Saturated fatty acid combined with lipopolysaccharide stimulates a strong inflammatory response in hepatocytes in vivo and in vitro

Abstract

Nonalcoholic fatty liver disease (NAFLD) is the most common chronic liver disease and consumption of high-fat diet (HFD) is a risk factor for NAFLD. The HFD not only increases intake of saturated fatty acid (SFA) but also induces metabolic endotoxemia, an HFD-associated increase in circulating lipopolysaccharide (LPS). Although it is known that SFA or LPS promote hepatic inflammation, a hallmark of NAFLD, it remains unclear how SFA in combination with LPS stimulates host inflammatory response in hepatocytes. In this study, we performed both in vivo and in vitro experiments to investigate the effect of SFA in combination with LPS on proinflammatory gene expression in hepatocytes. Our animal study showed that feeding low-density lipoprotein-deficient mice HFD enriched with SFA and injection of low-dose LPS cooperatively stimulated IL-6 expression in livers. To understand how SFA and LPS interact to promote IL-6 expression, our in vitro studies showed that palmitic acid (PA), a major SFA, and LPS exerted synergistic effect on the expression of IL-6 in hepatocytes. Furthermore, coculture of hepatocytes with macrophages resulted in a greater IL-6 expression than culture of hepatocytes without macrophages in response to the combination of PA and LPS. Finally, we observed that LPS and PA increased ceramide production by cooperatively stimulating ceramide de novo synthesis, which played an essential role in the synergistic stimulation of proinflammatory gene expression by LPS and PA. Taken together, this study showed that SFA in combination with LPS stimulated a strong inflammatory response in hepatocytes in vivo and in vitro.

INTRODUCTION

Nonalcoholic fatty liver disease (NAFLD) is the most common chronic liver disease characterized by accumulation of fat in liver cells in the absence of excessive alcohol intake (30). Consumption of Western diets plays an important role in NAFLD (47), and metabolic syndrome and type 2 diabetes are associated with NAFLD (36, 38). NAFLD is histologically categorized into nonalcoholic fatty liver, which is hepatic steatosis without inflammation and injury and nonalcoholic steatohepatitis (NASH), which is characterized by the presence of hepatic steatosis with inflammation and necrosis (5). It has been well documented that NASH is associated with cardiovascular disease (CVD) and is considered as an independent predictor for the risk of future CVD events (48). CVD but not liver steatohepatitis is the major cause of death in patients with NASH (9). The potential molecular mediators linking NASH with CVD include proatherogenic molecules such as low-density lipoprotein (LDL), fibrinogen, plasminogen activator inhibitor-1, and proinflammatory cytokines (8, 49).

In recent years, studies have shown that NASH involves a complex interaction between high-fat diet (HFD), host genetics, and immunological factors (35), and the hallmarks of NASH are hepatic inflammation, injury, and fibrosis (2). The pathological features of livers with NASH include increased infiltration of immune cells, such as monocytes, macrophages, T-lymphocytes, and neutrophils, and activation of Kupffer cells and stellate cells (31). Furthermore, studies have shown that innate immunity plays an essential role in the pathogenesis of NASH as pathogen-associated molecular patterns such as bacterial lipopolysaccharide (LPS) activate immune cells and trigger inflammatory signaling cascades, leading to hepatic inflammation and injury (2).

LPS, a constituent of the outer membrane of Gram-negative bacteria, activates Toll-like receptor (TLR)4 to elicit inflammatory signaling (4). It has been shown that patients with type 2 diabetes or metabolic syndrome have metabolic endotoxemia, a condition referring to an increased circulating LPS as a result of HFD-increased intestinal permeability, favoring translocation of gut microbiome-derived LPS to the bloodstream (13, 32). When circulating LPS reaches the liver, it activates TLR4-mediated inflammatory signaling cascades in Kupffer cells and other types of cells and triggers proinflammatory gene expression, leading to hepatic inflammation, injury, and fibrosis (25).

In addition to LPS, HFD-related increase in free saturated fatty acid (SFA) also contributes to NASH (10, 55). Diets enriched with SFA are harmful as they increase both liver fat and insulin resistance. In addition, SFA has been shown to promote hepatocellular dysfunction by activating endoplasmic reticulum stress, dysregulating mitochondrial metabolism and increasing reactive oxygen species accumulation (18). Accumulation of SFA in liver also results in hepatocyte apoptosis (42).

HFD-induced metabolic endotoxemia and HFD-related fat uptake may increase both LPS and SFA in bloodstream in patients with type 2 diabetes or metabolic syndrome (13, 55). Therefore, it is important to elucidate the interaction between LPS and SFA on inflammation-associated diseases such as atherosclerosis and NASH. Recently, we reported that LPS and SFA cooperatively increased atherosclerosis in LDL receptor-deficient (LDLR^−/−) mice, an animal model for human atherosclerosis (21). However, it remains largely unknown how the interaction between LPS and SFA affects hepatic inflammation, which is well known to contribute to atherosclerosis (48). Given the crucial role of hepatic inflammation in systemic inflammation and atherosclerosis, it is important to determine how immunological and metabolic factors interact to alter hepatic inflammatory response for a better understanding of the pathogenesis of atherosclerosis in type 2 diabetes and metabolic syndrome.

In this study, we used LDLR^−/− mice to investigate the effect of LPS and SFA on proinflammatory cytokine expression in hepatocytes. We also performed in vitro study to elucidate the involvement of sphingolipid metabolism in the regulation of hepatic inflammation by LPS and SFA.

MATERIALS AND METHODS

Animals, diets, and treatment.

Male LDLR^−/− mice were purchased from Jackson Laboratory (Bar Harbor, ME) and housed at the animal facility of the Veterans Affairs Medical Center in Charleston, SC. The male mice were used in this study because it has been shown that female mice are protected against HFD-induced metabolic syndrome (34). The animal protocol was approved by the Institutional Animal Care and Use Committee. All mice were maintained on a 12-h light/dark cycle in a pathogen-free environment and had ad libitum access to water and food. Mice were fed different diets (Table 1) purchased from Envigo RMS, Inc. (Indianapolis, IN), including low-fat diet, which is similar to the chow diet, HFD with low content of the major SFA palmitic acid [PA (LP-HFD)], or HFD with high content of PA (HP-HFD) for 20 wk. The coconut-based LP-HFD and lard-based HP-HFD contain the same amount of protein (23.5%), carbohydrate (27.3%), and fat (34.3%). LP-HFD and HP-HFD contain 25% and 40% of long-chain SFA, respectively. Furthermore, LPS-HFD and HP-HFD contain 2.8% and 8% of PA, respectively (33). During the last 12 wk, half the mice received intraperitoneal injection of LPS (Escherichia coli serotype 055:B5, Sigma, St. Louis, MO), 25 μg in 200 μl of PBS per mouse, once a week whereas the other half received an equivalent amount of PBS, the vehicle for LPS. After the treatments, blood metabolic parameters and livers in six groups (6–9 mice per group) were analyzed. The dose of LPS and the administration by weekly intraperitoneal injection were reported to be effective to induce atherosclerosis by the previous studies (26, 52). We have also used this method in our published studies (21, 22).

Table 1.

Saturated fat and palmitic acid contents in different diets

Ingredients	LFD (TD. 2018)	LP-HFD (TD. 08500)	HP-HFD (TD. 06414)
Protein, % of weight	18.4	23.5	23.5
Fat, % of weight	6.2	34.3 (Coconut-based)	34.3 (Lard-based)
Carbohydrate, % of weight	44.2	27.3	27.3
Calories from protein, % kcal	24.0	18.4	18.4
Calories from fat, % kcal	18.0	60.3	60.3
Calories from carbohydrate, % kcal	58.0	21.3	21.3
Long-chain saturated fatty acid, % of total fatty acid	0.9	27.0	40.0
Long-chain monounsaturated fatty acid, % of total fatty acid	1.3	2.0	47.0
Long-chain polyunsaturated fatty acid, % of total fatty acid	3.4	5.5	16.0
Palmitic acid, % of total fatty acid	0.7	2.8	8.0

HFD, high-fat diet; HP-HFD; HFD with high content of palmitic acid; LDF, low-fat diet; LP-HFD, HFD with low content of palmitic acid. The links for the detailed information of three diets: TD. 2018: https://www.envigo.com/resources/data-sheets/2018-datasheet-0915.pdf, TD. 08500: https://www.envigo.com/resources/data-sheets/08500.pdf, TD. 06414: https://www.envigo.com/resources/data-sheets/06414.pdf.

Metabolic measurements.

Blood samples were obtained under the fasted condition and glucose level was determined using a Precision QID glucometer (MediSense, Inc., Bedford, MA). Serum cholesterol and triglycerides were assayed using Cholestech LDX Lipid monitoring System (Fisher Scientific, Pittsburgh PA). Serum fatty acids were determined using the EnzyChrom free fatty acid kit (BioAssay Systems, Hayward, CA). Serum fasting insulin was assayed using the Ultra Sensitive Insulin ELISA Kit (Crystal Chem, Inc., Downers Grove, IL). Fasting whole-body insulin sensitivity was estimated with the homeostasis model assessment of insulin resistance according to the formula [fasting plasma glucose (mg/dl) × fasting plasma insulin (μU/ml)]/405 (56).

Immunohistochemical analysis of protein expression.

Liver tissues were fixed in 4% paraformaldehyde for 10 min, and frozen sections were made using a cryostate. Immunohistochemical analysis was performed as described previously (23) with anti-F4/80 (cat. no. MCA-497, Bio-Rad Laboratories, Inc., Hercules, CA), anti-IL-6 (cat. no. ab-6672, Abcam, Cambridge, MA), anti- GPR40 (cat. no. sc-32905, Santa Cruz Biotechnology, Dallas, TX), or anti-CD36 (cat. no. NB-400-144, Novus Biologicals, LLC., Littleton, CO) antibodies. Counterstaining was performed with hematoxylin. Photomicrographs of tissue sections were taken using an Olympus BX53 digital microscope with Cellsens digital image software (Olympus American, Inc., Center Valley, PA), and the positively immunostained area was quantified with Image-Pro Plus 6 (Media Cybernetics, Rockville, MD).

Oil red O and picric Sirius red staining.

For oil red O staining, the frozen sections were fixed with 10% formalin for 10 min, placed in 60% isopropyl alcohol, and stained in 0.5% Oil Red O solution for 10 min. The slides were transferred to 60% isopropyl alcohol, rinsed in distilled water, and processed for hematoxylin counter-staining. For picric Sirius red staining, the sections were fixed with 10% formalin for 10 min, incubated with a 0.1% Sirius Red solution dissolved in aqueous saturated picric acid for 1 h, washed in acidified water (0.5% acetic acid), dehydrated, and mounted on slides. Sections were then examined using a Polarizing microscope.

Cell culture and treatment.

The mouse primary hepatocytes were purchased from Thermo Fisher Scientific, Inc. (Miami, OK) and cultured with DMEM/F-12K medium containing 10% heat-inactivated fetal calf serum (HyClone, Logan, UT) by following the instructions from the manufacturer. The murine macrophage cell line RAW264.7 has been used extensively in the investigations of the role of macrophages in inflammation-related diseases (37). RAW264.7 cells were purchased from the American Type Culture Collection (Manassas, VA) and grown in DMEM (American Type Culture Collection) supplemented with 10% heat-inactivated fetal calf serum. The cells were maintained in a 37°C, 90% relative humidity, 5% CO₂ environment. For cell treatment, LPS from E. coli was used. The LPS was highly purified by phenol extraction and gel filtration chromatography and was cell-culture tested. PA (Sigma) used in this study was bovine serum albumin-free (40). To prepare PA, PA was dissolved in 0.1 N NaOH and 70% ethanol at 70°C to make PA solution at a concentration of 50 mM. The solution was kept at 55°C for 10 min, mixed, and brought to room temperature. For coculture, hepatocytes and RAW264.7 macrophages were grown in 12-well Corning Transwell plates (Fisher Scientific, Pittsburgh, PA), a noncontact coculture system, with 2 compartments separated by a polycarbonate membrane with 0.4-μm pores. Hepatocytes were grown at the lower compartment and RAW264.7 macrophages were cultured in the upper compartment. To treat cells in the Transwell plates, LPS and PA were added to the culture medium in both upper and lower compartments at the same time to ensure equal exposure of hepatocytes and macrophages to LPS and PA. In all experiments unless otherwise specified, hepatocytes were treated with 5 ng/ml of LPS, 100 μM of PA, or both 5 ng/ml of LPS and 100 μM of PA, and RAW264.7 cells were treated with 1 ng/ml of LPS, 100 μM of PA, or both 1 ng/ml of LPS and 100 μM of PA. After the treatment, culture medium was collected for quantification of IL-6 protein, and cells were harvested for quantification of IL-6 mRNA.

Enzyme-linked immunosorbent assay.

IL-6 and TNFα in medium were quantified using sandwich ELISA kits according to the protocol provided by the manufacturer (Biolegend, San Diego, CA).

Real-time PCR.

Real-time PCR was performed as described previously (17). The Beacon designer software (PREMIER Biosoft International, Palo Alto, CA) was used for primer designing (mouse IL-6: 5′ primer sequence, TGGAGTCACAGAAGGAGTGGCTAAG; 3′ primer sequence, TCTGACCACAGTGAGGAATGTCCAC. Primers were synthesized by Integrated DNA Technologies, Inc. (Coralville, IA). Mouse glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as a control (5′ primer sequence, CTGAGTACGTCGTGGAGTC; 3′ primer sequence, AAATGAGCCCCAGCCTTC). Data were analyzed with the iCycler iQ software. The average starting quantity of fluorescence units was used for analysis. Quantification was calculated using the starting quantity of targeted cDNA relative to that of GAPDH cDNA in the same sample.

PCR arrays.

First-strand cDNA was synthesized from RNA using RT² First Strand Kit (SuperArray Bioscience Corp., Frederick, MD). Mouse TLR pathway-focused PCR Arrays (SuperArray Bioscience Corp.) were performed using 2× SuperArray RT² qPCR master mix and the first strand cDNA by following the instructions from the manufacturer.

Immunoblotting.

Cytoplasmic protein was isolated as described previously (22). The concentration of protein was determined using a protein assay kit (Bio-Rad). Fifty micrograms of protein from each sample were electrophoresed in a 10% polyacrylamide gel. After transferring proteins to a polyvinylidene fluoride membrane, immunoblotting was performed using antibodies against GPR40, CD36 (cat. no. sc-32905 and sc-7309, Santa Cruz Biotechnology, Inc.), or GAPDH (cat. no. 2118, Cell Signaling Technology, Inc., Danvers, MA). The proteins were visualized by incubating the membrane with chemiluminescence reagent (NEN Life Science Products, Boston, MA) for 1 min and exposing the membrane to X-ray films for 1–30 min.

Lipidomics.

Hepatocytes were collected, fortified with internal standards, extracted with ethyl acetate/isopropyl alcohol/water (60:30:10, vol/vol/vol), evaporated to dryness, and reconstituted in 100 μl of methanol. Simultaneous ESI/MS/MS analyses of sphingoid bases, sphingoid base 1-phosphates, ceramides (CERs), and sphingomyelins (SMs) were performed on a Thermo Finnigan TSQ 7000 triple quadrupole mass spectrometer operating in a multiple reaction monitoring positive ionization mode. The phosphate contents of the lipid extracts were used to normalize the MS measurements of sphingolipids. The phosphate contents of the lipid extracts were measured with a standard curve analysis and a colorimetric assay of ashed phosphate (50).

Apoptosis studies.

To assess cell apoptosis, mono-oligo-nucleosomes (histone-associated DNA fragments) were quantitated with the Cell Death Detection ELISA kit (Sigma-Aldrich, St. Louis, MO) by following the manufacturer’s instructions. Briefly, hepatocytes were incubated for 18 h with palmitate in the absence or presence of LPS. After the incubation, cells were lysed, and the cytosol protein was transferred to the wells of a streptavidin-coated plate supplied by the manufacturer. A mixture of antihistone-biotin and anti-DNA-peroxidase antibodies was added to the cell lysate and incubated for 2 h. After washing, 2,2′-azinobis-3-ethyl-benzothiazoline-6-sulfonic acid substrate was added to each well. Absorbance at 405 nm was measured with a VersaMax microplate reader (Molecular Devices, Sunnyvale, CA).

Statistical analysis.

GraphPad Instat statistical software (Version 5.0) (GraphPad Software, Inc., La Jolla, CA) was used for statistical analysis. The one-way analysis of variance (ANOVA) was used to compare the data between multiple groups. To determine the statistical significance of differences between two experimental groups, parametric analysis using Student’s t-test was performed for data with normal distribution and nonparametric analysis using Mann-Whitney test was performed for data without normal distribution. A value of P < 0.05 was considered significant.

RESULTS

Effect of HFD with or without LPS on metabolic parameters.

The assays on the metabolic parameters (Table 2) at the end of the study showed that either LP-HFD or HP-HFD increased bodyweight, cholesterol, LDL, insulin, and insulin resistance, but HP-HFD increased more bodyweight, insulin, and insulin resistance than LP-HFD. It appeared that HP-HFD also increased more total cholesterol, LDL, and triglyceride than LP-HFD, but the differences are not statistically significant. LPS alone slightly increased glucose, insulin, and insulin resistance, but the addition of LPS to HFD reduced HFD-induced body weight, insulin, and insulin resistance.

Table 2.

Metabolic parameters of mice fed with different diets and treated with or without LPS

	LFD without LPS	LFD with LPS	LP-HFD without LPS	LP-HFD with LPS	HP-HFD without LPS	HP-HFD with LPS
Mouse number	7	7	6	9	6	8
Body weight, g	31 ± 2	31 ± 3	40 ± 4a	38 ± 4a	49 ± 5a,b	42 ± 7a
Glucose, mg/dl	191 ± 9	238 ± 12c	257 ± 23c	204 ± 7	212 ± 12	216 ± 13
Cholesterol, mg/dl	355 ± 19	391 ± 27	522 ± 40d	532 ± 28d	659 ± 87d	567 ± 82d
LDL, mg/dl	230 ± 20	265 ± 23	363 ± 43e	369 ± 33e	444 ± 140e	465 ± 87e
HDL, mg/dl	90 ± 3	76 ± 4	122 ± 11f	91 ± 5	94 ± 5	105 ± 11
Triglyceride, mg/dl	164 ± 13	182 ± 9	179 ± 15	268 ± 34g	241 ± 35g	188 ± 43
Free fatty acid, mg/dl	424 ± 37	413 ± 39	490 ± 29	516 ± 23	401 ± 40	370 ± 26
Insulin, ng/ml	0.30 ± 0.08	1.23 ± 0.8	1.45 ± 0.32h	0.73 ± 0.13	6.09 ± 1.50h,i	2.24 ± 0.77
HOMA-IR	3.5 ± 0.9	18.1 ± 12.3j	23.0 ± 5.1k	9.2 ± 1.6	79.7 ± 19.6l	29.9 ± 10.3

HFD, high-fat diet; HOMA-IR, homeostasis model assessment of insulin resistance; HP-HFD; HFD with high content of palmitic acid; LDF, low-fat diet; LP-HFD, HFD with low content of palmitic acid.

^aP < 0.01 vs. LFD without or with LPS;

^bP < 0.01 vs. LP-HFP without or with LPS;

^cP < 0.05 vs. LFD without LPS;

^dP < 0.05 vs. LFD without or with LPS;

^eP < 0.05 vs. LFD without LPS;

^fP < 0.05 vs. LFD without LPS or LP-HFD with LPS;

^gP < 0.05 vs. LFD without LPS;

^hP < 0.05 vs. LFD without LPS;

ⁱP < 0.05 vs. LP-HFD without LPS;

^jP < 0.05 vs. LFD without LPS;

^kP < 0.05 vs. LFD without LPS;

^lP < 0.05 vs. LP-HFD without LPS.

HP-HFD in combination with LPS increases macrophage content and IL-6 expression in livers.

It is known that either SFA or LPS promotes hepatic inflammation (14, 27). To determine how the copresence of SFA and LPS affects hepatic inflammation, we first determined the effect of HP-HFD with LPS on macrophage content in livers using immunohistochemical staining of murine macrophage-specific marker F4/80 (20). Results showed that LPS or HP-HFD alone increased F4/80 expression in livers significantly, but LP-HFD slightly decreased it (Fig. 1, A–C). Interestingly, the combination of HP-HFD and LPS stimulated more F4/80 expression than HP-HFD alone, indicating that the combination of HP-HFD and LPS increased more macrophage infiltration into liver than HP-HFD alone. We then performed immunohistochemical staining of IL-6, a major proinflammatory cytokine involved in hepatic inflammation (28). Figure 1D showed that both hepatocytes and mononuclear cells expressed IL-6. Surprisingly, LPS alone had no effect on IL-6 expression but either LP-HFD or HP-HFD increased IL-6 expression (Fig. 1, D and E). Furthermore, the combination of HP-HFD and LPS but not the combination of LP-HFD and LPS increased higher IL-6 expression than LPS or HP-HFD alone (Fig. 1, D and E).

Fig. 1.

The effect of LPS and diets on hepatic inflammation. After mice were treated with LPS or vehicle PBS in combination with LFD, LP-HFD, or HP-HFD, livers were dissected and subjected to immunohistochemical staining of F4/80 to detect macrophages. Representative photomicrographs of hepatic tissue sections with F4/80 immunostaining for all 6 groups (A) and quantification of F4/80-positive staining area (C) are shown. As negative control, immunostaining using a control IgG (B) is shown. Also, representative photomicrographs of hepatic tissue sections with IL-6 immunostaining (D) and quantification of IL-6 positive staining area (E) are shown. Furthermore, liver tissue sections were stained with oil red O (F) and Sirius red (H). The photomicrographs and quantification of oil red O (G) and Sirius red positive area (I) are shown. The data presented are means ± SD (n = 6–9).

HP-HFD increases fat content and fibrosis in livers.

In addition to the studies on hepatic macrophage content and IL-6 expression, we also investigated the effect of HP-HFD in combination with LPS on fat accumulation and fibrosis in livers. Interestingly, results from oil red O staining showed that HP-HFD is much more potent than LP-HFD in inducing fat accumulation, but LPS reduced fat accumulation stimulated by HP-HFD (Fig. 1, F and G), indicating that PA but not LPS plays a critical role in fat accumulation. Results from Sirius red staining show that although LPS, LP-HFD, or HP-HFD induced collagen content, the addition of LPS to LP-HFD or HP-HFD further increased it (Fig. 1, H and I), indicating that HFD regardless of PA content and LPS cooperatively increased hepatic fibrosis.

LPS and PA synergistically stimulate proinflammatory gene expression in hepatocytes in vitro.

To understand how HP-HFD and LPS cooperatively stimulate IL-6 expression in livers, we performed in vitro studies in which mouse primary hepatocytes were treated with LPS, PA, or LPS plus PA. Results from the time course study showed that LPS stimulated IL-6 secretion whereas PA had no effect, but PA robustly augmented LPS-stimulated IL-6 secretion during 12–24 h (Fig. 2A). Consistently, the quantification of IL-6 mRNA using real-time PCR also showed that LPS stimulated IL-6 mRNA expression at 4 h whereas PA had no effect, but PA enhanced LPS-induced IL-6 mRNA expression during 12–24 h (Fig. 2B). These findings clearly demonstrated that PA boosted the stimulatory effect of LPS on IL-6 secretion and mRNA expression.

Fig. 2.

LPS in combination with PA increases proinflammatory cytokine expression in hepatocytes. Hepatocytes were treated LPS, PA or LPS plus PA for different times as indicated and IL-6 secreted into the medium (A) and IL-6 mRNA (B) at each time point were quantified using ELISA and real-time PCR, respectively. PA, palmitic acid.

To determine further the effect of LPS plus PA on proinflammatory gene expression in hepatocytes, we profiled the hepatic gene expression in response to LPS plus PA using a TLR signaling pathway PCR array. Results showed that although either LPS or PA stimulated the expression of many genes, the combination of LPS and PA synergistically upregulated a number of proinflammatory molecules, including MCP-1, CD86, CSF-3, IL-1α, IL-1β, IL-6, and Cox-2 (Table 3). For example, although LPS and PA increased IL-1α mRNA expression by 14.08- and 3.66-fold, respectively, the combination of LPS and PA increased IL-1α expression by 123.81-fold.

Table 3.

The synergistic effect of LPS and PA on proinflammatory gene expression in hepatocyte

	Ct				Fold change by
Genes	Control	LPS	PA	LPS + PA	LPS	PA	LPS + PA
MCP-1	26.03	21.05	23.79	19.95	31.59	4.71	67.62
CD14	21.04	19.90	20.07	19.34	2.20	1.97	3.25
CD80	29.54	29.57	28.11	27.70	0.98	2.70	3.59
CD86	30.70	29.95	29.38	27.42	1.68	2.51	9.71
CLEC4E	27.07	24.65	25.03	23.72	5.36	4.09	10.14
CSF-2	31.60	26.93	31.02	27.22	25.43	1.50	20.80
CSF-3	32.51	28.16	31.84	26.89	20.33	1.59	48.97
CXCL10	23.85	21.47	22.01	20.39	5.19	3.59	10.98
FOS	30.39	30.13	29.74	28.91	1.20	1.57	2.79
IFNB1	32.90	32.28	31.19	30.90	1.54	3.26	3.99
IL-10	33.08	32.39	32.25	31.29	1.61	1.78	3.46
IL-1α	32.56	28.75	30.69	25.61	14.08	3.66	123.81
IL-1β	32.17	29.86	31.84	27.07	4.95	1.26	34.38
IL-1R1	32.88	31.94	30.99	31.44	1.91	3.71	2.70
IL-2	33.58	31.88	31.26	32.29	3.24	5.00	2.44
IL-6	31.71	28.61	31.40	26.32	8.58	1.24	41.78
Irak2	28.36	27.17	28.45	27.17	2.27	0.94	2.28
Lta	32.66	32.37	31.76	30.89	1.22	1.86	3.41
Mapk8ip3	31.39	30.05	30.25	29.67	2.53	2.22	3.31
NFKB2	31.18	30.24	30.17	29.27	1.92	2.00	3.75
NFKBIA	22.62	20.67	22.06	21.15	3.86	1.48	2.76
NFKBIB	30.89	30.15	30.55	29.68	1.67	1.26	2.30
Peli1	27.11	26.40	27.07	25.93	1.64	1.03	2.27
Ppara	33.29	32.40	32.99	31.36	1.86	1.23	3.81
COX-2	28.85	25.59	26.97	22.99	9.59	3.69	58.19
Ticam1	31.74	31.88	30.85	30.50	0.91	1.86	2.37
Tlr1	29.61	28.78	28.55	28.27	1.77	2.08	2.52

Hepatocytes were treated with 1 ng/ml of LPS, 100 µM of PA, or both for 24 h. For control, hepatocytes were not cultured without LPS and PA. After treatment, cells were harvested and RNA was isolated from duplicate samples, combined, and subjected to PCR array study as described in methods. The GAPDH mRNA was used as a housekeeping gene for control. To compare the gene expression in cells treated with LPS, PA, or LPS + PA with that in control cells, ΔCt was first calculated by the following formula: ΔCt = Ct in LPS, PA, or LPS + PA-treated cells − Ct in control cells. The gene expression in cells treated with LPS, PA, or LPS + PA was presented as the fold of the control gene expression and calculated as 2^ΔCt. CLEC4E, C- Type lectin domain family 4 member E; CSF2, colony stimulating factor 2; Ct, cycle threshold; CXCL10, C-X-C motif chemokine 10; IFNB1, interferon beta 1; IL-1R1, interleukin 1 receptor type 1; Irak2, interleukin-1 receptor-associated kinase 2; Lta, lymphotoxin alpha; PA, palmitic acid; Peli1, pellino E3 ubiquitin protein ligase 1; Ppara, peroxisome proliferator activated receptor alpha; Ticam1, toll like receptor adaptor molecule 1; Tlr1, Toll-like receptor 1; MAPK8IP3, mitogen-activated protein kinase 8 interacting protein 3; NFKB2, nuclear factor kappa B subunit 2; NFKBIA, NFKB inhibitor alpha.

Crosstalk between hepatocytes with macrophages further increases proinflammatory cytokine expression.

It is known that NAFLD is characterized by an increase in monocyte-derived macrophages in the liver (46). Under this circumstance, macrophages may cross talk with hepatocytes to upregulate proinflammatory gene expression. In this study, we cocultured hepatocytes with RAW264.7 macrophages in a two-chamber system without direct contact and determined the effect of the coculture of hepatocytes and macrophages on proinflammatory gene expression in both hepatocytes and macrophages in response to LPS, PA, or LPS plus PA. Results showed that although LPS and PA synergistically stimulated secretion of IL-6 (Fig. 3, A and B) and TNFα (Fig. 3, E and F) in the separate culture of hepatocytes or macrophages, the extent of the stimulation on IL-6 (Fig. 3, C and D) and TNFα (Fig. 3, G and H) secretion was further increased when hepatocytes and macrophages were cocultured. Quantification of IL-6 mRNA using real-time PCR showed that LPS plus PA stimulated a higher expression of IL-6 mRNA in either hepatocytes or macrophages in the coculture than that in the separated culture (Fig. 3, I and J).

Fig. 3.

Hepatocyte and RAW264.7 macrophages were cultured separately (A and B; E and F) or together (C and G) and exposed to LPS, PA, or LPS plus PA for 24 h. After the treatment, IL-6 (A–C) and TNFα (E–G) in culture medium were quantified using ELISA. The amount of IL-6 (D) and TNFα (H) secretion by hepatocytes, RAW264.7 macrophages or coculture of hepatocytes and RAW264.7 macrophages was compared. Hepatocytes cultured alone or cultured with RAW264.7 macrophages in the coculture system were exposed to LPS, PA, or LPS plus PA for 24 h. After the treatment, IL-6 mRNA was isolated and compared between hepatocytes cultured alone and those in the coculture (I) and between RAW264.7 macrophages cultured alone and those in the coculture (J). The data (mean ± SD) presented are representative of three experiments with similar results. PA, palmitic acid.

GPR40 and CD36 are involved in the upregulation of proinflammatory genes by LPS and PA.

To understand how PA boosted the stimulatory effect of LPS on proinflammatory gene expression in hepatocytes, we first determined the involvement of GPR40 and CD36 in the stimulation. GPR40, a long-chain fatty acid receptor, is expressed in pancreatic beta cells (15) and other types of cells, such as hepatocytes (19, 45). CD36, a scavenger receptor also known as fatty acid translocase (12) is expressed by a large number of cells, including hepatocytes (41). Our immunoblotting showed that both GPR40 and CD36 are expressed by hepatocytes in livers (Fig. 4, A and B) and cultured hepatocytes (Fig. 4C). Furthermore, GW1100 and sulfo-N-succinimidyl oleate, which are pharmacological inhibitors of GPR40 and CD36, respectively, inhibited IL-6 (Fig. 4, D and E) and MCP-1 secretion (Fig. 4F) stimulated by LPS or LPS plus PA, suggesting that both GPR40 and CD36 are involved in the upregulation of proinflammatory genes by LPS or the combination of LPS and PA.

Fig. 4.

The hepatic expression of GPR40 and CD36 in vivo and in vitro, and the involvement of GPR40 and CD36 in the stimulation of cytokine secretion by LPS or LPS plus PA. Immunostaining of GPR40 (A) and CD36 (B) in livers of mice fed high-fat diet. Immunoblotting of GPR40 and CD36 in cultured mouse hepatocytes (C). GAPDH was detected as control. Hepatocytes were treated with LPS, PA, or LPS plus PA in the absence or presence of 10 μM of GW1100 (D and F), a selective GPR40 antagonist, or 50 or 100 μM of sulfo-N-succinimidyl oleate (SSO) (E and F), a selective inhibitor of CD36, for 24 h. After the treatment, cytokine secreted into medium was quantified using ELISA. The data (mean ± SD) presented are representative of three experiments with similar results. GPR, G protein-coupled receptor; PA, palmitic acid.

CER de novo synthesis plays an essential role in the cooperative upregulation of proinflammatory genes by LPS and PA in hepatocytes.

To understand further how LPS and PA cooperate to promote gene expression, we focused on sphingolipid metabolism because it is known that treatment of cells with SFA such as PA or stearic acid increases CER production (17, 24). Results from lipidomic analysis showed that PA or the combination of LPS and PA but not LPS markedly increased total CER and C16- CER production (Fig. 5, A and B and Table 4). Results also showed that although PA increased dhC16-CER, the combination of LPS and PA increased more dhC16- CER (Fig. 5C and Table 4). Given that dhC16-CER is an intermediate during CER de novo synthesis (44), these results indicate that PA and LPS synergistically increased CER de novo synthesis. Interestingly, results showed that LPS increased sphingosine 1 phosphate (S1P) production and LPS in combination with PA further increased it (Fig. 5D and Table 4). Because hydrolysis of SM is a major pathway for CER production (17), we also analyzed the effect of LPS, PA, or LPS plus PA on SM hydrolysis. Results showed that LPS, PA, or LPS plus PA did not have significant effect on the hydrolysis of total SM (Fig. 5E) or C16-SM (Fig. 5F).

Fig. 5.

The effect of LPS, PA, or LPS plus PA on sphingolipid production in hepatocytes. Hepatocytes were treated with LPS, PA, or LPS plus PA for 12 h, and the cells were harvested for quantification of total CER (A), C16-CER (B), dhC16-CER (C), sphingosine 1 phosphate (D), total sphingomyelin (E), and C16-sphingomyelin (F) using Lipidomics. The data are mean ± SD of triplicate samples. CER, ceramide; PA, palmitic acid.

Table 4.

Effect of LPS, palmitate, or LPS + PA on CER and S1P levels

	C16-CER	C18-CER	C20-CER	C24-CER	C24:1-CER	dhC16-CER	S1P	Total CER
Control	2.265 ± 0.220	0.180 ± 0.021	0.170 ± 0.018	2.210 ± 0.283	1.948 ± 0.199	0.082 ± 0.006	0.019 ± 0.004	7.540 ± 0.810
LPS	2.539 ± 0.114	0.182 ± 0.016	0.150 ± 0.006	2.809 ± 0.160	1.811 ± 0.103	0.104 ± 0.002	0.060 ± 0.007	8.335 ± 0.444
PA	5.680 ± 0.404	0.520 ± 0.055	0.492 ± 0.061	2.252 ± 0.201	2.245 ± 0.048	0.132 ± 0.016	0.031 ± 0.001	12.608 ± 0.876
LPS + PA	5.638 ± 0.479	0.413 ± 0.053	0.437 ± 0.064	2.623 ± 0.126	2.182 ± 0.134	0.158 ± 0.012	0.093 ± 0.009	14.741 ± 1.070

Unit: pmole/nmole Pi. CER, ceramide; PA, palmitic acid; S1P, sphingosine 1 phosphate.

To determine if CER de novo synthesis is involved in the stimulation by LPS and PA of proinflammatory gene expression, we applied myriocin, a specific inhibitor for serine palmitoyltransferase (SPT). SPT is an enzyme involved in the condensation of serine and palmitoyl-CoA to generate 3-ketosphinganine, a rate-limiting reaction in CER de novo synthesis (16), and the effect of myriocin on the inhibition of CER synthesis has been well established (3, 27). Results showed that myriocin inhibited the stimulation of IL-6 secretion by LPS or LPS plus PA in a concentration-dependent manner (Fig. 6A). Time course study also showed that myriocin inhibited IL-6 secretion stimulated by LPS or LPS plus PA at 8, 18, and 24 h (Fig. 6B). Furthermore, study using quantitative real-time PCR showed that myriocin robustly inhibited the upregulation of IL-6 mRNA by LPS or LPS plus PA at 18 and 24 h (Fig. 6C), suggesting that CER de novo synthesis is involved in IL-6 transcription stimulated by LPS or LPS plus PA.

Fig. 6.

The involvement of the de novo synthesis of CER in IL-6 expression stimulated by LPS, PA or LPS plus PA. A: hepatocytes were treated with LPS, PA, or LPS plus PA in the absence or presence of 5 or 10 μM of myriocin, a selective inhibitor of serine palmitoyltransferase (SPT), for 24 h. After the treatment, IL-6 in culture medium was quantified. B: hepatocytes were treated with LPS, PA, or LPS plus PA in the absence or presence of 10 μM of myriocin for different times as indicated. After the treatment, IL-6 in culture medium was quantified. C: hepatocytes were treated with LPS, PA, or LPS plus PA in the absence or presence of 10 μM of myriocin for 18 or 24 h. After the treatment, IL-6 mRNA was quantified using real-time PCR. D: hepatocytes were treated with 5 or 10 ng/ml of LPS in the absence or presence of 100 μM of PA for 24 h. After the treatment, histone-associated DNA fragments were quantified as described in methods. The data (mean ± SD) presented are representative of three experiments with similar results. CER, ceramide; Ctl, control; PA, palmitic acid.

Effect of LPS and PA on hepatocyte apoptosis.

It is known that PA induces apoptosis in hepatocytes by increasing CER production (27, 54). Because our study showed that the combination of LPS and PA increased CER (Fig. 5A) and S1P (Fig. 5D), which is known to suppress CER-induced apoptosis (7) we determined the effect of LPS, PA, or LPS plus PA on hepatocyte apoptosis. Results showed that although PA strongly induced apoptosis, the addition of LPS to PA modestly reduced the degree of PA-induced apoptosis in a concentration-dependent manner (Fig. 6D).

DISCUSSION

The present study utilized both an animal model and cell cultures to test our hypothesis that SFA and LPS interact to stimulate a strong inflammatory response in hepatocytes. In the animal study, metabolic analysis showed that HP-HFD increased more body weight, insulin, and insulin resistance than LP-HFD. Because it is generally accepted that insulin resistance is closely associated with chronic inflammation (51), the above finding indicates that HP-HFD is more potent than LP-HFD in stimulating inflammation. Consistently, our immunohistochemical study showed that HP-HFD induced more macrophages infiltrating into livers than LP-HFD. Furthermore, our study also showed that the addition of LPS to HP-HFD led to a higher hepatic IL-6 expression than HP-HFD alone or the addition of LPS to LP-HFD (Fig. 1E). Our in vitro study with primary mouse hepatocytes showed that PA amplified LPS-stimulated expression of IL-6 and other proinflammatory genes, supporting our in vivo observation that HP-HFD and LPS act in concert to stimulate hepatic inflammation.

In addition to hepatocytes, our previous study has shown that PA also amplifies LPS-induced inflammatory gene expression in macrophages (17). Because it is known that the hepatocyte and macrophage are two major types of cells involved in hepatic inflammation (25), the amplification of LPS-induced inflammatory gene expression by PA in both hepatocytes and macrophages is likely to result in a strong upregulation of proinflammatory response in the liver. Furthermore, our studies with the coculture system indicate that hepatocytes and macrophages had a crosstalk in response to PA and LPS, resulting in a higher expression of proinflammatory cytokines such as IL-6 and TNFα (Fig. 3, D and H). In support of our findings, a previous study has shown that macrophages, including Kupffer cells play an important role in the initial response to LPS by producing proinflammatory cytokines, which subsequently stimulate hepatocytes to release more proinflammatory cytokines (25).

In our investigation on the mechanisms involved in the cooperative upregulation of proinflammatory gene expression in hepatocytes by LPS and PA, we focused on sphingolipid metabolism because it is known that sphingolipid metabolism is involved in gene expression (17, 39). We found that LPS and PA increased CER production in hepatocytes primarily by synergistically stimulating CER de novo synthesis. It is interesting to note that this finding is different from that in macrophages, in which LPS and PA increased CER production mainly by synergistically stimulating acid SM-mediated SM hydrolysis (17). Inconsistent with our observations, Martínez et al. (27) reported that SFAs such as myristic acid and PA increased CER de novo synthesis that in turn contributed to inflammation-associated lipotoxicity and steatohepatitis (22). However, our current study reported for the first time that LPS augmented PA-induced CER de novo synthesis in hepatocytes. Furthermore, we demonstrated that CER de novo synthesis plays an important role in the upregulation of proinflammatory gene expression by LPS and PA as we showed that myriocin, an SPT specific inhibitor, significantly inhibited the stimulatory effect of LPS and PA on IL-6 secretion and expression. The role of CER in the expression of proinflammatory cytokines stimulated by LPS and PA was also demonstrated by Schwartz et al. (40), who showed that the amplified LPS response by PA was blocked by the inhibitor of CER de novo synthesis but rescued by addition of cell-permeable CER to cells.

GPR40 is a G protein-coupled receptor specific for free long-chain fatty acids (15) whereas CD36 is a scavenger receptor for a number of ligands such as free long-chain fatty acids, oxidized low-density lipoprotein, collagen, and thrombospondin (1). It has been shown that hepatocytes express GPR40 and CD36 that interact with long-chain free fatty acids (45, 57). Suh et al. (45) reported that free fatty acids such as linoleic acid stimulated expression of Ca²⁺/phospholipase C, cytosolic phospholipase A2, and peroxisome proliferator-activated receptor through GPR40 in chicken hepatocytes. Wilson et al. (53) showed that hepatocyte-specific disruption of CD36 attenuated fatty liver and improved insulin sensitivity in HFD-fed mice, indicating that CD36 is involved in HFD-induced fat accumulation and insulin resistance in hepatocytes. All these findings indicate that the action of PA on hepatocytes is mediated by multiple receptors such as GPR40 and CD36. Our present study also showed that both GPR40 and CD36 were expressed by hepatocytes and the inhibitor for either GPR40 or CD36 inhibited expression of proinflammatory genes stimulated by LPS plus PA in hepatocytes, confirming that multiple receptors are involved in the augmentation by PA of LPS-triggered inflammatory signaling.

In addition to the finding that GPR40 or CD36 inhibition attenuated IL-6 secretion induced by LPS plus PA, it is interesting to find that GPR40 or CD36 inhibition also attenuated IL-6 secretion from hepatocytes stimulated by LPS without PA. These findings suggest that GPR40 or CD36 is associated with TLR4. To understand how CD36 interacts with TLR4 signaling, Stewart et al. (43) have shown that CD36 forms a heterodimer with TLR4. It is thus expected that targeting CD36 would interfere with TLR4 signaling elicited by LPS. Based on our finding, we postulated that like CD36, GPR40 also forms a complex with TLR4 on the cell surface. Further studies are necessary to define how GPR40 interacts with TLR4 and the role of GPR40-TLR4 interaction in the inflammatory signaling in hepatocytes.

It has been well documented that PA induces hepatic apoptosis (27, 29) via CER production (11, 27). However, given that the addition of LPS to PA not only increased CER but also S1P, which has antiapoptotic effect (7), it is interesting to determine how the addition of LPS to PA alters PA-induced hepatic apoptosis. Results showed that although PA strongly induced hepatic apoptosis, the addition of LPS to PA reduce the extent of PA-induced apoptosis. This finding is interesting as it indicates that the copresence of LPS and PA may reduce PA-induced hepatocyte death and thus maintains a high level of proinflammatory cytokine production from hepatocytes.

In this study, we investigated the combination effect of LPS and SFA on hepatic inflammation by feeding mice an HFD with high PA content (HP-HFD) and LPS injection. Although the study successfully demonstrated that LPS and HP-HFD cooperatively induced hepatic inflammation (Fig. 1, C and E), it also showed some results that are not in line with the clinical observations. First, the metabolic data showed that although LPS alone had no effect on bodyweight but increase insulin resistance as expected, the addition of LPS to HP-HFD reduced HP-HFD-induced body weight and insulin resistance (Table 2), which is different from the clinical observation that patients with metabolic endotoxemia have increased body weight and insulin resistance (6). Second, although LPS alone slightly increased fat content in liver, the addition of LPS to HP-HFD reduced HP-HFD-induced fat accumulation (Fig. 1G), which is different from the clinical observation that patients with metabolic endotoxemia have increased fat content in liver (6). Although the cause of these results remains unknown, it is likely associated with the different LPS deliveries in our animal models and the patients with metabolic endotoxemia. In our animal study, LPS was injected once a week but in patients with metabolic endotoxemia, LPS is translocated from the intestine to the bloodstream. The different LPS deliveries in our animal models and in patients with metabolic endotoxemia may elicit different systemic responses, leading to different effects on body weight, insulin resistance, and hepatic fat accumulation. Obviously, these results reflect the limitations of the animal model in this study.

Another limitation of this study is the lack of the complete fatty acid profiles of the HFDs (TD.08500 and TD.06414) although the contents of saturated, monounsaturated, and polyunsaturated fatty acids are available (Table 1). Obviously, the complete fatty acid profiles of the HFDs are important because they would allow us to compare the contents of each fatty acid between the HFDs and determine if fatty acids other than SFA may potentially play a role in the hepatic inflammation.

In conclusion, this study demonstrated that the simultaneous activation of TLR4 and GPR40/CD36 signaling by LPS and PA, respectively, cooperatively triggers a strong inflammatory response in hepatocytes. Moreover, PA and LPS cooperatively stimulate CER de novo synthesis, leading to an increase in CER, which further enhances proinflammatory gene expression (Fig. 7). Given that both PA and LPS are elevated in patients with obesity, type 2 diabetes, or metabolic syndrome, the findings from this study revealed a novel mechanism involved in the hepatic inflammation, which contributes to systemic inflammation and CVD in patients with obesity, type 2 diabetes, or metabolic syndrome.

Fig. 7.

Schematic diagram to show the proposed mechanism involved in the upregulation of proinflammatory gene expression in hepatocytes by LPS and PA. The simultaneous activation of TLR4 signaling and GPR40 or CD36 signaling by LPS and PA, respectively, cooperatively triggers a strong inflammatory response in hepatocytes. Moreover, PA and LPS cooperatively stimulate CER de novo synthesis, which increases CER production and further enhances proinflammatory gene expression. CER, ceramide; GPR, G protein-coupled receptor; PA, palmitic acid; TLR, toll-like receptor.

GRANTS

This work was supported by Merit Review Grant no. BX000855 from the Biomedical Laboratory Research and Development Program of the Department of Veterans Affairs and NIH grant no. DE-016353 (to Y. Huang). The work on sphingolipid analysis was supported in part by the Lipidomics Shared Resource, Hollings Cancer Center, Medical University of South Carolina (grant no. P30 CA138313), the Lipidomics Core in the SC Lipidomics and Pathobiology Centers of Biomedical Research Excellence (P20 RR017677), and the National Center for Research Resources and the Office of the Director of the NIH through Grant No. C-06-RR-018823.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

Y.H. conceived and designed research; Y.L., Z.L., and J.H.R. performed experiments; Y.L., Z.L., J.H.R., M.F.L.-V., and Y.H. analyzed data; M.F.L.-V., T.J.L., and Y.H. interpreted results of experiments; Y.L., Z.L., and J.H.R. prepared figures; M.F.L.-V., T.J.L., and Y.H. drafted manuscript; M.F.L.-V. and Y.H. edited and revised manuscript; Y.L., Z.L., J.H.R., M.F.L.-V., T.J.L., and Y.H. approved final version of manuscript.