- Email: [email protected]
Occurrence and diversity of lipids in modern coral skeletons
Zoology 113 (2010) 250–257
Contents lists available at ScienceDirect
Zoology journal homepage: www.elsevier.de/zool
Occurrence and diversity of lipids in modern coral skeletons Bastien Farre, Jean-Pierre Cuif, Yannicke Dauphin ∗ UMR IDES 8148, bat. 504, Université Paris XI, rue G. Clemenceau, F-91405 Orsay, France
a r t i c l e
i n f o
Article history: Received 9 June 2009 Received in revised form 18 November 2009 Accepted 19 November 2009 Keywords: Biomineralisation Intraskeletal lipids Coral microstructure
a b s t r a c t Coral skeletons are composite acellular structures, in which organic macromolecules are intimately associated with mineral phases. Previous studies focussed on proteins and sugars of the soluble organic matrices extracted from the skeletons. Here we report the occurrence of diverse lipids which were extracted from the aragonitic skeletons of seven modern coral species. Using thin layer chromatography, we show that these lipids differ in quantity and composition between the species. Higher proportions of sterols and sterol esters in skeleton extracts as compared to a much higher abundance of waxes and triglycerides in previously studied extracts from scleractinian soft tissues suggest a specific, although not yet determined, role in biomineralisation. The occurrence of intraskeletal lipids along with other organic components should also be taken into account when using coral skeletons as bone allografts, as well as in fossilisation processes. © 2010 Elsevier GmbH. All rights reserved.
1. Introduction Corals, the main reef builders in the tropical seas, are well known for the diversity of the calcareous skeletons built by their anatomically simple polyps. Although simple, these polyps have undergone a remarkable evolution regarding the colonial mode of life, leading to numerous variations in sizes and spatial relationships between the polyps. The skeletons are always produced by the calicoblastic epithelium (=basal part of the ectoderm) of the polyps and in close contact with it (Bourne, 1887, 1899; Clode and Marshall, 2002; Tambutté et al., 2007a). The organisation of the animals is reflected by the organisation of the calcareous skeletons, not only from a morphological point of view but also at the smaller scale of the spatial arrangement, structure and composition of the skeleton building units, i.e. the mineral fibres (Bourne, 1899). Coral skeletons are built from calcium carbonate belonging to the aragonite variety. Due to their long life span (specifically of the large colonial build-ups that can grow for centuries) and fixed positions, corals are largely used as biological archives for palaeoclimate reconstruction. Since this group is also remarkable for the intensity of the “vital effect” (Urey et al., 1951), i.e. the property of each species to record environmental variations with a specific correlation between isotopic fractionation and temperature changes, these biological archives have also been used empirically through the complex and very uncertain procedure of “calibrating” the coral species. A major improvement would result from an understanding
∗ Corresponding author. Tel.: +33 1 69 15 61 17; fax: +33 1 69 15 61 21. E-mail address: [email protected] (Y. Dauphin). 0944-2006/$ – see front matter © 2010 Elsevier GmbH. All rights reserved. doi:10.1016/j.zool.2009.11.004
of the factors influencing the crystallisation process, and notably the role of the organic matrix in which crystallisation occurs. For several decades it has been recognised that glycoprotein compounds are always associated with coral skeletons (Young, 1971; Mitterer, 1978). But only in recent times precise information has been collected about the biochemical control of crystallisation. Far from being crystals of pure aragonite, as commonly assumed (Bryan and Hill, 1941; Veis, 2005), species-specific associations of macromolecules are detectable within the crystal-like fibres (Mitterer, 1978; Cuif and Gautret, 1995; Cuif et al., 1999; Dauphin, 2001; Dauphin and Cuif, 1997; Puverel et al., 2005; Tambutté et al., 2007b). Atomic force microscopy has shown that the building units of the fibres are small round-shaped corpuscles irregularly coated with organic components (Cuif and Dauphin, 2005a,b). Information about the crystallisation process has been obtained by transmission electron microscopy (imaging and diffraction) (Johnston, 1977, 1979; Clode and Marshall, 2002). It has been shown that crystallisation occurs as the final step of mineralisation in the growth layers, the growth units of the fibres (Baronnet et al., 2008). These results emphasise the need for a better understanding of the complex organic matrices that form the hydrogel in which crystallisation occurs. Besides proteins and glucids, lipids are one of the major category of organic components. Lipids are structural components of cell membranes and important signalling molecules. They play a role in energy storage and in the synthesis of materials “through their ability to self-assemble, compartmentalise and template” (Collier and Messersmith, 2001). Two broad classes have been defined for chromatography purposes: simple lipids and complex lipids. Glycerophospholipids (also called phospholipids) are the primary constituents of biomembranes. Although (phospho)lipids are said
B. Farre et al. / Zoology 113 (2010) 250–257
251
Fig. 1. Morphological variety of coral skeletons. (A) Branching colonial Millepora (“fire coral”), with a porous skeleton and small corallites (inserts); Caribbean archipelago, Guadalupe Island. (B) Massive skeleton of Porites, with small adjacent corallites (insert); New Caledonia. (C) Symphyllia, medium-sized corallites with meandroid arrangement; New Caledonia. (D) Pocillopora, with small corallites (insert); Polynesia, Moorea Island. (E) Fungia, one of the biggest solitary corals; New Caledonia. (F) Solenosmilia, with distinct corallites (insert); Porcupine plateau, North-East Atlantic. (G) Caryophyllia, a solitary deep-sea coral; Biscay Gulf, Meriadzec plateau, West Atlantic.
to be involved in biomineralisation processes, their role in extracellular calcareous biominerals is not yet known. Despite some pioneer work, lipids in coral skeletons have so far not been actually investigated. Silliman (1846) described “waxlike” materials extracted from skeletons. Lipids were described in some modern species by Lester and Bergmann (1941), Young et al. (1971), and Meyers et al. (1978). However, while some analyses were done on decalcified skeletons, in other studies lipids were extracted using acetone or ether, therefore comparisons are difficult. Thus, although a number of analyses have shown the presence of lipids in the soft tissues (Harland et al., 1993; Ward, 1995), data on coral skeletons remain scarce. This study, based on several species of corals selected for their microstructural diversity and difference in mode of life, aims at providing a first set of data concerning the presence and diversity of skeletal lipids.
the Gulf of Biscaye, Meriadzec plateau (Fig. 1G) are representatives of the non-symbiotic metabolism.
2. Materials and methods
2.3.1. Scanning electron microscopy (SEM) Several fractured and polished sections were cut perpendicular and parallel to the surface of the septa and walls. Polished sections and some fractures were etched with various acids and enzymes to reveal microstructural features. Detailed procedures of sample preparation are given in the legend of Fig. 7. SEM observations were conducted using a Philips XL30 (Philips, Amsterdam, The Netherlands) at UMR IDES, Université Paris XI.
2.1. Materials Calcareous skeletons are built by several high level taxa of modern and fossil Cnidaria. Among them, we have studied aragonite skeletons, selected for the disparities of their skeleton shapes, phylogeny, metabolism and lifestyle, but all living in a modern marine environment. The Hydrozoa are represented by Millepora alicornis from Guadalupe (Fig. 1A), the skeletons of which are important contributors to tropical reefs. The other six species belong to the Scleractinia. They comprise three symbiotic colonial species: Porites lutea from New Caledonia (Fig. 1B), one of the most used skeletons in palaeoclimate reconstructions; Symphyllia, from New Caledonia, a colonial type with a restricted number of medium-sized polyps (Fig. 1C); and Pocillopora from Moorea island (Polynesia) with small polyps (Fig. 1D). The solitary Fungia repanda from New Caledonia is also a symbiotic coral (Fig. 1E). Two deep-sea corals, the colonial Solenosmilia variabilis from the Porcupine plateau, north-east Atlantic ocean (Fig. 1F) and the solitary Caryophyllia ambrosia from
2.2. Standards Commercial analytic grade standards were used for all experiments. A phospholipid mixture was used for polar lipids: l-␣-lysophosphatidylcholine, l-␣-phosphatidylcholine and l-␣phosphatidylinositol ammonium salt from soybean and l-␣phosphatidylethanolamine from Escherichia coli. Cholesterol was used as standard for sterols, triolein for triglycerides, oleic acid for free fatty acids, stearyl oleate for waxes and cholesteryl oleate for sterol esters. 2.3. Methods
2.3.2. Atomic force microscopy (AFM) AFM observations were obtained with a Dimension 3100 Nanoscope III (Veeco Instruments Inc., Plainview, NY, USA) housed at UMR IDES, Université Paris XI. Samples were imaged at room temperature and in air using tapping mode. Phase images were generated as a consequence of variations in material properties such as viscoelasticity, composition, adhesion, friction, etc. AFM has several advantages over SEM, e.g., samples do not require coatings. However, disadvantages are the small image size and a reduced depth of field in the order of micrometers. Thus, observation of fractured surfaces is difficult, and most samples have to be polished
252
B. Farre et al. / Zoology 113 (2010) 250–257
Fig. 2. FTIR spectra of standards: (A) cholesterol (lipid); (B) chitin; (C) bovine serum albumin (protein).
and cleaned. Depending upon the structures, different preparative processes were used (see details in the legend of Fig. 7). 2.3.3. Fourier transform infrared spectrometry (FTIR) For all species, FTIR spectra were first recorded on undecalcified powder, then on the extracted insoluble organic matrices of selected species, depending on the TLC results. All spectra were recorded at 4 cm−1 resolution with 64 scans with a strong Norton-Beer apodization on a Perkin-Elmer Model 1600 FTIR spectrometer (Perkin-Elmer Inc., Waltham, MA, USA) in the range from 4000 to 450 cm−1 . The spectrometer was equipped with a diffuse reflectance accessory, which permits diffuse reflectance infrared Fourier transform (DRIFT) measurements with high sensitivity for powders. Raw diffuse reflectance spectra appear different from their transmission equivalents, so a conversion was applied to compensate for these differences. All spectra were corrected using the Kubelka–Munk function. The system was
Fig. 3. FTIR spectra of non-decalcified coral skeletons.
B. Farre et al. / Zoology 113 (2010) 250–257
253
purged and permanently maintained under nitrogen to reduce atmospheric CO2 and H2 O absorption. A background spectrum was measured for pure KBr. Sample spectra were automatically ratioed against background to minimise CO2 and H2 O bands. To assess reproducibility, several spectra from the same sample were obtained and the results gave correlation coefficients higher than 95%. As insoluble organic matrices resulting from a decalcification process, two symbiotic samples (Fungia as a solitary genus and Porites as a colonial genus) and a non-symbiotic solitary genus (Caryophyllia) were used.
2.3.5. Extraction and purification of the insoluble organic matrix Powdered samples were immersed in 5 ml of Milli-Q water, then decalcified by progressive addition of 50% acetic acid so that the pH (automatically controlled with a titrimeter) was above 4. The entire extract was directly centrifuged at 21,000 × g for 15 min, which separated the supernatant (soluble) and precipitated (insoluble) fractions. The insoluble fraction was desalted by repeated centrifugations in Milli-Q water and lyophilised.
2.3.4. Removal of the polyps To remove tissues, polyps were decayed by a 3 h immersion in distilled water to destroy the cells. Removal of the remaining living tissues was done by using a water-jet, followed by immersion in 3% NaClO for 1 h to remove organic contaminants, sonicated for 5 min.
2.3.6. Lipid extractions After removal of the polyps and decontamination, the studied material was ground to a powder in an electric mortar. 9.1 g of each sample were soaked in warm chloroform/methanol (1:1, v/v) for three days under mild and constant stirring and sonicated every
Then the samples were rinsed with Milli-Q water and air-dried, then ground to powder.
Fig. 4. TLC lanes of lipid standards.
Fig. 5. TLC lanes of lipids extracted from the aragonitic coral skeletons, and a mixture of standards.
254
B. Farre et al. / Zoology 113 (2010) 250–257
day. The mixture was then centrifuged at 19,500 × g for 5 min to separate the powder and insoluble parts from the solvent-soluble part. The organic solvent-soluble part was finally concentrated by evaporation. 2.3.7. Thin layer chromatography (TLC) 100 l of the concentrated extracts were applied to 10 cm × 10 cm high performance thin layer chromatography (HPTLC) plates (nano-silica gel plates; Macherey-Nagel GmbH & Co. KG, Düren, Germany). A four-stage development process was used. TLC plates were first developed to their full height with pure n-hexane, then again to full height with pure benzene, and finally twice to half height with hexane: ether: acetic acid (70:30:1, by vol.). After drying, the plates were immersed in a mixture of phosphoric acid, 33% acetic acid, sulfuric acid and copper sulfate (5:5:0.5:90, by vol.) for 40 s, and heated at 110 ◦ C for 15 min. The positions were compared with individual standards (Fig. 2) and 10 l of a mixture of the commercial standards. 3. Results
Fig. 5 shows the diversity in quantity and composition of the lipids extracted from the coral skeletons. The samples of Fungia and Solenosmilia show numerous well-stained bands, whereas Pocillopora and Caryophyllia show only faint or no bands. The most common lipids are the strongly polar lipids (phospholipids), which represent a very heterogeneous group of various functions in the organism. They are found in every sample, although in some species in very low amounts (Pocillopora and Caryophyllia; Fig. 5). Sterols are also well represented, with a major (sometimes unique) spot around the cholesterol position for every specimen except Caryophyllia, which displays basically no lipids but the polar ones. Waxes (such as stearyl oleate) and steryl esters (cholesteryl oleate) are also present in every sample, but in very small amounts. They are rare even in the high-lipid samples (Fungia and Solenosmilia), nevertheless present. Free fatty acids (such as oleic acid) are only represented in the most lipid-rich samples, Solenosmilia and Fungia, although some traces may exist in less lipidic samples, but in too low amounts to be noticed. Triglycerides (triolein standard), mostly known as storage lipids, are only present in the two samples with the most lipids, Solenosmilia and Fungia.
3.1. Bulk composition: FTIR data Standards (Fig. 2): cholesterol, chitin and BSA show strong amide A bands. Amide I and amide II bands are strong in BSA and chitin spectra, whereas only a weak band at 1671 cm−1 is visible in cholesterol. Bands between 800 and 1100 cm−1 are strong in cholesterol (Fig. 2A), weak or absent in BSA spectrum. On the opposite, bands due to C–H stretching vibrations (2800–3000 cm−1 ) are stronger than those of amide A, amide I and amide II in cholesterol. Similar bands are present in polysaccharides (Fig. 2B) and proteins (Fig. 2C), but they are weaker than those of amide bands. They are usually said to be characteristic of lipids. All spectra of undecalcified samples show the organo-mineral composition of the coral skeletons, with bands (1 to 4) characteristic of aragonite (Fig. 3). The first part of the 2 doublet (858 cm−1 ) shows that the Sr content of aragonite is high (>7000 ppm). Bands are also present between 4000 and 1500 cm−1 , indicative of the presence of organic components. Amide A, amide I and amide II bands may be assigned to proteins, sugars or lipids. Wave numbers of amide A bands in Scleractinia are higher than 3350 cm−1 , whereas in the Hydrozoa sample (Millepora), this band is lower than 3300 cm−1 . Amide A intensities are variable: high in Millepora and Porites, low in other specimens. Weak bands in the 2800–3000 cm−1 region, usually assigned to lipids, but also present in chitin, are visible as shoulders in all the samples. However, these bands are identified as bands by the software in Fungia (2 bands), Solenosmilia and Caryophyllia (1 band) for Scleractinia, and Millepora (1 band). 3.2. Lipid components Standards were chromatographed separately (Fig. 4). Some resulted in a discrete band (triolein), others in large spots (stearyl oleate, cholesterol). Depending on the used quantity, the oleic acid band is large (Fig. 4), or small (Fig. 5). Standards were also mixed to estimate the ability of the used technique to differentiate a complex mixture (Fig. 5). Because of the overlap between the large band of oleic acid and that of cholesterol when large amounts were used, only a reduced quantity of oleic acid was used in the mixed standard. Cholesteryl oleate and stearyl oleate also show some overlap (Fig. 5). Despite these overlaps, discrete bands are obtained when using low amounts of individual products. Since the same amount of skeleton powder was used for each sample, the relative density of the spots is related to the relative proportion of the lipids.
Fig. 6. FTIR spectra of the insoluble organic matrices of three decalcified coral skeletons.
B. Farre et al. / Zoology 113 (2010) 250–257
The lipid content varies a lot from one sample to another, but some of the major components can be compared. It seems that the depth or life style (i.e. solitary or colonial) have no influence on the lipid yield of the coral. The two corals with the highest lipid content are also the most different: the single, symbiotic, shallow-water Fungia and the colonial, non-symbiotic, deep-sea Solenosmilia. Some lipids are, however, present in all the samples. These lipids are mainly polar lipids and sterol esters. A very polar compound also seems recurrent, although it is not present in Caryophyllia. Finally, despite the similarity of Solenosmilia and Fungia, the two corals display a lipid composition pattern very different from one another. The most relevant differences show up in the polar lipid
255
zone between cholesterol and non-migrating lipids, where they are both in high quantity and displaying a wide diversity of composition. From TLC analyses, Fungia is rich in lipid contents, whereas Caryophyllia is poor; Porites is in an intermediate position (Fig. 5). FTIR spectra of the insoluble matrices obtained from decalcification show strong differences in lipid contents (Fig. 6), despite the different extraction procedures. The intensity of lipid bands (between 2950 and 2850 cm−1 ) decreases from Fungia (Fig. 6A) to Caryophyllia (Fig. 6C), with Porites in the middle position (Fig. 6B). The decalcification process is easy and fast, while the lipid extraction is time consuming and complex. The above results suggest that FTIR spectra on decalcified insoluble organic matri-
Fig. 7. Diversity and similarities in the biocrystallisation of coral skeletons. SEM and AFM phase images. (A/B) Millepora. (A) Section showing the different sensitivity to etching of the centre (dark grey) and lateral skeleton areas with a layered organisation. Polished and etched with formic acid (1%, aqu.) for 35 s. (B) Nanoparticles in the fibres, polished and etched surface, formic acid (1%, aqu.) for 10 s, 20 ◦ C. (C/D) Porites. (C) Polished and etched sections showing the layered growth mode of the skeletal tissue. (D) Aligned nanoparticles in the fibres, polished and etched surface, formic acid (10%) for 7 s. (E) Pocillopora. Regular fibrous fan systems diverging from the centres of calcification (arrows). Differences in the distribution of etching-sensible zones result in the clearly distinct aspect of the fibrous bundles compared to the fibrous tissues of Porites (see panel C). Polished and etched with alcalase for 3 h at 38 ◦ C. (F/G) Fungia. (F) Growth edge of a septum in a Fungia skeleton. Etching of a section perpendicular to the growth direction reveals that formation of a septum can be a polyphased process. Fibres radiate from centres of calcification (e.g. 1, arrows) first creating cylindrical structures (2) which are further covered by layered tissue (3). Polished and etched with a mixture of alcalase (2/3) and NaOH 1 M (1/3) for 4 h at 30 ◦ C, then rinsed with water. (G) Section presented produces a rather homogenous fibrous tissue. Polished and etched with formic acid (1%, aqu.) for 45 s. (H/I) Caryophyllia. (H) Layering of the fibrous tissue of the Caryophyllia septum shows that growth of the fibre is a step-wise process, symmetrical on both sides of the dotted line built by the centres of calcification. (I) Composite nanoparticles in the fibres, polished and etched surface, Milli-Q water for 4 h, 20 ◦ C.
256
B. Farre et al. / Zoology 113 (2010) 250–257
ces may be used to estimate the presence and quantity of lipids in biominerals.
Although the bulk position of the different lipid compounds allows identification of their classes, their exact position may shift from the position of the standard used, due to differences of polarity: the analysed compounds are different from the standard, although they are of the same class. The most important shift is between the cholesterol position and the other mollusk sterols. It displays a long series of compounds, most of them more polar than cholesterol. Triglycerides are only present in noticeable amounts in two samples (Fungia and Solenosmilia), but neither of the samples displays the exact position of the triolein. Solenosmilia has two triglyceride bands, one more polar than that of triolein, and another less polar. The triglycerides of Fungia are a bit less polar than the used standard (triolein). The free fatty acids are, in most of the samples, quite similar to oleic acid, although a second, more polar stain, appears on the Fungia sample.
2005a,b). When submitted to identical etching conditions, the studied species reveal the diversity of the microstructural (Fig. 7A, C, E–H) and nanostructural patterns (Fig. 7B, D, I). This diversity in the sensitivity to etching conditions can be reasonably correlated to the diversity of the organic components, including the lipid diversity. The reported results concerning lipid components show that, in addition to the molecular diversity of proteins and polysaccharides, lipids also contribute to the complexity of the organic blends involved in the crystallisation mechanism producing coral skeletons. Lipid diversity may also contribute to explaining a series of observations not compatible with the postulated uniformity of coral skeletons. For instance, coral skeletons are a good material for allografts of bone, widely used in bone repair surgery (Guillemin et al., 1987, 1995). However, the guiding function of the coral graft in the bone reconstruction process appears to be dependent on the species used for grafting. Thus, with respect to both fine structure and biochemical composition, coral skeletons appear to be produced through a crystallisation process quite similar to the mechanism responsible for shell formation in molluscs, in contrast to what is commonly published.
4.2. Comparison of lipids in the soft and skeletal parts of corals
5. Conclusion
Most of the data concerning the lipids in the coral skeletons are dealing with the soft tissues. Yamashiro et al. (1999), for example, studied the lipid content of the soft parts of 15 Cnidaria, among them eleven symbiotic zooxanthellate corals. Although the analytical methods are not the same, the chromatograms show that the lipid contents of the soft and skeletal parts of a given coral differ. The most important differences show up in the distribution of waxes and sterol esters: the soft parts generally contain much more waxes than sterol esters, whereas the skeletons show almost no waxes, but a much higher yield of sterol esters. Even more significant is the comparison of the lipid pattern of Porites lutea, which has been analysed both in the soft parts (Yamashiro et al., 1999) and the skeleton (this study). The relative abundance of the different lipids strongly differs from one tissue to another: the dominant lipids in soft tissues are waxes and triglycerides, while in the skeleton, the major lipids are polar lipids and sterols, along with sterol esters. Fatty acids and sterols are almost equivalent. Other lipid classes are almost not represented. This suggests that the lipids found in the hard parts of the corals are not just trapped parts of the living organism. The relative increase of sterol and sterol esters suggests that the lipids in the skeleton may be traces of remnant hormonal signals. However, more research, and specifically analytical chemistry, will be necessary to precisely identify the compounds and their specific roles. It is now demonstrated that the lipid content varies from one species to another, but also according to environmental conditions. Other parameters have also been identified as factors of variation of the lipid content: health or diet mode (Koop et al., 2001; Yamashiro et al., 2001).
Although most biochemical analyses of biocarbonates in corals focussed on either proteins or sugars extracted from the soluble organic matrices (Falini et al., 1996; Dauphin and Cuif, 1997; Cuif et al., 1999, 2003; Gotliv et al., 2003; Dauphin et al., 2008), it appears that lipids are also present in sufficient amounts to be analysed in the cnidarian hard parts, as well as in mollusc shells. The combined analysis with TLC/FTIR highlighted the presence of cholesterol-based lipids, mostly sterol compounds, but also sterol esters. These compounds have to be further examined to be accurately identified, both structurally and in their usual role in the living organisms. However, the high proportion of sterols and sterol esters suggests that several steroids may be involved in the biomineralisation process. Some lipids, mainly polar lipids and sterol esters, are present in all the studied coral skeleton samples. Those lipids have already been observed in other biocarbonates (Rousseau et al., 2006; Farre and Dauphin, 2009). They may be either very common lipids or compounds involved in the biomineralisation mechanisms. A very polar compound also seems recurrent, although it is not present in Caryophyllia. Further analysis may show, via their structure and type, which roles these compounds may play.
4. Discussion 4.1. Comparison with standards
4.3. Correspondence between the diversity of the skeletal fine structures and the biochemical composition of their organic matrices Coral skeletons were long believed to be uniformly built by fibrous aragonite crystals (Pratz, 1882–1883), diverging from “centres of calcification” (Ogilvie, 1896; Vaughan and Wells, 1943; Wells, 1956). During the last decade, however, various methods have revealed that the fibres, commonly considered as single crystals, are actually composite structures (Cuif and Dauphin,
Acknowledgements This work has been supported by grants of the ANR (contract ANR-06-BLAN-0233, BioCristal project) and by the European Science Foundation (ESF) under the EUROCORES Programme EuroMinScI (BioCalc project, contract No. ERAS-CT-2003-980409 of the European Commission, DG Research, FP6). R. Morgan (UMR IDES) improved the English language. References Baronnet, A., Cuif, J.P., Dauphin, Y., Farre, B., Williams, C.T., 2008. Distribution of chemical components in coral skeletons provides evidence for an overall control of crystallization at a submicrometer scale. Geophys. Res. Abs. 10, 02602, 16077962/gra/EGU2007-A-02602. Bourne, G.C., 1887. The anatomy of the madreporian coral Fungia. Quart. J. Microsc. Sci. 27, 293–324. Bourne, G.C., 1899. Studies on the structure and formation of the calcareous skeleton of the Anthozoa. Quart. J. Microsc. Sci. 41, 499–541. Bryan, W.H., Hill, D., 1941. Spherulitic crystallization as a mechanism of skeletal growth in the hexacorals. Proc. R. Soc. Queensl. 52, 78–91.
B. Farre et al. / Zoology 113 (2010) 250–257 Clode, P.L., Marshall, A.T., 2002. Low temperature FESEM of the calcifying interface of a scleractinian coral. Tissue Cell 34, 187–198. Collier, J.H., Messersmith, P.B., 2001. Phospholipid strategies in biomineralization and biomaterials research. Annu. Rev. Mater. Res. 31, 237–263. Cuif, J.P., Dauphin, Y., 2005a. The environmental recording unit in coral skeletons – a synthesis of structural and chemical evidences for a biochemically driven, stepping-growth process in fibres. Biogeosciences 2, 61–73. Cuif, J.P., Dauphin, Y., 2005b. The two-step mode of growth in the scleractinian coral skeletons from the micrometre to the overall scale. J. Struct. Biol. 150, 319–331. Cuif, J.P., Gautret, P., 1995. Glucides et protéines de la matrice soluble des biocristaux de scléractiniaires Acroporides. C. R. Acad. Sci. Paris Sér. II a 320, 273–278. Cuif, J.P., Dauphin, Y., Freiwald, A., Gautret, P., Zibrowius, H., 1999. Biochemical markers of zooxanthellae symbiosis in soluble matrices of skeleton of 24 Scleractinia species. Comp. Biochem. Physiol. A123, 269–278. Cuif, J.P., Dauphin, Y., Doucet, J., Salomé, M., Susini, J., 2003. XANES mapping of organic sulfate in three scleractinian coral skeletons. Geochim. Cosmochim. Acta 67, 75–83. Dauphin, Y., 2001. Comparative studies of skeletal soluble matrices from some scleractinian corals and molluscs. Int. J. Biol. Macromol. 28, 293–304. Dauphin, Y., Cuif, J.P., 1997. Isoelectric properties of the soluble matrices in relation to the chemical composition of some scleractinian skeletons. Electrophoresis 18, 1180–1183. Dauphin, Y., Cuif, J.P., Williams, C.T., 2008. Soluble organic matrices of aragonitic skeletons of Merulinidae (Cnidaria, Anthozoa). Comp. Biochem. Physiol. B150, 10–22. Falini, G., Albeck, S., Weiner, S., Addadi, L., 1996. Control of aragonite or calcite polymorphism by mollusc shell macromolecules. Science 271, 67–69. Farre, B., Dauphin, Y., 2009. Lipids from the nacreous and prismatic layers of two Pteriomorpha mollusc shells. Comp. Biochem. Physiol. B152, 103–109. Gotliv, B., Addadi, L., Weiner, S., 2003. Mollusc shell acidic proteins: in search of individual functions. Chem. Biochem. 4, 522–529. Guillemin, G., Patat, J.L., Fournié, J., Chétail, M., 1987. The use of coral as a bone graft substitute. J. Biomed. Mater. Res. 21, 557–567. Guillemin, G., Patat, J.L., Meunier, A., 1995. Natural corals used as bone graft substitutes. In: Allemand, D., Cuif, J.P. (Eds.), Biomineralization 93. 7th International Symposium on Biomineralization, 4. Surgical Uses of Biominerals. Bulletin de l’Institut océanographique, Monaco, vol. 14, pp. 67–77. Harland, A.D., Navarro, J.C., Davies, S.P., Fixter, L.M., 1993. Lipids of some Caribbean and Red Sea corals: total lipid, wax esters, triglycerides and fatty acids. Mar. Biol. 117, 113–117. Johnston, I.S., 1977. Aspects of the structure of a skeletal organic matrix, and the process of skeletogenesis in the reef-coral Pocillopora damicornis. In: Proceedings of the Third International Coral Reef Symposium, University of Miami, pp. 447–453. Johnston, I.S., 1979. The organization of a structural organic matrix within the skeleton of a reef-building coral. Scan. Elect. Microsc. 2, 421–431. Koop, K., Booth, D., Broadbent, A.D., Brodie, J., Bucher, D., Capone, D., Coll, J., Dennison, W., Erdmann, M., Harrison, P., Hutchings, O., Jones, G.B., Larkum, A.W., O’Neil, J.,
257
Steven, A., Tentori, E., Ward, A., Williamson, J., Yellowlees, D., 2001. ENCORE: the effect of nutrient enrichment on coral reefs. Synthesis of results and conclusions. Mar. Poll. Bull. 42, 91–120. Lester, D., Bergmann, W., 1941. Contributions to the study of marine products. VI. The occurrence of cetyl palmitate in corals. J. Org. Chem. 6, 120–122. Meyers, P.A., Barak, J.E., Peters, E.C., 1978. Fatty acids composition of the Caribbean coral Manicina areolata. Bull. Mar. Sci. 28, 789–792. Mitterer, R.M., 1978. Amino acid composition and metal binding capability of the skeletal protein of corals. Bull. Mar. Sci. 28, 173–180. Ogilvie, M.M., 1896. Microscopic and systematic study of madreporarian types of corals. Proc. R. Soc. Lond. 59, 9–18. Pratz, E., 1882–1883. Über die verwandtschaftlichen Beziehungen einiger Korallengattungen mit hauptsächlicher Berücksichtigung ihrer Septalstructur. Palaeontographica 29, 81–122. Puverel, S., Tambutté, É., Pereira-Mouries, L., Zoccola, D., Allemand, D., Tambutté, S., 2005. Soluble organic matrix of two scleractinian corals: partial and comparative analysis. Comp. Biochem. Physiol. 141B, 480–487. Rousseau, M., Bedouet, L., Latie, E., Gasser, P., Le Ny, K., Lopez, E., 2006. Restoration of stratum corneum with nacre lipids. Comp. Biochem. Physiol. B145, 1–9. Silliman, B., 1846. On the chemical composition of the calcareous corals. Am. J. Sci. Arts 51, 189–199. Tambutté, É., Allemand, D., Zoccola, D., Meibom, A., Lotto, S., Caminiti, N., Tambutté, S., 2007a. Observations of the tissue–skeleton interface in the scleractinian coral Stylophora pistillata. Coral Reefs 26, 517–529. Tambutté, S., Tambutté, E., Zoccola, D., Allemand, D., 2007b. Organic matrix and biomineralization of scleractinian corals. In: Bäuerlein, E. (Ed.), Handbook of Biomineralization. Wiley, Weinheim, pp. 243–259. Urey, H.C., Lowenstam, H.A., Epstein, S., McKinney, C.R., 1951. Measurement of paleotemperatures and temperatures of the Upper Cretaceous of England, Denmark and the Southeastern United States. Geol. Soc. Am. Bull. 62, 399–416. Vaughan, T.W., Wells, J.W., 1943. Revision of the suborders, families, and genera of the Scleractinia. Geol. Soc. Am. Spec. Pap. 44, 1–363. Veis, A., 2005. A window on biomineralization. Science 307, 1419–1420. Ward, S., 1995. Two patterns of energy allocation for growth, reproduction and lipid storage in the scleractinian coral Pocillopora damicornis. Coral Reefs 14, 87–90. Wells, J.W., 1956. Scleractinia. In: Moore, R.C. (Ed.), Treatise on Invertebrate Paleontology. Part F. Coelenterata. Geological Society of America, Lawrence, pp. 328–344. Yamashiro, H., Oku, H., Higa, H., Chinen, I., Sakai, S., 1999. Composition of lipids, fatty acids and sterols in Okinawan corals. Comp. Biochem. Physiol. B122, 397–407. Yamashiro, H., Oku, H., Onaga, K., Iwasaki, H., Takara, K., 2001. Coral tumors store reduced level of lipids. J. Exp. Mar. Biol. Ecol. 265, 171–179. Young, S.D., 1971. Organic material from scleractinian coral skeletons. I. Variation in composition between several species. Comp. Biochem. Physiol. 40B, 113–120. Young, S.D., O’Connor, J.D., Muscatine, L., 1971. Organic material from scleractinian coral skeletons. II. Incorporation of 14 C into protein, chitin and lipid. Comp. Biochem. Physiol. 40B, 945–958.
Contents lists available at ScienceDirect
Zoology journal homepage: www.elsevier.de/zool
Occurrence and diversity of lipids in modern coral skeletons Bastien Farre, Jean-Pierre Cuif, Yannicke Dauphin ∗ UMR IDES 8148, bat. 504, Université Paris XI, rue G. Clemenceau, F-91405 Orsay, France
a r t i c l e
i n f o
Article history: Received 9 June 2009 Received in revised form 18 November 2009 Accepted 19 November 2009 Keywords: Biomineralisation Intraskeletal lipids Coral microstructure
a b s t r a c t Coral skeletons are composite acellular structures, in which organic macromolecules are intimately associated with mineral phases. Previous studies focussed on proteins and sugars of the soluble organic matrices extracted from the skeletons. Here we report the occurrence of diverse lipids which were extracted from the aragonitic skeletons of seven modern coral species. Using thin layer chromatography, we show that these lipids differ in quantity and composition between the species. Higher proportions of sterols and sterol esters in skeleton extracts as compared to a much higher abundance of waxes and triglycerides in previously studied extracts from scleractinian soft tissues suggest a specific, although not yet determined, role in biomineralisation. The occurrence of intraskeletal lipids along with other organic components should also be taken into account when using coral skeletons as bone allografts, as well as in fossilisation processes. © 2010 Elsevier GmbH. All rights reserved.
1. Introduction Corals, the main reef builders in the tropical seas, are well known for the diversity of the calcareous skeletons built by their anatomically simple polyps. Although simple, these polyps have undergone a remarkable evolution regarding the colonial mode of life, leading to numerous variations in sizes and spatial relationships between the polyps. The skeletons are always produced by the calicoblastic epithelium (=basal part of the ectoderm) of the polyps and in close contact with it (Bourne, 1887, 1899; Clode and Marshall, 2002; Tambutté et al., 2007a). The organisation of the animals is reflected by the organisation of the calcareous skeletons, not only from a morphological point of view but also at the smaller scale of the spatial arrangement, structure and composition of the skeleton building units, i.e. the mineral fibres (Bourne, 1899). Coral skeletons are built from calcium carbonate belonging to the aragonite variety. Due to their long life span (specifically of the large colonial build-ups that can grow for centuries) and fixed positions, corals are largely used as biological archives for palaeoclimate reconstruction. Since this group is also remarkable for the intensity of the “vital effect” (Urey et al., 1951), i.e. the property of each species to record environmental variations with a specific correlation between isotopic fractionation and temperature changes, these biological archives have also been used empirically through the complex and very uncertain procedure of “calibrating” the coral species. A major improvement would result from an understanding
∗ Corresponding author. Tel.: +33 1 69 15 61 17; fax: +33 1 69 15 61 21. E-mail address: [email protected] (Y. Dauphin). 0944-2006/$ – see front matter © 2010 Elsevier GmbH. All rights reserved. doi:10.1016/j.zool.2009.11.004
of the factors influencing the crystallisation process, and notably the role of the organic matrix in which crystallisation occurs. For several decades it has been recognised that glycoprotein compounds are always associated with coral skeletons (Young, 1971; Mitterer, 1978). But only in recent times precise information has been collected about the biochemical control of crystallisation. Far from being crystals of pure aragonite, as commonly assumed (Bryan and Hill, 1941; Veis, 2005), species-specific associations of macromolecules are detectable within the crystal-like fibres (Mitterer, 1978; Cuif and Gautret, 1995; Cuif et al., 1999; Dauphin, 2001; Dauphin and Cuif, 1997; Puverel et al., 2005; Tambutté et al., 2007b). Atomic force microscopy has shown that the building units of the fibres are small round-shaped corpuscles irregularly coated with organic components (Cuif and Dauphin, 2005a,b). Information about the crystallisation process has been obtained by transmission electron microscopy (imaging and diffraction) (Johnston, 1977, 1979; Clode and Marshall, 2002). It has been shown that crystallisation occurs as the final step of mineralisation in the growth layers, the growth units of the fibres (Baronnet et al., 2008). These results emphasise the need for a better understanding of the complex organic matrices that form the hydrogel in which crystallisation occurs. Besides proteins and glucids, lipids are one of the major category of organic components. Lipids are structural components of cell membranes and important signalling molecules. They play a role in energy storage and in the synthesis of materials “through their ability to self-assemble, compartmentalise and template” (Collier and Messersmith, 2001). Two broad classes have been defined for chromatography purposes: simple lipids and complex lipids. Glycerophospholipids (also called phospholipids) are the primary constituents of biomembranes. Although (phospho)lipids are said
B. Farre et al. / Zoology 113 (2010) 250–257
251
Fig. 1. Morphological variety of coral skeletons. (A) Branching colonial Millepora (“fire coral”), with a porous skeleton and small corallites (inserts); Caribbean archipelago, Guadalupe Island. (B) Massive skeleton of Porites, with small adjacent corallites (insert); New Caledonia. (C) Symphyllia, medium-sized corallites with meandroid arrangement; New Caledonia. (D) Pocillopora, with small corallites (insert); Polynesia, Moorea Island. (E) Fungia, one of the biggest solitary corals; New Caledonia. (F) Solenosmilia, with distinct corallites (insert); Porcupine plateau, North-East Atlantic. (G) Caryophyllia, a solitary deep-sea coral; Biscay Gulf, Meriadzec plateau, West Atlantic.
to be involved in biomineralisation processes, their role in extracellular calcareous biominerals is not yet known. Despite some pioneer work, lipids in coral skeletons have so far not been actually investigated. Silliman (1846) described “waxlike” materials extracted from skeletons. Lipids were described in some modern species by Lester and Bergmann (1941), Young et al. (1971), and Meyers et al. (1978). However, while some analyses were done on decalcified skeletons, in other studies lipids were extracted using acetone or ether, therefore comparisons are difficult. Thus, although a number of analyses have shown the presence of lipids in the soft tissues (Harland et al., 1993; Ward, 1995), data on coral skeletons remain scarce. This study, based on several species of corals selected for their microstructural diversity and difference in mode of life, aims at providing a first set of data concerning the presence and diversity of skeletal lipids.
the Gulf of Biscaye, Meriadzec plateau (Fig. 1G) are representatives of the non-symbiotic metabolism.
2. Materials and methods
2.3.1. Scanning electron microscopy (SEM) Several fractured and polished sections were cut perpendicular and parallel to the surface of the septa and walls. Polished sections and some fractures were etched with various acids and enzymes to reveal microstructural features. Detailed procedures of sample preparation are given in the legend of Fig. 7. SEM observations were conducted using a Philips XL30 (Philips, Amsterdam, The Netherlands) at UMR IDES, Université Paris XI.
2.1. Materials Calcareous skeletons are built by several high level taxa of modern and fossil Cnidaria. Among them, we have studied aragonite skeletons, selected for the disparities of their skeleton shapes, phylogeny, metabolism and lifestyle, but all living in a modern marine environment. The Hydrozoa are represented by Millepora alicornis from Guadalupe (Fig. 1A), the skeletons of which are important contributors to tropical reefs. The other six species belong to the Scleractinia. They comprise three symbiotic colonial species: Porites lutea from New Caledonia (Fig. 1B), one of the most used skeletons in palaeoclimate reconstructions; Symphyllia, from New Caledonia, a colonial type with a restricted number of medium-sized polyps (Fig. 1C); and Pocillopora from Moorea island (Polynesia) with small polyps (Fig. 1D). The solitary Fungia repanda from New Caledonia is also a symbiotic coral (Fig. 1E). Two deep-sea corals, the colonial Solenosmilia variabilis from the Porcupine plateau, north-east Atlantic ocean (Fig. 1F) and the solitary Caryophyllia ambrosia from
2.2. Standards Commercial analytic grade standards were used for all experiments. A phospholipid mixture was used for polar lipids: l-␣-lysophosphatidylcholine, l-␣-phosphatidylcholine and l-␣phosphatidylinositol ammonium salt from soybean and l-␣phosphatidylethanolamine from Escherichia coli. Cholesterol was used as standard for sterols, triolein for triglycerides, oleic acid for free fatty acids, stearyl oleate for waxes and cholesteryl oleate for sterol esters. 2.3. Methods
2.3.2. Atomic force microscopy (AFM) AFM observations were obtained with a Dimension 3100 Nanoscope III (Veeco Instruments Inc., Plainview, NY, USA) housed at UMR IDES, Université Paris XI. Samples were imaged at room temperature and in air using tapping mode. Phase images were generated as a consequence of variations in material properties such as viscoelasticity, composition, adhesion, friction, etc. AFM has several advantages over SEM, e.g., samples do not require coatings. However, disadvantages are the small image size and a reduced depth of field in the order of micrometers. Thus, observation of fractured surfaces is difficult, and most samples have to be polished
252
B. Farre et al. / Zoology 113 (2010) 250–257
Fig. 2. FTIR spectra of standards: (A) cholesterol (lipid); (B) chitin; (C) bovine serum albumin (protein).
and cleaned. Depending upon the structures, different preparative processes were used (see details in the legend of Fig. 7). 2.3.3. Fourier transform infrared spectrometry (FTIR) For all species, FTIR spectra were first recorded on undecalcified powder, then on the extracted insoluble organic matrices of selected species, depending on the TLC results. All spectra were recorded at 4 cm−1 resolution with 64 scans with a strong Norton-Beer apodization on a Perkin-Elmer Model 1600 FTIR spectrometer (Perkin-Elmer Inc., Waltham, MA, USA) in the range from 4000 to 450 cm−1 . The spectrometer was equipped with a diffuse reflectance accessory, which permits diffuse reflectance infrared Fourier transform (DRIFT) measurements with high sensitivity for powders. Raw diffuse reflectance spectra appear different from their transmission equivalents, so a conversion was applied to compensate for these differences. All spectra were corrected using the Kubelka–Munk function. The system was
Fig. 3. FTIR spectra of non-decalcified coral skeletons.
B. Farre et al. / Zoology 113 (2010) 250–257
253
purged and permanently maintained under nitrogen to reduce atmospheric CO2 and H2 O absorption. A background spectrum was measured for pure KBr. Sample spectra were automatically ratioed against background to minimise CO2 and H2 O bands. To assess reproducibility, several spectra from the same sample were obtained and the results gave correlation coefficients higher than 95%. As insoluble organic matrices resulting from a decalcification process, two symbiotic samples (Fungia as a solitary genus and Porites as a colonial genus) and a non-symbiotic solitary genus (Caryophyllia) were used.
2.3.5. Extraction and purification of the insoluble organic matrix Powdered samples were immersed in 5 ml of Milli-Q water, then decalcified by progressive addition of 50% acetic acid so that the pH (automatically controlled with a titrimeter) was above 4. The entire extract was directly centrifuged at 21,000 × g for 15 min, which separated the supernatant (soluble) and precipitated (insoluble) fractions. The insoluble fraction was desalted by repeated centrifugations in Milli-Q water and lyophilised.
2.3.4. Removal of the polyps To remove tissues, polyps were decayed by a 3 h immersion in distilled water to destroy the cells. Removal of the remaining living tissues was done by using a water-jet, followed by immersion in 3% NaClO for 1 h to remove organic contaminants, sonicated for 5 min.
2.3.6. Lipid extractions After removal of the polyps and decontamination, the studied material was ground to a powder in an electric mortar. 9.1 g of each sample were soaked in warm chloroform/methanol (1:1, v/v) for three days under mild and constant stirring and sonicated every
Then the samples were rinsed with Milli-Q water and air-dried, then ground to powder.
Fig. 4. TLC lanes of lipid standards.
Fig. 5. TLC lanes of lipids extracted from the aragonitic coral skeletons, and a mixture of standards.
254
B. Farre et al. / Zoology 113 (2010) 250–257
day. The mixture was then centrifuged at 19,500 × g for 5 min to separate the powder and insoluble parts from the solvent-soluble part. The organic solvent-soluble part was finally concentrated by evaporation. 2.3.7. Thin layer chromatography (TLC) 100 l of the concentrated extracts were applied to 10 cm × 10 cm high performance thin layer chromatography (HPTLC) plates (nano-silica gel plates; Macherey-Nagel GmbH & Co. KG, Düren, Germany). A four-stage development process was used. TLC plates were first developed to their full height with pure n-hexane, then again to full height with pure benzene, and finally twice to half height with hexane: ether: acetic acid (70:30:1, by vol.). After drying, the plates were immersed in a mixture of phosphoric acid, 33% acetic acid, sulfuric acid and copper sulfate (5:5:0.5:90, by vol.) for 40 s, and heated at 110 ◦ C for 15 min. The positions were compared with individual standards (Fig. 2) and 10 l of a mixture of the commercial standards. 3. Results
Fig. 5 shows the diversity in quantity and composition of the lipids extracted from the coral skeletons. The samples of Fungia and Solenosmilia show numerous well-stained bands, whereas Pocillopora and Caryophyllia show only faint or no bands. The most common lipids are the strongly polar lipids (phospholipids), which represent a very heterogeneous group of various functions in the organism. They are found in every sample, although in some species in very low amounts (Pocillopora and Caryophyllia; Fig. 5). Sterols are also well represented, with a major (sometimes unique) spot around the cholesterol position for every specimen except Caryophyllia, which displays basically no lipids but the polar ones. Waxes (such as stearyl oleate) and steryl esters (cholesteryl oleate) are also present in every sample, but in very small amounts. They are rare even in the high-lipid samples (Fungia and Solenosmilia), nevertheless present. Free fatty acids (such as oleic acid) are only represented in the most lipid-rich samples, Solenosmilia and Fungia, although some traces may exist in less lipidic samples, but in too low amounts to be noticed. Triglycerides (triolein standard), mostly known as storage lipids, are only present in the two samples with the most lipids, Solenosmilia and Fungia.
3.1. Bulk composition: FTIR data Standards (Fig. 2): cholesterol, chitin and BSA show strong amide A bands. Amide I and amide II bands are strong in BSA and chitin spectra, whereas only a weak band at 1671 cm−1 is visible in cholesterol. Bands between 800 and 1100 cm−1 are strong in cholesterol (Fig. 2A), weak or absent in BSA spectrum. On the opposite, bands due to C–H stretching vibrations (2800–3000 cm−1 ) are stronger than those of amide A, amide I and amide II in cholesterol. Similar bands are present in polysaccharides (Fig. 2B) and proteins (Fig. 2C), but they are weaker than those of amide bands. They are usually said to be characteristic of lipids. All spectra of undecalcified samples show the organo-mineral composition of the coral skeletons, with bands (1 to 4) characteristic of aragonite (Fig. 3). The first part of the 2 doublet (858 cm−1 ) shows that the Sr content of aragonite is high (>7000 ppm). Bands are also present between 4000 and 1500 cm−1 , indicative of the presence of organic components. Amide A, amide I and amide II bands may be assigned to proteins, sugars or lipids. Wave numbers of amide A bands in Scleractinia are higher than 3350 cm−1 , whereas in the Hydrozoa sample (Millepora), this band is lower than 3300 cm−1 . Amide A intensities are variable: high in Millepora and Porites, low in other specimens. Weak bands in the 2800–3000 cm−1 region, usually assigned to lipids, but also present in chitin, are visible as shoulders in all the samples. However, these bands are identified as bands by the software in Fungia (2 bands), Solenosmilia and Caryophyllia (1 band) for Scleractinia, and Millepora (1 band). 3.2. Lipid components Standards were chromatographed separately (Fig. 4). Some resulted in a discrete band (triolein), others in large spots (stearyl oleate, cholesterol). Depending on the used quantity, the oleic acid band is large (Fig. 4), or small (Fig. 5). Standards were also mixed to estimate the ability of the used technique to differentiate a complex mixture (Fig. 5). Because of the overlap between the large band of oleic acid and that of cholesterol when large amounts were used, only a reduced quantity of oleic acid was used in the mixed standard. Cholesteryl oleate and stearyl oleate also show some overlap (Fig. 5). Despite these overlaps, discrete bands are obtained when using low amounts of individual products. Since the same amount of skeleton powder was used for each sample, the relative density of the spots is related to the relative proportion of the lipids.
Fig. 6. FTIR spectra of the insoluble organic matrices of three decalcified coral skeletons.
B. Farre et al. / Zoology 113 (2010) 250–257
The lipid content varies a lot from one sample to another, but some of the major components can be compared. It seems that the depth or life style (i.e. solitary or colonial) have no influence on the lipid yield of the coral. The two corals with the highest lipid content are also the most different: the single, symbiotic, shallow-water Fungia and the colonial, non-symbiotic, deep-sea Solenosmilia. Some lipids are, however, present in all the samples. These lipids are mainly polar lipids and sterol esters. A very polar compound also seems recurrent, although it is not present in Caryophyllia. Finally, despite the similarity of Solenosmilia and Fungia, the two corals display a lipid composition pattern very different from one another. The most relevant differences show up in the polar lipid
255
zone between cholesterol and non-migrating lipids, where they are both in high quantity and displaying a wide diversity of composition. From TLC analyses, Fungia is rich in lipid contents, whereas Caryophyllia is poor; Porites is in an intermediate position (Fig. 5). FTIR spectra of the insoluble matrices obtained from decalcification show strong differences in lipid contents (Fig. 6), despite the different extraction procedures. The intensity of lipid bands (between 2950 and 2850 cm−1 ) decreases from Fungia (Fig. 6A) to Caryophyllia (Fig. 6C), with Porites in the middle position (Fig. 6B). The decalcification process is easy and fast, while the lipid extraction is time consuming and complex. The above results suggest that FTIR spectra on decalcified insoluble organic matri-
Fig. 7. Diversity and similarities in the biocrystallisation of coral skeletons. SEM and AFM phase images. (A/B) Millepora. (A) Section showing the different sensitivity to etching of the centre (dark grey) and lateral skeleton areas with a layered organisation. Polished and etched with formic acid (1%, aqu.) for 35 s. (B) Nanoparticles in the fibres, polished and etched surface, formic acid (1%, aqu.) for 10 s, 20 ◦ C. (C/D) Porites. (C) Polished and etched sections showing the layered growth mode of the skeletal tissue. (D) Aligned nanoparticles in the fibres, polished and etched surface, formic acid (10%) for 7 s. (E) Pocillopora. Regular fibrous fan systems diverging from the centres of calcification (arrows). Differences in the distribution of etching-sensible zones result in the clearly distinct aspect of the fibrous bundles compared to the fibrous tissues of Porites (see panel C). Polished and etched with alcalase for 3 h at 38 ◦ C. (F/G) Fungia. (F) Growth edge of a septum in a Fungia skeleton. Etching of a section perpendicular to the growth direction reveals that formation of a septum can be a polyphased process. Fibres radiate from centres of calcification (e.g. 1, arrows) first creating cylindrical structures (2) which are further covered by layered tissue (3). Polished and etched with a mixture of alcalase (2/3) and NaOH 1 M (1/3) for 4 h at 30 ◦ C, then rinsed with water. (G) Section presented produces a rather homogenous fibrous tissue. Polished and etched with formic acid (1%, aqu.) for 45 s. (H/I) Caryophyllia. (H) Layering of the fibrous tissue of the Caryophyllia septum shows that growth of the fibre is a step-wise process, symmetrical on both sides of the dotted line built by the centres of calcification. (I) Composite nanoparticles in the fibres, polished and etched surface, Milli-Q water for 4 h, 20 ◦ C.
256
B. Farre et al. / Zoology 113 (2010) 250–257
ces may be used to estimate the presence and quantity of lipids in biominerals.
Although the bulk position of the different lipid compounds allows identification of their classes, their exact position may shift from the position of the standard used, due to differences of polarity: the analysed compounds are different from the standard, although they are of the same class. The most important shift is between the cholesterol position and the other mollusk sterols. It displays a long series of compounds, most of them more polar than cholesterol. Triglycerides are only present in noticeable amounts in two samples (Fungia and Solenosmilia), but neither of the samples displays the exact position of the triolein. Solenosmilia has two triglyceride bands, one more polar than that of triolein, and another less polar. The triglycerides of Fungia are a bit less polar than the used standard (triolein). The free fatty acids are, in most of the samples, quite similar to oleic acid, although a second, more polar stain, appears on the Fungia sample.
2005a,b). When submitted to identical etching conditions, the studied species reveal the diversity of the microstructural (Fig. 7A, C, E–H) and nanostructural patterns (Fig. 7B, D, I). This diversity in the sensitivity to etching conditions can be reasonably correlated to the diversity of the organic components, including the lipid diversity. The reported results concerning lipid components show that, in addition to the molecular diversity of proteins and polysaccharides, lipids also contribute to the complexity of the organic blends involved in the crystallisation mechanism producing coral skeletons. Lipid diversity may also contribute to explaining a series of observations not compatible with the postulated uniformity of coral skeletons. For instance, coral skeletons are a good material for allografts of bone, widely used in bone repair surgery (Guillemin et al., 1987, 1995). However, the guiding function of the coral graft in the bone reconstruction process appears to be dependent on the species used for grafting. Thus, with respect to both fine structure and biochemical composition, coral skeletons appear to be produced through a crystallisation process quite similar to the mechanism responsible for shell formation in molluscs, in contrast to what is commonly published.
4.2. Comparison of lipids in the soft and skeletal parts of corals
5. Conclusion
Most of the data concerning the lipids in the coral skeletons are dealing with the soft tissues. Yamashiro et al. (1999), for example, studied the lipid content of the soft parts of 15 Cnidaria, among them eleven symbiotic zooxanthellate corals. Although the analytical methods are not the same, the chromatograms show that the lipid contents of the soft and skeletal parts of a given coral differ. The most important differences show up in the distribution of waxes and sterol esters: the soft parts generally contain much more waxes than sterol esters, whereas the skeletons show almost no waxes, but a much higher yield of sterol esters. Even more significant is the comparison of the lipid pattern of Porites lutea, which has been analysed both in the soft parts (Yamashiro et al., 1999) and the skeleton (this study). The relative abundance of the different lipids strongly differs from one tissue to another: the dominant lipids in soft tissues are waxes and triglycerides, while in the skeleton, the major lipids are polar lipids and sterols, along with sterol esters. Fatty acids and sterols are almost equivalent. Other lipid classes are almost not represented. This suggests that the lipids found in the hard parts of the corals are not just trapped parts of the living organism. The relative increase of sterol and sterol esters suggests that the lipids in the skeleton may be traces of remnant hormonal signals. However, more research, and specifically analytical chemistry, will be necessary to precisely identify the compounds and their specific roles. It is now demonstrated that the lipid content varies from one species to another, but also according to environmental conditions. Other parameters have also been identified as factors of variation of the lipid content: health or diet mode (Koop et al., 2001; Yamashiro et al., 2001).
Although most biochemical analyses of biocarbonates in corals focussed on either proteins or sugars extracted from the soluble organic matrices (Falini et al., 1996; Dauphin and Cuif, 1997; Cuif et al., 1999, 2003; Gotliv et al., 2003; Dauphin et al., 2008), it appears that lipids are also present in sufficient amounts to be analysed in the cnidarian hard parts, as well as in mollusc shells. The combined analysis with TLC/FTIR highlighted the presence of cholesterol-based lipids, mostly sterol compounds, but also sterol esters. These compounds have to be further examined to be accurately identified, both structurally and in their usual role in the living organisms. However, the high proportion of sterols and sterol esters suggests that several steroids may be involved in the biomineralisation process. Some lipids, mainly polar lipids and sterol esters, are present in all the studied coral skeleton samples. Those lipids have already been observed in other biocarbonates (Rousseau et al., 2006; Farre and Dauphin, 2009). They may be either very common lipids or compounds involved in the biomineralisation mechanisms. A very polar compound also seems recurrent, although it is not present in Caryophyllia. Further analysis may show, via their structure and type, which roles these compounds may play.
4. Discussion 4.1. Comparison with standards
4.3. Correspondence between the diversity of the skeletal fine structures and the biochemical composition of their organic matrices Coral skeletons were long believed to be uniformly built by fibrous aragonite crystals (Pratz, 1882–1883), diverging from “centres of calcification” (Ogilvie, 1896; Vaughan and Wells, 1943; Wells, 1956). During the last decade, however, various methods have revealed that the fibres, commonly considered as single crystals, are actually composite structures (Cuif and Dauphin,
Acknowledgements This work has been supported by grants of the ANR (contract ANR-06-BLAN-0233, BioCristal project) and by the European Science Foundation (ESF) under the EUROCORES Programme EuroMinScI (BioCalc project, contract No. ERAS-CT-2003-980409 of the European Commission, DG Research, FP6). R. Morgan (UMR IDES) improved the English language. References Baronnet, A., Cuif, J.P., Dauphin, Y., Farre, B., Williams, C.T., 2008. Distribution of chemical components in coral skeletons provides evidence for an overall control of crystallization at a submicrometer scale. Geophys. Res. Abs. 10, 02602, 16077962/gra/EGU2007-A-02602. Bourne, G.C., 1887. The anatomy of the madreporian coral Fungia. Quart. J. Microsc. Sci. 27, 293–324. Bourne, G.C., 1899. Studies on the structure and formation of the calcareous skeleton of the Anthozoa. Quart. J. Microsc. Sci. 41, 499–541. Bryan, W.H., Hill, D., 1941. Spherulitic crystallization as a mechanism of skeletal growth in the hexacorals. Proc. R. Soc. Queensl. 52, 78–91.
B. Farre et al. / Zoology 113 (2010) 250–257 Clode, P.L., Marshall, A.T., 2002. Low temperature FESEM of the calcifying interface of a scleractinian coral. Tissue Cell 34, 187–198. Collier, J.H., Messersmith, P.B., 2001. Phospholipid strategies in biomineralization and biomaterials research. Annu. Rev. Mater. Res. 31, 237–263. Cuif, J.P., Dauphin, Y., 2005a. The environmental recording unit in coral skeletons – a synthesis of structural and chemical evidences for a biochemically driven, stepping-growth process in fibres. Biogeosciences 2, 61–73. Cuif, J.P., Dauphin, Y., 2005b. The two-step mode of growth in the scleractinian coral skeletons from the micrometre to the overall scale. J. Struct. Biol. 150, 319–331. Cuif, J.P., Gautret, P., 1995. Glucides et protéines de la matrice soluble des biocristaux de scléractiniaires Acroporides. C. R. Acad. Sci. Paris Sér. II a 320, 273–278. Cuif, J.P., Dauphin, Y., Freiwald, A., Gautret, P., Zibrowius, H., 1999. Biochemical markers of zooxanthellae symbiosis in soluble matrices of skeleton of 24 Scleractinia species. Comp. Biochem. Physiol. A123, 269–278. Cuif, J.P., Dauphin, Y., Doucet, J., Salomé, M., Susini, J., 2003. XANES mapping of organic sulfate in three scleractinian coral skeletons. Geochim. Cosmochim. Acta 67, 75–83. Dauphin, Y., 2001. Comparative studies of skeletal soluble matrices from some scleractinian corals and molluscs. Int. J. Biol. Macromol. 28, 293–304. Dauphin, Y., Cuif, J.P., 1997. Isoelectric properties of the soluble matrices in relation to the chemical composition of some scleractinian skeletons. Electrophoresis 18, 1180–1183. Dauphin, Y., Cuif, J.P., Williams, C.T., 2008. Soluble organic matrices of aragonitic skeletons of Merulinidae (Cnidaria, Anthozoa). Comp. Biochem. Physiol. B150, 10–22. Falini, G., Albeck, S., Weiner, S., Addadi, L., 1996. Control of aragonite or calcite polymorphism by mollusc shell macromolecules. Science 271, 67–69. Farre, B., Dauphin, Y., 2009. Lipids from the nacreous and prismatic layers of two Pteriomorpha mollusc shells. Comp. Biochem. Physiol. B152, 103–109. Gotliv, B., Addadi, L., Weiner, S., 2003. Mollusc shell acidic proteins: in search of individual functions. Chem. Biochem. 4, 522–529. Guillemin, G., Patat, J.L., Fournié, J., Chétail, M., 1987. The use of coral as a bone graft substitute. J. Biomed. Mater. Res. 21, 557–567. Guillemin, G., Patat, J.L., Meunier, A., 1995. Natural corals used as bone graft substitutes. In: Allemand, D., Cuif, J.P. (Eds.), Biomineralization 93. 7th International Symposium on Biomineralization, 4. Surgical Uses of Biominerals. Bulletin de l’Institut océanographique, Monaco, vol. 14, pp. 67–77. Harland, A.D., Navarro, J.C., Davies, S.P., Fixter, L.M., 1993. Lipids of some Caribbean and Red Sea corals: total lipid, wax esters, triglycerides and fatty acids. Mar. Biol. 117, 113–117. Johnston, I.S., 1977. Aspects of the structure of a skeletal organic matrix, and the process of skeletogenesis in the reef-coral Pocillopora damicornis. In: Proceedings of the Third International Coral Reef Symposium, University of Miami, pp. 447–453. Johnston, I.S., 1979. The organization of a structural organic matrix within the skeleton of a reef-building coral. Scan. Elect. Microsc. 2, 421–431. Koop, K., Booth, D., Broadbent, A.D., Brodie, J., Bucher, D., Capone, D., Coll, J., Dennison, W., Erdmann, M., Harrison, P., Hutchings, O., Jones, G.B., Larkum, A.W., O’Neil, J.,
257
Steven, A., Tentori, E., Ward, A., Williamson, J., Yellowlees, D., 2001. ENCORE: the effect of nutrient enrichment on coral reefs. Synthesis of results and conclusions. Mar. Poll. Bull. 42, 91–120. Lester, D., Bergmann, W., 1941. Contributions to the study of marine products. VI. The occurrence of cetyl palmitate in corals. J. Org. Chem. 6, 120–122. Meyers, P.A., Barak, J.E., Peters, E.C., 1978. Fatty acids composition of the Caribbean coral Manicina areolata. Bull. Mar. Sci. 28, 789–792. Mitterer, R.M., 1978. Amino acid composition and metal binding capability of the skeletal protein of corals. Bull. Mar. Sci. 28, 173–180. Ogilvie, M.M., 1896. Microscopic and systematic study of madreporarian types of corals. Proc. R. Soc. Lond. 59, 9–18. Pratz, E., 1882–1883. Über die verwandtschaftlichen Beziehungen einiger Korallengattungen mit hauptsächlicher Berücksichtigung ihrer Septalstructur. Palaeontographica 29, 81–122. Puverel, S., Tambutté, É., Pereira-Mouries, L., Zoccola, D., Allemand, D., Tambutté, S., 2005. Soluble organic matrix of two scleractinian corals: partial and comparative analysis. Comp. Biochem. Physiol. 141B, 480–487. Rousseau, M., Bedouet, L., Latie, E., Gasser, P., Le Ny, K., Lopez, E., 2006. Restoration of stratum corneum with nacre lipids. Comp. Biochem. Physiol. B145, 1–9. Silliman, B., 1846. On the chemical composition of the calcareous corals. Am. J. Sci. Arts 51, 189–199. Tambutté, É., Allemand, D., Zoccola, D., Meibom, A., Lotto, S., Caminiti, N., Tambutté, S., 2007a. Observations of the tissue–skeleton interface in the scleractinian coral Stylophora pistillata. Coral Reefs 26, 517–529. Tambutté, S., Tambutté, E., Zoccola, D., Allemand, D., 2007b. Organic matrix and biomineralization of scleractinian corals. In: Bäuerlein, E. (Ed.), Handbook of Biomineralization. Wiley, Weinheim, pp. 243–259. Urey, H.C., Lowenstam, H.A., Epstein, S., McKinney, C.R., 1951. Measurement of paleotemperatures and temperatures of the Upper Cretaceous of England, Denmark and the Southeastern United States. Geol. Soc. Am. Bull. 62, 399–416. Vaughan, T.W., Wells, J.W., 1943. Revision of the suborders, families, and genera of the Scleractinia. Geol. Soc. Am. Spec. Pap. 44, 1–363. Veis, A., 2005. A window on biomineralization. Science 307, 1419–1420. Ward, S., 1995. Two patterns of energy allocation for growth, reproduction and lipid storage in the scleractinian coral Pocillopora damicornis. Coral Reefs 14, 87–90. Wells, J.W., 1956. Scleractinia. In: Moore, R.C. (Ed.), Treatise on Invertebrate Paleontology. Part F. Coelenterata. Geological Society of America, Lawrence, pp. 328–344. Yamashiro, H., Oku, H., Higa, H., Chinen, I., Sakai, S., 1999. Composition of lipids, fatty acids and sterols in Okinawan corals. Comp. Biochem. Physiol. B122, 397–407. Yamashiro, H., Oku, H., Onaga, K., Iwasaki, H., Takara, K., 2001. Coral tumors store reduced level of lipids. J. Exp. Mar. Biol. Ecol. 265, 171–179. Young, S.D., 1971. Organic material from scleractinian coral skeletons. I. Variation in composition between several species. Comp. Biochem. Physiol. 40B, 113–120. Young, S.D., O’Connor, J.D., Muscatine, L., 1971. Organic material from scleractinian coral skeletons. II. Incorporation of 14 C into protein, chitin and lipid. Comp. Biochem. Physiol. 40B, 945–958.