# Multigenerational effects of temperature exposure on upper thermal limit and mitochondrial functioning in medaka (Oryzias latipes) brain

Julie Morla, Mélissa Richard, Rémy Lassus, Nicolas Pichaud, Martin Daufresne, Arnaud Sentis

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Julie Morla, Mélissa Richard, Rémy Lassus, Nicolas Pichaud, Martin Daufresne, et al. Multi-generational effects of temperature exposure on upper thermal limit and mitochondrial functioning in medaka (Oryzias latipes) brain. Journal of Thermal Biology, 2025, 130, pp.104161. 10.1016/j.jtherbio.2025.104161. hal-05227658

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# Multigenerational effects of temperature exposure on upper thermal limit and mitochondrial functioning in medaka (*Oryzias latipes*) brain

Julie Morla<sup>a, *</sup> ◆, Mélissa Richard<sup>a</sup>, Rémy Lassus<sup>a</sup>, Nicolas Pichaud<sup>b</sup>, Martin Daufresne<sup>a, 1</sup>, Arnaud Sentis<sup>a, 1</sup>

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## ARTICLE INFO

Keywords:

Multigenerational exposure

Ectotherm

Temperature

CT<sub>max</sub>

Mitochondrial respiration

## ABSTRACT

Determining the mechanisms underpinning the thermal limits of organisms is essential to anticipate the impact of climate change on their survival. As mitochondrial activity is key for cellular ATP production, we aimed at investigating the link between mitochondrial dysfunction and the upper thermal limit (CT<sub>max</sub>) at which neuro-muscular functions are lost. We reared medaka fish (*Oryzias latipes*) over generations at 20 °C (COLD) and 30 °C (WARM) during five years and measured their CT<sub>max</sub> and their mitochondrial oxygen consumption rates (OCRs) at 25 °C and at their CT<sub>max</sub>. We found that multigenerational exposure at elevated temperature increases the CT<sub>max</sub> by 13 %. We observed for all fish a general decrease in the OCRs and for the RCR of complex-I between 25 °C and their CT<sub>max</sub> suggesting a link between CT<sub>max</sub> and cerebral mitochondrial performance. Finally, we found an increase in the contribution of the mitochondria complex II to OCRs at elevated temperature suggesting that this complex underlie flexibility in mitochondrial functioning at high temperature. Our study highlights the link between whole organism thermal tolerance and mitochondrial dysfunction and that multigenerational exposure can modify mitochondrial and thermal performance.

## 1. Introduction

Temperature is a fundamental environmental factor that significantly affects metabolic and physiological processes, through the increased speed of biochemical reactions in ectotherms. Global warming and the expected increase in the frequency of extreme thermal events (IPCC, 2023) will likely impose significant constraints on ectothermic organisms. To predict species vulnerability to climate change, thermal vulnerability indices such as safety margins and warming tolerance have increased in popularity (Clusella-Trullas et al., 2021). Notably, the Critical Thermal maximum (CT<sub>max</sub>; Martínez et al., 2016; O'Donnell et al., 2020) is considered as an excellent proxy of upper thermal tolerance as it may trigger a conserved behavioural response across a diversity of species (Lutterschmidt and Hutchison, 1997).

Following the Arrhenius thermodynamic response, the metabolism of ectotherms increases with warming, accelerating cellular respiratory and oxygen demand (Arrhenius, 1889; Hilton et al., 2010; Sommer and

Pörtner, 2004; Iftikar and Hickey, 2013; Willis et al., 2021). Moreover, rapid warming in fish disrupts molecular mechanisms such as reaction rates, protein structure, and membrane fluidity, which can lead to mitochondrial dysfunction and organismal failure. Thus, mitochondrial functions have been put forward as a crucial determinant of organism thermal tolerance.

Mitochondria are central to cellular respiration as they use oxygen to transduce energetic substrates (derived from nutrients) into adenosine triphosphate (ATP), needed to maintain essential body functions. Mitochondrial respiration is assessed as the oxygen consumption that supports ATP synthesis through oxidative phosphorylation (OXPHOS) and compensates for proton leakage in the absence of ATP synthesis (LEAK). Enzyme and protein complexes of the respiratory chain that allow cellular respiration are profoundly influenced by temperature (see appendix Fig. S1; Abele et al., 1998; Clotfelter et al., 2013; Somero et al., 2017). Previous studies reported that the different mitochondrial complexes have different thermal sensitivities and may play a role in thermal

<footnote><sup>a</sup> INRAE, Aix-Marseille Univ, UMR RECOVER, 3275 Route Cézanne, 13182, Aix-en-Provence, France</footnote>

<footnote><sup>b</sup> Department of Chemistry and Biochemistry, Université de Moncton, Moncton, NB, E1 A 3E9, Canada</footnote>

<footnote>* Corresponding author. INRAE, UMR RECOVER, Aix-en-Provence, France.</footnote>

<footnote>E-mail addresses: juliemorla96@hotmail.com (J. Morla), melissa.richard@etu.univ-littoral.fr (M. Richard), remy.lassus@inrae.fr (R. Lassus), nicolas.pichaud@moncton.ca (N. Pichaud), martin.daufresne@inrae.fr (M. Daufresne), arnaud.sentis@inrae.fr (A. Sentis).</footnote>

<footnote n="1">¹ these authors contributed equally to this work.</footnote>
adaptation of ectotherms (Blier et al., 2014; Chung and Schulte, 2020). In particular, the first complex (complex I) of the respiratory chain has been suggested to be a site of thermal mitochondrial injury after a critical heat exposure and could be the main determinant of heat tolerance (El-Wadawi and Bowler, 1996; Jørgensen et al., 2021; Menail et al., 2022; Iftikar et al., 2010, 2014; Iftikar and Hickey, 2013). However, it has also been shown that this heat-induced failure in mitochondrial oxygen consumption at the level of complex I is not necessarily linked to the capacity of complex I to oxidize NADH (the main reducing equivalent providing electrons to ETS), but rather from upstream tricarboxylic acid (TCA) cycle dehydrogenases producing NADH (Blanchard et al., 2024; Jørgensen et al., 2021) or even from mitochondrial transport of substrates allowing this production (Chung and Schulte, 2020). Mitochondria can also utilize other substrates, such as succinate via complex II, providing metabolic flexibility. In fact, albeit complex I-induced oxygen consumption has been shown to decline at high temperatures, increased mitochondrial oxygen consumption when succinate was provided via complex II was also observed in several instances (Jørgensen et al., 2021; Menail et al., 2022; Blanchard et al., 2024; Hraoui et al., 2020; Chung et al., 2017; Willis et al., 2021; Hickey et al., 2024). Thus, the study of these complexes can help to explain the thermal limits of ectotherms, which holds important significance in the context of the current climate warming and species adaptation.

According to Chung and Schulte (2020) data from many species suggest that there may be a potential connection between mitochondrial failure and the breakdown of higher-level processes, including brain dysfunction, cardiac failure, and whole-organism thermal limitations (Chung and Schulte, 2020; Ekström et al., 2016; Fangue et al., 2009; Harada et al., 2019; Hraoui et al., 2020; Martínez et al., 2016). Mitochondria may specifically mediate heart failure under heat stress and play a role in $CT_{max}$ (Ekström et al., 2016; Hunter-Manseau et al., 2019; Iftikar et al., 2014; Iftikar and Hickey, 2013; Pichaud et al., 2017; Willis et al., 2021). However, mitochondrial dysfunction could also underlie neuronal failure. Indeed, mitochondrial oxygen consumption measured in the brain could indicate greater mitochondrial thermal sensitivity compared to cardiac tissue in *Fundulus heteroclitus* (Chung et al., 2017). Neuronal system dysfunction induced by temperature have been suggested to constrain whole-organism thermal limits as brain failure occurs at the same temperature as the whole organism in different fish species (Ern et al., 2015; Somero and DeVries, 1967; Friedlander et al., 1976; Andreassen et al., 2022; Jutfelt et al., 2019). Recent studies have thus tested the hypothesis that the loss of equilibrium during acute warming is due to direct thermal effects on neuronal function in the brain (Temperature-Dependent deterioration of Electrical Excitability, TDEE hypothesis) (Robertson, 2004; Vornanen, 2020). While this hypothesis may contribute to explain whole-organism thermal limits, recent studies concluded that it is probably not the only mechanisms leading to heat failure (Willis et al., 2021; Ern et al., 2023).

Indeed, the link between mitochondrial failure and failure of higher-level processes is not well established and remains a controversial topic issue (Bilyk and DeVries, 2011; Clark et al., 2013; Ern et al., 2015; Schulte, 2015; Ørsted et al., 2022). Early studies showed little evidence of a functional link between the mitochondrial function and the whole organism thermal tolerance because in vitro rates of mitochondrial oxygen consumption were maintained at temperatures above the $CT_{max}$ (Somero et al., 1996; Somero, 2002; Pörtner, 2002; Chung and Schulte, 2020). However, using measures of mitochondrial oxygen consumption in presence of excess ADP (OXPHOS rate) as well as a proxy for mitochondrial efficiency (the Respiratory Control Ratio – RCR which integrates OXPHOS and LEAK respiration rates), Chung and Schulte (2020) reported a correlation between acute thermal limits of ectotherms and the acute limits of mitochondrial function. It is important to note here that mitochondrial functions are generally determined using excess substrate concentrations, which is not necessarily representative of *in vivo* metabolite concentration and availability.

To provide more evidence of this relationship, Ern et al. (2023)

suggested to preferentially use thermal acclimation to manipulate in concert the thermal tolerance of the mitochondrial components and the whole animal, a concept that has been recently applied in Drosophila (Blanchard et al., 2024). Several studies suggest that $CT_{max}$ varies with the exposure duration and may thus be enhanced after a period of acclimation to elevated temperature (Angilletta, 2001; Bilyk and DeVries, 2011). Some studies found that thermal acclimation can lead to plastic modifications of the structure and function of mitochondria in ectotherms (Guderley and St-Pierre, 2002; Seebacher et al., 2010). Warm acclimation typically reduces mitochondrial metabolism, mainly due to lower respiration rates, decreased mitochondrial density, changes in membrane composition, and temperature-dependent variations in enzyme activity (Ekström et al., 2016; Schulte, 2015; Yan and Xie, 2015; Chung et al., 2017; Morla et al., 2024; Blanchard et al., 2024). These adjustments may result both from reversible phenotypic plasticity, notably observable in mitochondria, and from genetic adaptation over generations (Guderley and St-Pierre, 2002; Seebacher et al., 2010) and can have an impact on population performance. In a recent study, Rank et al., (2020), reported that population of montane leaf beetle can exhibit latitudinal differences in mitochondrial and nuclear genotypes that are associated with differences in performance traits, suggesting that mitochondrial thermal limits can affect whole organism performances. According to Pichaud et al. (2017), the complexity of the thermal acclimation process makes it unclear which aspects of mitochondrial metabolism are affected by temperature, although it is crucial to predict and anticipate the impacts of rapid global warming on ectotherms. To go further, an effective strategy to identify the mechanisms underlying thermal tolerance is to study thermal responses after multiple generations of exposures. Multiple generations can produce lineages that differ in their innate thermal tolerances or in their thermal acclimation capacity (Morgan et al., 2020).

In this study, our objective was to determine multigenerational temperature exposure effects on (1) the upper critical thermal maximum ($CT_{max}$) of the whole organism medaka fish (*Oryzias latipes*), (2) the oxygen consumption rates (OCRs) in brain mitochondria and (3) a potential link between a dysfunction of brain mitochondria and whole organism $CT_{max}$. We hypothesized that (1) brain mitochondria can limit the whole organism $CT_{max}$ and (2) that $CT_{max}$ can increase after a multigenerational exposure at elevated temperature resulting in mitochondrial changes (Angilletta, 2001; Bilyk and DeVries, 2011; Somero and DeVries, 1967; Ern et al., 2015). To test our hypotheses, we assessed the $CT_{max}$ of medaka fish maintained over several generations to cold (20 °C) and warm (30 °C) temperatures by exposing fish of each thermal origin to a progressive increase in temperature until it loses its locomotor function. In a second step, we measured the mitochondrial OCRs of fish from the two lines at 25 °C and at their mean $CT_{max}$ focusing on differences between the contributions of the different mitochondrial complexes to respiration.

## 2. Materials & methods

### 2.1. Biological model and rearing conditions

The Japanese medaka fish (*Oryzias latipes*) is a small, amphidromous freshwater fish from Southeast Asia. This species is eurythermic with a thermal range from zero to 40 °C (Dhillon and Fox, 2007; Hirshfield, 1980; Shima and Mitani, 2004; Kirchen and West, 1976), and a thermal optimum of 25 °C (Kirchen and West, 1976; Hirshfield, 1980; Leaf et al., 2011). It has a short generation time of 3 months and can reach sexual maturity in about 10 weeks (Hirshfield, 1980), making it a good biological model for multigenerational experiments. In its native range, the medaka generally lives in small water bodies such as rice paddies and with important temperature fluctuations (Hirshfield, 1980). In shallow environments, adult medaka can support temperature as high as 43 °C (personal observation).

Medaka used in this study were reared over multiple generations at
either 20 °C (COLD fish) or 30 °C (WARM fish) under controlled
experimental conditions in climate-controlled chambers. Two lineages
by temperatures (W1, W2 for WARM fish and C1, C2 for COLD fish) were
derived from an initial F₀ population of 320 Medakas (with an equal sex
ratio) belonging to the CAB strain supplied by Carolina Biological Sup-
ply Company (USA) and provided by AMAGEN® (France). The F₀ gen-
eration was initially maintained at 25 °C before being gradually
acclimatized to 20 or 30 °C (one degree every 2 days). To establish each
new generation, eggs clutches were collected during the optimal
fecundity period and incubated at the parental temperature. We kept the
new-born fish in climate-controlled chambers, housed in five replicates
aquariums until reaching maturity, with approximately 30 adult fish per
tank. They were fed "ad libitum" with TétraMin® food.

## 2.2. Whole organism thermal critical maximum tolerance (CT max)

After 5 years of multigenerational exposure to 20 °C and 30 °C, we exposed fish from each thermal origin to a thermal tolerance dynamic test to estimate their CT<sub>max</sub>. We used 32 fish from the two different thermal origin (W1 & W2, C1 & C2). The average generation time for 20 °C exposure (i.e. C1 and C2) was 283.3 days, and for 30 °C exposure (i.e. W1 and W2) was 181.6 days. Specifically, we used 16 fish maintained at 20 °C for six generations (F<sub>6</sub>): 8 adults females (4 from C1 & 4 from C2) (mean age in days ± se: 215.1 ± 6.7; i.e. 4.302<sup>a</sup> days) and 8 adults males (4 from C1 & 4 from C2) (215.1 ± 6.7). We also used 16 fish maintained at 30 °C for ten generation (F<sub>10</sub>): 8 adults females (4 from W1 & 4 from W2) (122.6 ± 3.8 days; i.e. 3.678<sup>a</sup> days) and 8 adults males (4 from W1 & 4 from W2) (122.6 ± 3.8 days). Ages of the studied fish were chosen accordingly to female reproductive optimum in terms of clutch size (i.e. number of eggs laid per day and per female; *Loisel, 2019*). Although fish had different ages in days, their physiological ages in degree-days were relatively close (i.e. 3.678<sup>a</sup> days for WARM fish and 4.302<sup>a</sup> days for COLD fish).

The WARM and COLD fish used in the experiments differ in gener-
ation numbers (10 and 6, respectively) due to the shorter generation
time at 30 °C compared to 20 °C. This difference arises from the
warming conditions applied in the study, as elevated temperatures
accelerate growth and lead to earlier sexual maturity, resulting in
asynchronous generations between the two acclimation temperatures
(Alberto-Payet et al., 2022; Atkinson, 1994, 1995). To align the fish
generations, the experiment would have needed to run for additional
years.

In addition, to test if the CT<sub>max</sub> could influence mitochondrial
respiration, 16 control fish (i.e. 4 males and 4 females from each thermal
origin) were also maintained under the same laboratory conditions
except that they were not exposed to the CT<sub>max</sub> test.

To estimate the CT<sub>max</sub>, two water baths (WB) were used (WB1 and WB2) (see appendix Fig. S2). Each WB had a water pump and a water heater (1500 Watt). Within each WB, two aquariums were placed at each side of the WB (hereafter referred as to top and bottom positions T & B; Fig. S2). To reduce bias, the location of the fish in the aquariums (i. e. T or B) and in the WB (i.e. WB1 & WB2) was switched each day. The water for the experimental tanks was filtered using two mechanical filters (25 µm then 5 µm) and a UV filter to eliminate microorganisms. Two cameras were installed at the front and at the top of the water-baths to observe fish behaviour remotely. Before the CT<sub>max</sub> experiment, each fish was weighed, measured, and photographed.

For each thermal origin and lineage, pairs (one male, one female) were placed in an aquarium in the WB at their rearing temperature (20 °C or 30 °C). Control fish were placed in a recovery aquarium. We then exposed organisms to rapid temperature changes through a constant heating rate (~1.5 °C every 10 min) that allows deep body temperature to follow ambient temperatures without a significant time lag (Hutchison, 1961, 1976). Loss of equilibrium (LOE) was used as an end-point for this experiment (Ziegeweid et al., 2008). Fish were then immediately returned to a tank at their initial temperature to allow their

total recovery. Temperature was recorded continuously during the
CTmax trial. Another potential cause of metabolic stress is the decreased
oxygen concentration in hot waters, coupled with an increase in meta-
bolic demand. To verify that the oxygen concentration was not limiting,
we measured dissolved oxygen concentration before and after each
temperature rise in each aquarium using a fiber optic oxygen probe
(HACH - HQ40d). For the recovery phase, fish were fed twice a day and
monitored individually to ensure their recovery (normal behaviour,
weight gain, absence of lesions) over the following 20 days (see ap-
pendix Table. s1).

## 2.3. Measurement of mitochondrial oxygen consumption rates (OCR s)

After 20 days of recovery following the CT<sub>max</sub> experiment, we measured their mitochondrial OCR at 25 °C (Topt) and at mean CT<sub>max</sub> for each thermal origin (WARM and COLD), defined as CT<sub>max</sub>, to determine the effects of transgenerational long-term temperature on mitochondrial functioning. We used a full factorial experimental design (see appendix Fig. S3A,B) with (I) fish from the two ancestral temperatures (COLD or WARM) tested at (II) three assay temperatures: 25 °C (thermal optimum), CT<sub>max</sub> of COLD fish and CT<sub>max</sub> of WARM fish (obtained from the previous experiment). The 25 °C assay temperature was chosen to compare fish from both lines at a common non-stressful temperature representing the thermal optimum.

Medakas fish were euthanized by cranial percussion and subsequently measured (mm), weighed (mg), and dissected to collect cerebral tissue samples. Dissections were performed within 5 min post-euthanasia at 4 °C. For each fish, the brain was transferred in 0.5M Respiration Medium (MIR) (Pesta and Gnaiger, 2012), gently mechanically permeabilized with fine forceps and placed on an orbital shaker at 4 °C for 5 min. The permeabilized tissue was split into three different parts and weighed before measurements. Mitochondrial oxygen consumption rates (OCRs) (pmol O₂·s⁻¹·mg⁻¹ of tissue) were measured using three Oxygraph-2k high-resolution respirometers (Oroboros Instruments, Innsbruck, Austria), each equipped with two independent measurement chambers. For each fish, we tested simultaneously three assay temperatures (25 °C, CT<sub>max</sub> COLD fish, CT<sub>max</sub> WARM fish) with three Oroboros Oxygraph. To avoid pseudo-replication issues, we switched the temperature of the three Oroboros every day. The oxygen electrodes were calibrated at air-saturated MiR05 and zero oxygen following sodium dithionite addition for each tested temperature. OCRs was determined using a sequential substrate/inhibitor protocol (see Appendix Fig. S4).

Specifically, the permeabilized cerebral tissue sample was added to the oxygraph chamber and we immediately injected 5 mM Pyruvate, 2 mM Malate and 5 mM glutamate. These substrates allowed to determine the LEAK value for proton gradient created by electron transfer in complex I of the electron transport system, which represents the oxygen consumption compensating for the proton leak when ATP synthase is not active (non-phosphorylating state). Then, the oxidative phosphorylation at the level of complex I (OXPHOS-CI) was initiated by adding a 2.5 mM saturating concentration of ADP to couple the proton gradient created by electron transfer in CI to phosphorylation into ATP. Additional injection of cytochrome C (10 µM) was used to verify the outer mitochondrial membrane integrity (CYTO-C). The oxidative phosphorylation at the level of complex I + II (OXPHOS-CI + II) was measured with the addition of 10 mM succinate. Electron transport system capacity (ETS-CI + II) (reflecting the maximal mitochondrial respiration induced in an uncoupled state) was achieved with repeated additions of carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone 0.5 µM (FCCP, 1 + 1 + 1 µL). This was followed by inhibition of complexes I and III through the sequential addition of 0.5 µL rotenone and 2.5 µM Antimycin A, respectively. The resulting non-mitochondrial oxygen consumption obtained was subtracted from the previous rates to obtain only mitochondrial specific OCRs. The COX-activity flux was then reached by
adding 2 mM of Ascorbate and a stepwise addition (5 µL + 5 µL + ...) of 0.5 mM TMPD. Finally, we added (≥100 mM) sodium azide to inhibit the complex IV and correct for auto-oxidation of TMPD. Albeit auto-oxidation of TMPD depends on PO₂ and should therefore be measured at the same levels of oxygen concentration than those used for COX, we rationalized that this should not have significantly altered the correction applied for TMPD auto-oxidation due to consistent methodology performed for the different groups. The average flux value was retrieved using Datlab software (Oroboros Instruments) to determined OCRs. We obtain 5 OCRs (LEAK-CI, OXPHOS-CI, OXPHOS-CI + II, ETS-CI + II and COX) each representing distinct processes within the mitochondrial respiratory chain (Fig. S1; Fig. S4).

## 2.4. Mitochondrial ratios

The Respiratory Control Ratio of complex I (RCR.CI) reflects the coupling between electron transport and oxidative phosphorylation from substrates oxidised at the CI level (i.e. complex I supporting substrates such as pyruvate, malate and glutamate allowing the generation of NADH by the TCA cycle). This was calculated as the ratio of ATP-synthesizing mitochondrial respiration (OXPHOS-CI) over mitochondrial respiration in the absence of ATP-synthesizing (LEAK-CI) for the complex I (with CI supporting substrates, Eq. (1)) (Brand and Nicholls, 2011; Estabrook, 1967). The RCR.CI represents a single general measure of mitochondrial function. In permeabilized fish brain (as used in this study) RCR.CI has been shown to fall into the range of 1.4–7.5 during control conditions depending on species and thermal origins (Chung et al., 2017; Devaux et al., 2019, 2023).

$$ \text{RCR.CI} = \frac{\text{OXPHOS-CI}}{\text{LEAK-CI}} \qquad \text{Eq. 1} $$

We also calculated the substrate relative contribution ratio (SCR, Eq. (2)) (i.e. see Jørgensen et al., 2021) for OXPHOS state using rates measured before and after injection of succinate:

$$ \text{SCR} = \frac{\text{OXPHOS-CI} + \text{II} - \text{CYTO-C}}{\text{CYTO-C}} \qquad \text{Eq. 2} $$

Finally, for each fish, we calculated ratios of OXPHOS-CI, LEAK-CI and OXPHOS-CI + II values measured at their CTmax divided by 25 °C assay temperature values (i.e. OCR CTmax/OCR 25 °C) to characterize the fold change in mitochondrial oxygen consumption at the CTmax relative to the oxygen consumption at the optimal temperature.

## 2.5. Statistical analyses

Statistical analyses were performed using R software version 4.3.1 (R Foundation for Statistical Computing Platform, Free Software Foundation's, Boston, USA). For the CTmax experiment, we used a linear mixed effect model (LMM) to test the effects of fish ancestral temperatures (i.e. 20 °C or 30 °C), fish lineage, fish sex and their interactions on the CTmax values. The aquarium position nested in the experimental day (i.e. the same information as WB) were included as random effects, to account for potential spatial and temporal effects. Non-significant terms were excluded from the final analysis.

For OCRs at the mitochondrial level, we tested the effects of fish ancestral temperatures, assay temperatures, treatment (CTmax test or control fish), fish sex, fish lineage, and their interactions for LEAK-CI, OXPHOS-CI, OXPHOS-CI + II, ETS-CI + II and COX activity as well as on the RCR.CI and SCR using LMMs. The fish identification number and the couple pair code nested in the Date experiment were included as random factors, to account for the repeated measurements (three measurements for each fish) and for potential temporal effects. Non-significant terms were excluded from the final analysis. For each model, when the interaction was significant between the two experimental conditions tested, we divided the datasets by the experimental condition using LMMs.

We then compared OCRs between the COLD fish at their assay temperate CTmax (C-38.9) and the WARM fish at their assay temperature CTmax (W-43.8) using LMMs to better understand if their mitochondria function similarly when exposed to their respective CTmax temperatures.

We finally illustrated distribution pattern of the individual CTmax for each fish for OXPHOS-CI, LEAK-CI and OXPHOS-CI + II as a function of the mitochondrial OCRs at CTmax assay temperature. We also compared distribution pattern for the OCR at CTmax divided by the OCR at 25 °C for OXPHOS-CI, LEAK-CI & OXPHOS-CI + II as a function of CTmax whole-organism values for each thermal origin and tested the differences in the Ratio (flux CTM/flux 25 °C) between the two fish thermal origins using LMMs.

The application conditions for the LMMs were evaluated. The significance of the fixed model terms was tested using deviance analysis (Anova function from the car package), and non-significant variables or interactions (p > 0.05) were excluded from the final model. Linear mixed models (LMMs) were implemented using the lme4 package (Bates et al., 2015).

# 3. Results

## 3.1. Whole organism thermal critical maximum (CTmax)

The CTmax values (Fig. 1) were significantly influenced by fish ancestral temperature (Chisq = 163, df = 1, p < 0.001 ***, Fig. 1; see appendix Tab. S2). Specifically, WARM fish had a CTmax (43.8 °C ± 0.27) significantly higher than COLD fish (38.9 °C ± 0.32) representing a 12.6 % increase.

<figure><img src="image_4.png" /><figcaption>Fig. 1. Boxplot with Mean (red dots) of the CTmax (°C) according to fish ancestral temperature (20 for COLD fish & 30 for WARM fish) (N = 32). *** represents highly significant differences between WARM and COLD fish (P < 0.001).</figcaption></figure>
## 3.2. Mitochondrial oxygen consumption rates (OCR's)

LEAK-CI (Fig. 2a; see appendix Tab. S3, S4) was not influenced by the interaction between ancestral temperatures and assay temperatures (Chisq = 4.9, df = 2, p = 0.09). However, LEAK-CI OCR was significantly influenced by assay temperature (Chisq = 16.5, df = 2, p < 0.001 ***) through increased OCR with temperatures. We found no significant effect of ancestral temperatures on LEAK-CI (Chisq = 1.05, df = 1, p = 0.31).

OXPHOS-CI (Fig. 2b; Tables S3 and S4) was significantly influenced by the interaction between ancestral temperature and assay temperature (Chisq = 9.94, df = 2, p = 0.006 **). When we split the dataset by experimental conditions, COLD and WARM fish OXPHOS-CI state was significantly affected by assay temperature. For COLD fish, we observed a decrease with increased assay temperature (Chisq = 12.39, df = 2, p < 0.005**) and for WARM fish we observe an increase between 25 °C and 38.9 °C followed by a decrease between 38.9 °C and 43.8 °C (Chisq = 12.16, df = 2, p < 0.005**). Moreover, we found a significant difference between COLD and WARM fish at the 38.9 °C assay temperature (Chisq

= 4.8, df = 1, p < 0.05*) and at 43.8 °C (Chisq = 4.21, df = 1, p < 0.05*) but no difference for 25 °C (Chisq = 0.3, df = 1, p = 0.59).

OXPHOS-CI + II was significantly influenced by the interaction between ancestral temperature and assay temperature (Chisq = 6.14, df = 2, p = 0.05*) (Fig. 2c; Tables S3 and S4). When we split the dataset by experimental conditions, OXPHOS-CI + CII was significantly influenced by assay temperature for COLD and WARM fish. For COLD fish, we observed a decrease with increased assay temperature (Chisq = 6, df = 2, p = 0.05*), and for WARM fish, an increase between 25 °C and 38.9 °C followed by a decrease between 38.9 °C and 43.8 °C (Chisq = 21.3, df = 2, p < 0.001**). We found no significant difference between COLD and WARM fish for each assay temperature (25 °C: Chisq = 1e-04, df = 2, p = 0.99; 38.9 °C: Chisq = 1.47, df = 2, p = 0.23; 43.8 °C: Chisq = 0.42, df = 2, p = 0.52).

ETS-CI + II (Tables S3 and S4; Fig. 2d) was not significantly influenced by the interaction between ancestral temperature and assay temperature (Chisq = 5.93, df = 2, p = 0.051) even if we are close to the interaction and we are seeing similar trends to the OXPHOS fluxes. ETS-CI + II significantly decreased with assay temperature (Chisq = 46.33,

<figure><img src="image_5.png" /><figcaption>**Fig. 2.** (a) LEAK-CI, (b) OXPHOS-CI, (c) OXPHOS-CI + II, (d) ETS-CI + II, (e) COX, (f) RCR.CI and (g) SCR mean (±SE) oxygen consumption rates (OCR's) in picomol. s⁻¹.mg⁻¹ according to the ancestral temperature (COLD 20 °C fish in blue or WARM 30 °C fish in red) and assay temperature (25 °C, dots; 38.9 °C, triangles; 43.8 °C, square). * represents significant differences between WARM and COLD fish (P < 0.05).</figcaption></figure>
df = 2, p = p < 0.001 ***). Finally, we found no significant effect of
ancestral temperature on ETS-CI + II (Chisq = 0.36, df = 1, p = 0.55).

The COX-activity (Tables S3 and S4; Fig. 2e) was not significantly influenced by the interaction between ancestral temperature and assay temperature (Chisq = 3.48, df = 2, p = 0.18). This OCR significantly increased with assay temperature (Chisq = 23.4, df = 2, p = p < 0.001 ***). We also found a significant effect of ancestral temperature (Chisq = 11.5, df = 1, p < 0.001 ***).

For all OCRs, we observed a non-significant general trend towards lower fluxes for WARM fish at 25 °C than for COLD fish at 25 °C.

3.3. *Mitochondrial ratio*

The RCR.CI was generally low for all conditions tested (Table S3; Fig. 2f). It is however important to note that RCR measured using complex I substrates in permeabilized fish brain is highly dependent of species and has been reported to be as low as 1.4 during control conditions (Devaux et al., 2019, 2023). The RCR.CI measured here is also only taking into account complex I substrates which might not be optimal oxidative substrates to fuel mitochondrial respiration in medaka fish. The RCR.CI was not significantly affected by the sex and the treatment, but we observed a significant effect of lineage (see appendix Tab. S5). We found the same tendency for the two lineages through lower fluxes for COLD fish than WARM fish. This tendency was

non-significant for C1 and W1 but (W2) and COLD (C2) displayed sig-
nificant differences at 25 °C (Chisq = 7.7, df = 2, p = 0.005**) and
38.9 °C (Chisq = 5.2, df = 2, p = 0.03*) (see appendix Fig. S5). RCR.CI
was not significantly influenced by the interaction between ancestral
temperature and assay temperature (Chisq = 0.95, df = 2, p = 0.62), but
was significantly influenced by assay temperature (Chisq = 94.2, df = 2,
p < 0.001 ***) through a decrease with increased assay temperature. We
found no significant effect of ancestral temperature on the RCR.CI
(Chisq = 1.76, df = 1, p = 0.18).

The statistical interaction between ancestral temperature and assay
temperature was non-significant (Chisq = 1.13, df = 2, p = 0.57) for the
SCR ratio reflecting the succinate relative contribution to respiration
(Tables S2 and S5; Fig. 2g). However, the SCR was significantly influ-
enced by assay temperature (Chisq = 15.5, df = 2, p < 0.001 ***)
through an increase with assay temperature. We found no significant
effect of ancestral temperature on SCR (Chisq = 0.11, df = 1, p = 0.73).

3.4. OCRs comparisons between warm and cold fish at their respective $CT_{max}$

To highlight mitochondrial OCRs differences at CTmax temperature between thermal lineages, we compared OCRs between COLD fish at CTmax 38.9 °C and WARM fish at CTmax 43.8 °C (Table S3; Fig. 3). We found that OCRs did not significantly differ between COLD and WARM

<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Boxplot with mean (red dots with black text) of OCR in picomol.s⁻¹.mg⁻¹ at *CTmax* assay temperature for (a) LEAK-CI, (b) OXPHOS-CI, (c) OXPHOS-CI + II, (d) ETS-CI + II, (e) COX activity, (f) RCR.CI, and (g) SCR for each thermal origin (COLD in blue or WARM in red). \* represents significant differences between WARM and COLD fish (P < 0.05).</figcaption></figure>
fish at their respective $\overline{CTmax}$ for LEAK-CI (Fig. 3a) (Chisq = 0.91, df = 1, p = 0.34), OXPHOS-CI (Fig. 3b) (Chisq = 0.79, df = 1, p = 0.37), OXPHOS-CI + II (Fig. 3c) (Chisq = 0.25, df = 1, p = 0.62), ETS-CI + II (Fig. 3d) (Chisq = 1.17, df = 1, p = 0.28), RCR.CI (Fig. 3f) (Chisq = 3.49, df = 1, p = 0.38) and for SCR despite slightly higher SCR for WARM fish (Fig. 3g) (Chisq = 3.5, df = 1, p = 0.06). However, we found that COX activity significantly differed between COLD and WARM fish at their $\overline{CTmax}$ (Fig. 3e) (Chisq = 4.38, df = 1, p = 0.03*).

### 3.5. Links between mitochondrial OCRs and whole organism $CT_{max}$

For a better understanding of the link between whole organism $CT_{max}$ and mitochondrial OCRs at $\overline{CTmax}$ assay temperature, we plotted the $\overline{CTmax}$ assay temperature according to the whole-organism real $CT_{max}$ values for each fish against individual mitochondrial OCRs in *picomol.s⁻¹.mg⁻¹* (see appendix Fig. S6) as well as for relative OCRs (i.e. OCR $\overline{CTmax}$/OCR 25 °C; see appendix Fig. S7) $\overline{CTmax}$. We did not observe a distribution pattern of OCR or ratio values flux at $\overline{CTmax}$ assay temperature for OXPHOS-CI, LEAK-CI and OXPHOS-CI + II (Fig. S6 and Fig. S7). We also illustrated the fold change in mitochondrial OCR at $\overline{CTmax}$ temperature relative to the OCR at the optimal temperature (OCR $\overline{CTmax}$/OCR 25 °C ratio) for LEAK CI, OXPHOS CI & OXPHOS CI + II according to the whole-organism real $CT_{max}$ values for each fish from both thermal origins (Fig. 4). We found no significant difference in the Ratio (flux $\overline{CTmax}$/flux 25 °C) between the two fish thermal origins for LEAK-CI (Chisq = 0.02, df = 1, p = 0.9), OXPHOS-CI (Chisq = 0.10, df = 1, p = 0.74) and OXPHOS-CI + II (Chisq = 1.12, df = 1, p = 0.29) (Fig. 4).

## 4. Discussion

The aim of this study was to investigate whether acclimation over several generations (multigenerational exposure) of freshwater Medaka fish to an elevated, non-stressful temperature (30 °C) can result in changes in whole-organism thermal limits ($CT_{max}$) and brain mitochondrial functions. Moreover, we aimed at determining whether there is a link between whole-organism $CT_{max}$ and brain mitochondrial performance at elevated temperature in the same fish. These questions are motivated by studies showing that higher thermal tolerance can be improved after a period of exposure to elevated temperature (Bilyk and DeVries, 2011; Angilletta, 2001) and by studies showing that

mitochondrial neural dysfunction can limit the upper thermal tolerance of the entire organism (Somero and DeVries, 1967; Ern et al., 2015).

### 4.1. Whole organism upper thermal tolerance ($CT_{max}$)

In line with studies suggesting that the $CT_{max}$ varies with the exposure duration (Podrabsky and Somero, 2006; Bilyk and DeVries, 2011; Angilletta, 2001), we found that thermal acclimation over several generations increases the maximum thermal tolerance of the whole organism. Fish from multigenerational exposure at 20 °C (COLD fish) had a $\overline{CTmax}$ of 38.9 °C, compared to 43.8 °C for those exposed to 30 °C (WARM fish) representing a 12.6 % increase (Fig. 1).

Our results thus highlight the influence of an acclimation period spanning several generations (long-term exposure) on the thermal tolerance of the whole organism. Similar results have already been observed for studies performing single-generation exposure. For example, Fangue et al. (2006) obtained significant differences in $CT_{max}$ for common killifish (*Fundulus heteroclitus*) after 21 days of acclimation. Killifish acclimated at 5 °C had a $CT_{max}$ of 31 °C, while fish acclimated at 33 °C had an average $CT_{max}$ around 41 °C (representing a 10 °C increase). Li et al. (2015) also demonstrated that the $CT_{max}$ of marine medaka (*Oryzias melastigma*) larvae increases significantly with acclimation temperatures. Fish were subjected to stepwise temperature change at a rate of 1 °C/h, increasing or decreasing from a starting temperature of 25 °C (the control) to reach six target temperatures (12, 13, 15, 20, 28 and 32 °C) with a range of maximum thermal tolerance variation from 39.9 °C at 13 °C-42.8 °C at 32 °C. In their study, they found a $CT_{max}$ of 40.53 °C at 20 °C and a $CT_{max}$ of 42.56 °C at 30 °C (Li et al., 2015). Since exposition took place over several generations in our case, it is even possible that this difference in thermal tolerance reflects adaptation to different thermal regimes considering that temperature might be a driving force for species adaptation. Genetic and genomic data would however be essential to evaluate the relative contribution of phenotypic plasticity (acclimation) versus genetic adaptation for the observed changes in $CT_{max}$.

### 4.2. Mitochondrial performances

The link between acute warming and mitochondrial function is already well established. Several studies have shown that acute exposure to elevated (non-stressful) temperatures leads to an increase in mitochondrial respiration (Abele et al., 2002; Bielski et al., 1983; Brand,

<figure><img src="image_7.png" /><figcaption>**Fig. 4.** Boxplot of the Ratio (OCR $\overline{CTmax}$/OCR 25 °C) for (a) LEAK-CI, (b) OXPHOS-CI and (c) OXPHOS CI + II, according to fish thermal origin (i.e. ancestral temperature of 20 °C for COLD fish in blue or 30 °C for WARM fish in red).</figcaption></figure>
2000; Ricquier and Bouillaud, 1998; Roussel and Voituron, 2020). However, this increase in mitochondrial respiration can also lead to higher levels of reactive oxygen species (ROS) production at temperatures close to the thermal limit that can fasten ectotherms senescence and mortality (Christen et al., 2018; Woodward, 2009). In this study, we observed a general increase in OCRs for OXPHOS-CI, OXPHOS-CI + II and LEAK-CI states between assay temperatures of 25 °C and 38 °C for WARM fish (38 °C is considered as non-stressful because lower than $\overline{CTmax}$ value for WARM fish), confirming that assay elevated temperature exposure leads to an increase in oxygen consumption (Fig. 2abcd).

There is evidence of thermal mitochondrial plasticity in ectotherms following acclimation (Guderley and St-Pierre, 2002; Seebacher et al., 2010; Morla et al., 2024). Our results indicate a trend towards lower fluxes for WARM fish at their optimal temperature (25 °C assay temperature). Similar results have been observed previously in other ectotherms (Chung et al., 2017; Ekström et al., 2016; Khan et al., 2014; Schulte, 2015; Yan and Xie, 2015) and on medaka lineage after two years of temperature exposure (Morla et al., 2024). This decline in mitochondrial respiration with acclimation would allow counteracting the increasing oxygen consumption inherent to faster reaction rates at higher temperature and associated cellular damages (Abele et al., 2002; Hochachka and Somero, 2002).

Interestingly, regarding LEAK-CI, mitochondrial fluxes increased between assay temperatures of 25 °C and $\overline{CTmax}$, regardless of the thermal origin of fish (Fig. 2a). Previous studies have reported mechanisms allowing to reduce the metabolic cost at elevated temperatures by decreasing mitochondrial efficiency in term of ATP production through increasing proton leakage (LEAK) and decreasing the proton gradient for ATP production (Brand, 2000; Ricquier and Bouillaud, 1998; Skulachev, 1998). In our study, RCR.CI values are in line with previous fish brain studies (isolated mitochondria or permeabilized brain tissue), which typically report low oxygen fluxes and RCR under control conditions with complex I substrates (Cambier et al., 2012; Bourdineaud et al., 2013; Chung et al., 2017; Devaux et al., 2019; Willis et al., 2021). In the context of acute thermal challenge, these fluxes are impaired, leading to even lower RCR indicative of mitochondrial dysfunction (which is also highly species-dependent, Chung et al., 2017; Willis et al., 2021), thus consistent with our RCR.CI values. Moreover, considering that almost no increased oxygen consumption was detected after addition of cytochrome c, which attests of mitochondrial outer membrane integrity (results not shown), the low values for the RCR.CI may reflect a specific impairment of complex I at high temperature. Alternatively, this indicates that substrates used were not optimal to stimulate oxygen consumption in our species (at least at high temperature) consistent with what has been demonstrated in other freshwater species such as mussels (Hraoui et al., 2020). Nevertheless, our results showing that LEAK flux exceeds OXPHOS flux at high temperatures can indicate a profound collapse in substrate oxidation capacity via Complex I, insufficient to counteract proton leakage (Brand and Nicholls, 2011). Alternatively, this OXPHOS impairment may also be associated with collapse of dehydrogenases of the TCA cycle or mitochondrial substrate transport (i.e. pyruvate by the mitochondrial pyruvate carrier), leading to decreased NADH production and thus declined complex I-supported respiration. In any case, this breakdown likely causes an imbalance between ATP supply and demand, leading to reduced ATP/O efficiency at high temperature (Roussel et al., 2023). While the mitochondrial membrane potential remains elevated during the leak state, amplifying proton loss at high temperatures, it drops when ADP is available, enabling some ATP production. However, the persistently high proton leak significantly constrain ATP output. These observations raise concerns about whether ATP production can adequately meet the brain's energy demands under thermal stress, and whether increased oxygen consumption might be required to offset the loss in efficiency. Overall, our results suggest mitochondrial dysfunction, particularly involving Complex I-linked substrates, which could impair the efficiency of ATP production

(Willis et al., 2021). However, direct measurements of ATP output are still necessary to confirm this.

There was no difference in OXPHOS and LEAK between WARM and COLD fish at $\overline{CTmax}$ temperature (Fig. 3abcdfg), suggesting a convergence towards a similar response of the mitochondrial OCRs at temperature reaching the $\overline{CTmax}$. We observed neither a significant difference in the Ratio (OCR $\overline{CTmax}$/OCR 25 °C) between the two thermal origins of the fish (Fig. 4) suggesting limits in thermal flexibility at stressful elevated temperature for mitochondrial respiration.

However, our results concerning the relative contribution of complex II (SCR) (Fig. 2g) show an increase between assay temperatures of 25 °C and $\overline{CTmax}$ only for WARM fish, suggesting that the relative contribution of complex II at higher stressful temperatures is more important for these fish. This result suggests that WARM fish may benefit from increased succinate oxidation by complex II when temperatures approach the organism's thermal limits. Indeed, complex II appears to have excess capacity to contribute to the electron transport system (ETS). In the presence of saturated succinate and reduced electron input via complex I (either from impairment in NADH production by TCA dehydrogenases or NADH oxidation by complex I), this pathway could compensate for oxygen consumption (OCR). This suggests that, rather than a plastic or genetic response, it is perhaps this inherent metabolic flexibility of complex II that allows partial maintenance of mitochondrial function under extreme thermal conditions, when complex I becomes limiting (Jørgensen et al., 2021; Menail et al., 2022; Blanchard et al., 2024; El-Wadawi and Bowler, 1996; Chung et al., 2018; Martínez et al., 2016; Iftikar et al., 2010).

### 4.3. Links between mitochondrial performances and whole organism $CT_{max}$

Due to the thermal sensitivity of mitochondria, studies have suggested a link between mitochondrial neuronal dysfunction under thermal pressure and the failure of whole-organism (Somero and DeVries, 1967; Pörtner, 2002; Iftikar and Hickey, 2013; Iftikar et al., 2014; Ern et al., 2015). In this study, we did not observe a distribution pattern of ratio or OCR values (Figs. S6 and S7) as a function of relative $CT_{max}$ for each fish measured for LEAK-CI, OXPHOS-CI, and OXPHOS-CI + II.

Our results suggest a decrease (not always significant) in the flux of OXPHOS-CI, OXPHOS-CI + II and ETS-CI + II between assay temperatures of 25 °C and $\overline{CTmax}$, for WARM and COLD fish (Fig. 2bcd). Chung et al. (2017) found similar results for mitochondrial OCRs of fish acclimatized to 15 °C, which decreased significantly as they approached $\overline{CTmax}$, suggesting a link between whole-organism $CT_{max}$ and mitochondrial performance at critical temperature. A more important decrease was observed for WARM fish, suggesting greater thermal sensitivity (Fig. 2bcd). However, the link between whole-organism $CT_{max}$ and mitochondrial performance at elevated temperatures needs to be clarified. Our results also suggest that the mitochondria of COLD fish continue to function (flux >0) beyond average whole organism mean heat tolerance. In line with this finding, recent studies suggests that mitochondria are more tolerant than the whole organism (Chung and Schulte, 2020; Jørgensen et al., 2021; Syromyatnikov et al., 2019). However, mitochondrial responses to extreme temperatures may also affect other functions not assessed in this study, which could potentially explain our findings, such as ATP and ROS production, as well as membrane fluidity to name a few (Biederman et al., 2019; Roussel et al., 2023).

To conclude, our results indicate that 5 years of multigenerational exposure to elevated temperature significantly increases the $CT_{max}$ of individuals. Although cerebral mitochondria appear to continue functioning beyond the organism's average heat tolerance, the measurement of oxygen flux does not necessarily guarantee that ATP production is occurring efficiently. Thus, our results suggest a link (even tenuous) between whole-organism $CT_{max}$ and cerebral mitochondrial
performance, particularly in terms of complex I- supported respiration, highlighting the importance of brain mitochondria in the context of thermal tolerance. Finally, we suggest that WARM fish may exhibit mitochondrial flexibility at the level of complex II at temperatures reaching the thermal limit of the whole organism. Our study highlights the complexity of the relationship between mitochondrial thermal sensitivity and whole-organism thermal tolerance, as well as the fact that acclimation can modify mitochondrial and thermal performance, which can have tremendous impacts on natural populations considering the increased average environmental temperature as well as the frequencies and durations of heatwaves brought forth by climate change. These capacities are nevertheless rarely taken into account in species distribution models (Clusella-Trullas et al., 2021). Finally, thermal tolerances can vary with life stage or body size, we thus need more research to determine if the limiting mechanisms during acute warming also differ across species, life stages, and thermal history (Dahlke et al., 2020; Pörtner, 2021).

### CRediT authorship contribution statement

**Julie Morla:** Writing – review & editing, Writing – original draft, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Mélissa Richard:** Methodology, Formal analysis, Data curation. **Rémy Lassus:** Methodology, Data curation. **Nicolas Pichaud:** Writing – review & editing, Methodology. **Martin Daufresne:** Writing – review & editing, Validation, Supervision, Project administration, Methodology, Funding acquisition, Conceptualization. **Arnaud Sentis:** Writing – review & editing, Validation, Supervision, Project administration, Methodology, Funding acquisition, Formal analysis, Conceptualization.

### Ethical approval

Experimental protocols were approved and performed in agreement with the ethical permits #42422-2023040411317548 v3. The above-mentioned project has been ethically assessed by the ethics committee for animal experimentation n°071.

### Data accessibility statement

The data that support the findings of this study are openly available on ZENODO at https://doi.org/10.5281/zenodo.14900919.

### Funding sources

This work was funded by the project EcoTeBo (ANR-19-CE02-0001-01) from the French National Research Agency (ANR) and by the project CLIMAQUA from the PACA (Provence-Alpes-Côte d'Azur) region and the INRAE AQUA department.

### Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

### Acknowledgments & Funding

This work was funded by the project EcoTeBo (ANR-19-CE02-0001-01) from the French National Research Agency (ANR) and by the project CLIMAQUA from the PACA (Provence-Alpes-Côte d'Azur) region and the INRAE AQUA department. We thank D. Roussel and Y. Voituron for Oroboros equipment.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.jtherbio.2025.104161.

### Data availability

No

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