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Hypoxia-Inducible Factor 1-Mediated Inhibition of Peroxisome Proliferator-Activated Receptor Expression During Hypoxia¹

Center for Experimental Therapeutics and Reperfusion Injury, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA 02115

	Abstract

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Peroxisome proliferator-activated receptors (PPARs) are nuclear hormone-binding proteins that regulate transcriptional responses to peroxisome proliferators and structurally diverse fatty acids. PPARs have been implicated in a wide variety of functions, including lipid homeostasis and inflammatory responses. In this study, we examined the expression of PPAR- in response to ambient hypoxia. Initial studies using microarray analysis of intestinal epithelial mRNA revealed that hypoxia rapidly down-regulates PPAR- mRNA and protein in epithelial cells in vitro and in vivo. Subsequent studies revealed that the PPAR- gene bears a DNA consensus motif for the transcription factor hypoxia-inducible factor 1 (HIF-1). EMSA analysis revealed that ambient hypoxia induces HIF-1 binding to the HIF-1 consensus domain of PPAR- in parallel to HIF-1 nuclear accumulation, and antisense depletion of HIF-1 resulted in a loss of PPAR- down-regulation. The PPAR- ligand pirinixic acid (WY14643) functionally promoted IFN--induced ICAM-1 expression in normoxic epithelia, and this response was lost in cells pre-exposed to ambient hypoxia. Such results indicate that HIF-1-dependent down-regulation of PPAR- may provide an adaptive response to proinflammatory stimuli during cellular hypoxia. These studies provide unique insight into the regulation of PPAR- expression and, importantly, provide an example of a down-regulatory pathway mediated by HIF-1.

	Introduction

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The peroxisome proliferator-activated receptors (PPARs)³ are a family nuclear hormone-binding proteins with increasingly diverse functions as transcriptional regulators (1, 2). Three subtypes of PPARs have been described (, (), and ), each of which is encoded by a different gene, has unique tissue distribution patterns, and each has ligand-selective binding (1, 2). Recent studies have identified a role for PPARs in the control of inflammation. For example, PPAR- ligands were recently shown to inhibit NF-B activation and to diminish colonic inflammation in a mouse model of inflammatory bowel disease (3). Similarly, ligands which bind the PPAR- isoform may amplify or inhibit the expression of inflammation-related gene products such as cyclooxygenase 2 and IL-6 (4, 5, 6) and may regulate the duration of inflammatory responses through feedback pathways involving leukotriene B₄ (7).

In many disease states, hypoxia and inflammation occur coincidentally (8). A number of studies have indicated that tissue hypoxia associated with ongoing disease processes may amplify proinflammatory signals and, paradoxically, may significantly contribute to the resolution of ongoing inflammation (8). For this reason, it is imperative to understand how induction pathways of inflammation and hypoxia overlap. In addition, recent studies indicate that hypoxia can directly activate transcription. This latter response is exemplified by discovery of hypoxia-inducible factor 1 (HIF-1), a member of the rapidly growing Per-ARNT-Sim family of basic helix-loop-helix transcription factors (9, 10). Functional HIF-1 exists as an heterodimer, the activation of which is dependent upon stabilization of an O₂-dependent degradation domain of the subunit by the ubiquitin-proteasome pathway (11). Binding of HIF-1 to DNA consensus domains (5'-RCGTG-3') results in the transcriptional induction of HIF-1-bearing gene promoters (12). HIF-1 is widely expressed and recent studies indicate that consensus HIF-1-binding sequences exist in a number of genes (12). Transcriptional responses mediated by HIF-1 include those ascribed to a hypoxia-adaptive response (e.g., erythropoietin, vascular endothelial growth factor, glycolytic enzymes, etc.) (13). Less is known as to whether HIF-1 may function to directly regulate inflammatory events or whether HIF-1 may mediate negative (down-regulatory) pathways.

In the present studies, we identified a hypoxia-elicited down-regulation of PPAR- in intestinal epithelial cells. In parallel, these results identified a previously unappreciated binding site for HIF-1 on the antisense strand of the PPAR- gene. Down-regulation of PPAR- by hypoxia correlated with HIF-1 induction and functioned to protect epithelia from PPAR- agonist amplification of ICAM-1 induction. These studies are the first to define a down-regulatory pathway mediated by HIF-1 binding.

	Materials and Methods

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Cell culture

T84 intestinal epithelial cells (passages 67–85) were grown and maintained as confluent monolayers on collagen-coated permeable supports as previously described in detail (14). T84 monolayers were maintained on 0.33-cm² or 5-cm² ring-supported polycarbonate filters (Costar, Cambridge, MA), unless otherwise noted, and used 6–12 days after plating. The oral epithelial line (KB cells) were grown as described previously (15), and human microvascular endothelial cells were plated and cultured as described previously (16).

Monolayer exposure to hypoxia was performed as previously described (17). Growth medium was replaced with fresh, pre-equilibrated hypoxic medium and cells were placed in a humidified environment within the hypoxia chamber (Coy Laboratory Products, Ann Arbor, MI) and maintained at 37°C. Oxygen concentrations were maintained as indicated with the balance made up of nitrogen, 5% carbon dioxide, and water vapor. Normoxic controls were cells exposed to the same experimental protocols under conditions of atmospheric oxygen concentrations (21% O₂/5% CO₂ within a tissue culture incubator). Lactate dehydrogenase release into soluble supernatants was measured colorimetrically (Cytotox 96 Non-Radioactive Cytotoxicity Assay; Promega) according to the manufacturer’s protocol. Total cellular lactate dehydrogenase was determined from samples lysed in 0.5% Triton X-100.

Transcriptional analysis

The transcriptional profile of epithelial cells exposed to ambient hypoxia was assessed in RNA derived from control or hypoxic epithelia (T84 cells at 6 or 18 h hypoxia) using quantitative genechip expression arrays (Affymetrix, Santa Clara, CA) (18). RT-PCR analysis of mRNA levels was performed using DNase-treated total RNA as previously described (19). Briefly, single-stranded cDNA was synthesized from 1 µg of RNA (DNA Polymerase High Fidelity PCR System; Life Technologies, Grand Island, NY). The PCR for human PPAR- contained 1 µM each of the sense primer (5'-TCA TCA AGA AGA CGG AGT CG-3') and the antisense primer (5'-CGG TTA CCT ACA GCT CAG AC-3') in a total volume of 50 µl, resulting in a 211-bp fragment, and for PPAR- (sense, 5'-AGA CAA CAG ACA AAT CAC CAT-3' and antisense, 5'-AAG TTT GAG TTT GCT GTG AAG-3'), resulting in a 401-bp fragment. PCR were then visualized on a 1% agarose gel containing 5 µg/ml ethidium bromide. Human and mouse -actin expression was examined in identical conditions as an internal control (sense primer, 5'-TGA CGG GGT CAC CCA CAC TGT GCC CAT CTA-3' and antisense primer, 5'-CTA GAA GCA TTT GCG GTG GAC GAT GGA GGG-3') revealing a 661-bp amplified fragment. Where indicated, HIF-1 mRNA was examined by RT-PCR (sense primer, 5'-CTC AAA GTC GGA CAG CCT CA-3' and antisense primer, 5'- CCC TGC AGT AGG TTT CTG CT -3'), revealing a 460-bp amplified fragment.

Generation of nuclear lysates

For analysis of nuclear extracts, confluent monolayers of T84 cells on 100-mm petri dishes were washed in ice-cold PBS, lysed by incubation in 500 µl of buffer A (10 mM HEPES (pH 8.0), 1.5 mM MgCl₂, 10 mM KCl, 0.5 mM DTT, 200 mM sucrose, 0.5 mM PMSF, 1 µg of both leupeptin and aprotinin per ml, and 0.5% Nonidet P-40) for 5 min at 4°C. The crude nuclei released by lysis were collected by microcentrifugation (15 s). Nuclei were rinsed once in buffer A and resuspended in 100 µl of buffer C (20 mM HEPES (pH 7.9), 1.5 mM MgCl₂, 420 mM NaCl, 0.2 mM EDTA, 0.5 mM PMSF, 1.0 mM DTT, and 1 µg/ml of both leupeptin and aprotinin). Nuclei were incubated on a rocking platform at 4°C for 30 min and clarified by microcentrifugation for 5 min. Proteins were measured (detergent-compatible-protein assay; Bio-Rad, Hercules, CA). Samples (25 µg/lane, as indicated) of T84 cell lysates were separated by nonreducing SDS-PAGE, transferred to nitrocellulose, and blocked overnight in blocking buffer (250 mM NaCl, 0.02% Tween 20, 5% goat serum, and 3% BSA). For Western blotting, anti-HIF-1 (rabbit antipeptide polyclonal directed to sequence MVNEFKLELVEKLFA encoding aa 527 through 541 of HIF-1) (20), anti-PPAR- (Research Diagnostics, Flanders, NJ), or anti--actin (Santa Cruz Biotechnology, Santa Cruz, CA) was added for 3 h, blots were washed, and species-matched peroxidase-conjugated secondary Ab was added exactly as described previously (27). Labeled bands from washed blots were detected by ECL. Resulting bands were quantified from scanned images using NIH Image software (Bethesda, MD).

EMSA

Nuclear extracts of cells exposed to indicated experimental conditions were obtained as described above. The following synthetic oligonucleotide probes were synthesized (Sigma-Genosys, The Woodlands, TX) and used as probes in EMSAs; the hypoxic response enhancer (HRE)-like motif (bold) on the antisense strand at positions 832–836 relative to the transcription start site in the PPAR- gene (sense, 5'-CTG CCA GTG CAC GTC AGT GGA G-3' and antisense, 5'-GAC GGT CAC GTG CAG TCA CCT C-3'). Oligonucleotide probes for EMSA were digoxigenin-labeled according to the manufacturer’s instructions (gel shift kit; Boehringer Mannheim, Indianapolis, IN). Labeled oligonucleotides were incubated with nuclear lysates for 10 min at 37°C and separated by electrophoresis on a 6% nondenaturing polyacrylamide gel. DNA-protein complexes were transblotted to nylon membranes, probed with antidigoxigenin-peroxidase, and developed by ECL. For supershift analysis, protein-DNA complexes were incubated with anti-HIF-1 mAb (Transduction Laboratories, Lexington, KY) for 1 h at 4°C before electrophoresis. Controls consisted of free probe alone, excess unlabeled probe, and isotype-matched control Ab (anti-cAMP response element binding protein (CREB)-2; Santa Cruz Biotechnology).

HIF-1 antisense oligonucleotide treatment of epithelia

HIF-1 depletion in epithelial cells was accomplished by using antisense oligonucleotide loading as described previously (21). Antisense oligonucleotide treatment of subconfluent epithelial cells was done as described previously (21), with modification. T84 epithelial cells were washed in serum-free medium and then in medium containing 20 µg/ml Geneporter transfection reagent (Gene Therapy Systems, San Diego, CA) with 2 µg/ml HIF-1 antisense or sense oligonucleotide. Cells were incubated for 4 h at 37°C, then replaced with serum containing growth medium. Treated cells were exposed to hypoxia or normoxia for indicated periods of time. As indicated, PPAR- or HIF-1 mRNA were quantified by RT-PCR as described above (see Transcriptional analysis).

ICAM-1 surface ELISA

IFN--induced ICAM-1 cell surface expression was quantified using a cell surface ELISA as described before (22). Epithelial cells were grown and assayed for Ab binding following exposure to normoxia or hypoxia (24 h) in the presence or absence of IFN- and addition of pirinixic acid (WY14643; Chemsyn Science Laboratories, Lenexa, KS) for an additional 48 h as indicated. Cells were washed with HBSS (Sigma, St. Louis, MO), and blocked with medium for 30 min at 4°C. Anti-ICAM-1 mAb (clone P2A4 (23) obtained from the Developmental Studies Hybridoma Bank, Iowa City, IA) was added for 2 h at 4°C. After washing with HBSS, a peroxidase-conjugated sheep anti-mouse secondary Ab (Cappel, West Chester, PA) was added. Secondary Ab (1:1000 final dilution) was diluted in medium containing 10% FBS. After washing, plates were developed by addition of peroxidase substrate (2,2'-azinobis(3-ethylbenzthiazoline-6-sulfonic acid), 1 mM final concentration; Sigma) and read on a microtiter plate spectrophotometer at 405 nm (Molecular Devices, Menlo Park, CA). Controls consisted of medium only and secondary Ab only. Data are presented as the mean ± SEM OD at 405 nm (background subtracted).

Mouse hypoxia model in vivo

Six- to 8-wk-old wild-type Bl6/129 (Taconic Farms, Germantown, NY) were exposed to hypoxia (constant flow of 8% O₂, 92% N₂ in a sealed modular chamber) or ambient room air for 8 h (n = 3 per condition). At the end of the experiments, animals were immediately sacrificed and tissues were collected for mRNA or protein analysis. Human PPAR- PCR primers (both forward and reverse, corresponding to nts 1550–1759 of mouse RNA) were used for mRNA analysis, see Transcriptional analysis above). This protocol was in accordance with National Institutes of Health guidelines for use of live animals and was approved by the Institutional Animal Care and Use Committee at Brigham and Women’s Hospital and Harvard Medical School.

	Results

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Hypoxia rapidly down-regulates PPAR- mRNA

A transcriptional profiling approach was used to identify potential hypoxia-regulated gene expression in model epithelia (T84 cells). As shown in Fig. 1A, microarray analysis identified a time-dependent down-regulation of the PPAR- gene (2.1- and 8.9-fold decrease compared with control normoxia at 6- and 18-h hypoxia, respectively). Such analysis suggested this loss to be specific for PPAR-, since no apparent changes were evident with PPAR- expression (0.05-fold loss and 0.1-fold increase at 6- and 18-h hypoxia, respectively) and by microarray, PPAR- was not expressed in model epithelia (data not shown). As shown in Fig. 1B, RT-PCR analysis was used to verify these microarray results and revealed a time-dependent loss of PPAR-, but not PPAR-, mRNA expression in hypoxia. Similar results of PPAR-, but not PPAR-, loss were observed in an oral epithelial cell line (KB cells, 92 ± 7% loss of PPAR- mRNA at the 6-h period of hypoxia, n = 3, p < 0.01) and in endothelial cells (human dermal microvascular endothelia, 65 ± 10% loss of PPAR- mRNA at the 8-h period of hypoxia, n = 3, p < 0.05), suggesting that these observations are not specific for intestinal epithelia.

View larger version (36K): [in this window] [in a new window]   FIGURE 1. Hypoxia rapidly down-regulates PPAR- mRNA. Confluent epithelial monolayers were exposed to the indicated periods of ambient hypoxia (pO2, 20 torr) or normoxia (pO2,147 torr). A, Quantitative RNA microarray analysis of PPAR- and - in epithelial cells exposed to the indicated conditions. B, Total RNA was isolated and examined for indicated PPAR transcripts by RT-PCR. -Actin transcript was used as an internal standard. Data shown are representative of three separate experiments.

Loss of PPAR- is HIF-1 dependent

Prompted by these results, a search of the PPAR- gene sequence identified a previously unappreciated HIF-1 binding site at positions 832–836 (relative to the first methionine codon) on the antisense strand of PPAR- (DNA consensus motif 5'-ACGTG-3') (24). Consequently, we assessed the induction of HIF-1 and loss of PPAR- protein during subjection of epithelia to ambient hypoxia. First, we determined whether conditions of hypoxia induce HIF-1 in intestinal epithelia (Fig. 2A). Western blot analysis of nuclear lysates derived from hypoxic intestinal epithelia demonstrated detectable HIF-1 at 2 h and abundant HIF-1 by 4 h of hypoxia. -Actin was used as a loading control for these experiments. These data are consistent with previous studies suggesting that HIF-1 is expressed in most cell types during periods of hypoxia and localizes to the nucleus during hypoxia exposure (25). Parallel examination of PPAR- protein by Western blot revealed a time-dependent loss over the course of 12 h of exposure to hypoxia (by densitometry, 63 and 84% loss at 8- and 12-h hypoxia, respectively, compared with control). Exposure of normoxic epithelial cells to CoCl₂ (100 µM, 8 h), a well-recognized activator of HIF-1 (9), also resulted in a loss of PPAR- protein (72% decrease by densitometry), suggesting that other known activators of HIF-1 similarly diminish PPAR-. -Actin was used as a loading control for these experiments. Such data indicate a temporal induction of HIF-1 and loss of PPAR- protein during hypoxia.

View larger version (38K): [in this window] [in a new window]   FIGURE 2. Induction of HIF-1 and loss of PPAR- protein in hypoxia. Confluent T84 epithelial monolayers were exposed to the indicated conditions of normoxia, hypoxia, or the HIF-1 activator CoCl2 (100 µM for 8 h). Nuclear lysates were prepared and levels of HIF-1 or PPAR- were determined by Western blot analysis. -Actin was used as a loading control in each case. Data shown are representative of three separate experiments.

View larger version (48K): [in this window] [in a new window]   FIGURE 3. EMSA for PPAR- HRE. A, EMSA analysis was used to evaluate DNA-protein interactions using digoxigenin-labeled double-stranded oligonucleotide probes spanning positions +832 to +836 of the PPAR- exon containing the putative HRE on the antisense strand. B, Confluent epithelial monolayers were exposed to (left to right, as indicated) normoxia (0), 4-h hypoxia, 24-h hypoxia, or CoCl2 (100 µM, 4 h). Nuclear lysates were generated and mixed with labeled probe in the presence or absence of anti-HIF-1 mAb (+) or species-matched control (anti-CREB-2) or 100-fold excess unlabeled probe (XS cold). DNA-protein complexes were resolved on a 4% nondenaturing polyacrylamide gel followed by ECL. Data shown are representative of three separate experiments.

View larger version (33K): [in this window] [in a new window]   FIGURE 4. HIF-1 antisense diminishes the loss of PPAR-. Confluent T84 epithelial monolayers were loaded with sense (S) oligonucleotides, antisense (AS) oligonucleotides, or mock treated for 24 or 48 h, as indicated. A, HIF-1 mRNA levels were determined by RT-PCR. B, Oligonucleotide-treated cells were exposed to 24- or 48-h normoxia, and PPAR- mRNA levels were determined by RT-PCR. C, Mock-treated or oligonucleotide-treated cells were exposed to 24- or 48-h hypoxia, and PPAR mRNA levels were determined by RT-PCR. D, Mock-treated or sense/antisense oligonucleotide-treated cells were exposed to 24- or 48-h hypoxia, as indicated, and PPAR- protein levels were determined by Western blot analysis. -Actin was used as a loading control in each case. Data shown are representative of three experiments.

Functional loss of PPAR- in hypoxia

Given the findings described above, studies were undertaken to determine whether PPAR- was functional in epithelial cells and whether such functional responses were lost during hypoxia. Previous studies have suggested that the PPAR- ligands such as pirixinic acid (WY14643) may enhance proinflammatory gene expression in epithelial cells (4, 5). Thus, as a functional assay for PPAR- activation, we examined the induction of epithelial ICAM-1 by IFN- (22, 26) in the presence and absence of PPAR activators. As shown in Fig. 5, the PPAR- ligand pirinixic acid enhanced IFN--induced ICAM-1 expression in epithelial cells (Fig. 5, p < 0.025 by ANOVA). Consistent with previous studies (22, 26), ICAM-1 was not expressed to an appreciable amount in the absence of IFN-. Concentrations as low as 500 nM pirinixic acid significantly enhanced IFN--induced ICAM-1 induction (Fig. 5, p < 0.025). Similar experiments performed on epithelial cells pre-exposed to 24-h hypoxia revealed a loss of pirixinic acid influence (p = NS by ANOVA). Such findings indicate that PPAR- is functional in epithelial cells, and consistent with our biochemical evidence, hypoxia induces a loss of functional PPAR- in the setting of an inflammatory stimulus (IFN-).

View larger version (28K): [in this window] [in a new window]   FIGURE 5. Functional PPAR- in intestinal epithelia; loss with hypoxia. T84 epithelial cells were grown to confluence on permeable supports. Monolayers were preincubated with the indicated concentrations of pirinixic acid in the presence or absence of 1000 U/ml IFN- (as indicated). Monolayers were then exposed to normoxia ( ) or hypoxia () for 48 h, after which ICAM-1 expression was assessed by cell surface ELISA. Data are pooled from three separate experiments and presented as the mean ± SEM ICAM-1 expression.

Down-regulation of PPAR- in vivo

We next evaluated whether hypoxia elicits the loss of PPAR- in vivo. Similar to the human PPAR- gene HIF-1 binding site, the mouse PPAR- gene includes an identical HIF-1-binding motif on the antisense strand (DNA consensus 5'-ACGTG-3' located at positions 905–909 relative to the first methionine codon) as well as identical flanking regions around this site (27). Since PPAR- is expressed at high levels in mouse intestine (28) and our results using cultured intestinal epithelia (T84 cells) revealed a loss of PPAR-, we compared PPAR- mRNA and protein levels in intestinal tissue following mouse subjection to whole-body hypoxia (8% O₂, 92% N₂ for 8 h) or to ambient room air. A similar hypoxia model has been used to examine hypoxia-regulated gene products in a variety of tissues (29, 30, 31). As shown in Fig. 6, lumenal scrapings (enriched in epithelial cells) harvested from intestinal tissue derived from hypoxic mice revealed a significant decrease in both PPAR- mRNA (93 ± 17% decrease compared with normoxic control, n = 3 mice per condition, p < 0.01) and protein (71 ± 12% decrease compared with normoxic control, n = 3 per condition, p < 0.025). These data indicate the likelihood that similar HIF-1-mediated down-regulatory pathways exist in vivo.

View larger version (23K): [in this window] [in a new window]   FIGURE 6. Loss of PPAR- in vivo. Bl6/129 mice were subjected to hypoxia (constant flow of 8% O2, 92% N2 in a sealed modular chamber) or ambient room air for 8 h (n = 3 per condition). At the end of the experiments, animals were sacrificed and intestinal tissues (lumenal scrapings enriched in epithelial cells) were collected for mRNA or protein analysis. A, Human PPAR- and -actin PCR primers were used to assess mRNA levels in DNase-treated RNA. B, Protein lysates (25 µg/lane) were resolved by nonreduced SDS-PAGE and immunoblotted with anti-PPAR- or anti--actin Ab. Representative samples are from three mice in each condition.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

Previous studies have demonstrated that PPARs are multifunctional nuclear receptors which when activated by a variety of fatty acids and prostaglandins regulate gene transcription via localization to recognition sequences termed peroxisome proliferator response elements (PPREs) (2). It is unclear, however, how PPAR genes are transcriptionally regulated. In the present studies, several lines of evidence identify a HIF-1-mediated repressor activity for PPAR-. Initial insight was gained through a broad screen of epithelial genes by microarray analysis, a strategy that has proven beneficial for identifying hypoxia-regulated pathways in epithelial cells (32, 33). Subsequent studies identified a specificity for such responses (e.g., not evident for PPAR-), some degree of universality for this response (i.e., also apparent in oral epithelial cell line KB and in human microvascular endothelial cells), and the existence of a putative hypoxia response enhancer within the PPAR- gene. To date, defined HIF-1 regulatory pathways have been limited to induction of hypoxia-regulated genes (34), and we provide an example here of HIF-1-mediated repressor activity. Evidence for this HIF-1-mediated pathway includes 1) the existence of a consensus binding site for HIF-1, 2) recapitulation of this response with a known activator of HIF-1 (CoCl₂), 3) HIF-1 binding by EMSA, and 4) inhibition of this response in cells not expressing HIF-1 (antisense oligonucleotides).

It is not known how HIF-1 repressor activity manifests itself. It is possible that binding of the HIF-1 heterodimer to the antisense strand of some genes (e.g., PPAR-) is directional and, as such, results in a functional transcriptional repressor. This aspect of HIF-1 signaling has not been studied in detail. Alternatively, HIF-1 may associate with other hypoxia-modified transcriptional regulators to maintain PPAR- expression. For example, we have recently demonstrated that CREB is targeted for degradation under conditions of hypoxia (19, 33). Given that the human PPAR- gene sequence includes a previously unappreciated CREB binding site (DNA consensus 5'-TGACGGA-3' at positions 1309–1315 relative to the translation start site) (24) downstream from the HIF-1 binding site (nt positions 832–836) and HIF-1 DNA binding sites have been shown to interact with CREB protein (35, 36), a potential pathway could involve destabilization of PPAR- expression through the loss of CREB. Thus, details of PPAR- down-regulation by HIF-1 remain unclear and will require extensive additional studies.

HIF-1 is particularly interesting as a regulatory pathway for mucosal transcriptional responses. First, mucosal organs, such as the lung and intestine, support a rich underlying vasculature, and perturbations in blood flow can result in rapid and severe tissue hypoxia (8). Important in this regard, epithelial cells that line mucosal tissues are active participants in the inflammatory response, and hypoxia may contribute substantially to such ongoing inflammation (8). We demonstrate here that PPAR--selective ligands, such as pirixinic acid, enhance epithelial ICAM-1 induction, an inflammatory marker on intestinal epithelial cells in vitro and in vivo (22). These findings of enhanced ICAM-1 induction are consistent with one study in endothelial cells (37), although other evidence suggests that PPAR ligands may inhibit TNF-induced ICAM-1 expression (38). Our own analysis of the published 5' region of the ICAM-1 gene (39) identified a PPAR response element (PPRE)-like consensus sequence (2) (consensus AGGTCATGCATGCTTAGGT positioned 197–215 nts downstream of the TATA signal), providing at least the possibility that the ICAM-1 gene directly binds PPAR. Detailed studies are necessary to further define the identity of PPRE within the ICAM-1 gene. Alternatively, it is possible that PPAR- interacts with other inflammatory pathways (e.g., NF-B, mitogen-activated protein kinases) and that PPAR- activity on ICAM-1 is indirect, particularly since the activity of the PPAR- ligand required additional inflammatory stimuli (i.e., IFN-). Nonetheless, our data that PPAR- ligand-mediated induction of proinflammatory signals (e.g., ICAM-1) are consistent with previous observations that PPAR- ligands enhance cyclooxygenase 2 in colonic and corneal epithelial cells (4, 5), and such observations may indicate that PPAR- provides a proinflammatory signal to epithelial cells. Of note, the isoform of PPAR, which was not regulated by hypoxia (see Fig. 1), has recently been targeted as an anti-inflammatory therapeutic pathway for mucosal disorders such as Crohn’s disease (3). Second, it is possible that HIF-1-mediated PPAR- expression could regulate epithelial cancers, such as colon cancer. For instance, recent studies have indicated that induction of cyclooxygenase 2 may be rate limiting to the development of colon cancer (40, 41, 42), and that cyclooxygenase inhibitors may afford some protection in this regard (43). Along these same lines, a mounting body of evidence has demonstrated that HIF-1-mediated genes are crucial to the establishment and maintenance of solid tumors (44, 45). Our studies with PPAR- may provide additional insight into these pathways. Taken together, these results of HIF-1-mediated down-regulation of PPAR- may define a novel adaptive pathway to dampen mucosal inflammation and limit epithelial proliferation responses.

In summary, these results define a new pathway of PPAR- transcriptional regulation. Such results indicate that pathophysiologically relevant conditions (e.g., hypoxia) have the potential to directly impact PPAR signaling through regulation of the nuclear receptor. Moreover, these studies provide an example of a counterregulatory transcriptional pathway for HIF-1.

	Acknowledgments

 
We thank Kristin Synnestvedt for technical assistance.

	Footnotes

 
¹ This work was supported by National Institutes of Health Grant DK50189 and Project 3 of PO-1 DE13499 and by a grant from the Crohn’s and Colitis Foundation of America.

² Address correspondence and reprint requests to Dr. Sean P. Colgan, Center for Experimental Therapeutics and Reperfusion Injury, Brigham and Women’s Hospital, Thorn Building, Room 704, 20 Shattuck Street, Boston, MA 02115. E-mail address: colgan{at}zeus.bwh.harvard.edu

³ Abbreviations used in this paper: PPAR, peroxisome proliferator-activated receptor; HIF-1, hypoxia-inducible factor 1; CREB, cAMP response element binding protein; HRE, hypoxic response enhancer; PPRE, peroxisome proliferator response element.

Received for publication November 20, 2000. Accepted for publication April 11, 2001.

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