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BINDING SITE WITHIN THE PROXIMAL PROMOTER OF PXR
School of Biomedical and Molecular Sciences, University of Surrey, Guildford, Surrey, United Kingdom
(Received June 15, 2005; accepted October 14, 2005)
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Abstract |
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, and chicken ovalbumin upstream promoter transcription factor, commensurate with the high expression of PXR in liver. Furthermore, we identified putative binding sites for a number of ligand-activated transcription factors, suggesting that these factors may regulate PXR gene expression. Further analysis of this regulatory region has shown that transcriptional activation of PXR by peroxisome proliferator-activated receptor (PPAR) is via a binding site located approximately 1.3 kb upstream of the putative transcription start site, with ablation of this site preventing PPAR-mediated activation of PXR gene expression. We present a model of how regulation of PXR gene expression by ligand-activated transcription factors may play a central role in the body's response to xenobiotic exposure.
)], aiming to maintain the status quo and ensuring normal/homeostatic physiology. Proteins involved in this process include active transport pumps (e.g., MDR1 and OATP2) to regulate cellular influx/efflux of chemicals and phase I (e.g., cytochromes P450) and phase II (e.g., glutathione S-transferase) metabolic enzymes, which catalyze chemical alterations to increase rates of excretion (Plant, 2004). To respond effectively to fluxome alterations, a feedback mechanism exists whereby levels of drug transporters and metabolic enzymes are regulated by a superfamily of ligand-activated transcription factors (LATFs). These LATFs generally possess large ligand-binding domains and show promiscuity in their activation profile (Watkins et al., 2001). Due to the overlapping nature of these activation profiles and the complex chemical pool within the body at any one time, it is perhaps not surprising that an interaction network exists between these LATFs, with the sum of the interactions/activations elicited by a chemical determining the exact profile of transporters and/or drug-metabolizing enzymes activated to respond.
The pregnane X-receptor (PXR; alternate names SXR, PAR, or NR1I2) is an LATF that has emerged as a transcriptional activator of at least 40 genes, including several biologically important drug transporters and metabolic enzymes, including CYP3A4 (El-Sankary et al., 2000), CYP2B6 (Goodwin et al., 2001), GST-A2 (Falkner et al., 2001), and OATP2 and MDR1a (Maglich et al., 2002). This allows PXR to act as a xenobiotic/metabolite sensor, responding to alterations in the fluxome. Furthermore, evidence is beginning to accrue that PXR may function as a master xenobiotic/metabolite sensor, integrating inputs from other LATFs into the final output it places on the body. This integration is achieved through the interactions of these LATFs, with PXR at both the transcriptomic (Pascussi et al., 2000a) and proteomic (Ourlin et al., 2003) levels. To understand how the body responds to alterations in the fluxome, it is therefore imperative that we delineate this highly refined interaction network.
Whereas many studies have been undertaken on the transcriptional regulation of PXR target genes [e.g., CYP3A4 (El-Sankary et al., 2001, 2002)], little research has been directed at understanding the transcriptional regulation of PXR itself. Several chemicals have been shown to regulate PXR mRNA levels, both in primary human hepatocytes [dexamethasone (Pascussi et al., 2000a); lithocholic acid (Kliewer and Willson, 2002)] and rat liver [clofibrate, perfluorodecanoic acid, isoniazid, and troleandomycin (Zhang et al., 1999)]. As these chemicals are known ligands for other metabolite sensors (e.g., GR, farsenoid X receptor, liver X receptor, PPAR), this is consistent with PXR acting as a master metabolite sensor, coordinating body responses to changes in the fluxome. In the case of glucocorticoids such as dexamethasone, the reason behind such an interaction is clear; glucocorticoids are ligands for both GR and PXR (Ekins and Erickson, 2002) and are metabolized by PXR target genes [e.g., CYP3A4 (Gibson et al., 2002)]. Hence, increased levels of PXR will ultimately lead to increased metabolism of the stimulating glucocorticoid. Activation of PXR expression by bile acids such as lithocholic acids may also be explained, since they are ligands for the LATFs farsenoid X receptor and liver X receptor, which undergo protein/protein interactions with PXR (Edwards et al., 2002). By comparison, the activation of PXR gene expression by clofibrate and perfluorodecanoic acid, ligands for the fatty acid sensor PPAR, is more difficult to explain. These chemicals do not seem to be PXR ligands, nor does their metabolism seem to be dependent upon PXR target genes, although some more potent PPAR agonists do seem to be able to activate PXR. Such data are suggestive that the role of PXR as a master metabolite sensor extends beyond what is currently understood; delineation of the interaction network of metabolic-sensing LATFs will thus greatly increase our knowledge on this key biological molecule. Currently, several assays have been developed to measure PXR activation (Kawana et al., 2003; Vignati et al., 2004), and research into PXR target genes/pathways is extensive (Handschin and Meyer, 2003; Bhalla et al., 2004; Uppal et al., 2005). However, the majority of research on transcriptional activation of PXR has been descriptive (Zhang et al., 1999; Pascussi et al., 2000a,b; Kliewer and Willson, 2002) rather than mechanistic, and the work presented herein aims to provide novel mechanistic insights into the transcriptional regulation of the master xenobiotic/metabolite sensor PXR.
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Materials and Methods |
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). Putative binding sites for transcription factors were identified using a stringent search setting, with the matrix similarity at >0.75 and core similarity at >0.85 to minimize the identification of false positives. The matrix represents the DNA binding profile for individual or groups of transcription factors, with the matrix similarity being the quality of a match between the matrix and the input sequence. Core similarity represent the quality of a match between the core sequence of a matrix (the four most conserved position within a matrix) and the input sequence (Quandt et al., 1995). Chemicals. FuGENE 6 transfection reagent was purchased from Roche Diagnostics, Lewes, UK. Unless otherwise stated, all other chemicals were of molecular biology grade and obtained from Sigma Chemical Co. (Poole, Dorset, UK).
Plasmids. Expression plasmids for LATFs were kindly provided as follows: PXR, Dr. S. Kliewer (University of Texas, Dallas, TX); CAR, Prof. M. Negishi (National Institute of Environmental Health Sciences, Research Triangle Park, NC); VDR, Dr. R. Kim (Vanderbilt University, Nashville, TN); PPAR and GR, Dr. J. Tugwood (AstraZeneca, Macclesfield, Cheshire, UK); and RXR, Prof. P. Chambon (INSERM, Strasbourg, France).
Cell Culture. All cell culture medium and supplements were purchased from Invitrogen (Paisley, UK).
Primary human hepatocytes were obtained from the UK Human Tissue Bank (Leicester, UK) and cultured in Williams' E Medium [containing 2 mM L-glutamine, 10% heat-inactivated bovine serum, penicillin/streptomycin (50 U/ml and 50 µg/ml, respectively), and insulin (1 mg/ml)] in collagen-coated 24-well plates (BD Biosciences, San Jose, CA) at 3 x 105 cells/well. Cells were allowed to attach for 24 h and then exposed, in triplicate, for 48 h to 100 µM clofibrate or Wy-14,643.
The Huh7 human hepatocellular carcinoma cell line (Nakabayashi et al., 1982) was a kind gift from Dr. Steve Hood (GlaxoSmithKline, Ware, UK). All cells were routinely cultured in 75-cm2 vented tissue culture flasks (Nunc International, Leicestershire, UK), using minimal essential medium with Earle's salts supplemented with 1% nonessential amino acids, 2 mM L-glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin, and 10% fetal bovine serum. To maintain phenotypic consistency, Huh7 cells were only used for 3 weeks (approximately five passages) following recovery from liquid nitrogen.
Transfection. Huh7 cells were seeded into 96-well plates (Nunc International) at a concentration of 10,000 cells/well and incubated at 37°C for 24 h in a humidified container for attachment. FuGENE 6-mediated DNA cotransfections, using 75 ng/well PXR reporter gene construct, were performed as described previously (Goodwin et al., 1999), using serum-free medium for the 6-h transfection period; this was then replaced with fresh, complete medium for the remaining culture period. For cotransfection experiments, 25 ng/well of the expression plasmid for each ligand-activated transcription factor, or the empty expression plasmid as a control, was included in the transfection mix. Transfections were allowed to proceed for 48 h, and secretory alkaline phosphatase (SEAP) activity was measured.
Alkaline Phosphatase Activity Assay and Data Analysis. Aliquots of cell culture medium (25 µl/well) were transferred into 96-well optiplates (Canberra Harwell Ltd., Didcot, UK). Endogenous alkaline phosphatase activity was deactivated by heat treatment of the medium at 65°C for 30 min. SEAP activity was then assayed using the AURORA system (ICN, Thame, Oxfordshire, UK), according to the manufacturer's protocol. Chemiluminescent output was measured using a LumiCount automated plate reader (Wolf Laboratories, Pocklington, UK). SEAP activity following 48-h culture was calculated for both reporter constructs and blank, control, plasmid, and a fold induction relative to vehicle control calculated.
Quantitative PCR. Total RNA was extracted using the RNeasy Mini Kit (QIAGEN Ltd., Crawley, UK), quantified using RiboGreen (Invitrogen), and the 1 µg of total RNA was treated with RNase-free DNase I (Promega, Southampton, UK) at 37°C for 30 min. Following heat inactivation, cDNA was produced using Superscript II (Invitrogen) according to the manufacturer's protocol.
Quantitative PCR reactions were set up using FAM reporter dye/TAMRA quencher dye-labeled probes in conjunction with appropriate primer sets as given below (MWG Biotech, Milton Keynes, UK). Forward PXR primer, 5'-CGAGCTCCGCAGCATCA-3'; reverse PXR primer, 5'-TGTATGTCCTGGATGCGCA-3'; and PXR probe, 5'-FAM-TGCTCAGCACACCCAGCGGCT-TAMRA-3'. qPCR Rox Mastermix (Abgene, Epsom, UK) was used, and 25-µl reactions were set up according to the manufacturer's instructions; quantitative PCR results were quantified using the ABI proprietary software against a standard curve generated from human genomic DNA (Promega).
Site-Directed Mutagenesis. Specific mutations for disruption of the putative PPAR binding site were created using a PCR-based methodology. Primers were designed to amplify the fragments upstream and downstream of the putative PPAR binding site (PPRE), with alterations in the primer sequence producing the desired mutation (see below). Engineering of an AgeI restriction site into these mutant sequences allowed the two fragments to then be joined via ligation to form the full PXR proximal promoter sequence (shown in bold below). The second primer for each amplicon was derived from the SEAP plasmid, thus allowing easy cloning of the mutated construct in to the reporter gene system. All mutation constructs were sequenced on both strands to ensure that the desired mutations had been incorporated. Mutant A upstream fragment primer, 5'-CCATAGAGACCGGTCCTTTTTCCA-3'; mutant B upstream fragment primer, 5'-CAGCCATACCGGTCTGTCCTTTTT -3'; downstream fragment primer, 5'-AGGACAGACCGGTATGGCTGTGG-3'; SEAP upstream fragment primer, 5'-ATAAGGGATTTTGCCGATTTCGG-3'; and SEAP downstream fragment primer, 5'-CACAGGTAGGCCGTGGCTGTG-3'.
Preparation of Nuclear Extracts. Nuclear protein extracts were isolated according to the protocol of Dignam et al. (1983). Briefly, Huh7 cells were grown to approximately 90% confluence and then collected by trypsinization. Cells were pelleted by centrifugation (1300g for 5 min) and washed twice with phosphate-buffered saline. After the second wash, cells were resuspended in 5x packed cell volume of ice-cold phosphate-buffered saline. Cells were pelleted, resuspended in 2x packed cell volume of buffer A (10 mM Hepes-KOH, pH 7.9, 1.5 mM MgCl2, 10 mM KCl, and 0.5 mM DTT), and allowed to swell on ice for 10 min before disruption using a Dounce homogenizer (VWR International, Lutterworth, UK). Nuclei were pelleted (2000g for 15 min) and resuspended in 0.5x packed nuclear volume (homogenate volume-supernatant volume) of buffer C (25% glycerol, 20 mM Hepes-KOH, pH 7.9, 1.5 mM MgCl2, 0.2 mM EDTA, 20 mM NaCl, 0.5 mM DTT, and 0.5 mM PMSF); 0.5x packed nuclear volume of high-salt buffer (buffer C containing 1.2 M NaCl) was then added dropwise with swirling, and the suspension was homogenized with a Dounce homogenizer. The resulting homogenate was centrifuged at 16,000g for 30 min, and supernatant (nuclear protein) aliquots were stored at –80°C. Protein concentration was determined by a modification of the method of Stoscheck (1990) and integrity assessed by SDS-polyacrylamide gel electrophoresis. Each aliquot was taken through only three freeze/thaw cycles to maintain protein integrity.
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EMSA binding reactions were carried out at room temperature (22°C) and consisted of 10 µl of 2x binding buffer (40 mM Tris-HCl, pH 7.9, 100 mM NaCl, 20% glycerol, and 0.2 mM DTT), 1 µg of poly-dI:dC, and 1 to 10 µl of protein extracts (representing 4 to 40 µg of nuclear proteins) in a total volume of 20 µl. After 10-min incubation, 2 µl of oligomer probe was added, and the reaction was further incubated for 30 min, followed by separation by polyacrylamide electrophoresis. Competition experiments used between a 1x and 100x excess of unlabeled putative PPRE probe in addition to the labeled probe. In vitro-translated PPAR was produced using the PPAR expression plasmid coupled to the TNT T7 Quick system (Promega) according to the manufacturer's protocol.
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Results |
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). Figure 1 details those sites identified with a high match and hence represent good possibilities of representing functional sites. A TATAA box was identified 30 bp from the putative transcription site, consistent with previous studies that have indicated that approximately 25 to 30 bp upstream of the transcription start site is the optimal location for a TATAA box (Latchman, 2001). In addition, we have identified putative DNA/protein interaction sites for both auxiliary/tissue-specific transcription factors such as HNFs, C/EBP, and Sp1, and a large number of ligand activated transcription factors, including VDR, GR, PRE, and PPAR. This suggests that the PXR proximal promoter is a complex promoter with binding sites for many regulatory transcription factors, consistent with the paradigm of PXR as a master xenobiotic/metabolite sensor capable of responding to many different stimuli. Whereas identification of putative protein/DNA interaction sites by in silico data mining provides the basis for an investigative hypothesis, it does not form the basis for proven molecular mechanisms of action. Hence, we next examined the regulation of PXR using in vitro methodology. The 2.2-kb region of PXR proximal promoter was cloned from human genomic DNA and confirmed by sequencing, and a reporter gene construct was prepared. From this, nine daughter constructs were made (Fig. 2A). Transfection of these constructs into Huh7 cells was used to determine the input of each region into the basal expression of PXR. Figure 2B shows the basal reporter expression from each of these constructs, and from these, both positive and negative regions of regulation can be inferred. It can be seen that the 2.2 kb proximal to the PXR transcription start site has a number of both positive and negative regulatory regions, consistent with the observation of this region as a complex regulatory region.
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, and PPAR overexpression had a positive effect on PXR reporter expression when the whole 2.2-kb construct was used, suggesting that ligands for these receptors may act to increase PXR expression and hence activation of PXR target genes. However, it is interesting to note that overexpression of PXR itself, or CAR, significantly decreases basal expression of the PXR reporter construct, and indeed this suppression occurs across most of the tested deletion constructs (Fig. 3, A and B). This would suggest that PXR may act in a negative-feedback mode and prevent overexpression of both PXR and its target genes.
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ligands are also ligands for PXR itself (El-Sankary et al., 2001) and are metabolized in the body by protein products of PXR target genes (Gibson et al., 2002). By contrast, the large transcriptional activation of PXR gene expression by overexpression of the ligand-activated transcription factor PPAR is less readily understandable, since PPAR ligands have not previously been shown to be PXR ligands or to be metabolized by PXR target genes. In all cotransfection experiments carried out herein, it should be noted that no exogenous ligands are added, with activation of receptors via the endogenous ligand pool being presumed. Whereas this may, to some extent, limit the extent of the responses observed, it should not alter their effect/presence. Indeed, previous evidence has shown that many of the ligand-activated and liver-enriched transcription factors are expressed in Huh7, albeit at reduced levels compared with in vivo, supportive of the presence of all the necessary factors within these cells for functioning of these transcription factors (Phillips et al., 2005).
To further investigate this phenomenon, we examined which region of the PXR proximal promoter was involved in mediating activation by PPAR. Figure 4A shows that the region –1514 to –1321 bp (relative to the putative transcription start site) bounded by the –1.5 kb and –1.3-kb daughter constructs was significantly activated by overexpression of PPAR; examination of other fragments showed no significant induction, thus localizing the PPAR-mediated activation of PXR to this region (Fig. 4B). In addition, we examined the role of RXR, the heterodimerization partner of PPAR, in this response (Fig. 4B). Overexpression of both PPAR and RXR in the system resulted in an increased degree of activation of the 1.5-kb fragment (data not shown). Thus we have experimentally localized the PPAR-mediated activation of PXR to the –1514 to –1321 bp region, a localization that is consistent with the identification of a putative PPRE within this region (Fig. 2A).
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provides strong evidence that it is indeed PPAR that is the protein that interacts with this site in vitro. Finally, mutagenesis of the putative PPRE resulted in disrupted protein/DNA interactions (Fig. 4D) and reduced activation of the 1.5-kb reporter gene construct to PPAR overexpression (Fig. 4E). To examine whether the observed in vitro effects were likely to translate into an in vivo effect, we next examined the level of PXR transcripts in primary human hepatocytes exposed to the PPAR ligands clofibrate and Wy-14,643. As seen in Fig. 4F, clofibrate elicited a statistically significant increase in PXR transcript level, 189% of control levels, whereas no significant change was observed with Wy-14,643 (160% of control levels). For comparison, these changes are significantly less that that caused by dexamethasone (316% of control), a chemical previously shown to increase human PXR gene expression (Pascussi et al., 2000a), although the use of only a single dose cannot rule out the possibility of larger effects with increased doses. We have thus demonstrated that PPAR mediates its activation of PXR gene expression via a PPRE located –1346 bp upstream of the putative transcription start site and that this activation is mirrored in primary human hepatocytes.
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Discussion |
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). The ligand-activated transcription factors represent one of the best examples of such an interaction network, with the 48 members of this family found in humans (Zhang et al., 2004) interacting both at the transcriptional (Pascussi et al., 2000a) and protein (Edwards et al., 2002) levels. The outcomes of such interactions are 2-fold; first, it is possible for a single stimulus to elicit tissue-specific responses, activating a gene set to produce the most effective response in each tissue (Hartley et al., 2004). Second, that redundancy in the response system allows for a metabolic safety net, ensuring correct cellular homeostasis is maintained even under extreme external stimulation (Xie et al., 2000b). The PXR seems to be a prime candidate for this last role, with increasing evidence demonstrating that it can be stimulated (at both the gene and protein levels) by a wide range of both xenobiotic and endogenous chemicals (Lehmann et al., 1998; Kliewer and Willson, 2002). Based upon the findings described herein, we propose a model by which PXR may be regulated both by itself and other LATFs (Fig. 5). Xenobiotic exposure results in the activation of LATFs, which in turn may stimulate PXR gene expression. If the stimulating chemical is a ligand for PXR, then PXR is activated and two endpoints are seen: feedback inhibition of PXR gene expression and activation of PXR target gene expression. The latter would result in increased metabolism of the stimulating chemical and reducing its level. However, if the stimulating chemical is not a ligand for PXR, then we hypothesize that the increased expression of PXR would only result in increased activity of PXR if sufficient levels of endogenous PXR ligands were present within the cell. Activation of PXR target genes in this latter case would presumably not result in metabolism of the stimulating chemical but may have consequences for coexposed chemicals or endogenous metabolism.
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ligands such as clofibrate increase PXR transcript levels in rat hepatocytes (Zhang et al., 1999), despite the fact that molecular modeling suggests that clofibrate is not a ligand for PXR, with a theoretical EC50 in excess of 100 µM (D. Lewis, University of Surrey, Guildford, UK, personal communication). In this study, we have shown that there exists a molecular rationale for a transcriptional activation by PPAR, enabled by the presence of a functional PPRE within the proximal promoter of the human PXR gene. In addition, this suggests that transcriptional activation of PXR by nonligands such as clofibrate may be a cross-species event, since the initial observations were made in rat liver. This is of potential interest in terms of extrapolation from rodent models to the human situation and may go some way to explaining the apparent conflict between the high species selectivity of the PXR ligand binding domain (Xie et al., 2000a) and the cell type-specific responses observed in vivo (Barwick et al., 1996; Swales et al., 2003).
If the stimulating chemical is not a direct ligand of PXR, the question as to the biological relevance of such activations is raised. It is possible that such activation may be effectively a "bystander effect", whereby PXR gene activation is not the desired biological endpoint but merely a silent side effect of chemical stimulation (Butte, 2002; Cajiao et al., 2004; Jansen and Gerstein, 2004). However, as described in Fig. 5, an increase in PXR protein levels may result in increased PXR activation by endogenous ligands or coadministered chemicals. This increased biological activity could result in disruption of endogenous metabolic processes, leading to a loss of cellular homeostasis. Such events could therefore represent a mechanism by which adverse side effects could occur; indeed, these events may become increasingly relevant as the potency of chemicals against PXR-activating LATFs increases and hence their activation of PXR gene expression.
In summary, we have undertaken an examination of the proximal promoter of PXR and provide a molecular rationale for the activation of PXR gene expression by LATFs, including PPAR. We have proposed a model of PXR regulation whereby PXR levels are controlled through the activation of both PXR and other LATFs, placing PXR at the center of a regulatory network designed to sense, assimilate, and respond to a chemical stimulus.
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Footnotes |
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ABBREVIATIONS: LATF, ligand-activated transcription factor; PXR, pregnane X receptor; GR, glucocorticoid receptor ; PPAR, peroxisome proliferator-activated receptor ; kb, kilobase(s); CAR, constitutive active receptor; VDR, vitamin D receptor binding element; Wy-14,643, 4-chloro-6-(2,3-xylidino)-2-pyrimidinyl)thioacetic acid (pirinixic acid); SEAP, secretory alkaline phosphatase; PCR, polymerase chain reaction; PPRE, PPAR binding site; DTT, dithiothreitol; EMSA, electromobility shift assay; bp, base pair(s); HNF, hepatic nuclear factor; C/EBP, CAAT-enhancer binding protein ; ANOVA, analysis of variance.
Address correspondence to: Dr. Nick Plant, School of Biomedical and Molecular Sciences, University of Surrey, Guildford, Surrey, GU2 7XH, UK. E-mail: n.plant{at}surrey.ac.uk
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