- Email: [email protected]
RETRACTED: Peroxisome Proliferator-Activated Receptor α C ontrols Hepatic Heme Biosynthesis Through ALAS1
J. Mol. Biol. (2009) 388, 225–238
doi:10.1016/j.jmb.2009.03.024
Available online at www.sciencedirect.com
Peroxisome Proliferator-Activated Receptor α Controls Hepatic Heme Biosynthesis Through ALAS1
Department of Biochemistry, University of Kuopio, FIN70211 Kuopio, Finland 2
Nutrigenomics Consortium, Top Institute of Food and Nutrition, Wageningen University, 6701 HD Wageningen, The Netherlands 3
Metabolism and Genomics Group, Division of Human Nutrition, Wageningen University, 6701 HD Wageningen, The Netherlands 4
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Life Sciences Research Unit, Université du Luxembourg, 162A, Avenue de la Faïencerie, L-1511 Luxembourg, Luxembourg
Heme is an essential prosthetic group of proteins involved in oxygen transport, energy metabolism and nitric oxide production. ALAS1 (5aminolevulinate synthase) is the rate-limiting enzyme in heme synthesis in the liver and is highly regulated to adapt to the metabolic demand of the hepatocyte. In the present study, we describe human hepatic ALAS1 as a new direct target for the nuclear receptor peroxisome proliferator-activated receptor α (PPARα). In primary human hepatocytes and in HepG2 cells, PPARα agonists induced an increase in ALAS1 mRNA levels, which was abolished by PPARα silencing. These effects are mediated by two functional PPAR binding sites at positions − 9 and −2.3 kb relative to the ALAS1 transcription start site. PPARα ligand treatment also up-regulated the mRNA levels of the genes ALAD (5-aminolevulinate dehydratase), UROS (uroporphyrinogen III synthase), UROD (uroporphyrinogen decarboxylase), CPOX (coproporphyrinogen oxidase) and PPOX (protoporphyrinogen oxidase) encoding for enzymes controlling further steps in heme biosynthesis. In HepG2 cells treated with PPARα agonists and in mouse liver upon fasting, the association of PPARα, its partner retinoid X receptor, PPARγ coactivator 1α and activated RNA polymerase II with the transcription start site region of all six genes was increased, leading to higher levels of the metabolite heme. In conclusion, these data strongly support a role of PPARα in the regulation of human ALAS1 and of five additional genes of the pathway, consequently leading to increased heme synthesis.
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Tatjana Degenhardt 1 , Sami Väisänen 1 , Maryam Rakhshandehroo 2,3 , Sander Kersten 2,3 and Carsten Carlberg 1,4 ⁎
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Received 14 November 2008; received in revised form 5 March 2009; accepted 9 March 2009 Available online 14 March 2009 Edited by J. Karn
© 2009 Elsevier Ltd. All rights reserved.
Keywords: nuclear receptor; transcriptional regulation; chromatin; metabolism; PPARα
Introduction
Heme is an irreplaceable compound for mammalian life. It is an essential component of numerous heme-containing proteins that exhibit functions in mitochondrial respiration, hormone synthesis and
metabolism and nitric oxide synthesis.1–3 Heme biosynthesis in eukaryotic cells is composed of eight enzymatic steps, of which the first and the last three steps take place in the mitochondria, whereas the others occur in the cytoplasm. Eighty to ninety percent of the total heme in mammals is synthesized
*Corresponding author. E-mail address: [email protected]. Abbreviations used: ALAD, 5-aminolevulinate dehydratase; ALAS, 5-aminolevulinate synthase; BSA, bovine serum albumin; ChIP, chromatin immunoprecipitation; CPOX, coproporphyrinogen oxidase; CYP, cytochrome P450; DMEM, Dulbecco's modified Eagle's medium; DMSO, dimethyl sulfoxide; DR1, direct repeat spaced by one nucleotide; FBS, fetal bovine serum; FECH, ferrochelatase; GW7647, 2-(4-(2-(1-cyclohexanebutyl-3-cyclohexylureido)ethyl)phenylthio)-2methylpropionic acid; HMBS, hydroxymethylbilane synthase; PBS, phosphate-buffered saline; PGC-1α, PPARγ coactivator 1α; PPAR, peroxisome proliferator-activated receptor; pPol II, phosphorylated RNA polymerase II; PPOX, protoporphyrinogen oxidase; PPRE, PPAR response element; RE, response element; RPLP0, acidic riboprotein P0; RXR, retinoid X receptor; TSS, transcription start site; UROD, uroporphyrinogen decarboxylase; UROS, uroporphyrinogen III synthase; Wy14,643, pirinixic acid. 0022-2836/$ - see front matter © 2009 Elsevier Ltd. All rights reserved.
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number of direct inducible PPAR target genes have been identified;20,21 mostly, these genes represent different metabolic pathways. In humans, little knowledge is available regarding the regulation of whole pathways by PPARα.22 Usually, only a gene family or one single gene is examined, while whole pathways under the control of PPARs have not yet been intensively studied. An essential prerequisite for the direct modulation of transcription by PPAR ligands is the location of at least one activated PPAR protein close to the transcription start site (TSS) of the target gene. This is commonly achieved through the binding of PPARs to a specific sequence of double-stranded DNA, called a PPAR response element (PPRE), often located thousands of base pairs up- or downstream of a gene's TSS. A subsequent DNA-looping event links the activated PPAR–PPRE complex to the TSS.23 PPARs bind to DNA as heterodimers with the nuclear receptor retinoid X receptor (RXR).24 PPREs are formed by two hexameric motifs with the optimal AGGTCA core binding sequence in a direct repeat orientation with a spacing of one nucleotide (DR1), where PPAR occupies the 5′ motif.25 Binding of agonists to the PPARs causes a conformational change within their ligand-binding domain that results in an enhanced binding of co-activator proteins, such as PGC-1α.26,27 These co-activators link ligand-activated PPARs to enzymes displaying histone acetyltransferase activity that cause chromatin relaxation. In a subsequent step, ligandactivated PPARs rapidly exchange co-activator proteins for components of mediator complexes,28 which act as a bridge from the activated PPARs to the basal transcriptional machinery. In the present study, we investigated the role of PPARα as a direct regulator of ALAS1 and the whole heme biosynthesis pathway in primary hepatocytes and HepG2 cells. We identified that the genes ALAS1, ALAD (5-aminolevulinate dehydratase), UROS (uroporphyrinogen III synthase), UROD (uroporphyrinogen decarboxylase), CPOX (coproporphyrinogen oxidase) and PPOX (protoporphyrinogen oxidase) are direct PPARα targets, thereby connecting PPARα signaling and hepatic heme biosynthesis.
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in erythroid cells and is subsequently incorporated into hemoglobin. Regulation of heme biosynthesis in these cells involves the enzyme encoded by the erythroid-specific ALAS2 (5-aminolevulinate synthase 2) gene. In contrast, the housekeeping gene ALAS, also called ALAS1 or ALAS-N, is ubiquitously expressed, as all nucleated cells need to synthesize heme for their cytochrome P450 (CYP) enzymes. Most of the heme that is synthesized outside of erythroid cells is produced in the liver for various heme-containing enzymes, in particular for microsomal CYPs. In mitochondrial biogenesis, coordinated increase of heme synthesis and the synthesis of respiratory CYPs are prerequisites for a functional energy metabolism.4 Hereditary partial defects of enzymes involved in heme biosynthesis lead to rare metabolic diseases known as porphyrias. Inducible, acute hepatic porphyrias are characterized by attacks of neuropsychiatric dysfunction, precipitated by stimuli, such as fasting, alcohol, drugs and sex steroids.5,6 The detrimental effects of heme to the cell make a tight control of its biosynthesis essential. In non-erythroid cells, the rate of heme synthesis is controlled at its first enzymatic step. Accordingly, the ALAS1 gene is highly regulated in different cellular contexts to ensure adequate levels of heme.7–9 In recent years, progress has been made in understanding the regulation of ALAS1 in liver on a molecular level. Members of the nuclear receptor superfamily, such as the xenosensors pregnane X receptor and constitutive androstane receptor and the bile acidactivated farnesoid X receptor,10–13 have been shown to play a role in the regulation of ALAS1. Furthermore, the peroxisome proliferator-activated receptor (PPAR) γ co-activator 1α (PGC-1α), which is a key regulator in mitochondrial biogenesis and energy homeostasis, was shown to be important for a fasting response of ALAS1 by acting through the insulin-regulated transcription factor Forkhead O1A binding to the ALAS1 promoter,14 whereas the other genes of the pathway were not subject to studies. Nuclear receptors are transcription factors that have important roles in controlling cellular metabolism because many of them are activated by lipophilic ligands, including cholesterol, fatty acids and their metabolic derivatives.15 The three members of the PPAR subfamily, PPARα, PPARγ and PPARβ/δ, are thought to play a prominent role in the development of the metabolic syndrome since they are key regulators of lipid storage and catabolism.16 PPARα has been studied most intensively in the context of liver metabolism and is known to control hepatic fatty acid catabolism.17 PPARγ, which is highly expressed in adipose tissue, is a master regulator of adipogenesis. The widely expressed PPARβ/δ stimulates fatty acid oxidation, regulates hepatic very-low-density lipoprotein production and catabolism18 and is involved in more diverse actions, such as wound healing by governing keratinocyte differentiation.19 All three receptors are activated by (mainly polyunsaturated) fatty acids and various fatty acid derivatives, such as eicosanoids. In rodents, a large
Results PPARα-dependent regulation of the genes involved in hepatic heme biosynthesis To explore the regulation of the heme biosynthetic genes by PPARα, we used human primary hepatocytes from six donors that were treated for 6 and 24 h with 50 μM concentration of the PPARα-specific agonist Wy14,643 (pirinixic acid; Fig. 1a). RNA was extracted, and expression changes were monitored by quantitative real-time PCR. The genes ALAS1 (2.8-fold after 6 h, 1.5-fold after 24 h), ALAD (1.3-fold after 6 and 24 h), UROS (1.3-fold after 6 h, 1.2-fold after 24 h), CPOX (1.2-fold after 6 and 24 h) and PPOX (1.2-fold after 6 and 24 h) were significantly
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Fig. 1. Expression profiling of the eight genes involved in hepatic heme biosynthesis. (a) Human primary hepatocytes from six donors were stimulated with 50 μM concentration of the PPARα ligand Wy14,643. (b, c) HepG2 cells were stimulated with 100 nM concentration of the PPARα ligand GW7647 for 6 and 24 h. (d) HepG2 cells were transfected with either control siRNA or siRNA specific for PPARα and incubated for 48 h. RNA was extracted and quantitative real-time PCR was performed for the eight heme biosynthesis genes. Columns indicate the means of at least three independent treatments, and the bars represent standard deviations. Two-tailed, paired Student's t-tests were performed to determine the significance of the mRNA induction by PPARα agonist relative to solvent controls (a–c) and that of PPARα siRNA knockdown relative to control siRNA (d) (⁎p b 0.05, ⁎⁎p b 0.01, ⁎⁎⁎p b 0.001).
up-regulated, whereas no regulation was observed for HMBS (hydroxymethylbilane synthase), UROD and FECH (ferrochelatase).
Next, we asked the question of whether some of the eight genes of the hepatic heme biosynthesis are also PPARα targets in the human hepatocellular
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Fig. 2. Increase in heme biosynthesis. The hepatic heme biosynthesis pathway is illustrated. Enzymes that are encoded by PPARα target genes are depicted in red, whereas non-regulated genes are illustrated in blue (a). HepG2 cells were treated for 6, 8 and 24 h with GW7647 or DMSO. The cells were lysed, and heme concentration was determined and normalized to the total protein content (b). Columns indicate the means of at least three independent treatments, and the bars represent standard deviations. Two-tailed, paired Student's t-tests were performed to determine the significance of the induction of heme concentration by PPARα agonist relative to solvent controls (⁎p b 0.05).
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carcinoma cell line HepG2. For this purpose, HepG2 cells were treated for 6 or 24 h with 100 nM concentration of the PPARα ligand GW7647 [2-(4-(2(1-cyclohexanebutyl-3-cyclohexylureido)ethyl)phenylthio)-2-methylpropionic acid], RNA was extracted and expression changes were monitored using quantitative real-time PCR (Fig. 1b). The results confirmed the findings in primary hepatocytes (Fig. 1a); again, HMBS and FECH did not show any response at any time point, whereas ALAS1 (1.7fold after 6 h, 2.1-fold after 24 h), ALAD (1.2-fold after 6 h), UROS (1.5-fold after 6 h, 1.4-fold after 24 h), UROD (1.4-fold after 6 and 24 h), CPOX (1.5fold after 6 h, 1.3-fold after 24 h) and PPOX (1.8-fold after 6 h, 1.5-fold after 24 h) were significantly upregulated. In order to exclude the possibility that there may be specific effects of the two PPARα ligands, we also treated HepG2 cells with 50 μM Wy14,643 and observed a significant induction of the same six genes of the heme pathway (Fig. S1a). The lowest expressed genes, HMBS and FECH (Fig. S1b), did not respond to any PPAR agonist. To assess possible PPARα-independent ligand effects, we silenced PPARα in HepG2 cells with PPARα-specific siRNA oligonucleotides (Fig. 1c and d). After silencing PPARα, mRNA levels were 75% reduced, whereas the levels of PPARγ (10% reduction) and PPARβ/δ (35% reduction) were only slightly affected (Fig. S2a). On protein level, PPARα was reduced after siRNA treatment by about 50%, whereas PPARγ and PPARβ/δ were not affected (Fig. S2b). In the PPARα-silenced cells, we could not detect any changes of gene expression after 6 or 24 h of treatment with GW7647 (Fig. 1c), indicating that the observed effects are clearly linked to PPARαdependent gene regulation. Moreover, after PPARα silencing, the basal expression levels of ALAS1,
ALAD, UROS, UROD, CPOX and PPOX also were reduced by 60%, 37%, 40%, 20%, 33% and 36%, respectively (Fig. 1d), indicating an important role of PPARα in the regulation of these genes. In contrast, the genes HMBS and FECH were not significantly affected in their basal expression by silencing PPARα. Taken together, quantitative real-time PCR analysis of the hepatic heme biosynthesis pathway in primary human hepatocytes and in HepG2 cells suggested that the genes ALAS1, ALAD, UROS, UROD, CPOX and PPOX are likely to be direct PPARα targets. Increase in heme concentration An overview on the hepatic heme biosynthesis pathway is shown in Fig. 2a. To gain insight on whether the PPARα-dependent regulation of six of the eight genes leads in turn to an activation of the whole pathway, resulting into an increased production of heme, we treated HepG2 cells with PPARα agonist GW7647 or the solvent dimethyl sulfoxide (DMSO) and measured the heme concentration of the cells after ligand treatment. Intriguingly, already after 8 h we could detect a significant 1.5-fold increase in heme concentration that after prolonged treatment with ligand (1.6-fold after 24 h) still increased (Fig. 2b). In summary, the PPARα-dependent transcriptional control leads to an increase in heme levels in HepG2 cells. Conservation of PPREs in the heme biosynthetic genes Recently, we developed an in silico screening method for a reliable prediction of DR1-type
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Functional PPREs in the ALAS1 gene
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The rate-limiting enzyme ALAS1 displayed the strongest response to PPARα ligands and based on our in silico screening the ALAS1 gene carries seven putative PPREs in a distance of less than 10 kb from the TSS, of which three (RE2, RE4 and RE6) are conserved compared with the orthologous sequence in mouse (Fig. 3). In order to test the affinity of these REs for PPARs, we performed gel-shift assays with in vitro-translated PPARα–RXR heterodimers and the seven candidate PPREs of the ALAS1 gene (Fig. 5a). Compared with the established PPRE of the CPTI gene,31 PPARα–RXR heterodimers bound RE3 with 21.1% strength, RE1 with 10.2%, RE6 with 8.6% and RE2 with 3.3%, while they bound RE4, RE5 and RE7 with 1.3% or less. The functionality of the REs was further investigated using ChIP assays in HepG2 cells (Fig. 5b). Please note that RE4 and RE5 are so close to each other that they could not be distinguished. The TSS of the ALAS1 gene served as a positive control. Significant binding of PPARα was found in regions carrying RE1, RE2, RE4/5 and RE6, that of RXRα was found in regions of RE2, RE6 and RE7, that of PGC-1α was found in regions of RE1, RE2, RE4/5 and RE6 and that of pPol II was found in regions of RE1, RE4/5, RE6 and RE7. These association patterns on chromatin region show no direct correlation with the affinity of the PPREs that they carry, which suggests that in vitro binding affinity may be not the only criterion for the functionality of a PPAR-responsive region. However, the regions of the conserved RE2 and RE6, as well as the region of RE1, were found to be functional. In contrast, the highest affinity putative PPRE, RE3, appears to be not accessible to PPARα binding. Finally, we tested the functionality of the identified REs by reporter gene assays in transiently transfected HepG2 cells (Fig. 5c). The genomic regions investigated in ChIP assays (Fig. 5b) were fused with the thymidine kinase promoter driving the luciferase reporter gene. The genomic region of the human CPTI gene carrying the established PPRE served as a positive control. PPARα overexpression resulted for all genomic regions in increases of their basal activity, which were 2.0-fold even for the empty reporter gene vector, 5.3-fold for the region of the CPTI gene, 4.8-fold for the genomic region around ALAS1 RE1, 5.2-fold for RE2, 2.4-fold for RE3, 19.6-fold for RE4/5, 18.1-fold for RE6 and 15.2fold for RE7. However, most informative were the statistically significant increases after stimulation with GW7647, which were 2.1-fold for the CPTI region and 1.7-, 1.6- and 1.2-fold for regions 1, 2 and 7 of the ALAS1 gene, respectively. The three other ALAS1 regions did not show any significant ligand induction. Taken together, our in vitro analysis of the seven candidate PPREs of the ALAS1 gene indicated for four a reasonable affinity for PPARα–RXR, three of which (RE1, RE2 and RE6) could also be confirmed by ChIP analysis in HepG2 cells. Three of RE-
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PPREs.29,30 We now applied this method to the genomic regions 10 kb up- and downstream of the TSSs of the eight human heme biosynthesis genes (Fig. 3) and their mouse orthologs (data not shown). Within the 20 kb of analyzed sequence, each of the genes displayed between none (UROS) and seven (ALAS and PPOX) putative PPREs. A comparison with the respective mouse orthologs indicated that the eight genes contain 28 PPREs in humans and 23 PPREs in the mouse, of which 17 (74% and 61%, respectively) are conserved between both species. This suggests a high degree of conservation in the regulation of this pathway. The PPOX gene showed six conserved PPREs, the ALAS1 and UROD genes showed three, the ALAD gene showed two, the HMBS, CPOX and FECH genes showed one and only the UROS gene showed no conserved PPRE. This means that ALAS1, ALAD, UROD and PPOX are more likely PPAR target genes than HMBS, UROS, CPOX and FECH. Taken together, most of the genes responding to PPARα ligands and PPARα siRNA knockdown show a significantly increased number of potential PPAR binding sites in the genomic sequence 10 kb up- or downstream of their TSSs, compared with the unresponsive genes HMBS and FECH.
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Recruitment of PPARα to the TSS regions of the heme biosynthesis genes
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For an experimental validation of the prediction of PPAR binding to the regulatory regions of heme biosynthetic genes, we tested the binding of PPARα to the TSS for each of the eight genes using chromatin immunoprecipitation (ChIP) assays. In addition, binding of its heterodimeric partner RXRα, the common co-activator PGC-1α and activated RNA polymerase [phosphorylated RNA polymerase II (pPol II)] was assessed. Only active PPREs will loop together with their associated proteins to the TSS hosting Pol II, connecting the activated PPRE with the basal transcription machinery. Therefore, the association of the TSS with PPARα, RXRα and PGC-1α indicates that at least one of the putative PPREs of the respective genes was active. We treated HepG2 cells for 240 and 360 min with GW7647 and cross-linked the proteins to DNA using formaldehyde (Fig. 4). Quantification of ChIP assays by realtime PCR showed that PPARα enriched at both time points on the TSS regions of ALAS1, ALAD, UROS, UROD, CPOX and PPOX (Fig. 4a). In contrast, on the TSSs of the genes HMBS and FECH, no significant enrichment of PPARα binding was observed. RXRα (Fig. 4b), PGC-1α (Fig. 4c) and pPol II (Fig. 4d) essentially showed the same binding pattern—i.e., they were found on the TSSs of ALAS1, ALAD, UROS, UROD, CPOX and PPOX but not on those of HMBS and FECH. In summary, in HepG2 cells, PPARα, RXRα, PGC1α and pPol II associate with the TSS regions of the genes ALAS1, ALAD, UROS, UROD, CPOX and PPOX but not with those of the genes HMBS and FECH.
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Fig. 3 (legend on next page)
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Fasting-induced changes in PPARα TSS binding and heme concentration
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For experiments in mice, we fed or fasted wildtype mice and PPARα knockout mice for 24 h (three mice per group), removed their livers and immediately cross-linked the nuclear proteins to the genomic DNA (Fig. 6). Quantification of ChIP assays by real-time PCR showed that PPARα associated with the TSS regions of Alas1, Alad, Uros, Urod, Cpox and Ppox when PPARα wild-type mice were fasted (Fig. 6a). This shows that PPARαmediated transcription is stronger than the basal transcription of the heme biosynthesis pathway genes. However, the TSSs of Hmbs and Fech showed no significant enrichment of the receptor. The same binding pattern that was observed for RXRα (Fig. 6b), PGC-1α (Fig. 6c) and pPol II (Fig. 6d) binding could confirm these associations with the exception of the TSS of the Urod gene. To gain insight on whether the PPARα-dependent regulation of the heme biosynthetic genes leads in turn to an activation of the whole pathway, resulting into an increased production of heme, we measured heme concentration in livers from wild-type and PPARα null mice that were fed or fasted for 24 h. Small liver pieces were homogenized and heme concentration was measured. We observed a significant 2.1-fold increase in heme levels in the fasted wild-type mice compared with the fed wild-type mice (Fig. 6e). Although we found slightly higher heme levels in the fasted PPARα null mice compared with the fed null mice, we were able to observe a significant decrease in heme levels in the fasted PPARα null mice compared with the fasted wildtype mice. In summary, in mouse liver PPARα, RXRα, PGC1α and pPol II associate with the TSS regions of Alas1, Alad, Uros, Urod, Cpox and Ppox but not with those of Hmbs and Fech. Moreover, the heme levels in livers of fasted wild-type mice increase compared with fed mice and are also higher than those in fasted PPARα null mice.
the first enzyme in heme biosynthesis, ALAS1, by fasting through the co-activator PGC-1α.14 In addition, several other nuclear receptors, such as pregnane X receptor, constitutive androstane receptor and farnesoid X receptor, were suggested to regulate ALAS1 expression.10–13 In this study, we demonstrated that human hepatic ALAS1, as well as five others out of the additional seven genes of the hepatic heme biosynthesis pathway, is a novel direct target for PPARα. By quantitative real-time PCR using primary human hepatocytes and HepG2 cells, siRNA knockdown in HepG2 cells, in silico screening and ChIP assays in HepG2 cells and in mouse liver, we consistently demonstrated that the genes ALAS1, ALAD, UROS, UROD, CPOX and PPOX are PPARα target genes. In contrast, the genes HMBS and FECH carry only one conserved PPRE, which is apparently insufficient, since these two genes showed no sign of regulation by PPARα. Moreover, we showed that PPARα agonist treatment increases heme concentration in HepG2 cells and in fasted wild-type mice. The response to PPARα agonists was most prominent for the ALAS1 gene. We found seven putative PPREs within 10 kb of the TSS of this gene. Two of these PPREs (RE1 and RE2 at positions − 9 and − 2.3 kb relative to the ALAS1 TSS, respectively) showed significant in vitro binding for PPARα–RXR heterodimers as well as association of PPARα and PGC-1α to their chromatin regions and mediated ligand inducibility to their genomic regions in reporter gene assays. The other five putative sites show inconsistent effects and may not be functional. Many studies focus on the regulation of ALAS1 since it catalyzes the rate-limiting step in hepatic heme biosynthesis.9 The accumulation of the intermediates of the heme biosynthesis pathway is the biochemical hallmark of porphyria. It is known that different factors, including fasting, alcohol and drugs, can trigger porphyric attacks in individuals who carry a mutation in one of the enzymes of the heme pathway.5,6 There is high expression not only of ALAS1 but also of ALAD, UROS, UROD, CPOX and PPOX in order to avoid a too high (i.e., toxic) metabolite concentration in the cell upon fasting. Under normal physiological conditions, free heme levels are extremely low and tightly regulated because cellular toxicity can occur with increased cellular concentrations of unincorporated heme. In addition, it is known that treatment with phenobarbital or other prototypical drug inducers elevates heme concentrations in the liver to accommodate the increased levels of heme-dependent enzymes.33–35 In this study, we showed that fasting up-regulates heme concentrations in mouse liver in a PPARα-dependent way. Moreover, other studies demonstrated that this
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containing genomic regions (RE1, RE2 and RE7) were found to be ligand inducible in reporter gene assays. In conclusion, only RE1 and RE2 seem to be fully functional PPREs.
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Discussion
PPARα is emerging as one of the major components regulating liver metabolism. The role of PPARα in the adaptive response to fasting is well established.32 Previous studies showed regulation of
Fig. 3. In silico screening for PPREs in the heme biosynthesis genes in human. Overview of the eight genes of the heme biosynthesis pathway. The diagrams show 10 kb upstream and 10 kb downstream of the TSS (arrow), regions of repetitive sequence (white boxes) and exons (dark gray boxes). Putative REs (red boxes, conserved; blue boxes, non-conserved) were identified by in silico screening of the genomic sequences and are classified according to their degree of conservation between human and mouse.
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Fig. 4. Ligand-induced PPARα recruitment to the TSS of heme biosynthesis genes in HepG2 cells. Chromatin was extracted from HepG2 cells that had been treated with 100 nM GW7647 for 0, 240 and 360 min, and IPs were performed with antibodies against PPARα (a), RXRα (b), PGC-1α (c) and pPol II (d). Rabbit IgG was used as a specificity control. Real-time PCR was performed on reverse-cross-linked chromatin templates with primers specific to genomic regions of the eight human hepatic heme biosynthesis genes. Columns indicate the means of at least three independent treatments, and the bars represent standard deviations. Two-tailed, paired Student's t-tests were performed to determine the significance of the specific antibody enrichment by PPAR agonist relative to control IgG (⁎p b 0.05, ⁎⁎p b 0.01, ⁎⁎⁎p b 0.001).
is also the case for several heme-containing proteins.32 Both observations link the up-regulation of the heme biosynthesis upon fasting and an increase in heme concentration to the need of more heme for heme-containing enzymes, such as Cyp4a10, Cyp4a12 and Cyp4a1.32 This suggests that the regulation of the heme biosynthetic pathway
upon fasting is necessary to keep up with the demand of heme for heme-containing proteins involved in processes, such as fatty acid metabolism. PPARα is one of the most prominent regulators of fatty acid metabolism in liver, showing a clear need for the up-regulation of hepatic heme biosynthesis and to enable a functioning fatty acid metabolism.
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Cell culture experiments
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Materials and Methods
temperature (Table S2) and 40 s at 72 °C. Most of the primer sequences were obtained from the PrimerBank at Harvard University.36 The sequences of the gene-specific primer pairs for the human genes involved in heme biosynthesis and the internal control gene RPLP0 (acidic riboprotein P0) are listed in Table S2. PCR product quality was monitored using post-PCR melt curve analysis. Fold inductions were calculated using the formula 2−(ΔΔCt), where ΔΔCt is the ΔCt(PPARα ligand) − ΔCt(DMSO), ΔCt is Ct(gene X) − Ct(RPLP0) and Ct is the cycle at which the threshold is crossed.
HepG2 cells were grown overnight in medium containing charcoal-stripped FBS to 30% confluency. Cells were transfected using Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer's instructions. A total amount of 200 pmol of either control siRNA or a mixture of the three PPARα-specific siRNAs (Eurogentec, Liege, Belgium; for sequences, see Table S3) was transfected. Cells were incubated for 8 h, and then charcoal-stripped FBS was added and the transfection was continued for 48 h in total. Cell treatments and RNA extractions were carried out as described above.
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Taken together, the regulation of the heme biosynthesis pathway by PPARα offers an important insight in the mechanism—how the demand for more hemecontaining proteins upon fasting triggers the transcriptional production of genes involved in the heme biosynthesis pathway. In conclusion, our data demonstrate that the hepatic heme biosynthesis pathway in humans is under direct transcriptional control of PPARα, consequently leading to an increase in heme concentration. The six out of eight genes of the pathway are novel primary PPARα target genes. These also suggest a mechanism by which fasting or fibrate treatment may trigger acute porphyric attacks in susceptible patients.
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The human hepatocellular carcinoma cell line HepG2 was cultured in RPMI 1640 containing 10% fetal bovine serum (FBS), 2 mM L-glutamine, 0.1 mg/ml of streptomycin and 100 U/ml of penicillin in a humidified 95% air/ 5% CO2 incubator. Before use, FBS was stripped of lipophilic compounds, such as endogenous nuclear receptor ligands, by stirring it with 5% activated charcoal (Sigma-Aldrich) for 3 h at room temperature. Charcoal was then removed by centrifugation and sterile filtration. Prior to mRNA, chromatin extraction or transient transfections, cells were grown overnight in phenol red-free Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% charcoal-stripped FBS to reach a density of 50% to 60% confluency. Cells were then treated with either solvent (DMSO, final concentration = 0.1%) or 100 nM concentration of the PPARα agonist GW7647 (Alexis Biochemicals, San Diego, CA). Primary human hepatocytes and Hepatocyte Culture Medium BulletKit were purchased from Lonza Bioscience (Verviers, Belgium). Human hepatocytes were isolated from six donors (Table S1). Cells were plated on collagen-coated six-well plates. Upon arrival of the cells, the medium was discarded and was replaced by Hepatocyte Culture Medium. The next day, cells were incubated in fresh medium in the presence or absence of 50 μM Wy14,643 (ChemSyn Laboratories, Lenexa, KS) dissolved in DMSO for 6 or 24 h, followed by RNA isolation. RNA extraction and real-time quantitative PCR
Total RNA from HepG2 cells was extracted using a Mini RNA Isolation II kit (Zymo Research, HiSS Diagnostics, Freiburg, Germany), and cDNA synthesis was performed for 1 h at 37 °C using 1 μg of total RNA as a template, 100 pmol oligodT18 primer and 40 U of reverse transcriptase (Fermentas, Vilnius, Lithuania). For human primary hepatocytes, RNA was extracted using TRIzol reagent (Invitrogen) and 1 μg of RNA was reverse transcribed using iScript (BioRad, Hercules, CA). Realtime quantitative PCR was performed in an IQ-cycler (BioRad) using the dye SybrGreen I (Molecular Probes, Leiden, The Netherlands). Per reaction, 1 U of Hot Start Taq polymerase (Fermentas) and 3 mM MgCl2 were used and the PCR cycling conditions were as follows: 45 cycles of 30 s at 95 °C, 30 s at primer-specific annealing
Measurement of heme concentration
PPARα wild-type and null mice (n = 4 per group) were fed or fasted for 24 h, their livers were removed and frozen in liquid nitrogen. Small pieces of liver were homogenized in Reagent buffer, and heme concentration was measured using a QuantiChrom™ Heme Assay Kit at 414 nm (BioAssay Systems, Hayward, CA). HepG2 cells were grown overnight in phenol red-free DMEM supplemented with 10% charcoal-stripped FBS and treated for 0, 6, 8 and 24 h with 100 nM GW7647. Cells were washed with phosphate-buffered saline (PBS). The heme concentration was measured according to the manufacturer's instructions. The data were normalized to the total protein content of the sample using the BCA (bicinchoninic acid) protein assay (Pierce Biotechnology, Rockford, IL). In silico PPRE screening Genomic sequences spanning ± 10 kb around the TSSs of the human and mouse genes were extracted from the current database release for the human genome and the mouse genome. Putative PPREs were screened from the sequence files as described previously.29,30 Conservation of putative PPREs between human and mouse was checked using the program Align (MacMolly® Tetra package, Soft Gene GmbH, Bocholt, Germany). Repetitive sequences from both species were obtained using CENSOR.37 ChIP assays in HepG2 Nuclear proteins were cross-linked to genomic DNA by adding formaldehyde for 5 min directly to the medium to a final concentration of 1% at room temperature. Crosslinking was stopped by adding glycine to a final concentration of 0.125 M and incubating for 5 min at room temperature on a rocking platform. The medium was removed and the cells were washed twice with icecold PBS. The cells were collected by scraping into ice-cold PBS supplemented with a protease inhibitor cocktail
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centrifugation and the lysates were diluted 1:10 in ChIP wash buffer 1 (150 mM NaCl, 1% Triton X-100, 2 mM EDTA, 50 mM Tris–HCl, pH 8.1). The samples were centrifuged and the recovered chromatin solutions were incubated with 5 μl of indicated antibodies, 2.4 μl of sonicated salmon sperm (10 mg/ml) and 25 μl of bovine
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(Roche Diagnostics, Mannheim, Germany). After centrifugation, the cell pellets were resuspended in lysis buffer [1% SDS, 10 mM ethylenediaminetetraacetic acid (EDTA), protease inhibitors, 50 mM Tris–HCl, pH 8.1] and the lysates were sonicated to result in DNA fragments of 300 to 1000 bp in length. Cellular debris was removed by
Fig. 5 (legend on next page)
PPARα Controls Heme Biosynthesis
ChIP assays in mouse liver
inhibitors (Roche) for 10 min at room temperature. Lysates were sonicated to result in DNA fragments of 300 to 1000 bp in length. Cellular debris was removed by centrifugation and the lysates were diluted 1:10 in ChIP wash buffer 1. The recovered chromatin solutions were incubated with 5 μl of indicated antibodies (200 μg/ml), 2.4 μl of sonicated salmon sperm (10 mg/ml) and 25 μl BSA (10 mg/ml) to remove unspecific background overnight at 4 °C with rotation. The clean-up of the samples and recovery of DNA were performed as described above. PCR of chromatin templates
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For the genomic regions containing the TSSs of all eight human and mouse genes involved in heme biosynthesis and the putative REs of the ALAS1 gene, specific primer pairs were designed (Table S4), optimized and controlled by running PCRs with 25 ng of genomic DNA (input) as a template. When running immunoprecipitated DNA (output) as a template, the following PCR profile was used: pre-incubation for 5 min at 94 °C, 45 cycles of 30 s at 95 °C, 30 s at the temperature indicated for each primer pair (Table S4) and 30 s at 72 °C and one final incubation for 10 min at 72 °C. The primers were controlled using melt curve analysis and gel pictures to allow the detection of primer-specific products. The fold change relative to the non-specific IgG background was calculated. The fold inductions were calculated using the formula 2−(ΔCt), where ΔCt is Ct(specific antibody) − Ct(IgG control) and Ct is the cycle where the threshold is crossed.
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serum albumin (BSA) (10 mg/ml) to remove unspecific background overnight at 4 °C with rotation. The antibodies against PPARα (sc-9000), RXRα (sc-553), PGC-1α (sc-13067), pPol II (sc-13583) and control immunoglobulin G's (IgGs) (sc-2027) were obtained from Santa Cruz Biotechnologies. The immunocomplexes were collected with 25 μl of MagnaCell magnetic protein A agarose beads (Cortex Biochem, Madison, WI) for 1 h at room temperature with rotation. The magnetic beads were pre-blocked overnight in a solution containing ChIP wash buffer 1, protease inhibitors, BSA and sonicated salmon sperm in the same concentrations as used later in the immunocollection. Magnetic beads were collected using a magnetic rack (Qiagen) and were washed twice for 1 min with ChIP wash buffer 1 containing 1 mM PMSF, 5 min with ChIP wash buffer 2 (500 mM NaCl, 1% Triton X-100, 2 mM EDTA, 0.1% SDS, 20 mM Tris–HCl, pH 8.0, containing 1 mM freshly added PMSF), 5 min with ChIP wash buffer 3 (0.25 mM LiCl, 1% Nonidet P-40, 1% sodium deoxycholate, 1 mM EDTA, 10 mM Tris–HCl, pH 8.1) and finally twice for 1 min with TE buffer (1 mM EDTA, 10 mM Tris– HCl, pH 8.0). Then, 250 μl of elution buffer (10 mM EDTA, 0.5% SDS, 25 mM Tris–HCl, pH 7.5) was added and incubated for 30 min at 64 °C. The supernatant was removed and the beads were washed for 2 min with 250 μl of elution buffer. The supernatants were combined, and proteins were digested overnight at 64 °C using proteinase K (Fermentas, final concentration = 40 μg/ml). Genomic DNA fragments were recovered by phenol–chloroform extraction and a following ethanol precipitation.
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Mice were fed or fasted for 24 h (three mice per group). The mice were sacrificed, and their livers were excised and washed with PBS. The fresh livers were cut into four smaller pieces and placed into a PBS/formaldehyde solution (final formaldehyde concentration = 1%). Crosslinking was stopped by addition of glycine to a final concentration of 0.125 M and incubation for 5 min with rotation. The samples were centrifuged for 5 min at 700g at 4 °C to collect the liver pieces, and the supernatant was removed and washed once again with ice-cold PBS. Fresh PBS containing protease inhibitors (Roche) was added, and the tissue was disaggregated with a homogenizer Ultra Turrax T25 basic (Ika Werke, Staufen, Germany). The tissue was distributed into six tubes (2 ml each) and centrifuged for 5 min at 700g at 4 °C, and the supernatant was discarded. The homogenized tissue was resuspended and lysed with SDS lysis buffer containing protease
DNA constructs
Full-length cDNAs for human PPARα38 and human RXRα39 were subcloned into the T7/SV40 promoterdriven pSG5 expression vector (Stratagene). The same constructs were used for both T7 RNA polymerase-driven in vitro transcription/translation of the respective cDNAs and for viral promoter-driven overexpression in mammalian cells. Promoter regions of the ALAS1 gene and of the CPTI gene were cloned by PCR from human genomic DNA (primers listed in Table S4) and fused with the thymidine kinase promoter driving the firefly luciferase reporter gene. All constructs were verified by sequencing. Gel-shift assays In vitro-translated PPAR and RXR proteins were generated by coupled in vitro transcription/translation
Fig. 5. Functionality of PPREs in the ALAS1 gene. Gel-shift experiments were performed with the in vitro-translated PPARα, RXR alone or in combination, and in the presence of different 32P-labeled REs representing the seven candidate PPREs of the human ALAS1 gene and the human CPTI reference PPRE (a). Protein–DNA complexes were separated from free probe on non-denaturing 8% polyacrylamide gels. Representative gels are shown. PPAR–RXR heterodimer complex formation was quantified on an FLA-3000 reader relative to the reference PPRE. Numbers below the gels indicate the means of at least three independent gel-shift experiments. Standard deviations are shown in parentheses. NS indicates non-specific complexes. Chromatin was extracted from HepG2 cells that had been treated with 100 nM GW7647 for 0, 120 and 240 min, and IPs were performed with antibodies against PPARα, RXRα, PGC-1α and pPol II. Rabbit IgG was used as a specificity control. Real-time PCR was performed on reverse-cross-linked chromatin templates with primers specific to the TSS and six genomic regions containing the seven putative PPREs. Reporter gene assays were performed with extracts from HepG2 cells that were transiently transfected with luciferase reporter constructs containing the indicated REcontaining regions of the ALAS1 gene and with an expression vector for human PPARα (c). The established PPREcontaining region of the human CPTI gene served as positive control. Cells were treated for 16 h with either solvent (DMSO) or 100 nM GW7647. Relative luciferase activities are shown. Columns represent means of at least three experiments, and bars indicate standard deviations. Two-tailed Student's t-tests were performed to determine the significance of the specific antibody enrichment by PPAR agonist relative to control IgG (b) or effects mediated by PPAR overexpression and ligand induction relative to controls (c) (⁎p b 0.05, ⁎⁎p b 0.01, ⁎⁎⁎p b 0.001).
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Fig. 6 (legend on next page)
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5. Transfection and luciferase reporter gene assays
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HepG2 cells were seeded into six-well plates and grown overnight in phenol red-free DMEM supplemented with 10% charcoal-stripped FBS to 60% to 70% confluency. Polyethylenimine transfections were performed by incubating a reporter plasmid and the expression vector for human PPARα or empty pSG5 expression vector (1 μg each) with 10 μg of polyethylenimine (Sigma-Aldrich) in 100 μl of 150 mM NaCl for 15 min at room temperature. After dilution with 900 μl of phenol red-free DMEM, the mixture was added to the cells. Phenol red-free DMEM supplemented with 500 μl of 15% charcoal-stripped FBS, containing 100 mM GW7647 or DMSO, was added 8 h after transfection. The cells were lysed 16 h later using reporter gene lysis buffer (Roche). The constant light signal luciferase reporter gene assay was performed as recommended by the supplier (Perkin-Elmer, Groningen, The Netherlands). Luciferase activities were normalized with respect to protein concentration, and induction factors were calculated as the ratio of luciferase activity of ligand-stimulated cells to that of solvent controls.
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Acknowledgements
lian cellular functions: molecular, cellular, and pharmacological aspects. Pharmacol. Ther. 111, 327–345. Li, B., Holloszy, J. O. & Semenkovich, C. F. (1999). Respiratory uncoupling induces delta-aminolevulinate synthase expression through a nuclear respiratory factor-1-dependent mechanism in HeLa cells. J. Biol. Chem. 274, 17534–17540. Anderson, K. E., Bloomer, J. R., Bonkovsky, H. L., Kushner, J. P., Pierach, C. A., Pimstone, N. R. & Desnick, R. J. (2005). Recommendations for the diagnosis and treatment of the acute porphyrias. Ann. Intern. Med. 142, 439–450. Kauppinen, R. (2005). Porphyrias. Lancet, 365, 241–252. Kolluri, S., Sadlon, T. J., May, B. K. & Bonkovsky, H. L. (2005). Haem repression of the housekeeping 5aminolaevulinic acid synthase gene in the hepatoma cell line LMH. Biochem. J. 392, 173–180. May, B. K., Dogra, S. C., Sadlon, T. J., Bhasker, C. R., Cox, T. C. & Bottomley, S. S. (1995). Molecular regulation of heme biosynthesis in higher vertebrates. Prog. Nucleic Acid Res. Mol. Biol. 51, 1–51. Thunell, S., Harper, P. & Brun, A. (2000). Porphyrins, porphyrin metabolism and porphyrias: IV. Pathophysiology of erythyropoietic protoporphyria—diagnosis, care and monitoring of the patient. Scand. J. Clin. Lab. Invest. 60, 581–604. Fraser, D. J., Podvinec, M., Kaufmann, M. R. & Meyer, U. A. (2002). Drugs mediate the transcriptional activation of the 5-aminolevulinic acid synthase (ALAS1) gene via the chicken xenobiotic-sensing nuclear receptor (CXR). J. Biol. Chem. 277, 34717–34726. Fraser, D. J., Zumsteg, A. & Meyer, U. A. (2003). Nuclear receptors constitutive androstane receptor and pregnane X receptor activate a drug-responsive enhancer of the murine 5-aminolevulinic acid synthase gene. J. Biol. Chem. 278, 39392–39401. Podvinec, M., Handschin, C., Looser, R. & Meyer, U. A. (2004). Identification of the xenosensors regulating human 5-aminolevulinate synthase. Proc. Natl. Acad. Sci. USA, 101, 9127–9132. Peyer, A. K., Jung, D., Beer, M., Gnerre, C., Keogh, A., Stroka, D. et al. (2007). Regulation of human liver deltaaminolevulinic acid synthase by bile acids. Hepatology, 46, 1960–1970. Handschin, C., Lin, J., Rhee, J., Peyer, A. K., Chin, S., Wu, P. H. et al. (2005). Nutritional regulation of hepatic heme biosynthesis and porphyria through PGC-1alpha. Cell, 122, 505–515. Chawla, A., Repa, J. J., Evans, R. M. & Mangelsdorf, D. J. (2001). Nuclear receptors and lipid physiology: opening the X-files. Science, 294, 1866–1870. Willson, T. M., Brown, P. J., Sternbach, D. D. & Henke, B. R. (2000). The PPARs: from orphan receptors to drug discovery. J. Med. Chem. 43, 527–550. Mandard, S., Müller, M. & Kersten, S. (2004). Peroxisome proliferator-activated receptor alpha target genes. Cell. Mol. Life Sci. 61, 393–416. Akiyama, T. E., Lambert, G., Nicol, C. J., Matsusue, K., Peters, J. M., Brewer, H. B., Jr & Gonzalez, F. J. (2004).
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using their respective pSG5-based full-length cDNA expression constructs and rabbit reticulocyte lysate as recommended by the supplier (Promega). Protein batches were quantified by test translations in the presence of 35Slabeled methionine. Gel-shift assays were performed as described previously.29 The sequences of the PPREs are summarized in Table S5.
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This work was supported by grants from the Academy of Finland, the Juselius Foundation and the European Union (Marie Curie RTN NucSys).
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Supplementary data
Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j. jmb.2009.03.024
References
1. Atamna, H. (2004). Heme, iron, and the mitochondrial decay of ageing. Ageing Res. Rev. 3, 303–318. 2. Ponka, P. (1999). Cell biology of heme. Am. J. Med. Sci. 318, 241–256. 3. Tsiftsoglou, A. S., Tsamadou, A. I. & Papadopoulou, L. C. (2006). Heme as key regulator of major mamma-
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Fig. 6. Fasting-induced PPARα TSS association and heme synthesis in mouse liver. Male pure-bred Sv129 mice and PPARα null mice on an Sv129 background were fed or fasted for 24 h (three mice per group). The livers of sacrificed mice were excised and immediately cross-linked with formaldehyde. The chromatin of the livers was extracted, and IPs were performed with antibodies against PPARα (a), RXRα (b), PGC-1α (c) and pPol II (d). Rabbit IgG was used as a specificity control. Real-time PCR was performed on reverse-cross-linked chromatin templates with primers specific to genomic regions of the eight mouse hepatic heme biosynthesis genes. Columns indicate the fold enrichment compared with IgG control precipitations. Representative results from one mouse are shown. In the same set of fed or fasted mice, heme concentration (e) was determined as described in Fig. 2.
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vator complex interacts directly with nuclear receptors in a ligand-dependent fashion. Proc. Natl. Acad. Sci. USA, 95, 7939–7944. Degenhardt, T., Matilainen, M., Herzig, K. H., Dunlop, T. W. & Carlberg, C. (2006). The insulin-like growth factor-binding protein 1 gene is a primary target of peroxisome proliferator-activated receptors. J. Biol. Chem. 281, 39607–39619. Heinäniemi, M., Uski, J. O., Degenhardt, T. & Carlberg, C. (2007). Meta-analysis of primary target genes of peroxisome proliferator-activated receptors. Genome Biol. 8, R147. Mascaro, C., Acosta, E., Ortiz, J. A., Marrero, P. F., Hegardt, F. G. & Haro, D. (1998). Control of human muscle-type carnitine palmitoyltransferase I gene transcription by peroxisome proliferator-activated receptor. J. Biol. Chem. 273, 8560–8563. Rakhshandehroo, M., Sanderson, L. M., Matilainen, M., Stienstra, R., Carlberg, C., de Groot, P. J. et al. (2007). Comprehensive analysis of PPARalpha-dependent regulation of hepatic lipid metabolism by expression profiling. PPAR Res. 2007, 26839. Granick, S., Sinclair, P., Sassa, S. & Grieninger, G. (1975). Effects by heme, insulin, and serum albumin on heme and protein synthesis in chick embryo liver cells cultured in a chemically defined medium, and a spectrofluorometric assay for porphyrin composition. J. Biol. Chem. 250, 9215–9225. Lindberg, R. L., Porcher, C., Grandchamp, B., Ledermann, B., Burki, K., Brandner, S. et al. (1996). Porphobilinogen deaminase deficiency in mice causes a neuropathy resembling that of human hepatic porphyria. Nat. Genet. 12, 195–199. Sassa, S. (2002). The porphyrias. Photodermatol. Photoimmunol. Photomed. 18, 56–67. Wang, X. & Seed, B. (2003). A PCR primer bank for quantitative gene expression analysis. Nucleic Acids Res. 31, e154. Kohany, O., Gentles, A. J., Hankus, L. & Jurka, J. (2006). Annotation, submission and screening of repetitive elements in Repbase: RepbaseSubmitter and Censor. BMC Bioinformatics, 7, 474. Sher, T., Yi, H.-F., McBride, O. W. & Gonzalez, F. J. (1993). cDNA cloning, chromosomal mapping, and functional characterization of the human peroxisome proliferator activated receptor. Biochemistry, 32, 5598–5604. Mangelsdorf, D. J., Ong, E. S., Dyck, J. A. & Evans, R. M. (1990). Nuclear receptor that identifies a novel retinoic acid response pathway. Nature, 345, 224–229.
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Peroxisome proliferator-activated receptor beta/delta regulates very low density lipoprotein production and catabolism in mice on a Western diet. J. Biol. Chem. 279, 20874–20881. Di-Poi, N., Tan, N. S., Michalik, L., Wahli, W. & Desvergne, B. (2002). Antiapoptotic role of PPARβ in keratinocytes via transcriptional control of the Akt1 signaling pathway. Mol. Cell, 10, 721–733. Yamazaki, K., Kuromitsu, J. & Tanaka, I. (2002). Microarray analysis of gene expression changes in mouse liver induced by peroxisome proliferatoractivated receptor α agonists. Biochem. Biophys. Res. Commun. 290, 1114–1122. Frederiksen, K. S., Wulff, E. M., Sauerberg, P., Mogensen, J. P., Jeppesen, L. & Fleckner, J. (2004). Prediction of PPARα ligand-mediated physiological changes using gene expression profiles. J. Lipid Res. 45, 592–601. Vanden Heuvel, J. P., Kreder, D., Belda, B., Hannon, D. B., Nugent, C. A., Burns, K. A. & Taylor, M. J. (2003). Comprehensive analysis of gene expression in rat and human hepatoma cells exposed to the peroxisome proliferator WY14,643. Toxicol. Appl. Pharmacol. 188, 185–198. Kersten, S., Desvergne, B. & Wahli, W. (2000). Roles of PPARs in health and disease. Nature, 405, 421–424. Kliewer, S. A., Umesono, K., Noonan, D. J., Heyman, R. A. & Evans, R. M. (1992). Convergence of 9-cis retinoic acid and peroxisome proliferator signalling pathways through heterodimer formation of their receptors. Nature, 358, 771–774. Jpenberg, A., Jeannin, E., Wahli, W. & Desvergne, B. (1997). Polarity and specific sequence requirements of peroxisome proliferator-activated receptor (PPAR)/ retinoid X receptor heterodimer binding to DNA. A functional analysis of the malic enzyme gene PPAR response element. J. Biol. Chem. 272, 20108–20117. Vega, R. B., Huss, J. M. & Kelly, D. P. (2000). The coactivator PGC-1 cooperates with peroxisome proliferator-activated receptor α in transcriptional control of nuclear genes encoding mitochondrial fatty acid oxidation enzymes. Mol. Cell. Biol. 20, 1868–1876. Tudor, C., Feige, J. N., Pingali, H., Lohray, V. B., Wahli, W., Desvergne, B. et al. (2007). Association with coregulators is the major determinant governing peroxisome proliferator-activated receptor mobility in living cells. J. Biol. Chem. 282, 4417–4426. Yuan, C.-X., Ito, M., Fondell, J. D., Fu, Z.-Y. & Roeder, R. G. (1998). The TRAP220 component of a thyroid hormone receptor-associated protein (TRAP) coacti-
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Peroxisome Proliferator-Activated Receptor α Controls Hepatic Heme Biosynthesis Through ALAS1
Department of Biochemistry, University of Kuopio, FIN70211 Kuopio, Finland 2
Nutrigenomics Consortium, Top Institute of Food and Nutrition, Wageningen University, 6701 HD Wageningen, The Netherlands 3
Metabolism and Genomics Group, Division of Human Nutrition, Wageningen University, 6701 HD Wageningen, The Netherlands 4
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Life Sciences Research Unit, Université du Luxembourg, 162A, Avenue de la Faïencerie, L-1511 Luxembourg, Luxembourg
Heme is an essential prosthetic group of proteins involved in oxygen transport, energy metabolism and nitric oxide production. ALAS1 (5aminolevulinate synthase) is the rate-limiting enzyme in heme synthesis in the liver and is highly regulated to adapt to the metabolic demand of the hepatocyte. In the present study, we describe human hepatic ALAS1 as a new direct target for the nuclear receptor peroxisome proliferator-activated receptor α (PPARα). In primary human hepatocytes and in HepG2 cells, PPARα agonists induced an increase in ALAS1 mRNA levels, which was abolished by PPARα silencing. These effects are mediated by two functional PPAR binding sites at positions − 9 and −2.3 kb relative to the ALAS1 transcription start site. PPARα ligand treatment also up-regulated the mRNA levels of the genes ALAD (5-aminolevulinate dehydratase), UROS (uroporphyrinogen III synthase), UROD (uroporphyrinogen decarboxylase), CPOX (coproporphyrinogen oxidase) and PPOX (protoporphyrinogen oxidase) encoding for enzymes controlling further steps in heme biosynthesis. In HepG2 cells treated with PPARα agonists and in mouse liver upon fasting, the association of PPARα, its partner retinoid X receptor, PPARγ coactivator 1α and activated RNA polymerase II with the transcription start site region of all six genes was increased, leading to higher levels of the metabolite heme. In conclusion, these data strongly support a role of PPARα in the regulation of human ALAS1 and of five additional genes of the pathway, consequently leading to increased heme synthesis.
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Tatjana Degenhardt 1 , Sami Väisänen 1 , Maryam Rakhshandehroo 2,3 , Sander Kersten 2,3 and Carsten Carlberg 1,4 ⁎
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Received 14 November 2008; received in revised form 5 March 2009; accepted 9 March 2009 Available online 14 March 2009 Edited by J. Karn
© 2009 Elsevier Ltd. All rights reserved.
Keywords: nuclear receptor; transcriptional regulation; chromatin; metabolism; PPARα
Introduction
Heme is an irreplaceable compound for mammalian life. It is an essential component of numerous heme-containing proteins that exhibit functions in mitochondrial respiration, hormone synthesis and
metabolism and nitric oxide synthesis.1–3 Heme biosynthesis in eukaryotic cells is composed of eight enzymatic steps, of which the first and the last three steps take place in the mitochondria, whereas the others occur in the cytoplasm. Eighty to ninety percent of the total heme in mammals is synthesized
*Corresponding author. E-mail address: [email protected]. Abbreviations used: ALAD, 5-aminolevulinate dehydratase; ALAS, 5-aminolevulinate synthase; BSA, bovine serum albumin; ChIP, chromatin immunoprecipitation; CPOX, coproporphyrinogen oxidase; CYP, cytochrome P450; DMEM, Dulbecco's modified Eagle's medium; DMSO, dimethyl sulfoxide; DR1, direct repeat spaced by one nucleotide; FBS, fetal bovine serum; FECH, ferrochelatase; GW7647, 2-(4-(2-(1-cyclohexanebutyl-3-cyclohexylureido)ethyl)phenylthio)-2methylpropionic acid; HMBS, hydroxymethylbilane synthase; PBS, phosphate-buffered saline; PGC-1α, PPARγ coactivator 1α; PPAR, peroxisome proliferator-activated receptor; pPol II, phosphorylated RNA polymerase II; PPOX, protoporphyrinogen oxidase; PPRE, PPAR response element; RE, response element; RPLP0, acidic riboprotein P0; RXR, retinoid X receptor; TSS, transcription start site; UROD, uroporphyrinogen decarboxylase; UROS, uroporphyrinogen III synthase; Wy14,643, pirinixic acid. 0022-2836/$ - see front matter © 2009 Elsevier Ltd. All rights reserved.
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number of direct inducible PPAR target genes have been identified;20,21 mostly, these genes represent different metabolic pathways. In humans, little knowledge is available regarding the regulation of whole pathways by PPARα.22 Usually, only a gene family or one single gene is examined, while whole pathways under the control of PPARs have not yet been intensively studied. An essential prerequisite for the direct modulation of transcription by PPAR ligands is the location of at least one activated PPAR protein close to the transcription start site (TSS) of the target gene. This is commonly achieved through the binding of PPARs to a specific sequence of double-stranded DNA, called a PPAR response element (PPRE), often located thousands of base pairs up- or downstream of a gene's TSS. A subsequent DNA-looping event links the activated PPAR–PPRE complex to the TSS.23 PPARs bind to DNA as heterodimers with the nuclear receptor retinoid X receptor (RXR).24 PPREs are formed by two hexameric motifs with the optimal AGGTCA core binding sequence in a direct repeat orientation with a spacing of one nucleotide (DR1), where PPAR occupies the 5′ motif.25 Binding of agonists to the PPARs causes a conformational change within their ligand-binding domain that results in an enhanced binding of co-activator proteins, such as PGC-1α.26,27 These co-activators link ligand-activated PPARs to enzymes displaying histone acetyltransferase activity that cause chromatin relaxation. In a subsequent step, ligandactivated PPARs rapidly exchange co-activator proteins for components of mediator complexes,28 which act as a bridge from the activated PPARs to the basal transcriptional machinery. In the present study, we investigated the role of PPARα as a direct regulator of ALAS1 and the whole heme biosynthesis pathway in primary hepatocytes and HepG2 cells. We identified that the genes ALAS1, ALAD (5-aminolevulinate dehydratase), UROS (uroporphyrinogen III synthase), UROD (uroporphyrinogen decarboxylase), CPOX (coproporphyrinogen oxidase) and PPOX (protoporphyrinogen oxidase) are direct PPARα targets, thereby connecting PPARα signaling and hepatic heme biosynthesis.
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in erythroid cells and is subsequently incorporated into hemoglobin. Regulation of heme biosynthesis in these cells involves the enzyme encoded by the erythroid-specific ALAS2 (5-aminolevulinate synthase 2) gene. In contrast, the housekeeping gene ALAS, also called ALAS1 or ALAS-N, is ubiquitously expressed, as all nucleated cells need to synthesize heme for their cytochrome P450 (CYP) enzymes. Most of the heme that is synthesized outside of erythroid cells is produced in the liver for various heme-containing enzymes, in particular for microsomal CYPs. In mitochondrial biogenesis, coordinated increase of heme synthesis and the synthesis of respiratory CYPs are prerequisites for a functional energy metabolism.4 Hereditary partial defects of enzymes involved in heme biosynthesis lead to rare metabolic diseases known as porphyrias. Inducible, acute hepatic porphyrias are characterized by attacks of neuropsychiatric dysfunction, precipitated by stimuli, such as fasting, alcohol, drugs and sex steroids.5,6 The detrimental effects of heme to the cell make a tight control of its biosynthesis essential. In non-erythroid cells, the rate of heme synthesis is controlled at its first enzymatic step. Accordingly, the ALAS1 gene is highly regulated in different cellular contexts to ensure adequate levels of heme.7–9 In recent years, progress has been made in understanding the regulation of ALAS1 in liver on a molecular level. Members of the nuclear receptor superfamily, such as the xenosensors pregnane X receptor and constitutive androstane receptor and the bile acidactivated farnesoid X receptor,10–13 have been shown to play a role in the regulation of ALAS1. Furthermore, the peroxisome proliferator-activated receptor (PPAR) γ co-activator 1α (PGC-1α), which is a key regulator in mitochondrial biogenesis and energy homeostasis, was shown to be important for a fasting response of ALAS1 by acting through the insulin-regulated transcription factor Forkhead O1A binding to the ALAS1 promoter,14 whereas the other genes of the pathway were not subject to studies. Nuclear receptors are transcription factors that have important roles in controlling cellular metabolism because many of them are activated by lipophilic ligands, including cholesterol, fatty acids and their metabolic derivatives.15 The three members of the PPAR subfamily, PPARα, PPARγ and PPARβ/δ, are thought to play a prominent role in the development of the metabolic syndrome since they are key regulators of lipid storage and catabolism.16 PPARα has been studied most intensively in the context of liver metabolism and is known to control hepatic fatty acid catabolism.17 PPARγ, which is highly expressed in adipose tissue, is a master regulator of adipogenesis. The widely expressed PPARβ/δ stimulates fatty acid oxidation, regulates hepatic very-low-density lipoprotein production and catabolism18 and is involved in more diverse actions, such as wound healing by governing keratinocyte differentiation.19 All three receptors are activated by (mainly polyunsaturated) fatty acids and various fatty acid derivatives, such as eicosanoids. In rodents, a large
Results PPARα-dependent regulation of the genes involved in hepatic heme biosynthesis To explore the regulation of the heme biosynthetic genes by PPARα, we used human primary hepatocytes from six donors that were treated for 6 and 24 h with 50 μM concentration of the PPARα-specific agonist Wy14,643 (pirinixic acid; Fig. 1a). RNA was extracted, and expression changes were monitored by quantitative real-time PCR. The genes ALAS1 (2.8-fold after 6 h, 1.5-fold after 24 h), ALAD (1.3-fold after 6 and 24 h), UROS (1.3-fold after 6 h, 1.2-fold after 24 h), CPOX (1.2-fold after 6 and 24 h) and PPOX (1.2-fold after 6 and 24 h) were significantly
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Fig. 1. Expression profiling of the eight genes involved in hepatic heme biosynthesis. (a) Human primary hepatocytes from six donors were stimulated with 50 μM concentration of the PPARα ligand Wy14,643. (b, c) HepG2 cells were stimulated with 100 nM concentration of the PPARα ligand GW7647 for 6 and 24 h. (d) HepG2 cells were transfected with either control siRNA or siRNA specific for PPARα and incubated for 48 h. RNA was extracted and quantitative real-time PCR was performed for the eight heme biosynthesis genes. Columns indicate the means of at least three independent treatments, and the bars represent standard deviations. Two-tailed, paired Student's t-tests were performed to determine the significance of the mRNA induction by PPARα agonist relative to solvent controls (a–c) and that of PPARα siRNA knockdown relative to control siRNA (d) (⁎p b 0.05, ⁎⁎p b 0.01, ⁎⁎⁎p b 0.001).
up-regulated, whereas no regulation was observed for HMBS (hydroxymethylbilane synthase), UROD and FECH (ferrochelatase).
Next, we asked the question of whether some of the eight genes of the hepatic heme biosynthesis are also PPARα targets in the human hepatocellular
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Fig. 2. Increase in heme biosynthesis. The hepatic heme biosynthesis pathway is illustrated. Enzymes that are encoded by PPARα target genes are depicted in red, whereas non-regulated genes are illustrated in blue (a). HepG2 cells were treated for 6, 8 and 24 h with GW7647 or DMSO. The cells were lysed, and heme concentration was determined and normalized to the total protein content (b). Columns indicate the means of at least three independent treatments, and the bars represent standard deviations. Two-tailed, paired Student's t-tests were performed to determine the significance of the induction of heme concentration by PPARα agonist relative to solvent controls (⁎p b 0.05).
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carcinoma cell line HepG2. For this purpose, HepG2 cells were treated for 6 or 24 h with 100 nM concentration of the PPARα ligand GW7647 [2-(4-(2(1-cyclohexanebutyl-3-cyclohexylureido)ethyl)phenylthio)-2-methylpropionic acid], RNA was extracted and expression changes were monitored using quantitative real-time PCR (Fig. 1b). The results confirmed the findings in primary hepatocytes (Fig. 1a); again, HMBS and FECH did not show any response at any time point, whereas ALAS1 (1.7fold after 6 h, 2.1-fold after 24 h), ALAD (1.2-fold after 6 h), UROS (1.5-fold after 6 h, 1.4-fold after 24 h), UROD (1.4-fold after 6 and 24 h), CPOX (1.5fold after 6 h, 1.3-fold after 24 h) and PPOX (1.8-fold after 6 h, 1.5-fold after 24 h) were significantly upregulated. In order to exclude the possibility that there may be specific effects of the two PPARα ligands, we also treated HepG2 cells with 50 μM Wy14,643 and observed a significant induction of the same six genes of the heme pathway (Fig. S1a). The lowest expressed genes, HMBS and FECH (Fig. S1b), did not respond to any PPAR agonist. To assess possible PPARα-independent ligand effects, we silenced PPARα in HepG2 cells with PPARα-specific siRNA oligonucleotides (Fig. 1c and d). After silencing PPARα, mRNA levels were 75% reduced, whereas the levels of PPARγ (10% reduction) and PPARβ/δ (35% reduction) were only slightly affected (Fig. S2a). On protein level, PPARα was reduced after siRNA treatment by about 50%, whereas PPARγ and PPARβ/δ were not affected (Fig. S2b). In the PPARα-silenced cells, we could not detect any changes of gene expression after 6 or 24 h of treatment with GW7647 (Fig. 1c), indicating that the observed effects are clearly linked to PPARαdependent gene regulation. Moreover, after PPARα silencing, the basal expression levels of ALAS1,
ALAD, UROS, UROD, CPOX and PPOX also were reduced by 60%, 37%, 40%, 20%, 33% and 36%, respectively (Fig. 1d), indicating an important role of PPARα in the regulation of these genes. In contrast, the genes HMBS and FECH were not significantly affected in their basal expression by silencing PPARα. Taken together, quantitative real-time PCR analysis of the hepatic heme biosynthesis pathway in primary human hepatocytes and in HepG2 cells suggested that the genes ALAS1, ALAD, UROS, UROD, CPOX and PPOX are likely to be direct PPARα targets. Increase in heme concentration An overview on the hepatic heme biosynthesis pathway is shown in Fig. 2a. To gain insight on whether the PPARα-dependent regulation of six of the eight genes leads in turn to an activation of the whole pathway, resulting into an increased production of heme, we treated HepG2 cells with PPARα agonist GW7647 or the solvent dimethyl sulfoxide (DMSO) and measured the heme concentration of the cells after ligand treatment. Intriguingly, already after 8 h we could detect a significant 1.5-fold increase in heme concentration that after prolonged treatment with ligand (1.6-fold after 24 h) still increased (Fig. 2b). In summary, the PPARα-dependent transcriptional control leads to an increase in heme levels in HepG2 cells. Conservation of PPREs in the heme biosynthetic genes Recently, we developed an in silico screening method for a reliable prediction of DR1-type
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Functional PPREs in the ALAS1 gene
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The rate-limiting enzyme ALAS1 displayed the strongest response to PPARα ligands and based on our in silico screening the ALAS1 gene carries seven putative PPREs in a distance of less than 10 kb from the TSS, of which three (RE2, RE4 and RE6) are conserved compared with the orthologous sequence in mouse (Fig. 3). In order to test the affinity of these REs for PPARs, we performed gel-shift assays with in vitro-translated PPARα–RXR heterodimers and the seven candidate PPREs of the ALAS1 gene (Fig. 5a). Compared with the established PPRE of the CPTI gene,31 PPARα–RXR heterodimers bound RE3 with 21.1% strength, RE1 with 10.2%, RE6 with 8.6% and RE2 with 3.3%, while they bound RE4, RE5 and RE7 with 1.3% or less. The functionality of the REs was further investigated using ChIP assays in HepG2 cells (Fig. 5b). Please note that RE4 and RE5 are so close to each other that they could not be distinguished. The TSS of the ALAS1 gene served as a positive control. Significant binding of PPARα was found in regions carrying RE1, RE2, RE4/5 and RE6, that of RXRα was found in regions of RE2, RE6 and RE7, that of PGC-1α was found in regions of RE1, RE2, RE4/5 and RE6 and that of pPol II was found in regions of RE1, RE4/5, RE6 and RE7. These association patterns on chromatin region show no direct correlation with the affinity of the PPREs that they carry, which suggests that in vitro binding affinity may be not the only criterion for the functionality of a PPAR-responsive region. However, the regions of the conserved RE2 and RE6, as well as the region of RE1, were found to be functional. In contrast, the highest affinity putative PPRE, RE3, appears to be not accessible to PPARα binding. Finally, we tested the functionality of the identified REs by reporter gene assays in transiently transfected HepG2 cells (Fig. 5c). The genomic regions investigated in ChIP assays (Fig. 5b) were fused with the thymidine kinase promoter driving the luciferase reporter gene. The genomic region of the human CPTI gene carrying the established PPRE served as a positive control. PPARα overexpression resulted for all genomic regions in increases of their basal activity, which were 2.0-fold even for the empty reporter gene vector, 5.3-fold for the region of the CPTI gene, 4.8-fold for the genomic region around ALAS1 RE1, 5.2-fold for RE2, 2.4-fold for RE3, 19.6-fold for RE4/5, 18.1-fold for RE6 and 15.2fold for RE7. However, most informative were the statistically significant increases after stimulation with GW7647, which were 2.1-fold for the CPTI region and 1.7-, 1.6- and 1.2-fold for regions 1, 2 and 7 of the ALAS1 gene, respectively. The three other ALAS1 regions did not show any significant ligand induction. Taken together, our in vitro analysis of the seven candidate PPREs of the ALAS1 gene indicated for four a reasonable affinity for PPARα–RXR, three of which (RE1, RE2 and RE6) could also be confirmed by ChIP analysis in HepG2 cells. Three of RE-
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PPREs.29,30 We now applied this method to the genomic regions 10 kb up- and downstream of the TSSs of the eight human heme biosynthesis genes (Fig. 3) and their mouse orthologs (data not shown). Within the 20 kb of analyzed sequence, each of the genes displayed between none (UROS) and seven (ALAS and PPOX) putative PPREs. A comparison with the respective mouse orthologs indicated that the eight genes contain 28 PPREs in humans and 23 PPREs in the mouse, of which 17 (74% and 61%, respectively) are conserved between both species. This suggests a high degree of conservation in the regulation of this pathway. The PPOX gene showed six conserved PPREs, the ALAS1 and UROD genes showed three, the ALAD gene showed two, the HMBS, CPOX and FECH genes showed one and only the UROS gene showed no conserved PPRE. This means that ALAS1, ALAD, UROD and PPOX are more likely PPAR target genes than HMBS, UROS, CPOX and FECH. Taken together, most of the genes responding to PPARα ligands and PPARα siRNA knockdown show a significantly increased number of potential PPAR binding sites in the genomic sequence 10 kb up- or downstream of their TSSs, compared with the unresponsive genes HMBS and FECH.
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Recruitment of PPARα to the TSS regions of the heme biosynthesis genes
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For an experimental validation of the prediction of PPAR binding to the regulatory regions of heme biosynthetic genes, we tested the binding of PPARα to the TSS for each of the eight genes using chromatin immunoprecipitation (ChIP) assays. In addition, binding of its heterodimeric partner RXRα, the common co-activator PGC-1α and activated RNA polymerase [phosphorylated RNA polymerase II (pPol II)] was assessed. Only active PPREs will loop together with their associated proteins to the TSS hosting Pol II, connecting the activated PPRE with the basal transcription machinery. Therefore, the association of the TSS with PPARα, RXRα and PGC-1α indicates that at least one of the putative PPREs of the respective genes was active. We treated HepG2 cells for 240 and 360 min with GW7647 and cross-linked the proteins to DNA using formaldehyde (Fig. 4). Quantification of ChIP assays by realtime PCR showed that PPARα enriched at both time points on the TSS regions of ALAS1, ALAD, UROS, UROD, CPOX and PPOX (Fig. 4a). In contrast, on the TSSs of the genes HMBS and FECH, no significant enrichment of PPARα binding was observed. RXRα (Fig. 4b), PGC-1α (Fig. 4c) and pPol II (Fig. 4d) essentially showed the same binding pattern—i.e., they were found on the TSSs of ALAS1, ALAD, UROS, UROD, CPOX and PPOX but not on those of HMBS and FECH. In summary, in HepG2 cells, PPARα, RXRα, PGC1α and pPol II associate with the TSS regions of the genes ALAS1, ALAD, UROS, UROD, CPOX and PPOX but not with those of the genes HMBS and FECH.
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Fig. 3 (legend on next page)
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Fasting-induced changes in PPARα TSS binding and heme concentration
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For experiments in mice, we fed or fasted wildtype mice and PPARα knockout mice for 24 h (three mice per group), removed their livers and immediately cross-linked the nuclear proteins to the genomic DNA (Fig. 6). Quantification of ChIP assays by real-time PCR showed that PPARα associated with the TSS regions of Alas1, Alad, Uros, Urod, Cpox and Ppox when PPARα wild-type mice were fasted (Fig. 6a). This shows that PPARαmediated transcription is stronger than the basal transcription of the heme biosynthesis pathway genes. However, the TSSs of Hmbs and Fech showed no significant enrichment of the receptor. The same binding pattern that was observed for RXRα (Fig. 6b), PGC-1α (Fig. 6c) and pPol II (Fig. 6d) binding could confirm these associations with the exception of the TSS of the Urod gene. To gain insight on whether the PPARα-dependent regulation of the heme biosynthetic genes leads in turn to an activation of the whole pathway, resulting into an increased production of heme, we measured heme concentration in livers from wild-type and PPARα null mice that were fed or fasted for 24 h. Small liver pieces were homogenized and heme concentration was measured. We observed a significant 2.1-fold increase in heme levels in the fasted wild-type mice compared with the fed wild-type mice (Fig. 6e). Although we found slightly higher heme levels in the fasted PPARα null mice compared with the fed null mice, we were able to observe a significant decrease in heme levels in the fasted PPARα null mice compared with the fasted wildtype mice. In summary, in mouse liver PPARα, RXRα, PGC1α and pPol II associate with the TSS regions of Alas1, Alad, Uros, Urod, Cpox and Ppox but not with those of Hmbs and Fech. Moreover, the heme levels in livers of fasted wild-type mice increase compared with fed mice and are also higher than those in fasted PPARα null mice.
the first enzyme in heme biosynthesis, ALAS1, by fasting through the co-activator PGC-1α.14 In addition, several other nuclear receptors, such as pregnane X receptor, constitutive androstane receptor and farnesoid X receptor, were suggested to regulate ALAS1 expression.10–13 In this study, we demonstrated that human hepatic ALAS1, as well as five others out of the additional seven genes of the hepatic heme biosynthesis pathway, is a novel direct target for PPARα. By quantitative real-time PCR using primary human hepatocytes and HepG2 cells, siRNA knockdown in HepG2 cells, in silico screening and ChIP assays in HepG2 cells and in mouse liver, we consistently demonstrated that the genes ALAS1, ALAD, UROS, UROD, CPOX and PPOX are PPARα target genes. In contrast, the genes HMBS and FECH carry only one conserved PPRE, which is apparently insufficient, since these two genes showed no sign of regulation by PPARα. Moreover, we showed that PPARα agonist treatment increases heme concentration in HepG2 cells and in fasted wild-type mice. The response to PPARα agonists was most prominent for the ALAS1 gene. We found seven putative PPREs within 10 kb of the TSS of this gene. Two of these PPREs (RE1 and RE2 at positions − 9 and − 2.3 kb relative to the ALAS1 TSS, respectively) showed significant in vitro binding for PPARα–RXR heterodimers as well as association of PPARα and PGC-1α to their chromatin regions and mediated ligand inducibility to their genomic regions in reporter gene assays. The other five putative sites show inconsistent effects and may not be functional. Many studies focus on the regulation of ALAS1 since it catalyzes the rate-limiting step in hepatic heme biosynthesis.9 The accumulation of the intermediates of the heme biosynthesis pathway is the biochemical hallmark of porphyria. It is known that different factors, including fasting, alcohol and drugs, can trigger porphyric attacks in individuals who carry a mutation in one of the enzymes of the heme pathway.5,6 There is high expression not only of ALAS1 but also of ALAD, UROS, UROD, CPOX and PPOX in order to avoid a too high (i.e., toxic) metabolite concentration in the cell upon fasting. Under normal physiological conditions, free heme levels are extremely low and tightly regulated because cellular toxicity can occur with increased cellular concentrations of unincorporated heme. In addition, it is known that treatment with phenobarbital or other prototypical drug inducers elevates heme concentrations in the liver to accommodate the increased levels of heme-dependent enzymes.33–35 In this study, we showed that fasting up-regulates heme concentrations in mouse liver in a PPARα-dependent way. Moreover, other studies demonstrated that this
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containing genomic regions (RE1, RE2 and RE7) were found to be ligand inducible in reporter gene assays. In conclusion, only RE1 and RE2 seem to be fully functional PPREs.
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Discussion
PPARα is emerging as one of the major components regulating liver metabolism. The role of PPARα in the adaptive response to fasting is well established.32 Previous studies showed regulation of
Fig. 3. In silico screening for PPREs in the heme biosynthesis genes in human. Overview of the eight genes of the heme biosynthesis pathway. The diagrams show 10 kb upstream and 10 kb downstream of the TSS (arrow), regions of repetitive sequence (white boxes) and exons (dark gray boxes). Putative REs (red boxes, conserved; blue boxes, non-conserved) were identified by in silico screening of the genomic sequences and are classified according to their degree of conservation between human and mouse.
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Fig. 4. Ligand-induced PPARα recruitment to the TSS of heme biosynthesis genes in HepG2 cells. Chromatin was extracted from HepG2 cells that had been treated with 100 nM GW7647 for 0, 240 and 360 min, and IPs were performed with antibodies against PPARα (a), RXRα (b), PGC-1α (c) and pPol II (d). Rabbit IgG was used as a specificity control. Real-time PCR was performed on reverse-cross-linked chromatin templates with primers specific to genomic regions of the eight human hepatic heme biosynthesis genes. Columns indicate the means of at least three independent treatments, and the bars represent standard deviations. Two-tailed, paired Student's t-tests were performed to determine the significance of the specific antibody enrichment by PPAR agonist relative to control IgG (⁎p b 0.05, ⁎⁎p b 0.01, ⁎⁎⁎p b 0.001).
is also the case for several heme-containing proteins.32 Both observations link the up-regulation of the heme biosynthesis upon fasting and an increase in heme concentration to the need of more heme for heme-containing enzymes, such as Cyp4a10, Cyp4a12 and Cyp4a1.32 This suggests that the regulation of the heme biosynthetic pathway
upon fasting is necessary to keep up with the demand of heme for heme-containing proteins involved in processes, such as fatty acid metabolism. PPARα is one of the most prominent regulators of fatty acid metabolism in liver, showing a clear need for the up-regulation of hepatic heme biosynthesis and to enable a functioning fatty acid metabolism.
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Cell culture experiments
RNAi experiments
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Materials and Methods
temperature (Table S2) and 40 s at 72 °C. Most of the primer sequences were obtained from the PrimerBank at Harvard University.36 The sequences of the gene-specific primer pairs for the human genes involved in heme biosynthesis and the internal control gene RPLP0 (acidic riboprotein P0) are listed in Table S2. PCR product quality was monitored using post-PCR melt curve analysis. Fold inductions were calculated using the formula 2−(ΔΔCt), where ΔΔCt is the ΔCt(PPARα ligand) − ΔCt(DMSO), ΔCt is Ct(gene X) − Ct(RPLP0) and Ct is the cycle at which the threshold is crossed.
HepG2 cells were grown overnight in medium containing charcoal-stripped FBS to 30% confluency. Cells were transfected using Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer's instructions. A total amount of 200 pmol of either control siRNA or a mixture of the three PPARα-specific siRNAs (Eurogentec, Liege, Belgium; for sequences, see Table S3) was transfected. Cells were incubated for 8 h, and then charcoal-stripped FBS was added and the transfection was continued for 48 h in total. Cell treatments and RNA extractions were carried out as described above.
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Taken together, the regulation of the heme biosynthesis pathway by PPARα offers an important insight in the mechanism—how the demand for more hemecontaining proteins upon fasting triggers the transcriptional production of genes involved in the heme biosynthesis pathway. In conclusion, our data demonstrate that the hepatic heme biosynthesis pathway in humans is under direct transcriptional control of PPARα, consequently leading to an increase in heme concentration. The six out of eight genes of the pathway are novel primary PPARα target genes. These also suggest a mechanism by which fasting or fibrate treatment may trigger acute porphyric attacks in susceptible patients.
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The human hepatocellular carcinoma cell line HepG2 was cultured in RPMI 1640 containing 10% fetal bovine serum (FBS), 2 mM L-glutamine, 0.1 mg/ml of streptomycin and 100 U/ml of penicillin in a humidified 95% air/ 5% CO2 incubator. Before use, FBS was stripped of lipophilic compounds, such as endogenous nuclear receptor ligands, by stirring it with 5% activated charcoal (Sigma-Aldrich) for 3 h at room temperature. Charcoal was then removed by centrifugation and sterile filtration. Prior to mRNA, chromatin extraction or transient transfections, cells were grown overnight in phenol red-free Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% charcoal-stripped FBS to reach a density of 50% to 60% confluency. Cells were then treated with either solvent (DMSO, final concentration = 0.1%) or 100 nM concentration of the PPARα agonist GW7647 (Alexis Biochemicals, San Diego, CA). Primary human hepatocytes and Hepatocyte Culture Medium BulletKit were purchased from Lonza Bioscience (Verviers, Belgium). Human hepatocytes were isolated from six donors (Table S1). Cells were plated on collagen-coated six-well plates. Upon arrival of the cells, the medium was discarded and was replaced by Hepatocyte Culture Medium. The next day, cells were incubated in fresh medium in the presence or absence of 50 μM Wy14,643 (ChemSyn Laboratories, Lenexa, KS) dissolved in DMSO for 6 or 24 h, followed by RNA isolation. RNA extraction and real-time quantitative PCR
Total RNA from HepG2 cells was extracted using a Mini RNA Isolation II kit (Zymo Research, HiSS Diagnostics, Freiburg, Germany), and cDNA synthesis was performed for 1 h at 37 °C using 1 μg of total RNA as a template, 100 pmol oligodT18 primer and 40 U of reverse transcriptase (Fermentas, Vilnius, Lithuania). For human primary hepatocytes, RNA was extracted using TRIzol reagent (Invitrogen) and 1 μg of RNA was reverse transcribed using iScript (BioRad, Hercules, CA). Realtime quantitative PCR was performed in an IQ-cycler (BioRad) using the dye SybrGreen I (Molecular Probes, Leiden, The Netherlands). Per reaction, 1 U of Hot Start Taq polymerase (Fermentas) and 3 mM MgCl2 were used and the PCR cycling conditions were as follows: 45 cycles of 30 s at 95 °C, 30 s at primer-specific annealing
Measurement of heme concentration
PPARα wild-type and null mice (n = 4 per group) were fed or fasted for 24 h, their livers were removed and frozen in liquid nitrogen. Small pieces of liver were homogenized in Reagent buffer, and heme concentration was measured using a QuantiChrom™ Heme Assay Kit at 414 nm (BioAssay Systems, Hayward, CA). HepG2 cells were grown overnight in phenol red-free DMEM supplemented with 10% charcoal-stripped FBS and treated for 0, 6, 8 and 24 h with 100 nM GW7647. Cells were washed with phosphate-buffered saline (PBS). The heme concentration was measured according to the manufacturer's instructions. The data were normalized to the total protein content of the sample using the BCA (bicinchoninic acid) protein assay (Pierce Biotechnology, Rockford, IL). In silico PPRE screening Genomic sequences spanning ± 10 kb around the TSSs of the human and mouse genes were extracted from the current database release for the human genome and the mouse genome. Putative PPREs were screened from the sequence files as described previously.29,30 Conservation of putative PPREs between human and mouse was checked using the program Align (MacMolly® Tetra package, Soft Gene GmbH, Bocholt, Germany). Repetitive sequences from both species were obtained using CENSOR.37 ChIP assays in HepG2 Nuclear proteins were cross-linked to genomic DNA by adding formaldehyde for 5 min directly to the medium to a final concentration of 1% at room temperature. Crosslinking was stopped by adding glycine to a final concentration of 0.125 M and incubating for 5 min at room temperature on a rocking platform. The medium was removed and the cells were washed twice with icecold PBS. The cells were collected by scraping into ice-cold PBS supplemented with a protease inhibitor cocktail
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centrifugation and the lysates were diluted 1:10 in ChIP wash buffer 1 (150 mM NaCl, 1% Triton X-100, 2 mM EDTA, 50 mM Tris–HCl, pH 8.1). The samples were centrifuged and the recovered chromatin solutions were incubated with 5 μl of indicated antibodies, 2.4 μl of sonicated salmon sperm (10 mg/ml) and 25 μl of bovine
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(Roche Diagnostics, Mannheim, Germany). After centrifugation, the cell pellets were resuspended in lysis buffer [1% SDS, 10 mM ethylenediaminetetraacetic acid (EDTA), protease inhibitors, 50 mM Tris–HCl, pH 8.1] and the lysates were sonicated to result in DNA fragments of 300 to 1000 bp in length. Cellular debris was removed by
Fig. 5 (legend on next page)
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ChIP assays in mouse liver
inhibitors (Roche) for 10 min at room temperature. Lysates were sonicated to result in DNA fragments of 300 to 1000 bp in length. Cellular debris was removed by centrifugation and the lysates were diluted 1:10 in ChIP wash buffer 1. The recovered chromatin solutions were incubated with 5 μl of indicated antibodies (200 μg/ml), 2.4 μl of sonicated salmon sperm (10 mg/ml) and 25 μl BSA (10 mg/ml) to remove unspecific background overnight at 4 °C with rotation. The clean-up of the samples and recovery of DNA were performed as described above. PCR of chromatin templates
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For the genomic regions containing the TSSs of all eight human and mouse genes involved in heme biosynthesis and the putative REs of the ALAS1 gene, specific primer pairs were designed (Table S4), optimized and controlled by running PCRs with 25 ng of genomic DNA (input) as a template. When running immunoprecipitated DNA (output) as a template, the following PCR profile was used: pre-incubation for 5 min at 94 °C, 45 cycles of 30 s at 95 °C, 30 s at the temperature indicated for each primer pair (Table S4) and 30 s at 72 °C and one final incubation for 10 min at 72 °C. The primers were controlled using melt curve analysis and gel pictures to allow the detection of primer-specific products. The fold change relative to the non-specific IgG background was calculated. The fold inductions were calculated using the formula 2−(ΔCt), where ΔCt is Ct(specific antibody) − Ct(IgG control) and Ct is the cycle where the threshold is crossed.
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serum albumin (BSA) (10 mg/ml) to remove unspecific background overnight at 4 °C with rotation. The antibodies against PPARα (sc-9000), RXRα (sc-553), PGC-1α (sc-13067), pPol II (sc-13583) and control immunoglobulin G's (IgGs) (sc-2027) were obtained from Santa Cruz Biotechnologies. The immunocomplexes were collected with 25 μl of MagnaCell magnetic protein A agarose beads (Cortex Biochem, Madison, WI) for 1 h at room temperature with rotation. The magnetic beads were pre-blocked overnight in a solution containing ChIP wash buffer 1, protease inhibitors, BSA and sonicated salmon sperm in the same concentrations as used later in the immunocollection. Magnetic beads were collected using a magnetic rack (Qiagen) and were washed twice for 1 min with ChIP wash buffer 1 containing 1 mM PMSF, 5 min with ChIP wash buffer 2 (500 mM NaCl, 1% Triton X-100, 2 mM EDTA, 0.1% SDS, 20 mM Tris–HCl, pH 8.0, containing 1 mM freshly added PMSF), 5 min with ChIP wash buffer 3 (0.25 mM LiCl, 1% Nonidet P-40, 1% sodium deoxycholate, 1 mM EDTA, 10 mM Tris–HCl, pH 8.1) and finally twice for 1 min with TE buffer (1 mM EDTA, 10 mM Tris– HCl, pH 8.0). Then, 250 μl of elution buffer (10 mM EDTA, 0.5% SDS, 25 mM Tris–HCl, pH 7.5) was added and incubated for 30 min at 64 °C. The supernatant was removed and the beads were washed for 2 min with 250 μl of elution buffer. The supernatants were combined, and proteins were digested overnight at 64 °C using proteinase K (Fermentas, final concentration = 40 μg/ml). Genomic DNA fragments were recovered by phenol–chloroform extraction and a following ethanol precipitation.
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Mice were fed or fasted for 24 h (three mice per group). The mice were sacrificed, and their livers were excised and washed with PBS. The fresh livers were cut into four smaller pieces and placed into a PBS/formaldehyde solution (final formaldehyde concentration = 1%). Crosslinking was stopped by addition of glycine to a final concentration of 0.125 M and incubation for 5 min with rotation. The samples were centrifuged for 5 min at 700g at 4 °C to collect the liver pieces, and the supernatant was removed and washed once again with ice-cold PBS. Fresh PBS containing protease inhibitors (Roche) was added, and the tissue was disaggregated with a homogenizer Ultra Turrax T25 basic (Ika Werke, Staufen, Germany). The tissue was distributed into six tubes (2 ml each) and centrifuged for 5 min at 700g at 4 °C, and the supernatant was discarded. The homogenized tissue was resuspended and lysed with SDS lysis buffer containing protease
DNA constructs
Full-length cDNAs for human PPARα38 and human RXRα39 were subcloned into the T7/SV40 promoterdriven pSG5 expression vector (Stratagene). The same constructs were used for both T7 RNA polymerase-driven in vitro transcription/translation of the respective cDNAs and for viral promoter-driven overexpression in mammalian cells. Promoter regions of the ALAS1 gene and of the CPTI gene were cloned by PCR from human genomic DNA (primers listed in Table S4) and fused with the thymidine kinase promoter driving the firefly luciferase reporter gene. All constructs were verified by sequencing. Gel-shift assays In vitro-translated PPAR and RXR proteins were generated by coupled in vitro transcription/translation
Fig. 5. Functionality of PPREs in the ALAS1 gene. Gel-shift experiments were performed with the in vitro-translated PPARα, RXR alone or in combination, and in the presence of different 32P-labeled REs representing the seven candidate PPREs of the human ALAS1 gene and the human CPTI reference PPRE (a). Protein–DNA complexes were separated from free probe on non-denaturing 8% polyacrylamide gels. Representative gels are shown. PPAR–RXR heterodimer complex formation was quantified on an FLA-3000 reader relative to the reference PPRE. Numbers below the gels indicate the means of at least three independent gel-shift experiments. Standard deviations are shown in parentheses. NS indicates non-specific complexes. Chromatin was extracted from HepG2 cells that had been treated with 100 nM GW7647 for 0, 120 and 240 min, and IPs were performed with antibodies against PPARα, RXRα, PGC-1α and pPol II. Rabbit IgG was used as a specificity control. Real-time PCR was performed on reverse-cross-linked chromatin templates with primers specific to the TSS and six genomic regions containing the seven putative PPREs. Reporter gene assays were performed with extracts from HepG2 cells that were transiently transfected with luciferase reporter constructs containing the indicated REcontaining regions of the ALAS1 gene and with an expression vector for human PPARα (c). The established PPREcontaining region of the human CPTI gene served as positive control. Cells were treated for 16 h with either solvent (DMSO) or 100 nM GW7647. Relative luciferase activities are shown. Columns represent means of at least three experiments, and bars indicate standard deviations. Two-tailed Student's t-tests were performed to determine the significance of the specific antibody enrichment by PPAR agonist relative to control IgG (b) or effects mediated by PPAR overexpression and ligand induction relative to controls (c) (⁎p b 0.05, ⁎⁎p b 0.01, ⁎⁎⁎p b 0.001).
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Fig. 6 (legend on next page)
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5. Transfection and luciferase reporter gene assays
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HepG2 cells were seeded into six-well plates and grown overnight in phenol red-free DMEM supplemented with 10% charcoal-stripped FBS to 60% to 70% confluency. Polyethylenimine transfections were performed by incubating a reporter plasmid and the expression vector for human PPARα or empty pSG5 expression vector (1 μg each) with 10 μg of polyethylenimine (Sigma-Aldrich) in 100 μl of 150 mM NaCl for 15 min at room temperature. After dilution with 900 μl of phenol red-free DMEM, the mixture was added to the cells. Phenol red-free DMEM supplemented with 500 μl of 15% charcoal-stripped FBS, containing 100 mM GW7647 or DMSO, was added 8 h after transfection. The cells were lysed 16 h later using reporter gene lysis buffer (Roche). The constant light signal luciferase reporter gene assay was performed as recommended by the supplier (Perkin-Elmer, Groningen, The Netherlands). Luciferase activities were normalized with respect to protein concentration, and induction factors were calculated as the ratio of luciferase activity of ligand-stimulated cells to that of solvent controls.
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Acknowledgements
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using their respective pSG5-based full-length cDNA expression constructs and rabbit reticulocyte lysate as recommended by the supplier (Promega). Protein batches were quantified by test translations in the presence of 35Slabeled methionine. Gel-shift assays were performed as described previously.29 The sequences of the PPREs are summarized in Table S5.
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This work was supported by grants from the Academy of Finland, the Juselius Foundation and the European Union (Marie Curie RTN NucSys).
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Supplementary data
Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j. jmb.2009.03.024
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Fig. 6. Fasting-induced PPARα TSS association and heme synthesis in mouse liver. Male pure-bred Sv129 mice and PPARα null mice on an Sv129 background were fed or fasted for 24 h (three mice per group). The livers of sacrificed mice were excised and immediately cross-linked with formaldehyde. The chromatin of the livers was extracted, and IPs were performed with antibodies against PPARα (a), RXRα (b), PGC-1α (c) and pPol II (d). Rabbit IgG was used as a specificity control. Real-time PCR was performed on reverse-cross-linked chromatin templates with primers specific to genomic regions of the eight mouse hepatic heme biosynthesis genes. Columns indicate the fold enrichment compared with IgG control precipitations. Representative results from one mouse are shown. In the same set of fed or fasted mice, heme concentration (e) was determined as described in Fig. 2.
PPARα Controls Heme Biosynthesis
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