Conservation of ParaHox genes' function in patterning of the digestive tract of the marine gastropod Gibbula varia

Background Presence of all three ParaHox genes has been described in deuterostomes and lophotrochozoans, but to date one of these three genes, Xlox has not been reported from any ecdysozoan taxa and both Xlox and Gsx are absent in nematodes. There is evidence that the ParaHox genes were ancestrally a single chromosomal cluster. Colinear expression of the ParaHox genes in anterior, middle, and posterior tissues of several species studied so far suggest that these genes may be responsible for axial patterning of the digestive tract. So far, there are no data on expression of these genes in molluscs. Results We isolated the complete coding sequences of the three Gibbula varia ParaHox genes, and then tested their expression in larval and postlarval development. In Gibbula varia, the ParaHox genes participate in patterning of the digestive tract and are expressed in some cells of the neuroectoderm. The expression of these genes coincides with the gradual formation of the gut in the larva. Gva-Gsx patterns potential neural precursors of cerebral ganglia as well as of the apical sensory organ. During larval development this gene is involved in the formation of the mouth and during postlarval development it is expressed in the precursor cells involved in secretion of the radula, the odontoblasts. Gva-Xolx and Gva-Cdx are involved in gut patterning in the middle and posterior parts of digestive tract, respectively. Both genes are expressed in some ventral neuroectodermal cells; however the expression of Gva-Cdx fades in later larval stages while the expression of Gva-Xolx in these cells persists. Conclusions In Gibbula varia the ParaHox genes are expressed during anterior-posterior patterning of the digestive system. This colinearity is not easy to spot during early larval stages because the differentiated endothelial cells within the yolk permanently migrate to their destinations in the gut. After torsion, Gsx patterns the mouth and foregut, Xlox the midgut gland or digestive gland, and Cdx the hindgut. ParaHox genes of Gibbula are also expressed during specification of cerebral and ventral neuroectodermal cells. Our results provide additional support for the ancestral complexity of Gsx expression and its ancestral role in mouth patterning in protostomes, which was secondarily lost or simplified in some species.


Background
The three ParaHox genes, Gsx, Xlox, and Cdx, were first described as a gene cluster in the invertebrate chordate Branchiostoma floridae (amphioxus) by the elegant work of Brooke et al. 1998 [1]. ParaHox and Hox genes are believed to have evolved from a single ancient proto-Hox cluster composed of two to four genes prior to the divergence of cnidarians and bilaterians. Thus, they are con-sidered evolutionary sister (or paralogue) clusters [1][2][3][4][5][6][7]. Vectorial expression of the ParaHox genes in anterior, middle, and posterior tissues of amphioxus and its distinct similarities to vertebrate ParaHox gene expression suggest that these genes may be responsible for axial patterning of the digestive tract [1,3].

Expression of ParaHox genes in ecdysozoans
In ecdysozoans, Gsx expression has been documented in the insects Drosophila and Tribolium [30,31]. Insect Gsx (called ind) is expressed along a pair of medio-lateral neural columns and promotes neural precursor formation in the medial and intermediate columns of the CNS [30,31]. The central ParaHox gene, Xlox, is lost in all insect genomes sequenced to date. Caudal has been known as a posterior patterning gene in several arthropods during segmentation [32][33][34][35][36][37][38][39][40][41][42]. Cdx is also a posterior patterning gene in the nematode Caenorhabditis elegans. Here, this gene is called pal-1 and patterns the precursor cells of alae and rays in the posterior of the worm [43]. Gsx and Xlox orthologs are absent in the nematode [44].

Expression of ParaHox genes in lophotrochozoans
Within Lophotrochozoa, expression of the full complement of ParaHox genes has been described in the polychaetes Capitella teleta, Nereis virens, and Platynereis dumerilii [45][46][47][48]. In Capitella, Gsx is not expressed in the gut but in some neuroectoderm cells of the anterior brain [45]. This is very different from the expression of Gsx in the nereid polychaetes, Nereis virens and Platynereis dumerilii [46,48]. Nereid Gsx is first expressed in symmetrical bilateral domains in the dorso-medial episphere of the trochophore [46,48]. Later this gene is expressed during formation of the midgut and the posterior foregut in both nereids [46,48]. Xlox is expressed throughout the midgut in the polychaete Capitella [45]. This is also true for the Xlox genes, named Lox3 in the leeches Helobdella triserialis and Hirudo medicinalis [49,50]. Expression of Xlox is not reported in the nervous system of these annelids [45,49,50]. Nereid Xlox is also expressed in the midgut, but in contrast to Capitella and the leeches its expression is additionally detected in the CNS [46,48]. As in arthropods, Cdx is a posterior patterning gene in the annelids. However in Platynereis, Nereis, Tubifex, and Capitella there are both anterior and posterior expression domains of Cdx [45][46][47][48]51]. Capitella Cdx is expressed in the cerebral ganglia, Nvi-Cdx expression is detectable in the ventral nervous system, while Pdu-Cdx is not detected in the nervous system [45][46][47][48]. Expression of Cdx is also detected in more posterior parts of the gut [45][46][47][48]. Moreover, Cdx is expressed in the posterior ectodermal cells that form the pygidium epidermis of both nereids [46][47][48]. Additionally, Cdx expression can be traced in mesodermal cells in Capitella, Tubifex, and Platynereis [45,47,48,51].
Little is known of ParaHox genes in other Lophotrochozoa than annelids [52][53][54][55]. The only available data are on the Cdx gene expression during the early development of the marine limpet, Patella vulgata [53]. Pvu-Cdx is expressed at the onset of gastrulation in the ectodermal cells at the posterior edge of the blastopore and in the paired mesentoblasts [53]. During trochophore larval stage, PvuCdx is expressed in the posterior neurectoderm of the larva, as well as in part of the mesoderm [53]. Within Mollusca, a full complement of ParaHox genes has been shown for the chiton Nuttallochiton mirandus and the scallop Pecten maximus [54,55]. However the information is limited to partial homeobox sequences, whereas expression patterns of Gsx and Xlox or chromosomal organisation of ParaHox genes have not been reported yet in any mollusc species.

Ancestral role of ParaHox genes
Holland (2001) elaborated the hypothesis of the ancestral role of ParaHox genes proposed by the original work of Brooke et al. 1998 [1,3]. Holland's hypothesis proposes that the three ParaHox genes originated from the Proto Hox gene cluster and pattern anterior, middle and posterior gut regions in a colinear manner in basal animals [3]. According to this hypothesis, a link of Gsx and anterior gut development existed in basal animals. However, Gsx is not expressed in the anterior gut of deuterostomes. This is explained by the loss of the primary mouth and formation of a secondary mouth in deuterostomes [3].

Aim
Gastropoda is undoubtedly the most successful taxon of the Mollusca, embracing more than 80% of all mollusc species [56]. The vetigastropod Gibbula varia L. is a shallow subtidal top shell snail with encapsulated development. The lecithotrophic larval development is completed within the eggs. The juveniles leave the gelatinous egg masses only after metamorphosis. In order to elucidate the function of the ParaHox genes in molluscs and to gain broader insights into the evolution of the ParaHox genes in the Lophotrochozoa, we describe the sequences as well as expression patterns for all three ParaHox orthologues by whole mount in situ hybridiza-tion from embryonic through juvenile stages in the top shell Gibbula varia. This is the first report of expression patterns of the full ParaHox complement in a mollusc.

Gibbula varia life history
Gibbula varia is a dioecious species. The eggs fertilized via copulation are laid in gelatinous egg masses (additional file 1, Figure S1A, S1B, S1C). Development of the embryos and larvae takes place inside the egg capsule and takes about four days. The main stages of G. varia development are presented in Table 1. Epibolic gastrulation occurs by the micromeres rapidly spreading downwards and enclosing the macromeres. The blastopore, being wide at first, gradually becomes constricted at 10 to 12 hours post fertilization (hpf ), when the trochoblasts start to become ciliated. At 16 hpf, the prototroch is clearly visible as a circular ciliary band, separating the trochophore larva's episphere from the hyposphere (additional file 1, Figure S1D). There is no sign of apical cilia (apical tuft) at any stage in the development of the trochophore, although the pretrochal cells were observed to be smaller than those of the posttrochal region (additional file 1, Figure S1D). By this stage the blastopore gradually moves to a position just below the prototroch, forming the stomodeum. Simultaneously, the shell-gland invagination appears as a thin patch of cells gradually spreading over the dorsal region of the larva (additional file 1, Figure S1D). At 18 hpf, the late trochophore larva comprises a prototroch, the shell field surrounded by the mantle edge, and a pedal rudiment (additional file 1, Figure S1E). The late trochophore (24 hpf ) turns into an encapsulated pretorsional veliger larva by differentiation of the prototroch to a distinct velum (additional file 1, Figure S1F). The mantle fold and mantle cavity become visible mid-ventrally on the posterior surface of the pedal rudiment (additional file 1, Figure S1E, S1G). At 36 hpf, the pretorsional veliger has a velum, an apical organ marked by apical cilia (apical tuft), a mouth opening, and a pedal rudiment with the operculum anlage (additional file 1, Figure S1H). The first 90° of torsion take place between 36-48 hpf, presumably by contraction of the larval retractor (shell) muscle. This results in a 90° displacement of the mantle cavity to the right side, and, when viewed from the front, the foot and velum are rotated anti-clockwise in relation to the protoconch. The remaining part of torsion is completed within one day while the velum gradually becomes reduced in size and splits ventrally (additional file 1, Figure S1I). At 60 hpf, the operculum appears in the posttorsional veliger larva (additional file 1, Figure S1I). The radula and cephalic eyes appear about three days after fertilization. As the eyes form, the cephalic tentacles begin to appear as outgrowths of the prevelar surface. The juvenile hatches on the fourth day of development (about 96 hpf ) and after that mineralization of the shell begins. The animals become sexually mature after 11-12 months.

Development of gut in G. varia
The development of the digestive tract starts with the development of the stomodeum (future mouth opening) in the trochophore (additional file 1, Figure S1E). The mouth opens during the pretorsional veliger stage (additional file 1, Figure S1H, S1J) whereas the anus opens in the late posttorsional stage at the site of a few ciliated cells (anal markers). The development of the digestive tract is very similar to that described in G. cineraria and Haliotis tuberculata [57,58]. The digestive gland begins

Brief description of main features
Early Trochophore Larva (12 hpf) The pretrochal cells are smaller than the posttrochal cells; prototroch starts to form by cilliation of trochoblasts; shell gland starts to evaginate; foot rudiment and stomodaeum are not completely formed.
Pretorsional veliger larva (36-48 hpf) The mantle and mantle cavity form. The larva has a velum, apical organ marked by apical cilia, mouth opening, and pedal rudiment with anlage of operculum.
Post-torsional veliger larva (60 hpf) The mantle lies over the back of the head and the velum gradually splits ventrally, the operculum apears.
Metamorphotic (competence) stage (72 hpf) Eye rudiments and cephalic tentacles begin to form in the prevelar area. The anlage of the radula becomes visible.
Encapsulated juvenile Velum is completely lost; eyes and cerebral tentacles are formed.
Hatchling (96 hpf) The encapsulated juvenile hatches and shell mineralization begins.
to differentiate on the left side of the veliger just before torsion sets in [57,58]. The gut develops from differentiated endodermal cells initially scattered within the yolk in the pretorsional veliger. They later migrate to the yolk boundaries to form the definitive midgut in the posttorsional veliger [57,58]. Later, the hindgut develops from actively dividing cells of the digestive gland migrating to their final positions in the intestine [57,58]. The competent larva's digestive system comprises a mouth opening and a bipartite oesophagus (the anterior part immediately behind the buccal cavity is not effected by torsion, the mid oesophagus includes a portion affected by the torsion), a stomach with the digestive gland, the hindgut leading to the anus that opens into the mantle cavity over the back of the head (additional file 1, Figure S2A and S2B). The radula anlage is a ventral differentiation of the foregut where mesenchym cells aggregate. The radula teeth become visible in the competent larva at the distal end of the radula sheath (additional file 1, Figure S2A and S2B).

ParaHox gene sequences
The entire coding sequences for all three G. varia Para-Hox genes were isolated by a combination of 3' and 5' rapid amplification of cDNA ends (RACE, see Methods). Alignments of each G. varia ParaHox amino acid sequence to orthologs of other species are shown in additional file 2, Figure S3, S4, and S5. Beside the homeobox which is the main region of conservation between Para-Hox genes, further conserved domains are the N-terminal domain in Gsx, and the hexapeptide motifs just upstream of the homeodomains in both Xlox and Cdx (Additional file 2, Figure S3, S4, and S5). The classification of the G. varia ParaHox genes into their orthology groups is apparent from phylogenetic analyses ( Figure 1). The species names and accession number of the genes used in phylogenetic analysis are provided in additional file 2. Although the phylogenetic analysis clearly assigns the Gibbula paraHox genes to the Gsx, Xlox and Cdx classes with high support values, the internal grouping remains unclear.

ParaHox gene expression in the trochophore larva
We did not detect Gva-ParaHox transcripts by wholemount in situ hybridization (WMISH) in developmental stages before the trochophore stage. A scanning electron micrograph (SEM) of a late trochophore larva (18-24 hpf ) is shown in Figure 2A.
The expression pattern of Gva-Gsx is rather dynamic. The first signs of transcripts of Gva-Gsx are already detected at 12 hpf in early trochophore larvae, when a pair of intensive, bilateral expression domains appears in the dorso-medial episphere ( Figure 2B). When viewed from the anterior, each pair of expression domains appears to be composed of 4-5 Gva-Gsx-positive cells, presumably in the area of future cerebral ganglia ( Figure  2C). This pattern of expression continues in 18 hpf trochophores ( Figure 2D and 2E). Here, the pattern of expression becomes considerably more complex. In addition to the paired expression domains in the dorsomedial episphere, Gva-Gsx transcripts can now be detected in a pair of cells at the tip of the developing apical sensory organ ( Figure 2D and 2E). These two Gva-Gsx-positive cells at the tip of the apical organ do not bear any cilia or apical tuft in the trochophore stage of G. varia ( Figure 2F). The expression of Gva-Gsx in the apical sensory organ is restricted to two groups consisting of three sensory cells ( Figure 2G). Beside the expression in prospective neural or sensory tissues, Gva-Gsx transcripts are also detected around the stomodeum where they appear for the first time in trochophore 18 hpf in two intensely stained bilateral semicircular clusters located anteriorly at the sides of the mouth and a less intensely stained semicircular domain at the posterior part of the mouth ( Figure 2D and 2H). Figures 2H and 2I show the trochophore stomodaeum and Gva-Gsx expression around it at 18 hpf. About 24 hpf, Gva-Gsx is expressed in a complete circle around the stomodeum ( Figure 2J) and in three episphere domains: a pair of adjacent cells at the tip of the apical sensory organ, and two pairs of cell groups dorsolaterally marking presumptive sites of future cephalic neuroectodermal differentiation ( Figure 2K).
Gva-Xlox transcription begins later than Gva-Gsx expression. No expression is detectable until 24 hpf when Gva-Xlox transcripts appear in a group of cells located ventrally in the hyposphere and in a pair of symmetrical expression domains in the medio-ventral episphere of the trochophore larva ( Figure 2L and 2M). These symmetrical expression areas are located ventrally of the more intensely stained Gva-Gsx expression domains in the pretrochal area. Gva-Xlox is also expressed in the hyposphere in 8-9 cells forming a semicircle around the anal marker ( Figures 2N and 2O). These weakly stained Gva-Xlox-positive cells are probably part of ventral neuroectoderm.
Gva-Cdx transcripts are first detected in the early trochophore larva (12 hpf    and 2Q). Using Patella vulgata as a reference, the latter expression of Pvu-Cdx probably marks the left and right primary mesentoblasts (green arrows in Figure 2P and 2Q). Gva-Cdx-positive neuroectodermal cells are first observed as a patch of cells expressing this gene in varying intensities ( Figure 2P). Gradually they migrate to the boundary of the expression area ( Figure 2Q) so that they from a circle of Gva-Cdx-expressing cells around the anal marker at 24 hpf ( Figure 2R and 2S). The expression of Gva-Cdx around the anal marker at 24 hpf partly overlaps with the expression of Gva-Xlox in the ventral area at this stage, which is visible as a semicircle located ventrally around the anal marker ( Figure 2N and 2R).

ParaHox gene expression in the pretorsional veliger larva
The transcripts of all three Gva-ParaHox genes are detected almost simultaneously in the visceral mass area of the pretorsional veliger larva prior to torsion (36-48 hpf ), on the left side of the larva where the digestive gland is forming (Figure 3). At this stage, the velum forms a complete circle and a pair of apical tufts is observed in the velar area ( Figures 3A, B, and 3C). In addition to the apical tufts, there are "sensory cups" in the velar area. These are ciliated pockets embedded within the apical ganglion ( Figure 3B). The expression of Gva-Gsx observed in the area of the mouth opening and of the apical organ of the late trochophore larva ( Figure 2J and 2K) is retained in the pretorsional veliger ( Figure 3D). Gva-Gsx transcripts are also detected in the ventral part of the forming digestive gland in the left side of the visceral mass ( Figure 3D and 3E). Gva-Gsx-positive signals are further detected in the area of the mouth opening ( Figure 3E) and in five cells in the area of the apical organ ( Figure 3D), the two apical tuft cells ( Figure 3F), and the sensory cup cells (compare Figures 3B and 3D). Similar to Gva-Gsx, Gva-Xlox is expressed in the left side of the pretorsional veliger in the forming digestive gland ( Figure 3G). The expression area of Gva-Xlox is located in the ventral part of the digestive gland, more dorsally but partly overlapping Gva-Gsx expression ( Figure 3G and 3H). The expression pattern of Gva-Xlox detected on the ventral side of the episphere of the late trochophore larva ( Figure 2L and 2M red arrow heads) is lost in the pretorsional veliger stage ( Figure 3G and 3I). Additionally, five ectodermally derived Gva-Xlox-positive cells appear on the right side of the larva prior to torsion ( Figure 3I). Similar to the trochophore stage ( Figure 2M and 2O), these ectodermal cells form an incomplete circle and are presumably linked to the ventral nervous system ( Figure 3J). Gva-Cad is expressed weakly in the whole area of the nascent digestive gland of the pretorsional veliger larva ( Figure 3K). The intensity of expression is stronger in a few cells in the dorsal area of the visceral mass in the left side of the larva ( Figure 3K and 3L).

Expression of ParaHox genes in veliger and competent larvae
After torsion (60 hpf ), the velum reduces in size with a ventral split, and the mantle expands over the back of the head ( Figure 4A). As the digestive tract continues to develop in the posttorsional veliger larva, expression patterns of Gva-ParaHox become more elaborated. At this stage, Gva-Gsx expression in the ventral part of the digestive gland and in the area of the mouth opening persists ( Figure 4B and 4C). Sections reveal Gva-Gsx-positive cells at the ventral border of the area of yolk-filled cells ( Figure 4D). Gva-Gsx transcripts are further apparent as paired domains beneath the apical organ where the formation of the cerebral ganglia commences (Figures 4C  and 4D). At about three days post fertilization, expression of Gva-Gsx fades in the digestive gland. Instead, the gene is now expressed in the foregut around the area of the radula anlage ( Figure 4E and 4F). At metamorphosis, when the apical sensory organ starts to dissociate, Gva-Gsx continues to be expressed in the area of the cerebral ganglia ( Figure 4F). Gva-Xlox expression persists on the left side of the visceral mass from the pretorsional to the posttorsional stages ( Figure 4G and 4H). Sections through the left side of the larva reveal that these Gva-Xlox-positive cells are part of the developing digestive gland ( Figure 4J). Six or seven ectodermally-derived Gva-Xlox-positive cells are located in the ventral part of the visceral mass ( Figure 4G, H, and 4I). Gva-Cdx is mainly expressed in the newly formed hindgut and rectum, and weakly in the digestive gland ( Figures 4K and 4L).

Post-larval ParaHox gene expression
Serial section in situ hybridizations were used to trace the expression pattern of all three Gva-ParaHox in the hatchling (about four days after fertilization). No positive signals for Gva-Xlox and Gva-Cad transcripts are detected at this stage. Gva-Gsx is the only ParaHox gene that is expressed in the most posterior part of the radula sac during postlarval development ( Figure 5). The juvenile hatchling has a complete radula with the radula sheath, buccal musculature, and radula bolsters (also called odontoblastic cartilages, Figure 5A). The posterior end of the radula sac forms the odontoblastic cushion which consists of a single-layered epithelium arranged in a semicircle and protruding into the sac's lumen. The epithelial cells are produced by two separated dorsolateral mitotic centres at the end of the sac ( Figure 5B). Mitotic activity is scattered over the posterior area of odontoblastic cushions where the cells are small and undifferentiated. Towards the anterior of the cushions, the cells gradually elongate and form the tall odontoblastic epithelial cells ( Figure 5B). Gva-Gsx transcripts are mainly detected in the paired odontoblastic cushions at the base of the radula ( Figure 5C; the weak signal observed in the pedal area seems to be unspecific). Gva-Gsx is expressed both in undifferentiated cells located at the back of the cushions and in odontoblastic epithelial cells. No transcripts were detected in the cells separating the two halves of the odontoblastic cushion ( Figure 5D and 5E). The intensity of expression of Gva-Gsx diminishes gradually from posterior to anterior, i.e. from the undifferentiated cells to fully differentiated epithelial odontoblasts ( Figure 5E).

Is ParaHox gene expression colinear during patterning of gut?
It has been proposed that the origin of the three germ layered animals, the Bilateria, is associated with the innovation of several gene clusters of the ANTP family, with the Hox-cluster genes participating mainly in patterning of the neuroectoderm, the NK-cluster genes in formation of the mesodermal layers, and ParaHox in colinear regionalisation of the endoderm [1,3]. Of the animals studied to date, the chromosomal linkage of ParaHox genes has been shown only in amphioxus, mouse, and human [1,3]. The ParaHox genes are not linked in teleost fishes, the ascidian or the sea urchin [27,28,59]. The only description of the expression patterns of all three ParaHox genes for lophotrochozoans in relation to their genomic organi-sation is for the polychaete P. dumerilii [48]. Here, Gsx and Xlox are clustered and Cdx is separated, without clear evidence of colinear expression.
We were unable to detect clear colinear expression of ParaHox genes in Gibbula prior to torsion. If present, it is obscured by the permanent migration of cells from the digestive gland to their final positions in the gut, and by torsion processes. After torsion, however, a spatially colinear expression of ParaHox genes is obvious in the digestive system, with Gva-Gsx patterning the mouth opening and radula anlage, Gva-Xlox expressed in the midgut, and Gva-Cdx in the hindgut (Figure 4 and 6). Therefore, our results support Holland's hypothesis that ParaHox genes are involved in gut regionalization along the anterior-posterior body axis in protostomes [3].
There also seems to be a temporal colinearity in expression of ParaHox genes in the gradual formation of the digestive system. In the trochophore larva, development of the digestive system begins with the formation of the stomodeum involving Gva-Gsx expression only ( Figure  6). Gva-Xlox and Gva-Cdx are expressed at later stages in the more posterior parts of the gut. When the patterning of the gut is completed in the hatchling, expressions of Gva-Xlox and Gva-Cdx cease while Gva-Gsx continues to be involved in the patterning of the radula (Figure 6). During postlarval development, Gva-Gsx is expressed in the paired odontoblastic cushions of Gibbula ( Figure 5). The gradient of Gva-Gsx expression from posterior to anterior in the odontoblastic cushions suggests that this gene is associated with mitotic features of these cells and their ability to divide and replace the odontoblasts, rather than direct involvement in secretion of radula teeth.

Expression of ParaHox genes in cephalic neural and neurosensory cells
Gastropod larvae are well provisioned with multicellular sensory structures, but only the apical sensory organ is typically present in both plankton-feeding and nonplankton-feeding veligers [60]. This suggests that information detected by the apical sensory organ is important during the entire larval stage, regardless of the length of larval life or capacity for feeding. Moreover, the apical sensory organ disappears at metamorphosis in species . bc buccal cavity, cf ctenidial filaments, eso esophagus, frt forming radula teeth, ie inferior epithelium, j jaw, mf mantle fold, mo mouth opening, oc odontoblastic cushions, od odontoblasts, odc odontoblastic cartilage, op odontophores, op operculum, pf pedal folds, rm radula membrane, rn radula nerve, rt radula teeth, sc separating cells, sm shell matrix, tsm tensor muscle. where this has been studied [61]. Therefore the apical sensory organ has functions restricted to the larval stage. During larval development in Gibbula, Gva-Gsx exhibits a complex pattern of expression in potential cephalic neural cells and in the apical organ. This pattern shows distinct similarity to Pdu-Gsx expression in the trochophore stage in which Pdu-Gsx expression is detectable in flask-shaped sensory-neurosecretory cells in the medial forebrain [48]. Prior to torsion, the Gva-Gsx pattern is spotted in the paired apical tufts and several neurosecretory cells or sensory cups of the apical organ (Figure 3 and 6). This gene also appears to be involved in the formation of parts of the cerebral ganglia from the apical sensory organ in competent larvae. This compares well to the polychaetes Capitella, Nereis, and Platynereis, where Gsx is expressed in the cerebral ganglia [45,46,48].
Our results may lend further support to the theory of complex ancestral expression of Gsx that was secondarily simplified in several lineages. In addition to Gva-Gsx expression in the dorsal episphere of the trochophore, Gva-Xlox is detected in a pair of expression domains located more ventrally. It is possible that these cells contribute to neural cells of future cerebral ganglia. However this pattern of expression is transient and is lost in later developmental stages.

Possible expression of ParaHox genes in the trunk neuroectoderm
Expression of ParaHox genes in ventral or dorsal neuroectoderm has been demonstrated in several species. Within Lophotrochozoa Capl-Cdx is expressed in posterior neuroectodermal cells in the polychaete Capitella. In Platynereis, Pdu-Gsx is expressed in a central part of the larval ventral neuroectoderm in which somatic serotonergic neurons are identified [45,48]. Nereis is the only species studied so far in which all three ParaHox genes are known to be involved in patterning of the trunk neuroectoderm [46]. In Gibbula, Gva-Xlox and Gva-Cdx are expressed around the anal marker in the trochophore larvae. It has been shown that these cells express SoxB in the prospective neuroectoderm of the trunk in Patella [62]. Therefore, it is likely that these cells expressing Gva-Xlox and Gva-Cdx contribute to the trunk neuroectoderm. Temporary expression of Gva-Cdx in ventral neuroectoderm earlier during development, and expression of Gva-Xlox in overlapping regions at a later stage (Figure 6), may suggest that Gva-Cdx contributes to patterning of ventral neuroectoderm upstream of Gva-Xlox.

Hypothetical ancestral ParaHox gene expression
Comparative analyses across the animal kingdom show conservation of ParaHox gene expression domains in distinct tissues. Comparing Platynereis ParaHox gene expression to that of the orthologues in deuterostomes and ecdysozoans, Hui et al. 2009 confirmed Holland's hypothesis about the ancestral role of ParaHox genes, suggesting that the pattern of Gsx expression in the protostome-deuterostome ancestor was complex, with Gsx domains in several structures of the nervous system, and was secondarily reduced to small patches of expression in the anterior CNS in several lineages [48]. Holland's model further suggests that Gsx was expressed in the mouth region of the last bilaterian ancestor [3]. Lack of Gsx expression in the anterior gut of deuterostomes is explained by loss of the primary mouth and evolution of a new secondary mouth [3]. If this be the case, protostomes should maintain Gsx expression in anterior gut structures. Capitella results do not support such a model since CapI-Gsx expression is limited to a restricted region of the forming brain. The expression of Nvi-Gsh, Pdu-Gsx, and Gva-Gsx described here provides further support to the ancestral mouth patterning role of Gsx [46,48].
Xlox is expressed during midgut development in annelids [45,46,[48][49][50]. Pdu-Xlox and Nvi-Xlox are also expressed in the nervous system. In Gibbula, Gva-Xlox pattern is detected in the digestive gland and ventral neuroectoderm, and expression in potential cephalic nerve cells is transient. Therefore, our results provide additional support that the expression of Xlox may reflect an ancestral function in central regions of the gut as well as a role in the nervous system. If this hypothesis is true, however, it would once more imply secondary simplification and loss of neural Xlox expression in several lineages [45]. However, the possibility that ancestral Xlox expression was simple and has become more complicated in different lineages cannot be ruled out since Xlox is expressed in ventral neuroectoderm in Nereis and Gibbula, in addition to cerebral ganglia, but is lacking in all other protostomes studied to date [46].
Cdx shows a complex, dynamic pattern of expression in cells of the ectoderm, endoderm and possibly mesoderm, extending to extremely anterior regions in all annelids studied so far [45][46][47][48]51]. This anterior expression of Cdx was also recently described in the acoel flatworm, Convolutriloba longifissura [63]. ClCdx is expressed in the commissures posterior to the statocyst, following the paths of nerve tracks and extending anteriorly. ClCdx is also expressed in an area surrounding the eyes, forming direct connections to the brain commissure [63]. Cdx anterior expression seems to be the case in the limpet Patella as well, in which the gene is expressed in posterior ectoderm during gastrulation. The posterior ectodermal expression starts to fade in the trochophore, while expression extends anteriorly in the shape of an incomplete equatorial ring of ectodermal cells that corresponds to some cells of the prototroch [53]. Later in the young free swimming trochophore, Pvu-Cdx expression in the prototroch disappears. The gene is also transiently expressed in the stomodeum [53]. Gva-Cdx expression differs from that of Pvu-Cdx by being absent during gastrulation. In addition, we did not detect any sign of Gva-Cdx expression in the trochophore prototroch or stomodeum. In contrast, the detection of Cdx in mesentoblasts and in ectodermal cells situated on the posterior most part of the ventral side of the trochophore is a common feature in Gibbula and Patella. These are some of the cells that also express SoxB, a neurectodermal marker [62]. Therefore, Cdx seems to pattern the ventral neuroectoderm as well as mesentoblasts in gastropods. Anterior expression of Cdx was not observed during the larval development of Gibbula at any stage. This can be either interpreted as secondary loss of the anterior function of Cdx in Gibbula, or as a gain of function for this gene in several tissues in other species. The first possibility has been favoured since it can be explained by the separation of the gene from the cluster [48]. Nonetheless, variety in the pattern of expression of Cdx in different animals can serve as another example for the plasticity of gene expression during evolution. Whether the expression of the ParaHox genes in nervous systems is related to their function in the gut, i.e. innervation of different parts of the gut and/ or to feeding behaviour, awaits future research. Gene function experiments, therefore, would be desirable to give us better understanding of how these genes are employed.

Conclusions
The expression of ParaHox genes during anterior-posterior development of the digestive system (with Gsx patterning the mouth and foregut, Xlox patterning the midgut or digestive gland, and Cdx patterning the hindgut) suggests that these genes are involved in anterior-posterior specification of the G. varia gut. Our results support Holland's hypothesis that ParaHox genes are involved in gut regionalization and offer further support to the ancestral mouth patterning role of Gsx in protostomes. All three ParaHox genes of G. varia are involved in patterning of the nervous system. Gva-Gsx and Gva-Xlox are expressed in neural precursors of cerebral ganglia, the expression domain of these two genes does not coincide in the episphere and fades away in the case of Gva-Xlox in later larval stages. Additionally, Gva-Gsx patterns the neurosensory cells of the apical organ. Gva-Xlox and Gva-Cdx pattern the ventral neuroectoderm with Cdx possibly acting upstream of Xlox. During postlarval development, Gva-Gsx transcripts are detected in the precursor cells of odontoblasts at the base of the radula sac. This is probably a molluscan novelty related to radula evolution. Further research in other molluscan classes and use of experimental tools, e.g. RNAi, are required to improve our understanding of gene functions and enable a sound reconstruction of their ancestral role.

Snail culturing
The adults of Gibbula varia (L.) were collected in Crete, Greece and cultured in 150-200 liter aquariums in artificial sea water at 22°C (salinity 28°). Copulation was induced by lowering the salinity a few degrees by adding fresh water to the aquariums at 17°C (personal observation of Achim Meyer, The Johannes Gutenberg University of Mainz).

Cloning of ParaHox genes
DNA extraction was performed using the PeqGOLD Tissue DNA kit (PEQLAB Biotechnologie GmbH, Polling, Austria) according to the manufacturer's instructions. Homeobox fragments of ParaHox genes were obtained by polymerase chain reaction (PCR) from genomic DNA using Hox degenerate primers described previously [64,65]. These primers produce PCR amplification products that are mixtures of different fragments containing homeobox. The PCR fragments were purified using peq-GOLD MicroSpin Cycle-Pure Kit (PEQLAB Biotechnologie GmbH, Polling, Austria). Purified PCR products were cloned with the TOPO TA Cloning Kit (Invitrogen GmbH, Karlsruhe, Germany). In total 255 clones were sequenced and all eleven Hox genes (Samadi and Steiner, unpublished data) and the three ParaHox genes were recovered. RNA was extracted from blastula and gastrula stages, trochophore, veliger, and competent larvae, and encapsulated juveniles using RNeasy Mini Kit (QIAGEN Vertriebs GmbH, Vienna, Austria). The cDNA from each developmental stage was synthesized using SuperScript ® III reverse transcriptase (Invitrogen GmbH, Karlsruhe, Germany). The homeobox fragments were used to design primers for rapid amplification of cDNA ends (RACE). The RACE was performed with modifications according to Schramm et al. 2000 [66]. For further details on RACE protocol see supplementary data of [67]. The RACE products were cloned by the Topo-TA cloning kit (Invitrogen GmbH, Karlsruhe, Germany) and sequenced using a BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA, USA) and run on an ABI 3130xl DNA analyser automated capillary sequencer.

Orthology assignment and phylogenetic analyses
The initial orthology of the ParaHox genes was tested by searching against GenBank non-redundant protein databases using the BlastX algorithm. The genes were named Gva-Gsx, Gva-Xlox, and Gva-Cdx and deposited in Gen-Bank under accession numbers HM136802, HM136803, HM136804, respectively. Orthology assignment of the genes was made based on phylogenetic analysis. The phy-logenetic analyses were carried out using amino acid sequences. We compiled a ParaHox gene alignment including representatives of bilaterians. Sequences were aligned using the program ClustalX v.2.0.10. First the homeobox region was aligned, then, using the homeobox as an anchor, the flanking regions were aligned and subsequent trimming carried out manually. Bayesian inference on amino acid data using MrBayes version 3.1.1 was applied for orthology analysis, with 2 × 4 Markov chains under the Jones amino acid substitution model [68]. Chains were run for five million generations with a sampling frequency of 1000 generations and the burnin set to 5000 generations.

Whole-mount in situ hybridization
The Maxiscript T7 and SP6 RNA polymerase kit (Ambion, Austin, USA) was used to synthesize the sense and anti-sense probes that were labelled by the Dig RNA labelling kit (Roche Molecular Biochemicals, Vienna, Austria). WMISH was performed with few modifications after Lespinet et al. 2002 [69]. DIG-labelled riboprobes were detected colourimetrically with NBT/BCIP substrates. The details of modifications can be found in [67]. For WMISH, embryos were mounted in 70% glycerol and the expression patterns were documented. For serial-sectioned in situ hybridization, embryos were embedded in Epoxy resin after in situ hybridization according to the standard protocols, and sectioned with a microtome at a thickness of 2 μm. Sections were stained with Eosin using standard histological protocols.

Scanning-electron microscopy
Larvae were fixed in 4% paraformaldehyde (PFA) in 0.1 M saline phosphate buffer (PBS) for 4 h at room temperature or overnight at 4°C, washed three times for 15 min in PBS containing 0.1% sodium azide (NaN3), postfixed in osmium tetroxide (1% in distilled water for 2 h at room temperature), followed by three washes in distilled water, and dehydrated in a graded ethanol/acetone series. Drying was performed either by critical point dryer or chemical drying with HMDS (Hexamethyldisilazane). After drying, the samples were mounted on scanning electron microscopy (SEM) stubs, sputter-coated with gold, and observed with a LEO 1430VP scanning electron microscope.
Authors' contributions LS established the animal cultures, sequenced the ParaHox genes, performed WMISH experiments, and wrote the first draft of the manuscript. GS is responsible for the supervision of the project, the phylogenetic analyses, and editing of the manuscript. Both authors have read and approved the final manuscript.