Physiological and molecular responses of juvenile ...

CBA-10137; No of Pages 9 Comparative Biochemistry and Physiology, Part A xxx (2016) xxx–xxx

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Comparative Biochemistry and Physiology, Part A journal homepage: www.elsevier.com/locate/cbpa

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Yueyang Zhang a,b, Jennifer R. Loughery a, Christopher J. Martyniuk c, James D. Kieffer a,⁎

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Physiological and molecular responses of juvenile shortnose sturgeon (Acipenser brevirostrum) to thermal stress

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Article history: Received 13 June 2016 Received in revised form 13 October 2016 Accepted 19 October 2016 Available online xxxx

Department of Biological Sciences, University of New Brunswick, Saint John, New Brunswick, Canada Department of Biological Science, University of Alberta, Edmonton, AB, Canada Center for Environmental and Human Toxicology, and Department of Physiological Sciences, UF Genetics Institute, College of Veterinary Medicine, University of Florida, Gainesville, Florida, USA

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The shortnose sturgeon (Acipenser brevirostrum LeSueur, 1818) is a vulnerable species that is found along the eastern coast of North America. Little is known about temperature tolerance in this species and with a rapidly changing global climate, it becomes increasingly important to define the thermal tolerance of this species to better predict population distribution. Using a modified critical thermal maximum test (CTMax), the objectives of this study were to determine the impact of heating rate (0.1, 0.2 and 0.25 °C min−1) on the thermal tolerance, associated hematological responses, and oxygen consumption in juvenile sturgeon. In addition, transcripts associated with physiological stress and heat shock (i.e., heat shock proteins) were also measured. Heating rate did not alter the CTMax values of shortnose sturgeon. Neither heating rate nor thermal stress affected plasma sodium and chloride levels, nor the expression of transcripts that included catalase, glucocorticoid receptor, heat shock protein70 (hsp70), heat shock protein 90α (hsp90α) and cytochrome P450 1a (cyp1a). However, regardless of heating rate, thermal stress increased both plasma potassium and lactate concentrations. Glucose levels were increased at heating rates of 0.2 and 0.25 °C min−1, but not at 0.1 °C min−1. Overall, oxygen consumption rates increased with thermal stress, but the response patterns were not affected by heating rate. These data support the hypothesis that shortnose sturgeon can tolerate acute heat stress, as many physiological and molecular parameters measured here were non-responsive to the thermal stress. © 2016 Published by Elsevier Inc.

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Keywords: Acipenser brevirostrum Shortnose sturgeon Thermal tolerance Hematology Oxygen consumption Heat shock protein

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Water temperature is considered to be one of the most influential abiotic factors that affect the distribution, abundance, physiology and behavior of fishes (Fry, 1971; Beitinger and Lutterschmidt, 2011; Akhtar et al., 2013). The thermal stress response in fish is affected by both the duration and magnitude of the stressor, as well as genetic factors (reviewed in Barton, 2002; Kassahn et al., 2009). Fish are also subjected to considerable diurnal temperature changes (Rodnick et al., 2004). For example, the discharge of cooling water by power plants into a river can also rapidly increase the water temperature (Beitinger et al., 2000; Rajaguru, 2002). As a result, fish can be exposed to different rates of water temperature increases in their natural environment, which can result in a range of physiological and molecular responses to cope with the stressor (Basu et al., 2002; Comin et al., 2004; Mora and Maya, 2006). When water temperatures become too high and fish experience elevated temperatures over time, thermal stress can eventually overwhelm the physiological and genetic mechanisms needed to compensate for the negative effects of the thermal stress

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1. Introduction

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⁎ Corresponding author. E-mail address: [email protected] (J.D. Kieffer).

(reviewed in Iwama et al., 2004; Richardson et al., 1994; Lee et al., 2003). Temperature governs an organism's metabolism (Fry, 1971). Metabolism reflects an organism's energy expenditure and its ability to perform activities to survive environmental stressors (Davison, 1997; Zeng et al., 2010; Akhtar et al., 2013). Temperature elevates metabolic rates (e.g., often measured by changes in oxygen consumption) of fish (Eliason and Farrell, 2016) to a point where less energy can be invested into growth, locomotion, reproduction, and homeostasis (Atkinson, 1996; Person-Le Ruyet et al., 2004; Singer and Ballantyne, 2004). In the context of maintaining homeostasis, studies have shown that temperature modifies blood chemistry values, such as cortisol level, ion concentrations, total protein level, hemoglobin and hematocrit (Houston, 1997; Nikinmaa and Salama, 1998; Marco et al., 1999), thus the individual may invest significant energy into balancing these biochemical parameters. A significant body of literature reports on the relationship between temperature and physiology/molecular responses in teleost fishes (Crawshaw, 1977; Currie et al., 1998; Beitinger et al., 2000; Claireaux et al., 2006; Komoroske et al., 2015; Jesus et al., 2016), but limited data have been gathered on the relationship between temperature and physiology/molecular responses in ancient fish, including sturgeon

http://dx.doi.org/10.1016/j.cbpa.2016.10.009 1095-6433/© 2016 Published by Elsevier Inc.

Please cite this article as: Zhang, Y., et al., Physiological and molecular responses of juvenile shortnose sturgeon (Acipenser brevirostrum) to thermal stress, Comp. Biochem. Physiol., A (2016), http://dx.doi.org/10.1016/j.cbpa.2016.10.009

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Juvenile shortnose sturgeon were purchased from Acadian Sturgeon and Caviar (Carter's Point, Kingston, New Brunswick, Canada). Fish were held in three- 1 m diameter (160 L) cylindrical with flow-through aerated, de-chlorinated municipal freshwater at the University of New Brunswick (Saint John, N.B., Canada). The tanks were supplied with a flow-through of fresh (1 L min− 1 renewal rate), aerated, de-chlorinated, municipal water at 15 °C. Fish were fed twice daily to satiation (Corey Aquafeed: 1.5 mm optimum, 52% protein), but fish were fasted for 24 h prior to the experiments. Fish used in the study were acclimated to 15 °C for at least 4 weeks. A 12 h light: 12 h dark photoperiod was maintained throughout the study.

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2.3. Experiment 1: effect of heating rate on thermal tolerance, blood 154 parameters and molecular endpoints for stress 155 Thermal tolerance was determined using the modified critical 156 thermal maximum (CTMax) methodology as described in Zhang and 157 Kieffer (2014) and Spear and Kieffer (2016). 158

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Two subsets of shortnose sturgeon were used: 1) those used to determine the effect of heating rate on thermal tolerance, blood parameters and molecular responses to stress; and 2) those used to evaluate the effects of heating rate on oxygen consumption rates during the modified critical thermal maximum tests (see below for details).

2.3.1. Experimental setup CTMax experiments were conducted in an insulated, rectangular test tank (approximately 60 L). A heating reservoir (approximately 45 cm diameter, 56 cm height, and 90 L), which stood next to the test tank, was equipped with a 1000 W heater (Aquatic Ecosystems) and a 60 cm bio-weave air diffuser (Aquatic Ecosystems). The test tank was filled with water to a depth of 15 cm (~28 L). The heater in the reservoir was calibrated to increase the water temperature in the test tank by either 0.1 °C min− 1, 0.2 °C min− 1 or 0.25 °C min− 1. To maintain temperature within the setup, the water from the reservoir flowed to the test tank by gravity and was then pumped back to the heating reservoir via a submersible pump (Loligo Systems, Denmark). The submersible pump was isolated from the test chamber by a perforated black plexiglass shield. An electronic thermometer (Loligo Systems, Denmark) was placed at each end of the test tank to record the temperature within the test tank.

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2.3.2. CTMax test design A single juvenile shortnose sturgeon (~ 150 g, one-year-old) was placed in the insulated rectangular test tank with an air stone at 15 °C for 1 h to recover from handling stress (Zhang and Kieffer, 2014). For the control group (no thermal stress) (N = 10), fish were anaesthetized with 250 mg L−1 of MS222 (buffered in NaHC03) after the 1 h period (Zhang and Kieffer, 2014). Once fully anaesthetized, the fish was removed from the tank and the weight and length taken, and a blood sample was taken from the caudal vasculature using a lithium heparinized needle and syringe and stored for further analysis. The individual was then sacrificed by spinal severance and a liver sample (~ 0.04–0.05 g each) was collected for molecular study and snap frozen at − 80 °C. Following the 1 h period, the fish from the experimental group (thermal stress) (N = 10), were exposed to a thermal stress at a constant heating rate of either 0.1 °C min−1 (0.102 ± 0.0031 °C min−1), 0.2 °C min−1 (0.200 ± 0.003 °C min− 1) or 0.25 °C min−1 (0.247 ± 0.006 °C min−1; N = 10 fish for each heating rate). Beitinger et al. (2000) recommended heating rates ranging between 0.1 and 0.3 °C min−1 in CTMax tests for fish to avoid bias. A slow (0.1 °C min−1) and medium (0.2 °C min−1) heating rate were selected. However, the linear increasing temperature could not be maintained at high (0.3 °C min−1) heating rate throughout the entire experiment period. Therefore, 0.25 °C min−1 was selected. Loss of equilibrium (LOE) was the endpoint used for the critical thermal maximum test (Zhang and Kieffer, 2014), and was defined as the first time a fish was unable to remain in the dorso-ventrally upright position for a consecutive 10 s. Once the fish had reached LOE, the final temperature was recorded and was considered the CTMax temperature. Fish were then anaesthetized at the CTMax temperature and blood samples and liver were collected from the experimental fish as noted above for control fish. After each trial, the tanks were emptied and cleaned prior to the next experimental trial.

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(Cech and Doroshov, 2005; Allen et al., 2006; Sardella et al., 2008; Ziegeweid et al., 2008; Zhang and Kieffer, 2014). Sturgeons have existed for more than 200 million years (Robles et al., 2004) and are considered “living fossils” with most species now identified as threatened, vulnerable or endangered (Cech and Doroshov, 2005). Determining the thermal tolerance of a species is important for understanding its physiology and ecology, as well as predicting the long-term effects of climate change (Pörtner and Peck, 2010). This knowledge is required to improve management and conservation efforts for endangered species (Deslauriers et al., 2016). In addition, the molecular mechanisms that underlie heat stress tolerance are important to measure in order to better determine the range of responses involved in heat tolerance. Thermal stress has been shown to induce oxidative stress responses in a number of fish species (Airaksinen et al., 1998; Basu et al., 2002; Iwama et al., 2004; Barat et al., 2016). For example, heat shock proteins are critical for maintaining the proper structure and function of proteins within the cell and are often used as molecular indicators for general stress (Basu et al., 2002; Iwama et al., 2004; Ni et al., 2014). Moreover, higher water temperature has been shown to increase cortisol level in fish (Davis, 2004), which can bind to glucocorticoid receptors to regulate transcription of heat shock proteins (Iwama et al., 1999). Thus, transcripts related to cortisol and heat shock can be regulated by temperature. Oxidative stress is also affected by heat stress which promotes the generation of reactive oxygen species (ROS) in cells (Trenzado et al., 2006; Furné et al., 2009). This is followed by an increase in the production and activity of antioxidant enzymes, such as catalase to protect the cell (Valavanidis et al., 2006; Furné et al., 2009). Other transcripts such as those related to xenobiotic transformation (e.g. cyp1a) have also been measured as indicators of environmental stress. To gain a holistic perspective on the implications of thermal stress on aquatic organisms, an integrative approach involving multiple endpoints across biological levels of organization should be utilized. The current study used a modified CTMax test to investigate the effects of thermal stress at three different heating rates (0.1, 0.2 and 0.25 °C min− 1) on physiological and molecular parameters, such as thermal tolerance, hematological parameters, oxygen consumption rate, and expression levels of transcripts related to stress, in shortnose sturgeon acclimated to 15 °C. The goal was to elucidate physiological and/or molecular indictor(s) that may be related to thermal stress. The overarching hypothesis was that sturgeon exposed to a high temperature over a longer period would experience higher stress due to the higher energy demand. Based on this hypothesis, we predicted that: (1) thermal tolerance will be higher in sturgeon at the highest heating rate; (2) the most dramatic physiological and molecular responses related to heat stress would be observed at the highest heating rate; (3) Expression of genes involved in increasing the capacity of the fish to handle the thermal stress will increase.

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Whole blood samples were collected (see above) placed in 1.5 mL centrifuge tubes. Approximately, 100 μL blood samples were used for duplicate hematocrit determination (Penny and Kieffer, 2014). The remaining sample was centrifuged for 2 min at 10,000 rpm and the resulting plasma was decanted and placed into new labeled tubes. From the plasma, glucose was measured in duplicate using an OneTouch Ultra glucose meter (OneTouch Ultra 2; Code 25 test strips; www. onetouch.ca; Penny and Kieffer, 2014). The remaining plasma was frozen and stored at − 80 °C for later analysis. Sodium, potassium and chloride levels were measured using a Smartlyte Ion Analyzer (Diamond Diagnostics, U.S.A.). The plasma protein concentrations were measured by a standard colorimetric assay at 540 nm wavelength (Total Protein Reagent, Buiret Method & Protein Standard Set P5495; Sigma; www. sigmaaldrich.com; Penny and Kieffer, 2014). The plasma lactate concentrations were measured using a standard spectrophotometric assay at 540 nm wavelength (Lactate Reagent 735-10, Lactate Standard Solution 826-10 & Lactate Standards Set 735-11; Trinity Biotech; www.trinitybiotech.com; Zhang and Kieffer, 2014).

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RNA was isolated from frozen liver tissues using 1 mL TRIzol® reagent following the RNA isolation protocol (TRIzol® Reagent, Invitrogen, Burlington, ON, Canada). RNA pellets were reconstituted in 20–30 μL of RNA Secure™ (Ambion, Austin, TX, USA) following the manufacturer's protocol. RNA quality was measured on a NanoDrop 2000 (Thermo Scientific) and RNA integrity was evaluated on an Agilent 2100 Bioanalyzer (Agilent, Mississauga, ON, Canada). Samples with RNA integrity number (RIN) b6 were excluded from further analysis. The sample with the lowest RIN used in the study was 6 and the mean was 7.3 ± 1.2 SD.

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Real-time qPCR (qPCR) followed that outlined in Cowie et al. (2015). Briefly, genomic DNA was digested with TURBO DNA-free™ DNase (Ambion Austin, TX, USA). Approximately 2–5 μg of RNA was treated with 2 μL 10× TURBO DNase Buffer and 1 μL TURBO DNase for 20 min at 37 °C. Inactivation reagent (2 μL; Ambion Austin, TX, USA) was added to the sample to remove DNase and divalent cations. RNA quantity was measured on the NanoDrop 2000 (Thermo Scientific). Synthesis of 10 μL cDNA was generated from 1 μg total DNase treated RNA with 2 μL 5 × iScript Reaction Mix, 0.5 μL iScript Reverse Transcriptase and Ultra-pure water following the Bio-Rad iScript™ cDNA Synthesis Kit (Hercules, CA, USA). Samples were first diluted (1/40 dilution) in RNAse-DNAse free water prior to real-time PCR analysis. Primers (Table 1) were designed using Primer3 (Untergasser et al., 2012) and specificity was evaluated using PCR according to Cowie et al. (2015). All amplicons were verified as correct target genes by

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Table 1 Primers used to amplify shortnose sturgeon genes of interest. Provided in the table are the NCBI/UniProt accession identifiers, primer sequences (5′ to 3′), efficiency of primer pair (%E), and goodness of fit of linear regression for the relative standard curve.

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2.7. Experiment 2: effects of heating rate on oxygen consumption rates

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Juvenile shortnose sturgeon acclimated to 15 °C were randomly selected from its holding tank and placed into a custom built an intermittent flow respirometer system (3.57 L; Loligo Instruments) immersed in an ambient tank of water with an air stone at 15 °C. The respirometer system consisted of a recirculation pump, flush pump, flush out tube, temperature probe and oxygen electrode. The flush pump, which was controlled by a computer, replenished water inside the respirometer, and the recirculation pump remained on for the entire experiment to ensure mixing of water within the respirometer. During oxygen measurements, the flush pump was shut off while the recirculation pump continued running, making the system function like a closed respirometer (Kieffer et al., 2014). During this phase, the oxygen saturation level was measured by galvanic oxygen electrodes (Loligo Systems, Denmark). After a designated measurement period (set using the computer program), the flush pump was activated to restore the water oxygen saturation in the respirometer back to 100% saturation.

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Sanger Sequencing at the McGill University and Génome Québec Innovation Centre (Montréal, QB, Canada). Glucocorticoid receptor, hsp70 and hsp90 were used to assess the general stress response. Responses to oxidative stress were assessed by measuring cytochrome P450 1a (cyp1a) and catalase. Real-time PCR was performed using SsoFast™ EvaGreen® Supermix (BioRad, Mississauga, ON, Canada) and genes were assayed using the CFX96™ Real-Time-PCR Detection System (BioRad). Primers were tested for efficiency using a seven point dilution series. All standard curves were generated with pooled liver cDNA. The two-step thermal cycling parameters for the reaction were as follows: initial 1-cycle Taq polymerase activation at 95 °C for 30 s, followed by 40 cycles at 95 °C for 5 s, and annealing at the primer-specific temperature (58 °C for all primers) for 30 s. After 40 cycles, a DNA melt curve was generated starting at 65.0 to 95.0 °C with increment of 0.5 °C for 5 s. Four reference genes were evaluated for stability (β-actin, rps18, ef1 and gapdh (Vandesompele et al., 2002). An ANOVA was used to compare the mean Cq values among groups for each reference gene to assess if reference genes were different among groups (β-actin, P = 0.55; rps18, P = 0.50; ef1, P = 0.40; gapdh P = 0.92). Values for reference genes (Cq) did not significantly differ across treatments (one-way ANOVA) based on RNA input. The geometric mean of two transcripts (β-actin and ef1) was the most stable combination for baseline normalization based upon the target stability function in CFX Manager™ software (combined M-value = 1.05, mean CV = 0.36). Each qPCR plate included 4 samples prepared without reverse transcriptase (NRT) and 1 sample without cDNA template (NTC). Samples were analyzed in duplicate with the following biological replicates: N = 8 for control group, N = 8 for 0.1 °C/min, N = 8 for 0.2 °C min−1, N = 9 for 0.25 °C min− 1. Normalized gene expression was extracted using CFX Manager™ software using the relative ΔΔCq method (Hellemans et al., 2007). The qPCR analysis followed suggestions of the MIQE guidelines (Taylor et al., 2010).

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ATGCGTGCATTTATCAGATCA CATTGTCACCAACTGGGATGAC CAGAAGACCAGGTGGTGTC TGACTGTGCCGTCCTGAT GGAGATCCTACCAGCAGTCTA TTCACCATCCCTCACTGCAT GGAGAGTTCTACAAGAGCCTG TTGAAGATGGAATCTTTGAAGTGAAA GAGCGCATCCCAGAGAGG

TGTGGTAGCCGTTTCTCAG ACACGCAGCTCATTGTAGAAGGT GGCTGTGATTGGAAGGGTTT CTGTAAGGTGGCTCAGTGGA CTTGGCTCTTCAGACCCTCT ACAGCCAGGAAGAGGAAGAC TGAGAGGCAGGTCTTCAGAG CCCTGGTGATGGAGGTGTAA GTGGATGAAGGAGGGGAACA

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AF188395.1 HQ439362.1 (Roy et al., 2011) EF413586.1 HQ449562.1 JX104648.1 HQ439360.1 JN700180.1 AY880255.1 DQ493908.1


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The fish was held in the respirometer for 6 h to recover from handling stress and to adjust to the respirometer chamber. The mean oxygen consumption rate of the last 5 h (when values were considered stable) during the recovery period was used as the routine oxygen consumption (routine MO2) of each fish. After the recovery period, the fish was exposed to the modified CTMax test at a heating rate of either 0.1 °C min−1 (N = 8), 0.2 °C min−1 (N = 8) or 0.25 °C min−1 (N = 8). The oxygen level was recorded during the experiment (see Appendices Fig. A1). The oxygen consumption rate (MO2) of each individual fish was measured using intermittent flow respirometry (Kieffer et al., 2014) and the CTMax endpoint was a loss of equilibrium (Zhang and Kieffer, 2014). The mass-specific oxygen consumption rate (mg O2 kg− 1 h− 1) for each fish was calculated continuously by using the slope of oxygen decline (mg O2 L− 1), time (h), respirometer volume (L) and wet weight of the fish (kg) during the measurement period, using the following equation:

affect the hematocrit (one-way ANOVA, F3,36 = 2.49, P = 0.076). The mean (± S.D.) hematocrit was 26.6%(± 2.9%), 30.5% (± 3.9%), 28.4% (± 4.7%)and 26.6% (± 3.0%) in control, and fish exposed to thermal stresses of 0.1 °C min−1, 0.2 °C min−1 and 0.25 °C min−1, respectively. The protein levels of the three thermally stressed groups were not significantly different from control values (no thermal stress; one-way ANOVA, F3,36 = 0.90, P = 0.45). The protein averaged 2.3 g dL− 1, 2.5 g dL− 1, 2.1 g dL−1 and 2.2 g dL− 1 in control, 0.1 °C min−1, 0.2 °C min−1 and 0.25 °C min−1, respectively. Plasma K+ concentrations were significantly elevated following heating stress, regardless of the heating rate, compared to control fish (one-way ANOVA, F3,36 = 17.72, P b 0.001; Fig. 1A). Following thermal stress, plasma lactate levels increased nearly 12-fold in all fish regardless of heating rate (one-way ANOVA, F3,36 = 18.69, P b 0.001,

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The normality and homoscedasticity for the critical thermal maximum (CTMax) and blood parameters (e.g. hematocrit, lactate and protein. Sodium, potassium and chloride levels) were evaluated by Shapiro-Wilk and Breusch-Pagan tests, respectively. One-way ANOVA tests (α = 0.05) were used to examine the effects of heating rate on CTMax and blood parameters. If data did not meet the assumptions of the ANOVA, then an ANOVA on ranks was used. Holm Sidak post-hoc tests (α = 0.05) were used to compare mean values between groups when the overall model was significant. Two-way repeated measures ANOVA (α = 0.05) was used to determine the effect of heating rate on the oxygen consumption rate during the CTMax test (factors: heating rate and temperature when oxygen consumption was measured). All real-time PCR data (untransformed) were checked for normality using Kolmogorov–Smirnov test and the expression data were not normally distributed. Thus, the expression data were analyzed using a non-parametric Kruskal–Wallis, followed by a Nemenyi test to identify groups that were significantly different. Statistical significance was determined using a P value ≤0.05. R version 3.0.3 was used for statistical analysis and GraphPad Prism 6 was used for generating graphs.

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The CTMax of shortnose sturgeon acclimated to 15 °C was not affected by heating rate (one-way ANOVA, F2,27 = 0.81, P = 0.97). Overall, the mean (± standard error) of CTMax values across heating rates were 31.06 °C (±0.31), 31.1 °C (±0.68) and 30.95 °C (±0.26) at 0.1 °C min−1, 0.2 °C min−1 and 0.25 °C min−1, respectively. Plasma Na+ concentrations were not influenced by thermal stress (one-way ANOVA, F3,36 = 1.57, P = 0.21). The mean (± standard error) plasma Na+ concentrations were 122.4 meq L−1 (± 1.9), 124.4 meq L− 1 (± 2.4), 123.8 meq L− 1 (± 1.7) and 128.1 meq L− 1 (± 1.7) at control, 0.1 °C min− 1, 0.2 °C min− 1 and 0.25 °C min−1, respectively. Similarly, plasma Cl− concentrations were not influenced by thermal stress (one-way ANOVA, F3,36 = 1.39, P = 0.26). The average plasma Cl− concentrations were 109.3 meq L−1, 105.9 meq L−1, 106.8 meq L− 1 and 111.0 meq L− 1 at control, 0.1 °C min− 1, 0.2 °C min−1 and 0.25 °C min−1, respectively. The heating rate did not

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Fig. 1. Mean (± S.E.) plasma K+ (meq L−1) (A), plasma glucose (mmol L−1) (B) and lactate (mmol L−1) (C) of juvenile shortnose sturgeon in the control group (no thermal stress group, N = 10) and following a critical thermal maxima test (thermally stressed group, N = 10) at a heating rate of either 0.1 °C min−1, 0.2 °C min−1 or 0.25 °C min−1. Different letters indicate significant differences (P b 0.05; Holm Sidak multiple comparison).


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The expression levels in liver of six target genes, including catalase (cat, Kruskal–Wallis test, F3,29 = 1.17, P = 0.34), glucocorticoid receptor (gr, F3,29 = 0.26, P = 0.85), heat shock protein70 (hsp70, F3,29 = 0.88, P = 0.46), heat shock protein 90α (hsp90α, F3,29 = 0.69, P = 0.57) and cytochrome P450 1a (cyp1a, F3,29 = 0.81, P = 0.33) were not different among control and thermally stressed groups (Fig. 2).

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Fig. 3A represents the effect of heating rate on patterns of oxygen consumption when plotted in relation to experimental time. It took approximately 160 min, 80 min and 62 min from the beginning of the experiment to the endpoint in each CTMax test at the heating rates of

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No study to date has examined the physiological and molecular responses to thermal stress invoked by different heating rates in shortnose sturgeon. The CTMax values of shortnose sturgeon acclimated to 15 °C were not significantly different at the three heating rates. The CTMax value at heating rates of 0.1 °C min− 1 in the current study (31.06 °C ± 0.31) is comparable to that reported in a previous study (31.49 °C ± 0.91; F1,16 = 0.926, P = 0.350; Zhang and Kieffer, 2014), and for those of other sturgeon species (see Spear and Kieffer, 2016). These findings are also consistent with most studies on other fish species that used a similar range of heating rates (reviewed in Beitinger et al., 2000). However, some studies showed increased

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0.1 °C min−1, 0.2 °C min−1 and 0.25 °C min−1, respectively (Fig. 3A). When plotted in relation to temperature, oxygen consumption rates among groups of fish thermally stressed were not significantly affected by heating rate (repeated measures ANOVA, F2,29 = 1.89, P = 0.17; Fig. 3B). The routine MO2 (one-way ANOVA, F2,21 = 0.26, P = 0.77) and maximal MO2 (one-way ANOVA, F2,21 = 0.023, P = 0.92) were not different for all three thermally stressed groups (Table 2).

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Fig. 1B). Plasma glucose levels were significantly influenced by heating rate (one-way ANOVA, F3,36 = 6.02, P = 0.002; Fig. 1C). Specifically, the glucose levels were elevated in fish exposed to heating rates of 0.2 °C min−1 and 0.25 °C min−1compared to fish in the control group and those exposed to a heating rate of 0.1 °C min−1 (Fig. 1C).

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Fig. 2. Normalized gene expression of hsp70 (A), hsp90α (B), catalase (C), cytochrome P450 1a (D) and glucocorticoid receptor (E) in juvenile shortnose sturgeon liver from CTMax tests following heating rates of 0.1 °C min−1, 0.2 °C min−1, and 0.25 °C min−1, and the control group (no thermal stress). There were no differences between groups (P N 0.05, non-parametric Kruskal–Wallis test).


393 394 395 396 397

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Kieffer, 2014). During the CTMax trials, sturgeon became agitated and attempted to escape from the testing tank as temperature was increased towards their upper thermal limits. This intense, burst swimming activity is mainly supported by anaerobic metabolism (Kieffer, 2000). The accumulation of lactate in the blood from the anaerobic metabolism decreases the hemoglobin-oxygen affinity (Nikinmaa, 1990). Therefore, the increased energy expenditure and reduced oxygen affinity (Herbert et al., 2006) contributed to the activation of anaerobic respiration. High temperature may cause red blood cells to swell (Nikinmaa, 1990; Lang, 2012). Red blood cells release K+ into the plasma as a compensatory mechanism to regulate cell volume under thermal stress (Lang, 2012). This may explain why the plasma K+ concentration was significantly elevated in all three thermally stressed groups. However, the plasma Na + and Cl− concentrations in shortnose sturgeon were not significantly affected by thermal stress nor heating rate (Table 2). Oxygen consumption rate of sturgeon in the current study followed the same trend regardless of heating rate during the CTMax test, which suggests that oxygen consumption rate was only affected by the temperature, and not by the rate of temperature change. Tunnah et al. (2016) observe the similar trend in their study. The oxygen consumption of Atlantic salmon exposed to two thermal cycles with different rates of temperature change was similar at the end of the experiment (27 °C). Therefore, temperature appears to be the driving force for the oxygen consumption rate of fish. As elevated temperatures increase the metabolic rates (Brett, 1964; Clarke and Johnston, 1999; Das et al., 2004; Zaragoza et al., 2008; Akhtar et al., 2013; present study), fish also must increase the availability of blood glucose to provide enough fuel for aerobic and anaerobic respiration (Zaragoza et al., 2008; Zhang and Kieffer, 2014). The lower glucose level at 0.1 °C min− 1 may indicate the depletion of glycogen storage in the fish due to the longer exposure time, relative to the other two temperature stressed groups. Acute thermal stress during CTMax test did not result in a change in the expression of heat shock proteins 70 and 90α, catalase, glucocorticoid receptor, or cytochrome P450 1a. Studies have shown that heat shock proteins can be up-regulated when animals are exposed to thermal stress (reviewed in Iwama et al., 1999, Basu et al., 2002, Iwama et al., 2004). Recently, Linares-Casenave et al. (2013) showed that the mRNA levels of hsp60, hsp70 and hsp90 genes in larval green sturgeon at stage 45, when yolk-sac absorption occurs, were up-regulated when the fish were exposed to temperatures ranging from 20 to 28 °C for 24 h. Wang et al. (2013) also showed that the hsp70 response in multiple tissues of juvenile white sturgeon and juvenile green sturgeon (weight ~ 2.3 kg), were significantly higher than levels in the control group. Fish require oxygen to maintain normal whole-body functions, but an imbalance between antioxidant defenses and oxidative processes can result in a generation of reactive oxygen species (ROS) and tissue damage (Trenzado et al., 2006; Furné et al., 2009). Catalase and CYP1a are among the main antioxidant and oxidative stress reactive enzymes in fish (Furné et al., 2009; Madeira et al., 2013; Vinagre et al., 2014, Olsvik et al., 2013). Studies show that mRNA levels of liver cat of 1-year-old black porgy (Acanthopagrus schlegelii Bleeker, 1854)

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Fig. 3. Mean (± S.E.) oxygen consumption rates (MO2; mgO2 kg−1 h−1) of shortnose sturgeon (~150 g) at a heating rate of 0.1 °C min−1, 0.2°C min−1 or 0.25 °C min−1 during CTMax test over time (A) and over temperature (B). There were no differences between groups (P = 0.169, Repeated measures ANOVA).

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t2:1 t2:2

Table 2 Results from metabolic studies for CTMax test at three different heating rates, including mean and standard error (±S.E.) of routine MO2 and maximal MO2.

419 420 421 422 423 424 425 426 427 428

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thermal tolerance (i.e., increased CTMax values) of fish species at a very fast heating rate (more than 20 °C h−1, 0.33 °C min−1) or reduced tolerance at a heating rate lower than 6 °C h−1 (0.1 °C min− 1). The discrepancies between studies reflects the importance of using standard and/or recommended heating rates during thermal tests (Becker and Genoway, 1979; Beitinger et al., 2000), and careful consideration is required for the various inputs (acclimation temperature and body mass) in order to understand variability and processes underlying the thermal tolerance of fish. Even though the CTMax values are similar among the groups, thermal stress triggers some hematological changes in shortnose sturgeon. Lactate levels were increased in thermally stressed fish regardless of heating rate, emphasizing the importance of anaerobic metabolism during thermal stress (Walsh et al., 1998; Murchie et al., 2011; Zhang and Kieffer, 2014). A rise in water temperature leads to an increase in the aerobic metabolism of fish species (Walsh et al., 1998; Das et al., 2004; Akhtar et al., 2013). Once temperature reaches certain point, aerobic metabolism could no longer provide enough energy for the increasing demand, so shortnose sturgeon have to incorporate anaerobic metabolism; Kieffer et al., 2014; Zhang and

410

t2:3

Thermally stressed groups

t2:4 t2:5 t2:6 t2:7 t2:8

Routine MO2 (mgO2 kg−1 h−1) Maximal MO2 (mgO2 kg−1 h−1) Metabolic scope (mgO2 kg−1 h−1) Factorial aerobic scope

p value

0.1 °C/min

0.2 °C/min

0.25 °C/min

134.6(±12.1) 328.8(±17.2) 194.4(±12.4) 2.54(±0.18)

139.1(±9.9) 324.8(±13.2) 185.7(±12.9) 2.37(±0.14)

129.0(±9.8) 332.8(±14.7) 203.6(±13.3) 2.64(±0.16)

0.774 0.917 0.619 0.498


430 431 432 433 434 435 436 437 438 439 440 441 442 443 444 445 446 447 448 449 450 451 452 453 454 455 456 457 458 459 460 461 462 463 464 465 466 467 468 469 470 471 472 473 474 475 476 477 478 479 480 481 482 483 484


485 486

5. Conclusion

518

Acclimation temperature, body mass and heating rate are influential factors in determining the CTMax value in teleost fish (Beitinger et al., 2000). The current study investigated the effects of three heating rates at an acclimation temperature of 15 °C. The results from our previous study (Zhang and Kieffer, 2014) and current study suggest that acclimation temperature has the most profound effects on the thermal tolerance of shortnose sturgeon. Expression of the investigated genes and several blood variables (Na+, Cl+, Hct and blood protein) were not significantly affected by the heating rate in the current study. This may indicate that the duration, rather than the magnitude of the exposure to thermal stress, is more important for triggering physiological and molecular compensatory mechanisms to mitigate thermal stress in shortnose sturgeon.

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531 Q11 Uncited references

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Clark et al., 2013 Cox, 1974 Deslauriers and Kieffer, 2012 Pang et al., 2013 Werner et al., 2001 Acknowledgements

538 This research was supported by a Natural Sciences and Engineering 539 Q12 Research Council of Canada (NSERC) Discovery grant to J.D.K. 540 541 542

Support was also provided by the MADSAM fish group. All procedures followed the guidelines of animal use set out by the Canadian Council on Animal Care.

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Appendix A

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Fig. A1. Mean (± S.E.) of oxygen level (mgO2/L) in the water during experiment at a heating rate of 0.1 °C min−1, 0.2°Cmin−1 or 0.25 °C min−1.

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Appendix B. Supplementary data

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Supplementary data to this article can be found online at http://dx. 546 doi.org/10.1016/j.cbpa.2016.10.009. 547

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References

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increased with high temperature (An et al., 2010). Cyp1a transcript levels were elevated in immature Atlantic salmon (Salmo salar) 487 13–19 °C. These studies used much slower heating regimes and fish 488 were exposed to thermal stress for longer than 10 h compared to 489 60, 80 and 160 min in current study. The lack of change in transcripts 490 here may also reflect 1) a short exposure time to thermal stress that 491 may not have provided enough stress; 2) shortnose sturgeon may 492 use a different strategy to handle thermal stress (i.e., they may use 493 different heat shock proteins); and 3) proteins were available to mit494 igate the thermal stress, and did not require increased de novo tran495 scription. In addition, shortnose sturgeon is considered as octaploid 496 Q10 (8n) or dodecaploid (12n) (LeBreton and Beamish, 2004) and other 497 stress related genes or multiple isoforms may be constitutively 498 expressed to cope with acute thermal stress. Prolonged thermal ex499 posure used in other studies may trigger the upregulation of these 500 transcripts, to provide additional proteins for survival under high 501 temperature environment. 502 Many existing studies on CTMax of various fish species at heating 503 rates between 0.1 and 0.3 °C min−1 suggest heating rate has no impact 504 on the CTMax values (Iwama et al., 1999; Beitinger et al., 2000; An et al., 505 2010; Madeira et al., 2013; Vinagre et al., 2014). However, some studies 506 have reported molecular changes at very low heating rates (Iwama 507 et al., 1999; Madeira et al., 2013; Linares-Casenave et al., 2013; Vinagre 508 et al., 2014). Studies involving thermal cycling using various heating 509 rates have shown that the maximal MO2 and the MO2 at the end of 510 the experiment are not significantly affected by the heating rate 511 (Johnstone and Rahel, 2003; Tunnah et al., 2016). Thus, current research 512 suggests that the thermal tolerance of shortnose sturgeon and the asso513 ciated physiological/molecular responses may be closely related to the 514 thermal history (i.e., acclimation temperature), duration of the expo515 sure and how temperature approaches to the upper limit (e.g., linear in516 crease or thermal cycle), other than the rate of temperature increasing.

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