HS (heparan sulfate) proteoglycans are key regulators of vital processes in the body. HS chains with distinct sequences bind to various protein ligands, such as growth factors and morphogens, and thereby function as important regulators of protein gradient formation and signal transduction. HS is synthesized through the concerted action of many different ER (endoplasmic reticulum) and Golgi-resident enzymes. In higher organisms, many of these enzymes occur in multiple isoforms that differ in substrate specificity and spatial and temporal expression. In order to investigate how the structural complexity of HS has evolved, in the present study we focused on the starlet sea anemone (Nematostella vectensis), which belongs to the Anthozoa, which are considered to have retained many ancestral features. Members of all of the enzyme families involved in the generation and modification of HS were identified in Nematostella. Our results show that the enzymes are highly conserved throughout evolution, but the number of isoforms varies. Furthermore, the HS polymerases [Ext (exostosin) enzymes Ext1, Ext2 and Ext-like3] represent distinct subgroups, indicating that these three genes have already been present in the last common ancestor of Cnidaria and Bilateria. In situ hybridization showed up-regulation of certain enzymes in specific areas of the embryo at different developmental stages. The specific mRNA expression pattern of particular HS enzymes implies that they may play a specific role in HS modifications during larval development. Finally, biochemical analysis of Nematostella HS demonstrates that the sea anemone synthesizes a polysaccharide with a unique structure.
HSPGs [HS (heparan sulfate) proteoglycans] are ubiquitous macromolecules that are present on the cell surface and in the extracellular matrix. HSPGs consist of a protein core with at least one or more covalently attached HS chains . The HS chains interact with a vast number of protein ligands, such as proteases, extracellular matrix molecules, growth factors, cytokines, chemokines and morphogens . Such interactions regulate the stability and activity of the protein ligands and affect morphogen gradients. Therefore HSPGs play essential roles in a variety of developmental, morphogenic and pathogenic processes. The interactions between the negatively charged HS chains and protein ligands depend on complex patterns of negatively charged carboxyl and sulfate groups along the HS chains that can differ in composition between cell types and animal species (reviewed in ).
The expression and function of HS is well documented, however, most studies are from bilaterian model organisms and not much is known about more basal animal phyla. The Cnidaria is one of oldest metazoan phyla and is among the simplest animals that display epithelial tissue organization [4,5]. The starlet sea anemone (Nematostella vectensis) belongs to the Anthozoa, and is considered to have retained more ancestral features [6,7]. It has been established previously to be a versatile model organism that has become of increasing interest for use in studying developmental pathways. The availability of powerful molecular and genetic tools, the sequencing of the Nematostella genome and the accessibility of Nematostella development to laboratory manipulations make Nematostella an ideal organism for studies in evolutionary genomics [4,6]. The embryonic development of Nematostella includes the formation of a hollow blastula and gastrulation via invagination [8,9]. After a free-swimming planula stage, the larvae metamorphose into primary polyps with four tentacles, a gastric cavity and longitudinal mesenteries . To date, there have been no studies of HS structure or function in Nematostella. However, HS has previously been shown to be present in Hydra, which belongs to a more derived class of the Cnidarians, the Hydrozoa [11,12].
The fine structure of HS is generated during its biosynthesis in the Golgi apparatus by the concerted action of the HS biosynthetic enzymes (Figure 1). HS chains are elongated by the Ext (exostosin) family of glycosyltransferases and are initially composed of alternating GlcA (D-glucuronic acid) and GlcNAc (N-acetylglucosamine) units. During synthesis, the growing HS chain is modified by several enzymes: the C5-epimerase [that converts GlcA to IdoA (L-iduronic acid)] and sulfotransferases (that add sulfate groups at different positions) [13,14]. The extent of these reactions varies, resulting in enormous structural heterogeneity and the creation of different domains within HS chains. Thus the final molecular design of HS is characterized by domain structures that are highly modified and domains that are less modified or not modified at all (Figure 1) [1,13,15]. After synthesis, the HS chains are further modulated at the cell surface by a family of Sulfs (6-O-endosulfatases) .
Schematic illustration of the modifications of HS chains
In vertebrates, the enzymes involved in the synthesis and modification of HS can occur in several genetic isoforms that differ in substrate specificity and tissue expression (for reviews, see [14,17]). To date, two GlcA/GlcNAc transferases, four NDSTs (N-deacetylase/N-sulfotransferases), three 6OSTs (HS 6-O-sulfotransferases), seven 3OSTs and two Sulfs have been identified in mammals. In contrast, only one C5-epimerase and one 2OST have been identified in mammals. The isoform diversity and tissue expression patterns of the biosynthetic enzymes most probably influence the tissue-specific differences in the fine structure of HS and thus regulate HS–growth factor interactions in various developmental processes .
The requirements for sulfated HS chains for the development of metazoans is undisputable, as seen in Drosophila melanogaster, Caenorhabditis elegans, Mus musculus and other model organisms , but the evolutionary background to more subtle HS variations is still largely unknown. The enzymes involved in HS biosynthesis are highly conserved throughout evolution, but the number of isoforms differs between organisms. Has the structural complexity of HS evolved together with its biological function? Is the tissue variation in HS structure evidence of a tight regulation of HS biosynthesis, or does it reflect a lack of evolutionary pressure controlling HS biosynthetic enzyme expression and/or activity?
To be able to answer these questions structural information about HS is needed. Therefore we have studied the expression of HS biosynthetic enzymes in N. vectensis and characterized its HS chains. By performing in silico searches of the Nematostella genome and EST (expressed sequence tag) libraries we have identified members of all HS biosynthetic enzyme families and the cell surface HS Sulfs. Biochemical analysis of labelled polysaccharides isolated from Nematostella embryos revealed that the sea anemone synthesizes a polysaccharide that has many characteristics of vertebrate HS, but in addition also contains structures not commonly found in vertebrate HS.
Cloning of Nematostella HS biosynthetic enzymes
The human cDNA sequences of HS biosynthetic enzymes and Sulf were used for tBLASTN homology searches of a Nematostella EST library and the Nematostella genome browser (http://genome.jgi-psf.org/cgi-bin/runAlignment?db=Nemve1&advanced=1). The identities of the obtained Nematostella sequences with corresponding mammalian proteins were confirmed by homology searches against the non-redundant NCBI peptide database (http://www.ncbi.nlm.nih.gov/BLAST/). Primers were designed from the EST sequences and used to amplify cDNA fragments from oligo(dT)-primed first-strand cDNA derived from embryonic and larval Nematostella total RNA. The primers used for amplification are available by request. PCR was carried out using ExTaq polymerase (TaKaRa) and an annealing temperature of 56 °C. After the addition of overhangs using GoTaq (Promega), the PCR fragments were cloned into pGEM-T (Promega) or pCRII (Invitrogen) vectors and DNA sequenced.
Stage-specific RT-PCR (reverse transcription-PCR)
Total RNA was isolated from Nematostella embryos at different developmental stages using the RNeasy Kit (Qiagen) and genomic DNA was eliminated by digestion with DNaseI (Promega). Isolated RNA (0.7 μg) was used to synthesize stage-specific oligo(dT)-primed cDNAs using Superscript III reverse transcriptase (Invitrogen). After cDNA synthesis, the RNA templates were digested with RNaseH (New England Biolabs). Reverse-transcribed control cDNAs were prepared in parallel without reverse transcriptase.
In situ hybridization
Isolation of metabolically [35S]sulfate-labelled polysaccharides from Nematostella
Mixed stages of Nematostella embryos (1–4-days-old) were collected in one 100-mm-diameter plate and washed with fresh medium (33% autoclaved seawater) three times. For radiolabelling, the embryos were transferred to a new culture dish with 10 ml of medium containing 400 μCi/ml Na2[35S]O4 (1494 Ci/mmol; GE Healthcare). After incubation for 24 h at room temperature (21 °C), the culture medium was removed and frozen at −20 °C, and excess radioactive isotope was eliminated from the embryos by five successive washes in 10 ml of PBS, followed by centrifugation for 5 min at 800 g. The pelleted embryos were digested with 10 mg of a non-specific protease (P8811; Sigma) in 500 μl of digestion buffer [0.1 M Tris/HCl (pH 8.0) containing 0.002 M CaCl2] at 55 °C for approx. 16 h. Thereafter an additional aliquot (5 mg) of enzyme in 500 μl of digestion buffer was added and the digestion continued for ≥8 h. To release O-linked sugar chains (including HS) from the attached peptide resulting from protease digestion, the digest was treated with 0.5 M NaOH overnight at 4 °C and was then neutralized using 4 M HCl. The polysaccharides were recovered by gel chromatography on a column (1×200 cm) of Sephadex G-25 superfine (GE Healthcare) in 0.2 M ammonium bicarbonate and desalted by freeze-drying.
Isolation of non-labelled HS disaccharides from Nematostella
Approx. 400 mg of frozen Nematostella embryos from different embryonic stages were subjected to protease digestion [100 mg of protease in 20 ml of 0.1 M Tris/HCl (pH 8.0) and 0.002 M CaCl2], followed by alkali treatment as described above. After centrifugation (38000 g for 60 min), the supernatant was applied to a 4 ml column of DEAE–Sephacel (GE Healthcare) equilibrated with 50 mM Tris/HCl (pH 8.0) and 50 mM NaCl. The column was first washed with equilibrium buffer, then with 50 mM acetate buffer (pH 4.0) containing 50 mM NaCl, followed by 50 mM acetate buffer (pH 4.0) containing 150 mM NaCl. The pH was increased by washing with 50 mM Tris/HCl (pH 8.0) and 150 mM NaCl, and finally DEAE-bound material was eluted with 50 mM Tris/HCl (pH 8.0) and 1 M NaCl. Fractions (1 ml) were collected and analysed for uronic acid content by the carbazole reaction . Carbazole-positive fractions were pooled and desalted on a PD-10 column.
Chemical 3H-labelling of disaccharides from Nematostella, bovine intestinal HS and Ascidian heparin
Nematostella carbazole-positive material, bovine intestinal HS and Ascidian Heparin (Iduron) (approx. 25 μg each) were dried, deaminated with nitrous acid (pH 1.5) (cleavage at N-sulfated glucosamine residues), followed by reduction with 0.5 mCi NaB[3H]4 (24 Ci/mmol; GE Healthcare) to yield reducing-terminal 3H-labelled aManR (2,5-anhydro-D-mannitol) residues, followed by reaction with excess unlabelled NaBH4. The labelled oligosaccharides were separated from unincorporated radioactivity by gel chromatography on a column of Sephadex G-15 (1×170 cm) in 0.2 M ammonium bicarbonate. Nematostella-labelled oligosaccharides were further separated into disaccharides and larger oligosaccharides on a column of Sephadex G-50 Superfine (GE Healthcare) in 0.2 M ammonium bicarbonate. Fractions corresponding to disaccharides were pooled (see Supplementary Figures S1 and S2 at http://www.BiochemJ.org/bj/419/bj4190585add.htm) and desalted by freeze-drying.
Characterization of 35S-labelled HS from Nematostella
Labelled polysaccharide samples were applied to a 1 ml column of DEAE–Sephacel equilibrated with 0.05 M Tris/HCl (pH 7.5) and 0.05 M NaCl. The column was washed with >10 ml of equilibration buffer and eluted with a linear gradient (100 ml), ranging from 0.05 M to 1.2 M NaCl in 0.05 M Tris/HCl (pH 7.5), at a flow rate of 0.5 ml/min. In analytical runs, the labelled polysaccharide samples were mixed with an internal standard of Escherichia coli K5 capsular polysaccharide (0.6 mg), and an aliquot (50 μl) from each effluent fraction was subjected to uronic acid determination using the carbazole method . The remaining material was monitored for radioactivity. Preparative DEAE anion-exchange chromatography was performed using the same column without the added internal standards. Instead the column was calibrated with 14C-labelled K5 polysaccharide before the preparative run. The 14C-labelled K5 polysaccharide was prepared as described previously . 35S-labelled material was pooled as indicated in the legend to Figure 5 and desalted on PD-10 columns (Sephadex G-25, GE Healthcare) in 10% (v/v) ethanol. The size of Nematostella HS was analysed on a Superose 12 column (GE Healthcare).
Disaccharide compositional analysis of Nematostella HS
The disaccharides obtained after deamination were analysed by anion-exchange HPLC using a Whatman Partisil 10-SAX column eluted with stepwise increasing concentrations of aqueous KH2PO4 at a rate of 1 ml/min using HexA-[3H]aManR disaccharides, with O-sulfate groups in different positions, as reference compounds . The preparation of the Ascidian standard disaccharides is shown in Supplementary Figure S2. The purified disaccharide preparations were subjected to sequence analyses using bovine β-glucuronidase (which also contained α-iduronidase activity) (Sigma) and recombinant human iduronate 2-sulfatase (a gift from Professor John Hopwood, Women's and Children's Hospital, Adelaide, Australia) as described previously . The presence of α-iduronidase activity in the bovine β-glucuronidase from Sigma was determined by incubating the enzyme preparation with the appropriately labelled disaccharide substrates, GlcA–aManR6S (where 6S is 6-O-sulfate group) and IdoA–aManR6S. The enzyme preparation removed both the terminal GlcA (β-glucuronidase activity) and IdoA (α-iduronidase activity) and converted both types of disaccharide substrates to aManR6S (results not shown).
Isolation of related HS biosynthesis enzyme sequences from Nematostella
To identify Nematostella orthologues of the enzymes involved in HS biosynthesis, BLAST searches using human cDNA sequences were carried out using a Nematostella EST collection  and the Nematostella genome browser . Genes identified in this way included three EXT/EXTL (Ext-like) genes, three NDST genes (termed NDSTa–c), one C5 epimerase gene, two 2OST genes (termed 2OSTa and 2OSTb), one 3OST gene, one 6OST gene and one Sulf gene.
Using PCR, we cloned the partial or full-length cDNAs of the Nematostella HS biosynthesis enzymes. A comparison of the deduced amino acid sequence of the Nematostella 2OSTa protein with the corresponding sequences in mouse, Drosophila and C. elegans revealed that the amino acid sequence for 2OSTa is evolutionarily conserved (Figure 2A). Although relatively low sequence similarities were observed in the N-terminal region, the amino acid sequences of the central region containing the two putative binding sites for the sulfate donor PAPS (3′-phosphoadenosine 5′-phosphosulfate) are highly conserved. The predicted amino acid sequence for Nematostella Sulf is homologous to the mouse Sulfs throughout the N-terminal region, which includes the structural motifs required for the formation of the active site and the conserved cysteine residue (Figure 2B and Supplementary Figure S3 at http://www.BiochemJ.org/bj/419/bj4190585add.htm) that undergoes post-translational modification into formylglycine, thus forming the catalytic active enzyme  and the hydrophilic domain that characterizes the vertebrate and C. elegans Sulfs .
Amino acid sequence alignment among members of the HS 2OST (A) and HS 6-O endosulfatase (Sulf) (B) family
Phylogenetic analysis of the three NvNDST genes and the two 2OST genes showed that these genes most probably represent lineage-specific duplications, which are frequently observed in Nematostella [6,26]. In contrast, the three EXT/EXTL genes can be assigned to the EXT1, EXT2 and EXTL3 groups, indicating that these three genes have already been present in the last common ancestor of all eumetazoans. The phylogenetic trees for NDSTs, 2OSTs and EXT/EXTL can be found in Supplementary Figures S4–S6 (at http://www.BiochemJ.org/bj/419/bj4190585add.htm).
Temporal mRNA expression of HS biosynthetic enzymes
To investigate if HS biosynthetic enzymes are transcriptionally regulated during Nematostella embryonic development, we analysed the temporal expression profile of the HS biosynthetic enzymes and Sulf by RT-PCR on stage-specific RNA (Figure 3). With the exception of 2OSTb and NDSTc, mRNA transcripts for all enzymes were expressed at all developmental stages. Although the expression levels of EXT1, EXT2, NDSTa, NDSTb, C5 epimerase and 2OSTa were rather uniform at all stages, 6OST, 3OST and Sulf displayed elevated expression levels during blastula and gastrula stages. 2OSTb and NDSTc transcripts were not expressed during the unfertilized egg stage and commenced at the gastrula stage, suggesting that they are not maternally provided, unlike the other enzymes.
RT-PCR expression analysis of HS biosynthesis enzymes and the extracellular Sulf during different stages of Nematostella development
Spatial mRNA expression of HS biosynthetic enzymes by in situ hybridization
Whole-mount in situ hybridizations were carried out in order to analyse the spatial expression patterns of Nematostella HS biosynthesis enzymes. Diffuse and weak signals were detected for most enzymes throughout embryonic development (results not shown). However, EXT1, NDSTc and Sulf showed distinct and interesting expression patterns. Sulf was expressed uniformly until the onset of gastrulation, when it became enriched in the invaginating tissue that forms the endoderm (Figures 4A and 4B). The enhanced expression in the endoderm was maintained during the planula stage (Figure 4C). At the late planula stage, shortly before the onset of and during metamorphosis, Sulf expression becomes particularly strong in four endodermal spots at the oral end of the larvae, marking the sites of tentacle evagination (Figures 4D–4F). The strong expression in the tentacle endoderm was maintained also after metamorphosis into primary polyps (Figure 4G). Thus elevated expression of Sulf correlated with two prominent morphogenetic events: invagination of the endoderm and evagination of the tentacles.
Expression patterns of NvSulf, NvExt1 and NvNDSTc during Nematostella development
The expression patterns of EXT1 and NDSTc at the late planulae larvae stadium were also quite interesting. In addition to elevated expression in the endoderm, EXT1 showed a distinct expression in the aboral ectoderm (Figure 4H). This is the site where the major larval sense organ, the apical organ, forms. NDSTc also showed an elevated expression in the endoderm, but the expression was not uniform: it was stronger in the oral endoderm and decreased towards the aboral pole (Figure 4I).
Characterization of HS in Nematostella embryos
To analyse HS in Nematostella, mixed stage embryos were metabolically labelled with [35S]sulfate for 24 h. Labelled macromolecules were extracted by proteolytic digestion, followed by β-elimination, and separated from unincorporated [35S]sulfate by gel filtration. Approx. 17% of total labelled material was susceptible to chemical degradation specific for HS and the related polysaccharide heparin (results not shown).
The anionic properties of the isolated 35S-labelled macromolecules were analysed by anion-exchange chromatography. A major portion of the labelled material was eluted ahead of a non-sulfated polysaccharide standard, composed of a GlcA–GlcNAc polymer, identical to the unmodified mammalian HS, but approx. 17–18% eluted somewhat later than the standard polysaccharide (Figure 5A). The labelled material was pooled as indicated in Figure 5(A), desalted and subjected to depolymerization at N-sulfated GlcN residues. The P1 (pool 1) and P2 subfractions were resistant to degradation (results not shown) and were not further analysed, whereas the P3 subfraction was completely degraded to disaccharides (Figure 5B, lower panel), indicating that this fraction contained Nematostella HS. The size distribution of 35S-labelled Nematostella HS was determined by gel-filtration chromatography. Compared with the elution position of standard polysaccharides, the peak maximum for the sulfated chains corresponded to a mean molecular mass of ∼40 kDa (Figure 5B, upper panel).
Analysis of Nematostella HS chains
The disaccharides were analysed for composition by anion-exchange HPLC. Disaccharides from control bovine HS contains a mixture of variously O-sulfated species, typical for HS (Figure 6A), whereas the corresponding Nematostella disaccharides had a very different pattern, lacking the sulfated GlcA-containing disaccharide (disaccharide 1) and with a marked increase in trisulfated disaccharides, corresponding to more than 60% of total O-sulfated disaccharides compared with 30% in mammalian HS (Figure 6). A portion of the trisulfated disaccharides from Nematostella eluted in the position corresponding to IdoA2S–GlcNS6S (where NS is N-sulfate group) in the intact HS chain. However, the majority of the Nematostella trisulfated disaccharides eluted somewhat later than this standard disaccharide, indicating that this peak most probably contained at least two different disaccharides.
Disaccharide composition of N-sulfated domains in HS
To further characterize the trisulfated disaccharide X (indicated with an asterisk in Figure 6), 3H-labelled HS disaccharides were prepared from Nematostella embryos by radiolabelling of nitrous acid cleavage products with NaB[3H]4 as described in the Experimental section and previously . We then determined the number and position of the sulfate groups on disaccharide X by enzymatic digestion of the 3H-labelled disaccharides with IdoA-2-sulfatase and a commercial β-glucuronidase that also contained α-iduronidase activity. Upon digestion of the standard bovine disaccharides with the commercial bovine β-glucuronidase, we found that both peaks 1 and 2 (GlcA–GlcNS6S and IdoA–GlcNS6S respectively) in Figure 6(A) disappeared and instead a major peak was observed in the position of a mono-sulfated monosaccharide corresponding to GlcNS6S in the intact HS chain (results not shown). Hence this enzyme preparation removes both non-sulfated terminal GlcA and IdoA and is referred to as β-glucuronidase/α-iduronidase. After digestion with β-glucuronidase/α-iduronidase, Nematostella disaccharide 2 disappeared, indicating that this disaccharide corresponds to IdoA–GlcNS6S (Figure 6D). The trisulfated disaccharides were sensitive to iduronate 2-sulfatase and generated disaccharides 2 and 5 that eluted in the same positions as the authentic standards corresponding to IdoA–GlcNS6S and IdoA–GlcNS3S respectively (Figure 6E, see also Figure 7). The enzyme activity of the iduronate 2-sulfatase is significantly affected by the sugar unit adjacent to the IdoA residue and exhibits higher activity towards substrates with O-sulfation on the adjacent glucosamine unit . Therefore peak 3 (corresponding to IdoA2S–GlcNS) was only slightly reduced. When we treated the disaccharides with iduronate 2-sulfatase followed by β-glucuronidase/α-iduronidase, both peaks 2 and 5 were significantly decreased. Since the iduronate 2-sulfatase is specific for IdoA and does not remove sulfates from GlcA, the results of the present study demonstrate that the structure of disaccharide X was IdoA2S–GlcNS3S. This unusual disaccharide accounted for approximately two-thirds of the trisulfated disaccharides and 40% of the total sulfated disaccharides.
Structures of sulfated disaccharides identified in Nematostella
HS chains interact with a large number of proteins, including growth factors, and thereby function as potent and specific regulators of cell signalling in a variety of circumstances. HS sulfation patterns within the saccharide chains are critical for these interactions . Although the pattern may not be unique for each protein ligand, the cell- and tissue-specific regulation of HS sulfation patterns appears to be important for HS function [13,29]. Nematostella expresses FGFs (fibroblast growth factors), Wnts and other growth factors known to use HS as a co-receptor. One may speculate that HS structures exhibit a similar regulation during Nematostella development as in higher-evolved organisms. The individual HS biosynthesis enzymes that participate in the formation of the complex HS chains have been studied thoroughly in mammalian systems. However, not much is known about the evolution of the biosynthetic machinery or whether the enzymes involved in HS biosynthesis generate the same polysaccharide in simple non-bilaterian animals. We therefore decided to utilize the sea anemone Nematostella to combine an investigation of the expression profile of Nematostella HS biosynthetic enzymes with a structural analysis of Nematostella HS. We show that the transcripts for all members of the HS biosynthetic machinery and one Sulf enzyme were expressed in Nematostella. Two enzymes, NDST and 2OST, were expressed by three and two independent genes respectively. The three NDSTs clearly form a separate cluster, indicating that they arose by lineage-specific gene duplications. One of the two 2OSTs (Nv2OSTb) clusters with moderate support with HS 2OSTs from other organisms and Nv2OSTa, also shows similarity to the chondroitin sulfate 2OST (see Supplementary Figure S5). Since only a partial sequence is available for this gene, we cannot currently exclude that it represents a highly divergent 2OST involved in chondroitin sulfate biosynthesis. In both vertebrate and invertebrates, only one isoform of the 2OST has been described, whereas four different NDSTs are found in vertebrates. Whether both Nematostella 2OST transcripts encode active enzymes is not known at present. However, the restricted and developmentally regulated expression of Nv2OSTa and Nv2OSTb suggest specific roles for these two isoforms during Nematostella development. Interestingly, the single Sulf enzyme expressed by Nematostella is strongly up-regulated in the endoderm during two morphogenetic processes: the invagination of the endoderm during gastrulation and the evagination of the tentacles during metamorphosis. In both processes, Sulf expression overlaps with that of secreted growth factors; in the invaginating endoderm with two Nematostella homologues of BMPs (bone morphogenetic proteins) NvDpp (where Dpp is decapentaplegic) and NvBmp5–8 [30,31], and in the tentacle endoderm with NvWnt5 and NvWnt11 [32,33]. Although this co-expression is suggestive, it remains to be shown whether Sulf indeed affects signalling by these morphogens. Two mammalian Sulf enzymes have been identified recently [25,34]. Both are HS Sulfs and their enzymatic activities have been linked to the regulation of the activity of HS-dependent growth factors and morphogens [16,35,36]. Removal of 6-O-sulfates by the Sulf enzyme may, depending on the signalling molecule, either enhance or inhibit its biological activity.
Interestingly, in addition to low expression levels in the endoderm, there was a distinct expression of NvEXT1 in the aboral ectoderm where the major larval sense organ is located. The formation and maintenance of this organ requires FGF signalling, and NvFGFa1 and NvFGFa2 are expressed from a very early time point in this region [37,38]. At the planula stage (after the organ has formed), there are also BMPs and BMP antagonists expressed in the apical organ, but their function is not clear [19,30,31]. Based on the level of amino acid conservation, EXT1 and EXT2 and three additional proteins, designated EXTL1, EXTL2 and EXTL3, form the mammalian EXT family . In the Nematostella genome, we could identify three genes, NvEXT1, NvEXT2 and NvEXTL3, that were homologous to the mammalian EXT genes. In Drosophila, the same three Ext genes have been identified [40–42], whereas only the EXT1 and EXTL3 orthologues rib-1 and rib-2 respectively, exist in C. elegans [43,44]. Although several studies have established that EXT1, EXT2 and EXTL3 are involved in HS chain elongation [40–42,45–49], the functions of EXTL1 and EXTL2 remain unclear. Furthermore, both EXTL1 and EXTL2 seem to be absent in more basal animals, suggesting that they are not essential for HS synthesis.
One of the HS core proteins found in Nematostella, NvGlypican 4/6, is strongly expressed in the apical organ, although the domain is broader than that of NvEXT1, NvFGFa1 and NvFGFa2, NvDpp and NvBMP5–8 (F. Rentzsch and U. Technau, unpublished work). Furthermore, another of the glypican core proteins, NvGlypican 3/5, shows a similar expression in the endoderm to NvNDSTc, but has additional expression domains in the oral ectoderm and pharynx (F. Rentzsch and U. Technau, unpublished work).
The enzymes involved in HS biosynthesis are highly conserved although the number of isoenzymes is different in vertebrates and invertebrates. The more evolved the organisms are, the more complex the biosynthetic machinery appears to be, involving multiple isoforms of enzymes with apparently similar substrate specificities. Interestingly, however, increasing the complexity of the biosynthetic machinery does not necessarily produce more highly sulfated polysaccharides. In contrast, polysaccharides from several marine species, e.g. molluscs and ascidians, are characterized by highly sulfated complex structures [11,50]. Disaccharide analysis of HS isolated from Nematostella indicated that this polysaccharide contains an unusually high amount of IdoA2S–GlcNS3S (∼40% of total sulfated disacccharides). Small amounts of this disaccharide have been detected previously in bovine glomerular basement membrane (14%)  and Styela plicata (8–9%) .
HS structural variation could have evolved as a dynamic way for cells to regulate when and which signals they can receive, in the same way as cells express different types of cell surface receptors. Despite the effort of many investigators the regulatory mechanisms behind the heterogeneity of HS structure are still poorly understood. Further studies using Nematostella and other marine invertebrates should provide new information regarding the interplay between the different HS enzymes in creating HS structure diversity and the evolutionary significance of the HS sulfation pattern. For instance, it will be very interesting to determine whether IdoA2S–GlcNS3S modification of Nematostella HS leads to the formation of specific growth factor binding sites and to analyse the role of HS during Nematostella developmental patterning in vivo.
In summary, the results of the present study demonstrated that members of all of the enzyme families involved in the generation and modification of HS are present in Nematostella. The distinct expression patterns of HS synthesis and modification enzymes suggest that differential modification of HS contributes to patterning and morphogenesis in Nematostella, and were thus employed for these processes already at a very early evolutionary stage. Furthermore, biochemical analysis of Nematostella HS showed that the sulfation patterns of the chains are structurally unique. Our present results are a first step toward utilizing Nematostella as a tool for understanding the role of evolution in generating HS structural diversity.
We thank Dr John Hopwood (Women's and Children's Hospital, Adelaide, Australia) for providing the recombinant human iduronate 2-sulfatase.
bone morphogenetic protein
expressed sequence tag
fibroblast growth factor
- 2OST etc.
HS 2-O-sulfotransferase etc.
- P1 etc.
pool 1 etc.
- 2S etc.
2-O-sulfate group etc.
This work was supported by the University of Bergen; the Research Council of Norway [grant number 101936]; the Norwegian Cancer Society [grant number HS02-2007-0060]; and the Meltzer Foundation [grant number 480523] (to M. K.-G.). The work in the U. T. and F. R. laboratories was funded by the Sars Centre/Unifob AS (Bergen University Research Foundation) core budget.
These authors contributed equally to this work.
Present address: Boston Biomedical Research Institute, 64 Grove Street, Watertown, MA 02472, U.S.A.
Present address: Developmental Biology Section, Department of Theoretical Biology, University of Vienna, Althanstrasse 14, 1090 Vienna, Austria.