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Mol Endocrinol. Jan 2008; 22(1): 78–90.
Published online Sep 20, 2007. doi:  10.1210/me.2007-0298
PMCID: PMC2194635

Nuclear Receptor Hepatocyte Nuclear Factor 4α1 Competes with Oncoprotein c-Myc for Control of the p21/WAF1 Promoter


The dichotomy between cellular differentiation and proliferation is a fundamental aspect of both normal development and tumor progression; however, the molecular basis of this opposition is not well understood. To address this issue, we investigated the mechanism by which the nuclear receptor hepatocyte nuclear factor 4α1 (HNF4α1) regulates the expression of the human cyclin-dependent kinase inhibitor gene p21/WAF1 (CDKN1A). We found that HNF4α1, a transcription factor that plays a central role in differentiation in the liver, pancreas, and intestine, activates the expression of p21 primarily by interacting with promoter-bound Sp1 at both the proximal promoter region and at newly identified sites in a distal region (−2.4 kb). Although HNF4α1 also binds two additional regions containing putative HNF4α binding sites, HNF4α1 mutants deficient in DNA binding activate the p21 promoter to the same extent as wild-type HNF4α1, indicating that direct DNA binding by HNF4α1 is not necessary for p21 activation. We also observed an in vitro and in vivo interaction between HNF4α1 and c-Myc as well as a competition between these two transcription factors for interaction with promoter-bound Sp1 and regulation of p21. Finally, we show that c-Myc competes with HNF4α1 for control of apolipoprotein C3 (APOC3), a gene associated with the differentiated hepatic phenotype. These results suggest a general model by which a differentiation factor (HNF4α1) and a proliferation factor (c-Myc) may compete for control of genes involved in cell proliferation and differentiation.

A GENERALLY RECOGNIZED tenet of developmental biology is that differentiated cells are typically not undergoing rapid cell division. This is true not only during normal development but also during tumor progression. Although we currently have a fairly detailed understanding of the molecular mechanisms by which the individual processes of cellular differentiation and proliferation take place, the molecular mechanisms that promote differentiation over proliferation, or vice versa, are largely undefined.

A key player in both proliferation and differentiation is the cyclin-dependent kinase inhibitor gene p21/WAF1 (CDKN1A). p21 is a pleiotropic molecule that plays a role in several aspects of cell division and growth regulation, including mediation of cell cycle arrest, and the apoptotic response (1). p21 gene expression is also activated in several cell types during terminal differentiation (2,3,4,5,6,7,8,9,10). Regulation of the p21 gene is complex and involves a number of transcription factors that both positively and negatively regulate p21 expression. The tumor suppressor p53, ubiquitous factors Sp1, Sp3, activator protein 2 (AP2), and signal transducers and activators of transcription (STATs), tissue-specific factors CCAAT/enhancer binding protein-α (C/EBPα) and MyoD, and several nuclear receptors [retinoic acid (RAR), vitamin D (VDR), and androgen (AR) receptors] have all been shown to up-regulate p21 gene expression (10,11,12,13,14,15). In contrast, transcription factors associated with cell proliferation, such as the oncoprotein c-Myc, are known to down-regulate p21 gene expression (16,17).

Hepatocyte nuclear factor 4α (HNF4α), a member of the nuclear receptor superfamily of ligand-dependent transcription factors (NR2A1), has also recently been shown to activate the expression of the human p21 gene (18,19). HNF4α, found primarily in the liver, kidney, intestine/colon, and pancreas, is associated with cellular differentiation and maintenance of differentiated phenotypes. For example, through classical promoter analysis, HNF4α1 has been shown to regulate over 60 liver-specific genes via direct binding to specific DNA response elements as a homodimer. (There are multiple isoforms of HNF4α generated by alternative promoter usage and splicing; when a given isoform, such as HNF4α1, has been associated with a specific result, it is so noted.) Those HNF4α target genes include genes involved in xenobiotic and drug metabolism, glucose and lipid metabolism, nutrient transport, and blood maintenance (20). More recent studies using genome-wide expression profiling have shown that HNF4α also plays a role in the epithelial transformation of the liver by regulating the expression of cell adhesion genes (21) as well as a role in the mouse intestine and colon (22,23). Additional genome-wide analyses (ChIP-on-chip) have shown that HNF4α binds to the promoters of thousands of genes in primary human islet cells and hepatocytes, including the p21 gene (24,25). The importance of HNF4α as a master regulator of differentiation is underscored by its regulation of other key liver-enriched transcription factor genes (25,26) and its linkage to several human diseases including diabetes, hemophilia, and hepatitis (20,27).

In addition to its role in regulating genes responsible for differentiation and development, there is also an increasing amount of evidence suggesting that HNF4α might also play a role in regulating the cell cycle. Previous studies have shown that HNF4α1 expression is lower than normal or completely lost in hepatocellular carcinoma (HCC) cells (28,29). This loss of HNF4α1 expression may be a determinant in HCC progression because ectopic expression of HNF4α1 can cause tumors to revert to a less invasive, more differentiated, slow-growing phenotype (29). HNF4α1 expression has also been shown to reduce or inhibit proliferation of cells in culture (30,31,32). Furthermore, it has been reported that HNF4α1 overexpression can arrest cell cycle progression at the G1 phase (18). However, despite all of these links to the cell cycle, the mechanisms by which HNF4α1 regulates cell cycle progression and p21 expression have not been elucidated.

In contrast to HNF4α, oncoprotein c-Myc is associated primarily with cell proliferation. Not only is it expressed at high levels in early development and embryonic stem cells, but it is also frequently overexpressed in human tumors. Elevated levels of c-Myc are known to bring about continued cell-cycle progression (33,34) and cellular immortalization (35) as well as to block differentiation (36,37) and induce programmed cell death (apoptosis) (38). Indeed, overexpression of c-Myc can induce HCC in mice (39,40,41), whereas inhibition of c-Myc expression results in a loss of the transformed phenotype (42) and a rapid and sustained tumor regression and differentiation (43). c-Myc activates gene expression by heterodimerizing with Max (44,45) and binding E-boxes (CACGTG) in the promoter regions of target genes (46). In contrast, inhibition of transcription by c-Myc, such as on the p21 promoter, appears to be largely independent of Max (16,17,38,47,48). p21 has been identified as a c-Myc target by overexpression of c-Myc, which results in repression of p21 gene expression (17,48,49), and also by c-Myc knockdown using RNA interference (RNAi), which leads to increased p21 expression (50,51). Previous studies suggest multiple mechanisms of p21 repression by c-Myc, one of which includes a direct protein-protein interaction between c-Myc and Sp1/Sp3 in the proximal promoter region (−119 to +16 bp relative to the transcription start site, +1) (10,17).

Here, we show that HNF4α1 up regulates the expression of the human p21 gene and blocks cell proliferation in a p53-independent fashion. We also show that although HNF4α1 binds the p21 promoter in vivo at regions that contain putative HNF4α1 binding sites, most of the activation by HNF4α1 appears to be via interaction with Sp1 at sites both distal and proximal to +1. We also show for the first time that HNF4α1 interacts with c-Myc in vivo and in vitro and competes with it for occupancy of the Sp1 sites as well as for control of the p21 promoter. HNF4α1 and c-Myc also have opposing effects on a classical HNF4α target gene, apolipoprotein C3 (APOC3). Taken together, these observations suggest a mechanism that could partially explain the dichotomy between cellular differentiation and proliferation.


Ectopically Expressed HNF4α1 Increases Endogenous p21 mRNA and Protein Levels and Inhibits Cell Proliferation Independently of the p53 Pathway

Others previously reported that HNF4α1 activates human p21 promoter activity in a transient transfection assay (18). To determine whether HNF4α1 could stimulate the expression of the endogenous p21 gene, we used a Tet-on recombinant adenovirus system (Ad.HNF4α1) to overexpress wild-type (wt) HNF4α1 in a human colon cancer cell line, HCT116 (wt), that does not express endogenous HNF4α1. Immunoblot (IB) analysis showed that ectopically expressed HNF4α1 was first detected at 12 h after doxycycline induction followed by a time-dependent increase at 24 and 36 h (Fig. 1A1A,, top panel). Elevated levels of endogenous p21 protein were also observed at the 24- and 36-h time points (middle panel). This increase in p21 protein was accompanied by a small, but reproducible, increase in endogenous p21 mRNA expression at 20 h after doxycycline treatment (Fig. 1B1B).). Because p53 is also known to stimulate the expression of the p21 promoter and to be activated by adenovirus infection (52,53), we examined the levels of p53 protein in the Ad.HNF4α1-infected cells and found that p53 protein levels did not increase appreciably until 36 h (Fig. 1A1A,, lower panel). Furthermore, infection of the isogenic cell line HCT116 p53−/− with the Ad.HNF4α1 resulted in an increase in expression of the p21 mRNA at 20 and 24 h after doxycycline induction (Fig. 1C1C).). These results indicate that ectopically expressed HNF4α1 increases p21 gene expression in a p53-independent fashion. Consistent with this, using RNAi, we have observed a decrease in p21 gene expression when expression of the endogenous HNF4α gene is knocked down in the human hepatoblastoma/HCC cell line HepG2 (Yang, C., H. Liao, E. Bolotin, W. W. Hwang-Verslues, J. Evans, K. Ellrott, T. Jiang, and F. M. Sladek, manuscript in preparation) (see supplemental Fig. S1, published as supplemental data on The Endocrine Society’s Journals Online web site at http://mend.endojournals.org). This observation further supports the notion that HNF4α1 induces p21 expression.

Figure 1
Ectopic Expression of HNF4α1 Increases Endogenous Human p21 Gene Expression and Decreases Cell Proliferation Independently of p53

To determine whether HNF4α1 could also affect cell proliferation, HCT116 wt and p53−/− cells transiently transfected with an HNF4α1 expression vector were counted at 24 and 48 h after transfection. In both cell lines, the HNF4α1-transfected cells exhibited a slower growth rate than the mock-transfected cells (Fig. 11,, D and E), indicating that, in addition to increasing p21 gene expression, HNF4α1 also decreased cell proliferation in a p53-independent fashion. To confirm that the observed differences in cell counts were due to differences in proliferation, we performed bromodeoxyuridine (BrdU) ELISA in HCT116 wt cells grown under the same experimental conditions. Less BrdU incorporation was observed in the HNF4α1-transfected cells (Fig. 1F1F),), confirming that HNF4α1 expression decreases cell proliferation.

HNF4α1 Activates the p21 Promoter via Multiple Promoter Regions

To determine which portion of the p21 promoter is required for HNF4α1 activation, transient cotransfection assays with nine luciferase reporter constructs containing different portions of the human p21 promoter were performed in HEK293 cells (Fig. 22).). The results show that HNF4α1 activated a 2.4-kb fragment of the promoter (−2.4 kb construct) roughly 5-fold (Fig. 22).). Smaller fragments of the promoter (−1.6 kb, −1.3 kb, −100 bp, and −93 bp) were also activated by HNF4α1, although to a lesser degree (2- to 3-fold). Similar results were obtained using HepG2 cells (supplemental Fig. S2). Two of these fragments (−1.6 and −1.3 kb) contain potential HNF4α binding sites as determined by TRANSFAC (18) and HNF4 Motif Finder analysis (http://bioinfo.ucr.edu/~ebolotin/fuzzhtmlform.html). The potential HNF4α binding sites are 5′-ATTGGTTCAATGTCCAATT-3′ at −755 to −737, H4.117, and 5′-GAGGCAAAAGTCC-3′ at −980 to −968 (see supplemental Fig. S3 for entire sequence of the −2.4-kb promoter and location of transcription factor binding sites). However, these HNF4α binding sites did not appear to be required for HNF4α1 activation of the promoter. The minimal constructs (−100 and −93 bp) still exhibited a 2-fold activation by HNF4α1, which was similar to other constructs that retained the HNF4α binding sites (−1.6 and −1.3 kb). In contrast, much of the 5-fold activation of the −2.4-kb construct was lost when a distal region containing three putative Sp1 sites (as predicted by TRANSFAC, http://www.gene-regulation.com/pub/programs.html#match) was deleted (−2.15 and −2.0 kb).

Figure 2
HNF4α1 Activates Human p21 Promoter Activity

HNF4α1 Is Recruited to Regions of the p21 Promoter that Contain Sp1 Binding Sites

To further investigate which portions of the p21 promoter are important for HNF4α1 transactivation, we performed chromatin immunoprecipitation (ChIP) assays in HepG2 cells that express endogenous HNF4α1. Eight sets of PCR primers that yield 350-to 450-bp products spanning the full-length p21 promoter were designed (Fig. 3A3A;; see supplemental Table S1 and Fig. S3 for primer sequences and locations). The distal and proximal regions containing Sp1 sites were flanked by primer sets 1 and 2 and primer sets 7 and 8, respectively; primer sets 5 and 6 cover the two putative HNF4α binding sites. ChIP assays using an anti-Sp1 antibody confirmed Sp1 binding to both the distal (−2.5 to −2.1 kb) and the proximal (−200 to +44 bp) regions (Fig. 3B3B,, primer sets 1, 2, 7, and 8) but not to intervening regions (primer sets 3–6). Importantly, ChIP assays using the anti-HNF4α antibody also showed that endogenous HNF4α1 was associated with the distal and proximal Sp1 regions (Fig. 3C3C,, primer sets 1, 2, 7, and 8) as well as the regions containing the predicted HNF4α binding sites (primer sets 5 and 6). In contrast, there was no appreciable binding to the regions spanned by primer sets 3 and 4. These results suggest that HNF4α1 may be recruited to the p21 promoter via Sp1 as well as by binding directly to the promoter.

Figure 3
HNF4α1 Is Recruited to the p21 Promoter via Sp1

To further establish that HNF4α1 and Sp1 bind the same promoter regions, we performed ChIP assays using sequential IP (re-ChIP) with anti-HNF4α1 antibodies followed by anti-Sp1 antibodies and found that both the distal and proximal Sp1 regions were immunoprecipitated (Fig. 3C3C,, second ChIP, primer sets 1, 2 and 7, and 8, respectively). In contrast, the other promoter regions, including those with the putative HNF4α binding sites (primer sets 3–6), were not detected in the re-ChIP with the Sp1 antibodies. This result verifies that HNF4α1 and Sp1 are present on the same regions of the same p21 promoters. Because these regions do not contain any potential HNF4α1 binding sites and because others have shown previously that HNF4α1 can interact directly with Sp1 (54,55,56), we propose that Sp1 recruits HNF4α1 to the p21 promoter.

The detection of HNF4α binding at Sp1 sites suggested that direct DNA binding by HNF4α1 may not be required to activate the p21 promoter. To test this possibility, we repeated the transient cotransfection assay with the full-length p21 promoter (−2.4 kb) and two HNF4α1 mutants defective in DNA binding: HNF4α1.S78D, which introduces a negative charge in between the two zinc fingers (57), and HNF4α1.S304D, which introduces a negative charge in a charge clamp important for homodimerization (58) (Sun, K., K. Zhang, Y. Brelivet, D. Moras, and F. M. Sladek, manuscript in preparation). Importantly, both HNF4α1 mutants were able to activate the p21 promoter to the same extent as wt HNF4α1 (Fig. 3D3D).). In contrast, the DNA binding-defective mutants exhibited severely decreased activation of a heterologous HNF4α reporter construct that contains just four HNF4α binding sites and a TATA box (ApoB.-85.-47.E4.Luc, Fig. 3E3E).). To further assess whether HNF4α1 is directly recruited to the p21 promoter by Sp1, we used small interfering RNA (siRNA) to knock down Sp1 expression in HepG2 cells and performed an HNF4α ChIP assay. We found that when Sp1 expression was decreased, HNF4α1 recruitment to both the distal and proximal Sp1 regions (primer sets 1 and 8) was also decreased (Fig. 3F3F),), even though overall HNF4α1 protein levels remained constant (Fig. 3G3G).). In contrast, HNF4α1 recruitment to the region that contains HNF4α1 (but not Sp1) binding sites was not affected (Fig. 3F3F,, primer set 6). Taken together, these results support the notion that HNF4α1 activates the p21 promoter primarily via an interaction with Sp1, rather than by direct DNA binding.

Oncoprotein c-Myc Antagonizes HNF4α1-Mediated Activation of the p21 and Apolipoprotein C3 (ApoC3) Promoters

c-Myc promotes cell cycle progression by activating cell cycle-promoting genes and suppressing cell cycle/growth arrest genes, including p21 (59,60). It suppresses transcription of cell cycle arrest genes by two distinct mechanisms, one of which does not require DNA binding or interaction with Max but does require interaction with Sp1 (16,17). Because our results suggested that HNF4α1 is recruited to the p21 promoter via Sp1, we hypothesized that c-Myc might be able to antagonize HNF4α1 activation by competing with HNF4α1 for interaction with Sp1. To test this hypothesis, we first overexpressed c-Myc in HepG2 cells where endogenous HNF4α1 is expressed. We observed a significant suppression of the full-length p21 promoter (−2.4 kb) activity (Fig. 4A4Aa,a, lane 2 vs. lane 1). To determine whether the suppression observed in HepG2 cells was due to antagonism between c-Myc and HNF4α1, we then coexpressed HNF4α1 with increasing amounts of a c-Myc expression vector and determined the level of activation of the full-length p21 promoter in HEK293 cells, which do not express endogenous HNF4α1. As expected, HNF4α1 expression stimulated p21 promoter activity (Fig. 4Ab4Ab,, lane 2 vs. lane 1), whereas c-Myc expression repressed it (lane 3 vs. lane 1). Importantly, expression of c-Myc also blocked the induction of the p21 promoter by HNF4α1 (lane 4), although an increased HNF4α1/c-Myc ratio overcame the repression (lanes 5–8 vs. lane 3). To determine whether c-Myc could antagonize the ability of HNF4α1 to activate other promoters, we performed the competition experiment using the promoter of ApoC3 (APOC3), a classical HNF4α1 target gene (61). Just as with the p21 promoter, c-Myc inhibited the ability of HNF4α1 to activate the ApoC3 promoter (Fig. 4B4B),), suggesting that the competition between HNF4α1 and c-Myc may affect genes associated with differentiation, as well as proliferation.

Figure 4
c-Myc Antagonizes the Ability of HNF4α1 to Activate the p21 Promoter

To investigate the mechanism of the antagonistic effects of c-Myc and HNF4α1, we next asked whether c-Myc could interact with HNF4α1 in vivo. Co-IP assays using HepG2 cells (Fig. 5A5A,, top panel), which express endogenous c-Myc and HNF4α1, as well as HEK293 cells ectopically expressing Flag.c-Myc and HNF4α1 (Fig. 5A5A,, bottom panel), demonstrated that these two proteins do indeed interact in vivo. We also performed in vitro glutathione-S-transferase (GST) pull-down assays and verified that the interaction between c-Myc and HNF4α1 is a direct one and showed that c-Myc interacts with multiple regions of HNF4α1, including the DNA-binding domain (DBD) (Fig. 5B5B,, lane 2–4). Interestingly, c-Myc interacted well with the LBD/F, which contains the ligand-binding domain (LBD) and the 88-amino-acid C-terminal extension termed the F domain (lane 6) as well as the isolated F domain (lane 7). In contrast, it did not interact with the isolated LBD (lane 5). Because the LBD/F signal was stronger than that with the isolated F domain, this suggests that interplay between the LBD and the F domain may also be important for the c-Myc interaction.

Figure 5
c-Myc Interacts with HNF4α1 in Vivo and in Vitro and Is Associated with Sp1 and HNF4α Binding Sites in the p21 Promoter

We next verified that in the cell lines we were using, HCT116 wt and HepG2, ectopically expressed c-Myc was associated with the Sp1-binding regions of the p21 promoter (Fig. 55,, C and D, primer sets 1, 2 and 7, 8). Whereas c-Myc has been shown previously to bind the proximal Sp1 sites (17), this is the first report of its binding to the distal sites. Interestingly, in HepG2 cells, which contain endogenous HNF4α1, we also observed c-Myc binding to the region containing the two HNF4α binding sites (Fig. 5D5D,, primer sets 5 and 6). This binding was not observed in the HCT116 wt cells, which do not express endogenous HNF4α1 (Fig. 5C5C),), suggesting that HNF4α1 is required for c-Myc association with the promoter in regions 5 and 6. Moreover, the observation that HNF4α1 was associated only with the distal and proximal Sp1 regions in HCT116 wt cells and not the regions with the HNF4 sites further suggest that the dominant mechanism by which HNF4α1 activates p21 expression is via Sp1.

To demonstrate that c-Myc and HNF4α1 compete for regulation of the p21 promoter via the Sp1 sites, a competition ChIP assay was performed. We transiently expressed Flag.c-Myc in HepG2 cells and then immunoprecipitated endogenous HNF4α1 in the ChIP assay (Fig. 66).). If c-Myc competes HNF4α1 off the Sp1 sites of the p21 promoter, then we would expect to see a decrease in HNF4α1 binding in cells ectopically expressing Flag.c-Myc. A modest but reproducible competition between c-Myc and HNF4α1 was observed in the distal region of the promoter with both low (primer set 1) and high amounts of transfected c-Myc (asterisk, primer set 1). There was also a noticeable decrease in HNF4α1 binding to the proximal Sp1 region but only with the higher amount of c-Myc (primer set 8*). The discrepancy in the competition between the distal and proximal regions could be explained by the fact that there are a greater number of Sp1 sites in the proximal region compared with the distal region (six vs. three sites). Interestingly, in the presence of ectopically expressed c-Myc, there was an increase in the amount of HNF4α1 bound to the putative HNF4α binding regions (primer sets 5 and 6). The significance of this is not known but further supports the notion that c-Myc and HNF4α1 interact directly.

Figure 6
c-Myc Competes HNF4α Off the Distal and Proximal Sp1 Sites of the p21 Promoter


The results presented here demonstrate for the first time that ectopic expression of nuclear receptor HNF4α1 activates the human p21 (CDKN1A) promoter primarily through interaction with the ubiquitous transcription factor Sp1 and that the oncoprotein c-Myc competes with HNF4α1 for control of the p21 promoter. The mechanism of this competition appears to be multifaceted with Sp1 playing a central role (Fig. 77).). HNF4α1 bound to Sp1 sites in both the distal and proximal promoter regions, although the largest decrease in HNF4α1-mediated activation of the p21 promoter came when the distal Sp1 sites were deleted. Even though HNF4α1 was recruited to two of its own response elements, DNA binding-deficient mutants of HNF4α1 activated the p21 promoter to similar levels as the wt HNF4α1 (Fig. 3D3D).). These results, along with previous reports of direct physical and functional interaction with Sp1 on other HNF4α target genes (54,55,56,62,63), strongly suggest that HNF4α1 activates the p21 promoter by interacting with Sp1 at both the distal and proximal sites.

Figure 7
Proposed Mechanism Contributing to the Dichotomy between Differentiation and Proliferation

It has been previously reported that c-Myc inhibits the activation of the p21 promoter by multiple mechanisms, one of which also involves a direct protein-protein interaction with Sp1 in the proximal region (17). Here, we report for the first time that c-Myc is also tethered to Sp1 sites in the distal region (−2.4 kb). We also show that HNF4α1 and c-Myc interact in vivo and in vitro, both on the p21 promoter and in solution. We observed c-Myc bound to promoter regions containing HNF4α1 binding sites (but no canonical E boxes) but only under conditions in which HNF4α1 was also bound. In contrast, in the regions containing Sp1 sites, we observed less HNF4α1 on the promoter when c-Myc was present. Because the HNF4α1/c-Myc competition was observed on the proximal Sp1 sites only in the presence of higher amounts of c-Myc, we hypothesize that the larger number of Sp1 sites in the proximal region increases the opportunity for both HNF4α1 and c-Myc to bind Sp1 at the same time and thus makes the HNF4α1/c-Myc competition more difficult to observe at lower levels of c-Myc. Taken together, these findings suggest that there may be multiple mechanisms by which c-Myc blocks the HNF4α1-mediated activation of the p21 promoter: 1) it may directly compete off HNF4α1 from Sp1 bound to the p21 promoter; 2) it may sequester HNF4α1 in solution; and/or 3) it may bind to HNF4α1 on the promoter and block its ability to activate p21 expression. Additional studies will be required to determine the relative contributions of these different mechanisms.

In addition to competition for transcriptional control of a gene such as p21 that is involved in the regulation of proliferation, we also observed that c-Myc competes with HNF4α1 for control of the human APOC3, a gene associated with the differentiated phenotype. Similar to the p21 promoter, the ApoC3 promoter contains several HNF4α1 as well as multiple Sp1 sites but lacks canonical c-Myc binding sites. Because it has been shown previously that HNF4α1 and Sp1 synergistically activate the ApoC3 promoter (54,62,63), it is possible that c-Myc inhibits the transactivation ability of HNF4α1 on the ApoC3 promoter through a mechanism similar to that proposed here for the p21 promoter: competition for interaction with Sp1.

These observations raise the issue of whether other genes could also be affected by competition between HNF4α1 and c-Myc. ChIP-on-chip studies have shown that both HNF4α1 and c-Myc bind to many promoters in vivo that contain no identifiable binding sites for either factor (24,64,65,66). Because it is estimated that approximately 23% of human genes contain an Sp1 site in the −250- to +150-bp region alone (67), it is possible that HNF4α1 and c-Myc are associated with additional promoters through Sp1. These genes may be additional targets for competitive regulation by HNF4α1 and c-Myc. For example, the cell division cycle 25A gene (CDC25A, involved in promoting cell cycle progression) has been found in expression profiling studies to be up-regulated by c-Myc (http://www.myccancergene.org/index.asp) (68) and down-regulated by HNF4α1 in an RNAi knockdown study using HepG2 cells (Yang, C., H. Liao, E. Bolotin, W. W. Hwang-Verslues, J. Evans, K. Ellrott, T. Jiang, and F. M. Sladek, manuscript in preparation). The CDC25A promoter contains no canonical c-Myc binding sites and no clearly identifiable HNF4α1 binding sites; it does, however, contain several putative Sp1 sites (as determined by TRANSFAC analysis).

The competition described here between HNF4α1 and c-Myc via Sp1 for control of the expression of genes involved in both cellular proliferation and differentiation may have broader implications. Even though HNF4α1 is expressed in only a limited number of tissues, other nuclear receptors, expressed in other tissues, have also been found to bind and activate the p21 promoter (e.g. retinoic acid, vitamin D, and androgen receptors) (12,13,14,15) One of those receptors, the androgen receptor, has also been found to interact with Sp1 in vivo and to bind the proximal Sp1 sites on the p21 promoter (69). Because we have shown here that c-Myc interacts with the highly conserved DBD of HNF4α1, it might also interact with the DBD of other nuclear receptors. If c-Myc competes with these other nuclear receptors for control of gene expression, the competition described here between HNF4α1 and c-Myc could very well be part of a more general mechanism that underlies the dichotomy between differentiation and proliferation. Such a competition could also be relevant in determining the progression of carcinogenesis.



Full-length rat wt HNF4α1 (accession no. X57133) in pMT7 (pMT7.HNF4α1), the rat HNF4α1 DNA-binding mutants S78D and S304D, and GST.HNF4α fusion constructs have been previously described (57,70,71). The luciferase reporter constructs containing various portions of the human p21 promoter [−2.4 kb (p21P.FL), −2.15 kb (p21Δp53), −2.0 kb (p21PΔ400), −1.6 kb (p21PΔ800), −1.3 kb (p21PΔ1.1), −500 bp (p21PΔ1.9), −300 bp (p21PΔ2.1), −100 bp (p21PΔ2.3), and −93 bp (p21P93-S)] were generous gifts from Dr. Xiao-Fan Wang (Duke University Medical Center, Durham, NC) (72). The classical HNF4α reporter constructs ApoB.-85–47.E4.Luc and ApoC3.Luc have been previously described (71,73). Full-length mouse wt c-Myc cDNA fused on the N terminus to the Flag epitope in pCBS (Flag.c-Myc) and full-length human wt c-Myc cDNA in pRSET (pRSET.c-Myc) were kindly provided by Dr. Ernest Martinez (University of California, Riverside).

Cell Culture, Cell Extracts, and RNA Preparation

Human colorectal carcinoma cell lines HCT116 wt (a gift from Dr. Bert Vogelstein, Johns Hopkins University, Baltimore, MD), and HCT116 p53−/− (obtained from Dr. Xuan Liu, University of California, Riverside) were cultured in McCoy’s 5A growth medium supplemented with 10% Tet system-approved fetal bovine serum (BD Biosciences/Clontech, San Jose, CA) and antibiotics (100 U/ml penicillin and 100 μg/ml streptomycin). Human hepatoblastoma/HCC cell line HepG2 (HB-8065), human embryonic kidney cell line HEK293 (CRL-1573), and monkey kidney cell line COS-7 (CRL-1651) were purchased from American Type Culture Collection (Rockville, MD) and were cultured in DMEM growth medium supplemented with 10% fetal bovine serum, 1% nonessential amino acids, and antibiotics. All cells were grown at 37 C in 5% CO2. Whole-cell extracts were prepared by gentle scraping in a Nonidet P-40 (NP-40) lysis buffer [50 mm Tris-Cl (pH 8.0), 120 mm NaCl, 0.5% NP-40, 1 mm dithiothreitol, 2 μg/ml aprotinin, 2 μg/ml leupeptin] followed by centrifugation at 12,000 × g at 4 C. Nuclear extracts were prepared as previously described (74). RNA for RT-PCR analysis was extracted using TRIzol reagent as specified by the manufacturer (Invitrogen, Carlsbad, CA).

IB Analysis

IB analysis was performed after 10% SDS-PAGE as previously described (71) with overnight incubation of a 1:5000 dilution of an affinity-purified antibody to HNF4α1 (α-445, reacts with the very C terminus of human, rat, and mouse HNF4α1) (61), 0.5 μg/ml anti-p53 antibody (DO-1; Santa Cruz Biotechnology, Santa Cruz, CA), or a 1:200 dilution of anti-p21 antibody (ab7960; Abcam, Cambridge, MA) followed by a 1:5000 dilution of horseradish peroxidase (HRP)-conjugated goat anti-rabbit (GαR-HRP) or goat anti-mouse (GαM-HRP) antibodies (Jackson ImmunoResearch Laboratories, West Grove, PA). Signals were detected using enhanced chemical luminescence Western Blotting Detection Reagent Kit (GE Healthcare/Amersham, Piscataway, NJ). Protein concentration was determined by the Bradford assay (Bio-Rad, Hercules, CA) before loading and verified by Coomassie staining of the blot after transfer.

Cell Proliferation Assays

Cell proliferation was evaluated using cell counts and BrdU ELISA. HCT116 cells were plated in 12-well and 96-well plates at a density of 1 × 105 cells per well. At time 0 h (24 h after seeding), cells were transfected with or without pMT7.HNF4α1 using Lipofectamine 2000 (Invitrogen). The cells were then harvested and counted or subjected to BrdU incorporation at 24 and 48 h after transfection. BrdU ELISA was performed using BrdU Cell Proliferation Assay Kit (Calbiochem, San Diego, CA) following the manufacturer’s instructions.

Transient Transfection Assays

One day before transfection, HEK293, COS-7, or HepG2 cells were plated in six- or 12-well plates at a density of 0.5 × 106 or 2.4 × 105 cells per well, respectively. DNA mixtures containing pMT7.HNF4α1, Flag.c-Myc, CMV.β-gal, and reporter constructs were added using Lipofectamine 2000 as indicated. After 48 h, transfected cells were harvested, and luciferase and β-galactosidase (β-gal) activities were determined as previously described (75). Significant differences as determined by the Student’s t test (P < 0.05) are noted by asterisks.

Adenovirus Infection

Adenovirus expression constructs Adeno-X Tet-on (Ad.Tet-on) and rat Adeno-X-TRE/HNF4α1 (Ad.HNF4α1) were made using the Adeno-X Tet-on expression systems 2 kit (BD Biosciences/Clontech). The cDNA of rat wt HNF4α1 containing a vesicular stomatitis virus (VSV) tag at the C terminus was first subcloned from pCB6.HNF4α1.VSV (76) into the pDNR.CMV donor vector using EcoRI and XbaI and then transferred into the pLP.Adeno-X.TRE vector using the Cre-loxP recombination reaction. For adenovirus-mediated HNF4α1 expression, HCT116 cells were plated at a density such that more than 80% confluence was reached within 24 h. Cells were then infected with 35 multiplicity of infection (determined in HEK293 cells) of recombinant adenovirus (Ad.Tet-on and Ad.HNF4α1 at a 1:3 ratio) in a minimal amount of complete medium for 4 h after which time medium containing doxycycline to reach a final concentration of 5 μg/ml was added and the incubation continued until the time of harvest.


RT-PCR was performed using the Access RT-PCR System (Promega, Madison, WI). PCR was performed using 2 μl first strand cDNA and 27 cycles of amplification. Aliquots of each PCR were run on 2% agarose gels and visualized by ethidium bromide staining. Primers for RT-PCR were 5′-CGTACCACTGGCATCGTGAT-3′ (forward, + 480 nucleotides) and 5′-GTGTTGGCGTACAGGTCTTTG-3′ (reverse, + 951 nucleotides) for human β-actin (accession no. NM_001101) and 5′-GACACCACTGGAGGGTGACT-3′ (forward, + 262 nucleotides) and 5′-GGCGTTTGGAGTGGTAGAAA-3′ (reverse, + 560 nucleotides) for human p21 (accession no. NM_000389).

Co-IP Assay

For HepG2, 3 × 106 cells were seeded, and nuclear extracts (74) were prepared 48 h later for the co-IP. For HEK293, 3.75 × 106 cells were seeded in 150-mm plates 24 h before transfection with Flag.c-Myc and pMT7.HNF4α1 (12 μg each). Whole-cell lysates were prepared 30 h after transfection using the NP-40 lysis buffer. Seventy-five microliters of nuclear extract (HepG2 cells) or 1 ml of the crude whole-cell extract (HEK293 cells) was incubated with 5 μg anti-Myc (N262; Santa Cruz) for the HepG2 extracts, anti-Flag M2 (Sigma Chemical co., St. Louis, MO) for HEK293 extracts or control mouse IgG (Santa Cruz) antibodies at 4 C for 2 h. Then, 40 μl prewashed protein A beads (Pierce, Rockford, IL) were added to the mixture and incubated at 4 C overnight with gentle agitation. After extensive washing with a diluted NP-40 lysis buffer (0.1% NP-40), c-Myc-interacting proteins were eluted with SDS buffer and analyzed by IB analysis using the anti-HNF4α antibody (1:5000 dilution of α-445 followed by GαR-HRP or α-445 conjugated directly to HRP by the Peroxidase Labeling Kit from Roche Pharmaceuticals (Nutley, NJ).

GST Pull-Down Assay

In vitro protein-protein interaction assays were performed using GST and GST.HNF4α1 (FL, DBD, DBD/H, LBD, LBD/F, and F domains) fusion proteins as previously described (71,77). Two micrograms of GST protein were incubated at 4 C with 3 μl in vitro-translated 35S-labeled c-Myc (pRSET.c-Myc) protein produced using the rabbit reticulocyte lysate system (TNT; Promega). Incubation was performed for 4–8 h with gentle agitation. The beads were then extensively washed followed by elution with SDS buffer and detection of the labeled c-Myc by 10% SDS-PAGE followed by autoradiography.

ChIP Assay

One 150-mm plate of HepG2 or HCT116 cells (~80% confluent) was treated with 1% formaldehyde for 10 min at room temperature. Cross-linking was stopped by the addition of 0.125 m glycine (final concentration). Cells were harvested in cold PBS and lysed in ChIP sonication buffer [1% Triton X-100, 0.1% deoxycholate, 50 mm Tris-Cl (pH 8.0), 150 mm NaCl, 5 mm EDTA, 2 μg/ml aprotinin, 2 μg/ml leupeptin, 0.2 mm phenylmethylsulfonyl fluoride]. The DNA fragments were sonicated to an average size of 500 bp. IP were performed with anti-Sp1 (PEP2; Santa Cruz), anti-HNF4α (α-445), and anti-Flag (M2) and corresponding control (IgG) antibodies, and DNA-protein complexes were eluted in 1% SDS elution buffer (1% SDS, 0.1 m NaHCO3, 0.01 mg/ml herring sperm DNA). The cross-links were reversed by heating at 65 C overnight, proteins were digested by proteinase K (0.17 μg/μl; New England Biolabs, Ipswich, MA), and the DNA was extracted with phenol-chloroform, precipitated with ethanol, and dissolved in 100 μl Tris-EDTA buffer [10 mm Tris-Cl (pH 8.0), 1 mm EDTA]. PCR amplification (32–35 cycles) was performed with 2 μl template DNA and primers spanning the promoter regions of the human p21 gene (see supplemental Table S1 and Fig. S3 for sequence and location of primers). The products were analyzed by agarose gel electrophoresis and visualized by ethidium bromide staining. For sequential ChIP analysis (re-ChIP), cross-linked protein-DNA complexes eluted with the 1% SDS elution buffer after the first IP (α-445) were incubated at room temperature for 30 min and centrifuged at 12,000 × g for 1 min, and the supernatant was diluted 1:100 in ChIP sonication buffer and used for the second IP (anti-Sp1) in a manner identical to that of the first IP. After the second IP, the cross-links were reversed and the DNA analyzed as described above.

Sp1 RNA Interference

HepG2 cells plated the day before in six-well (2 × 105 cells per well) or 150-mm plates (3.5 × 106 cells per plate) were transfected with Sp1 or control siRNA (0.4 nmol per well or 2.4 nmol per 150-mm plate) using Lipofectamine 2000. After 48 h, transfected cells were harvested for IB and ChIP analyses. The target sequence of the Sp1 siRNA was 5′-AAAGCGCUUCAUGAGGAGUGA-3′, corresponding to nucleotides 2174–2193 of the human Sp1 cDNA (NM_138473.2) (Dharmacon, Lafayette, CO). The control siRNA target sequence was 5′-ATGGAAGAGATCAATACCAAA-3′ (QIAGEN, Valencia, CA).

Supplementary Material

[Supplemental Data]


We thank Drs. X. Wang, B. Vogelstein, X. Liu, E. Martinez, F. Faiola, and M. Weiss for providing plasmids and cell lines, Dr. K. Sun for S78D and S304D mutants, and Ms. K. Chellappa for GST fusion proteins. We also thank Dr. A. Grosovsky for providing equipment and Drs. J. Bachant and E. Martinez for thoughtful discussion.


This work was supported by National Institutes of Health Grant R01DK053892 to F.M.S. W.W.H-V. was supported by a fellowship from the University of California Toxic Substances Research and Teaching Program and a Grant-in-Aid of Research from the National Academy of Sciences, administered by Sigma Xi, The Scientific Research Society.

Disclosure Statement: The authors have nothing to disclose.

First Published Online September 20, 2007

Abbreviations: ApoC3, Apolipoprotein C3; BrdU, bromodeoxyuridine; ChIP, chromatin immunoprecipitation; DBD, DNA-binding domain; β-gal, β-galoctosidase; GST, glutathione-S-transferase; HCC, hepatocellular carcinoma; HNF4α, hepatocyte nuclear factor 4α; HRP, horseradish peroxidase; IB, immunoblot; LBD, ligand-binding domain; NP-40, Nonidet P-40; RNAi, RNA interference; siRNA, small interfering RNA; VSV, vesicular stomatitis virus; wt, wild type.


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