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. 2010 Jun 23;24(8):1626–1636. doi: 10.1210/me.2010-0117

Activation of the Farnesoid X Receptor Provides Protection against Acetaminophen-Induced Hepatic Toxicity

Florence Ying Lee 1,a, Thomas Quad de Aguiar Vallim 1,a, Hansook Kim Chong 1,a, Yanqiao Zhang 1, Yaping Liu 1, Stacey A Jones 1, Timothy F Osborne 1, Peter A Edwards 1
PMCID: PMC2940469  PMID: 20573685

Abstract

The nuclear receptor, farnesoid X receptor (FXR, NR1H4), is known to regulate cholesterol, bile acid, lipoprotein, and glucose metabolism. In the current study, we provide evidence to support a role for FXR in hepatoprotection from acetaminophen (APAP)-induced toxicity. Pharmacological activation of FXR induces the expression of several genes involved in phase II and phase III xenobiotic metabolism in wild-type, but not Fxr−/− mice. We used chromatin immunoprecipitation-based genome-wide response element analyses coupled with luciferase reporter assays to identify functional FXR response elements within promoters, introns, or intragenic regions of these genes. Consistent with the observed transcriptional changes, FXR gene dosage is positively correlated with the degree of protection from APAP-induced hepatotoxicity in vivo. Further, we demonstrate that pretreatment of wild-type mice with an FXR-specific agonist provides significant protection from APAP-induced hepatotoxicity. Based on these findings, we propose that FXR plays a role in hepatic xenobiotic metabolism and, when activated, provides hepatoprotection against toxins such as APAP.

FXR activation provides hepatoprotection against acetaminophen-induced toxicity.

Acetaminophen (APAP, also known as N-acetyl-p-aminophenol and paracetamol) is a widely used, over-the-counter antipyretic and analgesic. Nonetheless, intake of APAP over the recommended dose of 4 g/d can lead to severe liver damage (1). According to the U.S. Acute Liver Failure Study Group, APAP overdose is the leading cause of acute liver failures in the United States and most of Europe (2,3). Of patients suffering from acute liver failure caused by APAP overdose, 48% overdosed unintentionally without suicidal intent (2).

When taken at the recommended dose, APAP is metabolized in the liver mainly by UDP-glucuronosyltransferases (UGTs) and sulfotransferases (SULTs), with only a small amount being converted by hepatic cytochrome P450 (CYP) enzymes (CYP1A2, CYP2E1 and CYP3A4) into the cytotoxic and electrophilic reactive metabolite N-acetyl-p-quinoneimine (NAPQI). The small amounts of NAPQI generated are normally rapidly detoxified by glutathione (GSH) conjugation and cleared from the liver (4). However, excessive intake of APAP saturates both the glucuronidation and the sulfonation pathways, resulting in the formation of large amounts of the toxic NAPQI metabolite (4). Although the exact mechanism by which NAPQI causes liver damage is unclear, depletion of hepatic GSH during NAPQI elimination is believed to be a critical event that ultimately leads to centrilobular necrosis of hepatocytes and acute liver failure (5).

The nuclear receptor, farnesoid X receptor (FXR, NR1H4), is a bile acid-activated transcription factor that is expressed at high levels in the liver, intestine, kidney, and adrenal glands (6,7,8,9). FXR heterodimerizes with the retinoid X receptor (RXR) and binds to FXR response elements (FXREs) and, in the presence of specific agonists, activates transcription of target genes (10). FXR plays important roles in bile acid, cholesterol, lipoprotein, and glucose metabolism (reviewed in Refs. 10 and 11). It has also been shown to be important in maintaining an intact barrier function in the intestine (12), in preventing gall stone formation in susceptible mice and humans (13,14), and in attenuating cholestasis (15). These latter functions can be attributed to the regulation of a number of genes that affect bile acid detoxification and drug metabolism by FXR. These genes include phase I oxidation enzymes (e.g. CYP3A4) (16), phase II conjugation enzymes (e.g. UGT2B4 and SULT2A1) (17,18), and phase III transporters (e.g. human multidrug resistance-related protein 2, ABCB4) (19). Together, these studies suggest a possible function for FXR in xenobiotic metabolism.

In the present report, we explored the possible link between FXR and xenobiotic metabolism. To identify which drug metabolism enzymes might be regulated by FXR, we used three complementary approaches that involved infecting primary mouse hepatocytes with adenovirus expressing either 1) a constitutively active form of FXR (FXR-VP16), or 2) native FXR, or 3) treating wild-type and Fxr−/− mice with a specific FXR agonist. These approaches identified a number of phase II and phase III xenobiotic metabolism genes that were induced by FXR activation. To confirm that these genes were bona fide FXR-target genes, we used chromatin immunoprecipitation-based (ChIP) genome-wide response element analyses (e.g. ChIP-on-Chip and ChIP-Seq) to identify FXREs. Based on the established role of some of these genes in the metabolism of APAP, we then investigated the effect of loss of function and gain of function of FXR on APAP-induced hepatotoxicity. The data presented herein demonstrate that activation of FXR protects mice from toxic levels of APAP and that loss of FXR is associated with increased susceptibility to APAP toxicity.

Results

Activation of FXR induces expression of multiple phase II and phase III mRNAs

To identify hepatic genes that are induced by FXR, we infected mouse primary hepatocytes with adenovirus encoding either the transactivation domain of herpes virus viral protein (VP16), or individual human FXR isoforms (hFXRα1-α4) fused to VP16. The infected cells were also treated with 1 μm GlaxoSmithKline compound no. 4064 (GW4064), an FXR-specific synthetic agonist, for an additional 24 h to ensure high levels of target gene induction. Through gene-expression profiling, we discovered that FXR activation induced several genes involved in phase II (conjugation) and phase III (elimination) of xenobiotic metabolism. These included glutathione S-transferases (GSTs) (Gstα3, Gstα4, Gstμ1, Gstμ3), GSH metabolism-related genes (Gclm, Gpx1), sulfotransferases (Sult1a1 and Sult1a2), glucuronosyltransferase (Ugt1a1), (phase II) and transmembrane transporters (Abcb4 and Abcb11/Bsep) (phase III).

To validate that these observed transcriptional changes are physiological, we confirmed these microarray predictions in two additional models. First, we infected primary mouse hepatocytes from wild-type mice with adenovirus expressing mouse FXRα2 or green fluorescent protein (GFP). Adenovirus-driven expression of FXRα2 is necessary because endogenous FXR mRNA and protein levels decline more than 90% compared with the intact liver within 1 d of isolation of these primary hepatocytes (data not shown). After infection (24 h), the cells were treated with vehicle or GW4064 for an additional 24 h to ensure maximal target gene induction. Real-time quantitative PCR (RT-qPCR) analyses confirmed that mRNA levels of Gclm, Gstα3, Gstα4, Gpx1, Ugt1α1, Sult1a1, Abcb4, and Bsep were all significantly induced in cells that overexpressed FXR and had been treated with GW4604 (Fig. 1 and data not shown).

Figure 1.

Figure 1

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Activation of FXR leads to induction of phase II–III genes. Hepatocytes from wild-type mice were infected with either Ad-Fxrα2 (mouse) or Ad-Gfp for 24 h and treated with either vehicle or GW4064 for an additional 24 h. Total RNA was isolated, and gene expression was measured by RT-qPCR. These data are representative of three independent experiments. Statistics were analyzed using a one-way ANOVA and Dunnett’s post hoc test to compare treatments to GFP vehicle-treated samples. (n = 6; ***, P ≤ 0.001; **, P ≤ 0.01; *, P ≤ 0.05).

In a second model, we gavaged wild-type and Fxr−/− mice for 5 d with either vehicle or GW4064 and analyzed hepatic gene expression by RT-qPCR. Indeed most of the genes identified (see Fig. 1) were significantly induced after treatment of wild-type, but not Fxr−/− mice with GW4064 (Fig. 2 and data not shown). To ensure that the nuclear receptors pregnane X-receptor (PXR) and constitutive androstane receptor (CAR) were not activated under these conditions, we also measured mRNA levels for some of their target genes; the data indicated that a number of mRNAs including Cyp1a2 and Cyp2e1 were unchanged, consistent with a specific effect of GW4064 on FXR (data not shown). Bsep mRNA, a well-characterized FXR-target gene, served as a positive control in both models, although induction was more pronounced in the primary cells infected with FXR-expressing adenovirus. The low induction of FXR-target genes in uninfected cells is likely due, at least in part, to a significant decline in endogenous FXR levels in primary hepatocytes (data not shown). Together, these data demonstrate that activation of FXR induces the hepatic expression of a number of phase II and III genes known to be involved in metabolism of xenobiotics.

Figure 2.

Figure 2

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Pharmacological activation of FXR in vivo induces the expression of several phase II–III genes in wild-type (WT) but not Fxr−/− (KO) mice. Fxr+/+ or Fxr−/− mice were treated with either vehicle or GW4064 for 5 d. Total hepatic RNA was isolated, and gene expression was measured by RT-qPCR. Statistics were analyzed using a one-way ANOVA and Dunnett’s post hoc test to compare treatments with vehicle-treated Fxr+/+ animals (n = 5–7 mice per group; ***, P ≤ 0.0001; **, P ≤ 0.01; *, P ≤ 0.05).

Identification of a functional FXRE in the proximal promoter Gstα4 using ChIP-on-chip and luciferase reporter assays

To determine which of these phase II and III genes are direct targets of FXR we initially used ChIP-on-chip to identify potential FXREs that lie within 2.5 kb (2 kb upstream and 0.5 kb downstream) of the transcriptional start site of 25,000 mouse genes. Analysis of the data indicated that only Gstα4 of the phase II and III genes shown in Figs. 1 and 2 contained putative FXREs in the proximal promoter (data not shown). After narrowing down the region of the Gstα4 promoter, we used Nubiscan to identify putative FXREs. This approach identified two potential inverted repeats separated by one nucleotide (IR-1) that we termed the distal (dIR-1) and proximal (pIR-1) elements, respectively (Fig. 3).

Figure 3.

Figure 3

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FXR regulates the Gstα4 promoter through the proximal IR-1 (pIR-1) element. McA-RH7777 cells were transfected with luciferase reporter constructs under the control of wild-type (WT) or mutant (Mut) Gstα4 promoter, Shp, or empty promoters together with a plasmid encoding FXR and treated with either vehicle (DMSO) or GW4064 for 24 h. Cells were harvested and luciferase (Luc) activity was determined and corrected for β-galactosidase expression. These data are representative of at least three independent experiments, and each condition had six replicates. The pIR-1 is located at +29 to +41 bp relative to the transcriptional start site. Statistical analysis was carried out using a one-way ANOVA and Dunnett’s post hoc test to compare values with vehicle-treated controls (***, P ≤ 0.0001).

Based on these analyses, we generated a series of luciferase reporter genes under the control of the Gstα4 promoter containing wild-type or mutant IR-1 sequences (Fig. 3). These reporter genes were transiently transfected into the rat liver cell line McA-RH7777 in the absence or presence of a plasmid encoding FXR before treatment of the cells with GW4064 for 24 h. Importantly, McA-RH 7777 cells contain very low levels of endogenous FXR even though they were originally derived from rat liver (data not shown).

As shown in Fig. 3, the luciferase reporter gene driven by wild-type Gstα4 promoter was significantly induced in cells cotransfected with FXR and treated with GW4064 (Fig. 3, WT). A similar increase in luciferase activity was observed when the Gstα4 promoter contained a mutated distal IR-1 element (Fig. 3, Mut 1). This induction was abolished when the promoter contained mutations in either the proximal IR-1 or both IR-1 elements (Fig. 3, Mut 2 or Mut 1+2). A small heterodimer partner (Shp) promoter-luciferase reporter served as a positive control and was induced to a similar extent under these same conditions in an FXR- and GW4064-dependent manner (Fig. 3, Shp WT). These data identify Gstα4 as a new FXR-target gene with a functional FXRE in the proximal promoter.

Identification of additional FXREs in other phase II and III genes using ChIP-Seq and ChIP

The finding that analysis of data from ChIP-on-chip studies failed to identify FXREs in the proximal promoters of most FXR-regulated genes identified in Figs. 1 and 2 suggested that some or all of these genes might be regulated by FXREs that lie outside the proximal promoter. To identify such distal FXREs we initially performed a ChIP-Seq analysis using chromatin isolated from mouse liver essentially as described elsewhere (20), except we precipitated chromatin with an antibody raised against FXR or a control IgG. The DNA enriched by the antibodies was then used as a template for high-throughput sequence analysis (Solexa/Illumina Genome Analyzer). Analysis of the data identified more than 1600 putative FXREs (51).

Analysis of the ChIP-Seq data identified FXR-binding peaks (FXREs) that were located in the proximal promoters of Shp, that served as a positive control, and Gstα4 (Table 1 and Fig. 4). The latter observation is consistent with the identification of the functional FXRE in the Gstα4 promoter (Fig. 3). Additional analysis of the ChIP-Seq data identified FXREs that were located either in intragenic regions upstream of the transcriptional start site of many of the genes identified in Figs. 1 and 2, or were present within introns of these genes (Table 1). To validate these predictions, we next performed site-specific ChIP using chromatin isolated from the livers of wild-type and Fxr−/− mice and primers that flanked a representative subset of the predicted FXREs shown in Table 1. Analyses of the site-specific ChIP data obtained from livers of wild-type mice demonstrated that FXR was significantly enriched on the putative FXREs associated with Gclm, Gstα3, Gpx1, Abcb4, and Shp (positive control) (Fig. 5; compare WT mice with IgG vs. anti-FXR). In contrast, no enrichment was observed when chromatin from Fxr−/− mice was used (Fig. 5).

Table 1.

Summary of selected FXR-target genes identified by ChIP-Seq

Gene name Chr Peak start Peak stop Peak size (bp) No. of tags Strand Distance to TSS Peak location IR-1 sites (P < 0.001)
Gclm chr3 121909478 121909559 81 14 + 39031 Intergenic GCCAGCACTGGGGCCAATGAACTTTGTTGTTCT
Gstα3 chr1 21294403 21294550 147 10 + 63734 Intergenic CCACAAGAGGAATCCAATGGCCAGGTAAGGACT
Gstα4 chr9 78039608 78039960 352 31 + 164 Exon AACGACCTGCAGGACGGTGAACTTTCGCTCGCA
Gpx1 chr9 108244641 108244912 271 24 + 3035 Intergenic AACTTGCATTAGGTCATAGACCTTCTGAGAAGC
Ugt1a1 chr1 90109352 90109591 239 14 − 819 Exon ATACACACCTGGGATAGGGGCTTTTTCTGAAGG
Abcb4 chr5 8928533 8928876 343 30 + 34813 Intron TCGCAGAGAGAGGTCAATGATCTTTACAGGTTT
Shp chr4 133108683 133109257 574 36 + 621 Promoter GGTACAGCCTGGGTTAATGACCCTGTTTATGCA
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The genomic location for each peak is indicated by the peak start and peak stop. Distance to transcriptional start site (TSS) refers to the distance from the start of each peak to the TSS of the indicated gene. The promoter was defined as 2 kb of 5′-flanking DNA upstream of the TSS. Intergenic region refers to all locations other than the promoter, 5′-untranslated region (UTR), exon, intron, or 3′-UTR of upstream or downstream genes. The IR-1 is underlined. Chr, Chromosome. 

Figure 4.

Figure 4

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Representative snapshots of ChIP-Seq peaks for FXR-target genes. Shp (upper panel) and Gstα4 (lower panel), mapped onto University of California at Santa Cruz genome browser. Shown is their chromosomal location (chr 4, chr 9) according to the July 2007 Mouse Genome Assembly (mm9). Blue and red tags represent positive and negative strands, respectively, identified after DNA sequencing of the chromatin after immunoprecipitation using antibody to FXR or a control IgG. Each bar represents a distinct hit/sequence.

Figure 5.

Figure 5

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ChIP analysis confirms that the FXREs identified from ChIP-Seq bind FXR in vivo. Chromatin from livers of either Fxr+/+ or Fxr−/− mice was isolated and immunoprecipiated with either control IgG or a specific antibody to FXR. Quantification of enrichment was determined by quantitative PCR using specific primers for putative FXREs. Statistics were analyzed using a one-way ANOVA and Dunnett’s post hoc test to compare enrichment compared with wild-type (WT)-IgG samples (n = 5 mice/genotype; ***, P ≤ 0.001; **, P ≤ 0.01). KO, Knockout.

Functional characterization of putative FXREs

To date, almost all genes that are directly activated by FXR:RXR have been shown to contain FXREs within either the proximal promoter or intron 1 or 2, or in very few cases, in distant enhancers that are located greater than 10 kb from the transcriptional start site (11,21). Additionally, genome-wide binding studies for several other nuclear receptors have shown that these sites are distributed throughout the genome (22,23). Indeed, recent studies (24,51) have shown a broad distribution for FXR-binding sites throughout the genome (data not shown).

The findings that many of the FXREs identified in the current study are located from 10–60 kb from the transcriptional start site of a gene make the generation of endogenous promoter-luciferase reporter genes impractical. As an alternative approach, we inserted three copies of the putative FXREs of selective genes identified by ChIP-Seq and ChIP upstream of a minimal TK-luciferase reporter gene (Fig. 6). We considered that these FXREs likely regulated the expression of adjacent (upstream or downstream) genes (Abcb4, Gclm, Gpx1). As shown in Fig. 6, the putative FXREs from Abcb4, Gclm, and Gpx1 all induced luciferase activity in an FXR- and GW4064-dependent manner. The well-characterized FXRE in the Shp promoter served as a positive control (Fig. 6). Based on the data in Figs. 1, 2, 5, and 6, we conclude that these FXREs bind FXR in vivo and are sufficient to activate transcription of adjacent genes.

Figure 6.

Figure 6

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Validation of FXREs in phase II–III xenobiotic metabolism genes using luciferase reporters. McA-RH7777 cells were transfected with luciferase plasmids containing a minimal thymidine kinsase promoter and, where indicated, 3× FXREs from the annotated genes. Cells were cotransfected with plasmid expressing FXR and, where indicated, treated with vehicle or GW4064 for 24 h. Cells were harvested and luciferase activity was determined and corrected for differences in transfection efficiency after normalization to β-galactosidase activity. Experiments were carried out at least three times with six replicates for each condition. Statistical analysis was carried out using a one-way ANOVA and Dunnett’s post hoc test with values compared with vehicle-treated controls (***, P ≤ 0.001).

FXR provides protection from APAP-induced hepatotoxicity

The observation that activated FXR results in a broad induction of phase II and III genes (Figs. 1 and 2) suggested that FXR activation may increase the metabolism of certain xenobiotics and/or endogenous substrates. The finding that Gclm was induced by FXR (Figs. 1, 2, 5, and 6) also suggested that hepatic GSH levels may be increased through FXR-dependent pathways. Gclm functions as a modifier subunit of the glutamate cysteine ligase, the rate-limiting enzyme in GSH biosynthesis (25), and an increase in Gclm was sufficient to increase hepatic GSH biosynthesis (25,26). Interestingly, it has been proposed that APAP-dependent hepatotoxicity is due, at least in part, to depletion of hepatic GSH levels (27,28,29).

Based on these observations, we hypothesized that activation of FXR would provide some protection from the hepatotoxic effects of APAP overdose. To test this hypothesis, wild-type mice were pretreated for 5 d with vehicle or GW4064 before a single challenge of APAP (500 mg/kg). Analysis and quantification of the H&E-stained livers 24 h after APAP treatment show that GW4064 pretreatment significantly reduced the damaged area, from 53% in vehicle-treated mice to 28% (Fig. 7, A and B). Consistent with these findings, serum levels of alanine aminotransferase (ALT), aspartate aminotransferase (AST), and lactate dehydrogenase (LDH) were significantly reduced in the GW4064-treated compared with vehicle-treated mice (Fig. 7C). Interestingly, GW4064 pretreatment also resulted in a small but significant increase in hepatic GSH levels (Fig. 7D). These data clearly demonstrate that FXR activation provides hepatoprotection against a subsequent challenge with toxic levels of APAP.

Figure 7.

Figure 7

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Pretreatment of Fxr+/+ (WT) mice with GW4064 provides hepatoprotection from a subsequent APAP challenge. Fxr+/+ mice (n = 4–6 mice per group) were pretreated with vehicle (Veh) or GW4064 (GW) for 5 d as described in Materials and Methods and subsequently challenged with 500 mg/kg APAP for 24 h. A, Representative pictures of H&E-stained paraffin-embedded liver sections are shown. Original magnification, ×50. B, Quantification of damage induced by APAP. The area of damage was quantified in 16 randomly chosen fields taken from multiple H&E-stained paraffin-embedded liver sections (n = 4 mice/group) using ImageJ software. C, Serum levels of ALT, AST, and LDH were measured in the indicated groups (n = 4–6 mice/group). D, Hepatic GSH levels were measured as described in Materials and Methods with samples taken from Fxr+/+ mice after vehicle or GW4064 treatment (n = 6). Arrows mark damaged areas. All data are reported as mean ± sem; *, P < 0.05; **, P < 0.01; ***, P < 0.0001. WT, Wild type.

Wild-type, heterozygous, and Fxr−/− mice displayed gene dosage-dependent susceptibility to APAP-induced toxicity

Because activation of FXR provided significant hepatoprotection toward APAP-induced toxicity (Fig. 7), we hypothesized that reduced expression of FXR might result in increased hepatic damage in mice after an APAP challenge. As such, we challenged Fxr+/+, Fxr+/− and Fxr−/− mice with a sublethal dose of APAP (300 mg/kg) that has an intermediate effect on hepatic function (data not shown). Analysis of H&E-stained liver sections indicated that Fxr−/− mice sustained more severe and extensive liver damage than either Fxr+/+ or Fxr+/− mice (Fig. 8, A and B). In contrast, analysis of H&E-stained liver sections indicated that damage was similar in wild-type and Fxr heterozygous mice (Fig. 8B).

Figure 8.

Figure 8

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FXR gene dosage positively correlates with resistance toward APAP-induced hepatotoxicity. Fxr wild-type (WT), heterozygous (HET), and knockout (KO) mice (n = 3–7 mice/group) were challenged with 300 mg/kg of APAP for 24 h. A, Representative pictures of H&E-stained paraffin-embedded liver sections are shown. Original magnification, ×50. B, Quantification of damaged liver. For each treatment group, the area of liver damage was quantified in 12 (Fxr+/−) or 16 (Fxr+/+ and Fxr−/−) randomly chosen fields taken from the indicated mice (three to six mice per group) as described in Materials and Methods. C, Serum levels of ALT, AST, and LDH were measured in the indicated groups (n = 3–7). Arrows mark damaged areas. All data are reported as mean ± sem; *, P < 0.05; **, P < 0.01; ***, P < 0.0001.

Analysis of plasma levels of ALT, AST, and LDH provides a more sensitive measure of hepatic damage. As shown in Fig. 8C, all three proteins were significantly increased in the plasma of APAP-treated Fxr+/− as compared with Fxr+/+ mice. However, the highest levels of ALT, AST, and LDH were seen in the plasma of APAP-treated Fxr−/− mice (Fig. 8C). Together the data of Fig. 8 demonstrate that the susceptibility of mice to a challenge with nonlethal levels of APAP is dependent upon the Fxr gene dosage.

Discussion

Using wild-type and Fxr−/− mice, together with a combination of ChIP-on-chip, Chip-Seq, and site-directed ChIP, we have identified novel FXREs in enhancer regions that lie up to 60 kb upstream of transcription start sites of previously unidentified FXR-target genes. These FXR-target genes encode UGTs, GSTs, SULTs, transmembrane transporters, and a modifier of the rate-limiting enzyme of GSH biosynthesis (Figs. 1 and 2). As we have demonstrated here using the APAP-induced hepatotoxicity model, specific activation of FXR by GW4064 in wild-type, but not Fxr−/− mice, provides significant hepatoprotection. These data provide compelling evidence that the FXR-dependent induction of a combination of these genes is important for the regulation and hepatic conjugation and transport of toxic xenobiotics.

The genome-wide FXRE-mapping approach used here proved to be key in the validation of our microarray predictions, because among the novel FXR targets identified in this report, only Gstα4 contained an FXRE within the proximal promoter (Figs. 3 and 4 and Table 1). Furthermore, this ChIP-Seq approach identified a previously unknown FXRE located approximately 34.8-kb upstream of the transcriptional start site of the mouse Abcb4 gene. Although the human ABCB4 gene contains an FXRE in the proximal promoter (16), this element is not conserved in mice. These examples highlight the fact that the cis-acting enhancer elements can be located in distant inter- and intragenic region of a target gene. To prove that these newly identified FXREs associated with Gclm, Gpx-1, and Abcb4 are indeed functional, traditional luciferase reporter genes were constructed and are shown to be induced in response to FXR and GW4064. However, further studies will be required to prove definitively that the distantly located enhancer elements identified in the current study are functionally important to specific gene activation in vivo. The current studies are further supported by a recent study by Amador-Noguez et al. (30), who demonstrated that FXR was a likely regulator of many genes involved in xenobiotic metabolism in the long-lived Little mouse model (Ghrhlit/lit). Although no molecular mechanism was reported in these latter studies, the authors suggested the increased gene expression in xenobiotic metabolism genes in this particular mouse model required FXR and was independent of CAR and PXR (30).

APAP overdose can result in hepatotoxicity and death in both animals and humans (31). Because APAP is metabolized mainly in the liver, we decided to use the APAP-induced hepatotoxicity model to probe the physiological relevance of our newly identified FXR targets in phase II and phase III of hepatic xenobiotic metabolism. Using this model, we have demonstrated in mice that pretreatment with an FXR agonist provides significant protection from a subsequent challenge with toxic levels of APAP (Fig. 7). Additionally, we find that hepatic damage in response to APAP is inversely proportional to the copy number of FXR genes in mice (order of susceptibility is Fxr−/− > Fxr+/− > Fxr+/+) (Fig. 8). These data indicate that target genes of FXR are important in providing hepatoprotection from APAP ingestion. However, due to the fact that multiple phase II and III genes are induced after FXR activation (Figs. 1 and 2), and that APAP metabolism is a multistep process, it is not practical to determine which combination of the FXR-induced genes are critical in protecting the liver from the toxic effects of APAP.

Attempts to determine the relative importance of the individual phase II and III FXR-target genes in providing hepatoprotection from APAP is likely to be very complicated. Interestingly, several studies that used specific gene-knockout mice identified roles for some of these FXR-targets in hepatoprotection from various toxins. For instance, loss of Gstα4 in mice leads to increased susceptibility to paraquat and CCl4 liver toxicity (32,33). Similarly, loss of Gstα3 leads to increased susceptibility to damage by Aflatoxin B1, a major risk factor for hepatocellular carcinoma (34). The role of Gpx-1 ablation on APAP-induced toxicity has also been investigated, although no clear effect was reported (35). A more striking result was observed in Gclm-knockout mice; these mice were shown to be more susceptible to an APAP challenge (36). Perhaps more importantly, transgenic mice overexpressing Gclm are protected from APAP hepatic damage, when compared with wild-type mice (26). The importance of individual FXR-target genes, or combinations of these genes, in providing hepatoprotection from APAP is beyond the scope of the current investigation because it would require studies on multiple knockout mice. Taken together, these latter studies highlight the importance of a number of novel FXR-target genes identified in the current study in providing protection against hepatotoxins.

In addition to FXR, both the PXR and the CAR have been implicated in APAP metabolism (37,38,39,40). PXR and CAR have been deemed xenobiotic sensors because they regulate genes involved in all phases of drug metabolism (38). However, in contrast to the increased susceptibility of Fxr−/− mice to APAP, Car−/− (40) or Rxr−/− (41) mice were shown to be resistant to a challenge with APAP. However, the relationship between PXR and APAP-induced toxicity is less well understood because treatment of Pxr−/− mice with APAP has been reported to result in either increased (39) or decreased (37) hepatotoxicity. Given the difference in APAP susceptibility between Fxr−/−, Pxr−/−, and Car−/− mice, and the finding that activation of FXR does not directly induce genes involved in phase I metabolism of xenobiotics, as PXR and CAR do, we propose that FXR agonists protect mice from APAP-induced hepatotoxicity through a pathway independent of PXR and CAR. However, additional studies with multiple knockout mice would be required to definitively exclude any role for PXR and/or CAR in the hepatoprotection after activation of FXR.

Indeed, FXR has been shown previously to protect mice from another hepatotoxin, α-naphthylisothiocyanate (ANIT) (42). However, ANIT causes hepatic damage through a different mechanism from APAP. ANIT induces cholestasis as a result of the ABC transporter multidrug resistance-related protein 2 excreting a toxic GSH-conjugated ANIT metabolite across the canalicular membrane where it subsequently damages the cholangiocytes lining the bile ducts (43,44). Interestingly, an earlier study demonstrated that pretreatment of rats with the FXR agonist GW4064 provided partial protection from ANIT-induced hepatic necrosis and inflammatory cell infiltration (42). At the time, the hepatoprotective action of FXR was attributed to induction of organic anion transporters BSEP and ABCB4 (42). However, the current study suggests that FXR activation of phase II and III xenobiotic metabolism genes may also play a protective role in this model.

Finally, our study suggests that patients with compromised FXR activity, e.g. those with progressive familial intrahepatic cholestasis type 1 (45), may exhibit increased hepatic damage after ingestion of excess APAP. Whether decreased expression of FXR, as a result of primary or secondary hepatic injury, affects a patient’s ability to metabolize APAP or other xenobiotics is unknown and is clearly an area that warrants further investigation.

Materials and Methods

Materials

GW4064 was kindly provided by Drs. T. Willson and P. Maloney (GlaxoSmithKline, Research Triangle Park, NC) (46,47). APAP, dimethylsulfoxide (DMSO), 5,5′-dithiobis(2-nitrobenzoic acid), and 2-hydroxypropyl-β-cyclodextrin (CD) were purchased from Sigma-Aldrich (St. Louis, MO). Collagenase I was purchased from Worthington Biochemical Corp. (Freehold, NJ). Adenoviruses encoding either VP16 or FXRα1, -α2, -α3, or -α4 isoforms, as VP16 fusions, have been described previously (47). GFP and mouse FXRα2 adenoviruses were made using the AdEasy system according to the manufacturer’s instructions. CodeLink microarrays were manufactured by GE Healthcare Life Sciences (Piscataway, NJ; formerly Amersham).

Animals and treatments

Characterization of Fxr−/− mice on a C57BL/6 background have been previously reported (48). All animal experiments have been approved by and were performed in accordance to guidelines provided by the Animal Care and Research Advisory Board at University of California, Los Angeles. Hepatic mRNA levels were determined after a 5-d treatment of wild-type and Fxr−/− mice by oral gavage with either vehicle (CD) or GW4064 (30 mg/kg, twice daily) dissolved in CD. Where indicated, mice were pretreated with vehicle or GW4064 (30 mg/kg, twice daily) for 4.5 d before a single ip injection of APAP dissolved in basic PBS (pH 11) (300 or 500 mg/kg; as indicated in the specific legend). Tissues were analyzed 24 h later. All experiments were carried out with age- and sex-matched animals.

Histology

Livers were fixed in 10% neutral-buffered formalin and paraffin-embedded. H&E staining was carried out by the Tissue Procurement and Histology Core Laboratory in the Department of Pathology and Laboratory Medicine at UCLA following standard protocols. Liver damage area was quantified using the ImageJ software developed at National Institutes of Health.

Serum hepatic enzyme measurements and hepatic nonprotein thiol measurement

Serum ALT, AST, and LDH levels were measured as described elsewhere (42). GSH is considered the major nonprotein thiol species in the liver. Consequently, we determined hepatic nonprotein thiol as a measure of hepatic GSH level. Liver samples were prepared as described elsewhere (40). Hepatic nonprotein thiol determination was carried out following protocol developed by Pierce Biotechnology (Rockford, IL). Concentration of nonprotein thiols were calculated using a molar extinction coefficient of 14,150 m−1cm−1 for 5,5′-dithiobis(2-nitrobenzoic acid) (49).

Adenoviral infection of primary murine hepatocytes and microarray

Primary murine hepatocytes were isolated and maintained essentially as described (47). Monolayers of primary hepatocytes were maintained for 72 h before they were infected with 10 M.O.I. of adenovirus encoding either the transactivation domain of VP16 alone or individual FXR isoforms as FXR-VP16 fusions. The cells were treated 24 h after infection with either vehicle (DMSO) or GW4064 (1 μm) and RNA isolated after an additional 24 h. cDNA probes were synthesized from these mRNAs and hybridized onto CodeLink Microarrays (GE Healthcare Life Sciences) according to the manufacturer’s recommendation at the BioMedical Genomics Microarray Facility (BIOGEM) at University of California, San Diego. Validation studies were carried out in primary murine hepatocytes infected with either murine FXRα2 or GFP adenovirus for the same period of time and treated with vehicle and GW4064 (1 μm) in the same way as described above.

RT-qPCR analysis

RT-qPCR analyses were performed using the Roche Lightcycler 480 and Roche Lightcycler 480 SYBR Green Master Mix (Roche, Indianapolis, IN). Quantification was carried out using the efficiency corrected method (Roche) using primers specific for each gene (sequences available upon request). Gene expression values were normalized to 36B4 and expressed as fold changes from control. Statistics were performed using one-way ANOVA and Dunnett’s post hoc test, as indicated. Data are reported as mean ± sem.

Cell culture and transient transfection assays

McA-RH7777 cells were purchased from the American Type Culture Collection (ATCC, Manassas, VA)and maintained according to ATCC recommendations. Cells were seeded 24 h before transfection in 48-well dishes. The following day, cells were transfected with reporter constructs as indicated in the figure legends, for 6 h in OPTI-MEM (Invitrogen, Carsbad, CA). Transfection medium was then removed and cells were treated with either vehicle (DMSO) or GW4064 (1 μm) in medium containing 10% charcoal-inactivated serum for a further 24 h before being harvested. Luciferase activity was determined using Luciferase Assay System (Promega Corp., Madison, WI) and normalized to β-gal activity to correct for variations in transfection.

ChIP, ChIP-on-chip, and ChIP-Seq

ChIP assays for livers from 6-wk-old wild-type C57BL/6 male mice were performed as previously described (50) with a minor modification. Chromatin was harvested and subjected to an immunoselection process using antibodies against FXR (sc-13063; Santa Cruz Biotechnology, Inc., Santa Cruz, CA) or mouse IgG (Sigma, St. Louis, MO) as a control. After isolating the ChIP-enriched DNA, gene-specific enrichment for the known FXRE in the promoter of the target gene Shp, in the FXR chromatin relative to IgG control chromatin, was confirmed as described by Chong et al. (51).

DNA eluted after immunoenrichment was analyzed by high-throughput DNA sequencing using technology for the Solexa/Illumina genome analyzer. Library preparation and sequencing were performed by Ambry Genetics (Aliso Viejo, CA). The short-sequencing reads produced from the genome analyzer were mapped to a reference mouse genome using ELAND, allowing one mismatch. ChIP-seq data were further analyzed as previously described (20,51).

For ChIP-on-chip array hybridizations, mice were administered either vehicle or 50 mg/kg GW4064 dissolved in vehicle by oral gavage for 5 d. After ChIP, the FXR, IgG control, and input DNA for both control and GW4064-treated groups were prepared for hybridization to the 2.5-kb mouse promoter array (NimbleGen/Roche) using a random PCR amplification method according to the company’s protocol. The hybridization data were analyzed by using NimbleScan 2.3 and SignalMap software from NimbleGen/Roche. There were no statistically significant differences for peaks from either set of chromatin.

Footnotes

This work was supported by National Institutes of Health Grants HL30568 and HL68445 (to P.A.E.), a grant from the Laubisch Fund (to P.A.E.), and U.S. Public Health Service National Research Service Award GM07185 (to F.Y.L.), a Beginning Grant-in-Aid 0565173Y from American Heart Association (to Y.Z.), Grant DK70121 (to T.F.O.), and National Institutes of Health Training Grant T1507443 (to H.K.C.).

Present address for F.Y.L.: University of California, San Francisco, Department of Cellular and Molecular Pharmacology 1550 4th Street, Room 281, Box 2611 San Francisco, California 94143-2611.

Present address for Y.Z.: Northeastern Ohio Universities College of Medicine Integrative Medical Sciences, 4209 State Route 44, P.O. Box 95, Rootstown, Ohio 44272.

Disclosure Summary: The authors have nothing to disclose.

First Published Online June 23, 2010

Abbreviations: ALT, Alanine aminotransferase; ANIT, α-naphthylisothiocyanate; APAP, acetaminophen; AST, aspartate aminotransferase; BSEP, bile salt export pump; CAR, constitutive androstane receptor; ChIP, chromatin immunoprecipitation; CYP, cytochrome P450; FXR, farnesoid X receptor; FXRE, FXR response element; GCLM, glutamate cysteine ligase; GFP, green fluorescent protein; GSH, glutathione; GST, glutathione S-transferase; GW4064, GlaxoSmithKline compound no. 4064, synthetic FXR ligand; H&E, hematoxylin and eosin; LDH, lactate dehydrogenase; NAPQI, N-acetyl-p-quinoneimine; PXR, pregnane X receptor; RT-qPCR, real-time quantitative PCR; RXR, retinoid X receptor; SHP, small heterodimer partner; SULT, sulfotransferase family; UGT, UDP-glucuronosyltransferase; VP16, herpes virus viral protein 16.

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