# How short-term change in temperature or salinity affect cellular immune parameters of three-spined stickleback, Gasterosteus aculeatus?

Anne Bado-Nilles, Cleo Tebby, Alexandrine Pinet, Cyril Turiès, Jehan-Hervé Lignot, Jean-Marc Porcher

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Anne Bado-Nilles, Cleo Tebby, Alexandrine Pinet, Cyril Turiès, Jehan-Hervé Lignot, et al.. How short-term change in temperature or salinity affect cellular immune parameters of three-spined stickleback, Gasterosteus aculeatus?. Marine Environmental Research, 2025, 204, pp.106972. 10.1016/j.marenvres.2025.106972. ineris-04991258

HAL Id: ineris-04991258

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# How short-term change in temperature or salinity affect cellular immune parameters of three-spined stickleback, *Gasterosteus aculeatus*?

Anne Bado-Nilles<sup>a,*</sup>, Cleo Tebby<sup>b</sup>, Alexandrine Pinet<sup>a</sup>, Cyril Turiès<sup>a</sup>, Jehan-Hervé Lignot<sup>c</sup>, Jean-Marc Porcher<sup>a</sup>

## ARTICLE INFO

### Keywords:

*Field reference values*

Biomonitoring

Three-spined stickleback

Innate immunity

Temperature

Salinity

## ABSTRACT

Reference values for the non-specific immune response of stickleback have been developed to better understand the natural variability of the immunomarkers and to increase their relevance for the detection of environmental perturbations. However, under field conditions, temperature and salinity can vary from station to station and their influence on the reference ranges of the immunomarkers should therefore be quantified. To this end, adult sticklebacks were exposed either to different temperatures (from 12 to 18 °C) or to different salinities (from 0 to 30 g/L) for 21 days after 10 days of acclimatization. The results were then projected onto reference ranges to better determine the effect of temperature and salinity on the innate immune response. With the exception of leucocyte necrosis at higher temperature and respiratory burst at lower temperature, previously established reference ranges for immunomarkers of sticklebacks were suitable when variations in temperature and salinity were tested. Finally, this study argues for the possibility of using stickleback and its immune reference range in the field regardless of temperature and salinity, due to its relatively temperature and salinity independent innate immune response. Reference ranges for immunomarkers in stickleback could be a real added value to water quality diagnosis in biomonitoring programs in variable seasonal and geographical environmental contexts. Furthermore, these results confirm the rapid adaptability of sticklebacks to different variations in temperature and salinity without affecting their immunological parameters.

## 1. Introduction

Characterising reference values for biomarkers is helpful to account for natural variation in environmental diagnostics and to improve data interpretation (Burgeot et al., 2010; Coulaud et al., 2011; Marchand et al., 2019, 2020). In this way, the use of reference ranges seems to be a better way to understand the biological effects of contaminants in biomonitoring studies. To determine these reference ranges, a separate assessment of each confounding factor should be performed before being integrated into a larger model of biomarker responses (Coulaud et al., 2011; Hanson, 2011; Krell et al., 2011; Maltby et al., 2002; Marchand et al., 2019).

Because the immune response provides valuable information in the context of biomonitoring (Bado-Nilles et al. 2015a, 2015b; Monserrat et al., 2007; Rice et al., 1996; Zelikoff et al., 2002), reference ranges

have been developed for the three-spined stickleback cellular immune response including three confounding factors, season, sex and body size (Marchand et al., 2019). These reference ranges were then successfully tested within biomonitoring programmes using multiple uncontaminated and contaminated sites (Marchand et al. 2020, 2021) to include natural variability and to provide first hand assessment of whether this tool could be regarded as sensitive to known disturbances. The immunomarkers which were selected involved various activities of the phagocytosis function, which is central to the cellular immune process in fish (Biller and Takahashi, 2018; Bols et al., 2001; Neumann et al., 2001) and which has an interface between innate and adaptive responses (Biller and Takahashi, 2018). Furthermore, each part of the phagocytic mechanism, from phagocyte recruitment to pathogen destruction, has been described as an attractive immunomarker of field pollution in fish (Bado-Nilles et al. 2014a, 2015b; Betoulle et al., 2000; Bols et al., 2001;

<footnote><sup>a</sup> Institut national de l'environnement industriel et des risques, Université de Reims Champagne-Ardenne, Université Le Havre Normandie, Normandie Univ, UMR-1 02 SEBIO, 60550, Verneuil-en-Halatte, France</footnote>

<footnote><sup>b</sup> INERIS, Unit Experimental Toxicology and Modelling (TEAM), Parc Technologique Alata, BP 2, 60550 Verneuil-en-Halatte, France</footnote>

<footnote><sup>c</sup> MARBEC (MARine Biodiversity, Exploitation and Conservation), Univ Montpellier, CNRS, Ifremer, IRD, 34000, Montpellier, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: Anne.Bado-Nilles@ineris.fr (A. Bado-Nilles).</footnote>
Köhler, 1991; Köhler et al., 2002; Reynaud and Deschaux, 2006). Cellular mortality (apoptotic and necrotic cell death) is currently associated with respiratory burst and lysosomal modifications (Guicciardi et al., 2004; Holtzman, 1989; Kurz et al., 2008; Risso-de Faverney et al., 2001), which occur in phagocytes to degrade internalized pathogens. Similarly, a change in leucocyte distribution due to cellular mortality could also affect the phagocytic response (Scharsack et al., 2016). Nevertheless, to increase the robustness of the reference ranges developed by Marchand et al. (2019), external factors affecting the immune response should be included in the models. Among these, temperature, which is strongly related to seasonality, seems to be quite relevant (Bowden et al., 2007). Furthermore, due to the ability of stickleback to live in both marine and freshwater habitats (Wootton, 1984), the salinity is also quite relevant.

Although the acquired immune response appears to be more affected due to its slower reactivity (Ellis, 2001; Magnadóttir, 2006), temperature changes can alter innate parameters, particularly phagocytic activity (Magnadóttir, 2006). For example, Alcorn et al. (2002) showed a higher percentage of phagocytic kidney macrophages in sockeye salmon (*Oncorhynchus nerka*) reared at 8 °C compared to fish at 12 °C. In the same way, in two cyprinids (*Tinca tinca*, *Cyprinus carpio*), phagocytosis and respiratory burst activity are higher at 12 °C than at higher temperatures (>20 °C) (Collazos et al., 1994; Le Morvan et al., 1997), which maximises pathogen destruction. Using different parameters of the respiratory burst response (peak or integral values not adjusted or adjusted for phagocyte count), Buchtíková et al. (2011) also found a trend towards a negative relationship between respiratory burst activity and water temperature in common carp (*Cyprinus carpio*). Conversely, in channel catfish (*Ictalurus punctatus*), a reduction in phagocytic activity and respiratory burst adaptability is observed at 10 °C compared to a higher temperature (27 °C) (Ainsworth et al., 1991). In rainbow trout (*Oncorhynchus mykiss*), a reduction in respiratory burst and phagocytic activity is observed at lower temperature (5 °C) than at higher temperature (20 °C) (Nikoskelainen et al., 2004). At higher temperatures, an increased degradation of endocytosed/phagocytosed ligands by lysosomal activity (Larsen et al., 2018), especially due to the highest enzymatic activity, as shown with cod lysosomal α-mannosidase (Sørensen et al. 1998, 2001), could explain the continuity of phagocytosis activity. In sticklebacks, the immune response appears to be impaired at high temperature, from 18 to 24 °C, compared to relatively low temperatures (around 13 °C), mainly due to a marked decrease in respiratory burst activity (Dittmar et al., 2014). At a higher temperature (28 °C), attributed to the stress response, Dittmar et al. (2014) observed spleen enlargement due to increased immune cell turnover and respiratory burst increase. Similarly, when sticklebacks are stimulated with zymosan, respiratory burst is increased in stickleback maintained at 13 and 18 °C, but not at 24 °C (Scharsack et al., 2016).

Stickleback must also be able to rapidly adapt their physiological response to osmoregulatory contrasts due to opposing physiological challenges in salt or fresh water. This adaptative capacities of stickleback to saline variation explain the attractiveness of stickleback for analysis of evolutionary processes (Reid et al., 2021). Thus, many authors have attempted to determine the plastic physiological response of this species to salinity change. For example, Taugbøl et al. (2022) observed that an abrupt salinity transition affects the genomic response of stickleback, likely altering proteins and/or cell functions. Furthermore, Wang et al. (2014) and Gibbons et al. (2017) showed a different genomic response during salinity adaptation of native freshwater and anadromous saltwater stickleback, which were not only classified as salt-responsive genes, but also as genes involved in immune responses. In medaka (*Oryzias melastigma*), an immediate response in the gills is also detected, mainly in the adjustment of ion balance, immune and signal transduction expressed genes (Liang et al., 2021). In stickleback gills, proteomic responses confirmed that the maintenance of branchial osmoregulation during salinity changes is not only mediated by the regulation of osmoregulatory proteins, but also by proteins involved in

immunity (Li and Kültz, 2020). Therefore, salinity alters the immune response of fish. For example, transfer from freshwater to saltwater induces an increase in total white blood cell count, plasma lysozyme and head kidney macrophage respiratory burst activity in rainbow trout (*Oncorhynchus mykiss*) (Taylor et al., 2007) and of head kidney phagocytic activity in brown trout (*Salmo trutta*) (Marc et al., 1995). Different results are obtained in Nile tilapia (Oreochromis niloticus), with increased phagocytosis when salinity decreases and no effect when salinity increases (Choi et al., 2013). Nevertheless, after prolonged exposure to various salinity in catfish (*Pangasianodon hypophthalmus*), the abundance of proteins involved in the regulation of the phagocytic process is increased, such as formation of the phagocytic cup, maturation and acidification of the phagosome, activation of respiratory burst, production of iNOS and phagosome localisation (Schmitz et al., 2017). As previously described (Makrinos and Bowden, 2016), all these results show that salinity exposure tends to increase innate immune cells and, due to resource allocation, immune adaptive responses are often compromised or even suppressed if the stress is prolonged. These impairments in the immune response could clearly affect the response of fish to pathogens. For example, even though salinity fluctuations increase immune cell activity and proliferation, a delayed adaptive immune response is observed in the broadnose pipefish (*Syngnathus typhle*) during Vibrio infection, probably due to the investment of energy in osmoregulation processes that fail the immune response (Birrer et al., 2012). In addition, an hypoosmotic stress has suppressive immunoregulatory effects in *Scatophagus argus* stimulated by *Aeromonas hydrophila* infection (Lu et al., 2022).

The three-spined stickleback, *Gasterosteus aculeatus*, is an euryhaline fish species commonly used in ecotoxicological studies to assessed pollutant effects in vivo (Bado-Nilles et al., 2013; Jolly et al., 2009), and in situ (Bado-Nilles et al., 2015; Le Guernic et al., 2016; Sanchez et al., 2008b). Furthermore, stickleback has a wide repartition area, in temperate zones of northern hemisphere, and can live in highly disturbed streams (Ernst et al., 1991; Svecevičius, 2006). The ability of sticklebacks to adapt to a novel environment may be explained by their adaptability, which is certainly due to rapid changes in relevant phenotypes (Schulte et al., 2011) associated with the plasticity of gene expression (Morris et al., 2014) and/or with epigenetic modifications (Metzger and Schulte, 2017). These advantages make it a good sentinel species, interesting for environmental monitoring studies. Furthermore, seasonal variations of some biomarkers have already been studied in this species, including DNA integrity (Santos et al., 2013), biotransformation enzymes (Sanchez et al., 2008a), oxidative stress parameters (Sanchez et al., 2008a), neurotoxicity (Sanchez et al., 2008a), endocrine disruption markers (Sanchez et al., 2008a) and immune response (Marchand et al., 2019).

Characterising the remaining natural sources of biomarker variability could allow to discriminate responses induced by pollutant exposure and background noise. To correctly interpreted in situ assays for chemical water quality assessment, the natural immune plasticity induced by environmental confounding factors, such as temperature and salinity, should be improved. In this context, the stickleback is a relevant sentinel species for interpreting in situ observations in different environmental contexts, from freshwater to saltwater. For this reason, the Ineris Institute had developed active biomonitoring using standardized sticklebacks from its experimental nursery. Thus, the present work aims to determine how temperature (from 12 to 18 °C) and salinity (from 0 to 30 g/L) modified the immune responses of sticklebacks by using the reference ranges developed by Marchand et al. (2019) in different environmental contexts in time and space. As demonstrated by Allen and Wootton (1982), the temperature range in their natural habitat corresponded from 3 to 19 °C. In this way, we used temperature from 12 °C, which corresponds to minimal current temperature in field during caging (https://www.phresques.fr/), to 18 °C. Due to their plastic physiological response to salinity changes, the range of salinities was consistent with current salinities recorded in freshwater ecosystems and estuarine
conditions (Slaby et al., 2024). In addition, the selected salinities correspond to different physiological conditions, i.e. hyper-osmotic (10 g/L), iso-osmotic (15 g/L), hypo-osmotic (30 g/L).

## 2. Material and method

Each experiment was performed in accordance with the European Directive 2010/63/UE on the protection of animals used for scientific purposes at the INERIS facilities (registration number E60-769-02, APAFIS file n°16291).

### 2.1. Three-spined stickleback and exposure conditions

For the present work, all stickleback were obtained from a well-characterised population of the French National Institute for Industrial Environment and Risks (INERIS) (Verneuil-en-Halatte, Oise, France), maintained in an open-air tank with natural vegetation and macro-invertebrate communities. The stickleback origin is planned to be used continuously for monitoring purposes in the field and in the lab (Cant et al., 2022; Mit et al., 2023). Sticklebacks used were young of the year 2018. In October 2018, after monitoring of the health of the stickleback and confirming their disease-free status (VETOFISH, 13 220 Châteauneuf-les-Martigues, France), sticklebacks were transferred for acclimatization to 300 L laboratory opaque tanks with a continuous freshwater circulation system (16.5 ± 0.3 °C, 7.8 ± 0.1 pH, 284 ± 15 µS/cm, 100 ± 0.7% O², circadian cycle, pathogen free conditions). One week before the start of each experiment, the sex of the sticklebacks was determined using a discriminant model based on head morphology (de Kermoysan et al., 2013). The animals were fed, before and during experiment, daily with bloodworms (3% of body weight/day; Ocean Nutrition, Belgium). For each experiment, the presence or absence of acclimatization was determined according to the active biomonitoring protocol used by the Institute.

In February 2019, sticklebacks (n = 260, 47.7 ± 5.2 mm, 1.2 ± 0.4 g) were directly transferred to the different temperature for 21 days. Sticklebacks were randomly distributed in four 100 L tanks (7.8 ± 0.1 pH, 283 ± 17 µS/cm, 101 ± 0.7% O², circadian cycle) corresponding to the four tested temperatures (12.1 ± 0.1 °C, 13.7 ± 0.1 °C, 16.3 ± 0.1 °C, 18.1 ± 0.1 °C). As stickleback were kept at 16 °C during their growth in the laboratory, this temperature was defined as the control. Temperatures were maintained throughout the experiment using a TECO TK 500 Chiller Group. Each week (D0, D7, D14, D21), 20 sticklebacks per condition were sampled.

During exposure to different salinities, carried out in October 2019, stickleback (n = 160, 55.5 ± 3.7 mm, 2.6 ± 0.6 g) were acclimated for 10 days in four 100 L tank (16.5 ± 0.4 °C, 8.1 ± 0.1 pH, circadian cycle), corresponding to the four tested salinity, by gradually increasing the salinity at the rate of 1/10 of the final salinity to reach the desired concentrations (0 g/L, 10 g/L, 15 g/L, 30 g/L). At the end of the acclimatization period (D0), 20 sticklebacks per condition were sampled to measure the effect of acclimatization on the organisms. The remaining sticklebacks were then kept under exposure conditions for 21 days, with weekly water changes before sacrifice (D21).

For each experiment, 20 sticklebacks per condition and date were anaesthetised by balneation (tricaine methanesulphonate, 100 mg/L, Sigma) before cervical dislocation, then sticklebacks were measured, weighed and sexed. The spleen was removed to measure the immune response. During exposure to the different salinities only, an average of 5 µL of blood was collected per stickleback to determine blood osmolality using an Osmomat 030 osmometer (GONOTEC, Germany).

### 2.2. Leucocyte isolation and innate immune response analysis

The spleen filters plasma, traps blood-borne substances, and enriches the blood with new immune cells (Press and Evensen, 1999), and is therefore chosen to assess immunological properties. A splenic leucocyte

suspension was obtained by gently pressing the spleen through sterilised nylon mesh (40 µm, Sigma) with 5 mL Leibovitz 15 (L15) medium (Sigma) containing heparin-lithium (100 mg/L, Sigma), penicillin (500 mg/L, Sigma) and streptomycin (500 mg/L, Sigma). To eliminate any bias due to stress conditions of the stickleback prior to sacrifice, samples were stored at 4 °C for 12 h (Bado-Nilles et al., 2014b). A Malassez haemocytometer was then used to adjust the leucocyte concentration in the samples to 10⁶ cells/ml in L15 medium. Analyses were performed by flow cytometry on whole leucocytes using a CyAnTM ADP flow cytometer (Beckman Coulter). A total of 10 000 events per sample were analyzed after cell excitation with a 488 nm argon laser.

The percentages of each leucocyte subpopulation (lymphocytes and granulocyte-macrophages) were determined by measuring size (forward scatter, FSC) and complexity (size scatter, SSC) (Bado-Nilles et al., 2014). For ease of reading, only the granulocyte-macrophage percentages of total leucocyte populations are presented in figures and tables.

Double labelling with Yo-PRO®-1 (Invitrogen, final concentration: 3.14 mg/L) and propidium iodide (Invitrogen, final concentration: 5.01 mg/L) probes was used to assess the percentage of cell death. After 10 min incubation on ice in the dark, cellular fluorescence parameters were measured. Apoptotic and necrotic cells express green (FL1) and red (FL3) fluorescence respectively (Bado-Nilles et al., 2014).

Measurement of the leucocyte respiratory burst was performed using a modification of the technique of Chilmonczyk and Monge (1999) technique, adapted to stickleback. The determination of reactive oxygen species (ROS) production in unstimulated cells depends on the incorporation of 2',7'-dichlorodihydrofluorescein diacetate acetyl ester (H₂DCF-DA, Invitrogen, final concentration: 29.30 mg/L), a stable non-fluorescent molecule that is hydrolysed by cytosolic enzymes to dichlorodihydrofluorescein diacetate (DCFH). When leucocytes are stimulated with phorbol 12-myristate 13-acetate (PMA, Invitrogen, final concentration: 9.25 mg/L), an inducer of the respiratory burst (Ambrozova et al., 2011; Chadzinska et al., 2012) by direct activation of protein kinase C (Jin et al., 2021), H₂DCF-DA is hydrolysed by H₂O₂. Finally, the resulting DCFH is oxidised to the fluorescent dichlorofluorescein (DCF) to allow quantification by flow cytometry of unstimulated and stimulated cells in FL1 after 30 min incubation at room temperature. The respiratory burst index was determined as the ratio of the fluorescence of PMA-stimulated cells (H2DCF-DA plus PMA) to that of unstimulated cells (H2DCF-DA).

Phagocytic activity (percentage of leucocytes that have engulfed microspheres; fluorescence of at least three beads) was assessed after 1 h incubation in the dark at room temperature of a splenocyte suspension with fluorescent microspheres at a concentration of 2.7 x 10⁷ particles/mL (Fluorospheres® carboxylate-modified microspheres, diameter 1 µm, Invitrogen) (Gagnaire et al., 2014).

### 2.3. Blood osmotic pressures

Blood osmotic pressures were determined from 160 anaesthetised stickleback (20 sticklebacks per condition) previously maintained at different salinities. Blood samples were taken immediately from the heart of stickleback immediately after cervical dislocation, using heparinized insulin syringes. An average of 10 µl of blood was collected per stickleback. Blood osmolality (in mOsm.kg⁻¹) was then measured directly using a micro-osmometer (model 3300, Advanced Instruments, Needham Heights, MA, USA).

### 2.4. Statistical analysis

Statistical analysis was performed using R v.2.14.1 software (R Core Team, 2014). Normality was tested using Shapiro's test and homogeneity of variance using Levene's test (p ≤ 0.05). If the normality assumption was not met, data were log-transformed if this improved normality.

The reference ranges used are based on those developed by
Marchand et al. (2019). These reference ranges, based on ANCOVA results, has been successfully applied to laboratory, semi-field and field studies (Marchand et al. 2019, 2021). The reference ranges model biomarker levels based on linear combinations of stickleback length, sex and month.

Since the model developed previously was based on seasonal effects, in this work the temperature data were projected to February and the salinity data to October, in accordance with the season in which the experiments were carried out. Based on the previously developed statistical models, the evaluation of biomarker levels in case of temperature or salinity variation was based on two different statistical analyses. Firstly, for each sampling period, the differences between the conditions were tested either by ANOVA followed by a t-test with Holm's adjustment for multiple comparisons (parametric data, $p \le 0.05$) or by Kruskal-Wallis test followed by a Wilcoxon test with Holm's adjustment for multiple comparisons (non-parametric data, $p \le 0.05$). When the assumption of homoscedasticity was met across the 2 or 3 timepoints a two-way analysis of variance was performed on the data or log-transformed data in order to determine whether the pattern of immune responses relative to the various temperature or salinity conditions varied over time. Secondly, to determine whether temperature or salinity data fell within the February or October reference ranges, respectively, the percentage of data outside the reference ranges was calculated for each condition. Statistical significance of the number of data points outside reference range was evaluated using a unilateral binomial test compared to the expected 5%. Unilateral binomial tests ($p \le 0.05$) were performed at each sampling period to determine whether there was a significant number of out-of-range points of data collected

with various temperatures compared to the February reference range and of data collected with various salinity levels compared to the October reference range.

## 3. Results

### 3.1. Effects of temperature on immune parameters

The immune responses to the various temperature levels were not dependent on the timepoint for percentages of granulocytes-macrophages (log-transformed) and necrotic cells (log-transformed), and for phagocytosis efficiency. For these immune responses, the differences between temperature conditions that are observed are expected to be congruent between timepoints. A difference in profile was observed for percentage of apoptic cells in particular at 14 °C (log-transformed, 2-way anova, F-test of date:temperature interaction, $p < 0.05$). The strong heteroscedasticity in respiratory burst (including in log-transformed data) between timepoints and conditions prevented carrying out this statistical comparison.

After seven days of exposure to the various temperatures, only phagocytosis efficiency was modified. In fact, this parameter was significantly higher at 18 °C than at 12 °C and 14 °C, and only levels subsequent to exposure at 12 °C were lower than the control (16 °C) (Fig. 1). After 14 days of exposure, there was no difference in phagocytosis efficiency. This lack of effect is confirmed after 21 days of exposure by projection of the data within the February reference range (Fig. 2) with no significant percentage of error (0% at 12 °C, 5% from 14 °C to 18 °C, Table 1).

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Immune responses of stickleback in term of leucocyte distribution (granulocytes-macrophages, A), cellular mortality (apoptosis and necrotic cell (B et C, %)), respiratory burst (MFI, D) and phagocytosis activity (E) after seven (T7), 14 (T14) and 21-days (T21) of exposure to dissimilar water temperature (from 12 to 18 °C). ANOVA followed by a t-test with Holm's adjustment for multiple comparisons (parametric data, $p \le 0.05$) or Kruskal-Wallis test followed by a Wilcoxon test with Holm's adjustment for multiple comparisons (non-parametric data, $p \le 0.05$) were used to test statistical differences between temperature at each sampling date. Boxplots represent the values of 20 fish (n) including male (red) and female (green). For each parameter and date, difference between conditions was indicated by different letters ($p < 0.05$). (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Projection of temperature data after 21 days of exposure on the February reference ranges. For apoptotic cells (A), granulocyte-macrophage percentages (C), respiratory burst index (D) and phagocytosis activity (E), the lines represent the values predicted by the model and the prediction interval at 95 %. For necrotic cells (B), the full lines represent prediction intervals for female and the dotted lines represent prediction intervals for male. With MFI = Mean Fluorescent Intensity.</figcaption></figure>

Conversely, a greater effect on the other immune responses was observed after 14 days of exposure at different temperatures. The percentage of granulocytes-macrophages and the percentage of necrotic cells showed a different response only between 12 °C and 18 °C. Similarly, exposure at 12 °C could induce a reduction in the percentage of apoptotic cells compared to any other temperature exposure. The respiratory burst was clearly the more affected parameter, with a higher value at 12 °C compared to 14 °C and to 16 °C and 18 °C, the last two not being significantly different (Fig. 1).

After 21 days of exposure, each immune value in control temperature conditions (16 °C) was within the February reference range (Fig. 2), with no data outside the February reference range for apoptotic cells and granulocyte-macrophage percentage, and 5%, 10% and 15% outside the February reference ranges for phagocytic activity, respiratory burst

index and necrotic cells, respectively (Table 1). Furthermore, the difference between exposure conditions is somewhat less pronounced after 21 days of exposure. The response profile of granulocyte-macrophage percentages was similar at 14 and 21 days of exposure, with a significant reduction in granulocyte-macrophage percentages at 18 °C compared to 12 and 14 °C (Fig. 1). However, this discrepancy was not confirmed by the February reference ranges, as the 10 % of data outside the reference range at 18 °C were not statistically significant (Fig. 2, Table 1). In addition, no data were outside the reference range from 12 °C to 16 °C. After 21 days of maintenance at different temperatures, the percentage of splenic apoptotic leucocytes was different at 12 °C compared to 14 °C and 18 °C, but not compared to the control (16 °C). In addition, the proportions of data outside the reference range were not significant, with 0% error percentage from 12 °C to 16 °C and 5% at
<caption>**Table 1**</caption>

Percentage of data outside the February reference ranges for temperature (12 °C, 14 °C, 16 °C, 18 °C) and the October reference ranges for salinity (0 g/L, 10 g/L, 15 g/L, 30 g/L) condition. Significant deviation from the 5 % assumption error percentage was tested by unilateral binomial test. With *, ** and *** indicates significant difference at *p* ≤ 0.05, *p* ≤ 0.01 and *p* ≤ 0.001 levels respectively. The *p*-values of each condition were notified in parentheses.

<table><thead><tr><th rowspan="2"></th><th colspan="4">Temperature (°C)</th><th colspan="4">Salinity (g/L)</th></tr><tr><th>12</th><th>14</th><th>16</th><th>18</th><th>0</th><th>10</th><th>15</th><th>30</th></tr></thead><tbody><tr><td>Apoptotic cells (%)</td><td>0 (0.623)</td><td>0 (0.623)</td><td>0 (0.623)</td><td>5 (1)</td><td>5 (1)</td><td>0 (1)</td><td>10<br>(0.264)</td><td>0 (0.623)</td></tr><tr><td>Necrotic cells (%)</td><td>5 (1)</td><td>0 (0.623)</td><td>15<br>(0.0755)</td><td>25**<br>(2.57x10<sup>-3</sup>)</td><td>10<br>(0.264)</td><td>44***<br>(1.09x10<sup>-6</sup>)</td><td>10<br>(0.264)</td><td>20*<br>(0.0159)</td></tr><tr><td>Granulocytes-macrophages (%)</td><td>0 (0.623)</td><td>0 (0.623)</td><td>0 (0.623)</td><td>10 (0.264)</td><td>0 (0.623)</td><td>6 (0.603)</td><td>5 (1)</td><td>15 (0.0755)</td></tr><tr><td>Respiratory burst index<br>(Experimental units)</td><td>70*** (1.78x10<sup>-14</sup>)</td><td>70***<br>(1.78x10<sup>-14</sup>)</td><td>10 (0.264)</td><td>5 (1)</td><td>0 (0.623)</td><td>0 (1)</td><td>0 (0.623)</td><td>5 (1)</td></tr><tr><td>Phagocytic activity (%)</td><td>0 (0.623)</td><td>5 (1)</td><td>5 (1)</td><td>5 (1)</td><td>10<br>(0.264)</td><td>22* (0.0109)</td><td>5 (1)</td><td>5 (1)</td></tr></tbody></table>

18 °C (Fig. 2, Table 1). For necrotic cell percentage and respiratory burst, the difference between the control (16 °C) and the other temperature was increased. In fact, at 12 °C and 14 °C the percentage of necrotic cells was lower than at 16 °C (control) and 18 °C. Compared to the February reference range, levels of necrotic cells were significantly out of the reference range only at 18 °C (25%) (5% for 12 °C, 0% for 14 °C and 15% for 16 °C) (Fig. 2, Table 1). Conversely, the respiratory burst was significantly higher at 12 °C and 14 °C, which was confirmed by a significantly non nul out-of-range percentage compared to the February reference values (70% out-of-range) (Figs. 1–2, Table 1). Although the respiratory burst level at 18 °C was significantly lower than the control (16 °C) (Fig. 1), the percentage of data outside the February reference range was not significant, with 5% outside the range (Fig. 2, Table 1).

## 3.2. Effects of salinity on immune response and blood osmolarity

Fig. 3 suggests that all immune responses to salinity are different between T0 and T21. Even with the transformed values, significant differences in variance were observed between conditions at the different time points (Fig. 3), so we did not perform a two-way ANOVA. Only the log transformed necrotic cell counts were analyzed with a two-way ANOVA, which showed statistically significant differences between the response to salinity depending on the time point (date:salinity interaction F-test, *p* < 0.05).

After ten days of acclimatization by gradual increase of the salinity (T0), at the rate of 1/10 per day of the final salinity, many parameters were altered (Fig. 3). Only the percentage of apoptotic cells showed no difference between the salinity concentrations (10, 15 and 30 g/L) and the control value (0 g/L), even though at 10 g/L salinity the percentage of apoptotic cells had a higher value than at 30 g/L salinity (Fig. 3). The increase in salinity induced a proportional decrease in the percentage of granulocytes-macrophages, with a higher value at 0 g/L and a lower value at 30 g/L (Fig. 3). The respiratory burst index showed lower control data than at the other salinity levels (Fig. 3). The percentage of necrotic cells and the phagocytic activity showed similar values between 0 g/L (control) and 10 g/L; the percentages increased significantly at 15 g/L and even more at 30 g/L (Fig. 3). After these ten days of acclimatization, the osmotic pressures at 30 g/L and 15 g/L were similar to the control (0 g/L), whereas the 10 g/L had not yet reached the osmotic pressure of the control (Fig. 3).

After 21 days of exposure to the various salinity levels, we confirmed that the control condition (0 g/L) was within the October reference range (Fig. 4) with no significant error percentage (0% for the percentage of granulocytes-macrophages and respiratory burst index, 5% for apoptotic cells and 10% for necrotic cells and phagocytic activity, Table 1). After these 21 days of exposure, there is also less individual variability in the different responses (Fig. 3). For three parameters (apoptotic cells, percentage of granulocytes-macrophages and respiratory burst), the significant difference is not confirmed by projection on

the October reference range (Fig. 4). In fact, although the apoptotic cell levels were significantly lower at 15 g/L compared to control (Fig. 3), only 10% datapoints were out of range (Table 1). Similarly, the statistically significant difference obtained between the control condition and 15 g/L and 30 g/L for the percentages of granulocytes-macrophages (Fig. 3) did not imply a significant number of out-of-range points compared to the October reference range (Fig. 4) (0% for 0 g/L, 6% for 10 g/L, 5% for 15 g/L, 15% for 30 g/L, Table 1). The significant reduction in respiratory burst index at 30 g/L (Fig. 3) did not result in a significant number of out-of-range points compared to the October reference range (Table 1) (5% out of range). Regarding necrotic cells, the significant difference obtained at 10 g/L (Fig. 3) was confirmed by the large number of data outside the October reference range (44% out of range, Fig. 4, Table 1) and at 30 g/L a significant number of data points was also out of the reference range (20%) (Fig. 4, Table 1). Finally, although phagocytic activity was not significantly different between the control (0 g/L) and the other salinity levels (Fig. 3), a significant number of points were out of range at 10 g/L compared to the October reference range (22% out of range, Fig. 4, Table 1). The osmotic pressure was not significantly different between 15 g/L salinity and the control, whereas osmotic pressure was significantly higher at 10 g/L and 30 g/L salinity after 21 days of exposure (Fig. 3).

## 4. Discussion

To better understand the natural variability of the immunomarkers and to increase the relevance of the immunomarkers for the detection of environmental disturbances, some reference ranges of the non-specific immune response of stickleback have been developed. In this context, Marchand et al. (2019) had modelled the effect of three main founders (season, length and sex) under laboratory conditions. The statistical model was then tested in mesocosms and under different stream conditions (Marchand et al. 2020, 2021). However, under field conditions, temperature and salinity can vary from station to station, justifying the importance of determining the impact of these confounding factors on the immune response. During the experiments, fish density was changed after each sampling date. Nevertheless, stocking density is known to induce stress, which could differentially affect the immune response of fish (Ellison et al., 2020). For stickleback, Le Guernic et al. (2016) showed that the innate immune response of stickleback was not affected by stocking density. This finding was supported here by the lack of changes in the immune response in the control along the two experiments. Therefore, this confounding factor had no effect on this study.

Many authors have shown that temperature changes can modify innate parameters, especially phagocytic activity (Alcorn et al., 2002; Collazos et al., 1994; Le Morvan et al., 1997; Magnadóttir, 2006). In the present work, even though we did not acclimatise to the temperature tested, only the percentage of apoptosis showed some variation at 14 °C, all the other immune responses were independent of the time point,
<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

**Fig. 3.** Osmotic (mOsm/Kg, F) and immune responses (leucocyte distribution (granulocytes-macrophages %, A), cellular mortality (apoptosis and necrotic cell (%, B et C)), respiratory burst (MFI, D) and phagocytosis activity (%, E) of stickleback was collected after ten days of acclimatation (T0) and 21-days (T21) of exposure to dissimilar salinity (from 0 to 30 g/L). ANOVA followed by a t-test with Holm's adjustment for multiple comparisons (parametric data, p ≤ 0.05) or Kruskal-Wallis test followed by a Wilcoxon test with Holm's adjustment for multiple comparisons (non-parametric data, p ≤ 0.05) were used to test statistical differences between salinity at each sampling date. Boxplots represent the values of 20 fish (n) including male (red) and female (green). For each parameter and date, difference between conditions was indicated by different letters (p < 0.05). (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)
<figure><img src="image_12.png" /><figcaption>**Fig. 4.** Projection of salinity data after 21 days of exposure on the October reference ranges. For apoptotic cells (A), granulocyte-macrophage percentages (C), respiratory burst index (D) and phagocytosis activity (E), the lines represent the values predicted by the model and the prediction interval at 95 %. For necrotic cells (B), the full lines represent prediction intervals for female and the dotted lines represent prediction intervals for male. With MFI = Mean Fluorescent Intensity.</figcaption></figure>

regardless of the temperature used. After seven days of exposure to temperatures ranging from 12 °C to 18 °C, few effects were observed. Only the phagocytic activity was lower in cold water (12 °C and 14 °C) compared to the current laboratory temperature (16 °C) and to the higher temperature (18 °C), and after 14 days of exposure no more effects were detected. Similarly, exposure of juvenile cachama (*Colossoma macropomum*) to three temperatures (18, 28, 35 °C) for 21 days had no effect on immunological parameters such as bacterial killing by phagocytic cells and leucocyte distribution (Salazar-Lugo et al., 2009). Although Ainsworth et al. (1991) also observed a reduction in phagocytic activity at lower temperature (10 °C) compared to higher temperature (27 °C) in channel catfish, this result was obtained after 21 days of exposure, at which time no effect was observed here. Similarly, a reduction in phagocytic activity at lower temperature (5 °C) in rainbow

trout (*Oncorhynchus mykiss*) was obtained after 57 days of exposure (Nikoskelainen et al., 2004). However, these two authors did not provide information on the sampling period. As recommended by Collazos et al. (1994), the season is very important when working on the influence of temperature on the immune response. In line with this, the present work has determined the effect of temperature on phagocytic activity using a seasonal reference value. Furthermore, Collazos et al. (1994) suggest a good adaptation of the phagocytes to temperature variation, especially in winter, mainly due to a high efficiency of the phagocytic capacity of granulocytes. The present work confirms these results by the absence of significant temperature effect on this parameter, suggesting a good plasticity of the phagocytic activity of sticklebacks to temperature variation in winter. The other immune parameters are rather modified after 14 days of exposure. After 21 days of exposure,
this significant immune variation seems to be mainly related to a natural variation, as shown by the projection on the reference values. Although some authors have shown an increase in the percentage of granulocytes-macrophages in the blood or head kidney when the fish are kept at low temperatures (Alcorn et al., 2002; Kurata et al., 1997), no effect was observed in the present work, suggesting also a good plasticity of this parameter. Only the respiratory burst at 12 °C and 14 °C and the necrotic cells at 18 °C were significantly outside the reference ranges, with an increased response compared to the control (16 °C). This increase in respiratory burst response at the lowest experimental temperature was also obtained in sea bass (*Dicentrarchus labrax*) phagocytes (Angelidis et al., 1988). As previously shown (Collazos et al., 1994; Le Morvan et al., 1997), this increase in the respiratory burst index is due to a decrease in both the basal level of ROS and the level of PMA-activated ROS (Table S1). At very high temperatures, the immune response seems to be more affected. Indeed, heat stress (>20 °C for temperate species) seems to induce an overproduction of ROS resulting in a reduced phagocytic activity (Mahmoud et al., 2020), probably due to heat shock (Collazos et al., 1994). Similarly, in sticklebacks, temperatures from 18 to 28 °C impaired phagocytic activity, particularly due to a marked decrease in respiratory burst activity (Dittmar et al., 2014). Furthermore, after 24 °C, exposure to a fungal immune inducer, zymosan, does not stimulate the respiratory burst (Scharsack et al., 2016). However, this temperature range is not realistic in the field during winter. Nevertheless, the increase in basal ROS levels at higher temperatures (18 °C, Table S1), although within the reference range, could explain the significant increase in necrotic cells by direct action of ROS on the leucocyte membrane. For example, Prophete et al. (2006) have shown that elevated temperature mediates the production of superoxide and free radicals in fish. In conclusion, even if temperature has an effect on the cellular immune response of sticklebacks, it remains within the natural variation of these immune markers. In this way, the reference values developed for the cellular immune response of stickleback could be used within different temperature ranges. Due to the euryhaline characteristics of stickleback, the effect of salinity on the immune response needs to be better understood before this species can be used in biomonitoring studies involving sites with different salinities. In the present work, the 21 days of exposure to salt clearly demonstrate the good adaptation of stickleback to salt water in terms of innate immune response and osmolarity. In fact, the blood osmolarity of stickleback was lower than control values at 10 g/L after ten days of progressive acclimatization and higher at 10 and 30 g/L after 21 days of exposure. These results suggest an adaptive process allowing the survival of the fish during salt exposure, which seems to be more effective at high salinities (15 g/L, 30 g/L) than at low salinities (10 g/L). The regulation of blood osmolality involves a complex network of physiological and genetic pathways. In stickleback, the expression of 20 genes mapped to the same diuretic pathway is involved (Wang et al., 2014). Many of these genes are essential for ion regulation, facilitating transport by energy conversion or suppressing passive diffusion of ions. They are also directly involved in the construction of ion channels or ion pumps. For example, after exposing freshwater sticklebacks to a salinity of 11 g/L or 33 g/L for 30 days, these authors have demonstrated a downregulation of the expression of a serine/threonine protein kinase (SGK1) (Wang et al., 2014), which plays an important role in renal sodium excretion through phosphorylation of epithelial sodium channels that regulate sodium intake (Wulff et al., 2002). Similarly, higher expression of the Na+/H+ exchanger (SLC9a2) (Taugbøl et al., 2022) and reduced expression of the sodium/potassium-transporting ATPase subunit alpha-1 (ATP1A1) (McCairns and Bernatchez, 2010; Wang et al., 2014) have been shown in stickleback. Due to its ability to establish and maintain the electrochemical gradients of sodium and potassium across the plasma membrane, ATP1A1 is identified as a candidate gene for freshwater adaptation in stickleback (McCairns and Bernatchez, 2010). The expression of CLCNKA was down-regulated in the kidney, suggesting reabsorption of chloride from kidney cells into the blood, while

SLC12A1 and SLC12A3 genes, encoding the sodium-potassium-chloride cotransporter, were up-regulated to increase reabsorption of sodium chloride from blood into kidney cells (Wang et al., 2014). However, other physiological functions may also be affected by this change in salinity. Although freshwater sticklebacks are able to adjust their gene expression, a change of ~1500 transcripts suggests an acute energy cost when salinity changes, accompanied by an increase in stress-related genes (HSP family) and immune response through upregulation of protein tyrosine phosphatase receptor type C (PTPrc) (Taugbøl et al., 2022). In the same way, cytochrome P450 genes corresponding to CYP1a were upregulated in freshwater stickleback exposed to saltwater (Taugbøl et al., 2022). This biomarker is directly linked to general stress and immune responses (Lenoir et al., 2021). In addition, salinity change was known to disrupt the immune response. For example, in juvenile *Takifugu fasciatus*, total leucocyte count, serum protein, albumin and globulin concentrations, C3 concentration and lysozyme activity were significantly decreased with the increase of salinity to 15 g/L to 30 g/L (Wang et al., 2019). In the present work, after the 10 days of acclimatization, each innate immune response tested was significantly modified for each salt stress (i.e. modification of leucocyte distribution, increase of necrosis, respiratory burst and phagocytosis efficiency) and the inter-individual variability was significant. Three weeks of exposure of juvenile catfish to increase salinity (up to 10 g/L) followed by infection with *Edwardsiella ictaluri* affected the immune kidney proteome, particularly in expressed proteins related to leucocyte diapedesis, phagocytosis and respiratory burst activity (Schmitz et al., 2017). In brown trout, *Salmo trutta*, seawater transfer increases plasma growth hormone, which is associated with stimulation of macrophage functions such as phagocytic activity (Marc et al., 1995) and macrophage respiratory burst activity (Taylor et al., 2007). Similarly, in tilapia, *Oreochromis mossambicus*, transfer from freshwater to seawater (25 g/L) increases, among other things, phagocytic and respiratory burst activities (Jiang et al., 2008). However, after 21 days of exposure to stable salinity, inter-individual variability and effects were reduced. In fact, only necrosis activity at 10 and 30 g/L salt and phagocytic activity at 10 g/L salt were significantly affected compared to the reference range. This result suggests a good adaptability of freshwater stickleback to seawater and increases the attractiveness of using this species and their cellular immune reference values in a biomonitoring programme to compare freshwater and saltwater field sampling. As shown by Li and Kültz (2020), stickleback can adjust gill function to environmental salinity using its specific osmoregulatory gill proteins and KEGG pathways. These adaptive osmoregulatory mechanisms promote transepithelial ion transport, cellular osmoregulation and remodelling of the gill epithelium, which allows the maintenance of physiological functions such as the immune response (Li and Kültz, 2020).

In conclusion, as far as the non-pathogenic innate immune response is concerned, apart from leucocyte necrosis, the innate immune response of sticklebacks has a good capacity to adapt to changes in temperature and salinity. Nevertheless, the use of respiratory burst at low temperature should be interpreted with caution and a temperature-dependent reference range should be proposed for this parameter. In addition, the interactions between temperature and salinity should be carefully studied to better understand the effects on the immune response. The reference ranges proposed here must also be used with caution, as the stickleback used had similar lengths. Finally, for homogeneous length groups, this study argues for the possibility of using sticklebacks and their immunomarker reference ranges in the field regardless of temperature and salinity, due to their relatively temperature and salinity independent innate immune response. The use of a sentinel species that is less affected by co-factors and adapted to salinity variation is an added value for the development of active biomonitoring. In addition, these results could be very attractive for a better understanding of their genomic evolution, opening the door to a probable role of epigenetic modifications and/or a freshwater-adaptive allele in their immune adaptability.
## CRediT authorship contribution statement

**Anne Bado-Nilles:** Writing – original draft, Visualization, Validation, Supervision, Resources, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Conceptualization.
**Cleo Tebby:** Writing – original draft, Validation, Methodology, Formal analysis.
**Alexandrine Pinet:** Investigation.
**Cyril Turies:** Project administration, Methodology, Investigation.
**Jehan-Hervé Lignot:** Methodology, Investigation.
**Jean-Marc Porcher:** Supervision.

## Declaration of competing interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: Anne Bado-Nilles reports financial support was provided by French Agency for Biodiversity. If there are other authors, they declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgment

This work is funded by the French Ministry of Ecology and Sustainable Development (Programme 190 Ecotoxicology and Programme 181 DRC50) and by the French Biodiversity Agency (SASHIMI project). During the preparation of this work, we used DeepL Write in order to improve language and readability. We would like to thank VETOFISH (13 220 Châteauneuf-les-Martigues, France) for their monitoring of the health of the stickleback.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.marenvres.2025.106972.

## Data availability

Data will be made available on request.

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