Identification of two organic bands showing di ff erent chemical composition within the skeleton of Porites lutea : a confocal Raman microscopy study

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Introduction
Scleractinian corals are marine organisms that thrive most vigorously in clear tropical oceans, forming one of the most important marine ecosystems -coral reefs.A thin layer of coral polyps secrete an aragonitic skeleton beneath their basal ectoderm forming an intricate and complex exoskeleton, which represents a chronological layered archive (e.g.Lough and Barnes, 2000;Cohen and McConnaughey, 2003).The fact that environmental information is recorded within these layered structures makes corals an important archive for palaeo climate research.Even though coral reefs have been studied since Darwin's monography (Darwin, 1842) and successfully employed to learn about the past, the fundamental biomineralization processes behind their formation is still not fully understood.
The morphology of skeletal structures of corals represents the foundation of this investigation and will be explained in the following part, followed by a summery of the Figures

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Full different biomineralization concepts that have been developed to explain their formation.
Each single polyp consists of different skeletal elements: a columella in the center, septa that radiate away from the center, a wall (theca and epitheca) and dissepiment bordering the corallite to the lateral and basal end, respectively (Sorauf, 1972; Fig. 1).Adjacent corallites either share common walls or, if not, separated corallite walls are connected by skeletal elements forming the coenosteum.All of these macromorphological elements can be more or less well developed depending on the coral genus (Veron, 2000;Nothurft and Webb, 2007).A close-up of a corallite i.e. viewing individual macro-morphological skeletal elements (such as septa and columella), displays micromorphological features such as spines, granules and nodules, which structure walls, septal margins and septal faces (Sorauf, 1972;Fig. 1).The skeleton of the species studied in this investigation, Porites lutea, are relatively porous and less differentiated into marcromorphological elements (Fig. 1) than other species (Sorauf, 1972;Barnes and Devereux, 1988;Nothurft and Webb, 2007).The skeleton is formed of vertical rods (trabecluae) that are interconnected horizontally by bars called synapticulae or radi and each vertical rod terminates in pali or denticle (Fig. 1a and e) with several spines (Fig. 1c).Both macro-and micro-morphological elements -are composed of two building blocks (microstructural elements) first described by Ogilvie (1896): the centers of calcification (COC) and fibers.The COC -later described as early mineralization zone (EMZ) (Cuif et al., 2004) -build the scaffold for the coral skeleton ultimately responsible for the colony shape, while fibers represent the bulk of the skeleton (e.g.Cohen and McConnaughey, 2003;Nothdurft and Webb, 2007).These different skeletal morphological levels reflect the complex arrangement of coral skeletal elements and it is astonishing how precise they are repeated.
Even though early descriptive studies of coral skeletons focused on the classification of species they also led to the development of biomineralization concepts, of which the first date back to the end of the 19th century (e.g.Pratz, 1882;Stolarski, 2003;Cuif and Dauphin, 2005b).The first biomineralization concepts emphasize the Figures

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Full resemblence of microstructural elements as purely inorganic crystals.These growth concepts are based on a simple physico-chemical precipitation process, where centers act as "germs" and facilitate and direct the growth of aragonite crystals (Bryan and Hill, 1941;Barnes, 1970).Goreau (1959) observed a layer of organic compounds that he interpreted as a template for growth.Young (1971) was one of the first showing that proteins are part of the skeleton.Later scanning electron microscopy (SEM) revealed the incremental growth of coral fibers (Sorauf and Jell, 1977;Cuif and Dauphin, 1998).This insight, together with information from UV-fluorescence examinations, clearly demonstrated microstructural and chemical differences between fibers and centers of calcification (e.g.Cuif and Dauphin, 1998;Cuif et al., 1999).Since then the number of studies that addressed the distribution (Cuif and Dauphin, 1998;Gautret et al., 2000), composition (e.g.Dauphin and Cuif, 1997;Gautret et al., 1997;Puverel et al., 2005) and origin (Clode and Marshall, 2002;Puverel et al., 2005;Tambutte et al., 2007) of organic components within skeletal structures increased rapidly.The incremental growth of coral fibers is expressed in the alternation of organic-rich and organic-depleted growth lines and a layered distribution of trace elements (Meibom et al., 2004(Meibom et al., , 2007) ) as well as sulphated polysaccharides.To explain the formation of these layered structures Cuif and Dauphin (2005b) proposed a two-step mode of growth by introducing a sequential process acting at micrometer scale emphasizing an ectodermal control of biomineralization.The ecotoderm secretes an organic framework (where sulphated polysaccharides potentially play a major role) onto which minerals grow.The high fluctuation of Mg within fibers (Meibom et al., 2004) suggests that Mg might be incorporated to suppress crystal growth and end the growth cylce (Cuif and Dauphin, 2005b).Stolarski (2003) supported the alternating nature of growth in corals but challenged the proposition of differing temporal formation of EMZ and fibers.
In summary all these observations led to the perception that the formation of a coral skeleton involves the regulative control of dedicated cells/organic molecules during the different stages of growth, rather than "simple inorganic" precipitation of a mineral from a liquid (Cuif and Dauphin, 2005a, b).However, "simple" bio-inorganic models Introduction

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Full still prevail pointing out the morphological and compositional similarity of abiotic precipitated aragonite under cyclically varying saturation state and pH with skeletal fiber growth of coral microstructural elements (Holcomb et al., 2009).This analogy corresponds to the idea of a liquid filled space between epithelium and skeleton, where supersatuartion is maintained to induce crystallization (Furla et al., 2000;Adkins et al., 2003).A critical review of the shortcomings associated with the latter perception can be found in Cuif et al. (2012).
The recent biomineralization concepts explain processes on the micrometer length scale such as growth of fibers around the EMZ.However, they do not provide a linkage to macroscopic structures such as synapticulae joining skeletal elements in regular intervals (Fig. 1) or to temporal differences in formation of different skeletal elements and areas, such as secondary thickening of septa and the formation of dissepiments at the base of the polyp (Nothdurft and Webb, 2007).In coral skeletons, the distribution of the organic matrix -a term encompassing all organic compounds within coral skeletons (Tambutt é et al., 2011) -with respect to crystal arrangement has been thoroughly described and attributed a function in calcification (Cuif and Dauphin, 2005b).This skeletal organic matrix was shown to be tightly associated with the mineral phase down to submicrometer scale delineating nano-crystals (Cuif and Dauphin, 2005a;Stolarski and Mazur, 2005) and act as a pre-oriented organic framework that potentially directs the self-assembly of the nano-crystals to form crystal fibers (Cuif and Dauphin, 2005a).Several studies described the different functions for organic compounds extracted from the organic matrix of biominerals.These functions involve mechanisms such as the binding of Ca (Watanabe et al., 2003;Endo et al., 2004), stabilization of amorphous calcium carbonate (Addadi et al., 2003) or inhibition of growth (Wheeler and Sikes, 1984;Marin et al., 1996).According to Cuif et al. (2011, p. 177) "present knowledge of the distribution of organic compounds at the submicrometer scale completely refutes any (. . . ) accidental inclusion of organic compounds within skeletons".However, the exact function on site within the skeleton and the role in the formation Introduction

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Full of different structural elements (and potentially the three-dimensional arrangement of macromorphological skeletal elements) is still elusive.The high spatial resolution (sub-micrometer scale) of Confocal Raman Microscopy (CRM), has been demonstrated to be ideal to describe the structural relation between organic and inorganic phases in biogenic materials (e.g.Hild et al., 2008;Nehrke and Nouet, 2011;Neues et al., 2011;Nehrke et al., 2012).Recently, one study (Zhang et al., 2011) used CRM to map the skeleton of a blue coral (Helipora coerulea, Octocorallia).They mapped a small region providing a preliminary insight into the ability of CRM to relate the distribution of organic compounds to the mineral phase in coral skeletons.In this study we applied CRM in combination with Polarized Light Microscopy (PLM), SEM and Electron Micro Probe mapping (EMP) to the scleractinian coral species Porites lutea (Milne Edwards and Haime, 1860.The main focus was the organic matrix distribution in relation to the mineral phase, in particular the organicrich (ORGL) and organic-depleted (ODGL) growth lines (as first described in Cuif and Dauphin 1998) that form a growth layer (the so-called environment recording unit as described by e.g.Cuif and Dauphin 2005a) and how they correlate with the distribution of the elements Sr, Mg, and S. Furthermore it should be investigated how the information obtained by CRM can be compared to the information obtained by long established methods like PLM and SEM.

Coral sample
The scleractinian coral Porites lutea (Milne Edwards and Haime, 1860) was collected by SCUBA divers from an off-shore island in the Andaman Sea, Thailand in March 2011.Coral tissue was removed by submerging the specimen in a 5 % sodium hypochlorite solution for 24 h, and subsequently rinsing with de-ionised water and drying at 60 Full The flow chart in Fig. 2 provides an overview of samples (with sample ID) derived from this specimen and gives an outline of the preparation and measurements applied.After tissue removal, three longitudinal and one transversal block (Fig. 1) were cut out of the skeleton and all but one of the longitudinal skeletal blocks were embedded in an epoxy resin (Araldite 2020, Huntsmann).All samples were ground using HERMES water grinding papers (in the order P1200, P2400, and P4000) and polished with a Struers diamond suspension of 3 µm and finished with a 0.3 µm aluminum oxide suspension.Each grinding and polishing step was followed by rinsing the sample with de-ionised water as well as a short cleaning using an ultrasonic bath.To determine how the observed structures continue into deeper levels of the skeleton, a vertical skeletal rod (trabeculae) of one sample was mapped by CRM in three different levels.This was done by re-grinding and polishing after each mapping to remove ∼ 10 µm of the sample.

Confocal Raman microscopy
Raman mapping was done using a WITec alpha 300 R (WITec GmbH, Germany) CRM.Scans with a high spatial resolution were performed using a piezoelectric scanner table having a maximum scan range of 200 µm × 200 µm and a minimum step size of 4 nm lateral and 0.5 nm vertical.An ultra high throughput spectrometer (UHTS 300, WITec, Germany) equipped with en EMCCD camera was used with a grating of 600 grooves mm −1 , blazed at 500 nm.This set-up allows for a very short integration time (down to a few ms) and a spectral range from 0-3600 cm −1 or 0-4000 cm −1 depending on the wavelength used (532 nm or 488 nm, respectively).The Raman instrument could also be operated as a normal light microscope using transmitted and reflected light and was equipped with an polarizing filter (one before the sample -the polarizer and one after the sample -the analyzer).The analyzer can be used in the range from 500 to 800 nm allowing on the one hand the operation of the microscope for PLM and on the other hand for polarized Raman microscopy (at 532 nm).All Raman maps were obtained using a Nikon 100x (NA = 0.9) objective, with the polarizer Introduction

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Full set to 0 • and the analyzer to 90 • .The spectra during mapping were recorded with a step size of 0.5 µm and an integration time of 50 ms or 10 ms for 532 or 488 nm, respectively.Raman measurements of biogenic materials are often hindered by strong fluorescence, which overlay distinct Raman lines.However, as shown by e.g.Nehrke and Nouet (2011) for the shell of the snail Nerita undata, fluorescence intensity distribution of a region can be used as a proxy to map organic matrix distribution within biogenic minerals.Thus, we used the spectral range between 2400-2700 cm −1 to map the fluorescence intensity distribution across the sample.The spectral analysis and image processing was performed using the WITecProject software (verion 2.04, WITec GmbH, Germany).

Scanning electron microscopy
Prior to SEM analysis the samples were etched using 0.1 % formic acid and 3 % glutaraldehyd solution for 50 s to make microstructural features visible (Waite and Andersen, 1980; Cuif and Dauphin, 1998;Stolarski, 2003).After etching the sample was air dried, put on aluminum stabs, and sputter-coated using a plantinum-gold target.Scanning electron microscopic images were acquired at 10 kV or 15 kV and 1.7 µA filament current using a Philipps XL 30 Environmental Scanning Electron Microscope (ESEM).

Electron microprobe mapping
Electron Micro-Probe mapping (EMP: JXA-8200 JEOL, Geomar) was used to study minor and trace element distribution in relation to the structures determined by CRM.
The EMP maps were obtained by wavelength dispersive spectrometry (WDS) mode measuring simultaneously Mg (Ka, TAP), Sr (La, TAP) and S (Ka, PETH).The electron beam was focused to a spot size of 1 µm, accelerating voltage set to 15 kV and beam current to 20 nA.A step size of 1 µm as well as an accumulation time of 20 ms was used and the map was repeated to gather 10 accumulations of the selected area.Standards Introduction

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Full (Vulcanicglass -VG-2) were measured before mapping the sample to calculate concentrations of the trace elements.

Linear extension rate and growth layer thickness
The coral colony was stained in situ using alizarin red (15 mg l −1 , approx.16 h) three months before collection.Daily growth rates can be calculated from this staining experiment by measuring the distance from the staining line (visible in longitudinal sections cut along the major growth direction of corallites) to the colony outer surface with a caliper and dividing this distance by the number of days between staining and collection.Porites skeletons consist of vertical rods (trabeculae) that are built of EMZ and fibers (Fig. 3).The fibers are deposited to both sides of the EMZ and composed of incremental growth layers (Cuif and Dauphin 2005a).The growth layer thickness was measured perpendicular to the outer coral mineralizing epithelium and comprises a pair of an ORGL and ODGL (see Fig. 3).The vertical growth rate (determined from the staining experiment) was compared to growth layer thickness (determined using CRM fluorescence mapping).Differences in vertical and lateral extension rates result from the oblique arrangement of the growth layers with respect to the vertical line of EMZ.
Both growth layer thickness and the angle between growth layers were measured in all Raman maps that displayed the alternation of a ORGL and ODGL using the computer program imageJ.

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Full differences in crystallographic orientation of aragonite (Fig. 4) were visualized by using the relative peak intensity changes as shown by Nehrke and Nouet (2011) for the orientation of aragonite prisms in a gastropod shell.The images show the typical fan like fibers described by Cuif and Dauphin (2005b) for scleractinian coral skeletons using PLM (Fig. 4b and c).The intensity distribution of the major aragonite peak can be used to visualize changes in crystal orientations.The visualization of these orientations was possible in embedded as well as non-embedded polished samples (Fig. 5).
Hence, structural information of the mineral phase and relative orientation can be derived without preparation of thin section, as required for PLM.This allowed determining the distribution of organic compounds in relation to different structural elements as will be shown below.
Fluorescence intensity maps allow visualizing the distribution and arrangement of the organic matrix within coral skeletons, as revealed by means of acridine orange staining (Cuif and Dauphin, 1998;Cuif et al., 1999;Gautret et al., 2000;Stolarski, 2003).Both the distribution of incremental ORGL and the EMZ (e.g.Cuif and Dauphin, 1998) could be visualized using CRM (Fig. 6).The mapped organic matrix distribution was confirmed by a direct comparison with SEM micrographs after etching (Fig. 7).The superimposed maps of crystal orientation and fluorescence (Fig. 5 and 6) confirmed the synchronism in formation of growth layers between adjacent fibers (Cuif et al., 2004).Therefore, Raman mapping can simultaneously determine differences in crystallographic orientation, the distribution of incremental growth layers, and the location of the EMZ within corals without the alteration by staining.This improves the interpretation of data while reducing the number of different sample preparation steps and therewith the possibility of sample alteration.

Determination of growth rates and daily growth cycles
Based on the previously described results, fluorescence images can be used to derive information on growth layer thickness.As described in the Sect.2.5 a growth layer comprise of an ORGL and a ODGL here reflected by a high and a low fluorescence Introduction

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Full line.These two together illustrate the stepping growth mode of coral fibers (e.g.Cuif and Dauphin 2005b).The distances of growth layers were measured perpendicular to the growth lines as explained in Sect.2.5 (indicated by the position of the scale bar within fluorescence maps, e.g.Fig. 6a2) and equals to a distance of 3.9 ± 0.2 µm (mean ± SE; n = 18).These layers encompass on average an angle of 36 ± 2 • (mean ± SE; n = 16) and hence, a growth layer results in a mean vertical extension of 12 µm (see Fig. 3; calculation: growth layer thickness / sin (angle between growth layers / 2)).This vertical extension (often referred to as linear extension) was compared to the mean daily growth rate (in vertical direction) of this specimen of approx.30 µm day −1 .These data suggest that per day 2-3 growth cycles occur.Meibom et al. (2007) compared average linear extension rates to distances between high Mg bands determined by means of nanoSIMS measurements (on Porites sp.) and suggested that within a day up to five growth cycles could be deposited.In his measurements, one growth layer was between 2-5 µm wide, thus, both -cyclicity and growth layer thickness correspond to the values obtained in this study.Cuif et al. (2011) obtained 10-15 cycles per week (∼ 2 cycles per day) for Galaxea fascicularis from a consecutive calcein staining experiment.Thus, this study and the study by Meibom et al. (2007) observed a slightly higher number of cylces per day for Porites spp.than Cuif et al. (2011) for Galaxea fascicularis.Nothdurft and Webb (2007) observed that a vertical extension of 384 µm equals to 36 µm in lateral growth by tracking growth lines in a skeletal rod of Porites lobata.
This vertical distance was deposited within 12 days (derived from mean growth rates for this species; see Nothdurft and Webb 2007) and equals a lateral growth of 3 µm per day.Based on the growth rates determined in this study and the study by Meibom et al. (2007), 3 µm lateral growth corresponds to approximately one growth cycle per day.
The differences in growth cycles per day summarized here could be of various nature: (1) the coral was stressed from the staining process (Thebault et al., 2006;Houlbreque et al., 2009), (2) effect of the nutritional status (Cuif et al., 2011), (3) daily growth cycles are species-specific (Cuif et al., 2011), and/or (4) growth rates could differ depending on the location within the skeleton for wich they are determined.However, Introduction

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Full especially a nonlinear growth rate in lateral and longitudinal direction and/or different growth rates depending on the distance from the EMZ could explain the observed decrease in number of growth cylces when tracing growth lines over larger areas.Two types of ORGL could be detected in this study, which differ in thickness and intensity.From here on the wide growth lines showing increased fluorescence (indicated by a blue arrow within Raman fluorescence maps; Fig. 5-7) are referred to as ORGL 2 (Fig. 6B2, b) whereas the thinner incremental growth lines are referred to as ORGL 1 (Fig. 6B2, a).It could be shown that the ORGL 2 continue in deeper skeletal layers of a vertical rod (Fig. 8a-c).Moreover, these lines resemble the corals outer surface forming a three-dimensional organic envelope.Within a skeletal rod the location of this growth line is slightly changing (Fig. 8), in particular in regions where other skeletal elements, such as radi or synapticulae arise.Early mineralization zones are known to build the central skeletal framework directing growth.The organic envelopes described here indicate former outer surfaces and could potentially play a role as initiation points for interconnecting skeletal elements where new EMZ are formed.

Correlation between organic matrices and elemental composition
EMP mapping showed that Sr is slightly variable within the sample without a distinct pattern (Fig. 9).On the other hand, the Mg distribution displayed a banding pattern, which correlates with the fluorescence bands determined by CRM (Fig. 9).Mapping of S showed elevated concentrations in the areas representing the EMZ (Fig. 9).These findings qualitatively agree with previous studies (Cuif et al., 2003;Meibom et al., 2004Meibom et al., , 2007Meibom et al., , 2008)), even though the resolution of EMP is not comparable with high-resolution techniques like XANES or nanoSIMS (Cuif et al., 2003;Meibom et al., 2004).Therefore, a cyclic variation of S within fibers could not be resolved in this study.
As described above, two different types of ORGL were identified by CRM (named type one and type two).The ORGL 1 show elevated Mg concentrations whereas the ORGL 2 are depleted in Mg.In general it is assumed that in corals Mg is associated with organic compounds (Finch and Allison, 2008), which often also show increased S 8284 Introduction

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Full concentrations (Cuif et al., 2012).However, the growth lines of type two do not follow this pattern but show decreased Mg concentration and no pattern in S (Fig. 9).The low Mg ORGL 2 most likely correspond to the distinct low Mg lines described by Meibom et al. (2004, their  ).Therefore, they would represent a structural feature not restricted to the skeleton of P. lutea.Cuif and Dauphin (2005b) proposed that Mg incorporation in incremental growth lines (type one) functions to repress growth before another layer is added.The ORGL 2 identified in this study, however, are characterized by a decreased Mg concentration and therwith a potentially different function.

Growth patterns and cyclicity
Nothdurft and Webb (2007) pointed out that adjoining structural elements like dissepiment and septa are formed at different times.This temporal heterogeneity is not taken into account in existing growth models and therefore requires their adaptation above the microstructural level.The temporal heterogeneity in skeletal formation also becomes obvious when trying to track spines along the calice into deeper parts of the skeleton, where it appears that they are not present.Hence, different regulative processes must act to produce the spine-rich distal end of the corallites, the skeletal layer covering spines above dissepiments before the skeleton is cut off from the polyp as well as the formation of synapticulae at a regular interval.Position and shape of the ORGL 2 resemble either denticle finger-like structures, traces of former spines or previous skeletal surfaces.Hence, the ORGL 2 which differs in elemental composition and location compared to previously described ORGL 1 arrangements, could be involved in regulating three-dimensional growth patterns of micro-and macroarchitectural elements.We hypothesize that these ORGL 2 act as an "envelope" representing the outer surface at a certain time of deposition (Fig. 10c), before it becomes overgrown and covered by a next growth cycle.The temporal units 1-3 in Fig. 10c represent different growth cycles with 1 being the oldest and 3 the youngest deposited skeleton.Each growth cycle lasts about two weeks and ends with an ORGL 2 (Fig. 10c).Within Introduction

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Full this biweekly cycle elongation, infilling between spine tips and thickening of macromorphological elements takes place (Fig. 10d).This periodicity in growth agrees with the growth model proposed by Nothdurft and Webb (2007).They traced a growth band in their study (Nothdurft and Webb 2007;their Figs. 16c and 17b) that resembles the ORGL 2 described in this study (Fig. 10b) with a similar longitudinal extension spanning approx.12 days of growth.This observation supports the proposed mechanism of a three-dimensional structural regulation by ORGL 2 in Porites skeletal growth that follows a biweekly periodicity.
Cyclicity represents an ubiquitous characteristic of coral skeletal formation.The described cycles range from the cm-level displayed by seasonal density bands (e.g.Lough and Barnes, 2000; Fig. 1) down to the µm-level of daily/sub-daily formation of incremental growth layers (e.g.Meibom et al., 2007).High-resolution proxy studies detected a cyclicity of measured proxy values in the range of weekly to monthly periods (Meibom et al., 2003;Rollion-Bard et al., 2003;Cohen and Sohn, 2004;Allison et al., 2010Allison et al., , 2011)).In these studies the proxy relation (e.g.Sr / Ca) did not only depend on the target parameter (e.g.T ) but was potentially influenced by other parameters like: (1) metabolic changes involving spawning and larval release at a lunar periodicity (Meibom et al., 2003), (2) weekly tidal forcing overlying temperature forcing (Cohen and Sohn, 2004) and (3) a strong bi-weekly periodicity that was hypothesized to result from increased calcification required for the formation of synapticulae (Cohen and Sohn, 2004).The latter corresponds to the biweekly growth bands (ORGL2) described in this study.

Chemical characterization of organic compounds
So far we have used the fluorescence intensity determined by CRM as a "black box" for organic molecules.This concept is sufficient since the scope of this investigation is on structural information of coral skeletons in relation to three-dimensional growth patterns and not the characterization of organic compounds.The emphasis was laid on obtaining Raman maps of high spatial resolution (requiring a hundred thousand spectra per Introduction

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Full scan) hence, it was not possible to obtain spectra with high spectral resolution at the same time.However, it should be pointed out that CRM is much more powerful and can deliver more detailed information for the chemical characterization.Most studies that applied CRM on coral skeleton derived single spectra.Their aim was to check for different carbonate polymorphs (with a spectra range up to 1300 cm −1 ; e.g.Cuif and Dauphin, 1998;Clode et al., 2011) and only few studies focused on organic spectral lines (Perrin and Smith, 2007;Zhang et al., 2011).To demonstrate what kind of chemical information can be obtained, we analyzed the spectra of some areas showing increased fluorescence in more detail and compared the spectra to other studies using Raman spectroscopy on biominerals.Spectral lines not attributed to aragonite were detected (Fig. 11) corresponding to asymetric SO 2− 4 bend (646 cm −1 , Jolivet et al., 2008;Zhang et al., 2011), Amid III (1190-1310 cm −1 , Jolivet et al., 2008;Zhang et al., 2011), CH bands (2850-3080 cm −1 , Perrin and Smith, 2007;Jolivet et al., 2008) and OH group (3200-3400 cm −1 , Jolivet et al., 2008).However, there are some difficulties associated with Raman spectral studies of biominerals that need to be considered.In biominerals the mineral phase is generally dominating the spectra and to obtain the very weak organic bands, long integration times are necessary.Additionally, such samples often show high fluorescence that can lead to oversaturation of detectors when long integration times are used (which sometimes can be avoided by decreasing the intensity of the laser, which on the other hand also increases the integration time).Contaminations of the sample during sample preparation such as embedding in epoxy resins need to be considered and accounted for.We used non-embedded samples to reproduce banding pattern of skeletal fibers to exclude the possibility that during grinding and polishing abraded material accumulates predominately at the organic layers.Recent studies used Raman spectroscopy to characterize organic matrix in biominerals (Kaczorowska et al., 2003;Perrin and Smith, 2007;Jolivet et al., 2008;Zhang et al., 2008, 2011Nehrke and Nouet, 2011), but coral Raman spectroscopic analyses covering a spectral region between 100-3200 cm −1 are quite rare.However, organic compound bands were detected by all studies (Kaczorowska et al., 2003;Perrin and Smith, 2007;Introduction Conclusions References Tables Figures

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Full Zhang et al., 2011et al., , this study), although, Kaczorowska et al. (2003) ) compared the Raman spectra of scleractinian corals to pure aragonite and only detected organic bands for precious corals.In contrast, Perrin and Smith (2007, Lobophyllia corymbosa and Leptastrea purpurea) and this study (Porites lutea) both detected organic compounds within scleractinian coral skeletons.In fact, it was demonstrated the ability to observe both differences in quantity (not absolut but relative Fig. 5-7, Jolivet et al., 2008, for otholiths;Nehrke and Nouet, 2011, for molluscs) as well as quality (spatial distribution, Fig. 11a1 and b1, as well Perrin and Smith, 2007) by Raman spectroscopy.These observations have also been derived for other biominerals (Jolivet et al., 2008, for otholiths; Nehrke and Nouet 2011, for gastropods).Zhang et al. (2011) even discussed the potential to retrieve diurnal environmental variability of phosphorus by mapping the intensity distribution of the symmetric P-O stretch.Corals are well known for recording environmental parameter, however, high-resolution analysis revealed some constraints associated with the skeletal structural complexity (e.g.Meibom et al., 2004;Cuif and Dauphin, 2005b;Juillet-Leclerc et al., 2009;Rollion-Bard et al., 2010).Thus, the full capability of CRM to acquire environmental information needs further investigation.

Conclusions
In the present study we used CRM on P. lutea skeleton samples to demonstrated: 1. that two different types of organic-rich growth lines are present.One of these growth lines corresponds to the well known incremental growth lines.In contrast, the second type of organic-rich growth lines showed an elemental composition that differs from the incremental growth lines (low Mg concentration).
2. that mapping by CRM can be used to visualize the differences in crystal orientation, without the necessity to prepare thin sections.
3. that the layered distribution of organic matrices could be shown and simultaneously related to the orientation of fibers.Introduction

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Full 4. that CRM mapping can provide information on skeletal growth patterns by tracking growth lines within a skeletal rod (in the polished plane as well as in depth).
Position and shape of the second type of organic-rich growth line (within the text referred to as ORGL 2) resemble either denticle finger-like structure, traces of former spines or previous skeletal surfaces.Thus, we hypothesize that this growth lines are involved in the three-dimensional arrangement of skeletal macro-to micromorphological elements and represent the outer skeletal surface before another cycle of elongation, infilling and thickening of skeletal components continues.
5. and that Raman spectra of organic compounds can be measured.The latter represents a powerful possibility in the chemical characteriazation of organic compounds wihin coral skeletons.
Hence, high spatial resolution CRM mapping enables to track growth lines over large skeletal areas and relate skeletal enteties (EMZ and incremental growth layer) as well as the organic to the mineral phase.This simultaneously obtained information can definitively deepen our understanding of growth patterns in coral skeletons and improve growth concepts above micrometer scale.Organic compounds are hypothesized to play a crucial role in coral biomineraliatzion.Here we detected an organic-rich growth line that potentially regulates coral growth.EMP maps provided a first indication of how to complement CRM studies and added new information.Instruments of high spatial resolution (e.g.nanoSIMS, or XANES) are crucial in the investigation of biogenic materials but are restricted by the small area that can be mapped.Thus, a combination of these methods with with CRM is highly valueable.CRM allows to idientify the area of interest prior to measurements by these methods, which are often difficult to access and are cost intensive.

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Figure 1 :
Figure 1: Scanning electron microscopic (SEM) images and X-radiograph positive of Porites 3 lutea (Milne Edwards and Haime, 1860).A) Schematic representation of a Porites corallite 4 indicating the macromorphological elements.B) Distal view of ceroid corallites.C) Detail 5 showing the spines structuring septal walls.D) X-radiograph of the coral skeleton displaying 6 yearly density banding pattern.Arrow indicates view direction in B and rectangles show how 7 skeletal blocks were cut out of the skeleton to obtain transversal and longitudinal samples.E) 8 Scanning electron microscopic (SEM) image of a longitudinal section displaying skeletal 9 elements forming the corallite.10

Fig. 1 .Figure 2 :
Fig. 1.Scanning electron microscopic (SEM) images and X-radiograph positive of Porites lutea (Milne Edwards and Haime, 1860).(A) Schematic representation of a Porites corallite indicating the macromorphological elements.(B) Distal view of ceroid corallites.(C) Detail showing the spines structuring septal walls.(D) X-radiograph of the coral skeleton displaying yearly density banding pattern.Arrow indicates view direction in (B) and rectangles show how skeletal blocks were cut out of the skeleton to obtain transversal and longitudinal samples.(E) Scanning electron microscopic (SEM) image of a longitudinal section displaying skeletal elements forming the corallite.

Fig. 2 .Figure 3 :
Fig. 2. Flow chart displays the chronology of preparation and measurement steps for individual samples (ID).For confocal Raman measurements, the number of maps and laser wavelength used for each map are also given.(Within subsequent figures Raman maps are denoted by: ID map, e.g.#22 29MW #1).

Fig. 3 .
Fig. 3. Schematic representation of coral skeletal growth of a trabeculae (modified after Cuifand Dauphin, 2005a).Stepping growth mode of fibers results in growth layers of organic-rich (ORGL, dark grey) and organic-depleted growth lines (ODGL, light grey) to both sides of the early mineralization zone (EMZ).The growth layers are oblique and encompass an angel resulting in differences in vertical and lateral extension per growth cycle.

Fig. 5 .Figure 6
Fig. 5. Raman maps of longitudinal non-embedded (A; #10 29MW #1) and embedded (B: #40 29MW #1) thick sections of Porites lutea (Milne Edwards and Haime, 1860).(A4-B4) Reflected light images with insert showing the location of the Raman maps.The number of each Raman map (from sample A and B) indicates the spectral region used for mapping: (1) represents the intensity distribution of the symetric stretch of aragonite (1085 cm −1 ), (2) the fluorescence intensity distribution within the area and (3) shows the superposition of map (1) and (2).Blue arrows indicate repeated features of organic-rich growth lines (ORGL 2) of increased fluorescence detected within fluorescence maps.The scale bar within fluorescence maps dispays the skeletal extension for each growth cylce comprised of one high and one low fluorescence growth line.(Superimposed images derive from WITecProject software).

Fig. 6 .Figures 7 :
Fig. 6. (A-C) Raman maps of Porites lutea (Milne Edwards and Haime, 1860) skeleton elements in longitudinal thin section (#22 29MW #2-4).Numbers indicate the spectra region used for the maps: (1) represents the intensity distribution of the symetric stretch of aragonite (1085 cm −1 ), (2) the fluorescence intensity distribution and (3) shows the superposition of map (1) and (2).Blue arrows indicate repeated features of organic-rich growth lines (ORGL 2) of increased fluorescence detected within fluorescence maps.White arrow heads represent the oblique traces of the position of mineralizing epithelium forming layered growth increments with a high and a low fluorescence growth line comprising one growth cylce.The scale bar within fluorescence maps dispays the skeletal extension for each growth cylce.The two types of growth lines within fluorescence map are indicated in (B2) with (a) representing the incremental growth layers (organic-rich and organic-depleted growth lines) and (b) the high fluorescence growth lines (ORGL 2).(Superimposed images derive from WITecProject software).

Fig. 7 .Figure 9 :
Fig. 7. Raman maps of transveral thin section represent the intenstity distribution of the symetric stretch of aragonite (1085 cm −1 , A1, B1) and the fluorescence intensity distribution (A2, B2).(A3, B3) Scanning electron microscopic images were obtained after Raman mapping and etching of sample (A: #20 29MW #1, B: #20 29MW #2).Circle in the images illustrates same areas within the different maps for each row.Blue arrows indicate repeated features of organicrich growth lines (ORGL 2) of increased fluorescence detected within fluorescence maps.The scale bar within fluorescence maps dispays the skeletal extension for each growth cycle comprised of one high and one low fluorescence growth lines.Asterisk indicates same regions within the same row of maps.

Fig. 9 .
Fig. 9. Trace element concentrations (Mg, S, Sr) were mapped with electron microprobe.Mg distribution of the mapped area was superimposed on the respective fluorescence intensity distribution map derived from Raman mapping (A: Fig. 3b2, #22 29MW #3, B: Fig. 3c2, #22 29MW #4).On the right side of the superimposed images (Adobe Photoshop CS4), the trace element maps of S and Sr are displayed.