The structure of mollusc larval shells formed in the presence of the chitin synthase inhibitor Nikkomycin Z
© Schönitzer and Weiss; licensee BioMed Central Ltd. 2007
Received: 03 July 2007
Accepted: 06 November 2007
Published: 06 November 2007
Chitin self-assembly provides a dynamic extracellular biomineralization interface. The insoluble matrix of larval shells of the marine bivalve mollusc Mytilus galloprovincialis consists of chitinous material that is distributed and structured in relation to characteristic shell features. Mollusc shell chitin is synthesized via a complex transmembrane chitin synthase with an intracellular myosin motor domain.
Enzymatic mollusc chitin synthesis was investigated in vivo by using the small-molecule drug NikkomycinZ, a structural analogue to the sugar donor substrate UDP-N-acetyl-D-glucosamine (UDP-GlcNAc). The impact on mollusc shell formation was analyzed by binocular microscopy, polarized light video microscopy in vivo, and scanning electron microscopy data obtained from shell material formed in the presence of NikkomycinZ. The partial inhibition of chitin synthesis in vivo during larval development by NikkomycinZ (5 μM – 10 μM) dramatically alters the structure and thus the functionality of the larval shell at various growth fronts, such as the bivalve hinge and the shell's edges.
Provided that NikkomycinZ mainly affects chitin synthesis in molluscs, the presented data suggest that the mollusc chitin synthase fulfils an important enzymatic role in the coordinated formation of larval bivalve shells. It can be speculated that chitin synthesis bears the potential to contribute via signal transduction pathways to the implementation of hierarchical patterns into chitin mineral-composites such as prismatic, nacre, and crossed-lamellar shell types.
Molluscs were among the first organisms on earth that were able to produce highly organized calcium carbonate composite materials with unique structural features and remarkable materials properties [1, 2]. Even today, the mollusc shell surprises us with new concepts for understanding biomineralization processes .
Chitin, a linear homopolymer consisting of β-(1–4)-linked N-acetyl-D-glucosamine subunits, plays an important role in mollusc shell formation. The presence of chitin in mollusc shell matrices is well documented in the literature [4–6]. Recently, it has been demonstrated that chitin fulfils various structural tasks in the formation of larval shells of the bivalve mollusc Mytilus galloprovincialis . In the adult stage, the fibres of chitin and certain crystallographic axes of aragonite are aligned in mollusc nacre [8, 9]. Based on a cryo-TEM study, Levi-Kalisman and colleagues suggested that chitin is the ordered component of the decalcified nacre matrix, whereas a silk-like protein gel environs the chitin sheets . Mineralizing proteins are either attached to the core sheets or distributed within the silk like protein gel. Thus, the chitin and the silk together form a regular lamellar structure. Subsequently, certain mollusc shell protein fractions induce aragonite formation within this lamellar β-chitin and silk framework . The involvement of a transient amorphous mineral precursor phase in the formation of aragonite biominerals is currently discussed based on the presence of such a phase in calcite forming sea urchins and aragonite forming mollusc larvae [3, 12–14]. These observations raised new questions regarding the role of structural biopolymers such as chitin in the formation of shell microtextures . As recently discovered, the calcitic prismatic layer of the bivalve mollusc Atrina rigida (Pteriomorphia, Pinnidae) is the result of a structural interplay between chitin and the mineral phases .
Chitin synthases are transmembrane glycosyltransferases that are responsible for the enzymatic synthesis of chitin . The representatives of these enzymes in molluscs contain a N-terminal myosin motor head domain . For many years, the chitin synthases have been studied mainly in fungi [19–23]. Chitin synthases are localized either in the plasma membrane or in so-called chitosomes. Chitosomes are intracellular membrane vesicles that host the chitin synthase or its zymogenic precursor form and may contain preformed chitin in their lumen during vesicle transport prior to their fusion with the cytoplasmic membrane [24, 25].
Despite its strong ecological impact, the structure of the much more complex insect chitin synthase , which is closely related to the C-terminus of the mollusc enzyme , has attracted significant research interest in recent years . Mutagenesis experiments and RNAi approaches demonstrated that chitin does not only fulfil a structural role in the arthropod exoskeleton. In fact, it guides the development of invertebrates such as Drosophila, Tribolium, and Caenorhabditis [28–30].
There are several options in order to interfere chemically with the biosynthesis of chitin [31, 32]. One prominent example are small-molecule inhibitor drugs such as nucleosid-peptides that structurally imitate the UDP-activated chitin precursor substrate, UDP-N-acetylglucosamine (UDP-GlcNAc) and thus inhibit the chitin synthases of fungi and insects in a competitive manner [33–35]. Polyoxins, first time described in 1965, and Nikkomycins, first described in 1976, belong to this class of inhibitors that are produced by certain strains of Streptomyces, such as S. tendae and S. cacaoi [36, 37]. Remarkably, these nucleosid antibiotics did not interfere with protein synthesis (reviewed by ). Nevertheless, a broad profile of inhibitory potency, ranging from Ki = 0.02 μM to Ki = 3.5 μM, was reported for chitin synthases across different eukaryotic phyla in vitro . The commercially available chitin synthase inhibitor NikkomycinZ is well suited for non-invasive cell biological and biochemical studies in vivo: It is cell permeable due to active transport mechanisms that are employed by eukaryotic cells for the uptake of dipeptides. The fact that this substance is built from unusual amino acids counts for its resistance against intracellular proteolytic degradation [38–40].
In a previous study, we investigated the chitinous matrix of larval shells of the marine bivalve mollusc Mytilus galloprovincialis . The way how the chitin is distributed throughout the organic framework of the larval shell, how its distribution and structure change with the time-course of larval development, and the reflection of typical functional elements of larval shells such as the hinge teeth, the bivalve symmetry, and the radial and tangential shell growth front within the observed chitin pattern suggest an integrative functional role for chitin synthesis in biomineral composite formation, from the subcellular to the organism scale. These observations motivated us to study the biochemical process of chitin synthesis in relation to the formation of the larval shell. Here, we present the first polarized light microscopic and scanning electron microscopic data from shells of the marine bivalve mollusc Mytilus galloprovincialis that were cultivated in the presence of the chitin synthase inhibitor NikkomycinZ.
By using NikkomycinZ in vivo as a small-molecule inhibitor drug we demonstrated that even a partial inhibition of chitin synthesis interferes dramatically with the biosynthesis of larval mollusc shells at various hierarchical levels. The presented data suggest that chitin formation may provide one of the regulatory targets of "biologically controlled"  mineralization, thus forming a multi-link between the constantly developing mollusc shell and the mantle epithelial cells. This might lead to a novel understanding of the role of the epithelial cells that, at one and the same time, guide larval development, secrete the shell precursor components, and monitor the dynamics of the various shell formation interfaces.
We chose an in vivo approach for investigating the impact of chitin synthesis in mollusc larval shell formation. Therefore, we reared mollusc larvae in the presence of NikkomycinZ, a small-molecule drug that is well-known to inhibit enzymatic chitin synthesis in a competitive manner in vivo and in vitro. We used polarized light video microscopy for the in vivo investigations and scanning electron microscopy imaging of extracted shell material prepared from NikkomycinZ treated Mytilus galloprovincialis larvae of various age.
Shell development under the conditions of the test (control experiments)
Additional File 1: Video clip of spawning female. Demonstration of spawning female and eggs prior to fertilization. (MPG 2 MB)
Additional File 2: Video clip of spawning male. Demonstration of spawning male and sperm prior to fertilization. (MPG 4 MB)
Additional File 3: Video clip of fertilization procedure. Demonstration of fertilization procedure and collection of fertilized eggs. (MPG 2 MB)
Additional File 4: Video clip of large scale larvae culture (10 l). Demonstration of the culture conditions of larvae in artificial seawater at 10 l-scale. (MPG 365 KB)
Additional File 5: Video microscopy of 2-day larvae. Video microscopic (32× objective) demonstration of the behaviour of 2-day larvae. Note that the larvae are already in the early veliger stage and have a D-shaped shell. (MPG 3 MB)
Additional File 6: Video microscopy of 5-day larvae. Video microscopic (32× objective) demonstration of the behaviour of 5-day larvae. The size of the velum and the motility of the larvae increased. Note that the hinge of the larval shell became straight. (MPG 3 MB)
Additional File 7: Video microscopy of 8-day larvae. Video microscopic (32× objective) demonstration of the behaviour of 8-day larvae. The motility of the larvae is similar to 5-day larvae. The size of the larval shells slightly increased. Most of the shells are still in D-shape. (MPG 3 MB)
Additional File 8: Video microscopy of 12-day larvae. Video microscopic (20× objective) demonstration of the behaviour of 12-day larvae. Note that the size of the shell increased, and the larvae switched from the D-shape stage to the umbo stage of shell formation. (MPG 2 MB)
Additional File 9: Video microscopy of 31-day larvae. Video microscopic (20× objective) demonstration of the behaviour of 31-day larvae. Larvae are in different developmental stages, such as veliger larvae and pediveliger larvae. The latter are indicated by the developing foot and a degenerate, vanishing velum that is retracted into the shell. (MPG 3 MB)
Additional File 10: Video microscopy of 36-day larvae. Video microscopic (20× objective) demonstration of the behaviour of 36-day larvae. At this age, the velum disappeared, and metamorphosis into the adult was completed as indicated by the functional foot. (MPG 5 MB)
Statistical effects of NikkomycinZ treatment observed in larvae cultures
The effects of NikkomycinZ on the cultivation of larvae were evaluated by estimating the percentage of affected individuals per culture well. It was not possible to determine accurate numbers due to the high motility of the individuals. Based on values obtained from different experimenters, we estimated a deviation of ~20%. This deviation includes also effects of variable numbers of individuals per culture well. We defined test conditions to be appropriate when larvae in the control culture without NikkomycinZ represented viability and motility comparable to a regular 10 l scale culture, which allowed us to grow larvae until metamorphosis into the adult stage occurred. When NikkomycinZ was applied in concentrations of > 25 μM, most of the individuals did not survive for the duration of the test. When NikkomycinZ was applied in concentrations of less than 5 μM, no significant effects on viability or increased morphological changes were observed. We found NikkomycinZ suitable to be applied in concentrations of 5 μM and 10 μM in order to compromise between the detrimental effects on viability and inducing significant effects on shell formation.
As demonstrated in Fig. 2, the early larval stages are more affected by NikkomycinZ treatment than the older larvae. If NikkomycinZ is added to a 2-day culture, the survival rate is below 20% on the 8th day (Fig. 2a). A comparable decrease in the survival rate is obtained when 5-day larvae are treated with NikkomycinZ for seven days (Fig. 2b). 8-day larvae were not affected comparably as much on the average (Fig. 2c). About 50% more individuals than in the control culture died within seven days in the presence of NikkomycinZ. As shown in Fig. 2d, no significant decrease in survival rate was observed during the first three days of NikkomycinZ incubation in 12d old cultures, whereas all younger larval stages did show an effect after three days.
The following criteria were defined in order to classify shell phenotypes that were observed in NikkomycinZ treated cultures: undulated shell edge, bilaterally asymmetrical valves, lack of umbo stage after 12 days, extraordinary small shell relative to the size of the organism, no straight hinge line, increased transparency of the shell, fractured shell. A larval shell was considered affected by NikkomycinZ treatment if one or more of the described phenotypes were applicable. Only motile individuals were taken into account. Fig. 3 shows the percentages of individuals with abnormally developed shells grown in the presence of NikkomycinZ with respect to the criteria described above. With progressive incubation time, the NikkomycinZ treatment from day 2 on induced a steady increase of abnormally developed individuals in the range of three times higher than observed in the control culture (Fig. 3a). Similar results were obtained from cultures incubated from day 5 on. Three days of a treatment with 10 μM NikkomycinZ slightly increased the number of observed shell defects in the population (Fig. 3b). No significant effects of the NikkomycinZ treatment with respect to shell development were observed at the binocular microscope level at 80 × magnification for the older stages of 8–15 days and 12–15 days (Fig. 3c,d).
Morphological effects of NikkomycinZ treatment on the organism scale (> 100 μm)
Additional File 12: Video microscopy of 8-day larvae after 6 days of NikkomycinZ treatment. Video microscopic (32× objective) demonstration of the behaviour and phenotype of 8-day larvae after 6 days of treatment with the chitin synthase inhibitor NikkomycinZ. The overall motility of the larvae decreased as compared to untreated control cultures. Note that NikkomycinZ appears to have a dramatic effect on shell development. The size of the shells is comparable to 2-day larvae. The hinge is not a straight line, and shell edges appear irregular or undulated. (MPG 4 MB)
Additional File 13: Video microscopy of 12-day larvae after 7 days of NikkomycinZ treatment. Video microscopic (32× objective) demonstration of the behaviour and phenotype of 12-day larvae after 7 days of treatment with the chitin synthase inhibitor NikkomycinZ. The overall motility of the larvae decreased as compared to untreated control cultures. Note that NikkomycinZ appears to have a dramatic effect on shell development. The formation of the umbo was prevented (see additional file 8). The size of the shells is comparable to 5-day larvae. The most prominent features were curved hinges and malformed shell edges. Shells of some individuals were too small to host the organism completely. (MPG 2 MB)
Additional File 14: Video microscopy of 12-day larvae after 4 days of NikkomycinZ treatment. Video microscopic (32× objective) demonstration of the behaviour and phenotype of 12-day larvae treated from the 8th day on with the chitin synthase inhibitor NikkomycinZ. Even four days of treatment with NikkomycinZ have similar effects on shell development of individuals as described in additional file 13. Note that less affected individuals were highly motile and therefore out of focus in this data set. (MPG 2 MB)
Additional File 15: Video microscopy of 15-day larvae after 7 days of NikkomycinZ treatment. Video microscopic (32× objective) demonstration of the behaviour and phenotype of 15-day larvae treated from the 8th day on with the chitin synthase inhibitor NikkomycinZ. Even in later developmental stages, NikkomycinZ induced characteristic effects on the shell development of living individuals. Note that the previously straight hinge (see additional files 7 &8) appears now curved. This indicates that NikkomycinZ interferes with the remodelling of the hinge region. The fact that shells are much smaller than in the untreated control cultures (additional file 8) suggests that either the solubility of the newly formed shell is increased, or lateral shell growth is blocked by the chitin synthase inhibitor drug. Note that also shell remnants of larvae that died at undefined age are present in this data set. (MPG 5 MB)
Structural effects of NikkomycinZ treatment on the tissue scale (10 μm – 100 μm)
Ultrastructural effects of NikkomycinZ treatment on the subcellular to cellular scale (<1 μm – 10 μm)
Inner surface layer – lateral growth and thickening of the larval shell
These shells (Fig. 6d) also lack the granular layer, which usually covers the flake-like material on the inner shell surface (also compare Fig. 6c). The uncovered flakes (Fig. 6d) have evident interspaces between them, which also decrease towards the shell edge.
Outer surface layer – interference of NikkomycinZ with periostracum formation
Hinge formation and hinge ultrastructure
The influence of NikkomycinZ on the development of hinge teeth is strongly dependent on the time frame of NikkomycinZ application. In older developmental stages, hinge tooth formation is not completely inhibited by NikkomycinZ. However, we observed structural alterations that might predominantly affect the functionality of the youngest formed hinge teeth. For comparison, the hinge of a larval shell from the control culture without NikkomycinZ is shown in Fig. 9e. Even the youngest teeth in the centre of the hinge represented at least small protuberances. The transition region between the smaller (Fig. 9e, arrow) and the bigger hinge teeth is shown in detail in Fig. 9f. All teeth exhibited smoothly curved edges. Each tooth spans the whole cross-section (thickness) of the shell's hinge throughout the complete line. The picture changes, once NikkomycinZ was present in the culture medium during shell growth (Fig. 9g,h). It is obvious that especially the smaller hinge teeth (Fig. 9g) were not well formed. As observed more clearly at higher magnification (Fig. 9h, arrow indicates the zoom-in position in Fig. 9g), such hinge teeth did not span the whole thickness of the shell. The edges of the hinge teeth did not confine cubic elements. The same structural features applied to the bigger hinge teeth. The close-up view of the small hinge teeth (Fig. 9h) revealed that the building blocks of the hinge region in larvae grown in the presence of NikkomycinZ consisted of small, flat, and sharp-edged prismatic building blocks with a planar upper surface. Such sharp-edged building blocks were never observed in hinges of control larvae, which were smoothly curved-edged, and which were equipped with a fine-dispersed globular surface cover (Fig. 9b,f). These results indicate a direct interference of NikkomycinZ with the formation and remodelling of hinge teeth in the bivalve larvae of Mytilus galloprovincialis throughout development.
Effects of NikkomycinZ treatment on the crystallization and molecular self-assembly scale (Å – 100 nm)
Effects of NikkomycinZ treatment on the physical properties and functionality
The primary aim of this study was to analyze the impact of chitin synthesis on mollusc shell formation. If the cellular regulation of chitin synthesis is related to mollusc shell formation, the partial inhibition of enzymatic chitin synthesis by a competitive enzyme inhibitor such as NikkomycinZ is supposed to induce structural alterations in the shell. Since also mollusc larvae of the marine bivalve species Mytilus galloprovincialis express the gene for a myosin chitin synthase specifically in the shell forming tissue , and since larval shells of this species consist of considerable amounts of chitin , these larvae provide an ideal system for studying possible effects of NikkomycinZ on the development of mollusc shells. Especially effects related to the transformation of amorphous calcium carbonate (ACC) into aragonite can be observed more clearly during larval shell development , while providing the option to extrapolate the results for a better understanding of the adult shell formation process . The study of larval shell formation processes is of fundamental interest since both, the mineralogy and the structural organization of the larval shell, are evolutionary highly conserved among diverse bivalve taxa. The fact that a "classification of larvae by hinges results in a classification closely parallel to a classification of adults" as recognized by Rees in 1950  indicates that the development and biomineralization of the hinge (provinculum) is of special evolutionary interest. The results presented in this study indicate that NikkomycinZ predominantly interferes with the mineralization process at the hinge region, suggesting that the mollusc myosin chitin synthase may have evolved as an effective regulatory element between cell differentiation and tissue mineralization.
Chitin synthesizing enzymes are complex transmembrane proteins that, unlike other glycosyltransferases , have an evolutionary highly conserved amino acid motif in their intracellular active site, which is identical in unicellular fungi as well as in more complex organisms (arthropods, molluscs), and which is well-known to accept UDP-GlcNAc as the only substrate . Conceivably, chitin synthases are highly sensitive for the structural UDP-GlcNAc-analogue NikkomycinZ that inhibits chitin synthesis in a competitive manner and is therefore applied as a pesticide against fungi and arthropods . This drug was also used to test the biological impact of the insect chitin synthase and found to be lethal for first instar larvae at their transition to second instar at concentrations of > 1 μM . It is self-evident that NikkomycinZ can be applied for structural studies on mollusc shell formation in vivo in only sublethal concentrations. We determined a suitable working concentration of NikkomycinZ in the range of 5 μM to 10 μM for larvae of Mytilus galloprovincialis. It is important for the interpretation of the obtained results to keep in mind that chitin synthesis is not supposed to be completely inhibited in that concentrations range. The exact degree of enzyme inhibition can hardly be estimated in vivo, as the quantity and enzymatic activity of mollusc chitin synthases could not be determined, and the exact rate of NikkomycinZ uptake into the larval tissue by mollusc peptide transport systems is unknown. Nevertheless, a phenomenological comparison between larvae cultured in the absence and presence of NikkomycinZ revealed that even a partial inhibition of chitin synthesis at 5 μM – 10 μM inhibitor induced drastic structural alterations in the larval shells. Taking the limited stability of NikkomycinZ in the culture medium (sea water) into account , it can be assumed that the effective working concentration of NikkomycinZ continuously decreased from the start value at the time of sea water replacement to the next sea water exchange, allowing the animals to compensate temporarily the detrimental effects on cell metabolism. In fact, the calnexin and calreticulin pathways [54, 55] provide eukaryotic cells with effective mechanisms to circumvent detrimental effects that an activated sugar analogue such as NikkomycinZ might have on the posttranslational modification of glycoproteins . Still, it has to be taken into consideration that certain extracellular biomineralization proteins, which are typically glycosylated (see  for review), might have been affected by NikkomycinZ as well, and therefore may contribute to certain effects on shell formation observed in this study. Furthermore, an altered level of chitin deposition could well interfere with intracellular signal transduction pathways that regulate the expression of biomineralization genes.
We optimized the NikkomycinZ treatment of the mollusc larvae in order to maintain a survival rate as documented in Fig. 2. Due to the fact that also chitin synthesis was not intended to be completely inhibited, the culture may have been kept under conditions that allow proteins glycosylation to occur above the critical threshold. We observed that even those organisms were extremely vital that were only partly covered with a shell. Therefore, we concluded that chitin synthesis is the much more NikkomycinZ sensitive process. Schlüter dealt with the same problem many years ago , inspired by the observations of Holst and colleagues that Nikkomycin was predominantly lethal for arthropod larvae in the course of moulting . In-between the moulting cycles, glycoproteins must be equally important for the survival of the larvae. Obviously, the synthesis of glycoproteins was significantly less affected in these invertebrate animals. Schlüter observed Nikkomycin induced ultrastructural defects in the procuticular region in moulting beetle larvae. Thus, the enzymatic synthesis of chitin appears to be a rather specific target of the Nikkomycins.
One of the general problems of experiments with mollusc larvae cultures is that a 100% survival rate can never be achieved, due to the fact that with proceeding development, a genetic defect of any individual might become lethal or may at least lead to an abnormal phenotype. Therefore, the really significant effect of NikkomycinZ on mollusc larvae populations was that after a certain period of incubation time, none of the animals had a "healthy" phenotype such as observed in populations that have not been in contact with the drug. Malformed animals were not always completely detected as such, mainly due to the low resolution of binocular microscopy, and due to the high motility of the individual larvae and the movements of velar tissue and cilia (see also Additional Files 5, 6, 7, 8 and 12, 13, 14, 15). Therefore the numbers of abnormalities within a NikkomycinZ treated population may in reality be higher than estimated (see Fig. 2 and Fig. 3 for details). Certain malformations were detectable in a definite way only by using scanning electron microscopy, which was not applicable for an in vivo screening.
We regarded it important to study morphological effects of the NikkomycinZ treatment by video microscopy. This method allowed us to demonstrate the viability of the organism along with the induced morphological alterations. Our primary aim was to exclude those individuals from the analysis that were affected by NikkomycinZ in any respect other than shell formation. On the other hand, it is conceivable that malformed shells will also interfere with the regular development of a bivalved organism. Therefore, the criteria as described in the results section were defined in order to group the phenotypes observed only in the motile organisms. In later experimental stages, shells of lethargic organisms were also taken into consideration, if a certain morphological effect was exposed there more clearly.
During the time-course of this study, it was recognized that certain categories of malformations were visualized by video microscopy only in a later developmental stage (after a longer treatment with NikkomycinZ), whereas the same features were observed by SEM comparably earlier (after short treatment with NikkomycinZ). This phenomenon becomes especially clear in Figure 5, when comparing the respective age of specimens in SEM and video microscopy images. Undoubtedly, this is one of the limitations of the in vivo approach on shell formation. Nevertheless, it can be speculated that the number of individual shells affected by NikkomycinZ was even higher than suggested by the data presented in Figure 3.
Our observations presented here may also contribute to an implementation of mechanical signal transduction into mollusc shell formation concepts. The mechanical functionality of the hinge gets lost whenever the formation of hinge teeth (see Fig. 9) is prevented. This might influence the formation of the valves as well, if one assumes a mechanical feedback mechanism that is responsible for the regulation of shell formation. If forces during valve closure are responsible for a correct valve formation, a direct correlation with mineral deposition should be guaranteed. An asymmetry of the valves (Fig. 4b) or an irregularly shaped shell edge (Fig. 5a,b, Fig. 6b,d), hinge line (Fig. 4a, Fig. 5c, Fig. 11e), or hinge teeth (Fig. 9d,h) will inevitably prevent a correct force application. Consequently, this mechanical feedback at the organism's level will interfere with stimulating the correct mechanically induced cell answer such as deposition of multiple organic and inorganic shell precursor components that are necessary for correct valve structuring. As the mollusc myosin chitin synthase strongly interferes with the actin cytoskeleton (unpublished results), multiple bottom-up effects are expected to be induced once the central coordination gets lost. Thus, the chitin synthase inhibitor NikkomycinZ has obviously the ability to interfere directly with one of the central players of this biomechanical coordination process.
Our continued research on larval shell biomineralization in Mytilus galloprovincialis aims to address the issue of chitin synthesis and structural modifications in the context of regulatory cascades that coordinate shell formation and the development of the shell forming tissue within the whole body plan in vivo.
The small-molecule drug NikkomycinZ, a competitive chitin synthase inhibitor, was used at low concentrations (5–10 μM) for investigating the impact of chitin synthesis on mollusc larval shell formation in vivo. Being aware of the tremendous uncertainties that any in vivo approach bears, we observed dramatic structural alterations in mollusc larval shells by using polarized light video microscopy for the in vivo investigations and scanning electron microscopy imaging of extracted shell material prepared from NikkomycinZ treated Mytilus galloprovincialis larvae of various age. Provided that NikkomycinZ mainly affects chitin synthesis, our data obtained from different shell locations suggest that the mollusc chitin synthase fulfils an important enzymatic role in the coordinated formation of larval bivalve shells. It can be speculated that chitin synthesis bears the potential to contribute via signal transduction pathways to the implementation of hierarchical patterns into chitin mineral-composites such as prismatic, nacre, and crossed-lamellar shell types.
Mytilus galloprovincialis spawning and larvae culture
Larvae of Mytilus galloprovincialis were obtained after spontaneous spawning in the laboratory from February to April according to established protocols [61, 62]. Artificial seawater was prepared from Reef Crystals™ (Aquarium Systems, Sarrebourg, France) and kept at a density of 1.022 g/cm3 at 18°C. Adult animals of Mytilus galloprovincialis were obtained from Thaeron Fils (Riec, Belon, France). Animals were cleaned from algae, and thoroughly rinsed in tap water. Purified animals were put into a shallow tank containing 60 l of artificial seawater. After 2 h of recovery, 20 females were sorted out according to prespawning behavior and placed in a 40 l spawning tank. Animals were fed with 10 ml Aquatim™ phytoplankton (Kroon AQA® /Z+L, Langen, Germany), and spawning males were added for 1–2 min in order to induce the females to spawn. The release of eggs occurred spontaneously during the following hours (see Additional File 1). 5 spawning male animals (see Additional File 2) were added for 30 min to complete fertilization. The fertilized eggs were harvested by using a combination of nylon membranes with 20 μm and 40 μm mesh size (see Additional File 3). The success of fertilization and the removal of potential contaminants were checked microscopically. The fertilized eggs were placed in 10 l of artificial seawater with slight aeration (see Additional File 4). The day of fertilization was defined as day zero (0-day). The developing embryos were fed with 5 ml Aquatim™ phytoplankton the following day. Every second day, the water was exchanged completely in order to remove contaminants, non-developing embryos, and dead larvae by increasing the nylon mesh sizes in intervals of ~20 μm according to the average size of healthy individuals. Mollusc larvae were fed with 5 ml Aquatim™ phytoplankton in several portions per day. Larvae of various developmental stages were obtained. About 22 to 29 days after fertilization, larvae reached the pediveliger stage with subsequent metamorphosis into the adult.
Larvae culture in the presence of NikkomycinZ
Four series of experiments were performed, with each series covering selected developmental time frames of Mytilus galloprovincialis larvae. The time frames started 2, 5, 8, and 12 days after fertilization, and ended 5, 8, 12, and 15 days after fertilization, respectively. For each test culture, 100–200 Larvae were transferred from a 10l culture into 1 ml artificial seawater per well in a 12-well culture dish (Falcon, Heidelberg, Germany). NikkomycinZ (Sigma, Deisenhofen, Germany) was added in concentrations of 0 μM (control), 5 μM and 10 μM. Duplicate test cultures were performed. Every second day, the seawater was exchanged completely in order to remove contaminants and algal debris, and to avoid changes in salt concentration due to evaporation. NikkomycinZ was added freshly in the desired concentration for each test culture. Larvae were fed with 15 μl Aquatim® (Z+L/Kroonaqa, Langen, Germany) phytoplankton once a day. The larvae cultures were incubated at 18°C with a 12 h light cycle.
A series of 40 ml scale cultures of 2-day, 5-day, 8-day, and 12-day Mytilus galloprovincialis larvae in the presence of 0 μM (control), 5 μM and 10 μM NikkomycinZ was performed for preparative purposes. The cultures were incubated at 18°C with a 12 h light cycle and aerated by gently shaking. Each culture was fed daily with 500 μl Aquatim® phytoplankton. The seawater medium containing NikkomycinZ in the desired concentrations was exchanged every second day. Larvae were harvested after 3 days (earlier stages), and after about 10 days (later stages) of NikkomycinZ incubation on the 5th, 8th, 19th, and 22nd day after fertilization, respectively, and processed immediately or stored shock-frozen in liquid N2 at -70°C.
Evaluation of survival rate and phenotypes
Test cultures (1 ml scale) of Mytilus galloprovincialis larvae were frequently inspected using a stereomicroscope Wild M10 equipped with a Plan Apo 0,63× objective/10× zoom and a KL1500 electronic light source (Leica, Heerbrugg, Switzerland). The percentages of surviving and of abnormally developed larvae were estimated independently by up to three different persons with identical values within 10–20% deviation. The estimated deviation was thus defined as 20%.
Larval phenotypes were investigated directly in the 12-well culture dish using video microscopy. An Axiovert 35 microscope (Zeiss, Oberkochen, Germany) equipped with LD ACHROPLAN 32×/0.40 Ph2 and LD ACHROPLAN 20×/0.40 Korr Ph2 objectives was used. Images were recorded with a colour video camera TK-1070BE (JVC, Friedberg, Germany) in S-VHS quality (AG-6730 Time lapse recorder, Panasonic, Hamburg) and digitized (Scenalyzer live: Andreas Winter, Nassau, Bahamas; TMPGEnc 3.0 × Press: Pegasys, Tokyo, Japan; and Premiere 7.0: Adobe Systems, München, Germany).
Polarized Light Microscopy
Specimens were prepared on glass slides. Polarized light images were obtained by video microscopy as described in the previous section. The Axiovert 35 microscope (Zeiss, Oberkochen, Germany) was optionally equipped with crossed polarizers that were placed into the light path below and above the specimen. All images were taken by a constant light intensity of 11 arbitrary units.
Scanning Electron Microscopy
For structural investigations of larval shells by scanning electron microscopy, shock frozen samples were thawed at room temperature prior to use. Larvae were washed 2× in deionised water. Larval shells were purified from tissue in 1% – 2.5% sodium hypochlorite for 7 minutes and washed in deionised water several times. Larval shells were placed in a small drop of deionised water on top of an ethanol cleaned specimen holder (stainless steel) and dried in air and under vacuum. Specimens were sputter coated with gold 3× for 30 seconds at 1,0 kV, 10 mA, and 3·10-1bar using a Scancoat Six Sputter Coater (Boc Edwards, Kirchheim, Germany). A Leo 1530 FESEM scanning electron microscope (Zeiss, Oberkochen, Germany) was operated at 2.0 kV and 6 mm working distance. The Leo-32 V03.00h software and Adobe Photoshop (Adobe München, Germany) was used for image processing.
The authors thank Prof. Dr. M. Sumper for sustained scientific support. The authors acknowledge Prof. Dr. G. Haszprunar, and Dr. B. Ruthensteiner, Zoologische Staatssammlung, Munich, for their expert advice on mollusc larvae cultures. The authors thank Dr. S. Kaufmann for critically revising the manuscript. This work was supported by institutional funds from the University of Regensburg.
- Simkiss K, Wilbur KM: Biomineralization, Cell Biology and Mineral Deposition. New York , Academic Press; 1989.Google Scholar
- Lowenstam HA, Weiner S: On Biomineralization. New York , Oxford Univ. Press; 1989.Google Scholar
- Addadi L, Joester D, Nudelman F, Weiner S: Mollusk shell formation: A source of new concepts for understanding biomineralization processes. Chem Eur J 2006, 12(4):980–987. 10.1002/chem.200500980View ArticlePubMedGoogle Scholar
- Jeuniaux C: Chitine et chitinolyse. Paris , Masson; 1963.Google Scholar
- Peters W: Occurence of chitin in mollusca. Comp Biochem Physiol B 1972, 41: 541–550. 10.1016/0305-0491(72)90117-4Google Scholar
- Goffinet G, Jeuniaux C: Distribution et importance quantitative de la chitine dans les coquilles de mollusques. Cahiers Biol Mar 1979, 20: 341–349.Google Scholar
- Weiss IM, Schonitzer V: The distribution of chitin in larval shells of the bivalve mollusk Mytilus galloprovincialis. J Struct Biol 2006, 153(3):264–277. 10.1016/j.jsb.2005.11.006View ArticlePubMedGoogle Scholar
- Weiner S, Traub W: X-ray diffraction study of the insoluble organic matrix of mollusk shells. FEBS letters 1980, 111(2):311–316. 10.1016/0014-5793(80)80817-9View ArticleGoogle Scholar
- Weiner S, Talmon Y, Traub W: Electron diffraction of mollusk shell organic matrices and their relationship to the mineral phase. Int J Biol Macromol 1983, 5: 325–328. 10.1016/0141-8130(83)90055-7View ArticleGoogle Scholar
- Levi-Kalisman Y, Falini G, Addadi L, Weiner S: Structure of the nacreous organic matrix of a bivalve mollusk shell examined in the hydrated state using Cryo-TEM. J Struct Biol 2001, 135: 8–17. 10.1006/jsbi.2001.4372View ArticlePubMedGoogle Scholar
- Falini G, Albeck S, Weiner S, Addadi L: Control of aragonite or calcite polymorphism by mollusk shell macromolecules. Science 1996, 271: 67–69. 10.1126/science.271.5245.67View ArticleGoogle Scholar
- Beniash E, Aizenberg J, Addadi L, Weiner S: Amorphous calcium carbonate transforms into calcite during sea urchin larval spicule growth. Proc R Soc Lond B 1997, 264: 461–465. 10.1098/rspb.1997.0066View ArticleGoogle Scholar
- Politi Y, Arad T, Klein E, Weiner S, Addadi L: Sea urchin spine calcite forms via a transient amorphous calcium carbonate phase. Science 2004, 306(5699):1161–1164. 10.1126/science.1102289View ArticlePubMedGoogle Scholar
- Weiss IM, Tuross N, Addadi L, Weiner S: Mollusc larval shell formation: amorphous calcium carbonate is a precursor phase for aragonite. J Exp Zool 2002, 293(5):478–491. 10.1002/jez.90004View ArticlePubMedGoogle Scholar
- Bøggild OB: The shell structure of the mollusks. K Dan Vidensk Selsk Skr Naturvidensk Math Afd 1930, 9: 233–326.Google Scholar
- Nudelman F, Chen HH, Goldberg HA, Weiner S, Addadi L: Spiers Memorial Lecture. Lessons from biomineralization: comparing the growth strategies of mollusc shell prismatic and nacreous layers in Atrina rigida. Faraday Discuss 2007, 136: 9–25. 10.1039/b704418fView ArticlePubMedGoogle Scholar
- Glaser L, Brown DH: The synthesis of chitin in cell-free extracts of Neurospora crassa. J Biol Chem 1957, 228(2):729–742.PubMedGoogle Scholar
- Weiss IM, Schonitzer V, Eichner N, Sumper M: The chitin synthase involved in marine bivalve mollusk shell formation contains a myosin domain. FEBS Lett 2006, 580(7):1846–1852. 10.1016/j.febslet.2006.02.044View ArticlePubMedGoogle Scholar
- Cabib E: Chitin synthetase system from yeast. Methods Enzymol 1972, 28: 572–580.View ArticleGoogle Scholar
- Cabib E: The septation apparatus, a chitin-requiring machine in budding yeast. Arch Biochem Biophys 2004, 426(2):201–207. 10.1016/j.abb.2004.02.030View ArticlePubMedGoogle Scholar
- Ruiz-Herrera J, Martinez-Espinoza AD: Chitin biosynthesis and structural organization in vivo. In Chitin and Chitinases. Edited by: Jolles P, Muzzarelli RAA. Basel/Switzerland , Birkhaeuser Verlag; 1999:39–53.View ArticleGoogle Scholar
- Ruiz-Herrera JM, Gonzalez-Prieto J, Ruiz-Medrano R: Evolution and phylogenetic relationships of chitin synthases from yeasts and fungi. FEMS Yeast Research 2002, 1(4):247–256. 10.1111/j.1567-1364.2002.tb00042.xView ArticlePubMedGoogle Scholar
- Ruiz-Herrera J: Fungal Cell Wall: structure, synthesis, and assembly. New York , CRC; 1992.Google Scholar
- Ruiz-Herrera J, Bartnicki-Garcia S: Synthesis of cell wall microfibrils in vitro by a "soluble" chitin synthetase from Mucor rouxii. Science 1974, 186(4161):357–359. 10.1126/science.186.4161.357View ArticlePubMedGoogle Scholar
- Bartnicki-Garcia S: Chitosomes: past, present and future. FEMS Yeast Res 2006, 6(7):957–965. 10.1111/j.1567-1364.2006.00158.xView ArticlePubMedGoogle Scholar
- Tellam RL, Vuocolo T, Johnson SE, Jarmey J, Pearson RD: Insect chitin synthase cDNA sequence, gene organization and expression. Eur J Biochem 2000, 267(19):6025–6043. 10.1046/j.1432-1327.2000.01679.xView ArticlePubMedGoogle Scholar
- Merzendorfer H: Insect chitin synthases: a review. J Comp Physiol B 2006, 176: 1–15. 10.1007/s00360-005-0005-3View ArticlePubMedGoogle Scholar
- Moussian B, Schwarz H, Bartoszewski S, Nüsslein-Volhard C: Involvement of chitin in exoskeleton morphogenesis in Drosophila melanogaster. J Morphology 2005, 264(1):117–130. 10.1002/jmor.10324View ArticleGoogle Scholar
- Arakane Y, Hogenkamp DG, Zhu YC, Kramer KJ, Specht CA, Beeman RW, Kanost MR, Muthukrishnan S: Characterization of two chitin synthase genes of the red flour beetle, Tribolium castaneum, and alternate exon usage in one of the genes during development. Insect Biochem Mol Biol 2004, 34(3):291–304. 10.1016/j.ibmb.2003.11.004View ArticlePubMedGoogle Scholar
- Zhang Y, Foster JM, Nelson LS, Ma D, Carlow CK: The chitin synthase genes chs-1 and chs-2 are essential for C. elegans development and responsible for chitin deposition in the eggshell and pharynx, respectively. Dev Biol 2005, 285(2):330–339. 10.1016/j.ydbio.2005.06.037View ArticlePubMedGoogle Scholar
- Hector RF: Compounds active against cell walls of medically important fungi. Clin Microbiol Rev 1993, 6(1):1–21.PubMed CentralPubMedGoogle Scholar
- Cohen E: Chitin synthesis and inhibition: a revisit. Pest Manag Sci 2001, 57: 946–950. 10.1002/ps.363View ArticlePubMedGoogle Scholar
- Ruiz-Herrera J, San-Blas G: Chitin synthesis as target for antifungal drugs. Curr Drug Targets Infect Disord 2003, 3(1):77–91. 10.2174/1568005033342064View ArticlePubMedGoogle Scholar
- Cabib E: Differential inhibition of chitin synthetases 1 and 2 from Saccharomyces cerevisiae by polyoxin D and nikkomycins. Antimicrob Agents Chemother 1991, 35(1):170–173.PubMed CentralView ArticlePubMedGoogle Scholar
- Cohen E, Casida JE: Inhibition of Tribolium gut chitin synthetase. Pesticide Biochemistry and Physiology 1980, 13: 129–136. 10.1016/0048-3575(80)90064-4View ArticleGoogle Scholar
- Suzuki S, Isono K, Nagatsu J, Mizutani T, Kawashima Y, Mizuno T: A New Antibiotic, Polyoxin A. J Antibiot (Tokyo) 1965, 18: 131.Google Scholar
- Dähn U, Hagenmaier H, Höhn H, König WA, Wolf G, Zähner H: Metabolic products of microorganisms 154. Nikkomycin, a new inhibitor of fungal chitin synthesis. Arch Microbiol 1976, 107: 143–160. 10.1007/BF00446834View ArticlePubMedGoogle Scholar
- Yadan JC, Gonneau M, Sarthou P, Le Goffic F: Sensitivity to nikkomycin Z in Candida albicans: role of peptide permeases. J Bacteriol 1984, 160(3):884–888.PubMed CentralPubMedGoogle Scholar
- McCarthy PJ, Troke PF, Gull K: Mechanism of action of nikkomycin and the peptide transport system of Candida albicans. J Gen Microbiol 1985, 131(4):775–780.PubMedGoogle Scholar
- Decker H, Zahner H, Heitsch H, Konig WA, Fiedler HP: Structure-activity relationships of the nikkomycins. J Gen Microbiol 1991, 137(8):1805–1813.View ArticlePubMedGoogle Scholar
- Kniprath E: Ontogeny of the Molluscan Shell Field: a Review. Zoologica Scripta 1981, 10(1):61–79. 10.1111/j.1463-6409.1981.tb00485.xView ArticleGoogle Scholar
- Kniprath E: Larval development of the shell and the shell gland in Mytilus (Bivalvia). Development Genes and Evolution 1980, 188(3):201–204.Google Scholar
- Medaković D, Hrs-Brenko M, Popovié S, Gržeta B: X-ray diffraction study of the first larval shell of Ostrea edulis. Marine Biology 1989, 101(2):205–209. 10.1007/BF00391459View ArticleGoogle Scholar
- Medaković D: The calcification processes of larval, juvenile and adult shells of oysters (Ostrea edulis, Linnaeus) and mussels (Mytilus galloprovincialis, Lamarck). In Ruder Bosković Institute. Volume Ph.D.. Zagreb, Croatia , University of Zagreb; 1995.Google Scholar
- Wilt F: Development of calcareous skeletal elements in invertebrates. Differentiation 2003, 71(4–5):237–250. 10.1046/j.1432-0436.2003.7104501.xView ArticlePubMedGoogle Scholar
- Carriker MR, Palmer RE: Ultrastructural morphogenesis of prodissoconch and early dissoconch valves of the oyster Crassostrea virginica. Proc Natl Shellfish Assoc 1979, 69: 103–128.Google Scholar
- Waller TR: Functional morphology and development of veliger larvae of the European oyster, Ostrea edulis Linne. Smithsonian Contrib Zool 1981, 328: 1–70.View ArticleGoogle Scholar
- Rees CB: The interpretation and classification of lamellibranch larvae. Hull Bull Mar Ecol 1950, 3: 73–104.Google Scholar
- Redfearn P, Chanley P, Chanley M: Larval shell development of four species of New Zealand mussels: (Bivalvia, Mytilacea). New Zealand Journal of Marine & Freshwater Research 1986, 20(2):157–172.View ArticleGoogle Scholar
- Lowenstam HA: Minerals formed by organisms. Science 1981, 211: 1126–1131. 10.1126/science.7008198View ArticlePubMedGoogle Scholar
- Schmidt WJ: Die Bausteine des Tierkörpers in Polarisiertem Lichte. Bonn , Cohen; 1924.Google Scholar
- Coutinho PM, Deleury E, Davies GJ, Henrissat B: An Evolving Hierarchical Family Classification for Glycosyltransferases. J Mol Biol 2003, 328(2):307–317. 10.1016/S0022-2836(03)00307-3View ArticlePubMedGoogle Scholar
- Tokumura T, Horie T: Kinetics of nikkomycin Z degradation in aqueous solution and in plasma. Biol Pharm Bull 1997, 20(5):577–580.View ArticlePubMedGoogle Scholar
- Ellgaard L, Helenius A: Quality control in the endoplasmic reticulum. Nat Rev Mol Cell Biol 2003, 4(3):181–191. 10.1038/nrm1052View ArticlePubMedGoogle Scholar
- Williams DB: Beyond lectins: the calnexin/calreticulin chaperone system of the endoplasmic reticulum. J Cell Sci 2006, 119(Pt 4):615–623. 10.1242/jcs.02856View ArticlePubMedGoogle Scholar
- Lehle L, Tanner W: Protein glycosylation in yeast. New Comprehensive Biochemistry. In Glycoproteins. Volume 29a. Edited by: Montreuil J, Vliegenthart JFG, Schachter H. Amsterdam , Elsevier; 1995:475–509.View ArticleGoogle Scholar
- Weiss IM, Marin F: The Role of Enzymes in Biomineralization Processes. In Metal Ions in Life Sciences: Biomineralization: From Nature to Application. Volume 4. Edited by: Sigel A, Sigel H, Sigel RKO. West Sussex, UK , John Wiley & Sons; 2008:71–126.Google Scholar
- Schlüter U: Ultrastructural evidence for inhibition of chitin synthesis by nikkomycin. Development Genes and Evolution 1982, 191(3):205–207.Google Scholar
- Holst H, Malissiovas D, Schmutterer H: Wirkungen eines chitinsynthesehemmenden Antibiotikums auf Spinnmilben und Insekten. Mitt Dtsch Ges Allg Angew Ent 1978, 1: 143–146.Google Scholar
- Gotliv BA, Addadi L, Weiner S: Mollusk shell acidic proteins: In search of individual functions. ChemBioChem 2003, 4(6):522–529. 10.1002/cbic.200200548View ArticlePubMedGoogle Scholar
- Hoff FH, Snell TW: Plankton Culture Manual. 5th edition. Edited by: Neslen J. Dade City, FL, U.S.A. , Florida Aqua Farms, Inc.; 1987:162.Google Scholar
- Satuito CG, Natoyama K, Yamazaki M, Fusetani N: Larval development of the mussel Mytilus edulis galloprovincialis cultured under laboratory conditions. Fisheries Science 1994, 60(1):65–68.Google Scholar