Introduction

Coral reefs owe most of their high diversity and productivity to energetic relationships occurring in Scleractinian corals1. Coral symbiotic nature is characterized by a complex metabolism integration between the cnidarian host and its photosynthetic endosymbiont (Symbiodiniaceae, dinoflagellate)2,3. The coral physiology is thus determined by multiple metabolic traits4 from which respiration and photosynthesis are the utmost driving processes5. Coral bioenergetics has been investigated through gas exchange analysis thanks to the development of different systems which allow measurements in situ (e.g.6,7) or in laboratory (e.g.8,9). Coral respiration is largely fuelled by photosynthetically-derived carbon molecules10, by active heterotrophy11, and by oxygen released during photosynthesis12. It is carried out in both animal and algal mitochondria by the respiratory complexes (Complex I, II, III, IV and F1FO ATP synthase) which fulfils most of the cellular ATP demands. Host mitochondrial electron transport activity is directly related to respiratory substrates availability13. However, it has been suggested that the way in which corals consume storage molecules, as lipids, may be related to their sensitivity to bleaching conditions14. In the model sea anemone Exaiptasia pallida, thermal stress causes a decrease in gene expression of cytochrome c and ATP synthase subunit a, and ultimately to mitochondria degradation and apoptosis15. Cnidarian cellular physiology seems to be intimately linked to mitochondrial redox state16. Beyond this, the role of non-proton pumping (and consequently non-ATP yielding) alternative respiratory enzymes, such as alternative NADH dehydrogenases and alternative ubiquinol oxidase (AOX) which genes are present in the cnidarian genome17,18, is still unknown. Differently from the host, Symbiodiniaceae mitochondria lack the canonical NADH:ubiquinone oxidoreductase (Complex I) but possess an AOX19,20. Understanding the contribution of all these components to symbiont and host respiration will help to characterize the physiological responses of corals to natural oxygen variations in the reefs and its interaction with stress21.

Coral photosynthesis is carried out by Symbiodiniaceae algae only. Light absorption occurs in the thylakoid antenna system, which in peridinin plastid-containing algae as Symbiodiniaceae consists of a membrane embedded chlorophyll a-chlorophyll c2-peridinin protein complex (acpPC) and a water soluble peridinin-chlorophyll a-protein complex (PCP)22. These antennas transfer energy to photosystem II (PSII) and photosystem I (PSI) reaction centers, allowing charge separation, and a linear electron flow (LEF) from oxidation of water molecules at the PSII donor side to reduction of NADP+ at the PSI acceptor side. The intermediate Cytochrome b6f complex oxidises the plastoquinol pool (electron acceptor of PSII) and reduces cytochrome c6 (cyt c6; electron donor of PSI), meanwhile translocating protons to the lumenal side of the thylakoid. This transmembrane electrochemical proton gradient is then used by the CF1FO ATP synthase complex. The availability of NADPH and ATP for CO2 fixation depends largely on a complex balance between the different processes producing and consuming the electrochemical proton gradient, ATP and NADPH. An alternative cyclic electron flow (CEF), involving PSI and Cytochrome b6f, may work to balance the energetic requirements for CO2 fixation. CEF activity, by contributing to increase the electrochemical proton gradient, accomplishes two main functions that are driving the synthesis of extra ATP and inducing a non-photochemical quenching (NPQ) dissipation mechanism that may prevent PSII photoinhibition23. CEF has been poorly studied in Symbiodiniaceae and its role under steady physiological conditions, both in culture and in hospite, is not clear.

Carbon fixation is carried out by a type II Rubisco which is characterized by a low CO2-O2 selectivity factor, and requires the presence of an efficient CO2 concentrating mechanism24,25 largely controlled by the host26,27 that limits the occurrence of the photorespiratory pathway24. After accomplishing carbohydrate synthesis, an important amount of photosynthetic products is transported to the host28 thanks to the expression of membrane transporters29. The photosynthetic activity of corals is dynamic at different levels of time and space scales in coral reefs30, and it has been shown to be highly coupled with nutrients31 and water flow7. The delicate balance of symbiosis in corals can be disrupted as a consequence of affections in the photosynthetic activity. For instance, a decrease in the electron transfer at the PSII acceptor side in heat sensitive species, leads to damage in thylakoid membranes and potentially to bleaching32. Altogether, this points out the importance of studying Symbiodiniaceae photosynthesis in hospite.

At the cellular level, different regulatory mechanisms of photosynthetic electron flow help to balance the light energy absorbed and the energetic requirements for carbon fixation in microalgae33. The characterization of these mechanisms has been achieved through in vivo PSII-related chlorophyll a fluorescence34. This technique has shown that different Symbiodiniaceae isolates show diverse photobiological responses to light changes35. For instance, the presence of CEF has been suggested to occur in Symbiodinium genus (formerly Clade A) both free-living and in symbiosis36. The combination of chlorophyll a fluorescence and oxygen exchange measurements also showed that Symbiodinium sp. in culture activates a light-enhanced dark respiration during photosynthesis8. A more detailed characterization through the combination of various techniques (O2 exchange, PSII chlorophyll a fluorescence emission, PSI primary electron donor [P700] absorbance change in the far-red spectrum, and analysis of the thylakoid transmembrane electric field), revealed that an oxygen-dependent reduction at the acceptor side of PSI occurs as photoprotective mechanism in Symbiodinium isolates37,38.

Spectrophotometric measurements of PSI activity performed in the anemone model species Exaiptasia pallida harboring Breviolum sp. (formerly Clade B) revealed that exposure to high temperature can stimulate the CEF capacity23. However, on corals such studies are still scarce because: (1) their morphology is not suitable for most spectroscopic apparatus that accommodate 1 cm2 cuvette; (2) the thickness and nature of their calcareous skeleton makes coral branches or nubbins not homogenous spectroscopic objects, scattering and poorly transmitting detecting light; (3) actinic light cannot be delivered in a homogenous manner on all sides of the coral specimens even in simple systems, like in oximeter chambers.

The simultaneous monitoring of in vivo PSI and PSII activity in corals has been attempted by using the pulse-amplitude-modulation technique (Dual-PAM, Walz, Germany) which records variable chlorophyll a fluorescence and P700 photo-oxidation. Hoogenboom et al.39 analysed the laminar coral Turbinaria reniformis and concluded that PSI was not a major site for damage during thermal stress. They also suggested the existence of CEF at high light intensities because of a decoupling between PSII and PSI activity was observed, but no clear results were obtained from the branching coral S. pistillata. In contrast, a recent study using improved settings of the Dual-PAM in reflectance mode, in combination with chlorophyll a fluorescence kinetics, showed a strong sensitivity of PSI to a Calvin cycle inhibitor and thermal stress in the plate-like coral Pavona decussata40. This highlights the lack of knowledge on PSI activity in corals in response to environmental changes.

Coral skeleton is composed by aragonite depositions which has a low light absorption capacity and diffuses and scatters light through its structure41. Other works have used small coral fragments to better characterize biophysical properties of the coral skeleton41 or coral pigments through spectroscopic kinetics42. Consequently, coral fragments might be good spectroscopic objects but, despite this, they have not been studied by advanced biophysical and spectroscopic methods commonly applied for other photosynthetic organisms to study photosynthesis (and more generally bioenergetics). In this work we challenged thin and small coral fragments to classical physiological and highly specialized spectroscopic measurements of respiration and photosynthesis, and we aim to propose them as reliable sources of in vivo information.

Methods

Biological material

Coral colonies of Stylophora pistillata variety Milka were obtained from DeJong Marinelife (Netherlands), and maintained in a 300 L capacity aquarium filled with artificial seawater prepared at 34 PSU (Coral Pro Salt, Red Sea Fish LTD). Temperature was set to 26 °C and light was provided by a LED spot light (A360W tuna Blue, Kessil, USA) at 100 μmol photons m−2 s−1 in a light–dark cycle of 12 h:12 h. Corals were fed two to three times per week with freshly hatched artemia. Some measurements reported here were also conducted on coral colonies of Pachyseris speciosa, Pocillopora damicornis, Acropora formosa and Pavona cactus. These colonies were collected by SCUBA diving in the area of Malakal in the Palau Archipelago (7°19′28.3″ N 134°28′01.6″ E), during a sampling campaign conducted in January 2018. After collection, colonies were transferred to the facilities of the Palau International Coral Reef Center (PICRC) and suspended from a nylon wire inside a tank supplied with running filtered natural seawater. Temperature in the tank was 30 ± 1 °C and the maximum Photosynthetic Photon Flux Density (PPFD) at noon was about 100 μmol photons m−2 s−1.

Symbiodinium microadriaticum culture (CCMP 2467) originally isolated from shallow water coral S. pistillata from the Red Sea, was kindly provided by Prof. Oren Levy (Bar Ilan University, Israel). The culture was maintained in 250 mL Fernbach flasks containing 75 mL of artificial sea water (Coral Pro Salt, Red Sea Fish LTD) at 34 PSU and enriched with Guillard’s (F/2) Marine Water Enrichment Solution (Sigma-Aldrich). The culture was grown under cool white fluorescent lamps at 75 µmol photons m−2 s−1 on a 12 h:12 h light:dark cycle and it was subcultured weekly. For spectroscopic measurements cells were adjusted to 10 µg of total chlorophyll per mL and supplemented with 10% (w/v) of Ficoll to avoid sedimentation.

Coral fragment preparation

Square fragments of about 5 by 5 mm (Fig. 1a) were cut from various coral colonies with a Dremel 8220 (Dremel, USA) equipped with a diamond disk. For the branching corals S. pistillata and A. formosa, two transversal cuts were done close to the end of similar terminal branches, one to remove the growing white tip, and the other which determine the length of the fragment (i.e. approximately 5 mm). A third longitudinal cut determined the thickness of the fragment. The longitudinal cut in the shrubby coral P. damicornis was made in the skeleton regions thicker than 3 mm. Laminar corals P. speciosa and P. cactus were cut perpendicularly to the plane of the skeleton. This procedure was performed out of the aquarium and took less than 30 s per fragment, after which it was rapidly immersed in another sea water container to remove debris. Coral fragments were analysed within one hour after being cut (termed ‘day 0’) or thereafter maintained in the same aquarium conditions as their parent colony.

Figure 1
figure 1

Experimental setup and representative experiments on small and thin coral fragments. (a) Picture of recently obtained S. pistillata fragments settled back in the aquarium; the black bar indicates 5 mm. (b) Schematic representation of the setup used for simultaneous analyses of chlorophyll a fluorescence yield and oxygen exchange. (c) Disposition and orientation of a coral fragment into a 1 cm2 glass cuvette relative to the light guide and the detecting photodiode in the JTS-10 spectrophotometer. Drawing in (b) and (c) were created using PowerPoint of Microsoft Office 365 ProPlus. (d,e) Representative records of chlorophyll a fluorescence yield measurements and oxygen concentration changes. Each light step (represented by the upper bar with a grey scale) lasted three minutes and saturating pulses were given each 30 s (r.u. = relative units). (f) Representative record of Photosystem I primary donor (P700) photo-oxidation upon five seconds of illumination (210 µmol photons m−2 s−1) followed by a saturating pulse of light (SP), in presence of PSII inhibitor (DCMU at 20 µM).

Chlorophyll a fluorescence measurement

In vivo chlorophyll a fluorescence images were obtained by a SpeedZen imaging system (BeamBio/API, France). An array of blue LEDs (450–470 nm) was used to excite chlorophyll a during 400 μs, and images were obtained by a CCD camera (UI-3240CP-NIR-GL Rev.2, IDS, Obersulm, Germany) filtered by a long pass red filter (680 nm) (Fig. 1b). After incubating samples in darkness for at least 15 min, minimal chlorophyll a fluorescence yield (Fo) image was acquired. A saturating red actinic light pulse (3000 μmol photons m−2 s−1), provided by a LED array (660 nm), was applied for 200 ms and maximal chlorophyll a fluorescence yield (Fm) was acquired. Maximum quantum yield of photosystem II was calculated as Fv/Fm = (Fm − Fo)/Fm43.

Coral fragments were exposed to different PPFD (15, 68, 172, 354 and 755 μmol photons m−2 s−1, provided by the same red LED array than the saturating pulse) during 3 minutes and fluorescence (Fs) was recorded every 5 s (Fig. 1d). A saturating red actinic light pulse was applied to obtain the maximum fluorescence yield under light acclimation (Fm′) every 30 s, and the effective quantum yield of PSII at steady state was calculated as ϕPSII = (Fm′ − Fs)/Fm′. The relative electron transfer rate through PSII (rETR-PSII) was then calculated as ϕPSII x PPFD, and the non-photochemical quenching of chlorophyll a fluorescence (NPQ) as (Fm − Fm′)/Fm′43.

Fast induction curves of chlorophyll a fluorescence were recorded with a Handy PEA fluorometer (Hansatech Instruments, UK), by illuminating the sample during 10 s with a strong red light pulse (peak wavelength of 650 nm) of 2000 μmol photons m−2 s−1. This fluorescence rise is characterized by a sudden increase from a basal fluorescence level (O), with two intermediate inflections (J and I), to a maximum fluorescence level (P) commonly referred as OJIP transient (see Chapter 12 in34). O phase is taken at 50 μs, J appears at 2 ms, while I appears after 50 ms and P after 2 s in corals. Each coral fragment was set in the bottom of a standard spectrophotometer cuvette facing the head sensor and filled with 2 mL of artificial seawater and maintained in darkness for 15 min before the measurement.

Light-induced absorption changes

All measurements were performed with a JTS-10 spectrophotometer (BioLogic, France) piloted by a BeamBio/API (France) electronic device. When necessary, the final size of the coral fragments was adjusted to fit into standard 45 mm × 12.5 mm × 12.5 mm (H × W × D) glass spectrophotometer cuvette, i.e. with a pathlength of 10 mm. Each coral fragment was held in the cuvette with the side containing most polyps facing the light source and the skeleton exposed area (when applicable) in front of the detecting photodiode (Fig. 1c). This guaranteed a controlled illumination of the sample.

In the photosynthetic electron transport chain of peridinin chloroplast-harboring dinoflagellates cyt c6 is the electron donor to PSI instead of Plastocyanin (PC)44. For this reason, light-induced absorption changes of P700 were directly measured at 705 nm45 by applying μs detecting flashes during short intervals of dark pulses (as described in46). Continuous actinic light at different intensities (25, 85, 210, 380, 670 and 1400 μmol photons m−2 s−1) was provided during 5 s (3 min for field coral specimens) by a LED array (640 nm). When coral fragments were exposed to continuous light, a stable negative change in absorbance at 705 nm (due to the formation of oxidized P700)47 was obtained and recorded as Ps (Fig. 1f). A strong saturating pulse of light (9000 μmol photons m−2 s−1) was applied to obtain a more negative absorbance signal corresponding to maximum photo-oxidation of P700 as Pm’, while after a dark period of 1 s absorbance was recorded as Po. Maximum absorbance change (Pm) was estimated by adding the potent photosystem II inhibitor DCMU (3-(3,4-dichlorophenyl)-1,1-dimethylurea) at 20 μM (Fig. 1f). Photosystem I quantum yield was calculated as Y(I) = (Pm′ − Ps)/(Pm − Po), quantum yield of non-photochemical energy dissipation due to donor side limitation as Y(ND) = (Ps − Po)/(Pm − Po), and the quantum yield of non-photochemical energy dissipation due to acceptor side limitation as Y(NA) = (Pm − Pm′)/(Pm − Po)48. The relative electron transfer rate through PSI (rETR-PSI) was calculated as the product of Y(I) and PPFD.

Absorbance changes of some pigments in the green region of the visible spectrum occur in response to light-induced changes of the trans-thylakoid electric field. This so-called electrochromic shift (ECS) is a useful way to determine photosynthetic parameters such as stoichiometry of active PSI and PSII in photosynthetic eukaryotes49, including Symbiodiniaceae37. ECS spectra were determined by measuring light-absorption changes in response to 2 ms continuous illumination, each 10 nm using light band-pass filters from 480 to 600 nm, as previously described50.

The ratio between active PSII and PSI centers can be quantified because the ECS signal responds linearly to the intensity of the electric field37. ECS signal at 554 nm was followed by applying a saturating single-turnover flash of 5 ns provided by a Nd:YAG Laser (Minilite II, Continuum) and its amplitude was considered as PSI + PSII reaction centers contribution. PSII inhibitors DCMU (20 µM, from a stock solution dissolved in ethanol) and hydroxylamine (1 mM, from a stock solution dissolved in distilled water) were added simultaneously and incubated in darkness during 10 min, another single-turnover flash was given and the resulting ECS amplitude accounted for PSI contribution. Ten consecutive repetitions, separated by 5 s in darkness, were averaged per measurement.

Oxygen exchange

Oxygen consumption (respiration) or evolution (photosynthesis) were measured with an Oxygraph + System (Hansatech, UK). Oxygen concentration calibration was carried out at two points with 0.2 µm-filtered F/2 medium either saturated in oxygen after vigorous shaking, or depleted in oxygen by addition of sodium dithionite (Sigma-Aldrich).

Coral fragments were held suspended into a DW1 oximeter chamber (Hansatech, UK) by an inert piece of acetate sheet. This enabled the adequate stirring of the medium by the magnetic stir bar (Fig. 1b). Each fragment was dark acclimated for at least 15 min with the chamber opened before closing it to monitor oxygen consumption in the dark for 5 min. After this dark period, illumination was provided by the Chlorophyll a fluorescence imaging system, so during fluorescence recording oxygen concentration was simultaneously acquired (Fig. 1e).

Chlorophyll content and coral surface determination

Pigments were extracted by incubating coral fragments in cold methanol during at least 12 h in darkness at 4 °C. For S. microadriaticum, cells were resuspended in cold methanol and vortexed in presence of 500 µL of acid-washed and autoclaved glass beads (710–1180 µm; Sigma-Aldrich) for 2 min at 30 Hz with a TissueLyser II (QIAGEN). Pigment extracts were centrifuged at 16,000 g for 10 min to remove debris, and chlorophyll a and c2 concentrations were calculated according to51 from absorption measurements at 632 and 665 nm. Tissue-containing surface of coral fragments was calculated from conventional real color pictures of each fragment by using the software ImageJ (version 1.51j8).

Symbiodiniaceae genotyping

Small fragments of each species (n = 3) were removed from independent colonies and immediately processed or preserved in RNAlater (Invitrogen). Fresh or preserved fragments were washed with filtered artificial sea water and coral tissues were extracted by airbrushing the fragments into sterile PBS. DNA from the microalgal fraction was then extracted by using a DNeasy Plant mini kit (QIAGEN) according to the manufacturer’s instructions, with slight modifications. The PCR amplification of the ITS2 gene marker was performed using primers pair SYM_VAR_5.8S2 and SYM_VAR_REV as detailed in52. The amplicons library was constructed using the Nextera XT kit (Illumina, Inc., San Diego, CA, USA) and sequencing was carried out on the Illumina MiSeq platform with 2 × 250 bp read configuration. The ITS2-type profiles were obtained by using the SymPortal analytical framework53.

Results

Whole S. pistillata coral colonies containing S. microadriaticum exhibited a homogeneous pigmentation (as illustrated in Fig. 2a). Chlorophyll a fluorescence imaging indicated a maximum quantum yield of photosystem II (Fv/Fm) of 0.53 ± 0.03 (Fig. 2b,d). Small half-cylinder shape fragments were cut from these colonies. They were covered with about 35 mm2 of coral tissue with an average total chlorophyll content (Chl a + c2) of 30.5 ± 7.6 µg cm−2, a Chl a/c2 ratio of 2.39 ± 0.11 and 61 ± 11 polyps cm−2 (n = 16). These values remained stable up to 2 days after cutting and no visual sign of tissue necrosis was observed during this period (Fig. 2c). Although there was a positive correlation (R2 = 0.41, p = 0.007) between Fv/Fm and fragment thickness (from 1.5 to 5 mm), the slope of the linear regression is very small so that the difference between a 2 mm fragment and a 5 mm fragment is less than 6% (Fig. 2e). In addition, the average of Fv/Fm values of coral fragments is not significantly different to the Fv/Fm value of coral colonies (one-way ANOVA, F(3,20) = 2.14, P = 0.13) (Fig. 2d). The fast polyphasic chlorophyll a fluorescence rise (i.e. OJIP curves) was obtained from freshly cut coral fragments and from 1- or 2-days old fragments (Fig. 2f) and no changes in the relative amplitude or presence of phases were observed.

Figure 2
figure 2

Maximum photosynthetic quantum yield of photosystem II (Fv/Fm) is not altered in thin fragments of the coral S. pistillata. (a) Picture of one S. pistillata colony maintained in aquarium conditions. (b) False-color picture of Fv/Fm obtained by chlorophyll a fluorescence imaging of Fo and Fm. (c) Representative coral thin fragment monitored during two days and its related Fv/Fm false-color image. (d) Fv/Fm measured on the apical region of coral colonies where coral fragments were sampled and coral fragments monitored two days after sampling. (e) Fv/Fm of coral fragments of variable thickness after one day of sampling. (f) Polyphasic chlorophyll a fluorescence rise from dark adapted coral fragments was normalized to 0 at 50 µs (‘O’ phase) and to 1 at the plateau (‘P’ phase) (r.u. = relative units). The two typical inflections, phase J and I, are indicated (n = 6 for (d) and (f)).

To further test the good health of coral fragments, we determined respiratory and photosynthetic oxygen exchange rates from 3 mm thick fragments. Dark oxygen consumption (Rd) was on average 13.9 ± 2.1 nmol O2 cm−2 min−1 over the three days and remained stable (Fig. 3a, inset). Net oxygen evolution as a function of the light intensity (Fig. 3a) was obtained simultaneously with fluorescence-based calculation of relative electron transport rate of PSII (rETR-PSII) (Fig. 3b). Both parameters showed a similar light dependency as indicated by the fact that rETR-PSII linearly correlated to oxygen evolution rates (R2 = 0.98, p = 0.002) (Fig. 3c). Moreover, non-photochemical quenching (NPQ) capacity also remained stable (Fig. 3d).

Figure 3
figure 3

In vivo measurement of respiration and photosynthesis in S. pistillata coral fragments. (a) inset : Oxygen consumption in the dark (Rd, nmol O2 cm−2 min−1). (a,b) Light-dependent oxygen exchange and relative electron transport rate (rETR-PSII) were obtained simultaneously. (c) Relationship between rETR-PSII and oxygen evolution. (d) Non photochemical quenching of fluorescence (NPQ) is shown for three minutes light-acclimated coral fragments. Vertical bars in (ad) represent standard deviation, n = 6.

Altogether, these data indicated that the global photosynthetic and respiratory activities of the coral fragments remained stable after cutting. In addition, most of the coral fragments showed the same potential to regenerate the tissue and the coral skeleton, as already shown for fragments of similar size54.

We then tested the effect of two classical inhibitors on respiration: potassium cyanide (KCN), a potent inhibitor of mitochondrial cytochrome c oxidase complex (Complex IV), and salicylhydroxamic acid (SHAM), an inhibitor of mitochondrial alternative oxidase (AOX). Addition of 1 or 2 mM KCN reduced Rd to 50% after five minutes (Fig. 4a). Addition of 2 mM SHAM further decreased the remaining cyanide-insensitive oxygen consumption to 26% of total Rd (Fig. 4b), and higher concentrations of SHAM (4 or 6 mM) did not suppress this residual oxygen consumption (data not shown). In contrast, addition of SHAM up to 2 mM alone did not have an impact on Rd during the 20 min of exposure to the inhibitor (Fig. 4a).

Figure 4
figure 4

Impact of SHAM and KCN on dark respiration of S. pistillata coral fragments. (a) Respiratory rates of dark-adapted coral fragments were monitored up to 20 min in dark conditions, n = 3. Cyanide (KCN dissolved in deionized H2O) at 1 or 2 mM, n = 4. SHAM (dissolved in DMSO) at 0.5 or 2 mM, n = 4. DMSO at 2%, n = 4. DMSO at 2% plus KCN at 1 mM, n = 6. (b) Mitochondrial dark respiration rate (Rd) and the effect of SHAM 2 mM, KCN 1 mM or SHAM 2 mM + KCN 1 mM (KCN + SHAM). Respiratory rates were measured 5 min after the addition of inhibitors, n = 10.

We then tested the potential of coral fragments to monitor light-induced absorption changes of PSI primary electron donor (P700) in the far-red region of the visible spectrum (705 nm) (Fig. 1f). These absorption changes corresponding to P700 photo-oxidation are fully consistent with previous observations in plant leaves or algae48 and allowed us to calculate PSI quantum yield [Y(I)] and both PSI donor and acceptor side limitations [Y(ND) and Y(NA), respectively] at different light intensities (Fig. 5a). In addition, relative electron transport rates of PSI (rETR-PSI) were calculated from Y(I) values and a typical photosynthesis-light curve was obtained (Fig. 5b). We also observed a positive correlation (R2 = 0.40, p = 0.01) between absorbance changes upon a saturating pulse of light in presence of DCMU (Pm values) and chlorophyll a content per coral fragment (Fig. 5c). In contrast, thickness of coral fragments did not correlate (R2 = 0.06, p = 0.36) to the maximum amplitude of P700 absorbance change (Pm) (Fig. 5d). This indicates that the coral skeleton did not interfere significantly with our measurements.

Figure 5
figure 5

Light-dependent P700 oxidation measurement and PSI activity in S. pistillata coral fragments. (a) PSI quantum yield (Y(I), acceptor and donor side limitations (Y(NA) and Y(ND), respectively) as a function of PPFD). (b) Relative electron transport rate through PSI (rETR-PSI) calculated as the product of Y(I) and PPFD. (c,d) Maximum P700 absorbance change in the presence of 20 µM DCMU at high light (Pm) in function of its chlorophyll a content (c) or in function of its skeleton thickness (d). A correlation factor (R2) obtained from a linear trendline is shown for (c) and (d). n = 13 for (a) and (b).

We next quantified the fraction of active PSI and PSII centers in coral fragments by monitoring the electrochromic shift signal (ECS), a technique previously used in various algal species, including Symbiodiniaceae37,49. The overall shape of the ECS spectrum of S. pistillata was similar to the one of its isolated symbiotic algae (S. microadriaticum), with a maximum and a minimum observed at 510 nm and around 560 nm, respectively (Fig. 6a). The ratio between maximum (510 nm) and minimum (560 nm) values was smaller in coral fragments than in isolated algae. We then assessed the amplitude of the fast ECS (< 1 ms) at 554 nm in response to a saturating single turnover flash (Fig. 6b). Under these conditions, the amplitude of the ECS is proportional to the amount of active PSI + PSII (reviewed in49). Addition of Hydroxylamine (HA) and DCMU, two PSII inhibitors, halved the amplitude of the ECS both in coral fragments of S. pistillata and isolated S. microadriaticum (Fig. 6b), indicating that PSI to PSII ratio is about 1 in both cases.

Figure 6
figure 6

Electrochromic shift (ECS) signal from different Symbiodiniaceae sources. (a) ECS signal is compared between isolated and in hospite S. microadriaticum cells normalized to ECS at 510 nm (r.u. = relative units normalized to ECS signal at 510 nm). (b) PSI and PSII contribution to ECS signal at 554 nm was assessed by addition of PSII inhibitors hydroxylamine (HA, 1 mM) and DCMU (20 µM). Note the different scale magnitude for ECS signal between S. microadriaticum (left axis) and S. pistillata (right axis) acquisition. Time scale shows different units for negative and positive values. (c) ECS signal obtained from four different scleractinian corals species freshly sampled in the field (r.u. = relative units normalized to ECS signal at 510 nm). (d) True colour pictures of the coral fragments used to obtain ECS spectrum and a representative picture of the corals collected. The black bar stands for 5 mm. n = 3 for (a) and (b).

The four different coral species collected from Palau harbored the same Symbiodiniaceae genus (Durusdinium sp), which is a dominant algal genotype of inshore corals in this area55. For them, a similar ECS spectrum was obtained (Fig. 6c, d). It was not possible to test PSI:PSII stoichiometry in these coral species because a saturating flash laser was not available at that time of analysis.

Spectroscopic-based photosynthesis activity measurements allowed us to compare different parameters obtained from small coral fragments. In S. pistillata, rETR-PSII linearly correlated (R2 = 0.999, p = 0.0005) to rETR-PSI up to the minimum saturating light intensity (around 200 µmol photons m−2 s−1), after which PSI activity continued increasing compared to PSII (Fig. 7a). Corals from the field revealed different responses when PSI and PSII activities were compared (Fig. 7b). A linear response was obtained from the two branching corals A. formosa (R2 = 0.95, p = 0.004) and P. damicornis (R2 = 0.96, p = 0.003) along with the different light intensities tested, but the laminar corals P. cactus and P. speciosa exhibited a disrupted relationship when light intensity was higher than 200 µmol photons m−2 s−1.

Figure 7
figure 7

Relationship between rETR-PSI and rETR-PSII in S. pistillata (a) and in four reef-building corals species freshly collected in the field (b). Each light step was 3 min. n = 6 for (a), n = 5 for (b).

Discussion

Coral survival to fragmentation is a well-known and broadly documented phenomenon (e.g.56). In the case of S. pistillata, a robust scleractinian coral species, small nubbins survive after fragmentation as well54 and have been used in skeleton calcification observations57. In this work, we demonstrate that thin coral fragments (around 25 mm2) can be used to study various bioenergetics traits which so far have not been assessed in reef-building coral species.

The chlorophyll a fluorescence-based photosynthetic parameter Fv/Fm, which accounts for the maximum quantum yield of PSII, has been used to track damages on the photosynthetic capacity and health in photosynthetic organisms, including corals (e.g.58). On the other hand, OJIP chlorophyll fluorescence rise has been used to reveal alterations of the PSII reaction center in Symbiodiniaceae cultures59 and changes in the redox state of PSII acceptor side in corals60. The fact that we did not observe any significant modification neither of the Fv/Fm value nor in OJIP chlorophyll a fluorescence kinetics up to 48 h after fragmentation in small coral fragments (Fig. 2), indicates the absence of structural or functional disturbance in the photosynthetic electron transfer chain. Moreover, our coral fragments exhibited a photoprotection capacity (NPQ; Fig. 3d) comparable to values registered from branches of S. pistillata collected from shallow waters in the Red Sea61 and from colonies maintained in aquarium conditions for several years31. Respiration measured in our coral fragments (Rd = 13.9 ± 2.1 nmol O2 cm−2 min−1; Fig. 3a) agreed with previous studies on 2 cm long nubbins or 10 to 15 cm long branches from S. pistillata collected in the Red Sea (11 nmol ± 4 O2 cm−2 min−1 in average)12,62. Maximum gross photosynthetic rate (Pgmax = 33 ± 4.7 nmol O2 cm−2 min−1; Fig. 3a) was also similar to values measured in coral branches of this species either maintained in aquarium conditions or freshly collected from the sea (20–35 nmol O2 cm−2 min−1)31,62. Rd and Pgmax values can be used to roughly calculate an extrapolated daily photosynthesis to respiration ratio (Pgmax/Rd*2)63. In our thin coral fragments of S. pistillata this ratio is 1.23 ± 0.16, a value in good agreement with S. pistillata branches (10 to 15 cm long) collected in summer from the Gulf of Aqaba (1.4)56, or from nubbins (2 to 3 cm long) prepared from S. pistillata coral colonies grown in aquarium conditions (1.1 to 1.5)63. Similarities between our results and data from the literature show that values measured on small thin fragments give reliable information on energetic relationships as those measured from bigger pieces of corals.

Alternative electron transport pathways in mitochondria and chloroplasts are described as regulators of the redox poise in the main electron transfer chains, and more generally are known for their influence on the energetic state of the cell64. By measuring oxygen consumption rates in S. pistillata in presence of inhibitors (Fig. 4), we evidenced, for the first time, the presence of a cyanide-insensitive but SHAM-sensitive mitochondrial alternative oxidase (AOX) activity in coral. In several algal species, AOX contributes to dissipate excess of redox equivalents produced in the chloroplast during photosynthetic activity (i.e. NAD(P)H) (e.g.66). The capacity of the AOX pathway measured in S. pistillata was 24% of Rd and is in the range as values previously reported for different species of Symbiodiniaceae grown in culture20,65. The observed lack of effect of the AOX inhibitor (i.e. SHAM) addition to fully respiring coral fragments demonstrates a high capacity of mitochondrial respiration through Complex IV. However, further studies are needed to determine the extent of both the cnidarian and photosymbiont’s AOX contribution to whole holobiont respiration and to elucidate the role of AOX under control and stress conditions.

Besides, in several species of Symbiodiniaceae in culture, a high light-dependent oxygen uptake was found to occur at the acceptor side of PSI through a Mehler-type reaction37,38. As illustrated in Fig. 3, the linear relationship between rETR-PSII and oxygen evolution observed in S. pistillata suggests that the photosynthetic electron transfer rate in high light does not depend on an alternative electron pathway to oxygen reduction as it does in free living cultures (e.g.37,67). This result may be explained by a lower CO2 limitation for symbionts in hospite compared to those isolated in culture26,37, but does not exclude that a different strategy operates under high light intensities. In this respect, PSI-dependent CEF is another alternative photosynthetic pathway that has been suggested to play an important role in the energetic balance in Symbiodiniaceae23,36. The most recommended way to assess CEF in vivo is to compare PSI and PSII activities68 (Fig. 7). Despite the fact that calculation of absolute electron transport was not assessed69 in the present study, our results indicate the occurrence of CEF in high light in S. pistillata and in two laminar corals from the field (P. cactus and P. speciosa). In contrast, we found that light influenced at the same proportion the relative electron transport rate of both photosystems in A. formosa and P. damicornis, two branching coral species, indicating that CEF is neglectable in high light (Fig. 7). It is worth to notice that these last four corals species harbour the same Durusdinium genus. A previous study, based on chlorophyll a fluorescence Serial Irradiation Pulses (SIP) method, indicated the absence of CEF in isolates of Durusdinium genus (referred as Clade D)36. Altogether, this highlights that involvement of photosynthetic alternative electron flows may strongly differ between corals harboring Symbiodiniaceae from the same genus.

Contrastingly to PSII, the role of PSI on photoacclimation has just started to be revealed in Symbiodiniaceae dinoflagellates70. PSI has been the target of very few studies on coral photosynthesis due to technical constraints. An indirect determination made on freshly extracted algae from S. pistillata showed a higher PSI content in coral colonies in symbiosis with Symbiodinium sp. (referred as Clade A) than Cladocopium sp. (referred as Clade C)61, suggesting that PSI content can be determinant for coral photosynthesis adaptation to changing light regimes. Other studies, aiming to assess in vivo PSI activity in nubbins from other coral species were performed by following changes in light reflectance of 820 or 830 nm, and correcting the signal at 870 nm with a pulse-amplitude-modulated Dual-PAM instrument (Walz, Germany)39,40. With this method a correction at 870 nm is commonly implemented in PSI activity determination to deconvolute PC contribution in the far-red region48. However, due to the absence of PC in peridinin chloroplasts44 we consider that this correction is not necessary in photosynthetic corals, and light-induced absorption change at 705 nm can be used (Fig. 1f), as it commonly yields a high negative value in plants46 and green algae71. In the case of Symbiodiniaceae isolates, absorbance change at 705 nm has been used with no major differences in kinetics of P700 photo-oxidation when a correction at 740 nm was applied (personal observation37,38). Assuming that the molar attenuation coefficient of P700 at 705 nm of Symbiodiniaceae is in the range of values determined for P700 in green photosynthetic organisms (i.e. 64 mM−1 cm−1 in spinach72 and 105 mM−1 cm−1 in Chlamydomonas reinhardtii47), we estimated the number of P700 per chlorophyll a to be between 1/720 to 1/1180 in S. pistillata. This range of values is in good agreement with values reported for several Symbiodiniaceae isolates35,37. Despite missing a molar absorption coefficient of P700 for peridinin plastids, our results indicate that spectroscopy-based measurement in thin coral fragments is a reliable and accurate strategy to ascertain PSI activity of coral endosymbionts (e.g. Fig. 5).

As thin coral fragments enabled spectroscopy-based measurements, ECS spectrum could be acquired. The presence of an ECS signal allows a direct measure of the electric field across algal thylakoid membranes49. The ECS spectrum measured in S. pistillata showed a small ratio between maximum (510 nm) and minimum (560 nm) ECS values in comparison to Symbiodiniaceae in culture (Fig. 6a37). On the other hand, field coral specimens showed different ECS signal amplitudes at 554 nm compared to 510 nm, despite harbouring the same Symbiodiniaceae genera Durusdinium (Fig. 6c). In general, this difference can be attributed to pigment composition, distribution and orientation along the thylakoid membranes49, but we cannot exclude that other effects such as differences in coral tissue or skeleton optical properties contribute to this difference. ECS-based measurements of PSI to PSII active center ratio in coral fragments gave comparable values to those previously reported in Symbiodiniaceae isolates (Fig. 6b37,38).

In conclusion, in vivo monitoring of photosynthesis and respiration, as well as PSI and PSII activities, of S. pistillata coral fragments resulted in robust and reliable information. We thus consider that coral fragments are good spectroscopic objects to obtain reliable information about coral photosynthesis. This approach does not consume large amounts of coral colonies or nubbins allowing to limit the loss of living samples when destructive measures are taken. This work paves the way towards a better understanding of the role of bioenergetics processes in coral symbiosis establishment and persistence.