The peroxisome proliferator-activated receptor alpha (PPARα) is a fatty acid-activated transcription factor that governs a variety of biological processes. Little is known about the role of PPARα in the small intestine. Since this organ is frequently exposed to high levels of PPARα ligands via the diet, we set out to characterize the function of PPARα in small intestine using functional genomics experiments and bioinformatics tools. PPARα was expressed at high levels in both human and murine small intestine. Detailed analyses showed that PPARα was expressed most highly in villus cells of proximal jejunum. Microarray analyses of total tissue samples revealed, that in addition to genes involved in fatty acid and triacylglycerol metabolism, transcription factors and enzymes connected to sterol and bile acid metabolism, including FXR and SREBP1, were specifically induced. In contrast, genes involved in cell cycle and differentiation, apoptosis, and host defense were repressed by PPARα activation. Additional analyses showed that intestinal PPARα-dependent gene regulation occurred in villus cells. Functional implications of array results were corroborated by morphometric data. The repression of genes involved in proliferation and apoptosis was accompanied by a 22% increase in villus height and a 34% increase in villus area of wild-type animals treated with WY14643. This is the first report providing a comprehensive overview of processes under control of PPARα in the small intestine. We show that PPARα is an important transcriptional regulator in small intestine, which may be of importance for the development of novel foods and therapies for obesity and inflammatory bowel diseases.
- peroxisome proliferator-activated receptor alpha
- gene expression
- crypt-villus axis
- lipid absorption
- proximal-distal axis
the peroxisome proliferator-activated receptor (PPAR)-α is a ligand-activated transcription factor with diverse functions and is activated by a variety of synthetic compounds, including the lipid lowering fibrate drugs (35, 37). High-affinity natural ligands include eicosanoids, unsaturated as well as long-chain fatty acids, and their activated derivatives (acyl-CoA esters) (19, 25, 31, 32). In analogy with other nuclear receptors, PPARα forms obligate heterodimers with the retinoid X receptor and stimulates gene expression by binding to peroxisome proliferator response elements (PPRE) located in the regulatory domain of genes (37). PPARα is expressed in a variety of tissues including the small intestine (6, 16); however, its function has been almost exclusively studied in liver. In liver PPARα is critical for the coordinate transcriptional activation of genes involved in lipid catabolism, including cellular fatty acid uptake and activation, mitochondrial β-oxidation, peroxisomal fatty acid oxidation, ketone body synthesis, fatty acid elongation and desaturation, and apolipoprotein synthesis (35, 37). In addition, PPARα is an important regulator of the hepatic acute phase response. While the function of PPARα in liver is well studied, little is known about PPARα and PPARα target genes in nonhepatic tissues. This is especially true with respect to the role of PPARα in the small intestine, which has only been addressed in few studies (39, 40). Knowledge of the regulatory and physiological function of PPARα in the small intestine is of particular interest, since the average Western diet contains a high amount of triacylglycerols (5) that are hydrolyzed to monoacylglycerol and free fatty acids before entering the enterocyte (46). Consequently the small intestine is frequently exposed to high levels of PPARα ligands.
Therefore we set out to determine the role of PPARα in the small intestine. We first analyzed in detail the expression of PPARα throughout the small intestine and then evaluated the outcome of specific PPARα activation on small intestinal gene expression using microarrays and bioinformatics tools. This allowed the genome-wide identification of intestinal PPARα target genes and corresponding processes. We conclude that PPARα plays an important role in the regulation of intestinal function by governing diverse processes ranging from numerous metabolic pathways to the control of apoptosis and cell cycle.
MATERIALS AND METHODS
Pure bred wild-type (129S1/SvImJ) and PPARα-null (129S4/SvJae) mice (34) were purchased from Jackson Laboratories (Bar Harbor, ME) and bred at the animal facility of Wageningen University.
Mice were housed in a light- and temperature-controlled facility and had free access to water and standard laboratory chow (RMH-B; Hope Farms, Woerden, the Netherlands). All animal studies were approved by the Local Committee for Care and Use of Laboratory Animals.
Experimental design and tissue handling.
Four independent studies were performed. In all studies 4- to 5-mo old male wild-type and PPARα-null mice were used. Study A: Mice were fed chow or chow supplemented with 0.1% WY14643 (Chemsyn, Lenexa, KS) for 5 days (n = 6 mice per group). On the sixth day, mice were anaesthetized with a mixture of isoflurane (1.5%), nitrous oxide (70%), and oxygen (30%). Small intestines were excised and flushed with ice-cold PBS, and all subsequent tissue handling was performed on ice. Remaining fat and pancreatic tissue was carefully removed, and RNA was isolated from the complete full-length small intestine for microarray analysis. Study B: The above described experiment was repeated (n = 3 mice per group), except that after removal the small intestine was divided into 10 equal parts to study gene expression along the proximal-distal axis. Study C: Study A was repeated (n = 3–4 mice per group), except that after removal, the small intestine was inverted on a 0.75-mm-diameter rod, washed in ice-cold PBS, and divided into segments of 1 cm. Before continuing with the cell isolation protocol, we pooled segments of all animals within each experimental group. Fractions enriched in crypt or villus cells were isolated as described by Flint et al. (18). This isolation protocol was repeated 1 week later for the control group. Cell fractions were used for RNA isolation. Study D: The feeding experiment was repeated as described, except that in addition to WY14643 mice were fed chow supplemented with fenofibrate (0.1% wt/wt; Sigma, St. Louis, MO) for 5 days (n = 5 mice per group). RNA was isolated from the complete full-length small intestine for quantitative reverse-transcription polymerase chain reaction (qRT-PCR) analysis.
RNA isolation and quality control.
Total RNA was isolated from small intestinal samples using TRIzol reagent (Invitrogen, Breda, the Netherlands) according to the manufacturer's instructions. RNA was treated with DNase and purified using the SV total RNA isolation system (Promega, Leiden, the Netherlands). Concentrations and purity of RNA samples were determined on a NanoDrop ND-1000 spectrophotometer (Isogen, Maarssen, the Netherlands). RNA integrity was checked on an Agilent 2100 bioanalyzer (Agilent Technologies, Amsterdam, the Netherlands) with 6000 Nano Chips according to the manufacturer's instructions. RNA was judged as suitable for array hybridization only if samples exhibited intact bands corresponding to the 18S and 28S ribosomal RNA subunits and displayed no chromosomal peaks or RNA degradation products. Total RNA from human tissues (FirstChoice Human Total RNA Survey Panel) was obtained from Ambion (Austin, TX). Each tissue pool comprises RNA from three or four donors.
Affymetrix GeneChip oligoarray hybridization and scanning.
For microarray analyses, we used RNA isolated from the full-length small intestine. RNA was hybridized on an Affymetrix GeneChip Mouse Genome 430A array. This array detects 22,626 transcripts that represent ∼13,700 known genes. For each experimental group, three biological replicated were hybridized; thus in total 12 arrays were used. Detailed methods for the labeling and subsequent hybridizations to the arrays are described in the eukaryotic section of the GeneChip Expression Analysis Technical Manual, Revision 3, from Affymetrix (Santa Clara, CA). Array data have been submitted to the Gene Expression Omnibus, accession number GSE5475.
Analyses and functional interpretation of microarray data.
Scans of the Affymetrix arrays were processed using packages from the Bioconductor project (20). Expression levels of probe sets were summarized using the GC content-corrected robust multichip average algorithm (GCRMA) (62), where after differentially expressed probe sets were identified using Limma (51). P values were corrected for multiple testing using a false discovery rate (FDR) method (55). Probe sets that satisfied the criterion of FDR <5% (q value <0.05) were considered to be significantly regulated. Of these, probe sets that were also >1.5-fold changed in wild-type mice upon WY14643 treatment but were not changed in treated PPARα-knockout mice were designated PPARα regulated. Three complementary methods were applied to relate changes in gene expression to functional changes. One method is based on overrepresentation of Gene Ontology (GO) terms (33). Another approach, gene set enrichment analysis (GSEA), takes into account the broader context in which gene products function, namely in physically interacting networks, such as biochemical, metabolic, or signal transduction routes (57). Both applied methods have the advantage that it is unbiased, because no gene selection step is used, and a score is computed based on all genes in a GO term or gene set. In addition, biological interaction networks among PPARα regulated genes were identified using Ingenuity Pathways Analysis (IPA) (Ingenuity Systems, Redwood City, CA). Detailed descriptions of the applied methods are available in the supplemental text (supplemental_1).1
Single-stranded complementary DNA (cDNA) was synthesized from 1 μg of total RNA using the reverse-transcription system from Promega (Leiden, the Netherlands) according to the supplier's protocol. qRT-PCR was performed on a MyIQ thermal cycler (Bio-Rad, Veenendaal, the Netherlands) using Platinum Taq DNA polymerase (Invitrogen) and SYBR green (Molecular Probes, Leiden, the Netherlands). Most of the primer sequences were obtained from the PrimerBank at Harvard University (59). Primer sequences are listed in Table S1 of the supplemental data (supplemental_3). Samples were analyzed in duplicate and standardized to either cyclophilin or 18S expression. Expression levels in isolated villus cells were standardized to villin.
For histology studies a fifth, independent experiment was performed, exactly as described in study A. After removal, intestines were divided into three equal parts, which are referred to as duodenum, jejunum, and ileum, respectively. Each section was prepared using a “Swiss roll” technique (38) to evaluate the entire longitudinal section on one slide. Tissues were fixed by immersion in 4% PBS-buffered formaldehyde, processed in an automatic tissue processor, embedded in paraffin, sectioned at 5 μm, and stained with hematoxylin and eosin. Sections were examined on a CKX41 microscope (Olympus, Zoeterwoude, the Netherlands) equipped with calibrated DP software, version 3.2 (Olympus). This software was used to measure villus height, crypt depth, and villus area of 50 villi per section for each animal. Statistical analysis between groups was performed with ANOVA, followed by the least significant difference post hoc test.
PPARα expression in human and murine small intestine.
To ascertain whether PPARα may be functionally relevant in the small intestine, the expression of PPARα was measured in 20 human tissues by qRT-PCR. In humans the highest expression levels of PPARα were observed in kidney, followed by heart, small intestine, and liver (Fig. 1A). In mice, the expression of PPARα was slightly higher in liver compared with small intestine of SV129 mice, whereas RXRα expression was comparable between both tissues (Fig. 1B). These data suggest that PPARα may be functionally relevant in the small intestine.
Next, the expression levels of PPARα and selected genes along the crypt-villus and proximal-distal axes of the small intestine were examined. As expected, mRNA levels of intestinal alkaline phosphatase (IAP) and villin, two markers for differentiated absorptive epithelial cells (17, 49) were maximal in fraction 1 (Fig. 2). Conversely, expression of pancreatic lipase-related protein-2 (PNLIPRP2), a marker for Paneth cells located at the base of villi (53), peaked in fractions 6–8. Importantly, expression of PPARα declined from villus to crypt cell-enriched fractions, which was mimicked by fatty acid transport protein 4 (FATP4) and CD36/FAT, two proteins known to be highly expressed in enterocytes and involved in fatty acid uptake (9, 52). These data demonstrate that PPARα is predominantly expressed in differentiated enterocytes and colocalizes with other genes involved in fatty acid metabolism.
A similar relationship was observed along the proximal-distal axis. PPARα expression gradually increased from the duodenum throughout the distal jejunum and then decreased in ileum (Fig. 3). The same pattern of expression was observed for FATP4. Expression of liver-type fatty acid binding protein (L-FABP) peaked more proximally, whereas intestine-type fatty acid binding protein (I-FABP) expression was highest in the distal jejunum. As expected, the expression of IAP (Fig. 3A) and apical sodium-dependent bile salt transporter (ASBT) (Fig. 3B) was restricted either to the duodenum or terminal ileum, respectively (43, 61). Combined, these data demonstrate that PPARα expression is highest in jejunal villus cells.
Function of intestinal PPARα as assessed by transcriptome analyses.
To study the function of PPARα in the small intestine, wild-type and PPARα-null mice were treated with the synthetic PPARα agonist WY14643, followed by analyses of changes in global gene expression using Affymetrix MOE430A arrays. Results on the number of significantly regulated genes are summarized in Fig. 4. A complete list of regulated genes is available in supplemental_2. Additional qRT-PCR analyses were performed for selected genes, which confirmed the array results (Fig. 8 and supplemental_3). Under control conditions, expression levels of only 21 genes out of the ∼13,700 genes analyzed were significantly different between wild-type and PPARα-null mice [fold change (FC) >1,5; FDR <0.05]. Of these 21 genes, 16 genes were expressed at lower levels and five genes at higher levels in PPARα-null mice (Fig. 5). Most of these genes are known to be involved in lipid metabolism, and several have been identified as direct PPARα target genes in other tissues (37). In wild-type mice, activation of PPARα resulted in differential expression of 1,138 genes, of which only two, BC018473 [hitchhiker, Entrez Gene (EG) ID: 193217] and Abcb1a (Mdr1a, EG ID: 18671) were also altered in the PPARα-null mice upon WY14643 treatment (Fig. 4). Thus, in total 1,136 genes were PPARα-dependently regulated in the small intestine; PPARα activation resulted in increased mRNA levels of 567 genes, whereas 569 were repressed. To gain insight into the underlying biological phenomena affected by PPARα activation, a scoring-based resampling method was applied to identify significantly overrepresented Gene Ontology (GO) classes (33). As input the >22,000 t-test P values from the probe set comparisons across the two diets in wild-type mice were used. Classes of genes that changed most significantly are listed in Table 1. With respect to the concept “biological process,” terms on this top list were mainly descriptors for fatty acid and lipid metabolism. Other overrepresented GO classes included descriptors for immune system, cell proliferation and differentiation, and programmed cell death (Table 1), suggesting that PPARα is involved in the regulation of these processes in the small intestine.
A parallel GSEA was used to focus on groups of genes that comprise specific biochemical, metabolic, or signal transduction routes (57). This method allows the identification of up- or downregulated processes (Table 2). However, due to overlap in the source databases, several functions are represented multiple times. The outcome of GSEA was similar to that of the GO-based analysis. Remarkable, almost all increased gene sets correspond to metabolic processes, including fatty acids catabolism, mitochondrial oxidative metabolism, and several pathways that feed intermediates into these processes. Other processes of interest that were upregulated include genes related to steroid and bile acid metabolism. The cellular responses represented by the downregulated gene sets were much more diverse and did not include metabolic pathways. Various pleiotropic signal transduction routes were suppressed, and the functional outcomes were summarized as acting on immune system, cell proliferation, migration and differentiation, and apoptosis. The functional outcomes of these transcriptome analyses are summarized in Fig. 6. The figure underscores the role of PPARα as an important transcriptional regulator in small intestine, governing diverse processes ranging from apoptosis and cell cycle, to the immune response and numerous metabolic pathways.
Many genes involved in fatty acid catabolism are known to be direct PPARα target genes (37). However, the mechanisms by which PPARα activation results in downregulation of numerous genes are less well understood. We therefore used IPA to search for biological interaction networks. As input all 1136 PPARα-dependently regulated genes were used. Of these genes, 588 were eligible for network analysis. For the remaining 548 genes molecular interaction information was lacking. IPA computed 83 networks, of which five scored equally best, as judged by a statistical likelihood approach (8). The five networks with a score of 39 (P < 10−39) were combined to form a composite network representing the underlying biology of PPARα activation in small intestine (Fig. 7). This complex comprised a network of 175 unique genes and their interactions. All genes were responsive to PPARα activation, and every interaction between the genes was supported by published information. In addition, a right-tailed Fisher's exact test identified 23 canonical pathways significantly affected by PPARα activation (P < 0.05) (data not shown). All of these were also identified with GSEA, and six canonical pathways linked to the composite network (Fig. 7). Moreover, we identified six nodes that were central in connecting many of the changed genes. As expected, PPARα linked directly to most of the induced genes. The downregulated genes of the merged network all linked to MYC, caspase 3 (CASP3), major histocompatibility class (MHC) 2 transactivator (MHC2TA), epidermal growth factor receptor (EGFR), or lymphocyte-specific protein tyrosine kinase (LCK), which themselves were also PPARα-dependently downregulated. No direct interaction between PPARα and the other five central nodes could be identified. Since the maximum network size was limited to 35 genes, all genes presented in the composite network were regulated by PPARα. Yet this does not imply that all genes connected to the central nodes were regulated. For example, IPA knowledge base linked PPARα and MYC to 109 and 459 genes, respectively (data not shown). Of these, 35 (32%) respectively 60 (13%) genes were significantly regulated (FC >1.5, FDR <0.05) in the small intestine. Taken together, the generation of biological interaction networks identified five genes that may play a central role in mediating the pleiotropic suppressive effects of PPARα activation.
PPARα-dependent gene regulation occurs in villus cells.
To confirm that PPARα-dependent gene regulation occurred in the differentiated enterocytes, we isolated fractions enriched in villus cells from wild-type and PPARα-null mice treated with WY14643. The expression for selected PPARα target genes identified in the array analysis was then determined by qRT-PCR (Fig. 8, A–C). For comparison, microarray data and qRT-PCR results of total tissue are presented as well. Expression of “classical” PPARα target genes, cytochrome P450, family 4, subfamily a, polypeptide 10 (CYP4A10); enoyl-coenzyme A, hydratase/3-hydroxyacyl coenzyme A dehydrogenase (EHHADH); 2–4-dienoyl-coenzyme A reductase 2, peroxisomal (DECR2); and angiopoietin-like 4 (ANGPTL4) (28, 29, 34, 37) was increased by WY14643 in total tissue and villus cells from wild-type but not PPARα null mice (Fig. 8A). Similar expression patterns were observed for four putative intestinal PPARα target genes aldo-keto reductase family 1, member B8 (AKR1B8); glutamate oxaloacetate transaminase 2, mitochondrial (GOT2); farnesoid X receptor; retinoid X receptor (FXR); and diacylglycerol O-acyltransferase 2 (DGAT2), which were specifically induced by WY14643 treatment in total tissue and isolated fractions (Fig. 8B). Expression of PPARα itself was increased by WY14643 in total tissue and isolated fractions, indicating autoregulation of PPARα gene expression. Besides PPARα itself we also determined the expression levels of five other nodes identified by network analysis. Expression of CASP3, MHC2TA, EGFR, MYC, and LCK was PPARα-dependently downregulated (Fig. 8C). Thus, for all genes analyzed we observed great similarity in regulation in total tissue between microarray and qRT-PCR, as well as between total tissue and isolated villus cells. This again demonstrates the robustness of our microarray analysis.
PPARα activation by fenofibrate.
To study the specificity of PPARα activation, we compared the effects of WY14643 with the effects of fenofibrate another PPARα agonist. qRT-PCR was used to analyze expression of a set of classical and putative PPARα target genes as well as central nodes, which also were studied in isolated villus cells (Fig. 9). The effects of fenofibrate were comparable to those of WY14643, as both agonists induced the expression of PPARα itself and (putative) PPARα target genes and reduced the expression of the central nodes MHC2TA and CASP3. However, at the same concentration fenofibrate was less potent in activating or repressing genes compared with WY14643. Thus, we show that activation of PPARα by two different ligands does not result in qualitative differences in expression of a specific set of genes.
Morphometric changes of the villus of mice treated with WY14643.
One of the outcomes of the transcriptome analyses was that PPARα activation suppressed cell proliferation and apoptosis and enhanced cell differentiation. We hypothesized that morphologically this should result in elongated villi. We therefore prepared paraffin sections of control and treated intestines from wild-type and PPARα-null mice and studied gross intestinal morphology and height of the villi (Fig. 10, Table 3). Under basal conditions no differences were observed with respect to form, structure, and morphometric parameters between wild-type and PPARα-null mice. In wild-type mice, treatment with WY14643 significantly increased the height of the villus, whereas no effect was observed in PPARα-null mice. Crypt depth was not affected by WY14643 treatment in both groups. These data indicate that PPARα activation specifically increases villus height but does not alter crypt depth. The morphometric assessments are in line with the functional outcomes of the transcriptome analyses.
In this study we set out to determine the role of PPARα in the small intestine using genomics tools. We find that PPARα is very well expressed in the small intestine in both mouse and human. PPARα expression is highest in villus cells and peaks in the proximal jejunum. Activation of PPARα results in altered expression of a large set of genes involved in a variety of pathways, including intestinal lipid handling, cell cycle, differentiation, apoptosis, and host defense. These data suggest an important role for PPARα in the regulation of gene expression in the small intestine.
Under control (fed) conditions we observed few changes in gene expression between wild-type and PPARα null mice. Only 21 genes were significantly altered, most of which are involved in lipid metabolism. These observations are in accordance with numerous studies showing that the effect of PPARα deletion becomes mainly noticeable under conditions of metabolic stress, and does not a priori imply that the physiological role of PPARα is limited to lipid metabolism (14, 36, 41, 45).
Our finding that PPARα is expressed highest in villus cells from the jejunum is supported by crude tissue distribution studies performed in rats (6, 16). Intestinal PPARα expression thus coincides with the main anatomical location where long-chain fatty acids are digested, taken up, and secreted into the body, suggesting an important regulatory function of PPARα in these processes. Indeed, activation of PPARα resulted in the specific induction of genes involved in fatty acid uptake, binding, transport, and catabolism, as well as genes involved in triacylglycerol and glycerolipid metabolism, in the small intestine of wild-type but not PPARα-null mice. Although PPARα is known to regulate fatty acid metabolism in other organs (35, 37), the link between PPARα and regulation of lipid handling in the intestine is new. Hence, at the level of the enterocyte, PPARα serves as a fatty acid sensor that is part of a feed-forward mechanism in which fatty acids stimulate their own catabolism, storage, transfer through the enterocyte, and secretion as triacylglycerols. PPARα thus tightly controls the intracellular levels of these potential toxic compounds.
Next to genes involved in fatty acid and triacylglycerol metabolism, genes coding for transcription factors and enzymes connected with steroid (sterol) and bile acid metabolism, including FXR, SHP, and SREBP1 are specifically induced. This demonstrates that cross talk between PPARα and other lipid-regulated transcription factors also occurs in small intestine, in addition to liver (10, 26), and is functionally in line with the well-established roles of bile acids in absorption of dietary lipids (46). Moreover, since it was recently shown that intestinal FXR target genes are also involved in enteroprotection and inhibition of bacterial overgrowth (27), our data suggest that activation of PPARα might also influence epithelial barrier function.
In addition to PPARα itself, network analyses identified five central genes that connected many of the repressed genes. These central genes (MYC, EGFR, CASP3, LCK, and MHC2TA) themselves were also PPARα-dependently downregulated. The MYC gene encodes a multifunctional nuclear phosphoprotein and plays a role in cell cycle progression, apoptosis, cellular transformation, and differentiation. The results of the microarray study show that MYC and related genes are repressed in a PPARα-dependent manner, including cyclin D1 (CCND1), CASP3, nuclear factor-κB (NFkB), signal transducer and activator of transcription 1 (STAT1), and EGFR via platelet-derived growth factor receptor-β (PDGFRB). Inasmuch as MYC is known to repress major histocompatibility complex, class I-B (HLA-B); PDGFRB; and N-myc downstream-regulated gene 1 (NDRG1) expression (21, 44, 60), the observed upregulation of these genes by PPARα is likely mediated through repression of MYC. Our data for the first time connect PPARα with regulation of cell proliferation, differentiation, and apoptosis in the small intestine. Our results indicate that an important consequence of intestinal PPARα activation is blocking cells in transition to the G1-S checkpoint of the cell cycle, resulting in reduced proliferation and increased differentiation of cells (58). We speculate that the specific downregulation in villus cells of CASP3, a key enzyme in the apoptotic cascade (15), points to inhibition of apoptosis, as has been reported for other cell types (22, 42, 48, 63). Since cell shedding is strongly associated with apoptosis, this in turn may result in reduced shedding of cells from the villus tips (7, 47). Functional support of the gene expression data is provided by morphometric data showing a significant 22% increase in villus height, which is accompanied by a 34% increase in villus area. These findings were not observed in PPARα-null mice treated with WY14643. Combined, our data demonstrate that PPARα represses cell growth and apoptosis and stimulates cell differentiation, resulting in an increased number of mature absorptive enterocytes. We believe that this may be an important adaptation mechanism of the small intestine aimed at adjusting lipid absorptive capacity to increased dietary fats (i.e., hydrolyzed TAGs).
Our study connects PPARα with the immune system of the small intestine. Activation of PPARα suppresses complement activation, antigen presentation, and B-cell receptor signaling. It is known that PPARα-null mice have abnormally prolonged hepatic responses to inflammatory stimuli (13). In vascular cells the expression of interleukin-6, vascular cell adhesion molecule, and cyclooxygenase-2 in response to cytokine activation can be inhibited by PPARα ligands (11). In these cells PPARα ligands may inhibit the functional expression of NF-κB, in part by augmenting the expression of inhibitor of NF-κB (IκBα) (12). However, inhibitory effects of PPARα in the intestine as well as on the innate immune system have not been reported before. Although the lymphoid tissue of the gut is the primary system for host defense, it is known that intestinal epithelial cell expresses MHC2 molecules and can function as antigen-presenting cells, thus being capable of regulating mucosal T-cell responses (23, 24, 56). Moreover, there is evidence that some of the complement proteins, such as complement C3, are synthesized in enterocytes (1, 2). Network analyses showed that the repression of the positive “master” regulator MHC2TA by PPARα is likely responsible for the suppression of MHC2 gene transcription, whereas suppressed B-cell receptor signaling is linked to decreased expression of LCK (54). Taken together, our data show that PPARα influences the immune and inflammatory response in the intestine and support the possibility that enterocytes are involved in a local response to injury/inflammation at the epithelial surface. A repression of the inflammatory response in the intestine by PPARα might be therapeutically valuable for patients with inflammatory bowel disease. Although several studies suggest a link between PPARγ and inflammatory bowel disease (3, 4, 30, 50), hardly anything is known about the effect of PPARα.
In summary, by using a combination of functional genomics experiments and current bioinformatics tools, we are the first to identify the pathways and processes under control of PPARα in the small intestine. Our data provide new insight into the role of PPARα in the small intestine and may be of particular importance for the development of fortified foods and for prevention and therapies for treating obesity and inflammatory bowel diseases.
This work was supported by the Dutch Ministry of Economic Affairs through the innovation-oriented research program on genomics IOP-IGE01016. Additional support was obtained from the Wageningen Centre for Food sciences.
The authors thank Rene Bakker and Bert Weijers for excellent assistance with animal experiments.
↵1 The online version of this article contains supplemental material.
Address for reprint requests and other correspondence: M. Müller, Nutrition, Metabolism and Genomics Group, Div. of Human Nutrition, Wageningen Univ., PO Box 8129, NL-6700EV Wageningen, the Netherlands (e-mail:)
Article published online before print. See web site for date of publication (http://physiolgenomics.physiology.org).
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