Biol. Bull. 211: 193-202. (October 2006)
© 2006 Marine Biological Laboratory
Light-Enhanced Calcification and Dark Decalcification in Isolates of the Soft Coral Cladiella sp. During Tissue Recovery
E. Tentori1,* and
D. Allemand2
1 Central Queensland University, Bruce Highway, Rockhampton QLD 4702, Australia
2 Centre Scientifique de Monaco. Av Saint Martin MC 98000, Principaute de Monaco
* To whom correspondence should be addressed, at School of Biological Sciences, The University of Sydney, NSW 2006, Australia. E-mail: ttentori{at}bio.usyd.edu.au
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Abstract
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Light-enhanced calcification is a general characteristic of zooxanthellate corals, suggesting a link between calcification by the coral and photosynthesis by the zooxanthellae, but the relationship between zooxanthellae and coral hosts during this process has not been elucidated. We hypothesized that the effects of tissue injury on the coral fragments used in experiments studying calcification might obscure that link. To detect the effects of tissue injury on light-enhanced calcification, we measured calcification rates (sclerite formation) in the soft coral Cladiella sp. by the alkalinity anomaly method during a 36-day experiment following injury associated with coral fragmentation. In the 2 weeks after colony fragmentation, the calcification response did not show a relation with light intensity. The typical light-enhanced calcification pattern was not noticed until day 15 of tissue recovery. The calcification rate of this soft coral increased with light intensity and time of tissue recovery and was comparable to that of hard corals exposed to similar experimental conditions. However, Cladiella sp. decalcified in the dark. The diurnal calcification-decalcification cycles probably control sclerite size and shape.
Abbreviations: HL, high light • LL, low light • NL, no light
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Introduction
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Studies of the short-term effect of light on calcification rates of zooxanthellate scleractinian corals indicate that photosynthesis by symbiotic algae is involved in the process (see reviews by Barnes and Chalker, 1990; Gattuso et al., 1999). Kawaguti and Sakumoto (1948) inferred that calcification occurs in the light but not in the dark by measuring variations of levels of Ca2+ in the seawater in which they incubated the corals. Using 45Ca, Goreau (1959) also noted the favorable effect of light on coral calcification. Pearse and Muscatine (1971) introduced the concept of light-enhanced calcification in coral physiology and strongly suggested that zooxanthellae from the base of coral branches participate in the calcification of the tips of branches in Acropora cervicornis. Subsequent research has shown overwhelmingly that calcification rates are greater in the light than in the dark in various species of tropical and temperate scleractinian corals (Gattuso et al., 1999) and that calcification is biologically controlled. The process depends upon (1) the calcifying cells in the aboral epidermis; (2) the active transport of calcium and inorganic carbon from the animal cells to the skeleton; and (3) the organic matter as a nucleating center of calcium carbonate crystals. Symbiotic algae may supply the coral with the necessary organic carbon and metabolic energy, they may create the optimum chemical microenvironment for crystallization of CaCO3, or they may do both.
Scleractinian corals, coralline algae, and green algae are the main calcifying organisms of coral reefs. Symbiotic benthic foraminiferans and molluscs as well as bryozoans and echinoderms are also regarded as important reef calcifiers (Barnes and Chalker, 1990; Done et al., 1996; Langer et al., 1997; Gattuso et al., 1999; Yamano et al., 2000). The level of contribution by each of these taxa to reef carbonates varies with geographic region and is difficult to assess because different methods of study have been used in obtaining the data.
Soft corals (Octocorallia: Alcyonacea) are a major benthic component of Indo-Pacific reefs in particular, and they have a high level of ecological interaction through their production of secondary metabolites (Coll, 1992). Soft coral spiculite (the massive skeletons formed by sclerites cemented in aragonite) appears in fossil Quaternary coral reefs at various locations in the Indo-Pacific (Johnson and Risk, 1987; Accordi et al., 1989; Cabioch et al., 1995). Examples of spiculite in living Sinularia species are known (Konishi, 1981; Schumacher, 1997); however, today soft corals appear to provide only sand particles or cementing material.
The species-specific control of calcification in soft corals is implicit in the extensive use of sclerite morphology as key taxonomic characters (Bayer, 1981), but the underlying mineralization mechanisms in these corals remain largely unknown.
In a recent study (Tentori et al., 2004) on growth and calcification of the tropical soft coral Litophyton arboreum during 49 days of tissue recovery from colony fragmentation, it was found that (1) tissue injury can affect the results of the investigation, and (2) under laboratory-controlled conditions, L. arboreum has a calcification rate comparable to those of some hermatypic corals. However, comparison across species should be taken only as a guide, particularly in cases in which calcification rates were obtained in experiments performed immediately after cutting and relocating the coral fragments from the reef (Goreau, 1959; Barnes, 1985; Dennison and Barnes, 1988).
There are fundamental differences between calcification of hard (scleractinian) and soft (alcyonacean) corals. Hard corals deposit calcium carbonate as aragonite crystals underneath their tissues, forming the well-known complex geological structures; in contrast, soft corals deposit calcium carbonate as high-magnesium calcite in the form of individual microscopic sclerites within the tissues of the colony (Konishi, 1981). Nevertheless, given their close systematic relationship, geographic distribution, and common association with zooxanthellae, we expected to find similar mechanisms of calcification between tropical hard and soft corals. In particular, it was of interest to determine whether soft corals exhibit light-enhanced calcification and to know if this calcification is affected by tissue injury associated with the removal of fragments from the parent colony. Our study suggests that a diurnal pattern of calcification/decalcification in Cladiella sp. may be typical of soft corals.
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Materials and Methods
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The experimental organism
Cladiella sp., in common with many alcyoniids, grows as thick carpet-like colonies that can extend to several square meters on the coral reef substratum (E.T., pers. obs.). The top surface of these colonies has lobes and fingerlike projections that are about 7 cm long and composed of hundreds of polyps that open to the external medium through tiny mouths 1–2 mm diameter (Fig. 1a). When disturbed, the coral contracts, changing color from dark mauve-brown to gray. The skeleton of Cladiella consists of microscopic sclerites (Fig. 1b) distributed throughout the body, mainly within the mesoglea. The colonies used in this study were collected at 5–8 m depth in the Gulf of Aqaba, Jordan, and had been maintained in the aquarium of the Oceanographic Museum of Monaco for at least 3 years prior to this study. In this artificial environment Cladiella sp. is a relatively fast grower. Several fragments (5 to 6 cm in length) of one parent colony were cut on the same day and placed on plastic mesh trays in the experimental aquarium of the Centre Scientifique de Monaco under controlled conditions (semi-open circulation with a water exchange rate of 2% h–1, 38 psu salinity, 26 ± 1 °C; 220 µmol m–2 s–1; photoperiod of 12h:12h light/dark). The fragments displayed the expansion and color typical of intact colonies within 2 h of fragmentation. Survival was 100%.
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Figure 1. Cladiella sp. (a) Colony reared at the Centre Scientifique Monegasque. Scale bar = 5 cm. (b) Sclerites diversity; smallest sclerites are found in the polyps; larger and well-defined halters (dumbells) are more abundant toward the base and the interior of the colony. Scale bar = 25 µm. Figure 1a photographed by E. Tambutté and used with permission.
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Experimental setup
Nine colony fragments selected at random from the pool were incubated for 4 h (0900 to 1300 hours) under one of three light intensities: high light (HL), low light (LL), and no light (NL) corresponding to 390, 90, and 0 µmol m–2 s–1 (metal halide lamps, Phillips HQI-TS 400 W). Each fragment was incubated at 26 ± 1 °C in 70 ml of filtered seawater (0.45-µm Millipore, FSW). The incubation chamber consisted of a glass tube 3.5 cm in diameter, 8 cm tall, fitted with a plastic mesh floor under which a stirring bar was placed. Each tube was wrapped with the corresponding light filter. The set of incubating chambers was kept on a stirring plate in a water bath. A few minutes before the aquarium lights were switched on, we covered the plastic trays with light filters, allowing the Cladiella fragments to adapt to the experimental light intensity for 1 h before transferring them from the common tank into the individual incubation chambers. During incubation, the water was stirred with the magnet, providing constant and gentle movement of water around the coral (500 rpm); the incubation chamber was loosely covered with a plastic petri dish to avoid loss of water by evaporation. After incubation, the coral fragments were lightly dried on paper towels and their wet weight was measured before they were transferred back to the common tank. The same fragments were used throughout the experiment and exposed to the same light treatment so that individual responses could be followed for the 36 days of the experiment. Calcification rates were measured at 2, 9, 14, 19, 29, and 36 days (recovery times) after the corals were cut. All coral fragments were maintained in the same tank and under the same conditions between incubations.
Calcification measurements
The method employed to measure calcification rates is based on the alkalinity anomaly technique developed by Smith and Kinsey (1978); it uses a ratio of two equivalents of total alkalinity for each mole of CaCO3 precipitation. The ionic strength of the titrating acid (0.01 mol/l HCl) was adjusted by addition of high-grade NaCl (38.4 g l–1, in agreement with Mediterranean seawater). The pH meter was calibrated on the day of use against pH 7.00 (ORION 9109107) and 4.01 (ORION 910104). The pH solutions, incubation samples, and titrating acid were equilibrated to 25°C in a water bath before being used. From each incubation medium, three replicate samples of about 20 ml were analyzed (sample volume was calculated from the sample weighed to the nearest 0.01 g); a final volume ratio of SW/HCl of about 20:8.5 was used for titration. FSW samples were refrigerated for up to 24 h before analysis. Alkalinity determinations were performed with a Mettler DL70 automatic titrator. The total alkalinity (TA) was calculated as mEq l–1, from the slope of the curve HCl vol/pH within the range of pH 3.0 to 4.2, using the Gran equation corrected for sulfate and fluorides (Hansson and Jagner, 1973). The calcification rates were normalized by protein content, skeletal mass, and wet weight.
Measurement of proteins and skeletal mass
To follow changes of calcification rates through time in each one of the coral fragments, it was necessary to keep the specimens intact. Their protein and skeletal contents were measured at the end of the last experiment (day 36 of tissue recovery). The coral samples were covered with 20 ml of 1 mol/l NaOH and heated at 90 °C for 30 min, left to cool down for a few minutes, homogenized, and centrifuged at 12,000 x g, 22 °C, for 20 min. The supernatant was removed for protein analysis using Pierce MicroBCA reactant, measuring at 595 nm using beta gamma globulin standards. The NaOH-insoluble precipitate consisted of sclerites. The total skeletal weight was measured to the nearest 0.001 g from oven-dried samples (60 °C for 24 h).
To determine whether light intensity affected calcification rate, the responses to the three light treatments at each date of tissue recovery were analyzed by regression analysis. On day 29, only HL and LL samples were processed; these data were analysed by student’s t test.
To determine whether calcification rate was affected by time of tissue recovery, the data of each light treatment throughout the study were analyzed by one-way analysis of variance.
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Results
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Calcification rates
Light-enhanced calcification was detected in Cladiella sp. 15 days after major tissue injury, when the coral fragments had recovered (Fig. 2a). The calcification rate increased with time of recovery under the high light (HL) and low light (LL) treatments normalized per protein content (HL P = 0.0116, LL P = 0.0095). The no light (NL) treatment always resulted in decalcification; the decalcification rate decreased significantly with time of recovery (NL P = 0.0178). The variation of the calcification (or decalcification) responses within treatments tended to be smaller as the coral fragments recovered.
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Figure 2. Cladiella sp. Calcification rates through time of tissue recovery. Data normalized by (a) protein content; (b) sclerite dry weight, and (c) colony wet weight (mean ± SE). Values significantly different between light treatments are indicated by *P < 0.05 or **P<0.001.
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With different levels of significance, the same pattern of calcification rate was observed whether accretion of CaCO3 was normalized by protein content or by skeletal weight measured on day 36 of the study, or by coral wet weight measured on the day of each experiment (Fig. 2a–c).
Alkalinity and pH of the incubation medium
The seawater incubation medium had an alkalinity range of 2.549 to 2.594 mEq l–1 and a pH range of 7.979 to 8.256. After 4 h, according to the light intensity, with the coral fragments present, the seawater pH changed. The HL incubation medium had a higher pH than the original incubation medium (FSW). The NL incubation medium had a lower pH than FSW. The LL incubation medium showed the smallest changes of pH and tended to be lower than FSW (Fig. 3).
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Figure 3. Cladiella sp. pH of seawater at start (FSW) and end (HL, LL, NL) of incubation periods (mean ± SE). HL, high light; LL, low light; NL, no light.
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Wet weight, calcium carbonate and protein content
The colony fragments used were of similar dimensions. On completion of the study, their wet weight and protein content seemed well correlated, while the sclerite weight appeared to be independent (Table 1).
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Discussion
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Choice of normalizing parameter
Since the seminal work of Goreau (1959), protein content has been the most common parameter used for normalization of calcification estimates (see Appendix). The underlying assumptions are that protein represents biomass, that all biomass has the same calcifying potential, and that biomass runs parallel to the accretion surface of the skeleton. These assumptions may not apply to soft corals because a high proportion of their protein content is extracellular and because the sclerites are not homogenously distributed in the colony (Tentori et al., 2004).
Estimating calcification rates in terms of surface area has been attempted through a variety of methods (Appendix). Although these methods revealed valuable information, comparisons of such experimental data across research groups and calcifying taxa should be used with caution because the level of accuracy of surface area measurement does not match the level of accuracy of calcium carbonate measurement.
In the interests of comparing the calcification rates of Cladiella sp. to those of other calcifiers, and considering the impossibility of measuring the surface area of soft corals or their sclerites, we used protein as the normalizing parameter, keeping in mind that our data would underestimate the process. The analysis of protein content requires the destruction of the sample. The 36-day duration of the present study was a compromise between a time short enough to avoid significant changes in protein content but long enough to cover the required period of tissue recovery indicated in our previous study (Tentori et al., 2004).
Calcification
The calcification rates of several tropical and temperate calcifying organisms have been measured over the last few decades. The Appendix summarizes the investigations that employed protein, surface area, or both as normalizing parameters; the original data were recalculated for comparative purposes. The calcification rates fluctuate from 6.4 to 3680 and from 0.6 to 19.2 nmol CaCO3 mg prot–1 h–1 for tropical and temperate species, respectively. The wide range of variation is in part due to the diverse experimental settings. Under similar measurement conditions, the calcification rates of the soft corals Cladiella sp. and Litophyton arboreum (23 and 49 nmol CaCO3 mg prot–1 h–1, respectively) are within the range of scleractinian calcification rates. These calcification rates are remarkably high and unexpected, suggesting that high calcification rates in daytime are not an exclusive characteristic of reef-building organisms and supporting the observations reported by Goreau and Goreau (1959).
The calcification rates of Cladiella were measured on colony fragments maintained in the same aquarium conditions, the only difference being the incubation under high, low, or no light initiated an hour prior to the alkalinity measurements. The results indicate that, once recovered from tissue injury, the effect of light intensity on calcification is immediate.
Decalcification
Decrease of calcification rates in the dark is common in scleractinian corals, but decalcification is not (Gattuso et al., 1999). Kawaguti and Sakumoto (1948) noted "intake of Ca2+" in all corals exposed to light and "output of Ca2+" in all corals exposed to dark. They argued that the skeleton formation was favored by the alkaline pH (8.84 to 9.15) of the incubating medium, which was assumed to be result of photosynthesis by zooxanthellae in the coral; correspondingly, the drop in pH (8.00 to 7.80) in the dark was the reason for the "resolution of the skeleton [sic]." We interpret such intake and output as calcification and decalcification, respectively. Chisholm (2000) observed dark decalcification in coralline algae incubated at various depths, and explained this as a result of previous light exposure or the acidification caused by cell respiration. It is not clear how "previous light exposure" could cause decalcification. Our results agree with Chisholm’s latter explanation. It is also possible that the tissue recovery verified visually underwater was overestimated and that decalcification was due to tissue injury.
Although the calcification rates of soft corals in the light are comparable to those of some tropical scleractinian corals, the net calcification rate in a 24-h cycle would be much lower if dark decalcification was taken into account. Our preliminary experiments on Sarcophyton sp. and Sinularia sp. indicated that these corals also decalcify in the dark; they also showed that the level of calcification at a given light intensity is affected by the previous treatment.
Octocoral sclerites consist of calcite and organic matter. Their main role is traditionally seen as physical deterrence to predation (Coll, 1992; Van Alstyne et al., 1994; West, 1997). According to Bengston (2004), marine invertebrate skeletons with a poor organic matter structure are not the strongest. Bengston (2004) also states that the physiological cost of producing the mineral is smaller than that of producing the organic matrix. In this sense, and assuming that predation stress is reduced in the colder months in temperate waters, the loss of sclerite mass and sclerite organic matter during winter in the gorgonian Leptogorgia virgulata (Kingsley et al., 1990) fits the physical deterrence model. However, a daily cycle of sclerite formation as suggested in this study would be a physical adaptation of little value in tropical waters. Our decalcification results must be linked to a different mechanism—one that is possibly related to the 2-h cycles of movement of Ca2+ between seawater, tissues, and sclerites (Velimirov and King, 1979) in the gorgonian Eunicella papillosa.
Tissue recovery
The experimental data presented in the Appendix clearly confirm the light-enhanced calcification concept. Our study shows an additional aspect of calcification studies not considered before: the light-enhanced calcification is unmistakable only when the experimental coral fragments have had time to heal (Fig. 2a–c). The delayed response on calcification obtained in this study is in agreement with the delayed response on cellular growth observed in the soft coral Litophyton arboreum following tissue injury (Tentori et al., 2004). These findings suggest that other processes take over the metabolic energy reserves of the coral immediately after tissue injury.
Meszaros and Bigger (1999) investigated the mechanism of wound healing in the zooxanthellate gorgonian Plexaura fusifera and found that amoebocytes from the mesoglea accumulated at the wound site by migration rather than by cell division. These amoebocytes later differentiated into epithelial cells. Furthermore, while the process of wound healing was noticeable for 4 to 7 days, it took 2 weeks to complete. Tissue repair and metabolic energy status in scleractinian corals (Kramarsky-Winter and Loya, 2000; Houlbreque et al., 2003) could also explain the increased calcification rates of A. formosa after a one-week recovery period (Barnes, 1985) shown in the Appendix.
pH measurement
We detected a drop of pH and decalcification as the typical response to dark conditions (Fig. 3); however, corals under the high light treatment exhibited increases of pH and decalcification in the first few days of recovery (Fig. 2a–c). If the increase of pH indicates that photosynthesis was not interrupted in the high and low light treatments, the recorded decalcification results suggest that in order for carbonates to be deposited, metabolic energy needs to be available.
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Acknowledgments
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The Centre Scientifique de Monaco made available the research funds and infrastructure to conduct this work. Francesca Marubini helped us with the progress of the methodology employed. The Musée Océanographique de Monaco (Nadia Ounais and Pierre Gilles) kindly supplied the biological material. Eric Tambutté produced the photograph used in Figure 1. E. Tentori was supported by a study leave from Central Queensland University, Australia, and the hospitality of the CSM.
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Footnotes
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Received 10 August 2005; accepted 6 July 2006.
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Abstract
Introduction
