Introduction

Alvarez-Filip et al. (2009) showed that the structural complexity of coral reefs in the Caribbean region has tended to decline since the 1970s. The massive corals, which also provide progressively less shelter for other reef organisms, could disappear in 10 to 15 yrs. Other analyses in the Caribbean Basin demonstrated that the reef coral cover has decreased by 80% over the last three decades, and by nearly 40% in the French Antilles in recent years (Gardner et al. 2003; Bouchon et al. 2004, 2008a, b).

A fine examination of the dynamics of coral assemblages revealed a general decrease in the abundance of most species particularly of the species proposed to be the main builders of Caribbean reefs at the end of the last glaciation (i.e., Orbicella annularis, O. faveolata, Acropora palmata). In contrast, one species, Porites astreoides (Lamarck, 1816), which was among the dominant species of the Antillean coral assemblages from the early 1980s on, was seen to be more resistant than most other corals. For example, the coral cover in the Bay of the Grand Cul-de-Sac Marin in Guadeloupe Island (French Antilles) decreased from 42% in 1984 to 21% in 2018. In 1984, the two dominant species were Agaricia agaricites (9.9%) and Porites astreoides (8.7%), but in 2018, A. agaricites had dropped to 0.8%, while P. astreoides remained dominant and accounted for 8% of the coral assemblage. In fact, Porites astreoides has maintained its original abundance and today is the dominant species in most Caribbean reefs (Green et al. 2008; Javier Gonzalez-Barrios and Alvarez-Filip 2018). Porites astreoides was also shown to be resilient to experimental ocean acidification, to increases in temperature (Bove et al. 2019), and to one bleaching event (Levas et al. 2018), but appeared to be sensitive to successive heating events that can cause damage through bleaching (Grottoli et al. 2014; Schoepf et al. 2015).

One of the common features of reef scleractinian corals is that they harbor dinoflagellate algae belonging to the family Symbiodiniaceae (LaJeunesse et al. 2018). However, genera within this family may respond differently to environmental forcing factors (Franklin et al. 1996; Romaine et al. 1997; Robison and Warner 2006; Takahashi et al. 2009; Hennige et al. 2009; Ragni et al. 2010; Gregoire et al. 2017; Diaz-Almeyda et al. 2017). Under thermal and light stress reactive oxygen species (ROS) including superoxide (O2−) and singlet oxygen are produced in Symbiodiniaceae (Lesser 1996, 2006; Wooldridge and Done 2009; McGinty et al. 2012; Krueger et al. 2014) and can damage both the symbiotic Symbiodiniaceae and their hosts, and these two different organisms have developed specific mechanisms to mitigate the negative effects of these products (Krueger et al. 2014; Roberty et al. 2014; Rehman et al. 2016). Some mechanisms related to the transport of electrons in the photosynthetic apparatus have also been proposed (Roberty et al. 2014). Cyclic electron flow around photosystem I (PSI-CEF) and chlororespiration have been suggested to reduce ROS production in some Symbiodiniaceae genera hosted by scleractinian corals (Reynolds et al. 2008). These studies and many others explored whether photosynthesis and photoacclimation mechanisms are used to reduce the negative effects of high light intensity (Suggett et al. 2008; Roberty et al. 2014, 2015). However, oxidative stress has also been shown to occur in the dark, in the absence of photosynthetic activity (Tolleter et al. 2013; Gardner et al. 2017; Zhang et al. 2016; Diaz et al. 2016). Chlororespiration takes place in the dark or when the light intensity is very low. This mechanism is activated when the photosynthetic machinery is inoperative (Bennoun 1994; Nixon 2000) and, as mentioned above (Beardall et al. 2003), is known to reduce the production of damaging superoxide and hydroxyl radicals. The role of chlororespiration and which metabolic pathway is involved are still the subject of debate and many differences have been observed, for example between land plants and eukaryotic algae (Jans et al. 2008; Martin and Sabater 2010; Nawrocki et al. 2015). Several studies of Symbiodiniaceae reported a decrease in the maximum photochemical quantum yield of PSII (Fv/Fm) at night or during prolonged dark periods that can be explained by a reduction in the PQ pool caused by a chlororespiration-like reaction (Jones and Hoegh-Guldberg 2001; Hill and Ralph 2008; Reynolds et al. 2008).

The aim of the present work was to compare the physiological regulation of the electron transport chain of Symbiodiniaceae during the dark phase of Porites astreoides with that in nine other species of scleractinian corals sampled on Caribbean reefs. We evaluated the level of plastoquinone (PQ) reduction in these species and propose that it can be explained by a chlororespiration-like mechanism. We discuss the potential physiological and ecological benefits of such a mechanism.

Materials and methods

Measurements were made on 10 common species of Caribbean scleractinian corals: Acropora palmata, Agaricia agaricites, Orbicella annularis, O. faveolata, Porites astreoides, Porites divaricata, Porites furcata, Pseudodiploria clivosa, Pseudodiploria strigosa and Siderastrea siderea. PAM (Pulse Amplitude Modulated Fluorometer) measurements were performed between April 2016 and January 2019 in the Bay of the Grand Cul-de-Sac Marin (16° 21.651′ N; 61° 34.787′ W) in Guadeloupe Island (French Antilles). The morphometric characteristics of each species are listed in Table 1. Field measurements were performed at midday, and samples were collected between 9:00 a.m. and midday. Most field measurements were made, and samples were collected between a depth of 2 and 5 m (Table 1). Additional PAM measurements were made on Porites astreoides at depths of 10, 15 and 20 m. Sampled colonies were placed in collection nets underwater. Upon surfacing, the corals were transferred to an insulated ice-box containing seawater collected at the sampling site and transported to the marine laboratory in less than 90 min. Each colony was placed in seawater tanks containing 20 L of seawater.

Table 1 Colony form, calice distribution and size for each species, sampling depth and dates (month/year)
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Measurements in the seawater tanks

The water tanks were placed outside in the shade with air bubbling to avoid oxygen depletion, and the water was renewed at 12-h intervals. The temperature was maintained constant at the same temperature as that measured in the field (between 26.5 and 28.5 °C depending on the sampling date). Natural light was monitored using a PAR underwater sensor LI-192 (Li-COR Biosciences, Lincoln, USA) and a Diving PAM-PAR sensor (WalzTM, Effeltrich, Germany).

Light–dark transition measurements

In seawater tanks under natural light with a natural light cycle, PAM measurements were performed independently in triplicate for each species using an underwater Diving PAM (WalzTM, Effeltrich, Germany) (Ralph et al. 1998). Light–dark transitions were measured starting on the sampling day and continued the following day. The coral sample was excited by a weak red measuring light (1 µmol photons m−2 s−1, 650 nm, frequency 0.6 kHz) to record minimum fluorescence (Fo). The optical fiber (diameter 6 mm) of the PAM was positioned exactly 5 mm above the surface of the coral. The maximum fluorescence in the dark (Fm) or in the light (Fm′) measured at 10-min intervals was obtained using a multiple turnover flash method (Kromkamp and Forster 2003). We used a saturating white light pulse (0.6 s, 4000 µmol photons m−2 s−1) produced by a halogen bulb that allowed reduction of the QA, QB and part of plastoquinone (PQ) pools. Blank fluorescence was done on a bleached coral skeleton under water on the sampling sites before measurements in order to remove the potential noise signal due to reflectance and the background fluorescence signal of seawater. The background fluorescence was always very low (lower than 2% of the Fo signal). The maximum quantum yield, Fv/Fm, was calculated according to the following equation (Genty et al. 1989):

$$\frac{{F_{\text{v}} }}{{F_{\text{m}} }} = \frac{{\left( {F_{\text{m}} - F_{\text{o}} } \right)}}{{F_{\text{m}} }}$$
(1)

When measurements were made in the light, the effective quantum efficiency of PSII was computed as follows (Genty et al. 1989):

$$\frac{{F_{{{\text{q}}^{\prime} }} }}{{F_{{{\text{m}}^{\prime} }} }} = \frac{{\left( {F_{{{\text{m}}^{\prime} }} - F^{\prime } } \right)}}{{F_{{{\text{m}}^{\prime} }} }}$$
(2)

An index was calculated to estimate variation of the Fv/Fm. The index corresponds to the percentage of the Fv/Fm variation during the first 2 h of the dark period, where DK0 is the Fv/Fm at the beginning of the dark phase (i.e., the first Fv/Fm measurement after sunset when the PAR was below 1 µmol m−2 s−1), and DK2 h is the Fv/Fm after 2 h of darkness. An interval of 2 h was chosen because it corresponds to the average time needed to observe the lowest Fv/Fm value in Porites astreoides during the dark period.

$$\Delta {\text{YII}}_{{{ \hbox{max} }(2\;{\text{h}})}} = \left\{ {\frac{{\left( {\frac{{F_{\text{v}} }}{{F_{\text{m}} }}} \right)_{{{\text{DK}}_{0} }} - \left( {\frac{{F_{\text{v}} }}{{F_{\text{m}} }}} \right)_{{{\text{DK}}_{{2\;{\text{h}}}} }} }}{{\left( {\frac{{F_{v} }}{{F_{\text{m}} }}} \right)_{{{\text{DK}}_{0} }} }}} \right\} \times 100$$
(3)

The percentage variations of Fo and Fm called ΔFo(2 h) and ΔFm(2 h), respectively, were also calculated over a 2-h dark period according to:

$$\Delta F_{{{\text{O}}\;{\text{or}}\;{\text{m }}(2\;{\text{h}})}} = \left\{ {\frac{{\left( {F_{{{\text{o}}\;{\text{or}}\;{\text{m}}}} } \right)_{{{\text{DK}}_{0} }} - \left( {F_{{{\text{o}}\;{\text{or}}\;{\text{m}}}} } \right)_{{{\text{DK}}_{{2\;{\text{h}}}} }} }}{{\left( {F_{{{\text{o}}\;{\text{or}}\;{\text{m}}}} } \right)_{{{\text{DK}}_{0} }} }}} \right\} \times 100$$
(4)

The morphometric (Table 1) and optical properties of the tissues of the 10 tested species differ and may partially influence PAM measurements (Enriquez et al. 2017; Wangpraseurt et al. 2019). In this study, the variations between light and dark transition were compared. Consequently, even if the saturating pulse was not able to saturate all the microalgae all through the layer of tissue of the corals, this would not influence the observed photobiology dynamics. In order to easily compare the F, Fo and Fm′, Fm dynamics, the values were normalized to the maximum value of Fm. According to the manufacturer’s recommendations, the range of raw fluorescence was 200 to 500.

Serial irradiance pulses (SIP) fluorescence response

The serial irradiance pulses (SIP) fluorescence responses were recorded as proposed by Reynolds et al. (2008) to investigate the post-illumination behavior of the PQ pool. The SIP protocol was applied to the samples within 3 h of their arrival at the laboratory. Serial irradiance pulses (SIP) fluorescence responses were recorded independently in triplicate for each species. The same diving PAM as the one described above was used. The protocol was performed as follows: Scleractinian corals were acclimated to the dark for 1 h. Base line fluorescence was monitored using a modulated weak red measuring light (1 µmol photons m−2 s−1, 650 nm, frequency 0.6 kHz), and Fm was measured by a 1-s saturating flash (650 nm, 4000 µmol photons m−2 s−1) at 30 s intervals. The minimum fluorescence measured after a saturating flash corresponds to Fo′. An increase of Fo′ after the flash reveals a post-illumination PQ pool reduction (Reynolds et al. 2008; Roberty et al. 2014) while a stability of Fo′ indicates sustained linear electron transport activities between PSII and PSI.

Field measurements

Measurements were made using the same setup as described above in three independent colonies. The scleractinian corals were placed in the dark for Fv/Fm measurements using a simple homemade experimental device comprising a black rubber swimming cap with the PAM Fiber optic cable running through it. The swimming cap covered the coral and ensured it was completely in the dark without damaging it.

A field index was calculated to estimate variations of the Fv/Fm. The index corresponds to the percentage of the Fv/Fm variation during the first 3 min of the dark period, DK0 being the Fv/Fm at the beginning of the dark phase and DK3 min the Fv/Fm after 3 min of dark. A 3-min interval was chosen because of scuba diving constraints and because a significant decrease of Fv/Fm was already observed in Porites astreoides within this short dark period.

$$\Delta {\text{YII}}_{{{ \hbox{max} }(3\;{ \hbox{min} })}} = \left\{ {\frac{{\left( {\frac{{F_{\text{v}} }}{{F_{\text{m}} }}} \right)_{{{\text{DK}}_{0} }} - \left( {\frac{{F_{\text{v}} }}{{F_{\text{m}} }}} \right)_{{{\text{DK}}_{{3\;{ \hbox{min} }}} }} }}{{\left( {\frac{{F_{\text{v}} }}{{F_{\text{m}} }}} \right)_{{{\text{DK}}_{0} }} }}} \right\} \times 100$$
(5)

Data analyses

Analyses of variance (ANOVA) were followed by pairwise multiple comparison (Tukey’s test). Previously, a Kolmogorov–Smirnov normality test and an equal variance (homoscedasticity) test were applied. Differences were considered significant when the p value was less than 0.05. When ANOVA could not be performed, a Kruskal–Wallis one-way analysis of variance by ranks test was applied. All the plots were produced using SigmaPlot 12.5 (Systat Software Inc. Chicago, USA).

Results

Fo and Fm (or Fq′) measured on Porites astreoides and Pseudodiploria strigosa displayed very different dynamics during the light/dark transitions and during the night (Fig. 1). In Porites astreoides, in the dark, Fo increased significantly from 31.50 to 58.65 (slope test p < 0.001) whereas in Pseudodiploria strigosa, it decreased significantly from 34.68 to 28.65 (slope test p = 0.01). A decrease in Fo in the dark was also observed in the seven other species tested: Acropora palmata, Agaricia agaricites, Orbicella annularis, O. faveolata, Porites furcata, Pseudodiploria clivosa, and Siderastrea siderea. Interestingly, Porites divaricata showed an increase in Fo but the magnitude was much lower than in Porites astreoides (Supplementary Fig. 1). At the end of the light period, during the light dark transition, Pseudodiploria strigosa showed an increase in Fq′/Fm′ from 0.632 up to a maximum 0.661, which was reached during the dark phase (Fv/Fm). This increase in Fq′/Fm′ was due to a decrease in fluorescence (F) in the dark associated with a weak decrease in Fm′ and Fm. In contrast, in Porites astreoides, Fv/Fm dropped from 0.642 to reach a minimum 0.271 within 2 h after the night phase. This drop was mainly due to the increase in Fo. The light–dark transition was performed under natural light and with a natural light cycle, but the same results were observed when the samples were placed in the dark during the daylight period.

Fig. 1
figure 1

Dynamics of F (black circle), Fm or Fm′ (white circle) and of Fv/Fm or Fq′/Fm′ (white triangle) in one sample of Porites astreoides (a) and in one sample of Pseudodiploria strigosa (b) during light dark transitions (dotted line: PAR)

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Based on these observations, an index was built to estimate Fv/Fm dynamics. The ΔYIImax(2 h) highlights the singular behavior of Porites astreoides sampled in shallow water that showed a mean value of 47.79% while the average of the nine other species tested was 0.677% (Fig. 2). The mean ΔYIImax(2 h) value of the two other Porites was 3.29% and − 1.15% for P. divaricata and P. furcata, respectively. The mean ΔYIImax(2 h) value of Porites astreoides differed significantly (F(1,29) = 63.665, p < 0.001, Tukey’s test p < 0.001) from the other species tested, whereas no significant differences were observed among the nine other species.

Fig. 2
figure 2

ΔYIImax(2 h) (%) corresponding to the percentage of Fv/Fm variation during the first 2 h of the dark period, measured in triplicate (mean ± SD) in 10 species of Caribbean scleractinian corals. Data were obtained from samples collected at a depth of 1 to 5 m. Different lowercase letters indicate significantly different means (p < 0.05)

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The mean ΔFo(2 h) values of all the species were higher than zero, except Porites astreoides in which a major significant increase in Fo occurred during the night (Fig. 3). The increase in Fo during the night led to a very high negative value of ΔFo(2 h) (− 108.64% ± SD 21.48) significantly different from all other species (F(1,29) = 28.788, p < 0.001, Tukey’s test p < 0.001). In contrast, Agaricia agaricites and Porites furcata presented high positive values of, respectively, ΔFo(2 h) (21.22% ± SD 19.81) and (19.09% ± SD 15.58), indicating a marked decrease in Fo during the dark phase, but these species displayed high variability. Fm remained more stable than Fo even in Porites astreoides. ΔFm(2 h) was positive in all species and ranged between 5.01% (Orbicella faveolata) and 21.40% (Agaricia agaricites), indicating a decrease in Fm after 2 h of adaptation to the dark. No significant differences were observed in the relationship between the species tested (KW: χ2 = 6.110, df = 9, p = 0.729). This result highlights the fact that the ΔYIImax(2 h) dynamics is mainly controlled by the Fo dynamics in Porites astreoides.

Fig. 3
figure 3

ΔFo(2 h) (%) and ΔFm(2 h) (%) measured in triplicate in 10 species of Caribbean scleractinian corals after 2 h in the dark (mean ± SD). Data were obtained on samples taken collected at a depth of 1 to 5 m. Different lowercase letters indicate significantly different means (p < 0.05) between ΔFo(2 h) (%) values

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In order to confirm a reduction of the quinone pool of the PSII during the night, Porites astreoides was exposed to a moderate light source (150 µmol photons m−2 s−1) for 1 min, at two periods in the night before the PAM saturating flashes were produced (Fm measurement) (Fig. 4). An increase in Fv/Fm from 0.08 to 0.52 was observed in the first light period, and from 0.09 to 0.61 in the second light period. These increases were mainly due to the decrease in Fo as described above. Then, in the dark, following these two short light periods, Fv/Fm dropped to previous values. At the end of the night Fq′/Fm′ increased on exposure to sunlight and reached 0.69 at 7:45 a.m. (Fig. 4).

Fig. 4
figure 4

Effects of a short light phase during the dark period on Fv/Fm in one sample of Porites astreoides. (Dotted line: PAR)

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The ΔYIImax(3 min) measured in situ (Fig. 5) was in agreement with the trends observed in ΔYIImax(2 h) (Fig. 2). As observed for ΔYIImax(2 h), the mean value of ΔYIImax(3 min) of Porites astreoides was significantly higher than all the other species (F(1,29) = 32.472, p < 0.001, Tukey’s test p < 0.001). In all the species tested, except Acropora palmata (3.56% ± SD 3.30) and Porites furcata (2.59% ± SD 1.53), the ΔYIImax(3 min) was lower than the ΔYIImax(2 h). For example, the mean ΔYIImax(3 min) value of Porites astreoides was 32.22%, which corresponds to a decrease of 15.57 between ΔYIImax(2 h) and ΔYIImax(3 min). In all the other species, the average decrease between the two ΔYIImax calculated was 4.54. Except for the three species of the Porites genus and Acropora palmata, the other tested species had a negative ΔYIImax(3 min). Agaricia agaricites, presented the lowest mean value of ΔYIImax(3 min) (− 23.62% ± SD 6.51) which differed significantly from that in all the other species (F(1,29) = 32.472, p < 0.001, Tukey’s test p < 0.001) except S. siderea (− 11.70% ± SD 6.47) (Tukey’s test p = 0.064).

Fig. 5
figure 5

ΔYIImax(3 min) (%) corresponding to the percentage of Fv/Fm variation during the first 3 min of the dark period, measured in triplicate (mean ± SD) in 10 species of Caribbean scleractinian corals. Data come from samples collected at a depth of 1 to 5 m. Different lowercase letters indicate significantly different means (p < 0.05)

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Because Porites astreoides showed a particular physiological behavior in shallow water, additional measurements were made at other depths. ΔYIImax(3 min) of Porites astreoides was measured in situ at four different depths, 4, 10, 15 and 20 m (Fig. 6). The mean coefficient of light extinction (Kpar) in the sampling area was 0.07 m−1. No significant differences between ΔYIImax(3 min) values were observed as a function of depth (F(1,11) = 0.234, p = 0.870). Considering all depths, the mean value of ΔYIImax(3 min) was 32.71% ± SD 6.72.

Fig. 6
figure 6

ΔYIImax(3 min) (%) corresponding to the percentage variation in Fv/Fm during the first 3 min of the dark period, measured in situ in triplicate on Porites astreoides at four different depths (4, 10, 15, 20 m) (mean ± SD)

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The SIP protocol was applied to the dark adapted scleractinian corals selected in this work. We observed two types of responses (Fig. 7). After activation of the measuring light that led to the first increase in fluorescence in Porites astreoides, we observed a progressive increase in Fo′ that can be attributed to a reduction of the plastoquinone pool. More precisely, after the second saturating flash, a light-induced plastoquinone oxidation is observed leading to a decrease of Fo′ just after the flash, which was immediately followed by an increase of Fo′ due to PQ dark reduction. Conversely, in Pseudodiploria strigosa, after the first increase in fluorescence due to measuring light, Fo′ remained stable, pointing to an equilibrium between reduction and oxidation of the plastoquinone pool due to linear electron transport between PSII and PSI. We then classified the two kinds of response of PQ-reduction SIP response as “positive” or “neutral” (Table 2). The three species of Porites tested displayed AEF activity while all the other species tested showed a similar pattern to that of Pseudodiploria strigosa, indicating no PQ-reduction. A positive PQ-reduction SIT response was measured in Porites astreoides at depths of 4, 10, 15 and 20 m. The decrease in maximum fluorescence observed after several flashes for both types of SIP responses, can be attributed to non-phototochemical quenching regulations.

Fig. 7
figure 7

Examples of CEP-SIP responses in a sample of Porites astreoides (a) and in a sample of Pseudodiploria strigosa (b)

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Table 2 PQ-reduction-SIP responses in 10 species of Caribbean scleractinian corals
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Discussion

In the present study, various protocols were applied dealing with the physiological regulations of the electron transport chain of coral-associated Symbiodiniaceae with a focus on the light/dark transition. Measurements were made in situ and in freshly sampled corals placed in regulated seawater tanks, and no difference was observed between the two, indicating that mechanisms explored here were not affected by perturbations resulting from handling. The in situ protocols enabled wide screening of the physiological behavior of corals without being obliged to collect them, while the laboratory protocols allowed us to explore physiological mechanisms in more detail.

An archetypal behavior of the maximum photochemical quantum yield during the dark phase consists of an increase in this parameter, due to a decrease in Fo caused by oxidation of the PQ pool (Genty et al. 1989). In the present study, this behavior was observed in all the species tested except in Porites astreoides. In this species, the main result of our experiments was the marked variation in the maximum photochemical quantum yield after only a few minutes of dark exposure. The marked decrease in Fv/Fm was mainly due to the increase in Fo in the dark phase (Fig. 1), while Fm remained relatively stable.

There is only a limited literature on the subject of PSII fluorescence in coral during the dark period. In a previous work on three species of coral, Acropora millepora, Pocillopora damicornis and Pavona decussata from the Great Barrier Reef in Australia (Hill et al. 2012), it was shown that Fv/Fm display archetypal behavior, as we observed in nine of the coral species we tested. In another study on Porites astreoides in Belize (Caribbean Sea), Warner et al. (2006) observed no reduction in Fv/Fm in the dark phase. These authors mentioned that their measurements were made in the early morning close to sunrise, but in a preliminary work cited in the article, they did not report any differences between morning Fv/Fm values and those recorded approximately 1 h after sunset. The data we obtained from the same species showed that Fv/Fm increased at the beginning of sunset and that very low light led to a decrease in Fo due to rapid re-oxidation of the PQ pool (Fig. 1). Warner et al. (2006) probably did not observe the mechanism we observed because it disappears at very low light (below 10 µmol photons m−2 s−1) and is therefore already suppressed by sunrise. Another possible explanation for the apparent discrepancy between the two studies is the diversity within the Symbiodiniaceae complex. Several lineages of Symbiodiniaceae may be present in the host and may differ depending on environmental conditions, including depth (Rowan and Knowlton 1995; Iglesias-Prieto et al. 2004; Reynolds et al. 2008; LaJeunesse et al. 2010; Byler et al. 2013; Pochon et al. 2015). Nevertheless, several authors showed that the Symbiodiniaceae associated with Porites astreoides in the Caribbean Sea are quite homogenous in shallow water and are mainly represented by Symbiodinium ITS2 type A4 (Reynolds et al. 2008; Hauff et al. 2016). The homogeneity of the response we observed in P. astreoides suggests that the same Symbiodinium symbiont occurs in shallow waters and down to a depth of 20 m (Fig. 6). The complementary molecular analyses performed during the course of the present study demonstrated the presence of Symbiodinium ITS2 type A4 in our samples (Japaud 2017). ITS2 type A4 is commonly reported in symbiotic association with Porites astreoides and is considered to belong to the “shallow water specialist” lineage of Symbiodiniaceae (Rowan and Knowlton 1995; LaJeunesse 2002; Serrano et al. 2016).

The increase in Fo in the dark phase we measured can be attributed to the reduction of the PQ pool, which is typical of chlororespiration-like mechanisms, as already reported in isolated Symbiodiniaceae lineages in culture, and in coral samples (Jones and Hoegh-Guldberg 2001; Hill and Ralph 2008; Reynolds et al. 2008). To evaluate the proposed chlororespiration-like mechanism in the Caribbean coral species, we propose two simple indexes, ΔYIImax(2 h) and ΔYIImax(3 min). Both indexes highlight the particular PQ dynamics of Porites astreoides in the dark with a very high value of ΔYIImax. We assume these indexes can be used as indicators of chlororespiration-like mechanisms. We observed that the chlororespiration-like mechanism was triggered immediately after exposure to the dark, it appears to be independent of any circadian clock, and brief exposure to light can interrupt it (Fig. 4). Chlororespiration is due to the oxidation of carbon reserves, with oxygen as a terminal electron acceptor using photosynthetic electron transfer chain by sharing plastoquinone and some stromal electron carriers (NADPH dehydrogenase, cytochrome b6 fcomplex, terminal oxidase (PTOX)) (Nawrocki et al. 2019). This process, which behaves like an antagonist of the light reactions of the photosynthesis by consuming O2 and NADPH, is preserved in most photosynthetic lineages (Nawrocki et al. 2015).

Different enzymes are involved in chlororespiration in higher plants and algae, and two types of enzymes play a major role. The first, the NAD(P)H:PQ oxidoreductase presents two forms, called type I NADH dehydrogenase (NDH-1) and type II NADH dehydrogenase (NDH-2), that reduce PQ. NDH-2, a non-electrogenic enzyme (Peltier et al. 2016), was found in Symbiodiniaceae clade A (genus Symbiodinium) (Aihara et al. 2016). Because NDH-2 is a non-electrogenic enzyme, recent evidence suggests it is primarily involved in PSI-CEF rather than in chlororespiration (Peltier et al. 2016; Aihara et al. 2016). The second type of enzyme, PTOX (plastid terminal oxidase) takes up electrons from plastoquinol (PQH2) to reduce oxygen (Nawrocki et al. 2015). Two types of PTOX (PTOX 1 and PTOX 2) have been found in Prasinophyceae, Chlorophyceae, Bacillariophyceae and Rhodophyta. To our knowledge, no information is yet available concerning Symbiodiniaceae. However, Roberty et al. (2014) identified a PTOX-related sequence in the EST database of Symbiodiniaceae belonging to clade A (genus Symbiodinium) but their results suggest low PTOX activity. Houille-Vernes et al. (2011) and Nawrocki et al. (2019) showed that PTOX2 is the main oxidase involved in chlororespiration in Chlamydomonas reinhardtii and its absence from K.O. PTOX2 mutant led to the same type of fluorescence behavior as we observed in Porites astreoides. The reduced efficiency of PTOX 1 electron flows for oxidized PQH2 compared to those of PTOX2 entail a higher reduction of the PQ pool and a lower value Fv/Fm in the dark (Houille-Vernes et al. 2011; Nawrocki et al. 2019). Even if it is not yet possible to draw a parallel between studies performed on C. reinhardtii and our results, it is possible that the singular behavior of Porites astreoides we observed is due to a difference in PTOX function and/or activity between species of Symbiodiniaceae.

Almost all current studies on Symbiodiniaceae photobiology deal with photosynthesis and the regulation of photoacclimation mechanisms under high light intensity (Suggett et al. 2008; Roberty et al. 2014, 2015; Hill et al. 2012; Slavov et al. 2016). Beyond linear electron flow (LEF) and standard non-phototochemical quenching acclimation, AEF has been described in many articles and the absence of state transition regulation has been pointed out in Symbiodiniaceae (Hill et al. 2012; Kanazawa et al. 2014; Roberty et al. 2014, 2015). Like Reynolds et al. (2008), we found positive PQ-reduction SIP responses in the three species of the genus Porites we tested here. Reynolds et al. (2008) first attributed this regulation to PSI-CEF and chlororespiration, then Roberty et al. (2014) showed that the “Water-Water Cycle” (Mehler reaction) was the main AEF in Symbiodiniaceae lineages ITS2 type A1, B1, F1, enabling photoreduction of oxygen by PSI under high light intensity and moderate temperatures (Roberty et al. 2015). Therefore, the chlororespiration pathway was only proposed as a weak mechanism in the dissipation of energy during the light phase (Roberty et al. 2015).

At this stage, our study does not enable us to characterize the chlororespiration-like mechanism precisely, however, it does allow us to ask an important question: how does Porites astreoides benefit from a potential high reduction power during the night? Several hypotheses are possible but will require further experiments with different data than the ones obtained using PAM fluorometry.

In addition, the role of chlororespiration is still the subject of debate [for a review, see Nawrocki et al. (2015), and Peltier et al. (2016)]. Chlororespiration has been linked to nitrogen depletion in the green microalgae Chlamydomonas and Haematococcus (Peltier and Schmidt 1991; Scibilia et al. 2015; Saroussi et al. 2016, 2017), but there is no evidence of such a relationship in Symbiodiniaceae.

According to the literature, in the dark, chlororespiration maintains the redox poise of the photosynthetic electron transport chain thereby allowing more efficient photosynthesis upon re-illumination (Jones and Hoegh-Guldberg 2001; Nawrocki et al. 2019). This redox buffer could also play an indirect photoprotective role by promoting CEF and the ensuing photoprotective mechanisms (Nawrocki et al. 2015). Because our experiments were performed under natural light with a natural light cycle, we were not able to test this hypothesis, but future experiments using a controlled light cycle could provide relevant information regarding such a regulation.

Chlororespiration has also been shown to be one of the pathways involved in ROS-scavenging systems in chloroplasts that reduce the production of long-lived redox active compounds (Khorobrykh et al. 2020). Chlororespiration has been shown to be able to consume reactive oxygen species (ROS) and may reduce their production by reducing the O2 concentration in chloroplasts (Casano et al. 2000; Nixon 2000; Jones and Hoegh-Guldberg 2001, Beardall et al. 2003). ROS are mainly produced during the light phase but also in the dark, in the absence of photosynthetic activity (Tolleter et al. 2013; Diaz et al. 2016, Zhang et al. 2016). In coral, ROS are produced by the photosynthetic symbiont and the host, so diffusive transfer of ROS from one symbiotic partner to the other may occur (Nielsen et al. 2018; Zhang et al. 2016). In the presence of reduced PQ (PQH2), as we observed in our study on P. astreoides, PQH2 oxidation by a superoxide anion radical (a type of ROS) leads to the accumulation of H2O2 in the thylakoid membrane (Khorobrykh and Ivanov 2002; Khorobrykh et al. 2020). The mechanism of oxidation of PQH2 by a superoxide has not yet been directly explored in Symbiodiniaceae, but we found this to be consistent with observations of Zhang et al. (2016), who showed elevated production of H2O2 in P. astreoides in the dark. The chlororespiration-like mechanism we found could help reduce or balance the concentration of ROS produced by the symbiont and (or) originating in the host gastroderm.

The proposed antioxidant capacity of Porites astreoides in the dark phase will require further investigation considering the interactions between the photosynthetic symbiont and its host. However, it already allows us to explain the singular physiological mechanism found in Porites astreoides that probably enhances or at least contributes to its physiological resistance to current environmental changes. Our data favor the hypothesis that P. astreoides present peculiar physiological capabilities thanks to the symbiosis, which may enhance resilience among the Caribbean reef corals.