Disclosure: MB is employed by Metabolon and has equity interests in Metabolon. SC is employed by ZenBio. MM is employed by Metabolon and has equity interests in Metabolon. KL is employed by Metabolon and has equity interests in Metabolon. BC is employed by ZenBio.
Funding agencies: This study was supported by a grant from the National Institute of Health DK088430. Support for MB and SC was received from the North Carolina Biotechnology Center Industrial Fellowship Program.
A spectrum of disorders including simple steatosis, nonalcoholic steatohepatitis, fibrosis, and cirrhosis is described by nonalcoholic fatty liver disease (NAFLD). With the increased prevalence of obesity, and consequently NAFLD, there is a need for novel therapeutics in this area. To facilitate this effort, a cellular model of hepatic steatosis was developed using HepaRG cells and the resulting biochemical alterations were determined.
Design and Methods
Using global metabolomic profiling, by means of a novel metabolite extraction procedure, the metabolic profiles in response to the saturated fatty acid palmitate, and a mixture of saturated and unsaturated fatty acids, palmitate and oleate (1:2) were examined.
We observed elevated levels of the branched chain amino acids, tricarboxylic acid cycle intermediates, sphingosine and acylcarnitines, and reduced levels of carnitine in the steatotic HepaRG model with both palmitate and palmitate:oleate treatments. In addition, elevated levels of diacylglycerols and monoacylglycerols as well as altered bile acid metabolism were selectively displayed by palmitate-induced steatotic cells.
Biochemical changes in pathways important in the transition to hepatic steatosis including insulin resistance, altered mitochondrial metabolism, and oxidative stress are revealed by this global metabolomic approach. Moreover, the utility of this in vitro model for investigating the mechanisms of steatotic progression, insulin resistance, and lipotoxicity in NAFLD was demonstrated.
In parallel with the increasing occurrence of obesity and type 2 diabetes, nonalcoholic fatty liver disease (NAFLD) may be affecting up to 25% of the general population, including a significant portion of pediatric cases [1, 2]. Current predictions indicate that 40% of the population will be obese by 2025 with NAFLD becoming the leading cause of liver disease and need for liver transplantation . Although the exact mechanisms are not completely understood, excess lipid accumulation in hepatocytes is a primary event in the development of NAFLD.
Hepatic free fatty acids (FFAs) are derived from multiple sources including de novo synthesis, adipose tissue, and dietary triglycerides . In the liver, FFAs are converted to fatty acyl CoAs which are directed to specific metabolic pathways depending on energy demand and fatty acid composition. The predominant pathway involves conversion of fatty acyl CoAs into triglycerides and glycerophospholipids and results in the generation of lipid intermediates including diacylglycerols (DAGs). Alternatively, fatty acids can be directed toward the generation of sphingolipids or converted to acylcarnitines for transport into the mitochondria for β-oxidation.
Hepatic steatosis occurs when excess FFAs are transported to the liver and/or when increased de novo synthesis of triglycerides exceeds the ability of the liver to metabolize or secrete FFAs as very low density lipoprotein. The accumulation of excess FFA in hepatocytes may directly cause lipotoxicity by promoting oxidative stress and activation of inflammatory pathways or may sensitize hepatocytes to mitochondrial dysfunction, oxidative stress, and inflammatory cytokines and adipokines . These events will likely lead to simple steatosis and eventually progress to NASH.
NAFLD is frequently observed in patients with type 2 diabetes mellitus and is highly associated with insulin resistance. Although the contributing mechanisms involved in the relationship between NAFLD and insulin resistance are unknown, there are several proposed models. One prominent hypothesis suggests that underlying mitochondrial deficiency results in the accumulation of toxic lipid metabolites that can interfere with insulin signaling [6, 7]. An alternative hypothesis suggests that increased lipids cause mitochondrial overload, leading to the accumulation of lipid metabolites and reactive oxygen species (ROS) that subsequently cause mitochondrial dysfunction and an insulin-resistant state [7, 8]. Moreover, elevated ROS and associated reactive derivatives can deplete ATP, NAD, and glutathione (GSH) levels, causing oxidative stress, DNA, and protein damage and cell death . Studies using serum or liver biopsies from patients with NAFLD have explored metabolic characteristics of NAFLD, and observed altered GSH, bile acids (BAs), carnitine, branched chain amino acids (BCAA), and γ-glutamyl peptides, and mouse models of diet-induced obesity show similar overlap in serum- and liver-specific metabolites [8, 10].
Metabolomics has been used to identify biochemical alterations in cellular systems . The aim in this study was to establish a cell-based model of steatosis and apply an unbiased metabolomic approach to determine biochemical signatures associated with steatotic progression and insulin resistance. We achieved this goal by developing a novel, high-throughput method for extracting metabolites from adherent cells. With this method, termed Intact Sample Extraction (ISE), metabolites are extracted and processed for metabolomic analysis, leaving cells attached to the culture plate, preserving the cells for additional analysis and streamlining metabolite extraction. We first examined steatotic transition using the saturated fatty acid palmitate, which is associated with lipid-induced cell toxicity [18, 19]. In a separate experiment, we sought to understand the differences between steatosis induced by palmitate and a more physiologically relevant mixture of saturated and unsaturated fatty acids, palmitate and oleate (1:2). Using the ISE method, metabolites analyzed from the lipid-loaded HepaRG cells revealed key metabolic pathways associated with the progression of human NAFLD including insulin resistance, altered mitochondrial metabolism, and oxidative stress.
For the method comparison study, HEK293 cells were seeded at 4 × 104 cells per well in six-well plates and grown to confluence in EMEM (ATCC, Manassas, VA) supplemented with 10% fetal bovine serum (FBS), 1% Pen/Strep. For method scale-down studies, HepG2 cells were seeded in 6-, 12-, 24-, and 96-well formats and expanded to confluence for more than 48 h in Dulbecco's modified Eagle's medium supplemented with 10% FBS, glutaMAX I (Life Technologies, Grand Island, NY) and antibiotics. For each experiment, cells were seeded simultaneously using an appropriate number of wells for metabolomic studies and protein analysis. HepaRG cells (Biopredic International, Rennes, France) were seeded at 2.9 × 104 cells per cm2 in 12-well plates and differentiated as described previously .
Metabolite extraction and sample preparation
For method validation, four replicates of HEK293 cells were prepared using the conventional cell scraping and homogenization extraction method. Briefly, cells were rinsed once in PBS, dislodged using a cell scraper, and pelleted. Cell pellets were extracted as described previously .
Metabolites were extracted from cells at confluence for the method development (four replicates/condition), method scale down (five replicates/condition), palmitate time course (four replicates/condition), and palmitate:oleate (three replicates/condition) experiments. Except where indicated, metabolites were extracted using the ISE method, media were removed, and cells were rinsed once in PBS. At room temperature, 80% methanol (ultrapure water, 20%) was immediately added to the cells, and metabolites were extracted for 5 min. The methanol extract was removed and stored at −80°C until processing. Cells for the lipid time course experiment were rinsed briefly with PBS and fixed for staining as indicated below. Methanol extracts were evaporated to dryness under a stream of nitrogen gas at 40°C in a Turbovap LV evaporator (Zymark, Hopkinton, MA). The dried extracts were reconstituted in 550 μl methanol:water (80:20) containing recovery standards (D,L-2-fluorophenylglycine, D,L-4-chlorophenylalanine, tridecanoic acid, D6 cholesterol).
Global metabolomic profiling was carried out on three independent instrument platforms, one gas chromatography/mass spectrometry (GC/MS) and two ultra-high-performance liquid chromatography/tandem mass spectrometry (UHPLC/MS/MS2) platforms optimized for either basic or acidic species as described previously [21, 22]. Metabolites were identified by automated comparison of the ion features in the experimental samples to a reference library of chemical standard entries that included retention time, molecular weight (m/z), preferred adducts, and in-source fragments as well as their associated MS/MS2 spectra. This library allowed the rapid identification of metabolites in the sample with high confidence.
Preparation of fatty acids and lipid loading
Stock solutions of palmitate (200 mM) or oleate (100 mM) (Sigma, St. Louis, MO) were prepared in ethanol or 1.6 mM NaOH solution, respectively. Palmitate or a mixture of palmitate:oleate at a ratio of 1:2 was conjugated to albumin in 30% essentially fatty acid-free (FAF) BSA solution (Invitrogen, Grand Island, NY) and diluted to 10 mM in Williams E (Invitrogen). Differentiated cells were transferred to media containing 0.5 mM lipid or BSA alone for the indicated times, with media replaced daily.
HepaRG were loaded with 0.1-0.5 mM BSA-conjugated palmitate or palmitate:oleate (1:2) for 48 h. Samples were rinsed in PBS followed by the determination of triglyceride content using the triglyceride assay kit (ZenBio Inc., Research Triangle Park, NC). The optical density was read at 540 nm on a Spectramax (Molecular Devices, Sunnyvale, CA) plate reader, and the concentration of glycerol was determined using the following equation: 1 M triglyceride = 1 M glycerol + fatty acids. The mean concentration of triglyceride and standard deviation were calculated for each sample. A two-tailed t-test was used to calculate significant (P ≤ 0.05) differences between BSA and lipid-loaded samples at each concentration.
Oil Red O staining
After metabolite extraction, the cells were rinsed briefly in PBS and stained with Oil Red O. Cells were fixed for at least 1 h in 4% paraformaldehyde at 4°C and rinsed twice with water. Cells were then stained with 0.5% Oil Red O solution for 20 min at room temperature, washed two to three times in water and images were captured on a Ziess Axiovert microscope equipped with a digital camera and supporting IC capture software (The Imaging Source, Charlotte, NC).
Differentiated HepaRG cells were treated with BSA or palmitate (0.3 or 0.5 mM) for 72 h in triplicate. ROS production was analyzed using the Image-IT live reactive oxygen detection kit (Invitrogen) using the manufacturer's protocol. The mean and standard deviation were calculated and expressed as relative fluorescent units (RFUs). Significance was determined by two-tailed t-test, with P ≤ 0.05 considered significant.
Insulin-stimulated phosphorylation of AKT
Lipid-loaded HepaRG cells were placed in serum-free media for 8 h, followed by stimulation with recombinant human insulin (Roche Diagnostics, Indianapolis, IN) at 0.1, 1.0, 10, or 100 nM for 10 min. Samples were rinsed in PBS and lysed in RIPA buffer (150 mM NaCl, 50 mM Tris, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, and 1 mM EDTA) supplemented with complete protease inhibitor cocktail (Roche Diagnostics) and HALT phosphatase inhibitor cocktail (Thermo Scientific, Waltham, MA). Protein concentration was determined in the lysates using BCA protein assay reagent (Thermo Scientific). In brief, 50 μg of protein was separated on a 4-12% NUPAGE BIS-Tris gel (Life Technologies), transferred to nitrocellulose, and blocked in Tris-buffered saline, pH 7.5, 0.1% Tween-20, and 3% FAF BSA for 1 h. Membranes were rocked at 4°C overnight with phospho or total AKT primary antibody (Cell Signaling Technology, Danvers, MA) at a dilution of 1:1,000. The membranes were washed three times in TBST for 5 min followed by incubation with secondary antibody 1:3,000 in 5% milk in TBST (Cell Signaling Technology). After three TBST washes, membranes were incubated with Supersignal West Pico chemiluminescence detection reagent (Thermo Scientific), and luminescent signals were captured using a BioRad Gel Documentation system (BioRad laboratories, Hercules, CA).
Missing values for a given metabolite were imputed with the observed minimum detection value based on the assumption that they were below the limits of instrument detection sensitivity. All comparisons were performed using log-transformed data. All samples were normalized to protein as determined by BCA assay (described above). Welch's two sample t-tests or two-way ANOVA with contrasts (palmitate time course) were used for comparisons. Multiple comparisons were accounted for with the false discovery rate (FDR) method, and each FDR was estimated using q values. Biochemicals with P ≤ 0.05 and q ≤ 0.1 were considered statistically significant. For convenience of data visualization, raw area counts for each biochemical were rescaled by dividing the value for a specific biochemical in each sample by the median value for that specific biochemical. Comparisons are represented as fold of change values. Principal component analysis (PCA) was performed to characterize the metabolic differences between the palmitate and the control groups at each time point. For the PCA, each principal component is a linear combination of all metabolites. The coefficients for the first component are determined by those that maximize the variance. The coefficients for the second component are chosen to maximize the variance with the constraint that it is orthogonal to the first component. All statistical analyses were generated using Array Studio software. Array Studio, Array Viewer, and Array Server and all other Omicsoft products or service names are registered trademarks or trademarks of Omicsoft Corporation (Research Triangle Park, NC).
Method development and scale down
The ISE method reported here was first compared to a traditional homogenization extraction method of harvested pellets of HEK293 cells. We identified 182 metabolites using the ISE method and 163 using the harvested cell pellet extraction method; 161 metabolites were common to both methods (Supporting Information Figure 1A). The metabolites identified using the ISE method represent a broad range of biochemical classes (Supporting Information Figure 1B).
The ability to detect metabolites in multiwell formats was first determined using HepG2 cells plated in various well formats and analyzed by global metabolomics. The well sizes, volume of solvent used for extraction, and the number of cells seeded per well are summarized in Supporting Information Table 1. Using the ISE method and nontargeted biochemical profiling, 221 distinct metabolites, matching named structures in our chemical reference library, were identified from a confluent six-well plate of HepG2 cells (Supporting Information Table 1). Not surprising, the average levels of metabolites decreased as the size of the well and total cell number decreased (Supporting Information Table 1); likely owing to some metabolites to falling below the level of detection by the instrument. Nonetheless, 189 out of 221 metabolites (86%) were detected in the 96-well format when using this extraction method. Furthermore, the cells remained fixed to the plate, allowing them to be utilized in additional cell-staining procedures. With the ISE method, it is possible to perform global metabolomic analysis using higher throughput plate formats.
Table 1. Selected metabolites altered in palmitate- and palmitate:oleate-treated HepaRG cells at 8 and 48 h
Fold change relative to BSA control
Metabolite values are expressed as fold of change for the indicated condition relative to BSA control. Bold values are significant with P ≤ 0.05 and q ≤ 0.1.
Pantothenate and CoA metabolism
Glycolysis, gluconeogenesis, pyruvate metabolism
Valine, leucine, and isoleucine metabolism
Bile acid metabolism
Glutathione, reduced (GSH)
Glutathione, oxidized (GSSG)
Induction of steatosis in HepaRG cells
To establish conditions that induce steatosis, HepaRG cells were cultured in growth media supplemented with 0.1, 0.3, or 0.5 mM BSA-conjugated palmitate or palmitate:oleate (1:2), or with BSA alone for 48 h, and triglyceride levels were measured. Increased total triglyceride levels were observed in both lipid treatment conditions. These increases were significant for treatment with 0.3 and 0.5 mM lipid compared to BSA alone (Figure 1A). Using 0.5 mM fatty acid, triglyceride levels were similarly increased for both fatty acid treatments (2.9- and 3.1-fold increases compared to BSA control for palmitate and palmitate:oleate treatments, respectively) (Figure 1A). For subsequent experiments, 0.5 mM of fatty acid was used to induce steatosis.
Metabolite profiles of steatotic HepaRG cells
To capture the metabolic transition to steatosis, HepaRG cells were treated with BSA-conjugated palmitate or BSA alone over a 48-h time course. Lipid accumulation was visible in the palmitate-loaded samples within 8 h and continued to accumulate through the 48-h time point as determined by Oil Red O staining (Figure 1B). After metabolite extraction, global metabolomic profiling of the palmitate-loaded samples at 8, 16, 24, and 48 h identified a total of 169 biochemicals (Supporting Information Table 2). PCA was used as a way to visualize the samples based on their metabolic profiles. Information from the complete data set was reduced to two principal components. As shown in Figure 1C, palmitate-treated samples separated from BSA samples at all time points with the exception of time 0. These data imply that a clear metabolic signature of palmitate-induced steatosis can be obtained in HepaRG cells. In fact, we observed distinct metabolic profiles of palmitate-treated HepaRG cells during the transition to steatosis (Figures 1-5).
However, given the lipotoxic effects of prolonged exposure to saturated fatty acids, including palmitate, we performed an independent follow-up study using palmitate alone or a physiological mixture of saturated and unsaturated fatty acids (palmitate and oleate, 1:2) (Table 1). Based on the results of the palmitate time course, we selected 8- and 48-h treatment time points for the follow-up study. Although distinct metabolic profiles were observed for each steatotic treatment, both palmitate- and palmitate:oleate-treated HepaRG cells demonstrated changes in metabolites that were indicative of altered lipid and energy metabolism and insulin resistance. The results from both studies are presented in parallel below.
As expected from fatty acid loading, lipid metabolism was significantly altered in both metabolomic studies, particularly evident through observed changes in carnitine metabolism. Treatment with palmitate or palmitate:oleate led to reduced levels of carnitine (Figure 2A and Table 1), whereas acylcarnitines including butyrylcarnitine, propionylcarnitine, and oleoylcarnitine were significantly elevated (Figure 2B and Table 1). Acetyl CoA was also significantly elevated with both fatty acid treatments at 8 h (Table 1). However, reduced levels of coenzyme A were observed only with palmitate treatment at the 48-h time point (Figure 2C and Table 1).
Significant changes in energy metabolism were consistently observed between palmitate-treated and BSA control samples. Carbohydrate metabolism contributions to the tricarboxylic acid (TCA) cycle during the 48 h of palmitate treatment are shown in Figure 3. The product of anaerobic glycolysis, lactate, and the TCA cycle intermediates, cis-aconitate, fumarate, and malate were significantly higher in palmitate-loaded samples for all time points. Citrate was significantly elevated at 8, 16, and 24 h, and α-ketoglutarate was elevated at 16 and 48 h with palmitate treatment. Elevated lactate and TCA cycle intermediates were also observed in palmitate:oleate-treated cells at the 48-h time point (Table 1). The BCAAs, isoleucine, leucine, and valine, were significantly elevated in palmitate- and in palmitate:oleate-treated cells at 48 h (Figure 4A and Table 1), indicating possible anaplerotic contributions to the TCA cycle.
In palmitate-treated samples, DAG, and monoacylglycerols (MAG) were significantly elevated (Figure 4B, Supporting Information Table 2, and Table 1). The DAGs, 1,2-dipalmitoylglycerol, and 1,3-dipalmitoylglycerol were elevated within 8 h of palmitate treatment (Figure 4B and Table 1), whereas increased MAG levels were observed at 48 h (Supporting Information Table 2 and Table 1). In contrast, DAG and MAG levels were not significantly altered in palmitate:oleate-treated cells (Table 1). Levels of the ceramide precursor, sphingosine, were elevated in both palmitate- (Figure 4C and Table 1) and palmitate:oleate-treated cells (Table 1).
BAs are involved in glucose homeostasis through interaction with BA receptors FXR (Farnesoid X receptor) and TGR5 [23, 24]. The primary BA taurocholate was significantly reduced compared to BSA controls within 8 h of palmitate treatment in both studies (Figure 1D and Table 1). In contrast, taurocholate was unaltered by palmitate:oleate treatment (Table 1).
Biochemicals involved in GSH metabolism, including cystathionine and S-adenosylhomocysteine (SAH), were altered with palmitate treatment in both studies (Figure 5A, Table 1, and Supporting Information Table 2). Several other biochemicals of GSH metabolism were significant in the palmitate time course study. Beginning at 24 h, GSH levels in palmitate-treated samples were reduced compared to BSA and were significant at 48 h (Figure 5B). γ-Glutamylthreonine was transiently elevated by palmitate at 8- to 24-h time points, with γ-glutamylleucine elevated later at 16 and 24 h (Figure 5B). Overall, GSH metabolism was not significantly altered in HepaRG cells treated with palmitate:oleate.
To determine the extent of ROS generation in palmitate-treated cells, we labeled cells with carboxy-H2DCFDA fluorescein. This cell-permeable fluorogenic marker is readily oxidized by ROS into a fluorescent product. Cells were treated with BSA, 0.3 or 0.5 mM palmitate; tert-buytl hydroperoxide (100 μM, 1 h) was used as a positive control. Compared to BSA control samples, ROS were elevated 1.5-fold (P < 0.036) and 1.7-fold (P < 0.025) in the presence of 0.3 or 0.5 mM palmitate, respectively (Figure 5C). These data support the metabolic indications of oxidative stress in the palmitate-induced steatotic model as predicted by the metabolic signatures.
Lipid-induced decrease in insulin-stimulated phosphorylation of AKT
Elevated mitochondrial β-oxidation and increased oxidative and anaplerotic contributions to the TCA cycle are reported to generate lipid-derived metabolites and ROS in human NAFLD and hepatocyte cultures, decreasing the ability of insulin to stimulate proximal signaling events . To test the hypothesis of lipid-induced alterations in insulin signaling, BSA or palmitate-loaded HepaRG cells were treated with increasing concentrations of insulin (0.1-100 nM) for 10 min followed by analysis of total and phosphorylated AKT by immunoblotting. In BSA control samples, insulin stimulation resulted in the expected concentration-dependent increase in AKT phosphorylation at serine 473. In contrast, insulin-stimulated phosphorylation of AKT was reduced in palmitate-loaded cells by ∼ 30% at the highest concentrations (data not shown). Total AKT levels remained constant for all treatment conditions. This finding supports our metabolomic data, indicating an insulin-resistant phenotype in palmitate-loaded HepaRG cells.
In this study, we characterized a cell-based model of lipid-induced hepatic steatosis using HepaRG cells treated with palmitate alone or a combination of palmitate and oleate. We established a protocol to extract metabolites from intact cells in a multiwell format, identified metabolic signatures of steatotic progression induced by palmitate and palmitate:oleate, and validated several of these metabolic profiles with biological assays. Our data from two independent metabolomic studies revealed changes in metabolites related to energy metabolism, insulin resistance, and oxidative stress, which are key biochemical processes known to be involved in steatosis and NASH. Importantly, the distinct metabolic signatures identified using this in vitro cell model, including elevated levels of lipid metabolites, TCA cycle intermediates and BCAAs and reduced levels of carnitine and GSH, recapitulates biochemical alterations observed in serum and liver tissue from human NAFLD patients and in animal models of NAFLD [10, 14, 26].
Elevated DAG levels in liver biopsies from NAFLD patients and animal models of NAFLD are associated with hepatic lipotoxicity . Elevated DAG has also been implicated in the development of hepatic insulin resistance through activation of protein kinase C ε . We observed a significant accumulation of DAG and MAG metabolites in palmitate-treated HepaRG cells, consistent with the studies describing insulin-resistant and lipotoxic phenotypes in response to saturated fatty acids [18, 19, 33]. Metabolic indications of oxidative stress in palmitate-treated HepaRG cells were supported by ROS assays. In contrast, oleate protects against palmitate-induced lipotoxicity by targeting palmitate for triglyceride synthesis [33, 34]. In the palmitate:oleate-treated steatotic model, DAG and MAG levels were not significantly altered, suggesting that oleate is protective against accumulation of these lipid metabolites. However, elevated levels of sphingolipid suggest that the ceramide signaling pathway may be altered in palmitate:oleate-treated HepaRG cells. Ceramide levels are increased in insulin-resistant humans, and this has been attributed to ceramide impairment of insulin-stimulated phosphorylation of Akt . These data demonstrate that different fatty acids or combinations of fatty acids may induce insulin resistance through distinct mechanisms.
Elevated acetyl CoA and concomitant decrease in carnitine levels in the steatotic HepaRG cells suggest that processing of fatty acids through β-oxidation is occurring at early time points but may be limited with prolonged lipid treatment owing to eventual depletion of free carnitine. Carnitine supplementation has been explored for its ability to promote β-oxidation of fatty acids and reduce steatosis in rodent models . Without carnitine supplementation, fatty acids may be preferentially partitioned toward alternative metabolic pathways that could contribute to the generation of toxic lipid metabolites . Interestingly, palmitoylcarnitine was elevated in palmitate-treated cells, whereas both palmitoylcarnitine and oleoylcarnitine were elevated in palmitate:oleate conditions. These incompletely oxidized fatty acids further suggest altered β-oxidation. In a mouse model of NAFLD, metabolomic analysis revealed depletion of carnitine in liver tissue but not in serum . In a separate metabolomic study, elevated levels of circulating carnitine species including free carnitine, butyrylcarnitine, and methylbutyrylcarnitine, were observed in plasma from NAFLD patients . The results in liver tissue are consistent with our findings in the two steatotic cell treatment conditions, and emphasize that plasma/serum metabolites reflect the net result of all organs/tissues rather than organ/tissue-specific alterations.
A recent study analyzing mitochondrial oxidative metabolism in the liver of individuals with NAFLD demonstrates that elevated hepatic triglyceride is associated with higher flux in the TCA cycle and elevated anaplerotic TCA cycle contributions from pyruvate cycling and gluconeogenesis . In this study, we observed an increase in lactate and TCA cycle intermediates in response to palmitate, suggesting increased TCA cycle activity in this condition. This effect may be less severe or delayed in palmitate:oleate conditions as TCA cycle intermediates were not elevated until 48 h, and lactate was not significantly altered.
High levels of BCAAs, which include the essential amino acids valine, leucine, isoleucine, and their metabolites, are associated with insulin resistance [28, 37]. Elevated BCAAs have been observed in patients with obesity, type 2 diabetes, and NAFLD [10, 28, 37, 38] and are strong candidates in predicting insulin sensitivity  and monitoring disease progression and response to therapeutic intervention [28, 37]. Consistent with these clinical observations and with the altered TCA cycle activity in this model, we observed elevated levels of BCAAs in both fatty acid-loaded conditions, suggesting anaplerotic contributions to the TCA cycle. The BCAA levels in palmitate:oleate-treated cells increased at 48 h, which is consistent with the delayed TCA cycle effect in that condition. When taken in concert, the metabolic indications of impaired β-oxidation and increased TCA cycle activity through anaplerotic contributions suggest clear mitochondrial abnormalities in both fatty acid-loaded conditions.
In addition to their role in dietary lipid absorption and cholesterol homeostasis, BAs are signaling molecules that play a role in glucose and lipid metabolism . Unlike DAGs and ceramide, BA signaling has been shown to improve insulin sensitivity by modulating triglyceride, cholesterol, and glucose homeostasis . Lower levels of the BA taurocholate are associated with early stage insulin resistance in mice . Our observation of reduced taurocholate levels in palmitate-treated HepaRG cells may indicate the onset of an insulin-resistant phenotype. In contrast to our findings, increased levels of BAs were reported in steatotic liver tissues . These opposite effects may be owing to sample type differences (HepaRG cells used in our studies compared to human liver tissue) although additional studies are needed to clarify the role of BA signaling in this steatotic HepaRG cell model. Regardless, the data suggest that dysregulation of BA homeostasis is associated with insulin resistance and steatosis and may contribute to the pathogenesis of NAFLD.
In this study, we used a novel method to extract metabolites from intact cells and performed an unbiased metabolomic approach to examine global changes in hepatocyte-like cells during steatotic progression induced by fatty acids. With this approach, we identified key biochemical signatures related to steatosis and NAFLD, including altered mitochondrial metabolism, insulin resistance, and oxidative stress. These profiles support many of the reported biochemical alterations observed in animal models of NAFLD and importantly are consistent with metabolomic signatures obtained with clinically relevant human liver samples and serum from patients with varying degrees of fatty liver disease. Our results demonstrate the suitability and utility of this steatotic cell model to further investigate steatosis and insulin resistance and represent a novel platform for drug development and preclinical toxicity studies.
The authors thank the Metabolon platform team and chemical spectra analysts for contributing to data acquisition. The assistance of Jacob Wulff in statistical analysis of the data is gratefully acknowledged.