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Research Article
Future ocean warming may prove beneficial for the northern population of European seabass, but ocean acidification will not
Sarah Howald, Louise Cominassi, Nicolas LeBayon, Guy Claireaux, Felix C. Mark
Journal of Experimental Biology 2019 222: jeb213017 doi: 10.1242/jeb.213017 Published 11 November 2019

ABSTRACT

The world's oceans are acidifying and warming as a result of increasing atmospheric CO2 concentrations. The thermal tolerance of fish greatly depends on the cardiovascular ability to supply the tissues with oxygen. The highly oxygen-dependent heart mitochondria thus might play a key role in shaping an organism's tolerance to temperature. The present study aimed to investigate the effects of acute and chronic warming on the respiratory capacity of European sea bass (Dicentrarchus labrax L.) heart mitochondria. We hypothesized that acute warming would impair mitochondrial respiratory capacity, but be compensated for by life-time conditioning. Increasing PCO2 may additionally cause shifts in metabolic pathways by inhibiting several enzymes of the cellular energy metabolism. Among other shifts in metabolic pathways, acute warming of heart mitochondria of cold life-conditioned fish increased leak respiration rate, suggesting a lower aerobic capacity to synthesize ATP with acute warming. However, thermal conditioning increased mitochondrial functionality, e.g. higher respiratory control ratios in heart mitochondria of warm life-conditioned compared with cold life-conditioned fish. Exposure to high PCO2 synergistically amplified the effects of acute and long-term warming, but did not result in changes by itself. This high ability to maintain mitochondrial function under ocean acidification can be explained by the fact that seabass are generally able to acclimate to a variety of environmental conditions. Improved mitochondrial energy metabolism after warm conditioning could be due to the origin of this species in the warm waters of the Mediterranean. Our results also indicate that seabass are not yet fully adapted to the colder temperatures in their northern distribution range and might benefit from warmer temperatures in these latitudes.

INTRODUCTION

The increasing amount of atmospheric CO2 is working as a greenhouse gas, raising atmospheric temperatures and, as a consequence, also sea surface temperatures (ocean warming). At the same time, about a third of the atmospheric CO2 is taken up by the oceans, which leads to decreasing seawater pH through the formation of carbonic acid (ocean acidification). These processes together will lead to warmer and more acidic oceans, a trend that has already been observed over the last decades and is predicted to continue. Depending on the representative concentration pathway, the IPCC (2014) predicts that the partial pressure of CO2 (PCO2) will increase up to 1000 µatm above current values until the end of this century. Temperature projections for the same time span predict increases of up to 4°C at the coast of Brittany (Sheppard, 2004). Changes in water temperature have direct influence on the metabolic rate of ectothermic organisms, such as fish, with consequences for growth (Pörtner et al., 2007; Peck, 2002), reproductive success (for review, see Llopiz et al., 2014) and biogeography (Turner et al., 2009; Pörtner, 2006). The body of studies looking into the effects of the changing environment on marine ectothermic organisms is growing rapidly. However, only a small number of these studies have investigated the effects of ocean acidification and the combined effects of ocean acidification and warming on fish, with contrasting results between species as well as life stages (Kreiss et al., 2015; Heuer and Grosell, 2014; Pope et al., 2014; Bignami et al., 2013; Frommel et al., 2012). More investigation on a variety of fish species, on different life stages and under ecologically relevant PCO2 concentrations appears necessary (Pope et al., 2014).

The effects of temperature on fish metabolism have been investigated intensively (e.g. Johnson and Katavic, 1986; Mirkovic and Rombough, 1998; Bláquez et al., 1998; Farrell, 2002; Pörtner et al., 2007; Hilton et al., 2010; Strobel et al., 2012). Thermal sensitivity of fish is mainly dependent on the capacity of the cardiovascular system to supply the tissues with oxygen (Pörtner and Lannig, 2009). The heart is a highly aerobic tissue (Driedzic, 1992) and it is therefore believed that the capacity of the heart mitochondria to produce ATP plays a central role in defining thermal tolerance in fish. Although the subcellular processes are not yet fully understood, it has been suggested that the functionality of heart mitochondria determines the temperature of heart failure, and that heart thermal acclimation capacity is relatively limited to safeguard functionality (e.g. Chen and Knowlton, 2010; Chung et al., 2017; Iftikar and Hickey, 2013; Strobel et al., 2013a). Different mitochondrial processes have been shown to be impaired at elevated temperatures, as indicated by decreased oxidative phosphorylation, decreased ATP production efficiency and lost integrity of the protein complexes of the electron transport system (e.g. Fangue et al., 2009; Hilton et al., 2010; Mark et al., 2012; Iftikar and Hickey, 2013). Additionally, high temperatures increase the fluidity of mitochondrial membranes, potentially leading to increased proton leak through the inner mitochondrial membrane and decreased mitochondrial efficiency (Pörtner, 2012). Impaired mitochondrial metabolism thus might lead to alterations in cardiomyocyte ATP supply and consequently affect the performance of the cardiovascular system, which will ultimately determine the thermal sensitivity of the fish. Although juvenile and adult fish generally possess well-developed acid–base regulating mechanisms (for review, see Heuer and Grosell, 2014), increased ocean acidification may act as an additional stressor, e.g. by increasing the ATP demand for acid–base regulation. Consequently, well-functioning mitochondria are even more important if temperature and PCO2 are increased simultaneously. However, while the body of literature on the effects of temperature on mitochondrial function in fish is relatively large (e.g. Fangue et al., 2009; Shama et al., 2014, and references therein), only a handful of studies have investigated the effects of increased PCO2 on fish mitochondria and even fewer have combined ocean acidification and ocean warming (but see Strobel et al., 2013a; Leo et al., 2017). CO2 can freely diffuse out of the water into the extracellular, intracellular and mitochondrial space. Just as in the blood, decreased intracellular pH due to elevated PCO2 is buffered by actively raising intracellular bicarbonate levels (Strobel et al., 2013a). Mitochondrial membranes are permeable to CO2 but impermeable to bicarbonate (Arias-Hidalgo et al., 2016), resulting in elevated matrix bicarbonate levels under high CO2 (Pörtner and Sartoris, 1999; Strobel et al., 2012). Bicarbonate acts as a competitive inhibitor of mitochondrial citrate synthase and succinate dehydrogenase in rodents (Simpson, 1967; Wanders et al., 1983), which can be overcome by a compensatory increase in mitochondrial capacity following hypercapnia acclimation – although this has not yet been documented unequivocally for marine fish (Strobel et al., 2012; 2013a,b). Bicarbonate also acts as a pH-sensing molecule in marine fish and invertebrates (Tresguerres et al., 2014; Barott et al., 2017), with both upregulating and downregulating consequences on mitochondrial metabolism. As over the coming decades ocean temperature and PCO2 will rise hand in hand, it is important to determine their combined effects on mitochondrial metabolism to be able to take ecologically relevant conclusions. Furthermore, there is a relative lack of studies that focus on temperate, large and/or economically relevant species, while at the same time employing realistic PCO2 and exposure scenarios.

List of abbreviations

A
ambient PCO2 condition
C
cold life-conditioned group
CCO
cytochrome c oxidase
CI–IV
complex I–IV of the electron transport system
CTmax
critical thermal maximum
dd
degree days
dph
days post-hatching
LOmy
LEAK respiration with oligomycin
LME model
linear mixed effect model
P
OXPHOS respiration
RCP
representative concentration pathway
RCROmy
respiratory control ratio with oligomycin
W
warm life-conditioned group
Δ500
ambient PCO2+500 µatm acidification condition
Δ1000
ambient PCO2+1000 µatm acidification condition

In our study, we exposed European seabass, Dicentrarchus labrax (L.), from 3 days post-hatching (dph) for 7 months to two temperatures and three PCO2 conditions in a full-factorial design. Temperature and PCO2 conditions reflect the predictions of the IPCC for 2100 (IPCC, 2014). The European seabass is an important aquaculture species (160,000 metric tons in 2015), but also an important target in commercial as well as in recreational fishing activities (Bjørndal and Guillen, 2018). European seabass are distributed throughout the Mediterranean, the Black Sea and the North-Eastern Atlantic from Norway to Senegal in coastal waters from coastal lagoons and estuaries up to 100 m depth (Bjørndal and Guillen, 2018). It has a relatively high tolerance to different temperatures and salinities (Dalla Via et al., 1998; Claireaux and Lagardère, 1999). We studied the effect of ocean acidification and warming on mitochondria of juvenile seabass by determining mitochondrial respiratory capacity in permeabilized heart fibers of seabass juveniles that had been reared under the respective ocean acidification and warming conditions since hatching (7 months). We examined the effects of ocean acidification and warming on mitochondrial ATP-producing processes (OXPHOS respiration) and the counteracting proton leak (LEAK respiration), as well as their resulting ratio (the respiratory control ratio, RCR). An increased LEAK respiration, which is not compensated for by increased OXPHOS respiration, results in a drop in RCR and indicates that mitochondrial functionality and consequently mitochondrial capacity to produce ATP are impaired.

We hypothesized that acute warming would impair mitochondrial performance in juvenile seabass hearts, as LEAK respiration may increase with thermal deterioration of mitochondrial membranes. However, compensational processes after long-term and developmental thermal conditioning could include changes in mitochondrial membrane properties, which would reduce LEAK respiration rates and consequently restore RCR. Additionally, we wanted to fathom the capacity of seabass mitochondria to cope with ocean acidification, especially when combined with ocean warming. We hypothesized that the changes in intracellular PCO2 and bicarbonate concentration elicited by ocean acidification would affect mitochondrial metabolism, e.g. by inhibiting citrate synthase and succinate dehydrogenase, putting further pressure on the cellular energy metabolism.

MATERIALS AND METHODS

The present work was performed within Ifremer-Centre de Bretagne facilities (agreement number: B29-212-05). Experiments were conducted according to the ethics and guidelines of French law and legislated by the local ethics committee (Comité d'Ethique Finistérien en Experimentation Animal, CEFEA, registering code C2EA-74; authorization APAFIS 4341.03, permit number 2016120211505680.v3).

All chemicals were purchased from Sigma-Aldrich, except for tricaine methane sulfonate (MS-222), which was purchased from Pharma Q Limited.

Datasets of mitochondrial respiration and water conditions during rearing, as well as additional information on larval rearing are available online in PANGAEA (www.pangaea.de/).

Animals and experimental conditions

Animals

Larvae were obtained from the aquaculture facility Aquastream (Ploemeur-Lorient, France) at 2 dph (20 January 2016). Brood stock fish were caught in the sea off Morbihan, France. Four females (mean mass 4.5 kg) were crossed with 10 males (mean mass 2.4 kg), which spawned naturally using photothermal manipulation. Conditions in the aquaculture facility during breeding were as follows: 13°C, 35 psu, pH 7.6, 8 h 45 min of light followed by 15 h 15 min darkness. Spawning of eggs took place on 15 January 2016; larvae hatched on 18 January 2016 and were transported to our structures on 20 January 2016.

Larval rearing

Larval rearing was performed in a temperature-controlled room using black 35 l tanks initially stocked with ca. 5000 larvae per tank. Three replicate tanks were used for each temperature–PCO2 combination. Larvae were randomly distributed into the experimental tanks at 3 dph (21 January 2016). During the following 3 days, the temperature for the warm life condition was increased stepwise, 1°C during the first day and 2°C during each of the two subsequent days. The PCO2 conditions were applied directly after division into the experimental tanks. Starting at 7 dph (mouth opening), larvae were fed with live artemia, hatched from High HUFA Premium artemia cysts (Catvis, AE's-Hertogenbosch, The Netherlands). Artemia were fed to the larvae 24 h after rearing cysts in sea water up to 33 dph; afterwards, the artemia nauplii themselves were fed with cod liver oil and dry yeast after 24 h and fed to the larvae after 48 h. The artemia were transferred to the larval rearing tanks from two storage tanks (one for each temperature) with peristaltic pumps; their concentration in the tanks was maintained high during the day to allow ad libitum feeding; excess artemia left the tank via the waste water outflow. The 15 h photoperiod in the larval rearing room lasted from 07:00 h to 22:00 h; the light intensity increased progressively during the larval rearing period from total darkness to 96 lx (Table S1). Headlamps were used (set to the lowest light intensity) to allow us to work in the larval rearing facility. Larval mortality was 10–80%, regardless of the rearing condition (Table S2). The water surface was kept free of oily films using a protein skimmer. Larval density was reduced regularly during larval rearing as samples for other experiments were taken throughout the entire larval rearing period (at approximately 100, 300, 500, 700, 750 and 900 degree days, dd=dph×temperature in °C). At approximately 980 dd, the early juveniles were transferred from the larval to the juvenile rearing facility, corresponding to 50 dph and 65 dph for 20°C and 15°C rearing, respectively.

Juvenile rearing

As they reached juvenile stage, fish were moved from the larval rearing facilities to juvenile tanks at approximately 1000 dd (50 dph, 8 March 2016, and 65 dph, 23 March 2016, for warm and cold life conditions, respectively). Fish were counted per tank and all fish from one condition were pooled in one tank until the swim bladder test and separation into duplicate tanks at 1541 dd (78 dph, 5 April 2016) and 1301 dd (86 dph, 13 April 2016) for warm and cold life conditions, respectively. The swim bladder test was done to keep only the fish with developed swim bladders. Briefly, the fish were anesthetized and introduced into a test container with seawater with a salinity of 65 psu. Those fish floating at the surface were removed from the test container and placed into the rearing tanks for recovery. The juveniles were reared in round tanks with a volume of 0.67 m3 and a depth of 0.65 m. Mortality rates of 24.8–43.4% (Table S3) occurred between moving to the juvenile facility and the swim bladder test. During the first 5 days after moving to juvenile rearing facilities, the juveniles were fed artemia nauplii (48 h old and enhanced with cod liver oil and dry yeast) and commercial fish food. Commercial fish food (Neo Start) was fed daily and was adjusted in size (1–3) and amount during the juvenile rearing time, as recommended by the supplier (Le Gouessant, Lamballe, France). More precisely, fish were fed ad libitum until 19 August 2016; afterwards, food ratios were calculated every 3–4 weeks for each tank in respect to biomass and temperature of the tank using the formulae provided by Le Gouessant. The daily ration of food was supplied to the tanks by automatic feeders during the daytime; the fish were not fed during the night-time. Photoperiod was adjusted to natural conditions once a week, with slowly increasing light intensity in the juvenile rearing facilities during the first hour each morning. The tanks were cleaned daily after pH measurements. Water flow within the tanks was adjusted once a week, so that oxygen saturation levels were not below 90%, keeping equal flow-through rates in all tanks of one temperature.

Experimental conditions

The larvae and juveniles were reared under six different ocean acidification and warming scenarios, following the predictions of the IPCC (2014) for the next 130 years. The acidification conditions included three different PCO2: today's ambient situation in coastal waters of Brittany and the Bay of Brest (ambient group – A, approximately 650 µatm; see Pope et al., 2014; Duteil et al., 2016), a scenario according to the IPCC representative concentration pathway RCP 6.0, projecting a ΔPCO2 of 500 µatm to current values (Δ500, approximately 1150 µatm) and a scenario according to RCP8.5, projecting a ΔPCO2 of 1000 µatm (Δ1000, approximately 1700 µatm). All acidification conditions were crossed with two different temperatures to create a ‘cold’ (C) and a ‘warm’ (W) life condition scenario. In the cold life condition, larvae were reared at 15°C; juveniles in the cold life condition were reared at 15°C until ambient temperature in the Bay of Brest reached 15°C, and from there on juveniles were reared at ambient temperatures of the Bay of Brest (up to 18°C in 2016). The warm life condition mirrored these thermal profiles, but with an offset of plus 5°C (20–23°C). As larvae and post-larval juveniles would display different growth rates under the two different thermal scenarios, we adopted the concept of degree days (dd, see above) as the basis for comparison between these life conditions.

The sea water used in the aquaria was pumped in from the Bay of Brest from a depth of 20 m approximately 500 m from the coastline, passed through a sand filter (∼500 µm), heated (tungsten, Plate Heat Exchanger, Vicarb, Sweden), degassed using a column, filtered using a 2 µm membrane and finally UV sterilized (PZ50, 75 W, Ocene, Louvigné-du-Désert, France), ensuring high water quality. During larval and early juvenile rearing, the water supply for the acidified incubation tanks came from a central header tank, where the water PCO2 conditions were adjusted. The water pH was controlled by an IKS Aquastar system (iks Computer Systeme GmbH, Karlsbad, Germany), which continuously measured pH in one of the replicate tanks and opened a magnetic valve to bubble CO2 into the header tank when pH in the rearing tank became too high. Water exchange was set to 20 l h−1 until 12 dph and 25 l h−1 until the end of larval rearing. During juvenile rearing with higher water exchange rates, additional PVC columns were installed to control the pH in the rearing tanks. The water arrived at the top of the column and was pumped from the bottom of the column to the rearing tanks. The CO2-bubbling apparatus was installed at the bottom of the column and was adjusted by a flow control unit when pH deviated from the desired value. One column supplied both replicate tanks of each condition. Temperature and pH were checked each morning with a handheld WTW 3110 pH meter (with a WTW Sentix 41 electrode, NIST scale; both from Xylem Analytics Germany, Weilheim, Germany) before fish were fed. The pH meter as well as the IKS Aquastar system were calibrated daily with NIST-certified WTW technical buffers pH 4.01 and pH 7.00 (Xylem Analytics Germany). Total alkalinity was measured once a week following the protocol of Anderson and Robinson (1946) and Strickland and Parsons (1972): 50 ml of filtered tank water (200 µm nylon mesh) was mixed with 15 ml HCl (0.01 mol l−1) and pH was measured immediately. Total alkalinity was then calculated with the following formula:Embedded Image (1)where TA is total alkalinity (mol l−1), VHCl is the volume of HCl (l), cHCl is the concentration of HCl (mol l−1), Vsample is the volume of the sample (l), [H+] is hydrogen activity (10−pH) and γH+ is the hydrogen activity coefficient (here γH+ = 0.758).

The Microsoft Excel macro CO2sys (Lewis and Wallace, 1998) was used to calculate seawater carbonate chemistry, using the constants proposed by Mehrbach and colleagues, refitted by Dickson and Millero (see CO2sys). Oxygen saturation (WTW Oxi 340, Xylem Analytics Germany) and salinity (WTW LF325, Xylem Analytics Germany) were measured once a week together with total alkalinity, from juvenile stage onwards (see Table 1 for all water parameters).

View this table:
Table 1.

Water parameters during larval and juvenile phase of the 2016 batch

Mitochondrial respirometry

Measurements of mitochondrial respiration rates were performed from approximately 3700 to 4100 dd in all conditions (183–199 dph and 234–249 dph in warm and cold life conditions, respectively). Although the same age in degree days was chosen, the cold life-conditioned fish were significantly smaller than the warm life-conditioned fish [length: 8.72±0.09 cm and 9.59±0.09 mm, respectively, least squares (ls)means±s.e.m., P<0.001; and carcass mass: 10.00±0.45 g and 13.38±0.46 g, respectively, lsmeans±s.e.m., P<0.001, linear mixed effect models (LME)], resulting in smaller ventricle sizes (0.0105±0.0004 g and 0.0122±0.005 g, respectively, P<0.05, lsmeans±s.e.m., LME), as well as lower hepatosomatic indices (1.51±0.05 and 2.45±0.06, respectively, lsmeans±s.e.m., P<0.0001, LME). However, condition factor was not different in the two temperature conditions (1.51±0.01 and 1.48±0.01, respectively, lsmeans±s.e.m., P>0.1, LME) (see Table S4). PCO2 did not have an effect on fish size. Prior to the experiments, the fish were starved for 2 days. Two batches of eight fish each were processed per day. Juveniles were randomly caught from their tanks and anesthetized with MS-222. The concentration of the anesthetic was adjusted to reach a loss of equilibrium within less than 5 min, typically 0.2 g l−1. Mass, fork length and body length were directly determined with a precision balance (Mettler, Columbus, OH, USA) and a caliper, to the nearest 0.01 g and 0.01 mm, respectively. Afterwards, fish were killed by a cut through the neck, and the heart was completely dissected from the fish, followed by excavation of the ventricle. Excess blood was removed from the ventricle by cleaning it on blotting paper, prior to weighing (Sartorius, Göttingen, Germany). Tissue from a whole ventricle was used for respiration measurements in each respiration chamber of the oxygraphs and respiration rates were normalized to ventricle mass. The ventricle was stored in relaxing and biopsy preservation solution [BIOPS: 10 mmol l−1 Ca-EGTA (0.1 µmol l−1 free calcium), 20 mmol l−1 imidazole, 20 mmol l−1 taurine, 50 mmol l−1 K-Mes, 0.5 mmol l−1 DTT, 6.56 mmol l−1 MgCl2, 5.77 mmol l−1 ATP, 15 mmol l−1 phosphocreatine, pH 7.1; modified after Gnaiger et al., 2000] until all eight ventricles were dissected. The ventricles were manually frayed and were permeabilized on ice with saponin (50 µg ml−1) for 20 min on a shaking table (80 rpm), followed by two cleaning steps for 10 min at 80 rpm in modified mitochondrial respiration medium [MiR05: 160 mmol l−1 sucrose, 60 mmol l−1 potassium-lactobionate, 20 mmol l−1 taurine, 20 mmol l−1 Hepes, 10 mmol l−1 KH2PO4, 3 mmol l−1 MgCl2, 0.5 mmol l−1 EGTA, 1 g l−1 bovine albumin serum (fatty acid free), pH 7.45 at 15°C; modified after Fasching et al., 2014]. During the permeabilization step, the livers and the carcasses of the fish were weighed to calculate the hepatosomatic index (HSI) and condition factor (K) (see Table S4).

Mitochondrial respiration of the permeabilized heart fibers was measured using four Oroboros Oxygraph-2K respirometers with DatLab 6 software (Oroboros Instruments, Innsbruck, Austria). Permeabilized fibers have the advantage of resembling the living state as closely as possible, while still allowing control of the supply of substrates and inhibitors to the mitochondria (Saks et al., 1998; Pesta and Gnaiger, 2012). Measurements were conducted at 15 and 20°C for all treatments to determine the effect of acute temperature changes on mitochondrial metabolism in vitro. The oxygen sensors were calibrated in air-saturated buffer prior to the experiments and in oxygen-depleted buffer after each experiment. The measurements were done in MiR06 buffer (MiR05 buffer enriched with 280 U ml−1 catalase; Fasching et al., 2014) to allow for reoxygenation with H2O2. A standard substrate–uncoupler–inhibitor titration protocol was employed to measure the respiration rates of the different complexes: glutamate (10 mmol l−1), malate (2 mmol l−1) and pyruvate (10 mmol l−1) were used to measure LEAK respiration of complex I [L(n)CI; Table 2], followed by addition of ADP (2.5 mmol l−1) to measure OXPHOS respiration of complex I (PCI). Succinate (10 mmol l−1) and further ADP (two additions of 2.5 mmol l−1) resulted in OXPHOS respiration of complex I (CI) and complex II (CII) combined (P). Cytochrome c (0.01 mmol l−1) was used as a control for inner mitochondrial membrane integrity; measurements with increases of more than 10% following cytochrome c addition were not used for further analyses. Oligomycin (4 µg ml−1) was used to inhibit FOF1-ATPase, resulting in LEAK respiration (LOmy). Stepwise titration of FCCP (0.05 µl of 2 mmol l−1 stock solution per step) was used to uncouple the mitochondrial electron transport system and determine its maximum capacity. After uncoupling, CI, CII and complex III (CIII) were successively inhibited with rotenone (0.005 mmol l−1), malonate (5 mmol l−1) and antimycin A (0.0025 mmol l−1), respectively. Residual respiration after antimycin A addition was used to correct all mitochondrial respiration rates. Complex IV (CIV) capacity was then determined by addition of ascorbate (2 mmol l−1) and TMPD (0.5 mmol l−1). Oxygen levels were usually restored by addition of 2 µl H2O2 (3% stock solution) after the oligomycin and rotenone steps.

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Table 2.

Analyzed mitochondrial metabolic states (after Gnaiger, 2014) during the substrate–uncoupler–inhibitor titration protocol

A measure for OXPHOS respiration of CII (PCII) was calculated (PCII=P−PCI), although this measure will lead to lower respiration rates than direct determination of PCII respiration, when only substrates for CII are available (Gnaiger, 2009; Mark et al., 2012). For all complexes, the contribution of the respiration of the respective complex to OXPHOS respiration was calculated. This was also done for LOmy, which we tentatively termed the ‘LOmy fraction’, as a relative indicator of proton leak, despite the fact that membrane potential is potentially higher in LOmy than in natural state 4. Mitochondrial quality and efficiency were evaluated by calculating the respiratory control ratio (RCROmy=P×LOmy−1), which is an indicator of mitochondrial coupling (Gnaiger, 2009; Strobel et al., 2013a).

Statistical analysis

All statistics were performed with R (http://www.R-project.org/). Data were tested for outliers (Nalimov test), normality (Shapiro–Wilk's test, P>0.05) and homogeneity (Levene's test, P>0.05). Mitochondrial respiratory data were fitted to linear mixed effect models (LME model, ‘lme’ function of ‘nlme’ package; https://CRAN.R-project.org/package=nlme). Conditioning and assay temperature, as well as PCO2 and their interactions were included as fixed effects. Because of the significantly different sizes of the fish, fish mass was also included as a fixed effect, whereas the oxygraph chamber and the number of the run on that day were included as random effects. In case of heterogeneity of data, variance structures were included in the random part of the model; the best variance structure was chosen according to the lowest Akaike information criterion (AIC) values. Validity of linearity for PCO2 concentration and mass was cross-tested with generalized additive models (‘gam’ function of ‘mgcv’ package; Wood, 2017), as described in Zuur et al. (2009). If linearity was given, the LME model was chosen instead of the generalized additive model. If significant effects were detected in the LME models, post hoc Tukey tests were performed with the ‘lsmeans’ function (‘lsmeans’ package; Lenth, 2016). Significance for all statistical tests was set at P<0.05. All graphs are produced from the lsmeans data with the ‘ggplot2’ package (Wickham, 2016). All data are shown as lsmeans±s.e.m. Biometrical data were also tested with LME models, with PCO2, conditioning and assay temperature as fixed effects and origin tank as a random effect. Model validation was carried out in the same way as described above for mitochondrial respiratory data.

RESULTS

Effects of acute in vitro warming on mitochondrial function in cold life-conditioned fish

PCII (Fig. 3) respiration rates increased significantly with acute warming in the Δ1000 group (LME, P<0.05; Table 3). All other analyzed parameters were not affected by acute warming: P (Fig. 1), PCI (Fig. 2) and the relative contributions of CI and CII to P (Table 3; CI fraction: 53.4±3.3–61.1±3.3%, Fig. S1; and CII fraction: 38.8±3.3–46.5±3.3%, Fig. S2).

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Table 3.

F-values of fixed effects from the linear mixed models on mitochondrial respiration of permeabilized heart fibers of juvenile European seabass

Fig. 1.
Fig. 1.

Respiration rates of permeabilized heart ventricle of European seabass. Respiration rate (oxidative phosphorylation capacity, P) data are least squares (ls)means±s.e.m. Different letters indicate significant differences [linear mixed effects (LME), P<0.05]; blue: cold life-conditioned fish (C), orange: warm life-conditioned fish (W), light shading: cold assay temperature, dark shading: warm assay temperature. A, ambient PCO2; Δ500, ambient+500 µatm CO2; Δ1000, ambient+1000 µatm CO2. nC–A=16/15, nC–Δ500=15/16, nC–Δ1000=16/16, nW–A=16/15, nW–Δ500=17/11, nW–Δ1000=13/15, for cold/warm assay temperature, respectively.

Fig. 2.
Fig. 2.

OXPHOS respiration rate of complex I (PCI) in permeabilized heart ventricle fibers of European seabass. Data are lsmeans±s.e.m. Different letters indicate significant differences (LME, P<0.05); blue: cold life-conditioned fish (C), orange: warm life-conditioned fish (W), light shading: cold assay temperature, dark shading: warm assay temperature. A, ambient PCO2; Δ500, ambient+500 µatm CO2; Δ1000, ambient+1000 µatm CO2. nC–A=16/15, nC–Δ500=14/16, nC–Δ1000=16/16, nW–A=16/14, nW–Δ500=17/11, nW–Δ1000=13/15, for cold/warm assay temperature, respectively.

Fig. 3.
Fig. 3.

OXPHOS respiration rate of complex II (PCII) in permeabilized heart ventricle fibers of European seabass. Data are lsmeans±s.e.m. Different letters indicate significant differences (LME, P<0.05); blue: cold life-conditioned fish (C), orange: warm life-conditioned fish (W), light shading: cold assay temperature, dark shading: warm assay temperature. A, ambient PCO2, Δ500: ambient+500 µatm CO2, Δ1000: ambient+1000 µatm CO2. nC–A=15/15, nC–Δ500=14/16, nC–Δ1000=16/16, nW–A=15/14, nW–Δ500=16/11, nW–Δ1000=13/15, for cold/warm assay temperature, respectively.

CIV capacity increased with acute warming in both hypercapnia groups (LME, Δ500: P<0.05 and Δ1000: P<0.05; Table 3) but not in fish at ambient PCO2 (Fig. 4). However, this increase was not strong enough to change the relationship between CIV capacity and P (Fig. S3).

Fig. 4.
Fig. 4.

Complex IV (CIV) respiration rates of permeabilized heart ventricle fibers of European seabass. Data are lsmeans±s.e.m. Different letters indicate significant differences (LME, P<0.05); blue: cold life-conditioned fish (C), orange: warm life-conditioned fish (W), light shading: cold assay temperature, dark shading: warm assay temperature. A, ambient PCO2; Δ500: ambient+500 µatm CO2; Δ1000: ambient+1000 µatm CO2. nC–A=12/12, nC–Δ500=14/16, nC–Δ1000=15/16, nW–A=15/14, nW–Δ500=17/11, nW–Δ1000=11/13, for cold/warm assay temperature, respectively.

LOmy respiration rate increased significantly with acute warming in the A and Δ1000 treatments (LME, P<0.05 and P<0.01, respectively; Table 3 and Fig. 5). Assay temperature had a significant effect on the relative contribution of LOmy to P (LOmy fraction; Table 3). However, no specific differences were detected with the post hoc tests (Fig. S4). As PCO2 did not affect the LOmy fraction, we pooled data over PCO2 treatments, which emphasized that acute warming led to impaired mitochondria in the cold life-conditioned fish, as indicated by a significantly higher LOmy fraction under warm assay temperatures compared with cold assay temperatures (Table 4; LME, P<0.05).

Fig. 5.
Fig. 5.

LEAK respiration with oligomycin (LOmy) of permeabilized heart ventricle fibers of European seabass. Data are lsmeans±s.e.m. Different letters indicate significant differences (LME, P<0.05); blue: cold life-conditioned fish (C), orange: warm life-conditioned fish (W), light shading: cold assay temperature, dark shading: warm assay temperature. A, ambient PCO2; Δ500, ambient+500 µatm CO2; Δ1000, ambient+1000 µatm CO2. nC–A=16/15, nC–Δ500=15/16, nC–Δ1000=16/16, nW–A=16/15, nW–Δ500=17/11, nW–Δ1000=13/15, for cold/warm assay temperature, respectively.

View this table:
Table 4.

LOmy fraction of P respiration in permeabilized heart fibers of European seabass

Effects of acute in vitro cooling on mitochondrial function in warm life-conditioned fish

Acute cooling led to significantly decreased P and PCII respiration rates in the Δ1000 group (LME, P<0.01, Fig. 1 and P<0.01, Fig. 3, respectively). All other analyzed parameters were not affected by acute cooling: PCI (Fig. 2), CIV (Fig. 4), LOmy (Fig. 5), and the relative contributions of CI, CII, CIV and LOmy to P (Table 3; CI fraction: 50.2±3.2–57.5±3.3%, Fig. S1; CII fraction: 41.7±3.2–49.7±3.1%; Figs S2, S3 and S4).

Effects of long-term thermal conditioning on mitochondrial function

Warm life conditioning led to decreased P and PCI respiration rates in the A and Δ1000 group, either at both assay temperatures (PCI in the A group, Fig. 2, LME, P<0.01) or only at the cold assay temperature (P in the A and Δ1000 group, Fig. 1, LME, P<0.001 and P<0.01, respectively; and PCI in the Δ1000 group, LME, cold assay temperature, P<0.05).

No significant effects of warm life conditioning were observed for PCII (Fig. 3) and CIV capacity (Fig. 4) or the relative contributions of CI and CII to P respiration (Table 3). CIV capacity was 1.5–2 times higher than P respiration (Fig. S3) and did not differ between thermal life conditions.

LOmy respiration rate was significantly decreased in warm life-conditioned fish compared with cold life-conditioned fish in the A group (Fig. 5; LME, P<0.01 and P<0.0001, both assay temperatures, respectively) and in the Δ500 group (LME, P<0.0001, warm assay temperature).

Thermal life condition and assay temperature had significant effects on the relative contribution of LOmy to P (LOmy fraction; Table 3). As PCO2 did not affect the LOmy fraction, PCO2 treatment data were pooled. These data indicated by a significantly higher LOmy fraction in the warm compared with the cold assay temperature that acute warming led to impaired mitochondria in the cold life-conditioned fish, whereas the LOmy fraction did not differ between the thermal life history groups when compared at the respective conditioning temperature (Table 4; LME, P<0.05). Conditioning to warmer temperatures led to a significantly decreased LOmy fraction (Table 3).

Mitochondria were well coupled in all treatments (RCROmy>4), with significantly higher RCROmy in warm life-conditioned fish than in cold life-conditioned fish (LME, 9.43±1.38 and 6.46±1.33, respectively, P<0.05). There were no significant effects of assay temperature, PCO2 or any interaction terms on RCROmy (Table 3).

Effects of acclimation to different PCO2 on mitochondrial function

Elevated PCO2 alone did not have significant effects on any of the studied complexes and processes of the electron transport chain (Table 3). However, we found synergistic effects with temperature which became visible as interaction effects with life condition or assay temperature only in the Δ500 or Δ1000 fish, as specified above.

DISCUSSION

Mitochondrial functional capacities were examined in seabass juveniles raised in six combinations of three PCO2 and two temperature treatments. The data provide evidence that heart mitochondria of juvenile seabass can be impaired by acute warming, as observed in increased LOmy respiration rates, for example. In contrast, warm life conditioning increased mitochondrial efficiency in comparison to that of cold life-conditioned fish, as seen through increased RCROmy.

Ocean acidification did not affect mitochondrial functioning in juvenile seabass as a single factor, as indicated by no significant effects of PCO2 alone on mitochondrial capacities. However, ocean acidification intensified the effects of acute or long-term warming. This was most prominent in the high acidification warm life condition treatment, e.g. P respiration rates were only significantly affected by acute temperature change in the W–Δ1000 fish, due to the decrease of PCII in this group. CI, CIV and RCROmy were not affected by PCO2. This observation reflects the findings of previous studies in polar fish, where thermal effects on mitochondrial capacity were much more prominent than those of ocean acidification (e.g. Leo et al., 2017; Strobel et al., 2013b). However, the reduced ability of CII to cope with acute temperature changes in the W–Δ1000 fish is in agreement with other studies which found that CII was inhibited by elevated PCO2 in mammals and fish (Simpson, 1967; Wanders et al., 1983; Strobel et al., 2013a). In juvenile seabass, CII was only inhibited by high PCO2 in warm life-conditioned fish facing an acute temperature decrease. Therefore, juvenile seabass mitochondria appear generally able to cope with the inhibiting effect of high PCO2 on CII. Other studies suggested that mitochondria could employ anaplerotic mechanisms, such as decarboxylation of aspartate and glutamate, feeding into the Krebs cycle via oxaloacetate and oxoglutarate and stimulating CI with additional NADH-related substrates to overcome inhibitory effects of high PCO2 of CII (Langenbuch and Pörtner, 2002; Strobel et al., 2013a). However, if environmental temperature or PCO2 concentration further increase, decreased P respiration rates due to reduced CII respiration could occur at the respective habitat (conditioning) temperature and not only under acute temperature change.

Acute warming impairs heart mitochondria of cold life-conditioned juvenile seabass by increasing LOmy respiration, significantly so in the C–A and C–Δ1000 groups, as seen in the increased LOmy fraction. Increased leak respiration rates (LOmy as well as LOmy fraction) indicate decreasing mitochondrial membrane integrity, translating into less ATP produced for the same amount of oxygen consumed. Increases in mitochondrial enzyme activity and respiration rates of mitochondrial complexes, as well as P respiration rates with acute temperature increase have been shown in other fish species, e.g. Antarctic nototheniids, European perch and Atlantic cod (Strobel et al., 2013a; Ekström et al., 2017; Leo et al., 2017). Iftikar and Hickey (2013) showed in hearts of New Zealand triple fin fishes that compromised mitochondria at acutely elevated temperature will ultimately lead to heart failure. Consequently, as acute warming of only 5°C impaired mitochondria of the cold life-conditioned fish, it appears likely that cold life-conditioned juvenile seabass can suffer from heart metabolic deficiencies, if acute temperature changes exceed 5°C. This reduced tolerance to acute temperature increase in the cold life-conditioned fish seems to contradict the fact that European seabass are generally highly tolerant to a wide range of temperatures (Dalla Via et al., 1998; Claireaux and Lagardère, 1999). It also seems to contradict the high critical thermal maximum (CTmax) of European seabass (28.12±0.09 to 32.50±0.04°C in Mauduit et al., 2016; and 31.3±0.3°C in Anttila et al., 2017). However, in seabass acclimated to 17°C, Anttila et al. (2017) found arrhythmia occurred at around 22°C, although CTmax was above 30°C. Furthermore, the Arrhenius breakpoint temperature for maximum heart rate, which is connected to the thermal optimum of growth and aerobic scope, was 19.3±0.3°C and the temperature with the highest maximal heart rate, a measure of the thermal limits of cardiac function, was 21.8±0.4°C in these fish (Anttila et al., 2017). These findings support our conclusion that cold life-conditioned juvenile sea bass might be less able to cope with large acute temperature changes than their warm life-conditioned siblings.

While acute warming impairs the performance of juvenile seabass heart mitochondria, the warm life-conditioned fish showed higher mitochondrial functionality, indicating that the chosen cold life-conditioning temperature is not the optimal temperature for Atlantic juvenile seabass. The thermal biology of D. labrax has been the topic of several studies, although mainly on Mediterranean populations (e.g. Marangos et al., 1986; Koumoundouros et al., 2001; Lanari et al., 2002; Person-Le Ruyet et al., 2004; Dülger et al., 2012; but see Russel et al., 1996; Ayala et al., 2003; Gourtay et al., 2018). However, in contrast to the Mediterranean populations, which are exposed to higher habitat temperatures (typical annual range 13–29°C; Person-Le Ruyet et al., 2004), the Atlantic population experiences temperatures lower than 15°C for most of the year and mainly within the range 6–18°C along the coast of France up to the North Sea (Russel et al., 1996). Our fish are the offspring of fish caught in the Bay of Biscay. In these latitudes, spawning, egg development and larval hatching take place at temperatures of 8–13°C (Jennings and Pawson, 1992) and later life stages experience mainly temperatures between 6 and 18°C (Russel et al., 1996). Therefore, the temperature range we used for incubating the larvae in the cold life condition was slightly above the natural temperature range of seabass larvae from the chosen distribution area. However, for juvenile incubation the cold life condition temperature range of 15–18°C was well within the natural temperature range during summer. The temperature of the warm life-conditioned juveniles was consequently above the temperature range of their natural habitat in the Bay of Brest. Our study thus provides evidence that the seabass from the chosen population are not yet fully adapted to lower temperatures, as the warm life-conditioned juveniles displayed much better mitochondrial functionality than the cold life-conditioned animals, reflecting their evolutionary origin in warmer waters.

As a consequence of warm life conditioning, LOmy and P respiration rates were both significantly decreased, while RCROmy was significantly increased, which was the case in all treatments. The increased RCROmy in warm life-conditioned fish in comparison to cold life-conditioned fish indicates that although mitochondrial capacity was decreased in warm life-conditioned fish (decreased P), mitochondrial efficiency was increased (decreased LOmy and increased RCROmy). This could translate into higher growth rates: Shama et al. (2014) found that lower P capacity resulted in optimized metabolic rate that could generate higher scope for growth in sticklebacks acclimated to warmer temperatures. Additionally, in brown trout, lower food intake and failure to grow were correlated to high LOmy respiration rate and lower mitochondrial coupling in liver and muscle mitochondria (Salin et al., 2016a). In other words, individuals with more efficient oxidative phosphorylation tend to grow better than those with less efficient mitochondria. Our study reflects these findings: the warm life-conditioned juveniles showed higher RCROmy and lower P respiration rates, while being significantly larger than the cold life-conditioned fish, even when compared at equal age in degree days.

In our study, CIV or cytochrome c oxidase (CCO) activity was not affected by thermal life condition. As terminal electron acceptor of the electron transport system, CCO is important in aerobic respiration and was found to be the controlling site of mitochondrial respiration and ATP synthesis (Villani and Attardi, 2001; Gnaiger, 2009, 2012; Kadenbach et al., 2010). CCO generally displays excess capacity, especially in heart tissue (Gnaiger et al., 1998). In our study, CIV had excess capacity 1.5- to 2-fold higher than P respiration rate in all treatments, which is within the scope typically found in fish (1.5–3.2, Hilton et al., 2010; 1.8–2.7, Iftikar et al., 2015; 1.9–2.6, Salin et al., 2016b). Therefore CIV is not limiting the capacity of juvenile seabass mitochondria.

Conclusion

Although we used specimens originating from a northern population of seabass for this study, the results altogether indicate that the mitochondrial metabolism still supports (and favors) temperatures as found in Mediterranean specimens. Consequently, juvenile seabass in the North Atlantic might benefit from increased temperatures. Within the limits of this study, we also observed a high capacity to cope with ocean acidification, although this was less pronounced under ocean acidification and warming. The results of this study indicate that juvenile European seabass will be able to survive in an acidifying and warming ocean; however, there are further bottlenecks that may constrict their survival in a future climate. Firstly, other life stages, especially egg and larval stages, might be more vulnerable to temperature changes and increased PCO2; and secondly, other important traits, such as behavior or reproductive capacity and phenology might be affected differently by ocean acidification and warming. Consequently, other traits and life stages shall be analyzed in further studies.

Acknowledgements

We thank Hanna Scheuffele for her support in measuring mitochondrial respiration and Karine Salin for her critical comments. We acknowledge the technicians and researchers from the Laboratory of Adaptation, and Nutrition of Fish for their support in animal welfare during the weekends.

FOOTNOTES

  • Competing interests

    The authors declare no competing or financial interests.

  • Author contributions

    Conceptualization: S.H., G.C., F.C.M.; Methodology: S.H., N.L., G.C., F.C.M.; Validation: S.H., F.C.M.; Formal analysis: S.H., G.C., F.C.M.; Investigation: S.H., L.C., N.L., G.C., F.C.M.; Resources: G.C., F.C.M.; Data curation: F.C.M.; Writing - original draft: S.H., F.C.M.; Writing - review & editing: S.H., L.C., G.C., F.C.M.; Visualization: S.H.; Supervision: G.C., F.C.M.; Project administration: F.C.M.; Funding acquisition: G.C., F.C.M.

  • Funding

    This work was supported by the German Science Foundation (Deutsche Forschungsgemeinschaft, DFG) and part of the FITNESS project (DFG grants MA 4271/3-1 to F.C.M. and PE 1157/8-1 to M. Peck, University of Hamburg, Germany).

  • Data availability

    Datasets of mitochondrial respiration and water conditions during rearing, as well as additional information on larval rearing are available online from PANGAEA (www.pangaea.de).

  • Supplementary information

    Supplementary information available online at http://jeb.biologists.org/lookup/doi/10.1242/jeb.213017.supplemental

  • Received August 23, 2019.
  • Accepted October 1, 2019.

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Research Article
Future ocean warming may prove beneficial for the northern population of European seabass, but ocean acidification will not
Sarah Howald, Louise Cominassi, Nicolas LeBayon, Guy Claireaux, Felix C. Mark
Journal of Experimental Biology 2019 222: jeb213017 doi: 10.1242/jeb.213017 Published 11 November 2019
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