Mechanistic Toxicity Tests Based on an Adverse Outcome Pathway Network for Hepatic Steatosis

Abstract

Risk assessors use liver endpoints in rodent toxicology studies to assess the safety of chemical exposures. Yet, rodent endpoints may not accurately reflect human responses. For this reason and others, human-based in vitro models are being developed and anchored to adverse outcome pathways to better predict adverse human health outcomes. Here, a networked adverse outcome pathway-guided selection of biology-based assays for lipid uptake, lipid efflux, fatty acid oxidation, and lipid accumulation were developed. These assays were evaluated in a metabolically competent human hepatocyte cell model (HepaRG) exposed to compounds known to cause steatosis (amiodarone, cyclosporine A, and T0901317) or activate lipid metabolism pathways (troglitazone, Wyeth-14,643, and 22(R)-hydroxycholesterol). All of the chemicals activated at least one assay, however, only T0901317 and cyclosporin A dose-dependently increased lipid accumulation. T0901317 and cyclosporin A increased fatty acid uptake, decreased lipid efflux (inferred from apolipoprotein B100 levels), and increased fatty acid synthase protein levels. Using this biologically-based evaluation of key events regulating hepatic lipid levels, we demonstrated dysregulation of compensatory pathways that normally balance hepatic lipid levels. This approach may provide biological plausibility and data needed to increase confidence in linking in vitro-based measurements to chemical effects on adverse human health outcomes.

Hepatic steatosis is a pathological condition that can lead to liver failure, altered xenobiotic metabolism (Donato et al., 2006; Gomez-Lechon et al., 2009), and sensitization to endocrine disruption (Berlanga et al., 2014; Koo, 2013) including thyroid disease (Pacifico et al., 2013). Risk assessors and regulators (Kaiser et al., 2012) currently use a number of liver endpoints, including hepatic steatosis, from epidemiological and rodent toxicology studies to assess the safety of exogenous chemical exposures. Critically, rodent physiologic responses to chemical perturbation do not always reflect human responses. Differences in rodent behaviors, endocrine signaling, and metabolism confound attempts to translate findings from rodent toxicity studies to human pathophysiology (Bergen and Mersmann, 2005). Furthermore, traditional toxicity testing approaches are time and resource intensive and create a bottleneck to test the health effects of nearly 100000 registered chemicals of unknown toxicity (Toxic Substances Control Act registry, https://www.epa.gov/tsca-inventory/how-access-tsca-inventory; last accessed June 22, 2017). For these reasons, alternative in vitro human cell models are being developed to accelerate the pace of the chemical evaluation process. For example, the U.S. EPA launched the Toxicity Forecaster (ToxCast) program in 2007 (Dix et al., 2007). ToxCast uses high-throughput screening (HTS) and computational toxicology to evaluate large sets of chemicals for effects on a wide range of biological activity.

As the in vitro HTS toxicity testing paradigm has evolved to address the need to rapidly evaluate large numbers of chemicals, so have evaluations to frame this data in a biologically meaningful context. An example is the adverse outcome pathway (AOP) concept, which is related to the toxicity pathway and mode-of-action paradigms for toxicological risk assessment (Ankley et al., 2010). A primary driver of developing the AOP framework was to better interpret high-throughput data by connecting assay endpoints to a series of biological pathways that lead to an adverse outcome (AO) (Ankley et al., 2010; Villeneuve et al., 2014). We previously assembled an interconnected network of linear AOPs (ie, an AOP network) for hepatic steatosis that was characterized by 4 apical key events (KEs)—fatty acid uptake, efflux, synthesis, and oxidative metabolism (Angrish et al., 2016). Hepatic steatosis is not a direct chemical-receptor mediated effect and instead results from an imbalance of these interacting biological processes. For example, hepatic fat retention could result from both a chemical-mediated decrease in fatty acid breakdown by disrupting mitochondrial-mediated β-oxidative metabolism and decreasing very low-density lipoprotein (VLDL, a carrier of fatty acid-derived triglycerides) efflux, as observed in rodent liver with carbon tetrachloride exposure (Kaiser et al., 2012). Alternatively, an increase of fatty acid uptake alone, such as a transient increase in fatty acid in the blood due to a meal, may be balanced by normal responses mediated by the other compensatory mechanisms. Therefore, it may be important to evaluate the 4 KE of hepatic steatosis (fatty acid uptake, efflux, synthesis, and oxidative metabolism) to mechanistically understand the action of steatotic chemicals.

Given the large amount of scientific evidence on the mechanism, mode of action, potential serious human health impacts, and increased risk to chemical exposures associated with hepatic steatosis, we developed biologically based toxicity tests for steatotic KEs that, when unbalanced, can lead to the initiation and progression of the disease. Specifically, here we describe the development of assays for lipid uptake, lipid efflux, fatty acid oxidation (FAO), lipid accumulation, cytotoxicity, and gene expression. These assays were evaluated in the HepaRG human hepatocyte. This cell model overcomes the serious limitations of the commonly used hepatoma cell line, HepG2, that are metabolically incompetent, lack chemical transport function, and fail to adequately express transcription factors (Jackson et al., 2016; Kanebratt and Andersson, 2008; Tolosa et al., 2016). A defined chemical selection was used to evaluate the KE assay suite (Table 1). Three of the test chemicals had reported steatotic effects in various models: T0901317 (sulfonamide and liver X receptor [LXR] agonist), cyclosporin A (potent immunosuppressant, also a known cholestatic compound), and amiodarone (antiarrhythmic medication, also known to cause phospholipidosis in hepatocytes). Three other compounds were chosen that did not have strong evidence for hepatic steatosis effects but exhibited lipid metabolism-associated nuclear receptor activation that could perturb the test KE assays: 22(R)-hydroxycholesterol (an endogenous LXR agonist), Wyeth-14643 (a hepatotoxic peroxisome proliferator activated receptor [PPAR] α agonist), and troglitazone (a hepatotoxic PPARc agonist). The overall goal for developing a networked AOP-driven suite of assays for hepatic steatosis was to demonstrate a strategic in vitro chemical exposure evaluation by measuring KEs of compensatory pathways that, when sufficiently disrupted, could lead to an AO.

Table 1.

Test Chemical List

Chemical	Tested [mM]	General Description	Liver Mode of Action	Citations
amiodarone hydrochloride	5.0, 15.8, 50a	Antiarrhymatic medication used to treat cardiac arrhythmia. Steatotic and hepatotoxic.	Possible HNF4a, PPARa, PPARc, PXR, RXRa	Antherieu et al. (2011), Fromenty and Pessayre (1997), Tolosa et al. (2016), Vitins et al. (2014)
cyclosporin A	5.0, 15.8, 50	Immunosuppressant drug. Steatotic and hepatotoxic.	Oxidative Stress, mitochondrial dysfunction.	Donato et al. (2012), Rezzani (2006), Tolosa et al. (2016), Van Summeren et al. (2011)
T0901317	1.58, 5.0, 15.8	Upregulates genes associated with cholesterol and bile acid synthesis. Synthetic LXR agonist. Steatotic.	LXR, FXR, PXR	Grefhorst et al. (2002), Houck et al. (2004)
Troglitazone	1.0, 3.16, 10	Antidiabetic and anti-inflammatory drug removed from the market because it caused drug induced liver injury. Hepatotoxic.	PPARa and PPARc	Chojkier (2005), Jaeschke (2007)
Wyeth-14,643	5.0, 15.8, 50	Chemical developed by the pharmaceutical industry to lower serum cholesterol. Not used clinically. Hepatotoxic.	PPARa	Larter et al. (2012), Woods et al. (2007)
22(R)hydroxycholesterol	5.0, 15.8, 50	Endogenous steroid hormone biosynthetic intermediate and LXR agonist. Has not been observed to be steatotic nor hepatotoxic.	LXR, FXR	Deng et al. (2006), Hessvik et al. (2012), Larter et al. (2012), Woods et al. (2007)

^aOvert toxicity observed by microscopy and cytotoxicity assay.

MATERIALS AND METHODS

Chemicals and reagents.

Chemicals and test concentrations are listed in Table 1 and were purchased from Sigma-Aldrich (St Louis, Missouri). Oleic acid (CAS-No. 112–80-1), fatty acid free bovine serum albumin (CAS-No. 9048–46-8), Nile Red (CAS-No. 7385–67-3), and Hoechst 33342 (CAS-No. 23491–52-3) were purchased from Sigma-Aldrich. Chemical stock solutions were prepared in DMSO and diluted by serial 1/2 log increments. Oleic acid was dissolved in chloroform at 50mg/ml. The fatty acid synthase (FASN, Cat no. 31889) and vinculin rabbit (Cat no. 13901) antibodies were purchased from Cell Signaling Technology (Beverly, MA). BCA protein Assay kit (Cat no. 232250) was purchased from Thermo Scientific (Rockford, Illinois).

Cell culture and chemical treatment.

NoSpin HepaRG™ cells were purchased from Triangle Research Labs (Durham, North Carolina), thawed, suspended in thawing and plating medium (MH100 NoSpin HepaRG Base Medium and HepaRG™ Base Medium Supplement, Cat no. MH100; NoSpin HepaRG™ Media Thawing and Plating Additive, Cat no. MHTAP; Triangle Research Labs), and seeded at a density of 100000 cells per well into 96-well black wall, clear bottom tissue culture plates. Cells were cultured at 5% CO₂, 37°C and 85% humidity, and the medium was renewed every 48h according to the assay conditions described below. Cells were used in the free fatty acid uptake/fatty acid efflux assays or 200 ml RNeasy Lysis Buffer (RLT) lysis buffer (Qiagen RNeasy Kit; Valencia, CA) was added to each well and the plate was stored at −80°C until RNA isolation.

Lipid accumulation assay.

On the fifth day of culture, cells were preloaded with lipid by supplementing thawing and plating media with 50 μM oleic acid conjugated to bovine serum albumin at a molar ratio of 6:1 and incubated for an additional 24 h. On the sixth day of culture, cells were treated with chemical stocks (1000×) diluted in cell culture media and incubated at 5% CO₂, 37°C, and 85% humidity for 48h. After treatment the medium was removed and reserved at −80°C for lactate dehydrogenase (LDH) cytotoxicity and fatty acid efflux assays (Figure 1). Cells were washed with phosphate-buffered saline (PBS), fixed with 4% paraformaldehyde in PBS (sc-281692, Santa Cruz Biotechnology, Inc., Dallas Texas) for 20min, and washed twice with PBS. Nile Red (1mg/ml in methanol; to selectively stain for lipid accumulation) and Hoechst 33342 (5mg/ml in water; to stain for nuclei as a normalization metric) were diluted 1000× in PBS. One-hundred microliters of the mixture were added to each well, and the plate was incubated for 15min in the dark on a shaker. The plate was washed twice with 200 μl PBS. One-hundred microliters PBS were added to each well and the excitation/emission 510/580, 552/636, and 350/461nm were measured on a CLARIOstar microplate reader (BMG Labtech, Inc., Cary, North Carolina). Data were analyzed as described below. Images were also captured using a Nikon Eclipse Ti Microscope equipped with a 20× long objective, yellow fluorescent protein (excitation 500/20nm, DM 515nm, emission 535/30nm), tetramethylrhodamine (excitation 540/25nm, emission 605/55nm), and 4’,6-diamidino-2-phenylindole (excitation 360/40nm, emission 460–50nm) fluorescence filter cubes, and an Andor Zyla sCMOS camera. Images were acquired under the control of NIS-Elements software (Nikon Instruments, Inc., Melville, New York).

Figure 1.

Representative images of HepaRG cells treated with Wyeth-14,643 (50, 15.8, 5.0 μM), T0901317 (15.8, 5.0, 1.58 μM), troglitazone (10.0, 3.16, 1.0 μM), cyclosporin A (50, 15.8, 5.0 μM), 22(R)hydroxycholesterol (50, 15.8, 5.0 μM), amiodarone (50, 15.8, 5.0 μM), DMSO, and untreated. Fluorescence of Nile Red (to selectively stain for phospholipid rich environments (red) or lipid droplets (yellow); and Hoechst 33342 to stain for nuclei (blue) are included, 20× magnification.

LDH cytotoxicity assay.

Cytotoxicity was evaluated by LDH cytotoxicity assay (88953, ThermoFisher Scientific, Waltham, Massachusetts) according to the manufacturer’s protocol with slight modification. Cell culture medium was thawed and diluted 1:4 with plating media. Fifty microliters each of diluted test sample was mixed with 50 μl Reaction Mixture (included with kit) and incubated in the dark for 30min. Fifty microliters of Stop solution was added to each well and the absorbance of 490 and 680nm read on the CLARIOstar microplate reader. The background signal was removed (A490-A680) and test samples with overt cytotoxicity (see Supplementary Figure 1) were excluded from further analysis.

Fatty acid efflux assay.

Fatty acid efflux was inferred from apolipoprotein B100 (APOB100) levels. APOB100 is exclusively synthesized in the liver of humans and functions as a scaffold for lipids effluxed from the liver in VLDL particles. On the day of the assay, media was thawed and APOB100 levels were measured from 1:20 diluted media by ELISA according to the manufacturer’s protocol (Human Apolipoprotein B ELISA kit, Cat no. ab108807; Abcam, Cambridge, Massachusetts). Sample concentration (reported in mg/ ml) was calculated from the applied standard curve and corrected for dilution. Data were analyzed as described below.

Free fatty acid uptake assay.

Free fatty acid uptake assays (ab176768; Abcam, Cambridge, Massachusetts, USA) were performed according to the manufacturer’s protocol with a slight modification. On the sixth day of culture, lipids were depleted for 1h by incubating cells in completed base medium with MHIND supplement (Triangle Research Labs, LLC, Durham, North Carolina). Cells were treated with chemical stocks diluted in cell culture media and incubated at 5% CO₂, 37°C, and 85% humidity. After 2h of chemical treatment, cells were treated with a proprietary dodecanoic acid fluorescent fatty acid substrate (TF2-C12) in assay buffer and incubated an additional 2h at 5% CO₂, 37°C, and 85% humidity. The excitation/emission 485/515nm was read on a CLARIOstar monochromator.

FAO assay.

The oxygen consumption rate (OCR) due to oxidation of exogenous fatty acids was measured using a Seahorse XFe96 extracellular flux analyzer (Agilent Technologies, Santa Clara, California). HepaRG cells were suspended in thawing and plating media and seeded into collagen coated 96-well Seahorse microplates at 20000 cells/well (identified as the optimal cell density given the cell type with Seahorse cell culture plate well size and oligomycin/ carbonyl cyanide 4-[trifluoromethoxy] phenylhydrazone [FCCP] concentrations used). Media was renewed every 48h and after 96h culture (24 h before the assay) lipids were depleted by renewing with substrate limited media (Dulbecco’s Modified Eagle’s Medium supplemented with 0.5mM glucose, 1mM GlutaMAX, 0.5mM carnitine and 1% FBS, pH 7.4). Twenty-four hours later, chemicals were diluted in FAO assay media (KHB: 111mM NaCL, 4.7mM KCl, 1.25mM CaCl₂, 2mM MgSO₄, 1.2mM NaH₂PO₄) supplemented with 2.5mM glucose, 0.5mM carnitine, and 5mM HEPES on the day of the assay and adjusted to pH 7.4 at 37°C. Cells were washed one time with the FAO assay media and 135 μl assay media was added. Cells were degassed for 45min in a 37°C nonCO₂ incubator. The cell assay cartridge was loaded as follows: 15 μl 10× chemical stock (15.8 μM final) or the negative control etomixir (40 μM final) into Port A, 20 μl oligomycin (3.0 μM final) into Port B, 22 μl FCCP (2.5 μM final) into Port C, and 25 μl rotenone/antimycin A (AA) mix (2 or 4 μM final, respectively) into Port D. The cartridge was loaded into the flux analyzer and calibrated. Immediately before beginning the assay, 30 μl XF Palmitate-bovine serum albumin (BSA) FAO substrate or BSA control was added to the appropriate wells. The plate was loaded into the XFe analyzer and equilibrated for 12min. The run protocol was as follows: basal measurements, 3 repetitions of (mix 3min, wait 0min, measure 3min); inject port A, 3 repetitions of (mix 3min, wait 0min, measure 3min); inject port B, 3 repetitions of (mix 3min, wait 0min, measure 3min); inject port C, 3 repetitions of (mix 3min, wait 0min, measure 3min); inject port D, 3 repetitions of (mix 3min, wait 0min, measure 3min). There were a total of 15 time points and the assay duration was 1h and 42min. The data were baselined to the median rotenone/AA measurement at time point (T)15 of DMSO controls and scaled from 0 to 100. The basal threshold at T6 (after chemical injection) and maximal response at T12 (after FCCP injection) were analyzed as described below with the exception that the response threshold was set at 3 (basal, T6) and 1.5 (maximal response, T12) times the baseline median absolute deviation (BMAD) of vehicle-only (DMSO) controls.

Protein studies.

Total protein was extracted from HepaRG cells treated with DMSO, cyclosporin A (5, 15, and 50 μM), and T0901317 (1.58, 5, and 15.8 μM) in RIPA buffer and quantitated using the BCA protein assay kit. RIPA lysis buffer (Cat no. 040–483), Rabbit Master Kit for proteins between 66 and 440kDa (Cat no. PS-MK20), Wes capillary immunoassay instrument, and Compass analysis software were purchased from Protein Simple (San Jose, California). Protein levels of FASN (molecular weight of 273kDa) and vinculin (molecular weight of 124kDa) were measured from homogenates using Wes™ instrument-based capillary nano-immunoassay (ProteinSimple) according to the manufacturer’s protocol. Briefly, protein and antibody concentrations were optimized such that 0.200 and 0.075 μg/μl FASN and vinculin, respectively, and 1:50 primary antibody dilution were used in the assay. Vinculin was included as a loading control. Protein samples were heated at 95°C for 5min and then labeled with fluorescent master mix. Labeled protein samples, primary antibodies, and other reagents included in the rabbit master kit were loaded into a plate. The plate and the accompanying 25-well capillary were inserted into the Wes™ instrument and processed for 3h. Using the Compass analysis software, the peak area ratio of the FASN/vinculin were used to determine FASN protein levels in each sample. Data are reported as fold change (FC) relative to DMSO controls with n = 3.

Gene expression studies.

On the fifth day of culture, cells were preloaded with lipid by supplementing thawing and plating media with 50 μM oleic acid conjugated to bovine serum albumin at a molar ratio of 6:1 and incubated for an additional 24 h. On the sixth day of culture, cells were treated with chemical stocks (1000×) diluted in cell culture media and incubated at 5% CO₂, 37°C, and 85% humidity for 48 h. After treatment the medium was removed, 200 μl RLT lysis buffer was added, and samples were lysed by freeze thawing. An additional 150 μl RLT lysis buffer was added, and total RNA was isolated according to the manufacturer’s protocol with an additional on-column DNA digestion Qiagen® RNeasy® Mini Kit. Isolated RNA was dissolved in nuclease-free water, quantitated (A₂₆₀), and assessed for purity by determining the A₂₆₀/A₂₈₀ ratio using the NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington, Delaware), accepting ratios of >1.9. Quantitative reverse transcriptase PCR (RT-qPCR) of target genes was performed. Briefly, 100ng of total RNA was reverse transcribed using iScript™ cDNA Synthesis Kit as described by the manufacturer (Bio-Rad; Hercules, California). The cDNA was used as template in a 12 μl PCR reaction containing 1× PrimePCR™SYBR® Green Assay primers and SsoAdvanced™ Universal SYBRVR Green Supermix set-up using the QIAgility robotic workstation (Qiagen) for automated PCR setup. PrimePCR™SYBR® Green Assay primers included DGAT1 (assay qHsaCID0006734), DGAT2 (assay qHsaCED0036324), APOB (APOB100; assay qHsaCID0016796), CD36 (assay qHsaCID0011828), ACACA (assay qHsaCED0038035), PPARG (assay qHsaCID0011718), SREBF1 (assay qHsaCID0010452), FASN (assay qHsaCIP0026813), MTTP (assay qHsaCID009236), PP1A (assay qHsaCED0038620), NRL13 (assay qHsaCID0013633), CPT1A (qHsaCID0023721), GAPDH (assay qHsaCEP0041396), ACTB (assay qHsaCEP0036280), HPRT (assay qHsaCID0016375), and TBP (assay qHsaCID007122). PCR reactions were performed with the CFX384 Touch™ RT PCR Detection System according to the manufacturer’s protocol (Bio-Rad; Hercules, Californina). cDNAs were quantified according to the 2^−ΔΔCT method (Livak and Schmittgen, 2001) and normalized to the geometric mean of GAPDH, ACTB, and HPRT reference genes.

Data analysis.

All data were analyzed in R (R 3.1.1; R Foundation for Statistical Computing, Vienna, Austria). Fatty acid efflux, fatty acid uptake, and lipid accumulation data were normalized by setting the FC equal to the sample response over the median DMSO value. The response threshold was set at log₂FC ≥ 3 times the BMAD of the vehicle-only (DMSO) controls (Judson et al., 2015; Karmaus et al., 2016). Supplementary Materials 1–15 contained all raw assay results and R code for analysis of this data was provided in Supplementary File 16. Data were plotted in GraphPad Prism 6 (La Jolla, California) or Microsoft Excel (Redmond, Waltham) software.

RESULTS

Cell Viability

LDH is a cytosolic enzyme that is commonly used as an indicator of cytotoxicity. Overt cytotoxicity was not observed with the chemical concentrations tested, except with 50 μM amiodarone (see Supplementary Figure 1 and Supplementary Material 6) and this concentration was not further evaluated. These results were consistent with loss of nuclear staining by Hoechst 33342 in 50 μM amiodarone treated cells (Figure 1).

Hepatocellular Lipid Accumulation

Lipid accumulation was assessed by staining fixed cells with Nile Red and observed in hepatocyte-like cells (Figure 1). Nile Red is a lipophilic dye with emission spectra that shifts toward the blue spectrum as hydrophobicity increases (Greenspan and Fowler, 1985). In the presence of cell membranes and other phospholipid rich regions Nile Red emits red fluorescence (628 nm), whereas Nile Red emits yellow fluorescence (540 nm) in triglyceride rich lipid droplets. This shift in fluorescence was clearly seen in cells exposed to T0901317 and cyslosporin A. T0901317 caused marked perinuclear lipid droplet accumulation (yellow droplets, Figure 1). Cyclosporin A caused accumulation of both triglyceride rich lipid droplets and phospholipids that displaced the nucleus (yellow and red staining, Figure 1). Lipid accumulation was not observed by other chemical treatments and significant cell death was observed with an exposure of 50 μM amiodarone (Supplementary Figure 1). Quantitative analysis revealed that T0901317 (1.58, 5, and 15.8 μM) and cyclosporin A (15.8 and 50 μM) increased lipid droplet accumulation above the response threshold in a concentration responsive manner (Figure 2A). T0901317 increased phospholipid above the response threshold only at the highest concentration (15.8 μM), and cyclosporin A increased phospholipid at 15.8 and 50 μM in a concentration-responsive manner. Although near the established thresholds, Wyeth-14643 increased phospholipids at the highest dose, and 22-(R)-hydroxycholesterol decreased phospholipid levels at the highest and lowest concentrations tested (Figure 2B). Interestingly, 15.8 μM amiodarone induced the formation of red lipid droplets that displaced the nucleus (Figure 1). Although quantitative analysis demonstrated the effect did not exceed the response threshold, our observations were consistent with previous reports of amiodarone inducing phospholipidosis in HepaRG cells (Antherieu et al., 2011).

Figure 2.

Key biological event assessment in HepaRG cells utilizing assays for lipid droplet accumulation (A), phospholipid accumulation (B), fatty acid uptake (C), and fatty acid efflux (D). HepaRG cells were treated for 48hours with 1/2 log dilutions of 22(R)hydroxycholesterol, amiodarone, cyclosporine A, T0901317, troglitazone, Wyeth-14,643, and DMSO control. Lipid accumulation (A,B), fatty acid uptake (C), and APOB100 levels (fatty acid efflux, D) were assessed. Data are presented as the log₂ FC of test samples and were considered significant for log₂ FC ≥ 3 times the BMAD of DMSO controls (threshold). Bars are standard error, data points are median value, and data in the shaded region are within the threshold, reported from n=3–4 technical replicates.

Fatty Acid Uptake

Hepatocellular lipid accumulation depends on energy balance and, in particular, lipid flux. Therefore, the effects of the six test chemicals on hepatocellular fatty acid uptake were measured. This assay takes advantage of a bodipy labeled fatty acid that fluoresces when introduced into an intracellular environment. All chemicals tested had some effect on fatty acid uptake (Figure 2C). Specifically, Wyeth-14,643 (5, 15.8, and 50 μM), T0901317 (1.58 and 15.8 μM), cyclosporin A (5 and 15.8 μM), and 22(R)-hydroxycholesterol (15.85 and 50 μM) increased fatty acid uptake at levels exceeding, but near the response threshold. In contrast, troglitazone and amiodarone treatment decreased fatty acid uptake at the highest doses tested.

Fatty Acid Efflux

Hepatic lipids also accumulate when fatty acid efflux pathways are modified. Fatty acids primarily efflux from the liver as triglycerides attached to APOB100. APOB100 is the primary structural component of VLDL, lipid-rich particles secreted by hepatocytes into the circulation. Therefore, APOB100 levels were measured from the media to infer hepatocellular lipid efflux. Both T0901317 and cyclosporin A decreased APOB100 levels in the media of HepaRG cells (Figure 2D). This observed decrease was consistent with T0901317 and cyclosporin A induction of hepatic lipid accumulation (Figs. 1 and 2A).

Fatty Acid Oxidation

FAO is another mechanism to clear intracellular fatty acids. In hepatocytes, fatty acids are primarily broken down by mitochondrial beta-oxidative pathways that require the electron transport chain and cellular respiration (oxygen consumption) in eukaryotes. To assess this, FAO was measured using the Seahorse Mito Stress Test in the presence of the exogenous fatty acid palmitate. Media conditions were severely glucose limited and meant to favor fatty acids as the primary energy source. Etomixir inhibits carnitine palmitoyl transferase-1, an enzyme required for fatty acid transport across the mitochondrial membrane, was used as a negative control. As expected, Etomixir inhibited FAO (Figure 3). Of the chemicals evaluated, only Wyeth-14,643 (a PPARα agonist) had a median effect above the established response threshold. Although there were no treatment effects on basal respiration (Figure 3, T6), Wyeth-14,643 increased maximal mitochondrial respiration in the presence of exogenous fatty acids (Figure 3, T12).

Figure 3.

FAO was assessed in HepaRG cells treated with test compound (15.8 μM Wyeth-14,643, T0901317, cyclosporin A, 22(R)hydroxycholesterol, amiodarone, 10 μM troglitazone, 40 μM etomixir [negative control], and 0.01% DMSO) using the Seahorse Flux Analyzer XFe96 FAO Assay. The results are shown for chemical effects on OCR due to the oxidation of the exogenous fatty acid palmitate. Test compound, oligomycin, FCCP, and a mix of rotenone and AA were serially injected to measure chemical effects on adenosine triphosphate production, maximal respiration, and nonmitochondrial respiration. The data were baselined to the median rotenone/AA measurement at time point 15 of DMSO controls and scaled from 0 to 100. The basal threshold at T6 (after chemical injection) and maximal response at T12 (after FCCP injection) were analyzed to identify chemical effects on mitochondrial respiration outside of the response threshold (shaded region). Data reported from n=4 technical replicates.

Gene Expression Analysis

Increased hepatocellular lipid levels mediated by T0901317 and cyclosporin A were consistent with increased diacylglycerol O-acyltransferase 2 mRNA levels (DGAT2, Table 2), an enzyme required for cellular triglyceride biosynthesis (Yen et al., 2008). Consistent with troglitazone mediated decreases in fatty acid uptake, fatty acid transporter CD36 mRNA levels were also decreased by troglitazone in a concentration dependent manner (2.3-, 8.5-, and 43.7-fold; 1.0, 3.16, and 10 μM troglitazone, respectively) compared with DMSO controls (Table 2). Interestingly, T0901317 treatment increased APOB mRNA levels, in contrast to decreased media APOB100 protein levels. To indirectly examine whether or not hepatic fatty acids accumulated by increased synthesis, mRNA levels of genes key to the fatty acid synthetic process were measured. T0901317 and cyclosporin A decreased the expression of fatty acid synthesis genes ACACA, a rate-limiting substrate in fatty acid synthesis, and FASN, an enzyme that catalyzes the synthesis of fatty acids. T0901371 treatment decreased or increased the mRNA levels of PPARG depending on concentration and decreased sterol regulatory element binding transcription factor 1 (SREBF1) (Supplementary Table 1).

Table 2.

Median Log2 FC in Targeted Gene Expression Assays

		FA uptake	FA Efflux	FA Synthesis		TAG synthesis		FA oxidation	Lipid Droplet Accumulation?
Chemical	[μM]	CD36	APOB	ACACA	FASN	DGAT1	DGAT2	CPT1A
T0901317	1.58	↓2.33	↑1.52	↓2.20	↓8.35	↑1.40	↑1.92		Y
	5		↑1.95	↓2.18	↓8.87	↑1.55	↑2.90	↑1.83	Y
	15.8		↑6.25	↓2.41	↓2.16	↑2.57	↑10.22	↓1.49	Y
Troglitazone	1	↓2.33							N
	3.16	↓8.53							N
	10	↓43.73	↓1.43		↓1.36	↓1.28	↓1.58		N
22(R)-Hydroxycholesterol	5	↑2.03			↑1.56		↑1.54		N
	15.8	↑3.09			↑1.74				N
	50	↑2.07			↑2.04		↑1.47		N
Cyclosporin A	5		↓1.62	↓1.60	↓1.33	↓1.36			N
	15.8		↓1.56	↓1.72			↑3.41		Y
	50				↓1.39	↑1.61	↑4.86		Y
Wyeth-14,643	5					↑1.29	↑1.46		N
	50	↓3.20					↑1.51		N
Amiodarone	15.8	↓3.20	↑1.53			↑1.37			N

↓=FC<3 × BMAD, ↑=FC>3 × BMAD, gray shading for no change; Y, yes; N, no.

Protein Studies

Consistent with decreased extracellular (media) APOB100 protein measurements, intracellular APOB100 protein levels were also decreased by 1.35-fold with 15.8 μM T0901317 exposure (data not shown). These results suggested that T0901317 decreased efflux by limiting the protein scaffold (APOB100) needed to carry lipids out of the cell. This result was in contrast to increased APOB100 mRNA levels and suggested a T0901317-mediated posttranscriptional mechanism that lead to the decreased fatty acid efflux. Although fatty acid synthetic activity was not directly measured, FASN, the rate-limiting enzyme in fatty acid synthesis, was measured to infer activity. Cyclosporin A increased FASN protein levels approximately 2-fold, whereas T0901371 increased FASN protein 7- to 9-fold compared with DMSO controls (Figure 4). These results were in contrast to decreased FASN mRNA measurements (Supplementary Table 1) with T0901317 and cyclosporin A exposures and further supported posttranscriptional mechanisms leading to lipid accumulation.

Figure 4.

FASN protein levels were assessed from cyclosporin A and T0901317 treated HepaRG cell homogenate by an automated capillary immunoassay system. Bars represent FC relative to DMSO controls (dashed line). * for p < .05 by Mann Whitney test, n=3.

DISCUSSION

The goal of this study was to develop and evaluate in vitro assays for chemical evaluation based on a networked AOP for hepatic steatosis. The data from AOP-supported assays provide biological plausibility to link in vitro-based measurements after chemical exposures to adverse human health outcomes. Specifically, we evaluated a suite of assays that measured KE perturbations linked to a networked AOP for hepatic steatosis (Angrish et al., 2016). The underlying assumption was when biological pathways mediated by the KEs of the networked AOP were unbalanced, hepatic lipid accumulation could occur. These assays were evaluated with six compounds likely to impact these KE (lipid uptake, lipid efflux, FAO, and lipid accumulation) using an in vitro cryopreserved HepaRG model. The compounds were selected based on literature evidence of steatotic and/or hepatotoxic potential through various modes of action which were known to impact the KEs assessed (Table 1). The HepaRG cell line was selected as a model over other in vitro liver models because it was human-derived, less expensive than primary hepatocytes, less likely to include variable responses caused by inter-individual genetic differences, and maintained xenobiotic metabolism activity needed for toxicity testing (Jackson et al., 2016). Of note, HepaRG cells do differentiate into a co-culture system of hepatocyte and biliary-like cells in the presence of DMSO (Cerec et al., 2007). Our observations with Nile Red indicated lipid accumulation in the hepatocyte-like cells; however, we did not specifically verify cell type nor did we distinguish the specific contribution of hepatocyte or biliary-like cells to the responses measured. Furthermore, we did not assess the contribution of xenobiotic metabolism and resulting metabolites, which could have a major impact on the toxicodynamics of a chemical exposure; however, this is an area of current research in our laboratory.

Using an AOP-based targeted toxicity testing approach in a HepaRG model, we demonstrated that T0901317 and cyclosporin A caused significant lipid accumulation. These effects were mechanistically anchored using KE assays for fatty acid efflux, uptake, and synthesis. Although amiodarone treatments were previously shown to increase intracellular fatty acid levels in the HepaRG cell model (Antherieu et al., 2011; Tolosa et al., 2016), no significant lipid accumulation was identified after exposure in our testing conditions. These differences could be explained by both the time point evaluated and culture conditions used. Specifically, Antherieu et al. had not observed neutral lipid accumulation (indicated by Oil Red O stain) but noted phospholipidosis 24 h after a single exposure to 20 μM amiodarone exposure. Antherieu et al. observed lipid accumulation only after 14 days of repeated doses. Tolosa et al. measured an increase in intracellular neutral lipids 24 h after amiodarone exposure. However, Tolosa et al. evaluated responses after preincubation with fatty acid-free media, whereas we supplemented complete media with oleic acid 24 h prior to test chemical exposure. These differences in HepaRG culture conditions may shift the interplay between glucose and lipid metabolism that is amplified in the presence of a chemical such as amiodarone.

A key finding of this study was that hepatic steatosis was dependent on altering specific combinations of KEs above a certain threshold. For example, all 6 chemicals tested triggered activity above the established response threshold for at least one KE assay, but only 2 of those chemicals (T0901317 and cyclosporin A) induced lipid droplet accumulation in our model. This is likely explained by compensatory biological pathways that can be activated to maintain homeostasis. In vivo, lipid homeostasis is balanced by compensatory pathways that regulate fatty acid uptake, synthesis, efflux, and metabolism (Angrish et al., 2016). This implies there is an interconnected biological network regulating lipid levels rather than a linear path. If perturbed, these alternative biological pathways may be activated. For example, even though Wyeth-14643 increased fatty acid uptake in our HepaRG model, the observed increase in FAO likely maintained homeostasis. Likewise, multiple perturbed biological pathways may result in an AO. For T0901317 and cyclosporin A, increased uptake and decreased efflux likely tipped the balance to favor hepatic steatosis. Additionally, for T0901317, superimposing the KE assay with mRNA and protein level endpoints identified a posttranscriptional mechanism affecting APOB100 that lead to a decreased lipid efflux. These data may highlight the need for developing expert guided AOP networks, based on systems biology that can be used to scope a suite of assays that better predict the adverse effects of chemical exposures.

The steatosis AOP assays described here were developed with the premise that mechanistic and biology-based assays could add critical scientific information and data needed for the chemical evaluation process (Figure 5). Yet, the qualitative state of this AOP may limit its application to risk assessment. The chemical concentration and KE and/or AO response data could be used to estimate a benchmark dose for derivation of a point of departure (USEPA, 2012). However, the concentration-response curve created by this approach would likely model the AO (lipid accumulation measured by Nile Red fluorescence) as the response implicit of the other KEs. As indicated, the “lipid accumulation” outcome is dependent on the compensatory actions of more than one KE and therefore this model would lack explicit mechanistic detail added by the other biologically based KE assays. These ideas were recently alluded to by Conolly et al. (2017) who described the quantitative (q)AOP concept using aromatase inhibition as a case study. To create the qAOP, the authors modeled KEs and KE relationships (defined by quantitative relationships of exposure dose and duration) in silico. The qAOP was then applied to an ecological risk assessment, specifically addressing chemical exposure effects on fish population dynamics. This study demonstrated how AOP networks and qAOPs (driven by mechanistic and biological-based data) can use in vitro data and reduce overall uncertainty in the chemical risk assessment process.

Figure 5.

Summary of an AOP-based toxicity test for steatosis. An AOP network for hepatic steatosis was used to develop mechanistic toxicity tests for KEs. Leveraging scientific knowledge, mechanistic AOP-based toxicity tests, and dosimetry, the community may be able to rapidly and formally describe (with empirical data) causal relationships between exposure and health effects.

AOP networks are a critical step toward improving the weight of evidence for chemical hazard characterization. In the past decade, high-throughput in vitro evaluation of “toxicity pathways”, defined as “pathways that can lead to adverse health effects when sufficiently perturbed” (Rainham et al., 2010), and more recently linear AOPs (https://aopwiki.org/, accessed 06/22/17), have been useful prioritization tools. In order to evaluate chemical impacts on human health, however, these AOPs and toxicity pathways need to be networked to more clearly represent complex biological and ecological systems. For example, selective chemicals (ie, mediated by nuclear receptor activation) may lead to pleiotropic effects too far upstream of an apical endpoint to causally anchor that specific event to an AO. Therefore, a single in vitro assay may accurately predict chemical-mediated proximal effects directly dependent on receptor activation; however, this assay could fail to predict responses dependent on compensatory, adaptive, feedback, and other related mechanisms that ultimately impact apical KEs. AOP networks may help reduce the uncertainty in those predictions by using interconnected KEs or “biological landmarks” (such as lipid efflux and uptake) that can be causally linked to an AO. The utility of this approach is that a suite of biologically based assays can also provide the science-based analysis and data needed to predict responses from chemicals that act through less selective molecular interactions (Thomas et al., 2013). For example, LXR activation or changes in APOB mRNA levels by T0901317 alone are not sufficient evidence to infer hepatic steatosis. However, effects on LXR and APOB in combination with impaired lipid efflux and increased lipid uptake provides better mechanistic evidence to anchor T0901317 exposure to lipid accumulation. This is a critical concept in the evolution of predictive toxicology; by integrating expert opinion, experimental evidence, and AOPs, it is possible to organize massive data sets into groups or “AOP fingerprints” predictive of downstream KEs.

By design, AOP networks can be leveraged as guidance tools to organize chemical evaluation data that already exists to assess chemical risk for endpoints of regulatory concern. Rapid and defensible chemical assessment methods are necessary to evaluate the potential health hazards of the tens of thousands chemicals in commerce and the environment. The approach described in this study demonstrates the value of applying a biologically based framework to evaluate important endpoints. AOP-based testing batteries can be applied to both prioritize chemical evaluation and predict potential for human health and ecological adversity.