Polycyclic Aromatic Hydrocarbon and Hypoxia Exposures Result in Mitochondrial Dysfunction in Zebrafish

Abstract

Organisms are routinely subjected to a variety of environmental and chemical perturbations simultaneously. Often, multi-stressor exposures result in unpredictable toxicity that occurs through unidentified mechanisms. Here, we focus on polycyclic aromatic hydrocarbons (PAHs) and hypoxia, two environmental and physiological stressors that are known to co-occur in the environment. The aim of this study was to assess whether interactive mitochondrial dysfunction resulted from co-exposures of PAHs and hypoxia. Zebrafish embryos were co-exposed to non-teratogenic concentrations of an environmental PAH mixture and hypoxia beginning at 6 hpf for an acute period of 24 hours and afterwards were given either no recovery period, 45-minutes, 5-hours, or 18-hours of recovery time in clean conditions. Mitochondrial function and integrity were assessed through the use of both in ovo and in vitro assays. Hypoxia exposures resulted in drastic reductions in parameters relating to mitochondrial respiration, ATP turnover, and mitochondrial DNA integrity. PAH exposures affected ATP production and content, as well as mitochondrial membrane dynamics and lactate content. While PAH and hypoxia exposures caused a broad range of effects, there appeared to be very little interaction between the two stressors in the co-exposure group. However, because hypoxia significantly altered mitochondrial function, the possibility remains that these effects may limit an individual’s ability to respond to PAH toxicity and therefore could cause downstream interactive effects.

1. Introduction

Multi-stressor considerations are fundamentally important to the study of environmental health. In their natural environment, organisms are exposed to a suite of chemical contaminants, environmental perturbations, and physiological stressors. Frequently, these co-exposures result in unpredictable phenotypes and severity of toxicity (Holmstrup et al., 2010; Sokolova, 2013). Investigations into the mechanistic underpinnings of multi-stressor exposures are necessary to increase the environmental and physiological relevance of laboratory toxicity assessments (Holmstrup et al., 2010; Todgham and Stillman, 2013).

Here, we focus on an increasingly prevalent environmental stressor, hypoxia, and the ubiquitous class of organic compounds, polycyclic aromatic hydrocarbons (PAHs). These stressors regularly occur simultaneously in the environment, as with pollution of oxygen-depleted water bodies (Adcroft et al., 2010; Diaz and Rosenberg, 2008; Whitehead, 2013). Of the known reports on hypoxia and PAH mixture co-exposures, many have described additive and even synergistic toxicity for multiple developmental, molecular, reproductive, and transgenerational endpoints (Dasgupta et al., 2015, 2016; Fleming and Di Giulio, 2011; Hedgpeth and Griffitt, 2016; Jasperse et al., 2019; Negreiros et al., 2011; Simning et al., 2019). In juvenile and adult fish, the combination of stressors has also been found to result in reduced organism fitness and performance (Mager et al., 2018; Milinkovitch et al., 2019).

While the toxicological mechanisms for exposures to both stressors individually have been well-studied (Brette et al., 2014; Incardona et al., 2006, 2011; Richards, 2009; Timme-Laragy et al., 2007), there have been inconsistencies stemming from investigations into mechanistic underpinnings of interactions between the two. Many studies have focused on the potential for competition or cross-talk between the aryl hydrocarbon receptor (AHR) and hypoxia-inducible factor-1 (HIF-1) signaling pathways, with conflicting conclusions (see Vorrink and Domann, 2014). For example, the dominance of one pathway over another is not always apparent and can be altered depending on several factors, including AHR ligand, method of HIF-1a induction, experimental organism/cell line, and others. Previous work in our laboratory has also shown that synergistic toxicity results from combinations of PAH-type chemicals not expected to interact with hypoxia (i.e. fluoranthene and alpha-naphthoflavone, CYP1A inhibitors) in zebrafish larvae (Matson et al., 2008). The role of cross-talk between the AHR and HIF-1 pathways as a causal factor behind the resulting synergistic phenotypes has yet to be solidified and in addition, there may be other mechanisms underlying the toxicity associated with PAH and hypoxia co-exposures.

The overarching aim of this study was to explore under-examined physiological responses in which this compounding toxicity could occur, below concentrations which result in the well-studied teratogenic effects (Fleming and Di Giulio, 2011; Matson et al., 2008). Both PAH and hypoxia exposures cause mitochondrial dysfunction (i.e. disruption in energy production or metabolism) and a loss of mitochondrial integrity (i.e. interference with mitochondrial membrane dynamics, structure, or DNA) (Ballinger et al., 1996; Bansal et al., 2014; Ivanina et al., 2016; Jung et al., 2009; Li et al., 2003; Lindberg et al., 2017; Meyer et al., 2013; Solaini et al., 2010; Xia et al., 2004; Zhang et al., 2008). However, no known studies have focused on PAH-hypoxia interactions and the influence of co-exposures on mitochondrial function or bioenergetics in fish.

Bioenergetics endpoints are excellent indicators of the physiological significance of exposures and are sensitive to even slight alterations in organism fitness (Beyers et al., 2011; Sokolova et al., 2012). Any metabolic change could potentially be a consequence or cause of the interactive toxicity seen after co-exposures. Here, we hypothesized that due to the evident mitochondrial toxicity that results from both types of exposure, enhanced mitochondrial dysfunction could be an outcome observed during co-exposures of PAHs and hypoxia that do not cause overt teratogenicity. To test this hypothesis, we utilized zebrafish (Danio rerio), a model which allows for the unique opportunity to measure alterations in mitochondrial function in vivo directly after chemical or environmental perturbations (Kim et al., 2008; Stackley et al., 2011). We exposed zebrafish embryos to a non-teratogenic concentration of an environmental PAH mixture and hypoxia and assessed them for alterations in mitochondrial function and integrity. Separately, PAH and hypoxia exposures resulted in diverse mitochondrial responses. However, we observed limited evidence of mitochondrial interactions between the two stressors. Regardless, this work provides insights into the toxicological mechanisms and consequences underlying sub-teratogenic exposures to the individual stressors in vivo.

2. Materials and Methods

2.1. Fish Care and Handling

The Duke University Institutional Animal Care and Use Committee approved all care, handling, and techniques used during this work (Protocol # A139–16-06). Adult zebrafish (Ekkwill and MLS-EGFP) were maintained in the laboratory at approximately 28 °C with a 14:10 light:dark cycle. They were fed once per day with Artemia franciscana (Brine Shrimp Direct, Ogden, UT, USA) and once per day with Zeigler Adult Zebrafish Diet (Zeigler Bros., Inc., Gardners, PA, USA). During spawning, adults were placed in groups of 5 (2 males and 3 females) in breeding tanks. They were given one hour to breed after which embryos were collected and stored in an incubator at 28 °C until use. At approximately 5 hpf, embryos were screened for proper development and age-matched for exposures. Embryos were only used during exposures if they had a developed germ ring and were approaching the shield stage. This is crucial as developmental stage greatly influences cellular bioenergetics and embryonic oxygen consumption (Stackley et al., 2011).

2.2. Chemicals

Elizabeth River sediment extract (ERSE) was prepared previously according to the methods of Clark et al. (2013) and was characterized for PAH content by Fang et al. (2014). ERSE is a real-world, complex PAH mixture derived from creosote-contaminated sediment obtained from the Atlantic Wood Industries Superfund Site in the Elizabeth River (VA, USA; 36°48′27.2″ N, 76°17′38.1″ W). It was previously analyzed and found to contain approximately 5,073 ± 409 μg/L of 36 selected PAHs (Fang et al., 2014). Although the extract itself was not analyzed for presence of other chemicals or metals, the sediment from which it was derived was found to have limited levels of both 18 selected polychlorinated biphenyls and 8 metals (unpublished data). Dimethyl sulfoxide (DMSO), pronase, phenylthiourea (PTU), carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP), oligomycin, and sodium azide (NaN₃) were purchased from Sigma-Aldrich (St. Louis, MO, USA). Embryos were kept in 30% Danieau’s solution (58 mM NaCl, 0.7 mM KCl, 5 mM HEPES (pH 7.1–7.3), 0.4 mM MgSO₄, and 0.6 mM Ca(NO₃)₂). KCl, HEPES, and MgSO₄ were purchased from VWR International (Radnor, PA, USA), NaCl was purchased from Macron (Avantor Performance Materials Inc., Center Valley, PA, USA), tricaine was purchased from Thermo Fisher Scientific Inc. (Waltham, MA, USA), and Ca(NO₃)₂ was purchased from Alfa Aesar (Haverhill, MA, USA).

2.3. Embryonic Exposures and Staging

At 6 hpf, embryos were placed in pairs in 7.5 mL exposure media in glass scintillation vials (Wheaton, Millville, NJ, USA). Exposure media included a control solution (Danieau’s media) or a 0.5% (v/v) ERSE solution, a concentration that does not cause teratogenesis in zebrafish (Fleming and Di Giulio, 2011; Supplementary Table 1). Hypoxia exposures (5% O₂ air saturation, 2.56 ± 0.13 mg/L O₂ after 24-hour incubation) began at 6 hpf and were conducted in a Heracell VIOS 160i incubator (Thermo Fisher Scientific Inc.). To create and maintain oxygen conditions, nitrogen gas was bubbled into the sealed incubator. Air within the chamber reached 5% O₂ saturation within 45 minutes. Oxygen conditions in the exposure vials were monitored once every hour for 9 hours and then once every 5 hours until the 24-hour exposure time was complete (Supplementary Figure 1). Oxygen concentrations were measured with a Fibox 3 fiber optic oxygen probe and transmitter (PreSens Precision Sensing GmbH, Regensburg, Germany). Half of the embryos were placed in hypoxic conditions and half were placed in normoxic (7.64 ± 0.04 mg/L O₂ after 24-hour incubation) conditions. This resulted in 4 treatment groups: normoxia + control (CON), hypoxia + control (HYP), normoxia + 0.5% ERSE (ERSE), and hypoxia + 0.5% ERSE (H+E). Embryos were kept in exposure vials (2 embryos per vial) at 28 ˚C for 24 hours (until 30 hpf).

After the 24-hour exposure, embryos were removed from exposure conditions and either used immediately, flash frozen, or placed into small (3-inch diameter) glass petri dishes containing clean 65ppm solution or Danieau’s media (1 mL per embryo) for recovery periods. Time points included no recovery (0h), 45-minute recovery (45min), 5-hour recovery (5h), and 18-hour recovery (18h) periods. These periods correspond to 30 hpf, 30.75 hpf, 35 hpf, and 48 hpf, respectively. Due to calibration requirements of the Seahorse Extracellular Flux Analyzer, endpoints could not be collected at the 0h timepoint. The same is true for exposures of MLS-EGFP fish due to length of time required for dechorionation and plate setup.

Embryos were monitored at each recovery time point for developmental stage using the methods of Kimmel et al. (1995) as a guide. Assessments included presence of heartbeat and circulation, head trunk angle (HTA), yolk extension to yolk ball ratio (YE/YB), and standard length. Images of individual embryos were captured using a Nikon SMZ1500 and Digital Sight Camera (Nikon, Tokyo, Japan) and were measured with Fiji (ImageJ v2.0; Schindelin et al., 2012). Additionally, larvae were assessed for incidence of deformities including pericardial edema, yolk sac edema, yolk extension abnormalities, and delayed development.

2.4. Relative ATP Content and ADP/ATP Ratio Measurements

Measurements were taken using the ApoSENSOR™ ADP/ATP Ratio Assay Kit (Enzo® Life Sciences, Inc., Farmingdale, NY, USA). Methods were adapted for live zebrafish embryos from the kit instruction manual and Lu et al. (2011). All luminescence readings were measured with a FLUOstar® Optima spectrophotometer (BMG Labtech, Cary, NC, USA). Initial trials were performed to ensure that the ERSE solution and embryos alone were not auto-luminescent. A total of 2 replicate plates were completed per time point, with 5 wells per treatment in each plate (total n=10). Predicted values for additive interactions between hypoxia and ERSE in the H+E group were calculated by subtracting the observed control values for ATP content from both the observed hypoxia and ERSE values, then adding those together (i.e. (HYP-CON)+(ERSE-CON)=Expected). This provides us with a baseline mathematical expectation of additivity for comparisons to observed H+E values, although may not fully reflect biological additivity.

2.5. Embryonic Bioenergetic Assessment

Methods described below were adapted from Stackley et al. (2011). Using the Seahorse Extracellular Flux Analyzer (Agilent Technologies, Santa Clara, CA, USA), routine respiration of embryo pairs, reported as the oxygen consumption rate (OCR), was measured over a 250-minute period immediately after removal from exposures and placement into islet capture microplates. Each well contained 700 μL of normoxic 65 ppm ASW. Routine respiration measurements were recorded approximately once every 5 minutes (57 cycles of 1 minute mix, 1 minute wait, and 2 minute measure periods). Simultaneous measurements of extracellular acidification rate (ECAR), in zebrafish a means of quantifying Krebs Cycle activity, of embryo pairs were also taken under these conditions (Stackley et al., 2011). Based on the recovery time course, 3 time points were chosen throughout an extended recovery period to further analyze embryos for alterations in mitochondrial function and integrity due to exposures. For assessments with pharmacological agents, after the allotted recovery period, embryos were placed as pairs in islet plates with 525 μL of 65 ppm ASW. The plates were then immediately placed in the Seahorse instrument to begin trials with pharmacological agents. There were 3 replicate plates for routine OCR measurements, FCCP trials, and oligomycin trials and 6 for ECAR measurements, with 5 wells per treatment per plate (total n=15 and 30, respectively).

Table 1 describes the use of pharmacological agents and measurement specifics for bioenergetics testing. Table 2 describes the calculations necessary to quantify specific mitochondrial parameters. Supplementary Figure 2 depicts a representative trace of a control embryo during routine respiration measurements and after injections with FCCP, oligomycin, and NaN₃.

Table 1.

Pharmacological agents utilized during Seahorse Extracellular Flux Analyzer bioenergetics testing of zebrafish embryos.

Compound	Compound Mechanism of Action	Final Working Concentration	Measurement Cycles and Length
No addition	N/A	N/A	8 cycles; 1 minute mix, 1 minute wait, 2 minute measure
FCCP	Ionophore, un-coupler of oxidative phosphorylation, depolarization of mitochondrial membrane	2.5 μM	8 cycles; 1 minute mix, 1 minute wait, 2 minute measure
Oligomycin	ATP synthase inhibitor, blocks oxidative phosphorylation of ADP to ATP	9.4 μM	18 cycles; 1 minute mix, 1 minute wait, 2 minute measure
NaN3	Cytochrome c oxidase inhibitor, inhibits mitochondrial respiration	6.25 mM	25 cycles; 1 minute mix, 1 minute wait, 2 minute measure

Table 2.

Calculations completed to quantify mitochondrial and non-mitochondrial parameters of respiration.

Mitochondrial Parameter	Calculation
Routine Mitochondrial Respiration	Routine OCR minus NaN3inhibited OCR
OCR due to ATP Production	Routine OCR minus oligomycin-inhibited OCR
OCR due to Proton Leak	Oligomycin-inhibited OCR minus NaN3-inhibited OCR
Maximal Mitochondrial Respiration	FCCP-stimulated OCR minus NaN3-inhibited OCR
Spare Respiratory Capacity	FCCP-stimulated OCR minus routine OCR

2.6. Lactate Measurements

Sample preparation and analyses were conducted with instructions and materials from the Eton Bioscience L-Lactate Assay Kit I. For each recovery time point, embryos were placed as groups of 5 into 1.7 mL Eppendorf tubes. Fresh tissue was homogenized for 30 seconds with a hand-held homogenizer in 75 uL of 80% ethanol and flash frozen until analyzed for lactate content in duplicate (total n=10–12 per treatment). The assay was performed per manufacturer instructions (“solid sample” protocol) and lactate concentration was calculated using a standard curve. Results are reported as fold change due to inconsistencies between replicate plates most likely attributed to the use of different aliquots of kit materials during experimentation.

2.7. Mitochondrial Genome Integrity Assessments

For each recovery timepoint, embryos were placed in groups of 15 into 1.7 mL safe-lock tubes and flash frozen for DNA. Either 3 or 4 replicates of 3 samples per treatment group were completed per time point (total n=12–15). DNA was extracted from each sample using the phenol chloroform extraction method (Bogdanović et al., 2013). DNA content was quantified using a NanoDrop® ND-1000 Spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA) and then Quant-It™ PicoGreen™ dsDNA reagent (Thermo Fisher Scientific). Based on these measurements, samples were diluted to 3 ng/uL DNA in 1 x TE buffer (10 mM TrisHCl, pH 8.0 and 1 mM disodium EDTA). Mitochondrial DNA (mtDNA) damage and relative mitochondrial copy number were analyzed using the long amplicon-polymerase chain reaction (LA-PCR) method, described in detail in Gonzalez-Hunt et al. (2016). This assay is designed to measure gene-specific damage through targeted amplification of a “long” DNA fragment (approximately 10.3 kb). DNA that contains strand breaks, adducts or other types of lesions are amplified less due to mitochondrial DNA polymerase encountering them during the amplification process (Hunter et al., 2010). DNA damage of treated samples relative to control samples was quantified as lesion frequencies per 10 kb DNA through application of the Poisson distribution and normalization to mitochondrial genome copy number, obtained through QPCR amplification of a “short” DNA fragment (approximately 200 bp). Table 3 reports the primer sequences used during the long and short amplification processes, some of which have been published previously (Hunter et al., 2010; Gonzalez-Hunt et al., 2016).

Table 3.

List of primers used during the DNA damage and copy number assessments.

Target Amplicon	Forward Primer Sequence (5’−3’)	Reverse Primer Sequence (5’−3’)
Long Mitochondrial DNA Fragment	TTAAAGCCCCGAATCCAGGTGAGC	GAGATGTTCTCGGGTGTGGGATGG
Short Mitochondrial DNA Fragment	CGTTTACCCCAGATGCACCT	GTGCGATTGGTAGGGCGATA
Short Nuclear DNA Fragment	TGGATACCTGACCGAGAGCT	AGACAACTCTTACGGCTGGC

2.8. Mitochondrial Content

Assays utilized the mitochondrial localization sequence-enhanced green fluorescent protein (MLS-EGFP) transgenic zebrafish line which were genetically modified to tag cytochrome c oxidase subunit VII with enhanced GFP (Kim et al., 2008). Embryos were exposed as described above, with a supplement of 0.2 mM PTU to prevent pigmentation, and at the appropriate time point were placed into a 1 mg/mL pronase solution for dechorionation. Embryos were washed in clean media then were placed into a 0.02 mg/mL tricaine solution for anesthetization. Anesthetized embryos were transferred into individual wells of a glass-bottom, black 96-well plate for imaging. Fluorescence and brightfield images were recorded with a Keyence BZ-X710 microscope (Osaka, Japan). Ekkwill embryos were also imaged to control for any auto-fluorescence due to exposures. Total fluorescence was measured with Fiji software (ImageJ v2.0; Schindelin et al., 2012).

2.9. Data Analysis

All analyses were performed using GraphPad Prism 7.0b (GraphPad Software, Inc., La Jolla, CA, USA). All endpoints were analyzed for statistical significance using a three-way ANOVA to identify interactions between variables (see Supplementary Table 3 for results) and a two-way ANOVA and Tukey’s multiple comparisons test to assess differences between treatment groups within specific recovery time periods. The routine respiration time course was analyzed by an Area Under the Curve (AUC) analysis and subsequent two-way ANOVA and Tukey’s multiple comparisons test as a means to differentiate between the responses of treatment groups over an extended period of time, rather than by differences between individual time points. Each dataset was analyzed for significant differences between replicates using a one-way ANOVA and Tukey’s multiple comparisons test. This resulted in removal of only one dataset: replicate plate 6 of the 45min ECAR endpoint, where all four treatments produced different values from the same treatments in all 5 other replicate plates (all p<0.05), indicating a technical issue had occurred during this experiment. Datasets were also analyzed for normality using the Shapiro-Wilk test. If the data was found to follow a non-normal distribution, it was analyzed by the non-parametric Friedman’s test and Dunn’s test for multiple comparisons.

3. Results

3.1. Embryonic Staging

Regardless of treatment, all embryos displayed a heartbeat and weak circulation, as should be present by 30 hpf (equivalent to the 0h recovery time point). There were no observed trends across time points for HTA that indicated developmental delays or teratogenesis in embryos due to any treatment, although there were a few incidences of significantly altered measurements compared to controls (Supplementary Table 2). ERSE alone appeared to cause a slight increase in YE/YB ratio, although this was no longer apparent by 18h (Supplementary Table 2). No other consistent trends were reported for this endpoint. At all time points, hypoxia exposure resulted in reduced body length (p<0.05); however, there were no other indications of developmental delay in these embryos (as evidenced by no alterations in HTA, YE/YB ratio, and a visible heartbeat and circulation in all embryos), indicating that this is a phenotype of exposure rather than delay (Shang and Wu, 2004; Supplementary Table 2).

3.2. Relative ATP Content and ADP/ATP Ratio Measurements

Across the first 3 time points (0h-5h), there was a decrease in relative ATP content of HYP, ERSE, and H+E treatment groups compared to controls (Figure 1a). At all of these time points, the H+E groups had reduced ATP content compared to the CON groups (all p<0.05). The ERSE group was lower than CON at 0h (p<0.05). The H+E group had reduced ATP content compared to the HYP group at 0h (p<0.05). The predicted values for the H+E group support additive effects of hypoxia and PAH co-exposures at 45min and 5h (Supplementary Table 4); however, there were no statistically significant interaction terms after completion of a three-way ANOVA (Supplementary Table 3).

Figure 1.

Relative measurements of ATP and ADP in individual embryos across progressive recovery periods (0h, 45min, 5h, and 18h). a) ATP content, b) ratio of ADP/ATP. Different letters denote statistically significant differences between treatment groups within a recovery time point (p<0.05). Error bars indicate mean ± SEM (n=10 per treatment).

The only difference between the control and various treatment groups for the ADP/ATP ratio was at 0h (Figure 1b). The HYP and H+E groups had higher ADP/ATP ratios than the CON group (both p<0.05). There were no other differences at 45min, 5h or 18h, nor were there consistent trends in responses of ADP/ATP ratios across these time points.

3.3. Recovery of Oxygen Consumption Rates and Bioenergetic Assessments

As evident in the 250-minute respirometry time course, hypoxia exposures, with and without PAH co-exposures, caused a drastic reduction in OCR in zebrafish embryos, which recovered, first rapidly (0–45 minute period), then gradually over time (Figure 2). HYP and H+E treatments resulted in decreased AUC measurement of OCR over the 250-minute period compared to the CON and ERSE groups (all p<0.0001; Table 4). ERSE treatment resulted in slightly elevated AUC measurements, but the values were not statistically different from the CON group. The H+E group appeared to recover slightly faster than the HYP group over the 250-minute period which resulted in an elevated AUC measurement (p<0.05). For all treatment groups, there were increases in OCR visible over time, which is consistent with previous reports that embryonic OCR increases linearly throughout development (Stackley et al., 2011).

Figure 2.

Recovery of Oxygen Consumption Rates over Time. Oxygen consumption rate (OCR) of embryos over a 250-minute period immediately after removal from exposure conditions. Different letters indicate statistically significant differences between exposure groups (p<0.05). Error bars indicate mean ± SEM (n=15 per treatment and timepoint).

Table 4.

Results of area under the curve analyses for the recovery of oxygen consumption rates over time. Different letters indicate statistically significant differences between treatment groups (p<0.05).

Treatment	Area Under the Curve (pmol O2/embryo over a 250-minute period; mean ± SE)
Control	37,560.0 ± 792.1a
Hypoxia	26,174.0 ± 481.4c
0.5% ERSE	40,182.0 ± 1,074.0a
Hypoxia+ERSE	29,510.0 ± 668.0b

When compared with controls, exposure to hypoxia with and without ERSE resulted in altered total routine respiration, routine mitochondrial respiration, non-mitochondrial respiration, maximal mitochondrial respiration, spare respiratory capacity, OCR due to ATP turnover, and ECAR (Krebs Cycle activity) (all p<0.05; Figure 3). For the most part, drastic alterations due to hypoxia exposures were seen at the 45min period. These effects appear to normalize with lengthened recovery periods, and in some cases returned to levels indistinguishable from controls (i.e. total routine respiration, routine mitochondrial respiration, and ECAR; Figures 3a, b, and h). For the maximal mitochondrial respiration parameter, this trend was not observed; the only time point with alterations due to hypoxia exposures was 5h (Figure 3d).

Figure 3.

Changes in embryonic bioenergetics due to exposures and progressive recovery time (45min, 5h, and 18h recovery periods). Graphs represent OCR due to a) total routine respiration, b) routine mitochondrial respiration, c) non-mitochondrial respiration, d) maximal mitochondrial respiration, e) spare respiratory capacity, f) ATP turnover, g) proton leak, and h) ECAR. Different letters above bars indicate significant differences between treatment groups within a recovery time point (p<0.05). Error bars indicate mean ± SEM (n=15 per treatment and time point).

The H+E group only differed from the HYP group in measures of proton leak (Figure 3g), the only parameter where ERSE was the statistically significant factor at all three recovery periods (all p<0.01). Otherwise, when compared with controls, ERSE exposure only caused changes in OCR due to ATP turnover at 5h and 18h (both p<0.05; Figure 3f).

The only statistically significant interaction terms were found after analyses of routine mitochondrial respiration, with both the “oxygen level x PAH exposure” and “oxygen level × PAH exposure × time” factors highlighted as sources of variation (both p<0.05; Supplementary Table 3). This indicates that there may be interactions between PAH and hypoxia exposures that change over time and can be attributed to the combination of stressors rather than a single stressor alone. The variation of all other endpoints was attributed either to PAH exposure or hypoxia exposure as individual terms, indicating that there were unlikely to be interactions between PAH and hypoxia exposures leading to alterations in the responses of embryos for these endpoints.

3.4. Lactate Measurements

The HYP group had increased lactate content compared to the other treatment groups until the 18h recovery period (Figure 4; all p<0.05). ERSE exposures alone decreased embryonic lactate content compared to controls at the 0h and 18h time points (both p<0.05) and showed trending decreases at both 45min and 5h. The H+E groups appeared to have an intermediate response between the HYP and ERSE groups and never statistically differed from controls.

Figure 4.

Measurements of embryonic lactate content over progressive recovery periods (0h, 45min, 5h, and 18h). Different letters denote statistically significant differences between treatment groups within a recovery time point (p<0.05). Error bars indicate mean ± SEM (n=10–12 per treatment and timepoint).

3.5. Mitochondrial Genome Integrity

Except for the H+E group at 18h, none of the treatments caused alterations in mtDNA damage at any of the time points measured when compared to controls (Figure 5a). However, at both 45min and 5h, a similar trend of differences between the HYP and ERSE treatment groups were present, with ERSE causing higher mtDNA damage than hypoxia (p<0.05 at 45min) and the H+E group showing intermediate responses between the two. At 18h, the H+E group had a lower lesion frequency than all other treatment groups (all p<0.05); at this time point the “oxygen level x PAH exposure” interaction term was also significant (p<0.05; two-way ANOVA). At all time points, the HYP treatment group had higher copy number values than the CON and ERSE groups, (Figure 5b; all p<0.05). The H+E group had higher values than the CON group at all time points (all p<0.005) and the ERSE group at the first three time points (all p<0.05).

Figure 5.

Measurements of mitochondrial genome integrity over progressive recovery periods (0h, 45min, 5h, and 18h). a) mitochondrial DNA damage, b) relative mitochondrial copy number. Different letters denote statistically significant differences between treatment groups within a recovery time point (p<0.05). Error bars indicate mean ± SEM (n=12–15 per treatment and timepoint).

3.6. Mitochondrial Content

Hypoxia exposures reduced mitochondrial content of embryos across all recovery time points (Figure 6). However, the HYP group was only statistically different from controls at the 45min period (p<0.01). ERSE exposures alone appeared to slightly elevate mitochondrial content, although never above control levels.

Figure 6.

Measurements of mitochondrial content over progressive recovery periods (45min, 5h, and 18h). a) representative image of a fluorescent MLS-EGFP embryo at 35 hpf, b) mitochondrial content displayed as fold change relative to control embryos. Different letters denote statistically significant differences between treatment groups within a recovery time point (p<0.05). Error bars indicate mean ± SEM (n=15–20 per treatment and timepoint).

4. Discussion

4.1. PAH-Hypoxia Interactions

Initially, we hypothesized that co-exposures of PAHs and hypoxia would result in exacerbated mitochondrial toxicity, a potentially significant factor contributing to the overall toxicity of the combined exposures. Not only was there no evidence for this, there was also limited evidence for any interaction between the two stressors. Statistical interactions between hypoxia and ERSE were only observed with assessments of embryonic ATP content (Figure 1a). For most other endpoints, results for H+E embryos appeared similar to the outcomes of the individual treatment (HYP or ERSE) which caused the most drastic changes. For example, PAH exposures resulted in increases in OCR due to proton leak at all time points, which was seen in the H+E and ERSE groups, but not the HYP group (Figure 3g). Interestingly, effects that coincide rather than interact appear to predominately result from co-exposures.

4.2. Effects of PAHs on Mitochondrial Function

As we previously described in Fundulus heteroclitus embryos exposed to ERSE during development (Lindberg et al., 2017) and similar to what others have found after exposures to individual PAHs and PAH mixtures in a variety of fish species (compiled in Klinger et al., 2015), ERSE exposures stimulated routine respiration during recovery periods (Figure 2, Figure 3). An increase in routine metabolism could be due to a variety of factors, including inefficiencies in energy production and glycolysis or enhanced energy demand that may be necessary for xenobiotic metabolism. Here, both may play a role: increased OCR due to proton leak (Figure 3g) and decreased OCR due to ATP production (Figure 3f) indicate greater uncoupling of ATP synthesis with oxygen consumption, resulting in inefficient mitochondrial respiration (Divakaruni and Brand, 2011). Decreased lactate content (Figure 4), a result also seen after PAH exposures in cells (Gurbani et al., 2013) and whole organisms (Gagnon & Holdway, 1999; Jones et al., 2008), potentially caused by inhibition of lactate dehydrogenase (Gagnon & Holdway, 1999), may also indicate inefficiencies in anaerobic energy production. Meanwhile, declines in ATP content (Figure 1a) were apparent prior to alterations in OCR due to ATP production (Figure 3f). Embryos were possibly depleting available ATP stores to react to PAH exposures, likely through enhanced metabolic signaling pathways and enzyme activity (Klinger et al., 2015).

Interestingly, most of the effects observed after PAH exposures occurred during later stages in recovery (i.e. 5h and 18h), potentially due to PAH metabolism. Reactive metabolites may drive the toxicity associated with decreases in OCR due to ATP production and increases in mtDNA damage (Figures 3f and 6, respectively; Knecht et al., 2013; Meyer et al., 2013); parent compounds may drive the toxicity associated with increases in OCR due to proton leak (Figure 3g; Meyer et al., 2013). Although cytochrome P450s (CYPs) and other xenobiotic metabolizing enzymes are expressed in zebrafish embryos as early as 3 hpf, their activities are inconsistent throughout development (Goldstone et al., 2010). It is possible that some PAHs taken in by embryos during exposures were not metabolized until after 31 hpf, and therefore could not immediately cause overt toxicity.

4.3. Effects of Hypoxia on Mitochondrial Function

Hypoxia exposures resulted in alterations in almost every endpoint measured during experimentation. Zebrafish embryos responded similarly to other fish upon exposure to hypoxia-reoxygenation events, with corresponding increases and decreases in respiration dependent on oxygen levels and length of time in oxygenated conditions (Figure 2; Hughes, 1973), as well as increases in lactate content (Figure 4; Dunn & Hochachka, 1986; Genz et al., 2013; Richards et al., 2007; Speers-Roesch et al., 2012). However, unlike in adult fish (e.g. Genz et al., 2013), the respiration rates of hypoxia-exposed embryos never exceeded that of normoxic fish during recovery periods, indicating that embryos likely do not have the same capacity or kinetics to increase oxygen uptake compared to adults (Papandreou et al., 2006).

A reduction in both ECAR, here an indication of Krebs Cycle activity (Stackley, et al., 2011), and OCR values during and immediately after hypoxia exposures suggests that these embryos were in a quiescent state and were undergoing a gradual recovery from metabolic rate suppression (Boutilier and St-Pierre, 2000; Sokolova, 2013). However, they did appear to have the capacity to function at optimal levels, similar to CON embryos, as evident with measures of maximal respiration (Figure 3d). Similar to what has been found in cell lines exposed to hypoxia (Zhang et al., 2008), hypoxia-exposed embryos appeared to be bioenergetically operating at a level far lower than their biological limits (Brand & Nicholls, 2011).

With these short exposure times, routine respiration returned to levels indistinguishable from controls by 18h of recovery, although non-mitochondrial respiration rates did not. Likely, most of the newly-available oxygen was preferentially shunted towards oxidative phosphorylation over other cellular processes. This may be important considering how many xenobiotic metabolizing enzymes, including CYPs, require oxygen to function, and may play a role in PAH-hypoxia interactions in fish, although this was not further investigated here.

Considering previous findings (Dasgupta et al., 2016; Hammond and Giaccia, 2004; Lushchak et al., 2001; Wang et al., 2000), the lack of mtDNA lesions seen in hypoxia- and co-exposed embryos was initially surprising (Figure 5a). However, the substantial increases in mtDNA copy number (Figure 5b) likely explain our results. It is possible that these mild hypoxia exposures stimulated replication of new, undamaged mitochondrial genomes, thus resulting in a lack of induced lesions. The increases in mtDNA copy number, as well as the capacity of the organisms to induce maximal respiration to the same extent as control embryos, led us to hypothesize that hypoxia-exposed embryos were undergoing mitochondrial biogenesis. However, measures of mitochondrially-targeted protein expression actually indicated a decline in mitochondrial density within hypoxia-exposed embryos (Figure 6). These findings are in line with those of others (Solaini et al., 2010; Zhang et al., 2008) and potentially point to enhanced mitochondrial fragmentation in hypoxia-exposed embryos.

5. Conclusions

Although there was little evidence of interactions between hypoxia and PAHs in exposed embryos that resulted in enhanced mitochondrial dysfunction, embryos in the co-exposure group displayed adverse effects due to both types of exposure. Hypoxia exposures resulted in a sweeping range of effects, from alterations in mitochondrial genome copy number, to drastic reductions in mitochondrial respiratory parameters. PAH exposures appeared to have much more targeted effects on mitochondrial membrane dynamics which could affect ATP production. Effects caused by each individual stressor were seen in the co-exposure group. This indicates that they are more bioenergetically vulnerable than if they had been exposed to only one stressor, a key factor when considering the energetic requirements of developing organisms. Additionally, the lack of interaction seen in the endpoints measured here does not necessarily mean that mitochondrial dysfunction does not play a role in the toxicity seen during co-exposures. Hypoxia may cause such drastic changes in mitochondrial function that it affects how organisms respond to ERSE, potentially through toxification of PAHs. Future work will supplement our findings here as well as focus on other potential mechanisms of toxicity.

Highlights.

• Hypoxia exposures had severe effects on mitochondrial and non-mitochondrial respiration

• Hypoxia exposures increased mitochondrial genome copy number, but not mitochondrial content

• PAH exposures reduced lactate content and appeared to influence mitochondrial membrane dynamics

• Co-exposed embryos exhibited phenotypes due to both PAH and hypoxia exposures

• There were very few visible interactions between the two stressors

Abbreviations

PAHs

polycyclic aromatic hydrocarbons

FCCP

carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone

NaN₃

sodium azide

hpf

hours post fertilization

HTA

head trunk angle

YE/YB

yolk extension to yolk ball ratio

OCR

oxygen consumption rate

ECAR

extracellular acidification rate, a measure of Krebs Cycle activity

mtDNA

mitochondrial DNA

LA-QPCR

long amplicon-polymerase chain reaction

AUC

area under the curve

RLU

relative light units

ANOVA

analysis of variance

PTU

phenylthiourea

MLS-EGFP

mitochondrial localization sequence-enhanced green fluorescent protein

CON

refers to the control treatment group

HYP

refers to the hypoxia-alone treatment group

ERSE

refers to the 0.5% ERSE-alone treatment group

H+E

refers to the hypoxia+0.5% ERSE co-exposed treatment group

0h

refers to the groups given no recovery period

45min

refers to the groups given a 45-minute recovery period

5h

refers to the groups given a 5-hour recovery period

18h

refers to the groups given an 18-hour recovery period