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1545 Journal of Cell Science 111, 1545-1554 (1998) Printed in Great Britain © The Company of Biologists Limited 1998 JCS4535 Spinalin, a new glycine- and histidine-rich protein in spines of Hydra nematocysts Alexander W. Koch1, Thomas W. Holstein2, Carola Mala3, Eva Kurz3, Jürgen Engel1 and Charles N. David3,* 1Department of Biophysical Chemistry, Biozentrum, University of Basel, Klingelbergstrasse 70, 4056 Basel, Switzerland 2Cell and Developmental Biology, J. W. Goethe-University, Theodor-Stern-Kai 7/75, 60590 Frankfurt am Main, Germany 3Zoological Institut, University of Munich, Luisenstrasse 14, 80333 Munich, Germany *Author for correspondence (e-mail: [email protected]) Accepted 20 March; published on WWW 14 May 1998 SUMMARY Here we present the cloning, expression and immunocytochemical localization of a novel 24 kDa protein, designated spinalin, which is present in the spines and operculum of Hydra nematocysts. Spinalin cDNA clones were identified by in situ hybridization to differentiating nematocytes. Sequencing of a full-length clone revealed the presence of an N-terminal signal peptide, suggesting that the mature protein is sorted via the endoplasmic reticulum to the post-Golgi vacuole in which the nematocyst is formed. The N-terminal region of spinalin (154 residues) is very rich in glycines (48 residues) and histidines (33 residues). A central region of 35 residues contains 19 glycines, occurring mainly as pairs. For both regions a polyglycine-like structure is likely and this may be stabilized by hydrogen bond-mediated chain association. Similar sequences found in loricrins, cytokeratins and avian keratins are postulated to participate in formation of supramolecular structures. Spinalin is terminated by a basic region (6 lysines out of 15 residues) and an acidic region (9 glutamates and 9 aspartates out of 32 residues). Western blot analysis with a polyclonal antibody generated against a recombinant 19 kDa fragment of spinalin showed that spinalin is localized in nematocysts. Following dissociation of the nematocyst’s capsule wall with DTT, spinalin was found in the insoluble fraction containing spines and the operculum. Immunocytochemical analysis of developing nematocysts revealed that spinalin first appears in the matrix but then is transferred through the capsule wall at the end of morphogenesis to form spines on the external surface of the inverted tubule and the operculum. INTRODUCTION which punches a hole in the prey’s integument. Nematocyst discharge is driven by the very high osmotic pressure (15 MPa; Weber, 1989) in the resting capsule, which results from the high concentration of poly-γ-glutamic acid and the corresponding cations in the matrix (Weber, 1990, 1991). The high osmotic pressure in capsules and the extraordinary speed of the evagination process suggest that the capsule wall must have a high tensile strength. Although the structure of the wall is not known in detail, recent work has demonstrated the presence of unusual, small, collagen-like proteins in the wall (Kurz et al., 1991; Engel, 1997) and high-resolution electron microscopy has indicated that these are probably organized in collagen-like fibrils (Holstein et al., 1994). Nothing is known about the structure of the other parts of the nematocyst such as the tubule, spines or operculum, and no other nematocystspecific genes have been isolated or characterized yet. The aim of the present study was to investigate another nematocyst-specific protein, which we have named spinalin. Spinalin is not homologous to any protein in the databases but does have regions with partial homology to loricrins (Mehrel et al., 1990; Steinert et al., 1991) and avian keratins (Whitbread et al., 1991). Both proteins are involved in forming cellular Nematocytes (cnidocytes, ‘stinging cells’) are highly specialized cells characteristic of the phylum Cnidaria. The cells harbor a specialized organelle, the nematocyst, which serves various functions such as defense, the capture of prey and locomotion (Weill, 1934; Mariscal, 1974). Nematocysts consist of a capsule with a stiff wall and an internal tubule which is coiled, pleated and barbed with spines. More than 25 morphologically different nematocyst types have been described and all of them are able to ‘discharge’ after appropriate stimulation (Weill, 1934; Mariscal, 1974). During the discharge the long barbed tubule is everted in one of the fastest known processes in living systems (Tardent and Holstein, 1982). High-speed cinematography has revealed that the initial steps in the eversion of the tubule take less than 10 microseconds and that the spine-bearing part of the tubule is accelerated by up to 40,000 g (Holstein and Tardent, 1984). Spines serve different functions depending on the nematocyst type but all seem to have a high mechanical strength. In particular, stylets, the large spines of stenoteles, have to withstand extreme mechanical stress, since they form the tip Key words: Hydra, Nematocyst, Nematocyte, Spinalin. 1546 A. W. Koch and others products that tolerate high mechanical stress, i.e. the cornified epidermal cell envelope in the case of loricrins, and keratinized epithelial derivatives (claws, nails) in the case of avian keratins. A truncated form of spinalin, named spinalin19K, was recombinantly expressed in Escherichia coli. Using a polyclonal antibody generated against spinalin19K we were able to localize the mature protein to the spines and operculum of nematocyst capsules, both structures having high mechanical strength. MATERIALS AND METHODS Hydra strains and culture conditions Hydra magnipapillata strain sf-1 was used for all experiments. This strain contains temperature-sensitive interstitial cells, which can be removed by warming cultures to 28°C (Sugiyama and Fujisawa, l977). Mass cultures were maintained in M solution at 18°C (Loomis and Lenhoff, 1956) and fed daily with Artemia nauplii. Animals were starved for 24 hours before experiments. cDNA library and differential screening A cDNA library was prepared in lambda gt10 using poly(A)+ RNA isolated from cell fractions enriched for the interstitial cell lineage (Kurz et al., 1991). These cells are smaller than epithelial cells and can be enriched by low-speed sedimentation from suspensions of dissociated Hydra cells (Greber et al., 1992). The cDNA library was differentially screened using radioactively labeled cDNA probes prepared from poly(A)+ RNA isolated from temperature-shocked sf1 (epithelial cell probe) or from an interstitial cell-enriched fraction (interstitial cell probe) as previously described (Kurz et al., 1991). Approximately 13% of clones were interstitial cell-specific; 1% of these clones (6 of 590 isolated clones) represent the spinalin sequence described here. Molecular techniques Commonly used recombinant techniques such as gel electrophoresis, subcloning, growth of plasmids and bacteriophages, nucleic acid isolation, and DNA and RNA blot analysis were carried out according to standard protocols (Sambrook et al., 1989). DNA sequencing was done with the dideoxy chain termination method (Sanger et al., 1977) using sequence-specific primers. Sequence data were analyzed with software from the Wisconsin Package, Version 8.1 (Genetics Computer Group, Madison, WI 53711). The sequence data are available from EMBL/GenBank/DDBJ under accession number AF043907. Cell type separation Intact Hydra were enzymatically dissociated into single cells and various cell types were separated into size fractions using the counterflow centrifugation elutriation with a Beckman J6 centrifuge and the Beckman JE 5.0 elutriator (Greber et al., 1992). The following size fractions were collected: 4000, 3000, 2500, 2200, 1800, 1300 and 1100 rpm at 15 ml flow rate (fractions 1-7) and 1100 rpm at 30 ml flow rate (fraction 8). Cells were eluted with 150 ml in each fraction, and poly(A)+ RNA was isolated using a poly(A)+ isolation kit (Qiagen, 5911 KJ Venlo, Netherlands). Construction of spinalin expression vectors Spinalin cDNA cloned in a pUC19 vector (Stratagene, La Jolla, CA 92037) was used as a template for PCR amplification of spinalin fragments. Different fragments were cloned into the expression vectors pET-15b (Novagen, Madison, WI 53711) and pGEX (Pharmacia, 751 82 Uppsala, Sweden), but only the spinalin19K fragment was purified and further characterized (see Results). Plasmid stability tests were carried out according to the manufacturer’s suggestions (pET system manual, Novagen, Madison, WI 53711). To prepare spinalin19K, a DNA sequence coding for residues 18207 was generated by the use of a 5′ primer introducing a NdeI site (5′ end: CCC CAT ATG AGG CCA TGG GGA CCT GGA T ) and a 3′ primer introducing a stop codon and a BamHI site (3′ end: CCC GGA TCC TTA ATA AAC ATG TAG ACC ACC AAC TC). The amplified product was cloned into the NdeI/BamHI site of the pET15b expression vector, which contains an N-terminal histidine tag followed by a thrombin recognition site (Novagen, Madison, WI 53711). Recombinant expression and purification The pET-15b vector encoding spinalin19K was transformed into E. coli BL21(DE3) cells (Novagen, Madison, WI 53711) and expression of spinalin19K was induced by IPTG. Cells were lysed by sonication and the debris removed by centrifugation (13,000 g for 20 minutes). To purify histidine-tagged spinalin19K crude extract was chromatographed on a nickel sepharose column under denaturing conditions according to the manufacturer’s instructions (Novagen, Madison, WI 53711). Protein samples used for antibody generation were dialyzed against TBS buffer (20 mM Tris, 150 mM NaCl, pH 7.4), containing 2 M urea. Ultrafiltration was performed with Ultrafree-15 centrifugal filters (Millipore, Bedford, MA 01730-2271) having a nominal molecular mass limit of 30 kDa. Spinalin19K was recovered in the flowthrough, as judged by SDS-PAGE. To remove the histidine tag, thrombin digestion was carried out in cleavage buffer (20 mM Tris-HCl, pH 8.4, 150 mM NaCl, 2.5 mM CaCl2, containing 2 M urea) at room temperature for 4 hours using 5 i.u. thrombin per mg protein. Cleavage was monitored by SDS-PAGE as described by Laemmli (1970) using 12×13 cm slab gels. Protein bands were visualized by Coomassie Blue staining and molecular masses were estimated by comparing bands with molecular mass standards (Pharmacia, range 94-14.4 kDa). Antibodies and immunoblotting A polyclonal antibody directed against spinalin19K was generated in rabbits by Eurogentec Bel S.A. (4102 Ougrée, Belgium). Purified spinalin19K, dissolved in 20 mM Tris-HCl, pH 7.4, containing 2 M urea, was directly injected into rabbits. Immunization was carried out following a standard protocol suggested by the manufacturer, using 100 µg of the antigen per injection. For western blot analysis nematocyst proteins or total Hydra proteins were dissolved in SDS-sample buffer, separated by Tricine SDS-PAGE (Schägger and von Jagow, 1987), transferred to nitrocellulose membranes and incubated with spinalin19K antibody (1/1000 dilution) and a secondary goat anti-rabbit antibody coupled to horseradish peroxidase (Bio-Rad, Hercules, CA 94547). The filters were developed employing the ECL chemoluminescence system (Amersham, Arlington Heights, IL 60005). DTT treatment of isolated nematocysts Nematocysts were isolated from whole Hydra tissue as described previously (Kurz et al., 1991; Weber et al., 1987) and treated for 1 hour at room temperature in 100 mM Tris-HCl, pH 9.5, containing DTT at various concentrations ranging from 65 µM to 1 mM. Solubilized nematocyst proteins were separated from insoluble material by centrifugation (5 minutes at 14,000 g) and analyzed by Tricine SDS-PAGE (Schägger and von Jagow, 1987). Immunofluorescence microscopy Whole mount preparations Animals were relaxed in 2% urethane in M solution for 1 minute and fixed in Lavdovsky’s fixative for at least 24 hours (Technau and Holstein, 1995). After washing extensively in PBS buffer (pH 7.2) the polyps were incubated overnight in spinalin19K antibody (diluted 1:1000 in PBS/1% BSA/0.1% sodium azide). To visualize spinalin Spinalin, a new Hydra nematocyst protein 1547 antibody, polyps were incubated for 2 hours with an FITC-conjugated goat anti-rabbit antibody (Boehringer Mannheim Corp., D-68305 Mannheim, Germany) diluted 1:50 in PBS/1% BSA/0.1% sodium azide. Preparations of isolated capsules Nematocysts were isolated as described (Kurz et al., 1991; Weber, 1989). Nematocyst suspensions were induced to discharge on gelatincoated slides by using 0.1 N NaOH (30 minutes), then treated for 1 hour with 65 µM DTT (in 100 mM Tris-HCl, pH 9.5) in order to partly solubilize the spines, and were finally fixed in 4% formaldehyde. After washing extensively in PBS (pH 7.2), slides were incubated overnight in spinalin19K antibody (1:1000 in PBS/1% BSA/0.1% sodium azide), washed and incubated with FITC-conjugated goat anti-rabbit antibody (Boehringer; diluted 1:50 in PBS/1% BSA/0.1% sodium azide). Microscopy and photography All preparations were analyzed using a Zeiss Axiovert microscope equipped with epifluorescence, and photography was performed with Kodak Tungsten 320 (1200 ASA). Confocal laser scanning microscopy was performed using a Leica TCS NT confocal microscope. Electron microscopy Transmission electron microscopy (TEM) analysis was performed as previously described (Holstein, 1981). For fixation the specimens were chilled to 4°C for 1 hour, then double-fixed with 3.5% glutaraldehyde (2-3 hours) and with 1% osmium tetroxide (1-2 hours); the fixatives were buffered with 0.05 M collidine buffer (pH 7.2). Specimens were embedded in epon-araldite with propylene oxide as an infiltration solvent; serial sections (Reichert Ultracut microtome) were stained with uranyl acetate and lead acetate, and examined with a Zeiss EM 9S electron microscope. Scanning electron microscopy (SEM) was performed as previously described (Tardent and Holstein, 1982). Whole Hydra or isolated nematocysts were fixed in 2.5% glutaraldehyde solution containing 50 µM phosphate buffer, which was also used for the post-fixation in 12% OsO2. Samples were dehydrated through a graded series of acetone followed by critical-point drying. Specimens were sputtered with gold and analyzed with a Cambridge S-4 scanning electron microscope (this analysis was done in the laboratory of Prof. Pierre Tardent, Zoological Institute, University of Zürich). RESULTS Isolation of the nematocyte specific cDNA for spinalin We identified nematocyte specific cDNAs by in situ hybridization of individually picked clones. The clones were derived from a cDNA library prepared from cells enriched for the interstitial cell lineage. Interstitial cell-specific clones were identified by differentially screening the library with interstitial cell and epithelial cell probes. From a set of 590 interstitial cell-specific cDNA clones, we identified six clones that crosshybridized with each other and which hybridized in situ to clusters of differentiating nematocytes in the ectoderm throughout the body. In northern blots these clones hybridized to a 1000 bp transcript that was expressed in differentiating nematocytes but not in other cell types (Fig. 1). We have named the protein encoded by these cDNA clones spinalin, based on its localization in the spines of nematocysts (see below). Sequence analysis and putative domain organization of spinalin Fig. 2A shows the nucleotide and deduced amino acid Fig. 1. Northern blot analysis of spinalin expression in fractionated cells. Hydra cells were fractionated by elutriation centrifugation and 1 µg of poly(A)+ RNA from each fraction was applied to the gel. Spinalin transcripts are enriched in fraction 2, which contains small differentiating nematocytes and interstitial cells. The other fractions contain mainly nerve cells (1), large differentiating nematocytes (3/4), gland cells (5), ectodermal epithelial cells (6), endodermal epithelial cells (7) and battery cells (8). sequence of spinalin. There is a single open reading frame of 254 amino acids beginning at nucleotide 126. The start codon ATG is immediately followed by a G residue and preceded at position −3 by an A, which is in agreement with the consensus sequence for eukaryotic translation initiation sites (Kozak, 1987). No homology to other proteins in the database (SWISSPROT; Bairoch and Apweiler, 1997) was found by amino acid sequence comparisons using the entire spinalin sequence. Based on features of the primary sequence and similarities of parts of the spinalin sequence to known protein motifs we propose the domain organization shown in Fig. 2B. The first 17 amino acids fulfil the criteria for a signal peptide (Von Heijne, 1986), indicating that spinalin is synthesized on the endoplasmic reticulum and processed through the Golgi. The presence of the leader sequence is consistent with spinalin being a component of nematocysts, which are the major Golgi products of differentiating nematocytes (Slautterback and Fawcett, 1959; Holstein, 1981). Following removal of the signal peptide, the mature protein consists of 237 amino acids Table 1. Homologies of parts of the spinalin sequence to protein sequences of the SWISS-PROT database* Spinalin region† I I I II II Homology to % identity References Mouse type II keratin (aa 440-562) Human cytokeratin 9 (aa 476-612) Mouse loricrin (aa 154-293) Chick claw keratin (aa 75-104) Chick tropoelastin (aa 32-62) 36 Steinert et al. (1985) 36 Langbein et al. (1993) 29 Mehrel et al. (1990) 70 Whitbread et al. (1991) 48 Bressan et al. (1987) *The FASTA program (Pearson and Lipman, 1988) was used for computer homology searches (Wisconsin Package, Version 8.1, Genetics Computer Group) in the SWISS-PROT database (Bairoch and Apweiler, 1997). †Annotation according to Fig. 2B. 1548 A. W. Koch and others A TTTTTTTTTTTTTGTTGCAGGTGCGAAACACCACAAATGCGTCTCATTTTGCAGAAAGGATA 63 AAACTGTATTCAACAAAAGCAGCATAAGAAATATACTCGCGCCACTATCGGAGGTATTAA 123 AATATGGTGATCGCACAGGCTGCTTTAGTTCTACTTTTGGTTGCTGTTGATGCAAGGCCA 1 M V I A Q A A L V L L L V A V D A R P 183 TGGGGACCTGGATGTGCGGATGGTAGTTATGGTTATGGCGGATGCGGTCACCATCAAGCT 20 W G P G C A D G S Y G Y G G C G H H Q A 243 AACGGTTATGGCGGTGCACACCATGCTGCTGGTTGCTGCAACGGCCTAGCCCATGGCGGA 40 N G Y G G A H H A A G C C N G L A H G G 303 CATCACGGAGGAGCATACGGACAAGCTGCTCATCATGCCGGAGGATATGGTGGTGCATAC 60 H H G G A Y G Q A A H H A G G Y G G A Y 363 GGAGGCCATCATGAGGACCATGGAGAAGATGTTAAAGTAGAGACTCATGGTGTTCACCAT 80 G G H H E D H G E D V K V E T H G V H H 423 ATCGCATCTCATGGTGGTATCCATCATGAACATGGTGGACTTAGAGGAGGACATCATGGA 100 I A S H G G I H H E H G G L R G G H H G Fig. 3. Purification of spinalin19K monitored by SDS-PAGE. The recombinant protein was expressed in E. coli with an N-terminal histidine tag and purified on a nickel affinity column (lane 1). Samples used for antibody generation were further purified by ultrafiltration (lane 2). The histidine tag was removed by thrombin digestion to prepare spinalin19K (lane 3). 483 GCTTACGGAGGACATGGAGCTCATGAGGCAGGTAACTTTGGTCATGTTCATGGCGGACAC 120 A Y G G H G A H E A G N F G H V H G G H 543 TATGGATTAGGTGGAAATCATTATGGTGGCTCCGATTACGGACATGGACACCATGGTGGG 140 Y G L G G N H Y G G S D Y G H G H H G G 603 TATGTGCATGAGCCTTGTCACACTCCATGCCACACTGTTGGTGGATGTGGTGGATGTGGT 160 Y V H E P C H T P C H T V G G C G G C G 663 GGTTATGGACATCTTGGTGGTGTTGGTGGATATGGAACAGGTCTTGGTGGAGTTGGTGGA 180 G Y G H L G G V G G Y G T G L G G V G G 723 GTTGGTGGTCTACATGTTTATAAAAAAGAAACCGTGAAAGGAAAAAAAGTTCAACAAGTT 200 V G G L H V Y K K E T V K G K K V Q Q V 783 ACAAAAGCCGAGGAGGAGAATGAAGATGATGATGATGACGAATCTGAAGATGCTGAAGCT 220 T K A E E E N E D D D D D E S E D A E A 843 GAAAGTGGAAATAGTAGTGATGAAGACATGCCAAATGGAGGTGATTAAATTCAACCTGTA 240 E S G N S S D E D M P N G G D *** 903 AACTCATATATGTATATATAGAATTTTTATTTTGACAATGTAGCATTGTCCCTGTGAAAA 963 ATGGAATAAATTAATATAAAA B 17 R18 I II 154 35 V172 III 16 K207 E223 IV 32 D254 Fig. 2. Spinalin sequence and putative domain organization. (A) Nucleotide and deduced amino acid sequence of Hydra spinalin cDNA. The stop codon is indicated by asterisks. The glycine- and histidine-rich region is single-underlined and the putative polyglycine type II helix is double-underlined. Cysteines are indicated in bold letters. (B) Putative domain organization of spinalin. The first 17 amino acids constitute a signal peptide for translation on the endoplasmic reticulum. The boxed areas represent the proposed domain structure of the mature protein. Borders of the domains are indicated by arrows and the number for the first amino acid of a domain. The putative domains are: a glycine- and histidinerich region (I), a polyglycine type II helical region (II), a lysine-rich region (III) and an acid ‘tail’ (IV). Cysteine residues are indicated by solid vertical bars. and has a calculated molecular mass of 23.7 kDa. It can be divided into four distinct regions. The first part of the sequence (region I, 154 residues) is glycine- and histidine-rich (31 and 21 mol%, respectively). For this region, sequence homologies to mouse loricrin and the C-terminal regions of different cytokeratins were found (Table 1). Region I is followed by a sequence of 35 amino acids (region II) in which, with only few exceptions, two of three amino acids are glycines. This sequence could adopt a polyglycine type II (PGII) conformation with three residues (a, b, c) per turn, where large, hydrophobic residues are located in a positions and glycine residues in b and c positions (Ramachandran et al., 1966; Traub and Yonath, 1966; Lotz and Keith, 1971). This region reveals sequence homology to the glycine-rich regions of chick claw keratin and to chick elastin (Table 1). Region II is succeeded by a stretch of 16 amino acids (region III) with a high content of lysine residues and with weak homology to histone sequences (not shown). The last 32 residues (region IV) represent a region with a high content of negatively charged residues designated as the acid tail. Recombinant expression of spinalin fragments Full-length spinalin could not be overexpressed in E. coli. Although different expression vectors and different conditions were tried, plasmid instability was observed in all cases (see Materials and Methods), which is an indication that the spinalin gene product is toxic for the host cells. Since our aim was to prepare antibodies against spinalin, we also tried to express fragments of the spinalin sequence. Fragments that included the lysine-rich region (region III) could not be expressed in E. coli. However, a large fragment comprising regions I and II, designated spinalin19K, could be successfully overexpressed with a histidine tag for affinity purification. The protein was localized in inclusion bodies and could be purified under denaturing conditions using nickel sepharose affinity chromatography (Fig. 3, lane 1). The histidine-tagged spinalin19K was soluble in TBS buffer containing 2 M urea. It was further purified by ultrafiltration and directly used for antibody production (Fig. 3, lane 2). The histidine tag was cleaved by thrombin to prepare spinalin19K (Fig. 3, lane 3). Solubility tests revealed that spinalin19K tends to aggregate at urea concentrations below 2 M and pH higher than 5, but that the protein is partly soluble in 1 M urea at pH 7.4. Spinalin is present in mature nematocysts In western blots of total Hydra protein the spinalin19K antibody reacted specifically with a protein of apparent molecular mass 24 kDa, which is the expected size of the mature spinalin polypeptide (Fig. 4A, lane 1). Purified Spinalin, a new Hydra nematocyst protein 1549 Fig. 4. Western blot and SDSPAGE analysis of spinalin. (A) Western blot. Samples were solubilized in SDS sample buffer, separated by Tricine SDS-PAGE, transferred to nitrocellulose membranes and immunostained with rabbit anti-spinalin19K antibody. Lane 1, total Hydra protein (not DTT-treated); lane 2, purified nematocysts (DTTtreated); lane 3, supernatant of DTT-treated nematocysts; lane 4, pellet of DTT treated nematocysts. (B) Coomassie Blue-stained gel. Lanes 1- 4 correspond to the lanes in (A). Samples were treated in the same way but approximately five times more protein was applied to the gel for Coomassie Blue staining than for western blotting. nematocysts also contained spinalin (Fig. 4A, lane 2). To determine which structures in nematocysts contain spinalin, purified nematocysts were dissolved in 1 mM DTT under conditions that solubilized the capsule wall but left the stylets and operculum intact. SDS-PAGE confirmed that most nematocyst proteins were in the soluble fraction (Fig. 4B, lane 3) while about 10% of the nematocyst proteins remained insoluble and could be pelleted from the DTT suspension (Fig. 4B, lane 4). Western blotting revealed that the 24 kDa spinalin polypeptide was exclusively present in the pellet fraction (Fig. 4A, lane 4). Microscopic examination demonstrated the presence of stylets (particularly large spines formed in stenotele nematocysts) and opercula as well as amorphous debris in the DTT pellet (T. W. Holstein, unpublished data). These results indicate that spinalin constitutes a DTT-insoluble component of nematocysts and suggest that it is present in stylets or operculum. Localization of spinalin within developing nematocysts We used the spinalin19K antibody to localize spinalin in nematocytes at different stages of nematocyst morphogenesis. In whole mounts of Hydra, nests of differentiating nematocytes were immunostained throughout the gastric region (Fig. 5A). Higher magnification images showed that the antibody stained all nematocyte types and that staining was confined to the developing nematocysts. Nests of desmonemes and isorhizas stained more strongly than nests of stenoteles (Fig. 5B-D). Staining first appeared in immature nematocysts 2-4 µm in length and increased in intensity as cysts increased in size. Following synthesis of the capsule, the ‘preinverted’ tubule is formed as a long thin protrusion from the apical end of the capsule (Holstein, 1981). This preinverted tubule was also strongly stained by the spinalin antibody (Fig. 5B,C). In both the tubule and the capsule, spinalin staining appeared to be confined to the matrix, whereas the capsule wall was not stained. Following completion of the tubule, it inverts and is sequestered within the capsule (Holstein, 1981). Spinalin staining at this stage was still confined to the matrix. The lumen of the inverted tubule, which corresponds to space outside the capsule, was clearly unstained as revealed by appropriately oriented optical sections taken with a confocal microscope shortly after inversion of the tubule (not shown). During the next stages of capsule maturation spinalin staining disappeared progressively from the matrix, became concentrated along the inverted tubule, and then appeared to pass through the tubule wall to fill the lumen of the tubule. This process was not well visualized in the smaller nematocyst types but could be clearly seen in developing stenoteles because of their relatively large size. Fig. 5D shows a stenotele at this stage of development. The stylets (large spines) have developed within the lumen of the inverted tubule and can be seen to form a prominent structure in the center of the capsule. Optical sections of immunostained preparations, taken with a confocal microscope, demonstrated that the stylets are strongly stained by spinalin antibody and that staining has disappeared from the matrix of the capsule at this stage (Fig. 6). This suggests that spinalin is ‘transferred’ from the matrix of the capsule to the lumen of the inverted tubule where stylets develop. Shortly thereafter spinalin immunostaining disappeared completely. Spinalin protein was, however, still present, as shown by western blotting of mature nematocysts (Fig. 4A). TEM images of sections through the center of stenoteles reveal the structure of the stylets in more detail. Stylets appear to be formed as a series of laminar structures projecting into the lumen from the wall of the inverted tubule (Fig. 7A). These lamina are 20-25 nm thick and exhibit a distinct substructure with an outer electron-dense layer (4-5 nm thick) enclosing an inner textured core (15-20 nm thick). Transverse sections through the developing stylet apparatus indicate that the stylets and spines are formed in three sectors spaced around the inner circumference of the inverted tubule and that they consist of a clearly demarcated series of lamina (Fig. 7C). Continued development, however, leads to a dramatic condensation of the lamina to form the mature stylet structure shown in cross section in Fig. 7D and in the DIC image in Fig. 7B. This extreme condensation is consistent with the observation that mature stylets could be pelleted from suspensions of DTTdissociated capsules (see above). 1550 A. W. Koch and others Fig. 5. Immunolocalization of spinalin in Hydra. Whole mounts of Hydra were stained with spinalin19K polyclonal antibody. (A) Head and upper gastric region: nests of differentiating nematocytes are stained in the gastric region. (B) Nest of differentiating desmonemes: matrix and preinverted tubule (arrow) are stained. (C) Nest of differentiating isorhizas: matrix and preinverted tubule (arrow) are stained. (D) Nest of stenoteles at a late stage of morphogenesis after inversion of the tubule; only the operculum and stylet (arrow) are stained. Bars, 200 µm (A); 10 µm (B-D). similar to those observed in developing spines (Fig 7A,C,D). At later stages, these structures are no longer visible, probably due to condensation similar to that observed during spine formation. Fig. 6 shows a reconstruction of serial optical sections through two stenoteles at the stage of operculum formation. A ring of spinalin staining is present at the site of operculum formation (Fig. 6B), suggesting that spinalin was transferred from the capsule matrix through the wall of the inverted tubule to the site of operculum formation at the apical end of the capsule. This process would initially form a ring, which would fill up to become a cap as a result of further extrusion of spinalin. The cap is brightly stained at this stage but loses immunoreactivity as the capsules mature. Fig. 6. Confocal images of two stenotele nematocysts immunostained with spinalin antibody. (A) Apical end of nematocyst is oriented toward top. Stained structures correspond to operculum (top) and stylet (center). The matrix surrounding the stylet is unstained. (B) Nematocyst is tipped slightly, permitting a view into incomplete operculum. Bar, 5 µm. Operculum formation The operculum of nematocysts is formed after inversion of the tubule by a process which appears to be very similar to spine formation (Holstein, 1981). In particular, TEM images indicate the presence of laminar structures in the developing operculum Localization of spinalin in stylets/spines of discharged nematocysts Stylets and spines are most clearly visualized in SEM images of discharged nematocysts. Fig. 8 shows such images for desmonemes, stenoteles and isorhizas. The spines of desmonemes are long thin structures which become oriented down the central axis of the tight coil formed by the tubule following discharge of the nematocyst (Fig. 8A,B). Stenoteles have three large stylets and a series of smaller spines or lamellae located at the base of the everted tubule (Fig. 8C); the tubule appears to be free of spines (not shown). By comparison, in atrichous isorhizas the entire length of the tubule is covered with the small barbs or spines (Fig. 8D). Western blotting (Fig. 4A) indicated the presence of spinalin in mature nematocysts despite the lack of spinalin immunoreactivity. This suggests that spinalin epitopes are Spinalin, a new Hydra nematocyst protein 1551 Fig. 7. TEM of stylets and spines in stenotele nematocysts. (A) Differentiating stenotele shown in longitudinal section. Note the laminar structures consisting of distinct layers 20-25 nm thick. The approximate positions of the cross-sections shown in C and D are indicated. (B) DIC micrograph of mature stenotele nematocyst: the stylets are visible in the center of the capsule. (C) Cross section showing spines in a developing stenotele at the same stage as in A. (D) Cross section of a mature stenotele (equivalent to B) showing spines (spi) in the center and highly compacted stylets (sti) on the outside. Bars, 1 µm (A); 5 µm (B); 500 nm (C,D). masked during formation of spines. To test this idea we searched for a method to restore immunoreactivity to mature capsules and thus directly demonstrate the presence of spinalin in spines. We used discharged nematocysts for these experiments because they permit good visualization of spines. To reduce the extent of condensation of spinalin-containing structures we treated capsules with 0.1 M NaOH and 65 µM DTT (see Materials and Methods). Both treatments were insufficient to dissolve capsules but they did restore spinalin immunoreactivity. The stylets and spines of stenoteles were clearly stained; the capsule and tubule were not stained (Fig. 9A,A′). The tightly coiled tubule of discharged desmonemes was strongly stained by spinalin antibody; the capsule wall was unstained (Fig. 9B,B′). In isorhizas the entire length of the everted tubule was stained whereas the capsule wall was unstained (Fig. 9B,B′). Although the barbs were difficult to resolve by light microscopy, the pattern of immunoreactivity along the tubule of the discharged isorhiza suggests that only the barbs were stained. DISCUSSION Structure of nematocysts The cnidarian nematocyst is one of the most complex secretory products produced by an animal cell (Holstein, 1981; Hessinger and Lenhoff, 1988). Despite a wide diversity of morphological types, the basic structure of all nematocysts is the same: an inner and outer capsule wall, an inverted tubule armed with spines, and an operculum or opercular flaps. When 1552 A. W. Koch and others Fig. 8. SEM images of spines in discharged nematocysts. (A,B) Discharged desmonemes. Spines are long thin structures protruding from the center of the coiled tubule. The two desmonemes shown in (A) are wrapped around a bristle. (C) Discharged stenotele showing the apical end of the capsule and the base of the everted tubule. Three stylets project from the base of the tubule; distal to the stylets are lamellae or small spines. (D) Discharged holotrichous isorhiza. Small barbs or spines decorate the surface of the everted tubule; the capsule is located toward the bottom of the micrograph. Bars, 2 µm (A); 1.5 µm (B); 2 µm (C); 0.5 µm (D). the nematocyst discharges, the inverted tubule everts and the spines are exposed on the outer surface. SDS-PAGE analysis indicates that Hydra nematocysts consist of a complex mixture of proteins varying from 12 to 200 kDa in size (Kurz et al., 1991). Most of the proteins larger than 40 kDa can be removed from nematocysts by treatment with Pronase or dilute DTT. Capsules remain intact and are still refractile in phase contrast under these conditions, although the outer wall is removed by these treatments (Kurz et al., 1991). Atomic force microscopy has demonstrated that the inner wall consists of bundles of fibrils (Holstein et al., 1994) and it has been proposed that these fibrils are polymers of small 12-15 kDa collagen-like proteins (mini-collagens), which are major components of the capsule wall (Kurz et al., 1991; see also Fig. 9. Immunolocalization of spinalin in discharged nematocysts. Isolated nematocysts were treated with 0.1 N NaOH and 65 µM DTT to induce nematocyst discharge and partial denaturation of the spines. Samples were dried on slides and immunostained with spinalin antibody. Phase contrast micrographs (A,B) and immunofluorescence micrographs (A′,B′) show staining of the spines in a discharged stenotele (A,A′) and in two desmonemes and a holotrichous isorhiza (B,B′). No staining was observed with undischarged, isolated capsules (data not shown). Bar, 20 µm. Blanquet and Lenhoff, 1966). Mini-collagens are encoded by a family of nematocyte-specific genes which were first identified in Hydra but have more recently also been found in a reef-building coral, another member of the Cnidaria (Wang et al., 1995). The present experiments have identified a novel 24 kDa protein, spinalin, as an additional component of nematocysts and have shown that it is a constituent of stylets, spines and opercula. The open reading frame includes a signal peptide at the N terminus, indicating that spinalin is synthesized on the endoplasmic reticulum, processed through the Golgi and sorted to the developing nematocyst capsule. Immunocytochemistry with a polyclonal antibody prepared against a 19 kDa fragment of spinalin confirmed these conclusions. The antibody recognized a single protein of 24 kDa in extracts of whole Hydra and in extracts of purified nematocysts. When purified capsules were dissociated with DTT and separated by centrifugation into soluble and insoluble fractions, spinalin was only found in the insoluble fraction, which contained stylets Spinalin, a new Hydra nematocyst protein 1553 and opercula (Fig. 4). Immunofluorescence images confirmed that spinalin is a part of these structures (Figs 5, 6 and 9). Structure and function of spinalin Spinalin shows no overall similarity to other proteins in the databases. Nevertheless, parts of the sequence have features characteristic of particular secondary structures. The Nterminal half of spinalin shows sequence similarity to loricrins (Mehrel et al., 1990; Steinert et al., 1991) and to the glycinerich C-terminal parts of cytokeratins (Steinert et al., 1985, 1991). Loricrins are involved in shaping the cornified cell envelope of epidermal keratinocytes; the terminal parts of cytokeratins are important for bundling keratin filaments by non-covalent supramolecular interactions and by disulfide cross-linking (Steinert et al., 1991). Spinalin, loricrins and the bundling regions of cytokeratins share a high glycine content and other sequence similarities. For loricrins and the terminal parts of cytokeratins ‘glycine loop’ structures were postulated (Steinert et al., 1991). ‘Glycine loops’ are defined by the form X(Y)n, where X is an aromatic or long-chain aliphatic residue and Y is usually glycine, although polar residues like serine, asparagine, arginine and cysteine may also occur. The value of n can range from 1 to 35 and ‘glycine loop’ regions are thought to be formed when several such X(Y)n quasi repeats are tandemly arranged (Steinert et al., 1991). Region I of spinalin fulfils these defining characteristics for ‘glycine loop’ domains, although the high content of histidine in this region of spinalin is unusual. Very recently it has been proposed that related histidine- and glycine-rich regions may be involved in Zn2+ and pH-dependent supramolecular self-association (Coyne et al., 1997). Region II has an even higher glycine content (54%) and contains a large number of Gly-Gly-X triplets, where X stands for long chain aliphatic or aromatic residues. Such sequences can adopt a polyglycine II conformation, which is a left-handed helix with three residues per turn (Rich and Crick, 1961; Lotz and Keith, 1971; Navarro et al., 1995). Such Gly-Gly-X triplets occur in avian keratins, which are highly abundant in claws and scales (Whitbread et al., 1991). Both ‘glycine loop’ domains and polyglycine II helices have been proposed to associate by hydrogen bond formation and to play a role in organizing higher-order structures (Steinert et al., 1991; Traub and Yonath, 1966). In the case of spinalin, the polyglycine-like structure of regions I and II might be also stabilized by disulfide formation between the eight cysteines in these regions. The 19 kDa fragment of spinalin comprising regions I and II was found to be insoluble under physiological condidition. This behavior is similar to observations made on loricrins and cytokeratins and is consistent with the high association potential predicted from the spinalin19K sequence. At 1-2 M urea spinalin19K becomes soluble, although circular dichroism measurements (A. W. Koch, unpublished data) suggest that it is still partly folded under these conditions. Association of such partially folded structures could be the mechanism by which spinalin molecules form higher order structures. More conformational data, however, are needed to unravel the details of spinalin association. Electron micrographs (Fig. 7) suggest that spines are very densely packed structures. The observation that spinalin immunoreactivity disappears during spine formation also supports this conclusion. Tight packing and network formation are probably important for the function of spines. Stylets, the large spines in stenoteles, are needed to puncture the cuticle of prey organisms when capsules discharge (Tardent and Holstein, 1982), while spines in desmonemes and isorhizas appear to function as barbs binding the discharged nematocyst to prey or to the substrate. These results, together with observations on the structure and function of loricrins, cytokeratins and avian keratins (see above), suggest that polyglycine-like structures may be particularly well adapted to formation of organelles having a high mechanical strength. Morphogenesis of nematocyst capsules and formation of stylets Immunocytochemical staining of Hydra polyps demonstrated that spinalin first appears in developing nematocysts in the body column at an intermediate stage of differentiation. Spinalin is homogeneously distributed in the matrix at this stage and also extends into the tubule when it forms on the apical end of the capsule (Fig. 5B,C). Following inversion of the tubule into the capsule, spinalin becomes concentrated in the lumen of the inverted tubule, which corresponds topologically to the external surface of the everted tubule. Spines develop on this external surface (Holstein, 1981). At the apical end of the capsule spinalin also appears to be transferred from the matrix to the outside of the tubule in order to form the operculum. Upon further maturation, spinalin staining ceases in both the operculum and the spines, although western blot analysis demonstrated that spinalin continues to be present in these structures (Fig. 4A). We interpret this loss of immunoreactivity in the mature operculum and spines as an indication of extreme condensation of spinalin in these structures. This suggestion is supported by the observation that spinalin immunoreactivity could be restored by mildly denaturing conditions (Fig. 9). The mechanism by which spinalin is transferred through the tubule wall is not known. However, the wall at this stage is still ‘soft’, since it can be deformed during the process of fixation. At a slightly later stage the wall becomes rigid and is no longer deformed by fixation. At this stage the capsule wall also becomes impermeable to polypeptides since poly-γ-glutamate polymers are retained inside capsules (Weber, 1990). Even at this stage, however, the wall can and does stretch with the increase in osmotic pressure which results from the synthesis of poly-γ-glutamate in the matrix (Weber, 1990, T. W. Holstein and C. N. David, unpublished results). It is interesting to note that a similar pattern of staining has been observed for the H22 antigen during capsule formation. The monoclonal antibody H22 recognizes a 100 kDa protein in the outer capsule wall (Kurz et al., 1991). Like spinalin, the H22 antigen first appears in the matrix and then moves through the inner capsule wall to form the outer wall (T. W. Holstein, unpublished results). Some of the H22 antigen also appears to move through the wall of the tubule and to become localized in spines. The relationship between spinalin structure and spine morphogenesis requires further investigation. Although the spines of stenoteles, desmonemes and isorhizas are quite different in shape (Fig. 8), they all contain spinalin (Fig. 9). Whether other proteins are also present in spines and contribute to formation of these different shapes is not clear from the 1554 A. W. Koch and others present experiments. However, an attractive alternative hypothesis is that differences in the folding of the inverted tubule in different nematocyst types affect the transfer of spinalin across the tubule wall and hence the morphology of spines. Which molecular features of spinalin are important for spine morphogenesis is also unclear. At present we can only speculate that the highly charged regions III and IV might be involved in solubilization of the protein when transported to or located in the matrix while the glycine- and histidine-rich region I might be involved in pH- or Zn2+-dependent assembly of higher order structures. Changes in pH have indeed been observed during capsule morphogenesis (Gunzl, 1968) and could regulate such processes. We are very grateful to Dr A. Brancaccio for stimulating discussions and helpful suggestions. We thank Timo Zimmermann for help with image processing and Petra Snyder for technical assistance with the cell type separation and immunocytochemistry. This work was supported by the Deutsche Forschungsgemeinschaft and the Swiss National Science Foundation. REFERENCES Bairoch, A. and Apweiler, R. (1997). The SWISS-PROT protein sequence data bank and its supplement TrEMBL. Nucleic Acids Res. 25, 31-36. Blanquet, R. and Lenhoff, H. M. (1966). Disulfide-linked collagenous protein of nematocyst capsules. Science 145, 152-153. Bressan, G. M., Argos, P. and Stanley, K. K. (1987). Repeating structure of chick tropoelastin revealed by complementary DNA cloning. Biochemistry 26, 1497-1503. Coyne, K. J., Qin, X.-X. and Waite, J. H. (1997). Extensible collagen in mussel byssus: a natural block copolymer. Science 277, 1830-1832. Engel, J. (1997). Versatile collagens in invertebrates. Science 277, 1785-1786. Greber, M. J., David, C. N. and Holstein, T. W. (1992). A quantitative method for separation of living Hydra cells. Roux’s Arch. Dev. Biol. 201, 296-300. Gunzl, H. (1968). Über die Reifung der Nesselkapseln bei Dipurena reesi Vannucci (Hydrozoa). Zeitsch. f. Zellforsch. 89, 509-518. Hessinger, D. A. and Lenhoff, H. M. (1988). The Biology of Nematocysts. Academic Press, San Diego. 593 pp. Holstein, T. W. (1981). The morphogenesis of nematocytes in Hydra and Forskålia: An ultrastructural study. J. Ultrastruct. Res. 75, 276-290. Holstein, T. W. and Tardent, P. (1984). An ultrahigh-speed analysis of exocytosis: nematocyst discharge. Science 223, 830-832. Holstein, T. W., Benoit, M., von Herder, G., Wanner, G., David, C. N. and Gaub, H. E. (1994). Fibrous mini-collagens in Hydra nematocytes. Science 265, 402-404. Kozak, M. (1987). An analysis of 5′-noncoding sequences from 699 vertebrate messenger RNAs. Nucleic Acid Res. 15, 8125-8148. Kurz, E. M., Holstein, T. W., Petri, B. M., Engel, J. and David, C. N. (1991). Mini-collagens in Hydra nematocytes. J. Cell Biol. 115, 1159-1169. Langbein, L., Heid, H. W., Moll, I. and Franke, W. W. (1993). Molecular characterization of the body site-specific human epidermal cytokeratin 9: cDNA cloning, amino acid sequence, and tissue specificity of gene expression. Differentiation 55, 57-71. Laemmli, U. K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680-685. Loomis, W. F. and Lenhoff, H. M. (1956). Growth and sexual differentiation of Hydra in mass culture. J. Exp. Zool. 132, 555-574. Lotz, B. and Keith, H. D. (1971). The crystal structures of poly(LAla-GlyGly-Gly)II and poly(LAla-Gly-Gly)II. J. Mol. Biol. 61,195-200. Mariscal, R. N. (1974). Nematocysts. In Coelenterate Biology. Reviews and New Perspectives (ed. L. Muscatine and H. M. Lenhoff), pp. 129-178, Academic Press, New York. Mehrel, T., Hohl, D., Rothnagel, J. A., Longley, M. A., Bundman, D., Cheng, C., Lichti, U., Bisher, M. E., Steven, A. C., Steinert, P. M., Yuspa, S. H. and Roop, D. R. (1990). Identification of a major keratinocyte cell envelope protein, loricrin. Cell 61, 1103-1112. Navarro, E., Tereshko, V., Subirana, J. A. and Puiggali, J. (1995). Incorporation of diacids into the polyglycine II structure: model studies. Biopolymers 36, 711-722. Pearson, W. R. and Lipman, D. J. (1988). Improved tools for biological sequence comparison. Proc. Nat. Acad. Sci. USA 85, 2444-2448. Ramachandran, G. N., Sasisekharan, V. and Ramakrishan, C. (1966). Molecular structure of polyglycine II. Biochim. Biophys. Acta. 112, 168170. Rich, A. and Crick, F. H. C. (1961). The molecular structure of collagen. J. Mol. Biol. 3, 483-506. Sambrook, J., Fritsch, D. M. and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual, 2nd edition. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Sanger, F., Nicklen, S. and Coulson, A. R. (1977). DNA sequencing with chain termination inhibitors. Proc. Nat. Acad. Sci. USA 74, 5463-5467. Schägger, H. and von Jagow, G. (1987). Tricine-sodium dodecyl sulfatepolyacrylamide gel electrophoresis for the separation of proteins in the range from 1-100 kDa. Anal. Biochem. 166, 368-379. Slautterback, D. B. and Fawcett, D. W. (1959). The development of the cnidoblasts of Hydra: an electron microscopy study of cell differentiation. J. Biophys. Biochem. Cytol. 5, 441-451. Steinert, P. M., Parry, D. A., Idler, W. W., Johnson, L. D., Steven, A. C. and Roop, D. R. (1985). Amino acid sequences of mouse and human epidermal type II keratins of Mr 67,000 provide a systematic basis for the structural and functional diversity of the end domains of keratin intermediate filament subunits. J. Biol. Chem. 260, 7142-7149. Steinert, P. M., Mack, J. W., Korge, B. P., Gan, S.-Q., Haynes, S. R. and Steven, A. C. (1991). Glycine loops in proteins: their occurrence in certain intermediate filament chains, loricrins and single-stranded RNA binding proteins. Int. J. Biol. Macromol. 13, 130-139. Studier, F. W. and Moffatt, B.-M. (1986). Use of bacteriophage T7 polymerase to direct selective high-level expression of cloned genes. J. Mol. Biol. 189, 113-130. Sugiyama, T. and Fujisawa, T. (l977). Genetic analysis of developmental mechanisms in Hydra. Sexual reproduction of Hydra magnipapillata and isolation of mutants. Dev. Growth Diff. 19, 187-200. Tardent, P. and Holstein, T. W. (1982). Morphology and morphodynamics of stenotele nematocyst in Hydra attenuata. Cell Tissue Res. 224, 269-290. Technau, U. and Holstein, T. W. (1995). Head formation in Hydra is different at apical and basal levels. Development 121, 1273-1282. Traub, W. and Yonath, A. (1966). Polymers of tripeptides as collagen models. J. Mol. Biol. 16, 404-414. Von Heijne, G. (1986). A new method for predicting signal sequence cleavage sites. Nucleic Acids Res. 14, 4683-4690. Wang, W., Omori, M., Hayashibara, T., Shimoike, K., Hatta, M., Sugiyama, T. and Fujisawa, T. (1995). Isolation and purification of a minicollagen encoding a nematocyst capsule protein from a reef-building coral, Acropora donei. Gene 152, 195-200. Weber, J. (1989). Nematocysts (stinging capsules of Cnidaria) as Donnanpotential-dominated osmotic system. Eur. J. Biochem. 184, 465-476. Weber, J. (1990). Poly(γ-glutamic acid)s are the major constituents of nematocysts in Hydra (Hydrozoa, Cnidaria). J. Biol. Chem. 265, 9664-9669. Weber, J. (1991). A novel kind of polyanions as principal components of cnidarian nematocysts. Comp. Biochem. Physiol. 98A, 285-291. Weber, J., Klug, M. and Tardent, P. (1987). Some physical and chemical properties of purified nematocysts of Hydra attenuata Pall. (Hydrozoa, Cnidaria). Comp. Biochem. Physiol. 88B, 855-862. Weill, R. (1934). Contribution à l’étude des cnidaires et des leurs nématocystes. Trav. Sta. Zool. Wimereux 10-11, 1-701. Whitbread, L. A., Gregg, K. and Rogers, G. E. (1991). The structure and expression of a gene encoding chick claw keratin. Gene 101, 223-229.