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PLoS Biol. Nov 2008; 6(11): e278.
Published online Nov 18, 2008. doi:  10.1371/journal.pbio.0060278
PMCID: PMC2586386

A Novel Gene Family Controls Species-Specific Morphological Traits in Hydra

Nipam Patel, Academic Editor

Abstract

Understanding the molecular events that underlie the evolution of morphological diversity is a major challenge in biology. Here, to identify genes whose expression correlates with species-specific morphologies, we compared transcriptomes of two closely related Hydra species. We find that species-specific differences in tentacle formation correlate with expression of a taxonomically restricted gene encoding a small secreted protein. We show that gain of function induces changes in morphology that mirror the phenotypic differences observed between species. These results suggest that “novel” genes may be involved in the generation of species-specific morphological traits.

Author Summary

Closely related animal species share most of their genes, and only minor morphological differences allow us to tell them apart. The genetic basis for these differences may involve minor changes in the spatial and temporal activity of transcription factors—“regulator” genes—which are surprisingly conserved throughout the animal kingdom. However, every group of animals also has a small proportion of genes that are extremely variable among closely related species or even unique. Such genes are referred to as “novel,” “orphan,” or “taxonomically restricted.” Their functions and origins are often obscure. We have found that a family of novel genes is responsible for morphological differences between two closely related species of fresh water polyps called Hydra. A secreted protein encoded by a novel gene regulates the way in which tentacles develop. Our data indicate that novel genes may play a role in the creation of novel morphological features, thus representing one way how evolution works at the genus level. Appearance of novel genes may reflect evolutionary processes that allow animals to adapt in the best way to changing environmental conditions and new habitats.

Introduction

Understanding the molecular events that underlie the evolution of morphological diversity is a major challenge in biology. Comparative studies have demonstrated that morphological evolution occurs through alterations in development [15]. Recent advances in genome sequencing have shown that all eumetazoan animals, from cnidarians to humans, share a handful of highly conserved signal transduction pathways that, together with several hundred conserved transcription factors, make up a molecular toolkit common for all living beings [6,7]. It has been proposed that morphological traits are altered in evolution when these conserved components obtain a novel spatiotemporal mode of expression [8]. For example, pigmentation patterns in insects are thought to evolve by modifications in the cis-regulatory elements of pigmentation genes [911]. In finches, a relationship has been revealed between species-specific beak morphology and the spatiotemporal expression of calmodulin and bone morphogenetic protein 4 (BMP4) [12,13]. Supporting that observation, heterochronic manipulations of BMP4 expression during chick development can reproduce the different patterns observed among Darwin's finches [14]. Rewiring of regulatory networks is the equally well source for morphological changes on a large scale as well as for inter-specific or even intra-specific variations [15].

There is, however, one much less appreciated source for the creation of morphological novelties. All genome and expressed sequence tag (EST) projects to date in every taxonomic group studied so far have uncovered a substantial fraction of genes that are without known homologs [1618]. These “orphans” or “taxonomically restricted genes” (TRGs) are defined as being exclusively restricted to a particular taxonomic group [19]. For example, analysis of the phylum Nematoda has identified more than 20% of genes that were nematode-unique TRGs [20]. The draft genome of Ciona intestinalis revealed [21] that nearly one-fifth of the genes were orphans. A comparison between the genome sequences of Schizosaccharomyces pombe and Saccharomyces cerevisiae showed [22] about 14% of the predicted proteins to be unique to Sc. pombe and 19% unique to Sa. cerevisiae. In Drosophila, TRGs include indispensable regulators of development such as bicoid [23] and spätzle [24]. Recent comparative data on the genomes of 12 Drosophila species revealed that about 2.5% of genes are not present outside of the genus Drosophila and, therefore, have most likely arisen de novo [25]. An even larger proportion of lineage-specific genes have been detected in the genome of Tribolium [26]. In bacteria, the cumulative number of orphans identified does not appear to be leveling off, although hundreds of complete genome sequences have been already analyzed [18,19]. Models for evolution of TRGs have been proposed [27] and the significance of their evolutionary contribution to ecological adaptation has been postulated [19]. Despite this, TRGs are poorly studied and little understood, in large part because the lack of homology confounds attempts to determine the putative function of the protein.

Thus, although progress has been made towards understanding the molecular mechanisms controlling the evolution of morphology, many questions remain to be addressed: What genes are differentially expressed in two closely related species and how many? Do the differences occur predominantly in structural or regulatory genes? Do novel genes play a role in generation of morphological novelties? And, ultimately, what is the genetic basis of species-specific morphologies?

Here, we address these fundamental questions in a study designed to identify the transcriptional signatures of two closely related, yet morphologically distinct species of the basal metazoan Hydra. Freshwater polyps of the genus Hydra belong to one of the earliest branches in the animal tree of life, the Cnidaria (Figure 1A), and they represent the simplest animals at the tissue grade of organization. The body plan consists of only two cell layers and a limited number of cell types derived from three distinct stem cell lineages [28]. A head and tentacles are on one end and a foot on the opposite end of a single body axis. Tentacles are used for prey capture and are armed with several types of nematocytes. Proliferation occurs mostly asexually by budding, a process during which a secondary polyp forms in the lower gastric region of the parental animal. Morphological simplicity in cnidarians is accompanied by unexpected molecular complexity. With the continuously growing EST and genomic databases from cnidarians, evidence is accumulating for extensive conservation in gene content, structure and organization between cnidarians and vertebrates [6,7,2931]. At the same time, systematic approaches to isolate morphogenetically active molecules have identified numerous peptides in Hydra that are encoded by novel genes [32].

Figure 1
Variation in Morphological Traits among Different Hydra Species

Here we show that in Hydra, a family of novel genes, defined as genus-level TRGs, plays a significant role in controlling phenotypic features that are referred to as species-specific traits. Our data show that morphological diversity at the genus level can be generated through changes in the spatial and temporal deployment of genes that are not highly conserved across long evolutionary distances. We also propose that losses and duplications of those novel genes among closely related species may be one of the driving forces leading to morphological diversification in the genus Hydra.

Results

Species-Specific Traits in the Genus Hydra

Hydra species differ in morphology, development, physiology, and ecology [33,34]. As shown in Figure 1B–1F, Hydra species differ in the presence or absence of symbionts, in carotenoid metabolism, and in the presence or absence of a conspicuous stalk structure (labeled with an asterisk in Figure 1C). Hydra species vary in genome sizes, ranging from 380 to 1,450 Mbp. Hydra species also differ in the shapes and sizes of their nematocysts. For example, as schematically shown in Figure 1F, in Hydra oligactis in contrast to other Hydra species the inverted tube of holotrichous isorhiza is coiled irregularly. Other species-specific traits include a ring of neurons running circumferentially around the hypostome in H. oligactis but not in other Hydra species [35]. Hydra species can also be identified by the order in which tentacles arise on young buds. Whereas the mean number of tentacles per polyp does not differ in H. oligactis and H. magnipapillata (Figure 1G and and1H),1H), in H. oligactis two lateral tentacles arise early and are conspicuously longer during bud development (Figure 1I–1M) in contrast to H. magnipapillata and H. vulgaris, where tentacles arise synchronously and are all of the same length (Figure 1N–1R).

Identification of TRGs in H. oligactis and H. magnipapillata

To gain insight into the molecular mechanisms controlling phenotypic differences in the genus Hydra, we compared the transcriptomes of H. oligactis and H. magnipapillata by suppression subtractive hybridization (SSH) of complimentary DNAs (Figure 2A and and2B).2B). The goal of SSH was to identify quickly evolving genes that are present (and transcriptionally active) in one species and absent (or transcriptionally silent) in the other, as well as to uncover homologous genes that are expressed in both species but have considerably diverged sequences. As shown schematically in Figure 2A and and2B,2B, two subtractive hybridizations were performed using cDNA pools from H. oligactis and H. magnipapillata polyps, resulting in two cDNA libraries enriched for species-specific transcripts. As expected, a considerable proportion of clusters and singletons in each of the library had no homologs in the NCBI protein database (Figure 2A and and2B),2B), representing, therefore, genes putatively restricted to the genus Hydra (for clustering and sequence analysis see Tables S1S3 and Figure S1). The expression of these genes was analyzed by in situ hybridization. Genes expressed in taxonomically informative structures (tentacles, nematocytes, and stalk; see Figure S1) were selected for further analysis. Here, we focus on genes represented by clusters CL223 and CL87. Both genes show no similarity to any gene outside the genus Hydra. Both genes, however, are weakly similar (see Tables S1 and S2) to the Hym301 protein, which was identified earlier in a systematic approach to isolate morphogenetically active peptides in H. magnipapillata [36]. Interestingly, as shown in Figure 2C and and2D,2D, in the two species analyzed these Hym301-like genes have complementary expression patterns: whereas in H. oligactis expression is restricted to tentacles (Figure 2C), in H. magnipapillata Hym301-like transcripts are absent in tentacle tissue but abundant throughout the body column (Figure 2D). Because tentacle development is one of the major morphogenetic processes in Hydra, differs between H. oligactis and H. magnipapillata (Figure 1M and and1R),1R), and is under strong selective constraints as tentacles are the only structures by which polyps can take up food, we focused on the identified Hym301-like genes to investigate their function.

Figure 2
Identification of Species-Specific Genes in Hydra by SSH

Characterization of the TRG Family Hym301

Detailed analysis of the Hym301-like genes CL223 (in H. oligactis) and CL87 (in H. magnipapillata) revealed that they belong to a gene family that exhibits variation in gene number and expression patterns among Hydra species (Figure 3). In H. magnipapillata we discovered four CL223-related genes (see Materials and Methods for detail). Because one of them has been identified earlier [36] and was described as a gene Hym301 in H. magnipapillata [37], we named the four genes mHym301A to mHym301D. In H. magnipapillata, the genes mHym301A (Figure 3A), mHym301C (Figure 3C), and mHym301D (Figure 3D) are expressed in the head, whereas mHym301B (Figure 3B) is expressed along the body column. None of the four H. magnipapillata genes is expressed in the tentacles. In contrast, H. oligactis has only two Hym301-related genes, oHym301A and oHym301B, with both genes expressed in tentacles (Figure 3E and and3F).3F). In H. vulgaris AEP, only one gene is present (aepHym301A) and expressed in the basis of tentacles (Figure 3G). Expression of aepHym301A is considerably weaker than expression of the Hym301-like genes in the other two species (Figure 3I).

Figure 3
Analysis of the Taxonomically Restricted Gene Family Hym301 in Different Hydra Species

A phylogenetic tree based on the amino acid sequence alignment (Figure 3J) places the members of Hym301 gene family in three distinct groups, which fits with the phylogenetic tree of the genus Hydra [34] (see Figure 1A). Predicted proteins from H. oligactis and H. magnipapillata form two distinct clades, whereas the single protein from H. vulgaris AEP forms a separate clade (Figure 3J). The phylogeny clearly indicates that gene duplications occurred separately in the H. oligactis and H. magnipapillata lineages. In H. magnipapillata the available draft genome revealed that all four genes share common genomic organization, with four exons (Figure 3H). Interestingly, mHym301C and mHym301D are clustered in the genome within 70 kb (Figure 3H), suggesting a recent duplication event. Amino acid alignment (Figure 3K) indicates that all members of the gene family encode short proteins (87–106 amino acids) with a putative signal peptide sequence at the N terminus followed by a relatively conserved region. The C-terminal part of the protein is highly diverged, not only among species, but also among the different Hym301 proteins in H. oligactis and H. magnipapillata. In mHym301A the C terminus harbors the peptide (marked in orange in Figure 3K) reported previously [37] to affect tentacle formation in H. magnipapillata. According to the sequence information available, the genes of the Hym301 family have no homologs in eukaryotic organisms outside the genus Hydra. They are also not present in the genome of Nematostella, which, as a representative of the class Anthozoa, occupies a more basal position in the phylogenetic tree than Hydra. Therefore, we consider the members of the Hym301 family as TRGs, which have most likely originated within the class Hydrozoa and expanded in the genus Hydra.

Generation of Transgenic Hydra Overexpressing mHym301A

To explore the biological significance of the Hym301 gene family in vivo, we produced transgenic polyps that overexpress mHym301A in all ectodermal epithelial cells. As shown in Figure 4, we introduced the H. magnipapillata mHym301A gene into polyps of the closely related species H. vulgaris AEP, in which the expression level of the endogenous gene (aepHym301A) is low (see Figure 3G and and3I).3I). Hydra embryos were microinjected (see Materials and Methods for details) with an expression construct in which mHym301A including the sequence encoding the signal peptide was fused in frame to enhanced green fluorescent protein (eGFP) and under the control of the Hydra β-actin promoter (Figure 4A). In transgenic animals producing mHym301A:eGFP, secretion of the fusion protein was detectable during embryonic development in the egg shell of the embryo starting 3 d after injection (Figure 4B and and4C).4C). After hatching, fully transgenic animals producing mHym301A:eGFP in all their ectodermal epithelial cells were generated by asexual propagation of founder polyps (Figure 4D–4F). Figure 4G indicates that, in nontransgenic H. magnipapillata polyps, mHym301A transcripts are restricted to the head tissue and absent in tentacle cells. Transgenic line H. vulgaris AEPA14 was used in all further experiments. As a control, a transgenic line was produced (Ecto-1) using the same construct without the sequences encoding signal peptide and the mHym301A sequence. As shown in Figure 4I, the mHym301A:eGFP fusion protein is visually detectable by fluorescence microscopy in transgenic animals along the whole body column and in the tentacles. Fluorescence, however, is much weaker than in the control Ecto-1 line (Figure 4H). Confocal laser scanning microscopy shows (Fig 4J–4L) the presence of the mHym301A:eGFP fusion protein in secretory vesicles of ectodermal epithelial cells. In contrast, the eGFP protein in control Ecto-1 animals is not secreted and resides in the cytoplasm of epithelial cells (Figure S2). Thus, by a gain-of-function approach we generated H. vulgaris AEP transgenic animals that strongly overexpress the H. magnipapillata mHym301A gene in all ectodermal epithelial cells, including the tentacle cells. In these transgenic H. vulgaris AEP animals, the level of Hym301 expression is not only greatly enhanced compared with wild-type H. vulgaris AEP, but expression also extends to the tentacles—thereby mimicking oHym301A transcript localization in H. oligactis.

Figure 4
Transgenic H. vulgaris AEP Overexpressing mHym301A

Do these perturbations in Hym301 expression affect morphological traits of transgenic H. vulgaris AEPA14 polyps? To address this issue, we compared tentacle development in transgenic animals with that in wild-type H. vulgaris AEP, transgenic H. vulgaris AEP line Ecto-1, and wild-type H. magnipapillata and H. oligactis.

mHym301A Overexpression Affects Both the Speed of Tentacle Regeneration and the Pattern in Which Tentacles Arise

When a hydra's hypostome and tentacles are removed, the animal promptly develops new ones. “Head regeneration” is a morphallactic process involving the reorganization of existing tissue; it follows species-specific rules, with tentacles arising synchronously in H. magnipapillata and H. vulgaris but asynchronously in H. oligactis. Intact transgenic H. vulgaris AEPA14 animals exhibit normal morphology without any obvious disturbances (Figure 4I). The number of tentacles seems equal in nontransgenic H. vulgaris AEPwt, transgenic H. vulgaris AEPA14, and control transgenic line Ecto-1 animals (Figure 5A; see also Materials and Methods). Differences, however, become obvious when a polyp is bisected in the body column (Figure 5B) and a head regenerates at the apical end of the lower half within 96 h. Figure 5C indicates that in H. vulgaris AEPA14, tentacles begin to appear much earlier and in larger numbers than in nontransgenic H. vulgaris AEPwt or control transgenic line Ecto-1 (Table S4). Forty-two h after bisection, transgenic H. vulgaris AEPA14 had 4.2 ± 1.5 or 3.6 ± 1.61 tentacles compared with only 1.71 ± 1.24 in nontransgenic H. vulgaris AEPwt or 1.6 ± 1.64 in the transgenic Ecto-1 strain (Figure 5C). Figure 5D shows a representative transgenic animal, with well developed tentacles 42 h after decapitation; in the nontransgenic control (Figure 5I), tentacle formation is just at its beginning. By 66 h after decapitation, the mean number of tentacles in transgenic H. vulgaris AEPA14 was observed (Figure 5C) to vary from 5.6 ± 1.52 to 5.8 ± 1.44 compared with 3.68 ±1.51 in nontransgenic H. vulgaris AEPwt and 3.0 ± 1.64 in the control Ecto-1 line. The difference in tentacle number was maintained until 130 h after bisection (7.2 ± 1.3 in H. vulgaris AEPA14 versus 6.1 ± 1.05 and 5.7 ± 1.31 tentacles, respectively, in nontransgenic and Ecto-1 line controls).

Figure 5
Overexpression of mHym301A Alters Timing and Order of Tentacle Development during Regeneration

When examining regeneration in nontransgenic H. vulgaris AEPwt and transgenic H. vulgaris AEPA14, we noticed differences not only in the time kinetics of tentacle development, but also in the order in which tentacles arise on the regenerating tip (Figure 5D–5M). Unexpectedly, in transgenic H. vulgaris AEPA14 (Figure 5D–5F), in contrast to nontransgenic H. vulgaris AEPwt (Figure 5I–5K), tentacles arise asynchronously with two tentacles developing first. This mode of tentacle formation is similar to the order in which tentacles arise in regenerating H. oligactis polyps (Figure 5N–5P) and different from the mode of tentacle formation in the closely related species H. magnipapillata (Figure 5S–5U). Thus, overexpression of mHym301A in H. vulgaris AEP body column and tentacle cells (Figure 4I) causes reorganization of the order in which tentacles appear, from the H. vulgaris AEP pattern towards the H. oligactis-specific pattern. Given that in transgenic H. vulgaris AEPA14 as in H. oligactis polyps, mHym301A protein is present in tentacle cells (Figures 4I, I,5H5H and and5R),5R), these observations suggest that Hym301 genes are involved in determining the symmetry of tentacle formation.

To lend further credibility to this hypothesis, we next examined tentacle development during budding, a developmental process by which de novo head formation occurs in an adult hydra (Figure 6). As shown in Figure 6A–6D, in nontransgenic H. vulgaris AEPwt the pattern of tentacle formation is similar to that observed in H. magnipapillata (see Figure 1N–1Q) and different from that in H. oligactis (see Figure 1I–1L). In H. vulgaris AEPwt, five tentacle buds appear almost synchronously at the earliest stages of bud formation (arrowheads in Figure 6A) in a regular pentagonal pattern. In contrast, in transgenic H. vulgaris AEPA14 overexpressing mHym301A (Figure 6F–6I), the tentacles arise in a significantly larger number than in nontransgenic controls and in a slightly staggered sequence. As depicted in Figure 6G and and6H,6H, transgenic H. vulgaris AEPA14 develop six to seven tentacle buds in a rather irregular pattern, with some of the tentacles arising even outside the head in the upper part of the body column (open arrowhead in Figure 6H). Given that in adult polyps the number of tentacles is similar (varying from five to eight; Figure 5A) in nontransgenic and transgenic H. vulgaris AEP, the full set of tentacles appears to be established early in transgenic polyps and late (after detachment from the mother polyp) in nontransgenic controls. Taken together, these results substantiate the view that mHym301A is a molecule that affects the timing and order in which tentacles arise in Hydra.

Figure 6
Overexpression of mHym301A Alters Tentacle Development during Budding and Mimics a Mutant Phenotype of H. magnipapillata

Hym301 Expression in a Mutant with Altered Number of Tentacles

To further explore the role of Hym301 in tentacle development, we studied a H. magnipapillata mutant, whose number of tentacles is significantly increased [38]. As shown in Figure 6P, adult polyps of H. magnipapillata line E6 have on average seven to eight tentacles, whereas polyps of the H. magnipapillata wild-type line 105 have five to six tentacles. Analysis of tentacle formation during early stages of budding showed that in the mutant H. magnipapillata line E6 tentacles arise earlier and in significantly larger numbers (Figure 6K–6N) than in H. magnipapillata wild-type line 105 (Figure 1N–1Q). In E6 mutant six to seven tentacles appear simultaneously at very early stages of bud development (Figure 6L and and6M),6M), whereas in wild-type H. magnipapillata only four to five tentacles are present at this time point (Figure 1P).

Tentacle development in buds of H. magnipapillata mutant line E6 (Figure 6K–6N) strikingly parallels the pattern of tentacle development in transgenic H. vulgaris AEPA14 (Figure 6F–6I). Is Hym301 responsible for the alterations observed in H. magnipapillata mutant line E6? To examine this possibility we investigated the expression of mHym301A and mHym301B in H. magnipapillata mutant line E6 by in situ hybridization (Figure 6Q and and6R).6R). Given the results described above, we anticipated an increased level of mHym301A expression in the mutant line. As predicted, in situ hybridization showed that the expression domain of mHym301A (Figure 6Q) in the head is considerably enlarged in the mutant and extends down into the body column. In evaginating buds, the differences in Hym301 expression between wild-type and mutant polyps are even more striking. Whereas in buds of the same developmental stage of wild-type H. magnipapillata line 105 mHym301A expression is restricted to the distal part of the evagination, buds in H. magnipapillata mutant line E6 express mHym301A throughout the whole cylindrical bud protrusion (Figure 6Q). Interestingly (Figure 6R), the increased expression domain of mHym301A is accompanied by reduced expression of mHym301B. Taken together, these studies indicate that altered spatiotemporal control of Hym301 expression strongly correlates with altered tentacle development in H. magnipapillata mutant E6, and suggest that the taxonomically restricted gene family Hym301 is essential in head morphogenesis as it seems to ensure the development of tentacles in a correct number and species-specific order.

How are such novel genes incorporated into conserved signaling pathways? To address this question we examined whether mHym301A is interacting with conserved regulatory components such as transcription factor Aristaless (HyAlx). In H. magnipapillata, HyAlx is expressed exclusively in the tentacle zone, the lower part of the head from which tentacles emerge. As shown previously by expression and RNA interference (RNAi) analysis, HyAlx is directly involved in the specification of tissue for tentacle formation [39]. Figure 6 shows that in nontransgenic H. vulgaris AEPwt (Figure 6S–6U), the expression of HyAlx is similar to that described previously [39] in H. magnipapillata. However, in contrast to nontransgenic H. vulgaris AEPwt, in transgenic H. vulgaris AEPA14, the expression of HyAlx is drastically increased (Figure 6S–6U). Reverse transcription PCR (RT-PCR) suggests (Figure 6S) that in polyps overexpressing mHym301A, HyAlx transcripts are not only present in higher amounts in the head region compared with wild type, but are detectable even in the gastric region of AEPA14 animals. Given that overexpression of mHym301A causes an increased expression of HyAlx, interaction between novel gene mHym301A and a conserved transcription factor seems likely. Interestingly, our expression analysis places Hym301 upstream of HyAlx in the signaling cascade. Previously, Smith et al. have shown [39] that depletion of HyAlx RNA drastically delays tentacle appearance in buds, indicating that expression of HyAlx is required for this process. We conclude that in transgenic H. vulgaris AEPA14 the mHym301A-induced increase in the level of HyAlx is responsible for the alteration observed in tentacle development compared with controls.

Silencing of oHym301A in H. oligactis Interferes with Tentacle Formation

We have observed drastic disturbances in head morphogenesis, leading to an increased speed of tentacle formation and altered pattern of tentacle distribution in animals overexpressing mHym301A (Figures 5C–5M and 6A–6J). We next investigated the role of Hym301 in tentacle formation by silencing oHym301A expression in H. oligactis by RNAi.

To optimize the RNAi procedure, we first performed control experiments in which GFP expression was knocked down in H. vulgaris AEP Ecto-1 animals by electroporation of GFP double-stranded RNA (dsRNA). As shown in Figure 7A, introduction of GFP dsRNA causes complete disappearance of GFP protein in large patches of ectodermal epithelial cells covering 10–70% of the polyp's surface. Ecto-1 polyps electroporated with oHym301A dsRNA showed no depletion of GFP signal (Figure 7B).

Figure 7
Silencing of oHym301A in H. oligactis Interferes with Tentacle Formation

Next we examined the effect of oHym301A knock-down on the tentacle formation and morphology of H. oligactis. Polyps were electroporated with oHym301A dsRNA. Control polyps were electroporated with GFP dsRNA. The depletion of oHym301A was monitored by RT-PCR at various time points and by in situ hybridization 7 d after electroporation. As shown in Figure 7C, by introducing oHym301A dsRNA the expression of oHym301A was depleted by about 50% (7 d and 10 d in Figure 7C). Electroporation with GFP dsRNA had no effect on the expression of oHym301A (Figure 7C). The decrease of Hym301 transcript level 4 d after electroporation in GFP control is due to the loss of tentacles during the electroporation procedure (see Materials and Methods). After electroporation tentacles regenerate within 5 d and in control polyps the normal symmetrical pattern is reestablished.

Intriguingly, in H. oligactis polyps electroporated with oHym301A dsRNA, tentacle formation was found to be greatly disturbed. As shown in Figure 7D, 42.5% of polyps electroporated with oHym301A dsRNA developed one or two aberrant tentacles, which were considerably shorter than the others (labeled with arrowheads in Figure 7D). In control animals only 13.6% of polyps displayed abnormalities in tentacle morphology. Abnormal length of tentacles strongly correlates with the absence of oHym301A transcript as detected by in situ hybridization (Figure 7D). On the basis of the observation shown in Figure 7A, which indicates that RNAi depletion is restricted to patches of cells, we assume that shorter tentacles originate from areas of epithelial cells where oHym301A expression was effectively silenced. Taken together, the RNAi experiments indicate that in the absence of oHym301A mRNA tentacle development in Hydra is retarded.

Head regeneration experiments (Figure 7E) also show that H. oligactis polyps in which the oHym301A was silenced by RNAi develop new tentacles considerably more slowly than control polyps that were electroporated with GFP dsRNA (see Table S4). Differences in regeneration kinetics between H. oligactis knock-down and control animals are not as drastic as between animals of H. vulgaris AEP14 and Ecto-1 lines (Figure 5C). We assume that the mosaic distribution of cells with complete knock-down of oHym301A is the reason for this, as tentacles develop more slowly only in the areas in which oHym301A transcript was depleted.

In summary, oHym301A expression in tentacles is required for correct development in H. oligactis. Knock-down of oHym301A gene does not fully abolish tentacle formation, but slows it down considerably (Figure 7D and and7E).7E). Hence, a “novel” gene belonging to the Hym301 gene family is causally involved in the process of tentacle formation in Hydra.

Discussion

Our results delineate a role for novel gene family Hym301 in tentacle formation in Hydra. We show that the species-specific differences in the symmetry of tentacle formation between H. oligactis and H. magnipapillata may be due to the differences in the expression domains of TRGs. Although these results do not determine the precise genetic network responsible for the tentacle formation, they are consistent with previous observations [37,39] and make a significant prediction: conserved regulatory genes and signal transduction cascades alone may not be sufficient to explain the phenotypic differences observed between closely related organisms.

Previous studies have revealed that upstream developmental control mechanisms and regulatory pathways are conserved from Cnidaria to humans [6,7,2931]. Conserved genetic components and specific changes in cis- and trans-acting factors were thought to be sufficient for generating novelty [8,15]. So far, the evolutionary significance of TRGs has not been widely recognized [18,19]. However, it has been presumed earlier [4] that at least part of the resolution of the paradox—where the diversity comes from if the genes are highly conserved—may lie in lower levels of conservation of downstream genes. The sequencing of a large number of eukaryotic and bacterial genomes has uncovered an abundance of genes without homologs, classified [19] as TRGs and has shown that new genes have arisen in the genomes of every group of organisms studied so far including humans [4043]. Here we show that in a basal metazoan group of animals a family of TRGs controls morphological traits in closely related species. The data provide experimental support for the hypothesis [27] that novel genes are involved in specific ecological adaptations that change over time and that such genes serve as the raw material for microevolutionary divergence. The observations also extend earlier findings of an abundance of TRGs in organisms from prokaryotes [18,19] to animals. The observations show that regulatory evolution [8] may act not only by modifying expression domains of conserved genes, but also by spatial and temporal changes in the deployment of TRGs, and that TRGs can be integrated with conserved developmental regulators to form functional signaling cascades.

In Hydra it appears that the TRGs belonging to the Hym301 family are specifically required to control the speed of tentacle formation and arrangement of tentacles on buds and regenerating tips. Transgenic manipulations affecting the expression domains of Hym301 genes are responsible for profound effects in tentacle formation, mimicking evolutionary changes. The Hym301 gene family, therefore, is of special importance in Hydra. Given that Hym301 genes are without homologs in eukaryotic genomes outside Hydrozoa, they might have been specifically acquired in this animal group. An important step that remains to be demonstrated is the role of natural selection in fine tuning of expression of Hym301 genes or their gene regulators for this lineage-specific adaptation. Tentacles are the only structures allowing Hydra polyps to catch and take up food. Differences in their shape, number, and arrangement open different ecological niches and, therefore, are under strict selective constraints. Because pattern formation in Hydra is purely morphallactic and the amount of tissue available for the evagination of tentacles is limited, there are only two mechanisms that can produce different arranged tentacles at the developmental level: simultaneous generation of many short tentacles or early generation of few long tentacles before the other tentacles appear. It seems that two subgroups within the genus Hydra have adapted different species-specific strategies in this respect and that Hym301 genes play a key role in this morphogenetic process. In H. oligactis, expression of Hym301 genes in the tentacles correlates with formation of two long and functional tentacles before the other tentacles appear. In H. magnipapillata and H. vulgaris, which express Hym301 genes in the tentacle zone but not in the tentacles, four or five short tentacles are built simultaneously. It will be interesting to see whether these rather specific adaptations correlate with differences in the planktonic organisms on which polyps feed in nature.

Differences among H. oligactis, H. magnipapillata, and H. vulgaris extend to many other aspects of their morphology and physiology (Figure 1). Therefore, future research on these species may provide novel insights on how TRGs are involved in the evolution of the corresponding adaptive traits. Although it is difficult to generalize from this example, comparing morphogenetic processes in different Hydra species seems to promise new perspectives on how nature fine-tunes morphogenesis. Discovering not only the similarities but also the molecular differences between different organisms might yield intriguing clues in the mechanisms responsible for evolutionary changes.

Materials and Methods

Animals and culture conditions.

Experiments were carried out with H. oligactis, H. vulgaris strain AEP, H. magnipapillata strain 105, and H. magnipapillata strain E6. Transgenic animals were generated using H. vulgaris strain AEP [44]. Animals where cultured according to standard procedures at 18 °C.

Molecular techniques.

Nucleic acid isolation, cDNA synthesis, RT-PCR, cloning, and sequencing were done following standard protocols. RT-PCR was performed using the following primer sets: oHYM301A_F(40) and oHYM301A_R(144), AEP301A_F and AEP301A_R, gmHYMA_E1F and gmHYMA_E3R, Alx_F(422) and Alx_R(631) (HyAlx), GAPDH_F and GAPDG_R (GAPDH), ACTIN35 and ACTIN34 (β-actin). Sequencing was done with Li-COR 4200 and Li-COR 4300 sequencers and manually verified using e-Seq V2.0 and e-Seq V3.0 software. Primer sequences are shown in Table S5.

SSH and cDNA libraries.

For SSH, double-stranded cDNA was synthesized using 2 μg mRNA from H. oligactis and H. magnipapillata strain 105 polyps. SSH was performed in both directions using PCR-Select cDNA Subtraction Kit (Clontech) according to the manufacturer's protocol. Two subtractive libraries were generated: Kiel 6, enriched for H. oligactis specific transcripts and Kiel 7, enriched for H. magnipapillata specific transcripts (see Figure 2). cDNAs were cloned into pGEM-T vector (Promega) and transformed into DH5α Escherichia coli cells. Bacterial clones were picked into 384-well plates using Q-Pix robot and plasmid inserts were sequenced at the Washington University Genome Sequencing Centre (St. Louis, Missouri, United States). Raw sequences were submitted to NCBI (http://www.ncbi.nlm.nih.gov/) under accession numbers CV284311-CV284646 and CV2845050–CV285735 (Kiel 6) and CV284048-CV284310, CV284647-CV285049, and CV285736-CV286173 (Kiel 7).

Isolation of Hym301-like genes from H. magnipapillata, H. oligactis, and H. vulgaris AEP.

To identify all Hym301-like genes in H. magnipapillata, we searched Hydra ESTs at http://blast.ncbi.nlm.nih.gov/Blast.cgi (about 170,000 sequences) and single whole-genome shotgun reads deposited at CompaGen server in Kiel (http://www.compagen.org) using translated BLAST and the mHym301A and mHym301B protein sequences as queries. To isolate Hym301-like genes in H. oligactis we screened macroarrays containing 30,000 cDNA clones with the mixture of mHym301A, mHym301B and oHym301A probes using low stringency washing conditions. Out of eight positive clones, four were identical to the oHym301A cDNA already known from the SSH library, two represented transcripts of a new member of a gene family designated oHym301B, and the other two clones contained sequences not related to Hym301. To isolate Hym301-like genes in H. vulgaris AEP, we performed 3′ RACE PCR according to the previously published method using primer oHym301_F(32) (Table S5) directed against sequence highly conserved in both H. oligactis and H. magnipapillata. In three independent RACE experiments only one type of Hym301-related cDNA was identified, designated later as aepHym301A. Sequences of Hym301 genes were submitted to NCBI under accession numbers EU787492-EU787498.

Gene expression analysis.

For assessment of gene expression, whole-mount in situ hybridization was carried out as described previously [45].

Generation of transgenic H. vulgaris AEP expressing mHym301A:eGFP.

Transgenic founder polyps overexpressing the mHym301A:eGFP fusion protein under control of the β-actin promoter (construct ligE) were produced at the University of Kiel Transgenic Hydra Facility (http://www.transgenic-hydra.org/). Briefly, a 291 bp fragment of mHym301A coding for the full-length protein including the signal peptide was amplified from H. magnipapillata strain 105 cDNA using Platinum High Fidelity Taq polymerase (Invitrogen) and primers Hym301_F(2)Pst and Hym301_R(295)Pst. The cDNA was cloned into the modification of HoTG expression vector using the PstI cutting site (see Figure 4A). The resulting transfection construct was sequenced, plasmid DNA was purified using Qiagen MidiPrep Kit and injected into H. vulgaris AEP embryos as described previously [44]. Embryos began to express the reporter gene 2–3 d after injection. Founder transgenic animals bearing the mHym301A:eGFP construct started to hatch 14 d after microinjection. Out of 24 embryos injected with the LigE construct, two transgenic lines were generated. One of them (line A7) showed stable integration of mHym301A:eGFP in ectodermal and endodermal epithelial cell lineages. Another (line A14) showed integration of the construct in ectodermal epithelial cells only. Initial founder transgenic animals of A14 line were further expanded into a mass culture by clonal propagation by budding and used in all further experiments.

Sequence analysis.

TIGR Indices Clustering Tools [46] were used for clustering the sequences of Kiel 6 and Kiel 7 libraries. Nucleotide and translated BLAST engines at the NCBI server (http://blast.ncbi.nlm.nih.gov/Blast.cgi) were used for homology searches in public databases. The Seqtools program (S.W. Rasmussen, http://www.seqtools.dk) was used for sequence analysis and batch BLASTX and BLASTN searches. DNA and protein sequences were aligned with DnaMan Version 4.12. Phylogenetic tree analysis was performed by the neighbor-joining method with standard settings and a bootstrap value of 1,000 using the Mega 3.1 software. Local assemblies of H. magnipapillata genome were performed manually using single whole genome shotgun reads deposited at NCBI trace archive (http://www.ncbi.nlm.nih.gov/Traces/) and the Compagen server in Kiel (http://www.compagen.org).

Microscopic analysis.

Fluorescent images were taken on a Zeiss Axioscope fluorescence microscope with an Axiocam (Zeiss) digital camera. Confocal laser microscopy was done using a LEICA TCS SP1 CLS microscope. A Zeiss S420 microscope was used for scanning electron microscopy.

Gene silencing by RNAi in Hydra.

Double-stranded RNAs for GFP and oHym301A were synthesized using a MEGAscript RNAi Kit (Ambion). DNA templates for RNA synthesis were generated by PCR using primers T7_GFP_F, T7_GFP_R, o301Ai_T7_F, o301Ai_T7_R (see Table S5). DsRNA was introduced into Hydra polyps by electroporation as described previously [47] with minor modifications. For each experiment 30–60 polyps were placed into chilled electroporation cuvettes with a 4 mm gap (Peqlab). Polyps were washed twice with 1 ml sterile ice-cold Millipore water. Electroporation was carried out in 200 μl water containing 15 μg dsRNA. DsRNA was added to polyps just before electroporation and cuvettes were briefly shaken to insure regular distribution of animals and dsRNA. Bio-Rad Gene Pulser (Bio-Rad) was adjusted to an electric field strength of 0.5 kV/cm and 50-μF capacitance. Polyps were electroporated two times successively with each pulse lasting for about 10–12 ms. Immediately after the second pulse polyps were transferred into 10 ml ice-cold hydra medium that was supplemented with 20% hyperosmotic dissociation medium (DM) containing 6 mM CaCl2, 1.2 mM MgSO4, 3.6 mM KCl, 12.5 mM N-tris-[hydroxymethyl] methyl-2 aminoethanesulfonic acid, 6 mM sodium pyruvate, 6 mM sodium citrate, 6.0 mM glucose, and 50 mg/ml rifampicin, pH 6.9. Electroporation causes cell loss and tissue damage with all the animals losing their tentacles (note the absence of tentacles in Figure 7A and and7B,7B, B,33 d after electroporation). To facilitate recovery, polyps were kept at 10 °C for up to 3 d. Hydra medium supplemented with 20% DM was exchanged every 12 h; viable polyps were separated from cell debris and transferred into new Petri dishes. Thirty-six h after electroporation the medium was exchanged for standard hydra medium. Five d after electroporation polyps were fully recovered and used for experimentation. Survival rate for the electroporated polyps varied between 40% and 60%.

Ecto-1 transgenic line which expresses GFP in all ectodermal epithelial cells was used to directly visualize RNAi in vivo. Electroporation of Ecto-1 animals with GFP dsRNA allowed for the first time to observe the disappearance of the RNAi target protein in living Hydra polyps. Because of the high stability of the GFP protein, the first areas of completely GFP-negative epithelial cells become clearly visible 5 d after electroporation. Complete down-regulation of the target gene was observed in areas of cells that, in the best case, covered about 70% of the polyp surface. The RNAi effect is stable for the period of over 14 d (Figure 7A).

Supporting Information

Figure S1

Analysis of the SSH Library Enriched for H. oligactis–Specific Transcripts:

(A) BLASTN search of H. oligactis clusters and singletons (Kiel 6 library) against all H. magnipapillata ESTs. The cut-off value was set to E < 1e–10. Pie diagram shows the distribution of the SSH clusters and singletons according to their sequence identity to H. magnipapillata ESTs: 42% have sequence identity of 90–100% (possible false positives); 39% have sequence identity of 80–90% (transcripts of highly diverged genes); 19% have sequence identity below 80% (putative H. oligactis-specific genes).

(B–G) Whole mount in situ hybridization showing differences in the expression patterns of homologous genes between H. oligactis and H. magnipapillata. (B) A member of the Kazal-type family of proteinase inhibitors (CL67 in Table S1, CV284473) is not expressed in the foot (stalk) of H. oligactis, whereas its closest homolog (C) in H. magnipapillata (CL42 in Table S2, CV284784) is expressed in gland cells all over the body column, including the foot. (D,E) A novel secreted protein is expressed in ectodermal epithelial cells exclusively in the stalk (D) of H. oligactis (CL225 in Table S1, EU787491). This gene seems to be taxonomically restricted to H. oligactis as (1) no signal could be detected by in situ hybridization in H. magnipapillata (E) and (2) no homologous sequences are present among the 170,000 ESTs and in the genome of H. magnipapillata. (F,G) Hydra-specific genes identified by our approach include also genes expressed in developing nematocytes. Minicollagen-15 (EF624460)-like genes in H. oligactis (CL140, CV285608) and H. magnipapillata (CL173, EU787490) show different expression domains. The minicollagen-15 transcript is absent in the stalk structure of H. oligactis (F) and expands much further down to the foot in H. magnipapillata (G).

(274 KB PDF)

Figure S2

Ectodermal Epithelial Cells of H. vulgaris AEP and Cellular Localization of eGFP and mHym301A:eGFP in Transgenic Lines Ecto-1 and H. vulgaris AEPA14:

Semi-thin section in H. vulgaris AEP shows localization of vesicles on the periphery of ectodermal epithelial cells. In transgenic H. vulgaris AEPA14 polyps the mHym301A:eGFP fusion protein is located in vesicles in ectodermal epithelial cells. In control Ecto-1 polyps transformed with the same expression construct but lacking the mHym301A sequence, the eGFP reporter protein is localized in the cytoplasm. Vesicles appear as black holes on the green background.

(676 KB PDF)

Table S1

Kiel 6 BLASTX Search against the Nonredundant NCBI:

General information about clusters and singletons of H. oligactis-specific cDNA library (Kiel 6). Results of the BLASTX search against the nonredundant NCBI database. Sequences with E value ≥ 1e–5 were referred to as having no significant similarity to the proteins in NCBI database (potential TRGs). The clusters were numbered according to the amount of ESTs comprising them, with cluster 01 (CL01CONTIG1) being the largest. The consensus cluster sequences and singleton sequences are stored as a multi-sequence file (FASTA format) at the COMPAGEN server (http://compagen.zoologie.uni-kiel.de/retrieve.htm).

(78 KB PDF)

Table S2

Kiel 7 BLASTX Search against the Nonredundant NCBI:

General information about clusters and singletons of H. magnipapillata-specific cDNA library (Kiel 7). Results of the BLASTX search against the nonredundant NCBI database. Sequences with E value ≥ 1e–5 were referred to as having no significant similarity to the proteins in NCBI database (potential TRGs). The clusters were numbered according to the amount of ESTs comprising them, with cluster 01 (CL01CONTIG1) being the largest. The consensus cluster sequences and singleton sequences are stored as a multi-sequence file (FASTA format) at the COMPAGEN server (http://compagen.zoologie.uni-kiel.de/retrieve.htm).

(80 KB PDF)

Table S3

Kiel 6 BLASTN Search against Hydra ESTs:

Results of the BLASTN search of H. oligactis clusters and singletons from the Kiel 6 library against all H. magnipapillata ESTs. The cut-off value was set to E < 1e–10. Sequences with E value ≥ 1e–10 were referred to as having no significant similarity among H. magnipapillata ESTs (transcripts of potential H. oligactis-specific genes).

(68 KB PDF)

Table S4

Regeneration Experiments:

Mean number of tentacles per polyp at 42, 66, and 130 h after decapitation (mean ± standard deviation) in independent experiments (I, II, III, etc.).

(15 KB PDF)

Table S5

Oligonucleotide Primer Sequences:

(7 KB PDF)

Acknowledgments

We thank the members of the Bosch laboratory for discussion, Antje Thomas for help with microscopy; Jan Lohman, Ingrid Lohman, and Sebastian Fraune for comments on the manuscript; and T. Fujisawa and H. Shimizu for the gift of H. magnipapillata strain E6. We are grateful to Rene Augustin and Bjorn Spudy for the opportunity to use H. oligactis cDNA macroarrays. We also thank Lucinda Fulton and the Genome Sequencing Centre in St. Louis (US).

Abbreviations

dsRNA
double-stranded RNA
eGFP
enhanced green fluorescent protein
EST
expressed sequence tag
GAPDH
glyceraldehyde 3-phosphate dehydrogenase
GFP
green fluorescent protein
RNAi
RNA interference
SSH
suppression subtractive hybridization
TRG
taxonomically restricted gene

Footnotes

Author contributions. KK, FA-E, JW, GH, and TCGB conceived and designed the experiments. KK, FA-E, SS, JW, GH, and TCGB performed the experiments. KK, FA-E, SS, JW, GH, and TCGB analyzed the data. GH contributed reagents/materials/analysis tools. KK and TCGB wrote the paper.

Funding. Supported in part by grants from the Deutsche Forschungsgemeinschaft (DFG SFB 617-A1), and grants from the DFG Cluster of Excellence programs “The Future Ocean” and “Inflammation at Interfaces” (to TCGB).

Competing interests. The authors have declared that no competing interests exist.

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