P450 Gene Induction by Structurally Diverse Xenochemicals: Central Role of Nuclear Receptors CAR, PXR, and PPAR

Archives of Biochemistry and Biophysics Vol. 369, No. 1, September 1, pp. 11–23, 1999 Article ID abbi.1999.1351, available online at http://www.idealibrary.com on

MINIREVIEW P450 Gene Induction by Structurally Diverse Xenochemicals: Central Role of Nuclear Receptors CAR, PXR, and PPAR 1 David J. Waxman 2 Division of Cell and Molecular Biology, Department of Biology, Boston University, Boston, Massachusetts 02215

Received May 26, 1999

The biochemistry of foreign compound metabolism and the roles played by individual cytochrome P450 (CYP) enzymes in drug metabolism and in the toxification and detoxification of xenochemicals prevalent in the environment are important areas of molecular pharmacology and toxicology that have been widely studied over the past decade. Important advances in our understanding of the mechanisms through which foreign chemicals impact on these P450-dependent metabolic processes have been made during the past 2 years with several key discoveries relating to the mechanisms through which xenochemicals induce the expression of hepatic P450 enzymes. Roles for three “orphan” nuclear receptor superfamily members, designated CAR, PXR, and PPAR, in respectively mediating the induction of hepatic P450s belonging to families CYP2, CYP3, and CYP4 in response to the prototypical inducers phenobarbital (CAR), pregnenolone 16a-carbonitrile and rifampicin (PXR), and clofibric acid (PPAR) have now been established. Two other nuclear receptors, designated LXR and FXR, which are respectively activated by oxysterols and bile acids, also play a role in liver P450 expression, in this case regulation of P450 cholesterol 7a-hydroxylase, a key enzyme of bile acid biosynthesis. All five P450-regulatory nuclear receptors belong to the same nuclear receptor gene family (family NR1), share a common heterodimerization partner, retinoid X-receptor (RXR), and are subject to cross-talk interactions with other 1 This study was supported in part by NIH Grant ES07381/Superfund Basic Research Center at Boston University, with funding provided by the U.S. EPA. 2 Address correspondence to author at Department of Biology, 5 Cummington St., Boston, MA 02215. Fax: 617-353-7404. E-mail: [email protected].

0003-9861/99 $30.00 Copyright © 1999 by Academic Press All rights of reproduction in any form reserved.

nuclear receptors and with a broad range of other intracellular signaling pathways, including those activated by certain cytokines and growth factors. Endogenous ligands of each of those nuclear receptors have been identified and physiological receptor functions are emerging, leading to the proposal that these receptors may primarily serve to modulate hepatic P450 activity in response to endogenous dietary or hormonal stimuli. Accordingly, P450 induction by xenobiotics may in some cases lead to a perturbation of endogenous regulatory circuits with associated pathophysiological consequences. © 1999 Academic Press Key Words: P450 induction; nuclear receptors; CAR; PXR; PPAR.

Over the past decade there has been an enormous increase in the database of known P450 3 genes, which are found in a wide range of phyla and species (1). Mammals contain at least 17 distinct P450 gene families that together code for an estimated 50 – 60 individual P450 genes in any given species (2). Four of these P450 gene families (families CYPs 1– 4) code for 3

Abbreviations used: P450 or CYP, cytochrome P450; Ah receptor, aryl hydrocarbon receptor; CAR, constitutive androstane receptor; PB, phenobarbital; PXR, pregnane X receptor; PPAR, peroxisome proliferator-activated receptor; LXR, liver X receptor; FXR, farnesol X receptor (also known as BAR, bile acid receptor); TR, thyroid hormone receptor; RAR, retinoic acid receptor; PBRE, PB-responsive enhancer; NR1 and NR2, nuclear receptor binding sites 1 and 2; DR, direct repeat; ER, everted repeat; IR, inverted repeat; RXR, retinoid X receptor; SRC-1, steroid receptor coactivator-1; PCN, pregnenolone 16a-carbonitrile; PPC, peroxisome proliferator chemical; PPRE, PPC response element; JAK, Janus tyrosine kinase; STAT, signal transducer and activator of transcription; CPF, CYP7A promoter-binding factor; TNFa, tumor necrosis factor-a; IL, interleukin. 11

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MINIREVIEW TABLE I

CYP Induction Mediated by Nuclear Receptors

P450 inducing agents Polycyclic aromatic hydrocarbons Phenobarbital Dexamethasone Fibrate drugs Cholesterol Bile acids b Thyroid hormone a b

Prototypic responsive rat liver CYPs

Receptor

1A1, 1A2, 1B1 2B1, 2B2 3A1, 3A2, 3A23 4A1, 4A2, 4A3 7A1 7A1 P450 reductase

Ah receptor a CAR PXR PPARa LXRa FXR TR

PAS transcription family member, not a nuclear receptor. Inhibitors of CYP7A1 transcription.

liver-expressed enzymes that metabolize foreign compounds (drugs, environmental chemicals, and other xenobiotics) and endogenous lipophilic substrates. These P450 genes are regulated in a variety of ways and at multiple levels: they exhibit tissue-specific expression, are regulated by endogenous hormones and cytokines, and respond to structurally diverse foreign chemicals, which often increase P450 protein levels by stimulating P450 gene transcription initiation. This “P450 induction” response has a major impact on P450-dependent drug metabolism, pharmacokinetics, and drug– drug interactions, on the toxicity and carcinogenicity of foreign chemicals, and on the activity and disposition of endogenous hormones (3). P450s belonging to the other 13 mammalian P450 gene families (CYPs 5, 7, 8, 11, 17, 19, 21, 24, 26, 27, 39, 46, and 51) typically do not metabolize foreign chemicals. Rather, they metabolize endogenous substrates along physiologically important pathways: CYPs 5 and 8 are important for thromboxane and prostacyclin biosynthesis; CYPs 11, 17, 19, and 21 catalyze hydroxylation reactions required for steroid hormone biosynthesis from cholesterol; CYPs 7, 24, 27, and 51 catalyze hydroxylations required for the biosynthesis of bile acids, activated vitamin D3, and cholesterol; and CYP26 catalyzes the hydroxylation of retinoic acid, a step that may be important during development. These “biosynthetic” P450s are also subject to regulation in a tissue-specific manner and by endogenous hormones and other factors, as reviewed elsewhere (4). Many, but not all, genes belonging to families CYPs 1– 4 can be transcriptionally activated by foreign chemicals that induce P450 gene expression through one of four receptor-dependent mechanisms. As discussed in greater detail below, specific xenobiotic receptor proteins have now been identified as playing a key role in gene induction stimulated by each of the four mechanistically distinct classes of P450-inducing xenochemicals (Table I). One of these receptors, the Ah receptor,

is a helix-loop-helix protein that belongs to the PAS family of transcription factors and stimulates transcription of CYP1 genes. The Ah receptor becomes activated by binding an aromatic hydrocarbon ligand in the cytosol. The activated receptor then translocates to the nucleus, heterodimerizes with the nuclear factor Arnt, binds to DNA enhancer sequences (“xenobioticresponse elements”) found upstream of CYP1 and other Ah receptor-inducible genes, and stimulates target gene transcription. The overall process is conserved in many cell types and across species and accounts for the induction of CYP1 genes by a large number of polycyclic aromatic hydrocarbons, including several important environmental carcinogens found in auto exhaust and cigarette smoke. The molecular mechanism of CYP1 induction and the role of Ah receptor–Arnt in this process and in the toxicological effects of aromatic hydrocarbons (5, 6) are fairly well understood and have been reviewed elsewhere (7, 8). In contrast to the induction of CYP1 genes, the three other known xenobiotic induction mechanisms involve three distinct “orphan receptors” (i.e., receptors whose endogenous, physiological ligands are unknown) that belong to the nuclear receptor/steroid receptor superfamily (9 –11). These are (i) a novel orphan nuclear receptor, termed CAR, which was recently shown (12, 13) to mediate the widely studied induction of CYP2B genes by phenobarbital (PB) and many other “PB-like” lipophilic chemicals (14); (ii) the pregnane nuclear receptor PXR (also called PAR and SXR), which activates CYP3A genes in response to diverse chemicals, including certain natural and synthetic steroids (15–17); and (iii) the peroxisome proliferator-activated receptor PPAR (18), which mediates induction of the fatty acid hydroxylases of the CYP4A family by many acidic chemicals classified as “non-genotoxic” carcinogens and peroxisome proliferators (19). All three xenochemical receptors are most highly expressed in liver, where they are responsive to endogenous ligands. The discovery of endogenous ligands for CAR (adrostanes, which inhibit receptor activity), PXR (certain pregnenolone derivatives and other steroids), and PPAR (certain polyunsaturated long-chain fatty acids and their metabolites) (Table II) supports the proposed role for these receptors in modulating liver CYP expression in response to endogenous hormonal stimuli (127), in addition to their more obvious role in modulating liver capacity for foreign chemical metabolism by induction of cytochrome P450. Nuclear receptor-dependent modulation of liver P450 activity also occurs at the level of NADPH–P450 reductase, which provides electrons required for all microsomal P450 activities, and is regulated by thyroid hormone and the thyroid hormone receptor via multiple mechanisms (20, 21) (Table I). This review evaluates recent studies leading to the identification of these three nuclear receptors as key

MINIREVIEW TABLE II

Nuclear Receptors: Endogenous Ligands and DNA Response Elements in Target CYP Genes Nuclear receptor

Representative endogenous ligands a

AGGTCA-based DNA response element b

CAR PXR PPARa LXRa FXR TR

Androstanol, androstenol Pregnenolone, corticosterone Linoleic acid, arachidonic acid 24(S)-hydroxycholesterol Chenodeoxycholic acid Thyroid hormone

DR4 DR3, ER6 DR1 DR4 IR1 DR4

a Ligands shown stimulate receptor activity, with the exception of the CAR ligands, which are inhibitory. b Shown are the hexameric repeat motifs, including the indicated number of base pair spacing between repeats, that have been found in one or more CYP genes regulated by each receptor. Exception: IR1 motif shown for FXR was identified in an hsp27 ecdysone response element (126) and that for TR is derived from the rat P450 reductase gene (21). Other response element motifs found for TR include ER6, ER8, and IR0 (10). Also see Fig. 2.

mediators of the foreign chemical induction of liverexpressed P450 genes. The discovery that the nuclear receptors LXR and FXR play a key role in the physiological regulation of CYP7A-dependent bile acid biosynthesis is also discussed. Finally, the mechanisms of cross-talk between nuclear receptor-dependent P450 induction pathways and intracellular signaling pathways involving cytokines and endogenous hormones are considered (Fig. 1), and several important unresolved questions and areas for further study are identified. PB INDUCTION OF MAMMALIAN LIVER CYPS

Earlier studies of PB induction mechanisms have led to the following conclusions (14). 1. Certain liver P450 enzymes, e.g., rat CYP2B1, are highly inducible in vivo following treatment with PB, with increases of up to 50- to 100-fold occurring at the level of P450 protein, mRNA, and transcription initiation. PB induces several other P450 genes, and also many non-P450 genes, to a much lower extent (e.g., 2to 4-fold induction of CYP2C6 and CYP3A in the same rat liver model as CYP2B1) and perhaps by a distinct mechanism(s). Human CYP2B6 is also inducible by PB in liver cells, albeit to a much lesser extent than its rodent CYP2B counterparts (128). PB induction of CYP mRNA does not require new protein synthesis (22), despite earlier indications to the contrary. 2. PB may induce liver CYPs and related enzymes by multiple mechanisms, as indicated by the finding that lower doses of PB are required for induction of CYP2B1 compared to CYP3A1 mRNA in primary hepatocyte cultures (23) and by the posttranscriptional PB induc-

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tion of some P450 genes [e.g., mouse Cyp2a5 (24)]. The slower time course for PB induction reported for NADPH–P450 reductase (25) compared to CYP2B genes (26) also argues in favor of multiple induction mechanisms. 3. Evidence for a receptor-dependent induction mechanism includes the tissue specificity of PB induction (liver .. other tissues) and the finding of saturable dose–response curves and structure–activity relationships in the case of certain families of PB-like inducers (e.g., among PCB congeners). Because diverse chemicals with no obvious structural relationship other than their general lipophilicity can serve as PBlike inducers, PB and PB-like inducers may bind to a common receptor with a “sloppy fit” or an “elastic recognition site” (27). A similar paradigm may apply to the receptor-dependent mechanisms involved in the induction of CYP3A and CYP4A genes (see below). 4. CYP induction by PB is a regulated process: it exhibits tissue specificity, developmental and hormonal regulation, and sex dependence (e.g., female rats being less responsive than males). PB induction is also characterized by some striking strain differences that may support a genetic approach to further elucidate the genes and factors required for PB induction (14). 5. PB induction responses are seen in mammals, including humans, in avian species, and even in bacteria. Bacterial PB induction mechanisms have been elegantly elucidated at the molecular level and several central factors have been identified (28). However, it is now clear that the key feature of the bacterial PB induction mechanism, namely, removal of the repressor protein Bm3R1 from a 17-bp promoter regulatory sequence (“Barbie box”), does not serve as a paradigm for mammalian systems, despite the fact that a homologous promoter sequence can be found in several PB-

FIG. 1. CYP gene induction: Cross-talk between foreign chemical and endogenous regulator pathways.

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responsive mammalian genes, including CYP2B1 (29). This conclusion is most clearly established by deletion or mutation of the CYP2B promoter’s Barbie box-like sequence, which does not abolish PB responses when assayed in intact liver or cultured hepatic cells (30, 31). Nevertheless, there may be some conceptual similarity between the bacterial and mammalian induction mechanisms, in view of the recent discovery (discussed below) that PB-like inducers can activate the mammalian nuclear receptor CAR by reversing the repression (i.e., the inhibition) of this receptor’s transcriptional activity by certain endogenous steroids. IDENTIFICATION OF AN UPSTREAM PB-RESPONSIVE ENHANCER (PBRE) IN CYP2B GENES

Upstream but not proximal promoter CYP2B gene sequences are required for PB induction in vivo. Mice expressing rat CYP2B2 transgenes containing 19 kb of 59-flanking DNA exhibit the striking liver specificity and strong PB inducibility that is characteristic of the endogenous mouse Cyp2b10 gene, whereas mice expressing the identical CYP2B2 transgene, but containing only 800 bases of 59-proximal DNA, do not (32). Subsequent to this report, a highly PB-responsive primary rat hepatocyte culture system (33) was utilized in transient transfection studies to localize what has turned out to be a bonafide PB enhancer in the distal 59 region of CYP2B2 (nts 22318 to 22155) (34). This finding was soon confirmed using a novel in situ rat liver transfection assay (30) and later extended to mouse Cyp2b10, whose upstream PB-responsive enhancer (PBRE) is highly conserved with that of CYP2B2 (35). These CYP2B PBREs are now recognized as being composed of a central binding site for the nuclear factor NF1 flanked by two nuclear receptor binding sites, designated NR1 and NR2 (36). Each of the NR sites consists of two imperfect direct repeats spaced by 4 bps (“DR4 motif”) based on the half-site sequence AGGTCA [cf., (37)] (Fig. 2). At least one of the NR sites must be present to maintain PB inducibility. By contrast, whereas the NF1 site may be required for maximal PB induction, it is nonessential for the basic PB response, as shown by transfection studies (35, 38 – 40) and in transgenic mice (41). This conclusion is supported by the finding that the NR sites, but not the central NF1 site, are highly conserved between rodent and human CYP2B genes (13). Further studies established that a 51-bp Cyp2b10 fragment encompassing the PBRE, when linked to a heterologous promoter and transfected into primary mouse hepatocytes, confers reporter gene inducibility not only to PB, but also to a series of 16 structurally unrelated PB-like inducers in excellent rank order to their induction of the endogenous mouse Cyp2b10 gene (36). Thus, a distal PBRE located at 22.3 kb is, by

FIG. 2. Role of nuclear receptors in CYP gene induction. Shown is the structure of a xenobiotic receptor–RXR heterodimer (e.g., CAR– RXR, PXR–RXR, or PPAR–RXR) bound to two copies of a hexameric DNA response element based on the sequence AGGTCA spaced by X nucleotides. The hexameric repeat can be arranged as a DR, ER, or IR motif, as indicated. Naturally occurring DNA response elements for these nuclear receptors are generally imperfect repeats of the AGGTCA sequence. The extent to which retinoids that bind to RXR synergize with the xenobiotic in stimulating CYP gene transcription is uncertain.

itself, sufficient to confer a high degree of PB inducibility and to maintain the discrimination between PB-like inducers with widely differing potencies that is characteristic of liver cells. This important observation highlights the central nature of the PBRE and its associated binding proteins for the PB induction response. Moreover, the CYP2B2 PBRE region undergoes a PB-induced increase in nuclear protein binding, as demonstrated by in vivo footprinting, which is dependent on native chromatin structure (42). Thus, this region of DNA undergoes dynamic changes in intact liver in vivo in response to PB treatment. Other CYP2B regulatory elements that have been identified include proximal elements that strongly activate the promoter and negative element(s) within the region of 2971 to 2775 bp of Cyp2b10. These negative elements may help maintain the low basal promoter activity in the absence of inducers that characterizes the PB-inducible CYP2B genes (29, 40). CENTRAL ROLE OF THE “CONSTITUTIVELY ACTIVE RECEPTOR” CAR IN PB INDUCTION

A major advance in our understanding of the mechanism of PB induction was provided by the discovery of Negishi and co-workers that the liver-enriched orphan nuclear receptor CAR (43, 44) is the key regulated factor that interacts with PBRE and confers PB-inducible gene transcription (12). CAR binds to each of the PBRE NR sites as a heterodimer with the retinoid X receptor RXR, which serves as a common heterodimerization partner for many orphan nuclear receptors (10), including those involved in the induction of CYP3A (PXR) and CYP4A (PPAR) and the regulation of

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CYP7A (LXR, FXR) (see below). The (CAR–RXR)– PBRE binding interaction is functional, as demonstrated by the ability of a CAR expression plasmid to stimulate PBRE-dependent reporter gene activity following CAR transfection into HepG2 cells (12). Surprisingly, this CAR-dependent transcriptional stimulation does not require treatment of the cells with PB. By contrast, in liver in vivo, CAR–RXR DNA-binding activity is increased by PB at least several fold, as judged by gel mobility shift assay of liver nuclear extracts from PB-treated compared to untreated mice using an NR1 binding site probe. Moreover, PB-induced CAR activation precedes Cyp2b10 induction, as would be expected if these events are causally linked (12). In principle, two types of mechanisms could explain the observed activation of CAR DNA-binding by PB. (i) PB may increase CAR DNA-binding activity in liver nuclei, and thereby increase PBRE-dependent gene transcription, by stimulating translocation of CAR from the cytosol to the nucleus. Alternatively, (ii) CAR could be constitutively localized in the nucleus, in which case PB would need to activate the DNA-binding activity of the CAR–RXR heterodimer. Precedent for both types of receptor regulatory mechanisms (ligandinduced nuclear localization and ligand-activated DNA binding) is provided by other receptors within the nuclear receptor superfamily (45, 46). However, while these mechanisms may explain PB induction of CARdependent, PBRE-linked CYP2B gene transcription, they do not explain the observed PB independence of CAR-stimulated PBRE-reporter gene transcription seen in CAR-transfected HepG2 cells (12). Indeed, unlike most other nuclear receptors, CAR has been characterized as a constitutively active receptor, i.e., one that does not require ligand binding for transactivation (44). The fact that CAR is a constitutively active receptor would seem to be inconsistent with the requirement for PB treatment in vivo to activate liver CAR DNA-binding activity (12) and with the proposal that this receptor mediates the strong PB induction response that occurs in vivo. A possible resolution to this paradox was recently provided by Forman et al. (47), who discovered that CAR is actually a “constitutive androstane receptor” that interacts in a stereospecific manner with two steroids, androstanol (5a-androstan3a-ol) and androstenol (5a-androst-16-en-3a-ol). Unexpectedly, however, these androstanes inhibit CAR transcriptional activity by a unique mechanism that is reported to involve dissociation of the (CAR–RXR)– DNA complex from the nuclear receptor coactivator SRC-1 (47). SRC-1 is a widely expressed coactivator that serves as a general transducer between DNAbound nuclear receptors and the basal transcriptional machinery and is required for transcriptional stimulation by many nuclear receptors (48). Accordingly, in-

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hibitory adrostanes may bind directly to the DNAbound CAR–RXR heterodimer in a manner that alters the nuclear receptor complex’s conformation and prevents its interaction with SRC-1. This novel finding suggests the following model: in liver, where CAR is most highly expressed (44), endogenous inhibitory steroids related to androstanol and androstenol bind to CAR and maintain it in an inactive state. In the presence of PB or PB-like inducers, however, the binding of inhibitory androstanes to CAR is abolished and receptor activity is thereby derepressed. The intrinsic constitutive activity of CAR would thereby become manifest, leading to transactivation of CYP2B and other PBRE-regulated target genes. The observation that CAR can directly transactivate a PBRE-linked reporter gene in transfected cells in the absence of PB inducers (12) could thus be explained by the receptor’s constitutive activity and by the absence of inhibitory androstanes in the HepG2 cell transfection system. This model recently received strong experimental support from a study demonstrating that, in the presence of the CAR inhibitor adrostenol, PB and PB-like inducers transactivate a PBRE found 1.7 kb upstream of the human CYP2B6 gene in a CAR-dependent manner (13). Thus, in human HepG2 hepatoblastoma cells stably transfected with CAR, androstenol treatment inhibits endogenous CYP2B6 expression, while PB (5 mM) or the potent PB-like inducer TCPOBOP (25 nM) (49) reverses this inhibition leading to CYP2B6 induction. Several other PB-like inducers can also reverse the inhibitory effects of androstenol, resulting in a CAR-dependent derepression of CYP2B6 expression and also transactivation of a PBRE-linked reporter gene (13). CAR-DEPENDENT PB INDUCTION: UNANSWERED QUESTIONS

These exciting new findings raise many important questions about the mechanism of PB induction and the role of CAR in this process. 1. Do androstanes bind directly to CAR (or to a CAR–RXR heterodimer) to inhibit SRC-1 binding and CAR’s constitutive transcriptional activity? Do PB-like inducers reverse androstane inhibition by binding directly to CAR? Does this PB binding interaction involve direct competition leading to steroid displacement at the androstane binding site, or does PB bind to an allosteric site on the receptor? Does PB binding induce or stabilize a CAR conformation that facilitates functional interactions with SRC-1? CAR-binding studies carried out with the potent PB-like inducer TCPOBOP (49) may be useful in this regard. 2. By which mechanism(s) is CAR’s constitutive transcriptional activity suppressed in liver in the absence of PB or PB-like inducers? Androstanol and androste-

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nol, the most potent CAR inhibitory steroids identified thus far, may not fully explain this suppression, because the half-maximal CAR inhibitory activity of these steroids, 0.4 mM (47), exceeds the measured human plasma concentration by ;40-fold (adult males) or ;200-fold (adult females) (50). Conceivably, circulating levels of the CAR inhibitory androstanes could be much higher in the rat, which shows a more dramatic PB induction response than is seen in humans (14). Additionally, CAR suppression in vivo may involve more potent metabolites of these androstanes, perhaps including hydroxylated metabolites formed by liver CYP enzymes. Conceivably, some PB-like inducers may activate CAR indirectly, e.g., by binding to, and thereby inhibiting the catalytic activity of constitutively expressed cytochrome P450 enzymes that catalyze formation of CAR inhibitory hydroxylated androstanes. In an earlier review, we hypothesized one such mechanism: inhibition by PB of a cytochrome P450catalyzed hydroxylation reaction that activates a steroidal repressor of CYP2B1 (14). More detailed investigation, including (i) analysis of liver steroid profiles to identify CAR inhibitory molecules and (ii) characterization of the CAR inhibitory activity of liver microsomal metabolites of androstane and androstene could be carried out to test this hypothesis and to identify more potent endogenous CAR inhibitors. 3. What impact does phosphorylation/dephosphorylation have on CAR transcriptional activity? Numerous studies implicate phosphorylation/dephosphorylation in the regulation of PB induction, e.g., the potent inhibitory effects of the protein phosphatase inhibitor okadaic acid on PB induction (51, 52). Is CAR nuclear localization, DNA binding, or transcriptional activity modulated by reversible phosphorylation of CAR itself or of an associated component? Modulation of nuclear receptor activity (including that of PPAR; see below) through receptor phosphorylation reactions is well established (53) and provides an important precedent for this potential CAR regulatory mechanism. 4. Which of the many other liver PB induction responses are mediated by CAR? Are the effects of CAR limited to the highly inducible CYP2B genes or do the promoters of other, less highly PB-responsive liver CYP genes also contain PBREs that are transactivated by CAR? Further investigation of other PB-inducible genes involved in xenobiotic metabolism, including rat CYPs 2A1 and 2C6, NADPH–P450 reductase, epoxide hydrolase, and certain glutathione S-transferases and UDP– glucuronyltransferases is likely to be useful in this regard. Targeted disruption of the mouse CAR gene (CARb) will also be revealing with respect to the role of CAR in each of these gene induction responses and in the pleiotropic effects of PB-like inducers (e.g., liver enlargement, proliferation of smooth endoplasmic reticulum, tumor promotion).

5. What role does CAR play in the activation of CYP3A genes? This possibility is suggested by the intriguing finding that CAR–RXR heterodimers can bind to and transactivate an ER6 sequence (everted repeat, with 6-bp spacing) from the human CYP3A4 gene, which encompasses a CAR-like DR4 sequence motif. However, CAR induction of the endogenous HepG2 cell CYP3A4 gene was not shown in these studies (13). Nevertheless, this finding raises the possibility that PB may activate CYP3A and perhaps other PB-responsive genes by mechanisms that involve two receptors, CAR and PXR (see below). 6. Do other nuclear receptors with specificity for DR4 motifs, such as LXRa and TR (Table II), interact with CYP2B or other CAR-responsive elements to modulate (either to antagonize or enhance) their transcription? Thyroid hormone inhibition of PB-inducible rat CYP2B expression has been observed, albeit in a strain-dependent manner (54). 7. Do interindividual differences or species differences in the levels of CAR, in the expression of CAR isoforms or related genes within the same nuclear receptor subfamily (11), or in the synthesis or metabolism of CAR inhibitory androstanes contribute to interindividual or interspecies differences in CAR-dependent liver induction responses? Significant sex differences in plasma androstane levels have been reported for both humans and pigs (male .. female) (50), and these differences, as well as potential sex differences in adrostane metabolism, could contribute to the substantial sex differences in PB responsiveness seen in rodent species (55, 56). PXR: A NOVEL ORPHAN NUCLEAR RECEPTOR INVOLVED IN CYP3A INDUCTION

CYP3A4, the most abundantly expressed P450 in human liver, metabolizes a large number of structurally diverse drug substrates. This fact, together with the inducibility of this enzyme by numerous steroids, antibiotics, and other pharmacological agents, gives rise to many drug interactions involving CYP3A enzymes (57). Members of the CYP3A subfamily include 3A4, 3A5, and 3A7 (humans), 3A1, 3A2, and 3A23 (rats), and 3A6 (rabbits) (1). CYP3A genes are characterized by expression in liver and intestine, catalysis of steroid hormone and bile acid 6b-hydroxylation reactions, selective inhibition by the mechanism-based inactivator troleandomycin, and induction by a broad range of steroids and antibiotics. Early studies of rat liver CYP3A enzyme induction made the important, but seemingly paradoxical, observation that both glucocorticoids (such as dexamethasone) and anti-glucocorticoids (such as pregnenolone 16a-carbonitrile, PCN) induce these enzymes at the transcriptional level. Both this finding and the requirement for a rel-

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atively high glucocorticoid concentration for CYP3A induction were recognized as inconsistent with the classical glucocorticoid receptor playing a major role in the CYP3A induction response (58). A broad range of drugs, environmental chemicals, and steroids can serve as CYP3A inducers; however, important species differences in the induction response have been described (59). Most notably, while the rat, rabbit, and human CYP3A genes are all inducible by dexamethasone, the anti-glucocorticoid PCN is an efficacious CYP3A inducer in the rat but not in humans or rabbits. By contrast, the antibiotic rifampicin is an excellent CYP3A inducer in humans and rabbits, but not in the rat. Transfection studies carried out in rat and rabbit hepatocytes and utilizing CYP3A constructs containing dexamethasone-responsive regulatory elements derived from rat CYP3A23 (60 – 62), rabbit CYP3A6, and human CYP3A4 genes (63) demonstrated that the species-specific induction responses are not due to species differences in the individual CYP3A genes, but rather are a function of factor(s), most likely a receptor, provided by the host species liver cell (63). Moreover, the finding that the dexamethasone response element is composed of two imperfect repeats of a consensus nuclear receptor half-site sequence AG(G/T)TCA (62, 64) strongly suggested a role for a nuclear receptor, albeit one that is distinct from the glucocorticoid receptor. In the case of rat CYP3A23, the dexamethasone-responsive sequence contains a DR3 motif (direct repeat, separated by 3 bp; AGTTCA-N 3-AGTTCA) that is also present in rat CYP3A2, whereas in the human CYP3A4 gene, the response element contains an unusual ER6 motif (everted repeat, separated by 6 bp; TGAACT-N 6-AGGTCA) that is conserved in human CYP3A5 and rabbit CYP3A6 (63) (Table II). The nuclear receptor that binds to and transactivates the DR3-based dexamethasone response element of rat CYP3A23 was recently cloned from mouse liver and termed mouse pregnane X receptor (mPXR) based on its strong activation by steroids related to pregnenolone (15). mPXR can also be activated by the antiglucocorticoid PCN, and by several other CYP3A inducers, including dexamethasone. mPXR is most highly expressed in liver and intestine, tissues where CYP3A is also highly expressed. Characterization of a human homolog, variously designated hPXR (65), hPAR (17), and SXR (16), indicates that this receptor, as well as mPXR, heterodimerizes with RXR and efficiently transactivates either DR3 elements (present in rat CYP3A genes) or ER6 elements (present in human and rabbit CYP3A genes). However, mPXR and hPXR share only ;75% amino acid sequence identity in their COOH-terminal ligand-binding domain region (vs 96% identity between their DNA-binding domains), and this apparently results in significant differences in li-

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gand-binding specificities: hPXR but not mPXR is highly activated by xenochemicals that preferentially induce human CYP3A genes, such as rifampicin, while mPXR but not hPXR exhibits the strong response to PCN that characterizes mouse CYP3A gene induction. Thus, the species-dependent ligand specificity for CYP3A induction seen in vivo can be explained by the corresponding ligand specificity of each species’ PXR receptor. Other CYP3A inducers and PXR activators include anti-hormones belonging to several steroid classes, the organochlorine pesticides trans-nonachlor and chlordane, and various nonplanar chlorinated biphenyls (65, 66). Interestingly, PXR can also be activated by PB (65), suggesting that the effects of PB on liver CYP genes may be mediated by multiple receptors (e.g., CAR for CYP2B and PXR for CYP3A). This conclusion is consistent with the distinct dose–response curves that characterize PB induction of CYP2B and CYP3A genes in hepatocytes (ED 50 ;10 25 and ;10 24 M, respectively) (23) and with several other observations, noted above, that support the existence of multiple induction mechanisms. PXR AND CYP3A INDUCTION: AREAS FOR FURTHER STUDY

Important unresolved questions regarding the role of PXR in CYP gene induction include the following. 1. Are mPXR and hPXR encoded by orthologous genes, or do they represent a and b subtypes within the same nuclear receptor subfamily? This latter possibility is suggested by the relatively low amino acid conservation (;75% identity) within the ligand-binding domain of mPXR and hPXR, noted above, compared to the much higher sequence identity of true mouse/human nuclear receptor homologs (cf. 95% ligand-binding domain identity between mouse and human glucocorticoid receptor). However, further screening has not yielded other mouse or human PXR-related cDNAs, suggesting that mPXR and hPXR may represent unusually divergent orthologs that represent receptor adaptation to each species’ unique diet (16) or perhaps their distinct endogenous steroid profiles. 2. What unique structural features enable PXR to bind to both DR3 and ER6 AG(G/T)TCA sequence motifs? 3. Why are orphan receptors with steroidal ligands, namely CAR and PXR, utilized to mediate the liver cell’s responses to structurally diverse xenochemicals? In the case of CAR, the steroidal ligand inhibits receptor-dependent gene transcription, whereas with PXR the steroidal ligand exerts a stimulatory response. PPAR (see below) can also be activated by endogenous steroids, notably dehydroepiandrosterone 3b-sulfate (67), although direct binding of the steroid has not been established in that case (68). Does the utilization of the

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steroid receptor PXR to regulate liver P450 induction reflect its role as a broadly based “steroid and xenobiotic sensor” (69) whose biological function is to stimulate synthesis of CYP3A enzymes that catabolize circulating steroidal substrates? This possibility is supported by the responsiveness of PXR to steroids belonging to several distinct classes (pregnanes, estrogens, and corticoids) (16) and by the catalysis by many CYP3A enzymes of steroid 6b-hydroxylation reactions using a wide range of substrates, including androgens, corticoids, progestins, and bile acids [e.g., (70, 71)]. 4. Does PXR have a specific, high-affinity, and physiologically relevant natural ligand in either rodents or humans? Although direct binding of CYP3A steroidal inducers to PXR has not been demonstrated, this may reflect a relatively low binding affinity, which would be consistent with the proposed role of PXR as a sensor for a wide range of steroids and foreign chemicals. Indirect evidence for steroid–PXR binding interactions has been provided in studies using a coactivator–receptor ligand assay (72), where PXR activators induce an association between PXR’s ligand-binding domain and a fragment of the steroid receptor coactivator SRC-1 (15, 65). 5. Do retinoids synergize with PXR ligands to activate PXR–RXR heterodimers and stimulate CYP3A transcription, as can occur for PPAR–RXR heterodimers and CYP4A transcription (see below)? Similarly, do retinoids synergize with CAR activators to stimulate CYP2B transcription (Fig. 2)? 6. Do other nuclear receptors also transactivate CYP3A genes? This question is important in view of the CYP3A inductive effects of dexamethasone, a portion of which could be mediated by the classical glucocorticoid receptor, and in view of the potential for both CAR and PXR to bind to and transactivate the ER6 element of CYP3A4. Moreover, both CAR and PXR are classified within the same nuclear receptor subfamily [family NR1, subfamily I (11)], suggesting that other NR1I subfamily members might also contribute to the regulation of CAR- or PXR-like response elements upstream of CYP2B, CYP3A, or other xenobiotic inducible genes, particularly in extrahepatic tissues, where CAR and PXR expressions are greatly reduced compared to liver. Of note, 1a,25-dihydroxyvitamin D3, the physiological ligand for the vitamin D3 receptor, which also belongs to nuclear receptor subfamily NR1I (11), exhibits a specificity for activation of DR3based response elements (10) and can also stimulate CYP3A gene expression (73). 7. Does PXR transactivate CYP genes other than CYP3A? This possibility is suggested by the presence of putative PXR target sequences in several CYP2A, CYP2C, and CYP2E genes, as well as in the promoters of UDP– glucuronyltransferase 1A6 and NADPH–P450 reductase (16).

CYP4A INDUCTION BY PEROXISOME PROLIFERATOR CHEMICALS (PPCs): ROLE OF PPARa

CYP4A enzymes catalyze the oxygenation of biologically important fatty acids, including arachidonic acid and other eicosanoids. CYP4A gene expression can be transcriptionally activated in both liver and kidney by a range of acidic drugs and other xenobiotics, including hypolipidemic fibrate drugs, phthalate ester plasticizers used in the medical and the chemical industries, and various other environmental pollutants (74). These foreign chemical CYP4A inducers are classified as peroxisome proliferator chemicals (PPCs), since they markedly induce hepatic peroxisomal enzymes, which leads to a dramatic increase in both the size and the number of liver cell peroxisomes. The receptor protein responsible for CYP4A induction, peroxisomal enzyme induction, and hepatic peroxisome proliferation was first cloned in 1990 and is termed PPARa, peroxisome proliferator-activated receptor-a (18). The tissue distribution of PPARa, namely, liver . kidney . heart . other tissues, mirrors the PPC responsiveness of these tissues. CYP4A induction in liver and kidney and hepatic peroxisome proliferation are abolished in PPARa gene knockout mice (75), demonstrating the essential nature of PPARa in vivo for these responses. This finding is consistent with the presence in the 59-flank of CYP4A genes of functional PPC response elements (PPREs) that bind PPARa as a heterodimer with RXR and serve as functional enhancers with respect to the stimulation of CYP4A transcription by PPCs (76, 77). Upon binding a PPC or endogenous fatty acid ligand, PPARa, which is constitutively nuclear, displays enhanced binding to DNA as a PPARa–RXR heterodimer; this ligand-induced DNA binding is most readily apparent at physiological concentrations of receptor protein (78). In the case of the rabbit CYP4A6 gene, a strong PPRE enhancer localized to nts 2677 to 2644 contains four imperfect repeats of the nuclear receptor half-site AGGTCA. Half-sites 2 and 3 are arranged as a DR1 motif (imperfect hexameric direct repeat, spaced by 1 nt), similar to that found in other PPAR-activatable genes. This DR1 sequence is necessary but not sufficient for specific PPAR–RXR binding and transactivation, which additionally requires half-site 1, located on the immediate 59-flank of the DR1 (79). Half-site 1 enhances the binding of PPARa–RXR and also increases binding specificity by diminishing the binding of RXR homodimers and perhaps other competing nuclear receptor complexes with specificity for DR1 sequences (79, 80). PPARa–RXR complexes bound to the PPREs of CYP4A6 and other responsive genes are “permissive,” insofar as they can be synergistically activated by the combination of a PPC, which binds to PPARa, with 9-cis-retinoic acid or a related retinoid,

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which binds to RXR (76). By contrast, nuclear receptor–RXR heterodimers involving retinoic acid receptor (RAR) are nonpermissive, insofar as RAR allosterically blocks the binding of RXR-specific ligands and recruits the nuclear receptor corepressor N-CoR, leading to RAR–RXR repression of DR1-containing promoters (81). ROLE OF PPARa IN PPC-INDUCED HEPATOCARCINOGENESIS

Foreign chemical PPARa activators not only stimulate peroxisome proliferation in liver cells, but also induce hepatocellular carcinoma development by a non-genotoxic mechanism, i.e., one that does not involve direct DNA damage caused by PPCs or their metabolites. This hepatocarcinogenic response is abolished in mice deficient in PPARa (82), underscoring the central role of PPARa, as opposed to the other two mammalian PPARs (PPARg and PPARd), in PPC-induced hepatocarcinogenesis. Other toxic responses to PPCs, such as the kidney and testicular toxicities of the plasticizer di-(2-ethylhexyl)phthalate, are not prevented by disruption of the PPARa gene (83), raising the possibility that PPARg or PPARd may mediate these responses. The mechanistic basis for the hepatocarcinogenesis of PPCs is incompletely understood, but is hypothesized to involve several key factors and events (19). These include (i) transcriptional activation of lipidmetabolizing enzymes, including CYP4A, leading to the formation of DNA-damaging reduced oxygen species; and (ii) alteration of the balance between hepatocyte proliferation, which is stimulated by PPCs, and hepatocyte apoptosis, which is suppressed following PPC exposure (84). Suppression of apoptosis blocks a critical mechanism for elimination of genetically damaged cells prior to their clonal expansion, leading to fixation of mutations in initiated cells. Unlike rodents, humans exhibit very weak liver peroxisome proliferative responses, suggesting that the striking hepatocarcinogenic actions of PPCs seen in rats and mice may not be predictive of human health risks of PPC exposure. Factors that contribute to the weak response of human hepatocytes to PPCs include (i) the greatly reduced expression of PPARa in human liver compared to rat liver (85); (ii) the lower sensitivity of human PPARa compared to its rodent counterparts with respect to transactivation by at least some PPCs (86); and possibly (iii) the expression of a dominant-negative inhibitory human PPARa variant that is found in at least some individuals (84). Although PPARa may thus play a minor role in humans as a mediator of the deleterious effects of PPCs, it is quite possible that PPARg and PPARd, which are highly expressed in multiple human tissues, may be transactivated by a

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subset of PPCs in a manner that perturbs physiological pathways involving these receptors and elicits a pathophysiological response. CYP4A AND PPAR: FUTURE DIRECTIONS

While many of the molecular details regarding PPARa and its transcriptional activation of CYP4A and other target genes have now been elucidated, several important questions relating to the physiological role and toxicological impact of PPAR receptors remain to be answered. 1. What is the precise role of PPARa in lipid metabolism and homeostasis? Which of the numerous endogenous lipophilic substrates of CYP4A enzymes are key to these metabolic and regulatory processes? Is there any cross-talk or coregulation between PPARa-dependent fatty acid metabolism and LXR- and FXR-regulated cholesterol catabolism, mediated by CYP7A (see below)? 2. What structural features enable PPARa to bind a broad range of xenochemicals, in addition to structurally diverse endogenous fatty acids and related lipophilic substrates? Recent crystallographic analyses of the ligand-binding domains of PPARg and PPARd evidence a large ligand-binding pocket in comparison to other nuclear receptors (87– 89). This finding may have direct relevance for our understanding of the structural basis for the broad ligand specificity of PPARa and perhaps that of CAR and PXR as well. 3. Does PPARg or perhaps PPARd contribute to CYP4A gene expression? These PPARs do have the potential to strongly transactivate CYP4A PPREs (80). The involvement of PPARg or PPARd in basal or perhaps PPC-inducible CYP4A gene expression has not been explored, but is suggested by the PPARa-independent constitutive expression of CYP4A protein(s) seen in liver and kidney (68). Also of potential interest is the high level of PPARg expression in many human tissues (90), including adipose tissue, where many lipophilic foreign chemicals tend to accumulate and where PPARg exerts central regulatory control on adipogenesis (91). PPARg is also highly expressed in hematopoietic cells where it, along with PPARa (92), plays a role in monocyte and macrophage differentiation (93) and may regulate CYP4A enzyme expression. 4. To what extent is CYP4A induction or the induction of other PPAR-dependent responses (e.g., PPCstimulated hepatocarcinogenesis) modulated in vivo through cross-talk between PPAR and other nuclear receptors or other signaling molecules? Thyroid hormone, which suppresses hepatic peroxisome proliferative responses, exhibits inhibitory cross-talk with PPARa, in part via competition between thyroid hormone receptor and PPAR for their common heterodimerization partner RXR (94). Other examples of

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nuclear receptors inhibitory to PPARa activity include RAR, noted above, as well as LXRa, PPARd, and TAK1 (95–97). PPARa can also cross-talk with signaling pathways that do not involve nuclear receptors. Examples include the growth hormone-activated JAK/STAT signaling pathway, which inhibits PPAR transcriptional activity (98), and an insulin-induced serine kinase pathway that activates PPARa by phosphorylation of the receptor on a pair of NH 2-terminal serine residues (99). PPARa, in turn, exhibits striking inhibitory effects on these same hormone/cytokine signaling pathways, as demonstrated by the PPARa-dependent suppression that PPCs confer on growth hormone-regulated, STAT5-dependent (100, 101) liver P450s and other liver gene products (102). CYP7A-DEPENDENT CHOLESTEROL CATABOLISM: ROLE OF NUCLEAR RECEPTORS LXR AND FXR

CYP7A is a liver-specific P450 enzyme that catalyzes cholesterol 7a-hydroxylation, a metabolic reaction that has long been established as the first, and rate-limiting, step in the catabolism of cholesterol to bile acids. CYP7A and liver bile acid biosynthesis are highly regulated by multiple physiological factors, most notably cholesterol, which induces CYP7A activity (“feed-forward pathway”) and bile acids, which serve as end product inhibitors of CYP7A expression upon their return to the liver from the gut via the enterohepatic circulation (“feed-back inhibition pathway”) (103). This feedback inhibition step is an important target for pharmacological intervention to control hypercholesterolemia. This is exemplified by the therapeutic use of cholestyramine, an anion-exchange polystyrene resin that binds bile acids in the human gut and facilitates their elimination, thereby interrupting CYP7A feedback inhibition and ultimately driving increased hepatic uptake and catabolism of circulating cholesterol. The substantial medical importance of CYP7A has stimulated molecular studies of this gene and its regulation, leading to the identification of factors required for its liver-specific expression, diurnal regulation, and regulation by hormonal factors [e.g., (104 –106)]. Nuclear receptor-like promoter sequences with DR1 and DR5 motifs that can mediate bile acid downregulation of CYP7A transcriptional activity have also been identified (107); however, the putative bile acid receptor that interacts with these sequences has remained elusive. Recently, however, the nuclear receptor FXR, which was originally characterized as being weakly activated by farnesol derivatives (“farnesol X receptor”), was shown to bind to and be activated by physiologically relevant concentrations of bile acids (EC 50 ;5–10 mM), most notably the primary bile acid chenodeoxycholic acid and the secondary bile acids lithocholic acid and deoxycholic acid (108 –110). The natu-

rally occurring glycine and taurine conjugates of these bile acids are also efficient activators of FXR, provided that the cellular system used to assay receptor activity coexpresses a suitable bile acid transporter (108). FXR thus serves, and is more appropriately designated, as BAR (bile acid receptor). Interestingly, whereas bile acid-activated FXR suppresses CYP7A promoter activity, bile acid-activated FXR transcriptionally activates the intestinal bile acid-binding transporter gene IBAT, whose expression in vivo is induced by bile acids to facilitate bile acid transport across intestinal enterocytes (109). Whether FXR acts by binding to the CYP7A bile acid response elements identified earlier (107) and the molecular mechanisms by which bile acid-activated FXR suppresses CYP7A transcription, on the one hand, while activating IBAT transcription, on the other hand, are unknown. Precedent for such a dual effect of nuclear receptors on gene transcription is provided by the glucocorticoid receptor (111). The finding that CYP7A gene expression is downregulated by FXR is intriguing, in view of the earlier discovery that a closely related, liver-expressed, nuclear receptor, termed LXRa (liver X receptor-a), mediates oxysterol-stimulated induction of CYP7A gene expression (112, 113). The critical role of LXRa in this induction has been confirmed in mice with a disruption in the LXRa gene: these mice are defective in CYP7A induction in response to dietary cholesterol and accumulate very large amounts of cholesterol in the liver (114). It has been suggested (115) that LXRa may interact with yet another nuclear factor, termed CPF (CYP7A promoter-binding factor), that binds to the CYP7A promoter within 70 bp of the LXRa binding site and may play an important role in the liver-specific expression of CYP7A (116). Other studies suggest that the inhibitory actions of bile acid-activated FXR on CYP7A may involve antagonism of the stimulatory effects of LXRa on this gene (110). Further molecular details of the inhibitory actions of FXR on CYP7A gene transcription as they relate to the interaction of FXR with LXRa, CPF, and perhaps other nuclear factors are likely to emerge in the near future. CYTOKINE INHIBITION OF LIVER P450 INDUCTION

P450 induction in response to foreign chemical exposure is often associated with enhanced formation of toxic or chemically reactive metabolites that elicit pathophysiological responses, such as cell and tissue damage and tumorigenesis. Recent studies have demonstrated, however, that the induction of P450 enzymes by xenochemicals can be modulated by endogenous mediators, including cytokines, hormones, and growth factors that activate intracellular signaling pathways which cross-talk with the receptor-dependent pathways responsible for P450 induction (Fig. 1).

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Elucidation of the underlying mechanisms for these pathways for interaction between cytokines/hormones and xenochemicals may help to define the impact that these endogenous mediators have on cellular responses to, and the toxicological outcome of, foreign chemical exposure. Examples where xenobiotic induction of liver P450 enzymes can be downregulated by cytokines or endogenous hormones include the following: (i) inhibition of polycyclic aromatic hydrocarbon induction of CYP1 enzymes by tumor necrosis factor-a (TNFa) or by transforming growth factor-b (117, 118); (ii) interleukin-6 (IL-6) inhibition of PB-induced CYP2B expression in rat hepatocytes (119) and of rifampicin-induced CYP3A expression in human hepatocytes (117); and (iii) IL-1b and IL-6 inhibition of clofibrate-induced CYP4A expression (120). Numerous other studies document effects of TNFa, interferons a and g, and various interleukins, such as those which become activated during inflammation and infection, on both basal and foreign chemical-inducible liver P450s (121). Cytokine suppression of the xenobiotic-inducible P450 enzymes occurs primarily at the level of P450 gene transcription; however, the underlying molecular mechanisms for this suppression are largely uncharacterized. Key unanswered questions include (i) whether the inhibitory effects of cytokines on P450 induction involve JAK/STAT-dependent signaling (122, 123), which is a major cytokine-activated pathway in mammalian cells, or whether STAT-independent signaling pathways are involved; (ii) whether inhibitory cytokines exert direct effects on the P450 induction process through changes in the expression or activity of the relevant xenochemical receptors (AhR, CAR, PXR, and PPAR); (iii) whether the CYP induction inhibitory effects associated with TNFa or other cytokines are mediated by the cytokine-inducible transcription factor NFKB, which itself can exert inhibition cross-talk with nuclear receptors (124); and (iv) whether cytokine induction, or modulation, of other liver-expressed transcription factors, such as HNF-4 (itself an orphan nuclear receptor), C/EBPb or AP-1, may contribute to the observed inhibition of nuclear receptor-dependent P450 induction. Alternatively, the effects of cytokines on CYP induction could be a consequence of cytokine modulation of nuclear receptor coactivators or corepressors [cf. (125)]. Studies designed to answer these and related questions may uncover novel mechanisms through which cytokines and other local or systemic mediators regulate xenobiotic-responsive nuclear receptors and through them the expression of hepatic P450 enzymes most directly involved in drug metabolism and in the toxication and detoxification of structurally diverse environmental chemicals.

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REFERENCES 1. Nelson, D. R., Koymans, L., Kamataki, T., Stegeman, J. J., Feyereisen, R., Waxman, D. J., Waterman, M. R., Gotoh, O., Coon, M. J., Estabrook, R. W., Gunsalus, I. C., and Nebert, D. W. (1996) Pharmacogenetics 6, 1– 42. 2. Nelson, D. (1999) Arch. Biochem. Biophys. 369, 1–10. 3. Conney, A. H. (1982) Cancer Res. 42, 4875– 4917. 4. Jefcoate, C. R. (Ed.) (1996) in Advances in Molecular and Cell Biology, Vol. 14, pp. 1–379, JAI Press, Greenwich, CT. 5. Gonzalez, F. J., and Fernandez-Salguero, P. (1998) Drug Metab. Dispos. 26, 1194 –1198. 6. Wilson, C. L., and Safe, S. (1998) Toxicol. Pathol. 26, 657– 671. 7. Sogawa, K., and Fujii-Kuriyama, Y. (1997) J. Biochem. (Tokyo) 122, 1075–1079. 8. Hankinson, O. (1995) Annu. Rev. Pharmacol. Toxicol. 35, 307– 340. 9. Meier, C. A. (1997) J. Recept. Signal Transduct. Res. 17, 319 – 335. 10. Mangelsdorf, D. J., and Evans, R. M. (1995) Cell 83, 841– 850. 11. Nuclear Receptors Nomenclature Committee (1999) Cell 97, 161–163. 12. Honkakoski, P., Zelko, I., Sueyoshi, T., and Negishi, M. (1998) Mol. Cell. Biol. 18, 5652–5658. 13. Sueyoshi, T., Kawamoto, T., Zelko, I., Honkakoski, P., and Negishi, M. (1999) J. Biol. Chem. 274, 6043– 6046. 14. Waxman, D. J., and Azaroff, L. (1992) Biochem. J. 281, 577– 592. 15. Kliewer, S. A., Moore, J. T., Wade, L., Staudinger, J. L., Watson, M. A., Jones, S. A., McKee, D. D., Oliver, B. B., Willson, T. M., Zetterstrom, R. H., Perlmann, T., and Lehmann, J. M. (1998) Cell 92, 73– 82. 16. Blumberg, B., Sabbagh, W., Jr., Juguilon, H., Bolado, J., Jr., van Meter, C. M., Ong, E. S., and Evans, R. M. (1998) Genes Dev. 12, 3195–3205. 17. Bertilsson, G., Heidrich, J., Svensson, K., Asman, M., Jendeberg, L., Sydow-Backman, M., Ohlsson, R., Postlind, H., Blomquist, P., and Berkenstam, A. (1998) Proc. Natl. Acad. Sci. USA 95, 12208 –12213. 18. Issemann, I., and Green, S. (1990) Nature 347, 645– 650. 19. Gonzalez, F. J., Peters, J. M., and Cattley, R. C. (1998) J. Natl. Cancer Inst. 90, 1702–1709. 20. Ram, P. A., and Waxman, D. J. (1992) J. Biol. Chem. 267, 3294 –3301. 21. O’Leary, K. A., Li, H. C., Ram, P. A., McQuiddy, P., Waxman, D. J., and Kasper, C. B. (1997) Mol. Pharmacol. 52, 46 –53. 22. Sidhu, J. S., and Omiecinski, C. J. (1998) J. Biol. Chem. 273, 4769 – 4775. 23. Kocarek, T. A., Schuetz, E. G., and Guzelian, P. S. (1990) Mol. Pharmacol. 38, 440 – 444. 24. Aida, K., and Negishi, M. (1991) Biochemistry 30, 8041– 8045. 25. Hardwick, J. P., Gonzalez, F. J., and Kasper, C. B. (1983) J. Biol. Chem. 258, 8081– 8085. 26. Adesnik, M., and Atchison, M. (1986) CRC Crit. Rev. Biochem. 19, 247–305. 27. Okey, A. B. (1990) Pharmacol. Ther. 45, 241–298. 28. Liang, Q., Chen, L., and Fulco, A. J. (1998) Biochim. Biophys. Acta 1380, 183–197. 29. Honkakoski, P., and Negishi, M. (1998) J. Biochem. Mol. Toxicol. 12, 3–9.

22

MINIREVIEW

30. Park, Y., Li, H., and Kemper, B. (1996) J. Biol. Chem. 271, 23725–23728. 31. Honkakoski, P., Moore, R., Gynther, J., and Negishi, M. (1996) J. Biol. Chem. 271, 9746 –9753. 32. Ramsden, R., Sommer, K. M., and Omiecinski, C. J. (1993) J. Biol. Chem. 268, 21722–21726. 33. Waxman, D. J., Morrissey, J. J., Naik, S., and Jauregui, H. O. (1990) Biochem. J. 271, 113–119. 34. Trottier, E., Belzil, A., Stoltz, C., and Anderson, A. (1995) Gene 158, 263–268. 35. Honkakoski, P., and Negishi, M. (1997) J. Biol. Chem. 272, 14943–14949. 36. Honkakoski, P., Moore, R., Washburn, K. A., and Negishi, M. (1998) Mol. Pharmacol. 53, 597– 601. 37. Forman, B. M., and Evans, R. M. (1995) Ann. N.Y. Acad. Sci. 761, 29 –37. 38. Stoltz, C., Vachon, M. H., Trottier, E., Dubois, S., Paquet, Y., and Anderson, A. (1998) J. Biol. Chem. 273, 8528 – 8536. 39. Liu, S., Park, Y., Rivera-Rivera, I., Li, H., and Kemper, B. (1998) DNA Cell Biol. 17, 461– 470. 40. Kemper, B. (1998) Prog. Nucleic Acid Res. Mol. Biol. 61, 25– 65. 41. Ramsden, R., Beck, N. B., Sommer, K. M., and Omiecinski, C. J. (1999) Gene 228, 169 –179. 42. Kim, J., and Kemper, B. (1997) J. Biol. Chem. 272, 29423– 29425. 43. Baes, M., Gulick, T., Choi, H. S., Martinoli, M. G., Simha, D., and Moore, D. D. (1994) Mol. Cell. Biol. 14, 1544 –1551. 44. Choi, H. S., Chung, M., Tzameli, I., Simha, D., Lee, Y. K., Seol, W., and Moore, D. D. (1997) J. Biol. Chem. 272, 23565–23571. 45. Tsai, M. J., and O’Malley, B. W. (1994) Annu. Rev. Biochem. 63, 451– 486. 46. DeFranco, D. B. (1999) Cell Biochem. Biophys. 30, 1–24. 47. Forman, B. M., Tzameli, I., Choi, H. S., Chen, J., Simha, D., Seol, W., Evans, R. M., and Moore, D. D. (1998) Nature 395, 612– 615. 48. Edwards, D. P. (1999) Vitam. Horm. 55, 165–218. 49. Kende, A. S., Ebetino, F. H., Drendel, W. B., Sundaralingam, M., Glover, E., and Poland, A. (1985) Mol. Pharmacol. 28, 445– 453. 50. Bicknell, D. C., and Gower, D. B. (1976) J. Steroid Biochem. 7, 451– 455. 51. Sidhu, J. S., and Omiecinski, C. J. (1997) J. Pharmacol. Exp. Ther. 282, 1122–1129. 52. Honkakoski, P., and Negishi, M. (1998) Biochem. J. 330, 889 – 895. 53. Weigel, N. L. (1996) Biochem. J. 319, 657– 667. 54. Ganem, L. G., Trottier, E., Anderson, A., and Jefcoate, C. R. (1999) Toxicol. Appl. Pharmacol. 155, 32– 42. 55. Larsen, M. C., Brake, P. B., Parmar, D., and Jefcoate, C. R. (1994) Arch. Biochem. Biophys. 315, 24 –34. 56. Shapiro, B. H., Pampori, N. A., Lapenson, D. P., and Waxman, D. J. (1994) Arch. Biochem. Biophys. 312, 234 –239. 57. Thummel, K. E., and Wilkinson, G. R. (1998) Annu. Rev. Pharmacol. Toxicol. 38, 389 – 430. 58. Schuetz, E. G., and Guzelian, P. S. (1984) J. Biol. Chem. 259, 2007–2012. 59. Wrighton, S. A., Schuetz, E. G., Watkins, P. B., Maurel, P., Barwick, J., Bailey, B. S., Hartle, H. T., Young, B., and Guzelian, P. (1985) Mol. Pharmacol. 28, 312–321. 60. Huss, J. M., and Kasper, C. B. (1998) J. Biol. Chem. 273, 16155–16162.

61. Quattrochi, L. C., Yockey, C. B., Barwick, J. L., and Guzelian, P. S. (1998) Arch. Biochem. Biophys. 349, 251–260. 62. Huss, J. M., Wang, S. I., Astrom, A., McQuiddy, P., and Kasper, C. B. (1996) Proc. Natl. Acad. Sci. USA 93, 4666 – 4670. 63. Barwick, J. L., Quattrochi, L. C., Mills, A. S., Potenza, C., Tukey, R. H., and Guzelian, P. S. (1996) Mol. Pharmacol. 50, 10 –16. 64. Quattrochi, L. C., Mills, A. S., Barwick, J. L., Yockey, C. B., and Guzelian, P. S. (1995) J. Biol. Chem. 270, 28917–28923. 65. Lehmann, J. M., McKee, D. D., Watson, M. A., Willson, T. M., Moore, J. T., and Kliewer, S. A. (1998) J. Clin. Invest. 102, 1016 –1023. 66. Schuetz, E. G., Brimer, C., and Schuetz, J. D. (1998) Mol. Pharmacol. 54, 1113–1117. 67. Waxman, D. J. (1996) J. Endocrinol. 150(Suppl.), S129 –S147. 68. Peters, J. M., Zhou, Y. C., Ram, P. A., Lee, S. S., Gonzalez, F. J., and Waxman, D. J. (1996) Mol. Pharmacol. 50, 67–74. 69. Blumberg, B., and Evans, R. M. (1998) Genes Dev. 12, 3149 – 3155. 70. Waxman, D. J., Attisano, C., Guengerich, F. P., and Lapenson, D. P. (1988) Arch. Biochem. Biophys. 263, 424 – 436. 71. Chang, T. K., Teixeira, J., Gil, G., and Waxman, D. J. (1993) Biochem. J. 291, 429 – 433. 72. Krey, G., Braissant, O., L’Horset, F., Kalkhoven, E., Perroud, M., Parker, M. G., and Wahli, W. (1997) Mol. Endocrinol. 11, 779 –791. 73. Schmiedlin-Ren, P., Thummel, K. E., Fisher, J. M., Paine, M. F., Lown, K. S., and Watkins, P. B. (1997) Mol. Pharmacol. 51, 741–754. 74. Rao, M. S., and Reddy, J. K. (1987) Carcinogenesis 8, 631– 636. 75. Lee, S. S., Pineau, T., Drago, J., Lee, E. J., Owens, J. W., Kroetz, D. L., Fernandez-Salguero, P. M., Westphal, H., and Gonzalez, F. J. (1995) Mol. Cell. Biol. 15, 3012–3022. 76. Palmer, C. N., Hsu, M. H., Muerhoff, A. S., Griffin, K. J., and Johnson, E. F. (1994) J. Biol. Chem. 269, 18083–18089. 77. Aldridge, T. C., Tugwood, J. D., and Green, S. (1995) Biochem. J. 306, 473– 479. 78. Forman, B. M., Chen, J., and Evans, R. M. (1997) Proc. Natl. Acad. Sci. USA 94, 4312– 4317. 79. Palmer, C. N., Hsu, M. H., Griffin, H. J., and Johnson, E. F. (1995) J. Biol. Chem. 270, 16114 –16121. 80. Juge-Aubry, C., Pernin, A., Favez, T., Burger, A. G., Wahli, W., Meier, C. A., and Desvergne, B. (1997) J. Biol. Chem. 272, 25252–25259. 81. DiRenzo, J., Soderstrom, M., Kurokawa, R., Ogliastro, M. H., Ricote, M., Ingrey, S., Horlein, A., Rosenfeld, M. G., and Glass, C. K. (1997) Mol. Cell. Biol. 17, 2166 –2176. 82. Peters, J. M., Cattley, R. C., and Gonzalez, F. J. (1997) Carcinogenesis 18, 2029 –2033. 83. Ward, J. M., Peters, J. M., Perella, C. M., and Gonzalez, F. J. (1998) Toxicol. Pathol. 26, 240 –246. 84. Roberts, R. A., James, N. H., Woodyatt, N. J., Macdonald, N., and Tugwood, J. D. (1998) Carcinogenesis 19, 43– 48. 85. Palmer, C. N., Hsu, M. H., Griffin, K. J., Raucy, J. L., and Johnson, E. F. (1998) Mol. Pharmacol. 53, 14 –22. 86. Keller, H., Devchand, P. R., Perroud, M., and Wahli, W. (1997) Biol. Chem. 378, 651– 655. 87. Uppenberg, J., Svensson, C., Jaki, M., Bertilsson, G., Jendeberg, L., and Berkenstam, A. (1998) J. Biol. Chem. 273, 31108 – 31112.

MINIREVIEW 88. Nolte, R. T., Wisely, G. B., Westin, S., Cobb, J. E., Lambert, M. H., Kurokawa, R., Rosenfeld, M. G., Willson, T. M., Glass, C. K., and Milburn, M. V. (1998) Nature 395, 137–143. 89. Xu, H. E., Lambert, M. H., Montana, V. G., Parks, D. J., Blanchard, S. G., Brown, P. J., Sternbach, D. D., Lehmann, J. M., Wisely, G. B., Willson, T. M., Kliewer, S. A., and Milburn, M. V. (1999) Mol. Cell 3, 397– 403. 90. Vidal-Puig, A. J., Considine, R. V., Jimenez-Linan, M., Werman, A., Pories, W. J., Caro, J. F., and Flier, J. S. (1997) J. Clin. Invest. 99, 2416 –2422. 91. Brun, R. P., Kim, J. B., Hu, E., and Spiegelman, B. M. (1997) Curr. Opin. Lipidol. 8, 212–218. 92. Bronfman, M., Ponce, C., Rojas, S., Roth, A., Loyola, G., Vollrath, V., and Chianale, J. (1998) Eur. J. Cell Biol. 77, 214 –219. 93. Tontonoz, P., Nagy, L., Alvarez, J. G., Thomazy, V. A., and Evans, R. M. (1998) Cell 93, 241–252. 94. Miyamoto, T., Kaneko, A., Kakizawa, T., Yajima, H., Kamijo, K., Sekine, R., Hiramatsu, K., Nishii, Y., Hashimoto, T., and Hashizume, K. (1997) J. Biol. Chem. 272, 7752–7758. 95. Miyata, K. S., McCaw, S. E., Patel, H. V., Rachubinski, R. A., and Capone, J. P. (1996) J. Biol. Chem. 271, 9189 –9192. 96. Jow, L., and Mukherjee, R. (1995) J. Biol. Chem. 270, 3836 – 3840. 97. Yan, Z. H., Karam, W. G., Staudinger, J. L., Medvedev, A., Ghanayem, B. I., and Jetten, A. M. (1998) J. Biol. Chem. 273, 10948 –10957. 98. Zhou, Y. C., and Waxman, D. J. (1999) J. Biol. Chem. 274, 2672–2681. 99. Juge-Aubry, C. E., Hammar, E., Siegrist-Kaiser, C., Pernin, A., Takeshita, A., Chin, W. W., Burger, A. G., and Meier, C. A. (1999) J. Biol. Chem. 274, 10505–10510. 100. Waxman, D. J., Ram, P. A., Park, S. H., and Choi, H. K. (1995) J. Biol. Chem. 270, 13262–13270. 101. Park, S. H., Liu, X., Hennighausen, L., Davey, H. W., and Waxman, D. J. (1999) J. Biol. Chem. 274, 7421–7430. 102. Corton, J. C., Fan, L. Q., Brown, S., Anderson, S. P., Bocos, C., Cattley, R. C., Mode, A., and Gustafsson, J. A. (1998) Mol. Pharmacol. 54, 463– 473. 103. Russell, D. W., and Setchell, K. D. (1992) Biochemistry 31, 4737– 4749. 104. Cooper, A. D., Chen, J., Botelho-Yetkinler, M. J., Cao, Y., Taniguchi, T., and Levy-Wilson, B. (1997) J. Biol. Chem. 272, 3444 –3452. 105. Lee, Y. H., Alberta, J. A., Gonzalez, F. J., and Waxman, D. J. (1994) J. Biol. Chem. 269, 14681–14689. 106. Crestani, M., Stroup, D., and Chiang, J. Y. (1995) J. Lipid Res. 36, 2419 –2432. 107. Crestani, M., Sadeghpour, A., Stroup, D., Galli, G., and Chiang, J. Y. (1998) J. Lipid Res. 39, 2192–2200.

23

108. Parks, D. J., Blanchard, S. G., Bledsoe, R. K., Chandra, G., Consler, T. G., Kliewer, S. A., Stimmel, J. B., Willson, T. M., Zavacki, A. M., Moore, D. D., and Lehmann, J. M. (1999) Science 21, 1365–1368. 109. Makishima, M., Okamoto, A. Y., Repa, J. J., Tu, H., Learned, R. M., Luk, A., Hull, M. V., Lustig, K. D., Mangelsdorf, D. J., and Shan, B. (1999) Science 284, 1362–1365. 110. Wang, H., Chen, J., Hollister, K., Sowers, L. C., and Forman, B. M. (1999) Mol. Cell 3, 543–553. 111. Diamond, M. I., Miner, J. N., Yoshinaga, S. K., and Yamamoto, K. R. (1990) Science 249, 1266 –1272. 112. Janowski, B. A., Willy, P. J., Devi, T. R., Falck, J. R., and Mangelsdorf, D. J. (1996) Nature 383, 728 –731. 113. Lehmann, J. M., Kliewer, S. A., Moore, L. B., Smith-Oliver, T. A., Oliver, B. B., Su, J. L., Sundseth, S. S., Winegar, D. A., Blanchard, D. E., Spencer, T. A., and Willson, T. M. (1997) J. Biol. Chem. 272, 3137–3140. 114. Peet, D. J., Turley, S. D., Ma, W., Janowski, B. A., Lobaccaro, J. M., Hammer, R. E., and Mangelsdorf, D. J. (1998) Cell 93, 693–704. 115. Russell, D. W. (1999) Cell 28, 539 –542. 116. Nitta, M., Ku, S., Brown, C., Okamoto, A. Y., and Shan, B. (1999) Proc. Natl. Acad. Sci. USA 8, 6660 – 6665. 117. Muntane-Relat, J., Ourlin, J. C., Domergue, J., and Maurel, P. (1995) Hepatology 22, 1143–1153. 118. Dohr, O., Sinning, R., Vogel, C., Munzel, P., and Abel, J. (1997) Mol. Pharmacol. 51, 703–710. 119. Clark, M. A., Williams, J. F., Gottschall, P. E., and Wecker, L. (1996) Biochem. Pharmacol. 51, 701–706. 120. Parmentier, J. H., Schohn, H., Bronner, M., Ferrari, L., Batt, A. M., Dauca, M., and Kremers, P. (1997) Biochem. Pharmacol. 54, 889 – 898. 121. Morgan, E. T. (1997) Drug Metab. Rev. 29, 1129 –1188. 122. Darnell, J. E., Jr. (1997) Science 277, 1630 –1635. 123. Davey, H. W., Wilkins, R. J., and Waxman, D. J. (1999) Am. J. Hum. Genet., in press. 124. McKay, L. I., and Cidlowski, J. A. (1998) Mol. Endocrinol. 12, 45–56. 125. Yanagisawa, J., Yanagi, Y., Masuhiro, Y., Suzawa, M., Watanabe, M., Kashiwagi, K., Toriyabe, T., Kawabata, M., Miyazono, K., and Kato, S. (1999) Science 283, 131–135. 126. Forman, B. M., Goode, E., Chen, J., Oro, A. E., Bradley, D. J., Perlmann, T., Noonan, D. J., Burka, L. T., McMorris, T., Lamph, W. W., et al. (1995) Cell 81, 687– 693. 127. Nebert, D. W. (1991) Mol. Endocrinol. 5, 1203–1214. 128. Chang, T. K., Yu, L., Maurel, P., and Waxman, D. J. Cancer Res. 57, 1946 –1954.