/7⁺/msx⁺ columns in the onychophoran nerve cords.

These results are in line with selective stainings of specific neurons, including retrograde fills of the leg nerves, and immunolabeling of various neurotransmitters and neuromodulators, which did not provide any evidence for adjacent columns of medial serotonergic neurons, cholinergic motor neurons, lateral interneurons or sensory neurons in the onychophoran nerve cords [108,109,110]. Instead, most of the neuron types are located in largely overlapping areas [108]. For example, the somata of serotonergic neurons are not restricted to a medial area but are rather distributed in a random fashion in the ventromedial and ventral perikaryal layers [108, 109].

While the expression of NK2.2, NK6 and Msx in the ventral ectoderm of early developmental stages as well as in the nerve cords of late developmental stages might be correlated to the specification of neurons, their corresponding patterns in the ventral ectoderm of late developmental stages are not necessarily correlated with any neural structures. Interestingly, the double-paired patterns of NK6 and Msx in the ventromedial ectoderm correspond to the emergence of paired ectodermal thickenings which are the anlagen of the ventral and preventral organs [84, 94, 111]. Although these structures arise from the ventral ectoderm, they do not seem to contribute any cells to the central nervous system but rather persist as attachment sites for segmental limb muscles in adult onychophorans ([56, 84, 112, 113] but see ref [114] for an opposing view on a putative function of ventral organs in neurogenesis). Interestingly, similar patterns have been reported from Delta and Notch transcripts that are expressed in two pairs of bilaterally symmetric domains on the ventrum of each segment in embryos of Euperipatoides kanangrensis and E. rowelli [84, 114, 115]. This characteristic, double-paired pattern appears only after most neurons have been segregated from the neuroectoderm [84, 109, 116]. Thus, it has been concluded, that these Delta and Notch domains specify regions of the ectoderm that give rise to the ventral and preventral organs rather than neurogenic tissue in the onychophoran embryo [56, 84]. Likewise, our results indicate that the double-paired segmental Msx and NK6 domains in the ventral ectoderm might provide identity information for specific regions of the ectoderm that give rise to the ventral and preventral organs rather than neurogenic tissue because the nerve cords have already been segregated from the ectoderm at this developmental stage (cf. Ref. [56]).

Thus, NK6, Msx, NK2.2, pax6 and pax3/7 expression seems to be unrelated to the trunk neuroanatomy in onychophorans, which is similar to what has been reported from Xenacoelomorpha, Rotifera, Nemertea, Brachiopoda and Enteropneusta, and some Annelida [17]. Consequently, if a complex medial nerve cord with a distinct regionalization of specific neuron types was present in the nephrozoan ancestor, one would have to assume either multiple losses of the mediolateral pattern in several lineages (including nematodes and onychophorans; Fig. 17B), or a single loss in the ecdysozoan ancestor followed by a secondary gain of this pattern in the arthropod lineage (Fig. 17C). The latter scenario would further imply a homology of the medially condensed nerve cords in vertebrates and annelids, but also an independent evolution of medially condensed nerve cords in arthropods (Fig. 17C). Alternatively, a mediolateral regionalization of the nerve cords might have never existed in the panarthropod ancestor, which would support the hypothesis [17] of convergent evolution of a condensed, mediolaterally patterned nerve cord in vertebrates, arthropods and some annelids (Fig. 17A).

Conclusion

Our analysis of the NK cluster genes in the onychophoran E. rowelli revealed that these genes are involved in various developmental processes. Our major findings can be summarized as follows:

NK1, Msx and Lbx are expressed in a segment polarity-like pattern early in development of E. rowelli—a pattern that otherwise has only been reported from the annelid P. dumerilii. However, major differences in timing, position and extent of these patterns suggest considerable differences in segment patterning mechanisms between the two animal groups, which argue against a common origin of the segment polarity patterns of NK genes in onychophorans and annelids.
NK1, NK3, NK4, NK5, Lbx and Tlx are expressed in distinct domains in the developing limb musculature of E. rowelli, which corresponds to the involvement of these genes in somatic muscle development in other bilaterian taxa, suggesting that this role is conserved in Nephrozoa.
Our results on NK4 expression further provide evidence for a common origin of the dorsal heart in onychophorans and arthropods.
NK3, NK5, NK6, Lbx, Msx and NK2.2 are expressed in the developing nervous system of E. rowelli, which resembles the situation in other nephrozoans, indicating that these genes were involved in neural patterning in the last common ancestor of Nephrozoa. Furthermore, our results indicate that a staggered mediolateral expression of NK6, Msx and NK2.2 was either lost or was never present in Onychophora.

Methods

Specimens and sample preparation

Specimens of Euperipatoides rowelli Reid, 1996 (Peripatopsidae) were collected from decaying logs in the Tallaganda State Forest (35°26′S, 149°33′E, 954 m, New South Wales, Australia) in October 2011, January 2013 and October 2016. Permissions for specimen collection were obtained from the National Parks & Wildlife Service New South Wales (permit numbers: SL100159 and SL101720). Permissions for the export of specimens were obtained from the Department of Sustainability, Environment, Water, Population and Communities (permit numbers: PWSP104061, PWSP208163, and PWS2016-AU-001023). The animals were maintained in the laboratory as described previously [118]. Females were anesthetized with chloroform vapor for 10–20 s and the reproductive tracts were dissected and transferred to a physiological saline [119]. Embryos were dissected from the uteri, and the two membranes surrounding each embryo were removed using fine forceps. The embryos were staged according to Walker MH and Tait NN [53] with modifications described previously [55, 120]. After staging, the embryos were fixed overnight in 4% paraformaldehyde in phosphate-buffered saline (PBS; 0.1 M, pH 7.4), dehydrated in a graded methanol series and stored in absolute methanol at − 20 °C.

Identification of NK homologs

Library preparation and assembly of the embryonic transcriptomes from the onychophoran E. rowelli and the tardigrade Hypsibius exemplaris were performed as described previously [121, 122]. The latter species was commonly referred to as “Hypsibius dujardini (Doyère, 1840)” in the literature (e.g., [51, 67, 122,123,124,125,126]), but it was described recently [127] as Hypsibius exemplaris Gąsiorek et al., 2018. To identify the expressed homologs of NK homeobox genes in onychophorans and tardigrades, the transcriptome libraries of E. rowelli and H. exemplaris were screened by local tBLASTx searches [128]. For the gene Tlx, the previously published homolog clawless from Euperipatoides kanangrensis was used as a query [129]. For all other NK genes, previously published sequences from Drosophila melanogaster were retrieved from the online database HomeoDB (http://homeodb.zoo.ox.ac.uk/; accessed 25 Jan 2018) [130, 131] and used as queries. The retrieved putative NK gene sequences were then verified by reciprocal BLASTn and tBLASTx searches against the nr database of GenBank. Additional BLAST searches of the putative tardigrade NK homologs in two independently published genomes of H. exemplaris [see 50, 51] yielded nearly identical sequences. The sequences of identified NK homeobox genes from E. rowelli were made available in GenBank (Table 2).

Table 2 GenBank accession numbers of the identified NK homeobox genes from E. rowelli, and commonly used synonyms for each gene family

Full size table

Sequence alignment, phylogenetic analysis and short read mapping

The amino acid sequences of the homeodomain of NK genes of various model organisms were retrieved from the online database HomeoDB (http://homeodb.zoo.ox.ac.uk/; accessed 25 Jan 2018) [130, 131]. Furthermore, publicly available resources and databases (nr, TSA and EST databases of GenBank) as well as the genomes of the centipede Strigamia maritima [58], the common house spider Parasteatoda tepidariorum [132], the tardigrade Ramazzottius varieornatus [52] and the crustacean Daphnia pulex [133] were screened for putative NK homologs. The previously published NK sequences of the polychaete Platynereis dumerilii were retrieved from GenBank [19]. For the phylogenetic analysis, we generated a dataset with a total of 382 sequences containing the 60 amino acid positions of the homeobox of various metazoans using the online tool MAFFT version 7 [http://mafft.cbrc.jp/alignment/server/; 134] with the L-INS-I alignment strategy (Additional file 5). Afterward, a maximum likelihood analysis was performed using the Pthreads-SSE3 version of RAxML v8.2.10 [135] under a dataset-specific GTR substitution model. The best tree was obtained from 100 independent inferences and GAMMA correction of the final tree. Bootstrap values were calculated from 1000 pseudoreplicates using the rapid bootstrapping algorithm implemented in RAxML (Fig. 1 and Additional file 6). The phylogeny was visualized with iTol and edited with Adobe (San Jose, CA, USA) Illustrator CS 5.1.

The abundance of the obtained NK homologs was estimated using segemehl v0.1.7 [136] by mapping the raw sequence reads of various embryonic transcriptome libraries of E. rowelli [see 121] back to the NK sequences of E. rowelli allowing for a maximum of 5% mismatching nucleotides per read. Afterward, the relative abundance [measured in matched nucleotides per position and per giga base pair (Gb)] of the NK genes were normalized using the relative abundance of the ribosomal protein RPL31 as a reference (= 100%) and visualized as a bar graph (Fig. 3; Additional file 2). We have chosen RPL31 as reference since this gene shows the lowest variation [relative standard deviation (RSD) = 7.69%] among the studied transcriptomes as well as the tested putative reference genes (GAPDH and 44 ribosomal protein genes; Additional file 2). Furthermore, we analyzed the localization and orientation of NK genes in the publicly available genomes of Hypsibius exemplaris [see 50] and Ramazzottius varieornatus [see 52]. We checked the scaffold location, direction, distance as well as the number of other genes between each NK gene.

Nomenclature

The nomenclature of NK genes is confusing since a large variety of synonyms has been established for each NK gene in different animal groups. Thus, we decided to use the general gene family names to avoid further confusion (see table 1 in Ref. [7], table 2 in Ref. [21], and table 1 in Ref. [137] for summaries of NK family genes and their synonyms). The abbreviation “cll” was used previously [129] for the Tlx ortholog in the onychophoran Euperipatoides kanangrensis; we instead use the more general name/abbreviation “Tlx” for the corresponding ortholog in the closely related species E. rowelli.

RNA isolation, amplification of gene fragments, molecular cloning, probe preparation and whole-mount in situ hybridization

RNA isolation and cDNA synthesis were performed as described previously [11, 55, 138]. Fragments of NK1, NK3, NK4, NK5, NK6, Msx, Lbx and Tlx were amplified from the cDNA using specific primers (Table 3) and Phusion Green High-Fidelity DNA Polymerase (2 U/µl; Thermo Scientific, Waltham, MA, USA) to obtain PCR fragments with blunt ends. The gene fragments that were amplified from the cDNA (Table 3) were cloned into the pJet1.2/blunt cloning vector (Thermo Scientific). Digoxigenin-labeled RNA probes were prepared using the DIG RNA Labeling Kit SP6/T7 (Roche, Mannheim, Germany). In situ hybridization on whole embryos was performed as described previously [120] with the following modifications: (1) Stage V and VI embryos were digested in 25 µl of a chitinase/chymotrypsin solution (50 mg/ml chitinase and 15 mg/ml chymotrypsin in potassium phosphate buffer; Sigma-Aldrich, St. Louis, MO, USA) diluted in 1 ml PBST (PBS; 0.1 M; pH 7.4; with 0.1% Tween-20) prior to the pre-hybridization, followed by three 5 min washes in PBST; (2) after the staining reaction in NBT/BCIP solution and the postfixation with 4% PFA, the embryos were dehydrated in an increasing ethanol series (25%, 50%, 75%, 2 × 100% in PBST) and left in ethanol overnight; after rehydration in a decreasing ethanol series, the embryos were counterstained with the DNA-selective fluorescent dye 4′,6-diamidin-2-phenylindol (DAPI; Thermo Scientific, formerly Invitrogen; diluted 1:1000 in PBS) for 1 h and stored in 70% glycerol. Cross and sagittal sections were prepared manually using a razor blade.

Table 3 Specific primers used to amplify the transcripts of NK homeobox genes from cDNA of the onychophoran E. rowelli

Full size table

Controls were performed using the sense probes of each gene and the same protocol as described above. Early embryonic stages (0 to IV) did not show any labeling. Staining artifacts that have been reported to appear in cuticular structures in late developmental stages [139, 140] are absent in the sense probes treated with chitinase/chymotrypsin prior to hybridization. The only remaining unspecific labeling appears in the sclerotized jaws and claws of stage VII embryos. Thus, we conclude that the staining obtained with the antisense probes is specific for all genes.

Microscopy and image processing

The embryos were analyzed with the stereomicroscope Axio Zoom V16 (Carl Zeiss MicroImaging GmbH, Jena, Germany) equipped with an Axiocam 503 color digital camera (Carl Zeiss MicroImaging GmbH). Micrographs were taken at different focal planes and merged to single projections using the ZEN 2012 blue edition software version 1.1.2.0 (Carl Zeiss MicroImaging GmbH) or the Auto-Blend Layers function in Adobe Photoshop version CS 5.1. The micrographs were adjusted for brightness and contrast using Adobe Photoshop CS 5.1. Final panels and diagrams were designed using Adobe Illustrator CS 5.1.

References

1.
Hejnol A, Martindale MQ. Acoel development supports a simple planula-like urbilaterian. Philos Trans R Soc Lond B Biol Sci. 2008;363:1493–501.
PubMed PubMed Central Article Google Scholar
2.
Carroll SB, Grenier JK, Weatherbee SD. From DNA to diversity. molecular genetics and the evolution of animal design. Malden: Blackwell Publishing; 2005.
Google Scholar
3.
Shenk MA, Steele RE. A molecular snapshot of the metazoan ‘Eve’. Trends Biochem Sci. 1993;18:459–63.
PubMed Article CAS Google Scholar
4.
Balavoine G, Adoutte A. The segmented Urbilateria: a testable scenario. Integr Comp Biol. 2003;43:137–47.
PubMed Article Google Scholar
5.
Hirth F, Kammermeier L, Frei E, Walldorf U, Noll M, Reichert H. An urbilaterian origin of the tripartite brain: developmental genetic insights from Drosophila. Development. 2003;130:2365–73.
PubMed Article CAS Google Scholar
6.
Holland PWH. Evolution of homeobox genes. Wiley Interdiscip Rev Dev Biol. 2013;2:31–45.
PubMed Article CAS Google Scholar
7.
Ferrier DEK. Evolution of homeobox gene clusters in animals: the Giga-cluster and primary versus secondary clustering. Front Ecol Evol. 2016;4:36.
Article Google Scholar
8.
Arendt D, Nübler-Jung K. Inversion of dorsoventral axis? Nature. 1994;371:26.
PubMed Article CAS Google Scholar
9.
De Robertis EM, Sasai Y. A common plan for dorsoventral patterning in Bilateria. Nature. 1996;380:37–40.
PubMed Article Google Scholar
10.
Gehring WJ, Ikeo K. Pax6 mastering eye morphogenesis and eye evolution. Trends Genet. 1999;15:371–7.
PubMed Article CAS Google Scholar
11.
Franke FA, Schumann I, Hering L, Mayer G. Phylogenetic analysis and expression patterns of Pax genes in the onychophoran Euperipatoides rowelli reveal a novel bilaterian Pax subfamily. Evol Dev. 2015;17:3–20.
PubMed Article CAS Google Scholar
12.
Tautz D. Segmentation. Dev Cell. 2004;7:301–12.
PubMed Article CAS Google Scholar
13.
Reichert H. A tripartite organization of the urbilaterian brain: developmental genetic evidence from Drosophila. Brain Res Bull. 2005;66:494.
Article Google Scholar
14.
Patel NH, Ball EE, Goodman CS. Changing role of even-skipped during the evolution of insect pattern formation. Nature. 1992;357:339–42.
PubMed Article CAS Google Scholar
15.
Wu LH, Lengyel JA. Role of caudal in hindgut specification and gastrulation suggests homology between Drosophila amnioproctodeal invagination and vertebrate blastopore. Development. 1998;125:2433–42.
PubMed CAS Google Scholar
16.
Cannon JT, Vellutini BC, Smith J, Ronquist F, Jondelius U, Hejnol A. Xenacoelomorpha is the sister group to Nephrozoa. Nature. 2016;530:89–93.
PubMed Article CAS Google Scholar
17.
Martín-Durán JM, Pang K, Børve A, Lê HS, Furu A, Cannon JT, Jondelius U, Hejnol A. Convergent evolution of bilaterian nerve cords. Nature. 2018;553:45–50.
PubMed Article CAS Google Scholar
18.
Jagla K, Bellard M, Frasch M. A cluster of Drosophila homeobox genes involved in mesoderm differentiation programs. BioEssays. 2001;23:125–33.
PubMed Article CAS Google Scholar
19.
Saudemont A, Dray N, Hudry B, Le Gouar M, Vervoort M, Balavoine G. Complementary striped expression patterns of NK homeobox genes during segment formation in the annelid Platynereis. Dev Biol. 2008;317:430–43.
PubMed Article CAS Google Scholar
20.
Kim Y, Nirenberg M. Drosophila NK-homeobox genes. Proc Natl Acad Sci USA. 1989;86:7716–20.
PubMed Article CAS Google Scholar
21.
Holland PWH, Booth HAF, Bruford EA. Classification and nomenclature of all human homeobox genes. BMC Biol. 2007;5:47.
PubMed PubMed Central Article CAS Google Scholar
22.
Luke GN, Castro LFC, McLay K, Bird C, Coulson A, Holland PWH. Dispersal of NK homeobox gene clusters in amphioxus and humans. Proc Natl Acad Sci USA. 2003;100:5292–5.
PubMed Article CAS Google Scholar
23.
Chan C, Jayasekera S, Kao B, Páramo M, von Grotthuss M, Ranz JM. Remodelling of a homeobox gene cluster by multiple independent gene reunions in Drosophila. Nat Commun. 2015;6:6509.
PubMed Article CAS Google Scholar
24.
Butts T, Holland PWH, Ferrier DEK. The urbilaterian Super-Hox cluster. Trends Genet. 2008;24:259–62.
PubMed Article CAS Google Scholar
25.
Garcia-Fernandez J. The genesis and evolution of homeobox gene clusters. Nat Rev Genet. 2005;6:881–92.
PubMed Article CAS Google Scholar
26.
Larroux C, Fahey B, Degnan SM, Adamski M, Rokhsar DS, Degnan BM. The NK homeobox gene cluster predates the origin of Hox genes. Curr Biol. 2007;17:706–10.
PubMed Article CAS Google Scholar
27.
Evans SM. Vertebrate tinman homologues and cardiac differentiation. Semin Cell Dev Biol. 1999;10:73–83.
PubMed Article CAS Google Scholar
28.
Azpiazu N, Frasch M. tinman and bagpipe: two homeobox genes that determine cell fates in the dorsal mesoderm of Drosophila. Genes Dev. 1993;7:1325–40.
PubMed Article CAS Google Scholar
29.
Janssen R, Damen WGM. Diverged and conserved aspects of heart formation in a spider. Evol Dev. 2008;10:155–65.
PubMed Article CAS Google Scholar
30.
Bodmer R, Jan LY, Jan YN. A new homeobox-containing gene, msh-2, is transiently expressed early during mesoderm formation of Drosophila. Development. 1990;110:661–9.
PubMed CAS Google Scholar
31.
Xavier-Neto J, Castro RA, Sampaio AC, Azambuja AP, Castillo HA, Cravo RM, Simäes-Cost MS. Parallel avenues in the evolution of hearts and pumping organs. Cell Mol Life Sci. 2007;64:719–34.
PubMed Article CAS Google Scholar
32.
Simões-Costa MS, Vasconcelos M, Sampaio AC, Cravo RM, Linhares VL, Hochgreb T, Yan CYI, Davidson B, Xavier-Neto J. The evolutionary origin of cardiac chambers. Dev Biol. 2005;277:1–15.
PubMed Article CAS Google Scholar
33.
Olson EN. Gene regulatory networks in the evolution and development of the heart. Science. 2006;313:1922–7.
PubMed PubMed Central Article CAS Google Scholar
34.
Nielsen C. Animal evolution: interrelationships of the living phyla. 3rd ed. Oxford: Oxford University Press; 2012.
Google Scholar
35.
Bodmer R, Venkatesh TV. Heart development in Drosophila and vertebrates: conservation of molecular mechanisms. Dev Genet. 1998;22:181–6.
PubMed Article CAS Google Scholar
36.
Harvey RP. NK-2 Homeobox genes and heart development. Dev Biol. 1996;178:203–16.
PubMed Article CAS Google Scholar
37.
Jagla T, Bellard F, Lutz Y, Dretzen G, Bellard M, Jagla K. Ladybird determines cell fate decisions during diversification of Drosophila somatic muscles. Development. 1998;125:3699–708.
PubMed CAS Google Scholar
38.
Kanamoto T, Terada K, Yoshikawa H, Furukawa T. Cloning and expression pattern of lbx3, a novel chick homeobox gene. Gene Expr Patterns. 2006;6:241–6.
PubMed Article CAS Google Scholar
39.
Dietrich S, Schubert FR, Healy C, Sharpe PT, Lumsden A. Specification of the hypaxial musculature. Development. 1998;125:2235–49.
PubMed CAS Google Scholar
40.
Gross MK, Moran-Rivard L, Velasquez T, Nakatsu MN, Jagla K, Goulding M. Lbx1 is required for muscle precursor migration along a lateral pathway into the limb. Development. 2000;127:413–24.
PubMed CAS Google Scholar
41.
Brohmann H, Jagla K, Birchmeier C. The role of Lbx1 in migration of muscle precursor cells. Development. 2000;127:437–45.
PubMed CAS Google Scholar
42.
Kaul-Strehlow S, Urata M, Praher D, Wanninger A. Neuronal patterning of the tubular collar cord is highly conserved among enteropneusts but dissimilar to the chordate neural tube. Sci Rep. 2017;7:7003.
PubMed PubMed Central Article CAS Google Scholar
43.
Holland LZ, Carvalho JE, Escriva H, Laudet V, Schubert M, Shimeld SM, Yu J-K. Evolution of bilaterian central nervous systems: a single origin? EvoDevo. 2013;4:27.
PubMed PubMed Central Article Google Scholar
44.
Denes AS, Jékely G, Steinmetz PRH, Raible F, Snyman H, Prud’homme B, Ferrier DEK, Balavoine G, Arendt D. Molecular architecture of annelid nerve cord supports common origin of nervous system centralization in Bilateria. Cell. 2007;129:277–88.
PubMed Article CAS Google Scholar
45.
Arendt D, Tosches MA, Marlow H. From nerve net to nerve ring, nerve cord and brain—evolution of the nervous system. Nat Rev Neurosci. 2016;17:61–72.
PubMed Article CAS Google Scholar
46.
Arendt D, Denes AS, Jekely G, Tessmar-Raible K. The evolution of nervous system centralization. Philos Trans R Soc Lond B Biol Sci. 2008;363:1523–8.
PubMed PubMed Central Article Google Scholar
47.
Aguinaldo AMA, Turbeville JM, Linford LS, Rivera MC, Garey JR, Raff RA, Lake JA. Evidence for a clade of nematodes, arthropods and other moulting animals. Nature. 1997;387:489–93.
PubMed Article CAS Google Scholar
48.
Giribet G. Molecules, development and fossils in the study of metazoan evolution; Articulata versus Ecdysozoa revisited. Zoology. 2003;106:303–26.
PubMed Article CAS Google Scholar
49.
Giribet G, Edgecombe GD. Current understanding of Ecdysozoa and its internal phylogenetic relationships. Integr Comp Biol. 2017;57:455–66.
PubMed Article Google Scholar
50.
Arakawa K, Yoshida Y, Tomita M. Genome sequencing of a single tardigrade Hypsibius dujardini individual. Sci Data. 2016;3:160063.
PubMed PubMed Central Article CAS Google Scholar
51.
Koutsovoulos G, Kumar S, Laetsch DR, Stevens L, Daub J, Conlon C, Maroon H, Thomas F, Aboobaker AA, Blaxter M. No evidence for extensive horizontal gene transfer in the genome of the tardigrade Hypsibius dujardini. Proc Natl Acad Sci USA. 2016;113:5053–8.
PubMed Article CAS Google Scholar
52.
Hashimoto T, Horikawa DD, Saito Y, Kuwahara H, Kozuka-Hata H, Shin-I T, Minakuchi Y, Ohishi K, Motoyama A, Aizu T, et al. Extremotolerant tardigrade genome and improved radiotolerance of human cultured cells by tardigrade-unique protein. Nat Commun. 2016;7:12808.
PubMed PubMed Central Article CAS Google Scholar
53.
Walker MH, Tait NN. Studies on embryonic development and the reproductive cycle in ovoviviparous Australian Onychophora (Peripatopsidae). J Zool. 2004;264:333–54.
Article Google Scholar
54.
Eriksson BJ, Tait NN, Budd GE, Akam M. The involvement of engrailed and wingless during segmentation in the onychophoran Euperipatoides kanangrensis (Peripatopsidae: Onychophora) (Reid 1996). Dev Genes Evol. 2009;219:249–64.
PubMed Article Google Scholar
55.
Franke FA, Mayer G. Controversies surrounding segments and parasegments in Onychophora: insights from the expression patterns of four “segment polarity genes” in the peripatopsid Euperipatoides rowelli. PLoS ONE. 2014;9:e114383.
PubMed PubMed Central Article CAS Google Scholar
56.
Mayer G, Franke FA, Treffkorn S, Gross V, Oliveira IS. Onychophora. In: Wanninger A, editor. Evolutionary Developmental Biology of Invertebrates 3: Ecdysozoa I: Non-Tetraconata. Wien: Springer; 2015. p. 53–98.
Google Scholar
57.
Janssen R. A molecular view of onychophoran segmentation. Arthropod Struct Dev. 2017;46:341–53.
PubMed Article Google Scholar
58.
Chipman AD, Ferrier DEK, Brena C, Qu J, Hughes DST, Schröder R, Torres-Oliva M, Znassi N, Jiang H, Almeida FC, et al. The first myriapod genome sequence reveals conservative arthropod gene content and genome organisation in the centipede Strigamia maritima. PLoS Biol. 2014;12:e1002005.
PubMed PubMed Central Article CAS Google Scholar
59.
Takatori N, Butts T, Candiani S, Pestarino M, Ferrier DEK, Saiga H, Holland PWH. Comprehensive survey and classification of homeobox genes in the genome of amphioxus, Branchiostoma floridae. Dev Genes Evol. 2008;218:579–90.
PubMed Article CAS Google Scholar
60.
Di Z, Yu Y, Wu Y, Hao P, He Y, Zhao H, Li Y, Zhao G, Li X, Li W, Cao Z. Genome-wide analysis of homeobox genes from Mesobuthus martensii reveals Hox gene duplication in scorpions. Insect Biochem Mol Biol. 2015;61:25–33.
PubMed Article CAS Google Scholar
61.
Chai C-L, Zhang Z, Huang F-F, Wang X-Y, Yu Q-Y, Liu B-B, Tian T, Xia Q-Y, Lu C, Xiang Z-H. A genomewide survey of homeobox genes and identification of novel structure of the Hox cluster in the silkworm, Bombyx mori. Insect Biochem Mol Biol. 2008;38:1111–20.
PubMed Article CAS Google Scholar
62.
Smith Frank W, Boothby Thomas C, Giovannini I, Rebecchi L, Jockusch Elizabeth L, Goldstein B. The compact body plan of tardigrades evolved by the loss of a large body region. Curr Biol. 2016;26:224–9.
PubMed Article CAS Google Scholar
63.
Ferrier DEK, Minguillón C. Evolution of the Hox/ParaHox gene clusters. Int J Dev Biol. 2003;47:605–11.
PubMed CAS Google Scholar
64.
Ferrier DEK, Holland PWH. Ciona intestinalis ParaHox genes: evolution of Hox/ParaHox cluster integrity, developmental mode, and temporal colinearity. Mol Phylogenet Evol. 2002;24:412–7.
PubMed Article CAS Google Scholar
65.
van Auken K, Weaver DC, Edgar LG, Wood WB. Caenorhabditis elegans embryonic axial patterning requires two recently discovered posterior-group Hox genes. Proc Natl Acad Sci USA. 2000;97:4499–503.
PubMed Article Google Scholar
66.
Yoshida Y, Koutsovoulos G, Laetsch DR, Stevens L, Kumar S, Horikawa DD, Ishino K, Komine S, Kunieda T, Tomita M, et al. Comparative genomics of the tardigrades Hypsibius dujardini and Ramazzottius varieornatus. PLoS Biol. 2017;15:e2002266.
PubMed PubMed Central Article CAS Google Scholar
67.
Gross V, Treffkorn S, Mayer G. Tardigrada. In: Wanninger A, editor. Evolutionary developmental biology of invertebrates 3: Ecdysozoa I: non-tetraconata. Wien: Springer; 2015. p. 35–52.
Google Scholar
68.
Janssen R, Budd GE. Deciphering the onychophoran ‘segmentation gene cascade’: gene expression reveals limited involvement of pair rule gene orthologs in segmentation, but a highly conserved segment polarity gene network. Dev Biol. 2013;382:224–34.
PubMed Article CAS Google Scholar
69.
Damen WGM. Evolutionary conservation and divergence of the segmentation process in arthropods. Dev Dyn. 2007;236:1379–91.
PubMed Article CAS Google Scholar
70.
Prud’homme B, de Rosa R, Arendt D, Julien JF, Pajaziti R, Dorresteijn AWC, Adoutte A, Wittbrodt J, Balavoine G. Arthropod-like expression patterns of engrailed and wingless in the annelid Platynereis dumerilii suggest a role in segment formation. Curr Biol. 2003;13:1876–81.
PubMed Article CAS Google Scholar
71.
Jagla K, Jagla T, Heitzler P, Dretzen G, Bellard F, Bellard M. ladybird, a tandem of homeobox genes that maintain late wingless expression in terminal and dorsal epidermis of the Drosophila embryo. Development. 1997;124:91–100.
PubMed CAS Google Scholar
72.
Master VA, Kourakis Matthew J, Martindale Mark Q. Isolation, characterization, and expression of Le-msx, a maternally expressed member of the msx gene family from the glossiphoniid leech, Helobdella. Dev Dyn. 1996;207:404–19.
PubMed Article CAS Google Scholar
73.
Nardelli-Haefliger D, Shankland M. Lox10, a member of the NK-2 homeobox gene class, is expressed in a segmental pattern in the endoderm and in the cephalic nervous system of the leech Helobdella. Development. 1993;118:877–92.
PubMed CAS Google Scholar
74.
Wheeler SR, Carrico ML, Wilson BA, Skeath JB. The Tribolium columnar genes reveal conservation and plasticity in neural precursor patterning along the embryonic dorsal–ventral axis. Dev Biol. 2005;279:491–500.
PubMed Article CAS Google Scholar
75.
Döffinger C, Stollewerk A. How can conserved gene expression allow for variation? Lessons from the dorso-ventral patterning gene muscle segment homeobox. Dev Biol. 2010;345:105–16.
PubMed Article CAS Google Scholar
76.
Nose A, Isshiki T, Takeichi M. Regional specification of muscle progenitors in Drosophila: the role of the msh homeobox gene. Development. 1998;125:215–23.
PubMed CAS Google Scholar
77.
Dohrmann C, Azpiazu N, Frasch M. A new Drosophila homeo box gene is expressed in mesodermal precursor cells of distinct muscles during embryogenesis. Genes Dev. 1990;4:2098–111.
PubMed Article CAS Google Scholar
78.
Knirr S, Azpiazu N, Frasch M. The role of the NK-homeobox gene slouch (S59) in somatic muscle patterning. Development. 1999;126:4525.
PubMed CAS Google Scholar
79.
Lord PCW, Lin M-H, Hales KH, Storti RV. Normal expression and the effects of ectopic expression of the Drosophila muscle segment homeobox (msh) gene suggest a role in differentiation and patterning of embryonic muscles. Dev Biol. 1995;171:627–40.
PubMed Article CAS Google Scholar
80.
Houzelstein D, Auda-Boucher G, Cheraud Y, Rouaud T, Blanc I, Tajbakhsh S, Buckingham ME, Fontaine-Perus J, Robert B. The homeobox gene Msx1 is expressed in a subset of somites, and in muscle progenitor cells migrating into the forelimb. Development. 1999;126:2689–701.
PubMed CAS Google Scholar
81.
Houzelstein D, Cheraud Y, Auda-Boucher G, Fontaine-Perus J, Robert B. The expression of the homeobox gene Msx1 reveals two populations of dermal progenitor cells originating from the somites. Development. 2000;127:2155–64.
PubMed CAS Google Scholar
82.
Bendall AJ, Ding J, Hu G, Shen MM, Abate-Shen C. Msx1 antagonizes the myogenic activity of Pax3 in migrating limb muscle precursors. Development. 1999;126:4965–76.
PubMed CAS Google Scholar
83.
Oliveira IS, Mayer G. Apodemes associated with limbs support serial homology of claws and jaws in Onychophora (velvet worms). J Morphol. 2013;274:1180–90.
Article Google Scholar
84.
Oliveira IS, Tait NN, Strübing I, Mayer G. The role of ventral and preventral organs as attachment sites for segmental limb muscles in Onychophora. Front Zool. 2013;10:1–18.
Article Google Scholar
85.
Pflugfelder O. Onychophora. In: Czihak G, editor. Grosses zoologisches praktikum, vol. 13a. Stuttgart: Gustav Fischer; 1968. p. 1–42.
Google Scholar
86.
Hoyle G, Williams M. The musculature of Peripatus and its innervation. Philos Trans R Soc Lond B Biol Sci. 1980;288:481–510.
Article Google Scholar
87.
Birket-Smith SJR. The anatomy of the body wall of Onychophora. Zool Jahrb Abt Anat Ontog Tiere. 1974;93:123–54.
Google Scholar
88.
Manton SM. The evolution of arthropodan locomotory mechanisms. Part 11: Habits, morphology and evolution of the Uniramia (Onychophora, Myriapoda and Hexapoda) and comparisons with the Arachnida, together with a functional review of uniramian musculature. Zool J Linn Soc. 1973;53:257–375.
Article Google Scholar
89.
Snodgrass RE. Evolution of the Annelida, Onychophora and Arthropoda. Smithson Misc Coll. 1938;97:1–159.
Google Scholar
90.
Müller M, de Sena Oliveira I, Allner S, Ferstl S, Bidola P, Mechlem K, Fehringer A, Hehn L, Dierolf M, Achterhold K, et al. Myoanatomy of the velvet worm leg revealed by laboratory-based nanofocus X-ray source tomography. Proc Natl Acad Sci USA. 2017;114:12378–83.
PubMed Article CAS Google Scholar
91.
Mayer G. Nephridial development in the Onychophora and its bearing on the Articulata hypothesis. Org Divers Evol. 2005;5:40.
Google Scholar
92.
Storch V, Ruhberg H, Alberti G. Zur Ultrastruktur der Segmentalorgane der Peripatopsidae (Onychophora). Zool Jahrb Abt Anat Ontog Tiere. 1978;100:47–63.
Google Scholar
93.
Mayer G. Origin and differentiation of nephridia in the Onychophora provide no support for the Articulata. Zoomorphology. 2006;125:1–12.
Article Google Scholar
94.
Mayer G, Whitington PM. Neural development in Onychophora (velvet worms) suggests a step-wise evolution of segmentation in the nervous system of Panarthropoda. Dev Biol. 2009;335:263–75.
PubMed Article CAS Google Scholar
95.
Sedgwick A. The development of the Cape species of Peripatus. Part IV. The changes from stage G to birth. Q J Microsc Sci. 1888;28:373–96.
Google Scholar
96.
Sedgwick A. The development of the Cape species of Peripatus. Part III. On the changes from stage A to stage F. Q J Microsc Sci. 1887;27:467–550.
Google Scholar
97.
von Kennel J. Entwicklungsgeschichte von Peripatus edwardsii Blanch. und Peripatus torquatus n. sp. II. Theil. Arb Zool-Zootom Inst Würzburg. 1888;8:1–93.
Google Scholar
98.
Janssen R, Jörgensen M, Prpic NM, Budd Graham E. Aspects of dorso-ventral and proximo-distal limb patterning in onychophorans. Evol Dev. 2015;17:21–33.
PubMed Article CAS Google Scholar
99.
Schmidt-Rhaesa A. Tardigrades—are they really miniaturized dwarfs? Zool Anz. 2001;240:549–55.
Article Google Scholar
100.
Lints TJ, Parsons LM, Hartley L, Lyons I, Harvey RP. Nkx-2.5: a novel murine homeobox gene expressed in early heart progenitor cells and their myogenic descendants. Development. 1993;119:419.
PubMed CAS Google Scholar
101.
Evans SM, Yan W, Murillo MP, Ponce J, Papalopulu N. Tinman, a Drosophila homeobox gene required for heart and visceral mesoderm specification, may be represented by a family of genes in vertebrates: XNkx-2.3, a second vertebrate homologue of tinman. Development. 1995;121:3889–99.
PubMed CAS Google Scholar
102.
Wang W, Razy-Krajka F, Siu E, Ketcham A, Christiaen L. NK4 antagonizes Tbx1/10 to promote cardiac versus pharyngeal muscle fate in the ascidian second heart field. PLoS Biol. 2013;11:e1001725.
PubMed PubMed Central Article CAS Google Scholar
103.
Hanken J, Wake DB. Miniaturization of body size: organismal consequences and evolutionary significance. Annu Rev Ecol Syst. 1993;24:501–19.
Article Google Scholar
104.
Navet S, Bassaglia Y, Baratte S, Martin M, Bonnaud L. Somatic muscle development in Sepia officinalis (Cephalopoda–Mollusca): a new role for NK4. Dev Dyn. 2008;237:1944–51.
PubMed Article CAS Google Scholar
105.
Ramos C, Robert B. msh/Msx gene family in neural development. Trends Genet. 2005;21:624–32.
PubMed Article CAS Google Scholar
106.
Arendt D. Animal evolution: convergent nerve cords? Curr Biol. 2018;28:R225–7.
PubMed Article CAS Google Scholar
107.
Eriksson BJ, Samadi L, Schmid A. The expression pattern of the genes engrailed, pax6, otd and six3 with special respect to head and eye development in Euperipatoides kanangrensis Reid 1996 (Onychophora: Peripatopsidae). Dev Genes Evol. 2013;223:237–46.
PubMed PubMed Central Article Google Scholar
108.
Martin C, Gross V, Pflüger H-J, Stevenson PA, Mayer G. Assessing segmental versus non-segmental features in the ventral nervous system of onychophorans (velvet worms). BMC Evol Biol. 2017;17:3.
PubMed PubMed Central Article CAS Google Scholar
109.
Mayer G, Harzsch S. Immunolocalization of serotonin in Onychophora argues against segmental ganglia being an ancestral feature of arthropods. BMC Evol Biol. 2007;7:118.
PubMed PubMed Central Article CAS Google Scholar
110.
Mayer G. Onychophora. In: Schmidt-Rhaesa A, Harzsch S, Purschke G, editors. Structure and evolution of invertebrate nervous systems. Oxford: Oxford University Press; 2016. p. 390–401.
Google Scholar
111.
Whitington PM, Mayer G. The origins of the arthropod nervous system: insights from the Onychophora. Arthropod Struct Dev. 2011;40:193–209.
PubMed Article Google Scholar
112.
Mayer G, Kato C, Quast B, Chisholm RH, Landman KA, Quinn LM. Growth patterns in Onychophora (velvet worms): lack of a localised posterior proliferation zone. BMC Evol Biol. 2010;10:1–12.
Article CAS Google Scholar
113.
Mayer G, Whitington PM. Velvet worm development links myriapods with chelicerates. Proc R Soc B Biol Sci. 2009;276:3571–9.
Article Google Scholar
114.
Eriksson BJ, Stollewerk A. Expression patterns of neural genes in Euperipatoides kanangrensis suggest divergent evolution of onychophoran and euarthropod neurogenesis. Proc Natl Acad Sci USA. 2010;107:22576–81.
PubMed Article Google Scholar
115.
Eriksson BJ, Stollewerk A. The morphological and molecular processes of onychophoran brain development show unique features that are neither comparable to insects nor to chelicerates. Arthropod Struct Dev. 2010;39:478–90.
PubMed Article CAS Google Scholar
116.
Mayer G, Harzsch S. Distribution of serotonin in the trunk of Metaperipatus blainvillei (Onychophora, Peripatopsidae): implications for the evolution of the nervous system in Arthropoda. J Comp Neurol. 2008;507:1196–208.
PubMed Article Google Scholar
117.
Reid AL. Review of the Peripatopsidae (Onychophora) in Australia, with comments on peripatopsid relationships. Invertebr Taxon. 1996;10:663–936.
Article Google Scholar
118.
Baer A, Mayer G. Comparative anatomy of slime glands in Onychophora (velvet worms). J Morphol. 2012;273:1079–88.
PubMed Article Google Scholar
119.
Robson EA, Lockwood APM, Ralph R. Composition of the blood in Onychophora. Nature. 1966;209:533.
Article Google Scholar
120.
Treffkorn S, Mayer G. Expression of the decapentaplegic ortholog in embryos of the onychophoran Euperipatoides rowelli. Gene Expr Patterns. 2013;13:384–94.
PubMed Article CAS Google Scholar
121.
Hering L, Henze MJ, Kohler M, Bleidorn C, Leschke M, Nickel B, Meyer M, Kircher M, Sunnucks P, Mayer G. Opsins in Onychophora (velvet worms) suggest a single origin and subsequent diversification of visual pigments in arthropods. Mol Biol Evol. 2012;29:145.
Article CAS Google Scholar
122.
Hering L, Mayer G. Analysis of the Opsin repertoire in the tardigrade Hypsibius dujardini provides insights into the evolution of Opsin genes in Panarthropoda. Genome Biol Evol. 2014;6:2380–91.
PubMed PubMed Central Article CAS Google Scholar
123.
Boothby TC, Tenlen JR, Smith FW, Wang JR, Patanella KA, Osborne Nishimura E, Tintori SC, Li Q, Jones CD, Yandell M, et al. Evidence for extensive horizontal gene transfer from the draft genome of a tardigrade. Proc Natl Acad Sci USA. 2015;112:15976–81.
PubMed Article CAS Google Scholar
124.
Gross V, Mayer G. Neural development in the tardigrade Hypsibius dujardini based on anti-acetylated α-tubulin immunolabeling. EvoDevo. 2015;6:12.
PubMed PubMed Central Article CAS Google Scholar
125.
Gross V, Minich I, Mayer G. External morphogenesis of the tardigrade Hypsibius dujardini as revealed by scanning electron microscopy. J Morphol. 2017;278:563–73.
PubMed Article Google Scholar
126.
Gross V, Bährle R, Mayer G. Detection of cell proliferation in adults of the water bear Hypsibius dujardini (Tardigrada) via incorporation of a thymidine analog. Tissue Cell. 2018;51:77–83.
PubMed Article CAS Google Scholar
127.
Gąsiorek P, Stec D, Morek W, Michalczyk Ł. An integrative redescription of Hypsibius dujardini (Doyère, 1840), the nominal taxon for Hypsibioidea (Tardigrada: Eutardigrada). Zootaxa. 2018;4415:45–75.
Article Google Scholar
128.
Altschul SF, Madden TL, Schäffer AA, Zhang J, Zhang Z, Miller W, Lipman DJ. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 1997;25:3389–402.
PubMed PubMed Central Article CAS Google Scholar
129.
Oliveira M, Liedholm S, Lopez J, Lochte A, Pazio M, Martin J, Mörch P, Salakka S, York J, Yoshimoto A, Janssen R. Expression of arthropod distal limb-patterning genes in the onychophoran Euperipatoides kanangrensis. Dev Genes Evol. 2014;224:1–10.
Article Google Scholar
130.
Zhong Y-F, Butts T, Holland PWH. HomeoDB: a database of homeobox gene diversity. Evol Dev. 2008;10:516–8.
PubMed Article CAS Google Scholar
131.
Y-f Zhong. Holland PWH: homeoDB2: functional expansion of a comparative homeobox gene database for evolutionary developmental biology. Evol Dev. 2011;13:567–8.
Article Google Scholar
132.
Schwager EE, Sharma PP, Clarke T, Leite DJ, Wierschin T, Pechmann M, Akiyama-Oda Y, Esposito L, Bechsgaard J, Bilde T, et al. The house spider genome reveals an ancient whole-genome duplication during arachnid evolution. BMC Biol. 2017;15:62.
PubMed PubMed Central Article Google Scholar
133.
Colbourne JK, Pfrender ME, Gilbert D, Thomas WK, Tucker A, Oakley TH, Tokishita S, Aerts A, Arnold GJ, Basu MK, et al. The ecoresponsive genome of Daphnia pulex. Science. 2011;331:555–61.
PubMed PubMed Central Article CAS Google Scholar
134.
Katoh K, Standley DM. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol Biol Evol. 2013;30:772–80.
PubMed PubMed Central Article CAS Google Scholar
135.
Stamatakis A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics. 2014;30:1312–3.
PubMed PubMed Central Article CAS Google Scholar
136.
Hoffmann S, Otto C, Kurtz S, Sharma CM, Khaitovich P, Vogel J, Stadler PF, Hackermüller J. Fast mapping of short sequences with mismatches, insertions and deletions using index structures. PLoS Comput Biol. 2009;5:e1000502.
PubMed PubMed Central Article CAS Google Scholar
137.
Ryan JF, Burton PM, Mazza ME, Kwong GK, Mullikin JC, Finnerty JR. The cnidarian-bilaterian ancestor possessed at least 56 homeoboxes: evidence from the starlet sea anemone, Nematostella vectensis. Genome Biol. 2006;7:R64.
PubMed PubMed Central Article CAS Google Scholar
138.
Franke FA, Mayer G. Expression study of the hunchback ortholog in embryos of the onychophoran Euperipatoides rowelli. Dev Genes Evol. 2015;225:207–19.
PubMed Article Google Scholar
139.
Janssen R, Eriksson B, Tait N, Budd G. Onychophoran Hox genes and the evolution of arthropod Hox gene expression. Front Zool. 2014;11:22.
PubMed PubMed Central Article CAS Google Scholar
140.
Janssen R, Damen WGM, Budd GE. Expression of collier in the premandibular segment of myriapods: support for the traditional Atelocerata concept or a case of convergence? BMC Evol Biol. 2011;11:50.
PubMed PubMed Central Article Google Scholar
141.
Martin C, Mayer G. Neuronal tracing of oral nerves in a velvet worm—implications for the evolution of the ecdysozoan brain. Front Neuroanat. 2014;8:7.
PubMed PubMed Central Article Google Scholar

Download references

Authors’ contributions

GM and ST designed the research. ST conducted the experiments, assembled the NK gene alignment and wrote the first draft of the manuscript. LK assisted with in situ hybridization of NK4 and Tlx. LH conducted the phylogenetic and mapping analysis and helped analyzing the phylogenetic data. All authors read and approved the final manuscript.

Acknowledgements

We are thankful to members of the Mayer laboratory for their support with animal husbandry and to Franziska Anni Franke for her help with embryo dissection. We gratefully acknowledge Dave M. Rowell, Ivo de Sena Oliveira, Franziska Anni Franke and Michael Gerth for their assistance with collecting the specimens, Noel N. Tait for his help with permits, and Vladimir Gross for proofreading the article. The staffs of the National Parks & Wildlife Service New South Wales (Australia) and the Department of Sustainability, Environment, Water, Population and Communities (Australia) are gratefully acknowledged for providing the collection and export permits. The Open Access Office of the University of Kassel is thanked for financial support.

Competing interests

The authors declare that they have no competing interests.

Availability of data and materials

All data and material underlying the current study are publicly available or included in the additional files.

Consent for publication

Not applicable.

Ethics approval and consent to participate

Not applicable.

Funding

This study was supported by the Emmy Noether Programme of the German Research Foundation (DFG: Ma 4147/3-1) to GM.

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Author information

Affiliations

Department of Zoology, Institute of Biology, University of Kassel, Heinrich-Plett-Str. 40, 34132, Kassel, Germany
Sandra Treffkorn, Laura Kahnke, Lars Hering & Georg Mayer

Authors

Sandra Treffkorn
View author publications
You can also search for this author in PubMed Google Scholar
Laura Kahnke
View author publications
You can also search for this author in PubMed Google Scholar
Lars Hering
View author publications
You can also search for this author in PubMed Google Scholar
Georg Mayer
View author publications
You can also search for this author in PubMed Google Scholar

Corresponding author

Correspondence to Sandra Treffkorn.

Additional files

Additional file 1.

NK (blue) and NKL (orange) gene complements of different bilaterian species. Numbers indicate the number of genes, dashes indicate the absence thereof, numbers in brackets indicate the number of pseudogenes, question marks indicate missing data. The gene complements were retrieved from publicly available data as well as the sources specified below the table.

Additional file 2.

Comparative transcriptomic analysis of NK genes in the onychophoran E. rowelli showing the average number of matched nucleotides per position and GB (upper table) of all studied NK genes from different transcriptome libraries of partially pooled embryonic stages, and the relative expression level in comparison to the abundance of ribosomal protein RPL31 transcripts (lower table) obtained from the short read mapping analysis.

Additional file 3.

NK6.2 expression at consecutive developmental stages in embryos of the onychophoran E. rowelli. Developing limbs are numbered. Anterior is left in all images. A Stage II embryo in ventrolateral view. B Proctodeum of a stage II embryo in ventral view. Arrowheads indicate the expression around the proctodeum. C Stage III embryo in lateral view. Arrow indicates the decreasing intensity of the signal. D Developing jaw, slime papilla and first leg of a stage III embryo. Signals in the ventral ectodermal thickenings and ventral nerve cords in are indicated with asterisks and arrowheads, respectively. E Stage V embryo in ventral view. A weak expression is visible in the ventral nerve cords (arrowheads). Abbreviations: at, developing antenna; cl, cephalic lobe; de, dorsal extra-embryonic tissue; jw, developing jaw; js, jaw segment; po, proctodeum; sp, developing slime papilla; ss, slime papilla segment; ve, ventral extra-embryonic tissue. Scale bars: A, C: 200 µm; B, D: 100 µm: E: 500 µm.

Additional file 4.

Cross sections of stage IV embryos of the onychophoran E. rowelli showing the expression of NK6.1 (A, A’, B, B’), Msx (C, C’) and NK2.2 (D, D’) in the ventral ectoderm (empty arrowheads) and developing nerve cords (filled arrowheads). Developing nerve cord is indicated with dashed lines, developing neuropil is indicated with asterisks. Scale bars: A–C’: 50 µm; D, D’: 100 µm.

Additional file 5.

Alignment of the 382 NK homeodomains of various metazoan taxa that was used for the maximum-likelihood analysis.

Additional file 6.

Maximum-likelihood analysis of the NK cluster and NKL genes among metazoans. The ML phylogeny was computed using the Pthreads-SSE3 version of RAxML v8.2.10.

Rights and permissions

Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. 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.

Reprints and Permissions

About this article

Cite this article

Treffkorn, S., Kahnke, L., Hering, L. et al. Expression of NK cluster genes in the onychophoran Euperipatoides rowelli: implications for the evolution of NK family genes in nephrozoans. EvoDevo 9, 17 (2018). https://doi.org/10.1186/s13227-018-0105-2

Download citation

Received: 28 February 2018
Accepted: 06 July 2018
Published: 18 July 2018
DOI: https://doi.org/10.1186/s13227-018-0105-2

Keywords

Homeobox genes
NK genes
NK-linked genes
Gene expression
Velvet worms
Onychophora
Mesoderm development
Nephrozoa
Urbilaterian

Expression of NK cluster genes in the onychophoran Euperipatoides rowelli: implications for the evolution of NK family genes in nephrozoans

Abstract

Background

Results

Conclusion

Background

Results

Identification and analysis of NK and NKL homologs in onychophorans and tardigrades

NK/NKL repertoire in Onychophora

NK/NKL repertoire in Tardigrada

Expression of NK cluster genes in embryos of the onychophoran E. rowelli

Transcription levels of NK cluster genes and the NKL gene \(\text{NK2.2}\)

Expression of \(\text{NK1}\)

Expression of \(\text{NK3}\)

Expression of \(\text{NK4}\)

Expression of \(\text{NK5}\)

Expression of \(\text{NK6.1}\) and \(\text{NK6.2}\)

Expression of \(\text{Msx}\)

Expression of \(\text{Lbx}\)

Expression of \(\text{Tlx}\)

Expression of \(\text{NK2.2}\)

Discussion

NK and NKL gene repertoires in Onychophora and Tardigrada

Segment polarity-like expression patterns of NK1, Lbx and Msx in the onychophoran E. rowelli

Conserved expression patterns of NK cluster genes in the mesoderm of E. rowelli

Non-regionalized neural expression of NK cluster genes and the NKL gene NK2.2 in E. rowelli supports convergent evolution of bilaterian nerve cords

Conclusion

Methods

Specimens and sample preparation

Identification of NK homologs

Sequence alignment, phylogenetic analysis and short read mapping

Nomenclature

RNA isolation, amplification of gene fragments, molecular cloning, probe preparation and whole-mount in situ hybridization

Microscopy and image processing

References

Authors’ contributions

Acknowledgements

Competing interests

Availability of data and materials

Consent for publication

Ethics approval and consent to participate

Funding

Publisher’s Note

Author information

Affiliations

Corresponding author

Additional files

Additional file 1.

Additional file 2.

Additional file 3.

Additional file 4.

Additional file 5.

Additional file 6.

Rights and permissions

About this article

Cite this article

Share this article

Keywords

EvoDevo

Contact us